Oxygen regulation of vascular smooth muscle cell

proliferation and survival

by

Julie Basu Ray

A thesis submitted in conformity with the requirements

for the degree of doctor of philosophy

Institute of Medical Sciences

University of Toronto

© Copyright by Julie Basu Ray, 2009

ii

Oxygen regulation of vascular smooth muscle cell

proliferation and survival

Julie Basu Ray

Doctor of Philosophy

Institute of Medical Sciences

University of Toronto

2009

ABSTRACT

Arterial smooth muscle cells (SMCs) from the systemic and pulmonary circulations

experience a broad range of oxygen concentrations under physiological conditions. The hypoxic

response, however, has been inconsistent, with both enhanced proliferation and growth arrest

being reported. This variability precludes a definitive conclusion regarding the role of oxygen

tension in arterial disease.

In the first part of this study, we determined if hypoxia elicits different proliferative and

apoptotic responses in human aortic SMCs (HASMCs) incubated under conditions which do or

do not result in cellular ATP depletion and whether these effects are relevant to vascular

remodeling in vivo. Gene expression profiling was used to identify potential regulatory

pathways. In HASMCs incubated at 3% O2, proliferation and progression through G1/S

interphase are enhanced. Incubation at 1% O2 reduced proliferation, delayed G1/S transition,

increased apoptosis and cellular ATP levels were reduced. In aorta and mesenteric artery from

hypoxia exposed rats, both proliferation and apoptosis are increased after 48hrs. p53 and

p21expression is differentially affected in HASMCs incubated at 1% and 3% O2. Hypoxia

iii

induces a state of enhanced cell turnover, conferring the ability to remodel the vasculature in

response to changing tissue metabolic needs while avoiding the accumulation of mutations that

may lead to malignant transformation or abnormal vascular structure formation. A unifying

hypothesis in which events at the G1/S transition and apoptosis activation are coordinated by

effects on p53, p21, their downstream effector genes and regulatory factors is proposed.

Differences in the contractile responses of systemic and pulmonary arterial smooth

muscle cells to hypoxia are well studied. Differences in proliferation and survival are anticipated

because of differences in embryonal cell origin, oxygen concentrations within their respective

microenvironments and in cellular energetics but these responses have not been directly

compared.

In the second part of the study, human pulmonary arterial SMCs (HPASMCs)

proliferated at oxygen concentrations which inhibited cell growth in HASMCs. HPASMCs

survived and maintained their intracellular ATP levels at levels of hypoxia sufficient to deplete

ATP and induce apoptosis in HASMCs. In vivo studies in rats show proliferation and apoptosis

in main or branch PASMCs only after 7 days of hypoxia. VSMCs are able to proliferate under

hypoxic conditions as long as cellular ATP levels are maintained. HPASMCs have an enhanced

capacity to maintain cellular energy status compared to HASMCs and hence their viability is

preserved and the proliferative response predominates at lower oxygen concentrations.

iv

“I have become my own version of an optimist. If I can't make it through one door, I'll go

through another door - or I'll make a door. Something terrific will come no matter how dark the

present.”

Rabindranath Tagore

“Take up one idea. Make that one idea your life - think of it, dream of it, live on that idea.

Let the brain, muscles, nerves, every part of your body, be full of that idea, and just leave every

other idea alone. This is the way to success.”

Swami Vivekananda

v

ACKNOWLEDGMENTS

The path towards this work spans several years of research and it is a pleasure to thank the

many people who made this thesis possible. I acknowledge my debt to all those who have helped

along the way, who have been involved and contributed to the presented ideas and understanding

gained.

It is difficult to overstate my gratitude to my Ph.D. supervisor, Dr. Michael E Ward. With

his enthusiasm, his inspiration, and his great efforts to explain things clearly and simply, he

helped to make research fun for me. I am indebted to his continued encouragement and

invaluable suggestions especially during my thesis-writing period. He has been a wonderful

guide and a great teacher. I would also like to include my gratitude to Dr. Linda Penn, my co-

supervisor, and Dr. Philip Marsden who have provided support for this research all along the

way.

I am deeply indebted to my student colleagues at the Terrence Donnelley Research

Laboratories at the St. Michael’s Hospital, for providing a stimulating and fun environment in

which to learn and grow. I would specially like to thank Jeff He, Lakshmi Kugathasan, Massey

Rezai and Yupu Deng for their extremely valuable support, and insights. Many others who have

been involved also deserve recognition. It is, however, not possible to list them all here. Their

support in this effort is, however, greatly appreciated.

I would like to thank the many people who have initiated me into the rites of science - my

high school teachers at Calcutta Girls’ High School, my undergraduate teachers at Presidency

College, and my graduate teachers at the University of Calcutta, India.

vi

I wish to thank my friends in high school, college and university, and my friends in Toronto,

Buffalo and Boston, whose continued support helped me get through the difficult times, and for

all their emotional support, camaraderie, entertainment, and caring they provided.

Finally I want to thank my family. A special thought is devoted to my parents, Anil and

Jayanti Ghosh for a never-ending support. They bore me, raised me, supported me, taught me,

and loved me. I am indebted to my entire family for providing a loving environment for me.

The encouragement and support from my husband, Indranill, have always been a powerful

source of inspiration and energy.

Lastly, and most importantly, I thank from the bottom of my heart my son, Ishan, whose

adjustment and sacrifice of many a childhood demand and wish have helped me sail through to

this goal. To him I dedicate this thesis.

vii

CONTRIBUTIONS

The work presented in Chapter 2 has been published in Am J Physiol Heart Circ Physiol

Feb 2008; 294: H839 - H852. Basu Ray J, Arab S, Deng Y, Liu P, Penn L, Courtman DW, Ward

ME. Oxygen Regulation of Arterial Smooth Muscle Cell Proliferation and Survival. Permission

has been obtained from the American Physiological Society and all of the authors for inclusion

of the paper in the thesis.

As the first author of the publication, I contributed to study design, figure making and

manuscript writing. I performed all of the experiments and data analysis except Table 2.1:

Experiment and data analysis was done by Dr. Sara Arab from Dr. Peter Liu’s lab at the Toronto

Genomic Core Centre at the Hospital for Sick Children. Dr. Yupu Deng has helped with the

animal sacrifices for Figures 2.10 and 2.11 and confocal microscopic analysis of sections.

The work presented in Chapter 3 has been written into a manuscript and is expected to be

submitted for publication before December 2009. As the first author of this manuscript, I

contributed to study design, figure making and manuscript writing. I performed all of the

experiments and data analysis. Dr. Yupu Deng has helped with the animal sacrifices for Figures

3.7 and 3.8.

In the Supplement chapter, experiment design and data analysis for Figure S1 has been

done by Karen Ho from Dr. Philip A Marsden’s lab.

The research work has been funded by Canadian Institute of Health Research Grant and

Keenan Collaborative Research Award by Keenan Research Foundation, Toronto.

viii

TABLE OF CONTENTS

ABBREVIATIONS xi

LIST OF FIGURES xiii

LIST OF TABLES xvi

CHAPTER 1 Review of literature

1.1 Introduction 2

1.2 Systemic and pulmonary circulations 2

1.3 Vascular smooth muscle cells 5

1.4 Oxygen delivery 6

1.5 Physiological responses to hypoxia 9

1.5.1 Systemic responses 9

1.5.2 Regulation of cellular metabolism 10

1.6 Regulation of gene expression 11

1.6.1 Hypoxia-inducible factors 12

1.6.2 Regulation of HIF activity 14

1.6.3 HIFs as transcriptional regulators 17

1.6.4 HIF independent transcriptional activation 21

1.7 Hypoxic regulation of mRNA stability 23

1.8 Hypoxic repression of transcription 23

1.9 Hypoxic control of protein translation 25

1.10 Cell cycle and hypoxia 25

1.10.1 The mammalian cell cycle 25

1.10.2 Effects of hypoxia on cell cycle 30

1.11 Apoptosis and hypoxia 32

1.11.1 Apoptotic pathways 32

1.11.2 Regulation of apoptosis during hypoxia 38

1.11.2.1 Role of p53 in hypoxia-induced apoptosis 38

ix

1.11.2.2 Role of Bcl-2 family proteins 39

1.11.2.3 Role of PI3-kinase pathway 40

1.11.2.4 Role of electron transport chain inhibition 41

1.12 Oxygen sensing mechanisms 42

1.12.1 Evidence of heme as oxygen sensor 43

1.12.2 NAD(P)H oxidases 44

1.12.3 Mitochondria 47

1.13 Thesis objective 50

1.14 Aims and hypotheses 52

CHAPTER 2 Oxygen regulation of systemic arterial smooth muscle cell proliferation and

survival

2.1 Introduction 54

2.2 Materials and Methods

2.2.1 Antibodies and reagents 56

2.2.2 Cell culture studies 56

2.2.3 Cell counting 57

2.2.4 [3H]-thymidine incorporation 57

2.2.5 Ki67 protein levels 58

2.2.6 Annexin V- Propidium iodide labeling 58

2.2.7 Caspase activation 59

2.2.8 TUNEL 59

2.2.9 Cell cycle analysis 59

2.2.10 Mitochondrial membrane potential 60

2.2.11 Intracellular ATP concentration 60

2.2.12 Western blotting 60

2.2.13 Microarray analysis 61

2.2.14 In vivo apoptosis 63

2.2.15 In vivo proliferation 64

x

2.3 Results 66

2.4 Discussion 89

CHAPTER 3 Oxygen regulation of pulmonary arterial smooth muscle cell proliferation

and survival

3.1 Introduction 98

3.2 Materials and Methods

3.2.1 Antibodies and reagents 101

3.2.2 Cell Culture Studies 101

3.2.3 Cell counting 102

3.2.4 Cell cycle analysis / BrdU incorporation 102

3.2.5 Annexin V- Propidium iodide labeling 102

3.2.6 Caspase activation 103

3.2.7 Mitochondrial membrane depolarization 103

3.2.8 Measurement of Intracellular ATP concentration 103

3.2.9 Western blotting 104

3.2.10 In vivo apoptosis 105

3.2.11 In vivo proliferation 106

3.3 Results 108

3.4 Discussion 128

CHAPTER 4 General discussion and conclusions 135

CHAPTER 5 Future directions 143

Supplement 150

REFERENCES 158

xi

ABBREVIATIONS

ANOVA analysis of variance

APAF apoptotic protease activating factor

AP-1 activator protein 1

ARD1 arrest defective 1

ARNT aryl hydrocarbon receptor nuclear translocator

ATM ataxia telangiectasia mutated

ATP adenosine triphosphate

ATR ATM and rad3 related

Bcl B-cell leukemia/lymphoma

bHLH basic helix-loop-helix

BNIP BCL2/adenovirus E1B 19kD interacting protein like

BrdU Bromodeoxyuridine

CBP CREB binding protein

CCN Cyclin

CDK Cyclin dependent kinase

CDKI Cyclin dependent kinase inhibitor

CO carbon monoxide

Dec1 deleted in esophageal cancer 1

DPG 2,3-disphosphoglycerate

eIF eukaryotic initiation factor

ets-1 erythroblastosis virus E26 oncogene homolog-1

ET-1 endothelin-1

ETC electron transfer chain

FIH factor-inhibiting HIF

H2O2 hydrogen peroxide

HIF hypoxic inducible factor

HO heme oxygenase

HPV hypoxic pulmonary vasoconstriction

HRE hypoxia responsive element

HUVEC human umbilical vein endothelial cell

HVR hypoxic ventilatory response

IAP inhibitor of apoptosis

IRES internal ribosomal entry site

JC-1 5,5’,6,6’-tetrachloro-1,1’,3,3’-tetraethylbenzimidazolcarbocyanine iodide

Kv voltage activated potassium current

MAPK mitogen-activated protein kinase

MMP-2 matrix metalloproteinase-2

mTOR mammalian target of rapamycin

NLS nuclear localization signal

NO nitric oxide

ODD oxygen dependent degradation domain

ORP150 oxygen-regulated protein 150

xii

PERK PKR-like endoplasmic reticulum kinase

PHD prolyl hydroxylase

PI propidium iodide

pO2 partial pressure of oxygen

PPAR peroxisome proliferator-activated receptor gamma

ROS reactive oxygen species

SRC sarcoma (Schmidt-Ruppin A-2) viral oncogene homolog

TAD transactivation domain

TIF transcriptional intermediary factor 1

TIMP tissue inhibitors of matrix metalloproteinase

TNF- tumor necrosis factor

UTR untranslated region

VEGF vascular endothelial growth factor

VHL von Hippel-Lindau

VSMC vascular smooth muscle cell

xiii

LIST OF FIGURES

CHAPTER 1

Figure 1.1 Regional distribution of pO2 from the airways to the cytosol 7

Figure 1.2 Oxyhaemoglobin dissociation curve 8

Figure 1.3 HIF-1, HIF-2 and HIF-3 subunit structure 13

Figure 1.4 Regulation of HIF activity 16

Figure 1.5 Cell cycle phases and G1/S transition 30

Figure 1.6 Pathways of apoptosis 37

Figure 1.7 Heme sensor model 43

Figure 1.8 Structure of NAD(P)H oxidase 45

Figure 1.9 NAD(P)H oxidase as oxygen sensor 46

Figure 1.10 Mitochondrial electron transport chain 47

CHAPTER 2

Figure 2.1 (A) Effects of hypoxia on HASMC cell numbers. Effects of HASMC cell

numbers to PDGF-BB at (B) 1% O2 and (C) and 3% O2. 67

Figure 2.2 [3H]-Thymidine incorporation in HASMCs after incubation at (A) 1% O2 and

(B) 3% O2 compared with the normoxic cells. 70

Figure 2.3 The percentage of cells positive for the Ki67 antigen after incubation at (A) 1%

O2 and (B) at 3% O

2. 71

Figure 2.4 CDC6 (A and B) and MCM2 (C and D) protein levels after normoxic and

hypoxic (1% O2 and 3% O2) incubation. 72

xiv

Figure 2.5 Flow cytometric analysis of propidium iodide stained cells at (A) 1% O2 and

(B) 3% O2. 74

Figure 2.6 Apoptosis assays in HASMCs. (A) Annexin V/PI (B) Caspase activity and (C)

TUNEL. 75

Figure 2.7 Mitochondrial membrane depolarization after incubation at 1% O2 (A) and 3%

O2 (B). 78

Figure 2.8 Cellular ATP concentration after incubation at 1% O2 (A) and 3% O2 (B).

79

Figure 2.9 Nuclear levels of (A) HIF-1 , (B) p21 and (C) p53 after incubation of

HASMCs at normoxic and hypoxic (1% or 3% O2) conditions. 82

Figure 2.10 (A) PI staining of en face sections and (B) TUNEL in paraffin embedded

sections of normoxic and hypoxic rat aorta and mesenteric artery. (C)

Quantitative analysis 84

Figure 2.11 (A) Immunohistochemical staining of incorporated BrdU (B) Double staining

with TO-PRO-3 and (C) -Smooth Muscle Actin in paraffin embedded sections

of normoxic and hypoxic rat aorta and mesenteric artery (D) Quantitative

analysis of incorporated BrdU and TO-PRO-3 staining. 87

CHAPTER 3

Figure 3.1 Effects of hypoxia on human pulmonary artery smooth muscle (HPASMC) (A)

cell numbers and (B) viability 109

Figure 3.2 % BrdU incorporated cells in HPASMCs 111

Figure 3.3 (A,B) Annexin V/PI (C,D) Caspase activity and (E) JC-1 monomer formation

after incubation at 3%, 1% and 0% O2 113

xv

Figure 3.4 Cell cycle analysis of propidium iodide stained normoxic and hypoxic (3%, 1%

or 0% O2) HPASMC cells. 117

Figure 3.5 Nuclear levels of (A) p21, (B) p53 and (C) HIF-1 in HPASMCs at normoxic

and hypoxic (1% or 3% O2) conditions. 119

Figure 3.6 Cellular ATP concentrations in normoxic and hypoxic (3%, 1% or 0% O2)

HPASMC cells. 122

Figure 3.7 (A) PI staining of en face sections of normoxic and hypoxic rat pulmonary

artery and pulmonary artery branch, (B) Quantitative analysis of PI stained

cells, (C) TUNEL in paraffin-embedded sections of normoxic and hypoxic rat

pulmonary artery and pulmonary artery branch (D) Quantitative analysis of

TUNEL positive cells. 124

Figure 3.8 (A) Immunohistochemical staining of incorporated BrdU and -smooth muscle

actin in paraffin-embedded sections of pulmonary artery and pulmonary artery

branch from normoxic and hypoxic rats. (B) Quantitative analysis of BrdU

positive cells. 126

Supplement

Figure S1 Average fold change of mitochondrial DNA levels 152

Figure S2 Cytoplasmic levels of Phosphoglycerate kinase and Enolase protein in

HPASMCs (A, C) and HASMCs (B, D), lactate concentration after incubation

of HPASMCs (E) and HASMCs (F) under normoxic and hypoxic (3, 1 or 0%

O2) conditions. 153

xvi

LIST OF TABLES

Table 1.1 Hypoxia-inducible genes harboring HRE sequences 19

Table 2.1 Normalized expression of pro- and antiproliferative genes and pro- and

antiapoptotic genes under hypoxia. 80

Table 3.1 Influence of hypoxia on pulmonary artery smooth muscle cell proliferation.

100

Table 3.2 Medial wall thickness of pulmonary artery and pulmonary artery branch from

normoxic and hypoxia exposed rats. 127

Table 5.1 Phenotypic heterogeneity in pulmonary artery smooth muscle cells.

144

Chapter 1

1

CHAPTER 1

Review of literature

Chapter 1

2

1.1 Introduction

The efficient delivery of oxygen to the tissues of the body is required for aerobic ATP

production to support their metabolic activities and as a substrate in the synthesis of a number of

signaling molecules such as carbon monoxide and nitric oxide [1-3]. Inadequate oxygen supply

will impair the capacity to meet these needs and result in the failure of vital functions. Hypoxia

refers to conditions under which oxygen concentration becomes limiting for normal cellular

processes [4]. The oxygen concentration in the atmosphere is 20.9% (partial pressure ~ 160

mmHg at sea level). The cells that comprise the vascular wall, however, experience much lower

oxygen tensions (25 mm Hg at the preterminal arterioles) [5, 6] with even lower levels

(bordering on anoxia) reported in vessels affected by disease [7-9] . At levels of 3-5% oxygen

vascular cells are close to the hypoxic range, although oxygen availability is not yet limiting to

cellular viability or function. Any further decrease in oxygen levels, however, will trigger

hypoxia-induced responses, which includes regulation of both cell proliferation and/or cell

survival to alter the structure of the vessels [10, 11]. These are aimed at both enhancing the

capacity to utilize the available oxygen supply and, in the event that hypoxia is prolonged to

protect cell viability and function. The molecular mechanisms underlying these responses are

complex and remain poorly understood. Their elucidation will aid in the development of

therapeutic approaches to ameliorate the effects of hypoxia in diseases associated with reduced

systemic oxygen delivery.

1.2 Systemic and pulmonary circulations

Structural and functional differences between the systemic and pulmonary circulations

support their respective physiological functions. The pulmonary circulation is a low pressure

Chapter 1

3

system with a mean pressure of 10-22 mm Hg, compared to 70-105mm Hg in the systemic

circulation. The pulmonary vasculature is thin walled compared to the systemic circulation, and

contains much less vascular smooth muscle. In the systemic circulation, 75–80% of vascular

resistance is maintained by small muscular arterioles whilst resistance is relatively evenly

distributed throughout the normal pulmonary circulation.

Pulmonary vascular reactivity to endogenous and exogenous vasoconstrictors and to

hypoxia is influenced by the level of basal tone. Baseline vascular tone is low in the normal lung

but is enhanced during hypoxia due to mechanisms intrinsic to the smooth muscle and because of

the effects of locally released and circulating vasoactive mediators such as Endothelin-1,

vasoconstrictor prostaglandins, histamine and serotonin [12, 13]. In the systemic circulation

basal tone is maintained by tonic activity of the sympathetic nervous system. Sympathetic

innervation of the pulmonary circulation does exist and its activation has similar effects as in the

systemic circulation but contributes little to the maintenance of basal vasomotor tone.

Pulmonary arteries exhibit a vasoconstrictor response to hypoxia in contrast to the

vasodilator response to hypoxia exhibited by the systemic circulation [14, 15]. In the foetus, this

hypoxic pulmonary vasoconstriction (HPV) serves to increase pulmonary vascular resistance and

divert the circulation through the ductus arteriosus. As a result the foetal pulmonary circulation

only receives ~10% of the cardiac output unlike the situation after birth where exposure to

atmospheric oxygen fully dilates the pulmonary circulation which henceforth receives 100% of

the cardiac output. After birth, HPV is required for ventilation-perfusion matching. Despite its

clinical and physiological relevance the mechanism of HPV remains largely unresolved. Recent

Chapter 1

4

hypothesis for HPV proposes hypoxia-induced inactivation of voltage activated potassium (Kv)

channels in the pulmonary circulation [16].

The larger arteries in the vasculature provide little resistance to blood flow and therefore

serve as a rapid conduit for blood to travel. The walls of these vessels contain large amounts of

elastic and fibrous tissue. As the arteries branch into smaller arteries, the amount of elastic tissue

in the walls decreases while the amount of smooth muscle increases. Arteries less than 0.1mm in

diameter lose most of their elastic properties and are sometimes called muscular arteries. The

combination of stiffness and flexibility enables arteries to act as pressure reservoirs to ensure a

continual smooth flow of blood through the vasculature even when the heart is not pumping

blood. The arterioles are the blood vessels that provide the greatest resistance to blood flow. In

the systemic circuit, blood enters arterioles at an average pressure of about 90 mmHg and leaves

them at a pressure of about 40 mmHg. The walls of arterioles contain little elastic material but

have an abundance of circular smooth muscle that forms rings around the arterioles. Resistance

is regulated by the contraction and relaxation of the circular smooth muscle.

The arteries have two functions. One is to deliver an adequate supply of blood to

peripheral tissues and to smooth out pressure oscillations due to intermittent ventricular ejection.

The efficiency of conduit function is related to the width of the arteries and the almost constancy

of mean blood pressure along the arterial tree. Resistance arteries with an internal diameter of

150 m contribute significantly to total peripheral resistance and basal vascular tone. Resistance

arteries are continuously subjected to changes in mechanical forces (flow and pressure) that

regulate active vasomotion, fitting blood flow continuously to local demands. The fundamental

function of resistance-sized arteries is control of blood flow to the capillary beds, partly achieved

Chapter 1

5

by a putative pressure-sensing mechanism. Vascular remodeling is an adaptive process

occurring in response to long-lasting changes in arterial pressure or flow, and whose ultimate

effect tends to maintain the constancy of tensile and/or shear stresses. In response to blood

pressure increase, the luminal diameter in large conduit arteries is usually unchanged while width

of wall increases. In distal resistive arteries and arterioles, luminal diameter is reduced but

medial layer is not hypertrophied.

1.3 Vascular smooth muscle cells

The vascular smooth muscle cell (VSMC) in mature animals is highly specialized whose

principal function is contraction and regulation of vessel tone and diameter, blood pressure,

and

distribution of blood flow. SMCs within adult blood vessels proliferate at a low rate and express

a unique repertoire of contractile proteins, ion channels, and signaling molecules required for the

cell's contractile function [17, 18].

Three independent embryonic origins for VSMCs have been identified: (a) Vessels that

recruit SMCs from progenitors that originate in cardiac neural crest; (b) Coronary SMCs arise

from mesothelial cells that line villus-like projections of the proepicardial organ and (c) Vessels

that recruit SMCs from either lateral or splanchnic mesoderm depending on the position of a

particular vessel within the embryo [19-23]. SMCs are also recruited from endothelial cells and

from circulating multipotential stem cells at later stages of development and in adults [24-27].

The majority of VSMCs exhibit common properties regardless of their origins, however, certain

lineage-specific differences in growth and transcriptional responses to various cytokines and

other factors implicated in the progression of arterial diseases persist beyond the embryonic

period.

Chapter 1

6

Unlike either skeletal or cardiac muscles, that are terminally differentiated, VSMCs

within adult animals retain remarkable plasticity. The

ability of VSMCs to be plastic in their

growth responses is a key mechanism by which the vasculature responds to hemodynamic,

developmental, and injurious stimuli. Biological processes during which VMSC growth is vital

include vessel development, the vascular response to tissue injury, and vessel remodeling in

response to changes in tissue demand [28-32]. Pathological examples include atherosclerosis,

hypertension, restenosis post angioplasty, and vasculitis. In these situations, interactions between

endothelial cells and VSMC, as well as between VSMC and other cells (e.g., fibroblasts,

dendritic cells, and inflammatory cells) within the vessel wall, determine

the nature of the growth

response [33]. The role of SMCs is not a simple function of alterations in its growth state but

rather is a function of very complex changes in the differentiated state of the SMC

including

increased matrix production [34], production of various proteases [35], participation in chronic

inflammatory responses including production of inflammatory cytokines and

expression of

inflammatory cell markers [36, 37], altered contractility and expression of contractile proteins

[38]. On one hand, the plasticity exhibited by VSMCs prevents accumulation of replication

errors or mutations. On the other, the high degree of plasticity

exhibited by the VSMCs

predisposes the cells to abnormal environmental cues/signals that can lead to adverse phenotypic

switching and the acquisition of characteristics that can contribute to development and/or

progression of vascular disease.

1.4 Oxygen Delivery

The primary function of the cardiovascular system is the delivery of oxygen that we

breathe from the air to the cells that comprise the body. The partial pressure of oxygen (pO2) of

Chapter 1

7

dry air at sea level is ~160 mmHg (21/100 x 760=159.6) [39]. However, by the time the inspired

air reaches the trachea it has been warmed and humidified by the upper respiratory tract and,

taking water vapor pressure (47 mmHg) into account, the pO2 in the trachea while breathing air

is ~150 mmHg (19.7%). By the time the inspired gas has reached the alveoli, the pO2 has fallen

to about 100 mmHg (because of diffusion of oxygen and CO2 from and into the alveolar gas,

respectively. The median pO2 in systemic arteries is ~92 mmHg (12%), however, it falls to ~50

mmHg (6.6%) in arterioles and ~25 mmHg (3.3%) in precapillary arterioles and capillaries as a

result of transarterial wall oxygen diffusion [6, 40, 41]. Oxygen gradients exist across the aortic

wall where pO2 ranges from ~85 mmHg (11.2%) at the lumen to ~17mm Hg (2.2%) at a depth of

150 m [6].

Oxygen diffuses from the alveolus to the pulmonary capillary until the pO2 in the

capillary is equal to that in the alveolus. This process is normally complete by the time the blood

has passed one third of the way along the pulmonary capillary. Oxygen dissociates from

Figure 1.1 Regional distribution of pO2 from the airways to the cytosol.

Source: Ward J (2007) Oxygen sensors in context [39].

Chapter 1

8

haemoglobin in red blood cells to the tissues according to the oxyhaemoglobin dissociation

curve.

Sequential branching of the arteriolar tree forms microvessels of decreasing diameter,

which, in turn, increases the surface area per unit volume available for the diffusion of oxygen to

the tissue [40]. In any oxygen-consuming tissue, the rate of intravascular oxygen loss is inversely

related to arteriolar vessel diameter, thereby creating an intravascular longitudinal oxygen

gradient. The affinity with which oxygen binds to hemoglobin is also influenced by pH, carbon

monoxide (CO), temperature and erythrocyte 2, 3-disphosphoglycerate (DPG) concentration [42,

43]. Lastly, increasing capillary perfusion increases the capacity for oxygen extraction during

exercise [44, 45].

Figure 1.2 Oxyhaemoglobin dissociation curve for normal adult haemoglobin.

Source: http://www.anaesthesiamcq.com/downloads/odc.pdf

Chapter 1

9

1.5 Physiological responses to hypoxia

The tissue oxygen supply is regulated by the number and function of the blood vessels,

whereas the demand is regulated by the number of cells in the tissue and their rate of

metabolism. All nucleated cells in the body respond to reduced O2 availability, through a series

of coordinated responses in a time and oxygen concentration-dependent manner. Stimulus-

response pathways induced by hypoxia can be categorized as either acute or chronic. Acute

responses are of rapid onset and short-term duration, whereas chronic responses are of delayed

onset and long-term duration. This difference in kinetics reflect the underlying molecular

mechanisms: acute responses involve post-translational modifications of existing proteins that

alter their activity whereas chronic responses are comprised of transcriptional and post-

transcriptional events involving changes in gene expression that result in the synthesis of novel

proteins or increased synthesis of proteins already present in the cell.

1.5.1 Systemic responses

During acute hypoxic exposure, oxygen supply to essential organs is maintained by the

following: (i) the hypoxic ventilatory response increases the respiratory rate and tidal volume

[46]. In humans, this is almost solely due to depolarization of glomus cells in the carotid body

which leads to enhanced ventilation and increased alveolar oxygen concentrations [47], (ii) the

pulmonary vasculature O2 sensors initiate hypoxic pulmonary vasoconstriction (HPV) to increase

efficiency of gas exchange. Pulmonary arterial vasoconstriction directs blood to better

oxygenated regions of the lung while changes in bronchial and bronchiolar tone optimize the

distribution of gas flow within the lung. An increase in pulmonary arterial blood pressure forces

blood into greater numbers of alveolar capillaries than normal [22], (iii) activation of the

Chapter 1

10

sympathetic system increases oxygen extraction by increasing the heart rate and diverting

unnecessary blood flow away from organs such as the kidneys and splanchnic viscera toward the

essential organs like the heart and brain [43], (iv) Vessels in essential organs accommodate the

increased blood flow through both a sympathetically-mediated increase in arteriolar tone and the

release of vasodilators in areas of imbalance between metabolic demand and oxygen supply.

The sympathetic excitation results partly through chemoreceptor reflexes and partly through

altered baroreceptor function, (v) The O2 sensors in the vasculature of other tissues activate

expression of VEGF-1 to promote angiogenesis, (vi) O2 sensors in the kidney and liver activate

the expression of erythropoietin to up-regulate red blood cell mass to improve oxygen carrying

capacity.

1.5.2 Regulation of cellular metabolism:

1.5.2.1 Effect of hypoxia on mitochondria

Mitochondria are the seat of oxidative phosphorylation and the main source of high

energy phosphate bond molecules in normal cells. Studies on isolated mitochondria have shown

that limited oxygen availability inhibits the electron transport chain and increases the proton

leak, although phosphorylation is less affected [1]. The inhibition of the respiratory chain occurs

at pO2 levels high above the Km of cytochrome c oxidase, indicating that a

specific inhibitory

mechanism, still unknown, is switched on well before oxygen concentration by itself would limit

the activity of this enzyme [48].

1.5.2.2 Cellular adaptation to hypoxia

Chapter 1

11

At the cellular level, adaptation to hypoxia is brought about on one hand by increased

anaerobic glycolysis activity, and on the other hand by decreasing energy-consuming processes

[49, 50]. Ion-motive ATPases and protein synthesis are the dominant energy-consuming

processes of cells at normal metabolic rates, making up more than 90% of the ATP consumption

in rat skeletal muscle and 66% in rat thymocytes [50]. As energy becomes limiting, protein

synthesis and RNA/DNA synthesis are the first to be inhibited while Na+/K

+ pumping and Ca

2+

cycling are potentiated. This phenomenon, known as oxygen conformance, involves precise

regulatory mechanisms mostly at the level of translation initiation [51].

The switch between aerobic and anaerobic pathways of ATP regeneration during hypoxia

was first noted by Pasteur in the late 19th century, hence its name "Pasteur effect." Although

glycolysis is less efficient than oxidative phosphorylation in the generation of

ATP, in the

presence of sufficient glucose, glycolysis can sustain ATP production due to increases in the

activity of the glycolytic enzymes.

1.6 Regulation of Gene Expression

Faced with a hypoxic challenge, the early physiological responses include increased

ventilation and cardiac output, a switch from aerobic to anaerobic metabolism, improved

vascularization, and enhancement of the O2 carrying capacity of the blood. In the longer term,

these responses are reinforced by up-regulation of genes encoding factors which facilitate these

responses, such as (i) tyrosine hydroxylase, which is involved in dopamine synthesis in carotid

body type I cells; (ii) glycolytic enzymes and glucose transporters Glut-1 and Glut-4; (iii) VEGF,

PDGF which promote angiogenesis, and inducible NO synthase which increases

vasodilation;

Chapter 1

12

and (iv) erythropoietin and transferrin receptors that favor erythrocyte production [52]. This

transcriptional response is mediated in large part by the action of HIF-1 .

1.6.1 Hypoxia-inducible factors

Hypoxia increases nuclear translocation of a family of hypoxia inducible transcription

factors (HIFs) activates expression of genes participating in the compensatory mechanisms that

support cell survival in a potentially lethal microenvironment [53].

HIF transcription factors are composed of one of three alpha subunits (1 , 2 or 3 ), and

beta ( ) subunits. HIF-1 is also denoted as the aryl hydrocarbon receptor nuclear translocator

(ARNT) [54]. In the subunit, the basic helix-loop-helix (bHLH) and the Per Arnt Sim (PAS)

domains in the N-terminus are important for dimerization and DNA binding [55-57]. HIF-1

and HIF-2 proteins also contain two transactivation domains (TADs) in their C-terminal region.

Within the N-terminal TAD there is an oxygen-dependent degradation (ODD) domain that is

responsible for degradation of the subunit under normoxic conditions [58, 59]. The main

function of the C-terminal TAD is to recruit transcriptional coactivators such as CBP, p300,

SRC1 and TIF-2 [56, 60, 61].

Under hypoxic conditions, the HIF heterodimer (named HIF-1, -2 or -3) translocates to

the nucleus where it binds to a core DNA sequence (5-ACGTG-3) - the hypoxia-responsive

element (HRE), located in the promoter/enhancer regions of many hypoxia-regulated genes [53,

62].

Chapter 1

13

Of the three HIF- -subunits, HIF-1 is the best studied and characterized to date.

Nevertheless, the understanding of HIF-2 (also known as endothelial PAS domain protein 1,

EPAS-1) function has increased dramatically, whereas the most recently identified and

consequently less-well studied subunit is HIF-3 [63, 64]. Although the HIF-1 and -2 subunits

can bind to the same DNA motifs, they appear to control rather distinct biological functions.

Mouse knockout studies have shown the vital importance of HIF-1 for development and

survival, and that HIF-2-/-

mice have different phenotypes depending on their genetic

Chapter 1

14

background, thus illustrating the importance of HIF-2 [65, 66]. In addition, a recent study has

demonstrated that HIF-2 cannot functionally substitute for HIF-1 in embryonic stem cells

[67]. Whereas HIF-1 is expressed in virtually all cell types, HIF-2 exhibit a more restricted

expression pattern in endothelial cells and in catecholamine producing cells in the organs of

Zuckerkandl [68]. The biological function of HIF-3 is under investigation, but a HIF-3 splice

variant, denoted inhibitory PAS (IPAS), appears to function as a negative regulator of hypoxia-

inducible responses [69, 70].

HIF- is not controlled by oxygen levels and is found constitutively expressed in all cell

types [71]. By contrast, -subunit levels are under tight control. In response to changing oxygen

levels, the control of HIF- subunit expression is achieved by regulating protein level, although

other stimuli such as oncogene activation and cytokines can induce both transcription and protein

synthesis increases of the subunits [72].

1.6.2 Regulation of HIF activity

Nuclear levels of HIF- proteins, which are extremely low under normoxic conditions,

dramatically increase in response to hypoxia. The presence of HIF- subunits only in conditions

of low oxygen tension or after treatment with iron chelators had puzzled researchers until a

group of novel oxygen-dependent enzymes essential for regulating HIF- protein levels was

discovered. These are the prolyl hydroxylases (PHDs), of which four isoforms (PHD1, PHD2,

PHD3 and PHD4) have been identified at this time [73]. The PHD enzymes are members of the

2-oxoglutarate-dependent hydroxylase superfamily.

Hydroxylation of two proline residues (402 and/or 564) in the HIF- ODD domain by

PHDs serves as a recognition/binding site for the von Hippel-Lindau (pVHL) E3 ubiquitin ligase

Chapter 1

15

complex [73, 74]. Binding of pVHL targets HIF- for polyubiquitylation and subsequent

degradation by the 26S proteasome [75]. To obtain full transcriptional activity, HIFs must bind

to the HRE DNA sequence and recruit transcriptional co-factors. In the presence of oxygen,

asparagine 803 in the C terminal TAD gets hydroxylated by a HIF asparaginyl hydroxylase

called factor-inhibiting HIF (FIH-1) [76, 77]. This will silence the TAD domain by preventing

the binding of transcriptional co-activators CBP/p300. The enzymatic modifications effected by

the prolyl and the asparaginyl hydroxylases are dependent on oxygen and iron (Fe2+), explaining

the fact that HIF- subunits escape degradation in the absence of oxygen or iron [78-81].

Additional pathways for silencing HIF activity under normoxic conditions include

acetylation of lysine 532 in HIF-1 ODD domain by the arrest defective 1 (ARD1) acetylase

[82]. This modification increases the interaction with pVHL, resulting in enhanced degradation

of HIF-1 . The stability and activity of HIF- subunits may also be influenced by reactive

oxygen species, hydrogen peroxide as well as by growth factor- and cytokine-induced

phosphorylation [56, 83-86].

Chapter 1

16

1.6.3 HIFs as transcriptional regulators

Chapter 1

17

When the protein levels of HIF- increase, e.g. in response to hypoxia, it translocates to

the nucleus, dimerizes with the subunit and activates the transcription of a number of target

genes displaying an HRE motif (Table 1.1). Nuclear localization signal (NLS) domains in the

and subunits confer autonomous translocation into the nucleus [87]. One group of HIF-1 target

genes is involved in the adaptive response facilitating oxygen delivery to oxygen-deprived

tissues. These include the genes encoding erythropoietin, vascular endothelial growth factor-A

(VEGF-A) and the inducible NOS (iNOS) [62]. The erythropoietin (Epo) gene, was discovered

as the first true hypoxia-inducible gene in 1992 [53]. EPO stimulates red blood cell production

(erythropoiesis), thereby increasing oxygen delivery. Hypoxia also promotes iron uptake and

transport by increasing the expression of transferrin and the transferrin receptor [88, 89]. Another

well-known hypoxia-regulated gene is Vegf-a, which plays a crucial role in development and

growth of blood vessels [90, 91]. One of the VEGF receptors, encoded by the Vegfr-1 gene, is

also a direct HIF target, harboring an HRE motif [92]. The Vegfr-2 gene, which at first was

reported to lack HIF binding sites, has now been shown to be upregulated by HIF-2 [93].

Hypoxia also affects vascular tone and local blood flow by induction of vasoconstrictors, such as

endothelin-1 (ET-1) [94], or by increased expression of genes regulating vasodilation, such as

heme oxygenase-1 (HO-1) and iNOS [95, 96].

Another group of genes upregulated by HIF-1 acts to compensate for the loss of oxygen-

dependent metabolism in hypoxia. The increased expression of various glucose transporters and

glycolytic enzymes under hypoxic conditions, allows for oxygen-independent generation of ATP

(glycolysis). When oxygen levels fall to a critical point, metabolic switches turn off oxidative

phosphorylation and mitochondrial electron transport and instead oxygen-independent or

anaerobic energy production (glycolysis) is induced. In the glycolytic pathway, four ATP

Chapter 1

18

molecules are produced when glucose is metabolized to two molecules of pyruvate. As two ATP

molecules are consumed during this process, this leaves a net yield of two ATP. Compared to

aerobic conditions where pyruvate is further oxidized in the Kreb’s cycle and the net yield is 31

molecules of ATP, anaerobic glycolysis is much less efficient.

In addition to the “classic” hypoxia-inducible genes, that are direct transcriptional targets

of HIF, the response to low oxygen triggers expression of select micro RNAs (miRNAs), which

in turn down regulate specific genes. MicroRNAs are short non-coding transcripts. A wide set of

hypoxia-regulated miRs (HRMs) have been identified. Among them HIF plays an important

regulatory role for miR-210, 26 and 181. Studies have revealed a highly complex spectrum of

candidate targets of HRMs. These include key genes of the apoptotic pathway such as BID (miR-

23), BIM (miR-24); CASP3 (miR-30), CASP 7 (miR-23), APAF1 (miR-27), BAK1 (miR-26),

Bnip3L (miR-23). Conversely, antiapoptotic Bcl2 is a target of miR15 and 16. Another process

known to be affected by hypoxia is proliferation, since many cell types undergo cell cycle

slowdown or arrest during oxygen deprivation. A multitude of cell cycle genes are HRM targets,

a few examples being cdc25A (miR-21, miR-103/107), cyclin D2 (miR-26, miR-103/107),

cyclin E1 (miR-26), cyclin H (miR-23), cdk6 (miR-26, miR-103/107) [97-99]. HRMs miR-16,

miR-20, let-7b, miR-17-5p, miR-27, miR-106, miR-107, miR-193, miR-210, miR-320 and miR-

361 have been shown to target VEGF [100].

Table 1.1 Hypoxia-inducible genes harboring HRE sequences [62].

Chapter 1

19

Gene Function

Oxygen supply

1B-adrenergic receptor Vessel diameter

Adrenomedullin Vessel diameter

Atrial natriuretic peptide (ANP) Blood volume

Breast cancer resistance protein (BCRP) Heme binding

Endothelial nitric oxide synthase (eNOS) Vessel diameter

Endothelin-1 Vessel diameter

Erythropoietin Erythropoiesis

Ferrochelatase Heme synthesis

Heme oxygenase 1 Vessel diameter

Inducible nitric oxide synthase (iNOS) Vessel diameter

Leptin Metabolism/ Angiogenesis

Transferrin Iron transport

Transferrin receptor Iron transport

Plasminogen activator inhibitor-1 (PAI-1) Blood flow

Vascular endothelial growth factor-A (VEGF-A) Angiogenesis

VEGF-D Angiogenesis

VEGF receptor-1 (VEGFR-1) Angiogenesis

VEGFR-2 Angiogenesis

Cellular metabolism

Aldolase A Glycolysis

Carbonic anhydrase-9 (CA-9) pH regulation

Cytochrome P450 2C11 (CYP2C11) Metabolism

CYP3A6 Metabolism

CYP4B Eicosanoid synthesis

Enolase 1 Glycolysis

Glucose transporter 1 (Glut1) Glucose uptake

Glucokinase Glycolysis

Glutathione peroxidase-3 (GPx-3) Glutathione peroxidase

Glyceraldehyde-3-phosphate dehydrogenase (GAPDH) Glycolysis

Lactate dehydrogenase A Glycolysis

Multidrug resistance gene 1 (MDR1) Xenobiotic transporter

Phosphoenolpyruvate carboxykinase (PEPCK) Gluconeogenesis

Phosphofructokinase L (PFKL) Glycolysis

6-Phosphofructo-2-kinase/fructose-2,6-biphosphatase-3 Glycolysis

6-Phosphofructo-2-kinase/fructose-2,6-biphosphatase-4 Glycolysis

Phosphoglycerate kinase 1 (PGK1) Glycolysis

Table 1.1 Hypoxia-inducible genes harboring HRE sequences (contd)

Chapter 1

20

Gene Function

Cell growth and metabolism

Connective tissue growth factor (CTGF) Growth factor

Ecto-5’-nucleotidase (CD73) Intestinal barrier function

Endoglin TGF- coreceptor

Insulin growth factor binding protein-1 (IGFBP-1) Growth factor

Intestinal trefoil factor Intestinal barrier function Intestinal barrier function

Transforming growth factor- 3 (TGF- 3) Placenta development

Cell growth and apoptosis

CXCR4 Chemokine receptor

Bcl-2/E1B 19kDa interacting protein (BNip3) Pro-apoptotic

Met Proto-oncogene

Myeloid cell factor-1 (Mcl-1) Anti-apoptotic

Nip3 Pro-apoptotic

Noxa Pro-apoptotic

Nucleophosmin p53 inhibition

Nur77 Orphan steroid receptor

Serine/threonine protein phosphatase 5 (PP5) Anti-apoptotic

Stromal cell-derived factor-1 (SDF-1 or CXCL12) Chemokine

Telomerase reverse transcriptase (TERT) Telomere extension

Wilms’ tumor suppressor (Wt1) Tumor suppressor gene

Others

CD18 Leukocyte adhesion

Cited2/p35srj Transcription cofactor

Collagen prolyl 4-hydroxylase I Hydroxylase

DEC1 and DEC2 Transcription factors

Ets-1 Transcription factors

Furin Pro-protein convertase

Glucose-regulated protein 94 (GRP94) Chaperone

Inhibitor of differentiation/DNA binding protein 2 (ID2) Transcriptional repressor

Membrane type-1 matrix metalloproteinase (MT-1) Matrix metalloproteinase

Prolyl hydroxylase domain protein 2 and 3 (PHD2/PHD3) Oxygen sensing

Retrotransposon VL30 Retrotransposon

Chapter 1

21

Besides prolyl hydoxylases, stabilization and/or synthesis of HIF-1 under hypoxia is also

dependent on the activity of the PI-3 kinase/Akt pathway [84]. PI-3K inhibitors inhibits the

accumulation of HIF-1 in these conditions, while dominant negative

mutants for PI-3K or for

Akt decrease the hypoxia-induced overexpression of VEGF. Conversely, disruption of PTEN, a

phosphatidylinositol triphosphate phosphatase that inactivates Akt, leads to increased

VEGF

expression in normoxic cells [101]. Finally, growth factor- or cytokine-induced activation of

HIF-1 in normoxia results from an increased synthesis of HIF-1 which is also dependent on the

PI3K/Akt pathway. It remains unclear, however, how the PI-3K/Akt pathway interacts with the

prolyl hydroxylase-pVHL system to regulate HIF-1 protein level.

Post stabilization, redox status, dissociation from the chaperone hsp90, association with

co-activators like CBP/p300 or SRC-1 as well as phosphorylation are also required for full

transcriptional activity (36, 37). Hypoxia directly regulates the association of HIF-1 with the

coactivator CBP/p300. Similar to prolyl hydroxylase, an asparagyl hydroxylase, whose activity

strictly depends on

the presence of oxygen, hydroxylates HIF-1 carboxyl-terminal

transactivation domain on Asn 803. This modification prevents its association with CBP/p300

under normoxic conditions [76, 77].

1.6.4 HIF independent transcriptional activation

Although HIF-1 is a pivotal regulator of transcription in hypoxia, other transcription

factors induced in response to hypoxia include the early growth response protein (Egr-1),

Nuclear factor- B (NF- B) and activator-protein 1 (AP-1). The early growth response protein

(Egr-1) is a zinc finger nuclear phosphoprotein that is induced and activated within minutes of

oxygen deprivation [50]. Induction of Egr-1 DNA binding activity leads to activation of tissue

Chapter 1

22

factor gene transcription. In Egr-1 null mice, expression of tissue factor and intravascular fibrin

deposition were severely decreased after hypoxia. This general effect of Egr-1 activation extends

to the tissue factor (TF), VEGF, plasminogen-activator inhibitor (PAI), intracellular adhesion

molecule (ICAM), as well as several interleukins. The Egr-1 and HIF-1 pathways appear to be

initiated independently of each other, indicative of the separate role they each play in inducing

different facets of the adaptive response to hypoxia [102].

The p50-p60 heterodimer of Nuclear factor- B (NF- B) is induced both under hypoxia

and following re-oxygenation. To date the role of NF- B in mediating induction of hypoxia-

responsive genes is poorly understood. Studies by Koong et al and Imbert et al have suggested

hypoxia-induced activation of NF- B to occur via a mechanism involving tyrosine

phosphorylation of the upstream inhibitory subunit I B . One of the genes thought to be

regulated by NF- B in hypoxia is cyclooxygenase-2 (COX-2), which is induced in human

vascular endothelial cells by the binding of p65 to the NF- B consensus element in the COX-2

promoter [103-106].

The activator-protein 1 (AP-1) is a dimeric transcription factor comprising subunits from

the jun and fos multigene families. The DNA binding and transcriptional activity of AP-1 has

been demonstrated to be strongly induced by hypoxia [107]. A single cysteine residue in the

DNA-binding domain of fos and jun is responsible for this redox sensing and signaling.

Activation of VEGF [108], tyrosine hydroxylase [109], collagenase IV [110], endothelin-1 [111]

and c-jun [112] has been shown to correlate with the activation of AP-1. The hypoxic regulation

of AP-1 may be functionally distinct from other AP-1-inducing stresses, as it was shown that

ectopically expressed c-jun functionally cooperates with HIF-1 to regulate HRE-dependent

Chapter 1

23

reporter expression without binding to AP-1. Furthermore, induction of c-jun mRNA expression

and phosphorylation by prolonged hypoxia was dependent on HIF-1 [113].

1.7 Hypoxic regulation of mRNA stability

Both transcriptional activation and post-transcriptional mechanisms contribute to the

hypoxia-mediated regulation of gene expression. Hypoxic regulation of cis-acting regulatory

elements found at the 3-UTR of the hypoxia-responsive mRNAs are responsible for hypoxia-

specific message stabilization [114] and the most commonly described cis-acting sequences

include the AU-rich element (ARE), stem-loop element and pyrimidine-rich element. Hypoxia

has been known to specifically increase stability of the mRNAs of vascular endothelial growth

factor (VEGF), tyrosine hydroxylase (TH), glucose transporter (GLUT-1) and erythropoietin

(EPO) [115]. Hypoxic induction of VEGF occurs in a biphasic manner: initial activation via

transcriptional induction is followed by augmented mRNA stability mediated by binding of

specific proteins such as the heterogeneous nucleoprotein L (hNRP L) and the RNA binding

protein, HuR [116-118]. Likewise, as shown in pheochromocytoma-derived PC12 cells, the

hypoxia-dependent stabilization of the TH gene mRNA is due to a hypoxia-inducible protein

binding site (HIPBS) in the 3-UTR of the TH mRNA [119].

1.8 Hypoxic repression of transcription

In contrast to inducing the expression of specific genes, hypoxia can also result in

specific gene repression. Several proteins have been reported to contribute to transcriptional

repression in hypoxic cells. These include negative cofactor 2 (NC2), differentiated embryo

chondrocyte 1 (Dec 1), histone deacetylases (HDAC), mSin3a, and p53.

Chapter 1

24

NC2, a transcriptional repressor, activated in extracts from hypoxia-treated hepatoma

cells inhibits the formation of the pre-initiation complex through direct interaction with the

TATA binding protein (TBP). This interaction prevents access to the promoter by TFIIB, thus

preventing the formation of the RNA polymerase holoenzyme, and thereby blocks transcription

[120, 121]. Because of the large number of TATA-containing genes in the human genome, the

activation of NC2 may contribute to a global repression of transcription during hypoxia.

A more specific hypoxia-dependent transcriptional repressor is Dec1 (Stra13, Sharp2).

Dec1 is a member of the basic helix loop helix family of transcription factors and has been

demonstrated to repress expression through binding to E-box elements [122]. Hypoxia can

prevent differentiation or cause dedifferentiation in a number of cell types including adipocytes,

breast carcinomas and neuroblastomas [123, 124]. Hypoxic induction of Dec1 has been shown to

be able to block the expression of PPAR 2 in pre-adipocytes, blocking their differentiation

[125].

In addition to DNA binding by direct acting repressors, gene expression can be down

regulated by corepressors such as histone deacetylases (HDACs) that act to modify the local

chromatin. HDACs function as co-repressors through their association with transcription factors,

such as p53, which can recruit and target them to specific genes. Hypoxia has been shown to

elevate HDAC activity [126] in addition to increased interactions with factors like p53. p53

protein has been shown to be stabilized under severe hypoxia, but the protein does not activate

transcription of its typical target genes [127]. However, in addition to its transactivation property,

p53 can also repress target genes, and this activity is retained under hypoxia [128]. To repress

gene transcription, p53 selectively interacts with its known transcriptional co-repressors mSin3a

and HDAC1 in hypoxic cells [129]. In addition, studies show that hypoxic induction of this

Chapter 1

25

complex containing p53, HDAC1 and mSin3a represses genes such as stathmin and Map4,

proteins which play a role in microtubule organization and ultimately in G2/M phase growth

arrest [128-130].

1.9 Hypoxic control of protein translation

Regulation of gene expression by hypoxia may occur at a post-transcriptional level.

Phosphorylation of eIF2 by the endoplasmic reticulum associated kinase PERK, during severe

hypoxia, results in a global reduction in protein synthesis. The activation of the PERK kinase is a

recognized response to ER stress, and the blocking of new protein synthesis is a means of

reducing that stress. Koumenis et al also shows that cells deficient in this response have also

been shown to be more sensitive to hypoxia-induced toxicity [131, 132].

Mammalian cells respond to wide ranges of oxygen concentration through alterations in

both metabolic states and growth rates. Hypoxia alters cellular proliferation by regulating the cell

cycle as well as by programmed cell death or apoptosis. The following two sections (1.10 and

1.11) give an overview of the cell cycle and apoptosis and outlines the effects of hypoxia on each

of these processes.

1.10 Cell cycle and hypoxia

1.10.1 Overview of mammalian cell cycle

The mammalian cell cycle consists of four distinct phases: G1, S (synthesis), G2

(collectively known as interphase) and M (mitosis). In the interphase, the cell grows,

accumulating nutrients needed for mitosis and duplicating its DNA and in the M phase, the cell

Chapter 1

26

splits itself into two daughter cells. M phase is itself composed of two tightly coupled processes:

mitosis, in which the cell's chromosomes are divided between the two daughter cells, and

cytokinesis, in which the cell's cytoplasm divides forming distinct cells. Activation of each phase

is dependent on the proper progression and completion of the previous one. Cells that have

temporarily or reversibly stopped dividing are said to have entered a state of quiescence called

G0 phase.

The first phase within interphase, from the end of the previous M phase until the

beginning of DNA synthesis is called G1 (G indicating gap). This phase is marked by synthesis

of various enzymes that are required in S phase, mainly those needed for DNA replication. In the

S phase the chromosomes are replicated with each chromosome having two (sister) chromatids.

Rates of RNA transcription and protein synthesis are low during this phase, except for histone

production, most of which occurs during the S phase [133]. The cells then enter the G2 phase,

which lasts until the cell enters mitosis. Significant protein synthesis occurs during this phase,

mainly involving the production of microtubules, required during mitosis. Inhibition of protein

synthesis during G2 phase prevents the cell from undergoing mitosis. The M phase has been

broken down into several distinct phases, sequentially known as prophase, prometaphase,

metaphase, anaphase and telophase leading to cytokinesis.

Control mechanisms ensuring the fidelity of cell division are called the checkpoints.

These verify whether the processes at each phase of the cell cycle have been accurately

completed before progression into the next phase. DNA damage checkpoints sense DNA damage

both before the cell enters S phase (a G1 checkpoint) as well as after S phase (a G2 checkpoint).

Damage to DNA before the cell enters S phase inhibits the action of CDK2 thus stopping the

progression of the cell cycle until the damage can be repaired. In case of irreparable damage, the

Chapter 1

27

cell self-destructs by apoptosis. Damage to DNA after S phase (the G2 checkpoint), inhibits the

action of Cdk1 thus preventing the cell from proceeding from G2 to mitosis.

The first checkpoint is located before entry into S phase, making the key decision of

whether the cell should divide, delay division, or enter a resting stage. The G1 checkpoint

(restriction point) is where eukaryotes typically arrest the cell cycle if environmental conditions

make cell division impossible [134]. The restriction point is mainly controlled by action of the

CKI- p16 (CDK inhibitor p16). This protein inhibits CDK4/6 and ensures that it can no longer

interact with cyclin D1 to cause cell cycle progression. The second checkpoint is located at the

end of G2 phase, triggering the start of the M phase (mitosis). The CDKs associated with this

checkpoint are phosphorylated by the "Maturation promoting factor" (or Mitosis Promoting

Factor, MPF). The MPF activates the CDK in response to environmental conditions being right

for the cell and allows the cell to begin DNA replication. An activating phosphatase, Cdc25,

under favourable conditions removes the inhibitory phosphates present within the MPF complex.

However, DNA is frequently damaged prior to mitosis, and to prevent transmission of this

damage to daughter cells, the cell cycle is arrested via inactivation of the Cdc25 phosphatase (via

phosphorylation with other protein kinases). There are also spindle checkpoints that detect any

failure of spindle fibers to attach to kinetochores and arrest the cell in metaphase until all the

kinetochores are attached correctly (M checkpoint).

In differentiated mammalian cells, G1 to S progression is regulated by the

hypophosphorylated Rb gene or its related proteins, p107 and p130, which inhibit the expression

of genes required for entry into S phase by sequestering the E2F family of transcription factors.

During G1 phase the Rb/HDAC repressor complex binds to the E2F-DP1 transcription factors

inhibiting downstream transcription. Eukaryotic cell cycle progression is dependent, in part, on

Chapter 1

28

the tightly regulated activity of CDKs. CDK4/CDK6 and Cdk2 whose regulatory partners are the

D-type cyclins (D1, D2 and D3) and cyclin E, respectively, represent two different classes of G1-

specific CDKs whose activation is required for entry into S phase. Cyclin D/CDK4–CDK6

activity occurs in mid-late G1 phase, upstream of CDK2/cyclin E activity. The mitogenic activity

of CDKs are inhibited by cell cycle inhibitory proteins, including p15 (INK4B), p16 (INK4A),

p18 (INK4C), p19 (INK4D), p21 (CIP), p27 (Kip1) and p57 (Kip2).

Studies have suggested that cyclin D/CDK complexes also play a second non-catalytic

role in G1 progression by sequestering proteins of the Cip/Kip family, including p27 (Kip1) and

p21(Cip1), two potent inhibitors of CDK2 [135]. Binding of Cip/Kip proteins to cyclin

D1/CDK4 stabilizes the complex and facilitates its nuclear import [136]. Mitogen withdrawal

results in the disassembly of the cyclin D/CDKs and in addition mobilizes the latent pool of

p27Kip1, which blocks the activity of cyclin E/CDK2 and facilitates cell cycle exit. Murine

embryonic fibroblasts (MEFs) lacking p27 and p21 do not express D-type cyclins and have a

significant reduction in CDK activity, but continue to proliferate normally, suggesting that D-

type cyclins might not be essential for cell cycle progression, at least in a setting where Cip/Kip

proteins are absent [136]. Further studies have shown that activation of the cyclin D1/CDK4

complex occurs when quiescent p21/p27-null MEFs are stimulated to re-enter the cell cycle. In

addition, the ectopic expression of p34 SEI-1, a mitogen-induced CDK4 activator, increased the

levels of active cyclin D1/CDK4 complex in the absence of p21 and p27, suggesting that there

are several independent pathways to stimulate the assembly of the cyclin D1/CDK4 complex

[137]. More recent studies have highlighted the role of an additional cell cycle regulatory

mechanism at the G1 to S transition that is able to govern the initiation of histone gene

expression needed for packaging of newly replicated DNA [138]. This is commonly referred as

Chapter 1

29

the S point and is initiated by cyclin E/Cdk2-dependent phosphorylation of p220 NPAT and the

formation of a functional HiNF-p220 NPAT complex that controls H4 gene transcription.

The expression of cell cycle inhibitory protein, p21, is tightly controlled by the tumor

suppressor protein p53, through which this protein mediates the p53-dependent cell cycle G1

phase arrest in response to a variety of stress stimuli. p21 can interact with proliferating cell

nuclear antigen, a DNA polymerase accessory factor, and plays a regulatory role in S phase

DNA replication and DNA damage repair [139]. p21 has been reported to be specifically cleaved

by Casp-3 like caspases, leading to apoptosis [140].

p53 is a transcription factor which in humans is encoded by the TP53 gene. The three

main functions of p53 include (a) activation of DNA repair proteins when DNA has sustained

damage; (b) induction of growth arrest at G1/S by activation of p21 expression, to allow DNA

repair proteins time to fix the damage; and (c) intiation of apoptosis if the DNA damage proves

to be irreparable. p53 becomes activated in response to a variety of stress signals including

hypoxia. The half-life of p53 is increased causing p53 accumulation in stressed cells. Also a

conformational change causes p53 to function as a transcriptional regulator in these cells.

Phosphorylation of the p53 N-terminal domain makes it a traget of two groups of protein kinases,

namely the MAPK family (JNK1-3, ERK1-2, p38 MAPK), and the ATR, ATM, CHK1 and 2

kinases which are implicated in the genome integrity checkpoint. Oncogenes also stimulate p53

activation, mediated by p14ARF. In unstressed cells, p53 levels are kept low through its

continuous degradation. Mdm2 binds to p53, preventing its action and transports it from the

nucleus to the cytosol. Also Mdm2 acts as ubiquitin ligase and covalently attaches ubiquitin to

p53 leading to its proteasomal degradation. This is reversible and a ubiquitin specific protease,

USP7, can cleave ubiquitin off p53, preventing its proteasome-dependent degradation.

Chapter 1

30

Phosphorylation of the N-terminal end of p53 disrupts Mdm2-binding. Other proteins, such as

Pin1, are then recruited to p53 and induce a conformational change in p53 which prevents

Mdm2-binding even more. Phosphorylation also allows for binding of transcriptional

coactivators, like p300, which then acetylate the carboxy terminal end of p53, exposing the DNA

binding domain of p53, allowing it to activate or repress specific genes. Deacetylase enzymes,

such as Sirt1 and Sirt7, can deacetylate p53, leading to inhibition of apoptosis [141].

1.10.2 Effects of hypoxia on cell cycle

When cells are exposed to severe hypoxia cell cycle progression and DNA synthesis

rapidly cease. The induction of HIF-1 activation prevents G1/S transition through the action of

CKIs and the regulation of cyclin E expression [142, 143]. Expression of p21 and p27 is

increased transcriptionally in a HIF-1-dependent manner [142, 144]. Sustained expression of

Figure 1.5 Cell cycle phases and G1/S transition.

Source: Herrup K (2007) Cell cycle regulation.

Chapter 1

31

these CKIs is observed in wild-type cells, but not in HIF-1 null cells. These CKIs suppress

cyclin/CDK2 activity, and thus reduce the ratio of phosphorylated to dephosphorylated Rb

protein, resulting in cell cycle arrest at the G1/S interface [145, 146]. HIF-1 may also regulate

cyclin E protein levels; CCNE binds to CDK2 and modulates its kinase activity dependent upon

cell cycle phase [146]. It has been reported that hypoxic cells lacking HIF-1 displayed enhanced

and sustained accumulation of cyclin E, without any effect on CDK2 protein expression, relative

to wild-type cells. In accordance with changes in cyclin E expression, cyclin E/CDK2 kinase

activity in HIF-1 -deficient cells was also increased, resulting in somewhat retarded, but still

substantial, cell growth, even under hypoxia.

Hypoxia causes an increase in the CDKN1A mRNA in a p53-independent manner [127].

The number of cells in G1 phase in p53 null cultures is increased relative to wild-type cultures.

This change may be attributable to enhanced HIF-1 activity by inactivation of p53, rather than

the direct action of p53, because expression and transcriptional activity of p53 change little under

hypoxia. HIF-1 null, p53 wild-type cells do not show any hypoxia-induced G1 arrest. Rather,

S-phase entry is accelerated, indicating that HIF-1, but not p53, plays an essential role in the

regulation of cell cycle progression under hypoxia [147, 148]. Cells lacking functional copies of

both p53 and HIF-1 have been shown to display no change in the proportion of cells entering S-

phase, as was seen in HIF-1 null cells. These cells appear to lose the ability to sense and

respond to hypoxia. Collectively, these data strongly suggest that both transcription factors, HIF-

1 and p53, cooperate to regulate the cell cycle progression through distinct mechanisms, but HIF-

1 serves as the primary determinant for cell cycle regulation under hypoxia. Previous studies

have indicated that hypoxia-induced cell cycle arrest is accompanied by a decreased activity of

CDKs and Rb protein, leading to inhibition of cell cycle progression. Also cyclin G2, a negative

Chapter 1

32

regulator of cell cycle progression via binding with protein phosphatase 2A in certain cell types,

is induced by hypoxia through HIF-1 activation [149, 150]. Hypoxia-induced S phase-dependent

arrest is mediated by a rapid shutdown of DNA synthesis through a block to replicon initiation

[151]. This block persists as long as the cells are held hypoxic, and is signaled through the ATR

kinase [152, 153].

1.11 Apoptosis and hypoxia

1.11.1 Apoptotic pathways

Cells can activate an intracellular death program and “commit suicide” in a controlled

way, a process known as apoptosis. Programmed cell death (apoptosis) was first described in

1972 by Currie and colleagues [154]. Apoptotic cell death is important for the maintenance of

tissue homeostasis under physiological conditions as well as for pathogenesis during disease

states including myocardial infarct, neurodegenerative disorders, autoimmune diseases, and

cancer [155, 156]. Alternately, cells can die by an uncontrolled process known as necrosis.

Apoptosis can be induced by a variety of factors, including ligand activation of death receptors,

growth factor deprivation and hypoxia. Characteristics of apoptosis include chromatin

condensation, membrane blebbing, phosphatidylserine exposure, cytoplasmic shrinkage,

formation of apoptotic bodies, and DNA fragmentation. The apoptosis pathway is dependent

upon caspase activation. Caspases comprise an expanding family of cysteine proteases that exist

as inactive pro-enzymes in viable cells [157]. Activated caspases acquire the ability to cleave key

intracellular substrates as well as activate other caspases, resulting in the induction of a protease

cascade that can kill the cell. Caspase activation is an ATP dependent process and is sufficient to

induce all of the morphological features of apoptosis. In contrast, necrosis does not involve the

Chapter 1

33

activation of caspases and is not an energy dependent process [158]. Characteristics of necrosis

include organelle swelling and cell bursting, leading to an inflammatory response. This

inflammatory response does not occur under apoptotic conditions since apoptotic cells display

phagocytosis markers and are engulfed by neighboring cells [159].

There are two possible mechanisms of apoptosis - intrinsic and extrinsic [160]. The

critical regulators of the intrinsic pathway are the Bcl-2 family members [161]. The family can

be divided into three different groups based on Bcl-2 homology (BH) domains and function. The

anti-apoptotic members, such as Bcl-2 and Bcl-XL, typically have BH1 through BH4 domains.

The pro-apoptotic members can be divided into two groups. The first group consists of proteins

such as Bax and Bak that contain BH1, BH2 and BH3 domains. The second group consists of

proteins such as Bad and Bim that contain only BH3 domains. The BH domains have functional

and structural significance. Many members of this family, such as Bcl-2 and Bcl-XL, are

predominantly localized to the outer membrane of mitochondria, while others interact with

mitochondria indirectly. In response to a variety of apoptotic stimuli, pro-apoptotic Bcl-2 family

members (such as Bax or Bak) initiate the mitochondrial dependent apoptotic pathway by

causing a loss of outer mitochondrial membrane integrity [162]. This releases apoptogenic

proteins located in the intermembrane space of mitochondria, such as cytochrome c,

Smac/Diablo, and apoptosis inducing factor (AIF) into the cytosol [163, 164]. Cytochrome c, an

electron carrier within the respiratory chain, interacts directly with Apaf-1 in the cytoplasm

leading to the ATP dependent formation of a macromolecular complex known as the apoptosome

[163, 164]. This complex recruits and activates the aspartyl directed protease caspase-9.

Activated caspase-9 can activate additional caspase-9 molecules, as well as the downstream

caspases such as caspase-3 or -7, resulting in morphological features of apoptosis.

Chapter 1

34

Smac/DIABLO, another mitochondrial protein released into the cytosol in response to apoptotic

stimuli, promotes caspase activation by eliminating inhibitory of apoptosis protein (IAP)

function [165]. AIF induces a caspase independent cell death and is critical for developmental

apoptosis [166]. Anti-apoptotic members Bcl-2 and Bcl-XL inhibit mitochondrial dependent

apoptosis by preventing Bax or Bak from disrupting the integrity of the outer mitochondrial

membrane. Previous studies have shown that DNA damaging agents, serum deprivation, and

endoplasmic reticulum stress agents trigger apoptosis through the mitochondrial dependent

pathway. Fibroblasts from embryos of mice lacking either Bax and Bak genes or cells that over

express BcL-XL or Bcl-2 are resistant to these apoptotic agents [167]. The mechanisms by which

these apoptotic stimuli converge on Bax or Bak to activate mitochondrial dependent apoptosis

remain unknown.

The extrinsic pathway is initiated when a death ligand, such as FasL or TNF , interacts

with its cell surface receptor, Fas (CD95) or TNF receptor (TNFR1/2) [168]. This results in the

formation of a death-inducing signaling complex (DISC). The formation of DISC involves

adaptor proteins such as FADD (Fas-associating protein with death domain) or TRADD (TNF

receptor associating death domain) [169, 170]. These proteins are involved in the recruitment of

pro-caspase-8 and its subsequent proteolytic activation. A variety of cell types undergoing

apoptosis through this pathway show strong activation of caspase-8 and direct activation of

caspase-3 [171]. In contrast, other cell types initially display a weak activation of caspase-8,

which subsequently employs the mitochondria for amplification of the death signal. This process

occurs by the caspase-8 dependent cleavage of Bid, a pro-apoptotic factor [172, 173]. A

truncated Bid requires either Bax or Bak to induce the loss of outer mitochondrial membrane

Chapter 1

35

integrity leading to cytochrome c release and caspase-9 activation [174]. Thus, there is cross talk

between the extrinsic and intrinsic pathways through truncated Bid.

Several studies indicate that oxygen deprivation can induce apoptosis in a variety of cell

types. As long as cells have an adequate supply of ATP during oxygen deprivation, apoptosis can

be executed [175]. However, if cells are deprived of oxygen and glucose then cells undergo

necrosis. The requirement for ATP to execute apoptosis during oxygen deprivation is attributed

to energy dependent activation of caspases. Cells over-expressing the anti-apoptotic proteins Bcl-

2 or BcL-XL have been shown to prevent oxygen deprivation induced apoptosis by inhibiting the

release of cytochrome c from the mitochondria [176-178]. Fibroblasts from mice lacking both

Bax and Bak genes are resistant to oxygen deprivation induced apoptosis [175]. Furthermore, the

pro-apoptotic protein Bax translocates from the cytosol to the mitochondria during oxygen

deprivation [179]. Cytochrome c is released and caspase-9 is activated in oxygen-deprived cells

undergoing apoptosis. Cytochrome c is released independent of caspase activation since

cytochrome c is still released in the presence of the caspase inhibitor zVAD. Fibroblasts from

caspase-9 or Apaf-1 deficient mice transformed with c-Myc and H-ras are resistant to cell death

during oxygen deprivation [180]. Consistent with this, Bid null fibroblasts are able to undergo

apoptosis in response to oxygen deprivation indicating that the extrinsic pathway does not

contribute to oxygen deprivation induced apoptosis [181]. In Jurkat cell lines, hypoxia-induced

apoptosis was not affected by lack of caspase-8 or FADD, whereas overexpression of Bcl-2 or

expression of dominant-negative caspase-9 mutant rendered the cells resistant to hypoxia-

induced apoptosis [182]. Together, these results suggest that hypoxia-induced apoptosis mainly

relies on intrinsic, mitochondrial pathways.

Chapter 1

36

The mitochondria are the central organelle

in the intrinsic pathway. In some

circumstances, however, the endoplasmic reticulum (ER) or sarcoplasmic reticulum in muscle

cells plays an important role in the hypoxia-induced mitochondrial death pathway, as well as

mediating cell death independently of mitochondria. Although the mechanisms by which the ER

brings about cell death are poorly understood, increases in intracellular Ca

2+ appear

to be central.

ER Ca2+

stores are thought to be increased by Bax and Bak, which are located at ER, as well as

mitochondrial membranes [183, 184]. Increased ER Ca

2+ facilitates a more robust release of Ca

2+

into the cytoplasm on delivery of an apoptotic stimulus, and may activate several apoptotic

mechanisms. First, mitochondrial Ca

2+ overload can trigger mitochondrial permeability transition

pore (MPTP) opening and cytochrome c release [185]. Cytochrome

c binds the inositol 1,4,5-

trisphosphate (IP3) receptor, one of the ER Ca

2+ release channels, to further stimulate Ca

2+

release [186]. Second, increased intracellular Ca

2+ can activate calpain. Calpain can cleave Bid,

providing another mechanism for cytochrome c release. Calpain activation also causes cleavage

of procaspase-12 [187]. Caspase-12 has been shown in knockout mice to be required for

apoptosis induced specifically by ER stress [188]. Cleaved caspase-12 translocates to the

cytoplasm and activates caspase-9 independently of apoptosome formation [189, 190]. These

events provide a mitochondria-independent mechanism for ER-mediated apoptosis. Some signals

that activate the ER death pathways originate within this organelle itself, where a complex array

of pathways mediate the unfolded protein and other ER stress responses [191]. In addition,

given

their roles in carrying upstream apoptotic stimuli to Bax and Bak at the mitochondria, BH3-only

proteins would be anticipated to perform an analogous function in the ER pathway.

BH3-only

proteins Bik (Bcl-2–interacting killer) and Puma have been implicated in the ER death

pathway

[192]. It remains unclear, however, whether these proteins function to relay signals from the

Chapter 1

37

periphery to the ER and/or from the ER to mitochondria. However, upstream signals originating

in the extrinsic pathway are known to be linked with the ER by Bap31 (B-cell receptor–

associated protein 31), an integral ER membrane protein that is cleaved by caspase-8 resulting in

ER Ca2+

release [193].

The apoptotic pathways described above have been schematically represented below.

Figure 1.6 Pathways of apoptosis.

Source: Gupta S et al. (2006) Lessons learned from apoptosis.

Chapter 1

38

1.11.2 Regulation of apoptosis during hypoxia

Both pro-and anti-apoptotic genes have been shown to be regulated by hypoxia. The

molecular pathways triggering the apoptotic response to hypoxia are far from completely

understood. Both p53 and Bcl-2 family members are involved in hypoxic activation of apoptosis.

The PI-3 kinase pathway plays an important role during oxygen deprivation induced cell death.

Electron transport inhibition also regulates hypoxia induced apoptosis.

1.11.2.1 Role of p53 in hypoxia-induced apoptosis

The transcription factor p53 has been implicated in regulating oxygen-deprivation

induced apoptosis. p53 can induce the expression of apoptotic genes such as Bax, NOXA,

PUMA and PERP [194]. Oxygen deprivation leads to p53 protein stabilization [148]. Hypoxia

causes p53 interaction with transcriptional repressor mSin3A but not with the transcriptional

activator p300 [128]. Also hypoxia causes localization of p53 to the surface of the mitochondria

[195]. In addition, severe hypoxia causes p53 accumulation by down-regulating its negative

regulator mdm2 and activation by post-translational modifications [196]. Hansson and

colleagues demonstrated that the DNA-binding domain of p53 binds two specific motifs adjacent

to and within the ODD domain of HIF-1 , irrespective of HIF-1 hydroxylation status [197].

Moreover, HIF-1 null cells accumulate more p53 protein than wild-type cells at low oxygen

concentrations, suggesting the expression of HIF-1 to cause p53 protein accumulation in

hypoxic cells, and thus induce apoptosis through p53 activation [198]. The hypoxia-dependent

post-transcriptional phosphorylation of p53 is by the ATR kinase, presumably due to replication

arrest that occurs under hypoxia. Reoxygenation and the consequent DNA damage can then

activate the ATM kinase which maintains p53 phosphorylation [152]. Hypoxic stress results in

Chapter 1

39

the association of p53 with transcriptional co-repressors and not co-activators capable of

repressing anti-apoptotic molecules such as stathmin [128].

1.11.2.2 Role of Bcl-2 family proteins in hypoxia-induced apoptosis

Pro- and anti-apoptotic members of the Bcl-2 family play a role in hypoxia-induced

apoptosis [175, 181]. BH3-only proapoptotic proteins, PUMA and Noxa, and BNIP3 and

BNIP3L have emerged as potential initiators of apoptosis by low oxygen concentrations [199-

202]. HIF-1 has been shown to induce the expression of BNIP3 (formerly called Nip3), a pro-

apoptotic member of the Bcl-2 family. BNIP3 heterodimerizes with Bcl-2/BcL-XL at

both the mitochondrial and non-mitochondrial sites [202]. Removal of the BH3 domain does not

inhibit apoptotic activity of BNIP3; instead, the transmembrane domain is critical for its

function/activity.

A direct role for HIF-1 in regulating sensitivity to oxygen deprivation induced apoptosis

come from genetic studies using embryonic stem cells with HIF-1 deleted. HIF-1 null cells

show a decrease in apoptosis compared to wild-type cells during oxygen deprivation [203].

Wild-type cells exhibit a decrease in Bcl-2 protein levels and an increase in p53 levels. In

contrast, HIF-1 null cells display no changes in p53 or Bcl-2 protein levels during oxygen

deprivation. Also, pancreatic cancer cells with constitutive expression of HIF-1 are resistant to

apoptosis induced by oxygen deprivation when compared to cells without constitutive expression

of HIF-1 [204]. Murine hepatoma cell lines without a functional HIF-1 display no difference

in sensitivity to oxygen deprivation induced apoptosis. Thus, HIF-1 can have various effects on

oxygen deprivation induced apoptosis depending on the cell type.

Other anti-apoptotic proteins regulated by hypoxia include (i) ORP150, an ER associated

heat shock protein, whose overexpression has been shown to prevent the release of cytochrome c

Chapter 1

40

and cell death in neurons deprived of oxygen. The precise mechanism by which ORP-150

inhibits cytochrome c release remains unknown. Presumably, ORP-150 prevents activation of

pro-apoptotic Bcl-2 family members [205]; (ii) IAP-2, which inhibits caspase activity IAP-2 is

induced in response to oxygen deprivation in a HIF-1 independent mechanism [206, 207] and

(iii) RTP801, a recently identified and cloned HIF-1 target gene. Hypoxia induces RTP801 in a

HIF-1 dependent manner. Over expression of RTP801 under normal oxygen conditions is toxic

in neuron-like PC12 cells while it is protective against oxygen deprivation induced cell death in

dividing PC12 cells and MCF-7 cells [208].

1.11.2.3 Role of PI3-kinase pathway in hypoxia-induced apoptosis

The phosphoinositide-3 kinase (PI3K)/Akt pathway is a potent mediator of cell survival

signals [209]. Activated PI3K phosphorylates inositol phospholipids to phosphatidylinositol

(3,4,5)-triphosphate (PIP-3). The increase in PIP-3 at the plasma membrane recruits Akt via its

pleckstrin homology (PH) domain. Akt, activated upon phosphorylation, in turn, can

phosphorylate and inactivate several substrates including the pro-apoptotic Bcl-2 family member

Bad, the forkhead family transcription factor FKHRL1 and caspase-9 [210-212]. Akt can also

mediate cell survival through hexokinase, an enzyme involved in the first committed step in

glycolysis [213]. Glycolysis utilizes glucose in the anaerobic production of ATP, which is

necessary for cell survival. Both hexokinase II and the pro-apoptotic protein Bax can bind to

mitochondria at the site of voltage dependent anion channel (VDAC). Binding of hexokinase to

the VDAC can inhibit or block Bax binding, thereby preventing cytochrome c release and

subsequent activation of apoptosis [214]. The tumor suppressor PTEN is involved in the negative

regulation of the PI3K/Akt pathway [215]. PTEN is a dual specificity phosphatase, which is

capable of dephosphorylating inositol phospholipids. Thus, PTEN activation results in decreased

Chapter 1

41

levels of PIP-3, leading to decreased Akt activity and increased apoptosis. PTEN null mouse

embryonic fibroblasts exhibit constitutively elevated activity of Akt and display decreased

sensitivity to cell death in response to a number of apoptotic stimuli, including UV irradiation

[216].

In cardiac myocytes adenoviral gene transfer of activated Akt protects against oxygen

deprivation induced apoptosis in vitro [217]. In contrast, over expression of activated Akt in

Rat1a fibroblasts did not protect cells from oxygen deprivation induced apoptosis [175]. Other

studies have shown that oxygen deprivation leads to Akt activation in PTEN null glioblastoma

cell lines [218]. Expression of wild-type PTEN in these cell lines prevented Akt activation in

response to oxygen deprivation. However, expression of PTEN at highly elevated levels did not

alter sensitivity to oxygen deprivation induced apoptosis in PTEN null glioblastoma cell lines.

Thus, the ability of Akt to prevent cell death during oxygen deprivation might be restricted to

cell type.

1.11.2.4 Role of electron transport inhibition in hypoxia-induced apoptosis

Electron transfer through the respiratory chain is coupled to the directional movement of

protons across the inner mitochondrial membrane. This movement across the membrane

establishes an electrochemical potential that provides the thermodynamic driving force for the

F1F0-ATP synthase to generate ATP in the matrix. Hypoxia leads to an inhibition of the electron

transport chain at cytochrome c oxidase, resulting in a decrease in inner mitochondrial membrane

potential. This initial decrease in inner mitochondrial membrane potential due to electron

transport inhibition during oxygen deprivation is the trigger for Bax or Bak activation [179].

Furthermore, mitochondrial membrane potential decreases in response to hypoxia prior to

cytochrome c release [175]. Cells devoid of mitochondrial DNA (o cells) do not undergo cell

Chapter 1

42

death in response to oxygen deprivation. Mitochondrial DNA encodes 13 polypeptides, including

the three catalytic subunits of cytochrome c oxidase, whereas nuclear DNA encodes the pro-

apoptotic protein cytochrome c. Therefore, o cells do not have a functional electron transport

chain and must rely only on ATP derived from anaerobic glycolysis for survival and growth. The

mitochondrial dependent cell death pathway is intact in o cells, as shown by their ability to

undergo death in response to a variety of apoptotic stimuli such as doxorubicin, growth factor

withdrawal, and staurosporine treatment [219, 220]. The inability of o cells to inhibit the

electron transport chain during oxygen deprivation is one explanation as to why o cells are

resistant to oxygen deprivation induced cell death. An alternate explanation is that o cells have

adapted to glycolysis. The loss of mitochondrial generated ATP due to electron transport

inhibition during oxygen deprivation could lead to activation of Bax or Bak. Since o cells have

adapted to glycolysis and show no acute changes in mitochondrial ATP levels during oxygen

deprivation, these cells might be resistant to hypoxia induced cell death. Presently, it remains

unresolved whether adaptation to glycolysis or electron transport inhibition is sufficient to

prevent hypoxia induced apoptosis.

1.12 Oxygen sensing mechanisms

Cells generally respond to prolonged hypoxia by reducing energy consumption and up

regulating ATP producing pathways [221]. The universal existence of these homeostatic

processes implies that all cells have the ability to sense changes in oxygen concentration and

their persistence in all mammalian species attests to their importance in determining survival

[222, 223]. The specific tissue of the carotid body is comprised of groups of glomus cells,

enveloped by glial-type sustentacular cells, and innervated by sensory nerve fibers. These units

Chapter 1

43

sense arterial pO2 and respond to hypoxia by initiating a variety of responses, which include the

arterial chemoreflex, i.e., increasing firing activity in the carotid sinus nerve. Despite extensive

work to identify the “oxygen sensor” in tissues whose specific role is to regulate the response to

changes in oxygen supply such as the glomus cells of the carotid body, the molecular

mechanism(s) involved have yet to be identified. It is likely that there are multiple O2 sensors

that respond to various pO2 levels, are differentially distributed and subserve different cellular

processes [222]. Current hypotheses centre around a number of biomolecules that can bind to O2,

either as a reversible ligand, producing allosteric shifts of the sensor, as a substrate, capable of

direct oxidation of the sensor or enzymatically converted to reactive oxygen species (ROS)

which, in turn, mediate the action on the effector molecules. Accordingly, the following

candidates for the mechanism of oxygen sensing have been proposed.

1.12.1 Evidence of heme as an oxygen sensor

This ligand model is based on the observation that heme proteins, such as hemoglobin

and cytochrome c-oxidase, bind O2 reversibly and the resulting configurational changes are

involved in the O2-dependent regulation of ion transporters and enzyme activity [224]. A heme

based O2 sensor has been proposed in erythropoeitin secreting cells because their response to

hypoxia is reversed by carbon monoxide (CO) which binds with greater affinity to and displaces

Figure 1.7 Heme sensor model.

Source: Lopez-Barneo J (2001) Cellular mechanism of oxygen sensing.

Chapter 1

44

oxygen from heme groups and is mimicked by incubation of the cells with iron chelators or

cobalt, which interfere with heme synthesis or render heme unable to bind O2 [225]. The heme-

sensor model, however, lacks direct experimental support. Moreover, it is now known that CO

can interact directly with HIF-1 , preventing its dimerization, and that iron and cobalt have

opposing effects on the interaction between pVHL and HIF-1 providing alternate explanations

for the observations upon which the hypothesis is based. Although HIF-1 has a PAS domain

that could bind a heme group, no direct interaction of HIF-1 with O2 has been demonstrated

[226].

The enzymatic production of reactive oxygen species (ROS) is sensitive to oxygen

availability and, hence, is a potential signaling pathway for transmission of the hypoxic stimulus

[227]. Although ROS can be produced at numerous cell sites and organelles, two systems have

been investigated as potential O2 sensors are NADPH oxidases and the mitochondrial electron

transport chain.

1.12.2 NAD(P)H oxidases

This plasma membrane-associated, phagocytic oxidase (NOX2) system is a multisubunit

assembly consisting of a membrane-bound catalytic complex of gp91phox

and p22phox

subunits,

which together form a flavo-cytochrome b558, and a cytosolic regulatory component consisting

of p47phox

, p67phox

, as well as other regulatory units including the GTPases Rac-1 and Rac-2

[228]. In NOX1 and 3 other isoforms substitute for gp91phox

and possibly other sub-units but are

mechanistically similar to NOX2; however NOX4 apparently does not require the cytosolic sub-

units for activation [229].

Chapter 1

45

This “membrane model” proposes that NAD(P)H-derived electrons are shuttled to O2 by

NAD(P)H oxidase at a rapid rate during normoxia, causing superoxide production. Under

hypoxic conditions, limited oxygen supply decreases superoxide generation. Reduced H2O2

production and a shift of cytosolic redox pairs to the reduced state ensues [230]. In pulmonary

vascular smooth muscle, this has been proposed as the mechanism underlying hypoxic

pulmonary vasoconstriction. According to this model, hypoxia inactivates redox-dependent

membrane K+ channel, results in membrane depolarization, opening of voltage-dependent Ca

2+

channels, entry of Ca2+

, and vasoconstriction [231, 232]. The findings that O2-sensitive K+

currents are potentiated by exogenous H2O2 and by activation of the oxidase with phorbol esters,

and that regulation of K+ channels by O2 is abolished in the transgenic oxidase-deficient mouse

with a null gp91phox

alleles provide experimental support for this proposal [233, 234].

Figure 1.8 Structure of NAD(P)H oxidase

Source: Tracy A. (2006) Chronic granulomatous disease, our understanding.

Chapter 1

46

There is strong evidence that NOX2 and a reduction in ROS underlies the response to

hypoxia in neuroepithelial bodies, with consequent inhibition of K+ channels and depolarization,

an important component of this evidence being loss of O2 sensing in gp91phox

-deficient mice

[235-237]. In contrast, O2 sensing was retained in glomus cells, adrenomedullary chromaffin

cells and lung from mice lacking gp91phox

, effectively ruling out any role for NOX2 in these

tissues [238-241]. Although knockout of p47phox

did not abolish O2 sensing in either lung or

glomus cells, in the former it caused inhibition of a rapid transient phase of HPV, whilst in

glomus cells it enhanced O2 sensitivity and abolished the hypoxia-induced elevation in ROS

[233, 241]. It has been suggested that the latter represents an important modulatory role for a

non-phagocytic NOX in glomus cells [234]. NOX4 has been shown to confer O2 sensitivity upon

TASK-1 channels in a model cell system, and TASK-1 has been implicated in O2 sensing for

both PASMCs and glomus cells [242-244]. Thus, although viable as an O2 sensor for specific

cellular functions in select cell types, the persistence of robust hypoxic pulmonary

vasoconstriction in the gp91phox

knockout mouse argues against a universal role [235, 238, 239,

245, 246].

Figure 1.9 NAD(P)H oxidase as oxygen sensor.

Source: Lopez-Barneo J (2001) Cellular mechanism of oxygen sensing.

Chapter 1

47

1.12.3 Mitochondria

Mitochondria, the largest consumers of oxygen, play a vital role in determining the

cytosolic pO2 and the O2 gradient between alveoli and cytosol. They are also recognized as

important signaling organelles [247]. To generate ATP by oxidative phosphorylation,

mitochondria use oxygen as the final electron acceptor. Accordingly, the mitochondria are well

positioned as the loci of cellular oxygen sensing [247]. The electron transport chain (ETC) is a

multi-step redox process that occurs in the inner mitochondrial membrane.

Figure 1.10 Mitochondrial Electron Transport Chain.

Source: Fig. 21.30, Biochemistry, 2nd

edition, R. Garrett and C. Grisham, Saunders Publishing

Chapter 1

48

The Kreb's cycle and -oxidation of fatty acids generate reduced nicotinamide adenosine

dinucleotide (NADH) and flavin adenine dinucleotide (FADH2) which are oxidised by the

electron transport chain (ETC) in the mitochondrial inner membrane. The components of the

ETC are outlined in Figure 1.10. Briefly, oxidation of NADH in complex I and FADH2 in

complex II leads to transfer of 2 electrons to ubiquinone to form ubiquinol. This is reoxidized by

complex III (cytochrome bc1) in two stages. One electron is first removed by the Fe–S group

leaving ubisemiquinone, and transferred via cytochromes c1 and c to complex IV (cytochrome c

oxidase, COX). The ubisemiquinone left behind is reoxidised to ubiquinone by cytochrome bL,

which passes the remaining electron to cytochrome bH. This reduces ubiquinone first to

ubisemiquinone and then back to ubiquinol, which re-enters complex III. Oxidation of one

molecule of ubiquinol back to ubiquinone thus takes 2 cycles of complex III, with the sequential

transfer of its 2 electrons to cytochrome c. These electrons are finally and sequentially

transferred by cytochromes a and a3 in complex IV to O2 to form water. The operation of

complexes I, III and IV cause extrusion of protons thus generating the mitochondrial membrane

potential ( m) and proton gradient ( pH) which drive the F0F1 ATP synthetase (reviewed in

[248]) . The mechanisms of the ETC are such that at various points single electrons can be lost to

molecular O2 to form reactive O2 species (ROS) in the form of superoxide, primarily from

reduced flavins in complex I and ubisemiquinone in both Qo (intermembrane space) and Qi

(matrix) sides of complex III; as much as 3% of electron flux through the ETC may be

constitutively lost in this way [249]. Superoxide is rapidly dismuted to peroxide by cytosolic and

mitochondrial superoxide dismutase (CuZnSOD and MnSOD respectively).

There is a wide consensus that inhibitors of oxidative phosphorylation or procedures that

modify mitochondrial function strongly affect O2 sensing in PASMCs, glomus cells and

Chapter 1

49

adrenomedullary cells though not in neuroepithelial bodies [250-254]. Significant controversy

exists, however, concerning the signalling mechanisms that link mitochondrial function to the

effectors, and there are currently three main hypotheses of mitochondrial O2 sensing, involving

cytosolic redox state, reactive O2 species (ROS) and energy state respectively. Under normal

physiological conditions, the ETC is a significant source of reactive oxygen species (ROS) [249].

Increased ROS generation occurs under hypoxic condition, when limited oxygen availability

decreases the Vmax of cytochrome oxidase, causing iron to remain in the ferric (Fe3+

) state,

thereby inhibiting prolyl hydroxylase activity HIF- stabilization and transactivation [255].

ROS generation and hypoxia-responsive gene expression occur simultaneously in

cardiomyocytes and Hep3B cells incubated under hypoxic conditions and these effects are

abolished in mitochondria-deficient cells. Others report that hypoxic responses can be blocked

by proximal ETC inhibition and addition of exogenous scavengers of ROS [86]. Similarly,

overexpression of enzymes that reduce ROS levels such as catalase or glutathione peroxidase

inhibit hypoxic pulmonary vasoconstriction (HPV) and the hypoxia-induced elevation in

intracellular [Ca2+

] and reverses hypoxia-induced HIF activation [256, 257]. However, the exact

correlation between pO2 and ROS production, the downstream targets of ROS, and how ROS

regulate these targets at the molecular level remain to be established.

Chapter 1

50

1.13 Thesis objective

The pulmonary and systemic circulations have evolved to preserve blood and oxygen

supply to the tissues, and this is vital for survival in physiological as well as pathological

conditions. By contraction and relaxation, VSMCs alter the luminal diameter, which enables

blood vessels to maintain an appropriate blood pressure. VSMCs are also important in vessel

remodeling in hypoxia induced disease states, such as arteriosclerosis and pulmonary

hypertension. In these cases, hypoxia regulates both cell proliferation and/or cell survival to alter

the structure of the vessels. Thus VSMCs are suited not only for short-term regulation of the

vessel diameter, but also for long-term adaptation, via structural remodeling by changing cell

number.

The function of VSMCs, and their responses to hypoxia, the basis for several

cardiopulmonary disorders, has been mostly studied in relation to the function of their adjacent

structures – in particular to the neighboring endothelial cells and fibroblasts. The direct response

of VSMCs in either the systemic or the pulmonary circulation to varying oxygen concentrations

remains largely unknown. Besides, these VSMCs experience a broad range of oxygen tensions

across the vessel wall, with an oxygen concentration of about 11.2% at the lumen to about 2.2%

at a depth of 150 . Previous studies have shown hypoxia to act as a mitogen, or, in some other

studies to induce growth arrest and cell death. Lack of consistency thus precludes a unifying

hypothesis.

There are several structural and physiological differences between the systemic and

pulmonary circulations. The bioenergetic processes that maintain cellular integrity depend on a

continuous supply of oxygen and substrates. To achieve this function, vascular smooth muscle

cells (VSMCs), in both the circulations, have adapted to tolerate relatively prolonged periods of

Chapter 1

51

hypoxia whilst maintaining appropriate distribution and delivery of essential substrates to other

tissues. However, there is a contrast in their basic response to hypoxia. Whereas in the systemic

circulation vasodilatation to hypoxia (hypoxic systemic vasodilatation, HSV) improves blood

supply and substrate delivery to hypoxic tissue, in the lung hypoxic pulmonary artery

vasoconstriction (HPV) maintains ventilation/perfusion matching and arterial oxygenation.

These could lead to differences in the proliferative and survival response of VSMCs from the

systemic and pulmonary circulations.

Chapter 2 reports the responses of human aortic smooth muscle cells to varying degrees

of hypoxia and the oxygen regulation of their proliferation and survival. Chapter 3 studies human

pulmonary artery smooth muscle cells – their proliferative and survival response/s compared

under normoxic and hypoxic conditions. Finally, based on the responses of aortic and pulmonary

artery smooth muscle cells to varying oxygen concentrations, we propose a possible molecular

mechanism that accounts for the divergent responses of the two cell types to hypoxia.

Chapter 1

52

1.14 Aims and hypotheses

Aims:

Determine if hypoxia has different effects depending on degree to which cellular energy

status is compromised in HASMCs and HPASMCs.

Identify oxygen sensitive cell-cycle regulatory genes and potential regulatory pathways

through gene expression profiling.

Establish the relevance of the above in vitro findings to systemic vascular remodeling

during hypoxia in vivo.

Hypothesis 1:

HASMC proliferation is enhanced or inhibited depending on the severity of hypoxia.

Hypothesis 2:

Proliferation and survival of PASMCs under hypoxic conditions are determined by their

capacity to maintain cellular ATP levels and that differences between these cells and

those from the systemic circulation reflect this parameter.

Chapter 2

53

Chapter 2

Oxygen regulation of systemic arterial smooth muscle cell proliferation and survival

Chapter 2

54

2.1 Introduction

Arterial smooth muscle cells are exposed to a broad range of oxygen concentrations. In

the aortic wall O2 concentrations from 11.2% at the lumen to 2.2% at a depth of 150 m[5, 6]

have been recorded and longitudinal gradients of similar magnitude occur in the normal

microcirculation[40]. Regions of severe hypoxia ( 1%O2) exist subjacent to atherosclerotic

plaques[7] and in arteries of hypertensive[8], and diabetic[9] rodents. Smooth muscle cell

proliferation and survival are modulated by hypoxia [203, 258] leading to speculation about its

role in vasculogenesis [11], vascular remodeling [10] and atherosclerosis [54, 203, 258, 259].

The effects of hypoxic incubation on vascular smooth muscle cells in culture, however, have

been inconsistent; with both enhanced proliferation [258, 260, 261] and growth arrest and

apoptosis [142, 262-264] have been reported. More importantly, the effect of levels of

hypoxemia relevant to human cardiopulmonary disease on systemic arterial smooth muscle cell

turnover, in vivo, is unknown.

Inconsistencies notwithstanding, the experimental evidence supports an important role for

oxygen in regulating vascular smooth muscle cell growth and survival which, given their

physiological and clinical relevance, requires clarification. Since cell replication is highly

energy dependent [265] it is intuitive, though unproven, that hypoxia may have different effects

depending on the degree to which cellular energy status is compromised. Furthermore, although

a number of cell cycle regulatory genes are oxygen sensitive, discrepant results have prevented

the development of a unifying hypothesis that accounts for the divergent effects observed. This

study was carried out to determine if different responses are elicited in human aortic smooth

muscle cells subjected to hypoxic incubation under conditions which do or do not result in

cellular ATP depletion, whether these effects are relevant to vascular remodeling during hypoxia

Chapter 2

55

in vivo and to identify potential regulatory pathways using gene expression profiling in cells

exposed to conditions that elicit discordant responses.

Chapter 2

56

2.2 Materials and Methods

Antibodies and Reagents:

FITC (fluorescein isothiocyanate) -conjugated Ki67 antibody was obtained from Dako

(Glostrup, Denmark), Hypoxia Inducible Factor 1- alpha (HIF-1 ) antibody from Novus

Biologicals (Littleton, CO, USA) and Cell division cycle 6 (CDC6) antibody from Lab Vision

(Fremont, CA, USA). Mini chromosome maintenance 2 (MCM2) and p21 antibodies were from

BD Pharmingen (San Diego, CA, USA). p53 antibody was from Cell Signaling Technology

(Danvers, MA,USA) and Telomerase Reverse Transcriptase (TERT) antibody from Calbiochem

(San Diego, CA,USA). CaspACE FITC-VAD-fmk in situ marker and TUNEL kits were both

from Promega (Madison, WI, USA). JC-1 labeling kit, ATP bioluminescence assay kit and TO-

PRO-3 dye were purchased from Molecular Probes (Carlsbad, CA, USA). All other reagents

were from Sigma (St. Louis, MO, USA).

Cell Culture Studies:

Human aortic smooth muscle cells (HASMC, Cambrex Bio Science Walkersville, MD,

USA), were propagated to passage 6 in SMGM-2 medium (Cambrex) consisting of SmBM

medium supplemented with single aliquots of 0.1% insulin, 0.2% hFGF-B, 0.1% GA-1000

(Gentamicin and Amphotericin B) and 5% v/v FBS, 0.1% hEGF. Cells exposed to hypoxia were

placed in a humidified Plexiglas chamber (Billups Rothberg, San Diego, CA, USA) maintained

at 37°C and continuously flushed with gas mixtures containing 10%, 5%, 3% or 1% O2, 5% CO2,

balance N2. By convention, the term normoxic is applied to cells exposed to air/5% CO2 (culture

media O2 concentration = 20.5%) under otherwise identical conditions. Upon reaching 70%

confluence the media was changed and cells were incubated for a further 16 or 48 hrs under

Chapter 2

57

either normoxic or hypoxic conditions. Experiments were repeated three times using cells from

at least 2 human donors with 6 replicates per observation.

Cell Counting: After exposure to normoxia or hypoxia (10%, 5%, 3% or 1% O2) for 16 or

48 hours, cells were washed twice with HBSS and detached with 0.25% trypsin and 0.02%

EDTA. Cell number was determined by cell counting using a standard hemacytometer

(American Optical, Buffalo, NY, USA) and cell viability was assessed by Trypan Blue

exclusion. Initial cell counting studies indicate that the effects of 3% and 1% O2 differ

qualitatively (Figure 1A), therefore, further studies compared normoxic cells with cells incubated

at these O2 concentrations. The concentrations of dissolved O2 in the culture medium, measured

using the ISO2 dissolved oxygen meter (World Precision Instruments, Sarasota, FL, USA) were

20.5 ± 0.6%, 3.1 ± 0.4% O2 and 1.2 ± 0.3%O2, respectively under the three conditions. Steady

state oxygen concentrations were achieved within 30 minutes.

The effect of hypoxia on the response to platelet derived growth factor (PDGF) was

assessed in HASMCs incubated under normoxia for 48 h following which, the cell culture

medium was replaced with medium containing 0.5% FBS. 48 hours later, 10nM PDGF-BB

(R&D Systems, Minneapolis, MA, USA) or the same volume of diluent (100 l SmBM-0.5%

FBS) was added to the medium, and the cells were allowed to proliferate under either normoxic

or hypoxic (1% or 3% O2) conditions for 48 hours.

[3H]-Thymidine incorporation: HASMCs, at passage 6, were seeded at a density of 2x10

4

cells / well in Corning 24 well plates, and grown for 24 hours. The culture medium was then

removed and cells were incubated in 1% FBS in SMGM-2 media for another 24 hours. 1

microcurie of [3H]-Thymidine (specific activity 3.22TBq/mmol; Amersham) was added to each

Chapter 2

58

well and the cells were exposed to air, 1% O2 or 3% O2 for 16, 24, 48 or 72 hours. Following

each exposure, the cells were washed twice with phosphate buffered saline (PBS), and fixed with

ice-cold 10% (w/v) trichloroacetic acid (TCA) for 20 minutes. The resulting precipitate was

solubilized in 0.1 N NaOH (0.5 ml/well) at 37°C. Solubilized DNA was transferred into

scintillation vials and [3H]-Thymidine incorporation was quantified by scintillation counting

(Liquid Scintillation System, Beckman Instruments).

Ki67 protein: The presence of Ki67 was detected in cells grown under normoxic or hypoxic

(1% or 3% O2) conditions for 16 or 48 hours. Detached cells were centrifuged at 1500 rpm for

10 minutes at 20°C. The supernatant was removed, the pellet suspended in 200 l of membrane

shredding solution (99.5% v/v Ca2+

and Mg2+

free Dulbecco’s PBS, 0.5% v/v NP-40, 0.5 mM

Sodium-EDTA, 0.5% w/v BSA, 20μg/ml protease inhibitior, 0.2 mg/ml RNAse) and kept at

room temperature for 15 minutes. Samples were incubated with 10 l of antibody in the dark for

30 minutes and analyzed by flow cytometry (Model Epics Altra, Beckman Coulter, Fullerton,

CA, USA). A minimum of 10,000 events, per sample, was recorded and cell debris was excluded

by adjusting the forward light scatter threshold setting. The number of cells positive for Ki67

was calculated using CELLQuest software (Becton

Dickinson, St. Louis, MO, USA).

Annexin V/Propidium Iodide labeling: The Roche Annexin V–FLUOS staining kit was

used to detect phosphatidylserine externalization (a marker of apoptosis) in HASMC, exposed to

normoxia or hypoxia (3% or 1% O2) for 16 or 48 hrs. The cell suspension was centrifuged at

1500 rpm for 10 minutes at 4° C. The pellet was resuspended in 5 ml of cold PBS and

centrifuged again. The supernatant was removed and the pellet suspended in 100 l of Annexin

V- FLUOS labeling solution (20μl Annexin V-Fluos labeling reagent and 20μl Propidium Iodide

Chapter 2

59

(PI) solution per ml of incubation buffer) at 37° C. Labeled cells were analyzed by flow

cytometry and the numbers of cells positive for either Annexin-V, or PI, or both, were

calculated.

Caspase activation: Caspase activation was detected in HASMCs exposed to normoxia or

hypoxia (3% or 1% O2) for 16 or 48 hours. CaspACE FITC-VAD-fmk is a FITC conjugate of

the cell permeable inhibitor of caspases. This structure allows delivery of the inhibitor into the

cell where binding to activated caspase, serves as an in situ marker for apoptosis. About 2 106

cells were incubated with 100 M FITC-VAD-FMK at room temperature in the dark for 20 min.

Cells were then washed, resuspended in PBS and the percentage of cells positive for activated

caspase quantified by flow cytometry.

Terminal deoxynucleotidyl transferase-mediated dUTP nick-end-labeling (TUNEL): HASMCs

exposed to normoxia or hypoxia (3% or 1% O2) for 16 or 48 hours were centrifuged at 1500 rpm

for 10 minutes at 4° C. The supernatant was removed and the cells fixed in 1% w/v iced

paraformaldehyde for an hour. The cells were washed with PBS and 70% ethanol. DNA labeling

solution (TdT enzyme and FITC-dUTP, DeadEndTM

Fluorometric TUNEL System, Roche,

Basel, Switzerland) was added and samples kept at 37oC for an hour, after which they were

analyzed for DNA breaks by flow cytometry.

DNA Content/Cell Cycle Analysis: HASMCs exposed to normoxia or hypoxia (3% or 1% O2)

for 16 or 48 hours were trypsinized, pelleted, resuspended, and washed once in cold PBS. The

cells were fixed with 70% ethanol and maintained at 4°C for 60 min. Ethanol was washed out

and cells were resuspended in 1ml of PBS and 10 l RNAse, and incubated for 45 min at room

temperature. PI (10 l) was added and cells were incubated in dark at room temperature for 30

Chapter 2

60

min prior to analysis by flow cytometry. In each experiment, 10,000 cells were counted. The

amplitude of the fluorescent signal was analyzed to quantify DNA content in order to determine

the fraction of the cell population in each phase of the cell cycle (G0/G1, S, and G2/M).

Mitochondrial membrane potential: Depolarization of the mitochondrial membrane was

detected by a cytofluorometric method using the potential-sensitive probe 5,5', 6,6'-tetrachloro-

1,1',3,3'-tetraethyl-benzimidazolylcarbocyanine iodide (JC-1). The JC-1 monomer enters the

mitochondria at physiological membrane potentials [266] where, as a result of aggregation, its

emitted wavelength changes from 530 nm (green) to 590 nm (orange) when excited at 490 nm.

Disaggregation to the monomeric form during mitochondrial membrane depolarization is

detected as an increase in green emission. HASMCs exposed to normoxia or hypoxia (3% or 1%

O2) for 16 or 48 hours were incubated with JC-1 (10 g/ml) for 20 minutes in the dark, washed

and resuspended in 1 ml PBS. The percentage of cells positive for JC-1 monomers was

quantified by flow cytometry.

Intracellular ATP Concentration: HASMCs exposed to normoxia or hypoxia (3% or 1% O2)

for 16 or 48 hours were washed twice with ice-cold PBS and lysed by the addition of equal

volumes of 3.6% perchloric acid. Samples were centrifuged, and ATP concentrations in the

supernatants were

determined using an ATP bioluminescence assay (Molecular Probes),

according to the instructions provided by the manufacturer.

The photometer was set for a 5-s

delay period and a 5-s integration period. ATP levels were calculated using standard reference

solutions corrected for background luminescence.

Western blotting: Western analysis was used to quantify levels of the hypoxia-inducible

transcription factor HIF-1 , markers of cell division - CDC6[267] and MCM2[268], cell cycle

Chapter 2

61

regulatory proteins p21, p53 and telomerase subunit TERT in nuclear extracts from HASMCs

exposed to normoxia or hypoxia (3% or 1% O2) for 16 or 48 hours. Cells were lysed in buffer A

(10 mM HEPES [pH 7.8], 10 mM KCl, 0.1 Mm EDTA, 1 mM dithiothreitol [DTT], 0.1%

Nonidet P-40 [NP-40]) with protease inhibitors (5 μg of aprotinin per ml, 5 μg of pepstatin per

ml, 5 μg of leupeptin per ml, 0.5 mM Pefabloc, 1 mM phenylmethylsulfonyl fluoride) and

phosphatase inhibitors (10 mM sodium fluoride, 1mM sodium orthovanadate, and 20 mM -

glycerophosphate). Nuclear proteins were then extracted with buffer B (50 mM HEPES [pH 7.8],

420 mM KCl, 0.1 mM EDTA, 1 mM DTT, 5 mM MgCl2, 20% glycerol) containing both

protease and phosphatase inhibitors. Equal amounts of protein extracted from HASMCs,

incubated under normoxic and hypoxic (3% and 1% O2) conditions for 16 and 48 hrs were

loaded on 4-12% Tris-Glycine gels, separated by electrophoresis and transferred to

nitrocellulose. Membranes were blocked with 5% milk overnight, and probed with anti-CDC6

(1: 2000), anti-MCM2 (1: 2000), anti-HIF-1 (1:500), anti-p21 (1: 3000), anti-p53 (1: 1000) and

anti-TERT (1: 200). In all cases, protein concentration was determined by the Bradford assay

and appropriate volumes of extraction buffer to produce constant protein loading in each lane

were mixed with SDS loading buffer. Equality of protein loading and transfer efficiency were

corroborated by full-lane densitometry of the Ponceau red-stained membranes. Immunoblots

were probed with horseradish peroxidase (HRP)-donkey anti-rabbit IgG (1:1000 in blocking

buffer) and visualized by enhanced chemiluminescence (ECL Plus kit, Amersham Biosciences,

Buckinghamshire, UK). Band intensity was quantified by densitometry (Bio-Rad Laboratories,

Mississauga, ON, Canada).

Microarray analysis using Affymetrix Gene Chip hybridization: In three separate experiments

total RNA was isolated from HASMCs exposed to normoxia, 3% O2 and 1% O2 for 16 and 48

Chapter 2

62

hours using Trizol Reagent (GIBCO/BRL). The quality of RNA was assessed using an Agilent

2100 Bioanalyzer (version A.02.01S1232, Agilent Technologies). Hybridizations were

performed on the HG-U133A GeneChip Set with a total of 22,280 genes (Affymetrix, Santa

Clara, CA, USA). Samples were prepared for hybridization (6 hybridizations per experiment)

according to standard Affymetrix instructions and performed at the Toronto Genomic Core

Centre at the Hospital for Sick Children. Experimental design, gene lists, hierarchical trees, chip

hybridizations and statistical analyses were in compliance with the Minimum Information About

a Microarray Experiment (MIAME) guidelines[269]. Data obtained from the GCOS (GeneChip

Operating Software) analyses of the individual arrays were normalized using the RMA (Robust

Multi-chips Analysis) method. After filtering, 2 way-ANOVA (non equal variance) was

performed and differentially regulated genes were clustered using GeneSpring 7.0

(http://www.agilent.com). For details see the supplementary data (GEO Accession # GSE 4725).

Animal studies:

All protocols were in compliance with standards set by the Canadian Council of Animal

Care and were approved by the institutional animal care committee. Male Sprague-Dawley rats

(175–200g) were placed in a Plexiglas chamber into which the flow of air and nitrogen was

controlled independently. In preliminary experiments, the arterial pO2 in rats, breathing a gas

mixture containing 10% O2, averaged 38 Torr (range 35–42 Torr)[270]. Rats exposed to hypoxia

breathed a gas mixture containing 10% O2 for 48 hours. Normoxic control animals breathed

room air under otherwise identical conditions. Data from each animal were averaged to serve as

a single value for statistical analysis.

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63

Detection of Nonviable Aortic and Mesenteric Artery Smooth Muscle Cells: Nonviable

cells were detected by their failure to exclude propidium iodide. As a positive control

Lipopolysachharide (LPS) from Escherichia coli 055:B5 (0.1mg/kg body weight) was injected

into the right jugular vein 72 hours prior to sacrifice, to induce apoptosis. At the end of the

exposure period normoxic, LPS treated, and hypoxia exposed rats were anesthetized by

intramuscular injection of 0.08ml/kg xylazine (20mg/ml) plus 0.72ml/kg

ketamine (100mg/ml)

followed by intravenous injections of 5 mol/kg propidium iodide. After 15 minutes, an incision

was made at the left atrial appendage of the heart and flushed thoroughly with 300 ml PBS. En

face sections of aorta and mesenteric artery were perfusion-fixed with 3.7% paraformaldehyde

for 1 hour, washed with PBS and permeabilised with 0.2% Triton X-100. TO-PRO-3 was used

for nuclear counterstaining. Sections were then mounted on glass slides with glycerol/phosphate-

buffered saline, and viewed under a laser-scanning confocal microscope (BioRad Radiance,

Hercules, CA, USA). Cells

that did not exclude propidium iodide, as indicated by

nuclear

fluorescence, were considered nonviable.

Detection of apoptotic cells: DNA fragmentation in aortic and mesenteric artery sections

was detected by TUNEL (DeadEndTM

Fluorometric TUNEL System, Roche). Slides containing

paraffin-embedded sections (normoxic and hypoxic rat aorta and mesenteric artery) were

dewaxed, rehydrated, permeabilized with Proteinase K, preincubated with equilibration buffer

and incubated with labeling solution (rTDT and nucleotide mix with fluorescein labeled dUTP)

for 1hour at 37oC. The reaction was terminated by incubating samples in a stopping buffer for 30

minutes. After PBS washes and counterstaining with TO-PRO-3, the samples were mounted and

examined by laser confocal microscopy.

Chapter 2

64

Detection of proliferating cells: Bromodeoxyuridine (BrdU) was infused subcutaneously

using osmotic pumps (Model 2ML-2, Alza Corp, Palo Alto, CA). Pumps containing 0.32 g of

BrdU in 2 ml vehicle (0.4%DMSO) were implanted intrascapularly in normoxic and hypoxia-

exposed rats 48 hours prior to sacrifice. The rats received approximately 0.4mg BrdU/hour

delivered continuously. At the end of the labeling period thoracic aorta and mesenteric artery

segments were excised, fixed with paraformaldehyde, washed with PBS, dehydrated in graded

ethanol (70-100%), cleared in xylene, and embedded in paraffin. 5μm thick sections were cut on

an oscillating blade microtome (Leica, Wetzlar,

Germany) and placed on coated glass

microscope slides (Fisher Scientific, Pittsburgh, PA, USA). Dewaxed and rehydrated slides were

incubated with 2M HCl for 1hour at 37oC. The acid was neutralized by 0.1M borate buffer, pH

8.5. Following PBS washes, the slides were incubated with fluorescein labeled anti-BrdU

antibody (Roche) for 1hour at room temperature, protected from light. Slides were then washed

with PBS and mounted for microscopy.

Quantitative analysis: The number of TUNEL positive, BrdU and TO-PRO-3 labeled

cells were determined in 3 aortic and mesenteric artery sections from each of 6 animals from

each experimental group. Four to six randomly selected microscopic fields ranging from 90–300

cells per field were counted for each section and the number of TUNEL- or BrdU-positive nuclei

expressed as a percentage of the total number of nuclei. Nuclear density was measured was

determined using Image J analysis software (NIH 2002, http://rsb.info.nih.gov/ij) and expressed

as the number of nuclei per μm2 x 10

-4. Data from each animal in each group were averaged to

serve as a single value for statistical analysis. Apoptotic index was evaluated for PI and TUNEL

staining.

Chapter 2

65

Smooth muscle -actin staining: Aorta and mesenteric artery sections from normoxic and

hypoxic rats were embedded (OCT compound; Miles Scientific; Naperville, IL) and quickly

frozen in liquid nitrogen. Cryostat sections (100 thick) were stained overnight at

4°C with Cy3

conjugated monoclonal anti smooth muscle -actin (Sigma, St. Louis, MO, USA). Sections were

counterstained with TO-PRO-3, washed with PBS and viewed under the confocal microscope.

Data analysis: Data are presented as mean ± standard error of the mean of n observations

with P < 0.05 considered significant. The

significance of differences between individual means

was determined by two-tailed Student’s t test. Differences among multiple means were evaluated

by analysis of variance corrected for multiple measures where appropriate and, when overall

differences were detected, differences between individual means were evaluated post-hoc using

the Student Neuman - Keuls procedure.

Chapter 2

66

2.3 Results

The proliferative response of HASMCs during normoxic and hypoxic incubation

was evaluated by cell counting (Figure 2.1), [3H]-thymidine incorporation (Figure 2.2) and

nuclear levels of S phase proteins (Figure 2.3). As shown in figure 2.1A cell number is increased

after incubation at 5% O2 for 48 hours and after incubation at 3% O2 for 16 hours and 48 hours.

Incubation at 1% O2 for either 16 or 48 hours reduced the total cell number. Similarly, in rat

A7R5 smooth muscle cells, cell number increased 22.8± 6.5% and 31.4±7.3% after incubation at

3% O2 for 16 and 48hrs, respectively (p <0.05 vs. normoxic control, for both) and decreased by

38.8±5.9% and 49.7±8.6% after incubation at 1% O2 for 16 and 48hrs, respectively (p < 0.05 vs.

normoxic control, for both). Trypan blue was excluded (cells are viable) in 97.1±0.9% of

HASMCs after normoxic incubation and in 94±0.4% after 16hrs and 92.2±0.6% after 48 hrs of

incubation at 1% O2 (p < 0.05 vs. normoxic control values for both). HASMC viability did not

differ between normoxic cells and cells incubated at 3% O2.

HASMC (synchronized by prior incubation with 0.5% FBS) cell numbers were decreased

by incubation at 1% O2 in the absence and in the presence of PDGF after 16 (18.5±4.6% and

19.7±3.8% decrease from normoxic control values, respectively) and 48 hrs (24.4±4.5% and

26.7±2.9% decrease from normoxic control values, respectively, Figure 2.1B). Incubation of

HASMCs at 3% O2, either in the absence or presence of PDGF caused an increase in cell number

at both 16 (20.3±4.5% and 23.3±4.8% increase from normoxic control values, respectively) and

48 hrs (35.6±3.7% and 36.7±4.8% increase from normoxic control values, respectively, Figure

2.1C).

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67

Figure 2.1 (A) Effects of hypoxia on human aortic smooth muscle (HASMC) cell

numbers after incubation for 16 and 48 hours * p <0.05 vs. corresponding normoxic

control values, #,p <0.05 vs. corresponding 3% O2 values.

Chapter 2

68

Figure 2.1 Effects of hypoxia on the proliferative response of HASMC cell numbers to

10nM PDGF-BB after incubation at 1% O2 (B) and 3% O2 (C) for 16 and 48 hours. n=6 per

condition. * p<0.05 vs. corresponding normoxic control values.

Chapter 2

69

The effect of hypoxic incubation for up to 72 hours on [3H]-thymidine incorporation, in

HASMCs, is illustrated in Figure 2.2. Incubation at 1% O2 decreased (Figure 2.2A) whereas 3%

O2 increased (Figure 2.2B) the rate of [3H]-thymidine incorporation reflecting the effects of these

conditions on the rate of DNA synthesis.

The effects of hypoxia on other markers of cell proliferation, Ki67 (nuclear proliferation

marker), CDC6[267] and MCM2[268] (both form the prereplication complex at the initiation site

for DNA synthesis), were also studied (Figures 2.3 and 2.4). The percentage of cells staining

positive for Ki67 decreased after incubation at 1% O2, whereas 3% O2 had the opposite effect

(Figure 2.3 A and B). Similarly, CDC6 protein was decreased and increased in cells incubated at

1% and 3% O2, respectively (Figure 2.4, A and B) and MCM2 protein was reduced after

incubation at 1% O2 (Figure 2.4, C and D).

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Figure 2.2 (A) (3H)-Thymidine incorporation (counts per minute), in human aortic

smooth muscle cells is decreased after incubation at 1% O2 and (B) increased after

incubation at 3% O2 compared with the normoxic cells. n=6, * p <0.05 vs.

corresponding normoxic control values.

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71

Figure 2.3 The percentage of cells positive for the Ki67 antigen is decreased after

incubation at 1% O2 (A) and increased after incubation at 3% O

2 (B). * p <0.05 vs.

corresponding normoxic control values.

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72

Figure 2.4 CDC6 (A and B) and MCM2 (C and D) protein levels after normoxic and

hypoxic (1% O2 and 3% O2) incubation. 1% O2 decreased CDC6 protein level while CDC6

protein is increased in cells exposed to 3% O2. 1% O2 decreased MCM2 protein level with no

change in cells exposed to 3% O2. * p <0.05 vs. corresponding normoxic control values.

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To determine whether hypoxia alters HASMC proliferation through a general effect on

cell cycle progression or through mechanisms specific to a particular cell cycle phase or

checkpoint, DNA content was quantified by PI staining (Figure 2.5). After incubation at 1% O2

there is accumulation of G0/G1 phase cells and depletion of cells in S and G2/M phases (Figure

2.5A) indicating a delay in G1/S transition. Conversely, incubation at 3% O2 results in a decrease

in the percentage of cells in G0/G1 phase, an increase in S phase cells and a corresponding

increase in the percentage of cells undergoing mitosis (Figure 2.5B) suggesting acceleration of

progression through the G1/S interphase. 94±0.8% of cells remained viable (did not stain for

either Annexin V or PI) after normoxic incubation compared with 91.8±0.7%, and 83.7±0.8%

following incubation at 1% O2 for 16 and 48 hours, respectively (p < 0.05 vs. corresponding

normoxic control values for both). After incubation at 3% O2 the percentage of viable cells did

not differ from the normoxic control value.

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74

% cells

G0/G1 S G2/M

1% O2 48hrs 91.8* 5.1

* 3.1

1% O2 16hrs 85.1* 11.1

* 3.8

21% O2 80.2 14.8 5.0

% cells

G0/G1 S G2/M

3% O2 48hrs 67.6* 22.9

* 9.5

3% O2 16hrs 75.3* 16.9

* 7.8

21% O2 81.1 13.6 5.3

Figure 2.5 Flow cytometric analysis of propidium iodide stained cells incubated to 70%

confluence under normoxia, then incubated for a further 48 hours under normoxia or hypoxia (1%

or 3% O2) or for 32 hours under normoxia and 16 hours under hypoxia (1% or 3% O2).

(A) Incubation at 1% O2 increases the percentage of cells at the G0/G1 interphase compared with

the normoxic cells, whereas (B) incubation at 3% O2 increases the percentage of cells in G2/M at

16 and 48 hours, compared with the normoxic cells. n=6, * p<0.05 vs. corresponding normoxic

control values.

% cells in cell cycle stage

% cells in cell cycle stage

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75

Since differences in cell number may reflect changes in the rate of cell death as well as

proliferation, the prevalence of smooth muscle cell apoptosis after normoxic and hypoxic

incubation was assessed in HASMCs by Annexin V / PI staining and TUNEL (Figure 2.6). The

percentage of cells that stained with Annexin V-FITC only (early apoptotic cells) increased after

incubation at 1% O2 and decreased at 3% O2 compared to the normoxic cells (Figure 2.6A).

Similarly, caspase activity and the percentage of TUNEL positive cells are increased after

incubation at 1% O2 (Figure 2.6B and Figure 2.6C respectively).

Figure 2.6 (A) Annexin V/PI staining indicate that apoptosis increased after

incubation at 1% O2 and decreased at 3% O2. * p<0.05 vs. corresponding normoxic

control values.

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76

Figure 2.6 (B) Caspase activity and (C) TUNEL indicate that apoptosis is increased

after incubation at 1% O2. * p<0.05 vs. corresponding normoxic control values.

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77

Depolarization of the mitochondrial membrane both results from and contributes to

impairment of cellular ATP synthesis and is an early event in apoptotic cell death. Figure 2.7

illustrates the effect of hypoxic incubation on the percentage of cells positive for JC-1 monomers

(an index of mitochondrial membrane depolarization). In cells incubated at 1% O2 for 16 hours

there is an increase in the percentage of cells positive for JC-1, indicating mitochondrial

membrane depolarization, and a further increase after 48 hours (Figure 2.7 A). There was no

change in the percentage of JC-1 monomer-positive cells after incubation at 3% O2 (Figure 2.7

B).

As shown in Figure 2.8 A, there is a decrease in cellular ATP concentration in HASMCs

incubated for 16 hours at 1% O2 and a further decrease after 48 hours. Incubation at 3% O2 did

not alter cellular ATP concentration (Figure 2.8, B).

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78

Figure 2.7 Increased JC-1 monomer formation indicates mitochondrial membrane

depolarization after incubation at 1% O2 (A) but not at 3% O2 (B). * p<0.05 vs. corresponding

normoxic control values.

Chapter 2

79

Figure 2.8 Cellular ATP concentration is decreased after incubation at 1% O2 (A) but

not 3% O2 (B). * p <0.05 vs. corresponding normoxic control values.

Chapter 2

80

Gene expression profiling was used to identify oxygen regulated genes and, hence,

potential regulatory pathways. Complete gene expression data are presented in online (GEO

accession # GSE 4725). The fold changes from normoxic control values in the expression of

genes with known pro- and antiproliferative and pro- and antiapoptotic roles, which

demonstrated significant changes after hypoxic incubation, are presented in Table 2.1.

Table 2.1 Normalized expression of genes of pro- and antiproliferative genes and pro- and

antiapoptotic genes under normoxic and hypoxic conditions (1% O2 and 3% O2).

Genebank # Gene 1% O2 3% O2

16hrs 48hrs 16hrs 48hrs

Proproliferative genes

M63889 Fibroblast growth factor receptor 1 0.9 0.7 2.9 2.5

NM_00295

Replication protein A1

1.1 1.1 1.4 1.5

L14922

Replication factor C (activator 1) 1

0.9 1.0 1.4 1.5

NM_00125

Cell division cycle 20

0.5 0.4 0.8 1.3

NM_00479

Mitogen-activated protein kinase-activated

protein kinase 2

1.0 0.8 1.7 2.0

NM_00275

Mitogen-activated protein kinase 2

0.5 0.4 1.6 1.5

NM_00655

Mitogen-activated protein kinase kinase

kinase kinase 5

1.3 1.4 2.3 1.9

U29725

Mitogen-activated protein kinase 7

0.9 0.8 1.3 1.4

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81

Table 2.1 Normalized expression of genes of pro- and antiproliferative genes and pro- and

antiapoptotic genes under normoxic and hypoxic conditions (1% O2 and 3% O2) contd.

Genebank # Gene 1% O2 3% O2

16hrs 48hrs 16hrs 48hrs

Antiproliferative genes

NM_00039

Cyclin-dependent kinase inhibitor 1A, p21

1.4 2.6 0.9 0.7

NM_00561

Retinoblastoma-like 2 (p130)

1.5 1.7 1.0 1.0

NM_00154

Inhibitor of growth family, member 1-like 3.0 2.8 1.0 1.2

AA583044

Bone morphogenetic protein 2 2.9 3.4 0.9 0.8

NM_00447

Dual specificity phosphatase 1 1.7 1.6 1.1 1.0

U16996

Dual specificity phosphatase 5 1.5 1.6 1.0 0.9

AF013168 Tuberous sclerosis 1 1.3 1.3 0.7 0.4

Proapoptotic genes

AF307851

Tumor protein p53 3.3 4.0 0.8 0.8

NM_00436

Caspase 3 1.3 1.3 0.8 0.8

AB037736

Caspase 8 associated protein 2 1.9 2.1 1.3 1.3

AF310105

Caspase recruitment domain protein 7 1.5 1.4 0.9 0.9

NM_004760

Serine/threonine kinase 17a 1.3 1.3 0.9 0.9

NM_004226

Serine/threonine kinase 17b 1.5 1.4 1.1 1.1

NM_013229

Apoptotic protease activating factor 1.2 1.2 1.0 1.0

Antiapoptotic genes

AF041461

CASP8 and FADD-like apoptosis regulator 0.6 0.6 1.5 1.5

U72398

BCL2-antagonist of cell death 0.9 0.6 1.8 1.9

Chapter 2

82

Figure 8 presents the effects of hypoxic incubation on nuclear levels of HIF-1 , p21 and

p53. HIF-1 levels are increased to a similar extent after incubation at 1% and 3% O2. Levels

of p21 and p53 proteins, inhibitory regulators of cell cycle progression, are increased after

incubation at 1% O2 with no change after incubation at 3% O2. Levels of TERT protein, a

component of the telomerase complex, that increaseed and enhanced HASMC survival following

longer hypoxic epochs (>20 days), [271, 272] did not differ from the normoxic control values

after 16 and 48 hours of incubation at 1% and 3% O2 (data not shown).

Figure 2.9 (A) Nuclear levels of HIF-1 protein after incubation of HASMCs under

normoxic and hypoxic (1% or 3% O2) conditions. Solid bars represent protein levels after

normoxic incubation, open bars after incubation at 1% O2 and hatched bars after incubation at

3% O2. * p<0.05 vs. corresponding normoxic control values.

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83

Figure 2.9 Nuclear levels of p21 (B) and p53 (C) proteins after incubation of HASMCs

under normoxic and hypoxic (1% or 3% O2) conditions. Solid bars represent protein levels

after normoxic incubation, open bars after incubation at 1% O2 and hatched bars after

incubation at 3% O2. * p<0.05 vs. corresponding normoxic control values.

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84

As shown in Figure 2.10, exposure to hypoxia increased the number of propidium iodide

staining cells (Figure 2.10 A) with a corresponding increase in the number of TUNEL positive

nuclei (Figure 2.10 B) in both aortic and mesenteric artery sections.

Mes

ente

ric

arte

ry

Ao

rta

Normoxic Hypoxic

Ao

rta

M

esen

teri

c ar

tery

Figure 2.10 (A) Propidium iodide staining of en face sections in paraffin embedded

sections of normoxic and hypoxic (48hrs) rat aorta and mesenteric artery (40X

magnification).

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85

B

Figure 2.10 (B) TUNEL in paraffin embedded sections of normoxic and hypoxic

(48hrs) rat aorta and mesenteric artery (40X magnification). (C) Quantitative analysis

confirms increased cell death and apoptosis after hypoxic exposure.

Mes

ente

ric

arte

ry

Ao

rta

Normoxic Hypoxic

Ao

rta

M

esen

teri

c ar

tery

C

Aorta Mesenteric artery

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86

Figure 2.11 (panel A) shows that, in aortic and mesenteric artery sections from hypoxia

exposed rats, there is an increase in the number of BrdU labeled cells compared with the

normoxic control group. Compared to normoxic animals, aortic and mesenteric artery sections

from hypoxia-exposed rats also demonstrate increased medial nuclear density (Figure 2.11, panel

B). Smooth muscle -actin staining (Figure 2.11, Panel C) confirms that the increase in

cellularity reflects an increase in medial smooth muscle.

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87

Figure 2.11 (A) Immunohistochemical staining of incorporated BrdU in paraffin embedded

sections of aorta and mesenteric artery from normoxic and hypoxia exposed (48hrs) rats (20X

magnification). (B) Double staining with TO-PRO-3 and (C) -Smooth Muscle Actin in

paraffin embedded sections of normoxic and hypoxic (48hrs) rat aorta and mesenteric artery

shows increased cellularity and medial smooth muscle cell density after hypoxia (40X

magnification).

Aorta Mesenteric artery

Normoxic Normoxic Hypoxic Hypoxic

A

B

C

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88

Figure 2.11 (D) Quantitative analysis of incorporated BrdU and TO-PRO-3 staining

confirms increased proliferation (top) and nuclear density (bottom).

Nu

cle

ar

de

nsity

(Nu

mb

er

of

nu

cle

i p

er μ

m2 x

10

-4)

Chapter 2

89

2.4 Discussion

We found that in HASMCs in culture: 1) incubation at 3% O2 enhances, whereas 1% O2

inhibits DNA synthesis and the time-dependent increase in cell number; 2) incubation at 1% O2

is associated with accumulation of cells in G1 phase of the cell cycle whereas 3% O2 increased

the percentage of cells in S and G2/M; 3) the prevalence of apoptosis is increased after

incubation at 1% O2; 4) cellular ATP levels are reduced and the mitochondrial membrane is

depolarized after exposure to 1% but not 3% O2; and 5) pro-and antiproliferative and pro- and

antiapoptotic gene expression are tightly coordinated to effect directionally opposed responses

within a narrow range of oxygen concentrations. In aorta and mesenteric arteries of rats

breathing 10% O2 (arterial PO2 40 mmHg) both smooth muscle cell proliferation and apoptosis

are increased.

Hypoxia has been reported to enhance proliferation in pulmonary artery and aortic

smooth muscle cells [10, 258, 260, 261, 273-276]. In apparent contradiction, others have

observed that hypoxic stress inhibits growth of these same cells [142, 147, 263]. The results of

the current study show that cell numbers, [3H]-Thymidine incorporation, biochemical markers of

proliferation (Ki67[277], CDC6[267] and MCM2[268] protein levels) and the response to

PDGF-BB are increased and decreased after incubation at 3% O2 and 1% O2, respectively. We

conclude that hypoxia may either enhance or inhibit HASMC proliferation depending on its

severity. Rather than being due to experimental error or differences in cell origin, the

discrepancy in previous observations, therefore, reflects fundamental differences in the nature of

the cellular response elicited by varying degrees of hypoxic stress. The effect persists during and

is additive to the effects of extrinsic stimulation with PDGF, suggesting that the responses are

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90

mediated by nonconvergent pathways. Total cell number, however, reflects effects on both

proliferation and cell death, and differential modulation of these processes by hypoxia and/or

PDGF could account for these findings without inferring independent signaling mechanisms.

Following incubation at 1% O2 we found that the percentage of cells in G1 phase of the

cell cycle is increased, with a corresponding depletion of cells in G2/M and S phases indicating a

delay in progression through the G1/S interphase. In contrast, incubation at 3% O2 results in a

decrease in the number of cells in G1 phase, an increase DNA synthesis and in the percentage of

mitotic cells, consistent with acceleration of the G1/S transition. The most frequently reported

effect of hypoxia is delayed entry into S phase [142, 147, 148, 262-264, 278] as we observed in

cells incubated at 1% O2. Our finding that enhanced proliferation in cells incubated at 3% O2 is

also associated with an alteration in the rate of G1/S transition, albeit opposite in direction, is

novel. It suggests that the bidirectional effects of hypoxia are integrated by events occurring at

this checkpoint.

During transition from G1 to S phase cyclin-dependent kinases (cdks) phosphorylate the

retinoblastoma protein (Rb) displacing the E2F-1 transcription factor and activating expression

of S phase genes required for DNA synthesis [145]. Activity of the cdks is dependent on

association with their respective cyclins and regulated by endogenous cdk inhibitors (eg. p21 and

p27). As shown in Table 2.1, genes whose products act at the G1/S transition are differentially

regulated under the conditions studied. Both p53 and its transcriptional activation target p21,

which inhibit G1/S transition, are increased after incubation at 1% O2 and unchanged at 3% O2.

Moreover, genes with known roles in modulating p53/p21 activity demonstrate patterns

consistent with their expected functional effects: Expression levels of replication protein A1,

which binds p53 preventing activation of p21 transcription [279]; components of the

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91

extracellular signal regulated kinase cascade, which stimulate assembly of the cyclinD–CDK4/6

complex promoting G1/S progression and p27 degradation [280]; are enhanced after incubation

at 3% O2 and inhibited following incubation 1% O2. Replication factor C (activator 1) 1[281],

CDC6 [267] and MCM2 [268] are similarly affected. Conversely, expression of antiproliferative

genes, Inhibitor of Growth Family, member 1 like, which binds and enhances p53 activity [282],

Bone Marrow Morphogenetic Protein 2, which induces SMAD-mediated expression of p21 and

p27 [283] and down regulates antiapoptotic Bcl-xL expression [284], and the dual specificity

phosphatases 1 and 5, which deactivate ERKs [285], show the opposite pattern. Corresponding

changes in Tuberous Sclerosis Complex-1, a negative regulator of mRNA translation [286]

(Table 2.1), suggest a mechanism by which the effects of hypoxia on protein and DNA

replication may be coordinated.

The results of three independent assays (Annexin V/PI, TUNEL and mitochondrial

membrane depolarization) and the changes in pro- and antiapoptotic gene expression (Table 2.1)

in the present study indicate that the rate of apoptosis is increased in HASMCs incubated at 1%

O2. The reduced cell number observed under this condition therefore reflects, in part, an

enhanced rate of cell death. In fibroblasts and tumour cells, hypoxia of sufficient severity to

cause ATP depletion impairs DNA repair and the increase in apoptosis in this setting is

considered a protective mechanism to prevent the accumulation of hypoxia-induced mutations

[201, 203, 287]. The increased rate of smooth muscle cell apoptosis during hypoxic epochs

associated with reduced [ATP] i in the current study may serve a similar adaptive purpose. That

hypoxia induces apoptosis at oxygen concentrations recorded in arteries affected by aneurismal

dilatation and atherosclerotic plaques, [288] and that hypoxemia, in vivo, increases smooth

muscle apoptosis in the walls of systemic arteries supports the suggestion that it may play a

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92

pathogenic role in arterial smooth muscle cell dysfunction [270] and loss in patients with

cardiopulmonary disease and shock.

Apoptosis may be triggered in response to stimuli extrinsic or intrinsic to the affected

cell. Ligand binding to the TNF receptor superfamily such as Fas, through their association with

the Fas associated death domain (FADD) protein, results in assembly of the death inducing

signaling complex (DISC) which recruits and activates caspase-8. The intrinsic pathway is

activated when mitochondrial membrane depolarization releases cytochrome C into the

cytoplasm where it binds apoptotic protease activating factor-1 (Apaf-1) allowing it to activate

pro-caspase-9. In the final common pathway caspase-9 (intrinsic) and caspase-8 (extrinsic)

cleave and activate the effector protease, caspase-3[289]. p53, through interactions with Apaf-1

[180] and protective Bcl proteins, directly activates caspase-3 [290] and enhances permeability

of the outer mitochondrial membrane [291]. Complementary regulation by death inhibitors and

the balance between prosurvival and proapoptotic Bcl-2 family members[292] superimposes an

additional level of control. Our finding (Table 1) that the expression of components of both the

extrinsic (Caspase-8 associated protein 2, Casp8 and FADD-like apoptosis regulator) and

intrinsic (caspase recruitment domain protein 7, apoptosis-inducing serine/threonine kinase 17a

and b, BMP2, Apaf-1, Bcl2l1 (BCL2-antagonist of cell death) and p53 [148]) pathways as well

as Caspase-3 are oxygen regulated, therefore, indicates that stimuli originating within the cellular

microenvironment as well as the intrinsic response are important in hypoxic activation of the

apoptotic program.

Resistance to apoptosis has been reported in lung fibroblasts, A7r5 cells, rat kidney

proximal tubular cells and rat pheochromocytoma PC12 cells during hypoxic incubation [207,

Chapter 2

93

293, 294]. In the current study the percentage of cells staining positive for Annexin V was

decreased and the expression of antiapoptotic genes (CASP8 and FADD-like apoptosis regulator

[295] and Bcl2l1, Bcl2–antagonist of cell death [296]) was increased following incubation at 3%

O2. Consistent with previous results in other cell types, therefore, hypoxia, at levels above those

causing outright cytotoxicity, enhances smooth muscle cell survival. In the systemic circulation,

protection from apoptotic cell death may be important in neovasoformation in response to

oxygen delivery/requirement imbalance during development and in ischemic tissues where

investiture of new endothelial networks by regulatory smooth muscle cells must occur under

conditions of relative oxygen deficiency.

The mitochondria are the site of oxidative phosphorylation, however, they must

themselves consume ATP [297] to maintain the trans-mitochondrial membrane potential on

which electron transport coupling depends [298]. Failure of efficient ATP synthesis and,

consequently, ion-motive ATPase activity ultimately results in depolarization of the membrane,

further ATP depletion, Ca2+

influx, phospholipase and protease activation and the release of

apoptotic factors [299-301]. Little information is available concerning the degree of hypoxia

required to initiate programmed cell death and it may vary among cell types [302]. Our present

results show that incubation of HASMCs at 1% but not 3% O2 causes mitochondrial membrane

depolarization, therefore, the threshold between normal physiological functioning and cell

destruction is exceptionally narrow in these cells, particularly when compared to the range of

oxygen concentrations to which they are normally exposed [303]. Conflicts among previous

reports are undoubtedly attributable, at least in part, to the lack of appreciation of this and the

experimental rigor needed to separate these responses.

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94

Nuclear levels of HIF-1 were elevated to a similar extent in HASMCs incubated at 3%

and 1% O2 whereas the effects on HASMC proliferation were directionally opposed. This

suggests that HIF-1 independent regulatory mechanisms predominate. It would be unusual,

however, if the primary oxygen sensing mechanism in mammalian cells were not involved in

such an important response. Moreover, many cell cycle associated genes contain functional

hypoxia regulatory elements and HIF-1-regulated pathways that both enhance cell survival [304,

305] and, conversely, increase apoptotic cell death [147, 306] have been identified. A more

complex role than can be accounted for by changes in HIF-1 levels must therefore be proposed

in order to reconcile these observations. Differences between mRNA expression of the HIF-1

regulated genes [87, 307] represented on the Affymetrix HG-U133A array in cells incubated

under the two conditions (see supplemental data, GEO Accession # GSE 4725) support this

notion. This is not surprising since HIF-1 is subject to extensive post-translational

modification prior to nuclear translocation [56] and interacts with a multitude of coregulatory

factors [147, 306, 308] offering many sites at which its function may be differentially affected by

the two conditions.

During the prenatal period, the systemic circulation undergoes continuous restructuring in

response to the changing requirements of the developing tissues. In the mature circulation,

hypoxemia, due to cardiopulmonary disease, elicits responses which redistribute blood flow and

enhance the capacity for oxygen extraction [309-311]. As the duration of hypoxia increases,

however, systemic vascular smooth muscle and endothelial cell function are impaired [270, 312],

limiting the efficacy of the acute responses. Concurrent structural remodeling thus plays an

increasing role in maintaining the balance between oxygen delivery and metabolic demand[311].

Our current results indicate that this is facilitated by increases in the rates of both smooth muscle

Chapter 2

95

cell proliferation and death, a paradoxical state that will markedly enhance cell turnover. During

remodeling, vascular cell replication and removal must be tightly controlled to ensure a degree of

plasticity sufficient to achieve the required structural change while avoiding the accumulation of

mutations and malignant transformation or the formation of abnormal structures that exacerbate

circulatory dysfunction. The results of the present study indicate that the difference between

oxygen concentrations that enhance smooth muscle cell proliferation and those that impair

cellular energy status and trigger cell destruction is correspondingly small and well within the

transmural and longitudinal gradients known to exist in the systemic circulation.

Although both proliferation and apoptosis are enhanced in aortae and mesenteric arteries

from hypoxia-exposed rats, the net effect is an increase in medial smooth muscle. Prolonged

hypoxia of this severity results in a progressive loss of systemic arterial and arteriolar

contractility [270, 313] with consequent impairment of the sympathetically-mediated reflexes

that regulate blood flow distribution [314]. In this context, increased muscularity of the arterial

wall can be viewed as a compensatory adaptation that preserves the capacity to regulate the

systemic circulation. Vital organ function is highly intolerant of oxygen deprivation.

Accordingly, mechanisms linking vascular cell turnover and the capacity for rapid structural

change directly to oxygen concentration are required to avoid delays inherent in second

messenger signaling. Our results indicate that pro- and antiproliferative and pro- and

antiapoptotic gene expression are tightly coordinated to produce directionally opposed responses

effected at the level of G1/S transition and involvement of both intrinsic and extrinsic apoptotic

pathways. Further definition of the individual roles of these regulatory mechanisms will be

valuable in identifying therapeutic targets in conditions in which enhanced plasticity of the

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96

vasculature may be exploited to alleviate tissue oxygen deficiency or in which over exuberant

remodeling interferes with normal cardiovascular function.

Chapter 3

97

Chapter 3

Oxygen regulation of pulmonary artery smooth muscle cell

proliferation and survival

Chapter 3

98

3.1 Introduction

Hypoxic pulmonary arterial hypertension (HPAH) contributes to morbidity and

premature mortality in patients with cardiopulmonary diseases and responds poorly to treatment.

[315-319]. Increased smooth muscle in the pulmonary arterial wall is a hallmark of this

condition and contributes to the increase in resistance to blood flow [315, 316, 320-323]. The

factors determining SMC proliferation and survival in the pulmonary circulation and, hence,

susceptibility to and severity of HPAH are poorly understood. Studies to determine the

mechanisms regulating these processes are, therefore, of high priority.

Smooth muscle cells of diverse origin proliferate in response to moderate levels of

hypoxia (2-3% O2) [258, 323]. Proliferation and viability of systemic arterial SMCs are reduced,

however, when hypoxia is sufficiently severe that the capacity to maintain intracellular ATP

levels is impaired [323]. Pulmonary artery smooth muscle cells (PASMCs) differ from those in

systemic arteries in several important respects. In addition to differences in their contractile

responses to hypoxia (contraction in PASMCs vs. relaxation in systemic vascular SMCs), and

vasoactive factors which may reflect differences in membrane ion channel expression [14, 324,

325], PASMCs are metabolically distinct. Mitochondria from these cells are relatively

depolarized, display lower expression of proximal ETC components and a greater propensity to

generate ROS at reduced oxygen tensions [326]. This could affect the response to hypoxic stress

and would be particularly apparent in highly energy dependent responses such as proliferation.

In vivo, hypoxia causes initial PASMC proliferation in rodents followed by a return to

quiescence with continued hypoxic exposure under the combined influences of endothelium-

derived mediators, circulating neurohumoral factors, and the direct effects of the hypoxic

microenvironment [327-330]. In cultured PASMCs from various species proliferation has been

Chapter 3

99

reported to be enhanced [276, 329, 331-338], inhibited [10, 276, 329, 331, 339-341] or unaltered

[331, 335, 340, 341] during exposure to hypoxia of varying severity and duration (Table 3.1).

This lack of consistency in experimental design among earlier studies has precluded the

development of consensus regarding the direct role of ambient oxygen concentration. It has been

equally difficult, for similar reasons, to establish the threshold at which hypoxia triggers cell

death in PASMCs and whether this differs from that in SMCs derived from other tissues,

although mechanisms that might confer protection from apoptosis in PASMCs have been

proposed [284, 342-345]. The current study was, therefore, carried out to determine if: 1) the

effects of hypoxic incubation on human PASMC proliferation and survival differ from those in

SMCs from the systemic circulation; 2) these responses differ under conditions which do or do

not result in cellular ATP depletion; and 3) these effects are relevant to pulmonary vascular

remodeling during hypoxia in vivo.

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100

Table 3.1 Influence of hypoxia on PASMC proliferation.

First author

(Reference) Species O2 %

Duration of

hypoxia

(days)

Proliferation PASMC

location

Seeding

density

(cells/cm2)

FBS %

Yang (329)

Human

(bronchial

carcinoma)

0 2, 4, 6 Distal PA 10000 10

Eddahibi (339) Rat 0 1 Proximal PA 25000 0.2

Hassoun (340) 0 Proximal PA 5000 10

3,10 Proximal PA 5000 10

Cooper (276) Human 0 4 Not specified 5000 2

5 5000 1, 2, 5

Rose (10) Human,

rabbit

1 1 Not specified 4000 Serum free

Stotz (338) Rat 1 2 Microvascular 4000 5

Lu (336) Rat 2 1,2 Not specified 2500 5

Frid (331) Cow 3 2 Lower media, outer

media rounded

epitheloid cells

~ 10000 10

3 2 All medial cells ~ 10000 0.1

3 2 Middle media, outer

media spindle shaped

cells

~ 10000 10

Lanner (335) Pig 3 2, 3, 4 Main branch of PA 5200 5

3 Main branch of PA 5000 0.1

Tamm (350) Human

(peripheral

lung cancer)

3 Not specified 5

Preston (337) Rat 3 3-10 Lobar PA 2000-4000 10, 0.1

Dempsey (273) Cow 3 2, 3, 4 Not specified 25,000-

50000

0.1

Stiebellehner Cow 3 4 Distal PA 25000 10

(341) 3 Distal PA 25000 0.1

Ambalavalan

(332)

Pig 1, 2, 3, 5, 7, 10 3

(1, 3, 5, 7)

Not specified 10000 10

Benitz (333) Cow 3,6, 9 10 Not specified 2500 10

PASMC: Pulmonary artery smooth muscle cell, FBS: Foetal bovine serum (concentration in cell culture media),

: Increase in PASMC proliferation, : Decrease in PASMC proliferation, : No change in PASMC proliferation.

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101

3.2 Materials and methods

Antibodies and Reagents

Hypoxia inducible factor 1- (HIF-1 ) antibody was purchased from Novus Biologicals

(Littleton, CO), p21 antibody from BD Pharmingen (San Diego, CA) and p53 antibody from Cell

Signaling Technology (Danvers, MA). CaspACE FITC-VAD-fmk in situ marker and TUNEL

kits were both from Promega (Madison, WI). 5,5’,6,6’-Tetrachloro-1,1’,3,3’-tetraethyl-

benzimidazolylcarbocyanine iodide (JC-1) labeling kit, ATP bioluminescence assay kit, and TO-

PRO-3 dye were purchased from Molecular Probes (Carlsbad, CA). All other reagents were

from Sigma (St. Louis, MO).

Cell Culture Studies:

Human pulmonary artery smooth muscle cells (HPASMCs, Cambrex Bio Science

Walkersville, MD, USA) were propagated to passage 6 in SMGM-2 medium (Cambrex)

consisting of SmBM medium supplemented with single aliquots of 0.1% insulin, 0.2% hFGF-B,

0.1% GA-1000 (Gentamicin and Amphotericin B) and 5% v/v FBS, 0.1% hEGF. Upon reaching

70% confluence the media was changed and cells were incubated for a further 16 or 48 hrs under

either normoxic or hypoxic conditions. Cells exposed to hypoxia were placed in a humidified

Plexiglas chamber (Billups Rothberg, San Diego, CA, USA) maintained at 37°C and

continuously flushed with gas mixtures containing 10%, 5%, 3%, 1% or 0% O2, 5% CO2,

balance N2. Normoxic cells were exposed to air/5% CO2 under otherwise identical conditions.

The concentrations of dissolved O2 in the culture medium, measured using the ISO2 dissolved

oxygen meter (World Precision Instruments, Sarasota, FL, USA) were 20.5 ± 0.6%, 3.1 ± 0.4%

O2, 1.2 ± 0.3% O2 and 0.3±0.2% O2 when the chamber was flushed with air/5% CO2, 3% O2, 1%

O2 and 0% O2 gas mixtures, respectively. Steady state oxygen concentrations were achieved

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102

within 40 minutes in each case. Experiments were repeated three times using cells from at least 3

human donors with 6 replicates per observation.

Cell Counting: After exposure to normoxia or hypoxia (10%, 5%, 3%, 1% or 0% O2) for

16 or 48 hours, cells were washed twice with HBSS and detached with 0.25% trypsin and 0.02%

EDTA and cell number was determined using a Coulter counter (Beckman Coulter, Inc,

Fullerton, CA). Cell viability was assessed by Trypan Blue exclusion.

BrdU incorporation/Cell cycle analysis: Bromodeoxyuridine (BrdU), an analog of the DNA

precursor thymidine is incorporated into newly synthesized DNA. Propidium iodide (PI) staining

was simultaneously done to measure total DNA. Cells were pulse-labeled by the addition of

BrdU to the culture medium to a final concentration of 10 μM during the last 16hrs or 48 hrs of

normoxic or hypoxic incubation. At the end of the incubation period, cells were harvested and

fixed for an hour in 70% ethanol followed by denaturation of DNA with 0.1 M HCl and

neutralization with 0.1M borate buffer. Cells were then incubated for 45 min in a solution of PI

(2.5 μg/ml) and RNase A (50 μg/ml) and with a fluorescein isothiocyanate-conjugated

monoclonal anti-BrdU antibody (diluted 1:1000, Pharmingen, San Diego, CA). The percentage

of cells staining positive for BrdU uptake was determined by flow cytometry (BD FACScan flow

cytometer, BD Biosciences, NJ) using CellQuest Software.

Annexin V-Propidium Iodide labeling: To assess the effect of hypoxic incubation on

apoptosis, the Roche (Basel, Switzerland) annexin V-fluorescence (Fluos) staining kit was used

to detect phosphatidylserine externalization (an early event in apoptosis), and PI uptake (a

marker of cell death) in HPASMC exposed to normoxia or hypoxia (3%, 1% or 0% O2) for 16 or

48 h. Cell suspension was centrifuged at 1,500 rpm for 10 min at 4°C. The pellet was

resuspended in 5 ml of cold PBS and centrifuged again. The supernatant was removed and the

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103

pellet suspended in 100 l of annexin V-Fluos labeling solution (20 l annexin V-Fluos labeling

reagent and 20 l Propidium Iodide (PI) solution per milliliter of incubation buffer) at 37°C.

Labeled cells were analyzed using flow cytometry, and the number of cells positive for Annexin

V, PI, or both were calculated.

Caspase activation: Caspase activation was detected in HPASMCs exposed to normoxia or

hypoxia (3%, 1% or 0% O2) for 16 or 48 hours. CaspACE FITC-VAD-fmk is a FITC conjugate

of the cell permeable inhibitor of caspases. This structure allows delivery of the inhibitor into the

cell where binding to activated caspase, serves as an in situ marker for apoptosis. About 2 106

cells were incubated with 100 μM FITC-VAD-FMK at room temperature in the dark for 20 min.

Cells were then washed, resuspended in PBS and the percentage of cells positive for activated

caspase quantified by flow cytometry.

Mitochondrial membrane potential: Depolarization of the mitochondrial membrane, was

detected using the potential-sensitive probe JC-1. JC-1 monomers enter the mitochondria at

physiological membrane potentials (62), where, as a result of aggregation, the emitted

wavelength changes from 530 nm (green) to 590 nm (orange) when excited at 490 nm.

Disaggregation to the monomeric form during mitochondrial membrane depolarization is

detected as an increase in green emission. HPASMCs exposed to normoxia or hypoxia (3%, 1%

or 0% O2) for 16 or 48 h were incubated with JC-1 (10 g/ml) for 20 min in the dark, washed,

and resuspended in 1 ml PBS. The % cells positive for JC-1 monomer was assessed by flow

cytometry.

Intracellular ATP concentration: HPASMCs exposed to normoxia or hypoxia (3%, 1% or

0% O2) for 16 or 48 h were washed twice with ice-cold PBS and lysed by adding equal volumes

of 3.6% perchloric acid. Samples were centrifuged, and ATP concentrations in the supernatants

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104

were determined by using an ATP bioluminescence assay (Molecular Probes), according to the

instructions provided by the manufacturer. The photometer (Multiskan EX Microplate

Photometer, Thermo Labsystems, Philadelphia, PA) was set for a 5-s delay period and a 5-s

integration period. ATP levels were calculated by using standard reference solutions corrected

for background luminescence.

Western blot analysis: Western blot analysis was used to quantify levels of HIF-1 and

the cell-cycle regulatory proteins p21 and p53 in nuclear extracts from HPASMCs exposed to

normoxia or hypoxia (3%, 1% or 0% O2) for 16 or 48 h. Cells were lysed in buffer A [10 mM

HEPES (pH 7.8), 10 mM KCl, 0.1 mM EDTA, 1mM dithiothreitol (DTT), and 0.1% Nonidet P-

40 (NP-40)] with protease inhibitors (5 g/ml aprotinin, 5 g/ml pepstatin, 5 g/ml leupeptin, 0.5

mM Pefabloc, and 1 mM phenylmethylsulfonyl fluoride) and phosphatase inhibitors (10 mM

sodium fluoride, 1 mM sodium orthovanadate, and 20 mM glycerophosphate). Nuclear proteins

were then extracted with buffer B [50 mM HEPES (pH 7.8), 420 mM KCl, 0.1 mM EDTA, 1

mM DTT, 5 mM MgCl2, and 20% glycerol], containing both protease and phosphatase

inhibitors. Equal amounts of protein extracted from HPASMCs, incubated under normoxic and

hypoxic (3%, 1% and 0% O2) conditions for 16 and 48 h, were loaded on 4–12% Tris-glycine

gels, separated by electrophoresis and transferred to nitrocellulose. Membranes were blocked

with 5% milk overnight and probed with anti-HIF-1 (1:500), anti-p21 (1:3,000), anti-p53

(1:1,000). In all cases, protein concentration was determined by the Bradford assay, and

appropriate volumes of extraction buffer to produce constant protein loading in each lane were

mixed with SDS loading buffer. Equality of protein loading and transfer efficiency were

corroborated by full-lane densitometry of the Ponceau red-stained membranes. Immunoblots

were probed with horseradish peroxidase-donkey anti-rabbit IgG (1:1,000 in blocking buffer)

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and visualized by enhanced chemiluminescence (ECL Plus kit, Amersham Biosciences). Band

intensity was quantified by densitometry (Bio-Rad Laboratories, Mississauga, ON, Canada).

Animal Studies:

The effects of hypoxic exposure on pulmonary vascular smooth muscle cell proliferation and

apoptosis in vivo were assessed in male Sprague-Dawley rats (175–200 g). All protocols were in

compliance with standards set by the Canadian Council of Animal Care, and were approved by

the Institutional Animal Care Committee. Rats were placed in a Plexiglas chamber into which

the flow of air and nitrogen was controlled independently. Rats exposed to hypoxia breathed a

gas mixture containing 10% O2 for 2 days, 7 days and 14 days. Normoxic control animals

breathed room air under otherwise identical conditions. In preliminary experiments, the arterial

PO2 in rats breathing 10% O2 averaged 38 Torr (range, 35–42 Torr). At the end of the exposure

period right and left main and first branch pulmonary arteries were excised immediately after

decapitation, rinsed with PBS, fixed with paraformaldehyde, dehydrated in graded ethanol (70–

100% ), cleared in xylene, and embedded in paraffin. Sections (5 m thick) were cut on an

oscillating blade microtome (Leica, Wetzlar, Germany) and placed on coated glass microscope

slides (Fisher Scientific, Pittsburgh, PA).

Detection of apoptotic cells: DNA fragmentation was detected by TUNEL (DeadEnd

Fluorometric TUNEL System, Roche). Slides containing paraffin-embedded sections were

dewaxed, rehydrated, permeabilized with proteinase K, preincubated with equilibration buffer,

and incubated with labeling solution (rTdT and nucleotide mixed with fluorescein-labeled dUTP)

for 1 h at 37°C. The reaction was terminated by incubating samples in a stopping buffer for 30

min. After PBS washes and counterstaining with TO-PRO-3, the samples were mounted and

examined by laser confocal microscopy.

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Detection of proliferating cells: To assess the effects of hypoxia on pulmonary artery

smooth muscle cell proliferation in vivo, bromodeoxyuridine (BrdU) uptake by these cells was

assessed as a marker of de novo DNA synthesis. BrdU was infused subcutaneously using

osmotic pumps (model no. 2ML-2; Alza Corp, Palo Alto, CA). Pumps containing 0.32 g of BrdU

in 2 ml vehicle (0.4% DMSO) were implanted intrascapularly in normoxic and hypoxia-exposed

rats 2 days, 7 days and 14 days before euthanasia. The rats received 0.4 mg BrdU/h delivered

continuously. At the end of the labeling period, the right and left main and first branch

pulmonary arteries were excised, fixed and embedded in paraffin. Dewaxed and rehydrated

slides were incubated with 2 M HCl for 1 h at 37°C. The acid was neutralized by 0.1 M borate

buffer, pH 8.5. Following PBS washes, the slides were incubated with fluorescein-labeled anti-

BrdU antibody (Roche) for 1 h at room temperature, protected from light. Slides were then

washed with PBS and mounted for confocal microscopy.

Smooth muscle -actin staining: Immunohistochemical staining using anti alpha-smooth

muscle actin antibody was carried out on the same slides to confirm localization of TUNEL and

BrdU uptake to smooth muscle. Following TUNEL or BrdU labeling, sections were covered with

blocking buffer (5% goat serum with PBS containing 1% BSA) for 30 minutes, following which

they were stained with Cy3-conjugated monoclonal anti-smooth muscle -actin (1:200 in

blocking buffer, Sigma) for 2 hours. Sections were counterstained with TO-PRO-3, washed with

PBS, and viewed with a confocal microscope.

Quantitative analysis: The number of TUNEL-positive, BrdU- and TO-PRO-3-labeled

cells was determined in three sections from each of six animals from each experimental group.

Four to six randomly selected microscopic fields ranging from 90–300 cells per field were

counted for each section, and the number of TUNEL or BrdU-positive nuclei was expressed as a

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percentage of the total number of nuclei. The diameter of a vessel (distance between external

lamina) and the thickness of the tunica media (distance between the internal and external elastic

lamina) were measured in main and first branch pulmonary artery sections stained for smooth

muscle -actin, from digitized images using Image J analysis software (NIH 2002,

http://rsb.info.nih.gov/ij). Thickness was measured at equally spaced 10 points around the vessel

wall and the percentage of medial wall thickness expressed as:

% medial wall thickness = ([medial thickness x 2]/external diameter) x 100

Data from each animal in each group were averaged to serve as a single value for statistical

analysis.

Data Analysis: Data are presented as means ± SEM of n observations with P< 0.05

considered significant. The significance of differences between individual means was determined

by two-tailed Student’s t-test. Differences among multiple means were evaluated by analysis of

variance (ANOVA) corrected for multiple measures, where appropriate, and, when overall

differences were detected, differences between individual means were evaluated post hoc using

the Student-Newman-Keuls procedure.

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3.3 Results

Figure 3.1A illustrates the effect of normoxic and hypoxic incubation on human

pulmonary artery smooth muscle cell (HPASMC) number. After incubation at 5% O2 for 16 or

48 hours, there was a trend toward increased cell numbers which did not reach statistical

significance. After incubation at 3% O2 for 16 or 48 hours, cell number was significantly

increased, compared to the normoxic control condition. After incubation at 1% O2 for 16 hours

cell number was increased compared to the normoxic control value, whereas after 48hrs cell

number was reduced. Incubation at 0% O2 inhibited the increase in cell number at both 16 and

48hr. time points. As shown in Figure 3.1B, HPASMC viability (Trypan Blue exclusion) was

not significantly affected by incubation at 10, 5, 3 or 1% O2. Viability was decreased after

incubation at 0% O2 for 16 and 48 hours.

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Figure 3.1 (A) Effects of hypoxia on human pulmonary artery smooth muscle cell

(HPASMC) numbers after incubation for 16 and 48 h. n= 6 per condition. *P <0.05 vs.

corresponding normoxic control values.

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Figure 3.1 (B) Effects of hypoxia on human pulmonary artery smooth muscle cell

(HPASMC) viability after incubation for 16 and 48 h. n= 6 per condition. *P <0.05 vs.

corresponding normoxic control values.

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To confirm that PASMC proliferation was altered by hypoxic incubation the percentage

of cells that incorporated BrdU after incubation at 3%, 1% and 0% O2 for 16 and 48 hrs was

assessed. As shown in Figure 3.2, the percentage of BrdU positive cells is increased after

incubation at 3% O2 for both 16 and 48 hours and at 1% O2 for 16 hours. BrdU incorporation is

decreased after incubation at 1% O2 for 48 hours and at 0% O2 for both16 and 48 hours.

Figure 3.2 The % BrdU incorporated cells increased after incubation at 3% O2

for 16 and

48 h, and after incubation at 1% O2 for 16 h. BrdU incorporation decreased after incubation at

1% O2 for 48 h and after incubation at 0% O2

for 16 and 48 h. n= 6 per condition. *P <0.05

vs. corresponding normoxic control values.

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Since the effects of hypoxic incubation on PASMC cell numbers may also reflect

changes in the rate of cell death, the percentage of PASMCs staining positive for Annexin V,

Propidium iodide, activated caspases and JC-1 monomers was recorded after normoxic and

hypoxic incubation and are presented in Figures 3.3 A-E respectively. The percentage of viable

cells (staining negative for both Annexin V and PI) was unaffected by incubation at 3% and 1%

O2 but decreased significantly after incubation at 0% O2. Similarly, the percentage of cells

positive for activated caspases and for JC-1 monomers are unchanged after incubation at 3% or

1% O2 but increased after incubation at 0% O2 for 16 h and increased further after 48h.

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Pro

pid

ium

io

did

e

Pro

pid

ium

io

did

e

0% O2, 16hrs

Annexin V

Figure 3.3 (A) Representative flow cytometric plots for Annexin V/PI staining after

incubation at 0% O2 , 16 and 48 h, compared with the normoxic cells; n= 6. *P<0.05 vs.

corresponding normoxic control values.

21% O2, 16hrs

21% O2, 48hrs 0% O2, 48hrs

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Figure 3.3 (B) Bar graphs representing reduced number of Annexin V and PI negative cells

(live cells) after incubation at 3%, 1% and 0% O2 , 16 and 48 hours, compared with the normoxic

cells (open bars); n= 6. *P<0.05 vs. corresponding normoxic control values.

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Figure 3.3 (C) Representative flow cytometric plots for increased caspase activity after

incubation at 0% O2 , 16 and 48 hours, compared with the normoxic cells (open bars); n= 6.

*P<0.05 vs. corresponding normoxic control values.

0% O2, 16hrs

FITC-Caspase

21% O2, 16hrs

21% O2, 48hrs 0% O2, 48hrs

Counts

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D

E

Figure 3.3 (D) Increased caspase activity and (E) increased 5,5’,6,6’-tetrachloro-

1,1’,3,3’-tetraethyl-benzimidazolylcarbocyanine iodide (JC-1) monomer formation

(indicating mitochondrial membrane depolarization) after incubation at 0% O2 compared

with the normoxic cells (open bars); n=6. * P<0.05 vs. corresponding normoxic control

values.

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To determine if hypoxia exerts a general effect on cell cycle progression or affects a

specific cell cycle phase, DNA content was quantified by PI staining to assess the percentage of

cells in G0/G1, S and G2/M phases. As illustrated in figure 3.4, incubation of HPASMCs at 3%

O2 (16 and 48hrs) decreased the percentage of cells in G0/G1 phase and increased the percentage

of S-phase cells with a corresponding increase in the percentage of cells undergoing mitosis.

This suggests acceleration of progression through the G1/S interphase. A similar distribution is

observed after incubation at 1% O2 for 16 hrs. After incubation at 1% O2 for 48hrs and at 0% O2

for 16 and 48hrs, however, there is accumulation of G0/G1phase cells and depletion of cells in S

and G2/M phases, indicating a delay in transition through the G1/S interphase.

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Figure 3.4 Flow cytometric analysis of propidium iodide stained normoxic (N) and

hypoxic (3%, 1% or 0% O2) cells. Incubation at 3% O2 16 and 48 h and 1% O2

16 h

increases the percentage of cells in G2/M, whereas incubation at 1% O2 48 h and 0% O

2 16

and 48 h increases the percentage of cells at the G0/G1 interphase. n = 6, * P<0.05 vs.

corresponding normoxic control values.

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p53 and its transcriptional target p21 are inhibitory regulators of cell cycle progression

and are involved in hypoxia induced growth arrest in cancer cells [144]. Expression of these

nuclear proteins were therefore, used as biochemical indices of the proliferative response. HIF-

1 nuclear translocation reflects activation of the cellular response to hypoxia and its nuclear

levels were measured to confirm that the cells were responding, at, a molecular level, to the

hypoxic stimulus. Levels of both p21 (Figure 5A) and p53 (Figure 5B) are increased after

incubation at 0% O2 but did not differ significantly from the normoxic control values after

incubation at 3% O2 or 1% O2. Nuclear HIF-1 levels (Figure 5C) increased from the normoxic

values after incubation at 3%, 1% and 0% O2.

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Figure 3.5 (A) Nuclear levels of p21 protein after incubation of HPASMCs under

normoxic and hypoxic (3, 1 or 0% O2) conditions. n= 6. *P <0.05 vs. corresponding

normoxic control values.

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Figure 3.5 (B) Nuclear levels of p53 protein after incubation of HPASMCs under

normoxic and hypoxic (3, 1 or 0% O2) conditions. n= 6. *P <0.05 vs. corresponding

normoxic control values.

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Figure 3.5 (C) Nuclear levels of HIF-1 protein after incubation of HPASMCs under

normoxic and hypoxic (3, 1 or 0% O2) conditions. n= 6. *P <0.05 vs. corresponding

normoxic control values.

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As shown in Figure 3.6, intracellular ATP concentration remained unchanged in human

pulmonary artery smooth muscle cells incubated at 3% O2. After 48 hours of incubation at 1%

O2, ATP levels decreased (7.6 ± 2.8 % decrease from normoxic control value), however this

change did not reach statistical significance (p = 0.096). In cells incubated at 0% O2 cellular

ATP concentration was reduced after 16 h and decreased further after 48 h (17.38 ± 4.5% and

22.5 ± 3.8% decrease vs. corresponding normoxic control values, respectively, p < 0.05 for

both).

Figure 3.6 Cellular ATP concentration is decreased after incubation at 0% O2

compared with the normoxic cells (open bars). n= 6. *P<0.05 vs. corresponding normoxic

control values.

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The effects of hypoxic exposure on pulmonary artery smooth muscle cell apoptosis and

proliferation in rats was assessed to evaluate the effects of physiologically relevant levels of

hypoxia on the pulmonary vasculature in vivo. TUNEL staining of pulmonary artery sections

(Figure 3.7, A and B) was carried out after 2, 7 and 14 days of hypoxic exposure (10% FiO2). In

the main pulmonary artery and first branch pulmonary artery the apoptotic index was

significantly increased after 7 days of hypoxic exposure and returned to normoxic control values

after 14 days. Figure 3.8, A and B illustrate the effects of in vivo hypoxia for 2, 7 and 14 days on

thickness of the pulmonary artery smooth muscle layer and BrdU incorporation. There is

maximum BrdU incorporation after 7 days with a subsequent decrease in the number of BrdU

staining cells to normoxic levels after 14 days, mirroring the effects of apoptosis. As illustrated

in Table 3.2, medial wall thickness was increased after 7 days and remained at this level after 14

days of continuous hypoxic exposure.

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Normoxic Hypoxic

Figure 3.7 (A) Representative images of TUNEL in paraffin-embedded sections of

normoxic and hypoxic (2 days, 7 days, 14 days) rat pulmonary artery and pulmonary artery

branch (x40 magnification).

Pu

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nar

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y

(Mai

n)

Pu

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y a

rter

y

bra

nch

2 days 7 days 14 days

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Figure 3.7 (B) Quantitative analysis of TUNEL positive cells; n=6 rats per group. *P<0.05

vs. corresponding normoxic control values.

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127

Normoxic Hypoxic

Figure 3.8 (A) Representative images of immunohistochemical staining of incorporated

BrdU (green) and -smooth muscle actin (red) in paraffin-embedded sections of pulmonary

artery and pulmonary artery branch from normoxic and hypoxia (2 days, 7 days, 14 days)

exposed rats (x40 magnification).

2 days 7 days 14 days

Pu

lmo

nar

y a

rter

y

(Mai

n)

Pu

lmo

nar

y a

rter

y

bra

nch

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Table 3.1 Medial wall thickness (% ) of pulmonary artery and pulmonary artery branch

from normoxic and hypoxia (2 days, 7 days, 14 days) exposed rats; n=6 rats per group. *P<0.05

vs. corresponding normoxic control values.

Main Pulmonary artery Branch pulmonary artery

Normoxic 5.8±0.04 4.9±0.03

2 day hypoxic 6.1±0.03 5.1±0.06

7 day hypoxic 10.4±0.9* 9.4±0.8*

14 day hypoxic 11.1±0.1* 9.9±0.5*

Figure 3.8 (B) Quantitative analysis of BrdU positive cells; n=6 rats per group. *P<0.05

vs. corresponding normoxic control values.

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3.4 Discussion

The main findings of this study are that: in HPASMCs in culture 1) incubation at 3% O2

for 16 and 48hrs and at 1% O2 for 16hrs increases cell number DNA synthesis and the

percentage of cells in S and G2/M phases of the cell cycle; 2) proliferation and DNA synthesis

are inhibited after incubation at 1% O2 for 48hrs and at 0% O2 for 16 and 48hrs with a

corresponding accumulation of cells in G1 phase; 3) the prevalence of apoptosis and nuclear

levels of the antiproliferative, proapoptotic factors p21 and p53 are unaffected by incubation at

3% and 1% O2 but are increased after incubation at 0% O2; and 4) cellular ATP levels are

reduced, and the mitochondrial membrane is depolarized after incubation at 0% O2. In the

pulmonary artery of rats breathing 10% O2 (arterial pO2 40 Torr), smooth muscle cell

proliferation and apoptosis are increased during the first seven days of hypoxic exposure with the

net effect being accumulation of smooth muscle in the vessel wall.

A hallmark of hypoxic pulmonary hypertension is medial thickening in larger pulmonary

arteries and muscularization of distal arteries not normally invested with smooth muscle [346,

347]. Hypoxia has long been known to stimulate PASMC proliferation in vivo although in most

models, including that used in the current study, this is a transient effect being maximal after 7

days and then subsiding [348]. The mechanisms, by which the proliferative response is activated

and, subsequently, regulated, are unresolved [349].

Hypoxia affects PASMC proliferation even in the absence of other cell types; therefore,

although mitogens and cytokines from surrounding cells may modulate the response in vivo, both

the sensory and effector mechanisms that underlie these effects are localized to the smooth

muscle cell [323]. Studies of the influence of hypoxia on PASMC proliferation in vitro have

yielded contradictory results [273, 276, 329, 331-338, 340, 341, 350]. Consequently, it remains

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130

unclear whether hypoxia has a direct mitogenic effect on PASMCs or acts as a comitogen acting

in concert with hypoxia-induced release of proproliferative factors from adjacent endothelial

cells or fibroblasts. Studies in which a positive correlation between acute hypoxia and PASMC

proliferation was observed tended to use moderate levels of hypoxia (3–5% O2) [276, 332-334,

336, 338, 350], whereas studies in which hypoxia caused a decrease in PASMC proliferation

were carried out under conditions of severe hypoxia or anoxia (< 2% O2) [329, 339, 340]. For

example, Ambalavan et al. showed

that swine proximal PASMC proliferated at oxygen

concentrations of

5–10% [273, 332], whereas, Stotz et al.

showed a 5–10% increase in

proliferation rates of rat pulmonary microvascular SMC at 1% O2 [338, 341]. Our current

findings reconcile these discrepancies since they demonstrate that, rather than being due to

variations in experimental conditions or species, hypoxia of differing severity may exert

opposing effects depending on the capacity of these cells to maintain cellular energy status under

the specific conditions studied.

Previous studies have shown hypoxia induced G1 arrest in cell cycle of HPASMCs to be

mediated by CDK inhibitor p21. In contrast, others have reported that cell cycle arrest at late G1

is caused by p27 expression under severe hypoxia [142, 351-353]. Li et al found that the oxygen-

dependent checkpoint of the cell cycle is controlled by p27 expression, and that cAMP signaling

also interferes with the cell cycle and p27 expression [354]. However, the precise mechanisms

and interactions between the pathways activated by hypoxia, as well as the antiproliferative

effects of p27 or p21 during hypoxic exposure in HPASMCs remain uncertain [355]. Our finding

that HPASMC incubation at 1% O2 for 48hrs and 0% O2 for 16 and 48hrs, causes G1/S arrest is

in line with those of others [258]. In addition, we also show that the directionally opposed effects

of hypoxia on HPASMCs, proliferation or growth arrest, are integrated by events at G1/S.

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131

Proliferation of HPASMCs requires these normally quiescent cells to enter the cell cycle. The

most important molecular process for cell cycle progression is retinoblastoma protein

phosphorylation by cyclin-dependent kinase (CDK)-cyclin complexes, and CDK activities are

mainly regulated by CDK inhibitors such as p21 and p27 [356].

In HPASMCs, both p53 and its transcriptional activation target p21, which inhibits G1/S

transition, are increased after incubation at 1% O2 48hrs and at 0% O2, but remain unchanged at

3% O2 and at 1% O2 16hrs. HIF levels are also increased in HPASMCs post incubation at 3, 1

and 0% O2, although the cell number increased at 1% O2 16 hrs and cell number decreased at 1%

O2 48hrs. This indicates HIF-1 -independent regulatory mechanisms causing the directionally

opposed proliferative responses of hypoxia. HIF-1 is the major transcription factor induced

under hypoxia, and is known to regulate pathways that both enhance cell survival as well as

induce cell death. Many cell cycle-associated genes containing functional hypoxia regulatory

elements have been identified [304-306]. Considering these, it is unlikely that there is no HIF-1

involvement at all in affecting the proliferative responses of HASMC or HPASMC. Further

studies are needed to understand the possible interaction of HIF-1 with other coregulatory

molecules.

Apoptosis plays an important role in cell number control in various tissues

and organs by

balancing cell growth and multiplication. The remodeling in pulmonary vascular structure is

mainly caused by imbalanced proliferation and apoptosis in pulmonary artery smooth muscle

cells [357-359]. An increase in PASMC proliferation and a decrease in PASMC apoptosis could

concurrently mediate thickening of the pulmonary vasculature, which subsequently reduces the

lumen diameter of pulmonary arteries, increasing pulmonary vascular resistance. It has been

demonstrated that increased PASMC proliferation and/or inhibited PASMC apoptosis both

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132

contribute to induce pulmonary vascular medial thickening [360, 361], and this acquisition of

resistance to apoptosis along with increased rates of VSMC proliferation appear to be necessary

for neointima formation [362-365]. The role of p53 in mediating VSMC apoptosis has been

proposed [366-368], though p53 alone does not induce apoptosis in either normal human or rat

VSMCs in vitro or in vivo unless the cells are primed to die [369-371], or massive expression is

induced via adenovirus vectors [372]. The precise mechanisms

involved in the regulation of

PASMC proliferation and apoptosis in PAH are still incompletely understood.

The precise level

of hypoxia that induces apoptosis in HPASMCs, has also not been defined.

The results of our apoptosis assays (Figure 3.3 A-E) show hypoxia induced cell death in

HPASMCs only after incubation at 0% O2. Viability is maintained at 3% O2 and at 1% O2.

Although the cell number is reduced in HPASMCs after incubation at 1% O2 48hrs, there is no

change in apoptosis compared to normoxic condition. Together these results suggest that hypoxia

in HPASMCs enhances smooth muscle cell survival, at levels above those that cause

cytotoxicity. In the pulmonary circulation, this protection from apoptotic cell death and enhanced

smooth muscle cell proliferation at moderate levels of hypoxia may be important in

neovasoformation during development and in ischemic tissues, where growth of new endothelial

networks by regulatory smooth muscle cells must occur under conditions of relative oxygen

deficiency. Depolarization of the mitochondrial membrane only at 0% O2 where apoptosis is

increased suggests the involvement of the mitochondria mediated intrinsic pathway of apoptosis.

Besides, the enhanced expression of p53 (at 0% O2) could also play a role in inducing PASMC

apoptosis.

When the ratio of energy supply to energy demand decreases, as during severe hypoxia,

homeostatic mechanisms attempt to match ATP production to ATP utilization [373, 374]. The

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133

hypothesis that changes in energy state signal pulmonary vasomotor responses to hypoxia has

been considered by many investigators, but the role of energy state in these responses remains

unclear

[375]. In fibroblasts and tumor cells, severe hypoxia causes ATP depletion thus

impairing DNA repair, but the role of intracellular ATP concentration ([ATP]i) in determining

the hypoxic response of HAPSMCs has not been studied earlier [376, 377]. [ATP]i in HPASMCs

is maintained at normoxic levels after incubation at 3 and 1% O2 and reduced after incubation at

0% O2. ATP depletion is concurrent with reduced cell number and enhanced apoptosis and

depolarization of the membrane potential suggesting that under hypoxia, HPASMCs continue to

proliferate and/or survive as long as the [ATP]i is maintained. However, if the level of hypoxia is

sufficient to cause depletion in intracellular ATP, the HPASMCs can no longer maintain the

mitochondrial membrane potential, and the cells apoptose.

As the duration of hypoxia is prolonged permanent structural remodeling occurs in the

vasculature. This reflects changes in structure and biochemical phenotype of all of the cells that

compose the pulmonary arteries. Universally observed is medial thickening and the appearance

of SMC- like (based on -actin staining) cells in previously nonmuscularized vessels. Our in vivo

data supports previous findings. In the current study, proliferation and apoptosis are both

enhanced in the main and branch pulmonary artery after 2 days and 7 days of hypoxia and

decreases after 14 days. This concurrent increase in cell proliferation as well as programmed cell

death confers plasticity to the vessel wall, whereby required structural change is enabled while

avoiding accumulation of mutations or abnormal vascular structure formation.

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134

Chapter 4

General Discussion and Conclusions

Chapter 4

135

VSMC proliferation has been recognized as

central to the pathology of several

cardiovascular diseases, such as atherosclerosis and hypertension. Hypoxia is an important

regulator of physiologic processes, including erythropoiesis, angiogenesis, and glycolysis. In the

vasculature, chronic hypoxia has been shown to cause proliferation of VSMCs, leading to vessel

wall

remodeling, a key pathophysiologic component of pulmonary hypertension. The

mechanisms by which hypoxia regulates VSMC growth include direct cell cycle-specific effects,

as well as indirect effects, via the regulation of VSMC mitogen production by neighboring

endothelial cells. Hypoxia triggers a cellular adaptive response that

is primarily mediated by the

transcription factor hypoxia-inducible factor 1 (HIF-1). Expression of HIF-1 target genes serves

to maintain cellular homeostasis. Transcriptional activation of

hypoxia-responsive genes

represents one major component of the vascular cell hypoxic response; however, the mechanisms

regulating VSMC proliferation and survival in the vessel wall – systemic or pulmonary, under

hypoxia, remain to be elucidated.

The normoxic pulmonary circulation is vasodilated and accommodates the entire cardiac

output at much lower pressures than the systemic circulation. During hypoxia, the pulmonary

arteries constrict, whereas systemic arteries, such as the aorta dilate. The mechanism of this

opposing control of tone between the two vascular beds is unknown. Although the response of

each bed to hypoxia is significantly modulated by the endothelium, the mechanism for the

opposing responses to hypoxia appear to lie within the VSMCs. Hypoxia increases intracellular

Ca2+

and contracts PASMCs; in contrast, SMCs from systemic arteries display decreased

intracellular Ca2+

and relax in response to hypoxia. The present series of investigations compares

the oxygen regulation of HASMC and HPASMC proliferation and survival.

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136

In both HASMCs and HPASMCs, incubation at various oxygen concentrations caused

directionally opposite responses – proliferation and growth arrest along with or without

apoptosis, depending upon the ability of these cells to maintain intracellular ATP concentration.

3% O2 incubation in both HPASMCs and HASMCs trigger a proliferative response without any

change in viability compared to normoxic cells. 0% O2 incubation diminishes cell numbers and

enhances apoptosis in both HPASMCs and HASMCs. However, the difference in the hypoxic

response of these two cell types is in the response at 1% O2 incubation: (i) In HASMCs cell

number is reduced after 16hrs and even more after 48hrs, while in HPASMCs, cell number

increases after 16hrs and decreases only after 48hrs; (ii) Incubation at 1% O2 in HASMCs, at

both 16 and 48hrs, reduces cell viability, increases apoptosis, depolarizes the mitochondrial

membrane potential and depletes intracellular ATP concentration. But in HPAMCs even though

the number of cells was reduced at 1% O2 for 48hrs, the viability along with mitochondrial

membrane potential and intracellular ATP concentration is maintained. Our study has shown that

hypoxia can increase or decrease cell number within the same cell type (systemic or pulmonary

smooth muscle cells), and the fundamental differences in response depend on the severity of

hypoxia that the cell is exposed to. The above responses also suggest that HPASMCs are better

adapted to reduced oxygen concentrations than their systemic counterpart.

In HASMCs an oxygen concentration of 1% O2 16hrs was sufficient to cause a delay in

cell cycle progression, with increased cells in G1/S phase. In HPASMCs, incubation at 1% O2 for

16 hrs caused an opposite effect – there is enhanced progression of cell cycle, with increased

cells in G2/M and S phases. But even in HPASMCs, incubation at 1% O2 for 48 hrs causes a

delay in cell cycle progression, with increased cells in G1/S phase. Together the data suggest that

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137

in both HASMCs and in HPAMCs, the bidirectional effects of hypoxia are integrated by events

occurring at the G1/S checkpoint, though the mechanism/s remain unclear.

In HASMCs at levels where hypoxia causes a decrease in cell number (1% and 0% O2)

our results have shown apoptosis to be a contributing factor towards the decrease. However, in

HPASMCs, at 1% O2 for 48hrs, no apoptosis is observed though there is a decrease in cell

number at this point. These results along with viability studies in HPASMCs, suggest that

oxygen concentrations which are capable of inducing apoptosis in systemic arterial smooth

muscle cells are not “sufficiently hypoxic” to induce cell death in pulmonary SMCs. In both the

pulmonary and the systemic circulation, protection from apoptotic cell death and enhanced

smooth muscle cell proliferation at moderate levels of hypoxia may be important in

neovasoformation during development and in ischemic tissues, where investiture of new

endothelial networks by regulatory smooth muscle cells must occur under conditions of relative

oxygen deficiency. Possibly since pulmonary circulation carries deoxygenated blood, the smooth

muscle cells of this region are genetically primed to survive under oxygen concentrations that are

otherwise toxic to the systemic smooth muscle cells. Reduced sensitivity of pulmonary smooth

muscle cells to hypoxia induced cell death provides the pulmonary vasculature an added

advantage. Dead cells are generally replaced by fibrous or connective tissue, which would have

otherwise impaired gas exchange and caused an early onset of pulmonary hypertension.

Enhanced apoptosis in both HASMCs and HPASMCs under severe hypoxia is also an adaptive

response that helps remove replication errors and prevents accumulation of hypoxia-induced

mutations.

Chapter 4

138

Conclusions

Systemic vascular smooth muscle cells experience a broad range of oxygen tensions

under physiological (90 mmHg in aortic lumen; 20 mmHg at 150 m depth; 24 mmHg in

terminal arterioles) and pathological (atherosclerosis, <7 mmHg) conditions. In earlier studies

hypoxia has had an inconsistent effect on smooth muscle cell proliferation and survival. This

lack of consistency has prevented the development of a unifying hypothesis regarding oxygen

regulation of these processes in diseases in which arterial remodeling plays a significant

pathophysiological role. Since cell replication is highly energy dependent it is intuitive, though

unproven, that hypoxia may have different effects depending on the degree to which cellular

energy status is compromised. On the other hand, many cell cycle associated genes are known to

be induced at levels of hypoxia at which inhibition of ATP synthesis would not be predicted and

a regulatory role beyond mere cytotoxicity merits consideration.

This study was carried out to determine if different proliferative and apoptotic responses

are elicited in human aortic smooth muscle cells (HASMCs) and human pulmonary artery

smooth muscle cells (HPASMCs) subjected to hypoxic incubation under conditions which do or

do not result in cellular ATP depletion, whether these effects are relevant to vascular remodeling

during hypoxia in vivo and to identify potential regulatory pathways using gene expression

profiling in cells exposed to conditions that elicit discordant responses.

The novel findings of our study in HASMCs, presented in Chapter 2, are:

1. Hypoxia may enhance or inhibit HASMC proliferation and differences in the cellular

response are elicited by varying degrees of hypoxic stress.

Chapter 4

139

2. The difference between O2 concentrations that enhance SMC proliferation and those that

impair cellular energy status and trigger cell destruction is small.

3. Pro- and antiproliferative and pro- and antiapoptotic gene expression are tightly

coordinated in HASMCs during hypoxic incubation to produce directionally opposed

responses, effected at the level of G1/S transition.

4. Our in vivo studies show increases in both SMC proliferation and death. This enhanced

cell turnover confers plasticity sufficient to enable the required structural change while

avoiding accumulation of mutations or abnormal vascular structure formation.

5. The net effect is an increased muscularity of the arterial wall, which is possibly a

compensatory adaptation that preserves the capacity to regulate the systemic circulation.

Hypoxic pulmonary arterial hypertension occurs in many cardiopulmonary diseases.

Increased smooth muscle cell proliferation in the pulmonary arterial wall is a hallmark of this

condition. The mechanism/s regulating these processes are yet unknown. The novel findings of

our study in HPASMCs, presented in Chapter 3, are:

1. HPASMCs proliferated at O2 concentrations which inhibited cell growth in HASMCs.

2. HPASMCs did not undergo apoptosis, maintained intracellular ATP concentration and

mitochondrial membrane potential at O2 concentrations which were cytotoxic to

HASMCs.

Together, our data suggest that the response to hypoxia, in either HASMCs or

HPASMCs, depends on whether the hypoxic stress is sufficient to impair cellular energy status.

In HPASMCs, ATP is preserved at a lower O2 concentration and this permits the cells to

maintain viability and proliferation at more severe levels of hypoxia than are tolerated by

HASMCs. The differences in the hypoxic response could be a result of adaptation to exposure to

Chapter 4

140

the deoxygenated blood that the pulmonary circulation carries or due to differences in the

embryonic origins of the cells that comprise the two circulations.

In vivo, hypoxia induces a state of enhanced cell turnover and, hence, plasticity. Vital

organ function is highly intolerant of oxygen deprivation. Accordingly, mechanisms linking

vascular cell turnover and the capacity for rapid structural change directly to oxygen

concentration are required to avoid delays inherent in second messenger signaling.

Our in vitro studies show that systemic and pulmonary artery smooth muscle cells

respond differently to hypoxia. HPASMCs are able to proliferate and remain viable at oxygen

concentrations that induce growth arrest and/or apoptosis and in HASMCs. However, under

reduced oxygen concentrations in both the cell types, systemic or pulmonary, cell viability is

maintained until intracellular ATP levels are depleted. Our data do not address the

mechanism(s) by which information about the site of origin of SMCs from systemic and

pulmonary circulations is preserved from generation to generation (passage to passage) in in

vitro cultures. A possible explanation for the different responses to hypoxia of the SMC when

coming from two different sources (systemic vs. pulmonary) could be attributed to their diverse

embryological origin. Arteries cranial to the heart are mostly products of the paired aortic arches,

which course axially within the branchial arches, thus interconnecting the ventral aorta with

paired dorsal aortas. The fourth pair of embryonic aortic arches becomes the aortic arch in the

adult while the fifth pair forms the pulmonary arteries and ductus arteriosus. The dorsal aortas

fuse into the single descending aorta [378].

During development of multicellular eukaryotic organisms, differences in genetic

programming cause the cells to differentiate into different types, which perform different

functions, and respond differently to the environmental stimuli. Thus, genes are silenced or

Chapter 4

141

activated in an epigenetically heritable fashion, giving cells a "memory" which determines their

phenotype over subsequent cell divisions. This implies that in order to grow and maintain a

specific, lineage-restricted state, such gene expression configurations need to be transmitted to

daughter cells, without mutations of the DNA. Two types of epigenetic modifications can

regulate chromatin conformation, thereby also regulating the transcriptional activity or silencing

of specific genomic regions. These include (1) DNA methylation at cytosine residues of CpG

dinucleotides in gene promoters, transposons, and imprinting control regions. In most cases,

DNA methylation is associated with gene repression; and (2) histone modifications in the

chromatin organization, such as methylation and acetylation, affecting the N-terminal tail of

histones [379].

Studies to define which mechanisms are active in distinguishing the responses of

PASMCs from SMCs derived from the systemic circulation are now indicated. Insight into the

molecular mechanisms underlying these responses would aid in the development of

pharmacological tools to modulate vascular remodeling in disease states.

Chapter 5

142

Chapter 5

Future directions

Chapter 5

143

Our studies in HASMCs (chapter 2) and HPASMCs (chapter 3) have shown that (i)

incubation of these cells at varying oxygen concentrations cause divergent responses – cell

proliferation and enhanced cell survival on one hand and growth arrest and/or apoptosis at

further reduced oxygen concentrations; (ii) cell viability under hypoxia remains unaffected as

long as the cells are able to maintain their intracellular ATP concentration. Oxygen

concentrations, at which intracellular ATP levels are diminished, cause the cells to undergo cell

cycle growth arrest (at the G1/S interphase) and cell apoptosis; (iii) HPASMCs proliferated at

oxygen concentrations which inhibited cell growth in HASMCs and (iv) HPASMCs did not

undergo apoptosis and maintained their intracellular ATP levels and mitochondrial membrane

potential at oxygen concentrations that are sufficient to deplete ATP and induce apoptosis in

HASMCs. Based on these, it could be proposed that in HPASMCs, ATP is preserved at a lower

O2 concentration to enable the cells to maintain their viability, compared to HASMCs. In other

words this difference/s in energetics in HASMCs and HPASMCs may play a key role in

determining oxygen regulation of their proliferation and survival.

The pulmonary circulation is adapted to function at a lower oxygen concentration

compared to the systemic circulation. There is greater heterogeneity in the phenotype of

pulmonary vascular SMCs compared to the systemic SMCs [325]. This varies with the size and

location of the pulmonary artery (Table 5.1). K+ channels are differentially distributed and there

are cell-specific differences in the endothelin-1 (ET-1) system [380]. The vasoconstrictor

response to ET-1 is mediated by ETA receptors in large pulmonary arteries but by ETB-like

receptors in smaller muscular pulmonary arteries [381].

Chapter 5

144

Table 5.1 Phenotypic heterogeneity in pulmonary vascular smooth muscle cells.

Feature Main Pulmonary artery smooth muscle cells Distal Pulmonary artery

smooth muscle cells

Location

Subendothelial Middle

Media

Outer

media

3000μ 1500μ 100-

150μ

L1 L2 L3 L4 L3 L4

Morphology Round, small,

irregularly

shaped ,

interspersed in

fragmented

elastin

Elongated,

spindle shaped,

oriented

circumferent-

ially between

well developed

and continuous

elastic lamellae

Large, spindle

shaped cells,

arranged in

compact cell

clusters, oriented

longitudinally

within vessel

wall devoid of

elastic lamellae

Small, spindle

shaped cells,

oriented

circumferent-

ially in

intersitial

areas between

L3 large cell

clusters

Same as Main

Pulmonary

Artery

-SMA

Calponin

- + + - + - + +

SM-MHC - + + - + - + +

Metavinculin

Desmin

- + + - + - + +

Caldesmon - + + - + - ND ND

Hypoxia ++ - - ++ - - - -

SMA: Smooth muscle actin, MHC: Myosin heavy chain,

+: Positive, -: Negative, ND: Not detected, ++: Increase in proliferation.

Chapter 5

145

Significant mitochondrial diversity exists between pulmonary and systemic arteries, with

direct imaging of rat vascular SMCs showing that PASMC mitochondria are more depolarized

than ASMC mitochondria. Compared to systemic arteries, pulmonary artery mitochondria have

higher rates of superoxide anion and H2O2 production, lower expression of proximal electron

transport chain components and higher expression of mitochondrial superoxide dismutase [326].

A deficiency in oxidative metabolism under hypoxic conditions will cause a cell to resort to

glycolytic ATP production. Increased activity of rate-limiting glycolytic enzymes has been found

in fibroblast, endothelial and kidney cell lines cultured under hypoxia.

Considering these, a relevant question to address in future would be whether metabolic

differences between systemic and pulmonary artery smooth muscle cells determine the

differences in oxygen regulation of their proliferation and survival. It is possible that HPASMCs

possess a greater aerobic capacity and/or an enhanced anaerobic capacity under hypoxia that

allows these cells to maintain the intracellular ATP concentration and survive at levels of

hypoxia that are cytotoxic to HASMCs.

Preliminary studies were conducted to assess both the aerobic and anaerobic capacities of

HASMCs and HPASMCs (Supplement). Real-time PCR was performed on DNA extracted from

normoxic and hypoxic HASMCs and HPASMCs for three nuclear-specific genes in the promoter

region, eNOS (endothelial nitric oxide synthase) promoter, iNOS (inducible nitric oxide

synthase) promoter, VCAM1 (vascular cell adhesion molecule 1) promoter, and three

mitochondrial-specific genes, mtTRNL1 (mitochondrially encoded tRNA leucine 1), CO2

(cytochrome c oxidase II), and Dloop (non-coding region in mtDNA controlling DNA

replication). Mitochondrial copy number was assessed as an index of oxidative phosphorylative

capacity in these cells (Supplementary Figure 1). To measure anaerobic glycolysis, protein

Chapter 5

146

levels of two key glycolytic enzymes namely Enolase and Phosphoglycerate kinase (PGK)

(Supplementary Figure 2 A-D) and lactate concentration (Supplementary Figure 2 E and F) have

been compared in normoxic and hypoxic HPASMCs and HASMCs.

The effect of hypoxic incubation on mitochondrial DNA levels is illustrated in

Supplementary Figure 1. In both HPASMCs and HASMCs, mitochondrial DNA levels decrease

after hypoxic incubation. The decline correlates with the severity of the hypoxic exposure and is

similar in the two cell types. Copy number values generated from standard curves (using

plasmid DNA and genomic DNA) were also compared and similar results were obtained.

Each human cell contains several hundreds to thousands of mitochondria

and each

mitochondrion has 2 to 10 copies of mitochondrial DNA (mtDNA). The copy number of mtDNA

may reflect the abundance of mitochondria in a human cell [382]. Mitochondrial biogenesis is a

highly regulated process and occurs on a regular basis in healthy cells, where it is controlled

by

the nuclear genome. Alteration of mitochondrial biogenesis and increased expression of nuclear

genes encoding mitochondrial proteins are responses triggered by mitochondrial dysfunction

or

high energy demands found in pathophysiological conditions [383, 384]. The decrease in

mtDNA copy number has been observed in lung cancer, hepatocellular carcinoma and gastric

cancers [385, 386]. Rapidly increased neuronal mitochondrial biogenesis has been observed after

hypoxic-ischemic brain injury [387]. Mitochondrial biogenesis has not been studied earlier in

HASMCs or HPAMSCs after incubation under various oxygen concentrations. Our results show

that in both HASMCs and HPASMCs, there is a decrease in mitochondria. Since oxygen serves

as a substrate in aerobic ATP production, it is quite obvious that under limited oxygen supply

both the cell types would choose to reduce their aerobic capacity, and possibly switch their

energy supply from aerobic metabolism to anaerobic glycolysis. The characteristics, including

Chapter 5

147

intron-less, without binding to histones, and inefficient mtDNA proof-reading and DNA repair

system, render mtDNA more susceptible to oxidative damage than nuclear DNA [388]. Reduced

mitochondria number could also be an adaptive response in both the cell types to prevent

accumulation of mutations.

ATP is the immediate source of metabolic energy whose hydrolysis causes protein

synthesis, muscle contraction and ion transport across cell membrane. In the outer mitochondrial

membrane, ATP phosphorylates creatine (Cr) by the creatine kinase (CPK) reaction:

[ATP] + [Cr] [PCr] + [ADP] + [H+]

The phosphocreatine (PCr) shuttle carries the high energy bonds from the mitochondria to the

sites of utilization where the CPK reaction is reversed (Lohman reaction) forming ATP and Cr

[389]. Future studies should assess the ratios of PCr ([PCr]) or inorganic phosphate ([Pi])

concentration to [ATP] or creatine ([Cr]) concentration

([PCr]/[ATP], [Pi]/[ATP], and

[PCr]/[Cr]), phosphorylation potential

( = [ATP]/[ADP][Pi], where [ADP] is ADP

concentration). It is possible that HPASMCs have a greater potential to maintain its ATP through

the Lohman reaction as above.

Hypoxia is known to up regulate expression of several glycolytic enzymes to enhance

anaerobic ATP production. Protein levels of glycolytic enzymes, PGK1 and enolase are

increased after hypoxic incubation, in both HPASMCs (Suppl Fig 2 A and C) and HASMCs

(Suppl Fig 2 B and D), suggesting enhanced glycolytic ATP production under

hypoxic

conditions in both cell types. Glycolysis in the cytoplasm produces the intermediate metabolite

pyruvate. Under aerobic conditions, pyruvate is converted to acetyl CoA to enter the Kreb’s

cycle. Under anaerobic conditions, pyruvate is converted by lactate dehydrogenase to lactic acid.

In aqueous solutions, lactic acid dissociates to lactate and H+. These H

+ can be used in the

Chapter 5

148

production of ATP by oxidative phosphorylation. Impairment of oxidative pathways, however,

during lactate production results in a net gain of H+ and acidosis occurs [390].

The lactate levels reflect the balance between lactate production and clearance. The

normal plasma lactate concentration is 0.3–1.3 mmol litre-1

, while concentrations >5 mmol litre-1

are considered high enough to cause acidosis. In vascular smooth muscle, changes in intracellular

pH (pHi) can alter membrane potential, calcium homeostasis, and myosin light chain kinase

activity [391-393]. Altered pHi has been shown to change vasomotor tone in both systemic and

pulmonary arteries and responses to hypoxia in isolated lungs [394-400]. Lactate concentrations

are increased under hypoxia in both HASMCs and HPASMCs. Incubation at 1% O2 causes

lactate to accumulate more (>5 mmol litre-1

) in HASMCs than HPASMCs. A possible

explanation could be that HPASMCs possess a better ability to prevent lactate accumulation and

thereby acidosis under hypoxic conditions. Likely, this excess lactate accumulation, beginning at

higher oxygen concentrations in HASMCs (1% O2) compared to that in HPASMCs (0% O2) that

make the aortic cells more vulnerable to acidosis induced cell apoptosis than the pulmonary

cells. Future studies will include determining the lactate/pyruvate ratio and rate of extracellular

acidification in both HASMCs and HPASMCs.

Factors other than enhanced lactic acid production might contribute to the fall of arterial

pHi during hypoxia. Decreased mitochondrial electron transport and proton

pumping could lead

directly to cytoplasmic acidification [401]. Na

+-H

+ exchange, an important component of pHi

regulation in vascular smooth muscle, depends on activity of the Na

+-K

+ pump, which requires

energy for operation [391, 402, 403]. Thus under hypoxia when ATP concentrations are

depleted, Na+-K

+-ATPase activity could be limited, which would reduce the transmembrane

sodium gradient, and decrease acid extrusion via Na

+-H

+ exchange. In HPASMCs, [ATP]i is

Chapter 5

149

maintained at oxygen concentrations that are sufficient to deplete ATP levels in HASMCs.

Hence in HPASMCs, Na+-H

+ exchange is maintained under hypoxia and onset of acidosis

delayed compared to HASMCs.

Alternatively, HPASMCs could genetically possess a greater ability to down regulate

ATP utilization in the face of hypoxia or a cytochrome oxidase of greater

oxygen affinity [404].

The two cell types might also differ in their glucose uptake. Role of glucose transporters and

metabolic sensitivity to various modulators of glycolytic enzymes need to be assessed. Further

investigation will determine which of these (or other) explanations is

correct. Insight into these

mechanisms could lead to the development of pharmacological tools for use in the treatment of

diseases associated with hypoxia.

Supplement

150

Supplement

Materials and Methods

Antibodies and Reagents

Phosphoglycerate kinase (PGK) and Enolase antibodies were purchased from Santa Cruz

Biotechnology, Inc (Santa Cruz, CA). Lactate assay kit was obtained from BioVision (Mountain

View, CA). All other reagents were from Sigma (St. Louis, MO).

Mitochondrial DNA content: Mitochondrial DNA (normalized for nuclear DNA) was quantified

as a marker of mitochondria number. HASMCs and HPASMCs were exposed to 48 hours of

normoxic or hypoxic (3%, 1%, and 0% O2) incubation. Cells were harvested with 1ml of lysis

buffer per plate (1M Tris (pH 8.0), 0.5M EDTA (pH 8.0), 10mg/ml RNase A, 10% SDS).

Samples were incubated at 37°C for 2 hours, proteinase K (20mg/ml) was added and the lysate

incubated at 50°C overnight. DNA was extracted using phenol/chloroform extraction and

ethanol-precipitated overnight. DNA was pelleted instead of spooled to ensure a maximum yield

including both nuclear and mitochondrial DNA, and resuspended in 30-80μl TE buffer.

Real-time Polymerase Chain Reaction (PCR): Primer pairs specific for three Human

mitochondrial genes and three nuclear chromosome genes were chosen for PCR, and cloned into

the pCR®

II plasmid using the TA Cloning®

Kit Dual Promoter (Invitrogen; Carlsbad, CA). The

three mitochondrial genes chosen were: tRNA leucine 1 (mtTRNL1: sense mt3212F 5’-

CACCCAAGAACAGGGTTTGT-3’; antisense mt3319R 5’-TGGCCATGGGTATGTTGTTAA-

3’); cytochrome c oxidase II (CO2: sense CO2F 5’-CCCCACATTAGGCTTAAAAACAGAT-

3’; antisense CO2R 5’-TATACCCCCGGTCGTGTAGCGGT-3’); and non-coding region in

tDNA controlling DNA replication (Dloop: sense DloopF 5’-

Supplement

151

TATCTTTTGGCGGTATGCACTTTTAACAGT-3’; antisense DloopR 5’-

TGATGAGATTAGTAGTATGGGAGTGG-3’). Primers for the three nuclear genes were

chosen to amplify genomic sequences in their promoter regions The three nuclear genes chosen

were inducible nitric oxide synthase (iNOS: sense hiNOStaqman 5’-

TGAAGAGGCACCACACAGAGT-3; antisense hiNOStaqman3’-

TGGTTTCCAAAGGGAGTGTCC-5’), endothelial nitric oxide synthase (eNOS: sense

heNOStaqman 5’- GTGGAGCTGAGGCTTTAGAGC-3’; antisense heNOStaqman 3’-

TTTCCTTAGGAAGAGGGAGGG-5’) and vascular cell adhesion molecule 1 (VCAM1: sense

5’- ACTTGGCTGGGTGTCTGTTA -3’; antisense VCAM 3’-

GCGGAGTGAAATAGAAAGTC -5’). The cloned plasmid sequences were verified by DNA

sequencing (The Centre for Applied Genomics; Toronto, ON), amplified by maxiprep and used

as plasmid standards for real-time PCR.

DNA concentrations were quantified and diluted to a concentration of approximately 500

copies of nuclear DNA per μl. Real-time PCR was performed on the DNA samples in triplicate.

Real-time PCR settings were 95°C for 10mins (Step I); 95°C for 15s, 60°C for 1 min (Step II x

40 cycles); 95°C for 15s, 60°C for 15s, 95°C for 15s (Step III). SYBR®

Green I dye (Applied

Biosystems; Foster City, CA) was used to detect the PCR products, and the specificities of the

amplicons were verified by comparing the Tm for each amplicon, which were consistent across

the different experiments. The number of copies of target sequence was determined by

comparison with standard curves generated by both plasmid DNA and genomic DNA serial

dilutions. Copy number values from mitochondrial genes were divided by that of nuclear genes

to normalize the amount of mtDNA to the total number of cells, for each cell type under each

condition. The amount of mtDNA for each hypoxic condition was divided by the amount of

Supplement

152

mtDNA in the normoxic control cells to determine the average fold change of mtDNA levels.

Lactate assay: Lactate is oxidized by lactate oxidase to generate a product, which interacts with

the lactate probe (provided by manufacturer) to produce fluorescence (at Exitation/Emission =

535/590 nm). Samples were prepared in50 μl/well with Lactate Assay Buffer in a 96-well plate.

A standard reference curve was used, corrected for background fluorescence for calculating

lactate concentrations, according to the instructions provided by the manufacturer.

Western blot analysis: Western blot analysis was used to quantify levels of glycolytic enzymes

Phosphoglycerate kinase and Enolase in cytoplasmic extracts from HPASMCs and HASMCs

exposed to normoxia or hypoxia (3%, 1% or 0% O2) for 16 or 48 h. Cells were lysed in buffer A

[10 mM HEPES (pH 7.8), 10 mM KCl, 0.1 mM EDTA, 1mM dithiothreitol (DTT), and 0.1%

Nonidet P-40 (NP-40)] with protease inhibitors (5 g/ml aprotinin, 5 g/ml pepstatin, 5 g/ml

leupeptin, 0.5 mM Pefabloc, and 1 mM phenylmethylsulfonyl fluoride) and phosphatase

inhibitors (10 mM sodium fluoride, 1 mM sodium orthovanadate, and 20 mM glycerophosphate).

Nuclear proteins were then extracted with buffer B [50 mM HEPES (pH 7.8), 420 mM KCl, 0.1

mM EDTA, 1 mM DTT, 5 mM MgCl2, and 20% glycerol], containing both protease and

phosphatase inhibitors. Equal amounts of protein extracted from HPASMCs and HASMCs,

incubated under normoxic and hypoxic (3%, 1% and 0% O2) conditions for 16 and 48 h, were

loaded on 4–12% Tris-glycine gels, separated by electrophoresis and transferred to

nitrocellulose. Membranes were blocked with 5% milk overnight and probed with anti-

Phosphoglycerate kinase (1:1000) and anti-Enolase (1:500). In all cases, protein concentration

was determined by the Bradford assay, and appropriate volumes of extraction buffer to produce

constant protein loading in each lane were mixed with SDS loading buffer. Equality of protein

loading and transfer efficiency were corroborated by full-lane densitometry of the Ponceau red-

Supplement

153

stained membranes. Immunoblots were probed with horseradish peroxidase-donkey anti-rabbit

IgG (1:1,000 in blocking buffer) and visualized by enhanced chemiluminescence (ECL Plus kit,

Amersham Biosciences). Band intensity was quantified by densitometry (Bio-Rad Laboratories,

Mississauga, ON, Canada).

Supplement

154

Figure S1 Average fold change of mtDNA levels in two cell types (HASMC and

HPASMC) under different hypoxic conditions (3%, 1%, and 0% O2). Primer pairs specific

for three human mitochondrial genes (tRNA leucine 1, cytochrome oxidase II, Dloop) and

three nuclear chromosome genes (iNOS, eNOS, VCAM1) were chosen for PCR, and cloned

into the pCR®

II plasmid. Copy number determined using plasmid DNA as standards.

Results were averaged from 3 independent experiments.

O2 concentration O2 concentration

HPASMC

HASMC

Supplement

155

HPASMC A.

B. HASMC

Figure S2 Cytoplasmic levels of Phosphoglycerate kinase protein in HPASMCs (A)

and HASMCs (B) after incubation under normoxic and hypoxic (3, 1 or 0% O2) conditions.

n= 6. *P <0.05 vs. corresponding normoxic control values.

Supplement

156

C. HPASMC

D. HASMC

Figure S2 Cytoplasmic levels of Enolase protein in HPASMCs (C) and HASMCs (D)

after incubation under normoxic and hypoxic (3, 1 or 0% O2) conditions. n= 6. *P <0.05 vs.

corresponding normoxic control values.

Supplement

157

Figure S2 Lactate concentration (mmol / L) after incubation of HPASMCs (E) and

HASMCs (F) under normoxic and hypoxic (3, 1 or 0% O2) conditions. n= 6. *P <0.05 vs.

corresponding normoxic control values.

E. HPASMCs

F. HASMCs

Lac

tate

co

nce

ntr

atio

n

(mm

ol

/L)

Lac

tate

co

nce

ntr

atio

n

(mm

ol

/L)

References

158

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