i

The Role of Vascular Matrix Metalloproteinase-2 and Heme Oxygenase-2 in Mediating the Response to

Hypoxia

by

Jeff ZiJian He

A thesis submitted in conformity with the requirements for the degree of doctor of philosophy

Department of Laboratory Medicine and Pathobiology University of Toronto

© Copyright by Jeff ZiJian He, 2009

ii

The Role of Vascular Matrix Metalloproteinase-2 and Heme

Oxygenase-2 in Mediating the Response to Hypoxia

Jeff ZiJian He

Doctor of Philosophy

Department of Laboratory Medicine and Pathobiology University of Toronto

2009

ABSTRACT Systemic hypoxia frequently occurs in patients with cardiopulmonary diseases.

Maintenance of vascular reactivity and endothelial viability is essential to preserving

oxygen delivery in these patients. The role of matrix metalloproteinase-2 (MMP-2) and

heme oxygenase-2 (HO-2) in the vascular response to hypoxia were investigated. In the

first part of the thesis, the role of MMP-2 in regulating systemic arterial contraction after

prolonged hypoxia was investigated. MMP-2 inhibition with cyclic peptide

CTTHWGFTLC (CTT) reduced phenylephrine (PE)-induced contraction in aortae and

mesenteric arteries harvested from rats exposed to hypoxia for 7 d. Responses to PE

were reduced in MMP-2-/- mice exposed to hypoxia for 7 d compared to wild-type

controls. CTT reduced contraction induced by big endothelin-1 (big ET-1) in aortae

harvested from rats exposed to hypoxia. Increased contraction to big ET-1 after hypoxia

was observed in wild-type controls, but not MMP-2-/- mice. Rat aortic MMP-2 and

MT1-MMP protein levels and MMP activity were increased after 7 d of hypoxia. Rat

aortic MMP-2 and MT1-MMP mRNA levels were increased in the deep medial vascular

iii

smooth muscle. These results suggest that hypoxic induction of MMP-2 activity

potentiates contraction in systemic conduit and resistance arteries through proteolytic

activation of big ET-1.

The second part of the thesis investigated oxygen regulation of HO-2 protein and

whether it plays a role in preserving endothelial cell viability during hypoxia. HO-2, but

not HO-1, protein level was maintained during hypoxia in human endothelial cells

through enhanced translation of HO-2 transcripts. Inhibition of HO-2 expression

increased the production of reactive oxygen species, decreased mitochondrial membrane

potential, and enhanced apoptotic cell death and activated caspases during hypoxia, but

not during normoxia. These data indicate that HO-2 is translationally regulated and

important in maintaining endothelial viability and function during hypoxia.

In summary, the thesis demonstrates the importance of MMP-2 and HO-2 in

preserving vascular function during prolonged systemic hypoxia. These enzymatic

pathways may, therefore, represent novel therapeutic targets that may be exploited to

ameliorate the effects of hypoxia in patients with cardiopulmonary disease.

iv

ACKNOWLEDGMENTS

I would like to thank all of the people who have helped and inspired me during my

doctoral study.

I especially want to thank my supervisor, Dr. Michael Ward, for his guidance during my

research. He has provided me with encouragement, sound advice, good teaching, and

lots of good ideas. I would have been lost without him. I want to thank my co-

supervisor, Dr. Philip Marsden, for constantly challenging me to look beyond the

obvious and ensuring that I am knowledgeable in the field. I also want to thank my

advisory members for their constructive criticism, advice, and continuous support.

My deepest gratitude goes to my parents, Ying Wong and Wing Ming Ho, for their

unflagging love and support throughout my life; this dissertation is simply impossible

without them. I am thankful everyday that they are part of my life.

I am indebted to my many student colleagues for providing a stimulating and fun

environment in which to learn and grow. I am especially grateful to Ben Lai, Shirley

Mei, Diana Wong, Julie Basu-Ray, Orisha Yacyshyn, Lakshmi Kugathasan for all the

emotional support, entertainment, and caring they provided. I look forward to their

company in the next phase of my life.

v

TABLE OF CONTENTS

ACKNOWLEDGMENTS iv

TABLE OF CONTENTS v

LIST OF TABLES viii

LIST OF FIGURES ix

ABBREVIATIONS xii

PREFACE xiv

CHAPTER 1 Introduction

1.1 Oxygen Delivery 2

1.2 Mechanisms of Oxygen Sensing 5

1.3 Regulation of Gene Expression by Hypoxia 6

1.4 Physiological Responses to Hypoxia 11

1.5 The Endothelium 14

1.6 Matrix Metalloproteinase-2 17

1.6.1 Matrix Metalloproteinase-2 Protein Structure 17

1.6.2 Regulation of Matrix Metalloproteinase-2 Activity 18

1.7 Endothelin 20

1.7.1 Activation of Endothelin-1 21

1.7.2 Endothelin Receptor Signalling 22

1.8 Heme Oxygenase 23

1.8.1 Functions of Heme Oxygenase-2 24

1.8.2 Structure and Expression of Heme Oxygenase-2 27

1.8.3 Regulation of Heme Oxygenase Protein Expression 29

1.9 Thesis Objectives 31

vi

CHAPTER 2 Induction of matrix metalloproteinase-2 enhances systemic arterial

contraction after hypoxia

2.1 Introduction 34

2.2 Materials and Methods

2.2.1 Exposure to Hypoxia 36

2.2.2 Chemicals/Antibodies 36

2.2.3 Aortic and Mesenteric Contractile Responses 37

2.2.4 Immunohistochemistry 38

2.2.5 Western Blots 38

2.2.6 Gelatin Zymography 39

2.2.7 MMP and ECE Activity 40

2.2.8 Aortic MMP-2, MT1-MMP, TIMPs 1 to 4 mRNA Levels 40

2.2.9 Contractile Responses in Aorta of MMP-2-/- and

MMP-2+/+ Mice 41

2.3 Results 43

2.4 Discussion 65

CHAPTER 3 Enhanced translation of HO-2 transcripts preserves human

endothelial cell viablility during prolonged hypoxia

3.1 Introduction 72

3.2 Materials and Methods

3.2.1 Chemicals and Reagents 73

3.2.2 Cell Culture Studies 73

3.2.3 Quantitative Real Time PCR 74

3.2.4 Western Blotting 75

3.2.5 3H-uridine and 3H-leucine Incorporation 75

3.2.6 35S-methionine Incorporation 76

3.2.7 Polysome Profiling 77

3.2.8 Measurement of Intracellular Reactive Oxygen Species 77

vii

3.2.9 Mitochondrial Membrane Depolarization 77

3.2.10 Annexin V/Propidium Iodide Labeling 78

3.2.11 Total Caspase Activation 78

3.3 Results 79

3.4 Discussion 99

CHAPTER 4 Perspective 106

REFERENCES 112

viii

LIST OF TABLES

Table 1. Effect of MMP inhibition on maximum contraction and EC50 values

during PE and big ET-1-induced rat aortic contraction 46

Table 2. Maximum contraction and EC50 values during PE- induced contraction and response to big ET-1 in MMP-2-/- and MMP-2+/+ mice 64

ix

LIST OF FIGURES

CHAPTER 1

Figure 1.1 Regional distribution of pO2 from the airways to the cytosol 3

Figure 1.2 Oxygen dissociation curve from adult haemoglobin 3

Figure 1.3 Oxygen regulation of HIF activity 7

Figure 1.4 Control of translation initiation during hypoxia 9

Figure 1.5 Domain structure of matrix metalloproteinase-2 18

Figure 1.6 Generation of endothelin-1 22

Figure 1.7 Metabolism of heme by heme oxygenase 24

Figure 1.8 Structural organization of the human HO-1 and HO-2 gene 28

CHAPTER 2

Figure 2.1 Concentration-response relationships for phenylephrine in aortic rings

and mesenteric arteries and concentration response relationships for

big ET-1 in aortic rings from normoxic rats and rats exposed to

hypoxia for 7 d 44

Figure 2.2 Immunohistochemistry for MMP-2 on aortic sections from normoxic

rats and rats exposed to hypoxia for 7 d 47

Figure 2.3 Aortic MMP-2 protein levels in normoxic rats and rats exposed to

hypoxia for 16 h, 48 h, and 7 d 49

Figure 2.4 MMP and ECE activity in aortae from normoxic rats and rats exposed

to hypoxia for 7 d 51

Figure 2.5 Levels of MT1-MMP, TIMP-1, -2, -3, and -4 proteins in aortae

from normoxic rats and rats exposed to hypoxia for 16 and 48 h

and 7 d 52

Figure 2.6 Levels of MMP-2, MT1-MMP, TIMP-1, -2, -3, -4 mRNAs in

aortae from normoxic rats and rats exposed to hypoxia for 16 h, 48 h,

and 7 d 56

x

Figure 2.7 MMP-2 and MT1-MMP mRNA levels in endothelial, sub-endothelial

VSMC, and deep medial VSMC from aortae of normoxic rats and from

rats exposed to hypoxia for 7 d 60

Figure 2.8 Concentration-response relationship for phenylephrine and contractile

response to big ET-1 in in aortic rings from MMP-2+/+ and MMP-2-/-

mice exposed to normoxia or hypoxia for 7 d 63

CHAPTER 3

Figure 3.1 HO-1 and HO-2 mRNA and protein levels in HUVEC exposed to

normoxia or 1% oxygen for either 16 or 48 h 81

Figure 3.2 HO-1 and HO-2 protein levels in HAEC and EPC exposed to normoxia

or 1% oxygen for 16 h 83

Figure 3.3 Rate of 3H-uridine (A) and 3H-leucine (B) incorporation into RNA and

protein of HUVEC exposed to normoxia or hypoxia for 16 or 48 h. (C)

Rate of 35S-methionine incorporation into HO-1 and HO-2 protein of

HUVECs exposed to normoxia or hypoxia for 16 h. (D) Quantification

of the abundance of HO-2 mRNA in various polysome fractions from

HUVEC exposed to normoxia or hypoxia for 6 or 24 h 85

Figure 3.4 ROS levels in HUVEC transfected with scrambled or HO-2 siRNA

exposed to normoxia or hypoxia for 48 h, or exposed to normoxia or

hypoxia for 16 h treated with TNF-α or H2O2 88

Figure 3.5 Mitochondrial membrane potential in HUVEC transfected with

scrambled or HO-2 siRNA exposed to normoxia or hypoxia for 48 h,

or exposed to normoxia or hypoxia for 16 h treated with TNF-α or

H2O2 91

Figure 3.6 Cell death in HUVEC transfected with scrambled or HO-2 siRNA

exposed to normoxia or hypoxia for 48 h, or exposed to normoxia or

hypoxia for 16 h treated with TNF-α or H2O2 93

xi

Figure 3.7 Total activated caspase level in HUVEC transfected with scrambled or

HO-2 siRNA exposed to normoxia or hypoxia for 48 h, or exposed to

normoxia or hypoxia for 16 h treated with TNF-α or H2O2 95

Figure 3.8 Cell death and total activated caspase level in HAEC transfected with

scrambled or HO-2 siRNA exposed to normoxic or hypoxia for 48 h or

or exposed to normoxia or hypoxia for 16 h treated with TNF-α or

H2O2 97

xii

ABBREVIATIONS

ANOVA analysis of variance ARD1 arrest defective 1 ATF activating transcription factor ATP adenosine triphosphate bHLH basic helix-loop-helix CGRP calcitonin gene related peptide CRC concentration response curve CO carbon monoxide CTT CTTHWGFTLC DCF 2’,7’-dichlorofluorescein ECE endothelin converting enzyme eIF eukaryotic initiation factor EPC endothelial progenitor cell ET-1 endothelin-1 ETAR endothelin type A receptor ETBR endothelin type B receptor ETC electron transfer chain FIH factor-inhibiting HIF H2O2 hydrogen peroxide HAEC human aortic endothelial cell HIF hypoxic inducible factor HO heme oxygenase HRM heme regulatory motif HUVEC human umbilical vein endothelial cell HVR hypoxic ventilatory response IRES internal ribosomal entry site JC-1 5,5’,6,6’-tetrachloro-1,1’,3,3’-tetraethylbenzimidazolcarbocyanine iodide MAPK mitogen-activated protein kinase MARE Maf recognition element miRNA microRNA MMP-2 matrix metalloproteinase-2 MT1-MMP membrane type-1 matrix metalloproteinase mTOR mammalian target of rapamycin NO nitric oxide ODD oxygen dependent degradation domain PE phenylephrine PERK PKR-like endoplasmic reticulum kinase PHD prolyl hydroxylase PI propidium iodide pO2 partial pressure of oxygen ROS reactive oxygen species TIMP tissue inhibitors of matrix metalloproteinase

xiii

TNF-α tumor necrosis factor α UTR untranslated region uORF upstream open reading frame VSMC vascular smooth muscle cell

xiv

PREFACE

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

292:H684-H693, 2007. Jeff Z. He, Adrian Quan, Yi Xu, Hwee Teoh, Guilin Wang,

Jason E. Fish, Brent M. Steer, Shigeyoshi Itohara, Philip A. Marsden, Sandra T. Davidge

and Michael E. Ward. Induction of matrix metalloproteinase-2 enhances systemic

arterial contraction after hypoxia. Permission has been obtained from the American

Physiological Society and all of the authors for inclusion of the paper in the thesis and

for the National Library to make use of 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

i) Figure 2.1 B: Adrian Quan measured the contractile response from

mesenteric arteries

ii) Figure 2.3: Experiment was done by Yi Xu from Dr. Davidge’s lab

iii) Figure 2.6 and 2.7: mRNA extraction and measurements were done by

Jason Fish and Brent Steer from Dr. Marsden’s lab

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

be submitted for publication before December 2008. 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 except

Figure 3.2: Blood outgrowth endothelial cells were a gift from Dr. Courtman

Figure 3.3 D: Experiment was done by members of Dr. Marsden’s Lab

Chapter 1

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CHAPTER 1

Introduction

Chapter 1

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In mammalian cells, oxygen serves as the terminal electron acceptor in the

mitochondria during generation of adenosine triphosphate (ATP) and as substrate for

numerous enzymes, such as heme oxygenases and prolyl hydroxylases.[1, 2] Depending

on the metabolic activity and the distance away from vessels, cells experience a variety

of oxygen concentrations under physiological conditions. Hypoxia is thus a relative

term, and is most usefully described as a condition in which normal tissue function is

inhibited due to insufficient oxygen supply. Reductions in systemic oxygen delivery

occur commonly in patients suffering from cardiopulmonary diseases, hemorrhagic

shock, or sepsis and in normal individuals during ascent to high altitudes.[3-5] The

molecular mechanisms that underlie the physiologic adaptations to hypoxia and its

pathophysiological consequences are complex and incompletely understood.

Investigation into these mechanisms will lead to the development of strategies to

mitigate the effects of hypoxia in these conditions. The role of vascular matrix

metalloproteinase-2 (MMP-2) and heme oxygenase-2 (HO-2) in the adaptive response to

hypoxia were investigated in the current study.

1.1 Oxygen Delivery

The delivery of oxygen to tissues is one of the main functions performed by the

cardiovascular system (Figure 1.1). The partial pressure of oxygen (pO2) in inspired air

at sea level is 21 kPa (~150 mmHg).[6] Oxygen extraction at the terminal airways and

CO2 accumulation in the alveolar gas decreases the alveolar pO2 to 14 kPa (~100

mmHg). At the alveoli, oxygen diffuses down its pressure gradient into the pulmonary

capillaries where it binds to haemoglobins in red blood cells. The median pO2 in

Chapter 1

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systemic arteries is 13 kPa (~92 mmHg), falling to 7 kPa (~50 mmHg) in arterioles and

~3-4 kPa (~25 mmHg) in precapillary arterioles and capillaries.[6, 7]

Oxygen is off-loaded from haemoglobin in red blood cells to tissues as it travels

through the cardiovascular system according to the oxygen-haemoglobin dissociation

curve (Figure 1.2). A substantial amount of oxygen diffuses out from the lumen of the

arterioles.[7, 8] The rate of diffusion increases from the larger to the smaller arterioles,

Figure 1.1 Regional distribution of pO2 from the airways to the cytosol. Source: Ward J (2007) Oxygen sensors in context.

Figure 1.2 Oxygen dissociation curve from adult haemoglobin. Source: http://www.anaesthesiamcq.com/downloads/odc.pdf

Chapter 1

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consistent with the lower velocity of blood in the smaller arterioles and their larger

surface area-to-volume ratio.[7, 9] Because the pO2 gradient between capillaries and the

tissue interstitium is small and pO2 actually increases in the venular circulation as blood

moves from the collecting venules to the larger veins, the capillary bed contributes

relatively little oxygen to tissues under resting conditions. The capillary circulation’s

function at rest appears predominantly to extract from the tissue byproducts of

metabolism and participate in oxygen exchange primarily when the tissue is active.

The amount of oxygen delivered to tissues is determined mainly by bulk blood

flow, haemoglobin saturation and the sympathetic nervous system. The cardiovascular

system constantly matches tissue oxygen supply to tissue oxygen demand by adjusting

these parameters. Oxygen supply to tissues could be enhanced by increasing vessel

diameter through the release of vasodilators (nitric oxide) and/or inhibition of

vasoconstrictors (endothelin-1). Increased capillary perfusion augments oxygen delivery

during exercise.[7] Oxygen delivery to tissues could also be increased by decreasing the

oxygen affinity for haemoglobin, as occurs during increases in body temperature or

carbon dioxide concentration. Other factors that are known to alter the affinity of

oxygen binding to haemoglobin include pH, carbon monoxide, and 2,3-

disphosphoglycerate. Lastly, activation of the adrenergic nervous system increases

oxygen delivery by optimizing of blood flow among different vascular beds to increase

oxygen extraction.[10, 11] The role of vascular MMP-2 in the contractile response to

adrenoceptor stimulation after exposure to hypoxia was investigated in the current study.

Chapter 1

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1.2 Mechanisms of oxygen sensing

Cells generally respond to hypoxia, especially if it is prolonged, by reducing

energy use and upregulating energetically efficient ATP producing pathways.[12] The

existence of these homeostatic processes in every cell implies that all cells have the

ability to sense changes in oxygen concentration. Changes in oxygen concentration are

detected by O2 sensors, which in turn regulate the activity of the effectors that determine

the modifications of specific cellular functions.[13] Despite recent progress in the

characterization of the cellular effectors of hypoxia, understanding of this process

remains incomplete.

Mitochondria have long been considered as a potential O2 sensing site since they

are the site of oxidative phosphorylation and electron transport and their activity is

altered by changes in oxygen concentration.[1, 14] Single electrons are lost to molecular

O2 to form ROS at various points in the electron transfer chain (ETC). The tendency for

the ETC to generate ROS increases during hypoxia because the Vmax of the cytochorome

oxidase is reversibly decreased due to reduced O2 availability.[1, 14] The increase in

ROS levels provides a favourable environment for iron to be in the Fe3+ state which,

along with decreased available oxygen, results in rapid inhibition of prolyl hydroxylase

activity leading to HIF-α stabilization and transactivation of hypoxia-inducible gene

expression. In addition to the mitochondria, plasma membrane-associated NAD(P)H

regulates intracellular ROS levels and function as an oxygen sensor in some cells.[15]

Other potential oxygen sensors that do not regulate intracellular ROS levels include

heme containing enzymes such as heme oxygenase and enzymes that utilize oxygen as

Chapter 1

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substrate, such as proly hydroxylase.[6, 16] It is now recognised that a number of

mechanisms with different sensitivities may effectively act as O2 sensors for different

cellular processes in different cell types.

1.3 Regulation of gene expression by hypoxia

Hypoxia activates a variety of complex pathways with the ultimate aim of

reinstating oxygen homeostasis. The identification of the hypoxia inducible factors

(HIFs) has greatly advanced our understanding of gene regulation during hypoxia.[17]

The HIFs are a family of transcription factors which are heterodimers containing of α

and β subunits.[18] They are characterized by the presence of basic helix-loop-helix

(bHLH) and Per/ARNT/Sim (PAS) domains. HIF-α, but not HIF-β, subunits possess an

oxygen-dependent degradation domain (ODD), rendering these proteins labile in the

presence of oxygen.[19]

HIF mediates a large number of critical responses that restore tissue oxygenation

and limit tissue injury.[14, 20] HIF activity is primarily controlled via the stability of

the alpha subunit (Figure 1.3). Under physiological conditions, prolyl hydroxylases

(PHD) catalyse hydroxylation of the HIF-α subunit on proline residues 402 and/or 564

within the ODD.[21] Upon hydroxylation, HIF-α binds to the VHL protein, which is

the recognition component of an E3 ubiquitin-protein ligase. Binding of VHL targets

HIF-α for ubiquitination and subsequent degradation by the 26S proteasome.[22]

Binding of HIF-α to VHL is enhanced by acetylation of HIF-α at lysine 532 in the ODD

by arrest defective 1 (ARD1).[23] In addition to PHDs and ARD1, HIF-α activity is

Chapter 1

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Figure 1.3 Oxygen regulation of HIF activity. Source: Schmid et. al. (2006) Lights on for Low Oxygen: A Noninvasive Mouse Model Useful for Sensing Oxygen Deficiency

regulated by factor-inhibiting HIF (FIH), which hydrolyzes asparagine 803 located in

the C-transactivation domains.[24, 25] Hydroxylation of asparagine 803 prevents

binding of p300/cAMP-response element-binding protein and reduces the transcriptional

activity of HIF-1 during normoxic conditions. During hypoxia, activities of both PHD

and FIH are inhibited.[16] Inhibition of HIF-α hydroxylation prevents HIF-α binding to

VHL, leading to stabilization and accumulation of HIF-α in the cytoplasm. The

stabilized HIF-α enters the nucleus to dimerize with HIF-β and activates transcription of

genes that contain hypoxic response element (5’-RCGTG-3’) in their promoter or

enhancer regions.[17, 26] Apart from the HIF family, hypoxia activates a number of

other important transcription factors, including nuclear factor κB, activator protien-1,

and p53.[27]

Normoxia

Hypoxia

HIF-1 accumulation + transcriptional activity

HIF-1 inactivation+ degradation

Chapter 1

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Whereas HIFs enable long-term cellular survival, a variety of HIF-independent

pathways promote ATP conservation by limiting energy-consuming processes.[14, 28]

Protein synthesis is the second costliest cellular process in terms of ATP demand and is

tightly linked to cellular oxygen availability.[29-33] In human cells, protein synthesis

drops to less than 50% of that in normoxic cells during hypoxic incubation (<1% O2)

due to inhibition of translation and later through effects on transcription.[34-37]

Translation of mRNA transcripts occurs in three stages: initiation, elongation, and

termination. Initiation is the most complex step and is tightly controlled by a number of

eukaryotic initiation factors (eIF).[38] The two central mechanisms for regulating

translation initiation are the assembly of eIF4F (comprised of eIF4A, eIF4E, and eIF4G

subunits) and the phosphorylation status of the eIF2α subunit.[39] Binding of the eIF4F

complex to the mRNA 5’cap structure is essential for initiating cap dependent

translation.[40, 41] Phosphorylation of the eIF2α subunit regulates the availability of

the ternary eIF2-GTP-Met-tRNA complex upon which 60S ribosomal subunit

recruitment depends.[12]

During acute hypoxia, global protein translation initiation is inhibited by

phosphorylation of eIF2α, preventing the exchange of GDP for GTP and sequestering

eIF2B in an inactive complex (Figure 1.4).[35, 42] Phosphorylation of eIF2α can be

achieved by several different kinases: interferon-inducible double-stranded RNA-

activated kinase, heme-regulated inhibitor of translation, kinase activated by nutrient

starvation, and PKR-like endoplasmic reticulum kinases (PERK).[43, 44] PERK is the

main kinase responsible for eIF2α phosphorylation following acute hypoxia.[36]

Chapter 1

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During prolonged hypoxia, cap-dependent protein translation initiation is

inhibited by preventing eIF4F complex formation (Figure 1.4).[35] A family of eIF4E-

binding proteins (4E-BPs) competes with eIF4G for binding to limited amounts of

eIF4E. Binding affinity of 4E-BPs to eIF4E is regulated by mammalian target of

rapamycin (mTOR). mTOR-mediated phosphorylation of 4E-BPs reduces the affinitiy

of 4E-BP binding to eIF4E.[45] Hypoxia inhibits mTOR activity, leading to

hypophosphorylation of 4E-BP and its binding to eIF4E.[38, 46, 47]

Logic dictates that mechanisms must exist to ensure that proteins which comprise

of the phenotypic modulation on which hypoxic adaptation depends are synthesized in

the face of global suppression of cap-dependent mRNA translation. Polysome profile

comparisons of cells cultured under normoxia and hypoxia have identified mRNA

Figure 1.4 Control of translation initiation during hypoxia. Source: Van Den Beuken et al. (2006) Translational Control of Gene Expression During hypoxia

Chapter 1

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transcripts that remain actively translated during hypoxia.[48-50] Translationally

regulated transcripts show an increased association with polysomes during hypoxia

compared with normoxia. These genes generally have long and GC rich nucleotide

sequences in their 5’ untranslated region (UTR) and significant secondary structures that

impede ribosomal scanning for the initiation codon.[12] One of the genes regulated at

the translational level during hypoxia is activating transcription factor 4 (ATF4).[48, 51]

The 5’-UTR of ATF4 contains two conserved upstream open reading frames (uORF).

Under normoxia, ATF4 protein synthesis is repressed because most ribosomes that

translate the first uORF reinitiate translation at the second uORF rather than at the ATF4

ORF. During hypoxia, ribosome reintiation is delayed by low ternary complex

availability. The delay increases the proportion of ribosomes that scan through the

second uORF and reinitiate, instead, at the ATF4 ORF. The presence of uORF,

therefore, represents an RNA element that can stimulate translation during hypoxia.

Other genes containing uORFs and are selectively translated when eIF2α is

phosphorylated include CHOP and GADD34.[35, 52]

In an environment of low eIF4F availability, a competitive advantage over other

mRNAs for ribosome binding could be conferred through activation of transcripts

containing a 5’-UTR lacking secondary structure that obviate the need to engage the

helicase activity of the eIF4F complex. This was identified as the operative mechanism

regulating expression of the hypoxia inducible nNOS variant.[27] Another gene with

extensive variation in the 5’-UTR that influences its translational efficiency is Dicer.[53]

Enhanced protein translation during hypoxia could also occur for transcripts containing

Chapter 1

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internal ribosomal entry sites (IRES) within the untranslated regions of the mRNA.[28]

IRES increases translation by facilitating direct ribosome binding independent of

formation of eIF4F at the cap, which is inhibited during hypoxia. Translation of the

angiogenic factor Tie2 transcript is enhanced through its IRES activity at the 5’-UTR

during hypoxia in human umbilical vein endothelial cells (HUVECs).[54] MicroRNAs

(miRNA) are small noncoding RNAs that regulate gene expression post transcriptionally

by preventing initiation or elongation.[55] Selective relief from micro-RNA mediated

suppression could be another mechanism that selectively enhances translation during

hypoxia. For example, the cationic amino acid transporter 1 translation under stress is

enhanced due to relief of micro-RNA mediated suppression by miR-122 in human

hepatic cell lines.[56] In the current study, the effect of hypoxia on HO-2 translation in

human endothelial cells was investigated.

1.4 Physiological responses to hypoxia

Oxygen supply to essential organs during acute systemic hypoxia is initially

maintained through three mechanisms. First, ventilation is increased through increases

in respiratory rate and tidal volume.[57] This response, termed the hypoxic ventilatory

response (HVR), in humans is almost solely due to depolarization of glomus cells in the

carotid body.[16] Second, alveolar oxygen uptake is enhanced by improved ventilation-

perfusion matching. This response is mediated by hypoxia-induced modulation of

pulmonary arterial and bronchial smooth muscle tone.[58] Pulmonary arterial

vasoconstriction redirects blood to better oxygenated regions of the lung while changes

in bronchial and bronchiolar tone optimize the distribution of gas flow within the lung.

Chapter 1

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Third, oxygen delivery to essential organs is maintained by activation of the sympathetic

system and release of local vasodilators.

Activation of the 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 heart and brain. Schlichtig et al.

demonstrated that redistribution of blood flow improves oxygen delivery during

progressive hemorrhage.[11] Cain demonstrated that vigourous vasoconstrictor

sympathetic tone during hypoxia increased survival time by promoting greater O2

extraction.[10] 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.

Hypoxic vasodilation is particularly well manifested in coronary and cerebral vessels

where it ensures sufficient oxygen to support the activity of the working myocardium

and the neuronal activity of the brain.[3]

Conduit vessels are also subject to dual regulatory mechanisms. Sympathetic

responses regulate blood flow among organs while local paracrine responses modulate

tone in response to regional environmental conditions.[59] Contractile response to

adrenoceptor stimulation is impaired in conduit and resistance vessels from animals

exposed to prolonged systemic hypoxia in vivo.[60-64] Hu et al. demonstrated that

chronic hypoxia attenuates coupling efficiency of α1-adrenoceptors to inositol 1,4,5-

trisphosphate synthesis in the uterine artery.[65] Others have shown impaired smooth

muscle activation in aorta of rats after prolonged exposure to hypoxia.[62, 63] These

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results suggest that as the duration of systemic hypoxia is prolonged, as occurs in

patients suffering from cardiovascular and pulmonary diseases, shock, or sepsis, the

sympathetically mediated responses to hypoxia are impaired and lead to reduced oxygen

extraction. In this setting, the endothelium plays an increasingly important role in

preserving the capacity to regulate the systemic circulation by releasing vasoconstrictor

substances which maintain vasoreactivity.[61, 64] In the current work, the role of

matrix metalloproteinase-2 in preserving vascular reactivity and the role of heme

oxygenase-2 in maintaining endothelial viability during prolonged exposure to hypoxia

are, therefore, investigated.

Chapter 1

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1.5 The endothelium

The endothelium consists of a single layer of cells lining the luminal side of all

blood vessels. Under physiological conditions, the endothelium works in concert with

other cells in the vessel wall to maintain vascular homeostasis by 1) maintaining a non-

thrombogenic blood-tissue interface; (2) regulating leukocyte adhesion/migration; 3)

participating in the regulation of vascular tone; and 4) producing cytokines and other

paracrine signalling molecules in response to external stimuli.[66]

In addition to its homeostatic functions, the endothelium constitutes an active

gate for the passage of oxygen into the interstitium.[67, 68] The endothelium consumes

oxygen at a rate one to two orders of magnitudes higher than is observed in most tissues

(about 0.1 ml O2/min/cm3 for arterioles) and alterations in its metabolic activity can

affect the gradient in oxygen concentration that determines tissue oxygen delivery.[7,

68] As a result, induction of vasoconstriction lowers tissue pO2 and increases vessel

wall oxygen consumption, whereas vasodilation is associated with a decrease in the

arteriolar vessel wall oxygen concentration gradient with a concomitant increase in

tissue pO2. [7] This effect compounds the detrimental effect of excessive

vasoconstriction on tissue perfusion and contributes to the impairment of viability and

function during hypoxia by further reducing tissue oxygen delivery.

Because of their location, endothelial cells are directly and frequently subjected

to changes in oxygen tension. In comparison to other cell types, endothelial cells are

relatively resistant to hypoxia-induced cell death.[69, 70] However, hypoxia triggers

profound changes in endothelial phenotype leading to disruption of the homeostatic

balance in coagulation, vascular permeability and vascular tone. These changes, if

Chapter 1

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uncontrolled, result in failure of microcirculatory regulation of oxygen extraction, local

inflammation, thrombosis and eventually organ failure and death. Endothelial injury and

dysfunction occur in many disease processes, including diabetes, atherosclerosis,

systemic and pulmonary hypertension, and inflammatory syndromes.[71]

Regulation of blood flow is one of the essential functions of the endothelium,

especially during hypoxia. Under normal physiological conditions, the vasoconstricting

and vasodilating substances produced by endothelium are in dynamic balance and

influence each other through multiple feedback mechanisms.[72] Vasodilating

substances released by the endothelium include CO, NO, prostacylin, and endothelium-

derived hyperpolarizing factor. Vasoconstricting factors released by the endothelium

include prostagladins (Thromboxane A2 and Prostaglandin F2α) and endothelins. Upon

exposure to hypoxia, the endothelium initiates a rapid but transient vasoconstriction

followed by relaxation. As the duration of hypoxic exposure is prolonged, smooth

muscle contractility is reduced and endothelial function is dramatically altered in both

conduit and resistance vessels.[60-62, 64, 73-75] Specifically, the inhibitory effect of the

endothelium on basal myogenic tone is lost and endothelium-dependent relaxation is

impaired. Furthermore, the contractile responses to α-agonist, KCl, and increased

transmural pressure are enhanced in endothelium-intact compared to denuded vessels,

indicating an excess endothelial vasoconstrictor over vasodilator release. Superoxide

ion, endothelin-1 and vasoconstrictor prostanoids are some of the factors implicated in

endothelium-dependent vasoconstriction during hypoxia.[69]

Chapter 1

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Changes in the endothelial function during hypoxia have both adaptive and

pathophysiological consequences. On one hand, since systemic vasoreactivity to

adrenoceptor stimulation is impaired following hypoxia,[60, 64] and since the reflexes

that redistribute blood flow and preserve vital organ oxygen supply are sympathetically

mediated,[76-78] they represent an important compensatory response that preserves the

capacity to regulate the circulation in the early phases of hypoxic stress. On the other

hand, impairment of the endothelium dependent mechanisms that optimize tissue

perfusion and adjust arteriolar tone in response to hypoxia[79-81] will compromise the

capacity to match blood flow to metabolic demand in vital vascular beds. Hence,

understanding pathways that regulate vascular reactivity during prolonged hypoxia is

important in understanding the pathophysiology of diseases associated with tissue

hypoxia and the development of effective treatment strategies. In the studies described

in this thesis, the role of two important pathways of endothelium-dependent regulation

of vascular function, matrix metalloproteinase-2 and heme oxygenase-2, are

investigated.

Chapter 1

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1.6 Matrix metalloproteinase-2

Matrix Metalloproeinase-2 (MMP-2) is produced constitutively by a wide range

of cell types, including endothelial cells and vascular smooth muscle cells.[82] Its

expression and activity increases during hypoxia.[83] MMP-2 metabolizes a wide

variety of matrix proteins, including gelatin, type I, IV and V collagens, elastin, and

vitronectin.[84] Based on its ability to degrade extracellular matrix proteins, MMP-2 is

thought to play a major role in vascular remodelling and angiogenesis during

hypoxia.[84-91] A broader biological role for MMP-2 has become apparent as novel

MMP-2 substrates are identified.[84, 92] For example, MMP-2 catalyzed degradation of

contractile proteins contributes to contractile dysfunction after ischemia/reperfusion

injury and dampens inflammatory responses due to the effects of its activity on

monocyte chemotactic protein-3.[83, 93-95] MMP-2 also regulates vascular tone

through modification of the activity of vasoactive molecules. MMP-2 catalyzed

inactivation of the potent vasodilatory neuropeptide calcitonin gene-related peptide

(CGRP) and activation of big endothelin-1 (ET-1) may both contribute to altered

vasoreactivity during hypoxic exposure [96, 97]. Therefore, hypoxic regulation of

vascular MMP-2 expression and activity and the role it plays in altered contractile

responses observed in systemic conduit and resistance arteries after prolonged hypoxia

in vivo was investigated.

1.6.1 Matrix Metalloproteinase-2 protein structure

Matrix metalloproteinases are a family of 28 related zinc-containing

endopeptidases that share structural similarities, but differ from each other in their

Chapter 1

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expression profiles and substrates.[82] MMPs are classified on the basis of substrate

specificity, sequence similarity , and domain organization into collagenases,

stromelysins, gelatinases, and membrane type metalloproteinases.[82] The two

gelatinases in the MMP family are MMP-2 and MMP-9. The activated MMP-2 (64

kDa) protein contains a hemopexin/vitronectin-like domain that is connected to the

catalytic domain by a hinge region (Figure 1.5). The hemopexin domain contains four

repeats with the first and last repeat linked by a disulfide bond. It influences substrate

specificity and binding of tissue inhibitors of matrix metalloproteinase (TIMP) to MMP-

2.[82] The catalytic domain contains three head-to-tail cysteine-rich repeats resembling

the collagen-binding type II repeats of fibronectin. The cysteine-rich repeats are

required for MMP-2 to bind and cleave collagen and elastin.[82]

1.6.2 Regulation of matrix metalloproteinase-2 activity

Like all MMPs, MMP-2 is secreted as ab inactive proenzyme.[86] Although

MMP-2 is constitutively released, its activity is tightly controlled posttranslationally

through a unique mechanism of proenzyme activation and interaction with its

endogenous inhibitors.[98] Unlike other MMPs, MMP-2 is refractory to activation by

Figure 1.5 Domain structure of matrix metalloproteinase-2. Source: Schulz R. (2007) Intracellular targets of matrix metalloproteinase-2 in cardiac disease: rationale and therapeutic approaches

Chapter 1

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serine proteinases and is instead, activated at the cell surface by membrane type-matrix

metalloproteinases (MT-MMP).[82] The activity of activated MMP-2 is inhibited by

TIMPs and the broad spectrum proteinase inhibitor α2-macroglobulin. Therefore, the

overall activity of MMP-2 depends not only on its abundance but also on the relative

bioactivity of its activators and inhibitors.

MMP-2 is produced and secreted as a 72 kDa proenzyme called proMMP-2.[85]

Activation of proMMP-2 occurs through a multistep pathway involving MT1-MMP and

TIMP-2.[98, 99] Initially, a cell surface MT1-MMP binds to the N-terminal domain of

TIMP-2. The C-terminal domain of the bound TIMP-2 then acts as a receptor for the

hemopexin domain of proMMP-2. Once proMMP-2 is tethered to TIMP-2, an adjacent

MT1-MMP that is free of TIMP-2 cleaves portions of the MMP-2 prodomain.

Following the initial cleavage by MT1-MMP, the residual portion of the MMP-2

propeptide is removed by another MMP-2 molecule to yield the fully active form of

MMP-2 that is 64 kDa in size.[98, 99]

In addition to proteolytic activation, reactive oxygen and nitrogen species are

known to activate MMP-2 without proteolytic removal of the autoinhibitory propeptide

domain.[84] The cysteine residues of the propeptide domain are highly sensitive to

changes in the redox environment and may exist in one of several oxidation states

depending on the type and level of oxidative challenge. Human recombinant 72 kDa

MMP-2 has been show to be activated by very low concentrations of peroxynitrite and

this occurred without evidence for the formation of the lower-molecular-weight 64 kDa

enzyme.[84]

Chapter 1

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1.7 Endothelin

The endothelins are a family of three small peptides, endothelin-1 (ET-1),

endothelin-2, and endothelin-3, that are central to regulating vascular function.[100]

ET-1, ET-2, and ET-3 are encoded by distinct genes located on chromosomes 6, 1, and

20, respectively which encode their respective propeptides (big ET-1, -2 and -3) which

require proteolytic activation to generate the vasoactive peptides.[101] ET-1 is

recognized as the major isoform of relevance in human cardiovascular physiology and

pathophysiology and regulation of vascular tone is one of the major functions of ET-

1.[100, 102-105] Altered expression/activity of ET-1 is involved in the development of

diseases such as hypertension, atherosclerosis, and vasospasm.[100]

Previous results suggest that ET-1 plays a central role in the adaptation to

hypoxia. Hypoxia increases circulating ET-1 concentrations, as well as its production

by cultured endothelial cells and rat arteries in vitro.[106-110] ET-1 has been

demonstrated to potentiate vascular responses that enhance pulmonary gas exchange and

tissue oxygen extraction,[61, 111, 112] however, the predominant mechanism of big

ET-1 activation in the systemic vasculature during hypoxia is unknown. Accordingly,

the current study was undertaken to investigate the contribution of ECE and MMP-2 in

the activation of big ET-1 in systemic arteries of rats exposed to prolonged hypoxia.

ET-1 acts primarily as a local hormone in an autocrine and paracrine

fashion.[113] It has been shown to be secreted primarily into the basolateral

compartment and not into the apical compartment.[114] Thus, plasma concentrations of

ET-1 are typically very low, ranging between 1 and 5 pM and may not be representative

of locally released ET-1 at its site of action.[115, 116] The effects of endothelins are

Chapter 1

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mediated primarily by two receptor subtypes ETA and ETB. In the systemic and

pulmonary vessels under physiological condidtions, endothelin type A receptor (ETAR)

are located primarily on vascular smooth muscle cells while endothelin type B receptor

(ETBR) are expressed on both vascular endothelial and smooth muscle cells. Both

receptors expressed on vascular smooth muscle cells mediate vasoconstriction while

endothelial ETBR activate the release of vasodilating factors, such as prostacyclin or

nitric oxide.[117] The net effect produced by ET-1, whether vasoconstriction or

vasodilation, is determined by the balance between ETAR and ETBR expression and

localisation.

1.7.1 Activation of big endothelin-1

Human endothelin-1 protein is encoded by the preproendothelin-1 mRNA.[118]

Removal of the signal peptide from preproET-1 in the rough endoplasmic reticulum

yields the ~200 amino acid proET-1 (Figure 1.6).[119] ProET-1 is, in turn, cleaved by a

furin-like convertase to release the precursor peptide big ET-1 which has minimal

biological activity and requires further proteolytic activation. The primary pathway for

big ET-1 activation is cleavage by the type II integral membrane zinc metalloproteinases

endothelin converting enzymes (ECE) 1 & 2.[120] These enzymes catalyse hydrolysis

of the Trp21-Val22/Ile22 bond of big ET-1 to release the 21 amino acid active peptide

termed ET-1[1-21]. Other enzymes also cleave big ET-1, including chymase, and MMP-

2. MMP-2 cleavage of big ET-1 at Gly32-Leu33 generates the isopeptide ET-1[1-32] with

enhanced potency compared to ET-1[1-21].[96, 121]

Chapter 1

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1.7.2 Endothelin receptor signalling

Endothelin-1 exerts its effects by activating cell-surface ETAR or ETBR.[122]

ETAR and ETBR are members of the G-protein-coupled receptor superfamily containing

seven hydrophobic transmembrane domains, an intracytoplasmatic C terminus and an

extracellular N terminus. Binding of ET-1 to ETAR or ETBR stimulates phospholipase C

activity through a pertussis toxin-insensitive G protein that is coupled to the ET receptor

intracellular domain. Phospholipase C hydrolyzes phosphatidyl inositol bisphosphate

into diacylglycerol and inositol triphosphate, leading to increase intracellular Ca2+ levels

and signalling through the RAF/MEK/MAPK pathway to activate genes that promote

cell growth and mitogenesis.[123-127] Increases in intracellular Ca2+ leads to SMC

contraction and activation of eNOS in endothelial cells.

Figure 1.6 Generation of endothelin-1.

preproET-1 proET-1furin like endopeptidase

Chapter 1

- 23 -

1.8 Heme Oxygenase

Given the multifaceted actions of ET-1 and consequences of excessive ET-1

production, numerous mechanisms have evolved to modulate the local bioavailability

and potency of ET-1. Another enzyme in the vasculature that modulates ET-1

bioavailability and potency is heme oxygenase.[74, 128-130] Heme oxygenases (HO)

are the only enzymes that catalyze the oxidative breakdown of heme into biliverdin IXα,

carbon monoxide (CO), and ferrous iron (Fe2+).[131, 132] Biliverdin IXα is

subsequently reduced to bilirubin IXα by biliverdin IXα reductase (Figure 1.7).[133]

Biliverdin/bilirubin possesses the capacity to suppress intracellular concentrations of the

reactive oxygen species (ROS) that regulate ET-1 precursor mRNA expression through

its antioxidant properties.[134] CO mimics many NO functions including cGMP-

dependent and –independent inhibition of agonist-induced vascular smooth muscle

contraction.[135] HO activity decreases ET-1-mediated potentiation of contraction to α-

adrenoceptor stimulation in aorta of rats exposed to hypoxia.[74] Taken together, these

findings suggest that HO-2 is an important enzyme in regulating vascular function in

systemic vessels during hypoxia.

The HO enzymatic activity requires three moles of molecular oxygen per heme

molecule oxidized, and the reducing equivalents from NAD(P)H.[136] Bilirubin

produced in the process is excreted from cells and passes through the blood in

association with serum proteins such as albumin to the liver where it is excreted in bile.

CO binds to haemoglobin to form carboxyhemoglobin, which is transported to the lung

Chapter 1

- 24 -

and is excreted in exhaled air. Iron is mainly transported to the bone marrow where it is

reused for heme biosynthesis and erythropoiesis.

1.7.1 Functions of heme oxygenase-2

By virtue of its catalytic action, heme oxygenase 2 regulates intracellular

concentrations of heme, CO, biliverdin/bilirubin, and ferrous iron, each of which has

important cellular functions.[135] Investigations into the function of heme oxygenases

have revealed that heme oxygenase activity possesses antioxidant, vasodilatory, and

antiapoptotic properties. These findings suggest that HO activity may ameliorate the

deleterious effects of hypoxia on endothelial function. Therefore, the role of HO-2 in

preserving endothelial cell viability during prolonged hypoxic exposure was investigated

during my studies.

The antioxidant function of HO-2 is supported by the findings that HO-2-/- mice

are more susceptible to hyperoxic lung damage, streptozotocin-induced renal

dysfunction, and intracerebral hemorrhage than wild-type mice.[137-139] Furthermore,

Figure 1.7 Metabolism of heme by heme oxygenase. Source: Ryter et al. (2005) Heme Oxygenase-1/Carbon Monoxide: From Basic Science to Therapeutic Applications.

Chapter 1

- 25 -

overexpression of the catalytically inactive HO-2 protein protects against H2O2-induced

oxidative stress.[140] Heme oxygenase exerts its antioxidative effects by reducing

levels of the oxidative heme molecules and by increasing antioxidant bilirubin and

ferritin levels. Unsequestered or free heme is a potent oxidant.[141, 142] Together with

iron, heme catalyzes free radical reactions to create ROS that damage DNA and proteins.

Heme oxygenases are the only known enzymes that can degrade heme and thus play a

critical role in heme homeostasis. In addition to metabolising heme, HO-2 can also

reduce intracellular heme levels by sequestering heme at its heme binding motifs, which

is not present in HO-1.[143] Bilirubin exerts its antioxidant effect through a repeated

process by which it is oxidized to biliverdin and then recycled back to bilirubin by

biliverdin reductase.[144] At micromolar concentrations in cell culture media, bilirubin

protects against cytotoxicity induced by H2O2 and/or enzymatically generated ROS in

endothelial and vascular smooth muscle cells.[145, 146] The antioxidant protection of

HO with respect to ferrous iron is facilitated by ferritin. The enhancement of ferritin

synthesis induced by ferrous iron increases the iron storage capacity of the cell as well as

protecting cells from oxidative stress.[147, 148]

The vasodilatory role of HO-2 has been demonstrated in numerous studies.

Govindaraju et al. has shown that HO-2 contributes to impaired contractile responses in

systemic arteries of rats exposed to hypoxia.[74] HO-2 has also been shown to regulate

blood flow in the uteroplancental vascular system.[149, 150] The vasodilatory effects of

heme oxygenase-2 are mainly mediated through the effects of CO, although bilirubin

could play an indirect role.[151-154] CO is a stable non-radical that binds the heme iron

moiety in a number of hemoproteins and metalloenzymes. It exerts its vasodilatory

Chapter 1

- 26 -

properties through several mechanisms. Binding of CO to the heme iron of soluble

guanlylate cyclase stimulates its enzymatic activity and increases cGMP

production.[153] Other mechanisms of CO-mediated vasodilation include activation of

calcium-dependent potassium channels and/or blocking expression of endothelial-

derived vasoconstrictors, such as ET-1.[128, 129, 155, 156] Bilirubin may regulate

vascular tone by suppressing intracellular concentrations of the ROS that serve as

second messengers in signalling the response to a number of contractile agonists,

including ET-1. During hypoxia, increases in ET-1 and HO-2 immunoreactivity are

localized to the endothelium [74] and HO inhibition increases the contractile response to

phenylephrine and ET-1 in endothelium-intact, but not –denuded aortic rings from

hypoxia-exposed rats.[74]

An anti-apoptotic role for HO-2 was demonstrated in studies in which inhibition

of HO-2 protein levels was shown to increase apoptosis induced by TNF-α, glutamate,

or hydrogen peroxide.[140, 157-159] The anti-apoptotic properties of CO are mediated

mainly through activation of mitogen-activated protein kinases (MAPK).[160, 161]

MAPK are a family of Ser/Thr protein kinases that are activated in response to a variety

of stimuli.[162] The three major MAPK signalling pathways identified in mammalian

cells include extracellular signal-regulated protein kinase, p38 MAPK, and c-Jun NH2-

terminal protein kinase.[162] The inhibitory effect of CO on TNF-α induced apoptosis

was abolished with a p38 MAPK dominant negative mutant in endothelial cells,

implying a critical role for the p38 MAPK pathway.[163] In addition to directly

Chapter 1

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activating anti-apoptotic pathways, HO activity could be cytoprotective through its anti-

oxidant and anti-inflammatory effects.

1.7.2 Structure and expression heme oxygenase-2

Heme oxygenases consist of two structurally related isozymes representing

products of distinct genes. The human HO-1 and HO-2 genes are localized to

chromosome 22q12 and 16q13.3, respectively (Figure 1.8).[164] Rat HO-1 and HO-2

gene share similar organization into five exons and four introns, but only share 43% in

amino acid homology.[165, 166] A highly conserved sequence of 24 amino acid

residues has been identified in common to both HO-1 and HO-2.[167] Both HO-1 and

HO-2 also share similar hydrophobic regions at the extreme COOH terminus that serve

to anchor the protein in cellular membranes. Although HO-1 lacks Cys residues, HO-2

contain three Cys-Pro sequences in regions that have been proposed to contain heme

regulatory (or responsive) motifs (HRM) centered at Cys127, Cys265, and Cys282.[143]

Interactions between heme and the HRM have been proposed to control the activity or

stability of several regulatory proteins, including the transcriptional repressor Bach1 and

eukaryotic initiation factor-2α kinase. Therefore, these motifs could regulate HO-2

activity and provide additional heme binding sites that function to maintain intracellular

free heme level and act as a sink for CO.

Chapter 1

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HO-1 is the inducible form of heme oxygenase with a high level of expression in

the spleen and other tissues that degrade senescent red blood cells, including specialized

reticuloendothelial cells of the liver and bone marrow.[135] In most other tissue not

directly involved in erythrocyte or haemoglobin metabolism, HO-1 typically occurs at

low to undetectable levels under basal conditions but responds to rapid transcriptional

activation by diverse chemical and phycial stimuli. HO-2 is constitutively expressed.

HO-2 is expressed in abundance in the testes, but the protein is also found ubiquitously

in other tissues including central nervous system, vasculature, and the gut.[2] Both HO-

1 and HO-2 catalyze the same biochemical reaction with similar substrate specificity and

co-factor requirements. However, in a comparative analysis of rat HO-1 and HO-2,

differential properties with respect to enzyme kinetics and substrate Km values have

been reported, as well as differences in thermostability.[168, 169]

Figure 1.8 Structural organization of the human HO-1 (top) and HO-2 (bottom) gene. Source: Shibahara et al. (2007) Hypoxia and heme oxygenases: oxygen sensing and regulation of expression.

Chapter 1

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1.7.3 Regulation of heme oxygenase protein expression

Protein expression of HO-1 and HO-2 is differentially regulated. Human HO-1

is mainly regulated at the level of transcription. Its promoter contains one copy of the

functional Maf recognition element (MARE) immediately downstream from the

cadmium-responsive element (Figure 1.8).[170] HO-1 expression is regulated by

members of the small Maf family, which are basic region leucine zipper proteins that

can function as transcriptional activators or repressors. NF-E2-related factor-2 functions

as a transcriptional activator of HO-1 by forming a heterodimer with a member of the

Maf family, whereas Bach1 heterodimerizes with MafK to inhibit HO-1

expression.[171, 172]

Analysis of the genomic sequences 5’ to the rat and human HO-2 genes reveals

no regulatory elements corresponding to transcription factors known to participate in the

hypoxic or other cellular stress responses (Figure 1.8).[165, 173-175] The two

noticeable features that exist in the organization of the human HO-2 gene are the

presence of a potential bidirectional promoter and a large intron 1 of ~30 kb.[176] The

HO-2 gene and the gene encoding HSCARG of unknown function appear to share a

common promoter region. Similar to other bidirectional promoters, the HO-2 gene

promoter lacks the TATA box and contains GC-rich sequences.[177] The exon 1 of the

HO-2 gene encodes the 5’untranslated region of the HO-2 mRNA. Diversity of HO-2 5’

UTR sequences has been documented previously in the rodent, suggesting the use of

alternate transcription initiation sites (and promoters).[174, 178]

Despite the lack of evidence for transcriptional control, HO-2 protein expression

is not entirely constitutive. Development stage-specific changes in HO-2 protein level

Chapter 1

- 30 -

have been reported and cigarette smoke increases the number of lung cells expressing

HO-2 protein.[179-181] HO-2 protein increases 1.3 fold in the heart of mice exposed to

hypoxia for 28 days compared with age-matched normoxic controls.[182] Spatial and

temporal dissociation between HO-2 protein and mRNA expression have been noted in

rodent brain and testis and changes in HO-2 protein levels have been detected without

changes in HO-2 mRNA during hypoxia.[165, 173-175] These findings strongly

suggest that HO-2 protein expression is regulated mainly at the posttranscriptional level.

Therefore, posttranscriptional regulation of HO-2 protein expression during prolonged

hypoxic exposure was investigated in the studies described in the current work.

Chapter 1

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1.8 Thesis Objectives

Aim 1:

• To determine the role that altered matrix metalloproteinase-2 expression and

activity play in the changes in contractile responses observed in systemic conduit

and resistance arteries after prolonged hypoxia in vivo

Hypothesis 1:

• Increased matrix metalloproteinase-2 activity potentiates contraction in systemic

conduit and resistance arteries after hypoxia in vivo by proteolytic activation of

big endothelin-1

Rationale:

• Endothelin-1 protein is increased in aorta of rats exposed to hypoxia without a

concomitant increase in preproET-1 mRNA expression.

• Increases in rat aortic ECE-1 expression, the activator of endothelin-1, could not

be detected

• Matrix metalloproteinase-2 and its activator, MT1-MMP have been shown to be

increased during hypoxic exposure.

Chapter 1

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Aim 2:

• To determine if hypoxia alters HO-2 expression through effects on protein

translation and whether HO-2 preserves endothelial cell viability during

concurrent hypoxic and oxidative stress in human endothelial cells.

Hypothesis 2:

• Heme oxygenase-2 translation is enhanced during hypoxia in human endothelial

cells.

Rationale:

• Enhanced HO-2 protein expression has been detected in rat aortic endothelium

without a concomitant increase in HO-2 mRNA expression.

• Discordance between HO-2 protein level and mRNA expression has previously

been identified.

Hypothesis 3:

• Endothelial HO-2 expression during hypoxia preserves endothelial viability.

Rationale

• Products of the HO-catalyzed reaction have properties that favours enhanced

endothelial cell survival.

• HO-2 is the predominate HO isoform expressed in the endothelium.

• HO-1 protein and mRNA expression are decreased by hypoxia in endothelial

cells.

Chapter 2

- 33 -

CHAPTER 2

Induction of Matrix Metalloproteinase-2 Enhances Systemic Arterial

Contraction After Hypoxia

Chapter 2

- 34 -

2.1 Introduction

Hypoxia is frequently observed in patients with cardiopulmonary diseases and in

normal subjects at high altitude. Studies done to date have primarily focused on the

short term systemic circulatory responses which redistribute blood flow [183] and

enhance the capacity for oxygen extraction [184] in these conditions. As the duration of

hypoxia increases, however, systemic vascular smooth muscle and endothelial function

are impaired, limiting the efficacy of the acute responses [61, 62, 64, 185] while

concurrent structural remodelling plays an increasing role in maintaining the balance

between oxygen delivery and metabolic demand [183]. Although the clinical and

physiological relevance of these longer term effects on vascular function are being

increasingly recognized [186], the mechanisms that mediate them remain unknown.

Matrix metalloproteinase-2 (MMP-2) is a zinc-dependent proteinase secreted by

both endothelial and smooth muscle cells, and its expression is increased in regions of

matrix turnover and remodelling [187]. Recently, it was discovered that MMP-2 can

mediate the posttranslational modification of several vasoactive peptides [96, 97, 188],

suggesting that it has a vasoregulatory role as well. MMP-2 protein and mRNA levels

are increased after hypoxic incubation in endothelial cells [83]. Its activating protease,

membrane type 1-matrix metalloproteinase (MT1-MMP), and its endogenous inhibitors,

tissue inhibitors of matrix metalloproteinase (TIMPs), are also oxygen-regulated in some

cell types [189-191]. If hypoxia induces a functionally significant increase in MMP-2

expression in systemic arteries, modulation of MMP-2 activity may, in addition to

mediating structural adaptations, contribute to the changes in vascular tone that occur

during prolonged reductions in oxygen delivery. This study was, therefore, carried out

Chapter 2

- 35 -

to determine the role that altered MMP-2 expression and activity play in the changes in

contractile responses observed in systemic conduit and resistance arteries after

prolonged hypoxia in vivo.

Chapter 2

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2.2 Materials and Methods

Exposure to Hypoxia: All protocols were in compliance with standards set by the

Canadian Council on Animal Care and were approved by the institutional animal care

committee. Male Sprague-Dawley rats (200–250 g) and C56/B16J mice (20-25 g) were

placed in a Plexiglas chamber into which the flow of air and nitrogen was controlled

independently. In preliminary experiments, the arterial PO2 averaged 38 Torr (range

35–42 Torr) in rats breathing a gas mixture containing 10% O2 [64] and 38.1 Torr

(range 35-40 Torr) in mice breathing 8% O2,

Rats and mice exposed to hypoxia breathed gas mixtures containing 10% or 8%

oxygen, respectively for 16 h, 48 h, or 7 d. Normoxic control animals breathed room air

under otherwise identical conditions. At the end of the exposure period, rats were

decapitated and mice scarified by cervical dislocation. Thoracic aortae were excised, cut

into 4 mm segments and mounted in tissue bath myographs (Radnoti Glass Technology

Inc.), frozen in liquid nitrogen, or fixed in 10% paraformaldehyde. Rat mesenteric

arteries (100-200 µm internal diameter) were either mounted in wire myographs (Living

Systems) or frozen in liquid nitrogen for later protein extraction.

Chemicals/Antibodies: The cyclic peptide MMP-2/9 antagonist, CTTHWGFTLC

(CTT) was purchased from Calbiochem. Polyclonal MMP-2- and TIMP-1 to 3-specific

antibodies and monoclonal MT1-MMP-specific antibody were purchased from

Chemicon. Polyclonal TIMP-4-specific antibody was obtained from Biomol Research

Laboratories. Mca-RPPGFSAFK(Dnp)-OH and Mca-PLGL-Dpa-AR-NH were from R

& D systems. ECE-1-specific polyclonal antibody was from Zymed Laboratories.

Histostain and PicoPureTM RNA Isolation Kit were purchased from Arcturus Bioscience

Chapter 2

- 37 -

Inc. SYBR green universal master mix was from Applied Biosystems. Primers and

Superscript II were from Invitrogen. All other reagents were from Sigma.

Rat Studies

Aortic and Mesenteric Artery Contractile Responses: Rat aortic segments were

mounted in tissue bath myographs and mesenteric arterial segments in wire myographs

containing Krebs-Henseleit solution (KHS) composed of (in mmol/l) 120 NaCl, 25

NaHCO3, 11.1 glucose, 4.76 KCl, 1.18 MgSO4, 1.18 KH2PO4, 2.5 CaCl2 aerated with

95% O2-5% CO2 at 37°C. Thoracic aortae and mesenteric arteries were equilibrated in

warmed aerated KHS for 1 hour under a resting tension of 2 g or 500 mg, respectively,

before drug-induced changes in tension were monitored. When necessary, the

endothelium was removed by gentle abrasion of the luminal surface. The failure of

acetylcholine (1 µmol/l) to elicit relaxation after contraction with phenylephrine (1

µmol/l) was taken as evidence of functional endothelial ablation [185].

Redistribution of blood flow during hypoxia is mediated by neurohumoral

stimulation of α-adrenoceptors [76]. Cumulative concentration-response curves (CRCs)

for the α1-adrenoceptor agonist phenylephrine (PE, 1 nmol/l to 10 µmol/l) were,

therefore, generated in endothelium-intact aortic rings. MMP-2 cleaves big ET-1 to

release the potent vasoconstrictor ET-1[1-32] [96]. Since big ET-1 itself has minimal

biological activity prior to proteolytic activation and since ECE-1 protein levels and

ECE activity are unchanged after hypoxia (see below), the contractile response to big

ET-1 was used as a bioassay for changes in vascular MMP-2 activity. To eliminate the

confounding influence of endogenous endothelium-derived ET-1 [96], CRCs for rat big

Chapter 2

- 38 -

ET-1 (1 nmol/l to 300 nmol/l) were generated in endothelium-denuded aortic rings from

normoxic rats and rats exposed to hypoxia for 7 d after 45 min incubation with, and in

the continuous presence of CTT (30 µmol/l) or vehicle.

To assess the effect of hypoxia on contractile response of resistance vessels,

CRCs for PE (10 nmol/l to 100 µmol/l) were generated in mesenteric arteries from rats

exposed to normoxia or hypoxia for 7 d after 30 min incubation with and in the

continuous presence of CTT (10 µmol/l) or vehicle.

Immunohistochemistry: Paraffin embedded sections (5 µm) of rat aortae from

normoxic rats and rats exposed to hypoxia for 7 d were analyzed using MMP-2 specific

polyclonal antibodies as described [185]. Slides processed in an identical manner,

except incubated with non-specific rabbit IgG instead of primary antibody, served as

negative controls.

Western Blots: Thoracic aortic proteins from rats exposed to normoxia or to

hypoxia for 16 h, 48 h, and 7 d, and mesenteric arteries proteins from rats exposed to

normoxia or to hypoxia for 7 d were extracted in 1% SDS, 0.001 mol/l sodium

orthovanadate, and 0.01 mol/l Tris (pH 7.4). After protein concentrations in aortic and

mesenteric arterial extracts were determined by the Lowry method, total proteins (60 µg

for MMP-2 and MT1-MMP, 40µg for TIMPs 1-4, 100 µg for ECE-1) were resolved by

4-12% SDS-PAGE (Helixx Technologies Inc) and transferred to nitrocellulose. MT1-

MMP membranes were blocked in 3% milk-0.1% tween tris-buffered saline (TTBS).

All other membranes were blocked in 5% milk-TTBS. Membranes were then incubated

for 3 h at room temperature with goat polyclonal anti-MMP-2 (1:100), 1 hour at room

Chapter 2

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temperature with rabbit polyclonal anti-TIMP-1 (1:2500) or anti-TIMP-4 (1:10000), or

overnight at 4°C with rabbit polyclonal anti-ECE-1 (1:400), anti-TIMP-2 (1:2500), anti-

TIMP-3 (1:2500), or mouse monoclonal anti-MT1-MMP (1:400). Immunoblots were

probed with horseradish peroxidase (HRP)-donkey anti-goat IgG (1:4000 for MMP-2) or

HRP-anti-rabbit IgG (1:4000 for ECE-1, TIMP-1, TIMP-2, TIMP-3, TIMP-4) and

visualized by enhanced chemiluminescence (Amersham Biosciences). HRP-goat anti-

mouse IgG (1:1000) was used to probe for MT1-MMP and the resulting bands were

visualized by chemiluminescence (Sigma). Bands were quantified by densitometry.

Samples from normoxic and hypoxic groups were paired on each gel to control for inter-

experimental variation. Protein loading and transfer efficiency were corroborated

following transfer, using full-lane densitometry of the Ponceau red-stained membranes.

Gelatin Zymography: Aortae from normoxic rats and rats exposed to hypoxia for

16 h, 48 h, and 7 d were extracted with 10 mmol/l Tris-HCl (pH 7.5) extraction buffer.

Zymography was performed using 7.5% SDS-PAGE with co-polymerized gelatin (2

mg/ml) as substrate. At the end of each run, gels were washed with 2.5% Triton X-100

and incubated for 48 h in an enzyme assay buffer (50 mmol/l Tris, pH 7.0, 5 mmol/l

CaCl2, 0.15 mol/l NaCl, 0.05% Na3N) to allow for the development of enzyme activity

bands. Gels were stained with 0.05% Coomassie brilliant blue G-250 in a mixture of

methanol: acetic acid: water (2.5:1:6.5) and de-stained in 4% methanol with 8% acetic

acid. The gelatinolytic activities were detected as transparent bands against the

background of Coomassie brilliant blue-stained gelatin. Gels were scanned using Fluor-

S Multi-Imager (Bio-Rad) and analyzed for pro- and activated MMP-2 (72 and 64 kDa

bands, respectively).

Chapter 2

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MMP and ECE Activity: To further ensure that the change in the response to big

ET-1 was not attributable to a change in endothelin converting enzyme (ECE) activity,

total MMP and ECE activities were measured in aortae from normoxic rats and rats

exposed to hypoxia for 7 d. Thoracic aortic proteins (50 µg) from normoxic rats and rats

exposed to hypoxia for 7 d were incubated for 1 hour at 37° C with either 20 µmol/l of

fluorogenic MMP substrate Mca-PLGL-Dpa-AR-NH [192] in 100 µl of reaction

mixture (pH 7.5) composed of (in mmol/l): 50 Tris-HCl, 150 NaCl, and 1 CaCl2 or with

20 µmol/l of fluorogenic ECE substrate Mca-RPPGFSAFK(Dnp)-OH [193] in 100 µl of

reaction mixture (pH 6.0) composed of (in mmol/l): 100 MES and 200 NaCl. Blanks

containing the substrate dissolved in assay buffer were analyzed in parallel. Increases in

fluorescence as a result of substrate cleavage were continuously measured using a

fluorescence plate reader (Thermo Labsystems). Samples were run in triplicate and final

values were derived by subtracting the blank reading from the raw data.

Aortic MMP-2, MT1-MMP, TIMPs 1 to 4 mRNA levels: Total aortic RNA was

isolated as previously described [185]. In addition, pure populations of aortic

endothelial cells were isolated from aortae of rats exposed to normoxia or hypoxia for 7

d using the Hautchen technique [194]. Pure cell populations of vascular smooth muscle

cells from immediately below the endothelial cell layer (subendothelial VSMC) or from

deep within the media of the vessel (deep medial VSMC, Figure 2.7) were obtained

using the PixCell IITM Laser Capture Microdissection System according to the

manufacturer’s instructions and RNA was extracted using the PicoPureTM RNA Isolation

Kit. Aortic levels of specific mRNAs were measured by quantitative real-time RT-PCR

Chapter 2

- 41 -

(ABI PRISM 7900 HT, Applied Biosystems and the SYBR Green detection system

[185]) using the following primers: MMP-2 (sense 5’-ACA CTG GGA CCT GTC ACT

CC-3’, antisense 5’-ACA CGG CAT CAA TCT TTT CC-3’); MT1-MMP (sense 5’-

TCC TGC TCC CCC TGC TCA CG, antisense 5’-GTG ACT GGG GTG AGC GTT

GTG T-3’); TIMP-1 (sense 5’-GGA TAT GTC CAC AAG TCC CAG AAC C-3’,

antisense 5’-TTA TGC CAG GGA ACC AGG AAG C-3’); TIMP-2 (sense 5’-GGC

CAA AGC AGT GAG CGA GAA -3’, antisense 5’-GGA GGG GGC CGT GTA GAT

AAA T-3’); TIMP-3 (sense 5’-CCC TTT GGC ACT CTG GTC TAC ACT A-3’,

antisense 5’- AGG CCA CAG AGA CTT TCA GAG GCT-3’); and TIMP-4 (sense 5’-

TAC ACG CCA TTT GAC TCT TCT CTC TG-3’, antisense 5’-CCT CCC AGG GCT

CAA TGT AGT TG-3’). 18S (sense 5’-GAC GAT CAG ATA CCG TCG TAG TTC-

3’, antisense 5’-GTT TCA GCT TTG CAA CCA TAC TCC-3’) and TATA binding

protein (sense 5’-CCC CTA TCA CTC CTG CCA CAC C-3’, antisense 5’-CGC AGT

TGT TCG TGG CTC TCT T-3’) transcripts were used as control genes for

normalization and the average change in the target gene with respect to 18S and TATA

binding protein was determined.

Studies in MMP-2-/- and MMP-2+/+ mice

Animals: To corroborate the results of the pharmacological studies described

above, aortic contraction was assessed in mice deficient in MMP-2. The MMP-2+/- mice

on the C57/Bl6J background previously described [195] were interbred to generate

MMP-2 knockout (MMP-2-/-) and littermate control (MMP-2+/+) groups. Mouse

genotypes were assessed by polymerase chain reaction of genomic DNA. Primers for

wild-type alleles were located in exon-1 (5’-CAA CGA TGG AGG CAC GAG TG-3’

Chapter 2

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and 5’-GCC GGG GAA CTT GAT CAT GG-3’), and primers for the mutant allele were

located in the neo cassette (5’-CTT GGG TGG AGA GGC TAT TC-3’ and 5’-AGG

TGA GAT GAC AGG AGA TC-3’).

Aortic Contractile Responses: Mouse aortic segments were equilibrated in

warmed KHS aerated with 95% O2-5% CO2 at 37°C for 1 hour under a resting tension of

1g before drug-induced changes in tension were monitored. As in rat aortic segments the

endothelium was removed by lumenal abrasion and the success of endothelial ablation

assessed by acetylcholine-induced relaxation of phenylephrine-induced contraction

[185]. CRCs were generated for PE (1 nmol/l to 10 µmol/l) in endothelium-intact

thoracic aortic rings, and the contractile response to human big ET-1 (100 nmol/L) was

determined in endothelium-denuded aortic rings from MMP-2+/+ and MMP-2-/- mice

exposed to normoxia or hypoxia for 7d.

Data Analysis

Results are presented as mean ± S.E.M. for n number of animals with P<0.05

representing statistical significance. Paired means were compared 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, individual means were compared post-hoc using

Dunnet's test.

Chapter 2

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2.3 Results

Rat Studies

Aortic and Mesenteric Artery Contractile Responses: Figure 2.1A shows the

concentration-response relationship for PE in endothelium-intact aortic rings from

normoxic rats and rats exposed to hypoxia for 7 d in the presence or absence of CTT (30

µmol/l). Inhibition of MMP-2 decreased the maximum tension generated during PE-

induced contraction in aortic rings from rats exposed to hypoxia (Table 1), but had no

effect in rings from normoxic rats. Figure 2.1B illustrates the response of endothelium-

intact rat mesenteric artery segments to PE in the presence or absence of CTT. As in

aortic rings, CTT had no effect on the response of mesenteric arteries from normoxic

rats but reduced contraction in those from hypoxia-exposed animals (Table 1).

The contractile responses to big ET-1 in endothelium-denuded aortic rings from

rats exposed to normoxia or hypoxia for 7 d in the presence or absence of CTT (30

µmol/l) are illustrated in Figure 2.1C. Maximal contractions achieved in rings from

hypoxic animals were higher than those in the normoxic controls (Table 1). This

hypoxia-dependent augmentation of big ET-1 mediated contraction was abolished by

CTT (Table 1). At this concentration, CTT had no effect on ET-1-induced contraction

in aortic rings from either normoxic or hypoxia-exposed rats (data not shown),

indicating that its effect is mediated by inhibition of big ET-1 conversion.

Chapter 2

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Figure 2.1 (A) Concentration-response relationships for phenylephrine (PE) in the

presence and absence of the MMP-2/9 inhibitor CTTHWGFTLC (CTT, 30 µmol/l) in

endothelium-intact aortic rings from normoxic rats and rats exposed to hypoxia for 7 d

(n = 8 per group). (B) Concentration-response relationships for PE in the presence and

absence of CTT (10 µmol/l) in endothelium-intact mesenteric arteries from normoxic

rats and rats exposed to hypoxia for 7 d (n = 5-6 per group).

A Endothelium-Intact Rat Aorta

B Endothelium-Intact Rat Mesenteric Arteriole

Chapter 2

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Figure 2.1 (C) Concentration response relationships for big ET-1 in the presence and

absence of CTT (30 µmol/l) in endothelium-denuded aortic rings from normoxic rats and

rats exposed to hypoxia for 7 d (n = 10 per group). *P<0.05 for differences between

CTT-treated and -untreated group. †P<0.05 for difference from corresponding

normoxic control group.

C Endothelium-Denuded Rat Aorta

Chapter 2

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Table 1: Effect of MMP inhibition on maximum contraction and EC50 values

during PE and big ET-1 big ET-1-induced rat aortic contraction.

Values are means ± SEM; *P<0.05 for differences between CTT-treated and -untreated

group. †P<0.05 for difference from corresponding normoxic control group.

Maximum Tension (g/mg dry weight)

Rat Vessels Treatment Nor. Hyp. (7 d)

PE KHS 2.89 ± 0.10 3.41 ± 0.21† Aortae

CTT (30 µmol/L) 3.11 ± 0.17 2.73 ± 0.19*

KHS 678.15 ± 52.20 911.93 ± 76.51† Mesenteric Arteries CTT (30 µmol/L) 748.95 ± 47.11 715.32 ± 101.21*

Big ET-1 KHS 1.35 ± 0.08 1.77 ± 0.10† Aorta

CTT (30 µmol/L) 1.31 ± 0.18 1.28 ± 0.12*

-log EC50, mol/L Rat

Vessels Treatment Nor. Hyp. (7 d) PE

KHS 7.412 ± 0.05 7.63 ± 0.09 Aortae CTT (30 µmol/L) 7.32 ± 0.09 7.43 ± 0.10

KHS 5.81 ± 0.07 5.79 ± 0.09 Mesenteric

Arteries CTT (30 µmol/L) 5.73 ± 0.08 5.64 ± 0.02 Big ET-1

KHS 7.25 ± 0.05 7.25 ± 0.03 Aorta CTT (30 µmol/L) 7.21 ± 0.08 6.92 ± 0.17

Chapter 2

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Immunohistochemistry: Figure 2.2 depicts representative immunohistochemical

staining for MMP-2 in aortic sections from normoxic rats (Figure 2.2A) and rats

exposed to hypoxia for 7 d (Figure 2.2C), along with the respective negative controls

(Figures 2.2B and 2.2D). Although MMP-2 protein was detected in both the intima and

media of the thoracic aorta from normoxic and hypoxic animals, staining was more

intense in the hypoxic group with no apparent inhomogeneity in its distribution across

the aortic wall.

Figure 2.2 Immunohistochemistry for MMP-2 on aortic sections from normoxic rats

and rats exposed to hypoxia for 7 d. Immunoreactivity (brown diaminobenzidine

staining) is apparent in the aortic endothelium and media layer of both the normoxic (A)

and hypoxic (C) groups, but not in the normoxic (B) or hypoxic (D) negative controls

(40X objective).

Chapter 2

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MMP-2 and ECE Protein and Activity Levels: Western analysis shown in Figure

2.3A indicates that rat aortic proMMP-2 (72 kDa) protein levels increased after

prolonged hypoxia. These differences reached statistical significance after 48 h and 7 d

of hypoxia. Activated MMP-2 (64 kDa) protein levels were also found to be elevated

with increasing duration of hypoxia, achieving statistical significance after 7 d. Aortic

MMP-2 activity, as determined by gelatin zymography, was also significantly greater

after 7 d of hypoxia compared to the normoxic control group (Figure 2.3B). No bands

corresponding to the expected molecular weight of MMP-9 were detected in these

samples, suggesting that MMP-2 is the predominant source of gelatinase activity.

Figure 2.3C illustrates that proMMP-2 protein is also increased in mesenteric arteries

from rats exposed to hypoxia for 7 d compared to the normoxic animals. Protein

concentrations obtained from these small vessels was insufficient to quantify levels of

the cleaved (activated) form. Aortic ECE-1 protein levels did not differ between

normoxic and hypoxia-exposed rats (data not shown).

The results of fluorometric assays of total MMP and ECE activities are presented in

figure 2.4. MMP activity was higher in aortae from hypoxia-exposed rats compared to

the normoxic group (Figure 2.4A), whereas ECE activity was unchanged (Figure 2.4B).

Chapter 2

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Figure 2.3 (A) Aortic MMP-2 protein levels in normoxic rats and rats exposed to

hypoxia for 16 h, 48 h, and 7 d (n = 9 per group). (B) Gelatin zymography showing the

levels of activated MMP-2 in aortae of normoxic rats and from rats exposed to hypoxia

for 16 h, 48 h, and 7 d (n = 6 per group). *P<0.05 for differences from corresponding

normoxic group.

Chapter 2

- 50 -

Figure 2.3 (C) MMP-2 protein levels in mesenteric arteries from normoxic rats and

rats exposed to hypoxia for 7 d (n = 8 per group). *P<0.05 for differences from

corresponding normoxic group.

Chapter 2

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Figure 2.4 MMP (A; n = 10) and ECE (B; n=9) activity in aortae from normoxic rats

and rats exposed to hypoxia for 7 d. *P<0.05 for differences from corresponding

normoxic group.

Chapter 2

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MT1-MMP and TIMPs Protein Levels: MMP-2 activity is regulated by its activator

protease MT1-MMP, and its tissue inhibitors TIMPs. Western analysis demonstrated

that rat aortic MT1-MMP protein levels increased progressively with increasing duration

of hypoxic exposure, reaching statistical significance after 7 d (Figure 2.5A). Although

aortic levels of TIMPs 1 to 4 exhibited an upward trend, these changes did not reach

statistical significance during the same period of hypoxic exposure (Figures 2.5B-2.5E).

Figure 2.5 Levels of MT1-MMP (A) proteins in aortae from normoxic rats and rats

exposed to hypoxia for 16 h, 48 h, and 7 d (n = 6 per group). *P<0.05 for differences

from corresponding normoxic group.

Chapter 2

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Figure 2.5 Levels of TIMP-1 (B) and 2 (C) proteins in aortae from normoxic rats

and rats exposed to hypoxia for 16 h, 48 h, and 7 d (n = 6 per group). *P<0.05 for

differences from corresponding normoxic group.

Chapter 2

- 54 -

Figure 2.5 Levels of TIMP-3 (D) and 4 (E) proteins in aortae from normoxic rats

and rats exposed to hypoxia for 16 h, 48 h, and 7 d (n = 6 per group). *P<0.05 for

differences from corresponding normoxic group.

Chapter 2

- 55 -

MMP-2, MT1-MMP and TIMPs mRNA Levels: Figure 2.6 illustrates MMP-2, MT1-

MMP and TIMPs 1 to 4 mRNA levels in aortae from normoxic rats and rats exposed to

hypoxia for 16 h, 48 h, or 7 d. After exposure to hypoxia for 7 d, MMP-2 and MT1-

MMP mRNA levels are increased compared to the normoxic control group. An increase

in levels of TIMPs -1 to -3 mRNA was observed after 7 d of hypoxia while TIMP-4

mRNA expression was upregulated at the earlier time points (16 h and 48 h) as well.

Chapter 2

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Figure 2.6 Levels of MMP-2 (A) and MT1-MMP (B) mRNAs in aortae from

normoxic rats and rats exposed to hypoxia for 16 h, 48 h, and 7 d (n = 7 per group).

*P<0.05 for differences from corresponding normoxic group.

Chapter 2

- 57 -

Figure 2.6 Levels of TIMP-1 (C) and TIMP-2 (D) mRNAs in aortae from normoxic

rats and rats exposed to hypoxia for 16 h, 48 h, and 7 d (n = 7 per group). *P<0.05 for

differences from corresponding normoxic group.

Chapter 2

- 58 -

Figure 2.6 Levels of TIMP-3 (E) and TIMP-4 (F) mRNAs in aortae from normoxic

rats and rats exposed to hypoxia for 16 h, 48 h, and 7 d (n = 7 per group). *P<0.05 for

differences from corresponding normoxic group.

Chapter 2

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MMP-2 and MT1-MMP mRNA Levels: To identify the cell type responsible for the

observed increase in MMP-2 and MT1-MMP expression, MMP-2 and MT1-MMP

mRNAs were quantified using RNA extracted from pure populations of aortic

endothelial cells, sub-endothelial vascular smooth muscle cells (VSMC) and deep

medial VSMCs (Figure 2.7A-C). MMP-2 (Figure 2.7D) and MT1-MMP mRNA levels

(Figure 2.7E) are increased in deep medial aortic VSMCs from rats exposed to hypoxia

for 7 d, whereas, no change was observed in endothelial or subintimal smooth muscle

cells.

Chapter 2

- 60 -

Figure 2.7 (A-C) Illustration of aortic regions where cells were collected for mRNA

extraction.

Chapter 2

- 61 -

Figure 2.7 MMP-2 (D) and MT1-MMP (E) mRNA levels in endothelial, sub-

endothelial VSMC, and deep medial VSMC from aortae of normoxic rats and from rats

exposed to hypoxia for 7 d (n = 6 per group). *P<0.05 for differences from

corresponding normoxic group.

Chapter 2

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Studies in MMP-2-/- and MMP-2+/+ Mice

Aortic Contractile Responses: Figure 2.8A presents the concentration-response

relationships for PE in endothelium-intact aortic rings from mice exposed to normoxia

or hypoxia for 7 d. In contrast to rats, hypoxia did not enhance the response to PE in

MMP-2+/+ mice possibly reflecting differences in the adaptive response to hypoxia

between the two species. Nevertheless, after hypoxic exposure, the effect of MMP-2

deletion in mice mimics the effect of MMP inhibition in rats in that, after hypoxia, the

maximum tensions generated during PE-induced contraction are reduced in MMP-2-/-

compared to their MMP-2+/+ littermate controls (Table 2). The responses of

endothelium-denuded aortic rings from MMP-2-/- and MMP-2+/+ mice to big ET-1 (100

nmol/l) are illustrated in figure 2.8B. Similar to the results obtained in rat aortae,

contraction to big ET-1 is greater in aortic segments from MMP-2+/+ mice exposed to

hypoxia than in segments from the corresponding normoxic control group (Table 2). In

contrast, the aortic response to big ET-1 in normoxic and hypoxia-exposed MMP-2-/-

mice do not differ.

Chapter 2

- 63 -

Figure 2.8 (A) Concentration-response relationship for phenylephrine in

endothelium intact aortic rings from MMP-2 deficient (MMP-2-/-) and littermate control

(MMP-2+/+) mice exposed to normoxia or hypoxia for 7 d (n = 8-9 per group). *P<0.05

for differences between MMP-2+/+ and MMP-2-/- groups. (B) Contractile response to big

ET-1 (100 nmol/l) in endothelium-denuded aortic rings from MMP-2+/+ and MMP-2-/-

mice exposed to normoxia or hypoxia for 7 d (n = 9 for each group). *P<0.05 for

difference from corresponding normoxic group.

Chapter 2

- 64 -

Table 2: Maximum contraction and EC50 values during PE- induced contraction

and response to big ET-1 (100 nmol/l) in MMP-2-/- and MMP-2+/+ mice.

Values are means ± SEM; *P<0.05 for differences between MMP-2+/+ and MMP-2-/- groups. †P<0.05 for difference from corresponding normoxic control group.

Maximum Tension (mg) Mouse Vessels Genotype Nor. Hyp. (7 d) PE

+/+ 462.11 ± 73.77 429.95 ± 51.27 Aortae -/- 444.42 ± 61.66 328.84 ± 31.17*

Big ET-1 +/+ 67.49 ± 11.32 172.62 ± 25.33† Aortae -/- 98.50 ± 10.30 125.18 ± 12.57

-log EC50, mol/L Mouse Vessels Genotype Nor. Hyp. (7 d) PE

+/+ 7.85 ± 0.17 8.03 ± 0.15 Aortae -/- 7.50 ± 0.17 8.04 ± 0.24

Big ET-1 +/+ n/a n/a Aortae -/- n/a n/a

Chapter 2

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2.4 Discussion

The results of this study show that after prolonged hypoxia: 1) MMP-2 inhibition

(rat) or deletion (mouse) reduces aortic and mesenteric arterial contraction to

phenylephrine; 2) the aortic contractile response to big ET-1 is enhanced in rats and

mice through an MMP-2-dependent mechanism; 3) MMP-2 protein levels in rat aortae

and mesenteric arteries, and MMP activity in rat aortae are increased; 4) aortic MT1-

MMP protein levels are increased; 5) aortic MMP-2, MT-1 MMP and TIMPs 1 to 4

mRNA levels are increased; and 6) the increase in rat aortic MMP-2 and MT1-MMP

mRNA expression is localized to the deep medial vascular smooth muscle.

The 23 MMPs identified to date are divided, based on substrate preference, into

collagenases, gelatinases, stromelysins, matrilysins and membrane-type MMPs. MMP-2

and MMP-9 are the gelatinases which efficiently degrade collagen type IV [82] and,

hence, are involved in the restructuring of vascular basement membranes. A broader

biological role for MMP-2 has become apparent with the recognition that its substrates

also include a number of vasoregulatory peptides [96, 188, 196-199]. Inactivation of

vasodilators (calcitonin gene-related peptide [97] and adrenomedullin [188]) and

activation or release of vasoconstrictors (big endothelin-1 (ET-1) [96], heparin binding

epidermal growth factors (HB-EGF) [197, 199], and integrin binding RGD peptides

[200]) all contribute to its vasoactive effects. Although the relative importance of each

of these pathways and the possible existence of others remains to be explored, our

present results emphasize that the net effect of increased MMP-2 activity in the systemic

circulation, as occurs after hypoxia, is to potentiate vascular smooth muscle contraction.

Chapter 2

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In regions affected by arterial insufficiency, MMP-2-mediated enhancement of

vascular contraction may be maladaptive since it will exacerbate ischemic injury.

During global hypoxia, however, the primary defensive vascular response depends on

the capacity of the adrenergic nervous system to regulate the regional distribution of

blood flow and oxygen extraction [76]. In previous studies in rats, systemic arterial

smooth muscle contraction to adrenoceptor stimulation is impaired after 48 hours of

hypoxia due to induction of heme oxygenase and nitric oxide synthase expression and

inhibition of myosin phosphorylation [61-63, 74, 185]. Hence, the ability to target

oxygen delivery to areas of greatest metabolic demand [76, 78, 183] is impaired. Our

present results indicate that after 7 days of hypoxia, rat aortic and mesenteric arterial

contraction are increased and that this is concomitant with and dependent on enhanced

MMP-2 activity. In this setting, therefore, upregulation of vascular MMP-2 provides a

mechanism to reinforce adrenergic regulation in the period during which maintenance of

oxygen delivery to vital organs is mediated by changes in vascular tone prior to the

structural change on which the redistribution of blood flow will ultimately depend.

In the rat aorta, MMP-2 and endothelin converting enzyme-1 (ECE-1) are the

major enzymes that convert big ET-1, the inactive prohormone, into the active

vasoconstrictor ET-1. Activation of big ET-1 by ECE-1 releases ET-1 [1-21] whereas

cleavage at Gly32-Leu33 by MMP-2 generates an isopeptide ET-1 [1-32] with enhanced

potency at the smooth muscle ETA receptor [96]. In the current study, the maximum

contraction that could be elicited by big ET-1 was increased after hypoxia. Since this

was reversed by MMP inhibition or MMP-2 deletion, MMP-2-mediated formation of the

Chapter 2

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more potent isopeptide ET-1 [1-32] appears to be a prominent pathway for big ET-1

conversion during prolonged hypoxia.

MMP-2 is secreted as a zymogen whose activity is regulated by its activating

protease MT1-MMP [187] and the endogenous tissue inhibitors of MMPs (TIMPs) [82].

An increase in MT1-MMP relative to the TIMPs is, therefore, requisite to any significant

enhancement of bioactivity. The possibility that MT1-MMP activity may be regulated

by an oxygen-sensitive mechanism has been suggested previously. MT1-MMP protein

is increased after hypoxic incubation in HepG2 cells and in the myocardium after

ischemia-reperfusion injury [201, 202] and the intracellular proprotein convertase furin,

responsible for its activation, is transcriptionally regulated by Hypoxia Inducible Factor-

1 [201]. Our current results confirm the functional relevance of these findings in the

systemic circulation and demonstrate that upregulation of MT1-MMP in the aorta

parallels the expression of its substrate, MMP-2. We also considered the possibility that

TIMPs may be oxygen regulated in order to provide an additional level of control. In

cultured cells, TIMP-1 and -2 have been observed to increase [189, 203, 204], decrease

[83, 205, 206], or remain unchanged [207] after hypoxic incubation. Our results confirm

that aortic levels of TIMPs mRNAs are sensitive to oxygen tension in vivo. However, in

contrast to MT1-MMP and MMP-2, changes in TIMP protein levels did not reach

statistical significance. Since MMP-2 activity was increased, this suggests that elevated

TIMP expression is insufficient to offset the increase in MT1-MMP and MMP-2.

Nevertheless, the fact that the expression of these endogenous inhibitors is hypoxia

inducible suggests that they may provide an important negative feedback mechanism in

some conditions.

Chapter 2

- 68 -

The genes encoding MMP-2 and MT1-MMP contain consensus binding elements

for a number of hypoxia inducible transcription factors [191, 208], and their

transcriptional regulation by oxygen tension would be anticipated. Nevertheless, the

results of previous studies in cultured cells, are in conflict on this point [83, 203, 209-

211]. Our present results provide both pharmacological and biochemical evidence that,

in vivo, the expression of MMP-2 and MT1-MMP is upregulated in the systemic

circulation after hypoxia as a result of increases in their steady state mRNA levels and

that this change in systemic vascular cell phenotype is functionally relevant. Vascular

cells experience a broad range of oxygen tensions under physiological conditions. In the

aorta, oxygen concentrations from 11.2% (90 mm Hg) at the luminal surface to 2.2% (20

mmHg) at a depth of 150µm [212] are reported and longitudinal gradients of similar

magnitude occur in the microcirculation [213]. Since the severity of the hypoxic

stimulus varies significantly across the aortic wall, production of MMP-2 and MT1-

MMP would also be expected to demonstrate regional heterogeneity.

Immunohistochemical analysis may be confounded because MMP-2 is secreted and,

hence, distributed in the intracellular space across the aortic wall (see Figure 2.2).

Accordingly, we evaluated the regional expression of MMP-2 and MT1-MMP mRNA

and found that these transcripts are selectively enriched in VSMC located deep within

the aortic media, the most hypoxic region of the tissue [214]. This supports our

hypothesis that the expression of MMP-2 correlates with the severity of the hypoxic

stimulus and suggests that proteolytic activation of MMP-2 proenzyme occurs as it is

produced in the deep medial layer.

Chapter 2

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It is well recognized that changes in vascular tone precede the structural

alterations that occur when changes in blood flow persist chronically [200, 215, 216].

Such remodelling of the circulation is important in adapting the mature circulation to

chronic changes in tissue perfusion as well as arterial growth to meet the changing blood

flow demands of developing peripheral tissues. A role for MMP-2 in this process is

supported by observations that MMP-2-/- mice demonstrate impaired angiogenesis [90]

and that inhibitors of MMP reduce the pathological structural remodelling that

accompanies monocrotaline-induced pulmonary arterial hypertension [217]. Hypoxia is

a potent stimulus for both changes in vascular tone and structural remodelling in the

systemic circulation [79, 218]. The results of the current study indicate that vascular

MMP-2 levels and activity are tightly regulated by oxygen tension and, hence, represent

a pivotal regulatory pathway by which the acute vascular responses associated with

hypoxia may be integrated with the longer term structural changes in both conduit and

resistance arteries. Further investigation to determine the specific roles of MMP-2 and

each of its newly identified substrates will advance our understanding of the

pathobiology of this process in cardiopulmonary diseases and offer new therapeutic

targets in their management.

Chapter 3

- 70 -

During exposure to prolonged hypoxia, maintenance of vascular reactivity is

essential to ensuring adequate oxygen supply to vital organs.[10, 76, 184, 219] In the

previous chapter, increased vascular MMP-2 was demonstrated to induce

vasoconstriction in systemic conduit and resistance arteries of rats exposed to hypoxia

for 7 days. This is mediated through cleavage of big ET-1 by vascular MMP-2 to

release the vasoconstrictor ET-1[1-32].

Given the multifaceted actions of ET-1 and consequences of excessive ET-1

production, numerous mechanisms have evolved to modulate the local bioavailability

and potency of ET-1. Another enzyme in the vasculature that modulates ET-1

bioavailability and potency is HO-2. HO-2 catalyzes the degradation of heme to release

CO, biliverdin/bilirubin, and ferrous iron. Biliverdin/bilirubin possesses the capacity to

suppress intracellular concentrations of the reactive oxygen species (ROS) that regulate

ET-1 precursor mRNA expression through its antioxidant properties. CO mimics many

NO functions including cGMP-dependent and –independent inhibition of agonist-

induced vascular smooth muscle contraction. Govindaraju et al. demonstrated that

enhanced endothelial HO-2 protein expression reduces aortic reactivity in rats exposed

to hypoxia. Taken together, these findings suggest that HO-2 is another important

enzyme in regulating vascular function in systemic vessels during hypoxia.

Accordingly, hypoxic regulation of HO-2 protein expression and the physiological

effects of HO-2 activity during exposure to prolonged hypoxia are investigated.

Chapter 3

- 71 -

CHAPTER 3

Enhanced translation of HO-2 transcripts preserves human endothelial

cell viablility during prolonged hypoxia

Chapter 3

- 72 -

3.1 Introduction

Hypoxia is frequently observed in patients with shock, cardiopulmonary diseases

and in normal subjects at high altitudes. The compensatory mechanisms that preserve

blood flow to vital organs under these conditions are, in part, dependent on the release of

endothelium derived vasoregulatory factors.[64, 79, 185] During prolonged exposure to

hypoxia, endothelial function is impaired due to changes in endothelial phenotype and

cell death, and the adaptive responses that they mediate are compromised.[61, 64, 79]

Investigation of mechanisms that preserve endothelial cell survival and function in this

setting is, therefore, needed in order to develop therapeutic strategies to mitigate the

effects of hypoxia in patients with disorders associated reduced oxygen delivery.

Heme oxygenases (HO) are the rate limiting enzymes in the heme catabolic

pathway that cleaves heme to release carbon monoxide (CO), biliverdin, and ferrous

iron.[135] These products possess anti-apoptotic, anti-oxidant, and anti-inflammatory

properties, and so, may ameliorate the deleterious effects of hypoxia on endothelial

function. HO-1 and HO-2 are the heme oxygenase isoforms identified in the

endothelium. HO-1 expression is suppressed during hypoxia in human endothelial cells

and this is mediated by increased expression of the transcription repressor Bach1.[172]

Although a protective role for HO-2 has been suggested in other conditions associated

with impaired endothelium-dependent vasoregulation (diabetes and ischemia), the effect

of hypoxia on endothelial HO-2 expression and its functional role are unknown.[74, 137,

157] The current study was, therefore, carried out to test the hypothesis that HO-2

expression is oxygen regulated in human endothelial cells and to determine whether it

plays a role in preserving endothelial cell viability during hypoxic stress.

Chapter 3

- 73 -

3.2 Materials and Methods

Chemicals and Reagents: The Protein and RNA isolation system (PARIS) kit and

SYBR green universal master mix was purchased from Applied Biosystems (Foster City,

CA). The cDNA Synthesis Kit and DC Protein Quantifiation kit were from BioRad

laboratories (Hercules, CA). Anti-HO-1 and HO-2 antibodies were from Assay

Designs (Ann Arbor, MI). HO-2 siRNA, scrambled siRNA, and siRNA transfection

reagents were from Santa Cruz Biotechnology Inc (Santa Cruz, CA). 3H-uridine, 3H-

leucine and 35S-methionine were purchased from Perkin Elmer Life Science (Waltham,

MA). Carboxy-5-(6)-chloromethyl-2’,7’-dichlorodihydrofluorescein diacetate (carboxy-

H2DCFDA), MitoProbe JC-1 Assay Kit, and custom primers were from Invitrogen

(Carlsbad, CA). Annexin V–FLUOS staining kit was purchased from Roche and the

CaspACE FITC-VAD-FMK in situ marker were from Promega (Madison, WI). All

other chemicals were from Sigma (St. Louis, MO).

Cell Culture Studies: Pooled human umbilical vein endothelial cells (HUVEC) and

human aortic endothelial cells (HAEC) were purchased from Lonza (Basel, Switzerland)

and cultured in EGM-2 medium according to manufacture’s instructions. Human blood

outgrowth endothelial cells were derived from peripheral blood as described.[220]

Healthy volunteers underwent a mononuclear cell collection (100 ml, 3-8% hematocrit)

procedure on a Cobe Spectra (Gambro, BCT, Denver CO). Samples were cultured in

EGM-2MV medium containing 20% human serum in tissue culture flasks pre-coated

with fibronectin (10µg/mL) for 7 to 10 days to obtain pure population of blood

outgrowth EC. After which, these cells were cultured for an additional 3 passages

Chapter 3

- 74 -

before being used in experiments. HUVEC at passages 3-5 and HAEC at passages 6-7

were used.

For siRNA transfections, HUVEC (105 cells/cm2) were seeded in antibiotic free

EGM-2 media in 60 mm dishes for 16-24 hours and transfected with human HO-2

siRNA (sc-35556) or non-specific control siRNA (sc-37007) using siRNA Transfection

Reagent following the manufacturer’s instructions. Transfected cells were replated in

either 60 mm dishes or 12 well plates after 24 h. Cells exposed to hypoxia were grown

to 70% confluence and transferred, after changing the medium, to a humidified Plexiglas

chamber maintained at 37°C and continuously flushed with gas composed of 1% O2/5%

CO2/balanced N2. Normoxic control cells were exposed to air/5% CO2/balanced N2

under otherwise identical conditions.

Quantitative Real Time PCR: Total RNA was extracted from HUVEC exposed to

either normoxia or hypoxia for 16 or 48 h using the PARIS kit and reverse transcribed

(1µg) with the cDNA synthesis kit containing random primers. All quantitative RT-

PCR analyses of were performed in triplicate using the ABI PRISM 7900 HT sequence

detection system (Applied Biosystems, Foster City, CA) with SYBR® green technology.

Levels of heme oxygenase 1 and 2 cDNA were detected using the following primers:

HO-1 (sense 5’- GTC CGC AAC CCG ACA G -3’, antisense 5’- ACC AGC TTG AAG

CCG TCT C -3’, exon 1/2); HO-2 (sense 5’- CCC TGG ACC TGA ACA TGA A -3’,

antisense 5’- ACC CAT CCT CCA AGG TCT C -3’, exon 4/5). The exponential

portion of the amplification curve for 1000 copies of target amplicon passed through the

cycle threshold (CT1000) at 24.34 ± 0.63 cycles for HO-1 and 24.35 ± 0.30 cycles for

Chapter 3

- 75 -

HO-2. 28S (sense 5’- TTG AAA ATC CGG GGG AGA G -3’, antisense 5’- ACA TTG

TTC CAA CAT GCC AG -3’) transcripts were used as controls for normalization.

Results obtained under each experimental condition were compared with their own

corresponding normoxic control values.

Western Blotting: Total protein was extracted from HUVEC, HAEC, and human

blood outgrowth endothelial cells exposed to either normoxia or hypoxia for 16 h or 48 h

using the PARIS kit according to the manufacturer’s instructions. After protein

concentrations were determined by the Lowry method, total proteins (20µg/lane) were

resolved by 8-16% SDS-PAGE and transferred to nitrocellulose. To detect HO-1 and

HO-2 protein, membranes were blocked in 5% milk-TTBS and incubated with rabbit

polyclonal anti-HO-1 (1:2000) or anti-HO-2 (1:2000) antibody overnight follow by

peroxidase conjugated anti-rabbit IgG (1:2500). HO-1 and HO-2 protein was

normalized to β-actin detected by reprobing the membranes with anti-β-actin

monoclonal antibodies (1:40,000, Sigma). The immunocomplexes were visualized with

the ECL plus kit purchased from GE HealthCare (Uppsala, Sweden) and quantified by

digital densitometry using the Quantity One software provided by BioRad laboratories.

Results obtained under each experimental condition were compared with their own

corresponding normoxic controls.

3H-uridine and 3H-leucine Incorporation: 3H-uridine and 3H-leucine incorporation

was performed as previously described.[221] HUVEC plated in 12-well plates were

incubated under either normoxia or hypoxia for 16 h or 48 h. 3H-uridine or 3H-leucine

were added to media (10 µCi/well) for the last 15, 30, 45, or 60 minutes of normoxic or

Chapter 3

- 76 -

hypoxic exposure. Cells were washed with cold PBS, incubated in 10% trichloroacetic

acid for 20 minutes, washed 3 times with 100% ethanol, and dried in oven at 45°C. The

residues were dissolved in 0.3 N NaOH for 20 min and neutralized with 0.3 N HCl. The

resultant mixture (400 µl) from each well was added to 5ml of scintillation fluid and

radioactivity in each sample was counted in a liquid scintillation counter. The rate of 3H-

leucine or 3H-uridine incorporation is represented by the slope of the radioactivity-

incubation time relationship.

35S-methionine Incorporation: HUVEC were exposed to either normoxia or

hypoxia for 16 h and incubated in methionine-free RPMI 1640 medium supplemented

with 10% fetal calf serum and 35S-methionine (10 µCi/ml) for 2 additional hours. Total

proteins were extracted in RIPA buffer (50 mM Tris, 150 mM NaCl, 50 mM NaF, 1 mM

Na Orthovanadate, 5 mM Benzamidine, 1 mM EDTA, 1% Igepal CA630, 0.5% Sodium

Deoxycholate, and 0.1% SDS) and quantified by the Lowry method. Cell lysates (500

µg), precleared with protein G-Sepharose, are then immunoprecipitated using rabbit

polyclonal anti-HO-1 (1:50) or HO-2 antibodies (1:50) overnight. The resulting

immunoprecipitates were separated by 10% SDS-PAGE and bands were visualized by

autoradiography.

Polysome Profiling: As decribed previously,[222] HUVEC were exposed to either

normoxic or hypoxia for 6 or 24 h. At the end of exposure periods, HUVEC were

washed with PBS containing 100µg/ml cycloheximide and lysed using 200µl of lysis

buffer (100 mM KCl, 5 mM MgCl2, 10 mM HEPES, pH 7.4, 100 mg/ml cycloheximide,

and 1000 units/ml RNAseOUT). After centrifugation to remove cell debris,

Chapter 3

- 77 -

supernatants were layered onto a sucrose gradient (15% - 45%) and centrifuged for 2

hours at 35,000 rpm. A programmable density gradient fraction collector was used to

divide the gradient into 15 fractions so that HO-2 mRNA from each fraction could be

measured using quantitative real-time PCR.

Measurement of Intracellular Reactive Oxygen Species (ROS): Intracellular ROS

levels were measured in intact HUVEC using carboxy-H2DCFDA. This method is

based on the oxidation of non-fluorescent carboxy-H2DCFDA resulting in the formation

of the fluorescent compound 2’,7’-dichlorofluorescein (DCF). The fluorescence

generated by DCF is proportional to the rate of carboxy-H2DCFDA oxidation, which is

in turn indicative of the cellular oxidizing activity and intracellular ROS levels.

HUVEC grown to 70% confluence in 12-well plates were incubated with water, TNF-

α (10 ng/ml), or H2O2 (100 µM), prior to exposure to normoxia or hypoxia for 16 or 48

h. Cells were washed twice in HBSS and then incubated in HBSS containing of

carboxy-H2DCFDA (15 µM) for 30 min in the dark at 37 degrees. After rinsing with

HBSS once to remove free probe, fluorescence (Ex484/Em518 nm) from each well was

measured using the Fluroskan Ascent & FL fluorescent plate reader (Thermo Fisher,

Pittsburgh, PA). Cell number in each well was counted using the Z2 Coulter particle

count and size analyzer (Beckman Coulter, Fullerton, CA) so that fluorescence could be

normalized to cell number.

Mitochondrial Membrane Depolarization: Depolarization of the mitochondrial

membrane was detected using the cationic dye, 5,5’,6,6’-tetrachloro-1,1’,3,3’-

tetraethylbenzimidazolcarbocyanine Iodide (JC-1). JC-1 localizes to and aggregates

Chapter 3

- 78 -

within the mitochondria in proportion to mitochondrial membrane potential, emitting red

fluorescence. When the mitochondrial membrane depolarizes, JC-1 leaks into the

cytoplasm and forms monomers that emit green fluorescence. The ratio of red to green

fluorescence is an index of mitochondrial membrane depolarization. HUVEC grown to

70% confluence in 12-well plates were incubated with water, TNF-α (10 ng/ml), or

H2O2 (100 µM), prior to exposure to normoxia or hypoxia for 16 or 48 h. At the end of

the exposure period, HUVEC were incubated with PBS containing JC-1 (2 µM) for 15

min in the dark. Cells were then washed once with PBS and fluorescence

(Ex485/Em518nm (green) and Ex544/Em590nm (red)) was measured using the

Fluroskan Ascent & FL fluorescent plate reader (Thermo Fisher, Pittsburgh, PA).

Annexin V/Propidium Iodide labeling: The Roche Annexin V–FLUOS staining kit

was used to detect phosphatidylserine externalization (a marker of apoptosis) in HUVEC

and HAEC exposed to normoxia or hypoxia for 16 or 48 h. HUVEC and HAEC treated

with TNF-α or H2O2 were exposed to normoxia or hypoxia for 16 h. Cells in the media

were included in the sample. After trypsinization, cells were washed once with PBS

before addition of 100 µl of labeling solution that contains 2µl Annexin V-Fluos

labeling reagent and 2µl Propidium Iodide (PI) solution. Labeled cells were analyzed

using the cytomicsTM FC 500 flow cytometer (Beckman Coulter, Fullerton, CA).

Total Caspase Activation: Total caspase activation was measured in HUVEC and

HAEC exposed to normoxia or hypoxia for 16 or 48 h. HUVEC and HAEC treated with

TNF-α or H2O2 were exposed to normoxia or hypoxia. CaspACE FITC-VAD-FMK is a

FITC conjugate of the cell permeable inhibitor of caspases. This structure allows

Chapter 3

- 79 -

delivery of the inhibitor into the cell where binding to activated caspase serves as an in

situ marker for apoptosis. After trypsinization, cells were suspended in PBS containing

FITC-VAD-FMK (1 µM) at room temperature in the dark for 20 min. Cells were then

washed, resuspended in PBS, and analyzed using the cytomicsTM FC 500 flow cytometer

(Beckman Coulter, Fullerton, CA).

Data Analysis: Results are presented as mean ± S.E.M. for n number of

independent experiments with P<0.05 representing statistical significance. 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 3

- 80 -

3.3 Results

Effect of hypoxia on HO-1 and HO-2 mRNA and protein expression in systemic

vascular endothelial cells.

The effects of hypoxia on the expression of HO-1 and HO-2 mRNA and protein

were compared in HUVEC incubated under normoxic or hypoxic conditions for 16 and

48 h. After 16 and 48 h of hypoxic exposure, HO-1 mRNA is reduced to 27.82 ± 1.8 %

and 29.14 ± 10% of the corresponding normoxic control values, respectively (Figure

3.1A). HO-2 mRNA expression was decreased (42.93 ± 7.52 % of normoxic control,

Figure 3.1B) after 16 h of hypoxia and returned to the normoxic control level after 48 h.

HO-1 protein (Figure 3.1C) is reduced after 16 h and 48 h of hypoxia (79.57 ± 2.11%

and 64.93 ± 5.75% of normoxic control values, respectively) whereas HO-2 protein

levels were unaltered (Figure 3.1D). To corroborate this finding in other systemic

vascular endothelial cells, HO-1 and HO-2 proteins were also measured in HAEC and

human blood outgrowth EC exposed to normoxia or hypoxia for 16 h. Blood outgrowth

ECs were characterized by the expression of cell-surface markers. These cells stained

positive for CD34, KDR, VEGFR2, CD146, and CD31 and negative for CD14 and

CD45. As shown in Figure 3.2, hypoxia decreased HO-1 protein levels in both HAEC

and human blood orgin EC (38.50 ± 8.47% and 59.1 ± 8.85% of normoxic control

values, respectively) but, as in HUVEC, HO-2 was unaltered.

Chapter 3

- 81 -

Figure 3.1 HO-1 and HO-2 mRNA (A and B) in HUVEC exposed to 1% oxygen for

either 16 or 48 h. mRNA data are normalized to 28S rRNA. Bars represent means ±

S.E.M. n = 6 independent experiments, *P<0.05 for differences from corresponding

normoxic control.

A

16hrs 48hrs0

25

50

75

100

125 21% O21% O2

HO

-1 m

RNA

/ 28

S rR

NA(%

of N

orm

oxic

Con

trol

)

* *

B

16hrs 48hrs0

25

50

75

100

125

*

1% O2

21% O2

HO

-2 m

RNA

/ 28

S rR

NA(%

of N

orm

oxic

Con

trol

)

*

Chapter 3

- 82 -

16hrs 48hrs0

25

50

75

100

125

150

Duration of Hypoxia (1% O2)

*

HO-2

β-actin

1% O2

21% O2

HO

-2 /

β-a

ctin

Pro

tein

Lev

els

(% o

f Nor

mox

ic C

ontr

ol)

16hrs 48hrs0

25

50

75

100

125

150 21% O21% O2

Duration of Hypoxia (1% O2)

*

HO-1

β-actin

HO

-1 /

β-a

ctin

Pro

tein

Lev

els

(% o

f Nor

mox

ic C

ontr

ol)

* *

Figure 3.1 HO-1 and HO-2 mRNA protein levels (C and D) in HUVEC exposed to

1% oxygen for either 16 or 48 h. Protein data are normalized to β-actin. Bars represent

means ± S.E.M. n = 6 independent experiments, *P<0.05 for differences from

corresponding normoxic control.

C

D

Chapter 3

- 83 -

30405060708090

100110

21% O21% O2

HO-1β-actin

*

*

HO

-1 /

β-ac

tin P

rote

in L

evel

s(%

of N

orm

oxic

Con

trol

)

HAEC Blood Outgrowth EC

30405060708090

100110

21% O21% O2

HO-2

β-actin

HO

-2 /

β-ac

tin P

rote

in L

evel

s(%

of N

orm

oxic

Con

trol

)

HAEC Blood Outgrowth EC

Figure 3.2 (A) Representative blots of HO-1 and HO-2 protein in HAEC and human

blood outgrowth EC exposed to normoxia or hypoxia for 16 h. (B) Quantification of

HO-1 and HO-2 protein in HAEC and human blood outgrowth EC exposed to normoxia

or hypoxia for 16 h. Bars represent means ± S.E.M. n = 4 independent experiments,

*P<0.05 for differences from corresponding normoxic control.

A

B

Chapter 3

- 84 -

Effect of hypoxia on the translation of HO-2 transcripts in systemic vascular

endothelial cells.

In order to compare the effects of hypoxia on HO-1 and HO-2 protein synthesis

with its non-selective effects on total cellular mRNA and protein synthesis, 3H-uridine

and 3H-leucine incorporation were assessed in HUVEC after 16 and 48 h of hypoxic

incubation and 35S-methionine incorporation into HO-1 and HO-2 protein was measured

in HUVEC after exposure to hypoxia for 16 h (Fig. 3). Figure 3.3A illustrates that RNA

synthesis is decreased to 37.93 ± 3.71% and 28.78 ± 4.88% of normoxic control values,

respectively, after 16 and 48 h of hypoxic exposure. Protein synthesis is reduced to 56.6

± 2.77% at 16 h and 34.80 ± 2.97% at 48 h of the normoxic control value (Figure 3.3B).

Synthesis of HO-2 protein is less affected by hypoxia than HO-1 protein synthesis

(74.80 ± 7.80% vs. 47.01 ± 6.55% of normoxic control values, respectively). The

relative preservation of HO-2 protein synthesis, despite the 57% reduction in steady state

mRNA level and 43% reduction in total protein synthesis, suggests that HO-2 protein

expression is preserved during hypoxia, possibly through enhanced translation and/or

reduced protein degradation. Hypoxic incubation for 16 h with cycloheximide, an

inhibitor of RNA translation, reduced HO-2 protein levels. This suggests that translation

of HO-2 is important in maintaining HO-2 protein levels during hypoxia (data not

shown). To demonstrate the effect of hypoxia on HO-2 translation, ribosmal association

of HO-2 mRNA was assessed by polysome profiling. As shown in Figure 3.3D, after 6

h of hypoxia, HO-2 mRNA transcripts are located in higher polysome fractions which,

together with the results of the metabolic labelling and immunoprecipitation studies

supports enhanced translation of HO-2 transcripts during hypoxia.

Chapter 3

- 85 -

Normoxia 16 h 48 h0

25

50

75

100

Hypoxia (1% O2)

3 H-u

ridin

e In

corp

orat

ion

(% N

orm

oxic

Con

trol

)

**

Normoxia 16 h 48 h0

25

50

75

100

Hypoxia (1% O2)

**

3 H-le

ucin

e In

corp

orat

ion

(% N

orm

oxic

Con

trol

)

Figure 3.3 Rate of 3H-uridine (A) and 3H-leucine (B) incorporation into RNA and

protein of HUVEC exposed to normoxia or hypoxia for 16 or 48 h Bars represent means

± S.E.M. n = 4 independent experiments, *P<0.05 for differences from corresponding

normoxic control.

A

B

Chapter 3

- 86 -

HO-1 HO-20

25

50

75

100

125

*

21% O2, 16 h1% O2, 16 h

*35

S-m

ethi

onin

e In

corp

orat

ion

(% o

f Nor

mox

ic C

ontr

ol)

3 4 5 6 7 8 9 10 11 12 13 14 150.0

0.5

1.0

1.5

2.0

2.5

3.0

3.5 21% O2,6 h

1% O2, 6 h

Fraction Number

Rela

tive

Copi

es o

f HO

-2 T

rans

crip

t(p

er c

opy

in n

orm

oxic

frac

tion)

Figure 3.3 (C) Rate of 35S-methionine incorporation into HO-1 and HO-2 protein of

HUVECs exposed to normoxia or hypoxia for 16 h. Bars represent means ± S.E.M. n =

4 independent experiments, *P<0.05 for differences from corresponding normoxic

control. (D) Quantification of the abundance of HO-2 mRNA in various polysome

fractions from HUVEC exposed to normoxia or hypoxia for 6 h.

C

D

Chapter 3

- 87 -

Effect of decreased HO-2 protein level on ROS production in human endothelial

cells during hypoxia with or without treatment with TNF-α or H2O2

HO-2 protects cells from oxidative stress by reducing intracellular concentrations

of heme (a pro-oxidant) and by increasing levels of bilirubin and ferritin, which are

potent anti-oxidants. To determine the importance of HO-2 in protecting cells from

oxidative stress during hypoxia, we compared reactive oxygen specie (ROS) levels in

HUVEC transfected with scrambled or HO-2 siRNA exposed to normoxia or hypoxia

for 16 or 48 h, and also in HUVEC treated with TNF-α or H2O2 exposed to normoxia or

hypoxia for 16 h. Inhibition of HO-2 protein expression had no significant effects on

HO-1 protein expression in any of the conditions tested (data not shown). As illustrated

in figure 3.4B, exposure to hypoxia for 16 or 48 h increases ROS levels in HUVEC.

Compared to the scrambled siRNA control, inhibition of HO-2 expression increases

ROS levels in HUVEC exposed to hypoxia for 48h, but had no effect in cells exposed to

hypoxia for 16 h or in cells incubated under normoxic conditions. In HUVEC treated

with either TNF-α or H2O2, inhibition of HO-2 expression increases ROS levels in cells

exposed to hypoxia for 16 h (figure 3.4C).

Chapter 3

- 88 -

Scrambled siRNA HO-2 siRNA0

25

50

75

100H

O-2

/β

-act

in P

rote

in L

evel

s(%

of N

orm

oxic

Con

trol

)

*

HO-2

β-actin

21% O2 1% O2 21% O2 1% O20

50

100

150

200

250Scrambled siRNAHO-2 siRNA

16 h 48 h

†*†

DCF

Flu

ores

cenc

e/10

6 cel

ls(%

of N

orm

oxic

Con

trol

)

††

Figure 3.4 (A) Representative blots and quantification of HO-2 protein in HUVEC

transfected with scrambled or HO-2 siRNA. (B) ROS levels in HUVEC transfected

with scrambled or HO-2 siRNA exposed to normoxia or hypoxia for 16 or 48 h. Bars

represent means ± S.E.M. n = 6 independent experiments, *P<0.05 for differences

between with or without inhibition of HO-2 protein. †P<0.05 for differences between

hypoxia and corresponding normoxic control.

A

B

Chapter 3

- 89 -

21% O2 1% O2 21% O2 1% O20

50

100

150

200

250Scrambled siRNAHO-2 siRNA

H2O2 (100µM)TNF-α (10ng/ml)

**

DCF

Fluo

resc

ence

/106

cells

(% o

f Nor

mox

ic C

ontr

ol)

††

Figure 3.4 (C) ROS levels in HUVEC transfected with scrambled or HO-2 siRNA

exposed to normoxia or hypoxia for 16 h treated with TNF-α or H2O2. Bars represent

means ± S.E.M. n = 6 independent experiments, *P<0.05 for differences between with

or without inhibition of HO-2 protein. †P<0.05 for differences between hypoxia and

corresponding normoxic control.

C

Chapter 3

- 90 -

Inhibition of HO-2 protein expression decreases mitochondrial membrane potential

in human endothelial cell during hypoxia in the presence and absence of TNF-α or

H2O2

Both hypoxia and increased intracellular ROS levels may trigger programmed

cell death (apoptosis) or necrosis depending on their severity/magnitude. To determine

the functional significance of preservation of HO-2 levels during hypoxia, the effect of

HO-2 expression inhibition using HO-2 siRNA on HUVEC viability during hypoxia in

the presence or absence of TNF-α or H2O2 was assessed. Mitochondrial membrane

depolarization is an early event in the intrinsic apoptotic pathway activated by hypoxia.

The ratio of JC-1 aggregates/JC-1 monomer, which is a measure of mitochondrial

membrane potential (13), is decreased during hypoxic exposure (Fig. 5). Inhibition of

HO-2 expression further reduced mitochondrial membrane potential after exposure to

hypoxia for 48 h (Fig. 5A), suggesting that HO-2 protein or its activity increases the

capacity to maintain mitochondrial membrane potential during hypoxia. In HUVEC

treated with TNF-α or H2O2, inhibition of HO-2 enhanced mitochondrial membrane

depolarization in both normoxic and hypoxic cells (Figure 3.5B).

Chapter 3

- 91 -

21% O2 1% O2 21% O2 1% O20

25

50

75

100

16 h 48 h

Scrambled siRNAHO-2 siRNA

*

† †

JC-1

Agg

rega

tes/

JC-1

Mon

omer

s(%

of N

orm

oxic

Con

trol

)

21% O2 1% O2 21% O2 1% O20

25

50

75

100

TNF-α H2O2

Scrambled siRNAHO-2 siRNA

*

**

† †

JC-1

Agg

rega

tes/

JC-1

Mon

omer

s(%

of N

orm

oxic

Con

trol

)

Figure 3.5 (A) Mitochondrial membrane potential in HUVEC transfected with

scrambled or HO-2 siRNA exposed to normoxia or hypoxia for 16 or 48 h. (B)

Mitochondrial membrane potential in HUVEC transfected with scrambled or HO-2

siRNA exposed to normoxia or hypoxia for 16 and treated with TNF-α, or H2O2. Bars

represent means ± S.E.M. n = 6 independent experiments, *P<0.05 for differences

between with or without inhibition of HO-2 protein. †P<0.05 for differences between

hypoxia and corresponding normoxic control.

A

B

Chapter 3

- 92 -

HO-2 preserves human endothelial cell viability during hypoxia in the presence and

absence of TNF-α or H2O2

Annexin V/PI double staining was used to detect externalization of

phosphatidylserine, an early event in apoptosis, and cell membrane permeabilization, an

indicator of cell death. Representative plots of annexin V/PI staining are shown in Fig

6A. Non-viable cells are cells that are stained by annexin V and/or PI (Figure 3.6A).

Figure 3.6B demonstrates that exposure of HUVEC to hypoxia for 48 h increases cell

death and that cell death is further increased by inhibition of HO-2 expression. In

HUVEC treated with TNF-α or H2O2, HO-2 expression inhibition had no effect on

normoxic cells and increases cell death after hypoxic exposure for 16 h (Figure 3.6C).

To confirm that the effect of decreased HO-2 activity on cell viability is mediated by

inhibition of apoptosis, caspase activation was assessed in HUVEC exposed to normoxia

or hypoxia for 16 or 48 h and in HUVEC treated with TNF-α or H2O2 exposed to

normoxia or hypoxia for 16 h (Figure 3.7). In HUVEC where HO-2 protein expression

is inhibited, exposure to hypoxia for 48 h increased activated caspase by 1.5 fold

compare to scrambled siRNA control (Figure 3.7A). Total activated caspase is also

increased in HUVEC treated with TNF-α or H2O2 and exposed to hypoxia for 16 h

(Figure 3.7B).

Chapter 3

- 93 -

Annexin V

Scrambled siRNA, 21% O2, 48 h HO-2 siRNA 21% O2, 48 h

Scrambled siRNA, 1% O2, 48 h HO-2 siRNA 1% O2, 48 h

Pro

pidi

um Io

dide

Figure 3.6 (A) Representative annexin V/PI staining plots of HUVEC transfected

with scrambled or HO-2 siRNA exposed to normoxia or hypoxia for 48 h.

A

Chapter 3

- 94 -

21% O2 1% O2 21% O2 1% O20

255075

100125150175200

16 h 48 h

Scrambled siRNAHO-2 siRNA *

†

Cell

Deat

h(%

of N

orm

oxic

Con

trol

) †

21% O2 1% O2 21% O2 1% O20

255075

100125150175200

Scrambled siRNAHO-2 siRNA

H2O2 (100µM)TNF-α (10ng/ml)

**

Cell

Deat

h(%

of N

orm

oxic

Con

trol

)

Figure 3.6 The amount of cell death in HUVEC transfected with scrambled or HO-2

siRNA exposed to normoxia or hypoxia for 16 or 48 h (B) or exposed to normoxia or

hypoxia for 16 h in the presence of TNF-α, or H2O2 (C). Bars represent means ± S.E.M.

n = 6 independent experiments, *P<0.05 for differences between with or without

inhibition of HO-2 protein. †P<0.05 for differences between hypoxia and corresponding

normoxic control.

C

B

Chapter 3

- 95 -

21% O2 1% O2 21% O2 1% O20

50

100

150

200Scrambled siRNAHO-2 siRNA

H2O2 (100µM)TNF-α (10ng/ml)

**

Tota

l Act

ivat

ed C

aspa

se(%

of N

orm

oxic

Con

trol

)

†

Figure 3.7 Total activated caspase level in HUVEC transfected with scrambled or

HO-2 siRNA exposed to normoxia or hypoxia for 16 or 48 h or exposed to normoxia (A)

or hypoxia for 16 h and treated with TNF-α or H2O2 (B). Bars represent means ±

S.E.M. n = 5 independent experiments, *P<0.05 for differences between with or without

inhibition of HO-2 protein. †P<0.05 for differences between hypoxia and corresponding

normoxic control.

21% O2 1% O2 21% O2 1% O20

50

100

150

200

Scrambled siRNAHO-2 siRNA

16 h 48 h

*†

Tota

l Act

ivat

ed C

aspa

se(%

of N

orm

oxic

Con

trol

) †

A

B

Chapter 3

- 96 -

HO-2 preserves human endothelial cell viability during hypoxia in the presence and

absence of TNF-α or H2O2

To confirm the cytoprotective effect of HO-2 protein during hypoxia is not

specific to HUVEC, annexin V/PI staining and total caspase activation were assessed in

HAEC exposed to normoxic or hypoxia for 48 h and in HAEC treated with TNF-α or

H2O2 exposed to normoxia or hypoxia for 16 h. As observed in HUVEC, inhibition of

HO-2 increased cell death in HAEC exposed to hypoxia for 48 h (Figure 3.8A) and in

HAEC exposed to hypoxia for 16 h treated with TNF-α or H2O2 (Figure 3.8C). When

HO-2 expression was suppressed, total activated caspase increased in HAEC exposed to

hypoxia for 48 h and in HAEC treated with TNF-α or H2O2 and exposed to hypoxia for

16 h (Figure 3.8B and Figure 3.8D).

Chapter 3

- 97 -

21% O2 1% O20

50

100

150

200

250

300 Scrambled siRNA, 48 h

HO-2 siRNA, 48 h **

Cell

Deat

h(%

of N

orm

oxic

Con

trol

)

21% O2 1% O20

50

100

150

200

250

300 Scrambled siRNA, 48 hHO-2 siRNA, 48 h

**

Tota

l Act

ivat

ed C

aspa

se(%

of N

orm

oxic

Con

trol

)

Figure 3.8 Cell death (A) and total activated caspase level (B) in HAEC transfected

with scrambled or HO-2 siRNA exposed to normoxic or hypoxia for 48 h. Bars

represent means ± S.E.M. n = 5 independent experiments, *P<0.05 for differences

between with or without inhibition of HO-2 protein.

B

A

Chapter 3

- 98 -

21% O2 1% O2 21% O2 1% O20

255075

100125150175200

H2O2 (100µM)TNF-α (10ng/ml)

Scrambled siRNA, 16 hHO-2 siRNA, 16 h

**

**

Cell

Deat

h(%

of N

orm

oxic

Con

trol

)

21% O2 1% O2 21% O2 1% O20

50

100

150

200

250

H2O2 (100µM)TNF-α (10ng/ml)

Scrambled siRNA, 16 hHO-2 siRNA, 16 h

**

*

*

Tota

l Act

ivat

ed C

aspa

se(%

of N

orm

oxic

Con

trol

)

Figure 3.8 Cell death (C) and total activated caspase level (D) in HAEC transfected

with scrambled or HO-2 siRNA exposed to normoxia or hypoxia for 16 h and treated

with TNF−α or H2O2. Bars represent means ± S.E.M. n = 5 independent experiments,

*P<0.05 for differences between with or without inhibition of HO-2 protein.

C

D

Chapter 3

- 99 -

DISCUSSION

The results of this study show that in human endothelial cells incubated under

hypoxic conditions: 1) HO-1 mRNA and protein levels are decreased; 2) HO-2 protein

level is unaltered despite a 40% reduction in HO-2 mRNA expression and 50%

reduction in total protein synthesis; 3) HO-2 protein level is maintained through

enhanced translation of HO-2 transcripts; and 4) inhibition of HO-2 expression increases

production of reactive oxygen species, decreases mitochondrial membrane potential and

enhances apoptotic cell death.

Previous studies indicate cell type- and inter-species differences in the regulation

of HO expression by hypoxia. Aortic HO-2 protein is increased in rats exposed to

hypoxia,[74] remains unchanged in cultured rat aortic smooth muscle cells and human

cytotrophoblast[223, 224] and is decreased in Jurkat, K562, and YN-1 cells after

hypoxic incubation.[176] HO-1 mRNA and protein levels decrease in HUVEC, human

astrocytes, and human coronary artery endothelial cells[172, 225] but increase in bovine

aortic and rat pulmonary artery endothelial cells and in human fibroblasts and smooth

muscle cells after hypoxic incubation.[226-229] In HUVEC, decreased expression of

HO-1 after hypoxia is mediated by induction of the transcription repressor Bach1.[172]

The current study, the first to directly compare the effects of hypoxia on HO-1 and HO-2

expression in human endothelial cells, demonstrates that although both HO-1 and HO-2

mRNA levels are decreased, HO-2, but not HO-1 protein level remains unchanged. HO-

1 and HO-2 protein levels are, therefore, differentially regulated by oxygen tension and

HO-2 is the predominant isoform present under these conditions.

Chapter 3

- 100 -

HO-1 is primarily regulated transcriptionally and the genomic sequences 5’ to its

coding region contain cis-acting response elements that bind transcription factors

including HIF-1α, AP1, SP1, as well as the heme response element GC-

NNNGTCA.[135] In contrast, the 5’-flanking region of the HO-2 gene contains no

regulatory elements corresponding to transcription factors known to participate in the

response to hypoxia.[165, 173-175] Not surprisingly, therefore, hypoxic incubation did

not result in increased HO-2 mRNA levels in the current study, or in any of the cell

culture systems in which it has previously been evaluated.[223, 230] Nevertheless, HO-

2 expression is not entirely constitutive. Development stage-specific changes in HO-2

protein levels have been reported and HO-2 protein is increased in the aortic

endothelium of rats exposed to hypoxia without a corresponding increase in HO-2

mRNA.[74, 181, 231] Similarly, spatial and temporal dissociation between HO-2

protein and mRNA expression have been noted in the rodent brain and testis.[165, 174,

181] Our current results, therefore, reconcile these observations by demonstrating that

HO-2 expression is regulated at the post transcriptional level.

Hypoxia results in decreased cap-dependent translation due to increased

formation of the eIF4E/4E-BP1 inhibitory complex and increased phosphorylation of

eIF2F-α.[35] When hypoxia is severe, or prolonged, transcription is also inhibited and

mRNA levels decrease, as observed in the present study. Accordingly, maintenance of

protein levels under these conditions requires enhanced translation of existing mRNA

transcripts and/or reduced degradation of protein. Our current observation that HO-2

transcripts are localized to larger polysome fractions after hypoxic incubation (Figure

Chapter 3

- 101 -

3D) supports enhanced translation as an important mechanism by which HO-2 protein

levels are preserved. Consistent with this conclusion, we observed that the reduction in

HO-2 protein synthesis after exposure to hypoxia for 16 h is small relative to the

decreases in HO-2 mRNA levels and the rate of total protein synthesis. Furthermore,

HO-2 protein levels were decreased after hypoxic incubation in HUVEC treated with the

protein translation inhibitor cycloheximide, but not with the RNA synthesis inhibitor

actinomycin D or the proteasome inhibitor epoximycin. In other transcripts for which

this has been described, structural features that enhance cap dependent (nNOS) and cap-

independent (Tie-2) ribosomal association have been identified in their 5’ untranslated

regions.[54, 185] Using the BD Marathon-Ready human testis cDNA library, Zhang et

al. has demonstrated that HO-2 transcription is initiated from multiple sites.[176] Thus,

translation of HO-2 could be enhanced through selective transcriptional activation of a

promoter that produces more efficiently translated mRNA species during hypoxia, a

mechanism we have previously shown to mediate hypoxic enhancement of nNOS

expression in vascular smooth muscle.[185] In view of the current findings, therefore,

further examination of HO-2 mRNA structure and its functional relevance in the

regulation of HO-2 protein expression during hypoxia are now warranted.

Oxidant injury occurs when there is an imbalance between the formation of ROS

and the antioxidant capacity of the cell and is implicated in the pathogenesis of organ

dysfunction in diseases associated with reduced oxygen delivery.[232, 233] Previous

studies demonstrate that HO-2 is a component of the endogenous cell defence against

oxidative stress; HO-2 gene deletion increases hemin-induced injury in astrocytes and

sensitizes cerebral vascular endothelial cells to glutamate and TNF-α induced

Chapter 3

- 102 -

apoptosis.[157, 158, 234] The results of the current study confirm the essential role that

HO-2 plays in oxidant stress defence in human endothelial cells exposed to hypoxia

since inhibition of its expression increases intracellular ROS levels after 48 hours of

hypoxic incubation (Figure 4B). Compensation of HO activity by increased HO-1

expression was not observed. To corroborate the conclusion that HO-2 is important in

modulating oxidant stress, the effect of inhibition of HO-2 expression on the response to

other oxidative stimuli (TNF-α or H2O2) was also evaluated. In cells deficient in HO-2,

significant increases in ROS levels were observed only after hypoxic incubation. Its role,

relative to other defence mechanisms, therefore, is specifically enhanced during hypoxia.

HO activity is required for catabolism the prooxidant heme and for production of

bilirubin, a scavenger of superoxide and peroxyl radicals.[135] HO-2 also plays a

specific role in regulating intracellular free iron which increases the generation of

reactive hydroxyl radicals through the Fenton reaction.[138, 235] Since HO-1

expression is inhibited by hypoxia, HO-2 becomes the predominant isoform under these

conditions and a significant mechanism of defence against oxidant stress and hypoxic

injury.

Apoptosis may be triggered in response to stimuli extrinsic or intrinsic to the

affected cell. Hypoxia-induced apoptosis occurs mainly through the intrinsic

pathway.[236, 237] The lack of oxygen limits ATP synthesis required for maintenance

of the mitochondrial membrane potential. Depolarization of the mitochondrial

membrane potential elevates cytoplasmic ROS levels and further inhibits oxidative ATP

synthesis because the electromotive force for electron transport is reduced. This, in turn,

Chapter 3

- 103 -

activates Bax or Bak and leads to cytochrome C release into the cytosol, caspase

activation and chromatin fragmentation.[238] In the current study, we demonstrate that

mitochondrial membrane potential is decreased after exposure to hypoxia for 16 h,

whereas increase in cell death (annexin V/PI staining) and total activated caspase

activity was detected only after 48 h. These results support involvement of the intrinsic

pathway since mitochondrial membrane depolarization precedes caspase activation.

HO-2 protects against apoptotic cell death induced by TNF-α and glutamate in

cerebrovascular endothelial cells and by hydrogen peroxide in HEK cells.[140, 157-159]

In the present study, inhibition of HO-2 expression exacerbated mitochondrial

membrane depolarization and increased cell death and activated caspase levels in

hypoxic, but not normoxic human endothelial cells. When cells where concomitantly

treated with TNF-α or H2O2, the anti-apoptotic effect of HO-2 was detected after

exposure to hypoxia for 16 h, instead of 48 h. These results indicate that HO-2 also

protects against hypoxia-induced apoptosis in human endothelial cells, and plays an

even greater role in preserving cell viability during concomitant oxidative stress induced

by TNF-α or H2O2, perhaps because generalized inhibition of protein synthesis by

hypoxia suppresses the expression of components of conventional cytoprotective

pathways.

Consistent with previous studies, our present results support a central role for

reactive oxygen species in triggering apopototic endothelial death induced by hypoxia,

TNF-α and H2O2. Elevated ROS levels induce apoptosis through activating the JNK-

cJUN pathway and/or damaging mitochondrial membrane integrity resulting in reduced

Chapter 3

- 104 -

mitochondrial membrane potential.[239] These may be abrogated by the antioxidative

effects of HO-2, however, several other mechanisms have been invoked. For example,

CO inhibits TNF-α-induced apoptosis by activation of p38 MAPK pathway.[161] In

addition HO-2 protects cell viability through mechanism(s) separate from its role in

heme degradation since transfection of HEK cells with a catalytically inactive HO-2

mutant protects against oxidative injury, although the mechanism of protection is

unknown.[140]

Although HO-2 and HO-1 catalyze the same reaction, the differences between

these enzymes could provide insight into the advantages of maintaining HO-2, but not

HO-1 during prolonged hypoxia. In HEK cells transfected with plasmids containing

either HO-1 or HO-2 and treated with hydrogen peroxide, HO-2 was found to colocalize

with its cofactor NAPH-cytochrome P450 reductase in the microsomal fraction, whereas

HO-1 was more widely dispersed.[159] Accordingly, HO-2 may provide a more

efficient pathway for heme degradation, hence greater cytoprotective capacity, due to its

subcellular localization in association with this co-factor. Additionally, HO-2 contains

Cys-Pro repeats, termed heme regulatory domains, not present in HO-1 that provide

heme binding sites distinct from the heme catalytic domain.[135] During hypoxia or

ischemia injury, large amounts of prooxidant heme are release by cells undergoing

necrosis or apoptosis. HO-2, but not HO-1, could sequester this excess free heme.

Finally, differential regulation of the expression of these isoforms enables fine control of

the antioxidant capacity of the endothelium; by maintaining HO-2 and downregulating

Chapter 3

- 105 -

HO-1 during hypoxia, endothelial cells reserve the capacity to increase HO activity in

response to additional stress.

Previously, we identified a role for HO-2 in preserving endothelium-dependent

modulation of vasoconstrictor responses to endothelin-1 and phenylephrine in rats

exposed to prolonged hypoxia.[74] HO-2 knockout mice exhibit hypoxemia and

myocardial hypertrophy while breathing room air, indicating that HO-2 contributes to

pulmonary ventilation-perfusion matching.[240] Our current results highlight the

importance of HO-2 in modulating endothelial cell apoptosis which is a prominent

feature in a variety of diseases including atherosclerosis, ischemia/reperfusion injury,

and transplantation. Accumulating evidence, therefore, supports a central role for HO-2

in the cardiopulmonary adaptation to hypoxia and in the pathophysiology of disorders in

which endothelial injury contributes to vascular dysfunction. Accordingly it represents a

potential novel target for therapeutic intervention.

Chapter 4

- 106 -

CHAPTER 4

Perspective

Chapter 4

- 107 -

Hypoxia occurs in a variety of cardiopulmonary diseases and in normal

individuals during ascent to high altitude. In response to acute hypoxia, oxygen delivery

to vital organs is maintained through adrenergically mediated sympathetic responses and

endothelial release of vasoactive peptides.[77] As hypoxic exposure is prolonged,

sympathetic regulation of vascular tone is impaired due to reduced contractile response

to adrenergic stimulation.[62, 64] Studies in rat aorta indicate that enhanced targeting of

type 1 phosphatase activity to the contractile myofilaments and increased expression of

the inhibitory thin-filament proteins caldesmon and calponin may contribute.[62, 63]

The impairment of vascular smooth muscle contractility is partially compensated by

alteration in the function of the endothelium, in that it becomes an agency of

vasoconstrictors as opposed to its normal role as a source of vasorelaxing factors.[60,

61]

Endothelin-1 is a potent vasoconstrictor released by the endothelium and

previous studies have shown that ET-1 plays a central role in the adaptation to

hypoxia.[61, 76, 111, 112, 241] The increased vascular ET-1 production during hypoxia

potentiates vascular reactivity and enhances oxygen extraction. However, the

mechanism of its activation in the vasculature has not been fully investigated.

Classically, ET-1 is produced by the cleavage of big ET-1 by ECE-1.[242] Recently,

vascular MMP-2 mediated cleavage of big ET-1 was found to release a vasoconstrictive

ET-1 isopeptide (ET-1[1-32]).[96] MMP-2 also regulates vascular tone by inactivating the

vasodilators CGRP and adrenomudulin.[97, 188] Given that hypoxia increases MMP-2

production and activation,[83] MMP-2 has the potential to play a significant role in

regulating vascular reactivity during hypoxia.

Chapter 4

- 108 -

In the studies presented in chapter 2, the vasoregulatory role of vascular MMP-2 was

investigated, along with the effect of prolonged exposure to hypoxia in vivo on vascular

MMP-2 production and activity. The novel findings of this study are:

1) Vascular MMP-2 mediates vasoconstriction in systemic conduit and resistance

vessels of rats exposed to hypoxia for 7 days.

2) Vascular MMP-2 mediated activation of Big ET-1 is a prominent mechanism of

regulation of vascular reactivity during hypoxia

3) Hypoxia increases vascular MMP-2 and MT1-MMP protein levels without

altering TIMPs 1-4 protein levels

4) Hypoxia induces MMP-2 and MT1-MMP mRNA expression in the deep medial

vascular smooth muscle

It is well recognized that changes in vascular tone precede the structural alterations

that occur when changes in blood flow persist chronically, as occurs during prolonged

hypoxia.[215, 216] Such remodeling of the circulation is important in adapting the

mature circulation to chronic changes in tissue perfusion. Given vascular MMP-2’s

vasoregulatory role and its role in basement membrane degradation, hypoxic activation

of vascular MMP-2 represents a pivotal pathway by which the acute vascular responses

to hypoxia may be integrated with the longer-term structural changes in both conduit and

resistance arteries. MMP-2 activation alters the activity of a number of vasoregulatory

peptides in addition to big ET-1.[96, 198, 199, 243] Although the present study

demonstrated that activation of big ET-1 plays a role, the relative importance of other

pathways remains to be explored. Given that ET-1 also promotes proliferation and

inhibits apoptosis of endothelial and smooth muscle cells,[100] investigation into other

Chapter 4

- 109 -

physiological roles of MMP-2 mediated release of ET-1[1-32] during hypoxia is

warranted. Expression of MMP-2 and big ET-1 is also be observed during conditions of

tissue injury, inflammation and cancer.[244-246] It is also important, therefore, to

elucidate the effects of MMP-2 mediated activation of big ET-1, and whether it is

influenced by tissue oxygenation, in these pathophysiological settings. Selective

inhibition of MMP-2 may represent a new pharmacological strategy for regulating

vascular reactivity and remodeling in pathological conditions.

Heme oxygenase-2 is another vasoregualtory enzyme with the potential to

regulate the production and potency of ET-1.[74] HO-2 increases in the endothelium

and alters aortic reactivity after exposure to hypoxia in rats. Based on the properties of

its products (CO and biliverdin) endothelial HO-2 may play additional roles, potentially

acting to increase endothelial cell viability and reduce inflammatory responses.[135]

Hypoxia reduces HO-1 mRNA and protein levels in human endothelial cells,[172, 225]

and it is likely that HO-2 is the dominate HO enzyme in these cells during hypoxia.

Accordingly we proposed that it may play a meaningful role in preserving endothelial

function in conditions associated with reduced oxygen delivery. In the absence of active

transcriptional regulation others have suggested that HO-2 protein expression may be

regulated posttranscriptionally.[173, 179, 181] In the studies presented in chapter 3,

therefore, oxygen regulation of HO-2 in human endothelial cells and its role in

preserving endothelial cell viability during hypoxic stress was investigated. The novel

findings are:

1) HO-2 protein level is unaltered despite a 40% reduction in HO-2 mRNA

expression and 50% reduction in total protein synthesis

Chapter 4

- 110 -

2) Hypoxia enhances translation of HO-2 transcripts

3) Inhibition of HO-2 protein expression increases production of reactive oxygen

species, decreases mitochondrial membrane potential and enhances apoptotic cell

death.

These results demonstrate that HO-2, but not HO-1, protein level is selectively

maintained in human endothelial cells during hypoxia through enhanced translation of

HO-2 transcripts. In some cases, a competitive advantage over other mRNAs for

ribosome binding is conferred through activation of an alternate promoter that drives

expression of an mRNA containing a 5’ UTR lacking secondary structure. This

mechanism regulates the expression of the hypoxia inducible nNOS variant.[185] In

other cases, the presence of an internal ribsosomal entry site in the 5’ UTR that enables

cap-independent translation increases translation in situations where cap-dependent

translation is inhibited, such as during hypoxia.[37, 247] In view of these findings,

therefore, further examination of HO-2 mRNA structure and its functional relevance in

the regulation of HO-2 protein expression during hypoxia are now warranted.

In the studies described in chapter 3, HO-2 protein was found to be anti-

oxidative and anti-apoptotic in human endothelial cells, suggesting that HO-2 is

essential in maintaining endothelial integrity in conditions associated with hypoxia.

Further study to assess the pathophysiological relevance of this effect in vivo is now

warranted. HO-2 could maintain endothelial cell viability through the effects of

bilirubin, carbon monoxide, ferritin, or other unidentified intracellular signaling

pathways. Therefore, the molecular mechanisms mediating the effects on HO-2 in the

current setting remain to be identified. Lastly, given that HO-2 activity reduces ROS

Chapter 4

- 111 -

levels in human endothelial cells during hypoxic exposure, and ROS increase both

MMP-2 and ET-1 mRNA expression, future studies will investigate the link between

HO-2 and oxygen regulation of MMP-2 expression and ET-1 bioavailability.

In conclusion, my work has revealed that vascular MMP-2 and HO-2 play

important roles in the adaptive response to hypoxia. Further investigation into signaling

pathways altered by these enzymes could lead to the development of novel therapeutic

strategies to mitigate the effects of hypoxia in patients with disorders associated reduced

oxygen delivery.

- 112 -

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