FOLK MEDICINE-

Cratoxylum cochinchinense

ANTIOXIDANT

BUT

CYTOTOXIC

TANG SOON YEW

(BSc (Hons), UNSW, Australia)

A thesis submitted for the degree of

Department of Biochemistry

NATIONAL UNIVERSITY OF SINGAPORE

2005 ACKNOWLEDGEMENTS

Firstly, special thanks to my Family for their support throughout this study.

I also wish to thank my supervisors, Professor Barry Halliwell and A/P Matthew

Whiteman, for their invaluable guidance and advice as well as patience throughout this

study.

Special thanks to Professor Yong Eu Leong and Dr. Yap Sook Peng from the

Department of Obstetrics and Gynecology, Faculty of Medicine, National University of

Singapore for supplying us the Chinese medicinal extracts. This project would not be possible without these extracts, especially Cratoxylum cochinchinense which was

harvested from Sook Peng’s garden in Malaysia.

My sincere thanks to Professor Sit Kim Ping, Department of Biochemistry, Faculty of

Medicine, National University of Singapore for her generous gifts of a number of cell lines used in this study.

My sincere thanks also extend to members of the and Oxidants Research

Group, in one way or the other, they have been a part of making this thesis possible.

Members of the group include Dr. Peng Zhao Feng, Siau Jia Ling, Dr. Andrew

Jenner, Lim Kok Seong, Wang Huang Song, Sherry Huang, Long Lee Hua, Chua

Siew Hwa, Dr. Wong Boon Seng and Dr. Jetty Lee. Last but not least, many thanks to my friends Chai Phing Chian, Ng Kian Hong, Dr.

Mirjam Nordling, members of the Flow Cytometry and Confocal Units and ……

ii List of journal publications

Tang, S. Y., Whiteman, M., Jenner, A., Peng, Z. F., and Halliwell, B. (2004)

Mechanism of cell death induced by an extract of Cratoxylum

cochinchinense (YCT) in Jurkat T cells: the role of and

calcium. Free Radic. Biol. Med. 36:1588-1611.

Tang, S. Y., Whiteman, M., Peng, Z. F., Jenner, A., Yong, E. L., and Halliwell, B.

(2004) Characterization of antioxidant and antiglycation properties and isolation of

active ingredients from traditional chinese medicines. Free Radic. Biol. Med.

36:1575-1587.

International conference attended during 2001-2005

1st Asia Pacific Conference and Exhibition on Anti-Aging Medicine 2002: From

Molecular Mechanisms to Therapies. June 23-26.

The 2nd International Conference on Natural Products 2002.

2st Asia Pacific Conference and Exhibition on Anti-Aging Medicine 2003: Diet, Disease,

and Lifestyle. Sep 08-11.

The 3nd International Conference on Natural Products- A Must for Human Survival. Oct

23-25, 2004.

ICCAM (International Congress on Complementary and Alternative Medicines) Herbal

Medicines: Ancient Cures, Modern Science. Feb 26-28, 2005.

International Networking for Young Scientists Symposium. Cancer Biology-Cell

Apoptosis. Mar 1-2, 2005.

iii TABLES OF CONTENTS

Page

Acknowledgements i

List of journal publications iii

International conference attended during 2001-2005 iii

Table of contents iv

Abstracts x

List of tables xii

List of figures xiii

List of abbreviations and keywords xvi

CHAPTER 1. INTRODUCTION

1.1. Introduction 1

1.2. Traditional Chinese medicines 2

1.3. Reactive oxygen species 4

1.3.1. Hypochlorous acid 7

1.3.2. Peroxynitrite 9

1.4. Oxidative stress 10

1.5. Antioxidant defence system 11

1.6. Antioxidant and polyphenolic compounds 13

1.7. Reactive oxygen species in cell signaling 15

1.7.1. Introduction 15

iv 1.7.2. Cellular state 17

1.8. Calcium and its role in cell signaling and cell death 19

1.8.1. Introduction 19

1.8.2. Cellular calcium homeostasis 19

1.8.3. Roles of endoplasmic reticulum in cytosolic calcium homeostasis 21

1.8.4. Role of mitochondria in cytosolic calcium homeostasis 22

1.8.5. Calcium and oxidative stress 22

1.8.6. Calcium and cytotoxicity 23

1.9. Mitochondria 25

1.9.1. Mitochondrial structure and function 25

1.9.2. Mitochondrial reactive oxygen species 26

1.10. Cell death 27

1.10.1. Introduction 27

1.10.2. Differences between apoptosis and necrosis 28

1.10.3. Mitochondrial permeability transition and cell death 34

1.10.4. Calcium overload and mitochondrial permeability transition 37

1.10.5. Bcl-2 proteins and cell death 38

1.11. Plasma membrane NADH reductase/ Plasma membrane redox system 39

1.12. Aims of this study 40

CHAPTER 2. EXPERIMENTAL PROCEDURES

2.1. Materials 41

v 2.2. Methods 43

2.2.1. Extract preparation 43

2.2.2. ABTS assay 43

2.2.3. Ascorbate-iron induced lipid peroxidation 44

2.2.4. Total phenolic content 45

2.2.5. Scavenging of DPPH• (2,2-Diphenyl-1-picrylhydrazyl) 45

•- 2.2.6. Superoxide anion (O2 ) scavenging effect 45

2.2.7. Xanthine oxidase (XO) activity 46

2.2.8. Iron-binding activity 46

2.2.9. Non-enzymatic protein glycation 47

2.2.10. Inhibition of hypochlorous acid-induced DNA damage 47

2.2.11. Isolation, purification, and identification of active ingredient

from Cratoxylum cochinchinense (WN) 49

2.2.12. Bleomycin-iron dependent DNA damage 50

2.2.13. Peroxynitrite scavenging assays 50

2.2.13.1. Synthesis of Peroxynitrite 50

2.2.13.2. Assessment of pyrogallol red (PR) bleaching by

peroxynitrite (ONOO-) 51

2.2.13.3. Measurement of tyrosine nitration 51

2.2.14. Cell culture 52

2.2.15. Cell counting 52

2.2.16. Isolation of human lymphocytes 53

2.2.17. Assessment of cell viability 53

vi 2.2.17.1. MTT assay 54

2.2.17.2. Trypan blue exclusion assay 54

2.2.18. Treatment of Jurkat T cells with YCT 55

2.2.19. O2 electrode assay 55

2.2.20. Cell cycle analysis using flow cytometry 56

2.2.21. Annexin V-FITC and propidium iodide staining of Jurkat T cells 56

2.2.22. Release of lactate dehydrogenase 57

2.2.23. Assessment of cellular and nuclear morphology 57

2.2.24. Caspase 3 and 9 activities 58

2.2.25. Western blot analysis 59

2.2.26. Internucleosomal DNA fragmentation assay 60

•- 2.2.27. Cytofluorimetric measurement of superoxide radicals (O2 ) and

other reactive oxygen species (ROS), lipid peroxidation (LPO),

2+ 2+ 2+ 2+ intracellular Ca ([Ca ]i), mitochondrial Ca ([Ca ]m), mitochondrial

+ transmembrane potential (ΔΨm), intracellular potassium ([k ]i) and

+ sodium ([Na ]i) 60

2.2.28. Multiwell plate reader measurement of lipid peroxidation (LPO) and

mitochondrial transmembrane potential (ΔΨm) 63

2.2.29. Intracellular ATP determination 63

2.2.30. Intracellular measurement 64

2.2.31. Cytochrome c levels 64

2.2.32. Kinetic study of rise in intracellular Ca2+ with Fluo3/ AM 66

vii 2.3. Data analysis 66

CHAPTER 3. RESULTS AND DISCUSSION

RESULTS

3.1. scavenging by TCM extracts 67

3.2. Inhibition of ascorbate-iron induced phospholipid peroxidation 70

3.3. Correlation between TEAC values, total phenolic content and DPPH

reducing ability 71

3.4. Superoxide scavenging and xanthine oxidase inhibition 72

3.5. Iron-binding activity 73

3.6. Scavenging of hypochlorous acid (HOCl); protection against HOCl-mediated

DNA damage 74

3.7. Inhibition of protein glycation 75

3.8. Purification and identification of ‘active’ ingredient from extract WN 76

3.9. Low pro-oxidant effect of YCT extract 79

3.10. Inhibition of PR bleaching and 3-nitrotyrosine formation by YCT 80

3.11. A comparison of our semi-purified extract (YCT) with pure mangiferin 83

DISCUSSION 86

CHAPTER 4. RESULTS AND DISCUSSION

RESULTS

viii 4.1. Cytotoxicity of YCT by MTT assay and propidium iodide staining 93

4.2. YCT-induced loss of viability in Jurkat T cells but not in normal lymphocytes 97

4.3. Changes in cellular morphology 99

4.4. YCT-induced phosphatidylserine exposure and LDH release 101

4.5. Caspase activities and internucleosomal DNA fragmentation 105

4.6. YCT induces rapid oxidative stress 108

4.7. YCT-induced lipid peroxidation in Jurkat T cells 112

2+ 4.8. Mitochondrial Ca overloading and ΔΨm dissipation 115

2+ 4.9. Effects of ruthenium red on mitochondrial Ca and ΔΨm 118

4.10. YCT-mediated changes in cytochrome c and ATP levels 120

4.11. Intracellular glutathione content 124

4.12. YCT-mediated mobilization of extracellular Ca2+ 127

4.13. Ca2+ influx through a non-selective cation channel? 131

4.14. The relationship between ROS generation and Ca2+ influx 133

4.15. Ferricyanide inhibit YCT-induced apoptotic signals 24 h after treatment 135

DISCUSSION 138

CHAPTER 5. CONCLUSION 143

CHAPTER 6. REFERENCES 146

APPENDIX A

ix ABSTRACT

There is considerable interest in the isolation of more potent antioxidant

compounds to treat diseases involving oxidative stress. Thirty-three Traditional Chinese

Medicine (TCM) extracts were examined for their antioxidant activity using the ABTS

(2,2’-azinobis[3-ethylbenzothiazoline-6-sulphonate]) assay. Five extracts with high

activity (Cratoxylum cochinchinense, Cortex magnoliae officinalis, Psoralea corylifolia

L, Curculigo orchioides Gaertn, and Glycyrrhiza uralensis Fisch.) were selected for

further characterization. C. cochinchinense out-performed other extracts in most of the assays tested except phospholipid peroxidation inhibition, where P. corylifolia L showed a higher activity. C. cochinchinense was particularly potent in inhibiting the formation of advanced glycation end products on proteins and strongly inhibited hypochlorous acid- induced DNA damage. We attempted to isolate the active ingredients from C. cochinchinense and obtained an extract (YCT) containing at least 90% mangiferin (MGF) as identified by HPLC and mass spectrometry. However, YCT showed significantly higher activity in assays of phospholipid peroxidation, inhibition of protein glycation,

•− − superoxide (O2 ) and peroxynitrite (ONOO ) scavenging as compared to MGF, suggesting that the non-mangiferin constituents of YCT contribute to its additional antioxidant activities. In cell culture experiments, we show that YCT is selectively toxic to certain cell types and investigated the mechanisms of this toxicity in Jurkat T cells. By flow cytometric analyses, we show that YCT causes intense oxidative stress and a rise in cytosolic Ca2+. This is followed by a rise in mitochondrial Ca2+, release of cytochrome c,

collapse of ∆ψm, and fall in ATP levels, and eventually cell death. The mechanism(s) of

x intense oxidative stress may involve a plasma membrane redox system since cell death is inhibited by potassium ferricyanide, which is an extracellular electron acceptor. Cell death has some features of apoptosis (propidium iodide staining, externalization of phosphatidylserine, limited caspase-3 and -9 activities), but there was no internucleosomal DNA fragmentation.

xi LIST OF TABLES

Table Title Page

1.1. Examples of reactive species 5

1.2. Half-lives of oxygen radicals and related species 7

1.3. Antioxidant defence in biological systems 12

1.4. Comparison of apoptosis, necrosis and paraptopsis 31

3.1. Trolox equivalent antioxidant capacity (TEAC) values of TCM extracts 67

• 3.2. Superoxide (O2 ¯) scavenging and xanthine oxidase (XO) inhibitory effects

of TCM extracts 72

3.3. Inhibition of HOCl induced DNA damage by Cratoxylum cochinchinense 74

3.4. A comparison of the effects of YCT with MGF 83

3.5. A comparison of the cytotoxicity of YCT extract with MGF on Jurkat T

cells and colorectal adenocarcinoma (HT29) cells 85

4.1. YCT-induced changes in HL60 and Jurkat T cells 96

4.2. Generation of H2O2 (µM) in cell culture medium (RPMI) 109

xii LIST OF FIGURES

Figure Title Page

1.1. Structure of mangiferin (1,3,6,7-tetrahydroxyxanthone-C2-β-D-glucoside) 4

1.2. Schematic representation of Ca2+ regulation in cells 20

1.3. Extrinsic and intrinsic pathways of cell death in mammalian cells 33

1.4. Model for caspase activation by mitochondria 35

3.1. Radical scavenging by TCM extracts 69

3.2. Inhibition of ascorbate-iron induced phospholipid peroxidation 70

3.3. Correlation between TEAC values and total phenolic content 71

3.4. Iron-binding activity of TCM extracts 73

3.5. Inhibition of protein glycation (advanced glycation end products (AGE)

formation) by TCM 75

3.6. Identification of mangiferin (MGF) by high performance liquid

chromatography (HPLC-PDA) 77

3.7. Identification of mangiferin by liquid chromatography-mass spectrometry

(LC-MS) 78

3.8. Pro-oxidant effect of YCT as compared to Trolox in causing DNA damage 79

3.9. Inhibition of PR bleaching by YCT and MGF 80

3.10. Inhibition of tyrosine nitration by YCT and MGF 82

3.11. A comparison of the cytotoxicity of YCT with MGF on human hepatoma

cells (HepG2) by MTT assay 85

4.1. Effects of YCT on cells as shown by MTT assay 96

xiii 4.2. YCT-induced loss of viability in Jurkat T cells 97

4.3. Changes in morphology of Jurkat T cells after YCT treatment 100

4.4. Phosphatidylserine exposure (PS) and release of lactate

dehydrogenase (LDH) 102

4.5. YCT-induced activation of caspase-3, -7 and -9 but lack of

internucleosomal DNA fragmentation 106

4.6. Reactive oxygen species (ROS) generation by YCT in Jurkat T cells 109

4.7. YCT-induced superoxide generation in Jurkat T cells 112

4.8. Lipid peroxidation (LPO) induced by YCT in Jurkat T cells 114

2+ 4.9. Mitochondrial Ca levels and ΔΨm in Jurkat T cells 117

2+ 4.10. Effects of ruthenium red (RuR) on mitochondrial Ca , ΔΨm dissipation,

cell arrest and cell death 118

4.11. YCT-mediated changes of cytochrome c and ATP levels in Jurkat T cells 122

4.12. YCT-mediated loss of cellular GSH and the effects of inhibitor on ROS

generation and cell death 125

4.13. YCT-mediated rise in cytosolic Ca2+ in Jurkat T cells through a

non-selective cation channel 127

4.14. Relationships between YCT-mediated ROS generation, Ca2+ influx and lipid

peroxidation (LPO) in Jurkat T cells 135

4.15. Effects of an extracellular electron acceptor, potassium ferricyanide, on YCT-

induced cell death 137

4.16. Effects of potassium ferricyanide on YCT-induced cell death in

xiv HepG2 cells 137

5.1. Diagram illustrating the mechanism(s) of YCT-induced cell death in

Jurkat T cells 145

xv LIST OF ABBREVIATIONS AND KEYWORDS

∆ψm-mitochondrial transmembrane potential

ABTS-2,2’-azinobis[3-ethylbenzothiazoline-6-sulphonate]

AGEs-advanced glycation end products

BHT-butylated hydroxytoluene

BSA-bovine serum albumin

BSTFA-N, O, bis(trimethylsilyl)trifluoroacetamide

DPPH-2,2-Diphenyl-1-picrylhydrazyl

FA-flufenamic acid

GAE-gallic acid equivalents

HOCl-hypochlorous acid

LDH-lactate dehydrogenase

LPO-lipid peroxidation

MGF-mangiferin

NAC-N-acetyl-L-cysteine

NBT-nitroblue tetrazolium chloride

ONOO− -peroxynitrite

PBS-phosphate buffered saline

PMRS-plasma membrane redox system

PR-pyrogallol red

xvi PS-phosphatidylserine

ROS-reactive oxygen species

RuR-ruthenium red

TCM-traditional Chinese medicines

TEAC-Trolox equivalents antioxidant capacity tR-retention time

YCT-Cratoxylum cochinchinense extract

Keywords⎯TCM, Cratoxylum cochinchinense, mangiferin, lipid peroxidation, advanced glycation end products, DNA damage, peroxynitrite, 3-nitrotyrosine, oxidative stress,

Ca2+ influx, mitochondrial depolarization, potassium ferricyanide, plasma membrane redox system.

xvii CHAPTER 1 INTRODUCTION

1.1 Introduction

There is a growing interest in the use of natural products, including herbs and plants, as consumer awareness of their beneficial health effects increases. As late as the

1930s, as high as 90% of the medicines prescribed by doctors or sold over the counter were herbal in origin (Chevallier, 1996).

The popularity of herbal medicine returned after the thalidomide tragedy in 1962 in Britain and Germany, when 3000 deformed babies were born to mothers who had taken a sedative chemical medicine during pregnancy (Chevallier, 1996). Moreover, the high cost of Western medicines has encouraged people to rely on herbs for better health, especially in developing countries (Cardellina, 2002). The poor state of health in Western societies, despite huge health care investment by the government, is another factor that caused people to change their attitude towards herbal medicines. Herbal-based therapies are now recommended for the treatment of several chronic conditions and degenerative disorders where modern medicines have proved inadequate (Iwu and Gbodossou, 2000).

To date, Chinese herbal medicines constitute multi-billion-dollar industries worldwide and more than 1500 herbal products are available in the market as dietary supplements or phytomedicines (Wang and Ren, 2002).

1 1.2 Traditional Chinese medicines

Traditional Chinese medicinal (TCM) herbs are some of the oldest alternative and complementary medicines that have long been used to treat a variety of disorders and improve general health (Wang and Ren, 2002). The origin of TCM is based on the experience accumulated from Chinese people using the TCM to maintain health and to treat diseases for more than 2000 years (Cheng, 2000). The system of TCM is based on two theories that govern good health and longevity, namely yin and yang, and the five elements (van Wyk and Wink, 2004). Yin and yang represent opposites that complement each other, such as cold and hot, and wet and dry. The five elements (i.e. earth, metal, water, wood, and fire) however, represent the spleen, lungs, kidneys, liver, and heart, respectively. The five elements are also linked to our emotions and tastes, and the seasons and climates of our surroundings (van Wyk and Wink, 2004).

Traditional medicine does not cure chronic diseases directly but it tries to restore the body to a normal state by balancing the five elements in our body and to grant vital energy, which has both yin and yang aspects. Thus, TCM practioners usually feel the person’s pulse and look at the tongues and eyes to diagnose the conditions. An imbalance between stress and protective elements in vivo is suggested to play a major role in various diseases (Halliwell and Gutteridge, 1999). Therefore, TCM may play a role in disease prevention rather than healing acute illnesses. One such example is Ginkgo biloba. Other than having a long history as a traditional Chinese medicine, the standardized Ginkgo leaves extracts are also popular as a phytomedicine in Europe and as a dietary supplement

2 in the USA (Wang and Ren, 2002). Some clinical studies have suggested that ginkgo

extracts possess therapeutic activity in a variety of conditions including Alzheimer's

disease, poor memory, age-related dementias, occlusion of ocular blood flow, and the

prevention of altitude sickness (McKenna et al., 2001).

Other functions of herbal extracts reported include antidiabetic (Ichike et al.,

1998), antimicrobial (Lin et al., 1995; Sindambiwe et al., 1999; Duffy and Power, 2001),

antiviral (Wang, 2000; Sindambiwe et al., 1999), anti−inflammatory, antiallergic (Di

Carlo et al., 1999), immunosuppressive, immunostimulatory (Wilasrusmee et al., 2002a;

Wilasrusmee et al., 2002b) and cancer chemoprevention effects (Lee et al., 2001). For example, Cratoxylum cochinchinense is a small genus of Southeast Asia trees belonging to the Guttiferae (Bennett and Lee, 1989). It is used in traditional medicine for many purposes (Bremness, 1994). Bennett et al. (1993), Sia et al. (1995) and Nguyen and

Harrison (1999) have described the triterpenoids, tocotrienols and xanthones constituents from the bark of Cratoxylum cochinchinense. Mangiferin is a member of the C- glucosylxanthone family that has been found in certain ferns and in over hundred species of higher plants, including Cratoxylum species (Bennett and Lee, 1989). Protective effects of mangiferin from different sources against reactive oxygen species (ROS)- related damage both in vitro and in vivo have been documented (Nunez-Selles et al.,

2002). Fig. 1.1 shows the chemical structure of mangiferin (Nott and Robets, 1967).

3

Fig. 1.1. Structure of mangiferin (1,3,6,7−tetrahydroxyxanthone−C2−β−D−glucoside)

Moreover, herbs and spices have also been used for other purposes such as food preservation against lipid oxidation (Finley and Given, 1986), nutrition, flavorings, beverages, dyeing, smoking, cosmetics and other industrial uses (Zheng and Wang,

2001).

1.3 Reactive oxygen species

Free radicals and other reactive species of oxygen (ROS), nitrogen (RNS) and chlorine (RCS) are generated in vivo by various mechanisms including aerobic metabolism, inflammatory responses and exposure to ionizing radiation (Halliwell and

Gutteridge, 1999). Elevated levels of these reactive species are associated with many pathophysiological events (Sies, 1997). Reactive oxygen species (ROS) is a collective term given to reactive non-radical and radical species containing oxygen that include , hypochlorous acid, singlet oxygen and hydroxyl radical. Other examples of reactive species are shown in Table 1.1 (Halliwell and Whiteman, 2004).

4 Table 1.1. Examples of reactive species

Free radicals Non-radicals

Reactive oxygen species (ROS)

•- Superoxide, O2 Hydrogen peroxide, H2O2

Hydroxyl, OH• Hypobromous acid, HOBr

• Hypochlorous acid, HOCl Hydroperoxyl, HO2

• Ozone O3 Peroxyl, RO2 Singlet oxygen (O 1∆g) Alkoxyl, RO• 2

•- Organic peroxides, ROOH Carbonate, CO3 - •- Peroxynitrite, ONOO Carbon dioxide, CO2 Peroxynitrous acid, ONOOH

Reactive nitrogen species (RNS)

• Nitric oxide, NO Nitrous acid, HNO2

• + Nitrogen dioxide, NO2 Nitrosyl cation, NO

Nitroxyl anion, NO-

Dinitrogen tetroxide, N2O4

Dinitrogen trioxide, N2O3

Peroxynitrite, ONOO-

Peroxynitrous acid, ONOOH

+ Nitronium (nitryl) cation, NO2

Alkyl peroxynitrites, ROONO

Nitryl (nitronium) chloride, NO2Cl

5 Reactive chlorine species (RCS)

Atomic chlorine, Cl• Hypochlorous acid, HOCl

Nitryl (nitronium) chloride, NO2Cl

Chloramines

Chlorine gas, Cl2

A free radical may be defined as any species that contains one or more unpaired

electrons (Halliwell and Gutteridge, 1999). The formation of free radicals is ubiquitous.

For example, in the process of oxygen reduction to form water, superoxide anion radical

and hydroxyl radical are formed from oxygen reduction by one and three electrons,

respectively (Halliwell and Gutteridge, 1999). A two electron-reduction process of

oxygen leads to the generation of hydrogen peroxide, which is a non-radical reactive

species. The half-lives of reactive species are an important factor in determining the

damages they can cause to biomolecules (Sies, 1997). For example, the reactions of

hydroxyl radical are diffusion limited because it usually reacts instantaneously with the

target molecules at the site of radical generation and thus, the damage caused by hydroxyl radical is unavoidable and is dealt with by repair processes. On the contrary, peroxyl radical is relative stable with a longer half-life that allows its reactions to occur at other target sites. The longer half-life of peroxyl radical also allows it to be intercepted by antioxidants. Table 1.2 shows the half-lives of oxygen radicals and related species (Pryor,

1986).

6 Table 1.2. Half-lives of oxygen radicals and related species

Radical Half-life (37°C)

HO• (hydroxyl radical) 10-9 sec

RO• (alkoxyl radical) 10-6 sec

ROO• (peroxyl radical) 7 sec

L• (linolenyl radical) 10-8 sec

H2O2 (hydrogen peroxide) Stable; enzymatic reduction

• - O2 (superoxide anion radical) Spontaneous and enzymatic dismutation

1 -6 O2 (singlet oxygen) 10 sec (depends on solvent)

Q- • (semiquinone radical) Days

NO• (nitric oxide radical) 1-10 sec

ONOO- (peroxynitrite) 0.05- 1 sec

For the purposes of this thesis, only hypochlorous acid (HOCl) and peroxynitrite (ONOO-

) will be briefly introduced.

1.3.1 Hypochlorous acid

There is much interest in the chemistry of oxidative damage induced by activated polymorphonuclear cells such as neutrophils. Activated neutrophils contain the enzyme myeloperoxidase (MPO), which is released at the site of inflammation from the azurophilic granules of these cells. MPO catalyses the formation of powerful oxidizing

7 and chlorinating agent, hypochlorous acid (HOCl) from hydrogen peroxide (H2O2) and chloride ions (Cl-) (equation 1) (Weiss et al., 1982; Weiss, 1989).

MPO - + H2O2 + Cl + H ------> H2O + HOCl (1)

Weiss et al. (1982) have estimated that about 40% of the H2O2 generated by activated neutrophils is used to form HOCl. At pH 7.4, HOCl is approximately 50% ionized to

OCl¯ as it has a pKa value of 7.46 (Morris, 1966). Hence, “hypochlorous acid” commonly refers to a mixture of HOCl and OCl¯. HOCl is both a one- and two-electron

oxidant (Koppenol, 1994) and has been shown to oxidize plasma membrane sulphydryl

groups (Schraufstatter et al., 1990), to inactivate α1-antiproteinase (Aruoma et al., 1989), and to react with glutathione, ascorbate and nucleotides (Folkes et al., 1995; Albrich et

al., 1981; Prütz, 1996) in mammalian systems. Moreover, HOCl can potentiate H2O2- mediated DNA damage by inactivation of DNA repair enzymes such as poly(ADP- ribose) polymerase (Van Rensburg et al., 1991; 1992; Pero et al., 1996) and damage

isolated calf-thymus DNA directly, causing extensive oxidative damage to pyrimidines

but less damage to the purines (Whiteman et al., 1997). HOCl is also capable of

chlorinating tyrosyl residues to form 3-chlorotyrosine, which may be used as a biomarker

for HOCl-mediated reaction in vivo (Kettle, 1996). In addition, HOCl has also been

shown to generate hydroxyl radicals on reaction with superoxide (Candeias et al., 1993) or with iron (II) salts (Candeias et al., 1994).

8

1.3.2 Peroxynitrite

Investigators’ interest on nitric oxide (•NO) arises from the very fast reaction of

• •− − NO with superoxide radicals (O2 ), forming peroxynitrite (ONOO ). Peroxynitrite is a

• •− non-radical because the unpaired electrons on NO and O2 have combined to form a covalent bond (Beckman et al., 1994). The reaction rate for the formation of peroxynitrite has been determined to be 6.7 ± 0.9 x 109 M-1. s-1 (Huie and Padmaja, 1993).

Peroxynitrite protonates and breaks down under physiological conditions to give highly

reactive oxidizing and nitrating species. The remarkable stability of peroxynitrite as an

anion in alkaline solution is probably a result of it’s being folded in the cis-conformation

(Tsai et al., 1994). Addition of peroxynitrite has been shown to inactivate aconitase

(Castro et al., 1994; Hausladen and Fridovich, 1994), to inhibit electron carriers such as

succinate dehydrogenase and NADH dehydrogenase in the mitochondrial electron

transport (Radi et al., 1994) and generate oxidized products from a range of biomolecules

(Beckman et al., 1990). Although peroxynitrite has a half-life of less than one second

under physiological conditions, it is able to diffuse distances equal to cellular diameter

(Beckman, 1996); its protonated form (peroxynitrous acid, ONOOH) can easily decay to

• + give strong oxidants such as nitrogen dioxide ( NO2), nitryl cation (NO2 ) and hydroxyl radicals (Beckman and Koppenol, 1996; Greenacre and Ischiropoulos, 2001). The toxicity of these reactive species is associated with numerous human diseases, including cardiovascular, neurodegenerative, cancer, and inflammatory diseases (Greenacre and

Ischiropoulos, 2001).

9 The reaction of peroxynitrite with carbon dioxide is of great importance because

bicarbonate is the main buffering system in vivo (Lymar and Hurst, 1995). Bicarbonate anion is present at a concentration of approximately 25 mM in the blood plasma and 12 mM in intracellular fluids (Denicola et al., 1996). Carbon dioxide can react rapidly with the peroxynitrite anion and thus modulate the oxidative activity of peroxynitrite. This reaction can be interrupted by a synthetic seleno-compound with peroxidase activity, ebselen, which reacts with peroxynitrite nearly 30 times faster than carbon dioxide

(Squadrito and Pryor, 1998). Natural water soluble antioxidants such as urate, ascorbate and glutathione at tissue concentrations are not able to compete with carbon dioxide for peroxynitrite. Furthermore, free and protein bound tyrosine undergo nitration when exposed to peroxynitrite at pH 7.4, forming 3-nitrotyrosine as one product. Peroxynitrite- mediated tyrosine nitration has been shown to be accelerated by superoxide dismutase, myeloperoxidase and horseradish peroxidase (Ischiropoulos et al., 1992; Sampson et al.,

1996). Nitration has been suggested to be a “marker” of peroxynitrite-dependent damage in vivo (Beckman et al., 1994) but in fact may be a general marker of damage by RNS

(Halliwell, 1997a; Greenacre and Ischiropoulos, 2001).

1.4 Oxidative stress

ROS play multiple roles in vivo. For examples, ROS are involved in energy production, phagocytosis during foreign invasion, regulation of cell growth and intercellular signalling (Halliwell, 1997b). Conversely, many reactive species have also been shown to cause oxidative stress and damage to biological molecules that include

10 DNA (Spencer et al., 2000), proteins (Vinson and Howard, 1996; Davies, 2005) and

lipids (Sevanian and Ursini, 2000). Oxidative stress may be characterized by a shift of the cellular redox status to a more oxidized state. Sies (1997) defined oxidative stress as “a disturbance in the pro-oxidant-antioxidant balance in favour of the former, leading to potential damage”. With increasing evidence from experimental, clinical and epidemiological studies on free radical-induced oxidative damage in a variety of diseases such as cardiovascular, cancer, and other neurodegenerative diseases (Halliwell, 1999;

2000; 2001; 2002), the role of antioxidants has received much attention. In other words, any interventions that can decrease or prevent damages caused by any of the reactive species on biomolecules should be beneficial. One such example is antioxidant treatment that may serve to intercept a damaging species.

1.5 Antioxidant defence systems

Many important antioxidant defence systems have evolved to detoxify reactive species generated from our daily aerobic life (Halliwell, 1996; Sies, 1991). Generally, the defence systems can be classified into enzymatic and non-enzymatic. Enzymatic defence systems include superoxide dismutases (this include both the mitochondrial MnSOD, and

CuZnSOD) and H2O2-removing enzymes, such as glutathione peroxidase,

peroxiredoxins, catalase, and other hemoprotein peroxidases. The non-enzymatic systems

include lipophilic and hydrophilic antioxidants, including phenolic compounds. Table 1.3

shows some examples of the defence systems (Sies, 1991). In addition, GSSG reductase

and conjugation enzymes also form part of the antioxidant defence system. The various

11 defences are complementary to each other because they act against different species in

• different cellular compartments. For example, SOD converts O2 ¯ to H2O2, which, in turn, is converted by catalase and peroxidases to H2O. However, despite these antioxidant defence systems, some ROS still escape to cause damage to biomolecules and repair systems may be needed to back up the inadequacies of antioxidant defences (Halliwell and Gutteridge, 1999). Noguchi et al. (2000) have classified antioxidant defence systems into preventative (which suppress the formation of free radicals), radical scavenging

(which suppresses chain initiation and / or breaking chain propagation reactions), repair and de novo, and lastly adaptation (where oxidative stress induces the formation and delivery of antioxidant to the radical generation site).

Table 1.3. Antioxidant defence in biological systems

Enzymatic Non-enzymatic

Superoxide dismutase α-Tocopherol

GSH peroxidases Ascorbate

Peroxiredoxins Glutathione

Catalase Polyphenolic compounds

Ancillary enzymes Artificial antioxidants (BHA and BHT)

-GSSG reductase β-Carotene

-Conjugation enzymes (GSH S- Urate

transferases, sulfonyltransferase, UDP- Plasma proteins

glucuronyltransferase)

12 1.6 Antioxidants and polyphenolic compounds

An antioxidant is defined as ‘any substance that, when present at low concentrations as compared with those of an oxidizable substrate, significantly delays or prevents oxidation of that substrate’ (Halliwell and Gutteridge, 1999). According to

Halliwell and Gutteridge (1999), mechanisms of antioxidant action can include (1) suppressing ROS formation either by inhibition of enzymes or chelating trace elements involved in free radical production; (2) direct scavenging of ROS; and (3) upregulating or protecting antioxidant defences. Direct scavenging of ROS may function by virtue of the capability of antioxidants to donate hydrogen to stabilize reactive and unstable free radicals (Rice−Evans et al., 1996).

The ability of phenolic compounds to chelate transition metals has also been suggested to make a contribution to their antioxidant properties, particularly in Fenton type of reactions (Mira et al., 2002). Iron and copper are powerful promoters of free radical reactions and hence their availability in vivo is tightly controlled (Halliwell and

Gutteridge, 1986). However, transition metal ions are not always safely bound in “non- catalytic” forms, therefore, the metal ions binding properties of phenolic compounds may play a role in the prevention of initiation of chain reactions, particularly iron and copper ions that are involved in lipid peroxidation and DNA fragmentation (Sies, 1993). For instance, (ascorbate), a well known antioxidant, can turn pro-oxidant in vitro when mixed with transition metal ions (Halliwell, 1996).

13 As well as scavenging free radicals, polyphenolic compounds in medicinal plant

extracts may have an effect on the defence system. Lee et al. (2003) demonstrated the

protective effects of medicinal plant extracts in H2O2-induced apoptosis of Chinese hamster lung fibroblast (V79-4) cells. Cells treated with various extracts showed increased activities of SOD, catalase and glutathione peroxidase, hence, these medicinal plant extracts might act by up-regulation of the intracellular defence system against exogenous reactive species. Back in 1977, Hassan and Fridovich showed the induction of catalase and SOD in microorganisms, such as Escherichia coli or Salmonella

typhimurium by treatment with H2O2.

The properties of herbal extracts may be related to its antioxidant actions because

they are rich sources of phenolic compounds (Cai et al., 2004; Zheng and Wang, 2001).

For examples, Zheng and Wang (2001) reported a linear relationship between oxygen

radical absorbance capacity (ORAC) values and total phenolic contents of medicinal

herbs (R = 0.919) and culinary herbs (R = 0.986). Liu and Ng (2000) examined twelve

selected folk medicines and reported that the aqueous extracts of Coptis chinensis,

Paeonia suffruticosa, Prunella vulgaris and Senecio scandens exhibited potent inhibitory

effects against rat erythrocyte hemolysis, and lipid peroxidation in rat kidney and brain

homogenates. The protective effects of these herbs may be attributed to their

antioxidative and free radical scavenging activities because they were shown to be good

scavengers for superoxide anions and hydroxyl radicals. Moreover, Ng et al. (2000)

studied the antioxidative activity of a variety of flavonoids, lignans, an alkaloid, a

bisbenzyl, coumarins and terpenes isolated from Chinese herbs. They concluded that the

14 aromatic hydroxyl groups in the structure are very important for antioxidative effects of these compounds.

A diet rich in fruits and vegetables has been shown to decrease the susceptibility of low-density lipoproteins (LDL) to ex vivo peroxidation (Hininger et al., 1997). Fruits and vegetables are also rich sources of phenolic compounds.

One may ask- how is the yin-yang theory in traditional medicinal practice associated with oxidative stress? The relationship between yin-yang and antioxidation- oxidation of some TCMs was demonstrated by Ou et al. (2003). According to the authors, the yin-tonic herbs contained higher phenolic contents as compared to the yang- tonic herbs and thus, the former herbs showed stronger antioxidant activities than the latter yang-tonic herbs as tested with oxygen radical absorbance capacity assay (ORAC).

Therefore, Ou et al. (2003) concluded that the yin in TCM represents antioxidation process, whereas yang represents the oxidation process. Of course, ORAC activity in vitro need not necessarily bear any relation to antioxidant activity of a herb in vivo.

1.7 Reactive oxygen species in cell signaling

1.7.1 Introduction

All multicellular organisms depend on a complex network of extracellular and intracellular signals to regulate cell-to-cell communication in various physiological

15 processes including cell proliferation and cell death (Thannickal and Fanburg, 2000).

Extracellular signals usually interact with cell surface receptors to transmit signals into

the cytosol. These intracellular signals are then relayed or amplified by second

messengers, leading to the activation of transcription factors that regulate the expression

of specific genes essential for important cellular functions.

Cells possess multiple sites for ROS production (Halliwell and Gutteridge, 1999;

Thannickal and Fanburg, 2000). It is now clear that not only are ROS capable of causing

gross damage to cellular constituents such as proteins, DNA, and lipids (Halliwell and

Gutteridge, 1999), they can also act as mediators or signaling molecules in diverse

physiological processes (Burdon, 1995; Hensley et al., 2000). For examples, arachidonic

acid metabolism by lipoxygenases leading to the formation of leukotriene has been

reported to generate ROS (Lim et al., 1983). Moreover, exogenous reactive species have

been demonstrated to lead to the activation of ERK MAPK pathways that are mitogen-

and stress-activated signaling events (Stevenson et al., 1994; Guyton et al., 1996;

Milligan et al., 1998). Like 12-O-tetradecanoylphorbol 13-acetate, both radiation and

H2O2 were shown to activate the MAP kinase through the production of ROS in NIH-3T3 cells (Stevenson et al., 1994). Butylated hydroxytoluene hydroperoxide, a skin tumor promoter, was shown to stimulate a rapid and significant rise in MAPK-related proteins and its effects were abolished in the presence of N-acetyl-L-cysteine and o- phenanthroline, suggesting a metal-catalysed ROS formation event (Guyton et al., 1996).

The roles of MAPKs in cellular redox regulation have aroused great interest and may act upstream at the level of growth factor receptors.

16 As signaling mediators, ROS are present ubiquitously within all cell types; they

are easily synthesized, readily activated, highly diffusible, and are easily degraded when

a mission has been fulfilled (Gabbita et al., 2000).

1.7.2 Cellular redox state

ROS have diverse cellular targets (Halliwell and Gutteridge, 1999). The

mechanisms of action of ROS signaling can be characterized into alternations of 1)

intracellular redox state and 2) redox state of specific proteins (Thannickal and Fanburg,

2000). For the purposes of this thesis, only the first one will be briefly discussed. The redox state of a cell may be expressed as the ratio of the concentration of reducing equivalents to the concentration of oxidizing equivalents (Chance et al., 1979). The cytosol is normally maintained under reducing conditions by intracellular thiols such as glutathione (GSH) and thioredoxin (TRX). Both GSH and TRX peroxidases remove hydrogen peroxide (H2O2) and lipid peroxides in normal cells or cells under stress. The oxidized GSSG and TRX-S2 in the cells is then recycled by glutathione and thioredoxin reductase, respectively, to maintain a high ratios of GSH/ GSSG and TRX-(SH)2/ TRX-

S2. Peroxiredoxins also help in the removal of H2O2 (Halliwell and Gutteridge, 1999).

Equations (1) and (2) show the action of GSH in removing H2O2.

Glutathione peroxidase

2 GSH + H2O2 ------> 2 H2O + GSSG (1)

Glutathione reductase GSSG + NADPH + H+ ------> 2 GSH + NADP+ (2)

17 High ratios of oxidized to reduced glutathione and thioredoxin can result from a depletion of endogenous antioxidant defence components or an increase in the production of ROS due to various environmental oxidants such as ionizing and UV radiation, heat

• • shock and toxic agents. Endogenously generated ROS such as O2 ¯, H2O2, and OH as a consequence of infection, cell injury, or a dysfunction of cellular defense and detoxifying systems due to aging process can also disturb cellular redox status (Ermak and Davies,

2001). Kane et al. (1993) have demonstrated a rapid increase in ROS and lipid peroxides in cells depleted of GSH; and cells depleted of GSH show decreased proliferation capability (Das et al., 1992) but are usually still viable although highly sensitive to insults. Mayer and Nobel (1994) reported the inhibition of ROS induced activation of caspases by antioxidants such as N-acetyl-L-cysteine (NAC) added externally to augment the cellular antioxidant pool and counteract the actions of ROS. Furthermore, overexpression of endogenous antioxidant defence systems such as catalase, MnSOD and

Cu/Zn SOD can counteract the detrimental effects of reactive species-induced ROS- associated cell death (Greenlund et al., 1995; Kiningham et al., 1999). Indeed, evidence is accumulating from experiments using antioxidants, showing that ROS act upstream of many cell death events such as mitochondrial membrane depolarization, caspase activation and nuclear fragmentation (Fleury et al., 2002). Changes in cellular redox status may also result in the activation of pro-apoptotic signaling proteins such as JNK

(Saeki et al., 2002).

18 1.8 Calcium and its role in cell signaling and cell death

1.8.1 Introduction

A well-regulated intercellular and intracellular calcium (Ca2+) signaling is crucial

for both survival and death in biological organisms. In its insoluble form, Ca2+ is the

major structural constituents of bones and teeth, whereas in its soluble form, it plays

important roles such as membrane stabilizer, cofactor for proteins, electric charge carrier,

and diffusible intracellular second messenger (Carafoli, 1987). As a second messenger,

Ca2+ is involved in the process of fertilization and some of the cell cycle events at the

beginning of life and during early development, respectively (Clapham, 1995). Ca2+ is also required for the regulation of processes such as muscle contraction, exocytosis, energy metabolism, chemotaxis and synaptic plasticity during learning and memory

(Stryer, 1995; Calpham, 1995). Within cells, Ca2+ is primarily a constituent of

phospholipids, proteins, and nucleic acids or sequestered in organelles such as

endoplasmic reticulum (ER), mitochondria and nucleus; only 0.1 % of the total cellular

Ca2+ content in the cytosol is in its free form (Kass and Orrenius, 1999).

1.8.2 Cellular calcium homeostasis

The regulation of Ca2+ homeostasis within a cell is achieved by a close

coordination of a number of channels and pumps on the plasma membrane and

organelles, and intracellular Ca2+-binding proteins (Fig. 1.2).

19

Fig. 1.2. Schematic representation of Ca2+ regulation in cells. Ca2+ influx through Ca2+ channels (1) or via the sodium-calcium exchanger (2). Once inside the cell, Ca2+ can be removed to the extracellular environment, primarily by the plasma membrane Ca2+- ATPase (3) but also by the sodium-calcium exchanger (2). In addition, Ca2+ can interact with Ca2+-binding proteins or become sequestered into the ER, M and N. The ER contains the SERCA, which translocates Ca2+ from the cytosol into the ER lumen (4), whereas mitochondria take up Ca2+ through the uniporter driven by mitochondrial potential (5). Abbreviations: ER, endoplasmic reticulum; M, mitochondria; N, nucleus, SERCA, sarcoplasmic-endoplasmic reticulum Ca2+-ATPase. (Taken from Kass and Orrenius, 1990)

Cytosolic Ca2+ concentration is maintained at a narrow range of around 0.02 to

0.1 µM, whereas extracellular Ca2+ concentration is in the millimolar range (Clapham,

1995). Ca2+ can gain access into the cytosol through receptor-operated, voltage-gated and store-operated Ca2+ channels, sodium-calcium exchangers and non-selective cation

channels (Kass and Orrenius, 1990; Lewis, 2001). To avoid being flooded by Ca2+ from

20 the extracellular environment, the plasma membrane is impermeable to Ca2+ except through these channels, where opening is highly regulated. In addition, the cell is equipped with plasma membrane Ca2+-ATPase pumps (PMCAs) that remove excess Ca2+ from the cytosol. Additionally, the sodium-calcium exchanger can operate in a reverse mode to remove Ca2+ in exchange for sodium under ischemia/ reperfusion conditions.

Furthermore, Ca2+ can interact with proteins such as calmodulin, calsequestrin and

calreticulin or become sequestered into ER, mitochondria, or nucleus for storage. This

also serves to avoid prolonged elevations of cytosolic Ca2+ which may cause irreversible damage to cells as occurs during cardiac or cerebral ischaemia (Trump and Berezesky,

1995).

1.8.3 Roles of endoplasmic reticulum in cytosolic calcium homeostasis

The ER is the largest store of Ca2+ in a cell and its internal Ca2+ concentration

may be in the millimolar range (Montero et al., 1995). The ER is also involved in the folding, modifying and sorting of newly synthesized proteins (Orrenius et al., 2003). Ca2+ levels in the ER is regulated by sarcoplasmic reticulum Ca2+-ATPase-type pumps

(SERCAs) and ligand-operated receptors such as inositol 1,4,5-triphosphate- (InsP3Rs) and ryanodine- (RYRs) receptors. SERCA translocates Ca2+ into the ER from the cytosol

2+ whereas InsP3R and RYR open channels and release Ca back to the cytosol after

2+ activation by ligand InsP3 and ryanodine, respectively. Overloading and depletion of Ca of the ER Ca2+ pool in response to various stimuli can affect the functions of ER in protein folding and hence, generating ER stress that may lead to cell elimination by apoptosis (Orrenius et al., 2003).

21 1.8.4 Role of mitochondria in cytosolic calcium homeostasis

The mitochondria possess a high capacity but low affinity for Ca2+ (Krieger and

Duchen, 2002). Under physiological conditions, mitochondrial Ca2+ concentration

2+ 2+ 2+ ([Ca ]m) is in equilibrium with the cytosolic Ca concentration ([Ca ]i) (Rizzuto et al.,

1994; Somlyo et al., 1985). Mitochondria take up Ca2+ from the cytosol mainly through the uniporter and the rapid mode (RaM) of Ca2+ import pathway and release through the export pathways, which can either be Na+-dependent or -independent depending on cell types (Gunter et al., 2000). Release of mitochondrial Ca2+ can also occur by a reversal in operation of the uniporter and transient opening of the mitochondrial permeability transition pore (Kass and Orrenius, 1999). It is believed that mitochondria and ER work together to monitor and control Ca2+ homeostasis of cells (Rizzuto et al., 1998).

Mitochondria have been shown to serve as a buffering system removing Ca2+ from the

2+ cytosol when [Ca ]i rises above 500 nM (Duchen, 1999), suggesting that the mitochondria take up Ca2+ only when the ER can no longer cope with the prolonged

2+ 2+ elevation of [Ca ]i. The uptake of Ca by mitochondria stimulates matrix

dehydrogenases that are involved in NADH production and hence, mitochondrial energy

metabolism (Griffiths et al., 1997; Brandes and Bers, 2002).

1.8.5 Calcium and oxidative stress

2+ A rapid rise in [Ca ]i is a common phenomenon in various cell types when they are exposed to reactive species (ROS/ RNS/ RCS) (Roveri et al., 1992; Wang and Joseph,

2+ 2+ 2000). This increase in [Ca ]i can be a result of the influx of extracellular Ca or release from internal stores (i.e. ER or mitochondria), depending on the type and dosage of

22 reactive species used in the systems. For examples, low levels of hydrogen peroxide

2+ 2+ induced transient increase in [Ca ]i, whereas at higher levels, sustained elevated [Ca ]i was observed (Hampton and Orrenius, 1997). The release of Ca2+ from the ER due to the

• action of inositol 1,4,5-triphosphate (InsP3) can be enhanced by O2 ¯ generated by

xanthine oxidase/ hypoxanthine system in endothelial cells (Graier et al., 1998).

• Moreover, O2 ¯ and H2O2 can inhibit the action of SERCA pumps leading to the accumulation of Ca2+ in the cytosol (Grover and Samson, 1988; Grover et al., 1992).

Although mitochondria can serve as a buffering system in removing Ca2+ from the

cytosol during a prolonged Ca2+ influx (Duchen, 1999), mitochondrial Ca2+ overload may

in turn increase the rate of mitochondrial ROS generation (Dugan et al., 1995). In

addition, it has been shown that caspase 3 activated in apoptotic cells can perturb Ca2+

+ 2+ homeostasis by cleaving InsP3 receptor (type I), PMCA and Na - Ca exchanger leading to a loss of function. As a consequence, more intense oxidative stress was generated in the apoptotic cells due to Ca2+ overloading (Hirota et al., 1999; Schwab et al., 2002).

1.8.6 Calcium and cytotoxicity

The detrimental effects of a prolonged elevation of Ca2+ on cell survival were first observed in the 1980s (Jewell et al., 1982; Nicotera et al., 1986). From these works, the perturbation of Ca2+ homeostasis was caused by the inhibition of Ca2+ transport

mechanisms including the PMCAs and SERCAs by cytotoxic agents such as

thapsigargin, cyclopiazonic acid, and 2,5-di(tert-butyl)hydroquinone that act primarily on

the ER (Kass et al., 1989; Suzuki et al., 1992). More direct evidence of cell death due to

2+ 2+ an elevated [Ca ]i was shown when Ca ionophore A23187 was found to induce

23 apoptosis in thymocytes (McConkey et al., 1989); thymocytes apoptosis induced by thapsigargin was abrogated in Ca2+-free medium or with the used of Ca2+ chelators such

as EGTA or BAPTA/ AM (Jiang et al., 1994; Macho et al., 2004). The mechanism of

tributyltin-induced thymocyte apoptosis has been shown to occur in a manner similar to

that of thapsigargin (Aw et al., 1990). The consequent activation of proteases (i.e.

2+ calpains and caspases), phospholipases, and endonucleases due to the elevated [Ca ]i can lead ultimately to cell death by both apoptosis or necrosis (Kass and Orrenius, 1999).

However, the importance of these proteins in cell death signaling is unclear. For examples, the contribution of calpains to apoptosis appears to be limited to certain cell types (Squier and Cohen, 1997; Nath et al., 1996; Jordan et al., 1997) and both calpain and caspases can cleave the same proteins leading to activation or inactivation of their function. These proteins may work together to accelearate the cell death events (Orrenius et al., 2003).

Cellular effects of Ca2+ are complex; it affects multiple targets (e.g. calcineurin, nitric oxide synthases, endonuclease, phospholipases, transglutaminases, and proteases

(caspase, calpain and serine)). Ca2+ trafficking, both entry from extracellular

environment and release from intracellular organelles or translocation from cytosol to

external space or intracellular stores, in response to various endogenous and exogenous

stimuli, is also complex. The effect of reactive species on Ca2+ signaling depends on the type of reactive species, their dosages, and the duration of exposure. The cross-talk between Ca2+-activated cellular signals add complexity to studies of its roles in cell death.

24 1.9 Mitochondria

1.9.1 Mitochondrial structure and function

Mitochondria are central to the maintenance of cell function and viability. They

are the “power house” of all the eukaryotic cells, except red blood cells and terminal

keratinocytes (Fuller and Shields, 1998). It is in the mitochondria where energy is

generated in the form of adenosine triphosphate (ATP) for cellular activities. In addition,

mitochondria have been recognized to play an important role in Ca2+ homeostasis and cellular Ca2+ signaling (Thomas et al., 1996). Mitochondria vary considerably in size (~

0.5- 2 µm) and shape, and they are flexible and mobile and are usually found at sites in the cells where energy consumption is high.

Based on the images obtained from an electron microscopy (Frey et al., 2002), the mitochondrion is bound by a smooth outer membrane and an inner membrane. The outer mitochondrial membrane contains multiple copies of porin, which are channels for passage of molecules with a molecular weight of approximately 5,000 Da. The inner membrane has a different structure; it is folded into numerous cristae, which greatly increases its surface area. The lumen surrounded by the inner mitochondrial compartment is called the matrix, which contains various soluble enzymes of the tricarboxylic acid cycle, as well as substrates, cofactors and inorganic ions. Moreover, the inner membrane also contains proteins that are responsible for the generation of ATP from the electron transfer chain by oxidative phosphorylation. Electrons are carried from complexes I and

II to complex III by coenzyme Q, and from complex III to complex IV by the peripheral

25 membrane protein cytochrome c. The inner membrane also contains the F1F0-ATP

synthase, and other transport proteins that regulate passage of metabolites such as ATP,

ADP, pyruvate, Ca2+ and phosphate.

The inner membrane is also highly resistant to proton leakage, leading to the buildup of a membrane potential estimated at ~150-200 mV negative with respect to the cyotsol (Gottlieb, 2000); the F1F0-ATP synthase uses the proton gradient generated from the flow of the protons from the intermembrane space into the matrix to generate ATP.

1.9.2 Mitochondrial reactive oxygen species

In the mitochondrial respiratory chain, the majority of electrons are involved in

the reduction of molecular oxygen in the terminal cytochrome oxidase (complex IV),

forming water molecules (H2O). However, some of the electrons can escape from the respiratory chain and perform one-electron reduction of oxygen, forming superoxide

• • anion (O2 ¯), and H2O2 by superoxide dismutation. Hydroxyl radical (OH ) can be

2+ generated from the cleavage of the O-O bond in H2O2 by Fe in Fenton reaction

• (Halliwell and Gutteridge, 1999) and the reaction between O2 ¯ and nitric oxide produces

ONOO¯ (Beckman et al., 1994).

Mitochondrial ROS can cause irreversible inhibition of mitochondrial respiration

and oxidative damage to mitochondrial components, leading to mitochondrial

dysfunction and eventually cell death (Halliwell, 2001). Mitochondrial membrane

proteins and polyunsaturated fatty acids (PUFA) components are the major targets of

26 mitochondrial ROS (Fleury et al., 2002). ROS affect cysteine residues (sulfhydryl

groups), causing intramolecular cross-linkings and formation of protein aggregates. OH• can attack PUFAs to initiate lipid peroxidation which is then propagated by the formation of peroxyl- and alkoxyl radicals. As a result, mitochondrial outer membrane permeabilization occurs and the release of mitochondrial matrix proteins that orchestrate programmed cell death.

In fact, mitochondria are widely thought to be the major source of oxygen free radicals in all cells except macrophages and neutrophils (Krieger and Duchen, 2002).

Other sources of ROS production include NADPH cytochrome P450 reductase of endoplasmic reticulum, NAD(P)H oxidase within the plasma membrane, xanthine oxidase, lipoxygenase, and cyclooxygenase (Thannickal and Fanburg, 2000; Halliwell and Gutteridge, 1999; Gabbita et al., 2000).

1.10 Cell death

1.10.1 Introduction

The study of cell death mechanisms is one of the most popular areas in

biomedical research. The recognition of cells resistant to die under unfavorable and stressful conditions such as exposure to anticancer agents or irradiation marked the importance of cell death in both tumorigenesis and the development of resistance to anticancer drug treatment (Hanahan and Weinberg, 2000, Green and Evan, 2002).

27 Understanding the mechanisms of cell death has implications for the design of novel

therapeutic agents for diseases such as cancer and neurodegeneration.

1.10.2 Differences between apoptosis and necrosis

Apoptosis is also known as a “cell suicide” programme that plays multiple and

important roles in embryonic development, immune system function and the maintenance of tissue homeostasis in multicellular organisms (Ellis et al., 1991). Apoptosis in mammalian cells is dependent on the activation of a family of cysteine proteases with aspartate-specificity known as the caspases that are expressed as inactive zymogens

(procaspases) in the cytoplasm (Alnemri et al., 1996). Over a dozen caspases have been identified in humans and the activation of procaspases to active caspases can be brought about by oligomerization (e.g. initiator caspase such as caspase-8) or cleavage after the aspartic acid residues (e.g. effector caspases such as caspase-3, -6 and -7) (Earnshaw et al., 1999; Hengartner, 2000).

Some of the morphological changes that have been reported from cells undergoing apoptosis include nuclear/ cytoplasmic condensation, membrane protrusions, fragmentation of nuclear contents and the resulting formation of “apoptotic bodies” (Kerr et al., 1972). Necrotic cells, however, show different morphological changes from apoptotic cells. Irreversible swelling of cells is usually the first evidence of cells undergoing necrosis, and then the plasma membrane collapses and the necrotic cells rapidly lyse (Proskuryakov et al., 2003). The consequent release of noxious intracellular constituents into the extracellular space may induce an inflammatory response (Orrenius

28 et al., 2003). Apoptotic cells are characterized by externalization of phosphatidylserine

residues in the plasma membrane, activation of caspases, production of ROS, reduction in

the mitochondrial transmembrane potential, and DNA degradation into internucleosomal

fragments, forming a DNA ladder (~ 200 bp) on electrophoresis (Martin et al., 1995;

Zamzami et al., 1996a, 1996b; Wyllie et al., 1984). However, some of these biochemical markers such as the activation of caspases and oligonucleosomal DNA fragmentation in apoptotic cells may or may not present in necrotic cells. Nevertheless, both apoptotic and necrotic cells will be digested by neighboring cells or phagocytes after showing the “eat

me” message (Henson et al., 2001; Shacter et al., 2000). It is externalization of phosphatidylserine on the plasma membrane of apoptotic cells and destruction of plasma membrane of necrotic cells that label them for removal by phagocytes. Leist et al. (1997) has demonstrated the importance of cellular ATP for a successful orchestration of the apoptotic cascades and suggested that cellular ATP contents may be critical in deciding the fate of cells.

Necrosis may seem unfavorable as compared to apoptosis due to a series of unregulated events occurring after extensive insult to the cells. However, necrosis may be important in the processes such as the negative selection of lymphocytes (Holler et al.,

2000; Doerfler et al., 2000), tissue renewal (Barkla and Gibson, 1999) and embryogenesis (Chautan et al., 1999). Reports that have shown the involvement of necrotic cell death during pathological processes include, to name a few, cultured human chondrocytes undergo necrotic death after treatment with cytokines and lipopolysaccharide (Blanco et al., 1995); 4-hydroxynonenal, a product of lipid

29 peroxidation, induced necrotic cell death in neuronal cells (Kruman and Mattson, 1999); tumor necrosis factor-mediated liver injury (Kunstel et al., 1999); pathogen invasion such as various viruses, bacteria, and protozoa (Ehlers et al., 1999); H2O2 can cause both apoptosis and necrosis depending on the dosage and the type of cells used in an experiment (Orrenius and Hampton, 1997; Palomba et al., 1999); and radiation- or toxic agents-induced DNA damage could indirectly cause necrosis by ATP depletion (Ha and

Snyder, 1999).

It has become clear that the morphological and biochemical extremes of apoptosis and necrosis are not sufficient to characterize all types of cell death programmes. Several reports have shown that the anti-apoptotic protein bcl-2 protects cells not only from apoptosis but also from necrosis (Shimizu et al., 1996). Moreover, several apoptotic features such as plasma membrane blebbling, phosphatidylserine exposure and mitochondrial damage may also occur in necrotic cells (Orrenius et al., 2003). Based on the morphology of cells undergoing cell death, Leist and Jäättelä (2001) classified them into apoptosis, apoptosis-like, and necrosis-like PCD. Sperandio et al. (2000) have described an alternative non-apoptotic form of programmed cell death that named as

“paraptosis”. They also summarized some of the features of apoptosis, necrosis, and paraptosis as shown in Table 1.4.

30 Table 1.4 Comparison of apoptosis, necrosis and paraptosis

Apoptosis Necrosis Paraptosis

Morphology Nuclear fragmentation + Sometimes Chromatin condensation + ± Apoptotic bodies + Cytoplasmic vacuolation + + Mitochondrial swelling Sometimes + Late Genomic effect TUNEL + Usually Internucleosomal DNA + Sometimes fragmentation Caspase activity DEVD-cleaving activity + Caspase-3 processing + PARP cleavage + (85-kDa + (50- to 62-kDa fragment) fragments) Inhibition by: zVAD.fmk + BAF + p35 + xiap + Bcl-xL + Usually Actinomycin D Sometimes + Cycloheximide Sometimes +

DEVD, Asp-Glu-Val-Asp; PARP, poly(ADP-ribose) polymerase. Modified from Sperandio et al. (2000)

In general, apoptotic cell death can be triggered by extrinsic (receptor-mediated)

or intrinsic (mitochondria-mediated) signaling pathways. Extrinsic pathway is activated

by ligation of death receptors on cell surface such as Fas and tumor necrosis factor

leading to the formation of the death-inducing signaling complex (DISC), which activates

31 pro-caspase 8. Active caspase 8 activates pro-caspase 3 in type I cells, which results in

the cleavage of target proteins. In type II cells, caspase 8 cleaves Bid, which in turn,

induces the translocation of Bax or Bak into the mitochondrial outer membrane. This is

followed by the release of cytochrome c, which forms a cytosolic apoptosome complex

comprising apoptosis inducing factor-1 and pro-caspase 9, in the presence of dATP. For

intrinsic pathway, apoptotic stimuli function directly or indirectly on the mitochondria, resulting in the fall of mitochondrial membrane potential and release of cytochrome c that

forms the apoptosome complex. This pathway may be mediated by apoptosis inducing

factor and endonuclease G in the absence of caspase activation (Green and Reed, 1998;

Orrenius et al., 2003). Figure 1.3 shows the extrinsic and intrinsic signaling pathways of

cell death (Orrenius et al., 2003).

32

Fig. 1.3. Extrinsic and intrinsic pathways of cell death in mammalian cells. The extrinsic pathway is induced by the formation of DISC after ligation of receptor specific ligand. The DISC in turn activates procaspase 8. In type I cells, caspase 8 activates procaspase 3 directly. In type II cells, Bcl-2 proteins such as Bid, Bax and Bak are activated and translocated to the mitochondria. This is followed by the formation of an apoptosome consisting of the released cytochrome c, Apaf-1, and procaspase 9 in the presence of dATP. Intrinsic pathway, death stimuli function directly or indirectly on the mitochondria, resulting in the formation of an apoptosome complex. This pathway is regulated by Bcl-2 family proteins, inhibitor of apoptosis proteins (IAPs) and IAPs’ inhibitors smad and Omi. Putative HLA-DR-associated protein (PHAP) facilitates the activation of caspases by the apoptosome while prothymosin-α (Pro-T) negatively regulate caspase 9 activation by inhibiting the formation of the apoptosome. This pathway might also operate by the release of apoptosis inducing factor (AIF) and endonuclease G (EndoG) from the mitochondria. (Modified from Orrenius et al., 2003)

33 1.10.3 Mitochondrial permeability transition and cell death

Mitochondria are critical in the control of cell death. Cell death involving the

mitochondrial transmembrane potential (∆ψm) usually starts with the accumulation of effector molecules (e.g. Ca2+) into the mitochondria. Overloading of these effector

molecules leads to a decrease in ∆ψm due to the opening of the mitochondrial permeability transition (MPT) pore and eventually the release of proteins (i.e. cytochrome c, smad, adenylate kinase 2, and apoptosis-inducing factor) from the intermembrane space into the cytoplasm that are capable of activating nucleases and proteases (Kroemer and Reed, 2000). Figure 1.4 shows a schematic diagram for the activation of caspase by mitochondria leading to cell death. Clofibrate (a peroxisome proliferator), is an example of an agent that leads to mitochondrial dysfunction and cell death (Qu et al., 2001).

The reduction of ∆ψm can be determined indirectly using lipophilic cationic fluorochromes such as 3,3’dihexyloxacarbocyanine iodide (DiOC6(3)) or 5,5’,6,6’- tetrachloro-1,1’,3,3’-tetraethylbenzimidazolylcarbocyanine iodide (JC-1), which accumulate in the mitochondrial matrix due to the ∆ψm. A reduction in the fluorescence intensity of the cationic dye (DiOC6(3)) or a shift in the emission spectrum from red to green (JC-1), as measured by cytofluorometry or confocal microscopy, is then interpreted as an indication of the loss of ∆ψm (Bernardi et al., 1999).

34

Fig. 1.4. Model for caspase activation by mitochondria. Multiple stimuli such as Bax, oxidants, Ca2+ overload, and active caspases can trigger mitochondria to release caspase- activating proteins, among which are cytochrome c (solid circles) and possibly other proteins such as AIF and intramitochondrial caspases (open circles). Cells in which

mitochondria have ruptured may die by necrosis due to the collapse of ∆ψm, production of ROS, and declining ATP production. (Taken from Green and Reed, 1998)

35 The MPT pore is a large proteinaceous complex that is voltage sensitive. The

structure and composition of the MPT pore remain unknown, but its constituents include

both the inner membrane proteins, such as adenine nucleotide translocator (ANT), and

the outer membrane protein porin which is a voltage-dependent anion channel (VDAC),

and ANT-interacting mitochondrial cyclophilin D (a target of cyclosporin A). However,

it is believed that the MPT pore complex also contains the peripheral benzodiazepine

receptor, creatine kinase, hexokinase II, Bax and Bcl-2 proteins (Crompton 1999;

Shimizu et al., 1999).

Based on in vitro experiments on purified mitochondria or proteins reconstituted into artificial membranes, there are at least four models that can be used to explain the involvement of MPT pore in the reduction of ∆ψm.

In the first model, the influx of pro-apoptotic molecules such as Ca2+ and agents that oxidize –SH groups cause ANT to form a non-specific pore allowing diffusion of ions within the matrix and intermembrane space of mitochondria. Osmotic matrix swelling due to volume dysregulation of the mitochondria permeabilizes or ruptures the inner- and outer-mitochondrial membrane because the inner-mitochondrial membrane with its folded cristae has a larger surface area than the outer-mitochondrial membrane.

As a result, ∆ψm dissipation and uncoupling of the mitochondrial respiratory chain occur

(Crompton, 1999). In the second model, the VDAC accounts for the primary permeabilization of the outer-mitochondrial membrane leading to the release of intermembrane space proteins such as cytochrome c and apoptosis inducing factor. The

36 inner mitochondrial membrane may remain intact. This event is enhanced in the presence

of Bax and inhibited by Bcl-2 in vitro (Shimizu et al., 1999). However, Bax is unlikely be part of the VDAC pore complex because pore formation is inhibited by Koenig’s polyanion, a VDAC inhibitor. In the third model, oligomeric bax is the primary pore former in the outer-mitochondria membrane (Eskes et al., 1998; Eskes et al., 2000) allowing translocation of proteins independently of the VDAC. The activity of the autonomous protein-translocating channel may be enhanced by Bid or t-Bid. In the last model, the VDAC controls ∆ψm through its functional and physical contact with ANT by

a different mechanism. VADC is a voltage-sensitive channel and hence, any apoptosis-

associated decrease in inner-mitochondrial membrane potential would result in the

opening of the VDAC.

1.10.4 Calcium overload and mitochondrial permeability transition

Although the molecular details of MPT pore are still not well defined, the

conditions that trigger MPT pore opening have been studied extensively, among which

are oxidative stress, overloading of mitochondria with Ca2+, oxidation of pyridine nucleotide and thiol groups, alkalinization, and low transmembrane potential (Van den

Dobbelsteen et al., 1996; Cai and Jones, 1998; Kass and Orrenius, 1999). Ca2+ ionophores, thapsigargin, neurotoxins, and peroxynitrite are some of the agents that trigger apoptosis in various cell types, involving Ca2+-mediated MPT (Korge and Weiss,

1999; Akao et al., 2002; Schweizer and Richter, 1996). Ruthenium red, an inhibitor of the uniporter and rapid mode of mitochondrial Ca2+ uptake pathway, inhibits apoptosis in

L929 fibroblasts, suggesting that mitochondrial Ca2+ influx is associated with apoptosis

37 (Van den Dobbelsteen et al., 1996). Cyclosporin A, an inhibitor of cyclophilin D, is able

to inhibit the opening of MPT pore and thus the release of mitochondrial pro-apoptotic

proteins and low-molecular mass compounds of less than 1.5 kDa, including Ca2+ (Weis

et al., 1994). In fact, the Ca2+-cardiolipin interaction might be an early and important step in Ca2+-mediated opening of MPT pore and the release of cytochrome c into the cytosol

(Orrenius et al., 2003). This is then followed by the formation of an apoptosome complex

consisting of cytochrome c, Apaf-1 and pro-caspase 9 in the presence of dATP for the

induction of apoptosis (Ott et al., 2002).

1.10.5 Bcl-2 proteins and cell death

Evidence is accumulating that pro-apoptotic Bcl-2 family proteins such as Bax,

Bak, and t-Bid translocate to the mitochondria in response to apoptotic stimuli. Both Bax

and Bak are capable of forming pores on the outer-mitochondrial membrane as dimers or

oligomers, leading to permeabilization of outer-mitochondrial membrane and the release

of pro-apoptotic mitochondrial proteins. These events may be stimulated by Bid cleavage

product, t-Bid (Eskes et al., 2000). Conversely, anti-apoptotic Bcl-2 proteins such as Bcl-

2 and Bcl-XL serve to protect against the opening of the MPT pore of the mitochondria by binding to pro-apoptotic proteins (Antonsson, 1997).

However, more experiments are needed to define the relationship between mitochondria and cell death. For examples, Yang et al. (1997) showed the release of cytochrome c in the absence of mitochondria depolarization; and the effects of inhibitors

38 on MPT pore opening, including cyclosporin A (which bind cyclophilin D associated

with the ANT) and bongkrekic acid (which inhibits the ANT), appear to be limited to

certain apoptotic processes (Zamzami et al., 1996a, Andreyev and Fiskum, 1999).

1.11 Plasma membrane redox system

Plasma membrane redox systems (PMRS) appears ubiquitous in living cells

(Medina, 1998) and have been proposed by some to play a vital role in the regulation of

the internal redox equilibrium in response to external stimuli (Crane et al., 1990). One of

the functions of the PMRS may be the oxidation of electron donors such as NADH and/

or NADPH, and the transfer of the resulting electrons to external electron acceptors via

coenzyme Q10 (CoQ) (Crane et al., 1990; Lawen and Baker, 2000; Morré and Morré,

2003; de Grey, 2003). To do that the PMRS has to be in a privileged position to interconnect the intra- and extracellular components. Some of these extracellular electron acceptors include molecular oxygen, ferricyanide and ascorbate (Lawen and Baker,

2000). The term “plasma membrane NADH reductase/ or NADH-oxidoreductase

(PMOR)” is also found in the literature and both PMRS and PMOR are generally referring to the same system.

39 1.12 Aims of this study

There is a lot of interest in natural products as the quest for more active and novel

antioxidant ingredients continues. TCM make-up a significant proportion of the total

consumption of herbal medicine around the world. They are rich sources of phenolic

compounds and these phenolic compounds may be related to their alleged antioxidant

effects. Therefore, we first tested the hypothesis that TCM (available from local

medicinal shops) possess strong antioxidant activities as characterize using various

established and reliable antioxidant-based assays. We also attempted to identify the

constituents that may be responsible for the antioxidant activities from Cratoxylum cochinchinense which outperformed other TCM extracts. C. cochinchinense is a small genus of Southeast Asian trees belonging to the Guttiferae family (Bennett and Lee,

1989). It is used in traditional medicine for many purposes (Bremness, 1994). However, many health claims pertaining to C. cochinchinense are not supported by scientific studies and are myths that passed down from previous generations. Triterpenoids, tocotrienols and xanthones are some of the constituents in the bark of C. cochinchinense

(Bennett et al., 1993; Sia et al., 1995, Nguyen and Harrison, 1999). However, there has been limited research on the biological properties of C. cochinchinense extract and its isolated constituents. Therefore, we examined the effects of C. cochinchinense extract

(YCT), which contains largely mangiferin, on a range of cell types to test whether a potent antioxidant also posseses cytoprotective effect. We are also interested to find out the modes of action of YCT extract-induced cell death in Jurkat T cells.

40 CHAPTER 2

EXPERIMENTAL PROCEDURES

2.1 Materials (parts extracted from and code of Traditional Chinese Medicines are in parenthesis)

Traditional Chinese Medicines (TCM) extracts used in this study include:

Anemarrhena asphodelodies Bunge (rhizome, AA), Angelica sinensis (root, AS),

Cistanche deserticola Y. C. Ma (stem, RS), Concha margaritifera Argyi (leaves, J),

Cortex magnoliae officinalis (bark, D), Cratoxylum cochinchinense (leaves, WN),

Curculigo orchioides Gaertn (rhizome, SY), Cuscuta chinensis Lam (seed, CC),

Dioscorea opposite Thumb (rhizome, DA), Epimedium sagittatum (leaf, ES), Flow

carthami (flower, O), Folium artennsiae (leaves, I), Frutus alpinae oxyphyllae (fruit,

YZ50), Frutus annantii (fruit, F), Gentiana scabra (root, LT), Glycyrrhiza uralensis Fish

(bark, E), Herba schizonepetae (root, A), Lycium chinense (fruit, LB), Morinda officinalis How (root, MO), Polygonatum kingianum (rhizome, PK), Polygonum nultiflorum Thumb (root, PM), Psoralea corylifolia L (fruit, PC), Radix achyranthis bidentatae (root, M), Radix astragali (root, B), Radix dipsaci (root, X), Radix paeoniae alba (root, H), Radix rehmanniae (root, HZ), Rhizoma atraetylodis marcocephalae

(rhizome, K), Rhizoma chuanxiong (rhizome, G), Rhizoma corydali (rhizome, L),

Rhizoma cyperi (rhizome, N), Rhizoma seu radix notoperygi (rhizome, C), and

Schisandra chinensis (fruit, MS).

41 All reagents were purchased from Sigma-Aldrich (St. Louis, MO, USA) unless

otherwise stated. Foetal bovine serum (FBS) was from Gibco-BRL (Gaithersburg, MD,

USA). Annexin V-FITC Apoptosis Detection Kit I and K2-EDTA coated vacutainer tubes were from BD Biosciences Pharmingen (Franklin Lakes, NJ, USA). Baicalein

monohydrate (BM), benzenemethanol (REV5901), 9,12-Octadecadiynoic acid (ODYA),

oleyloxyethyl phosphorylcholine, methyl arachidonyl fluorophosphonate (MAFP), and

bromoenol lactone (BL) were from Cayman Chemical (Ann Abor, MI, USA).

Dihydroethidium (DHE), fluo-3-acetoxymethyl ester (Fluo3/ AM), Caspase 9 substrate

(Ac-LEHD-AFC), anti-procaspase 3 and 7 antibodies were from Calbiochem (La Jolla,

CA, USA). Cytochrome c monoclonal antibody was from Chemicon (Temecula, CA,

USA). Succinimidyl ester (C11-Bodipy581/591), 9,11,13,15-octadecatetraenoic acid (cis-

parinaric acid), rhod-2/ acetoxymethyl ester (Rhod-2/ AM), tetramethylrhodamine methyl

ester (TMRM), 5,5',6,6'-tetrachloro-1,1',3,3'-tetraethylbenzimidazolylcarbocyanine iodide

(JC-1), 3,6-diamino-9-(2-(methoxycarbonyl) phenyl (Rhod 123), pluronic acid F-127,

potassium-binding benzofuran isophthalate acetoxymethyl ester (PBFI/ AM) and sodium-

binding benzofuran isophthalate acetoxymethyl ester (SBFI/ AM) were obtained from

Molecular Probes Inc. (Eugene, OR, USA). Caspase 3 substrate (Ac-DEVD-AFC) and

DC Protein assay kit were obtained from Bio-Rad (Hercules, CA, USA). Caspase-3

inhibitor (Z-DEVD-FMK) and -9 inhibitor (Z-LEHD-FMK) were obtained from BD

Biosciences (Franklin Lakes, NJ, USA). Protease inhibitor cocktail tablets were obtained

from Roche (Basel, Switzerland). BCA Protein Assay Kit and Mitochondria isolation kit

(Cat No. 89874) were purchased from Pierce (Rockford, IL, USA).

42 2.2 Methods

2.2.1 Extract preparation

TCM were purchased from local medicinal stores except WN, which was supplied

by Miss YAP Sook Peng (Dept. of Obstetrics & Gynecology, Faculty of Medicine,

National University of Singapore, Singapore). TCM extracts were prepared as described

in Peng et al. (2003) with slight modification. Dried herbs were blended to a fine powder

and extracted 3 times (30 min each with continuous stirring at room temperature, 25 °C) with 80% (v/v) methanol (herbs: methanol = 1: 50, w/v). No heating step was included.

Filtrates were combined and concentrated using a rotary evaporator. The concentrate was then partitioned with hexane (ratio 1: 3, v/v) to remove all the chlorophyll and lipophilic substances, followed by freeze-drying. The freeze-dried TCM powder was kept at −20

°C. Fresh samples were completely dissolved in 80% ethanol (v/v) for each assay. With the exception of the DPPH• reduction assay, final concentrations of ethanol in all the assays were 0.4% (v/v).

2.2.2 ABTS assay

This was carried out as described by Re et al. (1999). 2,2’-azinobis[3-

ethylbenzothiazoline-6-sulphonate] (ABTS) in water (7 mM final concentration) was

oxidized using potassium persulphate (2.45 mM final concentration) for at least 12 h in

the dark. The ABTS •+ solution was diluted to an absorbance of 0.8 ± 0.05 at 734 nm

(Beckman UV-VIS spectrophotometer model DU640B) with phosphate buffered saline

(PBS) (10 mM, pH 7.4). Absorbance was measured 3 min after initial mixing of TCM

extracts of different concentrations or Trolox standard with 1 ml of ABTS •+ solution.

43 Trolox was used as a reference standard. Antioxidant properties of TCM extracts were

expressed as Trolox equivalent antioxidant capacity (TEAC), calculated from the gradient of the plot of the percentage inhibition of absorbance versus concentration of the

TCM extract (at least 3 different concentrations tested in the assay giving a linear response) was divided by the gradient of the plot for Trolox.

2.2.3 Ascorbate-iron induced lipid peroxidation

Peroxidation of bovine brain extract was performed as described by Gutteridge and Halliwell (1990). Briefly, bovine brain extract (BBE, 100 mg) was dissolved with phosphate buffered saline (PBS, pH 7.4, 20 ml), and sonicated in an ice bath until dissolved. The BBE (0.2 ml) was pre-incubated with TCM extracts in the presence of

PBS buffer (0.5 ml) and FeCl3 (1 mM, 0.1 ml). Lipid peroxidation was started by adding ascorbate (1 mM, 0.1 ml) and the mixture was then incubated for 1 h at 37 °C. The reaction was stopped by adding butylated hydroxytoluene (BHT, 2% (w/v) in 95% (v/v) ethanol, 50 µl), followed by addition of trichloroacetic acid (TCA, 2.8% (w/v), 1 ml) and

2−thiobarbituric acid (TBA, 1% (w/v), 1 ml). The mixture was heated at 80 °C for 20 min in a water bath. The (TBA)2-MDA chromogen formed was measured at 532 nm after extraction into 1-butanol (2 ml) using a SpectraMax190 microplate reader (Molecular

Devices, Sunnyvale, CA, USA). Control sample contained 80% ethanol (final concentration was 0.4% v/v) as carrier and results were expressed as % of control.

44 2.2.4 Total phenolic content

Total phenolic content of extracts was assessed by using Folin-Ciocalteu’s phenol

reagent as described by Singleton et al. (1999). The extracts (100 µl) were mixed with

Folin-Ciocalteu’s phenol reagent (0.2 ml), water (2 ml), and Na2CO3 (15% (w/v), 1 ml) and absorbance at 765 nm was measured 2 h after incubation at room temperature using the microplate reader specified above. Gallic acid was used as a reference standard and the total phenolics were expressed as mg/ ml gallic acid equivalents (GAE).

2.2.5 Scavenging of DPPH• (2,2-Diphenyl-1-picrylhydrazyl)

Scavenging activity was determined as described by Gao et al. (2000). DPPH

solution (200 µM in 80% ethanol, v/v) was mixed with an equal volume of extracts and

the absorbance at 520 nm was measured after 20 min at room temperature using the

microplate reader. Results were expressed as percentage of control (100%) and the

concentration of extracts required to cause a 50% decrease in the absorbance was calculated (IC50).

•− 2.2.6 Superoxide anion (O2 ) scavenging effect

O2•− was generated by the hypoxanthine/ xanthine oxidase system as described

by Halliwell (1985). Briefly, extracts were pre-incubated at room temperature with the

reaction mixture comprising Na2EDTA (15 mM, 20 µl) in buffer (50 mM KH2PO4/ KOH, pH 7.4), nitroblue tetrazolium chloride (NBT, 0.6 mM, 50 µl) and hypoxanthine (3 mM

in 50 mM KOH, 30 µl). The reaction was initiated by addition of xanthine oxidase

solution (1 U in 10 ml buffer, 50 µl). The change in absorbance of NBT due to its

45 reduction by O2•− was detected at 560 nm using the microplate reader. Concentrations of extracts that decreased NBT reduction by 50% were determined as the IC50 value. The final concentration of ethanol in the system was 0.5% (v/v) and did not affect the assay.

2.2.7 Xanthine oxidase (XO) activity

This assay was conducted as reported by Sweeney (2001). Mixtures of extracts,

Na2HPO4−KH2PO4 buffer (66.67 mM, pH 7.5, 1.3 ml) and xanthine oxidase solution

(0.28 U/ ml in 66.67 mM phosphate buffer, 0.2 ml) were incubated at 30 °C for 10 min before the reaction was initiated by adding the xanthine solution (0.15 mM in H2O, 1.5

ml). Absorbance at 295 nm was measured each minute for 10 min using the microplate

reader and the concentration of extracts needed to cause a 50% inhibition of XO activity

was determined (IC50). Allopurinol (a XO inhibitor, 10 µM) was used as a positive

control. The final concentration of ethanol in the system was 0.15% (v/v) and did not

affect the assay.

2.2.8 Iron-binding activity

Fe2+-binding activity was measured according to the method of Simada et al.

(1992). Briefly, extracts (2 ml) were mixed with hexamine buffer (10 mM, 2 ml)

containing KCl (10 mM) and FeSO4 (3 mM), and then tetramethyl murexide (TMM, 1 mM, 0.2 ml) was added to bind free iron. TMM and TMM−Fe2+ complex show absorption maxima at 530 and 485 nm respectively. The absorbance ratio A485/ A530 was measured using the SpectraMax190 microplate reader. Fe2+-binding activity was determined by comparing the values obtained with a freshly prepared standard curve of

46 2+ FeSO4. Fe is relatively unstable and therefore, all the reagents used were kept on ice and the addition of reagents to the 96-well plate were carried out on top of a bucket of ice to minimize the possible oxidation of Fe2+ to Fe3+ during the course of the experiment.

2.2.9 Non−enzymatic protein glycation

Experiments were performed as described by Vinson and Howard (1996). Briefly,

bovine serum albumin (BSA, 10 mg/ml) in phosphate buffer (50 mM, pH 7.4) containing

0.02% (w/v) sodium azide was pre-incubated with TCM extracts at various

concentrations for 30 min at room temperature (25 °C). Sodium azide was used to prevent microbial growth during the incubation period. Glucose (25 mM) and fructose

(25 mM) solutions were added to the reaction mixture and incubated at 37 °C for 2 weeks. Fluorescence was determined using a Gemini Fluorescence microplate reader

(Molecular Devices) with an excitation wavelength of 350 nm and an emission wavelength of 450 nm. Results were expressed as percentage inhibition of formation of the glycated protein.

2.2.10 Inhibition of hypochlorous acid-Induced DNA damage

This was measured as described by Whiteman et al. (1999). Hypochlorous acid

(HOCl) concentration was quantified spectrophotometrically at 290 nm before use (ε =

350 M-1 cm-1 at pH 12) (Morris, 1966). Calf thymus DNA (1.0 mg/ml) in phosphate buffer (100 mM, pH 7.4) was pre-incubated with TCM extract at various concentrations for 30 min. HOCl (500 µM) was added to the reaction mixture, followed by incubation at

37 °C for 1 h. DNA samples were then dialyzed against distilled water for 24 h.

47 DNA concentration after dialysis was measured spectrophotometrically at 260 nm

(A2601.0 = 50 µg/ ml). Aliquots of 100 µg DNA were freeze−dried after addition of respective internal standards (0.5 nmole: 6-azathymine and 2,6-diaminopurine), followed

by hydrolysis with formic acid (0.5 ml, 60% v/v) and heating at 140−160 °C for 45 min

in evacuated glass tubes. Samples were derivatized under a nitrogen atmosphere in capped hypovials after freeze−drying under reduced pressure. Derivatization solution was a mixture of ethanethiol, acetonitrile, and BSTFA (1% (v/v) TMCS) in the ratio of 1:3:16

(v/v/v) at room temperature (25 °C) for at least 2 h (Jenner et al., 1998). Oxidized DNA bases were analysed by GC-MS (Hewlett-Packard, 6890 gas chromatograph interfaced with a Hewlett-Packard Mass Selective Detector 5973).

Quantification of modified bases was achieved by applying the following formula:

Concentration (nmol/ mg of DNA) = A/ AIST x [IST] x (1/ K) where A = peak area of product, AIST = peak area of the internal standard, K= relative molar response factor for each damaged base, and [IST] = concentration of internal standard (5 nmol/ mg of DNA). K constants can be calculated from the slopes of the calibration curves prepared using known concentrations of internal standards and

authentic compounds.

48 2.2.11 Isolation, purification, and identification of active ingredients from Cratoxylum

cochinchinense (WN)

Crude extracts of WN were eluted into four fractions using 20, 30, 35, and 100%

methanol through a C18 column (Supelco, PA, USA). Each fraction was examined using

the ABTS assay for its antioxidant activities and then analyzed using HPLC−Photodiode

Array Detector (Agilent 1100 Series) fitted with a Nucleosil5 C18 column (HPLC

Technology, MC, UK, 25cm x 4.6 mm) to identify the spectra of active ingredients. The

mobile phase consisted of H3PO4 (1.5% (v/v) in water, solvent A) and acetonitrile

(solvent B). Various gradients of solvents A and B were applied at a flow rate of 1.0 ml/

min. Identification of active ingredients was carried out by comparison of retention times

(tR), absorbance spectra and mass spectra with authentic compounds. Conditions for

LC−MS (Finnigan-LCQ) were: C18 column, mobile phase: TFA (0.1% (v/v) in water) and acetonitrile (60% (v/v) in water), gradient elution with flow rate at 1.0 ml/ min, ionization source: atmospheric pressure chemical ionization.

The following assays were carried out using the semi-purified extract (YCT) dissolved in 80% (v/v) ethanol. Since the purity of YCT is below 99%, we cannot consider it as pure compound. YCT is used from now on to represent the semi-purified extract and mangiferin (MGF) for its major ingredient. MGF was dissolved in DMSO due to its poor solubility in 80% (v/v) ethanol. Other compounds (~ 10%) in YCT extract may help mangiferin to dissolve better than the high purity mangiferin from Sigma-Aldrich.

Controls with either 80% (v/v) ethanol or DMSO were included in all experiments.

49 2.2.12 Bleomycin−iron dependent DNA damage

Assay was performed as described by Aruoma (1991, 1993). Briefly, calf thymus

DNA solution (0.5 mg/ ml) was added to a phosphate buffer (KH2PO4−KOH, 30 mM, pH

7.0) containing bleomycin sulfate (Sigma, 1.5 U/ml), MgCl2.6H2O (5 mM) and

FeCl3.6H2O (25 µM). Extracts were added to initiate pro−oxidation effects, and the mixture was incubated at 37°C for 1 h. The reaction was stopped by adding ethylenediaminetetra acetic acid (EDTA, 0.1 M), and incubated at 80°C for 20 min after the addition of thiobarbituric acid (TBA, 1 % w/v) and HCl (2.8 % v/v). The (TBA)2-

MDA chromogen formed was measured at 532 nm using the SpectraMax190 microplate reader.

2.2.13 Peroxynitrite scavenging assays

The synthesis of peroxynitrite and bleaching of pyrogallol red (PR) was performed as described by Beckman (1994) and Balavoine and Geletii (1999), respectively.

2.2.13.1 Synthesis of peroxynitrite.

In brief, an acidic solution (HCl, 0.6 M) of H2O2 (0.7 M) was mixed with NaNO2

(0.6 M) on ice for one second and the reaction quenched with ice-cold NaOH (1.2 M).

Any residual H2O2 was removed by mixing with granular MnO2. The stock solution was filtered to remove granular MnO2 and then frozen overnight at -20°C. The top layer of the solution was collected for the experiment. Concentrations of stock ONOO− were determined in 1.0 M NaOH before each experiment at 302 nm using a molar absorption

50 coefficient of 1670 M-1cm-1. ONOO− solutions were kept on ice for no longer than 30 minutes before each experiment.

2.2.13.2 Assessment of pyrogallol red bleaching by peroxynitrite (ONOO−)

PR (100 µM final concentration) was dissolved in K2HPO4−KH2PO4 buffer (100

mM, pH 7.4). Compounds to be tested were added into the PR solution, with or without

the addition of sodium bicarbonate (25 mM), and incubated at room temperature for 10

min, followed by the addition of ONOO− (100 µM) and the mixture was vortexed

immediately for 10 sec. The decrease in absorbance at 542 nm was determined using a

microplate reader. Final concentrations of ethanol and DMSO were 0.5% (v/v) and had

no effect on the assay.

2.2.13.3 Measurement of tyrosine nitration

This was performed as described by Kaur and Halliwell (1994). Briefly, D,

L−tyrosine (10 mM) was prepared by dissolving the required amount in water (9.5 ml)

with KOH (10% (w/v), 250 µl) followed by phosphoric acid (5% (v/v), 250 µl). Tyrosine

solution (1 mM, 0.1 ml) together with compound (0.1 ml, YCT in 80% ethanol and MGF

in DMSO) to be tested was added to a glass test tube containing phosphate buffer (100

mM K2HPO4−KH2PO4, pH 7.4, 0.795 ml) and incubated in a water bath at 37 °C for 15 min. The effect of NaHCO3 addition (25 mM final concentration) was tested.

Peroxynitrite (200mM, 5 µl) was added and the tube was vortexed for 15 sec and incubated for a further 15 min. Final concentration of peroxynitrite in the reaction mixture was 1 mM. Measurement of 3−nitrotyrosine employed a Spherisorb 5 µm ODS2

51 C18 column (Wellington House, Cheshire, UK) with a guard column (Hibar from BDH,

Poole, UK). An isocratic elution was applied and the mobile phase was a mixture of

KH2PO4−H3PO4 (50 mM, pH 3.01) and methanol (100%) (80: 20, v/v) at a flow rate of 1

ml/ min. The UV detector was set at 274 nm. Concentrations of tyrosine and

3−nitrotyrosine were determined by comparing the values (area under the curve) obtained

with freshly prepared standard curves. Final concentrations of ethanol and DMSO were

0.5% (v/v) and had no effect on the assay.

2.2.14 Cell culture

Human foetal liver (HFL), human embryonic kidney (HEK), Madin-Darby canine

kidney (MDCK), human hepatoma (HepG2) and Chang liver cells (CL) were generous

gifts from Professor Sit Kim Ping (Dept. of Biochemistry, Faculty of Medicine, National

University of Singapore, Singapore). Human articular chondrocytes (HAC), human chondrosarcoma (HTB94), human colorectal adenocarcinoma (HT29), rat pheochromocytoma (PC12), acute promyelocytic leukemia (HL60) and T lymphocyte

(Jurkat T cells) were obtained from the American Type Culture Collection. HL60 and

Jurkat cells were cultured in RPMI 1640, others were cultured in DMEM with 1% (v/v) penicillin, 10% (v/v) FBS, 5% CO2 at 37 °C. The cells were maintained in the logarithmic growth phase by routine passage every 2-3 days.

2.2.15 Cell counting

Trypan blue stain (0.4% w/v in phosphate buffered-solution) and the hemocytometer were used to determine total cell counts and viable cell number to

52 facilitate cell seeding. The principle of staining is based on the ability of live cells to

exclude dye staining whereas dead cells will be stained. Briefly, a cell suspension (200

µl) was mixed with 50 µl of trypan blue solution, and then a small volume (~5 µl) of

trypan blue-cell suspension mixture was transferred to both chambers of the

hemocytometer with a cover-slip in place. Live cells without the dye were counted in the

ten squares (2 at the center and 8 at the corner). The number of cells per millilitre =

average count pre square multiplied by 104 (each square contains 0.0001 millilitre) and

multiplied by dilution factor.

2.2.16 Isolation of human lymphocytes

Venous blood from healthy volunteers was collected in K2-EDTA coated vacutainer tubes, and mononuclear cells were isolated by density gradient centrifugation using Ficoll-Hypaque as described in Vasant et al. (2001). These peripheral mononuclear cells were cultured in RPMI 1640 medium with supplementation as described above plus

20 µg/ ml of interleukin 2 (IL-2). IL-2 was added immediately after isolation and treatment with YCT was started 1h later with IL-2 in the media for up to 48 h. The proportion of T cells to B cells in the isolated normal lymphocytes was estimated to be 9:

1 using the Coulter Epics Elite flow cytometer (ESP, Miami, Fl) with standard beads

(calibrated sizes) as reference.

2.2.17 Assessment of Cell Viability

We thought YCT extract, which contains at least 90 % mangiferin, was pure enough to be used as pure mangiferin as from Sigma-Aldrich. However, it was a mistake because the purity of YCT was below 99 %. Therefore, we changed the unit of the

53 concentrations of YCT from µM (25, 50, 100, and 150) to µg/ ml (10.56, 21.12, 42.23,

and 63.35). We used these concentrations to study the concentration-dependent

cytotoxicity of YCT on a wide range of cell types. The lower concentrations of 10.56 and

21.12 µg/ ml were not significantly toxic to most cell lines tested. Therefore, their results

are not shown in most of the figures and tables of Chapter 4.

2.2.17.1 MTT assay

Cell viability was measured by the MTT (3-(4,5-dimethyl-2-yl)-2,5-

diphenyltetrazolium bromide) method (Hansen, Nielsen and Berg, 1989). Cells in

DMEM were seeded overnight at a density of 1.5 x 104 cells per well in 96-well plates.

After exposure to YCT dissolved in fresh DMEM for various time points, 250 µl of MTT

(0.5 mg/ ml final concentration) dissolved in DMEM was added. Cells were incubated at

37 °C in the dark for 1h and then MTT was removed. 250 µl of DMSO was added to solubilize the formazan formed. Absorbance at 550 nm was determined using a microplate reader (SpectraMax190, Molecular Devices) after shaking in the dark for 15 min. Ethanol at the final concentration present (0.2%, v/v) had no effect on cell viability.

2.2.17.2 Trypan Blue Exclusion Assay

Jurkat T cells were plated in 24-well plates at a density of 5 x 105/ ml in RPMI

1640 medium with and without YCT (dissolved in 80% (v/v) ethanol) at different

concentrations. Duplicate hemocytometer counts of duplicate culture dishes were

performed at 0, 6, 12, 36, and 48 h of exposure to YCT. Results are expressed as the

percent change in trypan blue positive cells as compared with control (80% (v/v) ethanol

54 as carrier). Ethanol at the final concentration present (0.2%, v/v) had no significant effect

on cell viability.

2.2.18 Treatment of Jurkat T Cells with YCT

Jurkat T cells in RPMI 1640 medium were seeded at a density of 5 x 105 cells per

well in 6-well plates. After exposure to YCT at various concentrations (10.56, 21.12,

42.23, and 63.35 µg/ ml) for various time points, cells were harvested by centrifugation

(2000 rpm, 5 min) using a desktop centrifuge (Centrifuge 5417C, fixed angle rotor,

eppendorf), washed twice with phosphate buffered saline (PBS) and used immediately for

various analyses.

2.2.19 O2 electrode assay A Hansatech oxygen electrode (Hansatech, UK) was used for the indirect

measurement of H2O2 generated in the cell culture media as described by Long et al.

(1999). Briefly, the electrode was stabilized for 1 h with air saturated phosphate-buffered saline (PBS, pH 7.4) in the chamber. The buffer was then removed and washed twice with distilled water before replacing by 1.5 ml of culture medium in the absence or presence of YCT extract. Cell culture media pre-treated with YCT extract for 24 h, both with and without Jurkat T cells, in an incubator (37 ºC) were also tested for H2O2

generation. 100 µl of catalase solution in PBS was injected through the top of the

chamber. The deflection was followed by the recorder pen until it came back to the

starting point. Concentrations of H2O2 were determined from a freshly prepared standard curve of H2O2 prepared in PBS.

55 2.2.20 Cell cycle analysis using flow cytometry

YCT-induced loss of viability in Jurkat T cells and human peripheral lymphocytes

was measured as described in Ko et al. (2000) using the Coulter Epics Elite flow

cytometer (ESP, Miami, Fl). Briefly, Jurkat T cells were exposed to YCT for 1, 3, 5, 7,

12, 24 and 48 h. Human lymphocytes were exposed to YCT for 12, 24 and 48 h. Cells

were harvested and washed as described in the section Treatment of cells with YCT; and

fixed with 2 ml fixing solution consisting of PBS (200 µl) and 70% (v/v) ethanol (1800

µl) overnight. Fixed cells were harvested by centrifugation (2000 rpm, 5 min) using a

desktop centrifuge (Centrifuge 5417C, fixed angle rotor, eppendorf) and stained with 0.5

ml propidium iodide (PI) solution at 37 °C for at least 15 min. The PI solution consisted

of 1% (v/v) Triton-X-100 in PBS (1 ml), 10 mg/ ml RNAse A, 2.5 mg/ ml PI solution in

water (0.08 ml), and PBS (8.76 ml). The percentages of cells in subG1 (dead cells) and

G2/M (arrested cells) were calculated using WinMDI 2.8 software (Scripps Institute, La

Jolla, CA). Ethanol at the final concentration present (0.2%, v/v) did not affect cell

viability.

2.2.21 Annexin V-FITC and propidium iodide staining of Jurkat T cells

This was performed according to the experimental protocol from the supplier.

Briefly, treated Jurkat T cells were stained with 5 µl of Annexin V-FITC and 5 µl of propidium iodide (PI) after washing. Cells were gently vortexed for 5 sec followed by incubation for 15 min at room temperature (25 °C) in the dark. Analysis was performed using a flow cytometer (Becton Dickinson FACS Vantage SE, USA) (BD) within one hour. Results were expressed as percentage of cells with increased annexin V-FITC and

56 PI binding. Final concentration of ethanol in the system was 0.2% (v/v) and did not affect

cell viability.

2.2.22 Release of Lactate Dehydrogenase

Media were collected after centrifugation (2000 rpm, 5 min) using the desktop

centrifuge and cell pellets after washing were lysed by incubating for 10 min with 0.1%

(v/v) Triton X-100 at 37 °C. Cells were observed under the microscope to confirm

complete lysis. Lactate dehydrogenase (LDH) activity in the media and lysate were

measured with a commercially available kit (Sigma, LD-50) using a SpectraMax190

microplate reader (Molecular Devices) at 340 nm. Results were compared as a percentage

of LDH activity released from cells that had been lysed with 0.1% (v/v) Triton X-100 for

10 min at 37 °C. Ethanol at the final concentration present (0.2%, v/v) did not affect cell viability.

2.2.23 Assessment of cellular and nuclear morphology

Cellular morphology was viewed under an inverted microscope (Carl Zeiss,

Axiovert 25CFL) fitted with a digital camera (Nikon, CoolPix995). Nuclear morphology was assessed as described in Weis et al. (1995). Briefly, treated cells were harvested and washed twice with ice-cold PBS. Cells were then fixed in 4% (w/v) paraformaldehyde for

30 min at room temperature (25 °C). Fixed cells were washed and labelled with Hoechst

33258 (Molecular Probes, 0.5 µg/ ml) for 5 min at room temperature (25 °C) in the dark.

Cells were mounted with coverslips before viewing with an upright microscope

(Olympus BX51) using the filter cubes (UMWU2) for wavelength of 360 nm.

57 2.2.24 Caspase 3 and 9 activities

Cytosolic caspase 3 and 9 activities were determined according to the

experimental protocol from the supplier (Bio-Rad, Hercules, CA). Briefly, treated Jurkat

T cells (5 x 105 cells/ ml) were harvested and lysed by the addition of 100 µl lysis buffer after washing. The lysis buffer consisted of reaction buffer (100 mM HEPES, pH 7.4, 2 mM EDTA, 0.1% (w/v) CHAPS, 5 mM DTT) and a cocktail of protease inhibitors.

Complete lysis of the cells was confirmed by checking under the microscope after three freeze-thaw cycles (5 min in a -80 °C deep freezer followed by 5 min in a 37 °C water bath). Supernatants were collected by centrifugation at 13, 800 rpm at 4 °C for 15 min using the desktop centrifuge (Centrifuge 5417C, fixed angle rotor, eppendorf) and then used immediately for analysis or kept at -80 °C. Cell lysate (10 µl ~5 µg protein) was added to a reaction mixture containing water (80 µl), reaction buffer (100 µl), and caspase 3 fluorogenic substrate Ac-Asp-Glu-Val-Asp-AFC (Ac-DEVD-AFC, 10 µl), followed by incubation for 15 min at 37 °C. Fluorescence was determined using a Gemini

Fluorescence microplate reader with excitation and emission wavelengths of 390 and 520 nm, respectively, every 5 min for 30 min. The 7-amino-4-trifluoromethyl coumarin

(AFC) -substrate conjugate usually emits blue light (λmax = 400 nm) and cleavage of the substrate by the appropriate caspase liberates AFC, which fluoresce yellow-green light

(λmax = 505 nm). We used Ex = 390 nm and Em = 520 nm for the detection of caspase 3 activity because it gave a better signal under our experimental conditions. Determination of caspase 9 was performed in the same manner as caspase 3. The substrate used for caspase 9 assay was Ac-Leu-Glu-His-Asp-AFC (Ac-LEHD-AFC). Product formation was determined at excitation and emission wavelength of 400 and 505 nm, respectively,

58 every 5 min for 30 min. Caspase activity was expressed as relative fluorescence units per

mg protein. Results were expressed as fold increase as compared with control. Ethanol

present at the final concentration (0.2%, v/v) did not affect cell viability.

2.2.25 Western blot analysis

Experiments were conducted as described in Zhivotovsky et al. (1999). Briefly,

control and treated cells were washed and resuspended in buffer A (100 mM sucrose,

1mM EGTA, 20mM MOPS, pH 7.4, 5% (v/v) Percoll, 0.01% (w/v) digitonin, and a cocktail of protease inhibitors). Cells were centrifuged at 13,800 rpm at 4 °C for 15 min using the desktop centrifuge (Centrifuge 5417C, fixed angle rotor, eppendorf) to remove unbroken cells, nuclei, mitochondria and other organelles after incubation on ice for 15 min. The resulting supernatant was designated as the cytosolic fraction. The protein concentration of the supernatant was determined with the DC Protein assay kit (Bio-Rad) and 20 µg protein was boiled for 10 min before loading in a 12.5% (v/v) SDS-PAGE gel.

The separated proteins were transferred to nitrocellulose membranes and probed with antibodies against procaspase-3 and -7 and cytochrome c followed by the appropriate

HRP-conjugated secondary antibodies. Detection was by enhanced chemiluminescence

(Pierce). For cytochrome c, treated Jurkat T cells were fractionated using a mitochondria isolation kit (Cat No. 89874, Pierce) according to the supplier’s instructions. 40 µg of proteins from both the cytosolic and mitochondria fractions were loaded in a 15% (v/v) gel.

59 2.2.26 Internucleosomal DNA fragmentation assay

DNA isolation was conducted as described in Spencer et al. (2000). Briefly,

treated cells were washed and lysed with lysis buffer (0.1 M NaCl, 20 mM Tris-base pH

8.0 containing 10 mM EDTA and 0.5% (w/v) SDS) on ice for 10 min. Cell lysate was

treated with 15 U/ ml RNAse A and 10 U/ ml RNAse T1 for 1h at 37 °C. This was

followed by 2 h with proteinase K (30 U/ ml) at 37 °C. Protein was precipitated with 6 M

NaCl and removed by centrifugation at 4000 rpm for 15 min using a swinging bucket

centrifuge (Hitachi). DNA was precipitated with ice-cold ethanol, washed, and dissolved

in TBE buffer (0.09 M Tris base, 0.09 M boric acid, 2 mM EDTA, pH 8.3). The purity of

DNA was estimated by absorbance measurement ratio (260 /280 nm) between 1.7-1.9.

DNA was then loaded on 1% (w/v), agarose gels and electrophoresed as described in

Frisch and Francis (1994).

•− 2.2.27 Cytofluorimetric measurement of superoxide radicals (O2 ) and other reactive

2+ 2+ oxygen species (ROS), lipid peroxidation (LPO), intracellular Ca ([Ca ]i),

2+ 2+ mitochondrial Ca ([Ca ]m), mitochondrial transmembrane potential (∆Ψm),

+ + intracellular potassium ([K ]i) and sodium ([Na ]i)

Selective dyes used in experiments included JC-1 and DCFDA (Mukherjee et al.,

2002), Fluo3/ AM and DHE (Macho et al., 1997), C11-Bodipy581/591 (Pap et al., 1999,

Drummen et al., 2002), TMRM (Wong and Cortopassi, 2002), Rhod 123 (Bestwick and

Milne, 2001), Rhod-2/ AM (Babcock et al., 1997), PBFI/ AM and SBFI/ AM (McCarthy

and Cotter, 1997). 2’,7’-dichlorodihydrofluorescein diacetate (DCFDA) is deacetylated

by intracellular esterases forming a nonfluorescent product dichlorofluorescin (DCFH)

60 that can be converted by reactive species into 2’,7’-dichlorofluorescein (DCF), which

fluoresces at around 525 nm when excited at around 488 nm (Halliwell and Whiteman,

2004). Dihydroethidium (DHE) is a redox-sensitive probe frequently used to detect

• • intracellular O2 ¯. It is the oxidized product from the reaction between DHE and O2 ¯, oxyethidium (Oxy-E), which intercalates into nuclear DNA and fluoresces at around 580 nm when excited at around 500 nm (Fink et al., 2004). Rhod 123, JC-1 and TMRM are

some of the probes that used to study mitochondrial transmembrane potential (Δψm).

These probes are membrane-permeant and positively charged, and tend to accumulate in

the mitochondria, remained there by Δψm. Rhod 123 and TMRM fluoresce at around 536 and 590 nm when excited at around 500 and 550 nm, respectively. Loss of fluorescence is an indication of membrane depolarization. JC-1 is a ratiometric fluorescent cationic

dye that is concentrated by mitochondria in proportion to their Δψm. The dye exists as a monomer that fluoresces green (535 nm) and forms J-aggregates, which fluoresces red

(595 nm) in the mitochondria when excited at around 488 nm. Loss of Δψm can be calculated from the ratio of the reading at 595 nm (red fluorescence) to the reading at 535 nm (green fluorescence) (595: 535 ratio). C11-Bodipy581/591 is a lipophilic fluorescent ratio-probe suitable for the determination of lipid peroxidation (Drummen et al., 2002). It is insensitive to the changes in pH and has a good photo-stability. Non-oxidized and oxidized C11-Bodipy581/591 fluoresces red (~ 595 nm) and green (~ 520 nm), respectively, when excited at around 488 nm. Fluo3/ AM and Rhod-2/ AM are both Ca2+-sensitive probes for measuring intracellular and mitochondrial Ca2+, respectively. These probes

61 exhibit large fluorescence intensity increases (> 100-fold) upon binding Ca2+. Fluo3-Ca2+ fluoresces at around 526 nm when excited at around 505 nm. Rhod-2/ AM is a cationic probe that accumulates readily into mitochondria and fluoresces red (~ 595 nm) when excited at around 550 nm. SBFI/ AM and PBFI/ AM are fluorescent indicators for sodium and potassium, respectively. The AM ester is hydrolysed within the cell to yield the Na+- or K+-sensitive free acid, which fluoresces at around 510 nm when excited at

340 and 380 nm.

Briefly, stock solution of dyes in DMSO were first dissolved in either serum free

RPMI 1640 medium or PBS (for Fluo3/ AM and Rhod-2/ AM only) and loaded into the treated cells after cell washing. Cells were harvested by centrifugation (2000 rpm, 5 min) using the desktop centrifuge (Centrifuge 5417C, fixed angle rotor, eppendorf) after incubation at 37 °C for 30 min or 1h (for DCFDA, PBFI/ AM and SBFI/ AM only) in the dark. Cells were washed with PBS to remove excess dye, resuspended in 500 µl PBS and kept on ice until flow cytometry analysis (Coulter). Analysis was usually finished within

30 min. Samples were checked before and after each analysis to ensure that there were no significant changes to the distribution of intracellular fluorescent dyes. Final concentrations of each dye loaded were 5 µM for DCFDA, 2 µM for DHE and Fluo3/

AM, 1 µM for C11-Bodipy581/591, JC-1, Rhod 123, PBFI/ AM, and SBFI/ AM, 500 nM for Rhod-2/ AM and 50 nM for TMRM. For PBFI/ AM and SBFI/ AM, stock solutions were freshly prepared by mixing equal volume of the dye and 20% (w/v) pluronic acid F-

127. For TMRM, cells were permeabilized with digitonin (2 µg/ ml) for 5 min on ice,

followed by washing with PBS to remove excess dye and digitonin. The final

62 concentrations of DMSO (0.1%, v/v), pluronic acid F-127 (0.1%, v/v), and digitonin (2

µg/ ml) had no effect on cell viability.

2.2.28 Multiwell plate reader measurement of lipid peroxidation (LPO) and

mitochondrial transmembrane potential (∆Ψm)

LPO was also determined with cis-parinaric acid (Hedley and Chow, 1992; Tiku

et al., 2000) and ∆Ψm with JC-1 (Qu et al., 2001) and TMRM (Bestwick and Milne,

2001) using a Gemini Fluorescence microplate reader (Molecular Devices). Procedures were essentially the same as for flow cytometry measurements. Cells suspended in PBS after washing were aliquoted in duplicates into a 96-well black plate and fluorescence was determined. The excitation (Ex) and emission (Em) wavelengths were cis-parinaric acid (Ex: 325 nm Em: 405 nm), JC-1 (Ex: 488 nm Em: 535/ 590 nm), and TMRM (Ex:

550 nm Em: 590 nm). Cis-parinaric acid is a natural polyunsaturated fatty acid that contains four conjugated double bonds, which render it naturally fluorescent. It loses its fluorescence when oxidized by ROS. Caution should be exercised when handle cis- parinaric acid to avoid oxidative and photolysis degradation of stock solution of cis- parinaric acid, which was prepared in 100 % ethanol and stored at -20 ºC under nitrogen gas.

2.2.29 Intracellular ATP determination

Jurkat T cells were pelleted by centrifugation (2000 rpm, 5 min) using the desktop centrifuge (Centrifuge 5417C, fixed angle rotor, eppendorf). Cell pellets were washed twice with ice-cold PBS, followed by the addition of 200 µl of ice-cold trichloroacetic acid (TCA) (6.5%, w/v) and left on ice for 10 min. The TCA extract was removed and

63 used immediately for analysis of GSH and ATP or stored at -80 °C. ATP was measured

as described in Kalbhem and Koch (1967). Briefly, 3 µl of TCA extract was incubated with 200 µl of sodium arsenite buffer (26.67 mM MgSO4.7H2O, 33.33 mM KH2PO4, and

33.33 mM Na2HAsO4.7H2O, pH 7.4) at room temperature. Luminescence was measured

immediately after the addition of 10 µl of firefly lantern extract for 10 sec using a LUMI-

ONE luminometer (Trans Orchid Enterprises, Tampa, FL). Concentrations of ATP were

determined from a freshly prepared standard curve of ATP. Protein concentration was

measured by the BCA Protein Assay Kit after addition of 100 µl NaOH (1 M) to

solubilize cellular protein.

2.2.30 Intracellular glutathione measurement

GSH was measured according to Hissin and Hilf (1976). 7.5 µl of TCA extract (as described in the ATP determination section) was added to 96-well plates followed by the addition of 227.5 µl of KH2PO4-KOH buffer (100 mM, pH 10.0), and 15 µl of freshly

prepared o-phthaldialdehyde (10 mg/ml in methanol). Plates were incubated in the dark at

room temperature (25 °C) with gentle shaking for 25 min, followed by measurement

using a Gemini Fluorescence plate reader at excitation 350 nm and emission 420 nm.

Concentrations of GSH were determined from a freshly prepared standard curve of GSH.

2.2.31 Cytochrome c levels

Cytosolic cytochrome c was determined according to the experimental protocol

from the supplier (Cat. No. APT200, CHEMICON Int., Temecula, CA). Briefly, Jurkat T

cells at a density of 5x106 cells/ ml were harvested after treatment with YCT and washed

64 with ice cold PBS three times. The cells were resuspended with 1 ml of Buffer I (10 mM

Tris-HCl, pH= 7.5, 0.3 M sucrose, protease inhibitor cocktail tablet (1 tablet per 10 ml of

Buffer I)). Cells suspension was then sonicated (Misonix Sonicator XL2020, Structure

Probe, PA, USA) on ice with the following conditions: output control = 1.5, total time =

3 min, pulse on time = 30 sec, pulse off time = 5 sec. Cells were observed under the

microscope to confirm complete lysis. Cell lysates were then centrifuged at 9,700 rpm for

60 min at 4 °C using the desktop centrifuge (Centrifuge 5417C, fixed angle rotor,

eppendorf). The supernatant (cytosolic fraction) was collected and used for analysis

immediately or kept at -80 °C. The pelleted cell fragments/ organelles were resuspended

with 500 µl of Buffer II (10 mM Tris-HCl, pH 7.5, 1 % (v/v) Triton X-100, 150 mM

NaCl, protease inhibitor cocktail tablet (1 tablet per 10 ml of Buffer I)) (Roche, Basel,

Switzerland) followed by sonication (Misonix Sonicator XL2020, Structure Probe, PA,

USA) on ice with the following conditions: output control = 2, total time = 4 min, pulse on time = 30 sec, pulse off time = 5 sec. Cell lysates were then centrifuged at 9,700 rpm for 30 min at 4 °C using the desktop centrifuge (Centrifuge 5417C, fixed angle rotor, eppendorf). The supernatant was collected and used as the mitochondrial fraction.

Analysis of cytochrome c was conducted following the instructions provided by the manufacturer. Absorbance was determined at 450 nm using the SpectraMax190 microplate reader. Concentrations of cytochrome c were determined from a freshly prepared standard curve of cytochrome c. Ethanol at the final concentration present

(0.2%, v/v) did not affect cell viability.

65 2.2.32 Kinetic study of rise in intracellular Ca2+ with Fluo3/ AM

Experiments were conducted as described in Jiang et al. (1994). Jurkat T cells (1

x 107 cells/ ml) were harvested and washed twice with PBS, followed by loading with

Fluo3/AM dye to a final concentration of 5 µM in RPMI 1640 medium (with 5% (v/v)

FBS) at 37 °C for 30 min. Cells were then washed free of extracellular Fluo3/ AM dye and resuspended in PBS supplemented with 5 mM sucrose and 1 mM CaCl2. Final cell

concentration was 2 x 106 /ml and cells were kept on ice until used. Fluorescence was

determined at room temperature using a Perkin-Elmer LS-50B fluorescence

spectrophotometer with excitation and emission wavelengths of 506 and 526 nm,

respectively. 2 ml of cell suspension was loaded into a quartz cuvette and measurement

was started to obtain a basal fluorescence level of the cells. Treatment with YCT was

initiated after 1 min and the increase in fluorescence was recorded for 5 min. Where

inhibitors were used, cells were pre-treated with various concentrations of inhibitors for

10 min before YCT exposure.

2.3 Data Analysis

The mean values were calculated from data taken from at least 3 separate

experiments (in triplicate) conducted on separate days using freshly prepared reagents

unless otherwise stated. Where significance testing was performed, an independent t-test

(Students; 2 populations) was used; * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## <

0.01, ###p < 0.001. No gating of cell population was done for the analysis of the cytofluorimetric data using the WinMDI 2.8 software (Scripps Institute, La Jolla, CA).

66 CHAPTER 3 RESULTS

3.1 Radical scavenging by TCM extracts Thirty−three TCM extracts (see methods section) were screened for their antioxidant activity using the ABTS assay and their TEAC values are shown in table 3.1. Three different concentrations of each extract were used in the assay to check that the scavenging activity was proportional to the amount of extract added. And, the TEAC was calculated by dividing the gradient of TCM extracts from the plot of % inhibition of absorbance vs. concentration by the gradient of the plot for Trolox.

Table 3.1. Trolox equivalent antioxidant capacity (TEAC) values of TCM extracts Code of TEAC (mM) Code of TEAC (mM) Code of TEAC (mM) TCM* Mean ± s.d. TCM Mean ± s.d TCM Mean ± s.d A 0.03 ± 0.01 M 0.01 ± 0.00 LT 0.06 ± 0.01 B 0.07 ± 0.01 N 0.06 ± 0.00 RS 0.04 ± 0.00 C 0.15 ± 0.01 O 0.02 ± 0.00 SY# 0.28 ± 0.01 D# 0.67 ± 0.04 CC 0.08 ± 0.00 PM 0.10 ± 0.01 E# 0.18 ± 0.01 ES 0.16 ± 0.01 AS 0.02 ± 0.00 F 0.25 ± 0.03 PC# 0.24 ± 0.00 MS 0.07 ± 0.00 G 0.06 ± 0.00 WN# 0.23 ± 0.01 HZ 0.06 ± 0.00 H 0.26 ± 0.00 DA 0.06 ± 0.01 X 0.18 ± 0.00 I 0.11 ± 0.01 MO 0.01 ± 0.01 YZ50 0.11 ± 0.01 J 0.04 ± 0.01 LB 0.06 ± 0.01 Trolox 1.00 ± 0.00 K 0.04 ± 0.00 AA 0.16 ± 0.02 L 0.13 ± 0.00 PK 0.03 ± 0.00 •+ *1.0 ml of diluted ABTS solution (A734nm = 0.8 ± 0.05) was added to 5 µl of TCM extracts of three different concentrations (Trolox: 0.5, 1.0, 2.5 µg/ ml; other TCM extracts: 5, 10, 20 µg/ ml) as described in Experimental Procedures. Absorbance reading was taken 3 min after initial mixing. Trolox was used as a standard. TEAC was calculated by dividing the gradient of TCM extract obtained from the plot of % inhibition of absorbance vs. concentration by the gradient of the plot for Trolox. Results are expressed as mean ± s.d. of at least three independent experiments performed in triplicate. # TCM extracts were selected for further testing using our laboratory extraction protocol.

67 Fig. 3.1A shows a typical trolox standard curve. The five extracts with the highest

TEAC values (Table 3.1. D (0.67 ± 0.04), E (0.18 ± 0.01), PC (0.24 ± 0.00), WN (0.23 ±

0.01), and SY (0.28 ± 0.01)) were selected for further testing using our laboratory extraction protocol. The differences in extraction protocol between laboratories may explain the different TEAC values obtained for some of the extracts. For example, we have an additional step of removing the chlorophyll contents of the TCM extracts by partitioning with hexane in our extraction protocol. As shown in Fig. 3.1C, extract WN obtained using our extraction protocol showed higher antioxidant activity than Trolox.

Reaction of Trolox with ABTS•+ was complete after 3 min but the decolorization with extracts continued for at least 1 hour (Fig. 3.1B, only extract WN is shown), suggesting that the extracts may contain a mixture of slow-and fast-reacting species (Long et al.,

2000). However, for screening purposes, we retained measurements at a fixed time of 3 min. Ethanol at the final concentration present (0.4%, v/v) did not reduce the ABTS absorbance at 734 nm.

A 50 y = 9.7326x R2 = 0.997

) 40 m 734 n

30 ease ce at ecr

n 20 D a b % r o s

b 10 a (

0 0123456

Trolox Conc. (µg/mL)

68

B

Trolox WN 1 m n

4 0.8 3 7 0.6 at ce

n 0.4 a b r 0.2 so b

A 0 02345678910152025304560 Time (min)

C *** 1.55 1.60 1.40 1.20 1.00 1.00 0.80 *** *** ***

AC (mM) 0.60 0.36 *** 0.23 0.22 TE 0.40 0.09 0.20 0.00 Trolox D E PC WN SY

Fig. 3.1. Radical scavenging by TCM extracts. (A) Concentration-response curve for the absorbance at 734 nm for ABTS•+ as a function of concentration of standard Trolox solution dissolved in 80% ethanol. (B) Time-course of ABTS•+ depletion in the presence of Trolox standard and extract WN. Data are mean ± s.d., n≥ 3. (C) Trolox equivalent antioxidant capacity (TEAC) values of selected TCM extracts. 1.0 ml of diluted ABTS•+ solution (A734nm = 0.8 ± 0.05) was added to 5 µl of TCM extracts of three different concentrations (Trolox: 0.5, 1.0, 2.5 µg/ ml; WN: 0.65, 1.25, 2.5 µg/ ml; D, E, PC, SY: 5, 10, 20 µg/ ml) as described in Experimental Procedures. Absorbance reading was taken 3 min after initial mixing. Trolox was used as a standard. Results are expressed as mean ± s.d. of at least three independent experiments performed in triplicate. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with Trolox.

69 3.2 Inhibition of ascorbate-iron induced phospholipid peroxidation

*** 80 64

n 49 *** o

i 60 31 40 29 *** % Inhibit 20 4 *** 0 0 Trolox D E PC WN SY

Fig. 3.2. Inhibition of ascorbate-iron induced phospholipid peroxidation. The formation of (TBA)2-MDA chromogen from lipid peroxidation was measured at 532 nm. Trolox and TCM extracts were tested at final concentration of 5 µg/ ml. Experiments were conducted as described in Experimental Procedures. Results are expressed as mean ± s.d. of at least four independent experiments performed in triplicate. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with Trolox.

The percentage inhibition of ascorbate−iron induced phospholipid peroxidation by

Trolox and TCM extracts was determined (Fig. 3.2). Extracts PC and WN at final

concentrations of 5 µg/ ml exerted more powerful inhibitory effects than Trolox (p <

0.001). Extract PC performed better than WN in inhibiting lipid peroxidation (p < 0.05).

Ethanol at the final concentration present (0.4%, v/v) did not inhibit phospholipid peroxidation.

In a buffer containing 1 mM FeCl3 (final concentration) and at a pH of 7.0, the

Ksp (solubility product constant) of Fe(OH)x would have been surpassed. However, the

final concentration of FeCl3 in our assay studied is 100 µM. Further studies are needed to test whether the Ksp of Fe(OH)x is surpassed at this concentration.

70 3.3 Correlation between TEAC values, total phenolic content and DPPH reducing ability A

0.70 y = 0.0617x 0.60 2 0.50 R = 0.9952

ance at 0.40

b 0.30 765 nm

sor 0.20

b 0.10 A 0.00 01234567891011

Gallic acid conc. (µg/ mL)

B C 500 10

8

400 ) L m ) g g/ /

g 6

300 m ( m ( 50 PPH E C D 4

200 E GA / 1

n l 100 2

0 0 00.511.52 0.0 0.5 1.0 1.5 2.0 TEAC (mM) TEAC (mM)

Fig. 3.3. Correlation between TEAC values and total phenolic content. (A) Concentration-response curve for the absorbance at 765 nm for reduced Folin-Ciocalteu’s reagent as a function of concentration of standard gallic acid solution. (B and C) Correlation between TEAC values of selected TCM extracts (Cratoxylum cochinchinense, Cortex magnoliae officinalis, Psoralea corylifolia L, Curculigo orchioides Gaertn, and Glycyrrhiza uralensis Fisch.) and total phenolic content (Gallic acid equivalents, GAE) (B, r= 0.997, p< 0.001) and DPPH (C, r= 0.982, p< 0.001). Data are representative of at least three independent experiments performed in triplicate.

Fig. 3.3A shows a standard curve of gallic acid freshly prepared for an approximate estimation of total phenolic contents of TCM extracts. As shown in Fig.

3.3B and C, the TEAC values of the extracts and their total phenolic contents were well

71 correlated (Fig. 3.3B, r= 0.997, p < 0.001), indicating that the phenolic compounds make

a major contribution to the antioxidant capacity of the extracts. However, it should be

noted that Folin-Ciocalteu’s phenol reagent gives only an approximate estimate of the

total phenolic content of TCM extracts. The extracts also performed well in an organic

phase to reduce DPPH free radical to an extent highly correlated with their TEAC values

(Fig. 3.3C, r= 0.982, p < 0.001). Ethanol at the final concentrations of 0.4% (v/v) had no radical scavenging effects in these assays.

3.4 Superoxide scavenging and xanthine oxidase inhibition

•− Table 3.2. Superoxide (O2 ) scavenging and xanthine oxidase (XO) inhibitory effects of TCM extracts.

IC50 (µg/ ml) •− Extracts O2 Xanthine Oxidase

D > 25 > 600 E > 25 > 600 PC > 25 > 300 WN 15± 3*** > 600 SY > 25 > 600 Results were recorded as IC50. Experiments were conducted as described in Experimental Procedures. Results shown are mean ± s. d. of at least 3 independent experiments performed in triplicate. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with other TCM extracts.

As shown in Table 3.2, most TCM extracts did not scavenge superoxide radicals

at a significant rate, with IC50 values above 25 µg/ ml. However, extract WN showed an

IC50 of 14.95 ± 2.95 µg/ ml. It is important in such experiments to check for possible

interference with the assay by inhibition of XO (Halliwell, 1995). However, the IC50s for

XO were above 600 µg/ ml, except that PC at a concentration higher than 300 µg/ ml precipitated in the reaction mixture and could not be tested at a higher concentration.

72 Solvent controls were included and had no significant effect on NBT reduction or

xanthine oxidase activity.

3.5 Iron-binding activity

D E PC WN SY 70

60 ) %

( 50 y t i v i

t 40 c

30 ng a

ndi 20 bi on-

r 10 I

0 0 0.25 0.5 0.75 1 -10

-20 Extract concentration (mg/ ml)

Fig. 3.4. Iron-binding activity of TCM extracts. Results shown are mean ± s.d. of at least three independent experiments performed in triplicate.

Although plant derived polyphenols are good antioxidants in nature, some plant phenols can also interact with transition metals to produce pro-oxidant effects in vitro.

Ferrous ion is one of the most powerful pro-oxidant reported (Halliwell and Gutteridge,

1999). Tetramethyl murexide (TMM) is a chelating reagent, showing an absorption maxiumum at 530 nm, and formed a complex with free Fe2+ (TMM-Fe2+) that absorbs at

485 nm. Therefore, TCM with strong iron-binding activity could reduce the absorbance

2+ 2+ ratio of A485/ A530 (TMM-Fe / TMM) and the participation of Fe in the Fenton

reaction. Extract WN achieved about 50% binding of Fe2+ at relatively low concentration of 0.1 mg/ ml and remained up to 1 mg/ ml. Other extracts bound less than 40% Fe2+ at

73 the concentration of 1 mg/ ml. Experimental errors that can arise from the oxidation of

Fe2+ to Fe3+ was minimized by performing the experiment on top of a bucket of ice.

3.6 Scavenging of hypochlorous acid (HOCl); Protection against HOCl-mediated DNA damage

Table 3.3. Inhibition of HOCl induced DNA damage by Cratoxylum cochinchinense (WN). Oxidized base Control DNA + HOCl DNA + HOCl + WN 0.125 mg/ ml 2.5 mg/ ml 5-(OH, Me) Hydantoin 0.15 ± 0.12 0.25 ± 0.04a 0.12 ± 0.08a 0.11 ± 0.08a 5-OH Uracil 0.08 ± 0.03 16.62 ± 1.18b 0.16 ± 0.04c 0.09 ± 0.02a 5-(OH, Me) Uracil 0.22 ± 0.05 1.06 ± 0.21b 0.22 ± 0.06a 0.21 ± 0.07a 5-OH Cytosine 0.10 ± 0.01 10.55 ± 3.01b 0.24 ± 0.09a 0.16 ± 0.08a 5-Cl Uracil 0.03 ± 0.02 12.96 ± 2.60b 0.30 ± 0.06c 0.15 ± 0.05d 5-Formyl Uracil 0.70 ± 0.15 14.02 ± 2.62b 0.46 ± 0.05c 0.64 ± 0.13a Thymine Glycol 0.66 ± 0.07 25.67 ± 6.79b 0.53 ± 0.14a 0.50 ± 0.29a FAPY Adenine 1.03 ± 0.21 1.36 ± 0.60a 0.96 ± 0.22a 0.83 ± 0.12a 8-OH Adenine 0.07 ± 0.01 2.41 ± 0.39b 0.11 ± 0.02c 0.07 ± 0.01a 2-OH Adenine 0.07 ± 0.02 0.16 ± 0.02b 0.05 ± 0.02a 0.04 ± 0.03a FAPY Guanine 2.35 ± 0.93 24.25 ± 6.60b 1.76 ± 0.39a 0.80 ± 0.01c 8-OH Guanine 0.25 ± 0.09 4.21 ± 0.39b 0.17 ± 0.05ac 0.11 ± 0.04c DNA (1.0 mg/ ml) was oxidized using HOCl (500 µM) for 1 h at 37 °C. The samples were then dialyzed against water for 24 h, freeze dried, and hydrolyzed with 60% (v/v) formic acid. Oxidized DNA base products were analyzed by GC−MS after derivatization for at least 2 h. Different superscripts in the same row indicate significant differences (p < 0.01) as compared with control. Results are expressed as mean ± s.d. of 3 or more separate determinations performed in triplicate and expressed as nmol/ mg DNA.

Incubation of DNA with HOCl (500 µM) at pH 7.4 for 1 h at 37 °C led to significant increases (Table 3.3, p < 0.01) in the levels of a wide range of DNA base of modification products, except 5-(OH, Me) hydantoin and FAPY adenine. The increases in thymine glycol and FAPY guanine were greatest, followed by 5−hydroxyuracil,

74 5−formyluracil, 5−chlorouracil, and 5−hydroxycytosine (Whiteman et al., 1997). When

DNA was pre-incubated with extract WN at different concentrations (0.125 and 0.25 mg/

ml) DNA base product formation decreased significantly (p < 0.01) in a dose dependent

manner. WN itself did not change levels of modified bases in DNA. Ethanol at the final

concentration present (0.5%, v/v) had no effect on HOCl-mediated DNA damage.

3.7 Inhibition of protein glycation

D E WN SY

120 *** ** 100 80 60 40 Inhibition % 20 0 2.5 5 6.25 Conc (µg/ ml)

Fig. 3.5. Inhibition of protein glycation (advanced glycation end products (AGE) formation) by TCM. Extract PC could not be tested in this assay due to its strong fluorescence at the excitation maximum of 350 nm. Trolox enhanced the formation of AGE (data not shown). Experiments were conducted as described in Experimental Procedures. Results are expressed as mean ± s.d. of three independent experiments performed in quadruplicate. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with other TCM extracts.

Non-enzymatic glycation of proteins with reducing sugars and the subsequent transition metal catalysed oxidations leads to the formation of "advanced glycation

endproducts" (AGEs) are one of the consequences of hyperglycaemia (Vinson and

Howard, 1996). The accumulation of AGE on long-lived proteins such as collagen, eye

lens crystalline and β-amyloid plaques in Alzheimer’s disease (AD) may contribute

75 additionally to oxidative stress in the AD brain (Loske et al., 1998). We therefore

examined the protective effect of TCM on AGE formation (Fig. 3.5). Extract WN was

able to achieve more than 90% inhibition against AGE formation in vitro at a

concentration of 2.5 µg/ ml. Its effect was significantly higher than the other extracts at concentrations of 2.5 and 5 µg/ ml. Extract D at a concentration of 6.25 µg/ ml was as potent as WN. Ethanol at the final concentration present (0.4%, v/v) did not affect AGE formation.

3.8 Purification and identification of ‘active’ ingredient from extract WN

More than 90% of the major compound in the extract of WN was present in the

35% (v/v) methanol fraction. This fraction possessed high antioxidant activity as

determined by the ABTS assay (data not shown). Initially, the ‘active’ compound was

identified as a C−glucosylxanthone (mangiferin,

1,3,6,7−tetrahydroxyxanthone−C2−β−D−glucoside, Fig. 1.1) by comparison with an

authentic compound from Sigma using the HPLC−PDA (Agilent 1100 Series) and

LC−MS (Finnigan−LCQ). In our experimental conditions, tR for pure mangiferin (MGF)

and the major peak in the semi-purified extract (now referred to as YCT) were 12.45

(Fig. 3.6A) and 12.46 min (Fig. 3.6B), respectively. Fig. 3.6C shows the chromatogram

of a mixture of MGF and YCT (ratio 1: 1, v/v) with a tR of 12.46 min and similar UV

spectra (Fig. 3.6D). With LC-MS, both MGF and YCT show a tR of 8.29 min and the same molecular ions (unfragmented ion Mw: 423.1 and fragmented ions Mw: 405.2 and

303.4) (Fig. 3.7A & B).

76

DAD1 B, Sig=254,16 Ref=off (TSY\YCT-MG~1\15M4.D) DAD1 E, Sig=320,16 Ref=off (TSY\YCT-MG~1\15M4.D) 5 Norm. 4 Fig. 3.6A 12.45 12.45 600

500

400

300

200

100 12.616 12.616 12.967 21.934 23.484 21.325 3.177 11.838 4.632 22.668 20.384 23.222 3.844 0 3.176

0 2.5 5 7.5 10 12.5 15 17.5 20 22.5 min DAD1 B, Sig=254,16 Ref=off (TSY\YCT-MG~1\15Y4.D) DAD1 E, Sig=320,16 Ref=off (TSY\YCT-MG~1\15Y4.D) 7 Norm. 7 Fig. 3.6B 12.45 350 12.45

300

250

200

150

100

50 21.935 12.979 12.595 13.006 3.712 3.709 10.760 23.490 12.078 12.143 12.747 12.744 21.336 12.835 21.948 22.688 23.219 15.941 20.121 3.229 11.853 22.683 4.411 10.757 15.162 15.941 20.376 16.351 23.194 22.908 23.382 22.201 22.375 0 21.524 3.223

0 2.5 5 7.5 10 12.5 15 17.5 20 22.5 min DAD1 B, Sig=254,16 Ref=off (TSY\YCT-MG~1\15MY8.D) DAD1 E, Sig=320,16 Ref=off (TSY\YCT-MG~1\15MY8.D) 6 Norm. 6 Fig. 3.6C 12.44 12.44 800

700

600

500

400

300

200

100 12.916 12.610 12.610 21.929 12.996 3.184 10.624 23.494 2.506 21.321 21.942 12.722 2.500 3.182 12.348 3.110 6.732 20.318 15.109 22.678 15.921 23.219 22.867 23.380 0 22.207

0 2.5 5 7.5 10 12.5 15 17.5 20 22.5 min DAD1, 12.450 (964 mAU,Apx) of 15MY8.D DAD1, 12.459 (807 mAU,Apx) of 15Y4.D MGF+YCT DAD1, 12.456 (847 mAU,Apx) of 15M4.D Fig. 3.6D Norm. MGF

800

600 YCT

400

200

0 200 225 250 275 300 325 350 375 nm

Fig. 3.6. Identification of mangiferin (MGF) by high performance liquid chromatography (HPLC-PDA). Chromatogram of authentic MGF (A), YCT (B), a mixture of MGF and YCT (C), and UV spectra of MGF, YCT, and MGF + YCT (D). Retention time (tR) of MGF, YCT extract and the mixture of MGF and YCT are 12.45, 12.46, and 12.46 min, respectively. UV spectra of MGF and YCT were monitored at 254 and 320 nm. Conditions of identification were as described in the Experimental Procedures. Data are representative of three separate experiments.

77 8.29 423.1 Fig. 3.7A-MGF

405.2

303.4

8.29 423.1 Fig. 3.7B-YCT

405.2

303.4

Fig. 3.7. Identification of mangiferin by liquid chromatography- mass spectrometry (LC- MS). A- an authentic mangiferin purchased from Sigma-Aldrich (St. Louis, MO, USA). Left- retention time (tR): 8.29 min; right- Molecular weight (Mw) of unfragmented ion: 423.1, fragmented ions: 405.2 and 303.4. B- YCT extracted from C. cochinchinense. Left- tR: 8.29 min; right- Mw of unfragmented ion: 423.1, fragmented ions: 405.2 and 303.4. Conditions of identification processes were as described in the Experimental Procedures. Data are representative of two separate experiments.

78 3.9 Low pro-oxidant effect of YCT extract

40 YCT Trolox 35

30 e g

a 25 m

Da 20

15 DNA

% 10

5

0 0 50 100 150 200 250 300 Conc. (µg/ ml)

Fig. 3.8. Pro−oxidant effect of YCT compared to Trolox in causing DNA damage. MGF was not determined due to the interference of DMSO on DNA at high temperature. Results are mean ± s.d. of three independent experiments performed in triplicate.

The antibiotic bleomycin binds iron ions and the resulting bleomycin-iron

complex is capable of degrading DNA in the presence of O2 and a reducing agent

(Gutteridge et al., 1981). The bleomycin-Fe3+ complex by itself cannot cause DNA

damage (Aruoma, 1991). Therefore, a positive pro-oxidant effect is obtained when a

compound under test is able to reduce the bound Fe3+ to Fe2+ form which damages DNA.

Incubating DNA in the presence of bleomycin-Fe3+ for 1 h at 37°C with YCT at concentrations up to 250 µg/ ml did not induce more than 5% DNA damage as compared to control (ethanol (80%) alone). The estimation of DNA damage was based on the measurement of the absorbance at 532 nm of (TBA)2-MDA chromogen formed at high

temperature under an acidic condition. Trolox, on the other hand, showed dose-dependent

DNA damage of up to 40% at a concentration of 62.57 µg/ ml. Therefore, YCT is a

compound with potent antioxidant activity but low pro-oxidant effect in this assay.

79 3.10 Inhibition of PR bleaching and 3-nitrotyrosine formation by YCT

- A HCO3 (25 mM) 1.2

R 1.0 P f m o 0.8

ce YCT 2 n

n 0.6 a b

r 0.4 at 54 so

b 0.2 A 0.0 PR PR + PR PR + PR + ONONOO-OO- ONOOOO-- ONONOO-OO-

B YCT YCT+ HCO3−- HCO3 MGF MGF+ HCO3−- HCO3 100 *** 80 *** ng 60 *** ) on of i *** *** (% 40 *** *** ***

Inhibit *** PR bleachi 20 ***

0 0 20406080100120 Conc (µg/ml)

− Fig. 3.9. Inhibition of PR bleaching by YCT and MGF. (A) Effects of HCO3 (25 mM) on ONOO− -mediated PR bleaching. (B) Effects of YCT and MGF on the inhibition of pyrogallol red (PR) bleaching by peroxynitrite (ONOO−, 100 µM) in the absence and − presence of 25 mM HCO3 . Experiments were conducted as described in Experimental Procedures. Results shown are mean ± s.d. of at least 3 independent experiments performed in triplicate. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with MGF in the presence or absence of bicarbonate. The presence of bicarbonate did not affect the activity of YCT.

Carbon dioxide is present at sufficient concentration in vivo to react rapidly with peroxynitrite and this can modulate reactions of ONOO− (Denicola et al., 1996;

Ketsawatsakul et al., 2000). Therefore, we investigated the protective effects of our semi-

80 purified extract (YCT) on ONOO−-mediated PR bleaching and 3-nitrotyrosine formation

in the presence and absence of added bicarbonate. As shown in Fig. 3.9A, 100 µM of

ONOO− was able to reduce the absorbance at 542 nm of PR from 0.95 ± 0.14 to 0.26 ±

− 0.06 (n= 5) and the presence of 25 mM of HCO3 did not significantly affect the

bleaching effect of ONOO− (absorbance of PR dropped from 0.93 ± 0.07, n= 5 to 0.26 ±

0.06, n= 5). The presence of YCT or MGF deminished the decrease in absorbance of PR.

As shown in Fig. 3.9B, YCT was able to inhibit ONOO−- mediated bleaching of PR by

50% at a concentration of about 50 µg/ ml and its action was not affected by the presence of bicarbonate.

The addition of 1 mM of ONOO− to 1 mM of tyrosine led to the formation of 91 ±

11 µM of 3-nitrotyrosine under our experimental conditions (Fig. 3.10A). The presence

− of HCO3 (25 mM) significantly increased the formation of 3-nitrotyrosine to 247 ± 14

µM (n= 4) as previously reported by others (Gow et al., 1996; Ketsawatsakul et al., 2000;

Whiteman et al., 2002). These amounts of 3-nitrotyrosine formed were considered as

100% for the calculation of the inhibition of 3-nitrotyrosine formation in the presence of

YCT and MGF. As shown in Fig. 3.10B, YCT significantly inhibited 3-nitrotyrosine formation, virtually completely at a concentration of 85 µg/ ml in the absence of added bicarbonate. However, more YCT was required to achieve the same effect when 25 mM of bicarbonate was added. Ethanol at the final concentration present (0.4%, v/v) decreased ONOO−-mediated bleaching of PR or formation of 3-nitrotyrosine by ≤ 4%.

81 - A 300 + HCO3

) *** M 250 (µ e 200 n i s o

r 150 ty

o 100 tr i N

- 50 3 0 ONOO- ONOO-

B − YCT YCT+ HHCOCO33- − MGF MGF+ HHCOCO 3 - 100 a a

e 80 ) n l i o s r o

nt 60 a yr o a ***

C *** 2 Tr 40 of O b % N a ( a 3- 20 *** *** b a *** *** 0 a 0 20 40 60 80 100 120 140 160 180 200 220

Conc. (µg/ ml)

− Fig. 3.10. Inhibition of tyrosine nitration by YCT and MGF. (A) Effects of HCO3 (25 mM) on ONOO−-mediated 3-nitrotyrosine formation. (B) Effects of YCT and MGF on ONOO−-mediated 3-nitrotyrosine formation; effects of bicarbonate were also tested. Experiments were conducted as described in Experimental Procedures. Results shown are mean ± s.d. of at least four independent experiments performed in duplicate. a Comparison between YCT and MGF. b Comparison between YCT and MGF in the − *** − presence of HCO3 . Comparison between YCT and YCT+ HCO3 . Same superscript at each concentration indicate significant different (a and b, p < 0.01, ***, p < 0.001).

82 3.11 A comparison of our semi-purified extract (YCT) with pure mangiferin

Table 3.4. A comparison of the effects of YCT with MGF@

Sample % Inhibition (IC50) µg/ ml % Inhibition at sample concentration of 2.5 µg/ ml •− Phospholipid O2 Xanthine oxidase Formation of AGE peroxidation scavenging (XO) YCT 6.9 ± 0.2 5.6 ± 0.4*** 36.4 ± 5.1 93.8 ± 1.7* MGF > 16.9 *** 36.5 ± 1.3 > 42.2 72.7 ± 13.6 (86.4 ± 3.1 µM) Trolox 7.0 ± 0.6 ND ND ND @ •− For assays of ascorbate iron-induced phospholipid peroxidation, O2 scavenging and XO activity, results are expressed as the IC50, the concentration of the compound that inhibited 50% of (TBA)2-MDA chromogen formation, reduction of NBT or uric acid production, respectively. Protein glycation is expressed as % inhibition of AGE formation at the concentration of 2.5 µg/ ml relative to the control. Experiments were conducted as described in Experimental Procedures. Results are mean ± s. d. of at least 3 independent experiments performed in triplicate. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with Trolox (phospholipid peroxidation) and MGF (other assays). ND~ not determined.

YCT was compared with mangiferin (MGF) and Trolox for their protective effects against ascorbate-iron induced phospholipid peroxidation. The IC50 value of YCT was the same that of Trolox, whereas MGF was less inhibitory, showing a significantly

•− higher IC50 (Table 3.4). Moreover, YCT was able to scavenge O2 generated by the xanthine-xanthine oxidase system with an IC50 value of 5.60 ± 0.37 µg/ ml, which was

significantly lower (p< 0.001) than that of MGF, IC50 of 36.50 ± 1.29 µg/ ml (Table 3.4).

However, the IC50 values of YCT and MGF for XO inhibition were 36.41 ± 5.07 and >

42.23 µg/ ml, respectively. YCT at a concentration of 5.60 ± 0.37 µg/ ml did not inhibit

XO activity more than 5% (data not shown). Furthermore, YCT was as potent as WN

83 (Fig. 3.5) in inhibiting the formation of AGE on proteins and its effect was significantly

higher as compared with MGF (p< 0.05, Table 3.4).

With respect to ONOO− -mediated bleaching of PR, the inhibitory effect of YCT

− was much greater than that of MGF (Fig. 3.9B) whether or not HCO3 was added. MGF was unable to decrease PR bleaching by 50% at the highest concentration tested in the experiment. Furthermore, YCT inhibited 3-nitrotyrosine formation more strongly than

− MGF especially in the absence of added HCO3 (Fig. 3.10B). The final concentration of

DMSO (0.5%, v/v) used to dissolve MGF inhibited ONOO−-mediated bleaching of PR or formation of 3-nitrotyrosine by ≤ 5%.

In cell culture experiments, we used Jurkat T cells, HT29 and HepG2 cell lines as

examples to study the differences in cytotoxicity between YCT and MGF. YCT was

found to be more cytotoxic as compared to MGF to Jurkat T cells, HT29 and HepG2 cells

(Table 3.5 and Fig. 3.11). Incubating Jurkat T cells with YCT and MGF at a

concentration of 63.35 µg/ ml for 24 h induced 22.2 ± 2.4 (p < 0.001) and 10.7 ± 2.1% (p

< 0.01) cell death (% cells under SubG1), respectively (Table 3.5). With the same

concentration, YCT induced 17.3 ± 6.2 % (p < 0.01) cell death in HT29 cells but MGF

was relatively non-toxic. When the effects of YCT and MGF were tested on HepG2 cell

using the MTT assay, YCT was found to reduce cell viability in a concentration- and

time-dependent manner whereas MGF only reduced the viability of HepG2 cells by about

10% at a concentration of 63.35 µg/ ml after 12 h, with no change up to 48 h (Fig. 3.11).

84 Table 3.5 A comparison of the cytotoxicity of YCT extract with MGF on Jurkat T cells and colorectal adenocarcinoma (HT29) cellsa Jurkat T cells HT29

(% cells at SubG1) (% cells at SubG1)

Control 5.8 ± 1.6 6.5 ± 4.1

YCT (63.35 µg/ ml) 22.2 ± 2.4*** 17.3 ± 6.2**

MGF (63.35 µg/ ml) 10.7 ± 2.1** 5.5 ± 1.3

aCells were treated with YCT and MGF at a concentration of 63.35 µg/ ml for 24 h. Cell cycle analysis was determined after staining cells fixed overnight with ethanol (70% v/v) with PI solution as outlined under Experimental Procedures. The percentage of cells under SubG1 (death cells) was presented in the table. Data are means ± SD of three or more separate experiments. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with control.

Control Control A B 42.23 µg/ ml 42.23 µg/ ml 120 63.35 µg/ ml 120 63.35 µg/ ml 100

y l) y l)

o 100 ilit r 80 o t ilit *** r * * t n n o *** o

C 60 ll Viab

*** C * * ll Viab e % ( e % 80 C ( 40 C *** 20 60 0 12243648 0 12243648 Time (h) Time (h)

Fig. 3.11 A comparison of the cytotoxicity of YCT with MGF on human hepatoma cells (HepG2) by MTT assay. Viability of HepG2 cells was determined after exposure to YCT (A) and MGF (B) at different concentrations for various time points by MTT assay. Experiments were conducted as described under Experimental Procedures. Data points are means ± SD of at least three independent experiments performed in five replicates. * p < 0.05, **p < 0.01, ***p < 0.001 as compared with control.

85 DISCUSSION

Antioxidant ability may help to explain the alleged beneficial health effects of

TCM, since reactive oxygen/ nitrogen/ chlorine species contribute to the pathology of

multiple human diseases (Halliwell and Gutteridge, 1999; Halliwell, 1999). Herbal

extracts, including TCM, would be expected to possess antioxidant properties because

they are rich sources of phenolic compounds (Zheng and Wang, 2001). Phenolic

compounds have been reported to function as antioxidants by virtue of their capability to

donate hydrogen to stabilize reactive and unstable free radicals (Rice-Evans et al., 1996).

Indeed, our results indicate that phenolic compounds may make a major contribution to

the antioxidant capacity of the TCM extracts examined; extract WN (Cratoxylum

cochinchinense) is particularly active in tests of total antioxidant activity. The extracts are likely to contain multiple antioxidant entities that react with ABTS•+ at variable rates.

Furthermore, at least some of the antioxidants present also performed well in an organic

phase using DPPH free radical as a detector. However, the inhibitory effect of extract

WN against lipid peroxidation in liposomes was not as good as PC (Psoralea corylifolia

L). This might be explained by the differences in the affinity of the active antioxidant

compounds toward the lipid and aqueous phases as well as the interface. Extract PC may

contain more lipophilic antioxidant components, which enter the liposomal bilayer more

readily. Extract WN also possess higher iron-binding activity as compared to the other

extracts.

86 Xanthine oxidase catalyses the oxidation of hypoxanthine to xanthine and then to

•− uric acid in the presence of O2 to yield O2 and H2O2 (Halliwell and Gutteridge, 1999). It has been reported that plant extracts inhibit xanthine oxidase (Sweeney et al., 2001) and

•− have O2 scavenging activity (Kirby and Schmidt, 1997; Lin et al., 1995). Our results

•− demonstrate that WN is the only extract with O2 scavenging activity (and that a modest one) but does not significantly affect XO activity, confirming that its inhibitory effect on

•− NBT reduction is due to O2 scavenging. This is an important control frequently omitted

from published studies.

The pattern of oxidized DNA base products arising from attack of different

reactive species has been shown to be different (Spencer et al., 2000; Whiteman et al.,

1997). For example, the formation of hypochlorous acid (HOCl) by activated neutrophils

in defence against microorganisms may cause DNA damage. In the presence of

superoxide anion radicals, HOCl may generate hydroxyl radicals that are highly reactive

and unstable (Candeias et al., 1994). Formation of nitryl chloride (NO2Cl) from HOCl

− and NO2 (in saliva and foods) has been demonstrated to damage calf thymus DNA

(Spencer et al., 2000; Whiteman et al., 1997). Our data show the ability of TCM to protect against DNA damage in vitro by HOCl.

Peroxynitrite (ONOO−), a highly reactive oxidizing and nitrating nitrogen species,

• •− is formed in vivo by the interaction of nitric oxide ( NO) and superoxide radicals (O2 )

(Beckman et al., 1994). Although peroxynitrite has a half−life of less than one second under physiological conditions, its protonated form (peroxynitrous acid, ONOOH) can

87 • + easily decay to give strong oxidants such as nitrogen dioxide ( NO2), nitryl cation (NO2 ) and hydroxyl radicals (•OH) (Beckman and Koppenol, 1996; Greenacre and

Ischiropoulos, 2001). These reactive species have gained much interest due to their

toxicity associated with numerous human diseases, including cardiovascular,

neurodegenerative, cancer, and inflammatory diseases (Greenacre and Ischiropoulos,

2001). Bicarbonate anion (HCO3¯) is present at a concentration of approximately 25 mM in the blood plasma and 12 mM in intracellular fluids (Denicola et al., 1996). Carbon dioxide can react rapidly with the peroxynitrite anion and thus modulate the oxidative activity of peroxynitrite. Furthermore, free and protein bound tyrosine undergo nitration when exposed to peroxynitrite at pH 7.4, forming 3-nitrotyrosine. This reaction in the biological system has been suggested to be a “marker” of peroxynitrite−dependent damage in vivo (Beckman et al., 1994). Recent data have provided evidence for protection against peroxynitrite toxicity by endogenous and exogenous radical scavengers

(Ketsawatsakul et al., 2000; Pannala et al., 1997; Heijnen et al., 2001; Schroeder et al.,

2001; Whiteman et al., 2002). Our results show that bicarbonate affected the amount of

nitration as well as the inhibition by YCT on ONOO−-mediated formation of 3-

nitrotyrosine, but not on the bleaching of PR. Most of the HCO3¯ in the plasma and cytosol exists as carbon dioxide with a normal plasma concentration of about 1.3 mM

(Denico et al., 1996). The formation of nitrosoperoxycarbonate anion (ONOOCO2¯)

− from the reaction of ONOO with HCO3¯/CO2 and ONOOCO2¯-derived intermediates,

• such as CO3¯ and •NO2 can also modulate the oxidative activity of ONOO¯. Amino acid

− tyrosine seemed to be more susceptible as a target molecule for ONOO / HCO3¯- mediated oxidation as compared with PR under our experimental conditions. These data

88 demonstrate the importance of using different assays to study ONOO¯-scavenging

activity of antioxidant compounds in the presence of HCO3¯/CO2 (Whiteman et al.,

2002).

The semi-purified extract (YCT) contains at least 90% mangiferin (MGF) as

identified using the HPLC-PDA and LC-MS systems. MGF has been reported to have

numerous pharmacological activities, including immunomodulatory activities

(Chattopadhyay et al., 1987), antioxidant activity (Moreira et al., 2001) and to act as a

potent biological drug with antitumor and antiviral effects (Guha et al., 1996). Recently,

•− MGF was reported to scavenge O2 generated by the xanthine-xanthine oxidase system with an IC50 of 8.2 ± 0.72 µM without inhibiting the XO activity at 1-100 µM (Leiro et al., 2003). However, this IC50 value does not correlate with the figure (Fig. 1A) as shown

in the article (Appendix A). The IC50 value should fall between 10 and 100 µM as shown

•− in the figure. Our results show that YCT was a better O2 scavenger than MGF. When

express as µM, MGF showed an IC50 value of 86.4 ± 3.1 µM (Table 3.4) under our

experimental conditions. With respect to ONOO¯- mediated formation of 3-nitrotyrosine

and PR bleaching, YCT was more effective than MGF as a scavenger of ONOO¯,

ONOOH, and/ or its decomposition species, whether or not HCO3¯ was added. YCT extract also has a low pro-oxidant ability as tested with bleomycin-Fe3+dependent DNA damage.

Protein modifications leading to the formation of AGEs are one of the consequences of hyperglycaemia (Vinson and Howard, 1996). The contribution of AGEs

89 towards aging and Alzheimer’s disease has received considerable attention (Smith et al.,

1994); and free radicals have been shown to participate in AGE formation (Chase et al.,

1991). YCT effectively inhibited the formation of AGEs. However, the mechanism of inhibition by YCT needs further investigation.

Extract WN, largely mangiferin, has efficient free radical scavenging activity, inhibits phospholipid peroxidation and HOCl-induced DNA damage, as well as scavenging ONOO− and inhibiting the formation of AGEs. It may serve important pharmacological roles related to the prevention of atherosclerotic lesions and coronary hearty diseases, which are oxidative stress related. However, YCT is more active than

MGF in several assays (Table 3.4). This may be explained by the present of extra compounds in YCT as indicated by peaks at tR around 3.5, 12.5 and 22.5 min (Fig. 3.6B) and some MS ion peaks at higher molecular weight (Fig. 3.7B) not seen with pure MGF.

Thus, YCT may contain other polyphenols (≤ 10%), which contribute to its ‘extra’

antioxidant activities as compared to MGF in the assays of phospholipid peroxidation,

•− − O2 scavenging, AGE formation on proteins and ONOO -mediated PR bleaching and 3-

nitrotyrosine formation. Results obtained from cell culture experiments also show that

YCT is more cytotoxic than MGF to HT29, HepG2 and Jurkat T cells and thus, support

the above statement. This may be because such compounds are highly-active antioxidants

and/or because they synergistically interact with mangiferin. Work to isolate and identify

them is continuing.

90 Pharmacokinetics of mangiferin in rat plasma after oral administration of Zi-Shen

pill and Suanzaoren decoction have been studied (Dai et al., 2004; Li and Bai, 2005). Zi-

Shen pill is prepared by mixing Rhizoma anemarrhenae (main ingredient), cortex

phellodendri, and Cortex cinnamomi with honey. Mangiferin, the major active

constituent of Rhizoma anemarrhenae and also in Zi-Shen pill may be used as a marker

compound to characterize this pill. The limit of quantitation (LOQ), the maximum plasma

concentration (Cmax), the time to reach peak concentration (tmax) and the apparent

elimination half-life (t1/2) of mangiferin in rat plasma after oral administration of the pill at a dosage of 19g/ kg were 0.163 µg/ ml, 5.187 ± 2.8 µg/ ml, 5 h and 12.2 h, respectively

(Dai et al., 2004). For Suanzaoren decoction, the LOQ, Cmax, tmax, and t1/2 of mangiferin

in the rat plasma were 0.536 µg/ ml, 10.5 ± 2.2 µg/ ml, 5.8 ± 0.4 h and 5.0 ± 0.3 h,

respectively (Li and Bai, 2005). Moreover, Lai et al. (2003) reported that free mangiferin

was detectable in the blood but not in the bile and brain dialysates of rats by

microdialysis coupled with microbore high-performance liquid chromatography and

tandem mass spectrometry. These authors reported a linear pharmacokinetic of free

mangiferin in rats blood following its intravenous administration at doses of 10-30 mg/ kg.

Several pharmacologic studies of mangiferin and mangiferin containing extracts such as Mangifera indica L. (Vimang®), Anemarrhena asphodeloides Bunge (Ichiki et

al., 1998; Miura et al., 2001), Bombax ceiba (Dar et al., 2005) and Salacia reticulata

(Yoshikawa et al., 2002a; Yoshikawa et al., 2002b) have been reported. For examples,

Mangifera indica L. extract showed significant antioxidative effects in cholesterol-fed

91 experimental rats by stimulating the activities of free radical scavenging enzymes (Anila

and Vijayalakshmi, 2003) and protective effects against 12-O-tetradecanoylphorbol-13-

acetate (TPA)-induced oxidative damage in serum, liver, brain as well as the production

of reactive oxygen species by peritoneal macrophages (Sánchez et al., 2000). Mangiferin isolated from Mangifera indica L. also exhibited antidiabetic and antihyperglycemic

activities by significantly lower fasting plasma glucose level in streptozotocin (STZ)-

diabetic rats and improved oral glucose tolerance in glucose-loaded normal rats,

respectively. Mangiferin also decreases the level of glycosylated haemoglobin in STZ-

diabetic rats and ameliorates STZ-induced cardiac and renal toxicity. Furthermore, mangiferin showed antihyperlipidemic and antiatherogenic activities in STZ-diabetic rats

(Muruganandan et al., 2002, Muruganandan et al., 2005) and chemopreventive effect in azoxymethane-induced colon carcinogenesis in rats (Yoshimi et al., 2001). However, in vivo study with Cratoxylum cochinchinense extract is very limited.

92 CHAPTER 4

RESULTS

4.1 Cytotoxicity of YCT by MTT assay and propidium iodide staining

YCT is an extract isolated on the basis of measurements of total antioxidant activity (Tang et al., 2004). However, antioxidants can sometimes be cytotoxic agents

(Wiseman and Halliwell, 1993). The cytotoxicity of YCT was tested on adherent cells and cells in suspension at different concentrations for up to 48 h using the MTT assay and propidium iodide staining, respectively. As shown in Fig. 4.1, YCT did not result in significant toxicity to normal cells of CL, MDCK and HAC and tumour cells of PC12 and HTB94. Interestingly, YCT shows a trend to modest pro-proliferative effects on

HAC, HTB94 and PC12 cells. However, this could be possibly an effect of an increase in cell metabolism after YCT treatment because we did not observe an increase in cell number as determined by trypan blue assay. Viability of CL and MDCK cells was equal or greater than 80% after 48 h exposure. However, the viability of other cell lines was decreased by YCT at concentrations of 42.23 and 63.35 µg/ ml although HEK cells were only affected at the higher concentration. Liver cell lines (HFL and HepG2) were most susceptible to YCT with viability reduced in a concentration- and time-dependent manner. After 48 h of exposure to YCT (63.35 µg/ ml), viability of HLF and HepG2 cells was decreased to 10 and 20%, respectively. This suggests that YCT might be hepatotoxic.

For suspension cells, the cell cycle was studied after fixing cells overnight with

70% ethanol using propidium iodide solution. In HL60, YCT induced 46.8 ± 1.1% cell

93 arrest as compared with control of 20.3 ± 2.0 % at 12 h (Table 4.1, p< 0.001). Incubating

Jurkat T cells, the human leukaemic T-lymphoblast cell line, in the presence of YCT also

induced cell arrest, reaching a maximum of 38.2 ± 4.8% as compared with control of 17.7

± 2.0% after 12 h and continuing up to 24 h (Table 4.1). The percentage of non-viable

cells increased significantly after 24 h and more than 60% of the cells were non-viable

after 48 h. Therefore, the mechanisms of YCT-induced cell death were studied in detail using Jurkat T cells as a model.

We noticed that there was substantial cell death after 24 h exposure to YCT extract. Therefore, for flow cytometic assay, only viable cells were collected for analysis by discriminating the death cells. Other assays that involved using cell lysates such as cytochrome c colorimetric and ATP assays, we normalized the reading with protein content.

94 Control 10.56 µg/ ml Fig. 4.1 21.12 µg/ ml 42.23 µg/ ml 63.35 µg/ ml

120

120 ) y ) y l l t 100

100 it o o r ili 80 il b H 80 ntr H ab i o ont 60 F V 60 E l C

C 40 l ll Via

L e e % 20 (% 40 K C ( C 0 20 012243648 012243648 Time (h)

110 140

) y ) l y l

it 120 100 o il r o t ilit r M n b ab

C i 100 o a

i 90 D ont V l C

L l C e 80 C ll V (% C e % 80

( K C 60 70 0 12243648 0 12243648

160 140

y 130 y l) l) it o o il 140 r 120 P ilit r t b t n b a

n C H i a

o 110 i o 120 V

C 1

A ll 100 C e ll V % ( e C 2 % C 90

( 100 C 80 80 012243648 0 12243648

120 120

y l) ) y

l H 100 H o lit it o i r il

110 t

T b 80 e n a ntr ab i i o o B 60 p V C l C 100 ll V l 9 G e e

% 40 (% ( C 4 C 2 90 20 0 12243648 0 12243648 Time (h)

140 y l)

it 120 o il r t

b 100 H n a i o 80 T V c l l 60 2 % e

( 40 C 9 20 0 12243648 Time (h)

95 Fig. 4.1 Effects of YCT on cells as shown by MTT assay. Viability of normal (HFL: human foetal liver, HEK: human embryonic kidney, CL: Chang liver, MDCK: Madin- Darby canine kidney, HAC: human articular chondrocytes) and cancer cell lines (PC12: rat pheochromocytoma, HTB94: human chondrosarcoma, HepG2: human hepatoma, HT29: colorectal adenocarcinoma) exposed to different concentrations of YCT for various time points as assessed by the MTT assay. Experiments were conducted as described in Experimental Procedures. Data points are mean ± s.d. of at least four independent experiments performed in five replicates.

Table 4.1 YCT-induced changes in HL60 and Jurkat T cells. HL60 Jurkat cells Time Sample SubG1 G2/ M SubG1 G2/ M (h) (M1) (M2) (M1) (M2) 12 Control 1.2±0.1 20.3±2.0 4.7±1.9 17.7±2.0 YCT 4.3±0.2 46.8±1.1*** 8.1±3.6 38.2±4.8*** 24 Control 1.5±0.3 20.0±1.8 5.0±2.3 19.6±2.4 YCT 22.4±1.6*** 32.8±0.7*** 22.2±2.4*** 38.1±4.1*** 48 Control 1.7±0.1 21±0.5 3.8±1.3 16.6±1.1 YCT 22.7±2.6*** 24.6±1.5 60.2±6.1*** 11.2±1.5

Cells were treated with 63.35 µg/ ml YCT for 12, 24 and 48 h. Cell cycle analysis was determined after staining cells fixed overnight with 70% (v/v) ethanol with PI solution as outlined under Experimental Procedures. The percentages of cell death and cell arrest are represented by SubG1 and G2/ M, respectively. Data are mean ± s.d. of three or more separate experiments. * p< 0.05, **p < 0.01, ***p < 0.001.

96 4.2 YCT-induced loss of viability in Jurkat T cells but not in normal lymphocytes

YCT-induced loss of viability in Jurkat T cells was time-dependent (Fig. 4.2A).

Loss of viability increased gradually after 7 h exposure to YCT and more than 60% of the

cells had lost viability after 48 h. However, YCT induced only about 10% loss of

viability in peripheral lymphocytes isolated from healthy volunteers after 48 h as

determined by cell cycle analysis (Fig. 4.2B). Trypan blue exclusion studies confirmed

decreased viability of Jurkat T cells after 12 h exposure to YCT at concentrations of

42.23 and 63.35 µg/ ml (Fig. 4.2C). However, control Jurkat T cells proliferated after 12

h and continued up to 48 h as compared with control at time zero.

Figure 4.2

A 70 Control YCT (63.35 µg/ ml) *** 60

) 50 1 G

b 40 u S (

s 30 ll

e *** C 20 %

10

0 0 1020304050 Time (h)

Fig. 4.2 YCT-induced loss of viability in Jurkat T cells. (A) Kinetics of loss of viability as assayed by cell cycle analysis of Jurkat T cells exposed to 63.35 µg/ ml of YCT. Fixed cells were analyzed using the flow cytometer after staining with PI solution. At least 104 cells were collected for flow cytometry analysis. * p< 0.05, **p < 0.01, ***p < 0.001.

97

17.1±2.0 B 3.5±1.8 4.7±1.9 12h 38.2±4.8***

3.9±1.9 8.1±3.6

5.0±2.3 19.6±2.4 24h 3.9±1.3 22.2±2.4*** 38.1±4.1***

10.0±2.0***

3.8±1.3 16.6±1.1

5.6±1.5 60.2±6.1*** 48h 11.2±1.5 12.3±2.8***

Peripheral Jurkat cells lymphocytes

Fig. 4.2 YCT-induced loss of viability in Jurkat T cells. (B) YCT-induced cell arrest and loss of viability in Jurkat T cells as compared to peripheral lymphocytes isolated from healthy volunteers. Where appropriate, the percentages of cells in each subpopulation (SubG1 (M1)-loss of viability, G2/ M (M2)-cell arrest) are indicated. Values above and below the marker are representative of control and treated (63.35 µg/ ml) samples, respectively. At least 104 cells were collected in each for flow cytometry analysis. * p< 0.05, **p < 0.01, ***p < 0.001.

98 C

Control YCT (42.23 µg/ ml) YCT (63.35 µg/ ml)

200 *** *** *** 150 ll e C

le 100

ab *** i

V *** *** % 50

*** *** 0 *** 0 6 12 18 24 30 36 42 48

Incubation Time (h)

Fig. 4.2 YCT-induced loss of viability in Jurkat T cells. (C) Cell viability of Jurkat T cells using trypan blue exclusion assay. For statistical analysis, YCT treatments were compared with control at each time point. For control, comparison was based on time zero. Experiments were conducted as described in Experimental Procedures. Data are representative of at least 3 independent experiments. * p< 0.05, **p < 0.01, ***p < 0.001.

4.3 Changes in cellular morphology

Exposure of Jurkat T cells to 63.35 µg/ ml YCT for 48 h resulted in the appearance of what seemed to be apoptotic bodies (Fig. 4.3A, arrow). By contrast, Jurkat

T cells in the absence of YCT proliferated (Fig. 4.2C and Fig. 4.3A-control). Analysis by laser confocal microscopy revealed chromatin condensation and nuclear shrinkage (Fig.

4.3B, arrow) in cells stained with Hoechst 33258. The control cells showed normal

nuclear morphology.

99 Figure 4.3

Control A YCT (63.35 µg/ ml)

10.0 µm

Control B YCT (63.35 µg/ ml)

10.0 µm

Fig. 4.3 Changes in morphology of Jurkat T cells after YCT treatment. (A) Cells after exposure to 63.35 µg/ ml YCT for 48 h were viewed under an inverted microscope. Arrows show apoptotic-like bodies. (B) Treated cells were washed, fixed with paraformaldehyde (4%, w/v), and stained with Hoechst 33258 (0.5 µg/ ml) before examining using an upright microscope as outlined under the Experimental Procedures. Arrow shows a cell with condensed and fragmented nucleus. Bar, 10 µm.

100 4.4 YCT-induced phosphatidylserine exposure and LDH release

Propidium iodide (PI) staining is generally accepted as a late apoptotic indicator.

Phosphatidylserine (PS) externalization, however, is an early event during apoptosis

(Martin et al., 1995). We therefore tested the exposure of PS using annexin V-FITC in combination with PI by two-colour flow cytometry. As shown in Figure 4.4A, 24.2 ±

6.5% cells showed higher annexin V-FITC binding as early as 1h after YCT exposure as compared with control. The percentage of cells with annexin V-FITC binding increased to 40.4 ± 2.6% at 3 h at 37 °C. The percentage of cell stained with PI, however, remained below 10% at 3 h, indicating that PS exposure occurs earlier.

101 Figure 4.4

A

0 h

1 h

3 h

Fig. 4.4 Phosphatidylserine exposure (PS) and release of lactate dehydrogenase (LDH). (A) Cells exposed to 63.35 µg/ ml YCT were stained with annexin V-FITC and propidium iodide (PI) for the differentiation between early and late apoptotic cells. At least 104 cells were collected for analysis. Experiments were performed as described in Experimental Procedures. Data are representative of at least 3 independent experiments.

102 We also investigated the release of LDH into the medium which is used as an

index of loss of cell membrane integrity. YCT-mediated LDH release was not significant

at 12 h at either concentration tested (Fig. 4.4B). However, LDH release was significant

(p< 0.001) at 63.35 µg/ ml after 24 h and increased at 48 h.

B

25 Control YCT (42.23 µg/ ml) 20 YCT (63.35 µg/ ml) ***

d e

as 15 e l

e *** ***

H R 10 D

% L 5

0 12 h 24 h 48 h

Fig. 4.4 (B) The release of LDH into the medium was determined with a commercial kit (Sigma, LD−50) as an indicator of membrane integrity. Cells lysed with Triton X-100 were considered as 100% LDH released in the medium. Experiments were performed as described in Experimental Procedures. Data are representative of at least 3 independent experiments. * p< 0.05, **p < 0.01, ***p < 0.001.

Pre-treatment of Jurkat T cells with caspase 3 inhibitor (Z-DEVD-FMK, 25 µM)

and caspase 9 inhibitor (Z-LEHD-FMK, 25 µM) for 1h before YCT exposure for 1h did

not decrease annexin V-FITC fluorescence (Fig. 4.4C). Moreover, these inhibitors had no

effect on the loss of viability of Jurkat T cells 12 h after YCT treatment (63.35 µg/ ml,

Fig. 4.4D).

103 C 250

g 200 *** n i l) d o r n t

n 150 Bi o V C f n i 100 x o e % n ( 50 An

0

l) T ) T ol M) M r m µ C C / Y Y ont + C µg 25 (25 µ nh nh + 35 nh ( i nh i i 3 i 9 63. 9 ( C C T C3 C YC

D 25 ) 1 20 *** G

ub

S 15 (

s l l

e 10 C 5 % of 0 ) ) ) ol l T r m µM C µM CT / Y Y ont + + C µg 25 25 nh nh 35 i i inh ( 3 inh ( 9 (63. 3 C 9 C T C C C Y

Fig. 4.4 (C & D) Effects of caspase-3 and -9 inhibitors on annexin V binding (C) and cell death (D) induced by YCT. Jurkat T cells were pretreated with caspase-3 inhibitor (Z- DEVD-FMK, 25 µM) and -9 inhibitor (Z-LEHD-FMK, 25 µM) for 1h before exposure to YCT for 1h (annexin V binding) and 12 h (cell cycle analysis) at 37 °C. At least 104 cells were collected for flow cytometry analysis. Experiments were performed as described in Experimental Procedures. Data points represent mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitors were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

104 4.5 Caspase activities and internucleosomal DNA fragmentation

The activity of caspase 9 in Jurkat T cells was significantly higher (p< 0.05) as compared with control at 12 h and 24 h after YCT treatment (42.23 and 63.35 µg/ ml) but not at 48 h (Fig. 4.5A). There was no significant difference between 12 and 24 h at both concentrations tested. Similar results were obtained for caspase 3 activity (Fig. 4.5B).

Western blot results of procaspase-3 and -7 did not show a decrease in intensity of proteins of both procaspases, indicating limited activation of caspase-3 and -7 (Fig.

4.5C). Furthermore, YCT-treated cells did not undergo internucleosomal DNA laddering as assessed by agarose gel electrophoresis of genomic DNA (Fig. 4.5D), but rather formed a DNA smear.

105 Figure 4.5

A Control 5 YCT (42.23 µg/ ml) y

t YCT (63.35 µg/ ml) ) 4 vi e i * s

a * e

r 3 c n * * ase 9 act d i 2 ol sp f ( a

C 1 ** *** 0 12 h 24 h 48 h

B 5 Control YCT (42.23 µg/ ml)

y 4

t YCT (63.35 µg/ ml) vi i 3 ease) cr n i 2 *** ase 3 act d l ** ** ** o sp f ( a

C 1

0 12 h 24 h 48 h

Fig. 4.5 YCT-induced activation of caspase-3, -7 and -9 but lack of internucleosomal DNA fragmentation. (A) Activity of caspase-9 as assessed with substrate Ac-LEHD-AFC using the SpectraMax190 microplate reader at excitation and emission wavelength of 400 and 505 nm, respectively. (B) Caspase-3 activity as assessed with substrate Ac-DEVD- AFC at excitation and emission wavelength of 390 and 520 nm, respectively. The unit of caspase activity was expressed as relative fluorescence unit per mg protein. Results were expressed as fold increase as compared with control. Results in (A) and (B) are mean ± s.d. of at least three independent experiments. * p< 0.05, **p < 0.01, ***p < 0.001.

106 C

12h 24h 48h

/ ml) / ml) / ml) / ml) / ml) / ml) g g g g g g µ µ µ µ µ µ 5 5 3 3 5 3 3 3 2 2 3 2

l 3.

3. 2.

2. 3. 2. l l o r o (6 (6 (4 o (4 (6 (4 r r ont ont ont C YCT C YCT YCT C YCT YCT

YCT Pro-caspase-3 32 kDa

β-actin

Pro-caspase-7 36 kDa

β-actin

D

12h 24h 48h

Fig. 4.5 (C) Cytosolic procaspase-3 and -7 in M (Kb) C Y C Y C Y Jurkat T cells treated with YCT (42.23 and 63.35 µg/ ml) for 12, 24, and 48 h. Western blot results using equivalent amount of protein (20 µg) from

each sample loaded onto each lane. (D) Agarose 10 8

6 gel DNA fragmentation. 1 µg of DNA isolated 5 4 from treated Jurkat T cells was loaded into 1% 3 2.5 2 (w/v) agarose gels and electrophoresed for 1-1.5

1.5 h at 100 V. Marker (M) used was 1Kb DNA

ladder (Promega, Madison, WI). C, control; Y, 1

YCT (63.35 µg/ ml). Experiments were conducted as described in Experimental Procedures. Data in (C) and (D) are representative of two independent experiments.

107 4.6 YCT induces rapid oxidative stress

YCT-induced generation of intracellular reactive oxygen species (ROS) was

monitored by the fluorescence emission of DCFDA. DCFDA is hydrolyzed by

intracellular esterases to a non-fluorescent product (DCFH) that is readily oxidized to

highly fluorescent 2’,7’-dichlorofluorescein (DCF) by ROS such as peroxyl, alkoxyl,

• • • NO2 , carbonate (CO3 ¯) and OH radicals (Halliwell and Whiteman, 2004). YCT induced

a rapid increase in DCF fluorescence within the first few minutes, reaching a maximum

at 30 min (Fig. 4.6A). 5 min after YCT exposure DCF fluorescence increased rapidly to

1.99 ± 0.6 RFI. The fluorescence remained at this level up to 24 h (data not shown).

Some phenolics have been reported to react with cell culture media to generate H2O2

(Long et al., 2000), but measurement of H2O2 showed that less than 5 µM H2O2 was detected in any of the media used after 1, 2 and 24 h exposure to YCT at 37 °C (Table

4.2). Hence, the effect of YCT is not an artefact of pro-oxidant events occurring in the cell culture media.

Figure 4.6

A 5 YCT (63.35 µg/ ml)

4 e c ) I

F 3 scen e R r n o u ea

l 2 f (m F C

D 1

0 0 10 20 30 40 50 60 70 80 90 100 110 120 130 140 150 160 170 180 190 200 210 220 230 240 250 260 270 280 290 300 Time (min)

108

B 10 9 ### 8 7 ) 6 *** se lls 5 e ea

c 4 *** cr +

### n 3 S i

d 2 l RO

o 1 f ( 0

l l) l) ) T ) T ) T ) T ro CT M C C M / m /m Y YC u YC g Y Y mM 1 ont + t+ (1 6+ 10 uM + C at 100 µ ) ( T+ I ( )6 N T M P PI 1000U C t ( Ro C I D D ( o IM 63.35 µ t R CN e( ( e( F Ca F CT K3 Y K3

Fig. 4.6 Reactive oxygen species (ROS) generation by YCT in Jurkat T cells. (A) Kinetics of YCT (63.35 µg/ ml) induced ROS generation in Jurkat T cells. ROS was determined using the flow cytometer with DCFDA (B) Effects of catalase, rotenone, potassium ferricyanide, indomethacin and diphenylene iodonium on ROS generation. Cells were pretreated with catalase (1000 U/ml) for 1h, and 30 min for rotenone (Rot, 100 µM), potassium ferricyanide (1 mM), indomethacin (IMT, 10 µM) and diphenylene iodonium (DPI, 1 µM) before exposure to YCT for 1h at 37 °C. At least 104 cells were collected for flow cytometry analysis. Experiments were performed as described in Experimental Procedures. Data points represent mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitors were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

Table 4.2 Generation of H2O2 (µM) in cell culture medium (RPMI) Without cells (37 °C) With Jurkat T cells (37 °C) Incubation [H2O2] in [H2O2] in medium + [H2O2] in [H2O2] in medium + time medium YCT (63.35 µg/ ml) medium YCT (63.35 µg/ ml) (µM) (µM) (µM) (µM) (h) 1 < 5 < 5 < 5 < 5 2 < 5 < 5 < 5 < 5 24 < 5 < 5 < 5 < 5

Note:. YCT extract at a final concentration of 63.35 µg/ ml was added to culture medium with and without Jurkat T cells and incubated at 37 ºC for various times. H2O2 concentration was then measured by O2 electrode assay as describe in Experimental Procedures. Data points represent mean of at least three independent experiments.

109 As mentioned before DCFDA detects a wide range of ROS such as peroxyl,

• • • alkoxyl, NO2 , and CO3 ¯ radicals, including species derived from H2O2 such as OH

(Halliwell and Whiteman, 2004). However, catalase did not reduce DCF fluorescence after exposure to YCT (Fig. 4.6B) and thus H2O2 was unlikely to be the origin of ROS induced by YCT. Externally-added catalase can decrease intracellular H2O2 levels

(Halliwell, 2003). In order to target the source of ROS, we treated the Jurkat T cells with ferricyanide (1 mM), a cell-impermeant extracellular electron acceptor that interacts with plasma membrane redox systems (Martinus et al., 1993), rotenone (100 µM), a specific

inhibitor of the complex I of the mitochondrial respiratory chain (Li et al., 2003), indomethacin (10 µM), an inhibitor of cyclooxygenase (Laneuville et al., 1994), and diphenylene iodonium (1 µM), an inhibitor of NADPH oxidases (Hampton and

Winterbourn, 1995). As shown in Figure 4.6B, ferricyanide but not rotenone, indomethacin and diphenyl iodonium, was able to decrease YCT-induced generation of

ROS significantly (p< 0.001). Caution should be exercised when using rotenone and

diphenylene iodonium because they can lead to oxidative stress (Li et al., 2003; Carriere

et al., 2003). Li and colleagues have shown that diphenylene iodonium could induce

•− mitochondrial O2 generation at a concentration as low as 1 µM for 10 min in HL60 cell line. Indeed, diphenylene iodonium did raise ROS production in control cells but the effect of rotenone on the DCFDA fluorescence was very small (Fig. 4.6B).

•− We also studied the accumulation of intracellular O2 using dihydroethidium

(DHE) (Halliwell and Whiteman, 2004; Zhao et al., 2003). It is now known that the oxidized product, oxyethidium, fluoresces at around 590 nm when excited at around 500

110 nm. An increase in fluorescence, presumably due to oxidation of dihydroethidium by

•− O2 , occurred within 5 min after exposure of Jurkat T cells to YCT (Fig. 4.7A).

Fluorescence remained at this level up to 12 h, started to increase rapidly up to 24 h and

remained at the higher level until at least 48 h. The presence of ferricyanide but not

rotenone and diphenyl iodonium significantly (p< 0.001) reduced the increased ethidium

fluorescence mediated by YCT (Fig. 4.7B). However, unlike the DCFDA studies (Fig.

4.6B, rotenone had no effect but diphenylene iodonium did raise ROS production), both

•− rotenone and diphenylene iodonium alone caused increases of O2 as compared to

control, suggesting that these agents might accelerate electron leakage from the

mitochondrial respiratory chain of Jurkat T cells (Li et al., 2003; Carriere et al., 2003). It

•− is thus uncertain what their effects on O2 generation by YCT.

There was no significant difference between the concentration of YCT at 42.23 and 63.35 µg/ ml on rise in ROS and superoxide anion radicals in Jurkat T cells as determined with DCFDA and DHE fluorescent probes. Therefore, the results of 42.23 µg/ ml YCT are not shown in Fig. 4.6A and 4.7A. Furthermore, we used the higher YCT concentration (63.35 µg/ ml) to study the effects of inhibitors because the changes due to

YCT on intracellular responses at 1h were more obvious as compared to the effects of

YCT at 42.23 µg/ ml.

111 Figure 4.7

A 10 YCT (63.35YCT µ(63.g/ m35l) µg/ ml) 9 + 8 ce 7

scen 6 e

r s ease) l o 5 u cr l cel n f 4 I m (X

u 3 i d

i 2 h t

E 1 0 0 3 6 9 12 15 18 21 24 27 30 33 36 39 42 45 48

Time (h)

B

4 lls e

c 3 *** +

*** e

e) 2 *** nc e c eas ### s

cr 1 e n *** i uor d l

l 0 f ) ) ) )

(fo l o l M T T M T r m C M C um / YC Y 1m Y (1µ ont µg ( + t+ + di C 6 )6 200 µ I ) Ro PI 35 N N t ( D DP hi C o

t C (63. e( e( R

E 3F 3F CT K K Y Fig. 4.7 YCT-induced superoxide generation in Jurkat T cells. (A) A time course •− relationship between O2 generation and YCT exposure. (B) Effects of potassium ferricyanide (1 mM), rotenone (Rot, 200 µM) and diphenylene iodonium (DPI, 1 µM) on •− O2 generation at 1h. Inhibitors were added into the cells for 30 min before YCT (63.35 µg/ ml) exposure for 1h at 37 °C. At least 104 cells were collected in each for analysis. Experiments were conducted as described in Experimental Procedures. Data points represent mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitors were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

4.7 YCT induced lipid peroxidation in Jurkat T cells

Intense oxidative stress can induce oxidative damage (Halliwell and Gutteridge,

1999). We therefore also measured lipid peroxidation using C11-BODIPY 581/591 and cis-

112 parinaric acid, probes that have a high affinity for the plasma membrane (Maier et al.,

2002). The unusual stability of the lipophilic C11-BODIPY 581/591 fluorescent dye makes

it a better probe as compared to cis-parinaric acid in the determination of lipid

peroxidation. The non-oxidized and oxidized products of C11-BODIPY 581/591 fluoresce red (~ 595 nm) and green (~ 520 nm), respectively, when excited at around 488 nm.

Exposure of Jurkat T cells to YCT resulted in a time- and concentration-dependent increase of lipid peroxides (Fig. 4.8A). The increase in lipid peroxidation as measured by

C11-BODIPY 581/591 probe was two-fold 1h after exposure to YCT (63.35 µg/ ml) and

remained at this level until about 7 h. The fluorescence increased rapidly again after 7 h

and remained elevated up to 48 h. Cis-parinaric acid is a natural polyunsaturated fatty

acid that readily incorporates into membranes and loses its fluorescence when oxidized

by ROS (Hedley and Chow, 1992). Its fluorescence intensity was monitored at excitation

of 325 nm and emission at 405 nm. A significant decrease in cis-parinaric acid

fluorescence was observed 5 h after treatment with YCT (63.35 µg/ ml) and continued to

decrease until 24 h, with no further change at 48 h (Fig. 4.8B). This assay was carried out

in the dark to reduce the possible photo-oxidation effect on cis-parinaric acid.

Figure 4.8 YCT (42.23 µg/ ml) A 10 YCT (63.35 µg/ ml) 9 8 cells +

7 e c ) e n s

e 6 sc

e 5 crea r in uo

l 4 ld o d f f 3 (

Re 2 n/ e e

r 1 G 0 0 4 8 12 16 20 24 28 32 36 40 44 48

Time (h)

113

B YCT (42.23 µg/ ml) 120 YCT (63.35 µg/ ml)

d 100

aci 80

c *** *** i *** U) *** *** ar 60 *** *** n i RF ar % *** *** (

p 40 - s 20 Ci

0 0 4 8 12162024283236404448

Time (h)

Fig. 4.8 Lipid peroxidation (LPO) induced by YCT in Jurkat T cells. (A) Time course and concentration studies of LPO with C11-BODIPY 581/591 fluorescent dye by flow cytometry and (B) Cis-parinaric acid (10 µM) using a Gemini Fluorescence microplate reader (Molecular Devices). * p< 0.05, **p < 0.01, ***p < 0.001.

The effects of various inhibitors of phospholipases (Tithof et al., 2002; Lu et al.,

2001) and lipoxygenases (Gao et al., 1996; Hogaboam et al., 1992; Vanderhoek and

Lands, 1973) on lipid peroxidation (LPO) induced by YCT were studied. Phospholipase inhibitors such as methyl arachidonyl fluorophosphonate (MAFP, 1 and 2 µM),

bromoenol lactone (BL, 2.5 and 5 µM), and oleyloxyethyl phosphorylcholine (OP, 5 and

10 µM) did not reduce the increased C11-BODIPY 581/591 fluorescence induced by YCT; instead, they caused significant increases in the fluorescence intensities (Fig. 4.8C).

Therefore, some of these agents might cause oxidative stress in Jurkat T cells, for examples BL and MAFP. However, incubation of Jurkat T cells with butylated hydroxytoluene (BHT), a synthetic antioxidant, decreased the fluorescence somewhat, as did different lipoxygenase inhibitors (baicalein monohydrate (BM), ODYA, and REV

5901 are inhibitors of lipoxygenase 12, 15, and 5, respectively) (Fig. 4.8C). However, we

114 interpret these data with caution since we do not exclude the possibility that some

lipoxygenase inhibitors may exhibit antioxidant properties in addition to direct inhibitory

actions against lipoxygenases.

C

#

5 # lls

e # ### ## ## c + 4

ce *** 3 ### ### ### ### ease) escen r

cr ** ** o n 2 u i l d l f y o f

p 1 ( i d o

B 0

11- ) l l) ) T ) ) T ) T ) T ) T C o T r CT M) C M M C M C M M CT M) CT M) C C / m µM Y µ C µ µ CT µ µ µM g µ Y µ Y Y 1 Y 2 Y Y ont 5 1 + Y 1 + .5 + 5 + 5 µ + Y + C T+ ( (2 + Y ( v 2 ( ( P+ P+ ( P 10 T ( A A v ( P P ( P ( 35 u H H BM Y BL BL BL F O P OP B BM Y Re Re AF AF AF O 3. B BL M M O 6 OD OD MA M T ( C Y

Fig. 4.8 (C) Effects of BHT, lipoxygenase inhibitors (BM, ODYA, and Rev are inhibitors of lipoxygenases 12, 15 and 5, respectively) and phospholipase inhibitors (bromoenol lactone (BL), MAFP and oleyloxyethyl phosphorylcholine (OP)) on LPO. Cells were pretreated with the inhibitors for 30 min before YCT exposure for 1h at 37 °C. At least 104 cells were collected for flow cytometry analysis. Experiments were performed as described in Experimental Procedures. Data points represent mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitors were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

2+ 4.8 Mitochondrial Ca overloading and ∆ψm dissipation

Rhod-2/ AM is a positively charged fluorescent dye that accumulates readily into the mitochondria. It exhibits a strong fluorescence upon binding Ca2+ at around 595 nm

when excited at around 550 nm. The use of Rhod-2/ AM fluorescent dye to measure

mitochondrial Ca2+ showed a concentration- and time-dependent increase in the number

of cells positive for Rhod-2 fluorescence (Fig. 4.9A). The increase at 6 h after YCT

(63.35 µg/ ml) exposure was five-fold as compared with control (p< 0.001), and

115 continued to increase to about twenty-fold at 24 h. This was accompanied by a rapid

decline in tetramethylrhodamine methyl ester (TMRM) fluorescence between 12 and 24

h, indicating a loss of mitochondrial transmembrane potential (∆ψm) (Fig. 4.9B). Like

Rhod-2, TMRM is also positively charged and accumulates in the mitochondria and is retained there by ∆ψm. Mitochondrial dysfunction may lead to a loss of TMRM

fluorescence in permeabilized cells.

Rhod 123 is a cell-permeant, cationic, fluorescent dye that is readily sequestered

by cells with active mitochondria and thus can be used to monitor ∆ψm of cells (Scaduto and Grotyohann, 1999). Another commonly used dye JC-1 is a fluorescent dye that is concentrated by respiring mitochondria. It exists as a monomer that fluoresces green and forms J-aggregates with red fluorescence at low and high ∆ψm, respectively (Reers et al.,

1991). The decrease in ∆ψm as determined with Rhod 123 and JC-1 occurred after 12 h

exposure to YCT and was concentration- and time-dependent (Fig. 4.9C & 4.9D).

Interestingly, modest increases in ∆ψm were detected at 6 and 12 h at both concentrations of YCT as determined with JC-1 dye, but not with the other probes. We do not know the reason for this. It may be an early phenomenon as a result of Ca2+ influx into the mitochondria.

Furthermore, we think it is important to study the response of YCT treatment after

12 h because the fall in mitochondrial transmembrane potential occurred after 12 h as shown in Fig. 4.9. Moreover, cell viability at 12 h is 89 ± 12 % which is not significantly different as compared to control (Fig. 4.2).

116 Figure 4.9

A C 40 Control Control e

c YCT (42.23 µg/ ml) YCT (42.23 µg/ ml) *** n e

c 70 YCT (63.35 µg/ ml) s 30

) YCT (63.35 µg/ ml) e e

lls 60 *** s e a

uor *** c l ll) e 50 r +

*** e f

c 20 *** 3 C 2

f 40 in *** o ld

hod 2 30 % o ( f *** R ( 10 *** *** *** hod 1 20 ***

R *** 10

of s 0 0 Los 0 6 12 24 48 6 12182430364248 Time (h) Time (h) B D YCT (42.23 µg/ ml) YCT (42.23 µg/ ml) 160 YCT (63.35 µg/ ml) YCT (63.35 µg/ ml) * ) 120 ) 140 l ) o l r ** ce o 535 120 r n 100 ont **

ont 100 C 585/ * esce c o r

*** i * of o 80 of at

80 *** U u l U R f F (

FR 60 *** 1

- R % RM 60 *** (

JC 40 M % *** *** ( T 20 40 0 6 12 18 24 30 36 42 48 0 6 12 18 24 30 36 42 48 Time (h) Time (h)

2+ Fig. 4.9 Mitochondrial Ca levels and ∆ψm in Jurkat T cells. (A) Kinetics of rise in mitochondrial Ca2+ of Jurkat T cells exposed to YCT as determined with Rhod-2/ AM fluorescent dye loaded to a final concentration of 500 nM for 30 min at 37 °C. (B) Loss of ∆ψm as determined with TMRM (50 nM) using the Gemini Fluorescence microplate reader. (C) and (D) Time course study of ∆ψm with Rhod 123 and JC-1 using the flow cytometer and Gemini Fluorescence microplate reader, respectively. At least 104 cells were collected for flow cytometry analysis. Experiments were conducted as described in Experimental Procedures. Results are mean ± s.d. of at least three independent experiments. * p< 0.05, **p < 0.01, ***p < 0.001.

117 2+ 4.9 Effects of ruthenium red on mitochondrial Ca and ∆ψm

Ruthenium red (RuR) is an inhibitor of mitochondrial Ca2+ uniporter through which Ca2+ passes from the cytosol into the mitochondria (Gunter et al., 2000). Under

certain conditions, the uniporter may act differently to allow the release of Ca2+ back to the cytosol. Jurkat T cells treated with YCT in the presence of RuR (25 µM) showed

2+ decreased mitochondrial Ca levels (Fig. 4.10A) and smaller falls in ∆ψm (Fig. 4.10B).

However, RuR did not significantly prevent YCT-mediated cell arrest at 7 h or loss of viability at 24 h (Fig. 4.10C).

Figure 4.10 Fig. 4.10 Effects of ruthenium red A Control (RuR, 25 µM) on mitochondrial Ca2+ 40 YCT (63.35 µg/ ml) (A), ∆ψm dissipation (B), cell arrest RuR (25 µM)+ YCT and cell death (C). Jurkat T cells were ls pre-incubated with RuR for 30 min 30 cel + before YCT exposure for 24 h. At ce least 104 cells were collected for flow ease) 20 *** cr cytometry analysis. Experiments were escen r in o

d conducted as described in u l l f o ### f

( Experimental Procedures. Results are

2 10 d

o mean ± s.d. of at least three h

R independent experiments. For 0 statistical analysis, * YCT and 24 h inhibitor alone was compared with control; # YCT in the presence of inhibitor were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

118

B 12 Control YCT (63.35 µg/ ml) 10 RuR (25 µM) RuR+ YCT ce) 8 *** escen ls r

o 6 ease) u cel l cr + f

### n 1 i al -

t 4 ld o JC o f /T ( 2 een r G ( 0 24 h

C

G2/M Sub G1 50

*** 40

*** lls 30 e C f 20 % o

10

0

rol l) M) nt / m µ CT o g 5 + Y C 5 µ (2 R .3 R Ru (63 Ru T YC

119 4.10 YCT-mediated changes in cytochrome c and ATP levels

Fig. 4.11A shows a calibration curve for the determination of cytochrome c. A significant increase of cytochrome c was observed in the cytosol 12, 24 and 48 h after exposure to YCT (63.35 µg/ ml) (Fig. 4.11B) as compared with control. The decrease in mitochondrial cytochrome c was significant only after 48 h. Surprisingly, there appeared to be a rise in mitochondrial cytochrome c at 12 h (p< 0.01) in YCT-exposed cells, and the reason for this requires further studies. The time-dependent release of mitochondrial cytochrome c to the cytosol was also detected by western blot (Fig. 4.11C). The presence of potassium ferricyanide stopped the release of cytochrome c, suggesting that the collapse of ∆ψm (and possibly the opening of the MPT pore) and the subsequent release of cytochrome c is a downstream event. Cellular ATP levels were significantly lower at

24 h but not 12 h after incubation with YCT (Fig. 4.11E), probably due to mitochondrial depolarization as determined with TMRM, Rhod 123 and JC-1 (Fig. 4.9B, 4.9C, 4.9D).

The drop in ATP levels in control cells at 24 and 48 h as compared to 12 h was unexpected and might indicate a decrement of nutrients in the medium to support ATP generation. A calibration curve freshly prepared for the determination of ATP is shown in

Fig. 4.11D.

120

Figure 4.11

A 2.0 y = 0.1746x 2 1.5 R = 0.9865 m n 0

45 1.0

t a

D 0.5 O

0.0 0246810 Cytochrome c conc (ng/ ml)

B Cytosolic cytochrome c Mitochondrial cytochrome c Control Control 800 YCT (63.35 µg / ml) 1500 YCT (63.35 µg/ ml)

1200 ** ) ) **

600 n n c c i i

*** e e e t e *** 900 ot m m o o pr r pro

r 400

h g h

g **

c 600 c o o m m t t / g/ g 200 Cy Cy n

n 300 ( (

0 0 12 24 48 12 24 48

Incubation Time (h) Incubation Time (h)

121

YCT Extract (63.35 µg/ ml) C K3Fe(CN)6 (1 mM) Time (h) C 1 3 6 10 20 C 10

Cyto solic 12kD cytochrom e c

β-actin

12kD Mitochondrial cytochrome c

Cox IV

Fig. 4.11 YCT-mediated changes of cytochrome c and ATP levels in Jurkat T cells. (A) A calibration curve of cytochrome c. (B) Cytosolic and mitochondrial cytochrome c contents were determined with a commercial ELISA kit. Concentrations of cytochrome c were determined from the standard curve of cytochrome c and results were expressed as ng/ mg protein from two independent experiments each performed in duplicate. (C) Cytosolic and mitochondrial cytochrome c were also evaluated by Western blotting using cytochrome c monoclonal antibody. Jurkat T cells after treatment were fractionated using mitochondrial fractionation kit according to the supplier’s protocol. Equal loading of protein samples (40 µg) was determined using β-actin and COX-IV for cytosolic and mitochondrial fractions, respectively. Western blot data are representative of at least three independent experiments. * p< 0.05, **p < 0.01, ***p < 0.001.

122 D 150000

.) y = 2665.3x

.u 2

a R = 0.9982 ( e

c 100000 n e c s e min

u 50000 L y l f e r i F

0 010203040

ATP conc. (nmol/ ml)

E Control YCT (42.23 µg/ ml) 20.00 YCT (63.35 µg/ ml) 16.00 n i e ot

] 12.00 pr P * g T *** [A

m ** / 8.00 *** ol

nm 4.00

0.00 12 h 24 h 48 h

Fig. 4.11 (D) A typical calibration curve freshly prepared for each measurement of cellular ATP. (E) ATP levels were determined with a luminescence assay and results were expressed as nmol ATP/ mg protein. Experiments were conducted as described in Experimental Procedures. Results are mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT at different concentrations were compared with control. * p< 0.05, **p < 0.01, ***p < 0.001.

123 4.11 Intracellular glutathione content

Changes in reduced glutathione (GSH) are closely related to apoptosis (Macho et

al., 1997). The level of GSH after YCT exposure was not significantly changed for up to

48 h as compared with control (Fig.4.12B), indicating that YCT-induced loss of viability is not associated with an early fall in GSH. Like cellular ATP, the drop in GSH levels in control cells at 24 and 48 h as compared to 12 h might indicate a decrement in essential nutrients in the medium to replenish GSH pool. Incubating cells with N-acetyl-L-cysteine

(NAC, 1 mM) for 1h raised GSH levels in control cells as well as cells treated with YCT for 12 h (Fig. 4.12C) but did not inhibit YCT-induced ROS generation at 1h or cell death at 12 h (Fig. 4.12D and E). Therefore, the fall in GSH does not cause cell death but accompanies it. Intracellular GSH was determined from a freshly prepared calibration curve of GSH as shown in Fig. 4.12A.

Figure 4.12

A 500 y = 12.661x 2 y R = 0.9986 t 400 i s n .) e t .u n a 300 i ( ce nm

0 cen 2 200 s 4 e t r a o u l

F 100

0 0 5 10 15 20 25 30

GSH conc. (nmol/ ml)

124

Control YCT (42.23 µg/ ml) B YCT (63.35 µg/ ml) ETP (20 µM) 18 16 n) i

t e 14 n ot e 12 ** pr

ont 10 g

C 8 m / H l 6 S o ** G

m 4 n

( 2 *** 0 12 h 24 h 48 h

Fig. 4.12 YCT-mediated loss of cellular GSH and the effects of inhibitor on ROS generation and cell death. (A) A typical calibration curve for the determination of GSH. (B) A concentration- and time-dependent loss of cellular GSH in Jurkat T cells after YCT exposure. Etoposide (20 µM) was used as a positive control.

C

YCT+ NAC *** 12h YCT (65.35 µg/ml)

NA C ( 1 mM) *** 1h Control

01020304050 GSH content (nmol/ mg protein)

125 4 D *** )

3 se lls ea e 2 c cr + n S i

d 1 l RO (fo 0 ) ol l ) T tr m M C n g/ m Y o µ 1 + C ( C 35 C A . A N 63 N ( T C Y

E 16 14 ) 12 *** G1 -

b 10 u 8 (S

lls 6 e

c 4 % 2 0

ol l) ) T tr m M C n g/ m Y o µ 1 + C ( C 35 C A . A N 63 N ( T C Y

Fig. 4.12 Effects of N-acetyl-L-cysteine (NAC) on cellular GSH (C) and YCT-induced increased in ROS (D) and cell death (E). Jurkat T cells were pre-treated with NAC (1 mM) for 30 min followed by the addition of YCT (63.35 µg/ ml) for 1 h and 12 h at 37 ºC for ROS and cell cycle analyses, respectively. At least 104 cells were collected for flow cytometry analysis. Experiments were conducted as described in Experimental Procedures. Results are mean ± s.d. of at least three independent experiments. * YCT and inhibitor alone were compared with control; # YCT in the presence of inhibitors was compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

126 4.12 YCT-mediated mobilization of extracellular Ca2+

Oxidative stress and changes in intracellular Ca2+ are intimately related (Halliwell

and Gutteridge, 1999). We therefore used the Ca2+-sensitive Fluo3/ AM fluorescent dye

to monitor the rise of intracellular Ca2+ by flow cytometry and spectrofluorimetry. The

binding of Fluo3 to Ca2+ exhibits a strong fluorescence at around 526 nm when excited around 506 nm.

Exposure of Jurkat T cells to YCT induced a time-dependent rise in intracellular

Ca2+. The fluorescence intensity increased rapidly 5 min after exposure to YCT and

continued elevated for at least 7 h (Fig. 4.13A). Moreover, the rise in intracellular Ca2+ showed a bimodal phenomenon at 3 h after YCT treatment. We do not know the reason for this. It may be a phenomenon of the release of Ca2+ from intracellular stores such as the ER and mitochondria.

Figure 4.13

A

YCT (63.35 µg/ ml) 5

e c 4 ) I F escen r R 3 o u l ean 2 3 f (M o u l

F 1

0 0 25 50 75 100 125 150 175 200 225 250 275 300 325 350 375 400 425 450 Time (min)

Fig. 4.13 YCT-mediated rise in cytosolic Ca2+ in Jurkat T cells through a non-selective cation channel. (A) Time course of intracellular Ca2+ levels determined using the flow cytometer with Fluo3/AM fluorescent dye.

127 To further confirm the rise in intracellular Ca2+ mediated by YCT, we studied the

increase of fluorescence using a spectrofluorometer. Jurkat T cells were pre-loaded with

Fluo3/ AM fluorescent dye to a final concentration of 5 µM before YCT treatment. The

increase in fluorescence was detected 60 sec after YCT was added to the cells suspended in PBS containing 1 mM CaCl2 as shown in Figure 4.13B (top trace). Experiments were also carried out in CaCl2-free PBS (Fig. 4.13D). Little increase in fluorescence was

observed when YCT was added to cells suspended in CaCl2-free PBS. However, a rapid increase and decrease in fluorescence respectively were detected when CaCl2 (1 mM) and

EGTA (2 mM) were added to the cell suspension. No changes in fluorescence were

observed in cells without added Fluo3/ AM fluorescent dye when CaCl2 and EGTA were added (data not shown). Our data suggest that the rise in Ca2+ is due to influx from the

medium and that pre-treatment with YCT might cause the opening of ion channels

(possibly due to the accumulation of ROS), which allow rapid influx of Ca2+ if extracellular Ca2+ is present.

128 B

YCT

YCT

FA 100µM

200µM

300µM Fluo3 fluorescence intensity

Time (sec)

Fig. 4.13 (B) Kinetic study of Ca2+ C mobilization using a Perkin-Elmer LS-50B 4 lls spectrofluorometer. Jurkat T cells were e

c ***

+ preloaded with Flou3/ AM fluorescent dye )

3 e

se before YCT exposure. Flufenamic acid (FA), c n ea

e 2 a non-selective cation channel blocker was c

cr ### s n

e tested at various concentrations for its i r 1 2+ d

l inhibitory effect on the rise of cytosolic Ca . uo l (fo f 0 (C) Effect of FA (20 µM) and indomethacin (IMT, 10 µM) on cytosolic Ca2+ using the uo3 l ) ) T ) T ro M M Fl t ml C C µ Y flow cytometer. Jurkat T cells were on g/ Y 0 C µ 20 µ + 1 + ( ( T pretreated with FA (20 µM) for 30 min 35 A FA M 3. F MT I 6 I ( before YCT exposure for 1h at 37 °C. * p< T # C Y 0.05, **p < 0.01, ***p < 0.001 or p< 0.05, ## < 0.01, ###p < 0.001.

129 D

EGTA

YCT CaCl2 Fluo3 fluorescence intensity

Time (sec)

Fig. 4.13 (D) Kinetic study on Ca2+ mobilization in Jurkat T cells suspended in PBS 2+ (Ca free). YCT was added, followed by CaCl2 (1 mM) and subsequently the addition of Ca2+ chelator EGTA (2 mM). At least 104 cells were collected for flow cytometry analysis. Experiments were performed as described in Experimental Procedures. Results are mean ± s.d. of at least three independent experiments. For B and D, data are representative of three independent experiments. For statistical analysis, * YCT and inhibitor alone were compared with control; # YCT in the presence of inhibitor were compared with YCT alone.

130 4.13 Ca2+ influx through a non-selective cation channel?

To gain insight into the channels that might be involved in the rise in Ca2+, we

investigated the effect of YCT on Jurkat T cells in the presence of various channel

blockers. As shown in Figure 4.13B, Ca2+ influx was attenuated by flufenamic acid (FA),

a non-selective cation channel blocker (Ghamari-Langroudi and Bourque, 2002), in a

concentration-dependent manner. The increase in fluorescence was stopped by FA (300

µM) added 10 min before exposure to YCT. This suggests that YCT activated the non-

selective cation channels of Jurkat T cells, leading to Ca2+ influx. When the effect of FA was studied using a flow cytometer, a much lower concentration of 20 µM was needed to reduce Ca2+ influx (p< 0.001) mediated by YCT (Fig. 4.13C). Indomethacin at a

concentration of 10 µM did not inhit Ca2+ influx.

In contrast to flufenamic acid, the voltage-dependent Ca2+ channel and Na+/ Ca2+ exchanger blockers did not stop YCT-induced Ca2+ influx, suggesting that these ion channels are not involved. Cytofluorimetric studies on channel blockers of voltage- dependent Ca2+ channel (i.e. nifedipine and nicardipine) and Na+/ Ca2+ exchanger (i.e.

bepridil and benzamil) showed no inhibition of Ca2+ influx as determined with Fluo3/

+ 2+ AM fluorescent dye (Fig. 4.13D). The involvement of Na / Ca exchanger and KATP channels in YCT-induced cell death was excluded because no significant changes of

+ + [Na ]i and [K ]i were detected with SBFI and PBFI fluorescent dyes, respectively, as

determined by flow cytometry (Fig. 4.13E).

131 D s l l 80 ###

ce 70 ### + ### 60

ce *** 50 n ### e

) 40

c # *** *

s 30

(% *** e 20 *

uor 10 l 0 l ) ) T ) T ) T ) T ) T ) T ) ) o M T M T r ml C C C M C C C

uo3 f µM µM µ µ / µM Y Y Y µ Y µM YC µM YC Y Y ont g 0 + + 0 0 + 0 + + + C (5 F F (50 C+ 0 C+ 1 5 10 Z 50 Z Fl 100 1 ( ( ( ( NI ( NI NI NI l EP il EP il N il N ne ine idi B id B 63.35 µ pi ne ip ine ( r r am BE am BE ( di pi d ip p z z e di r d ep n n T f e a r B Be e e C Ni f c a B B Y Ni Ni ic N

E PBFI SBFI

y t

i 160 ) ns 140 l e t o

r 120 n i

e 100 ont c nc 80 e c of 60 s % e

( 40 20 uor

Fl 0 0 5 10 20 30 60 180 300 720 1440 2880 Incubation time (min)

Fig. 4.13 (D) Effects of voltage-dependent Ca2+ channel and Na+/ Ca2+ exchanger blockers on YCT-induced Ca2+ influx. Jurkat T cells were pre-treated with nifedipine and nicardipine (50 and 100 µM) and bepridil and benzamil (10 and 50 µM) for 30 min followed by the addition of YCT (63.35 µg/ ml) for 3 h at 37 ºC. (E) A time course study of cellular changes in potassium and sodium using PBFI and SBFI fluorescent dyes, respectively. At least 104 cells were collected for flow cytometry analysis. Experiments were conducted as described in Experimental Procedures. Results are mean ± s.d. of at least three independent experiments. * YCT and inhibitor alone were compared with control; # YCT in the presence of inhibitors was compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

132 4.14 The relationship between ROS generation and Ca2+ influx

We investigated the relationships between ROS generation, lipid peroxidation

(LPO) and the increase in Ca2+ influx upon YCT treatment. Incubating cells with flufenamic acid (FA, 20 µM) for 30 min before exposure to YCT for 1h led to significant decreases in intracellular Ca2+ (Fig. 4.14B) but has no significant effect on ROS (Fig.

4.14A) or LPO (Fig. 4.14C). When cells were treated with YCT in PBS (Ca2+ free) for 15 min, the increase in DCF fluorescence was significantly higher as compared with control

(Fig. 4.14D, p< 0.001). It follows that the YCT-induced increase in ROS is not dependent on the influx of Ca2+. The antioxidant, BHT (5 µM) decreases LPO markedly and ROS somewhat but does not decrease Ca2+ influx, suggesting that the Ca2+ influx is probably not caused by LPO . Potassium ferricyanide, when added into the medium reduces the intracellular Ca2+ and ROS (p< 0.001) and LPO (p< 0.05), significantly. The concentrations of FA, BHT and potassium ferricyanide used in these assays did not affect cell viability.

133 Figure 4.14

4 A ***

s l 3 ### ### cel

crease) +

n 2 S i d l RO o f ( 1

0

+ ###

5 e c 4 ***

ease) s l escen 3 cr r n u cel i o l d

l 2

f ### ### o 3 f u (

o *** ** l 1 F 0 s l

l 5 ce + 4

ce *** #

se) 3

cen ** s ea *** e r cr 2 o n

u ### i l d

f * l 1 y o f p ( i d 0 Bo - l l) ) ) T ) T o T M µM µM YC m YC YC ontr g/ m + 1 + + C11 C µ 20 6 (5 FA 6 ( T HT .35 A ( N) B 3 F N) (C BH (C Fe T (6 Fe 3 C K Y K3

134 D 50 ***

ls 40

cel ***

30

+ S

O 20 R f 10

% o 0 ) ol l r BS CT / m P Y ont g C S+ PB 63.35 µ ( T C Y

Fig. 4.14 Relationships between YCT-mediated ROS generation (A), Ca2+ influx (B) and LPO (C) in Jurkat T cells. Compounds studied (flufenamic acid (FA, 20 µM), potassium ferricyanide (K3Fe(CN)6, 1 mM) and butylated hydroxytoluene (BHT, 5 µM)) were added into the cell suspension for 30 min before YCT (63.35 µg/ ml) exposure for 1h at 37 °C. (D) YCT-induced ROS generation in Jurkat T cells in Ca2+-free PBS. Jurkat T cells were incubated with YCT (63.35 µg/ ml) for 15 min and ROS generation was determined using the flow cytometer with DCFDA fluorescent dye. At least 104 cells were collected for flow cytometry analysis. Experiments were performed as described in Experimental Procedures. Results are mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitors were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

4.15 Ferricyanide inhibit YCT-induced apoptotic signals 24 h after treatment

We measured parameters of cell injury and death in the presence of potassium

ferricyanide using the flow cytometer. Annexin V-FITC binding was tested at 3 h

•− incubation with YCT. As shown in Fig. 4.15, all parameters examined (ROS, O2 , C11-

Bodipy581/591, cis-parinaric acid, Fluo3, Rhod2, TMRM, Rhod 123, JC-1, annexin V and cell death) were diminished significantly by the presence of ferricyanide (1 mM) added

30 min before YCT treatment. The concentration of potassium ferricyanide used in these experiments has been found to have no quenching effect on fluorescent dyes.

135 Figure 4.15

lls

5 s 10 7 l e l

) *** c ce +

se *** 6 + e a 4 *** 8 e c e

) 5 e) cr e en enc n s c c I a 3 ### 6 s ### eas 4 e X e r r r es ( c ### c s l uo l l

In 3 uor In l 4

2 f X f ce X y (

( *** + p 2 um S di o

1 di 2 B

hi 1 RO t 1- E 0 0 1 0 C e

120 lls 10 *** 8 e lls enc c

e *** c +

100 c

e ### + 8 es e) ) l e

e) 6 ### o enc uor 80 eas *** c enc fl

r 6 eas c d ontr r i es

c 60 es

4 Inc

a ### Inc our X c

of C 4 i ( X uor ** 40 ( % ( nar i 2 2 uo3 fl hod2 fl 20 l par R F -

s 0 0 0 Ci

120 60 6 e) s s s l l o

100 enc e l c ce + ### *** *** )

e l es ) c o enc

80 s 40 l n l c e uor

e 4 ntr *** fl c es o

s C e) f

C 60 e tal r our o o ### o of eas u T r l

40 (% 20 % f c ( n

M ### I

een/ 2 R r 20 X ( *** G hod 123 fl TM

( *** R s

0 0 l l

e c

+

60 40 1 0 *** *** - l l) )

JC o

) M r m CT

s m Y

l l 50 ont g/ 1 + ) e C ( 6

1 6 C 30 35 µ ) N) N C

of 63. e(

40 (C ( F ub G Fe %

S CT 3 K3 Y K ( s

30 l 20 ng ( l i e d

n ### h C bi 20 V eat

n 10 * i

10 ** ### D % nnex A 0 0 l o l) ) T ol l) ) T r m M C r M t / m t / m m C on g Y on g Y C µ (1 6+ C µ (1 + 5 6 ) 5 6 )6 .3 ) N .3 ) N 3 N (C 3 N C 6 (C e 6 (C e( ( e F ( e T F 3 T F 3F C 3 K C 3 K Y K Y K

136 Fig. 4.15 Effect of an extracellular electron acceptor, potassium ferricyanide, on YCT- induced cell death. Jurkat T cells were pre-treated with potassium ferricyanide for 30 min before exposure to YCT for 24 h. For annexin V-FITC binding, cells were treated for 3 h with YCT. Experiments were performed as described in the Experimental Procedures. At least 104 cells were collected for flow cytometry analysis. Results are mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitors were compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

Fig. 4.16

50 )

1 ***

G 40 b u

S 30 ( ### lls

e 20 C f 10

% o 0

l ) T ro ml M) nt g/ m YC o µ C (1 6+ 35 )6 N) 3. N 6 C ( (C e( e F T F C K3 Y K3

Fig. 4.16. Effect of potassium ferricyanide on YCT-induced cell death in HepG2 cells. HepG2 cells were pre-treated with potassium ferricyanide for 30 min before exposure to YCT (63.35 µg/ ml) for 24 h. Experiments were performed as described in the Experimental Procedures. At least 104 cells were collected for flow cytometry analysis. Results are mean ± s.d. of at least three independent experiments. For statistical analysis, * YCT and inhibitors alone were compared with control; # YCT in the presence of inhibitor was compared with YCT alone. * p< 0.05, **p < 0.01, ***p < 0.001 or # p< 0.05, ## < 0.01, ###p < 0.001.

137 DISCUSSION

It is frequently assumed that antioxidants are always beneficial. Here we show

that an antioxidant extract from Cratoxylum cochinchinense (YCT) induces cell death in

certain cell types. Although we have focussed on Jurkat T cells here, mechanisms of cell

death induced in tumour liver cells (HepG2) appear to be similar and are also suppressed

by potassium ferricyanide (Fig. 4.16). Apoptosis is a highly regulated cell death

programme that is induced in cells by a suicide response due either to unfavourable growth conditions or exposure to pro-apoptotic external stimuli (Halliwell and

Gutteridge, 1999; Petit et al., 1995). Some characteristics of apoptotic cells include exposure of PS on the outer plasma membrane (Martin et al., 1995), changes in mitochondrial activity, activation of caspases, chromatin condensation, DNA laddering, and cell shrinkage (Tiku et al., 2000). Eventually, apoptotic cells are removed by neighbouring cells without release of cellular content into the extracellular space (Long et al., 2000). YCT was found to induce loss of viability in a range of cells (i.e. HEK, HFL,

HepG2, HT29, HL60 and Jurkat cells) as determined by both MTT and SubG1 assays but not in normal lymphocytes. Its ability to target Jurkat T cells but not IL-2 exposed normal lymphocytes might be of therapeutic interest, but its effects on liver cells suggest a potential problem with hepatotoxicity.

We studied the mechanism(s) of YCT-induced cell death in Jurkat T cells. The data are consistent with the occurrence of apoptosis by several assays such as exposure of

PS, release of cytochrome c, collapse of ∆ψm and chromatin condensation. Nonetheless, loss of cell viability occurred with limited activities of caspase-9 (1-2 fold), and -3 (1

138 fold), and no internucleosomal DNA fragmentation. Moreover, the presence of caspase

inhibitors did not block the loss of cell viability. Similar programmed cell death

phenomena have been reported (Jambrina et al., 2003; Sperandio et al., 2000) and are

termed ‘paraptopsis’. The limited activities of caspase-3, -7 and -9 might be explained by

•− the highly oxidizing environment (i.e. YCT-induced O2 and lipid peroxides), which

could cause their inactivation (Chandra et al., 2000). H2O2 has been reported to suppress

the activation and activity of caspases, possibly through oxidation of the cysteine residues

(Nicholson et al., 1995; Nobel et al., 1997; Hampton and Orrenius, 1997). The failure to

detect internucleosomal DNA fragmentation also indicates that what we observe is not

“pure” apoptosis.

ROS are produced in all mammalian cells due partly to normal cellular

metabolism and also when cells are exposed to certain exogenous agents (Finkel and

Holbrook, 2000). ROS production is also closely related to apoptosis (Macho et al., 1997;

Macho et al., 1999; Dirsch et al., 1998; Macho et al., 2003). Our results show that YCT rapidly induced both intense oxidative stress (ROS) and a rise in intracellular Ca2+.

However, the rise in ROS was independent of the increase in intracellular Ca2+. Thus, incubating cells with YCT in Ca2+ free buffer did not decrease DCF fluorescence. In fact,

YCT increased the DCF fluorescence. In addition, flufenamic acid blocked the rise in intracellular Ca2+ but did not inhibit the rise in ROS, suggesting that Ca2+ influx was apparently not due to lipid peroxidation. Inhibition at complex I of the mitochondrial

•− electron transport chain with rotenone did not prevent O2 and other ROS generation as

determined with DCFDA and DHE fluorescent probes. Ferricyanide, however, decreased

139 the rise in intracellular ROS, Ca2+ and cell death. Thus, the rapid ROS generation may be mediated by effects on the plasma membrane redox system (PMRS) and does not originate from the mitochondria. For example, in another study (Macho et al., 2003) inhibition of the PMRS with capsaicin caused redirection of the normal electron flow across the plasma membrane, favouring one-electron processes that generate ROS. YCT may act in the same way. The presence of ferricyanide might serve to accept the redirected electrons and alleviate ROS accumulation (Macho et al., 2003).

In recent years, the ability of PMRS to transfer electrons from cytoplasmic

NADH to external electron acceptors (i.e. oxygen, ferricyanide, transferrin and ascorbate) has been suggested to play an important regulatory role in the intracellular redox equilibrium (Macho et al., 2003; deGrey, 2003). Cells lacking an intact mitochondrial electron transport chain have been reported to remain viable by upregulation of the

PMRS (Larm et al., 1994). The PMRS seems to play an essential role in the control of cell proliferation and differentiation (Crane et al., 1990; Crane et al., 1995). Plasma membrane NAD(P)H oxidase (NOX) proteins are involved in hydroquinone (NAD(P)H) oxidation and protein disulfide-thiol interchange processes (Morré, 1998; Morré et al.,

2002). The presence of both CNOX (constitutive) and tNOX (tumour-associated) plasma membrane oxidases have been reported in cancer cells and tissues (Wang et al., 2001) with the former contributing to 40-60% of the activity and the latter accounting for the remainder (Morré, 1998). It seems likely that YCT may target a redox system in leukaemia cells that is inactive or less active in normal lymphocytes. It is tempting to

140 hypothesize that the redox system might also account for the different cytotoxicity of

YCT to different cells.

Mitochondria play a pivotal role in regulating cellular Ca2+ levels (Smaili et al.,

2000; Kroemer and Reed, 2000). Unregulated Ca2+ influx induced by external stimuli can

lead to mitochondrial Ca2+ overloading, which may cause loss of cell viability (Jambrina

et al., 2002; Crane et al., 1990). In this study, a rise in cytosolic Ca2+ was detected as early as 5 min after YCT exposure, but mitochondria depolarized only 12 h later. We suggest that oxidative stress induced by YCT causes a rise in Ca2+, leading to

mitochondrial Ca2+ overloading, release of cytochrome c (possibly due to mitochondrial permeability transition, MPT), loss of ∆ψm and decreased ATP levels. The inhibitory

2+ effects of RuR on rise in mitochondrial Ca and loss of ∆ψm but not on cell death suggesting that the mitochondrial events are secondary to the accumulation of ROS. The

MPT causes the inner mitochondrial membrane to become leaky, allowing solutes less than 1,500 Da to pass through the inner mitochondrial membrane (Kroemer and Reed,

2000; Cai and Jones, 1999). The mechanism of action of YCT appears different from toxins such as clofibrate which appear to target the mitochondria first (Qu et al., 2001).

Our results with CaCl2-free PBS and EGTA demonstrated that YCT induces the influx of extracellular Ca2+ in Jurkat T cells. FA, a non-selective cation channel blocker,

was shown to inhibit YCT-induced Ca2+ influx, but this seems unlikely to be a direct

interaction or activation by YCT of the non-selective cation channels of Jurkat cells by

YCT, since Ca2+ influx was abrogated with ferricyanide. Moreover, peripheral

141 lymphocytes should have undergone the same events as Jurkat T cells if YCT were to

activate the non-selective cation channels leading to Ca2+ influx, since these channels are

present in normal lymphocytes. Possible inhibitory effects of FA on cyclooxygenases do

not seem relevant since indomethacin (IMT) had no effect on rise of intracellular ROS

(Fig. 4.6B) and Ca2+ (Fig. 4.13C) as determined by flow cytometry. In addition, voltage-

2+ + 2+ dependent Ca channels, Na / Ca exchanger and KATP channels were not involved in

YCT-induced Ca2+ influx in Jurkat T cells. Indeed, voltage-dependent Ca2+ channels are

not expressed at a significant level in T cells (Lewis and Cahalan, 1995).

142 CHAPTER 5

CONCLUSION

Our data show that TCM may be a rich source of potentially-novel antioxidant compounds for the treatment of human diseases. However, we should be careful about the safety of antioxidant compounds that is assessed by ‘antioxidant’ screening using simple in vitro assays. Generally, there are two common methods for TCM preparation

(Chevallier, 1996). A mixture of Chinese medicines is prepared by decoction with water

for several hours and should be consumed while the extract is still warm. The other

method involves soaking different Chinese medicines with alcoholic liquor of various

alcoholic contents for several months in an airtight container. The ‘mature’ Chinese

medicinal extract prepared by this method can then be consumed on a daily basis in small

volume. This is believed to improve overall health of a person. Therefore, the data we

have collected using 80% ethanol as a carrier for TCM extracts in our experiments might

be relevant in vivo. However, the bioavailability of TCM compounds and their ability to

decrease oxidative damage needs further investigation using biomarkers.

In regard to the effects of YCT on various cell types, our data illustrate the

importance of studying multiple cell types before drawing conclusions about the

cytotoxicity (or lack of it) of a natural product. Our data also provide evidence for a

possible involvement of PMRS in YCT-induced cell death which is selective towards

certain cell types. We believe that (Fig. 5.1) the accumulation of ROS and intense

oxidative stress due to an effect on the PMRS is followed by a rapid Ca2+ influx through

143 non-selective cation channels that is prevented with FA. The oxidative stress occurs

rapidly and may lead to Ca2+ channel opening. The rise in cytosolic Ca2+ leads to a

2+ secondary rise in mitochondrial Ca , which when excessive causes a fall in ∆ψm, MPT, cytochrome c release and a drop in ATP. YCT-induced cell death events were prevented

in the presence of ferricyanide suggesting a distinct cell death mechanism from the one

induced by a depletion of intracellular GSH. Indeed, depletion of GSH did not occur significantly and NAC did not decrease the loss in cell viability. Our data suggest that the

PMRS may be a target for therapeutic drug design if it is at elevated levels in tumour cells.

The pathways in Fig. 5.1 are not new; they are mainly down stream processes of some toxic agents. It is the inhibition of the plasma membrane redox system by an effect of a natural product (e.g. YCT extract) that is not commonly reported in the literature.

144 Figure 5.1

YCT extract

K3Fe(CN)6

Extracellular Plasma membrane redox system Ca2+

Accumulation of ROS Non-selective (e.g. O •-, LOOH) cation channels 2

2+ ? Ca

FA Mitochondria ?

Scramblase 2+ Mitochondrial Ca activation overloading Phosphatidylserine externalization ?

Mitochondrial Phagocytosis depolarization Release of cytochrome c

ROS Apoptosome formation ? Caspase 3, 7 Caspase 9

Cell death

Fig. 5.1 Diagram illustrating the mechanism(s) of YCT-induced cell death in Jurkat T •- cells. Exposure of YCT causes intense intracellular oxidative stress (i.e. O2 , LOOH) and a rapid Ca2+ influx through a non-selective cation channel. Flufenamic acid (FA) inhibit Ca2+ influx in a concentration-dependent manner. Cytochrome c release is preceded by 2+ mitochondrial Ca overloading and probably also a loss of ∆ψm. ROS may inhibits the activation of caspase-3, -7 and -9. The presence of potassium ferricyanide inhibits YCT- induced cell death cascades suggesting a distinct cell death mechanism.

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181 APPENDIX A