ABSTRACT

ZENG, HAINIAN. Development-dependent Formation and Metabolism of and Proanthocyanidins in Acer Species. (Under the direction of David Danehower, William Hoffmann, Jenny Xiang and De-yu Xie).

Anthocyanins are one of the richest pigments, which belong to compounds in kingdom. They have many biological and ecological functions. Over the past many years, numerous efforts have been made to determine the biosynthetic pathway of anthocyanins and also to identify several regulatory proteins mainly in and of model and crop plants. However, many questions concerning the metabolism of anthocyanins in foliage remains unsolved. One example is “How can developmental processes impact on accumulation patterns of anthocyanins in ”. In this study, we choose several cultivars from one of the most popular ornamental plants Acer palmatum Thunb. to understand the mechanism of developmental changes of pigmentation in . Several other species were also analyzed. We propose that the metabolism of anthocyanins play an essential role in such changes. We use an integrated approach of phytochemistry and metabolic profiling to determine the biosynthesis and metabolism of anthocyanins and their impacts on foliage color. Proanthocyanidin analysis was carried out as well to determine their relationship to both production and foliar coloration. We have found that even for green leaves with no/trace amount of detectable anthocyanins, the biosynthetic pathway of /proanthocyanidin is still activated. Our results indicate that metabolic channeling directing the anthocyanidin pathway to the proanthocyanidin biosynthesis plays a very important role in pigmentation pattern change along developmental processes.

Development-dependent Formation and Metabolism of Anthocyanins and Proanthocyanidins in Acer Species

by Hainian Zeng

A thesis submitted to the Graduate Faculty of North Carolina State University in partial fulfillment of the requirements for the degree of Master of Science

Plant Biology

Raleigh, North Carolina

2009

APPROVED BY:

______Deyu Xie Jenny Xiang Committee Chair

______William Hoffmann David Danehower

DEDICATION

This thesis is dedicated to my dearest grandma, Yinchun Zhang, who passed away recently.

ii BIOGRAPHY

The first day I set my foot off the Airbus and onto the land of America, I knew it would be a

life change. However, the thing I didn’t realize back then, is that this change has been so

dramatic since that first day of air sickness and jet lag. From a metropolis to a countryside

city. From Shanghai, my beloved hometown for more than 20 years, to, Raleigh, the capital city of North Carolina, one of the biggest agricultural states in the United States. This is big.

Even one year later, sometimes I would still be wondering about where I was and why I was here. From time to time I was startled to see some many local people around and couldn’t help but mumbling to myself: I was in America.

How time flies! Now it’s been three years, such doubt doesn’t go away but become stronger.

In those three years, ups and downs are the main melody. Sometime I felt determined and I could see the bright future just ahead of me. Sometimes I felt lost and lonely, alienated myself from the outside due to serious depression. Sometimes I got sick and feared about the medical bill showing hundreds of dollars balance. Sometimes I demanded myself to be a man when facing my grandma’s past. Yes, this is life, but is it? At last, I find out what I want the most. To be with my family and friends. To fight against injustice and evil. To live a life whenever and wherever I can be myself, whenever and wherever I am in charge of my own life. At least, I won’t regret it. This is the whole point of being alive, isn’t it? Not to regret!

Thanks to these three years’ life experience, I have re-founded my goal and grasped the hang of how to lead a life. To be the king of my own “land”!

iii ACKNOWLEDGEMENTS

I want to give my sincere appreciation and to my major advisor, Dr. Deyu Xie. He has been a wonderful mentor and advisor for these three years. Additionally, he takes care of his graduate students like his own children and treats us with respect as friends. To me, it’s not overrated to say that he is a father-like idol. The only regret I can’t let it go is that I haven’t live up to Dr. Xie’s expectations mainly due to my own weaknesses and mistakes. I’d like to give my deep gratitude to my other committee members, Drs. Bill Hoffmann, Jenny Xiang and David Danehower. They kindly gave me a lot of instructions and suggestions on my project, as well as their generous help with my life. I can still recall the time that Dr.

Hoffmann gave me a ride back to my dorm, and Thanksgiving holidays I was spending in Dr.

Xiang’s house as well as the knowledge of phytochemistry passed on to me by Dr.

Danehower. Here I want to show my genuine gratefulness to all of my advisors for their understanding, forgiveness and noble gestures. I may be far away from being a good graduate student, but they have been continuously working with me as great advisors. And again, I’d like to apologize for my immature behaviors from time to time. I hope I can obtain your forgiveness some day. Besides my advisors, I’d like to thank our lab manager, Dr. Lili Zhou for her help with my research and her care about me. Dr. Zhou is a “mother” to us and I wish she can be with her family soon and lead a happy life ever after. Of course I would like to thank my labmates, Mingzhu, Fatima and Patrick. Without their inspiring ideas, contribution to my research and encouragements, I wouldn’t have gone this far. And last but not least, salutes to my colleagues in PBGSA for their unsinkable friendship as well.

iv TABLE OF CONTENTS

LIST OF TABLES ...... xi

LIST OF FIGURES ...... xii

CHAPTER 1 – LITERATURE REVIEW ...... 1

1.1 Plant ...... 3

Introduction ...... 3

Biosynthesis of flavonoids ...... 5

Ecological and physiological significances ...... 8

Nutraceutical significances ...... 10

Taxonomic significances ...... 10

1.2 Anthocyanins and ...... 12

Introduction ...... 12

Types of anthocyanins and anthocyanidins ...... 13

Properties of anthocyanins ...... 15

Biosynthesis of anthocyanins ...... 17

Ecological significances ...... 19

Nutraceutical significances ...... 24

1.3 Proanthocyanidins ...... 25

Introduction ...... 25

Types of proanthocyanidins...... 25

Biosynthesis of proanthocyanidins ...... 29

Ecological significances ...... 30

Nutraceutical significances ...... 31

v CHAPTER 2 - FOLIAR ANTHOCYANINS AND PROANTHOCYANIDINS IN SIX

ORNAMENTAL VARIETIES OF ACER PALMATUM ...... 32

2.1 Introduction ...... 32

2.2 Materials and methods ...... 39

2.2.1 Standard chemicals ...... 39 2.2.2 Plant materials ...... 39 2.2.3 Leaf collection ...... 42 2.2.4 Methods ...... 44 Extraction and measurement of anthocyanins ...... 44

Hydrolysis of anthocyanins, measurement of ABS and identification of anthocyanidins ...... 45

Extraction and analysis of proanthocyanidins ...... 46

LC-MS analysis ...... 48

Histological localization of proanthocyanidins ...... 49

Statistical analysis ...... 50

2.3 Results ...... 50

2.3.1 Analyses of anthocyanins and proanthocyanidins in leaves of Hubb’s Red Willow ...... 50 Difference of pigmentation in shaded and unshaded leaves ...... 50

Absorbance measurement of total anthocyanins ...... 51

Absorbance measurement of proanthocyanidins ...... 54

LC-MS analysis of anthocyanins and anthocyanidins ...... 55

LC-MS analysis of proanthocyanidins ...... 58

Histological staining with DMACA ...... 62

2.3.2 Analysis of anthocyanins and proanthocyanidins in leaves of Shaina ...... 66 Difference of pigmentation in shaded and unshaded leaves ...... 66

Absorbance measurement of total anthocyanins ...... 69

vi Absorbance measurement of proanthocyanidins ...... 69

LC-MS analysis of anthocyanins and anthocyanidins ...... 69

LC-MS analysis of proanthocyanidins ...... 72

Histological staining with DMACA ...... 76

2.3.3 Analysis of anthocyanins and proanthocyanidins in leaves of Hagoromo ...... 77 Red pigmentation and anthocyanin levels ...... 77

Absorbance measurement of proanthocyanidins ...... 78

LC-MS analysis of anthocyanins and anthocyanidins ...... 81

LC-MS analysis of proanthocyanidins ...... 83

Histological staining with DMACA ...... 87

2.3.4 Analysis of anthocyanins and proanthocyanidins in leaves of Dissectum Atropurpureum ...... 88 Property of foliar red pigmentation pattern ...... 88

Absorbance measurement of total anthocyanins ...... 89

Absorbance measurement of proanthocyanidins ...... 89

LC-MS analysis of anthocyanins and anthocyanidins ...... 89

LC-MS analysis of proanthocyanidins ...... 94

Histological staining with DMACA ...... 97

2.3.5 Analysis of anthocyanins and proanthocyanidins in leaves of Okushimo ...... 99

Green pigmentation ...... 99

Analysis of anthocyanins ...... 99

LC-MS analysis of proanthocyanidins ...... 103

Histological staining with DMACA ...... 106

2.3.6 Analysis of anthocyanins and proanthocyanidins in leaves of Oridono nishiki ...... 107

Green pigmentation ...... 107

vii Analysis of anthocyanins ...... 108

LC-MS analysis of proanthocyanidins ...... 112

Histological staining with DMACA ...... 114

2.4 Discussions ...... 116

2.4.1 Different pigmentation patterns in leaves of ornamental A. palmatum Thunb. cultivars are appropriate to investigate anthocyanin metabolism ...... 116 2.4.2 Anthocyanin level corresponds to leaf coloration pattern ...... 118 2.4.3 Development-dependent properties of anthocyanin pigmentation patterns in Acer cultivars ...... 119 2.4.4 The formation of proanthocyanidin leads to less red pigmentation ...... 122 2.4.5 Qualitative and quantitative changes contribute to total anthocyanin level change ...... 128 2.4.6 Levels of proanthocyanidins change in response to development ...... 132 2.4.7 Anthocyanin biosynthetic pathway is always activated ...... 136 2.4.8 Is proanthocyanidin formation a consequence of metabolic channeling resulting from genetic breeding? ...... 138 2.4.9 Metabolic profiling of anthocyanins and proanthocyanidins may help establish a key to A. palmatum Thunb. cultivars ...... 141 2.5 Conclusions and future prospects ...... 143

CHAPTER 3 – ANTHOCYANINS AND PROANTHOCYANIDINS METABOLISM

IN SPRING JUVENILE LEAVES OF SEVERAL SPECIES ..... 146

3.1 Introduction ...... 146

3.2 Materials and Methods ...... 147

3.2.1 Plant materials ...... 147 3.2.1 Methods ...... 149 3.3 Results ...... 150

3.3.1 Acer rubrum ...... 150 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 150

LC-MS analysis of anthocyanins and anthocyanidins ...... 153

viii 3.3.2 Acer x freemanii ”Celzam” ...... 155 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 155

LC-MS analysis of anthocyanins and anthocyanidins ...... 157

3.3.3 Acer longipes ...... 160 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 160

LC-MS analysis of anthocyanins and anthocyanidins ...... 163

3.3.4 Acer henryi ...... 166 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 166

LC-MS analysis of anthocyanins and anthocyanidins ...... 170

3.3.5 ...... 172 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 172

LC-MS analysis of anthocyanins and anthocyanidins ...... 175

3.3.6 Acer negundo"Kelly’s Gold" ...... 178 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 178

LC-MS analysis of anthocyanins and anthocyanidins ...... 179

3.3.7 Prunus padus ...... 182 Leaf pigmentation, anthocyanins and proanthocyanidins ...... 182

LC-MS analysis of anthocyanins and anthocyanidins ...... 186

3.4 Summary ...... 187

REFERENCES ...... 192

APPENDICES ...... 206

Appendix A...... 207

Appendix B...... 208

Appendix C...... 210

Appendix D...... 211

ix Appendix E...... 212

x LIST OF TABLES

Table 1. Naturally occurring anthocyanidins ...... 14

Table 2. A summary shows height, environment and location of 6 cultivars selected in this

work ...... 40

Table 3. List of cultivars selected in this work ...... 41

Table 4. Ratio of ABS values (ABS2/ABS1) of hydrolyzed samples ...... 54

Table 5. Height, and location of species selected in this work ...... 148

Table 6. List of species selected in this work ...... 149

Table 7. Estimation of anthocyanin content level ...... 189

Table 8. Estimation of proanthocyanidin content level ...... 190

Table 9. Ratio of ABS values (ABS2/ABS1) of hydrolyzed samples ...... 190

Table 10. Estimation of anthocyanin, proanthocyanidin content levels and ratio of ABS

values (ABS2/ABS1) of hydrolyzed samples from Prunus padus...... 191

xi LIST OF FIGURES

CHAPTER 1 – LITERATURE REVIEW

Fig 1. Backbone structures of some flavonoids ...... 4

Fig 2. Scheme of the flavonoid biosynthetic pathway ...... 7

Fig 3. General structures of anthocyanins and anthocyanidins ...... 13

Fig 4. Anthocyanins chemical forms depending on pH and degradation reaction for

anthocyanins ...... 16

Fig 5. Structures of different proanthocyanidins ...... 27

CHAPTER 2 - DEVELOPMENT-DEPENDENT FORMATION AND METABOLISM

OF ANTHOCYANINS IN ACER PALMATUM SPECIES

Fig 6. Summary of phylogenetic relationships for angiosperms ...... 38

Fig 7. Leaf phenotype of A. palmatum 'Hubb's Red Willow' and measurement of

anthocyanins and proanthocyanidins ...... 52

Fig 8. LC chromatogram of A. palmatum 'Hubb's Red Willow' ...... 56

Fig 9. Major anthocyanins of A. palmatum 'Hubb's Red Willow' ...... 58

Fig 10. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins

from leaves of A. palmatum 'Hubb's Red Willow' ...... 59

Fig 11. Major proanthocyanidins of A. palmatum 'Hubb's Red Willow' ...... 60

Fig 12. Trend of proanthocyanidins level change of A. palmatum 'Hubb's Red Willow' ...... 61

Fig 13. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Hubb's

Red Willow' ...... 62

Fig 14. Histological staining of proanthocyanidins in a mature leaf of A. palmatum 'Hubb's

Red Willow’ ...... 63

xii Fig 15. Microscopic images showing histological staining of proanthocyanidins in a mature green leaf of A. palmatum 'Hubb's Red Willow’ ...... 64

Fig 16. Microscopic images showing histological staining of proanthocyanidins in a young red leaf of A. palmatum 'Hubb's Red Willow’ ...... 65

Fig 17. Leaf phenotype of A. palmatum 'Shaina' and measurement of anthocyanins and proanthocyanidins ...... 67

Fig 18. LC chromatogram of A. palmatum 'Shaina' ...... 70

Fig 19. Major anthocyanins of A. palmatum 'Shaina' ...... 71

Fig 20. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Shaina' ...... 73

Fig 21. Major proanthocyanidins of A. palmatum 'Shaina' ...... 74

Fig 22. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Shaina'

...... 75

Fig 23. Trend of proanthocyanidins level change of A. palmatum 'Shaina' ...... 76

Fig 24. Histological staining of proanthocyanidins in a mature leaf of A. palmatum 'Shaina'

...... 77

Fig 25. Leaf phenotype of A. palmatum 'Hagoromo' and measurement of anthocyanins and proanthocyanidin ...... 79

Fig 26. LC chromatogram of A. palmatum 'Hagoromo' ...... 81

Fig 27. Major anthocyanins of A. palmatum 'Hagoromo' ...... 82

Fig 28. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Hagoromo' ...... 84

Fig 29. Major proanthocyanidins of A. palmatum 'Hagoromo' ...... 85

Fig 30. Trend of proanthocyanidins level change of A. palmatum 'Hagoromo' ...... 86

xiii Fig 31. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum

'Hagoromo' ...... 87

Fig 32. Histological staining of proanthocyanidins in a mature leaf of A. palmatum

'Hagoromo' ...... 88

Fig 33. Leaf phenotype of A. palmatum Dissectum Atropurpureum and measurement of anthocyanins and proanthocyanidins ...... 90

Fig 34. LC chromatogram of A. palmatum Dissectum Atropurpureu ...... 92

Fig 35. Major anthocyanins of A. palmatum Dissectum Atropurpureum ...... 93

Fig 36. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of A. palmatum Dissectum Atropurpureum ...... 94

Fig 37. Major proanthocyanidins of A. palmatum Dissectum Atropurpureum ...... 95

Fig 38. Trend of proanthocyanidins level change of A. palmatum Dissectum Atropurpureum

...... 96

Fig 39. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum

Dissectum Atropurpureum ...... 97

Fig 40. Histological staining of proanthocyanidins in a mature leaf of A. palmatum

Dissectum Atropurpureum ...... 98

Fig 41. Leaf phenotype of A. palmatum 'Okushimo' and measurement of anthocyanins and proanthocyanidins ...... 100

Fig 42. LC chromatogram of A. palmatum 'Okushimo' ...... 102

Fig 43. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Okushimo' ...... 103

Fig 44. Major proanthocyanidins of A. palmatum 'Okushimo' ...... 104

Fig 45. Trend of proanthocyanidins level change of A. palmatum 'Okushimo' ...... 105

xiv Fig 46. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum

'Okushimo' ...... 106

Fig 47. Histological staining of proanthocyanidins in a mature leaf of A. palmatum

‘Okushimo’ ...... 107

Fig 48. Leaf phenotype of A. palmatum 'Oridono nishik' and measurement of anthocyanins

and proanthocyanidins ...... 109

Fig 49. LC chromatogram of A. palmatum 'Oridono nishik' ...... 111

Fig 50. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins

from leaves of A. palmatum 'Oridono nishiki' ...... 111

Fig 51. Major proanthocyanidins of A. palmatum 'Oridono nishiki' ...... 112

Fig 52. Trend of proanthocyanidins level change of A. palmatum 'Oridono nishiki' ...... 113

Fig 53. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Oridono

nishiki' ...... 114

Fig 54. Histological staining of proanthocyanidins in a mature leaf of A. palmatum ‘Oridono

nishik’ ...... 115

Fig 55. Trend of anthocyanin and proanthocyanidin level change ...... 124

Fig 56. Trend of proanthocyanidin level change ...... 135

CHAPTER 3 - ANTHOCYANINS AND PROANTHOCYANIDINS METABOLISM IN

SPRING JUVENILE LEAVES OF SEVERAL DECIDUOUS TREE SPECIES

Fig 57. Leaf phenotype of Acer rubrum and measurement of anthocyanins and

proanthocyanidins ...... 151

Fig 58. LC chromatogram of Acer rubrum ...... 154

Fig 59. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins

from leaves of Acer rubrum ...... 155

xv Fig 60. Leaf phenotype of Acer x freemanii and measurement of anthocyanins and proanthocyanidins ...... 156

Fig 61. LC chromatogram of Acer x freemanii ...... 158

Fig 62. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of Acer x freemanii ...... 159

Fig 63. Leaf phenotype of Acer longipes and measurement of anthocyanins and proanthocyanidins ...... 161

Fig 64. LC chromatogram of Acer longipes ...... 163

Fig 65. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of Acer longipes ...... 165

Fig 66. Leaf phenotype of Acer henryi ...... 167

Fig 67. Measurement of anthocyanins and proanthocyanidins in Acer henryi leaves ...... 169

Fig 68. LC chromatogram of Acer henryi ...... 170

Fig 69. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of Acer henryi ...... 171

Fig 70. Leaf phenotype of Acer truncatum and measurement of anthocyanins and proanthocyanidins ...... 173

Fig 71. LC chromatogram of Acer truncatum ...... 175

Fig 72. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins from leaves of Acer truncatum ...... 177

Fig 73. Leaf phenotype of Acer negundo and measurement of anthocyanins and proanthocyanidins ...... 178

Fig 74. LC chromatogram of Acer negundo ...... 180

xvi Fig 75. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins

from leaves of Acer negundo ...... 181

Fig 76. Leaf phenotype of Prunus padus ...... 183

Fig 77. Measurement of anthocyanins and proanthocyanidins in Prunus padus leaves ...... 185

Fig 78. LC chromatogram of Prunus padus ...... 186

Fig 79. LC analysis of anthocyanidins released from butanol-HCl hydrolysis of anthocyanins

from leaves of Prunus padus ...... 187

xvii CHAPTER 1

LITERATURE REVIEW

Plant flavonoids are the largest group of plant phenolics. They exist in all of fern and plants investigated thus far. They have numerous biological and ecological functions in plants. Many of them show strong anti-microbial activities to protect plants from pathogens.

And many of them have strong antioxidative activities protecting against UV-radiation and other oxidant stress to cells and tissues. Particularly, the presences of plant flavonoids in reduce the UV-induced damage to embryos. Several publications summarized genetics and regulation of flavonoid biosynthesis (Koes et al., 2005; Winkel-Shirley, 2001) and also

ecological as well as physiological functions of flavonoids (Koes et al., 1994; Rolfe, 1988;

Shirley, 1996; Stenlid, 1963, 1968, 1970; Winkel-Shirley, 2001). Recently, plant flavonoids

(e.g., quercetin) have been found to be involved in the function of plant auxins (Jacobs and

Rubery, 1988; Peer and Murphy, 2007). These numerous functions are hypothesized to be the

consequence of plant evolution and coevolution (Koes et al., 1994; Shirley, 1996; Stafford,

1991).

Anthocyanins are one of the richest classes of pigments in plant kingdom. They are one of

the final products of plant flavonoids. Their existence in plants gives rise to pink, red, blue

and purple coloration in tissues (Grotewold, 2006; Mol et al., 1998). Over the past 100 years,

a large number of studies have demonstrated that the formation of anthocyanins in flowers

1 attracts pollinators and their presence in fruits and seeds appeals to animal dispensers

(Grotewold, 2006; Mol et al., 1998). Anthocyanins can absorb UV radiation and visible light,

thus protecting against UV-induced damage as well as damage induced by high light

intensity to cells and tissues (Close and Beadle, 2003; Lee and Gould, 2002b; Steyn et al.,

2002). In addition, anthocyanins are important taxonomic pigments for the systematic and taxonomic study of plants (Bate-Smith, 1962, 1968; Delendick, 1990; Ji et al., 1992b;

Nishiura et al., 1971).

Proanthocyanidins are another group of plant flavonoid end products. This group of plant flavonoids consists of oligomeric or polymeric flavanols. Like anthocyanins, they are also widely produced in most of families of seed plants and ferns (Dixon et al., 2005; Xie and

Dixon, 2005). The ecological functions of proanthocyanidins include anti-oxidative, anti- herbivore and anti-pathogens activities (Aron and Kennedy, 2008; Dixon et al., 2005; Peters and Constabel, 2002).

In this review, I briefly introduce the ecological activities, structures, and biosynthesis of plant flavonoids, anthocyanins, and proanthocyanidins, which are common plant secondary metabolites in Acer species of my research projects.

2 1.1 Plant Flavonoids

Introduction

Plant flavonoids are a large group of natural products prevalent in all investigated families of

angiosperms, gymnosperms and ferns. By 2004, approximately 9000 different flavonoids had been found from plants and this number is predicted to increase when more survey studies

will be carried out for more plant species (Williams and Grayer, 2004).

Major groups of plant flavonoids include chalcones, aurones, phlobaphenes, stilbenes,

flavones, flavanones, dihydroflavonols, flavonols, flavan-3,4-diols, isoflavonoids,

neoflavonoids, anthocyanins, flavan-3-ols and proanthocyanidins (Weiss, 1980). The major classes include chalcones, flavones, flavonols, anthocyanins, and proanthocyanidins

(condensed tannins) which are present in almost all higher plants investigated (Winkel-

Shirley, 2001). Their chemical structures are shown in Figure 1.

All plant flavonoids (with the exception of stilbene) share the same main molecular, C6-C3-

C6 (C15), which is formed from condensation of two substrates, 4-coumaroyl-CoA derived

from the general shikimic pathway, and three malonyl-CoAs derived from the malonic acid

pathway.

3

Fig 1. Backbone structures of some flavonoids. A: chalcone; B: flavone; C: flavanone; D: flavonol; E: flavan‐3‐ ol; F: isoflavonoids; G: aurones; H: neoflavonoids; I:) flavan‐3,4‐diols; J:) stilbene. * indicates positions as stereocenters.

4 Different groups of plant flavonoids are derived from chalcone structure by enzymatic

modifications including, a) saturation of the central heteroatomic pyran ring (ring C), b)

reduction or dehydrogenation of the C-ring; c) C-glycosylation, methylation, acetylation,

prenylation; d) O-glycosylation, methylation, acetylation, prenylation, gallylation etc; e)

deoxygenation; f) condensation (Maciej Stobiecki, 2005; Maike Petersen, 1999). Such

multiplicity leads to the large number of structurally diverse plant flavonoids as a group of

natural products.

Biosynthesis of flavonoids

The biosynthesis of flavonoids has been extensively studied in model plants, e.g. maize (Zea mays), snapdragon (Antirrhinum majus), petunia (Petunia hybrida) and Arabidopsis. The

application of integrative functional genomics approaches to these model plants has enhanced our understanding of the pathways and genetic regulation of plant flavonoid biosynthesis (Holton and Cornish, 1995; Mol et al., 1998; Winkel-Shirley, 2001). These networks of the pathways are summarized in Figure 2. Flavones, flavonols, anthocyanins and proanthocyanidins are four major groups of end products of the flavonoid pathway. Recent biochemical and genetic studies have shown that proanthocyanidins specifically seem to be

“the final frontier of the flavonoid pathway” (Dixon et al., 2005).

Chalcone, the first C15 (C6-C3-C6) intermediate, is formed from condensation of one 4-

coumaryol-CoA and three molecules of malonyl-CoA and is catalyzed by chalcone synthase

5 (CHS). There are two major types of chalcones, trihydroxychalcone and

tetrahydroxychalcone, the former leading to production of isoflavonoids whereas the other gives rise to the production of flavanoes, flavonols, flavandiols, anthocyanins and proanthocyanidins (condensed tannins). Tetrahydroxychalcone can undergo isomerization to form naringenin, which belongs to the group of flavanones with a closed hetero ring (ring C) catalyzed by chalcone isomerase (CHI). Naringenin is converted to dihydrokaempferol

(DHK) by flavanone 3-hydroxylase (F3H). DHK can be subsequently hydroxylated by either flavonoid 3’-hydroxylase (F3’H) or flavonoid 3’, 5’-hydroxylase (F3’5’H) to produce dihydroquercetin (DHQ) or dihydromyricetin (DHM) respectively. DHK, DHQ and DHM are 3-OH-flavanones, also known as dihydroflavonols. They undergo further reduction to form flavan-3,4-diols (flavandiols), which are also called as leucoanthocyanidins, catalyzed by dihydroflavonol 4-reductase (DFR). Those flavandiols can be diverted into two directions, one to the production of proanthocyanidins (condensed tannins), and the other to the

production of anthocyanidins by means of anthocyanidin synthase (ANS) (Heller, 1993;

Holton and Cornish, 1995; Koes et al., 1994; Maike Petersen, 1999; Shirley, 1996; Stafford,

1991; Winkel-Shirley, 2001). Figure 2 is a revised schematic diagram of major branch

pathways of the plant flavonoid biosynthesis including recently uncovered biosynthesis of

proanthocyanidins. This diagram is derived from previous work (Springob et al., 2003).

6

Fig 2. Scheme of the flavonoid biosynthetic pathway. ACCase, acetyl‐CoA carboxylase; PAL, phenylalanine ammonia‐lyase; C4H, cinnamate 4‐hydroxylase; 4CL, 4‐coumarate:CoA ligase; CHS, chalcone synthase; CHKR, chalcone ketide reductase; STS, stilbene synthase; CHI, chalcone isomerase; FS, flavone synthase; IFS, isoflavone synthase; F3H, flavanone 3‐hydroxylase; F3’H, flavonoid 3‐hydroxylase; F3’5’H, flavonoid 3,5‐ hydroxylase; FLS, flavonol synthase; DFR, dihydroflavonol 4‐reductase; LAR, leucoanthocyanidin reductase; ANS, anthocyanidin synthase; ANR, anthocyanidin reductase; GT, glucosyltransferase; ACT, anthocyanin acyltransferase; MAT, malonyltransferase. This figure is reproduced and updated with recent discovery of ANR based on original figure 1 (Springob et al., 2003). Stilbene and stilbene synthase are also added into the pathway.

7 Ecological and physiological significances

Flavonoid compounds have numerous biological and ecological functions in plants (Seigler,

1998). For starters, most flavonoids have very high absorption coefficients for UV radiation.

This UV-absorption feature, along with the fact that flavonoids are frequently localized in

vacuoles int the epidermal cells and sub-epidermal cells, provides plants with good UV

protection for mesophyll cells containing the photosynthetic apparatus that lies underneath epidermal cells. Thus the dominant presumed function of flavonoids is to serve as a UV filter for photoprotection.

Nevertheless, Stafford (1991) disagreed with such perception based on the possible evolution processes of both the flavonoid biosynthetic pathway and enzymatic efficiency. Instead, she proposed the very initial function for flavonoids as internal physiological regulators or

chemical messengers when those compounds arose during the early emergence of the first

land plants (Stafford, 1991).

Reports which support her hypothesis are that flavonoids serve as inhibitors of ATP

formation in plant mitochondria and also as either inhibitors or cofactors of IAA oxidase

involved in IAA degradation process (Stenlid, 1963, 1968, 1970). Moreover, Jacobs and

Rubery (1988) found a specific group of flavonoids which are able to compete with a

synthetic compound - naphthylphthalamic acid (NPA) for binding to its receptor, thus acting

like natural auxin transport regulators through affecting polar auxin transport (Jacobs and

8 Rubery, 1988). However, there is insufficient evidence to support a role for flavonoids as

chemical signals or regulators within intact plants, as all the data to date uses in vitro tests.

The phenomena that scientists have observed may not reflect the true conditions in vivo.

Such inconsistence might result from the possibility that plants, especially higher plants have developed more sophisticated regulatory mechanisms, including signaling pathways and regulation of gene expression, thereby eliminating the need to maintain flavonoids as regulators or chemical messengers in plants after millions of years of evolution. Another possibility is that, due to compartmentalization of production and accumulation of flavonoid compounds within certain cell types of specific tissues and organs, those effects and action may never occur in reality within intact plants due to inaccessibility to their targets.

Isoflavonoids (e.g. genistein and daidzein) also play a very important role in symbiotic interactions between legumes and nitrogen-fixing bacterium to form nodules (Debelle et al.,

2001; Denarie et al., 1992; Gottfert, 1993; Lorkiewicz, 1997; Rolfe, 1988; Schultze and

Kondorosi, 1998; Spaink, 1994; van Rhijn and Vanderleyden, 1995).

Another well-known physiological function of flavonoids is the recruitment of pollinators and seed dispersers through pigmentation in flowers, fruits and seeds. Flavonoids also have key roles in male fertility of some species, in defense as antimicrobial agents and feeding deterrents to herbivores (Winkel-Shirley, 2001). Many flavonoid compounds including anthocyanins and proanthocyanidins have antioxidative activities due to their chemical nature

9 and those flavonoids may protect plants from oxidation-induced damage in stressful environment (Chalker-Scott, 1999; Close and Beadle, 2003).

Nutraceutical significances

Flavonoids are major nutraceutical compounds (Lin and Weng, 2008). Due to their structures and chemical features, flavonoid compounds can act as potent antioxidants. So far, more than

14,000 research articles about the antioxidant activity of flavonoids from different sources can be retrieved from PubMed database (search conducted in June, 2009). Attributable to such antioxidant abilities, uptake of flavonoids from diets can protect many cells, cellular organelles and molecules from auto-oxidation. Flavonoids have also been recorded to have both carcinogenic activity and anti-mutagenic effects (Beier, 1989; Macgregor, 1986). Many studies have shown that flavonoid compounds have multiple beneficial effects for human health including anti-inflammatory, anti-allergic, antiviral, antimicrobial and anti-hepatotoxic activities (Chen et al., 1986; Gabor, 1986; Middleton et al., 1986; Selway, 1986; Vlietinck et al., 1986; Wagner, 1986).

Taxonomic significances

Since flavonoids compounds are widely distributed in all plants, and also have a wide variety of structures with modifications, they became a widely-used group of chemical markers for the purpose of chemotaxonomy about 20 to 30 years ago (Vickery, 1981). Numerous analytic methods have been developed. These methods include paper chromatography, thin-layer

10 chromatography, one- or two- dimensional chromatography etc., which are quick approaches

to survey flavonoids from different plant sources. In addition, direct visual detection or

detection under UV spectrophotometers for flavonoids without advanced equipment endows

flavonoids with more accessibility as useful chemotaxonomic markers for taxonomists in

those days (Mabry, 1970; Vickery, 1981). Bate-Smith conducted an extensive survey of

phenolic compounds including flavonoids in terms of taxonomic significance for both

monocotyledon and dicotyledons plants, and revealed that certain substitution patterns of

compounds were correlated with the plants’ taxonomic relationships, which maybe a

consequence of general evolutionary processes (Bate-Smith, 1962, 1968). Two well-known

representative examples of the taxonomic significance of flavonoids are isoflavonoids and

anthocyanins versus betacyanins. Isoflavonoids were long recognized as almost mainly produced in Fabaceae (bean family) species, although recently more and more isoflavonoids have been discovered from non-legume plants (Lapcik, 2007; Mackova et al., 2006).

Anthocyanins, as a group of natural pigments, are mutually exclusive to betacyanins, which only occur in Caryophyllales (Stafford, 1994). Flavonoid distribution patterns have been used to determine the parents of cultivars in crops or ornamental plants which have undergone numerous steps of hybridization and backcrossing. For example, such methods have been demonstrated to be an efficient approach to determine the type of cultivar or trace the origin of Citrus varieties (Nishiura et al., 1971; Nogata et al., 2006). Flavonoids including anthocyanins and proanthocyanidins can still be of significant use for chemotaxonomy to test and/or corroborate current plant phylogeny, not to mention the valuable information of plant

11 metabolism evolution. However, the power of flavanoids as phylogenetic/taxonomic markers

is limited due to many reasons regarding variability, stability, detectibility and incomplete

sampling. They can be explored more by using the modern techniques in metabolomic

profiling.

1.2 Anthocyanins and anthocyanidins

Introduction

Anthocyanins are one of the most ubiquitous pigments in the plant kingdom besides

chlorophyll and carotenoids. They are generally characterized by their blue color in solution

with higher pH, red color in acidic solution and purple in solution at near-neutral pH. As

early as the 17th century, scientists began to discover the presence of anthocyanins as red pigments in plants but not until 19th century did the term “anthocyanin” come into existence.

“Anthocyanin” originates from the Greek words antho () and kyanos (blue) likely due to its presence in and contribution to the blue coloration of flowers (Lee and Gould, 2002a).

Anthocyanins (Figure 3A), in terms of more formal chemical nomenclature, are glycosides of poly-hydroxy and poly-methoxy derivatives of 2-phenylbenzopyrylium or in short, flavylium salts (Kong et al., 2003). Anthocyanidins, are the aglycones of anthocyanins. Anthocyanins, as end products of the flavonoid pathway, have similar modification patterns to most flavonoids, including hydroxylation, methylation, C-glycosylation directly to carbon atom of the anthocyanidin skeleton, O-glycosylation of hydroxyl groups, actylation with either

12 aliphatic or aromatic acids, prenylation and sulfation (Maciej Stobiecki, 2005; Maike

Petersen, 1999). The types, position and degree of glycosylation also vary for different anthocyanins (Kong et al., 2003). Furthermore, each sugar (i.e. glycosyl-) can be acylated with a variety of different phenolic or aliphatic acids to form more anthocyanins based on various combinations of those modifications (Williams and Grayer, 2004).

Fig 3. General structures of anthocyanins and anthocyanidins. A: anthocyanins; B: anthocyanidins. R1‐R7 are often H, OH, OCH3 or glycosyl. R refers to substitution patterns for different anthocyanidins and R is often H, OH or OCH3.

Types of anthocyanins and anthocyanidins

New structures of anthocyanins have been continuously discovered over the past two decades

(Harborne and Williams, 1995, 1998, 2001; Williams and Grayer, 2004). More than 450

anthocyanins were reported from natural sources until 2004 (Harborne and Williams, 1998,

2001; Mazza, 1993; Williams and Grayer, 2004).

13 All those anthocyanins are derived from just a few aglycone anthocyanidins. Kong et al

(Kong et al., 2003) summarized from previous studies that there were 17 known naturally occurring anthocyanidins including three 3-deoxyanthocyanidins (Figure 3B, Table 1).

Table 1. Naturally occurring anthocyanidins. This table is reproduced from the original Table 1 (Kong et al., 2003). Substitution pattern Name Abbreviation 3 5 6 7 3’ 4’ 5’ Color Ap H OH H OH H OH H Orange Au OH OH OH OH H OH H Orange Cp OH OMe H OH OMe OH OMe Bluish-red Cy OH OH H OH OH OH H Orange-red Dp OH OH H OH OH OH OH Bluish-red Eu OH OMe H OH OMe OH OH Bluish-red Hs OH OH H OMe OMe OH OMe Bluish-red 6-Hydroxycyanidin 6OHCy OH OH OH OH OH OH H Red Lt H OH H OH OH OH H Orange Mv OH OH H OH OMe OH OMe Bluish-red 5-Methylcyanidin 5-MCy OH OMe H OH OH OH H Orange-red Pg OH OH H OH H OH H Orange Pn OH OH H OH OMe OH H Orange-red Pt OH OH H OH OMe OH OH Bluish-red Pl OH OMe H OH OH OH OH Bluish-red Rs OH OH H OMe OMe OH H Red Tr H OH H OH OH OH OH Red

Based on statistics, only six of the 22 naturally occurring anthocyanidins are common in higher plants – cyanidin (Cy), delphinidin (Dp), malvidin (Mv), pelargonidin (Pg), peonidin

(Pn) and petunidin (Pt). Three of them (Mv, Pn and Pt) are methylated anthocyanidins whilst the other three (Cy, Dp and Pg) are non-methylated ones, which actually are the most widespread in plant kingdom (Kong et al., 2003). The most common anthocyanidin glycosides consist of four groups: 3-monosides, 3-biosides, 3,5-diglycosides and 3,7-

14 diglycosides. Among all, cyanidin 3-glucoside is the most widespread anthocyanin in nature

(Kong et al., 2003).

There are five newly discovered aglycones, three of which were included in the review of

Williams and Grayer in 2004, including 6,7,3’-trihydroxy-5,4’-dimethoxyflavylium,

6,7,3’,4’-tetrahydroxy-5-methoxy-flavylium and rosacyanin B (Williams and Grayer, 2004).

The first two new anthocyanidins, 6,7,3’-trihydroxy-5,4’-dimethoxyflavylium, 6,7,3’,4’-

tetrahydroxy-5-methoxy-flavylium were isolated from Arrabidaea chica leaves

(Bignoniaceae) along with another two 3-deoxyanthocyanidins 6,7-dihydroxy-5,4’- dimethoxy-flavylium and 6,7,4’-trihydroxy-5-methoxyflavylium, also known as “Carajurin” and “Carajurone” respectively (Devia et al., 2002; Zorn et al., 2001). Rosacyanin B is a novel

violet pigment and was isolated from the petals of Rosa hybrida. It was the first C-4

substituted anthocyanidin characterized from plants (Fukui et al., 2002).

Properties of anthocyanins

Different pH conditions can change chemical forms of anthocyanins. For example,

anthocyanins with blue color results from the formation of an anhydro base of certain

anthocyanins (Vickery, 1981). It’s now believed and clarified that color change corresponds

to structure change (Chen and Hrazdina, 2005; Jurd and Geissman, 1963). Castaneda-Ovando

et al (2009) summarized that the forms of anthocyanins are dependent on pH values, which is

diagramed in Figure 4 (Castaneda-Ovando et al., 2009). At pH=1, the flavylium cation (red

15 color) is the predominant species. When pH increases to the range of 2 and 4, the quinoidal blue species become the predominant form. At pH values between 5 and 6, the colored forms

of anthocyanins turn into colorless species. With the rise of pH to 7 and higher, the

anthocyanins are more readily degraded than at low pH (Castaneda-Ovando et al., 2009).

Fig 4. Anthocyanins chemical forms depending on pH and degradation reaction for anthocyanins. Where R1 = H or saccharide, R2 and R3 = H or Methyl. (Castaneda‐Ovando et al., 2009)

16 Different purified anthocyanins/anthocyanidins have different colors in vitro under acidic

conditions. The most abundant and widespread anthocyanidins: cyanidin, delphinidin and

pelargonidin (Kong et al., 2003) have different chromatic features. Such differences come

from the hydroxyl group substitution of the ring B. With an increase in the hydroxylation

pattern (Pelargonidin

occurs (de Freitas and Mateus, 2006). In other words, the visible wavelength of maximum

absorption will be shifted to higher values as more hydroxyl groups are added in the ring B

(de Freitas and Mateus, 2006).

Biosynthesis of anthocyanins

Numerous efforts have been made to determine the enzymes involved in the biosynthetic

pathway of anthocyanins and also to identify regulatory proteins, mainly in the flowers and

fruits of model plants (including Arabidopsis thaliana, Antirrhinum majus and Petunia hybrida) and crop plants such as maize (Allan et al., 2008; Dooner et al., 1991; Ferrer et al.,

2008; Gonzalez et al., 2008; Holton and Cornish, 1995; Springob et al., 2003). Aglycone anthocyanidins and anthocyanin glycosides are end products of the flavonoid biosynthetic pathway, thus they share the same pathway catalyzed by a series of enzymes including CHS,

CHI, F3H (F3’H, F3’5’H) and DFR (Figure 2). Anthocyanidin synthase (ANS), also known as leucoanthocyanidin dioxygenase (LDOX), is the most indispensable enzyme because it is the key enzyme to produce anthocyanidins (Figure 2).

17 The biochemical reaction mechanism of ANS was not studied until 2001. Nakajima et al

(2001) conducted parallel enzymatic experiments with snapdragon, petunia, torenia and maize ANSs to determine their catalytic ability to form anthocyanidins (Nakajima et al.,

2001). These authors proposed a possible in vivo biochemical mechanism for the conversion of leucoanthocyanidin to anthocyanidin catalyzed by ANS. These steps involve an ANS-

catalyzed conversion of leucoanthocyanidin to 2-flaven-3,4-diol. This product would

spontaneously undergoes isomerization to 3-flaven-3,3-diol, which is a pseudobase. A

glycosylation process follows by means of 3-O-glucosyltransferase (3-GT) resulting in the formation of a 3-O-glucosylated pseudobase under the neutral conditions (pH=7) found in cytosol. This 3-O-glucosylated pseudobase might be transported into vacuoles and converted to the colored form flavylium cation due to the acidic conditions (pH=5) in vacuoles

(Nakajima et al., 2001). The structure of anthocyanidin synthase was later determined with

Arabidopsis thaliana ANS. AtANS is a small enzyme with 112 amino acids. Its structure and

functional mechanism was just recently elucidated and has been further studied since then

(Welford et al., 2005; Wilmouth et al., 2002).

The aforementioned 3-O-glucosyntransferase, belongs to another crucial group of structural

enzymes – glycosyltransferases, which modify and also stabilize labile anthocyanidins

through glycosylation. After glycosylation at the 3-O position, further potential modifications

include the addition of a second glycosyl, acyl, methyl and prenyl groups. 3-GT cDNA

homologs have been cloned and characterized from several plant species including Gentiana

18 triflora (Tanaka et al., 1996), Perilla frutescens (Gong et al., 1997), Petunia hybdrida

(Yamazaki et al., 2002) and Vitis vinifera (Ford et al., 1998). All these 3-GTs have a broad substrate specificity to catalyze 3-O-glycosylation of both flavonols and anthocyanidins. In comparison, anthocyanin 5-O-glucosyltransferase (5-GT) has relatively strict substrate specificity. Several 5-GT homologs have also been discovered from different species

(Yamazaki et al., 1999; Yamazaki et al., 2002). Another important group of enzymes involved in later steps of anthocyanin synthesis and modifications are acyltransferases (ACT).

Similar to 5-GTs, ACTs might also work downstream of 3-GTs. Usually ACTs modify anthocyanins with either aromatic or aliphatic organic acids through an acylation reaction, which can contribute to intra- and/or inter-molecular stacking for the stabilization of anthocyanin molecules. Such molecular stacking, also known as co-pigmentation, can result in color changes due to bathochromic effect (Springob et al., 2003).

Ecological significances

Anthocyanins are very important naturally occurring colorants which impart colorful phenotypes to plants. These colored pigments are long believed to be involved in the attraction of pollinators to flowers and the attraction of dispersers to seeds and fruits

(Winkel-Shirley, 2001). Evidence available to support these hypotheses includes “ Flag

Hypothesis” proposed by Stiles (1982). These researchers hypothesized that the brilliant leaf colors in the autumn were attractive to birds or other dispersers to ripe fruits thus promoting dispersal of propagules and enhancing the colonization success of the species (Stiles, 1982).

19 Another theory is the “Defensive Signal Hypothesis” (Hamilton and Brown, 2001), which proposes that anthocyanins function in plant defense, most likely as warning signs for herbivores (e.g. aphids) and pathogens. Several reports have been published to support this hypothesis (Archetti, 2000, 2008; Archetti and Brown, 2004; Archetti and Leather, 2005;

Chittka and Doring, 2007; Hamilton and Brown, 2001; Lev-Yadun and Gould, 2007;

Yamazaki, 2008). Even though it is a very inviting idea, there is difficulty in designing experiments to prove such a hypothesis. Apart from those hypothesized functions, anthocyanins may have other ecological and physiological significances for plants, which have been extensively investigated for a long time.

The first proposed ecological function of foliar anthocyanins is photoprotection (Gould et al.,

1995). As is known, plants are autotrophic organisms which make use of sunlight to produce

energy and nutrients through photosynthesis. However, utilization of sunlight does not

necessarily mean that more sunlight is better. On the contrary, high light intensity can cause

damage to both subcellular organelles and even the whole plant system. When the rate of

photon absorption exceeds the rate of photon utilization by photosynthesis, then excess light

intensity can cause “self-destruction” of the photosynthetic (PS) apparatus by increasing the tendency for PSII to transfer energy to molecular oxygen resulting in damage to the PS apparatus itself (Asada, 1999). Plants adopt mechanical and/or biochemical methods to minimize or dissipate the excess photons as heat or maintain stable and continuous turnover of photosynthetic apparatus in order to minimize possible damage to PSII. The xanthophyll

20 cycle, antioxidant activity and external foliar waxes are three well-characterized mechanisms

of photoprotection (Close and Beadle, 2003). However, when under stress or during periods

of developmental change, such protective or compensative strategies have not been fully

established or efficiently functional. Plants must have mechanisms to serve as sunscreen to

reduce incident visual light intensity; meanwhile this sunscreen should minimize the

interference of normal photosynthesis processes for growth and development (Dominy et al.,

2002; Gould et al., 2002b; Lee, 2002a; Lee and Gould, 2002a, b; Steyn et al., 2002).

Anthocyanins are unique among the flavonoids in their ability to absorb light within the visible spectrum. Such absorption in visible light can protect PS apparatus by directly shielding chloroplasts from bright light (Lee and Gould, 2002b). Additionally, most anthocyanins have peak absorbance in the blue-green-yellow region of the spectrum (500-

550nm). Such strong absorbance complements the absorbance spectrum of chlorophylls, which could reduce the interference of photosynthesis. Those characteristics suggest a potential light-screening function for anthocyanins. In order to fully carry out this light- attenuating function, anthocyanins have been postulated to exist at the upper leaf surface including upper epidermal cell layer or palisade cells (Lee, 2002b). Moreover, anthocyanins must be widely distributed in order to occupy almost the entire sun-facing surface. In plant cells, the vacuole is the only organelle that takes up almost all intracellular space. It is therefore reasonable to expect anthocyanins to be localized in the vacuoles of cells of the adaxial leaf surface or cells near the sun-facing surface, especially for newly expanding

21 leaves (Hughes et al., 2007), senescing leaves or leaves experiencing stress (Feild et al.,

2001; Lee, 2002a; Lee and Gould, 2002b) such as suboptimal water availability or cool but not freezing temperature. Those leaves, as mentioned above, are possibly deficient in effective and mature light-screening mechanism compared to mature and normal leaf, which is in less need of alternative sun-screening strategy. Studies have been carried out to determine the distribution of anthocyanins in leaves (Lee, 2002b) and the relationships between anthocyanin and chlorophyll levels (Feild et al., 2001; Lee, 2002a; Lee et al., 1987;

Lee and Collins, 2001; Lee and Gould, 2002a, b). These studies support the role of anthocyanins as providing photoprotection from visible light.

The second hypothesis for the possible function of foliar anthocyanins is osmotic adjustment.

It has been long assumed that anthocyanins may confer resistance to water stress by osmotic adjustment of the vacuolar sap where they are stored and deposited. The vacuole could serve as osmotic regulators by means of its dissolved contents affecting changes of water potential.

When plants are exposed to drought, heat, cold or high saline conditions, foliar anthocyanins are often synthesized de novo as a response to these adverse conditions to ameliorate drought stress. Evidence of this function has been reviewed (Chalker-Scott, 1999). The transient nature of foliar anthocyanin production and accumulation may allow plants to respond quickly and temporarily to environmental variability on metabolic levels rather than through more permanent structural modifications on developmental levels. Despite some evidence supporting this hypothesis, Close et al (2003) argued for the possibility that the production

22 and accumulation of anthocyanins may simply result from cold- or drought- induced

photoinhibition rather than for the sake of osmotic adjustment (Close and Beadle, 2003).

However, the question remains open. Anthocyanins may play important roles for both

photoprotecion and osmotic adjustment considering the fact that such resourcefulness would

be more efficient than one single function.

Anthocyanins can act as antioxidants as well in flowering plants (Kong et al., 2003) due to their reductive properties. Based on the chemical structure and characteristics of anthocyanins, the high activity of reducing power has been ascribed primarily to the two ortho-related hydroxyl groups on the ring B of the anthocyanin molecule, the presence of O+ in the C ring, and the numbers and positions of free hydroxyl groups (RiceEvans et al.,

1996). Thus, anthocyanins have an inherent potential to protect plant cells from the effects of oxidative damage (Gould et al., 2002a). However, such in vitro evidence though seemingly plausible, does not necessarily conclude the same condition in vivo. Antioxidant functions have to be confirmed and implemented with activity and effect in the proper subcellular compartments where oxidative damage would occur. So far, there is no sufficient evidence that could prove so.

Anthocyanins are now assumed to play important roles in ecophysiological interactions among plants and environment and they are no longer considered to be extravagant waste products of metabolism. Although many potential functions of anthocyanins continue to be

23 debated, the multiplicity of foliar anthocyanin roles in vitro has been demonstrated. It remains difficult to determine the primary role of anthocyanins with so many confounding elements. Thus, investigation of anthocyanins dependent on the reference system, i.e. species, developmental stage or specific biotic and abiotic stress is a more scientific and reasonable way to elucidate the real function of anthocyanins in plants (Manetas, 2006).

Nutraceutical significances

In addition to those ecological and physiological significances, anthocyanins are also well- known for their anti-oxidant characteristics. For the last two decades, anthocyanins have been identified and documented in numerous research papers and reviews as potential food supplements for the benefits of human wellness or prevention of cancer (Fimognari et al.,

2008; Wang and Stoner, 2008), heart health (Mazza, 2007) and even for prevention of Type

II diabetes (Ghosh and Konishi, 2007). Based on the review by Kong et al (2003), anthocyanins have pharmacological properties, including their effectiveness in reversing age- related deficits, their reduction of tissue inflammation, their ability to suppress tumors, their protective effect against carbon tetrachloride-induced liver injury, their potential antimutagenicity, their application in controlling oxidative stress during pregnancies complicated by intrauterine growth retardation, their ability to inhibit epidermal growth- factor receptor (EFGR) to inhibit the growth of tumor cells, their antioxidant properties to scavenge free radicals, their abilities to interact with DNA and stabilize and protect DNA from oxidation etc (Kong et al., 2003).

24 1.3 Proanthocyanidins

Introduction

Proanthocyanidins (condensed tannins) form a very special group of flavonoids considering

their large molecular weights as polymers built up with flavan-3-ol monomer units. They

have been known in history as tanning agents for animal skins due to their abilities of precipitating proteins and they also provide astringency taste in teas, wines and fruit juices

(Dixon et al., 2005). The biosynthesis of proanthocyanidins shares all the upstream steps leading to flavon-3,4-diols (Stafford, 1989) with anthocyanin biosynthesis.

Proanthocyanidins are usually deposited into vacuoles of plant cells after biosynthesis by

means of transportation just like anthocyanins. Mostly in woody flowering plants,

proanthocyanidins make up a considerable proportion of the dry weight of plants.

Proanthocyanidins are often concentrated in epidermal tissues, a similar phenomena to

anthocyanins. Colorless or yellowish proanthocyanidins can produce anthocyanidins under

hydrolysis. Such chemical change, as well as color change due to production of colored

anthocyanidins during hydrolysis, has been documented for a very long time (Bate-Smith,

1968; Batesmith, 1954; Lawrence et al., 1938; Robinson and Robinson, 1931).

Types of proanthocyanidins

Proanthocyanidins are widely distributed in the plant kingdom, but are particularly common

in conifers (Seigler, 1998). They are commonly known as either oligomeric or polymeric end

25 products of the flavonoid biosynthetic pathway (Dixon et al., 2005). So far, there are possibly

around 1000 proanthocyanidins which have been isolated and characterized (personal correspondence with Dr. Xie). All those structures differ in the type of linkages between flavonoid units, hydroxylation patterns, stereochemistry of carbons 2, 3, and 4 of the pyran ring, and the presence or absence of additional substitution groups (Lewis, 1989), and also the degree of polymerization is a very important parameter for proanthocyanidins (Xie and

Dixon, 2005). All proanthocyanidins consist of two parts; one is starter unit, the other one is extension unit. Proanthocyanidins can only have one starter unit, whilst the number of extension units has no certain limitation of this polymeric group of chemicals. The most abundant monomers in proanthocyanidins are (+)-catechin and (-)-epicatechin, with 2,3- trans-stereochemistry and 2,3-cis stereochemistry respectively. According to statistic analysis of naturally occurring proanthocyanidins B from previous literature, Xie et al (2005) found that 2,3-cis-(-)-epicatechin is the predominant extension unit among oligomeric

proanthocyanidins, and also a common starter unit as its stereo-isomer 2,3-trans-(+)-catechin

(Xie and Dixon, 2005). In short, these two stereochemically different monomers can serve as

either starter or extension units. Many plants have been found to contain mixtures of dimeric

proanthocyanidins (Seigler, 1998). Proanthocyanidin dimers B (Figure 4, (3-6)) (Dixon et al.,

2005) is one dimeric group of proanthocyanidins formed up with different arrangements and

combinations of (+)-catechin and (-)-epicatechin. Since all of them are derived merely from

cyanidin-related substrates, i.e. (+)-catechin and (-)-epicatechin, this group is known as

26 procyanidin, with four members in it. The linkage between monomeric units in this group is between the 4-position of the ‘upper’

Fig 5. Structures of different proanthocyanidins. Proanthocyanidins are oligomeric or polymeric end products of the flavonoid biosynthetic pathway. They are present in the fruits, bark, leaves and seeds of many plants, where they provide protection against predation. At the same time they give flavor and astringency to beverages such as wine, fruit juices and teas, and are increasingly recognized as having beneficial effects on human health. The presence of proanthocyanidins is also a major quality factor for forage crops. The past 2 years have seen important breakthroughs in our understanding of the biosynthesis of the building blocks of proanthocyanidins, the flavan‐3‐ols (+)‐catechin and (‐)‐epicatechin. However, virtually nothing is known about the ways in which these units are assembled into the corresponding oligomers in vivo. Molecular genetic approaches are leading to an understanding of the regulatory genes that control proanthocyanidin biosynthesis, and this information, together with increased knowledge of the enzymes specific for the pathway, will facilitate the genetic engineering of plants for introduction of value‐ added nutraceutical and forage quality traits. (Dixon et al., 2005) (Figure continues on next page.)

27

28 unit and the 8-position of the ‘lower’ unit (Figure 4). There is another special linkage

between both C2 and C4 of the ‘upper’ unit and the oxygen at C7 and positions 6 or 8 of the

‘lower’ unit, which occurs in procyanidin A (Figure 4, (12-13)). Figure 4 shows structures of

different flavan-3-ol building blocks (i.e. monomer) of proanthocyanidins and examples of

proanthocyanidins in dimer, oligomer and polymer forms. For the sake of convenience, 2,3-

cis-(-)-epicatechin is termed as epicatechin, 2,3-trans-(+)-catechin as catechin and

proanthocyanidins mentioned afterwards include both flavan-3-ols and other proanthocyanidin dimers/oligomers.

Biosynthesis of proanthocyanidins

It has been well established that proanthocyanidins come from the polymerization of flavan-

3-ol monomers, which are products of a branch pathway of anthocyanin biosynthesis since flavan-3-ols are synthesized from anthocyanidin precursors, i.e. flavan-3,4-diols

(leucoanthocyanidins), through leucoanthocyanidin reductase (LAR) to produce flavan-3-ols with 2,3-trans stereochemistry (Shirley, 1996; Winkel-Shirley, 2001). Flavan-3,4-diols (i.e. flavan-diols), also known as leucoanthocyanidins, have been long recognized as monomeric

proanthocyanidins or precursors for the biosynthesis of proanthocyanidins, because flavan-

3,4-diols have the same chemical reactions as many proanthocyanidins, which is the conversion of colorless flavan-3,4-diols into colored corresponding anthocyanidins under hot

acidic hydrolysis (Batesmith, 1954; Lawrence et al., 1938; Robinson and Robinson, 1931)

although flavan-3,4-diols do not possess tanning properties to precipitate proteins or tan

29 animal skins (Seigler, 1998). With the accumulation of genetic data and enzymological studies, current understanding of the biosynthesis of proanthocyanidins has been progressed.

Years ago, however, most models of proanthocyanidins biosynthesis could not decode the paradox that leucoanthocyanidins, as precursors of extension units of proanthocyanidins, have only 2,3-trans- stereochemistry whereas extension units with 2,3-cis- stereochemistry are widely present in the plant kingdom (Xie and Dixon, 2005). In other words, there must be another biosynthetic pathway in existence besides the branched LAR pathway from anthocyanin biosynthesis so that the 2,3-cis forms of flavan-3-ols can be produced and integrated into proanthocyanidins. Xie et al (2003) cloned two homologous BANYLUS genes

from Arabidopsis thaliana and Medicago truncatula and found that both genes encode

anthocyanidin reductase (ANS) which has the ability to convert achiral cyanidin to 2,3-cis-(-)

-epicatechin (Xie et al., 2003). Thus Xie (2005) proposed a modified model for the

biosynthesis of proanthocyanidins, which solves the problem of “missing” stereochemistry in

proanthocyanidins (Xie and Dixon, 2005; Xie et al., 2003).

Ecological significances

Proanthocyanidins are widespread throughout the plant kingdom. Mostly they are

accumulated and stored in vacuoles of cells in many different organs and tissues to provide

protection against predation (Xie and Dixon, 2005). Such deterrent traits can protect plants

against microbial pathogens, insects, pests and larger herbivores (Dixon et al., 2005). High

levels of monomeric flavan-3-ols such as (-)-epicatechin in plants have been reported to

30 impart intact plants with resistance to fungal attack (Ardi et al., 1998). Proanthocyanidins have another special property of binding metals. Such a property might be of benefit to some plants with high levels of proanthocyanidins, for their tolerance against heavy metals in soil

(Kamachi et al., 2005; Stoutjesdijk et al., 2001).

Nutraceutical significances

Proanthocyanidins are now attracting more attention due to their beneficial effects on human health (Dixon et al., 2005). Flavan-3-ols have been reported to exhibit several health benefits by acting as antioxidant, anticarcinogen, cardiopreventive, antimicrobial, anti-viral, and neuro-protective agents (Aron and Kennedy, 2008). Numerous reports (Abe et al., 2000;

Ahmad et al., 2000; Avramovic et al., 1999; Bagchi et al., 2000; Bagchi et al., 2003; Dauer et al., 2003; Deters et al., 2001; Foo et al., 2000; Kandil et al., 2002; Lin et al., 2002; Lou et al.,

1999; Noreen et al., 1998; Pataki et al., 2002; Serafini et al., 2003; Subarnas and Wagner,

2000; Vinson et al., 2002) and several reviews (Aron and Kennedy, 2008; Dixon et al., 2005) illustrate the potential health-protective effect of proanthocyanidins and flavan-3-ol monomers, including anticancer, antioxidant, immunomodulatory, antimutagenic effects etc

(Dixon et al., 2005).

31 CHAPTER 2

FOLIAR ANTHOCYANINS AND PROANTHOCYANIDINS IN SIX ORNAMENTAL

VARIETIES OF ACER PALMATUM

2.1 Introduction

Plant flavonoids are a large group of natural products prevalent in all investigated families of angiosperms, gymnosperms and ferns. By 2004, approximately 9000 different flavonoids had been found from plants and this number is predicted to increase when more survey studies

will be carried out for more plant species (Williams and Grayer, 2004). Major groups of plant

flavonoids include chalcones, flavones, flavonols, anthocyanins, and proanthocyanidins

(condensed tannins) which are present in almost all the higher plants (Weiss, 1980; Winkel-

Shirley, 2001). Anthocyanins, together with carotenoids, xanthophylls and other flavonoids, have been considered the major pigments responsible for the seasonal color changes of autumn senescing leaves (Archetti, 2000; Lee, 2002a). It has long been presumed that in green leaves where no anthocyanins are present, the anthocyanidin pathway may not be fully active thus no anthocyanidins are produced. A very similar and well known phenomenon is the mutual exclusion with betalains and anthocyanins (Grotewold, 2006). A hypothesis was proposed to explain the absence of anthocyanins in this mutual exclusion that anthocyanidin

synthase (ANS), as the last step in the anthocyanidin pathway, may not be functional by any

means (Clement and Mabry, 1996; Forkmann, 1994; Stafford, 1994) in spite of the presence

32 of other flavonoids including proanthocyanidins which have been detected in betalain-

producing plants (Giannasi, 1980). Systematists reconstructed the phylogenetic relationships

of the families in Caryophyllales based on the presence or absence of anthocyanins (in this

case, equivalent to anthocyanidins) and also on the assumption that presence/absence of

anthocyanins alone could reflect the evolutionary status of the anthocyanidin pathway (Soltis,

2005). With the recent discovery of a new biosynthetic pathway of proanthocyanidins from

anthocyanidins (Xie et al., 2003), it is highly possible that in green leaves, this new pathway

is preferred over the traditionally known pathway in which anthocyanidins are converted to anthocyanins through a series of modifications. However, there have been relatively limited numbers of detailed studies on the metabolism of foliar anthocyanins and proanthocyanidins, let alone the researches on the relationship between anthocyanins and proanthocyanidins in response to leaf ontogeny. Evidence of anthocyanidins and anthocyanidin-derived proanthocyanidins in mature green leaves would rewrite our long standing belief on the loss-

of-function of one or more steps in the anthocyanidin pathway in green leaves and would

shed new light on our understanding of the anthocyanidin pathway dynamics.

The study of anthocyanins can be dated back more than 200 years ago (Lee and Gould,

2002a). The early studies included surveys of anthocyanins in flowers and leaves of many

different plant species including Acer species (Lawrence et al., 1938; Price and Sturgess,

1938; Robinson and Robinson, 1931, 1932, 1933, 1934). Acer was traditionally classified in

the family Aceraceae, but has been recently placed in in the group

33 based on angiosperm phylogeny (Figure 6) (Bremer et al., 2003; Judd, 2007; Soltis et al.,

2000; Soltis et al., 1999). This switch was based on a recent phylogenetic analysis

(Harrington et al., 2005). Maple plants were chosen to extract anthocyanins for elucidation of

their structures. In spite of technique limitation, Price and Sturgess (1938) identified cyanidin

3-monoside in leaves of Acer cappadocicum var. rubrum and cyanidin 3-pentoseglycoside in

both A. ginnala and A. palmatum var. septumlobum elegans (Price and Sturgess, 1938). They

noted the differences in the sugar moieties of anthocyanins between young leaves in spring

and mature leaves in fall. They also suggested that these sugar moiety differences might be associated with the transient red coloration appearing in young leaves and with the appearance of autumnal red pigmentation in mature senescing leaves. Structural analysis of anthocyanins in Acer species started in the 1970s (Ishikura, 1975; Ishikura and Sugahara,

1979). Ishikura (1972) hypothesized that the anthocyanin pigmentation in leaves might be associated with foliar developmental stages (Ishikura, 1972). Ishikura (1972) found that leaves of A. palmatum Thunb. var. palmatum and A. buergerianum Miq produced 3- monoglucoside and 3-rutinoside of cyanidin in spring, none of the two anthocyanins in summer, and only the former in the autumn. In the leaves of A. palmatum Thunb. var.

amoenum (Carr.) Ohwi, he similarly found 3-monoglucoside and 3-rutinoside of cyanidin in

the spring and only 3-monoglucoside cyanidin in the autumn. These findings led Ishikura

(1972) to hypothesize that the anthocyanins produced in young leaves were different from

those produced in the autumn leaves (Ishikura, 1972). Further, Ishikura (1976) discovered

that the accumulation of reducing sugar in senescing leaves was positively correlated to a

34 vigorous formation of anthocyanin in autumn leaves (Ishikura, 1976). Since 1990, the

application of 1HNMR, 1H -1H COSY, FAB mass spectrometry techniques promoted

identification of several new anthocyanins from Acer species. Two acylated anthocyanins,

cyanidin 3-O-[2''-O-(galloyl)-beta-D-glucoside] and 3-O-[2''-O-(galloyl)-6''-O-(alpha-L-

rhamnosyl)-beta-D-glucoside] were reported and isolated from red fresh spring leaves of A. platanoides L. cv Crimson Sentry and A. palmatum Thunb. cv Beni-komachi, respectively

(Ji et al., 1992a). In addition, cyanidin 3-(2",3"-digalloyl-beta-glucopyranoside) (a diacylated anthocyanin), cyanidin 3-(2"-galloyl-beta-glucopyranoside) and cyanidin 3-beta- glucopyranoside were determined from extracts of the red leaves of A. platanoides cv

Crimson King by 2D-NMR (Fossen and Andersen, 1999). More chemotaxonomy studies were conducted to understand anthocyanins in both spring newly developing sprout and autumn colored leaves of 111 taxa in Acer. These surveys showed that the anthocyanin profile was more complex in spring leaves than in autumn leaves. Eleven anthocyanins were identified from newly sprouted leaves in spring whereas only six anthocyanins were isolated from red leaves in autumn. Furthermore, seventeen pigmentation patterns of anthocyanins were found in spring leaves but only eleven patterns existed in autumn leaves (Ji et al.,

1992b). All these evidence indicates that red pigmentation pattern in juvenile leaves may not be a simple reverse process of autumn reddening in senescing leaves. Pigmentation pattern change in spring juvenile leaves may be more complicated due to involvement of leaf ontogeny, physiological and ecological requirements, which further suggests a more complex metabolic regulation of anthocyanins production, accumulation and disappearance.

35 Delendick (1990) reported that procyanidin dimers were common among the surveyed Acer species, but prodelphinidin dimers were only found in the leaves of several species under the section Macrantha, A.negundo and A. monspessulanum (Delendick, 1990). In addition,

Ishikura (1976) reported that the level of (+)-catechin increased rapidly in young leaves

(growing from March to July/August) and then remained constant (Ishikura, 1976). However, at present, there are no detailed studies of proanthocyanidins in response to leaf ontogeny on

A. palmatum species. Nor the relationship between anthocyanins and proanthocyanidins or their contributions to pigmentation pattern has been investigated. No enzymatic assay has been carried out in to test ANR activities. Our study is to fill those gaps and to set up a model plant (i.e. A. palmatum) for the investigation of metabolism of foliar anthocyanins and proanthocyanidins.

A. palmatum is one of the most economically valuable Acer species grown in landscapes worldwide. Genetic breeding efforts particularly in Japan over the past three centuries have developed more than 250 cultivars from wild-type A. palmatum (Vertrees, 1978). These varieties are characterized by different leaf morphologies, pigmentation patterns, tree crown topology, or resistance to pathogens to meet landscape designing. Those varieties with various red pigmentation patterns in leaves, which were resulted from genetic modification, are useful materials to understand anthocyanin biosynthesis and catabolism. Our goal of the present study is to understand the mechanisms of anthocyanin metabolism leading to dynamic leaf coloration of Acer palmatum during leaf ontogeny. Our specific objectives are:

36 1) to profile foliar anthocyanins associated with pigmentations in leaf ontogeny and 2) to understand the metabolic and molecular mechanisms leading to the formation of different red pigmentation patterns. Our hypothesis is that the color changes from red/purple/pink to green during the ontogeny of leaves may result from the metabolic channeling of anthocyanidins to proanthocyanidins.

In the present study, we measure the content levels of anthocyanins and proanthocyanidins in leaves of six cultivars of Japanese maple (Acer palmatum Thunb.) at different developmental stages. Metabolic profiles of anthocyanins (if any) and proanthocyanidins were also analyzed by LC-MS. Development-dependent trends of both anthocyanins (if any) and proanthocyanidins were described and the relationship between those two groups of flavonoids was established. A hypothesis of metabolic channeling from anthocyanidins to proanthocyanidins (i.e. (-)-epicatechin and procyanidin B2 etc.) was proposed. ANR activity was tested by enzyme assays to confirm with our hypothesis. Microscopic and histological analyses were used for visual detection of proanthocyanidins. We found that even for green mature leaves with no/trace amount of detectable anthocyanins, the biosynthetic pathway of anthocyanidin was still activated. Unlike red juvenile leaves, most anthocyanidins were converted to proanthocyanidins in green leaves rather than to anthocyanins. Our evidence strongly indicates that metabolic channeling directing the anthocyanin pathway to the proanthocyanidin biosynthesis may play a very important role in pigmentation pattern change along leaf ontogeny.

37

Fig 6. Summary of phylogenetic relationships for angiosperms. This phylogeny is inferred from analysis of rbcL, atpB and 18S rDNA sequences; the jackknife consensus tree (for groups receiving >50% support) is shown. under eurosids II includes genus Acer. (Soltis et al., 2000)

38 2.2 Materials and Methods

2.21 Standard chemicals

Pelargonidin (CAS Number: 134-04-3 Catalog Number: 020912S Chemical Name:

PELARGONIDIN CHLORIDE with HPLC), cyanidin (CAS Number: 528-58-5 Catalog

Number: 020909S Chemical Name: CYANIDIN CHLORIDE with HPLC), and delphinidin

(CAS Number: 528-53-0 Catalog Number: 020904S Chemical Name: DELPHINIDIN

CHLORIDE with HPLC) standards were purchased from Indofine Company (Hillsborough,

NJ) and stored at -80°C before use. Peonidin-3-glucoside chloride (CAS Number: 6906-39-4

Catalog Number: 42008 Chemical Name: Peonidin 3-O-glucoside chloride BioChemika, ≥

97.0% HPLC) and keracyanin chloride (CAS Number: 18719-76-1 Catalog Number: 36428

Chemical Name: Keracyanin chloride purum, ≥98.0% HPLC Synonym: Cyanidin-3-O- rutinoside chloride) were purchased from Sigma (Louis, MO). Catechin, epicatechin,

procyanidin B1 and procyanidin B2 from Sigma were also purchased and used in this study.

2.2.2 Plant materials

Six varieties of A. palmatum used for this study are A. palmatum 'Hubb's Red Willow', A. palmatum 'Shaina', A. palmatum 'Hagoromo', A. palmatum Dissectum Atropurpureum, A. palmatum 'Okushimo' and A. palmatum 'Oridono nishiki'. All plants are growing in the JC

39 Raulston Arboretum, at North Carolina State University, Raleigh, North Carolina. The properties of height and foliar pigmentation are briefly summarized in tables 2 and 3. All cultivars have palmate opposite leaves.

These six varieties represent three distinguished leaf pigmentation patterns during ontogeny.

Hereafter, we define the three patterns as red-green (RG), red (R) and green (G).

Table 2. A summary shows height, environment and location of 6 cultivars selected in this work. *Locations for Acer are bedding numbers where those maples are planted in JC Raulston Arboretum, as a part of the Department of Horticulture Science at North Carolina State University. Numbers after scientific name of each cultivar are reference numbers to be used for the sake of convenience. Botanical Name (#Reference) Common Name Height Environment Location* purple Not fully exposed to A. palmatum 'Hubb's Red Willow'-1 narrowleaf ~1m S03 light, partially shaded Japanese maple dwarf purple-leaf Not fully exposed to A. palmatum 'Shaina'-2 ~0.8m Su7 Japanese maple light, partially shaded angel's dress A. palmatum 'Hagoromo'-3 ~2m Fully exposed to light D30 Japanese maple A. palmatum Dissectum red lace-leaf ~1.8m Fully exposed to light D01 Atropurpureum-4 Japanese maple tube-leaf Not fully exposed to A. palmatum 'Okushimo'-5 ~1.6m E35 Japanese maple light, partially shaded Variegated A. palmatum 'Oridono nishiki'-6 chameleon ~2m Fully exposed to light W20 Japanese maple

The RG pattern is characterized by a pigmentation of leaves from red to green color during ontogeny of leaves. The newly developing leaves are red due to the accumulation of anthocyanins. During the expansions of leaves, the red color is gradually replaced by green

40 color. The fully expanded mature leaves are phenotypically less red or green. Unshaded leaves of A. palmatum 'Hubb's Red Willow' and A. palmatum 'Shaina' have the RG pattern

(Table 3). All of leaves on A. palmatum 'Hagoromo' have this pattern (Table 3).

The R pattern is featured by an obvious red pigmentation through the whole ontogeny of leaves. Anthocyanin pigmentation in immature and mature stages of leaves is phenotypically similar. A. palmatum Dissectum Atropurpureum has this R pattern (Table 3).

Table 3. List of cultivars selected in this work. *Developmental sequence on the twig ‐ their sample numbers in the experiment. Numbers after scientific name of each cultivar are reference numbers to be used for the sake of convenience. Pigmentation Botanical Name (#Reference) Pattern of Collection of leaf pairs * leaves RG 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P-4), 5th(P-5), 6th(P-6), 7th(P-7) A. palmatum 'Hubb's Red Willow'-1 VRG Mixture of shaded leaves(SH-L)

RG 1st(P-1), 2nd(P-2), 3rd(P-3) A. palmatum 'Shaina'-2 VRG Mixture of shaded leaves(SH-L) 1st(P-1), 2nd(P-2), 3rd(P-3), 5th(P- A. palmatum 'Hagoromo'-3 RG 5), 7th(P-7) 1st(P-1), 2nd(P-2), 3rd(P-3), 4th A. palmatum Dissectum Atropurpureum-4 R (P-4), 5th(P-5) 1st(P-1), 2nd(P-2), 3rd(P-3), 5th(P- A. palmatum 'Okushimo'-5 G 5), 7th(P-7), 9th(P-9) A. palmatum 'Oridono nishiki'-6 G 1st(P-1), 2nd(P-2)

41 The G pattern is characterized by an obvious green pigmentation during ontogeny of leaves.

From immature to mature stages of leaves, the color is constitutively green. Sometimes, the

newly developing young leaves may have inconspicuously red coloration. A. palmatum

'Okushimo' and A. palmatum 'Oridono nishiki' have this G pattern (Table 3).

In addition, there is the forth pattern. This pattern is characterized by a variable red and green

coloration (VRG). This VRG pattern mainly occurs in shaded leaves inside the crown or

bush. The shaded young leaves are not as red as unshaded leaves exposed to sunlight. Most

of the time, the coloration of shaded leaves is green. Shaded leaves of both A. palmatum

'Hubb's Red Willow' and A. palmatum 'Shaina' have this VRG pattern.

2.2.3 Leaf collection

Early stage, immature (not fully expanded) and mature (fully expanded) leaves from the twig

were collected from plants growing at JC Raulston Arboretum, North Carolina State

University, Raleigh NC from late April to early May, 2008. No petioles were included.

Leaves collected for this experiment were healthy without damage caused by pathogens and insects. Leaf materials were immediately frozen in liquid nitrogen after excision from twigs

and then stored in -80°C degree freezer until further use.

42 Leaf samples collected from different nodes of a twig were carefully labeled to show their

spatial location and relative developmental stages. Pairs of palmate leaves are oppositely

growing on twigs of six A. palmatum Thunb. cultivars. Each new twig developed from those

cultivars in spring had numerous nodes, each of which had a pair of palmate leaves. The node

immediately next to the shoot apex meristem of twigs had a pair of the youngest leaves. In

this experiment, we numerated this node as node 1 and the pair of leaves as pair 1 (P-1). In

turn, after this node, other leaf pairs on the same twig are numerated as pair 2, 3, 4, 5, 6, 7, 8

etc (P-2 ~ P-8) in order. Therefore, pair 1 represents the youngest leaves. The bigger of the

number was, the more mature the leaves were.

We collected unshaded leaves from at least 5 twigs from one single plant and then pooled

those leaves from the same numerated node (e.g. node 1) together in order to obtain enough material for each node. All pair 1 leaves collected from the node 1 were termed as P-1. In turn, other sample leaves were termed as P-2, P-3, P-4 and P-5 etc (Table 4).

Shaded leaves (VRG) from the two cultivars 'Hubb's Red Willow'-1 and 'Shaina'-2 were collected as controls. We collected 5 pairs of leaves from each twig and pooled all pairs of leaves together as one single sample, termed as SH-L. Shaded leaves are very important controls in this study due to several reasons: 1) shaded leaves have similar phenotype to mature leaves, thus can be used for confirmation with mature leaves; 2) shaded leaves have

43 relatively less exposure to light, thus can be used in comparison to non-shaded leaves; 3)

shaded leaves are one unique group of leaves which should be included in our study.

The cultivar A. palmatum 'Oridono nishiki' was different from the other 5 cultivars. We noticed that most of new twigs developed from late spring were not healthy and lacked in many nodes. Thus we only collected leaves based on size. Only two sizes of leaves were collected in spite of their original positions on the twigs, big size as pair 1 (P-1) and small size as pair 2 (P-2). Both pairs are mixed leaf samples, thus can be considered to have similar composition.

2.2.4 Methods

Extraction and measurement of anthocyanins

Samples were homogenized into fine powder in liquid nitrogen in a mortar. One hundred mg

of fine powder was weighed into a 1.5ml centrifuge tube frozen in liquid nitrogen. Two

hundred and fifty μl of 0.1% HCl in methanol (100%) used as the extraction solvent was

added into each tube. The tubes were vigorously vortexed followed by 15 minutes of

sonication, and then further kept at 4°C in a refrigerator overnight. The tubes were

centrifuged at 10,000 rpm for 30 minutes at 4°C and the supernatant containing anthocyanins

was transferred into clean centrifuge tubes. The pellet was suspended in 250 ml of the same

44 extraction solution and this extraction was repeated once. The second supernatant was

combined with the first one. Thus, for each sample, the final extraction volume was 0.5 ml.

In order to measure levels of total anthocyanins, chlorophyll was removed from the extracts.

The 0.5 ml extract was dried by evaporating methanol from the tubes in a speed vacuum at room temperature. The pellet was re-suspended in 0.5 ml of 0.1% HCl-metanol : MiniQ H2O

(1:1). One hundred µl of chloroform was added into the tubes and then the mixture was

vigorously vortexed, followed by centrifuging the tubes at 10,000rpm for 10 minutes at 4°C.

The upper phase of methanol-H2O containing anthocyanins was transferred to a new tube.

The step was repeated once. The methanol-H2O phase of 200 μl was pipetted into a new tube,

which was stored at -20oC for measurement of anthocyanins.

Anthocyanin absorbance (ABS) values were recorded at the wavelength of 530nm on a UV-

spectrometer (Heλios-Y,Thermo Spectronic). 0.1% HCl methanol-H2O solution was used as

a background control. The volume of extracts used for ABS measurement was 0.1 ml. The

ABS value of 100 mg fresh weight sample was used to estimate the level of anthocyanins due

to lacking standards of an anthocyanin mixture.

Hydrolysis of anthocyanins, measurement of ABS and identification of anthocyanidins

Fifty µl of the methanol-H2O extraction was added into 450 µl of solvent consisting of n-

butanol (100%): HCl (36%) (95:5, v/v) contained in a 1.5 ml polypropylene centrifuge tube.

45 The mixture was vigorously vortexed and 0.1 ml out of the volume was used to measure an

ABS value at 550nm on the same UV spectrometer as described above. The background control used was 0.1ml of the n-butanol: HCl mixture. This ABS value obtained from measurement was designated as ABS1 (pre-hydrolysis ABS) for samples prior to hydrolysis.

The remaining 0.4ml of the mixture was then boiled for 1 hr and then cooled to room temperature. One hundred µl of the boiled mixture was used to measure a second ABS value, designated as ABS2 (post hydrolysis ABS) for samples after hydrolysis. The difference of

ABS (ΔABS) was obtained by subtracting ABS2 with ABS1. The background control used was also 0.1ml of the n-butanol: HCl mixture.

Hydrolyzed samples of 0.3ml were then dried by means of a speed vacuum at room temperature. The residue was dissolved in 100 µl of 0.1% HCl methanol (0.1% HCl in LC–

MS grade methanol) and centrifuged at a speed of 12,000rpm for 10 min. The supernatant was transferred into a new clean tube for identification of core anthocyanidins through LC-

UV-ESI-MS analysis described below. The volume of injection was 20ul. Standards included pelargonidin, cyanidin, delphinidin, peoni-din-3-glucoside chloride, and keracyanin chloride(cyanidin-3-o-rutinoside).

Extraction and analysis of proanthocyanidins

One hundred mg of fresh homogenized tissue was weighed into a 1.5mL centrifuge tube for the extraction of metabolites by adding 500µl acetone deionized water (70:30). For each

46 sample, triplicates were extracted. The tubes were vigorously vortexed and sonicated for 15

minutes followed by being kept at 4°C in a refrigerator for one hour. The tubes were then

centrifuged at 10,000 rpm for 30 minutes at 4°C and the supernatant was transferred into

clean centrifuge tubes. This extraction step was repeated once by suspending the pellets in another 500µl of 70% acetone. These two acetone extractions were combined into the same tube and then dried at room temperature by using a continuous flow of nitrogen gas. The residues were suspended in 500µl autoclaved MiniQ water. Then, 100µl of chloroform was added into the tubes followed by vigorously mixing the tubes. Tubes were centrifuged at

10,000 rpm for 10 min at 4°C. The lower chloroform phase was pipetted out and deposited into a waste container. This step was repeated once by adding another 100µl of chloroform to eliminate most chlorophyll and other lypophilic compounds. The remained water phase and interphase were extracted by adding 500µl ethyl acetate into the tubes. The tubes were vigorously vortexed to mix the two solvents completely and then centrifuged at 10,000 rpm for 5 min. The supernatant ethyl acetate phase was pipetted into a new clean centrifuge tube.

This ethyl acetate extraction step was repeated once. These two ethyl acetate extractions were combined in one tube and dried by using nitrogen gas. The residue was suspended in 250µl methanol (LC–MS grade) and the tube was centrifuged at a speed of 12,000 rpm for 10 min.

The supernatant was transferred into a new clean tube and was stored at -20°C until use.

Twenty µl of the methanol extract was used for identification of proanthocyanidin molecules through LC-UV-ESI-MS analysis described below. Authentic standards used as positive control included catechin, epicatechin, procyanidin B1 and B2.

47 Butanol-HCl hydrolysis of proanthocyanidins produces anthocyanidins. This method is

commonly used to measure proanthocyanidins at 550nm (Porter, 1989). Fifty µl of the

methanol extract was mixed with 450 ul of a mixture solvent consisting of n-butanol (100%):

HCl (36%) (95:5, v/v) in a 1.5 ml centrifuge tube. The tube was vigorously vortexed and then

boiled for 1hr. After boiled samples were cooled to room temperature, they were dried by

means of a continuous flow of nitrogen gas. The remained residue was suspended in 100µl of

0.1% HCl methanol (0.1% HCl in LC–MS grade methanol) and the tubes were centrifuged at

a speed of 12,000 rpm for 10min. The supernatant was transferred into a new clean tube and stored at 20°C for LC-UV-ESI-MS analysis described below. For each sample, the volume of

injection was 20μl. Authentic standards used as positive controls included pelargonidin, cyanidin and delphinidin.

LC-MS analysis

The analyses of anthocyanins, anthocyanidins, flavan-3-ols and proanthocyanidins were carried out by using high performance liquid chromatography/mass spectrometry on 2010EV

LC/UV/ESI /MS instrument (Shimadzu) (Zhou et al., 2008). The analytical separation column used for this experiment was the reversed phase Eclipse XDB-C18 (250 mm x 4.6 mm, 5 μm, Agilent). Two mobile phase solvents used to elute metabolites were 1% acetic acid in water (solvent A) (HPLC grade acetic acid and LC-MS grade water) and 100% acetonitrile (solvent B) (LC-MS grade). A gradient solvent program was designed to separate metabolites. The program was composed of ratios of solvent A to B, 90:10 (0-5 min), 90: 10

48 to 88: 12 (5-10 min), 88: 12 to 80: 20 (10-20 min), 80: 20 to 75: 25 (20-30 min), 75: 25 to 65:

35 (30-35 min), 65: 35 to 60: 40 (35-40 min), 60: 40 to 50: 50 (40-55 min), 50: 50 to 10: 90

(55-60 min), then followed by 10 min washing using 10% solvent B. The flow rate was 0.4

ml·min-1 and the injection volume for each was 20μl. The UV spectrum was recorded

from190 nm to 800 nm. The total ion chromatograms were recorded from 0 to 60 minutes by

using MS detection with positive electrospray ionization and MS spectrum was scanned and

stored from m/z of 120 to 1600 at speed of 1000 amu per second. A wavelength channel set to 530nm was used to detect, qualify and quantify anthocyanin and anthocyanidin compounds. The other wavelength channel at 280nm was used to detect, qualify and quantify flavan-3-ols and proanthocyanidin compounds.

Histological localization of proanthocyanidins

Cellular localization of anthocyanins was examined by microscopy. Fresh leaves were crossly dissected by hand. Cross sections of leaves were mounted on a slide for microscopic examination.

Fully expanded leaves were collected in July 2009 and then washed with deionized water.

Half of the leaves were cut into small pieces. Both incised leaves and intact leaves were completely immersed in 50ml of chloroform : methanol (2:1, V/V) for 30 min at room

temperature. This step removed surface wax from leaves (Bakker et al., 1998). Wax-removed

leaves were stained by freshly prepared 0.1% (w/v) dimethylaminocinnamaldehyde

49 (DMACA) dissolved in ethanol : 6M HCl (1:1) for 30 minutes (Porter, 1989; Xie et al.,

2003). Leaf materials were then washed three times with deionized water and then extracted

with acetic acid: ethanol (1:3, V/V) to remove chlorophyll. Acetic: ethanol was then replaced

by deionized water. In addition, leaves without staining in DMACA solution and leaves

staining in DMACA (-) solution were used as controls.

Statistical analysis

Statistical analysis for ABS of anthocyanins, anthocyanidins and peak area values of LC and

MS chromatograms from different samples were performed using the one-way ANOVA F-

test and the student t-test by JMP 7 (NCSU MLA license).

2.3 RESULTS

2.3.1 Analyses of anthocyanins and proanthocyanidins in leaves of Hubb’s Red Willow

Difference of pigmentation in shaded and unshaded leaves

A. palmatum 'Hubb's Red Willow' has two types of leaf pigmentation patterns on the same

tree. One type of red pigmentation phenotype is from red gradually to green on unshaded leaves. This type is termed as red-to-green (RG) pattern hereafter. This RG type leaves are characterized by high anthocyanin pigmentation during the early stage (unexpanded fully) of their development; then gradual loss of the red pigmentation during their expansion; finally

50 complete loss of red pigmentation when fully expanded (Figure 7A). The other type of

pigmentation is variable red and green pigmentation on shaded leaves (Figure 7A). This type

is termed as VRG.

Absorbance measurement of total anthocyanins

The levels of anthocyanins were significantly higher in first four pairs of leaves (1st, 2nd, 3rd and 4th) than in the 5th, 6th and 7th pairs of leaves (Figure 7). The level of anthocyanins was significantly higher in the 3rd pair of leaves than other pairs of leaves. The level of

anthocyanins in the 3rd pair of leaves was 5 times higher than that in the 7th pair of leaves.

From the 4th pair of leaves, the levels of anthocyanins started to reduce dramatically (P<0.05).

The ABS unit in shaded leaves was the lowest.

51

Fig 7. Leaf phenotype of A. palmatum 'Hubb's Red Willow' and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A’) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. Sample marked with asterisk doesn’t have enough replicates hence no error bar nor statistic analysis. (Figure continues on next page.)

52

53 Absorbance measurement of proanthocyanidins

The presence of proanthocyanidins in the shaded leaves and three pairs of unshaded leaves

(2nd, 3rd, and 5th) was examined by butanol-HCl hydrolysis and absorbance analysis. After the

methanol extracts were boiled in butanol-HCl for 1 hour, not only extracts from highly red

pigmented leaves produced red color in butanol-HCl solvent, but also extracts from dark

green leaves including both shaded and unshaded ones released red pigments into solvent.

Significant increases of ABS values were observed for all five analyzed samples (Figure 7C

and Table 4). From the 2nd to 5th pairs of leaves, the delta-ABS values of extracts dramatically increased. The ABS values of the extracts from the shaded leaves and the 5th unshaded ones were increased approximately 13 and 2.8 folds respectively (Figure 7 and

Table 4). These results indicated that proanthocyanidins were produced in leaves.

Table 4. Ratio of ABS values (ABS2/ABS1) of hydrolyzed samples. Ratio is calculated between average ABS2 and ABS1 of leaf samples from six cultivars of Acer palmatum Thunb. Letters after the cultivar reference number in the column of cultivar refer to four different coloration patterns mentioned before. No data for sample #4 or #6 for all cultivars. NA means samples not analyzed. A. palmatum 'Hubb's Red Willow'‐1; A. palmatum 'Shaina'‐2; A. palmatum 'Hagoromo'‐3; A. palmatum Dissectum Atropurpureum‐4; A. palmatum 'Okushimo'‐5; A. palmatum 'Oridono nishiki'‐6. Cultivar SH-L 1 2 3 5 7 9 1 (RG) 12.6 NA 1.3 1.4 2.8 NA NA 2 (RG) 11.4 1.4 NA 2.1 NA NA NA 3 (RG) NA 2.0 NA 6.2 NA 20.1 NA 4 (R) NA 1.4 NA NA 1.7 NA NA 5 (G) NA 41.5 NA NA NA NA 58.9 6 (G) NA 145.2 NA NA NA NA NA

LC-MS analysis of anthocyanins and anthocyanidins

54 We used LC-MS to analyze anthocyanins and proanthocyanidins in the 1st, 2nd, 3rd, and 5th pairs of leaves and shaded leaves (SH-L). Anthocyanins peaks were recorded at 530 nm.

Seven major anthocyanin peaks are labeled (Figure 8). Anthocyanin peaks 1 and 2 were present in these four pairs of unshaded leaves.

Obvious differences in anthocyanin profiles were observed in these five samples analyzed.

The chromatographic profiles for the 1st pair of leaves consisted of three major peaks 1, 2,

and 3. Anthocyanin peak profiles in the 2nd and 3rd pairs of leaves were similar, consisting of

1, 2, and 4. Major anthocyanin peak profiles in the 5th pair of leaves consisted of 1, 2, and 4.

In addition, three small peaks were observed from the 5th pair of leaves and marked as peak 5,

6 and 7(Figure 8D). Only a small peak 1 was observed from shaded leaves.

The height and area values of peak 2 dramatically decreased from the 1st to 5th pairs of leaves.

These values of peak 1 were similar in the 1st, 2nd and 3rd pairs of leaves, and then showed an obvious decrease trend from the 3rd to 5th pairs of leaves (Figure 8).

55

Fig 8. LC chromatogram of A. palmatum 'Hubb's Red Willow' . Major anthocyanin peaks were marked with different numbers. Only peak 1 and 2 are universally present in all cultivars with red pigmentation. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 3rd pair of leaves; D: the 5th pair of leaves; E: shaded leaves. Numerals 1‐7 are used to label major peaks detected at 530nm. Total peak area values are listed at right bottom corner of each chromatogram.

56 We performed LC-MS analysis to characterize these anthocyanins peaks. Positive electron

spay was used to ionize these anthocyanins. We found that peak 1 was composed of two

anthocyanins, which were separated by liquid chromatography. Their mass was 449 and 595

[m/z]+ respectively. Based on an anthocyanin library we developed and previous reports

(Chang and Giannasi, 1991; Delendick, 1990; Fossen and Andersen, 1999; Ishikura, 1976; Ji

et al., 1992a; Ji et al., 1992b), we predicted that they were cyanidin-3-glucoside and

cyanidin-3-rutinoside. Peak 2 also consisted of two anthocyanin molecules. Their mass was

601 and 747 [m/z]+ respectively. These two anthocyanins were predicted to be cyanidin 3-O-

[2’’-O-(galloyl)-β-D-glucoside] and cyanidin 3-O-[2’’-O-(galloyl)-6’’-O-(α-L-rhamnosyl)-β-

D-glucoside] (Figure 9).

A complete hydrolysis of anthocyanins via boiling extracts in butanol-HCl released anthocyanidins. LC-MS analysis showed cyanidin as the main core anthocyanidin produced in leaves (Figure 10). In addition, the area peak values of cyanidin were compared for extracts from the 2nd, 3rd and 5th pairs of leaves and shaded leaves. There was an obvious

decreasing trend of cyanidin levels from the 2nd to 5th pairs of leaves (Figure 10), which

positively related the trends of total ABS values (Figure 7) and total peak area values (Figure

8) of anthocyanins.

57

Fig 9. Major anthocyanins of A. palmatum 'Hubb's Red Willow'. A: Four selected ion chromatograms show ion fragments of 449, 595, 601 and 747. B: Four mass spectra show ion fragments of 449, 595, 601 and 747. The specific m/z is enclosed in a rectangle. The anthocyanin profiling was obtained from LC‐MS. In the chromatogram, in order to align four peaks representing four ions together, the scale for the heights of the peaks (m/z=601, 747) is magnified five‐fold and six‐fold respectively. A1 and A2 refer to two major peaks in the LC chromatogram.

LC-MS analysis of proanthocyanidins

We used a reverse-phase column to separate and a positive electron spray to ionize proanthocyanidins. This LC-MS analysis showed the presence of one monomer, one dimer, and one trimer. Based on standard samples, we identified that the monomer was epicatechin characterized by a mass of 291 [m/z]+; the dimer was procyanidin B2 with a

58

Fig 10. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Hubb's Red Willow'. A: the 2nd pair of leaves; B: the 3rd pair of leaves; C: the 5th pair of leaves; D: the shaded leaves. Numbers listed beside peak cyn are average values of peak area from triplicates. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1and An 2.

59 mass of 579 [m/z]+. The trimer has a mass of 867 [m/z]+, which is predicted to be a trimer

(Figure 11).

Fig 11. Major proanthocyanidins of A. palmatum 'Hubb's Red Willow'. A: Three selected ion chromatograms show ion fragments of 291, 579 and 867. B: Mass spectrum shows ion fragment of 291. C: Mass spectrum shows ion fragment of 579. D: Mass spectrum shows ion fragment of 867. The specific m/z is enclosed in a rectangle. The proanthocyanidin profiling was obtained from LC‐MS. In the chromatogram, in order to align peaks representing three ions together, the scale for the heights of the peak (m/z=867) is magnified fivefold.

60 Relative levels of these three proanthocyanidins in extracts from five pairs of leaves respectively were compared. The levels of proanthocyanidins were expressed with peak area values obtained from mass chromatogram analysis. A trend line for each compound level change and total sum change was charted (Figure 12) and such change will be further discussed later.

Fig 12. Trend of proanthocyanidins level change of A. palmatum 'Hubb's Red Willow'. Numbers below x‐axis refer to sample numbers. Average values ± standard deviation are presented.

61 Butanol-HCl hydrolysis was carried out via boiling ethyl acetate extracts of proanthocyanidins for one hr. LC-MS analysis identified cyanidin as the main compound from this hydrolysis (Figure 13). This result indicated that the extension units of proanthocyanidins in leaves were mainly composed of epicatechin or catechin.

Fig 13. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Hubb's Red Willow'. A: LC chromatogram of anthocyanidins. B: LC chromatogram of anthocyanidin controls. (cyn = cyanidin, pg = pelargonidin, dp = delphinidin)

Histological staining with DMACA

Cellular localization of proanthocyanidins was analyzed by staining fully expanded mature

leaves collected in summer with DMACA. This reagent reacts with proanthocyanidins to

62 provide blue coloration, which allow both visional and microscopic observations. We found that wounded sites of petioles and blades showed considerable blue coloration (Figure 14).

Fig 14. Histological staining of proanthocyanidins in a mature leaf of A. palmatum 'Hubb's Red Willow’. A: Leaf without staining as control; B: Leaf stained with DMACA(‐) solution; C: Leaf stained with DMACA(+) solution. D: Close view of incised cuts stained with DMACA(+) solution. Arrows indicate blue color.

63

Fig 15. Microscopic images showing histological staining of proanthocyanidins in a mature green leaf of A. palmatum 'Hubb's Red Willow’. A: Leaf without staining as control; B: Leaf stained with DMACA(‐) solution; C: Leaf stained with DMACA(+) solution.

Microscopic images were taken for hand-crossed leaf samples. In young red leaves, anthocyanins were mostly localized in palisade parenchyma cells while proanthocyanidins were shown to be present in both palisade parenchyma and spongy mesophyll cells (Figure

16). However, no anthocyanins or proanthocyanidins were found to be localized in epidermal cells. In mature green leaves, little anthocyanins were found in the whole cross section.

Proanthocyanidins were mostly present in spongy mesophyll rather than palisade parenchyma (Figure 15). No anthocyanins or proanthocyanidins were found to be localized in epidermal cells either.

64

Fig 16. Microscopic images showing histological staining of proanthocyanidins in a young red leaf of A. palmatum 'Hubb's Red Willow’. A and B: Leaf without staining as control; C and D: Leaf stained with DMACA(‐) solution; E and F: Leaf stained with DMACA(+) solution.

65 2.3.2 Analysis of anthocyanins and proanthocyanidins in leaves of Shaina

Difference of pigmentation in shaded and unshaded leaves

A. palmatum 'Shaina' has two types of leaf pigmentation patterns on the same tree as A.

palmatum 'Hubb's Red Willow'. The unshaded leaves are highly pigmented by red

anthocyanins in immature leaves. During leaf development and maturation, unshaded leaves

gradually lose this red pigmentation. Thus the unshaded leaves have a RG pigmentation

patter. The shaded leaves are green with limited red pigmentation (Figure 17A).

66

Fig 17. Leaf phenotype of A. palmatum 'Shaina' and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

67

68 Absorbance measurement of total anthocyanins

Levels of anthocyanins were significantly higher in the 1st and 2nd pairs of leaves than in the

3rd pairs of leaves (P<0.05). The ABS value of anthocyanins in extracts of the shaded leaves

(SH-L) was very low (Figure 17B), nearly 60 times less than that of in the 2nd pair (P-2) of

leaves (Figure 17B).

Absorbance measurement of proanthocyanidins

The presence of proanthocyanidins in the shaded leaves and two pairs of unshaded leaves (1st and 3rd) was examined by butanol-HCl hydrolysis and ABS measurement at 550 nm. After

the methanol extracts were boiled in butanol-HCl for 1 hr, significant increases of ABS

values were also observed for these samples (Figure 17C). The ABS values of the extracts

from the shaded and the 3rd pair of leaves were increased approximately 11 and 2.1 fold

respectively (Figure 17C and Table 4). These data indicated that proanthocyanidins were

produced in leaves.

LC-MS analysis of anthocyanins and anthocyanidins

Three major peaks (1, 2 and 3) of anthocyanins were observed in methanol extracts of the 1st,

3rd pairs of unshaded leaves and shaded leaves (SH-L) by LC analysis. The height and area values of peak 2 were significantly reduced during leaf maturation (Figure 18). The area value of peak 2 in the 3rd pairs of leaves was about 59 fold less than that in the 1st pairs of

69

Fig 18. LC chromatogram of A. palmatum 'Shaina'. Major anthocyanin peaks were marked with different numbers. A: the 1st pair of leaves; B: the 3rd pair of leaves; C: the shaded leaves. Only peak 1 and 2 are universally present in all cultivars with red pigmentation. Numerals 1‐9 are used to label major peaks detected at 530nm. Total peak area values are listed at right bottom corner of each chromatogram.

leaves. The levels of the peak 1 were similar in the 1st and 3rd leaves analyzed. It was interesting that five other peaks (peaks 4~9) of anthocyanins between retention times of 11-

17 min were observed in methanol extracts of the 3rd pair of leaves with increasing levels

70 compared to the 1st pair of leaves. Peaks 1 and 2 were also detected from methanol extracts of shaded leaves (Figure 18).

Fig 19. Major anthocyanins of A. palmatum 'Shaina'. A: Four selected ion chromatograms show ion fragments of 449, 595, 601 and 747. B: Four mass spectra show ion fragments of 449, 595, 601 and 747. The specific m/z is enclosed in a rectangle. The anthocyanin profiling was obtained from LC‐MS. A1 and A2 refer to two major peaks in the LC chromatogram.

MS analysis was carried out to characterize these anthocyanin peaks. Actually, the peak 1 consisted of two anthocyanins. Their masses were 449 and 595 [m/z]+. These two anthocyanins were predicted to be cyanidin 3-glucoside and cyanidin 3-rutinoside. Peak 2

71 also consisted of two anthocyanins characterized by two masses of 601 and 747 [m/z]+, which were predicated to be cyanidin 3-O-[2’’-O-(galloyl)-β-D-glucoside] and cyanidin 3-O-

[2’’-O-(galloyl)-6’’-O-(α-L-rhamnosyl)-β-D-glucoside] (Figure 19).

Butanol-HCl hydrolysis and LC-MS analysis identified cyanidin which formed the main core

structure of anthocyanins (Figure 20). In addition, two minor unidentified anthocyanidins

were found between retention times of 45-50 min. As leaves matured, the level of cyanidin

was significantly reduced. Peak area of cyanidin in the 3rd pair of leaves was reduced to only

60% of that in the 1st pair of leaves. In contrast, the levels of two unidentified anthocyanidins

were increased (Figure 20).

LC-MS analysis of proanthocyanidins

In order to demonstrate whether or not proanthocyanidins are produced in leaves of this

variety, we extracted proanthocyanidins and analyzed extracts as described above. Based on

authentic standards, LC-MS analysis identified epicatechin (m/z=291) and procyanidin B2

(m/z=579). In addition, a trimeric procyanidin with a mass of 867 [m/z]+ was found (Figure

21).

72

Fig 20. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Shaina'. A: the 1st pair of leaves; B: the 3rd pair of leaves; C: the shaded leaves. Numbers listed beside peak cyn are average values of peak area from triplicates. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1and An 2.

73

Fig 21. Major proanthocyanidins of A. palmatum 'Shaina'. A: Three selected ion chromatograms show ion fragments of 291, 579 and 867. B: Mass spectrum shows ion fragment of 291. C: Mass spectrum shows ion fragment of 579. D: Mass spectrum shows ion fragment of 867. The specific m/z is enclosed in a rectangle. The proanthocyanidin profiling was obtained from LC‐MS.

In order to characterize the extension units of proanthocyanidins, we carried out butanol-HCl hydrolysis of acetone-ethyl acetate extracts. LC-MS analysis identified cyanidin as the major compound produced from hydrolysis, indicating that the main extension units were either

74 epicatechin or catechin, which is consistent with hydrolysis data resulting from methanol extracts (Figure 20 and 22).

Area values of epicatechin, procyanidin, and the trimeric procyanidin were obtained from chromatography analysis. A trend of values for these three compounds was thus obtained to compare their abundance in each pair of leaves and shaded leaves (Figure 23).

Fig 22. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Shaina'. A: LC chromatogram of anthocyanidins. B: LC chromatogram of anthocyanidin controls. (cyn = cyanidin, pg = pelargonidin, dp = delphinidin)

75

Fig 23. Trend of proanthocyanidins level change of A. palmatum 'Shaina'. Average values ± standard deviation are presented. Numbers below x‐axis refer to sample numbers.

Histological staining with DMACA

In order to further demonstrate the presence of PAs leaves, fully expanded mature leaves

collected in summer were stained with DMACA, which react with PAs to form blue coloration in tissues. Petioles and wounded sites of leaves showed strong blue coloration,

demonstrating the existence of PAs (Figure 24).

76

Fig 24. Histological staining of proanthocyanidins in a mature leaf of A. palmatum 'Shaina'. A: Stained leaf and unstained leaf as control; B: Leaf stained with DMACA (‐) solution; C: Leaf stained with DMACA (+) solution. D: Close view of incised cuts stained with DMACA(+) solution. Arrows indicate blue color.

2.3.3 Analysis of anthocyanins and proanthocyanidins in leaves of Hagoromo

Red pigmentation and anthocyanin levels

A. palmatum 'Hagoromo' has only RG pigmentation pattern on leaves. Young leaves are red,

whereas fully expanded mature leaves are green. We observed that the loss of red

pigmentation in leaves was a gradual process during ontogeny from early stage to full

77 maturation. We chose five pairs of leaves (1st, 2nd, 3rd, 5th, and 7th) collected from several

twigs to characterize this development-dependant change (Figure 25A). A clear trend was obtained to show that the levels of anthocyanins significantly decreased from the 1st pair to the 7th pair of leaves. The level of anthocyanins in the 1st pair of leaves was 6 times higher

than in the 7th pair of leaves, which hardly show red pigmentation (Figure 25A).

Absorbance measurement of proanthocyanidins

The presence of proanthocyanidins in three pairs of leaves (1st, 3rd, and 7th) was examined by

butanol-HCl hydrolysis and absorbance analysis at 550 nm. After hydrolysis, significant

increases of ABS values were observed for all three samples (Figure 25C). The ABS values

of the extracts from two pairs of leaf samples (3rd and 7th) were increased approximately 2.1

and 11.4 fold respectively (Figure 25C and Table 4). The increase of ABS values resulted

from anthocyanidins produced from hydrolysis of proanthocyanidins. These data indicated

that proanthocyanidins were produced in leaves of this cultivar.

78

Fig 25. Leaf phenotype of A. palmatum 'Hagoromo' and measurement of anthocyanins and proanthocyanidin. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

79

80 LC-MS analysis of anthocyanins and anthocyanidins

Extracts of three pairs of leaves (1st, 3rd and 7th) were analyzed by LC-MS.

Fig 26. LC chromatogram of A. palmatum 'Hagoromo'. Major anthocyanin peaks were marked with different numbers. Only peak 1 and 2 are universally present in all cultivars with red pigmentation. A: the 1st pair of leave; B: the 3rd pair of leaves; C: the 7th pair of leaves. Numerals 1‐4 are used to label major peaks detected at 530nm. Total peak area values are listed at right bottom corner of each chromatogram.

81 LC chromatograms showed that height and area values of anthocyanins significantly

decreased from the 1st through 3rd and 7th pairs of leaves. Like in other analyzed cultivars, peak 1 and 2 were two dominant anthocyanins in this cultivar (Figure 26).

Fig 27. Major anthocyanins of A. palmatum 'Hagoromo'. A: Four selected ion chromatograms show ion fragments of 449, 595, 601 and 747. B: Four mass spectra show ion fragments of 449, 595, 601 and 747. The specific m/z is enclosed in a rectangle. The anthocyanin profiling was obtained from LC‐MS. A1 and A2 refer to two major peaks in the LC chromatogram.

ESI-MS analyses also identified that peak 1 of anthocyanins consisted of cyanidin 3-

glucoside, cyanidin 3-rutinoside. In addition, peak 2 of anthocyanins consisted of cyanidin 3-

82 O-[2’’-O-(galloyl)-β-D-glucoside] and cyanidin 3-O-[2’’-O-(galloyl)-6’’-O-(α-L- rhamnosyl)-β-D-glucoside] (Figure 27).

LC analysis showed different profiles of anthocyanidins from extracts of the three pairs of leaves. The main anthocyanidin in the 1st pair of leaves was cyanidin. In addition, the second

obvious peak of anthocyanidin was observed. In the 3rd pair of leaves, in addition to cyanidin,

there were 4 major additional peaks. In the 7th pair of leaves, cyanidin was not the main

anthocyanidins. In contrast, two other peaks of anthocyanidins showed higher peak area

values than cyanidin (Figure 28). Peaks around and after 45min retention time are unknown

due to lack of standards.

LC-MS analysis of proanthocyanidins

LC-MS analysis was performed to characterize monomers and dimers of proanthocyanidins.

Based on MS spectrum and our standards, we identified catechin (m/z=291), epicatechin

(m/z=291), and procyanidin B1 (m/z=579) (Figure 29). A trend line of each proanthocyanidin compound level change and total sum change was also charted (Figure 30).

83

Fig 28. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Hagoromo'. A: the 1st pair of leaves; B: the 3rd pair of leaves; C: the 7th pair of leaves. Numbers listed beside peak cyn are average values of peak area from triplicates. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1 to An10.

84

Fig 29. Major proanthocyanidins of A. palmatum 'Hagoromo'. A: Two selected ion chromatograms show ion fragments of 291 and 579. B: Mass spectrum shows ion fragment of 291 as epicatechin. C: Mass spectrum shows ion fragment of 291 as catechin. D: Mass spectrum shows ion fragment of 579. The specific m/z is enclosed in a rectangle. The proanthocyanidin profiling was obtained from LC‐MS.

85

Fig 30. Trend of proanthocyanidins level change of A. palmatum 'Hagoromo'. Average values ± standard deviation are presented. Numbers below x‐axis refer to sample numbers.

Just like previous cultivars, cyanidin was also found to be the only identifiable anthocyanidin converted from extension units of proanthocyanidins through hydrolysis, which is consistent with hydrolysis data resulting from methanol extracts (Figure 31).

86

Fig 31. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Hagoromo'. A: LC chromatogram of anthocyanidins. B: LC chromatogram of anthocyanidin controls. (cyn = cyanidin, pg = pelargonidin, dp = delphinidin)

Histological staining with DMACA

DMACA staining was also carried out to detect the presence of proanthocyanidins in this cultivar. Petioles and wounded sites and leaf blades show blue color (Figure 32).

87

Fig 32. Histological staining of proanthocyanidins in a mature leaf of A. palmatum 'Hagoromo'. A: Stained leaf and unstained leaf as control; B: Leaf stained with DMACA(‐) solution; C: Leaf stained with DMACA(+) solution. D: Close view of incised cuts stained with DMACA(+) solution. Arrows indicate blue color.

2.3.4 Analysis of anthocyanins and proanthocyanidins in leaves of Dissectum Atropurpureum

Property of foliar red pigmentation pattern

Shaded and unshaded leaves of A. palmatum Dissectum Atropurpureum are characterized by red/purple pigmentation through the whole ontogeny. Therefore, the pigmentation pattern belongs to “R” (red). In addition, this red pigmentation of leaves is similar from spring through fall (Figure 33A).

88 Absorbance measurement of total anthocyanins

Five pairs of leaves were collected from unshaded twigs and then were numerated as 1, 2, 3,

4 and 5 (Figure 33A). Levels of anthocyanins were measured by ABS analysis at 530nm and showed similar anthocyanin levels in these five pairs of leaves (Figure 33B).

Absorbance measurement of proanthocyanidins

Although there is no dramatic increase of ABS values in terms of increasing fold after butanol-HCl hydrolysis (Figure 33C, Table 4), these data still indicated that

proanthocyanidins were produced in leaves.

LC-MS analysis of anthocyanins and anthocyanidins

In order to investigate what contributes to the change of anthocyanin levels at different

developmental sequences, we selected the 1st and 5th samples for LC analysis.

HPLC analysis showed that anthocyanins extracts from these five pairs of leaves included one major peak labeled as 1 and a minor peak labeled as 2 (Figure 34). In terms of peak area, the values of these two peaks were very similar from the 1st pair of leaves through the 5th pair

of leaves.

89

Fig 33. Leaf phenotype of A. palmatum Dissectum Atropurpureum and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

90

91

Fig 34. LC chromatogram of A. palmatum Dissectum Atropurpureu. Major anthocyanin peaks were marked with different numbers. Only peak 1 and 2 are universally present in all cultivars with red pigmentation. A: the 1st pair of leave; B: the 5th pair of leaves. Numerals 1 and 2 are used to label major peaks detected at 530nm. Total peak area values are listed at right bottom corner of each chromatogram.

MS analysis identified that the anthocyanin peak 1 consisted of cyanidin 3-glucoside and cyanidin 3-rutinoside; the anthocyanin peak 2 included cyanidin 3-O-[2’’-O-(galloyl)-β-D- glucoside] and cyanidin 3-O-[2’’-O-(galloyl)-6’’-O-(α-L-rhamnosyl)-β-D-glucoside] (Figure

35).

92

Fig 35. Major anthocyanins of A. palmatum Dissectum Atropurpureum. A: Four selected ion chromatograms show ion fragments of 449, 595, 601 and 747. B: Four mass spectra show ion fragments of 449, 595, 601 and 747. The specific m/z is enclosed in a rectangle. The anthocyanin profiling was obtained from LC‐MS. In the chromatogram, in order to align four peaks representing four ions together, the scale for the heights of the peaks (m/z=601, 747) is magnified fivefold. A1 and A2 refer to two major peaks in the LC chromatogram.

Butanol-HCl hydrolysis was carried out to release core anthocyanidin molecules. LC-MS analysis showed cyanidin as the main anthocyanidin from hydrolysis. In addition, estimation of cyanidin peak area indicated that the levels of the compound were similar in these 5 pairs of leaves (Figure 36).

93

Fig 36. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of A. palmatum Dissectum Atropurpureum. A: the 1st pair of leaves; B: the 5th pair of leaves. Numbers listed beside peak cyn are average values of peak area from triplicates. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1and An 2.

LC-MS analysis of proanthocyanidins

The LC-MS analyses of proanthocyanidins extracts identified epicatechin (m/z=291) and procyanidin B2 (m/z=579) (Figure 37). Their abundance is quite similar to each other. A

94 trend line of those two compounds’ level change and total sum change was charted (Figure

38).

Fig 37. Major proanthocyanidins of A. palmatum Dissectum Atropurpureum. A: Two selected ion chromatograms show ion fragments of 291 and 579. B: Mass spectrum shows ion fragment of 291. C: Mass spectrum shows ion fragment of 579. The specific m/z is enclosed in a rectangle. The proanthocyanidin profiling was obtained from LC‐MS.

95

Fig 38. Trend of proanthocyanidins level change of A. palmatum Dissectum Atropurpureum. Average values ± standard deviation are presented. Numbers below x‐axis refer to sample numbers.

Butanol-HCl hydrolysis of proanthocyanidins was carried out to characterize molecular

properties. LC-MS analysis identified cyanidin as the major compound, indicating that the extension units of proanthocyanidins were either catechin or epicatechin (Figure 39).

96

Fig 39. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum Dissectum Atropurpureum. A: LC chromatogram of anthocyanidins. B: LC chromatogram of anthocyanidin controls. (cyn = cyanidin, pg = pelargonidin, dp = delphinidin)

Histological staining with DMACA

DMACA straining of leaf tissues was carried out to histologically localize proanthocyanidins.

Fully expanded mature leaves collected in summer 2008 were stained with DMACA for visualization of proanthocyanidins. Petioles, mid-ribs and incised cuts showed blue color

(Figure 40).

97

Fig 40. Histological staining of proanthocyanidins in a mature leaf of A. palmatum Dissectum Atropurpureum. A: Stained leaf and unstained leaf as control; B: Leaf stained with DMACA(‐) solution; C: Leaf stained with DMACA(+) solution. D: Close view of incised cuts stained with DMACA(+) solution. Arrows indicate blue color.

98 2.3.5 Analysis of anthocyanins and proanthocyanidins in leaves of Okushimo

Green pigmentation

Leaves of A. palmatum ' Okushimo' have a pigmentation pattern “Green” (G). The leaf color during the whole ontology is greenish from early growth to fully mature stages. All leaf blades are hardly pigmented by red anthocyanins (Figure 41A).

Analysis of anthocyanins

Six pairs of leaves numerated as 1, 2, 3, 5, 7, and 9 were collected from several twigs. These six pairs of leaves were representing six stages of leaf ontology, from immature to mature; although they appear to be similar phenotypically (Figure 41A).

Absorbance values of methanol extractions from all pairs of leaves analyzed at 530 nm was very low (Figure 41B). In comparison, we found that ABS values of 7 and 9 pairs of leaves were higher than other pairs. In addition, HLPC analysis did not show anthocyanin peaks at

530 nm (Figure 42). These results indicate the blade of leaves don’t accumulate anthocyanins.

99

Fig 41. Leaf phenotype of A. palmatum 'Okushimo' and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

100

101

Fig 42. LC chromatogram of A. palmatum 'Okushimo'. A: the 1st pair of leave; B: the 9th pair of leaves. No anthocyanin peaks were detected at 530nm.

Butanol-HCl hydrolysis of extracts from 1st and 9th pairs of leaves was carried out to further

analyze anthocyanins. This hydrolysis released a large amount of red pigment in reagents.

Absorbance recorded at 550 nm showed that ABS values after hydrolysis increased

approximately 40 and 60 fold respectively as those prior to hydrolysis (Figure 41C and Table

4).

HPLC-MS analysis further showed that cyanidin was the main compound resulting from this

hydrolysis. In addition, six peaks of anthocyanidins were observed from chromatogram

recorded at 530 nm (Figure 43). These anthocyanidins were suggested from

proanthocyanidins extracted in methanol.

102

Fig 43. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Okushimo'. A: the 1st pair of leaves; B: the 9th pair of leaves. Numbers listed beside peak cyn are average values of peak area from triplicates. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1 to An 5.

LC-MS analysis of proanthocyanidins

LC-ESI-MS analysis identified epicatechin (m/z=291), catechin (m/z=291), and procyanidin

B2 (m/z=579) from acetone-ethyl acetate extracts of leaves (Figure 44). Epicatechin appeared to be the dominant monomer of proanthocyanidins. Total area of all present peaks

103 of interest was calculated and their development-dependent change can be referred to Figure

45.

Fig 44. Major proanthocyanidins of A. palmatum 'Okushimo'. A: Two selected ion chromatograms show ion fragments of 291 and 579. B: Mass spectrum shows ion fragment of 291 as epicatechin. C: Mass spectrum shows ion fragment of 291 as catechin. D: Mass spectrum shows ion fragment of 579 as procyanidin B2. The specific m/z is enclosed in a rectangle. The proanthocyanidin profiling was obtained from LC‐MS.

104

Fig 45. Trend of proanthocyanidins level change of A. palmatum 'Okushimo'. Numbers below x‐axis refer to sample numbers. Average values ± standard deviation are presented.

Butanol-HCl analysis was also carried out to demonstrate the presence of proanthocyanidins in leaves. HPLC analysis demonstrated that cyanidin was the main compound produced from hydrolysis, indicating procyanidin the major proanthocyanidins in leaves (Figure 46).

105

Fig 46. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Okushimo'. A: LC chromatogram of anthocyanidins. B: LC chromatogram of anthocyanidin controls. (cyn = cyanidin, pg = pelargonidin, dp = delphinidin)

Histological staining with DMACA

In order to further confirm the accumulation of proanthocyanidins, fully expanded mature leaves collected in summer were stained with DMACA for visualization of proanthocyanidins. Petioles showed strong blue coloration. Incised sites of blades also showed strong blue color (Figure 47).

106

Fig 47. Histological staining of proanthocyanidins in a mature leaf of A. palmatum ‘Okushimo’. A: Stained leaf and unstained leaf as control; B: Leaf stained with DMACA(‐) solution; C: Leaf stained with DMACA(+) solution. D: Close view of incised cuts stained with DMACA(+) solution. Arrows indicate blue color.

2.3.6 Analysis of anthocyanins and proanthocyanidins in leaves of Oridono nishiki

Green pigmentation

Leaves of A. palmatum 'Oridono nishiki' are greenish, but hardly pigmented during ontogeny

by red anthocyanins (Figure 48A). We collected two sizes of leaves from different position

nodes of twigs. These two pairs of leaves are mixture of leaves even though their sizes are

107 different (Figure 48A). Thus we can consider them to be the same samples representing this

cultivar as a whole.

Analysis of anthocyanins

Absorbance values of methanol extracts were recorded at 530 nm but were very low for these

two pairs of leaves. Statistical analysis showed difference between ABS values of extracts

from these two pairs of leaves (Figure 48B) (P<0.05).

LC-MS analysis was also carried out to determine whether or not anthocyanins were present in methanol extracts. We did not observe anthocyanins peaks recorded from 515-535 nm, indicating that no these red pigments were produced in blade (Figure 49).

P-1 sample was then hydrolyzed and significant increase of ABS values was detected (Figure

48C). The ABS value of the extract was increased approximately 145 fold (Figure 48C and

Table 4). These data also strongly indicated that proanthocyanidins were produced in leaves in this cultivar instead of anthocyanins.

108

Fig 48. Leaf phenotype of A. palmatum 'Oridono nishik' and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

109

110

Fig 49. LC chromatogram of A. palmatum 'Oridono nishik'. No anthocyanin peaks were detected at 530nm.

Since there is little anthocyanin present in this cultivar, no anthocyanin peaks were detected at 530nm. However, anthocyanidins were analyzed after hydrolysis. Cyanidin was also the major anthocyanidin in this cultivar (Figure 50). There were other five unidentifiable anthocyanidins at 530nm.

Fig 50. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of A. palmatum 'Oridono nishiki'. Numbers listed beside peak cyn are average values of peak area from triplicates. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1to An 5.

111

Fig 51. Major proanthocyanidins of A. palmatum 'Oridono nishiki'. A: Two selected ion chromatograms show ion fragments of 291 and 579. B: Mass spectrum shows ion fragment of 291 as epicatechin. C: Mass spectrum shows ion fragment of 291 as catechin. D: Mass spectrum shows ion fragment of 579 as B2. The specific m/z is enclosed in a rectangle. The proanthocyanidin profiling was obtained from LC‐MS.

LC-MS analysis of proanthocyanidins

The LC-MS analysis of acetone-ethyl acetate extracts of leaves identified of epicatechin

(m/z=291) catechin (m/z=291), and procyanidin B2 (m/z=579). Epicatechin appeared to be

112 the dominant monomers of proanthocyanidins in leaves (Figure 51). Total area of all present peaks of interest was calculated and their abundance was charted in Figure 52.

Fig 52. Trend of proanthocyanidins level change of A. palmatum 'Oridono nishiki'. Numbers below x‐axis refer to sample numbers.

Butanol:HCl hydrolysis of acetone-ethyl acetate extracts of leaves was carried out to characterize extension units of proanthocyanidins. The main red pigment compound produced from this hydrolysis was cyanidin (Figure 52). This result indicated that procyanidins were mainly produced in leaves.

113

Fig 53. Major anthocyanidins from hydrolyzed proanthocyanidins of A. palmatum 'Oridono nishiki'. A: LC chromatogram of anthocyanidins. B: LC chromatogram of anthocyanidin controls. (cyn = cyanidin, pg = pelargonidin, dp = delphinidin)

Histological staining of proanthocyanidins

In order to further confirm whether this cultivar produces and accumulates proanthocyanidins, fully expanded mature leaves collected in summer were stained with DMACA for visualization of proanthocyanidins. Petioles and incised cuts show considerable blue staining while leaf blades have obvious blue dots and patches (Figure 54).

114

Fig 54. Histological staining of proanthocyanidins in a mature leaf of A. palmatum ‘Oridono nishik’. A: Stained leaf and unstained leaf as control; B: Leaf stained with DMACA(‐) solution; C: Leaf stained with DMACA(+) solution. D: Close view of incised cuts stained with DMACA(+) solution. Arrows indicate blue color.

115 2.4 Discussions

2.4.1 Different pigmentation patterns in leaves of ornamental A. palmatum Thunb. cultivars are appropriate to investigate anthocyanin metabolism

Anthocyanins are one large group of natural pigments which give diverse pigmentation patterns in leaves, flower and fruits (Close and Beadle, 2003; Holton and Cornish, 1995;

Winkel-Shirley, 2001). The formation of anthocyanins has been proposed to provide numerous advantageous ecological functions for plants to tolerate multiple stresses, e.g. absorbance of strong irradiative light, drought and low temperature, etc. (Asada, 1999;

Chalker-Scott, 1999, 2002; Close and Beadle, 2003; Dominy et al., 2002; Feild et al., 2001;

Gould, 2004; Hoch et al., 2001; Lee, 2002a, b; Lev-Yadun and Gould, 2007; Manetas, 2006;

Ougham et al., 2005; Yamazaki, 2008). Anthocyanins are also natural anti-oxidative compounds which may reduce oxidative damage to plants (Castaneda-Ovando et al., 2009;

Kong et al., 2003). The biosynthetic pathway of anthocyanins has been intensively investigated and all of the pathway genes have been cloned from numerous plant species

(Dooner et al., 1991; Grotewold, 2006; Holton and Cornish, 1995; Springob et al., 2003).

However, little has been understood on the development-dependent formation and loss of foliar anthocyanins in flowering plants. The formation, accumulation, and catabolism of anthocyanins in plant tissues have been strongly suggested to be closely associated with

116 development, environmental changes, and auto-oxidative or enzymatic degradation (Oren-

Shamir, 2009).

I used six ornamental cultivars to understand their metabolic difference of anthocyanins, which can directly change pigmentation patterns of leaves. Acer species are of ornamental significance due to their beautiful leaf morphology and red coloration in fall (Le Hardÿ de

Beaulieu, 2003; van Gelderen et al., 1994; Vertrees, 1978). Firstly, six A. palmatum cultivars are featured by four types of foliar pigmentation patterns (Table 4) and are grown in JC

Raulston Arboretum (NCSU). Secondly, these six cultivars resulted from genetic breeding and their pigmentation patterns are stable (van Gelderen et al., 1994). Therefore, they are appropriate materials to compare anthocyanin profiles, which provide red pigmentation in leaves as maturation.

In addition, these cultivars are appropriate to understand metabolic mechanisms of how and why leaves switch on or off the biosynthesis of anthocyanins as they mature. As shown in results and following up observation, the foliar pigmentation patterns (RG, R and G) of these cultivars are very stable, at least throughout 3 years in a row of our observation. Especially, the RG pattern is development-associated as maturation. Comparative analysis of these anthocyanin profiles and other flavonoids at different developmental stages can provide metabolic evidence to show whether or not RG patterns result from metabolic channeling to different metabolites.

117 2.4.2 Anthocyanin level corresponds to leaf coloration pattern

It is well understood that profiles of secondary metabolites of plants vary with different seasons, growing locations and developmental stages (Wink, 1999). The presence or absence of anthocyanins in leaves is an important physiological mark to define phase changes of leaf and development of plants (Hughes et al., 2007; Schmitzer et al., 2009; Steyn et al., 2004;

Vaknin et al., 2005). A common phenomenon of leaf pigmentation pattern changes is that

red/pink/purple coloration of juvenile leaves fades as maturation in many plants including woody Acer species (Hughes et al., 2007; Lee and Collins, 2001; Price and Sturgess, 1938).

Absorbance measurement and HPLC analysis clearly demonstrated that the levels of total anthocyanins formed the foundation of leaf pigmentation patterns. Furthermore, levels of

anthocyanins in RG patterns were strongly leaf development-dependent. The RG type of

unshaded leaves in both A. palmatum 'Hubb's Red Willow' and A. palmatum 'Shaina'

produced high levels of anthocyanins during early stages of leaf development; however, these leaves reduced levels of the red pigment during maturation (Figure 7 and 17). Levels of anthocyanins in A. palmatum 'Hagoromo' showed a clear decreasing trend of red pigmentation as leaves matured (Figure 25). When leaves were completed fully expanded and mature, levels of anthocyanins were very low. Leaves of A. palmatum Dissectum

Atropurpureum were always red and featured by a red pigmentation pattern as matured.

Levels of anthocyanins at different development stages of leaf development were similar

118 (Figure 33). Green leaves of A. palmatum 'Okushimo' and A. palmatum 'Oridono nishiki'

either produced trace levels of anthocyanins or did not make anthocyanins at all (Figure 41 and 48).

2.4.3 Development-dependent properties of anthocyanin pigmentation patterns in Acer

cultivars

A. palmatum 'Hubb's Red Willow' and A. palmatum 'Shaina' have two types of foliar

anthocyanin pigmentation patterns, RG in unshaded leaves and VRG in shaded leaves.

Developmental dependence of anthocyanin coloration was obvious in unshaded leaves of

both cultivars (Figure 7A and Figure 17A).

For A. palmatum 'Hubb's Red Willow', an increasing trend of anthocyanin levels from the first pair of leaves (the youngest leaves) through the 3rd pair of leaves. We observed that the

levels of anthocyanins in 3rd pairs of leaves was higher than in the 1st and 2nd pairs of leaves

(P<0.05) (Figure 7B). From the 4th pair of leaves, the levels of anthocyanins dramatically

decreased as leaves matured. HPLC analysis also showed decrease of area values of major

peaks (Figure 8). The 1st pair of leaves was visually redder that the 3rd pair of leaves. This

might be due to less chlorophyll pigment in the youngest leaves. We also observed this

phenotype in other species. A similar trend of anthocyanin levels were obtained in A.

palmatum 'Shaina'.

119 A. palmatum 'Hagoromo' also showed an obvious decrease trend of red pigmentation from

young leaves to mature leaves (Figure 25A). And mature leaves, starting from the 3rd sample, turn to green with little trace of red pigmentation. Consistently, a very clear decreasing trend of anthocyanin level shows up based upon spectrometric analysis (Figure 25B). Moreover, there is no sign of increase of anthocyanin level before decrease like A. palmatum 'Hubb's

Red Willow'. Anthocyanin level continues to decrease from the very young leaves to more mature ones, and at the meanwhile leaves seem greener possibly due to maturation and development of chlorophylls (Figure 25A).

A. palmatum Dissectum Atropurpureum has only red leaves and those leaves have similar coloration and phenotype. Thus, similar anthocyanin levels for all samples are expected

(Figure 33B). Leaves of the 5th pair are very similar to the rest of leaves, although they are

not shown in the picture (Figure 33A).

A. palmatum 'Okushimo' has only green leaves with inconspicuous or little red coloration

(Figure 41A). Those leaves have similar coloration and phenotype. For A. palmatum

'Oridono nishiki', since leaves are not collected as the other cultivars, but based on size instead, the samples from this cultivar can not be compared to others for development- dependent coloration pattern. However, it is feasible to consider the 1st and 2nd pairs of leaves

as the same mixture of A. palmatum 'Oridono nishiki' leaves because they were collected

from many different developmental sequences on twigs and pooled together. And based on

120 visual observation, most healthy leaves on A. palmatum 'Oridono nishiki' are green without

evident red pigmentation (Figure 48A). Anthocyanin levels for both cultivars tell the same

story. ABS values, approximately equivalent to anthocyanin levels, are no greater than 0.20,

although some of those samples have significant differences in anthocyanin levels among

each other (Figure 41B and Figure 48B). Such low values mean no anthocyanins in the

sample since they are even lower than ABS values of self-shaded leaves A. palmatum

'Hubb's Red Willow' and A. palmatum 'Shaina' (Figure 7B and Figure 17B).

To sum up, we find out that there is a development-dependent trend of anthocyanin level

change, as well as leaf coloration pattern change for some cultivars (1, 2, and 3) in our

research. For cultivars 4 and 5, development-dependent trend of leaf coloration pattern and anthocyanin level still exist, but in a way of either invariable accumulation or lack of anthocyanins. However, whether or not anthocyanins are present or the content level of anthocyanins is changed, the anthocyanin level corresponds well to leaf coloration pattern.

Such dramatic discrepancy of leaf pigmentation patterns and anthocyanin contents among

those cultivars are very likely due to manual selection, which results in mutations in

regulatory and/or structural genes in anthocyanin/proanthocyanidin pathway, hence alteration

in metabolic profiling of those compounds.

121 2.4.4 The formation of proanthocyanidin leads to less red pigmentation

Proanthocyanidins, which share most biosynthetic steps as well as substrates with anthocyanins, were recently discovered to be closely related to anthocyanidins, aglycones of

anthocyanins through the enzymatic reaction of anthocyanidin reductase in vivo (Xie et al.,

2003). Epi-falvan-3-ols (especially 2,3-cis-(-)epicatechin), as monomers of proanthocyanidins, which serve as predominant components for extension unit as well as a common starter unit (Xie and Dixon, 2005), are derived from anthocyanidins through enzymatic reduction. Proanthocyanidins, just like anthocyanins, play an important role in the interactions between plants and their environment, most likely as deterrents against herbivore

(Peters and Constabel, 2002). Such deterrent traits can protect plants against microbial

pathogens, insects, pests and larger herbivores (Dixon et al., 2005).

However, the relationship between anthocyanins and proanthocyanidins and impacts of such

relationship on the pigmentation pattern of leaves in plants in response to development has

not attracted more attention since the revelation of enzymatic role of anthocyanidin reductase

(Xie et al., 2003). Let alone the fact that most studies have been conducted on autumn

senescing leaves rather than spring developing leaves.

Since leaf coloration pattern shows a clear development-dependent trend, we selected several

samples from each cultivar to investigate both anthocyanin and proanthocyanidin level

122 through hot acid hydrolysis previously described (Porter, 1989; Xie et al., 2003). ABS of

post-hydrolyzed samples should be higher than that of pre-hydrolyzed samples due to the

conversion of certain compounds, including proanthocyanidins (if any), to anthocyanidins,

which are aglycones of anthocyanins. Another reason for the increase of ABS at 550nm after

hydrolysis may directly result from the conversion of anthocyanins to anthocyanidins, which

according to previous research, would lead to a bathochromic shift of maximum ABS from a

short wavelength to a longer wavelength, closer to 550nm (de Freitas and Mateus, 2006).

In order to estimate proanthocyanidin contents, P-2, P-3, P-5 and SH-L samples from cultivar

1, P-1, P-3 and SH-L samples from cultivar 2, P-1, P-3 and P-7 samples from cultivar 3, P-1

and P-5 samples from cultivar 4, P-1 and P-9 samples from cultivar 5, and P-1 sample from

cultivar 6 were selected for hydrolysis and following analysis. Seemingly, proanthocyanidin level, unlike anthocyanin level, has little direct indication of leaf pigmentation pattern, and

vice versa (Figure 7, 17, 25, 33, 41 and 48 C). The proanthocyanidin levels for selected

samples were estimated through subtraction of ABS1 (pre-) from ABS2 (post-) of leaf

samples prior to and after hydrolysis. Since the estimation of proanthocyanidin levels come

from hydrolyzed samples (with or without anthocyanins), we relate proanthocyanidin level to

anthocyanin level to see whether there is any correlation among those two groups of

flavonoid compounds in terms of leaf pigmentation pattern.

123

Fig 55. Trend of anthocyanin and proanthocyanidin level change. Average values ± standard deviation are presented. Values (ABS units/reaction) are calculated from differences between ABS values of pre‐ and post (post‐ minus pre‐) hydrolysis of samples from six cultivars of Acer palmatum Thunb. Dots in diamond represent ABS of pre‐ hydrolysis samples, dots in square ABS of post‐ hydrolysis samples and dots in triangle ABS of estimated proanthocyanidin level. Numbers below x‐axis refer to cultivar reference number and #sample. 1‐0 and 2‐0 refer to SH‐L samples. A: A. palmatum 'Hubb's Red Willow'‐1; B: A. palmatum 'Shaina'‐ 2; C: A. palmatum 'Hagoromo'‐3; D: A. palmatum Dissectum Atropurpureum‐4; E: A. palmatum 'Okushimo'‐5; F: A. palmatum 'Oridono nishiki'‐6.

124 From our data, we can clearly see that proanthocyanidin levels increase toward maturation, in

most cases, the opposite trend of anthocyanin levels (Figure 7, 17, 25, 33, 41, 48 B and C),

which corresponds to the decrease of red coloration in leaf phenotype. This phenomenon is

true for all cultivars 1-5 if shaded SH-L sample is not taken into account. Even for cultivar 1-

A. palmatum 'Hubb's Red Willow' , SH-L sample still has the maximum average ABS (i.e. highest level of proanthocyanidin), which is not statistically different from the 5th pair of leaves (P-5) though (also the highest level of proanthocyanidin), while SH-L sample has the lowest anthocyanin level among all samples in cultivar 1 (Figure 7B). For cultivar 2- A. palmatum 'Shaina', SH-L sample has very low level of proanthocyanidin based on our data.

However, the ratio of ABS values (SH-L sample) is actually the highest among three selected samples for cultivar 2 (Table 4). Considering the precision of the spectrometer being used, this ratio is probably not that accurate. Even though, the ratio of ABS values still reflects the exactly same increasing trend of proanthocyanidin contents, just like the estimated proanthocyanidin levels. Figure 55 shows a more straightforward picture of such relationship between contents level of proanthocyanidin/anthocyanin and leaf pigmentation pattern. We found out that anthocyanin levels corresponds well to leaf coloration, which in other words, show an obvious development-dependent trend, in most cases (Figure 55: A~D blue lines), a gradual decreasing one. However, on the contrary, the development-dependent trend of proanthocyanidin level change is exactly the opposite (Figure 55: A~B green lines). During the process of maturation, young leaves with high level of anthocyanins tend to reduce anthocyanin contents while increasing proanthocyanidin contents towards mature leaves.

125 Mature leaves with relatively lower level of anthocyanins tend to have high levels of

proanthocyanidin compared to their younger counterparts (Figure 55). This concomitant

change of anthocyanin and proanthocyanidin levels was also observed from a transgenic

tobacco with ectopic expression of a Medicago BAN, leading to accumulation of

proanthocyanidins with reduction in anthocyanin levels (Xie et al., 2003). This phenomenon

may result from the competition of limited substrates between anthocyanin and

proanthocyanidin biosynthesis. Nevertheless, proanthocyanidin levels alone can not tell whether leaves have more or less red pigmentation. Since proanthocyanidins and anthocyanins share the same precursors (i.e. anthocyanidins), a competition for substrates may occur between the production of colorless proanthocyanidins and colored anthocyanins.

It is anthocyanin level that connects proanthocyanidin level to leaf coloration pattern. High

level of proanthocyanidins, along with low level of anthocyanins, leads to less red

pigmentation within one cultivar.

In order to determine the extension units of proanthocyanidins and also approximately

quantify proanthocyanidin contents, butanol-HCl hydrolysis (Porter, 1989) was carried out

for both proanthocyanidin acetone-ethyl acetate extracts and anthocyanin methanol extracts.

Our data showed that cyanidin was the only identifiable anthocyanidin converted from

extension units of proanthocyanidins through hydrolysis, which is consistent with hydrolysis

data resulting from methanol extracts (Figure 13, 22, 31, 39, 46 and 53). Such consistency

indicates that only cyanidin-derived compounds (including anthocyanins and/or

126 proanthocyanidins) were present in our samples. Moreover, contents level change of cyanidin derived from both proanthocyanidins and anthocyanin extracts were charted in Appendix B for comparison. As for cultivar 1~4, change of cyanidin content levels is consistent with that of anthocyanin content levels (Appendix B: A~D) and cyanidin content levels are positively correlated to anthocyanin content levels, which corresponds to leaf pigmentation pattern, while for cultivar 5, cyanidin contents are consistent with proanthocyanidin contents instead of anthocyanin (Appendix B: E) given the fact that there is trace amount of anthocyanins.

However, based on hydrolysis results from proanthocyanidin extracts, development- dependent patterns are less obvious, even equivocal to previous data. Such ambiguity could be possibly due to the reason that only one replicate has been tested. Nevertheless, such data still suggests that increase of ABS values after hydrolysis may mostly come from anthocyanidins derived from anthocyanins rather than anthocyanidins converted from proanthocyanidin extension units, especially for cultivar 1, 2 and 4 which possess a large amount of anthocyanins (Appendix B: A, B and D). In other words, even though proanthocyanidins contents are produced and accumulated differently in abundance according to different stages of development, hydrolysis of proanthocyanidins will not have huge impact on the cyanidin level change no matter what (Appendix B: F, G and I) because anthocyanins contribute to most of the cyanidin production (Appendix B: A, B and D) while only extension units of proanthocyanidins would be converted to corresponding anthocyanidins (Porter, 1989). This indication is further supported by the dramatic different order of magnitude of cyanidin production from hydrolysis of methanol and acetone-ethyl

127 acetate extracts (Appendix B: y axis scale of A, B, D, F, G and I) and also the coherent

relationship between anthocyanin level and cyanidin level (Figure 9, 19, 27 and 353). A. palmatum 'Okushimo' and A. palmatum 'Oridono nishiki' (G) also has cyanidin as major anthocyanidin although both cultivars have trace amount of anthocyanin contents. However,

levels of cyanidin (Figure 42 and 49) correspond to proanthocyanidin levels. Thus, those high levels of cyanidin should come from the conversion of proanthocyanidin due to hot acid hydrolysis. Those results suggest that production and accumulation of proanthocyanidin would lead to relatively less red pigmentation within one plant.

2.4.5 Qualitative and quantitative changes contribute to total anthocyanin level change

In order to investigate what contributes to the change of anthocyanin levels at different

developmental sequences, we selected the 1st, 2nd, 3rd, 5th and SH-L samples from cultivar 1,

1st, 3rd and SH-L samples from cultivar 2, 1st, 3rd and 7th samples from cultivar 3, 1st and 5th samples from cultivar 4 for LC analysis. Samples from cultivars 5 and 6 were also included although there were no detectable anthocyanin peaks at 530nm due to their low anthocyanin levels. LC chromatography showed that the alteration of anthocyanin levels resulted from either qualitative or quantitative change or both, while qualitative changes probably stem from structural changes such as modification or degradation, for which we do not have data to further investigate.

128 LC chromatogram of almost all cultivars (1-4) samples showed two identifiable major peaks, marked uniformly as peak 1 and peak 2 in all LC figures (Figure 8, 18, 26 and 34). Peaks marked with numbers other than 1 and 2 have no comparability among samples of different cultivars. Although most samples possess peak 1 and 2, their relative abundance is different from each other. Some samples have peak 1 as the dominant anthocyanin peak while others have both as dominant peaks. Peaks marked with other numbers except for 1 and 2 and peaks observable from LC chromatogram but not being marked are unidentifiable in our research.

The development-dependent trend of anthocyanin levels of A. palmatum 'Hubb's Red Willow'

(RG) overall is gradual decreasing. From the 2nd sample to 3rd and 5th samples, with the developmental maturation process going on, anthocyanin level is decreasing basipetally, from the young pair of leaves at the apex of a twig to the mature pair of leaves below. Major peaks, peak 1 and 2 represent the dominant anthocyanins in this cultivar when the leaf samples have the highest anthocyanin level. However, peak 1 and 2 are not decreasing at the same pace (Figure 8). They contribute differently to the decrease of total anthocyanin levels which is impacted by development. Such decrease is a quantitative alteration. In addition, there are some unidentified peaks forming up and disappearing later including peak 3 and peak 4. Those peaks also contribute to the alteration of anthocyanin contents and it is defined as qualitative change. SH-L sample has only a tiny peak 1, which corresponds appropriately with the green leaf coloration due to lack of anthocyanin.

129 Alteration of anthocyanin level in cultivar A. palmatum 'Shaina' (RG) has a very similar pattern with that in A. palmatum 'Hubb's Red Willow'. There is a development-dependent decreasing trend of anthocyanin levels as well. The dominant anthocyanins in this cultivar are also anthocyanins represented by peak 1 and peak 2 (Figure 18). Quantitative and qualitative changes are also the major cause for the decreasing level of anthocyanins towards maturation. Abundance of peak 2 decreases much faster and dramatically than peak 1, which contributes more to the reduction of anthocyanin level. There is also emergence and disappearance of unidentified minor peaks, which is counted as qualitative change of anthocyanin level. For SH-L sample, both major peaks, peak 1 and 2 have dropped to a very low level, which reflects the pattern of leaf coloration quite well.

A. palmatum 'Hagoromo' (RG), has two major peaks, 1 and 2. However, for this cultivar, alteration of anthocyanin change is rather due to quantitative change than qualitative change

(Figure 26). There is no new minor peak showing up during the process of development.

Only abundance of all peaks is continuously decreasing and finally leads to the disappearance of peak 3 and 4, and leaves peak 1 and 2 at a limited level, which corresponds to the phenotype of mature green leaf without trace of red coloration.

A. palmatum Dissectum Atropurpureum (R), on the contrary, is very different from the three cultivars mentioned above. Since anthocyanin levels keep stable along development and maturation, the major peaks, peak 1 and 2 doesn’t have much change (Figure 34). Thus for

130 this cultivar, the development-dependent trend of anthocyanin level is actually constitutively

stable. Neither obvious quantitative nor qualitative alteration is detected.

From hydrolyzed data, only cyanidin was found in all cultivars, indicating that this group of cultivars, even original A. palmatum, may not have the capability to produce other anthocyanidins except cyanidin. Those findings are consistent with previous reports

(Delendick, 1990; Ji et al., 1992a; Schmitzer et al., 2009; Sibley et al., 1999). Some unidentifiable anthocyanin-like and anthocyanidin-like compounds from LC-MS (Channel

530nm) analysis were also found in our study as previously reported (Ji et al., 1992b).

Based on LC-MS analysis and calculation of peak area, a chart for abundance of anthocyanin

peak 1 and 2 in samples analyzed in each cultivar is made and compiled together for

comparison (Appendix A.). For cultivars 1-4, a development-dependent decreasing trend of

both peak 1 and 2 is quite obvious, although the slope of such decrease is different among

each other cultivar and even among two peaks (Appendix A). Abundance of peak 1 and 2

does not always share the same decreasing magnitude, thus peak 1 and 2 might not

corresponds equally to the same developmental regulation. However, the ultimate outcome

for both peaks is to reduce their abundance to trace or zero amount in cultivars 1-3. As for

cultivar 4, since the leaves show similar red coloration in spring with disregard of

development, peak 1 and 2 are present the whole time, with peak 1 as the predominant peak

131 and peak 2 with far less abundance. Even though, a slight decreasing trend of abundance for both peaks is still detectable (Appendix A Chart 4).

Thus, along with LC-MS data of anthocyanins, we conclude that all cultivars with obvious red pigmentations have four species of anthocyanins: cyanidin 3-glucoside, cyanidin 3- rutinoside, cyanidin 3-O-[2’’-O-(galloyl)-β-D-glucoside] (i.e. cyanidin 3-galloylglucoside) and cyanidin 3-O-[2’’-O-(galloyl)-6’’-O-(α-L-rhamnosyl)-β-D-glucoside] (i.e. cyanidin 3- galloylrutinoside), which form up two major peaks detected in LC chromatogram. Among different samples of different cultivars, the abundance of those anthocyanins in terms of peak area is altered in accordance with total anthocyanin level change and leaf coloration pattern in a development-dependent way, although in cultivar 4, total anthocyanins level tends to stay stable. Such development-dependent reduction of anthocyanin contents results from both quantitative change of major peaks 1 and 2 and also from qualitative change of some unidentifiable peaks in LC chromatogram.

2.4.6 Levels of proanthocyanidins change in response to development

In order to confirm that proanthocyanidins are produced and accumulated in maple leaves in spring, we extracted proanthocyanidins (if there are any) and analyzed extracts qualitatively and quantitatively through two ways. One is to identify proanthocyanidin constituents (if there are any) in our samples with LC-MS, and the other is to hydrolyze proanthocyanidins

132 (if there are any) in hot acid butanol to characterize their hydrolysates as anthocyanidins in order to confirm with previous hot acid hydrolysis of methanol extracts. Such data has not been reported from A. palmatum Thunb. cultivars to our best knowledge.

The LC-MS analyses of proanthocyanidins (including flavan-3-ols) showed three major proanthocyanidin compounds (m/z=291, 579 and 867) almost present in all our samples.

They are speculated as epicatechin (m/z=291), procyanidin B2 (m/z=579) and a trimer

(m/z=867). Catechin, which is a stereoisomeric counterpart of epicatechin, derived from leucoanthocyanidins instead of anthocyanidins (Tanner et al., 2003), was detected in only half of the cultivars (3, 5 and 6). This discrepancy is really surprising even though changes of metabolic profiling resulting from manual selection and cultivation of ornamental plants are expected. However, due to the unavailability of wild type A. palmatum plant, we are unable

to determine which hypothesis might be true, either the production of (epi)catechin as a gain-

of-function or as a loss-of-function in different cultivars compared to the wild type. Although the assumption of the wild type being able to produce both stereo-isomers of flavan-3-ols would be the most parsimonious evolution situation in this case and long-term cultivation of

A. palmatum leads to loss-of-function of producing catechin in certain cultivars, we hardly

have any solid evidence to prove that. Even though, such data still suggests that plant

metabolism is as likely to be changed through manual selection as natural selection. Among

all the proanthocyanidins and flavan-3-ols, epicatechin and/or procyanidin B2 appear to be

the dominant proanthocyanidins in most cases. Thus, abundance of epicatechin and

133 procyanidin B2 contributes to most of the total proanthocyanidin contents we can determine

and characterize from leaf samples.

In order to study the alteration of proanthocyanidin levels according to development, mass peak area for each determined compound (m/z: epicatechin-291, catechin-291, Procyanidin

B2-579 and trimer-867) was calculated automatically and checked manually. Total area of all

present peaks of interest was calculated. Due to the fact that standard for trimer is not

available in our study, only cultivar 1 and 2 were considered to possess this trimer based on

mass spectrum and retention time (see Figure 56A and B). A trend line of each compound

level change and total summation change was charted and compiled for all cultivars for the

sake of comparison (Figure 56). Within each cultivar, the development-dependent trends for

all detectible single proanthocyanidin (including flavan-3-ols) and the total amount are

surprisingly coherent, unlike the discrepant change levels of peak 1 and 2 in anthocyanin

chromatograph (Appendix A). Such consistency may suggest that those proanthocyanidins

detected in our study should be regulated by similar mechanism as a response to development,

which, given the fact that all of them are derived from proanthocyanidin biosynthetic

pathway, is quite reasonable.

134

Fig 56. Trend of proanthocyanidin level change. Average values ± standard deviation are presented (except F). Lines in dark blue, red, violet and green refer to mass peak area of epicatechin, Porcyanidin B2, catechin and trimer respectively. Lines in light blue refer to total proanthocyanidin level by adding up all single mass peak area values of interest. Only lines representing detectible compounds are shown. Numbers below x‐ axis refer to cultivar reference number and sample number. 1‐0 and 2‐0 refer to SH‐L samples. A: A. palmatum 'Hubb's Red Willow'‐1; B: A. palmatum 'Shaina'‐2; C: A. palmatum 'Hagoromo'‐3; D: A. palmatum Dissectum Atropurpureum‐4; E: A. palmatum 'Okushimo'‐5; F: A. palmatum 'Oridono nishiki'‐6.

135 2.4.7 Anthocyanidin biosynthetic pathway is always activated

Along the progress to clarify anthocyanidin biosynthetic pathway, dozens of methods have been developed and applied to detect anthocyanins, anthocyanidins and proanthocyanidins.

Robinson and Robinson (1931) insisted on the efficacy of boiling for greater reliance in

detecting “anthocyanins” without any genetic information (Robinson and Robinson, 1931).

Although some researchers questioned Robinson’s method (Delendick, 1990), determination of anthocyanidins from extracts through boiling is a classic method to date because boiling with hot acid (butanol/HCl assay) will produce anthocyanidins from anthocyanins and proanthocyanidins (Dalzell and Kerven, 1998). Very few researchers realized the importance of boiling due to their assumption that no anthocyanins present equals to no anthocyanidins being produced in the whole biosynthetic pathway, or even inactivation of the anthocyanidin pathway. However, with the recent discovery of ANR/BAN gene encoding anthocyanidin reductase, which converts anthocyanidins to their corresponding 2,3-cis-flavan-3-ols (Xie et al., 2003), it became evident that a complete functional anthocyanidin biosynthetic pathway is required for a plant to produce proanthocyanidins consisting of 2,3-cis-flavan-3-ols (e.g. epicatechin and procyanidin B2) derived from anthocyanidins by means of anthocyanidin reductase (Xie et al., 2006). Thus, boiling is the right choice to detect proanthocyanidins as well as anthocyanidin biosynthetic pathway.

In this study, we tested both anthocyanins and proanthocyanidins, terminal products from the flavonoid biosynthetic pathway, and found out that the anthocyanidin biosynthetic pathway is

136 always activated even though there is trace/little amount of anthocyanins detectable. In the

cases of cultivar 5 and 6, those two cultivars have green foliar coloration pattern with almost

no detectable anthocyanins. However, large amounts of proanthocyanidins can be extracted

from both cultivars, which can be detected through “boiling” experiment for their conversion

to corresponding anthocyanidins. Although Delendick (1990) brought up the possible false positive conversion of anthocyanidins from leucoanthocyanidins under hot acid hydrolysis

(Delendick, 1990), there is no success in isolating and detecting leucocyanidins, leucopelargonidins and leucodelphinidins from proanthocyanidin-producing plants so far

(Xie and Dixon, 2005). Thus anthocyanidins (i.e. , and delphinidins)

which were produced from boiling must come from proanthocyanidins which indicates that

proanthocyanidin biosynthesis, which is a branch pathway of anthocyanin biosynthesis (Xie

and Dixon, 2005), is activated and further suggests that the whole anthocyanidin biosynthetic pathway, at least to the step of anthocyanidin production is activated so as to provide

substrates for the synthesis and accumulation of proanthocyanidins in plants. Similar

situation occurs in mature green leaves in cultivars 1-3. Those leaf samples show green

coloration pattern with a trace amount of anthocyanins. However, the proanthocyanidin

content in those leaf samples (except SH-L sample from cultivar 2) are among the highest,

which suggests that the anthocyanidin biosynthetic pathway, along with proanthocyanidin

pathway are highly activated and functional. As for the exception (Figure 17B), it is quite

likely that shaded leaves (SH-L) shut down the whole anthocyanidin pathway along with

proanthocyanidin pathway, leading to both low levels of anthocyanins and proanthocyanidins.

137 The shaded leaves in cultivar 1, contrarily, have a highly activated anthocyanidin biosynthetic pathway, which totally leads to the production of proanthocyanidins though

(Figure 7). The question is, if anthocyanidin biosynthetic pathway is activated and functional, why do the leaves not show red pigmentation instead of proanthocyanidin accumulation?

2.4.8 Is proanthocyanidin formation a consequence of metabolic channeling resulting from genetic breeding?

Metabolic channeling was proposed to interpret that multiple proteins might form a complex to biosynthesize plant natural products (e.g. flavonoids) in plants (Stafford, 1974). Several reviews discussed metabolic channeling in plant cells (Koes et al., 2005; Stafford, 1991;

Winkel-Shirley, 2001). Plant natural products (e.g. anthocyanins) are synthesized through multiple steps catalyzed by numerous enzymes. Metabolic channeling is considered a defensive strategy of plants to protect against autonomous self-toxic compounds. In addition, a protein complex may be formed to efficiently utilize extremely unstable intermediates to form final products of certain pathways. The difficulty to isolate natural flavan-3,4-diols, e.g. leucopelargonidin, leucocyanidin and leucodelphinidin, which are intermediates of proanthocyanidins and anthocyanins (Dixon et al., 2005; Haslam, 1975), may be due to the presence of such a metabolic channeling in plant cells. Furthermore, metabolic channeling is involved in multiple branch pathways, e.g. the flavonoid pathway (including anthocyanin and

138 proanthocyanidin pathways), which compete for substrates thus balance metabolic fluxes in

cells (Winkel-Shirley, 2001).

In this study, I found the relevance of the reduction of anthocyanins levels versus the

formation of proanthocyanidins in RG pattern leaves. Green leaves without red pigmentation

in our research but still producing and accumulating proanthocyanidins can serve as evidence

to support the concept of multienzyme complex. For cultivar 5 and 6, leaf samples are always

green without detectable anthocyanins by LC-MS (Figure 42 and 49). However,

proanthocyanidins in both samples are very high, and in cultivar 6, the 1st sample (Figure

48C) actually has the highest amount of proanthocyanidins among all samples, with

corresponding high level of cyanidin after hydrolysis as well. Such high levels of

proanthocyanidin with no detectable anthocyanin indicate that all the substrates of the anthocyanidin pathway are channeled to the proanthocyanidin pathway, resulting in the

production of proanthocyanidins and lack of anthocyanin accumulation. There is another

possibility that due to manual selection and cultivation, genes involved in anthocyanin

biosynthesis are degeneratively mutated hence loss of functions in leaves. But even though

this is the case, it fails to explain how proanthocyanidins (including flavan-3-ols) are

produced whereas anthocyanins are not, considering the fact that proanthocyanidin pathway

is a branch pathway and takes anthocyanidins as substrates for biosynthesis (Dixon et al.,

2005; Xie and Dixon, 2005; Xie et al., 2003; Xie et al., 2006). Previous studies also showed

that mutations in anthocyanidin synthase (ANS/LDOX) would result in a deficiency in

139 proanthocyanidin accumulation in Arabidopsis (Abrahams et al., 2003; Abrahams et al., 2002;

Xie et al., 2003). Thus there ought to be a high level of regulatory mechanism for flux control at the interface between the anthocyanin and proanthocyanidin pathways (Xie and Dixon,

2005).

Green mature leaf samples from cultivar 1-3 are different from green leaves of cultivar 5 and

6 because their younger counterpart leaves produce anthocyanins in high levels, which means green mature leaves in cultivar 1-3 should have the ability to produce anthocyanins, but somehow fail to do so. While for younger leaves from cultivar 1-3, very few produce proanthocyanidin in a large quantity at this early developmental stage, but with the development progressing, the proanthocyanidin contents increase concomitantly with the decrease of anthocyanin level, which suggests some alteration of metabolic channeling from mainly the production of anthocyanins, to both anthocyanins and proanthocyanidins in between, and finally to mainly proanthocyanidins in accordance with development. The model of multienzyme complexes (i.e. metabolon) in the biosynthesis of anthocyanins and proanthocyanidins reviewed recently (Jorgensen et al., 2005; Winkel, 2004) can be applied to explaining our results. Since the anthocyanidin pathway is activated in our leaf samples at different developmental stages, a switch of metabolic channeling by means of multienzyme complexes may occur, turning the production of anthocyanin to proanthocyanidins. Such development-dependent alteration may result from higher expression or more active BAN competing for anthocyanidin substrates, or from lower expression or less active 3-GT and

140 following enzymes for the final production of anthocyanins. It may also be simply due to the association of BAN to the hypothesized multienzyme during the process of development.

Thus, as for the leaf samples in cultivar 5 and 6, a tight association between the multienzyme complex of key enzymes involved in flavonoid biosynthesis and a highly active BAN competing for anthocyanidin substrates is present at all developmental stages. For leaf samples in cultivar 4, anthocyanins are more abundant than proanthocyanidins, indicating that metabolic channeling is more likely to occur for the production of anthocyanins instead of proanthocyanidins. This switch can be presumed as the underlying mechanism of development-dependent leaf coloration change and also the reason for the change of metabolic profiling of anthocyanins and proanthocyanidins in leaves. Considering the possible ecophysiological functions of anthocyanins and proanthocyanidins, there may be a gradual development-dependent function alteration from photoprotection as a sunlight filter

(anthocyanins) to defensive mechanism as deterrent compounds (proanthocyanidins) in developing leaves in spring.

2.4.9 Metabolic profiling of anthocyanins and proanthocyanidins may help establish a key to

A. palmatum Thunb. cultivars

Chemical data has been adopted for plant taxonomy for many years, termed as chemotaxonomy or chemosystematics. One recent review can be referred for the history and evolution of chemosystematics (Reynolds, 2007). Flavonoids, as well as other secondary

141 metabolites have been continuously used for the purpose of plant taxonomy up till now.

Some examples are listed here (Jensen et al., 1988; Nishiura et al., 1971; Nogata et al., 2006;

Williams, 1978; Williams et al., 1974). Anthocyanins, anthocyanidins and proanthocyanidins,

which are products from flavonoid biosynthetic pathway, serve as important chemical

markers for plant chemotaxonomy such as family Aceraceae (Chang and Giannasi, 1991;

Delendick, 1990; Ji et al., 1992b). In our study, due to limitation of our equipment and lack

of standards, we were only able to identify one anthocyanidin, four anthocyanins and four

proanthocyanidins in all. Thus, we establish a key to A. palmatum Thunb. cultivars based on dichotomy in terms of metabolic profiling of anthocyanins and proanthocyanidins (Appendix

D). Detailed reports of anthocyanin and proanthocyanidin for each cultivar can be referred to

Appendix E.

In summary, we propose that the metabolism of anthocyanins play an essential role in development-dependent leaf coloration changes. We used an integrated approach of phytochemistry, and metabolic profiling to determine the biosynthesis and metabolism of anthocyanins and their impacts on foliage color. Anthocyanins were extracted from different developmental stages of leaves for six Acer palmatum cultivars, which have obviously different accumulation patterns of anthocyanins in spring. Spectrometric analysis showed four major development-dependent trends of anthocyanin levels in leaves growing in spring:

1) gradual decrease for unshaded leaves and trace amount for self-shaded leaves; 2) gradual decrease; 3) constitutive accumulation; 4) lacking anthocyanins. HPLC-MS analysis showed

142 that the alteration of anthocyanin levels resulted from either qualitative or quantitative

change or from both changes, while qualitative changes probably stem from structure changes or modification or degradation. Proanthocyanidin analysis was carried out as well to determine their relationship with both anthocyanin production and foliar coloration. We have

found that even for green leaves with no/trace amount of detectable anthocyanins, the

biosynthetic pathway of anthocyanidin/proanthocyanidin is still activated. Epicatechin,

catechin, Procyanidin B2 and one unidentified timer are characterized from different

cultivars. And our results indicate that metabolic channeling directing the anthocyanidin

pathway to the proanthocyanidin biosynthesis plays a very important role in pigmentation

pattern change along developmental processes. Additionally, such metabolic channeling may

be a development-dependent response, steering the functions of those specific secondary

metabolites from photoprotection to defensive deterrents.

2.5 Conclusions and future prospects

Considerable progress has been made recently in our understanding of anthocyanin and

proanthocyanidin biosynthesis as well as their regulation. However, many important

questions remain to be answered concerning ecophysiological roles of anthocyanin and

proanthocyanidin in the interactions between plants and their environment, as well as their

roles in plant growth and development per se. Here we give some prospects for future studies

143 so that one day in the near future we can fully understand the underlying mechanisms of

regulatory biosynthesis of anthocyanin and proanthocyanidin for plants.

• Given the fact that most of work that has been done in our research is much more focused

on phytochemical and metabolic levels, researches on genetic and enzymatic/biochemical

levels should be conducted to prove our hypothesis.

• Structural and regulatory genes involved in anthocyanin/proanthocyanidin biosynthesis

should be cloned and characterized in maples. Their expression profile should be tested

so that the gene expression profile data can be related to phytochemical data.

• Crude mixture of enzymes from different maples should be extracted and enzymatic

experiment should be carried out to test their catalytic abilities of color change reaction

(i.e. anthocyanidin to flavan-3-ols etc.) as a protein complex.

• Samples from different seasons, such as summer and autumn, should be included in the

future in comparison to spring data. Thus one can integrate seasonal change as one of the

factors into interactions between plants and their environment.

• Natural species, as well as some other cultivars with certain pigmentation patterns should

be investigated in the future to either confirm the data and interpretation obtained from

this study.

• With more and more commercially available standards, more compounds from flavonoid

biosynthetic pathway can be identified and determined from maples and other plants, so

144 that the whole network of flavonoid biosynthesis can be discussed in terms of

ecophysiological functions.

The prospects listed above will provide more information from genetics, genomics and

metabolomics. Armed with such knowledge will finally perfect our understanding of the ecophysiological roles of anthocyanins and proanthocyanidins being played in the interaction of plants and their living environment. Such knowledge can also shed some light on physiological and ecological functions of other secondary metabolites in the plant kingdom and offer a possible “manual” to genetically manipulate plants for the benefits of human being.

145 CHAPTER 3

ANTHOCYANINS AND PROANTHOCYANIDINS METABOLISM IN SPRING

JUVENILE LEAVES OF SEVERAL DECIDUOUS TREE SPECIES

3.1 Introduction

From our previous study elaborated in Chapter 2, the data we obtained from Acer palmatum

Thunb. cultivars indicates that:

• Anthocyanin level corresponds to leaf coloration pattern;

• The formation of proanthocyanidin leads to less red pigmentation and such level

change is possibly due to metabolic channeling of anthocyanidins to

proanthocyanidin production which plays a major role in leaf pigmentation pattern

change;

• Both qualitative and quantitative changes contribute to total anthocyanin level

change;

• Levels of proanthocyanidins as well as anthocyanins change through development;

• Anthocyanidin biosynthetic pathway is always activated even there is no detectible

anthocyanin present;

• A gradual alteration of ecophysiological function may be occurring from

photoprotection as a sunlight filter (anthocyanins) to defensive mechanism as

146 deterrent compounds (proanthocyanidins) in juvenile developing leaves in spring in a

development-dependent fashion.

However, all of those indications were interpreted based on our data retrieved from six A. palmatum cultivars, which might probably cause bias in the results, hence further misinterpretation. In order to confirm our previous speculations and test whether those speculations are prevailing in other plant species, even across the plant kingdom. We selected seven deciduous tree species including six maples and one Prunus spp. as our experimental objects. We adopted the same approaches described in Chapter 2 including analyses of anthocyanins and proanthocyanidins by means of spectrometry and HPLC. This study aims at a larger goal to expand our speculations and results to other maple species, or even to species beyond Aceraceae.

3.2 Materials and Methods

3.2.1 Plant materials

All materials were grown and collected in JC Raulston Arboretum from late April to early

May to ensure that similar growth conditions have been applied to all plants. The properties of height and foliar pigmentation are briefly summarized in tables 5 and 6. All cultivars have palmate opposite leaves.

147 Table 5. Height, taxonomy and location of species selected in this work. Numbers after scientific name of each species are reference numbers to be used for the sake of convenience. The reference numbers start from 7 following after six cultivars being studied in Chapter 2. Botanical Name (#Reference) Height Taxonomy1 Common Name Location2 Acer rubrum #7 ~5m Rubra maple D30 Celebration® Acer ×freemanii 'Celzam' 3#8 ~5m Rubra D30 Freeman maple Acer longipes #9 ~2.5m Platanoidea D32 Acer henryi #10 ~3m Negundo Chinese box-elder W12

Acer truncatum #11 ~7m Platanoidea Shantung maple E38e Acer negundo 'Kelly's Gold' #12 ~3m Negundo golden box-elder D32

Prunus padus ‘Red Ball’ #13 ~2m Rosaceae N/A NCSU

All materials are collected in the same manner described in Chapter 2 except for Prunus padus. This species had two types of twigs at the time of collection, one bearing young green leaves and the other bearing mature fully expanded leaves. Thus leaves from both types of twigs were collected according to their nodes numbers. Young green leaves were designated to Y-1, Y-2, Y-3 and Y-4. Fully expanded leaves were designated as the other species in the format of P-1 to P-9.

1 The names listed under Column “Taxonomy” for Acer are their taxonomic ranks as in cluster/section. Rosaceae is the family Purnus padux belongs to.

2 Locations for Acer are bedding numbers where those maples are planted in JC Raulston Arboretum, as a part of the Department of Horticulture Science at North Carolina State University, while for Prunus, it is located along Yarborough St., at NCSU main campus.

3 Acer xfreemanii is artificially hybridized between A. rubrum x A. saccharinum by Freeman, a plant breeder at Arnold Arboretum. This maple exists also occasionally in nature, where the two parent grow in close proximity. Both parents belong to section Rubra.

148 Table 6. List of species selected in this work. *Developmental sequence on the twig ‐ their sample numbers in the experiment. Numbers after scientific name of each cultivar are reference numbers to be used for the sake of convenience. Botanical Name (#Reference) Pattern Collection Manner* 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P- Acer rubrum #7 RG 4), 7th(P-7), 8th(P-8); 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P- RG Acer ×freemanii 'Celzam' #8 4) and shaded leaves (SH-L); 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P- Acer longipes #9 RG 4), 6th(P-6) 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P- Acer henryi #10 RG 4), 5th(P-5), 6th(P-6) 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P- Acer truncatum #11 RG 4), 5th(P-5) and shaded leaves (SH-L); 1st(P-1), 2nd(P-2), 3rd(P-3), 4th (P- Acer negundo 'Kelly's Gold' #12 G/Y 4), 5th(P-5), 7th(P-7) Green young twig: 1st(Y-1), 2nd(Y-2), 3rd(Y-3), 4th(Y-4) and st nd Prunus padus ‘Red Ball’ #13 GR Mature twig 1 (P-1), 2 (P-2), 3rd(P-3), 4th (P-4), 5th(P-5), 6th(P- 6), 7th(P-7), 8th(P-8), 9th(P-9);

3.2.2 Methods

The methods used here are the same as described in Chapter 2. Please refer to 2.2.3 for details.

3.3 Results

149

3.3.1 Acer rubrum

Leaf pigmentation, anthocyanins and proanthocyanidins

Leaf pigmentation of Acer rubrum is red to green (RG). Six pairs of leaves including 1st, 2nd,

3rd, 4th, 7th and 8th were collected. Measurement of ABS values at 530nm was conducted to

estimate anthocyanin levels. Extraction samples were also hydrolyzed for estimation of

proanthocyanidin levels at 550nm (Figure 57). Levels of anthocyanins correspond to foliar

pigmentation and but the proanthocyanidin levels appear stable. Detailed statistics can be

referred to Table 7, 8 and 9.

150

Fig 57. Leaf phenotype of Acer rubrum and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐ HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

151

152 LC-MS analysis of anthocyanins and anthocyanidins

We selected several leaf samples including the 1st, 2nd, 3rd and 8th samples, for liquid

chromatography analysis. Two major anthocyanin peaks were detected at 530nm (Figure 58).

Those samples were also hydrolyzed with hot acid to release the core anthocyanidin. Figure

57 shows the level change of anthocyanidins after hydrolysis in each sample. Anthocyanidin

levels of the 1st and 2nd pairs of leaves are consistent with their anthocyanin levels. However,

there is only trace anthocyanidin for the 3rd and 8th pairs of leaves (Figure 59).

153

Fig 58. LC chromatogram of Acer rubrum . Major anthocyanin peaks were marked with different numbers. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 3rd pair of leaves; D: the 8th pair of leaves. Numerals 1‐4 are used to label major peaks detected at 530nm.

154

Fig 59. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Acer rubrum. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 3rd pair of leaves; D: the 8th pair of leaves. Cyn=cyanidin. (Other unidentifiable anthocyanidin peaks are marked as An 1and An 2.)

3.3.2 Acer x freemanii ”Celzam”

Leaf pigmentation, anthocyanins and proanthocyanidins

This species has slightly red pigmented young leaves. When the leaves become more mature and their sizes increase, red pigmentation fades away. Even the red pigmentation is visually observable, low ABS values at 530nm were recorded. Compared to anthocyanin levels, proanthocyanidin levels in the 1st, 2nd and shaded leaves were higher. Detailed statistics can

be referred to Table 7, 8 and 9. Figure 60 shows the leaf phenotype and measurements of

both anthocyanins and proanthocyanidins.

155

Fig 60. Leaf phenotype of Acer x freemanii and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

156 LC-MS analysis of anthocyanins and anthocyanidins

Liquid chromatography detected two major anthocyanin peaks at 530nm in the 1st and 2nd pair of leaves. In the shaded leaves, no anthocyanin peaks were detected, which was consistent with low ABS values measured at 530nm (Figure 61).

Butanol-HCl hydrolysis was conducted in the 1st, 2nd and shaded leaf extraction samples. All

three major anthocyanidins, delphinidin, cyanidin and pelargonidin were detected along with

three more unidentifiable anthocyanidins in the 1st and 2nd pairs of leaves. However, in the

shaded leaves, no delphinidin, cyanidin or pelargonidin were detected (Figure 62).

157

Fig 61. LC chromatogram of Acer x freemanii. Major anthocyanin peaks were marked with different numbers. A: the 1st pair of leave; B: the 2nd pair of leaves; C: shaded leaves. Numerals 1‐3 are used to label major peaks detected at 530nm.

158

Fig 62. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Acer x freemanii. A A: the 1st pair of leave; B: the 2nd pair of leaves; C: shaded leaves. Dp=delphinidin. Cyn=cyanidin. Pg=pelargonidin. Other unidentifiable anthocyanidin peaks are marked as An 1, An2 and An 3.

159 3.3.3 Acer longipes

Leaf pigmentation, anthocyanins and proanthocyanidins

Acer longipes has red young leaves and dark green mature leaves. Measurement of ABS

values at 530nm corresponds to their leaf pigmentation. An anthocyanin level of the 1st pair of leaves is slightly less than that of the 2nd pair of leaves. When leaves become mature and

fully expanded, red pigmentation fades away and the anthocyanin level turns to very low.

However, hydrolysis analysis showed that even though anthocyanins were hardly produced

in the 6th pair of leaves, proanthocyanidin level was the highest among all samples. Such result indicated that proanthocyanidins were produced and accumulated in mature leaves of

Acer longipes even though no anthocyanins were produced. Detailed statistics can be referred to Table 7, 8 and 9. Figure 63 shows the leaf phenotype and measurements of both anthocyanins and proanthocyanidins.

160

Fig 63. Leaf phenotype of Acer longipes and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐ HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

161

162 LC-MS analysis of anthocyanins and anthocyanidins

The 1st, 2nd and 6th samples were analyzed by LC. Anthocyanin profiles were obtained. For

the 1st and 2nd samples, anthocyanin profiles were almost identical, while for the 6th sample,

no anthocyanin peaks were detected (Figure 64).

Fig 64. LC chromatogram of Acer longipes. Major anthocyanin peaks were marked with different numbers. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 6th pair of leaves. Numerals 1‐6 are used to label major peaks detected at 530nm.

163 After hydrolysis, all three samples showed complex anthocyanidin profiles. For the 1st and

2nd pairs of leaves, their anthocyanidin profiles were almost similar to each other just like

their anthocyanin profiles. For the 6th sample, delphinidin, cyanidin and pelargonidin were

also detected just like the 1st and 2nd samples, although with different quantity. Unidentifiable

anthocyanidins were also found in all samples. The 6th sample had more unidentifiable

anthocyanidins than the 1st and 2nd ones, which could probably result from proanthocyanidin

hydrolysis (Figure 65).

164

Fig 65. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Acer longipes. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 6th pair of leaves. Dp=delphinidin. Cyn=cyanidin. Pg=pelargonidin. Other unidentifiable anthocyanidin peaks are marked as An 1 to An 5.

165 3.3.4 Acer henryi

Leaf pigmentation, anthocyanins and proanthocyanidins

Young leaves of Acer henryi have strong red pigmentation. When leaves become mature, red pigmentation would disappear and leaves would turn to dark green. Measurement of anthocyanins showed that anthocyanin levels corresponded to leaf pigmentation. With the maturation of leaves, anthocyanin levels dramatically decreased. At the same time, proanthocyanidins seemed almost stable. Detailed statistics can be referred to Table 7, 8 and

9. Figure 66 and 67 show the leaf phenotype and measurements of both anthocyanins and proanthocyanidins of A. henryi.

166

Fig 66. Leaf phenotype of Acer henryi. Leaf coloration patterns from early young leaves through fully expanded mature leaves. A: the 1st and 2nd pairs of leaves; B: the 3rd and 4th pairs of leaves; C: the 5th and 6th pairs of leaves. (Figure continues on next page.)

167

168

Fig 67. Measurement of anthocyanins and proanthocyanidins in Acer henryi leaves. A: Measurement of ABS values of anthocyanins at 530nm; B: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐ HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart A and B.

169 LC-MS analysis of anthocyanins and anthocyanidins

The 1st, 3rd and 5th samples were chosen for LC analysis. Two major peaks were detected in

the 1st and 3rd samples with two other unidentifiable anthocyanins. In the 5th sample, peak 1 at 530nm was tiny, which indicated that there was almost no anthocyanin present in the 5th pair of leaves (Figure 68).

Fig 68. LC chromatogram of Acer henryi. Major anthocyanin peaks were marked with different numbers. A: the 1st pair of leave; B: the 3rd pair of leaves; C: the 5th pair of leaves. Numerals 1‐4 are used to label major peaks detected at 530nm.

170

Fig 69. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Acer henryi. A: the 1st pair of leave; B: the 3rd pair of leaves; C: the 5th pair of leaves. Cyn=cyanidin. Other unidentifiable anthocyanidin peaks are marked as An 1 and An2.

171 After hydrolysis, cyanidin was found to be the major anthocyanidin in both 1st and 3rd pairs of leaves. No cyanidin was detected in the mature 5th pair of leaves at 530nm although two

unidentifiable anthocyanidins were present in all three samples (Figure 69).

3.3.5 Acer truncatum

Leaf pigmentation, anthocyanins and proanthocyanidins

This species have dark green mature leaves and slightly red young leaves. Estimation of

anthocyanins with the help of ABS measurement showed that anthocyanin levels decreased during leaf maturation. Leaves with the highest amount of anthocyanins were the youngest

leaves. However, a dramatic increase of ABS values at 550nm was detected after hydrolysis

for the 5th pair of leaves even though there was trace anthocyanins produced. Such increase

indicated that proanthocyanidins were produced and accumulated in the mature leaves with

large quantities (Figure 70). Detailed statistics can be referred to Table 7, 8 and 9. Figure 70

shows the leaf phenotype and measurements of both anthocyanins and proanthocyanidins.

172

Fig 70. Leaf phenotype of Acer truncatum and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐ HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C. (Figure continues on next page.)

173

174 LC-MS analysis of anthocyanins and anthocyanidins

Extraction samples of the 1st, 2nd and 5th pairs of leaves were analyzed by LC. One major

anthocyanin peak was detected for the 1st and 2nd samples, while for the 5th sample, no

anthocyanin peaks were shown on LC chromatogram (Figure 71).

Fig 71. LC chromatogram of Acer truncatum. Major anthocyanin peaks were marked with different numbers. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 5th pair of leaves. Numerals 1‐3 are used to label major peaks detected at 530nm.

175 Butanol-HCl hydrolysis was conducted for all three samples to characterize the core

anthocyanidins. Cyanidin was found out to be the major anthocyanidin in our samples while

there was another unidentifiable anthocyanidin peak at 530nm. The abundance of cyanidin

corresponded to anthocyanin levels for the 1st and 2nd samples. However, for the 5th sample, abundance of cyanidin was the highest among all, which indicated that mature leaves produced proanthocyanidins, especially those with extension units as epicatechin or catechin

(Figure 72).

176

Fig 72. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Acer truncatum. A: the 1st pair of leave; B: the 2nd pair of leaves; C: the 5th pair of leaves. Cyn=cyanidin. One unidentifiable anthocyanidin peak is marked as An 1.

177 3.3.6 Acer negundo"Kelly’s Gold"

Leaf pigmentation, anthocyanins and proanthocyanidins

Fig 73. Leaf phenotype of Acer negundo and measurement of anthocyanins and proanthocyanidins. A: Leaf coloration patterns from early young leaves through fully expanded mature leaves; B: Measurement of ABS values of anthocyanins at 530nm; C: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐ HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart B and C.

178 This species appeared green yellow so we categorized its leaves with green-yellow (GY)

pigmentation pattern. Although young leaves showed slightly red pigmentation, very low

ABS values at 530nm were recorded. However, ABS values increased after hot acid

hydrolysis, which indicated that there might be proanthocyanidins produced in its leaves

despite their nodes number (Figure 73). Detailed statistics can be referred to Table 7, 8 and 9.

LC-MS analysis of anthocyanins and anthocyanidins

The 1st, 3rd and 5th samples were chosen for LC analysis even though low ABS values

suggested no anthocyanins. From LC chromatography, no anthocyanin peaks were detected for any samples, which corroborated the possibility that no anthocyanins were produced

(Figure 74).

However, after hydrolysis, cyanidin was detected for all samples as the major anthocyanidin.

Abundance of cyanidin was decreasing from the 1st sample to 3rd and then reached lowest in

the 5th sample (Figure 75). Such decrease could be related to estimated proanthocyanidin

levels (Figure 73D).

179

Fig 74. LC chromatogram of Acer negundo. No anthocyanin peaks were detected at 530nm. A: the 1st pair of leave; B: the 3rd pair of leaves; C: the 5th pair of leaves.

180

Fig 75. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Acer negundo. A: the 1st pair of leave; B: the 3rd pair of leaves; C: the 5th pair of leaves. Cyn=cyanidin.

181 3.3.7 Prunus padus

Leaf pigmentation, anthocyanins and proanthocyanidins

This cultivated species has totally different pigmentation pattern. In the early spring, the

plant produces twigs bearing green unexpanded leaves. With the time goes by, green

unexpanded leaves become more and more purple (Figure 76). Such pigmentation pattern is a

reverse process compared to most species in this thesis, which is from red to green (RG).

Thus we designated leaves of this species as green to red (GR). Detailed statistics can be referred to Table 7, 8 and 10.

Not fully expanded green leaves were collected from several twigs and pooled together and were designated as Y-1 to Y-4. Twigs with only fully expanded mature leaves were taken and leaves were collected, pooled together and designated as P-1 to P-9. Leaves from P-1 to

P-9 showed a clear trend of red/purple pigmentation along maturation. Thus, a similar increasing trend of anthocyanin levels were observed based on ABS values at 530nm (Figure

77).

After hydrolysis, all samples had a dramatic increase of ABS values at 550nm. However, there was no clear trend for such increase (Figure 77).

182

Fig 76. Leaf phenotype of Prunus padus. Leaf coloration patterns from early young leaves through fully expanded mature leaves. A: the 1st to 4th pairs of unexpanded leaves on young green twigs; B: the 1st, 2nd and 3rd pairs of fully expanded leaves on other twigs; C: the 4th and 5th pairs of fully expanded leaves; D: the 6th and 7th pairs of fully expanded leaves; E: the 8th and 9th pairs of fully expanded leaves. (Figure continues on next page.)

183

184

Fig 77. Measurement of anthocyanins and proanthocyanidins in Prunus padus leaves. A: Measurement of ABS values of anthocyanins at 530nm; B: ABS values of samples at 550nm before (Pre) and after (Post) butanol‐HCl hydrolysis and their differences (delta) as estimated levels of proanthocyanidins (PAs); Arrow in picture A indicates the nodes for each pair of leaves. P‐1, P‐2 etc are sample numbers for different leaf pairs, which corresponds to those on the x‐axis in chart B and C. Bars marked with different letters are significantly different (P<0.05) and values decrease alphabetically. Letters in lower case (e.g. =a), letters in lower case with apostrophe (e.g. =a’) and letters in upper case (e.g. =A) are only comparable within their own set. Standard deviation (based on three replicates) is depicted as error bars in chart A and B.

185 LC-MS analysis of anthocyanins and anthocyanidins

In order to characterize anthocyanins and the core anthocyanidin in this species, the 1st and

3rd pairs of young green leaves and the 1st, 4th, 7th and 9th pairs of fully expanded mature

leaves were used for LC-MS analysis. Only one anthocyanin peak was detected at 530nm for

all samples, with an increasing trend of abundance toward maturation. Such trend was

consistent to anthocyanin levels in those samples (Figure 78).

Fig 78. LC chromatogram of Prunus padus. One anthocyanin peak was detected and marked as 1. A: the 1st pair of unexpanded young leaves; B: the 3rd pair of unexpanded young leaves; C: the 1st pair of fully expanded leaves; D: the 4th pair of fully expanded leaves; E: the 7th pair of fully expanded leaves; F: the 9th pair of fully expanded leaves. Numerals 1 is used to label this major peak detected at 530nm.

Cyanidin along with another unidentifiable anthocyanidin was detected after hydrolysis.

There seems no clear increasing or decreasing development-dependent trend (Figure 79).

186

Fig 79. LC analysis of anthocyanidins released from butanol‐HCl hydrolysis of anthocyanins from leaves of Prunus padus. A: the 1st pair of unexpanded young leaves; B: the 3rd pair of unexpanded young leaves; C: the 1st pair of fully expanded leaves; D: the 4th pair of fully expanded leaves; E: the 7th pair of fully expanded leaves; F: the 9th pair of fully expanded leaves. Cyn=cyanidin. One unidentifiable anthocyanidin peak is marked as An 1.

3.4 Summary

Based on the data presented in this chapter, we found out that it confirms with several conclusions in Chapter 2, which are:

• Anthocyanin level corresponds to leaf coloration pattern;

187 • Both qualitative and quantitative changes contribute to total anthocyanin level

change;

• Levels of proanthocyanidins as well as anthocyanins change in response to

development;

However, we found out that the development-dependent trend of proanthocyanidins was not

that clear for the species selected in Chapter 3 even though anthocyanin levels had increasing

/decreasing development-dependent trends just like those cultivars of A. palmatum in Chapter

2. And since we did not conduct detailed experiments on proanthocyanidins, we can not

corroborate the data and interpretations derived from those results in Chapter 2. More

thorough and detailed survey is required for more species to confirm the hypotheses we

proposed in Chapter 2.

188 Table 7. Estimation of anthocyanin content level. Average values ± standard deviation are presented. Values (ABS units/100mg FW) are measured at 530nm for leaf samples from six Acer species. Different letters after standard deviation mean statistical differences of estimated anthocyanins contents level among different samples in each cultivar and values decrease alphabetically. NA means samples not analyzed. Acer rubrum‐7; Acer ×freemanii 'Celzam'‐8; Acer longipes‐9; Acer henryi‐10; Acer truncatum‐11; Acer negundo 'Kelly's Gold'‐12. Cultivar SH-L P-1 P-2 P-3 P-4 P-5 P-6 P-7 P-8 7 NA 4.98±0.52b 6.75±0.41a 1.81±0.01c 0.77±0.05d NA NA 0.10±0.01d 0.07±0.00d 8 0.05±0.00c 0.13±0.00b 0.28±0.01a 0.05±0.00c 0.03±0.00d NA NA NA NA 9 NA 0.45±0.01b 0.53±0.01a 0.10±0.00c 0.08±0.00d NA 0.08±0.00cd NA NA 10 NA 6.71±0.17a 2.72±0.20b 2.15±0.02c 0.36±0.01d 0.22±0.01d 0.13±0.00d NA NA 11 0.07±0.00c 0.35±0.01a 0.12±0.01b 0.06±0.00c 0.06±0.00c 0.07±0.00c NA NA NA 12 NA 0.01±0.01b 0.02±0.00ab 0.02±0.01b 0.02±0.00b 0.02±0.00ab NA 0.03±0.01a NA

189 Table 8. Estimation of proanthocyanidin content level. Average values ± standard deviation are presented. Values (ABS units/reaction) are calculated from subtraction of ABS1 (pre‐) from ABS2 (post‐) of leaf samples from six Acer species. Different letters after standard deviation mean statistical differences of estimated anthocyanins contents level among different samples in each cultivar and values decrease alphabetically. NA means samples not analyzed. No data for P‐4 or P‐7 was analyzed thus they were not listed here. Acer rubrum‐7; Acer ×freemanii 'Celzam'‐8; Acer longipes‐9; Acer henryi‐10; Acer truncatum‐11; Acer negundo 'Kelly's Gold'‐12. Cultivar SH-L P-1 P-2 P-3 P-5 P-6 P-8 7 NA 0.68±0.09a 0.60±0.05ab 0.47±0.01b NA NA 0.66±0.03a 8 0.25±0.01b 0.59±0.02a 0.52±0.10a NA NA NA NA 9 NA 0.11±0.01b 0.17±0.01b NA NA 0.87±0.04a NA 10 NA 0.28±0.01b NA 0.43±0.02a 0.39±0.02a NA NA 11 NA 0.15±0.02c 0.37±0.03b NA 4.27±0.04a NA NA 12 NA 0.08±0.00a NA 0.05±0.01ab 0.05±0.01b NA NA

Table 9. Ratio of ABS values (ABS2/ABS1) of hydrolyzed samples. Ratio is calculated between average ABS2 and ABS1 of leaf samples from six Acer species. NA means samples not analyzed. No data for P‐4 or P‐7 was analyzed thus they were not listed here. Acer rubrum‐7; Acer ×freemanii 'Celzam'‐8; Acer longipes‐9; Acer henryi‐10; Acer truncatum‐11; Acer negundo 'Kelly's Gold'‐12. Cultivar SH-L P-1 P-2 P-3 P-5 P-6 P-8 7 NA 1.90 1.58 2.19 NA NA 8.45 8 18.42 14.78 7.55 NA NA NA NA 9 NA 1.90 1.97 NA NA 37.77 NA 10 NA 1.33 NA 3.26 10.23 NA NA 11 NA 1.90 7.64 NA 201.17 NA NA 12 NA 48.80 NA 11.00 13.67 NA NA

190 Table 10. Estimation of anthocyanin, proanthocyanidin content levels and ratio of ABS values (ABS2/ABS1) of hydrolyzed samples from Prunus padus. Average values ± standard deviation are presented. Values (ABS units/100mg FW) are measured at 530nm for anthocyanin content level and values (ABS units/reaction) are calculated from subtraction of ABS1 (pre‐) from ABS2 (post‐) at 550nm for proanthocyanidin content level. Ratio is calculated between average ABS2 and ABS1. #Sample Anthocyanin Proanthocyanidin Ratio of ABS values Y-1 0.05±0.01 3.02±0.56b 207.02 Y-2 0.07±0.00 NA NA Y-3 0.06±0.00 1.44±0.07c 60.26 Y-4 0.05±0.00 NA NA P-1 0.09±0.00 4.44±0.03a 82.20 P-2 0.09±0.00 NA NA P-3 0.11±0.00 NA NA P-4 0.14±0.01 3.38±0.07b 19.11 P-5 0.15±0.00 NA NA P-6 0.19±0.01 NA NA P-7 0.20±0.00 3.04±0.48b 9.55 P-8 0.26±0.01 NA NA P-9 0.52±0.01 2.89±0.35b 5.14

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205 APPENDICES

Appendix A. Abundance of peak 1 and 2 in different leaf samples of four cultivars ...... 207

Appendix B. Trends of cyanidin level in different leaf samples of five cultivars ...... 208

Appendix C. Proanthocyanidin standards ...... 210

Appendix D. Key to A. palmatum Thunb. cultivars ...... 211

Appendix E. Summary of anthocyanin and proanthocyanidin reports for A. palmatum Thunb. cultivars ...... 212

206

Appendix A. Abundance of peak 1 and 2 in different leaf samples of four cultivars. Average values ± standard deviations are presented as error bars. Values are automatically calculated from LC peak area integration program from four cultivars of Acer palmatum Thunb possessing detectible anthocyanins. Numbers below x‐axis refer to cultivar reference number and #sample. Reference number: A. palmatum 'Hubb's Red Willow'‐1; A. palmatum 'Shaina'‐2; A. palmatum 'Hagoromo'‐3; A. palmatum Dissectum Atropurpureum¬‐4. A. palmatum 'Okushimo'‐5; A. palmatum 'Oridono nishiki'‐6 were not analyzed due to lack of detectible anthocyanins. Lines in blue represent abundance of peak 1 and lines in red abundance of peak 2. There is no unit for LC peak area, while “millions” along y‐axis just simplify the large value of LC peak area.

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Appendix B. Trends of cyanidin level in different leaf samples of five cultivars. Average values ± standard deviations are presented as error bars. Values are automatically calculated from LC peak area integration program from five cultivars of Acer palmatum Thunb. Numbers below x‐axis refer to cultivar reference number and #sample. Reference number: A. palmatum 'Hubb's Red Willow'‐1; A. palmatum 'Shaina'‐2; A. palmatum 'Hagoromo'‐3; A. palmatum Dissectum Atropurpureum¬‐4; A. palmatum 'Okushimo'‐5. Mixed samples of A. palmatum 'Oridono nishiki'‐6 were not charted. Lines in blue represent cyanidin levels detected and calculated under 530nm wavelength and lines in red refers to values obtained from 550nm wavelength for comparison. There is no unit for LC peak area, while “millions” along y‐axis just simplify the large value of LC peak area. (Figure continues on next page.)

208

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Appendix C. Proanthocyanidin standards. A~E: ion chromatograms of four proanthocyanidin standards and one trimer proanthocyanidin compound; F~J: mass spectra of corresponding compound. From A to E, compounds are epicatechin, catechin, procyanidin B1, procyanidin B2 and trimer. From F to J, mass spectra to be used for determination of those same compounds in A to E. Circled mass (m/z) numbers in F to J refer to molecular weights or fragments specific for certain compound peaks under positive ionization.

210 Appendix D. Key to A. palmatum Thunb. cultivars

1. Cultivars have trace or no amount of anthocyanins.

2. Cultivars have higher level of catechin than that of procyanidin B2 (Okushimo)

2’ Cultivars have lower level of catechin than that of procyanidin B2 (Oridono nishiki)

1’ Cultivars have detectible anthocyanins.

3. Cultivars have almost stable production of anthocyanins in juvenile developing leaves

(Dissectum Atropurpureum)

3’ Cultivars have declining production of anthocyanins in juvenile developing leaves towards maturation.

4. Cultivars have cyanidin 3-glucoside and 3-rutinoside (peak 1) as their dominant anthocyanins in the youngest developing leaves (Hubb's Red Willow)

4’ Cultivars have similar level of cyanidin 3-glucoside/ 3-rutinoside (peak 1) and cyanidin

3-galloylglucoside/ 3-galloylrutinoside (peak 2) in the youngest developing leaves.

5. Cultivars have both catechin and epicatechin as flavan-3-ols (Hagoromo)

5’ Cultivars only have epicatechin as flavan-3-ols (Shaina)

211 Anthocyanins Proanthocyanidins A. palmatum Thunb. Peak 1 Peak 2 Catechin Epicatechin Procyanidin B2 Trimer Hubb's Red Willow-1 +++ + - +++ + + Shaina-2 + + - + +++ + Hagoromo-3 + + + +++ + - Dissectum Atropurpureum-4 +++ + - +++ +++ - Okushimo-5 - - + +++ + - Oridono nishiki-6 - - + +++ + -

Appendix E. Summary of anthocyanin and proanthocyanidin reports for A. palmatum Thunb. cultivars. +/‐ means presence/absence respectively. +++ means that this compound or peak is the dominant one. Peak 1 consists of cyanidin 3‐glucoside and 3‐rutinoside while peak 2 consists of cyanidin 3‐galloylglucoside nd 3‐ galloylrutinoside. A. palmatum 'Hubb's Red Willow'‐1; A. palmatum 'Shaina'‐2; A. palmatum 'Hagoromo'‐3; A. palmatum Dissectum Atropurpureum¬‐4; A. palmatum 'Okushimo'‐5; A. palmatum 'Oridono nishiki'‐6.

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