View Highlighting the Importance of a Flexible and Multidisciplinary Approach,” J

INVESTIGATING CARDIAC METABOLISM IN BARTH SYNDROME

USING INDUCED PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES

ERICA M. FATICA

Bachelor of Science in Pharmaceutical Science

Cleveland State University

May 2014

Submitted in partial fulfillment of requirements for the degree

DOCTOR OF PHILOSOPHY IN CLINICAL-BIOANALYTICAL CHEMISTRY

at the

CLEVELAND STATE UNIVERSITY

May 2019

We hereby approve this dissertation For Erica Marie Fatica Candidate for the Doctor of Philosophy in Clinical-Bioanalytical Chemistry degree for the Department of Chemistry And CLEVELAND STATE UNIVERSITY’S College of Graduate Studies by

Committee Chairperson, Yana Sandlers Department of Chemistry, April 18th, 2019

Committee Member, Dr. David Anderson Department of Chemistry, April 18th, 2019

Committee Member, Dr. Michael Kalafatis Department of Chemistry, April 18th, 2019

Committee Member, Dr. Christine Moravec Department of Biological, Geological and Environmental Sciences, April 18th, 2019

Committee Member, Dr. Aimin Zhou Department of Chemistry, April 18th, 2019

April 18th, 2019 Date of Defense

Dedicated to my parents/biggest fans

In loving memory of Angela Grandillo, whose dedication to never stop learning will forever inspire me

ACKNOWLEDGEMENTS

This degree would not have been possible without the amazing people who have supported me along the way. First, I would like to sincerely thank my advisor, Dr. Yana

Sandlers, for taking me into her research group and for being the most supportive, understanding, and wonderful mentor I could have ever asked for. Thank you for all the opportunities you gave me which allowed me to grow as a scientist and as an individual.

Thank you for always believing in me and for encouraging me to have more confidence in myself. I would not have been able to go this far without you.

I would also like to thank my truly amazing dissertation committee members.

Without Dr. David Anderson’s guidance during my undergraduate work, I would not have stayed at Cleveland State to continue my studies. Thank you for your continued support, care, and guidance in all things clinical chemistry. I am extremely grateful to have had such a wonderful and encouraging female role model in Dr. Christine Moravec.

I am thankful to Dr. Aimin Zhou, who has always been the most kind person you could hope to know. Thank you to Dr. Michael Kalafatis for always having my back and supporting me.

A special thank you to Dr. Vania DePaoli for always being there for me in every aspect of my life and for looking out for me. I will miss our chats in your office. Also, thanks to Dr. DiBello for being so sweet and supportive, encouraging and inspiring. You are a truly great educator and person, and I hope to carry those qualities with me.

I also would like to acknowledge my labmates, especially Rohan Shah for being with me every step of the way (and buying us cookies) and Igor Radzikh, for being my right-hand man. Igor, without you, many things would not have been possible. Thank you

also to Ryan, Jared, and Jill for helping me along the way. Many thanks to SooYeon

Kang, for all her hard work and assistance.

Thank you to my mom, Elia Iafelice, who literally kept me alive during these five years. Without you, I probably would not have eaten for five years. Thank you for putting up with me and for reminding me to take care of myself sometimes. Thank you to my dad, Ron, for always being interested in my work and for helping me troubleshoot technological things. Thank you to my brother, Marco, for helping me understand how to do some of the complicated math stuff that I used for my research project. Your and dad’s ability to understand and work with completely new information never ceases to amaze me.

Also thank you to Liz Grandillo, the woman who looks like she could be my mom and acts like my sister, gives me cute hand-me down clothes, lets me visit her in New

York to raid her closet, and does not understand anything about cardiac function, except for having occasional chest pains (and thanks for dictating your own acknowledgement).

To my best friend, Alana, and our daily chats. Thanks for escorting me in and out of school with your ridiculous stories. Thank you, Alwilleed, for supporting me in every way, reminding me to stay tough, and for bringing me so much joy.

To everyone else who helped me over the past five years, especially my classmates, your moral support and love helped carry me through. Your presence in my life made a difference and I am truly grateful.

INVESTIGATING CARDIAC METABOLISM IN BARTH SYNDROME

USING INDUCED PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES

ERICA M. FATICA

ABSTRACT

Barth syndrome (BTHS) is an X-linked genetic disorder characterized by cardiomyopathy, neutropenia, skeletal muscle weakness, and 3-methylglutaconic aciduria

(Sandlers, et al., 2016). The mortality rate of BTHS patients is high during infancy and childhood due to sudden cardiac death. Despite the severity of this disease, there is a lack of targeted therapeutics which can be used to ameliorate symptoms and prolong the lives of BTHS patients. A major obstacle for the discovery of new therapeutic targets is poor understanding of the mechanisms of cardiac pathogenesis and downstream metabolic effects. To overcome this barrier, we developed a model of BTHS, using human-induced pluripotent stem cells (iPSCs), with or without the BTHS-causative TAZ mutation, to produce functional cardiomyocytes (iPS-CMs). iPS-CMs recapitulate the human donor genotype, reproducing the complex metabolic conditions of the hearts of affected individuals, and permitting investigation of molecular and metabolic mechanisms. We further applied stable isotope-labeled energy substrates and mass spectrometric analyses to our TAZ-mutant iPS-CM model to dynamically trace the fates of substrates through metabolic pathways, including fatty acid oxidation, glucose oxidation, and select anaplerotic pathways into the citric acid cycle (CAC). The studies herein provided novel insight into downstream metabolic differences in energy substrate metabolism between

vi control and TAZ-mutant iPS-CMs, revealing alterations in several pathways which can be further explored to uncover potential therapeutic targets for BTHS patients.

vii TABLE OF CONTENTS

Page ABSTRACT………………………………………………………………………...……vi

LIST OF FIGURES……………………………………………………………………. xxi

CHAPTER

I. INTRODUCTION……………………………………………………………... 1

1.1 Barth Syndrome……………………………………………..…..…… 1

1.2 Tafazzin and Cardiolipin…………………………………...... 2

1.3 Current Treatment Strategies for Barth Syndrome Patients………..... 4

1.4 Dilated Cardiomyopathy………………………………….……...…... 6

1.5 Cardiac Metabolism…………………………………………...….….. 6

1.6 Cardiac Metabolism in Pathophysiological Conditions……...…….… 9

1.7 Cardiac Anaplerosis……………………………………...……….… 12

1.8 Therapeutic Strategies for Modulating Cardiac Metabolism…...…... 15

1.9 Induced Pluripotent Stem Cells………………………..………….... 16

1.10 Stable Isotope Metabolomics………………………...……………. 19

1.11 References………………………………………...……………….. 22

II. ESTABLISHING A MODEL OF BARTH SYNDROME USING INDUCED

PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES…..……. 29

2.1 Abstract…………………………………………...…………….…... 29

2.2 Introduction…………………………………...…………….………. 30

2.3 Methods…………………………………...………………….…...… 35

2.4 Results………………………………...……….……….…...... 39

2.5 Discussion……………………………...……………………….…... 44

viii 2.6 References…………………………..…………………………....… 51

III. STABLE ISOTOPE TRACING STUDY IN INDUCED PLURIPOTENT

STEM CELL-DERIVED CARDIOMYOCYTES TO INVESTIGATE

METABOLIC ALTERATIONS IN BARTH SYNDROME………………. 51

3.1 Abstract…………………………………………...…………….…... 51

3.2 Introduction…………………………………..……….……………. 52

3.3 Methods…………………………………..……………………....… 54

3.4 Results………………………………..……………….…...... 60

3.5 Discussion………………………...……………………………….... 72

3.6 References……………………...………………………...... … 77

IV. ANAPLEROTIC PATHWAYS OF THE CITRIC ACID CYCLE IN TAZ-

MUTANT INDUCED PLURIPOTENT STEM CELL-DERIVED

CARDIOMYOCYTES…………..……………………………………...…. 81

4.1 Abstract…………………..………………………….……………... 81

4.2 Introduction…………..……………………………………….……. 82

4.3 Materials and Methods…………..………….…………...…………. 86

4.4 Results………………………..……………………….…...... 89

4.5 Discussion………………..………………………………….……... 99

4.6 References…………..………………………………….……….… 105

V. CALCIUM CHANNEL EXPRESSION IN TAZ-MUTANT INDUCED

PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES..…….. 109

5.1 Abstract………………………………………..………….………. 109

5.2 Introduction to Calcium Channel Proteins…………..….……….... 110

ix 5.3 Materials and Methods………………………………………….… 114

5.4 Results……………………………………..………….…………... 116

5.5 Discussion…………………………..…………………………….. 122

5.6 References……………………..………………………………..… 128

VI. GENERATION OF INDUCED PLURIPOTENT STEM CELL-DERIVED

CARDIOMYOCYTES IN 3D ON A 384-MICROPILLAR PLATE……... 133

6.1 Abstract………………………………..…...…………...…………. 133

6.2 Introduction……………………………....………………………... 134

6.3 Methods…………………………...... ……...………….…………... 138

6.4 Results………………………………………….…….……………. 143

6.5 Discussion……………………………………….………………… 155

6.6 References………………………………….………….…………... 161

VII. CONCLUSION ……...…………………………………………...... 164

7.1 References…………………………...………………….…………. 166

APPENDICES

A. CORRECTION OF MASS ISOTOPOMER DISTRIBUTIONS FOR

NATURAL ISOTOPIC ABUNDANCE……………..………………….... 169

B. GAS CHROMATOGRAPHY-MASS SPECTROMETRY METHODS…. 177

x LIST OF FIGURES

Figure Page

1.1 Cardiac Energy Substrates ...... 8

1.2 ATP Yield and Oxygen Consumption...... 11

1.3 Anaplerotic Pathways of the Citric Acid Cycle...... 14

1.4 Generation of Induced Pluripotent Stem Cells from Adult Somatic Cells...... 18

1.5 Mass Isotopomers of a 4-Carbon Molecule ...... 21

2.1 Cardiogenesis via Temporal Modulation of the Wnt Pathway...... 34

2.2 Small Molecule Differentiation Workflow...... 37

2.3 Evaluation of iPSCs...... 40

2.4 Immunocytochemistry of iPS-CMs ...... 42

2.5 Immunocytochemistry of iPS-CMs...... 43

3.1 Glucose and Palmitate Consumption...... 61

3.2 Mass Isotopomer Distributions...... 63

3.3 M2 Labeling of CAC Intermediates from 13C-Palmitate...... 65

3.4 Levels of CAC Intermediates...... 67

3.5 Fatty Acid Binding Proteins...... 69

3.6 Fatty Acids and Acyl Carnitines...... 71

4.1 Major Anaplerotic Pathways of the CAC...... 85

4.2 Glutamine as a Carbon Source for the Citric Acid Cycle...... 90

4.3 Mass Isotopomer Distributions of Citric Acid Cycle Intermediates and Aspartate. 92

4.4 M2 and M3 Enrichment Patterns...... 94

4.5 Arginine and Proline...... 96

xi 4.6 Levels of Anaplerotic Amino Acids in iPS-CMs...... 98

5.1 Calcium Handling ...... 112

5.2 α-Actinin Immunostaining...... 117

5.3 SERCA2a2, RYR2, and NCX mRNA...... 119

5.4 SERCA2a2 Protein Expression...... 121

6.1 Renderings of the 384-Pillar Plate...... 137

6.2 Structure of MitoQ ...... 141

6.3 iPSC Spheroid Formation on 384-Pillar Plates ...... 145

6.4 Immunostaining of 3D iPS-CMs...... 147

6.5 ATP Levels with MitoQ Treatment ...... 149

6.6 Untargeted Analysis of Metabolites from 3D iPS-CMs ...... 151

6.7 Measurement of Acyl Carnitines by LC-MS/MS ...... 153

6.8 Detection of CAC Intermediates by LC-MS/MS...... 154

6.9 Determining Natural Abundance...... 171

A. 1 Mass Isotopomers of Malate...... 175

xii

CHAPTER I

INTRODUCTION

1.1 Barth Syndrome

Barth syndrome (BTHS) is an X-linked, multisystem genetic disorder characterized by cardiomyopathy, neutropenia, skeletal muscle weakness, growth delay, lactic acidosis, and 3-methylglutaconic aciduria [1]. BTHS is associated with a high rate of mortality during infancy and childhood, usually due either to heart failure or overwhelming bacterial infection. Studies have shown that around 70% of BTHS patients were diagnosed after they had died. These retrospectively-diagnosed patients were only tested for the disease after a brother or male relative was diagnosed with BTHS [2]. In contrast, the mortality rate is 10% for BTHS-affected individuals who are diagnosed prospectively and begin a treatment regimen before the onset of severe symptoms, illustrating how critical it is to diagnose and treat this disease as early as possible in the patient’s life [2]. Despite the importance of making an early diagnosis, there is an average delay of about 3 years between the time that a child first presents with symptoms to the time a diagnosis is made [3].

Part of the reason BTHS is difficult to diagnose is due in part to lack of awareness of this disease and its clinical presentation. BTHS was first described in 1983 by Dr.

Peter Barth, who noted a triad of symptoms including cardiomyopathy, skeletal muscle weakness, and neutropenia [4]. During clinical workup, the cardiac features observed in

BTHS patients are often misdiagnosed as idiopathic dilated cardiomyopathy, or bacterial- induced cardiomyopathy [5][6]. Additionally, BTHS has been identified as a previously unknown cause of recurrent miscarriage and stillbirth [7]. Furthermore, BTHS has a notoriously wide and diverse phenotype. The severity of cardiomyopathy is highly variable, with 6% of BTHS patients showing no signs of cardiomyopathy at all [3]. The same is true of neutropenia, which may be persistent, intermittent, or not present at initial presentation. Thus, although the prevalence of BTHS is estimated to be 1/300,000 live births, recent work suggests that the disease is under-diagnosed [1].

1.2 Tafazzin and Cardiolipin

BTHS is caused exclusively by mutations in the TAZ gene, located at chromosome Xq28, which codes for the mitochondrial phospholipid acyltransferase, tafazzin [6][8]. Gene sequencing has revealed over 120 mutations in the TAZ gene, including missense mutations, deletions, insertions, and in some cases, large exon or whole gene deletions [9]. Gene sequencing of the 11 exons of the TAZ gene can be performed to confirm a suspected diagnosis of BTHS. Interestingly, genotype-phenotype studies have not revealed any correlation between a specific mutation and severity of symptoms, and intrafamilial variability is common, even for relatives with the same TAZ mutation. This was perhaps best demonstrated in a case study published by Ronvelia et al., in which a patient was diagnosed with BTHS at age 45 after left ventricular noncompaction was discovered [10]. The patient had a history of muscle weakness, but

2 did not display other characteristic symptoms of BTHS, such as recurrent infection. The patient’s great nephew, carrying the same TAZ mutation, was diagnosed with failure to thrive, lactic acidosis, decreased feeding, and dilated cardiomyopathy with congestive heart failure by three months of age, eventually requiring cardiac transplantation by eleven months of age. The wide range of clinical presentations and lack of genotype- phenotype correlations continues to present a challenge for understanding the progression of the disease, giving accurate clinical prognoses, and developing effective treatments for

BTHS patients.

The functional role of tafazzin is to catalyze the transacylation of cardiolipin, one of the principle structural phospholipids of the inner mitochondrial membrane which is highly expressed in cardiac and skeletal muscle [11]. Unlike most phospholipids which are composed of two hydrophobic acyl chains and one hydrophilic head group, cardiolipin has a unique structure consisting of four acyl chains and two head groups, which give the molecule a conical shape [12]. The predominant cardiolipin species in oxidative tissues is tetralinoleoyl cardiolipin (18:2 CL) [13]. Conversely, unremodeled cardiolipin, known as monolysocardiolipin (MLCL), has three saturated acyl chains and one hydroxyl group. The ratio of monolysocardiolipin to tetralinoleoyl cardiolipin

(MLCL/18:2 CL) is a highly sensitive and specific measurement which can be used in the diagnosis of BTHS as an alternative to genetic testing [14].

Cardiolipin plays a critical role in mitochondrial energetics and adenosine triphosphate (ATP) production, as the inner mitochondrial membrane hosts the fundamental machinery of the electron transport chain and interacts with proteins integral to oxidative phosphorylation. Cardiolipin is also involved in various phases of apoptosis

3 through its interaction with cell death-inducing proteins such as cytochrome c, as well as mitochondrial dynamics such as fission, fusion, and mitophagy [15][16]. Defective or absent tafazzin results in serious alterations of cardiolipin content and chain composition.

The ultimate manifestation of the many possible TAZ gene mutations is excessive accumulation of nascent cardiolipin species and a decline in tetralinoleoyl cardiolipin.

The resulting abnormalities in mitochondrial structure and function are central to BTHS, but have also been implicated in diabetes, heart failure, non-alcoholic fatty liver disease, neurodegeneration, and aging [16][17][18]. Therefore, although the TAZ mutation itself is rare, alterations in biochemical pathways of mitochondrial energy metabolism as a result of compromised tetralinoleoyl cardiolipin content may be paralleled in major clinical pathologies.

1.3 Current Treatment Strategies for Barth Syndrome Patients

There is currently no definitive treatment for BTHS. Supportive management of the cardiac features of BTHS includes various standard treatments of heart failure, such as β-blockers, angiotensin-converting enzyme inhibitors, diuretics, and cardiac glycosides

[19]. There are no comprehensive studies available that evaluate the effectiveness of these drugs in BTHS patients, thus they are prescribed on a case-by-case basis, guided by case reports and expert clinical opinions [19][20]. These medications seem generally effective in prolonging the lives of BTHS patients beyond childhood, but no medication has shown to be particularly advantageous. In fact, in some instances, cardiac status managed by these therapeutics had been stable, but suddenly declined after months of therapy for unidentified reasons [16]. In this scenario, BTHS patients often require

4 cardiac transplantation. About 70% of BTHS patients who receive cardiac transplantation do so in the first few years of life [9].

Besides heart failure medications, dietary supplementation has been investigated as a means to rescue the cardiolipin profile of BTHS patients. Supplementation with either L-carnitine, panthotenic acid, or arginine have been given, as these molecules have been implicated as metabolic derangements in BTHS. However, these compounds have demonstrated minimal benefits in vitro, and little published data is available in vivo.

Several studies, including a landmark publication by Wang et al., report that supplementation with linoleic acid may benefit patients based on in vitro data [17].

Linoleic acid, a precursor of cardiolipin synthesis, was able to restore sarcomere structure in an induced pluripotent stem cell-derived cardiomyocyte model [17], and reestablished cardiolipin levels in fibroblasts obtained from BTHS patients [21]. Whether linoleic acid would be effective in mitigating the symptoms of BTHS patients remains to be evaluated in vivo.

In an innovative attempt to rescue the tetralinoleoyl cardiolipin profile, Makaryan et al. succeeded in delivering cardiolipin to cells using cardiolipin nanodisks, which successfully attenuated apoptosis in an HL60 myeloid progenitor cell TAZ-knockdown model [22]. Ikon et. al demonstrated that this in vitro effect was not reproduced in vivo, and thus reported that restoring the cardiolipin profile using cardiolipin nanoparticles is not recommended as a viable option for treating BTHS patients [23]. Thus, much work still needs to be done to identify a specific and effective treatment for BTHS patients.

1.4 Dilated Cardiomyopathy

Dilated cardiomyopathy (DCM) is one of the most prominent and severe features of BTHS, manifesting in the first year of life for 70% of patients, and often necessitating cardiac transplantation before the age of five [1]. DCM is a progressive heart disease associated with heart failure and risk of sudden death [24]. In the US alone, the cost burden of DCM management is estimated to be over 4 billion dollars annually [25]. DCM is characterized by thinning and stretching of the ventricular wall, reducing the heart’s ability to pump blood [26]. In the general population, 30-50% of patients with DCM have an inherited form of the disease. Inherited DCM can be caused by mutations in genes affecting a wide range of cardiac features, such as sarcomeric structure, nuclear proteins, ion channels, or cytoskeletal proteins [27]. Gene mutations which result in metabolic dysregulation, especially those which affect mitochondrial oxidative metabolism such as in BTHS, can also lead to the development of dilated cardiomyopathy [28].

Besides DCM, BTHS patients may also present with other cardiovascular complications including left ventricular non-compaction in 50% of patients, hypertrophic cardiomyopathy, long QT syndrome, endocardial fibroelastosis, and ventricular arrhythmia [9]. Risk of ventricular arrhythmia and sudden death in BTHS occur seemingly independent of the degree of DCM [16].

1.5 Cardiac Metabolism

The heart must meet an enormous energy demand to sustain contractile function.

Accordingly, the heart is highly adaptive and can utilize various fuel sources, including fatty acids, carbohydrates, amino acids, and ketone bodies to produce the required 6

6 kilograms of ATP per day (Figure 1-1) [29]. The healthy adult heart relies primarily on β- oxidation of fatty acids to produce about 70-90% of the required ATP [30]. The remaining 10-30% of ATP is generated mainly from glucose and lactate, with lesser amounts coming from ketone bodies and amino acids [31]. Mitochondria play a vital role in cardiac energy metabolism, serving as the site of production for 95% of the ATP made by the adult heart [31].

i. Glucose Utilization

Upon entry into the cell, glucose is rapidly and irreversibly phosphorylated to glucose-6-phosphate, a precursor for several key metabolic pathways. The major catabolic pathway is glycolysis, which takes place in the cytosol and results in the production of pyruvate, 2 molecules of NADH, and a net of 2 ATP. Under normal physiological conditions, glycolysis in the heart is only responsible for about 5% of ATP production [29]. As pyruvate is further oxidized in the mitochondria through the citric acid cycle (CAC), reducing equivalents NADH and FADH2 are generated, carrying electrons to the electron transport chain, where the bulk of ATP is generated (Figure 1-1

A).

Other pathways of glucose-6-phosphate metabolism include the pentose phosphate pathway (PPP), the hexosamine biosynthetic pathway (HBP), or glycogen synthesis. The

PPP generates important precursors to DNA and RNA synthesis, as well as the electron carrier, nicotinamide adenine dinucleotide phosphate (NADPH). NAPDH plays an important role in preventing oxidative stress, is necessary for fatty acid synthesis, and is needed for pyruvate carboxylation into malate by malic enzyme. The HBP is critical for the formation of uridine diphosphate N-acetyl-glucosamine (UDP-GlcNAc), which is

Figure 1-1: Cardiac Energy Substrates

Energy substrates in A) the healthy adult heart and B) in various pathophysiological conditions. Adapted from [29].

8 used for the post-translational modification of proteins. Alterations in either of these accessory pathways, therefore, may contribute to the development of cardiac disease.

ii. Fatty Acid Oxidation

Free fatty acids are taken up into the cell by various receptors, including fatty acid binding proteins (FABPs), fatty acid translocase (FAT/CD36), and fatty acid transport protein (FATP) [32]. Upon cell entrance, free fatty acids are esterified in the cytosol to fatty acyl-CoAs. Short and medium chain fatty acids can cross into the mitochondria, while long-chain fatty acids need to be converted to acylcarnitines to cross into the mitochondrial matrix [33]. The enzyme carnitine palmitoyl transferase I esterifies the long-chain fatty acid to its corresponding acylcarnitine species, which can then enter the mitochondrial matrix. A second enzyme, carnitine palmitoyl transferase II, converts the long-chain acylcarnitine back to a fatty acyl-CoA. Once in the mitochondrial matrix, fatty acyl-CoAs undergo successive steps of β-oxidation to form acetyl-CoA which enters

CAC. During the process of β-oxidation, the reducing equivalents NADH and FADH2 are also formed.

1.6 Cardiac Metabolism in Pathophysiological Conditions

Decreased ATP levels are a biochemical hallmark of cardiac diseases. In many pathophysiological conditions such as ischemic heart failure, diabetic cardiomyopathy, idiopathic dilated cardiomyopathy, and various mitochondrial diseases, the oxidative capacity of mitochondria becomes decreased [34][35]. This is generally accompanied by the heart reverting to a glycolytic state, mimicking the energy metabolism of the fetal heart. In this scenario, fatty acid oxidation decreases, while the rate of glycolysis

9 increases, with subsequent uncoupling from complete glucose oxidation (Figure 1-1B)

[34][36][37].

Although complete oxidation of one molecule of palmitate yields 129 ATP compared to complete oxidation of one molecule of glucose which yields 36 ATP, glucose oxidation consumes less oxygen per ATP generated (Figure 1-2) [38]. Thus, the shift away from fatty acid metabolism may represent a protective mechanism to optimize cardiac efficiency, although whether this shift is an adaptive response or a causative factor in the development of heart failure is still not well understood [35][39].

A recent theory postulates that the DCM observed in BTHS manifests as a result of metabolic remodeling [40]. Absence of tafazzin activity results in cardiolipin molecular species heterogeneity, increased levels of monolysocardiolipin (MLCL), lower cardiolipin (CL) abundance, and a subsequent increase in the ratio of MLCL/CL [41]. In the mitochondria of skeletal muscle and cardiac tissue, these alterations in cardiolipin content perturb the inner membrane integrity, compromising electron transport chain structure and function and, subsequently, aerobic respiration. Decreased electron flow from reduced NADH ubiquinone oxidoreductase activity leads to a buildup of NADH in the matrix space, causing negative feedback inhibition of key CAC enzymes. As CAC activity slows, pyruvate generated by glycolysis is diverted to lactic acid, which purportedly reduces ATP availability for contractile work (Figure 1-1). The authors propose that when there is an increased demand for energy, the heart attempts to compensate for the lack of ATP by metabolic and structural remodeling, ultimately resulting in cardiomyopathy. The theory put forth by the authors is largely speculative, as little data were shown to support their claims. While an increase in NADH has been

ATP Yield Oxygen Consumed ATP/Oxygen Substrate (mol ATP/mol (mol ATP/mol (mol O/mol substrate) substrate) O) Glucose 36 12 3.0 Lactate 18 6 3.0 Palmitate 129 50 2.6 Pyruvate 15 6 2.5

Figure 1-2: ATP Yield and Oxygen Consumption.

Theoretical ATP yield and oxygen required to fully oxidize various carbon sources. Adapted from [38].

11 demonstrated, there is very little published evidence to date that support the hypothesis of

CAC enzyme inhibition. Furthermore, there is no evidence that pyruvate conversion to lactate occurs at a different metabolic rate, and if so, what the fate of lactic acid may be.

Therefore, an understanding of the metabolic consequences of the TAZ mutation in heart and muscle is warranted. Our work herein aims to elucidate metabolic and molecular mechanisms underlying the pathophysiology of BTHS.

1.7 Cardiac Anaplerosis

Besides fatty acid and glucose oxidation to acetyl-CoA, cardiomyocytes employ other metabolic conversions to generate CAC intermediates at various stages of the CAC.

These “refilling” processes are known as anaplerotic reactions. Besides playing a crucial role in normal cardiac physiology, anaplerosis also has implications in conditions which present with decreased ATP production, such as cardiomyopathy [3][4][5]. Despite the significance of anaplerosis in the heart, there is a wide gap in knowledge regarding the activity, regulatory mechanisms, and pathophysiological changes that occur in anaplerotic reactions due to lack of sensitive techniques for in vivo measurements and unavailability of models which fully recapitulate the metabolic complexity of the heart

[44].

The importance maintaining a balanced pool of CAC intermediates is illustrated by the flexibility of the cell to utilize numerous metabolites to replenish the CAC at various stages (Figure 1-3). Metabolites with anaplerotic activity include propionyl-CoA and its precursors (branched-chain amino acids, odd-chain fatty acids, C5 ketone bodies), a wide range of amino acids, and pyruvate [45][46]. The conversion of glutamine into α-

12 ketoglutarate is an anaplerotic pathway that is important in the kidney and proliferating cells [47][48]. The conversion of α-ketoglutarate to succinate produces one molecule of

ATP and NADH via substrate-level phosphorylation without contributing to intracellular acidification [49][50]. In normal cardiac metabolism, this pathway has not been shown to play a significant role [51]. In various heart conditions, however, it has been established that glutamine has cardioprotective benefits, although it is unclear whether this is through direct contributions to the CAC or alternate, non-energy producing pathways such as the pentose phosphate pathway or the hexosamine biosynthetic pathway [49][51].

Besides α-ketoglutarate precursors, additional amino acids act as anaplerotic substrates for other steps of the CAC [52]. The breakdown of amino acids to compensate for the energy deficit of BTHS patients has been proposed as an explanation of skeletal muscle wasting, one of the predominant clinical features of BTHS [53]. The use of amino acids as CAC precursors in BTHS was further supported by the finding of elevated whole-body proteolysis in a small cohort of five BTHS patients compared to matched healthy controls [54]. However, there is a need for robust evidence regarding the hypothesis that proteolysis seen in BTHS patients is a mechanism for correcting aberrant

CAC function by diverting amino acids into the CAC.

Another major anaplerotic pathway is pyruvate carboxylation, whereby pyruvate is carboxylated to malate or oxaloacetate via malic enzyme or pyruvate carboxylase, respectively. Alterations of cardiac pyruvate carboxylation has been demonstrated in several animal models [43][55][56] and was shown to cause rapid contractile dysfunction

[57]. Despite the suggested importance of this pathway in cardiac disease pathogenesis, to date, no studies have investigated potential modifications of this pathway in BTHS.

Figure 1-3: Anaplerotic Pathways of the Citric Acid Cycle.

Main anaplerotic pathways shown by blue arrows. ME: Malic Enzyme; PC:

Pyruvate Carboxylase.

To better characterize energy-producing pathways in BTHS, it is necessary to look beyond glucose/fatty acid metabolism and investigate the effect of TAZ mutations on anaplerotic reactions.

1.8 Therapeutic Strategies for Modulating Cardiac Metabolism

Current experimental and pre-clinical data suggest that modulating aberrant energy metabolism can be useful in treating pathophysiological heart conditions [58]. To utilize the full oxidative capacity of the heart, therapies which inhibit or partially inhibit fatty acid oxidation to indirectly increase glucose oxidation have been investigated. This strategy has yielded mixed results [58][59]. Inhibitors of thiolase I, trimetazidine and ranolazine, have been shown to be clinically effective in preliminary trials of patients with heart failure and idiopathic dilated cardiomyopathy [60]. Thiolase I catalyzes the last step of β-oxidation, thus its inhibition decreases fatty acid oxidation while indirectly boosting glucose oxidation. In some cases, inhibition of fatty acid oxidation exacerbated heart failure [37]. It is still under debate whether inhibiting an ATP-producing pathway in the already-compromised heart is a viable treatment strategy.

Another emerging therapeutic strategy proposes to improve cardiac energetics by restoring the coupling between glycolysis and glucose oxidation [61]. Increasing glucose oxidation could help restore cardiac efficiency, as supported by numerous studies

[31][37]. The use of dichloroacetate to indirectly activate the pyruvate dehydrogenase enzyme complex facilitated flux through the CAC in heart failure patients but was associated with severe neurotoxicity. Further studies to identify and test novel, less toxic glucose oxidation stimulators are needed [31]. Current literature calls for patient-specific

15 therapeutics depending on the type and stage of disease to ensure that a proper balance between fatty acid and glucose oxidation is met.

Besides directly stimulating energy metabolism, indirect strategies to increase

ATP production are currently being investigated. Istaroxime, a modulator of calcium

ATPases, showed promise in a phase II clinical trial in heart failure patients [62]. The tetrapeptide drug, elamipretide (Bendavia), stabilizes cardiolipin in the inner mitochondrial membrane and assists in reducing the formation of reactive oxygen species

[63]. Animal studies with elamipretide demonstrated improved mitochondrial energetics

[64][65][66] and increased ATP synthesis [67]. This drug is currently under phase II clinical testing for patients with reperfusion injury [68]. Because it specifically acts upon cardiolipin, elamipretide may be particularly advantageous for BTHS patients.

Therapeutics which indirectly target ATP production may circumvent the challenging task of pharmacologically maintaining the optimal balance between fatty acid oxidation and glucose oxidation.

1.9 Induced Pluripotent Stem Cells

Previous studies of inherited cardiomyopathies have been mostly limited to mouse models, which have greatly expanded knowledge of cardiac metabolism and heart disease

[16]. However, important parameters related to cardiac action potentials, contractility, and cardiac metabolism vary between species [69]. For example, the resting heart rate in mouse models is 10 times faster than in humans [70], and studies of mouse substrains have shown differences in cardiac metabolic enzyme activity and calcium handling [50].

To eliminate species-specific differences, human cardiomyocytes would be advantageous

16 for studying disease mechanisms, identifying drug targets, and performing drug screening. However, obtaining adult cardiomyocytes for in vitro studies depends on invasive biopsy procedures which result in a limited number of non-proliferative cells, and are difficult to obtain from patients with rare diseases [71].

Human induced pluripotent stem cells (iPSCs), first described in 2007, are a type of stem cell which are generated from mature human somatic cells by forced expression of a set of four transcription factors (Figure 1-4) [72]. Like human embryonic stem cells, iPSCs have the unique capacity for self-renewal and can be differentiated into any somatic cell type [73]. However, iPSCs are not associated with the same ethical issues that prevent human embryonic stem cells from widespread use. Furthermore, iPSCs can be generated from a donor of any age. This is especially advantageous when studying rare inherited diseases, in which case, the availability of embryonic stem cells from disease patients are limited or nonexistent. iPSCs represent a powerful and revolutionary model which enables the generation of a theoretically limitless amount cellular material which, importantly, retains patient genotype [73]. This enables investigations of the resulting cardiac disease phenotype, the study of underlying mechanisms of cardiovascular disease, and the ability to test therapeutic agents for the treatment or management of cardiovascular disease on patient-specific cardiomyocytes [74]. Notably, advancements in genetic engineering allow insertion of a known disease-causative gene into a healthy cell line, enabling investigations of patient-specific, genetic cardiomyopathies with isogenic controls for the first time [75][76]. In this way, iPS-CMs have great potential for disease modeling and patient-specific drug development and

Figure 1-4: Generation of Induced Pluripotent Stem Cells from Adult Somatic Cells.

18 screening. iPSC-derived cardiomyocytes with disease-causative mutations recapitulate disease genotypes, reproducing the complex metabolic condition in the hearts of affected individuals and exhibit many of the characteristics of in vivo cardiomyocytes, including syncytial and contractile activities, ion channels, receptors and transporters. This has successfully been demonstrated by iPS-CM models of catecholaminergic polymorphic ventricular tachycardia (RYR2 mutation) [77], hypertrophic cardiomyopathy (MYH7 mutation) [78], dilated cardiomyopathy (TNNT2 mutation) [70], arrhythmogenic right ventricular dysplasia (PKP2 mutation) [37] [38].

1.10 Stable Isotope Metabolomics

Of the four major “omics” fields, metabolomics is the closest to phenotype characterization. Since metabolites are the final products of the biological response to stimuli, changes in metabolite levels are more dynamically related to disease pathogenesis [30][81]. Metabolites are a diverse class of low molecular weight compounds, which includes sugars, amino acids, organic acids, and lipids. Because metabolic pathways are tightly controlled and interconnected, changes in steady-state concentrations may be very small, even if flux through those pathways is drastically altered [82]. Thus, simple endpoint measurements of metabolite levels do not necessarily reflect changes that occur in dynamic cellular metabolism [81]. Additionally, multiple compounds may contribute to the formation of the same metabolite, precluding the determination of the origin of the produced metabolite. A final disadvantage to steady state measurements of metabolite levels is the inability to determine whether a metabolite

19 concentration is increased because of excessive formation, or because of a block in the downstream pathway resulting in accumulation of the metabolite [83].

As interest in metabolomics continues to grow, the use of stable isotopes in metabolic studies are emerging as a valuable resource for overcoming some of the drawbacks of traditional metabolomics methodologies. Stable isotopes have increased biocompatibility and are safer than radioactive labeling [82]. Relevant stable isotopes for tracer studies include 2H, 13C, 15N, and 18O. The use of these stable isotope-labeled metabolites allows the fate of the metabolite to be traced through metabolic pathways.

Stable isotope labeling can be used for absolute quantification or for determining isotopic enrichment into a given metabolite, known as the mass isotope distribution (MID).

Positional isomers of a given compound vary only in the position of labeled atom(s) present, while isotopologues are isomers which vary in the number of labeled atom(s) present, and are commonly referred to as “mass isotopomers” (Figure 1-5).

Positional isotopomers are resolved by NMR, while mass isotopomers can be easily differentiated by mass spectrometry. The MID is used to describe the extent of labeling of a compound and takes all mass isotopomers into consideration. MIDs can be used to identify pathways and metabolite partitioning, independent of metabolite concentrations

[84]. MID analysis can be combined with compound concentrations, as well as uptake and secretion rates of relevant metabolites for a more complete picture of metabolic processes under investigation. Finally, parallel experiments in which all parameters (such as media composition and compound concentrations) are held constant but the labeled tracer is varied can provide complementary information, thus increasing confidence in biological conclusions.

Figure 1-5: Mass Isotopomers of a 4-Carbon Molecule

12C (open circles) or 13C (filled circles) atoms.

1.11 References

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CHAPTER II

ESTABLISHING A MODEL OF BARTH SYNDROME USING INDUCED PLURIPOTENT

STEM CELL-DERIVED CARDIOMYOCYTES

2.1 Abstract

Despite the utility of mouse models in expanding the knowledge of cardiac disease pathogenesis in Barth syndrome (BTHS) [1], there are critical parameters of cardiac functionality which vary between mouse and human cardiomyocytes, precluding the understanding of BTHS cardiac phenotype and, consequently, the development of efficient, targeted therapeutics [2][3]. Human induced pluripotent stem cells differentiated into cardiomyocytes (iPS-CMs) recapitulate the donor genotype, reproducing the complex metabolic conditions of the affected individuals’ heart [4].

These characteristics make iPS-CMs a powerful model to delineate molecular mechanisms underlying the cardiac phenotype of BTHS. The aim of the present study was to optimize a small molecule differentiation protocol to obtain functional, mature ventricular-like iPS-CMs. Our differentiation protocol led to the generation of iPS-CMs which express cardiac specific markers such as adult-type cardiac tropinin (TNNI3) and

α-actinin. Immunostaining revealed organized sarcomeric structure and the presence of z-

29 bands. iPS-CMs in culture displayed heterogenic morphologies, including rounded, triangular, and rod-shaped cells, and included a mix of mono- and multi-nucleated cells.

Here, we describe a robust differentiation protocol of ~80% efficiency which generated iPS-CMs positive for ventricle-specific markers. The use of isogenic cell lines eliminates confounding factors of inherent genetic variability, thus increasing confidence that any observed biochemical and molecular differences are due exclusively to the TAZ mutation. Long-term cell culture up to 45 days after differentiation by our protocol produced functional control and TAZ-mutant iPS-CMs.

2.2 Introduction

Since the identification of TAZ gene mutations as the cause of BTHS [5], several cellular and animal-based models of BTHS have been developed. Early models in yeast provided important initial insight into the phenotypic diversity observed in different pathogenic TAZ mutations. Yeast models also revealed biochemical features of BTHS, including the characteristic increase in the ratio of monolysocardiolipin to tetralinoleoylcardiolipin [6], and provided the first evidence that TAZ mutations result in mitochondrial dysfunction and increased production of reactive oxygen species (ROS)

[7]. Cellular models in fibroblasts [8], lymphoblasts [9], neutrophils [10], neonatal ventricular fibroblasts [11], and cardiac myocytes [12] further revealed mitochondrial dysfunction, pointing to lower mitochondrial membrane potential, decreased respiration rates in conjunction with diminished enzymatic activity in respiratory complexes, and alterations in respiratory supercomplex structure. Animal models in Drosophilia, zebrafish, and mice established a causative relationship between TAZ deficiency and

30 clinical symptoms seen in BTHS patients, including muscle weakness, abnormal cardiac development, and early mortality [13][14][15]. Subsequent mouse models played an integral role in further describing TAZ-induced cardiomyopathy, both in early development and adulthood [16][17], and aberrant interactions of the electron transport chain with several fatty acid oxidation enzymes [18].

TAZ knockdown mouse models of BTHS were instrumental in implicating TAZ mutations in cardiac dysfunction. However, these models are not entirely suitable for modeling the functional characteristics of cardiomyocytes, such as metabolism, electrophysiology, and calcium handling, due to notable species-specific differences, precluding the understanding of BTHS and, consequently, the development of specific and effective therapeutics [19]. The discovery and development of induced pluripotent stem cells circumvents this barrier. Human induced pluripotent stem cells (iPSCs), first described in 2007, are a type of stem cell generated from mature human somatic cells by forced expression of a set of four transcription factors [20]. iPSCs have the unique capacity for self-renewal and can be differentiated into somatic cell types, including cardiomyocytes [21]. Cardiomyocytes derived from iPSCs (iPS-CMs) recapitulate the donor genotype, reproducing the complex metabolic conditions of the affected individuals’ heart. Thus, the ability to generate patient-specific iPSC-derived cardiomyocytes represents a revolutionary way to study cardiac metabolism in genetic heart diseases. Moreover, these characteristics make iPS-CMs a powerful model to delineate molecular mechanisms underlying the cardiac phenotype of BTHS and provide the ability to test therapeutic agents for the treatment or management of BTHS on patient- specific cardiomyocytes. Human iPS-CMs have already been successfully used in vitro to

31 model inherited cardiomyopathies, including BTHS, arrhythmogenic right ventricular dysplasia, hypertrophic cardiomyopathy, and dilated cardiomyopathy [2][22][23].

iPSC lines bearing various TAZ mutations have been developed via reprogramming of healthy patient fibroblasts to iPSCs and subsequent genetic engineering of TAZ-mutant iPSCs using CRISPR-Cas9 gene editing [24]. These lines have been used by Wang et al to generate iPS-CMs by early, less efficient differentiation protocols [24]. Since the advent of differentiation of iPSCs to iPS-CMs in 2008, protocols claiming to indiscriminately differentiate multiple iPSC lines with high efficiency have been published [25][26]. Many of these protocols rely on the temporal modulation of signaling pathways using small molecules to generate specific subtypes of cardiomyocytes including atrial-like [27] and ventricular-like [28] cardiomyocytes, with more mature phenotypes and morphologies [29][30].

The aim of the present study was to optimize a small molecule differentiation protocol for our specific cell lines, in order to establish high yield populations of control and TAZ-iPS-CMs for further metabolic studies. Three iPS cell lines were used. The first iPSC line was reprogrammed from fibroblasts of a healthy patient, hereby referred to as the “control”. Next, an isogenic cell line was engineered using CRISPR-Cas9 gene editing bearing the TAZ1 mutation frameshift c.517delG (hereby referred to as TAZ1).

The third line bears the TAZ mutation c.821insAAGCTAACCATGG, hereby referred to as TAZ2. For differentiation of iPSCs to iPS-CMs, we optimized a protocol reporting

>90% efficiency of differentiation [25]. This protocol utilizes two small molecules to induce cardiogenesis via temporal modulation of the Wnt/β-catenin pathway, CHIR99021 and IWR1 (Figure 2-6). CHIR99021 is an aminopyrimidine derivative that inhibits

32 glycogen synthase kinase 3 (GSK3), allowing β-catenin to regulate transcription of genes promoting cardiac mesoderm formation. IWR-1 is a potent Wnt-inhibitor, resulting in the degradation of β-catenin by GSK3, promoting cardiomyocyte specification.

Figure 2-6. Cardiogenesis via Temporal Modulation of the Wnt Pathway.

2.3 Methods

i. Cell Lines

Three human induced pluripotent stem cell lines (iPSCs) were obtained as a kind donation from Dr. William T. Pu (Harvard Stem Cell Institute). iPSCs (Coriell

GM23248), referred to herein as “control”, were reprogrammed in Dr. Pu’s laboratory from healthy donor fibroblasts. The TAZ1 iPSC line was established from control iPSCs by introducing a c.517delG frameshift mutation in the TAZ1 gene using Cas9-mediated genome editing [24]. TAZ2 iPSCs were gene edited to have the TAZ mutation TAZ mutation c.821insAAGCTAACCATGG.

Cell Line Mutation Control Wild-type TAZ1 C.517delG TAZ2 c.821insAAGCTAACCATGG

ii. Induced Pluripotent Stem Cell Culture

iPSCs were cultured on plates coated with 1:200 Matrigel in serum-free, chemically defined Essential 8 Flex Medium (Thermo, A2858501). Cells were maintained at 37°C with 5% CO2. For standard proliferation, cells were passed at a 1:12 dilution once they reached 80% confluence, Cells were detached using 0.48mM EDTA tetrasodium in phosphate buffered saline (PBS) with a 3- to 5- minute incubation period at 37°C.

35 iii. Confirmation of iPSC Pluripotency

To evaluate the quality of iPSCs, the expression of key pluripotency markers

SSEA4 and OCT2 was confirmed via an immunocytochemistry kit according to the manufacturer’s instructions (Pluripotent Stem Cell Immunocytochemistry Kit,

Invitrogen) (Figure 2-8 B, C). iv. Differentiation of iPSCs to Cardiomyocytes.

Once iPSCs reached passage 20 and colonies grew to 90- 95% confluency, they were seeded for differentiation. Differentiation was carried out via modulation of the

Wnt/β catenin pathway by an optimized small molecule protocol with modifications [31]

(Figure 2-7). To induce differentiation (Differentiation Day 0), RPMI-1640 medium with

1X B27 supplement minus insulin (Thermo, A1895601) and 9μM CHIR99021 (Cayman,

13122) was added to cells for 24 hours. Next, RPMI-B27 minus insulin was given for another 24 hours. On Differentiation Day 2, media was replaced with RPMI-B27 minus insulin and 10μM IWR1-endo (Cayman, 13659) for 48 hours. On Differentiation Day 4, a serological pipette was used to gently remove spent media to avoid excessive detachment of cells. Then, fresh RPMI-B27 minus insulin and 10μM IWR1 was added for 24 hours.

After IWR1 treatment, fresh RPMI-B27 minus insulin was added for 48 hours, once again using a serological pipette to remove media. Finally, on Differentiation Day 7,

RPMI-1640 with B27 supplement with insulin was added to cells and refreshed every other day.

Figure 2-7: Small Molecule Differentiation Workflow.

v. Cardiomyocyte Purification

To metabolically enrich iPS-CM cultures, cells were purified for 48 hours by incubation in glucose-depleted, lactate- (Sigma, cat.no. L7022, 4mM) enriched media 16 days post differentiation. After lactate purification, cell culture medium was changed back to RPMI-1640 with B27 supplement with insulin for cell culture maintenance, until cells were harvested after 45 days. vi. iPS-CM Immunocytochemistry

20 days post-differentiation, iPS-CMs were replated onto Matrigel-coated coverslips before immunostaining. To replate, trypsin was applied to the cells for 10 minutes. RPMI-1640 with B27 supplement with insulin and 5% FBS was used to make a cell suspension. The suspension was triturated thoroughly to singularize the cells. Cells were seeded at a 1:500 dilution onto the coverslips. After 24 hours, spent media was replaced with RPMI-1640 with B27 supplement with insulin, which was used to maintain iPS-CMs until day 45. On day 45, cells were washed twice with 1x PBS and fixed with

4% formaldehyde in PBS for 10 minutes at room temperature. Coverslips were washed once more with PBS, then maintained at 30 minutes at room temperature for permeabilization with 0.5% TritonX (Sigma, cat.no. T8787) and blocked with 3% bovine serum albumin in 1x PBS. Primary antibodies anti-TNNI3 (Invitrogen, PA1-86820)

1:750, IRX4 (Invitrogen, PA5-40481) 1:750, and anti-α-actinin (Invitrogen, MA1-22863)

1:500 were diluted in 3% bovine serum albumin in 1x PBS. Cells were incubated with primary antibodies at 4°C overnight, and then were washed 3 times with 1x PBS. Next, cells were incubated with secondary antibodies CF543 (Biotium, 20308) 1:250,

AlexFluor® 555 (Invitrogen, A27017) 1:200, or AlexaFluor® 488 (Invitrogen, A11059

38 and A11055) 1:200 for one hour at room temperature, followed by three washes with 1x

PBS for five minutes. Coverslips were mounted onto microscope slides using EverBrite

Mounting Medium with DAPI (Biotium). Cells were visualized with Nikon A1R confocal microscope. vii. Statistical Analysis.

The data are presented as the mean ± the standard deviation from multiple samples. Significance was tested with the paired two tailed t-test using GraphPad calculator. A p value ≤ 0.05 was considered as significant.

2.4 Results

iPSC Pluripotency. Induced pluripotent stem cell morphology is critical for monitoring the quality of the culture. iPSC lines were evaluated for critical morphological features after being proliferated in culture and prior to being used for differentiation. Control and TAZ1 colonies were densely packed (Figure 2-8 A) with well-defined edges, and individual cells demonstrated a high nucleus-to-cytosol ratio.

Importantly, cell cultures which displayed spontaneous differentiation or poor morphology in greater than 5% of colonies were disqualified to be used for differentiation. To further evaluate pluripotency, iPSCs were stained with key pluripotency markers SSEA4 and OCT4 by immunostaining (Figure 2-8 B). The nuclear marker OCT4 was expressed in all cells, while the cell surface marker SSEA4 displayed clearly-defined borders on the colonies.

Figure 2-8: Evaluation of iPSCs.

A) Representative image of control and TAZ1 iPSC colonies by bright field microscopy; 20X magnification. Immunocytochemistry of pluripotency markers Oct4

(red) and SSEA4 (green) at B) 10X magnification and C) 60X magnification. Scale bars

= B) 200 microns and C) 50 microns. Nuclei stained by DAPI (blue).

Differentiation of iPSCs to Cardiomyocytes. To obtain functional, beating cardiomyocytes from induced pluripotent stem cells, an optimized small molecule differentiation protocol was developed. As described above, temporal modulation of the small molecules CHIR99021 and IWR1 in RPMI-1640 medium supplemented with B27 minus insulin efficiently induced cardiac differentiation. Cells began to beat after 11 days of differentiation. Metabolic purification was performed with 4mM lactate medium without glucose to positively select for cardiomyocytes, effectively eliminating any non- cardiomyocyte cell types in the culture [32]. iPS-CMs were maintained in culture for 45 days to allow them sufficient time to mature.

Control and TAZ1-iPS-CMs were positive for key cardiac markers after immunostaining. The presence of the adult isoform of troponin, cardiac troponin I

(TNNI3) indicates a presence of adult like iPS-CM (Figure 2-9). Organized sarcomeric structure as shown by α-actinin staining and presence of z-bands further suggests a relatively mature iPS-CM phenotype (Figure 2-9). Control and TAZ1-iPS-CMs were positive for IRX4 (Figure 2-10). Expression of iroquois-class homeodomain protein

(IRX4) [33] indicates the presence of ventricular-like cell subtype.

iPS-CMs in culture displayed a mix of morphologies, including rounded, triangular, and rod-shaped cells, and included a mix of mono- and multi-nucleated cells

(Figure 2-9, Figure 2-10).

Figure 2-9. Immunocytochemistry of iPS-CMs

Representative images of iPS-CMs stained positive for α-actinin (green), TNNI3

(red), and DAPI (blue). 60X magnification. Scale bar = 20 microns

Figure 2-10: Immunocytochemistry of iPS-CMs.

Representative images of iPS-CMs stained positive for TNNI3 (green) and IRX4

(red). The nuclear marker DAPI is shown in blue. 60X magnification. Scale bar = 20 microns

2.5 Discussion

Although it is well-established that mutations in the TAZ gene induce cardiac pathogenesis in BTHS, the actual mechanisms of disease onset and progression remain poorly understood. Cellular and animal models of BTHS were fundamental to providing early insight into the mechanisms of the disease, such as establishing a causative role of

TAZ mutations in mitochondrial dysfunction and production of ROS [1]. However, TAZ- mutant iPS-CMs enable investigations of molecular mechanisms of BTHS in cells of human origin and provide the ability to test therapeutic agents for the treatment or management of BTHS on patient-specific cardiomyocytes.

For our studies, three iPSC lines were donated from the Harvard Stem Cell

Institute. The pluripotency of these lines was previously established [24], however, iPSCs are susceptible to spontaneous differentiation, thus losing their ability to be induced toward a specific somatic cell type. Shear force, the presence of antibiotics, changes in media composition, changes in temperature or CO2 levels, or inconsistent passaging are among some of the factors that can lead to spontaneous differentiation [33][34][35].

Earlier formulations of iPSC media were comprised of growth factors, thus introducing batch-to-batch variability and increasing the chance of spontaneous differentiation [36].

In these studies, we expanded control and TAZ1 iPSC cultures in serum-free, fully defined medium. iPSCs cultured in our laboratory showed proper rounded morphology, with individual cells demonstrating a characteristic high nucleus-to-cytosol ratio. iPSC colonies were densely packed with well-defined edges. >99% of iPSCs in culture were positive for markers of pluripotency, including the cell surface marker SSEA4 and the

44 nuclear marker OCT4. These results demonstrate that the iPSC lines were successfully re-established into culture in our laboratory under optimized cell culture conditions.

BTHS patients present with dilated cardiomyopathy, a disease which presents with ventricular dilation. Thus, we sought a differentiation method to produce ventricular-like iPS-CMs. Several signaling pathways have emerged as the primary determinants for cardiac specification, thus many research groups have attempted to modulate one or several of these pathways using a number of techniques. Many of these protocols failed to generate a homogenous population of a specific cardiac-subtype, suffered from poor differentiation efficiency, and were largely cell-line dependent.

However, several recent publications report high-efficiency differentiation of >80% cardiomyocytes resulting in enriched populations of ventricular cardiomyocytes [25][26].

In this work, we chose to modify one such protocol due to its chemically-defined nature and reported high yields of ventricular-like iPS-CMs [31]. We optimized this small molecule protocol to our cell lines, based on modulation of the Wnt/β-catenin signaling pathway. By temporal addition of the GSK3 inhibitor, CHIR99021, followed by addition of the Wnt-inhibitor IWR1, we observed robust differentiation with up to 80% efficiency across several iPSC lines. Spontaneously beating cardiomyocytes appeared about ten days after differentiation was initiated (cell line-dependent).

Once functional cardiomyocytes were obtained, they were purified with lactate to enrich the population of cardiomyocytes due to the unique ability of cardiomyocytes to utilize lactate as a source of energy [32]. We next maintained iPS-CMs in culture for forty-five days to allow the cells to develop a more mature phenotype, as immature iPS-

CMs have been found to more closely resemble the gene expression and metabolic

45 phenotype of fetal cardiomyocytes [37]. Maturation of iPS-CMs has been shown to increase through long-term cell culturing [29]. Maturation status is typically evaluated by gene expression, protein composition and organization of the sarcomere, respiratory capacity/oxidative redox potential, and electrophysiological parameters. Here, we performed immunostaining for sarcomeric proteins to determine iPS-CM maturity and cardiac-subtype.

Immunostaining revealed that control and TAZ1 iPS-CMs express key cardiac markers. The presence of the adult isoform of troponin, cardiac troponin I (TNNI3) indicates presence of mature iPS-CMs in culture, while organized sarcomeric structure as shown by α-actinin staining and presence of z-bands further suggests cardiomyocyte maturity. iPS-CMs displayed heterogenic morphologies, including rounded, triangular, and rod-shaped cells, and included a mix of mono- and multi-nucleated cells (Figure 2-9).

Adult cardiomyocytes have a characteristic rod shape and are generally multinucleated.

Thus, iPS-CMs did not appear to achieve full maturation, which is a known limitation of iPS-CM models and a major, ongoing challenge in the field [38]. While a variety of physical [39], biochemical [37][40], and tissue-engineering techniques [41] have been applied to generate mature iPS-CMs, no single technique, or combination of techniques, is yet able to produce iPS-CMs which are completely identical to human adult cardiomyocytes. An additional discussion of this topic can be found in Chapter VI.

Control and TAZ1-iPS-CMs expressed iroquois-class homeodomain protein

(IRX4), indicating the presence of a ventricular-like cell subtype [31]. While immunocytochemistry is a reliable qualitative technique to examine the presence of important myocardial proteins, quantitative approaches such as mRNA expression and

46 flow cytometry are better suited for assessing cardiomyocyte subtypes, and thus should be considered in future studies for a more robust assessment of cardiac subtype.

Here, we optimized a differentiation protocol of ~80% efficiency which generated a high yield of ventricular-like iPS-CMs. The use of two isogenic cell lines eliminates confounding factors of inherent genetic variability, thus increasing confidence that any observed biochemical and molecular differences are due exclusively to the TAZ1 mutation, whereas the use of two TAZ mutants, TAZ1 and TAZ2, allow investigations of genotype-phenotype correlations. Our data show that iPSCs maintained pluripotent potential, an important condition signifying that cells are able to undergo differentiation to the somatic cell type of choice. Furthermore, immunocytochemistry of beating iPS-

CMs revealed organized sarcomeric structure, providing evidence of a relatively mature phenotype. This work resulted in successful production of control and TAZ-mutant iPS-

CMs for further studies of the cardiac features of BTHS.

2.6 References

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vol. 12, no. 1, pp. 127–137, 2013. [33] K. G. Chen, B. S. Mallon, R. D. G. McKay, and P. G. Robey, “Human pluripotent stem cell culture: considerations for maintenance, expansion, and therapeutics.,” Cell Stem Cell, vol. 14, no. 1, pp. 13–26, Jan. 2014. [34] G. S. Belinsky and S. D. Antic, “Mild hypothermia inhibits differentiation of human embryonic and induced pluripotent stem cells.,” Biotechniques, vol. 55, no. 2, pp. 79–82, Aug. 2013. [35] M. Serra, C. Brito, C. Correia, and P. M. Alves, “Process engineering of human pluripotent stem cells for clinical application,” Trends Biotechnol., vol. 30, no. 6, pp. 350–359, Jun. 2012. [36] G. Chen et al., “Chemically defined conditions for human iPSC derivation and culture.,” Nat. Methods, vol. 8, no. 5, pp. 424–9, May 2011. [37] C. Correia et al., “Distinct carbon sources affect structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells.,” Sci. Rep., vol. 7, no. 1, p. 8590, Dec. 2017. [38] T. J. Kolanowski, C. L. Antos, and K. Guan, “Making human cardiomyocytes up to date: Derivation, maturation state and perspectives,” Int. J. Cardiol., vol. 241, pp. 379–386, Aug. 2017. [39] E. Tzatzalos, O. J. Abilez, P. Shukla, and J. C. Wu, “Engineered heart tissues and induced pluripotent stem cells: Macro- and microstructures for disease modeling, drug screening, and translational studies.,” Adv. Drug Deliv. Rev., vol. 96, pp. 234–244, Jan. 2016. [40] F. B. Bedada et al., “Tri-iodo-l-thyronine promotes the maturation of human cardiomyocytes-derived from induced pluripotent stem cells.,” J. Mol. Cell. Cardiol., vol. 12, no. 1, pp. 296–304, 2017. [41] C. P. Jackman, A. L. Carlson, and N. Bursac, “Dynamic culture yields engineered myocardium with near-adult functional output,” Biomaterials, vol. 111, p. 66, 2016.

CHAPTER III

STABLE ISOTOPE TRACING STUDY IN INDUCED PLURIPOTENT STEM CELL-

DERIVED CARDIOMYOCYTES TO INVESTIGATE METABOLIC ALTERATIONS IN

BARTH SYNDROME

3.1 Abstract

Barth syndrome (BTHS) is an X-linked recessive multisystem disorder associated with cardiomyopathy, neutropenia, exercise intolerance, sudden cardiac death, skeletal muscle weakness, recurrent bacterial infections, and growth delay. BTHS is caused by mutations in the TAZ gene (TAZ, G 4.5, OMIM 300394) that encodes for the acyltransferase tafazzin. This protein is highly expressed in the heart and plays a significant role in cardiolipin remodeling. Heart disease is the major clinical manifestation of BTHS with a high incidence in early life. There is no specific treatment for the cardiac implications of BTHS and clinical management is largely supportive, using standard heart failure medications. Although the genetic basis of BTHS and the resulting tetralinoleoyl cardiolipin deficiency in BTHS-affected individuals are well- established, downstream metabolic changes in the heart remain to be elucidated. There is, therefore, a critical need to delineate TAZ-induced metabolic changes in the heart during

51 the course of the disease. In the absence of such knowledge, the development of effective cardiac therapies for BTHS will likely remain difficult.

Our study aimed to characterize TAZ-induced metabolic perturbations in the heart. This was achieved by metabolic profiling, targeted gene expression analysis, and studies with 13C-labeled substrates in an induced pluripotent stem cell-derived cardiomyocyte (iPS-CM) model of BTHS. We found evidence of perturbations in fatty acid levels and gene expression in our TAZ-iPS-CM model. Our data also demonstrate that a TAZ mutation induces metabolic alterations in pathways related to energy production and causes a shift in the preferred carbon energy substrate from fatty acids to glucose.

3.2 Introduction

Barth syndrome (BTHS) is an X-linked recessive multisystem disorder associated with cardiomyopathy, neutropenia, exercise intolerance, sudden cardiac death, skeletal muscle weakness, recurrent bacterial infections, and growth delay [1][2]. The approximated BTHS prevalence of 1/300,000-400,000 live births is likely underestimated since the disorder is substantially under-diagnosed. BTHS is caused by mutations in the

TAZ gene (TAZ, G 4.5, OMIM 300394) [3], which encodes for the acyltransferase tafazzin. This protein is involved in the remodeling of tetralinoleoyl cardiolipin, a mitochondrial membrane-associated phospholipid with a significant role in mitochondrial-related processes, especially in the heart [4]. Due to the impaired ratio of monolysocardiolipin to tetralinoleoylcardiolipin ratio in Barth syndrome [5], mitochondria from various cell types in humans and mice demonstrate morphological

52 abnormalities [6][7]. BTHS-affected individuals present with metabolic alterations that include lactic acidosis, 3-methylglutaconic aciduria [8], low plasma arginine [2], and a severe deficiency of mitochondrial tetralinoleoylcardiolipin [1][9]. The manifestation of cardiac disease is the major clinical feature of BTHS [10], with a high incidence in early life and, subsequently is a leading cause of death in infants. Affected individuals can present with hypertrophic cardiomyopathy (HCM), dilated cardiomyopathy (DCM) with endocardial fibroelastosis (EFE) [11], left ventricular non-compaction (LVNC), ventricular arrhythmia [12], sudden cardiac death, or prolonged QTc interval. Since the mechanisms of cardiac disease pathogenesis in BTHS are poorly understood, there is currently no specific treatment for the cardiac features of the disease. At the same time, there is renewed interest in utilizing metabolic therapies to treat heart diseases by regulating energy metabolism, supported by experimental [13][14] and pre-clinical data

[15][16][17]. Therefore, an understanding of the effect of TAZ mutations on cardiac metabolism has important implications for BTHS treatment.

In many types of heart disease, inadequate supply of ATP for contractile function and decreased cardiac efficiency are associated with altered mitochondrial energy metabolism [2][18]. Fatty acids are the main carbon source for ATP synthesis in the healthy adult heart. Cellular free fatty acid uptake is facilitated by transporters and to a lesser extent by diffusion across the cellular membrane. Upon entrance into the cell, long chain fatty acids such as palmitate (C16) are rapidly activated to form acyl-CoAs in the cytosol and then are converted to acylcarnitines to enter mitochondria through the carnitine shuttle. However, a shift from fatty acid oxidation to glucose utilization has been observed in the progression of many forms of heart disease [4][5]. This is generally

53 accompanied by the heart reverting to a glycolytic state, mimicking the energy metabolism of the fetal heart. In this scenario, glycolysis may become uncoupled from glucose oxidation, subsequently affecting fatty acid oxidation, as the two oxidative processes are intricately linked [19]. However, whether the shift is adaptive or causative in the development of heart failure and the specific mechanisms of these alterations are still poorly understood. Furthermore, the downstream metabolic effects are not well defined, presenting a major obstacle for the discovery of new therapeutic targets.

Our study aimed to investigate the metabolic consequences of TAZ mutations

(frameshift c.517delG; c.821insAAGCTAACCATGG) in an iPS-CM model of Barth syndrome. This was achieved by experimental workflows with stable isotopic tracers, targeted metabolic profiling and targeted gene expression. We hypothesized that as a result of impaired mitochondrial structure, a defect in TAZ affects energy production- related pathways in cardiomyocytes. The proposed studies provide novel insight into downstream metabolic differences in energy substrate metabolism between control and

TAZ-mutant iPS-CMs using stable isotope tracing. We report our findings herein.

3.3 Materials and Methods.

i. Cell Culture

Control, TAZ1, and TAZ2 iPSCs were differentiated to functional iPS- cardiomyocytes as described previously (See Chapter II).

ii. Carbon Tracer Experiments

45 days post-differentiation, RPMI-1640 medium was aspirated from beating iPS-

CMs. Cells were washed quickly with 1x D-PBS and medium was substituted with a

13 custom-prepared RPMI-1640 with 10mM C6-glucose with 0.4mM palmitic acid-BSA or

10mM glucose with 0.4mM 13C-palmitic acid-BSA. Prior to tracer experiments, palmitic acid was conjugated to bovine serum albumin (BSA) depleted from any fatty acids or lipids by charcoal purification [20]. iii. Cell Harvesting and Metabolite Extraction

At the end of the tracing period, the tracer media was aspirated and iPS-CMs were washed two times with cold 1x D-PBS and quickly washed once with cold water.

Metabolism quenching was achieved by the addition of a cold acetonitrile-water solution

(2ml:1.75ml per well). Cells were scraped, and lysates were transferred to tubes, followed by the addition of 2mL chloroform. Cell lysates were centrifuged at 5000rpm for 15 minutes. The resulting polar and nonpolar phases were separated from the intermediate protein layer. Polar and nonpolar phases were dried under a nitrogen stream at room temperature for downstream metabolic analyses, while the isolated protein layer was retained for total protein measurement. iv. Glucose Uptake

To analyze glucose uptake, cell media aliquots were collected during the tracer experiments. Galactose (10μL of 1mM) was added as an internal standard for glucose.

Metabolites were extracted from media aliquots using 1:1 methanol/chloroform. Samples were rocked for 10 minutes, then centrifuged at 5000rpm for 5 minutes. The resulting

55 polar phase was separated from the non-polar phase and dried down under N2 for downstream glucose analysis.

v. Palmitate Uptake

To analyze palmitate uptake, cell media aliquots were collected during the tracer experiments. Methylated-C17 was used as the internal standard for palmitate-BSA.

Metabolites were extracted from media aliquots using 1:1 methanol/chloroform. Samples were rocked for 10 minutes, then centrifuged at 5000rpm for 5 minutes. The resulting nonpolar phase was separated from the polar phase, adjusted to pH 10 with NaOH, and dried down under N2. 0.25mL hexane and 0.25mL BF3/MeOH were added to derivatize samples for 30 minutes at 80°C. After derivatization, pH of samples was adjusted to 3 with HCl. 150uL saturated NaCl and 100uL hexane were added and then samples were centrifuged at 5000rpm for 5 minutes. The top phase was removed and dried under N2 to ensure the sample was free of water, resuspended in 100μL hexane, transferred to a glass vial, and injected via GC-MS. vi. Derivatization and GC-MS Parameters for Cellular Metabolite Analysis

Citric acid cycle intermediates, amino acids, fatty acids, and glucose were derivatized and analyzed via GC-MS according to optimized parameters for each class of metabolites. Derivatization details for each class are as follows. Full GC-MS parameters, including the m/z ions monitored, can be found in Appendix B.

Citric Acid Cycle Intermediates

Dried cell extracts were derivatized at 80°C for 60 minutes with 20mg/mL methoxyamine hydrochloride (Sigma, 226904) in pyridine followed by derivatization

56 with N-methyl-N-tert-butyldimethylsilyltrifluoroacetamide (Sigma, 375934) at 70°C for

45 minutes. GC-MS analysis was carried out with a 5977 GC-MS (Agilent) operated in

EI mode and equipped with an HP-5ms, 30m column (Agilent). Retention times and fragmentation mass spectra of all metabolites were confirmed with commercially available standards and the NIST library. Unlabeled (M0) ions monitored through the method are presented in Appendix B.

Amino Acid Analysis

Dried cell extracts were derivatized at 80°C for 60 minutes with 20mg/mL methoxyamine hydrochloride (Sigma, 226904) in pyridine followed by derivatization with N-methyl-N-tert-butyldimethylsilyltrifluoroacetamide (Sigma, 375934) at 70°C for

45 minutes.

Untargeted Fatty Acid Analysis

Dried cell extracts were derivatized at 80°C for 60 minutes with 20mg/mL methoxyamine hydrochloride (Sigma, 226904) in pyridine followed by derivatization with N,O-Bis(trimethylsilyl)trifluoroacetamide (Sigma, B-023) at 70°C for 30 minutes.

Glucose Assay

Dried media extracts were derivatized using 0.2M hydroxylamine hydrochloride

(Sigma, 159417) at 90°C for 40 minutes, followed by derivatization with acetic anhydride at 90°C for 60 min. The derivatized samples were dried down under N2 and resuspended in ethyl acetate. Samples were analyzed by GC-MS using a SIM method to monitor for m/z 314, 315, 316, 317, 318, 319, 320, and 321 for all mass isotopomers of glucose, and m/z 314 for galactose.

vii. Calculation of Metabolite Levels

Metabolite levels were normalized to the reference standard tricarballylic acid

(Sigma, T53503; 10μl of 1mM) and then to the total protein amount of each cell pellet as

measured by the Pierce™ BCA Protein Assay (Thermo). Mass spectra were used to

calculate the ratio of peak areas of target metabolites to the reference standard. The

calculated ratios were used for crossover analysis to compare metabolite levels in TAZ

mutant vs control cells. In crossover analysis, the relative level of a given metabolite in

the control group is always expressed as 100%.

Relative level =

Average [(Peak area of metabolite)/ (Peak area of reference compound)]TAZ

Average [(Peak area of metabolite)/ (Peak area of reference compound)]control

viii. Mass Isotope Distribution Analysis

Mass isotopomers are molecules with the same molecular structure which differ

only by the number of 13C atoms present, resulting in different molecular weights which

can be resolved by mass spectrometry. For example, M2-Citrate signifies that two out of

six carbons are labeled with 13C. Peak areas of each mass isotopomer were integrated

from GC-MS chromatograms and corrected for natural abundance of all elements

contained in the analyzed molecule (See Appendix A). Percent enrichment of a given

mass isotopomer is expressed as the percent fraction of that specific isotopomer to the

sum of all isotopomers, including the unlabeled component, M0.

Mx % Enrichment of Mx = ∑ M0,M1,M2,…Mn

ix. Gene Expression Analysis

RNA from cells in independent biological triplicates was extracted using RNAzol

(Molecular Research Center, Inc) according to the manufacturer's instructions. RNA quality was assessed by NanoDrop. cDNA was generated using GeneAmp RNA PCR

Core Kit (Applied Biosystems). Real-time qPCR was carried out using TaqMan Assays- on-Demand Probe technology (Applied Biosystems). The following probes were used

(Thermo-Fisher): FABP4 Hs01086177_m1 and FABP3 Hs00997360_m1. 18s rRNA

(4352930, Thermo-Fisher) was used as a reference gene. Relative expression levels were calculated as 2−ΔΔCT.

x. Measurement of Acyl Carnitines

A mixture of labeled acyl carnitines was used as an internal standard for endogenous acyl carnitines (Cambridge Isotope Laboratories, cat.no NSK-B). Dried cell extracts were derivatized using 3M HCl-n-butanol for 30 minutes at 65°C. After derivatization, the samples were cooled to room temperature and dried down again. The samples were reconstituted in 100μL of acetonitrile with 0.1% formic acid. Separation of acyl carnitines was performed on a UHPLC (Shimadzu Nexera) using a C18 column

(Waters XBridge) held at 40°C with a flow rate of 0.6mL/minute. The composition of mobile phases was 0.1 % formic acid in water (Mobile Phase A) and 0.1% formic acid in acetonitrile (Mobile Phase B). The following gradient elution was applied:

% Mobile Phase Time B (minutes) 20% 1 37% 8 100% 22

Gradient elution was followed by re-equilibration at starting conditions for 4 minutes.

After chromatographic separation, compounds were infused onto the mass spectrometer

(ABSciex QTRAP 5500) and monitored for the parent ion transition of 85 Da. xi. Statistical Analysis

Data are presented as the mean ± standard deviation from multiple samples.

Significance was tested with paired two tailed t-test using GraphPad calculator. A difference of p ≤ 0.05 was considered significant.

3.4 Results.

Alterations in Substrate Preference: Healthy adult cardiomyocytes derive a significant amount of ATP from fatty acid catabolism. However, to meet the high energy demand of the heart, cardiomyocytes can shift their reliance on fatty acids to other carbon sources in response to developing cardiac pathologies. To determine the effect of TAZ1

13 on carbon source selection, we incubated iPS-CMs in 10mM C6-glucose with 0.4mM

13 unlabeled palmitate or 10mM glucose medium with 0.4mM C16-palmitate and measured the disappearance of 13C-tracers from the media after 8 hours. TAZ1-iPS-CMs consumed

3.21mM + 0.367 glucose, which is 1.5 times more than the control iPS-CMs (1.98mM +

0.101; p = 0.0177). TAZ1-iPS-CMs consumed about 20% less palmitate than control cells (0.158mM + 0.021 in TAZ1 versus 0.202mM + 0.009 in control; p = 0.0323).

4.00 0.2500 * 3.50 0.2000 3.00 * 2.50 0.1500

2.00

mM mM 1.50 0.1000 1.00 0.0500 0.50 0.00 0.0000 Glucose Palmitate

Control TAZ1 Control TAZ1

Figure 3-11: Glucose and Palmitate Consumption.

13 13 Concentration (mM) of C6-glucose and C16-palmitate consumed after 8 hours.

p < 0.05

Contribution of 13C-Carbon Sources to the CAC. We further analyzed incorporation of 13C from glucose and palmitate into their downstream metabolites. 13C- labeled glucose is metabolized via glycolysis to yield M3 pyruvate, which further yields

M2 acetyl-CoA after pyruvate decarboxylation. Unlabeled palmitate undergoes β- oxidation and yields only M0 acetyl-CoA. The total acetyl-CoA pool is then condensed with unlabeled oxaloacetate to produce M0 and M2 citrate (Figure 3-12A) and downstream 13C-labeled CAC intermediates. We analyzed the fraction of M2 labeled citrate as an indicator of glucose contribution to the CAC. M2 citrate was 24.1% + 0.0038 in the control, 26.8% + 0.0031 in TAZ1 (p = 0.0046), and 28.7% + 0.0026 (p = 0.0061) in TAZ2-iPS-CMs (Figure 3-12A). The higher percent fraction of M2 citrate in TAZ-iPS-

CMs suggests increased reliance on glucose for CAC replenishment and suggests a shift toward glucose as the preferred carbon source. The percent enrichment of M3 mass isotopomers of fumarate and malate are significantly decreased in TAZ1 and TAZ2 iPS-

CMs (Figure 3-12B)

Figure 3-12. Mass Isotopomer Distributions.

A) M2 Labeling and (B) mass isotopomer distributions (MID) of CAC intermediates from 13C-glucose. n= 6 (control, TAZ1); n = 3 (TAZ2)

3.3 Alterations in substrate preference: [1,2,3,413C]-labeled palmitate is metabolized through β-oxidation to yield M2 and M0 acetyl CoA, while supplied unlabeled glucose is simultaneously converted to pyruvate and further yields only M0 acetyl CoA (Figure 3-13). The total acetyl CoA pool is then condensed with unlabeled oxaloacetate to produce M0 and M2 citrate (Figure 3-13) and downstream 13C labeled

CAC intermediates. We analyzed the fraction of M2 labeled citrate as an indicator of palmitate flux through the β oxidation pathway. The lower percent fraction of M2 citrate in TAZ-iPS-CM indicates decreased palmitate utilization for CAC replenishment and suggests a shift away from palmitate as a carbon source.

Figure 3-13: M2 Labeling of CAC Intermediates from 13C-Palmitate.

The % fraction of M2 compared to the total pool of isotopomers in control (red bars) and TAZ1 (blue bars) iPS-CMs (n=3).

3.4 Alterations in CAC Intermediates. Given the essential role of the CAC for energy metabolism, we further analyzed cellular levels of CAC intermediates and found alterations in succinate, fumarate and malate pools (Figure 3-14). The abundance/mg of succinate in TAZ1-iPS-CM was increased to 0.31 + 0.012 compared to the control value of 0.18 + 0.005 (p < 0.05). There was a decrease in the levels of fumarate and malate in

TAZ1-iPS-CM, although only the decrease in fumarate was statistically significant.

Fumarate abundance/mg decreased from 5.24 + 0.377 in the control to 2.71 + 0.265 in

TAZ1-iPS-CMs (p < 0.005), whereas malate decreased from 1.80 + 0.260 in control to

1.34 + 0.304 in TAZ1-iPS-CMs.

Figure 3-14: Levels of CAC Intermediates.

Relative levels of CAC intermediates normalized to total protein amount (n=6).

*p < 0.05, **p < 0.005.

Disturbances in Fatty Acid Metabolism: Fatty acids are the main carbon source for ATP synthesis in the healthy adult heart. Cellular free fatty acid uptake is facilitated by transporters and to a lesser extent by diffusion across the cellular membrane. We selectively performed a gene expression assay for two isoforms of fatty acid transporters.

Fatty acid binding protein (FABP) isoforms FABP3 and FABP4 encode for heart and adipocyte proteins, respectively. Due to the reported association between FABP4 levels to cardiac function, we measured expression levels of both FABP4 and FABP3 in TAZ1- iPS-CM and matching isogenic control cells. The data indicate an increase in FABP4 mRNA expression levels (Figure 3-15B). Next, we performed Western blot analysis of

FABP4 to analyze protein expression levels (Figure 3-15C). In agreement with mRNA,

FABP4 protein expression levels were elevated in TAZ1-iPS-CMs.

Figure 3-15: Fatty Acid Binding Proteins.

(A) FABP3 and (B) FABP4 mRNA expression levels in control and TAZ1 iPS-

CMs. *p ≤0.05 and (C) Representative FABP4 Western Blot with GAPDH as a loading control. n=3.

Disturbances in Fatty Acid Metabolism: We further investigated fatty acid metabolism in TAZ-iPS-CMs by measuring intracellular levels of fatty acids by GC-MS and acyl carnitines by LC-MS/MS. TAZ-iPS-CMs exhibited an accumulation of free fatty acids (Figure 3-16A) including C14(myristic acid), C16 (palmitic acid) and C18 (stearic acid). Furthermore, we observed increased levels of long chain acyl carnitines (Figure

3-16B), including C14 (myristoyl-carnitine), C16 (palmitoyl-carnitine) and C18

(stearoyl-carnitine).

Figure 3-16 Fatty Acids and Acyl Carnitines.

(A) Levels of fatty acids in control, TAZ1, and TAZ2 iPS-CMs (n=3) and (B)

levels of acyl carnitines in control and TAZ1 iPS-CMs (n=3).

3.5 Discussion

Dysregulation of cardiac energy metabolism is recognized as a characteristic biochemical feature in various heart diseases. In this study, we investigated the metabolic consequences of two TAZ (c.517delG; c.821insAAGCTAACCATGG) mutations in an iPS-CM model of Barth syndrome.

Alteration in substrate preference: The metabolic flexibility of the heart allows a shift of reliance from one carbon source to another in response to developing pathologies [10][11]. In the healthy adult heart, most of the energy in the form of ATP derives from fatty acid β-oxidation [23][24]. We hypothesized that the TAZ mutation induces a carbon fuel shift from fatty acids to the glucose. To test this hypothesis, we performed parallel experiments using either labeled glucose with unlabeled palmitate or labeled palmitate with unlabeled glucose.

13 We introduced 10mM C6-glucose into cell media over a period of 8 hours to monitor glucose consumption by iPS-CMs. TAZ1-iPS-CMs consumed 1.6 times more glucose than control cells. We further analyzed 13C incorporation in the downstream

CAC intermediates. One notable finding was that the percent enrichment of M2 citrate in

TAZ1 and TAZ2-iPS-CMs was higher than the control, further suggesting that TAZ-iPS-

CMs have increased reliance on glucose as a fuel source for the CAC. Another important finding is the markedly decreased fraction of M3 malate and M3 fumarate in TAZ-iPS-

CMs, suggesting alterations in pyruvate carboxylation. A complete discussion of the significance of this finding is discussed in Chapter IV.

Next, we introduced labeled palmitate into cell media and collected aliquots after

8 hours of tracing. The data demonstrate that TAZ-iPS-CMs consumed 20% less

72 palmitate in 8 hours than control iPS-CMs, supporting the hypothesis that there is a shift

13 from fatty acid oxidation in the presence of the TAZ mutation. Using [1,2,3,4- C4]- palmitate as a tracer, we further analyzed 13C incorporation in the downstream CAC intermediates. Palmitate contribution for citrate synthesis was 14% and 11.5% in control- iPS-CM and TAZ1-iPS-CM respectively (Figure 3-13).

The observed reduction in palmitate oxidation as reflected by the lower pool of

13 M2 citrate produced from the [1,2,3,4 C4]-palmitate in TAZ1-iPS-CM can be attributed to metabolic adaptation toward glucose utilization. This is further supported by the observed increase in glucose oxidation as reflected by the higher pool of M2 glucose in

TAZ1-iPS-CM. The reliance on glucose as a carbon source improves oxygen efficiency per each generated ATP molecule (Figure 1-2). Whereas a full oxidation of one palmitate molecule requires 50 moles of atomic oxygen, a full oxidation of one molecule of glucose requires only 12 moles of oxygen [25]. The downstream consequence of this observed metabolic shift is also reflected in the accumulation of free fatty acids and acyl carnitines

(Figure 3-16), further contributing to the cellular functional impairments [26].

Alterations in the CAC: Of particular importance to cardiac energy metabolism is CAC function. A constant pool of CAC intermediates is required to produce adequate amounts of NADH and FADH2 under normal physiological conditions, thus there is a balance between production and efflux of all CAC intermediates. TAZ defects induce mitochondrial dysfunction [27] and impair this balance, leading to alterations in metabolic fates. Analysis of the levels of CAC intermediates in iPS-CMs indicates a statistically significant increase in the total succinate pool in TAZ1-iPS-CMs (Figure

3-14). This increase may be associated with cardiolipin-dependent destabilization of

73 respiratory chain complexes. Succinate dehydrogenase (SDH) is a component of the respiratory chain Complex II. In the presence of monolysocardiolipin (MLCL), efficient electron transfer across respiratory complexes is compromised due to loss of structural integrity [27]. Moreover, defective SDH could contribute to cardiac function by preventing full oxidation of glucose and fatty acids. In fact, Dudek et al. reported a decrease in SDH activity and protein expression levels in BTHS patient iPS-CMs and cardiomyocytes isolated from a mouse model of BTHS [28]. Loss of SDH activity or expression could explain the buildup of succinate and decrease of fumarate and malate observed in our TAZ-iPS-CM model. Although the decrease in malate levels was not statistically significant, this could be a result of various metabolic pathways which can replenish malate, such as pyruvate carboxylation via malic enzyme or the conversion of aspartate to malate. Anaplerotic pathways of the CAC are discussed in Chapter IV.

Alterations in fatty acid metabolism. The cellular uptake of fatty acids is facilitated by fatty acids binding proteins (FABPs). The FABP4 isoform is a member of the intracellular lipid binding protein family (iLBP). The iLBP family consists of small

(~15 kDa) soluble proteins that serve as modulators of intracellular lipid homeostasis by regulating long chain fatty acid transport in the nuclear and extra-nuclear cell compartments. The FABP4 isoform is predominantly expressed in adipose tissue, but also circulates in human plasma [15][16]. Aside from adipose tissue the heart has the highest FABP4 protein expression level [31]. It is involved in intracellular lipid trafficking and has a role in the development of insulin resistance, atherosclerosis, and inflammatory processes [32]. In accordance with recent studies, FABP4 over-expression in mouse adipocytes leads to the decreased expression of mitochondrial fatty acid

74 oxidation genes and reduced activities of mitochondrial complexes I and III [33]. A transgenic mouse study showed that FABP4 over-expression promotes the development of cardiac hypertrophy under pressure overload [31]. Moreover, human FABP4 was shown a detrimental effect on rat cardiomyocyte contraction [34] in vitro through acutely depressed shortening amplitude and intracellular systolic peak Ca2+. Increased FABP4 expression in TAZ1-iPS-CM (Figure 3-15) suggests that FABP4 plays a role in TAZ- induced cardiac dysfunction, although the exact mechanism of FABP4 in pathophysiological involvement in BTHS has not yet been investigated. Interestingly, despite the fact that FABPs are considered cytosolic proteins, FABP4 interacts in vitro with cardiolipin-containing membranes [35]. FABP4 secretion is stimulated by an increase in intracellular calcium in adipocytes [36], thus over expression of FABP4 in

TAZ-iPS-CM can be correlated to impaired calcium handling, which is discussed in detail in Chapter V. Alternatively, FABP4 overexpression can be associated with free fatty acid increase and elevated acyl carnitines (Figure 3-16), which can lead to decreased cardiac efficiency through a number of possible mechanisms [24] including the rise of intracellular calcium [37].

Barth syndrome is manifested biochemically by low tetralinoleoyl cardiolipin content and an impaired monolysocardiolipin/tetralinoleoyl cardiolipin (MLCL/18:2 CL) ratio. The uniformly substituted (18:2) cardiolipin plays a pivotal role in the structural organization of the mitochondrial membrane [38] and is essential to acylcarnitine translocase activity [39]. Accumulation of fatty acids and long chain acylcarnitines observed in TAZ1-iPS-CM is in agreement with alterations in fatty acid oxidative metabolism [40]. The increased levels are especially striking for C14, C16 and C18 acyl

75 carnitine species with fold of change 1.7, 2.5 and 2.3 respectively (Figure 3-16 B).

Interestingly, the corresponding fatty acid species C14, C16, and C18 are also elevated.

To the best of our knowledge, elevated circulating acylcarnitine levels have never been reported in BTHS individuals. Here, we report an increase in cellular levels of fatty acids and acylcarnitines that may have a detrimental effect on the electrophysiology of cardiomyocytes [41][42]–[45].

In summary, our findings reveal that the TAZ mutation induces metabolic alterations in energy production pathways and causes a shift from fatty acids to glucose as the preferred carbon substrate. In light of TAZ deficiency-induced impaired mitochondrial function, the increased reliance on glucose can be attributed to the improvement of oxygen efficiency per each generated ATP molecule. The exact functional significance of metabolic remodeling in Barth syndrome remains to be elucidated.

Study limitations: It should be noted that more than 120 different Barth syndrome- causative TAZ mutations have been identified, whereas our study focused only on two specific TAZ mutations. There are no established genotype/phenotype correlations and large phenotypic variation is present within Barth syndrome affected individuals. Further studies are needed to assess metabolic and molecular changes in cells carrying different mutations in the TAZ gene.

3.6 References

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CHAPTER IV

ANAPLEROTIC PATHWAYS OF THE CITRIC ACID CYCLE IN TAZ-MUTANT

INDUCED PLURIPOTENT STEM CELL-DERIVED CARDIOMYOCYTES

4.1 Abstract

Citric acid cycle (CAC) intermediates are recycled in a set of oxidation reactions that generate cellular energy but may also exit the CAC for use in biosynthetic reactions.

To ensure efficient energy production, various pathways are in place to balance CAC intermediate efflux. These so-called anaplerotic pathways serve to maintain constant levels of CAC intermediates. Anaplerosis is recognized as a crucial aspect of normal cardiac physiology, however, alterations in anaplerotic pathways have not been investigated in BTHS. This gap in understanding of an important cardiac energy process precludes the development of therapeutics which can improve cardiac function in BTHS patients.

13 13 In this study, we utilized labeled C5-glutamine and C6-glucose tracers to assess the role of glutamine anaplerosis and pyruvate carboxylation to replenish the CAC in

TAZ-iPS-CMs, respectively. Furthermore, we applied mass spectrometry to profile additional anaplerotic amino acids in TAZ-iPS-CMs. Tracer studies with labeled glutamine showed that in the absence of glucose and fatty acids, control and TAZ-iPS-

CMs can metabolize glutamine as a carbon source substrate to replenish the CAC. 13C tracing also revealed that glutamine-derived α-ketoglutarate was metabolized through two distinct pathways; oxidative metabolism and reductive carboxylation pathways, with a slight increase in reductive carboxylation in TAZ-iPS-CMs. The overall contribution of glutamine to α-ketoglutarate was not significantly different between control and TAZ- iPS-CMs. Stable isotope tracing with glucose displayed a 50% decrease in M3 malate, one product of pyruvate carboxylation, in TAZ-mutant cells. Profiling of amino acids also revealed a lower concentration of succinyl-CoA amino acid precursors in TAZ-iPS-

CMs. These data indicate that reduced anaplerosis in TAZ-iPS-CMs could contribute to deficient ATP production in BTHS.

4.2 Introduction

Besides participating in the step-wise oxidation reactions which are necessary for energy production, most citric acid cycle (CAC) intermediates also act as precursors for biosynthetic pathways [1]. To balance the efflux of CAC intermediates, various metabolic reactions serve to replenish the CAC pool. The process of “refilling” the CAC is termed anaplerosis [2]. Anaplerosis is recognized as a crucial aspect of normal cardiac physiology and is also important in replenishing the CAC pool in conditions of aberrant energy metabolism which cause imbalanced levels of CAC intermediates [3][4][5].

Despite the significance of anaplerosis in the heart, a deep understanding of activity and regulation of the specific reactions is lacking [6].

Anaplerotic substrates include propionyl-CoA and its precursors (odd-chain fatty acids, C5 ketone bodies), a range of amino acids, and pyruvate [7][8]. The conversion of

82 glutamine into α-ketoglutarate is an anaplerotic pathway that is important in the kidney and proliferating cells [9][10]. Glucogenic amino acids histidine, proline, arginine, and glutamate can also be metabolized to α-ketoglutarate. Upon CAC entry, α-ketoglutarate is then converted to succinate, producing one molecule of ATP and NADH via substrate- level phosphorylation without contributing to intracellular acidification [11]. This pathway of replenishing the CAC at the α-ketoglutarate level has not been shown to play a significant role in the healthy heart [12]. However, in cardiac pathophysiological conditions when oxygen becomes limited, glutamine as a carbon source becomes more important [13]. Glutamine has demonstrated cardioprotective benefits through its role as an antioxidant and by promotion of mitochondrial respiration in postischaemic reperfusion and other heart conditions [12][14][15]. However, it is less clear whether this is through direct contributions to the CAC, or alternate, non-energy producing pathways such as the pentose phosphate pathway and the hexosamine biosynthetic pathway

[12][16].

Another major anaplerotic pathway is pyruvate carboxylation. Pyruvate can be carboxylated to form oxaloacetate or malate via pyruvate carboxylase or malic enzyme, respectively. Dysfunction of pyruvate carboxylation was shown to lead to rapid decline in cardiac function [17]. Moreover, anaplerotic flux via pyruvate carboxylation was found to be increased in hypertrophied rat and mice hearts [5][18][19]. These studies suggest the importance of this pathway in cardiac pathophysiology.

Besides α-ketoglutarate precursors, additional amino acids act as anaplerotic substrates for other steps of the CAC (Figure 4-17) [1]. The breakdown of amino acids to compensate for the energy deficit of BTHS patients has been postulated considering that

83 one of the predominant clinical features of BTHS is skeletal muscle wasting and furthermore, as elevated whole-body proteolysis was revealed in a small cohort of five

BTHS patients compared to matched healthy controls [20][21]. However, there is very little evidence to support this claim, thus the role of amino acids as anaplerotic precursors in BTHS remains to be determined.

The aim of this study was to elucidate the effect of two TAZ mutations on anaplerotic pathways to provide a better understanding of cardiac disease progression and develop potential therapeutic strategies for BTHS patients. A main goal of this study was to investigate the use of glutamine as a carbon source by TAZ1-iPS-CMs to determine the effect of the TAZ mutation on anaplerosis at the α-ketoglutarate level. We also

13 13 analyzed MID distributions of CAC intermediates after C incorporation from C6- glucose to determine how the TAZ mutation affects the production of CAC intermediates via pyruvate carboxylation. Finally, because amino acids serve as anaplerotic precursors for many CAC intermediates, we used GC-MS and LC-MS/MS to profile selected anaplerotic amino acids in TAZ1-iPS-CMs. We hypothesized that TAZ-iPS-CMs utilize anaplerotic pathways to support CAC function and compensate for decreased energy production due to mitochondrial dysfunction.

Figure 4-17: Major Anaplerotic Pathways of the CAC.

Malic enzyme (ME) and pyruvate carboxylase (PC) catalyze the carboxylation of pyruvate to malate and oxaloacetate, respectively.

4.3 Materials and Methods.

i. iPS-CM Culture

Three cells lines were utilized in this study; control, TAZ1, and TAZ2. iPS-CMs were differentiated as described previously (See Chapter II).

ii. Stable Isotope Labeling

13 iPS-CMs were given RPMI 1640 medium with either 0.5mM C5-glutamine and

13 5mM galactose (without glucose or fatty acids) for 12 hours, or 10mM C6-glucose and

0.4mM palmitate conjugated to bovine serum albumin for 8 hours. Prior to labeling, spent media was removed and cells were washed twice with 1x D-PBS. Tracer media was quickly pipetted onto to cells to begin the labeling period. iii. Metabolite Extraction

After stable isotope labeling, tracer media was aspirated, and cells were washed once with 1x D-PBS and twice with water. Metabolism was quenched by the addition of cold acetonitrile and water (1mL:0.75mL per well). Cells were scraped and transferred to tubes, followed by chloroform addition (1mL). To separate polar and nonpolar phases, cell lysates were centrifuged at 5000rpm for 10 minutes. Polar phases were dried under a stream of nitrogen at room temperature. iv. GC-MS Analysis

After drying, cell extracts were derivatized with methoxyamine (Sigma, cat. no.

226904) in pyridine (20mg/ml, 40μl) for 60 minutes at 80°C, followed by N-methyl-N- tert-butyldimethylsilyltrifluoroacetamide derivatization at 75°C for 45 minutes (Sigma, cat. no. 375934). GC-MS analysis was carried out by GC-MS (Agilent 5977) operated in

EI mode and equipped with an HP-5ms/30 m column (Agilent). Retention times and fragmentation mass spectra of all metabolites were confirmed by an in-house library created from commercially-available standards. Unlabeled (M0) CAC ions and amino acid ions monitored through this method and their retention times are presented in

Appendix B. The GC program was as follows: initial oven temperature was 80°C and held for 3 minutes, then ramped 150°C/minute up to 305°C and held for 3 minutes, for a total run time of 21 minutes.

v. LC-MS/MS Analysis of Arginine

A mixture of labeled amino acids was used as an internal standard for endogenous amino acids (Cambridge Isotope Laboratories, cat.no NSK-A). Dried cell extracts were derivatized using 60μL of 3N hydrochloric acid-n-butanol (Sigma, cat. no. 87472) for 30 minutes at 65°C. Samples were cooled to room temperature and dried again under a stream of nitrogen. Samples were reconstituted in 100μL of mobile phase (90% acetonitrile in water, 0.1% formic acid), then analyzed by flow injection (FIA)MS/MS.

Chemoview 2.2 software (SCIEX) was used to perform data analysis. vi. Calculation of Amino Acid Levels

Amino acid levels were normalized to the reference standard tricarballylic acid

(Sigma, T53503; 10μl of 1mM, m/z 461) and then to the total protein amount of each cell pellet as measured by the Pierce™ BCA Protein Assay Kit (Thermo). Mass spectra were used to calculate the ratio of peak areas of target metabolites to the reference standard.

The calculated ratios were used for crossover analysis to compare arginine and proline

levels in TAZ-mutant vs control cells. In crossover analysis, the relative level of a given

metabolite in the control group is expressed as 100%.

Relative level =

Average [(Peak area of metabolite)/ (Peak area of reference compound)]TAZ

Average [(Peak area of metabolite)/ (Peak area of reference compound)]control

vii. Mass Isotopomer Distribution Analysis

Peak areas of each mass isotopomer were obtained from GC-MS chromatograms

and corrected for natural abundance of all elements contained in the derivatized

compound (See Appendix A). The mass isotopomer distribution (MID) is expressed as

the percent fraction of a specific isotopomer to the sum of all isotopomers of that

compound, including the unlabeled component, M0.

퐶표푟푟푒푐푡푒푑 퐴푟푒푎 표푓 푀푥 푀퐼퐷 = ∑ 푀0 − 푀푛

viii. Statistical Analysis

Data are expressed as the mean + standard deviation. Results were considered

statistically significant when p < 0.05 using the Student’s t-test.

4.4 Results.

Glutamine Anaplerosis into the Citric Acid Cycle. To determine the effect of the

TAZ mutation on glutamine anaplerosis into the CAC, control and TAZ1-mutant iPS-

13 CMs were given 0.5 mM C5-glutamine and 5mM galactose in media free of glucose and fatty acids for 12 hours. In the absence of the heart’s preferred energy sources, both

13 control and TAZ1-mutant cells took up C5-glutamine (Figure 4-18A), which made up

43.9% + 0.87% in control and 37.2% + 1.71% in TAZ1 of the total glutamine pool in cells. Glutamine entry into the CAC produces M5 α-ketoglutarate. Entry of M5 α- ketoglutarate into the CAC produces M4 labeled succinate, fumarate, malate, and citrate via the canonical oxidative pathway (Figure 4-18B), or M5-labeled citrate if reductive carboxylation occurs (Figure 4-18B).

Figure 4-18. Glutamine as a Carbon Source for the Citric Acid Cycle.

A) Percent fraction of mass isotopomers of glutamine in control and TAZ1-iPS- CMs. (B) Representative figure of the observed 13C enrichment patterns after labeling 13 with 0.5mM C5-glutamine and 5mM galactose. Glutamine enters the citric acid cycle at the α-ketoglutarate level, which proceeds through the oxidative pathway (OX) or undergoes reductive carboxylation (RC) to form citrate. Closed circles = 13C; Open circles = 12C.

Reductive Versus Oxidative Metabolism of α-Ketoglutarate. Both control and

13 13 TAZ1-mutant cells produced C-labeled CAC intermediates from C5-glutamine (Figure

4-19A) in addition to M4-labeled aspartate (Figure 4-19B). Mass isotopomer distributions

(MIDs) were calculated for the CAC intermediates αKG, succinate, fumarate, malate, and citrate. Both control and TAZ1 mutant iPS-CMs produced M4 isotopomers of succinate,

13 fumarate, malate, and citrate, representing oxidation of C5-α-ketoglutarate derived from

13 13 C5-glutamine (Figure 4-19A). C5-glutamine contribution to M5 α-ketoglutarate was

10% in both the control and TAZ-iPS-CMs. There was no significant difference in the percent of M5 α-ketoglutarate or the M4 isotopomers in fumarate or malate. The percent fraction of M4 succinate was decreased in TAZ1-iPS-CMs (6.05% + 0.8%) compared to the control (7.72% + 0.5%; p = 0.0476).

M5 citrate is formed via reductive carboxylation of glutamine-derived M5 α- ketoglutarate. MID revealed a fraction of M5 citrate in both control and disease, which was slightly elevated in TAZ-iPS-CM (4.62% + 0.09% in the control and 4.98% + 0.13% in TAZ, p < 0.05) (Figure 4-19C).

A 13C Distribution into CAC B Asparate 14% MID Intermediates Control 12% 8.0% BTHH 7.0% 10% 6.0% ** 8% 5.0% * 6% 4.0% * 3.0%

% of Total Pool Total of % 4% 2.0%

2% 1.0%

0% 0.0% aKG SuccinateFumarate Malate Citrate Citrate M4 M5 M4 M4 M4 M4 M5 Control

Figure 4-19 Mass Isotopomer Distributions of Citric Acid Cycle Intermediates and

Aspartate.

A) 13C distribution into citric acid cycle intermediates following 12 hours of

0.5mM 13C5-glutamine labeling. M4 succinate, fumarate, malate, and citrate represent the oxidative pathway, while M5 citrate derives from reductive carboxylation (p < 0.05). B)

Percent fraction of M4 aspartate (p < 0.005).

Pyruvate Anaplerosis. To assess the effect of TAZ mutations on pyruvate carboxylation and pyruvate oxidation, iPS-CMs were incubated with 10mM 13C-glucose for 8 hours, then MIDs of CAC intermediates were calculated. The fraction of the M2 isotopomer of citrate represents pyruvate oxidation through pyruvate dehydrogenase, while the fraction of the M3 isotopomer of citrate represents pyruvate carboxylation to oxaloacetate and malate.

M3 malate enrichment was found to be 40.5% + 1.5%, 23.4% + 0.6%, and 19.7%

+ 0.5% in control, TAZ1, and TAZ2, respectively (p < 0.0001) (Figure 4-20 A, C). In comparison, M2 malate was calculated as 12% + 1.0%, 13.6% + 0.5%, and 13.6% +

0.2% for control, TAZ1, and TAZ2, respectively, which does not differ by statistical significance. The ratio of M3/M2 malate was decreased by 50% in TAZ-iPS-CMs compared to the control (p = 0.0001).

The enrichment patterns of M3 fumarate were similar to that of M3 malate, with a decrease in M3 labeled in TAZ1 and TAZ2 iPS-CMs (Figure 4-20 C). M3 fumarate was found to be 41.9% + 2.0%, 18.4% + 0.5%, and 19.0% + 0.4% in control, TAZ1, and

TAZ2, respectively (p value < 0.0001).

The difference in percent enrichment of any given M2 isotopomer formed by pyruvate oxidation ranges from 0.1 to 4.6% between the control, TAZ1, and TAZ2 iPS-

CMs. M2 citrate is elevated in both TAZ1 and TAZ2 iPS-CMs compared to the control

(24.1% + 0.4% for control, 26.8% + 0.3% for TAZ1, 28.7% + 0.3% for TAZ2; p value <

0.0007) (Figure 4-20 D)

A Malate MID B Fumarate MID

70% 70% 60% 60% 50% 50% 40% 40% 30% 30% *** 20% *** 20% ****** 10% 10% 0% 0%

Percent Fraction of TotalPool ofFraction Percent M0 M2 M3 M0 M2 M3 Percent Fraaction of TotalPool Fraactionof Percent

Control TAZ1 TAZ2 Control TAZ1 TAZ2

Figure 4-20: M2 and M3 Enrichment Patterns.

A) Malate M0, M2, and M3 from iPS-CMs after labeling with 13C-glucose. (B)

Fumarate M0, M2, and M3 from iPS-CMs after labeling with 13C-glucose. (C) M3 MIDs for citrate, α-ketoglutarate, succinate, fumarate, and malate. (D) M2 MIDs for citrate, α- ketoglutarate, succinate, fumarate, and malate.

Arginine and Proline Levels. Decreased plasma arginine and increased plasma proline are consistent findings in studies of BTHS patients [22][23][24]. We measured levels of arginine and one downstream metabolite, phosphocreatine, in control and

TAZ1-iPS-CMs under standard culture conditions using LC-MS/MS. Proline levels were measured by GC-MS from control and TAZ1-iPS-CMs under standard culture conditions or after 12 hours of glucose deprivation. To compare phenotypic trends of iPS-CM amino acid levels to patient plasma amino acid levels from published studies, the relative levels of arginine and proline in TAZ1-iPS-CMs or BTHS patient plasma were calculated against the control, expressed as 100% (Figure 4-21A). In all studies, arginine levels were decreased. Relative levels of arginine ranged from 43% to 61% of the control levels

(Figure 4-21B). Proline levels in TAZ1-iPS-CMs were 96% of the control levels under standard culture conditions. However, under conditions of glucose-deprivation, proline levels increased to 296% of the control value. In published studies of patient plasma, the relative percent of proline ranged from 177% to 280%. The concentration of proline in

TAZ1-iPS-CMs was 0.44 + 0.08 μM/mg protein in standard conditions and 0.45 + 0.07

μM/mg protein in glucose-free conditions, whereas control iPS-CMs were 0.46 + 0.07

μM/mg protein in standard conditions and 0.15 + 0.02 μM/mg protein in glucose-free conditions (p = 0.0086). Phosphocreatine was decreased by more than 50% in TAZ1- mutant cells (Figure 4-21D). Phosphocreatine was measured as 19.2 + 2.39 μmol/g protein in control and 8.12 mol/g protein in control and 8.12 + 0.78 μmol/g protein in

TAZ1-iPS-CMs (p = 0.0478).

A Amino Acid Concentrations Arginine Proline Current iPS-CM Study Control TAZ Control TAZ Non-starved 5.02 3.05 0.46 0.44 Glucose-Starved not measured not measured 0.15 0.45

Plasma Studies Control BTHS Control BTHS Cade et al 2013† 100 50 190 280 Riguad et al 2013† 68 29 no data no data Vernon et al 2014 69.8 42.9 164.7 291.1 B Relative Percent of Amino Acid Levels in BTHS Compared to Control Arginine Proline Current iPS-CM Study Non-starved 60% 96% Glucose-Starved not measured 296% Plasma Studies Cade et al 2013† ~50% 280% Riguad et al 2013† 43% data not published Vernon et al 2014 61% 177% C D 350% 25.0 296% 300% 20.0 250% 15.0 200% 150% 10.0 * 100% 96% 100% 100% 100% Relative PercentRelative 60% 5.0

50% proteinmicromole/g Proline (Standard) Proline (Glucose- Arginine‡ 0.0 free) Phosphocreatine Control TAZ Control TAZ1

Figure 4-21: Arginine and Proline.

Arginine and proline concentrations (μM/mg protein) measured in iPS-CMs compared to concentrations published in studies of BTHS patients’ plasma (μmol/L). (B) Relative levels of arginine and proline in TAZ-iPS-CMs or BTHS patients compared to control levels. Control levels are expressed as 100%. (C) Bar graphs expressing the relative percent of arginine and proline from iPS-CMs. † = approximate levels. ‡ arginine was measured by LC-MS/MS; proline was measured by GC-MS. (D) Levels of phosphocreatine in iPS-CMs (p < 0.05).

Amino Acid Levels. GC-MS was used to analyze levels of amino acids in TAZ1- iPS-CMs which can serve as anaplerotic precursors. Fumarate precursors phenylalanine and tyrosine were decreased, but the difference was not significant. Only amino acids which are linked to succinyl-CoA anaplerosis differed by statistical significance. A 20% decrease was observed in TAZ-mutant cells for valine (p = 0.0071) and methionine (p =

0.0029).

3.50

3.00 Control 2.50 TAZ

2.00

1.50

Abundance/mg protein Abundance/mg 1.00 *

0.50 **

0.00

Valine

Serine

Glycine

Tyrosine

Aspartate

Isoleucine

Threonine

Asparagine

Methionine Phenylalanine

Fumarate Oxaloacetate Pyruvate Succinyl-CoA

Figure 4-22: Levels of Anaplerotic Amino Acids in iPS-CMs.

Phenylalanine and tyrosine are fumarate precursors. Aspartate and asparagine are oxaloacetate precursors. Glycine and serine are pyruvate precursors. Threonine, methionine, valine, and isoleucine are all precursors to succinyl-CoA. * = p < 0.05; ** = p < 0.005

4.5 Discussion

Glucose-independent glutamine metabolism. Fatty acids and glucose are the preferred carbon sources of the healthy heart [11]. The utilization of these substrates is often altered during cardiac pathogenesis to compensate for the increased demand for

ATP for contractile function [25]. BTHS presents with abnormal energy production as a result of impaired mitochondrial structure and is associated with a decoupling of glycolysis from glucose oxidation [26]. Under these conditions, anaplerotic pathways may function to restore flux through the CAC, bypassing reactions that may be impaired and generating high-energy intermediates through substrate-level phosphorylation. Thus, understanding the ability of anaplerosis to improve ATP production could provide novel therapeutic strategies to improve cardiac status in BTHS patients. However, there is a limited understanding of the role of anaplerotic pathways in cardiac pathologies. The primary goal of this study was to elucidate anaplerotic pathways utilized by control and

13 TAZ-iPS-CMs. Specifically, we used a C5-glutamine tracer to determine the extent of glutamine anaplerosis to the CAC. We also assessed pyruvate carboxylation, a major

13 anaplerotic pathway, using a C6-glucose tracer.

Glutamine is an important biological molecule as it replenishes the CAC, provides carbons for purine and pyrimidine biosynthesis, and many other roles [27][28]. Once glutamine enters the cell, it can act as an anaplerotic precursor of α-ketoglutarate by undergoing conversion to glutamate. Glutamate is metabolized to α-ketoglutarate by transamination or deamination [29]. α-ketoglutarate can then enter two different metabolic pathways; oxidation through the canonical CAC or reductive carboxylation

[30][31]. Glutamine metabolism through reductive carboxylation plays an important role

99 in cancer metabolism, but has also been described previously as a non-significant source of citrate and lipogenic carbon in mammalian cells, such as the kidney and intestine

[32][34][35]. However, there is little evidence demonstrating activity of this pathway in the healthy or diseased heart [12]. To determine the effect of the TAZ1 mutation on

13 glutamine metabolism, we supplemented cells with 0.5 mM C5-glutamine and 5mM galactose in glucose-free and fatty acid-depleted cell media. To compensate for the

13 absence of glucose and fatty acids, control and TAZ1-mutant iPS-CMs metabolized C5- glutamine and produced M4 isotopomers of fumarate, malate, and citrate via oxidative

13 13 CAC. The M4 isotopomers represent oxidation of C5-α-ketoglutarate derived from C5- glutamine (Figure 4-18). There was no significant difference in the percent enrichment of

M5 α-ketoglutarate or the M4 isotopomers of fumarate and malate between control and

TAZ-iPS-CMs, suggesting that flux through the oxidative pathways of the CAC is not affected by the TAZ mutation under the given conditions.

We also found a fraction of M5 citrate in both control and TAZ-iPS-CM lines, providing evidence that glutamine-derived α-ketoglutarate is metabolized via reductive carboxylation (Figure 4-19). The reductive glutaminolysis reaction involves the addition

13 of unlabeled CO2 to C-labeled M5 αKG, resulting in the formation of M5 citrate

[30][32][34]. Under hypoxic stress or impaired mitochondrial function, pyruvate synthesis via PDH is impaired or inhibited, decreasing the amount of acetyl-CoA available for condensation with oxaloacetate for citrate synthesis [33]. In these conditions, reductive carboxylation of glutamine-derived α-ketoglutarate becomes an important pathway for lipogenic citrate [31][32]. There was an increase in M5 citrate in

100

TAZ1-mutant iPS-CMs, which supports the hypothesis that reductive carboxylation is increased in conditions of impaired mitochondrial function.

13 In the absence of glucose and fatty acids, C5-glutamine contributed to the formation of M4 aspartate, which was observed in both control and TAZ1-iPS-CM. The fraction of M4-aspartate was reduced in TAZ1-iPS-CM by 30%, suggesting decreased synthesis of aspartate from glutamine catabolism in glucose-free conditions. This finding is supported by decreased levels of aspartate observed in TAZ1-iPS-CM amino acid profiling (Figure 4-22). This has important implications for energy production, as aspartate is a key metabolite in the malate-aspartate shuttle which essentially functions to carry electrons from reducing equivalents into the mitochondrial matrix [35]. In the event of insufficient aspartate availability, it is plausible that suboptimal electron transfer to the

ETC could occur. Thus, further investigations regarding aspartate synthesis, availability, and function in TAZ-mutant iPS-CMs are warranted.

Pyruvate Anaplerosis. Pyruvate can be converted to lactate in the cytosol, oxidized to form acetyl-CoA, or carboxylated to form oxaloacetate or malate via pyruvate carboxylase or malic enzyme, respectively [30]. Pyruvate carboxylation is an important anaplerotic pathway in the healthy heart [36] and disruption of pyruvate anaplerosis has been shown to cause contractile dysfunction [17][37]. During cardiac disease pathogenesis, alterations in the balance between fatty acid oxidation and glucose oxidation may further result in dysregulation of pyruvate carboxylation. To determine the effect of TAZ mutations on pyruvate carboxylation, we supplemented cells with 10mM

13 C6-glucose media. CAC intermediates generated through pyruvate oxidation are represented by M2 labeling, while pyruvate carboxylation produced M3-labeled CAC

101 intermediates. The enrichment of M3 malate and fumarate in TAZ1 and TAZ2-iPS-CM was decreased by about 50% from the control. This marked decrease in pyruvate carboxylation to malate may be related to the fact that malic enzyme requires NADPH to catalyze the reaction. NADPH is used in fatty acid chain elongation, nucleic acid synthesis, and producing reduced glutathione, an essential cellular antioxidant [3].

Because BTHS is associated with the formation of reactive oxygen species due to mitochondrial dysfunction [38], TAZ-iPS-CMs may preferentially use NADPH to protect against oxidative stress. The similar trends in M3 enrichment patterns between control and TAZ mutants is an evidence of the reverse activity of fumarase catalyzing the reversible reaction of fumarate from malate [39]. M3 oxaloacetate formed via pyruvate carboxylation or from M3 malate oxidation condenses with acetyl-CoA to form M3 citrate (Figure 4-18). M3 citrate in both TAZ-mutant iPS-CM lines under study was decreased 1.4-fold compared to the control, suggesting attenuation of CAC flux.

Arginine as an anaplerotic precursor. A consistent finding across several studies of BTHS patients is reduced plasma arginine [19][29][22]. It has been proposed that low levels of arginine may supplement the CAC via conversion to glutamate [21].

Accordingly, arginine supplementation has been used clinically as a dietary supplement for BTHS patients, although no published studies evaluating its therapeutic effect or its role as an anaplerotic precursor exist. We measured arginine by LC-MS/MS and found that arginine levels were decreased by 40% in TAZ1-iPS-CMs. This marked decrease in arginine levels is consistent with observations in patient plasma samples. In addition to arginine, histidine, proline, and glutamine can also be converted to glutamate for anaplerotic entry into the CAC. Because glutamine is converted directly to glutamate in

102

13 one step, we chose to assess its contribution to the CAC by using a C5-glutamine tracer.

We found that the enrichment of M5 α-ketoglutarate was 10% in both the control and the

TAZ1 mutation. Thus, even in the absence of the major fuel sources glucose and fatty acids, the conversion of glutamine to α-ketoglutarate was not significantly altered between the two cell lines. This finding does not support the notion that α-ketoglutarate anaplerosis is elevated in TAZ1-iPS-CMs, leaving the importance of this anaplerotic entry point unclear. Probing cells directly with a 13C-arginine tracer is thus necessary to determine if arginine flux is increased in TAZ1-iPS-CMs. An alternate explanation for consistent arginine depletion in BTHS patients is that arginine is an important precursor for creatine synthesis [40]. Creatine is phosphorylated by creatine kinase to form phosphocreatine, an essential carrier of high-energy phosphates for rapid ATP production. Our data reveal that phosphocreatine levels are decreased by more than 50%

(Figure 4-21D) while previous studies of BTHS have reported that enzyme levels of creatine kinase are mildly elevated [41]. Taken together, these results suggest that arginine may be redirected toward creatine synthesis in an attempt to correct dysregulation of the creatine-phosphocreatine balance in BTHS.

Amino acids as anaplerotic precursors. To obtain a profile of select amino acids that may play a role in anaplerosis in TAZ-iPS-CMs, we performed GC-MS analysis. We found proline levels were significantly elevated under glucose-free conditions, which is consistent with plasma findings in BTHS patients (Figure 4-21). An increase in proline levels has been reported in several other diseases with mitochondrial dysfunction as well

[42][43][44]. However, there is no clear explanation for this finding. Levels of other amino acids profiled were not significantly different in TAZ1, except for several amino

103 acids which serve as anaplerotic precursors to succinyl-CoA (Figure 4-22). Methionine, valine, and isoleucine were decreased by >15% in TAZ. This is supported by the decreased percent enrichment of M4 succinate observed in glucose-free conditions in

TAZ1-iPS-CMs (Figure 4-18). Entry of unlabeled metabolites at the succinyl-CoA level would increase the amount of M0 succinate, thus diluting the percent of M2 succinate in total pool. This suggests that succinyl-CoA, but not α-ketoglutarate, is an important anaplerotic entry point in TAZ-mutant iPS-CMs. The rapid conversion of succinyl-CoA to succinate involves the substrate-level phosphorylation of GDP to GTP, thus producing energy without contributing to intracellular acidification [45]. Therefore, anaplerosis through this entry-point represents a potentially useful mechanism for energy production in the presence of mitochondrial dysfunction.

Conclusions. The goal of this study was to assess the contribution of alternate metabolites to the CAC beyond fatty acid and glucose oxidation in TAZ-iPS-CMs. We hypothesized that TAZ1-iPS-CMs would utilize glutamine as a carbon source for the

CAC. Tracer studies with labeled glutamine revealed that despite mitochondrial dysfunction, in the absence of glucose and fatty acids, TAZ1-iPS-CMs can rely on glutamine as a carbon source substrate to replenish the CAC. 13C tracing also revealed that glutamine-derived α-ketoglutarate can be metabolized by two pathways in both healthy and TAZ1-mutant iPS-CMs; canonical oxidative metabolism or alternative reductive carboxylation to form citrate.

Our data also reveal major alterations in pyruvate carboxylation in TAZ1 and

TAZ2-iPS-CMs. The 50% decrease in the ratio of M3/M2 malate demonstrates that flux to form malate, either directly (from pyruvate) or indirectly (from pyruvate to

104 oxaloacetate to malate) is attenuated in TAZ-iPS-CMs. Currently, it is unclear which pathway is affected due to several limitations of this study. First, oxaloacetate is unable to be measured by GC-MS. Because OAA and malate are both produced by pyruvate carboxylation but by two different enzymes, enrichment patterns of OAA would be useful for elucidating alterations in enzyme activity. Furthermore, enrichment patterns alone are not able to delineate metabolite flux through any given pathway. To calculate flux, acetyl-CoA would also need to be measured in addition to OAA [46]. This would allow for a more detailed comparison of pyruvate oxidation versus pyruvate carboxylation in TAZ1 and TAZ2.

Finally, profiling of amino acids revealed a potential role for succinyl-CoA precursors to support energy production in TAZ, as methionine, valine, and isoleucine were decreased in TAZ1-iPS-CMs. Tracer studies at the acetyl-CoA level are needed to determine if the decrease in the levels of these amino acids is due to their catabolism into the CAC.

4.6 References

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[5] N. Sorokina et al., “Recruitment of Compensatory Pathways to Sustain Oxidative Flux With Reduced Carnitine Palmitoyltransferase I Activity Characterizes Inefficiency in Energy Metabolism in Hypertrophied Hearts,” Circulation, vol. 115, no. 15, pp. 2033–2041, Apr. 2007. [6] C. Des Rosiers, F. Labarthe, S. G. Lloyd, and J. C. Chatham, “Cardiac anaplerosis in health and disease: food for thought,” Cardiovasc. Res., vol. 90, no. 2, pp. 210– 219, May 2011. [7] H. Brunengraber, R. A. Ibarra, G.-F. Zhang, S. Deng, V. E. Anderson, and Q. Li, “Multiple Mass Isotopomer Tracing of Acetyl-CoA Metabolism in Langendorff- perfused Rat Hearts,” J. Biol. Chem., vol. 290, no. 13, pp. 8121–8132, 2015. [8] H. Brunengraber and C. R. Roe, “Anaplerotic molecules: Current and future,” J. Inherit. Metab. Dis., vol. 29, no. 2–3, pp. 327–331, 2006. [9] B. Vercoutère, D. Durozard, G. Baverel, and G. Martin, “Complexity of glutamine metabolism in kidney tubules from fed and fasted rats.,” Biochem. J., vol. 378, no. Pt 2, pp. 485–95, Mar. 2004. [10] M. G. Vander Heiden, L. C. Cantley, and C. B. Thompson, “Understanding the Warburg Effect: The Metabolic Requirements of Cell Proliferation,” Science (80-. )., vol. 324, no. 5930, pp. 1029–1033, May 2009. [11] H. Taegtmeyer et al., Assessing Cardiac Metabolism, vol. 118, no. 10. 2016. [12] B. Lauzier et al., “HHS Public Access,” pp. 92–100, 2015. [13] G. Marazzi, S. Rosanio, G. Caminiti, F. S. Dioguardi, and G. Mercuro, “The role of amino acids in the modulation of cardiac metabolism during ischemia and heart failure.,” Curr. Pharm. Des., vol. 14, no. 25, pp. 2592–604, 2008. [14] T. T. Nielsen, N. B. Stottrup, B. Lofgren, and H. E. Botker, “Metabolic fingerprint of ischaemic cardioprotection: importance of the malate-aspartate shuttle,” Cardiovasc. Res., vol. 91, no. 3, pp. 382–391, Aug. 2011. [15] S. L. Badole, G. B. Jangam, S. M. Chaudhari, A. E. Ghule, and A. A. Zanwar, “L- Glutamine Supplementation Prevents the Development of Experimental Diabetic Cardiomyopathy in Streptozotocin-Nicotinamide Induced Diabetic Rats,” PLoS One, vol. 9, no. 3, p. e92697, Mar. 2014. [16] K. J. Drake, V. Y. Sidorov, O. P. Mcguinness, D. H. Wasserman, and J. P. Wikswo, “Amino acids as metabolic substrates during cardiac ischemia,” Exp. Biol. Med., vol. 237, no. 12, pp. 1369–1378, Dec. 2012. [17] K. E. Sundqvist, J. K. Hiltunen, and I. E. Hassinen, “Pyruvate carboxylation in the rat heart. Role of biotin-dependent enzymes.,” Biochem. J., vol. 257, no. 3, pp. 913–6, Feb. 1989. [18] K. M. Pound et al., “Substrate–Enzyme Competition Attenuates Upregulated Anaplerotic Flux Through Malic Enzyme in Hypertrophied Rat Heart and Restores Triacylglyceride Content,” Circ. Res., vol. 104, no. 6, pp. 805–812, Mar. 2009. [19] S. C. Kolwicz et al., “Cardiac-specific deletion of acetyl CoA carboxylase 2

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CHAPTER V

CALCIUM CHANNEL EXPRESSION IN TAZ-MUTANT INDUCED PLURIPOTENT STEM

CELL-DERIVED CARDIOMYOCYTES

5.1 Abstract

Barth syndrome (TAZ, G4.5, OMIM 300394) is clinically manifested by a variety of cardiac pathophysiologies, including ventricular arrhythmia. Cardiomyocyte contraction and relaxation are tightly linked to calcium (Ca2+) transients. Defects in Ca2+ cycling can result in improper contraction or arrhythmia. We measured the expression of key calcium handling genes encoding excitation-contraction coupling in an induced pluripotent stem cell derived-cardiomyocyte model (iPS-CM) of Barth syndrome

(BTHS). Our data reveal involvement of calcium homeostasis genes in the TAZ1-iPS-

CM phenotype and indicate that disrupted calcium cycling may contribute to the cardiac complications of BTHS patients. Targeting the expression of these genes or proteins could represent novel therapeutic targets for optimizing cardiac function in BTHS- affected individuals.

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5.2 Introduction to Calcium Channel Proteins

Barth syndrome (BTHS) is an X-linked recessive disorder associated with cardiomyopathy, sudden cardiac death, neutropenia, exercise intolerance, skeletal muscle weakness, recurrent bacterial infections, and growth delay [1]. The manifestation of cardiac disease is the leading cause of mortality in BTHS patients. BTHS patients can present with multiple cardiovascular complications, including left ventricular noncompaction, hypertrophic cardiomyopathy, endocardial fibroelastosis, prolonged QTc interval, and ventricular arrhythmia, with the most prominent and severe presentation being dilated cardiomyopathy (DCM), manifesting in the first year of life for 70% of patients [1][2][3]. Although DCM is well-documented in BTHS patients, recent studies provide evidence that arrhythmia is also a prevalent and significant clinical manifestation, such as the case study by Spencer et al. (2005) which revealed that 43% of adolescent patients had documented ventricular arrhythmia, independent of the severity of cardiac dysfunction or dilation [4].

Currently, there is no specific therapy for the cardiac manifestations of BTHS, as the pathogenesis of the disease is poorly understood. Supportive management includes various standard treatments of heart failure, such as β-blockers, angiotensin-converting enzyme inhibitors, diuretics, and cardiac glycosides [5]. Although these medications are generally effective, cardiac failure may occur suddenly, resulting in the need for a heart transplantation [1]. The use of antiarrhythmic medication as a prophylactic measure has not been investigated for BTHS patients [6]. To prevent or mitigate cardiac dysfunction, it is critically important to elucidate precise mechanisms of cardiac dysregulation and arrhythmia underlying the cardiac pathophysiology of BTHS.

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Cardiomyocyte contraction and relaxation are tightly linked to calcium (Ca2+) transients [7]. Defects in Ca2+ cycling can result in improper contraction or arrhythmia.

Contraction is initiated by an action potential along the sarcolemma of a cardiomyocyte, which stimulates an influx of Ca2+ via gated L-type Ca2+ channels along the membrane. In a process known as Ca2+-induced Ca2+-release, Ca2+ is released into the cell from the sarcoplasmic reticulum via ryanodine receptors (RYR2), resulting in contraction. For relaxation to occur, cytosolic Ca2+ concentrations must be restored to pre-excitation levels

[8]. One mechanism for Ca2+ removal is reuptake via the sarcoendoplasmic reticulum

Ca2+ ATPase (SERCA2a), encoded by the gene ATP2A2. SERCA2a plays a fundamental role in regulating cytosolic Ca2+ concentrations via reuptake of Ca2+ into the sarcoplasmic reticulum. Ca2+ is also eliminated from the cytosolic space via the Na+/Ca2+ exchanger

(NCX; gene name SLC8A1).

RYR2, SERCA2a, and NCX are the primary channels responsible for Ca2+ homeostasis within the cardiomyocyte (Figure 5-23) [9]. Altered expression or improper function of these three genes have been shown to play a role in pathogenesis of various cardiac diseases. Decreased SERCA2a expression and activity has been described in heart failure, cardiac hypertrophy, and DCM [10][11][12]. Several studies reported that dysregulation of Ca2+ homeostasis as a result of RYR2 mutations lead to arrhythmogenic right ventricular dysplasia [13][14], while aberrant NCX activity and expression has been identified as a contributing factor in heart failure, ischemia, and diabetic cardiomyopathy

[15][16].

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Figure 5-23: Calcium Handling

Ca2+ enters the sarcolemma via L-type Ca2+channel after stimulation by an action potential. This influx of Ca2+ triggers the release of Ca2+ from RYR2, which allows the sarcomere to contract. Removal of Ca2+ from the intracellular space via uptake by

SERCA2a or efflux through NCX results in relaxation. Modified from [20].

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It has been proposed that ventricular arrhythmia seen in BTHS patients [17] could be a result of disturbed Ca2+ homeostasis, the key process underlying excitation-contraction coupling [18]. In addition to its essential role in systolic and diastolic functions, Ca2+ is critically important for mitochondrial signaling, including stimulation of key citric acid cycle dehydrogenases, prevention of excessive ROS production, and ROS elimination

[19]. The interplay between the multiple functions of Ca2+ is especially important in

BTHS, as it is associated with mitochondrial energetic abnormalities, oxidative stress, and contractile function defects [18][20][21].

Human induced pluripotent stem cells (iPSCs) have been established as a useful tool for in vitro modeling of cardiac channelopathies, including familial hypertrophic cardiomyopathy and arrhythmogenic right ventricular dysplasia [23][24].

Cardiomyocytes derived from iPSCs maintain important phenotypic characteristics, including contractile function, action potentials, and the expression of ion channels, receptors, and transporters [25][26][27][28], in contrast to transgenic animal models in which notable species-specific differences exist. This enables more precise investigations of molecular mechanisms of the cardiac features of BTHS. The objective of this study was to examine the effect of the TAZ1 mutation on key calcium regulatory genes involved in the contraction cycle of an induced pluripotent stem cell-derived cardiomyocyte (iPS-CM) model of BTHS. Elucidation of the molecular mechanisms underlying regulation of calcium homeostasis in BTHS may lead to the development of new therapeutic approaches for this disease.

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5.3 Materials and Methods.

i. Differentiation

iPS-CMs were differentiated from iPSCs carrying the TAZ1 mutation (frameshift c.517delG) [29] and an isogenic control as described in Chapter II. Prior to differentiation, iPSCs were analyzed for the pluripotency markers SSEA4 and OCT2.

Differentiation was carried out via modulation of the Wnt/β catenin pathway by an optimized small molecule protocol [30]. Following differentiation, iPS-CM colonies were purified for 48 hours by incubation in glucose-depleted, lactate (4mM) enriched media and were harvested after 45 days.

ii. iPS-CM Immunocytochemistry

Immunostaining of iPS-CMs for α-actinin was performed as described previously

(See Chapter II). iii. Quantitative Real-Time PCR

Total RNA from independent biological triplicates was extracted using Trizol according to the manufacturer's instructions. RNA quality was assessed by NanoDrop. cDNA was synthesized using GeneAmp RNA PCR Core Kit (Applied Biosystems). Real- time qPCR was carried out using TaqMan Assays-on-Demand Probe technology

(Applied Biosystems). The following probes were used (Thermo): ATP2A2

Hs00544877_m1, SLC8a1 Hs1062258_m1, RYR2 Hs00181461_m1. 18s rRNA

(4352930, Applied Biosystems) was used as a reference gene. Relative expression levels were calculated as 2−ΔΔCT.

114 iv. Western Blot

Protein was extracted with RIPA buffer supplemented with protease and phosphatase inhibitors (Halt™ Protease Inhibitor Cocktail and Halt Phosphatase Inhibitor

Cocktail; Fisher Scientific). Samples were resolved on a 10% (w/v) acrylamide gel and transferred onto a nitrocellulose membrane. 5% non-fat dry milk was used for blocking.

Membranes were incubated overnight with primary antibodies (1:1000), washed, and incubated with HRP-conjugated secondary antibodies. Proteins were detected by exposure to ECL and visualized on XR-B x-ray film. Band intensities were quantified using ImageJ 1.40g software (Wayen Rasband, NIH, USA). Primary antibody anti-

SERCA2A antibody was purchased from Abcam (ab2861) and anti-β-actin antibody was purchased from Santa Cruz (sc-47778).

v. Statistical analysis

Data are expressed as mean ± SEM. Results were considered statistically significant when p < 0.05 using the Student's t-test.

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5.4 Results:

Sarcomeric Structure. Calcium binding and release in the sarcomere is responsible for cardiomyocyte contraction and relaxation. To investigate the role of calcium handling proteins in BTHS, induced pluripotent stem cells bearing a TAZ1 mutation were differentiated to cardiomyocytes. iPS-CMs exhibited sustained beating and displayed sarcomeric organization as indicated by the presence of regularly-spaced z- bands after α-actinin immunostaining (Figure 5-24). Immunostaining for the adult isoform of cardiac troponin, TNNI3, also pointed to regular organization of sarcomeric thin filaments (Figure 2-4 and 2-5).

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Figure 5-24: α-Actinin Immunostaining.

Anti-α-actinin staining demonstrates the presence of sarcomeric organization in

TAZ1-iPS-CM. Green = α-actinin; Blue = DAPI. 60X magnification.

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Calcium Channel Gene Expression. After 45 days in culture, beating iPS-CMs were harvested to analyze mRNA levels of the genes encoding SERCA2A, RYR2 and

NCX1 (gene names ATP2A2, RYR2 and SLC8a1 respectively), using real-time PCR. The data show that levels of all three genes are notably decreased in TAZ1-iPS-CMs compared to the isogenic control (Figure 5-25). Specifically, TAZ1-iPS-CMs displayed a four-fold decrease in ATP2A2 (p<0.05) and a five-fold decrease in RYR2 mRNA expression (p<0.0005) compared to the isogenic control. Notably, SLC8a1 exhibited a twelve-fold decrease in mRNA expression (p<0.0005).

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Average Relative Protein Relative Gene Sample N mRNA SEM p value Target mRNA Range Level Control 5 1.00 - 3.56 0.572 2.04 0.0265 SERCA2a ATP2A2 TAZ 5 0.05 - 0.65 0.111 0.459 Control 5 0.97 - 1.23 0.078 0.988 0.0003 RYR2 RYR2 TAZ 3 0.09 - 0.26 0.05 0.176 Control 3 0.86 - 1.06 0.059 0.975 0.0001 NCX SLC8A1 TAZ 3 0.053 - 0.099 0.014 0.081

Figure 5-25: SERCA2a2, RYR2, and NCX mRNA.

Relative mRNA expression levels normalized to 18S of three key calcium channels in iPS-cardiomyocytes. ATP2A2 encodes for sarcoendoplasmic reticulum Ca2+

ATPase (SERCA2a) and SLC8A1 encodes for Na+/Ca2+ exchanger (NCX1). RYR =

Ryanodine Receptor. Data presented + standard error of the mean (SEM). * = p < 0.05;

*** = p < 0.0005.

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SERCA2a Protein Expression. To measure the levels of SERCA2a protein, beating iPS-CMs were harvested, lysed, and analyzed by Western blotting in independent triplicates. In contrast to the significant difference observed in ATP2A2 mRNA expression level, SERCA2a protein expression in TAZ1-iPS-CMs was mildly and insignificantly decreased. Semi-quantitation of protein bands by densitometry showed a relative expression of 0.738 + 0.091 for control iPS-CM and 0.626 + 0.018 (p = 0.264) for TAZ1-iPS-CM when normalized to the band intensity of the housekeeping gene β- actin (Figure 5-26).

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SERCA2a 0.900 0.800 0.700

0.600

Actin - β 0.500 0.400 0.300

ODSERCA/ 0.200 0.100 0.000 Control TAZ

Figure 5-26: SERCA2a2 Protein Expression

Semi-quantitative and qualitative SERCA2a expression levels by Western blotting. Β-actin was used as a loading control. Data presented + standard error of the mean (SEM). p > 0.05.

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5.5 Discussion

Although much work has been done to characterize the clinical heterogeneity of

BTHS, the mechanisms underlying the progression of cardiac disease in BTHS patients remain poorly understood. Wang et al demonstrated that TAZ1-iPS-CMs display defective contractility, which is thought to be at least partly a result of impaired sarcomere assembly [9]. In their model, ATP availability was not shown to play a role in contractile dysfunction, suggesting that an alternate mechanism, not energy depletion, affects contractility. Alternatively, it has been proposed that defective cardiolipin could hinder signaling between the sarcoplasmic reticulum and the mitochondria in BTHS, resulting in dysfunctional calcium (Ca2+) handling, and thus, aberrant contractility [31].

In fact, impaired sarcoplasmic reticulum Ca2+ handling has been implicated in various cardiac diseases [32]. The alteration in the expression or activity of proteins involved in cytosolic and sarcoplasmic reticulum Ca2+ handling is now recognized as a major contributor to numerous cardiac pathophysiologies including disrupted contraction and relaxation, dilated cardiomyopathy, cardiac hypertrophy, arrhythmogenesis, and heart failure [33]. Ca2+ also plays a role in mediating energy production and antioxidative capacity in mitochondria [34][35]. Since BTHS presents with various forms of heart disease and multiple mitochondrial dysfunctions, calcium-mediated processes may play a role in disease pathogenesis. However, to the best of our knowledge, no research groups have published data to support these hypotheses. This study aimed to establish how the

TAZ1 mutation affects the expression of key channels involved in excitation-contraction coupling and Ca2+ homeostasis, providing novel insight into potential therapeutic targets for BTHS.

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Ryanodine receptor 2 (RYR2), sarco/endoplasmic reticulum Ca2+ ATPase 2a

(SERCA2a), and the sodium-calcium exchanger (NCX1) are the primary channels responsible for Ca2+ homeostasis within the cardiomyocyte [9]. Altered expression or improper function of these three genes have been shown to play a role in pathogenesis of various cardiac diseases. To assess the effect of TAZ1 mutation c.517delG on expression of these genes, mRNA from isogenic control and TAZ1-mutant iPS-CMs was analyzed after 45 days in culture.

In our model, we found that TAZ1-iPS-CM exhibit decreased SERCA2a (gene name ATP2a2) expression at the mRNA level (Figure 5-25). Reduced Ca2+ efflux from the cytosol as a result of decreased SERCA2a activity prevents relaxation and consequently leads to a decrease in cardiomyocyte contraction [36]. Thus, targeting

SERCA2a gene expression has been a proposed therapeutic strategy for modulating heart failure and cardiomyopathy [37][38][39]. In an iPS-CM model of DCM (TNNT2

R173W), Sun et. al demonstrated that over-expression of SERCA2a enhanced cardiomyocyte contractility. We propose that targeting SERCA2a expression may be a potential mechanism to improve cardiomyocyte contractility in the TAZ1-iPS-CM model.

It is well-documented in some heart failure conditions, there is a decrease in sarcoplasmic reticulum Ca2+ uptake related to decreased SERCA2a activity or expression

[10][11][12]. Additionally, SERCA2a protein is a major consumer of ATP in the heart.

Thus, we further examined SERCA2a protein expression in order to establish how the

TAZ1 mutation might impact this protein target. As indicated by Western blot, expression of SERCA2a protein in TAZ1-iPS-CM was lower than in control cells (Fig.1e), however, the difference was not found to be significant. Although SERCA2a protein does not seem

123 to support the results of SERCA2a (ATP2a2) mRNA levels, it is important to note that only protein expression levels, but not activity levels, were measured. Thus, studies which determine SERCA2a protein activity and Ca2+ handling in TAZ1-iPS-CM are warranted.

The observed reduction of SERCA2A mRNA expression in TAZ1-iPS-CM is consistent with data reported for other cardiac functional abnormalities such as heart failure, hypertrophy, and dilated cardiomyopathy, and suggests that low SERCA2A expression contributes to the contractile dysfunction [33] observed in TAZ1-iPS-CMs

[29]. The functional consequences of low NCX1 mRNA levels in TAZ1-iPS-CMs are more controversial, as NCX1 expression in different cardiac pathologies is correlated to impaired Ca2+ handling, but the data regarding its relative expression are not consistent.

Multiple studies in human models of heart failure report increased or unchanged NCX1 expression [40][41]. In adult rat cardiomyocytes, the downregulation of NCX1 leads to a decrease in Ca2+ influx and efflux [42],[9] and overexpression of the gene in postinfarction rat myocytes rescued contractile dysfunction [43]. In our model, NCX mRNA was found to be significantly downregulated twelve-fold in TAZ1-iPS-CMs. To make sense of these discrepancies, NCX1 expression has to be considered with the associated changes in Na+/Ca2+ transients as well as RYR2 and SERCA2a expression, which work in tandem with NCX1 and strongly affect cytoplasmic Ca2+ levels. Moreover, NCX1 protein expression and activity levels should also be evaluated. NCX1 typically operates in the forward mode to promote Ca2+ efflux but may also function in the reverse mode to import Ca2+ to the intracellular space. Presently, it is unclear how NCX1 function is

124 affected by the TAZ1 mutation and further studies that include calcium transients and sodium currents need to be performed.

TAZ1-iPS-CMs exhibited low RYR2 mRNA expression levels. RYR2 plays a significant role in regulating heart rate by releasing Ca2+ from the SR to cytosol [44].

Numerous groups have studied RYR2 mRNA expression in heart failure, however, results have not been consistent [45][46]. Decreased SR Ca2+ related to RYR2 is generally thought to be a result of ‘leaky’ RYR2 channels, or dysfunction of its regulatory components such as S100A1, calmodulin, and FKBP12.6 [47][48][49]. It is well-established that RYRs are sensitive to oxidation status, which can cause either activation or inactivation of the RYR2 protein [47]. Given that excessive ROS production was measured in TAZ1-iPS-CM (Wang et al), it is plausible that ROS status may play a role in influencing the expression of this gene. Decreased RYR2 mRNA expression observed in TAZ1-iPS-CM may also be associated with an alteration of the frequency and amplitude of cytosolic and mitochondrial Ca2+ signals [50], simultaneously causing metabolic disturbances [51]. In addition to expression levels, measuring Ca2+ transients would be useful to determine if ‘leaky’ RYR2 channels are a factor in TAZ1-induced cardiomyopathy.

Given the downregulation of SERCA2a, RYR2, and NCX1 mRNA expression, our findings suggest that the cardiac contractility defects observed in BTHS may be due to impaired Ca2+ handling. Several mechanisms could explain impaired Ca2+ handling based on these results, including insufficient reuptake of Ca2+ into the SR, preventing relaxation. Similarly, downregulated NCX1, resulting in decreased Ca2+ efflux, could cause high concentrations of intracellular Ca2+. Further studies are needed to understand

125 how gene and protein expression of these Ca2+ channels are affected by TAZ mutations.

Here, we have demonstrated that TAZ1 mutation c.517delG causes downregulation in the gene expression of important protein channels involved Ca2+ homeostasis, including

SERCA2a, RYR2, and NCX1, representing novel therapeutic targets for the cardiac symptoms of BTHS.

Because of the high risk of cardiac death associated with BTHS, identification of precise targets for potential treatment strategies are needed. Current therapeutic strategies for the cardiac features of BTHS are mostly limited to standard heart failure medications, such as β-blockers, angiotensin-converting enzyme inhibitors, diuretics, and cardiac glycosides [5]. These medications seem generally effective in prolonging the lives of

BTHS patients, but no medication has shown to be particularly advantageous and sudden cardiac death may occur despite months of stability [1]. Because BTHS presents with various forms of heart disease and multiple mitochondrial dysfunctions, calcium- mediated processes represent new potential therapeutic targets.

Therapeutics which aim to restore calcium handling in the sarcoplasmic reticulum are currently being investigated for other types of heart diseases. SERCA2a gene therapy has shown promise in reducing the risk of arrhythmias and heart failure in preclinical trials [52][53]. Istaroxime, a modulator of calcium ATPases, showed promise in a phase

II clinical trial in heart failure patients [54][55]. Istaroxime increases SERCA2a activity, promoting calcium reuptake into the sarcoplasmic reticulum [56]. An additional advantage to this therapeutic agent is that stimulating SERCA2a activity indirectly benefits ATP utilization, as less ATP is needed for SERCA2a function compared to Ca2+

126 removal via NCX1 [57]. These therapeutic agents could be considered in future studies to investigate their effects on Ca2+ channel expression and Ca2+ transients in TAZ-iPS-CMs.

One of the limitations of this study is that we have focused on only one specific

TAZ1 mutation (c.517delG), while 120 Barth syndrome-causative TAZ mutations have been identified. Protein expression of RYR2 and NCX1 and Ca2+ transient studies in vitro are also warranted to further explore TAZ-induced functional abnormalities resulting in the cardiac phenotype of BTHS.

Another limitation of this study, and virtually all past and present studies using iPS-CMs, is that iPS-CM models are not yet able to reach the full maturation status of human adult cardiomyocytes [58]. Ca2+ handling kinetics in iPS-CMs are slower than adult cardiomyocytes, more closely resembling a fetal phenotype [28]. In the present study, many iPS-CMs displayed an elongated shape which resembles that of adult cardiomyocytes, while a subset of iPS-CMs had a more rounded, immature morphology.

Regardless of morphology, iPS-CMs in the present study displayed sarcomeric organization as indicated by α-actinin staining. Despite universal ongoing challenges to obtain a more mature phenotype, iPS-CMs are regarded as a valuable model for in vitro modeling of cardiac diseases [59]. Moreover, the use of an isogenic control in these studies gives confidence that any observed effects are due exclusively to the TAZ1 mutation.

This study revealed alterations in the expression of key components of the excitation-contraction coupling process in the presence of a TAZ1 mutation for the first time. Taken together, our data indicate that a mutation in the TAZ1 gene leads to alterations in genes that encode for calcium cycling regulatory proteins, and

127 consequently, point to the involvement of these genes and disrupted calcium cycling in the cardiac phenotype of BTHS. Our data are in agreement with multiple studies demonstrating alterations in expression of Ca2+ homeostasis genes in a variety of cardiac dysfunctions [60], implicating these genes and related proteins as new targets to explore for the treatment of BTHS.

5.6 References

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CHAPTER VI

GENERATION OF INDUCED PLURIPOTENT STEM CELL-DERIVED

CARDIOMYOCYTES IN 3D ON A 384-MICROPILLAR PLATE

6.1 Abstract

Despite the global burden of heart disease incidence and mortality, there has been a decline in the development and FDA-approval of cardiovascular therapeutics over the past two decades. One contributing factor is the lack of in vitro models which accurately predict cardiotoxicity of drug candidates. Many in vitro models rely on 2D, monolayer cells which are unable to accurately mimic the complexity of biological systems, thus providing a poor predictive measure when translated to in vivo studies.

Efficient differentiation methods have been described for producing iPS-CMs of specific subtypes with high efficiency, but many protocols have been developed in the 2D format, which poses a significant challenge to the application of induced pluripotent stem cell-derived cardiomyocytes (iPS-CMs) for disease modeling and drug discovery.

Culturing cells in 3D may allow maturation of iPS-CMs from a fetal to an adult-like phenotype. Thus, the aim of this study was to develop a 3D platform for generation of mature iPS-CMs which demonstrate enhanced structural and functional parameters

133 compared to 2D cells. Here, we describe a novel system for generating induced pluripotent stem cell-derived cardiomyocytes in 3D on a 384-micropillar chip.

6.2 Introduction

Heart disease is one of the leading causes of morbidity and mortality worldwide

[1]. Despite the prevalence and severity of heart disease, the rate of drugs receiving FDA approval has declined. Part of the reason for the lack of new therapeutics reaching the market is unexpected cardiotoxicity, which was responsible for 28% of all drug withdrawals in the U.S [2]. This includes drugs for cardiovascular disease, as well as any drug causing unexpected cardiotoxicity. Thus, besides identifying novel therapeutics for heart disease, there is a need for a reliable screening platform to evaluate the safety and efficacy of any drug candidate hoping to receive FDA clearance [3].

The development of induced pluripotent stem cell-derived cardiomyocyte (iPS-

CMs) technology has ushered in new hopes and expectations for in vitro modeling of diseases [4]. Due to advancements in genomic editing, iPS-CMs enable the study of familial cardiomyopathies with isogenic controls. This has greatly expanded the possibilities for drug discovery and risk assessment [5]. However, 2D monolayer differentiation protocols have become more popular than 3D cultures due to ease of handling [3]. These monolayer cell cultures often fail to reliably mimic in vivo responses in response to drug candidates, due to lack of 3D organization, altered interactions with cell matrices, minimized cell-to-cell interactions, and other differences in the cellular microenvironment [6]. Thus, 2D monolayers have poor predictive value for evaluating risk of cardiotoxicity, partially contributing to the failure of drugs in vivo which showed promise in vitro. Moreover, despite much progress, no single method exists which can

134 produce iPS-CMs which achieve the structural and functional maturity of human adult cardiomyocytes [7][8].

A few studies have demonstrated the potential for improving iPS-CM maturity using 3D culture methods [9][10]. Previous attempts to establish 3D iPS-CM cultures include the use of iPSC embryoid body formation or various bioengineering strategies, such as fibrin-based engineered heart tissues (EHTs), cardiac microtissues (CMTs), and cardiac biowires [3]. Use of embryoid bodies is associated with a lack of reproducibility in iPS-CM differentiation, due to difficulty in generating uniformly-sized embryoid bodies [11]. Small embryoid bodies are susceptible to cell death, while large embryoid bodies may develop necrotic cores, thus, irregular embryoid body sizes introduces biological variability [12]. Currently available bioengineering platforms for 3D cell cultures seem to improve size homogeneity but lack microscalability, and thus are associated with high costs due to large volumes of materials and reagents needed [13].

Moreover, this precludes their usefulness for high throughput screening and high content imaging (HCI) assays. HCI grants the ability to assess multiple phenotypic parameters such as cell viability, mitochondrial integrity, membrane integrity, ATP levels, DNA damage, and oxidative stress in a high-throughput format using a number of fluorescent dyes or fluorescent probes, thus allowing improved characterization and quantitation of drug toxicity [14].

The primary goal of this study was to generate a 3D iPS-CM platform which could produce mature iPS-CMs. We utilized 384-micropillar plates with sidewalls

(Figure 6-27) using hydrogels to encapsulate cells via two bioprinting methods. 384- micropillar plates require microliter volumes of cells and reagents, thus minimizing costs.

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Importantly, 384-micropillar plates can be sandwiched with standard 384-well plates, allowing for high-throughput (HTP) and high-content imaging (HCI) analyses.

Here, we describe successful miniaturization of 3D iPS-CM culture on a 384- micropillar plate. The use of ultra-low attachment plates enabled the generation of spheroids of uniform size and shape, thus improving homogeneity across the system. We also demonstrate the potential of our platform to be used for HCI assays and metabolomics using mass spectroscopic techniques. We hypothesize that this system will enable the production of mature iPS-CMs, which can ultimately be used for HTP drug discovery and assessments of cardiotoxicity.

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Figure 6-27: Renderings of the 384-Pillar Plate.

(A) Top view of the 384-pillar plate with side walls. Inset zoomed to show details of the pillar structure. (B) Side view of the 384-pillar plate with side walls.

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6.3 Methods

i. Monolayer Cell Culture

Three cell lines were used in this study. The control and TAZ1 lines are isogenic, while the WT2 is a control line from an independent healthy donor. Before miniaturizing cell cultures, iPSCs were first proliferated in monolayers as described previously (See

Chapter II). At passage 20, cells were detached from the plate using 0.48mM EDTA tetrasodium in phosphate buffered saline (PBS). Cells were resuspended in Essential 8

Flex media, and the resulting cell suspension was used to plate cells onto 384-pillar microchips with side walls or into ultra-low attachment 384-well plates.

Line Name Source Mutation Control Harvard Medical Center/Fibroblasts/Male Wild-type WT2 Stanford Medical Center/Fibroblasts/Male Wild-type TAZ1 Harvard Medical Center/Fibroblasts/Engineered C.517delG

ii. iPSC Encapsulation onto a 384-Sidewall Pillar Chip- “Direct Method”

384-sidewall pillar chips were prepared ahead of cell suspension preparation. To prepare the 384-sidewall pillar chip, the chip was first coated with 0.01% polymaleic anhydride alt‑1‑octadecene (Sigma, 419117) and dried for 4 hours. Next, 3 μL of

0.0033% of poly-L-lysine and 25mM calcium chloride were added to the chip and dried overnight. 3μL of a mixture of 1:2 Matrigel and PBS was printed onto each pillar of the chip and dried overnight. A new cell suspension of 3 × 106 cells/mL was made in 3 mg/mL Matrigel, alginate, or a combination of Matrigel and alginate were used to resuspend cells after spinning down the suspension and removing Essential 8 Flex media.

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3 μL of the cell-hydrogel suspension was printed onto each pillar of a 384-pillar chip using a bioprinter (S+ Microarrayer). The 384-sidewall pillar chip was sandwiched into a

384-well plate containing Essential 8 Flex Media. Cells were incubated for 10 days to allow for proliferation into spheroids, changing the media every other day until differentiation was initiated. iii. iPSC Spheroid Formation in ULA Plates- “ULA Method”

384-pillar chips with sidewalls were prepared ahead of cell suspension preparation. To prepare the 384-pillar chip, the chip was first coated with 0.01% polymaleic anhydride alt‑1‑octadecene and dried for 4 hours, then coated with 4μL of

5mg/mL Matrigel and dried overnight. Before transferring cells onto the microchip,

3x106 cells/well were seeded into a Corning® 384-well round bottom ultra-low attachment (ULA) plate in 80 µL media and incubated for 7 days, changing the media every 2 days to allow for spheroid formation. To transfer spheroids from the ULA plate to the 384-pillar plate, the 2.5mg/mL Matrigel-coated plate was stamped with the ULA plate, inverted, and incubated at room temperature for 25 minutes to allow for gelation.

After gelation, the 384-pillar chip was stamped with a 384-well plate containing Essential

8 Flex Media and incubated further at 37 ºC for 3 days before differentiation was initiated. iv. 3D Differentiation

Differentiation of iPSCs on 384-pillar plates with side walls was carried out as described previously. Briefly, to induce cardiac differentiation, cells were cultured in

RPMI-1640 medium supplemented with B27 minus insulin for 8 days. Cells were treated with 9μM CHIR 99021 (Cayman, 13122) on differentiation day 0 for 24 hours. Cells

139 were treated with 10μM IWR1-endo (Cayman, 13659). On differentiation days 2 and 4 for 48 and 24 hours, respectively. On differentiation day 16, iPS-CMs were purified with

4mM lactate for 48 hours. iPS-CMs were maintained in RPMI-1640 medium supplemented with B27 insulin for 40 days.

v. Immunostaining

To stain spheroids for cardiac markers, cells were first fixed with 4% formaldehyde for 30 minutes, then permeabilized with 0.5% triton-X for 60 minutes. 3%

BSA in D-PBS was used to block cells for 60 minutes. Primary antibodies anti-TNNI3

(Invitrogen, PA1-86820) 1:750, IRX4 (Invitrogen, PA5-40481) 1:750, and anti-α-actinin

(Invitrogen, MA1-22863) 1:500 were diluted in 0.5% TritonX, 3% bovine serum albumin in 1x PBS. Cells were incubated with primary antibodies at 4°C overnight, and then were washed 3 times with 1x D-PBS. Cells were incubated with secondary antibodies

AlexFluor® 555 (Invitrogen, A27017) 1:200, or AlexaFluor® 488 (Invitrogen, A11059 and A11055) 1:200 for one hour at room temperature, followed by three washes with 1x

PBS for five minutes. NucBlue™ Fixed Cell ReadyProbes™ (Thermo, R37606) was added into the last wash. Cells were visualized using an Olympus CKX53 Inverted

Microscope with a PhotoFluor LM-75 light source (89 North Inc.) vi. Treatment with MitoQ and ATP Assay

MitoQ is an antioxidant compound which targets oxidative stress in the mitochondria (Figure 6-28). Due to its lipophilic nature, MitoQ is able to cross the phospholipid bilayer of the cell and enter the mitochondrial matrix, which is a major source of reactive oxidative species [15]. Using MitoQ as a model compound for drug screening, we measured ATP levels in three cell lines.

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Figure 6-28: Structure of MitoQ

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After 40 days post-differentiation, iPS-CM spheroids were treated with 5μM

mitoQ (Figure 6-28) or 5 days. After 5 days of the treatment, the iPS-CM 384-sidewall

pillar plate was sandwiched with a 384-well plate containing 50μL of CellTiter-Glo

Reagent (kit, Promega, G7570) to measure ATP levels. The sandwiched plates were

placed on a shaker for two minutes to induce cell lysis, then incubated at room

temperature for ten minutes to stabilize luminescent signals. The luminescence was

measured with an automated fluorescence reader (S+ Scanner) at an emission wavelength

of 560nm.

vii. Harvesting iPS-CMs from the 384-Sidewall Plate

iPS-CMs were manually detached from individual pillars and placed into organic

solvents for metabolite extraction. To increase sensitivity of mass spectrometric assays

iPS-CMs from 6 pillars were combined into a mixture of (1:0.75:2mL) acetonitrile,

water, and chloroform. After centrifugation at 5000rpm, three phases were formed. Polar

and nonpolar phases were combined and dried down under N2 for subsequent LC-MS/MS

or GC-MS analysis, while the intermediate protein phase was retained for protein

quantification. viii. GC-MS Profiling of Metabolites

Dried cell extracts were derivatized with 60μL of 20mg/mL methoxyamine

(Sigma, 226904) in pyridine for 60 minutes at 80°C, followed by N,O-

Bis(trimethylsilyl)trifluoroacetamide (BSTFA), 40 μL (Regis Technologies, cat.no. 1-

270121-200) for 30 minutes at 70°C. Samples were analyzed via EI-GC-MS in a non-

targeted fashion using a scan method for m/z 50-650 Da (See “Untargeted Metabolite

Analysis” in Appendix B for full method parameters). AMDIS software with the Fiehn

142 library (Agilent) was used for metabolite identification. Compounds with a net match score of 80% or greater compared to the reference library [16] were considered as positive hits. ix. LC-MS/MS Profiling of Acyl Carnitines

Dried cell extracts were derivatized using 3M HCl-n-butanol for 30 minutes at

65°C. After derivatization, the samples were cooled to room temperature and dried down again. The samples were reconstituted in mobile phase composed of 80% acetonitrile,

20% water, and 0.1% formic acid. 7μL of derivatized sample was introduced into the mass spectrometer through the inline filter by direct flow infusion. The flow rate was held at 50μL/min. for a total run time of 1.8 minutes. Tandem mass spectrometry was set up to scan in high throughput mode for the acyl carnitine precursor ion m/z 85. Data analysis was performed using Chemoview 2.2 software.

x. Statistical Analysis.

The data are presented as the mean ± the standard deviation or the standard error of the mean from multiple samples. Significance was tested with the paired two tailed t- test using GraphPad calculator. A p value < 0.05 was considered as significant.

6.4 Results

Comparison of 384-Well Seeding Methods. To determine an effective protocol for miniaturization of iPSC into 3D and subsequent differentiation to iPS-CMs in 3D, several hydrogel compositions were first tested. iPSCs from monolayer cultures were suspended and printed into alginate, 1:1 alginate/Matrigel, or Matrigel hydrogels. 24

143 hours after seeding, spots from alginate and alginate/Matrigel pillars detached from the

384-pillar plate (data not shown), thus hydrogels containing alginate were not used for subsequent studies. Next, two methods were tested for development of 3D iPSC spheroids and efficiency of iPS-CM differentiation. Cells were either directly encapsulated into Matrigel hydrogels using a microarray printer (the “Direct Method”) or were aggregated in ultra-low attachment plates for one week prior to transfer to 384- micropillar plates (the “ULA Method”). iPSCs directly bioprinted onto micropillars exhibited variable densities, distribution of cells, and 3D morphology. Cells which were allowed to aggregate at the bottom of ULA plates displayed uniform spheroid size and shape. Both the Direct and ULA plating methods produced beating cells, which continued to beat for up to 40 days post-differentiation. The overall beating ratio of pillars using the

Direct Method was less than 10%. The differentiation efficiency improved to 20-25% of pillars across iPS-CMs from spheroids from all three cell lines which were first formed on ULA plates by the ULA Method.

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Figure 6-29: iPSC Spheroid Formation on 384-Pillar Plates

iPSCs were seeded onto 384-pillar plates directly from 2D culture (Direct Printing

Method) or transferred onto 384-pillar plates after aggregation on ultra-low attachment plates (ULA method).

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Differentiation and Immunostaining. To obtain functional, beating cardiomyocytes from spheroids, the small molecule protocol which was optimized for differentiation of 2D cultures was applied to our 3D model. As described previously, temporal modulation of the small molecules CHIR 99021 and IWR1 in RPMI-1640 medium supplemented with B27 minus insulin efficiently induced cardiac differentiation.

Cells began to beat spontaneously and maintaining beating for up to 40 days in culture.

To evaluate expression of key cardiac markers, control and TAZ1 iPS-CMs were immunostained for TNNI3 and α-actinin. Cells were positive for these markers, although microstructure was not able to be visualized (Figure 6-30).

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Figure 6-30 Immunostaining of 3D iPS-CMs.

Representative images of iPS-CMs stained positive for α-actinin (green), TNNI3

(red), and DAPI (blue). 20X magnification.

147

ATP Assay with drug treatment. To test the potential of for performing high content imaging analysis using this 3D iPS-CM platform, iPS-CMs from the two control lines and one disease line were treated with 5μM MitoQ in DMSO or DMSO alone for five days starting 40 days post-differentiation. After 5 days of treatment, CellTiter-Glo

Cell Viability Assay (Promega, 7570) was used in order to measure the luminescent signal with a plate reader. Luminescent signal is proportional to the amount of ATP present. Basal luminescence in the untreated Control and WT2 were higher than in TAZ- iPS-CMs (Figure 6-32). Treatment with MitoQ did not have a noticeable effect on the control line WT2 iPS-CMs, with the luminescence increasingly slightly from 459.9 +

33.5 relative light units (RLU) untreated to 471.5 + 41.9 RLU with treatment. Cell viability of iPS-CMs from the Control line increased by 34% from 363.5 + 19.3 RLU to

550.3 + 55.5 RLU (p < 0.005**). While luminescence of TAZ1 iPS-CMs without treatment was 250.4 + 19.0 RLU, treatment with MitoQ restored ATP levels to 356.9 +

23.5 RLU, near the levels of the untreated Control, which is isogenic to the TAZ1 line (p

= 0.001***).

148

ATP Levels 700 ** 600 500 400 *** 300 200

Luminescence (RLU) Luminescence 100 0 WT2 Control TAZ1 Untreated Mito-Q

Figure 6-31: ATP Levels with MitoQ Treatment

ATP levels in WT2, Control, and TAZ1 iPS-CMs with or without MitoQ treatment. n = 24. (**p < 0.005; ***p < 0.0005)

149

Untargeted Metabolic Profiling Using GC-MS. To test the sensitivity of the GC-

MS platform to the lower concentration of metabolites extracted from iPS-CMs cultured on 384-micropillars, six pillars were combined into one sample, then metabolites were extracted, derivatized, and analyzed via EI-GC-MS. Qualitative analysis identified only

22 compounds with a net score of 80% or greater compared to the reference library, including lactate, pyruvate, nine amino acids, and five fatty acids (Figure 6-32). Of the citric acid cycle intermediates, only fumarate was identified.

150

Figure 6-32: Untargeted Analysis of Metabolites from 3D iPS-CMs

(1) Pyruvate (2) Lactate (3) Glycolic Acid (4) Urea (5) Fumarate (6) L-leucine (7) DL-isoleucine (8) L-mimosine (9) Valine (10) Glycine (11) Serine (12) Threonine (13) Hydroquinone (14) Aspartic Acid (15) Capric Acid (16) Glutamic Acid (17) Lauric Acid (18) D-glucose (19) Palmitic Acid (20) Allo-inositol (21) Oleic Acid (22) Stearic Acid

151

LC-MS/MS Profiling of Acyl Carnitines and CAC Intermediates. Because of its greater sensitivity to measure low concentrations of compounds than GC-MS, we next tested the ability of LC-MS/MS to measure metabolites extracted from iPS-CMs on 384- micropillars. About 60 species of acyl carnitines were detected from iPS-CMs via LC-

MS/MS, including short-, medium-, long-, and very long-chain acyl carnitines and many unsaturated, hydroxylated, and dicarboxylic derivatives of various acyl carnitine species.

Concentrations of the detected acyl carnitines from iPS-CMs ranged from 0.04nM up to

90nM (. A complementary LC-MS/MS method was used to analyze citric acid cycle

(CAC) intermediates. Citrate, aconitate, isocitrate, α-ketoglutarate, succinate, fumarate, and malate were detected (Figure 6-34) (oxaloacetate was not included in the method).

Fumarate was measured in the lowest abundance at 0.00055 ng/μL, with citrate being the highest at 7.16 ng/μL.

152

2 WT2 1.5 Control TAZ1 1

0.5

Concetration (nM) Concetration 0 C2 C3-DC C4 C4-OH C5 C5:1 C6 C8 C10:3 -0.5

10 8 6 4 2 0 Concentration (nM) Concentration -2

8 7 6 5 4 3 2 1 0 Concentration (nM) Concentration -1

Figure 6-33: Measurement of Acyl Carnitines by LC-MS/MS

(A) Short and medium chain, (B) long chain, and (C) very long chain acyl carnitines detected in iPS-CMs

153

Figure 6-34: Detection of CAC Intermediates by LC-MS/MS

154

6.5 Discussion

Currently, the major challenge in iPS-CM technology is the production of mature cardiomyocytes which possess structural and functional characteristics resembling human adult cardiomyocytes. A simple method for increasing iPS-CM maturity is long-term culturing of cells >45 days [17]. More elaborate attempts for inducing cardiomyocyte maturation have centered on physical, biochemical, or tissue-engineering techniques [18].

Physical methods attempt to reproduce physical conditions of in vivo cardiomyocytes by modulating electrophysiological stimuli [19], substrate stiffness [20], magnetic field [21], or mechanical stretching of cells [22]. Biochemical stimulation to improve Ca2+ handling, cell morphology, and contractile function have been described. The thyroid hormone, triiodothyronine promotes maturity of morphological, molecular, and functional characteristics of iPS-CMs [23]. β-adrenergic receptor agonists improve Ca2+ fluxes and sarcomeric organization [8]. Finally, culturing cells in galactose and fatty acid medium free of glucose was shown to successfully induce the transition from fetal-like glycolytic metabolism to adult-like oxidative metabolism [24]. Engineering techniques include culturing cells on patterned substrates, hydrogels, cell sheets, or microfluidic devices in order to promote cardiomyocyte maturation [11][25]. Despite much progress, as of yet, no single method exists which can produce iPS-CMs which achieve the structural and functional maturity of human adult cardiomyocytes [26].

Besides immaturity, 2D cell cultures often fail to reliably mimic in vivo responses after treatment with drug candidates due to lack of 3D organization and microenvironment [13]. In order to develop a platform which would produce mature iPS-

CMs which could ultimately be used in HTP drug screenings or cardiotoxicity assays, we

155 sought to incorporate iPS-CMs into a miniaturized, 3D format. Previous attempts to establish 3D iPS-CM cultures include the use of iPSC embryoid body formation or various bioengineering strategies [27]. Bioengineered platforms for 3D cell cultures lack microscalability, precluding their usefulness for high throughput screening and are associated with high costs. Moreover, it is difficult to control the 3D microenvironment using these platforms. To address these limitations, we first tested three biomimetic hydrogel compositions to use as scaffolds for iPSC cultures, including alginate alone, alginate/Matrigel, or Matrigel alone. Hydrogels are polymers with high water content which possess structural similarity to the extracellular matrices of various body tissues

[28], and can be modified to create the desired microenvironment by controlling shape, size, stiffness, and porosity of the scaffold [29]. Alginate has been shown to support cell growth and proliferation [30], while Matrigel is a mixture of basement membrane proteins and growth factors, reproducing a degree of the complexity of cellular microenvironments [31]. Cell spots which were printed in alginate and alginate/Matrigel hydrogels detached from the micropillar plate 1-2 days after printing, and thus any hydrogel compositions containing alginate were not considered for further studies. Cell spot attachment with Matrigel remained at >95% in long-term cell culture, thus Matrigel was used to further optimize our 3D micropillar system.

Next, we sought to form spheroids of homogenous size and shape, as uniformity of culture conditions is necessary to reproducibly differentiate iPSCs to iPS-CMs. iPSCs from 2D monolayer cultures were applied onto 384-micropillar plates in two ways: either by directly bioprinting cells (Direct Method) or by first aggregating cells in an ultra-low attachment (ULA) plate for one week before transfer to micropillars (ULA Method). In

156 the Direct Method, a microarray cell printer was used to print cells mixed with Matrigel onto pillars with sidewalls. In the ULA method, cells from 2D culture were first allowed to aggregate in a ULA plate before being transferred to micropillar plates. Despite the automation of the microarray cell printer, cell spots which were directly printed onto the pillar displayed heterogenous cell densities, spheroid numbers, and spheroid shapes from pillar to pillar. In comparison, spheroids formed by the ULA method exhibited high uniformity. Each well of the ULA plate contained one distinct spheroid of uniform size and shape, which was preserved after transfer to the micropillar. Moreover, this homogeneity was maintained after iPSC spheroids on the micropillar plate were allowed to proliferate for two weeks, demonstrating further ability to reproducibly create uniform spheroid size and shape.

Spheroids from the direct printing method and from the ULA method were next subjected to cardiac differentiation. Beating cells were obtained from both methods for all control and disease lines, however, the efficiency of differentiation varied. Only 10% of pillars in the direct method exhibited spontaneous beating. This improved to 24% of pillars using the ULA method, demonstrating the advantage of working with a homogenous population of spheroids. However, a differentiation efficiency of 24% is relatively low compared to current protocols which report >80% efficiencies. Therefore, the differentiation protocol which was optimized for our 2D monolayer cultures was unable to achieve the same effect in 3D. The concentration of small molecules CHIR and

IWR1 may need to be increased to properly penetrate the 3D structure of spheroids.

Additionally, iPS-CMs were immunostained for α-actinin and cardiac troponin (TNNI3).

Although cells were positive for these markers, microstructure it was not possible to

157 resolve the microstructure of sarcomeric filaments or bands. Thus, to visualize structural maturation, iPS-CMs from pillars would need to be singularized and reseeded onto coverslips prior to immunostaining. This is a laborious process which would not be easily amenable to high-throughput. In light of this, alternate approaches such as PCR or flow cytometry should be considered, as these would allow for quantification of gene or protein expression levels, thus providing more robust, quantitative analysis of iPS-CM phenotype and maturity [32][33].

To understand the efficacy and toxicity of drug candidates, the ability to assess multiple phenotypic parameters is highly valuable. High content imaging (HCI) assays can provide such information, allowing for the visualization of characteristics such as cell viability, mitochondrial integrity, membrane integrity, ATP levels, DNA damage, and oxidative stress in a high-throughput format using a number of fluorescent dyes or fluorescent probes [14]. In this study, we chose to measure ATP levels using the

CellTiter-Glo Cell Viability Assay after treatment with an antioxidant drug candidate

MitoQ. MitoQ is an antioxidant which is used to improve mitochondrial oxidative capacity by mimicking the endogenous mitochondrial compound, coenzyme Q [15]. The

CellTiter-Glo assay revealed that ATP levels in both untreated control lines were greater than TAZ1 iPS-CMs by >1.5-fold, consistent with the increased energy demand in cardiomyopathy. MitoQ treatment rescued ATP levels of TAZ1 to the levels seen in the untreated, isogenic Control line. Luminescence from the CellTiter-Glo assay was measured from the 384-pillar plate in under 20 minutes. However, one drawback to this method is high background fluorescence, thus the additional step of image processing to eliminate background fluorescence needs to be considered. Additionally, this pilot assay

158 was tested using iPS-CMs from the direct printing method, which was shown to produce colonies and spheroids of wildly different sizes, thus introducing a large degree of biological variation. To obtain robust data, future HCI assays will be performed using spheroids generated via the ULA method. Nevertheless, these results demonstrate the feasibility of performing HCI assays on a 3D iPS-CM platform. Ultimately, this system has the potential to be used to test many drug concentrations, thus determining dose- response curves and IC50 of various drug candidates.

Besides high content imaging, measuring cardio-metabolic endpoints in response to potential cardiac therapeutics in a high-throughput manner could provide important insight for cardiac drug evaluation. This is especially true given the renewed interest in metabolic therapies as treatments for heart disease [34][35]. We applied an untargeted metabolic GC-MS method to search for metabolites against a compound library containing thousands of compounds. After combining six pillars into one sample, only 22 compounds were able to be identified through this method. Qualitative analysis identified only 22 compounds with a net score of 80% or greater compared to the reference library, including lactate, pyruvate, nine amino acids, and five fatty acids (Figure 6-32). Of the citric acid cycle (CAC) intermediates, only fumarate was identified. Lactate and pyruvate could serve as useful metabolic endpoints. However, GC-MS was not sensitive enough to identify a meaningful number of compounds from individual 384-pillars. Furthermore, the workflow for metabolite extraction and derivatization is labor-intensive and low- throughput.

Next, we turned to LC-MS/MS methods which were previously developed in our laboratory to measure CAC and acyl carnitines from iPS-CM micropillar samples.

159

Because of the increased sensitivity of LC-MS/MS, we were able to detect very low concentrations of analytes down to 0.4nM, including 60 species of acyl carnitines and all

CAC intermediates (except oxaloacetate, which is not included in the method). Sample preparation for LC-MS/MS by these methods requires extraction and derivatization techniques. However, various research groups have demonstrated methods by which 384- well plates can be paired to LC-MS/MS for enzymatic assays via matrix-assisted laser desorption ionization or direct ionization of samples directly from the plate, without complicated extraction and derivatization steps [36][37][38]. Future studies using such ionization methods combined with our 384-pillar 3D iPS-CM platform could enable high-throughput metabolic screening of metabolic endpoints to assess drug toxicity.

Reliable toxicity and efficacy studies of drug candidates are needed to identify novel lead compounds, however, iPS-CM maturation remains a major challenge in the application of iPS-CMs in drug and toxicity screens [3]. Compared to 3D cell cultures,

2D cell cultures are unable to accurately reproduce the complexity of in vivo characteristics. Moreover, several research groups have demonstrated that 3D iPS-CMs can contribute to metabolic, structural, and functional maturity of iPS-CMs [9][10]. The primary objective of the present work was to establish a miniaturized, 3D iPS-CM platform to generate mature iPS-CMs which more closely mimic the complex microenvironment and molecular responses of cardiomyocytes in vivo. Here, we successfully miniaturized iPSC culture and differentiation in a 384-micropillar format using hydrogels to encapsulate cells. Using ULA plates, we were able to generate spheroids of uniform size and shape, thus improving homogeneity across all 384 wells in a relatively-cost effective manner. Differentiation of iPSCs to iPS-CMs in 3D produced

160 spontaneously beating cells, although future work is needed to optimize the efficiency of differentiation. Notably, unlike other 3D systems, the ability to generate homogenous spheroids in a 384-pillar format makes our platform amenable to high throughput workflows, thus carrying the potential for drug candidate screening and phenotypic endpoints as indicators of toxicity.

6.6 References

[1] D. Mozaffarian et al., “Heart Disease and Stroke Statistics—2016 Update,” Circulation, vol. 133, no. 4, Jan. 2016. [2] J. K. Gwathmey, K. Tsaioun, and R. J. Hajjar, “Cardionomics: a new integrative approach for screening cardiotoxicity of drug candidates,” Expert Opin. Drug Metab. Toxicol., vol. 5, no. 6, pp. 647–660, Jun. 2009. [3] C. Denning et al., “Cardiomyocytes from human pluripotent stem cells: From laboratory curiosity to industrial biomedical platform,” Biochim. Biophys. Acta - Mol. Cell Res., vol. 1863, no. 7, pp. 1728–1748, 2016. [4] M. F. Hoes, N. Bomer, and P. van der Meer, “Concise Review: The Current State of Human In Vitro Cardiac Disease Modeling: A Focus on Gene Editing and Tissue Engineering,” Stem Cells Transl. Med., vol. 8, no. 1, pp. 66–74, Jan. 2019. [5] L. Guo et al., “Estimating the risk of drug-induced proarrhythmia using human induced pluripotent stem cell-derived cardiomyocytes.,” Toxicol. Sci., vol. 123, no. 1, pp. 281–9, Sep. 2011. [6] P. Joshi, S. Y. Kang, A. Datar, and M. Y. Lee, “High-Throughput Assessment of Mechanistic Toxicity of Chemicals in Miniaturized 3D Cell Culture,” Curr. Protoc. Toxicol., vol. 79, no. 1, pp. 1–22, 2019. [7] F. B. Bedada, M. Wheelwright, and J. M. Metzger, “Maturation status of sarcomere structure and function in human iPSC-derived cardiac myocytes.,” Biochim. Biophys. Acta, vol. 1863, no. 7 Pt B, pp. 1829–38, Jul. 2016. [8] T. J. Kolanowski, C. L. Antos, and K. Guan, “Making human cardiomyocytes up to date: Derivation, maturation state and perspectives,” Int. J. Cardiol., vol. 241, pp. 379–386, Aug. 2017. [9] C. Correia et al., “3D aggregate culture improves metabolic maturation of human pluripotent stem cell derived cardiomyocytes,” Biotechnol. Bioeng., vol. 115, no. 3, pp. 630–644, 2018. [10] A. H. Fong et al., “Three-Dimensional Adult Cardiac Extracellular Matrix Promotes Maturation of Human Induced Pluripotent Stem Cell-Derived

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Cardiomyocytes.,” Tissue Eng. Part A, vol. 22, no. 15–16, pp. 1016–25, 2016. [11] G. Pettinato, X. Wen, and N. Zhang, “Formation of well-defined embryoid bodies from dissociated human induced pluripotent stem cells using microfabricated cell- repellent microwell arrays,” Sci. Rep., vol. 4, 2014. [12] R. L. Carpenedo, C. Y. Sargent, and T. C. Mcdevitt, “Rotary Suspension Culture Enhances the Efficiency, Yield, and Homogeneity of Embryoid Body Differentiation,” Stem Cells, vol. 25, pp. 2224–2234, 2007. [13] A. Datar, P. Joshi, and M. Y. Lee, “Biocompatible hydrogels for microarray cell printing and encapsulation,” Biosensors, vol. 5, no. 4, pp. 647–663, 2015. [14] P. Joshi, A. Datar, K. N. Yu, S. Y. Kang, and M. Y. Lee, “High-content imaging assays on a miniaturized 3D cell culture platform,” Toxicol. Vitr., vol. 50, no. March, pp. 147–159, 2018. [15] J. S. Tauskela, “MitoQ--a mitochondria-targeted antioxidant.,” IDrugs, vol. 10, no. 6, pp. 399–412, Jun. 2007. [16] T. Kind et al., “FiehnLib: Mass Spectral and Retention Index Libraries for Metabolomics Based on Quadrupole and Time-of-Flight Gas Chromatography/Mass Spectrometry,” Anal. Chem., vol. 81, no. 24, pp. 10038– 10048, Dec. 2009. [17] K. Ronaldson-Bouchard et al., “Advanced maturation of human cardiac tissue grown from pluripotent stem cells.,” Nature, vol. 556, no. 7700, pp. 239–243, 2018. [18] X. Yang, L. Pabon, and C. E. Murry, “Engineering Adolescence,” Circ. Res., vol. 114, no. 3, pp. 511–523, Jan. 2014. [19] Y.-C. Chan et al., “Electrical Stimulation Promotes Maturation of Cardiomyocytes Derived from Human Embryonic Stem Cells,” J. Cardiovasc. Transl. Res., vol. 6, no. 6, pp. 989–999, Dec. 2013. [20] J. G. Jacot, A. D. McCulloch, and J. H. Omens, “Substrate Stiffness Affects the Functional Maturation of Neonatal Rat Ventricular Myocytes,” Biophys. J., vol. 95, no. 7, pp. 3479–3487, Oct. 2008. [21] M. Cornacchione et al., “β-Adrenergic response is counteracted by extremely-low- frequency pulsed electromagnetic fields in beating cardiomyocytes,” J. Mol. Cell. Cardiol., vol. 98, pp. 146–158, Sep. 2016. [22] O. J. Abilez et al., “Passive Stretch Induces Structural and Functional Maturation of Engineered Heart Muscle as Predicted by Computational Modeling,” Stem Cells, vol. 36, no. 2, pp. 265–277, Feb. 2018. [23] F. B. Bedada et al., “Tri-iodo-l-thyronine promotes the maturation of human cardiomyocytes-derived from induced pluripotent stem cells.,” J. Mol. Cell. Cardiol., vol. 12, no. 1, pp. 296–304, 2017. [24] A. Heinick et al., “Distinct carbon sources affect structural and functional maturation of cardiomyocytes derived from human pluripotent stem cells,”

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Biotechnol. Bioeng., vol. 12, no. 1, pp. 1–17, 2017. [25] C. Zuppinger, “3D culture for cardiac cells,” Biochim. Biophys. Acta - Mol. Cell Res., vol. 1863, no. 7, pp. 1873–1881, Jul. 2016. [26] G. J. Scuderi and J. Butcher, “Naturally Engineered Maturation of Cardiomyocytes.,” Front. cell Dev. Biol., vol. 5, p. 50, 2017. [27] Q. Gu, E. Tomaskovic-Crook, G. G. Wallace, and J. M. Crook, “3D Bioprinting Human Induced Pluripotent Stem Cell Constructs for In Situ Cell Proliferation and Successive Multilineage Differentiation,” Adv. Healthc. Mater., vol. 6, no. 17, pp. 1–11, 2017. [28] J. Zhu and R. E. Marchant, “Design properties of hydrogel tissue-engineering scaffolds.,” Expert Rev. Med. Devices, vol. 8, no. 5, pp. 607–26, Sep. 2011. [29] Q. L. Loh and C. Choong, “Three-dimensional scaffolds for tissue engineering applications: role of porosity and pore size.,” Tissue Eng. Part B. Rev., vol. 19, no. 6, pp. 485–502, Dec. 2013. [30] M. M. Capeling et al., “Nonadhesive Alginate Hydrogels Support Growth of Pluripotent Stem Cell-Derived Intestinal Organoids,” Stem Cell Reports, vol. 12, no. 2, pp. 381–394, Feb. 2019. [31] S. R. Caliari and J. A. Burdick, “A practical guide to hydrogels for cell culture.,” Nat. Methods, vol. 13, no. 5, pp. 405–14, 2016. [32] S. Bhattacharya et al., “High Efficiency Differentiation of Human Pluripotent Stem Cells to Cardiomyocytes and Characterization by Flow Cytometry,” J. Vis. Exp., no. 91, p. 52010, Sep. 2014. [33] A. Heinick et al., “Universal Cardiac Induction of Human Pluripotent Stem Cells in Two and Three-Dimensional Formats: Implications for In Vitro Maturation,” Stem Cells, vol. 33, no. 5, pp. 1456–1469, 2015. [34] C. Des Rosiers, F. Labarthe, S. G. Lloyd, and J. C. Chatham, “Cardiac anaplerosis in health and disease: food for thought,” Cardiovasc. Res., vol. 90, no. 2, pp. 210– 219, May 2011. [35] H. Taegtmeyer et al., “Assessing Cardiac Metabolism,” Circ. Res., vol. 118, no. 10, pp. 1659–1701, May 2016. [36] L. Qiao et al., “Electrostatic Spray Ionization from 384-Well Microtiter Plates for Mass Spectrometry Analysis-Based Enzyme Assay and Drug Metabolism Screening,” Anal. Chem., vol. 89, no. 11, pp. 5983–5990, 2017. [37] K. L. Kurita, E. Glassey, and R. G. Linington, “Integration of high-content screening and untargeted metabolomics for comprehensive functional annotation of natural product libraries,” Proc. Natl. Acad. Sci., vol. 112, no. 39, pp. 11999– 12004, 2015. [38] S. J. Stout et al., “Mass Spectrometric Techniques for Label-free High-Throughput Screening in Drug Discovery,” Anal. Chem., vol. 79, no. 21, pp. 8207–8213, 2007.

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CHAPTER VII

CONCLUSION

The manifestation of cardiac disease is the major clinical feature of Barth

Syndrome (BTHS), with a high incidence in early life and, subsequently is a leading cause of death in infants and young children affected by this disease [1]. Despite the severity of this BTHS, there is a lack of targeted therapeutics which can be used to ameliorate symptoms and prolong the lives of these patients.

To better characterize metabolic and molecular alterations that occur in the presence of BTHS-causative TAZ mutations, we first sought to develop a model of

BTHS using induced pluripotent stem cell-derived cardiomyocytes (iPS-CMs), which have shown great promise in modeling inherited cardiomyopathies [2][3][4]. Our work resulted in successful production of isogenic control and TAZ-mutant iPS-CMs for further studies of the cardiac features of BTHS, as well as a second line carrying a second

TAZ mutation. These three lines were used in stable isotope tracer studies, which demonstrated that TAZ deficiency induces metabolic alterations in energy production related pathways and shifts a preferable carbon substrate from fatty acids to glucose. We also identified significant alterations in pyruvate carboxylation, an anaplerotic pathway

164 which is partly responsible for preserving the balance of CAC intermediates, and subsequently, maintaining energy production. Labeling patterns of succinate and decreases in amino acid levels further implicate aberrations of anaplerosis at the succinyl-

CoA level. The exact functional significance of the metabolic remodeling in Barth syndrome still remains to be elucidated. Future stable isotope tracer studies should be performed to probe pyruvate carboxylation and amino acid anaplerosis at the level of succinyl-CoA.

We also presented evidence that key calcium homeostasis genes are involved in the TAZ-iPS-CM phenotype. These data support our hypothesis that disrupted calcium homeostasis contributes to the cardiac phenotype of BTHS. In addition to its role in excitation-contraction cycle, calcium also plays a role in mediating energy production and antioxidant capacity in mitochondria [5]. Those processes are highly relevant in phenotypic descriptions of BTHS patients, thus calcium-mediated cellular processes need to be further explored for their role in the pathogenesis of BTHS.

Maturation of iPS-CMs to a mature phenotype more closely resembling human adult cardiomyocytes continues to pose a major challenge to the field of disease modeling

[6]. 3D iPS-CM cultures have the potential for increased structural and function maturity, and the ability to more accurately mimic in vivo conditions compared to their 2D counterparts [7]. However, reproducible generation of 3D cultures remains difficult [8].

In preliminary work, we have established a miniaturized, 3D model of iPS-CMs using hydrogels and cellular aggregation to generate iPS-CM spheroids of uniform shape and size, representing an important step toward differentiation reproducibility. The use of a

384-micropillar plate with sidewalls is further advantageous over existing 3D systems, as

165 it is more cost-effective and is amenable to high throughput and high content imaging assays. Future work will center around improving efficiency of differentiation on this novel platform.

Overall, our findings reveal that the TAZ mutation induces alterations in molecular and metabolic pathways linked to ATP production. As an inadequate supply of

ATP for contractile function results in decreased cardiac efficiency, these sites of alterations represent important potential targets for the restoration of cardiac function in

BTHS. Further studies are needed to assess metabolic and molecular changes in cells carrying additional mutations in the TAZ gene.

7.1 References

[1] S. L. N. Clarke et al., “Barth syndrome,” Orphanet J. Rare Dis., vol. 8, no. 1, pp. 1–17, 2013. [2] G. Wang et al., “Modeling the mitochondrial cardiomyopathy of Barth syndrome with induced pluripotent stem cell and heart-on-chip technologies,” Nat. Med., vol. 20, no. 6, pp. 616–623, 2014. [3] N. Sun et al., “Patient-specific induced pluripotent stem cells as a model for familial dilated cardiomyopathy.,” Sci. Transl. Med., vol. 4, no. 130, p. 130ra47, Apr. 2012. [4] C. Kim et al., “Studying arrhythmogenic right ventricular dysplasia with patient- specific iPSCs,” Nature, vol. 494, no. 7435, pp. 105–110, Feb. 2013. [5] D. A. Eisner, J. L. Caldwell, K. Kistamás, and A. W. Trafford, “Calcium and Excitation-Contraction Coupling in the Heart,” Circ. Res., vol. 121, no. 2, pp. 181– 195, Jul. 2017. [6] C. Denning et al., “Cardiomyocytes from human pluripotent stem cells: From laboratory curiosity to industrial biomedical platform,” Biochim. Biophys. Acta - Mol. Cell Res., vol. 1863, no. 7, pp. 1728–1748, 2016. [7] C. Correia et al., “3D aggregate culture improves metabolic maturation of human pluripotent stem cell derived cardiomyocytes,” Biotechnol. Bioeng., vol. 115, no. 3, pp. 630–644, 2018. [8] M. Vinci et al., “Advances in establishment and analysis of three-dimensional tumor spheroid-based functional assays for target validation and drug evaluation.,”

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BMC Biol., vol. 10, p. 29, Mar. 2012.

167

APPENDIX

168

APPENDIX A

CORRECTION OF MASS ISOTOPOMER DISTRIBUTIONS FOR NATURAL ISOTOPIC

ABUNDANCE

A.1 Introduction

13C stable isotope tracer studies are becoming increasingly utilized in the field of metabolomics for delineating substrate production and consumption through metabolic pathways. Due to its ability to measure mass isotopomers, mass spectrometry (MS) is one of the primary analytical techniques used in stable isotope tracer studies [1]. The mass isotopomer information collected by MS enables 13C mass isotopomer distribution (MID) analysis, which takes into account the relative percent of each mass isotopomer in a given compound.

In order to accurately assess the extent of 13C contribution from the selected stable isotope tracer, it is necessary to correct MS data for the presence of naturally-occurring isotopes [2]. As 13C has a natural abundance of 1.1%, uncorrected data would overestimate the fraction of the M1 isotopomer in any given compound [3]. This error is further compounded when the number of carbons in the molecule’s formula increases.

Moreover, metabolite derivatization required for GC-MS analysis introduces additional atoms with naturally-occurring heavy isotopes, such as the case with trimethylsilyl (TMS;

C3H9Si) and tert-butyldimethylsilyl (TBDMS; C5H15Si) derivatives. To correct MS data, a matrix-based correction model has been described in the literature [1]. In this work, we have applied the described mathematical formulas to construct a correction matrix for

169 iPS-CM, using unlabeled iPS-CM to determine the naturally-occurring MID for analytes of interest.

A.2 Methods and Results

i. Cell Harvesting and Metabolite Extraction

Cells were harvested as previously described (Chapter$). Extracts were dried under a nitrogen stream at room temperature, then derivatized using methoxyamine hydrochloride in pyridine followed by derivatization with N-methyl-N-tert- butyldimethylsilyltrifluoroacetamide to form TBDMS derivatives. GC-MS analysis was carried out with a 5977 GC-MS (Agilent) operated in EI mode and equipped with an HP-

5MS column (Agilent). Peak areas for each M0-Mn ion of a given analyte were integrated using Agilent 5977 Data Analysis software with RTE Integrator.

ii. Construction of Correction Matrices

The following steps describe the steps followed to construct correction matrices for analytes from harvested iPS-CMs, including lactate, pyruvate, succinate, fumarate, malate, α-ketoglutarate, citrate, isocitrate, and glutamate. The steps as shown here use malate as an example.

Step 1: Integrate the peak areas for all ions M0-Mn of the compound (Figure

A-1). Malate has 4 carbons, and therefore, has 5 possible mass isotopomers; 419 (M0),

420 (M1), 421 (M2), 422 (M3), and 423 (M4).

170

Mass Isotopomer Peak Area (Uncorrected)

M0 353179 M1 128669 M2 62568 M3 15159 M4 3946

Figure A-1 Determining Natural Abundance.

Step 1: Integration of mass isotopomers of malate. (A) Representative chromatogram of unlabeled malate. (B) Peak areas of each mass isotopomer of malate,

M0-M4.

171

Step 2: Determine the relative mass isotopomer abundance of each mass isotopomer to M0 by dividing the peak area of a given isotopomer to the sum of all M0-

Mn ions, according to the following formula:

푃푒푎푘 퐴푟푒푎 표푓 푀x 푅푒푙푎푡푖푣푒 퐴푏푢푛푑푎푛푐푒 = 푃푒푎푘 푎푟푒푎 표푓 푀0

Relative Abundance to Mass Isotopomer Peak Area (Uncorrected) M0 M0 353179 1 M1 128669 0.362 M2 62568 0.178 M3 15159 0.062 M4 3946 0.0515

Step 3: Construct the correction matrix by filling in the rows of the matrix for incrementally-labeled 13-C malate, based on the mass isotopomer abundances calculated in Step 2.

Malate Correction Matrix 419 420 421 422 423 Unlabeled 1 0.362 0.178 0.043 0.011 13 C1 0 1 0.362 0.178 0.043 13 C2 0 0 1 0.362 0.178 13 C3 0 0 0 1 0.362 13 C4 0 0 0 0 1

172

Step 4: Construct the inverse correction matrix (CM-1). To do this in Excel, use the MINVERSE function on the CM to calculate the CM-1:

Malate Inverse Correction Matrix 1 -0.362 -0.047 0.038 -0.002 0 1 -0.362 -0.047 0.038 0 0 1 -0.362 -0.047 0 0 0 1 -0.362 0 0 0 0 1

iii. Using the Correction Matrix to Correct Peak Areas

Step 5: Integrate the peak areas of the analyte of interest which has been enriched by stable isotope tracing. Use the Excel function MMULT to return the matrix product of

-1 the inverse correction matrix (CM ) and the uncorrected peak areas (AU). The subsequent results are the corrected areas (AC), which can be used to calculate mass isotope distribution.

−1 퐴퐶 = 퐴푈 ∙ 퐶푀

M0 M1 M2 M3 M4

Uncorrected 1022763 449250 456778 498405 303122 Corrected 1022763 79202 246487 351388 117161

173 iv. Calculating the Mass Isotopomer Distribution of Labeled Metabolites

Step 6: Calculate the mass isotopomer distribution of a metabolite from a labeling experiment using the corrected peak areas. To calculate MID, divide the peak area of a given isotopologue by the sum of all peak areas (Figure A-2). Any corrected peak values which are negative are not included in this sum.

퐶표푟푟푒푐푡푒푑 퐴푟푒푎 표푓 푀푥 푀퐼퐷 = ∑ 푀0 − 푀푛

A.3 Discussion

Accurate determination of the distribution of 13C stable isotope tracers depends upon the correction of mass spectral peak areas to account for natural isotopic abundance.

As 13C makes up 1.1% of all carbon, uncorrected peak areas lead to an overestimation of the M1 mass isotopomer. As seen in Figure A-2, after 8 hours of tracing with 5mM 13C6- glucose, M1 malate extracted from control iPS-CMs was calculated to be 16% before correction. This value is 4 times higher than the corrected value of 4%. Additionally, the enrichment of M4 malate was calculated as 11% before correction and 6% after correction. Therefore, this illustrates how interpreting uncorrected data could lead to inaccurate biological conclusions.

174

MID of Malate after 13C Tracing 60% 56% 50% 40% 37% 30% 19% 16% 17% 18% 20% 14%

11% % Enrichment % 10% 4% 6% 0% M0 M1 M2 M3 M4 Uncorrected Corrected

Figure A-2: Mass Isotopomers of Malate.

13 MID in control iPS-CMs after labeling with 10mM C6-glucose for 8 hours.

175

A.4 References

[1] C. Des Rosiers, S. Lloyd, B. Comte, and J. C. Chatham, “A critical perspective of the use of13C-isotopomer analysis by GCMS and NMR as applied to cardiac metabolism,” Metab. Eng., vol. 6, no. 1, pp. 44–58, 2004. [2] J. C. Portais, P. Millard, F. Letisse, and S. Sokol, “IsoCor: Correcting MS data in isotope labeling experiments,” Bioinformatics, vol. 28, no. 9, pp. 1294–1296, 2012. [3] H. Brunengraber, C. Des Rosiers, C. A. Fernandez, S. F. Previs, and F. David, “Correction of13C Mass Isotopomer Distributions for Natural Stable Isotope Abundance,” J. Mass Spectrom., vol. 31, no. 3, pp. 255–262, 2002.

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APPENDIX B

GAS CHROMATOGRAPHY-MASS SPECTROMETRY METHODS

i. Instrumentation

GC: Agilent 7890B MS: Agilent 5977A Column: HP-5MS 5% Phenyl Methyl Siloxane

ii. Analytical Conditions

Amino Acid Analysis

177

Glucose Analysis

GC Derivatization: Aldonitrile/Pentaacetylation Injection volume: 1μL Injection mode: Splitless Carrier gas: He Time program: Initial temp. 80°C; hold 3 min. Ramp 15°C/min. up to 180°C Ramp 5°C/min. up to 205°C Ramp 1°C/min. up to 212°C Ramp 15°C/min. up to 310°C Hold 310°C; hold 5 min. MS Ion Source: Electron Ionization Ion Source Temp.: 280°C MS Quad. Temp.: 150°C Solvent Delay: 3.5 min. Acquisition Mode: SIM SIM Ions: m/z 191, 314, 315, 316, 317, 318, 319, 320, 321

178

Palmitic Acid Analysis

GC Derivatization: Methylation Injection volume: 2μL Injection mode: Splitless Carrier gas: He Time program: Initial temp. 100°C; hold 1 min. Ramp 8°C/min. up to 300°C Hold 300°C; hold 2 min. MS Ion Source: Electron Ionization Ion Source Temp.: 280°C MS Quad. Temp.: 150°C Solvent Delay: 4 min. Acquisition Mode: SIM SIM Ions: m/z 239, 241, 242, 243, 253, 255, 270, 274, 284, 286

179

Organic Acids Analysis

GC Derivatization: Methoxymation/tert-Butyldimethylsilylation Injection volume: 1μL Injection mode: Splitless Carrier gas: He Time program: Initial temp. 80°C; hold 3 min. Ramp 15°C/min. up to 205°C Ramp 10°C/min. up to 220°C Ramp 15°C/min. up to 305°C Hold 305°C; hold 3 min. MS Ion Source: Electron Ionization Ion Source Temp.: 280°C MS Quad. Temp.: 150°C Solvent Delay: 5.8 min. Acquisition Mode: SIM Group 1 Time Window: 5.8 min. – 12.5 min. Group 1 SIM Ions: m/z 174, 175, 176, 177, 233, 234, 235, 236, 237, 260, 261, 262, 263, 264, 265, 287, 288, 289, 290, 291, 292, 293, 419, 420, 421, 422, 423, 459, 460, 461, 462, 463, 464, 465, 466 Group 2 Time Window: 12.5 min. – 21 min. Group 2 SIM Ions: 272, 273, 274, 275, 276, 277, 288, 289, 290, 291, 292, 293, 329, 330, 331, 332, 333, 334, 335, 346, 347, 348, 349, 350, 351, 419, 420, 421, 422, 423, 431, 432, 433, 434, 435, 436, 437, 459, 460, 461, 462, 463, 464, 465

180

Untargeted Metabolite Analysis

GC Derivatization: Methoxymation/trimethylsilylation Injection volume: 1μL Injection mode: Splitless Carrier gas: He Time program: Initial temp. 80°C; hold 2 min. Ramp 5°C/min. up to 325°C Hold 325°C; hold 10 min. MS Ion Source: Electron Ionization Ion Source Temp.: 280°C MS Quad. Temp.: 150°C Solvent Delay: 5.5 min. Acquisition Mode: Scan Scan Range: m/z 50 – 650

181 iii. Fragment Ions and Retention Times

Fatty Acid-TMS Derivatives Compound m/z Retention Time (min.) Lauric Acid 257 26.97 Myristic Acid 285 30.57 Palmitoleic Acid 311 33.54 Palmitic Acid 313 34.00 Heptadecanoic Acid 327 35.63 Oleic Acid 339 36.88 Stearic Acid 341 37.24 Cholesterol-TMS Derivative Cholesterol 368 46.57

Amino Acid-tBDMS Derivatives Compound m/z Retention Time (min.) Alanine 260 10.38 Asparagine 417 15.58 Aspartic Acid 390 14.70 Cysteine 406 20.49 Glycine 246 10.56 Histidine 440 17.15 Isoleucine 302 11.88 Leucine 302 11.64 Lysine 431 16.00 Methionine 320 13.54 Ornithine 417 12.81 Phenylalanine 308 14.35 Proline 286 12.16 Serine 364 12.68 Threonine 417 12.06 Tyrosine 466 12.69 Tryptophan 375 17.67 Valine 260 11.32

Methylated Fatty Acid Derivatives Compound m/z Retention Time (min.) 270 (M0), Palmitic Acid 14.84 286 (M16) Heptadecanoic Acid 284 16.12

Organic Acid-tBDMS Derivatives Compound m/z (M0) Retention Time (min.) Pyruvate 174 7.22 Lactate 261 9.26 Succinate 289 11.32 Fumarate 287 11.54 α-Ketoglutarate 346, 431 13.19 Malate 419 13.91 Tricarballylic Acid 461 15.01 Citrate 357, 431 16.95 Isocitrate 403, 431 17.05

Hexose-Aldonitrile/Pentaacetylation Derivatives Compound m/z Retention Time (min.) Glucose (M0) 191, 314 15.22 Galactose 314 15.50

182