THE ADAPTIVE ROLE OF NEURONAL NITRIC OXIDE SYNTHASE IN MAINTAINING OXYGEN HOMEOSTASIS

DURING ACUTE ANEMIA

by

Albert King-Yeung Tsui

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

Department of Physiology University of Toronto

© Copyright by Albert King-Yeung Tsui, 2012

The Adaptive Role of Neuronal Nitric Oxide Synthase in

Maintaining Oxygen Homeostasis during Acute Anemia

Albert King-Yeung Tsui

Doctor of Philosophy

Department of Physiology University of Toronto

2012

Abstract

Mammals are well adapted to respond to changes in ambient oxygen concentration (O2) by

activating homeostatic physiological and cellular responses which maintain cell function and

survival. Although anemia has been associated with increased mortality in a number of clinical

settings, surprisingly little is known about how anemia affects tissue PO2 and hypoxia signaling.

Because nitric oxide synthases (NOSs) figure prominently in the cellular response to acute

hypoxia, we define the effects of NOS deficiency in acute anemia. Unlike wildtype (WT),

endothelial NOS (eNOS) and inducible NOS (iNOS) deficient mice, only neuronal NOS (nNOS)

deficient mice (nNOS-/-) demonstrated increased mortality during acute anemia. With respect to

global tissue O2 delivery, anemia did not increase cardiac output (CO) or reduce systemic

vascular resistance (SVR) in nNOS -/- mice. At the cellular level, anemia increased expression of

HIF-1α and HIF-responsive mRNA levels (EPO, VEGF, GLUT1, PDK) in the brain of WT, but

not nNOS-/- mice. These date suggest that nNOS contributed to cardiovascular and cellular

mechanisms which maintain oxygen homeostasis in anemia. To confirm the physiological

relevance of these findings in a whole animal model of anemia, we utilized transgenic animals

which express a reporter HIF-α(ODD)-luciferase chimeric protein. Using this model, we

confirmed that nNOS is essential for anemia-induced increases in HIF-α protein stability in vivo

ii

in real-time whole animal images and brain tissue. With respect to the mechanism, nNOS-

derived NO is known to affect S-nitrosylation of specific proteins, which may interfere with HIF-

α and von Hippal Lindau protein (pVHL) interaction. Utilizing the biotin switch assay, we

demonstrated that anemia caused a time-dependent increase in S-nitrosylation of pVHL in brain

tissue from WT but not nNOS-/- mice. In addition, anemia also leads to a decrease in S-

nitrosoglutathione (GSNO) reductase protein expression, an important enzyme responsible for

de-nitrosylation of proteins. The combination of increased nNOS expression and decreased

GSNO reductase expression would favor prolonged S-nitrosylation of proteins during anemia.

These findings identify nNOS effects on the HIF/pVHL signaling pathway as critically important

in the physiological responses to anemia in vivo. By contrast, after exposure to acute hypoxia,

nNOS-/- mice survived longer, retained the ability to regulate CO and SVR, and increased brain

HIF-α protein levels and HIF-responsive mRNA transcripts. This comparative assessment

provided essential mechanistic insight into the unexpected and striking difference between

anemia and hypoxia. Understanding the adaptive responses to acute anemia will help to define

novel therapeutic strategies for anemic patients.

iii

Acknowledgments

I would like to thank my supervisor, Dr. Gregory Hare, for his dedicated support,

patience, and mentorship on my doctoral studies throughout the years. His encouragement, good

advice, and teaching had allowed me to excel as a researcher and an individual. I would also like

to thank my co-supervisor, Dr. Philip Marsden, for his continued guidance during my research.

His ideas and scientific approaches helped developed my ability as a critical thinker and make

my PhD training years enjoyable. I am extremely grateful to the members who served on my

supervisory committee, Dr. David Mazer and Dr. Lee Adamson, who have supported my thesis

work from the beginning to the end. I wish to thank my collaborators, Dr. Scott Heximer, Dr.

Steffen-Sebastian Bolz, Dr. Kim Connelly, Dr. Mark Henkelman, for their resources and

insightful ideas to my research work.

I would also like to extend my thanks to past and present members in the laboratories of

Dr. Hare and Dr. Marsden. I am especially grateful to Elaine Liu, Anya McLaren, David Ho,

Matthew Yan, Apurva Shirodkar, Tania Petruzziello, Brent Steer, Mostafa El-Beheiry, Tenille

Ragoonanan, Kevin Chen, Neil Dattani, Namhee Kim, Tina Hu and Keith Lee for all their

scientific critique, emotional support, care, and entertainment they provided throughout the

years.

Lastly, my deepest gratitude goes to my parents, Philip Tsui and Christina Au-Yeung,

and my sister, Joyce Tsui, for their unconditional love, care and support throughout my life. This

dissertation would be impossible without their continuous encouragement.

iv

Contributions

The work presented in Chapter 2 and 3 has been published in Tsui AKY et al., Priming of

hypoxia inducible factor by neuronal nitric oxide synthase is essential for adaptive responses in

severe anemia. Proc Natl Acad Sci U S A. 2011 Oct 18;108(42):17544-9. As a first author of the

publication, I contributed to experimental design, data generation, figure construction, and

manuscript writing. I am grateful to acknowledge the contribution of the following people to help

with the paper and this thesis along the way:

Chapter 2:

Survival curve on isoflurance in mice (Figure 2.2) was performed by Mostafa El-Behiery.

Measurement of mean arterial pressure from femoral artery (Figure 2.3 and 2.5) was performed

together with Jenny Zhang in Dr. Scott Heximer’s laboratory. Microvascular brain oxygen

tension study (Figure 2.9 and 2.10) was performed by Neil Dattani. Cardiac output measured by

pressure-volume loops (Figure 2.4) were performed together with Dr. MD Golam Kabir and

Jean-Francois Desjardins in Dr. Kim Connelly’s laboratory. Assessment of vascular reactivity in

mesenteric resistance vessels (Figure 2.7) was performed by Dr. Darcy Lidington in Dr. Steffen

Bolz’s laboratory.

Chapter 3:

Kevin Chen performed some of the Western blots under my supervision (Figure 3.2 and 3.3).

Immunofluoresence staining (Figure 3.4) was performed by Dr. Elaine Liu. Technical support in

primer design and PCR was provided by Britta Knight, David Ho, and Matthew Yan in Dr.

Philip Marsden’s laboratory. Keith Lee aided in HIF-α(ODD) luciferase imaging (Figure 3.9 and

3.11) and performed the Hb threshold study (Figure 3.13). Biotin switch experiments (Figure

3.14 and 3.15) and GSNOR protein measurement (Figure 3.16) were performed in collaboration

with David Ho in Dr. Philip Marsden’s laboratory.

v

Table of Contents

Acknowledgments iv Contributions v Table of Contents vi List of Tables ix List of Figures x Abbreviations xiii Chapter 1: Introduction 1.1: Oxygen – The Element of Life 2

1.1.1: Importance of O2 to complex organisms 2 1.1.2: Oxygen transport in the body 3 1.1.3: Acute oxygen sensing in mammals 7

1.2: Anemia 12 1.2.1: Do all types of anemia share a common mechanism for morbidity and

mortality? 13 1.2.2: Risks of anemia 13 1.2.3: Traditional view of the physiological adaptation to acute anemia 16 1.2.4: Tissue oxygenation in acute anemia 21 1.2.5: Cellular responses to anemia 22 1.3: Hypoxia 24 1.3.1 Physiological adaptation to hypoxia 26 1.4: Nitric oxide (NO) 29

1.4.1: Nitric oxide synthases (NOS) 31 1.4.2: Targets of NO 32

Heme 32 S-nitrosylation of cysteine thiols 33

NO storage as nitrite and nitrate 36 1.5: Neuronal nitric oxide synthase (nNOS) 37

1.5.1: Biochemistry of nNOS 37 1.5.2: Regulation of the nNOS gene 38 mRNA diversity 38 Post-translational modification 41 1.5.3: Role of nNOS in different tissues 43 Brain 44

vi

Heart 44 Kidneys 46 Smooth muscle 46 Skeletal muscle 47 1.5.4: Is nNOS a friend or foe? 48 1.6.5: nNOS knockout mice 50

1.6: Hypoxia-inducible factor (HIF) 52

1.6.1: Discovery of HIF-1 52 1.6.2: Isoforms of HIF-α 52 1.6.3: Tissue HIF-α protein expression in vivo 54 1.6.4: Posttranslational regulation of HIF-α expression in oxygen-dependent manner Hydroxylation/ubiquitination 54 SUMOlyation 55 Acetylation 56 Phosphorlyation 56 1.6.5: Regulation of HIF-1α expression by NO in normoxia 59 1.6.6: Regulation of HIF-1α expression by NO in hypoxia 62 1.6.7: O2-independent HIF-a regulation 62 1.6.8: Other non-hypoxic activator of HIF expression 63 1.6.9: HIF-mediated adaptive responses 64 1.6.10: Clinical implication of mutation in PHD/VHL/HIF pathway 66

1.7: Thesis Objective 69 General hypothesis 69 Sub-hypothesis 69 Aims 70 1.8: References 71 Chapter 2: nNOS regulates adaptive cardiovascular responses in acute anemia 2.1: Introduction 90 2.2: Methods and Materials 92 2.2.1: Mouse Model of Acute Hemodilutional Anemia 92 2.2.2: Mouse Model of Acute Systemic Hypoxia 93 2.2.3: Co-Oximetry and Blood Gas Analysis 93 2.2.4: Mean Arterial Pressure 93 2.2.5: Carotid Blood Flow 93 2.2.6: Echocardiography 93 2.2.7: Pressure-volume Loops 94 2.2.8: Vascular Reactivity 95

2.2.9: Microvascular Tissue Oxygen Tension 96 2.2.10: Data Analysis 97

2.3: Results 2.3.1: nNOS is protective in acute anemia 98

2.3.2: nNOS regulates the cardiovascular responses in acute anemia 101

vii

2.3.3: nNOS limits the reduction in global oxygen delivery during acute anemia 108 2.3.4: Regulation of brain PO2 in acute anemia is independent of nNOS 108 2.3.5: nNOS did not impact co-oximetry results in anemia 112 2.3.6: nNOS did not affect blood gas status in anemia 112 2.3.7: nNOS may mediate increase in methemoglobin levels in anemia 114

2.4: Summary of main findings 116 2.5: Reference 118 Chapter 3: nNOS is required to increase HIF-1α protein expression in severe anemia 3.1: Introduction 121 3.2: Methods and Materials 123 3.2.1: Acute hemodilutional anemia 123 3.2.2: Acute hypoxia 123

3.2.3: Western Blot 123 3.2.4: Immunofluorescence staining 124 3.2.5: Quantitative Real-Time PCR 124

3.2.6: Plasma EPO measurement by enzyme linked immunosorbent assay 127 3.2.7: Breeding and Genotyping Strategy of HIF-(ODD)-luciferase mice and nNOS-/- mice 127 3.2.8: In Vivo Bioluminescent Imaging 129 3.2.9: In Vitro Luciferase Assay 129 3.2.10: Biotin Switch Assay 130 3.2.11: Statistical Anaylsis 131

3.3: Results 3.3.1: nNOS promotes HIF-1a protein expression in the brain of anemic animals 132 3.3.2: nNOS regulates HIF-dependent genes in anemia 136 3.3.3: nNOS did not influence plasma erythropoietin level in anemia 139 3.3.4: HIF-α(ODD)-luciferase mice in anemia 141 3.3.5: nNOS is required for total body HIF-1α activity increase in acute anemia 143 3.3.6: nNOS is required to increase HIF-1α in the anemic brain 146 3.3.7: Whole body and tissue HIF lucifearse activity at different Hb threshold 149 3.3.8: nNOS leads to S-nitrosylation of pVHL may be responsible for HIF-1α stabilization in anemia 151 3.3.9: Anemia reduces the level of GSNO reductase 154

3.4: Summary of main findings 156 3.5: Reference 157 Chapter 4: Discussion and future direction 160

Mechanisms of survival in anemia 163 Traditional physiological mechanisms of survival during anemia 163 nNOS-dependent mechanisms of survival during anemia 164 Stabilization of HIF-1α during anemia by non-hypoxic and hypoxic mechanisms 171 A novel pathway for SNO signaling in anemia 177

References 181

viii

List of Tables

Table 1.1: Basal O2 extraction of different organs.

Table 2.1: Co-oximetry and blood gas analysis from anemic and hypoxic mice.

Table 3.1: Primers used in real-time PCR Table 3.2: PCR primers used for genotype of HIF-α(ODD)-luciferase and nNOS

ix

List of Figures

Chapter 1: Figure 1.1: Oxygen dissociation curve (ODC). Figure 1.2: Oxygen molecule from inspired air to mitochondria. Figure 1.3: Oxygen gradient from blood to mitochondria in the brain. Figure 1.4: Different arterial to microvascular brain tissue PO2 gradients in anemia and

hypoxia Figure 1.5: Regulation of glucose metabolism in normoxia (aerobic) and hypoxia (anaerobic). Figure 1.6: Biological actions of nitric oxide (NO) Figure 1.7: The GSNO reductase (GSNOR)/GSH system. Figure 1.8: Schematic diagram of normoxic and hypoxic induction of nNOS transcript Figure 1.9: Regulation of HIF-α expression by hydroxylation in normoxia and hypoxia. Figure 1.10: Summary of the possible mechanisms by which NO can stabilize HIF-α in

normoxia by disrupting the degradation pathway. Chapter 2: Figure 2.1: Survival curve in acute anemia and hypoxia of WT and NOS-/- mice. Figure 2.2: Isoflurane survival curve of WT, nNOS-/- and eNOS-/- mice. Figure 2.3: Echocardiography measurements of the cardiovascular responses to acute

hemodilutional anemia in mice. Figure 2.4: Pressure-volume loop measurement of cardiovascular responses in mice at

moderate anemia (~90g/L). Figure 2.5: Cardiovascular responses to acute hypoxia in mice by echocardiography. Figure 2.6: Percent changes of cardiac output (CO), systemic vascular resistance (SVR), and

mean arterial pressure (MAP) relative to baseline/normoxia in anemic and hypoxic mice.

x

Figure 2.7: Phenylephrine responses and myogenic tone assessments under control and anemia conditions for WT and nNOS-/- mice.

Figure 2.8: Global oxygen delivery in acute anemia and hypoxia. Figure 2.9: Measurement of carotid blood flow and brain microvascular oxygen tension in

acutely anemic mice Figure 2.10: Measurement of carotid blood flow and brain microvascular oxygen tension in

mice exposed to normoxia and hypoxia. Figure 2.11: MetHb levels of anemic and hypoxic WT and nNOS-/- mice. Chapter 3: Figure 3.1: HIF-ODD-luciferase and nNOS genotype by PCR and visualized on agarose gel

stained with ethidium bromide. Figure 3.2: Assessment of nNOS, HIF-1α and HIF-2α protein expression in the brain of WT

and nNOS-/- in 1hr, 6hr and 24hr following hemodilution Figure 3.3: Assessment of nNOS, HIF-1α and HIF-2α protein expression in the brain of WT

and nNOS-/- in normoxia (21% O2) and hypoxia (6% O2 for 6hrs). Figure 3.4: Immunofluorescence staining of nNOS and HIF-1α in anemic and hypoxic brain Figure 3.5: Assessment of HIF-dependent mRNA expression in the brain of WT and nNOS-/-

mice 1, 6, and 24hrs anemia by real-time PCR. Figure 3.6: Assessment of HIF-dependent mRNA expression in the brian of normoxic (21%

O2) and hypoxic (6% O2 for 6hrs) WT and nNOS-/- mice by real-time PCR. Figure 3.7: Plasma EPO level in 6hr anemic mice. Figure 3.8: Whole body HIF-1α expression in anemic mice. Figure 3.9: In vivo bioluminescent imaging of anemic WT and nNOS-/- mice in dorsal and

ventral views Figure 3.10: In vivo bioluminescent imaging of hypoxic (6% O2 for 6hrs) WT and nNOS-/-

mice in dorsal and ventral views Figure 3.11: In vitro luciferase measurement of the tissue luciferase activity at baseline and

anemia in the brain, kidney, heart, and liver.

xi

Figure 3.12: In vitro luciferase measurement of the tissue luciferase activity at normoxia (21% O2) and hypoxia (6% O2) in the brain, kidney, heart, and liver.

Figure 3.13: HIF-luciferase activity in whole body and different organs at various Hb threshold

6hrs after hemodilution Figure 3.14: S-nitrosylated (SNO) pVHL level is increased in acute anemic brain in nNOS-

dependent fashion. Figure 3.15: Changes in other SNO-protein levels were not detected in anemic brain samples. Figure 3.16: GSNOR protein levels in acute anemia and hypoxia and mortality curve of WT

and GSNOR-/- mice during acute anemia Chapter 4: Figure 4.1: Non-hypoxic and hypoxic HIF-1α stabilization in acute anemia and systemic

hypoxia. Figure 4.2: S-nitrosylation of pVHL stabilization of HIF-1α in non-hypoxic manner.

xii

Abbreviations

ANGII Angiotensin II ARD1 Acetyltransferase arrest defective protein 1 ARNT Aryl hydrocarbon receptor nuclear translocator / HIF-1β ATP Adenosine triphosphate bHLH Basic helix-loop-helix cGMP Cyclic guanosine monophosphate CO Cardiac output CPK Creatine phosphokinase C-TAD C-terminus transactivation domain CXCR7 Chemokine receptor 7 DNA Deoxyribonucleic acid DFO Desforoximine EDRF Endothelial derived relaxing factor EDTA Ethylenediaminetetraacetic acid eNOS Endothelial nitric oxide synthase EPAS1 Endothelial PAS domain protein 1 / HIF-2α EPO Erythropoietin Fe2+ Iron (II) Fe3+ Iron (III) FIH Factor inhibiting HIF GAPDH Glyceraldehydes-3-phosphate dehydrogenase GLUT1 Glucose transporter 1 GSH Glutathione GSNO S-nitrosoglutathione GSNOR S-nitrosoglutathione reductase Hb Hemoglobin concentration Hct Hematocrit HEPES 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid HIF Hypoxia-inducible factor HRE Hypoxia response element iNOS Inducible nitric oxide synthase Km Michaelis constant LVEDV Left ventricular end diastolic volume LVSDV Left ventricular end systolic volume MAP Mean arterial pressure MetHb Methemoglobin MI Myocardial infarction MLC Myosin light chain MMTS Methyl methanethiosulfonate mRNA Messenger ribonucleic acid NADH Nicotinamide adenine dinucleotide NADPH Nicotinamide adenine dinucleotide phosphate NMDA N-methyl-D-aspartic acid

xiii

nNOS Neuronal nitric oxide synthase NO Nitric oxide NOSIP NOS interacting protein NTS Nucleus tractus solitarii O2 Oxygen O2

- Superoxide OONO- Peroxynitrite ODC Oxygen dissociation curve ODD Oxygen degradation domain ORF Open reading frame PCO2 Partial pressure of carbon dioxide PDK1 Pyruvate dehydrognase kinase 1 PGK Phosphoglycerate kinase PLB Phospholamin B PHD Prolyl hydroxylase domain PLG Placental growth factor PMCA Plasma membrane Ca2+/calmodulin-dependent Ca2+ ATPase PO2 Partial pressure of oxygen pVHL Von Hippel Lindau protein PVN Paraventricular nucleus ROS Reactive oxygen species RyR2 Ryanodine receptor S1P Sphingosine-1-phosphate SDF1 Stromal-derived factor 1 sGC Soluble guanlyl cyclase SENP Sentirin/SUMO-specific protease SERCA2a SR calcium pump SIRT1 Sirtuin 1 SNO S-nitrosothiol SNO-Hb S-nitrosohemoglobin SR Sacroplasmic reticulum SUMO Small ubiqutin-like modifier SVR Systemic vascular resistance TGF Tubuloglomerular feedback UTR Untranslated region VEGF Vascular endothelial growth factor

xiv

Chapter 1

Chapter 1:

Introduction

1

Chapter 1

1.1: Oxygen – The Element of Life

Since its discovery by Joseph Priestley, Carl Wilhelm Scheele and Antoine

Lavoisier in the 18th century, our understanding of elemental oxygen (O2) and its

important role in biology has continued to evolve. Oxygen is required for the survival of

aerobic cells and organisms, due to its ability to facilitate efficient production of ATP

from nutritional substrates. Oxygen levels that are too low (hypoxia) or too high

(hyperoxia) can both result in deleterious effects, thus tight regulation of O2 is essential at

the cellular level.

1.1.1: Importance of O2 to complex organisms:

Before the advent of atmospheric oxygen on Earth around 1-2 billion years ago,

anaerobic organisms predominated. With the evolution of single-celled organisms which

utilized solar energy and water to produce oxygen by photosynthesis, the pathway was

paved for development of complex aerobic organisms (metazoans). The ability of oxygen

to facilitate the efficient utilization of substrates to produce cellular energy (ATP) was

essential for metazoans development. At first, unicellular species obtained adequate

levels of oxygen via simple diffusion. However, with further evolutionary development,

sophisticated system for delivering oxygen to remote tissues was required: including an

efficient pump (heart), an effective means of distribution (circulatory system), and an

efficient oxygen carrying mechanism (hemoglobin; Hb).

The importance of Hb as an efficient oxygen carrying molecule is emphasized by

the fact that it is found throughout the plants and animal kingdoms, including eubacteria,

unicellular eukaryotes. Its ubiquitous presence within organisms and conserved amino

acid sequence suggested that the Hb genes are descended from an ancient, common

2

Chapter 1

ancestor (1). The importance of Hb in oxygen delivery will be further discussed in the

subsequent sections by defining the consequences of Hb deficiency (anemia). As stated

by Singel and Stamler (2), “a general principle of physiology holds that cells precisely

regulate their primary function. For red blood cell (RBC), this primary function is

delivery of O2 to tissues”

1.1.2: Oxygen transport in the body

Additionally, metazoans have developed a sophisticated network to regulate

oxygen delivery during times of increased oxygen demand (e.g. exercise) or reduced

oxygen availability (e.g. anemia or hypoxia): the cardiovascular system. During times of

decreased oxygen supply, adequate delivery of oxygen to tissue is facilitated by increase

in pump efficiency (cardiac output; CO) and reduced systemic vascular resistance (SVR).

The resultant integrated response enhances the delivery of oxygen to tissues in a

hierarchal manner such that organs of primary importance of survival (i.e. brain and

heart) are favored, possibly by tissue-specific vasodilation to increase organ blood flow.

Indeed, it is the tissues’ needs for oxygen that finally regulates tissue perfusion in

vascular beds. As stated by Guyton: “the single factor most responsible for the significant

linkage between metabolic rate and cardiac output is the tissue need for oxygen”(3)

Within red blood cells, hemoglobin is the key element which provides the high

oxygen carrying capacity of blood and maintains oxygen delivery to tissues, as denoted

by the equation: CaO2 = (Hb x 1.39 x SaO2) + (0.003 x PaO2), where CaO2 is arterial blood

oxygen content, Hb is hemoglobin (g/dL), SaO2 is oxygen saturation (%), PaO2 is partial

pressure of oxygen (arterial; mmHg). This determines that the majority of O2 in the blood

is bound to hemoglobin (~99.9%), and very little is dissolved as free oxygen in plasma

3

Chapter 1

(<0.01%). Thus, at atmospheric conditions, CaO2 is determined primarily by the amount

of O2 bound to hemoglobin (oxygen saturation; SaO2) and not by oxygen dissolved in the

plasma (PaO2), which serves as an interface between Hb and the cells.

The ability to globally distribute O2 is regulated primarily by cardiac output, CaO2

and peripheral blood flow. The control of O2 delivery in each tissue is determined by its

oxygen requirements. Organs with high oxygen consumption (e.g. brain and heart)

require higher rates O2 delivery to sustain tissue metabolic requirements. Mechanisms

which regulate tissue blood flow are focused on small resistance arteries and capillaries.

At this level, perfusion is controlled by regulating the diameter of these resistance vessels

via mechanisms which are both extrinsic and intrinsic to the vasculature. Extrinsic

mechanisms includes vasodilators, such as nitric oxide, can increase tissue blood flow,

whereas vasoconstrictors, such as endothelin-1, decrease tissue perfusion (4).

At resting conditions, on average only 25% of the O2 is extracted from Hb at the

tissue level as demonstrated by the mixed venous blood saturation (SvO2) of 75%. This

limited extraction is based, in part, on the physiological conditions of oxygen extraction

from Hb which is cooperative such that the first oxygen molecule is extracted with

greater ease than subsequent oxygen molecules (5). This ability to efficiently load and

offload oxygen molecule on Hb is essential for optimal oxygen delivery to tissues, and is

illustrated by the characteristic sigmoidal shape of the oxygen dissociation curve (ODC)

(Figure 1.1). The affinity of oxygen for Hb can be altered by factors such as changes in

body temperature, pH, carbon dioxide and 2,3-disphosphoglycerate (2,3-DPG). Left-shift

of ODC leads to increased oxygen loading in the lungs, whereas right shifting favors

oxygen offloading in the peripheral tissues. Of interest, anemia increases the production

4

Chapter 1

of 2,3-DPG in RBC, thereby the ODC is right shifted and O2 delivery to tissues is more

favorable (6). In addition, O2 extraction during anemia increases very rapidly, suggesting

a rapid right shifting of ODC, by alternate mechanisms, which facilitate increased O2

unloading during anemia (7). This results in decreased oxyhemoglobin saturation (i.e.

deoxyhemoglobin), favoring the tense (T) configuration of Hb, exposure of S-

nitrosothiols (SNO) on Hb and its subsequent transfer to other proteins. This mechanism

of NO release from Hb by O2 level had been proposed as a means to increase blood flow

in vascular beds (8, 9).

In the pulmonary circulation, the opposite occur to favor O2 loading onto Hb. In

the lung, NO can bind to Hb and re-form S-nitrosohemoglobin (SNO-Hb) such that the

ODC is left-shifted to facilitate higher affinity of O2 on oxygenated Hb and increased O2

uptake in pulmonary circulation. After O2 is picked up by Hb at the lungs, it is carried by

the circulation to the tissues. The blood PO2 reduces from large to small vessels. For

example, recent oxygen quantitative measurements in the brain have demonstrated that

there is a continuous oxygen gradient from the arteriole (80mmHg) to the capillary (60-

30mmHg) (10-12), and then the interstitial tissue (~30mmHg) (13), into the cells

(20mmHg) (14, 15) and finally to the mitochondria (~10-15mmHg) for oxidative

phosphorylation, where generation of ATP occurs aerobically (16). This measurement of

intracellular PO2 is higher than previously reported (~2mmHg) by numerous

experimental studies (17). However, more recent accurate quantitative measurement

demonstrated that the cellular PO2 where the mitochondria exist is near 10-15mmHg.

This enables the mitochondria to function in physiological range, and perform function

such as oxygen sensing (10, 13, 15, 18).

5

Chapter 1

6

Figure 1.1: Oxygen dissociation curve (ODC). Changes in pH, 2,3-DPG and temperature can affect the dissociation of oxygen to Hb. Left shift of ODC leads to increased oxygen loading, whereas right shift results in increased oxygen off-loading. Despite studies have demonstrated that nitric oxide (NO) species (right shift with FeNOHb (19), and left shift with SNO-Hb (20)) can affect the ODC curve, these NO species are not likely to meaningfully influence the ODC in vivo. Rather, the O2 is the principle regulator for interaction between NO and Hb. (Modified from: http://www.bio.davidson.edu/Courses/anphys/1999/Dickens/Oxygendissociation.htm)

Chapter 1

1.1.3: Acute oxygen sensing in mammals

The importance of oxygen to all aerobic organisms (single cell to mammals) is

indicated by the development of multiple redundant mechanisms for detecting reduced

tissue PO2. Candidate mechanisms include the mitochondria which has long considered

as a potential site for oxygen sensing through the process of oxidative phosphorylation

and electron transport: a process which can be altered by changes in oxygen level (21).

Reactive oxygen species (ROS) produced in the mitochondria are increased in hypoxia

primarily by complex III (22). High ROS levels provides favorable environment for the

oxidation of ferrous iron (Fe2+ to Fe3+), resulting in prolyl hydroxylase domain (PHD)

inhibition and thus leading to hypoxia-inducible factor (HIF)-α stabilization and increase

transcription of hypoxic genes to improve O2 delivery (23).

In addition, the early evolution (600 million years) and ubiquitous presence of the

HIF/PHD pathway in all cells, both single and complex organisms, suggests that

detection of cellular or tissue hypoxia was a requirement for survival and development of

the earliest aerobic species (24). The strong conservation of the HIF pathway throughout

evolution emphasizes its importance in aerobic organisms, including mammals. The

importance of oxygen sensing is emphasized by the redundancy of mechanisms designed

for the detection of hypoxia in the body. In mammals and humans, these mechanisms

include hypoxia sensors at the level of the organ (kidney), organelle (chemoreceptors),

and cell (hypoxia inducible factor [HIF] and prolyl hydroxylase [PHD]). Together, they

make up a specialized homeostatic oxygen-sensing system which detect reduced O2

tension and trigger adaptive responses.

7

Chapter 1

Kidney as an oxygen sensor at the organ level

For example, at the organ level, the kidneys can act as a sensitive hypoxia sensor

that detects small decreases in PO2 levels associated with anemia (10, 25, 26). The

resultant production of HIF-dependent erythropoietin (EPO) to restore the Hb level

demonstrates the exquisite sensitivity of this mechanism.

Chemoreceptors and resistance arteries as oxygen sensor at organelle level

At the level of the chemoreceptor, anemia is known to activate aortic

chemoreceptors in response to a reduction in Hb concentration (27). In addition,

hypoxemia (PaO2 < 60mmHg) is detected by carotid body chemoreceptor (28). These

mechanisms trigger neurosecretion of acetylcholine, catecholamines and dopamine,

increase action potential in the carotid-sinus nerve, and activate the respiratory center in

the medulla oblongata of the brain (28). In addition, decreased oxygen level in inspired

air can be sensed by airway neuroepithelial bodies to centrally control respiration (29).

Overall, the activation of chemoreceptors and neuroepithelial bodies in response to

reduced O2 level stimulates respiration. In addition, the resistance artery may also

function as an oxygen sensing organelle, where the vascular smooth muscle acts as the

oxygen sensor. For example, hypoxic pulmonary vasoconstriction, a mechanism to

maximize ventilation and perfusion matching in the lungs, is mediated in part by changes

in smooth muscle membrane depolarization which leads to myosin light chain (MLC)

phosphorylation (28), and reduction in eNOS-derived NO level in hypoxic pulmonary

vasculature (30, 31). By contrast, in peripheral vascular beds, blood vessels dilate in

response to hypoxia. This is well demonstrated in cerebral and coronary vessels (32, 33),

8

Chapter 1

whereby increased perfusion and oxygen delivery is mediated by the KATP channels.

Under hypoxic conditions where ATP levels are reduced, opening of KATP channels leads

to hyperpolarization and relaxation of vascular smooth muscle cells (32). In addition,

vasodilators are involved in response to hypoxia, such as nitric oxide (NO), CO2,

adenosine, and prostaglandin (33). Also, smooth muscle neuronal nitric oxide synthase

(nNOS) is activated in response to low oxygen that produced NO to cause hypoxic

vasodilation (34).

PHD/HIF cellular pathway as oxygen sensor at tissue level

At the level of tissue, the ability of all cells to act as hypoxia sensor is provided by

the presence of hypoxia inducible factor (HIF). Prolyl hydroxylase domain (PHD) of HIF

is regarded as an important oxygen sensor (35). When O2 is plentiful, PHD uses oxygen

and α-ketoglutarate to hydroxylate proline residues located in oxygen-dependent

degradation domain of HIF-α, thereby favoring interaction with von Hippel Lindau

protein (pVHL), and thus HIF-α is ubiquitinated and degraded. In hypoxic cells, the

deprivation of oxygen results in reduced PHD2 activity, thus pVHL cannot not bind to

non-hydroxylated HIF-α, and hence, HIF-α escapes the degradation pathway and

stabilized. Accumulation of HIF-α results in activation of genes that involve in

erythropoiesis, angiogenesis, energy metabolism, and cell proliferation to increase

oxygen delivery. Biochemical studies revealed that PHDs have high Km to oxygen (~230-

250 µM) (36), suggesting that only a small drop in oxygen would render inhibition of

PHD enzymatic activity. Activation of these cellular pathways ensures adequate supply

of O2 to tissues.

9

Chapter 1

Red blood cell as a potential oxygen sensor

Extensive experimental evidence has demonstrated that Hb itself may act as a

form of hypoxia sensor to increase blood flow in hypoxic tissues. This mechanism is

facilitated by the close proximity of Hb to vascular smooth muscle cell within resistance

arteries. As early as 1996, the role of red blood cell (RBC) and Hb in hypoxic

vasodilation has been suggested. The SNO-Hb hypothesis proposed by Stamler et al.

suggested that as Hb O2 saturation decreases, Hb changes conformation, thus the NO

groups that binds to cysteine thiol in β-globin chain (S-nitrosothiol) of Hb is exposed to

solvents such as glutathione to become S-nitrosoglutathione (8, 37). The release of NO

groups leads to vasodilation. An alternate hypothesis has suggested the role of Hb as

nitrite reductase that the reduction of nitrite to NO is more favorable during Hb

deoxygenation (38-40). Thus, Hb likely plays a central role in mediating hypoxic

vasodilation. By such a mechanism, deoxyhemoglobin increases NO bioavailability and

causes peripheral vasorelaxation in hypoxic vascular beds to ensure adequate tissue O2

delivery.

The importance of nitric oxide synthases (NOSs) as necessary sources of NO to

prime the SNO-Hb system requires consideration. Although all three NOSs may

contribute to vascular NO sources of SNO, experimental evidence favors eNOS and

nNOS as important sources of NO. With respect to eNOS expression, anemia and

hypoxia result in unchanged (14) or decreased (30, 31) level of eNOS, respectively. This

would suggest that eNOS is not an important source of NO during anemia. By contrast,

nNOS, a relatively new candidate for cardiovascular regulation, is increased in the

10

Chapter 1

cardiovascular system and the central nervous system during both anemia and hypoxia

(14, 34, 41). Therefore, we explored the specific role of nNOS as a mediator of vascular

NO and SNO protein modification in a model of oxygen substrate limitation (anemia)

11

Chapter 1

1.2: Anemia

Anemia is a worldwide health problem that affects 1.6 billion people and has a

global prevalence in general population of 25% as of a report published by World Health

Organization (WHO) in 2005 (42). Anemia is defined as a reduction in hemoglobin

concentration (Hb) or decrease amount of red blood cell (RBC). Since oxygen is

transported by Hb throughout the body, anemia leads to a decreased in oxygen carrying

capacity of blood, and, ultimately, this reduces the delivery of oxygen to tissues. The

need of oxygen is very crucial to all living organisms that rely on aerobic metabolism.

In humans, since baseline Hb varies with age, gender and ethnic groups, WHO

defines anemia as the Hb threshold value at 2 standard deviation from the mean or the

2.5th percentile of the normal distribution of the healthy population (normally, <130g/L

for men and <120g/L for women). Anemia is caused by a diverse number of

pathophysiological conditions including genetic defects, nutritional deficiency, infectious

diseases, malignancy, chronic inflammatory diseases and acute blood loss and fluid

resuscitation (hemodilution), secondary to trauma and surgery. Regardless of the type of

anemia, a common mechanism of anemia-induced morbidity and mortality is believed to

be a reduction in tissue oxygen delivery, and subsequent hypoxic organ injury. A more

complete understanding of hypoxia signaling suggest that S-nitrosothiols may be an

additional “non hypoxic” mechanism by which mammals adapt to anemia. The relative

importance of limited oxygen delivery (tissue hypoxia) and other “non-hypoxic”

mechanisms (S-nitrosothiol signaling) will be evaluated with respect to the

pathophysiology of anemia-induced mortality throughout the thesis.

12

Chapter 1

1.2.1: Do all types of anemia share a common mechanism for morbidity and mortality?

Although all forms of anemia are associated with increased mortality (renal

disease, cancer, sickle cell, trauma, surgery), it is unclear whether anemia is merely one

aspect of multi-organ failure in a series of complex diseases, or whether anemia leads to

decrease in oxygen delivery and hypoxic cell injury and death. Indeed, some authors hold

the extreme position that anemia may be protective and adaptive (43). If the primary

mechanism for anemia induced morbidity or mortality is the limitation of oxygen

delivery, then all forms of anemia should predict mortality at similar Hb thresholds.

Unfortunately, such a simple relationship does not exist. For example, in chronic renal

failure, patients can survive for prolonged times with relatively low Hb values.

Paradoxically, correction of anemia with erythropoietin stimulating agents (ESAs) does

not appear to improve mortality in this patient population (44-46). Thus, increasing blood

oxygen content and tissue oxygen delivery did not impact outcome. By contrast,

transfusion of children with sickle cell anemia significantly reduces the incidence of

stroke in these patients (47). These results suggest that increasing blood oxygen content

does improve tissue oxygen delivery, thereby reducing hypoxic brain injury. A clearer

understanding of the mechanisms involve will be required to unravel the mechanism of

organ injury.

1.2.2: Risks of Anemia

Anemia is associated with increased risk of morbidity and mortality in patients

with HIV(48), malaria (49, 50), sickle cell (51) and surgical patients especially those with

cardiovascular diseases (52, 53), renal disease (54-56), cancer (57, 58), critically illness

13

Chapter 1

(59, 60), and healthy mothers (61, 62). Although treatment of anemia often involves

blood transfusion and erythropoiesis stimulating agents, such as darbepoietin and

erythropoietin, it is now appreciated that these therapies do not necessarily improve

morbidity and mortality (44-46, 63, 64). These findings underscore the needs to define

the underlying physiological and molecular mechanisms in anemia such that problems in

anemic patients can be addressed.

Risks of brain injury

The brain is unique in that it represents a vital organ with relatively high oxygen

requirement in which a small injury can have disproportional adverse effect. In any

setting, brain injury is associated with an unexpectedly large increase in mortality (65-

68). The link between anemia, oxygen delivery, organ function and organ injury has been

most closely determined with respect to the brain. Acute and chronic anemia are

associated with cognitive decline and learning disabilities, suggesting that the associated

inadequacy of oxygen delivery impairs neuronal function (69-71). In addition, anemia

from multiple etiologies is associated with brain injury. For example, children with sickle

cell anemia who have high cerebral blood flow have an increase incidence of stroke,

which can be prevented by blood transfusion (71). Severe malaria result in neurological

injury (cerebral malaria), which is associated with a high mortality rate (72). Finally, the

lowest Hb associated with hemodilution on CBP is associated with increased incidence of

stroke that is proportional with reduced Hb concentration (66, 67).

14

Chapter 1

Risks of renal failure

Treatment of patients with chronic renal insufficiencies did not demonstrate

preserved renal function once anemia is corrected (45, 46). However, acute reducing in

Hb is associated with increased renal failure. An increase in the incidence of acute renal

failure has also been demonstrated in surgical patients who undergo hemodilution (54-56,

73). During hemodilution, experimental models demonstrated that the kidney becomes

hypoxic at much higher hemoglobin concentrations than the brain or heart (10, 26, 74).

This may partially explain the increased incidence of renal failure in these anemic

patients.

Risks of myocardial injury

During anemia, the myocardium maybe at particular risk for ischemic hypoxic

injury because it is the only organ in which O2 consumption is clearly increased as a

result of increased heart rate, contractility and cardiac output to maintain global O2

delivery during anemia (75). This may explain the large number of clinical studies which

have demonstrated that anemia is a predictor of mortality in patients with congestive

heart failure (76-79). In addition, perioperative patients treated with β-blocker therapy

which limit the heart rate response to anemia have been shown to have an increase in

myocardial infarction. This may relate to the impact on β-adrenergic blocker on coronary

perfusion, however the mechanism is as of yet undetermined. Recently, clinical data is

emerging that acute perioperative anemia may lead to an increased incidence of

myocardial infarction (80). The mechanism may include impaired coronary vasodilation

15

Chapter 1

(reduced coronary reserve) during acute anemia (75, 81). In addition, endothelial function

is impaired in anemia associated with sickle cell disease and hemodilution. The

mechanism of injury appears to be mediated by impairing nitric oxide (NO)

bioavailability (82-85).

Collectively, acute and chronic anemia are associated with increased mortality

and vital organ injury. A causal relationship in anemia, reduced oxygen delivery and

increase morbidity and mortality remains to be established. Therefore, it is important to

understand the adaptive physiological and cellular mechanisms which enable mammals to

tolerate anemia.

1.2.3: Traditional view of the physiological adaptation to acute anemia Cardiovascular adaptation to anemia

During acute anemia, the sudden drop in blood oxygen content may be sensed at

multiple redundant levels. For example, anemia causes an increase in cellular HIF

expression, raising the possibility that every cell could act as a HIF sensor during anemia.

This has always been assumed to be hypoxia-mediated. Importantly, this thesis directly

tests this hypothesis. At the level of the organelle, efferent nerve activity from aortic

chemoreceptors increases in proportion with the degree of anemia (86, 87). By contrast,

studies have not been able to demonstrate an increase in carotid body chemoreceptors

(27). In addition, organs such as the kidneys are viewed as traditional sites of hypoxia

sensing. This may be facilitated by the fact that the EPO-producing cells located in the

kidney. In addition, the kidney becomes hypoxic earlier during anemia (10), suggesting it

may sense reduced Hb concentration and tissue hypoxia at an earlier stage of anemia.

16

Chapter 1

Once tissue hypoxia is sensed, the associated efferent signals may trigger central

neuronal response which activates the adrenergic nervous system and increase

sympathetic activity (27). This mechanism is known to contribute to increase cardiac

output (CO), stroke volume (SV) and heart rate (HR) during anemia, as demonstrated by

cardiac denervation experiment (87). The composite effect is to increase CO and

maintain mean arterial pressure (MAP) until very low Hb levels. In addition, active and

passive mechanisms lead to the associated reduction in systemic vascular resistance

(SVR), which further promote O2 delivery to tissues (10, 88, 89). These cardiovascular

responses are consistently observed in humans and animals and act to maintain a global

balance of O2 delivery and consumption (69, 75). These mechanisms have been credited

with maintaining adequate O2 delivery on a global level. Measurements of whole body

oxygen consumption (AV difference) demonstrate that overall oxygen utilization is

maintained until very low Hb level (near 50g/L) (90). These types of studies define the

critical oxygen delivery as the Hb concentration below which oxygen consumption falls.

This concept demonstrates that O2 delivery is maintained. However, they do not explain

the incidence of mortality and morbidity at higher Hb level. Assessment of O2 delivery

and consumption has not clearly determined the kinetics of O2 delivery at the

microcirculatory level.

Another important component of the cardiovascular component to anemia is that

blood flow is not distributed equally to every organ. Indeed, organs receive preferential

blood flow in a hierarchal manner. Organs with high O2 extraction (e.g. brain and heart)

require higher rates of O2 delivery to sustain tissue metabolic requirements would receive

17

Chapter 1

larger blood flow (Table 1.1). This observation suggested that blood viscosity may not be

the primary driving force to increase vital organ blood flow.

Organ Basal O2 extraction (%)

Brain ~34%

Heart ~68%

Kidney ~16%

Portal system (GI tract, pancreas, spleen, liver)

~21%

Table 1.1: Basal O2 extraction of different organs. Values adapted from Wolff CB 2007 (91).

Respiratory adaptation to anemia

Anemia stimulates respiration, resulting in increased minute ventilation. In

addition, NO-mediated mechanisms improve ventilation and perfusion matching leading

to a characteristic rise in PaO2 and SaO2 (92, 93). These responses ensure that optimal

SaO2 is maintained at a time of reduced Hb to maximize blood oxygen content.

Metabolic adaptation to anemia

In addition, alteration in the metabolic requirement helps to preserve the overall

balance of systemic oxygen supply and demand during anemia (69, 75). For example,

increase in myocardial oxygen utilization results in reduction in tissue O2 consumption in

other organs (75, 90). While O2 consumption is maintained in the brain (94), it decreases

18

Chapter 1

in the kidney during anemia (26). Such mechanisms are activated in hypoxia (95), and

may contribute to adaptive changes to promote survival in acute anemia. These

mechanisms are dependent, in part, on HIF-mediated metabolic responses (96). The

activation of these adaptive responses ensures tissue O2 homeostasis and the balance of

O2 supply and demand is optimized during anemic stress (Figure 1.2).

Increased tissue oxygen extraction during anemia

Experimental studies in humans and animals demonstrated that global and tissue

specific O2 delivery is maintained during acute anemia, in part, through increased

systemic oxygen extraction (7, 69). This mechanism may be less important to organs with

high basal oxygen extraction rates, such as the heart, which are more dependent on

increased flow to match increased oxygen consumption during anemia (91). By contrast,

oxygen extraction in the brain increases from about 30% at baseline to near 50% during

anemia under experimental conditions (7, 10). This increase in oxygen extraction likely

depends upon at least three important factors: 1) A right shift of the oxyhemoglobin

dissociation curve to reduced Hb oxygen affinity which may occur by a number of

mediators including, decreased pH, increased 2,3-DPG and NO-mediated signaling

events (5, 9); 2) increased tissue blood flow which favors increased oxygen diffusion into

the tissue (25); and 3) Increased capillary recruitment and density may limit the diffusion

distance to cells during anemia (97). The primary goal of these mechanisms is to

facilitate oxygen diffusion from the microcirculation to the tissues, thereby sustaining

mitochondrial oxidative phosphorylation (aerobic respiration).

19

Chapter 1

Figure 1.2: Oxygen molecule from inspired air to mitochondria. Oxygen from the air (~150 mmHg) is inspired by the lungs (~100 mmHg), binds to hemoglobin (Hb) in the blood and propels to the rest of the body by the heart. Oxygen in the microvasculature (~30-60 mmHg) (10-13) is diffused through gradient to tissue (≥ 30mmHg) (14, 15), cell (10-20 mmHg) and, ultimately, mitochondria (17) for generation of ATP. Note that the oxygen level is reduced at each step, emphasizing the gradient of oxygen from inspired air to single cell. In anemia, a decreased in Hb in blood leads to reduced blood O2 content, which is sensed at the level of tissue, organelles (aortic chemoreceptor), and organs (kidneys). This activates responses to maintain tissue O2 delivery.

20

Chapter 1

1.2.4: Tissue Oxygenation in Acute Anemia

Quantitative non-invasive measurements of microvascular tissue oxygen tension

in anemia models demonstrated that acute anemia reduces microvascular PO2 in the

brain, heart, kidneys, intestine and muscle (7, 10, 26, 74, 98). Reduced tissue PO2

occurred in a hierarchical manner in which organs that are critically important for

survival (heart, brain) receive preferential oxygen delivery and maintain tissue

“normoxia” at very low Hb levels (~35-50g/L) (7, 10, 74). This is consistent with the

relatively disproportional increases in heart and brain blood flow during acute

hemodilutional anemia (7, 99). Conversely, less vital organs (kidney, intestine) become

hypoxic at much higher Hb levels (~60-70g/L) (7, 10, 74).

Experimental studies have focused on the brain, a vital organ with high metabolic

requirement which is at risk of injury during acute and chronic anemia. With earlier

proposition suggested that anemia increases cerebral and coronary blood flow and tissue

PO2 (43), many clinicians regarded this as “luxury” perfusion and therefore proposed

that acute hemodilutional anemia is beneficial. Attempt to demonstrate this concept was

reported in studies with the use of acute hemodilution as a strategy for blood conservation

(100), treatment of stroke (101), and acceptance of low hemoglobin threshold (102).

However, the failure to improve patient outcome in these studies suggested that the

beneficial effects is outweighed by the risks of hemodilution. Thus, it is important to

assess tissue oxygen tension in anemia.

21

Chapter 1

1.2.5: Cellular Response to Anemia

Despite the physiological and cardiovascular adaptation to acute hemodilutional

anemia are well studied, paucity of studies had assessed the molecular responses to

anemia. In animal models of both acute and chronic anemia, a common sets of hypoxic

genes are activated, including neuronal nitric oxide synthase (nNOS), hypoxia-inducible

factors (HIF-α), erythropoietin (EPO), and vascular endothelial growth factor (VEGF),

inducible nitric oxide synthase (iNOS), and glucose transporter-1 (GLUT1) (14, 41, 97,

103, 104). Upregulation of these cellular responses may optimize tissue oxygen delivery.

Thus, it is of interest to study how anemia affects hypoxia signaling to gain insight into

the molecular mechanisms of acute anemia. Chapter 3 will focus on this aspect.

22

Chapter 1

Figure 1.3: Oxygen gradient from blood to mitochondria in the brain. Using various methodology to assess oxygen level, arteriole has a PO2 of ~80mmHg, red blood cell (RBC) has a PO2 of ~60 mmHg (10-12), which is lowered to ~30 mmHg in the plasma (13). At the brain tissue, PO2 is approximately 20 mmHg (14, 15). PO2 is the lowest in the mitochondria and can range from 10 – 15 mmHg (17). The gradient of oxygen allows oxygen to diffuse from RBC to mitochondria for the production of ATP. In acute anemia (Hb ~60-80g/L), decreased in blood O2 content leads to reduced tissue PO2. This triggers adaptive mechanisms to maintain O2 homeostasis, including right-shift of ODC curve to favor oxygen release from Hb (5, 6), activation of cardiovascular responses and cellular responses (HIF).

23

Chapter 1

1.3 Hypoxia

Hypoxia is a pathological condition in which the body or tissue is deprived of O2.

This usually occurs when O2 supply is inadequate to meet its demand for normal cellular

process. By definition, hypoxia refers to a decrease partial pressure of O2 (PO2).

Hypoxemia refers to a low partial pressure in the blood (PaO2 <60 mmHg), whereas

tissue hypoxia refers to a lower than normal PO2 in the tissues. Hypoxemia and

subsequent tissue hypoxia can be the result of reduced fractional inspired oxygen content

(FIO2) which can occur during anesthesia when supply gas mixture is lower than required

or at high altitude in which low barometric pressure reduces the PaO2. Additional causes

of hypoxemia (PaO2 <60 mmHg) are alveolar hypoventilation, impaired

ventilation/perfusion (V/Q) matching, shunting the blood from the right to left side of

circulation bypassing the pulmonary circulation, and finally the impairment of O2

diffusion from the alveolus to the capillary at the lung. Hemoglobinopathies (e.g.

carboxyhemoglobin and methemoglobin) has also been thought to be sources of

hypoxemia. However, these conditions are more correctly considered to be disorders of

oxyhemoglobin binding and low oxyhemoglobin saturation (i.e. oxyHb/ [oxyHb +

deoxyHb + carboxyHb + metHb]) as the PaO2 can be well above 100mmHg. At the

tissue level, hypoxemia results in inadequate tissue oxygen delivery. In the blood,

decreased CaO2 in hypoxia is characterized by reduced PaO2 and SaO2, but maintained

Hb. Unlike hypoxia, anemia decreased CaO2 by reduced Hb level but PaO2 and SaO2 are

maintained. The reduced Hb level limits the amount of O2 that can be carried in the blood

and subsequently transported to tissues. Thus, both hypoxia (hypoxemia) and anemia

(low Hb) can cause tissue hypoxia but potentially by different mechanisms. The existence

24

Chapter 1

of different O2 gradients from arterial blood to tissue to cell between anemia and hypoxia

(Figure 1.4) may activate different sets of adaptive responses, which will be further

investigated in Chapter 2 and Chapter 3.

Figure 1.4: Different arterial to microvascular brain tissue PO2 gradients in anemia and hypoxia.

25

Chapter 1

1.3.1: Physiological adaptations to hypoxia

The ability of mammals to maintain oxygen homeostasis is essential in survival.

Aerobic organisms require O2 to produce energy in the form of ATP (Figure 1.5).

However, in low O2 conditions, cells produce ATP very inefficiently via anaerobic

metabolism, and lactate is generated as a by-product. Reduced O2 supply in systemic

hypoxia triggers various acute and chronic adaptive responses in an attempt to match O2

consumption delivery in the body. These responses may vary depending on the species,

duration, and severity of hypoxia.

In general, the acute responses are of rapid onset and of short duration, whereas

the chronic responses are delayed in onset and longer in duration. These responses to

hypoxia can be activated by several O2 sensing systems: 1) carotid chemoreceptor

activation leads to increase ventilation through increases in respiratory rate and tidal

volume, which is termed hypoxic ventilatory response (105). In addition, a newer model

of hypoxic ventilation postulated that Hb can also release molecules derived from nitric

oxide (NO) to stimulate neuronal control of ventilation in hypoxia (106). 2) The

pulmonary vasculature is modified to downregulate hypoxic pulmonary vasoconstriction.

This physiological mechanism is derived to limit perfusion of hypoxic alveoli and to

improve V/Q matching under physiological condition (107). However, at high altitude

where PaO2 is low, all alveoli become hypoxic, leading to excessive pulmonary

vasoconstriction. Thus, in altitude, the adaptive response is to downregulate this

mechanism. 3) Peripheral vasculature stimulates expression of VEGF and its receptors

(108) to promote angiogenesis, especially in the heart (109, 110) and brain (111). 4)

Kidney increases EPO expression to initiate the process of increase in RBC mass and Hb

26

Chapter 1

levels (112-114). 5) At the tissue level, metabolic switch from oxidative phosphorylation

to glycolysis can be initiated by increased HIF expression to maximize ATP production

per O2 and per mole of glucose (Figure 1.5) (35). These acute responses provide a quick

onset, but short duration, for the body to temporary maintain O2 delivery in hypoxia.

As hypoxia persists, the acute changes cannot be sustained and thus a different set

of adaptation is initiated for prolonged hypoxia. The characteristic difference between

acute and chronic response to hypoxia highlight the underlying molecular mechanisms.

Acute responses involve post-translational modification of existing proteins (such as HIF)

to affect its activity, whereas chronic responses involve the regulation of genes at the

transcriptional and post-transcriptional level. In prolonged hypoxia, inhibition of global

protein synthesis occurred to conserve ATP (115, 116). Although induction of HIF

response genes occurs rapidly, the processes of making blood vessels (angiogenesis;

VEGF) or a red blood cell (erythropoiesis; EPO) require days to complete. In addition,

genetic reprogramming at the metabolic level can alter fuel preference in prolonged

hypoxia (117). Thus, these chronic responses are activated to sustain cellular O2 delivery

in prolonged hypoxia. Remarkably, highlanders, such as Andean, Himalayan and

Tibetans, are well adapted to high altitude hypoxia due to the ability to reduce acute

effects of hypoxia (anaerobic metabolism) (116, 117) and their altered genetics

(discussed in section 1.6.10). Collectively, these acute and chronic responses are directed

to maintain the organisms’ survival in limited O2 environment.

27

Chapter 1

28

Figure 1.5. Regulation of glucose metabolism in normoxia (aerobic) and hypoxia (anaerobic). Glucose is metabolized to pyruvate by glycolytic enzymes. In the presence of oxygen (normoxia), pyruvate is converted to acetyl CoA by pyruvate dehydrogenase (PDH). Tricarboxylic acid cycle (TCA) is initiated in mitochondria, and electrons generated in TCA are transported to electron transport chain (ETC). This uses molecular oxygen to generate 38 mol ATP per 1 mol glucose. In settings of oxygen deprivation (hypoxia), pyruvate dehydrogenase kinase (PDK1) inhibits PDH, and lactose dehydrogenase A (LDHA) converts pyruvate to lactate. The shift of pyruvate away from the TCA that yield a net of 2 mol ATP per 1 mol glucose. Modified from Semenza GL. New England Journal Medicine, 2011.

Chapter 1

1.4 Nitric Oxide

Nitric oxide (NO) is a small, freely diffusible gas that is synthesized by NO

synthases (NOS) from L-arginine. This reaction required NADPH and molecular oxygen

to yield NO and L-citrilline as products. Citrilline can also be recycled to arginine to re-

enter the reaction. The discovery that NO is endothelium-derived relaxing factor (EDRF)

laid the ground to elucidate the mechanism of NO as a vasodilator by activating

guanylate cyclase (118). In 1998, Drs. Furchgott, Ignarro, and Murad were awarded the

Nobel Prize in Physiology and Medicine for their discoveries on “nitric oxide as a

signaling molecule in the cardiovascular system”. Not only does NO play an important

role in the cardiovascular system, but it is also essential to other parts of the body, such as

the respiratory, renal, reproduction, inflammation, and central nervous systems (119). NO

can be either protective or harmful, which depends on the quantity produced, local

environment, stimulus, and the NOS isoform it is derived from (120). Both the increase

and decrease NO level is attributed to pathological conditions, suggesting tight regulation

of NO is essential. For instance, impaired NO production is associated with

cardiovascular diseases and endothelial dysfunction. The use of NO donors to treat

ischemic heart disease, heart failure and hypertension greatly improves symptom of these

patients (121). In contrast, excessive production of NO can also lead to pathological

conditions, such as ischemic stroke, septic shock, and asthma. NOS inhibitors may be

useful in reducing the severity of the disease, but all NOS inhibitors lack specificity to

target the specific NOS isoforms and cell type. Interestingly, mortality increased

significantly despite blocking NOS activity in septic patients to correct blood pressure,

highlighting that NO may have beneficial effects in other tissues during septic shock

29

Chapter 1

(122, 123). Under physiological conditions, NO can scavenge free radicals such as

superoxide (O2-) to reduce the toxicity. But in pathophysiological state such as sepsis,

excess amounts of O2- reacts with NO to produce highly reactive species such as

peroxynitrite (OONO-), leading to oxidative damages through tyrosine protein nitration

(124)(Figure 1.6). Therefore, tight control of NO in the body is important to maintain

normal biological function.

Figure 1.6: Biological actions of nitric oxide (NO). 1) NO can bind to heme of soluble guanlyl cyclase (sGC) to exert vasodilative effects of NO. 2) NO can bind to cysteine of the globin portion at Cysβ93 of hemoglobin (SNO-Hb). During deoxygenation, Hb preloaded with SNO will be released to react with other solvents. 3) NO can also form SNO protein at cysteine residues to alter its protein function. 4) NO can be stored as nitrite (NO2

-) or nitrate (NO3-), which can release NO and generate methemoglobin

(MetHb) with deoxyhemoglobin. This chemical pathway may not be physiological relevance in vivo as suggested by other authors(125). 5) NO can react with superoxide (O2

-) to form peroxynitrite (ONOO-), which can result tyrosine nitration to damage cells.

30

Chapter 1

1.4.1. Nitric oxide synthases (NOS) There are three isoforms of NOS in mammals: neuronal NOS (nNOS, NOS1),

inducible NOS (iNOS, NOS2), and endothelial NOS (eNOS, NOS3). They are capable to

catalyze the reaction from L-arginine to produce NO. Each of the NOS enzymes require

tetrahydrobiopterin (BH4), heme, flavin mononucleotide (FMN), flavin adenine

dinucleotide (FAD) and nicotinamide adenine dinucleotide phosphate (NADPH) as co-

factors for the reaction. The NOS isoforms are unique to each other with respect to their

cell type specific localization and external factors for gene regulation. nNOS is found in a

variety of cell types, including neurons, vascular smooth muscle, skeletal muscle, and

cardiomyocytes. eNOS is expressed abundantly in the endothelium. iNOS is mostly

found in macrophages. In general, nNOS and eNOS are basally expressed and their

activation is calcium dependent, whereas iNOS expression is mainly induced and not

dependent on calcium level. Recently, it had been demonstrated that nNOS expression

can also be induced upon exposure to hypoxia (34). Despite the requirement of oxygen as

substrate, the sensitivity of each NOS isoform to oxygen is very different (Km for O2:

nNOS = 350μM, eNOS=4 μM, iNOS=130 μM) (126). The higher Km for O2 in nNOS

suggested that its enzymatic activity is more sensitive to a small drop of oxygen tension,

whereas the lower Km for oxygen in eNOS implied that eNOS enzymatic activity is

affected only with a bigger decrease in oxygen level. Furthermore, protein localization is

also important for nNOS and eNOS activity. NOS interacting protein (NOSIP) interacts

with nNOS (127), while both NOSIP and eNOS traffick inducer (NOSTRIN) can bind to

eNOS protein (128, 129), which negatively regulate NOS activity.

31

Chapter 1

1.4.2 Targets of NO Heme

The binding of NO to ferrous (Fe2+) heme proteins is the most potent action of

NO (Figure 1.6). A classic target for NO to heme is the activation of soluble guanlyl

cyclase (sGC) in vascular smooth muscle. NO binds to the heme group and activates

sGC, which stimulates the production of cyclic guanosine 3’5’-monophosphate (cGMP)

(130). Increased cGMP levels lead to smooth muscle relaxation, which is important in

blood pressure regulation. As a signaling molecule, cGMP-mediated vasodilation occurs

in multiple ways: 1) it inhibits calcium entry to the cell and decreases intracellular

calcium concentration, leading to reduced vascular smooth muscle contraction; 2) it

activates potassium channels, resulting in hyperpolarization and relaxation; and 3) it

promotes phosphorylation and activation of protein kinases that activates myosin light

chain phosphatase, which in turn leads to relaxation of vascular smooth muscle. The

importance of a vasodilative role for NO/cGMP pathway is highlighted by the fact that

acetylcholine, which is a potent vasodilator, was found to induce smooth muscle

relaxation via NO/cGMP dependent mechanisms.

In addition, one common physiologic target for NO is Hb, which is widely known

as a NO scavenger / carrier in the body. Under physiological conditions, NO can bind

with the heme pockets of deoxyhemoglobin to form iron-nitrosyl-hemoglobin. In

addition, ferrous heme of hemoglobin can be oxidized in a reaction involving NO to form

nitrate and methemoglobin as a byproduct, which cannot carry oxygen because the heme

group is oxidized to ferric heme (Fe3+).

32

Chapter 1

S-nitrosylation of cysteine thiols Other than binding to heme proteins, NO can form interaction with cysteine thiol

(S-nitrosylation) to transducer NO bioactivity (Figure 1.5). S-nitrosylation of various

proteins possesses important physiological implications. S-nitrosylation of specific

proteins may change gene expression and their functions. For example, S-nitrosylation of

G-protein-coupled receptor kinase 2 (GRK2) inhibits GRK2 phosphorylation activity,

thereby preventing the loss of beta adrenergic receptor signaling in vivo (131).

Interestingly, in neurons, the activity of NMDA receptor is associated with localized NO

production, which can S-nitrosylate the receptor, downregulate receptor activity and

affect cellular signal transduction (132-134). In cardiomyocytes, nNOS-mediated S-

nitrosylation of ryanodine receptor is important for preserving normal calcium cycling in

the sacroplasmic reticulum and retaining normal excitation-contraction in cardiac

muscles (135, 136). Furthermore, S-nitrosylation of proteins such as hypoxia-inducible

factors (HIF) (137, 138), H-ras (139), glyceraldehydes-3-phosphate dehydrogenase

(GAPDH) (140, 141) and β-arrestin (142) can alter their expression levels. Collectively,

S-nitrosylation is an important player in regulating biological processes in various

physiological situations.

Besides NO binding to the heme group of Hb, NO can also interact with the thiol

group cysteine within the β-globin chain of Hb (Cysβ93) to form S-nitrosohemoglobin

(SNO-Hb) (Figure 1.6). As the red blood cell preloaded with SNO-Hb travels to tissues

where deoxygenation occurs, Hb changes structural confirmation from relaxed

(oxygenated) state to tense (deoxygenated) state such that SNO is exposed to solvents

such as glutathione to form S-nitrosoglutathione (GSNO) (8, 143). This signaling

pathway provides a means of rapid response in the vasculature, thereby regulating local

33

Chapter 1

blood flow and oxygen delivery to maintain oxygen homeostasis. In addition to blood

flow regulation, SNO-Hb may also control ventilatory responses to hypoxia by the

enzyme γ-glutamyl transpeptidase that cleaves S-nitrosoglutathione (GSNO) to

CysGlyNO (106). Collectively, SNO-Hb plays key roles in the respiratory cycle; from

oxygen uptake in the lungs, to oxygen delivery to tissues, and to the drive to breathe.

The level of protein S-nitrosylation depends on the both the rates of nitrosylation

and denitrosylation. One denitrosylase system that is physiologically relevant is GSNO

reductase (GSNOR) system, which comprises glutathione (GSH) and GSNOR (Figure

1.7). GSNO is a main form of low-molecular-weight S-nitrosothiol (SNO), involved in

transnitrosylation to nitrosylate other proteins, and can be formed by reaction between

NO and GSH. SNO protein can be denitrosylated by GSH, thus forming a reduced thiol

and GSNO. In turn, GSNO is reduced by GSNOR to GSNHOH. Thus, the GSNOR/GSH

system reduces the level of SNO proteins by driving the equilibrium of SNO proteins

towards GSNO (144). In vivo studies demonstrated that mice deficient in GSNOR had

increased SNO levels in red blood cell and S-nitrosylated protein levels (138, 145).

Therefore, GSNOR/GSH can denitrosylate protein indirectly through reduction of GSNO

levels. Another important denitrosylase is the thioredoxin (Trx) system, which consists of

both Trx and Trx reductase (TrxR), and is a major protein disulphide reductase system in

all living organisms (146). Reports suggest that Trx can mediate denitrosylation by

promoting GSNO breakdown (147), interacting directly with SNO (148), forming a

disulphide linkage intermediate between substrate and Trx (149), and transiently transfer

nitrosylation to Trx (transnitrosylation) (148). Its physiological importance is

34

Chapter 1

demonstrated by denitrosylating SNO-caspase-3, thereby inhibiting caspase-3 activity,

and suggesting a role of NO in anti-apoptosis (149, 150).

Figure 1.7: The GSNO reductase (GSNOR)/ GSH system. Adapted from Benhar M et al Nat Rev 2009.

35

Chapter 1

NO storage as nitrite and nitrate

Potential sources of nitrate (NO3-) and nitrite (NO2

-) include the diet. Nitrite may

also be formed when NO is oxidized by multi-copper oxidase and ceruloplasmin (151). It

has been proposed that NO can be produced from nitrite and nitrate (Figure 1.6). This

may occur by a mechanism in which nitrite is converted to bioactive NO in hypoxic

vascular tissue by deoxyhemoglobin (HbFe2+) with methemoglobin (HbFe3+) Hb as a

byproduct (38, 152, 153). Indeed, methemoglobin may also serve as an intermediate to

produce SNO-Hb (154, 155). NO produced from nitrite in this way may contribute to

hypoxic signaling and vasodilation (38, 156). However, this biochemical process of

nitrite may not be physiologically relevant in vivo (125). This mechanism of hypoxic

vasodilation differs from that proposed by Stamler and colleagues. Although the

mechanism by which the RBC generates NO in hypoxic vascular beds is debated (157-

162), the concept of Hb as a NO shuttle was first proposed by Stamler. Other

investigators have provided further evidence for physiological regulation of hypoxic

vasodilation (9, 163).

36

Chapter 1

1.5: Neuronal nitric oxide synthase (nNOS)

The first mammalian NOS was isolated from the brain and termed neuronal NOS

due to its localization in neurons (164, 165). Unexpectedly, nNOS is also constitutively

expressed in a variety of cell types including neurons, skeletal muscle, smooth muscle,

and cardiac muscle. The activity of nNOS is calcium dependent, and its expression is

traditionally regarded to be constitutively expressed. nNOS expression can also be

induced by external stimulus, such as hypoxia.

1.5.1: Biochemistry of nNOS

The predicted molecular weight for nNOS is 160 kDa. nNOS is active as a dimer

and dimerization requires tetrahydrobiopterin (BH4), heme, and L-arginine binding.

nNOS monomer contains an oxygenase domain (N-terminus) and reductase domain (C-

terminus), which is flanked by the calmodulin binding domain. The oxygenase domain

contains binding sites for substrate L-arginine, BH4, cytochrome P-450-type heme, and

zinc which facilitates nNOS dimerization. The reductase domain contains binding sites

for NADPH, FMN and FAD. Electrons donated by NADPH can transfer via FAD and

FMN to heme, which can be facilitated by calcium/calmodulin binding. In general, active

nNOS homodimer catalyzes the oxidation of L-arginine to produce L-citrilline and NO.

Enzyme activity of nNOS is regulated by intracellular Ca2+ levels. In neurons,

activation of NMDA receptor leads to Ca2+ influx (166). This increase in free Ca2+

promotes CaM binding to nNOS and stimulates enzymatic activity. This quick response

requires close localization to the NMDA receptor by PDZ/GLGF domain, which is

encoded by exon 2 and located in the N-terminus of nNOS protein. The PDZ/GLGF

37

Chapter 1

domain binds to post-synaptic density PSD-95 and PSD93, which is bind to NMDA

receptor. Thus, membrane bound nNOS protein is in close proximity to NMDA receptor.

1.5.2: Regulation of the nNOS gene mRNA diversity

The genomic arrangement is highly conserved across evolution, dating back to

mollusks (167). nNOS is a complex gene consists of 29 exons in humans and 23 exons in

mouse. Transcription initiates at exon 2 and terminates at the last exon, with numerous

exon 1 splice variants expressed in a tissue specific manner (165, 168). To produce a

functional protein, nNOS gene encoded in the DNA is first transcribed to RNA, which is

then translated into protein. The regulation of nNOS gene expression is very complex and

can be controlled at different levels. At the mRNA level, diversity can exist within the 5’-

untranslated region (5’-UTR) of nNOS mRNA. For instance, nNOS contains ten different

examples of exon 1 variants (each regulated by a distinct promoter and expressed in

tissue-specific manner) that are spliced to a common downstream exon 2. Because

translation initiation codon AUG is located in exon 2, this diversity of exon 1 variants

does not affect the structure of the encoded full-length nNOS protein, despite varying

transcript length (165). Furthermore, other than alternate promoter usage, sequence

diversity within 5’-UTR can also arise from an alternatively spliced 89-nucleotide exon,

which is between the varied exon 1 examples and the common exon 2. This alternatively

spliced exon has been shown to repress translation efficiency of nNOS mRNA (169).

Therefore, the degree of sequence diversity at the 5’-UTR of nNOS provide examples of

the complexity of the nNOS human gene.

38

Chapter 1

Since nNOS transcripts produced from each of the unique exon 1 examples are

long and inefficiently translated, a faster response would be required for cells to adapt

quickly to rapid changes in the local environment. The ability to produce shortened

mRNA transcripts could lead to mRNAs that are more efficiently translated into proteins.

For example, hypoxia stimulates increases in nNOS mRNA and protein levels in various

tissues, such as aorta, mesenteric arterioles, brain and kidneys (34). Importantly, the

increased transcription of the gene in hypoxic cells and tissues does not reflect

transcription from the same promoter regions implicated in constitutive nNOS

expression. Rather, a novel, hypoxia-inducible promoter is activated, just upstream of

exon 2 (Figure 1.8). This promoter is located more than 100 kilobases downstream of the

genomic regions where the constitutively expressed exon 1 variants are transcribed from.

When transcription is initiated, a shortened nNOS mRNA transcript containing an

alternative 5’-UTR is produced. Under hypoxic conditions, nNOS mRNA transcript

containing this alternative 5’-UTR are very efficiently translated into protein, bypassing

the long exon 1 transcripts, which are very poorly translated (168). The in vivo relevance

of this process was demonstrated in a transgenic mouse line that contains a nNOS exon 2

promoter fused with a LacZ reporter. Hypoxia results in increased β-galactosidase

staining (indicative of promoter/reporter activity) in the brain and kidneys. Importantly,

there is no β-galactosidase activity in normal tissues (34), which confirm the inducibility

of this promoter at exon 2 by hypoxia. These studies highlighted that apart from the

constitutive nNOS expression via unique exon 1 variants spliced to a common exon 2,

full-length nNOS protein can be produced via promoter upstream of exon 2 so that nNOS

39

Chapter 1

transcripts are generated with short 5’-UTRs that are very efficiently translated. Thus,

this results in rapid induction of nNOS protein expression in response to external stimuli.

Figure 1.8: Schematic diagram of normoxic and hypoxic induction of nNOS transcript. Hypoxia induces a shortened 5’-UTR nNOS transcript that is efficiently translated into protein. In normal conditions (upper panel), nNOS transcription is initiated (arrowhead) at an upstream regions and encodes a long first exon that is spliced to a common exon 2, where translation initiates. This produces a nNOS mRNA transcript with a long, structured 5’-UTR that is poorly translated into protein. Under hypoxic conditions (lower panel), nNOS transcription initiates (arrowhead) at downstream regions just upstream of regions encoding exon 2. This generates a mRNA transcript containing a shortened 5’-UTR that is very efficiently translated into protein. Open box denotes 5’-UTR. Shaded box denotes open reading frame (ORF).

Published in a book review: Marsden, Philip A, Newton, Derek C, and Tsui, Albert K Y(Apr 2011) Alternative Processing: Neuronal Nitric Oxide Synthase. In: eLS. John Wiley & Sons Ltd, Chichester. http://www.els.net [doi: 10.1002/9780470015902.a0005040.pub2]

40

Chapter 1

Not only does sequence diversity occurs within 5’-UTR of nNOS transcript, many

alternative splicing events also exist within the open reading frame (ORF). These

processes would alter the amino acid sequence and the protein product structure. For

example, cassette insertion of adding 102 bp between exon 16 and 17 results in the

addition of 34 amino acids near the calmodulin binding region (170). This altered protein

is termed nNOSμ, which is important in cardiac and skeletal muscle development.

Additionally, deletion can occur at exon 2, which contains the translation initiation codon

AUG (171, 172). The loss of translation initiation codon prompts translation initiation at

other sites, such as at the CUG codon in exon 1 in mouse. This could give rise to a

modified protein named nNOSβ. Furthermore, translation may be initiated at a

downstream AUG codon at exon 5, which results in a truncated protein designated as

nNOSγ. Interestingly, nNOSγ is homologous to the testis-specific nNOS variant

(TnNOS), which is expressed in Leydig cells and important in male reproductive system

(173, 174). Collectively, the complexity and diversity of nNOS gene regulation at the

mRNA level set the stage for the tight control of nNOS expression in various biological

settings.

Post-translational modification

Post-translational modification of nNOS protein is mostly affected via the

phosphorylation status of nNOS by specific protein kinases and phosphatases, such as

PKA, calmodulin-dependent kinases (CaMK), PKC, and phosphatase 1. Studies in rat

cortical neurons, for example, demonstrated that phosphorylation of nNOS by CaMKII at

serine residual 847 (Ser847) reduces nNOS activity by inhibiting calcium-calmodulin

binding. By contrast, protein phosphatase-1 reduces phosphorylation at the same serine

41

Chapter 1

site can lead to increased nNOS activity (175). Furthermore, phosphorylation of nNOS

by Akt at serine residual 1412 (Ser1412) activates nNOS by NMDA receptor, whereas

dephosphorylation results in reduced nNOS activity (176). Of clinical significance, the

phosphorylation status of nNOS may be important in the pathogenesis of brain injuries

and diseases, such as cerebral ischemia (177) and hypoxia (178).

As a cytosolic protein, nNOS can also bind to proteins at the membrane to exert

biological functions. Protein-protein interactions are important for regulating localization

of nNOS and its enzymatic activity. The N-terminal PDZ domain of nNOS binds to the

postynaptic density protein PSD-95 and is connected with the NMDA glutamate receptor

that permits calcium level and accounts for the activation of nNOS by NMDA receptor

stimulation (179). Interestingly, the attachment of PSD-95 to the membrane is regulated

by dynamic palmitoylation and nitrosylation (180). Palmitoylated PSD-95 at cysteine 3

and 5 is associated with the membrane. NMDA stimulation promotes depalmitoylation,

and simultaneously, Ca2+ entry via NMDA receptor to activate nNOS and produce NO.

Subsequently, NO generated can nitrosylate PSD-95, preventing re-palmitoylation and

association to the membrane. Thus, nNOS-mediated nitrosylation can regulate its own

activity. Furthermore, the muscle isoform of phosphofructokinase (PFK-M) also bind to

the PDZ domain of nNOS in neurons and skeletal muscle (181). PFK-M is abundant in

neurons expressing nNOS. The product of PFK, fructose-1,6-bisphosphoate, may

neuroprotective effects. Another adaptor protein called CAPON (C-terminal PDZ ligand

of nNOS) can compete with PSD-95 for interaction with nNOS (182) and is needed for

the formation of ternary complex by interacting with Dexras1, which is a brain-enriched

member of the Ras family of small G proteins (183). Additionally, skeletal nNOS can

42

Chapter 1

interact with syntrophin and localized to the sacrolemmal membrane of the

neuromuscular junction (184). Also, NOSIP can interact with nNOS in the synaptic

spines, which inhibits nNOS activity (127). Furthermore, calcium-calmodulin binding of

nNOS triggers nNOS enzymatic activation, which, for example, can be achieved by

NMDA receptor activation in the neurons (185).

Importantly, heme insertion into nNOS, as well as other NOSs, is a key step in

NOS assembly and enzyme activity. Heme binding to nNOS monomer is required for the

formation of a catalytically active nNOS homodimer (186). This process of heme

insertion into heme proteins (such as NOS and Hb) can be inhibited by NO (187).

Specifically, Stuehr and colleagues recently demonstrated that GAPDH can deliver heme

to heme target proteins, such as iNOS (188). S-nitrosylation of GAPDH can inhibit heme

insertion and thus NOS activity. Importantly, GAPDH can bind to all NOS isoforms

(188), and thus may also enhance heme delivery to nNOS and eNOS. Collectively, these

studies demonstrated that NO can self-regulate NOS activity by controlling the insertion

of heme to the NOS proteins. In summary, regulation of nNOS expression and activity

can occur at multiple levels via mechanisms that involve transcriptional, translational,

post-translational, and/or cellular localization.

1.5.3: Role of nNOS-derived NO in different tissues

Since nNOS is expressed in many different tissues such as the brain, kidneys,

heart, smooth muscle, and skeletal muscle, nNOS-derived NO contributes to a diverse of

biological functions in a tissue specific fashion.

43

Chapter 1

Brain In the brain, nNOS is abundantly expressed in neurons, where it is important in

mediating synaptic plasticity and neuronal signaling (189). By inhibiting the transporters

of monoamines, nNOS-derived NO can increase the extracellular concentrations of

monoamines, including dopamine, noradrenaline and serotonin, and prolong their lifetime

and effects around the synapse (190). nNOS-derived NO can also contribute to the neural

regulation of the cardiovascular system. nNOS is found in the neurons and fibers of

nucleus tractus solitarii (NTS), where the cardiovascular centers are located in the brain.

It has demonstrated that injection of nNOS antisense oligonucleotides into NTS to inhibit

nNOS expression resulted in reduced systemic blood pressure and heart rate (191). In

addition, nNOS neurons found in the paraventricular nucleus (PVN) of the hypothalamus

can suppress sympathetic outflow of the kidneys (192). These studies suggested that not

only does nNOS is important in maintaining the integrity of brain function, but it also is

essential in controlling the central nervous system to repress the cardiovascular and renal

system.

Heart

In “healthy” heart, nNOS is found to be localized to the sacroplasmic reticulum

(SR) of cardiomyocyte (193). The role of nNOS in regulating Ca2+ level is complex and

controversial, and may involve a number of mechanisms, including Ca2+ ATPase,

ryanodine receptor, L-type Ca2+ channel, and phospholamin B (PLB). Studies using

cardiomyocytes from nNOS-/- mice revealed that nNOS-derived NO inhibit the activity

of the SR calcium pump (SERCA2a), thereby depleting Ca2+ storage in SR for

subsequent contractions (194). Regulation of SERCA2a by nNOS could be due to

modulation of phospholamban B phosphorylation status (195). In addition, nNOS is

44

Chapter 1

found in close proximity to ryanodine receptor (RyR2) in the SR (193, 194). Activation

of ryanodine receptor by nNOS leads to calcium release from SR via channel

phosphorylation, which results in increased myocardial contraction (196). Also, nNOS

can inhibit L-type Ca2+ channel, which leads to reduced Ca2+ influx to the cell (197).

Subsequently, decreases in Ca2+ entry leads to reduced Ca2+ -induced Ca2+ release by

RyR2, which results in decreased contraction and improved relaxation. Furthermore, it

has been suggested that nNOS may increase the activity of Na+/K+ ATPase pump, and

thus influence intracellular Ca2+ flux by altering the activity of Na+/Ca2+ exchanger (198).

Overall, from studies in pharmacological inhibition and genetic ablation of nNOS at basal

conditions, nNOS-derived NO has negative or neutral inotropic and a positive lusitropic

effect in left ventricular myocardiocyte (194, 197, 199, 200). In contrast, nNOS is shown

to be important in positive inotropic response under β-adrenergic stimulation, which is

impaired in nNOS-/- mice (200, 201). However, the exact mechanism on cardiomyocyte

response of nNOS in β-adrenergic stimulation has not been clearly shown.

In “failing” heart, nNOS is translocated from SR to sacrolemmenal, which is

evident by the increased interaction with caveolin-3 (202) and decreased physical

association with RyR2 (203) in rat model (202) and human failing myocardium (204).

This nNOS translocation in failing heart could affect myocardial contractility. Increased

nNOS-derived NO in caveolin may further enhance the inhibition of L-type Ca2+ channel,

in part via S-nitrosylation (205), while decreased nNOS-derived NO from SR may

disrupt RyR2 or SERCA2a activity and affect Ca2+ storage. Furthermore, nNOS can

inhibit xanothine oxidoreductase (XOR) activity, which may promote survival following

myocardial infarction (206). Taken together, the role cardiac nNOS in Ca2+ cycling is

45

Chapter 1

complex in that it affects multiple targets and can be differentially regulated in basal,

stimulated and diseased states.

Kidneys

In the kidneys, nNOS protein expression is highly present in the macula densa

cells, and it blunts tubuloglomerular feedback (TGF) responses as shown in rats, rabbits

and mice studies using pharmacological inhibitors of nNOS (207-210) by mechanisms

not clearly understood. Potential mechanisms of nNOS-derived NO blunt TGF include:

1) NO inhibition of NaCl entry via decreasing the activity of Na-K-2Cl cotransporter in

the macula densa (211); 2) inhibition of 5’-ectonucleotidase that leads to reduction in

adenosine levels (a mediator of TGF response) (212) and; 3) macula densa nNOS-derived

NO diffuses to smooth muscle cells, activates cGMP and dilates afferent arteriole (213).

Also, nNOS is found in the principle cells of the collecting ducts, which may inhibit

sodium and water reabsorption (213, 214). Furthermore, nNOS expressed in the proximal

tubule enhances fluid and HCO3- absorption, as well as regulates the acid-base balance

(215). Finally, nNOS is found in non-adrenergic, non-cholinergic neurons of renal

arteries (216), which is important for regulation of autonomic response in the kidneys.

Overall, nNOS-derived NO contributes to a diverse, yet specialized, function of the

kidney that is important to normal renal physiology.

Smooth muscle

Identification of nNOS expression in smooth muscle suggested that nNOS-

derived NO can also participate in vascular homeostasis (217, 218). Smooth muscle

nNOS is localized to caveolin-1, where it is closely associated with plasma membrane

Ca2+/calmodulin-dependent Ca2+ ATPase (PMCA) via PDZ domain (219). Activation of

46

Chapter 1

PMCA4b leads to extrusion, and thus reduction, of intracellular Ca2+, resulting in the

attenuation of Ca2+ responsive nNOS-derived NO and reduced cGMP levels (220, 221).

This results in reduced phosphorlyation of PKG and MLCK, which leads to inhibition of

smooth muscle relaxation. Therefore, regulation of vasomotor response by nNOS

happens by controlling Ca2+ concentration in cGMP-dependent manner. Furthermore,

there are evidence that nNOS may play a role in the myogenic response of small

resistance arteries, which is characterized by a constriction of the vessel after an increase

of transmural pressure and a dilation of the vessel after a decrease of transmural pressure.

For example, an increase in transmural pressure would stimulate stretch-activated Ca2+

channel, leading to increase intracellular Ca2+ concentration (221). This results in the

production of 20-hydroxyeicosatertaenoic acid (20-HETE) by smooth muscle cells,

which enhances vasoconstriction by inhibiting Ca2+ dependent K+ (Kca) channels. The

synthesis of 20-HETE further leads to vasoconstriction by the activation of L-type Ca2+

channel, activation of Rho kinase, and phosphorylation of MLC (222, 223). Since nNOS-

derived NO inhibits the formation of 20-HETE leading to vasodilation (220, 221, 224),

nNOS can modulate smooth muscle tone via a cGMP-independent mechanism that is

consistent with the vasodilative role of NO.

Skeletal muscle

In skeletal muscle, nNOS is bound to the dystrophin-associated protein complex

by the interaction between nNOS PDZ domain and α-syntrophin, where nNOS is

localized to the plasma membrane (225). nNOS in skeletal muscle has many functions,

including regulation of muscle blood flow, glucose homeostasis, muscle mass and

fatigue. During exercise when oxygen demand is increased, muscle contraction stimulates

47

Chapter 1

sacrolemmal nNOS-derived NO production, which diffuses out of the muscle fibres to

dilate blood vessels (226, 227). Skeletal muscle nNOS may also stimulate glucose

uptake by increasing glucose transporter 4 (GLUT4) translocation to plasma membrane

during exercise (228). Also, exercised-induced muscle fatigue occurs in muscular

dystrophy patients when sacrolemmal nNOS is displaced (i.e. not at the plasma

membrane) (227) or genetically deleted (229). An increase in skeletal muscle NO-cGMP

signaling had shown to improve dystrophic muscle phenotype by reducing muscle

degeneration, inflammation and increase exercise tolerance (230-233). Taken together,

nNOS is important in maintaining the integrity of skeletal muscle function during

exercise.

1.5.4: Is nNOS a friend or foe?

Experimental evidences suggested that nNOS is a two-edged sword: It exerts

protective role in physiological conditions when its expression is tightly controlled;

however, nNOS is detrimental when it is overexpressed in pathophysiological settings.

The use of nNOS-/- mice allows investigators to determine the role of nNOS in each

experimental model.

nNOS as a friend

nNOS expression, whether it is constitutive or induced, is important for normal

physiology in mammals. Using transgenic animal models, it has been reported that nNOS

contributes to protective effects in many disease conditions. In myocardial infarction

(MI) model, mice deficient of nNOS have worsened outcome (LV remodeling,

arrhythmia) and increased mortality following MI, suggesting protective roles for nNOS

48

Chapter 1

in heart conditions (136, 201, 206). Furthermore, pharmacological and genetic ablation of

nNOS demonstrated impaired spatial performance in the Morris water maze, suggesting

important role of nNOS in learning and memory (234, 235). Additionally, nNOS-/- mice

are known to possess pyloric stenosis as characterized by enlarged stomach,

demonstrating a physiological role of nNOS in the digestive tract (236). Furthermore,

loss of skeletal muscle nNOS result in muscle fatigue and resemble muscular dystrophic

pathology, indicating nNOS is important in normal muscle physiology (229-231).

In mouse model of atherosclerosis using a carotid artery ligation methodology, nNOS-/-

mice demonstrated increased neointimal formation and vascular remodeling (237). In

another atherosclerosis model (apo E deficient), mice lacking both apoE and nNOS result

in increase in plaque formation relative to mice lacking apoE alone, demonstrating nNOS

may protect against atherosclerosis pathology (238). Overall, these mutant mouse studies

showed evidence that nNOS is protective in certain conditions.

nNOS as a foe

Given that tight regulation of nNOS is beneficial, overexpression of nNOS have

been shown to produce deleterious effects. In focal ischemia, mouse model demonstrated

that nNOS is overly activated and associated with neural damage of ischemic stroke, as

evident that nNOS-/- mice have small infarct size following stroke (239, 240).

Anoxia/hypoxia plays a major role to the mechanism of nNOS overexpression in stroke.

In focal ischemia, lack of blood flow results in oxygen and glucose deprivation, causing

an excessive synaptic release of glutamate. Glutamate excitotoxicity results in the

activation of NMDA receptors, leading to Ca2+ influx to the cell and consequently

activates Ca2+ sensitive enzymes, one of which is nNOS. NO radical that is produced

49

Chapter 1

from overexpression of nNOS will react with superoxide to form peroxynitrite, leading to

neurotoxicity and increased infarct size. Furthermore, a transgenic mouse model of

conditional myocardial nNOS overexpression was shown to reduce myocardial

contractility and produce failing heart (241), suggesting the role of nNOS in

cardiomyocyte relaxation. Overall, unregulated nNOS expression results in detrimental

effects.

1.5.5: nNOS knockout mice

The first nNOS knockout mouse was generated by homologous recombination

and targeted deletion of exon 2 in nNOS gene (236). It was noted that this nNOS

knockout mouse was viable and fertile, and the male possess aggressive behavior. The

most apparent phenotype of these mice was enlarged stomach and abnormalities in

pyloric sphincter, which closely assemble pyloric stenosis in humans (236). Deletion of

nNOS exon 2 results in approximately 5% residual nNOS activity in the brain, which was

initially attributed to the contribution of other NOS isoforms. However, later work

identified that truncated nNOS proteins (nNOSβ, nNOSγ) produced by alternative splice

variants and alternative promoter usage account for the residual nNOS activity (168, 172,

236). The role of nNOS in many disease models have utilized this nNOS knockout mouse

line. In focal ischemia, for instance, it was found that nNOS-deficient mice had smaller

infarct size compared to wildtype mice, implying that nNOS is harmful in ischemia

(239). In addition, utilizing this nNOS knockout model revealed important role for nNOS

in heart (136, 200, 201, 206), skeletal muscle (229, 231, 232), and blood vessel biology

(237, 238).

50

Chapter 1

Following the generation of nNOS exon 2 knockout mice, a new line of nNOS

knockout mice lacking exon 6 had been created (242). Exon 6 encodes for heme-binding

domain of nNOS, which is responsible for the catalysis of arginine to NO and citrilline

(165, 243). In this new line, nNOS activity was completely abolished, however, the

consequence of exon 6 deletion results in hypoganadism and infertility (242). Using this

knockout mouse line, the role of nNOS splice variants in NO signaling have recently

been explored in skeletal muscle (231). However, due to infertility of these mice and

practical feasibility of the project, studies using nNOS knockout mice lacking exon 6

cannot be easily explored. Thus, nNOS knockout mice lacking exon 2 will be utilized in

this project.

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Chapter 1

1.6: Hypoxia-inducible factor

1.6.1. Discovery of Hypoxia-Inducible Factor-1:

Hypoxia-inducible factor-1 (HIF-1) is a heterodimeric transcription factor that

consists of α and β subunits. HIF-1 is first discovered by revealing a hypoxia-inducible

enhancer in the 3’ end of human erythropoietin (EPO) gene (244). This resulted in the

identification of a DNA sequence termed hypoxia response element (HRE) (5’ –

(A/G)CGTC - 3’) that is common to many genes responsive to hypoxia. Subsequently,

HIF-1α was isolated, cloned and found to bind to HRE that mediates hypoxia-induced

transcription. Amazingly, HIF pathway is evolutionary conserved and appeared before

the development of metazoan in ~600 million years ago, indicating the importance of this

pathway on the survival of organisms to adapt to changes in oxygen level that exist for

the past 600 million years.

1.6.2. Isoforms of Hypoxia-Inducible Factor -α

There are three isoforms of α subunits: HIF-1α, and 2α and HIF-3α. HIF-1α and

HIF-2α are more closely related in terms of homology and are responsive to changes in

oxygen tension. HIF-2α is also known as endothelial PAS domain protein 1 (EPAS-1).

HIF-3α is more distant, less well studied isoform and may play a role in inhibiting HIF-

1α by consisting a spliced variant termed inhibitory PAS domain protein (245-247).

Generally, HIF-α can be regulated by multi-step process involving changes in activity,

abundance, mRNA splicing and subcellular localization, but post-translational

modification (e.g. hydroxylation) of HIF-α plays a dominant role in regulating HIF-α

expression, especially in oxygen-related environment (114). The amino terminal of HIF-

1α and -2α consists of the basic-Helix-Loop-Helix (bHLH)-Per-ARNT-Sim (bHLH-

52

Chapter 1

PAS) protein domain that is required for dimerization and DNA binding in the nucleus

with HIF-1β, which is also known as aryl hydrocarbon receptor nuclear translocator

(ARNT). HIF-1α and HIF-2α also have an oxygen-dependent degradation domain

(ODD) that contains two proline residues for hydroxylation to regulate protein stability in

response to changes in oxygen level (248, 249). Furthermore, it has a carboxyl terminal

transactivation domain (TAD) that has the capability to bind the transcriptional

coactivator p300/CBP. Despite the structural similarities between HIF-1α and HIF-2α,

recent data demonstrated that the two isoforms appear to have their distinct as well as

common target genes. For example, HIF-1α mainly activates genes responsible for the

glycolytic pathways, whereas HIF-2α can specifically activate genes in cell survival and

proliferation (250). With HIF-1α is expressed ubiquitously in most tissues whereas HIF-

2α expression is cell specific (e.g. endothelium, astrocytes, liver), their responses may be

different depending on the stress conditions and local requirement for oxygen. In

addition, Hu et al., demonstrated that the target genes specificity by HIF-1α or HIF-2α

was not due to selective DNA binding at the HRE, but appeared to involve the

transactivation domain, specifically at N-terminus TAD (251). Other co-factors binding

to this region may confer selectivity between HIF-1α or HIF-2α. For instance, an ETS

transcription factor, ELK, is shown to interact with HIF-2α, but not HIF-1α, to activate

HIF-2α specific genes (251). Interestingly, HIF-2α, but not HIF-1α, is found to contain an

iron response element in its 5’-untranslated region (UTR). It is proposed that this is to

limit HIF-2α expression during iron deficiency, thus restricting unproductive excessive

erythropoiesis (i.e. deformation of RBC in iron deficiency) (252). Despite structural

similarities, distinct functions are found in the different HIF-α isoforms.

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Chapter 1

1.6.3: Tissue HIF-α protein expression in vivo

Since tissue oxygen tension is different in various tissues, it is not surprising that

the expression of HIF-α protein also varies as its expression is tightly linked with local

tissue oxygenation. Mouse studies have demonstrated that basal HIF-α level is low in the

well oxygenated organs (brain and heart) compared to kidneys and skeletal muscles at

rest (21% O2) (253, 254). The response to hypoxia also greatly varies. Brain HIF-1α

protein accumulation occurred as early as 1hr with modest hypoxia (18% O2), and further

increased as inspired oxygen was severely reduced. In contrast, hypoxic organs such as

kidneys and liver do not register increase in HIF-1α expression until hypoxia is severe

(6% O2) (253). This is biologically important because a quick adaptive response to a

small drop in oxygen ensure oxygen delivery to be maintained for the brain is vital for

survival. In contrast, hypoxia is only registered in the kidneys at very low oxygen level

suggested a role for kidney as a hypoxia sensor to release erythropoietin.

1.6.4. Post-translational Regulation of HIF-α Expression in Oxygen-Dependent Manner: The HIF-α subunits are primarily regulated by various post-transcriptional mechanisms: Hydroxlyation/ubiquitination

During normoxia, HIF-α protein is synthesized and subjected to hydroxylation on

proline residues by HIF prolyl hydroxylase domain (PHD), which requires oxygen and α-

ketoglutarate as substrates (Figure 1.9). Hydroxylated HIF-α recruits von Hippel Lindau

protein (pVHL)-E3 ubiquitin ligase complex to bind, leading to HIF-α ubiquitination, and

subsequent proteosomal degradation. Concurrently, factors inhibiting HIF (FIH) is

activated to hydroxylate the asparagine residue at the carboxyl transactivation domain (C-

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TAD), which blocks the interaction with coactivator p300/CBP. Thus, HIF-α stability and

transcriptional activity are negatively regulated by hydroxylation.

During hypoxia, HIF-α is stabilized and mainly regulated at the post-translational

level (Figure 1.9). As a result of substrate (oxygen) deprivation, PHD activity and

hydroxylation reaction is inhibited, thus increasing HIF-α stability. Hypoxia can also lead

to increased in reactive oxygen species (ROS) levels in the mitochondria, which can

inhibit PHD activity by reducing its Fe2+ catalytic site (255). Accumulation of HIF-α

translocate into the nucleus, where it binds to and dimerizes with nuclear HIF-1β to

increase transcription of HRE genes such as VEGF and EPO. In addition to the

hydroxylation at proline residues by PHD, hydroxylation status at asparagine residue by

factors inhibiting HIF (FIH) at the C-TAD region can affect transactivation of HIF-1α. In

hypoxia, FIH activity is reduced, thus co-activators (p300, CREB) are allowed to bind to

HIF-α. This helps to promote transcriptional activation of target genes that contain the

HRE sequence (96).

SUMOlyation

Besides hydroxylation and ubiquitination, there are other posttranslational

modifications of HIF-α that can affect its protein level in hypoxia, one of which is the

Small Ubiqutin-like Modifier (SUMO). It has been recently demonstrated that

SUMOlyation is involved to regulate HIF-1α in hypoxia (256). In hypoxia, hydroxylated

HIF-α can be degraded by pVHL-dependent pathway in the cytoplasm. Hypoxia can also

induce nuclear translocation of non-hydroxylated HIF-α and SUMOlyation of HIF-α.

SUMOylated HIF-α can be recognized by pVHL-dependent degradation in a

hydroxylation-independent manner in the nucleus. The presence of Sentrin/SUMO-

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specific proteases (SENPs) are required to de-SUMOlyate HIF-1α, which removes the

VHL-dependent degradation signal, and allows HIF-1α to be stabilized in hypoxia.

This modification on HIF-α represents an alternative way to degrade nuclear HIF-α.

Interestingly, mice lacking SENP1 develop anemia due to the inability to stabilize HIF-α,

suggesting that SUMO can de-stabilize HIF-α in vivo (256).

Acetylation

In addition, acetylation can serve as a secondary signal for HIF stabilization. In

normoxia, acetylation of lysine residues in the ODD of HIF-1α by acetyltransferase arrest

defective protein -1 (ARD1) enhances interaction of HIF-1α with pVHL, and HIF-1α

degradation (257). In contrast, hypoxia can deacetylate, for example, HIF-2α via sirtuin 1

(SIRT1), resulting in activation of HIF-2α target genes such as superoxide dismutase 2,

VEGFA and EPO (258). Opposite to HIF-2α, SIRT1 deacetylates and inactivates HIF-1α

by blocking p300 recruitment and consequently repressing HIF-1 activity and HIF target

genes (259). During hypoxia, SIRT1 level is decreased, thus allowing the acetylation and

activation of HIF-1α to occur. These results suggested a complex interaction between

SIRT1 and the two HIF-α isoforms.

Phosphorlyation

Furthermore, phosphorlyation of HIF-α can serve as an enhancer of

tranasactivation. Phosphorlyation of HIF-α by p42/44 Mitogen-Activated Protein Kinase

(MAPK) can promote nuclear accumulation and increase transcritptional activity by

increase preferential binding with HIF-1β (260). However, whether the phosphorlyation

status of HIF-α is influenced by hypoxia is still unclear.

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S-nitrosylation S-nitrosylation is a common post-translational modification of protein by NO. S-

nitrosylation of the HIF pathway will be discussed in detail in the next section.

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Figure 1.9: Regulation of HIF-α expression by hydroxylation in normoxia and hypoxia. In normoxia, prolyl hydroxylase (PHD) hydroxylate proline (P) residue of HIF-a protein in the cytosol. This results in the recruitment of pVHL complex, and HIF-α protein becomes ubiquitinated and subsequently degraded. In addition, factors inhibiting HIF (FIH) hydroxylates the asparagine residue (N) such that co-activators cannot bind. In hypoxia, prolyl and asparaginlyl hydroxylation does not occur. This stabilizes HIF-α in the cytosol, thus HIF-α is translocated into the nucleus and binds to HIF-1β. Binding of co-activators allow activation of transcription of genes containing hypoxia response element (HRE). Modified from Ratcliffe P.J. J of Clinical Investigation, 2007.

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1.6.5. Regulation of HIF-1α Expression by Nitric Oxide in Normoxia:

NO had been reported to affect HIF-α expression in different ways during

normoxic and hypoxic conditions in vitro. In normoxia, NO can stabilize HIF-1α by three

ways: 1) inhibiting PHD activity (261, 262), 2) S-nitrosylating pVHL (263), or 3) S-

nitrosylating HIF-1α (137, 138, 264) (Figure 1.10).

In vitro studies demonstrated that NO can inhibit PHD activity at normoxic

condition (261, 262). Biochemical analysis revealed that PHD requires iron (II) for its

activity in addition to the substrate oxygen and α-ketoglutarate. It is well known that NO

can bind to and modulate the activity of Fe(II)-containing proteins, such as soluble

guanlyl cyclase. Hence, NO has the ability to compete with molecular oxygen for binding

site to Fe(II) of PHD to reduce PHD enzymatic activity, thus HIF-α is stabilized.

S-nitrosylation, a process of adding NO group to cysteine residues, is another

mechanism for HIF stabilization by NO during normoxia. NO has been shown to S-

nitrosylate pVHL (263) and/or HIF-α (137, 138) under exogenous NO treatment to

stabilize HIF-α. When pVHL or HIF-α is S-nitrosylated, interaction between the two

proteins is prevented, thus HIF-α is allowed to escape pVHL-dependent degradation

pathway and accumulated. Additionally, NO can affect the transactivation status of HIF-

1α. It had been reported that NO donor can inhibit FIH activity (265) by competing with

oxygen for Fe(II) binding site, which blocks the binding of co-activators (p300/CBP) to

HIF-1α and inhibits HIF-α transactivation.

Most studies demonstrated the normoxic stabilization of HIF-α by NO occurred

with exogenous NO to cell culture conditions. Thus, the source of NO for this effect is

relatively novel in vivo. Li et al demonstrated that iNOS-derived NO can S-nitrosylate

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and stabilize HIF-1α in mouse cancer model (137). In a hind limb ischemia model,

Schleicher et al showed that NO derived from eNOS inhibits PHD activity, thereby HIF-

1α is accumulated (266). Lima et al demonstrated that deletion of S-nitrosoglutathione

(GSNO) reductase in mice increase SNO levels and resulted in normoxic myocardial

HIF-1α S-nitrosylation and protein stabilization following myocardial infarction (138).

However, in vivo evidence of normoxic HIF-stabilization by NO in normal physiology is

still very limited and warrant for further investigation.

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Figure 1.10: Summary of the possible mechanisms by which NO can stabilize HIF-α in normoxia by disrupting the degradation pathway. These mechanisms include: (A) inhibition of PHD activity, (B) S-nitrosylation of pVHL, and (C) S-nitrosylation of HIF-α.

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1.6.6. Regulation of HIF-α Expression by Nitric Oxide in Hypoxia: In hypoxia, the mechanisms by which NO degrades HIF-α appear to be

completely different. HIF-1α is usually stabilized in hypoxic condition by escaping the

pVHL-dependent degradation pathway. However, in vitro studies have demonstrated that

the addition of NO donor resulted in HIF-α de-stabilization during hypoxia (267),

suggesting a differential role for NO on HIF-α in normoxic and hypoxic conditions. This

occurs by which NO inhibits cytochrome C oxidase, thus reducing mitochondrial

respiration (267). Since oxygen is being consumed to produce ATP in mitochondrial

respiration, inhibiting this reaction would increase oxygen availability in the cytosol to

activate PHDs. In turn, HIF-1α is hydroxylated and de-stabilized. The cell may fail to

recognize low oxygen in a hypoxic environment. Further reducing ATP production

during hypoxia may result in injury to tissue with high energy demand (e.g. heart) (268),

and this may exemplify a possible pathophysiologic role for NO in hypoxia.

1.6.7: O2-independent HIF-a regulation HIF-a can also be regulated by oxygen-independent pathway, and it appears to be

more relevant to cancer biology. RACK1 can interact with Elongin C, which recruit E3-

ubiquitin-protein ligase complex. RACK1 can thus substitute VHL to ubiqutinate HIF-1α

in an oxygen-independent manner (269). RACK1 is found to compete with heat shock

protein (HSP90) at the PAS domain of HIF-1α, thus reducing the stability of HIF-1α

(269). Besides regulating the degradation pathway, HIF-1α protein synthesis can also be

controlled, which is demonstrated in MCF7 human breast cancer cell (270). It was shown

that activation of PI3K phosphorylates and activates Akt, which can phosphorylates and

activates mTOR. This leads to phosphorylation of p70 S6 kinases and eIF-4E binding

protein 1, both are regulators of translation. Activation of p70 S6 kinases results in

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ribosomal protein S6 phosphorylation. Phosphorylated eIF-4E binding protein 1 blocks

its ability to interact with and inhibit eIF-4E. Overall, these actions of mTOR can

increase the rate of translation of HIF-1α.

1.6.8: Other non-hypoxic activators for HIF-1α Expression Besides hypoxia and NO, there are many stimuli that can stabilize HIF-α under

normal oxygen environment, including ANGII and S1P. These signaling molecules can

modulate HIF-α expression independent of oxygen level

Angiotensin II (Ang II) is a hormone converted from angiotensin I by angiotensin

converting enzyme. AngII is a vasoconstrictor in the afferent and efferent arterioles in the

kidneys, and is a potent regulator of blood pressure. AngII has recently been shown to

induce HIF-1α stability in vascular smooth muscle cells (271). In vitro studies have

demonstrated that Ang II mediates the production of hydrogen peroxide that results in

ascorbate depletion, thereby leading to PHD activity inhibition and subsequently HIF-1α

accumulation under normoxic condition. In addition, Ang II can also induce HIF-1α

expression in VSMCs through transcriptional and translational mechanisms (272).

However, these studies have only been demonstrated in cell culture system in one cell

type, and will be required to be confirmed in an integrative whole animal model.

Sphingosine-1-phosphate (S1P) is a bioactive lipid mediator important in the

vascular system, such as regulation of angiogenesis, vascular reactivity and permeability.

S1P is recently identified as a novel non-hypoxic activator for HIF-α in vascular cells via

transcriptional and VHL-independent mechanisms (273). However, these experiments

were performed in cell culture system, and it is required to be confirmed in vivo settings.

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Desforoximine (DFO) is a drug used clinically to treat iron intoxication (274). In

cell culture studies, it is commonly used to mimic the effects of hypoxia (275). This

compound can accumulate HIF-a independent of oxygen level by acting as an iron

chelator. As iron is an essential substrate for PHD enzymatic activity, DFO stabilizes

HIF-a by limiting iron and reducing PHD activity, thus stabilizing HIF-α in the absence

of hypoxia.

1.6.9. HIF-mediated adaptive responses

Stabilization of HIF-α subunits result in transactivation of hypoxia-responsive

genes to compensate for the low oxygen environment, thus leading to improved oxygen

delivery. With similar structure between HIF-1α and HIF-2α, they can target activation of

a set of common genes, but also appear to activate distinct sets of genes by themselves

(250). Classic examples of adaptive cellular pathways activated by the HIF include

angiogenesis (VEGF), erythropoiesis (EPO), iron metabolism (transferrin), glucose and

energy metabolism (GLUT1, PDK1), and cytokine response (SDF1, CXCR4).

Transgenic mouse models of HIF-α have expanded our understanding on the

physiological impact of HIF in vivo. Genetic ablation of HIF-1α results in embryonic

lethality due to failure of vascularization (276) and HIF-1α heterozygous (+/-) mice

developed impaired angiogenesis upon artery occlusion (277) and impaired physiological

responses to chronic hypoxia (278), suggesting HIF-1 control angiogenesis and hypoxic

responses at multiple level. As cells grow and proliferate during development, the

increase in oxygen demand causes HIF activity to increase, which activates transcription

of genes encoding angiogenic growth factors. Classic examples include vascular

endothelial growth factor (VEGF), placental growth factor (PLGF), stromal derived

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factor (SDF-1), which bind to their respective receptors on endothelial, smooth muscle,

progenitor, and mesenchmyal stem cells for angiogenic signaling activation. HIF-1 also

activates the receptors, such as CXCR4 (receptor for SDF-1) and VEGF receptors

(VEGFR1).

In addition, conditional HIF-2α, but not HIF-1α, knockout results in anemia,

suggesting HIF-2α has a major effect in adult erythropoiesis in vivo (279, 280). In

contrast to the initial reports that EPO is activated by HIF-1α in vitro (244), it is now

appreciated that HIF-2α is the major HIF-α isoform responsible for renal and hepatic

erythropoiesis in vivo (280). Since iron is required for proper erythropoiesis, HIF also

regulate genes important for iron intestinal absorption (hepcidin) (281) and iron transport

to bone marrow for hemoglobin synthesis (ceruloplasmin, transferrin) (282, 283). Thus,

the ability for HIF-α to activate erythropoiesis and iron production leads to increase

oxygen carrying capacity in hypoxia.

In normoxia, the catabolism of glucose to pyruvate is normally taken up through

the Kreb cycle and electrons are transferred to the respiratory chain to generate ATP. In

hypoxia, the lack of oxygen results in a shift from oxidative to non-oxidative form of

glucose catabolism and ATP generation, namely anaerobic metabolism. HIF-1α is a

central mediator for this metabolic switch in cellular adaptation to hypoxia by enhancing

glycolysis, while reducing oxygen consumption in oxidative phosphorylation when

oxygen is limited. For example, hypoxic activation of HIF leads to increase transcription

of glucose transporters (GLUT1), which facilitate glucose uptake and glycolysis (284).

During hypoxia, HIF-1α also mediate the increases in glucose flux by activating enzymes

involved in glycolysis, such as phosphofructose kinase (PFK-1) and phosphoglycerate

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kinase (PGK1), and facilitate pyruvate conversion to lactate by activating lactate

dehydrogenase A (LDHA) (285). Concurrently, pyruvate dehydrogenase kinase (PDK1)

is activated to shunt the conversion of pyruvate away from the entry to Kreb cycle and

mitochondria (286). Hence, the conversion of pyruvate to lactate is favored, and ATP is

produced anaerobically. Induction of PDK1 can also repress mitochondrial function and

oxygen consumption (287). The overall goal of HIF-1 mediated metabolic adaptation to

hypoxia is the increase of glycolysis and decrease in oxygen utilization by inhibiting

respiration.

Although it is currently unclear as to why some genes are HIF-1α or HIF-2α

specific, relative abundance, cell type localization and stimulus dependent of each HIF-α

isoforms may be the contributing factors (250). For example in cell metabolism, glucose

consumption and glycolysis are primarily regulated by HIF-1α, whereas fatty acid storage

is promoted by HIF-2α (288). In general, the ultimate goal of regulated HIF response is to

mediate gene expressions that are helpful in stressful environment.

1.6.10: Clinical implication of the PHD/VHL/HIF pathway

There are human diseases which are associated with defects in the PHD-VHL-

HIF pathway. Patients with Von Hippel-Lindeau (VHL) disease have a mutated VHL

tumor suppressor gene. It is a rare, autosomal dominant disease that develops highly

vascular tumors in specific tissues including angiomatosis, hemangioblastomas, and renal

cell carcinoma (289). Loss of VHL results in constitutive HIF-α expression that leads to

abnormal expression of HIF target genes, which promotes tumor formation. Unlike most

VHL mutations which normally result in tumor formation, mutation in 598 C>T does not

form tumor, but rather, it leads to the development of an autosomal recessive disease in

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Chuvash polycythemia. The 598C>T mutation results in the replacement of tryptophan

for arginine at codon 200 in humans of pVHL which participates in pVHL-HIF-α

interaction (290). By definition, patients with Chuvash polycythemia are characterized by

increased hemoglobin level, elevated serum EPO, VEGF and plasminogen activator

inhibitor-1 levels (290), and are associated with higher risk of hemorrhage, thrombosis,

and mortality (291, 292). Primary polycythemia can be due to factors intrinsic to red cell

precursors, whereas secondary polycythemia can be caused by increase production of

EPO. Mouse models have demonstrated that HIF-2α signaling is the major HIF-α isoform

responsible for the phenotypes in this disorder, such as splenic erythropoiesis (293) and

pulmonary hypertension (294). Similarly, mutations at the HIF2A or PHD2 gene also

result in familial erythropoiesis. These patients either have mutation of glycine by

tryptophan at amino acid 537 in HIF2A that results in a gain of function (295), or a

substitution mutation of proline by arginine at amino acid 317 of PHD2 protein (296).

These mutations lead to increased HIF-2α and subsequently inappropriate elevation of

HIF response genes, most notably EPO. Additionally, polymorphisms in HIF-1α are

linked with heart conditions. A single nucleotide (C to T) change in residue 582 of HIF-

1α at exon 12 from proline to serine is associated among patients with the absence of

coronary collaterals in coronary artery diseases (297). Other polymorphisms in HIF-1α

gene are related to more stable angina (298). Identification of mutations in the

PHD/VHL/HIF axis could provide therapeutic treatments to the associated diseases.

The normal response to high altitude (chronic) hypoxia for low-land population is

an excessive increase of RBC mass and Hb level. Not only does elevation of Hb cannot

correct the problem of ambient hypoxia, it also worsened the crisis due to hyperviscosity,

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which can lead to increased vascular resistance and compromised tissue oxygenation

(299). Remarkably, the evolutionary adaptation of Tibetan population to habitat in high

altitude (4000m above sea level; PO2 ~110 mmHg) for the past 10000-20000 years is

associated with normal Hb level (116, 300). Single nucleotide polymorphisms (SNP) of

the HIF-2α and PHD2 gene were identified in Tibetan highlanders compared to

lowlanders. (300-303). These variants may result in reduced activity of HIF-2α and

PHD2, the net effect would change the level of HIF-2α and reduce erythropoiesis, and

hence, normal Hb (35). Despite reduced arterial oxygenation (304), Tibetan highlanders

have maintained aerobic metabolism that is due to NO-dependent increased in blood flow

(116, 305) and ventilation (306). Therefore, natural selection of important genes allows

highlanders to adapt to high altitude and survive in low oxygen environment.

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1.7: Thesis Objective

Previous studies in our laboratory demonstrated that brain nNOS protein

expression was increased in acute anemia (14, 41). However, the role of nNOS is

uncertain in this anemic condition. In Chapter 2, the role of nNOS on survival will be

assessed. Also, the cardiovascular effects in anemia will be studied in mice to provide a

physiological explanation of the role of nNOS in anemia. Besides nNOS, our laboratory

had also reported increased in HIF-1α protein expression in anemic rat brain (41). In vitro

studies have demonstrated that NO can stabilize HIF-α protein in oxygen-rich

environment, whereas NO can degrade HIF-α protein in oxygen-lacking conditions. The

in vivo evidence of the effects of NO and HIF-α is currently lacking. Thus in Chapter 3,

we will investigate the mechanism by which nNOS affect hypoxia signalling by assessing

HIF-1α expression in anemia. Lastly, the similarities and divergence between anemia and

hypoxia will be demonstrated at the cardiovascular and cellular level throughout the

thesis when appropriate in Chapters 2 and 3.

General Hypothesis:

nNOS contributes to adaptive cardiovascular and cellular mechanisms which

support survival during acute anemia

Sub-hypotheses:

1) nNOS supports survival in acute anemia

2) nNOS increases cardiovascular responses in acute anemia

3) nNOS primes hypoxia signaling (HIF) in acute anemia

4) Cardiovascular and cellular responses to anemia is different than hypoxia

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Aims

1) To determine the role of nNOS in survival and cardiovascular responses in mouse

model of acute hemodilutional anemia

2) To assess the cellular role and the mechanism of nNOS in hypoxia signaling and

HIF-α in acute anemia

3) To compare and contrast the survival, cardiovascular adaptation and molecular

responses between acute anemia and hypoxia

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1.8: References

1. Hardison R (1998) Hemoglobins from bacteria to man: evolution of different patterns of gene expression. J Exp Biol 201(Pt 8):1099-1117.

2. Singel DJ & Stamler JS (2005) Chemical physiology of blood flow regulation by red blood cells: the role of nitric oxide and S-nitrosohemoglobin. Annu.Rev.Physiol. 67:99-145.:99-145.

3. Shepherd AP, Granger HJ, Smith EE, & Guyton AC (1973) Local control of tissue oxygen delivery and its contribution to the regulation of cardiac output. Am J Physiol 225(3):747-755.

4. Schumacker PT & Samsel RW (1989) Oxygen delivery and uptake by peripheral tissues: physiology and pathophysiology. Crit Care Clin 5(2):255-269.

5. Sladen RN (1981) The oxyhemoglobin dissociation curve. Int Anesthesiol Clin 19(3):39-70.

6. Finch CA & Lenfant C (1972) Oxygen transport in man. N Engl J Med 286(8):407-415.

7. van Bommel J, et al. (2002) Intestinal and cerebral oxygenation during severe isovolemic hemodilution and subsequent hyperoxic ventilation in a pig model. Anesthesiology 97(3):660-670.

8. Jia L, Bonaventura C, Bonaventura J, & Stamler JS (1996) S-nitrosohaemoglobin: a dynamic activity of blood involved in vascular control. Nature 380(6571):221-226.

9. Doctor A, et al. (2005) Hemoglobin conformation couples erythrocyte S-nitrosothiol content to O2 gradients. Proc.Natl.Acad.Sci.U.S.A 102(16):5709-5714.

10. Ragoonanan TE, et al. (2009) Metoprolol reduces cerebral tissue oxygen tension after acute hemodilution in rats. Anesthesiology 111(5):988-1000.

11. El Beheiry MH, et al. (2011) Metoprolol impairs resistance artery function in mice. J Appl Physiol .

12. Yaseen MA, et al. (2011) Microvascular oxygen tension and flow measurements in rodent cerebral cortex during baseline conditions and functional activation. J Cereb Blood Flow Metab 31(4):1051-1063.

13. Lecoq J, et al. (2011) Simultaneous two-photon imaging of oxygen and blood flow in deep cerebral vessels. Nat Med 17(7):893-898.

14. Hare GM, et al. (2003) Hemodilutional anemia is associated with increased cerebral neuronal nitric oxide synthase gene expression. J Appl Physiol 94(5):2058-2067.

15. Sakadzic S, et al. (2010) Two-photon high-resolution measurement of partial pressure of oxygen in cerebral vasculature and tissue. Nat Methods 7(9):755-759.

16. Ward JP (2008) Oxygen sensors in context. Biochim Biophys Acta 1777(1):1-14. 17. Wheaton WW & Chandel NS (2011) Hypoxia. 2. Hypoxia regulates cellular

metabolism. Am J Physiol Cell Physiol 300(3):C385-393.

71

Chapter 1

18. Vinogradov SA, Lo LW, & Wilson DF (1999) Dendritic polyglutamic porphoryns: Probing porphyrin protection by oxygen-dependent quenching of phosphoresence. Chem Eur J. 5:1338-1347.

19. Stepuro TL & Zinchuk VV (2006) Nitric oxide effect on the hemoglobin-oxygen affinity. J Physiol Pharmacol 57(1):29-38.

20. McMahon TJ, Exton SA, Bonaventura J, Singel DJ, & Solomon SJ (2000) Functional coupling of oxygen binding and vasoactivity in S-nitrosohemoglobin. J.Biol.Chem. 275(22):16738-16745.

21. Bell EL, Emerling BM, & Chandel NS (2005) Mitochondrial regulation of oxygen sensing. Mitochondrion 5(5):322-332.

22. Chandel NS, et al. (1998) Mitochondrial reactive oxygen species trigger hypoxia-induced transcription. Proc Natl Acad Sci U S A 95(20):11715-11720.

23. Cash TP, Pan Y, & Simon MC (2007) Reactive oxygen species and cellular oxygen sensing. Free Radic Biol Med 43(9):1219-1225.

24. Taylor CT & McElwain JC (2010) Ancient atmospheres and the evolution of oxygen sensing via the hypoxia-inducible factor in metazoans. Physiology (Bethesda) 25(5):272-279.

25. Halperin ML, Cheema-Dhadli S, Lin SH, & Kamel KS (2006) Properties permitting the renal cortex to be the oxygen sensor for the release of erythropoietin: clinical implications. Clin J Am Soc Nephrol 1(5):1049-1053.

26. Johannes T, Mik EG, Nohe B, Unertl KE, & Ince C (2006) Acute Decrease in Renal Microvascular PO2 during Acute Normovolemic Hemodilution. Am.J.Physiol Renal Physiol. .

27. Hatcher JD, Chiu LK, & Jennings DB (1978) Anemia as a stimulus to aortic and carotid chemoreceptors in the cat. J.Appl.Physiol. 44(5):696-702.

28. Weir EK, Lopez-Barneo J, Buckler KJ, & Archer SL (2005) Acute oxygen-sensing mechanisms. N Engl J Med 353(19):2042-2055.

29. Michiels C (2004) Physiological and pathological responses to hypoxia. Am.J.Pathol. 164(6):1875-1882.

30. McQuillan LP, Leung GK, Marsden PA, Kostyk SK, & Kourembanas S (1994) Hypoxia inhibits expression of eNOS via transcriptional and posttranscriptional mechanisms. [erratum appears in 1995 Jun;268(6 Pt 3):section H followi; Am J Physiol 1995 Feb;268(2 Pt 2):section H following table of contents]. American Journal of Physiology 267(5:Pt 2):t-7.

31. Fish JE, et al. (2007) Hypoxia-inducible expression of a natural cis-antisense transcript inhibits endothelial nitric-oxide synthase. J Biol Chem 282(21):15652-15666.

32. Dart C & Standen NB (1995) Activation of ATP-dependent K+ channels by hypoxia in smooth muscle cells isolated from the pig coronary artery. J Physiol 483 ( Pt 1):29-39.

33. Pearce WJ (1995) Mechanisms of hypoxic cerebral vasodilatation. Pharmacol.Ther. 65(1):75-91.

34. Ward ME, et al. (2005) Hypoxia induces a functionally significant and translationally efficient neuronal NO synthase mRNA variant. J Clin Invest 115(11):3128-3139.

72

Chapter 1

35. Semenza GL (2011) Oxygen sensing, homeostasis, and disease. N Engl J Med 365(6):537-547.

36. Hirsila M, Koivunen P, Gunzler V, Kivirikko KI, & Myllyharju J (2003) Characterization of the human prolyl 4-hydroxylases that modify the hypoxia-inducible factor. J Biol Chem 278(33):30772-30780.

37. Stamler JS, et al. (1997) Blood flow regulation by S-nitrosohemoglobin in the physiological oxygen gradient. Science 276(5321):2034-2037.

38. Cosby K, et al. (2003) Nitrite reduction to nitric oxide by deoxyhemoglobin vasodilates the human circulation. Nat.Med. 9(12):1498-1505.

39. Huang Z, et al. (2005) Enzymatic function of hemoglobin as a nitrite reductase that produces NO under allosteric control. J Clin Invest 115(8):2099-2107.

40. Nagababu E, Ramasamy S, Abernethy DR, & Rifkind JM (2003) Active nitric oxide produced in the red cell under hypoxic conditions by deoxyhemoglobin-mediated nitrite reduction. J Biol Chem 278(47):46349-46356.

41. McLaren AT, et al. (2007) Increased expression of HIF-1alpha, nNOS, and VEGF in the cerebral cortex of anemic rats. Am J Physiol Regul Integr Comp Physiol 292(1):R403-414.

42. Anonymous (Worldwide prevalence of anemia 1993-2005: WHO global database on anemia. eds de Benoist B, Mclean E, Egli I, & Cogswell M (World Health Organization).

43. Shen H, Greene AS, Stein EA, & Hudetz AG (2002) Functional cerebral hyperemia is unaffected by isovolemic hemodilution. Anesthesiology 96(1):142-147.

44. Pfeffer MA, et al. (2009) A trial of darbepoetin alfa in type 2 diabetes and chronic kidney disease. N Engl J Med 361(21):2019-2032.

45. Singh AK, et al. (2006) Correction of anemia with epoetin alfa in chronic kidney disease. N Engl J Med 355(20):2085-2098.

46. Drueke TB, et al. (2006) Normalization of hemoglobin level in patients with chronic kidney disease and anemia. N Engl J Med 355(20):2071-2084.

47. Adams RJ, et al. (1998) Prevention of a first stroke by transfusions in children with sickle cell anemia and abnormal results on transcranial Doppler ultrasonography. N.Engl.J Med. 339(1):5-11.

48. Adesina O, et al. (2011) Risk of anaemia in HIV positive pregnant women in Ibadan, south west Nigeria. Afr J Med Med Sci 40(1):67-73.

49. Calis JC, et al. (2008) Severe anemia in Malawian children. N Engl J Med 358(9):888-899.

50. Ekvall H (2003) Malaria and anemia. Curr Opin Hematol 10(2):108-114. 51. Platt OS, et al. (1994) Mortality in sickle cell disease. Life expectancy and risk

factors for early death. N Engl J Med 330(23):1639-1644. 52. Sabatine MS, et al. (2005) Association of hemoglobin levels with clinical

outcomes in acute coronary syndromes. Circulation. 111(16):2042-2049. 53. Al FN, et al. (2002) Effect of anemia on 1-year mortality in patients with acute

myocardial infarction. Am Heart J 144(4):636-641. 54. Karkouti K, Wijeysundera DN, & Beattie WS (2008) Risk associated with

preoperative anemia in cardiac surgery: a multicenter cohort study. Circulation. 117(4):478-484.

73

Chapter 1

55. Kulier A, et al. (2007) Impact of preoperative anemia on outcome in patients undergoing coronary artery bypass graft surgery. Circulation. 116(5):471-479.

56. Karkouti K, et al. (2005) Hemodilution during cardiopulmonary bypass is an independent risk factor for acute renal failure in adult cardiac surgery. J.Thorac.Cardiovasc.Surg. 129(2):391-400.

57. Kapp DS, Fischer D, Gutierrez E, Kohorn EI, & Schwartz PE (1983) Pretreatment prognostic factors in carcinoma of the uterine cervix: a multivariable analysis of the effect of age, stage, histology and blood counts on survival. Int J Radiat Oncol Biol Phys 9(4):445-455.

58. Grogan M, et al. (1999) The importance of hemoglobin levels during radiotherapy for carcinoma of the cervix. Cancer 86(8):1528-1536.

59. Steyerberg EW, et al. (2008) Predicting Outcome after Traumatic Brain Injury: Development and International Validation of Prognostic Scores Based on Admission Characteristics. PLoS.Med. 5(8):e165.

60. McIntyre LA, et al. (2006) Effect of a liberal versus restrictive transfusion strategy on mortality in patients with moderate to severe head injury. Neurocrit Care 5(1):4-9.

61. Chowdhury ME, et al. (2007) Determinants of reduction in maternal mortality in Matlab, Bangladesh: a 30-year cohort study. Lancet 370(9595):1320-1328.

62. Roberts CL, et al. (2009) Trends in adverse maternal outcomes during childbirth: a population-based study of severe maternal morbidity. BMC.Pregnancy.Childbirth. 9:7.

63. Bennett CL, et al. (2008) Venous thromboembolism and mortality associated with recombinant erythropoietin and darbepoetin administration for the treatment of cancer-associated anemia. JAMA 299(8):914-924.

64. Hebert PC, et al. (1999) A multicenter, randomized, controlled clinical trial of transfusion requirements in critical care. Transfusion Requirements in Critical Care Investigators, Canadian Critical Care Trials Group. N.Engl.J.Med. 340(6):409-417.

65. Devereaux PJ, et al. (2008) Effects of extended-release metoprolol succinate in patients undergoing non-cardiac surgery (POISE trial): a randomised controlled trial. Lancet 371(9627):1839-1847.

66. Habib RH, et al. (2003) Adverse effects of low hematocrit during cardiopulmonary bypass in the adult: should current practice be changed? J.Thorac.Cardiovasc.Surg. 125(6):1438-1450.

67. Karkouti K, et al. (2005) Low hematocrit during cardiopulmonary bypass is associated with increased risk of perioperative stroke in cardiac surgery. Ann.Thorac.Surg. 80(4):1381-1387.

68. McIntyre LA, et al. (2006) Effect of a liberal versus restrictive transfusion strategy on mortality in patients with moderate to severe head injury. Neurocrit.Care. 5(1):4-9.

69. Weiskopf RB, et al. (1998) Human cardiovascular and metabolic response to acute, severe isovolemic anemia. JAMA. 279(3):217-221.

70. Weiskopf RB, et al. (2000) Acute severe isovolemic anemia impairs cognitive function and memory in humans. Anesthesiology 92(6):1646-1652.

74

Chapter 1

71. Clarke SE, et al. (2008) Effect of intermittent preventive treatment of malaria on health and education in schoolchildren: a cluster-randomised, double-blind, placebo-controlled trial. Lancet. 372(9633):127-138.

72. Scantlebury N, et al. (2011) White matter integrity and core cognitive function in children diagnosed with sickle cell disease. J Pediatr Hematol Oncol 33(3):163-171.

73. Swaminathan M, et al. (2003) The association of lowest hematocrit during cardiopulmonary bypass with acute renal injury after coronary artery bypass surgery. Ann.Thorac.Surg. 76(3):784-791.

74. van Bommel J, Siegemund M, Henny Ch P, & Ince C (2008) Heart, kidney, and intestine have different tolerances for anemia. Transl Res 151(2):110-117.

75. Murray JF & Rapaport E (1972) Coronary blood flow and myocardial metabolism in acute experimental anaemia. Cardiovasc Res 6(4):360-367.

76. Peterson PN, et al. (2010) Association of longitudinal measures of hemoglobin and outcomes after hospitalization for heart failure. Am Heart J 159(1):81-89.

77. von Haehling S, et al. (2010) Anaemia is an independent predictor of death in patients hospitalized for acute heart failure. Clin Res Cardiol 99(2):107-113.

78. Kimura H, et al. (2010) Cardio-renal interaction: impact of renal function and anemia on the outcome of chronic heart failure. Heart Vessels 25(4):306-312.

79. Saraiva F, et al. (2011) Anemia: only a marker or an independent predictor of mortality in advanced heart failure? Rev Port Cardiol 30(5):515-535.

80. Beattie WS, et al. (2010) Acute surgical anemia influences the cardioprotective effects of beta-blockade: a single-center, propensity-matched cohort study. Anesthesiology 112(1):25-33.

81. Gao F, et al. (2010) Both beta1- and beta2-adrenoceptors contribute to feedforward coronary resistance vessel dilation during exercise. Am J Physiol Heart Circ Physiol 298(3):H921-929.

82. Belhassen L, et al. (2001) Endothelial dysfunction in patients with sickle cell disease is related to selective impairment of shear stress-mediated vasodilation. Blood 97(6):1584-1589.

83. Kaul DK, Liu XD, Fabry ME, & Nagel RL (2000) Impaired nitric oxide-mediated vasodilation in transgenic sickle mouse. Am J Physiol Heart Circ Physiol 278(6):H1799-1806.

84. Gladwin MT & Kato GJ (2005) Cardiopulmonary complications of sickle cell disease: role of nitric oxide and hemolytic anemia. Hematology Am Soc Hematol Educ Program:51-57.

85. Wang P, Ba ZF, & Chaudry IH (1993) Endothelial cell dysfunction occurs very early following trauma-hemorrhage and persists despite fluid resuscitation. Am J Physiol 265(3 Pt 2):H973-979.

86. Szlyk PC, King C, Jennings DB, Cain SM, & Chapler CK (1984) The role of aortic chemoreceptors during acute anemia. Can J Physiol Pharmacol 62(5):519-523.

87. Glick G, Plauth WH, Jr., & Braunwald E (1964) Role of the autonomic nervous system in the circulatory response to acutely induced anemia in unanesthetized dogs. Journal of Clinical Investigation 43:2112-24.:2112-2124.

75

Chapter 1

88. Eichelbronner O, D'Almeida M, Sielenkamper A, Sibbald WJ, & Chin-Yee IH (2002) Increasing P(50) does not improve DO(2CRIT) or systemic VO(2) in severe anemia. Am.J.Physiol Heart Circ.Physiol 283(1):H92-101.

89. Ickx BE, Rigolet M, & Van Der Linden PJ (2000) Cardiovascular and metabolic response to acute normovolemic anemia. Effects of anesthesia. Anesthesiology 93(4):1011-1016.

90. von Restorff W, Hofling B, Holtz J, & Bassenge E (1975) Effect of increased blood fluidity through hemodilution on coronary circulation at rest and during exercise in dogs. Pflugers Arch 357(1-2):15-24.

91. Wolff CB (2007) Normal cardiac output, oxygen delivery and oxygen extraction. Adv Exp Med Biol 599:169-182.

92. Deem S, et al. (1999) Mechanisms of improvement in pulmonary gas exchange during isovolemic hemodilution. J.Appl.Physiol 87(1):132-141.

93. Sproule BJ, Mitchell JH, & Miller WF (1960) Cardiopulmonary physiological responses to heavy exercise in patients with anemia. J Clin Invest 39:378-388.

94. Johannsson H & Siesjo BK (1974) Blood flow and oxygen consumption in the rat brain in dilutional anemia. Acta Physiol Scand. 91(1):136-138.

95. Matsuoka T, Saiki C, & Mortola JP (1994) Metabolic and ventilatory responses to anemic hypoxia in conscious rats. J.Appl.Physiol 77(3):1067-1072.

96. Semenza GL (2009) Regulation of oxygen homeostasis by hypoxia-inducible factor 1. Physiology (Bethesda) 24:97-106.

97. El Hasnaoui-Saadani R, et al. (2009) Cerebral adaptations to chronic anemia in a model of erythropoietin-deficient mice exposed to hypoxia. Am J Physiol Regul.Integr.Comp Physiol. 296(3):R801-R811.

98. Barker MC, Golub AS, & Pittman RN (2007) Erythrocyte-associated transients in capillary PO2: an isovolemic hemodilution study in the rat spinotrapezius muscle. Am J Physiol Heart Circ Physiol 292(5):H2540-2549.

99. Fan FC, Chen RY, Schuessler GB, & Chien S (1980) Effects of hematocrit variations on regional hemodynamics and oxygen transport in the dog. Am.J.Physiol 238(4):H545-522.

100. Monk TG, et al. (1997) Acute normovolemic hemodilution can replace preoperative autologous blood donation as a standard of care for autologous blood procurement in radical prostatectomy. Anesth.Analg. 85(5):953-958.

101. Anonymous (1987) Multicenter trial of hemodilution in acute ischemic stroke. I. Results in the total patient population. Scandinavian Stroke Study Group. Stroke. 18(4):691-699.

102. Surgenor SD, et al. (2006) Intraoperative red blood cell transfusion during coronary artery bypass graft surgery increases the risk of postoperative low-output heart failure. Circulation.114(1 Suppl):I43-8.

103. Li M, et al. (2010) Acute anemia elicits cognitive dysfunction and evidence of cerebral cellular hypoxia in older rats with systemic hypertension. Anesthesiology 113(4):845-858.

104. Briet F, et al. (2009) Cerebral cortical gene expression in acutely anemic rats: a microarray analysis. Can J Anaesth 56(12):921-934.

76

Chapter 1

105. Tansey EA (2008) Teaching the physiology of adaptation to hypoxic stress with the aid of a classic paper on high altitude by Houston and Riley. Adv Physiol Educ 32(1):11-17.

106. Lipton AJ, et al. (2001) S-nitrosothiols signal the ventilatory response to hypoxia. Nature. 413(6852):171-174.

107. E AM (2007) Hypoxic pulmonary vasoconstriction. Essays Biochem 43:61-76. 108. Forsythe JA, et al. (1996) Activation of vascular endothelial growth factor gene

transcription by hypoxia-inducible factor 1. Mol.Cell Biol. 16(9):4604-4613. 109. Hashimoto E, et al. (1994) Rapid induction of vascular endothelial growth factor

expression by transient ischemia in rat heart. Am J Physiol 267(5 Pt 2):H1948-1954.

110. Ladoux A & Frelin C (1993) Hypoxia is a strong inducer of vascular endothelial growth factor mRNA expression in the heart. Biochem Biophys Res Commun 195(2):1005-1010.

111. Harik N, et al. (1996) Time-course and reversibility of the hypoxia-induced alterations in cerebral vascularity and cerebral capillary glucose transporter density. Brain Res. 737(1-2):335-338.

112. Goldberg MA, Dunning SP, & Bunn HF (1988) Regulation of the erythropoietin gene: evidence that the oxygen sensor is a heme protein. Science 242(4884):1412-1415.

113. Maxwell PH, Pugh CW, & Ratcliffe PJ (1993) Inducible operation of the erythropoietin 3' enhancer in multiple cell lines: evidence for a widespread oxygen-sensing mechanism. Proc.Natl.Acad.Sci.U.S.A 90(6):2423-2427.

114. Wang GL, Jiang BH, Rue EA, & Semenza GL (1995) Hypoxia-inducible factor 1 is a basic-helix-loop-helix-PAS heterodimer regulated by cellular O2 tension. Proc Natl Acad Sci U S A 92(12):5510-5514.

115. Liu L & Simon MC (2004) Regulation of transcription and translation by hypoxia. Cancer Biol Ther 3(6):492-497.

116. Erzurum SC, et al. (2007) Higher blood flow and circulating NO products offset high-altitude hypoxia among Tibetans. Proc.Natl.Acad.Sci.U.S.A. 104(45):17593-17598.

117. Hochachka PW (1998) Mechanism and evolution of hypoxia-tolerance in humans. J.Exp.Biol. 201(Pt 8):1243-1254.

118. Ignarro LJ, Buga GM, Wood KS, Byrns RE, & Chaudhuri G (1987) Endothelium-derived relaxing factor produced and released from artery and vein is nitric oxide. Proc Natl Acad Sci U S A 84(24):9265-9269.

119. Moncada S & Higgs EA (2006) The discovery of nitric oxide and its role in vascular biology. Br J Pharmacol 147 Suppl 1:S193-201.

120. Thippeswamy T, McKay JS, Quinn JP, & Morris R (2006) Nitric oxide, a biological double-faced janus--is this good or bad? Histol Histopathol 21(4):445-458.

121. Ignarro LJ, Napoli C, & Loscalzo J (2002) Nitric oxide donors and cardiovascular agents modulating the bioactivity of nitric oxide: an overview. Circ Res 90(1):21-28.

77

Chapter 1

122. Alexander JH, et al. (2007) Effect of tilarginine acetate in patients with acute myocardial infarction and cardiogenic shock: the TRIUMPH randomized controlled trial. JAMA 297(15):1657-1666.

123. Lopez A, et al. (2004) Multiple-center, randomized, placebo-controlled, double-blind study of the nitric oxide synthase inhibitor 546C88: effect on survival in patients with septic shock. Crit Care Med. 32(1):21-30.

124. Beckman JS, Beckman TW, Chen J, Marshall PA, & Freeman BA (1990) Apparent hydroxyl radical production by peroxynitrite: implications for endothelial injury from nitric oxide and superoxide. Proc Natl Acad Sci U S A 87(4):1620-1624.

125. Marozkina N, Gaston B, & Doctor A (2008) Transnitrosation signals oxyhemoglobin desaturation. Circ Res 103(5):441-443.

126. Stuehr DJ, Santolini J, Wang ZQ, Wei CC, & Adak S (2004) Update on mechanism and catalytic regulation in the NO synthases. J Biol Chem 279(35):36167-36170.

127. Dreyer J, et al. (2004) Nitric oxide synthase (NOS)-interacting protein interacts with neuronal NOS and regulates its distribution and activity. J Neurosci 24(46):10454-10465.

128. Dedio J, et al. (2001) NOSIP, a novel modulator of endothelial nitric oxide synthase activity. FASEB J 15(1):79-89.

129. Zimmermann K, et al. (2002) NOSTRIN: a protein modulating nitric oxide release and subcellular distribution of endothelial nitric oxide synthase. Proc Natl Acad Sci U S A 99(26):17167-17172.

130. Arnold DB & Heintz N (1997) A calcium responsive element that regulates expression of two calcium binding proteins in Purkinje cells. Proc Natl Acad Sci U S A 94(16):8842-8847.

131. Whalen EJ, et al. (2007) Regulation of beta-adrenergic receptor signaling by S-nitrosylation of G-protein-coupled receptor kinase 2. Cell 129(3):511-522.

132. Lipton SA, et al. (1993) A redox-based mechanism for the neuroprotective and neurodestructive effects of nitric oxide and related nitroso-compounds. Nature 364(6438):626-632.

133. Jaffrey SR, Erdjument-Bromage H, Ferris CD, Tempst P, & Snyder SH (2001) Protein S-nitrosylation: a physiological signal for neuronal nitric oxide. Nat.Cell Biol. 3(2):193-197.

134. Takahashi H, et al. (2007) Hypoxia enhances S-nitrosylation-mediated NMDA receptor inhibition via a thiol oxygen sensor motif. Neuron 53(1):53-64.

135. Gonzalez DR, Beigi F, Treuer AV, & Hare JM (2007) Deficient ryanodine receptor S-nitrosylation increases sarcoplasmic reticulum calcium leak and arrhythmogenesis in cardiomyocytes. Proc Natl Acad Sci U S A 104(51):20612-20617.

136. Burger DE, et al. (2009) Neuronal nitric oxide synthase protects against myocardial infarction-induced ventricular arrhythmia and mortality in mice. Circulation 120(14):1345-1354.

137. Li F, et al. (2007) Regulation of HIF-1alpha stability through S-nitrosylation. Mol.Cell. 26(1):63-74.

78

Chapter 1

138. Lima B, et al. (2009) Endogenous S-nitrosothiols protect against myocardial injury. Proc Natl Acad Sci U S A 106(15):6297-6302.

139. Lander HM, Ogiste JS, Pearce SF, Levi R, & Novogrodsky A (1995) Nitric oxide-stimulated guanine nucleotide exchange on p21ras. J Biol Chem 270(13):7017-7020.

140. Molina y Vedia L, et al. (1992) Nitric oxide-induced S-nitrosylation of glyceraldehyde-3-phosphate dehydrogenase inhibits enzymatic activity and increases endogenous ADP-ribosylation. J Biol Chem 267(35):24929-24932.

141. Kornberg MD, et al. (2010) GAPDH mediates nitrosylation of nuclear proteins. Nat Cell Biol 12(11):1094-1100.

142. Ozawa K, et al. (2008) S-nitrosylation of beta-arrestin regulates beta-adrenergic receptor trafficking. Mol Cell 31(3):395-405.

143. Sonveaux P, Lobysheva II, Feron O, & McMahon TJ (2007) Transport and peripheral bioactivities of nitrogen oxides carried by red blood cell hemoglobin: role in oxygen delivery. Physiology (Bethesda.) 22:97-112.

144. Liu L, et al. (2001) A metabolic enzyme for S-nitrosothiol conserved from bacteria to humans. Nature 410(6827):490-494.

145. Liu L, et al. (2004) Essential roles of S-nitrosothiols in vascular homeostasis and endotoxic shock. Cell 116(4):617-628.

146. Lillig CH & Holmgren A (2007) Thioredoxin and related molecules--from biology to health and disease. Antioxid Redox Signal 9(1):25-47.

147. Nikitovic D & Holmgren A (1996) S-nitrosoglutathione is cleaved by the thioredoxin system with liberation of glutathione and redox regulating nitric oxide. J Biol Chem 271(32):19180-19185.

148. Stoyanovsky DA, et al. (2005) Thioredoxin and lipoic acid catalyze the denitrosation of low molecular weight and protein S-nitrosothiols. J Am Chem Soc 127(45):15815-15823.

149. Benhar M, Forrester MT, Hess DT, & Stamler JS (2008) Regulated protein denitrosylation by cytosolic and mitochondrial thioredoxins. Science 320(5879):1050-1054.

150. Mannick JB, et al. (1999) Fas-induced caspase denitrosylation. Science 284(5414):651-654.

151. Shiva S, et al. (2006) Ceruloplasmin is a NO oxidase and nitrite synthase that determines endocrine NO homeostasis. Nat Chem Biol 2(9):486-493.

152. Doyle MP, Pickering RA, DeWeert TM, Hoekstra JW, & Pater D (1981) Kinetics and mechanism of the oxidation of human deoxyhemoglobin by nitrites. J Biol Chem 256(23):12393-12398.

153. Brooks J (1937) The Action of Nitrite on Hemoglobin in the Absense of Oxygen. Proceedings of the Royal Society of London 123(832):368-382.

154. Angelo M, Singel DJ, & Stamler JS (2006) An S-nitrosothiol (SNO) synthase function of hemoglobin that utilizes nitrite as a substrate. Proc Natl Acad Sci U S A 103(22):8366-8371.

155. Luchsinger BP, et al. (2003) Routes to S-nitroso-hemoglobin formation with heme redox and preferential reactivity in the beta subunits. Proc Natl Acad Sci U S A 100(2):461-466.

79

Chapter 1

156. Crawford JH, et al. (2006) Hypoxia, red blood cells, and nitrite regulate NO-dependent hypoxic vasodilation. Blood 107(2):566-574.

157. Gladwin MT, Lancaster JR, Jr., Freeman BA, & Schechter AN (2003) Nitric oxide's reactions with hemoglobin: a view through the SNO-storm. Nat.Med. 9(5):496-500.

158. Gladwin MT (2006) Role of the red blood cell in nitric oxide homeostasis and hypoxic vasodilation. Adv.Exp.Med.Biol. 588:189-205.

159. Diesen DL, Hess DT, & Stamler JS (2008) Hypoxic vasodilation by red blood cells: evidence for an s-nitrosothiol-based signal. Circ Res 103(5):545-553.

160. Isbell TS, et al. (2008) SNO-hemoglobin is not essential for red blood cell-dependent hypoxic vasodilation. Nat Med 14(7):773-777.

161. Stamler JS, Singel DJ, & Piantadosi CA (2008) SNO-hemoglobin and hypoxic vasodilation. Nat Med 14(10):1008-1009; author reply 1009-1010.

162. Palmer LA, Doctor A, & Gaston B (2008) SNO-hemoglobin and hypoxic vasodilation. Nat Med 14(10):1009; author reply 1009-1010.

163. Bin JP, et al. (2006) Effects of nitroglycerin on erythrocyte rheology and oxygen unloading: novel role of S-nitrosohemoglobin in relieving myocardial ischemia. Circulation 113(21):2502-2508.

164. Wang Y & Marsden PA (1995) Nitric oxide synthases: gene structure and regulation. Adv Pharmacol 34:71-90.

165. Wang Y, Newton DC, & Marsden PA (1999) Neuronal NOS: gene structure, mRNA diversity, and functional relevance. Crit Rev.Neurobiol. 13(1):21-43.

166. Bredt DS & Snyder SH (1989) Nitric oxide mediates glutamate-linked enhancement of cGMP levels in the cerebellum. Proc Natl Acad Sci U S A 86(22):9030-9033.

167. Matsuo R, Misawa K, & Ito E (2008) Genomic structure of nitric oxide synthase in the terrestrial slug is highly conserved. Gene 415(1-2):74-81.

168. Wang Y, et al. (1999) RNA diversity has profound effects on the translation of neuronal nitric oxide synthase. Proc Natl Acad Sci U S A 96(21):12150-12155.

169. Newton DC, et al. (2003) Translational regulation of human neuronal nitric-oxide synthase by an alternatively spliced 5'-untranslated region leader exon. J Biol Chem 278(1):636-644.

170. Silvagno F, Xia H, & Bredt DS (1996) Neuronal nitric-oxide synthase-mu, an alternatively spliced isoform expressed in differentiated skeletal muscle. J Biol Chem 271(19):11204-11208.

171. Brenman JE, et al. (1996) Interaction of nitric oxide synthase with the postsynaptic density protein PSD-95 and alpha1-syntrophin mediated by PDZ domains. Cell 84(5):757-767.

172. Eliasson MJ, Blackshaw S, Schell MJ, & Snyder SH (1997) Neuronal nitric oxide synthase alternatively spliced forms: prominent functional localizations in the brain. Proc Natl Acad Sci U S A 94(7):3396-3401.

173. Wang Y, Goligorsky MS, Lin M, Wilcox JN, & Marsden PA (1997) A novel, testis-specific mRNA transcript encoding an NH2-terminal truncated nitric-oxide synthase. J Biol Chem 272(17):11392-11401.

80

Chapter 1

174. Wang Y, et al. (2002) An alternative promoter of the human neuronal nitric oxide synthase gene is expressed specifically in Leydig cells. Am J Pathol 160(1):369-380.

175. Rameau GA, Chiu LY, & Ziff EB (2004) Bidirectional regulation of neuronal nitric-oxide synthase phosphorylation at serine 847 by the N-methyl-D-aspartate receptor. J Biol.Chem. 279(14):14307-14314.

176. Rameau GA, et al. (2007) Biphasic coupling of neuronal nitric oxide synthase phosphorylation to the NMDA receptor regulates AMPA receptor trafficking and neuronal cell death. Journal of Neuroscience 27(13):3445-3455.

177. Wang WW, et al. (2010) Transduced PDZ1 domain of PSD-95 decreases Src phosphorylation and increases nNOS (Ser847) phosphorylation contributing to neuroprotection after cerebral ischemia. Brain Res 1328:162-170.

178. Mishra OP, Ashraf QM, & Delivoria-Papadopoulos M (2009) Tyrosine phosphorylation of neuronal nitric oxide synthase (nNOS) during hypoxia in the cerebral cortex of newborn piglets: the role of nitric oxide. Neurosci Lett 462(1):64-67.

179. Christopherson KS, Hillier BJ, Lim WA, & Bredt DS (1999) PSD-95 assembles a ternary complex with the N-methyl-D-aspartic acid receptor and a bivalent neuronal NO synthase PDZ domain. J Biol Chem 274(39):27467-27473.

180. Ho GP, et al. (2011) S-nitrosylation and S-palmitoylation reciprocally regulate synaptic targeting of PSD-95. Neuron 71(1):131-141.

181. Firestein BL & Bredt DS (1999) Interaction of neuronal nitric-oxide synthase and phosphofructokinase-M. J.Biol.Chem. 274(15):10545-10550.

182. Jaffrey SR, Snowman AM, Eliasson MJ, Cohen NA, & Snyder SH (1998) CAPON: a protein associated with neuronal nitric oxide synthase that regulates its interactions with PSD95. Neuron 20(1):115-124.

183. Fang M, et al. (2000) Dexras1: a G protein specifically coupled to neuronal nitric oxide synthase via CAPON. Neuron 28(1):183-193.

184. Govers R & Oess S (2004) To NO or not to NO: 'where?' is the question. Histol Histopathol 19(2):585-605.

185. Dawson VL, Kizushi VM, Huang PL, Snyder SH, & Dawson TM (1996) Resistance to neurotoxicity in cortical cultures from neuronal nitric oxide synthase-deficient mice. J.Neurosci. 16(8):2479-2487.

186. Bender AT, Nakatsuka M, & Osawa Y (2000) Heme insertion, assembly, and activation of apo-neuronal nitric-oxide synthase in vitro. J Biol Chem 275(34):26018-26023.

187. Waheed SM, et al. (2010) Nitric oxide blocks cellular heme insertion into a broad range of heme proteins. Free Radic Biol Med 48(11):1548-1558.

188. Chakravarti R, Aulak KS, Fox PL, & Stuehr DJ (2010) GAPDH regulates cellular heme insertion into inducible nitric oxide synthase. Proc Natl Acad Sci U S A 107(42):18004-18009.

189. Paakkari I & Lindsberg P (1995) Nitric oxide in the central nervous system. Ann Med 27(3):369-377.

190. Kiss JP & Vizi ES (2001) Nitric oxide: a novel link between synaptic and nonsynaptic transmission. Trends Neurosci 24(4):211-215.

81

Chapter 1

191. Wang Y, Liu XF, Cornish KG, Zucker IH, & Patel KP (2005) Effects of nNOS antisense in the paraventricular nucleus on blood pressure and heart rate in rats with heart failure. Am.J.Physiol Heart Circ.Physiol. 288(1):H205-H213.

192. Li YF, Roy SK, Channon KM, Zucker IH, & Patel KP (2002) Effect of in vivo gene transfer of nNOS in the PVN on renal nerve discharge in rats. Am J Physiol Heart Circ Physiol 282(2):H594-601.

193. Xu KY, Huso DL, Dawson TM, Bredt DS, & Becker LC (1999) Nitric oxide synthase in cardiac sarcoplasmic reticulum. Proc.Natl.Acad.Sci.U.S.A. %19;96(2):657-662.

194. Sears CE, et al. (2003) Cardiac neuronal nitric oxide synthase isoform regulates myocardial contraction and calcium handling. Circ.Res. 92(5):e52-e59.

195. Wang H, et al. (2008) Neuronal nitric oxide synthase signaling within cardiac myocytes targets phospholamban. Am J Physiol Cell Physiol 294(6):C1566-1575.

196. Lim G, Venetucci L, Eisner DA, & Casadei B (2008) Does nitric oxide modulate cardiac ryanodine receptor function? Implications for excitation-contraction coupling. Cardiovasc Res 77(2):256-264.

197. Ashley EA, Sears CE, Bryant SM, Watkins HC, & Casadei B (2002) Cardiac nitric oxide synthase 1 regulates basal and beta-adrenergic contractility in murine ventricular myocytes. Circulation. 105(25):3011-3016.

198. Zhou L, et al. (2002) Lack of nitric oxide synthase depresses ion transporting enzyme function in cardiac muscle. Biochem Biophys Res Commun 294(5):1030-1035.

199. Khan SA, et al. (2004) Neuronal nitric oxide synthase negatively regulates xanthine oxidoreductase inhibition of cardiac excitation-contraction coupling. Proc Natl Acad Sci U S A 101(45):15944-15948.

200. Barouch LA, et al. (2002) Nitric oxide regulates the heart by spatial confinement of nitric oxide synthase isoforms. Nature. 416(6878):337-339.

201. Dawson D, et al. (2005) nNOS gene deletion exacerbates pathological left ventricular remodeling and functional deterioration after myocardial infarction. Circulation 112(24):3729-3737.

202. Bendall JK, et al. (2004) Role of myocardial neuronal nitric oxide synthase-derived nitric oxide in beta-adrenergic hyporesponsiveness after myocardial infarction-induced heart failure in rat. Circulation 110(16):2368-2375.

203. Damy T, et al. (2003) Up-regulation of cardiac nitric oxide synthase 1-derived nitric oxide after myocardial infarction in senescent rats. FASEB J 17(13):1934-1936.

204. Damy T, et al. (2004) Increased neuronal nitric oxide synthase-derived NO production in the failing human heart. Lancet 363(9418):1365-1367.

205. Sun J, et al. (2006) Hypercontractile female hearts exhibit increased S-nitrosylation of the L-type Ca2+ channel alpha1 subunit and reduced ischemia/reperfusion injury. Circ Res 98(3):403-411.

206. Saraiva RM, et al. (2005) Deficiency of neuronal nitric oxide synthase increases mortality and cardiac remodeling after myocardial infarction: role of nitroso-redox equilibrium. Circulation 112(22):3415-3422.

207. Ren YL, Garvin JL, Ito S, & Carretero OA (2001) Role of neuronal nitric oxide synthase in the macula densa. Kidney Int 60(5):1676-1683.

82

Chapter 1

208. Vallon V, et al. (2001) Feedback control of glomerular vascular tone in neuronal nitric oxide synthase knockout mice. J Am Soc Nephrol 12(8):1599-1606.

209. Wilcox CS, et al. (1992) Nitric oxide synthase in macula densa regulates glomerular capillary pressure. Proc Natl Acad Sci U S A 89(24):11993-11997.

210. Thorup C, Erik A, & Persson G (1996) Macula densa derived nitric oxide in regulation of glomerular capillary pressure. Kidney Int 49(2):430-436.

211. Kovacs G, et al. (2003) Neuronal nitric oxide synthase: its role and regulation in macula densa cells. J Am Soc Nephrol 14(10):2475-2483.

212. Siegfried G, Amiel C, & Friedlander G (1996) Inhibition of ecto-5'-nucleotidase by nitric oxide donors. Implications in renal epithelial cells. J Biol Chem 271(9):4659-4664.

213. Ortiz PA & Garvin JL (2003) Cardiovascular and renal control in NOS-deficient mouse models. Am.J.Physiol Regul.Integr.Comp Physiol 284(3):R628-R638.

214. Wang X, et al. (1998) Neuronal nitric oxide synthase is expressed in principal cell of collecting duct. Am J Physiol 275(3 Pt 2):F395-399.

215. Wang T, Inglis FM, & Kalb RG (2000) Defective fluid and HCO(3)(-) absorption in proximal tubule of neuronal nitric oxide synthase-knockout mice. Am J Physiol Renal Physiol 279(3):F518-524.

216. Bachmann S, Bosse HM, & Mundel P (1995) Topography of nitric oxide synthesis by localizing constitutive NO synthases in mammalian kidney. Am J Physiol 268(5 Pt 2):F885-898.

217. Boulanger CM, et al. (1998) Neuronal nitric oxide synthase is expressed in rat vascular smooth muscle cells: activation by angiotensin II in hypertension. Circ Res 83(12):1271-1278.

218. Schwarz PM, Kleinert H, & Forstermann U (1999) Potential functional significance of brain-type and muscle-type nitric oxide synthase I expressed in adventitia and media of rat aorta. Arterioscler Thromb Vasc Biol 19(11):2584-2590.

219. Schuh K, Uldrijan S, Telkamp M, Rothlein N, & Neyses L (2001) The plasmamembrane calmodulin-dependent calcium pump: a major regulator of nitric oxide synthase I. J Cell Biol 155(2):201-205.

220. Gros R, et al. (2003) Plasma membrane calcium ATPase overexpression in arterial smooth muscle increases vasomotor responsiveness and blood pressure. Circ Res 93(7):614-621.

221. Fleming I (2003) Brain in the brawn: the neuronal nitric oxide synthase as a regulator of myogenic tone. Circ Res 93(7):586-588.

222. Harder DR, Lange AR, Gebremedhin D, Birks EK, & Roman RJ (1997) Cytochrome P450 metabolites of arachidonic acid as intracellular signaling molecules in vascular tissue. J Vasc Res 34(3):237-243.

223. Randriamboavonjy V, Busse R, & Fleming I (2003) 20-HETE-induced contraction of small coronary arteries depends on the activation of Rho-kinase. Hypertension 41(3 Pt 2):801-806.

224. Liu X, et al. (2008) Interaction of nitric oxide, 20-HETE, and EETs during functional hyperemia in whisker barrel cortex. Am J Physiol Heart Circ Physiol 295(2):H619-631.

83

Chapter 1

225. Brenman JE, Chao DS, Xia H, Aldape K, & Bredt DS (1995) Nitric oxide synthase complexed with dystrophin and absent from skeletal muscle sarcolemma in Duchenne muscular dystrophy. Cell 82(5):743-752.

226. Thomas GD, Shaul PW, Yuhanna IS, Froehner SC, & Adams ME (2003) Vasomodulation by skeletal muscle-derived nitric oxide requires alpha-syntrophin-mediated sarcolemmal localization of neuronal Nitric oxide synthase. Circ Res 92(5):554-560.

227. Thomas GD, et al. (1998) Impaired metabolic modulation of alpha-adrenergic vasoconstriction in dystrophin-deficient skeletal muscle. Proc Natl Acad Sci U S A 95(25):15090-15095.

228. Ross RM, Wadley GD, Clark MG, Rattigan S, & McConell GK (2007) Local nitric oxide synthase inhibition reduces skeletal muscle glucose uptake but not capillary blood flow during in situ muscle contraction in rats. Diabetes 56(12):2885-2892.

229. Percival JM, Anderson KN, Gregorevic P, Chamberlain JS, & Froehner SC (2008) Functional deficits in nNOSmu-deficient skeletal muscle: myopathy in nNOS knockout mice. PLoS One 3(10):e3387.

230. Kobayashi YM, et al. (2008) Sarcolemma-localized nNOS is required to maintain activity after mild exercise. Nature 456(7221):511-515.

231. Percival JM, Anderson KN, Huang P, Adams ME, & Froehner SC (2010) Golgi and sarcolemmal neuronal NOS differentially regulate contraction-induced fatigue and vasoconstriction in exercising mouse skeletal muscle. J Clin Invest 120(3):816-826.

232. Lai Y, et al. (2009) Dystrophins carrying spectrin-like repeats 16 and 17 anchor nNOS to the sarcolemma and enhance exercise performance in a mouse model of muscular dystrophy. J Clin Invest 119(3):624-635.

233. Brunelli S, et al. (2007) Nitric oxide release combined with nonsteroidal antiinflammatory activity prevents muscular dystrophy pathology and enhances stem cell therapy. Proc Natl Acad Sci U S A 104(1):264-269.

234. Xu X, Russell T, Bazner J, & Hamilton J (2001) NMDA receptor antagonist AP5 and nitric oxide synthase inhibitor 7-NI affect different phases of learning and memory in goldfish. Brain Res 889(1-2):274-277.

235. Weitzdoerfer R, et al. (2004) Neuronal nitric oxide synthase knock-out mice show impaired cognitive performance. Nitric Oxide 10(3):130-140.

236. Huang PL, Dawson TM, Bredt DS, Snyder SH, & Fishman MC (1993) Targeted disruption of the neuronal nitric oxide synthase gene. Cell. 75(7):1273-1286.

237. Morishita T, et al. (2002) Vasculoprotective roles of neuronal nitric oxide synthase. FASEB J 16(14):1994-1996.

238. Kuhlencordt PJ, et al. (2006) Atheroprotective effects of neuronal nitric oxide synthase in apolipoprotein e knockout mice. Arterioscler Thromb Vasc Biol 26(7):1539-1544.

239. Eliasson MJ, et al. (1999) Neuronal nitric oxide synthase activation and peroxynitrite formation in ischemic stroke linked to neural damage. J Neurosci 19(14):5910-5918.

240. Huang PL (1999) Neuronal and endothelial nitric oxide synthase gene knockout mice. Braz.J.Med.Biol.Res. 32(11):1353-1359.

84

Chapter 1

241. Burkard N, et al. (2007) Conditional neuronal nitric oxide synthase overexpression impairs myocardial contractility. Circ.Res. 100(3):e32-e44.

242. Gyurko R, Leupen S, & Huang PL (2002) Deletion of exon 6 of the neuronal nitric oxide synthase gene in mice results in hypogonadism and infertility. Endocrinology 143(7):2767-2774.

243. Hall AV, et al. (1994) Structural organization of the human neuronal nitric oxide synthase gene (NOS1). J.Biol.Chem. 269(52):33082-33090.

244. Wang GL & Semenza GL (1993) General involvement of hypoxia-inducible factor 1 in transcriptional response to hypoxia. Proc.Natl.Acad.Sci.U.S.A 90(9):4304-4308.

245. Makino Y, et al. (2001) Inhibitory PAS domain protein is a negative regulator of hypoxia-inducible gene expression. Nature 414(6863):550-554.

246. Makino Y, et al. (2007) Transcriptional up-regulation of inhibitory PAS domain protein gene expression by hypoxia-inducible factor 1 (HIF-1): a negative feedback regulatory circuit in HIF-1-mediated signaling in hypoxic cells. J Biol Chem 282(19):14073-14082.

247. Maynard MA, et al. (2005) Human HIF-3alpha4 is a dominant-negative regulator of HIF-1 and is down-regulated in renal cell carcinoma. FASEB J 19(11):1396-1406.

248. Huang LE, Gu J, Schau M, & Bunn HF (1998) Regulation of hypoxia-inducible factor 1alpha is mediated by an O2-dependent degradation domain via the ubiquitin-proteasome pathway. Proc Natl Acad Sci U S A 95(14):7987-7992.

249. Kaelin WG, Jr. & Ratcliffe PJ (2008) Oxygen sensing by metazoans: the central role of the HIF hydroxylase pathway. Mol.Cell 30(4):393-402.

250. Ratcliffe PJ (2007) HIF-1 and HIF-2: working alone or together in hypoxia? J Clin.Invest. 117(4):862-865.

251. Hu CJ, Sataur A, Wang L, Chen H, & Simon MC (2007) The N-terminal transactivation domain confers target gene specificity of hypoxia-inducible factors HIF-1alpha and HIF-2alpha. Mol Biol Cell 18(11):4528-4542.

252. Sanchez M, Galy B, Muckenthaler MU, & Hentze MW (2007) Iron-regulatory proteins limit hypoxia-inducible factor-2alpha expression in iron deficiency. Nat Struct Mol Biol 14(5):420-426.

253. Stroka DM, et al. (2001) HIF-1 is expressed in normoxic tissue and displays an organ-specific regulation under systemic hypoxia. FASEB J 15(13):2445-2453.

254. Safran M, et al. (2006) Mouse model for noninvasive imaging of HIF prolyl hydroxylase activity: assessment of an oral agent that stimulates erythropoietin production. Proc.Natl.Acad.Sci.U.S.A. 103(1):105-110.

255. Hamanaka RB & Chandel NS (2009) Mitochondrial reactive oxygen species regulate hypoxic signaling. Curr Opin Cell Biol 21(6):894-899.

256. Cheng J, Kang X, Zhang S, & Yeh ET (2007) SUMO-specific protease 1 is essential for stabilization of HIF1alpha during hypoxia. Cell 131(3):584-595.

257. Jeong JW, et al. (2002) Regulation and destabilization of HIF-1alpha by ARD1-mediated acetylation. Cell 111(5):709-720.

258. Dioum EM, et al. (2009) Regulation of hypoxia-inducible factor 2alpha signaling by the stress-responsive deacetylase sirtuin 1. Science 324(5932):1289-1293.

85

Chapter 1

259. Lim JH, et al. (2010) Sirtuin 1 modulates cellular responses to hypoxia by deacetylating hypoxia-inducible factor 1alpha. Mol Cell 38(6):864-878.

260. Mylonis I, et al. (2006) Identification of MAPK phosphorylation sites and their role in the localization and activity of hypoxia-inducible factor-1alpha. J Biol Chem 281(44):33095-33106.

261. Metzen E, Zhou J, Jelkmann W, Fandrey J, & Brune B (2003) Nitric oxide impairs normoxic degradation of HIF-1alpha by inhibition of prolyl hydroxylases. Mol.Biol.Cell. 14(8):3470-3481.

262. Berchner-Pfannschmidt U, Yamac H, Trinidad B, & Fandrey J (2007) Nitric oxide modulates oxygen sensing by hypoxia-inducible factor 1-dependent induction of prolyl hydroxylase 2. J Biol.Chem. 282(3):1788-1796.

263. Palmer LA, et al. (2007) S-nitrosothiols signal hypoxia-mimetic vascular pathology. J Clin Invest 117(9):2592-2601.

264. Sumbayev VV, Budde A, Zhou J, & Brune B (2003) HIF-1 alpha protein as a target for S-nitrosation. FEBS Lett 535(1-3):106-112.

265. Park YK, et al. (2008) Nitric oxide donor, (+/-)-S-nitroso-N-acetylpenicillamine, stabilizes transactive hypoxia-inducible factor-1alpha by inhibiting von Hippel-Lindau recruitment and asparagine hydroxylation. Mol Pharmacol 74(1):236-245.

266. Schleicher M, et al. (2009) The Akt1-eNOS axis illustrates the specificity of kinase-substrate relationships in vivo. Sci Signal 2(82):ra41.

267. Hagen T, Taylor CT, Lam F, & Moncada S (2003) Redistribution of intracellular oxygen in hypoxia by nitric oxide: effect on HIF1alpha. Science 302(5652):1975-1978.

268. Brown GC & Borutaite V (2007) Nitric oxide and mitochondrial respiration in the heart. Cardiovasc Res 75(2):283-290.

269. Liu YV, et al. (2007) RACK1 competes with HSP90 for binding to HIF-1alpha and is required for O(2)-independent and HSP90 inhibitor-induced degradation of HIF-1alpha. Mol Cell 25(2):207-217.

270. Laughner E, Taghavi P, Chiles K, Mahon PC, & Semenza GL (2001) HER2 (neu) signaling increases the rate of hypoxia-inducible factor 1alpha (HIF-1alpha) synthesis: novel mechanism for HIF-1-mediated vascular endothelial growth factor expression. Mol Cell Biol 21(12):3995-4004.

271. Page EL, Chan DA, Giaccia AJ, Levine M, & Richard DE (2008) Hypoxia-inducible Factor-1{alpha} Stabilization in Nonhypoxic Conditions: Role of Oxidation and Intracellular Ascorbate Depletion. Mol.Biol.Cell 19(1):86-94.

272. Page EL, Robitaille GA, Pouyssegur J, & Richard DE (2002) Induction of hypoxia-inducible factor-1alpha by transcriptional and translational mechanisms. J Biol Chem 277(50):48403-48409.

273. Michaud MD, Robitaille GA, Gratton JP, & Richard DE (2009) Sphingosine-1-phosphate: a novel nonhypoxic activator of hypoxia-inducible factor-1 in vascular cells. Arterioscler Thromb Vasc Biol 29(6):902-908.

274. Porter JB, et al. (2005) Recent insights into interactions of deferoxamine with cellular and plasma iron pools: Implications for clinical use. Ann N Y Acad Sci 1054:155-168.

86

Chapter 1

275. Ren X, Dorrington KL, Maxwell PH, & Robbins PA (2000) Effects of desferrioxamine on serum erythropoietin and ventilatory sensitivity to hypoxia in humans. J Appl Physiol 89(2):680-686.

276. Iyer NV, et al. (1998) Cellular and developmental control of O2 homeostasis by hypoxia-inducible factor 1 alpha. Genes Dev 12(2):149-162.

277. Bosch-Marce M, et al. (2007) Effects of aging and hypoxia-inducible factor-1 activity on angiogenic cell mobilization and recovery of perfusion after limb ischemia. Circ Res 101(12):1310-1318.

278. Yu AY, et al. (1999) Impaired physiological responses to chronic hypoxia in mice partially deficient for hypoxia-inducible factor 1alpha. J Clin Invest 103(5):691-696.

279. Gruber M, et al. (2007) Acute postnatal ablation of Hif-2alpha results in anemia. Proc.Natl.Acad.Sci.U.S.A. 104(7):2301-2306.

280. Rankin EB, et al. (2007) Hypoxia-inducible factor-2 (HIF-2) regulates hepatic erythropoietin in vivo. J Clin.Invest. 117(4):1068-1077.

281. Peyssonnaux C, et al. (2007) Regulation of iron homeostasis by the hypoxia-inducible transcription factors (HIFs). J Clin Invest 117(7):1926-1932.

282. Seshadri V, Fox PL, & Mukhopadhyay CK (2002) Dual role of insulin in transcriptional regulation of the acute phase reactant ceruloplasmin. J Biol Chem 277(31):27903-27911.

283. Rolfs A, Kvietikova I, Gassmann M, & Wenger RH (1997) Oxygen-regulated transferrin expression is mediated by hypoxia-inducible factor-1. J Biol Chem 272(32):20055-20062.

284. Ouiddir A, Planes C, Fernandes I, VanHesse A, & Clerici C (1999) Hypoxia upregulates activity and expression of the glucose transporter GLUT1 in alveolar epithelial cells. Am J Respir Cell Mol Biol 21(6):710-718.

285. Hu CJ, et al. (2006) Differential regulation of the transcriptional activities of hypoxia-inducible factor 1 alpha (HIF-1alpha) and HIF-2alpha in stem cells. Mol Cell Biol 26(9):3514-3526.

286. Kim JW, Tchernyshyov I, Semenza GL, & Dang CV (2006) HIF-1-mediated expression of pyruvate dehydrogenase kinase: a metabolic switch required for cellular adaptation to hypoxia. Cell Metab. 3(3):177-185.

287. Papandreou I, Cairns RA, Fontana L, Lim AL, & Denko NC (2006) HIF-1 mediates adaptation to hypoxia by actively downregulating mitochondrial oxygen consumption. Cell Metab 3(3):187-197.

288. Majmundar AJ, Wong WJ, & Simon MC (2010) Hypoxia-inducible factors and the response to hypoxic stress. Mol Cell 40(2):294-309.

289. Kaelin WG, Jr. (2002) Molecular basis of the VHL hereditary cancer syndrome. Nat Rev Cancer 2(9):673-682.

290. Ang SO, et al. (2002) Disruption of oxygen homeostasis underlies congenital Chuvash polycythemia. Nat Genet 32(4):614-621.

291. Gordeuk VR, et al. (2004) Congenital disorder of oxygen sensing: association of the homozygous Chuvash polycythemia VHL mutation with thrombosis and vascular abnormalities but not tumors. Blood 103(10):3924-3932.

292. Sergeyeva A, et al. (1997) Congenital polycythemia in Chuvashia. Blood 89(6):2148-2154.

87

Chapter 1

88

293. Hickey MM, Lam JC, Bezman NA, Rathmell WK, & Simon MC (2007) von Hippel-Lindau mutation in mice recapitulates Chuvash polycythemia via hypoxia-inducible factor-2alpha signaling and splenic erythropoiesis. J Clin Invest 117(12):3879-3889.

294. Hickey MM, et al. (2010) The von Hippel-Lindau Chuvash mutation promotes pulmonary hypertension and fibrosis in mice. J Clin Invest 120(3):827-839.

295. Percy MJ, et al. (2008) A gain-of-function mutation in the HIF2A gene in familial erythrocytosis. N Engl J Med 358(2):162-168.

296. Percy MJ, et al. (2006) A family with erythrocytosis establishes a role for prolyl hydroxylase domain protein 2 in oxygen homeostasis. Proc Natl Acad Sci U S A 103(3):654-659.

297. Resar JR, et al. (2005) Hypoxia-inducible factor 1alpha polymorphism and coronary collaterals in patients with ischemic heart disease. Chest 128(2):787-791.

298. Hlatky MA, et al. (2007) Polymorphisms in hypoxia inducible factor 1 and the initial clinical presentation of coronary disease. Am Heart J 154(6):1035-1042.

299. Storz JF (2010) Evolution. Genes for high altitudes. Science 329(5987):40-41. 300. Beall CM, et al. (2010) Natural selection on EPAS1 (HIF2alpha) associated with

low hemoglobin concentration in Tibetan highlanders. Proc Natl Acad Sci U S A 107(25):11459-11464.

301. Yi X, et al. (2010) Sequencing of 50 human exomes reveals adaptation to high altitude. Science 329(5987):75-78.

302. Bigham A, et al. (2010) Identifying signatures of natural selection in Tibetan and Andean populations using dense genome scan data. PLoS Genet 6(9).

303. Simonson TS, et al. (2010) Genetic evidence for high-altitude adaptation in Tibet. Science 329(5987):72-75.

304. Beall CM (2007) Two routes to functional adaptation: Tibetan and Andean high-altitude natives. Proc Natl Acad Sci U S A 104 Suppl 1:8655-8660.

305. Beall CM, et al. (2001) Pulmonary nitric oxide in mountain dwellers. Nature 414(6862):411-412.

306. Zhuang J, et al. (1993) Autonomic regulation of heart rate response to exercise in Tibetan and Han residents of Lhasa (3,658 m). J Appl Physiol 75(5):1968-1973.

Chapter 2

Chapter 2:

nNOS regulates adaptive cardiovascular responses in acute anemia

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2.1: Introduction

Activation of adaptive responses to acute anemia is essential to organisms’

survival, and failure to quickly adapt to environmental changes may result in death. In

mammals, the physiological responses to acute anemia includes increased cardiac output

(CO), stroke volume (SV), heart rate (HR) and minute ventilation, coupled with reduced

systemic vascular resistance, maintained blood pressure and increased tissue oxygen

extraction. The ultimate goal of these acute responses is to optimize tissue oxygen

delivery. Importantly, vital organs (brain and heart) are preferentially perfused compared

to the less vital organs (intestines and skin) in anemia (1-4). This redistribution of blood

flow in acute anemia is an adaptive response to ensure adequate oxygen delivery to vital

organs to meet the high oxygen demand, and the survival of the organism during stress

conditions. Not surprisingly, these responses to hemodilutional anemia are well preserved

in many species, as demonstrated in humans, sheeps, dogs, cats, and rats (1-8). Despite

that these adaptive responses to acute anemia in mammals have been well studied, little is

known about how cellular pathways can regulate these adaptive responses. NO is

implicated in a variety of physiological roles, such as blood flow (9, 10), cardiac

relaxation (11, 12), respiration (13), neurotransmission (14), and renal function (15, 16)

and protein modification by S-nitrosylation (17). Since our laboratory and others have

demonstrated that brain nNOS protein expression is increased in animal model of anemia,

both acutely and chronically (18-21), it is of interest to investigate the role of nNOS in

the regulation of these responses. Surprisingly, there are currently very limited studies in

assessing the physiological responses to acute hemodilutional anemia in mouse model.

Clearly, using mouse as a model is preferred primarily because of the advantage in

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utilizing transgenic mice to study the function of a particular gene of interest. Therefore,

this chapter will investigate the differential role of nNOS in regulating cardiovascular

responses between acute anemia. Understanding the physiological role of nNOS in

anemia may help develop novel therapies to treat anemic patients

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2.2: Methods and Materials

2.2.1: Mouse Model of Hemodilutional Anemia

All animal protocols were approved by the institutional animal care committee

and in compliance with the standards set by the Canadian Council on Animal Care.

Spontaneously breathing male mice (WT, nNOS-/-, eNOS-/- and iNOS-/- of C57BL/6J

background, Jackson’s Laboratory) were anesthetized in 1.5% isoflurane with 21%

oxygen. Stepwise normovolemic hemodilution was performed by blood collection (0.2

ml) from tail nick, with an equal volume infusion of pentastarch (Bristol-Myers Squibb)

via a 27 gauge butterfly needle in the tail vein. Hemodilution was performed until

mortality (acute study) or a target hemoglobin concentration of near 50g/L was reached

(recovery study). In the acute study, mortality was assessed by the cessation of breathing.

In all physiological experiments, the body temperature of mice was maintained near

37oC.

2.2.2: Mouse Model of Acute Systemic Hypoxia

In mortality studies, mice were exposed to decreasing oxygen at 5% increments

from 21% to 5% oxygen (balance with nitrogen) under 1.5% isoflurane anesthesia. Mice

were kept in 5% oxygen until mortality, as assessed by breathing cessation. In

physiological studies, mice were exposed to normoxia (21% O2) or hypoxia (15% O2) for

10 minutes. The actual fractional inspired O2 (FIO2) was monitored by gas analyzer

(Ohmeda) throughout the experiment.

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2.2.3: Co-Oximetry and Blood Gas Analysis

Co-oximetery and blood gas machine (Radiometer, Denmark) was used to

measure hemoglobin concentration, oxygen saturation, oxygen content, methemoglobin,

pH, pCO2, pO2, and lactate. Blood was collected into a microtubes pre-coated with

heparin (Stardt). In the acute anemia study, blood was collected and measured in each

step of hemodilution. In the hypoxia study, blood gas was measured before and after 10

minutes exposure of 15% oxygen.

2.2.4: Mean Arterial Pressure

Spontaneously breathing mice were anesthetized in 1.5% isoflurane supplied with

21% O2. Mean arterial pressure (MAP) was measured continuously in acute anemia and

hypoxia (15% O2 for 10 min) studies via the femoral arterial line connected with a

pressure transducer. Data was collected by PowerLab system (ADInstrument).

2.2.5: Carotid Blood Flow

A doppler flow probe (0.75mm, Transonic Systems) was placed around the left

common carotid artery for flow measurement, and was connected to a flowmeter (TS420,

Transonic System Inc, Ithaca, New York). Data was acquired continuously by PowerLab

system (ADInstrument)

2.2.6: Echocardiography

Cardiac output was measured by ultrasound biomicroscope (Vevo 770). After the

removal of hair, an ultrasound probe was placed on the chest to obtain images from the

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heart. Electrocardiogram (EKG) was monitored continuously. Spontaneously breathing

mice were anesthetized in 1.5% isoflurane supplied with 21% O2. In anemia study,

echocardiography was measured at baseline, during and post-hemodilution (0, 1, 3, 7

days). In hypoxia study, mice were exposed to 15% oxygen for up to 40 minutes, and

echocardiography was measured at 10 min intervals. The long axis of the left ventricular

image was obtained. The end-systolic and end-diastolic areas were traced, which were

converted to volumes. Stroke volume was computed by the difference between end-

systolic and end-diastolic volumes. The product of stroke volume and heart rate yields

cardiac output.

2.2.7: Pressure-volume loops

Catheterization to the heart was performed as described previously (22). In brief,

mice were placed on a warming pad (37˚C), intubated, and ventilated using positive

pressure ventilation using 2% isoflurane. Mice were secured in a recumbent position and

the right jugular vein was cannulated. Pressure was calibrated after warming the catheter

in 0.9% NaCl at 37˚C for 30 minutes (#FT112B Scisense Inc, London, Canada). The

right internal carotid was then identified and ligated cranially. A 1.2F miniaturized

combined conductance catheter-micro-manometer (Scisense, London, Ontario) was

inserted into the right carotid artery and advanced into the left ventricle until stable

pressure volume loops were obtained. After 10 minutes, a steady state was achieved. All

steady state loops were obtained with the ventilator turned off for 5 -10 seconds with the

animal apnoeic. Using the pressure conductance data, a range of real-time functional

parameters were then measured using the ADVantage systemTM (Scisence Inc, Canada).

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These included: end diastolic pressure (EDP), end systolic pressure (ESP), end diastolic

volume (EDV), end systolic volume (ESV), cardiac output (CO), and systemic vascular

resistance (SVR).

2.2.8: Vascular Reactivity

Myogenic tone and phenylephrine-stimulated vasomotor responses were assessed

in isolated murine mesenteric resistance arteries, as described previously (23). Mice were

fully anaesthetized and then euthanized by cervical dislocation. The mesentery was

removed and placed in ice-cold MOPS buffer containing (in [mmol/L]): 145 NaCl, 4.7

KCl, 3.0 CaCl2, 1.17 MgSO4 •7H2O, 1.2 NaH2PO4 •2H2O, 2.0 pyruvate, 0.02 EDTA,

3.0 MOPS (3-morpholinopropanesulfonic acid), and 5.0 glucose. Mesenteric resistance

arteries were carefully dissected away from the mesentery, cannulated onto glass

micropipettes, stretched to their in vivo lengths and pressurized to 45mmHg. The vessels

were then warmed to 37oC; transmural pressure was increased to 60mmHg at this point.

All functional experiments were conducted in MOPS buffered saline at 37oC with no

perfusion.

To measure myogenic tone, vessels were subjected to step-wise increases in

transmural pressure (20mmHg increments) from 20-100mmHg. At each pressure step,

vessel diameter (diaactive) was measured once a steady state was reached (3-5 min).

Myogenic tone was calculated as the percent constriction in relation to the maximal

diameter at each respective transmural pressure: tone (% of diamax) = [(diamax-

diaactive)/diamax]x100, where diaactive is the vessel diameter in MOPS containing Ca2+ and

diamax is the maximal diameter in Ca2+-free MOPS.

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Vasomotor responses to increasing concentrations of phenylephrine (10-9 to 10-5

mol/L) used the tone same calculation, only in this case, diaactive represents the vessel

diameter at steady state following application of the given concentration of phenylephrine

and diamax represents the maximal diameter (measured under Ca2+-free conditions) at

60mmHg (vasomotor responses were measured at 60mmHg transmural pressure).

2.2.9: Microvascular Tissue Oxygen Tension

Oxygen tension in the blood within the microvasculature of the brain was

measured during hemodilution using Oxyphor G2 phosphorescence methodology as

previously described (1, 24, 25). This method utilizes oxygen dependent quenching of

phosphorescence to provide a quantitative measure of tissue oxygen tension in the

blood. This method is highly selective for oxygen in biological systems. It is not affected

by other molecular species, such as carbon monoxide, nitric oxide, CO2, H2S, at their

physiological concentrations or by changes in cerebral blood flow. Anesthetized (1.5 %

isoflurane; 21% O2) spontaneously breathing animals were placed in a steriotaxic frame

and two burr holes were performed over the parieto-temperal cerebral cortex, lateral to

the sagital sinus. Excitation and receiving light guides (2 mm core diameter) were

positioned such that the light passed through the cerebral cortex and deeper brain

structures. Mice were injected intravascularly with Oxyphor G2 (0.1 mg in 20 glutamate

dendrimer of Pd-tetra-(4-carboxyphenyl) tetrabenzoporphyrin) and the oxygen pressure

(by phosphorescence lifetime) measured continuously by a PMOD 5000 (Oxygen

Enterprises, Philadelphia, PA, USA).

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2.2.10: Data Analysis

Data were expressed as mean ± SEM. A two-way repeated measure ANOVA was used to

assess changes in mean values between strain (WT vs nNOS-/-) and treatment (baseline vs

anemia). In all cases, a post hoc Tukey test was used. For Kaplan Myer survival curve,

Mantel-Cox log-rank test followed by chi-square test was used to determine the

differences in survival outcome. Significance was assigned at a P<0.05.

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2.3: Results

2.3.1: nNOS is protective in acute anemia

To determine the role of each NOS isoforms in acute anemia, isoflurane

anesthetized WT and NOS-deficient mice were hemodiluted until mortality. We found

that only nNOS-/- mice died earlier and at a higher Hb concentration compared to all other

strains, suggesting nNOS promotes survival in acute hemodilutional anemia (Figure 2.1).

To assess whether tissue hypoxia was the reason that nNOS-/- mice died earlier, mice

were subjected to reducing FIO2 until 5% oxygen under isoflurane anesthesia.

Paradoxically, nNOS-/- mice survived longer relative to WT counterparts, indicating that

the lack of nNOS promotes survival in systemic hypoxia (Figure 2.1). These findings

suggested that a differential role of nNOS in anemia and hypoxia.

To exclude the possibility that the mortality difference was affected by isoflurane

anesthesia, increasing dose of isoflurane concentration was tested in WT and NOS-/- mice

at 21% O2. Importantly, there was no mortality observed at 1.5% isoflurane, which was

the level used in anemic and hypoxic mice (Figure 2.2). At the high end of isoflurane

concentration at 4 - 5%, nNOS-/- mice died earlier and at lower end-tidal isoflurane

concentration compared to WT (p<0.01; Mantel-Cox LogRank Test). Thus, this

suggested that nNOS-/- mice appear to be more sensitive to high level of isoflurane

anesthetic, which agrees with a previous report (26). This also implied that mortality in

anemic and hypoxic mouse studies was not due to lower level of isoflurane anesthesia

exposure.

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99

Figure 2.1: Survival curve in acute anemia and hypoxia of WT and NOS-/- mice. (A) Anesthetized (1.5% isoflurane) mice were hemodiluted until mortality. The median terminal Hb level for mice: WT (26g/L), nNOS-/- (36g/L), eNOS-/- (28 g/L), iNOS-/- (23g/L) (B) Anesthetized (1.5% isoflurane) mice were exposed to decreasing inspired O2 concentration from 21% to 5% at an increment of 5% O2 for 10 min each. No mortality was noted above 5% O2 under isoflurane. The median survival time at 5 % O2 for mice: WT (3.5 min), nNOS-/- (10 min), eNOS-/- (10 min) and iNOS-/- (5 min). Mortality was defined by breathing cessation.

Chapter 2

Figure 2.2: Isoflurane survival curve of WT, nNOS-/- and eNOS-/- mice. Increasing isoflurane level at an 0.5% increment for 10 min each was supplied to mice at 21% O2. Mortality was noted by breathing cessation. The median survival isoflurane concentration for mice: WT (4.7%), nNOS-/- (4.5%), and eNOS-/- (4.6%)

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2.3.2: nNOS regulates the cardiovascular responses in acute anemia

To determine the possible mechanism of increased mortality in anemic mice,

cardiovascular response was assessed by two methods (echocardiography and pressure-

volume loop). Data obtained from echocardiography demonstrated that moderate

(~90g/L) and severe anemia (~50g/L) resulted in a graded increase in CO, SV and HR in

WT mice (Figure 2.3). Increased LVEDV indicated improved left ventricular filling

(preload) during acute anemia. MAP measured from femoral artery was maintained

during hemodilution, confirming that hypotension did not occur (Figure 2.3). Calculation

of SVR by dividing MAP by CO demonstrated that SVR was reduced by 36% in severely

anemic WT mice (Figure 2.6). Similar CO and SVR results were confirmed by pressure-

volume loop techniques in a separate group of anemic mice (Figure 2.4). These

cardiovascular responses to acute hemodilutional anemia in our mouse model were

similar to other species studied (1, 2, 6, 7, 27, 28). To determine the role of nNOS in the

cardiovascular response to acute anemia, mice lacking the full length nNOS protein

(nNOS-/-) were hemodiluted and compared to WT mice. Surprisingly, despite increase

HR and maintained MAP that were similar to WT anemic mice, nNOS-/- anemic mice

failed to increase CO, SV and LVEDV. The similar increase in HR in anemic WT and

nNOS-/- mice suggested activation of sympathetic system in acute anemia. The reduction

in SVR was also smaller (10%) compared to WT mice (36%), suggesting a lack of

vasodilation in nNOS deficiency abolished increased CO response. To determine whether

the decrease in SVR was due to intrinsic factors, reactivity studies on isolated mesenteric

resistance artery of anemic WT and nNOS-/- mice demonstrated that anemia did not

impact the phenylephrine response or the myogenic tone (Figure 2.7). These results from

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isolated vessels suggested that reduced SVR in anemia is due to extrinsic factors to the

resistance vessels, such as SNO protein. Collectively, these data suggested that nNOS

contributes to increase CO response by increasing preload and reducing SVR during

acute anemia.

Unlike acute anemia, the cardiovascular responses were dramatically different in

mice exposed to acute hypoxia (15% O2). In WT mice, hypoxia resulted in increased HR

but a drop in MAP (Figure 2.5). However, CO and SV remained unchanged. A 13%

reduction in MAP and constant CO resulted in decreased SVR by 15% in hypoxia (Figure

2.6). Systemic hypotension may occur in hypoxia due to the release of systemic

vasodilators (29). In contrast, hypoxic nNOS-/- mice had an increased CO, SV, HR, a

slight reduction in LVEDV and maintained MAP. This led to a further reduction in SVR

(23%) compared to WT (15%) (Figure 2.6). These results suggested that nNOS

contributes to different cardiovascular responses hypoxia relative to anemia, which may

contribute to the observed mortality patterns. Interestingly, a common observation

between the two conditions was that mice that had increased CO and decreased SVR had

increased survival (i.e. WT anemic mice and nNOS-/- hypoxic mice).

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Figure 2.3: Echocardiography measurements of the cardiovascular responses to acute hemodilutional anemia in mice. (A) Cardiac output, (B) stroke volume, (C) heart rate, (D) mean arterial pressure, and (E) end-diastolic volume were measured in anemic WT (white bar) and nNOS-/- (black bar) mice (n = 6 / group). (F) Systemic vascular resistance was derived by dividing mean arterial pressure and cardiac output *p<0.05 vs baseline; #p<0.05 vs WT

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Figure 2.4: Pressure-volume loop measurement of cardiovascular responses in mice at moderate anemia (~90g/L). (A) Cardiac output, (B) stroke volume, (C) mean arterial pressure, (D) systemic vascular resistance were measured in anemic WT (white bar) and nNOS-/- (black bar) mice (n = 6-7 / group). *p<0.05 vs baseline; #p<0.05 vs WT

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Figure 2.5: Cardiovascular responses to acute hypoxia in mice by echocardiography. (A) Cardiac output, (B) stroke volume, (C) heart rate, (D) mean arterial pressure, and (E) end-diastolic volume were measured in normoxic (21% O2) and hypoxic (15% O2) WT (white bar) and nNOS-/- (black bar) mice (n = 6 / group). (F) Systemic vascular resistance was derived by dividing mean arterial pressure and cardiac output. *p<0.05 vs baseline; #p<0.05 vs WT

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Figure 2.6: Percent changes of cardiac output (CO), systemic vascular resistance (SVR), and mean arterial pressure (MAP) relative to baseline/normoxia in anemic and hypoxic mice.

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Figure 2.7: Phenylephrine responses (top) and myogenic tone assessments (bottom) under control and anemia conditions for WT (left) and nNOS-/- mice (right). With the exception of a slight (but significant) increase in constriction at 10-7 mol/L phenylephrine, anemia did not impact phenylephrine responses in WT mice. EC50 values in WT mice were not different (con = 5.2 ± 1.4 x10-7 mol/L, n=5; anemic = 2.3 ± 4.3 x10-

7 mol/L, n=6; P = N.S.). Phenylephrine responses in nNOS-/- mice were not different (EC50 con = 4.7 ± 0.8 x10-7 mol/L, n=6; anemic = 4.1 ± 1.3 x10-7 mol/L, n=5; P = N.S.). Anemia did not alter myogenic tone in WT mice. Anemia elicited a slight (but significant) reduction in myogenic tone observed at 40mmHg, although myogenic tone was not altered at any other transmural pressure.

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2.3.3: nNOS limits the reduction in global oxygen delivery during acute anemia Global oxygen delivery, a function of CO and blood O2 content, was reduced in

proportion with the severity of anemia. In severe anemia (50g/L), the decrease in global

oxygen delivery was more dramatic in nNOS-/- mice which failed to increase CO, relative

to WT mice (Figure 2.8).

In acute hypoxia, increased CO in nNOS-/- mice was accompanied by an increased

global oxygen delivery. In contrast, failure to increase CO response in hypoxic WT mice

resulted in a reduced in global oxygen delivery (Figure 2.8).

2.3.4: Regulation of microvascular brain PO2 in acute anemia is independent of nNOS

Due to the striking contrast in the cardiovascular responses to acute anemia

between WT and nNOS-/- mice, we investigated whether nNOS affects brain oxygen level

differently. In acute anemia, the increase in carotid blood flow and decrease in

microvascular brain PO2 were comparable in nNOS-/- mice despite a lack of CO response,

relative to WT mice (Figure 2.9). These measurements suggested that reduction in brain

PO2 and increase in brain perfusion did not depend on nNOS genotype during anemia.

Similarly, acute hypoxia (15% O2) leads to comparable reduction in brain PO2

regardless of nNOS genotype, with a lack of increase in carotid blood flow (Figure 2.10).

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Figure 2.8: Global oxygen delivery in acute anemia and hypoxia. Global oxygen delivery was calculated from the product of cardiac output and blood O2 content. In acute anemia study, WT (white bar) and nNOS-/- (black bar) mice were hemodiluted to Hb near 90g/L (moderate) and 50g/L (severe) (n=6/group). In acute hypoxia study, mice were exposed to 21% O2 (normoxia) or 15% O2 (hypoxia) (n=6/group). *p<0.05 vs baseline; #p<0.05 vs WT.

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Figure 2.9: Measurement of carotid blood flow (A) and brain microvascular oxygen tension (B) in acutely anemic mice (n=6/group). *p<0.05 vs baseline

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Figure 2.10: Measurement of carotid blood flow (A) and brain microvascular oxygen tension (B) in mice exposed to normoxia (21% O2) and hypoxia (15% O2) (n=6/group). *p<0.05 vs baseline

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2.3.5: nNOS did not impact co-oximetry results in anemia

In acute anemia, Hb concentration was stepwise reduced to near 90g/L (moderate

anemia) and 50g/L (severe anemia) (Table 2.1). Blood O2 content were reduced similarly

in WT and nNOS-/- mice as expected. Hb saturation increased in anemic mice, and this

effect was not affected by nNOS.

In acute hypoxia, there were no changes in Hb concentration in WT and nNOS-/-

mice (Table 2.1). Blood O2 content was decreased similarly in both strains. Interestingly,

WT mice had a more profound reduction in blood oxygen saturation compared to nNOS-/-

mice in hypoxia.

2.3.6: nNOS did not affect blood gas status in anemia

In WT and nNOS-/- anemic mice, pH was increased similarly with severe anemia

(Table 2.1). PaO2 was similar at baseline and increased during hemodilution. PaCO2 was

reduced similarly with severe anemia in both strains of mice.

In acute hypoxia (15% O2), pH was not statistically different between normoxic

and hypoxic WT and nNOS-/- mice (Table 2.1). PaO2 was similarly reduced in hypoxic

exposure for WT and nNOS-/- mice. There was a trend for reduced PaCO2 in WT mice

exposed to hypoxia, but did not reach statistically significance, whereas the reduction

PaCO2 was significant in hypoxic nNOS-/- mice.

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Anemia Hypoxia

Mouse Strain

Baseline (~130g/L)

Moderate (~90g/L)

Severe (~50g/L)

Normoxia (21% O2)

Hypoxia (15% O2)

Hb (g/L) WT 130.4 ± 2.1 91.9 ± 1.4 * 52.6 ± 0.8 * 140.6 ± 1.6 139.2 ± 1.9 nNOS-/- 134.0 ± 2.6 84.5 ± 0.9 * 52.5 ± 0.9 * 143.4 ± 3.3 137.0 ± 5.4

SaO2 (%) WT 92.6 ± 1.0 97.3 ± 0.5 * 99.3 ± 0.4 * 94.5 ± 2.5 75.2 ± 2.5 * nNOS-/- 93.7 ± 1.1 97.9 ± 0.7 * 99.4± 0.2 * 95.3 ± 1.0 86.7 ± 1.0 *†

O2 Content (mM)

WT 6.9 ± 0.1 5.0 ± 0.1 * 3.0 ± 0.1 * 7.9 ± 0.1 6.3 ± 0.5 * nNOS-/- 7.2 ± 0.2 4.9 ± 0.1 * 3.0 ± 0.1 * 8.3 ± 0.2 7.1 ± 0.3 *†

pH WT 7.28 ± 0.02 7.32 ± 0.02 7.38 ± 0.01 * 7.32 ± 0.03 7.38 ± 0.03 nNOS-/- 7.28 ± 0.03 7.33 ± 0.03 7.35 ± 0.02 * 7.35 ± 0.04 7.41 ± 0.03

PaCO2 (mmHg)

WT 40.6 ± 2.0 38.9 ± 1.5 32.8 ± 1.1 * 33.8 ± 2.9 29.0 ± 3.1 nNOS-/- 36.1 ± 2.7 33.5 ± 2.0 29.3 ± 1.3 * 28.5 ± 1.0 22.0 ± 0.9 *

PaO2 (mmHg)

WT 98.1 ± 4.2 112.5 ± 4.3 * 129.6 ± 4.5 * 101.4 ± 8.1 57.8 ± 1.9 * nNOS-/- 107.2 ± 4.3 125.4 ± 5.6 129.0 ± 6.6 * 97.06 ± 2.9 64.4 ± 2.9 *

Data are represented as mean ± SEM. *, p<0.05 vs Baseline or Normoxia; †, p<0.05 vs WT; Hb, Hemoglobin concentration (g/L); SaO2, Oxygen saturation of arterial blood (%); PaCO2, partial pressure of carbon dioxide in arterial blood; PaO2, partial pressure of oxygen in arterial blood;

Table 2.1: Co-oximetry and blood gas analysis from anemic and hypoxic mice.

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2.3.7: nNOS may mediate increase in methemoglobin (MetHb) levels in anemia

MetHb, an oxidized form of Hb, is produced in reaction with NO. MetHb level

was increased in WT anemic mice, but the increase MetHb level was lowered in nNOS-/-

mice (Figure 2.11). These results suggested that nNOS may mediate, in part, increase

MetHb level in acute anemia

In contrast to anemia, MetHb was reduced similarly in WT and nNOS-/- hypoxic

mice. (Figure 2.11).

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Figure 2.11: MetHb levels of anemic and hypoxic WT and nNOS-/- mice. In anemia study, MetHb was measured at baseline (120 – 140 g/L), moderate (80 – 100 g/L) and severe (40 – 60 g/L) anemia. In hypoxia study, measurement was made in normoxia (21% O2) and hypoxia (15% O2) (n>6 / group). *p<0.05 vs normoxia; #p<0.05 vs WT.

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2.4: Summary of main findings:

This study has demonstrated several novel findings which illustrate that nNOS is

critically important to the integrative physiological response to acute anemia. First,

increased mortality in anemic nNOS-/- mice suggested that nNOS is protective in anemia.

Second, nNOS-/- mice failed to regulate CO and SVR in a manner that optimizes tissue

perfusion suggesting that regulation of cardiovascular response may increase survival in

anemia. Assessment of isolated resistance arteries did not demonstrate a vasodilatory

phenotype in response to anemia. This suggested that the reduction in SVR was mediated

by nNOS-dependent factors that were extrinsic to the resistance artery in anemia, such as

SNO-Hb. Third, nNOS-/- mice had a more drastic reduction in global oxygen delivery as

a result of the inability to increase CO. Despite these differences in cardiovascular

function, cerebral tissue PO2 decrease comparably in anemic mice regardless of nNOS

genotype. These results demonstrated that the mechanism by which nNOS contributed to

improved survival in anemia could be due to the activation of these adaptive

cardiovascular responses which optimize tissue perfusion and O2 delivery.

The study also highlights previously unappreciated differences between acute

anemia and acute hypoxia. First, unlike acute anemia, nNOS was detrimental to survival

in acute hypoxia as demonstrated that nNOS-/- survived longer. Second, the ability of

hypoxic nNOS-/- mice to regulate CO and SVR to optimize tissue blood flow and increase

global oxygen delivery may contribute to survival. Third, hypoxia leads to reduced blood

oxygen content that is due to reduced SaO2 and PaO2 while Hb levels is unchanged, but

anemia reduces blood oxygen content by decreased Hb levels while SaO2 and PaO2 values

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are increased. These differences demonstrated fundamental differences between the two

conditions, and discussion will be expanded in Chapter 4.

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2.5. References: 1. Ragoonanan TE, et al. (2009) Metoprolol reduces cerebral tissue oxygen tension

after acute hemodilution in rats. Anesthesiology 111(5):988-1000. 2. van Bommel J, et al. (2002) Intestinal and cerebral oxygenation during severe

isovolemic hemodilution and subsequent hyperoxic ventilation in a pig model. Anesthesiology 97(3):660-670.

3. Grupp I, Grupp G, Holmes JC, & Fowler NO (1972) Regional blood flow in anemia. J Appl Physiol 33(4):456-461.

4. Fan FC, Chen RY, Schuessler GB, & Chien S (1980) Effects of hematocrit variations on regional hemodynamics and oxygen transport in the dog. Am.J.Physiol 238(4):H545-522.

5. Eichelbronner O, D'Almeida M, Sielenkamper A, Sibbald WJ, & Chin-Yee IH (2002) Increasing P(50) does not improve DO(2CRIT) or systemic VO(2) in severe anemia. Am.J.Physiol Heart Circ.Physiol 283(1):H92-101.

6. Ickx BE, Rigolet M, & Van Der Linden PJ (2000) Cardiovascular and metabolic response to acute normovolemic anemia. Effects of anesthesia. Anesthesiology 93(4):1011-1016.

7. Lovegrove TD, Gowdey CW, & Stevenson JA (1957) Sympathoadrenal system and response of heart to acute exchange anemia. Circ Res 5(6):659-663.

8. Weiskopf RB, et al. (1998) Human cardiovascular and metabolic response to acute, severe isovolemic anemia. JAMA. 279(3):217-221.

9. Hudetz AG, Shen H, & Kampine JP (1998) Nitric oxide from neuronal NOS plays critical role in cerebral capillary flow response to hypoxia. Am.J.Physiol 274(3 Pt 2):H982-H989.

10. Hudetz AG, Wood JD, & Kampine JP (2000) 7-Nitroindazole impedes erythrocyte flow response to isovolemic hemodilution in the cerebral capillary circulation. J.Cereb.Blood Flow Metab 20(2):220-224.

11. Sears CE, et al. (2003) Cardiac neuronal nitric oxide synthase isoform regulates myocardial contraction and calcium handling. Circ.Res. 92(5):e52-e59.

12. Barouch LA, et al. (2002) Nitric oxide regulates the heart by spatial confinement of nitric oxide synthase isoforms. Nature. 416(6878):337-339.

13. Kline DD, Yang T, Huang PL, & Prabhakar NR (1998) Altered respiratory responses to hypoxia in mutant mice deficient in neuronal nitric oxide synthase. J Physiol 511 ( Pt 1):273-287.

14. Paakkari I & Lindsberg P (1995) Nitric oxide in the central nervous system. Ann Med 27(3):369-377.

15. Ortiz PA & Garvin JL (2003) Cardiovascular and renal control in NOS-deficient mouse models. Am.J.Physiol Regul.Integr.Comp Physiol 284(3):R628-R638.

16. Wang T, Inglis FM, & Kalb RG (2000) Defective fluid and HCO(3)(-) absorption in proximal tubule of neuronal nitric oxide synthase-knockout mice. Am J Physiol Renal Physiol 279(3):F518-524.

17. Jaffrey SR, Erdjument-Bromage H, Ferris CD, Tempst P, & Snyder SH (2001) Protein S-nitrosylation: a physiological signal for neuronal nitric oxide. Nat.Cell Biol. 3(2):193-197.

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18. McLaren AT, et al. (2007) Increased expression of HIF-1alpha, nNOS, and VEGF in the cerebral cortex of anemic rats. Am J Physiol Regul Integr Comp Physiol 292(1):R403-414.

19. Hare GM, et al. (2003) Hemodilutional anemia is associated with increased cerebral neuronal nitric oxide synthase gene expression. J Appl Physiol 94(5):2058-2067.

20. El Hasnaoui-Saadani R, et al. (2009) Cerebral adaptations to chronic anemia in a model of erythropoietin-deficient mice exposed to hypoxia. Am J Physiol Regul.Integr.Comp Physiol. 296(3):R801-R811.

21. Ward ME, et al. (2005) Hypoxia induces a functionally significant and translationally efficient neuronal NO synthase mRNA variant. J Clin Invest 115(11):3128-3139.

22. Yuen DA, et al. (2010) Culture-modified bone marrow cells attenuate cardiac and renal injury in a chronic kidney disease rat model via a novel antifibrotic mechanism. PLoS One 5(3):e9543.

23. Hoefer J, et al. (2010) Sphingosine-1-phosphate-dependent activation of p38 MAPK maintains elevated peripheral resistance in heart failure through increased myogenic vasoconstriction. Circ Res 107(7):923-933.

24. Vinogradov SA, Lo LW, & Wilson DF (1999) Dendritic polyglutamic porphoryns: Probing porphyrin protection by oxygen-dependent quenching of phosphoresence. Chem Eur J. 5:1338-1347.

25. Rietveld IB, Kim E, & Vinogradov SA (2003) Dendrimers with tetrabenzoporphyrin cores: near infrared phosphors for in vivo oxygen imaging. Tetrahedron 59:3821-3831.

26. Engelhardt T, Lowe PR, Galley HF, & Webster NR (2006) Inhibition of neuronal nitric oxide synthase reduces isoflurane MAC and motor activity even in nNOS knockout mice. Br J Anaesth 96(3):361-366.

27. Habler OP, et al. (1996) The effect of acute normovolemic hemodilution (ANH) on myocardial contractility in anesthetized dogs. Anesth.Analg. 83(3):451-458.

28. Nozaki J, Kitahata H, Tanaka K, Kawahito S, & Oshita S (2002) The effects of acute normovolemic hemodilution on left ventricular systolic and diastolic function in the absence or presence of beta-adrenergic blockade in dogs. Anesth.Analg. 94(5):1120-1126, table.

29. Pearce WJ (1995) Mechanisms of hypoxic cerebral vasodilatation. Pharmacol Ther 65(1):75-91.

Chapter 3

Chapter 3:

nNOS is required to increase HIF-1α protein expression in severe anemia

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3.1: Introduction

Anemia is a global health problem which is associated with increased mortality.

Current understanding of the cellular responses in anemia is poorly understood. Though

the treatment of anemia often involves transfusion and erythropoiesis stimulating agent,

such as darbepoietin and erythropoietin, it is now appreciated that such treatment does

not improve morbidity and mortality (1-3). These findings suggested the need to define

the underlying physiological and molecular mechanisms in anemia such that problems in

anemic patients can be addressed. Our laboratory had previously demonstrated an

increase in hypoxic gene expression, with nNOS and HIF-1α being the most robustly

expressed molecules in the brain of anemic rodents (4, 5). In addition, others have

demonstrated that disrupting the hypoxia/HIF-α pathway results in anemia (6, 7). Thus, it

is important to investigate the targets that regulate the HIF-α pathway in anemia.

NO is a molecule that affects a diverse aspect of physiological responses, such as

the regulation of neuronal (8), vascular tone (9), renal function (10), and modification of

gene expression by S-nitrosylation (11). S-nitrosylation of Hb (SNO-Hb) has been

implicated as an important physiological regulator of hypoxia-induced vasodilation (12)

and ventilation (13). In the last decade, in vitro studies have demonstrated an association

between NO and HIF-1α. NO can stabilize HIF-1α in normoxic conditions, whereas NO

can degrade HIF-1α in hypoxic settings. In normoxia, HIF-1α is usually degraded, but

NO can stabilize HIF-α by a number of mechanisms, including a) inhibition of PHD

activity (14, 15) , b) S-nitrosylation of pVHL (16) and c) S-nitrosylation of HIF-1α (17-

19). All of these actions would prevent the interaction between pVHL and HIF-1α, thus

HIF-1α protein escapes the degradation pathway and become stabilized. Accumulation of

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cytosolic HIF-1α protein translocates to the nucleus and binds with HIF-1β. The resultant

formation of HIF-1 complex can activate transcription of genes containing hypoxia-

response elements (HRE), such as VEGF, EPO and GLUT1, which are important

molecules that can lead to increased tissue oxygen delivery. Conversely, NO can degrade

HIF-1α in hypoxia due to inhibition of mitochondrial respiration by NO, thus reducing

the subcellular oxygen consumption and redistribute oxygen elsewhere (20). However,

there are very limited in vivo evidence for this relationship (18), thus warrant for further

investigation.

To date, there are three NOS isoforms identified that can produce NO in

mammals. The sources of NO that regulate HIF-1α protein levels can be derived from

any of the NOS isoforms, and it is dependent on the stimulus and the cell type that the

NOS enzymes express. For example, iNOS-derived NO had been shown to S-nitrosylate

HIF-1α, and thus stabilizing HIF-1α protein expression in macrophage in a cancer model

(17). In ischemic endothelial cells, eNOS-derived NO can stabilize HIF-1α by inhibiting

PHD activity (21). In addition, regulating the degradation of NO availability by S-

nitrosoglutathione reductase (GSNOR) had been demonstrated to stabilize HIF-1α in the

heart via an in vivo myocardial infarction model (18). Since the role of nNOS-derived

NO in regulating HIF-1α expression is currently lacking in an in vivo model, this study is,

therefore, carried out to investigate the cellular function of nNOS in a mouse model of

acute hemodilutional anemia. Identifying the molecular targets in anemia and hypoxia

may help diagnosis and develop novel therapeutic interventions for patients.

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3.2: Methods and Materials

3.2.1: Acute hemodilutional anemia

All animal use protocols were in accordance of the standards set by the Canadian

Council of Animal Care and approved at the institutional animal care committee.

Spontaneously breathing WT and nNOS-/- mice (Jackson’s Laboratory) were anesthetized

in 1.5% isoflurane in 21% O2 and hemodiluted with pentastarch via tail vein until a target

Hb concentration of near 50g/L was reached. Mice were recovered from anesthesia.

Tissues were harvested 1, 6, and 24 hours following hemodilution for assessment of RNA

levels by real-time PCR and protein expression by Western blot.

3.2.2: Acute hypoxia

Mice were exposed to 6% O2 for 6 hours in a sealed, air-tight chamber with

access to food and water. Tissues were harvested immediately following hypoxia

exposure for RNA and protein measurement.

3.2.3: Western Blot

Tissues were excised from control and anemic (1, 6, 24 hours) mice, snap frozen

by liquid nitrogen and stored at -80ºC. Tissue were homogenized using a polytron

(Beckman) in buffer A (20mM HEPES pH 7.5, 1.5mM MgCl2, 0.2mM EDTA, 100mM

NaCl, 0.5mM PMSF, 1μg/ml Leupeptin, 0.2mM DTT), and centrifuged at 10000g for

30min in 4ºC. Total protein lysate was obtained by mixing the supernatant with buffer B

(buffer A + 40% v/v Glycerol). Protein samples were quantified by Lowry assay

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(Biorad), aliquoted and stored at -80ºC. 20μg of protein from mouse brain were separated

on a 7.5% SDS-polyacrylamide gel, transferred onto nitrocellulose membrane, and

transfer efficiency was verified by Ponceau red-stained membranes. Membranes were

blocked with 5% milk, and probed with monoclonal nNOS (BD Bioscences; 140kDa),

polyclonal HIF-1α (R & D; 120kDa), HIF-2α (Novus Biologicals; 120kDa) and GSNOR

(Cell Signalling) antibodies. Immunoblots were probed with appropriate secondary HRP

antibodies. Immunoreactive bands were detected by enhanced chemiluminescence

(Sigma) and quantified by densitometry using ImageJ software. α-tubulin (sigma) was

used as loading control.

3.2.4: Immunofluorescence staining

24hr following anemia (Hb ~ 50g/L), animals were anesthetized and gravity

perfused via cardiac puncture with 4% formaldehyde. Tissues were extracted and placed

in 4% formaldehyde for blocking. Immunofluroescence was performed on 10-μm fixed

tissue sections (4% paraformaldehyde) by incubating slides overnight at 4°C with

diluents of specific primary monoclonal antibodies for neuronal nitric oxide synthase

(nNOS) (BD Biosciences), and HIF-1α (Novus Biologicals) and specific binding detected

with an appropriately labeled secondary antibody. Microscopy was performed utilizing

fluorescent and confocal microscopes (Nikon ECLIPSE 90i, Bio-Rad Radiance 2100).

3.2.5: Quantitative Real-Time PCR

RNA was extracted from tissue samples by a handheld homogenizer using the

guandidium thiocynaide / phenol chloroform methodology as previously described (22).

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1μg of RNA was used to synthesize first strand cDNA using random primers and

Superscript III® Reverse Transcriptase (Invitrogen). RNase H was used to remove RNA

during cDNA synthesis. A known amount of exogenous luciferase was added into the

sample to control for RNA extraction and first strand cDNA synthesis.

Real-time PCR was performed using an ABI PRISM 7900HT (Applied

Biosystems) using SYBR green detection system. Reactions were performed in triplicate.

To quantify copy number, serial dilution of plasmids corresponding to the target gene

was used to construct standard curve. Target genes were corrected for efficiencies of

RNA extraction and first strand cDNA synthesis as indicated by exogenous luciferase

measurement. Data were converted to copy number, normalized to baseline, and

corrected for luciferase efficiency. Primers used for real-time PCR are listed in Table 3.1.

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Gene Sense Primer (5’ – 3’) Antisense Primer (5’ – 3’) Exon Ct 1000

EPO

CAC CCT GCT GCT TTT ACT CT

AAC CCA TCG TGA CAT TTT CT

3, 4 28.3

VEGF

ACC GCG AGG CAG CTT GAG TTA

ACC GCC TTG GCT TGT CAC AT

6, 7 25.5

PDK1

GTC GCC ACT CTC CAT GAA G

TGG GGT CCT GAG AAG ATT ATC

1, 2 25.9

GLUT1

SDF1

AGC CCT GCT ACA GTG TAT CCT CAA GGT CGT CGC CGT GCT G

CCG ACC CTC TTC TTT CAT CT CGT TGG CTC TGG CGA TGT GG

5, 6 1, 2

25.3 22.3

Luciferase ACT CCT CTG GAT CT CTG GTC

GTA ATC CTG AAG GCT CCT CA

n/a 28.3

Table 3.1: Primers used in real-time PCR. Gene-specific primers were used for mRNA measurement. The sequence of sense and antisense primers (5’-3’) was listed. Primers were designed to span exons to avoid the detection of endogenous DNA. The cycle threshold value for 1000 copies (Ct1000) was also listed as a reference.

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3.2.6: Plasma EPO measurement by enzyme linked immunosorbent assay (ELISA)

Blood from tail nick were collected into microtubes pre-coated with heparin. Plasma

samples were obtained by centrifugation of blood samples for 20 min at 4oC. According

to manufacturer’s instruction (Quantikine), EPO ELISA was performed on plasma

samples obtained from WT and nNOS-/- mice at baseline and 6hrs anemia.

3.2.7: Breeding and Genotyping Strategy of HIF-(ODD)-luciferase mice and nNOS-/- mice

A breeding pair of HIF-α(ODD)-luciferase mice was purchased from Jackson’s

Laboratory (23) . To genotype of HIF-α(ODD)-luciferase construct, mouse tail was

digested in Proteinase K, and DNA was extracted by phenol/chloroform methodology.

PCR was performed for 35 cycles. The PCR products were run out on 1.5% agarose gel

with ethidium bromide. Two sets of primers were used to identify the wildtype and of

HIF-α(ODD)-luciferase allele (Table 3.2). HIF-α(ODD)-luciferase negative has a PCR

product at 410bp. HIF-(ODD)-luciferase positive has a PCR product at 420bp (Figure

3.1).

To generate nNOS-/- mice with HIF-α(ODD)-luciferase construct, male

heterozygous HIF-(ODD)-luciferase positive mice were crossbred with nNOS-/- mice

(Jackson’s laboratory). To detect nNOS genotype, multiplex primers were used. Sense

primer was located at intron 1 of nNOS (Table 3.2). Two antisense primers were located

at nNOS intron 1 and PGK-neo. PCR product of 211bp corresponds to nNOS+/+, 258bp

corresponds to nNOS-/-, while bands at 211bp and 258bp corresponds to nNOS+/- (Figure

3.7). Only nNOS+/+ and nNOS-/- were used for experiments.

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Figure 3.1: HIF-ODD-luciferase and nNOS genotype by PCR and visualized on agarose gel stained with ethidium bromide.

Genotype Description Sense primer (5’-3’) Antisense primer (5’-3’)

HIF-(ODD)-

Luciferase

HIF-(ODD) Negative

GGA GCG GGA GAA ATG GAT ATG

CGT GAT CTG CAA CTC CAG TC

HIF-(ODD) Positive

CGG TAT CGT AGA GTC GAG GCC

GGT AGT GGT GGC ATT AGC AGT AG

nNOS nNOS positive (Intron 1)

CAG GGG GTT AGC TGG ACT CTT C

ACG GCT CCG ACT GTT ACA CTG

nNOS knockout (PGK-Neo)

GCT CAT TCC TCC CAC TCA TGA TCT

Table 3.2: PCR primers used for genotype of HIF-α(ODD)-luciferase and nNOS

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3.2.8: In Vivo Bioluminescent Imaging

To detect luciferase expression in HIF-α(ODD)-luciferase mice, D-luciferin

(50mg/kg; i.p.) was injected to male hemizygous mice. Ten minutes later, mice were

anesthetized in 1.5% isoflurane with 21% oxygen and placed in a light-tight chamber

equipped with IVIS imaging camera (Xenogen 300). Dorsal and ventral images of the

mouse were taken separately. Photons were collected for 10 sec, and images were

obtained by using LIVING IMAGE software (Xenogen) and IGOR image analysis

software. Images were standardized with the same colour range bar for each image

position. Total body radiance was measured in each image for quantification of luciferase

signal and normalized to the animal’s baseline value.

3.2.9: In Vitro Luciferase Assay

Hemizygous HIF-α(ODD)-luciferase mice were hemodiluted to 50g/L and

recovered for 6 hours, or exposed to hypoxia (15% O2, 6 hours). Organs were harvested

and immediately snap frozen in liquid nitrogen. Tissues were grounded with a mortal and

pestle. 400μl lysis buffer (Promega) were used with repeated freeze thaw cycle. Protein

concentration was determined using Lowery Assay (Biorad). Triplicate 10ul samples

were mixed with 100ul of Lucifearse Assay Reagent (Promega) in wells of a 96-well

plate. Luciferase activity was measured by FLUORstar Optima. Data were normalized to

the amount of protein in input sample.

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3.2.10: Biotin Switch Assay

Biotin switch assay was performed on mouse brain as previously described (11).

Briefly, brain tissues were grinded into powder using a mortal and pestle in liquid

nitrogen, and homogenized in HEN buffer (1M HEPES pH 7.7, 0.5M EDTA, 1mM

Neocuproine). Supernatant was extracted and protein concentration was measured by

BCA assay. Thiol was blocked by MMTS in blocking buffer at 50 C for 20 min with

frequent vortex. MMTS was removed by acetone precipitation at -20 C. Samples were

resuspended in HENS (HEN + 25% SDS) buffer. To biotinylate the nitrosothiols,

ascorbate and biotin-HPDP were added to the samples and incubated at room temperature

in the dark for 1 hour. Acetone was added to remove biotin-HPDP and pellets were

resuspended in HENS buffer. 2 volumes of neutralization buffer was added. To purify the

biotinylated proteins, strepavidin-agarose resin (sigma) was incubated with the samples

for 1hr at room temperature. Beads were washed with neutralizing buffer + NaCl, and

recovered with elution buffer. Sample buffer and reducing agents were added to the

biotinylated samples for Western blot analysis. In vitro nitrosylation was performed to

make positive and negative control. Brain homogenates were treated with 40uM of

GSNO (positive control) or GSH (negative control) and incubated for 20 min at room

temperature. Micro Bio-Spin P6 column (Biorad) was used to remove GSNO or GSH

using HEN buffer. These controls undergo the same condition in biotin switch assay as

described previously.

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3.2.11: Data Analysis

Statistical analysis was performed using SigmaPlot 11 (Systat Inc). Mantal-Cox

Log-Rank test was used to analyze the mortality studies. Physiological, RNA, and protein

data were analyzed independently. Data were assessed by a two-way ANOVA for time,

group, and interaction effects. When a significant interaction effect was observed, post

hoc analysis was performed using a Tukey test. All protein band densities were

normalized to the corresponding α-tubulin band density and reported as the relative to the

corresponding control. Comparison of two means was performed using a Student's t-test.

Bonferroni correction for multiple comparisons was utilized when indicated. Data are

presented as means ± SEM and a value of P < 0.05 was taken to be significant.

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3.3: Results

3.3.1: nNOS promotes HIF-1α protein expression in the brain of anemic animals

Figure 3.2 demonstrated brain nNOS and HIF-1α protein were increased 2 and 3

fold, respectively, in anemic WT mice (50g/L) at 6 and 24 hours relative to the time

control, as similar with previous studies (4). The increases in HIF-1α protein level was

not observed in the brain of nNOS-/- mice. Importantly, nNOS protein was not detected in

nNOS-/- brain homogenates, confirming that the full length nNOS protein was absent in

nNOS-deficient mice. There were no changes in HIF-2α protein expression in anemic

brain samples. α-tubulin was used to serve as loading control. These results suggested

that nNOS is required to increase HIF-1α expression in anemia.

In hypoxic (6% O2; 6hr) brain samples, HIF-1α and HIF-2α protein levels were

similarly increased in WT and nNOS-/- mice (Figure 3.3). This suggested that nNOS is

not important in HIF-α expression in hypoxic conditions. Importantly, basal HIF-α

protein levels were not different between brain samples of WT and nNOS-/- mice.

Immunohistological staining of brain section confirmed that nNOS and HIF-1α

expression were increased 24hrs after anemia (Figure 3.4). Increased HIF-1α expression

occurred at perivascular regions in anemic WT mice, but not evident in anemic nNOS-/-

mice.

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Figure 3.2: Assessment of nNOS (A), HIF-1α (B) and HIF-2α (C) protein expression in the brain of WT and nNOS-/- in 1hr, 6hr and 24hr following hemodilution. Protein level was normalized to non-anemic time control (n=6/group). *p<0.05 vs control

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Figure 3.3: Assessment of nNOS (A), HIF-1α (B) and HIF-2α (C) protein expression in the brain of WT and nNOS-/- in normoxia (21% O2) and hypoxia (6% O2 for 6hrs) (n=6 per group). *p<0.05 vs normoxia

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Figure 3.4: Immunofluorescence staining of nNOS and HIF-1α in anemic and hypoxic brain. (A) nNOS and HIF-1α staining in the cerebral cortex of WT mice 24hrs following anemia and hypoxia (n=3/group). White arrows indicated perivascular HIF-1α expression. (B) HIF-1α staining of anemic WT and nNOS-/- mouse cerebral cortex demonstrated the absence of perivascular HIF-1α staining in nNOS-/-. Scale bar = 25 μm.

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3.3.2: nNOS regulates HIF-dependent genes in anemia

Figure 3.4 demonstrated that nNOS is required to increase several HIF-dependent

genes in the brain of anemic mice. There was a time-dependent increase in steady state

mRNA transcript levels in anemic samples. HIF target genes responsible for

erythropoiesis (EPO), angiogenesis (VEGF), energy metabolism (GLUT1, PDK1) and

inflammation (SDF1, CXCR7) were all upregulated in brain samples from WT anemic

mice at 6 and/or 24 hrs following anemia. These increases were abolished in brain

homogenates of anemic nNOS-/- mice. These results suggested that nNOS is required to

regulate HIF-dependent genes in anemia.

In hypoxic brain homogenates, VEGF, GLUT1, PDK1 mRNA expressions were

increased in a statistically significant manner in WT mice (Figure 3.5). Other HIF-

dependent genes such as EPO, SDF1 and CXCR7 mRNA expressions had a tendency to

increase at 6hr hypoxia but did not reach statistically significance. In nNOS-/- mice,

VEGF mRNA expression was also increased in hypoxia, and there was a tendency that

for GLUT1 and CXCR7 to increase. Interestingly, EPO, PDK1 and SDF1 mRNA

expression did not increase in hypoxic nNOS-/- brain samples.

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Figure 3.5: Assessment of HIF-dependent mRNA expression in the brain of WT and nNOS-/- mice 1, 6, and 24hrs anemia by real-time PCR (n=6/group). *p<0.05 vs baseline

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Figure 3.6: Assessment of HIF-dependent mRNA expression in the brain of normoxic (21% O2) and hypoxic (6% O2 for 6hrs) WT and nNOS-/- mice by real-time PCR (n=6 per group). *p<0.05 vs normoxia

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3.3.3: nNOS did not influence plasma erythropoietin level in anemia

To understand whether EPO level is regulated by nNOS in response to anemia,

plasma EPO was measured in WT and nNOS-/- mice at 6hrs after hemodilution to

Hb~50g/L. Plasma EPO level increased in both WT and nNOS-/- mice (Figure 3.7),

suggesting nNOS does not play important role in the systemic EPO level primarily

produced by the kidneys. A correlation between plasma EPO and Hb concentration

demonstrated that EPO level increased similarly when Hb below 100g/L.

To determine the Hb threshold at which increase systemic EPO level occur,

WT mice were hemodiluted to a certain Hb threshold level, and plasma EPO was

assessed after 6hrs. It was found that plasma EPO level increased at Hb ~ 90g/L and

70g/L and increased further at 50g/L

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A

B

C

Figure 3.7: Plasma EPO level in 6hr anemic mice. (A) EPO level in WT and nNOS-/- anemic (Hb ~ 50g/L) mice. (B) Correlation of plasma EPO level with Hb concentration in WT and nNOS-/- mice (C) Plasma EPO level at different Hb threshold level in WT mice.

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3.3.4: HIF-α(ODD)-luciferase mice in anemia

To assess whole body HIF expression, HIF-α(ODD)-luciferase mice were used

(23). These mice contain a ubiquitously expressed ROSA26-targeted transgene driven by

a promoter that directs the transcription of a mRNA encoding a chimeric protein. The

protein contains firefly luciferase fused in frame with the 123 amino acid ODD of the

human HIF-1α subunit (amino acid 530 to 652). This ODD contains a prolyl residue of

HIF-1α that provides a cellular locus for oxygen-dependent hydroxylation and

subsequent proteosomal degradation. Thus, this chimeric protein is regulated by changes

in O2 tension similar to the native HIF-1α protein. In the absence of O2, the HIF-α(ODD)-

luciferase protein is stabilized and the luciferase enzymatic activity can be assessed by

dynamic real-time whole animal imaging.

Figure 3.8 demonstrated increased whole body luciferase signals at 6 and 24

hours following hemodilution when Hb concentration was 53 ± 3 g/L and 64 ± 3 g/L,

respectively. Increase in luciferase signals was more prominent near the kidneys and liver

regions. The signals returned to normal by day 3 and day 7 when Hb concentration was

87 ± 4 g/L and 125 ± 3 g/L, respectively.

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Figure 3.8: Whole body HIF-1α expression in anemic mice. (A) In vivo bioluminescent imaging of HIF-ODD-Luciferase mice at baseline and anemia of both dorsally and ventrally (representative from 6 mice). (B) Relationship between total body radiance and hemoglobin concentration of anemic mice up to 7 days after recovery. *p< 0.05 vs baseline

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3.3.5: nNOS is required for total body HIF-1α activity increase in acute anemia

In anemic WT mice, in vivo bioluminescent imaging of both dorsal and ventral

images demonstrated that anemia (6hrs) increases total body HIF-α luciferase expression,

most notably at the kidneys and liver regions (Figure 3.9), consistent with Figure 3.8. By

contrast, nNOS deficiency failed to increase total body HIF-α expression in anemic mice.

This suggested that nNOS is required to increase whole body HIF-1α expression in

anemia. Importantly, there was no significant different in basal total body HIF-α

expression between WT and nNOS-/- mice.

When WT and nNOS-/- mice were subjected to hypoxia (6% O2) for 6hrs, they

demonstrated increased HIF-α luciferase expression in whole body (Figure 3.10). This

suggested that nNOS did not impact whole body HIF-1α expression in hypoxia.

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Figure 3.9: Representative images from in vivo bioluminescent imaging of anemic WT and nNOS-/- mice in dorsal (A) and ventral (B) views. Total body radiance (represents whole body HIF expression) is plotted in the graphs on the right (n=6-8 per group). *p<0.05 vs baseline; #p<0.05 vs WT

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Figure 3.10: In vivo bioluminescent imaging of hypoxic (6% O2 for 6hrs) WT and nNOS-/- mice in dorsal (A) and ventral (B) views. Total body radiance is plotted in the graphs on the right (n=6 mice/ group). *p<0.05 vs baseline; #p<0.05 vs WT

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3.3.6: nNOS is required to increase HIF-1α in the anemic brain

Since the skull blocks the detection of luciferase signals, in vitro luciferase assay

was performed in the brain and other organs (kidneys, heart and liver) to quantitatively

assess luciferase activity. In anemia, the increase in HIF-α luciferase expression was

demonstrated in the brain, kidney, liver, and heart 6hrs after anemia in WT mice (Figure

3.11). Consistent with previous protein and RNA data, HIF-1α luciferase expression did

not increase in the brain of anemic nNOS-/- mice. Interestingly, HIF-1α luciferase

expression in the heart and kidney were increased in anemic nNOS-/- mice. Finally, basal

liver HIF-α levels seem higher in nNOS-/-, and it was not increased further with anemia.

Unfortunately, HIF-1α luciferase expression was not measured in other tissues (eg. gut,

skin, skeletal muscle, smooth muscle, etc). The HIF levels in these untested tissues may

contribute to the abolished whole body HIF expression in anemic nNOS-/- mice.

In hypoxic WT and nNOS-/- mice, HIF-1α luciferase expression increased in all

organs studied (brain, heart, kidney, liver) (Figure 3.12). This suggested that nNOS is not

essential to the hypoxic induction of HIF-1α. Interestingly, the increase in HIF-1α was

smaller in the liver of hypoxic nNOS-/- mice (Figure 3.12), suggesting nNOS deficiency

may have an impact in HIF-1α induction.

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Figure 3.11: In vitro tissue luciferase measurement of the tissue luciferase activity at baseline and anemia in the brain (A), kidney (B), heart (C), and liver (D). Data were normalized to protein concentration (n=6 / group). *p<0.05 vs baseline; #p<0.05 vs WT.

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Figure 3.12: In vitro tissue luciferase measurement of the tissue luciferase activity at normoxia (21% O2) and hypoxia (6% O2) in the brain (A), kidney (B), heart (C), and liver (D). Data were normalized to protein concentration (n=5-6 mice per group). *p<0.05 vs baseline; #p<0.05 vs WT.

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3.3.7: Whole body and tissue HIF luciferase activity at different Hb threshold level

To determine at which Hb level HIF is activated, WT HIFα-(ODD) Luciferase

mice were hemodiluted to different Hb threshold and assessed after 6hrs. Increased whole

body HIF luciferase was only detected at Hb ~50g/L (Figure 3.13). Interestingly,

different organs have unique patterns for the increase in HIF luciferase levels. For

example, brain HIF levels reduced at 90g/L, returned to baseline at 70g/L and increased

further at 50g/L. HIF levels in kidneys did not increase until Hb reached 50g/L, whereas

liver HIF levels increased significantly at 70g/L and 50g/L.

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Figure 3.13: HIF-luciferase activity in whole body and different organs at various hemoglobin threshold 6hrs after hemodilution.

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3.3.8: nNOS leads to S-nitrosylation of pVHL may be responsible for HIF-1α

stabilization in anemia

To determine the mechanism by which HIF-1α protein in the brain can be

stabilized in acute anemia, assessment of S-nitrosylated proteins were carried out using

biotin switch assay. We found that S-nitrosylated pVHL protein level was increased in 6

and 24hrs anemic WT mouse brain (Figure 3.14). Additionally, there was no change in

SNO-pVHL levels in anemic nNOS-/- brain. These changes were not observed in hypoxic

brain. Furthermore, anemia did not induce increases in S-nitrosylation to other proteins

known to be S-nitrosylated, such as HIF-1α, GAPDH and creatine phosphokinase (CPK)

(Figure 3.15). These results suggested that nNOS-dependent S-nitrosylation of pVHL can

be one of the mechanisms to stabilize HIF-1α protein in acute anemia.

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Figure 3.14: S-nitrosylated (SNO) pVHL level is increased in acute anemic brain in nNOS-dependent fashion. [A] SNO-pVHL level in control (c), 1, 6, 24 hr anemic brain as assessed by performing biotin switch assay (n=4/group). [B] Assessment of SNO-pVHL levels in WT and nNOS-/- (KO) brain samples in 6hrs anemia and hypoxia (n=5/group). Total pVHL levels were not changed. GSH and GSNO serve as negative and positive control, respectively, to biotin switch assay. Data was normalized to control and expressed as mean ± SEM. Ctrl denotes control; A denotes 6hr anemia; Norm denotes normoxia; Hyp denotes 6hr hypoxia.. *p<0.05 vs control.

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Figure 3.15: Changes in other SNO-protein levels were not detected in anemic brain samples. (A) S-nitrosylated creatine phosphokinase (SNO-CPK) and glyceraldehyde pyruvate dehydrogenase (SNO-GAPDH) levels were measured in anemic brains samples. No changes were found in total CPK and GAPDH protein levels. (B) SNO-HIF-1a was assessed in anemic and hypoxic (6hr) samples. Same concentration of protein was used in biotin switch assay, which denotes input. Data was normalized to control and expressed as mean ± SEM.

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3.3.9: Anemia reduces the level of GSNO reductase To determine whether denitrosylation status was affected by acute anemia,

GSNOR protein level was assessed in brain samples. In acute anemia, GSNOR protein

was reduced as early as 1hr and persisted until 24hrs (Figure 3.16). In acute hypoxia,

GSNOR protein was decreased at 6hrs, and returned to baseline in 24hrs. Earlier reports

demonstrated that GSNOR may promote heart HIF-1α expression in a myocardial

infarction model (18). To determine whether GSNOR plays a role in anemia, GSNOR-/-

mice were hemodiluted until mortality. There was no difference in anemia-induced

mortality relative to WT mice, despite a trend for survival advantage in GSNOR-/- mice.

These results demonstrated that anemia suppresses a de-nitrosylation pathway, which

may be an important mechanism for increased SNO-protein to promote survival.

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A) B) Figure 3.16: (A) GSNOR protein levels in acute anemia and hypoxia. (B) Mortality curve of WT and GSNOR-/- mice during acute anemia

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3.4: Summary of main findings:

The novel finding in this research demonstrated that nNOS is critically important

in HIF-1α protein stabilization in anemic brain despite a comparable decrease in brain

PO2 between anemic WT and nNOS-/- mice. First, nNOS is required for the increase in

brain HIF-1α protein levels in anemia as demonstrated that nNOS-/- mice cannot increase

HIF-1α protein in vivo. Second, nNOS is needed to increase HIF-dependent mRNA

transcripts, such as EPO, VEGF, and GLUT1, to optimize tissue oxygen delivery in

response to anemia. Third, the ability of nNOS to stabilize HIF-1α in anemia appears to

be specific to the brain. This is demonstrated by the similar response between anemic WT

and nNOS-/- mice in systemic EPO levels and the in vivo increase of HIF-1α luciferase

expression in the heart, kidney, and liver. Fourth, the Hb threshold for increase in HIF-1α

occurred in tissue-specific manner such that brain HIF increased at lower Hb level

whereas liver HIF increased at higher Hb level. Fifth, the mechanism by which nNOS

stabilizes HIF-1α protein in anemia can be due to increased nNOS-dependent S-

nitrosylation of pVHL, which may help HIF-1α protein to escape the degradation

pathway. Finally, anemia reduces brain GSNOR protein levels, and may suppress the de-

nitrosylation pathway to increase SNO protein. This may be adaptive as demonstrated

that anemic GSNOR-/- mice had a tendency to increase survival relative to WT mice.

Unlike anemia, nNOS did not have an impact on increased brain HIF-1α protein

expression and HIF-dependent mRNA transcripts in hypoxia. Also, the increase in HIF-

1α luciferase expression in other organs (heart, kidney, liver) did not depend on nNOS.

Overall, these results highlight the differences in cellular responses between anemia and

hypoxia.

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3.5. Reference:

1. Pfeffer MA, et al. (2009) A trial of darbepoetin alfa in type 2 diabetes and chronic

kidney disease. N Engl J Med 361(21):2019-2032. 2. Bennett CL, et al. (2008) Venous thromboembolism and mortality associated with

recombinant erythropoietin and darbepoetin administration for the treatment of cancer-associated anemia. JAMA 299(8):914-924.

3. Hebert PC, et al. (1999) A multicenter, randomized, controlled clinical trial of transfusion requirements in critical care. Transfusion Requirements in Critical Care Investigators, Canadian Critical Care Trials Group. N.Engl.J.Med. 340(6):409-417.

4. McLaren AT, et al. (2007) Increased expression of HIF-1alpha, nNOS, and VEGF in the cerebral cortex of anemic rats. Am J Physiol Regul Integr Comp Physiol 292(1):R403-414.

5. Hare GM, et al. (2003) Hemodilutional anemia is associated with increased cerebral neuronal nitric oxide synthase gene expression. J Appl Physiol 94(5):2058-2067.

6. Gruber M, et al. (2007) Acute postnatal ablation of Hif-2alpha results in anemia. Proc.Natl.Acad.Sci.U.S.A. 104(7):2301-2306.

7. Cheng J, Kang X, Zhang S, & Yeh ET (2007) SUMO-specific protease 1 is essential for stabilization of HIF1alpha during hypoxia. Cell 131(3):584-595.

8. Nathan C & Xie QW (1994) Nitric oxide synthases: roles, tolls, and controls. Cell 78(6):915-918.

9. Boulanger CM, et al. (1998) Neuronal nitric oxide synthase is expressed in rat vascular smooth muscle cells: activation by angiotensin II in hypertension. Circ Res 83(12):1271-1278.

10. Cervenka L, et al. (2002) Role of nNOS in regulation of renal function in hypertensive Ren-2 transgenic rats. Physiol Res 51(6):571-580.

11. Jaffrey SR, Erdjument-Bromage H, Ferris CD, Tempst P, & Snyder SH (2001) Protein S-nitrosylation: a physiological signal for neuronal nitric oxide. Nat.Cell Biol. 3(2):193-197.

12. Jia L, Bonaventura C, Bonaventura J, & Stamler JS (1996) S-nitrosohaemoglobin: a dynamic activity of blood involved in vascular control. Nature 380(6571):221-226.

13. Lipton AJ, et al. (2001) S-nitrosothiols signal the ventilatory response to hypoxia. Nature. 413(6852):171-174.

14. Metzen E, Zhou J, Jelkmann W, Fandrey J, & Brune B (2003) Nitric oxide impairs normoxic degradation of HIF-1alpha by inhibition of prolyl hydroxylases. Mol.Biol.Cell. 14(8):3470-3481.

15. Berchner-Pfannschmidt U, Yamac H, Trinidad B, & Fandrey J (2007) Nitric oxide modulates oxygen sensing by hypoxia-inducible factor 1-dependent induction of prolyl hydroxylase 2. J Biol.Chem. 282(3):1788-1796.

16. Palmer LA, et al. (2007) S-nitrosothiols signal hypoxia-mimetic vascular pathology. J Clin Invest 117(9):2592-2601.

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158

17. Li F, et al. (2007) Regulation of HIF-1alpha stability through S-nitrosylation. Mol.Cell. 26(1):63-74.

18. Lima B, et al. (2009) Endogenous S-nitrosothiols protect against myocardial injury. Proc Natl Acad Sci U S A 106(15):6297-6302.

19. Sumbayev VV, Budde A, Zhou J, & Brune B (2003) HIF-1 alpha protein as a target for S-nitrosation. FEBS Lett 535(1-3):106-112.

20. Hagen T, Taylor CT, Lam F, & Moncada S (2003) Redistribution of intracellular oxygen in hypoxia by nitric oxide: effect on HIF1alpha. Science 302(5652):1975-1978.

21. Schleicher M, et al. (2009) The Akt1-eNOS axis illustrates the specificity of kinase-substrate relationships in vivo. Sci Signal 2(82):ra41.

22. Ward ME, et al. (2005) Hypoxia induces a functionally significant and translationally efficient neuronal NO synthase mRNA variant. J Clin Invest 115(11):3128-3139.

23. Safran M, et al. (2006) Mouse model for noninvasive imaging of HIF prolyl hydroxylase activity: assessment of an oral agent that stimulates erythropoietin production. Proc.Natl.Acad.Sci.U.S.A. 103(1):105-110.

Chapter 4

Chapter 4:

Discussion and future direction

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Chapter 4

My research has demonstrated several novel fundamental findings which illustrate

that nNOS is critically important to the integrative physiological and cellular response to

acute anemia. First, nNOS is protective during anemia as demonstrated by the increased

mortality in anemic nNOS-/- mice. Whereas nNOS replete mice have a mean lethal Hb

concentration that is comparable to human (~25g/L) (1), nNOS -/- mice died at a higher

lethal Hb concentration (~35g/L). Second, anemic nNOS-/- mice cannot regulate their CO

or SVR in a manner that optimizes tissue perfusion. Acute changes in cardiovascular

function require NO mediated protein modifications (S-nitrosothiols) to improve

myocardial blood flow and optimize cardiac function (2). Thus, in the absence of nNOS,

anemic mice may have died because of inadequate SNO protein modification, which may

be responsible for cardiovascular adaptation.

Despite these differences in cardiovascular function, surprisingly, cerebral tissue

PO2 decreases comparably in anemic mice regardless of nNOS genotype. At comparable

reduction in tissue PO2, the cellular response to anemia was very different. Anemia led to

an increase in HIF-1α protein level and expression of HIF-dependent mRNA transcripts

in the brain of WT mice. These cellular responses may support cytoprotective,

angiogenic, and/or metabolic adaptation to optimize tissue oxygen delivery and

utilization in acute anemia. In marked contrast, anemic nNOS-/- mice did not increase

brain HIF-1α protein levels or HIF-dependent mRNA transcripts. The lack of the HIF

response to anemia may have impaired the ability of nNOS-/- mice to adapt to the anemic

conditions. To confirm that these changes were biologically relevant, we utilized real-

time imaging of transgenic animals that expressed a HIF-1α luciferase chimeric protein.

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Using this model, we confirmed that nNOS-derived NO is key for HIF-1α stabilization

during anemia in a whole animal model. The mechanism by which HIF-1α stabilization

occurred in anemia included nNOS-mediated S-nitrosylation of pVHL to generate SNO-

pVHL as detected by the biotin switch assay. Previous studies have demonstrated that S-

nitrosylation of pVHL cannot interact with HIF-α in vitro (3), thus allowing HIF-1α to

escape degradation. For the first time, we have demonstrated that this phenomenon also

occurred in vivo during anemia. The mechanism appeared to be specific to pVHL during

anemia, as other S-nitrosylation target proteins did not demonstrate increase in S-

nitrosothiol (SNO) modification. In addition to the effect of nNOS-mediated SNO-

protein modification, we assess the importance of denitrosylation reactions by measuring

GSNOR protein levels. Anemia led to a decrease in GSNOR protein levels in the brain,

possibly contributing to maintain high S-nitrosothiols levels. The combination of

increased S-nitrosylation and reduced denitrosylation may favor the kinetics of prolonged

SNO protein modification during anemia. These changes in SNO proteins likely mediate

important adaptive physiological and cellular responses to anemia.

In contrast to the clear protective role of nNOS during anemia, we were surprised

to demonstrate a divergent phenotype in nNOS-/- mice exposed to acute hypoxia. Striking

differences between anemia and hypoxia occurred in nNOS-/- mice. First, nNOS-/- mice

died earlier during anemia but survived longer during hypoxia. Second, nNOS-/- mice did

not regulate CO and SVR during anemia but were able to do so after exposure to acute

hypoxia. Third, nNOS-/- mice did not demonstrate evidence of HIF-1α protein

stabilization during anemia but nNOS genotype did not influence the strong HIF response

to hypoxia. Finally, S-nitrosylation of pVHL contributed to HIF-1α stabilization during

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anemia but was not required for HIF-1α stabilization during hypoxia. Thus, assessment of

the phenotypic responses to anemia and hypoxia in nNOS-/- mice demonstrated

completely divergent patterns, supporting the previously under-appreciated concept that

anemia and hypoxia are different. In other words, these findings provide essential

mechanistic insight into the differences between anemia and hypoxia.

From an evolutionary perspective, adaptive physiological and cellular responses

must have enabled mammals to tolerate and survive episodes of acute and severe blood

loss. As such, human physiology is particularly well adapted to support the associated

reduction in Hb concentrations and O2 carrying capacity. For example, humans can

survive extremely low Hb concentration (Hb ~ 7 g/L) if supported by maximal medical

therapy (4). Furthermore, examples of battlefield trauma demonstrated survival under

adverse conditions of extremely low Hb (5). Despite these remarkable examples of

humans’ survival at low Hb, there may be a significant degree of survivor bias in these

reported cases since the denominator (patients who died from low Hb) may be unknown

or unreported. This brings us back to a review of the literature defining of the impact of

low Hb on mortality in surgical patients. Perhaps the best patient population to help

answer this question includes patients undergoing surgical procedures who will not

accept blood transfusion therapy (i.e. Jehovah’s Witness patients) (1, 6-8). Studies in

these patients demonstrate a progressive increase in mortality with progressive blood loss

and fluid resuscitation (hemodilution). This study estimates a 2.1-fold increase (95% CI,

1.7-2.6) mortality for every 10g/L decrease in Hb (1). In addition, patients undergoing

heart surgery demonstrate an increase in mortality when their pre-operative hemoglobin

drops below 120g/L (9, 10) and their inter-operative Hb < 70g/L during cardiopulmonary

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bypass (11). Finally, a newer retrospective study suggested even mild to moderate anemia

poses increased risk of mortality in surgical patients (12). These examples outline the

importance of anemia as a risk factor for increased mortality in a number of different

patient populations. The magnitude of anemia-induced mortality may be far greater as

anemia is a predictor of adverse outcome in a broad outcome of clinical conditions,

including chronic heart failure (13), subarachnoid hemorrhage (14), neurotrauma (15),

and acute coronary syndrome (16). Therefore, an understanding of the mechanism of

anemia-induced mortality is required.

Mechanisms of survival in anemia

During anemia, a proportionate increase in CO and reduced SVR acts to maintain

a global balance of O2 delivery and consumption (17, 18). The mechanisms by which this

response occur are multiple and likely include: 1) sensing of reduced O2 delivery at the

tissue level (19, 20); 2) activation of the sympathetic nervous system (20, 21); 3) a

coordinated increase in CO and reduced SVR by direct (local tissue hypoxia) and remote

(perivascular innervations) mechanisms (22, 23); 4) local intrinsic vascular responses

(24); and 5) regulation of microvascular tone by circulating mediators (SNO-Hb, nitrite)

(25, 26). The composite response is to promote O2 delivery to vital tissues (27-29).

Traditional physiological mechanisms of survival during anemia

As reviewed in the Introduction (in 1.2.3), the traditional view of the

physiological adaptation to anemia is demonstrated in numerous studies beginning as

early as 1940s (30). Collectively, these studies demonstrated that anemia is sensed by the

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aortic chemoreceptor to activate the sympathetic nervous system which regulates increase

in CO, SV and HR (18-21, 29). These responses are activated to increase CO and maintain

MAP. Reduction in SVR by intrinsic and extrinsic mechanisms promotes further O2

delivery to tissues (27-29). To ensure that adequate O2 is delivered to vital organs,

redirection of blood flow by organ-specific vasodilation occurs. This response permits

organs with high metabolic rate for O2 (e.g. brain and heart) to receive a disproportional

increased in blood flow relative to increase CO, whereas lower blood flow is directed to

other “less” vital organs (e.g. kidney, liver, intestine) (31-33). The resultant effect leads

to maintained tissue oxygenation at the brain and heart, while reduction in tissue PO2 in

organs below the diaphragm. The overall pattern suggested that organs of the greatest

importance for survival would preferentially receive the limited blood O2 content (17). In

addition, organ specific changes in O2 consumption in anemia may help to balance

oxygen demand with its reduced supply (34-36). These mechanisms deal largely with

optimizing tissue O2 delivery. They do not fully explain adaptation to optimize O2

utilization at the cellular level. Therefore, a more complete understanding of O2

homeostasis is required.

nNOS-dependent mechanisms of survival during anemia

The traditional view of the physiological mechanisms to anemia only considers

responses in the context of optimizing O2 delivery. To further understand the integrated

physiology in response to anemia, we also need to consider from the perspective of the

adaptive cellular responses and the regulation of O2 homeostasis. In this regard, NO plays

a central role in regulating physiological responses to changes in O2 level (37). In

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particular, nNOS is well suited in the regulatory response to anemia because: 1) nNOS

confers protection in anemic mice; 2) nNOS protein expression is increased in animal

models during anemia (24, 29, 38); 3) relative to eNOS and iNOS, the high requirement

of nNOS activity for O2 makes this NOS isoform able to function during anemia where

PO2 values are maintained (39); and 4) nNOS is implicated in a number of important

cardiovascular responses (40-42).

Brain

nNOS facilitates synaptic transmission and controls neuronal regulation of HR

and MAP (43, 44). It also can regulate neurovascular coupling and control of cerebral

blood flow in specific regions of the brain in defined experimental conditions (45, 46).

During anemia, increased cerebral blood flow is an adaptive response to optimize brain

tissue PO2 (29, 31), which could be regulated by nNOS (45). Surprisingly in our anemia

model, none of these parameters was influenced in nNOS-/- mice, as we demonstrated a

small decrease in brain PO2 despite an increase carotid blood flow in anemia that were

comparable between anemic nNOS replete and deplete mice. Expectedly, redundant

mechanisms must exist to ensure adequate brain tissue oxygenation in anemia despite the

loss of nNOS. In this regard, the central nervous system is considered a key component

since it controls all systemic functions.

At this maintained brain PO2 level in anemia, an additional mechanism may be

required to activate hypoxia signaling. The existence of the HIF pathway which

originated near 600 million years ago is a primitive response to changes in O2 level which

is conserved across species through evolution (47). This pathway may be more important

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to survival in species where their behaviors and living habitats experience large

fluctuation in O2 levels (e.g. fish). However, mammals exposed to higher PO2 may have

evolved to have a HIF pathway that had been modified to respond to a small change in

PO2, such as anemia. Thus, a cellular response of nNOS mediated SNO-pVHL to

stabilize HIF-1α may be beneficial to a small reduction in PO2 in anemic mammals.

The activation of gene transcription through the HIF pathway may be a sustained

adaptive response to increase survival in anemia. For example, increased VEGF mRNA

expression in the neurons and astrocytes (24, 48) may promote neuronal survival,

angiogenesis (49-51), and capillary density to favor optimal oxygen delivery to the brain

in anemia (52). Evidence of increased VEGF expression and enhanced angiogenesis has

also been demonstrated in response to anemia (53, 54). Therefore, the increase in VEGF

expression observed in the brain of anemic WT mice, but not nNOS-/- mice, may

represent activation of proangiogenic and/or neuroprotective mechanisms directed at

optimizing cerebral oxygen delivery and neuronal viability. Conversely, increased VEGF

may have deleterious effects, including increased vascular permeability as demonstrated

during ischemia (55, 56). Further studies are required to assess the adaptive/maladaptive

role of VEGF in the anemic brain.

To survive in low O2 condition in which substrates are deprived, cells must be

able to utilize existing resources in an efficient manner. Increased metabolic pathway

(e.g. GLUT1, PDK1) is one way to optimize cellular oxygen utilization in anemia.

GLUT1, selectively localized to the blood brain barrier (BBB), facilitates glucose

transport from the blood to the brain across BBB for metabolism (57). Increased PDK1

can reduce mitochondrial oxygen consumption by shunting pyruvate away from the

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mitochondria and maintain ATP production (58, 59). The resultant effects lead to

increased glycolysis, while inhibiting fatty acid oxidation. Interestingly,

phosphofructokinase is physically associated with nNOS and may exert neuroprotection,

but its importance is not clear (60). In anemia, nNOS-mediated metabolic responses may

help to increase glucose flux, and block oxygen utilization which results in increase

oxygen availability.

A hallmark response to anemia is increased EPO level. In the brain, EPO has a

non-hematopoietic function which may contribute to neuroprotective effects in anemia

(61, 62). Neuroprotection of EPO may occur by the binding of EPO to neuronal EPO

receptor which activates JAK2, which can then activate other signaling pathway such as

PI3K and STAT5 (61, 63). This response may suppress cell death by promoting anti-

apoptotic genes and/or inhibiting caspases (64). Recently, a study demonstrated that

nNOS-derived NO induces EPO via HIF-1 pathway to prevent axonal degeneration in

glial cells (65). Collectively, these studies suggested that nNOS may protect against

neuronal cell damage by increased brain EPO levels in anemia.

Cardiac response in anemia

To maintain adequate tissue perfusion during anemia, the heart is a central organ

that requires special attention. A characteristic feature of the heart is that it is the only

organ in the body with high basal O2 extraction rate (~70%), which suggests that it must

rely heavily on increase in blood flow to increase metabolic demand in response to

anemia. For example, myocardial O2 consumption can be tripled while coronary blood

flow can increase up to six-fold during anemia (17, 32, 36). This provides the extra

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amount of O2 supply to accommodate for the higher metabolic requirement to increase

CO during anemia. The ability to increase CO in anemia is vital to maintain organ

oxygenation during anemia, as experimental studies demonstrated that abolished HR and

CO effects by β-blockade leads to severe tissue hypoxia (29). Other than increase O2

delivery to enhance heart function, cellular mechanisms may also play an important role

to modulate cardiac performance.

In this regard, a direct impact of nNOS can affect contraction and relaxation of

cardiac myocytes by regulating intracellular calcium levels (40). For instance, nNOS

deletion has been associated with increased L-type Ca2+ current, decreased

phospholamban phosphorylation and enhanced myocyte shortening (40-42). In addition,

nNOS-derived NO can undergo S-nitrosylation of proteins involved in Ca2+ cycling (L-

type Ca2+ channel, ryanodine receptor channel, SERCA) (66-68), which may lead to

increased (SERCA) (69) or decreased (L-type Ca2+ and ryanodine receptor channel)

(69, 70) activity. The resultant effect of nNOS deletion leads to impaired relaxation of

heart. This would correlate with diastolic dysfunction and inability of the myocardium to

relax during anemia. This mechanism could explain the lack of an increase in diastolic

volume, SV and CO in anemic nNOS-/- mice. However, CO data obtained in hypoxic

nNOS-/- mice refute this argument. During hypoxia, there is an unexpected increase in SV

and CO in nNOS-/- mice. This suggests that the cardiomyocyte of these mice retained the

ability to relax. Thus additional cardiovascular mechanisms must be explored.

the

In condition of a decreased myocardial O2 delivery, such as in anemia, the heart

must adapt quickly and find alternate ways to efficiently use the limited resources. In this

limited O2 environment, a metabolic switch away from fatty acid oxidation to glycolysis

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is favored. This response would reduce cellular O2 consumption without compromising

ATP production. Activation of nNOS-dependent, HIF-1α-mediated PDK1 expression in

anemia can tip the balance from oxidative to glycolytic metabolism by shunting pyruvate

away from the mitochondria (58, 59). In addition, increased GLUT1 expression and S-

nitrosylation of a number of glycolytic enzymes (hexokinase, enolase, creatine kinase,

GAPDH) in cardiomyocytes may facilitate glucose transport and optimize glycolytic

utilization the heart during anemia (71). The combined effects could lead to increased

glucose metabolism to lactate, which generates ATP by minimizing O2 consumption.

Although we have not measured these metabolic enzymes in the heart, we have evidence

of increased GLUT1 and PDK1 mRNA transcripts in the brain of anemic WT but not

nNOS-/- mice. These responses suggested that a nNOS-dependent metabolic switch to

glycolysis occurred in anemia, which may also occur in the heart to sustain cardiac

function. In this way, improved cardiac function by optimizing myocardial O2 delivery

and consumption in nNOS replete mice may enhance its survival in anemia.

Response of resistance artery in anemia

The understanding of resistance artery function in a whole animal model is

complex because SVR is derived from measurements of CO and MAP. For example, the

extremely well characterized and proportional increase in CO during progressive anemia

occurs with an overall maintenance of MAP meaning that calculated SVR must decrease.

In a whole animal model, it is difficult to determine which is the primary driving force of

this response. Inhibition of CO by denervation or β-blockade prevents both the increase

in CO and associated reduction in SVR during anemia (21, 29). Alternatively, Guyton et

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al would argue that anemia causes a “passive” reduction in SVR which leads to an

increase in venous filling of the heart (preload) and by the Starling relationship a

subsequent increase in CO (22). Both of these arguments are circular and do not allow for

the independent assessment of cardiac and vascular components. Therefore, we assessed

the impact of anemia on vascular tone in isolated resistance arteries from WT and nNOS

deficient mice. Although we expected to see a reduction in tone during anemia in nNOS

replete vessels, no difference was observed between the tone in anemic nNOS-/- mice. If

anything, the anemic vessels appear to have more tone, thus an intrinsic vascular

mechanism does not explain our results. This may be due to the fact that that mesenteric

beds are not the best experimental model, and that cerebral mesenteric arteries may be

preferred. Of interest, Ward et al demonstrated that hypoxia did indeed upregulate nNOS

in vascular smooth muscle cells (37). Thus by comparison the vascular response to

anemia and hypoxia is different.

Respiratory response to anemia

Anemia is sensed by the aortic chemoreceptors and triggered immediate

respiratory response to optimize O2 delivery. The characteristic increase in PaO2 and SaO2

during anemia suggested that improved ventilation/perfusion matching occurs in the

lungs. This response is, in part, due to eNOS-dependent vasodilation of pulmonary

vessels (72). In addition, centrally controlled respiration can occur via SNO which

involves the transfer of SNO signal from blood to brainstem (73). The mechanism occurs

by which SNO is transferred from SNO-Hb to glutathione, which can then

transnitrosylate to target proteins at the brainstem to regulate breathing. The deletion of a

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key enzyme (γ-glutamyl transpeptidase) involved in SNO transfer inhibits ventilation

supports that SNO indeed stimulates central respiration. Despite studies which

demonstrated that NO is important in controlling respiration, there is a lack of evidence

to support a role for nNOS in regulating adaptive respiratory responses in anemia. Our

results failed to show an impact for nNOS on the blood gases during anemia, as changes

in PaO2 and PaCO2 were similar regardless of nNOS genotype. Although it seems

reasonable that a role for increase nNOS in the brain during anemia may affect SNO

levels to control respiration in a similar manner, currently no clear experimental evidence

has identified the source of SNO for this mechanism. For these reasons, activation of

respiratory response in anemia may not be dependent on nNOS. Thus, nNOS may be

more important in mediating other adaptive responses, such as the cardiovascular system,

to protect mice during anemia.

Stabilization of HIF-1α during anemia by non- hypoxic and hypoxic mechanisms

The traditional view of HIF-α regulation includes transcriptional (non-hypoxic)

(74, 75), and post-translational mechanisms (PHD/pVHL) (76). In our anemia model, we

observed HIF-1α protein increased in the brain during anemia but mRNA level for HIF-

1α did not increase (24). This suggested that post-translational mechanism of HIF-1α is

important in anemia. Our data supported the possibility that during anemia, HIF-1α is

stabilized by traditional reduction in tissue PO2 and subsequent inhibition of PHD activity

in some organs (kidney, liver). However, in the brain, we observed a pattern of HIF-1α

stabilization that was more dependent on nNOS than changes in tissue PO2. To expand

upon this, in the brain of WT and nNOS-/- mice, there was a comparable, but small drop

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in tissue PO2 (baseline ~70mmHg to anemic ~40mmHg). Although this small change in

PO2 might be expected to result in HIF-1α stabilization, our data suggest another

explanation. In this tissue condition, nNOS is critical to HIF-1α stabilization. In other

words, our Western blots and mRNA data demonstrate that HIF-1α protein and mRNA

responses only increase in response to this small drop in PO2 only if nNOS is present and

active. The ability of low PO2 to stabilize HIF-1α independent of nNOS was

demonstrated in our animals exposed to hypoxia. Ironically, at a comparable small

reduction in tissue PO2, both WT and nNOS-/- mice demonstrated an increase in HIF-1α

after hypoxia exposure. These data demonstrate that a comparable reduction in PO2 by

anemia and hypoxia have different biological effects in the brain. The importance of

nNOS in the brain during anemia versus hypoxia may be due to the relatively high PaO2

in the vasculature of anemic mice (Figure 4.1). The importance of these findings is

emphasized at a systemic level in vivo by demonstrating the same differential effect of

nNOS on HIF-1α stabilization during anemia but not hypoxia in HIF-1α (ODD)-

luciferase mice. In conclusion, in the brain of anemic mice, the presence of nNOS

determines whether HIF-1α will be stabilized or not. In our model, this occurs

independent of PO2 differences, suggesting non-hypoxic mechanisms (Figure 4.2).

The importance of non-hypoxic nNOS mediated HIF-1α stabilization appears to

be specific to the brain because we observed strikingly different result with respect to the

kidney. Although we did not measure kidney tissue PO2, our lab and others have

demonstrated a much larger decrease in tissue PO2 during anemia relative to the brain

(29, 77). However, we measured systemic EPO as a surrogate of tissue hypoxia during

anemia. Of interest, there was a comparable Hb threshold for increased systemic EPO

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level in WT and nNOS-/- mice. With respect to HIF-1α stabilization, unlike the brain, the

kidney demonstrated an increase in HIF-1α protein levels in nNOS-/- mice relative to WT.

This may reflect the attenuation in cardiac output and reduction in global O2 delivery in

nNOS-/- mice that did not seem to affect cerebral perfusion, but may have profoundly

affected perfusion in other organs like kidney and liver. During hypoxia, again,

comparable increases in HIF-1α were observed in both nNOS replete and deplete mice.

In other words, nNOS phenotype did not negatively affect HIF-1α expression in the

kidney during hypoxia. Indeed, the lack of nNOS was associated with higher HIF-1α

expression during anemia in nNOS-/- mice. Therefore, SNO modification of pVHL,

although not measured, was likely not responsible. Thus, in the kidney of anemic mice, it

appears that HIF-1α stabilization is not dependent on nNOS, but dependent on PO2.

The Hb threshold data further emphasize HIF-1α stabilization during anemia. We

assessed anemic mice 6hrs after hemodilution at three different Hb thresholds (90g/L,

70g/L and 50g/L). Whole body luciferase activity demonstrated a stepwise increase in

HIF-1α luciferase signal with each progressive drop in Hb concentration. However, there

was a different HIF-1α pattern at the organ level. At Hb of 90g/L, brain HIF-1α

decreased, while kidney and liver HIF-1α remained stable. At 70g/L, brain HIF-1α

returned to baseline, kidney HIF-1α remained stable, while liver HIF-1α increased. At

50g/L, an increase in brain, kidney, and liver HIF-1α level was observed. These organ-

specific HIF-1α patterns may reflect the relative contribution between nNOS and tissue

PO2 at the different Hb levels. Assessment of the tissue PO2 at these Hb thresholds will

be useful to delineate the non-hypoxic or hypoxic mechanism of organ-specific HIF-1α

stabilization.

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Since O2 is a substrate for nNOS enzymatic activity, different tissue PO2 levels

may explain the observation that nNOS is critically required for HIF-1α stabilization in

the brain but not kidney. Of note, since nNOS is highly abundant in the brain, it may have

a higher functional importance compared to the kidneys in which nNOS expression is

lower (78). Also, biochemical studies revealed that nNOS enzymatic activity depends

heavily on O2 (apparent nNOS Km for O2 of 350µM) (39). This suggested that relatively

“higher” PO2 in the brain during anemia may create a favorable condition to increase

nNOS activity such that it is required for HIF-1α stabilization in this O2 condition.

However, the “lower” PO2 in the kidney may lead to inhibition of nNOS activity. Thus,

increase kidney HIF-1α expression is likely due to tissue hypoxia, but not nNOS. This

nNOS-independent, hypoxic stabilization of HIF-1α in the kidney is further supported by

our data that an increase in HIF-1α protein level is comparable in all tissues in hypoxic

WT and nNOS-/- mice. Therefore, nNOS is critically important for normal enzymatic

activity only in favorable PO2 conditions to stabilize HIF-1α level, such as in the brain

during anemia (non-hypoxic). At low tissue PO2, nNOS activity is not favored, thus HIF-

1α is stabilized by hypoxic mechanisms (i.e. inhibition of PHDs) (Figure 4.1).

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Figure 4.1: Non-hypoxic and hypoxic HIF-1α stabilization in acute anemia and systemic hypoxia. In anemia, high PaO2 and SaO2 results in high O2 gradient from lumen to tissue. The extraction of O2 from Hb allows SNO-Hb to be exposed and contribute to NO pool. At the tissue and vascular smooth muscle (VSM) compartments, increased nNOS activity (favourable O2 environment) results in a gradient of NO from the tissue to lumen. nNOS-derived NO may increase HIF-1α stabilization in non-hypoxic mechanism. In systemic hypoxia, low PaO2 and SaO2 reduced the O2 gradient from lumen to tissue. Also, SNO-Hb may be depleted due to low SaO2 (i.e. Hb in T structure and SNO cannot be loaded on Hb). The combined effects may result in reduced tissue O2 extraction (gradient). Because nNOS activity is inhibited in low O2 environment, HIF-1α stabilization in hypoxia is thus due to low O2 by prolyl hydroxylation inhibition rather than nNOS.

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Figure 4.2: S-nitrosylation of pVHL stabilization of HIF-1α in non-hypoxic manner. Figure adapted from Marsden PA, JCI 2007

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A novel pathway for SNO signaling in anemia

In a whole intact animal model, we demonstrated that nNOS can regulate

increased CO and reduced SVR in anemia. However, our assessment of isolated

resistance mesenteric arteries did not demonstrate a different vasodilatory phenotype in

anemic WT and nNOS-/- mice. This observation suggested that the reduction in SVR was

likely mediated by nNOS-dependent factors that were extrinsic to the resistance artery.

One such attractive candidate can be SNO-Hb.

As early as 1996, Stamler et al. has demonstrated the importance of red blood

cells (RBC) in the regulated delivery of NO to hypoxic vasculature. They had implicated

a key role for SNO-Hb in hypoxic vasodilation (25). RBCs are endogenously “preloaded”

with bioactive NO for delivery to vessels in conditions of oxyhemoglobin desaturation

(79, 80). In this model, an important cysteine residue within the β-globin chain of Hb

(Cysβ93) was biologically relevant to the control of local blood flow. They proposed that

SNO-Hb transports NO to the microcirculation. As O2 concentration decreases, the

release of O2 by SNO-Hb promotes a conformational change on Hb (T-structure) such

that the SNO on Cysβ93 exposes to solvent. This facilitates the transfer of NO groups to

other thiols such as glutathione and/or the red blood cell membrane protein AE1. Anemia

causes a right shift of the oxyhemoglobin dissociation curve which would favour O2

release from Hb (81). Thus, the change to T structure (deoxygenated) Hb in anemia may

facilitate NO release from SNO-Hb, which can improve NO/SNO signalling. The

increase in SNO content can promote a variety of physiological responses, such as

vasodilation (25), ventilation (73), and SNO-modification of proteins (3, 82-84).

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An alternative mechanism on hypoxic vasodilation had been proposed by other

investigators. Gladwin and colleagues reported that the vasodilative effects of nitrite

infusion into the circulation was associated with an increase in NO formation (26).

According to the reaction between nitrite and Hb, they suggested that as hemoglobin

oxygen saturation decreases, Hb can act as a nitrite reductase to convert nitrite to NO

(26, 85). Nitrite can be stored in tissues/plasma during normoxia, and can be reduced

back to NO during hypoxia. This dynamic regulation of NO may contribute to the

NO/SNO content in anemia.

overall

Regardless of the source, the importance of NO to regulate blood flow and modify

SNO proteins during deoxygenation is undisputable. Traditionally, eNOS has been

considered the primarily source of SNO “preloaded” on Hb to form SNO-Hb at the

pulmonary vasculature (3, 86). Our research demonstrated that tissue (neuronal or

vascular smooth muscle) nNOS can also be a source to prime SNO proteins (such as

pVHL) to stabilize HIF-1α in anemia. In addition, anemia can also affect the S-

nitrosylation equilibrium by reducing tissue GSNOR level. The composite effects lead to

increased tissue SNO content. Thus, in anemia, an NO gradient may occur from the

tissues and diffuse across to RBC to increase SNO-Hb level in the vasculature. Indeed,

our data that anemia causes increased in MetHb level supported increased NO content in

the blood. This process appears to be nNOS dependent, as the increase in MetHb level is

reduced in anemic nNOS-/- mice. In fact, experimental evidence suggested that MetHb

can form SNO-Hb (87, 88). This source of tissue NO could be due to increased tissue

nNOS/NO or nitrite reduction in anemia. Nonetheless, if the “spill over” from tissue

nNOS/NO to vasculature is true in vivo, our results might represent a novel mechanism

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by which tissue produce NO/SNO can regulate its local blood flow and SNO content.

Likewise, tissue SNO can be transported by SNO-Hb or specific proteins (89) to remote

areas where NO production is not favorable. This response has been implicated to the

control of central respiration (73) and may possibly have the potential to affect the central

control of cardiovascular function.

To delineate whether SNO derived from blood or tissue compartment is

responsible for the protective effect in anemia, survival studies with infusion of N-

acetylcysteine (NAC) to anemic WT and nNOS-/- mice may be desirable. NAC can act as

a bait molecule to decrease SNO content in the blood (3). If SNO level in blood confers

protection in anemia, we would expect an increased mortality in NAC-treated anemic

WT and nNOS-/- mice compared to non-treated mice because an important SNO signaling

in the blood is eliminated by NAC. If tissue SNO level is the primary source of

protection, we would expect that NAC treatment had no effect on the increased mortality

pattern in anemic nNOS-/- mice relative to WT mice. However, if both tissue and blood

SNO levels are important, we would expect a further increase in mortality in NAC-

treated nNOS-/- mice (deplete of tissue and blood NO) compared to non-treated or WT

mice during anemia. Additionally, measurement of SNO-Hb during anemia would also

be preferable. These experiments would provide further mechanistic insight into the role

of tissue nNOS and SNO signaling pathway in anemia.

Despite SNO-modification has been demonstrated to confer protection in acute

myocardial ischemic injury (84), however not all SNO signalling reactions are adaptive.

For example, increased SNO level in GSNOR-/- mice leads to increased morality during

sepsis (90). Also, the effect of the antioxidant NAC led to pulmonary arterial

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hypertension in a manner similar to chronic hypoxia (3). Therefore, future studies are

required to determine the adaptive or maladaptive effects of SNO in the setting of acute

anemia. By defining the importance of nNOS and nNOS-mediated signaling pathways,

novel therapies may be developed that could enhance survival in patients exposed to

acute blood loss and anemia, such as in battlefield and civilian trauma, and surgery.

In summary, we have provided novel evidence that both nNOS-dependent acute

and sustain mechanisms may contribute to survival in response to acute anemia. Acute

changes include nNOS mediated physiological adaptation in the brain, heart, and vessels

and rapid SNO modifications to optimize O2 delivery and utilization during anemia. After

these rapid responses, sustained mechanisms are activated, such as increased gene

transcription, to induce longer lasting effects and balance O2 delivery and utilization in

response to reduced blood O2 content. These mechanisms occur during anemia but not

hypoxia, suggesting that the manner in which nNOS contributes to maintaining O2

homeostasis is specific and critically important to anemia. Understanding these

mechanisms will help us advance in developing therapeutic treatments for anemic

patients.

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References: 1. Shander A, Javidroozi M, Ozawa S, & Hare G (2011) The prevalence and

association with transfusion, intensive care unit stay and mortality of pre-operative anaemia in a cohort of cardiac surgery patients. British Journal of Anesthesia.

2. Bin JP, et al. (2006) Effects of nitroglycerin on erythrocyte rheology and oxygen unloading: novel role of S-nitrosohemoglobin in relieving myocardial ischemia. Circulation 113(21):2502-2508.

3. Palmer LA, et al. (2007) S-nitrosothiols signal hypoxia-mimetic vascular pathology. J Clin Invest 117(9):2592-2601.

4. Dai J, Tu W, Yang Z, & Lin R (2010) Case report: intraoperative management of extreme hemodilution in a patient with a severed axillary artery. Anesth Analg 111(5):1204-1206.

5. Beilman GJ & Blondet JJ (2009) Near-infrared spectroscopy-derived tissue oxygen saturation in battlefield injuries: a case series report. World J Emerg Surg 4:25.

6. Carson JL, Noveck H, Berlin JA, & Gould SA (2002) Mortality and morbidity in patients with very low postoperative Hb levels who decline blood transfusion. Transfusion 42(7):812-818.

7. Carson JL, et al. (1996) Effect of anaemia and cardiovascular disease on surgical mortality and morbidity. Lancet 348(9034):1055-1060.

8. Carson JL, Poses RM, Spence RK, & Bonavita G (1988) Severity of anaemia and operative mortality and morbidity. Lancet 1(8588):727-729.

9. Kulier A, et al. (2007) Impact of preoperative anemia on outcome in patients undergoing coronary artery bypass graft surgery. Circulation. 116(5):471-479.

10. Karkouti K, Wijeysundera DN, & Beattie WS (2008) Risk associated with preoperative anemia in cardiac surgery: a multicenter cohort study. Circulation. 117(4):478-484.

11. Habib RH, et al. (2003) Adverse effects of low hematocrit during cardiopulmonary bypass in the adult: should current practice be changed? J.Thorac.Cardiovasc.Surg. 125(6):1438-1450.

12. Musallam KM, et al. (2011) Preoperative anaemia and postoperative outcomes in non-cardiac surgery: a retrospective cohort study. Lancet 378(9800):1396-1407.

13. Kimura H, et al. (2010) Cardio-renal interaction: impact of renal function and anemia on the outcome of chronic heart failure. Heart Vessels 25(4):306-312.

14. Diedler J, et al. (2010) Low hemoglobin is associated with poor functional outcome after non-traumatic, supratentorial intracerebral hemorrhage. Crit Care 14(2):R63.

15. McIntyre LA, et al. (2006) Effect of a liberal versus restrictive transfusion strategy on mortality in patients with moderate to severe head injury. Neurocrit.Care. 5(1):4-9.

16. Hebert PC, et al. (2001) Is a low transfusion threshold safe in critically ill patients with cardiovascular diseases? Crit Care Med. 29(2):227-234.

17. Murray JF & Rapaport E (1972) Coronary blood flow and myocardial metabolism in acute experimental anaemia. Cardiovasc Res 6(4):360-367.

181

Chapter 4

18. Weiskopf RB, et al. (1998) Human cardiovascular and metabolic response to acute, severe isovolemic anemia. JAMA. 279(3):217-221.

19. Hatcher JD, Chiu LK, & Jennings DB (1978) Anemia as a stimulus to aortic and carotid chemoreceptors in the cat. J.Appl.Physiol. 44(5):696-702.

20. Szlyk PC, King C, Jennings DB, Cain SM, & Chapler CK (1984) The role of aortic chemoreceptors during acute anemia. Can J Physiol Pharmacol 62(5):519-523.

21. Glick G, Plauth WH, Jr., & Braunwald E (1964) Role of the autonomic nervous system in the circulatory response to acutely induced anemia in unanesthetized dogs. Journal of Clinical Investigation 43:2112-24.:2112-2124.

22. Guyton AC & Richardson TQ (1961) Effect of hematocrit on venous return. Circ Res 9:157-164.

23. Chapler CK & Cain SM (1986) The physiologic reserve in oxygen carrying capacity: studies in experimental hemodilution. Canadian Journal of Physiology & Pharmacology. 64(1):7-12.

24. McLaren AT, et al. (2007) Increased expression of HIF-1alpha, nNOS, and VEGF in the cerebral cortex of anemic rats. Am J Physiol Regul Integr Comp Physiol 292(1):R403-414.

25. Jia L, Bonaventura C, Bonaventura J, & Stamler JS (1996) S-nitrosohaemoglobin: a dynamic activity of blood involved in vascular control. Nature 380(6571):221-226.

26. Cosby K, et al. (2003) Nitrite reduction to nitric oxide by deoxyhemoglobin vasodilates the human circulation. Nat.Med. 9(12):1498-1505.

27. Eichelbronner O, D'Almeida M, Sielenkamper A, Sibbald WJ, & Chin-Yee IH (2002) Increasing P(50) does not improve DO(2CRIT) or systemic VO(2) in severe anemia. Am.J.Physiol Heart Circ.Physiol 283(1):H92-101.

28. Ickx BE, Rigolet M, & Van Der Linden PJ (2000) Cardiovascular and metabolic response to acute normovolemic anemia. Effects of anesthesia. Anesthesiology 93(4):1011-1016.

29. Ragoonanan TE, et al. (2009) Metoprolol reduces cerebral tissue oxygen tension after acute hemodilution in rats. Anesthesiology 111(5):988-1000.

30. Brannon ES, Merrill AJ, Warren JV, & Stead EA (1945) The Cardiac Output in Patients with Chronic Anemia as Measured by the Technique of Right Atrial Catheterization. J Clin Invest 24(3):332-336.

31. van Bommel J, et al. (2002) Intestinal and cerebral oxygenation during severe isovolemic hemodilution and subsequent hyperoxic ventilation in a pig model. Anesthesiology 97(3):660-670.

32. Fan FC, Chen RY, Schuessler GB, & Chien S (1980) Effects of hematocrit variations on regional hemodynamics and oxygen transport in the dog. Am.J.Physiol 238(4):H545-522.

33. Grupp I, Grupp G, Holmes JC, & Fowler NO (1972) Regional blood flow in anemia. J Appl Physiol 33(4):456-461.

34. Johannsson H & Siesjo BK (1974) Blood flow and oxygen consumption in the rat brain in dilutional anemia. Acta Physiol Scand. 91(1):136-138.

182

Chapter 4

35. Johannes T, Mik EG, Nohe B, Unertl KE, & Ince C (2006) Acute Decrease in Renal Microvascular PO2 during Acute Normovolemic Hemodilution. Am.J.Physiol Renal Physiol. .

36. von Restorff W, Hofling B, Holtz J, & Bassenge E (1975) Effect of increased blood fluidity through hemodilution on coronary circulation at rest and during exercise in dogs. Pflugers Arch 357(1-2):15-24.

37. Ward ME, et al. (2005) Hypoxia induces a functionally significant and translationally efficient neuronal NO synthase mRNA variant. J Clin Invest 115(11):3128-3139.

38. Hare GM, et al. (2003) Hemodilutional anemia is associated with increased cerebral neuronal nitric oxide synthase gene expression. J Appl Physiol 94(5):2058-2067.

39. Stuehr DJ, Santolini J, Wang ZQ, Wei CC, & Adak S (2004) Update on mechanism and catalytic regulation in the NO synthases. J Biol Chem 279(35):36167-36170.

40. Sears CE, et al. (2003) Cardiac neuronal nitric oxide synthase isoform regulates myocardial contraction and calcium handling. Circ.Res. 92(5):e52-e59.

41. Ashley EA, Sears CE, Bryant SM, Watkins HC, & Casadei B (2002) Cardiac nitric oxide synthase 1 regulates basal and beta-adrenergic contractility in murine ventricular myocytes. Circulation. 105(25):3011-3016.

42. Zhang YH, et al. (2008) Reduced phospholamban phosphorylation is associated with impaired relaxation in left ventricular myocytes from neuronal NO synthase-deficient mice. Circ Res 102(2):242-249.

43. Wang Y, Liu XF, Cornish KG, Zucker IH, & Patel KP (2005) Effects of nNOS antisense in the paraventricular nucleus on blood pressure and heart rate in rats with heart failure. Am.J.Physiol Heart Circ.Physiol. 288(1):H205-H213.

44. Danson EJ, et al. (2004) Impaired regulation of neuronal nitric oxide synthase and heart rate during exercise in mice lacking one nNOS allele. J Physiol 558(Pt 3):963-974.

45. Hudetz AG, Wood JD, & Kampine JP (2000) 7-Nitroindazole impedes erythrocyte flow response to isovolemic hemodilution in the cerebral capillary circulation. J.Cereb.Blood Flow Metab 20(2):220-224.

46. Yang G, Zhang Y, Ross ME, & Iadecola C (2003) Attenuation of activity-induced increases in cerebellar blood flow in mice lacking neuronal nitric oxide synthase. American Journal of Physiology - Heart & Circulatory Physiology. 285(1):H298-H304.

47. Taylor CT & McElwain JC (2010) Ancient atmospheres and the evolution of oxygen sensing via the hypoxia-inducible factor in metazoans. Physiology (Bethesda) 25(5):272-279.

48. Marti HH & Risau W (1998) Systemic hypoxia changes the organ-specific distribution of vascular endothelial growth factor and its receptors. Proc.Natl.Acad.Sci.U.S.A 95(26):15809-15814.

49. Shweiki D, Itin A, Soffer D, & Keshet E (1992) Vascular endothelial growth factor induced by hypoxia may mediate hypoxia-initiated angiogenesis. Nature 359(6398):843-845.

183

Chapter 4

50. Cao L, et al. (2004) VEGF links hippocampal activity with neurogenesis, learning and memory. Nat.Genet. 36(8):827-835.

51. Marti HJ, et al. (2000) Hypoxia-induced vascular endothelial growth factor expression precedes neovascularization after cerebral ischemia. Am J Pathol 156(3):965-976.

52. Pichiule P & LaManna JC (2002) Angiopoietin-2 and rat brain capillary remodeling during adaptation and deadaptation to prolonged mild hypoxia. J.Appl.Physiol 93(3):1131-1139.

53. Dunst J, et al. (2002) Anemia and elevated systemic levels of vascular endothelial growth factor (VEGF). Strahlentherapie und Onkologie. 178(8):436-441.

54. Rakusan K, Cicutti N, & Kolar F (2001) Effect of anemia on cardiac function, microvascular structure, and capillary hematocrit in rat hearts. American Journal of Physiology - Heart & Circulatory Physiology. 280(3):H1407-H1414.

55. Schoch HJ, Fischer S, & Marti HH (2002) Hypoxia-induced vascular endothelial growth factor expression causes vascular leakage in the brain. Brain 125(Pt 11):2549-2557.

56. Zhang ZG, et al. (2000) VEGF enhances angiogenesis and promotes blood-brain barrier leakage in the ischemic brain. Journal of Clinical Investigation. 106(7):829-838.

57. Pardridge WM, Boado RJ, & Farrell CR (1990) Brain-type glucose transporter (GLUT-1) is selectively localized to the blood-brain barrier. Studies with quantitative western blotting and in situ hybridization. J Biol Chem 265(29):18035-18040.

58. Kim JW, Tchernyshyov I, Semenza GL, & Dang CV (2006) HIF-1-mediated expression of pyruvate dehydrogenase kinase: a metabolic switch required for cellular adaptation to hypoxia. Cell Metab. 3(3):177-185.

59. Papandreou I, Cairns RA, Fontana L, Lim AL, & Denko NC (2006) HIF-1 mediates adaptation to hypoxia by actively downregulating mitochondrial oxygen consumption. Cell Metab 3(3):187-197.

60. Firestein BL & Bredt DS (1999) Interaction of neuronal nitric-oxide synthase and phosphofructokinase-M. J.Biol.Chem. 274(15):10545-10550.

61. Digicaylioglu M & Lipton SA (2001) Erythropoietin-mediated neuroprotection involves cross-talk between Jak2 and NF-kappaB signalling cascades. Nature 412(6847):641-647.

62. Sakanaka M, et al. (1998) In vivo evidence that erythropoietin protects neurons from ischemic damage. Proc.Natl.Acad.Sci.U.S.A 95(8):4635-4640.

63. Ihle JN (1995) Cytokine receptor signalling. Nature 377(6550):591-594. 64. Silva M, et al. (1996) Erythropoietin can promote erythroid progenitor survival by

repressing apoptosis through Bcl-XL and Bcl-2. Blood 88(5):1576-1582. 65. Keswani SC, et al. (2011) Nitric oxide prevents axonal degeneration by inducing

HIF-1-dependent expression of erythropoietin. Proc Natl Acad Sci U S A 108(12):4986-4990.

66. Hare JM (2003) Nitric oxide and excitation-contraction coupling. J Mol Cell Cardiol 35(7):719-729.

184

Chapter 4

67. Sun J, et al. (2006) Hypercontractile female hearts exhibit increased S-nitrosylation of the L-type Ca2+ channel alpha1 subunit and reduced ischemia/reperfusion injury. Circ Res 98(3):403-411.

68. Burger DE, et al. (2009) Neuronal nitric oxide synthase protects against myocardial infarction-induced ventricular arrhythmia and mortality in mice. Circulation 120(14):1345-1354.

69. Sun J, Morgan M, Shen RF, Steenbergen C, & Murphy E (2007) Preconditioning results in S-nitrosylation of proteins involved in regulation of mitochondrial energetics and calcium transport. Circ Res 101(11):1155-1163.

70. Gonzalez DR, Beigi F, Treuer AV, & Hare JM (2007) Deficient ryanodine receptor S-nitrosylation increases sarcoplasmic reticulum calcium leak and arrhythmogenesis in cardiomyocytes. Proc Natl Acad Sci U S A 104(51):20612-20617.

71. Shi Q, Feng J, Qu H, & Cheng YY (2008) A proteomic study of S-nitrosylation in the rat cardiac proteins in vitro. Biol Pharm Bull 31(8):1536-1540.

72. Deem S, et al. (1999) Mechanisms of improvement in pulmonary gas exchange during isovolemic hemodilution. J.Appl.Physiol 87(1):132-141.

73. Lipton AJ, et al. (2001) S-nitrosothiols signal the ventilatory response to hypoxia. Nature. 413(6852):171-174.

74. Page EL, Chan DA, Giaccia AJ, Levine M, & Richard DE (2008) Hypoxia-inducible Factor-1{alpha} Stabilization in Nonhypoxic Conditions: Role of Oxidation and Intracellular Ascorbate Depletion. Mol.Biol.Cell 19(1):86-94.

75. Page EL, Robitaille GA, Pouyssegur J, & Richard DE (2002) Induction of hypoxia-inducible factor-1alpha by transcriptional and translational mechanisms. J Biol Chem 277(50):48403-48409.

76. Semenza GL (2009) Regulation of oxygen homeostasis by hypoxia-inducible factor 1. Physiology (Bethesda) 24:97-106.

77. van Bommel J, Siegemund M, Henny Ch P, & Ince C (2008) Heart, kidney, and intestine have different tolerances for anemia. Transl Res 151(2):110-117.

78. Schild L, Jaroscakova I, Lendeckel U, Wolf G, & Keilhoff G (2006) Neuronal nitric oxide synthase controls enzyme activity pattern of mitochondria and lipid metabolism. FASEB J 20(1):145-147.

79. Jia L, Bonaventura C, Bonaventura J, & Stamler JS (1996) S-nitrosohaemoglobin: a dynamic activity of blood involved in vascular control. Nature 380(6571):221-226.

80. Marozkina N, Gaston B, & Doctor A (2008) Transnitrosation signals oxyhemoglobin desaturation. Circ Res 103(5):441-443.

81. Torrance J, et al. (1970) Intraerythrocytic adaptation to anemia. N Engl J Med 283(4):165-169.

82. Whalen EJ, et al. (2007) Regulation of beta-adrenergic receptor signaling by S-nitrosylation of G-protein-coupled receptor kinase 2. Cell 129(3):511-522.

83. Jaffrey SR, Erdjument-Bromage H, Ferris CD, Tempst P, & Snyder SH (2001) Protein S-nitrosylation: a physiological signal for neuronal nitric oxide. Nat.Cell Biol. 3(2):193-197.

84. Lima B, et al. (2009) Endogenous S-nitrosothiols protect against myocardial injury. Proc Natl Acad Sci U S A 106(15):6297-6302.

185

Chapter 4

186

85. Crawford JH, et al. (2006) Hypoxia, red blood cells, and nitrite regulate NO-dependent hypoxic vasodilation. Blood 107(2):566-574.

86. Schleicher M, et al. (2009) The Akt1-eNOS axis illustrates the specificity of kinase-substrate relationships in vivo. Sci Signal 2(82):ra41.

87. Luchsinger BP, et al. (2003) Routes to S-nitroso-hemoglobin formation with heme redox and preferential reactivity in the beta subunits. Proc Natl Acad Sci U S A 100(2):461-466.

88. Angelo M, Singel DJ, & Stamler JS (2006) An S-nitrosothiol (SNO) synthase function of hemoglobin that utilizes nitrite as a substrate. Proc Natl Acad Sci U S A 103(22):8366-8371.

89. Kornberg MD, et al. (2010) GAPDH mediates nitrosylation of nuclear proteins. Nat Cell Biol 12(11):1094-1100.

90. Liu L, et al. (2004) Essential roles of S-nitrosothiols in vascular homeostasis and endotoxic shock. Cell 116(4):617-628.