REDOX REGULATION OF ESTROGEN RECEPTOR ALPHA AND SODIUM HYDROGEN EXCHANGER 1 GENE EXPRESSION BY HYDROGEN PEROXIDE
CHUA SUI HUAY BSc (HONOURS)
A THESIS SUBMITTED FOR THE DEGREE OF DOCTOR OF PHILOSOPHY
NUS GRADUATE SCHOOL FOR INTEGRATIVE SCIENCES AND ENGINEERING NATIONAL UNIVERSITY OF SINGAPORE 2011
ACKNOWLEDGEMENTS
I want to express my heartfelt gratitude for my supervisor Associate Professor Marie-
Veronique Clement, Department of Biochemistry. I am thankful that she allowed me to join the lab during my honors year and for continuing my graduate study under her.
A/P Clement was not just a supervisor, but a mentor who is always willing to listen, to encourage and to share her vast knowledge. I would like to thank her for her constant encouragement, patience and faith in my abilities to complete this study.
I would also like to thank my co-supervisor Dr Alan Kumar for his guidance and support throughout the course of my graduate study in NUS. I am grateful for
Professor Shazib Pervaiz for being on my thesis advisory committee and for his positive feedback on the direction of my project.
My warmest thanks to my lab mates in Biochemistry department, for making the long days in lab enjoyable. Special thanks to San Min and Jeya who helped me to proof- read my thesis. I also want to thank Mui Khin for helping me with all the ordering of reagents for my project.
Thank you NUMI girls for being my great lab mates and friends. Thanks for being there to cheer me on and also for being part of my sister gang during my wedding. I will certainly miss those days that we hung out together.
Finally, my deepest gratitude to my family for their encouragement and support throughout the years, and a special thanks to Zong Hao, who has stood by me through the good and bad times.
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SUMMARY
There is mounting evidence that implicates reactive oxygen species (ROS) as an important signaling molecule in various physiological processes. In contrast to the wealth of information on gene up-regulation by ROS, little attention has been paid to the down-regulation of gene expression by ROS. In this thesis, we have studied how two important genes, estrogen receptor alpha (ER α) and sodium hydrogen exchanger
1 (NHE1) were redox-regulated by H 2O2.
In the first part of the study, the regulation of ER α by oxidative stress induced by the
exposure of MCF7 cells to H 2O2 was investigated. The data supported that ER α
protein is down-regulated when exposed to oxidative stress. The down-regulation of
ER α protein occurs not through proteosomal degradation pathway, but via the decrease in ER α mRNA level. We found that Akt, MAPK and caspases were not
involved in the down-regulation of ER α by H 2O2. Instead, H2O2 inhibition of ER α
expression involves an oxidation event that is reversible by the addition of reducing
agent, DTT. We also demonstrated that 15d-PGJ2 suppressed ER α expression via the
production of ROS. ER α down-regulation resulted in decreased ER α-responsive gene
expression and impaired estrogen signaling. These effects could have contributed to
the growth arrest observed in H2O2 treated MCF7 cells.
In the second part of the study, the down-regulation of NHE1 by H2O2 was examined.
We found that the minimal promoter region required for full transcription activation
lies on the 0.15kb of NHE1 promoter. This region is also regulated by H2O2. The regulation of NHE1 by H2O2 is similar to the regulation of ER α. H2O2 targets NHE1 at the mRNA level in a reversible manner. The down-regulation involves an oxidation mechanism which is reversible by reducing agents DTT and BME. The drug 15d-
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PGJ2 was also shown to down-regulate NHE1 promoter activity via the production of
ROS. Finally, the data suggested that AP2 is not the transcription factor for NHE1 and is not a target in H 2O2 mediated down-regulation of NHE1.
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TABLE OF CONTENTS
ACKNOWLEDGEMENTS …………………………………………………..………II
SUMMARY………………………………………………………………………….III
TABLE OF CONTENTS……………………...……………………………………...V
LIST OF TABLES…………………………………………………………………....X
LIST OF FIGURES………………………………………………..………………...XI
ABBREVIATIONS……………………………………………………………...…XVI
PUBLICATIONS AND PRESENTATIONS……………………………………...XIX
CHAPTER 1: INTRODUCTION ...... 1
1.1 FREE RADICALS, REACTIVE SPECIES AND REDOX BALANCE ...... 1 1.1.1 Reactive oxygen species ...... 1 1.1.2 The antioxidant system ...... 5 1.1.3 Hydrogen peroxide as a signaling molecule ...... 7
1.2 REDOX REGULATION OF GENE EXPRESSION ...... 10 1.2.1 Transcriptional regulation ...... 10 1.2.2 mRNA stability ...... 13 1.2.3 Direct oxidative modification ...... 14 1.2.4 Regulation of protein turnover ...... 16
1.3 ESTROGEN RECEPTOR ...... 17 1.3.1 Estrogen and estrogen receptors: Introduction ...... 17 1.3.2 Estrogen receptor and genomic signaling ...... 20 1.3.3 Membrane and cytoplasmic ER signaling ...... 21 1.3.4 Regulation of estrogen receptors ...... 23 1.3.5 Estrogen and estrogen receptors in human breast cancer ...... 27
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1.4 THE SODIUM HYDROGEN EXCHANGER 1 ...... 29 1.4.1 pH regulation in the cells ...... 29 1.4.2 Mammalian Na +/H + exchanger ...... 30 1.4.3 NHE1: Basic structure ...... 31 1.4.4 Physiological and pathological roles of NHE1 ...... 33 1.4.5 Regulation of NHE1 ...... 37 1.4.6 Regulation of NHE1 gene transcription ...... 39
1.5 AIM OF STUDY ...... 41
CHAPTER 2: MATERIALS AND METHODS ...... 43
2.1 MATERIALS ...... 43 2.1.1 Chemicals and reagents...... 43 2.1.2 Antibodies ...... 44 2.1.3 Plasmids ...... 44 2.1.4 Cell lines and cell culture ...... 45
2.2 METHODS...... 47
2.2.1 Treatment of cells with hydrogen peroxide (H 2O2) and other compounds………………………………………………………………………47 2.2.2 Morphology studies ...... 47 2.2.3 Crystal violet assay ...... 47 2.2.4 Plasmid transfection ...... 48 2.2.5 Small interfering RNA (siRNA) transfection ...... 48 2.2.6 Protein concentration determination ...... 49 2.2.7 SDS-PAGE and western blot ...... 50 2.2.8 Caspase assay ...... 51 2.2.9 Single and dual luciferase assay ...... 52 2.2.10 Chloramphenicol acetyl transferase (CAT) assay...... 53 2.2.11 RNA isolation ...... 54 2.2.12 Reverse transcription (RT) and real-time chain polymerase reaction (PCR)……………………………………………………………………………54 2.2.13 Intracellular ROS measurement by CM-H2DCFDA ...... 55 2.2.14 Nuclear-cytoplasmic fractionation ...... 55 2.2.15 Electromobility shift assay (EMSA) ...... 56
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2.2.16 Statistical analysis ...... 57
CHAPTER 3A: RESULTS – REDOX REGULATION OF ER α ...... 58
3A.1 H2O2 AND THE EXPRESSION OF ER α PROTEIN...... 58
3A.1.1 Morphology and growth of MCF7 cells upon exposure to H 2O2 ...... 58
3A.1.2 Effect of H 2O2 on the expression of ER protein ...... 61
3A.1.3 Effects of estrogen on H 2O2 down-regulation of ER α ...... 66
3A.1.4 Time-dependent regulation of ER α protein by H 2O2 ...... 68 3A.1.5 Chronic ROS leads to continuous suppression of ER α ...... 70 3A.1.6 ROS involved in the down-regulation of ER α expression ...... 71 3A.1.7 Effect of peroxynitrite on the expression of ER α protein ...... 73
3A.2 DOWN-REGULATION OF ER α DOES NOT INVOLVE THE PROTEOSOMAL DEGRADATION PATHWAY ...... 74
3A.3 THE DOWN-REGULATION OF ER α BY H 2O2 IS VIA DECREASE IN ER α mRNA LEVEL ...... 78
3A.3.1 Effects of H 2O2 on the expression of ER α mRNA level...... 78
3A.3.2 New protein synthesis is not required for H 2O2-induced down-regulation of ER α mRNA...... 81
3A.4 MECHANISM INVOLVED IN H 2O2-INDUCED DOWN-REGULATION OF ER α ...... 82
3A.4.1 Role of Akt activation in H 2O2 down-regulation of ER α ...... 82
3A.4.2 Role of MAPK activation in H 2O2 down-regulation of ER α ...... 84
3A.4.3 Role of caspases in H 2O2 down-regulation of ER α ...... 87
3A.4.4 Oxidation is involved in the down-regulation of ER α by H 2O2 ...... 89
3A.5 EFFECTS OF ER α DOWN-REGULATION ON ER RESPONSE GENES.……………………………………………………………………………94
3A.5.1 Down-regulation of ER α by H 2O2 inhibits estrogen response element (ERE) activity ...... 94
3A.5.2 H2O2 down-regulates ER α response genes ...... 96
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3A.6 15-DEOXY-DELTA-12,14-PROSTAGLANDIN J2 (15D-PGJ2) DOWN- REGULATION OF ER α IS VIA ROS ...... 99 3A.6.1 Morphology and growth of MCF7 cells exposed to 15d-PGJ2 ...... 99 3A.6.2 15d-PGJ2 down-regulates ER α at the protein and mRNA level ...... 101 3A.6.3 15d-PGJ2 produces ROS in MCF7 cells ...... 103 3A.6.4 Scavenging of ROS by NAC restores cell growth and ER α expression..…...………………………………………………………………..107
CHAPTER 3B: RESULTS – REDOX REGULATION OF NHE1……………...…110
3B.1 DOWN-REGULATION OF NHE1 BY H 2O2 ...... 110 3B.1.1 Identification of crucial region in the promoter of NHE1 ...... 110
3B.1.2 H2O2 down-regulates 0.15kb proximal NHE1 promoter activity ...... 114 3B.1.3 Oxidation is involved in the down-regulation of 0.15kb proximal NHE1 promoter activity and endogenous mRNA level by H 2O2 ...... 118 3B.1.4 Peroxynitrite down-regulates 0.15kb NHE1 promoter activity ...... 124
3B.1.5 Role of caspases in H 2O2-mediated down-regulation of 0.15kb NHE1 proximal promoter activity ...... 125 3B.1.6 15d-PGJ2 down-regulates 0.15kb NHE1 promoter activity and NHE1 mRNA level via ROS production ...... 129
3B.1.7 H2O2 down-regulates NHE1 expression in MCF7 breast cancer cells ……………………………………………………………………………135
3B.2 TRANSCRIPTION FACTOR INVOLVED IN H 2O2 DOWN-REGULATION OF NHE1 EXPRESSION ...... 141 3B.2.1 Analysis of the region between 0.15kb and 0.12kb of NHE1 promoter...……………………………………………………………………..141 3B.2.2 Transcription factor AP2, SP1 and COUP ...... 145 3B.2.3 Investigation of AP2 as transcription factor for NHE1 ...... 148
CHAPTER 4: DISCUSSION………………………………………………………. 162
4.1 REDOX REGULATION OF ER α ...... 162 4.1.1 Regulation of ER α expression and function by oxidative stress ...... 163
4.1.2 Oxidation as the mechanism mediating H 2O2-induced down-regulation of ER α…………………………………………………………………………170 4.1.3 Significances of ER α as a redox regulated gene ...... 172 4.1.4 ROS-induced suppression of ER α gene expression: Possible development of ER α-negative breast cancer phenotype ...... 178 viii
4.2 REDOX REGULATION OF NHE1 ...... 181
4.2.1 H2O2 regulates an important region of NHE1 promoter ...... 181 4.2.2 Targeting NHE1 expression through ROS-producing agents ……….184 4.2.3 AP2 is a transcription factor of NHE1: True or false? ...... 186
4.3 ER α AND NHE1: TWO IMPORTANT MEDIATORS OF CANCER CELL SURVIVAL ...... 189
4.3.1 H2O2 regulates ER α and NHE1 in similar ways ...... 189 4.3.2 Relevance of ER α and NHE1 expression as prognosis markers in breast cancer…………………………………………………………………………..191
4.4 FUTURE WORK ...... 192
4.4.1 To determine if H 2O2 affect the stability of ER α mRNA ...... 192 4.4.2 To investigate the mechanism of ROS-producing compounds in suppression of ER α expression...... …193 4.4.3 To identify the transcription factor that regulates NHE1 transcription……………………………………………………………………194
4.5 MODEL AND CONCLUSION ...... 195
APPENDICES ...... 197
REFERENCES ...... 202
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LIST OF TABLES
Table I: Characteristics of H 2O2 as second messenger ...... 9
Table 1: Software used in the analysis of the 40 bp of 0.15kb NHE1 promoter. .. 143
Table 2: Transcription factors predicted to bind to the 40 bp nucleotide sequence of
0.15kb NHE1 promoter...... 144
Table 3: List of compounds and growth factors found to down-regulate ER α and produce ROS...... 177
Table 4: Similarities between the down-regulation of ER α and NHE1 expression by
H2O2……………… ...... 190
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LIST OF FIGURES
Figure I: Pathways of reactive oxygen species (ROS) production and clearance ..... 2
Figure II: Sites of superoxide formation in the respiratory chain ...... 3
Figure III: Assembly of Nox2 in phagocytic cell ...... 5
Figure IV: The redox-cycling reactions involved in the catalytic removal of hydrogen peroxide by the glutathione peroxidase and thioredoxin peroxidase systems ...... 7
Figure V: Domain structure representation of human ER α and ER β isoforms ...... 19
Figure VI: Topology of NHE1 isoform and its regulatory elements ...... 32
Figure 1: MCF cell survival upon H 2O2 treatment ...... 60
Figure 2: H 2O2 induces concentration dependent down-regulation of ER α ...... 62
Figure 3: Specificity of ER α antibody ...... 63
Figure 4: H 2O2 down-regulates ER α in both MCF7 and T47D cells ...... 64
Figure 5: H 2O2 induces down-regulation of both nucleus and cytosolic fractions of
ER α………………………………………………………………………………...65
Figure 6: ER α is down-regulated in all serum conditions ...... 67
Figure 7: H 2O2 induces a time dependent down-regulation of ER α ...... 69
Figure 8: H 2O2-induced down-regulation of ER α is reversible ...... 70
Figure 9: Chronic oxidative stress lead to ER α-negative phenomenon ...... 71
Figure 10: Hydroxyl radical not involved in down-regulation of ER α by H 2O2 ..... 72
Figure 11: Peroxynitrite down-regulates ER α ...... 73
Figure 12: Effective concentration of proteosomal inhibition by MG132 ...... 75 xi
Figure 13: H 2O2-induced decrease in ER α is not due to proteosomal degradation . 77
Figure 14: H 2O2 induces time dependent down-regulation of ER α mRNA level ... 80
Figure 15: H 2O2 induces reversible down-regulation of ER α mRNA level ...... 80
Figure 16: Protein synthesis is not required in H 2O2 down-regulation of ER α mRNA level ...... 81
Figure 17: Inhibition of Akt activation does not prevent ER α down-regulation by
H2O2………………………………………………………………………………..83
Figure 18: H 2O2 activates MAPK ...... 85
Figure 19: The inhibition of ERK 1/2 activation does not prevent ER α down- regulation by H 2O2 ...... 85
Figure 20: Inhibition of JNK activation does not prevent ER α down-regulation by
H2O2...... …………………………………………………………………….……...86
Figure 21: H 2O2 does not activate caspase activity ...... 88
Figure 22: Reducing agent DTT prevents the down-regulation of ER α by H 2O2 ... 91
Figure 23: Thiol-oxidant diamide induces down-regulation of ER α ...... 93
Figure 24: H 2O2 inhibits total ERE activity induced by E2 ...... 95
Figure 25: H 2O2 decreases ER α response genes, PR and c-Myc expression ...... 97
Figure 26: H 2O2 decreases the ability of estrogen to induce c-Myc expression ...... 98
Figure 27: MCF cell survival upon 15d-PGJ2 treatment ...... 100
Figure 28: 15d-PGJ2 down-regulates ER α protein and mRNA levels ...... 102
Figure 29: 15d-PGJ2 produces ROS in MCF7 cells ...... 104
Figure 30: NAC scavenges 15d-PGJ2-induced ROS ...... 106
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Figure 31: Scavenging of ROS by NAC restores ER α expression ...... 108
Figure 32: Summary of the down-regulation of ER α by H 2O2 ...... 109
Figure 33: H 2O2 down-regulates NHE1 full length promoter activity ...... 111
Figure 34: 0.15kb promoter is basal promoter required for full NHE1 promoter activity ...... 113
Figure 35: H 2O2 down-regulates NHE1 mRNA in a concentration- and time- dependent manner ...... 115
Figure 36: H 2O2 down-regulates NHE1 0.15kb promoter in a concentration- and time-dependent manner ...... 117
Figure 37: Scavenging ROS with NAC prevents H 2O2-induced down-regulation of
0.15kb NHE1 promoter activity and NHE1 mRNA level ...... 119
Figure 38: Reducing agent DTT prevents H 2O2-induced down-regulation of 0.15kb
NHE1 promoter activity and NHE1 mRNA level ...... 120
Figure 39: Reducing agent BME prevents H 2O2-induced down-regulation of 0.15kb
NHE1 promoter activity and NHE1 mRNA level ...... 121
Figure 40: Thiol-oxidant diamide down-regulates NHE1 0.15kb promoter activity and NHE1 mRNA level ...... 123
Figure 41: Peroxynitrite down-regulates 0.15kb NHE1 promoter activity ...... 124
Figure 42: H 2O2 treatment activates caspase 2 and 3 in NIH 3T3 cells ...... 126
Figure 43: Pan caspase inhibitor zVADfmk does not prevent H 2O2-induced down- regulation of the 0.15kb NHE1 promoter ...... 128
Figure 44: 15d-PGJ2 down-regulates 0.15kb NHE1 promoter activity and NHE1 mRNA level ...... 130
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Figure 45: 15d-PGJ2 produces ROS in NIH 3T3 cells ...... 131
Figure 46: Scavenging ROS produced by 15d-PGJ2 with NAC restores 0.15kb
NHE1 promoter activity...... 134
Figure 47: H 2O2 and diamide down-regulate NHE1 mRNA level ...... 136
Figure 48: 0.15kb region of NHE1 promoter is highly conserved and important for
NHE1 transcription ...... 138
Figure 49: H 2O2 prevents binding of transcription factor(s) to NHE1 promoter region……………………………………………………………………………..140
Figure 50: Redox regulation of NHE1 transcription at the 0.15kb NHE1 promoter………………………………………………………………………….141
Figure 51: Predicted transcription factor binding sites ...... 144
Figure 52: AP2 α isoform is present in NIH 3T3 cells ...... 151
Figure 53: AP2 α is present in the nucleus in NIH 3T3 cells ...... 152
Figure 54: Over-expressing AP2 α does not affect NHE1 expression ...... 155
Figure 55: Over-expressing AP2 dominant negative does not affect NHE1 expression ...... 158
Figure 56: Silencing AP2 α does not affect NHE1 expression ...... 161
Figure 57: Model of the down-regulation of ER α and NHE1 by H 2O2 ...... 195
Appendix A: PPRE present in human ER α promoter A ...... 197
Appendix B: PPRE present in human and mouse NHE1 promoter ...... 199
Appendix C: siRNA from two companies failed to knockdown SP1 protein expression………………………………………………………………………...200
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Appendix D: NHE1 levels in different breast cell lines ...... 200
Appendix E: The expression level of NHE1 in tumor tissues is associated with clinical outcome ...... 201
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ABBREVIATIONS
15d-PGJ2 15-Deoxy-Delta-12,14-prostaglandin J2
AF Activation function
AFC 7-amino-4-trifluoromethyl coumarin
AP-1 Activator protein 1
AP2 Activator protein 2
ARE Antioxidant response element
ATP Adenosine triphosphate bp Base pair
BME β-mercaptoethanol
BSO Buthionine sulfoximine
BSA Bovine serum albumin
CAII Carbonic anhydrase II
CaM Calmodulin
CAT Chloramphenicol acetyl transferase
Cdk2 Cyclin-A-cyclin dependent kinase 2
CHP Calcineurin homologus protein
Chx Cycloheximide
CM-H2DCFDA 5-(and-6)-chloromethyl-2´,7´-dichlorodihydrofluorescein diacetate acetyl ester
COMT Catehol O-methyltransferase
COUP-TF Chicken ovalbumin upstream promoter-transcription factors
CS FBS Charcoal-stripped FBS
DMEM Dulbecco’s Modified Eagle’s Medium
DMSO Dimethylsulfoxide
DMTU Dimethylthiourea
DTT Dithiothreitol
E2 Estradiol
ER Estrogen receptor
ERE Estrogen response element
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EGF Epithelial growth factor
EMSA Electromobility shift assay eNOS Endothelial nitric oxide synthase
ERK Extracellular-signal-regulated kinase
ERM Ezrin, radixin and moesin
ETC Electron transport chain
FACS Fluorescence-activated cell sorting
FBS Fetal bovine serum
GSH Glutathione
GSSG Glutathione disulfide
Gpx Glutathione peroxidase
HIF Hypoxia-inducible factor
HO-1 Heme oxygenase-1
HuR Human antigen R
HRP Horse radish peroxidase
IκB Inhibitors of kappa B
IKK IκB kinase
IL-6 Interleukin -6
Keap1 Kelch-like ECH-associating protein 1
KGF Keratinocyte growth factor
kb Kilo base pair
MAPK Mitogen-activated protein kinase
MIP-1a Macrophage inflammatory protein-1a
MSR Macrophage scavenger receptor
NAC N-Acetyl-L-cysteine
NF κB Nuclear factor kappa B
NHE Sodium hydrogen exchanger
NIK Nck-interacting kinase
Non-silencing siRNA NS siRNA
Nox NADPH oxidase
O-GlcNAc O linked N-acetylglucosamine
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PBS Phosphate buffered saline
PDGF Platelet-derived growth factor pHi Intracellular pH
PKA Protein kinase A
PKC Protein kinase C
PMSF Phenylmethanesulfonyl fluoride
PP1 Phosphatase protein phosphatase 1
PP2A Protein phosphatase 2A
PPAR γ Peroxisome proliferator-activated receptor gamma
PPRE Peroxisome proliferator response element
Prx Peroxiredoxins
PTK Protein tyrosine kinase
PTP Protein tyrosine phosphatase
RAR-1 Retinoic acid receptor α-1
ROS Reactive oxygen species
RPMI-1640 Roswell Park Memorial Institute-1640
Rsk p90 ribosomal S6 kinase
TBST Tris-buffered saline tween
TNF α Tumor necrosis factor α
TRE TPA response element
SDS Sodium dodecyl sulphate
SOD Superoxide dismutase
SP1 Specificity Protein 1
Trx Thioredoxin
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PUBLICATIONS AND PRESENTATIONS
PUBLICATIONS: Chua SH and Clément MV. Oxidative suppression of ER α expression: A possible mechanism of action by ROS producing drugs (Manuscript in preparation).
POSTER PRESENTATIONS: Chua SH , Kumar AP and Clément MV. Role of Transcription Factor, AP2 in + + H2O2-mediated Repression of the Na /H exchanger 1 (NHE1) Gene Expression. Presented at 1st Biochemistry Student Symposium. Clinical Research Centre, National University of Singapore (2008).
Chua SH , Kumar AP and Clément MV. Role of Transcription Factor, AP2 in + + H2O2-mediated Repression of the Na /H exchanger 1 (NHE1) Gene Expression. Presented at XIV Biennial Meeting of the Society for Free Radical Research International Conference. Beijing, China (2008).
Chua SH and Clément MV. H2O2-induced growth arrest and down-regulation of ER α. Presented at 3rd Biochemistry Student Symposium. Clinical Research Centre, National University of Singapore (2010).
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CHAPTER 1: INTRODUCTION
1.1 FREE RADICALS, REACTIVE SPECIES AND REDOX BALANCE
1.1.1 Reactive oxygen species
Reactive oxygen species (ROS) are a group of molecules generated in the cell from the partial reduction of oxygen (O 2) as byproducts of aerobic metabolism. The radical
·- · species includes superoxide anion (O2 ) and hydroxyl radical (OH ), while hydrogen
·- peroxide (H 2O2) is the main non-radical species. The O2 , the precursor of most other
3 ROS, is formed from the reduction of triplet-state molecular oxygen ( O2) in a process mediated by membrane NADPH oxidase (Nox), xanthine oxidase, or leakage from the mitochondria electron transport chain (Dröge, 2002). In the cells, superoxide
·- dismutase (SOD) dismutates O2 into H 2O2 (Deby and Goutier, 1990), and H 2O2 can
be converted into highly reactive OH · in the presence of reduced transition metal such
as ferrous iron (Fe 2+ ) through the Fenton reaction :
2+ 3+ Fe + H 2O2 Fe + OH · + OH
OH · is highly reactive and is toxic to the cell due to its propensity to react and cause damage to a wide range of cellular components, including DNA, proteins, and lipids
· (Stinefelt et al., 2005). Other than forming OH , H2O2 can also be converted to water
(H 2O) through catalase, glutathione peroxidase (GPx) or peroxiredoxins (Prx) (Day,
· ·- 2009). In addition to forming H 2O2 and OH , O 2 also reacts with nitrite oxide (NO ·) to from a highly reactive oxidant, the peroxynitrite (ONOO -) (Jourd'heuil et al., 2001).
Figure I illustrates the pathways by which ROS is produced and cleared in the cells.
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Figure I: Pathways of reactive oxygen species (ROS) production and clearance. Glutathione (GSH), glutathione disulfide (GSSG). Adapted from (Dröge, 2002).
1.1.1.1 Mitochondria electron transport chain
The mitochondria electron transport chain (ETC) is an important source of ROS production in most mammalian cells (Turrens, 2003; Balaban et al., 2005; Andreyev et al., 2005). The ETC, found at the inner membrane of the mitochondria, is a multi- complex system which utilizes oxygen to generate energy in the form of adenosine triphosphate (ATP). It is composed of a series of electron carriers (flavoproteins, iron- sulfur proteins, ubiquinone and cytochromes) arranged into four complexes (Liu et al.,
2002). The ETC complex I accepts electrons from NADH and passes them through flavin and iron-sulfur centers to ubiquinone (Buchanan and Walker, 1996). Complex
II uses succinate as substrate and provides electrons to ubiquinone. Complex III accepts electrons from ubiquinone and passes them on to cytochrome c (Trumpower,
1990). Complex IV transfer electrons from cytochrome c to O 2, producing H 2O. In the
·- ·- mitochondria, O 2 is produced by the one-electron reduction of O 2 and O2 mediates oxidative chain reactions (Turrens, 2003). Figure II shows the sites of O2 production
·- at the ETC. Approximately 2% of O 2 is converted into O 2 through the ETC (Chance
2
·- et al., 1979). Complex I and III are shown to be the major sites of O 2 production by
the ETC (Turrens and Boveris, 1980; Turrens et al., 1985; Liu et al., 2002). While
·- complex II has been shown to be capable of producing O 2 under certain circumstance of dysfunction, complex IV is not known to release ROS (Senoo-
Matsuda et al., 2001; Lemarie et al., 2010). The ROS produced by the ETC contributes not only to damage in the mitochondria but is also crucial in the redox signaling of the cell.
Figure II: Sites of superoxide formation in the respiratory chain. Figure adapted from (Turrens, 2003)
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1.1.1.2 NADPH oxidase
Another important contributor of ROS in the cells is the membrane bound NADPH oxidase complex. Initially discovered in phagocytic cells, the NADPH oxidase (Nox2) is found to be responsible for oxidative burst to fight against invading microorganisms
(Nauseef, 2004). In phagocytes, the complex is made up of a membrane-spanning heterodimeric protein (flavocytochrome b588) which comprises pg91phox and p22phox; and cytosolic proteins p47phox, p67phox, p40phox and small GTP-binding protein Rac (Rac1 and Rac2) (Babior, 1999). Upon stimulation, the cytosolic components of the complex are recruited to the flavocytochrome b588 to catalyze the
·- ·- conversion of O 2 to O 2 (Groeger et al., 2009). Some of this O2 is converted to H 2O2
by Cu/Zn SOD in the cytosol. Figure III shows the assembly of Nox 2 in phagocytic
cell. Similar NADPH oxidase systems have also been found in non phagocytic cells.
Nox1 is the homolog of gp91phox and is found to be expressed in colon epithelial and
other various cell types (Takeya et al., 2006). Noxo1 and Noxa1 are homolog of
p47phox and p67phox respectively (Bánfi et al., 2003; Geiszt et al., 2003; Cheng and
Lambeth, 2004). Nox1 complexes with p22phox and interacts with Noxo1 and Noxa1
·- to produce O 2 (Ambasta et al., 2004; Sumimoto et al., 2004). Other forms of non-
phagocytic NADPH oxidases that exist include three Nox complexes (Nox3-Nox5)
and two dual oxidases (Duox1 and Duox2) (Groeger et al., 2009). The exact
composition of these non-phagocytic NADPH oxidases and the mechanism of ROS
production have not yet been completely elucidated. However, it is known that the
activation of signaling pathways involving cytokine receptors, G-protein coupled
receptors, receptor tyrosines and serine/threonine kinases can lead to ROS production
from these Nox complexes (Sauer et al., 2001). Nox-produced ROS can also play a
role in pathologies such as nephropathy, cardiovascular diseases, liver fibrosis and
4 pulmonary diseases (Li and Shah, 2003; Dworakowski et al., 2006; Minicis and
Brenner, 2007; Pantano et al., 2007).
Figure III: Assembly of Nox2 in phagocytic cell. Adapted from (Groeger et al., 2009).
1.1.2 The antioxidant system
The amount of reactive species in the cell is determined by the rate of their production and the rate of clearance by various antioxidant enzymes and compounds in the cell
(Dröge, 2002). Antioxidants are defined as substances that are able to compete with other oxidizable substrates at a relatively low concentrations, thus slowing down or preventing the oxidation of those substrates (Halliwell and Gutteridge, 2007). While transient increase in ROS plays a role in redox signaling, excessive level or chronic production of ROS will lead to oxidative stress (Jones, 2008). High level of ROS in the cell can cause damage to a wide range of molecules including lipid peroxidation,
DNA adduct formation or strand breaks, and oxidation of proteins which inhibits their functions. In mammalian cells, antioxidant defense systems are present to maintain the redox balance and reduce the damage caused by ROS. The SOD system plays a
5
·- ·- key role in the removal of excessive O 2 from the cell. It converts O 2 into H 2O2.
There are three SOD isoforms found in mammalian cells: cytosolic Cu/Zn SOD
(SOD1), mitochondria Mn SOD (SOD2), and extracellular SOD (EC SOD) (Choung
et al., 2004). H 2O2 in the cells are in turn removed mainly by three groups of enzymes: glutathione peroxidase (Gpx) system, thioredoxin peroxidase (peroxiredoxins) system and catalase (Dringen and Hamprecht, 1997) . While catalase catalyses the direct decomposition of H 2O2 to O 2 and H 2O, Gpx removes H 2O2 by coupling its reduction to H 2O with the oxidation of glutathione (GSH) to glutathione disulfide (GSSG).
GSSG is reduced back to GSH by NADPH-dependent glutathione reductase. The
thioredoxin peroxidase removes H 2O2 by coupling its reduction to H 2O with the oxidation of thioredoxin (Trx). Oxidized form of Trx is reduced back by NADPH dependent thioredoxin reductase (Veal et al., 2007). Figure IV illustrates the removal of H 2O2 by glutathione peroxidase and thioredoxin peroxidase systems. Several other non-enzymatic compounds also participate in the removal of ROS in the cell and they include ascorbate, α-tocopherol, β-carotene and GSH (Dröge, 2002).
6
Figure IV: The redox-cycling reactions involved in the catalytic removal of hydrogen peroxide by the glutathione peroxidase and thioredoxin peroxidase systems. Glutathione peroxidases (Gpx), peroxiredoxins (Prx) selenocysteine (SeH), cysteine (SH), glutathione (GSH) disulphide glutathione (GSSG), thioredoxin (Trx), sulfenic acid (SOH), and sulfinic acid (SOOH). Adapted from (Veal et al., 2007).
1.1.3 Hydrogen peroxide as a signaling molecule
It has been established in recent years that H 2O2 is an important regulator of signal
transduction in addition to its cytotoxic activity. H 2O2 is produced in response to a
variety of extracellular stimuli and the inhibition of H 2O2 production leads to the
attenuation of downstream cell signaling of the stimuli (Rhee, 2006; D'Autréaux and
Toledano, 2007). Stimulation of human epidermoid carcinoma cells with epithelial
growth factor (EGF) transiently increased H 2O2 in the cells. This H 2O2 production is
important for the EGF-induced tyrosine phosphorylation of various cellular proteins
including EGF receptor and phospholipase C-γ1(Bae et al., 1997). Scavenging of
H2O2 produced in response to platelet-derived growth factor (PDGF) inhibits cellular signaling such as tyrosine phosphorylation, mitogen-activated protein kinase stimulation, DNA synthesis, and chemotaxis in rat vascular smooth muscle cells
(Sundaresan et al., 1995).
7
Upon oxidative stress or stimulation with growth factors there is an increase in tyrosine phosphorylation level of numerous proteins (Schieven et al., 1993; Schieven et al., 1993; Nakamura et al., 1993; Hardwick and Sefton, 1995; Rhee, 2006). (Woo et al., 2010). H2O2 can promote tyrosine phosphorylation by activating protein tyrosine kinases (PTKs). One such example is the activation of the tyrosine kinase Src through oxidation at two cysteine residues by H 2O2 upon cell attachment to extracellular matrix (Giannoni et al., 2005). The activation of PTKs alone is however insufficient to increase steady-state protein tyrosine phosphorylation level in the cells. The concurrent inhibition of protein tyrosine phosphatases (PTPs) is required (Woo et al.,
2010). H 2O2 can inactivate PTPs by oxidizing the catalytic cysteine residue of these
enzymes, thus inhibiting their activities (Chiarugi and Cirri, 2003; Tonks, 2005; Rhee
et al., 2005). It was recently shown that the high level of H2O2 required in the cell to inhibit PTPs was achieved by the transient phosphorylation and inactivation of peroxiredoxin I at tyrosine-194 following stimulation of growth factor or immune receptors
H2O2 is also known to be involved in the activation of MAPK cascade. Angiotensin II induces the production of H 2O2 and activates ERK 1/2 and p38 MAPK to promote hypertrophy in vascular smooth muscle cells (Ushio-Fukai et al., 1998). More recently,
H2O2 have been shown to activate p38 MAPK to regulate the translocation of fibroblast growth factor 1 (FGF1) into the cytosol and nucleus (Sørensen et al., 2008).
The serine/threonine kinase protein kinase C (PKC) can be activated by H 2O2 through
tyrosine phosphorylation of the PKC catalytic domain (Konishi et al., 1997; Lin and
Takemoto, 2005). The cytosolic Ca 2+ level also plays an important role in signal
2+ transduction in the cell (Clapham, 2007). The level of Ca can be modulated by H 2O2
through mobilization of intracellular Ca 2+ store or through the influx of extracellular
8
Ca 2+ (Roveri et al., 1992; Doan et al., 1994; Hecquet et al., 2008; Giambelluca and
Gende, 2008). In summary, H 2O2 fulfils five criteria to act as a signaling molecule or second messenger in cells and this is summarized in Table I.
Table I: Characteristics of H 2O2 as second messenger. Summarized from (Forman, 2007)
Characteristics of second messenger H2O2 as second messenger Increases in concentration occur through: H2O2 increases in concentration through enzymatic generation by oxidoreductases and •− 1) enzymatic generation DuOXs and from dismutation of O 2
2) regulated release into the cytosol from sites of higher concentration through channels
Decreases in concentration through: H2O2 decreases through enzymatic degradation catalyzed by catalase, 1) enzymatic degradation glutathione peroxidases and peroxiredoxins
2) restoration of the concentration gradients by the action of pumps
3) diffusion from the cell enhanced by reaction or binding of the second messenger in another cell
Intracellular concentration rises and falls H2O2 concentration rises and falls within a within a short period short period from a steady state of nanomolar
Specific in action Specific in action
Gradients of their concentration from their H2O2 react within a few molecular diameters origin to where they are either degraded or of its site of production with its target effecter sequestered determines where they are due to high distribution of glutathione effective peroxidases and peroxiredoxins throughout the cell.
9
1.2 REDOX REGULATION OF GENE EXPRESSION
The regulation of gene expression is the control of the level and timing of appearance
of the functional product of a gene. A shift in the redox balance in the cell due to
either increase in ROS production or diminished antioxidant capacity can affect
cellular components. Redox system can regulate the expression of a gene through its
transcription, mRNA stability, posttranslational modifications, and protein stability
(Trachootham et al., 2008).
1.2.1 Transcriptional regulation
The regulation of gene expression starts at the transcriptional level, and is a tightly
regulated process. In response to changes in the redox environment, various genes are
up-regulated to enable the cells to cope with the changes. This is mediated through
redox control of transcription factors. Some of the better studied redox responsive
transcription factors are NF κB, AP1 and Nrf2.
1.2.1.1 NF κB
Nuclear factor kappa B (NF κB) is a redox sensitive factor that is involved in
immunity, inflammation, cell proliferation, development, and cell survival
(Trachootham et al., 2008). In mammals, the NF κB consist of five members: p50, p52, p65 (RelA), c-Rel, and Rel-B. These members can form various combinations of homo- and heterodimers. Normally, NF κB is sequestered in the cytosol by inhibitors of kappa B (I κB) (Hayden and Ghosh, 2004). Upon oxidative stress, I κB kinase (IKK)
is activated and phosphorylates I κB, which leads to the release of NF κB from I κB.
NF κB can then translocate to the nucleus and bind to the promoter region of target
10
genes. In response to oxidative stress, NF κB up-regulates anti-apoptotic genes such as
Bcl-xL, A1/Bfl-1, FLIP L, IAPs and TRAF1. It also up-regulates Gadd45, which inhibits JNK-induced cell death and it activates antioxidants genes such as MnSOD and ferritin heavy chain (Karin and Lin, 2002).
1.2.1.2 AP-1
The AP-1 transcription factor belongs to the leucine zipper domain family of protein that includes Fos and Jun (Curran, 1992). AP-1 proteins form heterodimers and bind to their DNA target sequence-12-O-tetradecanoyl phorbol-13-acetate (TPA) response element (TRE) (Risse et al., 1989). AP-1 controls the expression of large number of genes by binding to the TRE in the promoters regions of these genes (Vollgraf et al.,
1999; Toualbi et al., 2007). AP-1 activation could lead to either cell survival/proliferation or cell death depending on the circumstance of activation
(Shaulian and Karin, 2002). The activity of AP-1 is controlled at the transcriptional, posttranscriptional and posttranslational level (Hunter and Karin, 1992; Karin, 1995).
The activity and expression of AP-1 are both inducible by H 2O2 (Li and Spector, 1997;
Chung et al., 2002). Oxidative stress can activate c-Jun via phosphorylation at Ser63
and Ser73 by activated JNK (Karin, 1995; Li et al., 2004). Oxidative stress can also
promote the activation of AP-1 through histone acetylation by inhibiting histone
deacetylases (Rahman et al., 2002; Rahman, 2003).
11
1.2.1.3 Nrf2
Nrf2 is a key transcription factor involved in the protection against oxidative stress.
Activation of Nrf2 induces transactivation of many antioxidant genes, phase II detoxification enzymes, heat shock proteins and glutathione-synthesis enzymes (Itoh et al., 1997; Kobayashi and Yamamoto, 2005). Under normal conditions, Nrf2 is found in the cytosol where it interacts with Kelch-like ECH-associating protein 1
(Keap1) (Itoh et al., 1999). During oxidative stress, multiple cysteine residues of
Keap1 are oxidized which results in the dissociation of Keap1 from Nrf2 (Dinkova-
Kostova et al., 2002). Other than oxidizing Keap1, oxidants can also activate Nrf2 through phosphorylation of PKC and PERK. The free Nrf2 translocates into the nucleus and forms heterodimer with small Maf proteins (Motohashi et al., 2004). This complex then binds to antioxidant response element (ARE) of target genes promoter to initiate their transcription (Tanito et al., 2007).
1.2.1.4 Regulation of transcription by redox modulation of DNA binding domain
Many transcription factors contain redox-sensitive cysteine residues at their DNA binding domains (Haddad, 2002). The oxidation of these critical cysteine residues often leads to the inactivation of their DNA binding activity, thereby inhibiting target gene transcription (Turpaev, 2002). The reduced form of NF κB is required for its binding to DNA. Oxidation of Cys62, a redox sensitive site on p50 abolishes its DNA binding ability and inactivates downstream gene transcription (Toledano and Leonard,
1991; Mitomo et al., 1994). NF κB has also been shown to be inactivated by glutathionylation (Pineda-Molina et al., 2001) and S-nitrosylation (Marshall and
Stamler, 2001). Nitric oxide has been shown to inhibit DNA binding of AP-1 through
12
S-glutathionylation (Klatt et al., 1999). Hypoxia inducible factor-1 (HIF-1) DNA- binding activity is shown sensitive to oxidizing reagents diamide and H 2O2 and the alkylating agent N-ethylmaleimide (Wang et al., 1995; Nikinmaa et al., 2004). Other transcription factors that are modulated by redox regulation at their DNA binding domain include estrogen receptor (ER) (Hayashi et al., 1997);Weitsman et al., 2009),
Activating Protein 2 (AP2) (Huang and Domann, 1998), and Specificity Protein (SP1)
(Bloomfield et al., 2003; Ammendola et al., 1994).
1.2.2 mRNA stability
The regulation of mRNA stability is a major control point in gene expression. The stability of mRNA can be regulated through redox-mediated mechanisms. The stability of an mRNA depends on the interaction between RNA-binding proteins and structural elements of the mRNA (Guhaniyogi and Brewer, 2001). RNA binding protein human antigen R (HuR) is found mainly in the nucleus. However, upon exposure to oxidative stress triggers such as H 2O2, arsenite, or UV radiation, it translocates to the cytoplasm, and binds to AU-rich elements in the 3' untranslated regions (UTR) of mRNA. This increases the half-life of many mRNA encoding stress-response genes (Brennan and Steitz, 2001). Snail is a protein that plays a fundamental role in the induction of a phenotypic change called epithelial to mesenchymal transition (EMT). It has been shown that H 2O2 can increase Snail protein level by stabilizing Snail mRNA (Dong et al., 2007). The association of HuR to Snail mRNA was suggested to play a major role in H 2O2-induced Snail mRNA stability. H2O2 can also stabilize MKP-1 mRNA and increase the association of MKP-
1 mRNA with the translational machinery (Kuwano et al., 2008). It does so by increasing the association of MKP-1 mRNA with RNA binding proteins HuR and
13
NF90 and decreasing the association of MKP-1 mRNA with translation repressor
TIAR and TIA-1. Macrophage inflammatory protein-1a (MIP-1a) mRNA half-life
was markedly increased after H 2O2 treatment (Shi et al., 1996). H2O2 has also been implicated in increased activity of macrophage scavenger receptor (MSR) by stabilizing MSR-I mRNA (De et al., 1998).
Conversely, ROS has also been shown to decrease mRNA stability. Resistin was reported to decrease endothelial nitric oxide synthase (eNOS) mRNA stability through production of superoxide in the cells (Chen et al., 2010). H 2O2 and intracellular oxidative stress can destabilize the mRNA of ceruloplasmin (Cp), a copper-containing protein, by affecting the RNA protein complex formation at the 3’-UTR of Cp mRNA
(Tapryal et al., 2009). H2O2 has recently been shown to decrease mRNA stability of transcription factor atf1 in Schizosaccharomyces pombe cells (Day and Veal, 2010).
1.2.3 Direct oxidative modification
Oxidative modification of proteins is an important redox regulation of protein function at the posttranslational level (England and Cotter, 2005). Mild to moderate levels of oxidative stress can induce reversible modifications such as glutathionylation
(Ghezzi, 2005), S-nitrosylation (Sun et al., 2006) and disulfide bond formation
(O'Brian and Chu, 2005) on cysteine residue of proteins. These modifications by mild oxidative stress lead to structural and functional changes of the proteins.
Glutathionylation is the formation of mixed disulfides between GSH and proteins. In most incidences, glutathionylation leads to the inactivation of a protein’s function. For example, glutathionylation of transcription factors such as Jun and NF κB inhibit their
DNA binding ability (Klatt et al., 1999; Pineda-Molina et al., 2001). Glutathionylation could be a way to protect the protein from more irreversible forms of oxidation that
14
would lead to permanent inactivation. S-nitrosylation is the attachment of an NO
moiety to the cysteine residue of a protein (Sun et al., 2006). S-nitrosylation has also
been proposed as a way to protect proteins from further oxidation during oxidative
stress. The activity of proteins may either be enhanced by S-nitrosylation (eg. p21 ras ,
Akt1, and thioredoxin) (Lander et al., 1997; Haendeler et al., 2002; Lu et al., 2005) or
suppressed by it (eg. caspase and methioine adenosyl) (Mannick et al., 1999; Pérez-
Mato et al., 1999). The cysteine residue of some protein tyrosine phosphatases (PTPs)
such as PTEN, Cdc25 and low molecular weight-PTPs can form disulphide bond with
a nearby cysteine when oxidized by H 2O2 (Caselli et al., 1998; Savitsky and Finkel,
2002; Kwon et al., 2004). Severe oxidative stress can result in more damaging modifications, such as the formation of sulfenic acid, sulfinic acid, and sulfonic acid
(Poole et al., 2004). Besides cysteine residue, tyrosine residue is also a target for redox modification. The oxidation of tyrosine to nitrotyrosine by reactive nitrogen species (RNS) causes the protein to lose its phosphorylation site (Trachootham et al.,
2008). Kinases such as PI3K, PKC and p38 MAPK are targets of tyrosine nitration, leading to their inactivation (Hellberg et al., 1998; Knapp et al., 2001; Webster et al.,
2006).
Another more severe form of modification is the carbonylation of proteins which can occur either directly through the oxidation of amino acid side chains (lysine, arginine, threonine and proline) or indirectly through amino acid interacting with oxidation products of lipids and sugars (Stadtman and Berlett, 1991; Schneider et al., 2001).
Carbonylation of proteins is a form of irreversible modification which often leads to loss of protein function (Dalle-Donne et al., 2006). Examples of such proteins are
ANT, Hsp and Bcl2 (England and Cotter, 2005). As carbonylated protein is easily detectable, it is frequently used as an indicator of the oxidative damage to the cell.
15
While moderately carbonylated proteins are degraded via the proteosomal pathway, heavily carbonylated proteins form aggregates that can inhibit proteosome function
(Dalle-Donne et al., 2006).
1.2.4 Regulation of protein turnover
The half-life of the protein affects the duration it can execute its function in the cell.
The more stable a protein is, the longer its effects are in the cell. The rate of protein turnover can also be subjected to redox regulation. The Rac1 protein is involved in the production of superoxide via NADPH oxidase. One study found that as a way to control ROS production in the cell, Rac-1-induced superoxide production activates a feedback loop where Rac-1 protein turnover is enhanced via increased degradation by the proteosome (Kovacic et al., 2001). Superoxide has also been shown to promote apoptosis in cells by increasing the turnover of anti-apoptotic protein Bcl2 through ubiquitin-proteosomal pathway (Azad et al., 2008). HIF is a transcription factor that controls the transcription of a number of genes during hypoxia (Brahimi-Horn and
Pouysségur, 2007). Under normoxia, HIF α, the subset of HIF protein is continuously synthesized but is also rapidly degraded via the ubiquitin, proteosomal system. ROS generated at the mitochondria have been shown to stabilize HIF α under hypoxic conditions by decreasing protein turnover (Chandel et al., 2000). In some cell types, growth factors have also been shown to stabilize HIF α during normoxia through the production of ROS (Richard et al., 2000).
16
1.3 ESTROGEN RECEPTOR
1.3.1 Estrogen and estrogen receptors: Introduction
Estrogens are steroid hormones produced primarily in the ovaries of pre-menopausal,
non-pregnant women. They are converted from testosterone to estradiol (E2) by an
enzyme aromatase, a member of the cytochrome P450 superfamily. Although E2 is
the primary estrogen in the body, it can be broken down into its less potent
metabolites estrone and estriol. In post-menopausal women and men, estrogen is
synthesized in mesenchymal cells of adipose tissue, osteoblasts and chondrocytes of
bone, the vascular endothelium and aortic smooth muscle cells, and numerous sites in the brain (Simpson and Davis, 2001). Other than playing important roles in the monthly estrus cycle of pre-menopausal women, estrogens are essential for various physiological processes in tissues and organs such as bone, urinary tract, brain, uterus, prostate and breast.
Estrogen exerts its effects primarily via binding to estrogen receptor (ER) which consist of two members, ER α and ER β. The DNA coding for ER α was cloned in 1985
(Walter et al., 1985). In the following decade, this receptor was thought to be the only
receptor mediating the physiological effect of estrogens. It was only in 1996 that a
second and novel estrogen receptor, ER β was discovered in rat prostate (Kuiper et al.,
1996). ER β was subsequently identified in human and mouse tissues (Mosselman et
al., 1996; Tremblay et al., 1997). Human ER α and ER β contains 595 and 530 amino
acids respectively and are coded by different genes (Green et al., 1986; Ogawa et al.,
1998). ER α gene is found on chromosome 6q25.1 while ER β gene is found on
chromosome 14q23.2. ER α is the major form of estrogen receptor in uterus, liver, adipose, skeletal muscle, pituitary gland and hypothalamus while ER β is the major
17 form in ovary, testis, prostate, lung and other parts of the brain. ER α and ER β colocalize in mammary glands, bone, uterus, central nervous system and cardiovascular system (Nilsson et al., 2001).
Estrogen receptor belongs to a class of nuclear receptors which are transcription factors targeted by small lipid-soluble molecules such as steroid hormones, thyroid hormones, retinoids, vitamin D3 and bile acids (Mangelsdorf et al., 1995). Like all nuclear receptors, ER can be divided into five functional domains. The A/B domain of the estrogen receptor, which is also known as activation function (AF-1) activates gene transcription of target genes through the recruitment of co-regulators
(Tzukerman et al., 1994). C domain is the DNA-binding domain (DBD) which is rich in basic amino acids. It recognizes specific DNA sequence known as estrogen response element (ERE) and binds to ERE via its two zinc-stabilized DNA-binding fingers (Kumar et al., 1987). D domain, which also harbors the nuclear localization signal, acts as a hinge between C and E domains. E domain is a hydrophobic domain that binds to ligands. Embedded within E domain is the dimerization region and a second transcriptional activation function (AF-2). F domain exerts a complex modulatory role on activity of the receptor and interaction with other proteins
(Montano et al., 1995; Koide et al., 2007). Human F domain contains a PEST region which is enriched in proline (P), glutamic acid (E), serine (S) and threonine (T).
PEST sequence had been suggested to be a signal for rapid degradation of proteins
(Rogers et al., 1986; Hirai et al., 1991). Thus, the F domain might also play a role in
ER turnover.
Structurally, the two ER receptors are very similar to each other particularly in the
DNA-binding domain (97% homology) and the c-terminal ligand-binding domain (47% homology). The major difference is at the A/B and F domains with only 18%
18 homology. This is an indication that the two receptors most likely interact with different set of co-factors upon activation (see Figure V for structure of ER α and
ER β). Despite considerable variation in sequence of the ligand-binding domain, both
ER α and ER β bind to estrogen with similar high affinity with dissociation constant of
0.1 nM (Kuiper et al., 1998). The major difference in the ligand binding of these two
receptors lies in having different affinities for various compounds. In addition, one
compound could also evoke differential transcriptional response in the two receptors.
For example, tamoxifen it is a mixed agonist and antagonist for ER α but is an antagonist for ER β (Watanabe et al., 1997; Barkhem et al., 1998). Phytoestrogens
such as genistein, coumestrol and zearalenon, and environmental estrogenic compounds for instance nonylphenol, bisphenol A, o, p'-DDT, and 2',4',6'-trichloro-4-
biphenylol have also been shown to compete with E2 in their binding to ER, abide
with lower affinities (Kuiper et al., 1998).
Figure V: Domain structure representation of human ER α and ER β isoforms . Domains (labeled A–F), amino acid sequence numbering, AF-1 and AF-2, and percentage homology between the two isoforms in different regions, including the DBD and LBD, are shown. Figure adapted from (Kong et al., 2003)
19
1.3.2 Estrogen receptor and genomic signaling
There are several pools of ER that exist in the cell. They can be found in the cytosol,
nucleus, plasma membrane and mitochondria (Nadal et al., 2001; Ivanova et al., 2009;
Jazbutyte et al., 2009). In the non-ligand bound state, ER is found to complex with
heat shock proteins, immunophiline and p23. (Pettersson and Gustafsson, 2001). For
genomic signaling, ER dissociates from chaperone proteins and dimerizes upon
binding to its ligand. The ligand-receptor complex moves into the nucleus and binds
to ERE of target genes. The ERE consensus sequence contains a palindromic
sequence which is an inverted repeat 5'GGTCAnnnTGACC-3' where n represents any
nucleotide. It should be noted that many EREs contain variations from the consensus
sequence (Klein-Hitpass et al., 1986; Klinge et al., 1997; Cheskis et al., 2007). The
binding of different ligands cause ER to assume different conformations. The LBD is
formed by a 12 α-helices (Helices 1-12) with a hydrophobic ligand-binding pocket.
The AF-2 in the LBD of ER is composed of amino acids in helix 3, 4, 5 and 12. The position of helix 12 changes when ER binds to ligands and therefore is the key in discriminating between agonists and antagonists of ER. For instance, when E2 binds to ER, helix 12 is positioned over the ligand pocket and forms the surface for recruitment and interaction with cofactors. In contrast, when ER binds to an antagonist, such as tamoxifen, helix 12 occupies the hydrophobic groove formed by helix 3, 4 and 5. This position prevents the recruitment of co-activators but encourages the recruitment of co-repressors instead. (Dickson and Stancel, 2000).
Besides binding directly to DNA and activating gene transcription, ER could activate gene transcription indirectly via interacting with other transcription factors. The interaction between ER α and the c-rel subunit of NF κB prevents NFκB from binding to interleukin-6 (IL-6) promoter and thus inhibiting the expression of IL-6 (Galien and
20
Garcia, 1997). ER α physically interacts with SP1 to activate retinoic acid receptor α-1
(RAR-1) gene transcription through the binding of ER-SP1 complex on SP1 site on
RAR-1 promoter (Sun et al., 1998; Zou et al., 1999). ER also interacts with the fos/jun
transcription complex on AP1 sites to modulate gene transcription depending on the
ligand and ER subtypes (Webb et al., 1995; Paech et al., 1997; Webb et al., 1999). For
instance, in the presence of ER α, typical agonists such as E2 and antagonist tamoxifen function as agonists in the AP1 pathway. Raloxifene acts as a partial agonist. In the presence of ER β, tamoxifen and raloxifene behave like agonist while E2 acts as an
antagonist, inhibiting the actions of tamoxifen and raloxifene (Paech et al., 1997).
ER can also be activated in a ligand-independent manner through phosphorylation of
the various serine and tyrosine residues in the AF-1 and AF-2 domains. Downstream
signaling pathway of peptide growth factor, PKA-activating agents, neurotransmitters,
and cyclins can mediate ligand-independent transcription of ER through
phosphorylation (Cenni and Picard, 1999). The phosphorylation regulates the
dimerization of the ER and recruitment of various co-activators to the AF-1 and AF-2
domains depending on the phosphorylation sites.
1.3.3 Membrane and cytoplasmic ER signaling
Rapid estrogen signaling from the plasma membrane was first identified forty years
ago (Szego and Davis, 1967; Pietras and Szego, 1977). Other than binding to nucleus
ER and activating transcriptional activities, estrogens can modulate intracellular
signaling through secondary messengers. Approximately 5-10% of the total ER is
found at the plasma membrane (Levin, 2009). This include both ER α and ER β,
however, the localization of the different subtypes depends on cell type. The nature of
21
the plasma membrane located ER is found to be the same as the classical nuclear ER α
and ER β. Endothelial cells from mice that are knockout of the classical ER α and ER β
also lost of all estrogen binding at the plasma membrane (Pedram et al., 2006).
Membrane proteins that bind to estrogen-conjugated beads in affinity chromatography
were identified by mass spectrometry to be the classical ER α, and were identical to
the nuclear ER α. (Pedram et al., 2006).
Many signaling cascades have been described for membrane ER and estrogen. These include activation of MAPK (Endoh et al., 1997; Song et al., 2002; Mize et al., 2003) and PI3K/Akt pathway (Haynes et al., 2000; Haynes et al., 2003), induction of ion channel fluxes (Rosenfeld et al., 2000; Coiret et al., 2005), GPCR-mediated second messenger generation of cAMP and calcium (Stefano et al., 2000; Wyckoff et al.,
2001; Wade and Dorsa, 2003), as well as stimulation of growth factor receptors
(Razandi et al., 2003). When E2 binds to membrane ER, the initiation of signal transduction triggers downstream signaling cascades that contribute to important functions such as cell growth and survival, migration, and new blood vessel formation
(Levin, 2002).
22
1.3.4 Regulation of Estrogen Receptors
1.3.4.1 Phosphorylation
ER α has been shown to be phosphorylated at multiple sites upon either binding to E2 or through the actions of second messenger signaling pathways (Washburn et al.,
1991; Denton et al., 1992; Joel et al., 1995). Phosphorylation sites are located at both amino- and carboxy-terminals of ER α. There are four serine located at the AF-1 of
ER α, (Ser104, Ser106, Ser118 and Ser167) that can be phosphorylated. The
phosphorylation of these serine residues in the AF-1 domain influences the
recruitment of co-activators (Lavinsky et al., 1998; Endoh et al., 1999; Tremblay et al.,
1999). Ser118 is the major site phosphorylated in response to E2; although Ser104
and Ser106 are also phosphorylated to a lesser extent (Joel et al., 1995). Upon
activation of MAPK by EGF, Ser118 is preferentially phosphorylated in ER α but
Ser104 and Ser106 remain unphosphorylated (Joel et al., 1998). TFIIH cyclin-
dependent kinase has also been shown to phosphorylate Ser118 (Chen et al., 2000).
On the other hand, cyclin-dependent kinase 2 (Cdk2) complex can phosphorylate
Ser104 and Ser106 but not Ser118 (Rogatsky et al., 1999). The downstream kinase of
MAPK, p90 ribosomal S6 kinase (Rsk) has been shown to phosphorylate Ser167 (Joel
et al., 1998; Frödin and Gammeltoft, 1999). Ser167 can also be phosphorylated by
casein kinase II and Akt (Arnold et al., 1995; Martin et al., 2000; Campbell et al.,
2001). In the presence of E2, ER α is phosphorylated by protein kinase A (PKA) on
Ser236 within the DNA binding domain which facilitates the dimerization of the receptor (Campbell et al., 2001). Two Scr tyrosine kinases, p60c-src and p56lck are shown to phosphorylate ER α at tyrosine-537 located in the LBD (Arnold et al., 1995).
The phosphorylation of tyrosine is however not required for the binding of E2 or
23
activation of transcription. Instead, it is important in the conformational change of the
LBD that can affect the ligand binding kinetics and the stability of ligand-bound
receptor (Yudt et al., 1999). Phosphorylation of ERβ is less well studied. One study suggested that Ser60 of mouse ER β is phosphorylated in response to activation of
MAPK pathway (Joel et al., 1995).
1.3.4.2 Glycosylation
A large number of proteins in the cells such as transcription factors, cytoskeletal proteins, nuclear pore proteins, oncogene product, and tumor suppressors can be modified by O-linked N-acetylglucosamine ( O-GlcNAc) (Hart, 1997; Haltiwanger et
al., 1997). Glycosylation, like phosphorylation, is a dynamic process with O-GlcNAc
displaying features essential for a role in signal transduction in the cell. For instance,
the attachment and removal of O-GlcNAc from proteins are tightly regulated processes and the half-life of O-GlcNAc moiety has been shown to be much shorter than that of the modified protein backbone (Roquemore et al., 1996). ER α from murine, bovine, and human sources is shown to be modified by O-GlcNAc and that a major O-GlcNAc site (Thr575) occurs within a PEST sequence of the F domain
(Jiang and Hart, 1997). Additional glycosylation sites have been found where are
Ser10 and Thr50 in the AF-1 domain near the N-terminus of murine ER α (Cheng and
Hart, 2000). O-GlcNAc glycosylation has also been found for murine ER β at Ser16 which is in the proximity of the AF-1 domain and within a region with a high PEST score (Cheng et al., 2000). The O-GlcNAc glycosylation may play a role in ER turnover and modulate the transcriptional activity of ER protein.
24
1.3.4.3 Acetylation
Modification via acetylation provides another layer of regulation of the estrogen
receptor. ER α can interact with co-activators that modify histone acetyl groups. It had been shown that ER α can interact with p300 which acetylates the receptor directly at the lysine residues Lys266 and Lys268. Acetylation of ER α by p300 enhances the
DNA binding activity of the receptor, leading to increase in ligand-dependent transcriptional activity (Kim et al., 2006). However, in another study, the acetylation of ER α within the hinge/ligand binding domain could attenuate estrogen-dependent
transregulation (Wang et al., 2001). Acetylation appears to act as a modulation of
ligand-dependent gene regulatory activity of ER α.
1.3.4.4 Ubiquitination
The ubiquitination-proteosomal pathway is a major system for the degradation of
proteins with short half-life in eukaryotic cells (Haas and Siepmann, 1997; Pickart,
1997). The covalent attachment of ubiquitin proteins, to lysine residues of targeted
proteins to form polyubiquitin is necessary for the subsequent proteosomal
degradation. Ubiquitination involves three classes of enzymes: E1 ubiquitin-activating
enzyme (UBA), E2 ubiquititin-conjugating enzyme (UBCs) and E3 ubiquiting-protein
ligases. Other than targeting proteins for degradation, ubiquitination also serves to
direct cellular localization of proteins (Chen et al., 1996; Kim and Maniatis, 1996).
Ubiquitination of ER has been observed in rat uterus (Pakdel et al., 1993), and the
stability of ER is affected by ligand binding. In the absence of E2, the half-life of ER
is about 5 days. Upon binding to E2, the half-life of ER shortens to 3-4 hours (Pakdel
et al., 1993; Nirmala and Thampan, 1995). Ubiquitin-proteosomal degradation of
ligand bound ER allows the E2-ER signal to be rapidly turned off once its desired
25
effects on the cells is achieved. Misfolded ER has also been shown to undergo
ubiquitination and proteosomal degradation to safeguard the quality of ER in the cells
(Tateishi et al., 2004).
1.3.4.5 Oxidation
Many cellular processes such as protein phosphorylation and binding of transcription
factors to promoters are driven by oxidation and antioxidant homeostasis (Sen and
Packer, 1996). Redox regulation of transcription factors had been identified for AP-1,
NF κB, SP1 and glucocorticoid receptor. The transcriptional activity of ER is influenced by its redox state which could be regulated by cellular antioxidant thioredoxin (Trx) (Hayashi et al., 1997). ER dependent gene pS2 and ERE-driven acetyltransferase (CAT) activity were found to be down-regulated in presence of
H2O2 and were recovered by the increasing intracellular Trx (Hayashi et al., 1997).
Cancer cells often have high levels of ROS and this altered redox homeostasis have an impact on the transcription activity of ER. ER extracted from a third of ER positive primary breast tumors are unable to bind to the consensus sequence of ERE (Liang et al., 1998). Treating the tumor extract with reducing agent could partially restore the
ER binding suggesting that these ERs were in the oxidized form in the primary tumors.
The DBD of ER contains two (Cys)(4)-liganded zinc finger motifs which function to stabilize the DNA-binding recognition helix and flexible helix-supported dimerization loop. The oxidation of the zinc finger in the DBD of ER prevents dimerization and hence DNA binding leading to down-regulation of ER downstream genes (Whittal et al., 2000; Atsriku et al., 2007). The redox state in cells may be an important mechanism for post-transcriptional regulation of ER function, thereby playing a critical role in regulating the transcriptional activity of ER.
26
1.3.5 Estrogen and estrogen receptors in human breast cancer
Breast cancer or malignant breast neoplasm is cancer originating from breast tissue, most commonly beginning either in the milk ducts (ductal carcinoma) or in the milk producing glands (lobular carcinoma). Breast cancer is the most common cancer found in women from industrialized countries and is the second biggest killer after lung cancer (Jemal et al., 2009). The importance of estrogen in regulating breast cancer growth was demonstrated more than a hundred years ago through the inhibitory effects of ovariectomy on patients with metastatic breast cancer (Beatson,
1896). Estrogen has been suggested to be important in breast cancer through non- receptor- and receptor-mediated mechanisms. The non-receptor-mediated mechanism includes a cytochrome P450 (CYP)-mediated metabolic activation, which elicits direct genotoxic effects by increasing mutation rate and inducing aneuploidy (Russo et al., 2003). Estrogen is metabolized in body by several cytochrome P-450 enzymes in phase I metabolism. Estrogen can be metabolized in the body to form catechol estrogen. Normally, catechol estrogen is detoxified by phase II enzyme catehol O- methyltransferase (COMT). Insufficient metabolism by COMT can generate estrogen
3,4-qinone, a electrophilic reactive intermediate that can form unstable adducts with adenine and guanine in the DNA, leading to depurination and mutations (Chakravarti et al., 2001; Devanesan et al., 2001; Yue et al., 2003). Mismatch repair or mis- replication of depurinated sites could lead to carcinogenesis. The reduction of estrogen quinones involves redox cycling with ROS generation that could further damage cellular lipids and DNA leading to carcinogenesis (Nutter et al., 1994; Seacat et al., 1997; Lavigne et al., 2001).
Deregulation of expression and activity of estrogen receptor contribute to the development and progression of breast cancer (Nilsson et al., 2001). High levels of
27
ER promote growth of estrogen-dependent tumors by increasing ER-dependent gene
transcription and non genomic functions of ER that promotes cell proliferation.
Although ER β is important in estrogen functioning, ER α appeared to play a more
predominant role in mammary tissues. While ER α is found only in 6 to 10% of
normal breast epithelial cells; 60% of primary breast cancer are ER α-positive and
their growth is often stimulated by estrogen (Dickson and Lippman, 1988; Jacquemier
et al., 1990). The over-expression of ER α in breast cancer is suggested to be due to amplification of the ER α gene. A tissue microarray analysis of more than 2,000
clinical breast cancer samples showed ER α gene amplification in 20.6% of breast cancers (Holst et al., 2007). ER α gene amplication was also found in benign and
precancerous breast disease, suggesting that amplification of ER α gene is a common and early genetic alteration in proliferative breast disease and breast cancer. Gene amplification alone, however, cannot explain all cases involving high ER α protein
levels. Other possible mechanisms include altered regulation of ER α transcription,
mRNA stability or protein turnover (Fowler and Alarid, 2007).
ER α status is important in the clinical setting as a prognostic tool as well as a therapeutic target. Currently, biopsy samples from breast cancer patients are routinely screened for the presence of estrogen receptor (ER), progesterone receptor (PR) and oncoprotein HER2. ER α status is the most important biomarker in breast cancer, because it provides clinicians with information on whether the patient will respond to endocrine treatment (Weigel and Dowsett, 2010). ER α-positive cells generally have a
better prognosis. ER α-positive tumor uses E2 as its main growth stimulus thereby
making ER α a direct target for endocrine therapy.
28
1.4 THE SODIUM HYDROGEN EXCHANGER 1
1.4.1 pH regulation in the cells
Intracellular pH (pHi) is a tightly regulated parameter in the cell as many cellular
processes function within a narrow range of pH. Large fluctuations of pHi can affect
the normal structure and functioning of proteins, gene expression, ion channels and
other processes. The normal cytosolic pHi in the cells is in the range of 6.99-7.2
(Gillies et al., 2002). Cells constantly produce H + ion through its metabolic activity or leakage from acidic compartments in the cells. Several transport proteins are located on the plasma membrane to transport the acids or bases across the cell. Two
+ - mechanisms are employed: either H ion is transported out of the cell or HCO 3 is imported into the cell to neutralize the H + ions in the cytosol. There are three main
families of exchangers that are present at the plasma membrane of cells. The first is
Na +/H + exchanger (NHE) which extrude H + ion out of the cell and import a Na + ion in.
- The second family of membrane transporters imports HCO 3 into the cell. They
+ - - - + - include Na / HCO 3 cotransporter (NBC), Cl / HCO 3 exchanger (AE), Na driven Cl /
- + - - HCO 3 exchanger (NDAE), and Na independent Cl / HCO 3 exchanger. The third family of membrane transporters is the H +-ATPase (Putney et al., 2002). With the
exception of H +-ATPase, none of the membrane transporters has a direct requirement
for ATP. Of all the membrane transporters, the NHE family plays a key role in
maintaining the pHi of cells upon acidification in virtually all cell types (Malo and
Fliegel, 2006). The first member, NHE1 is found to be expressed ubiquitously and is
suggested to be the main pH regulator in transformed cells (Harguindey et al., 2005).
29
1.4.2 Mammalian Na +/H + exchanger
In 1982, Pouyssegur and his group identified a “Na +/H + exchanger system (NHE)” that played an important role in the regulation of pHi (Pouyssgur et al., 1982). NHE is a family of glycoproteins that exchanges a H+ ion out of the cell for a Na + ion across
the membrane in a 1:1 stoichiometry. In mammalian cells, the NHE family consists of
9 members (NHE1 – NHE9). The gene coding for first member, NHE1, was cloned in
1989 by Sardet et al and was found to be expressed ubiquitously on the plasma
membrane of virtually all cell types. NHE1 is referred to as the “housekeeping”
isoform while the other isoforms have more restricted localization (Sardet et al., 1989;
Malo and Fliegel, 2006). The specific localization of NHE1 on the plasma membrane
varies depending on cell type. In fibroblast, NHE1 is mostly found on the
lamellipodia where it is involved in migration and anchoring (Denker et al., 2000;
Denker and Barber, 2002). In epithelial cells, NHE1 is distributed to the basolateral
membrane (Biemesderfer et al., 1992; Orlowski and Grinstein, 2004). In the
myocardial tissue, NHE1 is found in the intercalated discs and along the transverse
tubular system (Petrecca et al., 1999). The rest of the NHE members are more
restricted in their tissue and subcellular distribution. NHE2 – NHE5 are located at the
plasma membrane but they are more tissue specific. NHE2 is an apical membrane
protein that shares 42% sequence identity with NHE1 (Wang et al., 1993). NHE2 is
found in the epithelial tissue of the intestinal tract (mainly in the jejunum, ileum and
colon), kidney (cortical thick ascending limbs, distal convoluted tubules, connecting
tubules and macula densa cells), skeletal muscle and testis (Malakooti et al., 1999).
NHE3 and NHE4 have 39% and 42% identity with NHE1 respectively (Orlowski et
al., 1992). Both NHE3 and NHE4 are highly expressed in the kidney and
gastrointestinal tract. In kidney, NHE3 is found in the proximal tubule and thick
30
ascending limb (Amemiya et al., 1995) whereas NHE4 has a more basolateral
epithelial distribution in the inner medullar (Bookstein et al., 1994). In the
gastrointestinal tract, NHE3 appears predominantly in the intestines while NHE4 is
found in the stomach. NHE5 which shares 39% sequence identity with NHE1 is found
exclusively in the brain. NHE6 to NHE9 are found in the membrane of intracellular
organelles where they are involved in the maintenance pH in these compartments.
NHE6 is expressed in the early recycling endosomes, NHE7 in the trans-Golgi
network, NHE8 in the mid- to trans-Golgi, and NHE9 in the late recycling endosomes
(Nakamura et al., 2005).
1.4.3 NHE1: Basic structure
NHE1 isoform is the best characterized amongst the NHE family members. NHE1
protein is made up of 815 amino acids. The first 500 amino acids makes the 12 α- helices transmembrane (TM) spanning segment while the remaining residues constitute the cytoplasmic tail which is the intracellular regulatory domain (Sardet et al., 1989). Both the N- and C- terminals of NHE1 are found in the cytosol. Two forms of NHE1 are consistently observed: a plasma membrane protein of 110 kDa that contains N- and O-linked oligosaccharide, and an 85 kDa protein that is found intracellularly which contains only the N-linked high-mannose oligosaccharides
(Malo and Fliegel, 2006). The TM domain of NHE1 is responsible for the exchange of intracellular H + for extracellular Na + in a 1:1 stoichiometry. (See Figure VI for the topological model of NHE1). Several TM domains of NHE1 are important for its function. The TM IV is critical for the function of NHE1; it is involved in the affinity of NHE1 for Na + and overall NHE1 activity. TM VI and VII are crucial for NHE1
31
activity while TM IX is important in conferring sensitivity to antagonists. TM XI, on
the other hand, is essential in targeting NHE1 to the cell surface (Fliegel, 2005).
The recombinant C-terminal of human NHE1 have been expressed and purified from
Escherichia coli . It is found to be made of 35% α-helix, 16% β-turn, and 49% random
coils. Using circular dichroism to analyze the various C-terminal fragments, the
membrane proximal region was predicted to be more structured and compact. The
more distal region has a more flexible and unstructured feature, possibly to allow for
the association with various kinases and regulatory proteins (Gebreselassie et al.,
1998; Li et al., 2003).
Figure VI: Topology of NHE1 isoform and its regulatory elements. Extracellular loops 1- 6 (EL1-EL6), intracellular loops 1-5 (IL1-IL5), amino acids glutamic acid (E) and aspartic acid (D). Regulatory elements and their approximate location of binding: calmodulin (CaM), calcineurin homologus protein (CHP), ezrin/radixin/moesin proteins (ERM), carbonic anhydrase isoform II (CAII), phosphatidylinositol bisphosphate (PIP2). Putative protein kinase that phosphorylate the Na +/ H+ exchanger are indicated: ERK 1/2, p90rsk, NIK, p160ROCK, p38. Figure adapted from (Malo and Fliegel, 2006)
32
1.4.4 Physiological and pathological roles of NHE1
1.4.4.1 pHi and volume regulation
The major function of NHE1 in the cell is the regulation of pHi and cell volume.
NHE1 activity is activated by the decrease in intracellular pH. When the pHi of the
cell reach a certain threshold, NHE1 will be activated to extrude H + ion out of the cell for a Na + ion into the cell, leading to intracellular alkalinization (Malo and Fliegel,
2006). This process is dependent on extracellular Na +, and the cell uses the energy
gradient of Na + to catalyze the electro-neutral exchange. NHE1 also regulates cell volume during osmotic shrinkage. During hypertonic stress, NHE1 is activated; bringing more Na + into the cell resulting in intracellular alkalinization. The increase in Na + in cells will draw more water into the cells due to obligatory osmosis
(Bianchini et al., 1995).
1.4.4.2 Cell proliferation and differentiation
It has been long established that the activation of NHE1 and increase in pHi are early response to mitogens (Pouysségur et al., 1984). NHE1 has a permissive, but not an obligatory role in cell proliferation. Although NHE1 deficient cells were able to proliferate in the presence of serum, the rate of proliferation was markedly increased in cells with NHE1 expression (Kapus et al., 1994). Inhibition of NHE1 activity with pharmacological inhibitors such as EIPA or HOE694 also attenuated growth factor induced cell proliferation (Delvaux et al., 1990; Horvat et al., 1992; Benedetti et al.,
2001). Although NHE1 has been associated with proliferation, the mechanism by which NHE1 exerts this function is less known. It has been shown that NHE1 and pHi
33 regulates the G2/M entry and transition. Cells that have a defective NHE1 which prevents ion translocation have impaired G2/M transition and delayed entry into S phase (Putney and Barber, 2003).
NHE1 also play a role in cellular differentiation. The inhibition of NHE1 with pharmacological inhibitor prevented the differentiation of P19 embryonal carcinoma
(Wang et al., 1997). NHE1 has recently been shown to be important in the differentiation of embryonic stem cells into cardiomyocytes where inhibition of
NHE1 prevented such differentiation (Li et al., 2009).
1.4.4.3 Cell migration
NHE1 is important in cell shape and cell migration by acting as a structural anchor for organization of cytoskeleton. In some cell types NHE1 is located at the lamellipodia where its C-terminal binds ezrin, radixin and moesin (ERM) protein to interact with actin filaments (Fliegel, 2005). Actin filaments play a pivotal role in determining the shape and motility of a cell and also associate with the dynamic extensions such as lamellipodia. Cells that express NHE1 which is defective in the C-terminal and cannot interact with ERM proteins have reduced actin organization, irregular shape, and reduced cell migration (Tominaga and Barber, 1998; Denker et al., 2000; Denker and
Barber, 2002).
34
1.4.4.4 NHE1 in heart disease
NHE1 is the main isoform found in myocardiocytes. It is responsible for half of total
proton efflux and plays a critical role in maintaining pHi homeostasis and contractility
(Fliegel et al., 1991; Grace et al., 1993). Although it plays an important role in cardiac
cell functioning, NHE1 has also been implicated in heart diseases. During ischemia,
where cardiac cells are cut off from oxygen, anaerobic glycolysis occurs, resulting in
the accumulation of H + in the cells. In the subsequent reperfusion, NHE1 is activated to pump out H + in exchange for Na + into the cells (Avkiran, 2001; Allen and Xiao,
2003). This large influx of Na + activates Na +/Ca 2+ exchanger and drives the exchange
Na + out of the cell for Ca 2+ into the cell. The built up of Ca 2+ in the cells is damaging
as it triggers various pathways leading to cell death. Inhibition of NHE1 using
pharmacological drugs such as cariporide, amiloride and EMD 85131 have been
shown to have a cardioprotective effect in animal models (Karmazyn, 1988; Scholz et
al., 1995; Gumina et al., 1998).
1.4.4.5 NHE1 and Cancer
NHE1 has also been found to play a role in the development and progression of
cancer. One hallmark of transformed cells is intracellular alkalinization where tumor
cells have a pHi of 7.12-7.65 as compared to 6.99 -7.2 for normal tissue (Gillies et al.,
2002). This intracellular alkalinization occurs early in the cancer formation. NHE1
was shown to be responsible for the increase in pHi as blocking NHE1 with amiloride
could reverse both pH change and transformed phenotype (Rich et al., 2000; McLean
et al., 2000; Reshkin et al., 2000). The activation of NHE1 in tumor cells is a result of
higher affinity of NHE1 for H + (Reshkin et al., 2000). This causes the set point of
NHE1 in the cells to be higher than the resting pHi leading to a constitutively active
35
NHE1 in tumor cells. The development of intracellular alkalinization is accompanied by increased DNA synthesis (Hagag et al., 1987; Ober and Pardee, 1987), cell cycle progression (Doppler et al., 1987; Siczkowski et al., 1994), substrate- and serum- independent growth (Reshkin et al., 2000).
In addition to establishing an alkaline intracellular environment, NHE1 has also been implicated in cancer cell migration. The acidic extracellular environment of tumor increases metastatic ability of cancer cells by promoting neoangiogenesis (Kurschat and Mauch, 2000), anchorage-independent growth (Gillies et al., 2002), genetic stability (Gillies et al., 2002) and invasion.
36
1.4.5 Regulation of NHE1
1.4.5.1 Phosphorylation
The cytosolic domain of NHE1 regulates the TM domain by being the target region of
kinases and binding site of regulatory proteins. Phosphorylation of the residues in the
cytoplasmic domain stimulates the activity of NHE1, causing it to be more active at a
higher pH. Kinases that have been shown to phosphorylate NHE1 include
extracellular-signal-regulated kinase 1/2 (ERK 1/2) (Denker et al., 2000; Denker and
Barber, 2002), p90 ribosomal S6 kinase (p90rsk) (Takahashi et al., 1999) Rho
associated kinase p160ROCK (Tominaga et al., 1998), Nck-interacting kinase (NIK)
(Yan et al., 2001) and Ca 2+ /calmodulin-dependent kinase II (CaMKII) (Fliegel et al.,
1992). The p38 kinase of the MAPK family can also phosphorylate NHE1 directly
(Khaled et al., 2001). Protein kinase C and D regulate the exchanger via mechanism other than direct phosphorylation (Haworth et al., 1999; Sauvage et al., 2000). Protein phosphatase 1 (PP1) and protein phosphatase 2A (PP2A) have been shown to dephosphorylate NHE1 (Misik et al., 2005; Snabaitis et al., 2006).
1.4.5.2 Regulatory Proteins
The cytoplasmic tail of NHE1 interacts with a variety of signaling molecules. Three calcium binding proteins have been shown to interact with NHE1: calmodulin (CaM) and calcineurin homologous protein (CHP) stimulate NHE1 activity while tescalcin acts to inhibit NHE1 activity. Other molecules that can bind to the cytoplasmic tail include carbonic anhydrase II (CAII), adaptor protein 14-3-3, PIP 2, HSP70 and ERM.
CaM binds to the cytoplasmic tail at two sites: a high affinity site located at amino acids 636-656 and a low affinity site at amino acids 657-700 (Bertrand et al., 1994).
37
The high affinity site function as auto-inhibitory domain for NHE1 and binding of
CaM/Ca 2+ to this site can abolish this inhibition (Wakabayashi et al., 1994). CHP binds to the amino acids located at 510-530. The association between CHP and NHE1 is crucial for NHE1 functioning, as NHE1 or CHP mutant that prevents their association drastically reduces NHE1 activity (Pang et al., 2004). Tescalcin is a protein structurally homologous to CHP, where it interacts with the final 180 amino acids of cytoplasmic tail of NHE1 to inhibit its activity (Mailänder et al., 2001; Li et al., 2003). CAII binds to amino acids 790-802 of NHE1 to enhance the activity of
NHE1 (Li et al., 2002; Li et al., 2006). Phosphorylation within amino acids 634-789 region increases the association between NHE1 and CAII, resulting in a more efficient proton removal from the cell. The binding of 14-3-3 to NHE1 tail is also dependent on phosphorylation of NHE1. The protein 14-3-3 prevents the dephosphorylation of Ser703, thereby stabilizing the active conformation of NHE1
(Lehoux et al., 2001). PIP 2 binds to NHE1 at two PIP 2 binding motifs at amino acids
513-520 and 556-564 and is required for optimum activity of NHE1 (Aharonovitz et al., 2000). HSP70 binds to the cytoplasmic domain of NHE1 to play a role in the folding and processing of the exchanger (Silva et al., 1995). The binding of ERM proteins to the cytoplasmic tail of NHE1 allows NHE1 to interact with actin filaments.
This is important for the localization of NHE1 to the lamellipodia and for NHE1- dependent organization of cortical actin filaments (Vaheri et al., 1997).
38
1.4.6 Regulation of NHE1 gene transcription
The human NHE1 gene is located on chromosome 1 p35-p36.1 and is approximately
70 kilo-bases (kb). The coding region is divided into 12 exons and 11 introns. The promoter region of human NHE1 has been described. The promoter region is 1377 bases and is made of a TATA box, four GC boxes, two CAAT boxes, three AP-1 sites, and a cyclic AMP response element (Miller et al., 1991). The regulation of NHE1 gene transcription in mouse has been widely studied. The analysis of mouse NHE1 promoter revealed several important regions that are binding sites for transcription factors of NHE1. Mitogens, in addition to regulating NHE1 activity also regulate
NHE1 promoter activity (Besson et al., 1998). Addition of serum to serum-starved cells was able to activate a 1.1kb proximal fragment of the mouse NHE1 promoter.
Other growth factors such as insulin, thrombin and EGF also increase the activity of
NHE1 promoter (Besson et al., 1998). Promoter analysis and gel mobility shift assays showed that a region between 0.9 and 1.1 Kb from the start site of the mouse NHE1 promoter is involved in mitogen stimulation of NHE1 transcription (Besson et al.,
1998). The chicken ovalbumin upstream promoter-transcription factors (COUP-TFs) are found to be involved the transcription of mouse NHE1 at the distal end of NHE1 promoter (Fernandez-Rachubinski and Fliegel, 2001). Nucleotides at −841 to
−800 base pair (bp) upstream of the start site bind to COUP-TFs and were protected in footprint analysis of the mouse promoter. Further study showed that nucleotides at -
829 to -824 are critical for COUP-TF binding. Thyroid hormone was found to stimulate mouse NHE1 transcription in addition to stimulating the activity of the protein. The −838 to −832 region is crucial for the binding of thyroid hormone to
NHE promoter (Li et al., 2002). A highly conserved poly(dA dT)-rich region at −155 to −169 bp appears to be important in regulation of expression of the NHE1 gene
39
(Yang et al., 1996). AP2 transcription factor was found to bind to the proximal promoter end at −95 to −111 bp from the start site of mouse NHE1 promoter (Dyck et al., 1995). Deletion of distal promoter sequence up to the AP2 binding site decreased promoter activity by 25% in both NIH 3T3 and p19 embryonal carcinoma (Dyck et al.,
1995; Dyck and Fliegel, 1995). Deletion of the AP2 site abolishes most of the promoter activity, suggesting that the AP2 site is important in the transcription of
NHE1.
40
1.5 AIM OF STUDY
There is mounting evidence that implicates ROS as an important signaling molecule
in various physiological processes. Our lab have previously established that the
regulation of tumor cell sensitivity to death stimulus is linked to the ratio of
·- superoxide (O 2 ) and hydrogen peroxide (H 2O2) (Clément and Pervaiz, 1999; Clément and Pervaiz, 2001; Pervaiz and Clément, 2002; Pervaiz and Clement, 2004). ER α
over-expression in breast tumor cells confers cells proliferative advantages and
resistance to apoptosis. ER α had been shown to be a redox regulated gene. Its transcriptional activity and DNA binding capacity were shown to be modulated by thioredoxin (Hayashi et al., 1997). Thiol-reacting oxidants (diamide, iodosobenzoate,
H2O2) or alkylator (iodoacetamide) produce a concentration-dependent loss in ER
DNA-binding capacity in MCF7 cells which is fully reversible by dithiothreitol (DTT) reduction (Liang et al., 1998). One study using glucose oxidase system suggested that
ER α protein itself might be redox regulated (Weitsman et al., 2009). The mechanism of this regulation is however unclear. The first aim of this project is to investigate the mechanistic role of H 2O2 in regulating ER α expression. The understanding of how
ER α is effected by ROS may have an impact on the development of new strategies for treatment and prevention of breast cancer. This involves analyzing the protein level of
ER α in H 2O2 treated MCF7 cells and establishing at which level ER α expression is being affected. The molecular mechanism of regulation together with the effects of altered ER α expression on downstream targets will be explored. Concluding the first
section is the study of how 15-Deoxy-Delta-12,14-prostaglandin J2 (15d-PGJ2),
affects ER α expression through the production of ROS intermediate.
Our group has shown that the redox regulation of cell survival in tumor could also be
associated with regulation of intracellular pH (pHi) (Pervaiz and Clément, 2002).
41
·- Central to this is the regulation of NHE1 expression. An increase in intracellular O 2 increases NHE1 promoter activity and protein expression which enhance cells’ resistance to apoptosis whereas addition of exogenous H 2O2 represses NHE1 promoter activity, protein expression and sensitizes cells to death triggers (Akram et al., 2006);
(Kumar et al., 2007). Further studies have shown that H 2O2 mediates down-regulation
of 1.1kb mouse NHE1 promoter via an oxidative and caspase dependent mechanism
(Kumar et al., 2007). The second aim of this project is to further investigate the
regulation of NHE1 expression by H 2O2. This involves identifying the response element of H 2O2 within the NHE1 promoter and studying how H 2O2 affects the promoter activity. The role of caspase in H 2O2 down-regulation of NHE1 promoter
activity will be explored. In addition, the effects of 15d-PGJ2 on NHE1 promoter
activity will be investigated. Finally, we will attempt to identify the transcription
factor targeted by H 2O2 in the down-regulation of NHE1 transcription.
42
CHAPTER 2: MATERIALS AND METHODS
2.1 MATERIALS
2.1.1 Chemicals and reagents
Dulbecco’s Modified Eagle’s Medium (DMEM), Roswell Park Memorial Institute-
1640 (RPMI-1640), Fetal Bovine Serum (FBS), Charcoal-stripped Fetal Bovine
Serum (CS-FBS), Phosphate Buffered Saline (PBS), Trypsin, and L-glutamine were from Hyclone (UT, USA). Gentamicin Sulphate was purchased from BioWhittaker
(Walkersville, MD). Geneticin (G418) and Phenol-free RPMI-1640 were supplied by
GIBCO (MD, USA). Aprotinin, Pepstatin A, Phenylmethanesulfonyl Fluoride
(PMSF), Leupeptin, Sodium Vanadate, Dimethylsulfoxide (DMSO), 1,10-
Phenanthroline monohydrate, LY294002, U0126, SP600125, MG132, GW9662,
Cycloheximide, Buthionine Sulfoximine (BSO), N-Acetyl-L-cysteine (NAC),
Estradiol (E2), Diamide, Dithiothreitol (DTT), β-mercaptoethanol (BME),
Dimethylthiourea (DMTU), Crystal Violet and Bovine Serum Albumin (BSA) were
supplied by Sigma-Aldrich (LO, USA). Peroxynitrite (ONOO-) was purchased from
Upstate (Temecula, CA, USA). Methanol, Hydrogen Peroxide (H 2O2) and Sodium
Dodecyl Sulphate (SDS) were purchased from Merck (Darmstadt, Germany). Caspase
tetra-peptide inhibitor, z-VAD-FMK was bought from R&D Systems (MN, USA).
15d-PGJ2 and Caspase Fluorogenic Substrates: Ac-VDVAD-AFC, Ac-DEVD-AFC,
Ac-VEID-AFC, Ac-IETD-AFC and Ac-LEHD-AFC were from Alexis Biochemical
(Lausen, Switzerland). 5-(and-6)-chloromethyl-2´,7´-dichlorodihydrofluorescein
diacetate acetyl ester (CM-H2DCFDA) was purchased from Molecular Probes
(Eugene, OR, USA). Coomassie PlusTM Protein assay reagent, RestoreTM Western
43
Blot Stripping Buffer, Supersignal West Pico Chemiluminescent Substrate are from
Pierce Biotechnology Rockford (IL, USA).
2.1.2 Antibodies
Antibodies for ER α (D-12), ER β (H-150), AP2 α (3B5) and Cu/Zn SOD (FL-154) were purchased from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Antibodies for Akt (C67E7), Phospho-Akt (S-473), p44/42 MAPK (ERK 1/2), Phospho- p44/42
MAPK (ERK 1/2), SAPK/JNK, Phospho-SAPK/JNK (T183/Y185) (81E11), c-Myc
(D84C12) XP (TM), and Progesterone Receptor A/B (C89F1) were purchased from
Cell Signaling Technology Inc (Beverly, MA). PARP antibody was purchased from
Clontech Laboratories (CA, USA). V5 tag antibody (E10) was purchased from
Abcam (MA, USA). NHE1 antibody was purchased from Chemicon International Inc
(CA, USA). SP1 antibody was purchased from Active Motif (CA, USA). β-actin monoclonal antibody was purchased from Sigma (LO, USA). Secondary antibodies: horseradish peroxidase (HRP)-conjugated goat anti-rabbit antibody was obtained from
DakoCytomation (Glostrup, Denmark) and HRP-conjugated goat anti-mouse was from Pierce Biotechnology (IL, USA).
2.1.3 Plasmids
Estrogen response element (ERE) reporter construct was bought commercially from
Clontech Laboratories (CA, USA). The luciferase reporter plasmid constructs of mouse NHE1 promoter: pXP-1.1, pXP-0.9, pXP 0.5, pXP-0.2, pXP-0.18, pXP-0.15, and pXP-0.12 were kindly provided by Dr. Larry Fliegel, Department of Biochemistry,
University of Alberta, Canada. Luciferase reporter plasmid constructs of human
NHE1 promoter: -1374/+16, -252/+16, and -92/+16 were kindly provided by Dr.
44
Alexey Kolyada, Department of Medicine, Tufts University School of Medicine,
Boston, USA. The mouse AP2 plasmid constructs: AP2 α, AP2 β, AP2 γ, AP2 δ and
AP2 ε were kindly provided by Dr. Bruce D. Gelb, Center for Molecular Cardiology,
Mount Sinai School of Medicine, USA. The AP2 dominant negative construct was kindly given by Dr. Frederick B. Domann, Department of Radiation Oncology, and
Holden Comprehensive Cancer Center, The University of Iowa, USA.
2.1.4 Cell lines and cell culture
All cell lines used in this project are adherent cells. Cells were grown in 37 οC
incubator supplied with 5 % CO 2. Cells were replenished with fresh medium every 3 to 4 days and split 1:5 whenever they reached 80 % confluency. For use in experiments, cells were detached from the flask using 0.125 % trypsin and pelleted by centrifugation at 1,600 rpm before resuspension in fresh medium. Cells were counted using haemocytometer prior to seeding.
Human breast cancer cells lines MCF7 and T47D were maintained in RPMI-1640 medium supplemented with 10 % fetal bovine serum (FBS), 2 mM L-glutamine and
0.05 mg/ml gentamycin sulfate. For experiments that test the effects of estrogens, cells were grown for one day in complete medium as above before changing to phenol-free RPMI 1640 supplemented with required concentration of charcoal- stripped FBS.
Wild type L6 rat myoblast cells were maintained in DMEM medium supplemented with 10 % FBS, 2 mM L-glutamine and 0.05 mg/ml gentamycin sulfate. L6 cells stably transfected with 1.1 kb of NHE1 promoter and NIH 3T3 mouse embryonic fibroblast stably transfected with 0.15 kb of NHE1 promoter were obtained from Dr.
45
Larry Fliegel, Department of Biochemistry, University of Alberta, Canada. These cells were grown similarly to L6 wild type cells with the addition of 0.25 mg/ml of genetacin to the medium.
46
2.2 METHODS
2.2.1 Treatment of cells with hydrogen peroxide (H 2O2) and other compounds
A stock solution of 10 mM H 2O2 was first prepared by diluting 30 % (v/v) H 2O2
solution with 1X PBS. Various concentrations of H 2O2 were prepared by diluting the
stock solution of 10 mM H 2O2 with 1X PBS. The H 2O2 solutions were then added to
medium to attain the indicated final concentrations. In general, seeding density of
0.08 x 10 6 cells/ well was used for 12-well plate, 0.25 x 10 6 cells/ 60 mm dish and 1.0 x 10 6 cells/ 100 mm dish.
Stock solutions of zVADfmk, U0126, LY294002, SP600125, MG132, GW9662 cycloheximide, and E2 were prepared in DMSO. 15d-PGJ2 was dissolved in ethanol.
Stock solutions of diamide, DTT, DMTU, BSO, and NAC were dissolved 1X PBS.
2.2.2 Morphology studies
The morphology of the cells was analyzed by taking photo using the Olympus camedia digital camera attached to the light microscope (Olympus CK2) at the magnification of 10X.
2.2.3 Crystal violet assay
Cells were seeded at 0.08 x 10 6 cells/ well in a 12-well plate for 2 days. After drug treatment the medium was removed from the wells and cells were washed with 1X
PBS. Five hundred microliters of crystal violet solution (0.75 % crystal violet, 50 % ethanol, 1.75 % formaldehyde and 0.25 % NaCl) was added to stain the cells in each well for 10 min. The wells were washed with deionized water a few times to remove
47
any traces of the crystal violet solution and left to air dry. The cells were lysed with
500 µl of 1 % SDS in PBS to release the crystal violet into solution. Fifty microliters of the cell lysate from each well was transferred into separate wells of a 96-well microtitre plate. Cell density was assessed by measuring cell associated crystal violet at 595 nm using Spectrofluoro Plus plate reader (TECAN, GmbH, Grodig, Austria).
2.2.4 Plasmid transfection
For promoter activity studies, 0.08 x 10 6 cells/ well were seeded in a 12-well plate.
For western blot analysis, 0.17 x 10 6 cells/ 60 mm dish were seeded. Cells were seeded for 18 h to 24 h prior to transfection. Overexpression transfection was carried out using SuperFect® Transfection Reagent (Qiagen, GmbH, Hilden, Germany) according to manufacturer’s instruction. Briefly, for one 60 mm tissue culture dish, plasmid DNA was mixed with 150 µl of serum-free, antibiotics-free DMEM and 15 µl
of SuperFect Reagent and incubated at room temperature for 10 min to allow for
SuperFect-DNA complex formation. Another 600 µl of DMEM, supplemented with
serum and antibiotic was then added to the mixture before introducing to cells. Cells
were exposed to the transfection mixture for 3h in the incubator. After 3 h, cells were
washed with 1X PBS and provided with fresh medium. Cells were normally harvested
at 48 h post transfected, unless stated otherwise.
2.2.5 Small interfering RNA (siRNA) transfection
For promoter activity studies, 0.08 x 10 6 cels/ well were seeded in a 12-well plate. For
western blot analysis, 0.17 x 10 6 cells/ 60 mm dish were seeded. Cells were seeded for
18 h to 24 h prior to transfection. siRNA transfection was carried using
Lipofectamine™ RNAiMAX Transfection Reagent (Invitrogen, Carlsbad, CA, USA)
48
according to the manufacturer’s instructions. For 60 mm tissue culture dish, siRNA
duplex was diluted with 375 µl of Opti-MEM ® I Reduced Serum Medium.
Lipofectamine RNAiMAX was also diluted with 375 µl of Opti-MEM ® I Reduced
Serum Medium. siRNA was combined with RNAiMAX and incubated at room
temperature for 20 min before adding to cells. Cells were transfected with the
transfection mixture overnight, after which cells were washed with 1X PBS and
replenished with fresh medium. Cells were normally harvested at 48 h post
transfection.
siRNA for ER α (sc-29305), AP2 α (sc-29697) and SP1 (sc-29488) were purchased
from Santa Cruz Biotechnology (Santa Cruz, CA, USA). siRNA for SP1 was also
purchased from (Thermo Scientific, IL, USA). A control siRNA that is non-
homologous to any known gene sequence was used as a negative control from
QIAGEN (Valencia, CA, USA).
2.2.6 Protein concentration determination
Protein concentration of cell lysate was determined by comparing absorbance reading
of samples with protein standards of known dilutions. Briefing, 200 µl of Coomassie
Blue (PIERCE, IL, USA) was added to 2 µl of cell lysate in a 96-well plate and
incubated at room temperature for 5 min. Absorbance was read at 595 nm using
Spectrofluoro Plus spectroflurometer (TECAN, GmbH, Grodig, Austria). Protein
standards of 2.5, 5, 10 and 20 µg/ml were prepared using bovine serum albumin
(PIERCE, IL, USA) and treated similarly to obtain a standard graph from the
absorbance readings. Protein concentration of the cell lysates were estimated from the
standard graph.
49
2.2.7 SDS-PAGE and western blot
After treatment, the medium was removed and cells were washed once with 1X PBS
before they were removed from the tissue culture plate by scrapping method and
resuspended in PBS. The cells were pelleted down by centrifugation at 12,000 rpm for
5 min and lysed in RIPA buffer (50 mM Tris at pH 7.5, 150 mM NaCl, 1% v/v
deoxycholic acid, 0.1% v/v SDS and 1 mM EDTA) containing protease inhibitors (1
mM PMSF, leupeptin, pepstain A, aprotinin) and 1 mM phosphatase inhibitor
ο Na 3VO 4. Fifty microgram of protein was incubated at 95 C for 5 min and resolved by
8 % sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) at 100
V in a Bio-Rad Mini-PROTEAN 3 Cell (CA, USA). For detection of NHE1 protein,
samples were heated at 37 οC instead. Kaleidoscope standard (Bio-Rad, Ca, USA)
was loaded to estimate protein size. Resolved proteins were transferred to a
nitrocellulose membrane (Pall Corporation) by wet transfer method using Bio-Rad
Mini Trans-Blot Electrophoretic Transfer Cell (CA, USA) at 370 mA for 1 h. The
membrane was blocked in 5 % (w/v) non-fat milk in Tris-buffered saline/ 0.1 (v/v)
Tween 20 (TBST) for 1 h. The membrane was washed three times with TBST and
probed with the respective 1 o antibody dissolved in BSA overnight at 4 οC. After washing off the primary antibody with TBST, respective horse radish peroxidase
(HRP) conjugated 2 o antibody in 5 % fat-free milk was incubated with the membrane for 1 h. Membranes were washed three times with TBST, specific protein-antibody interactions were detected by enhanced chemiluminscence using SuperSignal
Chemiluminescent Substrate (PIERCE, IL, USA) with Kodak Biomax MR X-ray film.
For re-probing of the same membrane for different proteins, blots were stripped with
Restore Western Blot Stripping Buffer and re-probed for another protein of interest.
50
The bands obtained were scanned in with EPSON Perfection 1250 and analyzed by
FujiFilm Multigauge V3.0. The band density for proteins was normalized against the
band density of β-actin.
2.2.8 Caspase assay
After treatment with H 2O2 for specific period of time, cells were scraped off the dish and transferred to a 15 ml falcon tube together with the medium. Cells were pelleted down by centrifugation at 2500 rpm for 5 min at 4 οC. Cells were washed once with
PBS and pelleted. The supernatant was removed and cells were resuspended with 100
µl of Cell Lysis Buffer (BD Pharmigen). The lysate was incubated on ice for 10 min and transferred to a 1.5 ml eppendorf tube. For caspase assay, cell lysates were centrifuged at 12000 rpm for 5 min at 4 οC to remove cell debris. Forty microliters of
each sample was transferred into a 96-well plate and mixed 50 µM of specific caspase
substrate (caspase 2 substrate: Ac-VDVAD-AFC, caspase 3/7 substrate: Ac-DEVD-
AFC, caspase 6 substrate: Ac-VEID-AFC, caspase 8 substrate: Ac-IETD-AFC, and
caspase 9 substrate: Ac-LEHD-AFC) and 40 µl of 2X reaction buffer (10 mM HEPES, pH 7.4, 2 mM EDTA, 6 mM DTT, 10 mM KCl and 1.5 mM MgCl2) supplemented with protease inhibitors (1 mM PMSF, 10 µg/ml Aprotinin, 20 µg/ml pepstatinA, 10
µg/ml leupeptin). Samples were incubated at 37 οC for 1 h and AFC fluorescence was read at an excitation wavelength of 400 nm and an emission wavelength of 505 nm using Spectrofluoro Plus spectroflurometer (TECAN, GmbH, Grodig, Austria).
Caspase activity was normalized against protein concentration of each sample and expressed as relative fluorescence unit per microgram of protein (RFU/ µg).
51
2.2.9 Single and dual luciferase assay
NHE1 promoter activity of stably transfected cells was assessed with a single-
luciferase assay kit (Promega, Madison, WI) according to manufacturer’s instructions.
Cells were grown in 12-well plate and subjected to treatment. After treatment, cells
were washed with 1X PBS and lysed with 100 µl of reporter lysis buffer at 4 ˚C and stored at -80 ˚C before assay. For the assay, 10 µl of the cell lysate was added to 50 µl
of the luciferase substrate. Bioluminescence generated was measured using a Sirius
luminometer (Berthold, Pforzheim, Germany). The luminescence readings obtained
were then normalized with the protein concentration of the cell extract using the
Coomassie PlusTM Protein assay reagent.
Dual luciferase assay was used to measure promoter activity of cells transiently
transfected NHE1 promoter-luciferase construct. Fifty thousand cells/ well were
seeded overnight in 12-well plate and 1 µg of NHE1 promoter linked to luciferase
gene construct was transfected into cells using SuperFect Transfection Reagent for 3 h.
As a transfection control, 0.1 µg of Renilla plasmid was co-transfected. After transfection, cells were washed with 1X PBS and given fresh medium and allowed to recover. Forty-eight hours post-transfection, cells were washed with 1X PBS and lysed with 100 µl of passive lysis buffer.
For transfection of ERE, MCF7 cells were seeded at 0.05 x 10 6 cells/well in 12-well
plate overnight. One microgram of ERE plasmid construct was transfected in each
well using SuperFect Transfection Reagent for 3 h. As a transfection control, 0.1 µg of Renilla plasmid was co-transfected. After transfection, cells were recovered overnight in phenol-free RPMI-1640 supplemented with 10 % CS-FBS. Cells were then treated with 100 µM of H2O2 for 16h before 10 nM of E2 was added for 2, 4, or 6
52
h. After treatment, cells were washed with 1X PBS and lysed with 100 µl of passive lysis buffer.
Dual-Luciferase® reporter assay system (Promega, Madison, WI, USA) was used measure the promoter activity of cells transiently transfected with luciferase plasmid constructs, according to the manufacturer’s manual.
2.2.10 Chloramphenicol acetyl transferase (CAT) assay
NHE1 promoter activity of the transiently transfected cells with CAT reporter plasmid constructs was assessed using an enzyme immunoassay to determine the expression of the CAT. For transfection, cells were seeded at 0.05 x 10 6 cells/ well in 12-well plate overnight. One microgram of human NHE1 plasmid construct was transfected in each well using SuperFect Transfection Reagent for 3 h. As a transfection control, 0.1 µg of Renilla plasmid was co-transfected. Cells were washed with 1X PBS and given fresh medium and allowed to recover. Forty-eight post-transfection, cells were harvested and the promoter activity was analyzed by CAT ELISA assay (Roche) according to manufacturer’s protocol. The levels of CAT protein were quantified using a CAT antigen capture enzyme-linked immunosorbent assay (ELISA) (Roche
Molecular Biochemicals).
To measure the renilla luciferase activity, 50 µl of stop and glow substrate was added to 10 µl of cell lysate. The bioluminescence generated was measured using Sirius
luminometer. All CAT quantifications were normalized to the Renilla luciferase
reading.
53
2.2.11 RNA isolation
Total RNA was isolated by using RNeasy Mini Kit (Qiagen, GmbH, Hilden, Germany)
according to manufacturer’s instructions. RNA concentration and purity was
determined using Nanodrop-1000 (Thermo Scientific, Wilmington, DE, USA).
2.2.12 Reverse transcription (RT) and real-time chain polymerase reaction
(PCR)
Two hundred nanogram of total RNA was reverse transcribed with 1.1U/ µl
MultiScribe™ reverse transcriptase in the presence of 1X RT buffer, 5mM MgCl 2,
425 µM of each dNTPs, 2 µM random hexamers, 0.35U/ µl RNase inhibitor, made up to 10 µl with sterile water. RT reaction was carried out in Mastercycler gradient
(Eppendoff, USA) at 25 οC for 10 min, followed by 37 οC for 60 min and a terminating step of 95 οC for 5 min.
For real time PCR, 200 ng of total RNA was transcribed as described above. For a 50
µl reaction, 5 µl of RT product was mixed with 1X TaqMan® Universal PCR Master mix, 2.5 µl of 20x specific TaqMan probe, 2.5 µl of 18S rRNA TaqMan probe, topped up to 50 µl with sterile water. A negative control for RT in which sterile water
replaced the RNA template was included. Another control where RT mix was
replaced with sterile water was included to check for DNA contamination. Real-time
PCR was carried out using 7500 Fast Real-Time PCR System (Applied Biosystems,
USA) with a protocol that consists of 50 οC for 2 min, 95 οC for 10 min, followed by
40 cycles of denaturing at 95 ο C for 15 sec and extension at 60 οC for 1 min. Results were analyzed using Sequence Detection Software version 1.3 provided by Applied
Biosystems. Relative gene expression was obtained after normalization with
54
endogenous 18S and determination of the difference in threshold cycle (Ct) between
treated and untreated cells using 2 -∆∆Ct method.
2.2.13 Intracellular ROS assessment by CM-H2DCFDA
Intracellular concentration of ROS was determined by staining cells with the redox
sensitive dye CM-H2DCFDA, which fluoresces under oxidation by H 2O2 and its free radical products. Briefly, cells were washed with PBS once and detached off the tissue culture dish by incubating with 0.125 % trypsin at 37 οC for 10 min. Trypsin
was neutralized with 500 µl of medium and the cell pellet was collected by pulse-spin
centrifugation. The cells were incubated with 5 µM of CM-H2DCFDA at 37 οC in the
dark for 20 min. Cells were then pelleted and washed with PBS to remove excess dye.
Cells were resuspended in 500 µl of PBS and filtered through a 41 µm filter before injected into the Coulter Epics Elite ESP Flow Cytometer. Ten thousands events were collected with the excitation wavelength of 495 nm and emission wavelength of 525 nm. No dye treatment and 300 µM of H 2O2 were used as negative and positive
controls respectively.
2.2.14 Nuclear-cytoplasmic fractionation
Fractionation of the nucleus and cytosol was carried out using NE-PER Nuclear and
Cytoplasmic Extraction Reagents (PIERCE, Thermo Fisher Scientific Inc, Rockford,
USA) according to manufacturer’s instructions. Briefly, cells were washed with 1X
PBS and scrapped off the plate with the help of a scrapper and transferred to a 15 ml
falcon tube. The amount of the various buffers added and duration of lysis was
performed according to manufacturer’s instructions.
55
2.2.15 Electromobility shift assay (EMSA)
The synthetic biotin-labeled oligonucleotide of the sequence 5’-bCGC GAT ACT
TCT TTC CCT CGG CGA CAG GGG CCG CTG CGC T-3’ and non-labeled reverse
complement 5’-AGC GCA GCG GCC CCT GTC GCC GAG GGA AAG AAG TAT
CGC G-3’ were made corresponding to the 5’ 0.15kb of human NHE1 promoter. The
oligonucleotides were heated up to 95 οC for 5 min and allowed to cool to room temperature for annealing. EMSA was performed using LightShift Chemiluminescent
EMSA Kit (Thermo Scientific, IL, USA). Five microgram of MCF7 nuclear extracts was incubated with H2O2 for 20 min at room temperature. For treatment with reducing agent, the protein reaction samples were further incubated with various concentrations of DTT for another 20 min. The protein reaction mixtures were subsequently incubated with biotin-labeled DNA oligonucleotides in the presence of 1 µg poly dIdC and 1X binding buffer (10 mM Tris, 50 mM KCl, 1 mM DTT, 2.5 % glycerol, 5 mM MgCl 2, 0.05 % NP-40) for 20 min at room temperature. The bound DNA-protein was separated from free oligonucleotides by gel electrophoresis in 5 % native polyacrylamide gels in 0.5X TBE buffer (45 mM Tris-borate, 1 mM EDTA).
Electrophoresis was conducted at a constant current of 100 V for 45 min. The protein-
DNA complex was transferred to a nylon membrane using by wet transfer method using Bio-Rad Mini Trans-Blot Electrophoretic Transfer Cell (CA, USA) at 380 mA for 30 min. The DNA was cross-linked to the nylon membrane at 120 mJ/ cm 2 using a
commercial UV-cross-linker instrument equipped with 254 nm bulk. The membrane
was treated according to the manufacturer’s instructions to prepare it to be detected by
chemiluminescence with Kodak Biomax MR X-ray film.
56
2.2.16 Statistical analysis
Statistical analysis was performed using algorithm of unpaired Student’s t-test from
Microsoft Excel software to compare between means of relevant conditions. P-value of less than 0.05 was considered significant.
57
CHAPTER 3A: RESULTS – REDOX REGULATION OF ER α
3A.1 H2O2 AND THE EXPRESSION OF ER α PROTEIN
Depending on its concentration, H 2O2 can have a plethora of effects on cells. Low
amount of H 2O2 has been shown to increase cell proliferation while moderate amount
of H 2O2 regulates cellular processes such as redox regulation of reactive cysteine on
transcription factors and protein kinases. On the extreme end, H 2O2 causes cell death either by apoptosis or necrosis. Different types of cells also have different resistance or antioxidant capacity against ROS challenge.
3A.1.1 Morphology and growth of MCF7 cells upon exposure to H 2O2
MCF7 is a breast cancer cell line derived in the Michigan Cancer Foundation in 1973 from a pleural effusion of a 69 year old Caucasian woman (Soule et al., 1973). MCF7
cell line was chosen to study the redox regulation of ER α as it expresses high level of
the receptor and has been used as a model of ER α action in large number of reports
(Pinzone et al., 2004).
Here, we tested the effects of increasing concentrations of H 2O2 on the morphology
and growth of MCF7 cells. There were some morphological changes in MCF7 cells
treated with H 2O2 for 24 h. Starting from 50 µM of H 2O2, cells appeared flatter and
less clustered from one another (Fig. 1a). Cell density also decreased with increasing
concentrations of H 2O2. Notably, density of cells treated with 200 µM of H2O2 looked
similar to cells at the time of trigger.
To assess the effects of H 2O2 challenge on the growth of MCF7 cells, the cells were treated with increasing concentrations of H 2O2 for 24 and 48 h. Cell density was
58
estimated by crystal violet staining. The time at which H 2O2 was added was regarded
as time 0 h. Cell growth was expressed as percentage relative to the time of treatment.
At lower concentrations, cells showed a decreased proliferation starting from 50 µM of H 2O2. H 2O2 induced a concentration-dependent inhibition of cell growth with total inhibition seen at 200 µM of H2O2 treatment (Fig. 1b). There was no statistical difference in the cell density when cells were treated with 200 µM of H 2O2 and allowed to grow for 24 and 48h as compared to time of treatment.
59
a)
Time of trigger Control 25 µM
50 µM 100 µM 200 µM
b)
350 * * 300 * 250 * * 200 * * * 24h 150 48h
of trigger) of 100
50
0 Cell density (% relative to time time (% to relative density Cell 0 25 50 100 200
H2O2 (µM) *: p-value<0.05 (relative to time of treatment)
Figure 1: MCF cell survival upon H 2O2 treatment. a) Morphology of MCF7 upon H 2O2 treatment. MCF7 cells were seeded for 2 days before being treated with increasing concentrations of H 2O2 for 24 h. b). Cell density and survival were estimated by staining the cells with crystal violet 24 h and 48 h after the treatment. Results were obtained from mean of 3 independent experiments done in duplicates +/- SE.
60
3A.1.2 Effect of H 2O2 on the expression of ER protein
ER α had been shown to be a redox regulated gene. Its transcriptional activity and
DNA binding capacity could be modulated by oxidants such as diamide, iodosobenzoate and H2O2. Next, the effects of oxidative stress in the form of H2O2 on
the expression on ER protein itself were investigated. The effects of H 2O2 on ER
subtypes were examined through a 48 h time-course period. Cells were treated with
increasing concentrations of H 2O2 (0 to 200 µM) for 24 and 48 h. The total cell
lysates were subjected to western blot using specific antibodies against ER α and ER β.
As with the inhibition of cell growth, H 2O2 exerted its effects on ER α in a concentration-dependent manner (Fig. 2a). Figure 2b shows that treatment of MCF7 cells with H 2O2 decreased ER α protein level to 73, 26 and 5 % of untreated cells with
50, 100 and 200 µM of H2O2 respectively in the first 24 h. A statistically significant
decrease in the expression of ER α protein was observed with 100 and 200 µM of
H2O2. After 48 h of treatment, ER α level returned to 95, 69 and 8 % of untreated cells
with 50, 100 and 200 µM of H 2O2 respectively. The restoration of ER α is statistically
significant for 100 µM of H2O2. The result suggests that the down-regulation of ER α
is reversible at concentration of 100 µM of H 2O2 or less. ER β level on the other hand was unaffected by H 2O2 treatment and stayed constant regardless of the concentration cells were exposure to.
61
a)
24h 48h
H2O2 µM 0 25 50 100 200 0 25 50 100 200
ER α -65 kDa
ER β -59 kDa
β-actin -43 kDa
b)
140
120 ** control) 2 100 O 2
80 *
60 24h 48h 40 *
(% relative relative H no to (% 20 * α * ER 0 0 25 50 100 200
H2O2 (µM)
*: p-value < 0.05 (relative to no H 2O2 control) **: p-value < 0.05
Figure 2: H2O2 induces concentration dependent down-regulation of ER α. a) MCF7 cells were treated with increasing concentrations of H 2O2 for 24 h and 48 h. Cell extracts were analyzed by western blot. b) Densitometry of ER α protein level (% relative to no H 2O2 control). Results shown are representative of 3 independent experiments.
62
An experiment was carried out to ascertain whether the ER α protein was specifically recognized by the antibody. MCF7 cells were transfected with either 50 nM control
(scrambled) siRNA or different concentrations of ERα specific siRNA. Results from
Figure 3 show that the ER α band disappeared when treated with ER α specific siRNA
from as low as 10 nM. This confirmed the ER α antibody specifically detects ER α
protein. 20nM 50nM 10nM α α α siER siER siER NS siRNA 50nM siRNA NS ER α -65 kDa
β-actin -43 kDa
Figure 3: Specificity of ER α antibody. MCF7 cells were grown overnight and transfected with ER α siRNA at different concentrations using Lipofectamine RNAiMAX reagent as described in the Materials and Methods section. Cells were harvested at 48 h post transfection.
63
To determine that the effect of H2O2 is not limited to one single cell line, another ER α
positive breast cancer cell, T47D was challenged with H2O2. Similar to MCF7 cells,
the ER α level of T47D was down-regulated by H 2O2 in a concentration-dependent
manner in 24 and 48 h (Fig. 4a and 4b). Thus, the result indicates that the repression
of ER α protein by H2O2 was not limited to MCF7 cells.
a)
MCF7 24h 48h
H2O2 µM 0 25 50 100 200 0 25 50 100 200 ER α -65 kDa
β-actin -43 kDa
b)
T47D 24h 48h
H2O2 µM 0 25 50 100 200 0 25 50 100 200
ER α -65 kDa
β-actin -43 kDa
Figure 4: H 2O2 down-regulates ER α in both MCF7 and T47D cells. a) MCF7 and b) T47D cells were treated with increasing concentrations of H 2O2 for 24 h and 48 h. Cell extracts were analyzed by western blot. Results shown are representative of 3 independent experiments.
64
When ER α is not bound to its ligand, it resides in the cytosol. It migrates into the nucleus once it binds to its ligand and homodimerizes. In a normal growing cell, ER α
can be found both in the cytosol and nucleus (Jazbutyte et al., 2009) as there will be
some estrogenic ligands found in the cell culture medium. As the results in Figure 2a
came from total cell lysates, the observed decrease in ER α is from a pool of both the cytosolic and nucleus fractions. In order to understand which fraction of ER α was
affected by H 2O2 treatment, a cytosolic and nuclear fractionation was carried out with
cells treated with or without 200 µM of H 2O2. Figure 5 shows that without treatment,
MCF7 cells indeed have both cytosolic and nucleus pools of ER α. When cells were
treated with H 2O2, both pools of ER α protein were down-regulated. The results
indicate that the down-regulation of ER α protein by H 2O2 is not specific to a particular cellular compartment.
Medium H2O2 200 µM
CCNN
PARP -116 kDa
ER α -65 kDa
Cu/Zn SOD -16 kDa
Figure 5: H 2O2 induces down-regulation of both nucleus and cytosolic fractions of ER α. MCF7 cells were seeded for 2 days and treated with or without 200 µM of H 2O2 for 24 h. Cells were subjected to fractionation of the nucleus and cytosol using NE-PER nuclear- cytoplasmic extraction reagents. C: cytosolic fraction, N: nucleus fraction. PARP is the nucleus marker, while Cu/Zn SOD is cytosolic marker.
65
3A.1.3 Effects of estrogen on H 2O2 down-regulation of ER α
Estrogen has been shown to be involved in the feedback mechanism in the down-
regulation of ER α. Stimulation of cells with E2 will lead to rapid degradation of intracellular ER α (Tateishi et al., 2004; Alarid et al., 1999; Nawaz et al., 1999). To
determine if estrogen or estrogen-like compounds play any role in the repression of
ER α by H 2O2, we tested the down-regulation of ER α by H 2O2 in different regular
serum conditions and also in phenol-free medium supplemented with charcoal-
stripped serum. Regular serum contains many growth factors including estrogens, and
the phenol added to medium also has estrogen-like functions (Berthois et al., 1986).
Treating regular serum with charcoal/ dextran removes steroid hormones and growth
factors including most of the estrogen (Cao et al., 2009). Thus, growing the cells in
phenol free medium supplemented with charcoal-stripped serum will deprive the cells
of exogenous estrogen and estrogen-like molecules. Figure 6a shows the effects of
100 µM of H2O2 on cells grown in regular RPMI containing phenol-red supplemented
with 10 %, 0.5 % and 0 % fetal bovine serum (FBS). Figure 6b shows the effects of
100 µM of H2O2 on cells grown in phenol-free RPMI supplemented with 10 %, 0.5 % and 0 % charcoal-stripped FBS. In all the conditions, H2O2 was able to down-regulate
ER α protein in 24 h but the expression of the protein was recovered by 48 h. The basal levels of ER α when cells were grown in regular medium and in estrogen deprived condition were also similar. These results show that the down-regulation of
ER α by H2O2 is independent of estrogen present in the medium and precluded the possibility of H2O2 increasing the affinity of ER α for estrogen, leading to enhanced
receptor degradation.
66
a)
10% regular FBS 0.5% regular FBS 0% regular FBS
24h 48h 24h 48h 24h 48h 0 100 0 100 0 100 H2O2 µM 0 100 0 100 0 100
-65 kDa ER α
β-actin -43 kDa
b)
10% CS FBS 0.5% CS FBS 0% CS FBS 24h 48h 24h 48h 24h 48h H2O2 µM 0 100 0 100 0 100 0 100 0 100 0 100
-65 kDa ER α
β-actin -43 kDa
Figure 6: ER α is down-regulated in all serum conditions. MCF7 cells were seeded overnight. On day two, the medium was changed to either a) regular RPMI containing different percentage of regular serum (FBS) or b) phenol-free RPMI containing different percentage of charcoal-stripped serum (CS-FBS). On day three, cells were treated with 100 µM of H 2O2 for 24 and 48 h. Cell extracts were analyzed by western blot.
67
3A.1.4 Time-dependent regulation of ER α protein by H 2O2
Using 100 and 200 µM of H 2O2, the study was extended to cover a wider time course to determine when ER α protein level starts to decrease. Figure 7a shows that 4 h following the exposure of MCF7 cells to H 2O2, ER α of treated and untreated cells
were similar. However, at 8 h after treatment, ER α protein level was decreased from
100 % to 71 % and 40 % in the presence of 100 and 200 µM of H2O2 respectively
(Fig. 7b). Consistent with Figure 2, the ER α protein level of cells treated with 100 µM of H 2O2 was higher at 48 h as compared to 24 h. At 24 h after 100 µM of H2O2
treatment, ER α protein level of treated cells was at 16 % compared to untreated cells
while the level increased to 70 %, 48 h after treatment. ER α protein level of cells
treated with 200 µM of H 2O2 remained low even at 48 h. This data suggests that the
down-regulation of ER α by H 2O2 is a reversible event. To determine if ER α down-
regulation is reversible with 200 µM of H 2O2, cells were incubated for a longer period of time, up to 96 h after H2O2 treatment. Results show that the down-regulation of
ER α by 200 µM of H 2O2 is also reversible. ER α level of cells treated with 200 µM of
H2O2 started to recover at 72 h after treatment (Fig. 8).
68
a)
2h 4h 8h 16h 24h 48h
H O M 2 2 µ 0 100 200 0100 200 0 100 200 0 100 200 0 100 200 0 100 200
ER α -65 kDa
β-actin -43 kDa
b)
140
120 ** control) 2
O 100 2
2h 80 * 4h 8h 60 16h * 24h 40 * 48h * * level (% relative no H (% to relative level 20
α * * * ER 0 100 200
H2O2 (µM)
*: p-value < 0.05 (relative to no H 2O2 control) **: p-value < 0.05
Figure 7: H2O2 induces a time-dependent down-regulation of ER α. a) MCF7 cells were treated with increasing concentrations of H 2O2 for various time-points. Cell extracts were analyzed by western blot. b) Densitometry of (a). Results shown are representative of 3 independent experiments.
69
Medium H2O2 100 µMH2O2 200
Hr 0 24 48 72 96 0 24 48 72 96 0 24 48 72 96
ER α -65 kDa
β-actin -43 kDa
Figure 8: H2O2-induced down-regulation of ER α is reversible. MCF7 cells were treated with 0, 100 and 200 µM of H 2O2 for various time-points. Cell extracts were analyzed by western blot. Results shown are representative of 3 independent experiments.
3A.1.5 Chronic ROS leads to continuous suppression of ER α
Deregulation of the ROS production system in cells and antioxidant defense
mechanism is a common characteristic of cancer cells (Burdon, 1995). ER α negative breast cancer cells were found to have elevated amount of ROS (Choi et al., 2010).
We have previously shown that H2O2 can induce down-regulation of ER α protein (Fig
2a). This repression of ER α expression is reversible when the cells were treated with a
one-time dose of H2O2 (Fig 8). Here, we hypothesize that chronic presence of H2O2 could lead to ER α expression being suppressed for a prolonged period of time. To test the hypothesis, one set of MCF7 cells were treated with 100 µM of H2O2 once and harvested at every 24 h to monitor the ER α protein level. Another set of cells was treated with 100 µM of H2O2 every 24 h. Results in Figure 9 show that treating cells with one bolus concentration of H 2O2 led to a transient decrease in ER α at 24 h which
started recovering at 48 h after treatment. The set of cells treated with the periodic
addition of H2O2 maintained low ER α level throughout the 96 h tested.
70
H2O2 100 µM x1 H2O2 100 µM every day
Hr 0 24 48 72 96 0 24 48 72 96
ER α
β-actin
Figure 9: Chronic oxidative stress lead to ER α-negative phenomenon. MCF7 cells were treated with 100 µM of H2O2 once or every 24 h for 96 h. Cells were harvested at 24 h time interval. Cell extracts were analyzed by western blot analysis. Results shown are representative of 3 independent experiments.
3A.1.6 ROS involved in the down-regulation of ER α expression
H2O2 in the presence of iron can undergo Fenton Reaction to form a more reactive
molecule known as hydroxyl radical (OH·).
2+ 3+ − Fe + H 2O2 → Fe + OH· + OH
This hydroxyl radical is very potent as it can oxidize various cellular components
including DNA, proteins, and lipids (Stinefelt et al., 2005). To determine if hydroxyl
radical is the agent down-regulating ER α, MCF7 cells were pretreated for 2 h with different concentration of DMTU, a scavenger of hydroxyl radicals and also H2O2 to a lesser extent (Fox, 1984). Cells were then treated with 200 µM of H2O2 for another
24h. Figure 10a shows that there was slight restoration of the ER α protein in cells
preincubated with DMTU. As DMTU scavenges mainly hydroxyl radicals, the
inability of DMTU to completely restore ER α protein level suggested that hydroxyl
radicals are not the main effecter molecule in the down-regulation of ER α by H2O2.
To form hydroxyl radical, Fe 2+ is required. 1, 10-phenanthroline, a potent scavenger of Fe 2+ was used to scavenge iron in the cells. MCF7 cells were preincubated for 1 h
with different concentrations of 1, 10-phenanthroline before being treated with 71
200 µM of H2O2 for another 24 h. Scavenging iron, the co-factor required for formation of hydroxyl radical, was unable to prevent the down-regulation of ER α by
H2O2 (Fig. 10b). The results again suggest that hydroxyl radicals are not involved in the down-regulation of ER α in cells treated with H2O2.
a)
- + + + + H2O2 200 µM DMTU mM - - 2 5 10
ER α -65kDa
β-actin -43kDa
b)
H O 200 µM - + + + + 2 2 1,10-phenanthroline µM- - 5 10 20
ER α -65kDa
β-actin -43kDa
Figure 10: Hydroxyl radical not involved in down-regulation of ER α by H 2O2. a) MCF7 cells were seeded for 2 days. Cells were left untreated or pretreated with increasing concentrations of DMTU for 2 h before exposure to 200 µM of H 2O2 for 24 h. b) Cells were pretreated with various different concentrations of 1,10-phenanthroline for 1 h before exposed to 200 µM of H 2O2 for 24 h. Cell extracts were analyzed by western blot.
72
3A.1.7 Effect of peroxynitrite on the expression of ER α protein
Peroxynitrite is an anion formed in vivo by the reaction of NO with superoxide and is a powerful oxidizing agent that can induce lipid peroxidation, oxidize sulfhydryls, and nitrates the aromatic residues of proteins (Koppenol et al., 1992; Beckman and
Koppenol, 1996). Although more reactive than H 2O2, it is also more unstable. The anion is stable at basic conditions but is protonated at physiological condition and has a half-life of about 1 second. Here, we tested whether extracellularly added peroxynitrite could also down-regulate ER α protein. Figure 11 shows a concentration- dependent down-regulation of ER α by peroxynitrite.
24h
Peroxynitrite µM 0 100 200 500 1000
ER α -65kDa
β-actin -43kDa
Figure 11: Peroxynitrite down-regulates ER α. MCF7 cells were treated with various concentrations of peroxynitrite for 24 h. Cell extracts were analyzed by western blot. Results shown are representative of 2 independent experiments.
73
3A.2 DOWN-REGULATION OF ER α DOES NOT INVOLVE THE
PROTEOSOMAL DEGRADATION PATHWAY
ER α has been shown to be actively degraded via proteosomal pathway upon activation by E2 (Liang and Nilsson, 2004). Unliganded ER α has also been shown to
undergo proteosomal degradation in the absence of E2 (Tateishi et al., 2004). Hence,
the decrease in the expression of ER α upon exposure of the cells to H 2O2 could be due
to the activation of proteosomal degradation pathway. To test this hypothesis, we
utilize MG132, a proteosomal inhibitor, to prevent active degradation of ER α by proteosome upon H 2O2 treatment. An experiment was set up to determine the effective concentration of MG132 required for effective proteosomal inhibition in MCF7. The c-Myc protein, which has a short half-life due to rapid and continuous degradation by proteosome, was used as a marker of proteosomal inhibition. MCF7 cells were preincubated with various concentrations of MG132 for 2 h, after which protein synthesis was stopped by adding 100 µg/ml of cycloheximide for another 2 h.
Figure 12a shows that the degradation of c-Myc was inhibited by as low as 1 µM of
MG132. Maximum c-Myc inhibition was however seen with 2 µM of MG132. Thus,
2 µM of MG132 was used to determine the role of proteosome in H2O2 down-
regulation of ER α.
74
a)
Cycloheximide - + + + + +
MG132 µM - - 1 2 5 10
c-Myc -62kDa
ER α -65kDa
β-actin -43kDa
b)
300
250
200
150 control) 100
50
c-Myc level (% relative to relative untreated (% level c-Myc 0 0 0 1 2 5 10
MG132 µM
Chx 100 µg/ml
Figure 12: Effective concentration of proteosomal inhibition by MG132. a) MCF7 cells were seeded for 2 days before preincubation with different concentrations of MG132 for 2 h. Protein synthesis was subsequently blocked with 100 µg/ml cycloheximide for 2 h. Cell extracts were analyzed by western blot. b) Densitometry of c-Myc protein level (% relative to untreated control).
75
To determine whether the early down-regulation of ERα protein was due to proteosomal degradation, MCF7 was treated with or without H2O2 for 8 h in the presence or absence of 2 µM MG132. The western blot analysis reveals that although
the MG132 was effective in blocking proteosomal degradation, as shown by the
accumulation of c-Myc protein in the MG132-treated only control, H2O2 was still able
to induce a down-regulation of ER α protein in the presence of the proteosome inhibitor (Fig. 13a). MG132 treatment was unable to reverse H2O2-mediated early down-regulation of ER α protein, which suggests that H2O2 does not alter early ER α
protein degradation. To determine whether sustained down-regulation of ER α protein by H2O2 involves the proteosomal pathway, MCF7 cells were first treated with either
0 or 200 µM of H 2O2 for 16 h. Two micromolar of MG132 was then added to one set
of the cells for 6 h while another set received DMSO as vehicle control. Figure 13b
shows that H 2O2 was still able to strongly down-regulate ER α protein in the presence of MG132, suggesting that sustained down-regulation of ER α does not involve
proteosomal pathway.
76
a)
Medium + MG132 2 µM
H2O2 µM 0200 0 200
ER α -65kDa
β-actin -43kDa
c-Myc -62kDa
b)
Medium + MG132 2 µM H O µM 2 2 0200 0 200
-65kDa ER α
-43kDa β-actin
c-Myc -62kDa
Figure 13: H2O2-induced decrease in ER α is not due to proteosomal degradation. a) MCF7 cells were seeded for 2 days before co-treated with or without 2 µM proteosome inhibitor MG132 and exposed to either 0 or 200 µM of H 2O2 for 8 h. b) MCF7 cells were seeded for 2 days before treatment with either 0 or 200 µM of H 2O2 for 16 h. Cells were then treated with 2 µM of proteosome inhibitor MG132 for another 6 h. Cell extracts were analyzed by western blot. Results shown are representative of 3 independent experiments.
77
3A.3 THE DOWN-REGULATION OF ER α BY H 2O2 IS VIA DECREASE IN
ER α mRNA LEVEL
3A.3.1 Effects of H 2O2 on the expression of ER α mRNA level
Since H2O2-mediated down-regulation of ER α does not involve the proteosomal
degradation pathway, we examined whether the effects could be due to the inhibition
of ER α mRNA transcript level. MCF7 cells were treated with increasing concentrations of H2O2 for 24 h and the level of ER α mRNA was monitored by real- time PCR analysis of total RNA using specific TaqMan primer probe for ER α. 18S
RNA was used as a loading control. As shown in Figure 14a, there was a concentration-dependent reduction in the level of ER α mRNA in cells treated with
H2O2. Cells treated with 50, 100 and 200 µM of H2O2 showed a statistically significant difference in the mRNA transcript level as compared to untreated control.
Given that ER α protein level was shown to decrease as early as 8 h after H2O2 treatment, the time when ER α mRNA level started to decline was assessed. Figure
14b shows that as early as 4 h into H2O2 treatment, ER α mRNA level reduced significant when compared to its untreated control. The decrease in ER α mRNA level accelerated from 8 h and the repression was sustained for up to 24 h after H2O2
addition. At 48 h post-treatment, ER α mRNA level started to increase slightly. The
western blot in Figure 14b shows that the decline in ER α mRNA level preceded the
down-regulation of ER α protein.
As shown earlier, the ER α protein level of cells treated with 100 and 200 µM of H2O2
was restored by 48 h and 72 h respectively (Fig. 8). To determine if it is due to
recovery of the ER α mRNA level, we treated the cells with H2O2 and observed the
mRNA level for up to 72 h. Mirroring the protein expression (Fig. 8), ER α mRNA
78 level of cells treated with 100 µM of H2O2 was significantly increased at 48 h (Fig.15).
And at 72 h, the ER α mRNA level of these cells was similar to that of untreated control. For cells treated with 200 µM of H2O2, mRNA level of ER α was still low at
48 h after treatment. This is consistent with the low ER α protein expression observed at 48 h post-treatment (Fig. 8). In line with the restoration of protein expression at 72h, there was significant increase in mRNA level for cells treated with 200 µM of H2O2 after 72 h (Fig. 15). This set of results provides strong evidence that the down- regulation of ER α protein by H2O2 was a direct result of decrease in ER α mRNA level.
a)
1.2
1 control) 2 O 2 0.8
* 0.6
0.4 * * 0.2 mRNA (Fold relative no H to relative (Fold mRNA α 0 ER 0 25 50 100 200 H O (µM) 2 2
*:p-value < 0.05 (relative to no H 2O2 control)
79 b)
H2O2 200 µM Hr 0 2 4 8 16 2448 1.20
2 ER α -65 kDa O 2 1.00 β-actin -43 kDa
0.80 * *
0.60 control) 0.40 * * mRNA (Fold relative no H to relative (Fold mRNA 0.20 α * ER 0.00 2 4 8 16 24 48
Time upon H 2O2 200 µM treatment
*:p-value < 0.05 (relative to no H 2O2 control)
Figure 14: H2O2 induces time dependent down-regulation of ER α mRNA level . a) MCF7 cells were seeded for 2 days before treated with increasing concentrations of H 2O2 for 24 h. ER α mRNA level was measured using realtime-PCR. b) MCF7 cells were treated with 200 µM of H 2O2 for different time-points. ER α mRNA level was measured using realtime- PCR and ER α protein level was determined by western blot analysis. Results are obtained from the mean of 3 independent experiments +/- SE.
* * 1.2
1
0.8
H2O2 0 H2O2 0 μM 0.6 H2O2H2O2 100100 μM H2O2H2O2 200200 μM
0.4
mRNA mRNA (Foldrelative untreated to control) 0.2 α ER 0 24 48 72
Hr upon H 2O2 treatment
*:p-value < 0.05
Figure 15: H2O2 induces reversible down-regulation of ER α mRNA level. MCF7 cells were treated with various concentrations of H 2O2 for 24, 48 and 72 h. ER α mRNA level was measured using realtime-PCR. Results are obtained from the mean of 3 independent experiments +/- SE.
80
3A.3.2 New protein synthesis is not required for H2O2-induced down-
regulation of ER α mRNA
To determine if the effect of H2O2 requires de novo protein synthesis, we assessed the ability of H2O2 to repress ER α mRNA level in the presence or absence of protein synthesis inhibitor cycloheximide. MCF7 cells were treated with either 0 or 200 µM
H2O2 with or without 100 µg/ml of cycloheximide co-incubation for 24 h. After 24 h
of treatment, total RNA was extracted and ER α mRNA was monitored by real-time
PCR analysis. The results in Figure 16 show that the inhibition of protein synthesis by
cycloheximide did not prevent H2O2-mediated decrease in the ER α mRNA level. This indicates that the down-regulation of ER α mRNA does not require new protein
synthesis. Blocking protein synthesis with cycloheximide alone led to a sharp
decrease in the mRNA level of ER α in MCF7 cells. The result shows that continuous protein synthesis is required to maintain basal levels of ER α mRNA.
1.2
1
0.8
0.6 control)
0.4
0.2 mRNA level level mRNA untreated to (Fold relative α ER 0
-chxMedium H2O2 0 µM -chxH2 OH2O22 200 200µM µM +chx H2O2+chx 0 µM+chx/H +chx H2O22O2 200 200 µM µM
Treatment
Figure 16: Protein synthesis is not required in H2O2 down-regulation of ER α mRNA level. MCF7 cells seeded for 2 days before treated with either 0 or 100 µg/ml cycloheximide in the presence of 0 or 200 µM of H2O2 for 24 h. ER α mRNA level was measured using realtime-PCR. Results shown are obtained from the mean of 3 independent experiments +/- SE.
81
3A.4 MECHANISM INVOLVED IN H 2O2-INDUCED DOWN-REGULATION
OF ER α
3A.4.1 Role of Akt activation in H 2O2 down-regulation of ER α
Keratinocyte growth factor (KGF) had been reported to down-regulate ER α protein
and mRNA expression level in MCF7 cells (Chang et al., 2009). The authors
identified that Akt was phosphorylated at Ser473 early upon KGF treatment. Blocking
the phosphorylation with PI3K inhibitor, LY294002, could prevent the down-
regulation of ER α protein and mRNA level by KGF. Thus, their results demonstrated that KGF regulated ER α expression through the PI3K/Akt pathway.
Many researchers have reported that H2O2 treatment could also induce phosphorylation of Akt (Martin et al., 2002; Hand and Craven, 2003; Urata et al.,
2006). We asked whether H2O2-mediated down-regulation of ER α could be via a
similar PI3K/Akt pathway. First, MCF7 cells were tested for Akt phosphorylation at
Ser473 residue upon exposure to H2O2. Figure 17a shows that Akt was rapidly
phosphorylated at Ser473 following the treatment of the cells with 200 µM of H2O2.
Within 5 min of H2O2 addition, Akt phosphorylation could be detected. The
phosphorylation peaked at 15 min before it started decreasing and became
undetectable at 60 min post-treatment. LY294002 is an inhibitor of PI3K, a kinase
upstream of Akt phosphorylation (Furuya et al., 2007). Pre-incubation of cells with
different concentrations of LY294002 for 30 min followed by treatment with 200 µM of H2O2 for 15 min resulted in inhibition of Akt phosphorylation in a concentration
dependent manner (Fig. 17b). Twenty micromolar of LY294002 was deemed
sufficient to completely inhibit phosphorylation of Akt caused by H2O2. Figure 17c
shows that Akt phosphorylation was prevented by LY294002 treatment. However,
82 this inhibition was unable to reduce the down-regulation of ER α by H2O2 (Fig. 17d).
Taken together, the results suggest that the repression of ER α by H2O2 is not via the
PI3K/Akt pathway.
a)
H2O2 200 µM, Min 0 5 10 15 30 60
P-Akt -60 kDa (S-473)
Total Akt -60 kDa
b)
LY294002 µM - - 10 20 30 - + + + + H2O2 200 µM P-Akt -60 kDa (S-473)
-60 kDa Total Akt c)
+ + LY294002 20 µM -- - + H2O2 200 µM - +
P-Akt -60 kDa (S-473)
Total Akt -60 kDa
d)
-- + + LY294002 20 µM
H2O2 200 µM - + - +
ER α -65 kDa
β-actin -43 kDa
Figure 17: Inhibition of Akt activation does not prevent ER α down-regulation by H2O2. a) MCF7 cells were seeded for 2 days before being treated with 200 µM of H 2O2 for 5 to 60 min. b) MCF7 cells were pre-incubate with PI3K inhibitor LY294002 for 30 min prior to treatment with 200 µM of H 2O2 for 15 min. c) MCF7 cells were preincubated with LY294002 for 30 min prior to the treatment with 200 µM of H 2O2 for 15 min d) 24 h. Cell lysates were analyzed by western blot. Results shown are representative of 2 independent experiments.
83
3A.4.2 Role of MAPK activation in H 2O2 down-regulation of ER α
Hyperactivation of MAPK has been shown to induce loss of ER α expression in breast cancer cells (Oh et al., 2001). As part of the attempt to define which signal transduction pathways might be involved in down-regulation of ER α mediated by
H2O2, we examined the phosphorylation status of the MAPK members ERK and JNK in MCF7 cells when exposed to 200 µM of H2O2. Figure 18 shows that the two
members of the MAPK were activated early upon the exposure of the cells to H2O2.
ERK 1/2 was activated as early as 10 min upon H2O2 treatment while JNK was starting to be activated after 30 min. Both the MAPK were strongly phosphorylated at
2 h after H2O2 addition.
To determine if their activation are involved in the down-regulation of ER α,
pharmacological inhibitors were used to inhibit their phosphorylation. To block ERK
1/2 activation, U0126 was used. U0126 blocks ERK’s upstream kinase MEK 1/2, thus,
preventing ERK phosphorylation. Twenty micromolar of U0126 was able to
completely inhibit H2O2 activation of ERK 1/2 at 2 h (Fig. 19a). However, blocking
ERK 1/2 activation could not prevent down-regulation of ER α by H2O2 (Fig. 19b &
19c). Next, SP600125 was used to inhibit activation of JNK. Figure 20a shows that
20 µM of SP600125 effectively inhibited phosphorylation of JNK. The inhibition of
JNK was also unable to prevent down-regulation of ER α by H2O2 (Fig. 20b & 20c).
These results show that the activation of the MAPK by H2O2 was not involved in the down-regulation of ER α protein.
84
H2O2 200 µM, Min 0 5 10 15 30 60 120 -42 kDa p-ERK -44 kDa
-42 kDa Total ERK -44 kDa
-45 kDa P-JNK -55 kDa
Total JNK -45 kDa -55 kDa
Figure 18: H2O2 activates MAPK. a) MCF7 cells were seeded for 2 days before treated with 200 µM of H 2O2 for 5 to 120 min. Cell lysates were analyzed by western blot. Results shown are representative of 3 independent experiments. a)
U0126 µM - - 10 20 30
H2O2 200 µM - + + + + -42 kDa p-ERK 1/2 -44 kDa
-42 kDa Total ERK 1/2 -44 kDa
b) U0126 20 µM -- + + H2O2 200 µM - + - + -42 kDa p-ERK 1/2 -44 kDa
Total ERK 1/2 -42 kDa -44 kDa
c) -- + + U0126 20 µM H2O2 200 µM - + - +
ER α -65 kDa
β-actin -43 kDa
Figure 19: The inhibition of ERK 1/2 activation does not prevent ER α down-regulation by H2O2. a) MCF7 cells were seeded for 2 days and preincubated with various concentrations of MEK inhibitor U0126 for 30 min before treated with 0 or 200 µM of H 2O2 for 2 h. b) MCF7 cells were treated with 0 or 20 µM U0126 for 30 min. Cells were then treated with 0 or 200 µM of H 2O2 for 2 h and c) 24 h. Cell lysates were analyzed by western blot. Results shown are representative of 2 independent experiments.
85
a)
SP600125 µM - - 10 20 30
H2O2 200 µM - + + + + -45 kDa p-JNK -55 kDa
Total JNK -45 kDa -55 kDa
b)
SP600125 20 µM -- + + H2O2 200 µM - + - + -45 kDa p-JNK -55 kDa
-45 kDa Total JNK -55 kDa
c)
SP600125 20 µM - - + +
H2O2 200 µM - + - +
ER α -65 kDa
β-actin -43 kDa
Figure 20: Inhibition of JNK activation does not prevent ER α down-regulation by H2O2. a) MCF7 cells were seeded for 2 days and preincubated with various concentrations of JNK inhibitor SP600125 for 30 min before treated with 0 or 200 µM of H 2O2 for 2 h. b) MCF7 cells were treated with 0 or 20 µM SP600125 for 30 min Cells were then treated with 0 or 200 µM of H 2O2 for 2 h and c) 24 h. Cell lysates were analyzed by western blot. Results shown are representative of 2 independent experiments.
86
3A.4.3 Role of caspases in H 2O2 down-regulation of ER α
Caspases are cysteine proteases traditionally known for their role in apoptosis and
inflammation (Launay et al., 2005; Li and Yuan, 2008). They can also be involved in
immune response, cell proliferation, cell differentiation, cell migration and cell shape
(Kuranaga and Miura, 2007). The activation of caspases has been shown to be
involved in the down-regulation of gene expression (Guilherme et al., 2009). H 2O2
has also been reported to activate caspases to repress gene expression (Ma et al., 2003;
Kumar et al., 2007). Here, we ask if caspase could be involved in the down-regulation
of ER α by H 2O2. Executioner caspases mainly caspase 2, caspase 6 and caspase 7
were tested for their activity in MCF7 upon treatment with H 2O2. Results from Figure
21a show that there was no activation of caspase 2, in fact, H2O2 induced a down-
regulation of caspase 2 activity at 24 h after treatment. H2O2 treatment of MCF7 also
led to a lower caspase 6 activity than the untreated control (Fig. 21b). There is no
caspase 3 in MCF7 due to a 47-base-pair deletion in exon 3 of the CASP-3 gene in this particular cell line (Jänicke et al., 1998). Caspase 7 is however present in MCF7.
Since caspase 3 and 7 have been reported to share similar substrate cleavage site
(Chandler et al., 1998), caspase 3 substrate was used to measure caspase 7 activity.
There was a slight activation of 0.2 fold compared to untreated control (Fig. 21c).
However, this increase was small and statistically insignificant. Taken together, the
results show that H 2O2 does not induce the activation of executioner caspases, therefore the role of caspase activation in the down-regulation of ER α protein and
mRNA level was ruled out.
87
a)
1.2 * 1
0.8
control) 0.6 2
O H2O2H2O2 0uM0 µM 2
H 0.4 H2O2H2O2 200uM200 µM
0.2 to no to
0
Caspase 2 relative activity (Fold 4 8 16 24
Hr upon H2O2 treatment
b)
1.2 ** 1
0.8
control) 0.6 2
O H2O2H2O 20uM0 µM 2
H 0.4 H2O2H2O 2200uM200 µM
0.2 to no no to
0
Caspase 6relative activity (Fold 4 8 16 24
Hr upon H2O2 treatment
c)
1.6 1.4 1.2 1
control) 0.8 2
O H2O2H2O2 0uM0 µM 2 0.6 H2O2H2O2 200uM200 µM 0.4
to noH to 0.2 0
Caspase 7 relative activity (Fold 4 8 16 24
Hr upon H2O2 treatment
*: p-value < 0.05
Figure 21: H 2O2 does not activate caspase activity. Cells were seeded for 2 days before exposed to 200 µM of H2O2 for various time points. Capase activity assay for a) caspase 2, b) caspase 6 and c) caspase 7 was carried out using the cell extracts according to the Materials and Methods section. Results are obtained from the mean of 3 independent experiments +/- SE.
88
3A.4.4 Oxidation is involved in the down-regulation of ER α by H 2O2
3A.4.4.1 H2O2 mediates down-regulation of ER α through reversible oxidation
H2O2 is a mild oxidizing agent which does not normally react with DNA, lipids and most proteins. However, H2O2 can act on downstream targets to modulate the activity
of enzymes (e.g. phosphatase and kinases) and transcription factors through selective
oxidation of the thiol (SH) group of cysteine residues (Poole and Nelson, 2008). The
oxidized cysteine residues can be reduced back by intracellular antioxidants such as
glutathione (GSH) or thioredoxin (Trx), or by reducing agents such dithiothreitol
(DTT) or β-mercapthoethanol (BME) (Halliwell and Gutteridge, 2007).
To study whether oxidation is part of the signaling mechanism involved in the down- regulation of ER α expression by H2O2, MCF7 were preincubated with different concentrations of reducing agent DTT. DTT is a strong reducing agent capable of reducing oxidized proteins intracellularly, due to its propensity to form a six- membered ring with an internal disulfide bond. MCF7 cells were preincubated with various concentrations of DTT for 2 h before being treated with 100 or 200 µM of
H2O2 for 24 h. Results show that DTT was able to reverse the down-regulation of
ER α protein by H2O2 at 0.5 mM (Fig. 22a and 22b). The experiment was repeated with 0.5 mM of DTT (Fig. 22c) and mRNA level of ER α was determined. Figure 22d shows that pretreating cells with DTT prevented the repression of ER α mRNA level
by H2O2. The results indicate that the down-regulation of ER α by H 2O2 is mediated through reversible oxidation of components in the ER α expression pathway.
89 a)
H2O2 100 µM - + + + + DTT mM - - 0.1 0.2 0.5
ER α -65 kDa
β-actin -43 kDa
b)
H2O2 200 µM - + + + + DTT mM - - 0.1 0.2 0.5
ER α -65 kDa
β-actin -43 kDa
c)
DTT 0.5 mM - - + +
H2O2 µM 0 200 0 200
ER α -65 kDa
β-actin -43 kDa
90
d)
1.2 *
1
0.8
0.6 H2O2H2O2 0 0µM
control) 0.4 H2O2 200 H2O2 200μM
0.2 mRNA (Fold relative to untreated untreated to relative (Fold mRNA
α 0
ER 0 0.5 DTT (mM)
*: p-value < 0.05
Figure 22: Reducing agent DTT prevents the down-regulation of ER α by H2O2. MCF7 cells were seeded for 2 days before preincubation with various concentrations of reducing agent, DTT for 2 h. Cells were then treated with a) 100 µM and b) 200 µM of H 2O2 for another 24 h before harvesting. Cell lysates were analyzed by western blot. MCF7 cells were preincubated with 0.5 mM of DTT for 2 h before treated with 200 µM of H 2O2 for another 24 h before harvesting for c) western blot analysis and d) realtime PCR analysis. Results shown are representative or obtained from the mean of 3 independent experiments +/-SE.
91
3A.4.4.2 Thiol-oxidant diamide mimics the effects of H 2O2 in down-regulating
ER α expression
Diamide is a thiol-oxidizing agent that modifies or cross-links cysteine sulfhydryl
groups to form cysteine disulfide bridges in proteins (Kosower et al., 1969; Kozlova
et al., 2002). Oxidation by diamide is reversible through reducing agent DTT or BME
(Ward et al., 2000). To further verify the oxidation-mediated down-regulation of ER α,
the thiol-oxidant diamide was used. MCF7 cells were treated with increasing
concentrations of diamide and the ER α protein level was assessed. Figure 23a shows that diamide decreased the level of ER α starting from 0.2 mM and became more
pronounced at 0.5 mM. Next, we then tested the mRNA levels of ER α with 0.5 mM of diamide and found that the decrease in ER α mRNA level was also observed (Fig.
23c). The results show that ER α expression is indeed regulated by a mechanism which is thiol-oxidation dependent.
a)
Diamide mM0 0.05 0.1 0.2 0.3 0.5
ER α -65 kDa
β-actin -43 kDa
92
b)
Diamide mM 0 0.5
ER α -65 kDa
β-actin -43 kDa
c)
* 1.2
1
0.8
0.6
0.4 untreated control) untreated mRNA (Fold relative to relative (Fold mRNA α 0.2 ER
0 0 0.5 Diamide (mM)
*: p-value < 0.05
Figure 23: Thiol-oxidant diamide induces down-regulation of ER α. a) MCF7 cells were seed for 2 days before being exposed to different concentrations of diamide 24 h. Cell extracts were analyzed by western blot. MCF7 cells were treated with 0.5 mM of diamide for 24 h cells extract analyzed by b) western blot and c) realtime PCR. Results shown are representative or obtained from the mean of 3 independent experiments +/-SE.
93
3A.5 EFFECTS OF ER α DOWN-REGULATION ON ER RESPONSE GENES.
It has been shown that in response to estrogen stimulation, ER α protein is rapidly down-regulated via proteosomal degradation pathway to limit the extent of the estrogen signaling (Tateishi et al., 2004; Alarid et al., 1999; Nawaz et al., 1999).
Estrogen response genes were however up-regulated several folds. In this section, we evaluated the impact of ER α down-regulation by H2O2 on estrogen-dependent responses.
3A.5.1 Down-regulation of ER α by H 2O2 inhibits estrogen response element (ERE) activity
In the first set of experiments, we studied the effects of H2O2 on the general transcriptional activity of ER. MCF7 cells were transiently transfected with a plasmid containing luciferase reporter gene whose expression is driven by an estrogen response element (ERE). Estrogen receptor, upon binding to estrogen will be recruited to the ERE and drive the transcription of the luciferase gene. The amount of luciferase was subsequently assessed and used to represent the level of ER activity in the cells.
In the study, MCF7 cells transfected with ERE luciferase plasmid were estrogen- deprived overnight after transfection by growing the cells in phenol-free RPMI supplemented with 10 % charcoal-stripped FBS (CS FBS). Cells were then treated with 100 µM of H2O2 for 16 h to down-regulate ER α. After H 2O2 treatment, cells were incubated with 10 nM of E2 or DMSO control for 2, 4 or 6 h before harvesting for luciferase assay. Western blot analysis in Figure 24a shows that ER α expression
level was decreased in the cells treated with 100 µM of H2O2 for 16 h. Notably, down- regulation of ER α by H2O2 did not affect the basal activity of ERE (Fig. 24b). This is expected as the cells were grown in CS FBS and were deprived of estrogen, thus, in
94 the presence or absence of ER α, there is minimum stimulation of ERE. However,
when E2 was added to cells expressing ER α, there was a strong activation of ERE,
about 12 folds increase in the activity at 6h (Fig. 24b). In comparison, H2O2-treated cells which have much lower levels of ER α had less than 3 folds increase in ERE
activity in response to E2 stimulation. The results show that the down-regulation of
ER α by H2O2 decreased the activation of ERE by estrogen. a) H O µM0 100 2 2 ER α -65 kDa β-actin -43 kDa b)
* 1800
1600 * 1400
1200
1000 2h after E2 * 800 4h after E2 6h after E2 600
400
ERE activity (Luciferaseactivity ERE RFU/renlla RFU) 200
0
MediumDMSOMedium+E2 DMSO+ E2 H2O2H2O2 100100uMµM H2O2H2O2 100uM100 µM +E2 + E2
Treatment
*: p-value < 0.05
Figure 24: H2O2 inhibits total ERE activity induced by E2. a) MCF7 cells were seeded for 1 day before changing to phenol-free RPMI supplemented with 10 % charcoal-stripped FBS. Cells were treated with 100 µM H 2O2 for 16 h and harvested for western blot analysis. b) MCF7 cells were seeded overnight and transfected with 1 µg of ERE luciferase plasmid together with 0.1 µg of renilla plasmid using SuperFect as described in the Materials and Methods section. Cells were recovered in phenol-free RPMI supplemented with 10 % charcoal-stripped FBS after 3 h of transfection. On day three, cells were treated with 100 µM of H2O2 for 16 h before exposure to 10 nM of E2 for 2, 4 or 6 h. Dual luciferase assay was carried out according to the Materials and Methods section. Results obtained are representative of 3 independent experiments.
95
3A.5.2 H2O2 down-regulates ER α response genes
Progesterone receptor (PR) and c-Myc gene expression have been reported to be
regulated by ER α (Kastner et al., 1990; Dubik and Shiu, 1988). To determine if the
expression of these two genes are under the control of by ER α in the MCF7 cells, cells were estrogen-deprived by growing overnight in phenol-free RPMI supplemented with 10 % CS FBS. Lane 1 of Figure 25a shows that PR and c-Myc protein were hardly expressed in the absence of estrogen. When E2 was added to the cells for 24 and 48 h, PR and c-Myc protein expression were increased (Fig. 25a). The effects of E2 on these two proteins were investigated through a time course analysis.
MCF7 cells were estrogen-deprived overnight before E2 was added for of 4, 8, 24 and
48 h. The c-Myc expression increased rapidly upon addition of E2. The protein level increased from 4 h and was sustained up to 24 h after which the expression declined at
48 h (Fig. 25b). On the other hand, PR protein expression responded slower to E2 treatment. The protein became detectable only at 24 h and increased up to the 48 h time-point tested (Fig. 25b). These results show that PR and c-Myc are indeed ER α
responsive genes. Next, we examined the effects of H2O2 on the level of PR and c-
Myc in MCF7 cells. Figure 25c shows that concentration-dependent down-regulation of ER α by H2O2 resulted in decreased expression of both PR and c-Myc. c-Myc
decreased more than PR and this is most probably due to a longer half-life of PR as
compared to c-Myc (Fig. 26a).
To determine the effects of H2O2 on the ability of E2 to stimulate estrogen response
genes, estrogen-deprived MCF7 cells were treated with 0, 100 and 200 µM of H2O2 for 16 h and were given either 10 nM E2 or DMSO for another 4 h before harvesting.
Western blot analysis shows that in the absence of E2, c-Myc protein level was low
(Fig. 26b). Addition of E2 induced c-Myc expression in control cells; but was greatly
96
reduced in H 2O2 treated cells (Fig. 26b). The results indicate that the down-regulation of ER α affected estrogen’s ability to induce ER α responsive gene expression.
a)
24h 48h 10nM E2 - + - +
ER α -65 kDa
PR -120 kDa
c-Myc -63 kDa
β-actin -43 kDa
b)
+10nM E2 Hr 04 8 24 48
PR -120 kDa
c-Myc -63 kDa
β-actin -43 kDa
c)
H2O2 µM 0 25 50 100 200
ER α -65 kDa
PR -120 kDa
c-Myc -63 kDa
β-actin -43 kDa
Figure 25: H2O2 decreases ER α response genes, PR and c-Myc expression. a) MCF7 cells were seeded overnight in complete medium. Cells were changed to phenol-free RPMI supplemented with 10 % charcoal-stripped FBS for another 2 days. Cells were exposed to 10 nM E2 for 24 and 48 h. b) MCF7 cells were seeded overnight in complete medium. Cells were changed to phenol-free RPMI supplemented with 10 % charcoal-stripped FBS for another 2 days. Cells were exposed to 10 nM E2 for various time-points. c) MCF7 cells were seeded for 2 days before treated with various concentrations of H 2O2 for 24 h. Cell extracts were analyzed by western blot.
97
a)
Cycloheximide 100 µg/ml
Hr 0 2 4 8 16 24 48
PR -120 kDa
c-Myc -63 kDa
β-actin -43 kDa
b)
Medium +E2 10nM
H2O2 µM 0 100 200 0 100 200
ER α -65 kDa
-63 kDa c-Myc
β-actin -43 kDa
Figure 26: H2O2 decreases the ability of estrogen to induce c-Myc expression. a) MCF7 cells were seeded for 2 days before treated with 100 µg/ml of cycloheximide. Cells were harvested at various time-points after treatment. b) MCF7 cells were seeded for overnight in complete medium. Cells were changed to phenol-free RPMI supplemented with 10 % charcoal-stripped FBS for 24 h before treated with H 2O2 for 16 h. Cells were then exposed to 10 nM E2 for 4 h. Cell extracts were analyzed by western blot. Results shown are representative of 2 independent experiments.
98
3A.6 15-DEOXY-DELTA-12,14-PROSTAGLANDIN J2 (15D-PGJ2) DOWN-
REGULATION OF ER α IS VIA ROS
We have shown in the previous sections that the addition of exogenous ROS such as
H2O2 and peroxynitrite can down-regulate ER α expression. Next, we want study the
physiological relevance of ROS in the down-regulating ER α, by treating the cells with
an agent that is known to produce ROS intracellularly. The compound 15-Deoxy-
Delta-12,14-prostaglandin J2 (15d-PGJ2) is a natural ligand for peroxisome
proliferator-activated receptor gamma (PPAR γ), a member of the nuclear receptor family (Forman et al., 1995; Murphy and Holder, 2000). Activation of PPAR γ is a multi-step process involving the binding of ligand to PPAR γ, heterodimerization with
retinoic X receptor and binding to PPAR γ response element. 15d-PGJ2 was shown to down-regulate ER α via a PPAR γ independent but proteosomal dependent pathway
(Lecomte et al., 2008; Qin et al., 2003). Other than its role as a PPAR γ ligand, reports
have shown that 15d-PGJ2 induces ROS in cells when added exogenously (Kim et al.,
2008; Kim et al., 2009; Shin et al., 2009). Here, we ask the question whether ROS
could be the intermediate that led to ER α down-regulation in MCF7 cells treated with
15d-PGJ2.
3A.6.1 Morphology and growth of MCF7 cells exposed to 15d-PGJ2
We tested the effects of increasing concentrations of 15d-PGJ2 on the morphology and growth of MCF7 cells. Figure 27a shows the morphology of MCF7 cells treated with 15d-PGJ2 after 24 h. Cells looked slightly enlarged and sparse compared to untreated control. Next, MCF7 cells were incubated with increasing concentrations of
15d-PGJ2 for 24 h and cell density was estimated by crystal violet staining. The time at which 15d-PGJ2 was added was regarded as time 0 h. Cell growth was expressed as
99 percentage relative to the time of treatment. 15d-PGJ2 exhibited a concentration- dependent inhibition of cell growth. Density of cells treated with 0, 5, 10 and 20 µM of 15d-PGJ2 for 24 h was at 202, 166, 121, 120 % respectively, relative to the time treatment (Fig. 27b).
a)
Medium 5 µM 10 µM 20 µM
b)
250.0
200.0 *
150.0 **
100.0
50.0
Cell density (% relative to time time (% to treatment) relative ofdensity Cell 0.0 0 5 10 20 15d-PGJ2 (µM)
*: p-value < 0.05 (relative to time of trigger)
Figure 27: MCF cell survival upon 15d-PGJ2 treatment. MCF7 cells were seeded for 2 days before treated with various concentrations of 15d-PGJ2 for 24 h. a) Morphology of the cells was captured under phase-contrast microscope at 10x magnifications. b) Cell density and survival were estimated by staining the cells with crystal violet 24 h after treatment. Results shown were obtained from the mean of 2 independent experiments +/- SE. 100
3A.6.2 15d-PGJ2 down-regulates ER α at the protein and mRNA level
Next, we determined the effects of 15d-PGJ2 on the expression of ER α at 24 h after
treatment. Western blot analysis showed that 15d-PGJ2 down-regulated ER α protein in a concentration-dependent manner. ER α protein level was decreased sharply with
5µM of 15d-PGJ2 (Fig. 28a). At higher concentrations of 10 and 20 µM of 15d-PGJ2,
ER α protein expression was completed abolished. Realtime analysis of ER α mRNA
shows a decrease in the mRNA level (Fig. 28b). The transcript level of ER α was
observed to be at 33, 5 and 10 % with 5, 10 and 20 µM of 15d-PGJ2 respectively after
24 h of treatment. To determine whether the 15d-PGJ2-induced down-regulation of
ER α was mediated by PPAR γ, cells were pretreated with PPAR γ antagonist GW9662
(1 to 10 µM) before exposure to 15-PGJ2. GW9662 is a potent, irreversible and
selective antagonist that prevents the activation of PPAR γ by covalently modifying a
cysteine residue in the ligand-binding site of PPARγ (Leesnitzer et al., 2002; Seargent
et al., 2004). Results show that in cells co-treated with GW9662 and 15d-PGJ2, the
reduction in the level of ER α protein still occurred as compared to those treated with
15d-PGJ2 alone (Fig. 28c). The results suggest that the down-regulation of ER α
mediated by 15d-PGJ2 was triggered by a PPAR γ-independent mechanism.
101 a)
15d-PGJ2 µM 0 5 10 20
ER α -65 kDa
β-actin -43 kDa
b)
1.2
1
0.8
0.6
control) * 0.4
0.2 *
mRNA (Fold relative to untreated untreated to relative (Fold mRNA * α 0 ER 0 5 10 20 15d-PGJ2 µM
*: p-value < 0.05 (relative to untreated control)
c) 15d-PGJ2 10 µM - + + + +
GW 9662 µM - - 1 5 10
ER α -65 kDa
β-actin -43 kDa
Figure 28: 15d-PGJ2 down-regulates ER α protein and mRNA levels. MCF7 cells were seeded for 2 days before treated with various concentrations of 15d-PGJ2 for 24 h a) ER α protein expression was determined by western blot while b) mRNA level was determined with realtime-PCR. c) MCF7 cells were preincubated with various concentrations of GW9662 for 30 min before treated with 10 µM of 15d-PGJ2. Results shown are obtained from the mean of 3 independent experiments +/- SE.
102
3A.6.3 15d-PGJ2 produces ROS in MCF7 cells
ROS produced by 15d-PGJ2 was measured by flow cytometry FITC channel using a redox sensitive dye CM-H2DCFDA which fluoresces under oxidation by intracellular
H2O2 and other reactive species. Intracellular ROS were quantified by determining the mean of DCF fluorescence intensity. ROS level increased rapidly when MCF7 cells were treated with 10 µM of 15d-PGJ2. ROS could be detected as early as 1 h after treatment and the increase in ROS was sustained for up to 4 h of treatment (Fig. 29).
Quantification of two experiments shows that 15d-PGJ2 induced an increase in intracellular ROS with an increase of 56 % at 1h, 43 % at 2 h, 102 % at 3 h and 65 % at 4 h relative to untreated control.
The increased in the DCF fluorescence, a measurement of intracellular oxidation and
ROS production, can also be represented as a shift in DCF fluorescence to the right of the x-axis of the Fluorescence-activated cell sorting (FACS) histogram. As a positive control, 1 mM of H2O2 was added to another set of MCF7 cells 1 h prior to harvesting of the cells. Figure 30a shows that both H 2O2 and 15d-PGJ2 treatment caused the
DCF fluorescence to shift to the right on the x-axis of the FACs histogram, indicating
increase ROS production. The rightward shift caused by 15d-PGJ2 was more than the
rightward shift induced by H 2O2, which signifies that 10 µM of 15d-PGJ2 resulted in
greater intracellular oxidation than 1 mM of H 2O2 added extracellularly.
Next, we tested the ability of antioxidant N-acetyl-cysteine (NAC) to scavenge the
ROS produced by treatment with 15d-PGJ2. The cells were preincubated with 0 or 5
mM of NAC for 2 h prior to treatment with 10 µM of 15d-PGJ2 for another 2 h. Flow
cytometry result shows that preincubating the cells with 5mM of NAC caused the
DCF fluorescence to shift to the left on the x-axis of the FACs histogram, indicating
decreased ROS production (Fig. 30b). Figure 30c shows the summary of the DCF
103 mean data from three repeated experiments. This set of results shows that NAC preincubation prevented ROS from being elevated in the cells treated with 15d-PGJ2.
250.0 *
200.0 * * * 150.0
100.0 untreated control) untreated
50.0 DCF fluorescence (% relative (% to relative fluorescence DCF
0.0 1 2 3 4 Hr upon 15d-PGJ2 addition
*: p-value < 0.05 (relative to untreated control)
Figure 29: 15d-PGJ2 produces ROS in MCF7 cells . MCF7 cells were seeded for 2 days before treated with 10 µM of 15d-PGJ2 for 1 to 4 h. ROS levels in cells were detected using flow cytometry with CM-H2DCFDA flurogenic probe according to the Materials and Methods section. Results shown are obtained from the mean of 2 independent experiments +/- SE.
104 a)
H2O2 1mM
15d-PGJ2 10 µM
Control
b)
15d-PGJ2 10 µM + 5mM NAC
15d-PGJ2 10 µM 5mM NAC
Control
105 c)
* 200 180 160 140 120 100 MediumMedium 80 15d-PGJ215d-PGJ2 10 µM 60 medium control) medium 40 20 DCF Fluorescence (% relative to relative (% Fluorescence DCF 0 0 5 NAC (mM)
*: p-value < 0.05
Figure 30: NAC scavenges 15d-PGJ2-induced ROS. a) Cells were treated with 10 µM of 15d-PGJ2 and 1 mM of H2O2 for 1 h. ROS levels in cells were detected using flow cytometry with CM-H2DCFDA flurogenic probe. b) Cells were preincubated with 0 or 5 mM of NAC for 2 h before treated with 15d-PGJ2 for 2 h. ROS levels in cells were detected using flow cytometry with CM-H2DCFDA flurogenic probe. c) Graphical representative of (b). Results shown are representative of 3 independent experiments.
106
3A.6.4 Scavenging of ROS by NAC restores cell growth and ER α
expression
To test if ROS was the intermediate involved in growth inhibition and the down-
regulation of ER α by 15d-PGJ2, NAC was used to scavenge the ROS produced by
15d-PGJ2. Figure 31a shows the morphology of cells treated with 15d-PGJ2 with or without NAC preincubation. The cells preincubated with NAC and then treated with
15d-PGJ2 have similar morphology and cell density compared to control cells. This shows that scavenging ROS by NAC could restore cell growth. In addition, ROS scavenging by NAC prevented 15d-PGJ2-mediated down-regulation of ERα protein
(Fig. 31b), as well as mRNA level (Fig. 31c).
In summary, 15d-PGJ2 induces repression of ER α expression at the protein and
mRNA level. Large amount of ROS was produced early upon treatment and
scavenging of the ROS could prevent decrease of ER α protein and mRNA level.
Taken together, the down-regulation of ER α by 15d-PGJ2 was mediated through ROS.
a)
Medium PGJ2 10 µM NAC 5 mM NAC 5mM + PGJ2 10 µM
107 b)
NAC 5 mM - - + + 15d-PGJ2 10 µM - + - +
ER α -65 kDa
β-actin -43 kDa
c)
1.2 *
1
0.8
0.6
PGJ2Medium 0 0.4 PGJ215d-PGJ2 10 10 µM
0.2 mRNA (Fold relative to medium medium control) to relative (Fold mRNA
α 0
ER 0 5 NAC (mM)
*: p-value < 0.05
Figure 31: Scavenging of ROS by NAC restores ER α expression. a) Cells were preincubated with either 0 or 5 mM of NAC for 2 h before treated with 10 µM of 15d-PGJ2 for 24 h. a) Morphology of the cells were captured under phase contrast microscope. b) ER α protein level was analyzed by western blot. c) ER α mRNA level was measured using realtime-PCR. Results shown are obtained from 3 independent experiments +/- SE.
108
Figure 32 summarized the findings regarding H2O2 down-regulation of ER α. We
found that H2O2 represses ER α expression through a reversible oxidation pathway.
Diamide, a thiol-oxidant could also inhibit ER α expression. Drugs that down-regulate
ER α could do so via the production of H2O2 or other ROS. The suppression of ER α
expression is reversible by preincubation with reducing agent DTT and by scavenging
ROS. Down-regulation of ER α protein at 8 h was preceded by decrease in ER α
mRNA levels at 4 h. The repression of ER α expression results in an inhibition of ER α
response gene expression which could have contributed to growth arrest observed.
15d-PGJ2 H O Drugs 2 2 DTT ROS NAC reversible-oxidation Diamide ?
Tumor growth ER mRNA (4h)
ER response genes ER protein (8h)
Figure 32: Summary of the down-regulation of ER α by H2O2.
109
CHAPTER 3B: RESULTS – REDOX REGULATION OF NHE1
3B.1 DOWN-REGULATION OF NHE1 BY H 2O2
In this section, we investigated another gene that has been shown to be down- regulated by H2O2. Previously, our lab has reported that H2O2 repress the activity of the 1.1kb mouse NHE1 promoter in L6 rat myoblast through an oxidation and caspase-dependent mechanism. In this section, we would like to determine the response element of H2O2 and address the mechanism involved in the down-
regulation of NHE1 promoter activity by H2O2.
3B.1.1 Identification of crucial region in the promoter of NHE1
We have previously established that NHE1 is a redox-regulated gene (Akram et al.,
2006; Kumar et al., 2007). Using L6 cells with a stably transfected full length 1.1kb of NHE1 promoter (L6 1.1kb cells) we were able to reproduce the results previously published. Figure 33 shows that the 1.1kb of NHE1 promoter activity was decreased in a concentration-dependent manner at 24 h after H2O2 treatment.
110 a)
120
100 *
80 * control) 2 O 2
H 60
40
relative to to no relative * 20 * * 1.1kb NHE1 promoter activity in L6 cells cells in L6promoter activity (% NHE1 1.1kb
0 0 10 25 50 100 200
H2O2 µM
Figure 33: H 2O2 down-regulates NHE1 full length promoter activity. L6 myoblasts stably transfected with 1.1kb NHE1 promoter were subjected to various concentrations 10, 25, 50, 100 and 200 µM of H2O2 for 24 h. Cells were harvested and assessed for NHE1 promoter activity using luciferase assay as described in Materials and Methods section. Results represent mean of four independent experiments done in duplicates +/- SE.
In order to understand which region of the NHE1 promoter is important for its transcription, NHE1 promoter constructs with different 5’ end deleted were transfected into L6 cells together with a plasmid encoding for renilla as a transfection control. Figure 34a shows the different 5’ deletion constructs and their corresponding names. Figure 34b illustrates the basic principle of the luciferase assay. Briefly, the promoter of NHE1 is linked upstream of a luciferase gene. The plasmid construct will be introduced into the cells via transfection. Any signal that leads to an increase in
NHE1 promoter activity will result in an increase of luciferase enzyme production, which can be quantitatively measured by luciferase assay (as described in Materials and Methods section).
111
Wild type L6 cells were transfected with various 5’ deletion constructs of NHE1 mouse promoter. The promoter activity of NHE1 for the different constructs was similar from 1.1kb up to 0.15kb (Fig. 34c). There was some increase in the promoter activity at the 0.18kb region suggesting that the region between 0.18kb and 0.15kb contains a possible enhancer element involved in the transcription of NHE1. The most striking feature was that removal of a stretch of 33 bp nucleotides from the 5’ end of the 0.15kb region resulted in an 85% decrease in the promoter activity. This indicates that the 33 bp region at the 5’ end of the 0.15kb promoter most likely contains important transcription factor binding site for NHE1 gene transcription in cells growing in 10% FBS.
a)
TATA Luciferase gene 1.1kb
TATA Luciferase gene 0.9kb
TATA Luciferase gene 0.5kb TATA Luciferase gene 0.2kb
TATA Luciferase gene 0.18kb
TATA Luciferase gene 0.15kb 0.12kb TATA Luciferase gene
b)
NHE1’s Trancription factors
TATA Luciferase gene Luciferase
Nucleus
cytosplasm
112 c)
160 *
140
120
100
80
60
40 1.1kb full length length full promoter) 1.1kb
20
NHE1 promoter activity to activity relative (% promoter NHE1 * 0 pXP-1.11.1pXP-0.9 0.9pXP-0.5 0.5pXP-0.2 0.2pXP-0.18 0.18pXP-0.15 0.15pXP-0.12 0.12
NHE1 promoter constructs
*: p-value < 0.0.5
Figure 34: 0.15kb promoter is basal promoter required for full NHE1 promoter activity. a) Different 5’ deletion constructs of mouse NHE1 promoter. b) Basic principle of luciferase assay. c) L6 wild type cells were transiently transfected with the pXP-1.1, pXP-0.9, pXP-0.2, pXP-0.18, pXP-0.15 and pXP-0.12 plasmids. Cells were co-transfected with renilla plasmid for normalization. Results are obtained from the mean of 3 independent experiments +/- SE.
113
3B.1.2 H2O2 down-regulates 0.15kb proximal NHE1 promoter activity
3B.1.2.1 Regulation of NHE1 mRNA level by H 2O2 in NIH 3T3 cells
Having determined that the minimal response element of the NHE1 promoter that
activates full activity lies on the 0.15kb promoter, we next use a cell line that have a
stably transfected 0.15kb mouse NHE1 promoter (NIH 3T3 0.15kb cells) to assess if
the down-regulation of NHE1 mRNA could be correlated with the inhibition of the
0.15kb region of the promoter.
First, we determine the effect of H2O2 on mRNA of NHE1 in NIH 3T3 0.15kb cells.
Figure 35a shows that in 24 h, H2O2 down-regulated NHE1 mRNA transcript level to
70 % and 50 % that of control in the presence of 35 µM and 50 µM of H2O2 respectively. In a time course analysis of the effect of H2O2, we found that the level of
NHE1 mRNA started decreasing at 15 h post treatment with 35 µM and 50 µM of
H2O2 (Fig. 35b).
a)
1.2 2 O
2 1
0.8 * * 0.6 control)
0.4
0.2 NHE1 mRNA (Fold relative noH relative (Fold mRNA NHE1
0 0 25 35 50 H2O2 (µM)
114 b)
1.4
1.2 control) 2 O 2 1
0.8 * * 5h * 0.6 9h * 15h 24h 0.4
0.2 NHE1 mRNA (Fold relative to relative noH (Fold mRNA NHE1 0 0 35 50
H2O2 (µM)
Figure 35: H2O2 down-regulates NHE1 mRNA in a concentration- and time-dependent manner . a) NIH 3T3 cells were treated with 25, 35 and 50 µM of H 2O2 for 24 h and harvested for mRNA isolation. b) NIH 3T3 cells were treated with 35 and 50 µM of H 2O2 for 5, 9, 15 and 24 h. Cells were harvested for mRNA isolation. The level of NHE1 mRNA was determined from realtime PCR. Results shown are obtained from the mean of 3 independent experiments +/- SE.
3B.1.2.2 Regulation of NHE1 0.15kb promoter by H 2O2 in NIH 3T3 cells
Next, we want to understand how the promoter activity of 0.15kb was affected by
H2O2 treatment. Results show that H2O2 down-regulated NHE1 0.15kb promoter activity in a concentration-dependent manner, from 100 % to 80, 37, 19, 15 and 14 % for 10, 25, 35, 50 and 100 µM of H2O2 respectively (Fig. 36a). The promoter activity
of NHE1 first decreased slightly at 1 h after H2O2 treatment, but it recovered at 3 h.
The promoter activity started to decrease again at 9 h and the repression was sustained up to 24 h (Fig. 36b). To determine if the down-regulation of NHE1 0.15kb promoter activity by H2O2 is reversible, cells were allowed to grow for up to 72 h after H2O2
treatment. Cells were collected at every 24 h interval and assessed for promoter
115 activity. The results in Figure 36c shows that activity of NHE1 promoter started to recover 48 h after treatment and continued to recover at 72 h. This set of results demonstrated that the 0.15kb NHE1 promoter is indeed down-regulated by H2O2 and this repression is reversible, similar to what is seen with NHE1 mRNA level.
a)
120
100 control) 2 O 2 80
60
* 40
* 20 ** 0.15kb NHE1 promoter activity 0.15kb activity NIH in NHE1promoter 3T3 cells (% relative relative no H (% to cells 3T3 0 0 10 25 35 50 100
H2O2 (µM)
b)
140
120 control) 2 100 O 2 * * 80
60
40 * ** 20
0 0 1 3 5 9 12 18 24 cells (% relative no H (% to relative cells
0.15kb NHE1 promoter activity in 3T3 promoter NIH activity NHE1 0.15kb Hr after treatment with 50 µM of H 2O2
116
c)
70.0
60.0 **
50.0 control) 2 O 2 40.0
30.0
20.0 (% relative relative H no to (% 10.0 0.15kb 3T3 activity NIH cells in NHE1promoter 0.0 24 48 72
Hr after treatment with 50 µM of H 2O2
*: p-value < 0.05 (relative to no H 2O2 control)
**: p-value < 0.05
Figure 36: H2O2 down-regulates NHE1 0.15kb promoter in a concentration- and time- dependent manner. a) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was treated with various concentrations of H 2O2 for 24 h. b) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was treated with 50 µM of H2O2 for various time-points. c) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was treated with 50 µM of H2O2 for 24, 48 and 72 h. Cells were assessed for NHE1 promoter activity using luciferase assay. Results shown are obtained from the mean of 3 independent experiments +/- SE.
117
3B.1.3 Oxidation is involved in the down-regulation of 0.15kb proximal
NHE1 promoter activity and endogenous mRNA level by H 2O2
3B.1.3.1 H2O2 mediates down-regulation of 0.15kb NHE1 promoter activity and mRNA level through reversible oxidation
Here, we determine the nature of the down-regulation of the 0.15kb promoter activity by H2O2. NIH 3T3 cells were preincubated with NAC, a direct ROS scavenger and
precursor of GSH, for 2 h before treatment with H2O2 for another 24 h. Preincubation
with NAC completely prevented the down-regulation of the 0.15kb promoter activity
at all concentrations used (Fig. 37a). The mRNA level of NHE1 was also restored by
NAC preincubation (Fig. 37b). Cells pretreated with reducing agent DTT have a
concentration-dependent restoration of 0.15kb NHE1 promoter activity, starting from
0.2 mM of DTT. DTT at concentration of 0.5 mM prevented the repression of 0.15kb
NHE1 promoter activity by up to 80% and it also prevented the down-regulation of
NHE1 mRNA level by H2O2 (Fig. 38a and 38b). We used another reducing agent
BME to test the hypothesis that down-regulation of NHE1 promoter activity and
mRNA level is indeed a reversible oxidation event. NIH 3T3 cells were preincubated
with BME for 2 h before being treated with H2O2 for another 24 h. Figure 39a shows
that BME prevented the repression of 0.15kb NHE promoter activity starting from
0.5mM. Promoter activity of cells was 60, 90 and 100 % for cell pretreated with 0.5, 1
and 2 mM of BME respectively. The mRNA level of NHE1 treated with 1 mM of
BME was prevented from down-regulated by H2O2 (Fig. 39b).
Taken together, this set of data showed that the down-regulation of 0.15kb NHE1
promoter activity and NHE1 mRNA level is a reversible oxidation event which can be
blocked by NAC or reducing agents.
118 a)
120.0 * * * 100.0 * *
80.0 control) 2 O 2 60.0 H2O2H2O2 50uM 50 µM
40.0
(% relative H relative no (% 20.0
0.0
0.15kb NHE1 promoter activity 0.15kb 3T3 NIH activity cells in NHE1 promoter 0 1 2 5 10 20 NAC (mM)
*:p-value< 0.05 (Relative to H 2O2 treated cells without NAC) b)
1.2 *
1
0.8 treated control) 2 0.6 O 2 H2O2Medium 0
0.4 H2O2H2O2 50 50 µM
0.2 relativenotoH NHE1 mRNA NHE1mRNA in NIH 3T3cells (Fold 0 0 5 NAC (mM)
*:p-value< 0.05
Figure 37: Scavenging ROS with NAC prevents H 2O2-induced down-regulation of 0.15kb NHE1 promoter activity and NHE1 mRNA level. a) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was preincubated with various concentrations of NAC for 2 h before treated with 50 µM H 2O2 for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay. b) NIH 3T3 cells were preincubated with 5 mM of NAC for 2 h before treated with 50 µM of H2O2 for 24 h. Cells were harvested for realtime PCR analysis. Results shown are obtained from the mean of 3 independent experiments +/- SE.
119 a)
100.0 * * 90.0
80.0 *
70.0 control) 2
O 60.0 2
50.0 H O 50 µM 40.0 H2O22 2 50uM 30.0
(% relative norelative H to (% 20.0
10.0
0.15kbNHE1 promoter activity 0.15kbNHE1 3T3activity NIH cells in promoter 0.0 0 0.05 0.1 0.2 0.5 1 DTT (mM)
*:p-value< 0.05 (Relative to H 2O2 treated cells without DTT) b)
1.2 * 1 control) 2
O 0.8 2
0.6 h2o2H2O 0 2 0µM
h2o2H2O 502 50 µM 0.4
0.2 NHE1 mRNA in NIH 3T3 cells 3T3NIH cells in mRNA NHE1 (Fold relative relative H no to (Fold 0 0 0.5 DTT (mM)
*:p-value< 0.05
Figure 38: Reducing agent DTT prevents H 2O2-induced down-regulation of 0.15kb NHE1 promoter activity and NHE1 mRNA level. a) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was preincubated with increasing concentrations of DTT for 2 h before treated with 50 µM of H2O2 for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay b) NIH 3T3 cells were preincubated with 0.5 mM of DTT for 2 h before treated with 50 µM of H2O2 for 24 h. Cells were harvested for realtime PCR analysis. Results shown are obtained from the mean of 3 independent experiments +/- SE.
120 a)
120.0 *
100.0 *
80.0 control) 2 * O 2 60.0
H2O2H2O2 50 µM 50uM 40.0
20.0 (% relative relative no H to (%
0.0
0.15kbNHE1 promoter activity 0.15kbNHE1 3T3 activity NIH c3lls in promoter 0 0.1 0.2 0.5 1 2 BME (mM)
*:p-value< 0.05 (Relative to H 2O2 treated cells without BME) b)
1.2 *
1 control) 2
O 0.8 2 H2O2H2O2 0 0µM H O 50 µM 0.6 H2O22 2 50
0.4
0.2 NHE1 mRNA in NIH 3T3 cells 3T3 NIH cells in mRNA NHE1
(Fold relative relative no H to (Fold 0 0BME mM 1
*:p-value< 0.05
Figure 39: Reducing agent BME prevents H 2O2-induced down-regulation of 0.15kb NHE1 promoter activity and NHE1 mRNA level. a) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was preincubated with increasing concentrations of BME for 2 h before treated with 50 µM of H2O2 for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay. b) NIH 3T3 cells were preincubated with 1 mM of BME for 2 h before treated with 50 µM of H2O2 for 24 h. Cells were harvested for realtime PCR analysis. Results shown are obtained from the mean of 3 independent experiments +/- SE.
121
3B.1.3.2 Thiol-oxidant diamide mimics the effects of H 2O2 to down-regulate
the 0.15kb NHE1 promoter activity and mRNA expression level
To further verify the oxidant-mediated down-regulation of NHE1, the thiol-oxidant diamide was used. Figure 40a showed a concentration-dependent decrease in 0.15kb
NHE1 promoter activity which starts from 150 µM of diamide. Promoter activity
decreased to 60 % and 30 % that of control when 200 and 250 µM of diamide was used respectively. In the presence of 200 µM diamide, the endogenous mRNA level
of NHE1 was also decreased by 40 % after 24 h of treatment (Fig. 40b).
a)
120
100
80 *
* 60 *
40 (% relative control) relative untreated to (% 20 NHE1 0.15kb promoter activity in 3T3promoter NIH activity cells 0.15kb NHE1 0 0 50 100 150 200 250 Diamide (µM)
*:p-value< 0.05 (Relative to no diamide treatment)
122 b)
1.2 *
1
0.8
0.6
0.4
0.2 relative untreated to control)
NHE1 mRNA NHE1mRNA in NIH 3T3cells (Fold 0 0 200 Diamide ( µM)
*:p-value< 0.05
Figure 40: Thiol-oxidant diamide down-regulates NHE1 0.15kb promoter activity and NHE1 mRNA level. a) NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was treated with increasing concentrations of diamide for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay. b) NIH 3T3 cells were treated to 200 µM of diamide for 24 h. Cells were harvested for realtime PCR analysis. Results showed are obtained from the mean of 3 independent experiments +/- SE.
123
3B.1.4 Peroxynitrite down-regulates 0.15kb NHE1 promoter activity
Peroxynitrite is a strong oxidant and can react directly with the electron-rich groups such as sulfhydryls, iron-sulfur centers and zinc-thiolates (Crow et al., 1995; Kuhn et al., 1999; Soum et al., 2003). Peroxynitrite has been shown to inhibit gene expression by inhibiting activation of NF κB (Soum et al., 2003). Here, we tested whether
extracellularly added peroxynitrite could also down-regulate the activity of 0.15kb
NHE1 promoter. Figure 41 shows a concentration repression of this promoter activity.
Thus, similar to H2O2, peroxynitrite also has a negative effect on 0.15kb NHE1 promoter activity.
120
100
80 *
60
40
*
20 (% relative control) relative no peroxynitrite to (% *
0.15 kb NHE1 promoter activity in 3T3 promoter NIH activity cells kbNHE1 0.15 * 0 0 100 200 500 1000 2000 Peroxynitrite (µM)
*:p-value< 0.05 (Relative to no peroxynitrite treated control)
Figure 41: Peroxynitrite down-regulates 0.15kb NHE1 promoter activity. NIH 3T3 cells stably transfected with 0.15kb NHE1 promoter was treated with various concentrations of peroxynitrite for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay. Results shown are obtained from the mean of 2 independent experiments +/- SE.
124
3B.1.5 Role of caspases in H2O2-mediated down-regulation of 0.15kb
NHE1 proximal promoter activity
In the report previously published by our lab (Kumar et al., 2007), caspase activation
was shown to be crucial for the sustained down-regulation of the 1.1kb NHE1
promoter activity. Next, we ask if the activation of caspases is required for the down-
regulation of 0.15kb NHE1 promoter activity. The caspase activities upon treatment
with H2O2 in NIH 3T3 0.15kb cells were first investigated. Cells treated with 25, 35
and 50 showed increased caspase 2 and 3 activities (Fig. 42a). With 50 µM of H2O2,
activity of caspase 2 was increased by 5 folds compared to untreated cells while
activity of caspase 3 was increased by 16 folds. Although caspase 9 activity was also
detected, it was at much lower level than caspase 2 and 3. Caspase 6 and caspase 8
were not activated at all. We proceeded to determine the temporal activation of
caspases. NIH 3T3 0.15kb cells were treated with 50 µM of H2O2 for 3, 5, 9, 12, 24
and 48 h, capase activities in the cells were assessed. The results show that caspase 2
and 3 were activated as early as 9 h and the activation was sustained for up to 24 h
(Fig. 42b). After 48 h, caspase 2 and 3 activities returned to that of control cells where
no H 2O2 was added.
125 a)
20.0
18.0 *
control) 16.0 2 H2O2 25uM
O H 2 O2 25 µM 2 H2O2 35uM 14.0 H 2 O2 35 µM H2O2 50uM * H 2 O2 50 µM 12.0
10.0 * 8.0 *
6.0 *
4.0 * * * 2.0
0.0
Caspase to relative activity no H (Fold 2 3 6 8 9 Caspase
*: p-value < 0.05 (relative to no H 2O2 control) b)
20.0
18.0
16.0 control) 2 O
2 14.0
12.0 Caspase 2 10.0 Caspase 3 Caspase 6 8.0 Caspase 8 6.0 Caspase 9
4.0
2.0
Caspase to relative activity no H (Fold 0.0 3 5 9 12 24 48
Hr upon treatment with 50 µM of H2O2
Figure 42: H2O2 treatment activates caspase 2 and 3 in NIH 3T3 cells. a) NIH 3T3 cells were treated with various concentrations 25, 35 and 50 µM of H 2O2 for 24 h before harvested for caspase assay. b) NIH 3T3 cells were treated with 50 µM of H2O2 for various time points ranging from 3 h to 48 h and harvested for caspase assay. Cell lysates were assayed for caspase 2, 3, 6, 8 and 9 activity using caspase fluorogenic substrates of caspases. Results shown are obtained from the mean of 3 independent experiments +/- SE.
126
As the activation of caspase was shown to be important in the sustained down- regulation of the full length 1.1kb NHE1 promoter activity (Kumar et al., 2007), we tested if caspase activation is also required for the inhibition of 0.15kb NHE1 promoter. NIH 3T3 0.15kb cells were preincubated with general caspase inhibitor zVADfmk for 2 h followed by treatment with 50 µM of H2O2 for another 24 h. The cells were then harvested for caspase activity assay and luciferase assay. Figure 43a shows that preincubation of cells with zVADfmk was able to completely inhibit the activation of caspase 2 activity induced by H 2O2 treatment. Caspase 3 activity was also completely inhibited with zVADfmk (Fig. 43b). Having established that zVAdfmk completely inhibits caspase 2 and caspase 3 activities, we the assessed the effect on the 0.15kb NHE1 promoter activity. Figure 43c shows that the inhibition of caspase activity by zVADfmk could not prevent H 2O2-induced down-regulation
0.15kb NHE1 promoter. The data suggests that while 1.1kb NHE1 promoter required
the activation of caspases to have sustained down-regulation of its activity, this was
not the case for 0.15kb promoter.
a)
4.5 4 3.5 3 2.5
2 0H 2 O2 0 µM H 2 O2 50 µM treated control) treated 1.5 50 2
O 1 2
H 0.5
Caspase no to activity 2 (Fold 0 0 100 zVADfmk (µM)
127
b)
10 9 8 7 6 5 0H O 0 µM 4 2 2 H 2 O2 50 µM treated control) treated 50
2 3 O
2 2 H 1
Caspase 3 activity no to activity 3 (Fold Caspase 0 0 100 zVADfmk (µM)
c)
120
100
H2O2H2O2 0 0µM 80 H2O2H2O2 50 50 µM
control) 60 2 O 2 40
no H no 20
0 in NIH 3T3 cells to NIH cells relative 3T3(% in 0.15kbNHE1 promoter activity 0.15kbNHE1 activity promoter 0 100 zVADfmk (µM)
Figure 43: Pan caspase inhibitor zVADfmk does not prevent H2O2-induced down- regulation of the 0.15kb NHE1 promoter. NIH 3T3 stably transfected with 0.15kb NHE1 promoter was preincubated with 100 µM of pan caspase inhibitor zVADfmk for 2 h before treated with 50 µM of H2O2 for 24 h. Cells lysates were analyzed for a) caspase 2 activity and b) caspase 3 activity using caspase fluorogenic substrates of caspases. c) Cell lysates were assessed for NHE1 promoter activity using luciferase assay. Results shown are obtained from the mean of 3 independent experiments +/- SE.
128
3B.1.6 15d-PGJ2 down-regulates 0.15kb NHE1 promoter activity and
NHE1 mRNA level via ROS production
In the previous chapter of this thesis, we showed that 15d-PGJ2 down-regulated ER α
protein and mRNA expression via ROS production. Here, we ask if 15d-PGJ2 could
also inhibit 0.15 kb NHE promoter activity in a ROS-dependent manner. NIH 3T3
0.15kb cells were treated with different concentrations of 15d-PGJ2 for 24 h, after
which the NHE1 promoter activity was assessed. Figure 44a shows that 0.15kb NHE1
promoter activity decrease from 100 % to 70 % and 34 % when the cells were treated
with 10 and 20 µM of 15d-PGJ2 respectively. At the mRNA level, 20 µM of 15d-
PGJ2 reduced the amount of NHE1 mRNA by 50 % in 24 h when compared to control (Fig. 44b).
a)
120
100
80 *
60
control) * 40
20 3T3 cells (% relative relative untreated (% to cells 3T3
0.15kb activity NIH in NHE1promoter 0 0 5 10 20 15d-PGJ2
*:p-value < 0.05 (relative to no 15d-PGJ2 control)
129
b)
1.2 *
1
0.8
0.6 control)
0.4
0.2 NHE1 mRNA (Fold relative untreated relative (Fold mRNA NHE1 0 0 20 15d-PGJ2
*:p-value < 0.05
Figure 44: 15d-PGJ2 down-regulates 0.15kb NHE1 promoter activity and NHE1 mRNA level. a) NIH 3T3 stably transfected with 0.15kb NHE1 promoter were treated with various concentrations of 15d-PGJ2 for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay. b) Cells were treated with 20 µM of 15d-PGJ2 for 24 h. NHE1 mRNA level was determined by realtime-PCR. Results shown are obtained from mean of 2 independent experiments +/- SE.
The increased in the DCF fluorescence, a measurement of intracellular oxidation and
ROS production, can be represented as a shift in DCF fluorescence to the right of the x-axis of the FACS histogram. The 15d-PGJ2 caused the DCF fluorescence to shift to the right on the x-axis of the FACS histogram. Increase in intracellular ROS was detected at 2 h and 4 h after drug treatment (Fig. 45a and 45b). The rightward shift produced by 15d-PGJ2 was comparable to the rightward shift obtained when cells were incubated with 1 mM of H 2O2. This indicates that the ROS produced by 15d-
PGJ2 in NIH 3T3 was relatively high.
130 a)
H2O2 1mM Control 15d-PGJ2 20 µM
b)
H2O2 1mM Control 15d-PGJ2 20 µM
Figure 45: 15d-PGJ2 produces ROS in NIH 3T3 cells. NIH 3T3 0.15kb cells treated with 20 µM of 15d-PGJ2 for a) 2 h and b) 4 h. ROS levels in cells were detected using flow cytometry with DCFDA flurogenic probe.
131
Next, we asked if the ROS produced is responsible for the repression of 0.15kb NHE1 promoter activity by 15d-PGJ2. To test this hypothesis, antioxidant NAC was used in the following experiment to scavenge the ROS produced by 15d-PGJ2. The cells were preincubated with 0 or 5 mM of NAC for 2 h prior to treating the cells with 20 µM of
15d-PGJ2 for another 2 h. Cells were then assessed for DCF fluorescence. Figure 46a showed that 15d-PGJ2 caused a right-ward shift on the x-axis of the FACs histogram, indicating increased ROS production. Preincubation of NAC prior to 15d-PGJ2 addition shifted the graph back to the left on the x-axis of the FACs histogram, signifying a decrease in ROS production. The graph on Figure 46b summarized the effects of NAC on ROS production upon addition of 15d-PGJ2. This set of results show that NAC preincubation prevented ROS from being elevated in the cells treated with 15d-PGJ2. The outcome of scavenging ROS with NAC was that it indeed prevented the down-regulation of 0.15kb NHE1 promoter activity by 15d-PGJ2. As shown in Figure 46c, the 0.15kb NHE1 promoter activity decrease by 80 % in cells treated with 15d-PGJ2 alone. However, upon pretreatment with NAC this decrease in
NHE1 promoter activity was prevented. In a nutshell, the data suggests that similar to what was previously shown in section 3A.6 on the mRNA expression of ER α, the down-regulation of 0.15kb NHE1 promoter activity by 15d-PGJ2 was associated with the production of ROS.
132 a)
5mM NAC 15d- PGJ2 20 µM + 5mM NAC
Control 15d-PGJ2 20 µM
b)
200 *
180
160
140
120
100 MediumMedium control) 80 15d-PGJ215d-PGJ2 20 20 µM 60
40
DCF fluoresence (% relative to medium medium to (% relative fluoresence DCF 20
0 0 5 NAC (mM)
133 c)
120 *
100
80
60 PGJ2Medium 0
PGJ215d-PGJ2 20 20 µM 40
20 cells (% relative to medium medium control) (% to relative cells 0.15kb 3T3 activity NIH in NHE1 promoter 0 0 5 NAC (mM)
*: p-value < 0.05
Figure 46: Scavenging ROS produced by 15d-PGJ2 with NAC restores 0.15kb NHE1 promoter activity. a) NIH 3T3 0.15kb cells were preincubated with 0 or 5 mM of NAC for 2 h before treated with 20 µM of 15d-PGJ2 for 2 h. ROS levels in cells were detected using flow cytometry with CM-H2DCFDA flurogenic probe. b) Graphical representation of (a). c) NIH 3T3 0.15kb cells were preincubated with 0 or 5 mM of NAC for 2 h before treated with 20 µM of 15d-PGJ2 for 24 h. Cells were harvested for NHE1 promoter activity using luciferase assay. Results shown are obtained from mean of 2 independent experiments +/- SE.
134
3B.1.7 H2O2 down-regulates NHE1 expression in MCF7 breast cancer
cells
Since NHE1 is often up-regulated in tumor cells and has been shown to be important in tumorigenesis, we next tested if the H2O2-mediated inhibition of NHE1 expression also occurs in cancer cells. Using MCF7 as a model, NHE1 mRNA level was determined after H2O2 treatment. Treating MCF7 cells with 200 µM of H2O2 reduced the mRNA level of NHE1 by 40% (Fig. 47a). Similarly, diamide could down-regulate the mRNA transcript of MCF7. An 80 % reduction in the mRNA level of NHE1 was observed in diamide treated cells (Fig. 47b). This set of results show that, similar to
NIH 3T3 cells, NHE1 expression is negatively regulated by H2O2 and diamide in
MCF7 cells.
a)
1.2 *
1
0.8
0.6 control) 0.4
0.2
NHE1 mRNA (Fold relative to relative untreated (Fold mRNA NHE1 0 0 200
H2O2 (µM)
135 b)
* 1.2
1
0.8
0.6 control) 0.4
0.2
0 NHE1 mRNA (Fold relative to relative untreated (Fold mRNA NHE1 0 0.5 Diamide (mM)
*: p-value < 0.05
Figure 47: H2O2 and diamide down-regulate NHE1 mRNA level. a) MCF7 cells were treated with 200 µM of H2O2 for 24 h before harvested for realtime-PCR analysis. b) MCF7 cells were treated with 0.5 mM of diamide for 24 h before harvested for realtime-PCR analysis. Results are representative or mean of 3 independent experiments +/- SE.
Different 5’ deletion constructs of the mouse NHE1 promoter was transfected into
MCF7 to determine the important region for NHE1 promoter activity (Fig. 48a). The mouse promoter activity of NHE1 is similar to the full length 1.1kb promoter up to the 0.15kb region. When the 5’ 33bp fragment was removed from the 0.15kb region, it resulted in an 80% decrease in promoter activity (Fig. 48b). This shows that the 33 bp region of DNA contained important transcription binding site for NHE1 gene expression in MCF7 cells. The result was similar when we used the human NHE1 promoter (Fig. 48c). The promoter constructs of the human NHE1 gene was linked to a Chloramphenicol Acetyl Transferase gene (CAT) instead of luciferase gene. The
promoter activity up to the 0.2kb was similar to that of the full length with no
136 statistical difference (Fig. 48d). Subsequent deletion of the 5’ end of the promoter resulted in a significant decrease in promoter activity.
Multiple sequence alignment of the 0.15 kb promoter sequence of various animal species shows a high conservation of the entire region (Fig. 48e). This evolutionary conservation suggested that important and common transcription factors bind to this stretch of sequences. a)
TATA Luciferase gene 1.1kb TATA Luciferase gene 0.9kb TATA Luciferase gene 0.5kb
TATA Luciferase gene 0.2kb
TATA Luciferase gene 0.18kb TATA Luciferase gene 0.15kb
TATALuciferase gene 0.12kb b)
140
120
100 *
80
60
full length length promoter) full 40
20 * NHE1 promoter activity pXP-1.1 to activity relative (% promoter NHE1 0 pXP-1.1pxp 1.1pXP-0.9 pxp 0.9pXP-0.5 pxp 0.5pXP-0.2 pxp 0.2pXP-0.18 pxp 0.18pXP-0.15 pxp 0.15pXP-0.12 pxp 0.12 NHE1 promoter constructs
*: p-value < 0.05 (relative to pxp1.1 promoter) c)
TATA CAT gene -1374/+16
TATA CAT gene -252/+16
TATACAT gene -92/+16
137 d)
160
140
120
100
80 construct) 60
* 40
20 NHE1 promoter activity (% relative to -1374/+16 -1374/+16 to relative activity (% promoter NHE1 0 -1374/+16 -252/+16 -92/+16 NHE1 promoter constructs
*: p-value < 0.05 (relative to -1374/+16 promoter)
e) Mouse Rat Human Dog Cow Chimpanzee Bush Baby Mouse lemur Macaque Hedgehog
Figure 48: 0.15kb region of NHE1 promoter is highly conserved and important for NHE1 transcription. a) Different 5’ deletion constructs of mouse NHE1 promoter. b) MCF7 cells were transiently transfected with different 5’ deletion constructs of mouse NHE1 promoter. Cells were co-transfected with renilla plasmid for normalization. Results are obtained from the mean of 4 independent experiments +/- SE. c) Different 5’ deletion constructs of human NHE1 promoter. d) MCF7 cells were transiently transfected with the human NHE1 promoter plasmids -1374/+16, -252/+16 and -92/+16. Cells were co-transfected with renilla plasmid for normalization. CAT assay was carried out to assess the promoter activities. Results are obtained from the mean of 3 independent experiments +/- SE. e) NHE1 promoter sequence from various species was extracted from Pubmed database and underwent multiple sequence alignment using ClustalW function in Jarview software. Results shown are alignment of the start of NHE1 0.15kb promoter to the TATA box.
138
Next, we ask if the stretch of 33 bp region at the 5’ end of the 0.15kb NHE1 promoter is the H2O2 response element in MCF7. Since this short region of the mouse promoter
and the human promoter are 85 % similar, these promoter regions would most likely
bind to similar transcription factors (Fig. 49a & 49b). Oligonucleotides corresponding
to this region of 0.15kb human NHE1 promoter were synthesized for the subsequent
EMSA experiment. MCF7 nucleus extract was pretreated with various concentrations
of H2O2 for 20 min before it was allowed to bind to the 40 bp oligonucleotides. Figure
49c shows that binding of transcription factors to the oligonucleotides decreased with increasing concentrations of H2O2. This could be reversed by adding DTT after H2O2
had been allowed to oxidize the nuclear extracts (Fig. 49d). Taken together, the results
showed that the 33 bp of DNA sequence at the 5’ end of the 0.15kb of NHE1
promoter is important for basal transcription of NHE1 gene and it is a target of H2O2. a)
cgtgacacttccttccctgggcgacaggggccg ctgcacc -Mouse cgcgatacttctttccctcggcgacaggggccg ctgcgct -Human
b)
139 c)
H2O2 mM
Oligos only 0 0.2 0.5 1 3 5
Shift
Free probe
d)
H2O2 1mM - DTT 2 mM 2 DTT 0.5 mM 0.5 DTT Untreated Oligosonly
Shift
Free probe
Figure 49: H 2O2 prevents binding of transcription factor(s) to NHE1 promoter region. a) 40 bp of region from 5’ 0.15kb of NHE1 promoter in mouse and human. Underlined sequence represents exact 33 bp region between 0.15kb and 0.12kb of NHE1 promoter. b) Alignment of the mouse and human 40 bp region from 0.15kb of NHE1 promoter c) MCF7 nuclear extract was preincubated with various concentrations of H 2O2 for 20 min before the mixture were co- incubated with oligonucleotides that correspond to the NHE1 human 40 bp region for another 20 min. EMSA was then carried out according to the Materials and Methods section. d) MCF7 nuclear extract was preincubated with various concentrations of H 2O2 for 20 min before DTT was added to the mixture for another 20 min. The mixture was finally co- incubated with oligonucleotides that correspond to the NHE1 human 40 bp region for 20 min before EMSA was carried out as described in the Material and Methods section.
140
3B.2 TRANSCRIPTION FACTOR INVOLVED IN H 2O2 DOWN-
REGULATION OF NHE1 EXPRESSION
3B.2.1 Analysis of the region between 0.15kb and 0.12kb of NHE1 promoter.
Previously in this section, H2O2 was shown to down-regulate 1.1kb NHE1 promoter
activity (Fig. 33) and the important region for NHE1 transcription was located in a 33
bp stretch of nucleotides at the 5’ end of 0.15kb NHE1 promoter (Fig. 34c). This
0.15kb NHE1 promoter activity is regulated by H 2O2 in a reversible oxidative manner
(Fig. 37 – Fig. 39) and the binding of transcription factors to this short region were
also shown to be redox regulated (Fig. 49c and Fig 49d). Figure 50 is a diagram
illustrating the idea behind the down-regulation of NHE1 by H2O2. Firstly, H2O2
enters the cell and oxidize transcription factor(s) or co-factor(s) involved in the
transcription of NHE1 gene expression. This then leads to the down-regulation of the
promoter activity and inhibition of NHE1 mRNA synthesis.
H2O2
TF NHE1
NHE1 mRNA
Figure 50: Redox regulation of NHE1 transcription at the 0.15kb NHE1 promoter. Diagram showing how H 2O2 targets transcription factor of NHE1 leading to down-regulation of NHE1 mRNA.
141
3B.2.1.1 Bioinformatics software used to predict transcription factors binding to the 40 bp of the 0.15kb NHE1 promoter
In this section, bioinformatics was used to identify the transcription factors that might potentially bind to the 33 base-pair nucleotides at the 5’ end of 0.15kb NHE1 promoter. We extended the search to 40 base-pair nucleotides at the 5’ end so as to not exclude factors that might be binding to the boundary nucleotides. Table 1 shows a list of bioinformatics software that was used to predict transcription factors binding to the 40 bp of DNA sequence.
AliBaba 2 is a tool for prediction of transcription factor binding sites by context dependent matrices generated from TRANSFAC 3.5 public sites (Grabe, 2002).
ConSite is a user-friendly, web-based tool for finding cis-regulatory elements in genomic sequences. Predictions are based on the integration of binding site prediction generated with high-quality transcription factor models and cross-species comparison filtering (phylogenetic footprinting) (Sandelin et al., 2004). ConSite uses the JASPAR database, an open-access and non-redundant collection of curated profiles (Sandelin et al., 2004).
TESS is a web tool for predicting transcription factor binding sites in DNA sequences.
It can identify binding sites using site or consensus strings and positional weight matrices from the TRANSFAC, JASPAR, IMD, and CBIL-GibbsMat database
(Schug, 2008).
Patch is a search data base for potential transcription factor binding sites with the pattern search program using TRANSFAC 6.0 public sites (contributed by Jochen
Striepe and Ell en Goessling)
142
Table 1: Software used in the analysis of the 40 bp of 0.15kb NHE1 promoter.
Transcription factor prediction software Database used AliBaba 2 TRANSFAC v3.5 ConSite JASPAR TESS TRANSFAC v6.0 JASPER IMD v1.1 CBIL/GibbsMat v1.1 Patch TRANSFAC v6.0
3B.2.1.2 Transcription factors predicted to bind to the 40 bp of 0.15kb NHE1 promoter
As shown in Table 2, AP2, SP1 and COUP were predicted to bind to the 40 bp of
NHE1 promoter region by at least two of the transcription factor prediction software.
The predicted sites of transcription binding are shown in Figure 51. The binding of
AP2 to DNA is flexible and the consensus sequences reported for AP2 are GCC N3
GGC, GCC N4 GGC and GCC N3/4 GGG (Mohibullah et al., 1999). However, the possible AP2 binding sites predicted by the various software did not match the above
AP2 consensus sequence. This could be due to the flexibility of AP2 binding sites found in various targeted genes. Indeed, Dyck have found the site GGCGACAGGG on the NHE1 promoter to be an AP2 binding site (Dyck et al., 1995). The consensus sequence of SP1 is GGGCGG or its variants (Silverman and Drazen, 2000; Honda et al., 1997). Two possible SP1 binding sites were found on the 40 bp of NHE1 promoter region: GGGCGA and GGGCCG. A potential COUP binding site
(GTGTCA) was also found on the 40 bp sequence. This predicted site differs from the consensus sequence for COUP, (A/G)G(G/T)TCA (Guo et al., 2001), by one nucleotide.
143
Table 2: Transcription factors predicted to bind to the 40 bp nucleotide sequence of 0.15kb NHE1 promoter.
Transcription factor predicted Software used
AP2 Data from Dyck, ConSite, Patch, AliBaba 2 and TESS
SP1 AliBaba 2 and Patch
COUP Patch and TESS
a) AP2 sites
AP2 consensus sequence: GCC N3 GGC GCC N4 GGC GCC N3/4 GGG b)
SP1sites
SP1 consensus sequence: GGGCGG c) COUP site
COUP consensus sequence (A/G)G(G/T)TCA
Figure 51: Predicted transcription factor binding sites. Predicted binding sites for a) AP2, b) SP1 and c) COUP transcription factor.
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3B.2.2 Transcription factor AP2, SP1 and COUP
3B.2.2.1 AP2
AP2 or Activating Protein 2 is a family of transcription factors that consist of 5 members in human and mice namely AP2 α, AP2 β, AP2 γ, AP2 δ and AP2ε. All AP2 proteins share a highly conserved helix-span-helix dimerization motif at the carboxyl terminus, followed by a central basic region and a less conserved domain rich in proline and glutamine at the amino terminus (Eckert et al., 2005). The proteins are able to form hetero- as well as homodimers. The helix-span-helix motif together with the basic region mediates DNA binding and the proline- and glutamine-rich region is responsible for transactivation. AP2 has been shown to be flexible in its binding to
DNA sequence. It has been shown to bind to GCCN 3GGC, GCCN 4GGC and
GCCN 3/4 GGG (Mohibullah et al., 1999; Hilger-Eversheim et al., 2000). Other than the consensus sequence, AP-2 proteins have been shown to bind to a range of G/C- rich elements with variable affinities (Mitchell et al., 1987). AP2 is involved in cell proliferation and differentiation (Duan and Clemmons, 1995; Gaubatz et al., 1995;
Newman et al., 2000)
AP2 was reported to bind to the proximal promoter end at −95 to −111 bp from the start site of mouse NHE1 promoter (Dyck et al., 1995). Deletion of distal promoter sequence up to AP2 binding site decreases promoter activity by 25% in both NIH 3T3 and p19 embryonal carcinoma (Dyck et al., 1995; Dyck and Fliegel, 1995). Deletion of the AP2 site abolishes most of the promoter activity, suggesting that the AP2 site is important in the transcription of NHE1.
145
3B.2.2.2 SP1
SP1 or Specificity Protein 1 is a transcription factor found ubiquitously expressed in mammalian cells. It belongs to the Specificity Protein/Krüppel-like Factor (SP/KLF) transcription factor family, which is characterized by the highly conserved DNA binding domain with three adjacent Cys 2His 2-type zinc fingers (Suske et al., 2005).
SP1 binds to GC rich region on gene promoters (Briggs et al., 1986). There are more
than 12,000 sites in the human genome found to be SP1 binding sites and these sites
are associated with various genes involved in almost all cellular processes (Cawley et
al., 2004). SP1 also transactivates synergistically with a large variety of transcription
factors by binding DNA cooperatively or by recruiting the basal transcription
machinery synergistically (Wierstra, 2008). So far, SP1 have not been shown to be a
transcription factor for NHE1.
3B.2.2.3 COUP-TF
COUP-TF or Chicken ovalbumin upstream promoter transcription factor belongs to
the orphan subfamily of the nuclear receptor superfamily of steroid/thyroid hormone
receptor. There are 2 COUP genes, COUP-TFI and COUP-TFII. Members of the
nuclear receptor superfamily share similar structures such as N-terminal ligand-
independent activation domain, a highly conserved DNA binding domain (DBD), a hinge region and a ligand binding domain (LBD) (Tsai and O'Malley, 1994). COUP-
TFs bind to a consensus sequence of (A/G)G(G/T)TCA (Guo et al., 2001). They are involved in regulation of embryonic development and neuronal cell fate determination
(Tsai and Tsai, 1997).
146
The COUP-TFs have been reported to be involved the transcription of mouse NHE1 at the distal end of NHE1 promoter (Fernandez-Rachubinski and Fliegel, 2001). Study showed that nucleotides at -829 to -824 are critical for COUP-TF binding.
3B.2.2.4 AP2 and SP1 are redox regulated
AP2 had been reported to be oxidizable by H 2O2 and diamide, which inhibited its
DNA binding activity to synthetic AP-2 oligonucleotides (Huang and Domann, 1998).
The oxidation of AP2 is reversible by reducing agents such as DTT and BME. The mechanism of action might be via the oxidation of reactive cysteine residues in the
AP2 DNA binding domain.
SP1 has also been reported to be a redox regulated gene. It was found that thioredoxin alone, or in conjunction with the full thioredoxin system can increase SP1 DNA binding activity in vitro to oligonucleotides containing the SP1 consensus sequence
(Bloomfield et al., 2003). Another report showed that when young mouse nuclear extracts were treated with H 2O2, there was a drastic decrease in the SP1 DNA-binding
activity which can be restored by the treatment with high concentrations of DTT
(Ammendola et al., 1994).
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3B.2.3 Investigation of AP2 as transcription factor for NHE1
Based on the fact that AP2 has been shown to be redox regulated and found to bind to the 33 bp region at the 5’ end of 0.15kb NHE1 promoter (Dyck et al., 1995; Huang and Domann, 1998), we hypothesize AP2 to be the target transcription factor involved in H2O2-induced down-regulation of 0.15kb NHE1 promoter. H2O2 could oxidize AP2 and prevent it from binding NHE1 promoter, thus resulting in the down-regulation of promoter activity and NHE1 mRNA level. In order to test this hypothesis; we first need to know which isoform of AP2 is present in NIH 3T3 cells.
Mouse AP2 expression plasmid for each of the five AP2 isoforms and the empty plasmid were obtained from Dr. Bruce D. Gelb, Center for Molecular Cardiology,
Mount Sinai School of Medicine, USA. The plasmids were transfected individually into NIH 3T3 cells. Cells transfected with the empty plasmid have no AP2 isoform over-expressed, but contains the endogenous AP2 isoform. As the AP2 cDNA have been cloned into pCDNA6V5His plasmid containing a V5 epitope tag, we can detect over-expression of the isoforms using V5 specific antibody in western blot. Figure
52a shows that the different isoforms of AP2 have been expressed in NIH 3T3 cells.
To determine which isoform is present endogenously, we transfected another set of
NIH 3T3 cells with the plasmids and analyzed the mRNA expression using realtime-
PCR. Five TaqMan probes which detect specific AP2 isoforms were obtained. For each of the TaqMan assay using a specific AP2 isoform probe, the sample from the cells over-expressing the particular isoform, and sample from the empty plasmid expressing cells were assayed. The AP2 plasmid-transfected sample acts as the positive control for that particular isoform being detected. The cells transfected with the empty plasmid acts as the test of which isoform of AP2 is endogenously present.
Figure 52b to Figure 52f show the amplification plots of realtime-PCR assay detecting
148 for AP2 α to AP2 ε mRNA respectively. Specific AP2 isoform mRNA was detected from the positive control sample in the individual assay, but no signal was picked up from the empty plasmid sample except in AP2 α assay. This set of data suggests that
only AP2 α is present endogenously in NIH 3T3 cells.
a)
AP2 isoform
Empty α γ δ ε β plasmid
-50 kDa AP2-V5
β-actin -43 kDa
b)
AP2 α +ve control
Sample No RNA control
No reverse transcription control
149 c)
AP2 β +ve control
No reverse transcription control
No RNA control
Sample
d)
AP2 γ +ve control
No reverse transcription control Sample
No RNA control
150 e)
AP2 δ +ve control
No reverse transcription control
Sample
No RNA control
f)
AP2 ε +ve control
Sample
No reverse transcription control
No RNA control
Figure 52: AP2 α isoform is present in NIH 3T3 cells. NIH 3T3 cells were transfected with 6 µg of either pCDNA 6.1 empty vector, AP2 α, AP2 β, AP2 γ, AP2 δ or AP2 ε using SuperFect according to protocol in the Material and Methods section. Cells were harvested for a) western blot analysis or b) AP2 α mRNA, c) AP2 β mRNA, d) AP2 γ mRNA, e) AP2 δ, and f) AP2 ε mRNA by realtime-PCR.
151
To determine if AP2 α is indeed expressed in NIH 3T3, anti-AP2 α antibody was used
to probe for AP2 α protein in western blot. AP2 α siRNA was used to knockdown
AP2 α to ensure the specificity of the antibody used. Figure 53a shows that AP2 α is expressed exclusively in the nucleus with very little to no AP2 α in the cytosol. The siRNA targeting AP2 α was able to knockdown AP2 α protein. Transfection of the V5-
tagged AP2 α in to NIH 3T3 resulted in an increase in AP2 α protein detected by the
AP2 α antibody. Silencing of AP2 α also resulted in the disappearing of the band (Fig.
53b). This set of experiment indicates the antibody and siRNA of AP2 α is specific.
a) Cytosol Nucleus
(20nM) (20nM) (10nM) (10nM) (5nM) (5nM) α α α α α α siAP2 siAP2 siAP2 siAP2 siAP2 siAP2 NS siRNA (20nM) siRNA NS NIH 3T3 no NIH transfection
(20nM) siRNA NS PARP -116 kDa
AP2 α -50kDa
Cu/Zn SOD -16kDa
b) g) µ g) µ (10nM) (6 α
α NS siRNA (10nM) siRNA NS siAP2 pCDNA 6.1 (6 6.1pCDNA AP2 No transfection No
PARP -116 kDa
AP2 α -50kDa
Figure 53: AP2 α is present in the nucleus in NIH 3T3 cells. a) AP2 α in NIH 3T3 was silenced with various concentration of AP2 α siRNA using Lipofectamine RNAiMAX reagent as described in the Materials and Methods section. Cells were harvested 48 h post transfection and fractionated into cytosolic and nuclear fraction using NE-PER kit as described in the Materials and Methods section. b) NIH 3T3 cells were over-expressed with 6 µg of either empty vector pCDNA 6.1 or AP2 α. AP2 α of another set of cells were silenced with 10 nM of AP2 α siRNA or control siRNA. Cells were harvested 48 h post transfection and nucleus was isolated using NE-PER kit.
152
Having established that AP2 α is expressed in NIH 3T3, the effects of over-expressing
AP2 α on NHE1 expression was examined. First, the amount of plasmid required to
achieve good expression of AP2 α was optimized. NIH 3T3 cells were transfected with
2, 4 or 6 µg of either empty plasmid or AP2 α plasmid for 48 h before harvesting for
western blot analysis. As shown in Figure 54a, 2 µg of AP2 α plasmid hardly increased the level of AP2 α in the cells while 4 and 6 µg of the plasmid gave large increase in AP2 α expression. However, at the protein level, NHE1 was not increased
both at the pre-mature form or the mature form at 48 h post-transfection. In order to
determine when AP2 α start to be expressed, 6 µg of AP2 α plasmid was transfected into NIH 3T3 cells and harvested at 24, 48 and 72 h post transfection. From Figure
54b, it was observed that at 24 h, the cell were already expressing high level of AP2 α
protein and it remained high up to 48 h before it started declining at 72 h post-
transfection. Since the cells have already been expressing high level of AP2 α from 24
h to 48 h, NHE1 would have been given the time to be expressed. However, NHE1
protein expression remained similar to that of empty plasmid up to 72 h post-
transfection. The 0.15kb NHE1 promoter was checked for any increase in activity.
Figure 56c represented over-expression control for Figure 54d and 54e. No increase in
the 0.15kb promoter activity was observed when AP2 α was over-expressed (Fig. 56d).
Although there was a slight increase in the level of NHE1 mRNA, it was statistically insignificant (Fig. 54e).
153 a) g g g µ µ µ g g g µ µ µ 4 6 2 α α α AP2 AP2 AP2 Empty vector vector 4 Empty Empty vector vector 6 Empty Empty vector vector 2 Empty
-100 kDa NHE1 -85 kDa
AP2 α -50 kDa
β-actin -43 kDa b)
Hr post transfection 24 48 72 α α α AP2 AP2 AP2 Empty vector Empty vector Empty Empty vector Empty -100 kDa NHE1 -85 kDa
AP2 α -50 kDa
β-actin -43 kDa
c) α AP2 Empty vector Empty
-100 kDa NHE1 -85 kDa
AP2 α -50 kDa
-43 kDa β-actin
154 d)
120
100
80
60
40
20 3T3 (% relative vector) to relative empty (% 3T3 0.15kb NHE1 promoter activity 0.15kb NIH activity in NHE1promoter 0 Emptyempty vector vector AP2AP2 alphaα Plasmid
e)
1.6
1.4
1.2
1
0.8
0.6 vector control) vector
0.4
0.2 NHE1 mRNA (Fold relative to relative empty (Fold mRNA NHE1
0 Emptyempty vector vector AP2AP2 alphaα Plasmid
Figure 54: Over-expressing AP2 α does not affect NHE1 expression. a) NIH 3T3 cells were transfected with 2, 4, or 6 µg of empty vector or AP2 α plasmid overnight as described in the materials and methods section. Cells were harvested 48 h post-transfection. b) NIH 3T3 cells were transfected with 6 µg of empty vector or AP2 α plasmid overnight and harvested at 24 h, 48 h or 72 h post-transfection. Cell lysates were analyzed by western blot. NIH 3T3 cells were transfected with 6 µg of empty vector or AP2 α plasmid overnight and harvested at 48 h post transfection. Cell lysates were analyzed by c) western blot, d) luciferase assay for NHE1 promoter activity and e) realtime PCR for NHE1 mRNA. Results shown are obtained from the mean of 3 independent experiments +/- SE. p-value> 0.05 for d) and e).
155
It was showed in Figure 54 that over-expression of AP2 α protein did not lead to an
increase in the amount NHE1 protein, mRNA level or the 0.15kb NHE1 promoter
activity. However, it is possible that other AP2-like proteins might be involved in the
transcription of NHE1. To test the hypothesis, a dominant negative form of AP2 α
(AP2 ∆) which has the activation domain deleted (residues 31-117) was used. This form of AP2 is able to bind to the AP2 consensus sequence but is not capable of recruiting other co-activators for transcription. When the dominant negative AP2 sits on the NHE1 promoter, other AP2-like transcription factors would not be able to bind to the DNA sequence. This should result in a decrease in NHE1 transcription. Figure
55a shows that the transfection of AP2 ∆ was efficient for 2, 4 and 6 µg of plasmid
tested. Using 6 µg of AP2 ∆, we tested the expression of AP2 ∆ and NHE1 over 72 h.
AP2 ∆ was expressed from 24 h and the level remained high up to the 72 h tested (Fig.
55b). The level of NHE1, both immature and mature forms, were however unaffected
by the over-expression of AP2 ∆. In addition, the 0.15kb NHE1 promoter activity was also similar to that of the empty plasmid control (Fig. 55d). Along the same line, the mRNA level of NHE1 was also not affected by the over-expression of AP2 ∆ (Fig.
55e).
156 a) g g g µ µ µ g g g µ µ µ 6 4 2 ∆ ∆ ∆ AP2 AP2 Empty vector vector 6 Empty Empty vector vector 4 Empty AP2 Empty vector vector 2 Empty -100 kDa NHE1 -85 kDa
AP2 α -50 kDa AP2 ∆
β-actin -43 kDa
b)
Hr post transfection 24 48 72 ∆ ∆ ∆ AP2 AP2 AP2 Empty vector Empty Empty vector Empty Empty vector Empty
-100 kDa NHE1 -85 kDa
AP2 α -50 kDa
AP2 ∆
-43 kDa β-actin c) ∆ AP2 Empty vector Empty
-100 kDa NHE1 -85 kDa
AP2 α -50 kDa
AP2 ∆
β-actin -43 kDa
157 d)
120
100
80
60
40
20 relative to empty vector vector to control) empty relative 0.15kb NHE1 promoter activity 0.15kb activity (% NHE1promoter
0 Emptyempty vector vector AP2AP2 -ve∆ Plasmid
e)
1.2
1
0.8
0.6
vector control) vector 0.4
0.2 NHE1 mRNA (Fold relative to relative empty (Fold mRNA NHE1
0 Emptyempty vector vectorAP2 AP2∆ -ve Plasmid
Figure 55: Over-expressing AP2 dominant negative does not affect NHE1 expression. a) NIH 3T3 cells were transfected with 2, 4, or 6 µg of empty vector or AP2 ∆ plasmid overnight as described in the materials and methods section. Cells were harvested 48 h post-transfection. b) NIH 3T3 cells were transfected with 6 µg of empty vector or AP2 ∆ plasmid overnight and harvested at 24 h, 48 h or 72 h post-transfection. Cell lysates were analyzed by western blot. NIH 3T3 cells were transfected with 6 µg of empty vector or AP2 ∆ plasmid overnight and harvested at 48 h post transfection. Cell lysates were analyzed by c) western blot, d) luciferase assay for NHE1 promoter activity and e) real-time PCR for NHE1 mRNA. Results shown are obtained from the mean of 3 independent experiments +/- SE. p-value> 0.05 for d) and e).
158
Over-expression of AP2 α has no effect on either the level of protein or mRNA of
NHE1 or on the promoter activity of NHE1. This could be due to the efficient
transcription by endogenous AP2 α or insufficient cofactors required to increase transcription. However, the expression of dominant negative AP2 was also unable to affect NHE1 transcription. This led us to speculate that AP2 or AP2-like transcription factor are in fact not involved in transcription of NHE1. To test this hypothesis, the endogenous AP2 α was silenced using siRNA and its effects on the level of NHE1 protein and mRNA, and 0.15kb NHE1 promoter activity was examined. Figure 56a shows the efficient silencing of AP2 α where more than 90% knockdown was observed. Protein level of NHE1 remained unchanged. The mRNA level of NHE1 when AP2 α was silenced was the same as that of control cells (Fig. 56b). The 0.15kb
NHE1 promoter was also unaffected by the depletion of AP2 α in the cells (Fig. 56c).
Finally, to show that AP2 α is not involved in the down-regulation of NHE1 by H 2O2,
we silenced AP2 α and treated the cells with H 2O2 for 24 h. Results showed that the promoter activity of AP2 α depleted cells was still down-regulated when treated with
H2O2 (Fig. 56d). This set of data confirmed that the target of H 2O2 is a transcription factor other than AP2 α.
From the bioinformatics analysis, SP1 and COUP have been identified as potential transcription factor for NHE1 in addition to AP2. It would be interesting to investigate if they are indeed able to regulate transcription of NHE1. Of particular interest would be SP1, as it was been reported to be a redox-regulated gene. Future work would involve verifying SP1 as the transcription factor for NHE1 and determining what role it plays in H 2O2-mediated repression of NHE1 promoter activity.
159 a) 10nM α siAP2 NS siRNA 10nM siRNA NS -100 kDa NHE1 -85 kDa
AP2 α -50 kDa
β-actin -43 kDa b)
120
100
80
60 siRNA)
40
20 NHE1 mRNA (% relative to relative control (% mRNA NHE1
0 NScosi siRNA 10nM 10nM siAP2AsiAP2 α 10nM siRNA
c)
140
120
100
80
60
40
20 3T3 cells (% relative siRNA) relative control (% to cells 3T3 0.15kb NHE1 promoter activity 0.15kb NIH activity in NHE1 promoter 0 NS siRNAcosi 10nM 10nM siAP2AsiAP2 α 10nM10nM siRNA
160 d)
120
100 control) 2 O 2 80
H2O2 0 60 H2O2 0 µM H2O2H2O2 5050 µM 40
20 0.15kbNHE1 NIH activity in promoter 3T3 cells (% relative norelative H (% to cells 3T3 0 NS siRNAcosi 10nM 10nM siAP2 siAP2Aα 10nM 10nM siRNA
Figure 56: Silencing AP2 α does not affect NHE1 expression. AP2 α in NIH 3T3 was silenced with 10 nM AP2 α siRNA or control siRNA. Cells were harvested at 48 h post transfection and cell lysates were analyzed by a) western blot, b) real-time PCR for NHE1 mRNA and c) luciferase assay for NHE1 promoter activity. d) AP2 α in NIH 3T3 was silenced with 10 nM AP2 α siRNA or control siRNA and cells were treated with 50 µM of H2O2 at 24 h post-transfection for another 24 h. NHE1 promoter activity using luciferase assay was assessed. Results shown are obtained from the mean of 3 independent experiments +/- SE. p- value> 0.05 for b), c), and d).
161
CHAPTER 4: DISCUSSION
4.1 REDOX REGULATION OF ER α
Recent world estimate revealed that breast cancer is the most common cancer that
affects women (Jemal et al., 2009), and it is the leading cause of death in women
between the ages of 35 and 45 (Stoica et al., 2000). Of all the breast cancer patients,
approximately 60% of them are ER α-positive and their tumor growth is often stimulated by estrogen through ER α (Dickson and Lippman, 1988; Jacquemier et al.,
1990). Other than being an important prognosis marker that correlates with better
survival rate, ER α is also the target of endocrine therapy (Stoica et al., 2000). The
clinical relevance of ER α makes it an important target to study. Although research on
ER α has been ongoing for the past 30 years, there are aspects of the receptor which have not been fully explored. One such aspect that we have identified is the regulation of ER α by ROS.
ROS have been found to be up-regulated and implicated in the pathogenesis of a variety of diseases including breast cancer. Many anti-tumorigenic drugs have also been shown to produce ROS. While ER α transcriptional activity has been reported to
be redox regulated, the effects of ROS on the expression of ER α remain poorly
understood. Intrigued by this gap in knowledge, we set out to investigate the
mechanistic role ROS, in the form of H 2O2, on the expression and function of ER α.
The findings of this study provide new insights to ER α signaling and give rise to new
strategies for the treatment of breast cancer.
162
4.1.1 Regulation of ER α expression and function by oxidative stress
4.1.1.1 ER α protein expression is specifically targeted by H 2O2/ ROS
ER α signaling can be controlled at the expression level or at the level of its role as a
transcription factor that binds ERE. Our experimental data provided evidence that
ER α expression, but not ERβ expression, is a target of H 2O2-meditated down- regulation in MCF7 breast cancer cells. The down-regulation of ER α protein can be
observed as early as 8 h after H2O2 treatment. And it occurs regardless of the availability of estrogen. This indicates that estrogen does not participate in the suppression of ER α by H 2O2. H2O2 was also shown to down-regulate ER α protein in
T47D, another breast cancer cell line, which suggest that H2O2-mediated suppression
of ER α is a general phenomenon in ER α-positive breast cancer cells.
Our data is in contrast with another report which looked into the response of ER to oxidative stress. Tamir et al did not find a down-regulation of ER α when MCF7 or
T47D were treated with H 2O2 but instead found an up-regulation of ERβ (Tamir et al.,
2002). The discrepancy in the results is most likely due to the difference in the
concentration of H 2O2 used in the two systems. The authors of that paper used a very
low concentration of 2.5 µM of H 2O2 whereas we found that suppression of ER α
expression occurs only when the cells are exposed to at least 50 µM of H 2O2. It should be noted that the concentrations of H 2O2 (100 and 200 µM) used in our present study was considered to be sub-lethal levels (Barnouin et al., 2002). In alignment with our findings, another research using glucose oxidase system which produces H 2O2 in the medium, reported down-regulation of ER α in 24 h (Weitsman et al., 2009).
However, the amount of H2O2 the cells were exposure to in Weitsman’s study was not specified, as glucose oxidase system produces H 2O2 in a continuous manner with the amount of H2O2 produced dependent on the units of enzyme used. Although it was 163 not known how ER α was down-regulated by glucose oxidase, proteosomal degradation was suggested as its mechanism of action. This hypothesis was tested by us and was found to be incorrect in our system.
Apart from H2O2, peroxynitrite was shown to be able to down-regulate ER α protein expression. Therefore, the down-regulation of ER α might be a general phenomenon that can be mediated by different species of ROS. It remains to be tested whether other species of ROS such as superoxide and hydroxyl radical are able to repress ER α
expression. Through these findings, we propound that chemotherapeutic agents that
produce H 2O2 or ROS might also be able to down-regulate ER α in breast cancer cells.
4.1.1.2 ER α mRNA level is regulated by H 2O2: A novel finding
ER α expression can be regulated at the level of protein degradation (Wijayaratne and
McDonnell, 2001; Wang et al., 2009), transcription (Huang et al., 2006; Sundar et al.,
2008) and mRNA stability (Saceda et al., 1998; Kenealy et al., 2000; Ing and Ott,
1999). We tested the hypothesis that the down-regulation of ER α by H2O2 is due to
proteosomal degradation, by inhibiting proteosome with MG132. However, inhibition
of the proteosome was unable to restore ER α protein level. Having ruled out increased
ER α protein degradation as the mechanism of ER α down-regulation by H2O2, we moved on to examine the abundance of ER α mRNA in H2O2 treated and untreated
cells. The data showed that the ER α mRNA level was drastically decreased as early as
4 h upon treatment with H2O2. This finding is significant as it is the first study to
demonstrate that ER α mRNA level is decreased in response to H2O2 treatment. In addition, it is very likely that the observed down-regulation of ER α protein by H2O2
was a result of the decrease in ER α mRNA level.
164
Besides ER α, H2O2 has also been shown to decrease the mRNA of other genes. One
group which studied the effect of oxidative stress on the plasma membrane Ca 2+
extrusion systems in hippocampal neurons found that Na +/Ca 2+ exchangers were
significantly down-regulated at the mRNA level after 2-3 h exposure to 300 µM H 2O2
(Kip and Strehler, 2007). Along the same line, the mRNA level of CdK2 is decreased
after oxidative stress in human diploid fibroblasts exposure to 150 µM of H2O2
(Frippiat et al., 2003). Furthermore, H2O2 could partially block the mRNA expression of CD28 in Jurkat T cells (Ma et al., 2003).
The down-regulation of ER α mRNA could be the result of either of the following two
possible mechanisms or a combination of both the mechanisms. The first mechanism
is that H2O2 suppresses the transcription of ER α. Apart from the basal transcription machinery, genes recruit sequence specific transcription factors to sit on the enhancer/silencer region of DNA adjacent to the regulated genes to promote or silence gene expression. These transcription factors often have activation domains that will recruit other co-regulators to help in regulation transcription (Latchman, 1997). The timing and amplitude of gene transcription depend on the type of transcription factors and co-regulators being recruited. H2O2 could be targeting the specific transcription factor or co-regulator involved in the transcription of ER α. This can happen via direct oxidation of these proteins, causing them not to bind to the DNA or not being able to be recruited to promote transcription (Ammendola et al., 1994; Huang and Adamson,
1993; Huang and Domann, 1998; Liang et al., 1998). The transcription factor or co- regulators could also be degraded in response to oxidative stress. With the ER α gene containing at least 7 promoters, the transcriptional regulation of ER α is highly
complex (Kos et al., 2001). Additional experiments such as cloning of these
165 promoters would be needed to determine if H2O2 down-regulate ER α through decreasing its transcriptional activity.
The turnover of the mRNA also plays an important role in the control of gene expression. In mammalian cells, the abundance of a particular mRNA can fluctuate drastically following a change in the mRNA half-life, without any alteration in transcription rates (Ross, 1995). Another possible mechanism that leads to decreased in ER α mRNA level is through decrease in the stability of the synthesized mRNA.
The half-life of ER α mRNA was previously determined to be 4-5 h (Saceda et al.,
1998; Kenealy et al., 2000). Treating cells with E2 causes the half-life of ER α mRNA to drop from 4 h to 40 min. Similarly, Trichostatin A and 5-Aza 2'deoxycytidine can also decrease ER α mRNA and protein levels by decreasing ER α mRNA stability
(Pryzbylkowski et al., 2008). An mRNA with a shorter half-life will be less abundant
at any given time, leading to lower translation and decreased protein expression.
4.1.1.3 The transient nature of H 2O2-induced down-regulation of ER α
expression
The down-regulation of ER α protein and mRNA level was not permanent but transient where the suppression lasted up to 24 h or 48 h depending on the concentration of H 2O2 the cells were exposed to. This is not surprising as the data showed that the growth of the cells was inhibited by H 2O2 but the cells were not dying.
These findings are reminiscent of 1α,25-dihydroxyvitamin D inhibition of ER α
transcription. ER α mRNA level was first down-regulated 5 h after 1α,25- dihydroxyvitamin D treatment and subsequently returned to that of untreated cells after 24 h (Swami et al., 2000). Such reversible down-regulation and subsequent re- expression of protein was also observed in the regulation of Bcl-xL expression by
166
H2O2 in cardiac myocytes (Valks et al., 2003). Bcl-xL in H 2O2 treated cardiac myocytes was down-regulated in the first 6 h of treatment. The protein was subsequently re-expressed through the increase in mRNA level.
The signaling mechanism that resulted in the down-regulation of ER α was not
permanent due to the treatment being a one-time exogenous addition. Thus, whatever
molecules that were affected by H 2O2 treatment will return to their basal state after some time. Higher concentration of H 2O2 elicited a greater response in the cell signaling pathway, resulting in a strong down-regulation and took longer for ER α to
be reexpressed. For example, H2O2 might oxidize the transcription factor of ER α
preventing it from binding to the ER α promoter thereby inhibiting ER α transcription.
This oxidized transcription factor might return to its original reduced state through the
cell’s antioxidant system after some time. Alternatively, the transcription factor might
get degraded upon H2O2 treatment but new transcription factor would be formed to replace the degraded one. As such, ER α transcription would be restored and new ER α
protein will be made.
167
4.1.1.4 Functional consequences of ER α down-regulation
4.1.1.4.A ER α-regulated genes response
The down-regulation of ER α expression can have different functional outcomes. For
example, the binding of E2 to ER α has been shown to decrease the level of the receptor through proteosomal degradation, while ER α response genes expressions are
increased. On the other hand, hypoxia also down-regulates ER α via proteosomal
degradation, however, the ER responsive gene, PS2, was found to be down-regulated
together with the loss ERE-driven reporter transcription (Stoner et al., 2001).
Consistent with the hypoxia treatment, H2O2 suppression of ER α expression led to the disruption of the estrogen responsiveness. This is observed in the loss of ERE-driven reporter plasmid transcription. In addition, PR and c-Myc were shown to be estrogen responsive genes and the down-regulation of ER α by H 2O2 resulted in the decrease of
their gene expression. Similarly, ER α down-regulation affected E2 signaling by decreasing E2-mediated c-Myc expression. Defective ER α DNA-binding activity has
been found in up to one-third of all ER α-positive breast cancers and is correlated with absence of PR expression (Scott et al., 1991). Taken together, our results indicated a functional ER α in the MCF7 cell line used and the down-regulation of ER α by H2O2
led to a loss of E2 responsiveness.
H2O2 was shown to decrease ER α but not ERβ, therefore, E2 should still be able to stimulate ERE through ERβ. On the contrary, the ERE activity in H 2O2 treated, ER α
depleted cells were much lower compared to untreated cells upon E2 stimulation. This
observation indicates that the ERβ activity upon E2 treatment is low in MCF7 cells
and most of the estrogen-responsiveness is mediated through ER α. The lower transactivation ability of ERβ has also been highlighted by several groups both on synthetic and endogenous genes (McInerney et al., 1998; Cowley and Parker, 1999).
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4.1.1.4.B Growth inhibition
It is well documented that the lost of ER α results in growth arrest of breast cancer
cells (Lu et al., 2003; Chow et al., 2004; Sundar et al., 2008). In conjunction with the
suppression of ER α expression, the growth of MCF7 was significantly inhibited by
H2O2 treatment. It is likely that the observed anti-proliferative effect of H 2O2 on
MCF7 cells is a functional consequence of ER α down-regulation involving the loss of
estrogen-mediated signaling. In breast cancers, over-expression of ER α is commonly
found and has been linked to enhanced proliferation and resistance to apoptosis. ER α
does so by promoting cells in G 0 phase to enter the cell cycle and hastening the G 1 to
S phase transition (Prall et al., 1997; Foster et al., 2001). ER α upon binding to
estrogen, also stimulates the expression of c-Myc which is a powerful oncoprotein
involved in cell proliferation, differentiation and apoptosis (Dubik and Shiu, 1988;
Osborne et al., 2004). Tolhurst et al recently provided evidence that ER α over- expression led to ligand-independent expression of the estrogen-regulated genes PR and pS2, and growth in the absence of estrogen (Tolhurst et al., 2010).
A few reports have shed light that the ratio of ER α: ERβ is important in controlling estrogen regulated proliferation in estrogen responsive cells (Covaleda et al., 2008;
Treeck et al., 2010) . For instance, the ratio of ER α: ERβ expression is higher in breast
tumors than in normal tissues due to lower expression of ERβ in tumor cells
(Lazennec et al., 2001). This high ER α: ERβ ratio found in breast cancer cells corresponded to a highly proliferative state (Frech et al., 2005). Conversely, low ER α:
ERβ ratio found in normal mammary cells and in cancer cells made to over-express
ERβ, correlated with an anti-proliferative state and an induction of terminal differentiation (Förster et al., 2002; Ström et al., 2004; Paruthiyil et al., 2004; Frech et al., 2005). Treatment of cells with H2O2 decreases the levels of ER α while the levels
169 of ERβ remained unchanged. This resulting low ER α: ERβ ratio could also have contributed to the anti-proliferative state in H2O2-treated cells.
4.1.2 Oxidation as the mechanism mediating H 2O2-induced down-regulation of
ER α
In the present study, pretreatment of the cells with reducing agent DTT prevented the down-regulation of ER α by H 2O2. Thus, the results suggest that H2O2 induced the
down-regulation of ER α through a reversible oxidation mechanism in MCF7 cells.
Our data with diamide mimicked the effects of H 2O2 which indicates the involvement of thiol oxidation of cysteine residue that might be present in molecules required for
ER α expression. Others have shown that a number of signaling molecules, including receptors (Suc et al., 1998; Liu et al., 1999; Uchida et al., 1999) and downstream elements such as G-proteins (Mallis et al., 2001), kinases (Ward et al., 1998;
Gopalakrishna and Jaken, 2000), phosphatases (Mahadev et al., 2001) and
transcription factors (Choi et al., 2001) are regulated by reversible oxidation of critical
cysteine residues (Eaton et al., 2003).
H2O2 could affect the signaling cascade of kinases and phosphatases, leading to alteration of phosphorylation status and activity of cellular proteins such as transcription factors and RNA stabilizing factors. For instance, upon treatment of cells with H 2O2, the small GTPase Ral is activated. Resulting from this is a JNK-dependent
phosphorylation of FOXO4 transcription factor which leads to the nuclear
translocation and activation of FOXO4 (Essers et al., 2004). On the other hand,
mRNA stabilizing factor, HuR has been shown to be regulated by PKC α-dependent phosphorylation. This posttranslational modification of HuR by phosphorylation represents a critical feature for HuR shuttling from nucleus to cytosol to increase
170
COX-2 mRNA stability (Doller et al., 2007). H 2O2 can either activate or inactivate
PKC α depending on the concentration of the oxidant (Gopalakrishna and Anderson,
1989; Abdala-Valencia and Cook-Mills, 2006). In the down-regulation of ER α
expression, one possible scenario is that H2O2 could affect the balance of the kinases
and phosphatases activities in the cell, leading to the inactivation of transcription
factors or RNA stabilizing factors involved in the expression of ER α. This would
decrease the steady-state ER α mRNA level in the cell.
H2O2 could also oxidize the critical cysteine residues on the DNA-binding domain of transcription factors, abolishing their DNA-binding abilities. Indeed, oxidation of
transcription factors provides a direct regulation of transcription of a particular gene.
For example, H 2O2 and diamide were found to abolish the DNA-binding activity of
heat-shock transcription factor through oxidizing critical residues within the DNA-
binding domain. This oxidation can be reversed by DTT and Trx (Jacquier-Sarlin and
Polla, 1996). Although the hypothesis remains to be tested, it is possible that H 2O2
targets the DNA binding domain of transcription factors regulating ER α transcription, resulting in the loss of ER α promoter activity. Along the same lines, AP2 γ has been reported to be a transcription factor for ER α that trans-activates the ER α gene in hormone-responsive tumors (deConinck et al., 1995; McPherson et al., 1997;
McPherson and Weigel, 1999; Schuur et al., 2001; Woodfield et al., 2007). SP1, together with USF-1 and ER α, have also been shown to be essential for ER α gene
transcription (deGraffenried et al., 2002; deGraffenried et al., 2004). As mentioned in
section 4.3.2.4 of the results section, both AP2 and SP1 had been shown to be redox-
regulated transcription factors. The oxidation of these transcription factors has also
been found to be reversible by DTT treatment. Therefore, further research is warrant
171 to establish if these transcription factors are involved in H 2O2-mediated down- regulation of ER α.
4.1.3 Significances of ER α as a redox regulated gene
4.1.3.1 Potential clinical relevance
Resistance to apoptosis is a major hindrance in the treatment of cancer, and proliferative factors such as ER α play an important role in this resistance (Lee and
Nam, 2008). The use of Selective Estrogen Receptor Modulators (SERMs),
particularly tamoxifen, which competes with E2 in binding to ER α have proven to be effective in the treatment of breast cancer (Gronemeyer et al., 2004). However, resistance to tamoxifen is a major obstacle in the management of breast cancer.
Intrinsic resistance occurs in approximately 40 % of ER α-positive tumors where the
cancer cells fail to initially response to tamoxifen treatment. Acquired resistance can
occur when changes in ER α signal transduction pathways convert inhibitory SERM
ER α complex to a growth stimulatory signals. (Jordan, 2004). Alterations in signal transduction pathways which lead to ligand-independent activation of ER α via
phosphorylation can also be involved in tamoxifen resistance (Pearce and Jordan,
2004). Most resistant breast tumors remain ER α-positive and frequently respond to alternative endocrine treatment, indicative of a continued role for ER α in breast cancer cell proliferation (Ali and Coombes, 2000). The problem of resistance has resulted in the search for and the development of diverse therapies designed to inhibit
ER α action.
The mechanism underlying many chemotherapeutic agents and ionizing radiation on
tumor cell kill is the production of ROS leading to oxidative stress (Wang and Yi,
172
2008). The data presented in this study showed that ROS is able to down-regulate
ER α expression in breast cancer cells. The ability of ROS to suppress total ER α in the
cancer cells is superior to tamoxifen’s action of just blocking the binding of estrogen
to the receptor, since depletion of ER α reduces both ligand-dependent and ligand- independent activation ER α activity. The regulation of ER α expression by ROS provides an alternative approach to target the proliferative actions of ER α in breast
cancer cells. One application of our findings is to target the expression of ER α though agents that produces ROS intracellularly in tumor cells.
It was suggested that cancer cells are more vulnerable to oxidative stress because they function with a heightened basal level of ROS mediated signaling. Other than decreasing the level of functional ER α as shown in this study, ROS can induce lipid peroxidation, DNA damage, and protein oxidation (Schumacker, 2006). ROS have been shown to target several proteins that are important in cell signaling events, including those involved in cell death or survival, i.e ., NF κB, AP-1, H-Ras, MAPK,
IP3 kinase, PKC-ε, Ras, p53, HIF-1, ASK-1, Bcl-2, caspases, JNK and p38 MAPK
(Gibellini et al., 2010). Furthermore, ROS were reported to induce apoptosis by
triggering mitochondrial permeability transition pore opening and release of
proapoptotic factors (Brenner and Grimm, 2006). Due to their multiple effects
affecting cell survival, ROS producing agents might be more efficacious than
conventional ER α targeting drugs such as aromatase inhibitor and fulvestrant in treating breast cancer.
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4.1.3.2 Inducing down-regulation of ER α through ROS producing agents
Many of the factors shown to down-regulate ER α have been reported to produce ROS
in cells. Table 3 shows compounds that have been found to down-regulate ER α and
the proposed mechanism of down-regulation. Artemisinin and oligomines have been
found to down-regulate ER α by inhibiting the promoter activity of ER α. Tumor
necrosis factor α (TNF α), epithelial growth factor (EGF), arsenic trioxide, genestein, resveratrol, and 1 α, 25-dihydroxyvitamin D could suppress ER α at the mRNA level.
(-)-Epigallocatechin-3-gallate (EGCG) inhibits ER α function while 15d-PGJ2 down- regulates ER α through the proteosomal degradation pathway. All the compounds mentioned above have been demonstrated to produce ROS in cells. However, the link between the down-regulation of ER α and ROS by these agents has never been explored.
In this thesis, the mechanism of how 15d-PGJ2 down-regulate ER α and its relationship with ROS was investigated. Previously, ER α was shown to be down- regulated via proteosome-dependent pathway upon treatment with 15d-PGJ2 (Qin et al., 2003). However, in that study, the restoration of ER α by MG132 was only 50 % even after using high concentration of the inhibitor. Therefore, proteosomal degradation alone cannot fully account for how ER α was down-regulated by 15d-
PGJ2. Our research indicates that the mRNA of ER α was drastically reduced in 24 h after 15d-PGJ2 treatment. This is the first report to show that ER α mRNA level is
affected by 15d-PGJ2. Thus, in addition to the protein degradation, 15d-PGJ2 can also
down-regulates ER α at the mRNA level. As the mRNA of ER α was almost
completely down-regulated at 24 h, we propose that the down-regulation of ER α at
the transcript level is the mechanism which maintains low ER α protein level at later
174 time-points while proteosomal degradation could be responsible for the early down- regulation of ER α.
15d-PGJ2 is a natural agonist of PPAR γ which binds to PPAR γ, resulting in the heterodimerization with retinoic X receptor and binding to PPAR γ response element.
This could lead to either the up-regulation or down-regulation of target gene expression (Fuenzalida et al., 2007; Burgermeister et al., 2003; Kumar et al., 2009).
One obvious question when using 15d-PGJ2 is whether the effects that we observed on ER α were due to the genomic regulation by the activation of PPAR γ. Utilizing the
computational analysis tool, Peroxisome Proliferator Response Element (PPRE)
Search program (Kumar et al., 2009), we looked for potential PPRE on the ER α
promoter A, a promoter selectively engaged by breast cancer cells (Grandien et al.,
1995). The results in Appendix A show that there are two potential PPAR γ binding sites on the ER α promoter. It is plausible that 15d-PGJ2 affects the expression of ER α
by activating PPAR γ to repress the promoter activity of ER α. However, the down- regulation of ER α still occurred in the presence of PPAR γ antagonist, GW9662. It is
known that 10 µM of GW9662, the maximum concentration used in our study, is
sufficient to completely block 15d-PGJ2-induced PPRE-driven luciferase activity in
MCF7 cells (Wang et al., 2006). And toxic effects were observed when GW9662 was
used at a higher concentration (Seargent et al., 2004). In line with our findings,
Lecomte et al also reported that PPAR γ was not directly involved in the down-
regulation of ER α by 15d-PGJ2 (Lecomte et al., 2008). Three PPAR γ antagonists
(T0070907, BADGE and GW9662) failed to block the suppression of ER α expression induced by PPAR γ ligands. Silencing of PPAR γ in MCF7 also could not prevent ER α
down-regulation mediated by 15d-PGJ2, thus, demonstrating a PPAR γ-independent
mechanism (Lecomte et al., 2008).
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In fact, we found that the effects of 15d-PGJ2 on ER α were mediated through
intracellular ROS production. The level of ROS was increased dramatically in cells
treated with 15d-PGJ2 and this is consistent with previous reports (Kim et al., 2008;
Kim et al., 2009; Shin et al., 2009). Scavenging the ROS with NAC restored the ER α
protein and mRNA level completely, implicating ROS to be the critical intermediate
involved in the down-regulation of ER α by 15d-PGJ2 in our system. The ROS produced by 15d-PGJ2 has been suggested to be generated through mitochondria and
NADPH oxidase activation (Shin et al., 2009). The use of catalase could abolish 15d-
PGJ2-induced production of ROS, which implicates H2O2 as the ROS generated (Shin et al., 2009). In addition to ROS, RNS was also significantly elevated with 15d-PGJ2 treatment (Lim et al., 2007). In the current study, the mechanisms for ROS production and the type of ROS produced by 15d-PGJ2 are not explored; further research to clarify these questions would be required. Suffice to say, H 2O2 has been shown to be produced in 15d-PGJ2 treated cells (Shin et al., 2009), and therefore, it could potentially be the ROS mediating the down-regulation of ER α in our system.
In summary, our results have demonstrated that ROS produced by drug treatment
could indeed lead to the down-regulation of ER α. 15d-PGJ2 is able to inhibit breast cancer cell growth and suppress ER α expression through ROS production. Other have also found 15d-PGJ2 to be effective in killing various cancer cell types, i.e., leukemia cells, prostate cancer cells, thyroid papillary cancer cells, colorectal cancer cells, glioma cancer cells and oral squamous cell carcinoma cells (Butler et al., 2000;
Nikitakis et al., 2002; Chen et al., 2002; Cho et al., 2006; Shin et al., 2009). Taken together, these observations suggest that 15d-PGJ2 could be a promising chemotherapeutic agent in the treatment of breast and other cancers.
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Table 3: List of compounds and growth factors found to down-regulate ER α and produce ROS.
Compound Source Mechanism of action Produce ROS Reference article ROS Artemisinin (Sundar et Down-regulates Yes (Meshnick et al., al., 2008) mRNA transcripts by 1993; Schmuck et inhibiting promoter of al., 2002; Efferth ER α and Oesch, 2004)
(-)- (Farabegoli Inhibits ER α function Yes (Nakazato et al., Epigallocatechi et al., 2007) 2005) n-3-gallate (EGCG) TNF α (Lee and Down-regulates Yes (Goossens et al., Nam, 2008) mRNA transcripts 1999; Kim et al., 2010) EGF (Stoica et al., Down-regulates Yes (Huo et al., 2009) 2000) mRNA transcripts
Oligoamines (Huang et al., Down-regulates Yes (Wang and Casero, 2006) mRNA transcripts by 2006; Lasbury et inhibiting promoter of al., 2007) ER α
Arsenic trioxide (Woo et al., Down-regulates Yes (Samikkannu et al., 2004) mRNA transcripts 2003; Wang and Casero, 2006) 15d-PGJ2 (Qin et al., Proteosomal Yes (Kim et al., 2008; 2003) degradation of ER α Kim et al., 2009; Shin et al., 2009) Genistein (Fritz et al., Down-regulates Yes (Sánchez et al., 2002) mRNA transcripts 2009)
Resveratrol (Bhat and Down-regulates Yes (Pozo-Guisado et Pezzuto, mRNA transcripts al., 2005; de and 2001) Villegas, 2007) 1α,25- (Swami et Down-regulates Yes (Koren et al., 2001) Dihydroxyvita al., 2000) mRNA transcripts min D
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4.1.4 ROS-induced suppression of ER α gene expression: Possible development of ER α-negative breast cancer phenotype
The majority of literature suggested that most breast cancers, including ER-negative breast cancers, arose from ER-positive or otherwise estrogen-responsive progenitor cells (Allred et al., 2004). Many breast cancers lost their ER α expression during the
progression of the cancer, and become ER α-negative. The transition from ER α- positive to ER α-negative phenotype has been thought to be through various pathways.
One important way that ER α-negative breast cancer could arise is through hypermethylation of CpG island on ERα gene promoter (Leader et al., 2006). In ER α- negative breast cell lines, DNA methyltransferase 1 (DNMT1) level and activity are increased between 2 to 10 folds. This is in conjunction with high levels of methylation observed in these cells (Ottaviano et al., 1994). Treatment of these cells with
inhibitors of DNA methyltransferases, such as 5-azacitidine or decitabine, could elicit
re-expression of functional ER α protein (Oh et al., 2001).
Another way to suppress ER α expression is the activation of histone deacetylases
(HDAC) which remove acetyl groups from lysine residues on histones H3 and H4.
This results in the formation of a compact structure that diminishes gene activity
(Pinzone et al., 2004). Treating ER α-negative cells with trichostatin A, an HDAC
inhibitor, resulted in re-expression of ER α mRNA in these cells in a time- and
concentration-dependent manner (Yang et al., 2000).
The hyperactivation of MAPK can also lead to loss of ER α in breast cancer cells (Oh
et al., 2001). Here, ER α-negative cells were found to have high levels of growth factor receptors such as epidermal growth factor receptor or c-erbB-2 which are down-stream targets of Raf-1. Inhibiting Raf signaling via treatment with the MEK
178 inhibitors PD 098059 or U0126 resulted in re-expression of ER α in these ER α- negative cells.
Interesting, we showed that chronic exposure of ER α-positive cells to H 2O2 could lead
to a sustained low expression of ER α. The ROS production system and antioxidant
defense of cancer cell are often deregulated, leading to the increase of ROS in cancer
cells as compared to normal cells. The increase of ROS in cancer cells is thought to be
involved in the tumor initiation, promotion and metastasis (Wang and Yi, 2008). On
the other hand, further aberrant de-regulation of the ROS production system in the
cells could increase the amount of ROS in the cells such that it becomes detrimental.
Treatment of cancer cells with chemotherapeutic agents could also expose the cells to
higher amount of ROS (Pelicano et al., 2003; Zhou et al., 2003; Goto et al., 2003;
Inayat-Hussain et al., 2002). Such chronic increase of ROS in breast cancer cells
could inhibit cell growth, down-regulate ER α expression and deprive cells of
proliferative estrogenic signaling pathway. However, due to high adaptability and fast
mutation rate of cancer cells, a sub-population of them might accumulate enough
changes in their cellular signaling pathways to overcome the limitations and take
advantage of the elevated ROS level. This group of cells would be ER α negative and capable of growing in the absence of estrogen.
One example to illustrate this idea is through the fulvestrant-resistance model.
Fulvestrant is a drug used in the treatment of breast cancer where it depletes the cell of ER α by inducing rapid degradation of the receptor (Dauvois et al., 1992). Like chronic treatment of cells with ROS, chronic treatment of cells with fulvestrant leads to low expression of ER α. One report has shown that breast cancer cells treated with
fulvestrant grew significantly slower then untreated cells in the initial stage. However,
after a three month period of growth suppression, these ER α-negative cells began to
179 proliferate in fulvestrant at rates similar to those of untreated wild-type cells
(McClelland et al., 2001). These cells were found to have increased expression of a
number of components involved in the EGFR/MAPK signaling pathway which allowed them to grow even in the absence of ER α and estrogen.
Our data suggest that chronic increase of intracellular ROS could be a plausible
mechanism for breast cancer cells to loss their ER α expression and become estrogen
independent during tumor progression. Through acquiring additional survival
mechanisms, such cells can thrive in an ER α-negative state and will no longer be sensitive to anti-estrogenic drugs that take advantage of cancer cells’ dependence on estrogen.
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4.2 REDOX REGULATION OF NHE1
It has been demonstrated that cellular redox status plays a role in maintaining intracellular pH (Pervaiz and Clément, 2002). Increase of H2O2 in the cell leads to
intracellular acidification and apoptosis in tumor cells (Clément et al., 1998). NHE1 is
an important exchanger involved in pHi homeostasis. In addition, NHE1 has been
shown to facilitate tumor development and maintain tumorigenic phenotype (Reshkin
et al., 2000). In recent years, we have identified regulation of NHE1 gene expression
as a possible mechanism involved in ROS-mediated regulation of tumor cell’s
response to apoptosis (Akram et al., 2006). Further studies have shown that H 2O2
mediates down-regulation of 1.1kb mouse NHE1 promoter via an oxidative and
caspase dependent mechanism (Kumar et al., 2007). In the present study, we provide
more evidence to show that H2O2 is the signaling molecule that induces repression of
NHE1 gene transcription.
4.2.1 H2O2 regulates an important region of NHE1 promoter
Our results showed that the 0.15kb region of NHE1 promoter was the minimal region
required for full transcriptional activity in both rat L6 myoblast and human MCF7
cells. This is reminiscent of previously published data where deletion of NHE1
promoter up to the 0.15kb region resulted in 25 % decrease in total promoter activity
in NIH 3T3 cells (Dyck et al., 1995). As shown by us and others, removing the 33 bp
of DNA from the 0.15kb NHE1 promoter decreased the promoter activity drastically
(Dyck et al., 1995; Dyck and Fliegel, 1995). In addition, we found high conservation
between the 0.15kb region of the NHE1 proximal promoter amongst various species.
This evolutionary conservation highlights the importance of this region for the
transcription of NHE1 and this region is likely to be regulated by similar set of
181 transcription factors. In this regard, Façanha et al have compared 500 bp of the porcine NHE1 promoter sequence with NHE1 promoter of different species. The similarity between the porcine, human, rabbit and mouse NHE1 promoter was 78, 76 and 75 % respectively (Façanha et al., 2000). In line with our conclusion, the authors deduced that broadly similar regulatory mechanisms for NHE1 transcription exist amongst the different mammalian species.
4.2.1.1 Pattern of regulation
As the 0.15kb of NHE1 promoter is important for the transcription of NHE1, its inhibition suggests that H2O2 is targeting crucial transcription factor of NHE1. H2O2
was shown to down-regulate the activity of the 0.15kb NHE1 promoter region sharply
at a low concentration of 50 µM after 24 h. The suppression of 0.15kb NHE1
promoter activity occurs in two phases. First, there was a slight inhibition at 1 h post
treatment followed by a return to basal state steadily up to 5 h. The promoter activity
then decreased from 5 h onwards until 24 h. Cellular antioxidants act as reducing
agents and protect the cell from damage induced by oxidative stress. An example of
such antioxidant is GSH. With the help of various GSH peroxidases, GSH functions
as a major thiol antioxidant to detoxify peroxides (Fratelli et al., 2005). The decrease
of the promoter activity in 1 h may represent the initial oxidation of transcription
factor at the 0.15kb promoter. As the amount of ROS introduced is not too high, the
oxidized proteins could be quickly reversed back by antioxidant systems in the cells.
However, H2O2 can activate other signaling cascade to result in a second round of
ROS production which our lab has recently found to be peroxynitrite (Chang, unpublished data). Indeed, when exogenous aqueous peroxynitrite was added to the cells, 0.15kb NHE1 promoter was down-regulated. Other than NHE1, peroxynitrite
182 has been shown to down-regulate neutrophil surface expression of L-selectin and up- regulate expression of CD11b in a concentration-dependent fashion (Zouki et al.,
2001).
4.2.1.2 Oxidative nature of H 2O2 in suppressing gene expression
Using thiol antioxidants (NAC), as well as chemical reducing agents, DTT and BME,
we investigated the nature of H2O2-mediated suppression of 0.15kb NHE1 promoter activity. Preincubation of cells with ROS scavenger NAC and reducing agents, DTT and BME, prevented the inhibition of 0.15kb NHE1 promoter activity by H 2O2. The results demonstrate the presence of an oxidation phase in the regulation of NHE1 transcriptional activity mediated by H2O2. This is similar to our previous study on the
full length 1.1kb NHE1 promoter, and it suggests that the same transcription factor is
targeted by H2O2 in the two promoters (Kumar et al., 2007). The data with thiol-
oxidizing agent diamide showed that it mimicked the effect of H 2O2 in suppressing
the 0.15kb promoter activity. Indeed, our experimental data indicates that the
inhibition of NHE1 promoter activity could involve the oxidation of thiol moiety of
cysteine residue that might be present in the transcription factors necessary for NHE1
expression.
The reversible oxidative inactivation of transcription factors has been proposed to be
important in cellular response to oxidative stress. Other than the transcription of
NHE1, human cytochrome P450 1A1 (CYP1A1) gene expression has also been
reported to be down-regulated by H 2O2 (Morel and Barouki, 1998). The transcription factor of CYP1A1, nuclear factor I (NFI), contains redox-sensitive cysteine residue at the DNA-binding domain which renders it liable to oxidative inhibition by H2O2
(Morel and Barouki, 2000). It has been suggested that intracellular GSH-dependent
183 pathway could reactivate the DNA-binding activity of NFI proteins during oxidant stress through the reduction of oxidation-sensitive cysteine residue on NFI
(Bandyopadhyay et al., 1998). Our results show that the down-regulation of the
0.15kb promoter activity by H2O2 lasted for 24 h, after which it started to recover.
This could signify the reversal of the thiol oxidation of NHE1 transcription factors by
cellular antioxidant system or new transcription factors being made to replace those
that were degraded.
4.2.2 Targeting NHE1 expression through ROS-producing agents
Our lab has previously published that 15d-PGJ2 down-regulated human NHE1
expression in a PPAR γ-dependent manner (Kumar et al., 2009). From computational analysis tool, Peroxisome Proliferator Response Element (PPRE) Search, human
NHE1 promoter was predicted to contain 2 PPAR γ binding sites. Using the same program, mouse NHE1 promoter was not found to contain any PPRE (Appendix B).
Interestingly, our experimental data showed that 15d-PGJ2 at 20 µM could down-
regulate NHE1 0.15kb promoter activity. It is logical to believe that the repression of
the mouse NHE1 promoter activity is PPAR γ independent. Indeed, ROS was subsequently found to mediate the down-regulation of the promoter activity. Our finding is in line with other studies which have documented that many effects of 15d-
PGJ2 is independent of its receptor PPAR γ, but rather rely mainly on intracellular
ROS produced (Kim et al., 2008; Kim et al., 2009; Shin et al., 2009; Lecomte et al.,
2008). For example, the addition of 15d-PGJ2 to rat vascular smooth muscle cells resulted in up-regulation of heme oxygenase-1 (HO-1) which cannot be blocked by
PPAR γ antagonist GW9662. Instead, NAC could inhibit the up-regulation of HO-1
184 induced by 15d-PGJ2 completely; suggesting a ROS-mediated activation of HO-1 expression (Lim et al., 2007).
Although our previously published data did not implicate ROS as a mechanism for
NHE1 down-regulation by 15d-PGJ2, we are finding evidences that ROS is produced and could also be involved in the down-regulation of NHE1 in human breast cancer cells (Zhou, unpublished data). Along the same lines, resveratrol, a polyphenolic compound found in red wine, has recently been reported to regulate the expression of
NHE1 by repressing its promoter activity (Jhumka et al., 2009). H 2O2 was found to be
produced in cells treated with resveratrol and is critical for the suppression of NHE1
gene expression. Taken together, treatment of cells with ROS-producing agents could
be an effective strategy to target the expression of NHE1.
Over-expression of NHE1 has been shown to facilitate tumor development and
maintenance of tumorigenic phenotype. The down-regulation of NHE1 could sensitize
human breast cancer cells to drug-induced apoptosis (Kumar et al., 2009). Therefore,
transcriptional repression of NHE1 could potentially be a promising approach to
breast cancer therapy. Using 15d-PGJ2, the expression of NHE1 would be suppressed
through both PPAR γ-dependent and ROS-dependent mechanisms. It could also
decrease the ER α expression through ROS, thus, diminishing the proliferative actions of estrogen. The ROS produced by 15d-PGJ2 can induce apoptosis in cancer cells that are already stressed by elevated oxidative cellular environment. ROS could also regulate various survival proteins such as Akt and JNK so that the cells are geared towards growth arrest and finally apoptosis (Shin et al., 2009). As 15d-PGJ2 targets multiple survival pathways in the cancer cells; the likelihood of tumor cells developing resistance to it would be reduced. As such, 15d-PGJ2 may be a good candidate for used as an adjuvant in chemotherapy.
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4.2.3 AP2 is a transcription factor of NHE1: True or false?
Out of the three possible transcription factors identified by the bioinformatics software as possible transcription factor for NHE1, AP2 was selected for further investigation. One main reason was because AP2 has been shown to bind to the
0.15kb NHE1 promoter in vitro (Dyck et al., 1995; Yang et al., 1996). The second reason being AP2 has been reported to be a redox regulated protein where H 2O2 could inhibit its DNA-binding activity to AP2 consensus sequence ((Huang and Domann,
1998). Consistent with Huang and Domann’s study, our results also indicate that reversible thiol oxidation of a transcription factor could be responsible for the inhibition of NHE1 gene expression upon treatment with H2O2. As AP2 has only been
shown to bind to NHE1 promoter in vitro we sought to determine if AP2 is indeed the
transcription factor for NHE1 in vivo in NIH 3T3 mouse embryonic cells stably expressing 0.15kb of NHE1 promoter.
Through mRNA detection, AP2 α was found to be the only AP2 isoform expressed in
NIH 3T3 cells. However, when AP2 α was knocked down in the cells by up to 90%, there was no change in the NHE1 promoter activity, mRNA expression level and protein level. Even when AP2 α was knocked down, H 2O2 could still suppress the
NHE1 0.15kb promoter activity. Over-expression of AP2 α also resulted in no change
in the NHE1 protein, mRNA level and promoter activity. These data strongly suggest
that AP2 α is not the transcription factor for NHE1 and it is not the target of H 2O2- mediated down-regulation of the 0.15kb promoter activity in our system. Supporting our data that AP2 is not the transcription factor for NHE1 in vivo comes from a study which examines NHE1 protein expression in transgenic mice lacking AP2 α. In it, 18-
day-old AP2 α heterozygote mice show no significant changes in NHE1 expression in
186 heart, lung, liver, kidney and brain, while AP2 α null mice showed a large increase in
NHE1 protein expression in the brain (Rieder and Fliegel, 2003).
The discrepancy between our data and the data from Dyck et al could due to differences in methods used to address the role of AP2 in NHE1 transcription. While our experiments were conducted within live cells, most of the experiments showing
AP2 as the transcription factor for NHE1 in Dyck’s study were in vitro setups. Using
EMSA, Dyck showed that purified human AP2 α and NIH 3T3 nuclear extract could
bind to the DNA within the 33 bp region between the 0.15kb and 0.12kb of the NHE1
promoter. Although this method could provide evidences that a specific protein can
bind to a specific sequence of DNA in certain conditions, the limitation of EMSA as a
technique for representing DNA–protein interactions in vivo, is that the physiological conditions will not be identical to those used in vitro (Smith and Humphries, 2009).
EMSA also does not take into account of the chromatin structure and the various modifications that take place on the DNA which could enhance or repress transcription of a particular gene. In order to supplement the EMSA data, a Chromatin immunoprecipitation (ChIP) assay would be the next logical step. The assay uses specific antibody that recognizes the transcription factor of interest to pull down all
DNA sequences that the transcription factor is binding to in the cell. Polymerase chain reaction (PCR) is then carried out to determine if the transcription factor binds to the specific DNA of interest. Having confirmed that a particular transcription factor binds to a certain stretch of promoter region in vivo , silencing of the transcription factor should be carried out to determine its effects on target gene expression.
187
The present study has revealed that AP2 might not be the transcription factor responsible for the transcription of NHE1. This is a significant finding as it means that the transcription factor crucial for NHE1 expression is still not discovered. Using bioinformatics, SP1 and COUP were identified as possible transcription factor candidates that sit on the 33 bp of DNA between the 0.15kb and 0.12kb of NHE1 promoter. Attempts have been made to verify SP1 as the transcription factor for
NHE1 since it has also been shown to be a redox regulated gene. However, different sets of siRNA purchased from 2 different companies failed to knockdown gene expression of SP1 (Appendix C). Therefore, future work should focus on getting effective siRNA for SP1 and COUP to verify if these 2 transcription factors are factors binding to the 33 bp 5’ region of 0.15kb NHE1 promoter.
188
4.3 ER α AND NHE1: TWO IMPORTANT MEDIATORS OF CANCER
CELL SURVIVAL
ER α and NHE1 are proteins that have distinct functions in the cells. ER α is a transcription factor that mediates the effects of estrogen while NHE1 is a pH regulator.
Although seemingly unrelated, these two proteins play important roles in the survival of tumor cells. ER α contribute to tumor cell survival by increasing cell proliferation
rate through shortening the overall cell cycle time, and enhancing tumor cell’s
resistance to chemotherapeutic drugs (Teixeira et al., 1995). NHE1 plays a role in
tumor cell survival by maintaining an alkaline intracellular pH which is unsuitable for
apoptosis to occur. Tumor cells also exploit the permissive effect of NHE1 on cell
proliferation where the constitutive activity of NHE1 allows cancer cells to progress
through cell cycle faster. Thus, both ER α and NHE1 promote cancer cell survival by
increasing cell proliferation and preventing apoptosis.
4.3.1 H2O2 regulates ER α and NHE1 in similar ways
In this study, the regulation of ER α and NHE1 by H2O2 were found to be similar in
various aspects and they are summarized in Table 4. First, both of the genes are
down-regulated at the mRNA level. The down-regulation of NHE1 mRNA was due to
H2O2 suppression of NHE1 promoter activity. While it is unclear whether the down- regulation of ER α mRNA is due to suppression of promoter activity or destabilization
of the mRNA, it is known that redox sensitive transcription factors are involved in
ER α transcription. Therefore, it is plausible to think that ER α promoter activity could
also be suppressed by H 2O2. Although the target of H 2O2 that leads to the repression of these two genes might or might not be the same molecule, it is clear that thiol oxidation is involved in the down-regulation both genes. Thus, the target would
189 contain reactive cysteine that is oxidizable by diamide. The oxidative repression of these two genes is reversible by reducing agent DTT. In the absence of reducing agent, the gene expression is inhibited for 24 h, and started to recover by 48 h. In addition, caspase was found not to be involved in the regulation of these two gene expression by H 2O2. Finally, both genes expression were suppressed by 15d-PGJ2 and this down- regulation is mediated through ROS production in the cells.
As proteins important in cancer survival, ER α has long been the target of endocrine
treatment in breast cancer therapy (Jordan, 2004), while NHE1 has been proposed to
be a potential chemotherapeutic target (Liu et al., 2008; Lauritzen et al., 2010). It is
interesting that the expression of these two proteins is regulated by H2O2 in a similar
manner. These results provide new insight into the mechanistic action of H 2O2 on
gene down-regulation and suggest that H 2O2 responsive genes are regulated via
common signaling pathways. Moreover, our research has given evidences that ROS-
producing chemotherapeutic agents could be used to target ER α and NHE1 expression
together to negate the survival advantage mediated by these two proteins. Clinically, a
drug with such combinatorial effects could be use as an adjuvant to enhance the
efficacy of currently available cancer treatments.
Table 4: Similarities between the down-regulation of ER α and NHE1 expression by H2O2. ER α NHE1
Down-regulation at mRNA Yes Yes level by H 2O2 Down-regulation is Yes Yes reversible Can be thiol-oxidized Yes Yes Reducing agent can reverse Yes Yes H2O2 induced down- regulation Caspase not involved in Yes Yes, in 0.15kb promoter down-regulation Down-regulated by 15d- Yes Yes PGJ2 through ROS
190
4.3.2 Relevance of ER α and NHE1 expression as prognosis markers in breast cancer.
Over 60 % of patients who have breast cancer express ER α and this group of patients
generally has better prognosis than the patients who do not express ER α. In addition to ER α, we propose that that NHE1 expression level might be a potential prognosis
marker for breast cancer patients. Some preliminary results from our lab suggest that
NHE1 expression level is increased in breast cells that are cancerous but not yet
aggressive. However, the aggressive tumor cells have very low levels of NHE1
compared to both non-tumorigenic cells and non-aggressive cancer cells. Appendix D
shows the results comparing the non-tumorigenic breast cells (184A1 and MCF10A),
the non-aggressive breast cancer cells (MCF7, SK-BR3, BT-474 and HCC-90) and
the aggressive breast cancer cells (MDA-231 and BT-594). Supporting this is clinical
data from 97 patients who have breast cancer (Appendix E). These patients were
followed up over a median of 6.4 years and study showed with statistical significance
that those who have high level NHE1 protein in their tumor fared better in terms of
survival than those who have low level NHE1 expression. When we factor in the ER α
status of these patients we found that ER α-positive, NHE1-positive patients seemed to fare better then ER α-negative, NHE1-negative patients. However, due to the small sample size the results are not yet statistically significant.
Studies have shown that NHE1 is important in the initiation and progression of cancer.
However, no report had yet associated the decrease in NHE1 with increased aggressiveness of breast cancer. Therefore, more research should be conducted to further verify the findings and understand the etiology for the loss of NHE1 in aggressive breast tumors.
191
4.4 FUTURE WORK
4.4.1 To determine if H 2O2 affect the stability of ER α mRNA
To further understand the molecular mechanism by which H2O2 suppress the gene
expression of ER α, it is crucial to determine what causes the ER α mRNA level to decrease upon treatment. As mentioned in the discussion section, it could be a result of either suppression of transcription or increased mRNA instability. Knowing at which level ER α expression is being affected would provide new directions for future research on H2O2-mediated ER α suppression. Due to the complexity of ER α
transcriptional regulation, it would be logical to approach this question by addressing
the mRNA instability of ER α after H 2O2 treatment. Through the process of elimination, if the mRNA stability of ER α is found not to be affected by H 2O2, one could deduce that H2O2 targets the transcription of ER α instead.
Attempts were made to determine if there is any changes in the mRNA stability upon
H2O2 treatment using classic transcription inhibitor Actinomycin D and 5,6-dichloro-
1-ß-D-ribobenzimidazole (DRB ). However, the MCF7 cells used in this study was
insensitive to treatment with the transcription inhibitors. When used alone or in
combination, Actinomycin D and DRB could not block the transcription of ER α in
MCF7 (results not shown). Thus, because of experimental constraint, we were not able to determine if the down-regulation of ER α mRNA level was due to a change in the half-life of the mRNA. To overcome this limitation, other methods of measuring mRNA half-life could be utilized. One way is to employ other pharmacological inhibitor such as α-Amanitin which binds to RNA polymerase II and III, preventing
incorporation of new ribonucleotides into newly synthesized RNA (Cassé et al., 1999;
Gong et al., 2004). Another approach would be to use pulse chase method where
radioactive nucleotides are incubated with cells to allow for incorporation of these
192 nucleotides into the RNA. The time-dependent decay of newly synthesized mRNA would then monitored (Ross, 1995; Chen et al., 2009).
4.4.2 To investigate the mechanism of ROS-producing compounds in suppression of ER α expression
In this study, we have shown that 15d-PGJ2 inhibits ER α expression through ROS generation. In order to further explore the pathway leading to this inhibition, the mechanism for ROS production by 15d-PGJ2 in MCF7 cells could be determined.
Although mitochondria and NADPH oxidase activation have been shown to be the source of ROS produced by 15d-PGJ2, it has not been proven in our system. Thus, specific inhibitors of the mitochondria complexes and NADPH oxidase could be tested for their effects on 15d-PGJ2-induced ROS generation in MCF7 cells. The findings obtained could help to ascertain if 15d-PGJ2 produces ROS through similar mechanisms in different cell types. In addition, the nature of ROS produced by 15d-
PGJ2 in MCF7 cells should also be investigated. The information derived from this work could be used to validate our hypothesis that H2O2 is involved in 15d-PGJ2-
mediated down-regulation of ER α expression.
Other than 15d-PGJ2, many compounds have been identified to down-regulate ER α
and shown to produce ROS (Table 3). Further experiments would be needed to test if
these agents also suppress ER α expression via ROS production. Should it be found that ROS is the main mechanism involved in the inhibition of ER α expression by these agents, it would provide supporting evidence that redox-regulation of ER α
expression is an important and relevant signaling pathway in the cells.
193
4.4.3 To identify the transcription factor that regulates NHE1 transcription
From the current research, AP2 was found not to be the transcription factor of NHE1.
As the 0.15kb NHE1 promoter is important for the transcription of NHE1, it would be of scientific merit to investigate the transcription factor that binds to this region. This work could yield new information for NHE1 transcriptional regulation and allow us to further validate the target involved in H2O2-mediated suppression of NHE1 promoter activity.
Other than verifying SP1 and COUP as transcription factors for NHE1 0.15kb promoter, another method to identify the transcription factor binding to the 33 bp region would be to conduct a protein-DNA binding by streptavidin-agarose pull down followed by identification using mass spectrometry (MS). In this method, oligonucleotides labeled with biotin are allowed to bind with nuclear extracts. The protein complex together with the oligonucleotides is pulled down using streptavidin- agarose beans. The complexes are subsequently dissociated and the individual
constituents are resolved by SDS–PAGE and revealed by staining. Proteins are then
be subjected to limited proteolytic cleavage and analyzed by matrix-assisted laser
desorption ionization-time of flight (MALDI-TOF) MS. Peptide mass fingerprinting
in this way allows identification of proteins by comparison with simulated digests of
expressed protein databases (Drewett et al., 2001). The advantage of this method is
that no prior knowledge of what might bind to the oligonucleotides is required. It is
also able to resolve and identify proteins in the complexes that are too large to be
resolved using EMSA. However, ChIP assay would still be required to confirm the
binding of the transcription factor intracellularly.
194
4.5 MODEL AND CONCLUSION
15d-PGJ2 H O Drugs 2 2
DTT ROS NAC
reversible-oxidation Diamide ?
NHE1 mRNA Tumor growth ER mRNA NHE1protein ER response genes ER protein
Figure 57: Model of the down-regulation of ER α and NHE1 by H2O2.
Taken together, our findings provided a new perspective to ER α as a redox-regulated protein. Previous research had shown that the ER α DNA-binding is subjected to redox modulation. The current study places the gene expression of ER α itself to be targeted
by H 2O2 and potentially by other reactive species. The similarities in the regulation of
ER α and NHE1 by H 2O2 suggested that there might be a common pathway in the
control of gene expression by H2O2. Further examination of the mechanisms leading
to the down-regulation of ER α mRNA level would provide better comprehension of
how H2O2 represses gene expression. The discovery that AP2 is not the transcription
factor for NHE1 is a major finding that would challenge the scientific community’s
understanding of NHE1 transcriptional regulation. Thus, it is paramount to investigate
the target involved in H2O2 repression of NHE1 gene expression. In the clinical aspect,
195 emerging evidences are showing that a number of anticancer agents function through the production of reactive species such as H2O2 to kill or suppress tumor cells growth.
Increase of ER α and NHE1 levels in breast cancer have been shown to be involved in
carcinogenesis and tumor growth. The targeting of ER α and NHE1 expression
together by ROS could represent a novel approach to shut down breast cancer cells’
growth machinery and prove more efficient in killing these cells.
196
APPENDICES a) >Human_ESR1_promoter ATGTGCGTGTGCATGTTTAACTTTTAATTCAGTTAAAAACTTTTTTCTATTT GTTTTTCATCTGGATATTTGATTCTGCATATCCTAGCCCAAGTGAACCGAG AAGATCGAGTTGTAGGACTAAAGGATAGACATGCAGAAATGCATTTTAAA AATCTGTTAGCTGGACCAGACCGACAATGTAACATAATTGCCAAAGCTTT GGTTCGTGACCTGAGGTTATGTTTGGTATGAAAAGGTCACATTTTATATTC AGTTTTCTGAAGTTTTGGTTGCATAACCAACCTGTGGAAGGCATGAACAC CCATGTGCGCCCTAACCAAAGGTTTTTCTGAATCATCCTTCACATGAGAAT TCCTAATGGGACCAAGTACAGTACTGTGGTCCAACATAAACACACAAGTC AGGCTGAGAGAATCTCAGAAGGTTGTGGAAGGGTCTATCTACTTTGGGAG CATTTTGCAGAGGAAGAAACTGAGGTCCTGGCAGGTTGCATTCTCCTGAT GGCAAAATGCAGCTCTTCCTATATGTATACCCTGAATCTCCGCCCCCTTCC CCTCAGATGCCCCCTGTCAGTTCCCCCAGCTGCTAAATATAGCTGTCTGTG GCTGGCTGCGTATGCAACCGCACACCCCATTCTATCTGCCCTATCTCGGTT ACAGTGTAGTCCTCCCCAGGGTCATCCTATGTACACACTACGTATTTCTAG CCAACGAGGAGGGGGAATCAAACAGAAAGAGAGACAAACAGAGATATAT CGGAGTCTGGCACGGGGCACATAAGGCAGCACATTAGAGAAAGCCGGCC CCTGGATCCGTCTTTCGCGTTTATTTTAAGCCCAGTCTTCCCTGGGCCACCT TTAGCAGATCCTCGTGCGCCCCCGCCCCCTGGCCGTGAAACTCAGCCTCTA TCCAGCAGCGACGACAAGTAAAGTAAAGTTCAGGGAAGCTGCTCTTTGGG ATCGCTCCAAATCGAGTTGTGCCTGGAGTGATGTTTAAGCCAATGTCAGG GCAAGGCAACAGTCCCTGGCCGTCCTCCAGCACCTTTGTA b)
Appendix A: PPRE present in human ER α promoter A. a) 1kb sequence of human ER α promoter A. b) Results of bioinformatics analysis of human ER α promoter A using PPRE Search software.
197 a) >Human_NHE1_promoter TTTAAGTTCCACTACTTAAAGCATGCCACTGTTGTGGGTTGAATTGTGTCC CGGCAAAAGAAGTTGAAGTCCTAATGCCCAGTGCCTATGAAGATGGACTA ATTAGGATGCAGTCTTTGAAGATGTTCAGGTTAAGATGAGGTAATTACGTT GGATTTCTAATCCAATGACTGGTGTGGAATCGCATATCAAGCTTTCCAGTG ATTCCATTGTACAGCCATGATCCCTTGAACCTCACCAATTTCAACCAAACT ATAGGTTCAAACCTTATAAAAAGGGGAAATCTGGCTGGGGGTTGTGGCTT ACCCCTGTAATCCCAGCACTTTGGGAGGCCGAGGCGGGTGGATCACCTGA GGTCAGGAGTTCGAGACCAGCCTGACCAACATGGTGAAACCCCATCTCTA CTAAAAATACAAAAATTAGCTGGACGTGGTGGTGGGTGCCAGTAATCCCA GCTACTCTGGAGGCTGAGGCAGGAGAATCGCTTGAATCCAGGAGGCAGA GGTTGCATTGAGCTGAGATGGCGCCACCGCACTCCAGCCTGGGCAACAAG AGCCAGACTCTATCTCAAAAAAAATAAAAATAAAAATAAAGTGTGGGCG GGGAATCTGGACGCAGAGACAGAGACACCAGGAGAACTCCATGGAATAC CAGATAGTCCTAACAAACCACTGGAAGGTAGGAGAAAGGCATGGGACAG ATTCTCCCTCATAGCTCTCAGCTGAAACCAACCCTGCCAACACCTAGATCC GACCTCCAGCCTCCAGAACTGTGAGACAATCAATTTCTGTTGTTGCAGCCA CCCAGTTTGGGGTGATACTTTGTTACGGCAGCCCTAGTAAGCAATACAAC TACTTGCATAGTAGCCAGGGGACTCTCTTCACCTGTTTCCTCATCTGTAAA AGTGGAATTGTAATAATGTGCCAGGGTGCATTCCAAATAGTTTACACGGA TTGTCTCAGTCATTACATCATCCCTCTGACATAGTCACTATTACTGTCTCTA CTTAACAGATGAGAAAGTTGTGAAACAGGTTAAGTAACTTGCTCAAGGTC ACACGGTAACTAAATACATAAACTAATAATACATTCTTCACAGGATTATT CGAAAGCCCTTATGAGACTGCAGATGTGGACGTGAAATCGTTTTGTAAGT AGTCGGCATTTTACTCGCGTTAGTGAGGTTCTCTGTATATTCAGGACTTTTT TTTTTTTTTTTTTTTGTCATCTCTGACTCTCCTTCCTCTTCCTACGCGATACT TCTTTCCCTCGGCGACAGGGGCCGCTGCGCTGGGCGGGTGCCGACGGTCT CTCTAGCCCGCCGCACCGGCTGCTCGCTGGTGCCTATAAGTGACAGCGCC GGGCTCAGCTAGGCTTCAGTCTGCTGCGGCC
b)
198 c) >Mouse_NHE1_promoter CAGATAATAAAAATCCTTCTTTTCTCTTTAAACCAGACAGACAGACAGAC AGATAGACAGACAGACAAAAACAATACCCAGCCTCTTACTTTTTGGTTGT TTCACTACTGTTATGAGTTGAAACACGCCCTTTCAAAAGAAGCTGAAGTC CTACCCCTGCCCACCCCTCAGTGCCCGAGAATGTGGCTTTATTTAGATGAG GGGTTTTGTTGTTGTTGTTGTTGTTTTAAATCTCTTTGAGACAGGGTCTCCC TACTGACCTCAGCCTGGTCTAGAACTCACTTTGTTACACAAGCTGACGTTG AGTTAAAAATTCCTCTTTCTTTAGTTTCCCAAATGCTGGAATTACAGTTTCC CACCGTTCGTGGCTCAGATTGGGTTTTTGAAGTTGATCAAGGTAAGGTACG CCATTAGGGCCAATTCCTAATTCAATTTGCTTGGAGTCCTTTTAAAGGGGG AAATTTGGACCTAAAGACGGATACAGGGAGGCTGGGGTTATGCCGCCACA GTCAAGGGAAGCCAATACATTGCCATCAAAGCACCTGTGGCTGGGATGCA GGAACGGATTTCAGCTCACACGGTCCTCAGCACAGGTCAATGCTTGGGAA CATACAGAGTGCAGATTTCTAGCCCATTTCATTCATTCATTTAACTTTATTT TGTGACACTGGGCATAGGAACCAATGGTAGACAAGCACTCTACCTCTGAA CCACGATCCCATCCCTTTCATTCTAAACATTTCTCATAGATCACCTCATTTA ACCATCTTGTTATCCTTTTGTTATAGCTATCCATCTCCACCTTACAGTTGAG AAAGCTGTGAAACAGAGAGGCTAAATAACTTGTTCCAAAGTCACATGCTA ACAAGAAAACACTTTGTAGATAGTACAGTATGTCACTCGTGTTAGATTGT ACTTTTTTTTTTTTTTTCCAATTTAGGTCTCGGCTTCCTCTTCTTACGTGACA CTTCCTTCCCTGGGCGACAGGGGCCGCTGCACCGCGCGGGCGCTGACAGG TCTCTCTAGCTTGCTGCGCCTGCTACTCGTTGGTGCCTATAAATGGCTGCG CTGTGCTCAGCGAGGCATCAGTCCGCTACCGCGGGACCC d)
Appendix B: PPRE present in human and mouse NHE1 promoter. a) 1.4kb sequence of human NHE1 promoter. b) Results of bioinformatics analysis of human NHE1 promoter using PPRE Search software. c) 1.1kb sequence of mouse NHE1 promoter. d) Results of bioinformatics analysis of mouse NHE1 promoter using PPRE Search software
199
Santa Cruz Dharmacon siSP1 10nM siSP1 20nM siSP1 50nM siSP1 10nM siSP1 NS siRNA 50nM siRNA NS 50nM siRNA NS siSP1 20nM siSP1 50nM siSP1
SP1 -105 kDa
β-actin -43 kDa
Appendix C: siRNA from two companies failed to knockdown SP1 protein expression. SP1 in MCF7 was silenced with various concentration of SP1 siRNA from Santa Cruz and Dharmacon using Lipofectamine RNAiMAX reagent as described in the Materials and Methods section. Cells were harvested 48 h post transfection. Cell lysates were analyzed by Western blot.
MDA-MB-231 MCF10A BT-474 BT-549 SK-BR3 HCC-70 184A1 MCF7
NHE-1 -100 kDa
-43 kDa βββ- actin
• MCF7 – non-aggressive • MDA-MB-231 – aggressive • SK-BR3 – non-aggressive • BT474 – non-aggressive • BT549 - aggressive • HCC70 – non-aggressive • MCF10A – non-tumorigenic • 184A1 – non-tumorigenic
Appendix D: NHE1 levels in different breast cell lines. NHE1 level of different breast cell lines were analysed using Western blot.
200 a)
b)
Appendix E: The expression level of NHE1 in tumor tissues is associated with clinical outcome. a) Overall survival curves between patients expressing NHE1 and patients not expressing NHE1. b) Overall survival curves amongst patients with different ER α and NHE1 status.
201
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