Growth arrest and caspase-dependent apoptosis induced by 5 ...ethesisarchive.library.tu.ac.th/thesis/2016/TU_2016_5411301012_5959_4158.pdf · activation and further indicating the

Ref. code: 25595411301012PXORef. code: 25595411301012PXORef. code: 25595411301012PXO

GROWTH ARREST AND CASPASE-DEPENDENT APOPTOSIS

INDUCED BY 5,6-DIHYDROXY-2,4-DIMETHOXY-9,10-

DIHYDROPHENANTHRENE DERIVED FROM

Dioscorea membranacea PIERRE IN HUMAN

LUNG ADENOCARCINOMA A549 CELLS

BY

MISS WIPADA DUANGPROMPO

A DISSERTATION SUBMITTED IN PARTIAL FULFILLMENT OF

THE REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY PROGRAM IN

BIOCHEMISTRY AND MOLECULAR BIOLOGY

FACULTY OF MEDICINE

THAMMASAT UNIVERSITY

ACADEMIC YEAR 2016

COPYRIGHT OF THAMMASAT UNIVERSITY


GROWTH ARREST AND CASPASE-DEPENDENT APOPTOSIS

INDUCED BY 5,6-DIHYDROXY-2,4-DIMETHOXY-9,10-

DIHYDROPHENANTHRENE DERIVED FROM

Dioscorea membranacea PIERRE IN HUMAN

LUNG ADENOCARCINOMA A549 CELLS

BY

MISS WIPADA DUANGPROMPO

A DISSERTATION SUBMITTED IN PARTIAL FULFILLMENT OF

THE REQUIREMENTS FOR THE DEGREE OF

DOCTOR OF PHILOSOPHY PROGRAM IN

BIOCHEMISTRY AND MOLECULAR BIOLOGY

FACULTY OF MEDICINE

THAMMASAT UNIVERSITY

ACADEMIC YEAR 2016

COPYRIGHT OF THAMMASAT UNIVERSITY


(1)

Dissertation Title GROWTH ARREST AND CASPASE-

DEPENDENT APOPTOSIS INDUCED BY

5,6-DIHYDROXY-2,4-DIMETHOXY-9,10-

DIHYDROPHENANTHRENE DERIVED

FROM Dioscorea membranacea PIERRE IN

HUMAN LUNG ADENOCARCINOMA A549

CELLS

Author Miss Wipada Duangprompo

Degree Doctor of Philosophy Program in Biochemistry

and Molecular Biology

Major Field/Faculty/University Biochemistry and Molecular Biology

Faculty of Medicine

Thammasat University

Dissertation Advisor

Dissertation Co-Advisor

Assistant Professor Pintusorn Hansakul, Ph.D.

Assistant Professor Kalaya Aree, Ph.D.

Academic Years 2016

ABSTRACT

An active compound of Dioscorea membranacea Pierre called 5,6-

dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene (HMP) has been shown to

possess the selective antiproliferative effect against human lung large cell carcinoma

COR-L23 cells. In this study, an adequate amount of HMP was isolated from D.

membranacea Pierre using column chromatography on silica gel 60 as the stationary

phase, and the column was eluted by gradient elution in increasing order of polarity.

The isolated compound was determined to be HMP by comparing its spectral data of

proton nuclear magnetic resonance (1H NMR) with those of previously isolated

compound and was further tested for cell-type-specific cytotoxicity in two main types

of lung cancer cell lines including non-small cell lung cancer (NSCLC) and small cell

lung cancer (SCLC) using the sulforhodamine B (SRB) assay. The results showed that

the validated HMP exhibited the most antiproliferative and cytotoxic effects on

adenocarcinoma cell line A549, one of the three main subtypes of NSCLC. Therefore,


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the aims of this study were to comprehensively investigate the antiproliferative and

cytotoxic effects through cell cycle arrest and apoptosis in human lung carcinoma A549

cells.

In the present study, the antiproliferative and cytotoxic effects of HMP were

analyzed by the SRB assay. Cell division, cell cycle distribution, membrane asymmetry

changes and intracellular ROS generation were each performed with different

fluorescent dyes including carboxyfluorescein succinimidyl ester (CFSE), propidium

iodide (PI), annexin V-FITC double staining and 2´,7´-dichlorofluorescein (DCF),

respectively, and they were then analyzed by flow cytometry. Cell cycle- and apoptosis-

related mRNA and proteins levels were measured by real-time PCR and western blot

analyses, respectively. The nuclear morphology of apoptotic cells stained with 4’, 6-

diamidino-2-phenylindole (DAPI) and DNA fragmentation were detected by

fluorescence microscopy and gel electrophoresis, respectively. The results showed that

HMP exerted strong antiproliferative (represented as IC50 = 9.37 µM and TGI = 54.81

µM) and cytotoxic effects (represented as LC50 = 94.01 µM) in A549 cells with the

highest selectivity index as compared with the human lung fibroblast cell line MRC-5.

Treatment of A549 cells with HMP induced a rapid arrest of cell division and halted

the cell cycle at G2/M phase through down-regulation of the expression levels of G2/M

regulatory proteins cdc25C, cdk1 and cyclin B1. Moreover, HMP treatment induced

early apoptotic cells with externalized phosphatidylserine and subsequent apoptotic

cells in the sub-G1 phase of the cell cycle and concurrent activation of caspase-3, whose

activity was completely abolished with pan-caspase inhibitor Z-VAD-fmk. Indeed, the

active form of caspase-3 was detected, and its actions were supported by the results of

cleavage of its target PARP and morphological alterations of apoptotic cell death such

as nuclear condensation, DNA fragmentation with accompanying DNA ladder

formation. In addition, HMP significantly increased the Bax/Bcl-2 mRNA and protein

ratios of proapoptotic, especially at 72 h of incubation, leading to subsequent caspase-9

activation and further indicating the induction of the intrinsic apoptotic pathway. Also,

HMP induced apoptosis via the extrinsic pathway by causing the proteolytic cleavage

of Bid. This study, furthermore, demonstrated that HMP could generate excessive

intracellular ROS, which was confirmed using ROS scavenger NAC. This inhibitor was

used to further study the involvement of ROS in HMP-induced apoptosis.


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In conclusion, this is the first molecular evidence of HMP that exerted its

anticancer actions through the induction of G2/M cell cycle arrest as well as the intrinsic

and extrinsic apoptotic pathways in A549 cells. These data support the potential role of

HMP as a cell-cycle arrest and apoptosis-inducing agent for treatment of NSCLC and

the use of D. membranacea Pierre in Thai traditional herbal remedies for cancer

treatment.

Keywords: Apoptosis, Anticancer effect, G2/M arrest, Dioscorea membranacea Pierre,

5,6-Dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene


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ACKNOWLEDGEMENTS

I would like to express my deepest and sincere gratitude to my advisor,

Assistant Professor Dr. Pintusorn Hansakul for her kindness in providing an opportunity

to be my advisor. I am also grateful for her valuable supervision, valuable suggestions,

supporting, encouragement, guidance and criticism throughout the course of my study.

I also would like to express my greatest appreciation and sincere gratitude to my co-

advisors, Assistant Professor Dr. Kalaya Aree for her valuable comments and

suggestions.

I wish to express my sincere appreciation to Associate Professor Dr. Treetip

Ratanavalachai, Dr. Saengsoon Charoenvilaisiri and Dr. Srisopa Ruangnoo for being

my external committee and for giving helpful suggestions.

I would like to thank Associate Professor Dr. Arunporn Itharat and Dr.

Pakakrong Thongdeeying, Department of Applied Thai Traditional Medicine, Faculty

of Medicine, Thammasat University for their help and suggestion on the laboratory

techniques in the part of plant extraction and HMP isolation.

I am grateful to all staffs and friends of the Faculty of Medicine, Thammasat

University, for their kind help and friendship.

My special thanks are extended to Mr. Suebkul Kanchanasuk, Mr. Worawat

Surarit, Miss Kedsara Junmakho and whom it concern to my study as I did not mention

for their kindness, help support and friendships during a time of the study.

Finally, I would like to express my sincere gratitude and appreciation to my

dear parents for their love, pushing up, cheerfulness, devoting and encouragement

throughout my life.

This research was mainly supported by National Research University Project

of Thailand, Office of the Higher Education Commission and Center of Excellence of

Applied Thai Traditional Medicine Research of Thammasat University. The authors

gratefully acknowledge the financial support provided by the Research Grants of

Thammasat University for Ph.D. students and TU Research Scholar, Contract No. 79/2558.

Miss Wipada Duangprompo


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TABLE OF CONTENTS

Page

ABSTRACT (1)

ACKNOWLEDGEMENTS (4)

LIST OF TABLES (10)

LIST OF FIGURES (11)

LIST OF ABBREVIATIONS (15)

CHAPTER 1 INTRODUCTION 1

1.1 Rational and Background 1

1.2 Aims of this study 3

1.2.1 Overall aims 3

1.2.2 Specific aims 3

1.3 Outcomes 4

CHAPTER 2 REVIEW OF LITERATURE 5

2.1 Lung cancer 5

2.1.1 Incidence and etiology 5

2.1.2 Pathology and staging of lung cancer 6

2.1.2.1 Non-small cell lung cancer (NSCLC) 6

2.1.2.2 Small cell lung cancer (SCLC) 10

2.1.3 Lung cancer treatment 11

2.1.4 Molecular and genetic aspects of lung cancer 11


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2.1.4.1 Proto-oncogenes 12

2.1.4.2 Tumor suppressor genes 13

2.2 Alteration of cell cycle in cancer cells 18

2.2.1 Normal cell cycle regulation and cell cycle checkpoint 18

2.2.1.1 Cell cycle regulation 19

(1) Regulation of G1 phase progression 22

(2) Regulation of S phase progression 24

(3) Regulation of G2 phase progression 25

(4) Regulation of M phase progression 25

2.2.1.2 Cell cycle checkpoints 26

(1) Restriction checkpoint 27

(2) Replication checkpoint 27

(3) Spindle checkpoint 27

(4) DNA damage checkpoint 28

2.2.2 Alterations of cell cycle regulation in cancer cells 28

2.2.2.1 Oncogenes 29

2.2.2.2 Deregulated tumor suppressor genes 31

2.3 Classification of cell death 33

2.3.1 Autophagy 33

2.3.2 Necrosis 35

2.3.3 Apoptosis 35

2.4 Alteration of apoptotic cell death in cancer cells 36

2.4.1 Apoptosis in normal cells 36

2.4.1.1 The mitochondrial pathway (or intrinsic pathway) 37

(1) Caspase-dependent apoptosis 38

(2) Caspase-independent apoptosis 39

2.4.1.2 The death receptor pathway (or extrinsic pathway) 39

2.4.2 Apoptosis in cancer cells 40

2.4.3 Reactive oxygen species (ROS) leading to apoptosis in cancer cells 40

2.5 Targeting for cancer treatment 43


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2.5.1 Targeting cell cycle regulators in cancer treatment 43

2.5.2 Targeting apoptosis in cancer treatment 45

2.6 Thai medicinal plants (Hua-Khao-Yen) 47

2.6.1 Dioscorea membranacea Pierre 47

2.6.1.1 General description 47

2.6.1.2 Biological activities 51

(1) Antiproliferative activity 51

(2) Anti-allergic activity 53

(3) Anti-HIV activity 54

(4) Antioxidant activity 54

(5) Anti-inflammatory activity 54

2.6.2 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene 55

CHAPTER 3 RESEARCH METHODOLOGY 56

3.1 Conceptual framework of this study 56

3.2 Extraction of Dioscorea membranacea Pierre 58

3.3 Isolation of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene 59

3.4 Cell culture 60

3.5 Growth inhibitory and cytotoxic effects 60

3.6 Cell proliferation by CFSE assay 62

3.7 Cell cycle analysis 64

3.8 Annexin-V/PI double staining assay 65

3.9 Caspase-3 activity assay 66

3.10 Real-time Quantitative PCR Analysis 67

3.11 Western blot analysis 69

3.12 Tubulin polymerization assay 70

3.13 DNA fragmentation assay 71

3.14 Nuclear staining with DAPI 72

3.15 Intracellular reactive oxygen species (ROS) measurement 73

3.16 Statistical analysis 74


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CHAPTER 4 RESULTS AND DISCUSSION 75

4.1 Extraction of Dioscorea membranacea Pierre 75

4.2 Isolation and purification of HMP 75

4.3 Antiproliferative and cytotoxic effects of HMP against 88

a panel of human lung cancer cell lines

4.4 Inhibitory effects of HMP on cell division 90

4.5 Effects of HMP on the cell cycle distribution 92

4.6 Effect of HMP on protein expression of cell cycle regulatory proteins 95

4.7 Effect of HMP on interfering microtubule formation 96

4.8 Effect of HMP on apoptosis induction in A549 cells 97

4.9 Effect of HMP on caspase-3 activity in A549 cells 100

4.10 Bax and Bcl-2 mRNA and protein expression levels 104

4.11 Effect of HMP on expression of active caspases and their targets 107

4.12 Effect of HMP on nuclear morphological changes 108

4.13 Effect of HMP on DNA fragmentation 111

4.14 Effect of HMP on the generation of intracellular ROS 112

CHAPTER 5 CONCLUSIONS AND RECOMMENDATIONS 117

5.1 Antiproliferative effect of HMP in A549 cells 117

5.2 Molecular mechanism underlying antiproliferative effect of HMP 118

5.3 Cytotoxic effects of HMP in A549 cells 120

5.4 Apoptosis underlying cytotoxic effects of HMP 120

5.5 Effects of HMP on the generation of intracellular ROS and 122

the relationship between enhanced ROS and apoptosis

REFERENCES 123

APPENDICES 143

APPENDIX A GROWTH CURVE 144


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APPENDIX B STANDARD CURVE FOR PROTEIN 150

DETERMINATION

APPENDIX C HPLC CHROMATOGRAMS 151

APPENDIX D FLOW CYTOMETRIC ANALYSIS 154

APPENDIX E REAGENTS FOR LABORATORY EXPERIMENTS 155

BIOGRAPHY 160


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LIST OF TABLES

Tables Page

2.1 The TNM staging system for lung cancer 8

2.2 Cancer genes and their functions found in lung cancer 15

2.3 Cyclin/CDKs complex are activated within specific phases of 20

the cell cycle.

4.1 The percent yield of the ethanolic extract of D. membranacea Pierre 75

4.2 The percent yield of HMP-1 isolated from D. membranacea Pierre 79

4.3 The percent yield of HMP-2 isolated from D. membranacea Pierre 81

4.4 1H NMR spectral data (500 MHz) of HMP-1, HMP-2 and previously 85

isolated 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene

4.5 The retention time, area under the curve and percentage area of HMP-1, 87

HMP-2 and 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene

analyzed by HPLC at wavelengths 254 and 270 nm.

4.6 Antiproliferative effects of HMP on a panel of human cell lines 89

4.7 The percentages of HMP-treated cells in each phase of cell cycle 94

4.8 The percentages of cells in the respective quadrants 99

4.9 The percentages of cells in each phase of cell cycle 102

4.10 The percentages of cells in each phase of cell cycle 116

25 µM HMP treatment for 24 h.

E-1 Recipes for resolving and stacking gels 157


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LIST OF FIGURES

Figures Page

2.1 Signal transduction pathway that promotes cell division 13

2.2 Phases of the cell cycle. 19

2.3 Changes in cyclins during the cell cycle. 20

2.4 Regulation of cdk-cyclin complex by phosphorylation and 21

dephosphorylation

2.5 A schematic representation of various changes in the activity of 23

cyclin-cdk complex during the cell cycle

2.6 The cell cycle checkpoint 26

2.7 The mechanisms that lead to the conversion of proto-oncogenes to 30

oncogenes

2.8 The mechanisms that lead to the deregulation of 32

tumor suppressor genes

2.9 Characteristics of autophagy, apoptosis and necrosis 34

2.10 A schematic representation of intrinsic and extrinsic pathways of 37

Apoptosis

2.11 JNK/p38 MAPK signaling pathways, apoptosis pathway and multiple 42

molecular targets of plant-derived agents

2.12 The characteristics of D. membranacea Pierre 48

2.13 Dioscorea membranacea Pierre (Male plant) 49

2.14 Dioscorea membranacea Pierre (Female plant) 50

2.15 Chemical structures of isolated compounds from the rhizomes of 52

D. membranacea Pierre

2.16 Chemical structures of isolated compounds from the rhizomes of 53

D. membranacea Pierre

2.17 The structure of 5,6-dihydroxy-2,4-dimethoxy-9,10- 55

dihydrophenanthrene

3.1 The physical characteristics of the rhizome of D. membranacea Pierre 58

3.2 Formation of fluorescent compound CFSE by intracellular esterase 63


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3.3 DNA content distribution during the various phases of the cell cycle 65

obtained by flow cytometric analysis

3.4 Dot plot analysis by Annexin V-FITC/PI double staining 66

3.5 DNA fragmentation analysis 72

3.6 Formation of fluorescent compound DCF by ROS and RNS 74

4.1 TLC analysis of the 14 combined fractions of D. membranacea Pierre 76

extract obtained from the first silica gel column chromatography

4.2 TLC analysis of the odd numbered fractions, ranging from 25-99, 77

eluted from the second silica gel column chromatography

4.3 TLC analysis of the 3 groups of the combined fractions eluted from 77

the second silica gel column chromatography

4.4 The schematic flow chart for isolation of clearly separated bands 78

using a TLC glass plate

4.5 TLC isolation of the combined fractions in group 2 79

4.6 TLC analysis of the odd numbered fractions, ranging from 1-51, 80

eluted from the third silica gel column chromatography

4.7 TLC analysis for checking the purity of HMP-2 in three different 81

solvent systems of varying polarity

4.8 1H NMR spectrum of HMP-1 in deuterated chloroform (CDCl3) 82

4.9 1H NMR spectrum of HMP-2 in deuterated chloroform (CDCl3) 83

4.10 1H NMR spectrum of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10- 84

dihydrophenanthrene in deuterated chloroform (CDCl3)

4.11 HPLC chromatograms of HMP-1, HMP-2 and previously isolated 86

5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene

4.12 Effects of HMP on antiproliferative and cytotoxic activities 89

in A549 cells

4.13 Antiproliferative effects of HMP on A549 cells 91

4.14 Effects of HMP on cell cycle distribution in A549 cells 93

4.15 Effects of HMP on protein levels of cdc25C, cdk1 and cyclin B1 95

in A549 cells

4.16 Effect of HMP on in vitro tubulin polymerization 96

4.17 Effects of HMP on apoptotic induction in A549 cells 98


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4.18 Inhibitory effects of Z-VAD-fmk on sub-G1 populations 101

4.19 Effects of HMP on caspase-3 activity in A549 cells 103

4.20 The quantification of relative mRNA levels of Bax and Bcl-2 105

in A549 cells using Real-time PCR

4.21 Effects of HMP on protein expression of Bax and Bcl-2 in A549 cells 106

4.22 Effects of HMP on expressions of apoptotic proteins in A549 cells 108

4.23 Effects of HMP on nuclear morphological changes by DAPI staining 109

under bright-field microscopy (400x magnification)

4.24 Effects of HMP on nuclear morphological changes by DAPI staining 110

under fluorescent microscopy (400x magnification)

4.25 Effect of HMP on DNA fragmentation of A549 cells 111

4.26 Effect of HMP on ROS production in A549 cells treated with 113

25 µM HMP at different incubation times

4.27 Effect of the ROS scavenger NAC on ROS production in A549 cells 114

4.28 Inhibitory effects of NAC on sub-G1 populations 115

A-1 Growth curve of human lung carcinoma cell line A549 144

in 96-well plates

A.2 Growth curve of human lung squamous carcinoma cell line NCI-H226 145

in 96-well plates

A-3 Growth curve of human large cell lung cancer line COR-L23 146

in 96-well plates

A-4 Growth curve of human small cell lung cancer cell line NCI-H1688 147

in 96-well plates

A-5 Growth curve of human lung fibroblast cell line MRC-5 148

in 96-well plates

A-6 Growth curve of human lung carcinoma cell line A549 149

in 24-well plates

B-1 Standard curve for protein determination by Bradford’s method 150

B-2 Standard curve for protein determination by BCA Assay 150

C-1 HPLC chromatogram of HMP-1 151

C-2 HPLC chromatogram of HMP-2 152


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C-3 HPLC chromatogram of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10- 153

Dihydrophenanthrene

D-1 Flow cytometric analysis of the DNA from A549 cells treated with 154

NAC alone at different concentrations (0.1, 1 and 5 mM) for 72 h.


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LIST OF ABBREVIATIONS

Symbols/Abbreviations Terms

α

β

%

µg

µl

µM

°C

APS

AUC

bp

BCA

BSA

CAD

CDCl3

Cdc25C

Cdk1 CFSE

CO2

CT

DAPI

DCF

DCFH-DA DMSO

DNase

DNA

cDNA

et al.

Alpha

Beta

Percent

Microgram

Microliter

Micromolar

Degree Celsius

Ammonium per sulfate

Area under the curve

Base pair

Bicinchoninic acid

Bovine serum albumin

Caspase-activated deoxyribonuclease

Deuterochloroform

Cell Division Cycle 25C

Cyclin-dependent kinase 1

Carboxyfluorescein succinimidyl

Carbon dioxide

Comparative threshold

4′,6-diamidino-2-phenylindole

2,7-dichlorofluorescein

2´,7´-dichlorofluorescein diacetate

Dimethyl sulphoxide

Deoxyribonuclease

Deoxyribonucleic acid

Complementary deoxyribonucleic acid

et alibi, and others


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FBS

FITC

g

h

HMP

HPLC

i.e.

IAPs

ICAD

LC50

M

mAU

mg

ml

mm

mM

mole

MOMP

mRNA

M.W.

nm

NAC

NMR

1H NMR

NSCLC

O.D.

PARP

PBS

Fetal bovine serum

Fluorescein isothiocyanate

Gram

Hour

5,6-dihydroxy-2,4-dimethoxy-9,10-

dihydrophenanthrene

High-performance liquid chromatography

id est (Latin), that is or in other words

Inhibitor apoptotic proteins

Inhibitor of caspase-activated

deoxyribonuclease

50% lethal concentration

Molar (concentration)

Milli- absorbance units

Milligram

Milliliter

Millimeter

Millimolar

Mole

Mitochondrial outer membrane

permeabilization

Messenger ribonucleic acid

Molecular weight

Nanometer

N-acetylcysteine

Nuclear magnetic resonance

Proton nuclear magnetic resonance

Non-small cell lung cancer

Optical density

Poly (ADP-ribose) polymerase

Phosphate buffer saline


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PCR

PI

PS

PVDF

ROS

RNA

Rpm

Rf

RT

RT-PCR

RQ

SCLC

SD

SDS

SDS-PAGE

SI

SRB

TBS

TBST

TCA

TLC

TGI

UV

w/w

Polymerase chain reaction

Propidium iodide

Phosphatidylserine

Polyvinylidene fluoride

Reactive oxygen species

Ribonucleic acid

Revolutions Per Minute

Retention factor

Retention time

Reverse transcription polymerase chain

reaction

Relative quantitation

Small cell lung cancer

Standard deviation

Sodium dodecyl sulfate

Sodium dodecyl sulfate polyacrylamide

gel electrophoresis

Selectivity index

Sulphorhodamine B

Tris-buffered saline

Tris-buffered saline with Tween20

Trichloroacetic acid

Thin-layer chromatography

Total growth inhibition

Ultraviolet

weight per weight


1

CHAPTER 1

INTRODUCTION

1.1 Rational and Background

Lung cancer, one of the most common cancers, has become increasingly a

significant health problem in the world. The treatment options such as surgery,

radiotherapy, chemotherapy, and targeted therapy are currently being used depending

on the type and stage of lung cancer. Although chemotherapy treatment is the most

common regimen to treat patients, it has many unpleasant side effects such as bone

marrow suppression, gastrointestinal problems (nausea, vomiting, diarrhea), alopecia

(or hair loss) and others (Chun, Garrett, & Vail, 2007). Moreover, chemotherapy

resistance continues to be a major problem for lung cancer treatment. For this reason,

the search for new anticancer agents with increased safety and efficacy, and with

affordable price, is one of the most effective strategies to overcome the limitation of

currently available chemotherapeutic drugs. These new anticancer agents must exert

potent and specific cytotoxicity as well as their actions at the molecular level should be

clearly understood.

As cancer cells acquire defects in cell cycle control and apoptosis, new

promising anticancer agents should thus be able to potently block cell division via cell

cycle arrest (Choi, Lim do, & Park, 2009; Choi & Yoo, 2012) and concurrently restore

apoptosis towards normality (Wong, 2011). Over the years, induction of cell cycle

arrest and apoptosis has emerged as the major mechanisms by which active anticancer

agents act to inhibit the growth of cancer cells and eliminate them (Feng et al., 2011).

Therefore, investigating whether agents exert their cytotoxic actions through the

induction of cell cycle arrest and apoptosis appears to be a powerful strategy to obtain

effective agents for the development of chemotherapeutic drugs.

Many studies have shown that plant-derived compounds mediating cell

cycle arrest and apoptosis have increasingly attracted scientific interest, such as

curcumin (Curcuma longa) (Tan et al., 2006; Lee, Lee, & Kim, 2009; Wu et al., 2010;

Cheng et al., 2016), shikonin (Lithospermum erythrorhizon) (Wu et al., 2004; Gong &


2

Li, 2011; Tian, Li, & Gao, 2015), magnolol (Magnolia officinalis) (Zhou et al.,2013;

Li et al., 2015), genistein (soybean) (Ouyang et al., 2009; Zhang et al., 2013),

resveratrol (red grape skins) (Aziz, Nihal, Fu, Jarrard, & Ahmad, 2006; Gogada et al.,

2011) and so forth. Moreover, some of them have the effects on the generation and

accumulation of intracellular reactive oxygen species (ROS). The excessive ROS

production leads to the activation of mitochondria-mediated apoptosis pathway (Gong

& Li, 2011; Singh, Zaidi, Shyam, Sharma, & Balapure, 2012; Qui et al., 2015).

Dioscorea membranacea Pierre, also called Hua-Khao-Yen-Tai in Thai, is

one of Thai medicinal plants, which has long been used to prepare Thai traditional

medicine for cancer treatment (Itharat, Singchangchai, & Ratanasuwan, 1998;

Subchareon, 1998). Previous studies have shown that the ethanolic extract of D.

membranacea Pierre and its active compounds exhibited high cytotoxic activity against

a panel of human cancer cell lines (Itharat et al., 2003; Itharat et al., 2004; Itharat et al.,

2007; Itharat, Thongdeeying, & Ruangnoo, 2014). Among these active compounds,

dioscorealide B (Saekoo, Dechsukum, Graidist, & Itharat, 2010; Saekoo, Graidist,

Leeanansaksiri, Dechsukum, & Itharat, 2010) and dioscoreanone (Hansakul, Aree,

Tanuchit, & Itharat, 2014) have been elucidated for their molecular mechanisms of

action. Itharat et al. (2014) have demonstrated that an active compound named 5,6-

dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene (HMP) exerted the selective

cytotoxic effects against human lung, breast and prostate cancer cell lines whereas it

was less toxic to the normal cell line. However, the molecular mechanisms underlying

its cytotoxic effect have not yet been studied.

Thus, this study we further investigated the antiproliferative effect of HMP

against a panel of different human lung cancer cell lines. A549 cell line, one of cell

lines displaying the most potent inhibitory effect with the highest selectivity index, was

chosen to investigate the molecular mechanisms underlying anticancer effect through

the induction of cell cycle arrest and apoptosis. In addition, the effects of HMP on the

generation and accumulation of intracellular reactive oxygen species (ROS) were

studied.


3

1.2 Aims of this study

1.2.1 Overall aims

To investigate molecular mechanisms underlying the anticancer activity of

HMP in human lung adenocarcinoma cell line A549 through cell cycle arrest and

apoptosis

1.2.2 Specific aims

The aims of this study were as follows:

1.2.2.1 To isolate 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenan

threne (HMP) from Dioscorea membranacea Pierre

1.2.2.2 To determine the antiproliferative effect of HMP on different

lung cancer cell lines as compared to the normal cell line

1.2.2.3 To determine the cytotoxic effect of HMP on human lung

carcinoma cell line A549

1.2.2.4 To investigate the antiproliferative activity of HMP through

the induction of cell cycle arrest in A549 cells

1.2.2.5 To investigate the cytotoxic activity of HMP through the

induction of apoptosis in A549 cells

1.2.2.6 To investigate the intracellular reactive oxygen species (ROS)

levels in HMP-treated A549 cells and further examine whether ROS is associated with

HMP-induced apoptosis


4

1.3 Outcomes

These research findings provided:

1.3.1 Knowledge on HMP-induced G2/M arrest through modulation of

specific regulatory proteins and mitotic spindle disruption in A549 cells.

1.3.2 Knowledge on HMP-induced apoptosis via caspase-dependent

pathway in A549 cells.

1.3.3 Data supporting the development of HMP as a novel anticancer

drug.

1.3.4 Data supporting the high economic value of D. membranacea

Pierre as a source of potential anticancer compounds.


5

CHAPTER 2

REVIEW OF LITERATURE

2.1 Lung cancer

2.1.1 Incidence and etiology

Lung cancer is one of the most important malignancies and the most

common cause of cancer death worldwide. In 2012, data from GLOBOCAN project

produced by the International Agency for Research on Cancer (IARC) have shown that

lung cancer is responsible for more cancer-related deaths than any other types of cancer

(Torre et al., 2015). The major reason that contributes to its high mortality rate is the

fact that a large proportion of these cases are diagnosed with the advanced or metastatic

stage. The data from American Cancer Society (2015) have revealed that more than

half (57%) of lung cancer are diagnosed at a distant stage, for which the 1- and 5-year

survival is 26% and 4%, respectively. The 5-year survival for small cell lung cancer

(6%) is lower than that for non-small cell (21%). Moreover, an estimated 1.8 million

new lung cancer cases occurred in both men and women, accounting for 13% of total

cancer diagnoses. The high prevalence of lung cancer is increasingly becoming a

significant health problem in many regions of the world (Torre et al., 2015).

In Thailand during 2004-2006, lung cancer is the second most common

cancer in males after liver cancer, and the fourth in females after cervix, breast, and

liver cancer (Müller-Hermelink et al., 2004; Sriplung et al., 2005). In 2002, Vatanasapt,

Sriamporn, & Vatanasapt have reported that the cancer incidence rate appears to depend

on the geographical regions. For example, lung cancer predominates in the northern

part of Thailand whereas liver cancer, especially cholangiocarcinoma, is high in the

Northeast.

A major risk factor for developing lung cancer is tobacco consumption

because tobacco contains a complex mixture of potent carcinogens, predominantly

polycyclic aromatic hydrocarbons derived from combustion of tars. Other known risk

factors for lung cancer include gender, occupation, diet, radon exposure and passive

smoking. Interactions between risk factors, and in particular with cigarette smoking,


6

may increase lung cancer risk significantly. Hereditary factors and genetic

susceptibility to lung cancer currently remain ill-defined. Recently, outdoor pollution

has also been determined to cause lung cancer (Codony-Servat, Verlicchi, & Rosell,

2016).

2.1.2 Pathology and staging of lung cancer

According to pathological type, lung cancer can be divided into two

histological groups: non-small cell lung cancer (NSCLC) and small cell lung cancer

(SCLC). This broad stratification reflects fundamental differences in tumor biology and

clinical behavior as well as underlies current treatment strategies.

2.1.2.1 Non-small cell lung cancer (NSCLC)

NSCLC is the most common type of lung cancer, which accounts for

80% of the cases, and it usually grows and spreads more slowly than SCLC. NSCLC

can be divided into three major groups: adenocarcinoma, squamous cell carcinoma and

large-cell lung carcinoma based on morphological and immunohistochemical (IHC)

features (Rekhtman, Ang, Sima, Travis, & Moreira, 2011; Travis & Rekhtman, 2011;

Kadota et al., 2015).

- Adenocarcinoma is the most common histologic subtype of lung

cancer and accounts for about 50 % of NSCLC and 38 % of newly diagnosed lung

cancers. It usually originates in the periphery of the lung (outer part of the lung). The

adenocarcinoma is defined by the World Health Organization (WHO) as a malignant

epithelial tumor with glandular differentiation or mucin production, showing acinar,

papillary, bronchioloalveolar or solid with mucin growth patterns or a mixture of these

patterns (Müller-Hermelink et al., 2004). However, the adenocarcinoma can present

diverse histological patterns, which can be intermixed in the same tumor including

lepidic, acinar, papillary, micropapillary, and solid patterns. Therefore, pneumocyte

marker expression like napsin A or thyroid transcription factor 1 (TTF1) is useful in the

identification of the adenocarcinoma in challenging cases (Travis et al., 2015). For

example, solid patterns of adenocarcinoma can be confused with squamous cell


7

carcinoma or large-cell lung carcinoma; the mucin production and immunohistochemical

expression of TTF-1 or napsin A can help in such diagnosis (Rodriguez-Canales, Parra-

Cuentas, & Wistuba, 2016).

- Squamous cell carcinoma represents for nearly 20 % of all lung

cancers, and it is usually found in a central location, arising in a main or lobar bronchus.

The squamous cell carcinoma is defined as a malignant epithelial tumor showing

keratinization and/or intercellular bridges that arise from the bronchial epithelium

(Müller-Hermelink et al., 2004). However, some squamous cell carcinoma may not

show such morphological features. Immunohistochemical tests including markers of

squamous cell differentiation such as p40 or p63 and cytokeratins 5/6 may be useful in

the identification of squamous cell carcinoma in difficult cases (Travis et al., 2015). For

example, a distinct entity is the basaloid squamous cell carcinoma, a poorly

differentiated malignant tumor without morphological features of squamous cell

differentiation which can be confused with small-cell lung carcinoma, but it is

characteristically positive for immunomarkers of squamous cell differentiation

including p40, p63, and cytokeratins 5/6, while TTF-1 is negative (Rodriguez-Canales

et al., 2016).

- Large-cell lung carcinoma accounts for about 3 % of all lung

cancers. The large-cell lung carcinoma is defined as undifferentiated non-small cell

carcinoma that lacks the cytologic and architectural features of small cell carcinoma

and glandular or squamous differentiation (Müller-Hermelink et al., 2004). Based on

immunohistochemistry, large-cell lung carcinoma may be positive for cytokeratins 5/6

but they are negative for TTF-1 and p40 (Travis et al., 2015; Rodriguez-Canales et al.,

2016).

The lung cancer staging system provides useful prognostic information

for patients and structures treatment plans for physicians. According to the International

Association for the Study of Lung Cancer (IASLC), the latest revision of tumor–node–

metastasis (TNM) staging, presented in the 7th edition of American Joint Committee on

Cancer (AJCC) is shown in Table 2.1.


8

Table 2.1 The TNM staging system for lung cancer (Modified from Kalemkerian,

2011)

Anatomic stage/

prognostic groups Tumor (T) lymph nodes (N) Metastasis (M)

Occult Carcinoma TX N0 M0

Stage 0 Tis N0 M0

Stage IA T1a N0 M0

T1b N0 M0

Stage IB T2a N0 M0

Stage IIA T2b N0 M0

T1a N1 M0

T1b N1 M0

T2a N1 M0

Stage IIB T2b N1 M0

T3 N0 M0

Stage IIIA T1a N2 M0

T1b N2 M0

T2a N2 M0

T2b N2 M0

T3 N1 M0

T3 N2 M0

T4 N0 M0

T4 N1 M0

Stage IIIB T1a N3 M0

T1b N3 M0

T2a N3 M0

T2b N3 M0

T3 N3 M0

T4 N2 M0

T4 N3 M0

Stage IV Any T Any N M1a

Any T Any N M1b

Note: TNM descriptions for staging lung cancer

Primary Tumor (T)

TX Primary tumor cannot be assessed, or tumor proven by the presence of malignant

cells in sputum or bronchial washings but not visualized by imaging or

bronchoscopy


9

T0 No evidence of primary tumor

Tis Carcinoma in situ

T1 Tumor 3 cm or less in greatest dimension, surrounded by lung or visceral pleura,

without bronchoscopic evidence of invasion more proximal than the lobar

bronchus (for example, not in the main bronchus)

T1a Tumor 2 cm or less in greatest dimension

T1b Tumor more than 2 cm but 3 cm or less in greatest dimension

T2 Tumor more than 3 cm but 7 cm or less or tumor with any of the following

features (T2 tumors with these features are classified T2a if 5 cm or less): involves

main bronchus, 2 cm or more distal to the carina; invades visceral pleura (PL1 or

PL2); associated with atelectasis or obstructive pneumonitis that extends to the

hilar region but does not involve the entire lung

T2a Tumor more than 3 cm but 5 cm or less in greatest dimension

T2b Tumor more than 5 cm but 7 cm or less in greatest dimension

T3 Tumor more than 7 cm or one that directly invades any of the following: parietal

pleural (PL3), chest wall (including superior sulcus tumors), diaphragm, phrenic

nerve, mediastinal pleura, parietal pericardium; or tumor in the main bronchus

less than 2 cm distal to the carina1 but without involvement of the carina; or

associated atelectasis or obstructive pneumonitis of the entire lung or separate

tumor nodule(s) in the same lobe

T4 Tumor of any size that invades any of the following: mediastinum, heart, great

vessels, trachea, recurrent laryngeal nerve, esophagus, vertebral body, carina,

separate tumor nodule(s) in a different ipsilateral lobe

Regional lymph nodes (N)

NX Regional lymph nodes cannot be assessed

N0 No regional lymph node metastases

N1 Metastasis in ipsilateral peribronchial and/or ipsilateral hilar lymph nodes and

intrapulmonary nodes, including involvement by direct extension

N2 Metastasis in ipsilateral mediastinal and/or subcarinal lymph node(s)

N3 Metastasis in contralateral mediastinal, contralateral hilar, ipsilateral or

contralateral scalene, or supraclavicular lymph node(s)

Distant Metastasis (M)

M0 No distant metastasis

M1 Distant metastasis

M1a Separate tumor nodule(s) in a contralateral lobe, tumor with pleural

nodules or malignant pleural (or pericardial) effusion

M1b Distant metastasis (in extrathoracic organs)


10

2.1.2.2 Small cell lung cancer (SCLC)

SCLC is a malignant epithelial tumor consisting of small cells with

almost no visible cytoplasm, ill-defined cell borders, finely granular nuclear chromatin,

and absent or inconspicuous nucleoli. The cells are round, oval, or spindle-shaped.

Nuclear molding is prominent. Necrosis is typically extensive and the mitotic count is

high (Müller-Hermelink et al., 2004). SCLC accounts for about 20% of all lung cancers,

and it is considered to be the most aggressive form of lung cancer that has a high

propensity for metastases and a poor prognosis. Comparable to other lung cancers,

SCLC has the highest association with tobacco smoking and almost never arising in the

absence of smoking history (Pesch et al., 2012). In addition, this tumor is now generally

considered as a neuroendocrine carcinoma (with small and large cell variants), and

immunohistochemical studies have consistently demonstrated characteristic

biomarkers, including calcitonin, gastrin-releasing peptide, L-dopa decarboxylase,

chromogranin, synaptophysin, and neuron-specific enolase. However, the precise cell

of origin for lung cancer is controversial, and a mosaic of cellular elements (including

NSCLC) is common in tumors with otherwise predominantly small-cell histology

(Macdonald, Ford, & Casson, 2004).

According to the Veterans’ Administration Lung Study Group

(VALSG) system, SCLC is generally divided into two stages, limited and extensive.

Limited disease (LD) is defined as a tumor that is confined to one hemithorax and

associated regional lymph nodes whereas extensive disease (ED) is defined as tumor

outside the confines of limited stage disease including patients with malignant

pericardial and pleural effusion (Bernhardt & Jalal, 2016). Recently, the IASLC has

proposed the revised TNM staging system, presented in the 7th edition of American

Joint Committee on Cancer (AJCC), which is integrated into the classification of SCLC

(Kalemkerian, 2011). For example, LD constitutes approximately 35-40 % of patients

and includes TNM stages I through III, and ED includes patients of TNM IV. The TNM

staging seems more accurate than the limited versus extensive stage in determining

prognosis, especially, in the earlier stages of the disease (Bernhardt & Jalal, 2016).


11

2.1.3 Lung cancer treatment

There are several types of lung cancer treatments such as surgery,

radiotherapy, chemotherapy and targeted therapy, either alone or in combination. In

order to eliminate abnormal cells, these treatments are selected based on the histological

types, stages of lung cancer, the patient’s general condition (Dobbelstein & Moll,

2014). For patients with SCLC, the standard treatment of limited disease includes

combination chemotherapy and radiotherapy. Also, surgery may play a role in TNM

stages I and II. In extensive disease, chemotherapy alone is the standard treatment.

However, despite the fact that patients initially have a good response to standard

treatment the vast majority relapse, with a 1-year survival rate of 40%, and 5-year

survival under 5% (Codony-Servat et al., 2016). These indicate that advances in the

treatment of SCLC remain non-satisfactory nowadays, and novel therapies are needed

to improve survival in this disease. For patients with NSCLC, the treatment options

such as surgery, radiotherapy, chemotherapy, targeted therapy or a combination of these

treatments are currently being used depending on the stage of cancer. In the early stage,

chemotherapy is often used as an adjuvant treatment, which is given after surgery or

radiation therapy to kill any remaining cancer cells (Domont, Soria, & Le Chevalier,

2005; Visbal, Leighl, Feld, & Shepherd, 2005). Chemotherapy is also used as

neoadjuvant therapy, which is given before surgery or radiation therapy to shrink tumor

(Choong & Vokes, 2005; Salvà & Felip, 2013). For later stage of cancers when surgery

is no longer an option, chemotherapy is often administered with simultaneous radiation

therapy. In addition, chemotherapy is used to treat recurrent cancer that comes back

after treatment or metastatic cancer that has spread to other parts of the body.

2.1.4 Molecular and genetic aspects of lung cancer

Lung carcinogenesis, like the development of other cancers, is a multistep

process involving the progressive accumulation of genetic and epigenetic alterations

that ultimately transform normal cells into neoplastic cells. Then, the neoplastic cells

can be benign tumors (non-cancerous) or malignant tumors (cancerous), which have

more aggressive characteristics than benign tumors. The common characteristics of


12

malignant cells that make them different from other normal cells include 1) self-

sufficiency of growth signals; 2) lack of sensitivity to anti-growth signals; 3) evasion

of apoptosis; 4) limitless replicative potential; 5) sustained angiogenesis; and 6) tissue

invasion and metastasis (Hanahan & Weinberg, 2011). Specific molecular alterations

that drive malignant progression involve mutations in genes that regulate cell

proliferation (Larsen & Minna, 2011). There are two broad classes of cancer-relevant

genes: proto-oncogenes and tumor suppressor genes.

2.1.4.1 Proto-oncogenes

In normal cells, proto-oncogenes are genes that control cell growth and

code for the proteins that provide a signal for stimulating cell division. Such proteins

can be classified into six groups based on functional properties as follows: 1) growth

factors (e.g. PDGF or EGF molecules); 2) growth factor receptors (e.g. PDGF receptor

or EGF receptor); 3) plasma membrane G proteins (e.g. Ras); 4) intracellular protein

kinases (e.g. Raf, MEK, MAPK); 5) transcription factors (e.g. Fos, Jun and Myc); and

6) cell cycle or cell death regulators (e.g. cyclin, cdk, bcl-2, Mdm2) (Kleinsmith, 2006),

as shown in Figure 2.1.

In cancer cells, the genetic changes found gain-of-function mutations

in proto-oncogenes. These changes cause proto-oncogenes to become oncogenes,

which produce a mutated protein that interferes cell proliferation, thereby leading to

uncontrollable cell division seen in cancer cells.


13

Figure 2.1 Signal transduction pathway that promotes cell division. (Modified from

Kleinsmith, 2006)

2.1.4.2 Tumor suppressor genes

In contrast to proto-oncogenes, tumor suppressor genes code for

negative regulator proteins that help prevent uncontrollable cell growth and promote

DNA repair and cell cycle checkpoint activation. Their normal functions are generally

to inhibit proliferation in response to certain signals such as DNA damage. The signal

is removed when the cell is fully equipped to proliferate. Tumor suppressor genes are

also broadly divided into two classes: gatekeeper genes and caretaker genes.

- Gatekeeper genes directly regulate cell growth by either inhibiting

cell proliferation or promoting apoptosis. Examples of gatekeeper genes include RB


14

involved in restriction point control; APC involved in Wnt signaling and p53 involved

in DNA damage response.

- Caretaker genes do not directly regulate cell growth. Instead,

inactivation of caretaker genes leads to genetic instability that indirectly promotes

proliferation by causing an increased rate of mutation. The genes that encode proteins

involved in DNA repair are classic examples of caretaker genes, such as ATM involved

in DNA damage response; BRCA1 and BRCA2 involved in double-strand break repair;

MLH1 and MSH2 involved in DNA mismatch repair; and XP-A involved in the

nucleotide excision repair pathway (Kleinsmith, 2006).

In cancer cells, the majority of genetic changes found loss-of-function

mutations in tumor suppressor genes. Such mutations contribute to the development of

cancer by inactivating the growth inhibitory function.

As illustrated in Table 2.2, the gene families implicated in lung

carcinogenesis include dominant oncogenes and tumor suppressor genes (Larsen &

Minna, 2011).


15

Table 2.2 Cancer genes and their functions found in lung cancer. (Modified from

Weber, 2007)

Genes Functions Mutation Common cancer type

1. Oncogenes

1.1 Growth factors

EGF Epidermal growth factor,

expressed in submaxillary

gland, targets epithelial/

mesenchymal/glial cells

Overexpression Breast cancer, lung

cancer

TGF-α Transforming growth

factor, expressed in

platelets targets epithelial/

mesenchymal/glial cells

Overexpression Breast

adenocarcinoma, lung

cancer

1.2 Growth factor receptors

ERBB Part of epidermal growth

factor receptor, receptor

protein tyrosine kinase

Point mutation

Amplification

Glioblastoma, breast

cancer, bladder cancer,

squamous cell lung

cancer, lung

adenocarcinoma,

head and neck cancer,

colon cancer

1.3 Signal transduction molecules associated with growth factor receptors

- Protein kinases

ERBB2

(Neu /Her-2)

Receptor protein tyrosine

kinase

Amplification,

point mutation

Neuroblastoma,

glioma, breast

adenocarcinoma,

ovarian cancer,

lung adenocarcinoma,

salivary gland cancer

- GTP-binding proteins

K-ras Guanine nucleotide-binding

protein GTPase

Point mutation

Translocation

Lung cancer, ovarian

cancer, colon cancer,

pancreas cancer


16

Genes Functions Mutation Common cancer type

- Transcription factors

c-myc Acts together with MAX,

sensitizes cells to CD95-

mediated apoptosis

Chromosomal

translocation,

Insertional

mutagenesis

Burkitt lymphoma,

leukemia, breast

cancer, stomach

cancer, lung cancer

L-myc Acts together with MAX Amplification Lung cancer

E2F

Dimer with DP1 initiates

transcription of S phase

genes

Point mutation

Lung cancer, breast

cancer

2. Tumor suppressor genes

2.1 Gatekeeper genes

(1) Signal transduction molecules associated with growth factor receptors

- Blockage of cyclin-cdk activity

CDKN1A

(waf1/cip1)

(p21CIP1)

Binding to and inhibition of

cdk2 and cdk4, activated by

p53, inhibits DNA synthesis

when complexed with

PCNA,

transcription induced by

STAT1

Leukemia, lung cancer

CDKN2A

(mts1)

Cyclin-dependent kinase

inhibitor 2A Multiple

Tumor Suppressor 1

Melanoma, lung

cancer,

medulloblastoma,

CDKN2B

(mts2)

(p15INK4b)

Inhibitor of cyclin-

dependent kinases

Acute lymphoblastic

leukemia, lung cancer,

melanoma, glioma

PPP2R1B

β form of the serine/

threonine Protein

Phosphatase 2A, down-

regulates MAP kinase

cascade, inhibits nuclear

telomerase activity

Lung cancer, colon

cancer


17

Genes Functions Mutation Cancers

- Transcription factors

p53

Transcription factor,

stimulates transcription of

p21, cell cycle regulator, is

phosphorylated by CDK and

Casein Kinase, induces

apoptosis via transport of

CD95 from the Golgi

complex

Osteosarcoma, breast

cancer, brain tumor,

Li-Fraumeni

syndrome, pancreas

carcinoma, small cell

lung cancer

Rb1 Negative regulation of

transcription factors E2F-

DP1, cell cycle regulation,

activity regulated by

phosphorylation (low in

G0/G1, high in G1/S)

Retinoblastoma,

osteosarcoma, small

cell lung cancer,

bladder cancer,

cervical carcinoma,

breast cancer, prostate

cancer

BRG1 Component of the SWI–SNF

chromatin remodeling

complex, inhibition of

proliferation through

interaction with RB

Prostate cancer, breast

cancer, lung cancer

(2) Function not grouped

S100 A2 Nuclear calcium-binding

protein

Breast cancer, lung

cancer

HIC-1 Located on chromosome

17p13.3, frequently

hypermethylated in cancer

Medulloblastoma,

lung cancer, colon

cancer

2.2 Caretaker genes

(1) Signal transduction molecules associated with growth factor receptors

- Inactivation of G-protein-GTP signal

RASSF1A RAS association domain

family 1 isoform A protein.

The encoded protein was

found to interact with DNA

repair protein XPA

Medulloblastoma,

nasopharyngeal

cancer, lung cancer


18

2.2 Alteration of cell cycle in cancer cells

2.2.1 Normal cell cycle regulation and cell cycle checkpoint

In multicellular organisms, cells divide into two daughter cells for growth

and replacement of dead cells. In normal cell division, a cell is stimulated by growth

factors to enter the cell cycle. The binding of a growth factor to its corresponding

receptors triggers a multistep cascade in which a series of signal transduction proteins

relay the signal throughout the cell. These signal transduction proteins are encoded by

proto-oncogenes and function to regulate cell growth and division (Kleinsmith, 2006).

The cell capable of undergoing division passes through the cell cycle, which is broadly

divided into three stages: interphase, mitosis, and cytokinesis. Interphase is the period

of cellular growth and DNA synthesis and is subdivided into three phases called G1

phase, S phase, and G2 phase. Mitosis or nuclear division is a continuous process and

is divided into five phases, namely prophase, prometaphase, metaphase, anaphase, and

telophase. These divided phases are based on progress made to a specific point in the

overall nuclear division. Cytokinesis or cytoplasmic division is the last stage that ends

with the separation into two daughter cells (Chandar & Viselli, 2012), as shown in

Figure 2.2. In the absence of growth factors, cells become quiescent, and cell division

is restrained by tumor suppressor proteins such as cdk inhibitors (CKIs), Rb, and so

forth.

Also, the cell-division cycle is strictly controlled by checkpoints located at

each phase of the cell cycle to verify whether the cells are ready to progress to the next

phase. The cell cycle checkpoints can be divided into four phases: restriction

checkpoint, DNA damage checkpoint, replication checkpoint, and spindle checkpoint

(Kleinsmith, 2006). These checkpoints are considered to be safety measures for the cell,

preventing the control system from dictating the start of another cell cycle event before

the previous one has finished, or before any damage to the cell has been properly

repaired.


19

Figure 2.2 Phases of the cell cycle. The cell cycle can be divided into three stages:

interphase, mitosis and cytokinesis. (Modified from Huber et al., 2013)

2.2.1.1 Cell cyle regulation

The cell cycle progression is strictly regulated by key regulatory proteins

known as cyclin-dependent kinases (cdks). Cdks are protein kinases, a class of enzymes

that regulate the activity of targeted protein molecules by catalyzing their

phosphorylation. However, cdks only exhibit protein kinase activity when they are

bound to their regulatory subunits, the cyclins. Each cdk is paired with a specific cyclin,


20

and the cyclins are made and degraded during specific points in the cell cycle (Figure

2.3 and Table 2.3). Thus, cell cycle progression is controlled by several Cdk-cyclin

complexes as follows: cdk4/6-cyclin D for G1 progression, cdk2-cyclin E for the G1-S

transition, cdk2-cyclin A for S-phase progression, and cdk1-cyclin B for entry into M-

phase (Nguyen & Jameson, 1998).

Table 2.3 Cyclin/CDKs complex are activated within specific phases of the cell cycle.

CDKs Cyclins Cell cycle phases

Cdk4 Cyclin D G1 phase

Cdk6 Cyclin D G1 phase

Cdk2 Cyclin E G1/S transition

Cdk2 Cyclin A S phase and G2 phase

Cdk1 (cdc2) Cyclin A G2/M phase

Cdk1 (cdc2) Cyclin B Mitosis

Figure 2.3 Changes in cyclins during the cell cycle. The levels of different cyclins are

depicted schematically. (Modified from Nguyen & Jameson, 1998)


21

Cyclin-cdk complexes are regulated in several ways, such as

phosphorylation and dephosphorylation, inhibitory proteins, proteolysis, as well as

subcellular localization.

One way to control the activity of cdk-cyclin complexes is phosphorylation.

The activity of the various cdk-cyclin complexes is controlled by reactions in which

cdk molecules are altered by phosphorylation and dephosphorylation. For example, at

the G2-to-M transition, the cdk1-cyclinB complex is initially inactive because of the

inhibitory phosphorylation of the cdk molecule by inhibiting kinase Wee1 on a

conserved tyrosine residue (Tyr15) or on an adjacent threonine residue (Thr14).

Although an activating phosphate group is added to a threonine residue (Thr161) by

cdk-activating kinase (CAK), the cdk remains inactive as long as the inhibitory

phosphate groups are present. The last step in the activation sequence is the removal of

the inhibiting phosphate by a specific enzyme called a protein phosphatase cdc25

(Gould & Forsburg, 2015), as shown in Figure 2.4.

Figure 2.4 Regulation of cdk1-cyclin complex by phosphorylation and dephosphorylation.

(Modified from Gould & Forsburg, 2015)

Conversely, whenever the cell cycle is in unfavorable conditions for

progression to the next phase, cdk inhibitors (CKIs) also regulate cdks to halt cell cycle

progression. Two classes of CKIs based on their structure and cdk specificity are

recognized. First, INK4 family members (e.g. p15 INK4b, p16INK4a, p18 INK4c and p19 INK4d)

specifically inhibit cdk4/6-cyclin D activity by binding to either cdk4 and cdk6, thereby


22

preventing association between cyclin D and its catalytic partner (Lim & Kaldis, 2013).

The other one is CIP/KIP family members (e.g. p21CIP1, p27KIP1 and p57 KIP2) that bind

and strongly block cdk-cyclin complexes (Lim & Kaldis, 2013). Most CKIs identified

so far act during G1 and/or S phase and block cell cycle progression until conditions

allow them to be overcome. For example, p16 CKIs present in G1 block the cell cycle

until enough G1 cyclins are synthesized to displace them from the G1 cdks.

In addition, the control of subcellular localization of cdk–cyclins and their

regulators is essential for proper cell-cycle coordination. One of the best-understood

examples is the regulation of cyclin B localization during interphase. During interphase,

cyclin B shuttles between the nucleus and the cytoplasm because constitutive nuclear

import is counteracted by rapid nuclear export, resulting that it is mainly located in the

cytoplasm (Hagting, Jackman, Simpson, & Pines, 1999). Just before the onset of

mitosis, cyclin B is phosphorylated in the cytoplasmic retention sequence (CRS),

leading to inactivation of nuclear export signals. Therefore, cyclin B accumulates in the

nucleus. It is possible that this type of regulation also serves to bring cdk complexes

into contact with their substrates (Gould & Forsburg, 2015). The periodic availability

of cyclins is a key mechanism of regulating the catalytic activity of cdk subunits.

Cyclins accumulate at certain periods of the cell cycle to activate their cdk partners. At

cell cycle transition points, cyclins become highly unstable and cyclin destruction

irreversibly compels the cell cycle forward. The abrupt instability of cyclins is due to

activation of ubiquitin ligases that target cyclins for proteasome-mediated degradation.

(1) Regulation of G1 phase progression

When a cell receives the proper signals that trigger the process of

cell division e.g. growth factors and cytokines, these signals lead to increased

expression of genes encoding proteins that regulate cell cycle progression through the

G1 phase. Cyclin D and cyclin E are two major classes of G1 cyclins. The cyclins,

however, have no effect on G1-S transition unless they form a complex with their cdk

partners as follows: cyclin D binds mainly to cdk4 and cdk6 whereas cyclin E binds to

cdk2. In early- to mid-G1 phase, cdk4/6-cyclin D complex hypophosphorylates the

retinoblastoma tumor suppressor protein (pRb) in which the hypophosphorylated pRb


23

also forms a complex with E2F family of transcription factors, resulting in inhibition

of the expression of genes required for entry into S phase, including cyclin E and cyclin

A. After progression through the cyclin D-dependent portion of the cell cycle, cyclin E

becomes activated by forming a complex with cdk2. The cdk2-cyclin E complex

triggers hyperphosphorylation of pRb, leading to the liberation of E2F to initiate the

transcription of genes needed for DNA replication (e.g. cdc6, ORC1 and the

minichromosome maintenance (MCM) proteins) and the progression into S phase (e.g.

cyclin E, cyclin A, cdk1 and cdc25A) (Xu, Sheppard, Peng, Yee, & Piwnica-Worms,

1994; Neganova & Lako, 2008; Foster, Yellen, Xu, & Saqcena, 2010). Moreover, the

activities of cyclin D- and cyclin E-dependent kinases are linked through members of

the CIP/KIP family of CKIs, including p21CIP1, p27KIP1 and p57 KIP2. These CKIs control

cell proliferation by binding to cyclin and cdk to block entry into S phase (Sherr &

McCormick, 2002) (Figure 2.5).

Figure 2.5 A schematic representation of various changes in the activity of cyclin-cdk

complex during the cell cycle. (Modified from Nguyen & Jameson, 1998)


24

(2) Regulation of S phase progression

Once cells enter S phase and begin DNA replication, cyclin E that

binds to cdk2 is rapidly degraded via the ubiquitin-dependent pathway, resulting in the

reduction of its kinase activity (Nguyen & Jameson, 1998; Hwang & Clurman, 2005).

However, the continued hyperphosphorylation of pRb allows the transcription of cyclin

A and cyclin B that required for subsequent phases of the cell cycle. Cyclin A has roles

in S phase progression, and it can form complex with either cdk2 or cdk1 under different

circumstances (Pagano, Pepperkok, Verde, Ansorge, & Draetta, 1992). In fact, although

cyclin A is synthesized and associated with cdk2 during the late G1 phase, its activity

is negatively regulated by inhibitory phosphorylation of cdk2 and also by the

association of CKIs, p21CIP1 and p27KIP1. Thus, the removal of inhibitory phosphates

from a cdk2 subunit of cdk2-cyclinA complex by the cdc25A phosphatases and the

degradation of CKIs by ubiquitin ligases are required for initiation of S phase.

Cdk2-cyclinA complex is required for the initiation of DNA

replication by the disassembly of pre-replication complex through the phosphorylation

of cdc6 proteins. Briefly, at the onset of S phase, a prereplication complex is formed on

DNA replication origins by the assembly of several factors, such as origin recognition

complex (ORC), cdc6, cdt1 and minichromosome maintenance (MCM) complex.

Several processes are responsible for activation of such prereplication complex. First,

cdk2-cyclinA complex phosphorylates cdc6. Phosphorylation is an inhibitory

modification of cdc6, and inactivated cdc6 gets ubiquitinated and degrades in

proteosome. Then, cdt1 becomes inhibited by geminin, which is an inhibitor of cdt1.

With cdc6 and cdt1 no longer bond, MCM protein can unwind the double-stranded

DNA, and DNA replication begins (Marín-García, 2011). In fact, although cyclin A is

synthesized and associated with cdk2 during the late G1 phase, its activity is negatively

regulated by inhibitory phosphorylation of cdk2 and also by the association with CKIs,

p21CIP1 and p27KIP1. Thus, the removal of inhibitory phosphates from cdk2 subunit of

cdk2-cyclin A complex by the cdc25A phosphatases and the degradation of CKIs by

ubiquitin ligases are required for initiation of S phase. Moreover, the cdk2-cyclin A

complex directly binds and phosphorylates E2F, thereby preventing the binding of E2F

to DNA and turning off the Gl/S phase genes that are no longer required once the DNA


25

replication has begun. The inactivation of E2F helps ensure cell cycle progression into

S phase and prevents reversion back to G1 phase (Nguyen & Jameson, 1998; Xu et al.,

1994). In late S phase, cyclin A also associates with cdk1 in which the cdk1-cyclin A

complex drives the transition between S phase and G2 phase (Pagano et al., 1992).

(3) Regulation of G2 phase progression

During G2 phase, the period between DNA synthesis and mitosis,

cdk1-cyclin A and cdk1-cyclin B complexes play an important role in G2 phase. The

accumulation of active cdk1-cyclin B, also known as maturation promoting factor

(MPF), is strictly dependent on cdk1-cyclin A activity. Briefly, the cdk1-cyclin A

complex phosphorylates cdh1, which is one of the substrate adaptor protein of the

anaphase-promoting complex (APC) that is an ubiquitin E3-ligase complex, leading to

preventing cdh1 from targeting cyclin B to the anaphase promoting complex (APC) for

ubiquitination and degradation. Active cdk1-cyclin B thus accumulates in the

cytoplasm, where it is thought to prepare structural components of the cell for the

upcoming cell division. The activity of cdk1-cyclin B complex is also controlled by

regulation of the nuclear transport of cyclin B. And the activity of cdk1 is regulated

positively by the phosphatase cdc25, which dephosphorylates tyrosine 14 and threonine

15, and negatively by the kinases Wee-1 and Myt-1, which phosphorylate these

residues. Myt-1 is cytoplasmic and phosphorylates threonine 14, while Wee-1 is

nuclear and phosphorylates tyrosine 15. Cdc25 is activated at the end of G2, leading to

permitting the cell to enter M phase (Weber, 2007).

(4) Regulation of M phase progression

In M phase, the activity of cdk1-cyclin B complex reorganizes the

microtubules and microfilaments, and phosphorylates proteins in the nuclear lamina,

resulting in the nuclear envelope breakdown, chromosome condensation, mitotic

spindle formation and fragmentation of the Golgi complex and endoplasmic reticulum.

Just before the breakdown of the nuclear membrane, the cdk1-cyclin B complex

translocates to the nucleus to target further substrates, including those that control the


26

shutdown of RNA polymerase III-mediated transcription. Finally, cyclin B and cyclin

A are rapidly degraded by the APC before the end of mitosis (Skaar & DeCaprio, 2006).

The APC is inactivated by the accumulation of G1 phase cdks (Nasmyth, 1996). The

mutual inhibition between APC and cdks explains how cells suppress mitotic cdk

activity during G1 and then establish a period with elevated kinase activity from S phase

until anaphase (Weber, 2007).

2.2.1.2 Cell cycle checkpoints

The cell cycle transition, which passes from one phase to another, is

regulated by checkpoints consisting of the restriction checkpoint, DNA damage

checkpoint, replication checkpoint, and spindle checkpoint (Kleinsmith, 2006), as

shown in Figure 2.6. These checkpoints monitor conditions within the cell and

transiently halt the cell cycle at various points for correction and repair (Elledge, 1996).

If cells cannot repair such damage, they are eliminated through apoptosis.

Figure 2.6 The cell cycle checkpoint. (Modified from Kleinsmith, 2006)


27

(1) Restriction checkpoint

Restriction checkpoint marks a key phase in the cell cycle, which

decides whether or not to proceed to mitosis. The cell cycle progression cannot begin

until the appropriate cellular growth has occurred during G1. Therefore, growth factors

are necessary to promote passage through the restriction point, which occurs in the late

G1 phase leading to S phase. Normally, retinoblastoma protein (pRb) functions as a

tumor suppressor to halt the cell cycle in the resting or G1 phase, by binding to a

transcription factor E2F. This restriction point is inactivated by cdk4/6-cyclin D

phosphorylation of pRb, with subsequent release of E2F that directs the synthesis of

proteins, allowing the cell cycle to proceed. Loss of restriction point control occurs in

many cancers and deregulates progression through the cell cycle.

(2) Replication checkpoint

Replication checkpoint is important for the integrity of the

genome. This checkpoint monitors the DNA replication during S phase to ensure that

DNA damage has been repaired or that DNA synthesis has been completed prior to

proceeding into M phase. For an entry into M phase, cdc25 removes inhibitory

phosphorylations from cdk1 to promote its activity. However, when the checkpoint is

engaged in response to DNA damage or incomplete DNA synthesis, cdc25 becomes

phosphorylated and is degraded, leading to no removal of the inhibitory

phosphorylations of cdk1. Thus, cdk1 remains inactive, thus preventing progression

through mitosis.

(3) Spindle checkpoint

Spindle checkpoint acts between the metaphase and anaphase

stages of mitosis. At the end of metaphase, the two sets of chromosomes are normally

lined up at the center of the mitotic spindle, and the anaphase-promoting complex

(APC) becomes active, triggering the onset of anaphase. Before chromosome

movement begins at the onset, the spindle checkpoint is invoked to make certain that


28

chromosomes are all properly attached to the mitotic spindle. If chromosomes are not

properly attached, a Mad-Bub protein complex is formed and subsequently inhibits the

APC by blocking its essential activators. For this reason, the cell cycle is temporarily

halted to allow the chromosomes to become attached properly and completely to the

spindle.

(4) DNA damage checkpoint

DNA damage checkpoint monitors DNA damage and halts the cell

cycle including late G1, S, and late G2 by inhibiting different cdk-cyclin complexes. In

this checkpoint, activated p53 protein plays a central role in which its accumulation in

response to DNA damage increases transcription of its target genes including p21. This

leads to inhibition of cdk-cyclin complexes in G1 and G2 phases and subsequent cell

cycle arrest, thus giving the cells time to repair DNA damage. Therefore, DNA damage

can be avoided before division to limit heritable mutation. If the damage cannot be

repaired, p53 may also trigger cell death by apoptosis.

2.2.2 Alterations of cell cycle regulation in cancer cells

In cancer cells, the accumulation of genetic alterations that involve cell

division leads to an unrestrained cell proliferation. A special subset of cancer-relevant

genes is represented by oncogenes and deregulated tumor suppressor genes. Activation

of proto-oncogenes to become oncogenes and/or inactivation of tumor suppressor genes

causes carcinogenesis. These genes encode proteins controlling cell growth,

proliferation and survival (e.g. growth factors, growth factor receptors, signal-

transduction proteins, transcription factors, pro- or anti-apoptotic proteins, cell-cycle

control proteins, and DNA repair proteins) and play roles in cancer induction (Lodish

et al., 2000).


29

2.2.2.1 Oncogenes

Oncogenes are mutated forms of the normal genes, proto-oncogenes.

The oncogenes contribute to converting a normal cell into a cancer cell, as these genes

encode proteins involved in growth signal transduction pathways, including growth

factor, growth factor receptors, proteins involved in signal transduction and nuclear

regulatory proteins (transcription factors) to be overactive. For example, the v-sis

oncogene of simian sarcoma virus, which encodes a growth factor homologous to

PDGF-β causes cells to overproduce growth factors, leading to stimulating cells to grow

(Fleming, Matsui, Molloy, Robbins, & Aaronson, 1989). Some oncogenes produce

either aberrant receptor proteins that release stimulatory signals into the cytoplasm even

when no growth factors are present in the environment or increased amount of receptor

proteins that results in increased signaling via the Ras-MAPK pathway, driving cellular

proliferation. For instance, the HER2/neu gene, which encodes transmembrane

receptors for growth factors, including EGFR, HER2, HER3, and HER4 (Burstein,

2005). Also, several proteins, which are encoded by oncogenes have their effect at the

cell membrane (e.g. the ras oncogene encodes guanine nucleotide-binding proteins

(G proteins), whereas some oncogenes act in the nucleus by binding to DNA. For

example, the myc oncogene encodes a transcription factor.

These oncogenes act as dominant genes because a mutation in only one

copy of the gene is sufficient carcinogenesis (Hunt & Dacic, 2008; Larsen & Minna,

2011). There are three main mechanisms that lead to the conversion of proto-oncogenes

to oncogenes, including mutations, gene amplification, and chromosomal

rearrangements as shown in Figure 2.7.


30

Figure 2.7 The mechanisms that lead to the conversion of proto-oncogenes to oncogenes.

(Modified from Lieberman, Marks, & Peet, 2012)


31

2.2.2.2 Deregulated tumor suppressor genes

Tumor suppressor genes are genes that are important in preventing

carcinogenesis. There are two major groups of tumor suppressor genes: gatekeeper

genes and caretaker genes (Kinzler & Vogelstein, 1997). Gatekeeper genes are

responsible for controlling or inhibiting cell proliferation by regulating the cell cycle

whereas caretaker genes are responsible for processes that ensure the integrity of the

genome by repairing DNA damage (Morris & Chan, 2015). However, the mutations of

these genes cause a loss or reduction in its function, leading to the induction of cancer.

For example, p53 is a well-known transcription factor that plays a crucial role in the

response of the cell to stress. The p53 is most often identified as a gatekeeper gene,

since it is directly involved in cell cycle regulation and cellular proliferation. Also, p53

is identified as a caretaker genes, as it involved in DNA repair mechanisms (Soussi &

Wiman, 2015). The mutations of this gene can grant cells with additive growth and

survival advantages, such as increased proliferation, evasion of apoptosis, and

chemoresistance (Rivlin, Brosh, Oren, & Rotter, 2011).

Tumor suppressor genes are recessive as they contribute to the

development of cancer when both copies of the gene are inactivated (Figure 2.8).

Genetic mechanisms driving to the loss of function of tumor suppressor genes can arise

at the nucleotide or chromosome level. These involve point mutation and losses of

genetic material by interstitial deletion, by the loss of arm or whole chromosome, by

unbalanced translocations, or passages in the homozygosity by mitotic recombination

or chromosome loss followed by reduplication (isodisomy) (Pierron, 2015).


32

Figure 2.8 The mechanisms that lead to the deregulation of tumor suppressor genes.

(Modified from Alberts et al., 2014)

In addition, cell cycle deregulation associated with cancer occurs through

mutation of genes encoding cdks, cyclins, cdk-activating enzymes, CKIs, cdk

substrates, and checkpoint proteins at different levels of the cell cycle (Vermeulen, Van

Bockstaele, & Berneman, 2003). Changes in levels and activity of these cell cycle

regulators result in uncontrolled cell division. For example, increased levels/activity of

positive regulators, e.g. cdks and cyclins, as well as decreased levels/activity of

inhibitors of the cell cycle, e.g. CKI, can promote cancer. Furthermore, cancer cells

often lose the cycle checkpoint integrity as a result of inactivation of CKIs and/or

overexpression of cdks and cyclins. Checkpoint dysfunction contributes unregulated

cell growth.


33

2.3 Classification of cell death

Cell death is a crucial process which plays an important role in controlling

development, homeostasis, and immune regulation of multicellular organisms. It also

protects the organism overall by removing all cells damaged by disease, aging,

infection, genetic mutation, and exposure to toxic agents (Saikumar & Venkatachalam,

2009). According to recommendations of the Nomenclature Committee on Cell Death

(NCCD) 2009, cell death can be classified according to its morphological appearance

e.g. apoptosis, necrosis and autophagy (Figure 2.9), enzymological criteria (with and

without the involvement of nucleases or of distinct classes of proteases, such as

caspases, calpains, cathepsins and transglutaminases), functional aspects (programmed

or accidental, physiological or pathological) or immunological characteristics

(immunogenic or non-immunogenic) (Kroemer et al., 2009).

2.3.1 Autophagy

Autophagy is a self-degradative physiological process that removes

unnecessary or dysfunctional cellular components through the actions of lysosomes.

Also, it is important for balancing sources of energy at critical times during

development and in response to nutrient starvation or other stresses (Parlato &

Mastroberardino, 2015). The specific morphological features of autophagy are defined

especially by transmission electron microscopy as follows: lack of chromatin

condensation and the presence of massive vacuolization of the cytoplasm. These

vacuoles, by definition, are two-membraned and contain degenerating cytoplasmic

organelles or cytosol. And the autophagic cell death has little or no association with

phagocytes (Kroemer et al., 2009).


34

Figure 2.9 Characteristics of autophagy, apoptosis and necrosis. (Modified from Tan,

Lu, Ji, & Mao, 2014)


35

2.3.2 Necrosis

Necrosis is a form of cell injury which results in the premature death of cells

in living tissue by autolysis, and it is defined as accidental cell death due to the actions

of external factors (e.g. infection, toxins, or trauma) on the cells or tissues that result in

a pathological process. The common features of necrosis are associated with cell

swelling, the rapid loss of membrane integrity and leak their intracellular components,

some of which serve as danger signals that stimulate inflammation (Rock & Kono,

2008). In contrast to apoptosis, necrotic cell death does not fragment into discrete.

2.3.3 Apoptosis

Apoptosis (or programmed cell death) is a mode of cell death that occurs to

remove unwanted cells, improperly functioning cells and injured cells without

damaging neighboring cells or inducing inflammation. Kerr, Wyllie, & Currie (1972)

have described the specific morphological features of cells undergoing apoptosis. The

features include chromatin aggregation, nuclear and cytoplasmic condensation, the

formation of membrane-bound vesicles (apoptotic bodies) which contain ribosomes,

morphologically intact mitochondria and nuclear material. Then, the apoptotic bodies

are rapidly recognized and phagocytized by either macrophages or adjacent epithelial

cells in which this phagocytic removal of apoptotic cells does not elicit an inflammatory

response (Travis et al., 2015). Also, the NCCD (2009) has formulated that apoptotic cell

death can occur with or without caspase activation (Kroemer et al., 2009).

Among three types of cell death, apoptosis is the most extensively

characterized mechanism for cancer cell killing as apoptosis is required to remove

abnormal cells without harming cells. Moreover, apoptosis has emerged as the major

mechanism by which anticancer agents act to eliminate cancer cells (Feng et al., 2011).

And the mechanisms and pathways of apoptosis are described in the topic 2.4.


36

2.4 Alteration of apoptotic cell death in cancer cells

2.4.1 Apoptosis in normal cells

Apoptosis or programmed cell death is a mode of cell death in normal

homeostasis to maintain cell populations in tissues during development and aging. It

also occurs as a defense mechanism such as in immune reactions or when cells are

damaged by diseases or toxic agents (Norbury & Hickson, 2001). There are two major

pathways of apoptotic cell death namely the death receptor pathway (extrinsic pathway)

and the mitochondrial pathway (intrinsic pathway) (Xu & Shi, 2007), as shown in

Figure 2.10. These two pathways are involved with several proteins especially a group

of caspase enzymes. Therefore, caspases are central components of the regulatory

mechanisms of the apoptotic pathway because they play a central role as both initiators

(caspases 8 and 9) and executioners (caspases 3, 6 and 7) (Ghavami et al., 2009), as

shown in Figure 2.10. Caspase-3 is the most common executioner among executioner

caspases. These caspases cause the disassembly of the genome by activating the

caspase-activated DNase (CAD), which preexisted in cells as an inactive complex with

the inhibitor of caspase-activated DNase (ICAD). The executioner caspases then cleave

ICAD, thus allowing CAD to degrade chromosomal DNA into oligonucleosomal

fragments. They also disable the normal DNA repair process by directly inactivating

two key proteins involved in maintaining genomic integrity, poly (ADP-ribose)

polymerase (PARP) and DNA-dependent protein kinase (DNA-PK). In addition, the

effector caspases cause nuclear shrinking and budding to form apoptotic bodies, as well

as the structural disassembly of the cell through the direct proteolysis of the

cytoskeleton and nuclear scaffold (Hsieh & Nguyen, 2005).


37

Figure 2.10 A schematic representation of intrinsic and extrinsic pathways of apoptosis.

(Modified from Ghavami et al., 2009)

2.4.1.1 The mitochondrial pathway (or intrinsic pathway)

The mitochondrial pathway or the intrinsic pathway is initiated in

response to DNA damage and involved with two major groups of Bcl-2 family proteins

based on their function (Green, 2015). The first group is antiapoptotic (also called pro-

survival) members that have four BH domains such as Bcl-2, Bcl-xL, Mcl-1, etc. The

second group is proapoptotic (or anti-survival) members, which are subdivided into two

groups: the proapoptotic proteins containing three BH domains (e.g. Bax and Bak) and

the proapoptotic BH3-only proteins carrying a single BH3 domain (e.g. Bim, Bad, Bid


38

and others). Different BH3-only proteins can elicit apoptosis by either inactivating pro-

survival functions of Bcl-2 and Bcl-xL or directly stimulating Bax and Bak. In cells that

are in healthy equilibrium, these proteins are located in the mitochondrial

intermembrane space, and antiapoptotic proteins (e.g. Bcl-2, Bcl-xL, Mcl-1) inhibits

proapoptotic functions of Bax/Bak, thus providing a pro-survival signal as the default

setting in the mitochondria.

In intrinsic pathway, p53, a tumor suppressor protein, is activated in

the cytosol in response to irreparable DNA damage and translocates to the nucleus

where it activates transcription of many proapoptotic proteins while simultaneously

repressing transcription of antiapoptotic proteins. As a result, the balance between pro-

and anti-apoptotic proteins shifts to favor proapoptotic proteins. In parallel, high

concentrations of p53 also translocate to mitochondria and bind to antiapoptotic

proteins, resulting in deactivated functions of these proteins (Strayer & Rubin, 2014).

Mitochondrial outer membrane permeabilization (MOMP) is a key

feature of the intrinsic pathway. During apoptosis, DNA damage–activated p53 triggers

increased levels of proapoptotic proteins, particularly BH3-only proteins. These

proteins inactivate anti-apoptotic proteins and also directly stimulate Bax and Bak,

which then oligomerize and form the pores, resulting in disruption of the outer

membranes of the mitochondria. Then, soluble proteins normally found in the space

between the inner and outer mitochondrial membranes are released, including

cytochrome c, apoptosis-inducing factor (AIF), endonuclease G (endoG), Smac/Diablo

and HtrA2/Omi. These mitochondrial proteins activate either caspase-dependent or

-independent cell death pathways (Saelens et al., 2004).

(1) Caspase-dependent apoptosis

In caspase-dependent apoptosis, following Bax/Bak forming pores

in the outer membrane, cytochrome c is released and then interacts with Apaf-1 to begin

the formation of the apoptosome. This apoptosome is a multiprotein complex

comprising Apaf-1, cytochrome c, and caspase-9, which functions to activate caspase-3

downstream of mitochondria in response to apoptotic signals In addition, Smac/Diablo

and HtrA2/Omi activate apoptosis by neutralizing the inhibitory activity of inhibitor


39

apoptotic proteins (IAPs) that inhibit caspases, leading to elevated activities of

executioner caspases-3, -6 and -7 (Chai et al., 2000; Du, Fang, Li, Li, & Wang, 2000).

The downstream caspases induce cleavage of protein kinases, cytoskeletal proteins,

DNA repair proteins, and inhibitory subunits of endonuclease as previously mentioned

(Kalimuthu & Se-Kwon, 2013).

(2) Caspase-independent apoptosis

In caspase-independent apoptosis, the soluble proteins including

AIF and endoG released from mitochondrial intermembrane space induce apoptosis in

a manner independent of caspase activities. AIF directly translocates to the nucleus and

triggers chromatin collapse and digestion into high molecular weight fragments. EndoG

also translocates to the nucleus where it cleaves nuclear chromatin to produce

oligonucleosomal DNA fragments (Bajt, Cover, Lemasters, & Jaeschke, 2006). In

addition, the use of caspase inhibitors, which block caspase-dependent apoptotic cell

death, cannot rescue these cells from caspase-independent apoptosis.

2.4.1.2 The death receptor pathway (or extrinsic pathway)

This pathway is triggered when specific death ligands of the TNF

family (e.g. TNF, Fas-ligand, TRAIL) engage their receptors (e.g. TNFR, Fas (also

called CD95 or APO-1), TRAIL-R1 and -R2 (also called DR4 and DR5), respectively

(Green, 2015). At the cell surface, death receptors become activated upon binding their

ligands. As a result, the cytoplasmic tails of these receptors bind the death domains of

docking proteins, to form a death-inducing signaling complex (DISC), leading to

subsequent stimulation of downstream procaspases-8 and -10, to become active forms.

In turn, these caspases activate executioner caspases-3, -6 and -7.

The extrinsic (death receptor) pathway intersects the intrinsic

(mitochondrial) pathway via caspase-8, which cleaves a cytoplasmic protein, Bid.

However, Bid is inactive unless it is proteolytically cleaved. Truncated Bid (tBid)

translocates to mitochondria and activates Bax and Bak to cause MOMP and apoptosis

via mitochondrial pathway (Ghavami et al., 2009; Green, 2015).


40

2.4.2 Apoptosis in cancer cells

Insufficient apoptosis can lead to the development of cancer. Indeed, the

mutated cells that escape the apoptotic control typically ignore normal cellular signals

and become more proliferative than normal, allowing them to change to neoplastic cells

and even malignant cells rapidly. These cells can acquire reduction in apoptosis or

resistance to apoptosis through the mechanisms as follows: 1) impaired death receptor

signaling, e.g. reduced expression of death receptors or signals as well as expression of

decoy receptor without death domain, 2) defects or mutations in p53, 3) reduced

expression and function of caspases, 4) increased expression of IAPs and 5) disrupted

balance of pro-apoptotic and anti-apoptotic proteins, e.g. overexpression of anti-

apoptotic proteins or/and underexpression of pro-apoptotic proteins (Wong, 2011).

2.4.3 Reactive oxygen species (ROS) leading to apoptosis in cancer cells

Reactive oxygen species (ROS) are chemically reactive molecules that

contain oxygen, and they are broadly classified into two groups: free radicals and

nonradicals. The free radicals, including superoxide anion (O2•−), hydroxyl radical

(•OH), hydroperoxyl radical (HO2•−), peroxyl radical (ROO•) and alkoxyl radical (RO•),

are molecules that possess one or more unpaired electrons in their outer orbital (Islam

& Shekhar, 2015). Therefore, they are highly reactive, as they can either donate an

electron to or accept an electron from other molecules to achieve stability (Lobo, Patil,

Phatak, & Chandra, 2010). The nonradicals, including hydrogen peroxide (H2O2),

hypochlorous acid (HOCl), hypobromous acid (HOBr), ozone (O3) and singlet oxygen

(1O2), are molecules that do not have unpaired electron(s) but are chemically reactive

to generate free radicals under certain conditions with or without enzymatic catalysis

(Trachootham, Alexandre, & Huang, 2009; Shi, Zhang, Zheng, & Pan, 2012). ROS are

generated by either endogenous or exogenous sources. The former sources include

oxidases, peroxidases and oxygenases. In the cells, these intracellular enzymes bind O2

and transfer single electrons to it via a metal. Before the reaction is complete, such

reaction may accidentally release free radical intermediates. The latter sources include

environmental pollution, radiation, cigarette smoking, certain foods and drugs


41

(Bhattacharyya, Chattopadhyay, Mitra, & Crowe, 2014). Normally, ROS are

effectively neutralized by the potent antioxidant system. However, when ROS are

produced beyond the antioxidant capacity of the cells, they promote genotoxic damage

and thereby cancer progression (Al-Khayal et al., 2017). Nevertheless, exogenous

administration of ROS, especially via the use of chemotherapeutic drugs, leading to

apoptotic induction has become a potential mechanism of action in eliminating cancer

cells (Ahn et al., 2014).

Several studies have shown that various anticancer drugs e.g. doxorubicin,

azidothymidine (AZT), cisplatin (Deavall, Martin, Horner, & Roberts, 2012) produced

ROS at excessive levels, resulting in irreparable DNA damage, subsequently leading to

apoptotic cell death. Moreover, recent studies have revealed that some of the plant

natural compounds e.g. resveratrol and curcumin (Singh et al., 2012), neferine

(Poornima, Quency, & Padma, 2013), capsaicin (Bu et al., 2015) as well as isoliensinine

(Zhang et al., 2015) induce apoptosis through ROS-mediated c-Jun N-terminal protein

kinase (JNK) and p38 MAPK pathways. These two kinases are stress-activated protein

kinases (MAPK). Both JNK and p38 MAPK are then activated through apoptosis

signal-regulating kinase-1 (Ask-1), whose activity is regulated by its interaction with

thioredoxin, which is a redox-regulated protein (Saitoh et al., 1998). Under normal

conditions, thioredoxin directly binds to the N-terminal noncatalytic region of Ask-1,

resulting in inhibiting kinase activity. However, in response to ROS, the oxidized form

of thioredoxin dissociates from Ask-1, thus allowing Ask-1 activation (Saitoh et al.,

1998; Liu & Min, 2002). Ask-1 signaling activates downstream MAPK kinases that

promote activation of JNK and p38 MAPK signaling pathways (Figure 2.11). Activated

JNK readily translocates to mitochondria to phosphorylate a tumor suppressor protein

p53, which activate proapoptotic proteins, such as members of the BH3-only subgroup

of the Bcl-2 family (e.g., Bid and Bim), or suppress the activity of antiapoptotic Bcl-2

and Bcl-xL proteins It is also associated with the overexpression of proapoptotic Bax,

leading to formation of Bax homodimers resulting in MOMP, the release of cytochrome

c from the inner mitochondrial membrane, apoptosome formation and finally induction

of apoptosis (Zhang, Humphreys, Sahu, Shi, & Srivastava, 2008; Liou & Storz, 2010).

However, in normal cells, levels of p53 are usually kept low by its association with a


42

protein called Mdm2, which binds p53 and transports it from the nucleus to the cytosol

for proteolytic degradation by the proteasome (Shadfan, Lopez-Pajares, & Yuan, 2012).

Figure 2.11 JNK/p38 MAPK signaling pathways, apoptosis pathway, and multiple

molecular targets of plant-derived agents.


43

2.5 Targeting for cancer treatment

2.5.1 Targeting cell cycle regulators in cancer treatment

The knowledge of the molecular mechanisms of cell cycle regulation is

important to control the aberrant proliferation including DNA replication and accurate

segregation of chromosomes to daughter cells, which are characteristically aberrant in

cancer cells (Ganem, Storchova, & Pellman, 2007). Many anticancer agents such as

flavopiridol (Shapiro & Harper, 1999), rifampicin (Zhuang et al., 2011), paclitaxel

(Choi & Yoo, 2012) can slow down cell division by inducing cell cycle arrest in the

G0/G1, S, or G2/M phases (Hsiao et al., 2014). They can target cdks, which are required

at different phases of the cell cycle, and inhibit the function of cdk by directly binding

the catalytic cdk subunit or indirectly targeting regulatory pathways governing cdk

activity (Schwartz & Shah, 2005). In addition, several studies have revealed that many

anticancer agents and plant-derived compounds exerted their cytotoxic effect through

arresting of the cell cycle in each phase, depending on their selectivity, as follows:

- Compounds that have been shown to arrest G0/G1 phase. For

example, curcumin derived from turmeric (Mukhopadhyay et al., 2002); tangeretin

derived from the peel of citrus fruits (Pan, Chen, Lin-Shiau, Ho, & Lin, 2002); honokiol

derived from Magnolia officinalis/grandiflora (Hahm & Singh, 2007); lycorine, a

natural alkaloid extracted from Amaryllidaceae (Li et al., 2012) and glycyrrhetinic acid

derived from glycyrrhiza (Zhu et al., 2015) have been shown to modulate the activities

of several key G1 regulatory proteins via down-regulation of cdks (e.g. cdk4, cdk6) and

cyclins (e.g. cyclin D, cyclin E) but up-regulation of the tumor suppressor protein p53.

- Compounds that have been shown to arrest S phase. For

example, crude seed extract of celery (Apium graveolens L) has been shown to down-

regulate cdk2 and cyclin A (Gao et al., 2011). Besides altering expression levels of

regulatory proteins in S phase, some anticancer agents are also incorporated directly

into DNA or RNA, and disrupt DNA synthesis during S phase, such as 5-fluorouracil,

Cytarabine, and Methotrexate, which inhibits thymidylate synthase, DNA polymerase

and dihydrofolate reductase, respectively (Payne & Miles, 2008).


44

- Compounds that have been shown to arrest G2/M phase. For

example, jaceosidin, a flavonoid present in plants of genus Artemisia (Khan, Rasul, Yi,

Zhong, & Ma, 2011); myricetin, a naturally occurring flavonol widely presented in

fruits, vegetables, tea, berries and red wine (Zhang, Zou, Xu, Shen, & Li, 2011); and

curcumin (Cheng et al., 2016) have been shown to down-regulate the expression levels

of regulatory proteins of G2/M phase such as cyclin B1 and cdk1 but up-regulate p53

and p21. Moreover, Ouyang et al., (2009) have shown that genistein, a major

isoflavonoid, can increase the phosphorylation and activation of checkpoint kinases

(Chk1 and Chk2), which results in the phosphorylation and inactivation of phosphatases

(cdc25C and cdc25A), and thereby the phosphorylation and inactivation of cdc2, which

arrests cells at G2/M phase. Besides altering expression levels of regulatory proteins in

G2/M phase, some anticancer agents can bind to tubulin and lead to disrupting the

spindle apparatus of the microtubules required for chromosome segregation in M phase

(Stanton, Gernert, Nettles, & Aneja, 2011). The microtubules consist of -tubulin and

ß-tubulin and carry out polymerization and depolymerization. Altering a dynamic

balance between polymerization and depolymerization is a target for cancer drug

development (Shin et al., 2008). Certain compounds can inhibit or halt cell cycle by

interfering with microtubules during mitosis or M phase. Examples of drugs in this

category are vinca alkaloids and taxanes, which interrupt cell division by agitation of

microtubule dynamics. Vinca alkaloid (e.g. vinblastine and vincristine) binds to the

unpolymerized tubulin molecules and prevents them from polymerizing into a growing

microtubule. Taxol binds to tubulin within assembled microtubules and prevents

disassembly (Castedo et al., 2004; Mollinedo & Gajate, 2003).


45

2.5.2 Targeting apoptosis in cancer treatment

The discovery of various types of defects (e.g. impaired death receptor

signaling, defects or mutations in p53, reduced expression and function of caspases,

increased expression of IAPs and disrupted the balance of proapoptotic and

antiapoptotic proteins) in the apoptotic pathways becomes an interesting target for

cancer treatment (Wong, 2011). Anticancer agents including plant-derived compounds

can restore the apoptotic signaling pathways towards normality to eliminate cancer cells

acquiring these defects, as follows:

- Some plant compounds have been reported to induce the

extrinsic apoptotic pathway by up-regulation of death receptors, e.g. Fas/CD95, DR4

and DR5 or their corresponding ligands, e.g. Fas ligand, TRAIL-R1, and -R2,

respectively. For example, triterpenediol, which comprises of isomeric mixture of 3α,

24-dihydroxyurs-12-ene and 3α, 24-dihydroxyolean-12-ene from Boswellia serrate

(Bhushan et al., 2007); γ-humulene derived from Emilia sonchifolia (Lan et al., 2011)

and curcumin (Jung et al., 2005) stimulate the clustering of DR4/DR5, TNF-R1 and

increasing of associated FADD protein levels, leading to caspase-8 and caspase-3

activation.

- Some plant compounds can restore the intrinsic apoptotic

pathway by altering the balance of pro- and antiapoptotic proteins. For example, butein,

a polyphenol derived from Dalbergia odorifera (Kim et al., 2001); curcumin derived

from Curcuma longa and the root extract of Salvia miltiorrhiza (Duval, Moreno-

Cuevas, Gonzalez-Garza, Rodriguez-Montalvo, & Cruz-Vega, 2014) as well as the

hexane extract of the leaves from Ferulago angulata (Karimian et al., 2014) have been

shown to decrease the levels of antiapoptotic proteins (e.g. Bcl-2, Bcl-xL) while it

increased the levels of proapoptotic proteins (e.g. Bax and Bak). These proapoptotic

proteins directly form pores in the mitochondrial outer membrane leading to the release

of cytochrome c. Cytochrome c induced apoptosis by activation of caspase 9 and 3,

respectively. In addition, some plant compounds such as flavokawain C, found in Kava

(Piper methysticum Forst), caused disruption of mitochondrial membrane potential,

leading to the release of AIF, Smac/DIABLO and cytochrome c from the mitochondria


46

(Phang, Karsani, Sethi, & Abd Malek, 2016). These proteins then activate various

caspases (-3, -8, -9) and subsequent PARP cleavage, leading to apoptotic cell death.

Furthermore, the action of several plant-derived compounds through high

levels of ROS production is an important mechanism to eliminate cancer cells. This

excessive ROS formation can induce oxidative stress, which affects DNA damage and

leads to cell death. Briefly, in response to DNA damage, p53 is stabilized and then acts

to regulate the expression of stress response genes in DNA repair, cell-cycle arrest, and

apoptosis, in order to suppress cancer cell proliferation (Circu & Aw, 2010). Recently,

some of plant natural compounds such as curcumin (an active ingredient of turmeric),

triterpenediol (an isomeric mixture of 3α, 24-dihydroxyurs-12-ene and 3α, 24-

dihydroxyolean-12-ene from Boswellia serrate), shikonin (a naphthoquinone isolated

from Lithospermum erythrorhizon), and grape seed extract (a complex mixture of

polyphenols known as proanthocyanidins) have been shown to induce cancer cell death

through massive ROS generation, leading to an induction of intrinsic (including

caspase-dependent and caspase-independent) and extrinsic pathways of apoptosis

(Derry, Raina, Agarwal, & Agarwal, 2013; Bhushan et al., 2007; Wu et al., 2010; Gong

& Li, 2011).


47

2.6 Thai medicinal plants (Hua-Khao-Yen)

Thai medicinal plants locally known as “Hua-Khao-Yen” have widely been

used to prepare Thai traditional medicine. Five species of Hua-Khao-Yen including

Dioscorea membranacea Pierre, Dioscorea birmanica Prain ex Burkill, Smilax

corbularia Kunth, Smilax glabra Roxb and Pygmaeopremna herbacea Roxb have been

extensively used by Thai folk doctors for the treatment of cancers, AIDS, septicemia,

inflammation and lymphatic diseases. The preparation of plant extracts for treating such

diseases in Thai traditional medicine is usually made by boiling in water or by soaking

in alcohol (Pongbunrod 1976; Tungtrongjit 1978). Among these species, D.

membranacea Pierre has been the most widely used to prepare Thai traditional anti-

cancer medicine. Itharat et al. (2004) have reported that its aqueous and ethanolic

extracts were more cytotoxic against cancer cell lines than other species.

2.6.1 Dioscorea membranacea Pierre

2.6.1.1 General description

Dioscorea membranacea Pierre, also called Hua-Khao-Yen-Tai, is a

member of the Dioscoreaceae family. This plant is scattered in open areas of mixed

deciduous forests to lower montane evergreen forests, often on limestone; 50-800 m,

and its distribution in Thailand, Vietnam, Laos, and Myanmar (Wilkin, 2009). The

general characteristics were described by Wilkin and Thapyai (Thapyai, 2004; Wilkin,

2009) as following: Climber to 10 m. rhizomes are 4-8 long by 1-5 cm wide, branching

and spreading, dark brown, shallowly horizontally buried, periderm hard but lacking

rigid, spine-like roots. Stems are 2-5 mm in diameter, twining to the left, unarmed.

Leaves simple, alternate, blades ovate to broadly so, membranous to thinly chartaceous

and usually translucent when dried, 7–9(–11)-veined, base shallowly to deeply cordate

apex acuminate, margins usually deeply 3-lobed, sometimes shallowly so or entire

(reproductive shoots only) or with additional lobes (towards stem base); petioles 2–13

cm long; cataphylls and bulbils absent; lateral nodal spines usually present on either

side of each node, sometimes absent (especially on distal shoots) or broken off in


48

specimen preparation, curved, hard and rather brittle, to 4 mm long. Inflorescences

spicate, pendent, axes sometimes finely tuberculate, tepals inserted on a cup-shaped

torus, erect, apices not recurved, fused for 1/2 to 2/3 of their length. Male inflorescences

simple or compound, compound inflorescences 1(–2) per axil, simple/partial

inflorescences (Pl. 17C) 1–2(–3) per axil, peduncles 2.2–25 mm long, axes 4–33.5 cm

long, with an apparently sessile cymule of (1–)2–3 flowers at each node; female

inflorescences 1(–2) per axil, simple. Male flowers are 0.8–1.4 mm in diam. at anthesis,

outer tepals 1.9–2.3 by 0.6–1, narrowly obovate to obovate-oblong, inner tepals 1.9–

2.2 by 0.6–0.8 mm, obovate to obovate-oblong, stamens 6. Capsules 18–21 by 25–40

mm. Seeds 3–6 by 5–7 mm, ovoid-lenticular; wings 17–18 by 15–17 mm, extending all

around seed margin. The characteristics of D. membranacea Pierre are shown in Figure

2.12, 2.13 and 2.14.

Figure 2.12 The characteristics of D. membranacea Pierre. (Original picture)


49

Figure 2.13 D. membranacea Pierre (Male plant). (Thapyai, 2004)

Note: A. Male fluorescence; B. primary bract; C. partial male inflorescence, showing

cymose pattern; D. cymular bract; E-I. male flower; E. l-section stamens and pistillode;

F. floral bract; G. bracteole; H. outer tepal; showing stamen adnation, I. inner tepal with

stamen adnation; J. rhizome.


50

Figure 2.14 D. membranacea Pierre (Female plant). (Thapyai, 2004)

Note: K. inflorescence; L–Q. female flower; L. side view; M. l-section (excluded ovary)

showing staminodes, style and stigmas; N. floral bract; O. bracteole; P–Q. outer and

inner tepal respectively, with staminode adnation; R. infructescence; I. mature capsule,

l-section showing seed position; K. seed.


51

2.6.1.2 Biological activities

Dioscorea membranacea Pierre has shown numerous biological

activities such as anticancer activity (Itharat et al., 2004; Itharat et al., 2014), anti-

allergic activity (Tewtrakul & Itharat, 2006), anti-HIV activity (Tewtrakul, Itharat, &

Rattanasuwan, 2006), antioxidant activity (Itharat et al., 2007) and anti-inflammatory

activity (Tewtrakul & Itharat, 2007).

(1) Antiproliferative activity

The previous study has shown that the crude ethanolic extract of

D. membranacea Pierre exhibited high cytotoxic activity against lung, breast and colon

cancer cell lines (COR-L23, MCF-7 and LS-174T, respectively) whereas its water

extracts exhibited high cytotoxic activity against breast and colon cell lines. Both the

extracts had no cytotoxic effects against keratinocyte normal cell line (SVK-14) using

the SRB assay (Itharat et al., 2004). As shown in Figure 2.15, nine compounds were

isolated by bioassay-guided fractionation from the ethanolic extract of rhizomes of

D. membranacea Pierre (Itharat 2002; Itharat et al., 2003). They include two

naphthofuranoxepins (e.g. dioscorealide A and dioscorealide B), one phenanthraquinone

(e.g. 1,4-phenanthraquinone or dioscoreanone), three steroids (e.g. β-sitosterol,

stigmasterol and β-sitosterol-3-O-β-D-glucopyranoside), three steroid sapogenins (e.g.

diosgenin 3-O-α-L-glucopyranosyl (1→2)- β-D-glucopyranoside, diosgenin 3-O-β-D-

glucopyranosyl (1→3)-β-D-glucopyranoside and diosgenin). Among these compounds,

dioscorealides B, dioscoreanone and diosgenin 3-O-α-L-rhamnopyranosyl (1→2)-β-D-

glucopyranoside exerted cytotoxic activity against lung, breast and colon cancer cell

lines (Itharat et al., 2003). Moreover, dioscorealide B (Saekoo, Dechsukum et al. 2010;

Saekoo, Graidist et al. 2010) and dioscoreanone (Hansakul et al., 2014) were studied

on the molecular mechanism underlying the anticancer activity in cancer cell lines

through induction of apoptosis.


52

Figure 2.15 Chemical structures of isolated compounds from the rhizomes of

D. membranacea Pierre. (Itharat 2002; Itharat et al., 2003)

dioscorealide A (1) dioscorealide B (2) dioscoreanone (3)

β-sitosterol (4) stigmasterol (5)

diosgenin 3-O-β-D-glucopyranosyl

(1→3)- β-D-glucopyranoside (6)

β-sitosterol-3-O-β-D-glucopyranoside (7)

diosgenin (9) diosgenin 3-O-α-L-glucopyranosyl

(1→2)- β-D-glucopyranoside (8)


53

In 2014, three dihydrophenanthrene compounds were subsequently

isolated from the ethanolic extract of rhizomes of D. membranacea Pierre including 5,6-

dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene, 5-hydroxy-2,4,6-trimethoxy-9,10-

dihydrophenanthrene and 5,6,2-trihydroxy-3,4-methoxy-9,10-dihydrophenanthrene

(Figure 2.16). One of them was 5,6-dihydroxy-2,4 dimethoxy-9,10-dihydrophenanthrene

that showed higher selective cytotoxicity against lung, breast and prostate cancer cell

lines (COR-L23, MCF-7 and PC3 cell lines, respectively) than other compounds using

SRB assay (Itharat et al., 2014).

Figure 2.16 Chemical structures of isolated compounds from the rhizomes of

D. membranacea Pierre. (1) 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene,

(2) 5-hydroxy-2,4,6-trimethoxy-9,10-dihydrophenanthrene and (3) 5,6,2-trihydroxy-

3,4-methoxy-9,10-dihydrophenanthrene. (Itharat et al., 2014)

(2) Anti-allergic activity

D. membranacea Pierre has been used in Thai traditional medicine

for treatment of allergy and allergy-related diseases as claimed by Thai folk doctors.

The previous study has shown that the ethanolic extract of D. membranacea Pierre

exhibited potent inhibitory activity against β -hexosaminidase release as a marker of

degranulation in rat basophilic leukemia mast cells (RBL-2H3). In addition, four

compounds isolated from this crude ethanolic extract including dioscorealide A,

dioscorealide B, dioscoreanone and diosgenin are suggested to be the active ingredients

of this plant as anti-allergic agents (Tewtrakul and Itharat 2006).

1

3

4a 5

7 8a

9

10a

10

4b 1

3

4a 5

7 8a

9

10a

10

4b 1

3

4a 5

7 8a

9

10a

10

4b

(1) (2) (3)


54

(3) Anti-HIV activity

Traditional doctors have used D. membranacea Pierre in various

traditional Thai herbal remedies for treating HIV-infected persons. In 2006, the

ethanolic and water extracts of five species of Hua-Khao-Yen were investigated for

their inhibitory effects against HIV-1 protease (HIV-PR) and HIV-1 integrase (HIV-1

IN) (Tewtrakul, Itharat, & Rattanasuwan, 2006). The results showed the ethanolic

extract of S. corbularia exerted the most potent activity against HIV-1 IN whereas that

of D. membranacea Pierre possessed HIV-1 PR inhibitory effect. This data suggest the

combined usage of both plants in AIDS treatment.

(4) Antioxidant activity

Among five species of Hua-Khao-Yen, the ethanolic extract of D.

membranacea Pierre rhizomes possessed highest antioxidant activity using the lipid

peroxidation of liposomes assay (Itharat, 2010). In addition, an active compound

isolated from the ethanolic extract of this plants such as dioscoreanone also showed the

highest antioxidant activity using DPPH assay (Itharat et al., 2007).

(5) Anti-inflammatory activity

The ethanolic extract of D. membranacea Pierre rhizomes has

shown the anti-inflammatory activity in the inhibition of lipopolysaccahride (LPS)-

induced nitric oxide production in RAW264.7 cell lines. In addition, three active

ingredients of D. membranacea Pierre such as diosgenyl-3-O-α-L-rhamnopyranosyl

(1→2)-β-D-glucopyranoside, dioscoreanone and dioscorealide B are also active

principles for NO inhibitory activity, and only dioscoreanone showed potent inhibitory

effect on TNF-α release (Tewtrakul & Itharat, 2007). The anti-inflammatory activity of

the aqueous and ethanolic extracts from the rhizomes of D. membranacea Pierre was

also studied in in vivo experiments using carrageenin-induced paw edema in rats. The

results demonstrated that oral administration of both extracts at the dose of 1600 mg/kg

significantly decreased the paw edema induced by carrageenin in rats, indicating their


55

anti-inflammatory activity (Reanmongkol, 2007). Such data support the use of D.

membranacea Pierre by Thai folk doctors for treatment of the inflammatory diseases.

2.6.2 5,6-Dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene

An active compound, 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophe

nanthrene, is isolated from the ethanolic extract of D. membranacea Pierre rhizomes

(Itharat et al., 2014), as shown in Figure 2.13. It is white to pale yellow solid. The

molecular formula of this compound is C16H16O4, and its molecular weight (M.W.) is

272.1049 g/mol. The chemical structure was shown in Figure 2.17. Moreover, in 2014,

Itharat et al. have demonstrated that this compound exhibited the highest cytotoxicity

activity on human large cell lung carcinoma cell line COR-L23, human breast cancer

cell line MCF-7 and human prostate cancer cell line PC-3 but less cytotoxicity activity

on the human lung fibroblast cell line MRC-5. However, the molecular mechanisms

underlying its cytotoxic effect have not yet been studied.

Figure 2.17 The structure of 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene.


56

CHAPTER 3

RESEARCH METHODOLOGY

3.1 Conceptual framework of this study

Isolation of HMP

- Column Chromatography

- Thin-Layer Chromatography (TLC)

Structural analysis of HMP

- Nuclear magnetic resonance (NMR)

Determination of antiproliferative effects of HMP against cell lines

- Sulforhodamine B (SRB) assay

Normal cell lines Non-small cell lung cancer Small cell lung cancer

- A549

- NCI-H226

- COR-L23

- NCI-H1688 - MRC-5

Extraction of D. membranacea Pierre rhizomes

- Maceration in 95% ethanol


57

The conceptual framework of this study (cont.)

Bax, Bcl-2

Caspase3, PARP

Bid, Caspase9

Bax, Bcl-2

Caspase 3 activity Kit

Z-VAD fmk

Estimation of cell proliferation dynamics

- CellTrace™ CFSE Cell Proliferation Kit

Assessment of growth inhibitory and cytotoxic effects

- Sulforhodamine B (SRB) assay

Determination of apoptotic cell death

Flow cytofluorometric analysis of cell cycle distribution

- Staining with Propidium Iodide (PI)

Cell cycle arrest (G2/M arrest) Apoptotic cell death (sub-G1)

Detection of PS on external

leaflet membrane of early

apoptotic cells

- Annexin V-FITC/PI

staining

Protein analysis (Western blotting)

- - cdc25, cdk1, cyclinB1

Mitotic spindle disruption

- Tubulin polymerization assay Kit

Apoptosis morphology

- DAPI Staining

Enzyme activity analysis RNA analysis

Protein analysis

NAC

ROS generation

NAC

Study on the anticancer effects of HMP on A549 cell line

General caspase inhibitor test

- Z-VAD fmk

DNA fragmentation

DNA ladder


58

3.2 Extraction of Dioscorea membranacea Pierre

Dioscorea membranacea Pierre (DM) collected from Phetchabun province,

Thailand was used in this study. Authentication of plant materials was carried out at the

Herbarium of the Department of Forestry, Bangkok, Thailand, where the herbarium

vouchers are deposited (SKP A062041305). Assoc Prof. Dr. Arunporn Itharat,

Department of Applied Thai Traditional Medicine, Faculty of Medicine, Thammasat

University kindly provided it. In the extraction procedure, the rhizomes of D.

membranacea Pierre were cleaned, cut into small pieces and dried at 50 ºC (Figure 3.1).

Dried plant material (204.8 g) was macerated in 95% ethanol for 3 days at room

temperature. The extract was filtered through filter paper, and the supernatant was

evaporated to dryness by a rotary evaporator. Maceration of the residue was repeated

two times. The percent yield of the ethanolic extract of D. membranacea Pierre was

calculated.

Figure 3.1 The physical characteristics of the rhizome of D. membranacea Pierre.

(Original picture)


59

3.3 Isolation of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene

Isolation procedures of an active ingredient of D. membranacea Pierre

namely 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene (HMP) were kindly

supplied by Dr. Srisopa Ruangnoo, Department of Applied Thai Traditional Medicine,

Faculty of Medicine, Thammasat University. Briefly, the crude ethanolic extract of D.

membranacea Pierre was subjected to column chromatography on silica gel 60 (0.040-

0.063 mm) (Merck, Germany) as the stationary phase. The column was eluted by

gradient elution in increasing order of polarity. Fractions were collected in a 15-ml tube,

analyzed by thin layer chromatography (TLC) using aluminium sheets coated with

silica gel 60 (Merck, Germany) and visualized with acidic anisaldehyde spray. The

fractions containing similar spots were combined and evaporated to dryness under

reduced pressure. The combined fractions containing HMP were further separated by

additional silica gel column chromatography and/or by silica gel TLC glass plates,

followed by silica gel TLC aluminum sheets for analysis. The combined fractions or

scraped bands that contained one spot corresponding to 5, 6-dihydroxy-2, 4-dimethoxy-

9, 10-dihydrophenanthrene was checked for its purity using silica gel TLC aluminium

sheets 60 in three different solvent systems of varying polarity and High-performance

liquid chromatography (HPLC) technique using an Agilent Technologies 1200 Series

HPLC system (Agilent Technologies, USA). For HPLC analysis, 1 mg/ml of the

compound was achieved on a C18 reversed-phase HPLC column (250 x 4.60 mm 5

micron) (Phenomenex, USA) using water (eluent A) and acetonitrile (eluent B) as

mobile phase with the following gradient: 70-55% A at 0-10 min, 55% A for 5 min, 55-

30% A at 15-30 min, 30% A for 5 min, 30-70% A at 35-37 min and 70% A for 8 min.

The flow rate was 1 ml/min, and the injection volume was 10 μl. The ultraviolet (UV)

detector was used to detect the peak area of such compound with the fixed wavelength

at 254 and 270 nm.

Moreover, the chemical structure of the isolated compound was sent to

Bioresources Research Laboratory, National Center for Genetic Engineering and

Biotechnology (BIOTEC), Pathumthani, Thailand for the nuclear magnetic resonance

(NMR) analysis. In this study, such compound was determined to be 5,6-dihydroxy-

2,4-dimethoxy-9,10-dihydrophenanthrene by comparing its spectral data of proton


60

nuclear magnetic resonance (1H NMR) with those of previously isolated compound

(Itharat et al., 2014). The stock solution of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-

dihydrophenanthrene was prepared in DMSO at a concentration of 10,000 µM. In all

experiments, the final concentration of DMSO was kept below 0.5%.

3.4 Cell culture

Four cell lines used in this study were obtained from American Type

Culture Collection (ATCC, USA) e.g. 2 subtypes of human non-small cell lung cancer

(NSCLC); human lung carcinoma cell line A549 and human lung squamous carcinoma

cell line NCI-H226 as well as human small cell lung cancer cell line NCI-H1688 and

from European Collection of Cell Cultures (ECACC, UK) e.g. 1 subtype of NSCLC;

human large cell lung cancer line COR-L23. One normal cell line was obtained from

CLS-cell line service (CLS; Eppelheim, Germany) e.g. human lung fibroblast cell line

MRC-5.

The NSCLC such as A549, NCI-H226 and COR-L23 were cultured in

RPMI-1640 supplemented with 10% fetal bovine serum (FBS). The SCLC in the form

of NCI-H1688 was cultured in modified RPMI-1640 medium (2 mM L-glutamine, 10

mM HEPES, 1 mM sodium pyruvate, 4,500 mg/L glucose, 2,000 mg/L sodium

bicarbonate and 10% FBS. MRC-5 cell line was cultured in DMEM : Ham’s F12

medium supplemented with 10% FBS.

3.5 Growth inhibitory and cytotoxic effects

Growth inhibitory and cytotoxic effects of HMP were measured by

sulforhodamine B (SRB) assay. Its principle is the measurement of the cellular protein

content of living cells, based on the ability of SRB to bind to basic amino acid residues

that are fixed to the culture plate bottoms under mild acidic condition. The cell numbers

are estimated indirectly by staining total cellular protein content of each well with SRB

dye and the protein-bound dye is extracted from cells under mildly basic condition. The

optical density (O.D.) is measured using a microplate reader (Voigt, 2005).


61

In this experiment, the optimal cell number for seeding in a 96-well plate

was determined using SRB assay because it is critical to ensure exponential cell growth

for the entire duration of the assay. Briefly, The various numbers of cells were seeded

in a 2-fold serial dilutions of cells, ranging from 800-12,800 cells/well, in 100 µl

medium and incubated overnight at 37 ºC with 5% CO2 to allow cell attachment prior

to adding 100 µl of complete medium and further incubated for 0, 24, 48, 72 and 96 h.

At each of the indicated time points, the cells were fixed with 100 µl of 10%

trichloroacetic acid (TCA) at least 1 h to overnight at 4 ºC, washed with distilled water

and stained with 50 µl of 0.4% (w/v) sulforhodamine B (SRB) for 30 min in the dark.

The excess dye was removed by washing with 1% (v/v) acetic acid and then dried at

room temperature for 24 h in the dark. The protein-bound dye was extracted from cells

with 100 µl of 10 mM Tris, pH 10. The optical density was measured at 570 nm using

a microplate reader (Bio Tex, USA). The absorbance values obtained from microplate

reader were plotted versus incubation times, indicating the growth rate of each cell type.

According to the exponential cell growth for the entire assay period and O.D. 1.5-2.0

at the end of the 72-h assay time, the optimal cell numbers of each cancer cell line were

showed in Appendix A.

For measurement the antiproliferative and cytotoxic effects of HMP, 100

l of cells was seeded at the indicated numbers in the bracket (A549, NCI-H226 and

COR-L23 = 3.2 x 103 cells/well as well as NCI-H1688 and MRC-5 = 12.8 x 103

cells/well) in 96-well plates. On the following day, 100 l of HMP at final

concentrations of 3.125, 6.25, 12.5 and 25µM was added to the tested wells while 100

l of complete media was added to the control wells. These cells were further incubated

at 37°C with 5% CO2 for 72 h, fixed with 100 μl of 10% TCA, washed with distilled

water and stained with 50 µl of 0.4% SRB for 30 min in the dark. The excess dye was

removed by washing with 1% acetic acid and then dried at room temperature for 24 h

in the dark. The protein-bound dye was extracted from cells with 100 µl of 10 mM Tris,

pH 10. The optical density was measured at 492 nm using a microplate reader (Bio Tex,

USA). The percentages of cell survival for each concentration are calculated using the

following formula:


62

% cell survival = [(T - T0)/(C - T0)] × 100, T ≥ T0

or = [(T - T0)/ T0] × 100, T < T0

T = Average O.D. of cells treated with HMP for 72 h

T0 = Average O.D. at 0 h

C = Average O.D of cells treated with only media for 72 h

Based on the formula, the percentage of cell survival can be greater than zero, zero or

less than zero. A dose-response curve is obtained by plotting the percent cell survival

versus HMP concentrations. The concentrations of HMP required for 50% growth

inhibition (IC50), total growth inhibition (TGI), and 50% loss of cells (lethal

concentration, LC50) relative to the untreated cells are obtained by interpolating from a

dose-response cubic spline curve using GraphPad Prism 4.0 Software (GraphPad

Software, Inc., USA).

In addition, the selectivity index (SI), indicating the safety of HMP for

anticancer therapy was calculated by obtaining the ratio of IC50 of non-cancerous cell

lines and cancerous cell lines (Prayong, Barusrux, & Weerapreeyakul, 2008).

3.6 Cell proliferation by CFSE assay

Carboxyfluorescein diacetate, succinimidyl ester (CFDA-SE) is a cell-

tracking dye used to label cells for examining their proliferative activity. Briefly,

CFDA-SE is colorless and non-fluorescent that diffuses passively into cells. Within the

cells, intracellular esterase cleaves its acetate groups to yield fluorescent

carboxyfluorescein succinimidyl ester (CFSE). The succinimidyl ester group reacts

with intracellular amines, thus forming fluorescent conjugates that are well retained in

the cell (Wang, Duan, Liu, Fang, & Tan, 2005) (Figure 3.2). A profile of sequential

halving of CFSE fluorescence intensity with each generation can be monitored,

allowing the visualization of the number of rounds of cell division. Inhibition of cell

division by any substance can thus be traced through changes in the CFSE profile.

The proliferative activity of A549 cells treated with or without HMP was

determined by carboxyfluorescein-succinimidyl ester (CFSE) (BD Bioscience, USA).


63

According to the manufacturer’s instructions, A549 cells were labeled with 10 mM

CFSE in the dark at 37ºC with 5% CO2 for 20 min, washed two times using phosphate

buffered saline (PBS) with 10% FBS, resuspended with RPMI medium to plate at a

density of 3.2 x 104 cells/well in 24-well plates. Following overnight incubation, these

cells were incubated with 50µM HMP for 24, 48 and 72 h whereas control cells were

incubated with media only. For parent cells, non-treated cells were immediately

analyzed. These stained cells were detected using FACSCalibur flow cytometer

(Becton Dickinson, USA) and analyzed for numbers of cell division, proliferation

index, and precursor frequency with ModFit LT 3.2 program (Verity Software House,

USA). Proliferation index is the sum of the cells in all generations divided by the

number of original parent cells. Precursor frequency is defined as the fraction of the

parent population that proliferated in response to HMP treatment.

Figure 3.2 Formation of fluorescent compound CFSE by intracellular esterase.


64

3.7 Cell cycle analysis

The analysis of cell cycle distribution was performed by flow cytometry to

distinguish cells in different phases of the cell cycle. Before analysis, the cells were

permeabilised and treated with propidium iodide (PI), which is a fluorescent dye that

stains DNA quantitatively. In this analysis, the fluorescence intensity of stained cells at

a certain wavelength, therefore, correlates with the amount of DNA that the cells

contain. Four distinct phases can be recognized in a proliferating cell population. For

instance, cells in the G1 and S phases containing one copy of DNA, therefore, have 1x

fluorescence intensity whereas G2 and M phases with two copies of DNA have 2x

fluorescence intensity (Figure 3.3). In addition, apoptotic cells can be observed as a

hypodiploid or sub-G1 peak in DNA histogram.

In this study, A549 cells were plated at a density of 3.2 x 104 cells/well in

24-well plates and then incubated with 25 M and 50 µM HMP for 24, 48 and 72 h at

37°C with 5% CO2. Control cells were incubated with media only. After treatment,

these cells were collected by trypsinization, fixed gently (drop by drop) in 7 ml of 80%

ethanol, and then stored at -20ºC overnight. Then, cells were washed with PBS and

stained with 0.5 ml PI/RNase staining buffer (BD bioscience, USA) for 30 min at room

temperature in the dark. These stained cells were determined using a FACSCalibur flow

cytometer (Becton Dickinson, USA) and analyzed for cell cycle phases with ModFit

LT 3.2 Software (Verity Software House, USA). Moreover, the cell cycle distribution

was determined in HMP-treated cells after pretreatment for 6 h with 1.56 to 50 M of

the pan-caspase inhibitor Z-VAD-fmk.


65

Figure 3.3 DNA content distribution during the various phases of the cell cycle

obtained by flow cytometric analysis.

3.8 Annexin-V/PI double staining assay

The discrimination between intact and apoptotic cells was monitored by

annexin V-FITC (apoptotic cell marker) and propidium iodide (PI) (death cell marker)

double staining using flow cytometer. In live cells, phospholipid phosphatidylserine

(PS) is found in the inner membrane leaflet and translocates to the external membrane

leaflet in early apoptotic cells (Vermes, Haanen, Steffens-Nakken, & Reutelingsperger,

1995), these early apoptotic cells can be identified by the Annexin V-FITC staining,

which binds specifically to this externalized PS. Moreover, double staining with

propidium iodide (PI) differentiates early apoptotic cells with the intact membrane

(annexin V+/PI-) from late apoptotic/necrotic cells with leaky membranes (annexin

V+/PI+). Flow cytometric analysis was performed to quantify these cell populations.

The represented scatter dot plots demonstrate viable cells located in lower left quadrant

(annexin V-/PI-), early apoptotic cells in the lower right (annexin V+/PI-), late

apoptotic/necrotic cells, in the upper right (annexin V+/PI+), as shown in Figure 3.4.

In the present study, A549 cells were seeded and treated with HMP as

described in cell cycle analysis. After treatment, cells were trypsinized, washed with

cold PBS and then monitored by double staining with Annexin V-FITC Apoptosis


66

Detection Kit I (BD Biosciences, USA) and propidium iodide (PI) (Invitrogen, USA)

using flow cytometry. Briefly, the collected cells were resuspended in 100 µl of binding

buffer containing 5 µl of Annexin V-FITC and 5 µl of PI, incubated for 20 min at room

temperature in the dark and then determined using flow cytometer (Becton Dickinson,

USA) and analyzed with CellQuest Software.

Figure 3.4 Dot plot analysis by Annexin V-FITC/PI double staining.

3.9 Caspase-3 activity assay

The activity of caspase-3 was detected using the CaspACETM Assay

System, Colorimetric Kit (Promega, USA). The kit comprises a colorimetric substrate

N-acetyl-Asp-Glu-Val-Asp-p-nitroaniline (Ac-DEVD-pNA) and a cell-permeable pan-

caspase inhibitor N-benzyloxycarbonyl-Val-Ala-Asp-fluoromethylketone (Z-VAD-

fmk) for measuring caspase-3 activity. Its principle is that caspase-3 specifically

cleaves at the C-terminal side of the aspartate residue of the amino acid sequence

DEVD (Asp-Glu-Val-Asp), resulting that the chromophore p-nitroaniline (pNA) is

released from the substrate. Free pNA produces a yellow color that is monitored by a

spectrophotometer at 405 nm. The amount of yellow color produced upon cleavage is


67

proportional to the amount of caspase-3 activity present in the sample. However, the

finding that the inhibition of the increased caspase-3 activity by using pan-caspase

inhibitor Z-VAD-fmk, which irreversibly binds to and blocks the cleavage site of the

caspases could confirm such increased activity. In this assay, the comparison of the

absorbance of pNA from treatment sample, in the absence of inhibitor and in the

presence of inhibitor, with an un-induced control allows determination of the fold

inhibit and increase in caspase-3 activity, respectively.

A549 cells were incubated with or without 25 µM HMP for 24, 48 and 72

hrs at 37°C with 5% CO2. According to the manufacturer’s protocol, after treatment,

cells were collected, lysed in the cold lysis buffer by freeze-thaw procedure and

incubated on ice for 20 min. The cell lysates were centrifuged at 15,000 x g for 20 min

at 4ºC and the supernatant fraction was collected. The protein content in the supernatant

was then determined by Bradford’s method.

In a 96-well plate, cell extracts with an equal amount of 80 g of total

protein were added to each reaction containing caspase assay buffer and specific

colorimetric substrate (Ac-DEVD-pNA) for caspase-3, gently mixed, incubated at 37°C

for 4 h and measured at 405 nm by a microplate reader (Bio Tex, USA). Moreover,

caspase-3 activity was measured in cell extracts treated with 25 µM HMP after

pretreatment for 6 h with 50 μM Z-VAD-fmk.

3.10 Real-time Quantitative PCR Analysis

The Bcl-2 family proteins such as proapoptotic Bax and antiapoptotic Bcl-2

proteins have been reported to regulate mitochondrial outer membrane

permeabilization (MOMP), which triggers apoptotic pathways (Suen, Norris, & Youle,

2008). To determine relative mRNA expression levels of these proteins, real-time PCR

was performed using TagMan® Gene Expression Assay.

A549 cells were treated with or without 25 μM HMP for 24, 48 and 72 h at

37°C with 5% CO2. After treatment, RNA was extracted from these cells using total RNA

extraction kit (Real Biotech Corporation, Taiwan) according to the manufacturer’s

protocol. Briefly, cell pellets were lysed with 400 µl of RB buffer, mixed and incubated

at room temperature for 5 min. The sample mixture was placed in the Filter column and


68

centrifuged at 13,000 rpm for 2 min. The RNA-containing supernatant, which was passed

through this column, was collected and further added 400 µl of 70% ethanol prior to

transferring into the RB column. Such column was centrifuged at 13,000 rpm for 2 min.

The filtrate was removed whereas the column was kept. Next, this column, which

contains RNA binding to its glass fiber matrix was washed with 400 µl of R-W1 buffer

and 600 µl of R-Wash buffer, respectively, by centrifugation at 13,000 rpm for 1 min.

Such column was further centrifuged at 13,000 rpm for 3 min to dry the column matrix.

After that, RNA was eluted from this column by adding 35-50 µl of RNase-free water,

centrifugation at 13,000 for 1 min. The purity and quantification of RNA were measured

using a NanoDrop™ 2000 Spectrophotometer (Thermo Scientific, USA).

Two hundred and fifty nanograms of total RNA were converted to single-

stranded cDNA using High Capacity cDNA Reverse Transcription Kit (Applied

Biosystems, USA) according to the manufacturer’s protocol. Briefly, one reaction

requires 25 µl, as following: 2.5 µl of 10x RT buffer, 1 µl of 25x dNTP, 2.5 µl of 10x

Random primers, 1.25 µl of Reverse transcriptase, 5.25 µl of RNase-free water and 12.5

µl of RNA sample (20 ng/µl). The reaction mixture was performed on a GeneAmp PCR

system 2700 thermocycler (Applied Biosystems, USA) as follows: 25°C for 10 min,

37°C for 120 min, 85°C for 5 min and 4°C hold.

Thirty nanograms of cDNA were used for real-time PCR amplification to

determine the mRNA expression levels of target genes using an Applied Biosystems

(ABI) StepOne™ and StepOne Plus™ Real-Time PCR System (The Applied

Biosystems, USA). The quantitative PCR was performed in duplicate using EXPRESS

qPCR Supermix, Universal (Invitrogen, USA) and commercially available primer/probe

sets, which are pre-designed FAM™ dye- labeled TaqMan® MGB (minor groove

binder) probe and primer sets (inventoried Taqman® Gene Expression Assays)

(Thermo Scientific, USA) for human Bcl-2, Bax, and GAPDH. Briefly, The total

reaction volume was 20 µl, as following: 10 µl of 2x qPCR, 0.4 µl of Rox dye, 1 µl of

20x primer/probe set, 6 µl of RNase-free water and 3 µl of cDNA (10 ng/µl). The

thermal cycling parameters are one cycle of 50°C for 2 min, 95°C for 10 min and 40

amplification cycles of 95°C for 15 sec and 60°C for 1 min. The relative quantification

of gene expression was performed using the comparative threshold (CT) method and


69

determined by relative quantitation (RQ) value according to the 2-∆∆CT method. The

following equation was used to calculate RQ value.

∆CT treated group = CT of the target gene – CT of the endogenous gene (GAPDH)

∆∆CT = ∆CT of the treated group- ∆CT of the control group

Relative quantitation (RQ) value = 2 -∆∆CT

To compare fold changes in expression levels between the control group

and treated groups, CT values of all targets in the treated group were normalized to those

of GADPH as a house keeping gene.

3.11 Western blot analysis

Western blotting is an important technique used to identify specific proteins

from a mixture of proteins. The technique uses three elements to accomplish this task:

1) separation by size, 2) transfer to a solid support and 3) marking target protein using

a proper primary and secondary antibody to visualize (Mahmood & Yang, 2012).

A549 cells were treated with or without 25 μM HMP at different time

periods at 37°C with 5% CO2. After treatment, A549 cells were harvested and washed

with PBS. The cell pellets were lysed in 50-100 µl of RIPA buffer (Pierce, USA)

containing protease inhibitors cocktail (EMD Millipore, USA). The protein contents

were measured using BCA Protein Assay Kit (Pierce, USA). Forty micrograms of the

protein samples mixed with Laemmli sample buffer (Bio-Rad, USA) and 3 µl of

prestained molecular weight marker (Kaleidoscope; Bio-Rad, USA) were separated by

SDS-PAGE (7.5% or 12%) at 100 V of constant voltage until the dye front reached the

bottom of the gel. Following electrophoresis, the proteins were transferred onto

polyvinylidene fluoride (PVDF) membrane at 300 mA of constant current/slab for 3 h

(12 % gel) or 9 h (7.5% gel). After protein transfer, the protein bands on PVDF

membrane were rapidly stained using Ponceau S Staining Solution and washed with

Tris-buffered saline (TBS) until clean or colorless. Next, nonspecific sites on the

membrane were blocked using Odyssey blocking buffer (LI-COR Biosciences) in TBS

(1:1) for 1 h at room temperature. After blocking, the membrane was incubated with


70

specific primary antibody [anti-cdc25C (Santa Cruz Biotechnology,USA), anti-

cyclinB1 and anti-Bid (EMD Millipore, Germany), anti-caspase9 (Upstate

Biotechnologies, Charlottesville, VA), cdk1, Bax, Bcl-2, caspase3, β-actin (Cell

Signaling Technology, USA), poly(ADP-ribose) polymerase (PARP) (BD Biosciences,

USA)] at room temperature overnight, and washed with Tris-buffered saline with

Tween20 (TBST) for 15 min (3 times). Finally, the membrane was incubated with the

fluorescently-labeled secondary antibody (LI-COR Biosciences) for 1 h at room

temperature and then washed with TBST for 15 min (3 times) and washed with TBS

for 10 min. Detection of each protein was performed using Odyssey Infrared Imaging

System, Western Blot Analysis (LI-COR Biosciences, USA).

3.12 Tubulin polymerization assay

Tubulin polymerization assay is used to determine the disruption of

microtubule formation, either by inhibiting polymerization or by preventing

depolymerization of tubulin, resulting in the cell-cycle arrest and cell death. The effect

of HMP on tubulin polymerization was analyzed using in vitro Tubulin Polymerization

Assay Kit (≥ 99% pure bovine tubulin), Catalog No. 17-10194 (EMD Millipore, UK).

The method determines light that is scattered by microtubules to an extent that is

proportional to the concentration of microtubule polymer. The resulting polymerization

curve is a representative of the three phases of microtubule polymerization, namely

nucleation, elongation, and steady state phases.

According to the manufacturer’s protocol, polymerization reactions were

performed in 96-well plate half area, which is a UV transparent plate. The

polymerization reactions occur in 70 µl final volumes, which contain 60 µl of 60 µM

tubulin in 1x PB-GTP, 9 μL of 1x PB-GTP solution and 1 μL of the test substance,

including 1,750 µM HMP, 700 µM nocodazole, 700 µM paclitaxel and 1x PB-GTP

solution, which was used as a control. The polymerization of tubulin was monitored by

measuring the turbidity variation (light scattering) every 15 seconds at 350 nm during

60 minutes using a microplate reader (Bio Tex, USA).


71

3.13 DNA fragmentation assay

DNA fragmentation assay was used to determine apoptotic cell death in

A549 cells because apoptotic endonucleases, Caspase-Activated DNase; CAD, in these

cells can cleave chromosomal DNA at internucleosomal linker sites, resulting in the

formation of ladder pattern at about 180-base pair intervals in agarose gel

electrophoresis (Figure 3.5).

A549 cells were treated with or without 25 μM HMP for 48 h at 37°C with

5% CO2. After treatment, the formation of DNA fragments was detected using

Apoptotic DNA Ladder Detection Kit (ab66090) (Abcam, USA). According to the

manufacturer’s protocol, the cells were lysed with TE lysis buffer and extracted using

5 μl of Enzyme A solution incubated at 37°C for 10 min as well as 5 μl of Enzyme B

solution incubated at 50°C for 30 min. Next, 5 μl of ammonium acetate solution was

added to the reaction mixture and the DNA was then precipitated using 50 μl of

isopropanol for 30 min at -20°C. The DNA pellets were centrifuged at 10,000 rpm for

10 min, washed with 1 ml of 70% ethanol, centrifuged at 10,000 rpm for 10 min,

removed trace ethanol and air dry for 30 min at room temperature. The resulting pellets

were dissolved in 30 μl of DNA suspension buffer. The extracted DNA was separated

by electrophoresis on 1.5% agarose gel at 130 V of constant voltage. The resulting DNA

fragments were stained with ethidium bromide for 30 min, washed with distilled water

for 15 min and visualized by UV transilluminator and recorded by gel document (Alpha

Innotech, USA).


72

Figure 3.5 DNA fragmentation analysis. Internucleosomal DNA cleavage, a hallmark

of apoptosis, is demonstrated by a characteristic “laddering” pattern on gel

electrophoresis.

3.14 Nuclear staining with DAPI

For the assessment of apoptosis, the morphological changes of nuclei were

visualized following DNA staining by the fluorescent dye, 4’,6-diamidino-2-

phenylindole (DAPI) (Sigma, USA). This dye is a blue fluorescent nucleic acid stain

that preferentially stains double-stranded DNA (dsDNA).

In this study, A549 cells were seeded at a density of 3.2 x 104 cells/well

into 4-well cell culture slides (SPL Life Sciences, Korea). After being seeded on culture

slides overnight, cells were treated with or without 25 and 50 µM HMP for 24, 48 and

72h. The cells were washed with PBS and fixed with 80% ethanol for 30 min at room

temperature. After fixation, the cells were washed 3 times with PBS and then stained

with 1 µg/ml of DAPI in PBS for 45 min at room temperature in the dark. The cells


73

were washed 3 times with PBS, and stained nuclei were visualized using a fluorescence

microscope within 45 min (Nikon Eclipse Ci, Japan).

3.15 Intracellular reactive oxygen species (ROS) measurement

Intracellular reactive oxygen species (ROS) level was determined using

2´,7´-dichlorofluorescein diacetate (DCFH-DA). DCFH-DA is a non-fluorescent dye

that diffuses passively into cells. Inside cells, the cellular esterases cleave the acetate

esters to non-fluorescent 2´,7´-dichlorofluorescin (DCFH). In the presence of ROS,

DCFH is rapidly oxidized to a fluorescent molecule 2´,7´-dichlorofluorescein (DCF)

(Figure 3.6). The fluorescence intensity can be evaluated quantitatively using flow

cytometric analysis (Dikalov, Griendling, & Harrison, 2007).

In this experiment, A549 cells either untreated or treated with 25, 50 and

100 μM HMP for 3, 6, 12, 24 h were incubated with 20 µM DCFH-DA for 30 min at

37 C in 5% CO2, washed, and resuspended in phosphate-buffered saline (PBS). Before

being analyzed by FACSCalibur flow cytometer (Becton Dickinson, USA), the cells

will be shortly stained with PI for live/dead cell discrimination. The median

fluorescence intensity will be quantitated by CellQuest software (Becton-Dickinson,

USA) analysis of the recorded histograms. Moreover, the inhibition of ROS generation

was performed by pretreatment N-acetylcysteine (NAC), widely known as ROS

scavengers, for 1 h prior to 25 µM HMP treatment for 12 h.


74

Figure 3.6 Formation of fluorescent compound DCF by ROS and RNS.

3.16 Statistical analysis

Data are presented as mean ± SD for the indicated number of independent

experiments. Statistical differences between the control and treated groups were

analyzed using Independent-Samples T Test. Statistical differences between treatment

groups were analyzed by one-way analysis of variance (ANOVA) followed by post hoc

analysis. P value < 0.05 was considered statistically significant (SPSS 20 for

Windows).


75

CHAPTER 4

RESULTS AND DISCUSSION

4.1 Extraction of Dioscorea membranacea Pierre

The percent yield of the ethanolic extract of D. membranacea Pierre was

3.57 as shown in Table 4.1.

Table 4.1 The percent yield of the ethanolic extract of D. membranacea Pierre.

Dried plant material (g) Crude extract (g) % (w/w) Yield

204.80 7.31 3.57

4.2 Isolation and purification of HMP

The crude ethanolic extract of D. membranacea Pierre (7.31 g) was

subjected to silica gel column chromatography and eluted by gradient elution in order

of increasing polarity [hexane–chloroform (9:1) 1L, hexane–chloroform (6:4) 1L,

hexane–chloroform (2:8) 1L, chloroform 500 ml, chloroform–methanol (9:1) 500 ml,

chloroform–methanol (1:1) 500 ml and methanol 500 ml, respectively]. Four hundred

and forty-two fractions (10 ml each) were collected in a 15-ml tube, and the odd

numbered fractions were spotted on TLC aluminium sheets precoated with silica gel

60. Spots on the TLC sheets were then detected under UV light and visualized with

acidic anisaldehyde spray. The fractions containing similar spots were combined, and

14 combined fractions were obtained and then spotted on a TCL aluminium sheet along

with 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene, a previously isolated

compound (Figure 4.1).


76

Figure 4.1 TLC analysis of the 14 combined fractions of D. membranacea Pierre

extract obtained from the first silica gel column chromatography. (Lane 1: fractions 1-

87, lane 2: fractions 88-117, lane 3: fractions 118-143, lane 4: fractions 144-167, lane

5: fractions 168-185, lane 6: fractions 186-207, lane 7: fractions 208-217, lane 8:

fractions 218-223, lane 9: fractions 224-245, lane 10: fractions 246-279, lane 11:

fractions 280-291, lane 12: fractions 292-317, lane 13: fractions 318-331, lane 14: fractions

332-422, and lane 15: 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene)

As the combined fractions 5 to 7 contained 5, 6-dihydroxy-2, 4-dimethoxy-

9, 10-dihydrophenanthrene, these fractions were then pooled, evaporated to dryness and

weighed. Six hundred and forty milligrams was obtained from these pooled fractions,

dissolved in hexane–chloroform (1:9) and subjected to the second silica gel column

chromatography. The column was eluted by gradient elution in order of increasing

polarity [hexane–chloroform (1:9) 1.5 L, chloroform 500 ml, chloroform–methanol

(1:1) 500 ml and methanol 500 ml, respectively]. Two hundred and seventy-two

fractions (3 ml each) were collected in a 15-ml tube, and the odd numbered fractions

were spotted on TLC aluminum sheets. Spots on the TLC sheets were then detected

under UV light and visualized with acidic anisaldehyde spray along with 5,6-

dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene. In Figure 4.2, TLC analysis

showed three groups of the fractions containing 5,6-dihydroxy-2,4-dimethoxy-9,10-

dihydrophenanthrene based on the characteristic features of chromatographic bands.


77

Figure 4.2 TLC analysis of the odd numbered fractions, ranging from 25-99, eluted

from the second silica gel column chromatography.

The fractions in each group were combined and spotted on a TCL

aluminium sheet (group 1: fractions 38-52, group 2: fractions 53-61 and group 3:

fractions 62-83) along with 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene

(Figure 4.3).

Figure 4.3 TLC analysis of the 3 groups of the combined fractions eluted from the

second silica gel column chromatography. (lane 1: group 1, lane 2: group 2, lane 3: 5,

6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene and lane 4: group 3)


78

The group 3 (lane 4) showed two clearly separated spots, and the upper spot

has the same retention factor (Rf) as the spot of 5,6-dihydroxy-2,4-dimethoxy-9,10-

dihydrophenanthrene (lane 3). The combined fractions in this group were then

evaporated to dryness and weighed. Then, 40 milligrams was obtained and further

isolated on a TLC glass plate coated with silica gel 60 (Merck, Germany) using ethyl

acetate-hexane (1:1) as the mobile phase. The plate was visualized under UV light, and

two bands were detected and marked using a pencil. The upper band was scraped from

a TLC plate, eluted, evaporated to dryness and weighed (Figure 4.4). Twelve point six

milligrams of an isolated compound was obtained and called HMP-1. The percent yield

was 0.172 % (w/w) (Table 4.2).

Figure 4.4 The schematic flow chart for isolation of clearly separated bands using a

TLC glass plate.


79

Table 4.2 The percent yield of HMP-1 isolated from D. membranacea Pierre extract.

Crude extract (g) HMP-1 (mg) % (w/w) Yield

7.31 12.6 0.172

As the combined fractions in group 2 had several bands, all of which were

not well separated, the group 2 was spotted on a TLC aluminium sheet in different

solvent systems. Among these systems, ethyl acetate–hexane (4:6) is appropriate

because more clearly separated bands were observed (Figure 4.5).

Figure 4.5 TLC isolation of the combined fractions in group 2. (lane 1: one drop of

group 2, lane 2: 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene and lane 3:

two drops of group 2)


80

Next, the combined fractions in group 2 were evaporated to dryness and

weighed. Eighty milligrams was obtained, further dissolved in ethyl acetate–hexane

(4:6) and subjected to the third silica gel column chromatography. The column was

eluted by gradient elution in order of increasing polarity [ethyl acetate–hexane (4:6)

400 ml, ethyl acetate–hexane (6:4) 100 ml, ethyl acetate–hexane (8:2) 100 ml, ethyl

acetate 100 ml and ethyl acetate–methanol (2:8) 100 ml, respectively]. Fifty-one

fractions (3 ml each) were collected in a 15-ml tube, and the odd numbered fractions

were spotted on TLC aluminum sheets. Spots on the TLC sheet were then detected

under UV light and visualized with acidic anisaldehyde spray along with 5,6-

dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene. In Figure 4.6, fractions 15-23

containing 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene were pooled,

evaporated to dryness and weighed. Forty-eight point seven milligrams was spotted on

TLC glass plates coated with silica gel 60 and partitioned in chloroform–hexane (9:1)

twice. The plate was visualized under UV light, and two bands were detected and

marked using a pencil. The lower band that are the desired compound was scraped from

a TLC plate, eluted, evaporated to dryness and weighed (data not shown). Thirty-seven

point five milligrams of an isolated compound was obtained and called HMP-2. The

percent yield was 0.513 % (w/w) (Table 4.3).

Figure 4.6 TLC analysis of the odd numbered fractions, ranging from 1-51, eluted from

the third silica gel column chromatography.


81

Table 4.3 The percent yield of HMP-2 isolated from D. membranacea Pierre extract.

Crude extract (g) HMP isolation (mg) % (w/w) Yield

7.31 37.5 0.513

The purity of compounds can be tested by running in three different solvent

systems. A pure compound should appear as a single spot whereas an impure compound

has two or more spots in a single lane. In this study, the purity of HMP-1 and HMP-2

was checked by separating them on a TLC aluminium sheet in three different solvent

systems of varying polarity [ethyl acetate–hexane (4:6), chloroform–methanol (9.8:0.2)

and chloroform–hexane (8:2)]. The results revealed a single spot in the systems as

shown in Figure 4.7, indicating that HMP-1 and HMP-2 are pure compounds.

Figure 4.7 TLC analysis for checking the purity of HMP-2 in three different solvent

systems of varying polarity. (lane 1: HMP-2 and lane 2: 5, 6-dihydroxy-2, 4-dimethoxy-

9, 10-dihydrophenanthrene)


82

1H NMR spectrum of HMP-1 and HMP-2 was further analyzed as shown

in Figures 4.8 and 4.9, respectively. These spectra were compared with the spectrum of

previously isolated 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene (Figure

4.10), and showed the same pattern (Table 4.4). These data strongly support that HMP-

1 and HMP-2 were 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene.

In addition, the purity of HMP-1 and HMP-2 was assessed using the HPLC

technique. The chromatograms demonstrated that the retention time (RT) of a major

peak in HMP-1 and in HMP-2 was similar to that of the previously isolated compound,

indicating that the two isolated compounds were 5,6-dihydroxy-2,4-dimethoxy-9,10-

dihydrophenanthrene (Figure 4.11 and Appendix C). The purity of a compound is the

ratio of the area under the main peak to the total area under all peaks. The results showed

that the purity of HMP-1 and HMP-2 was greater than 94% (Table 4.5).

Figure 4.8 1H NMR spectrum of HMP-1 in deuterated chloroform (CDCl3).


83

Figure 4.9 1H NMR spectrum of HMP-2 in deuterated chloroform (CDCl3).


84

Figure 4.10 1H NMR spectrum of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydro

phenanthrene in deuterated chloroform (CDCl3).


85

Table 4.4 1H NMR spectral data (500 MHz) of HMP-1, HMP-2 and previously

isolated 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydrophenanthrene.

Position

1H (multi, J in Hz)

HMP-1

HMP-2

previously isolated 5, 6-

dihydroxy-2, 4-

dimethoxy-9, 10-

dihydrophenanthrene

(Itharat et al., 2014)

1

2

3

4

4a

4b

5

6

7

8

8a

9

10

10a

5-OH

6-OH

2-OMe

4-Ome

6.61 (d, 2.2)

6.56 (d, 2.4)

-

-

-

-

-

6.84 (d, 7.9)

6.77 (d, 7.9)

-

2.64 (m)

2.71 (m)

-

8.25 (s)

5.99 (s)

3.86 (s)

3.99 (s)

6.61 (d, 2.4)

6.56 (d, 2.4)

-

-

-

-

-

6.83 (d, 7.9)

6.76 (d, 8.2)

-

2.63 (m)

2.71(m)

-

8.25 (s)

5.99 (s)

3.85 (s)

3.98 (s)

6.64 (d, 2.5)

6.57 (d, 2.5)

-

-

-

-

-

6.84 (d, 8.0)

6.77 (d, 8.0)

-

2.64 (m)

2.70 (m)

-

8.26 (s)

6.02 (s)

3.87 (s)

3.98 (s)


86

Figure 4.11 HPLC chromatograms of HMP-1, HMP-2 and previously isolated 5,6-

dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene.

Sig = 254 nm

Sig = 270 nm

HMP-1

HMP-1

Sig = 254 nm

Sig = 270 nm

HMP-2

HMP-2

Sig = 254 nm

Sig = 270 nm

5, 6-dihydroxy-2, 4-dimethoxy-

9, 10-dihydrophenanthrene

5, 6-dihydroxy-2, 4-dimethoxy-

9, 10-dihydrophenanthrene


87

Table 4.5 The retention time, area under the curve and percentage area of HMP-1,

HMP-2 and previously isolated 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene

analyzed by HPLC at wavelengths 254 and 270 nm.

Compounds Wavelength

(nm)

Retention

time (RT)

(min)

Area under the

curve (AUC)

(mAU*s)

Percentage

area

(%)

HMP-1

254 23.462 13,689.4 95.36

270 23.462 30,924.9 97.77

HMP-2

254 22.722 15,039.3 94.07

270 22.722 32,605.3 96.55

previously

isolated 5, 6-

dihydroxy-2, 4-

dimethoxy-9,

10-dihydro

phenanthrene

254 23.492 6,810.1 98.08

270 23.492 16,256.3 99.21


88

4.3 Antiproliferative and cytotoxic effects of HMP against a panel of human lung

cancer cell lines

The antiproliferative effect of HMP was examined in four human lung

cancer cell lines i.e. A549, COR-L23, NCI-H226 and NCI-H1688 as well as a normal

cell line i.e. MRC-5 using SRB assay. The results demonstrated that HMP induced a

dose-dependent inhibition of growth in these cell lines (data not shown). As illustrated

in Table 4.6, HMP showed marked growth inhibitory effects on four different lung

cancer cell lines with the mean IC50 values ranging from 9.22 to 15.99 M, as compared

to a normal cell line i.e. MRC-5 (>100 M). Moreover, HMP was the most effective

against A549 cell line and also showed the highest selectivity index (SI > 10.85) toward

this cell line relative to MRC-5. Thus, we further investigated antiproliferative and

cytotoxic activities of HMP in A549 cell line through 3 parameters including IC50, TGI

and LC50. The results demonstrated that HMP exhibited growth inhibition (represented

as IC50 and TGI values) and cell death induction (represented as LC50 values), as shown

in Figure 4.12. Paclitaxel was also used as a positive control as previously described

(Hansakul et al., 2014).


89

Table 4.6 Antiproliferative effects of HMP on a panel of human cell lines.

Type of cell lines IC50 ± S.D. (μM) SI

A549 9.22 ± 1.26 * > 10.85

NCI-H226 10.38 ± 0.19 * > 9.63

COR-L23 15.99 ± 1.11 * > 6.25

NCI-H1688 15.78 ± 1.31 * > 6.34

MRC-5 > 100

The data are expressed as the mean ± SD (n 3). Each experiment was performed in

triplicate. (IC50 = 50% growth inhibition, SI = selective index)

* Statistical significance (P < 0.05) versus MRC-5 cells

Figure 4.12 Effects of HMP on antiproliferative and cytotoxic activities in A549 cells.


90

4.4 Inhibitory effects of HMP on cell division

The antiproliferative activity of HMP in A549 cells was further studied

using CFSE assay that monitors the number of cell divisions over time. With each round

of cell division, the relative CFSE fluorescence intensity decreases by half. Fifty µM

HMP was chosen in this study as nearly 100% growth inhibition was observed as

described in the previous results. Using ModFit LT 3.2 program, flow cytometric

analysis of CFSE-labelled cells showed that HMP-treated cells for 24, 48 and 72 h were

mainly arrested in the second round as early as 24 h (Generation 2 depicted as an orange

color, Figure 4.13A). In contrast, untreated cells proceeded through many cycles of the

division with increased incubation times. Besides the number of cell divisions, the

software provides proliferation index and precursor frequency. As for the proliferation

index, which indicates the fold-expansion of the overall culture (Roederer, 2011), the

index values of HMP-treated cells for 24, 48 and 72 h slightly increased, ranging from

1.5 to 3. On the contrary, these of untreated cells (control) considerably elevated and

reached the maximum value of approximate 16 at 72 h (Figure 4.13B). For the precursor

frequency, which defines the fraction of the parent population that divided in response

to HMP (Roederer, 2011), these values of treated cells increased slightly over

incubation periods, thus indicating that the majority of parent cells did not divide.

Conversely, untreated parent cells divided vigorously, resulting in substantial increases

in the precursor frequency over time (Figure 4.13C). Altogether, these data indicated

that HMP was effective in inhibiting cell division.


91

Figure 4.13 Antiproliferative effects of HMP on A549 cells. (A) Representative

profiles of the sequential halving of CFSE fluorescence intensity of the parent cells at

0 h incubation (blue peak), untreated and 50 µM HMP-treated cells at 24, 48 and 72 h.

(B-C) Bar charts representing the proliferation index and precursor frequency. The data

are expressed as the mean ± SD (n 3). * P < 0.05 versus control at equal incubation

times.

B C


92

4.5 Effects of HMP on the cell cycle distribution

As HMP was effective in inhibiting cell division, we next determined the

effect of HMP on cell cycle distribution by performing cell cycle analysis. Flow

cytometry was used to analyze DNA contents in cell cycles of A549 cells, which were

treated with various concentrations of HMP e.g. 100 µM (nearly the LC50 value), 50

µM (nearly the TGI value) and 25 µM (the half TGI value) for 24, 48 and 72 h. As

illustrated in Figure 4.14 and Table 4.7, flow cytometric analysis of the DNA from

HMP-treated cells displayed a substantial increase in the percentage of cells in G2/M

phase at 24, 48 and 72 h as compared to the control at equal incubation periods.

However, the percentage of cells in G2/M phase was inversely proportional to

incubation time. As these G2/M phase cells decreased between 48-h and 72-h incubation

periods, the marked increase in the sub-G1 peak representing apoptotic cells was also

observed in a time-dependent manner. For each incubation period, the data also

revealed that cells treated with increasing concentrations of HMP caused the decreased

percentages of cells in G2/M phase, along with the dramatically increased percentages

in G1 phase. Such data indicated that exposure to the high concentrations of HMP

resulted more DNA degradation than that the low concentrations did, leading to the

reduction in fluorescence intensity of nuclei from G2/M to G1 phase in a dose-dependent

manner. All concentrations used in this study could markedly block the cell cycle at the

G2/M phase and subsequently induced apoptosis in A549 cells. Therefore, only 25 μM

HMP was selected for further studies on molecular aspects.


93

Figure 4.14 Effects of HMP on cell cycle distribution in A549 cells. Cell cycle analysis

of untreated and treated cells with 25, 50 and 100 µM for 24, 48 and 72 h was performed

using ModFit LT 3.2 program. Percentages of cells in G0/G1, S, G2/M and sub-G1

phases are represented as the mean of three independent experiments.

* P < 0.05 versus control at equal incubation periods.


94

Table 4.7 The percentages of HMP-treated cells in each phase of cell cycle.

Time Samples

% non-apoptosis cells % apoptosis

cells

G1 S G2/M Sub-G1

24 h

Control 66.66 ± 3.09 10.07 ± 0.55 22.44 ± 1.83 1.25 ± 0.38

25 µM HMP 9.11 ± 0.94* 2.12 ± 0.16* 79.44 ± 4.24* 7.01 ± 1.94*

50 µM HMP 13.55 ± 0.45* 2.26 ± 0.31* 75.74 ± 0.39* 8.56 ± 0.18*

100 µM HMP 36.53 ± 2.22* 5.66 ± 1.09* 48.94 ± 1.59* 9.07 ± 2.87*

48 h

Control 71.67 ± 3.81 10.40 ± 1.58 20.50 ± 2.54 0.89 ± 0.14

25 µM HMP 12.04 ± 1.52* 7.16 ± 1.91* 55.60 ± 2.40* 26.51 ± 2.33*

50 µM HMP 15.76 ± 1.28* 7.07 ± 0.78* 48.29 ± 4.71* 29.31 ± 2.96*

100 µM HMP 33.78 ± 0.90* 8.12 ± 0.16 35.19 ± 0.46* 23.36 ± 1.23*

72 h

Control 74.99 ± 2.07 7.96 ± 1.05 15.26 ± 1.69 1.52 ± 0.56

25 µM HMP 13.30 ± 1.41* 10.27 ± 2.87 43.65 ± 4.07* 30.50 ± 3.55*

50 µM HMP 15.61 ± 1.30* 9.03 ± 1.80 44.49 ± 2.81* 32.90 ± 6.92*

100 µM HMP 33.95 ± 2.67* 9.27 ± 0.44 33.13 ± 3.79* 24.13 ± 4.25*

The data are expressed as the mean ± SD (n 3).

* Statistical significance (P <0.05) versus control at equal incubation periods.


95

4.6 Effect of HMP on protein expression of cell cycle regulatory proteins

As HMP arrested the cells in the G2/M phase, we next examined whether it

manifested cell cycle distribution and subsequent inhibition of cell growth through

G2/M regulatory proteins. Therefore, the effect of HMP on the expression of the G2/M

regulatory proteins cdc25C, cdk1 and cyclin B1 was monitored by Western blot

analysis. A549 cells were either untreated or treated with 25 µM HMP for 12, 24, 48

and 72 h. As shown in Figure 4.15, the results demonstrated the basal expression levels

of constitutive cdc25C, cdk1 and cyclin B1 in untreated cells. Also, HMP treatment

induced down-regulation of cdc25C and cdk1 protein levels in a time-dependent

fashion. Interestingly, elevated levels of cyclin B1 protein were detected in A549 cells

after 12 and 24 h of HMP treatment and markedly decreased at 48 and 72 h. β-actin

levels served as an internal control and were unaffected under these conditions. These

results revealed that HMP decreased the expression levels of cdc25C, cdk1 and cyclin

B1, all of which are involved in G2 to M phase progression, thereby leading to the G2/M

arrest.

Figure 4.15 Effects of HMP on protein levels of cdc25C, cdk1 and cyclin B1 in A549

cells. The cell cycle regulatory proteins of these cells either untreated or treated with

25 µM HMP for 12, 24, 48 and 72 h were analyzed by Western blotting. Data are

representatives of three independent experiments.


96

4.7 Effect of HMP on interfering microtubule formation

In addition to the alteration of the G2/M regulatory proteins, we further

investigated whether HMP could inhibit microtubule assembly in vitro using tubulin

polymerization assay kit in a cell-free system. The effect of 25 μM HMP on tubulin

polymerization was monitored every 15 seconds at 350 nm using a microplate reader,

as shown in Figure 4.16. For comparison, parallel experiments were conducted with

paclitaxel (a microtubule stabilizer), nocodazole (a microtubule depolymerizer), and

untreated tubulin as a control. The results demonstrate that 25 μM HMP, which was

shown to markedly block cell cycle at G2/M phase in A549 cells, showed very similar

tubulin polymerization to the control, indicating that HMP did not disrupt in vitro

polymerization of tubulin into microtubules.

Figure 4.16 Effect of HMP on in vitro tubulin polymerization. Sixty µM tubulin

concentration was incubated with polymerization buffer in a 96-well half area plate at

37°C in the absence (control) and the presence of 25 µM HMP, 10 µM paclitaxel and

10 µM nocodazole. Data are representatives of four independent experiments.


97

4.8 Effect of HMP on apoptosis induction in A549 cells

As cell cycle analysis revealed the “sub-G1 peak” that represents apoptotic

cells in HMP-treated cells, annexin V/PI staining and flow cytometry were performed

to quantify early apoptotic cells. In these cells, phosphatidylserine (PS) is translocated

from the inner to the outer membrane leaflet and specifically binds to annexinV.

Moreover, double staining with PI differentiates early apoptotic cells with intact

membranes from late apoptotic/necrotic cells with leaky membranes. The represented

scatter dot plots demonstrate viable cells located in lower left quadrant (annexin V–/PI–),

early apoptotic cells in the lower right (annexin V+/PI–), late apoptotic/necrotic cells,

in the upper right (annexin V+/PI+).

In the present study, A549 cells were treated with or without various

concentrations of HMP for 24, 48 and 72 h. The results indicated that HMP induced a

decrease in the percentage of viable cells with a concomitant increase in the percentage

of early and late apoptotic cells in a time- and dose-dependent manner (Figure 4.17 and

Table 4.8). Therefore, such data strongly suggest a critical role of HMP in stimulating

apoptosis in A549 cells.


98

Figure 4.17 Effects of HMP on apoptotic induction in A549 cells. Apoptotic profiles

of untreated and treated cells with 25, 50 and 100 µM HMP for 24, 48 and 72 h were

performed using CellQuest Software. The percentages of cells in the respective

quadrants i.e. LL: Viable cells, LR: Early apoptotic cells, UR: Late apoptotic cells, UL:

Dead cells are indicated as the mean of three independent experiments. * P < 0.05

versus control at equal incubation periods.


99

Table 4.8 The percentages of cells in the respective quadrants. (i.e. LL: viable cells,

LR: early apoptotic cells, UR: late apoptotic cells, UL: dead cells)

Time samples

LL LR UR UL

% live cells

% early

apoptotic

cells

% late

apoptotic

cells

% dead

cells

24 h

Control 92.74 ± 2.17 3.13 ± 0.71 3.73 ± 1.39 0.41 ± 0.32

25 µM HMP 83.01 ± 4.08* 14.09 ± 2.15* 3.66 ± 1.01 0.81 ± 0.46

50 µM HMP 77.17 ± 5.19* 13.25 ± 2.02* 6.21 ± 2.44 1.44 ± 0.88

100 µM HMP 81.08 ± 1.36* 14.25 ± 1.68* 6.23 ± 2.67 1.21 ± 0.98

48 h

Control 91.76 ± 1.68 3.64 ± 1.23 4.06 ± 1.13 0.55 ± 0.18

25 µM HMP 74.19 ± 9.10* 26.26 ± 8.00* 5.54 ± 1.73 1.02 ± 0.59

50 µM HMP 60.83 ± 4.93* 28.37 ± 7.81* 9.21 ± 0.65* 1.85 ± 0.82

100 µM HMP 56.48 ± 7.50* 24.81 ± 2.32* 27.12 ± 4.62* 2.35 ± 0.53*

72 h

Control 93.22 ± 0.86 2.21 ± 0.82 3.29 ± 0.97 1.27 ± 0.47

25 µM HMP 65.47 ± 9.59* 25.24 ± 4.89* 10.41 ± 0.91* 3.41 ± 1.44*

50 µM HMP 54.51 ± 5.93* 25.12 ± 5.76* 15.95 ± 2.51* 3.97 ± 1.06*

100 µM HMP 39.74 ± 8.16* 21.05 ± 4.95* 48.36 ± 6.36* 4.65 ± 2.00*

The data are expressed as the mean ± SD (n 3). * Statistical significance (P <0.05)

versus control at equal incubation periods.


100

4.9 Effect of HMP on caspase-3 activity in A549 cells

Next, the involvement of caspase activation was examined because caspase

enzymes, a family of cysteine proteases, are central components of the machinery for

apoptosis (Earnshaw, Martins, & Kaufmann, 1999). A549 cells were pretreated with

the pan-caspase inhibitor, Z-VAD-fmk, at different concentrations (1.56, 6.25, 25 and

50 μM) for 6 h prior to 25 µM HMP treatment for an additional 72 h and the percentage

of cells in each phase of the cell cycle was shown in Figure 4.18A and Table 4.9. In this

study, Z-VAD-fmk-pretreated cells displayed a significant decrease in sub-G1

populations as compared with unpretreated cells. Moreover, the increased percentage

of apoptotic cells with the induction of HMP treatment was markedly inhibited by Z-

VAD-fmk in a concentration-dependent manner (Figure 4.18B). These results indicated

that HMP-induced apoptosis in A549 cells was dependent on the activation of caspase

enzymes.

As caspase-3 is the primary effector (executioner) caspase responsible for

much of the cellular degradation during apoptosis, we then evaluated changes of its

activity in 25 µM HMP-treated cells for 24, 48 and 72 h using the CaspACE™ Assay

System (Promega, USA). Following HMP treatment, the relative activity of caspase-3

was not significantly different from the control at 24 h, but was significantly increased

at 48 and 72 h. Its highest level was detected at 48 h (Figure 4.19A). In addition,

pretreatment with 50 µM Z-VAD-fmk 6 h prior to HMP treatment for 48 h completely

suppressed the highest relative activity of caspase-3 (Figure 4.19B), thus ascertaining

its involvement in apoptosis.


101

Figure 4.18 Inhibitory effects of Z-VAD-fmk on sub-G1 populations. (A) Representative

profiles of flow cytometric analysis. (B) Bar graphs representing the percentage of sub-

G1 peaks with the percent inhibition presented below. The data are expressed as the

mean ± SD (n 3). * P < 0.05 versus HMP-treated cells.

A

B


102

Table 4.9 The percentages of cells in each phase of cell cycle. The cells were pretreated

with Z-VAD-fmk at different concentrations for 6 h prior to 25 µM HMP treatment for

72 h.

Time Samples % non-apoptosis cells

%apoptosis

cells

G1 S G2/M Sub-G1

72 h

Control 67.98 ± 1.19 23.04 ± 1.63 8.98 ± 0.98 0.13 ± 0.17

25 µM HMP 3.25 ± 1.33 37.12 ± 4.48 59.63 ± 5.37 18.46 ± 3.75

25 µM HMP 3.91 ± 0.01 20.65 ± 1.62 75.45 ± 1.61 14.06 ± 3.21

1.56 µM Z-VAD fmk

25 µM HMP 3.53 ± 0.33 22.78 ± 0.74 73.70 ± 1.07 13.11 ± 2.98

6.25 µM Z-VAD fmk

25 µM HMP 3.83 ± 0.65 13.01 ± 2.38 83.16 ± 1.73 9.47 ± 2.62

25 µM Z-VAD fmk

25 µM HMP 3.71 ± 1.02 11.97 ± 3.43 84.33 ± 2.64 8.21 ± 1.20

50 µM Z-VAD fmk



103

Figure 4.19 Effects of HMP on caspase-3 activity in A549 cells. (A) The relative

caspase-3 activity of 25 µM HMP-treated cells for 24, 48 and 72 h. * (P <0.05) versus

untreated cells at equal incubation periods using Independent-samples t-test. For the

treated cell groups, bars with different lowercase letters are significantly different at p

<0.05 using One-way ANOVA. (B) The inhibition of the relative caspase-3 activity of

cells pre-incubated with 50 µM Z-VAD-fmk for 6 h prior to HMP treatment for 48 h as

compared to cells treated with HMP alone. * P < 0.05 versus HMP alone.

A

B


104

4.10 Bax and Bcl-2 mRNA and protein expression levels

The significant cell cycle arrest followed by apoptosis in HMP treatment

provide a powerful hint that the intrinsic mitochondrial pathway was likely activated.

The pathway is initiated by non-receptor-mediated intracellular signals, leading to an

increase in pro-apoptotic proteins e.g. Bax relative to antiapoptotic proteins e.g. Bcl-2

and subsequent mitochondrial outer membrane permeabilization (MOMP). This event

is regulated by the balance between the Bcl-2 family proteins that shift for pro-apoptotic

proteins (Bender and Martinou, 2013). Therefore, mRNA and protein levels of pro-

apoptotic Bax and antiapoptotic Bcl-2 were determined in A549 cells treated with or

without HMP.

The results showed that treatment with HMP significantly increased Bax

mRNA (Figure 4.20) and protein levels (Figure 4.21) relative to control in almost all

incubation periods. Although significant increases relative to control were also seen in

Bcl-2 expression, Bcl-2 mRNA levels were significantly lower than those of Bax at

48 h and 72 h (Figure 4.20), and Bcl-2 protein level was notably lower than that of Bax

at 72 h (Figure 4.21). Due to higher levels of Bax relative to Bcl-2, it appeared that

apoptotic activity of Bax may be more involved in HMP-stimulated intrinsic apoptosis

than the antiapoptotic activity of Bcl-2. Moreover, the results showed more than 2-fold

increases in Bax versus Bcl-2 mRNA levels, but not in their protein levels. This made

us speculate whether high levels of Bax protein may be more affected by rapid

degradation due to cell death. Altogether, these data indicated that HMP activated

apoptosis through the intrinsic mitochondrial pathway by increasing Bax expression

over Bcl-2.


105

Figure 4.20 The quantification of relative mRNA levels of Bax and Bcl-2 in A549 cells

using Real-time PCR. The cells were treated with or without 25 µM HMP for 24, 48

and 72 h. Data are expressed as mean ± SD (n = 3). * P < 0.05 versus control at equal

incubation times. # P < 0.05 versus Bax at equal incubation times.


106

Figure 4.21 Effects of HMP on protein expression of Bax and Bcl-2 in A549 cells. The

cells were treated with or without 25 µM HMP for 24, 48 and 72 h. (A) Protein levels

of Bax and Bcl-2 using Western blotting. (B) Bar graphs representing the relative band

intensities of Bax and Bcl-2. Data are representatives of three independent experiments.

β-actin was used as a loading control. * P < 0.05 versus control at equal incubation

times. # P < 0.05 versus Bax at equal incubation times.

A

B


107

4.11 Effect of HMP on expression of active caspases and their targets

As the previous experiment showed the highest caspase-3 activity at 48 h,

we further investigated the expression of the active form of caspase-3 and its well-

established substrate, PARP, using western blotting analysis at the same time. The

results showed that HMP induced the cleavage of procaspase-3 (35 kDa) into the active

form (19 and 17 kDa), as shown in Figure 4.22A. Similarly, the reduction of pro PARP

(113 kDa) was accompanied by the increase of cleaved PARP (89 kDa), as shown in

Figure 4.22B, indicating the action of cleaved caspase-3. These results confirmed that

HMP indeed induced apoptosis via the caspase-dependent apoptotic pathway.

Since HMP activated apoptosis through the intrinsic mitochondrial

pathway, leading to mitochondrial outer membrane permeabilization (MOMP), we

further studied the expression of cleaved caspase-9 as the crucial initiator caspase of

this pathway. Our results showed that HMP treatment for 48 h reduced expression of

procaspase-9, and its cleaved form was generated (89 kDa), as compared to the control

(Figure 4.22C). These data further confirmed that HMP induced apoptosis through

activation of the intrinsic pathway. However, caspase-8, the predominant initiator

caspase in the extrinsic pathway, also induces MOMP through the cleavage of cytosolic

Bid to truncated Bid (tBid). The tBid then translocates to mitochondria, leading to

MOMP, thus connecting the extrinsic pathway to the intrinsic mitochondrial pathway.

For these reasons, we then investigated whether HMP induced apoptosis via the

extrinsic pathway by testing the levels of Bid protein. The results showed that Bid in

HMP-treated cells was significantly lower than that in the control, indicating that HMP

triggered apoptosis via the extrinsic pathway.


108

Figure 4.22 Effects of HMP on expressions of apoptotic proteins in A549 cells. Cells

were treated with 25 µM HMP for 48 h. Control cells were treated with media only.

The cellular proteins of these cells were analyzed by Western blot. Levels of caspase-3

and cleaved caspase-3 (A), PARP and cleaved PARP (B) as well as caspase-9 and Bid

(C) are shown. β-actin was used as a loading control. The immunoblots shown are


4.12 Effect of HMP on nuclear morphological changes

To determine the nuclear morphological changes during apoptosis, DAPI

staining was performed. As shown in Figure 4.23 and 4.24, HMP-treated cells exhibited

chromatin condensation and nuclear fragmentation as compared to control. Such

characteristic features can be seen in apoptotic cells. It is noteworthy that the chromatin

condensation and nuclear fragmentation decreased with prolonged incubation periods

because of the possibility of losing floating cells (dead cell) in wash steps.

A B C


109

Figure 4.23 Effects of HMP on nuclear morphological changes by DAPI staining under

Bright-field microscopy (400x magnification); chromatin condensation (red arrows),

chromatin fragmentation (green arrows). Data are representatives of three independent

experiments.

Control 25 µM HMP

24 h

48 h

72 h


110

Figure 4.24 Effects of HMP on nuclear morphological changes by DAPI staining under

Fluorescent microscopy (400x magnification); chromatin condensation (red arrows),

chromatin fragmentation (green arrows). Data are representatives of three independent

experiments.

Control 25 µM HMP

24 h

48 h

72 h


111

4.13 Effect of HMP on DNA fragmentation

To confirm HMP-induced apoptosis, we also investigated DNA

fragmentation, which is one of the biochemical hallmarks of apoptosis. The DNA

fragmentation is associated with apoptotic endonucleases, which cleave chromosomal

DNA at internucleosomal linker sites, thereby resulting in the formation of ladder

pattern at about 180-base pair intervals in agarose gel electrophoresis. In the present

study, genomic DNA of A549 cells treated either with or without 25 µM HMP for 48

h was extracted using Apoptotic DNA Ladder Detection Kit and analyzed by 1.5%

agarose gel electrophoresis. As illustrated in Figure 4.25, the DNA fragmentation was

clearly observed in only HMP-treated cells, indicating that HMP indeed induced

apoptosis in A549 cells.

Figure 4.25 Effect of HMP on DNA fragmentation of A549 cells. DNA fragments of

cells treated with 25 µM HMP for 48 h were analyzed by 1.5% agarose gel

electrophoresis. Data are representatives of three independent experiments.


112

4.14 Effect of HMP on the generation of intracellular ROS

Many studies have shown that natural plant compounds can produce high

levels of ROS in cancer cells, leading to apoptosis (Gong & Li, 2011; Jo et al., 2012;

Singh et al., 2012). Therefore, we further investigated whether HMP-induced apoptosis

was associated with the increase of ROS generation in A549 cells. The cells treated

either with or without 25 µM HMP at different incubation times were performed using

DCF assay and analyzed by flow cytometry. The results revealed that HMP could

generate ROS in treated cells. As illustrated in Figure 4.26, the ROS accumulation

reached the highest level at 12 h and then slowly decreased after 24 h. To confirm ROS

generation, cells were pretreated with the ROS scavenger NAC at 0.1, 1 and 5 mM for

1 h prior to the addition of 25 µM HMP and further incubation for 12 h. The results

showed that NAC completely inhibited the highest level of ROS in a dose-dependent

manner (Figure 4.27). The results showed that NAC completely inhibited the highest

level of ROS in a dose-dependent manner. This results indicated that HMP induced the

increased ROS generation.

We next investigated whether ROS generation induced by HMP was

directly associated with apoptotic cell death. The changes in sub-G1 populations were

thus determined in the cells, which were pretreated with NAC at different

concentrations (0.1, 1 and 5 mM) for 1 h prior to 25 µM HMP treatment for 24 h. As

shown in Figure 4.28A, 4.28B and Table 4.10, there were no statistically significant

changes in sub-G1 populations of cells that were pretreated with or without NAC. These

data indicate that the increased ROS levels by HMP did not influence cell apoptosis.

We also demonstrated that NAC alone did not induce apoptotic cell death. Flow

cytometric analysis of the DNA from cells treated with NAC alone at different

concentrations for 72 h displayed very similar cell cycle distribution to the control

(Appendix D).


113

Figure 4.26 Effect of HMP on ROS production in A549 cells treated with 25 µM HMP

at different incubation times. Data are representatives of three independent

experiments.

Control

25 µM HMP 3 h 6 h 12 h

24 h 48 h 72 h


114

Figure 4.27 Effect of the ROS scavenger NAC on ROS production in A549 cells.

(Upper row) Cells were pretreated with NAC at 0.1, 1 and 5 mM for 1 h before the

addition of 25 µM HMP for 12 h relative to 25 µM HMP-treated cells. (Lower row)

Cells were treated with only NAC at different concentrations for 13 h. Data are


Control Control Control Control

25 µM HMP 25 µM HMP 25 µM HMP 25 µM HMP

0.1 mM NAC 1 mM NAC 5 mM NAC + + +

Control Control Control 0.1 mM NAC 1 mM NAC 5 mM NAC


115

Figure 4.28 Inhibitory effects of NAC on sub-G1 populations. A549 cells were

pretreated with NAC at different concentration (0.1, 1 and 5 mM) for 1 h prior to 25

µM HMP treatment for 24 h. (A) Representative profiles of flow cytometric analysis.

(B) Bar graphs representing the percentage of sub-G1 peaks with the percent inhibition

presented below. Data are expressed as the mean ± SD (n ≥ 3).

A

B


116

Table 4.10 The percentages of cells in each phase of cell cycle. The cells were

pretreated with NAC at different concentrations for 6 h prior to 25 µM HMP

treatment for 24 h.

Time Samples % non-apoptosis cells

%apoptosis

cells

G1 S G2/M Sub-G1

24 h

Control 58.80 ± 1.06 30.23 ± 0.85 10.97 ± 1.57 0.51 ± 0.44

25 µM HMP 10.04 ± 3.98 7.63 ± 2.85 82.33 ± 1.13 6.01 ± 2.33

25 µM HMP 10.74 ± 4.06 8.47 ± 2.23 80.79 ± 4.37 6.07 ± 1.93

0.1 mM NAC

25 µM HMP 9.87 ± 3.60 8.38 ± 2.75 81.75 ± 1.98 6.89 ± 1.08

1 mM NAC

25 µM HMP 10.49 ± 3.59 6.77 ± 3.77 82.74 ± 1.52 7.39 ± 1.33

5 mM NAC



117

CHAPTER 5

CONCLUSIONS AND RECOMMENDATIONS

The search for plant-derived compounds has been considered to be an

interesting subject in generating new anticancer agents with high safety and high

efficacy. Exploring the precise molecular mechanisms involved in their actions has

become an important approach for preclinical evaluation of anticancer agents and

subsequent development of anticancer drugs. For this reason, the screening of

anticancer agents through induction of cell cycle arrest and apoptosis appears to be a

powerful strategy for discovery of potent anticancer agents (Li et al., 2016). An active

compound called 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene (HMP)

derived from the rhizome ethanolic extract of D. membranacea Pierre had the highest

selectivity index against human lung large cell carcinoma COR-L23 cells using the

SRB assay (Itharat et al., 2014). In this study, effects of HMP on growth inhibition were

further tested in a panel of human lung cancer cell lines representing NSCLC and SCLC

compared to the well-defined human MRC-5 fibroblast line. The effects of HMP on

growth inhibition and cell death through induction of cell cycle arrest and apoptosis

was comprehensively investigated in the most responsive cell line, A549 human lung

carcinoma cell line.

5.1 Antiproliferative effect of HMP in A549 cells

Antiproliferative effect of the active compounds is investigated by IC50

values. The lower the IC50 values, the more potent a compound is. According to

National Cancer Institute (NCI) plant screening program, pure compounds with IC50

values of 4 μg/ml or less are considered to confer significant in vitro cytotoxic activity

(Phang, Malek, & Ibrahim, 2013). Also, SI values are used to define the specificity of

the compound for cancer cells. The higher the SI values, the more selective in killing

cancer cells a compound is, as opposed to normal cells. Furthermore, SI values greater

than 3.0 are considered to be significant (Mahavorasirikul, Viyanant, Chaijaroenkul,

Itharat, & Na-Bangchang, 2010). In this study, HMP exerted strong and selective


118

antiproliferative activity against the A549 cell line among other lung cancer cell types

because it had an IC50 value of 9.22 µM (or 2.51 µg/ml) for this cell line. This value

was below 4 μg/ml, the cut-off value in which any compound with this value or less is

considered to be potent. HMP also had the highest SI value of > 10.85 for the A549 cell

line versus the MRC-5 cell line. These data indicated the potential use of HMP as a

promising anticancer agent. Thus, we chose the A549 cell line to investigate the

molecular mechanism underlying antiproliferative and cytotoxic activities of HMP.

5.2 Molecular mechanism underlying antiproliferative effect of HMP

This study showed that HMP possessed strong antiproliferative activity

against A549 cells by inhibiting cell cycle arrest at G2/M phase. In this G2/M phase

transition, specific regulatory proteins, such as cdc25C, cdk1, and cyclin B1 play an

important role as follows; the cdc25C removes the inhibitory phosphates present on

cdk1, thus rendering cdk1-cyclin B1 complex active and ultimately accelerating the

transition from G2 into mitosis (M) phase (Sanchez, McElroy,& Spector, 2003;

Potapova, Daum, Byrd, & Gorbsky, 2009). Our results demonstrated that HMP

treatment downregulated the expression levels of these proteins. Such decreased levels

of these proteins could affect their activity. Indeed, decreased activity of cdc25C due to

its decreased protein levels leads to unremoved inhibitory phosphorylation of cdk1,

causing accumulation of an inactive cdk1-cyclin B1 complex (Singh et al., 2004). The

decreased proteins are possibly caused by either their enhanced degradation through

ubiquitin/proteasome pathway or their suppressed mRNA synthesis (Shabbeer et al.,

2013). The first postulate could be proved by the restoration of these decreased

regulatory proteins after pretreatment with proteasome inhibitors. The latter postulate

could be determined by detection of decreased mRNA levels of the corresponding

proteins. However, the reduction of cdc25C, cdk1, and cyclin B1 protein levels does

not exclusively cause G2/M arrest. For example, Singh et al. (2004) have shown that

sulforaphane (SFN)-induced decline in cdc25 protein levels was nearly fully blocked

in the presence of proteasome inhibitor lactacystin. Such cell cycle arrest, however,

turned out not to be significantly affected upon such restoration of cdc25C protein


119

levels using this inhibitor. Cytoplasmic translocation of cdc25C appeared to be the main

mechanism of SFN-induced cell cycle arrest.

Besides the decreased levels of these regulatory proteins, there are several

other mechanisms that cause G2/M phase arrest such as the increased levels of tumor

suppressor protein p53 and cdk inhibitory (CKI) p21 (Charrier-Savournin et al., 2004),

the inhibition of mitogen‐activated protein kinases (MAPK)/extracellular signal-

regulated kinases (ERK) signaling pathway (Yin et al., 2014), as well as the disruption

of microtubule assembly as well as the disruption of microtubule assembly (Chang, Yu,

Wu, Wang, & Liu, 2011). An example of a plant-derived compound that induced

excessive G2/M phase arrest is diallyl disulfide, a natural organosulfur compound

isolated from garlic. It caused a decline in protein expression levels of cyclin B1, cdc2,

p-cdc2, cdc25C and an increase in mRNA levels of p53 and p21. Also, this compound

inhibited cell proliferation in human esophageal squamous carcinoma ECA-109 cells

through the MEK-ERK signaling pathway (Yin et al., 2014). Therefore, it is possible

that the molecular mechanisms underlying the antiproliferative effect of HMP were

likely similar to those of diallyl disulfide in that more than one mechanism is involved.

Further studies are required to elucidate. In this study, the effect of HMP on the

disruption of microtubule assembly that caused G2/M arrest was also investigated. The

results turned out that HMP at 25 μM had no effect on microtubule assembly in a cell-

free system. However, its effect may be different in cell-based systems if HMP

indirectly disrupts the polymerization of microtubules in cells by affecting some

microtubule-regulatory proteins (Duangmano, Sae-Lim, Suksamrarn, Domann, &

Patmasiriwat, 2012). Additional research is necessary to fully define the mechanisms.

Given markedly increased G2/M arrest, there may be multiple molecular mechanisms

involved in the control of G2/M-phase progression.


120

5.3 Cytotoxic effects of HMP in A549 cells

Besides the antiproliferative effect, HMP also exerted the cytotoxic effect

against A549 cells by presenting the cell survival curves with parameter LC50, which

displays a significant net loss and the increased sub-G1 peak after HMP treatment. In

addition, such effect was also related to the induction of apoptotic cell death by the

presence of early apoptotic cells (annexinV+/PI–). Briefly, one of the biochemical

hallmarks of apoptosis is the translocation of PS to the outer plasma membrane where

annexin-V-FITC binds specifically to PS. Moreover, double staining with PI

differentiates early apoptotic cells with intact membrane from late apoptotic/necrotic

cells with leaky membranes and healthy cells (Vermes et al., 1995). Furthermore, HMP

treatment also presented the characteristic features of apoptosis, such as chromatin

condensation and nuclear fragmentation by DAPI staining as well as the presence of

DNA ladder by agarose gel electrophoresis.

5.4 Apoptosis underlying cytotoxic effects of HMP

In apoptosis, caspases are synthesized as inactive procaspases that need to

be proteolytically processed to generate the active enzymes in response to apoptotic

signals (Steller, 1998). There are two main apoptotic pathways including intrinsic and

extrinsic pathways. Both pathways finally converge on caspase-3, which can cleave

many key cellular proteins such as the inhibitor of caspase-activated DNase (ICAD),

PARP and other structural proteins, causing nuclear shrinking, budding to form

apoptotic bodies, cytoskeletal proteolysis, etc. (Porter & Janicke, 1999; Elmore, 2007).

In this study, our results revealed that the use of Z-VAD-fmk was efficient in blocking

apoptotic cell death in HMP-treated A549 cells, indicating that HMP induced apoptosis

through activation of caspases. In addition, we first demonstrated the effect of HMP on

caspase-3 activation through detection of both caspase-3 activity and its active

(cleaved) form. Following HMP treatment, the cleavage of PARP and the presence of

nuclear condensation, DNA fragmentation, and DNA apoptotic ladder were also

detected. These are considered as indicative of functional caspase-3 activation.


121

It is well established that p53 is a sensor of cellular stress and is a critical

activator of the intrinsic apoptosis pathway (Haupt, Berger, Goldberg, & Haupt, 2003).

Once activated in the cytosol, p53 translocates to the nucleus where it activates

transcription of many proapoptotic proteins of the Bcl-2 family while simultaneously

repressing that of antiapoptotic proteins (Chumakov, 2007). As a result, proapoptotic

proteins subsequently oligomerize and form the pores, leading to the disrupted

mitochondrial outer membrane, cytochrome c release into the cytoplasm, the

apoptosome formation, and subsequent activation of caspase-9 (Zou et al.,

2003). The present study clearly showed that HMP could achieve the intrinsic apoptosis

pathway through activation of procaspase-9. The finding that levels of proapoptotic Bax

protein, one of the downstream targets of p53, were increased through its upregulated

mRNA levels, additionally supports the involvement of p53 in the intrinsic pathway.

The presence of wild-type p53 in A549 cells could increase such possibility.

Unexpectedly, however, increased expression levels of Bcl-2 were detected in HMP

treatment. This implies that Bcl-2 was likely not a key protein in HMP-mediated

apoptosis. It is possible that other antiapoptotic proteins i.e. Bcl-xL may be more

involved in this action (Zhang & Rosdahl, 2006). Therefore, further studies are needed.

The extrinsic apoptotic pathway is triggered when specific death ligands

engage their receptors on the plasma membrane, leading to the activation of initiator

caspase-8 (Elmore, 2007). Subsequently, active caspase-8 can directly cleave and

activate caspase-3, or it can alternatively cleave its downstream target Bid. This

truncated Bid can then activate proapoptotic Bax and Bak proteins directly as well as

suppress the anti-apoptotic proteins at the mitochondria, causing MOMP and

propagating the intrinsic mitochondrial pathway (Kantari & Walczak, 2011). However,

it is noteworthy to mention that the extrinsic death signals cannot directly elevate the

mRNA levels of Bax and Bak. In this study, the cleavage of Bid was detected and

therefore, provided additional insight into the extrinsic apoptosis pathway mediated by

HMP.

In summary, for the first time, this study demonstrated that HMP exerted

anticancer activity through the induction of G2/M cell cycle arrest and apoptosis in

A549 cells. Similarly, several well-known plant-derived compounds such as curcumin


122

(Cheng et al., 2016) and baicalein (Mu et al., 2016) have been revealed to mediate cell

cycle arrest and apoptosis in recent years.

5.5 Effects of HMP on the generation of intracellular ROS and the relationship

between enhanced ROS and apoptosis

In addition, recent studies have shown that many plant-derived compounds

can produce excessive ROS leading to the induction of apoptosis in certain cancer cell

lines e.g., capsaicin (Ito et al., 2014), curcumin (Chang, Xing, & Yu, 2014), Plumbagin

(Tian et al., 2012), and so forth. The relationship between enhanced ROS and apoptosis

is that ROS can stimulate proapoptotic signaling molecules such as apoptosis signal-

regulating kinase 1 (ASK1), c-Jun-NH2-kinase (JNK), and p38 (Benhar, Dalyot,

Engelberg, & Levitzki, 2001; Tobiume et al., 2001), which then activate the p53 protein

pathway or engage the mitochondrial apoptotic cascade (Alexandre, Batteux, & Nicco,

2012), leading to apoptotic cell death. Our results showed that HMP induced ROS

production in a dose-dependent manner, and increased intracellular ROS levels were

completely abolished by the ROS scavenger, NAC, indicating that HMP indeed

induced elevation of ROS levels in A549 cells. However, the pretreatment with NAC

failed to inhibit the induction of apoptosis, indicating that this increased ROS was not

likely involved in HMP-mediated apoptosis.


123

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143

APPENDICES


144

APPENDIX A

GROWTH CURVE

Figure A-1 Growth curve of human lung carcinoma cell line A549 in 96-well plates.

The optimal cell numbers of this cell line were 3,200 cells/well. Data are expressed as

mean (n ≥ 3). Each experiment was performed in triplicate.

0.000

0.500

1.000

1.500

2.000

2.500

3.000

3.500

0 24 48 72 96

O.D

. 5

70

nm

Time (hours)

Growth curve (A549), 96 wells

12,800 cells/well

6,400 cells/well

3,200 cells/well

1,600 cells/well

800 cells/well


145

Figure A-2 Growth curve of human lung squamous carcinoma cell line NCI-H226 in

96-well plates. The optimal cell numbers of this cell line were 3,200 cells/well. Data

are expressed as mean (n ≥ 3). Each experiment was performed in triplicate.

0.000

0.500

1.000

1.500

2.000

2.500

3.000

3.500

0 24 48 72 96

O.D

. 5

70

nm

Time (hours)

Growth curve (NCI-H226), 96-well plate

12,800 cells/well

6,400 cells/well

3,200 cells/well

1,600 cells/well

800 cells/well


146

Figure A-3 Growth curve of human large cell lung cancer line COR-L23 in 96-well

plates. The optimal cell numbers of this cell line were 3,200 cells/well. Data are

expressed as mean (n ≥ 3). Each experiment was performed in triplicate.

0.000

0.500

1.000

1.500

2.000

2.500

3.000

3.500

0 24 48 72 96

O.D

. 5

70

nm

Time (hours)

Growth curve (COR-L23), 96-well plate

12,800 cells/well

6,400 cells/well

3,200 cells/well

1,600 cells/well

800 cells/well


147

Figure A-4 Growth curve of human small cell lung cancer cell line NCI-H1688 in 96-

well plates. The optimal cell numbers of this cell line were 6,400 cells/well. Data are

expressed as mean (n ≥ 3). Each experiment was performed in triplicate.

0.000

0.500

1.000

1.500

2.000

2.500

3.000

3.500

0 24 48 72 96

O.D

. 5

70

nm

Time (hours)

Growth curve (NCI-H1688), 96-well plate

12,800 cells/well

6,400 cells/well

3,200 cells/well

25,600 cells/well


148

Figure A-5 Growth curve of human lung fibroblast cell line MRC-5 in 96-well plates.



0.000

0.500

1.000

1.500

2.000

2.500

3.000

3.500

0 24 48 72 96

O.D

. 5

70

nm

Time (hours)

Growth curve (MRC-5), 96-well plate

12,800 cells/well

6,400 cells/well

3,200 cells/well

1,600 cells/well

800 cells/well


149

Figure A-6 Growth curve of human lung carcinoma cell line A549 in 24-well plates.



0.000

0.500

1.000

1.500

2.000

2.500

3.000

3.500

0 24 48 72 96

O.D

. 5

70

nm

Time (hours)

Growth curve (A549), 24-well plate

128,000 cells/well

64,000 cells/well

32,000 cells/well

16,000 cells/well

256,000 cells/well


150

APPENDIX B

STANDARD CURVE FOR PROTEIN DETERMINATION

Figure B-1 Standard curve for protein determination by Bradford’s method.

Figure B-2 Standard curve for protein determination by BCA Assay.

y = 0.9325x + 0.0962

R² = 0.9928

0.000

0.200

0.400

0.600

0.800

1.000

1.200

0.000 0.250 0.500 0.750 1.000 1.250

O.D

. 5

95

nm

BSA concentration (mg/ml)

BSA standard curve by Bradford's method

y = 1.1836x + 0.0715

R² = 0.9984

0.000

0.500

1.000

1.500

2.000

2.500

3.000

0.000 0.500 1.000 1.500 2.000 2.500

O.D

. 5

62 n

m

BSA concentration (mg/ml)

BSA standard curve by BCA assay


151

APPENDIX C

HPLC CHROMATOGRAMS

Figure C-1 HPLC chromatogram of HMP-1.


152

Figure C-2 HPLC chromatogram of HMP-2.


153

Figure C-3 HPLC chromatogram of 5, 6-dihydroxy-2, 4-dimethoxy-9, 10-dihydro

phenanthrene.


154

APPENDIX D

FLOW CYTOMETRIC ANALYSIS

Figure D-1 Flow cytometric analysis of the DNA from A549 cells treated with NAC

alone at different concentrations (0.1, 1 and 5 mM) for 72 h. Data are representatives

of three independent experiments.


155

APPENDIX E

REAGENTS FOR LABORATORY EXPERIMENTS

1. Reagents for cell culture

RPMI medium 1640

Dissolve 10.4 g of RPMI medium 1640 powder (Biochrom, Germany) and

2.0 g of sodium bicarbonate in distilled water to a final volume of 1,000 ml.

The medium was adjusted pH to 7.4 with 1 M HCl (hydrochloric acid) and

was then filtered through 0.22 micron of filter. The complete RPMI 1640

medium was mixed with 10% heat-inactivated FBS and was stored at 4°C.

Phosphate buffer saline (PBS) Solution

Dissolve 9.55 g of PBS powder without Ca2+, Mg2+ (Biochrom, Germany)

in 1,000 ml distilled water. PBS solution was then filtered through 0.22

micron of filter and stored at 4°C.

Trypsin-EDTA Solution

Dissolve 10 ml Trypsin (1:250)/EDTA (0.5/0.2 %) in 10x PBS without

Ca2+, Mg2+ (Biochrom, Germany) in 90 ml 1x PBS solution.

2. Reagents for SRB assay

1% glacial acetic acid

Glacial acetic acid 10.00 ml

Distilled water 990.00 ml

0.4% (w/v) Sulforhodamine B (SRB) solution

Dissolve 0.4 g of sulforhodamine B dye to a final volume of 100 ml with

1% glacial acetic acid and keep at room temperature.


156

10% TCA (trichloroacetic acid)

Dissolve 10 g of trichloroacetic acid to a final volume of 100 ml with

distilled water and keep at 4°C.

Tris-base [10 mM, pH 10]

Tris 605.50 mg


Adjust pH into 10.0, then add distilled water to a final volume 500 ml and

keep at room temperature.

3. Reagents for SDS-PAGE preparation

5x running buffer (1L)

0.25 M Tris 15.15 g

1.92 M glycine 72.00 g

1% SDS 5.00 g

Adjust to a final volume of 1,000 ml with deionized water and keep at 4ºC.

1x running buffer (working solution)

5x running buffer 200.00 ml


4x stacking gel buffer [0.5M tris-Hcl, pH 6.8, 100ml]

Tris 6.055 g

Deionized water 80.00 ml

Adjust pH into 6.8 and add deionized water to a final volume 100 ml

4x separating (resolving) gel buffer [1.5M tris-Hcl, pH 8.8, 200ml]

Tris 36.33 g


Adjust pH 8.8 and add deionized water to a final volume 200ml


157

10% APS (ammonium per sulfate)

Ammonium per sulfate 10.00 g


10% SDS (Sodium dodecyl sulfate)

SDS 10.00 g


Table E-1 Recipes for resolving and stacking gels

Reagents Resolving gel

Stacking

gel

12% 7.5% 4%

30% Acrylamide/Bis Solution,

29:1 (Bio-Rad) 4.0 ml 2.5 ml 660 µl

4x resolving buffer 2.5 ml 2.5 ml -

4x stacking buffer - - 1.26 ml

10% SDS 100 µl 100 µl 50 µl

Deionized water 3.35 ml 4.85 ml 3 ml

10% APS 50 µl 50 µl 25 µl

TEMED, (Bio-Rad) 5 µl 5 µl 5 µl

Total 10 ml 10 ml 5 ml

4. Reagents for Western blotting

5x TBS (Tris-Buffered Saline) solution

200 mM Tris 12.10 g

1.5 mM NaCl 40.30 g

Adjust pH to 7.5 with 1 M HCl and make volume up to 1 L with distilled

water. Keep at 4ºC.

1x TBS solution

5x TBS 200.00 ml



158

TBST (Tris-Buffered Saline Tween-20) solution

5x TBS 200.00 ml


0.1% Tween 20 1.00 ml

1x transfer blotting buffer (Prepare fresh buffer)

Tris 3.00 g

Glycine 14.40 g


Methanol 200.00 ml

Destaining solution

Methanol 200.00 ml

Glacial acetic acid 100.00 ml

H2O 700.00 ml

Keep at room temperature.

Stripping solution [2.5 mM glycine, 1-2% SDS]

Glycine 0.9 g

SDS 10.00 g

Adjust pH to 2.0 with 1 M HCl and make volume up to 500 ml with

distilled water. Keep at 4ºC.

5. Reagents of DAPI staining

20 mg/ml DAPI solution (Stock solution)

Dissolve 1 mg of DAPI (Sigma, Cat.No. 9542) in 50 µl distilled water.

1 µg/ml DAPI solution

DAPI staining solution (Stock 20 mg/ml) 1.00 µl



159

80% Ethanol

Absolute ethanol 80.00 ml


6. Reagents for DNA electrophoresis and staining solution

5x TBE buffer

Tris 53.90 g

Boric acid 27.50 g

EDTA 3.70 g

Add distilled water to a final volume of 1,000 ml

1x TBE buffer

10x TBE buffer 200.00 ml


1.5% agarose gel

Agarose gel 1.50 g

1x TBE buffer 100.00 ml

DNA Staining solution

Ethidium bromide (10 mg/ml) 10.00 µ1



160

BIOGRAPHY

Name Miss Wipada Duangprompo

Date of Birth January 20, 1983

Educational Attainment

2004: Bachelor of Science (Medical Technology)

Faculty of Associated Medical Science

Khon Kaen University, Thailand

2007: Master of Science (Parasitology)

Department of Parasitology

Faculty of Medicine

Khon Kaen University, Thailand

Scholarship 2013: Research Grants of Thammasat University

for Ph.D. students

2015: TU Research Scholar, Contract No.79/2558

Publications

1. Duangprompo W, Aree K, Itharat A, Hansakul P. Effects of 5,6-dihydroxy-

2,4-dimethoxy-9,10-dihydrophenanthrene on G2/M cell cycle arrest and

apoptosis in human lung carcinoma A549 cell. The American Journal of

Chinese Medicine. 2016; 44 (7):1473-1490.

Oral/Poster presentation

1. Duangprompo W, Hansakul P. Antiproliferative and Apoptotic Effects of a

Novel 9, 10-Dihydrophenanthrene Isolated from Dioscorea membranacea in

the Human Non-Small Cell Lung Cancer cell line A549. The 18th World

Congress on Clinical Nutrition (WCCN) "Agriculture, Food and Nutrition for

Health and Wellness" December 1-3, 2014: Ubon Ratchathani, Thailand.

(Oral presentation).

2. Duangprompo W, Aree K, Itharat A, Hansakul P. Anticancer Effects of a

Novel 9, 10-Dihydrophenanthrene Isolated from Dioscorea membranacea


161

Pierre in A549 cells. The 74th Annual Meeting of the Japanese Cancer

Association. October 8 –10, 2015: Nagoya Congress Center, Japan. (Poster

presentation).

3. Duangprompo W, Aree K, Itharat A, Hansakul P. Antiproliferative activity

of 5,6-dihydroxy-2,4-dimethoxy-9,10-dihydrophenanthrene (HMP) derived

from Dioscorea membranacea Pierre against A549 cells. The 5th

International Biochemistry and Molecular Biology Conference. May 26-27,

2016. B.P: Samila Beach Hotel, Songkhla, Thailand. (Poster presentation).

Documents

Growth arrest and caspase-dependent apoptosis induced by 5 ...ethesisarchive.library.tu.ac.th/thesis/2016/TU_2016_5411301012_5959_4158.pdf · activation and further indicating the