Investigating the Putative Mechanism and Functional Role of RGS5 Upregulation in Vascular Smooth Muscle Cells

Following Statin Treatment

by

Joobin Sattar

A thesis submitted in conformity with the requirements for the degree of Master of Science

Department of Physiology

University of Toronto

© Copyright by Joobin Sattar (2015)

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Investigating the Mechanism and Putative Functional Role of RGS5 Upregulation in Vascular Smooth Muscle Cells Following

Statin Treatment

Joobin Sattar

Master of Science

Department of Physiology University of Toronto

2015

Abstract

RGS5 within vascular smooth muscle cells (VSMCs) is capable of inhibiting G-

protein signaling involved in VSMC recruitment. Based on the suggested ability of statin

to upregulate RGS5, we investigated whether RGS5 could mediate the pleiotropic effects

of statins in VSMCs. Fluvastatin treatment of isolated VSMCs significantly increased the

expression of RGS5, while downregulating RGS2, RGS3, and RGS16. However, these

results were not observed ex vivo or in vivo within the aorta following fluvastatin

treatment. Fluvastatin also upregulated PPARδ and PPARγ, demonstrated transcriptional

regulators of RGS5 expression, in VSMCs. However, PPARδ and PPARγ antagonists did

not block fluvastatin-induced RGS5 upregulation and their agonists did not increase

RGS5 expression. Lastly, fluvastatin did not significantly reduce neointimal hyperplasia

following femoral artery injury in RGS5 WT and KO mice. These results suggest that

statins robustly upregulate RGS5 only in cultured synthetic VSMCs, with the

physiological role of this pleiotropic effect remaining to be elucidated.

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Acknowledgements

First and foremost, I want to express my sincere gratitude to my supervisor and

mentor for the past four years, Dr. Scott Heximer. You have patiently guided me from the

very beginning and allowed me to mature as a researcher as well as develop skills and

knowledge that will be important for the rest of my life. I really appreciate how

understanding you have been with me, especially with my incredibly loud voice. You

cultivated my passion in physiology both during my undergrad and masters, and I truly

could not ask for a better supervisor.

I would also like to thank both of my committee members, Dr. Steffen-Sebastian

Bolz and Dr. Peter Backx. Continuing from my undergraduate studies, you have both

guided me with your valuable insights, comments, suggestions, and continual support.

Your inputs were crucial to my development as a learner and future scholar. I also greatly

appreciate the accommodation and understanding that both of you have shown during

your international affairs.

I believe the Heximer lab is truly a family. To all of my lab mates, namely Joey,

Guillaume, Steph, and all the undergrads, thank you for making my time in the lab so

enjoyable. Whether it was fixing lab equipment with Joey, playing GAP-a-ball, or

discussing the excellence of the French culture, you guys made the lab a place that I did

not want to leave. No matter where I end up, I am sure we will maintain our close

friendships.

But our lab family would not function without the one and only Dr. Jenny Zhang,

helping and guiding us all. Jenny, thank you so much for all your guidance throughout

my research. Your exceptional surgical capabilities and various other dynamic skills have

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been of tremendous help to me. Specifically, I would like to thank you for all the

assistance and collaboration in our femoral artery injury experiments, especially

considering its level of time commitment.

Lastly, I would like to thank all of those who have supported me throughout my

research outside of the lab. To my parents, JJ, and close friends, I cannot express my

immense gratitude to your understanding, support, and encouragement, even during my

moments of frustration. I could not have done this without you and will never forget it!

GO RGS!!!

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Table of Contents

Abstract ............................................................................................................................... ii Acknowledgements ............................................................................................................ iii Table of Contents .................................................................................................................v List of Tables .................................................................................................................... vii List of Figures .................................................................................................................. viii List of abbreviations .......................................................................................................... ix 1. Introduction ....................................................................................................................1

1.1 Atherosclerosis ...................................................................................................1 1.1.1 Atherosclerosis Initiation ....................................................................1 1.1.2 Atherosclerosis Progression ................................................................2 1.1.3 Atherosclerosis Complications ...........................................................3 1.1.4 Treatment of Atherosclerosis ..............................................................5

1.2 G-Protein Signaling ...........................................................................................7 1.2.1 Heterotrimeric G-Protein Signaling Mechanism ................................8 1.2.2 RGS Proteins .....................................................................................10 1.2.3 RGS5 Function & Localization ........................................................13

1.3 Statins ...............................................................................................................20 1.3.1 Comparison of Different Types of Statins ........................................21 1.3.2 Pleiotropic Effects of Statins ............................................................22 1.3.3 Cardiovascular Pleiotropic Effects of Statins ...................................23

2. Rationale .......................................................................................................................26 3. Hypothesis and Objectives ..........................................................................................27 4. Materials and Methods ................................................................................................29

4.1 Experimental Animals .....................................................................................29 4.2 Isolation and Culturing of Primary VSMCs ....................................................29 4.3 Dose Response and Time Profile Analysis of Fluvastatin Treatment .............32 4.4 Ex Vivo Analysis of Fluvastatin Treatment .....................................................32 4.5 In Vivo Analysis of Fluvastatin Treatment ......................................................33 4.6 qRT-PCR Reaction and Analysis .....................................................................34 4.7 Femoral Artery Injury Model ...........................................................................36

4.7.1 Fluvastatin Treatment and Induction of Femoral Artery Injury .......36 4.7.2 Femoral Artery Isolation ...................................................................37

4.7.3 Femoral Artery Histological Analysis ..............................................38 4.8 Statistical Analysis ...........................................................................................38

5. Results ...........................................................................................................................40 5.1 Analysis of RGS Protein mRNA Expression in Cultured VSMCs Treated with Fluvastatin ..............................................................................................................40

5.1.1 Dose Response Analysis of RGS Protein Expression .......................40 5.1.2 Kinetic Analysis of RGS5 Upregulation Following Fluvastatin Treatment ...................................................................................................41

5.2 Analysis of RGS5 mRNA Expression in Isolated Vessels Treated with Fluvastatin ..............................................................................................................44 5.3 Effect of Fluvastatin on Aortic RGS Protein mRNA Expression In Vivo .......46

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5.4 The Putative Role of PPARs in Fluvastatin-Induced RGS5 Upregulation in Cultured VSMCs ...................................................................................................48

5.4.1 Changes in the Expression of PPAR Isoforms Following Fluvastatin Treatment ...................................................................................................48 5.4.2 The Effect of PPARδ and PPARγ Agonists and Antagonists on RGS5 Expression .......................................................................................49

5.5 The Potential Role of Fluvastatin-Induced RGS5 Upregulation on Neointimal Hyperplasia ............................................................................................................53

5.4.1 Body Weight and Tibia Length Analysis ..........................................54 5.4.2 The Role of RGS5 on Neointimal Hyperplasia Following Fluvastatin Treatment ...................................................................................................54

6. Discussion .....................................................................................................................59 6.1 Modulation of RGS5 Expression by Fluvastatin .............................................59 6.2 The Role of PPARs on Statin-Induced Upregulation of RGS5 .......................65 6.3 Investigating the Physiological Role of Fluvastatin-Induced RGS5 Upregulation ..........................................................................................................68 6.4 Limitations and Future Directions ...................................................................72 6.5 Conclusions and Significance ..........................................................................75

7. References .....................................................................................................................76

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List of Tables

4. Materials and Methods

Table 4-1. Primers used for qRT-PCR reactions ...............................................................39

5. Results

Table 5-1. Analysis of neointimal size in RGS5 WT and KO mice treated with vehicle or

fluvastatin ...........................................................................................................................58

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List of Figures

1. Introduction

Figure 1-1. Heterotrimeric G-Protein signaling and the role of RGS proteins .................19

Figure 1-2. The effect of statins on cholesterol synthesis and isoprenylation ..................25

3. Hypothesis and Objectives

Figure 3-1. Proposed mechanism and role of RGS5 upregulation following statin

treatment in VSMCs ..........................................................................................................28

4. Materials and Methods

Figure 4-1. Representative image of cultured VSMCs .....................................................31

5. Results

Figure 5-1. Relative fluvastatin-induced changes in the mRNA expression of R4 subfamily of RGS proteins in VSMCs ..............................................................................42 Figure 5-2. The time profile of fluvastatin-induced upregulation of RGS5 .....................43

Figure 5-3. The ex vivo effect of fluvastatin on RGS5 mRNA expression in isolated aortas ..................................................................................................................................45 Figure 5-4. The in vivo effect of fluvastatin on the mRNA expression of R4 RGS proteins ...............................................................................................................................47 Figure 5-5. The regulation of PPARs by fluvastatin in cultured VSMCs ........................51

Figure 5-6. The effect of PPARγ and PPARδ agonists and antagonists on RGS5 mRNA expression ..........................................................................................................................52 Figure 5-7. Body size analysis of mice on femoral artery injury protocol .......................56

Figure 5-8. Representative images of control and injured femoral arteries ......................57

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List of Abbreviations

ApoE Apolipoprotein E

AngII Angiotensin II

AT1 Angiotensin II Receptor Type 1

CABG Coronary Artery Bypass Graft

cDNA Complementary Deoxyribonucleic Acid

dKO Double Knock-Out

DMSO Dimethyl Sulfoxide

EC Endothelial Cell

ECM Extracellular Matrix

EEL External Elastic Lamina

ERK Extracellular Signal-Regulated Kinase

ET-1 Endothelin-1

EVG Elastica Van Gieson

GAP GTPase-Activating Protein

GAPDH Glyceraldehyde 3-Phosphate Dehydrogenase

GDP Guanosine Diphosphate

GPCR G-Protein Coupled Receptor

GTP Guanosine Triphosphate

HDL High-Density Lipoprotein

HMG-CoA 3-hydroxy-3-methylglutaryl-coenzyme A

IEL Internal Elastic Lamina

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KO Knock-Out

LDL Low-Density Lipoprotein

MAPK Mitogen-Activated Protein Kinase

M-CSF Macrophage Colony-Stimulating Factor

MMP Matrix Metalloproteinase

PDGF-BB Platelet-Derived Growth Factor-BB

PBS Phosphate-Buffered Saline

PPAR Peroxisome Proliferator-Activated Receptor

qRT-PCR Quantitative Real-Time Polymerase Chain Reaction

RGS Regulator of G-Protein Signaling

RTK Receptor Tyrosine Kinase

S1P Sphingosine-1-Phosphate

VSMC Vascular Smooth Muscle Cell

VCAM Vascular Cell Adhesion Molecule

WT Wild-Type

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1. Introduction

1.1 Atherosclerosis

Atherosclerosis is a chronic vascular disease that remains the leading cause of

mortality in both developed and developing countries, responsible for various

cardiovascular complications including stroke and myocardial infarction [1]. During

recent years, the increasing prevalence of several risk factors for atherosclerosis,

including diabetes and an increasingly aged population, has further exacerbated

healthcare issues related to this disease [2, 3]. Moreover, due to the complexity of the

pathophysiology of atherosclerosis, the treatment options currently available remain

limited [4]. This conundrum will mean health care teams will face severe challenges in

the future, with an accompanying increased burden on healthcare spending.

1.1.1 Atherosclerosis Initiation

Atherosclerosis is characterized by thickening of the intimal region of large

arteries in the form of an atherosclerotic lesion. Although the pathophysiology of

atherosclerosis is not fully understood, the process is thought to be initiated by the

accumulation of oxidized low-density lipoprotein (LDL) particles underneath the

endothelial lining of arterial walls [5]. Once in the subendothelial region, LDL particles

are also more prone to further pathogenic modifications, including oxidation, due to

interactions with the extracellular matrix (ECM) and exposure to oxidative byproducts

from vascular cells [6]. This accumulation of LDL triggers endothelial cells (ECs) to

induce an inflammatory response via production of adhesion molecules and pro-

1  

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inflammatory chemokines and cytokines, including selectins, vascular cell adhesion

molecule 1 (VCAM-1), and macrophage colony-stimulating factor (M-CSF) [7-9].

These changes allow circulating monocytes to bind to the endothelium, migrate

into the subendothelial region, and differentiate into macrophages. With their expression

of scavenger receptors, macrophages begin to engulf the oxidized LDL particles,

removing them from the intimal region [10]. High density lipoprotein (HDL) particles

produced in the liver are thought to remove cholesterol from macrophages, and are thus

believed to counteract the effects of LDL accumulation in the arterial wall [11].

However, with excessive deposition and phagocytosis of oxidized LDL, cholesterol esters

begin to aggregate within macrophages, which are incapable of eliminating them at a

corresponding rate, thus forming lipid-filled foams cells. The accumulation of these foam

cells produces fatty streaks in the subendothelial space, which precedes the formation of

an atheroma [12].

1.1.2 Atherosclerosis Progression

The developing foam cells have been shown to aggregate, forming the

atheromatous core of the plaque. Over time, many foam cells rupture, primarily due to

apoptosis, releasing intracellular lipid and cellular contents [13]. This initiates the

formation of the necrotic core within the atheroma, a highly inflammatory

microenvironment of the plaque. This core grows over time due to continuous formation

and rupture of surrounding foam cells, leading to exacerbation of the pathology. Around

this core is the shoulder region, which is characterized by accumulation of T-cells, mast

cells, and additional macrophages [1, 10].

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During atherosclerosis, inflammatory cells, including foam cells and T-cells, as

well as ECs secrete a variety of cytokines and growth factors, including Angiotensin II

(AngII), endothelin-1 (ET-1), and platelet-derived growth factor-BB (PDGF-BB).

Among the various effects exerted by such factors is vascular smooth muscle cell

(VSMC) recruitment, a process that I will focus on as part of this thesis [14, 15]. These

cytokines and growth factors cause VSMCs within the tunica media to switch from a

quiescent contractile phenotype to a synthetic phenotype, capable of migrating along

chemokine gradients to the sites of atheroma formation and contributing to atheroma

volume via proliferation [16].

The synthetic VSMCs can be found within the atheroma core, and can uptake

lipids, forming VSMC-derived foam cells [17]. These cells can in turn rupture similar to

those derived from macrophages, further contributing to the necrotic core and

atherosclerosis progression [18]. However, VSMCs are also localized to the shoulder

region of the plaque as well as on the forming fibrous "cap", which separates the

atheroma core from the arterial lumen. These VSMCs are crucial as they secrete a large

quantity of collagen and other ECM proteins. This forms a layer of connective tissue,

resulting in a protective fibrous cap over the atheroma and preventing exposure of plaque

content to the blood [15].

1.1.3 Complication of Atherosclerosis

Initially as the plaque grows in size, the enlargement may extrude towards the

tunica adventitia, causing a bulge on the exterior of the artery that may not alter blood

flow. However, the progression of plaque enlargement may also cause it to protrude into

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the interior of the vessel, causing narrowing of the lumen, a process known as stenosis of

the artery. This stenosis can also result from repeated rupture of the plaque, leading to

recurrent remodeling and narrowing of the lumen [5]. Stenosis is a progressive and

gradual pathology that can remain asymptomatic for decades. However, advanced lesions

may reach critical stenosis, in which the limit of blood flow to tissues downstream of the

plaque is severe enough to cause ischemia and damage to tissues [19].

Atherosclerotic plaques are also susceptible to rupture, exposing atheroma content

into the blood. The lipids, collagen, inflammatory cells, as well as necrotic tissue within

the plaque are highly thrombogenic, causing activation of the coagulation system. This

leads to the formation of the thrombus that can become dislodged in downstream

vasculature, leading to ischemia of corresponding tissues [20]. However, plaque

disruption has a spectrum of severity, ranging from clinically silent microscopic thrombi

present on the plaque surface to large dislodged thrombi formed from deep intimal tears

[21].

The risks of rupture of the plaque, as well as its severity, depend on several

factors. The high arterial blood pressure places a mechanical strain on stenotic lumens in

patients with atherosclerosis. This pressure, especially if elevated due to acute stress or

hypertension, can be a trigger for plaque rupture [22]. However, the primary determinant

of plaque rupture has been demonstrated to be the composition of the plaque. Stable

plaques have abundant VSMC and ECM content, making a primarily fibrotic core with a

thick calcified fibrous cap, which helps maintain the integrity of the plaque. In unstable

plaques that are prone to rupture, the core of the atheroma is primarily composed of lipids

and necrotic tissue surrounded by foam cells, with a thin susceptible fibrous cap.

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Macrophages and foam cells also secrete matrix metalloproteinase (MMP), enzymes that

degrade ECM produced by VSMCs and other cells, exacerbating the susceptibility of the

plaque to rupture [21, 23].

1.1.4 Treatment of Atherosclerosis

The primary modifiable factors of atherosclerosis are diet and daily habits,

including exercise and smoking [24]. Despite its prevalence and the extensive research

into its pathophysiology, the clinical treatments of atherosclerosis remain limited and

primarily targeted towards alleviating symptoms and managing complications. Better

treatment of atherosclerosis will require further understanding of the molecular and

cellular mechanisms mediating the progression of the disease. With these mechanistic

insights, future treatments can target important modulatory molecules and cells, and more

effectively inhibit the progression of atherosclerosis and its complications.

Statins, a class of lipid-lowering drugs, currently remain the most widely

prescribed pharmaceutical agent for patient with atherosclerosis (statins will be further

described in section 1.3) [25]. Several other pharmacological agents have been developed

for treating atherosclerosis, also principally directed at modulating circulating cholesterol

and lipid profiles. For instance, a drug known as ezetimibe is capable of lowering

circulating cholesterol levels by decreasing cholesterol absorption in the small intestine,

and is prescribed in combination with statin to achieve more effective LDL-lowering

effects [26].

Once the stenosis of a vessel caused by the atherosclerotic plaque has progressed

to significantly limit blood flow and perfusion of downstream tissues, viable options

  6  

include endovascular procedures such as angioplasty and stenting. Angioplasty involves

the insertion of a balloon catheter into the stenotic region, followed by its inflation,

causing expansion of the narrowed region [27]. Stenting, which can be effectively

combined with angioplasty, involves the placement of a stent in the region occluded by

the atherosclerotic plaque, thereby maintaining lumen integrity and enhancing blood

flow. More recently, in order to enhance the beneficial effects of this procedure, stents

have been modified to contain therapeutic drugs, including statins, for their local

administration to the vessel wall in the region of the plaque. Some of these drug-eluting

stents have been demonstrated to improve health outcomes for patients, including

reducing post-procedural complications [28, 29].

The most common complication of angioplasty and stenting is a recurrent

narrowing of the vessel, known as restenosis. However, this process differs in several

aspects from the stenosis resulting from atherosclerosis, as restenosis is primarily

mediated by VSMC migration and proliferation in the neointimal area of the diseased

artery. This is a physiological response following damage to the endothelial lining

resulting from angioplasty or stent placement. This neointimal hyperplasia of VSMCs

may reduce luminal size of the artery, thus causing a recurrent limitation of blood flow

[30].

However, the most common adverse event associated with atherosclerosis is the

ischemia of tissues following thromboembolism. Depending on the duration of ischemia,

significant damage can occur to downstream tissues. Patients presenting with abrupt

thromboembolism, including stroke and myocardial infarctions are placed on anti-

coagulants and thrombolytic agents, and may undergo angioplasty [31]. Subsequently,

  7  

more invasive techniques can be performed in an attempt to further treat the ischemic

event. For instance, a common procedure for atherosclerosis within the coronary arteries

is coronary artery bypass graft (CABG) surgery, in which an alternate blood flow is

supplied to the myocardial tissue, bypassing the obstructed portion of the coronary artery

[32, 33].

1.2 G-Protein Signaling in VSMC Recruitment

Under physiological conditions, VSMCs remain in a highly structured ECM,

primarily mediating the contractile function of arteries. In this state, the VSMCs contain a

large proportion of myofilaments and undergo minimal proliferation [34, 35]. However,

in response to signaling from the adjacent tissue, VSMCs are capable of undergoing

significant structural and functional changes into a dedifferentiated state. This includes a

decrease in the expression of contractile proteins, increased capability to migrate

according to chemokine gradients, augmented production of ECM, as well as higher

turnover rates due to increased proliferation and apoptosis [15, 36].

During atherosclerosis, adjacent to the plaque, the ECM around VSMCs within

the media is degraded by MMPs secreted by ECs, foam cells, as well as infiltrating

inflammatory cells [37]. These cells are also responsible for secreting the various growth

factors and cytokines that cause the VSMC phenotypic modulation, making the VSMCs

capable of migrating into the plaque and proliferating [38]. One of the crucial signaling

mechanisms involved in VSMC recruitment during atherosclerosis has been shown to be

G-protein signaling, responsible for transducing signaling from AngII, ET-1,

sphingosine-1-phosphate (S1P), as well as several other mediators of VSMC recruitment

  8  

[14, 15, 39, 40]. While the effects of these agonists varies based on the receptor subtype,

the majority of signaling responsible for VSMC recruitment has been shown to be

mediated by the Gαq as well as Gαi signaling pathways [41, 42].

Although the intracellular G-protein signaling pathways that induce the

phenotypic change in VSMCs has not been effectively elucidated, the downstream role of

extracellular signal-regulated kinase (ERK) 1/2 as a mediator of VSMC recruitment has

been well established [43]. The change in the phenotype of VSMCs is also accompanied

by significant changes in the transcriptional network. A primary mediator of the

transcriptional status of VSMCs is the transcription factor serum response factor (SRF)

and its transcriptional co-activator, myocardin, which promote the contractile phenotype

[44]. Following stimulation of VSMC recruitment resulting from vascular injury or

agonists such as PDGF-BB, a nuclear complex including Krupple-like factor-4/5

(KLF4/5) and ETS-like transcription factor-1 (Elk-1) displaces the SRF/myocardin

complex, suppressing transcription of contractile genes and upregulating proteins

involved in proliferation and migration [16, 45].

1.2.1 Heterotrimeric G-Protein Signaling

G-proteins are a large family of proteins capable of regulating intracellular

signaling by binding and hydrolyzing guanosine triphosphate (GTP). There are two

classes of G-proteins: small GTPases such as Rho and Rab, and larger G-proteins

commonly found in a heterotrimeric form associated with G-protein coupled receptors

(GPCRs) (this thesis will focus on the latter). GPCRs are a superfamily of seven-

transmembrane spanning receptors, and are the largest targets of pharmaceutical agents

  9  

[46]. All GPCRs contain an extracellular N-terminus, an intracellular C-terminus and

three inter-helical loops on each side of the membrane. However, the binding site of the

ligand to the GPCR varies based on the properties of the ligand [47].

Once the ligand binds the extracellular surface of the GPCR, the receptor

undergoes a conformational change that results in changes in the structure of the

intracellular domains. These conformational changes cause the dissociation of the

guanosine diphosphate (GDP) from the Gα subunit, which allows the formation of a

high-affinity receptor-G-protein complex. Once a GTP molecule binds to the Gα subunit,

the complex is destabilized, dissociating the Gα subunit as well as Gβγ dimer from the

receptor. In this activated form, both the GTP-bound Gα subunit and the Gβγ dimer are

capable of engaging downstream effectors, inducing intracellular effects (Figure 1-1).

The signaling is terminated by the intrinsic ability of the Gα-subunit to hydrolyze the

GTP back to GDP, causing it rebind the Gβγ dimer and deactivating the signaling

pathways [48].

The temporal regulation of G protein signaling is essential for the accurate

communication during all physiological processes. Overstimulation of G-protein

signaling pathways has been implicated in numerous pathophysiologies, including VSMC

recruitment during atherosclerosis progression [15, 49]. Therefore, the appropriate

termination of G-protein signaling is crucial, and the intrinsic rate of the Gα subunit to

hydrolyze the bound GTP is far too slow for effective physiological signaling. A large

family of GTPase-activating proteins (GAPs), known as regulator of G-protein signaling

(RGS) proteins, are able to significantly accelerate the hydrolysis of the GTP on the Gα

  10  

subunit and have also been suggested to inhibit downstream effectors of the Gα subunit.

They are therefore considered potent inhibitors of G-protein signaling [50].

1.2.2 RGS Proteins

All RGS proteins share the well-conserved RGS domain, a 120-130 amino acid

motif that directly binds the Gα subunit and reduces the activation energy of GTP

hydrolysis [51]. This can result in an up to 2000-fold increase in the hydrolysis rate above

the intrinsic rate of the Gα subunit, and reduces the activation time as well as amplitude

of G-protein signaling [52]. However, RGS proteins can have various other functional

domains, which may facilitate binding to other proteins as well as perform functions

independent of G-protein signaling [53]. According to this sequence homology

similarities and differences, the >36 members of the RGS family have been divided into

eight subfamilies [54]. Based on the prototypical member or descriptive name of the

subfamily, these groups are termed RZ, R4, R7, R12, RA, GEF, GRK, and SNX (also

arbitrarily termed A-H, respectively).

The R4 subfamily, which includes RGS1-RGS5, RGS8, RGS13, RGS16, RGS18,

and RGS21, contains the smallest RGS proteins, with a size of 20-30 kDa. With the

exception of RGS3, the only recognizable domain within this subfamily is an RGS

domain, which is flanked by short N- and C-terminus sequences. However, unlike the

RGS domain, these flanking regions, especially the amino terminus, are less conserved

among different R4 RGS proteins and are sites of post-translational modification and

protein-protein interactions. These sites have been demonstrated to be important in the

regulation of the function and localization of the different RGS proteins [55, 56].

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Moreover, differences in the C-terminus and N-terminus sequences may underlie

differences in the G-protein specificity and GAP activity among the different members of

the subfamily.

The members of the R4 subfamily are differentially expressed throughout various

tissues and cell types, and serve different functions during development and in adults. For

instance, RGS4, the prototypical member of the subfamily, is highly expressed in the

sinoatrial node, different regions of the brain, pancreatic β-cells, and several other tissues

and its crucial physiological role has been extensively investigated [57-60].

Correspondingly, deletion of the functional RGS4 gene in mice has been demonstrated to

lead to numerous pathophysiologies, including susceptibility to arrhythmias,

neurodegenerative disorders, as well as abnormalities in insulin secretion.

Conversely, the closely related RGS16 protein has been shown to be very highly

expressed in the liver as well as inflammatory cells, and its physiological roles, which

includes inhibition of hepatic fatty acid oxidation and monocyte activation, significantly

differs from that of RGS4 [61-63]. In this way, different members of the R4 subfamily,

despite their high level of sequence homology, act on G-protein signaling pathways of

diverse cells and serve different physiological functions. Nevertheless, most cells express

multiple members of the R4 RGS family. However, their function within these cells

differs based on their cellular localization, interactions with regulatory proteins, as well

as their specificity for different G-protein signaling pathways.

With the exception of RGS2, which is selective to Gαq signaling, all members of

the R4 subfamily are capable of regulating both Gαi and Gαq signaling pathways.

However, the potency of effect and the selectivity of R4 RGS proteins for different

  12  

receptors and signaling pathways can vary significantly. For instance, analysis of ERK

activation by stimulation of S1P receptor 1-3, ET-1 receptor type A, and AngII receptor

type 1 (AT1) in transfected HEK293 cells demonstrated vastly different levels of

inhibition by RGS1, RGS2, RGS3, and RGS4 [64]. Coupling between RGS proteins and

G-proteins as well as a variety of other regulatory proteins have been shown to mediate

the selectivity and differential level of inhibition by several RGS proteins [65]. For

instance, the N-terminus of the RGS4 protein has been shown to be not only important

for localization to the plasma membrane, but also to act synergistically with the RGS

domain to more selectively act on muscarinic receptors [66, 67]. However, the

mechanisms underlying the selectivity of R4 RGS proteins is not fully understood and

remains to be elucidated.

The expression profile and functional role of R4 RGS proteins within VSMCs has

been investigated by our lab as well as several other groups. Under physiological

conditions, RGS proteins are believed to act on signaling pathways that modulate VSMC

contraction and vessel tone. For instance, RGS2 has been shown to be well-expressed

within VSMCs, and capable of inhibiting Gαq-mediated vasoconstriction signaling

induced by AngII and other ligands [64, 68]. Consistently, the deletion of the functional

RGS2 gene in mice has been shown to result in hypertension as well as enhanced

response to vasoconstrictor signals [69].

However, investigating the expression and function of different R4 RGS proteins

within VSMCs has been limited by several challenges and conflicting results. For

instance, several groups have demonstrated inconsistent RGS4 mRNA expression levels

within VSMCs, with its physiological role in these cells remaining unclear [61, 64, 70].

  13  

These discrepancies may be due to differences in origin of VSMCs, as well as species-

dependent differences. Moreover, a study by Hendricks-Balls et al. (2008) has shown that

the expression of RGS2, RGS3, RGS4, RGS5, and RGS16 changes as the VSMCs are

isolated and passaged, adding another factor of complexity into the study of RGS proteins

within VSMCs [70]. Notwithstanding the controversy surrounding the expression of

some of the other R4 group proteins in VSMCs, RGS5 has been shown repeatedly to be

the most highly expressed RGS protein in VSMCs [71, 72].

1.2.3 RGS5 Structure, Localization, and Function

Similar to most members of the R4 RGS subfamily, RGS5 is composed of an

RGS domain flanked by short amino- and carboxy-terminus. The RGS5 gene, which has

been mapped to the q23 of chromosome 1 in humans, has been shown to code for a 181

amino acid sequence that is highly homologous with RGS4 and RGS16 [73, 74].

Accordingly, similar to RGS4 and RGS16, RGS5 mutants lacking the N-terminus are

almost exclusively localized to the cytosol. The RGS5 N-terminus is also involved in the

N-end degradation pathway, regulating the level of RGS5 within the cell [75].

Although the expression of RGS5 appears to be restricted primarily to pericytes

and VSMCs of arterial vessel beds, RGS5 has also been detected in several other tissues.

Previous whole-animal X-gal perfusion of RGS5-βLac/Neo KO mice from our lab

demonstrated extensive staining of most vessel beds, including those found in the brain as

well as the renal arteries. However, consistent with previous studies, lacZ staining was

also detected in several other tissues, including the heart and liver, indicating a possible

regulatory function in cardiac and hepatic function, respectively [61, 76].

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A study by Deng et al. (2012) analyzed RGS5 knockout (KO) mice placed on a

high fat diet and demonstrated higher circulating lipids, including increased triglycerol

and cholesterol levels, with an accompanying increased weight gain as compared to the

RGS5 wild-type (WT) mice, consistent with preliminary results from our lab [77]. This

was accompanied by an exacerbated metabolic dysfunction, including hyperglycemia,

hyperinsulinemia, and increased circulating inflammatory markers as compared to RGS5

WT mice. These results suggest a non-vascular role for RGS5, possibly through the

demonstrated regulation of G-protein signaling in the liver to control metabolic

homeostasis [78]. However, the molecular and cellular mechanisms underlying these

phenotypes remain to be elucidated.

Given the high-level of its expression within VSMCs, the vast majority of

research on RGS5 has focused on elucidating its role within these cells. Previous results

from our lab demonstrated a high-level of RGS5 expression throughout the major vessel

beds of neonatal mice, which has been shown to be primarily restricted to the VSMCs

and pericytes of maturing blood vessels [79-81]. With several of the vasculogenesis

pathways regulated by G-protein signaling, vascular RGS5 expression during

development is believed to be involved in angiogenesis and vascular maturation [82].

Consistently, the mRNA expression of RGS5 within pericytes has been shown to be

upregulated during angiogenesis and play a role in the pericyte-endothelial interactions

throughout vascular maturation [83]. Interestingly, however, global deletion of the RGS5

gene does not appear to affect vascular development and maturation in mice [84].

The role of RGS5 in vascular remodeling appears to not be restricted to neonatal

development, and has also been shown to play a role in vascular remodeling in adult

  15  

tissues exposed to pathophysiological conditions. A study by Arnold et al. (2014)

demonstrated that ligation-induced angiogenesis within the femoral artery caused an

increase in RGS5 expression, which is capable of shifting vascular G-protein signaling

from Gαq/11 to Gα12/13 to promote arteriogenesis [79]. This enhanced expression of RGS5

and its role in neovascularization of vascularized tissue has also been observed in tumor

angiogenesis, wound healing, and ovulation [85, 86].

Although the robust expression of RGS5 within VSMCs appears to persist in

adult mice, work from our lab has also demonstrated that there are epigenetic

mechanisms in place that promote differential expression between different vessel beds

[81]. LacZ staining of RGS5-βLac/Neo KO mice, later confirmed by quantitative real-

time polymerase chain reaction (qRT-PCR) analysis, showed that although there is a

similarly high level of RGS5 expression in all of the major vessel beds of neonatal mice,

this expression was selectively downregulated in some vascular beds due to increased

methylation of CpG dinucleotides in the RGS5 promoter.

Specifically, a high level of RGS5 expression was observed in the abdominal

aorta and femoral artery where GpG methylation of the RGS5 promoter was low,

whereas a low level of RGS5 expression was observed in the ascending aorta as well as

the carotid arteries where CpG methylation of the RGS5 promoter was high.

Interestingly, such vessel bed-specific epigenetic changes in adults were shown to

correspond to the embryologic origin of the VSMCs, thus creating an origin-specific

mosaic of RGS5 expression.

Several different groups have investigated the functional role of RGS5 in adult

VSMCs. Under physiologic conditions, VSMCs within the tunica media regulate blood

  16  

pressure in response to various hormonal and physical signals. With many of these

pathways mediated by or interconnected with G-protein signaling, the role of RGS5

within VSMCs has been studied in the context of blood pressure regulation [87-90]. A

study by Holobotovskyy et al. (2013) demonstrated that RGS5 KO mice develop

hypertension and produce larger increases in blood pressure following AngII treatment,

indicating hyper-responsiveness to vasoconstrictive signals [87]. Previous analysis of

RGS5 KO mice from our lab also shows a trend towards a slightly higher myogenic

response, possibly due to the inhibitory role of RGS5 on signaling pathways involved in

myogenic regulation [91].

Consistent with this proposed role in blood pressure regulation, RGS5 has been

suggested to mediate the anti-hypertensive effects of peroxisome proliferator-activated

receptors (PPARs), a family of nuclear receptor proteins with a variety of cardiovascular

effects [92]. Specifically, the RGS5 promoter has been shown to contain response

elements for the PPARδ and PPARγ isoforms, and treatment of arteries with agonists of

either PPARδ or PPARγ upregulates RGS5 mRNA expression [88-90]. This increase in

RGS5 expression was in turn suggested to inhibit vasoconstrictive signals induced by

AngII and ET-1. However, while the expression of the closely related RGS2 protein

within VSMCs has been shown to have an important role in blood pressure regulation,

the evidence for the function of RGS5 in vasoreactivity is not entirely clear, with our lab

and others demonstrating RGS5 KO mice to be normotensive and even hypotensive [84,

93, 94].

As mentioned previously, under pathophysiological conditions such as

atherosclerosis, VSMCs undergo significant phenotypic changes which are primarily

  17  

mediated by G-protein signaling and related pathways [95]. Therefore, RGS5 has been

hypothesized to play a role in inhibiting signaling pathways involved in VSMC

recruitment. Gunaje and colleagues have demonstrated that RGS5 knockdown in VSMCs

potentiated AngII-mediated activation of multiple downstream mitogen-activated protein

kinases (MAPKs) involved in VSMC recruitment [72]. Moreover, this reduction in RGS5

expression was shown to enhance VSMC migration induced by PDGF-BB,

demonstrating the role of RGS5 in inhibiting VSMCs recruitment induced by G-protein

signaling and interdependent pathways such as the receptor tyrosine kinase (RTK)

pathways. Consistently, RGS5 upregulation was shown to contribute to the therapeutic

effects of PPARδ on atherosclerosis progression [96].

In order to assess the in vivo role of RGS5 during atherosclerosis progression, we

previously generated RGS5/apolipoprotein E (ApoE) double KO (dKO) mice strain and

corresponding littermate control mice. Based on the ability of RGS5 to accelerate the

termination of signaling pathways in VSMCs that are crucial to atherosclerosis

progression, we hypothesized accelerated atherosclerosis progression in the dKO mice.

Throughout the 12 weeks of ad libitum high-cholesterol diet provided to induce

atherosclerosis, RGS5/ApoE dKO mice were demonstrated to have markedly reduced

survival compared to littermate RGS5 WT/ApoE KO controls (unpublished data).

Moreover, in accordance with our hypothesis, analysis of the thoracic aorta demonstrated

larger normalized Oil Red O (ORO)-stained atherosclerotic plaque size, suggesting that

deletion of the functional RGS5 gene causes acceleration of the progression of

atherosclerosis.

  18  

A recent study by Cheng et al. (2014) also assessed RGS5/ApoE dKO mouse

model, demonstrating a similar larger ORO-stained plaque size, along with a greater

necrotic core area [97]. Moreover, analysis of the plaque content of the dKO mice

showed decreased collagen and VSMC content, along with greater macrophage and lipid

burden as compared to ApoE KO littermates, indicating increased plaque instability. This

susceptibility to rupture may underlie the reduced survival observed in our results,

possibly due to thromboembolism in crucial tissues. Lastly, immunofluorescence and

western blot analysis demonstrated enhanced phosphorylation of ERK1/2 in RGS5/ApoE

dKO plaques as compared to ApoE KO littermate controls, consistent with the inhibitory

role of RGS5 on G-protein signaling pathways involved in atherosclerosis.

Consistent with an atheroprotective role of RGS5, microarray analysis of the

descending aorta by Liu et al. (2007) have demonstrated that pravastatin (a clinically

used statin) treatment of high-cholesterol fed ApoE KO mice caused an increase in the

mRNA expression of RGS5 [98]. In fact, the upregulation in RGS5 was the highest

among all non-cytoskeletal related proteins. However, the vascular cell targeted, along

with the mechanism and the functional role of the RGS5 upregulation was not identified.

  19  

Figure 1-1. Heterotrimeric G-protein signaling and the role of RGS proteins. Upon binding of the ligand to the GPCR, a conformational change in the receptor occurs which causes exchange of the GDP on the Gα subunit with a GTP molecule. This induces the dissociation of the Gα subunit from the Gβγ dimer, allowing both to engage downstream effectors and to induce intracellular signaling pathways. The signaling is terminated by the intrinsic ability of the Gα subunit to hydrolyze the GTP back to GDP, causing the G-proteins to re-associate. However, this rate is far too slow for physiological signaling and a family of proteins known as RGS proteins are capable of significantly accelerating GTP hydrolysis on the Gα subunit. These proteins have also been shown to inhibit downstream effectors of the G proteins. Therefore, RGS proteins are considered potent inhibitors of G-protein signaling.

  20  

1.3 Statins

Statins have emerged as the leading pharmaceutical treatment option for patient

with atherosclerosis. In fact, statins are among the most successful pharmaceutical drugs

in history, with atorvastatin (under the trade name Lipitor) established as the most sold

drug in history [99]. The first statin discovered was mevastatin, which was isolated from

the fungus Penicillium citrinum by Endo and colleagues [100]. Since then, several types

of synthetic statins have been isolated and developed, including the commonly prescribed

atorvastatin and rosuvastatin. All statins share the same pharmacophore, which can exist

in a configuration with a high level of structural similarity to 3-hydroxy-3-

methylglutaryl-coenzyme A (HMG-CoA), an intermediate molecule in the synthesis of

cholesterol [101].

Due to this similarity in structure, statins act as competitive inhibitors of the

enzyme HMG-CoA reductase, a rate-limiting enzyme in the cholesterol synthesis

pathway (Figure 1-2). In fact, the affinity of statins for the catalytic domain of HMG-

CoA reductase is 10,000 fold greater than HMG-CoA itself [102]. In this mechanism,

statins reduce endogenous production of cholesterol, especially in the liver, triggering

cholesterol-deprived cells to uptake circulating LDL particles [103]. This in turn leads to

LDL clearance from the blood, decreasing the risk of LDL accumulation in the

subendothelial region [104]. Accordingly, statin treatment has been shown to reduce

atherosclerosis progression and lower the risks associated with plaque development and

cardiovascular-related mortality [103, 105]

  21  

1.3.1 Comparison of Different Types of Statins

Certain types of statins, including lovastatin, pravastatin, and simvastatin have

been isolated from fungal fermentation, similar to mevastatin, while other statins such as

fluvastatin, atorvastatin, and rosuvastatin are synthetically derived [106]. Although

different types of statins act on the same enzyme, each statin has a unique

hydrophobicity, which alters the tissue permeability and pharmacokinetics of the drug.

This includes significant differences in the half-life, maximum plasma concentration,

bioavailability, and excretion routes among the various types of statins [107]. Although

both lipophilic statins (such as atorvastatin and simvastatin) and hydrophilic statins (such

as pravastatin and rosuvastatin) can readily readily enter hepatocytes through active and

passive diffusion, lipophilic statins have been shown to more selectively target the liver

[108, 109].

Moreover, with the exception of pravastatin, all statins are biotransformed by the

cytochrome P450 (CYP) isoenzyme system of the liver. However, different statins are

metabolized by different isoforms of CYP, leading to unique interactions with various

other agents that are also metabolized by the CYP system [110]. This includes

interactions with agents such as cyclosporine, warfarin, fibrates, and erythromycin [111-

114]. These differences in statin metabolism also cause further dissimilarities in the

effective plasma concentrations of statins, creating variances in the risk of statin-

mediated adverse events, including myopathy [107, 115].

  22  

1.3.2 Pleiotropic Effects of Statins

Although the cholesterol-reducing effects of statins remain a major therapeutic

mechanism, emerging data suggest that statins have a variety of important pleiotropic

effects independent of their effect on cholesterol synthesis and LDL levels [116]. The

vast majority of these effects are believed to be due to the reduction in isoprenoid

intermediates in the cholesterol synthesis pathway. As shown in Figure 1-2, isoprenoids,

which include farnesyl pyrophosphate and geranylgeranyl pyrophosphate, are derived

from mevalonate through a pathway independent of cholesterol synthesis [117].

These isoprenoid molecules are used in the post-translational modification of

several intracellular signaling molecules, including the small GTPases such as Ras, Rho,

and Rac [118]. This post-translational isoprenylation process is believed to facilitate

anchoring of the modified protein to membranes and has been shown to be crucial to their

intracellular trafficking and subsequent function. Therefore, by reducing isoprenoid

synthesis and altering isoprenylation, statins are believed to exert various effects

independent of lowering LDL levels. For instance, simvastatin-mediated reduction in

isoprenoid synthesis was shown to inhibit Rac1 activity in cardiac myocytes, resulting in

reduced levels of reactive oxygen species and resultant decreased cardiac hypertrophy

[119].

Due to the widespread isoprenylation and its consequent intracellular effects

throughout most tissues, statins are capable of exerting pleiotropic effects on numerous

physiological and pathophysiological processes [120]. Moreover, due to the

pharmacokinetic differences of statins, the pleiotropic effects can vary based on the type

of statin used. This includes therapeutic effects in the treatment of cancer based on the

  23  

ability of statins to inhibit angiogenesis as well as promote apoptosis [121]. Relatedly,

statins have been shown to have prominent anti-inflammatory effects, including

reductions in inflammatory serum markers, and decreased recruitment of macrophages

and T-cells [122]. Statins have even been suggested to exert therapeutic pleiotropic

effects on neuropathologies, including reducing the risk for Alzheimer’s and other types

of dementia [120, 123, 124].

1.3.3 Cardiovascular Pleiotropic Effects of Statins

Although statins exert diverse pleiotropic effects, the cardioprotective effects of

statins have been the best characterized and most extensively studied. While the

mechanism underlying these therapeutic effects of statins in the cardiovascular system

varies, the statin-induced activation of PPARs has been suggested to play an important

role [125]. For instance, atorvastatin treatment in aging rats has been demonstrated to

improve cardiac function, including reducing left ventricular hypertrophy and collagen

deposition, by increasing the expression of PPARs [126]. Similarly, rosuvastatin

treatment of obese mice normalized blood pressure independent of blood cholesterol

levels through upregulation of PPARγ [127].

Statins have also been shown to exert pleiotropic effects on VSMCs. The anti-

oxidant properties of statins have been shown to be an important mediator of many of the

therapeutic effects of statins on VSMCs. A study by Wassmann et al. (2001)

demonstrated that atorvastatin-mediated inhibition of isoprenoid synthesis in VSMCs

resulted in downregulation of AT1 and a decrease in reactive oxygen species [128]. This

protective effect towards oxidative stress has been suggested to be partially mediated by

  24  

statin-mediated activation of the nuclear factor-erythroid 2-related factor 2 (Nrf2)

transcription factor and the upregulation of anti-oxidants genes [129].

However, beyond these cytoprotective effects, statins have also been shown to

have therapeutic effects by regulating VSMC recruitment [109]. For instance, in vitro

treatment of isolated VSMCs with pitavastatin has been demonstrated to inhibit VSMC

migration and proliferation induced by AngII and PDGF-BB agonists [130]. Similarly, in

vivo assessment of VSMC recruitment using a neointimal hyperplasia model

demonstrated that statin treatment reduced VSMC migration and proliferation [131, 132].

These pleiotropic effects on VSMC recruitment have also been shown to contribute to the

atheroprotective effect of statins [133]. The mechanism underlying this inhibitory role of

statins on VSMC migration and proliferation has been suggested to involve activation of

PPARγ as well as attenuation of RhoA and ERK1/2 signaling [133, 134]. Nevertheless,

the molecular mechanisms are not completely understood and remain to be fully

elucidated.

  25  

 

 

 

 

 

 

 

 

 

 

 

 

 

 

 

Figure 1-2. The effect of statins on cholesterol synthesis and isoprenylation. The synthesis of cholesterol is initiated by the condensation of acetyl-CoA and acetoacetyl-CoA to produce HMG-CoA. The conversion of HMG-CoA to mevalonate is the rate-limiting step and catalyzed by HMG-CoA reductase. The pharmacophore of statins competitively and potently inhibits HMG-CoA reductase, thereby reducing production of mevalonate. The mevalonate, through several steps creates farnesyl-PP, which eventually leads to the formation of cholesterol. However, the farnesyl-PP can also be converted into geranylgeranyl-PP. Both of these molecules are post-translationally attached to various intracellular proteins in a process known as isoprenylation, which changes the localization as well as function of the protein. This cholesterol-independent effect is believed to be the primary underlying mechanism of the pleiotropic effects of statins.

  26  

2. Rationale

RGS5 is highly expressed in VSMCs and may be capable of modulating VSMC

recruitment and atherosclerosis progression by inhibiting G-protein signaling. Consistent

with the potentially atheroprotective role for RGS5 in VSMCs, Liu et al. (2007)

demonstrated that statin treatment upregulates RGS5 expression within the aorta. Among

the diverse list of pleiotropic effects demonstrated for statins, they have been shown to

inhibit VSMC migration and proliferation in vitro and in vivo, with the underlying

mechanisms not fully understood. We will assess the role of RGS5 in potentially

mediating some of the pleiotropic effects of statins in VSMCs.

Fluvastatin was selected as the statin for our studies based on its intermediate

hydrophobicity relative to other statins, as well as its high permeability to VSMCs. The

pharmacokinetics of fluvastatin in vivo has shown a high maximum plasma concentration

relative to other statins, as well as inactive metabolites [107]. Moreover, fluvastatin has

been demonstrated to exert diverse pleiotropic effects in VSMCs, including inhibition of

VSMC migration and proliferation.

  27  

3. Hypothesis and Objectives

In light of these observations, we hypothesize that statins increase the expression

of RGS5 in VSMCs through activation of PPARs (Figure 3-1). Furthermore we

hypothesize that the upregulation of RGS5 expression is sufficient to inhibit G-protein

signaling and VSMC recruitment such that these mechanisms contribute to the known

pleiotropic effects of statins. To test these hypotheses, we have designed three primary

objectives:

1. To characterize the regulation of RGS5 expression in VSMCs following

statin treatment in vitro, ex vivo, and in vivo.

2. To assess the activation of PPARs following statins treatment and its role in

the upregulation of RGS5.

3. To characterize the potential physiological role of an upregulation in RGS5

expression within VSMCs using a neointimal hyperplasia model.

  28  

Figure 3-1. Proposed mechanism and role of RGS5 upregulation following statin treatment in VSMCs. RGS5 has been shown to inhibit G-protein signaling induced by AngII, ET-1, and other GPCR agonists. It is also able to inhibit interdependent pathways, such as that induced by PDGF-BB, possibly by acting downstream of the Gα subunit. These agonists have been shown to cause VSMC migration and proliferation, which RGS5 has been suggested to inhibit. Therefore, we hypothesized that statins, by activating PPARδ and PPARγ, demonstrated transcriptional regulators of RGS5, will increase the expression of RGS5 in VSMCs. This upregulation in turn is hypothesized to inhibit G-protein signaling and interdependent pathways involved in VSMC recruitment, thus allowing RGS5 to contribute to the pleiotropic effect of statins in VSMCs.

GPCR  Agonists  

  29  

4. Materials and Methods

4.1 Experimental Animals

All experiments conformed to the Guide for the Care and Use of Laboratory

Animals published by the US National Institutes of Health (NIH Publication No. 85-23,

revised 1996), Institutional Guidelines and the Canadian Council on Animal Care. Mice

with global knockout of the functional RGS5 gene were obtained from Jackson

Laboratory (Bar Harbor, Maine, USA; http://www.informatics.jax.org/allele/key/40032).

Briefly, the heterozygous Rgs5tm1Dgen mouse strain was generated by inserting a LacZ-

neo cassette into exon 2 of the RGS5 gene, thus disrupting transcription of the functional

RGS5 protein. These mice were then backcrossed into heterozygotes C57BL/6 mice

(Charles River Laboratories, Wilmington, MA) for >8 generations.

Mice were housed in temperature- and humidity-controlled rooms with 12-hour

light-dark cycles in the Department of Comparative Medicine animal facility at the

University of Toronto. Unless otherwise stated, all mice were kept in standard vented

cages with ad libitum access to normal chow diet and water. RGS5 heterozygous mice

were bred to generate littermate RGS5 WT and KO mice. The genotypes of the mice

were assessed using the protocol provided from Jackson Laboratory.

4.2 Isolation and Culturing of Primary VSMCs

Primary aortic VSMC were isolated as described previously [135], with minor

modifications. For each isolation procedure, three 12-16 week-old female mice were

sacrificed using cervical dislocation. Briefly, the complete aorta (from the aortic root to

  30  

the bifurcation of the iliac arteries) was removed from each mouse and placed in isolation

media (Dulbecco’s modified Eagle’s medium (DMEM) containing 25 mM HEPES, pH

7.4, 0.2 U/mL penicillin, 0.2 µg/mL streptomycin, 1 mg/mL bovine serum albumin). The

surrounding adipose tissue was removed, and the aortas were placed in Digestion 1

(containing 200 U/mL of collagenase and 100 µg/mL of elastase in isolation media) for

30 minutes.

Subsequently, the aortas were washed again in isolation media and the tunica

adventitia was carefully removed. Aortas were then placed in Digestions 2 (containing

130 U/mL of collagenase and 100 µg/mL of elastase in isolation media) for 45 minutes

and washed in isolation media. Finally, the aortas were peeled into fine pieces using

sterile forceps and incubated in digestion 3 (containing 130 U/mL of collagenase and 100

µg/mL of elastase in isolation media) for 1 hour.

DMEM supplemented with 10% FBS was then added to the digestion media

containing the aortas, and the mixture was transferred to a conical tube. The plate was

washed with isolation media and added to the same tube. The mixture was centrifuged at

1000 RPM for 6 minutes, and the resulting cell pellet was resuspended in 2 mL of

Complete Media containing equal volumes of high glucose DMEM and F12 (Gibco),

10% fetal calf serum (Sigma), 1% penicillin-streptomycin (Gibco), and 50 ng/mL PDGF-

BB (Sigma). This standardized protocol in our laboratory has been previously shown to

yield >95% spindle-shaped VSMCs as determined by light microscopy (Figure 4-1) and

α-smooth muscle cell actin immunostaining. Unless otherwise stated, cells were cultured

and grown in Complete Media in humidified air containing 5% CO2 at 37 °C. All in vitro

  31  

experiments were performed on VSMCs passaged 4-7 times at >90% confluency using 6-

well plates (35 mm wells).

 

                                 

Figure 4-1. Representative image of cultured VSMCs. Image of passage 6 aortic VSMCs taken under light microscope at 20X magnification.

 

  32  

4.3 Dose Response and Time Profile Analysis of Fluvastatin Treatment

Two hours prior to drug treatment of VSMCs, the growth medium was exchanged

with 2 mL of fresh Complete Media. For dose response analysis, 2 µL of 1 mM

fluvastatin (Sigma-Aldrich), and 2 µL and 20 µL of 10 mM fluvastatin were added

directly to the Complete Media in order to obtain a final concentration of 1 µM, 10 µM,

and 100 µM, respectively, and incubated under normal culture conditions for 24 hours.

Vehicle-treated control VSMCs were treated with corresponding volumes of UltraPure

water (Invitrogen Life Technologies). The media was then removed, and the cells were

washed with 1X phosphate-buffered saline (PBS) and lysed in TRIzol reagent (Invitrogen

Life Technologies) to prepare mRNA samples for subsequent qRT-PCR analysis.

Similarly, in order to assess the time-dependency of the effect of fluvastatin treatment,

VSMCs were treated with 10 µM fluvastatin or vehicle for 4, 8, 12, 16, 20, 24 hours,

prior to TRIzol lysis and qRT-PCR analysis.

4.4 Ex Vivo Analysis of Fluvastatin Treatment

In order to assess the ex vivo effect of fluvastatin, 22-24 week-old male mice were

sacrificed using cervical dislocation and the full aorta (from the aortic root to the

bifurcation of the iliac arteries) was isolated. The aortas were then placed in Complete

Media lacking PDGF-BB and under the microscope, cleaned of all surrounding tissues.

To ensure maximal access of fluvastatin to the VSMCs within the tunica media, the

aortas were cut longitudinally in order to provide both luminal and external access to the

fluvastatin-treated media.

  33  

Subsequently, aortas were transferred into a 35 mm plate containing fresh

Complete Media (lacking PDGF-BB) containing either 10 µM fluvastatin or UltraPure

water as a control. The aortas were incubated using standard VSMC culture conditions

(5% CO2 at 37 °C) for 24 hours. Subsequently, the aortas were placed in 1 mL of TRIzol

reagent and lysed on ice using a Dounce tissue grinder (VWR International). The

mechanical lysing for each aorta was performed for 15 minutes and the homogenized

tissue in TRIzol was used for subsequent qRT-PCR analysis.

4.5 In Vivo Analysis of Fluvastatin Treatment

The effect of fluvastatin on aortic gene expression was assessed by providing 10

mg/kg/day of fluvastatin to male 11-12 weeks old RGS5 WT mice. Mice were housed

individually in cages not provided with direct water supply. The fluvastatin was dissolved

at a concentration of 0.033 mg/mL in autoclaved water (based on an average water intake

of 10 mL/day and an average body weight of 24 g) and provided ad libitum in bottles to

each mouse cage. In order to avoid any reduction in the water intake that may be caused

by changes in the taster of water due to the dissolved fluvastatin, 10 mg/mL of sucrose

(BioShop) was added to the fluvastatin-treated water as well as the vehicle-treated water.

Following 5 weeks of treatment, mice were sacrificed using cervical dislocation

and the portion of the aorta from the aortic root to the diaphragm was isolated. The aorta

was washed in 1X PBS, placed in 1 mL of TRIzol, and flash-frozen in liquid nitrogen for

subsequent RNA isolation. The tubes were then thawed, and the aortas were lysed with a

Dounce tissue grinder using the same procedure as that used for ex vivo homogenization.

  34  

Note that the in vivo assessment of the effect fluvastatin on the aorta was performed in

conjunction with the femoral artery injury experiments described in section 4.7.

4.6 qRT-PCR Reaction and Analysis

Total RNA was extracted from the TRIzol mixtures containing homogenized

VSMCs or aorta using the procedures outlined by Life Technologies

(http://tools.lifetechnologies.com/content/sfs/manuals/trizol_reagent.pdf) and dissolved in

UltraPure DNAse/RNAse-free distilled water (Invitrogen Life Technologies). The RNA

concentration was measured using a NanoDrop 1000 spectrophotometer (Thermo

Scientific) and depending on the final concentration obtained, 250, 500, or 2000 ng of the

total RNA was reverse transcribed with random hexamer primers using Superscript II kit

(Invitrogen Life Technologies) following the manufacturer's protocol. The generated

complimentary DNA (cDNA) was then diluted to 1 ng/µL for subsequent qRT-PCR.

All quantitative RT-PCR reaction was performed using ViiA 7 (Applied

BioSystems by Life Technologies) and the SYBR Green detection system. A 10 µL assay

system was used with each well containing 8 µL of reagents, including 1X SYBR Green

PCR Master Mix (Applied BioSystems by Life Technologies) and 150-900 nM forward

and reverse primers, and 2 µL of cDNA. The sequences of the forward and reverse

primers of each gene assessed are listed in Table 4-1. Melt curve analysis demonstrated a

singular peak for all primers used, indicating primer specificity and production of a single

amplicon. All samples were run in triplicates and a no reverse transcriptase and no

template control sample was run for each primer set. Glyceraldehyde 3-phosphate

  35  

dehydrogenase (GAPDH) and 18S ribosomal RNA served as housekeeping genes, and

were analyzed for each sample to serve as a normalizing control.

Data obtained from the qRT-PCR reaction were analyzed using the comparative

CT method (User Bulletin 2; Perkin Elmer Life and Analytical Sciences). Any clear

outlier CT value within the triplicates of each sample was excluded and an average was

taken to provide a mean CT for each sample. The ΔCT was determined by subtracting the

mean CT of the target gene from the mean CT of the housekeeping genes. The ΔΔCT was

then determined by subtracting the ΔCT of each sample from the calibrator sample of

each experiment, as indicated in the figure legends. Values are expressed in log scale, and

the relative mRNA levels were established by conversion to a linear value using 2−ΔΔCT.

Sample analysis for the determination of RGS5 mRNA expression in each sample within

the fluvastatin dose response experiment is shown below (data shown in Figure 5-1A).

Mean CTRGS5/sample = Average CTRGS5/sample values of triplicates

ΔCTRGS5/sample = Mean CTRGS5/sample - Mean CTGAPDH/sample

ΔΔCTRGS5/sample = ΔCTRGS5/sample - ΔCTRGS5/1 µM Vehicle

Fold Change in RGS5 mRNA Level = 2-ΔΔCT

  36  

4.7 Femoral Artery Injury Model

4.7.1 Fluvastatin Treatment and Induction of Femoral Artery Injury

Male RGS5 WT and KO mice (11-12 weeks old) were treated with 10 mg/kg/day

by providing 0.033 mg/mL fluvastatin in the autoclaved drinking water. Both fluvastatin-

treated water and corresponding control water contained 10 mg/mL sucrose in order to

avoid any alterations in drinking habits caused by drug addition. Mice were housed

individually and placed on one week of treatment prior to induction of injury. The

femoral artery injury was performed as previously described by Sata et al. (2000) and

others [136, 137]. Briefly, the left femoral artery and vein were exposed, isolated from

the femoral nerve, and looped both proximally and distally with 6-0 silk sutures for

temporary vascular control during the surgical procedures.

A small muscular branch on the femoral artery was isolated and looped

proximally with 6-0 sutures. The associated vein and connective tissue surrounding this

muscular branch were carefully removed and a transverse arterioctomy was performed

using iris spring scissors. In order to induce endothelial denudation and subsequent

neointimal hyperplasia, a straight wire guide (0.457 mm in diameter, C-SF-18-20 Cook

Medical) was carefully inserted for at least 5 mm into the femoral artery through the

incision site towards the iliac artery. The straight wire guide was kept in the artery for 1

minute, and carefully removed. The muscular branch was ligated by securing the silk

suture at the proximal portion. The sutures on the femoral artery were removed, allowing

restoration of blood flow. Finally, the skin incision was closed with 4-0 sutures, and the

mice were monitored post-surgery in order to ensure effective recovery.

  37  

The mice were then kept individually caged and on four more weeks of

fluvastatin or control treatment, prior to femoral artery isolation. Throughout the five

weeks of treatment, weekly weight measurements were made to indirectly assess relative

water intake based on body size as well as to ensure a healthy recovery following

surgery. Mice with significant and persistent reduction in body weight following surgery

were excluded from analysis.

4.7.2 Femoral Artery Isolation

Following the completion of treatment protocol, mice were sacrificed using

cervical dislocation. In order perfuse the femoral artery, the abdominal aorta was

cannulated with 0.991 mm diameter polyethylene tubing and the hepatic veins were cut to

allow perfusate emptying. Prior to the initiation of perfusion, the portion of the aorta

above the cannulation site (from the aortic root to the diaphragm) were removed, flash-

frozen in liquid nitrogen, and stored at -80 °C for subsequent qRT-PCR analysis (see

section 4.3).

The lower circulatory system was perfused with 1X PBS for 15 minutes, followed

by 15 minutes of perfusion fixation with 10% formalin (Sigma-Aldrich). Subsequently,

the left and right femoral arteries were carefully isolated (5-9 mm in length) and

removed, with the right femoral artery serving as non-injured control for each mouse.

The arteries were further fixed by being placed in 10% formalin for 24 hours and stored

in 70% ethanol for future histological analysis. The tibia length was measured in order to

assess relative body size, and used as another indirect indicator of relative water intake

based on animal size.

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4.7.3 Femoral Artery Histological Analysis

In order to perform morphometric analysis, the fixed femoral arteries were

embedded in paraffin blocks, cross-sectioned, and visualized by staining for elastin

(elastica van Gieson (EVG) staining). Images of the histology sections were obtained

using a Nikon Eclipse E600 microscope with an attached digital camera (model C4742‐

95‐12NRB, Hamamatsu, Inc.). The area analysis was performed using NIS Elements

Basic Research software (Nikon) and the neointimal area was determined by subtracting

the luminal area from the area within the internal elastic lamina (IEL). The area within

the tunica media was determined by subtracting the area within the IEL from the area

within the external elastic lamina (EEL), and the neointimal area was normalized to the

medial area. The exclusion criteria for the samples included significant thickening of the

medial layer or the absence of a clear boundary between the neointima and media. All

analysis was performed under experimental blinding, including the exclusion of samples.

4.8 Statistical Analysis

All values are expressed as mean ± standard error of the mean (SEM). Data were

analyzed using one-way or two-way ANOVA, as indicated in the figure legend, followed

by Bonferroni post-hoc test for pairwise comparisons. Statistical significance was defined

by a P-value less than 0.05. Power analysis as well as sample size determination was

performed using G*Power 3.1 software. All other calculations and statistical analyses

were performed using Microsoft Excel and GraphPad Prism version 5.0c software,

respectively.

  39  

Table 4-1. Primers used for qRT-PCR reactions

Gene Accession Number Forward Primer (5'è3') Reverse Primer (5'è3')

GAPDH M17851.1 TTCACCACCATGGAGAAGG CTCGTGGTTCACACCCATC

18S NR_003278.3 AGGAATTGACGGAAGGGCAC GGACATCTAAGGGCATCACA

RGS2 NM_009061 ATCAAGCCTTCTCCTGAGGAA GCCAGCAGTTCATCAAATGC

RGS3 AF215669 TCACACGCAATGGGAACCT GCCAGCTTATTCTTCATGTCCTT

RGS4 AB004315 GGGCTGAATCGTTGGAAAAC ATTCCGACTTCAGGAAAGCTTT

RGS5 NM_009063 GCGGAGAAGGCAAAGCAA GTGGTCAATGTTCACCTCTTTAGG

RGS16 NM_011267 CCTGGTACTTGCTACTCGCTTTT AGCACGTCGTGGAGAGGAT

PPARα NM_011144 TGGGGATGAAGAGGGCTGAG GGGGACTGCCGTTGTCTGT

PPARδ U10375 ACAGTGACCTGGCGCTCTTC TGGTGTCCTGGATGGCTTCT

PPARγ NM_011146 CAGGCTTGCTGAACGTGAAG GGAGCACCTTGGCGAACA

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5. Results

5.1 Analysis of RGS Protein mRNA Expression in Cultured VSMCs Treated with Fluvastatin 5.1.1 Dose Response Analysis of RGS Protein Expression

In order to assess the in vitro effects of statins on RGS5 expression, isolated

VSMCs were treated with 1 µM, 10 µM, or 100 µM fluvastatin or vehicle for 24 hours.

qRT-PCR analysis was performed to assess changes in the mRNA expression of RGS5 as

well as other closely-related members of the R4 RGS subfamily: RGS2, RGS3, and

RGS16. The effect of fluvastatin on the expression level of RGS4 was not assessed, as

RGS4 mRNA levels in cultured VSMCs were extremely low relative to that of other RGS

proteins. These observations were consistent with previous whole-animal X-gal perfusion

staining of RGS4-lacZ mice from our lab, which also demonstrated low expression of

RGS4 in the aorta.

Consistent with our hypothesis, RGS5 mRNA expression increased in cultured

VSMCs treated with fluvastatin for 24 hours. Although apparent increases were observed

at all doses of fluvastatin tested, only the 10 µM and 100 µM doses showed significant

upregulation of RGS5 mRNA expression (4.9-fold and 2.5-fold, respectively) compared

to vehicle-treated controls (Figure 5-1A). Notably, the extent of RGS5 upregulation at 10

µM fluvastatin was significantly greater than that observed for 100 µM fluvastatin. In

contrast to the marked increases for RGS5 mRNA expression, analysis of RGS2, RGS3,

and RGS16 showed a downregulation in their mRNA expression at both 10 µM and 100

µM fluvastatin treatment, but no significant change at the 1 µM concentration (Figure 5-

1B-D). These results suggest that among the RGS protein mRNAs analyzed in this study,

  41  

RGS5 displays the unique property of robust upregulation following fluvastatin

treatment. Additionally, based on the data from these dose response analyses, 10 µM was

selected as the optimal concentration for fluvastatin treatment in future experiments.

5.1.2 Kinetic Analysis of RGS5 Upregulation Following Fluvastatin Treatment

The kinetics of the fluvastatin-induced upregulation of RGS5 mRNA expression

was also assessed by qRT-PCR. VSMCs were treated with 10 µM fluvastatin or vehicle

(water) for 4, 8, 12, 16, 20, and 24 hours. As shown in Figure 5-2, the increase in the

RGS5 mRNA level following fluvastatin treatment was first observed at 16 hours, and

was maintained at the 20- and 24-hour time points. The level of upregulation at the 16-,

20-, and 24-hour time points were not found to be significantly different from one

another. Interestingly, the upregulation in RGS5 mRNA expression appears to occur

relatively suddenly between the 12-hour and 16-hour time points, as opposed to occurring

in a more linear pattern throughout the 24 hours of treatment. Based on these data, we

selected 24 hours of fluvastatin treatment as the standard duration of treatment in

subsequent experiments.

  42  

Figure 5-1. Relative fluvastatin-induced changes in the mRNA expression of R4 subfamily of RGS proteins in VSMCs. (A) VSMCs were treated with 1, 10, and 100 µM fluvastatin or vehicle for 24 hours. The relative mRNA levels of various RGS proteins were measured by qRT-PCR, with GAPDH mRNA levels used for normalization and 1 µM vehicle used as the calibrator sample. Fluvastatin treatment at 10 µM and 100 µM caused a significant upregulation of RGS5 as compared to controls, while (B-D) RGS2, RGS3 and RGS16 displayed a downregulation following fluvastatin treatment. (*P

  43  

   

Figure 5-2. The time profile of fluvastatin-induced upregulation of RGS5. VSMCs were treated with 10 µM fluvastatin or vehicle for different durations. The mRNA expression of RGS5 was measured by qRT-PCR, with 18S mRNA used for normalization and 4-hour vehicle used as the calibrator sample. This kinetics suggest that the fluvastatin-induced upregulation of RGS5 mRNA occurred between the 12- and 16-hour time points, which was maintained at similar levels at the 20- and 24-hour time points. (*P

  44  

5.2 Analysis of RGS5 mRNA Expression in Isolated Vessels Treated with Fluvastatin

The effect of fluvastatin on RGS5 expression was assessed on isolated vessels

treated ex vivo. The aortas from RGS5 WT mice were isolated, cut open longitudinally to

allow maximal access to fluvastatin, and incubated in culture media (without added

PDGF-BB) containing 10 µM fluvastatin or vehicle for 24 hours. Subsequently, the

aortas were lysed, and qRT-PCR was performed to measure changes in RGS5 expression

levels. Surprisingly, in contrast to the effects observed in cultured mouse VSMCs,

fluvastatin treatment did not significantly alter the mRNA expression of RGS5 in isolated

intact mouse aortas as compared to vehicle-treated controls (Figure 5-3A).

An important distinction between the in vitro analysis of VSMCs and the ex vivo

analysis of isolated aortas was presence of the PDGF-BB growth factor in the culture

medium since isolated mouse VSMCs require PDGF-BB to grow at optimal rates [138].

Based on the surprising lack of statin effect described above, we asked whether the

signaling induced by PDGF-BB in cultured VSMCs may be necessary for fluvastatin-

induced RGS5 upregulation. If true, this may explain the lack of change in RGS5 mRNA

expression following fluvastatin treatment of intact vessels ex vivo.

In order to assess this, cultured VSMCs were placed in culture media containing

or lacking 50 ng/mL PDGF-BB for 12 hours prior to a subsequent 24-hour incubation in

10 µM fluvastatin or vehicle. As shown in Figure 5-3B, fluvastatin induced a similar

level of RGS5 upregulation in both VSMCs preincubated with or without PDGF-BB,

indicating that signaling induced by PDGF-BB may not be necessary for the effect of

fluvastatin on RGS5 mRNA expression in VSMCs.

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                                                                 Figure 5-3. The ex vivo effect of fluvastatin and the role of PDGF-BB on RGS5 mRNA expression. (A) Aortas were isolated from mice and placed into culture media containing 10 µM fluvastatin or vehicle for 24 hours. The aortas were lysed and qRT-PCR analysis of RGS5 mRNA expression demonstrated no significant change following fluvastatin treatment. 18S mRNA level was used for normalization and vehicle-treated aortas were used as the calibrator sample. Statistical analysis was performed using the paired student t-test. n=3. (B) Isolated VSMCs were incubated for 12 hours in culture media either lacking or containing 50 ng/mL of PDGF-BB, and subsequently treated with 10 µM fluvastatin or vehicle. qRT-PCR analysis demonstrates a fluvastatin-induced upregulation of RGS5 mRNA in VSMCs cultured in media containing or lacking PDGF-BB, with no significant difference in the extent of upregulation. GAPDH mRNA was used for normalization and vehicle-treated VSMCs cultured in media with PDGF-BB were used as the calibrator sample. (*P

  46  

5.3 Effect of Fluvastatin on Aortic RGS Protein mRNA Expression In Vivo

The absence of statin effect on isolated vessels was unexpected, particularly in

light of work from other groups showing that RGS5 expression was increased in the

vessels of mice subjected to long-term administration of statins [98]. Based on previous

fluvastatin dose response in mice, RGS5 WT mice were provided with 10 mg/kg/day of

fluvastatin in the drinking water, with control mice provided with only water [132].

Following 5 weeks of treatment, the ascending and thoracic aortas were removed, lysed,

and the mRNA expression levels of various R4 RGS proteins were measured using qRT-

PCR. The mRNA expression of each RGS protein was normalized to the RGS2 vehicle-

treated control that was arbitrarily given an expression value of 1 in our system.

As shown in Figure 5-4, the mRNA expression of RGS5 was several fold higher

than those observed for RGS2, RGS3, and RGS16, consistent with previous studies

demonstrating RGS5 to be the mostly highly expressed RGS protein within VSMCs [72].

Although in vivo f