Implications on Nitric Oxide (No) Signalling

REDOX REGULATION OF ASYMMETRIC DIMETHYLARGININE (ADMA) METABOLISM: IMPLICATIONS ON NITRIC OXIDE (NO) SIGNALLING

SCOTT PRESTON FORBES

A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA

2010

To my mother and father

ACKNOWLEDGMENTS

First and foremost, I would like to thank my advisor Dr. AJ Cardounel. AJ has given me superb guidance to be able to achieve completion of my graduate education. In addition, AJ has been a source of inspiration through his hard work and dedication to science. AJ has a unique attitude towards science that I will never forget.

Additionally I would like to thank my dissertation committee: Dr. Glenn Walter, Dr. Maria

Grant, Dr. Paul Oh and Dr. Scott Powers. Thank you for your interest in my research project and for your valuable suggestions about my research.

Finally, I would like to thank everyone I have had the pleasure to work with in the

Cardounel lab at the University of Florida: Arthur Pope, Patrick Kearns and Kanchana

Karuppiah. I would also like to thank Dr. Jorge Guzman and Dr. Lawrence Druhan from The

Ohio State University.

TABLE OF CONTENTS

page

ACKNOWLEDGMENTS...... 4

LIST OF TABLES...... 7

LIST OF FIGURES ...... 8

ABSTRACT ...... 10

CHAPTER

1 INTRODUCTION...... 12

eNOS ...... 13 eNOS Regulation by Protein-Protein Interactions ...... 16 eNOS Regulation by Phosphorylation ...... 17 eNOS Regulation by Endogenous Methylarginines ...... 21 PRMT ...... 22 DDAH ...... 24 ADMA in Disease ...... 29

2 REDOX REGULATION OF ADMA METABOLISM: DDAH-1 ...... 34

Introduction ...... 34 Materials and Methods ...... 36 DDAH Expression and Purification ...... 36 DDAH Colorimetric Activity Assay ...... 37 DDAH Radioisotope Activity Assay...... 37 Proteomic Analysis of hDDAH-1: In Gel Digestion Manual ...... 38 Proteomic Analysis of hDDAH-1: Mass Spectrometry LTQ ...... 38 Results ...... 39 hDDAH-1 Protein Purification ...... 39 Enzyme Kinetics of hDDAH-1 ...... 40 pH Dependence of hDDAH-1 ...... 41 Effect of Oxidants on hDDAH-1 Activity ...... 41 Effect of 4-HNE on hDDAH-1 Activity ...... 42 Proteomic Analysis of hDDAH-1 ...... 43 Discussion ...... 44

3 REDOX REGULATION OF ADMA METABOLISM: DDAH-2 ...... 57

Introduction ...... 57 Materials and Methods ...... 59 DDAH Expression and Purification ...... 59 Immunodetection of hDDAH-2 ...... 59

Isolation and Denaturation of Inclusion Bodies ...... 60 Protein Refolding ...... 61 Cell Culture ...... 62 DDAH-1 and -2 Gene Silencing ...... 62 Assessment of mRNA Levels following DDAH Gene Silencing ...... 63 EPR Spectroscopy and Spin Trapping ...... 63 HPLC Method ...... 64 eNOS expression...... 64 Results ...... 64 hDDAH-2 Soluble Fraction Purification...... 64 hDDAH-2 Purification from Inclusion Bodies ...... 65 In vivo role of DDAH-2 on NO production ...... 66 Discussion ...... 67

4 REDOX REGULATION OF ADMA SYNTHSIS ...... 85

Introduction ...... 85 Materials and Methods ...... 86 Cell Culture ...... 86 EPR Spectroscopy and Spin Trapping ...... 87 PRMT Radioisotope Activity Assay ...... 87 Immunodetection of Nitro-Tyrosine ...... 88 Biotin Switch...... 89 Results ...... 90 Effect of PRMT Inhibitioin on Endothelial NO Production ...... 90 Effect of H2O2 on PRMT-1 Activity ...... 90 Effect of ONOO- on PRMT-1 Activity ...... 91 Effect of NO on PRMT-1 Activity ...... 92 Discussion ...... 93

5 DISCUSSION...... 102

WORKS CITED ...... 109

BIOGRAPHICAL SKETCH ...... 128

LIST OF TABLES

Table page

3-1 iFOLD Protein Refolding System 1, plate layout ...... 83

3-2 Solubility of refolded hDDAH-2 inclusion bodies ...... 84

LIST OF FIGURES

Figure page

2-1 hDDAH-1 purification...... 49

2-2 hDDAH-1 enzyme kinetics for ADMA...... 50

2-3 hDDAH-1 enzyme kinetics for L-NMMA...... 51

2-4 Effects of pH on hDDAH-1 enzyme activity...... 52

2-5 Effect of oxidants on hDDAH-1 activity...... 53

2-6 Effect of lipid hydroperoxides on hDDAH-1 activity...... 54

2-7 Modification at H15 on hDDAH-1 after exposure to 4-HNE...... 55

2-8 Modification at H173 on hDDAH-1 after exposure to 4-HNE...... 56

3-1 hDDAH-2 expression...... 71

3-2 hDDAH-2 purification from soluble fraction...... 72

3-3 DDAH activity of soluble fraction...... 73

3-4 Protein profile of isolated inclusion bodies...... 74

3-5 hDDAH-2 purification from inclusion bodies...... 75

3-6 DDAH activity of refolded hDDAH-2 from isolated inclusion bodies...... 76

3-7 Effects of DDAH gene silencing on DDAH mRNA expression...... 77

3-8 Effects of DDAH-1 gene silencing on endothelial cell NO production ...... 78

3-9 Effects of DDAH-2 gene silencing on endothelial cell NO production ...... 79

3-10 Effects of dual DDAH gene silencing on endothelial cell NO production...... 80

3-11 Effects of DDAH gene silencing on endothelial cell eNOS phosphorylation...... 81

3-12 Effects of DDAH gene silencing on endothelial cell intracellular ADMA concentrations...... 82

4-1 Effect of PRMT inhibition on endothelial NO production...... 96

4-2 Effect of H2O2 on PRMT-1 activity...... 97

4-3 Effect of ONOO- on PRMT-1 activity...... 98

4-4 Nitro-tyrosine formation on PRMT-1 after exposure to ONOO-...... 99

4-5 Effect of NO on PRMT-1 activity...... 100

4-6 S-nitrosylation of PRMT-1 after exposure to NO...... 101

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy

REDOX REGULATION OF ASYMMETRIC DIMETHYLARGININE (ADMA) METABOLISM: IMPLICATIONS ON NITRIC OXIDE (NO) SIGNALLING

Scott Preston Forbes

May 2010

Chair: Arturo Juan Cardounel Major: Medical Sciences— Physiology and Pharmacology

Nitric oxide (NO) is a critical regulator of a healthy vascular homeostasis. The endogenous nitric oxide synthase (NOS) inhibitor Asymmetric Dimethylarginine (ADMA) has been identified as an independent risk factor for cardiovascular disease. However, the mechanisms responsible for regulation of ADMA levels and their role in disease progression are unclear. A family of proteins known as protein arginine methyltransferases (PRMTs) are responsible for synthesis of ADMA by methylating arginine residues on proteins. Following proteolysis of these methylated proteins free ADMA in released into the cytosol. ADMA is metabolized by dimethylarginine dimethylaminohydrolase (DDAH). It is unclear how activity of

PRMT and DDAH are regulated in the disease state. In cardiovascular disease states levels of reactive oxygen species (ROS) and reactive nitrogen species (RNS) are elevated and they have been found to play a role in pathogenesis of cardiovascular disease. Therefore, we hypothesized that ROS and RNS may be involved in regulation of PRMT and DDAH activity.

Here we present findings that the activity of DDAH-1 and PRMT-1 are regulated by reactive species. Activity of purified recombinant hDDAH-1 is found to be dose-dependently inhibited by the lipid hydroperoxide 4-hydroxy-2-nonenal (4-HNE). In addition, the basic enzyme kinetic properties of purified recombinant hDDAH-1 are determined. Activity of

PRMT-1 is demonstrated to be increased by physiological levels of NO. Both of these findings

have important implications on regulation of ADMA levels and NO production.

Recombinant hDDAH-2 was purified, but it was found to lack ADMA metabolizing properties. This finding is consistent with other results from our lab, in which, silencing of

DDAH-2 resulted in decreased NO production but levels of ADMA were unchanged. Currently, the role of DDAH-2 in regulation of NO production is unclear, but it appears that DDAH-2 regulates NO production through an ADMA-independent mechanism.

These findings demonstrate that ADMA metabolism is regulated by redox mechanisms.

CHAPTER 1 INTRODUCTION

Cardiovascular disease is the number one cause of death in the United States. It is estimated that 81,100,000 people in the United States in 2006 had one or more forms of cardiovascular disease which include: high blood pressure, coronary heart disease, stroke and heart failure. In 2010, it is estimated that an American will die every minute from a coronary event. There are a number of underlying factors that lead to the development of cardiovascular disease, including: hypertension, smoking, high cholesterol, diabetes and obesity. The total direct and indirect cost of cardiovascular disease in the United States in 2010 is estimated to be

$503.2 billion. This estimate includes health care costs and loss of productivity due to morbidity and mortality [1]

One of the hallmarks of cardiovascular disease is endothelial dysfunction. Endothelial dysfunction is characterized by a loss in production of nitric oxide (NO) from the endothelium of

the vasculature. Reduced NO production results in impaired vasodilation, increased monocyte

adhesion, increased platelet aggregation, increased vascular smooth muscle cell proliferation

(VSMC), increased oxidation of low-density lipoprotein and increased superoxide elaboration.

These factors lead to plaque formation on the vascular wall and narrowing of the vessel. This

increases the blood pressure and work load on the heart. Thereby, the risk of a cardiac event is

increased.

NO is produced by the enzyme nitric oxide synthase (NOS) in a reaction in which L-

Arginine is oxidized to L-Citrulline and NO. Production of NO can be inhibited by the

endogenous methylarginine asymmetric dimethylarginine (ADMA). ADMA is a competitive

inhibitor of NOS and levels of ADMA have been found to be significantly increased in

cardiovascular disease and other disease states. A family of proteins known as protein arginine

methyltransferases (PRMT) are responsible for the formation of ADMA. PRMT methylates

arginine residues on protein and then during proteolysis free ADMA is released into the cytosol

where it can inhibit NO production from NOS. Free-ADMA is metabolized by dimethylarginine dimethylaminohydrolase (DDAH) to L-Citrulline and dimethylamine. In cardiovascular disease states, levels of ADMA have been found to be increased by impairment of DDAH activity and increased expression of PRMT. My dissertation is focused on redox regulation of ADMA metabolic pathways and their role in the pathogenesis of endothelial dysfunction.

eNOS

Nitric oxide synthase (NOS) plays a central role in the regulation of various biological

processes through the formation of the gaseous free radical nitric oxide [2]. Endogenous nitric

oxide (NO) acts as an essential signaling molecule and effector in cardiovascular, neuronal, and

immune systems [3-6]. In endothelial cells, NO is derived from the guanidino group of L-

arginine in a reaction catalyzed by eNOS [7]. eNOS is constitutively present in cells, however it

requires several cofactors for catalytic activity. Activation of eNOS requires Ca2+/calmodulin, hence NO production from eNOS is initiated and modulated by elevated intracellular free Ca2+

[8,9]. In addition, eNOS activity can be modulated post-transcriptionally by phosphorylation and protein-protein interactions.

eNOS structurally resembles nicotinamide adenine dinucleotide phosphate (NADPH) cytochrome P-450 reductase [10]. eNOS uses L-arginine, oxygen, and NADPH as substrates to

synthesize NO. Tetrahydrobiopterin (BH4), CaM, FAD, and FMN are the requisite cofactors for

this catalytic process [3,11]. eNOS has three domains that are necessary for catalytic activity the

reductase, oxygenase and Cam binding domain [12-14]. In the reductase domain, FAD and

FMN shuttle electrons donated by NAPH to the heme binding site of the oxygenase domain.

This transfer of electrons requires Ca2+/calmodulin binding at the CaM binding domain. The

oxygenase domain also contains binding sites for L-Arg and BH4 in addition to a heme. At the

oxygenase domain L-Arg is oxidized to NO and L-Cit.

The primary function of NO in the vasculature is to mediate relaxation of vascular

smooth muscle cells (VSMCs). NO is produced in the endothelium and diffuses through the cell

membrane to the VSMC layer where it binds to the heme of guanylate cyclase activating the

enzyme. Once activated, guanylate cyclase catalyzes the reaction of guanosine

triphosphate (GTP) to cyclic guanosine 3’5’-monophosphate (cGMP) and inorganic phosphate

[15]. cGMP then activates protein kinase G (PKG), which in turn phosphorylates myosin light

chain phosphatase. Once activated by phosphorylation, myosin light chain phosphatase

dephosphorylates myosin light chain, resulting in relaxation of VSMCs and relaxation of the

blood vessel.

In addition to vascular relaxation, NO also has anti-thrombotic, anti-proliferative, and anti-atherogenic effects. These effects are critical in the vascular response to injury wherein the vascular wall elicits a proliferative VSMC response resulting in increased intimal/media ratio and accompanying lumen loss. Narrowing of the lumen can result in occlusion of the vessel and subsequent cardiac events. The mechanism through which NO inhibits proliferation of VSMC involves regulation of VSMC cell cycle progression. NO halts VSMC growth in the S-phase of the cell cycle through down-regulation of cyclin-dependent kinase 2 (CDK-2) protein activity and cyclin-A gene transcription [16]. Thus, loss or attenuation of NO production would result in increased VSMC proliferation after vascular injury and narrowing of the vessel lumen.

NO also exhibits anti-atherogenic activity and reduced bioavailability is implicated in the progression of atherosclerosis. During atherogenesis, endothelial cells become activated through up-regulation of cellular adhesion molecules E and P selectin, vascular cell adhesion molecule-1

(VCAM-1) and inter-cellular adhesion molecule 1 (ICAM-1) and pro-inflammatory genes. The

chemokines monocyte chemotactic protein-1 (MCP-1) and interleukin-1 (IL-1) are up-regulated

as well. Increased expression of these genes results in increased leukocyte rolling and

attachment to the endothelium. NO inhibition of NF-κβ signaling attenuates increased

expression of VCAM-1 and ICAM-1 resulting in decreased monocyte attachment to the

endothelium [17,18].

Platelet activation and aggregation are also important factors in the progression of

atherosclerosis. Injury to the endothelium results in expression of pro-thrombotic proteins,

collagen, von Willebrand factor (vWF) and tissue factor (TF), on the vessel wall. Platelets become activated when exposed to collagen or vWF, and TF initiates platelet activation by facilitating the conversion of pro-thrombin to thrombin through activation of factor X. The activated form of factor X, factor Xa, is responsible for conversion of the zymogen pro-thrombin to the active enzyme thrombin. NO acts through a cGMP dependent mechanism to inhibit platelet aggregation by inhibiting intracellular Ca2+ mobilization [19].

In addition to the vascular protective effects elicited by NO, eNOS, through its oxidase acitivity can also contribute to endothelial dysfunction. For example, under conditions of substrate depletion or BH4 cofactor depletion eNOS becomes “uncoupled” and produces

- superoxide anion radical, O2 , instead of NO [20-22]. In this enzymatic state, NADPH

- - oxidation is uncoupled from NO synthesis and O2 is reduced to O2 [21,23]. Production of O2 by eNOS leads to reduced NO availability in two ways. First, eNOS production of NO is

- - decreased and O2 production is increased. Second, O2 acts as a free radical scavenger and

- - reacts with NO to form peroxynitrite (ONOO ), a potent oxidant. O2 also activates cellular signaling mechanisms that are often opposite of the effects of NO. In this regard, impaired eNOS

activity may contribute to endothelial dysfunction through two distinct but related events;

- reduced NO and increased O2 . eNOS Regulation by Protein-Protein Interactions

NO biosynthesis is under the regulatory influence of a variety of mechanisms ranging from transcriptional to post-transcriptional controls. Among these mechanisms, protein-protein interactions have been demonstrated to play an important role in NOS regulation in both health and disease. Among the most widely studied of these interactions, is the role of calmodulin in the regulation of NOS activity and NO biosynthesis. Calmodulin has been demonstrated to act as an allosteric activator of eNOS. eNOS is a calcium dependent enzymes and is inactive at resting calcium concentrations [8,24]. Elevations in intracellular calcium concentrations induce CAM binding resulting in activation of eNOS [25].

In addition to direct activation induced upon CaM binding, it has been demonstrated by several groups that caveolin binding by eNOS is regulated by CaM-eNOS interactions [26,27].

Caveolin-1, the predominant scaffolding protein of caveolae, has been shown to directly associate with and tonically inhibit eNOS in endothelial cells. Under resting conditions, eNOS associates with caveolin-1 and this interaction maintains NOS in an inactive state. Following the intracellular rise in calcium that accompanies agonist stimulation, CaM binds eNOS and caveolin dissociates form the enzyme resulting in an activated eNOS-CaM complex. As the agonist invoked stimulation subsides, intracellular calcium levels drop resulting in eNOS disassociation from CaM with subsequent binding to caveolin resulting in inactivation [26,27].

The family of heat shock proteins, specifically hsp90, represents another NOS associated protein which influences the catalytic activity of eNOS. Hsp90 is a highly abundant cytosolic protein known to serve as a molecular chaperone in protein folding and maturation events [28].

Hsp90 has been found to serve as an allosteric activator of eNOS [29,30]. Garcia-Cardena et al.

have demonstrated that vascular endothelial growth factor, histamine, and fluid shear stress

induce association of Hsp90 with eNOS and enhance eNOS activity. Blockade of Hsp90-

mediated signaling limits both agonist-induced NO production and vasorelaxation [29]. In addition, hsp90 and eNOS appear to form a multiprotein complex that includes the kinase Akt.

In support of this, stimulation of endothelial cells with vascular endothelial growth factor has been shown to trigger the recruitment of eNOS and Akt to the same domain of Hsp90, facilitating

Akt-driven phosphorylation of eNOS and promoting NO release [31].

eNOS Regulation by Phosphorylation

Phosphorylation is a major regulatory mechanism of protein function. Kinases are

responsible for phosphorylation of proteins, most commonly at serine, threonine or tyrosine

amino acid residues. Kinases are a large and diverse family of enzymes, and as would be

expected their actions are also highly varied. Upon phosphorylation, the activity of a protein can

be increased or inhibited. In addition, cellular compartmentalization of proteins can be regulated

by phosphorylation.

eNOS is phosphorylated in response to a variety of factors, including hormones, cell

stress, growth factors and other cellular signaling molecules. Depending on the site of

phosphorylation, eNOS activity can be increased or decreased with subsequent regulation of

eNOS derived NO production. The most extensively studied site of eNOS phosphorylation is

Ser1177 human sequence/Ser1179 bovine sequence.

Ser1177 phosphorylation rapidly increases the production of NO primarily through increasing electron flux through the reductase domain and by reducing calmodulin dissociation at

low calcium levels [32]. Typically found unphosphorylated in endothelial cells, exposure to

stimuli causes rapid phosphorylation of Ser1177. Fluid shear stress phosphorylates Ser1177 [33] through both Akt/PKB [34] and PKA pathways [35]. Akt phosphorylation of Ser1177 is mediated

through activation of phosphatidylinositol 3-kinase (PI(3)K), an upstream regulator of Akt

activity. Exposure of VEGF [36], estrogen [37], sphingosine-1-phosphate [38], and hydrogen

peroxide also induce Ser1177 phosphorylation through the PI(3)K/Akt pathway. Insulin has been found to induce Ser1177 phosphorylation through both the PI(3)K/Akt pathway [39] and through the AMP-activated protein kinase (AMPK) [40]. PKG has also been shown to phosphorylate

Ser1177 [41]. Bradykinin and histamine are G protein-coupled receptor agonists, and are responsible for CaM-dependent kinase II (CaMKII) phosphorylation of Ser1177 [42]. CamKII

Ser1177 phosphorylation is associated with increased intracellular Ca2+ levels, but PI(3)K/Akt

Ser1177 phosphorylation derived increases in NO production are believed to be independent of intracellular Ca2+ levels [32].

To date protein phosphatase 2A (PP2A) is the only phosphatase that has been shown to

dephosphorylate Ser1177 [43], through two known mechanisms. Exposure of phorbol 12-

myristate 13-acetate (PMA) has been shown to activate PP2A [44]. In addition, proteasome inhibition has been found to dephosphorylate Ser1177 through enhanced PP2A association with eNOS [45]. Ubiquitinated PP2A dephosphorylates Akt, then is translocated to the cell membrane where it dephosphorylates eNOS and thus NO production is decreased. A negative feedback mechanism between NO and Ser1177 phosphorylation exists. NO mediates cytosolic

depletion of Akt through caspases resulting in decreased Ser1177 phosphorylation [46]. When

- uncoupled, by depletion of BH4, eNOS produces superoxide. Production of O2 by eNOS has not been found to be regulated by Ser1177 phosphorylation [47] or phosphorylation at other sites on eNOS.

Thr495 human sequence/Thr497 bovine sequence is located within the CaM binding domain of eNOS. Thr495 is constitutively phosphorylated and phosphorylation of this residue

leads to a decrease in eNOS activity and decreased NO production by prevention of CaM

binding [42]. PKC has been shown to be responsible for Thr495 phosphorylation [42,48] through

PMA [44]. Dephosphorylation of Thr495 is necessary for eNOS activation and CaM binding.

The exact mechanism for Thr495 dephosphorylation is unclear. There is evidence to support that

protein phosphatase 1 (PP1) [44,49] PP2A [43] and calcineurin [49] are responsible for dephosphorylation of Thr495. Stimuli that increase intracellular Ca2+ levels, such as bradykinin

and histamine are associated with alteration of Thr495 phosphorylation, but increases in NO

production are not a result of only increased association of Ca2+ with CaM [28]. In addition,

there is evidence that eNOS is regulated by reciprocal dephosphorylation of Thr497 and phosphorylation of Ser1179. Using Thr497A and Thr497A/Ser1179D mutants, Lin et.al demonstrated that Thr497 phosphorylation status plays a critical role in the regulation of eNOS, with both mutants generating 2-3 times more superoxide anion than WT eNOS, and both basal and stimulated interactions of Thr497A/Ser1179D eNOS with hsp90 were reduced in co-

immunoprecipitation experiments. Thus, the phosphorylation/ dephosphorylation of Thr497 may

be an intrinsic switch mechanism that determines whether eNOS generates NO versus

superoxide in cells. These results demonstrate an interrelationship between eNOS

phosphorylation sites and association with Akt and heat shock protein 90 (hsp90) [31] and

suggests that multiple phosphorylation sites of eNOS must act in concert to maximize production

of NO or to fine tune production of NO.

Additional sites of eNOS phosphorylation have been identified but are poorly understood.

Ser114 human sequence/Ser116 bovine sequence is located in the oxygenase domain of eNOS.

Ser114 is basally phosphorylated and causes inhibition of eNOS similar to that observed with

Thr495. Fluid shear stress [50] and lysophosphatidic acid [28] have been shown to phosphorylate

Ser114, but the mechanism of phosphorylation is unclear. Gallis et al. speculate that a proline- directed protein kinase is responsible for phosphorylation of Ser114 due to a neighboring proline

residue. VEGF activates Ser114 dephosphorylation that is inhibited by cyclosporin A, suggesting that calcineurin may be responsible for dephosphorylation of Ser114; dephosphorylation of Ser114

results in increased production of NO by eNOS [51]. This action of VEGF is in contrast to its

Akt mediated phosphorylation of Ser1177 [36], supporting a complex relationship between eNOS

114 2+ phosphorylation sites. The proximity of Ser to the Zn and BH4 binding sites of eNOS

- suggest a possible role in eNOS dimerization and O2 production due to eNOS uncoupling, but this is yet to be shown.

Ser615 human sequence/Ser617 bovine sequence and Ser633 human sequence/Ser635 bovine sequence have also been shown to be phosphorylated. Both residues are located in the CaM auto-inhibitory loop within the FMN binding domain of eNOS. Ser615 is rapidly phosphorylated upon exposure to stimulus resulting in increased eNOS activity by increased Ca2+/CaM

association, Ser633 is then phosphorylated and eNOS activity is increased to a level similar to that caused by Ser1177 phosphorylation [52]. Bradykinin, ATP, and VEGF are responsible for

phosphorylation of both sites [52] and fluid shear stress causes Ser633 phosphorylation via PKA

(31). PKG has also been associated with Ser633 phosphorylation [41].

Tyrosine residues have been reported to be phosphorylated in primary cultured endothelial cells [53], but in passaged endothelial cell cultures tyrosine phosphorylation has not been detected [33,54]. Tyrosine phosphorylation may be involved in the association of eNOS

with scaffolding and regulatory proteins [53,55].

There is evidence to suggest that phosphorylation of eNOS can result in cellular

compartmentalization of eNOS. Bradykinin induced phosphorylation has been found to

translocate eNOS from the plasma membrane to the cytosol [54], but this is not a universal

finding [56]. Bradykinin stimulated tyrosine phosphorylation of eNOS appears to target eNOS

to the cytoskeleton [57]. Other research has found that hydrogen peroxide stimulated tyrosine phosphorylation of eNOS causes association with caveolin-1 [58]. Ser1177 phosphorylated eNOS

has been found to be active in two distinct subcellular locations, caveolin-1-enriched plasma membranes and in the membrane of the Golgi apparatus [59]. The effect of eNOS phosphorylation on translocation is poorly understood, but it is clear that P-eNOS is found and is active in multiple locations within the cell. eNOS Regulation by Endogenous Methylarginines

eNOS activity is also influenced by the methylated arginine derivatives, ADMA and L-

NMMA. The biological significance of these guanidino-methylated arginine derivitives has been known since the inhibitory actions of NG-methyl-L-arginine (L-NMMA) on macrophage induced cytotoxicity were first demonstrated [60]. It was subsequently realized that these effects were

mediated through inhibition of NO release. Levels of ADMA have been found to be

significantly increased in a number of diseases related to endothelial dysfunction including

hypertension, hyperlipidemia, diabetes mellitus and others [61-65]. In this regard, intracellular

accumulation of ADMA has been implicated as causative factor in the development of a variety

of pathological conditions through the impairment of the normal physiological functions of NO

[66-69].

ADMA and L-NMMA enter the cell through the y+ cationic amino acid (CAT) carrier

family [70]. Intracellulary, ADMA and L-NMMA compete with L-Arg for binding at the active

site of NOS. This competitive inhibition of NOS activity results in decreased production of NO.

Kinetic studies on purified eNOS have demonstrated that the Km of eNOS for L-Arg is 3.14 μM

-1 -1 with a Vmax of 0.14 μmol mg min , and the Ki values of ADMA and L-NMMA are 0.9 μM and

1.1μM respectively [71]. Under normal physiological levels of ~100 μM L-Arg and 3.6 μM

AMDA and 2.9 μM L-NMMA only about ~10% inhibition of eNOS would be expected.

However, under disease conditions plasma ADMA levels can increase 3- to 9-fold [65,67,72-76].

The multicenter Coronary Artery Risk Determination investigating the Influence of ADMA

Concentration (CARDIAC) study identified plasma ADMA levels as an independent risk factor

for coronary heart disease (CHD) [77]. In addition, accumulation of ADMA by inhibition of

DDAH-1 activity has been demonstrated to result in decreased NO production and increased

systemic vascular resistance and elevated systemic and pulmonary blood pressure [78]. ADMA

levels are controlled by the activity of PRMT and DDAH. The PRMT pathway is responsible

for synthesis of ADMA by methylation of Arginine residues on proteins and DDAH is the

primary metabolic pathway of ADMA.

PRMT

Protein arginine residues are methylated by a family of proteins known as PRMTs. Upon

proteolysis of these methylated proteins ADMA and L-NMMA are released into the cytoplasm

where they inhibit NO generation by NOS [79]. To date, ten mammalian isoforms of the PRMT

family have been identified. In mammalian cells, PRMTs have been classified into type I

(PRMT-1, -3, -4, -6 and -8) and type II (PRMT-5, -7, and FBXO-11), based on their specific catalytic activity. The enzymatic activity of PRMT-2 and PRMT-9 is unclear [80]. Type I and II

PRMTs both catalyze the formation of mono-methylarginine (MMA) from L-Arginine. In a second step, type I PRMTs produce ADMA while type II PRMTs catalyze the formation of symmetric dimetylarginine (SDMA) [81,82]. As with other post-translation modifications such as phosphorylation, arginine methylation regulates protein activity. Over the last 40 years, arginine methylation has been extensively studied in prokaryotes and eukaryotes revealing a pivotal role of this posttranslational modification in the regulation of a number of cellular

processes including; transcription, RNA metabolism and protein-protein interactions, thereby controlling cellular differentiation, proliferation, survival, and apoptosis [81].

The catalytic activity of PRMTs is characterized by the transfer of a methyl group from

S-adenosylmethionine (SAM) to the guanidino nitrogen of arginine [83]. Glycine-arginine-rich motifs on proteins are preferential targets for methylation by the majority of PRMTs [84].

PRMT-4 and PRMT-5 methylate arginine residues with proline-, glycine-, or methionine-rich domains [85]. Transfer of one methyl group results in the formation of MMA and transfer of two methyl groups can result in production of ADMA or SDMA. Although protein-arginine methylation has not been shown to be reversible, the Jumonji domain-containing 6 protein

(JMJD-6) has been shown to demethylate histone-arginine residues [86]. In addition, prior to proteolysis, protein-incorporated MMA can be converted to citrulline by peptidylarginine deiminase 4 (PAD-4) [87]. LSD-1 is potentially an enzyme that is capable of demethylating protein-arginine residues. LSD-1 has been shown to demethylate histone-lysine residues although it is unclear if it has arginine demethylation activity [88]. PRMTs are ubiquitously expressed, however, gene-splicing may allow for tissue specific expression [89]. Two isoforms of PRMT have been identified as essential for embryonic development. PRMT-1 is necessary for early postimplantation development and PRMT-1 knock-out mice die at approximately embryonic day 6.5 [90]. Pawlak et al. also showed that PRMT-1 is responsible for over 85% or total PRMT activity in embryonic stem cells. PRMT-4/CARM-1 knock-put mice die at birth and have disrupted estrogen-responsive gene expression [91].

The activity of PRMT has been found to be regulated by binding of non-substrate proteins. Binding of BTG-1 and BTG-2/TIS-21 has been found to increases activity of PRMT-1

[92]. PRMT-3 activity is inhibited by binding of the tumor suppressor DAL-1 [93]. PRMT-1

has the ability to form homodimers and deletion of the protein domain facilitating dimerization

results in loss of PRMT-1 enzymatic activity [94].

Levels of PRMT expression has been found to be altered in various disease states. Chen

et al. noted increased levels of PRMT-1 protein in diabetic rat retina which was reversed by

administration of benazepril or telmisartan [95]. This study also found that exposure of bovine

retinal capillary endothelial cells to high glucose levels resulted in increased protein expression

of PRMT-1. Exposure of high glucose causes increased level of ROS production suggesting a

possible role of ROS mediated PRMT-1 expression. Exposure of human umbilical vein

endothelial cells (HUVECs) to fluid shear stress of > or =15 dyne/cm2 resulted in a two-fold

increase in expression of PRMT-1 mRNA via activation of the nuclear factor (NF)-kappaB pathway [96].

DDAH

The majority of the body’s clearance of ADMA and L-NMMA occurs through metabolism by DDAH [97-99]. The remaining ~ 10% of clearance is through the urine. DDAH hydrolyses the reaction of ADMA to L-citrulline and dimethyamine and of L-NMMA to L- citrulline and monomethylamine. The third methylarginine, SDMA, is unable to be metabolized by DDAH due steric and electrostatic hindrance caused by having a methyl group on both of the nitrogen side of chains of SDMA [100]. DDAH contains an acidic binding pocket where the unmethylated nitrogen side chain of ADMA and L-NMMA enter to initiate catalysis. There are two isoforms of DDAH that have been identified in humans DDAH-1 and DDAH-2. Both isoforms consist of 285 amino acids and share 62% homology.

The first report of metabolism of ADMA was by Ogawa et al. in 1987 when they identified a new enzyme referred to as NG,NG-dimethylarginine dimethylaminohydrolase [101].

At this time the role of ADMA in inhibition of NOS activity was unclear and regulation of NO

production by DDAH was unknown. In 1996, the Vallance group implicated DDAH as being a

regulator of NO synthesis [102]. A second isoform, DDAH-2, was identified by Lieper and

Vallance in 1999 after the observation that DDAH-1 expression did not correlate to DDAH

activity [103].

Both isoforms of DDAH are predominately found in the cytosol of cells with some

DDAH-1 present in the cell membrane fraction of endothelial cells [104]. DDAH-1 is found to be highly expressed in the liver and kidney, is strongly expressed in the pancreas, forebrain, aorta, peritoneal neutrophils and macrophages, and is widely expressed in other tissues [105-

110]. Expression levels in fetal and adult tissue are equal [109]. The liver and kidney are major

metabolic sites of ADMA suggesting that DDAH-1 plays an important role in regulation of

plasma ADMA levels. DDAH-2 is found to be expressed predominately in the vascular

endothelium, with strong expression in the heart, placenta and kidneys [107,109]. DDAH-2 is

also expressed in the spleen, thymus, peripheral leukocytes, lymph nodes, and bone marrow which all also express iNOS [109]. DDAH-2 expression is high in all fetal tissues but expression

decreases and is less ubiquitous in adult tissue [109]. DDAH-2 has also been reported in the

nucleus of rat vascular smooth muscle cells [111]. However, the role of DDAH-2 in the nucleus

is currently unclear.

Several methods of post-transcriptional regulation of DDAH activity have been previously reported. DDAH-1 activity has been shown to be inhibited by Zn(II) with a dissociation constant of 4.2 nM [112]. Due to the tight binding of inhibitory Zn(II) to DDAH-1

it is necessary to utilize a Zn(II) chelating buffer such as Imidazole when performing enzymatic

activity experiments. Inhibition of DDAH-1 activity by NO has also been reported. Exposure of

bacterial DDAH to the NO donor 2-(N,N-dimethylamino)-diazenolate-2-oxide·Na

(DEANONOate) causes S-nitrosylation at cysteine 249, part of the catalytic triad of DDAH

[113]. This S-nitrosylation can be reversed by exposure of the S-nitrosylated DDAH to the reducing agent dithiothreitol (DTT). In addition, the Zn(II) free, but not Zn(II) bound form of

bovine DDAH-1 is S-nitrosylated at cysteine 221 and 273 by exposure to DEANONOate [114].

Detection of heterologously expressed human DDAH-2 in endothelial cells has been detected after stimulation of iNOS [113]. It is interesting to note that some disease states such as sepsis result in increased production of NO from iNOS. Inhibition of DDAH activity due to S- nitrosylation by NO may act as mechanism to inhibit overproduction of NO by preventing degradation of NOS inhibitors ADMA and L-NMMA.

Cellular studies utilizing endothelial cells or smooth muscle cells have identified other modulators of DDAH activity. The following have been reported to increase DDAH activity: probucol [115], taurine [116], insulin and adiponectin [117], pravastatin [118], estradiol

[119,120], interleukin-1 beta (IL-1β) [121], and fenofibrate [122]. The following have been

reported to decrease DDAH activity: vasoconstrictive coupling factor 6 (CF-6) [123],

lipopolysaccharide [124], glycosylated bovine serum albumin [125], the erythropoietin analogs, epoetin-β and darbepoetin- [126], high concentrations of glucose [127], oxidized low density lipoprotein (oxLDL) or tumor necrosis factor alpha (TNF- ) [128], S-nitroso-L-homocysteine

[129], and cytomegalovirus infection [130]. As many of the factors that alter DDAH activity are accompanied by increased levels of oxidative stress it has been speculated that free radicals can modulate DDAH activity. Later I will present work analyzing the role of reactive oxygen species (ROS) and reactive nitrogen species (RNS) on the activity of purified human recombinant DDAH-1.

In blood vessels, studies have reported higher levels of DDAH-2 as compared to DDAH-

1. mRNA analysis of rat mesenteric resistance vessels found 5.1-fold higher levels of DDAH-2

than DDAH-1 [131]. Analysis of human endothelioma cells found 10-fold higher expression of

DDAH-2 mRNA than DDAH-1 mRNA [131]. These findings suggest an important role of

DDAH-2 in the vasculature. In support, Wang et al. demonstrated that siRNA knockdown of

DDAH-2 resulted in almost complete inhibition of EDRF/NO response to acetylcholine in rat mesenteric blood vessels and an accompanying reduction in NO as detected by 4,5- diaminofluorescein acetoxymethyl ester. Although DDAH-2 plays an important role in

regulation of EDRF/NO response in the vasculature it appears as though this regulation is

ADMA-independent. Studies from our laboratory have found that siRNA silencing of DDAH-2

in endothelial cells reduces NO levels as detected by Fe-MGD spin trapping, but an

accompanying reduction in L-Arg/ADMA ratio is not observed [132]. In this study, siRNA

silencing of DDAH-1 resulted in decreased production of NO and an accompanying reduction in

the L-Arg/ADMA ratio. Cell media supplementation with L-Arg was able to partially overcome

the reduction in NO production with DDAH-1 silencing but not with DDAH-2, further

supporting the hypothesis that DDAH-2 elicits its NO regulatory effects independent of ADMA.

Other groups have found ADMA independent mechanisms of DDAH action. In the study

by Wang et al., DDAH-2 siRNA silencing resulted in decreased eNOS mRNA and protein

expression [131]. Two groups have found that DDAH-2 over-expression resulted in increased transcription of vascular endothelial growth factor (VEGF) [133,134]. Hasegawa et al. found increased VEGF transcription was caused by DDAH-2 binding to PKA and subsequent phosphorylation of the transcription factor specificity protein-1 (Sp-1). Sp-1 then translocates to

the nucleus where it binds to the promoter of VEGF and initiates transcription. DDAH-1 has

been reported to bind to nuerofibromin-1 (NF-1) and cause increased phosphorylation of NF-1 by PKA [135].

An intriguing study on the role of DDAH on human disease was the Kuopio Ischemic

Heart Disease Risk Factor Study. In this study of 1,609 middle-aged Finnish men six single

nucleotide polymorphisms (SNPs) were detected in the gene for DDAH-1 and one SNP was detected in the gene for DDAH-2. 13 individuals were identified with the DDAH-1 SNP and the occurrence of coronary heart disease was 50-fold higher in these individuals. In addition, about half of the relatives of individuals with the DDAH-1 SNP were found to have the mutation as well. Relatives with the DDAH-1 gene mutation had an increased prevalence of hypertension

[136].

In a recent study by Breckenridge et al., DDAH-1 and DDAH-2 transgenic gene deletion mice were developed and characterized. Homozygous DDAH-1 null embryos were generated at low frequency and were embryonic lethal. DDAH-1 -/- blastocysts were generated at about 5% frequency and were unable to undergo implantation suggesting a role of DDAH-1 in early embryogenesis or fertilization. DDAH-2 -/- mice were viable, fertile and had a normal lifespan

[137]. The triple eNOS/iNOS/nNOS knockout mouse is viable suggesting that the lethality of

DDAH-1 gene deletion is not NO-dependent [138]. Studies of DDAH-1 +/- transgenic mice produced viable animals that had increased plasma levels of ADMA [78]. In these animals; kidney, lung, and liver DDAH activity levels dropped approximately 50% and DDAH-2 expression was unchanged, suggesting that DDAH-2 is not the predominate route of ADMA metabolism in these tissues. The DDAH-1 +/- mice also showed characteristics of endothelial dysfunction. Vascular relaxation in response to acetylcholine was blunted suggesting impaired

EDRF/NO response. In addition, mean arterial blood pressure, systemic vascular resistance, and

right ventricular pressure were all significantly increased.

These findings show a role of DDAH and ADMA in endothelial dysfunction. DDAH is

involved in both ADMA-dependent and ADMA-independent actions in the progression of

cardiovascular disease states. In the next section the role of ADMA in disease states will be

more closely examined.

ADMA in Disease

Plasma levels of ADMA have been found to be elevated in a number of disease

conditions related to endothelial dysfunction such as hypertension, hyperlipidemia, diabetes

mellitus and others [61-65]. Initial reports in the field by Vallance et al. described increased

levels of ADMA in the plasma of patients with chronic renal failure [69]. Later, AMDA was

identified as an independent risk factor in cardiovascular disease states. Evaluation of plasma

ADMA levels from patients with CHD in the CARDIAC study identified ADMA as an

independent risk factor for CHD [77]. In addition, ADMA has been identified as a predictor of future cardiovascular events in patients undergoing percutaneous coronary intervention [139].

Indeed, Schnabel at al. identified high levels of baseline ADMA as an independent predictor of cardiovascular events in patients with coronary artery disease [75].

Additional studies have demonstrated an important role of the ADMA/L-Arg ratio in endothelial dysfunction. A study on patients with hypercholesterolemia found ADMA increases mononuclear cell adhesiveness and this relationship is inversely correlated with plasma

ADMA/L-Arg ratio [140]. Normalization of the plasma ADMA/L-Arg ratio with oral supplementation of L-Arg resulted in ablation of the increased mononuclear cell adhesiveness.

Mononuclear cell adhesion is an initial step in plaque formation on the vessel wall. In a study on regenerated endothelial cells following balloon denudation in rabbit carotid artery impaired

endothelial dependent relaxation and an increased ADMA/L-Arg ratio was noted in regenerated endothelial cells [141]. Addition of L-Arg prevented the impaired endothelial dependent relaxation.

Reports indicate that plasma ADMA levels increase from ~ 0.4 μM to ~ 0.8 μM in patients with pathological conditions such as pulmonary hypertension, coronary artery disease, diabetes, and hypertension [65,77,142-145]. However, levels of ~ 0.8 μM plasma ADMA are likely not high enough to regulate NO production from eNOS given that plasma L-Arg levels are

~ 100 μM. An explanation for impaired endothelial function in patients with elevated plasma

ADMA levels is that plasma ADMA reflects increased intracellular concentration of ADMA. To investigate the role of intracellular ADMA on eNOS function studies our group previously examined cellular uptake uptake of ADMA by BAECs [71]. In the absence of L-Arg, exposure of cells to 10 μM ADMA in the media resulted in intracellular levels of 68.4 μM ADMA.

Addition of 100 μM L-Arg to 10 μM ADMA in the media resulted in intracellular ADMA levels of 23.5 μM. These results indicate that endothelial cells concentrate ADMA and that increased plasma ADMA levels are indicative of even greater increases in intracellular ADMA levels.

Additional studies were conducted to determine if increased ADMA could result in impaired endothelial dependent relaxation. ADMA was found to dose dependently inhibit the Ach mediated relaxation response in rat aortic rings with a 52% reduction seen at 5 μM ADMA and a

95% reduction at 500 μM ADMA, in the absence of L-Arg. In the presence of 100 μM L-Arg,

ADMA impaired relaxation was improved with 7% inhibition at 10 μM and 84% inhibition at

500 μM ADMA.

In support of the role of ADMA and L-NMMA as endogenous inhibitors of NOS function, several studies have demonstrated that L-Arg supplementation enhances endothelial

function through increased NO bioavailability [146-149]. This effect could be due to increased

substrate availability. However, when one takes into consideration that the intracellular levels of

L-Arg are typically more than 50 times higher than the Km of the enzyme, increased NO

generation would not be expected with L-Arg supplementation [150]. This observation of

increased NO bioavailability with L-Arg supplementation has been termed the “arginine

paradox”. One hypothesis put forth to explain this paradox is that L-arg supplementation overcomes the endogenous inhibitory actions of cellular methylarginines [68,151-154]. In support, by directly measuring NO production from endothelial cells and plotting these values as a function of the experimentally measured intracellular ADMA/L-Arg or L-NMMA/L-Arg ratio,

the cellular kinetics of ADMA and L-NMMA inhibition were generated. When these values

were plotted against the curve for competitive inhibition, a strong correlation was observed

demonstrating that cellular inhibition closely follows what would be predicted from the Ki [71].

’ Based on the Ki s for ADMA and L-NMMA, it was then predicted that under normal

physiological conditions endogenous methylarginine levels would inhibit NO production by ∼10-

20%. These results demonstrate the importance of ADMA and L-NMMA in NOS regulation and

support a role for methylarginines in the “arginine paradox”. In addition to these basal effects,

methylarginine levels are known to increase in a variety of pathological settings. There are

numerous studies demonstrating that under pathological conditions methylarginine levels are

increased by 3 to 9 fold [67,68,72,75,155-158], which based on enzyme kinetic data, would be

expected to inhibit NO generation from NOS by 30 - 70% [71]. The equation for inhibition can

thus be extrapolated to studies aimed at measuring cellular methylarginine levels in disease and

allow for the prediction of the extent of NOS inhibition. Indeed, in vitro studies have

demonstrated that the methylarginines, ADMA and L-NMMA, dose dependently inhibit eNOS-

derived NO production from both isolated enzyme and cellular systems [71,150]. The results

obtained from these studies translate to the whole organ as it has been demonstrated that

methylarginines modulate vascular relaxation [71,159]. Measurements of intracellular methylarginine levels suggest that normal physiological concentrations only have a modest effect on NOS activity. However, under pathological conditions such as vascular injury and restenosis, where endogenous methylarginines are elevated, methylarginine-mediated NOS inhibition with secondary endothelial dysfunction has been observed [71,159]. These studies demonstrate that cellular methylarginines are elevated under pathological conditions and reach concentrations sufficient to inhibit NO production. Together, these results represent a major step forward in our understanding of the regulation, impact, and role of methylarginines, in disease.

Recent findings from our laboratory have demonstrated that the endogenous L-Arg

- derivatives, ADMA and L-NMMA, play an important role in the regulation of O2 release from eNOS. When eNOS is uncoupled through depletion of the cofactor BH4, the eNOS derived superoxide that is observed is significantly enhanced in the presence of both ADMA and L-

NMMA [160]. The mechanism for this methylargininie mediated enhancement of eNOS derived superoxide involves the heme center of the oxygenase domain. NADPH consumption studies demonstrate that binding of L-NMMA or ADMA to BH4-free eNOS increases NADPH

consumption rate and studies of midpoint potential reveal that methylarginine binding to NOS

increases the heme reduction potential and thereby facilitates electron transfer through the heme,

- resulting in increased O2 generation from the enzyme. These results would suggest that under

conditions of decreased BH4 bioavailabiltiy, as occurs under a variety of pathological conditions,

- methylarginines would shift the balance of NO and O2 production from eNOS to a largely oxidant producing enzyme.

- In support, a previous study found that L-NMMA induced enhanced O2 generation in

cardiac myocytes subjected to ischemia-reperfusion injury [161]. These results were previously

unexplained; however, ischemia-reperfusion has been shown to result in BH4 oxidation, so the

- increased O2 production induced by L-NMMA is likely NOS-derived and BH4 dependent. In the future, further studies will be needed to characterize the effects of endogenous levels of

- methyl-arginines on O2 production in cells and tissues, as well as the modulatory action of BH4

on methylarginine-NOS interactions in in vivo models of physiology and disease.

It is clear that NO is necessary for maintenance of a healthy vasculature. Endothelial dysfunction, characterized by a loss of NO bioavailability in the vasculature, is found in cardiovascular disease and plays a critical role in the pathological progression of the condition.

The PRMT-ADMA-DDAH axis is one critical factor controlling regulation of NO production.

There is strong evidence that levels of ADMA are elevated in cardiovascular disease and they are responsible for inhibition of NOS. However, it is currently unclear how the synthesis and degradation of ADMA is regulated in cardiovascular disease. Cardiovascular disease is accompanied by increased levels of oxidative stress and it has been hypothesized that oxidative stress can regulate ADMA levels. Therefore, studies were conducted to evaluate the role of oxidative stress in regulation of the PRMT-ADMA-DDAH axis. A better understanding of the factors responsible for regulation of the PRMT-ADMA-DDAH axis will provide valuable insight into the development of endothelial dysfunction and the progression of cardiovascular disease.

CHAPTER 2 REDOX REGULATION OF ADMA METABOLISM: DDAH-1

Introduction

Nitric oxide (NO) is an important regulator of vascular homeostasis. Endothelium

derived NO is responsible for vasodilatation and has anti-proliferative and anti-atherogenic

effects on the vascular wall [2,162,163]. Dysregulation of NO biosynthesis has been noted in

multiple disease states including hypercholesterolemia [164] and atherosclerosis [165]. A

growing volume of literature implicates the endogenous nitric oxide synthase (NOS) inhibitors;

asymmetric dimethyl arginine (ADMA) and NG–monomethyl arginine (L-NMMA) are responsible for the impaired NO production associated with these diseases [166,167]. These methylarginines are produced by the proteolysis of methylated arginine residues on proteins. Six isoforms of the protein-arginine methyl transferase’s (PRMTs) family of enzymes have been identified and are responsible for methylation of arginine residues on proteins. Free ADMA and

L-NMMA, formed during routine proteolysis are subsequently hydrolyzed by Dimethylarginine

Dimethylamino Hydrolase (DDAH) [99]. DDAH metabolizes ADMA to L-citrulline and dimethyamine, and L-NMMA to L-citrulline and monomethylamine. There are two isoforms of

DDAH with distinct tissue localization patterns [103]. DDAH-1 has been implicated as a critical regulator of methyarginine levels and NO production in the vasculature [78]. The activity of DDAH has been shown to be altered in a variety of disorders involving endothelial dysfunction. In chronic heart failure DDAH-1 activity decreases, but protein levels remain constant, suggesting that post-translational modifications are responsible for modulation of

DDAH activity [168]. Unfortunately, little is known with regard to the enzyme kinetics of hDDAH-1. To date, enzymatic studies of DDAH have utilized either recombinant bacterial

DDAH-1 or DDAH-1 partially purified from porcine brain homogenates. Results using these

preparations have determined that DDAH-1 hydrolyses ADMA at faster rate than L-NMMA

with reported Km values of 0.18 and 0.36 mM, respectively and is responsible for >90% of

ADMA metabolism [97,98,169]. DDAH-1 contains a Zn(II) binding site with endogenously

bound Zn(II) inhibiting the catalytic activity of DDAH [112]. In addition to the zinc regulatory

domain it has been demonstrated that NO inhibits DDAH activity through S-nitrosylation of

Cys249 located within the active site of DDAH [113].

Evidence suggests that DDAH-1 activity is inhibited in a variety of cardiovascular

diseases and that this inhibition contributes to impaired NO bioavailability [71,73,170].

Therefore, it is of critical importance to elucidate the pathways and determine the effector

molecules involved in DDAH-1 dysregulation. In atherosclerosis, levels of lipid hydroperoxides,

such as 4-hydroxy-2-nonenal (4-HNE) are elevated. A biologically active major aldehyde of

membrane lipid peroxidation, 4-HNE is formed during inflammation and oxidative stress and

can reach tissue concentrations of 10 µM to 5 mM [171]. Plasma levels of 4-HNE, normally 5-

10 µM, can increase by as much 50-fold under disease conditions [172]. 4-HNE is a highly

reactive aldehyde and has been shown to inhibit the function of multiple proteins mainly through

formation of Michael or Schiff base adducts [173]. A potent nucleophile, 4-HNE preferentially

reacts with cysteine and histidine residues on proteins to form Michael adducts [174]. Therefore,

in the present study we have carried out comprehensive evaluation hDDAH-1 enzyme kinetics

and determined the effects of the lipid peroxidation product, 4-HNE, on DDAH activity as we

have recently observed that 4-HNE inhibits endothelial cell NO production.

Materials and Methods

DDAH Expression and Purification

A bacterial expression system for human DDAH-1, pDEST17-hDDAH1 in BL21star E. coli, was generated using Gateway technology (Invitrogen). This expression system generates human DDAH-1 fused in frame to an N-terminal His6-tag. After growth and induction by

Isopropyl β-D-1-thiogalactopyranoside (IPTG) in 2 L of terrific broth, the bacterial pellet was resuspended in 160 mL lysis buffer consisting of MAC buffer A (20 mM phosphate, 500 mM

NaCl, 30 mM imidazole, pH7.4), with the addition of EDTA-free complete protease inhibitor

(Roche), 1 mM DTT, 1 mM MgCl, 10 ug DNase, 10 ug lysozyme. The cells were homogenized with two passes through an Emulsiflex C3 (Avestin) at 12 – 15 kpsi, and the insoluble material was removed by centrifugation at 4oC (1 hour at 48,000g). The supernatant was loaded onto a 1

mL HisTrap column (GE Bioscience), equilibrated in MAC buffer A. The column was washed

with 30 column volumes of 5% MAC buffer B (20 mM phosphate, 500 mM NaCl, 250 mM

imidazole, pH 7.4), and eluted with 100% MAC buffer B. The fractions containing hDDAH-1

were desalted and exchanged into 25 mM bis tris propane, pH 7.4 (IEX buffer A), using a

HiTrap desalting column (GE Bioscience). The desalted fractions were loaded onto a 1 mL

HiTrap Q sepharose column (GE Bioscience), and eluted with a linear gradient from 0 to 12%

IEX buffer B (IEX buffer A plus 1 M NaCl) over 30 column volumes. All chromatography was

performed at 4 °C using an AKTA purifier 10 (GE Biosciences). The fractions containing

hDDAH-1 were pooled and concentrated using ultrafiltration. The concentrated fractions were

brought up to 10% v/v glycerol and 50 mM sodium phosphate at pH 7.4, fast frozen, and stored

at –80 °C.

DDAH Colorimetric Activity Assay

A modified version of the method designed by Knipp and Vašák was designed [175]. For

kinetic measurements of enzyme activity, purified hDDAH (1.0 μg) was incubated for 5 min at

37 °C in 50 mM imidazole buffer (pH 7.5) in the presence of 1.0−1000 μM substrate with a total

reaction volume of 100 μL. Following incubation, 200 μL of freshly prepared COLDER was

added to each sample, and the plate was incubated for 15 min at 95 °C in an oven. The plate was

allowed to cool to room temperature and citrulline formation was measured at 540 nm. For the

determination of DDAH hydrolytic activity on peptide incorporated ADMA, 10 μg of hDDAH

was incubated in the presence of peptides (1 mM) containing 5 random amino acid sequences

with ADMA at the 3 position (xxADMAxx). Incubation of the reaction mixture (100 μL) was

carried out at 37 °C for 2 h. At the termination of the incubation period, DDAH was removed using a 3,000 MW centricon filter, and the peptide eluent was subjected to acid hydrolysis. Acid hydrolysis was carried out by adding an equal volume of concentrated HCl to the eluent followed by incubation at 105 °C for 18 h. Following acid hydrolysis, samples were neutralized and citrulline formation measured as described above. The standard curve for citrulline was linear with an R2 = 0.99.

DDAH Radioisotope Activity Assay

DDAH activity was measured from the conversion of L-[14C]NMMA to L-[14C]citrulline.

Purified hDDAH-1 was incubated with 4-HNE in 50 μL of PBS for 2 h at 37 °C in a water bath.

A final volume of 100 μL was reached by adding the reaction buffer (50 mM Imidazole, 20 μM

L-[14C]NMMA, 200 μM L-NMMA at pH 7.4) to each sample. The samples were then incubated

in a water bath at 37 °C for 1 h. Following the incubation, the reaction was stopped with 1 mL of

ice-cold stop buffer using 20 mM N-2-hydroxyethylpiperazine-N‘-2 ethanesulfonic acid

(HEPES) with 2 mM EDTA at pH 5.5. Separation of L-[14C]citrulline from L-[14C]NMMA was

performed using the cation-exchange resin Dowex AG50WX-8 (0.5 mL, Na+ form, Pharmacia).

The L-[14C]citrulline in the eluent was then determined using a liquid scintillation counter.

Proteomic Analysis of hDDAH-1: In Gel Digestion Manual

Gels were digested with sequencing grade trypsin from Promega (Madison WI) or sequencing grade chymotrypsin from Roche (Indianapolis, IN) using the Montage In-Gel

Digestion Kit from Millipore (Bedford, MA) following the manufacturer's recommended protocols. Briefly, bands were trimmed as close as possible to minimize background polyacrylamide material. Gel pieces were then washed in 50% methanol/5% acetic acid for 1 h.

The wash step is repeated once before gel pieces are dehydrated in acetonitrile. The gel bands were rehydrated and incubated with DTT solution (5 mg/mL in 100 mM ammonium bicarbonate) for 30 min prior to the addition of 15 mg/mL iodoacetamide in 100 mM ammonium bicarbonate solution. Iodoacetamide was incubated with the gel bands in the dark for 30 min before removed. The gel bands were washed again with cycles of acetonitrile and ammonium bicarbonate (100 mM) in 5 min increments. After the gels were dried in a speed vac, the protease was driven into the gel pieces by rehydrating them in 50 μL of sequencing grade modified trypsin or chymotrypsin at 20 μg/mL in 50 mM ammonium bicarbonate for 10 min.

Then, 20 μL of 50 mM ammonium bicarbonate was added to the gel bands, and the mixture was incubated at room temperature for overnight. The peptides were extracted from the polyacrylamide with 50% acetonitrile and 5% formic acid several times and pooled together.

The extracted pools were concentrated in a speed vac to 25 uL.

Proteomic Analysis of hDDAH-1: Mass Spectrometry LTQ

Capillary-liquid chromatography-nanospray tandem mass spectrometry

(NanoLC/MS/MS) was performed on a Thermo Finnigan LTQ mass spectrometer equipped with a nanospray source operated in positive ion mode. The LC system was a UltiMate Plus system

from LC-Packings A Dionex Co. (Sunnyvale, CA) with a Famos autosampler and Switchos

column switcher. Solvent A was water containing 50 mM acetic acid, and solvent B was

acetonitrile. Five microliters of each sample was first injected on to the trapping column (LC-

Packings A Dionex Co, Sunnyvale, CA) and washed with 50 mM acetic acid. The injector port was switched to inject, and the peptides were eluted off of the trap onto the column. A 5 cm 75

μm ID ProteoPep II C18 column (New Objective, Inc. Woburn, MA) packed directly in the nanospray tip was used for chromatographic separations. Peptides were eluted directly off the column into the LTQ system using a gradient of −80%2 B o ver 50 min, with a flow rate of 300 nL/min. The total run time was 60 min. The MS/MS was acquired according to standard conditions established in the lab. Briefly, a nanospray source operated with a spray voltage of 3

KV and a capillary temperature of 200 °C was used. The scan sequence of the mass spectrometer was based on the TopTen method; briefly, the analysis was programmed for a full scan recorded between 350 and 2000 Da, and a MS/MS scan to generate product ion spectra to determine amino acid sequence in consecutive instrument scans of the 10 most abundant peak in the spectrum. The CID fragmentation energy is set to 35%. Dynamic exclusion is enabled with a repeat count of 30 s, exclusion duration of 350 s, and a low mass width of 0.5 and high mass width of 1.50 Da.

Results hDDAH-1 Protein Purification

An E. coli bacterial expression system was used to produce the hDDAH-1 enzyme.

Following the lysis of bacterial cells, FPLC was used to purify hDDAH-1 from a crude protein homogenate. The homogenate was passed through a nickel metal affinity column to bind His- tagged hDDAH-1 followed by a desalting column and then anion exchange chromatography.

Purity was determined using SDS−PAGE and Coomassie Brilliant Blue staining. Results

demonstrated a 37 kDa band consistent with hDDAH-1 with greater than 95% purity (Figure 2-

1). The yield was approximately 1 mg/L. We found that the purified protein was prone to precipitation after a freeze/thaw cycle, but with the addition of glycerol and the phosphate buffer prior to freezing, there was no precipitation.

Enzyme Kinetics of hDDAH-1

The activity of hDDAH-1 was analyzed by colorimetric detection of citrulline formation from ADMA and NMMA as described previously [175]. DDAH contains a zinc binding domain, and catalytic activity is inhibited by endogenously bound Zn(II); thus, 50 mM imidazole buffer was used for activity assays. Our purified enzyme exhibited the known properties of

DDAH, hydrolyzing the NOS inhibitors ADMA and NMMA to citrulline and showing no activity toward SDMA. In addition, it has been hypothesized that protein methylation may serve as a signaling mechanism and that DDAH may regulate this process through protein demethylation. Therefore, experiments were performed to assess the ability of DDAH to hydrolyze peptide-incorporated methylarginines using random five amino acid polypeptides with

ADMA at position 3. Results demonstrated that DDAH activity is selective for the free methylarginines ADMA and L-NMMA and cannot hyrdrolyze peptide-incorporated methylarginines. Measurements of DDAH enzyme kinetics were performed using a microplate colorimetric activity assay, and the data points were fitted using the Michaelis−Menton equation.

The results demonstrated a Km of 68.7 μM and a Vmax of 356 nmols/mg/min when ADMA was

used as the substrate (Figure 2-2). Similar experiments were repeated with L-NMMA as the

substrate, and the results demonstrated a Km value of 53.6 and Vmax equal to 154 nmol/mg/min

(Figure 2-3). Our results differed from previous studies of DDAH kinetics in which the enzyme was purified from either Pseudomonas or porcine brain homogenates. These studies observed Km

values of 0.18 for ADMA and 0.36 mM for NMMA [176]. Thus, we are among the first to

determine the catalytic activity of human DDAH-1, and these values differ from the values

previously reported for DDAH-1 from other species.

pH Dependence of hDDAH-1

In order to determine the effects of pH on hDDAH-1, enzyme activity measurements

were performed in the presence of 25 mM bis-tris-propane; 25 mM bis-tris buffer with pH

ranging from 5.5 to 9.5. The enzyme was found to be active from pH 6 to 9.5 with maximal

activity observed at pH 8.5. (Figure 2-4). Acidic conditions were found to greatly decrease the

catalytic activity of hDDAH-1 with almost complete inhibition of enzyme activity at pH 5.5.

Maximal activity was measured at pH 8.5 demonstrating that under alkaline conditions activity of the enzyme is increased. This is in contrast to previous studies in which maximal activity was measured at acidic pH values. These findings have important relevance in disease states in which the cellular environment becomes acidic, such as myocardial ischemia, in which cellular pH has been demonstrated to drop to pH 5.5. These results would suggest that under these conditions, DDAH activity would be largely inhibited and may contribute to the endothelial dysfunction and impaired NO synthesis observed in reperfusion injury.

Effect of Oxidants on hDDAH-1 Activity

It has been suggested that the activity of DDAH-1 may be influenced by the redox environment and that exposure of the enzyme to reactive oxygen and nitrogen species may result in oxidant-induced post-translational modifications of critical cysteine residues resulting in loss of enzyme activity. As a result of decreased DDAH activity, methylarginine levels would be expected to rise and result in decreased NOS derived NO. Therefore, we carried out a series of

- studies aimed at determining the dose-dependent effects of NO, ONOO , •OH, and H2O2 on

DDAH activity. NO studies were carried out using the NO donor compound DEANONOate.

- For ONOO and H2O2 studies, the authentic oxidant was used, and in the case of H2O2,

experiments were performed in the presence of 100 μM DTPA to prevent Fenton reactions. The

•OH radical-generating system consisted of H2O2 (1−1000 μM) in the presence of Fe-NTA (20

μM), which has been shown to effectively redox cycle the Fe and result in efficient •OH generation. Exposure of hDDAH-1 to pathophysiological levels of reactive oxygen species was found to have little to no effect on the activity of the enzyme (Figure 2-5). A modest inhibition of 15% was observed following exposure to authentic ONOO- at concentrations ranging from 10 to 100 μM. Higher levels of inhibition were observed at supra-physiological concentrations (1 mM) of H2O2 (43%), ONOO− (25%), and •OH (47%) (Figure 2-5). As previously demonstrated, exposure of hDDAH-1 to NO resulted in significant inhibition at concentrations at or above 100

μM [113]. Overall, hDDAH-1 was found to be largely resistant to inhibition by physiological/pathologically relevant levels of reactive oxygen and reactive nitrogen species.

However, because DDAH activity has been shown to be inhibited in a variety of cardiovascular diseases associated with oxidant stress, we carried out further studies aimed at studying the effects of oxidatively modified lipids on enzyme activity.

Effect of 4-HNE on hDDAH-1 Activity

Among the proposed mechanisms for the impaired NOS activity observed in endothelial dysfunction associated with coronary artery disease are elevated levels of oxidatively modified lipids [177,178]. Polyunsaturated fats in cholesterol esters, phospholipids, and triglycerides are subjected to free radical-initiated oxidation. These polyunsaturated fatty acid peroxides can yield a variety of highly reactive smaller molecules such as the aldehyde 4-HNE upon further oxidative degradation. Therefore, studies were carried out in order to determine the dose- dependent effects of 4-HNE on hDDAH-1 activity. The results demonstrated that exposure of hDDAH-1 to 4-HNE (10 μM−1 mM) at 37 °C caused a dose-dependent inhibition of enzyme catalytic activity with 50% inhibition observed at 50 μM and near complete inhibition at 500 μM

(Figure 2-6). These results demonstrate that at pathologically relevant levels, 4-HNE

significantly inhibits DDAH activity. This observation represents a new mechanism for DDAH

regulation and would be expected to significantly inhibit NOS-derived NO production.

Proteomic Analysis of hDDAH-1

Sequence information from the MS/MS data was processed by converting the raw dta files into a merged file (.mgf) using MGF creator (merge.pl, a Perl script). The resulting mgf files were searched using Mascot Daemon by Matrix Science (Boston, MA). Data processing was performed following the guidelines in Molec. Cell. Proteomics (published online). Assigned

peaks have a minimum of 10 counts (S/N of 3). The mass accuracy of the precursor ions were

set to 2.0 Da, given that the data was acquired on an ion trap mass analyzer, and the fragment

mass accuracy was set to 0.5 Da. Considered modifications (variable) were methionine

oxidation and carbamidomethyl cysteine. Protein identifications were checked manually, and proteins with a Mascot score of 40 or higher with a minimum of two unique peptides from one protein having a −b or −y ion sequence tag of five residues or better were accepted. 4-HNE

modification was searched using PEAKS (Bioinformatics Solutions, Waterloo, ON Canada)

programs. A peptide modified with 4-HNE will result in a mass increase of 156.1150 Da on

cysteine and histidine residues, if compared with the unmodified peptides. Therefore, the

program was set to search peptides with a mass shift of 156.1150 Da as well as the unmodified

peptides. Cysteine carbamido-methylation and methionine oxidation were also considered as variable modification. The mass accuracy of the precursor ions were set to 2.1 Da, and the fragment mass accuracy was set to 0.5 Da. On the basis of the predicted m/z ratios, the results demonstrated that exposure of hDDAH-1 to 50 μM 4-HNE for 1 h resulted in Michael adduct formation at histidine residues 15 and 173 (Figure 2-7 and 2-8). Interestingly, His 173 lies

within the active site catalytic triad of DDAH-1, and mutation of this amino acid has been

demonstrated to result in the near complete loss of enzyme catalytic activity [179,180].

Discussion

Endothelium-derived NO has been demonstrated to function as a critical effector molecule in the maintenance of vascular function [181-183]. In the vasculature, NO is derived from the oxidation of L-arginine, catalyzed by the constitutively expressed enzyme, eNOS

[184,185]. Altered NO biosynthesis has been implicated in the pathogenesis of a variety of cardiovascular diseases including atherosclerosis, heart failure, renal failure, diabetes, preeclampsia, and pulmonary hypertension; it is possible that accumulation of the endogenous

NOS inhibitors, ADMA and NMMA are responsible for the reduced NO generation observed in these conditions [73,99,170]. ADMA and L-NMMA are endogenous inhibitors of NOS exerting their inhibitory effects through competitive inhibition of L-arginine binding and thus preventing

L-arginine from being oxidized by NOS to form NO. Metabolism of these endogenous methylarginines is carried out by DDAH an enzyme which hydrolyzes the conversion of ADMA

to L-citrulline and dimethylamine. Dysregulation of this enzyme has been implicated in a

variety of disorders associated with endothelial dysfunction [73,170]. To date, only a few

studies have been published regarding DDAH enzymology and regulation; these studies have

been carried out using either enzyme purified from pseudomonas or from porcine brain homogenates [113,114,186-188]. Despite the potential importance of this enzyme in NOS regulation there is a paucity of information regarding the kinetics and cellular regulation of the human isoform. In this regard, we have recently expressed and purified hDDAH-1 and have measured the precise kinetic parameters of this enzyme. Results from these studies demonstrated

Km values of 68.7 and 53.6 μM and Vmax values of 356 and 154 nmols/mg/min for ADMA and L-

NMMA, respectively. The maximal enzymatic activity correlates well with previously published

reports in which values of 350-400 nmols/mg/min were reported [189]. However, the Km values obtained differ from those previously published using DDAH purified from other species, in which values of 180-360 µM were measured [190]. This has important physiological

consequence if one considers that under normal conditions, total intracellular levels of

methylarginies in the endothelium are reported to be between 10-20 µM [71]. This would

suggest that under physiological conditions intracellular methylarginine levels are at or near the

Km of DDAH and as a result, loss of enzyme activity would likely result in significant

accumulation of endogenous methylarginines. This would be expected to have significant impact on NOS derived NO as we have previously demonstrated that normal intracellular levels of methylarginines are present at levels sufficient to basally inhibit NOS by 10-15 % [71]. Thus, inhibition of DDAH and subsequent methylarginine accumulation would be expected to result in significant NOS inhibition and may represent a novel mechanism for NOS regulation and may play a role in the pathophsyiology of endothelial dysfunction.

In addition to measuring the precise kinetic parameters of hDDAH-1, we also evaluated the effects of pH on enzyme activity. Studies were performed using BIS-TRIS-Propane/BIS-

TRIS buffer with the pH ranging from 5.5 to 9.5. Results demonstrated that hDDAH-1 was maximally active at pH 8.5 and that the activity decreased with pH and almost complete inhibition was observed at pH values less than 6. This is in contrast to previous studies in which maximal activity was measured at slightly acidic pH values [191]. These finding have important relevance in disease states in which the cellular environment becomes acidic such as myocardial ischemia in which cellular pH has been demonstrated to drop to pH 5.5 [192]. These results would suggest that under these conditions, DDAH activity would be largely inhibited and may

contribute to the endothelial dysfunction and impaired NO synthesis observed in reperfusion

injury.

Recently, Jiang et.al have demonstrated that exposure of endothelial cells to the antioxidant Probucol decreases ADMA levels and enhances DDAH activity [193]. These results suggest that DDAH activity may be modulated by oxidative stress. In this regard, regulation of cellular homeostasis through post-translational modification of proteins is one of the major responses to oxidative and nitrosative stress. Lysine, arginine, proline and threonine side chains can be oxidatively converted to reactive aldehydes or ketone groups (carbonylation) causing inactivation, crosslinking or protein breakdown. Proteins containing cysteine thiol groups are particularly susceptible to oxidation by free radicals, electrophiles, and NO donors. Oxidation of these critical thiol groups can increase or decrease the activity of proteins and represents not only a major mechanism of normal cell signaling but also a mechanism by which disease can interfere with protein function. Therefore, subsequent studies were carried out in order to determine the effects of various reactive oxygen and reactive nitrogen species on DDAH activity. Following exposure of purified hDDAH-1 to varying concentrations (1 µM-1 mM) of NO, ONOO-, •OH and H2O2, DDAH activity was measured. Exposure of hDDAH-1 to pathophysiological levels

of reactive oxygen and reactive nitrogen species was found to have only modest effects on

enzyme activity. A modest inhibition of 10-20 % was observed following exposure to 100 µM

- ONOO , •OH and H2O2. At concentrations of 1 mM, exposure to the reactive oxygen species

•OH and H2O2 resulted in a ~50% inhibition of DDAH activity, however, these levels do not represent physiologically or pathologically relevant concentrations. Consistent with previous findings, exposure of zinc-free hDDAH-1 to the NO donor DEANONOate resulted in significant inhibition [113,114]. Overall, hDDAH-1 was found to be largely resistant to inhibition by

physiological/pathologically relevant levels of reactive oxygen and reactive nitrogen species.

However, because DDAH activity has been shown to be inhibited in a variety of cardiovascular diseases associated with oxidant stress [73,99,170], we carried out further studies aimed at studying the effects of oxidatively modified lipids on enzyme activity.

Among the proposed mechanisms for the impaired NOS activity observed in endothelial dysfunction associated with coronary artery disease are elevated levels of oxidatively modified lipids [177,178]. Polyunsaturated fats in cholesterol esters, phospholipids and triglycerides are subjected to free radical initiated oxidation. These polyunsaturated fatty acid peroxides can yield a variety of highly reactive smaller molecules such as the aldehyde 4-HNE upon further oxidative degradation [194]. 4-HNE is a major biologically active aldehyde formed during lipid peroxidation of w6 polyunsaturated fatty acids which has been shown to accumulate in membranes at concentrations from 10 µm to 5 mM [195]. Therefore, studies were performed in order to determine the dose dependent effects of 4-HNE on hDDAH-1 activity. Results demonstrated that exposure to 4-HNE resulted in a dose dependent inhibition of hDDAH-1 activity with 15% inhibition observed at 10 µM and near complete inhibition at 500 µM 4-HNE.

Mass spec analysis was then performed in order to determine the mechanisms through which 4-

HNE elicits its effects. Previous studies have shown that exposure of proteins to 4-HNE can result in the formation of Michael addition adducts on cysteine and histidine residues [196-198].

Results from our studies demonstrated that exposure of hDDAH-1 to 50 µM 4-HNE for 1 hour resulted in a Michael adduct formation at histidine residues 15 and 173. This has important functional consequences as his 173 lies within the active site of DDAH-1 and mutation of this amino acid has been demonstrated to result in near complete loss of enzyme catalytic activity

[179,180]. The levels of 4-HNE used in this study represent concentrations of 4-HNE observed

under conditions of inflammation and oxidative stress and suggest that DDAH may play a critical role in mediating the endothelial dysfunction observed in these pathological conditions.

Lane 1 2 3 4 5 6 7

37 kDa

Figure 2-1. hDDAH-1 purification. SDS−PAGE (4−20%) gel electrophoresis and Coomassie stained eluents following column chromatographic separation of E. coli hDDAH-1 expression system. Lane 1, MW ladder; lane 2, crude protein homogenate; lanes −5,3 eluents following passage through His-Trap column and desalting column; lanes −7,6 eluents following passage through ion-exchange column. The eluents from lanes 6 and 7 were used for enzymatic studies.

400

300

200 Km= 68.7 µM Vmax= 356 nmols/mg/min

100 DDAH Activity (nmols/mg/min)

0 0.0 0.2 0.4 0.6 0.8 1.0

ADMA (mM)

Figure 2-2. hDDAH-1 enzyme kinetics for ADMA. Purified hDDAH-1 (10 μg/mL) was incubated in the presence of varying concentrations of ADMA (0.1−1000 μM). hDDAH-1 activity was measured by the conversion of ADMA to citrulline. The Km and Vmax were fitted using the Michaelis−Menton equation. For ADMA, the Km was found to be 68.7 μM, and the Vmax was found to be 356 nmols/mg/min. n = 6.

200

150

100 Km= 53.6 µM

Vmax= 154 nmols/mg/min

50 DDAH Activity (nmols/mg/min)

0 0.0 0.2 0.4 0.6 0.8 1.0

NMMA (mM)

Figure 2-3. hDDAH-1 enzyme kinetics for L-NMMA. Purified hDDAH-1 (10 μg/mL) was incubated in the presence of varying concentrations of L-NMMA (0.1−1000 μM). hDDAH-1 activity was measured by the conversion of ADMA to citrulline. The Km and Vmax were fitted using the Michaelis−Menton equation. For L -NMMA, the Km was found to be 53.6 μM, and the Vmax was found to be 154 nmols/mg/min. n = 6.

100

DDAH Activity (%) 20

0 5.5 6.0 6.5 7.0 7.5 8.0 8.5 9.0 9.5

Figure 2-4. Effects of pH on hDDAH-1 enzyme activity. Purified hDDAH-1 was incubated in the presence of 500 μM ADMA at varying pH (5.5−9.5). 25 mM Bis-Tris and 25 mM Bis-Tris propane was used as the buffering system. hDDAH-1 activity was measured by the conversion of ADMA to citrulline. The results represent % activity compared to pH 8.5. n = 3.

100 ●OH H2O2 ONOO- 80 NO

20 DDAH Activity (% Inhibition)

0 1 10 100 1000 Oxidant Concentration (µM)

Figure 2-5. Effect of oxidants on hDDAH-1 activity. Purified hDDAH-1 was exposed to varying - concentrations (1−1000 μM) of •OH, H2O2, ONOO , and NO. hDDAH-1 activity was measured by the conversion of ADMA to citrulline. n = 3

* 100 *

* 60 *

20 DDAH Activity (% Inhibition)

0 10 50 100 500 1000

4-HNE (µM)

Figure 2-6. Effect of lipid hydroperoxides on hDDAH-1 activity. Increasing concentrations of 4- HNE (10−1000 μM) were found to dose-dependently inhibit the activity of hDDAH- 1. hDDAH-1 activity was measured by the conversion of L-[14C]NMMA to L- [14C]citrulline. The results are presented as % inhibition compared to vehicle-treated hDDAH-1. * indicates significance at p < 0.05. n = 3.

13 b b b b b 20 A T H15 * A V V R y y y y y y Intensity X Intensity X

446.212+ M-H2O

369.312+ y5

444.24 735.33 y4 b6 737.42 175.00 274.19 373.19 y5 y1 y3 y2 537.22 172.97 636.36 b2 419.86 466.24 b4 2+ b3 b5

20 30 40 50 60 70 m/z

Figure 2-7. Modification at H15 on hDDAH-1 after exposure to 4-HNE. MS/MS spectra of a tryptic peptide generating the sequence b/y-ion series from the in-gel digest of the hDDAH-1 reacted with 4-HNE. The peptide observed at m/z 455.2+2 corresponds to the aa sequence 13−20 with H15 modified by 4-HNE with the y1−y 6 ions labled and the b2−b6 ions labeled.

150 b b b b b b b1 b1 b1 b1 b1 b1 b1 b1 b2 b2 175 G A E I L A D T F K D Y A V S T V P V A D G L H173 * L K y2 y1 y1 y1 y1 y1 y y y y y y y

Intensity X 5 Intensity X 10 1105.53 y9

553.56 1392.62 y3 y12 1314.442+ b24 2+ 2+ 1371.23 1395.43 698.32 b25 b13 b13 838.49 918.28 1204.73 y6 b9 y10 723.47 1324.41 y5 b12 1781.60 b17 747.812+ 909.57 1046.38 1494.46 y7 b14 670.26 b14 b10 2 555.28 1166.68 1305.60 b7 + y11 484.22 b6 b5 666.51 1008.54 1491.652 y4 1581.50 1682.58 371.12 771.3 y8 y13 603.052 1161.43 b15 b16 b4 + 4 b11

400 600 800 1000 1200 1400 1600 m/z

Figure 2-8. Modification at H173 on hDDAH-1 after exposure to 4-HNE. MS/MS spectra of a tryptic peptide generating the sequence b/y-ion series from the in-gel digest of the hDDAH-1 reacted with 4-HNE. The peptide observed at m/z 969.9+3 corresponds to the aa sequence 150−175 with H173 modified by 4-HNE with the y3−y13, y15, and y 20 ions labeled along with the corresponding b4−b17 and b24−b25 ions labeled.

CHAPTER 3 REDOX REGULATION OF ADMA METABOLISM: DDAH-2

Introduction

Endothelium-derived nitric oxide (NO) is a critical regulator of vascular function. NO is a potent vasodilator and is responsible for maintaining a healthy vasculature through its antiatherogenic and antiproliferative effects on the vascular wall. Impaired NO production from nitric oxide synthase (NOS) through accumulation of the endogenous NOS inhibitors asymmetric dimethylarginine (ADMA) and NG-methyl-l-arginine (L-NMMA) has been

implicated in the pathogenesis of cardiovascular diseases [2,199]. ADMA and L-NMMA are

generated through the proteolysis of various proteins with methylated arginine residues.

Methylation of arginine residues on proteins is due to the action of a family of proteins known as

protein arginine methyltransferases (PRMTs) [79]. Previous studies have implicated protein

arginine methylation as an important post-translation modification involved in regulation of

DNA transcription, protein function, and cell signaling [81,82]. Following proteolysis of

methylated proteins, free ADMA and L-NMMA are released into the cytosol of cells where they

inhibit NO production through competitive inhibition of NOS. Dimethylarginine

dimethylaminohydrolase (DDAH) metabolizes ADMA and L-NMMA to citrulline and

dimethylamine or monomethylamine respectively [73]. To date, two isoforms of DDAH have

been identified with unique tissue expression patterns. DDAH-1 is predominately expressed in

tissues with neuronal NOS expression such as the liver, kidney and brain [105,106,109,110].

DDAH-2 is predominately expressed in tissues with eNOS expression such as the vascular endothelium and kidney [107,109]. In disease states with endothelial dysfunction, impaired activity and reduced expression of both DDAH isoforms have been noted [64,71-

73,76,78,129,200,201]. It is believed that this impaired DDAH activity/expression is responsible

for accumulation of the endogenous NOS inhibitors ADMA and L-NMMA and subsequent decreased NO production. However, the specific role of each DDAH isoform in regulation of

NO production is currently unclear.

To investigate the individual role of each DDAH isoform, transgenic mice have been developed and siRNA studies have been performed. Overexpression of DDAH-1 in mice has been found to protect against cardiac transplant vasculopathy [72,201]. In addition, DDAH-1 overexpression has been demonstrated to inhibit ADMA-mediated endothelial dysfunction in cerebral arteries and enhance insulin sensitivity through a NO dependent manner [202,203].

Recently, Wang et al. utilized in vivo siRNA techniques to evaluate the role of each DDAH isoform in rats [131]. Silencing of DDAH-1 resulted in a 50% increase in plasma ADMA levels, but this increase was not associated with impaired endothelial dependent relaxation. Wang et al. also found that while in-vivo silencing of DDAH-2 had no effect on plasma ADMA levels, endothelial dependent relaxation was reduced by 40%. These findings suggest that plasma

ADMA levels are not involved in the regulation of endothelial-dependent relaxation, and that decreased levels of DDAH-2 are associated with impaired endothelial-dependent relaxation through a non ADMA-dependent manner. In hyperhomocysteinemia, levels of DDAH-2 are found to be decreased while plasma levels of ADMA are unchanged [204]. Evidence from our laboratory and others suggest that the primary function of DDAH-2 may not involve metabolism of ADMA and that many of the effects attributed to DDAH-2 may occur through a non ADMA- dependent manner. To more clearly define the role of DDAH-2 in methylarginine metabolism and endothelial function, we cloned and purified human recombinant DDAH-2 and characterized its enzymatic activity.

Materials and Methods

DDAH Expression and Purification

A bacterial expression system for human DDAH-2, pDEST17-hDDAH2 in BL21star E. coli, was generated using Gateway technology (Invitrogen). This expression system generates human DDAH-2 fused in frame to an N-terminal His6-tag. After growth and induction by

Isopropyl β-D-1-thiogalactopyranoside (IPTG) in 2 L of terrific broth, the bacterial pellet was resuspended in 160 mL lysis buffer consisting of MAC buffer A (20 mM phosphate, 500 mM

NaCl, 30 mM imidazole, pH7.4), with the addition of EDTA-free complete protease inhibitor

(Roche), 1 mM DTT, 1 mM MgCl, 10 ug DNase, 10 ug lysozyme. The cells were homogenized with two passes through an SLM Aminco French Press (SLM Instruments) at ~14 kpsi, and the insoluble material was removed by centrifugation at 4oC (1 hour at 48,000g). The supernatant was loaded onto a HisPrep FF 16/10 column (GE Bioscience), equilibrated in MAC buffer A.

The column was washed with 30 column volumes of 5% MAC buffer B (20 mM phosphate, 500 mM NaCl, 250 mM imidazole, pH 7.4), and eluted with a linear gradient from 5% MAC buffer

B to100% MAC buffer B over 90 minutes. All chromatography was performed at 4 °C using an

AKTA purifier 10 (GE Biosciences). The fractions containing hDDAH-2 were pooled and concentrated using ultrafiltration.

Immunodetection of hDDAH-2

Following IPTG induction, BL21star E. coli cells were collected by centrifugation and lysed as described above. The cell pellet was then homogenized using RIPA buffer containing sodium orthovanadate (2 mM), phenylmethylsulphonyl fluoride (1 mM), and protease inhibitor cocktail (Santa Cruz biotechnology). Following homogenization, the cell pellet was briefly sonicated 2 x 2 sec. Sample protein concentration was quantified by the Bradford assay. 3x sample buffer containing DTT was added to 40 μg of protein and boiled at 95°C for 3 minutes

and then spun down briefly and cooled for 2 minutes. The samples were then loaded on to a SDS

Tris-Glycine gradient gel 4-20% (Invitrogen) and run at 130V for 2 hours. The gel was then removed and the protein was transferred to a nitrocellulose membrane using the semi-dry transfer blot system (BioRad). Following transfer, the nitrocellulose membrane was blocked for

1 hour in Tris Buffer Saline and 0.05% Tween (TBST) with 7% milk powder. After blocking, the membrane was washed 3x for 5 minutes with TBST and then anti-DDAH-2 rabbit IgG obtained

from Dr. Renke Mass (Hamburg, Germany) was added at a 1:1000 dilution and incubated

overnight at 4°C. Following incubation with the primary antibody the membrane was washed 3x for 15 minutes with TBST and the secondary goat-anti rabbit hrp tag antibody diluted 1:2000 was added. After 1 hour of incubation at room temperature, detection was performed using an enhanced chemiluminescence kit (Amersham Biosciences).

DDAH Radioisotope Activity Assay

DDAH activity was measured from the conversion of L-[14C]NMMA to L-[14C]citrulline.

Purified hDDAH-2 was incubated with the reaction buffer (50 mM Imidazole, 20 μM L-

[14C]NMMA, 200 μM L-NMMA at pH 7.4) in a water bath at 37 °C for 1 h. Following the incubation, the reaction was stopped with 1 mL of ice-cold stop buffer consisting of 20 mM N-2- hydroxyethylpiperazine-N‘-2 ethanesulfonic acid (HEPES) with 2 mM EDTA at pH 5.5.

Separation of L-[14C]citrulline from L-[14C]NMMA was performed using the cation-exchange resin Dowex AG50WX-8 (0.5 mL, Na+ form, Pharmacia). The L-[14C]citrulline in the eluent was then determined using a liquid scintillation counter.

Isolation and Denaturation of Inclusion Bodies

hDDAH-2 overexpressing bacteria were induced and homogenized as described above.

Bacteria were lysed in resuspension buffer (50 mM Tris-HCl, 50 mM NaCl, 1 mM tris(2-

carboxyethyl)phosphine (TCEP), 0.5 mM ethylenediaminetetraacetic acid (EDTA), 5% glycerol,

pH 8.0) with the addition of 20 μL Lysonase per gram of cell pellet. Following cell lysis, Triton

X-100 was added to a final concentration of 1.0% (v/v). Cell homogenates were centrifuged at

8000 × g for 15 min at 10°C and the supernatant was discarded. To wash the isolated inclusion bodies, 10 mL of wash buffer (50 mM Tris-HCl, 50 mM NaCl, 1 mM TCEP, 0.5 mM EDTA, 5% glycerol, and 1% Triton® X-100, pH 8.0) was added per 1 g of cell homogenate and the sample was centrifuged at 8000 × g for 15 min at 10°C. The wash process was repeated twice. Two additional wash steps were performed with resuspension buffer. Denaturation of inclusion was achieved by addition of 10 mL denaturation buffer (50 mM Tris-HCl, 50 mM NaCl, 5 mM

TCEP,0.5 mM EDTA, 5% glycerol, pH 8.0) per 0.5 g inclusion body pellet. Inclusion body

pellets were disrupted by brief sonication and 1.75 mL 30% N-Lauroylsarcosine was added per

0.5 g of inclusion body pellet. The sample was then stirred at room temperature until the

solution became clear. Finally, the sample was centrifuged at 25,000 × g for 15 min at 4°C and

the pellet was discarded to remove any cell debris.

Protein Refolding

Inclusion body samples were dialyzed against 2 L of dialysis buffer (10 mM Tris-HCl,

0.05 mM EDTA, 0.1 mM TCEP, and 0.06% (w/v) 30% N-Lauroylsarcosine, pH 8.0) overnight at

4°C in 3,500 molecular weight cutoff dialysis tubing. Protein concentration of sample was then brought to 1 mg/mL by dilution with dialysis buffer. The sample was then added to the iFold

Protein Refolding System 1 kit (Novagen) protein refolding matrix and the matrix was incubated overnight at room temperature with shaking. The A340 of the refolded protein was measured with a SpectraMax M5 plate reader (Molecular Devices). Subseuquent refolding studies utilized refolding buffer (50 mM Tris-HCl, 500 mM L-Arginine, 12.5 mM methy-β-D-cyclodextrin, pH

7.5) for refolding of hDDAH-2 purified from inclusion bodies.

Cell Culture

Bovine aortic endothelial cells (BAECs) were purchased from Cell Systems and cultured

in Dulbecco's modified Eagle's medium (Sigma) containing 10% FBS, 1% NEAA, 0.2%

endothelial cell growth factor supplement, and 1% antibotic-antimyotic (Invitrogen) and

incubated at 37 °C, 5% CO2, 95% O2.

DDAH-1 and -2 Gene Silencing

21-bp siRNA nucleotide sequences targeting the coding sequences of DDAH-1

(GenBankTM accession number NM_001102201) and DDAH-2 (GenBankTM accession number

NM_001034704) were purchased from Ambion. Control cells received scrambled siRNA also purchased from Ambion. 400 μl of nuclease-free water was added to the dried oligonucleotides to obtain a final concentration of 100 μm. Transfections were performed using the lipid-

mediated transfection reagent RNAiMax (Invitrogen). The procedure was as follows. 240 nm or

5.0 μl of siRNA per well of a 6-well plate was diluted into 250 μl of OptiMEM (Invitrogen), and

5.0 μl of RNAiMax was diluted in 250 μl of Opti-MEM. The siRNA and RNAiMax were then combined into one Eppendorf tube and incubated at room temperature for 20 min. Following the

20-min incubation period, the RNAiMax-siRNA complexes were added to each well of a 6-well plate. The mixture was rocked back and forth to allow for coating of the entire well. BAECs were trypsinized and spun down at 200 × g for 4 min and then resuspended in 1.5 ml of

OptiMEM plus 10% minimum essential medium containing 10% FBS, 1.0% NEAA, 0.2% endothelial cell growth factor. The cells were then added on top of the RNAiMax-siRNA complexes and incubated at 37 °C, 5.0% CO2, 95% O2 for 6 h. After the 6-h incubation period,

1.0 ml of minimum essential medium containing 10% FBS, 1.0% NEAA, 0.2% endothelial cell growth factor was added. 24 h later, 1.0 ml of minimum essential medium containing 10% FBS,

1% NEAA, 0.2% endothelial cell growth factor was added. At 48 h, 2.0 ml of medium was

removed and replaced with fresh minimum essential medium containing 10% FBS, 1% NEAA,

0.2% endothelial cell growth factor, and the transfection was continued for an additional 24 h.

Assessment of mRNA Levels following DDAH Gene Silencing

Following the 72-h siRNA transduction period, BAECs were trypsinized and pelleted at

200 × g for 4 min. The cell pellet was then washed one time with PBS and centrifuged at 200 × g for an additional 4 min. The cell pellet was then homogenized in the lysis buffer. Following lysis, RNA was extracted using a Qiagen (Valencia, CA) RNAeasy minikit. cDNA was then isolated using the Invitrogen One Step reverse transcription-PCR kit. Semiquantitative PCR was performed in order to detect changes in mRNA expression following DDAH-1 or DDAH-2 gene silencing. Bovine primers for DDAH-1 forward (GAGGAAGGAGGCTGACATGA), DDAH-1 reverse (TTCAAGTGCAAAGCATCCAC), DDAH-2 forward

(CTAGCCAAAGCTCAGAGGGACAT), and DDAH-2 reverse

(TCAGTCAACACTGCCATTGCCCT) were purchased from Invitrogen.

EPR Spectroscopy and Spin Trapping

Spin trapping measurements of NO were performed using a Bruker E-scan spectrometer

with Fe2+-MGD (0.5 mM Fe2+, 5.0 mM MGD) as the spin trap [76,143]. For measurements of

NO produced by BAECs, cells were cultured as described above, and spin trapping experiments

were performed on cells grown in 6-well plates. Attached cells were studied because scraping or

enzymatic removal leads to injury and membrane damage with impaired NO generation. The

medium from ~1 × 106 cells attached to the surface of the 6-well plates was removed, and the cells were washed three times in KREBS and incubated at 37 °C, 5% CO2 in 0.2 ml of KRBES buffer containing the spin trap complex Fe2+-MGD, and the cells were stimulated with calcium

ionophore (1.0 μM). Subsequent measurements of NO production were performed following a

30-min incubation period. Spectra recorded from cellular preparations were obtained using the

following parameters: microwave power, 20 milliwatts; modulation amplitude, 3.00 G;

modulation frequency, 86 kHz.

HPLC Method

BAECs were collected from confluent 75-mm culture flasks and sonicated in PBS

followed by extraction using a cation exchange column. Samples were derivatized with O-

phthaldialdehyde and separated on a Supelco LC-DABS column (4.6 × 25 cm inner diameter, 5

μm particle size), and methylarginines were separated and detected using an ESA (Chelmsford,

MA) HPLC system with electrochemical detection at 400 mV. Homoarginine was added to the

homogenate as an internal standard to correct for the efficiency of extraction. The mobile phase

consisted of buffer A (50 mm KH2PO4, pH 7.0) and buffer B (acetonitrile/MeOH, 70:30) run at

room temperature with a flow rate of 1.3 ml/min. The following gradient method was used: 0–

10 min, 90% A; 10–40 min, a linear gradient from 90% A to 30% A [76,144]. eNOS expression

eNOS was detected using an anti-eNOS antibody purchased from Calbiochem (San Diego,

CA). The secondary antibody was goat anti-rabbit IgG-horseradish peroxidas, purchased from

Santa Cruz (Santa Cruz, CA). The secondary antibody was diluted 1:2,000. Western blot detection was performed using an enhanced chemiliumnesece kit purchased from Amersham

Biosciences (Piscataway, NJ).

Results

hDDAH-2 Soluble Fraction Purification

An E. coli bacterial expression system was generated to produce hDDAH-2 protein.

Initial studies were conducted to determine optimum expression levels of hDDAH-2 in the

bacterial expression system. Expression of hDDAH-2, as determined by immunodetection, was

found to be highest following six hours of IPTG induction at 25°C (Figure 3-1). Subsequent

purification studies utilized these conditions. Purification of hDDAH-2 was achieved through

use of nickel metal affinity chromatography (MAC) and fast protein liquid chromatography

(FPLC) techniques. Following elution from the MAC column protein purity was determined by

SDS-PAGE and Coomassie staining. Results demonstrated a ~30 kDa band (lane seven, eight and nine) consistent with hDDAH-2 (Figure 3-2). Removal of lower weight protein contaminates was achieved through concentration of hDDAH-2 by centrifugation with 15 kDA

MWCO filtration columns.

Following purification of hDDAH-2, enzyme activity studies were conducted. For these experiments, the DDAH radioisotope activity assay was used. Results found that IPTG induction of bacterial cells with the hDDAH-2 expression vector did not result in increased

DDAH activity levels in bacterial lysates (Figure 3-3). Purification of hDDAH-2 from the soluble fraction did not result in substantial yields of hDDAH-2 protein. Less then 25 μg of total protein was obtained from 1.0 L of bacterial culture. Expression of proteins in bacterial cellular systems can result in the formation of inclusion bodies, aggregates of the expressed protein that can not be purified from the soluble fraction of cells. Therefore, subsequent studies were conducted to determine if hDDAH-2 forms inclusion bodies when expressed in E. coli.. hDDAH-2 Purification from Inclusion Bodies

Inclusion bodies from hDDAH-2 expressing bacterial cells were isolated and Coomassie staining was used to determine the proportion of hDDAH-2 protein in the inclusion bodies compared to other proteins. hDDAH-2 was found to be the predominant protein in the inclusion bodies (Figure 3-4) indicating that significant levels of hDDAH-2 form aggregates when expressed in a bacterial system. It is likely that the low purification yields from the soluble fraction of hDDAH-2 expressing cells is due to the formation of inclusion bodies. Purification of

proteins from inclusion bodies requires the protein aggregates to be denatured and then refolded

by incubation with a refolding buffer.

To determine the optimum composition of the refolding buffer, the Novagen iFold

Protein Refolding System 1 kit was used. This assay tests for the ability of proteins to be properly refolded as measured by protein precipitation at A340. Isolated inclusion bodies from

cells expressing hDDAH-2 were exposed to 92 different refolding buffers (Table 3-1). Protein

refolding efficiency was determined by measurement of A340 values (Table 3-2). Several candidate refolding buffers were identified and 50 mM Tris-HCl, 500 mM L-Arginine, 12.5 mM methy-β-D-cyclodextrin, pH 7.5 was selected as the refolding buffer to be used in subsequent experiments based on A340 values. This buffer contains arginine which has been previously shown to assist in protein refolding and we hypothesized that the similarity in structure of arginine to ADMA may indicate that arginine would be able to successfully refold hDDAH-2.

Following protein refolding, hDDAH-2 was purified by MAC and purity was determined by SDS-PAGE and Coomassie staining. Results showed a band of ~30 kDa (Lane 4 and 5) consistent with hDDAH-2 and ~ 90% purity (Figure 3-5). Samples were pooled and concentrated by centrifugation. The activity of hDDAH-2 purified from inclusion bodies was determined by the DDAH radioisotope activity assay (Figure 3-6). The activity of purified hDDAH-2 was found to be no different than vehicle control samples. It is likely that the primary function of DDAH-2 is not metabolism of ADMA.

In vivo role of DDAH-2 on NO production

In order to test this hypothesis, we evaluated the effects of siRNA mediated DDAH silencing on endothelial NO production. BAEC’s were cultured in 6-well plates and transfected with siRNA against DDAH-1 (240 nM), DDAH-2 (240 nM), or both. The siRNA concentrations were chosen based on titer optimization, which demonstrated a robust reduction in DDAH

expression at these concentrations (Figure 3-7). At 48 hours post-transfection, EPR studies were

performed to measure NO production using the spin-trapping complex Fe-MGD. Results

demonstrated that DDAH-1 siRNA reduced calcium ionophore (1.0 µM) evoked NO production

by 27% (Figure 3-8) while DDAH-2 siRNA inhibited NO production by 57% (Figure 3-9). Dual silencing of both DDAH-1 and DDAH-2 did not produce an additive effect as inhibition was

55% (Figure 3-10). These experiments were then repeated in the presence of exogenous L-arg

(100 µM). Because ADMA is a competitive inhibitor of eNOS, L-arg supplementation should reverse the inhibition associated with increased ADMA. L-arginine did partially restore NO production in the DDAH-1 (Figure 3-8) silenced cells but had no effect on the DDAH-2 group

(Figure 3-9). The effects on NO observed with DDAH silencing were independent of changes in eNOS expression or phosphorylation as neither was altered with silencing (Figure 3-11). These results suggest that DDAH-1 effects are mediated, in part, through ADMA accumulation while

DDAH-2 effects are independent of ADMA mediated inhibition of eNOS. Subsequent experiments were performed using HPLC analysis to measure the intracellular methylarginine levels following DDAH-1 and DDAH-2 silencing. Results demonstrated that DDAH-1 silencing increased total ADMA by two-fold while DDAH-2 silencing had no effect (Figure 3-12). The observed increases in ADMA would not be expected to elicit the degree of NOS inhibition observed in the NO studies and suggests that DDAH-1 may be modulating NOS activity through both ADMA-dependent and independent mechanisms as well. Moreover, these results clearly indicate that DDAH is eliciting its eNOS inhibitory effects through ADMA-indpendent pathways.

Discussion

Plasma levels of ADMA have been shown to be elevated in a variety of disease states including hypertension, diabetes and cardiovascular disease [64,76]. The Coronary Artery Risk

Determination Investigating the Influence of ADMA Concentration (CARDIAC) study

demonstrated that ADMA is an independent risk factor for coronary artery disease and that

ADMA is a predictor of cardiovascular mortality in these patients [77]. Plasma ADMA levels

have been found to increase from ~0.4 μM to ~0.8 μM in patients with pathological conditions

such as pulmonary hypertension, coronary artery disease, diabetes, and hypertension [65,77,142-

145]. ADMA causes pathological progression of these disease states through inhibition of NO

production from NOS. Intracellular levels of ADMA regulate NO production through

competitive inhibition of NOS. The intracellular accumulation of ADMA is demonstrated in

restenotic lesions and ischemia reperfused myocardium. In these conditions, intracellular

ADMA levels have been found to be increased 3-4-fold [71,205]. The principal route of ADMA metabolism is by the enzyme DDAH. Over 80% of ADMA clearance is believed to be conducted by DDAH, but the individual role of each DDAH isoform is unclear [64].

To determine the role of each DDAH isoform enzyme kinetic studies have been conducted. We have demonstrated that hDDAH-1 has Km values of 68.7 and 53.6 μM and Vmax

values of 356 and 154 nmol/mg/min for ADMA and L-NMMA, respectively [206]. In enzyme kinetic studies of purified recombinant hDDAH-2, ADMA metabolizing properties of the enzyme were unable to be detected. Potential explanations for the lack of detection of ADMA metabolism by hDDAH-2 are: (1) the rate of ADMA metabolism by hDDAH-2 is drastically lower than the rate of ADMA metabolism by hDDAH-1; (2) DDAH-2 requires additional cofactors for activity; or (3) hDDAH-2 has no catalytic activity towards ADMA. The DDAH radioisotope activity assay used in this study is considered the “gold standard” for DDAH activity and is capable of detecting enzymatic rates as low as 10 nmols/mg/min for ADMA.

Based on the sensitivity of this assay, we can assume that any enzymatic activity exhibited by

DDAH-2 is at least 100 times less than DDAH-1. Moreover, studies were also carried out using

DDAH-2 concentrations in excess of 10 times those used for DDAH-1 studies. Given that no enzymatic activity was detected under any conditions tested, we can conclude that if DDAH-2 is infact an arginine demethylase its activity is at least 100 fold lower than that of DDAH-1. With regards to the requirement of additional cofactors or protein-protein interactions, we carried out studies with purified enzyme in the presence of endothelial cell homogenates. The rationale for this approach was that the endothelial homogenate would provide the necessary cofactors or accessory proteins needed for catalytic activity. Results demonstrated no difference in DDAH activity between the endothelial homogenate and the homogenate with purified recombinant

DDAH-2. Alternatively, in the literature there have been several recent reports that indicate

DDAH-1 and DDAH-2 have significantly different actions.

Wang et al. demonstrated different effects of the two DDAH isoforms through in vivo siRNA silencing techniques [131]. Silencing of DDAH-1 was found to increase plasma levels of

ADMA by 50%, but endothelium-dependent relaxation was not changed. Silencing of DDAH-2 was found to have no effect on plasma ADMA levels, but endothelium-dependent relaxation was reduced by 40%. These results suggest the DDAH-2 is able to regulate NO production through an ADMA independent mechanism. Studies from our laboratory have demonstrated similar differences in the function of the two DDAH isoforms by gene silencing in endothelial cells

[132]. Silencing of DDAH-1 resulted in a 27% decrease in NO bioavailability and a 48% reduction in the L-Arg/ADMA ratio. Silencing of DDAH-2 resulted in a 57% reduction in NO

bioavailbility, however the L-Arg/ADMA ratio was unchanged. These results provide additional

evidence that DDAH-2 regulates NO production in an ADMA independent mechanism. In this

study overexpression of the two DDAH isoforms in endothelial cells by adenovirus infection was

also investigated. Overexpression of DDAH-1 resulted in an almost two-fold increase in DDAH activity. However, overexpression of DDAH-2 did not result in an increase in DDAH activity levels despite an increase in DDAH-2 protein levels. Overexpression of both isoforms resulted in increased NO production, but decreased intracellular levels of ADMA were only noted in cells overexpressing DDAH-1 and not overexpressing DDAH-2. Perhaps the most convincing piece of evidence of the different functions of the two DDAH isoforms lays in mouse transgenic gene deletion studies [137]. DDAH-1 -/- mice do not progress through embryonic development, while

DDAH-2 -/- mice have a normal lifespan. Both isofroms were detected in the developing limb buds of mice in areas with high NOS activity, suggesting that DDAH-1 and DDAH-2 are involved in embryonic development through actions on NO regulation.

The lack of detection of ADMA metabolism by hDDAH-2 was likely due to the fact that the primary role of DDAH-2 is not metabolism of ADMA. Currently it is unclear what the primary function of DDAH-2 is, but our evidence suggests that DDAH-2 is involved in regulation of NO production through an ADMA-independent manner. Future studies are required to determine the precise molecular mechanisms through which DDAH-2 regulates NO production.

6 hr 12 hr

40 kDa

30 kDa

20 kDa

IPTG - + - +

Figure 3-1. hDDAH-2 expression. Levels of hDDAH-2 expression in the E. coli bacterial expression system were measured by immunodetection at 6 or 12 hours post IPTG induction.

1 2 3 4 5 6 7 8 9

37 kDA

25 kDA

Figure 3-2. hDDAH-2 purification from soluble fraction. SDS-PAGE (4-20%) gel electrophoresis and Coomassie stained eluents following chromatographic separation. Lane 1: MW ladder. Lanes 3-6: eluents following passage through MAC column. The eluents from lanes 7-9 were used for enzymatic studies after removal of low MW bands by filtration.

1000

800

600

400 DDAH Activity

200

0 C IPTG

Figure 3-3. DDAH activity of soluble fraction. DDAH activity of total cell homogenates from IPTG induced or vehicle treated E. coli cells with hDDAH-2 vector was measured by the conversion of L-[14C]NMMA to L-[14C]citrulline. Results represent mean ± SD. n = 3.

Lanes 1 2

37 kDA

25 kDA

Figure 3-4. Protein profile of isolated inclusion bodies. SDS-PAGE (4-20%) gel electrophoresis and Coomassie stained samples from isolated inclusion bodies of IPTG induced E. coli hDDAH-2 bacterial expression system cells. Lane 1: MW ladder. Lane 2: inclusion body sample.

Lanes 1 2 3 4 5

37 kDA

25 kDA

Figure 3-5. hDDAH-2 purification from inclusion bodies. SDS-PAGE (4-20%) gel electrophoresis and Coomassie stained eluents following chromatographic separation. Lane 1: MW ladder. Lanes 2-3: eluents following passage through MAC column. The eluents from lanes 4-5 were used for enzymatic studies.

5000

4000

3000

2000 DDAH Activity

1000

0 C DDAH-2

Figure 3-6. DDAH activity of refolded hDDAH-2 from isolated inclusion bodies. DDAH activity of purified hDDAH-2 enzyme or vehicle control was measured by the conversion of L-[14C]NMMA to L-[14C]citrulline. Results represent mean ± SD. n = 3.

siDDAH1 siDDAH2

60 nM control control 60 nM control 240 nM 240 nM

DDAH-2 DDAH-1

Figure 3-7. Effects of DDAH gene silencing on DDAH mRNA expression. DDAH mRNA expression was measured by semi quantitative PCR, and run on an agarose gel to determine differences in DDAH expression following siRNA treatment. Experimental groups consist of 60 nM siRNA (DDAH-1, DDAH-2) and 240 nM siRNA (DDAH-1 and DDAH-2)

4 *

µ 3 *

NO Generation ( Generation NO 1

1 - Arg Arg - - Control Control 1 + L siDDAH - Control + L + Control siDDAH

Figure 3-8. Effects of DDAH-1 gene silencing on endothelial cell NO production. NO generation from calcium ionophore A23187 (1.0 μM)-stimulated BAECs (1 × 106) was measured by EPR spin trapping with the Fe2+-MGD complex. Experimental groups consisted of scrambled siRNA (control) and siDDAH-1 (240 nM). These experiments were performed both in the presence and absence of L-arginine (100 μM). Results are means ± S.D. * indicates significance at p < 0.05 as compared with the respective control. n = 3.

M) 3 µ

2 * *

NO Generation ( 1

2 - Arg Arg - - Control Control 2 + L - siDDAH Control + L Control siDDAH

Figure 3-9. Effects of DDAH-2 gene silencing on endothelial cell NO production. NO generation from calcium ionophore A23187 (1.0 μM)-stimulated BAECs (1 × 106) was measured by EPR spin trapping with the Fe2+-MGD complex. Experimental groups consisted of scrambled siRNA (control) and siDDAH-2 (240 nM). These experiments were performed both in the presence and absence of L-arginine (100 μM). Results are means ± S.D. * indicates significance at p < 0.05 as compared with the respective control. n = 3.

µ 3 *

2 *

NO Generation ( 1

2 1 - Arg Arg - - - Control Control 2 + L - siDDAH Control + L Control siDDAH1

Figure 3-10. Effects of dual DDAH gene silencing on endothelial cell NO production. NO generation from calcium ionophore A23187 (1.0 μM)-stimulated BAECs (1 × 106) was measured by EPR spin trapping with the Fe2+-MGD complex. Experimental groups consisted of scrambled siRNA (control) and siDDAH-1/siDDAH-2 dual gene silencing. These experiments were performed both in the presence and absence of L- arginine (100 μM). Experimental groups consisted of scrambled siRNA (control) and siDDAH-2. Results are means ± S.D. * indicates significance at p < 0.05 as compared with the respective control. n = 3.

1 2 1/2 - - -

control siDDAH siDDAH siDDAH eNOS

p-eNOS (S-1179)

GAPDH

Figure 3-11. Effects of DDAH gene silencing on endothelial cell eNOS phosphorylation. eNOS phosphorylation at Serine-1179 from BAECs (1 × 106) was measured by immunodetection. Experimental groups consisted of scrambled siRNA (control), siDDAH-1 (240 nM), siDDAH-2 (240 nM), and siDDAH-1/siDDAH-2 (240 nM each) dual gene silencing.

ADMA 10 (pmoles/mg protein) 5

0 control siDDAH1 siDDAH2

Figure 3-12. Effects of DDAH gene silencing on endothelial cell intracellular ADMA concentrations. Intracellular ADMA concentrations from BAECs (1 × 106) were measured by HPLC. Experimental groups consisted of scrambled siRNA (control), siDDAH-1 (240 nM) and siDDAH-2 (240 nM). Results are means ± S.D. * indicates significance at p < 0.05. n = 3.

Table 3-1. iFOLD Protein Refolding System 1, plate layout. All plate wells contain 450 μl refolding buffer, with exception of wells F12 and H12.

Table 3-2. Solubility of refolded hDDAH-2 inclusion bodies. A340 values were used to determine precipitation of protein from refolded hDDAH-2 samples. Lower A340 values correspond to less protein precipitation and higher protein solubility. 1 2 3 4 5 6 7 8 9 10 11 12 A 0.291 0.329 0.059 0.051 0.351 0.055 0.333 0.294 0.338 0.247 0.046 0.318 B 0.335 0.303 .0347 0.046 0.334 0.038 0.034 0.049 0.043 0.091 0.360 0.357 C 0.365 0.055 0.377 0.378 0.056 0.369 0.083 0.087 0.052 0.323 0.361 0.039 D 0.297 0.058 0.352 0.044 0.363 0.038 0.034 0.367 0.267 0.046 0.366 0.352 E 0.306 0.063 0.394 0.059 0.047 0.363 0.052 0.302 0.050 0.036 0.373 0.047 F 0.341 0.044 0.366 0.043 0.363 0.354 0.040 0.302 0.387 0.041 0.366 0.041 G 0.360 0.051 0.296 0.089 0.060 0.383 0.055 0.281 0.304 0.052 0.368 0.036 H 0.334 0.045 0.352 0.048 0.042 0.296 0.040 0.350 0.301 0.049 0.344 0.044

CHAPTER 4 REDOX REGULATION OF ADMA SYNTHSIS

Introduction

Activity of nitric oxide synthase (NOS), the enzyme responsible for production of nitric oxide (NO) has been shown to be regulated through several different mechanisms. These include protein-protein interactions, phosphorylation and the endogenous methylarginines asymmetric dimethylarginine (ADMA) and L-NG-monomethylarginine (L-NMMA). Free

ADMA and L-NMMA have been demonstrated to be competitive inhibitors of NOS and their

levels are elevated in cardiovascular disease. Recently, interest into the role of protein arginine

methylation in regulation of endothelial function has grown as the protein arginine

methyltransferases (PRMTs) have been demonstrated to control a wide range of cellular

functions. Thus, it is possible PRMTs play a critical role in regulation of endothelial function through synthesis of ADMA and L-NMMA and through protein arginine methylation.

Synthesis of ADMA and L-NMMA is controlled by PRMTs [79]. There are two subgroups of PRMTs; type I and type II. Type I PRMTs (PRMT-1, -3, -4, -6, and -8) methylate

arginine residues on proteins to from L-NMMA then in a subsequent step another methyl group

is added to from ADMA. The function of type II PRMTs (PRMT-5, -7, and FBXO11) is the same as type I PRMTs except in the second step symmetric dimethylarginine (SDMA) is formed instead of ADMA [81,82]. SDMA has no known biologically activity while ADMA and L-

NMMA are competitive inhibitors of NOS. In the PRMT enzymatic reaction, a methyl group is donated by S-adenosylmethionine (SAM) and is transferred to the guanidino nitrogen of arginine

[83]. Following methylation of arginine residues on proteins by PRMT and proteolysis of these proteins free ADMA and L-NMMA are released into the cytosol where they can inhibit NOS activity.

Like other post-translational modifications arginine methylation has been demonstrated to have an important regulatory influence on a number of cellular processes. Protein-arginine methylation has been shown to modulate transcription, RNA metabolism and protein–protein interactions leading to control of cellular differentiation, proliferation, survival, and apoptosis

[81]. The importance of PRMT activity is demonstrated by the PRMT-1 knock-out mouse.

PRMT-1 -/- mice are unable to develop past embryonic day 6.5, due to impaired cellular metabolism [90].

In cardiovascular disease states expression of PRMT has been found to be increased. In diabetic rat retina protein levels of PRMT-1 were found to be greater than in non-diabetic retinas

[95]. Chen et al. also demonstrated increased expression of PRMT-1 in bovine retinal capillary endothelial cells after exposure to high levels of glucose. Hypertension induced by infusion of angiotensin II in rats resulted in increased expression of PRMT-1 in the lungs [207]. In addition, exposure of human umbilical vein endothelial cells to fluid shear stress resulted in an increase in

PRMT-1 mRNA expression levels via activation of the nuclear factor (NF)-kappaB pathway

[96]. Cardiovascular disease states such as diabetes are characterized by increased production of reactive oxygen species (ROS) and reactive nitrogen species (RNS). ROS and RNS have been previously demonstrated to alter protein enzymatic activity through post-translational redox reactions. The aim of this study is determine if ROS or RNS can regulate the enzymatic activity of PRMT-1.

Materials and Methods

Cell Culture

Bovine aortic endothelial cells (BAECs) and COS7 cells were purchased from Cell

Systems and cultured in Dulbecco's modified Eagle's medium (Sigma) containing 10% FBS, 1%

NEAA, 0.2% endothelial cell growth factor supplement, and 1% antibotic-antimyotic

(Invitrogen) and incubated at 37 °C, 5% CO2, 95% O2. COS7 cells were transfected with eNOS plasmid. Cells were treated with 10 μM adenosine dialdehyde (Adox, Sigma) or 1 mM mineral trioxide aggregate (MTA, Sigma) for 48 hours.

EPR Spectroscopy and Spin Trapping

Spin trapping measurements of NO were performed using a Bruker E-scan spectrometer with Fe2+-MGD (0.5 mM Fe2+, 5.0 mM MGD) as the spin trap [76,143]. For measurements of

NO produced by BAECs, cells were cultured as described above, and spin trapping experiments

were performed on cells grown in 6-well plates. Attached cells were studied because scraping or

enzymatic removal leads to injury and membrane damage with impaired NO generation. The

ionophore (1.0 μM). Subsequent measurements of NO production were performed following a

30-min incubation period. Spectra recorded from cellular preparations were obtained using the

following parameters: microwave power, 20 milliwatts; modulation amplitude, 3.00 G;

modulation frequency, 86 kHz.

PRMT Radioisotope Activity Assay

14 14 PRMT activity was measured by the incorporation of [ C]CH3 from [ C]SAM into

unmethylated histone proteins. PRMT-1 activity was conducted as follows: 1.0 μg rat PRMT-1

(Invitrogen), 20 μg unmethylated histone (Sigma), 50 μM [14C]SAM (GE Biosciences), in PBS for 1 hour at 30°C. Following incubation, the samples were run on SDS-PAGE (4-20%

Invitrogen) and the gel was stained with Coomassie. Histone bands were identified by molecular

weight and excised from the gel. Bands were homogenized in 0.1% SDS, 250 mM Tris, pH 7.7

and histone protein was eluted by centrifugation of sample using a Nanosep MF 0.2 μM

centrifugal device (Pall). Eluted samples were then collected and PRMT-1 activity was determined using a liquid scintillation counter. PRMT activity after exposure of PRMT-1 to

ROS or RNS was conducted as follows. Varying concentrations (1.0 nM-1.0 mM) of H2O2, and

ONOO- or (10 pM-10 μM) NO were incubated with 1.0 μg PRMT-1 for 10 minutes at room temperature. For H2O2, the authentic molecule was used and 100 μM diethylene triamine pentaacetic acid (DTPA) was added to prevent Fenton reactions. For ONOO- the authentic molecule was used and for NO the NO donor DETANONOate was used.

Immunodetection of Nitro-Tyrosine

PRMT-1 (1.0 μg) was exposed to varying concentrations (1.0 nM-1.0 mM) of ONOO- for

10 minutes at room temperature. 1x sample buffer containing DTT was added to the samples

and the samples were boiled at 95°C for 3 minutes and then spun down briefly and cooled for 2

minutes. The samples were then loaded on to a SDS Tris-Glycine gradient gel 4-20%

(Invitrogen) and run at 130V for 2 hours. The gel was then removed and the protein was

transferred to a nitrocellulose membrane using the semi-dry transfer blot system (BioRad).

Following transfer, the nitrocellulose membrane was blocked for 1 hour in Tris Buffer Saline and

0.05% Tween (TBST) with 7% milk powder. After blocking, the membrane was washed 3x for 5 minutes with TBST and then anti-Nitro-Tyrosine primary antibody (Upstate) was added at a

1:250 dilution and incubated overnight at 4°C. Following incubation with the primary antibody the membrane was washed 3x for 15 minutes with TBST and the secondary goat-anti mouse hrp tag antibody diluted 1:5000 was added. After 1 hour of incubation at room temperature, detection was preformed using an enhanced chemiluminescence kit (Amersham Biosciences).

Biotin Switch

The biotin switch assay was performed as described previously by Jaffrey and Snyder

using low-light conditions and opaque tubes [208]. Free thiols were blocked by methylation

with methyl methanethiosulfonate (Sigma). Unreacted methyl methanethiosulfonate were

removed by centrifugation in Micro Bio-Spin 6 Columns (Bio-Rad). Cysteine residues that are

S-nitrosylated by NO were converted to free thiols with sodium ascorbate (1 mM final

concentration), which does not alter the methylated thiols. The free thiols were then biotinylated with biotin-hexyl pyridyldithiopropionamide (HPDP) at 25°C for 1 h. Thus, the S-nitrosylated cysteines were switched for biotin. Due to the low protein concentrations of PRMT-1 it was necessary to perform an immuno-precipitation (IP) with anti-PRMT-1 antibody. Following IP of

PRMT-1, 1x sample loading buffer was added and the sample was loaded on to a SDS Tris-

Glycine gradient gel 4-20% (Invitrogen) and run at 130V for 2 hours. The gel was then removed and the protein was transferred to a nitrocellulose membrane using the semi dry transfer blot system (BioRad). Following transfer, the nitrocellulose membrane was blocked for 1 hour in

Tris Buffer Saline and 0.05% Tween (TBST) with 7% milk powder. After blocking, the membrane was washed 3x for 5 minutes with TBST and then anti-Biotin primary antibody

(Upstate) was added at a 1:500 dilution and incubated overnight at 4°C. Following incubation with the primary antibody the membrane was washed 3x for 15 minutes with TBST and the secondary goat-anti mouse hrp tag antibody diluted 1:5000 was added. After 1 hour of incubation at room temperature, detection was preformed using an enhanced chemiluminescence kit (Amersham Biosciences). Following exposure the nitrocellulose membrane was stripped with glycine stripping buffer (0.2M Glycine, 0.05% Tween-20, pH 2.5) 2x for 30 minutes. The membrane was then reprobed with anti-PRMT-1 primary antibody at a 1:1000 dilution overnight at 4°C.

Results

Effect of PRMT Inhibitioin on Endothelial NO Production

We undertook a series of studies to determine whether protein-arginine methylation was

involved in the regulation of endothelial NO production. BAEC’s were treated with the PRMT

inhibitor Adox (10 µM) for 48 hours and NO production measured by EPR. Results demonstrate

that inhibition of PRMT with Adox increased NO production by 29% (Figure 4-1) suggesting that PRMT does in fact regulate endothelial NOS. In further support of this observation, studies were repeated using eNOS transfected COS7 cells treated with the PRMT inhibitor MTA (1.0

mM) for 48 hours. In this model, PRMT inhibition resulted in a near two-fold increase in endothelial NO production (Figure 4-1). Western blot analysis of eNOS expression showed no significant difference with PRMT inhibition (data not shown). These studies suggest that eNOS may be a target of PRMT methylation and that this modification results in impaired enzyme activity. Given that increased PRMT expression and protein methylation have been observed in models of atherosclerosis, this protein modification may represents a novel mechanism for endothelial dysfunction.

Effect of H2O2 on PRMT-1 Activity

Levels of ROS have been demonstrated to increase in cardiovascular disease states.

Increased levels of ROS can alter the function of proteins through post-translational modifications. PRMT-1 is responsible for the synthesis of the endogenous NOS inhibitors

ADMA and L-NMMA and altered PRMT-1 activity could have important implications on

regulation of NO production. Initial experiments were conducted to determine if the ROS, H2O2,

can alter PRMT-1 activity. PRMT-1 (1.0 μg) was incubated with varying concentrations of

H2O2 (1.0 nM-1.0 mM) for 10 minutes before activity was determined by the incorporation of

14 14 [ C]CH3 from [ C]SAM into unmethylated histone residues. Results demonstrated that

exposure of PRMT-1 to H2O2 had no significant effect on PRMT-1 activity (Figure 4-2)

suggesting that this enzyme is not regulated by ROS. Nevertheless, many enzymes contain

regulatory cysteines or tyrosines which are susceptible to modification by RNS. Therefore, we

next investigated the role of RNS on the activity of PRMT-1.

Effect of ONOO- on PRMT-1 Activity

As with ROS, levels of RNS increase in cardiovascular disease states and can have important regulatory actions on proteins. ONOO- is a highly reactive nucleophile and can alter protein function by formation of nitro-tyrosine residues on proteins. To investigate the role of

RNS on PRMT-1 activity, PRMT-1 (1.0 μg) was incubated with varying concentrations of

ONOO- (1.0 nM-1.0 mM) for 10 minutes and activity was then determined by the incorporation

14 14 of [ C]CH3 from [ C]SAM into unmethylated histone residues. Results demonstrated that at

1.0 mM ONOO-, PRMT-1 activity was almost completely inhibited (Figure 4-3). However,

physiological levels of ONOO- are in the low micromolar range and it is unlikely that even uynder pathological conditions that this level of RNS formation could be achieved. In the physiologically relevant range of ONOO- there is a non-statistically significant trend toward increased PRMT-1 activity. Based on these results it does not appear that physiologically relevant levels on ONOO- can elicit a significant regulatory effect on PRMT-1 activity.

Nevertheless, additional studies were carried out to determine the post-translational modification that is responsible for inhibition of PRMT-1 activity at 1.0 mM ONOO-. Samples were prepared

for detection of nitro-tyrosine residues by western blot. Exposure of PRMT-1 to ONOO- was

found to dose-dependently increase formation of nitro-tyrosine residues on PRMT-1 (Figure 4-

4). The inability of nitro-tyrosine residues to inhibit PRMT-1 activity at low concentration of

ONOO- is likely due to the location of the nitro-tyrosine formation. At low concentrations of

ONOO- the tyrosine residues being modified are likely not important to the catalytics activity of

the enzyme. At high concentrations of ONOO- (1.0 mM) the number of tyrosine residues being modified is greatly increased and tyrosine residues important to substrate binding are possibly being altered resulting in inhibitioni of PRMT-1 activity.

Effect of NO on PRMT-1 Activity

NO is an important signaling molecule in the vasculature and has previously been demonstrated to alter the activity of enzymes through S-nitrosylatoin of critical cysteine residues on proteins. The effect of NO on PRMT-1 activity was determined by the incorporation of

14 14 [ C]CH3 from [ C]SAM into unmethylated histone residues after incubation of PRMT-1 (1.0

μg) with varying concentrations of the NO donor DEANONOATE (10 pM-10 µM). Results found that PRMT-1 activity was increased approximately three-fold after exposure to 10 µM NO

(Figure 4-5). In addition, at 1.0 μM, 10 μM and 1.0 nM NO, PRMT-1 activity was increased approximately two-fold. Increased PRMT-1 activity at 1.0 μM NO is of particular interest

because this concentration is within the range of NO levels that are physiologically relevant.

This regulation of PRMT-1 activity by NO presents a possible negative feedback mechanism

where increased production of NO could result in increased PRMT-1 activity and increased

production of ADMA and L-NMMA. ADMA and L-NMMA would then inhibit NOS activity

resulting in decreased production of NO.

NO has been demonstrated to post-translationally modify proteins by S-nitrosylation of

cysteine residues. The biotin switch method was used to detect for S-nitrosylation of PRMT-1 after exposure to NO. Immunodetection for biotin resulted in the appearance of a new band at

~45 kDA after exposure of PRMT-1 to 1.0 mM NO (Figure 4-6). This is consistent with the molecular weight for PRMT-1 and indicates that NO is able to regulate PRMT-1 activity through

S-nitrosylation.

Discussion

The gaseous molecule NO is critical for maintenance of a healthy vascular system [181-

183]. NO is generated by the oxidation of L-Arginine by eNOS [184,185]. Dysregulation of NO

production has been noted in the pathogenesis of multiple cardiovascular disease states such as

diabetes, heart failure and pulmonary hypertension. The endogenous methylarginines ADMA

and L-NMMA inhibit NO production through competitive inhibition of eNOS activity. Levels of

ADMA and L-NMMA are elevated in cardiovascular disease states and may be responsible for

the impaired NO production found in these conditions [65,77,142-145]. PRMTs are responsible

for the synthesis of ADMA and L-NMMA. Arginine residues on proteins are methylated by

PRMTs and subsequent proteolysis of these proteins results in the release of free ADMA and L-

NMMA into the cytosol of cells where it can inhibit NOS activity. One possible mechanism for

the increased levels of ADMA and L-NMMA found in cardiovascular disease states is increased

activity of PRMT. In the diabetic rat retina protein expression of PRMT-1 is elevated [95]. In

addition, levels of PRMT-1 are increased in the lungs of rats with angiotensin II induced

hypertension [207]. These findings suggest that PRMT-1 may be, at least in part, responsible for

elevated levels of ADMA and L-NMMA found in cardiovascular disease states.

Increased levels of ROS and RNS accompany cardiovascular disease states and have

been demonstrated to alter enzyme function. In this regard we analyzed the effect of ROS and

RNS on the activity of PRMT-1. First, the ROS H2O2 was tested and found to have no

significant effect on PRMT-1 activity even at supraphysiological levels. Suggesting that PRMT-

1 is highly resistant to altered activity by ROS. It is likely that increased levels of ROS found in

cardiovascular disease states are not responsible for increased accumulation of ADMA and L-

NMMA by increasing the activity of PRMT-1. However, it is possible that ROS can mediate increased ADMA and L-NMMA production by increasing the expression of PRMT-1. In

support, levels of ROS and protein expression of PRMT-1 are found to be elevated in diabetes

and angiotensin II induced hypertension [95,207].

As ROS were not found to affect the activity of PRMT-1, RNS were tested next. The

RNS ONOO- was found to inhibit PRMT-1 activity almost completely at supraphysiological

levels of 1.0 mM through formation of nitro-tyrosine residues. It is interesting to note that

formation of nitro-tyrosine residues on PRMT-1 was detected at physiological levels of ONOO-.

Therefore, PRMT-1 likely has a good deal of protection from inhibition by ONOO- at

physiological levels of ONOO-. In disease states with increased levels of ONOO-, such as

cardiovascular disease, production of ADMA and L-NMMA by PRMT-1 activity is not

attenuated by ONOO-. At physiological levels of ONOO- there is a non-statistically significant increase in PRMT-1 activity. Suggesting that in concert with other modulators of PRMT-1 activity ONOO- may be involved in increased activity of PRMT-1 and increased production of

ADMA and L-NMMA.

To determine if other RNS can affect PRMT-1 activity studies were conducted to determine the effect of NO on PRMT-1 activity. NO was found to increase PRMT-1 activity by

greater than three-fold at 10 µM NO. At concentrations of 1.0 μM, 100 nM and 1.0 nM NO

PRMT-1 activity was increased approximately two-fold. These findings are of particular interest

because the concentration of NO are within the physiological range. The biotin switch method

showed that exposure of PRMT-1 to NO results in S-nitrosylation of PRMT-1. Increased activity of PRMT-1 by NO presents a possible negative feedback mechanism by which increased production of NO would result in increased activity of PRMT-1 and therefore increased production of the NOS inhibitors ADMA and L-NMMA. In certain disease states, such as sepsis, production of NO is greatly increased and can result in hypotension. Increased activity of

PRMT-1 by NO would help regulate NO production by inhibiting further NO production by

accumulation of ADMA and L-NMMA.

NO has been previously demonstrated to regulate ADMA metabolism through a negative feedback mechanism. Lieper et al. showed NO inhibits DDAH-1 activity and impairs ADMA metabolism through S-nitrosylation of a Cysteine residue in the catalytic triad of DDAH-1 [113].

Impaired DDAH-1 activity results in increased levels of ADMA and inhibition of NO production from NOS. This finding is consistent with our results demonstrating that increased NO production can lead to increased formation of ADMA through elevated PRMT-1 activity. Both of these mechanisms result in an increase in levels of ADMA and inhibition of NOS, resulting in decreased NO production.

In addition to synthesis of ADMA, PRMT has important actions in regulation of transcription and cell cycle progression [209]. One of the pathologies associated with cardiovascular disease is proliferation of smooth muscle cells in the vascular wall leading to an increased intimal/media ratio and narrowing of the vessel. One the vascular protective mechanisms of NO is inhibition of smooth muscle cell proliferation by down-regulation of CDK-

2 protein activity and cyclin-A gene transcription [16]. It is possible that NO could exert anti- proliferative properties through regulation of PRMT activity. As NO was found to alter PRMT-1 activity in the physiological range of NO levels it is possible that NO could regulate gene transcription through PRMT. Further studies are required to determine if NO can regulate transcription of cellular growth genes through PRMT.

120 * *

40 O Production (arb. Units) N 0 Control Adox Control MTA

BAEC COS7

Figure 4-1. Effect of PRMT inhibition on endothelial NO production. BAECs and COS7 cells were treated with PRMT inhibitor Adox (10 µM) or MTA (1 mM) respectively. NO production was measured by EPR at 48 hours post-treatment. * indicates p<0.05. n = 3.

160

140

120

100

PRMT PRMT Activity (% of Control) 20

C 1 nM 1 µM 1 mM 10 µM 10 nM 100 µM 100 nM

H 2 O 2

Figure 4-2. Effect of H2O2 on PRMT-1 activity. Increasing concentrations of H2O2 (1.0 nM-1.0 mM) were exposed to PRMT-1. PRMT-1 activity was measured by the incorporation 14 14 of [ C]CH3 from [ C]SAM into unmethylated histone residues. The results are presented as % activity compared to vehicle-treated PRMT-1. * indicates significance at p < 0.05. n = 3

250

200

150

100

50 PRMT PRMT Activity (% of Control) * 0

C 1 nM 1 µM 1 mM 10 µM 10 nM 100 µM 100 nM - ONOO

Figure 4-3. Effect of ONOO- on PRMT-1 activity. Increasing concentrations of ONOO- (1.0 nM-1.0 mM) were exposed to PRMT-1. PRMT-1 activity was measured by the 14 14 incorporation of [ C]CH3 from [ C]SAM into unmethylated histone residues. The results are presented as % activity compared to vehicle-treated PRMT-1. * indicates significance at p < 0.05. n = 3

50 kdA

40 kdA

30 kdA

_ + 1 nM 1 µM 1 mM 10 µM 10 nM 100 µM 100 nM - ONOO

Figure 4-4. Nitro-tyrosine formation on PRMT-1 after exposure to ONOO-. Purified PRMT-1 was incubated in the presence of varying concentrations of ONOO- (1.0 nM-1.0 mM). Nitro-tyrosine formation was probed for by immunodetection with an anti-nitro- tyrosine antibody.

400 *

300 * *

200 *

100 PRMT PRMT Activity (% of Control)

0 C M M M M nM µM

0 n 1 0 p 1 µM 00 p 1 1 10 100 n 1 NO

Figure 4-5. Effect of NO on PRMT-1 activity. Increasing concentrations of NO (10 pM-10 μM) were exposed to PRMT-1. PRMT-1 activity was measured by the incorporation of 14 14 [ C]CH3 from [ C]SAM into unmethylated histone residues. The results are presented as % activity compared to vehicle-treated PRMT-1. * indicates significance at p < 0.05. n = 3

100

50 kdA PRMT-1 40 kdA

30 kdA

50 kdA IgG PRMT-1 40 kdA

30 kdA

_ 10 µM NO

Figure 4-6. S-nitrosylation of PRMT-1 after exposure to NO. Purified PRMT-1 was incubated in the presence of 10 µM NO. S-nitrosylation was detected by the biotin switch method.

101

CHAPTER 5 DISCUSSION

Altered NO biosynthesis has been implicated in the pathogenesis of cardiovascular disease and evidence from animal models and clinical studies suggest that accumulation of the endogenous nitric oxide synthase (NOS) inhibitors, asymmetric dimethylarginine (ADMA) and NG- monomethylarginine (NMMA) contribute to the reduced NO generation and disease pathogenesis

[61,62,64,71,73,138,157,209,210]. Asymmetric dimethylarginine (ADMA) plasma levels have been shown to be elevated in diseases related to endothelial dysfunction including hypertension, hyperlipidemia, diabetes mellitus, and others [61-65]. Moreover, it has been shown that ADMA predicts cardiovascular mortality in patients who have coronary heart disease (CHD). Recent evidence published from the multicenter Coronary Artery Risk Determination investigating the

Influence of ADMA Concentration (CARDIAC) study has indicated that ADMA is indeed an independent risk factor for CAD [138]. However, whether the increased risk associated with elevated ADMA is a direct result of NOS impairment is an area of controversy. Significant debate about the contribution of ADMA to the regulation of NOS-dependent NO production has been initiated.

In pathological conditions such as pulmonary hypertension, coronary artery disease, diabetes and hypertension, plasma ADMA levels have been shown to increase from an average of

~0.4 µM to ~0.8 µM [65,138,140-143]. Given that these values are at least 2 orders of magnitude lower than the plasma L-arg levels it is unlikely that elevated plasma ADMA can significantly regulate eNOS activity. It is more likely that elevated plasma ADMA levels reflect increased endothelial concentrations of ADMA. In support of this hypothesis, we and others have demonstrated that endothelial ADMA levels increase 3-4-fold in restenotic lesions and in the ischemia reperfused myocardium [71,205]. Based on the kinetics of cellular inhibition, these

102

concentrations of ADMA would be expected to elicit a 30-40% inhibition in NOS activity [71].

These studies however involve lesion specific increases in ADMA and are not associated with

increased plasma levels of ADMA and would not be expected to contribute to systemic

cardiovascular pathology. In this regard, research has focused on mechanisms leading to

endothelial ADMA accumulation and DDAH has been identified as the principal target responsible

for ADMA accumulation. In support, decreased DDAH expression/activity is evident in disease

states associated with endothelial dysfunction and is believed to be the mechanism responsible for

increased methylarginines and subsequent ADMA mediated eNOS impairment. It has been

estimated that more than 80% of ADMA is metabolized by DDAH [64]; however, it is unclear

which DDAH isoform represents the principal mathylarginine metabolizing enzyme. Currently

there are two known isoforms of DDAH each having different tissue specificity

[72,73,101,131,157,205,211]. DDAH-1 is thought to be associated with tissues that express high levels of neuronal nitric oxide synthase (nNOS), while DDAH-2 is thought be associated with tissues that express eNOS [103].

PCR and western blot analysis has revealed that the endothelium contains mRNA and protein for both DDAH-1 and DDAH-2. However, the biochemical properties and the contribution of each enzyme to the regulation of endothelial NO production has yet to be elucidated. Therefore, in order to assess the relative contribution of each isoform, we cloned and purified recombinant

DDAH-1 and DDAH-2 in an attempt to characterize the enzmyme kinetics of these two methylarginine metabolizing enzymes. Kinetic studies of rhDDAH-1 demonstrated Km values of

68.7 and 53.6 μM and Vmax values of 356 and 154 nmols/mg/min for ADMA and L-NMMA, respectively. This enzymatic activity was selective for free ADMA and L-NMMA and was incapable of hydrolyzing peptide incorporated methylarginines. Subsequent studies performed to

103

determine the effects of reactive oxygen and reactive nitrogen species on DDAH activity

demonstrated that low level oxidant exposure had little effect on enzyme activity and that

concentrations approaching 100 μM were needed to confer significant inhibition of hDDAH-1

activity. However, exposure of hDDAH-1 to the lipid oxidation product, 4-HNE, dose- dependently inhibited DDAH activity with 15% inhibition observed at 10 μM, 50% inhibition at

50 μM, and complete inhibition at 500 μM. Mass spectrometry analysis demonstrated that the mechanism of inhibition resulted from the formation of Michael adducts on His 173, which lies within the active site catalytic triad of hDDAH-1. These studies were performed with pathophysiologicaly relevant concentrations of this lipid peroxidation product and suggest that

DDAH activity can be impaired under conditions of increased oxidative stress. Because DDAH-1

is the primary enzyme involved in methylarginine metabolism, the loss of activity of this enzyme

would result in impaired NOS activity and reduced NO bioavailability.

With regards to DDAH-2, studies indicated no catalytic activity towards ADMA despite the fact that gene silencing of DDAH-2 in the endothelium significantly inhibited eNOS-derived NO production. These findings, together with the observation that DDAH-2 silencing has no effect on endothelial ADMA levels and NO inhibition was not reversed with L-arg supplementation led us

to hypothesize that DDAH-2 effects on the endothelium are indepednet of ADMA. The most

convincing evidence that DDAH may regulate cellular function through mechanisms independent

of ADMA-mediated NOS inhibition come from data on the DDAH-1 knockout mouse.

Homozygous null mice for DDAH-1 are embryonic lethal while the NOS triple knockout mice are

viable [157]. This provides strong evidence that DDAH effects are not limited to ADMA

dependent regulation of eNOS. Using DDAH-1 heterozygous mice, which are viable, Leiper et. al

demonstrated that reduced DDAH-1 activity leads to accumulation of plasma ADMA and a

104

reduction in NO signaling. These animals exhibited a 50% decrease in DDAH activity which was

associated with a 20% increase in plasma and tissue ADMA levels [157]. This in turn was

associated with vascular pathology, including endothelial dysfunction, increased systemic vascular

resistance and elevated systemic and pulmonary blood pressure. Given that the intracellular

concentrations of ADMA are 1-3 µM, it is unlikely that a 20% increase in ADMA could be

responsible for the 40% reduction in endothelial dependent relaxation observed with the DDAH+/-

mice. Moreover, the addition of exogenous L-arg to the organ chambers only partially restored the loss in endothelial relaxation [157]. These results further support the hypothesis that DDAH modulates endothelial function through both ADMA-NOS dependent pathways as well as independent. Although this represents an overall paradigm shift, it is not surprising given the lethality of the DDAH-1 knockout mouse.

Recent data generated from our laboratory further support his hypothesis and provides a potential mechanism for the ADMA independent effects of DDAH-2. Evidence suggests that loss of

DDAH activity increases protein-arginine methylation and this increased methylation contributes to

endothelial dysfunction. Although the demethylation of methylarginines is believed to be restricted

to free methylarginines, it is possible that DDAH-2 may function as the elusive protein-arginine demethylase. Nevertheless, using the ApoE+/- mouse model we observed that the decreased DDAH

activity observed in these mice was associated with significantly increased levels of protein-arginine

methylation [210]. In order to further examine the role of DDAH in endothelial dysfunction we

crossed DDAH-1 transgenic mice with ApoE+/- mice and examined DDAH activity, levels of

protein-arginine methylation and vascular reactivity. Result demonstrated that ApoE+/- mice had a

41% decrease in DDAH activity, a 20% reduction in endothelial dependent relaxation and increased

protein-arginine methylation. DDAH-1 over-expression in the ApoE+/- mice restored DDAH activity

105

and increased endothelial dependent relaxation. Moreover, increased DDAH-1 expression reduced protein methylation in the crossed mice. These studies provide further support that DDAH regulates eNOS activity and endothelial function through both ADMA-dependent and independent mechanisms. In this regard, elevated plasma ADMA may serve as a marker of impaired methylarginine metabolism and the pathology previously attributed to elevated ADMA may be manifested, at least in part, through altered activity of the enzymes involved in ADMA regulation, specifically DDAH and PRMT.

Over the last 40 years, arginine methylation has been extensively studied in prokaryotes and eukaryotes revealing a pivotal role of this posttranslational modification in the regulation of a number of cellular processes. Protein arginine methylation has been demonstrated to be involved in the modulation of transcription, RNA metabolism and protein-protein interactions, thereby controlling cellular differentiation, proliferation, survival, and apoptosis [81]. It is thus not surprising that dysregulation of protein methylation would play a role in the regulation of endothelial function. In support, we have demonstrated that pharmacological inhibition of PRMT-1 results in a significant increase in the levels of endothelial NO production. These findings suggest that arginine- methylation may exert an inhibitory effect on eNOS activity. Therefore, we undertook a series of studies to examine potential mechanisms involved in the regulation of PRMT activity. Specifically, we examined whether PRMT-1 activity was redox regulated by examining the effects of ROS and

RNS on enzymatic activity.

Although results demonstrated that PRMT-1 activity was largely resistant to ROS induced dysregulation, the enzyme was very sensitive to NO. Specifically, at physiologically relevant concentratins of NO, PRMT-1 activity was increased two-fold. Further mechanistic studies identified the likely mechanism which involves S-Nitorsylation as the biotin-switch assay

106

demonstrated increased formation of S-nitrosyl complexes on PRMT-1 following exposure to NO.

The physiological implications of this are quite intriguing and suggest that a negative feed-back loop

may exist between PRMT-1 and NOS wherein elevated NO production results in increased PRMT-1

activity which would be expected to increase levels of methylarginines. Increased levels of both

protein-incorporated and free-methylarginines are known to inhbit eNOS activity. Alternatively,

reduced NO bioavailability, as is known to occur in cardiovascular diseases, would be expected to

trigger a reduction in PRMT activity. Because PRMT is involved in the regulation of numerous

cellular processes, NO-dependent regulation of PRMT activity may explain many of the

pathophysioligcal manifestations of reduced NO bioavailability. These findings together with our

previously published studies demonstrating that loss of DDAH acitivty is associated with increased

protein methylation, provide evidence for the existence of a PRMT-ADMA-DDAH axis.

Although increased plasma levels of ADMA are associated with cardiovascular disease, it is the endothelial free-ADMA levels that have been implicated in the regulation of NOS activity. It is therefore surprising that, to date, there have been no studies examining the cellular kinetics of

ADMA synthesis and metabolism in the endothelium. It is generally accepted that PRMTs synthesize methylarginines on proteins using the methyl donor SAM and L-Arg as the terminal methyl acceptor. It is then believed that normal protein turnover releases free methylarginines which are then metabolized to citrulline by DDAH. In this regard, loss of DDAH activity has been implicated as the molecular trigger for ADMA accumulation and subsequent endothelial dysfunction. It is our hypothesis that there is cross-talk among these pathways and that the levels of both free and protein incorporated methylarginines play important roles in regulating endothelial function, including but not limited to eNOS regulation. In summary, dysregulation of the PRMT-

DDAH-ADMA axis has now been shown to contribute to the pathogenesis of several cardiovascular

107

disorders, in experimental animal models as well as human disease. Causal relationships between

dysregulated arginine-methylation and the initiation, progression, or therapy of disease, however, remain to be dissected. Future investigations into arginine-methylation and DDAH dynamics in

disease states are clearly needed in order elucidate the role of this post-translational modification in

the pathogenesis of cardiovascular disease.

108

WORKS CITED

[1] D. Lloyd-Jones, R.J. Adams, T.M. Brown, M. Carnethon, S. Dai, G. De Simone, T.B. Ferguson, E. Ford, K. Furie, C. Gillespie, A. Go, K. Greenlund, N. Haase, S. Hailpern, P.M. Ho, V. Howard, B. Kissela, S. Kittner, D. Lackland, L. Lisabeth, A. Marelli, M.M. McDermott, J. Meigs, D. Mozaffarian, M. Mussolino, G. Nichol, V. Roger, W. Rosamond, R. Sacco, P. Sorlie, R. Stafford, T. Thom, S. Wasserthiel-Smoller, N.D. Wong and J. Wylie-Rosett Heart Disease and Stroke Statistics--2010 Update. A Report From the American Heart Association, Circulation (2009).

[2] R.M. Palmer, A.G. Ferrige and S. Moncada Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor, Nature 327 (1987) 524-526.

[3] D.S. Bredt and S.H. Snyder Nitric oxide mediates glutamate-linked enhancement of cGMP levels in the cerebellum, Proc Natl Acad Sci U S A 86 (1989) 9030-9033.

[4] C.A. Gruetter, B.K. Barry, D.B. McNamara, D.Y. Gruetter, P.J. Kadowitz and L. Ignarro Relaxation of bovine coronary artery and activation of coronary arterial guanylate cyclase by nitric oxide, nitroprusside and a carcinogenic nitrosoamine, J Cyclic Nucleotide Res 5 (1979) 211-224.

[5] J.B. Hibbs, Jr., R.R. Taintor, Z. Vavrin and E.M. Rachlin Nitric oxide: a cytotoxic activated macrophage effector molecule, Biochem Biophys Res Commun 157 (1988) 87- 94.

[6] R.M. Palmer, A.G. Ferrige and S. Moncada Nitric oxide release accounts for the biological activity of endothelium-derived relaxing factor, Nature 327 (1987) 524-526.

[7] R.M. Palmer, D.D. Rees, D.S. Ashton and S. Moncada L-arginine is the physiological precursor for the formation of nitric oxide in endothelium-dependent relaxation, Biochem Biophys Res Commun 153 (1988) 1251-1256.

[8] D.S. Bredt and S.H. Snyder Isolation of nitric oxide synthetase, a calmodulin-requiring enzyme, Proc Natl Acad Sci U S A 87 (1990) 682-685.

[9] R. Busse and A. Mulsch Calcium-dependent nitric oxide synthesis in endothelial cytosol is mediated by calmodulin, FEBS Lett 265 (1990) 133-136.

[10] D.S. Bredt, P.M. Hwang, C.E. Glatt, C. Lowenstein, R.R. Reed and S.H. Snyder Cloned and expressed nitric oxide synthase structurally resembles cytochrome P-450 reductase, Nature 351 (1991) 714-718.

[11] H.H. Schmidt and F. Murad Purification and characterization of a human NO synthase, Biochem Biophys Res Commun 181 (1991) 1372-1377.

[12] B.S. Masters, K. McMillan, E.A. Sheta, J.S. Nishimura, L.J. Roman and P. Martasek Neuronal nitric oxide synthase, a modular enzyme formed by convergent evolution:

109

structure studies of a cysteine thiolate-liganded heme protein that hydroxylates L-arginine to produce NO. as a cellular signal, Faseb J 10 (1996) 552-558.

[13] B. Mayer and B. Hemmens Biosynthesis and action of nitric oxide in mammalian cells, Trends Biochem Sci 22 (1997) 477-481.

[14] D.J. Stuehr Structure-function aspects in the nitric oxide synthases, Annu Rev Pharmacol Toxicol 37 (1997) 339-359.

[15] W.P. Arnold, C.K. Mittal, S. Katsuki and F. Murad Nitric oxide activates guanylate cyclase and increases guanosine 3':5'-cyclic monophosphate levels in various tissue preparations, Proc Natl Acad Sci U S A 74 (1977) 3203-3207.

[16] K. Guo, V. Andres and K. Walsh Nitric oxide-induced downregulation of Cdk2 activity and cyclin A gene transcription in vascular smooth muscle cells, Circulation 97 (1998) 2066-2072.

[17] R. De Caterina, P. Libby, H.B. Peng, V.J. Thannickal, T.B. Rajavashisth, M.A. Gimbrone, Jr., W.S. Shin and J.K. Liao Nitric oxide decreases cytokine-induced endothelial activation. Nitric oxide selectively reduces endothelial expression of adhesion molecules and proinflammatory cytokines, J Clin Invest 96 (1995) 60-68.

[18] M. Takahashi, U. Ikeda, J. Masuyama, H. Funayama, S. Kano and K. Shimada Nitric oxide attenuates adhesion molecule expression in human endothelial cells, Cytokine 8 (1996) 817-821.

[19] D.R. Riddell and J.S. Owen Nitric oxide and platelet aggregation, Vitam Horm 57 (1999) 25-48.

[20] J. Vasquez-Vivar, B. Kalyanaraman, P. Martasek, N. Hogg, B.S. Masters, H. Karoui, P. Tordo and K.A. Pritchard, Jr. Superoxide generation by endothelial nitric oxide synthase: the influence of cofactors, Proc Natl Acad Sci U S A 95 (1998) 9220-9225.

[21] Y. Xia, V.L. Dawson, T.M. Dawson, S.H. Snyder and J.L. Zweier Nitric oxide synthase generates superoxide and nitric oxide in arginine-depleted cells leading to peroxynitrite- mediated cellular injury, Proc Natl Acad Sci U S A 93 (1996) 6770-6774.

[22] Y. Xia, A.L. Tsai, V. Berka and J.L. Zweier Superoxide generation from endothelial nitric-oxide synthase. A Ca2+/calmodulin-dependent and tetrahydrobiopterin regulatory process, J Biol Chem 273 (1998) 25804-25808.

[23] C.R. Nishida and P.R. Ortiz de Montellano Electron transfer and catalytic activity of nitric oxide synthases. Chimeric constructs of the neuronal, inducible, and endothelial isoforms, J Biol Chem 273 (1998) 5566-5571.

[24] A. Luckhoff, U. Pohl, A. Mulsch and R. Busse Differential role of extra- and intracellular calcium in the release of EDRF and prostacyclin from cultured endothelial cells, Br J Pharmacol 95 (1988) 189-196.

110

[25] R.C. Venema, H.S. Sayegh, J.D. Kent and D.G. Harrison Identification, characterization, and comparison of the calmodulin-binding domains of the endothelial and inducible nitric oxide synthases, J Biol Chem 271 (1996) 6435-6440.

[26] G. Garcia-Cardena, P. Martasek, B.S. Masters, P.M. Skidd, J. Couet, S. Li, M.P. Lisanti and W.C. Sessa Dissecting the interaction between nitric oxide synthase (NOS) and caveolin. Functional significance of the nos caveolin binding domain in vivo, J Biol Chem 272 (1997) 25437-25440.

[27] J.B. Michel, O. Feron, D. Sacks and T. Michel Reciprocal regulation of endothelial nitric- oxide synthase by Ca2+-calmodulin and caveolin, J Biol Chem 272 (1997) 15583-15586.

[28] I. Fleming and R. Busse Molecular mechanisms involved in the regulation of the endothelial nitric oxide synthase, Am J Physiol Regul Integr Comp Physiol 284 (2003) R1-12.

[29] G. Garcia-Cardena, R. Fan, V. Shah, R. Sorrentino, G. Cirino, A. Papapetropoulos and W.C. Sessa Dynamic activation of endothelial nitric oxide synthase by Hsp90, Nature 392 (1998) 821-824.

[30] J.P. Gratton, J. Fontana, D.S. O'Connor, G. Garcia-Cardena, T.J. McCabe and W.C. Sessa Reconstitution of an endothelial nitric-oxide synthase (eNOS), hsp90, and caveolin-1 complex in vitro. Evidence that hsp90 facilitates calmodulin stimulated displacement of eNOS from caveolin-1, J Biol Chem 275 (2000) 22268-22272.

[31] P.M. Bauer, D. Fulton, Y.C. Boo, G.P. Sorescu, B.E. Kemp, H. Jo and W.C. Sessa Compensatory phosphorylation and protein-protein interactions revealed by loss of function and gain of function mutants of multiple serine phosphorylation sites in endothelial nitric-oxide synthase, J Biol Chem 278 (2003) 14841-14849.

[32] T.J. McCabe, D. Fulton, L.J. Roman and W.C. Sessa Enhanced electron flux and reduced calmodulin dissociation may explain "calcium-independent" eNOS activation by phosphorylation, J Biol Chem 275 (2000) 6123-6128.

[33] M.A. Corson, N.L. James, S.E. Latta, R.M. Nerem, B.C. Berk and D.G. Harrison Phosphorylation of endothelial nitric oxide synthase in response to fluid shear stress, Circ Res 79 (1996) 984-991.

[34] S. Dimmeler, I. Fleming, B. Fisslthaler, C. Hermann, R. Busse and A.M. Zeiher Activation of nitric oxide synthase in endothelial cells by Akt-dependent phosphorylation, Nature 399 (1999) 601-605.

[35] Y.C. Boo, G. Sorescu, N. Boyd, I. Shiojima, K. Walsh, J. Du and H. Jo Shear stress stimulates phosphorylation of endothelial nitric-oxide synthase at Ser1179 by Akt- independent mechanisms: role of protein kinase A, J Biol Chem 277 (2002) 3388-3396.

111

[36] D. Fulton, J.P. Gratton, T.J. McCabe, J. Fontana, Y. Fujio, K. Walsh, T.F. Franke, A. Papapetropoulos and W.C. Sessa Regulation of endothelium-derived nitric oxide production by the protein kinase Akt, Nature 399 (1999) 597-601.

[37] K. Hisamoto, M. Ohmichi, H. Kurachi, J. Hayakawa, Y. Kanda, Y. Nishio, K. Adachi, K. Tasaka, E. Miyoshi, N. Fujiwara, N. Taniguchi and Y. Murata Estrogen induces the Akt- dependent activation of endothelial nitric-oxide synthase in vascular endothelial cells, J Biol Chem 276 (2001) 3459-3467.

[38] J. Igarashi and T. Michel Sphingosine 1-phosphate and isoform-specific activation of phosphoinositide 3-kinase beta. Evidence for divergence and convergence of receptor- regulated endothelial nitric-oxide synthase signaling pathways, J Biol Chem 276 (2001) 36281-36288.

[39] F. Kim, B. Gallis and M.A. Corson TNF-alpha inhibits flow and insulin signaling leading to NO production in aortic endothelial cells, Am J Physiol Cell Physiol 280 (2001) C1057-1065.

[40] I. Fleming, C. Schulz, B. Fichtlscherer, B.E. Kemp, B. Fisslthaler and R. Busse AMP- activated protein kinase (AMPK) regulates the insulin-induced activation of the nitric oxide synthase in human platelets, Thromb Haemost 90 (2003) 863-871.

[41] E. Butt, M. Bernhardt, A. Smolenski, P. Kotsonis, L.G. Frohlich, A. Sickmann, H.E. Meyer, S.M. Lohmann and H.H. Schmidt Endothelial nitric-oxide synthase (type III) is activated and becomes calcium independent upon phosphorylation by cyclic nucleotide- dependent protein kinases, J Biol Chem 275 (2000) 5179-5187.

[42] I. Fleming, B. Fisslthaler, S. Dimmeler, B.E. Kemp and R. Busse Phosphorylation of Thr(495) regulates Ca(2+)/calmodulin-dependent endothelial nitric oxide synthase activity, Circ Res 88 (2001) E68-75.

[43] D.M. Greif, R. Kou and T. Michel Site-specific dephosphorylation of endothelial nitric oxide synthase by protein phosphatase 2A: evidence for crosstalk between phosphorylation sites, Biochemistry 41 (2002) 15845-15853.

[44] B.J. Michell, Z. Chen, T. Tiganis, D. Stapleton, F. Katsis, D.A. Power, A.T. Sim and B.E. Kemp Coordinated control of endothelial nitric-oxide synthase phosphorylation by protein kinase C and the cAMP-dependent protein kinase, J Biol Chem 276 (2001) 17625-17628.

[45] Q. Wei and Y. Xia Proteasome inhibition down-regulates endothelial nitric-oxide synthase phosphorylation and function, J Biol Chem 281 (2006) 21652-21659.

[46] P.P. Kou R, and Michel T Phosphorylation of the endothelial isoform of nitric oxide synthase at serine 116: Identification of a novel path for eNOS regulation by lysophosphatidic acid, Circulation 104 (2001) 509.

112

[47] J. Whitsett, P. Martasek, H. Zhao, D.W. Schauer, K. Hatakeyama, B. Kalyanaraman and J. Vasquez-Vivar Endothelial cell superoxide anion radical generation is not dependent on endothelial nitric oxide synthase-serine 1179 phosphorylation and endothelial nitric oxide synthase dimer/monomer distribution, Free Radic Biol Med 40 (2006) 2056-2068.

[48] M. Matsubara, K. Titani and H. Taniguchi Interaction of calmodulin-binding domain peptides of nitric oxide synthase with membrane phospholipids: regulation by protein phosphorylation and Ca(2+)-calmodulin, Biochemistry 35 (1996) 14651-14658.

[49] M.B. Harris, H. Ju, V.J. Venema, H. Liang, R. Zou, B.J. Michell, Z.P. Chen, B.E. Kemp and R.C. Venema Reciprocal phosphorylation and regulation of endothelial nitric-oxide synthase in response to bradykinin stimulation, J Biol Chem 276 (2001) 16587-16591.

[50] B. Gallis, G.L. Corthals, D.R. Goodlett, H. Ueba, F. Kim, S.R. Presnell, D. Figeys, D.G. Harrison, B.C. Berk, R. Aebersold and M.A. Corson Identification of flow-dependent endothelial nitric-oxide synthase phosphorylation sites by mass spectrometry and regulation of phosphorylation and nitric oxide production by the phosphatidylinositol 3- kinase inhibitor LY294002, J Biol Chem 274 (1999) 30101-30108.

[51] R. Kou, D. Greif and T. Michel Dephosphorylation of endothelial nitric-oxide synthase by vascular endothelial growth factor. Implications for the vascular responses to cyclosporin A, J Biol Chem 277 (2002) 29669-29673.

[52] B.J. Michell, M.B. Harris, Z.P. Chen, H. Ju, V.J. Venema, M.A. Blackstone, W. Huang, R.C. Venema and B.E. Kemp Identification of regulatory sites of phosphorylation of the bovine endothelial nitric-oxide synthase at serine 617 and serine 635, J Biol Chem 277 (2002) 42344-42351.

[53] I. Fleming, J. Bauersachs, B. Fisslthaler and R. Busse Ca2+-independent activation of the endothelial nitric oxide synthase in response to tyrosine phosphatase inhibitors and fluid shear stress, Circ Res 82 (1998) 686-695.

[54] T. Michel, G.K. Li and L. Busconi Phosphorylation and subcellular translocation of endothelial nitric oxide synthase, Proc Natl Acad Sci U S A 90 (1993) 6252-6256.

[55] G.R. Hellermann, B.R. Flam, D.C. Eichler and L.P. Solomonson Stimulation of receptor- mediated nitric oxide production by vanadate, Arterioscler Thromb Vasc Biol 20 (2000) 2045-2050.

[56] V. Rizzo, D.P. McIntosh, P. Oh and J.E. Schnitzer In situ flow activates endothelial nitric oxide synthase in luminal caveolae of endothelium with rapid caveolin dissociation and calmodulin association, J Biol Chem 273 (1998) 34724-34729.

[57] V.J. Venema, M.B. Marrero and R.C. Venema Bradykinin-stimulated protein tyrosine phosphorylation promotes endothelial nitric oxide synthase translocation to the cytoskeleton, Biochem Biophys Res Commun 226 (1996) 703-710.

113

[58] G. Garcia-Cardena, R. Fan, D.F. Stern, J. Liu and W.C. Sessa Endothelial nitric oxide synthase is regulated by tyrosine phosphorylation and interacts with caveolin-1, J Biol Chem 271 (1996) 27237-27240.

[59] D. Fulton, J. Fontana, G. Sowa, J.P. Gratton, M. Lin, K.X. Li, B. Michell, B.E. Kemp, D. Rodman and W.C. Sessa Localization of endothelial nitric-oxide synthase phosphorylated on serine 1179 and nitric oxide in Golgi and plasma membrane defines the existence of two pools of active enzyme, J Biol Chem 277 (2002) 4277-4284.

[60] J.B. Hibbs, Jr., Z. Vavrin and R.R. Taintor L-arginine is required for expression of the activated macrophage effector mechanism causing selective metabolic inhibition in target cells, J Immunol 138 (1987) 550-565.

[61] R. Maas, K. Quitzau, E. Schwedhelm, L. Spieker, W. Rafflenbeul, A. Steenpass, T.F. Luscher and R.H. Boger Asymmetrical dimethylarginine (ADMA) and coronary endothelial function in patients with coronary artery disease and mild hypercholesterolemia, Atherosclerosis 191 (2007) 211-219.

[62] R. Maas, L. Dentz, E. Schwedhelm, W. Thoms, O. Kuss, N. Hiltmeyer, M. Haddad, T. Kloss, T. Standl and R.H. Boger Elevated plasma concentrations of the endogenous nitric oxide synthase inhibitor asymmetric dimethylarginine predict adverse events in patients undergoing noncardiac surgery, Crit Care Med 35 (2007) 1876-1881.

[63] G.I. Boger, T.K. Rudolph, R. Maas, E. Schwedhelm, E. Dumbadze, A. Bierend, R.A. Benndorf and R.H. Boger Asymmetric dimethylarginine determines the improvement of endothelium-dependent vasodilation by simvastatin: Effect of combination with oral L- arginine, J Am Coll Cardiol 49 (2007) 2274-2282.

[64] V. Achan, M. Broadhead, M. Malaki, G. Whitley, J. Leiper, R. MacAllister and P. Vallance Asymmetric dimethylarginine causes hypertension and cardiac dysfunction in humans and is actively metabolized by dimethylarginine dimethylaminohydrolase, Arterioscler Thromb Vasc Biol 23 (2003) 1455-1459.

[65] R.H. Boger, S.M. Bode-Boger, K. Sydow, D.D. Heistad and S.R. Lentz Plasma concentration of asymmetric dimethylarginine, an endogenous inhibitor of nitric oxide synthase, is elevated in monkeys with hyperhomocyst(e)inemia or hypercholesterolemia, Arterioscler Thromb Vasc Biol 20 (2000) 1557-1564.

[66] R.H. Boger, R. Maas, F. Schulze and E. Schwedhelm Elevated levels of asymmetric dimethylarginine (ADMA) as a marker of cardiovascular disease and mortality, Clin Chem Lab Med 43 (2005) 1124-1129.

[67] E.B. Marliss, S. Chevalier, R. Gougeon, J.A. Morais, M. Lamarche, O.A. Adegoke and G. Wu Elevations of plasma methylarginines in obesity and ageing are related to insulin sensitivity and rates of protein turnover, Diabetologia 49 (2006) 351-359.

114

[68] P. Vallance and J. Leiper Cardiovascular Biology of the Asymmetric Dimethylarginine:Dimethylarginine Dimethylaminohydrolase Pathway, Arterioscler Thromb Vasc Biol 22 (2004) 22.

[69] P. Vallance, A. Leone, A. Calver, J. Collier and S. Moncada Accumulation of an endogenous inhibitor of nitric oxide synthesis in chronic renal failure, Lancet 339 (1992) 572-575.

[70] E.I. Closs, F.Z. Basha, A. Habermeier and U. Forstermann Interference of L-arginine analogues with L-arginine transport mediated by the y+ carrier hCAT-2B, Nitric Oxide 1 (1997) 65-73.

[71] A.J. Cardounel, H. Cui, A. Samouilov, W. Johnson, P. Kearns, A.L. Tsai, V. Berka and J.L. Zweier Evidence for the pathophysiological role of endogenous methylarginines in regulation of endothelial NO production and vascular function, J Biol Chem 282 (2007) 879-887.

[72] M. Tanaka, K. Sydow, F. Gunawan, J. Jacobi, P.S. Tsao, R.C. Robbins and J.P. Cooke Dimethylarginine dimethylaminohydrolase overexpression suppresses graft coronary artery disease, Circulation 112 (2005) 1549-1556.

[73] P. Vallance and J. Leiper Cardiovascular biology of the asymmetric dimethylarginine:dimethylarginine dimethylaminohydrolase pathway, Arterioscler Thromb Vasc Biol 24 (2004) 1023-1030.

[74] R.H. Boger, S.M. Bode-Boger, A. Szuba, P.S. Tsao, J.R. Chan, O. Tangphao, T.F. Blaschke and J.P. Cooke Asymmetric dimethylarginine (ADMA): a novel risk factor for endothelial dysfunction: its role in hypercholesterolemia, Circulation 98 (1998) 1842- 1847.

[75] R. Schnabel, S. Blankenberg, E. Lubos, K.J. Lackner, H.J. Rupprecht, C. Espinola-Klein, N. Jachmann, F. Post, D. Peetz, C. Bickel, F. Cambien, L. Tiret and T. Munzel Asymmetric dimethylarginine and the risk of cardiovascular events and death in patients with coronary artery disease: results from the AtheroGene Study, Circ Res 97 (2005) e53- 59.

[76] K.Y. Lin, A. Ito, T. Asagami, P.S. Tsao, S. Adimoolam, M. Kimoto, H. Tsuji, G.M. Reaven and J.P. Cooke Impaired nitric oxide synthase pathway in diabetes mellitus: role of asymmetric dimethylarginine and dimethylarginine dimethylaminohydrolase, Circulation 106 (2002) 987-992.

[77] F. Schulze, H. Lenzen, C. Hanefeld, A. Bartling, K.J. Osterziel, L. Goudeva, C. Schmidt- Lucke, M. Kusus, R. Maas, E. Schwedhelm, D. Strodter, B.C. Simon, A. Mugge, W.G. Daniel, H. Tillmanns, B. Maisch, T. Streichert and R.H. Boger Asymmetric dimethylarginine is an independent risk factor for coronary heart disease: results from the multicenter Coronary Artery Risk Determination investigating the Influence of ADMA Concentration (CARDIAC) study, Am Heart J 152 (2006) 493 e491-498.

115

[78] J. Leiper, M. Nandi, B. Torondel, J. Murray-Rust, M. Malaki, B. O'Hara, S. Rossiter, S. Anthony, M. Madhani, D. Selwood, C. Smith, B. Wojciak-Stothard, A. Rudiger, R. Stidwill, N.Q. McDonald and P. Vallance Disruption of methylarginine metabolism impairs vascular homeostasis, Nat Med. 2007 Feb;13(2):198-203. Epub 2007 Feb 4.

[79] J. Leiper and P. Vallance Biological significance of endogenous methylarginines that inhibit nitric oxide synthases, Cardiovasc Res 43 (1999) 542-548.

[80] Y.H. Lee and M.R. Stallcup Minireview: protein arginine methylation of nonhistone proteins in transcriptional regulation, Mol Endocrinol 23 (2009) 425-433.

[81] M.T. Bedford and S. Richard Arginine methylation an emerging regulator of protein function, Mol Cell 18 (2005) 263-272.

[82] M.C. Boulanger, C. Liang, R.S. Russell, R. Lin, M.T. Bedford, M.A. Wainberg and S. Richard Methylation of Tat by PRMT6 regulates human immunodeficiency virus type 1 gene expression, J Virol 79 (2005) 124-131.

[83] J.D. Gary and S. Clarke RNA and protein interactions modulated by protein arginine methylation, Prog Nucleic Acid Res Mol Biol 61 (1998) 65-131.

[84] J. Najbauer, B.A. Johnson, A.L. Young and D.W. Aswad Peptides with sequences similar to glycine, arginine-rich motifs in proteins interacting with RNA are efficiently recognized by methyltransferase(s) modifying arginine in numerous proteins, J Biol Chem 268 (1993) 10501-10509.

[85] D. Cheng, J. Cote, S. Shaaban and M.T. Bedford The arginine methyltransferase CARM1 regulates the coupling of transcription and mRNA processing, Mol Cell 25 (2007) 71-83.

[86] B. Chang, Y. Chen, Y. Zhao and R.K. Bruick JMJD6 is a histone arginine demethylase, Science 318 (2007) 444-447.

[87] Y. Wang, J. Wysocka, J. Sayegh, Y.H. Lee, J.R. Perlin, L. Leonelli, L.S. Sonbuchner, C.H. McDonald, R.G. Cook, Y. Dou, R.G. Roeder, S. Clarke, M.R. Stallcup, C.D. Allis and S.A. Coonrod Human PAD4 regulates histone arginine methylation levels via demethylimination, Science 306 (2004) 279-283.

[88] Y. Shi, F. Lan, C. Matson, P. Mulligan, J.R. Whetstine, P.A. Cole, R.A. Casero and Y. Shi Histone demethylation mediated by the nuclear amine oxidase homolog LSD1, Cell 119 (2004) 941-953.

[89] A. Scorilas, M.H. Black, M. Talieri and E.P. Diamandis Genomic organization, physical mapping, and expression analysis of the human protein arginine methyltransferase 1 gene, Biochem Biophys Res Commun 278 (2000) 349-359.

[90] M.R. Pawlak, C.A. Scherer, J. Chen, M.J. Roshon and H.E. Ruley Arginine N- methyltransferase 1 is required for early postimplantation mouse development, but cells deficient in the enzyme are viable, Mol Cell Biol 20 (2000) 4859-4869.

116

[91] N. Yadav, J. Lee, J. Kim, J. Shen, M.C. Hu, C.M. Aldaz and M.T. Bedford Specific protein methylation defects and gene expression perturbations in coactivator-associated arginine methyltransferase 1-deficient mice, Proc Natl Acad Sci U S A 100 (2003) 6464- 6468.

[92] W.J. Lin, J.D. Gary, M.C. Yang, S. Clarke and H.R. Herschman The mammalian immediate-early TIS21 protein and the leukemia-associated BTG1 protein interact with a protein-arginine N-methyltransferase, J Biol Chem 271 (1996) 15034-15044.

[93] V. Singh, T.B. Miranda, W. Jiang, A. Frankel, M.E. Roemer, V.A. Robb, D.H. Gutmann, H.R. Herschman, S. Clarke and I.F. Newsham DAL-1/4.1B tumor suppressor interacts with protein arginine N-methyltransferase 3 (PRMT3) and inhibits its ability to methylate substrates in vitro and in vivo, Oncogene 23 (2004) 7761-7771.

[94] X. Zhang and X. Cheng Structure of the predominant protein arginine methyltransferase PRMT1 and analysis of its binding to substrate peptides, Structure 11 (2003) 509-520.

[95] Y. Chen, X. Xu, M. Sheng, X. Zhang, Q. Gu and Z. Zheng PRMT-1 and DDAHs- induced ADMA upregulation is involved in ROS- and RAS-mediated diabetic retinopathy, Exp Eye Res 89 (2009) 1028-1034.

[96] T. Osanai, M. Saitoh, S. Sasaki, H. Tomita, T. Matsunaga and K. Okumura Effect of shear stress on asymmetric dimethylarginine release from vascular endothelial cells, Hypertension 42 (2003) 985-990.

[97] J.R. McDermott Studies on the catabolism of Ng-methylarginine, Ng, Ng- dimethylarginine and Ng, Ng-dimethylarginine in the rabbit, Biochem J 154 (1976) 179- 184.

[98] T. Ogawa, M. Kimoto, H. Watanabe and K. Sasaoka Metabolism of NG,NG-and NG,N'G-dimethylarginine in rats, Arch Biochem Biophys 252 (1987) 526-537.

[99] C.T. Tran, J.M. Leiper and P. Vallance The DDAH/ADMA/NOS pathway, Atheroscler Suppl 4 (2003) 33-40.

[100] J. Murray-Rust, J. Leiper, M. McAlister, J. Phelan, S. Tilley, J. Santa Maria, P. Vallance and N. McDonald Structural insights into the hydrolysis of cellular nitric oxide synthase inhibitors by dimethylarginine dimethylaminohydrolase, Nat Struct Biol 8 (2001) 679- 683.

[101] T. Ogawa, M. Kimoto and K. Sasaoka Occurrence of a new enzyme catalyzing the direct conversion of NG,NG-dimethyl-L-arginine to L-citrulline in rats, Biochem Biophys Res Commun 148 (1987) 671-677.

[102] R.J. MacAllister, H. Parry, M. Kimoto, T. Ogawa, R.J. Russell, H. Hodson, G.S. Whitley and P. Vallance Regulation of nitric oxide synthesis by dimethylarginine dimethylaminohydrolase, Br J Pharmacol 119 (1996) 1533-1540.

117

[103] J.M. Leiper, J. Santa Maria, A. Chubb, R.J. MacAllister, I.G. Charles, G.S. Whitley and P. Vallance Identification of two human dimethylarginine dimethylaminohydrolases with distinct tissue distributions and homology with microbial arginine deiminases, Biochem J 343 Pt 1 (1999) 209-214.

[104] G.M. Birdsey, J.M. Leiper and P. Vallance Intracellular localization of dimethylarginine dimethylaminohydrolase overexpressed in an endothelial cell line, Acta Physiol Scand 168 (2000) 73-79.

[105] R.J. Nijveldt, T. Teerlink, M.P. Siroen, A.A. van Lambalgen, J.A. Rauwerda and P.A. van Leeuwen The liver is an important organ in the metabolism of asymmetrical dimethylarginine (ADMA), Clin Nutr 22 (2003) 17-22.

[106] R.J. Nijveldt, T. Teerlink, C. van Guldener, H.A. Prins, A.A. van Lambalgen, C.D. Stehouwer, J.A. Rauwerda and P.A. van Leeuwen Handling of asymmetrical dimethylarginine and symmetrical dimethylarginine by the rat kidney under basal conditions and during endotoxaemia, Nephrol Dial Transplant 18 (2003) 2542-2550.

[107] M.L. Onozato, A. Tojo, J. Leiper, T. Fujita, F. Palm and C.S. Wilcox Expression of NG,NG-dimethylarginine dimethylaminohydrolase and protein arginine N- methyltransferase isoforms in diabetic rat kidney: effects of angiotensin II receptor blockers, Diabetes 57 (2008) 172-180.

[108] A. Tojo, W.J. Welch, V. Bremer, M. Kimoto, K. Kimura, M. Omata, T. Ogawa, P. Vallance and C.S. Wilcox Colocalization of demethylating enzymes and NOS and functional effects of methylarginines in rat kidney, Kidney Int 52 (1997) 1593-1601.

[109] C.T. Tran, M.F. Fox, P. Vallance and J.M. Leiper Chromosomal localization, gene structure, and expression pattern of DDAH1: comparison with DDAH2 and implications for evolutionary origins, Genomics 68 (2000) 101-105.

[110] M. Kimoto, H. Tsuji, T. Ogawa and K. Sasaoka Detection of NG,NG-dimethylarginine dimethylaminohydrolase in the nitric oxide-generating systems of rats using monoclonal antibody, Arch Biochem Biophys 300 (1993) 657-662.

[111] D. Wang, W. Wang, P. Gill, W. Welch and C. Wilcox DDAH-2 regulates endothelial function, eNOS expression and nitric oxide activity in rat mesenteric resistance vessels (Abstract), Proc AHA 61st Annu High Blood Pressure Res Conf 152 (2007).

[112] M. Knipp, J.M. Charnock, C.D. Garner and M. Vasak Structural and functional characterization of the Zn(II) site in dimethylargininase-1 (DDAH-1) from bovine brain. Zn(II) release activates DDAH-1, J Biol Chem 276 (2001) 40449-40456.

[113] J. Leiper, J. Murray-Rust, N. McDonald and P. Vallance S-nitrosylation of dimethylarginine dimethylaminohydrolase regulates enzyme activity: further interactions between nitric oxide synthase and dimethylarginine dimethylaminohydrolase, Proc Natl Acad Sci U S A. 2002 Oct 15;99(21):13527-32. Epub 2002 Oct 7.

118

[114] M. Knipp, O. Braun, P.M. Gehrig, R. Sack and M. Vasak Zn(II)-free dimethylargininase- 1 (DDAH-1) is inhibited upon specific Cys-S-nitrosylation, J Biol Chem 278 (2003) 3410-3416.

[115] J.L. Jiang, X.H. Zhang, N.S. Li, W.Q. Rang, Y. Feng, C.P. Hu, Y.J. Li and H.W. Deng Probucol decreases asymmetrical dimethylarginine level by alternation of protein arginine methyltransferase I and dimethylarginine dimethylaminohydrolase activity, Cardiovasc Drugs Ther 20 (2006) 281-294.

[116] B. Tan, D.J. Jiang, H. Huang, S.J. Jia, J.L. Jiang, C.P. Hu and Y.J. Li Taurine protects against low-density lipoprotein-induced endothelial dysfunction by the DDAH/ADMA pathway, Vascul Pharmacol 46 (2007) 338-345.

[117] H.M. Eid, T. Lyberg, H. Arnesen and I. Seljeflot Insulin and adiponectin inhibit the TNFalpha-induced ADMA accumulation in human endothelial cells: the role of DDAH, Atherosclerosis 194 (2007) e1-8.

[118] Q.F. Yin and Y. Xiong Pravastatin restores DDAH activity and endothelium-dependent relaxation of rat aorta after exposure to glycated protein, J Cardiovasc Pharmacol 45 (2005) 525-532.

[119] D.P. Holden, J.E. Cartwright, S.S. Nussey and G.S. Whitley Estrogen stimulates dimethylarginine dimethylaminohydrolase activity and the metabolism of asymmetric dimethylarginine, Circulation 108 (2003) 1575-1580.

[120] E. Monsalve, P.J. Oviedo, M.A. Garcia-Perez, J.J. Tarin, A. Cano and C. Hermenegildo Estradiol counteracts oxidized LDL-induced asymmetric dimethylarginine production by cultured human endothelial cells, Cardiovasc Res 73 (2007) 66-72.

[121] S. Ueda, S. Kato, H. Matsuoka, M. Kimoto, S. Okuda, M. Morimatsu and T. Imaizumi Regulation of cytokine-induced nitric oxide synthesis by asymmetric dimethylarginine: role of dimethylarginine dimethylaminohydrolase, Circ Res 92 (2003) 226-233.

[122] T.L. Yang, M.F. Chen, B.L. Luo, Q.Y. Xie, J.L. Jiang and Y.J. Li Fenofibrate decreases asymmetric dimethylarginine level in cultured endothelial cells by inhibiting NF-kappaB activity, Naunyn Schmiedebergs Arch Pharmacol 371 (2005) 401-407.

[123] M. Tanaka, T. Osanai, R. Murakami, S. Sasaki, H. Tomita, N. Maeda, K. Satoh, K. Magota and K. Okumura Effect of vasoconstrictor coupling factor 6 on gene expression profile in human vascular endothelial cells: enhanced release of asymmetric dimethylarginine, J Hypertens 24 (2006) 489-497.

[124] H.Y. Xin, D.J. Jiang, S.J. Jia, K. Song, G.P. Wang, Y.J. Li and F.P. Chen Regulation by DDAH/ADMA pathway of lipopolysaccharide-induced tissue factor expression in endothelial cells, Thromb Haemost 97 (2007) 830-838.

[125] Q.F. Yin, S.H. Fu, P. He and Y. Xiong Dimethylarginine dimethylaminohydrolase inhibition and asymmetric dimethylarginine accumulation contribute to endothelial

119

dysfunction in rats exposed to glycosylated protein: effects of aminoguanidine, Atherosclerosis 190 (2007) 53-61.

[126] F. Scalera, J.T. Kielstein, J. Martens-Lobenhoffer, S.C. Postel, M. Tager and S.M. Bode- Boger Erythropoietin increases asymmetric dimethylarginine in endothelial cells: role of dimethylarginine dimethylaminohydrolase, J Am Soc Nephrol 16 (2005) 892-898.

[127] V. Sorrenti, F. Mazza, A. Campisi, L. Vanella, G. Li Volti and C. Di Giacomo High glucose-mediated imbalance of nitric oxide synthase and dimethylarginine dimethylaminohydrolase expression in endothelial cells, Curr Neurovasc Res 3 (2006) 49-54.

[128] A. Ito, P.S. Tsao, S. Adimoolam, M. Kimoto, T. Ogawa and J.P. Cooke Novel mechanism for endothelial dysfunction: dysregulation of dimethylarginine dimethylaminohydrolase, Circulation 99 (1999) 3092-3095.

[129] L. Hong and W. Fast Inhibition of human dimethylarginine dimethylaminohydrolase-1 by S-nitroso-L-homocysteine and hydrogen peroxide. Analysis, quantification, and implications for hyperhomocysteinemia, J Biol Chem 282 (2007) 34684-34692.

[130] M. Weis, T.N. Kledal, K.Y. Lin, S.N. Panchal, S.Z. Gao, H.A. Valantine, E.S. Mocarski and J.P. Cooke Cytomegalovirus infection impairs the nitric oxide synthase pathway: role of asymmetric dimethylarginine in transplant arteriosclerosis, Circulation 109 (2004) 500-505.

[131] D. Wang, P.S. Gill, T. Chabrashvili, M.L. Onozato, J. Raggio, M. Mendonca, K. Dennehy, M. Li, P. Modlinger, J. Leiper, P. Vallance, O. Adler, A. Leone, A. Tojo, W.J. Welch and C.S. Wilcox Isoform-specific regulation by N(G),N(G)-dimethylarginine dimethylaminohydrolase of rat serum asymmetric dimethylarginine and vascular endothelium-derived relaxing factor/NO, Circ Res 101 (2007) 627-635.

[132] A.J. Pope, K. Karrupiah, P.N. Kearns, Y. Xia and A.J. Cardounel Role of dimethylarginine dimethylaminohydrolases in the regulation of endothelial nitric oxide production, J Biol Chem 284 (2009) 35338-35347.

[133] C.L. Smith, G.M. Birdsey, S. Anthony, F.I. Arrigoni, J.M. Leiper and P. Vallance Dimethylarginine dimethylaminohydrolase activity modulates ADMA levels, VEGF expression, and cell phenotype, Biochem Biophys Res Commun 308 (2003) 984-989.

[134] K. Hasegawa, S. Wakino, T. Tanaka, M. Kimoto, S. Tatematsu, T. Kanda, K. Yoshioka, K. Homma, N. Sugano, M. Kurabayashi, T. Saruta and K. Hayashi Dimethylarginine dimethylaminohydrolase 2 increases vascular endothelial growth factor expression through Sp1 transcription factor in endothelial cells, Arterioscler Thromb Vasc Biol 26 (2006) 1488-1494.

[135] H. Tokuo, S. Yunoue, L. Feng, M. Kimoto, H. Tsuji, T. Ono, H. Saya and N. Araki Phosphorylation of neurofibromin by cAMP-dependent protein kinase is regulated via a

120

cellular association of N(G),N(G)-dimethylarginine dimethylaminohydrolase, FEBS Lett 494 (2001) 48-53.

[136] V.P. Valkonen, T.P. Tuomainen and R. Laaksonen DDAH gene and cardiovascular risk, Vasc Med 10 Suppl 1 (2005) S45-48.

[137] R.A. Breckenridge, P. Kelly, M. Nandi, P.J. Vallance, T.J. Ohun and J. Leiper A role for Dimethylarginine Dimethylaminohydrolase 1 (DDAH1) in mammalian development, Int J Dev Biol 54 215-220.

[138] T. Morishita, M. Tsutsui, H. Shimokawa, K. Sabanai, H. Tasaki, O. Suda, S. Nakata, A. Tanimoto, K.Y. Wang, Y. Ueta, Y. Sasaguri, Y. Nakashima and N. Yanagihara Nephrogenic diabetes insipidus in mice lacking all nitric oxide synthase isoforms, Proc Natl Acad Sci U S A 102 (2005) 10616-10621.

[139] T.M. Lu, Y.A. Ding, S.J. Lin, W.S. Lee and H.C. Tai Plasma levels of asymmetrical dimethylarginine and adverse cardiovascular events after percutaneous coronary intervention, Eur Heart J 24 (2003) 1912-1919.

[140] J.R. Chan, R.H. Boger, S.M. Bode-Boger, O. Tangphao, P.S. Tsao, T.F. Blaschke and J.P. Cooke Asymmetric dimethylarginine increases mononuclear cell adhesiveness in hypercholesterolemic humans, Arterioscler Thromb Vasc Biol 20 (2000) 1040-1046.

[141] H. Azuma, J. Sato, H. Hamasaki, A. Sugimoto, E. Isotani and S. Obayashi Accumulation of endogenous inhibitors for nitric oxide synthesis and decreased content of L-arginine in regenerated endothelial cells, Br J Pharmacol 115 (1995) 1001-1004.

[142] D. Zakrzewicz and O. Eickelberg From arginine methylation to ADMA: a novel mechanism with therapeutic potential in chronic lung diseases, BMC Pulm Med 9 (2009) 5.

[143] C. Zoccali, R. Maas, S. Cutrupi, P. Pizzini, P. Finocchiaro, F. Cambareri, V. Panuccio, C. Martorano, F. Schulze, G. Enia, G. Tripepi and R. Boger Asymmetric dimethyl-arginine (ADMA) response to inflammation in acute infections, Nephrol Dial Transplant 22 (2007) 801-806.

[144] P. Vallance and J. Leiper Asymmetric dimethylarginine and kidney disease--marker or mediator? J Am Soc Nephrol 16 (2005) 2254-2256.

[145] R. Maas, F. Schulze, J. Baumert, H. Lowel, K. Hamraz, E. Schwedhelm, W. Koenig and R.H. Boger Asymmetric dimethylarginine, smoking, and risk of coronary heart disease in apparently healthy men: prospective analysis from the population-based Monitoring of Trends and Determinants in Cardiovascular Disease/Kooperative Gesundheitsforschung in der Region Augsburg study and experimental data, Clin Chem 53 (2007) 693-701.

[146] W. Aji, S. Ravalli, M. Szabolcs, X.C. Jiang, R.R. Sciacca, R.E. Michler and P.J. Cannon L-arginine prevents xanthoma development and inhibits atherosclerosis in LDL receptor knockout mice, Circulation 95 (1997) 430-437.

121

[147] R.H. Boger, S.M. Bode-Boger, R.P. Brandes, L. Phivthong-ngam, M. Bohme, R. Nafe, A. Mugge and J.C. Frolich Dietary L-arginine reduces the progression of atherosclerosis in cholesterol-fed rabbits: comparison with lovastatin, Circulation 96 (1997) 1282-1290.

[148] X.J. Girerd, A.T. Hirsch, J.P. Cooke, V.J. Dzau and M.A. Creager L-arginine augments endothelium-dependent vasodilation in cholesterol-fed rabbits, Circ Res 67 (1990) 1301- 1308.

[149] Z. Wu, X. Qian, D. Cai, Y. Liu, Z. Li and J. He Long-term oral administration of L- arginine enhances endothelium-dependent vasorelaxation and inhibits neointimal thickening after endothelial denudation in rats, Chin Med J (Engl) 109 (1996) 592-598.

[150] A.J. Cardounel and J.L. Zweier Endogenous methylarginines regulate neuronal nitric- oxide synthase and prevent excitotoxic injury, J Biol Chem 277 (2002) 33995-34002. Epub 32002 Jun 33928.

[151] R.H. Boger Asymmetric dimethylarginine, an endogenous inhibitor of nitric oxide synthase, explains the "L-arginine paradox" and acts as a novel cardiovascular risk factor, J Nutr 134 (2004) 2842S-2847S; discussion 2853S.

[152] J.P. Cooke and R.K. Oka Atherogenesis and the arginine hypothesis, Curr Atheroscler Rep 3 (2001) 252-259.

[153] J. Leiper and P. Vallance Biological significance of endogenous methylarginines that inhibit nitric oxide synthases, Cardiovasc Res 43 (1999) 542-548.

[154] D. Tsikas, R.H. Boger, J. Sandmann, S.M. Bode-Boger and J.C. Frolich Endogenous nitric oxide synthase inhibitors are responsible for the L-arginine paradox, FEBS Lett 478 (2000) 1-3.

[155] R.H. Boger, S.M. Bode-Boger, K. Sydow, D.D. Heistad and S.R. Lentz Plasma concentration of asymmetric dimethylarginine, an endogenous inhibitor of nitric oxide synthase, is elevated in monkeys with hyperhomocyst(e)inemia or hypercholesterolemia, Arterioscler Thromb Vasc Biol 20 (2000) 1557-1564.

[156] R.H. Boger, S.M. Bode-Boger, A. Szuba, P.S. Tsao, J.R. Chan, O. Tangphao, T.F. Blaschke and J.P. Cooke Asymmetric dimethylarginine (ADMA): a novel risk factor for endothelial dysfunction: its role in hypercholesterolemia, Circulation 98 (1998) 1842- 1847.

[157] H. Lenzen, D. Tsikas and R.H. Boger Asymmetric dimethylarginine (ADMA) and the risk for coronary heart disease: the multicenter CARDIAC study, Eur J Clin Pharmacol 62 Suppl 13 (2006) 45-49.

[158] K.Y. Lin, A. Ito, T. Asagami, P.S. Tsao, S. Adimoolam, M. Kimoto, H. Tsuji, G.M. Reaven and J.P. Cooke Impaired nitric oxide synthase pathway in diabetes mellitus: role of asymmetric dimethylarginine and dimethylarginine dimethylaminohydrolase, Circulation 106 (2002) 987-992.

122

[159] J. Leiper, M. Nandi, B. Torondel, J. Murray-Rust, M. Malaki, B. O'Hara, S. Rossiter, S. Anthony, M. Madhani, D. Selwood, C. Smith, B. Wojciak-Stothard, A. Rudiger, R. Stidwill, N.Q. McDonald and P. Vallance Disruption of methylarginine metabolism impairs vascular homeostasis, Nat Med 13 (2007) 198-203.

[160] L.J. Druhan, S.P. Forbes, A.J. Pope, C.A. Chen, J.L. Zweier and A.J. Cardounel Regulation of eNOS-derived superoxide by endogenous methylarginines, Biochemistry 47 (2008) 7256-7263.

[161] T.L. Vanden Hoek, L.B. Becker, Z. Shao, C. Li and P.T. Schumacker Reactive oxygen species released from mitochondria during brief hypoxia induce preconditioning in cardiomyocytes, J Biol Chem 273 (1998) 18092-18098.

[162] M.W. Radomski and S. Moncada Regulation of vascular homeostasis by nitric oxide, Thromb Haemost 70 (1993) 36-41.

[163] R. Guerra, Jr., A.F. Brotherton, P.J. Goodwin, C.R. Clark, M.L. Armstrong and D.G. Harrison Mechanisms of abnormal endothelium-dependent vascular relaxation in atherosclerosis: implications for altered autocrine and paracrine functions of EDRF, Blood Vessels 26 (1989) 300-314.

[164] K.Y. Stokes, D. Cooper, A. Tailor and D.N. Granger Hypercholesterolemia promotes inflammation and microvascular dysfunction: role of nitric oxide and superoxide, Free Radic Biol Med 33 (2002) 1026-1036.

[165] C. Napoli and L.J. Ignarro Nitric oxide and atherosclerosis, Nitric Oxide 5 (2001) 88-97.

[166] H. Miyazaki, H. Matsuoka, J.P. Cooke, M. Usui, S. Ueda, S. Okuda and T. Imaizumi Endogenous nitric oxide synthase inhibitor: a novel marker of atherosclerosis, Circulation 99 (1999) 1141-1146.

[167] P.S. Tsao and J.P. Cooke Endothelial alterations in hypercholesterolemia: more than simply vasodilator dysfunction, J Cardiovasc Pharmacol 32 Suppl 3 (1998) S48-53.

[168] Y. Chen, Y. Li, P. Zhang, J.H. Traverse, M. Hou, X. Xu, M. Kimoto and R.J. Bache Dimethylarginine dimethylaminohydrolase and endothelial dysfunction in failing hearts, Am J Physiol Heart Circ Physiol. 2005 Nov;289(5):H2212-9. Epub 2005 Jul 15.

[169] C.T. Tran, J.M. Leiper and P. Vallance The DDAH/ADMA/NOS pathway, Atheroscler Suppl 4 (2003) 33-40.

[170] J. Leiper and P. Vallance Biological significance of endogenous methylarginines that inhibit nitric oxide synthases, Cardiovasc Res. 1999 Aug 15;43(3):542-8.

[171] P.V. Usatyuk, N.L. Parinandi and V. Natarajan Redox regulation of 4-hydroxy-2- nonenal-mediated endothelial barrier dysfunction by focal adhesion, adherens, and tight junction proteins, J Biol Chem. 2006 Nov 17;281(46):35554-66. Epub 2006 Sep 17.

123

[172] H. Esterbauer, R.J. Schaur and H. Zollner Chemistry and biochemistry of 4- hydroxynonenal, malonaldehyde and related aldehydes, Free Radic Biol Med. 1991;11(1):81-128.

[173] K. Uchida Role of reactive aldehyde in cardiovascular diseases, Free Radic Biol Med. 2000 Jun 15;28(12):1685-96.

[174] K. Uchida 4-Hydroxy-2-nonenal: a product and mediator of oxidative stress, Prog Lipid Res. 2003 Jul;42(4):318-43.

[175] M. Knipp and M. Vasak A colorimetric 96-well microtiter plate assay for the determination of enzymatically formed citrulline, Anal Biochem. 2000 Nov 15;286(2):257-64.

[176] T. Ogawa, M. Kimoto and K. Sasaoka Purification and properties of a new enzyme, NG,NG-dimethylarginine dimethylaminohydrolase, from rat kidney, J Biol Chem 264 (1989) 10205-10209.

[177] J.F. Keaney, Jr., A. Xu, D. Cunningham, T. Jackson, B. Frei and J.A. Vita Dietary probucol preserves endothelial function in cholesterol-fed rabbits by limiting vascular oxidative stress and superoxide generation, J Clin Invest 95 (1995) 2520-2529.

[178] S. Yla-Herttuala, W. Palinski, M.E. Rosenfeld, S. Parthasarathy, T.E. Carew, S. Butler, J.L. Witztum and D. Steinberg Evidence for the presence of oxidatively modified low density lipoprotein in atherosclerotic lesions of rabbit and man, J Clin Invest 84 (1989) 1086-1095.

[179] D. Frey, O. Braun, C. Briand, M. Vasak and M.G. Grutter Structure of the mammalian NOS regulator dimethylarginine dimethylaminohydrolase: A basis for the design of specific inhibitors, Structure 14 (2006) 901-911.

[180] E.M. Stone, A.L. Costello, D.L. Tierney and W. Fast Substrate-assisted cysteine deprotonation in the mechanism of dimethylargininase (DDAH) from Pseudomonas aeruginosa, Biochemistry 45 (2006) 5618-5630.

[181] W.P. Arnold, C.K. Mittal, S. Katsuki and F. Murad Nitric oxide activates guanylate cyclase and increases guanosine 3':5'-cyclic monophosphate levels in various tissue preparations, Proc Natl Acad Sci U S A 74 (1977) 3203-3207.

[182] D.Y. Gruetter, C.A. Gruetter, B.K. Barry, W.H. Baricos, A.L. Hyman, P.J. Kadowitz and L.J. Ignarro Activation of coronary arterial guanylate cyclase by nitric oxide, nitroprusside, and nitrosoguanidine--inhibition by calcium, lanthanum, and other cations, enhancement by thiols, Biochem Pharmacol 29 (1980) 2943-2950.

[183] W. Martin, G.M. Villani, D. Jothianandan and R.F. Furchgott Selective blockade of endothelium-dependent and glyceryl trinitrate-induced relaxation by hemoglobin and by methylene blue in the rabbit aorta, J Pharmacol Exp Ther 232 (1985) 708-716.

124

[184] M. Hecker, D.T. Walsh and J.R. Vane On the substrate specificity of nitric oxide synthase, FEBS Lett 294 (1991) 221-224.

[185] S.P. Janssens, A. Shimouchi, T. Quertermous, D.B. Bloch and K.D. Bloch Cloning and expression of a cDNA encoding human endothelium-derived relaxing factor/nitric oxide synthase, J Biol Chem 267 (1992) 14519-14522.

[186] D. Frey, O. Braun, C. Briand, M. Vasak and M.G. Grutter Structure of the mammalian NOS regulator dimethylarginine dimethylaminohydrolase: A basis for the design of specific inhibitors, Structure 14 (2006) 901-911.

[187] E.M. Stone, A.L. Costello, D.L. Tierney and W. Fast Substrate-assisted cysteine deprotonation in the mechanism of dimethylargininase (DDAH) from Pseudomonas aeruginosa, Biochemistry 45 (2006) 5618-5630.

[188] E.M. Stone, M.D. Person, N.J. Costello and W. Fast Characterization of a transient covalent adduct formed during dimethylarginine dimethylaminohydrolase catalysis, Biochemistry 44 (2005) 7069-7078.

[189] M. Kimoto, S. Miyatake, T. Sasagawa, H. Yamashita, M. Okita, T. Oka, T. Ogawa and H. Tsuji Purification, cDNA cloning and expression of human NG,NG-dimethylarginine dimethylaminohydrolase, Eur J Biochem 258 (1998) 863-868.

[190] T. Ogawa, M. Kimoto and K. Sasaoka Purification and properties of a new enzyme, NG,NG-dimethylarginine dimethylaminohydrolase, from rat kidney, J Biol Chem 264 (1989) 10205-10209.

[191] M. Knipp, J.M. Charnock, C.D. Garner and M. Vasak Structural and functional characterization of the Zn(II) site in dimethylargininase-1 (DDAH-1) from bovine brain. Zn(II) release activates DDAH-1, J Biol Chem 276 (2001) 40449-40456. Epub 42001 Aug 40423.

[192] J.L. Zweier, P. Wang, A. Samouilov and P. Kuppusamy Enzyme-independent formation of nitric oxide in biological tissues, Nat Med 1 (1995) 804-809.

[193] J.L. Jiang, X.H. Zhang, N.S. Li, W.Q. Rang, Y. Feng, C.P. Hu, Y.J. Li and H.W. Deng Probucol decreases asymmetrical dimethylarginine level by alternation of protein arginine methyltransferase I and dimethylarginine dimethylaminohydrolase activity, Cardiovasc Drugs Ther 20 (2006) 281-294.

[194] G. Jurgens, Q. Chen, H. Esterbauer, S. Mair, G. Ledinski and H.P. Dinges Immunostaining of human autopsy aortas with antibodies to modified apolipoprotein B and apoprotein(a), Arterioscler Thromb 13 (1993) 1689-1699.

[195] P.V. Usatyuk, N.L. Parinandi and V. Natarajan Redox regulation of 4-hydroxy-2- nonenal-mediated endothelial barrier dysfunction by focal adhesion, adherens, and tight junction proteins, J Biol Chem 281 (2006) 35554-35566. Epub 32006 Sep 35517.

125

[196] A. Bennaars-Eiden, L. Higgins, A.V. Hertzel, R.J. Kapphahn, D.A. Ferrington and D.A. Bernlohr Covalent modification of epithelial fatty acid-binding protein by 4- hydroxynonenal in vitro and in vivo. Evidence for a role in antioxidant biology, J Biol Chem 277 (2002) 50693-50702. Epub 52002 Oct 50616.

[197] B.P. Sampey, D.L. Carbone, J.A. Doorn, D.A. Drechsel and D.R. Petersen 4-Hydroxy-2- nonenal adduction of extracellular signal-regulated kinase (Erk) and the inhibition of hepatocyte Erk-Est-like protein-1-activating protein-1 signal transduction, Mol Pharmacol 71 (2007) 871-883. Epub 2006 Dec 2012.

[198] S.J. Siegel, J. Bieschke, E.T. Powers and J.W. Kelly The oxidative stress metabolite 4- hydroxynonenal promotes Alzheimer protofibril formation, Biochemistry 46 (2007) 1503-1510.

[199] T.M. Griffith, D.H. Edwards, M.J. Lewis, A.C. Newby and A.H. Henderson The nature of endothelium-derived vascular relaxant factor, Nature 308 (1984) 645-647.

[200] R.H. Boger, S.R. Lentz, S.M. Bode-Boger, H.R. Knapp and W.G. Haynes Elevation of asymmetrical dimethylarginine may mediate endothelial dysfunction during experimental hyperhomocyst(e)inaemia in humans, Clin Sci (Lond) 100 (2001) 161-167.

[201] J. Jacobi, K. Sydow, G. von Degenfeld, Y. Zhang, H. Dayoub, B. Wang, A.J. Patterson, M. Kimoto, H.M. Blau and J.P. Cooke Overexpression of dimethylarginine dimethylaminohydrolase reduces tissue asymmetric dimethylarginine levels and enhances angiogenesis, Circulation 111 (2005) 1431-1438.

[202] H. Dayoub, R.N. Rodionov, C. Lynch, J.P. Cooke, E. Arning, T. Bottiglieri, S.R. Lentz and F.M. Faraci Overexpression of dimethylarginine dimethylaminohydrolase inhibits asymmetric dimethylarginine-induced endothelial dysfunction in the cerebral circulation, Stroke 39 (2008) 180-184.

[203] K. Sydow, C.E. Mondon, J. Schrader, H. Konishi and J.P. Cooke Dimethylarginine dimethylaminohydrolase overexpression enhances insulin sensitivity, Arterioscler Thromb Vasc Biol 28 (2008) 692-697.

[204] S. Dayal, R.N. Rodionov, E. Arning, T. Bottiglieri, M. Kimoto, D.J. Murry, J.P. Cooke, F.M. Faraci and S.R. Lentz Tissue-specific downregulation of dimethylarginine dimethylaminohydrolase in hyperhomocysteinemia, Am J Physiol Heart Circ Physiol 295 (2008) H816-825.

[205] M.C. Stuhlinger, E. Conci, B.J. Haubner, E.M. Stocker, J. Schwaighofer, J.P. Cooke, P.S. Tsao, O. Pachinger and B. Metzler Asymmetric dimethyl L-arginine (ADMA) is a critical regulator of myocardial reperfusion injury, Cardiovasc Res 75 (2007) 417-425.

[206] S.P. Forbes, L.J. Druhan, J.E. Guzman, N. Parinandi, L. Zhang, K.B. Green-Church and A.J. Cardounel Mechanism of 4-HNE mediated inhibition of hDDAH-1: implications in no regulation, Biochemistry 47 (2008) 1819-1826.

126

[207] J.M. Sasser, N.C. Moningka, M.W. Cunningham, Jr., B.P. Croker and C. Baylis Asymmetric Dimethylarginine in Angiotensin II Induced Hypertension, Am J Physiol Regul Integr Comp Physiol (2009).

[208] S.R. Jaffrey and S.H. Snyder The biotin switch method for the detection of S-nitrosylated proteins, Sci STKE 2001 (2001) PL1.

[209] S.S. Wolf The protein arginine methyltransferase family: an update about function, new perspectives and the physiological role in humans, Cell Mol Life Sci 66 (2009) 2109- 2121.

[210] A.J. Pope, K. Karuppiah and A.J. Cardounel Role of the PRMT-DDAH-ADMA axis in the regulation of endothelial nitric oxide production, Pharmacol Res 60 (2009) 461-465.

127

BIOGRAPHICAL SKETCH

Scott Preston Forbes was born in 1981 in Morgantown, WV. He graduated with honors

from Morgantown High School in 2000. He then attended West Virginina University and

graduated cum laude in 2004 with a B.S. in biology. In June of 2005, he enrolled in the

Integrated Biomedical Science Graduate Program at The Ohio State University College of

Medicine. In January of 2006, he joined the laboratory of Dr. AJ Cardounel. In June of 2007, he relocated with his mentor to the University of Florida and joined the Interdisciplinary Program in

Biomedical Sciences where he obtained his doctorate of philosophy in May of 2010. During his graduate education he received a pre-doctoral fellowship award from the American Heart

Association. Scott has been the first author on two publications and co-author on two publications. In addition, he has presented his work at several conferences

128