t...
Liv: s v

Y! «1"

JIV

:1.

31' n»

.
{ix
:0

.e.

.!1
33:31.:

0.5 5.4;
9.3%....

h

t}
.l

..: é.

.6: ‘1!

h.......u..... F l a...»

a. .u... 39!:

.. 96...)

. (J I:

E. . l. ‘.
its»; ..

IEi Pia... IS.

= \ Cl‘u..."
In? if}.

. {:1
\
.uuchuw...
‘ “than...

10..

km

. I
:2.
o v

I. I...
t
I s
’2‘:

5

cqfimtwbvflhm
r t...

 

tithe"...
1.0:» .
{ah-b."

 

«I LIBRARY
Mich?" State

Umvelsuy

 

 

 

This is to certify that the
dissertation entitled

VASCULAR REACTIVE OXYGEN SPECIES METABOLISM
AND ITS INFLUENCE ON VENOMOTOR TONE AND
HYPERTENSION

presented by

IRINA THEODORA SZASZ

has been accepted towards fulfillment
of the requirements for the

Ph.D. degree in Microbiology and Molecular
Genetics

 

 

SWW (Maggi

‘ Major Professor’s Signature

Qfiq/zooq

Date

MSU is an Affirmative Action/Equal Opportunity Employer

 

_A_.—.—.--—--.-.--u---.—-.-.-.—.-.—._._.-.-p—.-a-n----a-u--n-n---—--

PLACE IN RETURN BOX to remove this checkout from your record.
TO AVOID FINES return on or before date due.
MAY BE RECALLED with earlier due date if requested.

 

DATE DUE

DATE DUE DATE DUE

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

5/08 K'IProj/AchrelelRC/DateDue indd

 

VASCULAR REACTIVE OXYGEN SPECIES METABOLISM AND ITS
INFLUENCE ON VENOMOTOR TONE AND HYPERTENSION

By

Irina Theodora Szasz

A DISSERTATION

Submitted to
Michigan State University
in partial fulfillment of the requirements
for the degree of

DOCTOR OF PHILOSOPHY
Microbiology and Molecular Genetics

2009

ABSTRACT

VASCULAR REACTIVE OXYGEN SPECIES METABOLISM AND ITS
INFLUENCE ON VENOMOTOR TONE AND HYPERTENSION

By

Irina Theodora Szasz

Reactive oxygen species (ROS) are signaling mediators with multiple effects on
vascular function. ROS and oxidative stress have been implicated in the
pathogenesis of cardiovascular diseases such as hypertension. Veins and
arteries both contribute to the regulation of blood pressure (BP). As compared to
arteries, far less is currently known about venous ROS metabolism and its
functional impact. We tested the hypothesis that the metabolism and handling of
ROS would be different in veins compared to arteries and that venomotor

function, as influenced by ROS, would play a role in experimental hypertension.

We observed higher superoxide and hydrogen peroxide (H2O2) levels, as well as
higher antioxidant capacity for H2O2, in rat venous compared to corresponding
arterial tissues. There were multiple differences in ROS metabolizing enzymes
between arteries and veins, such as a higher expression of DUOX1, CuZn SOD
and catalase in venous compared to arterial tissue. Xanthine oxidase (XO) had a
higher protein and mRNA expression, as well as higher activity in veins
compares to arteries. During deoxycorticosterone acetate (DOCA)-salt
hypertension, the higher expression of XO was maintained by veins, and there

were indications of an increase in venous, but not arterial XO activity. The

contractile impact of ROS was also higher in veins compared to arteries. XO
appeared to play little role in normal arterial contractile function, however it
seemed to contribute to normal venous ET—1-mediated contraction. In
norrnotensive rats, the endothelial function of arteries and veins was similar.
During DOCA-salt hypertension, arteries, but not veins, were affected by
endothelial dysfunction. XO did not appear to contribute to the arterial
hypertensive dysfunction by either ROS or urate production. We inhibited XO
activity in vivo using the XO inhibitor allopurinol, expecting as consequences a
lowering of venomotor tone and therefore an effect on BP depending on the
administration. timing: either a lowering of BP if allopurinol was administered
when DOCA—salt hypertension was established or an attenuation of hypertension
development if it was administered before the DOCA-salt treatment. We
monitored XO metabolites in the urine, serum and tissues of allopurinol-treated
animals, overall observing increases in X0 substrates and decreases in X0
products. Despite what we considered effective XO inhibition, we did not observe
any effect on BP in DOCA—salt hypertensive rats, regardless of the timing of this

intervention.

These results suggest there are differences in ROS metabolism and functional
impact inherent to the blood vessel type. Although we believe we excluded the
involvement of X0 in DOCA-salt hypertension pathogenesis, further studies are
needed to uncover the role of ROS in venous contractile function and

hypertension pathogenesis.

ACKNOWLEDGEMENT

Over the course of time I spent as a PhD student, I have had the privilege and

pleasure of working with and learning from a number of remarkable people.

I thank, first and foremost, my mentor, Dr. Stephanie Watts. I am a scientist
today because of her. She supported me in countless ways professionally and
personally. Her energy, generosity and honesty are an inspiration for the future
mentor I hope to become. There truly are not enough words to express my

gratitude for her. Thank you, Stephanie!

I thank Dr. Wei Ni and Dr. Keshari Thakali, who have both helped me in the

beginning of my PhD and have remained not only my models but also my friends.

I have the utmost respect for the rigorous scientific methods applied by our
technicians. I learned a lot of technical subtleties from Janice Thompson, who
also helped me immensely in the everyday life of the lab; and Robert Burnett
provided crucial help for a big part of the experiments contained within my thesis.

I am extremely grateful for their support.

I thank the other graduate and undergraduate students who are or have been in
the Watts lab, especially Patrick Davis, whose practical and moral support were
and are invaluable, and Becca Sattler-Leja, my first student, who probably taught
me at least as much as I taught her and who has also helped with a lot of

experiments contained within this thesis.

 

I owe the Department of Microbiology and Molecular Genetics my being here and
I consider myself privileged to have been admitted into its graduate program. I
am also grateful to Drs Gregory Fink, Andrea Amalfitano, Susan Conrad and
Richard Schwartz, who, as members of my guidance committee, have been
instrumental in providing me with a lot of valuable feed—back that helped refine

and strengthen my work.

I have benefitted scientifically from being a part of the PPG group at MSU and
collaborating in various ways with Dr. Galligan, Dr. Fink and the people in their
laboratories. I am also grateful to the Department of Pharmacology and
Toxicology in its entirety, which has been my happy research home throughout

these years.

I am honored to have received a predoctoral fellowship from the American Heart

Association, which has financially supported a large part of my PhD.

My special thanks go to Dr. Elizabeth Linder, my colleague, mentor and most
important, the best friend one could ever dream of. I am lucky to have met

Elizabeth and I hope to always have her in my life.

I would not exist without the infinite support of my friends and family: Miha
Marina, all my friends from around the world, and the entire Szasz family (Ivan,

Lili, Dan, Gina and Cristi).

TABLE OF CONTENTS

LIST OF TABLES ................................................................................................ viii
LIST OF FIGURES ................................................................................................ ix
LIST OF ABBREVIATIONS ................................................................................. xii
Introduction .......................................................................................................... 1

I. General characteristics of arteries and veins ...................................................... 1

II. The role of veins in blood pressure regulation and hypertension ...................... 7

III. Vascular ROS metabolism .............................................................................. 12
IV. R08 involvement in vascular function ............................................................ 26
V. Comparisons of arterial and venous ROS metabolism .................................... 28
VI. Oxidative stress and role of ROS in hypertension .......................................... 29
VII. Xanthine oxidase: role in cardiovascular disease .......................................... 31
Hypothesis ................... , ....................................................................................... 39
Methods ....................................................................... . ....................................... 41

I. Animal use ........................................................................................................ 41

ll. Basal superoxide levels by lucigenin-enhanced chemiluminescence
measurements ..................................................................................................... 44
Ill. H2O2 measurements ....................................................................................... 45
IV. Protein concentration determination ............................................................... 48
V. Western blot analysis ...................................................................................... 51

VI. lmmunohistochemistry of paraffin-embedded sections .................................. 52

VII. mRNA expression .......................................................................................... 53
VIII. XO activity assays ........................................................................................ 57
IX. Isolated tissue bath contractility ...................................................................... 58
X. Blood uric acid measurement .......................................................................... 59
XI. High performance liquid chromatography ....................................................... 60
XII. Data analysis .................................................................................................. 62
Results ................................................................................................................ 63
Specific aim 1 ....................................................................................................... 63
I. ROS production ................................................................................................ 63

II. ROS destruction ............................................................................................... 64
Ill. ROS metabolizing enzymes ............................................................................ 68
IV. ROS metabolism in DOCA-salt hypertension ................................................. 82
V. X0 in normal conditions and in DOCA-salt hypertension ................................ 83
Specific aim 2 ....................................................................................................... 91
l. Direct impact of ROS on contractile function of arteries and veins ................... 91

vi

 

ll. Impact of XO on arterial and venous contractile function in normal rats ........ 93
III. Impact of flavoenzymes on endothelin-1 induced contraction of aorta and

vena cava ............................................................................................................. 99 ’
IV. Arterial and venous endothelial function in normal rats and during DOCA-salt
hypertension ........................................................................................................ 99
V. Impact of XO on arterial endothelial function in DOCA-salt hypertension ..... 104
Specific aim 3 ..................................................................................................... 111
l. XO inhibition in the established DOCA—salt hypertension ............................... 111
ll. XO inhibition in the development of DOCA—salt hypertension ....................... 114
Ill. Dose-dependent effects of allopurinol .......................................................... 131
Discussion ....................................................................................................... 135
l. ROS metabolism in veins vs arteries .............................................................. 135
II. Impact of ROS on venous and arterial contractile function ............................ 147
III. Results of in vivo XO inhibition in DOCA—salt hypertension .......................... 154
IV. Conclusions .................................................................................................. 161
Limitations .......................................................................................................... 162
Perspectives ...................................................................................................... 164
References ....................................................................................................... 165

vii

i

LIST OF TABLES

Table 1. Enzymatic ROS producers and ROS destroyers ........................... 15
Table 2. Results of PCR array on oxidative stress and antioxidant defense
genes in rat aorta and vena cava ................................................... 78

viii

Figure 1.
Figure 2.

Figure 3.
Figure 4.
Figure 5.
Figure 6.
Figure 7.
Figure 8.
Figure 9.

Figure 10.
Figure 11.
Figure 12.
Figure 13.

Figure 14.
Figure 15.

Figure 16.

Figure 17.

Figure 18.

Figure 19.

LIST OF FIGURES

Images in this dissertation are presented in color.

Diagram of generic artery-vein pair ................................................. 5
Smooth muscle staining of rat aorta and vena cava: A. modified
trichrome Masson; B. d-actin ........................................................... 6
Diagram of blood pressure regulation ............................................ 11
Diagram of ROS metabolism .......................................................... 13
Diagram of purine catabolism ......................................................... 20
Diagram of ROS effects on vascular function ................................ 27
Diagram of working hypothesis ...................................................... 40
Standard curves for: A. H202; B. uric acid measurements ............ 47
Standard curves for protein assays: A. BCA; B. Lowry; C.
Bradford ......................................................................................... 50
HPLC tracing for X0 metabolites ................................................... 61
Basal superoxide production: A. aorta-VC; B. SMA-SMV .............. 65
Basal H202 production: A. aorta-VC; B. CA-JV .............................. 66
H202 production in the presence of DDC: A. aorta-VC; B. CA-
JV ................................................................................................... 67
Antioxidant capacity for H202 of aorta and VC ............................... 69

Protein expression of 4-HNE in aorta and VC: A.
lmmunohistochemistry; B. Western blotting ................................... 70

Protein expression of X0 in aorta and VC: A. Western blotting;
B. lmmunohistochemistry ............................................................... 74

Protein expression of DUOX1 in aorta and V0: A. Western blotting;
B. lmmunohistochemistry ............................................................... 75

Protein expression of CuZn SOD in aorta and VC: A. Western
blotting; B. lmmunohistochemistry ............................................. 76

Protein expression of catalase in aorta and VC: A. Western blotting;
B. lmmunohistochemistry ............................................................... 77

Figure 20.
Figure 21.

Figure 22.

Figure 23.

Figure 24.

Figure 25.
Figure 26.

Figure 27.

Figure 28.

Figure 29.

Figure 30.

Figure 31.

Figure 32.

Figure 33.

Figure 34.

Figure 35.

Figure 36.

Figure 37.

Aorta and VC mRNA expression of: A. DUOX1; B. DUOX2 .......... 79
H202 production in aorta, VC, CA, JV in the presence of ROS

enzyme inhibitors, as grouped by inhibitor ..................................... 80
H202 production in aorta, VC, CA, JV in the presence of ROS
enzyme inhibitors, as grouped by tissue ........................................ 81
Basal H202 production in tissues from DOCA-salt rats: A. aorta-VC;
B. CA-JV ......................................................................................... 85
Western blot analysis of ROS metabolizing enzymes in sham and
DOCA aorta and VC ....................................................................... 86
Normal and DOCA aorta and VC mRNA expression of X0 ........... 87

X0 activity in normal aorta and VC represented as: A. urate
production; B. H202 production ...................................................... 88

X0 activity in normal and DOCA tissues: A. urate producing X0
activity of aorta and VC; B. H202 producing X0 activity of aorta and

VC; 0. H202 producing X0 activity of CA and JV ........................... 89
Contractile response to xanthine/XO of aorta and V0 ................... 92
Contractile responses of aorta in the presence/absence of
allopurinol to: A. NE; B. ACh; C. Angll; D. ET-1 ............................. 94
Contractile responses of VC in the presence/absence of allopurinol
to: A. NE; B. ACh; C. Angll; D. ET-1 .............................................. 95
Protein expression of uricase in normal aorta and V0 .................. 97

Contractile effects of UA: A. direct contractile effects on aorta and
VC; B. effects on ACh-induced relaxation of aorta; C. effects on
ACh-induced relaxation of VC ........................................................ 98

Contractile response to ET-1 in the presence of DPI of: A. aorta; B.
VC ................................................................................................ 101

Contractile response of normal aorta and VC to ACh in the
presence of: A. vehicle; B. LNNA ................................................. 102

Contractile response of DOCA aorta and VC to ACh in the
presence of: A. vehicle; B. LNNA ................................................. 103

Contractile response of sham and DOCA aorta to ACh in the
presence/absence of allopurinol ................................................... 105

Contractile response of sham and DOCA aorta to ACh in the
presence/absence of UA .............................................................. 106

X

Figure 38.
Figure 39.
Figure 40.

Figure 41.

Figure 42.

Figure 43.

Figure 44.

Figure 45.

Figure 46.

Figure 47.

Figure 48.

Figure 49.

Figure 50.

Figure 51.

Figure 52.

Figure 53.

Figure 54.

Uric acid levels in blood of sham and DOCA rats ........................ 107
Protein expression of uricase in sham and DOCA aorta .............. 108
Study design for allopurinol effects in the established DOCA-salt
hypertension ................................................................................. 1 15
Urinary X0 metabolites in vehicle and allopurinol-treated rats in the
established DOCA-salt hypertension ........................................... 116
Serum XO metabolites in vehicle and allopurinol-treated rats in the
established DOCA-salt hypertension ........................................... 117
Tissue X0 metabolites in vehicle and allopurinol-treated rats in the
established DOCA-salt hypertension ........................................... 118
Allopurinol effects on BP in the established DOCA-salt
hypertension ................................................................................. 1 19

Study design for allopurinol effects in the development of DOCA-
salt hypertension .......................................................................... 123

Urinary X0 metabolites in vehicle and allopurinol-treated rats in the
development of DOCA-salt hypertension ..................................... 124

Urinary XO metabolites in vehicle and allopurinol-treated rats in the
development of DOCA-salt hypertension ..................................... 125

Urinary X0 metabolites in vehicle and allopurinol-treated rats in the
development of DOCA-salt hypertension ..................................... 126

Allopurinol effects on BP in the development of DOCA-salt
hypertension ................................................................................. 127

Hemodynamic parameters of vehicle and allopurinol-treated rats in
the development of DOCA-salt hypertension ............................... 128

Measures of target organ damage in vehicle and allopurinol-treated
rats in the development of DOCA-salt hypertension: A. heart
weight; B. kidney weight ............................................................... 129

Hematoxylin-eosin staining of aorta from vehicle and allopurinol-
treated rats in the development of DOCA-salt
hypertension ................................................................................. 130

Urinary XO metabolites in vehicle and allopurinol (50 and 100
mg/kg/day)-treated rats in normal rats ......................................... 133

Serum XO metabolites in vehicle and allopurinol (50 and 100
mg/kg/day)-treated rats in normal rats ......................................... 134

xi

 

4-HNE
ACh
Ang II
BCA
BH4
BP
BSA
CA
CO
COX
CVD
DBP
DDC
DOCA
DPI
DUOX
ET-1
H202
HPLC
HSV
HUVEC

JV

LN NA
LOX
MAP

LIST OF ABBREVIATIONS

4-hydroxy-nonenal
acetylcholine

angiotensin II

bicinchoninic acid
tetrahydrobiopterin

blood pressure

bovine serum albumin
carotid artery

cardiac output
cyclooxygenase
cardiovascular disease
diastolic blood pressure
diethyldithiocarbamate
deoxycorticosterone acetate
diphenyleniodonium

dual oxidase

endothelin 1

hydrogen peroxide

high performance liquid chromatography
human saphenous vein
human umbilical vein endothelial cell
internal mammary artery
jugular vein
Nw-Nitro-L-arginine
Iipoxygenase

mean arterial pressure

xii

MCFP
NADPH
NE
NOS
PGan
PE
PSS
ROS
RT—PCR
SBP
SEM
SHR
SMA
SMV
SOD
TPR
UA
VC
VSMC
XDH
XO

mean circulatory filling pressure
nicotinamide dinucleotide phosphate (reduced)
norepinephrine

nitric oxide synthase

prostaglandin F 2 alpha
phenylephrine

physiological salt solution

reactive oxygen species

real time polymerase chain reaction
systolic blood pressure

standard error of the means
spontaneous hypertensive rat
superior mesenteric artery

superior mesenteric vein
superoxide dismutase

total peripheral resistance

uric acid

vena cava

vascular smooth muscle cell
xanthine dehydrogenase

xanthine oxidase

xiii

INTRODUCTION

I. General characteristics of arteries and veins
A. Structure

Arteries and veins, two separate components of the vascular system, are
different structurally and functionally. While arteries carry oxygenated blood from
the heart to the peripheral tissues at a high pressure, therefore requiring a more
elastic and muscular structure, veins carry blood from the tissues back to the
heart at a low pressure, providing capacitance, therefore requiring more

distensible, less muscular walls (fig 1).

Both artery and vein are composed of similar layers: the innermost layer or the
tunica intima, containing endothelial cells; the tunica media, which is largely
composed of smooth muscle, elastin and collagen; and the outermost tunica
adventitia, containing mainly fibroblasts, collagen and elastin. Small blood
vessels called vasa vasorum integrate into the adventitia of larger vessels,

providing nutrients to the vascular wall itself.

Several characteristics, in addition to a different distribution and relative
abundance of these layers, distinguish arteries from veins. The delineation of the
three layers is more obvious in an artery compared to a vein. This is particularly
illustrated when viewing the thoracic vena cava vs the thoracic aorta from the
same rat (fig 2). The media of an artery, flanked by two elastic laminas, is

typically thicker than that of a vein, while the elastic component of a vein is

1

smaller compared to that of an artery, in favor of a much larger adventitial
component in the vein. The greater relative contribution of the smooth muscle
layer to the thickness of the vascular wall in arteries compared to veins can be
appreciated in figure 2A. These differences are confirmed by
immunohistochemical staining for d-actin, a smooth muscle marker (fig 2B).
Larger veins possess venous valves on the luminal side of the wall, which help
prevent backflow of blood. The cardiovascular system is not abruptly split into the
two components (arteries and veins). Rather, a gradual transition of tissue
characteristics ensues: from the heart to the large elastic arteries, then smaller
muscular arteries/arterioles to, finally, the capillary section, having just one
endothelial layer, and then back from the peripheral tissues, through less
muscular venules, to large capacitance veins possessing all the components of

the vessel wall and back to the heart (43).

B.Funcfion

Due to these structural differences, there are also inherent differences in the
contractility and synthetic properties of arteries and veins that can impact overall
cardiovascular function. These differences in contractility, however, may vary
according to vascular beds, similarly to the structural differences. The magnitude
of the contractile force developed by a vein in response to receptor-dependent
and independent agonists is smaller compared to that developed by an artery.

The time needed to reach half maximal contraction, a measure of response

speed, is shorter for a vein than for an artery (61) The capacity to relax in
response to certain agonists that induce the production of endothelium-derived
relaxant factors is decreased in veins compared to arteries, in some vascular
beds (70, 125, 131). Similarly, specific differences exist in the contractile
response of arteries and veins to a series of receptor-dependent agonists, the
best studied of them being endothelin-1 (ET-1), a potent, though not selective,
venoconstrictor. There seem to be fundamental arterial-venous differences in the
endothelin system, such as receptor expression and interaction (153), processing
of endothelin-related peptides (169) or functional responses to endothelin

peptides and ET receptor agonists (152, 167).

Different properties of arterial and venous grafts used in bypass surgery, leading
to different outcomes, have stimulated research on comparing these vessel types
and the factors that influence their long term patency. Venous smooth muscle
cells appear to have a higher growth rate compared to their arterial counterparts,
both in basal conditions (177) as well as in response to various mitogenic stimuli.
Endothelium function is also different in veins compared to arteries. Venous
endothelium produces less prostacyclin (101) and NO (130) than arterial
endothelium, and its overall response to atherogenic stimuli is different.
Endothelial intercellular junctions are generally tighter in arteries than in veins. In
venules, at the post-capillary level, the organization of these junctions is
especially loose, thus making this sector of vasculature the primary site of

inflammatory permeability (23).

These intrinsic properties ultimately reflect the difference in the gene expression
pattern of arteries and veins. The most prominent differences in basal gene
expression between arteries and veins can be seen in the signaling molecules
that regulate selective expression of Eph-B4 (receptor) in veins and ephrin-B2
(ligand) in arteries, creating a cell-cell interaction system that establishes arterial
and venous endothelial identity during vasculogenesis (179). Other differences
with significant potential for influencing specific vascular function have been
identified through the use of gene arrays (1, 24, 25, 112). Although most of these
studies focus on the endothelium and the vascular smooth muscle layers of
these blood vessels, fibroblasts and other components of the adventitia could
also play important roles in vascular function and pathogenesis. Our studies on
whole genome expression profiling of rat aorta and vena cava whole tissues
have indicated a potential anti-inflammatorylanti-apoptotic profile of venous

compared arterial tissue (145).

Several arterio-venous differences in properties may also be explained by the
differences in the characteristic environment each of these blood vessel types is
exposed to. The most obvious of these factors is the higher pressure regime
experienced by arteries compared to veins. The blood flow and the resulting
shear stress are also considerably higher in arteries compared to veins. Venous
blood, carrying metabolites and lymph products, is also less oxygenated than the

arterial blood, and characterized by a lower pH (43).

Tunica intima
Tunica media

 

 

Tunica adventitia

 

 

 

 

Carry blood away from
the heart to organs
and tissues

 

 

 

Figure 1. Diagram of generic artery and vein.

 

 

 

Carry blood away from
organs and tissues
back to the heart

 

 

smooth muscle

endothelium

 

Figure 2. A. Modified trichrome Masson staining of rat aorta (left) and vena cava
(right). B. Immunohistochemical staining for alpha-actin in rat aorta (left) and
vena cava (right). Blood vessel layers are indicated by arrows. The luminal side
is indicated by L.

C. Pathology

There are overall differences in the venous response to pathophysiological
conditions that commonly affect arteries, such as atherosclerosis or
hypertension, when veins do not undergo the same atheromatous, remodeling or

endothelial dysfunction changes observed in arteries (43, 168).

Almost all vascular research currently performed is focused on arteries, while
veins are generally overlooked when considering the contribution of the vascular
system to systemic pathophysiology. Yet, besides being a passive clearance
conduit of every organ and tissue, veins hold most of the circulating blood
(~70%). This volume can be actively moved by venous contraction or external
compression (121), making veins a potential contributor to the pathophysiology of
hypertension (detailed below). Venous grafts are used in coronary artery bypass
graft (CABG) surgery with different properties and outcomes compared to
arteries (101, 132). Additionally, veins also develop specific pathologies, such as
deep vein thrombosis and chronic venous insufficiency. Although they do not
constitute the particular focus of this work, all these facts point out the need for a

better understanding of the basic physiology of the venous system.

II. The role of veins in blood pressure regulation and hypertension

A. Hypertension epidemiology

Cardiovascular disease (CVD) is the leading cause of death in USA and globally
for both women and men, with an estimated 17 million deaths/year worldwide
(93). In USA, each year, CVD causes more deaths than the next four leading
causes of death combined (cancer, accidents, chronic lower respiratory diseases
and diabetes melittus). Examples of CVD include coronary heart disease,

cerebrovascular disease, hypertension and congestive heart failure (89).

Hypertension (high blood pressure) is defined as a systolic blood pressure (SBP)
higher than 140 mmHg and/or a diastolic blood pressure (DBP) higher than 90
mmHg. Currently, hypertension afflicts approximately a quarter of the adult
population (93). Due to the health risks associated with high blood pressure, a
condition called “pre-hypertension” was recently defined as a SBP between 120-
139 mmHg and/or a DBP of 80-89 mmHg. In the US, approximately 37% adult

population is prehypertensive (89).

Hypertension contributes to CVD not only by itself but also as a major risk factor
for other CVDs, such as coronary artery disease, heart failure, peripheral artery
disease and stroke. Thus, half of the patients with a first heart attack and two
thirds of those with a first stroke are hypertensive. Although largely
asympthomatic, hypertension — “the silent killer” — can lead to serious
complications of target organs (heart, brain, kidney, eye, peripheral arteries). In
USA alone, the total estimated costs for high blood pressure in 2009 are $ 73.4
billion (89). These statistics have remained almost unchanged in spite of
research efforts that have resulted in significant progress in hypertension therapy

in the last decades. The cause of hypertension is unknown in around 90% of

8

cases (essential hypertension), most likely a multitude of hereditary as well as

environmental factors contributing to its occurrence.

B. Mechanisms of blood pressure regulation

Blood pressure (BP) is a highly regulated physiological parameter, its control
being ensured by many complex systems and organs, such as the autonomic
nervous system, the kidney or the vascular system (fig 3). BP is a direct product
of two major cardiovascular parameters: cardiac output (C0) and total peripheral
resistance (T PR), an increase in either one or both leading to an increase in BP.
The vascular system plays a major role in this equation by changing its tone
(contraction) or structure (remodeling). Thus, the arterial side is responsible for
increases in TPR, while the venous side influences the CO. CO is also critically
influenced by the contractility of the heart and by the volume of blood, in its turn

controlled by the kidney.

C. Role of veins in hypertension

In most cases of established human hypertension there is an increase in TPR
without changes in the C0. Therefore the prevailing opinion in the field is that
hypertension is caused by changes in the tone and/or structure of small arteries
and arterioles, and the majority of hypertension research is being conducted on

renal, autonomic nervous system and arterial mechanisms of increases in TPR

(144). However, more recent reports point to the potentially critical role of veins
during the developmental stages of hypertension (39, 96, 111, 120, 174). The
vascular system can be viewed as a fluid holding system, the pressure of which
will be determined by both the total volume of fluid contained and by the
distensibility, or the fluid holding capacity of the system. With no changes in
blood volume or myocardium contractility, a key factor that determines C0 and
BP is the vascular capacitance, or the blood holding capacity of the vascular
system. Due to the structural differences discussed above, veins have a much
higher compliance compared to arteries (30 times higher), resulting also in a
much higher capacitance compared to arteries. Thus, ~70% of the total
circulating blood volume resides in veins, particularly veins in the splanchnic
region. The smallest changes in venomotor tone may therefore have a large
impact on this volume of blood (106), which thus shifted, is redistributed to the
central, less distensible, compartment of circulation (composed of larger veins
such as the vena cava), which then directly affects the “effective blood volume”
or the volume of blood contained in the arterial circulation, with immediate effects

on blood pressure (38, 91, 126).

There are several lines of evidence supporting the role of veins in hypertension.
Borderline, as well as early human hypertension (initial stages), are correlated
with increases in central blood volume and/or C0, as well as decreased venous
compliance (90, 91, 126). Venous capacitance is also decreased in the
established stage of human hypertension (90). Additionally, increases in mean

circulatory filling pressure (MCFP), which is a gross measure of venomotor tone

10

 

 

 

 

 

932:7. ncm
6:8. 26sz £095. m30cm>

 

 

 

 

 

 

 

 

  
 

L.

5688 cm_:omm> mE:_o> nooE

_ \

 

 

 

 

 

 

 

 

 

 

23386 95.: 3823.5

 

 

2

53m: mzocm> 3:38.28 E29835.

2 .\

oE:_o> 9.05m 32 tam...

/ \

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

8:82am ncm
2.3 26sz £095. _m:mt<

\

 

 

Sago 8:88

 

 

 

 

8:398. 3859.3 .38

 

 

/ \

 

2:365 uooE _m_cmt<

 

 

 

Figure 3. Diagram of blood pressure regulation systems and organs. The

potential places of modulation by reactive oxygen species are indicated with

dashed arrows.

11

and a direct determinant of CO at a constant blood volume, were observed
during established mineralocorticoid hypertension in rats (39), in the
developmental stages of hypertension in the spontaneous hypertensive rat (96),
as well as in Goldblatt renal hypertensive rats (174). These changes appear

causative, rather than adaptive, since the increase in MCFP preceded that in BP.

These data further support that the hemodynamic impact of venoconstriction on
blood pressure may be important. Thus, changes in venomotor tone in the initial
phases of hypertension may lead to a dynamic redistribution of blood that
triggers an increase in BP. Subsequently, arteries adapt to the new pressure
regime by remodeling towards a less elastic, more muscular and fibrous
structure, with increased TPR, that then drives the sustained increase in blood

pressure.

III. Vascular reactive oxygen species (ROS) metabolism
A. ROS chemistry

R08 are important vascular signaling molecules or mediators of oxidative stress.
They are, by definition, highly reactive intermediates of oxygen metabolism,
constantly being generated and destroyed by both environmental and
endogenous systems. Produced by a gradual reduction of molecular oxygen,
ROS include both unstable free radicals (chemical species having unpaired
electrons in their outermost shell) like the superoxide or the hydroxyl radical, and
longer-lived non-free radical oxidants like hydrogen peroxide (H202) (fig 4). A

12

88QO some? 9269...: - mzm

2556.8 - .0020

8:6 03E .. 02

20m 3925an - .001

.862 axes? - :0.

o :8ch ammo. E . Nof
vcoEm maxocwasm - ..NO ._0 wzm

comes Ease - No. x/

combs 5.329: - NO

        

.001

03320385? .
+I + -m NI +1 + -w ._0 .+I +IN + -0 OZ -0

O
of IQK Nof . K mo r K No

 

 

 

8:82 :95“. 88:98am EeEcgém
:0..an $_e>>¢8ax 883.5% £56.85 £20 boasame 6:26:02:
mommcongBomano

«Seneca 2.25520
8....st

Figure 4. Basic diagram of reactive oxygen species metabolism.

13

related family of reactive intermediates bearing importance in vascular biology is
the reactive nitrogen species (RNS), including nitric oxide (NO) and the series of

radicals produced after the interaction of the latter with the superoxide anion.

The delicate balance in the continuous cycle of ROS generation and inactivation
is maintained by enzymatic and non-enzymatic endogenous systems. We have
previously reviewed the subject of ROS metabolism in arteries and veins (146).
The various producers and destroyers of ROS are listed (table 1), but only the
most comprehensively studied ones will be detailed in the following section. Their
order does not bear any reflection of their relative importance, overall or

specifically in vascular tissues.

B. Cellular mechanisms of ROS production
1. Mitochondria

The mitochondrial respiratory chain is the main energy source for the cell.
Situated in the inner mitochondrial membrane, it catalyzes electron transfer using
over 80 peptides organized in four complexes. The transfer of electrons, shuttled
by coenzyme Q and cytochrome C, usually leads to the formation of ATP by a
fifth complex. However, a certain amount (1-2% in vitro) of electrons leak (40),
principally from complex Ill but also from complex I, generating superoxide (134,
138). The rate of mitochondrial ROS production, the levels of mitochondrial DNA
oxidative damage and the degree of membrane fatty acid unsaturation
(potentially a target of lipid peroxidation by R08) are all inversely linked to

14

 

ROS producers

ROS destroyers

 

Mitochondrial respiratory chain
Nox and Duox families

Xanthine oxidase

Superoxide dismutase family
Uncoupled nitric oxide synthase
Monoamine oxidase

Lipoxygenase

Cyclooxygenase

Cytochrome P450

Haber-Weiss and Fenton reactions
Environment (ionizing radiation, etc)

NO (reactive nitrogen species)

 

Superoxide dismutase family

Catalase

Glutathione system

Selenoproteins (glutathione peroxidase,
thioredoxin reductase, etc)

Peroxiredoxins

Antioxidant ROS scavengers (A,C, E

vitamins, ceruloplasmin, ubiquinone, uric acid,

bilirubin, etc)

N0

Uncoupling proteins (?)

Table 1. Reactive oxygen species sources and degradation systems.

15

maximum longevity in animals (5). These facts are among the evidence that
supports the free radical theory of aging. Since superoxide production is directly
dependent on the proton motive force, a feedback mechanism has been
proposed for the uncoupling proteins (UCP 1, 2 and 3). Activated by superoxide
and lipid peroxidation, these proteins seem to act by slightly reducing the proton
motive force and hence energy production as a trade-off for a decreased ROS

production from the mitochondrial complexes I and Ill (12, 30).

Several lines of evidence indicate mitochondria may be critically involved in
cardiovascular disease pathogenesis, by altered mechanisms related to
mitochondrial mediated ATP generation, apoptosis and, more recently, calcium
signaling (34). However the link between mitochondrial-mediated ROS
production and cardiovascular disease is still a subject of investigation.
Angiotensin II, famously stimulating superoxide generation through NADPH
oxidase with many related important cardiovascular implications, may also
activate superoxide release from mitochondria (73, 181). Potentially important
vascular roles of mitochondrial ROS are highlighted by reports such as those
suggesting that the protective effect of estradiol on cerebral vasculature
endothelial function is mediated by a decrease in mitochondrial ROS production

(26).

2. NADPH oxidases

The Nox family of NADPH oxidases is another major source of ROS. The classic

example is the phagocytic NADPH oxidase, a multisubunit enzyme involved in

16

host defense. The phagocytic NADPH oxidase is composed of two membrane-
bound catalytic subunits, Nox2 (formerly known as gp91p“°") and p22°“°", forming
the central flavocytochrome b553, and four cytosolic regulatory subunits, p47""°",
p40""°", p67p“°" and Rac (29). NADPH oxidase requires for its activation a series
of phosphorylation and translocation events, triggered by pathogen recognition
(22). Deliberate generation of ROS by the professional phagocyte during the
“oxidative burst” is a rapid and powerful weapon of defense against pathogens. A
genetic lack of NADPH oxidase activity in patients suffering from chronic
granulomatous disease (CGD), a condition characterized by recurrent, life-
threatening infections, illustrates the importance of the beneficial side of ROS
chemistry (29). Based on the homology with Nox2, several other members of the
human Nox family have been identified, each of them seemingly having different
activation requirements and expression patterns. Nox1, Nox3 and Nox4 are
more similar in structure to Nox2 and they all require at least p22""°" for
activation. Duox1 and Duox2 are Cay-activated dual oxidases with a C terminal
NADPH oxidase domain and an N terminal peroxidase domain. Nox5 is closer to
the Duox1/2 in structure and is also Ca2"-activated, but lacks the peroxidase

domain (140).

Several features of Nox enzymes expressed in blood vessels, that distinguish
them from the generic phagocyte NADPH oxidase (129), have made researchers
in the field collectively term them “the vascular oxidase”. Compared to
superoxide production from the phagocyte NADPH oxidase, vascular oxidase

basal superoxide production is significantly lower (less than 1% (59)). While

17

phagocyte NADPH oxidase activity is primarily inducible, vascular oxidase has a
constitutive activity that can be further increased by agonists like angiotensin II
(124). Although not fully clarified, it appears that Nox4 is the subunit responsible
for the constitutive vascular oxidase activity. The cellular site of superoxide
production by vascular oxidase appears to also be different: vascular oxidase
produced-superoxide has been repeatedly detected intracellularly (84). Finally,
the physiological role of superoxide production by the blood vessel cells is
distinct: instead of cytotoxic superoxide production as a defense mechanism
against pathogens, ROS released by the vascular oxidase participate in cell

signaling, consistent with their comparative low tissue levels (84).

Various endogenous and external stimuli modulate the NADPH oxidase subunits
expression and/or activity. Expression of one or more of these subunits is
upregulated in human umbilical vein endothelial cell (HUVEC) culture in
response to angiotensin II, ET-1, oxidized LDL, pulsatile shear stress and
phorbol myristate acetate (PMA) (27, 84, 95). They are conversely
downregulated by treatment with statins, PPAR agonists or estradiol (84, 123).
Angiotensin II, TGFB, TNFoc, serum, PDGF, PGF2a, PMA and LDL upregulated
various NADPH oxidase subunits expression in the case of cultured arterial
smooth muscle cell (84, 85). Long term treatment with AT1 receptor blockers

downregulated Nox2 mRNA expression in human artery biopsies (124).

In hypertension, the already increased vascular superoxide generation is further
increased with NADH or NADPH and lowered through treatment with the NADPH
oxidase inhibitor apocynin (6, 87). Aortic mRNA expression of p22'°“°" is

18

increased in the DOCA-salt (10) and SHR (180) models of hypertension. The
angiotensin II infusion hypertension model has increased expression of all
NADPH oxidase subunits (100). In the same model, inhibition of NADPH oxidase
activity by treatment with gp91 ds-tat, a chimeric peptide that blocks the

phox

association of p47 with Nox2, lead to a decrease in superoxide production

and attenuated the Angll-induced BP elevation (119).
3. Xanthine oxidase

Xanthine oxidoreductase (XOR) is an evolutionarily conserved housekeeping
gene, encoding for an enzyme that catalyzes the last steps of purine metabolism
(fig 5): the transformation of hypoxanthine and xanthine to uric acid, with
superoxide generated as by-product. XOR is a flavoprotein that possesses one
molybdopterin, two iron-sulfur groups and one FAD, functions as a 145 KDa
homodimer, and has ~90% homology from mouse to rat to human (11). There
are two isoforms of XOR, each of them utilizing different electron acceptors:
xanthine dehydrogenase (XDH), which requires NAD“; and xanthine oxidase
(X0), which requires molecular oxygen. XDH is convertible to X0 by reversible
sulfhydryl oxidation or by irreversible proteolytic modifications (105). Although
both isoforms have ROS generating potential, in vivo X0 is by far the more
important superoxide/H202 source, making XDH to XO conversion in situations
such as ischemia/reperfusion or inflammation of pathophysiological significance
(53). X0 and its potential pathophysiological role in cardiovascular disease are

discussed in greater detail below.

19

ATP

I

AfP
AMP deaminase

 

 

IMP < AMP GMP
, l Nucleotidase iNuc/eotidase
Adenosine Guanosine
. . Pun'ne nucleosr'dase
Purine nucleosidase Adenosrne deaminase p hosp horylase
phosphorylase _ _
> lnosrne Guanine

 

     
 
 

Inosine phosphorylase
BIN/I; Hypoxanthine _ .
NAD+ 02 Guanine deaminase
XDH XO
NADH 02--
:H> Xanthine
0;:NI / NAD‘ 02
XDH X0
ADH 02-.
HN NH Unc acrd
A I : o
o N N“ Un'case
H
O
HN Allantoin

044” NH
H )\
NH2

Figure 5. Purine catabolism. Structures of hypoxanthine, xanthine, uric acid and
allantoin are displayed on the left.

20

4. Nitric oxide synthase (NOS)

NOS, the enzyme responsible for N0 generation, has three isoforms: NOS1 (the
neuronal NOS), N082 (the inducible NOS) and N083 (the endothelial N08). In
physiological conditions, N08 catalyzes the transformation of L-arginine into L-
citrulline and NO, using several cofactors: NADPH, FAD, FMN and 5,6,7,8
tetrahydrobiopterin (BH4). However, if the enzyme is depleted of 8H4 or of L-
arginine, it becomes uncoupled, and transfers electrons to molecular oxygen
rather than the substrate L-arginine, producing superoxide (41). Furthermore,
interaction of superoxide with nitric oxide generates peroxynitrite, the second in
the family of reactive nitrogen species, capable of producing a cascade of

deleterious effects through oxidation, nitration and nitrosation of molecules (8).

The classical view on nitric oxide synthase isoforms is summarized in their
alternative names: the neuronal NOS (N081), and the endothelial NOS (N083)
have constitutive expression in neurons and endothelial cells, respectively, while
the inducible NOS (N082) is the only calcium independent, transcriptionally-
regulated isoforrn, typically found in macrophages (128). This paradigm has
changed considerably in recent years: all three isoforms have been identified in
arteries and veins, as well as in HUVEC culture. Furthermore, red blood cells
appear to express a membrane associated N083, capable of modulating
vascular tone (75). Recent studies have even suggested a mitochondrial-specific
N08 (80), although its existence has not been proven yet. To our knowledge, no

study has yet compared arteries and veins in terms of NOS isoforms expression.

21

Normal endothelial function, crucial in maintaining cardiovascular homeostasis,
depends, among others, on normal NOS functioning. A reduction in the arterial
endothelium-dependent vascular relaxation, defined as endothelial dysfunction,
has been documented in atherosclerosis and hypertension. Besides decreased
N0 bioavailability, a malfunctioning NOS can also influence vascular function by
becoming uncoupled, in the absence of L-arginine or BH4, with consecutive
production of superoxide and potentially peroxynitrite. Further uncoupling can
occur by oxidation of the existent BH4 (127). The role of BH4 depletion in N08
uncoupling and hypertension development is illustrated by the fact that treatment
of DOCA-salt hypertensive mice with BH4 leads, by recoupling of N08, to
lowering of BP (82). Uncoupling of N08 does not occur in p47p"°""' mice,
supporting the idea that NADPH oxidases are required for BH4 oxidation. There
is no clear evidence indicating that N08 uncoupling follows the same rules in

veins as it does in arteries.

C. Cellular mechanisms of ROS destruction
1. Superoxide dismutases (SODs)

A central role in the regulation of ROS levels is attributed to superoxide
dismutases, a family of enzymes responsible for superoxide breakdown, with the
resultant production of H202. This othenrvise spontaneous dismutation reaction
(at rates dependent on superoxide concentrations) is significantly accelerated by

SOD, the enzyme with the highest turnover number from all characterized

22

enzymes. There are three known SODs: the cytosolic CuZn SOD (80D1), an
unusually stable homodimer, the mitochondrial Mn 80D (80D2), functioning as a
tetramer, and the extracellular EC-SOD (80D3),’a tetramer with a C terminal
heparin binding region. There is a great body of evidence supporting the
beneficial role of SOD (31, 65, 94). Knock-out experiments showed neonatal
lethality of mice lacking Mn SOD and reduced lifespan and multiple abnormalities
in mice lacking CuZn SOD. Furthermore, overexpression studies of 80Ds
strongly suggest a protective role of these enzymes in many diseases as well as
in aging. Additionally, mutations in the 80D1 gene leading to the production of a
toxic variant of CuZn 80D are linked to 20-25% cases of familial amyotrophic
lateral sclerosis (FALS or Lou Gehrig’s disease), a fatal neurologic condition

(114).

The blood vessel wall of both arteries and veins expresses all three SODs. The
cytosolic CuZn 80D has ubiquitous and high expression throughout the vascular
layers. Mitochondrial Mn SOD is relatively less expressed compared to CuZn
80D and EC-SOD, but is also ubiquitous (31). Extracellular 80D, produced
largely by VSMC, is localized between arterial intima and media (139) and is
thought to contribute substantially to the total SOD activity in the vasculature. In
addition to its primary and important role in scavenging extracellular superoxide,
EC-SOD may also be expressed intracellularly and translocated to the nucleus
via its heparin-binding domain, which could function as a nuclear localization
signal (NLS) (109). Rats have lower vascular EC-SOD levels compared to other

species, due to a change in the amino acid sequence of the protein that leads to

23

lower heparin binding, potentially influencing the results of SOD expression

studies performed in this species (31).

Arterial expression and/or activity of CuZn SOD and Mn 80D increased in animal
models of hypertension (160, 161). EC-SOD deficient mice had higher blood
pressures in two hypertension models compared to the wild type animals (67).
Conversely, overexpression of EC-SOD improved vascular function in

hypertensive animals (17).
2. Catalase

Catalase is a homotetrameric heme-containing enzyme that catalyzes the
conversion of H202 into water and oxygen with one of the highest turnover rates
known in enzymology (~107 l/mollsec). Catalase is usually found in peroxisomes,
cellular organelles involved in multiple metabolism pathways, where it functions
in detoxification of H202 resulted from SOD action on superoxide (155). If not
otherwise inactivated, H202 may continue the ROS cascade with the formation of
the hydroxyl radical. The latter, generated through a reaction with transition
metals such as Fe2+ via the Fenton/Haber-Weiss chemistry (fig 3), is a highly
reactive radical that to our knowledge cannot be destroyed enzymatically. The
only protection from its dangerous oxidative potential is therefore left to

antioxidant scavengers and metal chelators.

Catalase, as well as other R08 enzymes, has been correlated with aging. There
is not a clear association between catalase function and cardiovascular disease.

Catalase mutations have been linked with hypertension (182). Mice

24

overexpressing catalase have shown evidence of cardioprotection from
adriamycin-induced ischemia-reperfusion (58), as well attenuation of BP
increases in response to norepinephrine and angiotensin II (176). However,

catalase null mice do not have an obvious cardiovascular phenotype (58).
3. Other ROS degrading systems

Another R08 consuming system is the glutathione redox cycle. Glutathione
peroxidase transfers electrons from the reduced form of glutathione to H202 with
the formation of water and oxygen; subsequently, the oxidized glutathione
disulfide is reduced by glutathione reductase. There are other selenoproteins
with similar activity to glutathione peroxidase, such as thioredoxin reductase or

selenoprotein P, all of which work as antioxidant enzymes.

Because of their widespread therapeutic use as antioxidants, endogenous ROS
scavengers such as vitamin C and E should also be noted. However, when
considering the antioxidant properties of such compounds, it should be
appreciated that they are not enzymes, therefore a new molecule is needed for
each superoxide anion that is scavenged. These vitamins are therefore poor
R08 scavengers. Clinical trials testing antioxidants on cardiovascular disease
patients have found no evidence of beneficial effects on BP or other common
cardiovascular end-points — the so-called “oxidative paradox” (76). However,
these trials were conducted with orally-administered, weak ROS scavengers only
active on superoxide, but not on H202. Numerous other factors (such as the

insufficient doses or their unknown intracellular concentration and activity) have

25

been overlooked. Most importantly, these trials included patients with chronic
cardiovascular disease, where increased R08 could already be a disease effect

rather than a pathogenetic factor.

IV. R08 involvement in vascular function

ROS mediate effects in all three blood vessel layers. At the endothelial level,
superoxide directly interacts with NO and decreases its bioavailability, while the
resultant peroxynitrite may oxidize BH4 leading to eNOS uncoupling, further
decrease in NO levels and increase in superoxide production (8, 127). At the
smooth muscle level, both superoxide and H202 may interfere with K+ and Ca2+
channel function, as well as with signaling pathways that modulate vascular
contraction: MAPK, PI3 kinase, and Rho kinase (2, 13). Additionally, ROS may
stimulate signaling pathways that mediate the expression of pro-inflammatory
molecules (such as NF-KB), as well increase the proliferation and migration cf
vascular smooth muscle cells (VSMCs), properties that become relevant in the
case of cardiovascular disease such as atherosclerosis and hypertension (18,
157). In the adventitia, ROS induce collagen production, as well as
metalloprotease activation, both these effects contributing to the arterial
remodeling observed in hypertension (54, 157). A summary diagram of vascular

effects of R08 is presented in figure 6.

26

        
   

3,3330% 0.:2 9.25.8.9.

+ .23 + and

USES
E2325
oEmmEoBm
85

22:85 -Nmuw ......

cozumzcou

ozuom

. ....u-x

  

 

 

 

     
         

     

eaten. ommumcamoca v.32

 

 

I “L34

:8

Become
5:898 6:88:66 3: m _m>mo_
cozoaooa 9:295 5.0m \_m>_>5m co_mmm.axo $30208 coaomzcoo ...o _. b 02
3323 33852.99: 239“..an bofiEEmchE . m - cozoca szo

+
E” / ...a/ n . rd;
GE“— Q>U< 05m> E=__¢:uoucm
/ mom

Figure 6. Diagram summarizing the effects of ROS on vascular function. +, -

respectively. The mechanisms responsible

for the vascular contraction in response to agonists relevant to this work (such as

norepinephine, angiotensin II or endothelin-1) are represented below.

represent stimulation and inhibition

27

V. Comparisons of arterial and venous ROS metabolism

Few studies have compared basal ROS production in arteries and veins, and
their conclusions were contradictory. Basal superoxide production, measured
through nitroblue tetrazolium reduction to formazan, was increased in porcine
venous grafts compared to arterial grafts (132). In contrast, using lucigenin-
enhanoed chemiluminescence, no difference in basal superoxide production was
found in rings from human internal mammary artery (IMA) compared to
saphenous vein (HSV) (51). These were however blood vessels from patients
with cardiovascular disease, and from two different vascular beds. The increase
in superoxide release, following the addition of the NOS inhibitor, L-NMMA, was
greater in human arteries (IMA) compared to veins (HSV) (52). This suggests a
greater basal NO production in arteries that contributes to the quenching of
superoxide in comparison to veins. Accordingly, basal peroxynitrite formation

was higher in IMA compared to HSV (52).

Besides this quantitative difference in ROS production between arteries and
veins, there is probably also different utilization of ROS by arteries and veins,
both in physiological cell signaling and in oxidative stress during vascular
pathogenesis. For instance, H202 modulates vascular tone, acting as a
contraction-inducing agent in some vascular beds and as a relaxant in others
(44, 150). There is a greater contraction to H202 in veins compared to arteries,

possibly reflecting a difference in K+ channel activity and Ca2+ influx (150).

28

Only one research group has systematically compared R08 sources in human
arteries and veins (Guzik et al (50-52)). These studies have shown that subunits
of NADPH oxidase (p22”"°", p47p“°" and p67°“°" proteins as well as p22°“°" and
Nox2) mRNA are present and increased in abundance in the human saphenous
vein (HSV) compared to the internal mammary artery (IMA). Nox4 mRNA
expression was higher in the IMA and Nox1 mRNA had similarly low expression
in both types of vessels. In a study employing the use of specific chemical
inhibitors, NADPH oxidase contribution to the total basal vascular superoxide
production was found to be more important in the case of HSV compared to the
IMA, while xanthine oxidase seemed to have a more significant contribution in
the case of IMA. No significant difference was found between these vessels in
terms of their CuZn SOD (cytosolic SOD isoform) and Mn SOD (mitochondrial

80D isoform) protein expression and activity.

Vl. Oxidative stress and role of ROS in hypertension

In certain amounts, R08 are normal constituents of cells, participating in
physiological signaling processes, the extent of which is yet to be determined.
However, increased ROS levels in the cell, resulting either from their
ovenrvhelming generation or impaired destruction, have a substantial impact on
normal cellular function. This imbalance between prooxidant and antioxidant
factors, defined as oxidative stress, can affect cellular homeostasis either

through direct oxidative damage of basic cellular components (proteins, lipids

29

and nucleic acids) or through the activation of various redox sensitive signaling
pathways leading to defective cellular function, aging, disease or apoptosis. ROS
involvement in cellular signaling has been reviewed extensively. In summary, a
series of major signaling pathways, such as MAPK, Pl3K/Akt, NF-KB, p53 and
the heat shock response can potentially be activated in response to R08 or
oxidative stress. In the vascular system, oxidative stress can impair function
through direct oxidative damage, endothelial dysfunction, decreased N0
bioavailability, impaired contractility, platelet aggregation and ROS mediated

inflammation, proliferation and remodeling (18, 54, 127, 157).

The presence of increased markers of oxidative stress (peroxidized lipids,
oxidized proteins, increased GSSG, 8-oxo-guanine, DNA breaks, etc) has been
identified in many pathophysiological situations. However, in most cases,
establishing whether oxidative stress plays a causal role or is a mere reflection of
the effects of the disease process itself on cellular function has proven to be a

difficult task.

As far as hypertension is concerned, a great body of evidence supports the idea
that R08 are involved in its pathogenesis. Increased markers of oxidative stress
are found in human hypertensive subjects as well as in various animal models of
hypertension (118, 149, 157). Treatment of these models with R08 scavengers
or SOD mimetics (57, 108, 173), inhibitors of NADPH oxidase (10, 119, 170),
inhibitors of xanthine oxidase (103, 163), BH4 (157), targeted gene delivery of
80D (35) or NADPH oxidase inhibitors (68, 170) normalizes BP or prevents the
development of hypertension and/or attenuates target organ damage.

30

Furthermore, genetic deficiency in ROS-generating enzymes protects some
animals from experimental hypertension (81), while lack of antioxidant capacity

causes increased hypertension in others (31, 149).

Vll. Xanthine oxidase: role in cardiovascular disease
A. X0 expression

X0 expression in blood vessels has been difficult to detect. Human small vessel
arterial endothelium showed X0 immunoreactivity (88), and X0 mRNA was
identified in cultured rat pulmonary arterial endothelial cells (28). Moreover,
measurable X0 activity has been detected in arteries and cultured endothelial
cells. However, X0 from the circulation can also bind to endothelial cells via
heparin binding sites (62). Therefore, there is still some controversy on whether
X0 is functional in the blood vessel or is only acquired through association with

blood.

B. X0 activity

X0 has previously been demonstrated as capable of generating R08 in normal
vascular tissues (9, 16). Increased X0 activity was observed in human
atherosclerotic arteries, as well as in several animal models of hypertension,
lending support to the idea of the contributing role of X0 in cardiovascular

disease. The spontaneously hypertensive rat (SHR) displayed increased renal

31

X0 activity during the development of hypertension (79). The same model
exhibited higher mesenteric artery X0 activity (141). Similarly, mesenteric artery
X0 activity was increased in the DOCA-salt model of hypertension (16). Both the
expression and the activity of X0 were increased in aorta of aging rats (104). The
increase in ROS production in cultured arterial endothelial cells in response to
shear stress has been attributed to X0 activity (98). Increased superoxide
production by X0 was observed in coronary arteries from patients with coronary

artery disease (136).

X0 effects in pathophysiology may be mediated by at least one other mechanism
that is not related to XO-mediated ROS production: the production of uric acid, a
molecule with controversial cardiovascular effects that can act both as an

antioxidant and as a deleterious by-product (see below).

C. Uric acid in vascular biology

Uric acid or urate (UA) is a small organic molecule formed as a result of XO
metabolism of xanthine. In most species, uric acid is further catabolized by
uricase (or urate oxidase) to allantoin, which is then readily excreted through
urine. Higher primates and humans, as a result of a mutation, do not express
uricase (178). In these species, uric acid is the final product of purine catabolism.
Uric acid has a very low solubility in water and can form crystals of monosodium
urate in certain conditions (high concentrations, acidic pH, presence of other

molecules) (135).

32

Therefore, uric acid was traditionally viewed as metabolically inert, yet capable of
exerting harmful effects as a result of crystallization, the most common
pathophysiological consequence of which is the development of arthritic gout
(135). Even in the absence of gout, studies have shown hyperuricemia to be a
risk factor for several cardiovascular diseases, associated with metabolic
syndrome and an independent predictor for kidney disease and all-cause

mortality (32, 66, 72).

Hyperuricemia can be experimentally induced in rodents by treatment with the
uricase inhibitor oxonic acid. This has lead to development of hypertension and
endothelial dysfunction, accompanied by renal fibrosis, increase in
juxtaglomerular renin and decrease in renal nitric oxide synthase (NOS)
expression, in the absence of crystal formation in the kidney (71, 97).
Additionally, in cultured bovine aortic endothelial cells, uric acid inhibited both the
basal and the vascular endothelial growth factor (VEGF)-stimulated NO
production (71). Taken together, these results suggest uric acid to be an overall

deleterious molecule.

However, there are several other lines of evidence pointing to the more
complicated and potentially beneficial roles of uric acid. In vitro, uric acid acts as
an antioxidant for superoxide, hydroxyl radical, H202 and other R08 (7).
lmportantly, these reactions occur at uric acid concentrations that are in the
range of low to normal human uricemia. The antioxidant effects of uric acid were
confirmed in various biological systems by studies demonstrating attenuation or

protection from oxidative stress in the presence of uric acid (7). These effects

33

might become especially relevant when considering the known correlation
between increased ROS and endothelial dysfunction (15). Uric acid may not only
react with superoxide, but also with peroxynitrite (resulted from the reaction
between superoxide and NO), decreasing the amount of peroxynitrite and giving
rise to a reaction product that has endothelium-independent vasorelaxant
properties (133). On the other hand, uric acid has been shown in vitro to be able
to react irreversibly with NO itself, quenching N0 in a manner similar to

superoxide (46).

In stark contrast with Khosla et al. (71), and giving support to the idea of a
beneficial role for uric acid, are studies performed in humans by Waring et al
(165, 166). Administering uric acid to patients with diabetes type I or smokers
with endothelial dysfunction lead to an increase in forearm blood flow in response
to acetylcholine (ACh), a measure of endothelial function, and a gross measure
of N0 bioavailability (166). Conversely, lowering uricemia in patients with
diabetes type II and associated endothelial dysfunction did not improve their

endothelial function (165).

To further complicate the image of uric acid roles in vascular biology, a urate
transporter (8LC22A12) was recently identified on the membrane of human
vascular smooth muscle cells (115). In this type of cells, uric acid was shown to
increase proliferation by increasing the expression of platelet-derived growth

factor (PDGF) (117) and activating the renin-angiotensin-system (20).

34

Thus it is unclear whether uric acid effects are overall beneficial or deleterious at
the level of the vascular system generally, and on endothelial function

specifically.

D. In vivo effects of X0 inhibition

The beneficial effects of in vivo treatments with X0 inhibitors constitute one of
the strongest pieces of evidence pointing to the role of X0 in cardiovascular
disease, and in particular, in hypertension. Various human and animal studies
have demonstrated beneficial effects of X0 inhibition in hypertension, coronary
artery disease or cardiomyopathies by lowering blood pressure, decreasing end

organ damage, improving endothelial function and heart function (110).

Pharmacological inhibition of X0 classically involves use of purine analogues,
such as allopurinol or oxypurinol. Allopurinol serves both as an inhibitor and as a
substrate (suicide inhibitor) for X0, which catalyzes its transformation to
oxypurinol. Allopurinol is also metabolized by the phosphoribosyltransferases
hypoxanthine-guanine phopshoribosyltransferase (HGPRT) and by orotate
phosphoribosyltransferase. Oxypurinol, the product of allopurinol metabolism,
has a longer half-life compared to allopurinol (22 h compared to approx 1-3 h for
allopurinol) as well as a lower K; (11). Another means of inhibiting X0 is by
tungsten diet. This approach however, leads to inhibition of all other
molybdenum-containing enzymes, such as aldehyde oxidase and sulfite oxidase

(74). More recently, febuxostat, a new inhibitor of X0 was approved for human

35

use for the management of gout. As it inhibits XO through a different mechanism,
febuxostat is devoid of non-specific effects that allopurinol or other purine
analogues may have, such as inhibition of purine nucleoside phosphorylase and
orotidine-5'-monophosphate decarboxylase, both also involved in nucleotide
catabolism (148). Unfortunately, no genetic models may be employed as a tool to

study of X0 functions, as X0 null mice die at week 6 (164).

Allopurinol treatment improved vascular function in diabetic rats (64), while
oxypurinol improved coronary and peripheral endothelial function in coronary
artery disease patients (3). Treatment with oxypurinol acutely decreased the
blood pressure in the SHR model, while it had no effect on the BP of normal
counterparts (103). Treatment with allopurinol (100 mg/kg/day) also decreased
systolic blood pressure (SBP) in the DOCA-salt hypertensive rat (163). A unique
clinical study seeking an association between allopurinol administration and BP
decrease has recently been published (33). Its authors observed a decrease in
SBP with allopurinol in newly diagnosed hypertensive adolescents that have

concomitant hyperuricemia.

Several other studies, however, have reported a lack of effects of X0 inhibition
on BP. Chronic treatment of SHR with allopurinol did not prevent the
development of hypertension in this model (78, 175). Allopurinol also could not
prevent development of glucocorticoid-induced hypertension (107) and L-NAME-

induced hypertension (69).

36

Thus the effects of X0 inhibition through allopurinol treatment on blood pressure

are overall mixed, appearing to be species and model-dependent.

37

ROS and oxidative stress are involved in cardiovascular diseases such as
hypertension. ROS effects on vascular function are varied and complex. Arterial
ROS production and the contribution of arterial oxidative stress to hypertension
have been extensively studied in experimental models of hypertension. However,
similar to other physiological characteristics of veins, little is known about the
venous generation and destruction of R08, as well as about their consequences
on venous function and how they might be different from the ones in arteries.
Although venomotor function may impact blood pressure regulation, it is unclear
whether alterations of this function mediated by R08 could contribute to

hypertension pathogenesis. We investigated this idea in the work described next.

Throughout this research, we used mostly the rat thoracic aorta and vena cava
tissues. They were chosen as model vessels to reflect the states of the arterial
and venous system respectively, as they are large enough to allow for
experimental manipulation in most protocols. Additionally, a large body of data
from literature already existed on these blood vessels, facilitating the progress of

this research.

The deoxycorticosterone acetate (DOCA)-salt model of hypertension was chosen
because it is a low renin, salt-sensitive (therefore vascular capacitance)-
dependent model, where increases in MCFP (thus venomotor tone) have been
observed. Moreover, both endothelin-1 (E T- 1 ), a potent venoconstrictor and R08
are known to play a role in this model. Furthermore, in this model, the increase in

BP occurs over a long period of time, allowing for mechanistic studies.

38

HYPOTHESIS

R08 metabolism and handling is different in veins compared to arteries.
Venomotor function, as influenced by R08, plays a role in experimental

hypertension.

Specific aims

1. Determine if there is a difference in ROS metabolism between arteries and

veins in basal conditions and in hypertension.

2. Determine if there is a difference in the impact of R08 on arterial versus

venous contractile function in basal conditions and in hypertension.

3. Determine whether venous function and blood pressure can be influenced

by changing ROS levels in vivo.

39

ROS sources 4-—> ROS destroyers

AIM 1 ROS metabolism

 

AIM 2 R08 function

venous -{chml
‘ ’. " i" 1r 1 "
l "U; . :Il ," .

, ,1 Smooth muscle contraction I; , x '

. i‘. I)“ ,, .I -
I . ¢ “ .. . V‘ ‘
: I: at ll 1.)."

In vivo effects

A'” 3 of ROS

Mean circulatory
filling pressure
T\>\\/\/I
é)

\~> Blood ressure/
b'/\\// \/ \3‘

Total peripheral resistance

Figure 7. Diagram of working hypothesis.

40

METHODS
I. Animal use

All animal procedures were approved and performed In accordance with
regulations of the Institutional Animal Care and Use Committee at Michigan State
University. Male Sprague-Dawley rats (225-250 g) were purchased from Charles
River Laboratories (Indianapolis, IN). Until euthanasia or surgical use, all rats
were kept in clear plastic boxes, with access to standard rat chow (Teklad®) and

tap/treatment water ad libitum.

A. Euthanasia

Rats were euthanized with pentobarbital (60-80 mglkg i.p.).

3. DOCA-salt hypertension

Rats were uninephrectomized under isoflurane anesthesia (0.8 Umin, to effect).
DOCA rats received a subcutaneous implant at the nape of the neck of a pellet of
200 mglkg deoxycorticosterone acetate (DOCA) and were given drinking water
SUpplemented with 1% NaCI and 0.2% KCI for a duration of 4 weeks. The sham
r ats were uninephrectomized (surgery control group) but were not implanted with
DOCA and were given tap water. Animals received analgesic (carprofen 5 mglkg
S.c., at the time of surgery and 2 days postop.) and antibiotic (enrofloxacin 10

rhglkg i.m. at the time of surgery) treatment. .
41

C. Blood pressure measurements
1. Tail-cuff measurement of systolic blood pressure

For in vitro tissue experiments performed as part of the first and second specific
aim, systolic BP (SBP) was measured at the end of the 4 week DOCA-salt
treatment to validate increase in SBP, using a standard tail-cuff method in the
SC1000 Single Channel System (Hatteras Instruments, Cary, NC). DOCA rats
SBPs were increased by at least 50 mmHg over sham SBPs (sham SBP ranging

from 100 to 120 mmHg, DOCA SBP ranging from 175 to 225 mmHg).
2. Radiotelemetric measurement of mean arterial pressure (MAP)

For in vivo experiments performed as part of the third specific aim, MAP was
measured through the use of radiotelemetry. Under isoflurane anesthesia (0.8
Umin, to effect), radiotelemeter devices (TA11PA-C40, Data Sciences
International, St. Paul, MN) with attached catheters with pressure-sensing tips
were implanted into a subcutaneous pouch in the abdominal wall through a 1-1.5
cm incision in the left inguinal area. Catheters were introduced into the left
femoral artery 3 to 5 mm distal to the level of the peritoneal wall, and the tip was
advanced to the abdominal aorta. Animals received analgesic (carprofen 5 mglkg
so, at the time of surgery and 2 days postop.) and antibiotic (enrofloxacin 10
mglkg i.m. at the time of surgery) treatment. Rats were allowed 3 days to recover
postoperatively, before the start of baseline recording. Parameters (systolic
pressure, diastolic pressure, mean arterial pressure, pulse pressure, heart rate

and activity) were recorded throughout the duration of the study, at a sampling

42

rate of 10 seconds each 10 minute interval (Dataquest ART 4.1, DSI). Data is

presented as mean 1 SEM of 24h-averaged parameters.

D. In vivo allopurinol administration

Allopurinol was administered orally at 50 mglkg/day mixed in the DOCA-salt
water, in the case of in vivo experiments for allopurinol effects on BP; and at 50
mglkg/day or 100 mglkg/day mixed in water, in the case of in vivo experiments
for dose-dependent effects of allopurinol on XO metabolites. Water intake was
monitored daily and allopurinol concentrations in the drinking water were varied

according to this intake, in order to keep the same desired dosage.

E. Sample collection

1. Tissue

At the time of sacrifice, tissues of interest were removed and gently cleaned of
outer adipose tissue and then used in one of the protocols described below.
Tissues used were: thoracic aorta, caudal vena cava (VC), carotid arteries,
jugular veins, superior mesenteric artery, superior mesenteric vein, heart, liver

and right kidney.

2. Urine

43

Urine samples were collected by temporarily housing rats in metabolic cages that
allow for separation of urine from feces and solid debris. Samples were collected
after 24 hour periods. The total volume of urine for 24 h was measured and used

for normalization.
3. Serum

Whole blood was collected into plastic tubes (non-heparin coated) by cardiac
puncture from deeply anesthetized rats (pentobarbital 60-80 mglkg i.p.). After 30
minutes at room temperature, clotted blood was spun (25009, 10 minutes, twice,
mechanically separating the clot from sides of tube if necessary between the two

centrifugations), and the supernatant (serum) was collected.

ll. Basal superoxide levels by lucigenin-enhanced chemiluminescence

measurements

Rings of aorta, VC, superior mesenteric artery and superior mesenteric vein were
cleaned of outer adipose tissue and incubated in a modified Krebs-HEPES buffer
(containing in mM: 20 HEPES, 119 NaCI, 4.6 KCI, 1.0 MgSO4-7H2O, 0.15
Na2HP04, 0.4 KH2PO4, 5 NaH003, 1.2 CaCl2, and 5.5 dextrose, pH=7.4) at 37°C
for 1h with diethyl-dithio-carbamate (DDC), a superoxide dismutase (SOD)
inhibitor. A TD 20/20 Luminometer (Turner Designs, Sunnyvale, CA) was used.
Lucigenin (5 pM, concentration previously shown to avoid redox cycling (102))
was added for 10 minutes, and 10 consecutive Iuminometer readings (each 30
seconds long) were taken. Tiron (4,5-dihydroxy-1,3-benzene disulfonic acid), a

44

superoxide scavenger, was then added for 15 minutes to assess background and
10 more consecutive readings were taken. Tissues were blotted dry and
weighed. The average of readings with tiron was subtracted from the average of
readings without tiron, and this difference was multiplied by 2 for a resulting
measure of luminescence/minute. This value was used as input for the following
equation, resulted from a standard curve of superoxide concentrations,

previously constructed with a cytochrome c reduction assay:

tum
min

weight (mg)

(5.95..

 

. )+o.1
superoxrde =

Results are presented as nmol superoxide/(min*mg tissue).

Ill. Hydrogen peroxide (H202) measurements
A. Basal H202 levels

H202 production was measured using an Amplex Red H202 assay kit (Molecular
Probes, lnvitrogen, Carlsbad, CA) according to manufacturer’s protocol. Aorta,
VC, carotid artery and jugular vein were cleaned of outer adipose tissue and
equilibrated for 1h at 37°C in the same modified Krebs-HEPES buffer (pH=7.4),
as detailed for superoxide measurements. They were then incubated with Ultra
Red working solution (100 uM) at 37°C for 1h. The supernatant was then
transferred to a 96 well plate and fluorescence emission was measured

(excitation=530 nm, emission=590 nm) on an Ascent Fluoroskan plate reader

45

(Thermo Fisher Scientific, Waltham, MA). For each experiment, a H202 standard
curve constructed on the same 96-well plate was incubated with Ultra Red
working solution (100 ttM) at 37°C at the same time with the tissues and was
used to determine H202 concentrations from samples (fig 8A). After each
experiment, tissue total protein was determined by the Lowry method and used

for normalization.

In other experiments, H202 measurements were made in the presence of the
superoxide dismutase inhibitor diethyl-dithio-carbamate (DDC), in which case the
latter was added at both incubation times (1h in modified Krebs-HEPES buffer

and 1h with Ultra Red), at a 10 mM concentration.

B. Antioxidant capacity for H202

Antioxidant capacity of aorta and VC was measured using the Amplex Red H202
assay as detailed above. Incubation of aorta and VC rings with Ultra Red working
solution (100 pM) was performed in the presence of 20 pM H202. H202 that was
quenched by tissue was determined by subtracting the fluorescence values of
samples incubated with 20 uM H202 from the fluorescence value of 20 pM H202
alone, and using the value obtained as input in the H202 standard curve. After
each experiment, tissue total protein was determined by the Lowry method and

used for normalization.

46

A- 200«

 

 

 

 

y= 7.762x+ 9.1662 ’.
150- R2 = 0.99708 , I
’I
E I”
m too~ .e’
’I
50- ,0’
.I
or I U 1 I I
O 5 10 15 20 25
3 1.5-
y = 0.0108X + 0.0075
R2 = 0.99923 .
1.0“ I’
s 1"
<5“ y’
I
0.5- , ’
I
I I
00-! r r . t . .
0 20 4o 60 80 100 120
uM uric acid

Figure 8. A. Representative standard curve of H202 for the Amplex Red H202
assay. Known concentrations of H202 are plotted against relative fluorescence
units (RFU) obtained at 530 nm excitation with 590 nm emission. B.
Representative standard curve of uric acid for the X0 activity assay of urate
production. Known concentrations of uric acid are plotted against absorbance at

292 nm.

47

C. Relative contribution of ROS metabolizing enzymes to basal H202 levels

The same protocol as for basal H202 production (lllA, detailed above) was used.
Inhibitors: apocynin (100 pM), allopurinol (100 pM), Nw—Nitro-L-arginine (LNNA,
100 pM), rotenone (10 pM), indomethacin (10 pM), ketorolac (10 mM),
diphenylene iodonium (DPI, 10 pM) were incubated with tissues for 1h at 37°C in
the modified Krebs-HEPES buffer, and again for 2h incubation at 37°C with Ultra
Red reagent. The 2h Ultra Red reagent incubation time was chosen as yielding
mid-range fluorescence signals from a time course study of H202 production
measurements in aorta, vena cava, carotid artery and jugular vein (30 min, 1h,
2h and 3h) (data not shown). The remainder of the H202 measurement protocol

was carried out as described above (lllA).

IV. Protein concentration determination
A. Bicinchoninic acid (BCA) protein assay

Protein measurements in the tissue homogenates used for western blotting were
made using the BCA assay (Sigma, St. Louis, MO) according to the
manufacturer’s protocol. Bovine serum albumin (BSA) protein standard was used
to construct a standard curve with known protein concentrations (fig 9A).
Samples and standards (5 pl q.s. to 100 pl with water) were incubated for 30 min
at 37°C with 2 ml working reagent (BCA:copper (ll) sulfate in a 50:1 ratio) and

analyzed by end-point spectrophotometry at 562 nm.

48

B. Lowry protein assay

Protein measurements in the tissues used for H202 measurements were made
using the Lowry method. Tissue samples were digested in 1M NaOH with
sonication to solubilize. Bovine serum albumin (BSA) protein standard was used
to construct a standard curve with known protein concentrations (fig 98).
Samples and standards (10 pl q.s. to 100 pl with 1M NaOH) were incubated for
10 min at RT with 1ml reagent A (sodium carbamate:KNa tartrate2CuSO4 in a
100:1:1 ratio), followed by addition of 100 pl reagent B (Folin-Ciocalteu phenol
reagentzwater in a 1:1 ratio) for 1h at RT. Samples were then placed on a 96-well

plate and analyzed by plate-reader end-point spectrophotometry at 750 nm.

C. Bradford protein assay

Protein measurements for tissue homogenates used for HPLC measurements
and the X0 activity assay for urate production were made using the Bradford
protein assay (Bio-rad, Hercules, CA) according to the manufacturer’s protocol.
Bovine serum albumin (BSA) protein standard was used to construct a standard
curve with known protein concentrations (fig 9C). Samples and standards (5 pl
q.s. to 100 pl with water) were incubated for 5 min at RT with 900 pl Bradford

working reagent and analyzed by end-point spectrophotometry at 595 nm.

49

 

 

 

 

 

 

0.25-
: 0.0102x + 0.0008
0.20- Y o
A' R2 = 0.99987 ,i’
N 0.15- ,1’
if x
0.10- ’1
’I
0.052 ’1
’0
0.0“ 1 I U I l
0 5 10 15 20 25
pg protein
400-
B.
300. y = 5.4286x + 43 ,o
R2 = I ’
s r ’ I
I
(e 200- ’ 1’
’ I
100- '
C I I 1
0 20 40 60
pg protein
C 2.5-
y = 0.0478x + 0.7193 ’ ,.
2.0‘ R2 = 0.9526 ’ I ’
I
. I
a 1.5- ’ , I ’
<t’ " ’
1.0‘ . ’
0‘
..4f
0.0 | I I I 1
0 10 20 30 40

pg protein

Figure 9. A. Representative standard curve of protein for the BCA protein assay.
Known concentrations of BSA are plotted against absorbance at 562 nm. B.
Representative standard curve of protein for the Lowry protein assay. Known
concentrations of BSA are plotted against absorbance at 750 nm. C.
Representative standard curve of protein for the Bradford protein assay. Known
concentrations of BSA are plotted against absorbance at 595 nm.

50

V. Western blot analysis
A. Protein isolation

Thoracic aorta and vena cava were removed from the animal, placed in
physiological salt solution (P88) containing (mM): NaCI, 130; KCI, 4.7; KH2PO4,
1.18; M98047H20, 1.17; NaHC03, 14.8; dextrose, 5.5; CaNa2EDTA, 0.03;
CaCl2, 1.6 (pH=7.2) and cleaned as described above. Tissues were quick frozen
in liquid nitrogen, pulverized with a liquid nitrogen cooled mortar and pestle and
solubilized in lysis buffer supplemented with protease inhibitors [0.5 M Tris HCI
(pH=6.8), 10% SDS, 10% glycerol, 0.5 mM phenylmethylsulfonylfluoride (PMSF),
10 pgl pl aprotinin, 10 pglpl leupeptin, 0.1 M orthovanadate). Homogenates
were centrifuged (11,0009 for 10 minutes, at 4°C) and supernatants were
collected and stored at -80°C. Protein concentration was determined using the

BCA assay (detailed above).

B. Western blotting

Thirty-five pg of protein from each sample and 10 pl positive control (rat kidney
medulla for X0, CuZn SOD, Rac1; rat kidney cortex for MnSOD and OxPhosl; rat
liver for 4-HNE and uricase; human endothelial cell Iysate for eNOS; and human
thyroid Iysate for DUOX1) were separated on SDS-polyacrylamide gels (7, 10 or
12%) and transferred to nitrocellulose, PVDF or Immobilon-FL membranes. The
membranes were blocked with 5% milk, 4% chick egg ovalbumin or Ll-COR
blocker for 3h at 4°C. They were then probed overnight at 4°C with specific
51

antibodies against X0 (Rockland, Gilbertsville, PA and Abcam, Cambridge, MA),
eNOS (BD Biosciences, San Jose, CA), Rac1 (Upstate, Millipore, Billerica, MA),
p47phox (Santa Cruz, CA), DUOX1 (Santa Cruz, CA), CuZn SOD (Calbiochem,
EMD Biosciences, San Diego, CA), Mn SOD (BD Biosciences, San Jose, CA),
catalase (Abcam, Cambridge, MA), 39 KDa subunit of OxPhos complex I
(Molecular Probes, Invitrogen, Carlsbad, CA), 4-HNE (Abcam) and uricase
(Santa Cruz, CA). The appropriate horseradish peroxidase-linked or fluorescent
secondary antibodies were added for 1h at 4°C, followed by visualization with
ECL or the infrared imaging scanner Odyssey (LI-COR, Lincoln, NE). To ensure
equal protein loading, gels were stained with Gel Code Blue (Pierce Chemical,
Rockford, IL), and blots were reprobed with smooth muscle o-actin antibody
(Oncogene, EMD Biosciences, San Diego, CA). Band density was quantified
using NIH Image version 1.63. Smooth muscle q-actin expression per equal
amount of total protein is significantly higher in aortic vs VC homogenates,
therefore unless comparing the same type of tissue (e.g. sham with DOCA aorta
samples), densitometry values were not normalized to d-actin, as this
manipulation would have skewed results biasing expression of proteins of

interest towards the VC.

Vl. lmmunohistochemistry of paraffin-embedded sections

Forrnalin-fixed, paraffin-embedded rat thoracic aorta and vena cava sections (8

pm) were washed twice in Histochoice Clearing Agent (Vector Laboratories,

52

Burlingame, CA) and 4 times in isopropanol for 3 min each to dewax. Rat liver
sections were used as positive control for 4-HNE staining. Sections were
unmasked by boiling them twice for 2 min each in Vector Antigen Unmasking
solution. Endogenous peroxidase activity was blocked (0.3% H202 in phosphate-
buffered saline, PBS) for 30 min. Sections were blocked for non-specific binding
by incubating for 30 min with competing serum (1.5% in PBS). In a humidified
chamber, sections were incubated overnight at 4 °C with competing serum
(negative control) or with specific antibodies against X0 (1:200, Rockland,
Gilbertsville, PA and 5 pglml, Abcam, Cambridge, MA), DUOX1 (1 pglml, Santa
Cruz, CA), CuZn SOD (5 pglml, Calbiochem, EMD Biosciences, San Diego, CA),
catalase (5 pglml, Abcam, Cambridge, MA), 4-HNE (1 pglml, Abcam). The
remaining steps were performed according to the manufacturer’s protocol (Vector
Laboratories, Burlingame, CA). Sections were washed 3 times in PBS and
incubated for 30 min with the appropriate biotinylated secondary antibody. After
washing 3 times in PBS and incubation for 30 min with Vectastain® ABC Elite
reagent, sections were exposed to diaminobenzidine (DAB)/ H202 and staining
was monitored and timed. Reactions were stopped by PBS. Sections were then
counterstained with hematoxylin, dried and mounted. Photographs were taken
using an inverted Nikon microscope with a Spot digital camera and the
MetaMorph software (Molecular Devices, MDS Analytical Technologies, Toronto,

Canada).

VII. mRNA expression

53

A. RNA isolation

Total RNA was isolated using the MELTTM Total Nucleic Acid Isolation System
(Ambion/Applied Biosystems, Austin, TX) with the magnetic stand (Ambion) for
96-well plates. Aorta and vena cava were quickly cleaned in chilled DEPC—
treated water. Sections of ~10 mg from each tissue were then cut into 4-6 smaller
pieces and placed in the MELT Buffer with MELT Enzyme mix and vortexed for
20 minutes at RT. Samples were then centrifuged (11,0009 for 3 min) and
supernatants were taken through the remainder of the protocol according to
manufacturer’s instructions. The optional DNase digestion step was performed
using Turbo DNase (Ambion). The final elution was carried out in 12.5 pl Elution

Buffer. RNA samples were stored at -80°C.

B. RNA quality and quantity assessment

Total RNA was quantified at 260 nm using a Nanodrop ND-1000
spectrophotometer (Nanodrop, Wilmington, DE). The purity of RNA was verified
by a 260/280 ratio of 2.0 :l: 0.25 and a 260/230 ratio of 1.8 :l: 0.15. The integrity of
RNA isolated from aorta and vena cava by the MELT method was initially verified
by electrophoresis of 1 pg RNA samples on agarose-MOPS-formaldehyde gels
and UV visualization of ethidium bromide stained bands. Sharp 288 and 18S
bands were observed in an approximate 2:1 ratio, with no signs of RNA
degradation. Since the approximate yield from one whole VC is 1 pg RNA, we did

not perform this analysis for RNA isolated for downstream procedures.

54

C. Reverse transcription

1. RT for real-time PCR

One pg total RNA was DNAse-treated (DNase I, Roche Applied Science,
Mannheim, Germany) and then reverse-transcribed using oligo (dT)12-18 primer,
dNTP mix and Superscript II reverse transcriptase according to the
manufacturer’s protocol (lnvitrogen, Carlsbad, CA). cDNA samples were stored

at -20°C.

2. RT for PCR arrays

To perform SuperArray PCR arrays (see below), equal amounts of total RNA
were reverse-transcribed using the RT’ First Strand Kit (SA Biosciences,

Frederick, MD) according to the manufacturer’s protocol.

D. Primers

Primers for rat xanthine dehydrogenase (XDH) (GenelD 29289, mRNA sequence
NM_017154) were designed using the Primer 3 software (122) (Whitehead
Institute, Cambridge, MA), and synthesized at the Macromolecular Structure,
Sequencing and Synthesis Facility at MSU; X0 forward 5’-
GCATGCCAGACCATACTGAA-B’; reverse 5’-AAATCCAGTI'GCGGACAAAC-3’.
Primers for DUOX1, DUOX2 and GAPDH were purchased from SuperArray (SA

Biosciences, Frederick, MD).

55

E. Real-time PCR

Relative quantification of X0 relative to GAPDH and of DUOX1 and DUOX2
relative to beta-2 microglobulin was performed using SYBR Green PCR Master
Mix (Applied Biosystems, Foster City, CA) and the respective primers (0.1 pM),
on a 7500 Real-Time PCR System (Applied Biosystems, Foster City, CA).
Baseline values were established on the amplification curve with fluorescence on
a linear scale during the initial cycles with no detectable amplification. The
threshold was established on the amplification curve with fluorescence on a
logarithmic scale, such that it crossed amplification curves in the lower 1 l3 of the
exponential amplification phase. A dissociation curve was performed at the end
of each run to ensure a single product was amplified. Average cycle threshold
(C.) values for the housekeeping gene were subtracted from average 0: values
for the gene of interest for a resulting AC. value. Quantification was not
performed using the typical AAC. quantification, as no tissue was considered
control in comparing aorta and VC mRNA expression. Data are reported as

mean :t SEM of 2'ACt for each tissue.
F. PCR arrays

The Oxidative Stress and Antioxidant Defense PCR array (SA Biosciences,
Frederick, MD) was used according to the manufacturer’s protocol. Aorta and V0
cDNA samples reverse-transcribed by the RT2 First Strand Kit were mixed with

RT2 qPCR Master Mix, added on the 96-well plates with pre-aliquoted primers

56

against oxidative stress genes (84 genes) and amplified using the 7500 Real-
Time PCR System (Applied Biosystems, Foster City, CA). Quantification of
mRNA expression was performed using the manufacturer Excel-based analysis
software, with beta-actin chosen for normalization from the six housekeeping

genes included in the PCR array.

VIII. Xanthine oxidase (X0) activity assays
A. X0 activity assay for urate production

Aorta and VC rings were cleaned in X0 assay buffer (50 mM K phosphate, 0.1
mM EDTA, 1 mM oxonic acid; pH=7.4), pulverized in liquid nitrogen, solubilized
in X0 lysis buffer (X0 assay buffer containing 10 mM DTT, 1 mM PMSF, 10 pgl
pl aprotinin, 10 pgl pl leupeptin, 0.1M orthovanadate), sonicated and centrifuged
(20,8009, 4°C, 30 min). Equal amounts of supernatant were then incubated with
no additions, with 100 pM xanthine, or with 100 pM xanthine and 100 pM
allopurinol at 37 °C for 30 min. Urate formation was measured
spectrophotometrically at 292 nm in quartz cuvettes. The amount of uric acid
produced was calculated using a uric acid standard curve (fig 88). Protein
concentration of homogenates was determined using the Bradford assay. X0
activity was reported as the allopurinol-inhibitable uric acid production,

normalized for protein.

57

B. XO activity assay for H202 production

X0 H202-producing activity was measured using the Amplex Red H202 assay as
detailed above for basal H202 production measurements (III A). Aorta and V0
rings were incubated with Ultra Red working solution (100 pM) in the presence of
100 pM xanthine, or in the presence of 100 pM xanthine and 100 pM allopurinol.
H202 concentrations from samples were calculated using a H202 standard curve
(fig 8A) and were normalized to the total protein content of samples determined
by the Lowry method. XO activity was determined by subtracting H202 production
of samples incubated with 100 pM xanthine (substrate) and 100 pM allopurinol
(inhibitor) from the H202 production of samples incubated with 100 pM xanthine

alone.

IX. Isolated tissue bath contractility

Rings of aorta and VC were removed and cleaned of outer adipose tissue in
physiological salt solution (PSS) containing (mM): NaCl, 130; KCI, 4.7; KH2PO4,
1.18; M98047H20, 1.17; NaHCO3, 14.8; dextrose, 5.5; CaNa2EDTA, 0.03;
CaCl2, 1.6 (pH=7.2). Rings were mounted in warmed (37°C), aerated (95% 02,
5% CO2) P88 in isolated tissue baths (30 or 50 ml) for measurements of
changes in isometric force (PowerLab, ADInstruments, Colorado Springs, CO).
Tissues were placed under optimal passive tension (previously determined as 49
for aorta and 19 for VC) and equilibrated for 1h with frequent buffer changes. The

initial contraction to 10 pM d-adrenergic agonist (PE-phenylephrine for aorta, NE-

58

norepinephrine for VC) was used to verify viability of tissues and normalize any
further isometric contractions. As previously described, the maximal contractions
induced by PE (10 pM) and NE (10 pM) are similar in aorta (169). However, PE
does not induce reproducible contractions of VC, and NE was used instead in
this tissue. Endothelial integrity was tested in each experiment by measuring
relaxation to ACh (10 pM) from the contraction to 20 pM PGF2a. ln inhibition
experiments, tissues were incubated in parallel with inhibitor (100 pM allopurinol,
100 pM LNNA, 10 pM DPI) or their vehicles for 1h prior to performing cumulative
concentration response curves to agonists (NE, Angll, ET-1 or ACh).
Concentration response curves to ACh (relaxation curves) were performed on
tissues contracted with 20 pM PGF2a. Cumulative concentration response curves
to uric acid (10'9-10“ M) were performed in the presence or absence of oxonic
acid (uricase inhibitor, 10 pM). In other experiments, after 1h incubation with uric
acid (100 pM) and oxonic acid (10 pM), allopurinol (XO inhibitor, 100 pM) or their
vehicles (NaOH and DMSO, respectively), tissues were contracted with 20 pM
PGF2GI and cumulative concentration response curves to ACh were constructed.
The contraction induced by PGF2a was not changed by the presence of uric acid,

oxonic acid, allopurinol or their vehicles.

X. Blood uric acid measurement

59

Arterial blood was collected by cardiac puncture at the time of sacrifice and uric acid
was determined using the UAsure meter and test strips (ApexBio, Taiwan) according

to the manufacturer’s protocol.

Xl. High performance liquid chromatography (HPLC)

Tissue samples (aorta, vena cava, carotid artery, jugular vein, liver, kidney, heart)
were collected at the time of sacrifice and cleaned in PSS as described above. They
were then quick frozen in liquid nitrogen and pulverized in a liquid nitrogen cooled
mortar and pestle. Homogenates were solubilized in 0.2 M acetic acid, sonicated
and centrifuged (14,0009 for 10 min at 4°C). Serum and urine samples were diluted
1:10 in 0.2 M acetic acid and centrifuged (14,0009 for 10 min at 4°C). Analysis of XO
metabolites from sample supernatants was performed using Waters HPLC (Waters,
Milford, MA) coupled with Photo Diode Array detection. Separation of analytes was
achieved on a Phenomenex Luna, 5 pm, C-18, 250 x 4.6 mm column. Mobile phase
was 0.02 sodium acetate (pH=4.5 with acetic acid). A gradient with methanol was
used to clear the column after elution of analytes. An example tracing demonstrating
separation of standards under these conditions is represented in figure 10. The
individual X0 metabolites were analyzed at their respective maximum wavelength
intensities (uric acid at 284 nm, xanthine at 266 nm, hypoxanthine at 249 nm,
allantoin, oxypurinol and allopurinol at 210 nm). Peak height values for each
metabolite obtained in the same retention time window as standards were used as

input into standard curves constructed with known analyte concentrations. X0

60

AU

 

 

 

I

3..
-) "3

y

l/

allantoin

 

uric acid 9 :2—

hypoxanthine —->

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

xanthine —>

(....
1?
L
allopurinol -t> /,.._~————
l
i

oxypurinol ->

 

T ‘ I Y Y I T Y

"l"'l"'l"
10.00 12.00 14.00 16.00 18.00

Figure 10. Representative high performance liquid chromatography (HPLC)
tracing of standards for allantoin, uric acid, xanthine, hypoxanthine, xanthine,
allopurinol and oxypurinol, as analyzed at

simultaneously.

61

254 nm to display all analytes

metabolites were normalized to total protein content of HPLC samples determined
by the Bradford method, with the exception of urine, for which XO metabolites were

normalized to the total urine volume for 24h.

XII. Data analysis

Data are presented as mean :l: SEM for the number of animals (N). Plotting and
statistical analysis of data was accomplished using GraphPad Prism 5 (GraphPad,
La Jolla, CA). When comparing groups, the appropriate Student’s t-test or ANOVA
analysis was performed. For in vivo experiments, a 2way repeated measures
ANOVA with Bonferroni post-hoc analysis was performed. For association between
uricemia and SBP, a non-parametric correlation (Spearman) analysis was

performed. In all cases, a p value of 50.05 was considered statistically significant.

62

RESULTS

Overall hypothesis: ROS metabolism and handling is different in veins
compared to arteries. Venomotor function, as influenced by R08, plays a

role in experimental hypertension.

Specific aim 1: Determine if there is a difference in ROS metabolism

between arteries and veins in basal conditions and in hypertension

Rationale: Associations have been described between increased ROS
production in arterial tissues and cardiovascular disease, although the precise
enzymatic sources of this increase and its pathogenetic role are still a matter of
debate. However, very little is known about ROS production and antioxidant
defense in venous physiology, as well as about the potential dysregulation of
venous ROS metabolism during diseases such as hypertension. We started this
research by investigating the normal levels of ROS in arterial and venous tissue,
their molecular sources, and corresponding changes during DOCA-salt

hypertension.

l. ROS production
A. Superoxide production

Using lucigenin-enhanced chemiluminescence, we measured basal levels of

superoxide anions in rings of aorta and vena cava (VC) from normal rats. We

63

observed a higher basal superoxide level in the VC compared to the aorta (fig
11A). Similarly, superoxide production was higher in venous than in the paired
arterial tissue, when using a different artery-vein pair, the superior mesenteric

artery (SMA) and vein (SMV) (fig 11B).

B. Hydrogen peroxide production

H202 production was measured using Amplex Red fluorescence in rat aorta and
V0 tissues (fig 12A). Normalized to protein content, basal H202 production was
higher in V0 compared to aorta. Similarly, H202 production was higher in venous
than in the paired arterial tissue, when using a different artery-vein pair, the
carotid artery and the jugular vein (fig 128). The H202 production of all tissues
was almost abolished by diethyI-dithio-carbamate (DDC), a superoxide
dismutase inhibitor, suggesting that the vast majority of vascular H202 is

superoxide-derived (fig 13).

Because the basal levels of ROS are the result of opposing activity of various
systems of ROS generation and destruction, these results suggest that the
normal rat veins have either a higher ROS production rate and/or a decreased

ROS destruction rate, compared to normal rat arteries.

ll. ROS destruction

A. Antioxidant capacity for H202

64

 

7:?
3
£3
2:

A O)
E
.5
E
L
O
E
5
0)
'U
'2
2
G)
O.
3
(D

B
§ 02
.33 '
a
U)
E 0.1
s
*E
'5 ,o
E 01
5
,8 0.0
X
9
8
3 0.0
(I)

 

SMA SMV

Figure 11. Basal superoxide production measured by lucigenin-enhanced
chemiluminescence in normal rat tissues: aorta (A0) and vena cava (VC) (A);
superior mesenteric artery (SMA) and vein (SMV) tissues (B). Data are
represented as mean :l: SEM for the respective N. * represents a statistically
significant (p<0.05) difference.

65

 

.E
A 9
9
Q.
o:
E-
N
O
N
I
2
C
B

nM H202 lug protein

 

CA JV

Figure 12. Basal hydrogen peroxide production measured by Amplex Red
fluorescence in normal rat tissues: aorta (A0) and vena cava (VC) (A); carotid
artery (CA) and jugular vein (JV) (B). Data are represented as mean :1: SEM for
the respective N. * represents a statistically significant (p<0.05) difference.

66

nM H202/pg protein

 

nM H202 lug protein
.0

 

CA CA+DDC JV JV+DDC

Figure 13. H202 production in basal conditions and in the presence of 10 mM
diethyl dithiocarbamate (DDC, superoxide dismutase inhibitor) after 1h in normal
rat aorta (A0) and vena cava (VC) (A) and carotid artery (CA) and jugular vein
(JV) (B). Bars represent means :t SEM for the respective N. * represents
statistically significant difference (p<0.05) between artery-vein, and # represents
statistically significant difference (p<0.05) between control-DDC.

67

As a measure of antioxidant capacity, we tested the ability of normal aorta and
VC to quench exogenous H202 by measuring the decrease in H202 concentration
in the presence of tissue, using the Amplex Red H202 assay. The VC consumed
significantly more exogenous H202 per amount of protein compared to the aorta

in the presence of 20 pM H202 (fig 14).

B. 4-hydroxy-nonenal

After generation, regardless of source, ROS that are not degraded or that do not
participate in cellular signaling may directly modify (oxidize) cellular components
like proteins, lipids and DNA. We have investigated the expression level of 4-
hydroxy-nonenal, a lipid peroxidation product typically used as a marker of
oxidative stress, in normal aorta and VC. The immunohistochemical localization
of 4-HNE was different in vascular cross-sections (fig 15A), and also appeared
qualitatively higher in VC than in aorta, suggesting more lipid peroxidation may
occur in the VC. The banding pattern of 4-HNE protein adducts in homogenates
(fig 158) also appeared to be different between aorta and VC, however in this
case the staining intensity was observed to be higher in aorta samples,

suggesting 4-HNE protein modifications may occur at a higher rate in the aorta.

Ill. ROS metabolizing enzymes

68

nM H202 quenched lug protein
A N (A) -§ 01
O O O 0 O

O

 

Figure 14. Antioxidant capacity for hydrogen peroxide (20 pM) of normal rat
aorta (A0) and vena cava (VC) tissues measured by Amplex Red fluorescence.
Data are represented as mean :l: SEM for the respective N. * represents a
statistically significant (p<0.05) difference.

69

 

A .. ..
‘3er x3.
“ :1." .
. “.151” ._
. 77$ ‘
N ‘K r .\
L j L M 6%, aorta
“‘ gin:
.‘;-..
‘n. ‘1‘
— f
I.
L .. -
I
1'11 f
, VC
1" ’
1 1':
- ,‘ _/.' q, '

 

 

Figure 15. A. Immunohistochemical staining of 4-HNE (+, right) in normal rat
aorta (top), vena cava (middle) and liver, used as positive control (bottom) as
compared to no primary controls (-, left). The lumen is indicated by “L". Bar
represents 100 pm. B. Western blot of 4-HNE protein adducts in aorta and vena
cava homogenates. using the rat liver band pattern as positive control (+).

70

We have dissected the potential explanations for the differences in basal ROS
levels between arteries and veins in three different ways: detecting differences in
the protein expression (A) or in the mRNA expression of various ROS
metabolizing enzymes (B), as well as pharmacologically inhibiting various
enzymes that produce ROS and measuring the resulting decrease in basal ROS

levels (C).
A. Protein expression of ROS metabolizing enzymes

We first analyzed protein expression of enzymes involved in the generation
and/or destruction of ROS. Western blots displayed in figures 16-19 (A panels)
were performed using protein isolates of aorta and vena cava from normal rats,
and were followed by band densitometry analysis. The expression of XO, a
cytosolic ROS generator (fig 16A), DUOX1, a membrane-bound ROS generator
recently cloned as a Nox family member (fig 17A), CuZn SOD, the cytoplasmic
isoforrn of superoxide dismutase (fig 18A) and catalase, a major H202-
metabolizing enzyme (fig 19A) was significantly increased in V0 compared to

aona.

No difference was observed between the aortic and VC protein expression of
other important sources of vascular ROS: the endothelial isoforrn of nitric oxide
synthase (eNOS); the mitochondrial isoforrn of superoxide dismutase (MnSOD);
the small GTPase component of NADPH oxidase, Rac1; and the 39 KDa subunit
of mitochondrial oxidative phosphorylation complex I, the most important

mitochondrial ROS production site, as well as prohibitin, a mitochondrial marker

71

(data not shown). Use of a number of other commercially available antibodies for
other ROS metabolizing enzymes was attempted without success or marked by
technical difficulties: against other subunits of NADPH oxidase (Nox2/gp91p“°",
p67p"°", p22"“°", p40""°"), other isoforms of NOS (iNOS, nNOS), cyclooxygenase

(COX2) and Iipoxygenase (5LOX).

Protein expression of XO, DUOX1, CuZn SOD and catalase in tissues was
confirmed by immunohistochemical analysis of rat aorta and VC cross-sections
(figures 16-19, respectively, B panels). Qualitatively, the X0 and catalase
staining appeared to be more intense and more diffuse in VC compared to aorta
sections. CuZn SOD staining was intense in both aorta and V0 sections, without

appearing to be qualitatively different.

B. mRNA expression of ROS metabolizing enzymes

We have screened for differences in mRNA expression of ROS metabolizing
enzymes with a PCR array approach, using the Oxidative Stress and Antioxidant
Defense PCR array (SA Biosciences) for RNA samples from aorta and VC from
normal rats. Table 2 lists the statistically significant observed differences in gene
expression among the 84 oxidative stress-related genes included in the PCR
array. mRNA expression of DUOX1, as well as DUOX2, was higher in normal VC
than aorta. mRNA expression of NOX4, another Nox family member and
important vascular ROS producer, was higher in aorta than in VC. By this

method, no significant difference was observed in catalase or superoxide

72

dismutase isoforms mRNA expression, while XO was not included. RT-PCR
experiments confirmed the higher mRNA expression of DUOX1 in VC compared

to aorta, while that of DUOX2 displayed a similar trend (fig 20).

C. Relative contribution of ROS producing enzymes to basal vascular ROS

levels

We also used a pharmacological approach to establish the relative contribution of
various enzymatic ROS producers to the total levels of ROS in basal conditions.
Using two rat artery-vein pairs: aorta - VC and carotid artery (CA) - jugular vein
(JV), we examined H202 production in the presence of inhibitors for NADPH
oxidase (100 pM apocynin), X0 (100 pM allopurinol), eNOS (100 pM LNNA),
mitochondrial complex I (10 pM rotenone), flavoenzymes (diphenyleniodonium,
DPI, 10 pM) and cyclooxygenase (COX, 10 pM indomethacin/1O mM ketorolac).
The results of these experiments were expressed as percentage of the basal
H202 level from the same tissue and same animal and are displayed by inhibitor
(fig 21 A-F) or by tissue (fig 22 A-D). No specific enzyme substrate was added at
any point in these experiments. Apocynin reduced basal H202 production of all
tissues, reaffirming the finding that NADPH oxidase is an important source of
ROS in vascular tissues. Rotenone and DPI reduced basal H202 levels in veins,
but not arteries, suggesting a relative higher dependence on mitochondria and
flavoenzymes in veins compared to arteries. Inhibition of NOS with LNNA did not

significantly alter basal H202 levels of any tissue, suggesting that NOS

73

A xanthine oxidase

in. __._z_.,-__
i l l " ILI
Ao VC +

 

 

.12 30% *
C
3
Q
“E’ 200-
.8
(I)
C
a:
g 100-
.15
E
0.
Ac VC
Aorta Vena cava
B - '
" l-l
/ L '/ L
. ' —
. ff
l I I.
., ,/

 

Figure 16. A. Western blot analysis of XO protein expression in normal rat aorta
(A0) and vena cava (VC) homogenates. + represents rat kidney medulla
homogenate, used as a positive control. Band densitometry quantification of 130
KDa band depicted above, represented as mean :l: SEM. * represents a
statistically significant (p<0.05) difference. B. Immunohistochemical staining of
normal rat aorta (left) and vena cava (right) sections, without primary antibody
(top) and with anti—X0 antibody (bottom). Scale bar represents 100 pm. “L"
indicates the luminal side.

74

DUOX1

...“..q...

 

 

l l ”—1
A0 VC +

g; 6000-
I; *
E
g 4000- N=4
.8
(I)
c
g
2‘ 2000-
.‘é’
%

0.

A0 VC
Aorta Vena cava
L L
» H
— _“
ll?)
12‘ :1
t \ ‘ J “hi/(’Tffi (+)
‘L I \:\\' a \ :L \Q
— — K \ 1

Figure 17. A. Western blot analysis of DUOX1 protein expression in normal rat
aorta (A0) and vena cava (VC) homogenates. + represents human thyroid
homogenate, used as a positive control. Band densitometry quantification of
DUOX1, represented as mean :l: SEM. * represents a statistically significant
(p<0.05) difference. 8. immunohistochemical staining of normal rat aorta (left)
and vena cava (right) sections, without primary antibody (top) and with anti-
DUOX1 antibody (bottom). Scale bar represents 100 pm. “L" indicates the

75

CuZn SOD

 

 

   

A ’- ------- -------- -
L l l I l_,l
A0 VC +
a 600‘ *
"3
N=8
E
E 400'
.8
(I)
C
o
g 200-
.2
g
0.
Ac VC

Aorta vena cava

 

Figure 18. A. Western blot analysis of CuZn SOD protein expression in normal
rat aorta (A0) and vena cava (VC) homogenates. + represents rat kidney medulla
homogenate, used as a positive control. Band densitometry quantification of 16
KDa band depicted above, represented as mean 1 SEM. * represents a
statistically significant (p<0.05) difference. B. Immunohistochemical staining of
normal rat aorta (left) and vena cava (right) sections, without primary antibody
(top) and with anti-CuZn SOD antibody (bottom). Scale bar represents 100 pm.
“L" indicates the luminal side.

76

catalase

 

 

  
 

 

 

l
A l--. 4i 1 J 8
Ac VC +
,9 600-
'E
3
E -
“5’ 400- ”‘8
:2
(I)
C
(D
g 200-
.‘é’
E
to
A0 vc

Aorta Vena cava

 

Figure 19. A. Western blot analysis of catalase protein expression in normal rat
aorta (A0) and vena cava (VC) homogenates. + represents rat kidney medulla
homogenate, used as a positive control. Band densitometry quantification of 65
KDa band depicted above, represented as mean :t SEM. * represents a
statistically significant (p<0.05) difference. B. immunohistochemical staining of
normal rat aorta (left) and vena cava (right) sections, without primary antibody
(top) and with anti-catalase antibody (bottom). Scale bar represents 100 pm. “L"
indicates the luminal side.

77

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

     

 

 

 

 

 

mom 3%
EN 08265 20.. 2...
EN c3938 cozmcfiEooom
EN _. .6858 cmmExo Ian—<2
5N o 032x93 0:25.320
5N mommEon 20.5820
md 5228sz EN E22 uofiozéumm new $862 :22 m_xm~<
new 3% Es. 82: chaéscoeéchm 2 5:55
EN 3. $8.6 :89 «x25 own gleam
m3 4% men E2358 2 a :52 Em:
N .203 o__omo§o __r_ao.5oz 5d N mwmc0m>xoocoE mEEmEoo c_>m_u_
mmd v .859: .8 27:3 .meu anm mad Av omexo Ido<zv $62
Nod :wnEoE .mm =Em¢ .oEmo 920w QNF 690693 Fuoufloommm 5m 0 E0650
N3. 3 ommgxo age :83 Nome 8%
5% @3553 our. . . . . ..1 .._____<
«>3 28> E 3: =._ once—J 3

Bon— 20....

 

Table 2. Results of oxidative stress and antioxidant defense genes PCR array,

performed on normal rat aorta and vena cava whole tissue RNA samples.

78

0.0015-

0.0010.

z-ACt

0.0005- *

 

0.0000‘

 

0.031

0.02'l

z-ACt

0.01d

 

 

0.00‘

Aorta VC

Figure 20. RT-PCR analysis of DUOX1 (A) and DUOX2 (B) mRNA expression in
normal rat aorta (A0) and vena cava (VC), expressed as change from beta-2-

microglobulin (2'delta Ct), and represented as mean 1 SEM for the respective N. *
represents a statistically significant (p<0.05) difference.

79

§
§

 

 

 

   

 

 

 

 

 

 

 

 

   

_ _ indomethacin
A § 150. apocynin E §
.0 .0
"5 "5
O O
1
I; it ‘
g 5,, a m
o. g 7/
Ac vc CA JV
n=4 n=4 n=4 n=4
3 200' . F 200-
5 allopurinol _ ketorolac
.3 150. g 150'
“g '6
g: 100‘ 8) 100 1
c .9
a E *
50-
50-
0‘ c r ~ . .
= - _ - Ac VC CA JV
n 2 n-4 n-4 n-4 n=2 n=4 n=2 n=4
c: 300!
_ _ rotenone
t t
.0 .D
“5 “5
g» 5?»
5 E
I
g: // 33
4
Ac VC CA JV
n=3 n=4 n=4 n=4
D- LNNA
3
a:
.0
“6
O
5’
c
E /
0
.. %

 

   

 

 

Ab VC CA. JV
n=4 n=6 n=5 n=5

Figure 21. H202 production in normal rat aorta (Ao), vena cava (VC), carotid
artery (CA) and jugular vein (JV) in the presence of ROS enzyme inhibitors,
represented as percentage of the basal H202 production for the respective tissue,
measured in the same animals. Bars represent means :l: SEM for the respective
N. * represents statistically significant decrease (p<0.05) compared to basal.

80

Percenmge ofbasal 1’
Percentage ofbasal OD

 

 

 

basal?
a 3
U
3'5

 

 

 

 

g 150‘
D
”5 ‘8
gam- §1oc
3 E
1 00- 50"
8 8
o- o-
{x .
0d Q9 00*
99 \\° «° 09

 

Figure 22. H202 production in normal rat aorta (A), VC (B), carotid artery (C) and
jugular vein (D) in the presence of ROS enzyme inhibitors, represented as
percentage of the basal H202 production for the respective tissue, measured in
the same animals. Bars represent means :I: SEM for the respective N. *
represents statistically significant decrease (p<0.05) compared to control (basal).

81

uncoupling in the vasculature does not account for a major portion of superoxide
production in normal conditions. Ketorolac reduced, but indomethacin had no
effect on vascular H202 levels, rendering effects of COX inhibition inconclusive.
Overall, the results of these experiments do not appear to overlap the results of

the protein or mRNA analysis of ROS metabolizers (see discussion).

IV. ROS metabolism in DOCA-salt hypertension

A. Basal H202 levels

H202 production was measured using Amplex Red fluorescence in vascular
tissues from DOCA-salt hypertensive rats (fig 23). Similarly to the normotensive
conditions, basal H202 production (normalized to protein content) was higher in
DOCA VC compared to DOCA aorta (fig 23A). The basal H202 production in the
DOCA jugular vein was however not statistically different from the one in the
DOCA carotid artery (fig 238). The numerical results of these experiments
cannot be directly compared to the ones in normotensive rats (fig 12), since the

animals in the two sets were not age-matched.

B. Protein expression of ROS metabolizing enzymes

We investigated the protein expression of major ROS metabolizing enzymes in
aorta and V0 tissues from sham normotensive and DOCA-salt hypertensive rats.

We used the same antibodies to detect protein expression as mentioned before

82

in the case of normal rats. The results obtained in sham normotensive rats were
similar to those obtained previously in normal rats (fig 24, lanes 1-4 and 11-14).
Protein expression of xanthine oxidase, catalase and CuZn SOD was higher in
DOCA VC compared to DOCA Ao, while protein expression of p47p"°" was higher
in DOCA Ao compared to DOCA VC (fig 24, lanes 6-9 and 16-19). Similarly to
the situation in normal rats, no difference was observed in expression of MnSOD,
Rac1 and eNOS between arterial and venous tissue from DOCA-salt

hypertensive rats.

V. Xanthine oxidase in normal conditions and in DOCA-salt hypertension
A. XO protein expression

As previously shown, protein expression was higher in VC compared to aorta
samples in both normotensive and DOCA-salt hypertensive rats (fig 16, 24).
These results were obtained using a polyclonal XO antibody (Rockland). To
further confirm the validity of these findings, we verified protein expression with a

different XO antibody (Abcam), with similar results (not shown).

B. XO mRNA expression

Because circulating XO can bind to vascular endothelium (see discussion), we
tested whether vascular X0 is also locally produced by performing RT-PCR for

X0 mRNA. lmportantly, XO mRNA was present in both aortic and VC samples.

83

In addition, relative quantification of XO to GAPDH mRNA (fig 25) showed that
XO mRNA expression was higher in the VC compared to the aorta. During
DOCA-salt hypertension, XO mRNA expression was maintained at a similar level
in both types of tissues, with higher expression in DOCA VC compared to DOCA

aona.

C. XO activity

XO protein activity was measured through two independent assays. First, the
formation of urate, the final product of XO-catalyzed reactions, was determined
spectrophotometrically from tissue homogenates, in the presence of the XO
substrate xanthine. The signal measured in the presence of the XO inhibitor
allopurinol (100 pM) was considered as non-specific and was subtracted from the
total. The resulting XO activity, expressed as the allopurinol-inhibitable uric acid
production in the presence of xanthine, was normalized to the total protein
content of samples. XO activity was higher in VC compared to aorta samples in

normal rats (fig 26 A).

Second, the formation of H202 from whole tissues in the presence of xanthine
was measured fluorometrically using the Amplex Red assay. Allopurinol was
again used to subtract the non—XO derived H202 formation. XO activity expressed
as allopurinol-inhibitable H202 production, was higher in V0 compared to aorta

(fig 26 B).

84

 

.E
A g
9
O.
0)
EL
ON
N
I
2
C
DOCA Ao DOCA V0
0.3-
B

0.2-i

0.1-

nM H202 lug protein

 

0.0-

 

DOCA CA DOCA JV

Figure 23. Basal hydrogen peroxide production measured by Amplex Red
fluorescence in tissues from DOCA-salt hypertensive rats: aorta (A0) and vena
cava (VC) (A); carotid artery (CA) and jugular vein (JV) (B). Data are represented

as mean :1: SEM for the respective N. * represents a statistically significant
(p<0.05) difference.

85

. _* e-’_'.':m‘.
. .-. ’3 I. w. . ‘
but. up ... .4. - - ~" ' '

 

 

 

 

X0 5 Q '- a. - - u h - .0 '4 ”
cataIase ...- .... W m
cuanOD ...... ...—... ....-- '- ...—- -— __ _ __ __._ .
-—a-v—-—- -— wW-—7;—- ;,.‘ ".33“ ‘T‘ “ ow a "W
p47phox . é
.... - ...:
hill-i." '_ I g... ... . .. a
91 .
eNOS 'irM“ - .. ~ - -- -~ -- ‘ .
I—Im nm “ -' -... -d
“if . . 1". ..;&4- ....-..-.-.- '44-: .aa-r Jae}. :ue: .11. m hhfiiirmr
MnSOD

 

Willi 'fi'ill"

 

 

12 3 4 6 7 8 9 11121314 16171819 21

sham DOCA sham DOCA

aorta vena cava +

Figure 24. Western blot analysis of ROS metabolizing enzymes in sham aorta

(lanes 1-4), DOCA aorta (lanes 6-9), sham vena cava (lanes 11-14) and DOCA
vena cava (16-19). Positive controls (+) were loaded on lane 21.

86

0.8 *

0'6 N=6-13

&

 

 

 

 

 

 

 

 

 

 

 

A0 VC DA DVC

Figure 25. RT-PCR analysis of XO mRNA expression in normal rat aorta (A0)
and vena cava (VC), and DOCA-salt rat aorta (DA) and vena cava (DVC),
expressed as change from GAPDH (2““3 Ct), represented as mean :l: SEM for
the respective N. * represents a statistically significant (p<0.05) difference

between artery-vein pair.
87

.E
e
A e
O.
O)
3.
\
To
6
(U
0
'C
3
2
C
8

nM H202 lug protein

 

Ao VC

Figure 26. A. Xanthine oxidase activity in normal rat aorta (A0) and vena cava
(VC) homogenates as allopurinol-inhibitable urate production. B. Xanthine
oxidase activity in rat aorta (A0) and vena cava (VC) whole tissues as allopurinol-
inhibitable hydrogen peroxide production. Data are represented as mean 1 SEM
for the respective N. * represents a statistically significant (p<0.05) difference.

88

 

 

 

 

A r.
2
O.
D
E-
'U
’5
(U
0
'C
3
2
C
An VC DA DVC
B
.E
Q
2
Q
U)
E‘-
o:
I
2
C
SA SVC DA DVC
C

nM H202 lug protein

 

SCA SJV DCA DJV

Figure 27. A. Xanthine oxidase activity in normal aorta (Ao), normal vena cava
(VC), DOCA aorta (DA) and DOCA vena cava (DVC) homogenates as
allopurinol-inhibitable urate production. B. Xanthine oxidase activity in sham
aorta (SA), sham vena cava (SVC), DOCA aorta (DA) and DOCA vena cava
(DVC) whole tissues as allopurinol-inhibitable hydrogen peroxide production. C.
Xanthine oxidase activity in sham carotid artery (SCA), sham jugular vein (SJV),
DOCA carotid artery (DCA) and DOCA jugular vein (DJV) whole tissues as
allopurinol-inhibitable hydrogen peroxide production. Data are represented as
mean :l: SEM for the respective N. * represents a statistically significant (p<0.05)
difference between artery-vein and # between sham-DOCA.

89

XO activity in aorta from DOCA-salt hypertensive rats was similar to that in
normotensive rat aorta in both activity assays (fig 27A-B). However XO activity in
DOCA VC was significantly increased compared to normal VC in the urate-based
assay (fig 27A) and displayed a similar trend in the H202-based assay (fig 278).
The XO activity in jugular vein was not different from that in carotid artery in both
sham and DOCA-salt hypertensive rats, however, the XO activity of both these
tissues appeared increased in DOCA-salt hypertensive compared to sham
normotensive rats, this increase being statistically significant in the case of the

jugular vein (fig 27C).

ffition s_ummary: Comparisons of ROS levels by arterial and venous tissue
have shown that the ROS production in veins was higher compared to
conesponding arteries. The H202-degrading capacity of veins was similariy
increased. There were multiple differences in ROS metabolizing enzymes
between arteries and veins. There was a higher expression of DUOX1, CuZn
SOD and catalase in venous compared to arterial tissue. The most consistent
findings were related to xanthine oxidase. XO had a higher protein and mRNA
expression, as well as higher activity in veins compares to arteries. During
DOCA-salt hypertension, the higher expression of X 0 was maintained by veins,
and there were strong indications of an increase in venous, but not arterial

activity.

90

Specific aim 2: Determine-if there is a difference in the impact of ROS on
arterial versus venous contractile function in basal conditions and in

hypertension

Rationale: ROS can influence vascular contraction, proliferation, remodeling and
inflammation. Similariy to general characteristics of ROS metabolism, the
functional impact of R08 is by far more extensively studied in arteries compared
to veins. When testing the venous role in hypertension, vascular contraction is
the most important function to be investigated. Since levels of ROS appeared
overall to be higher in veins compared to arteries, we next examined whether the
functional consequence of this difference impacts upon the vascular response to

contractile stimuli in these blood vessels.

I. Direct impact of ROS on contractile function of arteries and veins.

To compare the functional impact of ROS on rat aorta and VC tissues, we
exposed these vessels to extraneous superoxide in isolated tissue baths.
Endothelium-intact aorta and VC rings both contracted in a concentration-
dependent manner to exogenous superoxide, generated by the addition of
increasing amounts of XO to xanthine (200 pM) in the tissue bath (figure 28).
This contraction was larger in the VC compared to the aorta, when normalized to

an a adrenergic-mediated response.

91

 

  
 

 

 
 

 

 

 

 

c 250
g -0- aorta
g 200- -I- VC
c N=6-13
8 150-
2
3.
S 100.
DJ
E 50- .
D_ * * * * *
o\° 0 . .
-5 4 s -2 -1
log XO [U/ml]

Figure 28. Cumulative xanthine oxidase concentration-response curves in the
presence of 200 pM xanthine, of normal, endothelium-intact rat aorta (circles)
and vena cava (squares) rings, represented as percentage of the initial
contraction to 10 pM phenylephrine (PE) or norepinephrine (NE) for aorta and
VC, respectively. Points represent mean 1 SEM for the respective N. *
represents a statistically significant (p<0.05) difference.

92

A similarly larger contraction in the VC compared to the aorta was observed in
response to H202 in experiments previously published from our laboratory (see

discussion).

ll. Impact of XO on arterial and venous contractile function in normal rats
A. Impact of XO

To assess the potential involvement of XO-mediated ROS production on vascular
reactivity, we tested the effects of the XO inhibitor allopurinol on vascular
responses to several agonists in isolated tissue bath experiments (fig 29-30). We
used agonists that are important endogenous effectors of vascular tone and that
induce contractile responses in both aorta and VC: norepinephrine (NE) as a
generic alpha adrenergic-agonist (panels A); and angiotensin II (Angll) (panels
C) and endothelin-1 (ET-1) (panels D), peptides commonly associated with ROS
production in vascular tissues. We also used acetylcholine (ACh) Has a relaxing
agent and a general measure of endothelial function (panels B). We used the
same allopurinol concentration (100 pM) as in our previous in vitro XO activity
experiments. This concentration effectively reduced increases in X0 products
(urate and H202) in the presence of substrate (xanthine). Allopurinol did not
modify the contraction or relaxation of normal aorta rings to any of these agonists
(fig 29). In the vena cava (fig 30), allopurinol (100 pM) did not have an effect on
the NE-induced contraction or the ACh-induced relaxation (fig 30B), and the

trend for a decreased Angll-induced contraction in its presence was not

93

aorta

 

 

 

 

 

 

 

 

 

 

 

 

A 150 _ B: 120 .
C -o- vehicle 583 -0- vehicle
E -O- allopurinol g 100‘ -O- allopurinol
§ 100- § eo~
A c
a 0' 6°-
2 504 9- 40..
V 2
3f 3 20. N=8-9
3‘ 3

o. O c I U r r T

- -4 -10 -9 -a -7 -0 -5 -4

log ACh [M]
F- -0- vehicle 5 -0- vehicle
E 20- -O- allopurinol E -o- allopurinol
N=4 c

8 15- 8
a ,0, a
9 2
31' 5+ 3."
0" c.\"

01

 

 

 

 

 

-11 -10 -9 -8 -7 -6
logAngll[M]

 

Figure 29. Cumulative concentration-response curves to norepinephrine (A),
acetylcholine (B), angiotensin II (C) and endothelin-1 (D) of endothelium-intact
rat aorta rings in the presence of allopurinol (100 pM) (open symbols) or vehicle
(closed symbols), represented as percentage of contraction to 10 pM
phenylephrine (PE) or 20 (M PGFZG. Points represent mean 1: SEM for the
respective N. * represents a statistically significant (p<0.05) difference.

94

vena cava

 

 

 

 

 

 

 

 

 

 

 

 

 

 

A B
100 _ c 120 .
g -I- vehicle g '3' vehicle
g 00‘ 43- allopurinol g 100‘ ~0- allopurinol
C 80"
2 so- 3
a g so-
2 “" ‘1 4o-
? 20‘ E 20. N=4-5
,3 c:
0" I I o\ C I I I I I
-10 -a -6 -4 -10 -9 -8 -7 -6 -5 -4
|og NEIM] log ACh[M]
C300 D
- 800
% -l~ vehicle 5 ~l- vehicle
c E
200‘ c
E 8
a S
3.
g 100« g
I.“
z ”2’
39 o\°
0" — 1

 

 

 

 

~11 -10 -9 -8 -7
log Angll [M]

 

Figure 30. Cumulative concentration-response curves to norepinephrine (A),
acetylcholine (B), angiotensin II (C) and endothelin-1 (D) of endothelium-intact
rat vena cava in the presence of allopurinol (100 pM) (open symbols) or vehicle
(closed symbols), represented as percentage of contraction to 10 pM
norepinephrine (NE) or 20 pM PGFZO. Points represent mean :1: SEM for the
respective N. * represents a statistically significant (p<0.05) difference.

95

statistically significant (fig 30C). However, allopurinol significantly reduced the
maximal contraction to ET-1 (10'7 M) in the VC (fig 30D). As previously reported,
Angll and ET-1 were more efficacious and more potent in VC compared to aortic
rings, when reported as a percentage of the initial contraction to 10 pM NE (VC)

or PE (aorta).

B. Impact of uric acid

XO may also influence contractile function through its other product, uric acid.
Rodents express uricase, a urate-degrading enzyme. The expression of uricase,
identified as the 64 KDa band corresponding to a uricase dimer, was higher in

VC compared to aorta in normal rats (fig 31).

Uric acid may have multiple effects on vascular contractile function (see
Introduction). Because XO expression, as well as its uric acid producing activity,
was higher in the rat VC compared to aorta, we hypothesized that any direct
contractile effects of uric acid would be higher in the VC compared to the aorta.
To investigate the acute effects of uric acid on vascular contractile function, we
performed isolated tissue bath experiments using uric acid in combination with
oxonic acid, an inhibitor of uricase. Uric acid (10'9-104 M), alone or in
combination with oxonic acid (10 pM), did not induce contraction of endothelial-
intact rat aorta or VC rings (fig 32A). Uric acid (100 pM) in combination with

oxonic acid (1 0 pM) did not significantly change acetylcholine (ACh)-induced

96

('— 64 KDa

<— 32 KDa

 

Aorta VC
2500- *
2000-I
1 500-
1 000-

500-

arbitrary densitometry units

 

 

Figure 31. (top) Uricase protein expression in normal rat aorta and vena cava.
Positive control (+) for uricase expression is a rat liver Iysate. (bottom) Band
densitometry quantification of 64 KDa band depicted above. Represented as
mean :I: SEM for the respective N. * represents a statistically significant
difference (p<0.05).

97

 

 

    

    

or IUAI

      

-7.5 -7 6.5 «6 -5.5 - 4.5 4 3.5

H oxonic acid

(+) oxonic acid

 

 

 

 

 

‘” ‘ RA 40° 1
zoo J 200 <
.W s
o J g 0
2 3.2
400 4 -200
400 . og[UA] 400 -
-9 -s.s -7.s - -6.5 -6 -5.5 -s 4.5 4.3.5
B C
‘5 15c .5 150
8 8
g -0- Ac vehicle E -l- VC vehicle
§ 100* .0. Ac uric acid 8 100‘ {I- VC uric acid
a
t a
a 501 a 50‘
o N= 0 N=5
N m
39 T 32

 

 

 

 

 

 

i 0

d
O
. O

.5
O

39 -'s -'7 is :5 -4
log ACh[M]

39 -23 37 -e -15. -4
log ACh[M]

Figure 32. A. Tracings of cumulative concentration response curves to uric acid
(10'9 M to 3*104 M) in the presence (left) or absence (right) of oxonic acid (10
pM) in endothelium-intact normal rat vena cava (top) and aorta (bottom) rings. B,
C. Cumulative concentration response curves to acetylcholine (ACh, 10'9-10’5 M)
of endothelium-intact aorta (B) and vena cava (C) in the presence of vehicle
(closed symbols) or uric acid (100 pM) plus oxonic acid (10 pM) (open symbols).
Points represent means 1 SEM of % of contraction induced by 20 pM PGan.

98

relaxation curves of endothelial-intact rat aorta or VC rings contracted with 20 pM

PGan (fig 32B-C).

Ill. Impact of flavoenzymes on endothelin-1 induced contraction of aorta

and vena cava

Because we observed allopurinol inhibiting maximal contractions induced by ET-
1 in the vena cava, as well as higher expression of X0 and DUOX enzymes
(both flavoenzymes) in the vena cava compared to aorta, we investigated
whether a general flavoenzyme inhibitor would alter ET-1-induced contraction of
these tissues in isolated tissue bath experiments. Concentration response curves
to ET-1 were performed on endothelium-intact aorta and V0 rings in the
presence of diphenyleniodonium (DPI) or vehicle (fig 33). ET-1-induced
contraction was not altered in the aorta (fig 33A), however it was significantly
decreased in the VC upon exposure to DPl (fig 338), suggesting flavoenzymes

contribute to ET-1 mediated contraction of VC, but not aorta.

IV. Arterial and venous endothelial function in normal rats and during

DOCA-salt hypertension

Arterial endothelial function is characteristically impaired in human and animal
models of hypertension. There are multiple mechanisms behind the hypertensive

endothelial dysfunction, but the increased ROS production commonly observed

99

 

in hypertension is a generally accepted pathogenetic factor. The most obvious
argument supporting this idea is the fact that superoxide, the initiator of the ROS
cascade, directly reacts with NO, the endothelial-derived relaxing factor, thus
decreasing its bioavailability. The decreased NO levels and the impaired NO-
_ mediated relaxation are hallmarks of endothelial dysfunction. Because we
observed higher ROS production in veins compared to arteries (see aim1), we
hypothesized that venous endothelial function would be affected by hypertension

to at least a similar, if not a higher degree as compared to arteries.

We tested the endothelial function of aorta and V0 from normal and DOCA-salt
hypertensive rats in isolated tissue bath experiments by performing ACh-induced
relaxation curves on rings of tissues stably contracted with PGFZO (20 pM). In
normal conditions, VC relaxed in a similar fashion to ACh as the aorta (fig 34A).
To confirm the dependence of ACh-induced relaxation on NO production, we
performed the same experiment in the presence of LNNA, a NOS inhibitor (fig
34B). We observed complete abolishment of ACh-induced relaxation in the
presence of LNNA, suggesting that both aorta and VC ACh-mediated responses
are dependent on NOS function. In tissues from DOCA-salt hypertensive rats,
while we confirmed the characteristic hypertensive arterial endothelial
dysfunction, Le. a significant decrease in the ACh—induced relaxation in aorta
rings, we did not observe any loss in the endothelial function of VC with
hypertension (fig 35A). The ACh-mediated responses in tissues from DOCA-salt
rats appeared to maintain their dependence on NOS function, as evidenced by

their abolishment in the presence of LNNA (fig 35B).

100

 

 

 

 

 

 

 

 

A C 200 + A h'l
.0 N=6-7 ove Ice
E 150- -0- Ac DPI
C
8
’2‘ 100-
3.
,9
I 50*
D.
o\°
0 .
-11 -10 -9 -8 -7 -6
log ET-1 [M]
800 VC h'l
N=6_7 + ve Ice
3 {I- VC DPI

 

% NE (10 pM) contraction

 

 

O

-11 -10 -9 -8 -7 -6
log ET-1 [M]

Figure 33. Cumulative concentration response curves to endothelin-1 (ET-1,
10‘11-10'7 M) of endothelium-intact rat aorta (A) and vena cava (B) in the
presence of vehicle (closed symbols) or diphenyleniodonium (DPI, 10 pM) (open
symbols). Points represent means 1 SEM percentage of initial contraction
induced by 10 pM phenylephrine (PE) or norepinephrine (NE) for aorta and vena
cava, respectively.

101

 

 

 

 

 

 

 

 

 

c 120 .
% +onehicle
A g 100‘ -l— VC vehicle
5 80-
o
E 604
0
0- . N=4 -
2 40
:1. _-.
o 20' .
N '.
°\° o r I I I I l
-10-9-8-7-6-5-4 F
logACh[M]
r: 120
B 2% 09009000; 0 ”LNNA
g 100‘ {I- VC LNNA
§ 80‘
8
E 60-
(D
0- - N=4
2 40
3.
o 20'
N
°\° 0 I I I T T
-10-9 -8 -7 -6 -5 4
logACh [NI]

Figure 34. Cumulative concentration response curves to acetylcholine (ACh,
109-105’ M) of rat aorta (RA) and vena cava (RVC) rings from normal rats in the
presence of vehicle (A, closed symbols) or LNNA (100 pM) (B, open symbols).
Points represent means 1 SEM percentage of contraction induced by 20 uM
PGan.

102

 

 

 

 

 

 

 

 

: 120

.g -o— DOCAAo vehicle (N=5)
A g 100‘ + DOCAVC vehicle (N=4)

§ 80-

:3

LL 60" *

8

2 40'

1

o 20‘

N

°\° c I I l I I

-10-9-8-7-6-5-4
logACh[M]

c 123

.g -o- DOCAAoLNNA(N=6)
B g 100- m -El- DOCAVC LNNA(N=3)

§ 80‘

:5

E 60-

8

2 4o-

3.

o 20'

N

°\° c . f. . t

-10-9-8-7-6-5-4
logACh[M]

Figure 35. Cumulative concentration response curves to acetylcholine (ACh,
10'9-10'5 M) of rat aorta (RA) and vena cava (RVC) rings from DOCA-salt
hypertensive rats in the presence of vehicle (A, closed symbols) or LNNA (100
pM) (B, open symbols). Points represent means :I: SEM percentage of
contraction induced by 20 pM PGFZO.

103

V. Impact of X0 on arterial endothelial function in DOCA-salt hypertension

Because we did not observe endothelial dysfunction in the VC during DOCA-salt
hypertension, we pursued the following experiments only in aorta tissues from

sham normotensive and DOCA-salt hypertensive rats.
A. Impact of X0

To evaluate the potential impact of X0 on the aortic endothelial dysfunction, we
tested the effect of allopurinol on ACh-induced relaxation of sham and DOCA
aorta. Endothelial function, assessed as relaxation to ACh (10'9-10‘5 M) from 20
pM PGFZO-induoed contraction, was again expectedly impaired in aorta from
DOCA-salt hypertensive rats compared to normotensive sham controls (fig 36,
vehicle), as evidenced by a decreased relaxation response to ACh. Inhibition of
X0 enzyme activity with allopurinol (100 pM) did not significantly change ACh-
induced relaxation curves of sham and DOCA rat aorta rings contracted with 20
pM PGF2Cl (fig 36, allopurinol), suggesting that acute inhibition of both uric acid
and ROS production by X0 does not improve nor worsen DOCA-salt aorta

endothelial dysfunction.

B. Impact of uric acid

We performed the same experiment as above to explore the direct effects of uric

acid on the endothelial dysfunction in DOCA aorta. Addition of uric acid (100 pM)

104

 

 

120
-0- Sham Ao vehicle
5 ,5 100- -0- Sham Ao allopurinol
:09! g + DOCAAo vehicle
a. c 80‘ -l'_'|- DOCAAo all0purinol
a 8
‘O 60" _

8 g A N—6
o\ E 40-

20 . .

 

 

 

-1o 39 -2; -7 -6 -15 -4
logACh [M]

Figure 36. Cumulative concentration response curves to acetylcholine (ACh,
10'9-10'5 M) of aorta (Ao) rings from sham normotensive (circles) and DOCA-salt
hypertensive rats (squares) in the presence of vehicle (closed symbols) or
allopurinol (100 pM) (open symbols). Points represent means :l: SEM percentage
of contraction induced by 20 pM PGan.

105

 

120
-0- Sham Ao vehicle
5 ,8 100- -o- Sham Ao uric acid
E E + DOCAAo vehicle
89. c 80‘ -I:l- DOCAAouricacid
a 8
'o 60-

8 §
o\° '8 40. N=6

20

 

 

 

-1o -9 33 37 36 35 -4
logACh[M]

Figure 37. Cumulative concentration response curves to acetylcholine (ACh,
10'9-10‘5 M) of aorta (Ao) rings from sham normotensive (circles) and DOCA-salt
hypertensive rats (squares) in the presence of vehicle (closed symbols) or uric
acid (100 pM) plus oxonic acid (10 pM) (open symbols). Points represent means
:1: SEM percentage of contraction induced by 20 pM PGan.

106

 

60

 

E 40
3
0
(B
-‘.—’ 20
3

 

SHAM DOCA

Figure 38. Uric acid levels in arterial blood of individual sham normotensive
(closed symbols) and DOCA-salt hypertensive (open symbols) rats. Line
represents average uric acid for the respective N.

107

 

A

3 - ‘ i ‘
uricase ,

*(_64KDa

 
 

6 32 KDa

 

   

a-actin “-..—~— ~u-_.- 1 ( 44KDa
l l l— ; {L_l
Sham aorta DOCA aorta +
B
Q E
S 5
5‘ 5‘
E E
.9 .8
Q 2
3 g
2‘ a
.3 .E
E B
G N
Sham aorta DOCA aorta Sham aorta DOCA aorta

Figure 39. A. Uricase (upper panel) and control d-actin (lower panel) protein
expression in aorta from sham normotensive and DOCA-salt hypertensive rats.
Positive control (+) for uricase expression is a rat liver Iysate. B. Band
densitometry quantification of the 64 KDa band represented in the upper panel
(left) and represented as a percent of the densitometry quantification of q-actin
band represented in the lower panel (right).

108

in combination with oxonic acid (10 pM) did not significantly change the ACh-
induced relaxation curves of sham and DOCA rat aorta (fig 37), suggesting that
acute exposure to exogenous uric acid does not improve nor worsen DOCA-salt

aorta endothelial dysfunction.

We also explored the possibility of a chronic decrease in antioxidant uric acid
levels in the DOCA-salt hypertension model that would contribute to arterial
endothelial dysfunction. Uricemia was measured in sham normotensive and
DOCA-salt hypertensive rats. No significant changes were observed between the
two groups (figure 38). Moreover, no significant correlation was found between

the uric acid level and blood pressure of individual rats (not shown).

Since X0 expression and activity do not change in aorta during DOCA-salt
hypertension, suggesting that local aortic uric acid production is not changed, we
decided to also investigate the aortic protein expression of uricase, the urate-
degrading enzyme (fig 39). Quantification of the 64 KDa band densitometry itself
or expressed as percentage of the smooth muscle a-actin band densitometry

revealed no significant change between the sham and the DOCA aorta.

Section gummarv: The functional impact of ROS on contraction was higher in
veins compared to arteries. In basal conditions, X 0 appeared to play little role in
arterial contractile function, however it contributed to venous E T-1-mediated
contraction. In normal conditions, the endothelial function of arteries and veins

was similar. During DOCA-salt hypertension, arteries, but not veins, were

109

 

affected by endothelial dysfunction. X0 did not appear to contribute to the arterial

hypertensive dysfunction by either ROS or urate production.

110

Specific aim 3: Determine whether venous function and blood pressure can

be influenced by changing ROS levels in viva

Rationale: Our results showed that ROS levels were higher in venous tissue
compared to arterial tissue. X 0 may contribute differentially to those levels,
having a higher expression and activity as well as a greater influence on venous
contraction to agonists such as E T-1 . During DOCA-salt hypertension, venous,
but not arterial X0 activity was increased. Inhibition of ROS production was
shown to have preventative or ameliorating effects in various animal models of
hypertension. If the above are true, then inhibiting ROS, or particulariy X 0
activity in vivo would have a differential effect in arteries and veins, preferentially
lowering ROS levels in venous tissue over arterial tissue and diminishing the
contractile effects of E T-1 (and perhaps other agonists) in veins. This would
potentially lower venomotor tone, and therefore mean circulatory filling rpessure
(MCFP). lf MCFP is lowered by X0 inhibition and if venomotor tone is a key
parameter during the development of hypertension, then in vivo inhibition of X 0
should either reduce BP in the case of established hypertension or protect the

animals from increases in BP in the initial stages of experimental hypertension.

l. X0 inhibition in the established DOCA-salt hypertension

To inhibit X0 activity in vivo, we used allopurinol, administered orally, at 50
mglkg/day (in the drinking water). The dose/administration route chosen are the

most commonly used in rodent X0 research studies. We have also previously

111

attempted parenteral administration of allopurinol: subcutaneously (through
osmotic minipumps) and intravenously (data not shown). Due to the poor water
solubility of purine analogues and the adverse reactions seen with organic or
strong basic solvents, we chose the oral route. Additionally, this route mimics the
human administration of allopurinol. The chosen dose is a mid-range rodent

allopurinol dose that has been proven to inhibit X0 activity (see discussion).

To determine whether allopurinol would reduce BP of hypertensive animals, we
performed a study (fig 40) administering allopurinol (50 mglkg/day, orally) or
vehicle to DOCA-salt hypertensive rats. The drug and its vehicle were
administered for 7 days starting at 3 weeks after the DOCA-salt treatment was

initiated.

Blood pressure was monitored through radiotelemetry. Urine was collected at
three time points: before the initiation of DOCA-salt, after 2 weeks of DOCA-salt
treatment and at the end of the allopurinol administration. Serum and tissues
(aorta, vena cava, carotid artery, jugular vein, liver, kidney and heart) were
collected at the end of the study. Urine, serum and tissue samples were used for
HPLC measurements of uric acid and allantoin (products downstream of xanthine
oxidase), hypoxanthine and xanthine (substrates upstream of xanthine oxidase)

and allopurinol and oxypurinol (the drug and its active metabolite).

The results of the HPLC analysis of urine samples are displayed in figure 41. No
significant changes from baseline were observed in the urinary levels of uric acid

and allantoin after 2 weeks of DOCA treatment or at the end of allopurinol

112

‘5. l4“.‘., f .?

 

administration in either group; and there were no significant changes of uric acid
and allantoin between the allopurinol treated and the vehicle group at any time
point (fig 41, top). Urinary levels of hypoxanthine and xanthine in the two
treatment groups were similar at baseline and during DOCA-salt hypertension.
However, these levels increased substantially in the allopurinol-treated group at
the end of the treatment (fig 41, middle). Allopurinol and oxypurinol were
detected in the urine of allopurinol-treated rats at the end of the treatment (fig 41,

bottom).

The results of the HPLC analysis of serum samples collected at the end of the
experiment are displayed in figure 42. A clear trend for reduction of both uric acid
and allantoin was observed in the allopurinol-treated group, although at N=4 this
was only statistically significant in the case of allantoin (fig 42, top). Similarly to
urine samples, serum levels of hypoxanthine and xanthine were dramatically
increased in the allopurinol-treated group (fig 42, middle). Allopurinol and its
metabolite, oxypurinol, were detected only in serum samples of allopurinol-

treated rats (fig 42, bottom).

Tissue samples were analyzed for X0 metabolites through HPLC in a similar
fashion (fig 43). Besides unusually high levels of allantoin in the carotid artery,
allantoin was generally not detectable in tissue samples (fig 43A). Uric acid was
significantly decreased in all the four blood vessels (aorta, vena cava, carotid
artery, jugular vein) in the allopurinol-treated group (fig 438). Hypoxanthine
appeared increased in all samples in the allopurinol-treated group, statistical

significance being reached in the case of the aorta, vena cava, jugular vein,

113

 

kidney and the heart (fig 43C). No significant change was observed in tissue

xanthine levels (fig 43D).

Overall, the results of HPLC analysis indicated effective inhibition of X0
metabolism with subsequent rise in X0 substrate levels and decrease in X0

products in tissues of interest.

The mean arterial pressure (MAP) observed over the course of this experiment is
displayed in figure 44. The missing data points on this figure correspond to the
removal of rats from the close proximity of receivers and their placing in
metabolic cages necessary for urine collection (second timepoint, day 18).
Expectedly, BP rose significantly following DOCA-salt treatment in all animals,
reaching an average of 156 mmHg in the allopurinol group and 169 mmHg in the
vehicle after 3 weeks, at this timepoint MAP being not statistically significant
between the two treatment groups. Over the course of the 7—day allopurinol
treatment that followed, MAP continued to rise in the allopurinol-treated group,
following the same pattern observed in the vehicle group, with no statistically
significant changes. Similarly, no changes were observed in the other
hemodynamic parameters obtained through radiotelemetry (data not shown).
These data suggest that allopurinol did not alter the course of hypertension in the

established DOCA-salt model.

II. X0 inhibition in the development of DOCA-salt hypertension

114

cozom=oo

 

 

osmmz new 823 655

2..

 

 

 

cozoo=oo cozoo=oo
mat: 055
$233.5 on 9.5:. 8m cogcsaé
3558.6 rem <08 raw 58528
em 2 m_ Tame
62.2:qu <00n.

Figure 40. Study design for allopurinol effects on established DOCA-salt

hypertension.

 

115

 

 

 

 

 

 

 

80- _
A allantorn
«’6‘
'2 60-1
or
E.
.E 40-
.9
r:
g 20-
‘“ N=4
c I I I
dayO day 18 day 31
hypoxanthine
3': 0.4- ...
(U
:9
E’ 03- l
v I
a 02- x
" ' I
E ~=4 ,I
S I
0.1. ’
g I
..C 0.0‘ —W I
dayO day 18 day 31
10- .
I; allopunnol *
m 8-
2
o:
E 6-
5 I
E 4" N=4 I’
3 I
o. I
g a I
<0 ,’
O-l ¢
day 0 day 18 day 31

Figure 41. Urinary X0 metabolites in vehicle (closed symbols) and allopurinol
(open symbols)-treated rats, determined by HPLC: allantoin and uric acid (top,
X0 products); hypoxanthine and xanthine (middle, X0 substrates); allopurinol
and oxypurinol (bottom, X0 inhibitors), represented as mean :t SEM for N=4. *
represents a statistically significant difference (p<0.05) between treatment

groups.

uric acid (mg/day)

xanthine (mg/day)

oxypurinol (mg/day)

 

 

 

 

 

 

urine
4. uric acid
31
..... .6. .. - .. - -
2.
1.
N=4
C 1 I I
day 0 day 18 day 31
20- ,
xanthine
*
15'I l
101 I’I
N=4 I
5‘ 1’
’I
g. _Q #Ig :
day 0 day 18 day 31
25. oxypurinol *
20‘ l
I
15- ’
I
’I
10« N=4 ,
I
5‘ I,
I
0' Y ;
day 0 day 18 day 31

116

-o- VEHICLE
-o- ALLOPURINOL

-o- VEHICLE
~0- ALLOPURINOL

'0' VEHICLE
-0- ALLOPURINOL

_.‘ . ..n"!n.'fi
. ”gr-.1 .If'

 

 

 

 

serum

 

 

 

 

 

 

 

 

 

 

allantoin
100-
E .E
o
*5 80- g
b L.
a N=4 o.
a: so- or
z . g
.2 40- as
g E’
“5 20- 3
o- . ,
VEHICLE ALLOPURINOL
hypoxanthine
C .E
a °°' .9
E —*— .9.
60-
? 8
-— - C
5 40- N'4 g
C
Q E
g: 20- :3
.c 3
2 c . .
VEHICLE ALLOPURINOL
allopurinol
.c 03' _*_ .E
Q a
23. E
g) 0.2'l N =4 £01)
.2 E
g 01- a
g ' %
II o
C)
:1.
g o.c , 1 .
VEHICLE ALLOPURINOL

Figure 42. Serum X0 metabolites in vehicle (closed symbols) and allopurinol
(open symbols)-treated rats, determined by HPLC: allantoin and uric acid (top,
X0 products); hypoxanthine and xanthine (middle, X0 substrates); allopurinol
and oxypurinol (bottom, X0 inhibitors), represented as mean :I: SEM for N=4. *
represents a statistically significant difference (p<0.05) between treatment

groups.

117

 

 

uric acid

 

 

 

 

 

 

 

 

 

 

 

2501
200-
150-
N:
100-
0- I 1 I
VEHICLE ALLOPURINOL
1001 xanthine
*
80d _ _
N=4
60-
40-
20-
C I I
VEHICLE ALLOPURINOL
oxypurinol
25-
2od __*_..
N:
15-
1m
5.
c I I ’
VEHICLE ALLOPURINOL

 

fissue

 

 

 

 

 

 

 

 

 

 

 

A allantoin
5 - VEHICLE
{,3 N=4 D ALLOPURINOL
a , i
gt .1.

E
E
g
(B

2
33 1

AORTA vc CA J'\/ LIV'ER KIDI'VEYHEART
uric acid

:: VEHICLE

B .5 2 I _ 'III I
§ N-4 __ _.=_ D ALLOPURINOL
O.

i
(B
O
'C
3
3
Eh
AORTA vc CA JV LIVER KIDNEY HEART
c 2 hypoxanthine
.- 1 *

C % - VEHICLE
a N=4 l D ALLOPURINOL
% 500v *

C
-- 400<
‘2 3001
g *
a 2°” .
J; 1004 *
33 0. F: _
AORTA vc CA JV LIVER KIDNEY HEART
0! xanthine
100

D .E I VEHICLE
*3 D ALLOPURINOL
a. -

O)
E
5

C

Q

3

 

 

 

 

A0 RTA VC CA JV LIVER KIDNEY HEART

Figure 43. Tissue X0 metabolites in vehicle (closed symbols) and allopurinol
(open symbols)-treated rats, determined by HPLC: allantoin (A), uric acid (B),
hypoxanthine (C) and xanthine (D) represented as mean :I: SEM for N=4. *
represents a statistically significant difference (p<0.05) between treatment
groups.

118

 

 

 

 

 

 

 

DOCA allopurinol
surgery 50 mglkg/day
-0- VEHICLE
200‘ -o- ALLOPURINOL
8 180-1
E
g, 160-
n_ 140‘
§ 120-1
100- .
BC I I I I I I I I I I
0 3 6 91215182124273033

days

Figure 44. Blood pressure of vehicle (closed symbols) and allopurinol (open
symbols)-treated rats, measured by radiotelemetry. Timing of interventions is
represented by vertical lines. Data is represented as means :I: SEM of 24h
average mean arterial pressure (MAP) for N=4.

119

To determine whether allopurinol would attenuate the development of DOCA-salt
hypertension, we performed a second study (fig 45) administering allopurinol (50
mglkg/day, orally) or vehicle to rats 3 days before the start and throughout the

course of DOCA-salt treatment (4 weeks).

Blood pressure was monitored through radiotelemetry. Urine was collected at two
time points: before the initiation of allopurinol treatment and at the end of the 4
week DOCA-salt treatment. Serum and the same tissues (aorta, vena cava,
carotid artery, jugular vein, liver, kidney and heart) were collected at the end of
the study. Urine, serum and tissue samples were used for the same HPLC
measurements of uric acid and allantoin (products downstream of xanthine
oxidase), hypoxanthine and xanthine (substrates upstream of xanthine oxidase)
and allopurinol and oxypurinol (the drug and its active metabolite). Additionally,
the weight of heart and the right kidney was measured as a gross estimation of
hypertensive end-organ damage. Histological examination of cross-sections of

aorta was performed for qualitative assessment of hypertensive remodeling.

The results of the HPLC analysis of urine samples are displayed in figure 46.
Urinary levels of uric acid and allantoin were decreased at the end of the study
compared to baseline, however there was no difference between the vehicle and
the allopurinol treatment (fig 46, top). Urinary levels of hypoxanthine and
xanthine increased significantly in the allopurinol—treated group at the end of the
treatment (fig 46, middle). Allopurinol and oxypurinol were detected in the urine

of allopurinol-treated rats at the end of the treatment (fig 46, bottom).

120

 

The results of the HPLC analysis of serum samples collected at the end of the
experiment are displayed in figure 47. A trend for reduction in uric acid levels was
observed in the allopurinol-treated group, although this was not statistically
significant. Surprisingly, allantoin was increased in the allopurinol-treated rats (fig
47, top). Similarly to urine samples, serum levels of hypoxanthine and xanthine
were dramatically increased in the allopurinol-treated group (fig 47, middle).
Allopurinol itself was not detected, however oxypurinol, its metabolite, was

detected only in serum samples of allopurinol-treated rats (fig 47, bottom).

HPLC analysis of X0 metabolites in tissue samples is displayed in figure 48.
Allantoin was variably detectable in tissues (fig 48A). A statistically significant
decrease in uric acid levels was observed in aorta, vena cava, jugular vein, liver
and kidney, with a similar trend in all remaining tissues (fig 48B). A statistically
significant increase in hypoxanthine was observed in the vena cava and liver (fig
48C). As opposed to the 7-day treatment described previously, xanthine tissue
levels after the 4-week allopurinol treatment appeared overall decreased, this
trend becoming statistically significant in the case of aorta, carotid artery, liver,

kidney and heart (fig 48D).

Despite differences observed in serum and tissue levels of X0 metabolites
between the two experiments (7 day and 4 week allopurinol administration), the
overall results of HPLC analysis in this experiment suggested inhibition of X0

function by allopurinol treatment (see discussion).

121

 

The mean arterial pressure (MAP) observed over the course of this experiment is
displayed in figure 49. DOCA-salt treatment effectively raised the BP of rats in
both treatment groups. However, there was no significant difference in the MAP
of allopurinol-treated compared to vehicle treated rats. These data suggest that
allopurinol did not prevent and did not attenuate the development of DOCA-salt

hypertension.

The other hemodynamic parameters obtained by radiotelemetry are displayed in
figure 50. A trend for increased heart rate was observed in the allopurinol-treated
group (fig 508). Diastolic pressure was increased by allopurinol treatment (fig
500), and there was a statistically significant decrease in pulse pressure
(Ppu.se=Psy3-Pdias) (fig 50E), with potentially important implications (see
discussion). No difference was observed in activity (movement) between the two

groups (fig 50F).

Despite the lack of effects of allopurinol on BP, X0 inhibition may have protective
effects on end-target organs of hypertensive complications. To investigate this
possibility, we measured the heart (fig 51A) and kidney weight (fig 51B) at the
end of the treatment (gross evaluation of left ventricular and renal hypertrophy).
We observed no change in the heart weight and a surprising statistically
significant increase in kidney weight in the allopurinol-treated group (see
discussion). The histological characteristics of hematoxylin-eosin stained aorta
sections did not appear qualitatively different between the two treatment groups
(fig 52), suggesting that hypertensive aortic remodeling was not altered by

allopurinol treatment.

122

 

$239: on a
_oc_5ao__m tmuw c885. .5

5:02.00 $580.0...

03mm: new :5me .055

mm

9.3a SN 882.8
<08 tam 2::
m _ c _ 2%

 

<08 32.2:qu

 

Figure 45. Study design for allopurinol effects on the development of DOCA-salt

hypertension.

123

allantoin

allantoIn (mg/day)
3 § $ $

uric acid (mg/day)

N=6

 

 

before agar

hypoxanthine

 

 

 

 

 

A 0.6!
a I.
2
or
E, 0.4-
o
.E
‘2 02.
S N=6
é - 4
"7 o.c . .
before after
allopurinol
10- *
it?
E 84
o:
E a
3
o.
2 2' N=6
co
0- ¢
before alter

xanthine (mg/day)

oxypurinol (mg/day)

ufine

 

 

 

 

uric acid
8-
-0- VEHICLE
6- -o- ALLOPURINOL
4-1
2.
N=6
C I I
before after
xanthine
20H
* -o- VEHICLE
-0- ALLOPURINOL
15-
10-
5.
N=6
c Y _
before after
oxypurinol

 

 

-O- VEHICLE
'0- ALLOPURINOL

 

before after

Figure 46. Urinary X0 metabolites in vehicle (closed symbols) and allopurinol
(open symbols)-treated rats, determined by HPLC: allantoin and uric acid (top,
X0 products); hypoxanthine and xanthine (middle, X0 substrates); allopurinol
and oxypurinol (bottom, X0 inhibitors), represented as mean :t SEM for N=6. *
represents a statistically significant difference (p<0.05) between treatment

groups.

124

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

serum
allantoin uric acid
150- 80-
.E .E N:
m .9
E N' 9 60-l -—
0.100. - ‘ * D. T
0’ I 2’
E 3° 40-1
o o
E (U
E 50‘ g
a :I 20‘
2 3
0'1 I 0- '
VEHICLE ALLOPURINOL VEHICLE ALLOPURINOL
hypoxanthine xanthine
C
'5 40‘ ,, so-
.. E
,2, "‘ s
c: 30‘ N: g 60d N=6 *
B I
.E 2’
g 204 g 40:
:2 E
3 10d 9 20‘
g; s
3. I I C I I
VEHICLE ALLOPURINOL VEHICLE ALLOPURINOL
allopurinol oxypurinol
c c 30-
E E _*_
_. 0
g N-6 a 20‘ N=6
g E”
0 0
.g .g
3 3
o. o. 104
g E‘
t0 0
33 c 3’ c
VEHICLE ALLOPURINOL VEHICLE ALLOPURINOL

Figure 47. Serum X0 metabolites in vehicle (closed symbols) and allopurinol
(open symbols)-treated rats, determined by HPLC: allantoin and uric acid (top,
X0 products); hypoxanthine and xanthine (middle, X0 substrates); allopurinol
and oxypurinol (bottom, X0 inhibitors), represented as mean :I: SEM for N=6. *
represents a statistically significant difference (p<0.05) between treatment
groups.

125

fissue
2oo- .

A 5 allantom - VEHICLE
% D ALLOPURINOL
L 150- _

Q N—6
O)
E
E 100-
5
75 so- 1,,
2 III
c _ .1.

 

 

I V U

1
AORTA VC CA JV LIVER KIDNEY HEART

 

1500- . .
B 5 LII'IC acud - VEHICLE
g 1250- D ALLOPURINOL
'5.
i’
(U
U
'C
3
3

 

 

AORTA VC CA JV LIVER KIDNEY HEART

 

 

 

 

C ,
-- 12 .
C % hypoxanthIne III - VEHICLE
a 6 El ALLOPURINOL
E’
a:
.E
5
C
i
.C
E
AORTA vc CA JV LIVER KIDNEY HEART
200 .
D (H xanthine - VEHICLE
D ALLOPURINOL
1500-

1000-

500‘

ug xanthine/g protein

 

 

A0 RTA VC CA JV LIVER KIDNEY HEART

Figure 48. Tissue X0 metabolites in vehicle (closed symbols) and allopurinol
(open symbols)—treated rats, determined by HPLC: allantoin (A), uric acid (B),
hypoxanthine (C) and xanthine (D) represented as mean 1 SEM for N=6. *
represents a statistically significant difference (p<0.05) between treatment
groups.

126

 

 

 

 

 

allopurinol DOCA
50 mglkg/day surgery
180- r
-o- VEHICLE
150. '0- ALLOPURINOL
E
E 140-
E
g 120-
100-
8G I l I I T I I l I
0 3 6 9121518212427

days

Figure 49. Blood pressure of vehicle (closed symbols) and allopurinol (open
symbols)-treated rats, measured by radiotelemetry. Timing of interventions is
represented by vertical lines. Data is represented as means 1 SEM of 24h
average mean arterial pressure (MAP) for N=6.

127

 

 

 

 

 

 

 

 

 

 

 

 

   

 

 

 

 

   

 

 

 

 

MAP Heartrate
A “’01 B
1604 N: A
A C
a ._
E 8
g a
o:
2 I
BC I I I I I I I I I 25c I I I I I I I I I
0369121518212427 036912151821242730
days days
SBP DBP
C m D 160-
g a
I
E E
E E
% o.
m
(D 0
1°C I I I I I I I I I I BC I I I I I I I I I I
036912151821242730 036912151821242730
days days
E Pulse Pressure F activity
A 80.1 A
53’ a
E '=
g 60- g
o as
L.
3 40- E
i g
5 E‘
20- .
g .2
‘5
a m
c I I I I I I I I r I c I I r I T I I I I I
036912151821242730 036912151821242730
days days
-0- VEHICLE

‘0' ALLOPURINOL

 

Figure 50. Hemodynamic parameters of vehicle (closed symbols) and allopurinol
(open symbols)-treated rats: mean arterial pressure (A), heart rate (B), systolic
blood pressure (C), diastolic blood pressure (D), pulse pressure (E) and activity
(F), represented as means 1 SEM of 24h average values for N=6.

128

HEART WEIGHT
2.01

>

N=6

 

1.5'l

1.0'

wet weight (g)

0.5-l

 

 

 

 

0.0‘

 

I
VEHICLE ALLOPURINOL

KIDNEY WEIGHT

m
j
I-

N=6 —H-—

 

wet weight (g)

14

 

 

 

 

 

r
VEHICLE ALLOPURINOL

Figure 51. Heart (A) and right kidney (B) weight at the time of sacrifice of vehicle
(closed bars) and allopurinol (open bars)-treated rats. Represented as mean :I:

SEM for N=6. * represents a statistically significant (p<0.05) difference between
treatment groups.

129

 

 

— _
‘1'? an)! m
. . - l a A
I "

1 ‘ ‘
I
_ _
.v' ' ,I J;
l .1 a‘

.. .
I
~ I.
,_ g I

P.

or- A." +1.“; ~-..

- -! M a
. . .
._.,
4“
-fl'

...". ‘1‘- A
‘ - l

I
.l'

,a
f
w .

. _ I ‘ _

f u '1'.
,r " ..? ’7
_ II
:'

.— JV:
t

Figure 52. Hematoxylin-eosin cross-sections of aorta from vehicle (A) and
allopurinol (B)-treated rats. Bars represent 100 pm.

130

III. Dose-dependent effects of allopurinol

For all in vivo experiments, we have used the same dose of allopurinol (50
mglkg/day). This treatment, despite inhibiting X0 activity, did not lead to
significant changes in the BP of DOCA-salt treated rats. Because a published
study (see discussion) reported decreased SBP in the DOCA-salt model with a
higher dose of allopurinol (100 mglkg/day), we investigated whether this dose

would provide a better inhibition of X0.

We administered vehicle, 50 mglkg/day allopurinol or 100 mglkg/day allopurinol
to normal rats for 7 consecutive days. At the end of this treatment, we collected

urine and serum samples and performed HPLC analysis of X0 metabolites.

Urinary allantoin and uric acid were not significantly changed between the three
groups (fig 53 top). A statistically significant increase in urinary hypoxanthine and
xanthine compared to vehicle was observed in both allopurinol-treated groups
(fig 53 middle). Similarly, allopurinol and oxypurinol were significantly increased
in the treated groups (fig 53 bottom). The effects on hypoxanthine, xanthine,
allopurinol and oxypurinol appeared dose-dependent, however the difference in
these metabolites was not statistically significant between the two allopurinol

treated groups.

Serum levels of allantoin were surprisingly increased in the 100 mglkg/day group
Compared to vehicle, while there was a trend for a dose-dependent decrease in
uric acid levels in the allopurinol-treated groups compared to vehicle. This

131

decrease was statistically significant in the 100 mglkg/day allopurinol group (fig
54 top). Hypoxanthine and xanthine levels increased in both allopurinol
compared to vehicle treated groups (fig 54 middle). 0xypurinol increased in the
treated groups, while allopurinol was close’to the detection limit and not
significantly changed between the three treatment groups (fig 54 bottom). The
effects on hypoxanthine, xanthine, uric acid and oxypurinol appeared dose-
dependent, however the difference in these metabolites was not statistically

significant between the two allopurinol treated groups.

Section summary: We have inhibited X0 activity in vivo, expecting as
consequences a lowering of MCFP and therefore of BP in the DOCA-salt
hypertensive model. We have monitored X0 metabolites in the urine, serum and
tissues of allopurinol-treated animals, overall observing increases in X0
substrates and decreases in X0 products. Despite what we considered effective
X0 inhibition, we have not observed any effect on BP of DOCA-salt hypertensive

rats, regardless of the timing of this intervention.

132

urine

    
  
  

 

 

allantoin uric acid
3 I VEHICLE
E 1% N=3-4 D ALLOPURINOL 50 mglkg/day
% 73, ALLOPURINOL 100 mglkg/day
E .E.
.1: f8
.2 m
g .2
E ‘5
hypoxanthine xanthine
B - VEHICLE
E A D ALLOPURINOL 50 mglkg/day
% a; . ALLOPURINOL 100 mglkg/day
.E, ‘a’.
g E.
g .2
.E
S E
g S
.C
allo urinol o urinol
p xyp I VEHICLE
A A 1 D ALLOPURINOL 50 mglkg/day
5 5‘ t ALLOPURINOL 100 mglkg/day
B: B:
E, .E,
6 T)
a. o.
g s?
a o

 

Figure 53. Urinary X0 metabolites in vehicle (closed bars), 50 mglkg/day
allopurinol (open symbols), and 100 mglkg/day allopurinol (hatched symbols)-
treated rats, determined by HPLC: allantoin and uric acid (top, X0 products);
hypoxanthine and xanthine (middle, X0 substrates); allopurinol and oxypurinol
(bottom, X0 inhibitors), represented as mean :I: SEM for N=3-4. * represents a
statistically significant difference (p<0.05) compared to vehicle.

133

serum

allantoin ”"0 30”

  

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

- VEHICLE _
15o. : 40-1 N=3-4 D ALLOPURINOLSO mglkg/day

7% * 'é ALLOPURINOL 100 mglkg/day
'5 N=3-4 9 30-
3. 1m.) 0.
OI 9
E 7% 20-
5 so .0
a g 10"
9 33

o- o-

hypoxanthIne xanthIne - VEHICLE
% 40- * 4% D ALLOPURINOL so mglkg/day
g N=3-4 .g N=3-4 * . ALLOPURINOL 100 mglkglday
°° °°' E 3" l
B 3
S ‘
E 2* 2 2r
3 E
g 101 3 1m
2 U)
1
g c I I c I l
allopurinol oxypurinol I VEHICLE
025.1 10- D ALLOPURINOL 50 mgfitglday
5 ~ .E w. ALLOP RIN 100
2 N=3-4 g N 34 . I U 0L make/day
02m 8,. =

g ,_—L'.'_ g
a: on
B 0.15- % 6-
1;; 0.10. E 44
£1 0.05- % 2-
? 0111-i , g 0* f

Figure 54. Serum X0 metabolites in vehicle (Closed bars), 50 mglkg/day
allopurinol (open symbols), and 100 mglkg/day allopurinol (hatched symbols)-
treated rats, determined by HPLC: allantoin and uric acid (top, X0 products);
hypoxanthine and xanthine (middle, X0 substrates); allopurinol and oxypurinol
(bottom, X0 inhibitors), represented as mean 1 SEM for N=3-4. * represents a
statistically significant difference (p<0.05) compared to vehicle.

134

DISCUSSION

Aside from likely contributing to many aspects of normal vascular biology,
reactive oxygen species are thought to play an important role in cardiovascular
disease, and in particular, in hypertension. Veins and the contribution of
venomotor tone to blood pressure control represent an understudied research
area. In this study, we investigated the metabolism of ROS in veins as compared
to arteries, their contribution to vascular contraction function, and finally their
potential involvement in hypertension pathogenesis. A particular focus was
placed on the role of xanthine oxidase. The male Sprague-Dawley rat DOCA-salt
model of hypertension was used in these experiments, for reasons detailed in the

introduction section.

I. ROS metabolism in veins vs arteries
A. Basal levels of ROS

We started our comparison of venous and arterial ROS metabolism by
measuring the levels of two of the most important ROS: superoxide and H202.
Several distinctions can be made between these two species. The half-life of
superoxide is much shorter (ms) than that of H202 (minutes-hours). Superoxide
local subcellular concentrations are highly dependent on the local concentration

of superoxide dismutase ($00), the enzyme that will readily metabolize it to

135

H202. 0n the other hand, superoxide is charged and therefore cannot freely
cross the cell membrane, while the longer-lived H202 is membrane-penneable,
extracellular concentrations measured at equilibrium being thus similar to
intracellular ones. Both superoxide and H202 may critically influence vascular
functions, and especially the contractile function that was at the core of this

research.

We observed that in normal conditions, basal levels of both superoxide and H202
are higher in V0 compared to aorta (fig 11-12). We have also observed higher
superoxide levels in the superior mesenteric vein compared to the superior
mesenteric artery, and higher H202 levels in the jugular vein compared to the
carotid artery. Our results appear therefore to reflect a general artery-vein
difference, rather than a vascular bed-specific characteristic. This difference
could result from a higher rate of ROS production, a lower rate of ROS

degradation, or both, in veins versus arteries.

We have measured superoxide in the presence of SOD inhibition, so that only
differences in production, but not destruction of superoxide would be taken into
account. Additionally, the use of SOD inhibition for superoxide detection also
increases the magnitude of the superoxide signal, thus increasing the sensitivity
of this method. The majority of H202 being produced in basal conditions
appeared as superoxide-derived, as it was almost abolished by SOD inhibition
through DDC. A limitation regarding our use of DDC is the fact that this copper
chelator compound would only inhibit the two Cu-containing SOD isoforms, CuZn

SOD and EC-SOD, but not Mn SOD, the mitochondrial isofonn (55). However,

136

we observed no arterio-venous differences in Mn SOD, at least in terms of its
protein expression (fig 24). Few comparisons of basal ROS levels were reported
in the literature (see introduction section V). Thus, our study provides the only
direct comparison of paired rat blood vessels ROS production in basal

conditions.

An additional interesting arterio-venous comparison would be on ROS levels in
the presence of contractile agonists that typically also stimulate ROS production
(such as Angll or ET—1). In preliminary studies, we were not able to observe
increases in whole tissue H202 in the presence of agonists. Further studies
comparing agonist-stimulated ROS levels in arteries and veins may have to take
into account the potential limitations of our H202 measurements, that use whole

tissues and omit the addition of specific substrates.

B. ROS degradation

ROS production by living cells is normally counterbalanced by the activity of
several antioxidant defense mechanisms such as SODs, catalase or glutathione
peroxidases. In order to compare the status of these mechanisms between aorta
and V0, we exposed vessels to exogenous H202 and assessed how much of that
H202 is metabolized in the presence of tissue. We chose H202 and not
superoxide for this purpose because H202 is more stable, capable of freely
passing cellular membranes, easier to generate at definite concentrations and

also more reliable to measure. We used H202 at a high enough concentration to

137

produce sufficient fluorescence such that its decrease in the presence of tissue
would be measurable with our method. However this concentration (20 pM) may
not reflect a physiological situation (~2.5 pM in normal plasma (4)), but rather
mimic a state of oxidative stress or a localized increase in H202 concentrations
such as the ones observed during the neutrophil oxidative burst. We have
observed a higher consumption of H202 by the VC compared to the aorta,
indicating a higher activity of antioxidant defense mechanisms, and possibly
reflecting the increased ROS destruction in normal VC balancing the increased
ROS production. However, in our results, the V0 to aorta ratio of ROS generation
(~15 times greater) does not numerically match the ratio of ROS destruction (~4
times greater), raising the question whether this is indeed a perfect balance or
the V0 is in fact exposed to oxidant stress. This method of measuring antioxidant
capacity by exogenous H202 exposure does not discriminate among specific
antioxidant mechanisms, although catalase is likely to be more important than
glutathione peroxidase at high H202 concentrations due to its higher Km (181).
This would be consistent with the higher catalase expression that we observed in

V0 compared to aorta (fig 19).

C. ROS metabolizing enzymes

ROS metabolism is a highly complex biological pathway with many important
enzymatic and non-enzymatic players involved. We have attempted to

investigate the profile of ROS producers that may contribute to the differences

138

observed between arterial and venous ROS levels. We used three different

approaches, with only partially overlapping results.
1. Protein expression

The protein expression analysis performed here (fig 16-19) is by no means
complete. Due to unavailable tools, we could not confidently assess the
expression of different Nox isoforms and various subunits of NADPH oxidase, a
major ROS producer repeatedly proven as important in vascular biology. In
human vessels, protein as well as mRNA expression of NADPH oxidase subunits
was reported as significantly different between the saphenous vein and internal
mammary artery. Additionally, a literature report suggested enzymatic activity of
NADPH oxidase to be contributing to total superoxide levels in a higher degree in
the vein than in the artery, (51). Also absent from our results are an entire range
of other ROS metabolizing systems that may also contribute to the differences
observed in ROS levels such as: other mitochondrial ROS production sites
(complex lll), uncoupling proteins, other isoforms of NO synthase (nNOS, iNOS),
Iipoxygenase, cyclooxygenase, myeloperoxidase, heme oxygenase, the
extracellular isoform of SOD, glutathione peroxidases. A comparison of SOD
expression in human saphenous vein and internal mammary artery reported
similar expression of the cytosolic (CuZn SOD) and mitochondrial (MnSOD)
isoforms (50). To our knowledge however, there are no other studies comparing
the arterial and venous expression of eNOS, an important enzyme in vascular
biology, that when uncoupled becomes capable of producing superoxide. We did

not find differences in the expression of eNOS between aorta and VC, correlated

139

with the similar endothelium-dependent relaxation found in these tissues in
normal conditions (fig 34). However, from the differences we did observe in
protein expression between aorta and V0, only X0 and DUOX1, as superoxide
producers, would help explain the higher superoxide levels found in basal
conditions in veins. Furthermore CuZn SOD, a cytosolic enzyme that converts
superoxide to H202, and catalase, responsible for further degrading H202, were
both more heavily expressed in the vein, potentially correlated with the higher

superoxide/ H202 levels.

Any protein expression analysis does not entirely reflect protein activity in living
tissues. Therefore, to fully explore arterial versus venous ROS metabolizing

systems, more studies would be necessary.
2. mRNA expression

The PCR array used for this experiment queried only a small number of probes
quantifying mRNA expression of enzymes tested in the western blot analyses
discussed above. Of them, DUOX1 had a higher mRNA expression, correlating
with the increased protein expression. No difference was observed however in
the mRNA expression of catalase and any SOD isoform, while X0 was not
present on this array. The lack of correlation between protein and mRNA levels
using broad expression profiling methods was previously described (47, 49) and
may be attributed to posttranscriptional events, posttranslational modifications,
variations in mRNA half-life and in protein stability, as well as technical errors.

Both DUOX isoforms higher venous mRNA expression was confirmed by RT-

140

PCR analysis. Of potential interest was also the higher expression of Nox4 in
aorta compared to V0. This result however, even if correlated with a higher
protein expression and activity of Nox4 in arteries, would not help explain the

higher ROS levels observed in veins and was not explored further.
3. Pharmacological inhibition of basal H202 levels

We observed higher in vitro basal H202 levels in veins than in arteries, when
measured in whole tissues. A logical means of identifying the source of this
increase would then be to inhibit potential sources in the same settings and
assign contributions of various sources to these basal ROS levels. Literature
reports on whole tissue ROS production are scarce compared to those on cell
cultures or cell fractions and to our knowledge, this particular type of assay was
not previously attempted for tissue production of H202. Rather, most
investigations tend to assess the activity of one or other of ROS producers in the
presence of specific substrates (such as NADPH for NADPH oxidase activity). In
these experiments, we used tissues originating in the same animal to quantify
inhibition as a percentage of the respective basal level. We did not use specific
substrates for any of the enzymes assessed. Moreover, we did not use specific
inhibition of catalase to amplify the H202 signal in these experiments,
hypothesizing that a potentially higher venous catalase activity may be impacted
differentially by catalase inhibition, thus biasing the results towards more efficient
reductions in H202 levels in arteries. The basal H202 levels in the two arteries
used (aorta and carotid artery) were often not measurable, thus making inhibition

observations in arteries impossible and lowering the experimental statistical

141

significance. We chose drug concentrations to use based on multiple published
results demonstrating efficient in vitro enzyme inhibition. Apocynin decreased the
basal H202 levels in most tissues, suggesting that NADPH oxidase was a
significant source of basal vascular ROS, as noted by previous reports. However,
recent studies have questioned the specificity of apocynin action, which was
instead shown to have antioxidant effects by non-specifically scavenging ROS
(56). The same potential scavenging effect was reported for indomethacin (36), a
non-specific COX inhibitor, which in our hands had no effect on basal H202
levels. Ketorolac, another non-specific COX inhibitor, decreased venous H202
levels, thus rendering the COX inhibition experiments overall inconclusive. Based
on our X0 expression analyses, we were expecting to observe a significant
decrease in basal venous ROS levels with allopurinol. This however did not
occur for VC, but only for JV. The higher venous compared to arterial X0 activity
was however later shown in separate experiments, performed in the presence of
the X0 substrate, xanthine (fig 26). Surprisingly, venous H202 levels were
significantly decreased by rotenone, a mitochondrial'complex l inhibitor. A trend
for decreased venous H202 levels was observed with DPI, a flavoprotein
(NADPH oxidases including DUOX, xanthine oxidase and eNOS) inhibitor.
Inhibition of NOS with LNNA did not significantly alter basal H202 levels of any
tissue, suggesting that in normal conditions, superoxide production from
uncoupled eNOS is not a major phenomenon. Overall, because of the absence
of enzyme substrates and the lack of more specific inhibitors, we believe that the

conclusions to be drawn from these experiments should be limited. More in depth

142

analyses of in vitro H202 production are needed and may include future
interesting studies incorporating agonist-stimulation of H202 production in the

presence of the ROS enzyme substrates and inhibitors.

D. Oxidative stress in arteries vs veins

We have observed increased basal ROS levels in veins compared to arteries,
despite increased venous vs arterial antioxidant capacity. These data suggest
that production of ROS in veins is much higher than that in arteries. Also, these
results raise the question of the function or effects of the ROS that, regardless of
their molecular source, are being produced, but are not being degraded in these
blood vessels. When production exceeds degradation of ROS, leading to
accumulation of deleterious levels of ROS, the state of oxidative stress ensues.
However, measurable levels of ROS and markers of oxidative stress are also
present in normal conditions. We hypothesized that these markers may be
accordingly higher in veins compared to arteries. We used 4-HNE, a marker of
lipid peroxidation, in V0 versus aorta. We have observed a more intense general
4-HNE staining in the VC compared to the aorta. 4-HNE can also form covalent
bounds to proteins (113). However when attempting to quantify the 4-HNE
protein adducts through western blotting, these appeared to have a less intense
banding pattern in veins versus arteries. The use of other oxidation markers (see

introduction) may therefore be needed to allow stating a clear conclusion

143

regarding lipid peroxidation or general oxidative stress in normal veins vs

anenes.

E. ROS metabolism in DOCA-salt hypertension

Increases in ROS generation and markers of oxidative stress were reported in
virtually all forms of hypertension, human and animal models. Similar to the
situation in normal rats, little information is available on differences between the
arterial and the venous ROS metabolism during hypertension. We have observed
that the increased production of ROS by veins vs arteries is maintained in
DOCA-salt hypertension. However, we were not able to confirm the increased
basal H202 levels within the same type of tissue with hypertension in the absence
of substrates (e.g. sham compared to DOCA aorta). However, this does not
exclude the possibility that given specific substrates for various ROS enzymes,
ROS production would not be increased with hypertension. We would also
expect the response in tissue ROS production to the presence of contractile
agonists (such as Angll or ET-1) to be amplified in tissues from hypertensive .
animals. We have not observed a significant change in the protein expression
pattern of ROS producers and destroyers between sham normotensive and
DOCA-salt hypertensive rats in aorta or VC samples, suggesting that potential
changes observed between arteries and veins during hypertension would not be

supported by changes in protein expression of these ROS enzymes, and may be

144

attributed to changes in protein activity, as well as to changes in other mediators

of vascular function affected by hypertension.

F. Xanthine oxidase

A significant focus was placed on X0 throughout this study due to the in vitro
results of X0 expression and activity in venous vs arterial tissues (figs 16, 25-

27). Most of these results have been published (147).

Circulating X0 binds to glucosaminoglycans on the vascular endothelium, still
retaining specific activity (62). The results presented in figure 25 represent a
novel finding in that they demonstrate for the first time that blood vessels produce
the X0 mRNA locally, and that binding of circulatory X0 to endothelial cells is not
the only mechanism responsible for the detection of X0 activity in these tissues.
We observed a higher protein and mRNA expression, as well as a higher activity
of X0 in V0 than in aorta from normal rats. It should be noted here that our
mRNA and protein expression studies do not distinguish between xanthine
dehydrogenase (XDH) and X0, the two isoforms of xanthine oxidoreductase
(XOR). These isoforms are generated by posttranslational modifications (11),
therefore they have the same mRNA source; because of the small difference in
protein structure between XDH and X0, antibodies will also recognize both
isoforms. However, our X0 activity assays specifically (fig 26) assess the
enzyme activity that uses oxygen (X0), as they are performed in the absence of

NAD“, the substrate for XDH. The differences in X0 expression between the vein

145

and the artery could be mediated by a number of factors and processes
regulating X0 expression such as cytokines or oxygen tension (9). An interesting
report in bovine aortic endothelial cells, which appears to be consistent with our
findings, has implicated a feed-forward mechanism by which increases in H202
stimulate the conversion of XDH to XO, potentially leading to even higher H202

levels generated by X0 (99).

No effect on X0 expression or activity was observed with hypertension between
sham and DOCA aorta samples (fig 27). One study reported an increase in
xanthine-stimulated superoxide production in the DOCA aorta, implying an
increased X0 activity (163). We did not obtain results leading to the same
conclusion, although we used two separate specific measures of X0 activity in
the aorta. However, DOCA VC had a higher X0 activity (urate-producing) than
normal VC, despite the lack of protein expression change. DOCA jugular vein
had also a higher X0 activity (H202-producing) than sham jugular vein. These
results would support the idea that during hypertension, although overall
(XDH+X0) expression levels are similar, there is an increase in the X0 to XDH
ratio, perhaps caused by an increased XDH-X0 conversion, supporting

increases in X0 activity that are not paralleled by increases in expression.

It is important to also note the lack of numerical correlation between the two
activity assays. The stoichiometric relationship between the two X0 products
depends largely on the presence of other substrates, and is largely inclined

towards urate. Moreover, these two tests were performed in different types of

146

samples: we used tissue homogenates for the urate-based assay, and whole

tissues for the H202-based assay.

ll. Impact of ROS on venous and arterial contractile function
A. Nomal conditions
1. ROS-mediated contraction

We have observed that both the aorta and the VC from normal rats respond to
exogenous superoxide with contraction, and that this contraction is higher in V0
than in aorta, as a percentage of adrenergic responses. The combination of
xanthine with X0 has long been used as a laboratory tool to produce superoxide.
Some researchers have suggested that superoxide can enter living cells through
Cl’ channels (63). However, the accepted view in the ROS field is that because it
carries a negative charge, superoxide does not cross membranes. Therefore we
can assume that contraction induced by extracellular xanthine/X0 is mediated
either by extracellular superoxide or by H202 as the product of superoxide
degradation, a longer-lived molecule that can freely diffuse across membranes.
The contraction we observed with xanthine/X0 is indeed parallel to the H202
contraction we previously observed in the same tissues (150, 151). However, to
effectively claim this, experiments with xanthine/X0 should be performed in the
presence of catalase. The mechanisms for the direct contractile effects of ROS
are complex, involving among others an interference with endothelium-released
N0 (2, 13). In our experiments, removal of endothelium did not alter aortic

147

contraction to exogenous superoxide (data not shown). The fact that the
contraction induced by xanthine/X0 was higher in V0 would suggest that any
alteration of ROS metabolism is likely to have a more important impact on

venous rather than arterial contraction.

2. X0 impact on normal vascular contractile function

Smooth muscle contraction signaling pathways may be modulated by ROS in
many ways. Some contracting agents such as angiotensin II (Angll) are widely
known as being capable of inducing superoxide release (48). General ROS
scavengers, catalase or specific ROS enzyme inhibitors have variably decreased
the arterial contractile response induced by agonists such as NE, Angll or ET-1
(92, 137, 156). ET-1, a potent venoconstrictor, also stimulates superoxide
production. Similarly to the Ang Il-induced superoxide production, the mechanism
mostly responsible for this increase in superoxide release seems to be NADPH
oxidase activation (86). We asked specifically whether XO-produced ROS could
contribute to the normal contraction induced by NE, Angll or ET-1 and normal
relaxation induced by ACh. Inhibition of X0 activity with allopurinol did not alter
any of the aortic responses to these agonists (fig 29). However, the maximal ET-
1-mediated contraction of the VC was significantly decreased in the presence of
allopurinol. These data suggest that X0 contributes to ET-1-mediated
venoconstriction in a way that it does not in the aorta. Although we did not see
effects on Angll-induced contraction in our study, long-term exposure to Ang II

can also induce X0 activation in cultured endothelial cells (83).

148

X0 may theoretically influence endothelial function in at least one other way
besides ROS production, specifically through the production of uric acid. Uric
acid may impact endothelial function at least by the following mechanisms: a. by
directly interacting with NO (46) and therefore decreasing its bioavailability; b. by
interacting with superoxide (7) and therefore decreasing the amount of
superoxide available to quench N0, thus increasing N0 bioavailability; c. by
interacting with peroxynitrite, the reaction product of superoxide and NO, to form
a product shown to have vasorelaxant properties (133) or to decrease
peroxynitrite availability. Peroxynitrite, in its turn, was shown to decrease
guanylate cyclase activity and oxidize BH4, an important cofactor for eNOS
activity (127). Paradoxically, peroxynitrite also presents vasorelaxant properties
(172). Additionally, uric acid may have antioxidant or other effects intracellularly
in vascular smooth muscle cells, where it is taken up by a specific transporter

(115).

However, when we inhibited urate production with allopurinol, we did not observe
any changes in the ACh-induced relaxation of either aorta or VC from normal
rats. Moreover, uric acid itself did not induce any direct contractile effects on rat
aorta or VC and it did not modify ACh-induced relaxation in any of these tissues.
We used uric acid at concentrations previously proven to have antioxidant
properties in vitro, and within the range of normal uricemia (7). Additionally, we
used oxonic acid at a concentration proven to inhibit uricase activity (42),
therefore preventing degradation of uric acid. We have not tested whether uric

acid would directly induce relaxation of PGan—contracted rat aorta, but the lack

149

of direct vasodilatory effects of uric acid alone in rat aorta was previously

reported (133).

We also assessed protein expression of uricase in aorta and V0 as the 64 KDa
band observed in our samples. This molecular weight was previously observed
and attributed to a uricase dimer (162). We observed an increase in the VC
uricase expression, compared to aorta (fig 31). This finding correlates with our

other results showing a higher urate-producing activity in V0 compared to aorta.

Since contraction to ET-1 was diminished in the presence of allopurinol, and
expression of DUOX1 was increased in the VC, we tested the effects of a
general flavoenzyme inhibitor, DPI, which should block both X0 and DUOX1
activities. Unfortunately, no specific pharmacological inhibitor for DUOX1 is
available presently. As expected, we observed a significant decrease of the ET-
1-mediated contraction of VC, but not aorta. These data suggest that the
contribution of flavoenzymes such as X0 and DUOX1 to ET-1-mediated
contraction is higher in V0 than in aorta. These results do not however exclude
the contribution of any other Nox isoform or any other flavoprotein to ET-1-

induced contraction of these blood vessels.
3. Normal arterial and venous endothelial function

Published studies have demonstrated lower venous vs arterial NO production in
some vascular beds (130). Moreover, there is a widely known interaction
between superoxide and N0 with subsequent decreases in N0 bioavailability (8),

and in our experiments we observed higher venous than arterial superoxide

150

levels. In this context, we hypothesized that there would be decreased relaxation
to ACh in normal venous vs arterial tissue. However this was not the case, as
both aorta and VC relaxed similarly to ACh, and both fully depended on NOS

function for this effect (fig 34).

B. DOCA-salt hypertension
1. Arterial and venous endothelial function in hypertension

Arterial endothelial function is characteristically impaired in human and animal
models of hypertension (144). The DOCA-salt hypertensive rat is no exception,
DOCA aorta exhibiting endothelial dysfunction, as evidenced by an impaired
relaxation to ACh. DOCA VC, however, maintained its normal ACh-mediated
responses. Both aorta and VC were still fully dependent on NOS function for the
ACh-mediated relaxation (fig 35). The mechanisms behind the loss of arterial
endothelial function in hypertension extend beyond the obvious detrimental effect
of pressure increases. The fact that VC was not similarly affected to aorta in
hypertension may be attributed to V0 not being exposed to the same regime of
increased pressure that the aorta is. However a role may also be played by
differences in the anti-inflammatory or other properties of VC that enable it to
handle the changes in deleterious circulating mediators that may be associated

with hypertension.

2. X0 impact on arterial endothelial function in hypertension

151

 

Many reports show improvements in endothelial function in vitro and in vivo by
inhibition of X0 activity, sometimes independently of other effects (14) but mostly
in the context of lowering blood pressure or improving end-organ damage (110,
141). It is however impossible to distinguish between direct effects of allopurinol
on endothelial function and the effects of simultaneous lowering of blood
pressure. Additionally, in these studies, the effects of allopurinol on lowering uric
acid cannot be discerned from effects on lowering ROS/oxidative stress. A
convincing attempt was made in this direction in a clinical study which
established that ameliorating endothelial dysfunction in chronic heart failure with
allopurinol was accomplished by a uric acid-independent mechanism, since
lowering uric acid to similar levels with the uricosuric agent probenecid did not
have any effect on endothelial function (45). The authors therefore concluded
that endothelial function was improved by allopurinol because of a reduction in
XO-mediated ROS production. We observed no effect of allopurinol on the

DOCA-salt rat aorta endothelial dysfunction (fig 36).

Moreover, no effect on DOCA aorta ACh-induced relaxation was observed with
direct administration of uric acid, suggesting the lack of acute antioxidant effects
of uric acid (fig 37). Uric acid, administered acutely (over 1h) at a dose that
raised uricemia from normal to the hyperuricemic range, ameliorated the
endothelial dysfunction of smokers and patients with diabetes type I, without an
impact on the normal endothelial function of control subjects, although the serum
antioxidant capacity was equally increased in all groups after the administration

of uric acid (166). This would imply that the beneficial antioxidant effects of uric

152

acid may only become apparent when there is a pathophysiological excess of
oxidants and consequently endothelial dysfunction. We excluded uricase
degradation of uric acid in the rat tissue by co-administering oxonic acid, the
uricase inhibitor. However, besides the presence of uricase in rat tissues, other
species differences cannot be ruled out, such as differences in the presence
and/or function of specific urate transporters in rodents compared to humans.
Additionally, the endothelial dysfunction observed in type I diabetes (the
condition in which urate had beneficial effects) may be different from the
endothelial dysfunction associated with hypertension (the condition we studied),
in which pressure-associated shear stress and endothelial injury may be less
ROS-dependent. In vivo treatments with various nonspecific antioxidants have
been shown experimentally to improve hypertensive endothelial dysfunction
(157), however it is difficult to dissociate the direct effects of antioxidants on
endothelium from the effects of the accompanying reduction in blood pressure

upon antioxidant treatment.

One of our hypotheses regarding uric acid was that during DOCA-salt
hypertension uricemia is decreased and the chronic lack of adequate levels of
uric acid contributes to the endothelial dysfunction of hypertension. Uric acid
levels in blood however were similar in hypertensive compared to normotensive
rats (fig 38) and we could not establish any correlation between individual values
of uricemia and blood pressure. This is a known association in humans (77),
hypertensives having increased risk of gout and hyperuricemia increasing the

risk for hypertension. As previously explained, rodents have significantly lower

153

uricemia levels than humans, as a result of the presence of functional uricase,
the enzyme that degrades urate into allantoin, and that is not expressed in
humans as a result of a mutation (178). Similarly to the expression of X0, uricase
expression was also not changed in aorta from hypertensive compared to

normotensive rats (fig 39).

Therefore, contrary to our hypothesis for these experiments, uric acid and
inhibition of its main producer, X0, did not change the endothelial function of
aorta from either normotensive or hypertensive rats. Our results do not, however,
exclude the long-term involvement of uric acid in human vascular function
through either its antioxidant, mitogenic or still uncovered properties. In the
context of the beneficial vs. deleterious uric acid effects controversy, one has to
be left wondering whether the evolutionary loss of uricase in higher primates and
humans may be explained by hyperuricemia conferring a physiological

advantage.

Yet, even if proven effective on endothelial function or as a general antioxidant,
the desirability of any hyperuricemic treatment has to be questioned, given the
controversial evidence and the clear pathophysiological associations of

hyperuricemia in humans.

III. Results of in viva X0 inhibition in DOCA-salt hypertension

Given the in vitro differences seen in X0 expression and activity between veins
and arteries, the differences in the impact of X0 on contractile function, as well

154

as the increase observed in venous X0 activity during DOCA-salt hypertension,
we expected that adequate inhibition of X0 activity in vivo would have more
important effects on the venous than the arterial system. Based on the idea that
venomotor tone is a contributor to BP regulation, we hypothesized that if we
influenced venomotor tone significantly by decreasing venous XO-mediated ROS
levels and effects on veins, we would decrease BP during the established phase
of DOCA-salt hypertension or attenuate the increase in BP in the initial phase of

DOCA-salt hypertension.

We therefore administered allopurinol, the most common X0 inhibitor, to DOCA-
salt treated rats and compared the effects of this treatment with those of vehicle

treatment.

We chose a mid-range dose of allopurinol, previously proven to be effective in
inhibiting X0 activity in rats in vivo (37) and the oral route of administration, for

reasons explained above (see Results aim 3).

A. Validation of X0 inhibition

Rather than only performing serum urate measurements or performing X0
activity assays ex vivo on select tissues, such as the liver or our target tissues,
the veins, we chose a more comprehensive approach and investigated the
alterations in X0 metabolites caused by X0 inhibition in biological fluids (urine,

serum) and several vascular (arteries and veins pairs and the heart) and other

155

organs (liver and kidney). This approach however precluded simultaneous

measurements of ROS levels in veins following allopurinol treatment.

Allopurinol is administered to humans with hyperuricemia and gout with the
intention of lowering serum urate levels below the solubility limit of urate (6 mg/dl
or 360 pM). Until recently, allopurinol was the only X0 inhibitor approved for use
in humans. The allopurinol dose used in humans is low in comparison with the
one used in typical rodent research (approx 1.25 — 3.75 to as high as 10
mglkg/day in humans compared to 10-100 mglkg/day for rodents), especially
considering the aforementioned lack of uricase activity in humans and
comparatively higher basal serum urate levels. This is partly due to adverse
reactions observed at higher allopurinol doses (135). Successful treatment is
evaluated based on whether a decrease is observed in serum urate levels on an
individual basis. Statistically, this is variably achieved in ~20 up to 80% patients
with a typical dose of allopurinol (300 mg/day), and there is a high percentage of

non-compliance to treatment (171).

Allopurinol administration to rats is typically done without regard to
cardiovascular parameters such as BP. Several published reports do not contain
any type of validation for achieving X0 inhibition with the chosen dose of
allopurinol, while most show either a decrease in serum urate levels or in the X0

activity of liver (37, 107, 163).

A few studies reported analysis of urine or serum X0 metabolites by HPLC (19,

116, 159), however to our knowledge no such analysis was performed for tissues

156

from all0purinol-treated animals. We expected decreases in urate levels and
potential accumulations in xanthine/hypoxanthine, paralleling the findings in

human allopurinol administration.
1. Allopurinol administration in established DOCA-salt hypertension

When we administered allopurinol for 7 days at the end of the DOCA-salt
treatment, we observed overall no significant changes of urate and allantoin in
urine, a trend for their decrease in serum, as well as decreases in urate in tissue.
This was accompanied by highly significant increases in hypoxanthine and
xanthine, the X0 substrates in urine, and serum and of hypoxanthine in tissues.
We attribute the increases observed in hypoxanthine and xanthine to effective
blockade of X0 function, leading to accumulation of substrates. We validated the
drug administration and effect on X0 by also observing not only allopurinol, but
also oxypurinol in both urine and serum of treated rats (fig 41-43). The longer
half-life of oxypurinol is what likely explains the higher levels of oxypurinol

compared to allopurinol detected in all our analyses.

2. Allopurinol administration in the developments of DOCA-salt

hypertension

When administered from the beginning of DOCA-salt treatment, allopurinol had
slightly different effects on X0 metabolites as compared to the shorter (7 days)
administration. We still did not observe significant decreases in urine and serum
uric acid in allopurinol-treated rats, however urate levels decreased significantly

in tissues. Similarly to results from the 7-day treatment, the levels of substrates

157

 

(hypoxanthine and xanthine) increased significantly in serum, urine and tissues.
Contrary to the 7-day treatment results, when no significant changes were
observed in tissue xanthine levels, we now observed a significant increase in
xanthine levels in several tissues. A potential explanation for this phenomenon
that would also take into account the opposite effects with hypoxanthine would
be the feed-back on substrate levels in the context of long-term enzyme
inhibition. Theoretically this may be achieved by modulation exerted on guanine
deaminase, an enzyme that contributes to xanthine, but not hypoxanthine levels.
Treatment validation was again obtained by observing levels of allopurinol and,

more importantly, oxypurinol in urine and serum (fig 46-48).
3. Dose-dependency of allopurinol effects

Because we did not observe BP effects with 50 mglkg/day, as opposed to
another published study using DOCA-salt hypertensive rats (163) in which
decreases in SBP were achieved with 100 mglkg/day allopurinol, the question
arose as to whether the lack of effects in our study was due to insufficient X0
blockade and whether we could have inhibited X0 activity more effectively with a
higher dose of allopurinol. Before repeating BP studies in the DOCA-salt model
with the higher allopurinol dose, we examined the comparative effects on X0
metabolites of the two allopurinol doses: 50 and 100‘mg/kglday. Overall, these
experiments demonstrated that the increases observed in hypoxanthine,
xanthine, allopurinol and oxypurinol in urine and serum were dose-dependent.
Numerically, these results did not overlap with those obtained in BP studies,

probably due to the fact that BP studies were conducted in significantly older,

158

DOCA-salt treated rats. Serum urate decreased significantly with the higher
allopurinol dose, however this was not significantly different from the lower dose.
Moreover, the 100 mglkg/day dose was previously reported as renal toxic in rats
(142, 143), and this nephrotoxicity appeared enhanced by hypertension (158).
Additionally, the DOCA-salt model used in the study with positive allopurinol
results on BP (163) was different from ours, using a higher dose of DOCA (250
mglkg as opposed to our 200 mglkg). Moreover, measurements of BP in this
study were made using the tail-cuff method, far less precise than the
radiotelemetric one we employed. Therefore we concluded that repeating the BP
studies with a higher dose of allopurinol was not needed, as likely little effects
could be expected on BP with similar levels of X0 inhibition. However, that does
not exclude the benefits of potential experiments using a different, more specific

pharmacological inhibitor or another method for X0 inhibition.

B. Effects of allopurinol on BP in the DOCA-salt model of hypertension

We observed no effects on MAP with 50 mglkg/day allopurinol, regardless of the
timing of X0 inhibition. A significant decrease in pulse pressure was however
observed following allopurinol treatment. This hemodynamic parameter was
independently associated with cardiovascular risk in humans (154). The
decreased pulse pressure may therefore be a beneficial effect of allopurinol
treatment, highlighting a potentially important role of X0 aside from its direct

impact on BP. Other secondary parameters observed with allopurinol

159

 

administration in the development of DOCA-salt hypertension were either not
changed or the change was opposite to the one expected. More specifically,
there was an increase in DBP, no change in heart weight and no qualitative
changes in arterial remodeling (although quantitative comparisons of remodeling
were not made). Additionally, we observed a surprising increase in kidney
weight. We measured kidney weight as a gross measure of hypertensive
nephropathy, however this may also have changed due to adverse renal effects

of allopurinol (60, 142, 143).

The lack of effects on MAP with allopurinol treatment can have several
explanations. We believe we excluded the explanation of an inefficient blockade
of XO with the chosen dose of allopurinol by performing the additional studies
comparing 50 with 100 mglkg/day. However, due to additional specificity
questions regarding the use of allopurinol (148), this explanation would be fully
excluded only by inhibiting X0 with different means: another pharmacological
inhibitor, such as the newer, non-purine based febuxostat; or another method,
such as in vivo siRNA inhibition of X0, although the latter may be hampered by a
number of other factors that typically impact in vivo RNAi experiments.
Alternatively, even though X0 inhibition may have successfully decreased the
venous ROS levels in vivo, one or all of the hypotheses made based on in vitro
findings may prove wrong in vivo: either the contribution of X0 to overall ROS
levels, or that of ROS to venomotor tone, or potentially that of the venomotor
tone to BP control in the DOCA-salt hypertension model may have been

overestimated.

160

IV. Conclusions

Unlike most studies on the role of ROS in vascular biology and/or hypertension
pathogenesis, we took a venous-centric approach. We compared the metabolism
of ROS as well as the functional implications of ROS to vascular contractility in
veins vs arteries in normal conditions. We then tested the hypothesis that
inhibition of XO-mediated ROS production would lead to a protective effect on BP

in the DOCA-salt hypertensive rat model.

We observed an overall higher ROS environment in veins compared to arteries:
higher basal levels of superoxide and H202 were accompanied by higher
antioxidant capacity. Expression and/or activity of several ROS metabolizing
enzymes was higher in veins vs arteries: X0, DUOX1, CuZn SOD and catalase.
The functional impact of ROS was also higher in veins compared to arteries.
Aside for the contribution of X0 to venous, but not arterial ET-1-mediated
contraction, X0 and its metabolite, UA, did not appear to othenrvise impact
arterial or venous contractile function, and did not modify the arterial
hypertensive endothelial dysfunction. X0 inhibition with allopurinol in vivo did not
lead to a decrease in BP in the established DOCA-salt hypertension and did not
prevent or attenuate the increases in BP in the development of DOCA-salt

hypertension.

Although the involvement of ROS in cardiovascular disease in general and in

hypertension in particular has widely been accepted, the question of increases in

161

ROS as a cause or as an effect of pathological states still remains. Venous
function in normal conditions, as well as the involvement of venomotor tone in
hypertension pathogenesis are understudied subjects of research. Further
studies are needed to elucidate the role of ROS in normal venous system biology

as well as that of ROS metabolism on venomotor tone and BP control.

Limitations

A number of facts limit the conclusions that may be drawn from this research. In
most of these experiments, we used the aorta-VC pair for comparison. As often
the case in vascular research, different results on ROS metabolism may have
been obtained in other vascular beds. Additionally, the use of blood vessels from
the splanchnic circulation may especially be more relevant to blood pressure
regulation. Second, we used a blood pressure model that, as most hypertension
models, mimics a form of secondary hypertension, rather than the more often
encountered essential one. Moreover, by using an induced hypertension model,
the potential pathogenetic role of ROS may be obscured by the effects of the
increased ROS that are a result, rather than one of the causes of the
pathogenetic process. Third, when assigning a role in ROS metabolism or in
vascular contraction to various enzymes, we relied on inhibition through
pharmacological tools that generally may be less than perfect in their specificity
and effectiveness. This limitation becomes particularly important in the case of

allopurinol, on which we relied extensively for both our in vitro and in vivo studies.

162

 

Concerns associated with use of allopurinol and its metabolite oxypurinol include
potential scavenging activities towards hydroxyl radical (21), as well as the
previously mentioned off-target inhibitory effects on other enzymes, such as
purine nucleoside phosphorylase and orotidine—5'-monophosphate
decarboxylase, both also involved in nucleotide catabolism. Aside from being
normotensive, the sham rats we used as surgery control for their DOCA
hypertensive counterparts may be different from normal (untreated) Sprague-
Dawley rats in other respects that we have not controlled for. Several other
limitations are inherent to the methods for our experiments, the most obvious of
which may be the ones related to ROS measurements. Due to a number of
factors, one of which being the extremely short half-life of ROS, their
measurements are so often inconsistent and fraught with such difficulty, that the
consensus in the free radical field of study is that at least two different types of
measurements are mandatory for formulating any conclusion on ROS levels.
Another limitation that may set apart our results from those of other reports is the
fact that in generating them we did not use pure cell cultures, but rather whole
tissues and tissue homogenates. Although this approach brings the matter closer
to the physiological setting, interpreting results thus obtained is more difficult, as
they may reflect contributions by all the varied cell types as well as their

interactions in a tissue.

163

Perspectives

Although we invalidated our hypothesis regarding the role of X0 in the
established or developing DOCA-salt hypertension, we have by no means
excluded the important role of ROS in venous function or the not fully understood
role of veins in hypertension pathogenesis. X0 may be still involved in the
modulation of vascular contractile function and thus indirectly in blood pressure
regulation, and during hypertension it may still be responsible for specific
damage of target organs. Other ROS metabolizing enzymes may also provide
interesting alternatives, one of them being DUOX1, as evidenced by data from
our first aim. By comparing veins and arteries we observed many molecular and
functional differences. Findings such as these mandate further studies on
uncovering the characteristics that enable each blood vessel type to perform
optimally in its own environment, and that explain their response during

pathological situations like hypertension.

164

REFERENCES

1. Adams LD, Geary RL, McManus B, and Schwartz SM. A comparison of
aorta and vena cava medial message expression by cDNA array analysis

identifies a set of 68 consistently differentially expressed genes, all in aortic
media. Circ Res 87: 623-631, 2000.

2. Ardanaz N, and Pagano PJ. Hydrogen peroxide as a paracrine vascular
mediator: regulation and signaling leading to dysfunction. Exp Biol Med
(Maywood) 231: 237-251, 2006.

3. Baldus S, Koster R, Chumley P, Heitzer T, Rudolph V, 0stad MA,
Warnholtz A, Staude HJ, Thuneke F, Koss K, Berger J, Meinertz T, Freeman
BA, and Munzel T. Oxypurinol improves coronary and peripheral endothelial
function in patients with coronary artery disease. Free Radic Biol Med 39: 1184-
1 190, 2005.

4. Banerjee D, Madhusoodanan UK, Sharanabasappa M, Ghosh S, and
Jacob J. Measurement of plasma hydroperoxide concentration by FOX-1 assay
in conjunction with triphenylphosphine. Clin Chim Acta 337: 147-152, 2003.

5. Barja G. Free radicals and aging. Trends Neurosci 27: 595-600, 2004.

6. Barry-Lane PA, Patterson C, van der Merwe M, Hu Z, Holland SM, Yeh
ET, and Runge MS. p47phox is required for atherosclerotic lesion progression in
ApoE(-/-) mice. J Clin Invest 108: 1513-1522, 2001.

7. Becker BF. Towards the physiological function of uric acid. Free Radic
Biol Med 14: 615-631, 1993.

8. Beckman JS, and Koppenol WH. Nitric oxide, superoxide, and
peroxynitrite: the good, the bad, and ugly. Am J Physiol 271: 01424-1437, 1996.

9. Berry CE, and Hare JM. Xanthine oxidoreductase and cardiovascular
disease: molecular mechanisms and pathophysiological implications. J Physiol
555: 589-606, 2004.

10. Beswick RA, Dorrance AM, Leite R, and Webb RC. NADH/NADPH
oxidase and enhanced superoxide production in the mineralocorticoid
hypertensive rat. Hypertension 38: 1 107-11 11, 2001 .

11. Borges F, Fernandes E, and Roleira F. Progress towards the discovery
of xanthine oxidase inhibitors. Curr Med Chem 9: 195-217, 2002.

12. Brand MD, Affourtit C, Esteves TC, Green K, Lambert AJ, Miwa S,
Pakay JL, and Parker N. Mitochondrial superoxide: production, biological

165

effects, and activation of uncoupling proteins. Free Radic Biol Med 37: 755-767,
2004.

13. Buetler TM, Krauskopf A, and Ruegg UT. Role of superoxide as a
signaling molecule. News Physio! Sci 19: 120-123, 2004.

14. Butler R, Morris AD, Belch JJ, Hill A, and Struthers AD. Allopurinol
normalizes endothelial dysfunction in type 2 diabetics with mild hypertension.
Hypertension 35: 746-751, 2000.

15. Cai H, and Harrison DG. Endothelial dysfunction in cardiovascular
diseases: the role of oxidant stress. Circ Res 87: 840-844, 2000.

16. Callera GE, Tostes RC, Yogi A, Montezano AC, and Touyz RM.
Endothelin-1-induced oxidative stress in DOCA-salt hypertension involves
NADPH-oxidase—independent mechanisms. Clin Sci (Land) 110: 243-253, 2006.

17. Chu Y, Iida S, Lund DD, Weiss RM, DiBona GF, Watanabe Y, Faraci
FM, and Heistad DD. Gene transfer of extracellular superoxide dismutase
reduces arterial pressure in spontaneously hypertensive rats: role of heparin-
binding domain. Circ Res 92: 461-468, 2003.

18. Clempus RE, and Griendling KK. Reactive oxygen species signaling in
vascular smooth muscle cells. Cardiovasc Res 71: 216-225, 2006.

19. Cooper N, Khosravan R, Erdmann C, Fiene J, and Lee JW.
Quantification of uric acid, xanthine and hypoxanthine in human serum by HPLC
for pharmacodynamic studies. J Chromatogr B Analyt Techno! Biomed Life Sci
837: 1-10, 2006.

20. Corry DB, Eslami P, Yamamoto K, Nyby MD, Makino H, and Tuck ML.
Uric acid stimulates vascular smooth muscle cell proliferation and oxidative
stress via the vascular renin-angiotensin system. J Hypertens 26: 269-275, 2008.

21. Das DK, Engelman RM, Clement R, Otani H, Prasad MR, and Rao PS.
Role of xanthine oxidase inhibitor as free radical scavenger: a novel mechanism

of action of allopurinol and oxypurinol in myocardial salvage. Biochem Biophys
Res Commun 148: 314-319, 1987.

22. Decoursey TE, and Ligeti E. Regulation and termination of NADPH
oxidase activity. Cell Mol Life Sci 62: 2173-2193, 2005.

23. dela Paz NG, and D'Amore PA. Arterial versus venous endothelial cells.
Cell Tissue Res 335: 5—16, 2009. '

24. Deng DX, Spin JM, Tsalenko A, Vailaya A, Ben-Dor A, Yakhini Z, Tsao
P, Bruhn L, and Quertermous T. Molecular signatures determining coronary

166

artery and saphenous vein smooth muscle cell phenotypes: distinct responses to
stimuli. Arterioscler Thromb Vasc Biol 26: 1058-1065, 2006.

25. Deng DX, Tsalenko A, Vailaya A, Ben-Dor A, Kundu R, Estay I,
Tabibiazar R, Kincaid R, Yakhini Z, Bruhn L, and Quertermous T. Differences
in vascular bed disease susceptibility reflect differences in gene expression
response to atherogenic stimuli. Circ Res 98: 200-208, 2006.

26. Duckies SP, Krause DN, Stirone C, and Procaccio V. Estrogen and
mitochondria: a new paradigm for vascular protection? Mol Interv 6: 26-35, 2006.

27. Duerrschmidt N, Wippich N, Goettsch W, Broemme HJ, and
Morawietz H. Endothelin-1 induces NAD(P)H oxidase in human endothelial cells.
Biochem Biophys Res Commun 269: 713-717, 2000.

28. Dupont GP, Huecksteadt TP, Marshall BC, Ryan US, Michael JR, and
Hoidal JR. Regulation of xanthine dehydrogenase and xanthine oxidase activity
and gene expression in cultured rat pulmonary endothelial cells. J Clin Invest 89:
197—202, 1992.

29. EI-Benna J, Dang PM, Gougerot-Pocidalo MA, and Elbim C.
Phagocyte NADPH oxidase: a multicomponent enzyme essential for host
defenses. Arch Immunol Ther Exp (Warsz) 53: 199-206, 2005.

30. Esteves TC, and Brand MD. The reactions catalysed by the
mitochondrial uncoupling proteins UCP2 and UCP3. Biochim Biophys Acta 1709:
35-44, 2005.

31. Faraci FM, and Didion SP. Vascular protection: superoxide dismutase
isoforms in the vessel wall. Arterioscler Thromb Vasc Biol 24: 1367-1373, 2004.

32. Feig DI, Kang DH, and Johnson RJ. Uric acid and cardiovascular risk. N
Engl J Med 359: 1811-1821, 2008.

33. Feig DI, Soletsky B, and Johnson RJ. Effect of allopurinol on blood
pressure of adolescents with newly diagnosed essential hypertension: a
randomized trial. Jama 300: 924-932, 2008.

34. Feissner RF, Skalska J, Gaum WE, and Sheu SS. Crosstalk signaling
between mitochondrial Ca2+ and ROS. Front Biosci 14: 1197-1218, 2009.

35. Fennell JP, Brosnan MJ, Frater AJ, Hamilton CA, Alexander MY,
Nicklin SA, Heistad DD, Baker AH, and Dominiczak AF. Adenovirus-mediated
overexpression of extracellular superoxide dismutase improves endothelial
dysfunction in a rat model of hypertension. Gene Ther 9: 110-117, 2002.

36. Fernandes E, Costa D, Toste SA, Lima JL, and Reis S. In vitro
scavenging activity for reactive oxygen and nitrogen species by nonsteroidal anti-

167

inflammatory indole, pyrrole, and oxazole derivative drugs. Free Radic Biol Med
37: 1895-1905, 2004.

37. Fields M, Lewis CG, and Lure MD. Allopurinol, an inhibitor of xanthine
oxidase, reduces uric acid levels and modifies the signs associated with copper
deficiency in rats fed fructose. Free Radic Biol Med 20: 595-600, 1996.

38. Fink GD. Arthur C. Corcoran Memorial Lecture. Sympathetic activity,
vascular capacitance, and long-term regulation of arterial pressure. Hypertension
53: 307-312, 2009.

39. Fink GD, Johnson RJ, and Galligan JJ. Mechanisms of increased
venous smooth muscle tone in desoxycorticosterone acetate-salt hypertension.
Hypertension 35: 464-469, 2000.

40. Finkel T, and Holbrook NJ. Oxidants, oxidative stress and the biology of
ageing. Nature 408: 239-247, 2000.

41. Forstermann U, and Munzel T. Endothelial nitric oxide synthase in
vascular disease: from marvel to menace. Circulation 113: 1708-1714, 2006.

42. Fridovich l. The Competitive Inhibition of Uricase by Oxonate and by
Related Derivatives of S-Triazines. J Biol Chem 240: 2491-2494, 1965.

43. Ganong WF. Review of medical physiology. New York: McGraw-Hill
Medical, 2005.

44. Gao YJ, and Lee RM. Hydrogen peroxide is an endothelium-dependent
contracting factor in rat renal artery. BrJ Phannacol 146: 1061-1068, 2005.

45. George J, Carr E, Davies J, Belch JJ, and Struthers A. High-dose
allopurinol improves endothelial function by profoundly reducing vascular
oxidative stress and not by lowering uric acid. Circulation 114: 2508-2516, 2006.

46. Gersch C, Palii SP, Kim KM, Angerhofer A, Johnson RJ, and
Henderson GN. Inactivation of nitric oxide by uric acid. Nucleosides Nucleotides
Nucleic Acids 27: 967-978, 2008.

47. Greenbaum D, Colangelo C, Williams K, and Gerstein M. Comparing
protein abundance and mRNA expression levels on a genomic scale. Genome
Bio/4: 117, 2003.

48. Griendling KK, Minieri CA, Ollerenshaw JD, and Alexander RW.
Angiotensin II stimulates NADH and NADPH oxidase activity in cultured vascular
smooth muscle cells. Circ Res 74: 1141-1148, 1994.

49. Guo Y, Xiao P, Lei S, Deng F, Xiao GG, Liu Y, Chen X, Li L, Wu 8,
Chen Y, Jiang H, Tan L, Xie J, Zhu X, Liang S, and Deng H. How is mRNA

168

expression predictive for protein expression? A correlation study on human
circulating monocytes. Acta Biochim Biophys Sin (Shanghai) 40: 426-436, 2008.

50. Guzik TJ, Olszanecki R, Sadowski J, Kapelak B, Rudzinski P, Jopek
A, Kawczynska A, Ryszawa N, Loster J, Jawien J, Czesnikiewicz-Guzik M,
Channon KM, and Korbut R. Superoxide dismutase activity and expression in
human venous and arterial bypass graft vessels. J Physiol Phannacol 56: 313-
323, 2005.

51. Guzik TJ, Sadowski J, Kapelak B, Jopek A, Rudzinski P, Pillai R,
Korbut R, and Channon KM. Systemic regulation of vascular NAD(P)H oxidase

activity and nox isoform expression in human arteries and veins. Arterioscler
Thromb Vasc Biol 24: 1614-1620, 2004.

52. Guzik TJ, West NE, Pillai R, Taggart DP, and Channon KM. Nitric oxide
modulates superoxide release and peroxynitrite formation in human blood
vessels. Hypertension 39: 1088-1094, 2002.

53. Harrison R. Physiological roles of xanthine oxidoreductase. Drug Metab
Rev 36: 363-375, 2004.

54. Haurani MJ, and Pagano PJ. Adventitial fibroblast reactive oxygen
species as autacrine and paracrine mediators of remodeling: bellwether for
vascular disease? Cardiovasc Res 75: 679-689, 2007.

55. Heikkila RE, Cabbat FS, and Cohen G. In vivo inhibition of superoxide
dismutase in mice by diethyldithiocarbamate. J Biol Chem 251: 2182-2185, 1976.

56. Heumuller 8, Wind 8, Barbosa-Sicard E, Schmidt HH, Busse R,
Schroder K, and Brandes RP. Apocynin is not an inhibitor of vascular NADPH
oxidases but an antioxidant. Hypertension 51: 211-217, 2008.

57. Hisaki R, Fujita H, Saito F, and Kushiro T. Tempol attenuates the
development of hypertensive renal injury in Dahl salt-sensitive rats. Am J
Hypertens 18: 707-713, 2005.

58. Ho YS, Magnenat JL, Gargano M, and Cao J. The nature of antioxidant
defense mechanisms: a lesson from transgenic studies. Environ Health Perspect
106 Suppl 5: 1219-1228, 1998.

59. Hohler B, Holzapfel B, and Kummer W. NADPH oxidase subunits and
superoxide production in porcine pulmonary artery endothelial cells. Histochem
Cell Biol 114: 29-37, 2000.

60. Horiuchi H, Ota M, Nishimura S, Kaneko H, Kasahara Y, Ohta T, and
Komoriya K. Allopurinol induces renal toxicity by impairing pyrimidine
metabolism in mice. Life Sci 66: 2051-2070, 2000.

169

61. Hottenstein OD, and Kreulen DL. Comparison of the frequency
dependence of venous and arterial responses to sympathetic nerve stimulation in
guinea-pigs. JPhysi01384: 153-167, 1987.

62. Houston M, Estevez A, Chumley P, Asian M, Marklund S, Parks DA,
and Freeman BA. Binding of xanthine oxidase to vascular endothelium. Kinetic
characterization and oxidative impairment of nitric oxide-dependent signaling. J
Biol Chem 274: 4985-4994, 1999.

63. lkebuchi Y, Masumoto N, Tasaka K, Koike K, Kasahara K, Miyake A,
and Tanizawa O. Superoxide anion increases intracellular pH, intracellular free
calcium, and arachidonate release in human amnion cells. J Biol Chem 266:
13233-13237, 1991.

64. Inkster ME, Cotter MA, and Cameron NE. Treatment with the xanthine
oxidase inhibitor, allopurinol, improves nerve and vascular function in diabetic
rats. Eur J Phannacol 561: 63-71, 2007.

65. Johnson F, and Giulivi C. Superoxide dismutases and their impact upon
human health. Mol Aspects Med 26: 340-352, 2005.

66. Johnson RJ, Kang DH, Feig D, Kivlighn S, Kanellis J, Watanabe S,
Tuttle KR, Rodriguez-lturbe B, Herrera-Acosta J, and Mazzali M. Is there a
pathogenetic role for uric acid in hypertension and cardiovascular and renal
disease? Hypertension 41: 1183-1190, 2003.

67. Jung 0, Marklund SL, Xia N, Busse R, and Brandes RP. Inactivation of
extracellular superoxide dismutase contributes to the development of high-
volume hypertension. Arterioscler Thromb Vasc Biol 27: 470-477, 2007.

68. Jung 0, Schreiber JG, Geiger H, Pedrazzini T, Busse R, and Brandes
RP. gp91phox-containing NADPH oxidase mediates endothelial dysfunction in
renovascular hypertension. Circulation 109: 1795-1801, 2004.

69. Kasal DA, Neves MF, Oigman W, and Mandarim-de-Lacerda CA.
Allopurinol attenuates L-NAME induced cardiomyopathy comparable to blockade
of angiotensin receptor. Histol Histopathol 23: 1241-1248, 2008.

70. Kemp BK, Smolich JJ, and Cocks TM. Evidence for specific regional
patterns of responses to different vasoconstrictors and vasodilators in sheep
isolated pulmonary arteries and veins. BrJ Pharmacol121: 441-450, 1997.

71. Khosla UM, Zharikov S, Finch JL, Nakagawa T, Roncal C, Mu W,
Krotova K, Block ER, Prabhakar S, and Johnson RJ. Hyperuricemia induces
endothelial dysfunction. Kidney Int 67: 1739-1742, 2005.

72. Kim SY, De Vera MA, and Choi HK. Gout and mortality. Clin Exp
Rheumatol 26: S1 15-119, 2008.

170

73. Kimura S, Zhang GX, Nishiyama A, Shokoji T, Yao L, Fan YY,
Rahman M, Suzuki T, Maeta H, and Abe Y. Role of NAD(P)H oxidase- and
mitochondria-derived reactive oxygen species in cardioprotection of ischemic
reperfusion injury by angiotensin II. Hypertension 45: 860-866, 2005.

74. Kisker C, Schindelin H, and Rees DC. Molybdenum-cofactor-containing
enzymes: structure and mechanism. Annu Rev Biochem 66: 233-267, 1997.

75. Kleinbongard P, Schulz R, Rassaf T, Lauer T, Dejam A, Jax T,
Kumara I, Gharini P, Kabanova S, Ozuyaman B, Schnurch HG, Godecke A,
Weber AA, Robenek M, Robenek H, Bloch W, Rosen P, and Kelm M. Red
blood cells express a functional endothelial nitric oxide synthase. Blood 107:
2943-2951, 2006.

76. Kris-Etherton PM, Lichtenstein AH, Howard BV, Steinberg D, and
Witztum JL. Antioxidant vitamin supplements and cardiovascular disease.
Circulation 1 10: 637-641 , 2004.

77. Krishnan E, Kwoh CK, Schumacher HR, and Kuller L. Hyperuricemia
and incidence of hypertension among men without metabolic syndrome.
Hypertension 49: 298-303, 2007.

78. Laakso J, Mervaala E, Himberg JJ, Teravainen TL, Karppanen H,
Vapaatalo H, and Lapatto R. Increased kidney xanthine oxidoreductase activity
in salt-induced experimental hypertension. Hypertension 32: 902-906, 1998.

79. Laakso JT, Teravainen TL, Martelin E, Vaskonen T, and Lapatto R.
Renal xanthine oxidoreductase activity during development of hypertension in
spontaneously hypertensive rats. J Hypertens 22: 1333-1340, 2004.

80. Lacza Z, Pankotai E, and Busija DW. Mitochondrial nitric oxide
synthase: current concepts and controversies. Front Biosci 14: 4436-4443, 2009.

81. Landmesser U, Cai H, Dikalov S, McCann L, Hwang J, Jo H, Holland
SM, and Harrison DG. Role of p47(phox) in vascular oxidative stress and
hypertension caused by angiotensin II. Hypertension 40: 511-515, 2002.

82. Landmesser U, Dikalov S, Price SR, McCann L, Fukai T, Holland SM,
Mitch WE, and Harrison DG. Oxidation of tetrahydrobiopterin leads to
uncoupling of endothelial cell nitric oxide synthase in hypertension. J Clin Invest
111: 1201-1209, 2003.

83. Landmesser U, Spiekermann S, Preuss C, Sorrentino S, Fischer D,
Manes C, Mueller M, and Drexler H. Angiotensin II induces endothelial xanthine
oxidase activation: role for endothelial dysfunction in patients with coronary
disease. Arterioscler Thromb Vasc Biol 27: 943-948, 2007.

171

84. Lassegue B, and Clempus RE. Vascular NAD(P)H oxidases: specific
features, expression, and regulation. Am J Physiol Regul Integr Comp Physiol
285: R277-297, 2003.

85. Lassegue B, Sorescu D, Szocs K, Yin Q, Akers M, Zhang Y, Grant SL,
Lambeth JD, and Griendling KK. Novel gp91(phox) homologues in vascular
smooth muscle cells : nox1 mediates angiotensin lI-induced superoxide
formation and redox-sensitive signaling pathways. Circ Res 88: 888-894, 2001.

86. Li L, Fink GD, Watts SW, Northcott CA, Galligan JJ, Pagano PJ, and
Chen AF. Endothelin-1 increases vascular superoxide via endothelin(A)-NADPH
oxidase pathway in low-renin hypertension. Circulation 107: 1053-1058, 2003.

87. Li L, Watts SW, Banes AK, Galligan JJ, Fink GD, and Chen AF.
NADPH oxidase-derived superoxide augments endothelin-I-induced

venoconstriction in mineralocorticoid hypertension. Hypertension 42: 316-321,
2003.

88. Linder N, Rapola J, and Raivio KO. Cellular expression of xanthine
oxidoreductase protein in normal human tissues. Lab Invest 79: 967-974, 1999.

89. Lloyd-Jones D, Adams R, Carnethon M, De Simone G, Ferguson TB,
Flegal K, Ford E, Furie K, Go A, Greenlund K, Haase N, Hailpern 8, Ho M,
Howard V, Kissela B, Kittner S, Lackland D, Lisabeth L, Marelli A,
McDermott M, Meigs J, Mozaffarian D, Nichol G, O'Donnell C, Roger V,
Rosamond W, Sacco R, Sorlie P, Stafford R, Steinberger J, Thorn T,
WasserthieI-Smoller S, Wong N, Wylie-Rosett J, and Hong Y. Heart disease
and stroke statistics-2009 update: a report from the American Heart Association
Statistics Committee and Stroke Statistics Subcommittee. Circulation 119: e21-
181, 2009.

90. London GM, Safar ME, Weiss YA, Laurent S, and London AM. Renal
and systemic hemodynamics in borderline hypertension. Am J Hypertens 1:
1278-1308, 1988.

91. London GM, Safar ME, Weiss YA, and Simon CA. Total effective
compliance of the vascular bed in essential hypertension. Am Heart J 95: 325-
330, 1978.

92. Loomis ED, Sullivan JC, Osmond DA, Pollock DM, and Pollock JS.
Endothelin mediates superoxide production and vasoconstriction through
activation of NADPH oxidase and uncoupled nitric-oxide synthase in the rat
aorta. J Phannacol Exp Ther 315: 1058-1064, 2005.

93. Mackay J, Mensah GA, World Health Organization., and Centers for
Disease Control and Prevention (U.S.). The atlas of heart disease and stroke.
Geneva: World Health Organization, 2004, p. 112 p.

172

94. Macmillan-Crow LA, and Cruthirds DL. Invited review: manganese
superoxide dismutase in disease. Free Radic Res 34: 325-336, 2001.

95. Madamanchi NR, Vendrov A, and Runge MS. Oxidative stress and
vascular disease. Arterioscler Thromb Vasc Biol 25: 29-38, 2005.

96. Martin DS, Rodrigo MC, and Appelt CW. Venous tone in the
developmental stages of spontaneous hypertension. Hypertension 31: 139-144,
1998.

97. Mazzali M, Hughes J, Kim YG, Jefferson JA, Kang DH, Gordon KL,
Lan HY, Kivlighn S, and Johnson RJ. Elevated uric acid increases blood

pressure in the rat by a novel crystal-independent mechanism. Hypertension 38:
1101-1106, 2001.

98. McNaIIy JS, Davis ME, Giddens DP, Saha A, Hwang J, Dikalov S, Jo
H, and Harrison DG. Role of xanthine oxidoreductase and NAD(P)H oxidase in
endothelial superoxide production in response to oscillatory shear stress. Am J
Physiol Heart Circ Physiol 285: H2290-2297, 2003.

99. McNally JS, Saxena A, Cai H, Dikalov S, and Harrison DG. Regulation
of xanthine oxidoreductase protein expression by hydrogen peroxide and
calcium. Arterioscler Thromb Vasc Biol 25: 1623-1628, 2005.

100. Mollnau H, Wendt M, Szocs K, Lassegue B, Schulz E, Oelze M, Li H,
Bodenschatz M, August M, Kleschyov AL, Tsilimingas N, Walter U,
Forstermann U, Meinertz T, Griendling K, and Munzel T. Effects of
angiotensin II infusion on the expression and function of NAD(P)H oxidase and
components of nitric oxide/cGMP signaling. Circ Res 90: E58-65, 2002.

101. Motwani JG, and Topol EJ. Aortocoronary saphenous vein graft disease:
pathogenesis, predisposition, and prevention. Circulation 97: 916-931, 1998.

102. Munzel T, Afanas'ev IB, Kleschyov AL, and Harrison DG. Detection of
superoxide in vascular tissue. Arterioscler Thromb Vasc Biol 22: 1761-1768,
2002.

103. Nakazono K, Watanabe N, Matsuno K, Sasaki J, Sato T, and lnoue M.
Does superoxide underlie the pathogenesis of hypertension? Proc Natl Acad Sci
U SA 88: 10045-10048, 1991.

104. Newaz MA, Yousefipour Z, and Oyekan A. Oxidative stress-associated
vascular aging is xanthine oxidase-dependent but not NAD(P)H oxidase-
dependent. J Cardiovasc Phannacol 48: 88-94, 2006.

105. Nishino T, Okamoto K, Eger BT, Pai EF, and Nishino T. Mammalian

xanthine oxidoreductase - mechanism of transition from xanthine dehydrogenase
to xanthine oxidase. Febs J 275: 3278-3289, 2008.

173

106. Nyhof RA, Laine GA, Meininger GA, and Granger HJ. Splanchnic
circulation in hypertension. Fed Proc 42: 1690-1693, 1983.

107. Ong SL, Vickers JJ, Zhang Y, McKenzie KU, Walsh CE, and
Whitworth JA. Role of xanthine oxidase in dexamethasone-induced
hypertension in rats. Clin Exp Phannacol Physiol 34: 517-519, 2007.

108. Onuma S, and Nakanishi K. Superoxide dismustase mimetic tempol
decreases blood pressure by increasing renal medullary blood flow in
hyperinsulinemic—hypertensive rats. Metabolism 53: 1305-1308, 2004.

109. Ookawara T, Eguchi H, Nishimura M, Kizaki T, Eiji T, Saitoh D, Ohno
H, and Suzuki K. Effects of oxidative stress on the nuclear translocation of
extracellular superoxide dismutase. Biochem Biophys Res Commun 303: 914-
919, 2003.

110. Pacher P, Nivorozhkin A, and Szabo C. Therapeutic effects of xanthine
oxidase inhibitors: renaissance half a century after the discovery of allopurinol.
Phannacol Rev 58: 87-114, 2006.

111. Pan YJ, and Young DB. Experimental aldosterone hypertension in the
dog. Hypertension 4: 279-287, 1982.

112. Payeli SK, Latini R, Gebhard C, Patrignani A, Wagner U, Luscher TF,
and Tanner FC. Prothrombotic gene expression profile in vascular smooth
muscle cells of human saphenous vein, but not internal mammary artery.
Arterioscler Thromb Vasc Biol 28: 705-710, 2008.

113. Poli G, Biasi F, and Leonarduzzi G. 4-Hydroxynonenal-protein adducts:
A reliable biomarker of lipid oxidation in liver diseases. Mol Aspects Med 29: 67-
71, 2008.

114. Potter 82, and Valentine JS. The perplexing role of copper-zinc
superoxide dismutase in amyotrophic lateral sclerosis (Lou Gehrig's disease). J
Biol Inorg Chem 8: 373-380, 2003.

115. Price KL, Sautin YY, Long DA, Zhang L, Miyazaki H, Mu W, Endou H,
and Johnson RJ. Human vascular smooth muscle cells express a urate
transporter. J Am Soc Nephrol 17: 1791-1795, 2006.

116. Putterman GJ, Shaikh B, Hallmark MR, Sawyer CG, Hixson CV, and
Perini F. Simultaneous analysis of substrates, products, and inhibitors of
xanthine oxidase by high-pressure liquid chromatography and gas
chromatography. Anal Biochem 98: 18-26, 1979.

117. Rao GN, Corson MA, and Berk BC. Uric acid stimulates vascular smooth
muscle cell proliferation by increasing platelet-derived growth factor A-chain
expression. J Biol Chem 266: 8604-8608, 1991.

174

118. Redon J, Oliva MR, Tomes C, Giner V, Chaves J, Iradi A, and Saez
GT. Antioxidant activities and oxidative stress byproducts in human hypertension.
Hypertension 41: 1096-1101, 2003.

119. Rey FE, Cifuentes ME, Kiarash A, Quinn MT, and Pagano PJ. Novel
competitive inhibitor of NAD(P)H oxidase assembly attenuates vascular 0(2)(-)
and systolic blood pressure in mice. Circ Res 89: 408-414, 2001.

120. Ricksten SE, Yao T, and Thoren P. Peripheral and central vascular
compliances in conscious normotensive and spontaneously hypertensive rats.
Acta Physio/Scand112: 169-177, 1981.

121. Rothe CF. Reflex control of veins and vascular capacitance. Physiol Rev
63: 1281-1342, 1983.

122. Rozen S, and Skaletsky H. Primer3 on the W for general users and
for biologist programmers. Methods Mol Biol 132: 365-386, 2000.

123. Rueckschloss U, Galle J, Hall: J, Zerkowski HR, and Morawietz H.
Induction of NAD(P)H oxidase by oxidized low-density lipoprotein in human
endothelial cells: antioxidative potential of hydroxymethylglutaryl coenzyme A
reductase inhibitor therapy. Circulation 104: 1767-1772, 2001.

124. Rueckschloss U, Quinn MT, Holtz J, and Morawietz H. Dose-
dependent regulation of NAD(P)H oxidase expression by angiotensin II in human
endothelial cells: protective effect of angiotensin II type 1 receptor blockade in
patients with coronary artery disease. Arterioscler Thromb Vasc Biol 22: 1845-
1851, 2002.

125. Saetrum Opgaard O, and Edvinsson L. Effect of parasympathetic and
sensory transmitters on human epicardial coronary arteries and veins. Phannacol
Toxicol 78: 273-279, 1996.

126. Schmieder RE, Schobel HP, and Messerli FH. Central blood volume: a
determinant of early cardiac adaptation in arterial hypertension? J Am Coll
Cardiol 26: 1692-1698, 1995.

127. Schulz E, Jansen T, Wenzel P, Daiber A, and Munzel T. Nitric oxide,
tetrahydrobiopterin, oxidative stress, and endothelial dysfunction in hypertension.
Antioxid Redox Signal 10: 1115-1126, 2008.

128. Schulz R, Rassaf T, Massion PB, Kelm M, and Balligand JL. Recent
advances in the understanding of the role of nitric oxide in cardiovascular
homeostasis. Phannacol Ther108: 225-256, 2005.

129. Selemidis S, Sobey CG, Wingler K, Schmidt HH, and Drummond GR.

NADPH oxidases in the vasculature: molecular features, roles in disease and
pharmacological inhibition. Pharmacol Ther 120: 254-291, 2008.

175

130. Shapira 0M, Xu A, Aldea GS, Vita JA, Shemin RJ, and Keaney JF, Jr.
Enhanced nitric oxide-mediated vascular relaxation in radial artery compared

with internal mammary artery or saphenous vein. Circulation 100: "322-327,
1999.

131. Shi W, Eidelman DH, and Michel RP. Differential relaxant responses of
pulmonary arteries and veins in lung explants of guinea pigs. J Appl Physiol 83:
1476-1481, 1997.

132. Shi Y, Patel 8, Davenpeck KL, Niculescu R, Rodriguez E, Magno MG,
Orrnont ML, Mannion JD, and Zalewski A. Oxidative stress and lipid retention
in vascular grafts: comparison between venous and arterial conduits. Circulation
103: 2408-2413, 2001.

133. Skinner KA, White CR, Patel R, Tan S, Barnes 8, Kirk M, Darley-
Usmar V, and Parks DA. Nitrosation of uric acid by peroxynitrite. Formation of a
vasoactive nitric oxide donor. J Biol Chem 273: 24491-24497, 1998.

134. Smith RA, Kelso GF, Blaikie FH, Porteous CM, Ledgerwood EC,
Hughes G, James AM, Ross MF, Asin-Cayuela J, Cocheme HM, Filipovska
A, and Murphy MP. Using mitochondria-targeted molecules to study
mitochondrial radical production and its consequences. Biochem Soc Trans 31:
1295-1299, 2003.

135. So A. Developments in the scientific and clinical understanding of gout.
Arthritis Res Ther 10: 221, 2008.

136. Spiekermann S, Landmesser U, Dikalov S, Bredt M, Gamez G, Tatge
H, Reepschlager N, Hornig B, Drexler H, and Harrison DG. Electron spin
resonance characterization of vascular xanthine and NAD(P)H oxidase activity in
patients with coronary artery disease: relation to endothelium-dependent
vasodilation. Circulation 107: 1383-1389, 2003.

137. Srivastava P, Hegde LG, Patnaik GK, and Dikshit M. Role of
endothelial-derived reactive oxygen species and nitric oxide in norepinephrine-
induced rat aortic ring contractions. Phannacol Res 38: 265-274, 1998.

138. St-Pierre J, Buckingham JA, Roebuck SJ, and Brand MD. Topology of
superoxide production from different sites in the mitochondrial electron transport
chain. J Biol Chem 277: 44784-44790, 2002.

139. Stralin P, Karlsson K, Johansson BO, and Marklund SL. The
interstitium of the human arterial wall contains very large amounts of extracellular
superoxide dismutase. Arterioscler Thromb Vasc Biol 15: 2032-2036, 1995.

140. Sumimoto H, Miyano K, and Takeya R. Molecular composition and
regulation of the Nox family NAD(P)H oxidases. Biochem Biophys Res Commun
338: 677-686, 2005.

176

141. Suzuki H, DeLano FA, Parks DA, Jamshidi N, Granger DN, lshii H,
Suematsu M, Zweifach BW, and Schmid-Schonbein GW. Xanthine oxidase

activity associated with arterial blood pressure in spontaneously hypertensive
rats. Proc Natl Acad Sci U S A 95: 4754-4759, 1998.

142. Suzuki Y, Sato Y, Sudo J, and Tanabe T. Renal lipid peroxidation
induced by allopurinol-administration in rats. Dev Toxicol Environ Sci 14: 65-72,
1986. ‘

143. Suzuki Y, Sudo J, and Tanabe T. Allopurinol toxicity: its toxic organ-
specificity between the liver and the kidney in the rat. J Toxicol Sci 9: 343-351,
1984.

144. Swales JD. Textbook of hypertension. Oxford ; Boston: Blackwell
Scientific Publications, 1994, p. xvi, 1328 p., [1326] p. of plates.

145. Szasz T, Eddy S, Paulauskis J, Burnett R, Ellekilde M, lovanna JL,
and Watts SW. Differential Expression of Pancreatitis-Associated Protein and

Thrombospondins in Arterial versus Venous Tissues. J Vasc Res 46: 551-560,
2009.

146. Szasz T, Thakali K, Fink GD, and Watts SW. A comparison of arteries
and veins in oxidative stress: producers, destroyers, function, and disease. Exp
Biol Med (Maywood) 232: 27-37, 2007.

147. Szasz T, Thompson JM, and Watts SW. A comparison of reactive
oxygen species metabolism in the rat aorta and vena cava: focus on xanthine
oxidase. Am J Physiol Heart Circ Physiol 295: H1341-H1350, 2008.

148. Takano Y, Hase-Aoki K, Horiuchi H, Zhao L, Kasahara Y, Kondo S,
and Becker MA. Selectivity of febuxostat, a novel non-purine inhibitor of
xanthine oxidase/xanthine dehydrogenase. Life Sci 76: 1835-1847, 2005.

149. Tanito M, Nakamura H, Kwon YW, Teratani A, Masutani H, Shioji K,
Kishimoto C, Ohira A, Horie R, and Yodoi J. Enhanced oxidative stress and

impaired thioredoxin expression in spontaneously hypertensive rats. Antioxid
Redox Signal 6: 89-97, 2004.

150. Thakali K, Davenport L, Fink GD, and Watts SW. Pleiotropic effects of
hydrogen peroxide in arteries and veins from normotensive and hypertensive
rats. Hypertension 47: 482-487, 2006.

151. Thakali K, Demel SL, Fink GO, and Watts SW. Endothelin-1-induced
contraction in veins is independent of hydrogen peroxide. Am J Physiol Heart
Circ Physiol 289: H1115-1122, 2005.

152. Thakali K, Fink GD, and Watts SW. Arteries and veins desensitize
differently to endothelin. J Cardiovasc Phannacol 43: 387-393, 2004.

177

153. Thakali K, Galligan JJ, Fink GD, Gariepy CE, and Watts SW.
Pharmacological endothelin receptor interaction does not occur in veins from
ET(B) receptor deficient rats. Vascul Pharmacol 2008.

154. Thomas F, Blacher J, Benetos A, Safar ME, and Pannier B.
Cardiovascular risk as defined in the 2003 European blood pressure
classification: the assessment of an additional predictive value of pulse pressure
on mortality. J Hypertens 26: 1072-1077, 2008.

155. Titorenko VI, and Rachubinski RA. The peroxisome: orchestrating
important developmental decisions from inside the cell. J Cell Biol 164: 641-645,
2004.

156. Torrecillas G, Boyano-Adanez MC, Medina J, Parra T, Griera M,
Lopez-Ongil S, Arilla E, Rodriguez-Puyol M, and Rodriguez-Puyol D. The
role of hydrogen peroxide in the contractile response to angiotensin II. Mol
Phannacol 59: 104-112, 2001 .

157. Touyz RM, and Schiffrin EL. Reactive oxygen species in vascular
biology: implications in hypertension. Histochem Cell Biol 122: 339-352, 2004.

158. Trachtrnan H, Valderrama E, and Futterweit S. Nephrotoxicity of
allopurinol is enhanced in experimental hypertension. Hypertension 17: 194-202,
1991.

159. Turnheim K, Krivanek P, and Oberbauer R. Pharrnacokinetics and
pharmacodynamics of allopurinol in elderly and young subjects. Br J Clin
Phannacol 48: 501 -509, 1999.

160. Uddin M, Yang H, Shi M, Polley-Mandal M, and Guo Z. Elevation of
oxidative stress in the aorta of genetically hypertensive mice. Mech Ageing Dev
124: 811-817, 2003.

161. Ulker S, McMaster D, McKeown PP, and Bayraktutan U. Impaired
activities of antioxidant enzymes elicit endothelial dysfunction in spontaneous
hypertensive rats despite enhanced vascular nitric oxide generation. Cardiovasc
Res 59: 488-500, 2003.

162. Varela-Echavarria A, Montes de Oca-Luna R, and Barrera-Saldana
HA. Uricase protein sequences: conserved during vertebrate evolution but
absent in humans. Faseb J 2: 3092-3096, 1988.

163. Viel EC, Benkirane K, Javeshghani D, Touyz RM, and Schiffrin EL.
Xanthine oxidase and mitochondria contribute to vascular superoxide anion
generation in DOCA-salt hypertensive rats. Am J Physiol Heart Circ Physiol 295:
H281-288, 2008.

178

164. Vorbach C, Scriven A, and Capecchi MR. The housekeeping gene
xanthine oxidoreductase is necessary for milk fat droplet enveloping and
secretion: gene sharing in the lactating mammary gland. Genes Dev 16: 3223-
3235,2002.

165. Waring WS, McKnight JA, Webb DJ, and Maxwell SR. Lowering serum
urate does not improve endothelial function in patients with type 2 diabetes.
Diabetologia 50: 2572-2579, 2007.

166. Waring WS, McKnight JA, Webb DJ, and Maxwell SR. Uric acid
restores endothelial function in patients with type 1 diabetes and regular
smokers. Diabetes 55: 3127-3132, 2006.

167. Watts SW, Fink GD, Northcott CA, and Galligan JJ. Endothelin-1-
induced venous contraction is maintained in DOCA-salt hypertension; studies
with receptor agonists. Br J Phannacol 137: 69-79, 2002.

168. Watts SW, Rondelli C, Thakali K, Li X, Uhal B, Pervaiz MH, Watson
RE, and Fink GD. Morphological and biochemical characterization of remodeling
in aorta and vena cava of DOCA-salt hypertensive rats. Am J Physiol Heart Circ
Physiol 292: H2438-2448, 2007.

169. Watts SW, Thakali K, Smark C, Rondelli C, and Fink GD. Big ET-1
processing into vasoactive peptides in arteries and veins. Vascul Phannacol 47:
302-312, 2007.

170. Williams HC, and Griendling KK. NADPH oxidase inhibitors: new
antihypertensive agents? J Cardiovasc Phannacol 50: 9-16, 2007.

171. Wortmann RL. Recent advances in the management of gout and
hyperuricemia. Curr Opin Rheumatol 17: 319-324, 2005.

172. Wu M, Pritchard KA, Jr., Kaminski PM, Fayngersh RP, Hintze TH, and
Wolin MS. Involvement of nitric oxide and nitrosothiols in relaxation of pulmonary
arteries to peroxynitrite. Am J Physiol 266: H2108-2113, 1994.

173. Xu H, Fink GD, and Galligan JJ. Tempol lowers blood pressure and
sympathetic nerve activity but not vascular 02- in DOCA-salt rats. Hypertension
43: 329-334, 2004.

174. Yamamoto J, Trippodo NC, MacPhee AA, and Frohlich ED. Decreased
total venous capacity in Goldblatt hypertensive rats. Am J Physiol 240: H487-
492, 1981.

175. Yamamoto Y, Ogino K, Igawa G, Matsuura T, Kaetsu Y, Sugihara S,

Matsubara K, Miake J, Hamada T, Yoshida A, Igawa 0, Yamamoto T,
Shigemasa C, and Hisatome l. Allopurinol reduces neointimal hyperplasia in the

179

carotid artery ligation model in spontaneously hypertensive rats. Hypertens Res
29: 915-921, 2006.

176. Yang H, Shi M, VanRemmen H, Chen X, Vijg J, Richardson A, and
Guo Z. Reduction of pressor response to vasoconstrictor agents by
overexpression of catalase in mice. Am J Hypertens 16: 1-5, 2003.

177. Yang Z, Oemar BS, Carrel T, Kipfer B, Julmy F, and Luscher TF.
Different proliferative properties of smooth muscle cells of human arterial and
venous bypass vessels: role of PDGF receptors, mitogen-activated protein
kinase, and cyclin-dependent kinase inhibitors. Circulation 97: 181-187, 1998.

178. Yeldandi AV, Yeldandi V, Kumar S, Murthy CV, Wang XD, Alvares K,
Rao MS, and Reddy JK. Molecular evolution of the urate oxidase-encoding
gene in hominoid primates: nonsense mutations. Gene 109: 281-284, 1991.

179. You LR, Lin FJ, Lee CT, DeMayo FJ, Tsai MJ, and Tsai SY.
Suppression of Notch signalling by the COUP-TFII transcription factor regulates
vein identity. Nature 435: 98-104, 2005.

180. Zalba G, Beaumont FJ, San Jose G, Fortuno A, Fortuno MA, Etayo
JC, and Diez J. Vascular NADH/NADPH oxidase is involved in enhanced
superoxide production in spontaneously hypertensive rats. Hypertension 35:
1055-1061, 2000.

181. Zhang DX, and Gutterman DD. Mitochondrial reactive oxygen species-
mediated signaling in endothelial cells. Am J Physiol Heart Circ Physiol 292:
H2023-2031, 2007.

182. Zhou XF, Cui J, DeStefano AL, Chazaro I, Farrer LA, Manolis AJ,

Gavras H, and Baldwin CT. Polymorphisms in the promoter region of catalase
gene and essential hypertension. Dis Markers 21: 3-7, 2005.

180

 

  

A

9

     

MIIIIIIIIIINIIII IllIIIIIIIIIIIIIIIIIIIIIIEs
3 12 3 03063 0036