Early Detection of Physiologic and Functional Changes of Common Structural Protein Mutations Underlying Human Diseases by Mri

EARLY DETECTION OF PHYSIOLOGIC AND FUNCTIONAL CHANGES OF COMMON STRUCTURAL PROTEIN MUTATIONS UNDERLYING HUMAN DISEASES BY MRI

CANDIDA LAURA GOODNOUGH

Submitted in partial fulfillment of requirements

for the degree of Doctor of Philosophy

Thesis Advisor: Xin Yu, ScD

Department of Physiology & Biophysics

School of Medicine

Case Western Reserve University

May 2015

CASE WESTERN RESERVE UNIVERSITY SCHOOL OF GRADUATE STUDIES

We hereby approve the thesis/dissertation of

CANDIDA LAURA GOODNOUGH

candidate for the degree of Doctor of Philosophy

Committee Chair

William Schilling, PhD

Committee Member

George Dubyak, PhD

Committee Member

Chris Flask, PhD

Committee Member

Charles Hoppel, MD

Thesis Advisor

Xin Yu, ScD

Date of Defense

March 4, 2015

*We also certify that written approval has been obtained for any proprietary material contained therein.

Dedicated to Henry and Harrison – the two loves of my life i

TABLE OF CONTENTS

LIST OF TABLES...... V LIST OF FIGURES ...... VI LIST OF COMMONLY USED ABBREVIATIONS…………………………………….…VIII

ABSTRACT ...... 1

CHAPTER 1. INTRODUCTION AND SIGNIFICANCE ...... 3 SIGNIFICANCE ...... 3 HYPERTROPHIC CARDIOMYOPATHY ...... 6 INTRODUCTION ...... 6 CLINICAL PICTURE ...... 7 PATHOLOGY ...... 7 GENETICS ...... 8 PRESENTATION ...... 10 DIAGNOSIS ………………………………………………………………………… .11 TREATMENT ……………………………………………………………………….. 13 RESEARCH ADVANCES …………………………………………………………….. 15 DUCHENNE MUSCULAR DYSTROPHY ...... 18 INTRODUCTION ...... 18 CLINICAL PICTURE ...... 20 PATHOLOGY ...... 21 GENETICS ...... 22 PRESENTATION ...... 23 DIAGNOSIS ...... 25 TREATMENT ...... 27 RESEARCH ADVANCES ...... 27 MAGNETIC RESONANCE IMAGING ……………………………………………..……. 31 DISPLACEMENT ENCODING WITH STIMULATED ECHOES ...... 31 ARTERIAL SPIN LABELING ...... 33 DIFFUSION-WEIGHTED IMAGING ...... 33 REFERENCES ………………………………………..………………………………… 35

CHAPTER 2.CARDIAC MYOSIN BINDING PROTEIN C INSUFFICIENCY LEADS TO EARLY ONSET OF MECHANICAL DYSFUNCTION ...... 43 ABSTRACT: ...... 43 INTRODUCTION ...... 44 MATERIALS AND METHODS ...... 46 ANIMAL MODELS ...... 46 SKINNED FIBER EXPERIMENTS ...... 46 IN VIVO MRI MEASUREMENTS OF CARDIAC FUNCTION ...... 46 STATISTICAL ANALYSIS ...... 46

RESULTS ...... 47 MYOFILAMENT PROTEIN EXPRESSION AND PHOSPHORYLATION IN WT AND CMYBPC MUTANT MYOCARDIUM ...... 47 MECHANICAL PROPERTIES OF SKINNED MYOCARDIUM ISOLATED FROM WT AND CMYBPC MUTANT HEARTS ...... 50 IN VIVO ASSESSMENT OF CARDIAC MORPHOLOGY AND MECHANICAL FUNCTION .. 51 LV MORPHOLOGY AND CONTRACTILE FUNCTION ...... 51 MYOCARDIAL STRAIN ...... 53 VENTRICULAR TWIST AND TORSION...... 54 DISCUSSION ...... 57 MYOFILAMENT FUNCTION IN CMYBPC MUTANT MYOCARDIUM ...... 58 EFFECTS OF DECREASED CMYPBPC EXPRESSION ON IN VIVO MECHANICAL FUNCTION ...... 60 POTENTIAL FUNCTIONAL CONSEQUENCES OF CMYBPC HAPLOINSUFFICIENCY ...... 62 CLINICAL IMPLICATIONS ...... 63 LIMITATIONS ...... 64 CLINICAL PERSPECTIVE...... 65 REFERENCES: ...... 67

CHAPTER 3.COMPARISON OF VELOCITY VECTOR IMAGING ECHOCARDIOGRAPHY WITH MAGNETIC RESONANCE IMAGING IN MOUSE MODELS OF CARDIOMYOPATHY ...... 71 ABSTRACT ...... 71 INTRODUCTION ...... 72 MATERIALS AND METHODS ...... 74 EXPERIMENTAL ANIMALS ...... 74 MAGNETIC RESONANCE IMAGING ...... 74 ECHOCARDIOGRAPHY ...... 75 STATISTICAL ANALYSIS ...... 78 RESULTS ...... 79 ECHOCARDIOGRAPHIC AND MRI VOLUMETRIC VARIABLES ...... 79 GLOBAL STRAIN ...... 79 REGIONAL STRAIN ...... 81 INTERPRETATIVE VARIABILITY ...... 83 DISCUSSION ...... 83 LIMITATIONS ...... 86 CLINICAL PERSPECTIVE...... 88 REFERENCES ...... 90

CHAPTER 4. ARTERIAL SPIN LABELING-FAST IMAGING WITH STEADY-STATE FREE PRECESSION (ASL-FISP): A RAPID AND QUANTITATIVE PERFUSION TECHNIQUE FOR HIGH-FIELD MRI ...... 92 ABSTRACT ...... 92 INTRODUCTION ...... 93

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MATERIALS AND METHODS ...... 95 IN VIVO COMPARISON OF IMAGE ARTIFACTS AT 7T ...... 96 ASL-FISP PULSE SEQUENCE DESIGN ...... 96 INITIAL IN VIVO ASL-FISP PERFUSION ASSESSMENTS IN MOUSE BRAIN ...... 97 ADDITIONAL IN VIVO ASL-FISP EXPERIMENTS: MOUSE BRAIN ISCHEMIA STROKE MODEL AND HEALTHY MOUSE KIDNEYS ...... 101 RESULTS ...... 102 DISCUSSION ...... 108 REFERENCES ...... 113

CHAPTER 5.LACK OF DYSTROPHIN RESULTS IN ABNORMAL CEREBRAL DIFFUSION AND PERFUSION IN VIVO ...... 116 ABSTRACT ...... 116 INTRODUCTION ...... 117 MATERIALS AND METHODS ...... 120 ANIMAL MODELS ...... 120 PERFUSION AND DIFFUSION MRI ...... 120 ANALYSIS OF VASCULAR DENSITY BY IMMUNOCYTOCHEMISTRY ...... 122 ANALYSIS OF BBB LEAKAGE USING EVANS BLUE STAINING ...... 123 ANALYSIS OF VASCULAR DENSITY BY CRYO-IMAGING...... 123 STATISTICAL ANALYSIS ...... 124 RESULTS ...... 124 DECREASED CEREBRAL DIFFUSIVITY WITH DYSTROPHIN DISRUPTION ...... 124 ABNORMALITIES IN CEREBRAL PERFUSION IN AGED MDX MICE ...... 126 DISRUPTION OF BLOOD BRAIN BARRIER INTEGRITY IN MDX MICE ...... 126 ENHANCED CEREBRAL ARTERIOGENESIS IN AGED MDX MICE...... 127 DISCUSSION ...... 128 REFERENCES ...... 135

CHAPTER 6. FUTURE DIRECTIONS ...……………………………………….……….139 MECHANICAL DYSFUNCTION IN OTHER MODELS OF HCM ...... 139 RATIONALE ...... 139 EXPERIMENTAL STRATEGY ...... 140 INTERPRETATION ...... 140 POTENTIAL CONCERNS ...... 142 PREDICTIVE VALUE OF MECHANICAL DYSFUNCTION IN PRECLINICAL HCM … .... 142 RATIONALE ...... 142 EXPERIMENTAL STRATEGY ...... 143 INTERPRETATION ...... 144 POTENTIAL CONCERNS ...... 145 FUNCTIONAL CONSEQUENCE OF CEREBRAL ISCHEMIA IN MDX MICE ...... 146 RATIONALE ...... 146 EXPERIMENTAL STRATEGY ...... 148 INTERPRETATION ...... 150 POTENTIAL CONCERNS ...... 150

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FUNCTIONAL CONSEQUENCE OF HYPOPERFUSION IN DMD ...... 151 RATIONALE ...... 151 EXPERIMENTAL STRATEGY ...... 152 INTERPRETATION ...... 153 REFERENCES ...... 155

BIBLIOGRAPHY……………………………………………………………………..…160

LIST OF TABLES

TABLE 1.1 CASE REPORTS OF CEREBRAL INFARCTION IN DMD PATIENTS ...... 25

TABLE 1.2 OVERVIEW OF STRATEGIES FOR DUCHENNE MUSCULAR DYSTROPHY GENE THERAPY ...... 28 ------TABLE 2.1 STEADY STATE AND DYNAMIC MECHANICAL PROPERTIES OF SKINNED FIBERS ISLOATED FROM WT AND CMYBPC MUTANT MYOCARDIUM ...... 50

TABLE 2.2 STRETCH ACTIVATION PARAMETERS OF SKINNED FIBERS ISOLATED FROM WT, CMYBPC +/-, AND CMYBPC -/- MYOCARDIUM ...... 51

TABLE 2.3 CARDIAC MORPHOLOGY AND CARDIOVASCULAR FUNCTION OF WT, CMYBPC +/-, AND CMYBPC -/- MYOCARDIUM ...... 52 ------TABLE 3.1 ECHOCARDIOGRAPHIC VARIABLES ...... 80

TABLE 3.2 MRI VARIABLES ...... 80 ------TABLE 6.1 GENES RESPONSIBLE FOR HYPERTROPHIC CARDIOMYOPATHY ...... 139

TABLE 6.2 MOUSE AND HUMAN GENES IDENTIFIED IN HYPERTROPHIC CARDIOMYOPATHY ...... 141

TABLE 6.3 HYPERTROPHIC CARDIOMYOPATHY VS PHYSIOLOGIC ATHETIC HEART HYPERTROPHY ...... 145

LIST OF FIGURES

FIGURE 1.1 SCHEMATIC OF THE CARDIAC SARCOMERE ...... 6

FIGURE 1.2 HYPERTROPHIC CARDIOMYOPATHY ...... 8

FIGURE 1.3 PRIMARY TREATMENT STRATEGIES FOR PRESENTATIONS OF HYPERTROPHIC

CARDIOMYOPATHY ...... 13

FIGURE 1.4 OUTCOMES RELATED TO GENETIC STATUS IN HCM ...... 16

FIGURE 1.5 STUCTURAL NETWORK IN THE CARDIOMYOCYTE ...... 18

FIGURE 1.6 THE NEUROVASCULAR UNIT ...... 19

FIGURE 1.7 ENDOTHELIAL AND ASTROGLIAL CELL INTERACTION AT THE BLOOD-BRAIN

BARRIER ...... 20

FIGURE 1.8 PROPOSED PATHOPHYSIOLOGY OF DUCHENNE MUSCULAR DYSTROPHY ... 21

FIGURE 1.9 STAGES OF DUCHENNE MUSCULAR DYSTROPHY AND TREATMENT OPTIONS ...... 26

FIGURE 1.10 LEFT VENTRICULAR MECHANICS DURING CONTRACTION ...... 32 ------FIGURE 2.1 QUANTIFICATION OF CMYBPC EXPRESSION IN CMYBPC MYOCARDIUM….48

FIGURE 2.2 QUANTIFICATION OF MYOSIN HEAVY CHAIN ISOFORM EXPRESSION IN WT

AND CMYBPC MUTANT MYOCARDIUM ...... 49

FIGURE 2.3 MYOFIBRILLAR PROTEIN CONTENT AND PHOSPHORYLATION IN WT AND

CMYBPC MUTANT MYOCARDIUM ...... 49

FIGURE 2.4 STRETCH ACTIVATION KINETICS IN SKINNED MYOCARDIUM ...... 51

FIGURE 2.5 CARDIAC HYPERTROPHY ANALYSIS ...... 52

FIGURE 2.6 REPRESENTATIVE MRI PRINCIPAL STRAIN VECTOR DISPLACEMENT MAPS .. 53

FIGURE 2.7 MEASUREMENTS OF LEFT VENTRICULAR GLOBAL PRINCIPAL STRAINS ...... 54

FIGURE 2.8 ANALYSIS OF PRINCIPAL STRAIN RATES ...... 55

FIGURE 2.9 LEFT VENTRICULAR TWIST RELATIONSHIPS DURING SYSTOLE ...... 57

FIGURE 2.10 ANALYSIS OF LEFT VENTRICULAR TORSION ...... 58 ------FIGURE 3.1 DENSE MRI AND VELOCITY VECTOR IMAGING ECHOCARDIOGRAPHIC VECTOR

MAPS FOR REGIONAL STRAIN IN WT VS. KO MICE ...... 77

FIGURE 3.2 BLAND-ALTMAN PLOTS COMPARING MRI AND ECHO-DERIVED STRAIN ...... 81

FIGURE 3.3 REGIONAL RADIAL STRAIN ...... 82

FIGURE 3.4 REGIONAL CIRCUMFERENTIAL STRAIN ...... 83 ------

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FIGURE 4.1 SCHEMATIC DIAGRAM OF ASL-FISP ACQUISITION ...... 97

FIGURE 4.2 REPRESENTATIVE AXIAL MOUSE BRAIN IMAGES AT 7T USING SPIN ECHO, FISP,

TRUE FISP, AND EPI ACQUISITIONS ...... 102

FIGURE 4.3 REPRESENTATIVE ASL-FISP IMAGES OF HEALTHY C57/BL6 MOUSE BRAIN AT

7T ...... 103

FIGURE 4.4 MEAN C57/BL6 BRAIN PERFUSION FROM ASL-FISP AT 7T AND 9.4T ...... 105

FIGURE 4.5 ASL-FISP IMAGES OF A MIDDLE CEREBRAL ARTERYOCCLUSION MOUSE

MODEL OF STROKE AT 7T ...... 106

FIGURE 4.6 KIDNEY ASL-FISP IMAGES FROM HEALTHY C57/BL6 MOUSE AT 7T ...... 107 ------FIGURE 5.1 REPRESENTATIVE DIFFUSION IMAGES OF 2-MONTH OLD WT VS MDX AND 10-

MONTH OLD WT VS MDX ...... 124

FIGURE 5.2 REPRESENTATIVE PERFUSION IMAGES OF 2-MONTH OLD WT VS MDX AND 10-

MONTH OLD WT VS MDX ...... 125

FIGURE 5.3 REPRESENTATIVE IMAGES OF EVAN’S BLUE FLUORESCENCE ON AXIAL

SECTIONS OF FROZEN 2-MONTH OLD WT VS MDX AND 10-MONTH OLD WT VS MDX

BRAINS ...... 126

FIGURE 5.4 REPRESENTATIVE IMAGES OF INDIRECT IMMUNOFLUORESCENCE AGAINST

CD31/PECAM1 ON AXIAL SECTIONS OF FIXED, FROZEN 2-MONTH OLD WT VS MDX AND

10-MONTH OLD WT VS MDX BRAINS ...... 127

FIGURE 5.5 REPRESENTATIVE VIEWS OF 3D VESSEL RECONSTRUCTION OF 2-MONTH OLD

WT VS MDX AND 10-MONTH OLD WT VS MDX BRAINS ...... 128

FIGURE 5.6 COMPARISON OF CLASSIC MODEL AND PELL MODEL ...... 133 ------FIGURE 6.1 COGNITIVE TESTING FOR THE MOUSE ...... 153

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LIST OF COMMONLY USED ABBREVIATIONS

ADC ...... Apparent Diffusion Coefficient AQP4 ...... Aquaporin 4 ASL ...... Arterial Spin Labeling BBB ...... Blood-Brain Barrier CK ...... Creatine Kinase CMR ...... Cardiovascular Magnetic Resonance cMyBPC ...... Cardiac Myosin Binding Protein C CS ...... Circumferential Strain DENSE ...... Displacement Encoding with Stimulated Echoes DGC ...... Dystrophin-Glycoprotein Complex DMD ...... Duchenne Muscular Dystrophy DWI ...... Diffusion Weighted Imaging ECG ...... Electrocardiogram ECHO ...... Echocardiography EF ...... Ejection Fraction FISP ...... Fast Imaging with Steady State Free Precession HCM ...... Hypertrophic Cardiomyopathy ICD ...... Implantable Cardioverter-Defibrillator LV ...... Left Ventricular LVH ...... Left Ventricular Hypertrophy MCA ...... Middle Cerebral Artery MLP ...... Muscle Limb Protein MRI ...... Magnetic Resonance Imaging RS ...... Radial Strain VVI ...... Velocity Vector Imaging WT ...... Wild Type

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Early Detection of Physiologic and Functional Changes of Common Structural Protein Mutations Underlying Human Diseases by MRI

ABSTRACT

CANDIDA LAURA GOODNOUGH

The most common genetic disorders of skeletal and cardiac muscle result from mutations in structural proteins essential for normal tissue function. The ability to identify tissue dysfunction prior to the onset of overt clinical symptoms will provide better insight of physiology and treatment of pathology. Hypertrophic cardiomyopathy (HCM) is the single most common genetic disorder of the heart, resulting from mutations in sarcomeric proteins, and manifesting as sudden death or progressive heart failure. Characterization of early cardiac phenotypes and improved screening measures in HCM may reduce morbidity and mortality. Likewise,

Duchenne Muscular Dystrophy (DMD) is an X-linked progressive, invariably fatal disease of skeletal and cardiac muscle resulting from mutations in dystrophin, one of the largest genes in the genome. One of the largest causes of mortality in this cohort is dilated cardiomyopathy. There is also a need to identify extra-cardiac sequelae of dystrophin mutations as new therapies may dramatically improve the life expectancy of patients with DMD. Indeed, dystrophin is also necessary to maintain the blood- brain barrier and cerebrovascular disease is case-reportable in DMD patients. The goal of this work is to explore the physiologic consequences of aberrant structural proteins using in vivo magnetic resonance imaging (MRI). In particular, MRI techniques were 2

used to detect early pathologic changes in cardiovascular and vascular function prior

to the manifestation of overt symptoms in mouse models of HCM and DMD,

respectively. Here, we demonstrate that in a mouse model of HCM, displacement

encoding with stimulated echoes (DENSE) MRI detects left ventricle mechanical

dysfunction antecedent of pathologic hypertrophy. We also demonstrate that velocity

vector imaging echocardiography is comparable to DENSE MRI in identifying

mechanical dysfunction in cardiomyopathy. Further, we developed an arterial spin

labeling-fast imaging with steady-state free precession (ASL-FISP) MRI method as a

means to measure cerebral perfusion. Finally, in a mouse model of muscular

dystrophy, ASL detected decreased cerebral perfusion in setting of increased angiogenesis. Taken together, we demonstrate that the development of new imaging technologies not only might allow earlier diagnosis and thus more timely intervention,

but also may facilitate identification of subtle and clinically relevant phenotypes in

important disease states.

CHAPTER 1: INTRODUCTION SIGNIFICANCE Hypertrophic cardiomyopathy (HCM) is a common genetic myocardial

disorder that is the leading cause of sudden cardiac death in young athletes(Maron et al., 1996a). An epidemiological study demonstrated a prevalence of HCM in the general adult population (ages 23-35) of 1:500 people using an echocardiographic basis

of left ventricular hypertrophy (LVH) (Maron et al., 1995a). However, a large portion

of adults carrying a mutant gene for HCM (particularly cMyBPC) are undetected since

they do not yet exhibit overt LVH by echocardiography(Maron et al., 2001; Niimura

et al., 1998a). The most devastating consequence of undetected HCM is sudden death,

particularly following vigorous exercise in young adults. Sudden death was the

primary mode of death in 51% (ages 45±20 yrs) of a cohort of HCM patients, followed

by progressive heart failure (36%) and HCM-related stroke associated with atrial

fibrillation (13%)(Maron et al., 2000). There is therefore precedence for early detection

of cardiovascular dysfunction in HCM patients prior to the onset of symptoms or

LVH.

Traditionally, family members of HCM patients are screened with

echocardiography and electrocardiography (ECG) on an annual basis from the age of

12 to 18, and if no LVH is detected, families are typically reassured that no further

echocardiographic testing is necessary(Maron et al., 2004). Nevertheless, there are

some genetic variants of HCM that demonstrate incomplete penetrance and LVH is

not detected until later in adulthood(Maron et al., 2001), so there is a risk that these

patients may not be appropriately detected. The introduction of commercially 4

available genetic tests to detect the most common mutations in HCM has improved

risk stratification and identifies family members suitable for further testing(Bos et al.,

2009a). However, the uncertainty between the genotype-phenotype relationship still

warrants a better imaging technique to identify pre-clinical expression of the genetic

substrate.

Muscular dystrophies present another common cause of cardiomyopathy in the

younger patient population(Hermans et al., 2010; Towbin, 1998). In particular, dilated

and hypertrophic cardiomyopathy may be detectable as early as age 10 in Duchenne

muscular dystrophy (DMD) boys (Nigro et al., 1990). Congestive heart failure or

sudden death account for 10-20% mortality in DMD patients(American Academy of

Pediatrics Section on Cardiology and Cardiac Surgery, 2005). Early detection of

cardiac dysfunction is typically limited due to the extreme physical inactivity and thus

a lack of symptoms in young DMD boys. However, there has been improvement in

the quality of life and prolonged survival in DMD, which highlights the need to

diagnose preclinical cardiovascular dysfunction and initiate treatment earlier.

Abnormal myocardial strain detectable by cardiovascular magnetic resonance imaging

(cMRI) actually preceeds the onset of global functional deterioration(Ashford et al.,

2005; Li et al., 2009a). Subclinical cardiac dysfunction is an indication for initiation of

angiotensin-converting-enzyme (ACE) inhibitors, followed by β blockers and diuretics

as management of heart failure(Bushby et al., 2010a).

Although significant advancements have been made in the treatment of muscle

and pulmonary function(Eagle et al., 2002a; Finder et al., 2004; Simonds et al., 1998),

the progression of vascular disease needs further investigation in DMD patients. In 5

addition to development of cardiomyopathy and/or cardiac arrhythmias, there has

also been evidence of vascular dysfunction in the skeletal muscle and brains in a mouse

model of DMD(Nico et al., 2002; Straino et al., 2004) . Cerebrovascular integrity is

essential for cognitive function and remains poorly understood in DMD.

OUTLINE

This work is dedicated to the early detection of structural protein dysfunction

underlying human disease by MRI. I begin by introducing the clinical features of

Hypertrophic Cardiomyopathy and Duchenne Muscluar Dystrophy. Short

descriptions of the MRI techniques used in my studies will follow. Chapter 2 describes

our finding of early mechanical dysfunction in HCM, which was published in

Circulation Imaging in 2012 (Desjardins et al., 2012), and Chapter 3 compares two

different imaging techniques to quantify this dysfunction (Azam et al., 2012). Chapter

4 introduces the MRI method (Gao et al., 2014) that is used to quantify perfusion

deficits in a mouse model of DMD laid out in Chapter 5 (Goodnough et al., 2014). I

conclude my dissertation with future directions in studying structural protein

dysfunction underlying HCM and DMD.

HYPERTROPHIC CARDIOMYOPATHY

INTRODUCTION

Cardiac myocytes are composed of parallel arrangements of myofibrils which

are further divided into sarcomeres, the basic contractile unit of the heart muscle (Hoit

and Walsh, 2011). The sarcomere is a cytoskeletal structure made of thick and thin myofilaments. Two units of β- or α-myosin heavy chains and four myosin light chain molecules make up the thick filament, whereas the thin filament is composed of repeating actin molecules that associate with troponins T (cTnT), I (cTnI), and C

Figure 1.1: Schematic of the cardiac sarcomere

Adapted from https://physiology.case.edu/people/faculty/julian‐e‐stelzer/ 7

(cTnC). Cardiac myosin-binding protein C (cMyBPC) plays a role in the regulation of

actin-myosin interaction and cross-bridge kinetics (Figure 1.1).

CLINICAL PICTURE

Hypertrophic cardiomyopathy (HCM) is a disease caused by mutations in

genes that encode sarcomeric proteins. Hypertrophic cardiomyopathy (HCM) is primarily an inherited disease of the myocardium affecting young adults. It is characterized by left ventricular hypertrophy and myofibril disarray in the absence of

other causes, which leads to a reduced left ventricular cavity (Wigle et al., 1995). HCM

is frequently featured in the news as the leading cause of sudden death in young

athletes. The overall prevalence of HCM in the general population has been estimated

as 1:500. HCM affects many races and ethnicities, and occurs equally in males and

females (Maron et al., 1995b).

PATHOLOGY

The pathophysiology of HCM is diastolic dysfunction, left ventricular outflow

tract obstruction, mitral regurgitation, myocardial ischemia, and arrhythmias

(Ommen et al., 2011). The hypertrophy in HCM is classically asymmetric, affecting

the septum more so than the free wall. The largely septal hypertrophy decreases the

left ventricular cavity (Davies et al., 1974). HCM can be present with or without an

obstructive gradient, which impedes outflow during systolic contraction and is

intensified by increased contractility. Diastolic dysfunction arises from defects in

ventricular relaxation and chamber stiffness (Wigle et al., 1995). The outflow tract

obstruction, asymmetric hypertrophy, and delayed inactivation due to abnormal 8

Figure 1.2: Hypertrophic Cardiomyopathy

(A) Gross heart specimen from a 13‐year‐old male athlete demonstrating thickiening of the ventricular septum (VS) with respect to the left ventricular (LV) free wall. (B) Marked disarray of cardiac muscle cells with adjacent hypertrophied cells. (C) LV myocardium demonstrating markedly thickened walls dispersed within replacement fibrosis.

Adapted from Maron BJ. Hypertrophic cardiomyopathy. Current Probl Cardiol. 1993;18:637‐ 704.

intracellular calcium reuptake causes impaired ventricular relaxation. The

hypertrophy of the myocardium causes increased chamber stiffness and impairs

relaxation. Myocardial ischemia may also affect relaxation and chamber stiffness. As

a result, there is a compensatory increase of late diastolic filling during atrial systole.

Approximately one third of patients with symptomatic HCM exhibit signs of

obstructive gradients (Gersh et al., 2011). Microscopically, disordered myocytes swirl

and branch, interspersed with fibrosis in HCM tissue (Figure 1.2, (Maron, 1993)).

GENETICS

It is critical to complete a full family history in patients with suspected HCM.

In addition to identifying family members with a definitive diagnosis of HCM, it is 9

also beneficial to evaluate if any family members died suddenly at a young age, or

were involved in an otherwise unexplained car accident. Approximately one half of

HCM cases are identified by the typical Mendelian autosomal dominant fashion

(Stevenson and Loscalzo, 2012). Clinical genetic testing may be particularly useful for

identifying family members at risk for developing HCM, especially if there are no overt

signs of left ventricular hypertrophy. However, the pathogenic mutation must be

known for genetic testing to be of any utility, which is the case approximately 35% of

the time (Richard et al., 2003). Although genetic screening may be possible in the

future, the genetic heterogeneity requiring extensive resequencing and the high

percentage of unknown causal genes for HCM require further investigation.

Hypertrophic cardiomyopathy is thought to be the most common genetic

myocardial disorder, affecting approximately 1 in 500 people (Maron et al., 1995b).

HCM is a highly heterogeneous disorder, with several hundred genetic mutations

described. The most common genes affected in HCM encode for the β-myosin heavy

chain and cardiac myosin-binding protein C (cMyBPC), however, mutations in

cardiac troponin T and I, actin, titin, α-tropomyosin, and myosin light chains can also

lead to HCM (Richard et al., 2003). cMyBPC is a thick filament protein that modulates

actin-myosin interactions and thereby the rate of muscle contraction (Barefield and

Sadayappan, 2010; Carrier, 2007). Mutations in cMyBPC are among the most

common genetic causes of HCM, accounting for more than 40% of the total known

cases worldwide (Richard et al., 2003).

Nearly 200 known mutations of the gene encoding cMyBPC have been

identified as causative for HCM (Bonne et al., 1995; Harris et al., 2011; Watkins et al., 10

1995). The mutations can be deletions, insertions, missense, or splice site.

Alternatively, de novo mutations may cause sporadic cases. A large number of disease-

causing mutations in cMyBPC in humans are predicted to produce C-terminal

truncated proteins that are not incorporated into the sarcomere, resulting in cMyBPC

haploinsufficiency. The majority of mutations in the gene that encode cMyBPC are

heterozygous and are predicted to result in expression of truncated cMyBPC lacking

the C-terminal regions of the protein that binds to myosin and titin (Carrier et al.,

1997). However, analysis of myocardial biopsy samples from patients with cMyBPC

mutations demonstrate a reduction in the amount of full-length cMyBPC protein (van

Dijk et al., 2009; Jacques et al., 2008; Marston et al., 2009). Therefore, the allele

generating mutant cMyBPC effectively functions as a null allele, causing cMyBPC

haploinsufficiency. However, the mechanisms that link reduced cMyBPC levels in the

heart with the development and progression of HCM have remained elusive.

PRESENTATION

The characteristic hemodynamic defect in HCM is diastolic dysfunction, both

as a primary consequence of the disease and also secondary to hypertrophy, fibrosis,

and outflow obstruction. The ejection fraction and cardiac output are typically normal

in HCM, unless the patient undergoes vigorous exercise, where the ventricular filling

is inadequate to match the increased demand. The physical exam of an HCM patient

is consistent with left ventricular hypertrophy: bifid apical impulse and fourth heart

sound(Ommen et al., 2011). A physical exam of the HCM patient would also reveal a

harsh systolic crescendo-decrescendo murmur heard best at the left lower sternal

border. The murmur arises from left ventricular outflow turbulence/obstruction and 11

mitral regurgitation. The classic sign of HCM is an increase in the intensity of the

murmur with maneuvers that decrease ventricular volume, like the Valsalva

maneuver, and a decrease by increasing ventricular volume by squatting. It also common to hear a fourth heart sound as a result of decreased ventricular compliance.

HCM patients typically present between the ages of 20 and 40 years with dyspnea on exertion (Stevenson and Loscalzo, 2012). Dyspnea is the most common symptom in HCM patients, occurring in up to 90% of symptomatic patients. In addition, it is common for patients to complain of chest pain with or without exertion due to myocardial ischemia from an increase in demand. Conduction abnormalities may also produce palpitations, either from atrial fibrillation or ventricular arrhythmias. The majority of HCM patients are asymptomatic when they are diagnosed, however, dyspnea, angina, and/or syncope are the most common presentations.

DIAGNOSIS

The first manifestation of HCM may be sudden cardiac death from ventricular

tachycardia or fibrillation in young patients during a sporting event (Maron et al.,

1996b). Many younger individuals who carry disease-causing mutations in the

cMyBPC gene do not exhibit overt LVH, because increases in LV wall thickness are often only detectable with advanced age (Maron et al., 2004; Niimura et al., 1998a).

Given that these seemingly asymptomatic carriers are at risk for the development of

HCM and cardiac disease later in life, the diagnosis and treatment of these patients is a major clinical challenge. The majority of HCM patients are asymptomatic at the time of diagnosis by abnormal electrocardiogram, heart murmur, or screening 12

echocardiogram. An abnormal electrocardiogram is present in the majority of

symptomatic HCM patients (95%), demonstrating left axis deviation, left ventricular hypertrophy, prominent septal Q waves, and/or diffuse T-wave inversions (Frank and

Braunwald, 1968). A chest x-ray would demonstrate mild to moderate cardiac

silhouette enlargement, which is consistent with left ventricular and atrial

enlargement.

The gold standard for a diagnosis of HCM is increased left ventricular wall

thickness in the absence of other pathology by 2D and Doppler echocardiography

(Ommen et al., 2011). Two-dimensionally (2D)-directed echocardiography has

arguably become the leading method for assessing left ventricular (LV) function

because it is noninvasive, relatively cost-effective, widely available, and has short

acquisition and post-processing times that allow high through-put. However, 2D-

directed detection of LV dysfunction by conventional echocardiography (M-mode,

2D, Doppler) is considered a late manifestation of cardiac disease that lacks the

sensitivity to identify subclinical disease (Marwick et al., 2010; Stanton and Marwick,

2010). Cardiac magnetic resonance imaging (MRI) is also useful in determining the

site and extent of hypertrophy. MRI studies can be particularly helpful in patients with

equivocal echocardiographic results. With its high spatial resolution and superb

imaging quality, MRI can be used to better delineate subtle apical or segmental

hypertrophy. Cardiac catheterization is not a common study in HCM patients,

however, it may demonstrate left ventricular outflow obstruction if the

echocardiographic study is inconclusive.

TREATMENT

The therapy goals in HCM are to manage symptoms and prevent sudden death

(see Figure 1.3, (Maron and Maron, 2013)). Studies have been inconclusive regarding treatment of asymptomatic family members of HCM patients (Ommen et al., 2011).

Medical therapy is the first line of treatment in symptomatic HCM patients. Calcium-

channel blockers (primarily verapamil) and beta-adrenergic blockers are used to

prevent exertional dyspnea and chest pain. Oral anti-coagulation is indicated to prevent of embolic events in at-risk patients with atrial fibrillation. If medical management fails, particularly in patients with severe ventricular outflow obstruction, interventional myomectomies reduce the size of the septum. Similarly, ethanol- mediated ablation via catheterization may reduce the size of the septum by infarction.

Figure 1.3: Primary treatment strategies for presentations of hypertrophic cardiomyopathy.

Adapted from Maron, B.J., and Maron, M.S. (2013). Hypertrophic cardiomyopathy. Lancet 381, 242–255. 14

Although these procedures may improve symptoms, they have not been shown to

improve survival in HCM patients.

It is estimated that the incidence of sudden death in the HCM population is

around 1% (Maron et al., 1996b). Sudden cardiac death occurs due to ventricular

tachyarrhythmias that are caused by disarray of myocytes with interstitial fibrosis. The

risk factors for sudden death include: prior sustained ventricular tachyarrhythmias,

family history of sudden death, genetic mutations associated with sudden death, septal wall thickness >30 mm, recurrent syncope, and exertional hypotension. Patients with risk factors for sudden death are encouraged to have implantable cardioverter- defibrillators, and all patients with HCM are urged to minimize competitive sports and intense training.

There have been no randomized trials to evaluate treatment options in HCM patients, however, retrospective observational studies have been used to create general guidelines. In addition to counseling on treatment, it is recommended that the family of the patient undergo genetic testing since HCM is inherited in an autosomal dominant fashion. Screening echocardiography is typically performed in young first

degree relatives of the patient every 12-18 months and before competitive sport

activities. Screening of non-athlete adult relatives is currently recommended every five

years. If the particular mutation of the patient is known, genetic analysis is the

preferred method of screening in relatives. Competitive athletics are advised against

and mild aerobic exercises are permitted for a healthy lifestyle in HCM patients

(Ommen et al., 2011).

RESEARCH ADVANCES

There has been significant progress in describing the pathophysiology, genetics,

and treatment of HCM, however there is still much debate and uncertainty regarding

the diagnosis, natural history, and management of HCM (Maron and Maron, 2013).

The role of genetic testing in HCM is still poorly defined. It has been shown that the

occurrence of a positive genetic test for HCM is associated with greater incidence of

LV dysfunction (Figure 1.4, Olivotto et al., 2008), however pathogenic mutations are

only identified in fewer than 50% of clinically affected HCM patients (Bos et al.,

2009b; Maron et al., 2012a; Richard et al., 2003). This is largely due to significant

genetic heterogeneity, presence of mutations in non-analyzed sequences, incomplete

sensitivity of the screening procedure, or involvement of currently unidentified genes

(Richard et al., 2003). As the genetic analysis of patients with HCM expands, there

may be potential for more targeted therapies against the progression of LVH. A proof

of concept study has demonstrated that in vivo gene transfer of cMyBPC directly

injected into the myocardium of cMyBPC knock-out mice improved myofilament and

in vivo contractile function (Merkulov et al., 2012).

Clinical genetic testing in the 1990’s revealed a population of genotype-positive

phenotype-negative individuals (Rosenzweig et al., 1991), also titled preclinical HCM.

It has been challenging to define the best screening/diagnostic plan and resulting

management of genotype-positive phenotype-negative individuals (Maron et al.,

2011a), particularly due to the long period of follow-up necessary to make evidence-

based guidelines. 16

Figure 1.4: Outcomes related to genetic status in HCM. Relation of genetic status to the probability of cardiovascular death, nonfatal ischemic stroke, or progression to severe heart failure (New York Health Association [NYHA] functional classes III or IV)

Adapted from Olivotto et al. (2008). Myofilament Protein Gene Mutation Screening and Outcome of Patients With Hypertrophic Cardiomyopathy. Mayo Clinic Proceedings 83, 630–638.

Following genetic identification of HCM individuals, the phenotype is

characterized by diagnostic imaging. Traditionally, 2-dimensional echocardiography

is used to define the LV thickness, and thus if the patient is positive for a phenotype of

LVH. There have been several studies aiming to better characterize the subclinical

presentation of genotype-positive phenotype-negative individuals. Abnormalities such

as blood-filled myocardial crypts (Maron et al., 2012b), biomarkers of fibrosis (Ho et

al., 2010), and mitral leaf elongation (Germans et al., 2006; Maron et al., 2011b) have

all been described in the LV myocardium of subclinical HCM. The definition of a

positive phenotype has been expanded to include subclinical diastolic dysfunction in

the absence of overt LVH measured by Doppler imaging (Ho et al., 2002).

Cardiovascular MRI (CMR) is becoming the preferred imaging modality in describing

preclinical HCM given its better characterization of LVH as compared to

echocardiography(Germans et al., 2006; Maron et al., 2009b). In addition, contrast- 17

enhanced CMR has been used to characterize LV myocardial fibrosis in four unrelated

genotype-positive phenotype-negative HCM patients (Rowin et al., 2012).

The clinical management of genotype-positive phenotype-negative HCM

individuals remains problematic even as our understanding of the pathophysiology

improves. The identification of this subset of HCM patients only emerged in the past

two decades, so there is a lack of clinical data following their clinical course. In

particular, complex questions regarding abstinence from competitive sports will need

to be addressed, given that HCM is the most common cause of sudden death in young

athletes (Maron et al., 1996b, 2009a) and that there may be a decreased risk of death

from avoiding vigorous exercise (Pelliccia et al., 2005). There is also a need to evaluate

primary prevention of sudden death with ICDs in genotype-positive phenotype-

negative HCM patients. Although ICDs have been effective in terminating life-

threatening arrhythmias in HCM patients with marked LVH (Maron et al., 2007,

2013), there is no data regarding ICD placement in preclinical subjects. Thus, there is

a need to better understand and define the negative phenotype in genotype-positive

HCM patients.

DUCHENNE MUSCULAR DYSTROPHY INTRODUCTION

The dystrophin-glycoprotein complex (DGC) is thought to be responsible for maintaining the structure and morphology of myocytes. Dystrophin is a major actin- binding component of the dystrophin glycoprotein complex (DGC) and links cytoskeletal and membrane elements in the muscle (Tinsley et al., 1994). (Figure 1.5,

(Liew and Dzau, 2004)) The DGC provides stability to the sarcolemma, and disruption of the DGC may cause tears in the membrane, altered cell signaling, and ultimately muscle fiber necrosis.

Figure 1.5: Structural network in the cardiomyocyte.

The dystrophin‐glycoprotein complex (DGC) is believed to be involved in maintaining the structural integrity of the myocyte. The complex is comprised of dystrophin, dystrobrevin, dystroglycans, sarcoglycans, and syntrophins concentrated at costameres, which anchor and maintain spatial organization of myofibrils to the plasma membrane and are the sites of mechanical coupling between the extracellular matrix and sarcomere. Dystrophin binds to F‐actin to link the sarcomere and costamere.

Adapted from Liew, C.-C., and Dzau, V.J. (2004). Molecular genetics and genomics of heart failure. Nat Rev Genet 5, 811–825. 19

The DGC is also a necessary component of the neurovascular units in the brain (Figure 1.6,

(Abbott and Friedman, 2012)), which are formed by neurons, glia, and microvessels implicated in regulating cerebral blood flow

(Abbott et al., 2006). The blood- brain barrier (BBB), which is formed by the endothelial cells lining the Figure 1.6: The neurovascular unit. Schematic depiction of the components of the neurovascular cerebral microvessels, acts within unit: endothelial cells, pericytes, astrocytes, microglia, and neurons. the neurovascular unit to ensure the Adapted from Abbott, N.J., and Friedman, A. (2012). Overview and introduction: The blood–brain barrier in proper metabolic and physical health and disease. Epilepsia 53, 1–6. environment for neuronal function

(Mäe et al., 2011). The endothelial cells lining the vasculature of the brain are joined by tight junctions in order to prevent free diffusion of large substances from the blood into the brain parenchyma (Brightman and Reese, 1969; Reese and Karnovsky, 1967).

Dystrophin is necessary for anchoring of aquaporin 4 (AQP4) and tight-junctional components, which regulate the movement of water in the brain (Nico et al., 2004)

(Figure 1.7, (Wolburg et al., 2009).

Figure 1.7: Endothelial and astroglial cell interaction at the blood‐brain barrier. Schematic depiction of the components at the blood‐brain barrier (BBB). The dystrophin‐ dystroglycan complex (DGC) can be seen in the astrocytic endfoot (top), which anchors the water channel protein aquaphorin‐4 (AQP4). Adapted from Wolburg, H., Noell, S., Mack, A., Wolburg-Buchholz, K., and Fallier-Becker, P. (2009). Brain endothelial cells and the glio-vascular complex. Cell Tissue Res. 335, 75–96.

CLINICAL PICTURE

Duchenne’s muscular dystrophy (DMD) is an invariably fatal X-linked

recessive disorder in which the cytoskeletal protein dystrophin is defective and therefore progressively disrupts muscle fiber function. The loss of muscle strength in boys with Duchenne’s muscular dystrophy is progressive, and is more severe in the proximal limb (legs more so than arms) and neck flexor muscles. The incidence of

Duchenne’s dystrophy is approximately 1 in 3000-4000 live-born males (Ropper et al.,

2014).

PATHOLOGY

There are two primary theories proposed to elucidate the pathogenesis of DMD

(Figure 1.8 (Deconinck and Dan, 2007). The first is that a lack of dystrophin

destabilizes the sarcolemma and results in muscle fiber damage from repetitive

contractions, particularly eccentric contraction (Mokri and Engel, 1998; Pestronk et

al., 1982; Petrof et al., 1993). The dystrophin-associated protein complex form

costameres that anchor the cytoskeleton to the extracellular matrix (Campbell, 1995;

Rybakova et al., 2000), and its disruption leads to membrane fragility. Secondly,

Figure 1.8: Proposed pathophysiology of Duchenne Muscular Dystrophy.

Adapted from Deconinck, N., and Dan, B. (2007). Pathophysiology of Duchenne Muscular Dystrophy: Current Hypotheses. Pediatric Neurology 36, 1–7. 22

disruption of the DGC alters calcium homeostasis in the muscle cell(Hack et al., 1999;

Rafael et al., 2000). The first evidence of calcium dysregulation came from muscle

biopsies of DMD patients that demonstrated accumulation of calcium and hypercontracted fibers (Bodensteiner and Engel, 1978; Cullen and Fulthorpe, 1975;

Duncan, 1978). Prior studies have established that there is a larger influx of calcium

into the cell, likely involving the TRPC calcium channel (De Backer et al., 2002;

Franco and Lansman, 1990; Tutdibi et al., 1999; Vandebrouck et al., 2002). The sustained increase in intracellular calcium leads to activation of proteases that destroy membrane components and ultimately leads to cell death (Spencer and Tidball, 1996;

Spencer et al., 1995).

GENETICS

Duchenne’s muscular dystrophy is the result of a mutation of the gene that

encodes for the cytoskeletal protein, dystrophin. The dystrophin gene is one of the

largest in the human genome, larger than 2000 kb in size, and is on the short arm of

the X chromosome at Xp21. The disease is most commonly caused by a deletion, and

the size of the mutation does not correlate with the severity of symptoms (Amato and

Brown, 2012). There are several genetic mutations that may lead to DMD. Point

mutations (approximately 10% of DMD cases) may generate premature termination

codons that produce a truncated protein, or the point mutation may affect a critical

domain of the dystrophin protein. Deletion or insertion of a single nucleotide disrupt

the open reading frame of the messenger RNA (mRNA) for dystrophin, which

significantly alters the structure of the protein. In addition, exon-intron splice site

mutations can also alter the open reading frame of dystrophin mRNA. Reading-frame 23

alterations are the most common mutations of the dmd gene (Amato and Brown,

2012). Mutations that result in a partially functional dystrophin protein clinically lead to a milder form of the disease, known as Becker muscular dystrophy (Ropper et al.,

2014). Approximately 30% of cases do not have any family history of dystrophy, which represents the rate of spontaneous mutations. In rare cases, females may exhibit

symptoms of Duchenne’s muscular dystrophy. For example, a female may inherit only

one X chromosome (Turner syndrome) that carries the dystrophin mutation.

PRESENTATION

Duchenne’s muscular dystrophy is present at birth, however, it is not typically

diagnosed until the boy is between 3 and 5 years old (Ropper et al., 2014). Children

with Duchenne’s have difficulty playing, fall frequently, and will demonstrate

abnormal running and jumping. A classic sign is the Gowers’ maneuver, in which the

boy will need to use his hands to gradually climb up the shins, knees, and torso when

getting up from the floor. By the age of 6 years, boys will exhibit toe walking from

gastrocnemii weakness and contractures of the heel cords and iliotibial bands. A

lordotic posture results from weakness of the abdominal and paravertebral muscles. It

is typical for the boys to have enlarged gastrocnemii (pseudo-hypertrophy) that have a

firm and rubbery feel due to the replacement of muscle tissue with connective and

adipose tissue. Patients waddle when walking and have a wide stance when standing

from gluteus medius weakness and instability. By the age of 10, walking requires the

use of braces, and most boys are wheelchair-bound by the age of 12. Joint contractures

of hip flexion, knee, elbow, and wrist extension may become fixed and painful. 24

Tendon reflexes eventually diminish and are not present once the muscle has

atrophied.

In addition to diaphragmatic weakness, progressive scoliosis is typical and

impairs pulmonary function. Pulmonary infections may be fatal in Duchenne’s muscular dystrophy teenage boys and is a common cause of mortality. Aspiration of food and acute gastric dilation are also common causes of death. The life expectancy in DMD does not commonly extend beyond the third decade (Annexstad et al., 2014;

Eagle et al., 2002b; Kieny et al., 2013).

Cardiomyopathy is present in almost all Duchenne’s muscular dystrophy boys by the age of 18 (Nigro et al., 1990). Similar to the pathogenesis in skeletal muscle, the necrosis of cardiac muscle fibers triggers fibrosis. Cardiac involvement typically includes progressive atrioventricular block, arrhythmia, dilated cardiomyopathy, and akinesis/dyskinesis of the posterobasal wall of the left ventricle. An ECG may display prominent R waves in the right precordial leads, deep Q waves in the left precordial

and limb leads, and diminished P-wave amplitude.

Intellectual impairment is common, with boys testing on average one standard

deviation below the mean for intelligence quotient (IQ) and commonly affecting verbal performance more than anything else (Bresolin et al., 1994a). The etiology of cognitive

impairment is not well understood in DMD, however studies have suggested that

hypometabolism may have a role (Al-Qudah et al., 1990; Tracey et al., 1995, 1996a).

There is also evidence of cerebrovascular disease in young adult DMD patients,

demonstrated by case reports of cerebral infarctions (Table 1.1, (Tsakadze et al., 2011))

(Díaz Buschmann et al., 2004; Ikeniwa et al., 2006; Matsuishi et al., 1982; Tsakadze 25

et al., 2011). Cerebral infarction has been reported in 0.75% of DMD patients aged 16-

20 years (Hanajima and Kawai, 1996a), however this may be significantly

underestimated given the difficulty in detecting symptoms in setting of severe

dystrophy. It is thought that the etiology of cerebral infarction in DMD is due to

cardio-embolic events as a result of cardiomyopathy.

DIAGNOSIS

Serum CK levels are elevated 20-100 times the normal level in Duchenne’s

muscular dystrophy (Amato and Brown, 2012). An elevated CK may be detected at

birth, and will progressively increase until the end stages of the disease where the

patient is inactive and has lost significant muscle mass. A muscle biopsy (typically

from the gastrocnemius) from a Duchenne’s muscular dystrophy boy demonstrates

fibers of varying size, necrosis, regeneration, and connective tissue and adipose tissue

that has replaced muscle. An official diagnosis of Duchenne’s muscular dystrophy is

Table 1.1: Case reports of cerebral infarction in DMD patients.

Adapted from Tsakadze, N., Katzin, L.W., Krishnan, S., and Behrouz, R. (2011). Cerebral Infarction in Duchenne Muscular Dystrophy. Journal of Stroke and Cerebrovascular Diseases 20, 264–265. 26 made by Western blot analysis of the biopsy, which shows abnormal quantity and size of the dystrophin protein. Immunohistochemical analysis of the muscle may also

Figure 1.9: Stages of Duchenne Muscular Dystrophy and Treatment Options.

Adapted from Bushby, K., Finkel, R., Birnkrant, D.J., Case, L.E., Clemens, P.R., Cripe, L., Kaul, A., Kinnett, K., McDonald, C., Pandya, S., et al. (2010). Diagnosis and management of Duchenne muscular dystrophy, part 1: diagnosis, and pharmacological and psychosocial management. The Lancet Neurology 9, 77–93. 27

demonstrate dystrophin deficiency or deletion. An EMG shows fibrillations, positive waves, and low-amplitude and brief polyphasic motor unit potentials.

TREATMENT

The current treatment of DMD is multi-disciplinary and involves therapies

directed towards alleviating symptoms, including corticosteroids, respiratory, cardiac,

orthopedic, and rehabilitative interventions (Figure 1.9 (Bushby et al., 2010b).

Glucocorticoids are the standard of care and have been shown to delay the progression

of DMD for up to 2 years by improving muscle strength and function (Manzur et al.,

2008; Moxley et al., 2010). However, many patients decline this therapy given the

significant side effect profile, including weight gain and increased risk of fractures in a

population that is already at danger for falls. ACE inhibitors and beta-blockers are the

first-line agents for treating the progression of cardiomyopathy in DMD (Bushby et

al., 2010a).

RESEARCH ADVANCES

There has been significant emphasis on improving strategies for DMD gene

and pharmacologic therapies over the past decade. Animal models of DMD have been

invaluable for evaluating the efficacy of such investigations. The dystrophin-null mdx

mouse has been used as a mouse model of muscular dystrophy for over two decades

(Sicinski et al., 1989). In addition, skeletal or cardiac specific deficiency of dystrophin

has been studied in dogs. The GRMD (Golden Retriever Muscular Dystrophy) is the

most accurate animal model to study given its homology to the clinical disease

(Kornegay et al., 2012). 28

Table 1.2: Overview of strategies for Duchenne Muscular Dystrophy gene therapy.

Adapted from Van Deutekom, J.C.T., and van Ommen, G.-J.B. (2003). Advances in Duchenne muscular dystrophy gene therapy. Nat Rev Genet 4, 774–783.

There has been significant focus on investigating the use of gene therapy to

restore the defective dystrophin gene or induce dystrophin production by correcting

the underlying mutation as a treatment in DMD since it has the potential to be curative

(Table 1.2 (van Deutekom and van Ommen, 2003)(Foster et al., 2012; Konieczny et

al., 2013; Malik et al., 2012). Genetic strategies have included viral-mediated insertion

of either full-length or truncated dystrophin transgenes, antisense oligonucleotides to

induce exon skipping and reestablish the dystrophin reading frame, agents that read-

through stop codon mutations, and upregulating surrogate proteins such as utrophin

to replace dystrophin. Stem cell therapies have focused on restoring the depleted

muscle fibers and inducing a functional dystrophin gene. The observation that patients with Becker muscular dystrophy maintain ambulatory capabilities and typically do not exhibit symptoms until well into adulthood led researchers to believe that even the 29

partial restoration of functional dystrophin may be beneficial in DMD patients

(Bushby et al., 1993; Malhotra et al., 1988). It has been shown in mice and humans

that the minimum level of dystrophin necessary to prevent muscular dystrophy is 20-

30% (Sharp et al., 2011).

Nonsense mutations that result in premature termination codons (PTC) may

be addressed by administering molecules (gentamycin and ataluren) that allow a read-

through of the PTC in mRNA. Approximately 15% of DMD patients have nonsense

mutations from single nucleotide DNA polymorphisms that lead to premature in-

frame stop codons and would therefore benefit from development of these drugs (Dent

et al., 2005). Administration of ataluren to prevent nonsense mutations has been studied in both mdx mice and DMD patients, which was able to restore some

functional dystrophin expression and slow disease progression (Barton-Davis et al.,

1999; Hoffman and Connor, 2013; L et al., 2003; Peltz et al., 2013; Wagner et al.,

2001; Welch et al., 2007).

Disruption of the open reading frame is the primary source of DMD cases

(approximately 80%), either through deletions/insertions or splicing mutations. Exon skipping is a tactic to restore the reading frame of the dystrophin gene (van Deutekom and van Ommen, 2003; Foster et al., 2012). Specific antisense oligonucleotides

(AONs) are synthetic, short nucleic acid polymers that bind to the mutation site in the pre-mRNA to skip over the offending exon and restore the reading frame (Summerton and Weller, 1997). There have been great advancements in drug therapy for exon skipping in DMD patients. Trials are currently underway studying two antisense oligomer drugs (eteplirsen and drisapersen) that target a DMD gene hot spot (exon 30

51), however the drisapersen trial was suspended due to concerns of efficacy (Hoffman

and Connor, 2013).

Given the recent advances along multiple fronts for the treatment of DMD, it

is reasonable to assume that the median life expectancy of DMD will increase.

Accordingly, the prevention, diagnosis, treatment for extra-cardiac manifestations of

dystrophin dysfunction will likely take on new importance. Specifically, the role of

dystrophin in the pathophysiology of cerebrovascular disease is poorly defined and

awaits delineation with the use of advanced in vivo imaging.

Magnetic Resonance Imaging

DISPLACEMENT ENCODING WITH STIMULATED ECHOES

Cardiovascular magnetic resonance (CMR) provides a non-invasive and accurate representation of cardiovascular function. In particular, techniques have been

developed to measure the mechanics of myocardial wall motion, such as strain, twist,

and torsion. The myofibers of the heart are arranged in a right-handed helix in the

subendocardium and transition to a left-handed helix in the subepicardium. This results in LV wall thickening in the radial direction and shortening in the longitudinal direction during contraction (Sengupta et al., 2006). Myocardial strain can be described as the deformation of the myocardial wall, and is defined in three orthogonal

planes: radial, circumferential, longitudinal (Figure 1.10, Jiang and Yu, 2014). A

positive strain represents lengthening or thickening and a negative strain is

representative of shortening of the myocardium. Strain rate is characterized as the rate

of change of strain over the course of the cardiac cycle, and is frequently used as an

index of regional LV function (Edvardsen et al., 2006). CMR is also useful in

describing the wringing or twisting motion of the heart. The base rotates in a clockwise

direction and the apex in a counterclockwise direction when viewed from the apex

(Figure 1.10, Jiang and Yu, 2014), which is quantified as the twist angle. Torsion is

defined as the difference in the twist angle from the apical and basal slices normalized

by the distance between the two slices. Torsion and twist have been shown to be

sensitive to subtle myocardial systolic and diastolic dysfunction since they are affected

by myofiber orientation, contractility, and loading conditions (Burns et al., 2008;

Dong et al., 1999; Li et al., 2009b; Stuber et al., 1999). Displacement encoding with 32 stimulated echoes (DENSE) is one method that was developed by Aletras et al. in 1999 to provide high spatial density of tissue displacement via stimulated echoes (Aletras et al., 1999). The myocardial displacement is encoded directly in the phase of the stimulated echo, and can therefore provide myocardial strain and twist data. A representative displacement map is shown in (Figure 1.10, Jiang and Yu, 2014).

Figure 1.10: Left ventricular mechanics during contraction. (A) Myocardial wall strains during

diastole and systole in the Radial‐Circumferential‐Longitudinal (RLC) system: radial (ERR),

circumferential (ECC), and longitudinal (ELL) strains, and the principal system: along the

direction of greatest length increase in myofibers (E11), greatest length decrease in myofibers

(E22), and orthogonal to both E11 and E22 (E33). (B) LV wall torsional deformation during contraction. The apex twists counterclockwise and base twists clockwise viewing from apex to base. (C) Representative 2‐D displacement map at peak systole as measured by displacement encoding with stimulated echoes (DENSE).

Adapted from Jiang, K., and Yu, X. (2014). Quantification of regional myocardial wall motion by cardiovascular magnetic resonance. Quant Imaging Med Surg 4, 345–357. 33

ARTERIAL SPIN LABELING

Arterial spin labeling (ASL) MRI is a non-contrast perfusion MRI technique

that has been shown to provide quantitative assessments of tissue perfusion in several

applications (De Bazelaire et al., 2005; Katada et al., 2012; Mai and Berr, 1999;

Martirosian et al., 2004; Wong et al., 1997). In essence, ASL is similar to traditional

contrast perfusion techniques in that it “magnetically tags” arterial blood water,

leading to altered tissue longitudinal magnetization that is proportional to tissue

perfusion. ASL MRI techniques generate blood flow contrast between multiple images

using a wide variety of blood labeling methods (Detre and Alsop, 1999; Detre et al.,

1992; Edelman et al., 1994; Kim and Tsekos, 1997; Kwong et al., 1995; Williams et

al., 1992). A typical ASL MRI acquisition combines an ASL preparation phase,

followed by a rapid imaging readout to capture the blood flow-weighted contrast.

One major advantage of ASL over conventional dynamic contrast-enhanced

(DCE) MRI perfusion techniques is the lack of exogenous, and potentially toxic,

paramagnetic contrast agents, which is especially important for the imaging of patients

with chronic kidney diseases (Kuo et al., 2007). This attribute is also important for

longitudinal preclinical imaging applications, as multiple tail-vein injections and/or

catheterizations can cause local inflammation and necrosis, resulting in reduced access

to veins.

DIFFUSION-WEIGHTED MAGNETIC RESONANCE IMAGING

Diffusion-weighted MRI (DWI) is a technique that has become invaluable in

defining neurological disorders, particularly in the diagnosis and therapeutic

intervention of stroke (Le Bihan et al., 1986; Sevick et al., 1990; Warach et al., 1995; 34

Schellinger et al., 2003; Kloska et al., 2010). The signal intensity of a DW image is

representative of the Brownian motion of water molecules in the tissue, and the

calculated apparent diffusion coefficient (ADC) provides a means to quantify this

diffusion under physiologic and pathologic states. High ADC values are characteristic

of a tissue with relatively free water diffusion as opposed to tissue water with a

restricted environment (Bihan, 2007). Following cerebral ischemia, the metabolic

production of ATP is halted and therefore leads to sodium and calcium retention in

the cell from non-functioning ionic pumps (Ito et al., 1979). The osmotic influx of

water into the cell results in cytotoxic edema (Klatzo, 1987) that decreases the

available extracellular space for diffusion of water. This pathologic response to

ischemia is supported by an immediate decrease in ADC which occurs before any

significant changes in a T2-weighted image (Moseley et al., 1990; Kucharczyk et al.,

1991).

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CHAPTER 2: CARDIAC MYOSIN BINDING PROTEIN C INSUFFICIENCY

LEADS TO EARLY ONSET OF MECHANICAL DYSFUNCTION

ABSTRACT

Background—Decreased expression of cardiac myosin binding protein C (cMyBPC)

as a result of genetic mutations may contribute to the development of hypertrophic

cardiomyopathy (HCM); however, the mechanisms that link cMyBPC expression and

HCM development, especially contractile dysfunction, remain unclear. Methods and

Results — We evaluated cardiac mechanical function in vitro and in vivo in young

mice (8–10 weeks of age) carrying no functional cMyBPC alleles (cMyBPC−/−) or 1

functional cMyBPC allele (cMyBPC±). Skinned myocardium isolated from

cMyBPC−/− hearts displayed significant accelerations in stretch activation cross-bridge

kinetics. Cardiac MRI studies revealed severely depressed in vivo left ventricular (LV)

magnitude and rates of LV wall strain and torsion compared with wild-type (WT)

mice. Heterozygous cMyBPC± hearts expressed 23±5% less cMyBPC than WT hearts

but did not display overt hypertrophy. Skinned myocardium isolated from

cMyBPC± hearts displayed small accelerations in the rate of stretch induced cross-

bridge recruitment. MRI measurements revealed reductions in LV torsion and

circumferential strain, as well reduced circumferential strain rates in early systole and

diastole. Conclusions—Modest decreases in cMyBPC expression in the mouse heart

result in early-onset subtle changes in cross-bridge kinetics and in vivo LV mechanical

function, which could contribute to the development of HCM later in life.

INTRODUCTION

Hypertrophic cardiomyopathy (HCM) is one of the most commonly occurring

genetic myocardial disorders, affecting approximately 1 in 500 people (Maron et al.,

1995). Cardiac myosin binding protein C (cMyBPC) is a thick filament protein that

modulates actin-myosin interactions and thereby the rate of muscle contraction

(Carrier et al., 2007, Barefield et al., 2010). It is well established that the gene encoding

cMyBPC is one of the most common causes of inherited HCM, with nearly 200 known

mutations identified (Harris et al., 2011) since the first reported mutation in this gene

(Watkins et al., 1995, Bonne et al., 1995). However, many younger individuals who

carry disease-causing mutations in the cMyBPC gene do not exhibit overt LVH,

because increases in LV wall thickness are often only detectable with advanced age

(Maron et al., 2004, Niimura et al., 1998). Because these seemingly asymptomatic

carriers are at risk for the development of HCM and cardiac disease later in life, the

diagnosis and treatment of these patients is a major clinical challenge.

The majority of mutations in the gene that encode cMyBPC are heterozygous

and are predicted to result in expression of truncated cMyBPC lacking the C-terminal

regions of the protein that binds to myosin and titin (Carrier et al., 1997). However,

analysis of myocardial biopsy samples from patients with cMyBPC mutations have

not detected truncated cMyBPC, but rather a reduction in the amount of full-length

cMyBPC protein has been noted (Rottbauer et al., 1997, Moolman et al., 2000, van

Dijk et al., 2009, Marston et al., 2009, Jacques et al., 2008). It is therefore likely that

mutant cMyBPC mRNA or proteins are rapidly degraded by nonsense mediated

mRNA decay or the ubiquitin-proteosome system, thereby preventing mutant proteins 55

from incorporating into the sarcomere (Schlossarek et al., 2011). Therefore, the allele

generating mutant cMyBPC effectively functions as a null allele, causing cMyBPC

haploinsufficiency. However, the mechanisms that link reduced cMyBPC levels in the

heart with the development and progression of HCM have remained elusive.

Considering the functional importance of cMyBPC in regulating myofilament

contractile properties, it is reasonable to suppose that decreased cMyBPC expression

could affect in vivo mechanical function. In this regard, it has been shown that

homozygous cMyBPC knockout mice (cMyBPC−/−) with 2 null cMyBPC alleles (i.e.,

a model of pure insufficiency) display early-onset impairments in systolic and diastolic

contractile function and severe LVH (Harris et al., 2002, Palmer et al., 2004,

Nagayama et al., 2007). In contrast, heterozygous cMyBPC knockout mice

(cMyBPC±) with 1 null cMyBPC allele develop a phenotype later in life, displaying

modest hypertrophy despite preserved systolic and diastolic contractile function

(Carrier et al., 2004). Interestingly, cMyBPC± hearts expressed ≈25% less total

cMyBPC than aged-matched wild-type (WT) mice (Carrier et al., 2004), which is

similar to the amount of full length cMyBPC in patients with heterozygous cMyBPC

mutations (van Dijk et al., 2009, Marston et al., 2009). These results suggest that

modest decreases in cMyBPC expression in the heart may be sufficient to produce

cardiac dysfunction and/or LVH; however, it is has not been established if these

changes are related to altered cardiac mechanical performance.

Therefore, in the present study we examined the effects of variable cMyBPC

expression on in vitro and in vivo mechanical function in young (8–10 weeks of age)

cMyBPC−/− and cMyBPC± mice to determine if cMyBPC insufficiency can cause 56

mechanical dysfunction early in life. We used MRI to quantify both global and

regional mechanical indices such as LV twist, torsion, and principal strains, over the

whole cardiac cycle to observe subtle changes in mechanical function. Our goal was

to link cMyBPC expression and cross-bridge function with LV strain and torsion,

which are direct measures of myocardial wall deformation, to characterize the

functional consequences of cMyBPC insufficiency.

METHODS

Animal Models

Adult male cMyBPC null (cMyBPC−/−), heterozygous cMyBPC mice

(cMyBPC±), and WT mice of the SV/129 strain (8–10 weeks of age) were used in this

study (Harris et al., 2002). All procedures involving animal care and handling were

performed according to institutional guidelines set forth by the Animal Care and Use

Committee at Case Western Reserve University.

Skinned Fiber Experiments

Force-pCa relationships and stretch activation kinetics were measured in

skinned ventricular myocardium isolated from WT, cMyBPC−/−, and cMyBPC± hearts

as previously described (Chen et al., 2010, Stelzer et al., 2006). Myofibrillar protein

content and phosphorylation were determined in LV homogenates and skinned

myocardium as previously described (Chen et al., 2010, Stelzer et al., 2007).

In Vivo MRI Measurements of Cardiac Function

Two-dimensional LV myocardial motion was quantified at the apex, mid, and

basal levels using multiphase displacement encoding with stimulated-echo (DENSE)

MRI in a 9.4-T Bruker Biospec (Billerica, MA) horizontal scanner (Liu et al., 57

2006). LV twist, torsion, circumferential and radial strain, as well as torsion and strain

rates were quantified, using custom software as previously described (Zhong et al.,

2008).

Statistical Analysis

Cross-sectional areas of skinned preparations were calculated by measuring the

width of the mounted preparation and assuming a cylindrical cross section.

2+ Submaximal Ca -activated force (P) was expressed as a fraction of the force (Po)

generated at pCa 4.5, that is, P/Po. Rate constants of stretch-induced force decay (krel)

and development (kdf) were obtained by fitting the time course trace with a single

exponential. Data are reported as mean±SD or mean±SEM. Means for fiber data were

generated by averaging data for each variable for each mouse and then averaging the

means for all mice within a group. Approximately 3 fibers were studied from each

mouse. Comparisons of force-pCa relationships and stretch activation variables

between groups were done using a 1-way ANOVA and a Tukey-Kramer post hoc test.

Comparisons of strain and twist angles were independently analyzed at each

ventricular level by 1-way ANOVA and Tukey-Kramer test. Strain and torsion time

course data were evaluated by 1-way ANOVA with a Bonferroni adjustment for

multiple comparisons. Additionally, a separate analysis examining only peak rates of

torsion and strain in systole and diastole was performed using 1-way ANOVA and

Tukey-Kramer post hoc test. Statistical significance was established at a level

of P<0.05.

RESULTS 58

Myofilament Protein Expression and Phosphorylation in WT and cMyBPC

Mutant Myocardium

We examined the expression and phosphorylation status of myofilament

proteins in age-matched male WT, cMyBPC−/−, and cMyBPC± LV tissue

homogenates. As expected, cMyBPC was not detected in cMyBPC−/− hearts in either

Coomassie-stained gels or Western blots probed with a cMyBPC specific antibody

(Santa Cruz, CA); however, cMyBPC± hearts expressed 23±5% less cMyBPC than

WT hearts (P<0.05, Figure 2.1). Consistent with previous findings, (Harris et al.,

2002) expression of β-myosin heavy chain (MHC) was slightly elevated in

cMyBPC−/−hearts (16±3%, P<0.05) compared with WT hearts, but there was no

significant difference in MHC isoform expression between cMyBPC± and WT hearts

(Figure 2.2). No significant differences in the relative abundance or phosphorylation

status of other myofilament proteins in WT, cMyBPC−/−, and cMyBPC± myocardium

were detected (Figure 2.3). The cMyBPC content of skinned myocardium isolated

from cMyBPC± LV used for mechanical experiments was also quantified (Table 2.1)

and was nearly identical to the cMyBPC content measured in LV tissue homogenates

prepared from cMyBPC± hearts (ie, reduced 24±6% compared with WT).

Figure 2.1. Quantification of cMyBPC expression in cMyBPC+/- myocardium. A representative Western blot of whole tissue homogenates probed with a cMyBPC specific antibody. Lane 1, cMyBPC- /-, lane 2 cMyBPC+/-, and lane 3 WT. cMyBPC protein content was normalized to α-actinin content. On average cMyBPC+/- hearts expressed 23% less cMyBPC than WT hearts. 59

Figure 2.2. Quantification of myosin heavy chain (MHC) isoform expression in WT and cMyBPC mutant myocardium. A representative 6% SDS-PAGE coomassie stained gel of whole tissue homogenates prepared from: Lane 1, cMyBPC-/-, lane 2 cMyBPC+/-, and lane 3 WT, myocardium. α- MHC, alpha myosin heavy chain, β-MHC, beta myosin heavy chain. The β-MHC isoform was detected only in cMyBPC-/- myocardium.

Figure 2.3. Myofibrillar protein content and phosphorylation in WT and cMyBPC mutant myocardium. (A) Representative 4- 12% SDS-PAGE gel of myofibrillar proteins isolated from skinned myocardium from cMyBPC+/- (lanes 1 and 4), WT (lanes 2 and 5), and cMyBPC-/- (lanes 3 and 6) hearts and stained with coomassie (lanes 1-3) to detect total proteins and Pro-Q Diamond to detect phosphorylated proteins (lanes 4-6). cMyBPC; cardiac myosin binding protein C, TnT; troponin T, TnI; troponin I, RLC; regulatory light chain. (B-E) Slopes of phosphorylation signals from Pro-Q Diamond stained gels (n=6) determined from regression analysis of plots of area x average optical density versus protein loaded (µg) for (B) cMyBPC, (C) RLC, (D) TnT, and (E) TnI. Data are means ± SD. 60

Mechanical Properties of Skinned Myocardium Isolated From WT and cMyBPC

Mutant Hearts

The steady-state mechanical properties of skinned myocardium isolated from

WT, cMyBPC−/−, and cMyBPC± hearts are summarized in the Table 2.1. Skinned

preparations from the 3 groups of mice exhibited similar force-pCa relationships.

There were no differences in force generation at maximal and submaximal [Ca2+] or

in the steepness of the force-pCa relationship (Hill coefficient, nH). Consistent with

previous studies (Stelzer et al., 2007), cMyBPC−/− myocardium displayed dramatically

accelerated rates of stretch-induced force decay (krel) and delayed force development

2+ (kdf) at submaximal Ca activations as well as greater stretch-induced force decay

(P2 amplitude) and stretch activation amplitude (Pdf), compared with WT myocardium

(Figure 2.4). Skinned myocardium isolated from cMyBPC± hearts displayed small but

significant (21%) accelerated stretch-induced delayed force development (kdf) at

2+ submaximal Ca activations, but P2 and Pdf amplitude were not significantly different

compared with WT (Table 2.2).

Figure 2.4. (A) Stretch activation kinetics in skinned myocardium. The rates of A, delayed force development (kdf), and B, force relaxation (krel) after a stretch of 1% of muscle length, were recorded from wild-type (WT), cardiac myosin binding protein C (cMyBPC+/-), and cMyBPC-/- skinned myocardium activated with Ca2+ to a prestretch isometric force of ≈50% maximal. Data are mean ± SD. *Significantly different from WT.

2.2

In Vivo Assessment of Cardiac Morphology and Mechanical Function

LV Morphology and Contractile Function

Cardiac morphology and global contractile function analysis of WT, cMyBPC±, and cMyBPC−/− hearts are presented in Table 2.3. The cMyBPC−/− hearts displayed significant LV hypertrophy as demonstrated by increased end-diastolic wall thickness and decreased ejection fraction compared with WT controls. In contrast, 62

± 2.3 cMyBPC hearts displayed a

small increase in apical

thickening at end-diastole

compared with WT hearts

(Table 2.3, P<0.05), but

systolic function was

preserved, as indicated by

near-normal ejection fraction

values. Global hypertrophy

was seen in cMyBPC−/− hearts,

as demonstrated by greater

Figure 2.5. Cardiac hypertrophy analysis. (A) Heart-weight to body-weight ratios (HTWT/BWT), 8 hearts from each group. Relative abundance of mRNA of molecular markers of cardiac hypertrophy (values normalized to glyceraldehyde 3-phosphate dehydrogenase, GAPDH abundance): (B) Atrial natriuretic factor (ANF), (C) brain natriuretic factor (BNP), (D) alpha-skeletal actin (α-sk actin). Values are expressed as means ± SD. *Significantly different from WT. 63

heart weight–to–body-weight (HTWT/BWT) ratios compared with WT hearts. There

was no difference in HTWT/BWT ratio between cMyBPC± and WT hearts (Figure

2.5). Molecular markers of hypertrophy, including transcript expression of ANF,

BNP, and α-skeletal actin were also significantly upregulated in cMyBPC−/− hearts but

not in cMyBPC± hearts (Figure 2.5).

Myocardial Strain

Representative displacement maps for the in-plane motion of the WT,

cMyBPC±, and cMyBPC−/− groups in the mid-ventricle at peak systole are presented

in Figure 2.6. The overall reduction in displacement magnitude (Figure 2.6C) for strain

vectors displayed by cMyBPC−/− hearts compared with WT hearts is consistent with

significant contractile dysfunction.

Figure 2.6. Representative MRI principal strain vector displacement maps. Representative DENSE displacement maps at the mid-ventricle during peak systole for A, wild-type (WT); B, cardiac myosin binding protein C (cMyBPC)+/-; and C, cMyBPC-/-. The direction of radial (solid arrow) and circumferential strain (dashed arrow) is labeled in (A).

Mean LV strain was calculated at the apex, base, and mid-ventricle for each group.

There was a significant reduction in maximal LV radial and circumferential strains in

the cMyBPC−/− mice for all 3 ventricular levels (Figure 2.7), and there was a significant 64

reduction in maximal LV

circumferential strain at the

mid-LV of cMyBPC± mice

compared with WT mice

(Figure 2.7B, P<0.05).

Measurements of average

radial (Figure 2.8A

through 2.8C) and

circumferential strain rates

(Figure 2.8D through 2.8F)

were significantly reduced in

cMyBPC−/− mice throughout

the cardiac cycle. Reduced

early systolic strain rates were Figure 2.7. Measurements of left ventricular (LV) global principal strains. Global radial (A) and circumferential (B) strain observed in cMyBPC± mice at the base, mid, and apex in the LV of wild-type (WT), cardiac myosin binding protein C (cMyBPC)+/-; and cMyBPC-/- mice. Data are mean ± SE. *Significantly different from WT. at the mid-LV (Figure 2.8).

Analysis of peak strain rates revealed that cMyBPC± mice displayed lower peak radial

systolic strain rates in the base and mid-LV, and lower peak circumferential diastolic

strain rates in the mid-LV (Figure 2.8G through 2.8L).

Ventricular Twist and Torsion

Figure 2.9A illustrates the difference in twist (rotation) angles between WT,

cMyBPC±, and cMyBPC−/− mice at the apical, mid-LV, and basal levels during peak

systole. There was a clear decrease in overall rotation in cMyBPC−/− hearts. LV 65

Figure 2.8. Analysis of principal strain rates. Time course of radial (A through C) and circumferential (D through F) strain rates were quantified throughout the cardiac cycle, and peak radial (G through I) and circumferential (J through L) strain rates were quantified (from top to bottom) at the base, mid-ventricle, and apical segments in wild-type (WT), cardiac myosin binding protein C (cMyBPC)+/-; and cMyBPC-/- mice. Data are mean ± SE. *Significantly different from WT by 1-way ANOVA with Bonferroni (A through F) or Tukey-Kramer (G through L).

Figure 8 continued on next page. twisting occurred in the counterclockwise direction at the apex in the WT (10.12±1.9°) and cMyBPC± hearts (7.59±1.3°, P=NS), but minimal apical twisting was observed in cMyBPC−/− hearts (0.83±0.9°, P<0.05). Clockwise twisting was observed at the base in WT (−6.48±1.0°) and cMyBPC± hearts (−4.90±0.40°, P=NS), but minimal rotation occurred in cMyBPC−/− hearts (−1.39±0.84°, P<0.05). 66

The net twist, defined as the Figure 8 continued. difference between the twist

angles at the base and apex, is

plotted in Figure 2.9B. There

was a significant reduction in

net twist in

cMyBPC−/− (2.23±0.61°) and

cMyBPC± (12.49±1.08°)

hearts compared with WT

hearts (16.60±1.46°). In

particular, net twist between

the mid and basal LV

segments was reduced in

cMyBPC± hearts compared

with WT (Figure 2.9A,

3.24±2.0° and 5.98±4.5°,

respectively), although this

difference fell short of

statistical significance

(P=0.08).

There was a significant reduction in the magnitude of peak torsion generated by

cMyBPC−/− and cMyBPC± hearts compared with WT (50.8°/cm ±5.7): 8.9°/cm ±2.2

in cMyBPC−/− and 39.0°/cm ±2.4 in cMyBPC± (Figure 2.10A and 2.10C), and peak 67

systolic torsion occurred

earlier in cMyBPC−/− hearts.

Maximal systolic and

diastolic torsion rates were

lower in cMyBPC−/− hearts

(Figure 2.10B, 2.10D, 2.10E).

DISCUSSION

There is growing evidence

that decreased cMyBPC

expression may be a common

feature of HCM related to

mutations in the gene

encoding cMyBPC (van Dijk

Figure 2.9. Left ventricular twist relationships during systole. A, et al., 2009, Marston et al., Maximal twist angles (degrees) from the apex, mid-ventricle, and base during systole, and B, net twist angles (apex-base) 2009); however, the link during systole of the wild-type (WT), cardiac myosin binding protein C (cMyBPC)+/-; and cMyBPC-/- mice. Data are mean ± between cMyBPC SE. *Significantly different from WT. expression, cardiac mechanical function, and the development of HCM is not well

understood. We show that decreased cMyBPC expression in young cMyBPC± mice

leads to altered myofilament function and in vivo torsion and principal strains in the

absence of overt hypertrophy. Our data suggest that mechanical dysfunction exists in

preclinical cMyBPC related HCM.

Figure 2.10. Analysis of left ventricular torsion. Time course of ventricular torsion (A) and torsion rates (B) in 1 cardiac cycle for wild-type (WT), cardiac myosin binding protein C (cMyBPC)+/-; and cMyBPC- /- mice. Peak systolic torsion amplitude (C), peak rates of systolic torsion (D), and diastolic torsion (E) in WT, cMyBPC+/-; and cMyBPC-/- mice. Data are mean ± SE. *Significantly different from WT by 1-way ANOVA with Bonferroni (A and B) or Tukey-Kramer (C through E).

Myofilament Function in cMyBPC Mutant Myocardium

Levels of full-length cMyBPC have been shown to be decreased (24–33%) in patients with mutations in cMyBPC (van Dijk et al., 2009, Marston et al.,

2009). Similarly, levels of cMyBPC were decreased in heterozygous mouse models expressing 1 functional cMyBPC allele (the present study and Carrier et al., 2004).

Mutations in cMyBPC predicted to produce C-terminal truncated proteins are not 69

found in patients (Rottbauer et al., 1997, Moolman et al., 2000, van Dijk et al., 2009,

Marston et al., 2009, Jacques et al., 2008). The mutant mRNA and/or proteins are

thought to be unstable and are degraded before incorporation in the sarcomere

(Schlossarek et al., 2011), such that the mutant allele effectively acts as a null allele,

leading to cMyBPC haploinsufficiency.

Consistent with previous studies, we found that skinned myocardium isolated

from cMyBPC+/− (Harris et al., 2002) or cMyBPC−/− hearts (Stelzer et al., 2007) does

not display changes in Ca2+-sensitivity of force generation. This suggests that cMyBPC

content in the sarcomere does not directly determine myofilament Ca2+ sensitivity. A

recent study (van Dijk et al., 2009) showed that skinned myocytes isolated from

myectomy samples obtained from patients with HCM-causing cMyBPC mutations

displayed increases in the Ca2+ sensitivity of force generation. However, changes in

force generation did not correlate with cMyBPC expression and were likely a result of

the large decrease in troponin I (TnI) phosphorylation (84%) in myectomy samples

compared with control donor samples.

It is well established that skinned myocardium isolated from cMyBPC−/− hearts

displays significantly accelerated rates of cross-bridge kinetics (Stelzer et al., 2007,

Stelzer et al., 2006, Palmer et al., 2004) because myosin heads are in closer

juxtaposition to actin molecules, thereby enhancing the probability of acto-myosin

interactions (Colson et al., 2007). In the present study, we found that a relatively small

decrease in cMyBPC expression (24%) in cMyBPC± skinned myocardium produced

acceleration in the rate of delayed cross-bridge recruitment (kdf) after rapid stretch at

submaximal Ca2+ activations compared with WT skinned myocardium (Table 2.2). 70

Decreased cMyBPC content may result in nonuniform incorporation of cMyBPC into

the sarcomere and regional inhomogeneities in rates of force development. Thus,

sarcomeres with reduced levels of cMyBPC could develop force at a faster rate

compared with sarcomeres with normal cMyBPC content (Theis et al., 2009). This

would result in simultaneous shortening and stretch of different regions of the

sarcomere, which would disrupt the timing and synchronization of fiber shortening

during systole. Additionally, there may be a delay in force relaxation of regions that

are being actively stretched (stretch-activated) (Stelzer et al., 2007) by regions of the

sarcomere that are shortening, thereby prolonging relaxation in vivo (Harris et al.,

2002, Palmer et al., 2004, Nagayama et al., 2007).

Effects of Decreased cMyBPC Expression on In Vivo Mechanical Function

Studies of in vivo cardiac contractile function in cMyBPC−/− mice (Harris et

al., 2002, Palmer et al., 2004, Carrier et al., 2004, McConnell et al., 2001) have

demonstrated global systolic and diastolic dysfunction and severe LVH. However, in

vivo cardiac function and morphology of cMyBPC± hearts has been shown to be

normal at a young age (Harris et al., 2002, Carrier et al., 2004, McConnell et al.,

2001), with unexplained asymmetrical hypertrophy developing at 10–11 months of age

despite preserved systolic and diastolic function (Carrier et al., 2004). Cardiac MRI

can detect subtle changes in LV morphology and mechanical function, so we

examined LV motion indices of torsion and strain throughout the cardiac cycle in

cMyBPC−/− and cMyBPC± hearts. In this study, we used DENSE MRI to quantify

myocardial mechanics in the mouse heart, a technique that has been validated against

rotating phantoms or traditional MRI tagging protocols in mice and humans (Kim et 71

al., 2004, Gilson et al., 2004). We show that the lack of cMyBPC nearly abolished LV

torsion and twist in young mice. Furthermore, the magnitude of LV torsion and strain

are reduced and the pattern of LV mechanical function is altered in cMyBPC± hearts

in the absence of overt hypertrophy. The latter may be the early mechanical

manifestation of cMyBPC haploinsufficiency, which could contribute to the

progression and development of LVH with advanced age.

The counterclockwise rotation of the apex and the clockwise rotation of the

base (Streeter et al., 1969) produces the wringing motion that maximizes pumping of

blood into the circulation. We observed that the magnitude of LV torsion (Figure 2.10)

and net twist (Figure 2.9) were severely impaired in cMyBPC−/− hearts such that LV

systolic rotation was minimal. These results are consistent with overt hypertrophy and

fibrosis (Harris et al., 2002), which renders the cMyBPC−/− LV extremely stiff.

Additionally, myofiber architecture abnormalities disrupt the transmural progression

of the helical arrangement of subendocardial and subepicardial fibers (Wang et al.,

2010). This disruption may decrease the generation of mechanical torque by

subepicardial fibers, which propels the rotation of the LV during systole (Taber et al.,

1996). Interestingly, despite a significantly reduced magnitude of LV torsion in

cMyBPC−/− hearts, peak systolic torsion occurred earlier in systole compared with WT

hearts. The resulting shortening of the ejection phase in cMyBPC−/− hearts is consistent

with findings that cMyBPC is crucial for modulating the rate of pressure development

during isovolumic contraction and the period of systolic ejection (Nagayama et al.,

2007). 72

In contrast to severe mechanical dysfunction and hypertrophy in

cMyBPC−/−hearts, a modest 23% decrease in cMyBPC content in cMyBPC± hearts

resulted in a small but significant reduction in the magnitude of peak systolic torsion

(Figure 2.10). The decrease in the magnitude and duration of systolic torsion in

cMyBPC± hearts was due to a reduction in the overall net twist angle between the apex

and base (Figure 2.9). Specifically, there was a more prominent reduction in twist

between the mid-LV and the basal LV segments (Figure 2.9), although this difference

did not reach statistical significance (P=0.08). Because systolic rotation and fiber

shortening in the LV proceeds in an apex-to-base sequence (Sengupta et al., 2008), a

reduction in net twist could be due to global or regional abnormalities in

circumferential strain. Thus, the small reduction in circumferential strain at the mid-

LV in cMyBPC± hearts could impair the progression of mechanical rotation from the

apex to the base during systole resulting in diminished twist between the mid-LV and

basal-LV. These data are also consistent with the idea that cMyBPC is important for

prolongation of fiber shortening during late systole (Nagayama et al., 2007), perhaps

through continued rotation of the later-activated LV base. Accelerated rates of cross-

bridge kinetics and fiber shortening in the early isovolumic contraction phase

(Nagayama et al., 2007, Stelzer et al., 2007) in cMyBPC± hearts may disrupt the timing

of fiber shortening in later-activated regions such as the base, thereby reducing LV

rotation in late systole.

Slower systolic and diastolic circumferential strain rates (Germans et al., 2010, Ennis

et al., 2003) are a common feature in HCM carriers. In the present study,

cMyBPC−/− hearts displayed significant reductions in radial and circumferential strain 73

rates during systole and diastole compared with WT hearts (Figure 2.8), and

cMyBPC± hearts displayed lower early systolic and peak diastolic circumferential

strain rates at the mid-LV, and lower peak radial systolic strain rates in the base and

mid-LV (Figure 2.8). Diastolic dysfunction is thought to be an early manifestation of

impaired cardiac function in preclinical HCM (Ho et al., 2009). The early phase of LV

reverse rotation after systole is an important determinant of diastolic filling (Doucende

et al., 2010, Carasso et al., 2010), and decreases in diastolic mid-LV circumferential

strain rates in cMyBPC± hearts may result in a slowing of the rate of LV relaxation

and decreased diastolic filling, as has been noted previously in cMyBPC−/− hearts

(Harris et al., 2002, Palmer et al., 2004).

Potential Functional Consequences of cMyBPC Haploinsufficiency

Our data show that small reductions in cMyBPC content in the mouse heart,

similar to decreases reported in myocardial samples obtained from patients expressing

cMyBPC mutations, can significantly accelerate rates of cross-bridge kinetics and alter

myocardial mechanical function in vivo. Nonuniform incorporation of cMyBPC in

sarcomeres may result in regional inhomogeneities in rates of force development

within sarcomeres along with uncoordinated sarcomere shortening and stretching.

This can result in abnormal patterns of LV mechanical torsion in vivo that decrease

the efficiency of contraction and relaxation and thereby, chamber ejection and filling.

Furthermore, both increased ATP turnover due to accelerated cross-bridge cycling and

reduced LV mechanical efficiency would be expected to increase energy expenditure

by myocytes (Crilley et al., 2003, Timmer et al., 2010), which promotes apoptotic

pathways (Eijssen et al., 2008) that lead to decreased myocyte survival and 74

replacement of myocytes with fibrotic tissue (Morita et al., 2010). The induction of LV

fibrosis progressively stiffens the myocardium and impairs active fiber shortening and

stretch. Fibrotic myocardium has a reduced capacity to generate sufficient strain and

torsion to meet circulatory demands, which leads to compensatory LV hypertrophy.

Clinical Implications

It is possible that phenotypic unpredictability in contractile function and LVH,

as well as the age at which disease symptoms are apparent in patients expressing

cMyBPC mutations, is related to variability in overall levels of cMyBPC expression

(Morita et al., 2010). In addition, the regional distribution of cMyBPC expression in

the myocardium, such that patients with more significant cMyBPC loss present with

more severe dysfunction and LVH at an earlier age. Although mutations in the gene

encoding cMyBPC have typically been associated with late-onset cardiac dysfunction

and LVH, there is growing evidence that clinical symptoms of the disease are apparent

in a significant number of cMyBPC mutation carriers at a young age, including infancy

(Kaski et al., 2009, Rodriguez-Garcia et al., 2010). It is noteworthy that decreased

levels of cMyBPC have been reported in myocardial samples from young HCM

patients (20–30 years of age) (van Dijk et al., 2009, Marston et al., 2009) carrying

heterozygous cMyBPC mutations, which parallel our findings in young

cMyBPC± mice. Collectively, these data imply that clinical symptoms of cardiac

dysfunction related to mutations in cMyBPC, and perhaps mechanical dysfunction

specifically, manifest earlier than previously thought and should be screened for in

young relatives of HCM patients (Kaski et al., 2009). With the emergence of improved

cardiac MRI technology (Maron, 2009), it may now be possible to identify subtle 75

changes in mechanical function and hypertrophy in cMyBPC carriers at an earlier age,

thus significantly reducing risks for development of cardiac disease later in life.

Limitations

The mechanical manifestation and degree of cardiac hypertrophy in human

HCM is often diverse due to the underlying cause of the disease, which often involves

multiple contributing factors and may or may not involve mutations in sarcomeric

genes. The mechanical data presented here were collected from a homogeneous

population of mice in the absence of environmental modifiers that could impact

disease progression in humans. Therefore, the animal models of cardiac disease used

in this study may not fully reproduce all aspects of human disease. Furthermore, the

mice used in this study were deficient in cMyBPC, and thus our data may not be

representative of a general population of HCM in which cardiac dysfunction and

hypertrophy is not related to mutations in cMyBPC. For example, echocardiography

studies in a general cohort of HCM patients of unknown genotype revealed slightly

increased systolic circumferential strain compared with controls (Carasso et al., 2008),

whereas we observed lower circumferential strains in cMyBPC-deficient mice.

However, consistent with our data, lower systolic circumferential strains have been

shown in HCM patients with cMyBPC mutations (Germans et al., 2010). Thus, it can

be speculated that divergent cardiac mechanical function in different HCM

populations may reflect the underlying cause of the disease.

Clinical Perspective 76

Hypertrophic cardiomyopathy (HCM) is a disease caused by mutations in

genes that encode sarcomeric proteins. Mutations in cardiac myosin binding protein

C (cMyBPC) are among the most common causes genetically explained HCM, accounting for more than 40% of the total known cases worldwide. A large number of disease-causing mutations in cMyBPC in humans are predicted to produce C-terminal truncated proteins that are not incorporated into the sarcomere, resulting in cMyBPC haploinsufficiency, which reduces the total amount of cMyBPC in the sarcomere.

However, the link between decreased cMyBPC expression in the sarcomere and the development of cardiac disease is unclear. Our data demonstrate that a mouse model of cMyBPC haploinsufficiency with only 1 functional cMyBPC allele (cMyBPC+/−)

displays a 24% reduction in total cMyBPC expression and accelerated cross-bridge

kinetics, which underlies in vivo impairments in systolic and diastolic cardiac

mechanics measured by MRI. Importantly, reduced mechanical indices of torsion,

twist, and strain were apparent in young cMyBPC+/− mice 8–10 weeks of age,

suggesting that cardiac dysfunction in cMyBPC-related HCM is not necessarily late-

onset, as is commonly accepted, but may be detected early in life using high-resolution

cardiac MRI. Furthermore, in contrast to the severe mechanical dysfunction and cardiac hypertrophy displayed by cMyBPC-null mice, mechanical dysfunction in cMyBPC+/− mice was apparent before the development of overt cardiac hypertrophy, implicating altered mechanical function as the first link between reduced cMyBPC

expression in the sarcomere and the development of pathological hypertrophy and

cardiac disease.

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CHAPTER 3: COMPARISON OF VELOCITY VECTOR IMAGING

ECHOCARDIOGRAPHY WITH MAGNETIC RESONANCE IMAGING IN

MOUSE MODELS OF CARDIOMYOPATHY

ABSTRACT

Background—Myocardial strain imaging using echocardiography can be a cost-

effective method to quantify ventricular wall motion objectively, but few studies have

compared strain measured with echocardiography against magnetic resonance

imaging (MRI) in small animals.

Methods and Results—We compared circumferential strain (CS) and radial strain

(RS) measured with echocardiography (velocity vector imaging [VVI]) to

displacement encoding with stimulated-echo MRI in 2 mouse models of

cardiomyopathy. In 3-month-old mice with gene targeted deficiency of cardiac

myosin-binding protein-C (cMyBPC−/−, n=6) or muscle LIM protein (MLP−/−, n=6),

and wild-type mice (n=8), myocardial strains were measured at 3 cross-sectional levels

and averaged to obtain global strains. There was modest correlation between VVI and

MRI measured strains, with global CS yielding stronger correlation compared with

global RS (CS R2=0.4452 versus RS R2=0.2794, both P<0.05). Overall, strain

measured by VVI was more variable than MRI (P<0.05) and the limits of agreement

were slightly, but not significantly (P=0.14), closer for global CS than RS. Both VVI

and MRI strain measurements showed significantly lower global CS strain in the

knockout groups compared with the wild type. The VVI (but not MRI) CS strain

measurements were different between the 2 knockout groups (−14.5±3.8% versus

−6.6±4.0%, cMyBPC−/− versus MLP−/− respectively, <0.05). 82

Conclusions—Measurements of left ventricular CS and RS are feasible in small

animals using 2-dimensional echocardiography. VVI and MRI strain measurements

correlated modestly and the agreement between the modalities tended to be greater for

CS than RS. Although VVI and MRI strains were able to differentiate between wild-

type and knockout mice, only global CS VVI differentiated between the 2 models of

cardiomyopathy.

INTRODUCTION

Two-dimensionally (2D)-directed echocardiography has arguably become the

leading method for assessing left ventricular (LV) function in small animals because it

is noninvasive, relatively cost-effective, widely available, and has short acquisition and

post-processing times that allow high through-put. These techniques have proved

useful in determining both the interplay between altered cardiovascular gene

expression and compensatory physiologic regulation, as well as the effects of genetic,

surgical, and pharmacological interventions in rodent models of disease (Popovic et

al., 2007, Bachner-Hinenzon et al., 2010, Hoit, 2006).

However, 2D-directed detection of LV dysfunction by conventional

echocardiography (M-mode, 2D, Doppler) is considered a late manifestation of

cardiac disease that lacks the sensitivity to identify subclinical disease (Marwick et al.,

2010, Borg et al., 2009, Bauer et al., 2011, Stanton et al., 2010). Doppler- and 2D

echocardiographic-derived myocardial strain (or deformational) imaging are recent

advances in noninvasive cardiac imaging intended, in part, to overcome the limitations

that confound the traditional echocardiographic assessment of LV function. 83

Abnormalities of myocardial deformation are seen early in the development of a

variety of pathophysiologic states, and thus provide a sensitive means for detecting

global and regional myocardial dysfunction (Hoit, 2011). Compared with Doppler-

determined strain, 2D echocardiographic (2D speckle-tracking echocardiography

[STE]) strain is independent of the angle of ultrasound propagation, measures the 3

normal Lagrangian strains (radial strain [RS], circumferential strain [CS], and

longitudinal strain), and is less noisy, and more reproducible. 2D STE has been

previously described in humans for use in myocardial layer-specific deformation

analysis (Adamu et al., 2009), and more recently has been used for phenotyping mice

(Bauer et al., 2011). This imaging technique tracks speckle patterns generated by

interference between the ultrasound beam and myocardium on 2D echocardiographic

images (Chetboul et al., 2010). It allows a non–Doppler-based assessment of regional

myocardial motion, and provides information about segmental as well as global LV

function with greater sensitivity and specificity compared with conventional

echocardiography (Bauer et al., 2011, Cottrell et al., 2010, Rappaport et al.,

2006). Velocity Vector Imaging (VVI; Siemens Medical Solutions, Mountain View,

CA) is a novel imaging technique based on 2D STE, which incorporates speckle-

tracking and endocardial contour tracking that allows angle-independent

measurements of strain (Hoit, 2011, Bansal et al., 2008).

With its high spatial resolution and superb imaging quality, magnetic resonance

imaging (MRI)-based strain imaging is considered the accepted standard method for

measuring myocardial wall strain (Chetboul et al., 2010, Liu et al., 2006). However, it

is costly, time-consuming, and not widely available, precluding it from routine use in 84

the small animal research arena. Although other studies have compared 2D STE strain

measurements with MRI (Li et al., 2007), there are no studies to date that have directly

compared VVI strain measurements with MRI-based strain in small animal models of

cardiovascular disease. In this study, we report LV strain measurements in normal

mice and in mouse models of hypertrophic and dilated cardiomyopathy using VVI and

displacement encoding with stimulated-echo MRI, thus providing a direct comparison

of the strain measurements in pathological states.

METHODS

Experimental Animals

Adult male (8–10 weeks of age) mice with deficiency of either myosin-binding

protein-C, a thick filament-associated protein of the sarcomere (cMyBPC−/−; n=6), or

muscle LIM protein, a promoter of myogenic cytoskeleton differentiation (MLP−/−;

n=6), and their wild-type littermates of the SV/129 strain (wild-type [WT]; n=8) were

used in this study. These groups served as models of hypertrophic and dilated

cardiomyopathy, and controls, respectively (Arber et al., 1997, Harris et al., 2002).

Mice were placed on a standard mouse chow diet and water ad libitum, and

housed in a temperature-controlled environment with an alternating 12-hour

light/dark cycle. All procedures involving animal care and handling were performed

according to institutional guidelines set forth by the Animal Care and Use Committee

at Case Western Reserve University.

Magnetic Resonance Imaging 85

Animals were anesthetized with 2% isoflurane with supplemented O2 in an

isoflurane induction chamber, and then moved into the magnet and kept under

inhalation anesthesia with 1.5% isoflurane. With electrocardiogram and respiratory

gating, cardiac functional MRI studies were performed with a 9.4-T Bruker Biospec

(Billerica, MA) horizontal scanner using a volume coil. A series of scout images were

first acquired to obtain the horizontal long-axis image from which 3 LV short-axis

planes at basal, mid-ventricular, and apical levels were prescribed as perpendicular to

the LV long-axis with an interslice distance of 1.5 mm. Two-dimensional myocardial

motion was quantified using displacement encoding with stimulated-echo MRI, the

details of which has been described previously (Zhong et al., 2010). Data were

captured at a rate of 13 frames/cardiac cycle, resulting in a temporal resolution of ≈9

ms. Image processing (MRI reconstruction) and data analysis were performed offline

using custom-built software written in Matlab (MathWorks, Natick, MA). Data

acquisition and analysis required ≈3 to 3.5 hours and 1 per animal.

The epicardial and endocardial LV borders were traced using cine images to

calculate LV ejection fraction. A 2D displacement map was calculated by means of

vector addition of the displacement from 2 orthogonal directions to compute

Lagrangian strain tensors at the base, mid, and apex of the LV. The CS and RS

measurements at the base, mid, and apex of the LV were averaged to obtain global CS

and RS for each animal (Figure 3.1A).

Echocardiography

Echocardiographic studies were performed within 5 days of the MRI. Animals

were anesthetized with 2% isoflurane supplemented with O2 in an isoflurane induction 86

chamber and maintained with 1.5% isoflurane by nose cone. Echocardiography was

performed as previously described (Morgan et al., 2004). Acoustic capture B-mode

cine clips (120 Hz) were obtained with electrocardiographic gating using a Sequoia

ACUSON System (Siemens Medical Solutions, Mountain View, CA) with a 15-MHz

linear array transducer. Image processing and data analysis were performed offline

using Syngo Vector Imaging technology software (Siemens Medical Solutions,

Mountain View, CA).

2D-directed M-mode images from the mid-papillary short-axis were used to

calculate conventional measurements of the LV, which included the LV end-diastolic

diameter, end-systolic diameter, anterior and posterior wall thicknesses, and fractional

shortening. 87

B-mode clips were selected based on adequate visualization of the endocardial

border and the absence of image artifacts. The epicardial and endocardial LV borders

were manually traced and accurate tracking verified >3 cycles in the parasternal short-

Figure 3.1. Displacement encoding with stimulated-echo magnetic resonance imaging (A) and velocity vector imaging echocardiographic (B) vector maps for regional strain in wild type (left) vs knockout (KO) (right). The KO group has decreased strain compared with the wild type, demonstrated by smaller size of the vectors.

axis view at end-systole to calculate Lagrangian strain at the base, mid, and apex of

the LV. Peak CS and RS values for each segment of the LV were recorded and

averaged to obtain global strains for each animal (Figure 3.1B).

Data acquisition (3 short axes) and analysis (CS and RS) required ≈15 minutes per

animal.

To ensure good quality images for STE-based strain analyses, image acquisition

was performed at a high frame rate by using the smallest possible depth and sector 88

size. All image acquisitions and offline measurements were conducted by a single

investigator (SA) who was blinded to animal groups. Using a second investigator

(BDH), interobserver differences of peak regional systolic RS and CS were determined

from 20 randomly selected clips as 100× the difference between 2 observations divided

by the mean of the 2 observations. Intraobserver differences were determined similarly

from 20 randomly selected clips measured 10 days apart by a single investigator (AL).

Intra- and interreader coefficients of variation were estimated as the root mean square

of the coefficients of variations and intra- and interreader intraclass coefficients were

calculated using the method of Fleiss (Fleiss et al., 1996).

Statistical Analysis

The statistical analysis was performed using SigmaPlot 12.0 (Systat Software, Inc.,

San Jose, CA) and SAS (SAS Institute, Cary, NC) version 9.2 commercial software.

All values are expressed as mean±SD. Nondeformation echocardiographic and MRI

variables were compared with 1-way ANOVA with post hoc Tukey tests. Strain data

were analyzed using linear mixed models, fitting a separate model for the RS and CS

at 3 locations (base, mid, and apex) and a model each for global RS and CS. Thus, 8

models were fit; for each, Tukey multiple comparison procedure with a family-wise

error rate of 0.05 was performed for pairwise comparisons of the 6 groups defined by

the 3 genotypes, and for comparisons between VVI and MRI within a genotype. Bland-

Altman plots were constructed to illustrate the agreement between VVI and MRI strain

measurements. The Morgan-Pitman test (Wilcox, 1990) was used to test the

hypotheses that variability was greater for VVI than MRI, and variability was greater

for RS than CS. A P value (2-sided) <0.05 was considered as statistically significant. 89

RESULTS

Echocardiographic and MRI Volumetric Variables

The LV ejection fraction measured by MRI for the cMyBPC−/− and

MLP−/− groups (33±8% and 29±9%, respectively) was significantly lower than the WT

group (69±9%; P <0.05), and was similar between the 2 groups of knockout mice.

Similarly, the fractional shortening measured by 2D-directed M-mode

echocardiography was significantly lower for the cMyBPC−/− and MLP−/− groups

(38±6% and 19±5%, respectively) compared with the WT group (61±5%), but was

significantly lower in the MLP−/− than cMyBPC−/− group (Table 3.1). The LV EDD

and LV ESD were greater in the MLP−/− group and wall thicknesses were greater in

the cMyBPC−/−group. There were no differences in heart rate among the 3 groups of

mice.

Global Strain

The Bland-Altman limits were narrower for global CS compared with global

RS (Figure 3.2A and 3.2B; however, the difference was not statistically 90

significant P=0.14).

Additionally, there was

modest correlation

between VVI and MRI

measured strains, with

global CS yielding a

stronger correlation

compared with global RS

(CS R2=0.4452 versus RS

R2=0.2794, both P<0.05).

The global RS measured

by MRI was greater in the

wild-type (24.7±1.0%) than knockout mice (cMyBP-C−/−: 12.7±2.0%, MLP−/−:

14.8±3.2%). In contrast, the global RS measured by VVI in the WT mice (23.5±9.2%)

was statistically similar to that in the cMyBP-C−/−(16.4±4.6%), but significantly

greater than in the MLP−/− mice (6.8±4.0%). The global CS determined with either

VVI or MRI was

significantly lower in the

knockout than WT mice.

However, only VVI

showed significantly lower

CS strain in the 91

MLP−/− mice (−6.6±4.0%) than the cMyBP-C−/− mice (−14.5±3.8%) (Tables 3.1and 2).

In comparing the VVI and MRI, only global CS strains in the WT mice were

statistically different, with lower strains measured with MRI compared with VVI.

Variability of global CS and RS strains was greater for VVI than MRI (both P<0.05),

although variability was similar for global CS and RS (P=0.39).

Regional Strain

Values of regional RS

comparing VVI and

MRI were similar in

both the WT and

cMyBP-C−/− mice, and

only at the LV base of

MLP−/− mice were

values significantly

different when

comparing VVI and

MRI (Figure 3.3).

Regional strains

measured with MRI in

Figure 3.2. Bland-Altman plots comparing the magnetic both knockouts were resonance imaging (MRI)- and echo-derived strain. A, Circumferential strain measurements. B, Radial strain lower than those in the measurements. There is closer agreement between the MRI and echo imaging modalities for circumferential strain when compared with radial strain measurements. Upper and lower dotted lines are WT mice, whereas, the 95% limits of agreement. only regional RS 92

Figure 3.3. Regional radial strain: *P<0.05 echo vs magnetic resonance imaging (MRI), †P<0.05 wild type vs knockout (KO). ‡P<0.05 MyBPC-/- vs MLP-/- (echo only). Regional radial strain measurements using MRI and echo in the wild-type group are similar. Regional radial strain measurements by MRI in the KO groups are significantly lower compared with the wild-type group but more variable via echo. cMyBPC-/- indicates deficiency of cardiac myosin-binding protein-C; MLP-/-, deficiency of muscle LIM protein; and WT, wild-type.

measured with VVI were significantly lower in the MLP−/− than those in the WT mice.

When comparing between the 2 knockout groups, strains measured by VVI were

significantly lower in the MLP−/− than cMyBP-C−/−group at the base of the LV;

regional RS measured by MRI was similar between the 2 knockout groups.

Values of regional CS comparing VVI and MRI were similar in both the WT and

MLP−/− mice, and only at the mid LV base of cMyBP-C−/− mice were values

significantly different when comparing VVI and MRI. Regional CS measured by MRI

was lower in the cMyBP-C−/− knockout group compared with the WT group at the

mid and apex levels, and the MLP−/− knockouts at the mid LV. The regional CS

measured by VVI was also lower for both knockout groups compared with WT except

for regional strain at the mid LV in cMyBP-C−/− mice (Figure 3.4). Similar to regional 93

RS, VVI regional CS measurements (in this instance, mid LV) was significantly lower

in the MLP−/− group compared with cMyBP-C−/− group.

Figure 3.4. Regional circumferential strain: *P<0.05 echo vs magnetic resonance imaging (MRI), †P<0.05 wild type vs knockout (KO). ‡P<0.05 MyBPC-/- vs MLP-/- (echo only). Regional circumferential strain measurements using MRI and echo in the wild-type group are similar. Regional circumferential strain measurements by MRI in the KO groups are significantly lower compared with the wild-type group but more variable via echo. cMyBPC-/- indicates deficiency of cardiac myosin-binding protein-C; MLP-/- , deficiency of muscle LIM protein; and WT, wild-type.

Interpretative Variability

The interobserver differences for CS and RS were −3.5±14.7 and −3.9±21.9,

respectively. Intraobserver differences for CS and RS were 4.7±9.2% and 4.7±13.0%,

respectively. The intra- and interreader coefficients of variations were 9.6% and 15.3%

for RS and 7.2% and 10.4% for CS, respectively. The intra- and interreader intraclass

coefficients were 0.986 and 0.955 for RS and 0.990 and 0.932 for CS, respectively.

DISCUSSION

This is the first study comparing echocardiographic strain imaging using VVI and

MRI in the wild-type and genetically altered mouse. The principal results of this study

are (1) correlations between MRI and VVI strains are modest and are greater (and 94

agreement tends to be greater) for CS than RS; (2) strains measured with VVI are more

variable than those measured with MRI and the variability is similar for RS and CS;

and (3) VVI-measured strains can be used to rapidly phenotype mouse models of

cardiomyopathy. Although MRI is currently considered the accepted standard for

noninvasive myocardial strain imaging, in part because of the ability to evaluate strain

in 3 dimensions, long acquisition and postprocessing times, expense, and limited

availability preclude its routine use in the small animal laboratory. Thus,

echocardiographic strain is a potentially advantageous method for the objective

assessment of global and regional LV function and for high through-put phenotyping

of murine models (Bauer et al., 2011).

Bauer et al (Bauer et al., 2011) previously reported speckle-tracking strain

measurements in WT mice, which were higher than values obtained in our study. The

previous study used speckle-tracking algorithm supplied by VisualSonics (VevoStrain,

VisualSonics, Toronto Canada), whereas echocardiographic strain measurements in

our study were obtained by VVI algorithm supplied by Siemens Medical Solutions

(Syngo Vector Imaging technology software, Siemens Medical Solutions, Mountain

View CA) (Bauer et al., 2011). The proprietary speckle-tracking software from the 2

companies and the superior sampling rates in the VisualSonics software may account

for the difference in the strain measurements. Importantly, in this study, the VVI

strains are similar to those obtained with MRI and the latter measured in the WT mice

were similar to MRI strains previously reported (Liu et al., 2006).

Bansal et al (Bansal et al., 2008) previously reported validation of VVI strain with

harmonic phase MRI in humans with a very modest correlation between the 2 95

modalities, greater with CS than RS (CS R2=0.12 versus RS R2=0.005). Similarly, in

our study, the correlation between VVI and MRI strain was higher with CS than RS

(CS R2=0.4452 versus RS R2=0.2794), but both CS and RS correlations in our study

were much greater than in that validation study involving human subjects and were

similar to both Bansal et al (Bansal et al., 2008) (CS R2=0.397 and RS R2=0.348,

both P<0.05) and Cho et al22(CS R2=0.26 and RS R2=0.36) using a different speckle-

tracking algorithm, automated functional imaging (GE Medical Systems, Milwaukee

WI). Nevertheless, our correlations are disappointingly less robust than those reported

in a validation study in 5 mice after myocardial infarction and 2 control mice using

the VisualSonics instrument with EKV (CS R2=0.81 and RS R2=0.72), which acquires

data from multiple cardiac cycles and constructs an image sequence composed of >100

images per cardiac cycle (Li et al., 2007).

LV contraction is a complex process involving deformation resulting in shortening

in 3 normal directions; longitudinal, CS, and RS. Longitudinal strain, which is a

sensitive indicator of subendocardial fiber dysfunction and an early marker of ischemia

and increased wall stress, was not evaluated in this study as we were comparing

directly with MRI as it is measured in small animals in our institution. In addition,

advantages of strain imaging compared with conventional echocardiographic

parameters could not be demonstrated in this study, which was designed to compare

strain imaging with MRI in fully developed models of cardiomyopathy. In a previous

study in mice, longitudinal strains were sensitive to changes early after experimental

myocardial infarction and were able to predict LV remodeling; CS and RS were not

as consistent.6 Longitudinal strain is typically obtained from an apical view (although 96

the parasternal long-axis view was used by Bauer et al (Bauer et al., 2011)), which is

difficult to obtain reliably in small animals. CS and RS are more influenced by

transmural fiber dysfunction (especially the mid-myocardium), and are generally more

suited for identifying dysfunction in ventricles with reduced LV systolic function

(Geyer et al., 2010).

Both MRI and VVI strain measurements were able to differentiate between the WT

group and the genetic models of cardiomyopathy, reinforcing the potential usefulness

of strain measurements in mouse models of LV dysfunction. However, only VVI CS

strain differentiated between the 2 models of cardiomyopathy, which corresponded

with differences in their echocardiographic LV fractional shortening.

One potential difficulty is that VVI had greater variability in strain measurement

compared with MRI, and was particularly problematic in measurement of RS. This

may be because of the need to track both epicardial and endocardial borders, the

former more difficult to identify with a resultant decrease in accuracy. The greater

variability has been reported in previous studies (Bansal et al., 2008). Although MRI

RSs are generally more variable than CS, in this study, their variability was similar.

Limitations

Several limitations of this study merit consideration. First, for logistic reasons,

echocardiography and MRI were not performed on the same day for all the animals.

This may be responsible, in part, for the modest correlations between the 2 techniques.

Second, the echocardiographic images were obtained using 2D rather than 3D imaging

used in MRI; translation of the heart remains a problem using 2D acquisition methods 97

as error is introduced to strain measurements when the heart swings out of the imaging

plane. In addition, out-of-plane motion occurs because of rotation and motion of the

heart; as a result, only a portion of the real motion can be detected. Third, potential

sources of variation include the quality of the images obtained for strain analysis. High

quality of the 2D images at high frame rates is essential for accurate VVI strain

measurements. Moreover, poor quality of images may hinder the investigator’s ability

to perform tracing of the endocardium and epicardium which may affect the strain

values, thereby introducing another source of variability. Fourth, while temporal

resolution for the echocardiograms (120 Hz) was similar (8.3 ms) to the MRI, the

accuracy of tracking speckles may theoretically be jeopardized at these frame rates and

may have resulted in a significant underestimation of strain values; higher frame rates

were reported using different instrumentation and algorithms (Bauer et al., 2011, Li et

al., 2007). Finally, changes in the imaging angle of incidence can result in capturing

different fiber layers at different levels and may introduce variability, particularly

because we analyzed exclusively short views which are sensitive to small differences

in image angle (Bauer et al., 2011).

Conclusion

Despite these limitations, we demonstrate that measuring LV CS and RS is

feasible in small animals using 2D echocardiography with VVI. VVI and MRI strain

measurements correlated modestly and the correlation was greater (and agreement

tended to be greater) for CS than RS. VVI had greater variability in strain measurement

compared with MRI. Although global VVI and MRI strains were able to differentiate 98

between WT and knockout mice, only VVI strain differentiated between the 2 models

of cardiomyopathy.

Clinical Perspective

Although 2-dimensional echocardiographic strain imaging cost-effectively and

objectively quantifies ventricular wall motion, only 1 small study has directly

compared strain measured with echocardiography against magnetic resonance

imaging (MRI) in mice. Using a different speckle-tracking algorithm (velocity vector

imaging [VVI]), we compared circumferential (CS) and radial strain (RS) to

displacement encoding with stimulated-echo MRI in 2 genetic mouse models of

cardiomyopathy. CS and RS were measured in groups of wild-type mice and mice

with gene targeted deficiency of cardiac myosin-binding protein-C or muscle LIM

protein. There was modest correlation between VVI and MRI measured strains, with

global CS yielding a stronger correlation compared with global RS (CS R2=0.4452

versus RS R2=0.2794, both P<0.05). Overall, strain measured by VVI was more

variable than MRI and the limits of agreement were slightly, but not significantly,

closer for global CS than RS. Both VVI and MRI strain measurements showed

significantly lower global CS strain in the knockout groups compared with the wild-

type, but only the VVI CS strain measurements were different between the 2 knockout

groups. These data demonstrate that measurements of LV CS and RS are feasible in

mice using VVI, although correlations and agreement were modest. VVI may

complement conventional methods that objectively assess global and regional LV 99

function and be particularly useful for high through-put phenotyping of murine

models.

100

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102

CHAPTER 4: ARTERIAL SPIN LABELING-FAST IMAGING WITH

STEADY-STATE FREE PRECESSION (ASL-FISP): A RAPID AND

QUANTITATIVE PERFUSION TECHNIQUE FOR HIGH-FIELD MRI

ABSTRACT

Arterial spin labeling (ASL) is a valuable non-contrast perfusion MRI technique

with numerous clinical applications. Many previous ASL MRI studies have utilized

either echo-planar imaging (EPI) or true fast imaging with steady-state free precession

(true FISP) readouts, which are prone to off-resonance artifacts on high-field MRI scanners. We have developed a rapid ASL-FISP MRI acquisition for high-field

preclinical MRI scanners providing perfusion-weighted images with little or no

artifacts in less than 2 s. In this initial implementation, a flow-sensitive alternating

inversion recovery (FAIR) ASL preparation was combined with a rapid, centrically

encoded FISP readout. Validation studies on healthy C57/BL6 mice provided

consistent estimation of in vivo mouse brain perfusion at 7 and 9.4 T (249 ± 38 and

241 ± 17 mL/min/100 g, respectively). The utility of this method was further demonstrated in the detection of significant perfusion deficits in a C57/BL6 mouse model of ischemic stroke. Reasonable kidney perfusion estimates were also obtained for a healthy C57/BL6 mouse exhibiting differential perfusion in the renal cortex and medulla. Overall, the ASL-FISP technique provides a rapid and quantitative in vivo assessment of tissue perfusion for high-field MRI scanners with minimal image artifacts.

103

INTRODUCTION

Arterial spin labeling (ASL) MRI is a non-contrast perfusion MRI technique that

has been shown to provide quantitative assessments of tissue perfusion in multiple

clinical imaging applications, including brain (Wong et al., 2007, Brookes et al., 2007,

Boss et al., 2007), kidney (Karger et al., 2000, Martirosian et al., 2004, De Bazelaire

et a., Fenchel et al., 2006), lung (Mai et al., 1999, 2001, Wang et al., 2003, Bolar et al.,

2006, Martirosian et al., 2006) and liver (Katada et al., 2012). One major advantage of

ASL over conventional dynamic contrast-enhanced MRI perfusion techniques is the

lack of exogenous, and potentially toxic, paramagnetic contrast agents. The use of an endogenous blood signal to obtain tissue perfusion information is especially important for the imaging of patients with chronic kidney diseases, which can be a contraindication for gadolinium-based MRI perfusion methods (Kuo et al., 2007).

This attribute is also important for longitudinal preclinical imaging applications, as multiple tail-vein injections and/or catheterizations can cause local inflammation and necrosis, resulting in reduced access to veins.

ASL MRI techniques generate blood flow contrast between multiple images using a wide variety of blood labeling methods (Detre et al., 1992, Williams et al., 1992,

Edelman et al., 1994, Kwong et al., 1995, Kim et al., 1997, Detre et al., 1999). A typical ASL MRI acquisition combines an ASL preparation phase, followed by a rapid imaging readout to capture the blood flow-weighted contrast (Fig. 1). It is important to note that a large majority of the imaging developments have focused on the optimization of the preparation phase of the ASL acquisition. As a result, a wide variety of preclinical and clinical ASL MRI techniques have been developed. These 104

ASL techniques can be broadly grouped into continuous [CASL (Detre et al., 1992,

Williams et al., 1992, Detre et al., 1999) and pulsed [PASL (Edelman et al., 1994,

Kwong et al., 1995, Kim et al., 1997) categories. A hybrid of these two techniques, pseudo-continuous ASL (pCASL), has also been developed recently (Wong et al.,

2007, Wu et al., 2007, Duhamel et al., 2014). Each of these specialized techniques offers advantages for specific imaging applications. At the same time, many of these studies have utilized conventional imaging readouts, including fast low-angle shot

(FLASH) (Pell et al., 2004, Zuo et al., 2013), echo-planar imaging (EPI) (Wang et al.,

2004, Rajendran et al., 2013) and true fast imaging with steady-state free precession

(true FISP) (Martirosian et al., 2004, 2006, Esparza-Coss et al., 2010, Boss et al., 2006).

Unfortunately, these imaging readouts exhibit significant limitations for high-field

MRI applications. Specifically,B0 inhomogeneities result in increased

distortion/ghosting and banding artifacts from EPI and true FISP imaging techniques,

respectively, on high-field MRI scanners. These artifacts are particularly problematic

for body imaging applications, where cardiac and respiratory motion, as well as

adipose tissue, can make precise shimming difficult. In addition, the increase

in T1 magnetic relaxation times on high-field MRI scanners can result in spoiled gradient echo images with a lower signal-to-noise ratio (SNR) relative to these other imaging techniques. A lower SNR is especially problematic for ASL MRI techniques, as the differential blood flow signal is typically less than 10% of the mean tissue signal

(Petersen et al., 2006, Yongbi et al., 1999). Therefore, there is a need to develop a rapid

and robust ASL MRI imaging readout that is both sensitive to blood flow labeling from 105

the ASL preparation and also immune to B0 inhomogeneities and motion artifacts on

high-field MRI scanners.

Previously, our group has reported the development of a centrically encoded FISP

imaging technique which can be flexibly combined with conventional diffusion and

magnetization transfer (MT)/chemical exchange saturation transfer (CEST)

preparation methods to rapidly generate quantitative imaging data (Lu et al., 2012,

Shah et al., 2011). Importantly, the magnetic field gradients for the FISP readout

sequence are not completely refocused in either the slice select or frequency encoding

directions, resulting in greatly reduced off-resonance artifacts in comparison with EPI

and true FISP acquisitions. In addition, centric k-space encoding of the FISP readout

retains the image contrast generated in the diffusion and MT/CEST preparations.

Here, we describe our initial in vivo results from an ASL-FISP acquisition on 7- and

9.4-T Bruker Biospec (Bruker Inc., Billerica, MA, USA) small-animal MRI scanners.

For this initial study, our ASL-FISP implementation combines a flow-sensitive

alternating inversion recovery (FAIR) preparation scheme with our rapid centrically

encoded FISP imaging readout to generate ASL imaging data (Martisosian et al.,

2004, Kim et al., 1997, Boss et al., 2006, Gunther et al., 2001). The reproducibility of

this new ASL-FISP technique was evaluated in brains of healthy C57/BL6 mice. In

addition, we demonstrate the effectiveness of this technique for multiple imaging applications, including mouse models of ischemic stroke and healthy mouse kidneys.

METHODS 106

In vivo comparison of image artifacts at 7 T

Axial FISP images of a healthy C57/BL6 mouse brain were acquired on a 7-T

Bruker Biospec MRI scanner for comparison with conventional spin echo

(TR/TE = 2000/14 ms; one average; total scan time, 4 min 16 s), true FISP

(TR/TE = 4.0/2.0 ms; 20 averages; tip angle, 60°; total scan time, 11 s) and EPI

(TR/TE = 2500/30.8 ms; four segments; two averages; total scan time, 20 s) imaging readout methods to assess image artifacts.

ASL-FISP pulse sequence design

The ASL-FISP acquisition was implemented on Bruker Biospec 7- and 9.4-T

MRI scanners equipped with 400 mT/m gradient inserts. The ASL-FISP sequence was implemented on both MRI scanners to demonstrate the robustness of the methodology to artifacts on high-field MRI scanners. The ASL-FISP pulse sequence was developed by combining a FAIR preparation (slice-selective or non-selective inversion) with a centrically encoded FISP imaging readout to acquire all lines of k space following each

ASL preparation (Fig. 4.1). It should be noted that, although a FAIR scheme was

implemented herein, the ASL-FISP technique is adaptable to a variety of ASL

preparations. A hermite excitation radiofrequency (RF) pulse with a duration of 0.5 ms

and a tip angle of 60° were selected for this implementation. The high tip angle was

selected for this FISP acquisition to provide increased SNR in comparison with low

tip angles (data not shown).

Uniformity of the inversion pulse is essential for accurate and precise quantification of

tissue perfusion. Therefore, a hyperbolic secant adiabatic inversion pulse with a

duration of 3.0 ms was used for the FAIR inversion preparation. Magnetic field 107

gradient spoilers were applied after the inversion preparation to limit transverse

magnetization prior to the FISP imaging readout. The FISP imaging readout

(including 10 dummy scans) provided in vivo images in less than 2 s with relatively high

Figure 4.1. Schematic diagram of arterial spin labeling-fast imaging with steady-state free precession (ASL-FISP) acquisition. A flow-sensitive alternating inversion recovery (FAIR) ASL preparation is combined with a centrically encoded FISP acquisition. This acquisition is repeated for both a slice- selective inversion (with shaded gradient) and a non-selective inversion (without shaded gradient) to generate the perfusion contrast. Note that all lines of k space are acquired following a single ASL preparation. ADC, analog-to-digital converter. SNR in comparison with conventional spoiled gradient echo acquisitions, and

minimal off-resonance distortion and ghosting artifacts in comparison with EPI (Lu et

al., 2012, Shah et al., 2011). The FISP imaging readout also prevented banding

artifacts typical of balanced steady-state free precession (SSFP)/true FISP acquisitions

on high-field MRI scanners. The FISP imaging readout was also designed with centric

encoding to retain blood flow sensitivity, as well as the T1 weighting generated from

the FAIR inversion preparation.

Initial in vivo ASL-FISP perfusion assessments in mouse brains

All animal studies were conducted in accordance with approved Institutional

Animal Care and Use Committee protocols at Case Western Reserve University. The

ASL-FISP technique was initially evaluated by assessing brain perfusion in healthy

wild-type C57/BL6 mice (The Jackson Laboratory, Bar Harbor, ME, USA). Each 108

animal was anesthetized with 1–2% isoflurane with supplementary O2 and positioned

within a mouse volume coil (inner diameter, 35 mm) in a Bruker Biospec MRI

scanner. Each animal's body temperature was maintained at 35 ± 1 °C with a warmed

air control system. Respiratory triggering was performed through an MR-compatible, small-animal gating and control system (SA Instruments, Stony Brook, NY, USA).

Ten male C57/BL6 mice, aged 10 weeks, were scanned with the ASL-FISP imaging protocol at 7 T. Five of these mice were scanned with the ASL-FISP imaging protocol at 9.4 T at least 1 day prior to the 7-T scan for comparison. Rapid localizer scans were first used to identify an appropriate and consistent axial mid-brain imaging slice for the ASL-FISP protocol. After slice positioning, the single-slice ASL-FISP protocol consisted of three sequential scans: (1) ASL-FISP with a slice-selective inversion; (2)

ASL-FISP with non-selective inversion; and (3) FISP with no inversion preparation as a reference (M0) scan for blood flow calculation (Kim et al., 1997). An inversion delay time (TI) of 1420 ms was used for this initial implementation to generate sufficient blood flow contrast between the slice-selective and non-selective ASL-FISP images.

Other than the inversion preparation, the FISP imaging readout parameters were identical for these three acquisitions (centric encoding; TR/TE = 2.4/1.2 ms; matrix,

128 × 128; field of view, 3 cm × 3 cm; imaging slice thickness, 1.5 mm; tip angle, 60°).

These scans were repeated 60 times to determine the effects of image quality on the perfusion calculations. Although not required, for this initial implementation, an additional delay time of ~13 s was incorporated between each scan repetition to allow the magnetization to return to equilibrium (M0) between repetitions and to limit the

duty cycle on the magnetic field gradients. The total acquisition time for one ASL- 109

FISP repetition, including the ASL preparation (1420 ms), FISP imaging readout

(331 ms) and ~13-s delay, was 15 s.

For comparison, we also implemented an ASL-GRE acquisition by combining

an identical FAIR preparation with a gradient echo (GRE) imaging readout

(TR/TE = 5000/2 ms; five averages; tip angle, 90°). For this simplified ASL-GRE

acquisition, only one line of k space was acquired for each ASL preparation. All other

imaging parameters, including the field of view and resolution, were identical to the

ASL-FISP acquisitions described above. This conventional ASL-GRE acquisition was

evaluated on a separate single healthy C57/BL6 mouse brain at 7 T for direct comparison with the ASL-FISP results.

Following the ASL-FISP scans, a FISP-based Look–Locker acquisition was implemented to generate voxel-wise T1 maps according to previously described

techniques (Deichmann & Haase, 1992, Jakob et al., 2001). The Look–Locker

acquisition consisted of a non-selective adiabatic inversion, followed by 10 continuous

and sequential FISP image repetitions (linear encoding; tip angle, 10°;

TR/TE = 4.0/2.0 ms; 512 ms/image). The Look–Locker acquisition was repeated at

least 40 times to ensure accurate T1 relaxation time estimation. As for the ASL-FISP

acquisitions above, an ~7-s delay was introduced after each repetition to reduce the

duty cycle on the imaging gradients and to allow the magnetization to return to

equilibrium (M0). Voxel-wise T1 relaxation maps were then calculated according to

previously described methods (Deichmann & Haase, 1992, Jakob et al., 2001). The

geometry of the Look–Locker acquisition was identical to that of the ASL-FISP 110

acquisitions to ensure one-to-one correspondence between the ASL and T1 relaxation

data, and to enable direct calculation of the quantitative ASL maps described below.

Quantitative, voxel-wise perfusion maps were generated using previously described

methods for the FAIR ASL technique according to (Kwong et al., 1995, Kim et al.,

1997):

where f is the tissue perfusion in mL/min/100 g of tissue, NS, SS and M0 are the signal intensities for the non-selective, slice-selective and reference (M0) ASL-FISP images,

respectively, T1 is the longitudinal relaxation time calculated from the Look–Locker

acquisition, TI is the inversion time set at 1420 ms and the tissue–blood partition

coefficient (λ) was assumed to be 0.9 mL/g (Herscovitch et al., 1985). Perfusion maps were calculated using this equation for each ASL-FISP scan.

A region of interest (ROI) analysis was then used to calculate the mean perfusion over the entire mouse brain visible within the single ASL-FISP imaging slice.

It should be noted that these ROIs included both white and gray matter, whereas the ventricles were excluded from the analysis. A histogram analysis of the mouse brain

ROI was also used to calculate a threshold (mean ± two standard deviations) to limit the impact of large vessels (high perfusion values) on the calculation of the mean brain perfusion value for each animal, similar to previously described methods (Boss et al.,

2007, Martirosian et al., 2004). The mean brain perfusion values for each individual mouse were then used to calculate an overall group average and standard deviation at

7 and 9.4 T, respectively, for comparison. This ROI analysis was repeated for 20, 40 and 60 ASL-FISP averages at 7 and 9.4 T to perform an initial determination of the 111

effects of noise on the brain perfusion assessments. For all C57/BL6 mice, ASL-FISP

scans were obtained with an inversion slab thickness three times that of the FISP

imaging slice thickness to ensure a uniform inversion over the entire FISP imaging

slice. This factor of three was previously described by our group (Lu et al., 2012) and provides an effective balance between blood flow sensitivity and inversion uniformity.

One C57/BL6 mouse was scanned with inversion slab thickness/imaging slice thickness ratios of 1, 3, 6 and 10 to determine the effects of the relative inversion slab thickness on the perfusion results.

Additional in vivo ASL-FISP experiments: mouse brain ischemic stroke model and healthy mouse kidneys

The ASL-FISP acquisition and analysis protocol described above was performed on two additional C57/BL6 mice to evaluate: (1) brain perfusion in the context of known pathology (ischemic stroke); (2) kidney perfusion; and (3) muscle perfusion. Except for the number of signal averages (Durukan et al., 2009), all of the

imaging parameters used for these imaging studies were identical to those of the brain

ASL-FISP experiments described above.

An ASL-FISP study was performed on a mouse model of ischemic stroke to assess the method's sensitivity to a known pathophysiology. A C57/BL6 mouse (male,

8 weeks of age) was anesthetized with isoflurane. A transient stroke was initiated by inserting a 0.22-mm-diameter, silicone-coated filament (Doccol Corporation, Sharon,

MA, USA) into the right carotid artery, resulting in an occlusion of the middle cerebral

artery (MCAo), whilst monitoring cerebral blood flow using laser Doppler flowmetry

(Schmid-Elsaesser et al., 1998, Harada et al., 2005, Durukan & Tatlisumak, 2009). 112

The filament was removed after 1 h of occlusion to allow reperfusion as the animal recovered. At 24-h post-ictus, the animal was re-anesthetized for ASL-FISP scanning

as described above. A diffusion-weighted EPI image

(TR/TE = 5000/31 ms; b = 500 s/mm2) was also acquired to confirm the presence of

the infarct from the MCAo. A brain perfusion map and a mean perfusion value for the

whole brain within the imaging slice were calculated for comparison with healthy

C57/BL6 mice.

For the kidney and paraspinal muscle perfusion assessment, ASL-FISP was

performed on a healthy male 10-week-old C57/BL6 mouse. Respiratory triggering was

performed as described above. This study was performed as an initial demonstration

of the effectiveness of ASL-FISP for high-field body imaging applications. Following

the ASL-FISP acquisitions, an ROI analysis was used to calculate the mean renal

perfusion values for the left and right kidneys of the mouse and the mean perfusion

value of the paraspinal muscles.

RESULTS

Axial mouse brain images from spin echo, FISP, true FISP and EPI techniques at

7 T are shown in Fig. 4.2. Banding and ghosting/distortion artifacts are clearly visible

Figure 4.2. Representative axial mouse brain images at 7T using spin echo (a), fast imaging with steady- state free precession (FISP) (b), true FISP (c) and echo-planar imaging (EPI) (d) acquisitions. Note the similar lack of artifacts in the spin echo and FISP images in comparison with the true FISP (banding) and EPI (ghosting/distortion) images. 113 in the true FISP and EPI images, respectively. By comparison, the spin echo and FISP images show a lack of image artifacts, with the FISP images generated in less than one-tenth of the acquisition time (22 s versus 4 min 16 s). Representative brain ASL-

FISP image sets, T1 relaxation time maps and calculated perfusion maps are shown for a healthy C57/BL6 mouse at 7 T (n = 10) in Fig. 4.3. Qualitatively similar images

Figure 4.3. Representative arterial spin labeling-fast imaging with steady-state free precession (ASL- FISP) images of a healthy C57/BL6 mouse brain at 7T: (a) slice-selective (bright blood) ASL-FISP image (40 averages); (b) non-selective (dark blood) ASL-FISP image (40 averages); (c) M0 image (no inversion); (d) brain T1 map from Look-Locker acquisition; (e, f) perfusion map from ASL-FISP (e) and ASL-GRE (f) (in mL/min/100g of tissue). Note that the ASL-FISP and ASL-GRE perfusion maps are from different mice. GRE, gradient echo. 114

and maps were obtained from the mice at 9.4 T (n = 5, data not shown). The means

and standard deviations for the groups of healthy mice scanned at 7 and 9.4 T are

shown in Fig. 4.4a as a function of the number of ASL-FISP averages used to calculate the perfusion data (20, 40 and 60 averages). A two-tailed Student's t-test showed no

significant difference between the mean perfusion values at the two magnetic field strengths (p > 0.2) or for 20, 40 and 60 averages (p > 0.6). With 40 signal averages, the mean brain perfusion values for healthy C57/BL6 mice ranged from 210–

333 mL/min/100 g of tissue at 7 T to 222–268 mL/min/100 g of tissue at 9.4 T. The means ± standard deviations of the mouse brain perfusion (excluding ventricles) at 7 and 9.4 T were 249 ± 38 and 241 ± 17 mL/min/100 g of tissue, respectively.

A secondary ROI analysis of the 7-T mouse brain perfusion data showed differential perfusion values in the cortex (211 ± 30 mL/min/100 g) and thalamus

(288 ± 48 mL/min/100 g), which is consistent with previous studies (Muir et al.,

2008). A perfusion map from the ASL-GRE method (mean cerebral perfusion,

285 mL/min/100 g) is shown in Fig. 4.3f for comparison. The total imaging time for five averages was approximately 4 h. In order to make a valid comparison between

protocols, the time under anesthesia should be consistent to ensure similar perfusion

conditions. We chose to evaluate the ASL-GRE method with two averages so that the

imaging time was approximately the same as ASL-FISP. The mean cerebral perfusion

value from the ASL-FISP method (40 averages) was 249 ± 38 mL/min/100 g, which is quite reasonable compared with the cerebral perfusion value of 240 mL/min/100 g from the ASL-GRE method (two averages). 115

The mean brain perfusion values from a single C57/BL6 mouse using inversion

slab thickness/imaging slice thickness ratios of 1, 3, 6 and 10 and 5, 10, 20, 40 or 60

Figure 4.4. (a) Mean C57/BL6 mouse brain perfusion from the arterial spin labeling-fast imaging with steady-state free precession (ASL-FISP) method at 7T (gray) and 9.4T (white). Results are plotted as a function of the number of ASL-FISP averages. No significant differences in mean perfusion were observed for different numbers of averages (p>0.6) or field strengths (p>0.2). (b) Mean brain perfusion from a single C57/BL6 mouse at 7T as a function of the slice thickness ratio is increased from 1 to 3. The mean perfusion also appears to be more sensitive to the inversion slab thickness than to the number of ASL-FISP averages. ASL-FISP averages are shown in Fig. 4.4b. A large decrease in perfusion was observed

as the inversion slab thickness/imaging slice thickness ratio was increased from 1

(inversion slab thickness = imaging slice thickness) to 3, as a result of both increased uniformity of the inversion pulse over the entire imaging slice and reduced perfusion sensitivity. The mean brain perfusion values continued to decrease at a lower rate as the inversion slab thickness/imaging slice thickness ratio was increased from 3 to 6 and 10. In addition, only minimal variation was observed in the mean perfusion values as the number of ASL-FISP averages was reduced from 60 to 5 for each ratio.

Axial ASL images, T1 maps and brain perfusion maps for the MCAo model of

ischemic stroke in a mouse are shown in Fig. 4.5. The region of infarct on the right side of the brain (left side of the image) is clearly visible in the ASL-FISP images and

the corresponding perfusion map. It should be noted that a smaller secondary infarct 116 is visible on the contralateral side, most likely resulting from the extensive cytotoxic edema caused by the induced ischemic stroke. Overall, the mean brain perfusion for

Figure 4.5. Arterial spin labeling-fast imaging with steady-state free precession (ASL-FISP) images of a middle cerebral artery occlusion (MCAo) mouse model of stroke at 7T: (a) slice-selective (bright blood) ASL-FISP image; (b) non-selective (dark blood) ASL-FISP image; (c) M0 FISP image (no inversion); 2 (d) diffusion-weighted image (b=500s/mm ) showing right brain infarct; (e) Look-Locker T1 map; (f) perfusion map. The primary infarct is visible in the right brain in all images. A potential contralateral perfusion deficit is also observed in the perfusion map, but is less evident in T1 and diffusion-weighted images. 117

the MCAo mouse was measured to be 143 mL/min/100 g. Axial ASL images, T1 maps and perfusion maps for the C57/BL6 mouse kidneys are shown in

Fig. 4.6. As expected, the aorta and renal arteries show a very bright signal in the slice-

selective inversion ASL-FISP image (Fig. 4.6a), whereas the arterial blood signal is

Figure 4.6. Kidney arterial spin labeling-fast imaging with steady-state free precession (ASL-FISP) images from a healthy C57/BL6 mouse at 7T: (a) slice-selective (bright blood) ASL-FISP image; (b) non-selective (dark blood) ASL-FISP image; (c) M0 FISP image (no inversion); (d) Look-Locker T1 map; (e) perfusion map. Renal arteries (high perfusion) and renal medulla (low perfusion) are clearly visible in the perfusion map. greatly attenuated for the non-selective ASL-FISP image (Fig. 4.6b). The measured mean renal perfusion values for the left and right kidneys (cortex + medulla + pelvis) were 513 and 560 mL/min/100 g, respectively. In addition, the perfusion in the renal cortex was visibly increased relative to that in the renal medulla, as expected.

Importantly, mouse kidney perfusion was approximately twice that of brain perfusion, consistent with previous reports (Duhamel et al., 2008, 2014, Rajendran et al.., 2013,

Kober et al., 2008). The mean perfusion value in the paraspinal muscles was

81 mL/min/100 g, which was significantly lower than that of the kidneys, as expected (Duhamel et al., 2014).

118

DISCUSSION

In this study, initial in vivo ASL MRI results were obtained using a rapid and

artifact-resistant ASL-FISP acquisition on 7- and 9.4-T small-animal MRI scanners.

The ASL-FISP technique combines an inversion preparation (slice-selective or non- selective), followed by a centrically encoded FISP imaging readout, to provide ASL data with minimal image artifacts in comparison with conventional EPI and true FISP imaging readouts for ASL MRI acquisitions.

There are several important design features of the ASL-FISP acquisition. Most

importantly, the FISP imaging readout is applied with centrick-space encoding and

relatively few (i.e. 10) dummy scans. As shown previously, this centric encoding

approach minimizes the loss of perfusion sensitivity caused by additional RF pulses

and gradient lobes encountered in linear k-space encoding (Shah et al., 2011). A FISP

imaging readout was selected for this study instead of a balanced SSFP readout

primarily to limit well-known banding artifacts that are increased significantly on high-

field MRI scanners. An alternative approach using balanced SSFP would be to acquire

multiple images with different RF phase variation sequences to reconstruct a banding-

free image (Vasanawala et al., 2000, Cukur et al., 2008). This approach may offer

increased signal-to-noise, but may require additional image processing to remove the

banding artifacts. As a result, this alternative approach and direct comparison with

ASL-FISP were not explored for this initial study. Further studies are also needed to

directly compare the FISP and true FISP techniques, as the FISP flow sensitivity may

be altered by dephasing (Haacke et al., 1990). 119

Overall, the key advantage of the FISP imaging readout is that it provides

perfusion sensitivity with a short acquisition time (~2 s/image) with minimal artifacts

in comparison with EPI readouts. The ASL-FISP and ASL-GRE acquisitions

generated relatively similar perfusion maps (Fig. 4.3) and mean brain perfusion values.

However, the ASL-GRE method required an acquisition time that was more than 50 times that of the ASL-FISP method. In addition, the FISP imaging readout can easily be coupled with virtually any ASL preparation in order to meet the requirements for

specific imaging applications. For this initial implementation, a simple FAIR ASL

preparation was implemented with either a slice-selective or non-selective inversion

pulse. However, more complex ASL preparations, such as pCASL, could also be implemented in order to measure transit times and other important perfusion

parameters (Duhamel et al., 2014). The FISP imaging readout is particularly relevant

for FAIR acquisitions as the difference between the images with the slice-selective and

non-selective inversion is small relative to the M0 image (typically <10%). As a result,

FAIR ASL studies generally require numerous signal averages to obtain a reasonable

estimate of tissue perfusion. Therefore, the acquisition of all k-space lines following a

single ASL preparation (with minimal artifacts) using the FISP readout results in

practical imaging times to acquire sufficient signal averages. Although only single-slice

ASL-FISP results are presented in this initial study, multi-slice and/or three-

dimensional ASL-FISP implementations are possible, and may have significant

advantages for specific imaging applications.

Initial in vivo mouse brain perfusion studies demonstrated that the ASL-FISP

technique provides reasonable perfusion assessments on high-field MRI scanners. 120

ASL-FISP images in the healthy mouse brain and in an induced ischemic stroke model

show an expected lack of distortion and artifacts, and clearly delineated perfusion

deficits in the stroke model (Figs 4.3 and 4.5, respectively). Further, the kidney ASL-

FISP images and perfusion maps in Fig. 4.6 show differential perfusion between the renal cortex and medulla, as expected from previous studies (Martirosian et al., 2004,

Miles et al., 1991). In addition, the renal arteries are clearly visible in both the slice- selective inversion ASL-FISP images and the perfusion maps, demonstrating the flow sensitivity of the ASL-FISP technique. Overall, the reliability of the ASL-FISP technique is exhibited by the lack of differences in the perfusion results at 7 and 9.4 T

(Fig. 4.4a). Most importantly, the mouse kidney images shown in Fig. 4.6 demonstrate that the ASL-FISP technique can provide high-quality ASL data for rodent brain and body imaging applications on high-field MRI scanners with no distortion and artifacts.

The ASL-FISP technique presented herein also has several important limitations. One key observation of these initial results is that the mean perfusion values for mouse brains and kidneys shown here are dependent on the perfusion preparation scheme and inversion pulse design. It has already been shown in multiple studies that the relative thickness between the slice-selective inversion and the imaging readout is directly related to the resulting tissue perfusion estimate (Brookes et al.,

2007, Karger et al., 2000, Esparza-Coss et al., 2010, Yongbi et al., 1999). For example, a smaller inversion slab thickness (e.g. one times that of the imaging slice) will result in enhanced perfusion sensitivity and erroneously high perfusion estimates.

Conversely, a larger inversion slab thickness (e.g. six times that of the imaging slice) will result in reduced perfusion sensitivity. These results are reflected in Fig. 4.4b 121

which shows that an inversion slab thickness/imaging slice ratio ≥ 3 is needed to maintain reasonably consistent perfusion results. The perfusion results can also be influenced directly by the shape of the inversion pulse and the excitation pulses of the

FISP imaging readout. However, optimization of these pulses was beyond the scope of this initial technical development. Nevertheless, the results shown herein confirm that the ASL-FISP technique can sensitively differentiate normal tissue perfusion from pathology (e.g. ischemic stroke) and relative tissue perfusion levels (renal cortex versus renal medulla versus skeletal muscle). It is important to note that the ASL-

FISP technique may also be sensitive to the effects of pulsatility, which may be an underlying cause of the bright cerebrospinal fluid signal in the mouse brains.

Another observation of these initial ASL-FISP results is the trend towards lower perfusion values at 9.4 T. Although not statistically significant, this trend may

be caused, in part, by the reduced T2* relaxation times at 9.4 T. For mouse brain

imaging, this reduction in T2* can reduce the SNR of the SS, NS and M0 images,

especially in regions near the ear canals. Fortunately, this potential limitation can be

partially mitigated using shorter TEs which would reduce the deleterious T2* effects at

all field strengths. For the FISP acquisition, the reduction in TE would provide an

additional increase in SNR, as TR would also be reduced by the same percentage,

providing an increase in the coherent steady-state magnetization.

Conclusions

This study reports a rapid and quantitative ASL-FISP MRI technique for high-

field MRI scanners. For this initial study, the ASL-FISP technique combines a FAIR

ASL preparation with a rapid, centrically encoded FISP imaging readout to provide 122

perfusion-weighted images in less than 2 s with minimal image distortion, ghosting

and banding artifacts in comparison with EPI and balanced SSFP readouts.

Initialin vivo ASL-FISP perfusion results were obtained for healthy and ischemic stroke

C57/BL6 mouse brains at 7 T and healthy mouse brains at 9.4 T. As a demonstration

of the invulnerability of the ASL-FISP technique to off-resonance artifacts,

initial in vivo kidney ASL results were also obtained for a C57/BL6 mouse at 7 T. This

new technique provides an alternative method for many perfusion imaging

applications on high-field MRI scanners where off-resonance artifacts can severely

limit the use of EPI and balanced SSFP acquisitions.

123

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126

CHAPTER 5: LACK OF DYSTROPHIN RESULTS IN ABNORMAL

CEREBRAL DIFFUSION AND PERFUSION IN VIVO

ABSTRACT

Dystrophin, the main component of the dystrophin-glycoprotein complex, plays an

important role in maintaining the structural integrity of cells. It is also involved in the

formation of the blood-brain barrier (BBB). To elucidate the impact of dystrophin

disruption in vivo, we characterized changes in cerebral perfusion and diffusion in

dystrophin-deficient mice (mdx) by magnetic resonance imaging (MRI). Arterial spin

labeling (ASL) and diffusion-weighted MRI (DWI) studies were performed on 2-

month and 10-month mdx mice and their age-matched wildtype controls (WT). The

imaging results were correlated with Evan’s blue extravasation and vascular density

studies. The results show that dystrophin disruption significantly decreased the mean

cerebral diffusivity in both 2-month (7.38 ± 0.30 x 10- 4 mm2/s) and 10-month

(6.93 ± 0.53 x 10- 4 mm2/s) mdx mice as compared to WT (8.49 ± 0.24 x 10- 4,

8.24 ± 0.25 x 10- 4 mm2/s, respectively). There was also an 18% decrease in cerebral

perfusion in 10-month mdx mice as compared to WT, which was associated with

enhanced arteriogenesis. The reduction in water diffusivity in mdx mice is likely due

to an increase in cerebral edema or the existence of large molecules in the extracellular

space from a leaky BBB. The observation of decreased perfusion in the setting of

enhanced arteriogenesis may be caused by an increase of intracranial pressure from

cerebral edema. This study demonstrates the defects in water handling at the BBB and

consequently, abnormal perfusion associated with the absence of dystrophin. 127

INTRODUCTION

The blood-brain barrier (BBB) ensures the proper physical and metabolic

environment for brain function. Endothelial cells lining the vasculature of the brain

are joined by tight junctions in order to prevent free diffusion of large and hydrophilic

molecules, including water, from the blood into the brain parenchyma (Brightman and

Reese, 1969 and Reese and Karnovsky, 1967). A major role of the tight junctions is to

maintain the physiologic and biochemical environment necessary for neuronal firing.

The cytoskeleton is crucial in forming, anchoring, and maintaining the BBB (Zlokovic,

2008). Disruption of the BBB leads to increased permeability of large molecules, such

as albumin, into the CNS, causing neuronal dysfunction, edema, and a lower seizure

threshold (Abbott et al., 2006, Hawkins and Davis, 2005, Tomkins et al., 2008 and Van

Vliet et al., 2007).

Dystrophin, a major actin-binding component of the dystrophin glycoprotein

complex (DGC), links cytoskeletal and membrane elements in the muscle (Tinsley et

al., 1994) and brain (Lidov et al., 1990 and Lidov et al., 1993). Inherited deficiency of

functional dystrophin leads to Duchenne muscular dystrophy (DMD), an X-linked

recessive disease leading to progressive muscle degeneration. In addition to voluntary

muscle weakness, DMD patients suffer from ventilatory and cardiovascular failure at

the end stages of the disease. Functional dystrophin is also not expressed in the brains

of DMD patients (Hoffman et al., 1987). However, aside from the observation that

pediatric DMD patients have lower intelligence quotient (IQ) assessments than that of

healthy control children (Cotton et al., 2001 and Karagan, 1979), few studies have 128

characterized the structural and functional impact of dystrophin disruption in the

brain.

The dystrophin-null mdx mouse has been used as a mouse model of muscular

dystrophy for over two decades (Sicinski et al., 1989). Aside from the muscular

phenotype, cognitive defects have also been observed in mdx mice (Vaillend et al.,

1995). The tight junction proteins of the BBB were shown to be reduced and

dysfunctional in the mdx mouse brain, leading to a leaky BBB (Nico et al., 2003).

Moreover, enhanced expression of matrix-metalloproteinase (MMP)-2 and -9 in mdx

mouse brains was associated with increased expression of vascular endothelial growth

factor (VEGF, Nico et al., 2006). These in vitro observations provide the evidence of

altered brain structure associated with dystrophin deletion. A full understanding of the

role of dystrophin in maintaining the BBB and vascularization has yet to be studied in

vivo with modern imaging technologies which will likely support future clinical

investigations.

Arterial spin labeling (ASL) is a non-contrast MRI method that has made

significant contributions towards assessing tissue perfusion in vivo (Detre et al.,

1992, Williams et al., 1992, Edelman et al., 1994, Kwong et al., 1995, Kim and Tsekos,

1997, Wong et al., 1997, Pell et al., 1999 and Thomas, 2005). In ASL MRI, water

molecules are “magnetically tagged” in the blood, leading to altered tissue longitudinal

magnetization that is proportional to tissue perfusion. This method does not require

exogenous paramagnetic contrast agents as in conventional Dynamic Contrast

Enhanced (DCE) MRI perfusion techniques. Hence, ASL may eventually become the

preferred method for longitudinal imaging studies. In spite of its utility, ASL has yet 129

to be fully applied to understanding the pathophysiologic consequence of dystrophin

disruption with regard to water movement and perfusion in the brain.

Diffusion-weighted MRI (DWI) has been invaluable in defining neurological

disorders, particularly in the diagnosis of stroke and the assessment of therapeutic

interventions (Le Bihan et al., 1986, Kloska et al., 2010, Schellinger et al., 2003, Sevick

et al., 1990 and Warach et al., 1995). The signal intensity of a DW image reflects the

restrictions on Brownian motion of water molecules in the tissue, and the calculated

apparent diffusion coefficient (ADC) provides a means to quantify this diffusion under

physiologic and pathologic states. High ADC values are characteristic of tissue with

relatively free water diffusion, e.g., in extracellular space, as opposed to tissue water

with a restricted environment, e.g., in intracellular space (Le Bihan, 2007). Therefore,

the diffusion of water molecules as detected by DWI can be used to delineate the

neural structure, anatomy, and pathophysiology in the absence of dystrophin.

The goal of this study was to characterize the impact of dystrophin deletion on

physiological and structural changes in the brain using both in vivo and in vitro

methods. Cerebral perfusion and brain structure were evaluated by ASL and DWI,

respectively. These in vivo imaging results were compared with in vitro and ex vivo

studies of vascular density. The present study demonstrates the defects in perfusion

and diffusion associated with dystrophin disruption in mdx mice that can be

observed in vivo with MRI and the association of these in vivo imaging assessments

with histopathologic measures.

130

METHODS

Animal Models

Studies were performed on young (2-month, n = 10) and adult (10-month,

n = 10) male dystrophin-null (mdx) and wild-type (WT) mice of the C57/BL6 strain.

All mice were obtained from Jackson Laboratories (Bar Harbor, ME). All procedures

involving animal care and handling were performed according to institutional

guidelines set forth by the Animal Care and Use Committee at Case Western Reserve

University.

Perfusion and Diffusion MRI

Imaging studies were performed on a 7 T Bruker Biospec (Billerica, MA)

horizontal bore MRI scanner. Anesthesia was induced with 2% isoflurane with

supplemented O2 in an isoflurane induction chamber and maintained via nosecone

with 1.5% isoflurane once the animal was put in the magnet. The body temperature

was monitored and maintained at approximately 36 °C by blowing hot air into the

magnet through a feedback control system. Respiratory gating and monitoring was

performed through an MR-compatible small animal gating and monitoring system (SA

Instruments, Stony Brook, NY) to reduce motion artifacts during image acquisition.

Single-slice axial ASL brain images were acquired with a Flow-Sensitive Alternating

Inversion Recovery (FAIR) preparation sequence followed by a centrically-encoded

Fast Imaging in Steady Precession (FISP) imaging readout (Gao, et al., 2014).

Specifically, arterial spin labeling was accomplished by respiratory-triggered, slice-

selective (4.5 mm thickness) and non-selective (global) inversion of magnetization, 131

respectively. The inversion pulse used a hyperbolic secant adiabatic inversion pulse

with a duration of 3 ms, and the inversion thickness was set to three times that of the

excitation pulse to ensure a uniform inversion over the entire imaging slice. An

inversion efficiency value of 1 was assumed. The FISP acquisition (including 10

dummy scans) was implemented at 1420 ms following the inversion pulse. This delay

time allowed data acquisition to occur during the mid- to late-stage of the subsequent

breathing cycle. It is a balance between maximizing blood flow sensitivity and

minimizing respiratory motion artifacts. Imaging parameters are: flip angle, 60°; TR,

2.4 ms; TE, 1.2 ms; number of averages, 40; matrix size, 128x128; FOV, 30x30 mm2;

slice thickness, 1.5 mm. A proton density image (M0) with no inversion pulse was also

acquired using the same imaging parameters. A voxel-wise T1 map with the same

spatial resolution was also acquired with a FISP-based Look-Locker acquisition

consisting of a non-selective adiabatic inversion pulse followed by 10 continuous FISP

acquisitions with the following parameters: flip angle, 10°; TR, 4.0 ms; TE, 2.0 ms;

number of averages, 40. Image reconstruction and analysis were performed offline

using custom-built software written in Matlab (MathWorks, Natick, MA). Perfusion

maps were generated from the magnetization difference of slice-selective and non-

selective inversion images (ΔM), proton density image (M0), and the T1 maps (Gao, et

al., 2014). A blood/tissue partition coefficient (λ) of 0.9 ml/g was used in this study

(Herscovitch and Raichle, 1985).

Single-slice axial Diffusion-Weighted Images (DWI) were acquired with a

diffusion-weighted Echo-Planar Imaging (DW-EPI) acquisition. Diffusion weighting

was accomplished with two 4 ms (δ) diffusion-encoding gradients separated by 15 ms 132

(Δ), yielding a b value of 500 s/mm2 applied in the readout direction. The DW-EPI

images were acquired with the following parameters: TR, 5000; number of averages,

5; matrix, 128x128; FOV, 30 x 30 mm2; slice thickness, 1 mm. Images were

reconstructed and analyzed using custom-built software written in Matlab

(MathWorks, Natick, MA). Voxel-wise apparent diffusion coefficient (ADC) maps

were calculated by the established equation: ADC = ln(I0/I)/b, where I0 and I are the

signal intensity of the b = 0 s/mm2 and the b = 500 s/mm2 images, respectively.

Analysis of Vascular Density by Immunocytochemistry

WT (2 mo, n = 4; 10 mo, n = 5) and mdx (2 mo, n = 4; 10 mo, n = 5) mice

were deeply anesthetized with isoflurane after imaging. The brains were excised and

fixed in 4% paraformaldehyde for 24 h at 4 °C. The tissue was transferred to 5%

sucrose in PBS for 1 h followed by 20% sucrose overnight at 4 °C. Subsequently, the

brains were rapidly frozen in OCT media (Electron Microscopy Sciences, Hatfield,

PA) with liquid nitrogen. Tissues were sectioned at 10 μm slice thickness at the

location corresponding to the MR imaging slice. A total of 7 sections were obtained

from each brain. Indirect immunofluorescence against CD31 (PECAM-1, 1:100,

Sigma Aldrich, St. Louis, MO) was performed to assess cerebral vascular density.

DAPI (4’,6-Diamidino-2-phenylindole dihydrochloride, Sigma Aldrich, St. Louis,

MO) staining was used for nuclear identification. Stained tissue sections were

photographed for the DAPI (blue) and PECAM-1 (red) channels. The number of

PECAM-1 positive cells per field was quantified. All images were photographed at the

same exposure times and magnification. The obtained values from all animals were 133

averaged, and the results were expressed as the mean number of PECAM-1 positive

cells per field ± SD.

Analysis of BBB Leakage using Evans Blue Staining

WT (2 mo, n = 2; 10 mo, n = 2) and mdx (2 mo, n = 2; 10 mo, n = 2) mice

were injected intraperitoneally with a bolus of 0.1 mL/10 g Evans Blue (10 mg/ml,

Sigma Aldrich, St. Louis, MO). After 24 hours, the mice were deeply anesthetized

with isoflurane and the brains were excised and rapidly frozen in OCT media (Electron

Microscopy Sciences, Hatfield, PA). Tissues were sectioned at 10 μm slice thickness

at the location corresponding to the MR imaging slice. Evans Blue extravasation was

evaluated by fluorescence in WT versus mdx.

Analysis of Vascular Density by Cryo-Imaging

The remaining WT (2-month, n = 2; 10-month, n = 2) and mdx (2-month,

n = 2, 10-month, n = 2) mice were anesthetized with isoflurane and Fluorescein

isothiocyanate-dextran (FITC-dextran, 10 mg/ml, Sigma Aldrich, St. Louis, MO) was

injected intravenously at a dose of 200 mg/kg. After 10 minutes, the brains were

excised and rapidly frozen with liquid nitrogen in OCT media (Electron Microscopy

Sciences, Hatfield, PA) and mounted to the cryo-imaging system. The tissue was

sectioned in 25 μm slices and the bright-field and fluorescent images from the cryo-

imaging system were processed and analyzed with custom-built Matlab (Mathworks

Inc., Natick, MA) and AMIRA (Mercury Computer Systems, San Diego, CA)

software (Roy et al., 2009). The brain contour was traced manually from the bright-

field images, and a mask was generated. The mask was applied to fluorescent images, 134

and the 3D cerebral vasculature was reconstructed using thresholding and volume

rendering methods (Qutaish et al., 2012). The resolution for the reconstructed 3-D

images were 10.5x10.5x25 μm3.

Statistical Analysis

All data are reported as mean ± SD. A two-tailed, unpaired Student’s t-test was

performed to compare mdx and WT mice. Statistical significance was established at a

level of p < 0.05.

RESULTS

Decreased cerebral diffusivity with dystrophin disruption

Representative brain apparent diffusion coefficient (ADC) maps of 2- and 10-

month old mdx and WT mice are shown in Figs. 5.1a-d. The mean and standard

Figure 5.1. a-d. Representative diffusion images of 2-month-old WT (a), 2-month-old mdx (b), 10- month-old WT (c), 10-month-old mdx (d). e. Mean cerebral (excluding ventricles) diffusion in C57/BL6 WT (black) and mdx (white), respectively. There was a significant decrease in mean diffusion as compared to WT observed in the mdx brains in both the 2- and 10-month old mice (*p<0.0005). There was also a significant age-dependent decrease in diffusion in young versus aged WT and mdx mice, respectively (#p<0.05). Data are represented as mean ± SD. Color bar is uniform in each panel. 135 deviations for each group of mice are shown in Fig. 5.1e. Mean diffusivity in 2-month mdx mice was 7.38 ± 0.30 x 10- 4 mm2/s, which was significantly lower than that in age-matched WT mice (8.49 ± 0.24 x 10- 4 mm2/s, Fig. 5.1e, p < 0.0005). Further, 10- month mdx mice also exhibited a decrease in cerebral mean diffusivity as compared to the age-matched WT mice (6.93 ± 0.53 x 10- 4 versus 8.24 ± 0.25 x 10- 4 mm2/s, p < 0.0005). In addition, a 2-tailed Student’s t-test showed a significant age-related decrease in the mean ADC values of both the WT and mdx mice (p < 0.05).

Figure 5.2. a-d. Representative perfusion images of 2-month-old WT (a), 2-month-old mdx (b), 10- month-old WT (c), 10-month-old mdx (d). e-h. Representative T1 maps of 2-month-old WT (e), 2-month- old mdx (f), 10-month-old WT (g), 10-month-old mdx (h). i. Mean cerebral (excluding ventricles) perfusion from the ASL-FISP method in C57/BL6 WT (black) and mdx (white), respectively. There was a significant decrease in mean perfusion as compared to aged WT observed in the aged mdx mice (*p<0.005). There was no significant change in perfusion in young versus aged WT (p>0.3), respectively. Data are represented as mean ± SD. Color bar is uniform in each panel. 136

Abnormalities in Cerebral Perfusion in Aged mdx Mice

Representative cerebral perfusion maps for the 2 and 10 month old mdx and WT mice are shown in Figs. 5.2a-d. The mean and standard deviations for each group are shown in Fig. 5.2e. There was a 15% decrease in cerebral perfusion in 10 month mdx mice as compared to WT (Fig. 5.2e, p = 0.001), without any significant difference in mean cerebral T1 or M0 values (p = N.S.). The mean brain perfusion values for the 2-month

old mdx mice were slightly lower than for 2-month old WT mice. However, no

significant difference in cerebral perfusion was observed.

Disruption of Blood Brain Barrier Integrity in mdx Mice

To investigate the integrity of

the BBB in the absence of

dystrophin, we examined the

extravasation of Evans blue

from blood vessels

histologically. There was a

significant leakage of Evans

blue into the cerebrum of both

2- and 10-month mdx mice as

demonstrated in Figs. 5.3a,c. Figure 5.3. a-d. Representative images of Evan’s blue fluorescence on axial sections of frozen 2-month-old mdx (a), There were few vessels 2-month-old WT (b), 10-month-old mdx (c), 10-month-old WT (d). Note the extravasation of Evan’s blue (arrows) in the demonstrating Evans blue mdx mice. 137

extravasation in both young (2-month old) and aged (10-month old) WT mouse brains

(Figs. 5.3b,d).

Enhanced Cerebral Arteriogenesis in Aged mdx Mice

Representative PECAM-1 stained immunofluorescent images for the 2- and 10-

month mdx and WT mice are shown in Figs. 5.4a-d. There was no difference in the

amount of cerebral vasculature (number of PECAM-1 positive cells) between the

young mdx and WT mice (Fig. 5.4e). Interestingly, there was enhanced arteriogenesis

in the aged (10 month) mdx mice as compared to the controls, as demonstrated by a

13% increase in PECAM-1 positive cells (Fig. 5.4e, p < 0.05). Representative 3D vessel

reconstructions from the FITC-dextran cryoimaging are shown in Fig. 5.5.

Qualitatively, there were more vessels visible in the 2- and 10-month mdx mouse

brains as compared to WT.

Figure 5.4. a-d. Representative images of indirect immunofluorescence against CD31 (platelet endothelial cell adhesion molecule 1, PECAM-1) on axial sections of paraformaldehyde-fixed frozen 2- month-old WT (a), 2-month-old mdx (b), 10-month-old WT (c), 10-month-old mdx (d). e. Quantification of PECAM-1 positive cells in 2- and 10-month-old mdx and WT mice. Data are represented as mean ± SD. *p<0.05 versus WT.

138

Figure 5.5. a-l. Representative dorsal (top row), lateral (middle row), and rostral (bottom row) views of the 3D vessel reconstructions from 2-month-old WT (a-c), 2-month-old mdx (d-f), 10-month-old WT (g-i), and 10-month-old mdx (j-l).

DISCUSSION

In this study, we report the first in vivo evaluation of age-dependent alterations

in cerebral perfusion and diffusion in mdx mice. In addition to BBB disruption, we

have observed increased arteriogenesis in the cerebrum as a result of dystrophin

deletion. Traditional Evans Blue extravasation confirmed an interruption at the BBB

in both the 2- and 10-month mdx mice. In addition, in vivo DWI experiments

established an alteration of the normal cerebral microstructural and physiologic

environment in both 2- and 10-month mdx mice. Interestingly, decreased in vivo

perfusion was also present in the 10-month mdx mice despite increased arteriogenesis

observed in both cryoimaging and immunocytochemistry. 139

Consistent with the findings by immunocytochemistry, cryoimaging

demonstrates the extensive arteriogenesis in mdx throughout the entire brain as

compared to WT. Previously, enhanced arteriogenesis was observed in the hindlimbs

of mdx mice (Straino et al., 2004). It was also shown that deletion of dystrophin results

in enhanced expression of matrix-metalloproteinase (MMP)-2 and -9, which

ultimately leads to an increase of VEGF and VEGFR2 expression (Nico et al., 2006).

In addition to angiogenesis, increased VEGF activity may also contribute towards an

opening of tight junctions at the BBB and enhanced vascular permeability (Schoch et

al., 2002 and Zhang et al., 2002). The observed increase in arteriogenesis by both cryo-

imaging and immunocytochemistry in the current study is consistent with the

observations of enhanced VEGF activity in vitro (Nico et al., 2002). However, unlike

the qualitative results from cryoimaging, we did not see a difference in PECAM-1

expression in mdx versus WT mice at 2-months of age. The high resolution, but limited

sampling of the immunofluorescence experiments may better reflect the molecular

content of the tissue, whereas the 3D cryoimaging data are more representative of the

global macrovascular structure. As such, the global nature of cryoimaging as described

herein provides unique and complementary information for these types of

investigation.

The decreased cerebral ADC observed in the current study likely represents

cellular edema as a result of BBB disruption. BBB disruption in mdx mice due to

reduced expression of tight junction proteins has been reported previously (Nico et al.,

2003). Consequently, the osmotic influx of water into the cell causes cellular edema

and a decrease in extracellular space, leading to a decrease in ADC (Kucharczyk et 140

al., 1991 and Moseley et al., 1990). While it has been shown that aqp4 facilitates the

clearance of vasogenic edema as a result of BBB disruption (Papadopoulos et al.,

2004), the lack of properly functioning aqp4 channels at the BBB in mdx mice (Adams

et al., 2008 and Frigeri et al., 2001) may prevent the removal of water from the

cerebrum and therefore worsen cellular edema. Alternatively, a leaky BBB many also

lead to an increase of large molecules in the extracellular space in mdx mice, which

may also limit water diffusivity in the extracellular space. Therefore, the observed

ADC decrease in mdx brains may be the result of cellular edema, increased

extracellular protein content, enhanced vascular permeability, or a combination of

these possible mechanisms. The age-dependent reduction of ADC in the 10-month

WT as compared to 2-month is supported by prior studies, which describe decreasing

ADC with age likely due to cellular atrophy and an increase in cerebrospinal fluid

(CSF) compartments (Heiland et al., 2002).

To the best of our knowledge, this is the first study that demonstrated decreased

perfusion in the setting of increased angiogenesis and a chronically leaky BBB in the

mdx brain in vivo. This decrease in cerebral perfusion is likely the result of cerebral

edema, as observed by DWI. It has been established that the presence of cerebral

edema and/or angiogenesis in the fixed volume of the skull leads to an increase in

intracranial pressure (ICP, Adams and Ropper, 1997 and Mokri, 2001). Increased ICP

in this way could decrease cerebral perfusion pressure (CPP), which is the difference

between mean arterial pressure and ICP. The control of CPP ensures proper brain

function as decrease in CPP may lead to ischemia and an increase may contribute

towards raising the ICP (Powers, 1991 and Schumann et al., 1998). As a result of 141

increased ICP, the consequent decrease of CPP ultimately leads to an autoregulatory

decrease in cerebral blood flow. This decrease in perfusion in mdx mice may

compromise their ability to react to periods of hypoxia.

There are several limitations in this initial study aimed at investigating whether

there were overall changes in diffusion and perfusion in the brain of mdx mice that can

be detected by MRI. First, fast imaging sequences were used for both DWI and ASL

acquisitions. The voxel size and slice thicknesses used gave rise to partial volume

averaging that obscured the small white matter tracts in a mouse brain in vivo. Future

studies aimed at white/gray matter differentiation/evaluation will require

optimization of the imaging parameters or potentially using a different imaging

readout approach. Second, instead of performing a comprehensive diffusion tensor

imaging study, we performed DWI in order not to overtly extend the acquisition time.

While DWI is sensitive to structural changes at the cellular level (e.g., edema), a future

diffusion tensor study will be helpful to validate and refine our current findings.

Third, measurement of cerebral perfusion used a single inversion time (TI) and

the classic model that assumes equal T1 for blood and tissue (Detre et al., 1992, Kwong

et al., 1995 and Kim and Tsekos, 1997). Further, transit time is not considered in the

calculation. A more recent model by Pell et al incorporated both the transit time (δ)

and the T1 of the blood (T1a) and the tissue (T1app) separately (Pell et al., 1999). While

the method requires the quantification of both T1app and T1a by using multiple inversion

times, it allows more comprehensive assessment of several physiological parameters

that can impact blood flow. In calculating the cerebral blood flow (CBF), the classic

model and the Pell model used the following equations respectively: 142

where FC and FP are the two terms that the classic model and the Pell model differ.

These two terms are defined as:

And T1 and T1app are related by:

Hence, for a measured , the ratio of measured blood flow by the two methods is:

Using a transit time of 50 ms (Thomas, 2005), a blood T1 of 2.2 and 2.4 s at 7 T and

9.4 T, respectively (ex vivo measurement) (Dobre et al., 2007), and a tissue T1 of 1.4,

1.6, and 1.8 s respectively, the simulated differences between the two models with CBF

ranging from 100 to 400 ml/min/100 g are shown in Fig. 5.6. These results suggest

that at a field strength of 7 T, the difference between the classic model and the Pell 143

model was < 3% when tissue T1 was 1.8 s. However, the classic model overestimated

CBF by 12% to 16% when tissue T1 was 1.4 s (Figs. 5.6.a&b). In our current study,

measured tissue T1 was ~ 1.7 s. Hence the calculated CBF difference should be within

8% compared to the Pell model. At 9.4 T, there could be an up to 12% over-estimation

by classic model (Figs. 5.6.c&d).

Figure 5.6. Comparison of the classic model and the Pell model. (a) and (b). Simulated Fc and Fp, and CBFc/CBFP at 7T, respectively. (c) and (d). Simulated Fc and Fp, and CBFc/CBFP at 9.4T, respectively. Solid lines in (a) and (c) represent the classic model, while dotted lines represent the Pell model. Black, red, and blue represent tissue T1 of 1.4, 1.6, and 1.8 s, respectively.

In summary, we have observed abnormal cerebral diffusivity, enhanced

angiogenesis, and decreased cerebral perfusion in the mdx mouse. Our study is the 144

first to assess the consequence of dystrophin deletion on cerebral blood flow in the

mdx mouse model. In vivo mouse studies are essential to further the translational value

of these results. Although this work is a step towards clinical translation, better mouse

models of DMD need to be studied in further detail since the mdx mouse has a mild

phenotype in comparison to the clinical presentation of DMD. The leaky BBB and

decreased cerebral perfusion observed in mdx mice may contribute towards the

disruption of proper brain function observed in DMD patients. As the life expectancy

of DMD patients increases with improved therapies, developing an understanding of

the neurologic manifestations of the dystrophin-null phenotype will lead to better

patient management when symptoms emerge. In particular, DMD patients suffering

from cardiac arrhythmias will be at an increased risk of stroke, and thus understanding

dystrophin’s role at the BBB and response to hypoxia is critical.

145

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CHAPTER 6: FUTURE DIRECTIONS

1) Mechanical dysfunction in other models of HCM

RATIONALE

This work identified the potential of MRI in detecting early phenotypic changes

of one mouse model of HCM. If future studies demonstrate the utility of MRI in

sensitively predicting early phenotypes of other HCM genotypes in mice, then MRI

may represent a broadly applicable early detection tool in identification of HCM in

humans. The most predominant HCM-cau

sing mutant genes are β-myosin heavy chain, myosin-binding protein C, and

cardiac troponin T (Table 6.1, (Houston and Stevens, 2014)). In addition, cardiac

troponin I, regulatory and essential myosin light chains, titin, α-tropomyosin, α-actin,

and α-myosin heavy chain gene

mutations have all been identified as

causes of HCM (Houston and

Stevens, 2014). In fact, more than

1400 mutations have been identified,

which are largely unique to

individual families (Landstrom et al.,

2010a). There is considerable clinical Table 6.1: Genes responsible for hypertrophic cardiomyopathy heterogeneity of HCM given the Adapted from Houston, B.A., and Stevens, G.R. (2014). Hypertrophic cardiomyopathy: a review. Clin diversity of molecular defects, Med Insights Cardiol 8, 53–65. including age of onset, degree of 150

LVH, and clinical course (Maron, 2002). In order for DENSE MRI to be of clinical

value in detecting preclinical HCM, the pattern of LV mechanical dysfunction must

be described in each of the sarcomeric protein mutations.

EXPERIMENTAL STRATEGY

Adult mouse models for each of the sarcomeric protein mutations causing HCM will be obtained (see Table 6.2) with appropriate age-matched controls. The MRI DENSE protocol will be repeated as previously described in Chapter 2 for each of the mouse models of HCM at several time points to better characterize the progression of

phenotypic expression (ages 2 weeks, 8 weeks, 6 months, and 10 months). LV twist,

torsion, circumferential and radial train, as well as torsion and strain rates will be

quantified as previously described (Zhong et al., 2008). Comparisons of strain and

twist angles will be independently analyzed at each ventricular level by 1-way

ANOVA and Tukey- Kramer test. Strain and torsion time course data will be

evaluated by 1-way ANOVA with a Bonferroni adjustment for multiple comparisons.

Statistical significance will be established at a level of P<0.05.

INTERPRETATION

I anticipate that the mechanical indices of cardiac function will be affected in other

HCM-causing mutations as what was observed in cMyBPC (Desjardins et al., 2012).

Given that there is significant heterogeneity in the phenotypic expression and clinical

course of HCM, I suspect that there will be differences in the timing and/or severity

of strain and twist alterations. For example, mutations in troponin T have been shown 151 to have very little LVH but high rates of sudden death (Moolman et al., 1997), cMyBPC mutations have been associated with late-onset of HCM (Niimura et al.,

1998b), and there was a family with mutations in β-myosin heavy chain with severe

HCM and early onset of LVH. Therefore, troponin T mutated mice might

demonstrate decreased mechanical function even earlier than seen in cMyBPC mutated mice, and β-myosin heavy chain mutated mice would show the most severe Table 6.2: Mouse and human genes identified in hypertrophic cardiomyopathy HUMAN GENE MOUSE REFERENCE GENE CSRP3: cysteine and glycine-rich protein 3 (cardiac LIM Csrp3 (Knöll et al., 2010) protein) MYBPC3: myosin-binding protein C, cardiac Mybpc3 (Harris et al., 2002) MYH6: myosin, heavy chain 6, cardiac muscle, alpha Myh6 (Palmer et al., 2004) ACTC1: actin, alpha, cardiac muscle 1 Actc1 (Song et al., 2011) CALR3: calreticulin 3 Calr3 (Ikawa et al., 2011) CAV3: calveolin 3 Cav3 (Galbiati et al., 2001) JPH2: junctophilin 2 Jph2 (Takeshima et al., 2000) MYH7: myosin, heavy chain 7, cardiac muscle, beta Myh7 (Pandya et al., 2006) MYL2: myosin, light chain 2, regulatory, cardiac, slow Myl2 (Kazmierczak et al., 2012) MYLK2: myosin light chain kinase 2 Mykl2 (Zhi et al., 2005) MYOZ2: myozenin 2 Myoz2 (Ruggiero et al., 2013) PRKAG2: protein kinase, AMP-activated, gamma 2 non- Prkag2 (Arad et al., 2003) catalytic subunit TCAP: titin-cap Tcap (Markert et al., 2010) TNNI3: troponin I type 3 (cardiac) Tnni3 (Wang et al., 2012) TNNT2: troponin T type 2 (cardiac) Tnnt2 (Tardiff et al., 1998) TTN: titin Ttn (Gramlich et al., 2009) VCL: vinculin Vcl (Zemljic-Harpf et al., 2007) The bolded genes are the most common mutations causing HCM 152

deficit in strain and twist seen at an early age. In addition, there has been a gene dose-

dependent effect seen in humans (Olivotto et al., 2008; Richard et al., 2000) and mice

(Desjardins et al., 2012), suggesting that a similar effect will be seen in the proposed

study.

POTENTIAL CONCERNS

There is some evidence to suggest that stratifying the genetic etiology of HCM

is not clinically beneficial (Landstrom et al., 2010b). It was shown that myofilament-

positive HCM patients had a greater probability of LV dysfunction than those with

myofilament-negative HCM, however there was no difference in dysfunction between

MyBPC3 thick or thin filament affected individuals(Olivotto et al., 2008). Although

this study did not detect differences in function between two myofilament mutations,

I believe that a more comprehensive evaluation of several myofilament proteins will

generate differences in outcome and provide data on phenotype-genotype

relationships.

2) Predictive value of mechanical dysfunction in preclinical HCM

RATIONALE

The introduction of commercial genetic testing has expanded the recognition

of patients with HCM. However, it can be difficult to manage the gene-positive and

phenotype-negative patients. It is unclear if these patients would benefit from

treatment or even close monitoring. The data is not clear if gene-positive phenotype-

negative individuals are at an increased risk for sudden death or disease progression 153

(Maron et al., 2011a). There is a need for better diagnostic technology and long term

follow-up in this patient population.

The next step of this investigation is to translate the findings from our small

animal studies to a clinical trial. We will conduct a small prospective cohort study in

HCM families to test if MRI-detected mechanical dysfunction is superior to

echocardiogram-identified cardiac hypertrophy in pinpointing early and clinically

significant pathology. We first established that there are reduced mechanical indices

of torsion, twist, and strain apparent in a mouse model of HCM (Desjardins et al.,

2012). Furthermore, the mechanical dysfunction was apparent before the development

of overt cardiac hypertrophy, implicating altered mechanical function as the first link

of the development of pathological hypertrophy and cardiac disease.

EXPERIMENTAL STRATEGY

The future directions of this aim will require several phases. I would begin by obtaining Institutional Review Board (IRB) approval at my institution for a

prospective cohort clinical study. I would enroll the children (ages 12-18) of families

identified with HCM that are already being evaluated with annual echocardiograms,

but do not yet exhibit overt LVH (i.e. genotype-positive phenotype-negative). Most

phenotypes of HCM begin to emerge in young adults during accelerated adolescent

growth, which is typically complete by age 18 (Maron et al., 1986). My exclusion

criteria would include pregnancy, prior myocardial infarction, prior septal myectomy

or alcohol septal ablation, incessant ventricular arrhythmias, known cardiovascular

disease, hypertension, substance abuse, standard MRI exclusion criteria including; 154

claustrophobia, ferromagnetic material in the body (foreign body or implanted), and inability to lie still for one hour or more. Once enrolled, the subjects would undergo

DENSE MRI on the same timing schedule as the standard of care echocardiography

study. After imaging is completed, subjects will be followed for cardiovascular

outcomes.

In addition to the conventional markers of cardiac function, the LV strain, twist, and torsion would be determined for each subject at every time point. The primary outcome to be studied in this clinical trial would be whether DENSE MRI can predict defects in cardiovascular function prior to standard echocardiography.

This information would be correlated with the incidence of cardiac death, aborted sudden cardiac death, heart transplantation, left ventricular assist device placement, all-cause mortality, ventricular tachyarrhythmias, ventricular fibrillation or sustained ventricular tachycardia, hospitalization for heart failure, atrial fibrillation, and stroke.

In particular, I would focus on twist/torsion as markers of dysfunction. In our study of cMyBPC+/- mice, we detected a 15% reduction in net twist, and a 13% decrease in

maximal torsion (Desjardins et al., 2012). Prior work has established that LV torsion

is similar between mice and humans, and thus represents a uniform measure of LV

function across species (Henson et al., 2000). In order to detect a 5% decrease in net

twist angle with a power of 90% and type I error rate less than 5%, the necessary

sample size must be n=71.

INTERPRETATION

I anticipate that DENSE MRI would be more sensitive than standard

echocardiography in detecting early myocardial dysfunction in young adults with 155

HCM. Since LV torsion is a consistent marker of cardiovascular function between

mice and men (Henson et al., 2000), I expect that our findings of decreased LV torsion in a mouse model of HCM would translate clinically.

POTENTIAL CONCERNS

The proposed study requires a long period of follow-up with subjects and may

require several years to enroll a sufficient subject population. There may be difficulty

ensuring that subjects continue to follow with MRI for several years. In addition, the

prevalence of disease progression has not yet been established in the genotype-positive

phenotype-negative population with standard echocardiography. We may need to

adjust the sample size of the study if data emerges regarding the incidence of disease

progression in phenotype-negative individuals. There are several clinical trials

currently underway investigating the diagnosis (markers of fibrosis, echocardiography

vs CMR) and treatment (diltiazem) of preclinical HCM.

Table 6.3: Hypertrophic cardiomyopathy vs physiologic athletic heart hypertrophy.

Adapted from Maron, B.J., and Maron, M.S. (2013). Hypertrophic cardiomyopathy. Lancet 381, 242– 255.

156

Lastly, a potential cofounder of the study is if there is also physiologic

hypertrophy from athletic training (Table 6.3,(Maron and Maron, 2013) ). We have

previously shown in a mouse model of physiologic hypertrophy that there are

alterations from the normal LV strain pattern (Montano et al., 2013). There are clinical

trials dedicated to studying the cardiovascular response to athletic training in patients

with known HCM mutations, particularly because HCM is the most common cause of sudden death in trained competitive athletes (Maron et al., 1996b). The target population in the proposed study is 12-18 years old, so there may not be many subjects with intense athletic training at this point in their lives. Although the physiologic hypertrophy may add complexity to data interpretation, it will be important to characterize myocardial function in this population due to the concern for sudden death.

3) Functional consequence of cerebral ischemia in mdx mice

RATIONALE

The blood-brain barrier (BBB) ensures the proper physical and metabolic

environment for brain function. Endothelial cells lining the vasculature of the brain

are joined by tight junctions in order to prevent free diffusion of large substances from

the blood into the brain parenchyma (Brightman and Reese, 1969; Reese and

Karnovsky, 1967). Although the water content of the brain has been studied for

decades (BERING, 1954; Bradbury et al., 1972; EDSTROM et al., 1961), the exact mechanism by which water passes through the BBB remained elusive. Discovery of the water channel, aquaporin 4 (AQP4), has contributed greatly to the understanding

of water movement in the brain (Francesca and Rezzani, 2010; Nicchia et al., 2004; 157

Nico et al., 2001; Zador et al., 2009). In mice, AQP4 plays a prominent role in both

the formation and removal of edema (cytotoxic and vasogenic respectively) following

brain injury, such as in stroke or tumor formation (Hirt et al., 2009; Kaur et al., 2006;

Meng et al., 2004; Promeneur et al., 2013; Saadoun et al., 2002; Tait et al., 2010; Tang

et al., 2010; Tourdias et al., 2011; Vajda et al., 2002a; Verkman et al., 2006; Vizuete et

al., 1999; Yang et al., 2008; Zeynalov et al., 2008). The anchoring of AQP4 by the

dystrophin-glycoprotein complex (DGC) is vital for its proper function in the brain

(Adams et al., 2008; Amiry-Moghaddam et al., 2004; Bragg et al., 2006; Nico et al.,

2005; Perronnet and Vaillend, 2010; Vajda et al., 2002b; Waite et al., 2012). Thus,

dystrophin may be indispensable in physiologic blood brain barrier maintenance, and

perturbation of dystrophin could be predicted to result in pathologic dysregulation of

the blood brain barrier.

The role of dystrophin in maintaining the BBB in vivo has not yet been

described, despite the clinical benefit in understanding the pathophysiology of stroke in dystrophinopathy. First, dystrophinopathies are associated with cardiomyopathy and arrhythmias and therefore represent an increased risk of ischemic stroke in these patients (Atsumi et al., 2004; Finsterer, 2012; Finsterer and Stöllberger, 2010; Finsterer et al., 2012; Hanajima and Kawai, 1996b; Tsakadze et al., 2011). Secondly, as the life expectancy of DMD patients increases with improved therapies, developing an understanding of the neurologic manifestations of the dystrophin-null phenotype will lead to better patient management when symptoms emerge. A comprehensive assessment of edema development as a result of acute stroke in mice lacking 158

dystrophin using powerful MR techniques may represent a first step in targeting

therapies towards preventing or treating stroke in dystrophinopathy patients.

Future work of this project would test the function of dystrophin in preventing

vasogenic edema following ischemic insult. Ischemic stroke induced by middle

cerebral artery occlusion (MCAO) demonstrates two distinct phases of edema with

unique pathologies (Simard et al., 2007). Ischemic stroke is first characterized by

cytotoxic edema, which is characterized as the swelling of cells as a result of movement

of osmotically active molecules to the intracellular space. The lack of blood flow during ischemia leads to decline in ATP synthesis, which impairs Na+/K+ ATPase function. As a result, cellular accumulation of Na+, Ca2+, and lactate drives water into

the cell and produces significant cell swelling. Vasogenic edema causes further cell

damage at later time points due to the breakdown of the BBB (Simard et al., 2007).

Several mechanisms have been described to alter the integrity of the BBB, including

reverse pinocytosis (Castejón, 1984) and disruption of Ca2+ signaling (Brown and

Davis, 2002). Although dystrophin is an integral part of the cytoskeletal network that

supports regulation of water transport at the blood brain barrier, the function of

dystrophin in these edematous conditions has not yet been explored. My hypothesis is

that a lack of dystrophin will be detrimental during ischemic injury by exacerbating

the formation of vasogenic edema.

EXPERIMENTAL STRATEGY

A permanent MCAO protocol will be implemented to induce cerebral ischemia

in mice (Longa et al., 1989). In summary, mice will be anesthetized with 2% isoflurane

via nosecone and their temperature maintained with a heating pad. Ischemia will be 159

induced by inserting and tying a monofilament distal to the carotid bifurcation. The

cervical wound will be closed with sutures and the animal will be allowed to recover.

In the sham surgery, the carotid artery will be maneuvered as in MCAO, however the

arteriotomy and filament ligation will not be executed. Studies will be performed on

the following groups of 3 month old mice (n=10 for each group):

i. Sham-operated C57/BL6 control

ii. MCAO C57/BL6 control

iii. Sham-operated mdx

iv. MCAO mdx

The infarct size as a result of MCAO will be quantified in each group using a

T2-weighted image 24 hours after the surgery. A DWI protocol will be implemented

and images will be acquired at the following time points after MCAO/Sham surgery:

1, 4, 8, 24, and 72 hours. In addition, a neurologic deficit score adapted from (Hunter

et al., 2000) will be assigned 24 hours after MCAO by an observer blinded to the

genotype and surgical procedure. Once imaging studies are completed, brains will be fixed for histologic analysis (n=3 each group) by transcardiac perfusion of paraformaldehyde (4% in 0.1M sodium phosphate buffer), immediately stored at 4°C in fixative overnight and paraffin embedded for sectioning. Slices will be stained with hematoxylin/eosin, 2,3,5-triphenyltetrazolium chloride (TTC), or glial fibrillary acid protein (GFAP) using the avidin-biotin peroxidase complex method (Garcia et al.,

1993) to assess morphology and infarct size in each group. The infarct size (as measured by both histology and MRI) and mean ADC will be statistically compared 160

between the control and mdx groups using standard ANOVA methods with the

appropriate adjustments.

INTERPRETATION

I anticipate that the mdx group will experience a significantly worse prognosis

following MCAO as measured by diffusion MRI, histology, and neurologic deficit

scoring. I expect to find an increase in cerebral edema and slower resolution of fluid

accumulation in the mdx group following an ischemic insult. Together, the results would suggest that dystrophin-mediated BBB integrity is an essential component of the tissue response to ischemic injury in the mouse.

POTENTIAL CONCERNS

Defining the pathophysiologic role of dystrophin in ischemic stroke is a complex task. Dystrophin is necessary to anchor AQP4 to the astrocytic foot processes

(Nicchia et al., 2008a, 2008b; Vajda et al., 2002b). Aqp4 homozygous null mice do not

exhibit a phenotype under baseline conditions and have similar brain water content as

compared to wild-type in the absence of pathology (Tait et al., 2010). Under conditions

of cytotoxic edema, either by cerebral ischemia or water intoxication, the absence of

AQP4 is protective and reduces injury in both pathologic models (Manley et al., 2000).

In contrast, AQP4 deletion worsens cerebral swelling and neurologic function

following vasogenic edema due to the inability to remove water from the tissue

(Papadopoulos et al., 2004). 161

We described a decrease in cerebral ADC of mdx mice that likely represents

cellular edema as a result of BBB disruption in our prior work (Goodnough et al.,

2014). While it has been shown that AQP4 facilitates the clearance of vasogenic edema

as a result of BBB disruption (Papadopoulos et al., 2004), the lack of properly

functioning AQP4channels at the BBB in mdx mice (Adams et al., 2008; Frigeri et al.,

2002) may prevent the removal of water from the cerebrum and therefore worsen

cellular edema.

Ischemic stroke is first characterized by cytotoxic edema, and vasogenic edema

causes further cell damage at later time points (Simard et al., 2007). I anticipate that

there is vasogenic edema in mdx mice prior to MCAO that will exacerbate and

accelerate the ischemic injury since the homeostatic fluid balance is already

compromised. This result would suggest the dystrophin has a role in maintaining the

BBB outside of its role for anchoring proteins for AQP4. However, if I observe an

improvement in edema formation (particularly in the acute phase) after MCAO, it

would imply that the main function of dystrophin is the anchoring of AQP4 in the

astrocytic endfeet. In either outcome, the interpretation of dystrophin’s role in

maintaining the BBB during acute stroke will be novel.

4) Functional consequences of hypoperfusion in DMD

RATIONALE

It was observed early on that DMD boys have lower IQs than their age-

matched controls (Karagan, 1979); in fact, Duchenne described cognitive impairment

in his initial description of the disease (Duchenne, 1868). Similarly, there is also 162

evidence of cognitive impairment in the mdx murine model, particularly passive

avoidance learning (Muntoni et al., 1991) and newly learned information (Vaillend et

al., 1995). There is evidence of cortical atrophy and ventricular dilation in DMD

patients (Septien et al., 1991; Yoshioka et al., 1980), however there has not been a

study to establish the link between cognitive impairment and gross cerebral

abnormalities. In addition, there is evidence of hypo-metabolism in DMD (Bresolin et

al., 1994b; Tracey et al., 1995, 1996b) where there is a significant increase in the

cerebral Pi/ATP ratio in DMD patients and mdx mice.

Our prior work suggests that cerebral perfusion is compromised in mdx mice

(Goodnough et al., 2014). Although it has not yet been studied in the brain, there is

also evidence of vascular dysfunction in the skeletal muscle of DMD. Functional

muscle ischemia has been shown in mdx mice, which is due to a deficiency of nNOS

deficiency and thus contraction-induced modulation of sympathetic vasoconstriction

(Sander et al., 2000). Previously, enhanced arteriogenesis was observed in the

hindlimbs of mdx mice (Straino et al., 2004). Similarly, an increase of VEGF and

VEGFR2 expression results from deletion of dystrophin via enhanced expression of

matrix-metalloproteinase (MMP)-2 and -9 (Nico et al., 2006). In addition to

angiogenesis, increased VEGF activity may also contribute towards an opening of

tight junctions at the BBB and enhanced vascular permeability (Schoch et al., 2002;

Zhang et al., 2002).

EXPERIMENTAL STRATEGY

The aim of this study is to correlate decreased perfusion in mdx mice with cognitive functional deficits. I will correlate deficits in cerebral perfusion and 163

Figure 6.1: Cognitive testing for the mouse.

Adapted from Rodriguiz, R.M., and Wetsel, W.C. (2006). Assessments of Cognitive Deficits in Mutant Mice. In Animal Models of Cognitive Impairment, E.D. Levin, and J.J. Buccafusco, eds. (Boca Raton (FL): CRC Press),.

metabolism with the onset cognitive function over an extended time course in mdx

mice compared to wild-type (WT) controls. Mdx mice will be compared to WT mice

at (n=8 each group): 2 and 4 weeks, 2, 3, 4, 6, 8 and 10 months. At each time point,

the cerebral perfusion will be measured as described in (Gao et al., 2014), as well as

the cerebral Pi/ATP ratio using magnetic resonance spectroscopy (Tracey et al.,

1996b). Prior to beginning the study, I will complete a full battery of cognitive testing

(Figure 6.1, (Rodriguiz and Wetsel, 2006) on 10 month mdx mice (n=5) to determine

which paradigm of cognitive exams I will choose for the course of the study. Although

it has been shown that passive avoidance is diminished in mdx (Muntoni et al., 1991), a broad range of cognitive abilities and functions remain untested in mdx mice.

INTERPRETATION 164

I anticipate a gradual decline in perfusion, metabolism, and cognition in the mdx mice as compared to control. We observed a decrease in cerebral perfusion from

2 to 10 months in mdx mice, and the mean ADC was already decreased at 2 months

in mdx compared to WT (Goodnough et al., 2014). It is unclear if defects in perfusion will precede hypometabolism or vice versa. I predict that perfusion and metabolism are unrelated and will demonstrate independent time courses. Since adequate perfusion and metabolism are necessary for cognition, it is likely that their effects will be additive on cognitive function testing.

165

CHAPTER 6 REFERENCES Abbott, N.J., and Friedman, A. (2012). Overview and introduction: The blood–brain barrier in health and disease. Epilepsia 53, 1–6.