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PREFACE
This thesis is submitted for the degree of Philosophy Doctor (PhD) in Chemistry at Sudan University of Science and Technology, Khartoum. The work has been carried out at Department of Chemistry, Sudan University of Science and Technology, Khartoum and at Department of Drug Technology, Faculty of Medical Technology, Libya, Dernah, during the period 20082011. One aim of the present work was to isolate and determine the chemical structures of flavonoids in different Libyan plants, for documentation and chemotaxonomic considerations with respect to these flavonoids. The findings of new flavonoids will expand the diversity of structures, which may through further examinations display other chemical and biological properties than previously known. At the same time, extensive spectroscopic methods, especially 2D NMR techniques will be applied to fully characterized the isolated compounds. Another purpose of the present study is to evaluated the biological activities of the extracts and the characterization of the active principles from these Libyan medicinal plant species. Finally, the present investigation also tries to study the fragmentation pattern of flavonoids isolated from medicinal plants by applying the tandem ESI-MS technique.Chapter 1 gives an introduction to the thesis, chapter 2 presents the methods used in this work and chapter 3 gives the results and discussion.
Kutaiba I. Alzand April, 2011
v
ACKNOWLEDGMENTS
Countless thanks to Almighty Allah, Creator of all of us, Worthy of all praises, Who guides in darkness and helps in difficulties. All respect to His Holly Prophet (peace be upon him), who enables us to recognize our Creator. He, who does not thank to people, is not thankful to Allah and Holly Prophet (peace be upon him). This is very difficult job to include names of all who were involved directly or indirectly in the accomplishment of this work. First of all, I would like to express my most sincerely appreciation to Professor Mohamed A. Mohamed. I feel so fortunate and pleasant to have him as my advisor. Prof. Mohamed impressed me not only by his profound and intelligent knowledge in Chemistry and its related fields, but also by his generosity, wisdom, patience, and the capability to grasp the key points from millions of minutiae. I would not have brought this dissertation to reality without his countless encouragement, support and kind direction. Dr. Mohamed also brought me into an amazing world where I have learned many advanced techniques. His strict and serious attitudes on my work, extensive knowledge, eagerness to teach me updated techniques, as well as organization habits inspired me to dedicate me to the interesting field of Chemistry. It is a pleasure to express my sincere thanks and gratitude to my co-supervisor Dr. George Y. Karkis for his supervision and encouragement during this study. I am grateful to Sudan University of Sciences and Technology at Khartoum, College of Science, and the Department of Chemistry for giving me the opportunity to do graduate studies and providing me the necessary working facilities. This work was not possible to be completed without having such sophisticated laboratory facilities, which we have at Faculty of Medical Technology, Libya, Dernah. Therefore acknowledgement is due to Dr. Tawfik Alhasadi Director of the Faculty. I offer my sincere thanks to Dr. Selva Kumar (Faculty of Medical Technology) for his help carrying out the microbiological studies. Thanks are also due to Dr. Salah Surgwa (Omar Almuktar University) for identifying the plant materials I am grateful to my colleagues and friends for their support and their friendship; especially I want to thank Dr. Tawfik Alhasadi and Dr. Saad Alkaisy. Finally, I would like to express my deepest gratitude to my loving family, my father Ibrahim Alzand, my mother Maisoon Abid Alhamid, my sisters Enas Ibrahim, Jumana Ibrahim and Asil Ibrahim, my brother Huthepha Ibrahim, my wife Sundos Kamil, my sons Muslim Kutaiba and Mustafa Kutaiba, and particularly my lovely and clever daughter, Malak Kutaiba, they never stopped giving their love and support to me over these years, I really feel happy to have them in my family. Without all of the help mentioned above, this dissertation would not have become reality.
Kutaiba Alzand April, 2011 vi
TABLE OF CONTENTS PREFACE ………………………………………………………………………………......……… v ACKNOWLEDGEMENTS............................................................................................................... vi TABLE OF CONTENTS ................................................................................................................. vii LIST OF TABLES .............................................................................................................................xi LIST OF FIGURES .......................................................................................................................... xii LIST OF SYMBOLS AND ABBREVIATIONS …………………………………………........… xvi CHAPTAR ONE 1. INTRODUCTION ………………………………………………………………………………. 1 1.1. PRIMARY AND SECONDARY METABOLITES …………………………….…………. 1 1.2. FLAVONOIDS …………………………………………………………………….………. 3 1.2.1. CLASSIFICATION AND CHEMISTRY …………………………………...………. 3 1.2.2. BIOSYNTHESIS OF FLAVONOIDS ………………………………………….…… 6 1.2.3. BIOAVAILABILITY AND METABOLISM OF FLAVONOIDS …………..…..…. 8 1.2.4. THE MECHANISMS OF ACTION OF FLAVONOIDS ……………………..…… 10 1.2.4.1. Antioxidant Potential …………………………………………………...…. 10 1.2.4.2. Pro-Oxidant Effects ……………………………………………….….…… 11 1.2.4.3. Hormonal Properties …………………………………………………...….. 12 1.2.5. BIOLOGICAL AND PHARMACOLOGICAL ACTIVITIES OF FLAVONOIDS ………………………………………...…….… 14 1.2.5.1. Antiviral Activity ………………………...……………………………….. 14 1.2.5.2. Antifungal Activity ………………………………………………….…..… 14
1.2.5.3. Antimicrobial Activity ……………………………………………………. 14 1.2.5.4. Anti-Carcinogenic Activity ……………………………………………..… 15 1.2.5.5. Hepatoprotective Activity …………………………………………..…….. 15 1.2.5.6. Cardioprotective Activity ……………………………………………..….. 15 1.2.5.7. Neuroprotective Activity ………………………………………………..… 16 1.2.5.8. Antimalarial Activity ………………………………………………..…….. 16 1.2.5.9. Antileishmanial Activity ………………..……………………..………...…17 1.2.5.10. Antitrypanosomiasis Activity …………………………………………….. 18 1.2.5.11. Antiamebiasis Activity ………………………………………..………..… 18 1.3. ANALYTCIAL SEPARATION AND DETECTION METHODS FOR FLAVONOIDS …………………………………….…………………… 19 1.3.1. SAMPLE TREATMENT ………………………………………..……………..…… 19 vii
1.3.1.1. Analyte Isolation ……………………………………...……………………19 1.3.1.2. Solid-Phase Extraction ………………………………………………..…… 21 1.3.1.3. Matrix Solid-Phase Dispersion ………………………………………..…… 24 1.3.1.4. Solid-Phase Micro-Extraction ………………………………….………….. 25 1.3.2. SEPARATION AND DETECTION ………………….……………………………..26 1.3.2.1. Column Liquid Chromatography …………………………..….…………... 26 1.3.2.1.1. General ……………………………...……..………...…………. 26 1.3.2.1.2. Detection in LC ………………………………………………… 28 1.3.2.1.2.1. UV Absorbance Detection …………………….…… 28 1.3.2.1.2.2. Fluorescence Detection ……………………….…… 32 1.3.2.1.2.3. Electrochemical Detection ………………………… 34 1.3.2.1.3. LC-MS …………….…………………….………………………. 36 1.3.2.2. Less Common Methods …………………...……………………………….. 38 1.3.2.2.1. General …………………………………………..……………... 38 1.3.2.2.2. Gas Chromatography ………….…………………………...…… 39 1.3.2.2.3. Capillary Electrophoresis ………………………………….…… 41 1.3.2.2.4. Thin-layer Chromatography …………………………...………..45 1.3.3. IDENTIFICATION AND STRUCTURAL CHARACTERIZATION …………..….48 1.3.3.1. General ……………………………………………………….….…………48 1.3.3.2. LC-MS/MS ………………………………………………………..……….. 48 1.3.3.2.1. General ………………………………………………......……... 48 1.3.3.2.2. Fragmentation in PI Mode ……………………………………… 50 1.3.3.2.3. Fragmentation in NI Mode ………………………………...…… 53 1.3.3.2.4. Flavonoid–(di)glycosides ………………………………….……56 1.3.3.3. LC–NMR …………………………………………………………………... 58
1.4. PHARMACOGNOSY OF PISTACIA LENTISCUS ………….…………………...…….. 63
1.4.1. BOTANY AND TAXONOMY …………………………………………………… 63
1.4.1.1. The family Anacardiaceae ………………………………………...……… 64
1.4.1.2. The genus Pistacia …………………………….……………………….…65
1.4.1.3. The species Pistacia lentiscus L. ……………………………..………….. 65 1.4.2. TRADITIONAL MEDICINE ……………………………………………..………. 65 1.4.3. PHYTOCHEMISTRY ………………………………………………..……………. 66 1.4.4. PHARMACOLOGY …………………………………………………….…………. 66
1.4.5. TOXICOLOGY ………………………………………………………..…..…..…… 66
1.5. PHARMACOGNOSY OF JUNIPERUS PHOENICEA ………………..………..……….. 67 viii
1.5.1. BOTANY AND TAXONOMY …………………………………………..………… 67
1.5.1.1. The family Cupressaceae …………………………………….…..…….…. 68
1.5.1.2. The genus Juniperus …………………………………………..…………. 68
1.5.2. TRADITIONAL MEDICINE ………………………………………..………...….. 69
1.5.3. PHYTOCHEMISTRY …………………………………………………..………..... 69
1.5.4. PHARMACOLOGY ………………………………………………………..…..…..69
1.5.5. TOXICOLOGY …………………………………………………………....………. 70
1.6. PHARMACOGNOSY OF CUPRESSUS SEMPERVIRENS …………………..…...…… 71
1.6.1. BOTANY AND TAXONOMY …………………………………………………… 71
1.6.1.1. The family Cupressaceae ……………………………………..……..……. 72
1.6.1.2. The genus Cupressus ……………………………………………..………. 72
1.6.2. TRADITIONAL MEDICINE ……………………………………..…………...….. 72
1.6.3. PHYTOCHEMISTRY …………………………………………..…………...…….. 73
1.6.4. PHARMACOLOGY ……………………...………………..………………..……. 73
1.6.5. TOXICOLOGY …………………………………..…………………………..……. 73
1.5.1.3. The species Juniperus phoenicea L. …………………………..………….. 68
1.6.1.3. The species Cupressus sempervirens L. …………………………..…..….. 72
CHAPTAR TWO 2. EXPERIMENTAL …………………………………………………………………….………. 74
2.1. MATERIALS AND METHODS ……….…………………………………………..……. 74
2.1.1. PLANT MATERIALS ……………………..………………………………..……. 74 2.1.2. PHYTOCHEMICAL SCREENING ……………………...………………..……. 74 2.1.2.1. Preparation of Reagents for Phytochemical Screenin ………………...…. 74 2.1.2.1.1. Flavonoid Test Reagents …………………………....…..……. 74 2.1.2.1.2. Alkaloid Test Reagents …………….……………...…..……... 75 2.1.2.1.3. Tannins Test Reagents ……………………………….…..…… 75 2.1.2.2. Preparation of Aqueous Extract for Phytochemical Screening ……....…. 75 2.1.2.3. Preparation of Alcoholic Extract for Phytochemical Screening ……....... 75 2.1.2.4. Test for Tannins ……………………………………………...……..…… 75
2.1.2.5. Test for Steroids ………………………………………………....……… 75
2.1.2.6. Test for Flavonoids ……………………………………………..……….. 76 2.1.2.7. Test for Alkoloids …………………………………………...……..….... 76
2.1.2.8. Test for Terpenoids (Salkowski Test) ……………………...…...………. 76 2.1.2.9. Test for Saponins …………………………………………..……...…….. 76 ix
2.1.3. BIOLOGICAL ACTIVITIES OF FLAVONOIDS ……………………....……….. 77 2.1.3.1. Micro-Organism ……………………………………………….…..…….. 77 2.1.3.2. Antimicrobial Activity …………………………………………..…..…... 77 2.1.4. EXTRACTION AND ISOLATION …………………………………….....……... 77 2.1.4.1. Extraction of Flavonoids from the Leaves of Pistacia Lentiscus ...........… 77 2.1.4.2. Isolation of Flavonoids from the Leaves of Pistacia Lentiscus..……..….. 78 2.1.4.2.1. Thin-layer Chromatography of the Crude Products …….....…. 78 2.1.4.2.2. Preparative Thin-Layer Chromatography ……………….....… 78 2.1.4.3. Extraction of Flavonoids from the Barks of Pistacia Lentiscus ……..…... 78 2.1.4.4. Isolation of Flavonoids from the Barks of Pistacia Lentiscus ………….. 78 2.1.4.4.1. Thin-layer Chromatography of the Crude Products ..……...…..78 2.1.4.4.2. Preparative Thin-Layer Chromatography ……………………. 79 2.1.4.5. Extraction of Flavonoids from the Leaves of Juniperus Phoenicea …...... 79 2.1.4.6. Isolation of Flavonoids from the Leaves of Juniperus Phoenicea …..….. 79 2.1.4.6.1. Thin-Layer Chromatography of the Crude Products ……….… 79 2.1.4.6.2. Preparative Thin-Layer Chromatography ………………....…. 79 2.1.4.7. Extraction of Flavonoids from the Barks of Juniperus Phoenicea …….... 80 2.1.4.8. Isolation of Flavonoids from the Barks of Juniperus Phoenicea ……..…. 80
2.1.4.8.1. Thin-Layer Chromatography of the crude products ………...... 80 2.1.4.8.2. Preparative Thin-Layer Chromatography ……………………. 80 2.1.4.9. Extraction of Flavonoids from the Barks of Cupressus Sempervirens ...... 80 2.1.4.10. Isolation of Flavonoids from the Barks of Cupressus Sempervirens ........ 81
2.1.4.10.1. Thin-Layer Chromatography of the Crude Products ……...... 81 2.1.4.10.2. Preparative Thin-Layer Chromatography ………………..… 81
CHAPTAR THREE 3. RESULTS AND DISCUSSION ………………………………………..…………………..…. 82 REFERENCES …………………………………………………………………………...……… 139 x
LIST OF TABLES Table 1. Main Classes of Phenolic Compounds ……………………………………………...…… 2 Table 2. Selection of Recent Publications on LC Analysis of Flavonoids ……………………… 27 Table 3. Ȝmax of Some Representative Flavonoids of the Main Sub-Classes, with Examples of Each Class ………………………………………………….……….. 30 Table 4. Representative Studies on TLC of Flavonoid …………………………………….……. 45 Table 5. Fragment ions observed for the selected flavonoid classes in the PI mode ………...….. 51 Table 6. Fragment ions observed for the selected flavonoid classes in the NI mode ……………. 54 Table 7. LC–NMR studies of flavonoids …………………………………………………………… 59
Table 8. Qualitative Analysis of the Phytochemicals of the Medicinal Plants …………….…….. 83 Table 9. Antimicrobial Screening of Ethanolic Extracts of the Leaves of Juniperus phoenicea,Pistacia lentiscus and Cupressus sempervirens. ……………..…. 83
xi
LIST OF FIGURES Figure 1. Schematic of the major branch pathways of flavonoid biosynthesis …………………... 7 Figure 2. Strategies for the determination of flavonoids in biological fluids, beverages, plants and food ……………………………………………………...……. 20 Figure 3.
LC–UV265 of Merlot wine without and with molecular imprinted polymer-SPE (elution with acetonitrile) ………………….……………… 23
Figure 4. RPLC–UV250 and RPLC–FLU of an extract of T. pratense leaves………………..….. 29 Figure 5.
LC–DAD UV of an H. stoechas extract ………………………...……………………. 31
Figure 6. Comparison of detection methods for 3&,4&,5&-trimethoxyflavone in an extract of Flos primulae veris ……………………………………………...…… 32 Figure 7.
Fluorescence excitation and emission spectra of formononetin and ononin in methanol–water (1:1, v/v) at the pH values indicated ………………………..……. 33
Figure 8. Voltammogram and structure of rutin ………………………………………………… 35 Figure 9. MALDI(−)-TOF MS and the performance of four MALDI-TOF MS matrices …..….. 37 Figure 10. SIM-mode GC–MS chromatogram of a Mentha spicata extract after derivatization with methyl iodide ………………………………..……. 40 Figure 11. Total ion current and SIM-mode CE–ESI(−)–MS of naringenin in a phytomedicine................................................................................... 44 Figure 12. Two-dimentional (2D)-TLC of Flos sambuci extract on cyanopropyl-bonded silica…………………………………………………………47 Figure 13. RDA reaction mechanisms for the formation of 1,3A+ and 1,3B+ fragment ions for apigenin and luteolin ……………………….…………… 49 Figure 14.
Fragmentation pathways for flavonoids caused by cleavage of C-ring bonds….…… 50
Figure 15. Carbohydrate ion nomenclature for naringenin–7-O-neohesperidose (naringenin–7-O-rhamnosyl-(1ĺ4)-glucose) and naringenin–7-O-glucoside…….... 56 Figure 16. MS/MS spectra obtained for [M +H]+ ions ofnaringenin–7-O-neohesperidoside and (b) naringenin–7-O-rutinoside, using LC–ESI(+)– MS/MS………………...…… 57 Figure 17.
LC–UV254 chromatogram of Gentiana ottonis with the 1H NMR, DAD UV, MS and MS/MS spectra of swertisin……………………………………... 60
Figure 18. Structures of formononetin-7-O-ȕ-D-glucoside-6'-O-malonate and biochanin A-7-O-ȕD-glucoside-6'-O-malonate; formononetin-7-O-ȕ-D-glucoside-4'-O-malonate and 5-hydroxy-7-methoxyisoflavone-4&-O-ȕ-D-glucoside-4'-O-malonate…………….... 62 xii
Figure 19. Pistacia lentiscus L. …………………………………………………………………. 63 Figure 20.
Juniperus phoenicea L. …………………………………….…………………..….… 67
Figure 21. Cupressus sempervirens L………………………………………..……………..….… 71 Figure 22.
Inhibition zones of the EtOH extract of Pistacia lentiscus against Staphylococcus aureus…………………………………………………..….... 84
Figure 23.
Inhibition zone of the EtOH extract of Pistacia lentiscus against Escherichia coli…………………………………………………………..…… 84
Figure 24.
Inhibition zone of the EtOH extract of Pistacia lentiscus against Klebsiella pneumoniae………………………………………………..….…… 85
Figure 25.
Inhibition zones of the EtOH extract of Juniperus phoenicea against Staphylococcus aureus………………………………………………...…..….. 85
Figure 26.
Inhibition zone of the EtOH extract of Juniperus phoenicea against Escherichia coli. …………………………………………………………..….. 86
Figure 27.
Inhibition zone of the EtOH extract of Juniperus phoenicea against Klebsiella pneumoniae…………………………………………………..…… 86
Figure 28.
Inhibition zones of the EtOH extract of Cupressus sempervirens against Staphylococcus aureus……………………………………………………….. 87
Figure 29.
Inhibition zone of the EtOH extract of Cupressus sempervirens against Escherichia coli………………………………………………………….…… 87
Figure 30.
Inhibition zone of the EtOH extract of Cupressus sempervirens against Klebsiella pneumoniae……………………………………………...……..… 88
Figure 31.
IR (KBr, disc) spectrum of 1 ……………………………………………………..….. 89
Figure 32. UV spectrum of compound 1 …………………………………………………..….… 89 Figure 33.
Flavone and flavonol skeletons with benzoyl and cinnamoyl moieties………..……...90
Figure 34.
1
Figure 35.
13
Figure 36.
1
H-13C HMQC NMR spectrum of 1 in MeOH-d4. ……………………..………….… 94
Figure 37.
1
H-1H COSY NMR spectrum of compounds 1 in MeOH-d4 …………………..……. 95
Figure 38.
1
H-13C HMBC NMR spectrum of 1 in MeOH-d4………………………………….… 96
H NMR spectrum of compound 1 in MeOH-d4. ……………………………..….…. 92 C NMR spectrum of compound 1 in MeOH-d4………………………………..…... 93
Figure 39. Enlarged 1H-13C HMBC NMR spectrum of 1 in MeOH-d4…………………..…...… 97 Figure 40. ESI-MS2 spectrum of compounds 1 …………………………………………..…...… 97 Figure 41.
Fragmentation pathways to A and B fragments of compound 1 ……………….....… 98
Figure 42. Alternative pathway to B fragments observed as prominent peak at m/z 119 in the negative ion ESMS spectrum of compound 1. ……………….....… 98 Figure 43.
IR (KBr, disc) spectrum of 2 …………………………………………………....…… 99 xiii
Figure 44. UV spectrum of compound 2 ………………………………………………….…… 100 Figure 45.
1
Figure 46.
13
H NMR spectrum of compound 2 in MeOH-d4. ………………………….………. 101 C NMR spectrum of compound 2 in MeOH-d4. ……………….………………… 102
Figure 47.
1
H-13C HMQC NMR spectrum of 2 in MeOH-d4. …………………………………. 103
Figure 48.
1
H-1H COSY NMR spectrum of compounds 2 in MeOH-d4 ………………………. 104
Figure 49. Enlarged 1H-1H COSY NMR spectrum of compounds 2 in MeOH-d4 ……………. 105 Figure 50. Enlarged 1H-1H COSY NMR spectrum of compounds 2 in MeOH-d4. ……………. 105 Figure 51.
1
H-13C HMBC NMR spectrum of 2 in MeOH-d4. ………………...……………….. 106
Figure 52. Enlarged 1H-13C HMBC NMR spectrum of 2 in MeOH-d4. ……………………..… 107 Figure 53. ESI-MS2 spectrum of compounds 2 ………………………………………………... 108 Figure 54.
Fragmentation pathways to A and B fragments of compound 2 …………………... 108
Figure 55. Alternative pathway to B fragments observed as prominent peak at m/z 119 in the negative ion ESMS spectrum of compound 2…………………… 109 Figure 56.
IR (KBr, disc) spectrum of 3 ...................................................................................... 110
Figure 57. UV spectrum of compound 3 ………………………………………………………. 111 Figure 58.
1
H NMR spectrum of compound 3 in MeOH-d4. ………………………….……….. 112
Figure 59.
13
Figure 60.
1
C NMR spectrum of compound 3 in MeOH-d4 …………………………….……. 113
H-13C HMQC NMR spectrum of 3 in MeOH-d4. …………………………………. 114
Figure 61. Enlarged 1H-13C HMQC NMR spectrum of 3 in MeOH-d4. ………………...…….. 115 Figure 62. Enlarged 1H-13C HMQC NMR spectrum of 3 in MeOH-d4. ………………...…….. 115 Figure 63.
1
H-1H COSY NMR spectrum of compounds 3 in MeOH-d4. .................................... 116
Figure 64.
1
H-13C HMBC NMR spectrum of 3 in MeOH-d4 ………………………………….. 117
Figure 65. Enlarged 1H-13C HMBC NMR spectrum of 3 in MeOH-d4…………………...…….. 118 Figure 66. Enlarged 1H-13C HMBC NMR spectrum of 3 in MeOH-d4 ………………...……… 118 Figure 67. Enlarged 1H-13C HMBC NMR spectrum of 3 in MeOH-d4……………………....…. 119 Figure 68. ESI-MS2 spectrum of compounds 3 ………………………………………………... 119 Figure 69.
Important mass fragmentation of compound 3 ……………………………….……. 120
Figure 70.
IR (KBr, disc) spectrum of 4 ………………………………………………….....…. 121
Figure 71. UV spectrum of compound 4 ……………….…………………………………..….. 122 Figure 72.
1
Figure 73.
13
H NMR spectrum of compound 4 in MeOH-d4………………………….………... 123 C NMR spectrum of compound 4 in MeOH-d4…………………………….…..… 124
Figure 74.
1
H-13C HMQC NMR spectrum of 4 in MeOH-d4………………………………..…. 125
Figure 75.
1
H-1H COSY NMR spectrum of compounds 4 in MeOH-d4 ………………………. 126
Figure 76.
1
H-13C HMBC NMR spectrum of 4 in MeOH-d4 ……………………………….…. 127
Figure 77. ESI-MS2 spectrum of compounds 4 ……………………………………………....... 128 xiv
Figure 78.
Fragmentation pathways to A and B fragments of compound 4 ……………….….. 128
Figure 79. Alternative pathway to B fragments observed as prominent peak at m/z 119 in the negative ion ESMS spectrum of compound 4…………………… 129 Figure 80.
IR (KBr, disc) spectrum of 5 ……………………………………………………….. 130
Figure 81. UV spectrum of compound 5 ………………………………………….…………... 131 Figure 82. Figure 83.
1
H NMR spectrum of compound 5 in MeOH-d4. …………………………….…….. 132
13
C NMR spectrum of compound 5 in MeOH-d4. ……………………………..…... 133
Figure 84.
1
H-13C HMQC NMR spectrum of 5 in MeOH-d4. …………………………………. 134
Figure 85.
1
H-1H COSY NMR spectrum of compounds 5 in MeOH-d4 ………………………. 135
Figure 86.
1
H-13C HMBC NMR spectrum of 5 in MeOH-d4 ………………………………….. 136
Figure 87. ESI-MS2 spectrum of compounds 5 ………………………………………...……… 137 Figure 88.
Fragmentation pathways to A and B fragments of compound 5 …………………... 137
xv
LIST OF SYMBOLS AND ABBREVIATIONS
[Į]D
specific optical rotation
BuOH
butanol
br
broad
ºC
degree centigrade
įc
carbon-13 chemical shift
c
concentration
calcd
calculated
CE
capillary electrophoresis
CHCl3
chloroform
CID
collision-induced dissociation
cm
-1
reciprocal centimeters
13
carbon-13 nuclear magnetic resonance
COSY
COrrelative SpectroscopY
CPC
centrifugal partion chromatography
C NMR
1D, 2D
one- or two-dimentional
IT
ion-trap
d
doublet
į (ppm)
chemical shift (in parts per million)
DEPT
distortionless enhancement by polarization transfer
DMSO
dimethyl sulfoxide
Ķ
molar absorptivity
ECD ED
electron capture detection electrochemical detection
ERĮ
alpha estrogen receptor
ERȕ
beta estrogen receptor
ESI-MS
electrospray ionization - mass spectrometry
EtOAc
ethyl acetate
EtOH
ethanol
eV
electron volt
FDA
Food and Drug Administration
FID
flame ionization detection
FLU g
fluorescence gram(s) xvi
GC
gas chromatography
h
hour
įH
proton chemical shift
HMBC
heteronuclear multiple-bond connectivity spectroscopy
HMQC
heteronuclear multiple-bond quantum coherence spectroscopy
1
proton nuclear magnetic resonance
H NMR
HPLC
high-performance liquid chromatography
HR
high resolution
Hz
hertz
IC50
concentration that inhibits a response by 50% relative to a positive control
IT
ion-trap
ȳ
coupling constant
L
liter(s)
LC
liquid chromatography
LLE
liquid–liquid extraction
Ȝ (nm)
wavelength (in nanometers)
LP
lower phase of two-phase solvent mixture
M
molar concentration
[M-H]
-
[M+H]+ǜ
deprotonated molecule protonated molecule
[M]
molecular ion
max
maximum
MeOH
methanol
MeOH-d4
deuterated methanol (NMR solvent)
M
mol/liter
min
minute(s)
mg
10-3 gram(s) or milligram(s)
MPLC
medium-pressure liquid chromatography
ȝg
10-6 gram(s) or microgram(s)
ȝM
10-6 mol/liter
MHz
106 Hertz or megahertz
mL
10-3 liter(s) or milliliter(s)
MP
mobile phase in countercurrent chromatography
mp
melting point xvii
MS MS
mass spectrometry or mass spectrum 2
MSPD
tandem mass spectrometry (= MS/MS) matrix solid-phase extraction
m/z
mass-to-charge ratio
Ȟ
infrared absorption frequency
nm
nanometers or 10-9 meters
nM
10-9 mol/liter
NMR
Nuclear Magnetic Resonance
NOE
Nuclear Overhauser effect
NOESY
Nuclear Overhauser effect correlation spectroscopy
PDA
Photo Diode Array
ppm
parts per million (for MS, represents [experimental mass –
Q
quadrupole
theoretical mass] / [theoretical mass]) QqQ
triple-quadrupole
Rf
retention factor; migration distance of analyte as a fraction of distance to solvent
s
singlet
front in thin-layer chromatography SE
solvent extraction
Si
silica
SPE
solid-phase extraction
SPME
solid-phase micro-extraction
t
triplet
TOF
time-of-flight
tR
retention time
TLC
thin-layer chromatography
TMS
tetramethylsilane
UP
upper phase of two-phase solvent mixture
UV
ultraviolet
VLC
Vacuum Liquid Chromatography
v
volume
w
weight
xviii
1. INTRODUCTION 1.1. PRIMARY AND SECONDARY METABOLITES Plant chemicals are often classified as either primary or secondary metabolites. Primary metabolites are substances widely distributed in nature, occurring in one form or another in virtually all organisms. In higher plants such compounds are often concentrated in seeds and vegetative storage organs and are needed for physiological development because of their role in basic cell metabolism. (Examples: carbohydrates, lipids, proteins…)1. Secondary metabolites are compounds biosynthetically derived from primary metabolites. Contrary to primary metabolites these compounds are not ubiquitous in the living organisms that produce them nor are they necessarily expressed continuously. Although plants are better known as a source of secondary metabolites, bacteria, fungi and many marine organisms (sponges, tunicates, corals, and snails) are very interesting sources, too. Secondary metabolites, also known as natural products are not essential for normal growth, development or reproduction of an organism. In this sense they are "secondary". The function or importance of these compounds to the organism's development is usually of ecological nature as they are used as defence against predators (herbivores, pathogens etc.), for interspecies competition, and to 1,2
facilitate the reproductive processes . Secondary metabolites can be classified by their chemical structure or physical properties into one or more of the following groups: alkaloids, phenols, terpenoids, iridoids, steroids, saponins, polyletides, aliphatic, aromatic, and heteroaromatic organic acids, peptides, ethereals oils, resins and balsams 2. Phenolic compounds form one of the main classes of secondary metabolites with a large range of structures and functions, but generally possessing an aromatic ring bearing one or more hydroxyl substituents. This definition is not entirely satisfactory, however, since it inevitably includes compounds such as oestrone, the female sex hormone that is principally terpenoid in origin. For this reason, a definition based on metabolic origin is preferable, the plant phenols being regarded as those substances derived from the shikimate pathway and phenylpropanoid metabolism3. Natural polyphenols can range from simple molecules, such as phenolic acids, to highly polymerized compounds, such as tannins. They occur primarily in conjugated form, with one or more sugar residues linked to hydroxyl groups, although direct linkages of the sugar unit to an aromatic carbon atom also exist. The associated sugars can be present as monosaccharides, disaccharides, or even as oligosaccharides, and glucose is the most common sugar residue. Associations with other compounds, such as carboxylic and organic acids, amines, and lipids, and linkage with other phenols are also common4.
Table 1. Main Classes of Phenolic Compounds
Polyphenols can be divided into at least 10 different classes depending on their basic chemical structure5. Table 1 illustrates the basic chemical structure of the main polyphenolic compounds. In order to simplify the drawing of chemical structures, only the skeleton structures are shown in Tables 1, and there should be at least one hydroxyl group attached to the aromatic ring.
1.2. FLAVONOIDS 1.2.1. CLASSIFICATION AND CHEMISTRY Flavonoids are phenolic substances isolated from a wide range of vascular plants, and more than 8150 different flavonoids have been reported5. Flavonoids are located inside the cells or on the surface of various plant organs and have various functions in plants6. They act in plants as antioxidants, antimicrobials, photoreceptors, visual attractors, feeding repellents, and for light screening7. Many studies have shown that flavonoids exhibit biological and pharmacological activities, including antioxidant, cytotoxic, anticancer, antiviral, antibacterial, cardioprotective,
hepatoprotective,
neuroprotective, 8-12
antitrypanosomal and antiamebial properties
antimalarial,
antileishmanial,
. These biological and pharmacological
properties are usually attributed to their free radical scavenging efficacies, metal complexion capabilities, and their ability to bind to proteins with a high degree of specificity13. The basic flavonoid structure contains the flavan nucleus (1), which consists of 15 carbon atoms derived from a C6-C3-C6 skeleton. A flavonoid skeleton is composed of two aromatic rings (commonly designated as A and B), which are linked by a three-carbon chain. The connecting carbon chain combines with an oxygen to form a heterocyclic central or C-ring for most flavonoids with the exception of chalcones (2) in which the carbon chain between the A and B rings is linear14. The numbering scheme for chalcones differs from three-ring flavonoids in that the A ring, rather than the B ring carbons are labeled as prime.
2
C
A
B
B
A
ȕ
Į
2
1
2
Depending on the position of the linkage of the aromatic B-ring to the benzopyrano (chromano) moiety, this group of natural products may be divided into three classes: the flavonoids (2-phenylbenzopyrans) 1, isoflavonoids (3-phynylbenzopyrans) 3, and the neoflavonoids (4-phynylbenzopyrans) 4. These groups usually share a common chalcone precursor, and therefore are biogenetically and structurally related6.
2
2
Based on the degree of oxidation and saturation present in the heterocyclic C-ring, the flavonoids may be divided into the following groups: flavone (5), flavonol (6), flavanone (7), dihydroflavonol (8), flavan (1), flavan-3-ol ( ), flavan-4-ol (10), flavan-3,4-diol (11), and anthocyanidin (12).
O
O
O
O
5
6
O OH
O
7
2
2 2+
2+ 2
8
9
O
+
O
O OH
OH
OH
10
11
12
Natural products such as chalcones (2), dihydrochalcones (13), aurones (14) and auronols (15) also contain a C6-C3-C6 backbone and are considered to be minor flavonoids6.
B A
A
2 C
&+
13
A
2+
2 C
B
2
2
2
B
14
15
The flavonoids may be modified by hydroxylation, methoxylation, or O-glycosylation of hydroxyl groups as well as C-glycosylation directly to carbon atom of the flavonoid skeleton. In addition, alkyl groups (often prenyls) may be covalently attached to the flavonoid moieties, and sometimes additional rings are condensed to the basic skeleton of the flavonoid core. Flavonoid glycosides are frequently acylated with aliphatic or aromatic acid molecules. These derivatives are thermally labile and their isolation and further purification without partial degradation is difficult. Condensed tannins create a special group of flavonoid compounds formed by polymeric compounds built of flavan-3-ol units, and their molecular weights often exceeding 1,000 Da15. th
Up until the middle of the 20 century, flavonoids were believed to be waste products of plant primary metabolism, a notion that was soon abandoned based on research demonstrating the myriad functions of flavonoids in plant survival16. We now know the complex metabolic pathways used to synthesize flavonoids in plants-pathways that have evolved over millennia to provide a survival advantage to plants. Plants, clearly do not have the luxury of mobility and have had to evolve elaborate chemical mechanisms in order to defend themselves from various insults (such as UV radiation) and in order to attract those that will assist with their reproduction (bees, birds)17. Indeed, the complexity of the plant genome, which exceeds that of humans, is believed to have developed to maintain these immensely complex pathways of synthesis of novel chemicals17. Flavonoids are best known as the red, blue and purple pigments of flowering plants (due to the anthocyanidin sub-group) although the red pigment of some fruits can be due to carotenoids such as lycopene18. These pigments and the yellow pigments of flavones and flavanols are also responsible for the fall leaves coloration19. Because of the importance of color in pollinator attraction, flavonoids have an important role in plant reproduction. Flavonoids in plants also serve to protect from ultraviolet (UV) light owing to their high UV absorbance coefficients, and it has been suggested that this property of flavonoids was critical in the evolution of aquatic
plants to a terrestrial existence16. Flavonoids also possess other critical functions in defense against microorganisms and germination of pollen20,21. Each group of flavonoids possesses unique chemical properties and has a particular distribution in plants. Anthocyanins (glycosylated anthocyanidins) and proanthocyanidins (polymers that produce anthocyanidins when hydrolyzed) primarily provide color to flowering plants and fruit, and are therefore found in high concentrations in the skin of red grapes, red wine and berries. Flavan-3-ols, such as catechin, epicatechin gallate are colorless and are found in high concentrations in green tea. Isoflavones are only found in legumes (e.g. soy) and are therefore consumed in high quantities in regions of the world with high soy consumption. Flavanones are found in high levels in citrus fruits, while flavones are present in green leafy spices such as parsley, and flavonols are ubiquitous and found in most fruits and vegetables consumed in the human diet14,22. Interest in the health benefits of flavonoids stemmed from early research in 1936 by the Hungarian scientist Szent-Gyorgyi, who also incidentally discovered vitamin C. He isolated a substance, which he called citrin, from lemons that restored weakened capillaries to normal. This effect was not noted when vitamin C alone was administered. Citrin was later shown to be composed of the flavonoids hesperidin and eriodictyol23. Although citrin was also called vitamin P, this name was dropped in the 1950s after it was concluded that flavonoids did not fit the strict definition of a vitamin24. Despite not qualifying as vitamins, flavonoids have been shown in the last 72 years since Szent-Gyorgyi’s initial discovery to affect various aspects of human health not limited to their beneficial effects on capillary wall integrity.
1.2.2. BIOSYNTHESIS OF FLAVONOIDS Biosynthesis of flavonoids in plants is via a series of enzymatic steps starting with the aromatic amino acid phenylalanine and acetate19. The initial step in biosynthesis of all flavonoids is the condensation of 4-coumarate coenzyme A (shikimate derived, B ring) with three malonyl coenzyme A molecules (polyketid origin, A ring) to give 2', 4', 6', 4tetrahydroxychalcone, which is catalysed by the enzyme chalcone synthase25. The chalcone is then isomerised to the flavanone naringenin, a key indermediate, which can be converted to several end-products including aurones, isoflavonoids, flavones, flavonols, flavandiols, anthocyanins, condensed tannins, and phlobaphene pigments (Figure 1)25,26.
Figure 1. Schematic of the major branch pathways of flavonoid biosynthesis, starting with general phenylpropanoid metabolism and leading to the nine major subgroups: the colorless chalcones, aurones, isoflavonoids, flavones, flavonols, and flavandiols (gray boxes), and the anthocyanins, condensed tannins, and phlobaphene pigments (colored boxes). The first committed step is catalyzed by chalcone synthase (CHS), which uses malonyl CoA and 4-coumaroyl CoA as substrates. Enzyme names are abbreviated as follows: cinnamate-4-hydroxylase (C4H), chalcone isomerase (CHI), chalcone reductase (CHR), chalcone synthase (CHS), 4-coumaroyl:CoA-ligase (4CL), dihydroflavonol 4-reductase (DFR), 7,2'-dihydroxy, 4'methoxyisoflavanol dehydratase (DMID), flavanone 3-hydroxylase (F3H), flavone synthase (FSI and FSII), flavonoid 3' hydroxylase (F3'H) or flavonoid 3'5' hydroxylase (F3'5'H), isoflavone O-methyltransferase (IOMT), isoflavone reductase (IFR), isoflavone 2'-hydroxylase (I2'H), isoflavone synthase (IFS), leucoanthocyanidin dioxygenase (LDOX), leucoanthocyanidin reductase (LCR), O-methyltransferase (OMT), Phe ammonia-lyase (PAL), rhamnosyl transferase (RT), stilbene synthase (STS), UDPG-flavonoid glucosyl transferase (UFGT), and vestitone reductase (VR).
1.2.3. BIOAVAILABILITY AND METABOLISM OF FLAVONOIDS The absorption, metabolism and excretion of flavonoids is a complex process involving various structural modifications to the ingested flavonoid in multiple tissues and cellular compartments. Determining the bioavailability of flavonoids (the proportion of flavonoid found in blood or target tissue after ingestion) is critical to understanding the effects of flavonoids as chemopreventive agents. Although flavonoids have been shown to undergo extensive metabolism, this does not necessarily equate to biological inactivation of the compound. In many cases in pharmacology, the metabolized product is often more active than the parent compound. A case in point is morphine whose glucuronated metabolite is known to be a more potent opiate than the parent compound27. Flavonoids are predominantly absorbed in the small intestine, with only small amounts absorbed via the gastric mucosa28. The glycosylation state of the flavonoid greatly affects the mechanism of flavonoid absorption. Most flavonoids in nature exist as glycosides. Early studies suggested that flavonoid glycosides were not absorbed intact in the human gut (due to their high hydrophilicity). These findings have more recently been refuted29. Studies have since demonstrated that quercetin glycosides are not only absorbed, but that their absorption is actually enhanced compared to quercetin aglycone30. This absorption is believed to occur partly due to the action of sodium dependent glucose transporter (SGLT1)31. More commonly, however, the first step in absorption of flavonoid-glycosides is usually hydrolysis of the sugar moiety in the gut resulting in generation of the flavonoid aglycone32. This hydrolysis was initially assumed only to occur in the colon by bacteria since humans lack the necessary enzymes to hydrolyze the ȕ-glycoside linkages of flavonoid glycosides. However, recently is has become clear that a broad-specificity ȕ-glucosidase enzyme in enterocytes and lactase phloridzin hydrolase in the small intestine brush border can
hydrolyze these ȕ-glycoside
linkages33. Hydrolysis of flavonoid-glycosides has also been shown to occur in the oral cavity34. The flavonoid aglycones generated by hydrolysis of the sugar moiety are more lipophilic, and hence are more readily absorbed in the gut by passive diffusion. Flavonoids entering the colon undergo a similar hydrolysis by bacterial glucosidases29. Flavonoids that reach the colon undergo ring scission of the aromatic ring after sugar hydrolysis, resulting in simple phenolic compounds, which accounts for the low levels of flavonoid absorption from the colon35. The type of sugar moiety is an important determinant of absorption efficiency, with flavonoid-glucosides much more readily absorbed than flavonoid-rutinosides36. With respect to the flavonoid aglycones, methoxylated flavonoids as opposed to hydroxylated flavonoids are much more readily absorbed owing to their increased lipophicity, Flavonoids lack an active transporter, and as such they are absorbed via passive diffusion, a more efficient process in
hydrophobic flavonoids37. Other factors affecting flavonoid absorption include the protein content of food ingested with the flavonoid. Since flavonoids bind to proteins, flavonoid absorption will be attenuated until the protein is digested38. The biotransformation of flavonoids continues in the enterocytes. The main metabolic transformations include conjugation of glucuronic acid (glucuronidation), methylation and sulphation39. These conjugations are essentially phase II detoxification reactions resulting in increased molecular mass and improved solubility of the compound which enhances excretion of the compound in bile and urine40. Thus the enterocyte is an important site of flavonoid metabolism. Flavonoid aglycones that reach the circulation are bound to albumin. Interestingly, binding to albumin does not affect the antioxidant ability of flavonoids, an important point in terms of the
likely biological effect of absorbed
flavonoids41. Flavonoids entering the
circulation subsequently undergo phase II detoxification in the liver. Other transformations in the liver include the formation of flavonoid-glutathione adducts also resulting in enhanced excretion of the flavonoid39. Excretion of flavonoids occurs in urine as well as bile42. The kinetics of flavonoid absorption and metabolism in humans has been studied for quercetin and other flavonoids. Hollman et al. demonstrated that peak plasma levels of quercetin were reached in 2.9 hours in subjects consuming a meal of 333 grams fried onion43. The mean peak plasma level of quercetin was 196 ng/ml and the half-life of quercetin was 16.8 hours. This long half-life suggests that quercetin may accumulate with continued flavonoid administration. Importantly for purposes of chemoprevention, studies have also demonstrated the accumulation of flavonoids in various animal tissues44. Quercetin, a flavonol, is believed to have different absorption and kinetics to other flavonoid classes. Anthocyanins, in comparison, are absorbed poorly and rapidly excreted in urine. Citrus flavanones are well absorbed but have shorter plasma half lives. They also reach higher maximum concentrations than flavonols45. A significant degree of variability is therefore expected in the pharmacokinetic properties of different flavonoids. As with other ingested compounds, inter-individual variability is another important factor accounting for the pharmacokinetic properties of flavonoids in humans. Food preparation has variable effects on the bioavailability of flavonoids in the diet. Peeling, for example, greatly reduces flavonoid content since the peel contains a large proportion of flavonoids in fruits and vegetables46. The effect of cooking on flavonoid content has been examined for quercetin content in onions after cooking. When onions are boiled, flavonoids diffuse out to enter the broth, making the broth a rich source of flavonoids. Frying onions for 40 minutes did not alter total quercetin content. An increase in quercetin is noted on microwaving due to increased extractability47. Consumption of flavonoids and protein together,
while postulated to reduce the absorption of flavonoids, has not been shown to have an effect48. Thus, in general, flavonoid availability from food appears to be enhanced by cooking.
1.2.4. THE MECHANISMS OF ACTION OF FLAVONOIDS 1.2.4.1. Antioxidant Potential
The ability of flavonoids to function as antioxidants has given them an important place in the field of human health and medicine.Diets high in flavonoids, fruits, and vegetables are protective against a variety of diseases, particularly cardiovascular disease and some types of cancer49.The protective effects of flavonoids in biological systems are ascribed to their capacity to transfer free radical electrons, chelate metal catalysts50, activate antioxidant enzymes51, reduce alpha-tocopherol radicals52, and inhibit oxidases53.The flavones and catechins seem to be the most powerful flavonoids for protecting the body against reactive oxygen species54. Reactive oxygen species (ROS) are highly reactive molecules with both physiologic and pathologic roles. ROS can occur in the form of molecules with highly reactive unpaired electrons known as free radicals (e.g. superoxide, O2.-), or as non-radicals that are highly liable to form free radicals (e.g. hydrogen peroxide, H2O2). They can exist in the body as a result of deliberate synthesis (e.g. production by macrophages for bacterial killing), or as a result of accidental production by metabolic processes such cellular respiration in mitochondria, or via exogenous insults such as smoking55. Many ROS are not in themselves exceptionally reactive; however, in the presence of free heavy metal ions such as copper and iron they generate highly toxic radicals such as hydroxyl ions (OH.). ROS are highly damaging as they can attack lipids in cell membranes, proteins, carbohydrates, and DNA. The resulting oxidative damage may play a role in aging and chronic and degenerative diseases including cancer56-59. About 1-3% of the oxygen we breathe ultimately goes into making ROS, resulting in a huge burden of pro-oxidant free radicals that has to be effectively removed60. The human body relies on both endogenous and exogenous (dietary) anti-oxidant systems to buffer the effect of the ROS constantly produced by metabolic processes. Endogenous antioxidant systems include enzymes such as superoxide dismutase, which converts O2.- into H2O2, and glutathione peroxidase and catalase, that serve to remove H2O2. Non-enzymatic endogenous defense mechanisms have a significant anti-oxidant impact, including buffering by plasma urate and plasma protein thiols. Furthermore, the sequestration of heavy metal ions in binding proteins such as transferrin (iron) reduces the risk of formation of toxic hydroxyl radicals61. Despite these many levels of protection against ROS damage, endogenous antioxidant systems are incompletely efficient in elimination of all ROS, particularly with the added insult of various
environmental ROS from smoking, air pollution etc. Exogenous antioxidant supplementation, from dietary sources therefore has a critical role in the prevention of oxidative stress in human physiology61,62. Antioxidant phytochemicals constitute some of the most important exogenous defense antioxidants in mammalian physiology. Up until the mid-1990’s the dietary phytochemicals most prominently studied for their antioxidant properties were vitamin C, E and the carotenoids63. Polyphenols, which constitute a major group of plant chemicals, only gained interest for their antioxidant effects in the last decade. Flavonoids, the largest group of polyphenols found in nature, appear to be particularly potent antioxidants in vitro. The flavonoids, however, are not all equally effective, with definite structural requirements necessary for the greatest antioxidant effect. The presence of a 2,3 double bond in the C-ring, a catechol structure in the B-ring, and hydroxylation at position 3 and 5 of the A ring appear to impart increased redox potential64. The redox potential of quercetin was similar to ascorbic acid and greater than the redox potential of uric acid65. Direct scavenging of free radicals is one of the major mechanisms of antioxidant activity by flavonoids. The resulting aroxyl radical (Flavonoid-O.) is more stable than other ROS and gains further stability on reacting with a second radical to form a stable quinine structure66. Several other mechanisms of antioxidant activity of flavonoids have been proposed including scavenging of transition metal ions67, and inhibition of enzymes responsible for antioxidant production. In terms of the latter property, flavonoids have been shown to inhibit several prooxidant enzymes including xanthine oxidase68, glutathione S-transferase69, nitric oxide
synthase70, and NADH oxidase71 among others. 1.2.4.2.
Pro-Oxidant Effects
While flavonoids are best known for their anti-oxidant properties, it has been shown that under certain conditions, flavonoids may be pro-oxidant. This property has been proposed to account for several biological effects of flavonoids, such as apoptosis, that are induced in the setting of oxidative stress. These pro-oxidant effects have also been observed for other phenolic antioxidants including tocopherols, ascorbate, urate, curcumin and N-acetylcysteine. The balance between anti-oxidant and pro-oxidant effects of these compounds is dependent on several factors in the cellular environment, particularly on the presence of transition metal ions. The hydroxyl groups of flavonoids account for much of their antioxidant effect. After scavenging ROS, flavonoids are themselves oxidized, with a hydroxyl group now containing a free radical known as a phenoxyl radical. Some flavonoids possess a catechol structure in the B-ring. Oxidation of these flavonoids can result in a semi-quinone radical. The flavonoid semi
quinone can undergo further oxidation resulting in flavonoid quinone. Therefore different types of oxidation of flavonoids occur depending on their exact structure. In addition to scavenging of ROS, flavonoids can also be oxidized in other ways. These include oxidation by cellular peroxidases, or by auto-oxidation in the presence of oxygen- a process greatly accelerated in the presence of transition metal ions. The paradox of ‘antioxidant’ flavonoids is that in the process of scavenging ROS, they become pro-oxidant radicals themselves, albeit less reactive than the scavenged species. The flavonoid radicals never-the-less have undesirable properties. For example, in the presence of transition metals such as Cu+2, flavonoids undergo a series of redox reactions culminating in the genesis of damaging hydroxyl radicals72. Another mechanism involves the flavonoid-quinones that are the products of oxidation in catechol containing flavonoids. These are highly reactive to thiol groups, and result in the formation of flavonoid conjugates with thiol containing proteins such as glutathione. Interestingly, flavonoids that do not form flavonoid-quinones have also been shown to form thiol conjugates, highlighting the complexity of flavonoid chemistry that remains to be fully elucidated73. The pro-oxidant effects of flavonoids have been shown to result in DNA damage and lipid peroxidation in vitro. Pro-oxidant radicals have been demonstrated for several flavonoids including myricetin74, quercetin75,76, proanthocyanidins77, green tea catechins78,79, daidzein80 and baicalin81.
1.2.4.3.
Hormonal Properties
Together with the antioxidant effects of flavonoids, the hormonal, and particularly the estrogenic effects of flavonoids have garnered the greatest attention in flavonoid research over the past 50 years. The estrogenic properties of flavonoids first came to light in the 1950’s when it was observed that sheep grazing on red clover pastures had reduced breeding rates82. Red clover was found to contain several isoflavones and the estrogen-like properties of isoflavones were shown to account for fertility disturbances in animals feeding on red clover. As a result of their estrogen-like properties, isoflavones are also classed as phytoestrogens. Flavonoids from the flavone, flavonols, flavonone and chalcone classes are considerably weaker phytoestrogens than the isoflavones as determined by competitive binding assays83. The precise physiologic effects of flavonoids mediated by their binding to ERĮ and ERȕ are yet to be fully determined. This is an important area of future research because phytoestrogenic flavonoids are consumed in large amounts in the human diet. If the estrogenic effects of these compounds are predominantly pro-proliferative at low concentrations, flavonoids could potentially pose a health risk in terms of promoting hormone dependent cancers.
Flavonoids may also exert anti-estrogenic effects by various enzymatic mechanisms. Blocking the synthesis of estrogens by inhibiting aromatase is an established strategy in the treatment of breast cancer. Flavonoids have been shown to bind the active site of aromatase, and inhibit its function, with flavones and flavanones, rather than isoflavones having the greatest effect84,85. Other enzymes of note in estrogen metabolism include sulfatase and 17ȕ hydroxydteroid dehydrogenase, both of which result in activation on estradiol precursors, and which are inhibited by flavonoids86. The levels of sex hormone binding globulin (SHBG) by flavonoids is also of importance, as excess SHBG can bind estrogen reducing its effect87. The estrogenic effects of isoflavones in humans is supported by studies demonstrating altered menstrual cycle length in women consuming daily soy protein, a product rich in isoflavones88-90. The similarity in structure of flavonoids to all steroid hormones raises the possibility of ligand binding of flavonoids to other members of the nuclear steroid receptor family. Flavonoids have been shown to bind and activate a number of nuclear receptors including androgen91,92, progesterone91, thyroid93, and peroxisome proliferator-activated receptor PPARȖ93. Flavonoids functionally activate androgen receptor mediated transcription, resulting in increased PSA, a major downstream androgen receptor regulated gene. Apigenin, a flavone, was the most effective flavonoid in up-regulating PSA expression94. Interestingly, in a related study other flavonoids were shown to have precisely the opposite effect and inhibit PSA production. It was concluded that unlike the estrogenic effects of flavonoids, the effects on PSA production did not follow a structure-function relationship95. In addition to their sex steroid effects, flavonoids affect several other hormonal pathways. For example, several mechanisms have been described for the potentiation of the vitamin D pathway by genistein. This includes up-regulation of vitamin D receptor gene expression and activity96, and inhibition of enzymes (CYP24) that convert 1,25-vitamin D into less active metabolites97. The vitamin D pathway is increasingly implicated in chemoprevention of prostate cancer, and up-regulation of this pathway is a potentially useful synergistic property of flavonoids. Genistein has also been shown to have a stimulatory effect on insulin secretion in vitro (although this effect is not seen in vivo) and an inhibitory effect on leptin secretion in rats administered genistein98. Flavonoids also inhibit corticosteroid secretion in vitro and in vivo98. Finally, the goitrogenic activity of soy is well documented, especially in the setting of iodine deficiency. This effect is believed to be secondary to inhibition of thyroid peroxidase, a major metabolizing enzyme in thyroxine biosynthesis99. Overall, the interactions of flavonoids with steroid hormone pathways are highly complex. However, the effect of flavonoids on these pathways has been shown to ultimately cause alterations resulting in beneficial effects such as the negative regulation of proliferative stimuli. As with much of flavonoid research, the in vivo
effects of flavonoids on hormonal signaling require further study. This is highlighted in a recent review by Hamilton-Reeves et al, where the majority of intervention studies reviewed did not find a difference in circulating sex steroid hormone levels100. 1.2.5. BIOLOGICAL AND PHARMACOLOGICAL ACTIVITIES OF FLAVONOIDS 1.2.5.1. Antiviral Activity Naturally occurring flavonoids with antiviral activity have been recognized since the 1940s but only recently have attempts been made to make synthetic modifications of natural compounds to improve antiviral activity. Quercetin, morin, rutin, dihydroquercetin (taxifolin), apigenin, catechin, and hesperidine have been reported to possess antiviral activity against some of the 11 types of viruses101. The antiviral activity appears to be associated with the nonglycosidic compounds, and hydroxylation at the 3-position is apparently a prerequisite for antiviral activity. It has been found that flavonols are more active than flavones against Herpes simplex virus type 1102. Recently, a natural plant flavonoid polymer of molecular weight 2,100 daltons was found to have antiviral activity against two strains of type 1 Herpes simplex virus and type 2 Herpes simplex virus103. Because of the worldwide spread of HIV since the 1980s, the investigation of the antiviral activity of flavonoids has mainly focused on HIV104. There have appeared several recent reports on the anti-AIDS activity of flavonoids. Out of twenty eight flavonoids tested, the flavans were generally more effective than flavones and flavonones in the selective inhibition of HIV-1 and HIV-2 or similar immunodeficiency virus infections105.
1.2.5.2. Antifungal Activity A number of flavonoids isolated from the peelings of tangerine orange, when tested for fungistatic activity towards Deuterophoma tracheiphila were found to be active; nobiletin and langeritin exhibited strong and weak activities, respectively, while hesperidin could stimulate fungal growth slightly. Chlorflavonin was the first chlorine-containing flavonoid type antifungal antibiotic produced by strains of Aspergillus candidus106.
1.2.5.3. Antimicrobial Activity
Flavonoids are found to be effective antimicrobial agents against a wide array of microorganism. This is probably due to their ability to complex with extracellular and soluble proteins and also with the bacterial cell wall107. Phenolics present in plants are known to be toxic to microorganisms108. Antibacterial activity of leaf and stem bark of Pterocarpus santalinus was investigated for both gram-positive and gram-negative bacteria109. The stem
bark and leaf extracts showed inhibitory activity against a number of infectious microbial strains including Enterobacter aerogenes and Staphylococcus aureus. The broad spectrum antibacterial activity exhibited by Pterocarpus santalinus may be attributed to its richness in isoflavone glucosides110. Catechins, an important group of flavonoids, have been known to inhibit Vibrio cholerae111, Streptococcus mutans112, Shigella113 and other bacterial strains in vitro. The catechins have been found to inactivate the cholera toxin from V. cholerae107.
1.2.5.4. Anti-Carcinogenic Activity Soybean food comprises a significant portion of Asian diet, where the incidence of breast and prostate cancer is much less than that in the United States114. Studies have found that genistein, an isoflavone from soy can inhibit the growth of various cancer cell lines including leukemia, lymphoma, prostate, breast, lung and head and neck cancer cells115. Another isoflavone present in soy, biochanin A has been found to have cytotoxic effect on cell growth in the mammary carcinoma cell line MCF-7116, myeloid leukemia 117 and pancreatic tumor cells118. Citrus fruit flavonoids, tangeretin and nobiletin have also been shown to inhibit human breast cancer and human colon cancer119.
1.2.5.5. Hepatoprotective Activity Flavonoids have been found to possess hepatoprotective activity. In a study carried out to investigate the flavonoid derivatives silymarin, apigenin, quercetin, and naringenin, as putative therapeutic agents against microcrystin LR-induced hepatotoxicity, silymarin was found to be the most effective one120. The flavonoid, rutin and venoruton, showed regenerative and hepatoprotective effects in experimental cirrhosis121.
1.2.5.6. Cardioprotective Activity In the prevention of cardiovascular disease, many of the observed effects of flavonoids can therefore be attributed to their recognized antioxidant and radical scavenging properties, which may delay the onset of atherogenesis by reducing chemically and enzymatically mediated peroxidative reaction122. Studies suggest that epigallocatechin-3-gallate (EGCG) can suppress reactive oxygen species and thereby prevent the development of cardiac hypertrophy123. Endothelial dysfunction is the pathophysiologic principle involved in the initiation and progression of arteriosclerosis. Some flavonoids have been shown to relax endotheliumdenuded arteries. Genistein, one of the major isoflavones in soy protein, binds to estrogen receptor b124 and can elicit endothelium dependant vasorelaxation in vitro125 and in vivo126.
Other isoflavones such as dihydrodaidzeins have also been reported to enhance endothelial function127. Flavonoids have also been found to be good hypochlorite scavenger in vitro and could have favorable effects in diseases such as atherosclerosis128.
1.2.5.7.
Neuroprotective Activity
Neurodegenerative disorders are a heteregenous group of diseases of the nervous system, including the brain, spinal cord and peripheral nerves, which have different aetiologies. The multifactorial etiology of these diseases suggests that interventions having multiple targets such as flavonoids could have therapeutic potential for them. Moreover, dietary habits and antioxidants from diet can influence the incidence of neurodegenerative disorders such as Alzheimer and Parkinson's diseases129. Flavonoids exhibit biological effects such as anti-inflammatory, antioxidant and metal chelating properties, which augment their role in neuroprotection. Polyphenols such as EGCG, curcumin, extracts of blue berries and Scutellaria are known to help in Alzheimer’s disease (AD)130. In vitro studies show that green tea extract rich in catechins could protect neurons from the amyloid beta-induced damages in AD131. EGCG is also found to be of use in Amyotrophic lateral sclerosis (ALS)132,133 and Parkinson’s disease (PD)134. Extract of Scutellaria stem and polyphenols such as curcumin and naringenin also exhibit neuroprotection in PD135. Alzheimer's disease is characterized by chronic inflammation and oxidative damages in the brain. Curcumin posses antioxidative and anti-inflammatory properties and a protect against oxidative damages or suppress inflammatory damage136.
1.2.5.8. Antimalarial Activity
Licochalcone A, isolated from Glycyrrhiza inflata, was initially proposed as new antimalarial agent in 1994137, and since then, many chalcones, naturally occurring or synthesized, have been identified as potential antimalarial agents, using both molecular modelling and in vitro testing against active parasites. Crotaorixin from Crotalaria orixensis, medicagenin from Crotalaria medicagenia, crotaramosmin, crotaramin and crotin from Crotalaria ramosissima, five prenylated chalcones, were found as inhibitors of schizont maturation against P. falciparum138. Crotaorixin exhibited 100% inhibition of maturation of schizont stage. 2',6'-Dihydroxy-4'-methoxydihydrochalcone exhibited the most potent antiplasmodial activity against of P. falciparum139. Chalcones are not the only flavonoids that have displayed useable antimalarial activity. Several flavones, flavanones, isoflavones and flavonoids glycosides are also reported. Artocarpones A and B, artonin A, cycloheterophyllin, artoindonesianins E, R and A-2,
heterophyllin and heteroflavanone C, 9 flavonoids isolated from of Artocarpus champeden were found to possess strong antimalarial activity of P. falciparum140. Some biflavonoids have also been reported for their antimalarial activity. Sikokianin B and C from Wikstroemia indica141, lanaroflavone, bilobetin, ginkgetin, isoginkgetin and sciadopitysin, a series of biflavonoids isolated from Campnosperma panamensis142,143, Liquiritigenin and isoliquiritigenin isolated from Ochna integerrima144, showed interesting antimalarial activity against several and different chloroquine-resistant strains of P. falciparum.
1.2.5.9. Antileishmanial Activity Recently, a number of synthetic and naturally occurring chalcones with potential antileishmanial activity have been reported. Licochalcone A, one of the many flavonoids isolated from the roots of Chinese licorice, and some related chalcones were shown to inhibit the in vitro growth of both L. major and L. donovani promastigotes together with a remarkably ability to kill intracellular amastigote form of both parasites145. Licochalcone A is not the only chalcones known for its antileishmanial activity. A variety of chalcones was shown to exhibit different levels of activity against Leishmania parasites. A series of 20 chalcones isolated from different plants were screened for their in vitro leishmanicidal activity against extracellular promastigotes of L. donovani146. Electron microscopic studies showed that licochalcone A and some oxygenated chalcones altered the ultrastructure of L. major promastigote and amastigote mitochondria in a concentration-dependent manner and inhibited the respiration of the parasite, as shown by inhibition of O2 consumption and CO2 production by the parasites147,148. Other classes of flavonoids and flavonoid glycosides are also reported for their antileishmanial activity. When evaluated a series of flavonoids for their antileishmanial activity, the majority of the metabolites tested showed remarkable leishmanicidal potential149. Fisetin, 3hydroxyflavone, luteolin, and quercetin were the most
potent. 7,8-Dihydroxyflavone and
149
quercetin appeared to ameliorate parasitic infections . Numbers of flavonoid glycosides have also displayed strong antileishmanial activity against
different
strains
of
Leishmania
parasites.
Quercitrin
(quercetin
3-O-Ĵ-L-
rhamnopyranoside) , kaempferol 3-O-Ĵ-L-arabinopyranosyl (1ĺ2) Ĵ-L-rhamnopyranoside, 150
quercetin 3-O-Ĵ-L-arabinopyranosyl (1ĺ2) Ĵ-L- rhamnopyranoside and 4',5-dihydroxy-3',8dimethoxyflavone 7-O-ȕ-L-glucopyranoside151, isolated from Kalanchoe pinnata, were tested against L. amazonenis amastigotes in comparison with quercetin and afzelin.
1.2.5.10.
Antitrypanosomiasis Activity
Since the available trypanocidal drugs are unsatisfactory as they are often associated with severe side effects, several synthetic and naturally occurring flavonoids have been evaluated for their antitrypanosomal activities. In the survey of a large set of flavonoid aglycons and glycosides for their in vitro activities against T. brucei rhodesiense and T. cruzi, the best in vitro trypanocidal activity for T. brucei rhodesiense was exerted by 7,8-dihydroxyflavone, followed by 3-hydroxyflavone, rhamnetin, and 7,8,3',4'-tetrahydroxyflavone149. The activity against T. cruzi was rather moderate for many compounds. Generally, the tested compounds lacked cytotoxicity in vitro and in vivo149. The most active compound was quercetin 3-methylether, which also showed no blood lysis activity, thus represents a promising compound for use against T. cruzi in blood banks152. Pinobanksin, pinobanksin-3-acetate, pinocembrin, galangin, galangin-3-methylether, pillion, tectochrysin, luteolin, luteolin-7-methylether and quercetin-3-methylether, a series of flavonoids from Lychnophora pohlii, when evaluated against T. cruzi, displayed only moderate activities153. The trypanocidal effect of green tea catechins against two different developmental stages of T. cruzi is also reported. Catechin, epicatechin, gallocatechin, epigallocatechin, catechingallate, epicatechingallate, gallocatechingallate, and epigallocatechingallate were tested on the nonproliferative bloodstream trypomastigote and the intracellular replicative parasite forms154. All the tested compounds displayed not only an outstanding in vitro activity, but were also capable to lyse more than 50% of the parasites present in the blood of infected mice at the same concentrations154. The most active compounds were gallocatechingallate and epigallocatechingallate154.
1.2.5.11. Antiamebiasis Activity As result of their significant antiprotozoal activity, a handful of flavonoids have also been tested for their antiamoebic activity. Apigenin, xanthomicrol, galangin, kaempferol, quercetin, myricetin, tiliroside,hyperfine, reynoutrin, rutin, fisetin, 3,6-dimethoxykaempferol, 4ƍ,5,7trihydroxyflavanone, pinocembrin, pinostrobin, catechin, epicatechin and epigallocatechin were evaluated for their activity against E. histolytica.155 These compounds displayed different levels of activity. Epicatechin, epigallocatechin and kaempferol were the most potent compounds155.
1.3. ANALYTICAL SEPARATION AND DETECTION METHODS FOR FLAVONOIDS
This review focuses on separation and detection methods for flavonoids and their application to plants, food, drinks and biological fluids. The topics that will be discussed are sample treatment, column liquid chromatography (LC), but also methods such as gas chromatography (GC), capillary electrophoresis (CE) and thin-layer chromatography (TLC), various detection methods and structural characterization. Because of the increasing interest in structure elucidation of flavonoids, special attention will be devoted to the use of tandem-mass spectrometric (MS/MS) techniques for the characterization of several important sub-classes, and to the potential of combined diode-array UV (DAD UV), tandem-MS and nuclear magnetic resonance (NMR) detection for unambiguous identification. Emphasis will be on recent developments and trends. An important aspect of flavonoid analysis is whether to determine the target analytes in their various conjugated forms or as the aglycones. In biological fluids (serum, plasma and urine) flavonoids exist as glucuronide and sulphate conjugates. In most cases, only the total aglycone content is determined; therefore, a hydrolysis step is used. However, in plants, medicine and food products, researchers are usually interested in the intact conjugates. In that case, analyses become much more complicated, because the number of target analytes increases significantly: much more selective and sensitive analytical methods are now required. In Figure 2 the principal strategies for the determination of flavonoids in biological fluids, drinks, plants and food – the main sample types – are schematically depicted. The various steps in this flow chart will be considered in some detail below, with attention to both routine procedures and recent developments. Of course, in view of the complexity of the problem (almost) all analytical methods dealing with flavonoids include a high-performance separation method. The choice of the method depends on the sensitivity required for the purpose at hand, the complexity of the biological matrix – which is related to the time spent on sample pretreatment prior to analysis – the required chromatographic resolution and the preferred detection method.
1.3.1. SAMPLE TREATMENT 1.3.1.1. Analyte Isolation Over the years many sample pre-treatment methods have been developed to determine flavonoids in various sample types. There are three main types of flavonoid-containing matrices: plants, food and liquid samples such as biological fluids and drinks (cf. Fig. 2). The solid samples are usually first homogenized, which may be preceded by (freeze-)drying or freezing with liquid nitrogen. The next step is analyte isolation. For this purpose, solvent
extraction (SE) – which may be followed by solid-phase extraction (SPE) – is still the most widely used technique, mainly because of its ease of use and wide-ranging applicability. Soxhlet extraction is used less frequently to isolate flavonoids from solid samples. Liquid samples are usually first filtered and/or centrifuged, after which the sample is either directly injected into the separation system or, more often, the analytes are first isolated using liquid– liquid extraction (LLE) or SPE. As regards SE and Soxhlet, in most cases aqueous methanol or acetonitrile is used as solvent. In the case of LLE the extraction solvent usually is ethyl acetate or diethyl ether containing a small amount of acid. LLE is usually directed at the isolation of aglycones, while the other methods can have the isolation of both aglycones and conjugates as their goal. If aglycones are the target analytes, chemical hydrolysis is usually performed – with
Figure 2. Strategies for the determination of flavonoids in biological fluids, beverages, plants and food. Abbreviations: LLE, liquid–liquid extraction; SE, solvent extraction; MSPD, matrix solid-phase extraction; SPME, solid-phase micro-extraction; SPE, solid-phase extraction; GC, gas chromatography; LC, liquid chromatography; MS, mass spectrometry; MS/MS, tandem mass spectrometry; CE, capillary electrophoresis; TLC, thin layer chromatography; FID, flame ionization detection; ECD, electron capture detection; Q, quadrupole; QqQ, triple-quadrupole; IT, ion-trap, FLU, fluorescence; NMR, nuclear magnetic resonance; TOF, time-of-flight and ED, electrochemical detection.
hydrochloric acid or formic acid at elevated temperatures (80–100 ƕC) or by refluxing with acid in the presence of ethanol – but enzymatic hydrolysis with ȕ-glucuronidase or ȕ-glucosidase is also used157. If the interest is in the intact flavonoid-glycosides, hydrolysis should of course be prevented. This means that harsh extraction conditions and heating should be avoided. Furthermore, the activity of hydrolyzing enzymes that may be released during milling of plant material can be inhibited by addition of, e.g. tris(hydroxymethyl)aminomethane. For more detailed information, the reader is referred to papers by Robards and co-worker who recently, reviewed various sample preparation procedures for flavonoids158,159. In other reviews these procedures received attention in the context of particular application areas, e.g. soy food and human biological fluids160 and fruits161. Sample treatment by means of SPE, matrix solid-phase dispersion (MSPD) and solidphase micro-extraction (SPME) deserves some more attention. Although SPE is not a very new technique, it has only recently been applied in flavonoid analysis. Moreover, compared with traditional extraction methods, the techniques mentioned above can be easily automated, while solvent consumption is lower and analysis times are shorter.
1.3.1.2. Solid-Phase Extraction Non-selective SPE on, typically, alkyl-bonded silica or copolymer sorbents is widely used for analyte extraction and enrichment from aqueous samples and sample extracts—primarily in environmental, pharmaceutical and biomedical analysis. Its use in flavonoid analysis is, however, relatively new. In most cases the sorbent is C18-bonded silica and the sample solution and solvents are usually slightly acidified to prevent ionization of the flavonoids, which would reduce their retention. A recent example is the purification of methanolic extracts of olives162. After SE of the homogenized olives, the extract was evaporated to dryness, redissolved in water containing hydrochloric acid (pH 2) and loaded on a C18 sorbent. After washing with hexane to remove lipids, the flavonoids were eluted with pure methanol. Combining this procedure with liquid chromatography (LC)–ESI(+)–MS resulted in the identification of up to eleven phenolic compounds in 29 types of olives. These included several flavonoid-(di)glycosides. Another recent application is the determination of daidzein (16) and genistein (17) in plasma using LC–ESI(−)–MS/MS163. Two hundred and fifty microliters of plasma were diluted and acidified with 0.5% formic acid before application to a C18 sorbent. Dilution and acidification were required to obtain satisfactory recoveries (ca. 80%), probably due to reduced flavonoid–protein interaction. After elution with methanol, offline combination with the LC procedure gave limits of detection (LODs) of 3 and 9 ng/ml for genistein and daidzein, respectively, when using multiple reaction monitoring (MRM).
HO
2
2
2
HO
OH
16
OH
2
OH
17
In a less traditional application, SPE was used on-line after an LC separation but prior to MS and NMR detection, to effect enrichment of the analytes of interest in an oregano sample extract164. The use of expensive deuterated solvents during the LC separation could now be avoided – for LC a traditional acetonitrile – aqueous ammonium formate buffer gradient was used – but no solvent suppression was required since the flavonoids were eluted from the C18 sorbent with (a limited volume of) deuterated acetonitrile. A multiple trapping process was used to further concentrate the analytes and thereby reduce the acquisition time of the off-line NMR measurements. With this LC–monitoring UV–SPE/NMR–MS method five flavonoids, a phenolic acid and a monoterpene were identified in the oregano sample. For the analysis of a red clover extract a dual-SPE method was used165. The methanolic extract was subjected to SPE and fractions were collected and transferred to a second SPE sorbent. Three sorbents were tested and the pH and organic mole fraction of the aqueous organic solvent were varied. Optimum conditions were created by applying the extract to a C18 sorbent, washing with methanol–water (35:65, v/v) containing 2% acetic acid, and eluting with a methanol–water mixture with an organic-solvent proportion increasing from 0 to 90%, and containing 2% ammonium hydroxide. With the same sorbent as in the first step, in the second step a mixture of 80% methanol containing 2% ammonium hydroxide was used to completely elute all analytes from the sorbent. Unfortunately, no recovery data are provided in the paper to show the beneficial effect of the second SPE step. LC–ESI(+)–MS of the purified extract enabled the provisional identification of 49 flavonoids, including several acylated flavonoidglucosides. A relatively new SPE method uses a molecularly imprinted polymer (MIP) as the sorbent. MIPs, typically, are highly selective for the target analyte and usually have good mechanical and thermal stability166,167. A MIP was used to determine quercetin (18) in red wine166. The recovery was over 98% when using methanol containing 15% acetic acid or acetonitrile containing 10% aqueous triethylamine as eluent. Fig. 3 shows that use of the MIP greatly reduced the complexity of the LC chromatogram and enhanced the intensity of the quercetin peak. Unfortunately, the MIP was not fully selective for quercetin; structurally related
2+
2+
2
+2
2
+2
2+
2+
2
2+
+2
2+
2+
18
2
19
compounds such as morin (19) and (+)-catechin also showed affinity. A general disadvantage of MIP–SPE is that a specific MIP has to be designed for each application and that the method is, in principle, not applicable to other analytes. That is, it can only be used for target analysis and not for screening purposes, while that is the main objective of most flavonoid analyses.
Figure 3. LC–UV265 of Merlot wine without (2) and with (1) molecular imprinted polymer-SPE (elution with acetonitrile)166.
1.3.1.3. Matrix Solid-Phase Dispersion MSPD enables the extraction of analytes from samples homogeneously dispersed in a solid support, usually a C18- or C8-bonded silica. In this way, sample extraction and clean-up are
carried out simultaneously with, generally, good recoveries and precision. MSPD is
frequently used to determine pesticides in, e.g. fruits, vegetables, beverages and foods (e.g.168171
), but application to flavonoid analysis172,173 was reported only recently. For the LC–ESI(+)–
MS determination of isoflavone aglycones and glycosides in Radix astragali – the dried root of Astragalus membranaceus that is widely used in Chinese medicine – MSPD was compared to Soxhlet and ultrasonic extraction172. For MSPD C18-bonded silica was used and elution was carried out with methanol–water (9:1, v/v). Careful optimization of the eluent composition was needed to prevent co-extraction of interfering matrix components and reduced isoflavonoid yields. For the aglycones, MSPD gave the best extraction efficiency (mean recovery for formononetin (20), 83%), but for the glycosides Soxhlet gave better results (ononin (21): UV260 peak area ratio Soxhlet/MSPD, 4.3). However, Soxhlet extraction required 10-fold more sample and solvent and the extraction time was much longer. Ultrasonic extraction gave rather poor results (formononetin: UV260 peak area ratio ultrasonic/Soxhlet/MSPD, 1/3/4), especially for the aglycones.
+2
2
2
*OX2
2
2
2&+
2&+
20
21
A similar MSPD procedure was used to obtain analyte enrichment and sample clean-up for LC–NMR analysis of leaves of red clover173. This approach provided sufficiently high concentrations of the seven main isoflavones in these leaves to permit their unambiguous identification while using a mere 500 mg of sample. In this case, MSPD-based sample preparation has the disadvantage that it is somewhat more time-consuming than SE, and therefore more prone to (partial) hydrolysis in the case of flavonoid conjugates. Furthermore, compared with SE the extraction efficiency for the glucosides was found to be lower. Obviously, a systematic study of sorbent materials is urgently needed. Finally, a study on flavanones and xanthones in the root bark samples of M. pomifera should be mentioned174. SE, MSPD with a C18-bonded silica and, as a novel approach, MSPD
with sea sand were used. For SE, 150 mg dry root bark were soaked in dichloromethane (DCM) or methanol–water (9:1, v/v). For the MSPD procedures 150 mg of dry root bark were mixed with 600 mg of the C18 sorbent or sea sand and 2 ml hexane, packed into a column and eluted with DCM or methanol–water (9:1, v/v). The best results were obtained with the sea sand procedure, with C18-MSPD in second, and SE in last place: when using sea sand, the analyte responses in LC–UV were about 25% higher than with SE. This seems to suggest that, for this application, analyte losses due to incomplete extraction were more important than sample cleanup. In the root bark extracts five prenylated xanthones and two prenylated flavones were found. The LOD of one of these, macluraxanthone, was 3 ȝg/g. It has to be noted that the authors do not give any information on the sea sand. Apart from its function as a sample disruptor it would be expected to have sorbent properties in order to give better results than C18. The influence of an MSPD-type treatment on the (partial) hydrolysis of glycosides has not been studied in any of the quoted papers. This is an aspect that requires further attention.
1.3.1.4. Solid-Phase Micro-Extraction In SPME a fused-silica fibre coated with polyacrylate or polydimethylsiloxane as a stationary phase is used to extract analytes from a liquid or gaseous sample, or from the headspace above a liquid sample. As is true for SPE, the procedure can effect considerable analyte enrichment. SPME is a straightforward technique and organic solvent consumption is less than in SPE. On the other hand, because it is an equilibrium method, analyte recoveries can be quite low while extraction times frequently are as long as 60 min. SPME is generally combined with gas chromatography (GC) analysis for the extraction of (semi-)volatile organic compounds from environmental, biological and food samples175,176. SPME has also been coupled with LC to analyze non-volatile and/or polar compounds175, although this is, in our view, a rather unfortunate and laborious combination and one that is, in the case of flavonoids, not really required because of the many satisfactory alternatives. Nevertheless, two such examples for flavonoid analysis are discussed below177,178. Satterfield and Brodbelt177 used SPME to extract genistein (17) and daidzein (16) from human urine in combination with LC–ESI(+)–MS analysis. A Carbowax-templated poly(divinylbenzene) resin proved to be the best fibre type, with a 5-min extraction at pH 4 and a temperature of 35 ƕC. Addition of sodium chloride to aqueous standard solutions of genistein and daidzein gave lower recoveries and caused the formation of sodium ion adducts that interfered in selected reaction monitoring (SRM) ESI(+)–MS. The LODs were 25 and 3 pg/ml
for daidzein and genistein, respectively. Concentrations detected in urine 3 h after consumption of 35 g soy protein were 16 ng/ml for both analytes. To improve the robustness of the SPME procedure, Mitani et al.178 used an open-tubular fused-silica capillary column instead off a fibre. The authors determined the same two analytes in soybean foods using on-line in-tube SPME–LC–DAD UV. An in-tube approach enables automation and usually provides better performance characteristics than manual techniques. Optimum extraction conditions for standard solutions were obtained with 20 draw/eject cycles of 40 ȝl of sample using a porous-layer open-tubular capillary column; the total extractionplusdesorption time was 30 min. Analyte recoveries from spiked food were above 97% in all cases. Unfortunately, compared with the earlier study177, the LODs were about 50-fold higher, i.e. about 0.5 ng/ml. In the absence of any further interpretation of the results by the authors of the paper178, we do not know how to explain the outcome of their study. Admittedly, in the intube SPME study an extra hydrolysis step was used since, according to the authors, the hydrophilic-glucosides were difficult to adsorb to the capillary and only the aglycones could be satisfactorily extracted—but this cannot be considered to account for such a large difference. In the present instance, the poor performance was no real stumbling-block because the concentrations of the aglycones were in the 3–450 ȝg/g range.
1.3.2. SEPARATION AND DETECTION The present section will be mainly devoted to LC-based methods (Section 1.3.2.1) because these are by far the most important ones in flavonoid analysis (cf. Fig. 2). Less common procedures involving GC, capillary electrophoresis (CE) or thin-layer chromatography (TLC), will be discussed in Section 1.3.2.2.
1.3.2.1. Column Liquid Chromatography 1.3.2.1.1. General LC of flavonoids is usually carried out in the reversed-phase (RP) mode, on C8- or C18bonded silica columns. However, also other phases, such as silica, Sephadex and polyamide are used. Gradient elution is generally performed with binary solvent systems, i.e. with water containing acetate or formate buffer, and methanol or
acetonitrile as organic modifier.
Phosphate buffers are less popular than they used to be, mainly because of the dreaded contamination of ion sources when MS detection is used. LC is usually performed at room temperature, but temperatures up to 40 ƕC are sometimes recommended to reduce the time of analysis and because thermostated columns give more repeatable elution times. If the main aim
Table 2. Selection of Recent Publications on LC Analysis of Flavonoids
of the study is to determine the major flavonoids in a sample, run times of 0.5–1 h usually suffice to separate the five to ten compounds of interest (e.g.179,180). If, on the other hand, a more exhaustive separation of constituents is intended, run times of up to some 2 h may well be required. Under such conditions, some 30–50 compounds can easily be separated (and identified) in a single run, with many conjugates such as glycosides, malonates and acetates frequently being included (e.g.181,182). To quote two extremes, when using a special coated silica column, Huck et al.183 needed only 5 min to separate five main aglycones—while a striking exception on the high side is found in a 340-min run for the LC of isoflavones in soy sauces for pattern recognition analysis184. Table 2 summarizes some typical examples of LC separation conditions reported in the recent literature156. Scrutiny of the text and comparison of the eluent compositions and gradients used in the quoted, and also other, papers reveals that it is often difficult to find out how, and with which main goal, optimization was carried out. Moreover, more recent papers usually do not discuss why elution conditions were selected which differ from those in earlier studies. In several publications, instead of linear gradients, rather complicated gradient profiles are used, comprising several steps and applying various slopes, without any explanation. Obviously, trial-and-error often plays a rather large role. Two exceptions are briefly discussed below. For the analysis of phenolic compounds in beer with LC coupled to electrochemical detection (LC–ED), separation conditions were optimized for a standard mixture of several flavone aglycones and glycosides185. Eleven different stationary phases (all C18-bonded silicas) were compared with column dimensions of (100–250) mm x (2.0–4.6) mm I.D. The pH and gradient – using water and acetonitrile, with ammonium acetate and formic acid to adjust the pH – were optimized for each column. Acetonitrile was preferred to methanol, which often caused a high baseline noise. On the basis of the experimental evidence, four columns were
selected: gradient elutionwas done at pH 3.14 for all of these, but the gradient profiles were slightly different for each of them. Flow rates providing the best resolution and repeatability varied from 0.23 to 0.9 ml/min, probably because the column I.D.s varied from 2.1 to 4.6mm185. To our opinion, the variation of too many parameters makes it difficult to reach a proper conclusion regarding the (dis)advantages of the various columns. Rauha et al.186 studied the influence of the LC eluent composition on the ionization efficiency of five flavonoids in atmospheric pressure chemical ionization (APCI), ion-spray (IS) and atmospheric pressure photoionization (APPI) MS. The effects were found to be considerable. For example, in positive ion (PI) IS and APCI, using 0.4% formic acid as the aqueous component of the LC eluent yielded optimum ionization conditions in contrast with an ammonium acetate buffer of pH 4.0 in the case of negative ion (NI) IS and APCI. The largest effects were obtained for APPI, where pure water gave the best results—with the final choice being ca. 5mM ammonium acetate to create satisfactory LC behaviour of the analytes. With all techniques, the NI mode gave better results than the PI mode, mainly because of lower background noise. Analyte detectabilitywas of the same order of magnitude in all cases, with IS giving marginally better results. The effects of eluent composition on ionization in MS will be further discussed in Section 1.3.2.1.3. According to a recent review187, stereochemistry is not often discussed in the flavonoid literature. In 1991, a ȕ-cyclodextrin-bonded phase, Cyclobond I, was used in the reversed-phase and normal-phase mode to separate the 2R and 2S diastereomers of flavanone glycosides and benzoylated flavanone glycosides188. Other papers discussed the enantiomeric separation of flavanones189 and the diastereomeric separation of flavanone-glycosides190.
1.3.2.1.2. Detection in LC 1.3.2.1.2.1. UV Absorbane Detection All flavonoid aglycones contain at least one aromatic ring and, consequently, efficiently absorb UV light. The first maximum, which is found in the 240–285 nm range, is due to the Aring and the second maximum, which is in the 300–550 nm range, to the substitution pattern and conjugation of the C-ring191. Simple substituents such as methyl, methoxy and nondissociated hydroxyl groups generally effect only minor changes in the position of the absorption maxima. Already several decades ago, UV spectrophotometry was, therefore, a popular technique to detect and quantify flavonoid aglycones. More recently, UV detection became the preferred tool in LC-based analyses and, even today, LC with multiple-wavelength or diode-array UV detection is a fully satisfactory tool in studies dealing with, e.g. screening, quantification of the main aglycones and/or a provisional sub-group classification. It will be clear that what has been said above for the detection and characterization of the aglycones, is also true for their conjugates. Generally speaking, this facilitates the recognition
of so-called satellite sets, comprising aglycones, their glycosides, glycoside-malonates and, in some cases, glycoside-acetates (see Fig. 4). Unfortunately, most glycosides and acyl residues are poor chromophores; consequently no further distinguishing can be achieved by means of DAD UV detection. As regards the potential and limitation of satellite-set recognition, Fig. 4 shows that – while even the identification of two flavonoids from the same sub-class, the isoflavones – does not present a real problem, the UV spectra of the various members of each set (peaks 1–3, 7 and 4–6, 8) are mutually indistinguishable. Characteristic UV spectra of four of the main classes of flavonoids are shown in Table 3. Details on wavelengths of maximum absorption and molecular extinction coefficients, İ , for over 150 flavonoids can be found in an early work by Mabry et al.191. The general experience is that the spectra in ethanol, methanol and acetonitrile are essentially the same and that log İ max values of the main absorption band are on the order of 3.4–4.6. One should be aware of the fact that chromophores with ionizable groups will show pH dependency; this is also discussed below in the section on fluorescence detection. The spectra included in Table 3 and the Ȝmax data summarized in Table 3 clearly indicate that: (i) the various flavonoid sub-classes can indeed be provisionally distinguished from each other, i.e. that LC–DAD UV is an interesting complementary tool during structural characterization (see Section 1.3.3) and (ii) a limited number of monitoring wavelengths suffices for a general flavonoid screening: flavonoid detection is usually carried out at 250, 265, 290, 350, 370 and/or 400 nm (with an added wavelength in the 500–525 nm range if anthocyanidins are included187). The modest losses of analyte detectability caused by the selection of less than fully optimized detection wavelengths are generally considered acceptable. LODs down to 1–10 ng (injected mass) are repeatedly
Figure 4. (A) RPLC–UV250 and (B) RPLC–FLU (ex, 250 nm; em > 450 nm) of an extract of T. pratense leaves. Peak numbering: (1) an isomer of FGM, (2) formononetin–7-O-ȕ-D-glucoside, (3) formononetin–7O-ȕ-D-glucoside–6'-O-malonate (FGM), (4) an isomer of BGM, (5) biochanin A–7-O-ȕ-D-glucoside, (6) biochanin A–7-O-ȕ-D-glucoside–6'-O-malonate (BGM), (7) formononetin and (8) biochanin A.
Table 3. Ȝmax of Some Representative Flavonoids of the Main Sub-Classes, with Examples of Each Class.
reported192,183. This implies that, for an injection volume equivalent to 1 g or 1 ml of original sample, concentration LODs in the low ng/g range can be obtained. In real samples often much higher concentrations are encountered for the most abundant – and, consequently, most relevant – target aglycones, and analysis will not be too demanding.
2+ 2+
2
+2
2+
2+
2
+2
2
+2 2+
2+
2
22
2+
2
2+
23
2
24
In order to illustrate the general usefulness and application of LC–(DAD) UV the analysis of an H. stoechas extract193 is presented in Fig. 5. The DAD UV spectra shown as inserts A–M illustrate the widely differing spectral characteristics of various flavonoid subclasses. The spectra played an important role in the identification: for instance, peaks G and H have the same aglycone skeleton (naringenin (22)). Based on the mass spectrum (also recorded) peak J could either be a kaempferol (23) or a luteolin (24) conjugate, but the DAD UV spectrum was only consistent with the former. A similar study was reported on flavones and isoflavones in a G. tinctoria extract194.
Figure 5. LC–DAD UV of an H. stoechas extract ((Ȝ = 270 nm) with the spectra recorded at the peak apices as inserts A–M193.
1.3.2.1.2.2. Fluoresene Detection In flavonoid analysis, fluorescence detection is used only occasionally, because the number of flavonoids that exhibit native fluorescence is limited. For these compounds, LODs in LC and CE are typically about an order of magnitude lower than with UV detection. Moreover, their fluorescence facilitates selective detection in complex mixtures195. Classes of flavonoids that show native fluorescence include the isoflavones196, flavonoids with an OH group in the 3position, e.g. 3-hydroxyflavone197 and catechin [trans-3,3&,4&,5,7-pentahydroxyflavan]198, and methoxylated flavones, e.g. 3&,4&,5&-trimethoxyflavone199. As an example, Fig. 6 illustrates the selectivity of fluorescence detection for the determination of 3&,4&,5&-trimethoxyflavone in an extract of Flos primulae veris. Fluorescence detection (LOD (S/N = 3), 25 ȝg/l) was 10-fold more sensitive than UV detection. However, with MS in the selected ion monitoring (SIM) mode, detectability was even better (LOD, 5 ȝg/l)199. When fluorescence detection is used in combination with UV it offers the possibility to discriminate between fluorescent and nonfluorescent co-eluting compounds200,195.
Figure 6. Comparison of detection methods for 3&,4&,5&-trimethoxyflavone in an extract of Flos primulae veris: (a) RPLC–UV216; (b) RPLC–FLU (ex, 330 nm; em, 440 nm) and RPLC–ESI(+)–MS; in (c) and fullscan and (d) extracted ion chromatogram of m/z 312–314199.
The nature of the functional groups and their substitution pattern determine whether a particular flavonoid is fluorescent or not. For example, from amongst the isoflavones, only those that do not have an OH group in the 5-position show strong native fluorescence—as is true for compounds 1–3 and 7 in Fig. 4B. Such compounds exhibit large Stokes’ shifts, possibly due to a change of the molecular structure of the molecule from non-planar in the S0 state to planar in the S1 state, with an accompanying change in electric dipole moment. These large shifts create a high selectivity over impurity fluorescence from the matrix, since it enables the use of long emission wavelengths for detection196. The large Stokes’ shifts for formononetin (20) and its glucoside, ononin (21), are shown in Fig. 7. Interestingly, the fluorescence excitation and, though less so, emission spectra of formononetin also show a pH dependence; an extra band at 340 nm starts to come up at pH 6, the pKa value of the analyte. At higher pH the compound is predominantly in its anionic form. The shift in emission is much less pronounced, i.e. from 495 nm for the neutral molecule to 470 nm for the anion. As expected, for ononin no such effect is observed since the glucose substituent prevents ionization.
Figure 7. Fluorescence excitation and emission spectra of: (A) formononetin and (B) ononin in methanol– water (1:1, v/v) at the pH values indicated196.
As stated above, some flavonols also show native fluorescence. Here, the 3-OH group is involved in excited-state intramolecular proton transfer, which causes solvent-dependent dual emission, i.e. two emission bands show up of which the ratio depends on the solvent composition. This phenomenon has been studied extensively in the literature (see e.g.197,201,202) and will not be further discussed here. To extend the application range of fluorescence detection, derivatization has been used. For example, quercetin (18), kaempferol (23) and morin (19), with their 3-OH, 4-keto substituents, can form complexes with metal cations, some of which are highly fluorescent203,204. Hollman et al.203 used postcolumn derivatization for the determination of quercetin and kaempferol, based on the formation of fluorescent complexes with Al(III). LODs were found to be 0.15 and 0.05 ng/ml for quercetin and kaempferol, respectively. The method was used to study the bioavailability of quercetin from onions and apples in humans205. Plasma levels of quercetin of nine individuals were measured over a 36-h period. Peak levels in the plasma were reached within 0.7 h after ingestion of onions (220 ng/ml), 2.5 h after ingestion of apples (90 ng/ml) and 9 h after ingestion of quercetin rutinoside (rutin, 14) (90 ng/ml).
1.3.2.1.2.3. Eletrochemial Detection Since most flavonoids are electroactive due to the presence of phenolic groups, electrochemical detection can also be used. Although ED is not as sensitive as fluorescence detection, LODs can be quite low: for trans-resveratrol in rat blood an LOD of 2 ȝg/l was obtained using LC–multichannel-ED206. In a recent paper isoflavones in soybean food and human urine were determined by LC–coulometric array-ED207. LC–UV–MS was used for identification purposes. The coulometric electrode array detector consisted of a 6- ȝl flowthrough analytical cell containing an Ag/AgCl reference electrode, a platinum wire counterelectrode and eight porous graphite working electrodes (carbon paste). For standard solutions of daidzein (16) and genistein (17) the highest signal was found at 450mV, most likely corresponding to an oxidation signal. Calibration plots were linear for genistein but not for daidzein, probably due to saturation of the electrode surface by the analyte. Under optimum conditions (eluent: acetonitrile–acetate buffer) the LODs for daidzein and genistein were 400 pg/ml. In several soybean foods, the concentrations of daidzein and genistein were 20–200 and 60–300 ȝg/g, respectively (recoveries, 95–107%). In human urine, only the corresponding glucosides, daidzin (25) and genistin (26), were found; their concentrations were ca. 5 ȝg/g.
*OX2
2
2
2
*OX2
2+
25
2+
2
2+
26
Peyrat-Maillard et al.208 used RPLC–ED to evaluate the anti-oxidant activity of phenolic compounds, including eleven flavonoids, by measuring the accelerated auto-oxidation of methyl linoleate in anhydrous dodecane, under strongly oxidizing conditions. For all flavonoids, two maxima showed up in the peak area vs. potential voltammograms, as shown for rutin (27) in Fig. 8. The first maximum corresponds to the oxidation of the phenolic substituents on ring B, the second one probably comes from the other less oxidizable phenolic groups. For the flavonoids there was no clear linear relation between the anti-oxidant activity and the ED signal, since various structural parameters play a role. For instance, glycosylation of the 3hydroxyl group decreases the antioxidant or antiradical activity in flavonols, but not their electrochemical behaviour. The decrease can be attributed to the steric hindrance posed by the glucose group or to poor solubility in the alkane solvent used in these experiments. However, for most analytes, the antioxidant efficiency could be related to the value of the first maximum, which corresponds to the lowest energy required to donate an electron.
Figure 8. Voltammogram and structure of rutin. MDRP: maximal detector response. Modified from208.
1.3.2.1.3. LC-MS In flavonoid analysis, MS is the state-of-the-art detection technique in LC. In this section we confine ourselves to single-stage MS. In most cases single-stage MS is used in combination with UV detection to facilitate the confirmation of the identity of flavonoids in a sample with the help of standards and reference data. For the identification of unknowns, tandem mass spectrometry (MS/MS or MSn) is used—a technique that deserves a separate discussion, which is presented in Section 1.3.3.2. In the LC–MS of flavonoids – as in many other application areas – atmospheric pressure ionization interfaces, i.e. APCI and electrospray ionization (ESI), are used almost exclusively today. Both positive and negative ionization are applied. ESI is more frequently used in flavonoid analysis, but APCI is gaining in popularity and in some cases better responses are obtained in that mode209-211. According to most studies, for both APCI and ESI the NI mode provides best sensitivity. However, the PI mode should not be neglected, since useful complementary information is often obtained in studies dealing with the identification of unknowns. For the rest, one should be aware that, with all four modes of operation, analyte responses can vary considerably – and rather unexpectedly – from one sub-class to another, and even within one class209,186. In addition, the composition of the LC (gradient) eluent, its pH and the nature of the buffer components added, can have a distinct influence (e.g.209,212), as was discussed in Section 1.3.2.1.1. for a study by Rauha et al.186. In flavonoid analysis, the most common additives are acetic acid213, formic acid214, ammonium-acetate and ammoniumformate215,209. Trifluoroacetic acid has also been used216 although it is known to suppress the ionization due to ion-pairing and surface-tension effects. The mass spectra of flavonoids obtained with quadrupole and ion-trap instruments typically are closely similar, even though relative abundances of fragment ions and adducts do show differences209. Therefore, direct comparison of spectra obtained with these two instruments is allowed. The main advantage of an ion-trap instrument is the possibility to perform MSn experiments, which enables the confirmation of proposed reaction pathways for fragment ions209. Next to ESI and APCI, ionization techniques such as electron ionization (EI)217, chemical ionization (CI)218, fast atom bombardment (FAB)219, and matrix-assisted laser desorption ionization (MALDI)220-222 are also used. The potential of off-line MALDI-TOF MS for flavonoid analysis was explored recently for flavonoids in red wine and fruit juices220, soy products221, and onions and green tea222. Samples were pre-treated with SPE and preparative LC and the collected flavonoid- and anthocyanin-containing fractions were analyzed by MALDITOF MS. Of the various MALDI matrices tested, 2,4,6-trihydroxyacetophenone (THAP)
proved to be the best for flavonol-glycosides from red wine and fruit juices and for anthoyanins from onions and tea, whereas 2,5-dihydroxybenzoic acid (DHB) was the preferred matrix for isoflavones from soy products. Fig. 9 shows MALDI–MS spectra of daidzin (25) and genistin (26), recorded with four different matrices. In the spectra of all flavonoids, the only ions detected were the protonated molecules and the minor sodium and potassium adducts; for the glycosides, glycosidic cleavage was observed. Contrary to studies of flavonoid–glucoside– malonates with ESI– and APCI–MS173,223, in the MALDI-TOF mass spectra of the glucosidemalonates in the soy samples, no loss of the malonate moiety was found. Sample clean-up with SPE was invariably needed because the matrix components with masses between 700 and 900 Da suppressed flavonoid ionization. Anthocyanins in juice extracts showed a linear response with concentration in the range of 1–10 mg/l, but rather unexpectedly the relative responses of the anthocyanin-diglucosides were four times lower than those of the monoglucosides. According to the authors, quantification is possible by using an internal standard.
Figure 9. MALDI(−)-TOF MS and the performance of four MALDI-TOF MS matrices: (A) 2,5dihydroxybenzoic acid (DHB); (B) 2,4,6-trihydroxyacetophenone (THAP); (C) 2-(4-hydroxyphenylazo)benzoic acid (HABA) and (D) 3-aminoquinoline (3-AQ), for the isoflavones: M1, daidzin (6.3×10−4 M in 70% methanol) and M2, genistin (6.7×10−4 M in 70% methanol)221.
In LC–MS, sets of flavonoids with the identical aglycon mass and which comprise, e.g. the glucoside-malonate, the glucoside and the aglycone – also called satellite sets – are easily recognized. In T. pratense, for example, satellite sets of the main isoflavones formononetin (20) and biochanin A (28) were found, which consisted of the glucoside-malonate, an isomer, the glucoside and the aglycone. Thirteen of such satellite sets were found, comprising 40 isoflavones. It lies at hand to conclude that, when screening a plant extract for unknowns, knowledge of the presence of satellite sets224 simplifies target analysis. However, a note of warning should be added. The literature shows that, even within the same plant species, various authors find different flavonoid conjugates. For example, in the paper cited above mainly glucoside-malonates were reported, whereas in another study on flavonoids in T. pratense, also several glycoside-acetates were found165. In an LC–APCI(−)–MS study of four leguminous plant species, the flavonoid patterns were found to be widely different: in Trifolium pratense L. and Trifolium repens L., the main constituents were flavonoid glucoside-(di)malonates, while Trifolium dubium L. and L. corniculatus L. mainly contained flavonoid (di)glycosides. Remarkably, those observations markedly differed from those reported in other papers, which sometimes also differed from each other225,165. This may be (partly) due to differences in environmental factors, growth conditions, etc.—an aspect that, until now, has not been studied in any detail224.
2
+2
2+
2 2&+
28 The structural characterization of flavonoid-glycosides by means of LC–MS/MS will be discussed in Section 1.3.3.2.4.
1.3.2.2. Less Common Methods 1.3.2.2.1. General In this section three separation techniques will be considered, i.e. GC, CE and TLC, which are used less frequently than LC. The renewed attention for classical techniques such as GC and TLC can be called somewhat surprising. On the other hand, CE is a relatively novel technique, and has only been used for flavonoid analysis in the last 10 years. Techniques like high-speed counter-current chromatography (HSCCC) and supercritical fluid chromatography (SFC) are
not considered here. Their role in flavonoid analysis is limited and they are both discussed quite elaborately in the 2004 review of Tsao and Deng226, which deals with separation techniques for phytochemicals.
1.3.2.2.2. Gas Chromatography Gas chromatography was used for the analysis of flavonoids already in the early 1960s. In the first paper on this topic227, derivatized flavonoids were separated on a semi-preparative scale using a SE-30 silicone polymer column with subsequent thermal conductivity detection; fractions were collected for IR and UV–vis spectroscopy. After the introduction of LC, GC analysis of flavonoids became less prominent, but recently it received renewed attention (e.g.228230
, possibly because of the developments in high-temperature GC and the introduction of
improved derivatization procedures—topics that will be discussed below. However, most recent GC studies use conventional temperature programmes and derivatization methods. GC-based methods provide high resolution and low detection limits, but they are labourintensive because derivatization – in most cases directed at the formation of trimethylsilylether (TMS) derivatives – is unavoidable to increase the volatility of the flavonoids and to improve their thermal stability. It should be noted that for flavonoids with more than one hydroxyl substituent methylation may yield several derivatives, which makes quantification difficult. The separation conditions have not changed much since the early 1960s although, today, fused silica capillary columns are used instead of packed glass columns. Most recent papers on the GC analysis of flavonoids are in the biological and nutritional area, and focus on the flavonoid antioxidant activity, metabolism or taxonomy. Little attention is paid to method development and, to our opinion, LC would have been a better choice in many cases. Typically in GC, flavonoids are hydrolyzed and converted into their TMS derivatives, injected onto a non-polar DB-5 or DB-1 column in the split or splitless mode and separated with a linear 30–90 min temperature programme up to 300 ƕC. N,O-bis(trimethylsilyl)-trifluoroacetamide (BSTFA) and N-(tert-butyldimethylsilyl)-N-methyltrifluoroacetamide (TBDMS) are the most commonly used derivatizing agents, and EI–MS in the selected ion monitoring mode with a source temperature of up to 250 ƕC is often used for detection. The molecular ion, [M+H]+, and fragments formed by the loss of CH3 and/or CO and retro-Diels–Alder (RDA) reactions are typically used for detection. The MS fragmentation is discussed in Section 1.3.3.2. To determine the genistein (17) and daidzein (16) contents of fruits and nuts, freeze-dried samples were extracted with methanol and hydrolyzed with cellulase in acetate buffer. The aglycones were extracted with ethyl acetate, derivatized with TBDMS and subjected to GC–
MS231. Of the 80 samples, only 37 contained detectable amounts of the isoflavones, of which nine contained more than 100 ȝg/kg wet wt. The limit of quantification was 1 ȝg/kg. An improved derivatization procedure used in-vial derivatization–extraction for the GC–MS analysis of flavonoids and phenolic acids in various herbs230,232. Derivatization takes place under basic conditions so that the hydroxyl groups of the analytes will be deprotonated. The anionic nucleophiles are transferred to the organic phase as ion-pairs using a phasetransfer catalyst (PTC) and are next subjected to reaction with methyl iodide. Polymer-bound tri-nbutylmethylphosphonium chloride proved to be the best PTC. In the SIM mode, the LODs of the flavonoids in the extracts were 4–40 ng/ml. The GC–MS chromatogram of a Mentha spicata extract is shown in Fig. 10. Several recent papers use GC–MS to determine flavonoids in food and food supplements – such as soy products and fruit – and in serum to study the bioactivity and bioavailability of flavonoids and phyto-estrogens (e.g.228,231,233,234). An interesting aspect of the first paper cited is that isotope dilution GC–MS was used228. At a first glance, there seems to be hardly any challenge from an analytical point of view since the analyte concentrations are high in soy; the daidzein (16) and genistein (17) concentrations are even in the low mg/g range. However, after consumption of soy food only low levels of these isoflavones are found in the human body and the challenge is to determine such low concentrations and accurately correct for losses during the various sample-treatment steps. To give an example228, for the determination of phytoestrogens in human serum, samples were hydrolyzed with ȕ-glucuronidase, the aglycones were extracted with ethyl acetate and the phyto-estrogen fraction isolated on a Sephadex LH20 column, with subsequent derivatization with BSTFA. Deuterated internal standards were used
Figure 10. SIM-mode GC–MS chromatogram of a Mentha spicata extract after derivatization with methyl iodide. Flavonoid peak assignment: 9, naringenin, m/z 300; 10, galangin, m/z 311; 11, kaempferol, m/z 327 and 12, luteolin, m/z 328; the other peaks are phenolic acids230.
for the isotope dilution procedure, with SIM-mode detection. The concentrations of daidzein and genistein varied between 2 and 900 ng/ml (means, 80 and 160 ng/ml, respectively) for 42 human serum samples. A lack of analytical performance data prevents further evaluation. In conventional GC it is very difficult to analyze flavonoid glycosides even after derivatization. Therefore, Pereira et al.235 used high-temperature–high-resolution (HT–HR) GC–MS, with columns that can withstand temperatures up to 400 ƕC, for the glucoside hesperidin (29). Unfortunately, the LOD of hesperidin (29) in a standard solution was found to be as high as 50 mg/l with both cold on-column injection and splitless injection at 370 ƕC. Other disadvantages were that derivatization of hesperidin with BSTFA took 72 h before analysis and that the derivative showed severe peak tailing. Interestingly, HT–HRGC–MS with cold oncolumn injection has been used without derivatization to determine mono-isoprenylated flavonoid aglycones, in an extract of Vellozia graminifolia236. After fractionation by means of preparative LC and TLC screening, ‘positive’ fractions were combined and analyzed by HT– HRGC–MS. Six mono-isoprenylated flavonols were identified on the basis of their melting points and their MS, IR, 13C and 1H NMR spectra. 2+ 2&+ 2
*OX5KD2
2+
2
29 It will be obvious that, for the analysis of flavonoids, GC will not easily replace LC—and certainly not if emphasis is on both aglycones and glycosides. For such studies, derivatization is needed (and several derivatives may be formed for one analyte), even in HT– GC. More efficient in-vial derivatization and the low LODs of SIM-mode MS detection are interesting advantages. However, they clearly do not outweigh the rapidity of direct LC–MS(/MS) procedures and the possibility to easily screen samples for target analytes as well as unknowns.
1.3.2.2.3. Capillary Electrophoresis Most studies that use CE for the analysis of flavonoids are in the field of natural product research, including the analysis of plants237-239, vegetables240, herbs241 and other plant or fruitderived products242,243. The CE modes primarily used are capillary zone electrophoresis (CZE)
and micellar electrokinetic chromatography (MEKC) with, typically, a phosphate or borate buffer, capillaries of 50–100 ȝmI.D., voltages of 10–30 kV and 10–50 nl injection volumes. Detection is usually performed with UV, but also fluorescence237, ED241,244 and MS detectors are used243,245. The practical usefulness of CE for flavonoid analysis can be illustrated by discussing a recent paper on the performance of MEKC and CZE in some detail248. With 13 flavonoids as model compounds, emphasis was given to the influence of separation conditions and molecular structure on the electrophoretic behaviour of the flavonoids. The separation mechanisms of these two CE modes are fundamentally different. CZE is only applicable to charged analytes and the charge-to-size ratios determine the electrophoretic migration times. In MEKC one should distinguish between neutral and charged analytes. With the former group, separation is based on hydrophobicity, which affects the analyte partitioning between the aqueous (moving with the electro-osmotic flow) and the micellar phases (charged and migrating with a different velocity). For ionic analytes separation in MEKC is based on both the degree of ionization and the hydrophobicity249. The flavonoids that were studied – flavonols, flavanones, flavanonols and a flavone – all have at least one phenolic hydroxy group; ionization of this group primarily determines the electrophoretic mobility. The magnitude of the net negative charge on a particular flavonoid is determined by the position and number of these groups and by the pH of the buffer solution. In CZE, two buffers were tested and the pH was varied over a wide range, but no conditions could be found that enabled separation of all 13 flavonoids. With MEKC better results were obtained. The composition of the four-component MEKC buffer and the pH range were varied. The migration times of all flavonoids except the very hydrophilic anthocyanins, increased with increasing pH; optimum resolutionwas at pH 7.3. Apparently, the separation selectivity of MEKC is better since more molecular-structure parameters play a role than in CZE. These include the degree of saturation and the stereochemistry of the C-ring, alkyl substitution and the number and position of phenolic hydroxy groups, methylation and glycosylation of the hydroxy groups and the complexation of flavonoids with borate buffer248. To the best of our knowledge, this is the first study in which MEKC and CZE are compared for flavonoid analysis and in which the better resolution of the former mode is demonstrated. Unfortunately, the authors did not study any real-life samples. As regards real-life applications, Baggett et al.238 used MEKC for the profiling of isoflavonoids in legume root extracts and compared it to LC. The optimum MEKC electrolyte was a mixture of 25 mM boric acid, 60 mM SDS and 1.6% 1,2-hexanediol, pH 9. Sample-tosample and run-to-run repeatabilities for MEKC were very good if the capillary was cleaned between each injection. Most MEKC results correlated rather well with those obtained for LC, while the runs were about two-fold faster238. A disadvantage was that optimization of the MEKC separation was more critical than that of LC.
In another study, CZE and LC were compared for the determination of the aglycones genistein (17) and daidzein (16) and their glucosides and glucoside-acetates in food products. LC was found to provide some 10-fold better UV detectability (LODs, 0.01–0.03 mg/l) and was less dependent on matrix effects239. In addition, in CZE resolution and repeatability were poor. The advantage of five-fold shorter run times in CZE is, of course, completely off-set by these drawbacks. CZE was also combined with SPE to determine flavone and flavonol-glucosides and -aglycones in Flos lonicerae249. Concentrations were 15–660 ȝg/g, with recoveries of 94– 104% and LODs, 0.4–0.6 mg/l. Capillary electrochromatography (CEC) has also been used for flavonoid analysis. CEC was compared with LC for the analysis of hop acids and prenylated hop flavonoids250, and of polymethoxylated flavones in essential oils251. In both studies the capillary CEC column was packed with C18-bonded silica, and acetonitrile–TRIS buffer (10 mM, pH 7.8) was used for separation with UV detection. The hop extract was subjected to off-line SPE and analyzed at 30 kV and 30 ƕC. Ten hop acids and flavonoids could be identified on the basis of their UV spectra and retention times; these included two pairs of hop acid isomers. LC on a C18-bonded phase, and with acetonitrile–formic acid gradient elution gave the same elution order of all target analytes, but one pair of isomers was not fully separated. In the study on polymethoxyflavones251, five of these compounds present in mandarin oil were well separated by both CEC and LC; with CEC the retention times were slightly shorter and, somewhat surprisingly, in this study the retention order was the opposite of that found in LC. The samples were also analyzed with CEC in the normal-phase mode using an acetonitrile– isopropanol–hexane eluent. The retention orders in NP-CEC and RP–CEC were very similar – and so were the run times – but the resolution was better in the latter mode. Unfortunately the authors did not provide analytical performance data, which precludes a more detailed comparison of the CEC and LC methods. Only a few papers discuss the use of CE–MS for the determination of flavonoids243 and phenolic compounds252. This may indicate that the technique is not considered sufficiently robust and user-friendly by many researchers. In the CE–ESI(−)–MS study by Lafont et al.252, a standard mixture of eight phenolic compounds was analyzed. Admittedly, phenolic acids are no flavonoids, but since they are very similar compounds, they will be included in the present discussion. With SIM–MS the authors were able to identify all eight compounds based on their retention times and characteristic fragment ions ([M−H]−, loss of CO, CO2 and CH3) and obtained LODs of 0.1–40 ȝg/l. The authors state that it is easier than in LC to couple CE to an ESI interface since, because of the very low (nl/min) flow rate, no flow splitting is needed. However, for successful coupling of CE to MS, the ESI interface design plays an important role; the needle should be grounded and a voltage applied to the counter electrode. In a more recent paper243, CE–ESI(−)–MS was used for the determination of flavonoids in a
phytomedicine and compared with CE–UV. MS detection sensitivity for hesperetin and naringenin (LODs, 0.5 mg/l in SIM) was similar to that of UV254, while for biochanin A (28) the LOD was less satisfactory, i.e. 2 mg/l. On the other hand, CE–UV was not sufficiently selective to enable quantitation, and CE–MS in the SIM mode had to be used to determine the concentration of naringenin (22) in a liquid herbal drug at 6 mg/l (cf. Fig. 11). Surprisingly, in the CE–MS study quoted earlier252, the LODs were one to two orders of magnitude lower. We do not have an explanation, since the authors did not compare their results with the former study. At the present time, the future of CE for flavonoid analysis is, to our opinion, not too bright. Compared with LC, there is no dramatic difference of run times, and the limited consumption of sample and solvents (not particularly expensive in LC anyway) does not appear to have much impact. Moreover, the repeatability of retention /migration times still is better in LC than in CE. Probably, the most promising aspects to explore are the complementarity of CEC and MEKC separations to LC, and their robust interfacing with MS.
Figure 11. Total ion current (a) and SIM-mode (b) CE–ESI(−)–MS of naringenin in a phytomedicine. Capillary, fused silica, 60 cm, 50ȝm I.D.; buffer, 40mM NH4OAc (pH 9.5), 15% ACN (v/v); sheath liquid, 2mM NH4OAc in 2-propanol/water/triethylamine (80/20/0.1, v/v); sheath flow, 4 ȝl/min; voltage, 25 kV; current, 31 ȝA; temperature of the heated capillary, 150 ƕC243
Table 4. Representative Studies on TLC of Flavonoid
1.3.2.2.4. Thin-layer Chromatography Since the early 1960s, TLC has been used in flavonoid analysis. TLC is especially useful for the rapid screening of plant or medicinal extracts for pharmacologically active substances prior to detailed analysis by instrumental techniques such as LC–UV especially because many samples can be analyzed simultaneously. In most cases silica is used as stationary phase, and plates are developed with either a combination of 2-(diphenylboryoxo)ethylamine and polyethylene glycol or with AlCl3. Detection is mainly performed using UV light at 350–365 or 250–260 nm or with densitometry at the same wavelengths. At present, TLC still plays a distinct role in flavonoid analysis: a representative selection of the about 80 papers published in the last 5 years is summarized in Table 4156. Some of these are discussed below to illustrate the state-of-the-art. One interesting example is the separation of flavonoid glycosides and rosmarinic acid from Mentha piperita (peppermint) on HPTLC plates253. A variety of (modified) silica sorbents were
tested as well as many organic eluents, which ranged from n-hexane to esters, ethers and methanol.
For
six
standard
compounds,
the
best
separation
was
obtained
on
aminopropylbonded silica with acetone–acetic acid (85:15, v/v) as eluent; with C18-bonded silica and water–methanol (60:40, v/v) good results were also achieved. The six standard compounds were found in the peppermint extract, isolated by preparative LC and their structures determined by several spectroscopic identification techniques. Eriocitrin (30) was found to be the main flavonoid constituent; no quantification data were provided.
2+ 2+ 2
*OX5KD2
2+
2 30
Soczewinski et al.245 used double-development TLC to separate a flavonoid mixture containing nine glucosides and seven aglycones. In the first step, the more polar glycosides were separated using an eluent with high solvent strength. After solvent evaporation, the aglycones were separated in a second step in the same direction with another, relatively weak, eluent. In several recent papers255-258, numerical taxonomy is used to calculate the orthogonality of the retention factors of mixtures of flavonoids. On the basis of results calculated for 19 standard compounds and using two parameters, the optimum eluent composition to study flavonoids in red wine258 and in propolis257 was determined for 11 tertiary eluents. The success of the approach was demonstrated by the fact that, for the optimal eluent combination, up to 10 of the 19 standard compounds were identified in 14 propolis samples of different origin. Application to red wine was also successful: several phenolic compounds, including three flavonoids, could be identified258. Quantification generally is not a main goal of TLC studies. However, densitometry is used in several studies259,260. In one paper259, kaempferol (23) and quercetin (18) were determined in an extract of Ginko biloba leaves by scanning the HPTLC silica plates in the reflectance mode at 254 nm. Recoveries using standard addition were above 94%. The concentrations of kaempferol and quercetin in the extract were 7 and 14 mg/l, respectively. Janeczko et al.260 used a similar method to determine genistin (26) and daidzin (25) at 260 nm in various soy cultivars. The analytical performance data were fully satisfactory, possibly because the analyte concentrations were fairly high: genistin, 0.06–0.15%; and daidzin, 0.03–0.01%. In propolis, several flavonols, flavanones and phenolic acids were quantified using two-dimensional (2D)-TLC with densitometry at 254 and 366 nm261. Also here, good analytical performance data were obtained; concentrations were 90–1440 mg/l.
Wojciak-Kosior et al.262 used TLC combined with densitometry to study the hydrolysis of six flavonoid glycosides. The flavonoids were heated under reflux with HCl and analyzed every 15 min. The pseudo first-order hydrolysis rate constants varied between 1.7×10−2 and 1.1×10−2 min−1 for 3- and 7-glycosides; for 7-glycosides the hydrolysis was practically complete after 90–105 min. The hydrolysis mechanism of rutin (27), a diglycoside, was found to be more complicated. There are presumably two steps—a mechanism to be studied in the near future. Two-dimensional-TLC on cyanopropyl-bonded silica was used to separate eight flavonoids and three phenolic acids in Flos sambuci L.263. The first dimension was a normalphase separation for which seven binary eluents were tested, and the second one a reversedphase separation, studied by using three binary eluents. From amongst the 21 combinations, the three best ones all contained n-hexane in the first, and water in the second dimension. Fig.12 shows the results of a separation using the eluents 40% propan-2-ol in n-hexane in the first dimension and 50% aqueous 1,4-dioxane in the second dimension, although errors in migration direction in their graphs and compound numbering complicate the interpretation of their data. More than 12 spots can be discerned and nine flavonoids and three phenolic acids were (tentatively) identified in the Flos sambuci extract. In summary, several modes of TLC are still in vogue for flavonoid analysis. The emphasis is on screening for the main flavonoids in real-life samples. In most cases a close-to-standard protocol can be followed, but a newer method such as numerical taxonomy also deserves attention.
Figure 12. Two-dimentional (2D)-TLC of Flos sambuci extract on cyanopropyl-bonded silica. Spot assignment: 1, myricetin; 2, naringenin; 3, luteolin; 4, apigenin; 5, acacetin; 6, hyperoside; 7, quercetin; 8, naringin; 9, rutin; 10, hesperetin; 11, quercitrin; 12, astragalin. Eluents: first dimension, 40% propan-2-ol in n-hexane; second dimension, 50% aqueous 1,4-dioxane. UV detection at 366 nm after development with 2(diphenyl boryoxo)ethylamine and polyethylene glycol263.
1.3.3. IDENTIFICATION AND STRUCTURAL CHARACTERIZATION 1.3.3.1. General Today, LC–MS/MS is the most important technique for the identification of target flavonoids and the structural characterization of unknown members of this class of compounds. As regards target analysis, tandem-MS detection has largely replaced single-stage MS operation because of the much better selectivity and the wider-ranging information that can be obtained. Depending on the nature of the application, additional information is derived from LC retention behaviour, and UV absorbance – and, occasionally, FLU or ED – characteristics, due comparison being made with standard injections and/or tabulated reference data. In studies on the characterization of unknowns, a wide variety of LC–MS/MS techniques is usually applied next to LC–DAD UV for rapid class identification. In addition, LC–NMR often turns out to be an indispensable tool to arrive at an unambiguous structural characterization. In Section 1.3.3.2., attention will be devoted to the main fragmentation pathways for four major classes of flavonoids, i.e. flavones, isoflavones, flavonols and flavanones. In this context, the retro-Diels–Alder reaction, which is an important fragmentation reaction of flavonoids, will be discussed. RDA fragments are especially important for the structural characterization of aglycones and the aglycone part of flavonoid conjugates. Next, in Section 1.3.3.3., the on-line coupling of LC and NMR will be discussed, and its increasing importance for the characterization of flavonoid conjugates illustrated. This part of the text will also serve to demonstrate the complementary roles of, specifically, NMR- and MS/MS-based information. Off-line NMR, which has been used extensively for flavonoid analysis, is outside the scope of this review and will not be considered.
1.3.3.2. LC–MS/MS 1.3.3.2.1. General In order to facilitate discussions on the mass fragmentations of flavonoid aglycones, Ma et al.
264
proposed a nomenclature to unambiguously describe the resulting fragment ions (Fig. 13). In the PI mode, the ions that are formed after the cleavage of two bonds in the C-ring, are denoted i,jA+ and i,jB+, with ion A containing the A-ring and ion B, the B-ring. The indices i and j represent the C-ring bonds which are broken. When the NI mode is used, the ions are denoted i,j
A− and i,jB−, respectively. Ions derived from the fragment ions by the loss of a fragment X, are denoted [i,jA± −X] and [i,jB± −X], respectively. As mentioned above, an important fragmentation reaction of flavonoids is the RDA reaction, which may occur in six-membered cyclic structures containing a double bond and involves the relocation of three pairs of electrons in the cyclic ring. The net result of these rearrangements is the cleavage of two ı-bonds and the formation of two ʌ-bonds, for example,
cyclohexene will fragment into butadiene and ethylene. Two (complementary) fragments, and
1,3
1,3
A+
+
B , are formed and charge retention can occur on either side of the cleavages, as depicted
in Fig. 13A for the flavones apigenin (31) and luteolin (24)264. The C-ring cleavage product ions can be used to determine the number and nature of the substituents on the A and B-rings. For example, in the MS/MS spectra of apigenin and luteolin (the latter is shown in Fig. 13B), which 2+ 2
+2
2+
2
31 have [M +H]+ m/z 271 and 287, respectively, a 1,3A+ fragment ion shows up at m/z 153. The corresponding 1,3B+ ions are found at m/z 119 and 135, respectively. This indicates that the two compounds differ in the substitution of the B-ring, with luteolin having two OH groups, and apigenin only one264.
Figure 13. (A) RDA reaction mechanisms for the formation of 1,3A+ and 1,3B+ fragment ions for apigenin (R=H) and luteolin (R=OH). Each arrow represents relocation of one pair of electrons. (B) Low-energy MS/MS spectrum of luteolin219 .
When reading the more detailed discussions on MS fragmentation behaviour presented below, one should consider that, in the quoted studies, different instruments and operating conditions were often used. Fortunately, experience shows that the fragmentation pathways are largely independent of the ionization mode (ESI or APCI) and the type of instrument (triple quadrupole or ion trap) used209,186. On the other hand, significant differences do occur as regards the relative abundances of the various fragment ions. These are, therefore, not included in the discussions presented in Sections 1.3.3.2.2 –1.3.3.2.3 below.
1.3.3.2.2. Fragmentation in PI Mode Relevant information on the main fragment ions formed after cleavage of the C-ring of the selected flavonoid classes in the PI mode (cf. Fig. 14), is summarized in Table 5.
Figure 14. Fragmentation pathways for flavonoids caused by cleavage of C-ring bonds; (A) in both PI and NI: (A1) 1 and 3, (A2) 0 and 4; (B) in PI: (B1) 0 and 2, (B2) 1 and 4 and (C) in NI: (C1) 0 and 3, (C2) 1 and 2, (C3) 1 and 4, (C4) 2 and 4.
Table 5. Fragment ions observed for the selected flavonoid classes in the PI mode
RDA cleavage, which generates
1,3
A+ and
1,3
B+ fragment ions (Fig. 14, A1), is the most
important fragmentation pathway for flavanones, flavones and flavonols, but also occurs with isoflavones. The first three classes all show
1,3
A+ as the most prominent product ion, with the
flavones luteolin (24) and apigenin (31), the flavonol kaempferol (23)264, and the flavanones naringenin (22) and hesperetin (32)224 as typical examples. The 265
was even proposed as a diagnostic ion for flavones
1,3
B+ ion is also observed, and
; however, it is also formed with flavones
and a flavanone such as naringenin. Compounds having a methoxy substituent show relatively weaker RDA fragmentation264: in the spectra of acacetin (33) and chrysoeriol (34), the 1,3A+ ion typically has a less than 10% relative abundance264. The
1,3
A+ fragment ion was also reported
for isoxanthohumol – a chalcone with 6-prenyl substitution – and 6- and 2&+
2+
+2
2
2
2+
2&+
2&+ 2
+2
2+
32
2
33
2
+2
2+
2
34
8-prenylnaringenin, although with less than 20% relative abundance266. If a low collision energy of 30V was used, the mass spectra of prenylated flavones and flavonols showed [1,3A–C4H8]+ as the only product ion. In a study on the leaf surface flavonoids of Chrysothamnus, this characteristic loss of the isoprenyl substituent has been used for the target analysis of prenylflavonoids in an SRM procedure267.
For flavones and flavonols cleavage of the 0,2 bonds (Fig. 14, B1) is a common C-ring cleavage pathway. Various authors have reported the formation of the
0,2
B+ ion with relative
abundances ranging from 1 to 90%, e.g. for kaempferol (23), quercetin (18), myricetin (35), isorhamnetin (36), apigenin (31), luteolin (24), acacetin (33) and chrysoeriol (34)264,267. The corresponding
0,2
A+ ion may be used to distinguish flavonols since it does not occur in the
spectra of other classes of flavonoids264. Interestingly, ions due to the 0,2 cleavage reaction are not reported in the NI mode. 2&+
2+
2+
2+ 2
+2
2
+2 2+ 2+
2+
2
2+
2+
35
2
36
The 0,4 C-ring cleavage (Fig. 14, A2) is not often discussed in the literature. Only protonated flavones fragment via this pathway, viz. under low-energy FAB–CID conditions264. Consequently, the presence of 0,4B+ ions can be considered diagnostic for flavone aglycones. To the best of our knowledge, the cleavage of bonds 1 and 4 of the C-ring (Fig. 14, B2) has been reported only once, viz. to explain the m/z 147 ion (1,4A+) in the spectrum of naringenin (22)265. However, one should add that the m/z 147 fragment may also correspond to a 0,4B+ ion after loss of water. Other, generally less characteristic, fragments common to most flavonoids are those arising from the loss of H2O (18 Da), CO (28 Da), C2H2O (42 Da) and the successive loss of H2O and CO (46 Da). However, to the best of our knowledge (see Table 5), the fragments [M +H−42]+ (–C2H2O) and [M +H−46]+ (successive loss of H2O and CO), do not show up in the spectra of isoflavones. A loss of 68 Da (successive loss of C2H2O and C2H2), which involves a more complex fragmentation process, has also been observed, and has been suggested to be indicative for flavones266. However, there are exceptions to this rule264. Similarly, the presence of [M+H−56]+ (loss of 2 × CO) has been suggested as an ‘indicator’ for isoflavones209,218,266. However, the same fragment has also been observed for several flavonols264,265,268. As can be seen from Table 5, the product ion [M+H−15]+•, formed by the loss of a methyl radical,
is
prominent
in
many O-methylated
isoflavones,
flavones and flavonols
(e.g.209,218,267,269). In addition to the loss of a methyl radical, further loss of water or H2O-plusCO may occur, resulting in [M +H−33]+• and [M +H−61]+• fragments. In the mass spectra of 3
methoxyflavonoids, [M +H−15]+• was found to be accompanied by an [M +H−16]+ ion of equal or somewhat lower abundance. For this ion, the loss of CH4 was proposed by the formation of a furan ring involving C2& and the oxygen at C3 267.
1.3.3.2.3. Fragmentation in NI Mode Relevant information on fragment ions of the selected flavonoids observed in the NI mode, which has been studied more frequently than PI, is summarized in Table 6. The RDA C-ring cleavage of the 1,3 bonds, which creates 1,3A− and 1,3B− product ions (Fig. 14, A1), is the most important fragmentation pathway in the NI mode, as is also true for the PI mode. As Table 1.6 shows,
1,3
A− and
1,3
B− fragments are reported for many flavonoids. In the
mass spectra of the flavonols kaempferid, eriodyctiol (37), morin (19), quercetin (18) and rhamnetin270, and the prenylated flavonoids, 8- and 6- prenylnaringenin271 1,3
abundant fragment ions with abundances of the
1,3
A− and
1,3
1,3
A− were the most
B− in second place. In another study179, however, the relative
B− ions of luteolin (36) and genkwanin (38) were found to be
quite low (1–10%), and in a third one, kaempferol (23) did not show any RDA ions at all270. These mutual differences possibly reflect differences in at all270. These mutual differences possibly reflect differences in the experimental set-up and/or operating conditions. As for kaempferol, it has been reported that this compound shows little fragmentation up to a collision energy of about 25 eV271. However, in the study quoted above no fragmentationwas observed even at a collision energy of 30 eV. All of this shows that one has to be careful when attributing an ‘indicator role’ to specific fragments. 2+ 2+
2+ 2
+2
2+
0H2
2
2
2+
37
2
38
Another C-ring cleavage generates 0,3A− and/or 0,3B− fragments (Fig. 14, C1). In a study of 14 isoflavones, flavones and flavanones, 0,3B− fragments were observed only for the isoflavones daidzein () and genistein (); however, the same fragment was also observed for a flavone265. Based on the presence of m/z 135 and 148 ions, which, according to them, represented 0,3A− and [0,3B–CH3•]− fragments, respectively, Prasain et al.266 tentatively
Table 6. Fragment ions observed for the selected flavonoid classes in the NI mode
identified an unknown compound in a kudzu dietary supplement extract as 3&-methoxydaidzein. The authors referred to an earlier study of methoxylated flavones272 in which similar fragments were observed, but did not provide additional (NMR) data to prove that the methoxy group is in the 3& position. 0,4
A− and 0,4B− fragments (Fig. 14, A2) are observed at relatively low abundance for at least
some members of all classes of flavonoids discussed here (Table 6). A 265
270
observed for kaempferol (23) , for several flavonoids
0,4
A− fragment was
and for the flavone apigenin (31), the
flavonols quercetin (18) and kaempferid, and the flavonones eriodictyol, naringenin (22) and isosakurametin (39)179. In a study of several isoflavones and flavones, fragmentation of isoflavones mainly resulted in
0,4
B− ions, whereas for the flavones
1,3
A− was more prominent
(cf. Table 6). 0,4 C-ring cleavage was also proposed for a fragment ion of the flavanone isosakurametin179 : loss of CH3• from the deprotonated molecule was followed by a 0,4 C-ring cleavage to create [0,4B CH3•]−. The 1,2 C-ring cleavage (Fig. 14, C2) was suggested for the main ions (1,2A−) of the flavonols quercetin (18) (m/z 179) and fisetin (40) (m/z 163)179. The complementary ions (1,2B−, m/z 121 for both compounds) were also observed, but with much lower abundances. For quercetin, the same fragment ions were also reported in two other studies265,270, although they were not designated as 1,2A− and 1,2B− there. Because other flavonols that were studied did not
show 1,2 C-ring cleavage, the authors proposed that this pathway is specific for 3&,4&dihydroxyflavonols179. It is interesting to add that, in another paper270, m/z 121 in the spectrum of the 3&,4&- dihydroxyflavonol rhamnetin was attributed to
0,4
A− rather than
1,2
B−.
1,2
A−, but no
1,2
B−, ions have also been observed in the spectra of two isoflavones, formononetin (20) and biochanin A (2)209,245. 2+
2&+ 2
+2
2
+2
2+ 2+
2+
2
2+
39
2
40
The cleavage of bonds 1 and 4 (Fig. 14, C3) has been proposed to explain the formation of m/z 149, [1,4B + 2H]−, in the mass spectrum of apigenin (31). The assignment was made because few other structures were considered acceptable; MS3 experiments did not provide more information179. However, Hughes et al. proposed that the m/z 149 ion is either 0,4B− or 0,3B−. The authors favoured the latter cleavage, because in that case a subsequent loss of 2 × CO may occur, which had earlier been observed for the [M−H]− ion of galangin (3,5,7trihydroxyflavone)265. An m/z 149 ion has also been found in the spectrum of luteolin (24)270 . However, it is probably not the equivalent of the m/z 149 ion of apigenin (31): since luteolin (24) has an additional hydroxy substituent on the B-ring, the ion would have a higher mass. Next to the various C-ring cleavages, other fragmentations are also observed in the NI mode. To give an example, the loss of m/z 15 from the deprotonated molecule indicates the loss of a methyl radical, as was also observed in the PI mode. This product ion was reported for the isoflavones formononetin (20) and biochanin A (28)218,209, and also for two of their 7-Oglycosidic conjugates, ononin (21) and sissotrin (41)209. Hesperetin (32) and its 7-O-glycoside, hesperidin (29), also yielded the [M−H−15]− ion. The same fragment is prominent in the spectra of methoxylated flavonoids such as acacetin (33), isorhamnetin (36), rhamnetin and hesperetin (Table 6)270. It is interesting to note the absence of B-ring fragments in the MS/MS spectra of acacetin (methoxylated in the B-ring), hesperetin and isorhamnetin270. 2
Glu-O
2+
2
2&+
41
The loss of small fragments such as CO and H2O occurs in the NI as well as in the PI mode. For example, after cleavage of bonds 2 and 4, the deprotonated molecule loses two CO moieties but does not form 2,4A− and 2,4B− fragment ions. This fragmentation pathway was suggested to explain the m/z 213 ion of galangin265. Two further comments are: (i) as in the PI mode, [M−H−56]− only occurs for isoflavones and flavonols; (ii) the [M−H−72]− ion (successive loss of CO2 and CO) which, to the best of our knowledge, has not been reported in PI, was observed for all selected classes except the flavanones. 1.3.3.2.4. Flavonoid–(di)glycosides Flavonoids commonly occur as flavonoid-O-glycosides; the 3- and 7-hydroxyl groups are the typical glycosylation sites. Glucose is the most frequently found sugar moiety, with galactose, rhamnose, xylose and arabinose in second place. Flavonoid-diglycosides are also found in nature rather frequently, with rutinose (rhamnosyl-(1ĺ6)-glucose) and neohesperidose (rhamnosyl-(1ĺ2)-glucose) being the most common sugar moieties. Flavonoid-C-glycosides – in which the sugar is directly linked to the aglycone by a C—C bond – comprise flavonoidmono- and di-C-glycosides and O,C-diglycosides (with this group, the O-glycoside moiety is linked either to a hydroxyl group of the aglycone or to a hydroxyl group of the C-bound glycosyl residue). To date, C-glycosylation has only been found at the C-6 and C-8 positions of the flavonoid aglycone273. 2+ Glu-Rha-O
2
2
2+
42 The fragment ions for glycocon jugates are denoted according to Domon and Costello274. Fig. 15 shows examples for naringenin–7-O-neohesperidose (42) and naringenin–7-Oglucoside. Y represents the diglycoside, with fragments that contain the aglycone part being
Figure 15. Carbohydrate ion nomenclature for: (A) naringenin–7-O-neohesperidose (naringenin–7-Orhamnosyl-(1ĺ4)-glucose) and (B) naringenin–7-O-glucoside274.
denoted Y1 (loss of one glycose moiety) and Y0 (loss of two glycose moieties); the corresponding glycose fragments are denoted B1 and B0, respectively. Ions formed due to cleavage in the glycose ring, and which contain the aglycone part, are labelled k,lXj, where j is the number of the interglycosidic bonds broken, counting from the aglycone; the superscripts k and l indicate the interglycosidic bonds, with the glycosidic bond linking the glycose part to the aglycone being numbered 0. MS/MS of flavonoid-(di)glycosides is a useful tool to differentiate: (i) the 1ĺ2 and 1ĺ6 glycose linking types of diglycosides, and also to distinguish; (ii) Oglycosidic (3- O- and 7-O-) and (iii) C-glycosidic (6-C- and 8-C-) flavonoids. Recently, an extensive tutorial on the use of mass spectrometry in the structural analysis of flavonoids was published275. In this overview, much attention was devoted to the characterization of the various groups of flavonoid-(di)glycosides. The present text is therefore limited to a discussion of several recent studies. In a recent paper, the interglycosidic linking types and the types of O-glycosidic linkage of eight flavonol, flavone and flavanone diglycosides were studied by means of LC–(+)ESI– MS/MS276. As an example, in Fig. 16 the MS/MS spectra of naringenin–7-O-rutinoside (Orhamnosyl-(1ĺ6)-glucose) and naringenin–7-O-neohesperidose (O-rhamnosyl-(1ĺ2)-glucose, (42)) are shown; the Y1 ion is formed after loss of a rhamnose unit (146 Da), and the Y0 ion after further loss of a glucose unit (162 Da). The interglycosidic linking can be determined on the basis of the value of the ratio Y0¯ /Y1¯ : when [M +H−146]+ (Y0¯ ) > [M+H−(146+162)]+ (Y1¯ ), this indicates a 1ĺ6 linkage, while [M +H−146]+ < [M+H−(146 + 162)]+ indicates a 1ĺ2 linkage. The type of O-glycosidic linkage can be determined by the presence of the Y* ion, which corresponds to the loss of an internal glucose residue from the product ion ([M+H−162]+). This ion is only observed for flavonoid-7-O-rutinosides and 7-Oneohesperidosides, and not for the corresponding 3-O-linked types. In another study on flavonoidrutinosides and neohesperidosides similar conclusions were reached concerning linking types277.
Figure 16. MS/MS spectra obtained for [M +H]+ ions of (a) naringenin–7-O neohesperidoside and (b) naringenin–7-O-rutinoside, using LC–ESI(+)– MS/MS276.
Recently, the above results were used to determine the glycose linking type of isoflavone-diglycosides in a L. corniculatus L. extract using LC–APCI(−)–MS/MS224. Two flavonoid-O-diglycosides with the same mass were found. In the mass spectra of both compounds the sequential loss of a rhamnose (Y0¯ ) and glucose (Y1¯ ) was observed. On the basis of the value of the ratio Y0¯ /Y1¯ , and the absence of Y*, the two flavonoids were both identified as 3-O- rutinosides of kaempferol (23). In one of the isomers probably one or more – OH groups are shifted. Waridel et al.278 studied the MS/MS fragmentation of 6-C- and 8-C-flavonoid-glycosides. In full-scan MS, fragment ions were observed that were formed by 0,2 cleavage of the glycosidic ring (see Fig. 15B) containing the aglycone part of the flavonoid. Upon further fragmentation with MS/MS,
1,3
B+ and
0,2
B+ ions (cf. Section 1.3.3.2.2) were observed for both
the 6-C- and 8-C-isomers. However, 1,3A+ ions were found only for the latter isomers and could, therefore, be used as an indicator. In a kudzu extract several isoflavone-O- and C-glycosides were found using LC–ESI(+)–MS/MS266. [M+H−120]+ was found to be the diagnostic ion for the C-glycosides. This diagnostic ion was also used to identify two xanthone-C-glycosides in a mango peel extract using LC–ESI(+)–MS/MS279. In an LC–ESI(±)–MS/MS study of a Crataegus extract273 10 flavonoid-(di)glycosides, including two acetates, were identified based on earlier information regarding the main flavonoid constituents and MS/MS results for the determination of the sugar linking type. The sugar moieties were O-bound glucoside, glucopyranoside, galactopyranoside and rutinoside. Loss of one or more glycose moieties was observed, e.g. the sequential loss of rhamnose and glucose for the rutinoside rutin (27), and cleavage of the interglycosidic linkage generating 0,2X1 and
0,2
X0 fragments for vitexin–2&&-O-rhamnoside, vitexin–2&&-O-rhamnoside–acetate and
isovitexin–2&&-O-rhamnoside-acetate.
1.3.3.3. LC–NMR In recent years, on-line (though often stopped-flow) LC–NMR has attracted increasing attention in the field of natural product research. The main advantages (e.g. high information content, differentiation of isomers and substitution patterns) and disadvantages (low sensitivity, expensive instrumentation, long run times) have been discussed and highlighted in several recent reviews280,281 and there is no need to further consider them here. What should, however, be emphasized is that, while NMR detection is particularly powerful for the differentiation of isomers, sugar configurations and substitution patterns on aromatic ring systems, (tandem) MS techniques are needed to obtain information on, e.g., molecular mass and functional groups.
Table 7. LC–NMR studies of flavonoids
Moreover, for a comprehensive structural elucidation of a novel natural product, preparative isolation is often still necessary because in LC–NMR usually part of the 1H spectral region is lost and, moreover, LC–NMR in most cases does not provide the indispensable
13
C NMR
280
data . The reader is referred to the same review for a discussion on the merits and demerits of hypernated techniques (LC–NMR–MS and LC–NMR–MS/MS, with or without a UV detector) compared with two separate hyphenated, LC–NMR and LC–MS, systems. Table 7 illustrates the recent interest in LC–NMR for flavonoid analysis. In most of the cited publications, the stoppedflow mode was used to enable very long scan times to record the NMR spectra; scan times varied between 1 h and several days per chromatographic peak. An alternative is to use very low flow rates as was, e.g., done in a study on the flavonoid constituents of the roots of Erythrina vogelii where a flow rate of 0.1 ml/min was used282. In this study accurate mass data were acquired by means of LC–Q-TOF MS and several prenylated isoflavones and isoflavanones were identified in the root extract. In a study of a Gentiana ottonis extract, LC–NMR was combined with LC–DAD UV and LC–MS/MS, and DAD UV, MS/MS and NMR spectra of the main chromatographic peaks were obtained283. As an example, the identification of an unknown constituent in the extract is shown in Fig. 17. The UV spectrum showed the characteristics of a flavone (cf. Table 3). Based on characteristic fragments observed in the MS/MS spectrum, the compound was determined to be a 6-C glycoside with a monohydroxylated B-ring. The NMR spectrum provided the additional information for the unequivocal identification of the flavone as swertisin. The authors claim that the lowest detection level for LC–NMR was about 0.05 ȝmol per peak in the on-flow mode and that in the stopped-flow mode about 100-fold less material was required (but the acquisition times then were extremely long). To our opinion, these figures seem to be rather optimistic compared to the results reported in other studies.
The need to use several complementary techniques was also apparent in the analysis of a Hypericum perforatum extract. To identify its constutuents a combination of stopped-flow LC– NMR, LC–DAD UV and LC–ESI(−)–MS/MS was used215. The two partly co-eluting peaks of interest, hyperoside and isoquercitrin–which are the 3-O-galactoside and 3-O-glucoside of quercetin (18), respectively – could not be identified with LC–MS/MS only, because they have
Figure17. LC–UV254 chromatogram of Gentiana ottonis with the 1H NMR, DAD UV, MS and MS/MS spectra of swertisin (peak 33). TSP, thermospray interface283.
The need to use several complementary techniques was also apparent in the analysis of a Hypericum perforatum extract. To identify its constutuents a combination of stopped-flow LC– NMR, LC–DAD UV and LC–ESI(−)–MS/MS was used215. The two partly co-eluting peaks of interest, hyperoside and isoquercitrin–which are the 3-O-galactoside and 3-O-glucoside of quercetin (18), respectively – could not be identified with LC–MS/MS only, because they have the same molecular mass and show identical fragmentation behaviour. LC–NMR allowed unambiguous identification because of the differences in the spectra of the sugar moieties. Stopped-flow LC–NMR required scan times of several hours to record useful spectra for injected analyte masses of 10–50 ȝg. It is interesting to add that LC–MS/MS experiments in H2O and D2O were performed to determine the number of exchangeable protons in the molecules, which are not visible in NMR. This enabled the determination of the number of hydroxy groups of each constituent. Flavonoid-glycosides in apple peel were successfully identified by using the same combination of techniques as in the previous example157. Five quercetin (18) glycosides and one quercetin diglycoside were recognized with LC–MS/MS, but no complete structure elucidation was provided. The glycosidic nature of the flavonoids in the extract was determined with LC– NMR, because the different sugar moieties each have their typical resonances. The sugar linkage position was derived from a comparison with reference compounds. For the least abundant compound in the extract, rutin (27) (concentration 40 ȝg/ml), the time needed to record spectra with a reasonable signal-to-noise ratio for a 100 ȝl injection was approximately 1.5 h. LC–DAD/UV–SPE–NMR was used in combination with on-line radical scavenging detection for the identification of radical scavenging compounds in extracts of Rhaponticum carthamoides285. A combination of on-line recorded 1H NMR spectra, MS/MS fragmentation and exact mass data were used to determine basic structures and elemental composition, while HMBC experiments were performed off-line to determine the sugar-linking type, after trapping the compounds of interest up to three times on separate SPE cartridges and combining the eluates. Without any prior off-line chromatographic steps, five flavonoid-ȕ-glucopyranosides were identified, of which two had a 6'-O-acetyl group. In a recent study of de Rijke et al. on the flavonoid constituents of a red clover extract, stopped-flow LC–NMR and stand-alone NMR were used to identify structural isomers that could not be distinguished on the basis of MS/MS information173. By combining the information provided by MS/MS, 1H NMR, correlation spectroscopy (COSY) and nuclear overhauser enhancement spectroscopy (NOESY) spectra recorded for two sets of isoflavone– glucose–malonate isomers, not only the positions of the glucose moieties on the flavonoidaglycones, but also those of the malonate moieties on the glucose groups were determined. One set of isomers only differed in the substitution position of the malonate group on the glucoside
ring, but rather unexpectedly – because the two pairs of isomers were thought to be mutually closely similar – for the other set of isomers the position of the glucose group was also different. Their structures are shown in Fig. 18. High mg/l concentrations of the analytes had to be used to record satisfactory NMR spectra on a 400 MHz instrument (1052 scans per peak). In all of the studies quoted in Table 7 conventional-size LC was used. Currently, much effort is devoted to the development of micro- or even nano-LC–NMR. This is an attractive development since expensive deuterated solvents, which are required to suppress the eluent background signals in 1H NMR, can now be used more easily. Much attention is also paid to probe design, to reduce the problem of poor sensitivity. The recently developed cryoflow NMR probe
286
that cools the receiver coils to cryogenic temperatures to improve the signalto-noise
ratio of the NMR spectra has been applied for the analysis of an oregano extract164. Five flavonoids were identified using an LC–UV–SPE–NMR–MS set-up. According to Spraul et al.286 with cryoflow probes the analyte detectability is about four-fold better than with conventional probes or, alternatively, the scan time is 16-fold shorter for the same amount of sample. This approach, and also the use on-line preconcentration using LC–SPE–NMR, are likely to gain popularity in the future.
Figure 18. Structures of: (A) formononetin-7-O-ȕ-D-glucoside-6'-O-malonate (FGM, R=H) and biochanin A-7-O-ȕ-D-glucoside-6'-O-malonate (BGM, R=OH); (B) formononetin-7-O-ȕ-D-glucoside-4'-O-malonate (FGMi) and (C) 5-hydroxy-7- methoxyisoflavone-4&-O-ȕ-D-glucoside-4'-O-malonate (BGMi).
1.4. PHARMACOGNOSY OF PISTACIA LENTISCUS 1.4.1. BOTANY AND TAXONOMY The following lists the botanical name, synonyms, and the classification of Pistacia lentiscus. Botanical Nomenclature: Pistacia lentiscus L. Synonym: Pistacia lentiscus var. lentiscus. Plant classification: Kingdom: Plantae-Plants Subkingdom: Tracheobionta – Vascular plants Superdivision: Spermatophyta – Seed plants Division: Magnoliophyta – Flowering plants Class: Magnoliopsida– Dicotyledons Subclass: Rosidae Order: Sapindales Family: Anacardiaceae Genus: Pistacia Species: P. lentiscus
Figure 19. Pistacia lentiscus L.
1.4.1.1. The family Anacardiaceae
Anacardiaceae Lindl., the cashew family, includes more than 700 species in 82 genera that are primarily distributed pantropically. Some genera, however, extend into the temperate zone. Members of the family are cultivated throughout the world for their edible fruits and seeds, medicinal compounds, valuable timber, and landscape appeal. Some of the products of Anacardiaceae, including mangos (Mangifera indica L. and other species), pistachios (Pistacia vera L.), cashews (Anacardium occidentale L.), and pink peppercorns (Schinus terebinthifolia L.), are enjoyed worldwide while other notables such as the pantropical Spondias L. fruits, the marula of Africa (Sclerocarya birrea (A. Rich.) Hochst.), and the Neotropical fruits of Tapirira Aubl., are restricted to localized cultivation and consumption and are not generally transported The far distances to larger markets. Anacardiaceae includes primarily trees, shrubs, and lianas with resin canals and clear to milky sap. The leaves are estipulate and are usually alternate but may be simple or pinnately compound or rarely bi-pinnate (in Spondias bipinnata Airy Shaw and Forman). The flowers are generally not highly conspicuous but are distinctive in having an intrastaminal nectariferous disc and apotropous ovules (an ovule with a raphe that is ventral when ascending and dorsal when descending) that are pendulous and apically, laterally, or basally attached. Morphological fruit diversity is exceedingly high with a myriad of types found in the family. Although the majority of the family has drupaceous fruits, many of these are variously modified for different mechanisms of dispersal. Several other fruit types are also represented. Two genera, Anacardium L. and Semecarpus L. f., have an enlarged edible hypocarp subtending the drupe. One species of Anacardium, A. microsepalum Loesener, lacks the hypocarp and grows in the flooded forests of the Amazon where it may be fish dispersed287. Water dispersal has been reported or purported for three genera, Mangifera L., Poupartiopsis Randrianasolo ined., and Spondias. The variety of mechanisms for wind dispersal seen throughout tribes Anacardieae, Dobineae, and Rhoeae include subtending enlarged sepals (Astronium Jacq., Hermogenodendron ined., Loxostylis Spreng. Ex Reichb., Myracrodruon Allem., Parishia Hook. f.), subtending enlarged petals (Gluta, Swintonia), trichome-covered margins on a globose fruit (Actinocheita F. A. Barkley), trichome-covered margins on a flattened fruit (Blepharocarya F. Muell., Ochoterenaea F. A. Barkley), elm-like samaras encircled with a marginal wing (Campylopetalum Forman, Cardenasiodendron F. A. Barkley, Dobinea Buch.- Ham. ex D. Don, Laurophyllus Thunb., Pseudosmodingium Engl., Smodingium E. Mey.), samaroid fruits with a single wing (Faguetia March., Loxopterygium Hook. f., Schinopsis Engl.), dry syncarps (multiple fruit, Amphipterygium Schiede ex Standl. and Orthopterygium Hemsl.), dry achene-like fruit without a wing (Apterokarpos Rizzini), and elongated ciliate pedicles of sterile florets on broken segments of an inflorescence that function much like a tumbleweed (Cotinus Mill.). The dry utricle fruits of Pachycormus Coville are most likely wind dispersed but there is no direct report of this in the literature287.
1.4.1.2. The genus Pistacia The genus Pistacia belongs to the family Anacardiaceae. It contains ten species that are native to the Canary Islands, northwest Africa, southern Europe, central and western Asia, and North America (Mexico, Texas). They are shrubs and small trees growing to 5–15 m tall. The leaves are alternate, pinnately compound, and can be either evergreen or deciduous depending on species. All species are dioecious, but monoecious individuals of Pistacia atlantica have been noted288. The genus is estimated to be about 80 million years old289. Well known species in this genus include pistachio, terebinth, and Chinese pistache. Ancient Greek physicians, such as Hippocrates, Dioscorides, Theophrastos and Galenos have recommended use of mastic gum obtained from genus Pistacia for gastrointestinal disorders like gastralgia, dyspepsia and peptic ulcer290,291. species of Pistacia have been used in folk medicine as antiflammatory, antipyretic, antibacterial, antiviral, in treatment diarrhea and throat infection292-294. Essential oils of some Pistacia species consist of components such as cymene, linalool, caryophyllene, thujene, fenchene, sabinene, phellandrene, cineol, fenchone, borneol and terpineil295,296. 1.4.1.3. The species Pistacia lentiscus L. Pistacia lentiscus (Greek: ȝĮıIJȓȤĮ) (Mastic) is a dioecious evergreen shrub or small tree of the Pistacio genus growing to 1-8 m tall297 which is cultivated for its aromatic resin, mainly on the Greek island of Chios. Pistacia lentiscus is native throughout the Mediterranean region, from Morocco and Iberian peninsula in the west through southern France and Turkey to Iraq and Iran in the east. It is also native to the Canary Islands. The word mastic derives either from the Greek verb mastichein ("to gnash the teeth", origin of the English word masticate) or massein ("to chew"). Pistacia lentiscus leaves are green, leather-like, and oval. The small flowers grow in clusters and are reddish to green. The fruit is an orange-red drupe that ripens to black. Mastic is tapped from June to August by making numerous, longitudinal gouges in the tree bark. The transparent, yellow-green resin is collected every 15 days. If chewed, it becomes “plastic,” with a balsamic/turpentine-like odor and taste298.
1.4.2. TRADITIONAL MEDICINE
Pistacia lentiscus L (Mastic) is an evergreen shrub or small tree growing to 1 – 8 m tall297 with a long tradition in folk medicine since the ancients Greeks299. The aerial part has traditionally been used as a stimulant, for its diuretic properties, and to treat hypertension, coughs, sore throats, eczema, stomach aches, kidney stones and jaundice299,300. Mastic gum from Pistacia has been used by traditional healers for the relief of upper abdominal discomfort, stomachaches, dyspepsia and peptic ulcer301. In the past, P. lentiscus L. oil is used in several industrial application such as perfumery, food and pharmaceutical and it has been re-evaluated recently as a flavouring in alcoholic beverages and chewing gum302.
1.4.3. PHYTOCHEMISTRY The chemical composition of the essential oil of this plant reveals the presence of several main compounds: myrcene (19-25%)303, Į-pinene (16%)304, terpinen-4-ol (22%)304, į-3-carene (65%)305, myrcene, limonene, terpinen-4-ol, Į-pinene, ȕ-pinene, Į-phellandrene, sabinene, para-cymene and Ȗ-terpinene306.
1.4.4. PHARMACOLOGY The aerial parts seem to have no or only a weak antimicrobial activity against the Gram (-) and Gram (+) bacteria307. The antifungal activity appears to be much more interesting against the clinical yeast297 and the pathogenic agricultural fungi308. The essential oil of the resin proved to be very active against micro-organisms and fungi, whereas the oils from the leaves and the twigs showed a moderate activity against the bacteria and was completely inactive against the fungi309. The antioxidant properties of the leaves phenolic compounds were reported: they act as a scavenger of the 1,1-diphenyl-2-picrylhydrazyl (DPPH)310. It was also shown that the presence of gallic acid and its derivative, the 1, 2, 3, 4, 6-pentagalloylglucose in the fruits, play a protecting role against lipid peroxidation induced by H2O2 in K562 cell line311. The Chios mastic gum (CMG) is also known to contain compounds that inhibit the proliferation and induce the death of HCT116 human colon cancer cells in vitro312. The iron-induced lipid peroxidation in rat liver homogenates was suppressed by aqueous extracts, without affecting mitochondrial respiration in cultured HepG2 and PC12. This extract administered daily for 5 weeks to rats was shown to contain compound causing hepathotoxic effect313.
1.4.5. TOXICOLOGY Most toxicity related to mastic or source P. lentiscus involves allergic reactions. The plant pollen is a major source for allergic reactions314. The first report of immunological reactions to pollen extracts of Pistacia genus occurred in 1987. 68 A monographic review of mastic's chemistry, pharmacology, and toxicity is available. Children ingesting mastic may develop diarrhea315. A 13-week toxicity study in rats documented changes in hematological parameters including increased white blood cell and platelet counts. Increases in total proteins, albumin, and total cholesterol were also documented. Liver weights increased in a dose-dependent manner and decreased body weight was documented at high doses316. Interestingly, some studies report hepatoprotective effects317 from the aqueous extracts, while others identify hepatotoxic effects318.
1.5. PHARMACOGNOSY OF JUNIPERUS PHOENICEA 1.5.1. BOTANY AND TAXONOMY The following lists the botanical name, synonyms, and the classification of Juniperus phoenicea. Botanical Nomenclature: Juniperus phoenicea L. Synonym: Juniperus phoenicea var. phoenicea. Plant classification: Kingdom: Plantae-Plant Division: Pinophyta Class: Pinopsida Order: Pinales Family: Cupressaceae Genus: Juniperus Species: J. phoenicea
Figure 20. Juniperus phoenicea L.
1.5.1.1. The family Cupressaceae
A family of 19 genera and 130 species of trees and shrubs. Members differ from the Pinaceae in that the leaves and cone-scales are usually opposite or whorled and the ovules erect. The genera include callirtris (16 spp., Australasia), Thuja (5 spp., China, Japan and North America), Cupressus (15-20 spp.), Chamaecyparis (7 spp.), Juniperus (60 spp., northern hemisphere). Juniperus communis yields juniper berries and volatile oil (q.v); J. virginiana, the red cedar wood used for pencil; and J. Sabina, volatile oil of savin; J. oxycedrus, by destructive distillation, yields oil of cade, which was formerly much used in veterinary work. This tar-like oil contains cadinene and phenols. Various diterpenes and flavonoids of the family have been studied319.
1.5.1.2. The genus Juniperus
The genus Juniperus includes 60 to 70 species of aromatic evergreen plants native to northern Europe, Africa, Asia and North America. The plant bear blue or reddish fruits variously described as berries or berry-like cones. Junipers are widely used as ornamental trees. The cone is a small green berry during its first year or growth and turn blue-black during the second year. The small fowers bloom from May to June.
1.5.1.3. The species Juniperus phoenicea L.
Juniperus phoenicea L., (Phoenicean Juniper or Arâr) is an evergreen coniferous shrub or small tree occurring athroughout the Mediterranean region, from Morocco and Portugal east to Turkey and Egypt, and also on Madeira and the Canary Islands, and on the mountains of western Saudi Arabia near the Red Sea and grows up to 10 m in height; it can be either prostate or erect. Its preferred habitat
is heath, moorland and chalk downs, but is also found as
undergrowth in mixed open forests. It is particularly common in pastures where sheep graze as they eat the berries and distribute the seeds in their faeces. As its botanical name suggests, Juniperus phoenicea often occurs in groups. The bark is choclate-brown tinged with red. The leaves, 5-20 mm long, are needle-like and stalkless, occurring in whorls of three, and are pale green below and dark shiny green on the other three sides. The male plant bears 1 cm long cones, the female a much smaller one; the fruit about 1 cm in diameter appears on the female plant. Initially green, it turns purplish-black with a grayish bloom in the second and third year and has a triangular indentation at the apex. Flowering takes place in April and May and the fruit ripen in September and October of the following year.
1.5.2. TRADITIONAL MEDICINE
Juniper berries (the mature female cone) have long been used as flavoring agents in foods and alcoholic beverages such as gin. Production by the Apothecaries and other historical uses for gin has been reported320. Gin's original preparation used Juniper for the
treatment of
kidney ailments. The berries also serve as seasoning, for pickling of meats and as flavoring for liqueurs and bitters. Other uses include perfumery and cosmetics. Oil of Juniper, also known as oil of Sabinal, is used for catgut ligatures321. Juniper tar is also used for its gin-like flavor and in perfumery. In herbal medicine, Juniper oil has been used as a carminative and as a steam inhalant in the management of bronchitis. It has also been used to control arthritis.
1.5.3. PHYTOCHEMISTRY
Juniper berries contain about 2% volatile oil, juniperin, risins (about 10%), proteins and formic, acetic and malic acids. In addition, fatty acids, sterols and terpenes content has also been analyzed by gas chromatography, obtained from extract of ripe and unripe Juniper berries322. The dried ripe fruits contain oil of juniper, pinene, cadinenes, camphene and a number of other diterpene acids. The volatile oil is composed of more than 50% monoterpenes (pinene, myrcene, sabinene) with many minor constituents. Steam distillation of the berries yields mono- and sesquiterpenes from the Oil323. In other studies, isolation, chemical characterization and composition of the essential oil of Juniper are described, revealing 23 compounds324. Isolates of dimeric proanthocyanidins (tannin-producing), from the bark extracts of Juniperus phoenicea have also been reported325, along with the polyprenols in the juniper pine needles325. Two neolignan glycosides (junipercomnosides A and B) have been isolated from the aerial parts of Juniperus phoenicea along with two known neolignan glycosides and seven flavonoid glycosides326.
1.5.4. PHARMACOLOGY Juniper berry oil has been used as diuretic. This activity is most likely due to the presence of terpinen-4-ol, which is known to increase renal glomerular filtration rate327. As a result Juniper berries are often found in herbal diuretic products. The effects of juniper berry oil on urinary tract disease have also been reported328. Juniper has been used in phytotherapy and cosmetics in the eastern Mediterranean region329. Reported therapeutic uses of juniper include juniper baths for the treatment of neurasthenic neurosis3301. and management of scalp psoriasis331. In traditional Swedish medicine, Juniperus phoenicea has been used to treat wounds and
inflammatory diseases. A study evaluates its inhibitory activity on prostaglandin biosynthesis and platelet activating factor (PAF)-induced exocytosis in vitro332. Dried berries of juniper and juniper decoction have been evaluated into recent animal studies. Results support hypoglycemic activity in streptozotocin-induced diabetic mice333. Berry extracts increase uterine tone, Anti-implantation/anti-fertility activity has been also reported in female rats by three similar studies, with one study reported 60% to 70% efficacy334. In another study, the antioxidant effects of juniper are reported335. In veterinary medicine, treatment of psoroptic mange in sheep with extract of Juniperus phoenicea has been reported336. Recently extracts of Juniperus phoenicea L. have have been evaluated for their inhibitory activity on human plate-type 12(S)-lipoxygenase [12(S)-LOX]. The methylene chloride extract and the ethyl acetate extract of Juniperus phoenicea showed a significant inhibition on the production of 12(S)-HETE [12(S)-hydroxy-5,8,10,14-eicosatetraenoic acid] at 100 ȝg/mL (54.0 ± 6.73, 66.2 ± 4.03 and 76.2 ± 3.36 %, respectively). From the methylene chloride extract of the wood, cryptojaponol and ȕ-sitosterol were isolated with inhibitory activity of 100 ȝg/mL (54.0 ± 2.80% [IC50 = 257.5 ȝM] and 25.0 ± 2.15%, respectively. In addition, a lipid fraction containing unsaturated fatty acids contributed to the in vitro activity of the crude extract337.
1.5.5. TOXICOLOGY
Adverse effects due to Juniperus phoenicea in humans are generally of an allergic nature. These include occupational allergy affecting the skin and respiratory tract338, through a sensitivity to airborne juniper pollen339. Two reports state that Chinese, Japanese and Fillipinos tend to be more sensitive to juniper pollens than Caucasains340. Juniper and other related pollens affect 13% to 36% of patients with pollen allergies341. Epidermal contact with juniper tar (eg, preparation for psorasis treatment) can cause potentially carcinogenic DNA damage in human tissue342. Single large doses of juniper berries may cause catharsis, and repeated large doses may be associated with convulsions and renal damag321. Kidney irritation from juniper oil is examined in one report that relates this effect to 1-terpinen-4-ol content328. Because Juniper berries are known to exert their diuretic effect by irritating the renal tissues, products containing Juniper should not be used by individual with reduced renal function. The Juniper oil can induce gastric irritation and may induce diarrhea. Therefore, its use is limited to low conventrations (less than 0.01%) as a beverage flavor. Juniper tar has an oral lethal dose of 8.014 mg/Kg in the rat321.
1.6. PHARMACOGNOSY OF CUPRESSUS SEMPERVIRENS
1.6.1. BOTANY AND TAXONOMY The following lists the botanical name, synonyms, and the classification of Cupressus sempervirens. Botanical Nomenclature: Cupressus sempervirens L. Synonym: Cupressus pyramidalis, Cupressus horizontalis Plant classification: Kingdom: Plantae-Plants Subkingdom: Tracheobionta – Vascular plants Superdivision: Spermatophyta – Seed plants Division: Coniferophyta – Conifers Class: Pinopsida Subclass: Rosidae Order: Pinales Family: Cupressaceae Genus: Cupressus Species: Cupressus sempervirens L.
Figure 21. Cupressus sempervirens L.
1.6.1.1. The family Cupressaceae
The family Cupressaceae was already discussed in Section 1.5.1.1. 1.6.1.2. The genus Cupressus The genus Cupressus includes 16 to 25 species of evergreen trees or large shrubs, growing to 5-40 m tall native to northern Europe, Africa, Asia and North America. The leaves are scalelike, 2-6 mm long, arranged in opposite decussate pairs, and persist for 3–5 years. On young plants up to 1–2 years old, the leaves are needle-like, 5-15 mm long. The cones are 8-40 mm long, globose or ovoid with 4-14 scales arranged in opposite decussate pairs; they are mature in 18–24 months from pollination. The seeds are small, 4-7 mm long, with two narrow wings, one along each side of the seed.
1.6.1.3. The species Cupressus sempervirens L. Cupressus sempervirens, the Mediterranean Cypress (also known as Italian, Tuscan, or Graveyard Cypress, or Pencil Pine) is a species of cypress native to the eastern Mediterranean region, in northeast Libya, southeast Greece (Crete, Rhodes), southern Turkey, Cyprus, Northern Egypt, western Syria, Lebanon, Israel, Malta, Italy, western Jordan, and also a disjunct population in Iran. It is a medium-sized evergreen tree to 35 m (115 ft) tall, with a conic crown with level branches and variably loosely hanging branchlets343. It is very longlived, with some trees reported to be over 1,000 years old. The foliage grows in dense sprays, dark green in colour. The leaves are scale-like, 2-5 mm long, and produced on rounded (not flattened) shoots. The seed cones are ovoid or oblong, 25-40 mm long, with 10-14 scales, green at first, maturing brown about 20–24 months after pollination. The male cones are 3-5 mm long, and release pollen in late winter. It is moderately susceptible to cypress canker, caused by the fungus Seiridium cardinale, and can suffer extensive dieback where this disease is common. The species name sempervirens comes from the Latin for 'evergreen'.
1.6.2. TRADITIONAL MEDICINE
Cupressus sempervirens L. (Cupressaceae) is a tree widely distributed in the Mediterranean region344. Its leaves, cones and young branches play an important role in traditional medicine. For many years this plant has been used as an anthelmintic, antipyretic, antirheumatic, antihemorrhoidal, antidiarrhoeic, astringent, balsamic, vaso-constrictive, antiinflammatory, hair tonic, the fruits of the plant were used traditionally for curing diabetes and as antiseptic345.
Taken internally it used in the treatment of whooping cough, the spitting up of blood, spasmodic, cough, cold, flu and sore throats, while applied externally as a lotion or a diluted essential oil (using an oil such as almond), it astringes varicose veins and hemorrhoids, tightening up the blood vessels, A foot bath of the cones is used to cleans the feet and counte excessive346. An essential oil from the leaves and cones is used in aromatherapy. Its key word is Astringent347. There are other uses of C. sempervirens like: cosmetic (an essential oil distilled from the shoots is used in perfumery and soap making348.
1.6.3. PHYTOCHEMISTRY C. sempervirens (Fig. 21) belongs to the family Cupressaceae349. C. sempervirens is rich in flavonoids constituents such as cupressuflavone, amenoflavone, rutin, quercitrin, quercetin, myricitrin350. Some of phenolic compounds (anthocyanidin, catechines flavones, flavonols and isoflavones) tannins (ellagic acid, gallic acid, phenyl isopropanoids, caffeic acid, coumaric acid, ferulic acid) lignans, catchol. Essential oil of C. sempervirens cones contains 49 components with Į-pinene (48.2%) and 3-carene (19.1%) as the main components351.
1.6.4. PHARMACOLOGY Virostatic activity of C. sempervirens with the help of the immune system by blocking virus entrance in host cells is previously reported352. There seems to be an increasing possibility of finding biological activity among plants with recorded medicinal uses rather than plants randomly selected353. The antiviral activity of ethanol extracts derived from leaves and fruits of C. semipervirens, C. semipervirens var. horizontalis and C. semipervirens cv. Cereiformis on HSV-1 in cultured HeLa cells were investigated354.Results showed that all three plants have antiviral activity against HSV-1 virus. The most active extract was the obtained extract from C. semipervirens. Among the different parts of this medicinal plant tested, the fruit’s extract appeared to possess the strongest anti- HSV activity.
1.6.5. TOXICOLOGY Aqueous preparations and hydroalcoholic extracts of the cones and leaves do not present toxicity; the essential oil, however, should be used with precautions. Aqueous preparations and hydroalcoholic extracts of the cones and leaves do not present toxicity; the essential oil, however, should be used with precautions348.
Chapter 2 2. EXPERIMENTAL
All melting points were determined on a Yanaco micromelting point apparatus and are uncorrected.UV spectra (Shimadzu UV-1203) were recorded in MeOH, whereas IR spectra (Nicolet 510P FT-IR) were obtained as a KBr disk film. 1H NMR (Bruker AM-500,500 MHz) and
13
C NMR (Bruker AC-200, 75 MHz) spectra were acquired in MeOH-d4 with TMS as
internal standard, whereas EIMS (Shimadzu QP-5000/Gc-17A/DI-50) and HREIMS (VG-ZABVSEQ) were recorded at 70 eV (ionizing potential) using a direct inlet system. To monitor the preparative separations, analytical thin-layer chromatography (TLC) was performed at room temperature on precoated 0.25 mm thick silica gel 60 F254 glass plates (20 x 20 cm). Chromatograms were visualized after drying (i) by UV light and (ii) by a phenol specific spray reagent, FeCl3 (3% in dry ethanol). All other chemicals and reagents were analytical grade.
2.1. MATERIALS AND METHODS 2.1.1. PLANT MATERIALS The leaves, barks and roots of Juniperus phoenicea,Pistacia lentiscus and Cupressus sempervirens were collected from Jebal Akhdar mountains (Libya), in February 2008. The plants were identified by the Botany Department, Omar Almuktar University, and voucher specimens were deposited in the herbarium of that Department. The plant samples were airdried and ground into uniform powder using a Thomas-Willey milling machine. 2.1.2. PHYTOCHEMICAL SCREENING Chemical tests were carried out on the aqueous and alcoholic extracts and on the powdered specimens using standard procedures to identify the constituents as described by Sofowara355, Trease and Evans356 and Harborne357. 2.1.2.1. Preparation of Reagents for Phytochemical Screening 2.1.2.1.1. Flavonoid Test Reagents abc-
Aluminium chloride solution: (1g) of aluminium chloride was dissolved in (100 mL) methanol. Potassium hydroxide solution: (1g) of potassium hydroxide was dissolved in (100 mL) water. Ammonia solution: (20 mL) of concentrated ammonia was diluted to (100 mL) water.
2.1.2.1.2. Alkaloid Test Reagents Modified Dragendroff's reagent (Kraut reagent): Stock solution A: (0.85 g) of bismuth nitrate was dissolved in (10 mL) acetic acid and (40 mL) of water was added. Stock solution B: (8.0 g) of potassium iodide was dissolved in (20 mL) water.
2.1.2.1.3. Tannins Test Reagents Ferric chloride solution: (0.1 g) of ferric chloride was dissolved in (100 mL) water.
2.1.2.2.
Preparation of Aqueous Extract for Phytochemical Screening
The aqueous extract of each sample was prepared by extracting 100 g of dried powdered samples by 500 ml of distilled water for 12 h. The extracts were filtered.
2.1.2.3.
Preparation of Alcoholic Extract for Phytochemical Screening
The alcoholic extract of each sample was prepared by extracting 100 g of dried powdered samples by 500 ml of (95%) ethanol for 5 days at room temperator. The extracts were filtered and the ethanol was removed by evaporation under reduced pressure at relatively low temperature (0+@P]
2
>0+@P]
2+
$ P]
2
&+ % P]
Figure 69. Important mass fragmentation of compound 3
On the basis of the above spectral evidences, the structure of compound 3 was deduced to be 1-(3,4-dihydro-3,5,7-trihydroxy-2,2-dimethyl-2H-1-benzopyran-6-yl)-3-(4-hydroxyphenyl)2-propen-1-one.
2+
2
+E +D
2+
ȕ Į
2+
2+
2
Compound 4 was isolated as a yellow crystalline needles from ethanolic extract of the barks of Juniperus phoenicea. The ethanolic extract of the plant was subjected to preparative TLC on silica gel and the TLC plates were developed with the solvent syste n-BuOH–HOAc– H2O, (3:1:1) to obtain compound 4. The structure of compound 4 was elucidated on the basis of MS, IR, UV, 1H NMR and 13C NMR spectroscopic data, including 2D NMR experiments. The IR spectrum of compound 4 (Fig. 70) showed characteristic absorption bands at ȣ (KBr) 3375 (OH), 2952 (CH-stretching), 1622 (C=O), 1525, 14769 and 1433 (C=C, Ar), 1270 (C-O) cm-1. The presence of conjugated carbonyl at 1622 cm-1 indicated that compound 4 belongs to: flavones, flavonols, chalcones or aurones. The UV spectrum of compound 4 in MeOH (Fig. 71) showed characteristic intense absorption band at 369 (Band I) and a diminished absorption band near 240 (Band II) nm, indicating the presence of a chalcone skelton358.
Figure 70. IR (KBr, disc) spectrum of 4
)LJXUH89VSHFWUXPRIFRPSRXQG
The chalcone skeltone of compound 4 can easily be distinguished from the remaining classes of flavonoids by the two set of doublets at įH 7.79 and 7.68 (each 1H, d, J=15.5 Hz) in the 1H-NMR spectrum and its corresponding carbons resonating at įC 128.4 and 143.5 in the 13
C-NMR spectrum, assigned to Į- and ȕ-carbon atoms respectively359-361. The 1H NMR spectrum (Fig. 72) of compound 4 showed a number of signals characteristic
of chalcone and 2-hydroxy-3-methyl-3-butenyl moieties361-363. The chalcone skelton of 4 was identified by a characteristic two 1H doublets with same coupling constants (J = 15.5 Hz) resonating at į 7.79 and 7.68 assigned to the olefinic Į- and ȕ-protons respectively. Two 2H doublets resonating at į 7.5 (J2,3/6,5 = 8.5 Hz) and 6.83 (J3,2/5,6 = 8.5 Hz) are due to the aromatic H-2,6 and H-3,5, respectively, of the B-ring and their coupling constants showed that they are ortho-coupled. A singlet at į 3.90 (s, 3H) was assigned to a methoxy group. Another singlet resonating at į 6.03 (s, 1H) was assigned to H-5'. The presence of a 2-hydroxy-3-methyl-3butenyl moiety was identified by the geminal protons of the C-4ƍƍ terminal double bond resonated at į 4.79 and 4.70 as two singlets, while the protons of the C-1ƍƍ methylene group resonated separately as a double doublet at į 2.96 (H-1ƍƍa, J = 13.6 and 5.5 Hz) and 2.83 (H1ƍƍb, J = 13.6 and 7.5 Hz). The oxymethine proton of the moiety was observed as a doublet (J = 8.9 Hz) at į 4.33, and the methyl group protons (H-5ƍƍ) resonated at į 1.80. The
13
C-NMR spectrum (Fig. 73) of compound 4 showed resonances for twenty one
carbon atoms with two methyl, two methylene, eight methine and nine quaternary carbons in the molecule. The chalcone skeleton of 4 was further supported by the presence of two characteristics olefinic carbons resonating at įC 128.4 (Į-C) and 143.5 (ȕ-C). The downfield signal at įC 194.1 was assigned to the ketonic carbonyl carbon. The downfield chemical shift of
the C=O ketonic carbon indicated the presence of an O-methy group at the adjacent C-6' position in A-ring361. The other signals at įC 106.4 (C-1'), 166.7 (C-2'), 106.7 (C-3'), 164.9 (C4'), 92.1 (C-5'), 162.9 (C-6'), 29.8 (C-1"), 76.7 (C-2''), 148.8 (C-3"), 110.9 (C-4") and 17.9 (C5") further supported the presence of a chalcone skeleton and 2-hydroxy-3-methyl-3-butenyl substituted A-ring. The carbons of the aromatic B-ring resonated at įC 125.8 (C-1), 131.3 (C2,6), 116.9 (C-3,5) and 161.1 (C-4), indicated a C-4 substituted B-ring. The methyl carbon signal at įC 56.2 (C-6') indicated the presence of O-methyl group in the molecule. The complete 13
C-NMR chemical shift data for 4, when compared with the reported data, indicated that the
aglycone is a chalcone having 2-hydroxy-3-methyl-3-butenyl as a side chain365. The complete 13
C-NMR and multiplicity data of compound 4 are presented in Figure 73.
1H H-Į H-ȕ H-2,6 H-3,5 H-5' H-4"a H-4"b H-2" 6'-OMe H-1Ǝa H-1"b CH3-5"
ppm 7.79 7.68 7.50 6.83 6.03 4.79 4.70 4.33 3.90 2.96 2.51 1.80
Mult (J) d (15.5) d (15.5) d (8.5) d (8.5) s s s dd (7.8, 5.5) s dd (13.6, 5.5) dd (13.6, 7.5) s
Integ 1H 1H 2H 2H 1H 1H 1H 1H 3H 1H 1H 3H
+E
+2 +2
+E +D
+D
2
2+
2+
ȕ ǂ
2
Figure 72. 1H NMR spectrum of compound 4 in MeOH-d4.
13C C-ȕ C-Į C=O C-1' C-6' C-5' C-4' C-3' C-2' C-1 C-2,6 C-3,5 C-4 C-1" C-2" C-3" C-4" C-5" 6'-OMe MeOH TMS
ppm 143.5 128.4 194.1 106.4 162.9 92.1 164.9 106.7 166.7 125.8 131.3 116.9 161.1 29.8 76.7 148.8 110.9 17.9 56.2 49.O3 0.0
HMQC HMBC X H-Į;H-2.6 X H-Į;H-ȕ;H-3,5 H-Į;H-ȕ, H-5' H-5' H-5';6'-OMe H-5';H1"a,b H-1"a,b;H-2"; H-5' H-1"a,b H-ȕ X H-2',6'; H-ȕ X H-3,5; H-2',6' H-2,6; H-3,5 X H-2" X H-4"a,b; H1"a,b; Me-5" Me-5";H-2"; H-1"a,b X Me-5"; H-2" X H-4"a,b;H-2" X
+E
+2 +2
+D +E +D
2
2+
13
2+
ȕ ǂ
2
Figure 73.
C NMR spectrum of compound 4 in MeOH-d4.
The HMQC technique was used to establish the direct one-bond 1H-13C connectivities. The protons of rings A, B and C i.e. H-5' (įH 6.03), Į-H (įH 7.79), ȕ-H (įH 7.68), H-2,6 (įH 7.50) and H-3,5 (įH 6.83) showed one-bond correlations with C-5' (įC 92.1), Į-C (įC 128.4), ȕC (įC 143.5), C-2,6 (įC 131.3) and C-3,5 (įC 116.9), respectively (Fig. 74). Protons signals of the prenyl moiety, i.e. H-1"a (įH 2.96), H-1"b (įH 2.78), H-2" (įH 4.33), H-4"a (įH 4.79), H-4"b (įH 4.70) and H-5" (įH 1.80) showed direct connectivities with C-1" (įC 29.8), C-2" (įC 76.7), C-4" (įC 110.9) and C-5" (įC 17.9), respectively.
Figure 74. 1H-13C HMQC NMR spectrum of 4 in MeOH-d4
1
H-1H COSY NMR spectrum of compounds 4 showed coupling between the doublets of
H-2,6 (įH 7.50), H-3,5 (įH 6.83) of aromatic B-ring (Fig. 75). The Į-H (įH 7.79) showed correlation with the ȕ-H (įH 7.68). Interactions of the C-1" methylene protons (įH 2.96 and 2.78) with the C-2" methine proton (įH 4.33) was observed. The latter (H-2") showed coupling with the C-5" methyl protons (įH 1.80). Intractions of the C-4" methylene protons (įH 4.79 and 4.70) with the C-5" methyl protons (įH 1.80) was observed.
Figure 75. 1H-1H COSY NMR spectrum of compounds 4 in MeOH-d4
Structure 4 was finally assembled with the help on Heteronuclear Multiple Bond Connectivity (HMBC) experiment (Fig. 76). The H-5' (įH 5.93) showed two-bond couplings with C-4' (įC 164.9) and C-6' (įC 162.9), three-bond couplings with C-3' (įC 106.7) and C-1' (įC 106.4). The H-2,6 (įH 7.50) of aromatic B-ring showed coupling with C-2,6 (įC 131.3), C-3,5 (įC 116.9), C-4 (įC 161.1) and ȕ-C (įC 143.5), while H-3,5 signal (įH 6.83) was found to be coupled with C-3,5 (įC 116.9), C-4 (įC 161.1) and Į-C (įC 128.4). The Į-H (įH 7.79) showed coupling with Į-C (įC 128.4), ȕ-C (įC 143.5) and C=O (įC 194.1), while H-ȕ (įH 7.68) signal was found to be coupled with C-2,6 (įC 131.3), C-Į (įC 128.4), C-1 (įC 125.8) and C=O (įC
194.1). The O-methyl protons (įH 3.90) were found to be coupled with C-6' (įC 162.9), indicating that the O-methyl group was present at C-6' of ring A. The attachment of the 2hydroxy-3-methyl-3-butenyl moiety at C-3' of ring A was inferred from the HMBC interaction of C-1" methylene protons H-1"a,b (įH 2.96, 2.78 ) with the C-3' signal (įC 106.7) of the aglycone. The H-1"a,b (įH 2.96, 2.78) showed heteronuclear ineractions with C-3' (įC106.7), C-2' (įC166.7), C-4' (įC 164.9), C-2" (įC 69.495) and C-3" (įC 148.8). The H-2" (įH 4.33) showed coupling with C-3' (įC 106.7), C-1" (įC 29.8), C-3" (įC 148.8), C-4" (įC 110.9) and C5" (įC 17.9), while Me-5" (įH 1.80) signal was found to be coupled with C-2" (įC 76.7), C-3" (įC 148.8) and C-4" (įC 110.9). The H-4"a,b (įH 4.79, 4.70) showed heteronuclear ineractions with C-2" (įC 76.7) and C-5" (įC17.9).
Figure 76. 1H-13C HMBC NMR spectrum of 4 in MeOH-d4.
The high-resolution mass spectrum of 4 (Fig. 77) showed a [M-1]- ion at 369.1421 corresponding to a molecular formula of C21H22O6. The electrospray ionization mass spectrum (ESI-MS) of 4 showed several fragments characteristic of a prenylated chalcone skeleton. The major peaks at m/z 249 (A fragment) and 119 (B fragment) were due to the cleavage of ring C through a retro-Diels Alder mechanism (Fig. 78), and indicated the presence of a 2-hydroxy-3methyl-3-butenyl moiety and methoxy group on ring A and a hydroxyl group on ring B of the aglycone362,363.
Figure 77. ESI-MS2 spectrum of compounds 4
2+ +2
2+
+2
2+
2
+2
2
2
2+
+2
2
2
2
2
& 2
2
2
2
2
& 2
$ P]
>0+@P]
>0+@P]
2
2 +2
+2 2
+2
+2
2
2
2
2
2
2
&+
% P]
Figure 78. Fragmentation pathways to A and B fragments of compound 4
2
2
2 +2
+2
2
2+
2
2+
2+
2
2
2
2
2
&+
% P]
Figure 79. Alternative pathway to B fragments observed as prominent peak at m/z 119 in the negative ion ESMS spectrum of compound 4
On the basis of the above spectral evidences, the structure of compound 4 was deduced to be 1-[2,4-dihydroxy-3-(2-hydroxy-3-methyl-3-butenyl)-6-methoxyphenyl]-3-(4-hydroxyphenyl)2-propen-1- one.
2+
+2 2+
+2
2
2
Compound 5 was isolated as a faint yellow powder from ethanolic extract of the barks of Cupressus sempervirens. The ethanolic extract of the plant was subjected to preparative TLC on silica gel and the TLC plates were developed with the solvent system CH3Cl3-MeOH, (96:4) to obtain compound 5. The structure of compound 5 was elucidated on the basis of MS, IR, UV, 1 H NMR and 13C NMR spectroscopic data, including 2D NMR experiments. The IR spectrum of compound 5 (Fig. 80) showed characteristic absorption bands at Ȟ (KBr) 3412 (OH), 2922 (CH-stretching), 1631 (C=O), 1550, 1513, 1469 and 1437 (C=C, Ar), 1271 (C-O) cm-1. The presence of carbonyl at 1631 cm-1 indicated that compound 5 belongs to: flavones, flavonols, isoflavone, flavanone dihydroflavonols, chalcones, dihydrochalcones or aurones. The various flavonoid classes can be recognized from each other by their UV spectra. Isoflavones, flavanones and dihydroflavonols all give similar UV spectra as a result of their having little or no conjugation between the A-and B-rings. They are all readily distinguished from flavones and flavonols by their UV spectra, which typically exhibit an intense Band II absorption with only a shoulder or low intensity peak representing band I299. The Band II absorption of isoflavones usually occurs in the region 245 – 270 nm. Both flavanones and dihydrofavonols have their major absorption peak (Band II) in the range 270 – 295 nm and are therefore clearly distinguished from the spectra of isoflavones (which have their Band II peaks between 245 and 270 nm)358. .
Figure 80. IR (KBr, disc) spectrum of 5
The UV spectrum of compound 5 in MeOH (Fig. 81) showed characteristic intense absorption band at 292 (Band II) with only a shoulder or low intensity peak representing Band I, indicating the presence of a flavanone or dihydroflavonol skelton358.
NXWDLEDBNLDB 899LVLEOH VSHFWUXP
Figure 81. UV spectrum of compound 5 1
The H NMR spectrum (Fig. 82) of compound 5 showed a number of signals characteristic of flavanone and isopreny moieties302-304. The flavanone skelton of 5 was identified by the signal for the C-2 proton appeared as quartet (two doublets Jcis = 3.1 Hz, Jtrans = 12.8 Hz) at į 5.29 ppm as a result of the coupling of the C-2 proton with the two protons at C-3299. The C-3 protons couple with each other (J = 17.0 Hz) in addition to their spin-spin intraction with the C2 proton, thus giving rise to two overlapping quartets at į 3.10 (H-3a, J = 17.6 and 12.8.5 Hz) and 2.67 (H-3b, J = 17.0 and 3.1 Hz). Two of the signals of each quartet , however, are weak and are often not observed (Fig. 3.60). Two 2H doublets resonating at į 7.31 (J2',3'/6',5' = 8.5 Hz) and 6.81 (J3',2'/5',6' = 8.5 Hz) are due to the aromatic H-2',6' and H-3',5', respectively, of the B-ring and their coupling constant showed that they are ortho-coupled. A singlet resonating at į 5.93 (s, 1H) was assigned to H-8. The presence of a prenyl (isopentenyl) unit was indicated by typical chemical shifts and J values364. The H-2' olefinic proton signal of isopentenyl unit was observed as a triplet at į 5.18 (1H, t, J = 7.2 Hz), while the two methyl signals typically appeared as singlets at į 1.75 and 1.65 (each 3H, s). The signal corresponding to the methylene CH2-1' protons appeared as one broad isochronic (2H) doublet at į 3.20 with J = 7.2 Hz.
1H H-2,6 H-3,5 H-8 H-2 H-2" CH2-1" H-3a H-3b CH3-4" CH3-5"
ppm Mult (J) 7.31 d (8.5) 6.81 d (8.5) 5.93 s 5.29 dd(12.8, 3.1) 5.18 t (7.2) 3.20 br-d 3.10 dd (17.0, 12.8) 2.67 dd (17.0, 3.1) 1.75 s 1.65 s
Integ 2H 2H 1H 1H 1H 2H 1H 1H 3H 3H
+2
2+
D
2
D
2
2+
+E
+D
Figure
82. 1H NMR spectrum of compound 5 in MeOH-d4
The 13C-NMR spectrum (Fig. 83) of compound 5 showed resonances for twenty carbon atoms with two methyl, two methylene, seven methine and nine quaternary carbons in the molecule. The flavanone skeleton of 5 was further supported by the presence of two characteristics methine carbon at įC 80.3 (C-2), a methylene carbon at įC 44.02 (C-3) and a carbonyl carbon at įC 198.1 (C-4), 143.5 (ȕ-C). The downfield chemical shift of the C=O ketonic carbon indicated the presence of a hydroxyl group at the adjacent C-5 position in A-ring 302 . The other signals at įC 106.0 (C-4a), 166.2 (C-8a), 109.1 (C-8), 163.2 (C-7), 96.4 (C-6), 158.94 (C-5), 22.52 (C-1"), 124.0 (C-2''), 131.6 (C-3"), 25.9 (C-4") and 17.9 (C-5") further supported the presence of a flavanone skeleton and isoprenyl group substituted A-ring. Similarly the carbons of the aromatic B-ring resonated at įC 131.4 (C-1'), 128.9 (C-2',6'), 116.3
(C-3',5') and 158.9 (C-4'), indicating a C-4' substituted B-ring. The complete
13
C-NMR
chemical shift data for 5, when compared with the reported data, indicated that the aglycone was a flavanone having isoprenyl as a side chain365. The complete data of compound 5 are presented in Figure 83.
13C C=O C-8a C-7 C-5 C-4' C-3" C-1' C-2',6' C-2" C-3',5' C-8 C-4a C-6 C-2 MeOH C-3 C-4" C-1" C-5" TMS
ppm 198.1 166.2 163.2 161.6 158.9 131.6 131.4 128.9 124.0 116.3 109.1 106.0 96.4 80.3 49.0 44.0 25.9 22.5 17.9 0.0
HMQC x x x x x x x
HMBC H-Į; H-ȕ H-1" H-5', H-1" OCH3-6'; H-5' H-3,5; H-2,6 H-4",5" H- Į; H-3,5 H-ȕ; H-2,6 H1"; Me-4",5" H-2,6; H-3,5 H-5', H-1" H-5' CH3-5" CH3-4"
13
C-NMR and multiplicity
+2
2+
D
2
D
2+
2
+E
+D
Figure 83.
13
C NMR spectrum of compound 5 in MeOH-d4.
The HMQC technique was used to establish the direct one-bond 1H-13C connectivities. The protons of rings A, B and C i.e. H-6 (įH 5.93), H-2 (įH 5.29), H-3a,b (įH 3.10, 2.67), H-2',6' (įH 7.31) and
H-3',5' (įH 6.81) showed one-bond correlations with C-6 (įC 96.45), C-2 (įC
80.3), C-3 (įC 44.0), C-2,6 (įC 128.96) and C-3,5 (įC 116.3), respectively (Fig. 84). Similarly protons signals of the isopentenyl moiety, i.e. H-1" (įH 3.20), H-2" (įH 5.18), H-4" (įH 1.75) and H-5" (įH 1.65) also showed direct connectivities with C-1" (įC 22.5), C-2" (įC 124.0), C-4" (įC 25.99) and C-5" (įC 17.9), respectively.
Figure 84. 1H-13C HMQC NMR spectrum of 5 in MeOH-d4
1
H-1H COSY NMR spectrum of compounds 5 showed coupling between the doublets of
H-2',6' (įH 7.31), H-3',5' (įH 6.81) of aromatic B-ring (Fig. 85). Interaction of the C-1" methylene protons (įH 3.20) with the C-2" methine proton (įH 5.20) was observed. The latter (H-2") also showed coupling with the C-4" methyl protons (įH 1.76), which was in turn coupled with the C-5" methyl protons (įH 1.65). The C-1" methylene protons showed correlation with the C-4" methyl protons (įH 1.76), which was in turn coupled with the C-5" methyl protons (įH 1.65).
Figure 85. 1H-1H COSY NMR spectrum of compounds 5 in MeOH-d4
Structure 5 was finally assembled with the help on Heteronuclear Multiple Bond Connectivity (HMBC) experiment (Fig. 86). The H-6 (įH 5.93) showed two-bond coupling with C-7 (įC 163.2) and C-5 (įC 158.9), three-bond coupling with C-8 (įC 109.1) and C-4a (įC 106.0). The H-2',6' (įH 7.31) of aromatic B-ring B showed coupling with C-2',6' (įC 128.9), C3',5' (įC 116.3), C-4' (įC 158.9) and C-2 (įC 80.3), while H-3',5' signal (įH 6.81) was found to be coupled with C-3',5' (įC 116.3), C-4' (įC 158.9) and C-2',6' (įC 128.9). The H-3a (įH 3.10) showed coupling with C-2 (įC 80.44), C-1' (įC 131.4), C-2',6' (įC 128.9), C=O (įC 198.1), C-4a (įC 106.0) and C-8a (įC 166.1), while H-3b (įH 2.67) signal was found to be coupled with C-3 (įC 44.0). The H-2 (įH 5.29) showed coupling with C-1' (įC 131.6). The attachment of the
isopentenyl moiety at C-8 of ring A was established by the HMBC interaction of methylene H1" (įH 3.20) with C-8 (įC 109.4) of aglycone. The H-1" (įH 3.20) showed two-bond couplings with C-8 (įC 109.4) and C-2" (įC 124.0), and three-bond couplings with C-8a (įC 166.1), C-7 (įC 163.2) and C-3" (įC 131.6). The Me-4" (įH 1.75) showed couplings with C-3" (įC 131.6) and C-1" (įC 22.5), while Me-5" (įH 1.65) signal was found to be coupled with C-3" (įC 131.6) and C-1" (įC 22.5). Some observations in the HMBC spectrum included the following: Ɣ There were no correlations observed with C-3. Ɣ A C-8/H-8 crosspeak appears as a "doublet" in the F1 direction.
Figure 86. 1H-13C HMBC NMR spectrum of 5 in MeOH-d4
The high-resolution mass spectrum of 5 (Fig. 87) showed a [M-1]- ion at 339.1226 corresponding to a molecular formula of C20H20O5. The electrospray ionization mass spectrum (ESI-MS) of 5 showed several fragments characteristic of an isoprenelyted flavanone skeleton. The major peaks at m/z 219 (A fragment) and 119 (B fragment) were due to the cleavage of ring C through a retro-Diels Alder mechanism (Fig. 88), and indicated the presence of an isoprenelyl unit and two hydroxy groups on ring A and a hydroxyl group on ring B of the aglycone362,363.
Figure 87. ESI-MS2 spectrum of compounds 5 2+
2
2
2
2+
2
&
2 >0@ P]
2
2
2+
2+
2 &
2
2+
$ P]
2 2
+2
2
2
+2
2+
2
2+
2
&+
% P]
Figure 88. Fragmentation pathways to A and B fragments of compound 5
2
On the basis of the above spectral evidences, the structure of compound 5 was deduced to be 6-prenylnaringenin.
REFERENCES 1. Balandrin, F.M., Klocke, J.A., Wurtele, E.S., and Bollinger, W.H., Sciences,228, 1154 (1995). 2. Dewick, P.M., Medicinal Natural Products. A Biosynthetic Approach, 2nd ed., John Wiley & Sons, West Sussex 2001. 3. Robard, K., Prenzler, P. D., Tucker, G., and Glover, W., Food Chemistry, 66, 401 (1999). 4. Bravo, L., Nutrition reviews, 56, 317 (1998). 5. Harborne, J.B., General procedures and measurement of total phenolics, in Methods in Plant Biochemistry, Vol. 1, Plant Phenolics, Harborne, J.B., Ed., Academic Press, London, 1989, chap. 1. 6. Marais, J.P.J., Deavours, B., Dixon, R.A., and Ferreira, D., The stereochemistry of flavonoids, in The Science of Flavonoids, Grotewold, E., Ed., Springer Science, New York, 2006, chap. 1. 7. Pieatta, G.P., Flavonoids as antioxidant, J. Nat. Prod., 63, 1035 (2000). 8. Harborne, J.B., and Williams, C.A., Phytochemistry, 55, 481 (2000). 9. Nowakowska, Z., Eur. J. Med. Chem., 42, 125 (2007). 10. Williams, C.A., Harborne, J.B., Geiger, H., and Hoult, J.R.S., Phytochemistry, 51, 417 (1999). 11. Weimann, C., Goransson, U., Ponprayoon-Claeson, P., Bhlin, L., Rimpler, H., and Heinrich, M., J. Pharm. Pharmacol., 54, 99 (2002). 12. Havsteen, B.H., Pharmacol. Therapeutics, 96, 67 (2000). 13. Snow, R.W., Guerra, C.A., Noor, A.M., Myint, H.Y., and Hay, S.I., Nature, 434, 214 (2005). 14. Beecher, G.R., J. Nutr., 133, 3248S (2003). 15. Stobiecki, M., Kachlicki, P., Isolation and identification of flavonoids, in The Science of Flavonoids, Grotewold, E., Ed., Springer Science, New York, 2006, chap. 2. 16. Andersen, Ø.M., and Markham, K.R., Flavonoids: Chemistry,Biochemistryand Applications, CRC Press: Boca Raton, 2006. 17. Prasain, J.K., and Barnes, S., Mol. Pharm., 4, 846 (2007). 18. Timberlake, C.F., and Henry, B.S., Endeavour, 10, 31 (1986). 19. Winkel-Shirley, B., Plant physiology 126, 485 (2001). 20. Taylor, L.P., and Grotewold, E.,Current opinion in plant biology, 8, 317 (2005). 21. Treutter, D., Plant biology, 7, 581 (2005). 22. Harnly, J.M., Doherty, R.F., and Beecher, G.R., Journal of agriculturaland food chemistry, 54, 9966 (2006). 23. Bentsath, A., St.Rusznyak, I., and Szent-Gyorgyi, A., Nature, 138 (1936). 24.
Harborne,
J.B.,
The
Flavonoids:
Advances
and Hall, London, 1994.
in
Research
Since
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