SLC44A1 Transport of Choline and Ethanolamine in Disease

Adrian Taylor

A Thesis presented to The University of Guelph

In partial fulfilment of requirements for the degree of Doctor of Philosophy in Human Health and Nutritional Sciences

Guelph, Ontario, Canada

ABSTRACT

SLC44A1 TRANSPORT OF CHOLINE AND ETHANOLAMINE IN DISEASE

Adrian Taylor Advisor(s): University of Guelph, 2019 Marica Bakovic

Choline and ethanolamine are important molecules required for the de novo synthesis of phosphatidylcholine (PC) and phosphatidylethanolamine (PE) via the Kennedy pathway. Additionally, these two molecules are vital for maintaining both muscular and neurological function. The goal of this thesis was to gain insight into PC and PE metabolism with the use of unique metabolic disturbances ranging from obesity and genetic mutations in neurodegenerative disease.

Firstly, the protective effects of choline supplementation on muscular function were investigated within the Pcyt2+/- mouse model. In Pcyt2+/- mice, substrate flow through the CDP-ethanolamine branch of the Kennedy pathway was diminished resulting in triacylglycerol (TAG) accumulation and obesity. Supplemental choline improved muscle function by altering the expression of genes devoted to reducing TAG synthesis and restoring energy homeostasis.

With this new insight about the role of choline in regulating metabolism, the cellular uptake mechanism of choline was then analyzed. Skin fibroblasts from two patients with homozygous mutations in the SLC44A1 gene suffering from Neurodegeneration with

Brain Iron Accumulation (NBIA) were utilized. In these fibroblasts, SLC44A1 expression and choline uptake were drastically diminished. Moreover, PC levels were unaffected while PE levels were diminished relative to control, an indication of perturbed phospholipid homeostasis. Within this model of choline transport dysfunction, choline supplementation moderately improved phospholipid homeostasis.

In the literature, there are data which suggest that the same transporter might facilitate choline and ethanolamine uptake. This prompted us to examine the role of SLC44A1 in ethanolamine transport. SLC44A1 was overexpressed in cells to understand its influence in ethanolamine uptake. Moreover, cells with defects in SLC44A1 mediated transport were utilized to characterize ethanolamine uptake kinetics. Additionally, choline and ethanolamine appeared to compete for uptake while antibody inhibition of

SLC44A1 diminished PC and PE levels. Lastly, it was determined that ethanolamine uptake occurs in a pH dependent manner. SLC44A1 mediated uptake has previously been characterized as being pH dependent, which was further evidence implicating

SLC44A1 in ethanolamine transport.

This thesis solidifies the metabolic relationships between the CDP-choline and CDP- ethanolamine pathways and the connection between choline and ethanolamine in regulating lipid homeostasis.

ACKNOWLEDGEMENTS

To begin, I would like to thank my advisor Dr. Marica Bakovic for the opportunity to work in her lab. I love how well you synthesize ideas and I appreciate the intensity that you demonstrate in your work. I am also thankful for the opportunity to assume a leadership role within the lab throughout my time as a graduate student.

I would also like to thank my advisory committee Dr. David Ma and Dr. Kelly Meckling for their support and feedback throughout this journey.

Also, thank you to my PhD qualifying examination committee, Dr. David Ma, Dr. Alison

Duncan and Dr. Adronie Verbrugghe. This was a very enjoyable experience for me and you all helped me develop a stronger appreciation for the interplay between different fields within nutritional sciences.

Thank you to the collaborators who made this thesis possible. This includes (but is definitely not limited to) Audric Moses of the Women and Children’s Health Research

Institute at the University of Alberta, Michael Leadley of the Analytical Facility for

Bioactive Molecules at the Hospital for Sick Children and Dr. Vern Dolinsky at the

Children’s Hospital Research Institute of Manitoba for their lipidomics expertise. Also, a big thank you to Dr. Laila Schenkel for her help with the mouse project. This study definitely does not happen without you. Thank you to our collaborators Christina

Fagerberg and Felix Distelmaier in Europe for sharing the skin fibroblasts with us and their contributions with the NBIA study. Also, thank you to the patients who served as the inspiration for the inspiration for the NBIA study. v

Lab mates of the Bakovic lab! It’s incredibly heartwarming to say that the lab has developed a ‘family’ feel to it. I really appreciate that the lab is a place where everyone can freely discuss any issues, scientific or not so scientific.

Thank you to everyone in HHNS! I’m thrilled to have been a part of a highly interactive department with intramural teams, outings to Blue Jays games, Christmas parties and a bunch of other things. A special thank you to my long-time house/apartment mates Elie

Chamoun and Will Peppler for all the fun conversations and the life chats.

Thank you Guelph! I first came here for the Biomedical Toxicology program in

2008...and I stayed here until 2019. There are many good reasons for that and I feel blessed for that happening to me.

Thank you to everyone back home. I appreciate your blue collar, ‘salt of the Earth’ ways. There’s never a bad time to hang out, which was music to my ears as I weaved my way through my grad school journey.

To conclude, thank you to my girlfriend Carly for being smart and patient and witty and understanding all the way through grad school. Lastly, thank you to my mom and dad for the funny and thoughtful conversations around a dinner table. They’re honestly some of my fondest memories and it really made coming home for the holidays that much more awesome.

TABLE OF CONTENTS

Abstract…………………………………………………………………………………………...ii

Acknowledgements ...... iv

Table of Contents ...... ivi

List of Tables ...... xi

List of Figures ...... xii

List of Abbreviations ...... xiv

Studies ...... xix

CHAPTER 1: INTRODUCTION ...... 1

1.1 Significance of choline and ethanolamine ...... 1

1.2 Choline and ethanolamine formation and their implications in membrane phospholipid biogenesis ...... 2

1.3 Choline oxidation to betaine ...... 5

1.4 Choline and betaine protect against fatty liver and insulin resistance ...... 8

1.5 Phospholipids in skeletal muscle and mitochondrial function ...... 12

1.6 The role of choline and epigenetics in pregnancy and gestation ...... 14

1.7 Neuroprotective roles of choline ...... 17

1.8 Interplay between fatty acids and phospholipids in modulating membrane biophysics ...... 19

CHAPTER 2: HYPOTHESES, RATIONALE AND OBJECTIVES ...... 21

CHAPTER 3 RATIONALE ...... 24

CHAPTER 3: ADAPTATIONS TO EXCESS CHOLINE IN INSULIN RESISTANT AND PCYT2 DEFICIENT SKELETAL MUSCLE ...... 25

3.1 Abstract ...... 25

3.2 Introduction ...... 26 xi

3.3 Methods ...... 28

3.3.1 Animals and choline treatments ...... 28

3.3.2 Analysis of phospholipids and cholesterol ...... 29

3.3.3 Analysis of DAG and TAG ...... 30

3.3.4 Analysis of muscle glycogen ...... 30

3.3.5 Immunoblotting ...... 31

3.3.6 Analysis of gene expression ...... 32

3.3.7 Statistical analysis ...... 35

3.4 Results ...... 35

3.4.1 Choline affects membrane composition and reduces TAG accumulation in Pcyt2+/- muscle ...... 35

3.4.2 Choline helps restore Pcyt2+/- muscle glycogen ...... 39

3.4.3 Choline does not modify phospholipid gene expression ...... 39

3.4.4 Choline downregulates Pcyt2+/- muscle lipogenesis ...... 40

3.4.5 Choline improves AMPK activity and FA oxidation in Pcyt2+/- muscle glycogen ...... 42

3.4.6 Choline improves nutrient signaling in Pcyt2+/- muscle ...... 43

3.5 Discussion ...... 47

CHAPTER 4 RATIONALE ...... 52

CHAPTER 4:THE CHOLINE TRANSPORTER LIKE PROTEIN 1 IS INTEGRAL FOR MAINTAINING PHOSPHOLIPID AND FATTY ACID HOMEOSTASIS ...... 53

4.1 Abstract ...... 53

4.2 Introduction ...... 54

4.3 Methods ...... 56

4.3.1 Whole exome sequencing and Sanger sequencing ...... 56

viii

4.3.2 Isolation and treatment of skin fibroblasts ...... 58

4.3.3 RNA Isolation and RT-PCR ...... 59

4.3.4 Immunoblotting ...... 60

4.3.5 Choline uptake ...... 61

4.3.6 Electron microscopy ...... 62

4.3.7 Iron determination with ferrozine ...... 63

4.3.8 Pulse and pulse-chase radiolabeling with [3H]-choline and [14C]- ethanolamine...... 63

3 14 4.3.9 Equilibrium radiolabeling with [ H]-glycerol and [ C]-(CH3)-methionine .... 64

4.3.10 Phospholipase D activity assay ...... 65

4.3.11 Analysis of membrane phospholipids and fatty acids ...... 65

4.3.12 Data analysis ...... 66

4.4 Results ...... 66

4.4.1 Clinical description ...... 66

4.4.2 Cerebral MRI scans ...... 70

4.4.3 Genetic results ...... 70

4.4.4 Histological investigations ...... 72

4.4.5 M1 mutant produces a truncated CTL1 protein while M2 mutant results in no protein being produced ...... 74

4.4.6 Choline uptake in mutant fibroblasts is reduced but improves with choline treatment ...... 75

4.4.7 Altered expression of other choline transporters in mutant fibroblasts ...... 78

4.4.8 Changes in phospholipid gene expression as a likely measure to preserve membrane PC ...... 80

4.4.9 Increased expression of lipogenic genes and reduced expression of fatty acid oxidation genes as a response to deficiency in choline transport ...... 82 ix

4.4.10 Preservation of mutant cell PC results in deficiency in other membrane phospholipids ...... 84

4.4.11 Membrane phospholipid profiles are greatly modified in mutant cells ...... 88

4.5 Discussion ...... 91

CHAPTER 5 RATIONALE ...... 96

CHAPTER 5: THE NOVEL ROLE OF SLC44A1/CTL1 IN ETHANOLAMINE TRANSPORT ...... 97

5.1 Abstract ...... 97

5.2 Introduction ...... 98

5.3 Methods ...... 100

5.3.1 Maintenance of cell lines ...... 100

5.3.2 Drug treatments in cancer cells ...... 101

5.3.3 Radiolabeling and PC and PE determination ...... 101

5.3.4 Kinetics of ethanolamine transport ...... 102

5.3.5 Choline and ethanolamine uptake ...... 102

5.3.6 RNA extraction and RT-PCR ...... 103

5.3.7 Expression of Myc-tagged rSLC44A1 cDNA and SLC44A2 siRNA in fibroblasts ...... 104

5.3.8 Immunoblotting ...... 105

5.3.9 Mitochondrial isolation ...... 105

5.3.10 Lipid analysis ...... 106

5.3.11 Pcyt2 activity assay ...... 106

5.3.12 Statistical analysis ...... 107

5.4 Results ...... 107

5.4.1 CTL1 inhibition with HC-3 and CTL1 antibody on PC and PE synthesis . 107

5.4.2 Kinetics of ethanolamine uptake and effects of extracellular pH and Na+ on ethanolamine uptake ...... 108 x

5.4.3 Effects of various compounds on CTL1 mediated uptake ...... 112

5.4.4 mRNA expression of CTL transporters and Kennedy pathway genes ..... 114

5.4.5 Effects of CTL1 overexpression on ethanolamine uptake ...... 116

5.4.6 Effects of CTL2 knockdown on ethanolamine uptake ...... 117

5.4.7 Lipid analysis in Control and M2 cells ...... 120

5.5 Discussion ...... 121

CHAPTER 6: INTEGRATIVE DISCUSSION ...... 125

6.1 Choline treatment in regulating lipid metabolism ...... 125

6.2 Consequences of CTL1/SLC44A1 deficiency ...... 127

6.3 Regulation of ethanolamine transport by CTL1/SLC44A1 ...... 129

6.4 Integration of CDP-choline and CDP-ethanolamine pathways ...... 130

6.5 Conclusion and future directions ...... 132

References ...... 134

LIST OF TABLES

Table 3.1: Primer sequences and melting temperatures (TM) for polymerase chain reaction (PCR) using mouse RNA...... 34

Table 4.1: Primer sequences and melting temperatures (TM) for polymerase chain reaction (PCR) using human fibroblast RNA...... 60

Table 5.1: Primer sequences and melting temperatures (TM) for polymerase chain reaction (PCR) using human fibroblast RNA...... 104

xiv

LIST OF FIGURES

Figure 1.1: Choline and ethanolamine in the Kennedy pathway...... 4

Figure 1.2: Choline and the interplay between the de novo synthesis, transsulfuration and mitochondrial oxidative pathways...... 7

Figure 3.1: Choline modifies muscle glycerolipid content...... 37

Figure 3.2: Choline modifies muscle TAG and DAG content and glycogen storage. .... 38

Figure 3.3: Choline positively affects the expression of genes involved in lipogenesis and lipolysis...... 41

Figure 3.4: Choline has a positive effect on the expression of key players in fatty acid oxidation...... 43

Figure 3.5: Choline reduces PKCα and restored muscle insulin signaling...... 45

Figure 3.6: Choline reduces mTOR activation and increases expression of key players in autophagy...... 46

Figure 4.1: Photos of the individuals, pedigrees and brain MRI studies...... 71

Figure 4.2: Ultrastructure and iron load of cultured skin fibroblasts...... 73

Figure 4.3: Topology and transport function of the native and mutated CTL1 proteins. 76

Figure 4.4: Choline stimulates CTL1 transport and protein abundance in M1 cells...... 77

Figure 4.5: mRNA expression of choline related transporters is dysregulated in mutant fibroblasts...... 79

Figure 4.6: mRNA expression of phospholipid genes is dysregulated in mutant fibroblasts...... 81

Figure 4.7: mRNA expression of fatty acid genes is dysregulated in mutant fibroblasts...... 83

Figure 4.8: PC and PE de novo biosynthesis and turnover ...... 86

Figure 4.9: Analysis of equilibrium phospholipid levels ...... 87

Figure 4.10: Glycerolipid subclasses and fatty acid composition are modified in mutant cells ...... 90

Figure 5.1: CTL1 inhibition with HC-3 and CTL1 antibody on PC and PE synthesis ... 108

xiii

Figure 5.2: Kinetics of ethanolamine uptake ...... 110

Figure 5.3: Effects of extracellular pH and Na+ on ethanolamine uptake ...... 111

Figure 5.4: Effects of various compounds on CTL1 mediated uptake ...... 113

Figure 5.5: mRNA expression of CTL transporters and Kennedy pathway genes and Pcyt2 activity assay ...... 115

Figure 5.6: Effects of CTL1 overexpression on ethanolamine uptake ...... 117

Figure 5.7: Characteristics of mitochondrial ethanolamine transport ...... 119

Figure 5.8: Lipid analysis in Control and M2 cells ...... 121

xiv

LIST OF ABBREVIATIONS

5,10-MTHF: N5, N10-methyleneTHF

AA: arachidonic acid

ACC: acetyl-CoA carboxylase

Ach: acetylcholine

AD: Alzheimer’s disease

ADP: adenosine diphosphate

AI: adequate intake

Akt: protein kinase B

AMPK: 5’ adenosine monophosphate-activated protein kinase

ANT: adenine nucleotide translocase

ATGL: adipose triglyceride lipase

ATP: adenosine triphosphate

BADH: betaine aldehyde dehydrogenase

BHMT: betaine homocysteine methyltransferase

CDP-choline: cytidine 5’-diphosphocholine

CE: cholesterol ester

CEPT: choline ethanolamine phosphotransferase 1

CHDH: choline dehydrogenase

CHT1: high-affinity choline transporter

CK: choline kinase

CoA: coenzyme A

CPT1: carnitine palmitoyltransferase I

CTL1: choline transporter-like 1

xvi

CVD: cardiovascular disease

DAG: diacylglycerol

DGAT: diacylglycerol acyltransferase

DHA: docosahexaenoic acid

DMG: dimethylglycine

DNA: deoxyribonucleic acid

DTT: dithiothreitol

EDTA: ethylenediaminetetraacetic acid

EK: ethanolamine kinase

ELOVL: fatty acid elongase

EPA: eicosapentaenoic acid

ER: endoplasmic reticulum

ERG: electroretinography

FA: fatty acid

FAS: fatty acid synthase

FC: free cholesterol

FMO3: flavin monooxygenase 3

GAPDH: glyceraldehyde-3-phosphate dehydrogenase

GPC: glycerophosphocholine

GSH: glutathione

GSK3: glycogen synthase kinase 3

HC-3: hemicholinium-3

HFD: high fat diet

Hcy: homocysteine

HPLC: high performance liquid chromatography xvii

HSL: hormone sensitive lipase

IR: insulin resistance

IRS1: insulin receptor substrate 1

LC3: microtubule-associated protein light chain 3

LPL: lipoprotein lipase

MAM: mitochondrial associated ER membranes

MAT: methionine adenosyl transferase

MPAN: mitochondrial membrane protein-associated neurodegeneration

MRI: magnetic resonance imaging

MTHFR: N5, N10-methyleneTHF reductase mTOR: mammalian target of rapamycin

NAFLD: non-alcoholic fatty liver disease

NBIA: neurodegeneration with brain iron accumulation

NHANES: National Health and Nutrition Examination Survey

NHE: sodium/proton exchanger

OCTN: organic cation transporter

PA: phosphatidic acid

PBS-T: phosphate buffered saline-tween 20

PC: phosphatidylcholine

PCho: phosphocholine

Pcyt1: choline-phosphate cytidylyltransferase 1

Pcyt2: CTP:phosphoethanolamine cytidylyltransferase 2

PDH: pyruvate dehydrogenase

PE: phosphatidylethanolamine

PEMT: phosphatidylethanolamine N-methyltransferase xvii

PG: phosphatidylglycerol

PGC-1α: peroxisome proliferator-activated receptor gamma coactivator 1-alpha

Pi: inorganic phosphate

PI: phosphatidylinositol

PI3K: phosphoinositide 3-kinase

PKCα: protein kinase C alpha

PLA: phospholipase A

PLD: phospholipase D

POTS: postural orthostatic tachycardia syndrome

PPARα: peroxisome proliferator-activated receptor alpha

PPARγ: peroxisome proliferator-activated receptor gamma

PS: phosphatidylserine

PSD: phosphatidylserine decarboxylase

PSS1: phosphatidylserine synthase 1

PSS2: phosphatidylserine synthase 2

PUFA: polyunsaturated fatty acid

SAH: s-adenosylhomocysteine

SAHH: s-adenosylhomocysteine hydrolase

SAM: s-adenosylmethionine

SCD1: stearoyl-CoA desaturase-1

SCP2: sterol carrier protein 2

SHMT: serine hydroxymethyltransferase

SIRT1: sirtuin 1

SLC44A1: solute carrier family 44 subfamily A member 1

SLC5A7: solute carrier family 5 subfamily A member 7 xviii

SM: sphingomyelin

SNP: single nucleotide polymorphism

SREBP1: sterol regulatory element-binding protein 1

T2D: type 2 diabetes

TAG: triacylglycerol

TCA: tricarboxylic acid

THF: tetrahydrofolate

TLC: thin layer chromatography

TMA: trimethylamine

TMAO: trimethylamine N-oxide

VLCFA: very long chain fatty acids

VLDL: very low density lipoprotein

xix

STUDIES

This thesis is based on the following studies:

Chapter 1: Taylor A, Michel V, Bakovic M. (2019). Metabolic and physiologic significance of choline formation and metabolism.

Chapter 3: Taylor A, Schenkel, LC, Yokich M, Bakovic M. (2017). Adaptations to

Excess Choline in Insulin Resistant and Pcyt2 Deficient Skeletal Muscle. Biochemistry and Cell Biology. DOI: 10.1139/bcb-2016-0105.

Chapter 4: Fagerberg CR, Taylor A, Distelmaier F, Kibaek M, Wieczorek D, Schroder

HD, Larsen, MJ, Jamra, RA, Gade, E, Markovic, L, Klee, D, Brady, L, Tarnopolsky, M,

Agarwal, P, Dolinsky V, Bakovic M. (2019). A new type of Neurodegeneration with Brain

Iron Accumulation is linked to the Choline Transporter Like 1 Protein. Submitted to

Brain.

Chapter 5: Taylor A, Grapentine, S, Ichhpuniani J, Bakovic M. (2019). The novel role of

SLC44A1/CTL1 in ethanolamine transport.

CHAPTER 1: INTRODUCTION

1.1 Significance of choline and ethanolamine

Choline is an essential nutrient involved in membrane lipid formation, neurotransmission and one carbon metabolism. In addition to free choline, most dietary choline is produced from the phospholipid phosphatidylcholine (PC) (1). Choline Adequate Intake (AI) of 550 mg/day for men, 425 mg/day for women, 450 mg/day for pregnant women and 550 mg/day for lactating women and has been considered a required dietary nutrient by the

US Institute of Medicine’s Food and Nutrition Board since 1998. However, numerous studies have reported that 4 in 5 Americans are not reaching the AI for choline. The

Framingham Heart Study reported that Americans consumed on average 203 mg/day of choline whereas the Nurses’ Health Study and the Atherosclerosis Risk in Communities study reported 293 and 217 mg/day choline intake respectively, far below the AI value

(2, 3, 4, 5). Choline is most abundant in eggs, meat and dairy products (6). To help improve choline intake, choline rich cereal based functional foods (7), milk (8) and goats

(9) have been developed.

Choline deficiency is implicated in fatty liver (10), insulin resistance (11) and muscle damage in humans (12) and rodents (13). Humans who have mutations in PCYT1A, the gene that encodes the rate limiting enzyme for PC synthesis via the CDP-choline pathway, have been shown to have severe insulin resistance and lipodystrophy (14). In mice, choline has been shown to be particularly important during fetal development as gestational deficiency leads to memory and learning deficits later in life (15).

Ethanolamine is essential for life and is typically found in circulation at a concentration of approximately 50 μM (16, 17). In addition, ethanolamine is a structural component of mammalian cell membranes as a constituent of the abundant membrane phospholipid phosphatidylethanolamine (PE) (16). Moreover, PE is critical for maintaining mitochondrial membrane morphology (18) by generating negative membrane curvature that allows cristae to have their long, tubular structure (19). Aside from mitochondrial biogenesis, the implications of ethanolamine are quite diverse: it has been used for cancer diagnostics (20), is likely involved in the pathophysiology of bipolar disorder (21) and modulates cell proliferation (22). Ethanolamine can also serve as a sole carbon and nitrogen source for bacteria (23) and is readily found in cosmetics as a wetting agent

(24).

1.2 Choline and ethanolamine formation and their implications in membrane phospholipid biogenesis

Choline and ethanolamine are critical components of PC and PE respectively (25, 26,

27). For the purposes of phospholipid biosynthesis, choline is transported into the cell via the choline transporter-like protein 1 (CTL1). The uptake mechanism for ethanolamine is unknown but past reports have found that choline and ethanolamine compete with one another for uptake (28, 29), hinting that CTL1 can facilitate ethanolamine uptake. De novo synthesis of PC and PE occur via the Kennedy pathway

(Figure 1.1): Upon entry into the cell, choline and ethanolamine are quickly

phosphorylated in the cytoplasm by choline kinase (CK) and ethanolamine kinase (EK) to generate phosphocholine and phosphoethanolamine (PCho and PEtn). PCho is then coupled with CTP by the CTP:phosphocholine cytidylyltransferase (CCT/Pcyt1) to generate CDP-choline and Pi. PEtn is coupled with CTP by the

CTP:phosphoethanolamine cytidylyltransferase (ECT/Pcyt2) to generate CDP- ethanolamine and Pi. In the final step, CDP-choline and CDP-ethanolamine are condensed with diacylglycerol (DAG) by CDP-choline:DAG phosphotransferase and

CDP-ethanolamine:DAG phosphotransferase in the endoplasmic reticulum (ER) to yield

PC and PE. In the liver, PC is additionally generated from PE by a 3-step methylation with S-adenosylmethionine (SAM) catalyzed by phosphatidylethanolamine (PE) N- methyltransferase (PEMT) (30). In mammals, the PEMT pathway is vital for PC formation to prevent the development of choline deficiencies when demands for choline are high, such as during embryonic development, pregnancy, and lactation (31).

Moreover, PEMT is important for VLDL particle formation and secretion from the liver

(32). PEMT formed PC is enriched with n-3 PUFA, which is crucial for providing fetuses with n-3 PUFA for proper developmental programming (33) and the optimal function of neurological tissues (34, 35, 36).

Figure 1.1: Choline and ethanolamine in the Kennedy pathway

Choline and ethanolamine are released from PC and PE by phospholipases and by phospholipid exchange mechanisms. Degradation of PC and PE by phospholipases is a critical element of cell signaling and choline homeostasis (37). PC is converted into glycerophosphocholine (GPC), a prominent acetylcholine precursor and osmolyte via

PLA2 (38) while PE is converted to glycerophosphoethanolamine. PLD1 and PLD2 cleave PC and PE between the phosphate moiety and choline or ethanolamine head group to generate free choline or ethanolamine and phosphatidic acid (PA). In mice, PA has been shown to be utilized for DAG formation and directly regulates signaling pathways such as the mTOR pathway (39). Moreover, in genetic disorders with choline deficiencies such as postural orthostatic tachycardia syndrome (POTS), there is an

increase in PLD activity apparently in an attempt to generate more intracellular choline when choline transport is reduced (40).

Free choline is also released at the mitochondria associated ER membranes (MAM) when PC is converted into phosphatidylserine (PS) by PS synthase 1 (PSS1) but the exact utilization of the choline formed at the MAM is not known. PE also exchanges its ethanolamine head group with serine to produce PS via PS synthase 2 (PSS2) (41).

Mitochondrial PS can be decarboxylated to generate PE by PS decarboxylase (PSD)

(41, 42). This newly made PE can be transported from mitochondria and further methylated by S-adenosylmethionine (SAM) in the liver by the PEMT pathway, to recycle PC (and intracellular choline) (43). As a result, the PE phospholipid head group exchange mechanisms and PS decarboxylation pathway also serve to generate additional pools of choline, the fate of which is not clearly established.

1.3 Choline oxidation to betaine

Aside from its role in PC synthesis, choline is oxidized to betaine, which is a prominent methyl group donor in one-carbon metabolism and SAM formation (Figure 2). In the liver and kidney, choline is oxidized to betaine by two poorly characterized enzymes, choline dehydrogenase (CHDH) and betaine aldehyde dehydrogenase (BADH), located within the mitochondrial matrix. Betaine remethylates homocysteine (Hcy) to generate methionine, and this reaction is carried out by betaine homocysteine methyltransferase

(BHMT) when dimethylglycine (DMG) is released. Furthermore, it is important to note

that the betaine product DMG is also a methyl group donor for mitochondrial tetrahydrofolate (THF) methylation to 5 methyl-THF and the completion of the sarcosine-glycine-serine cycle (Figure 2) that takes part both in the mitochondria and in cytosol. As a result, the production and availability of betaine is important for optimal one carbon metabolism in three ways: i.) the methyl groups of betaine can be donated into the one carbon cycle for SAM production ii.) betaine also supports the folate cycle via mitochondrial methylation of THF to generate 5-methylTHF and iii.) betaine is also capable of supplying the methyl donor serine to generate N5, N10-methyleneTHF (5,10-

MTHF). Therefore, differently from folate, betaine is a direct provider of methyl groups for Hcy methylation to methionine. In the folate cycle, serine hydroxymethyltransferase

(SHMT) catalyzes the donation of a methyl group from serine to synthesize 5, 10-MTHF from THF (44, 45, 46), which can then be reduced to 5-methylTHF by N5, N10- methyleneTHF reductase (MTHFR). Paradoxically, the metabolic dependence of the folate cycle from mitochondrial choline/betaine metabolism is not well understood.

Moreover, its significance is not well addressed when it comes to human nutrition and epigenetic consequences for human health. SAM is synthesized by the condensation of methionine and adenosine by methionine adenosyl transferase (MAT). SAM is the only methyl group donor implicated in multiple metabolic reactions including DNA methylation, neurotransmitter synthesis and PC synthesis (47). Due to the methylation reaction, SAM is converted into S-adenosylhomocysteine (SAH) which is then hydrolyzed to Hcy by SAH hydrolase (SAHH) (Figure 2). Hcy can also be converted into

cysteine via transsulfuration to be incorporated into glutathione (GSH) which is one of the most important intracellular reducing agents.

Figure 1.2: Choline and the interplay between the de novo synthesis, transsulfuration and mitochondrial oxidative pathways

In mice, methyl donor supplementation in vivo has also been shown to positively affect one carbon metabolism to correct perturbations in FA oxidation and energy balance which are evident in obesity. In mice with excess fat accumulation, 4 weeks of choline supplementation increased plasma glycerol and glycine, which are products of increased lipolysis and choline oxidation respectively (40). Moreover, plasma creatine

and sarcosine were reduced, further indicating an increase in homocysteine/methionine cycling to breakdown choline (40). In this model, SAM was more devoted to maintaining cellular PC balance, which is often perturbed in obesity, instead of guanidinoacetate methylation for creatine synthesis. In the skeletal muscle of mice, the main beneficial effects of supplemental choline included the reduction of energy stored as free FA, DAG and TAG and increasing energy utilization (76). Mice with excess fat accumulation supplemented with choline or supplemented for 8 weeks with betaine showed increased oxidative demethylation of these methyl donors (40). Oxidative demethylation of choline and betaine is beneficial as this requires energy in the form of reducing equivalents produced from succinate and α-ketoglutarate in the TCA cycle to be broken down (40).

In essence, choline and betaine breakdown increased metabolic demand, thereby decreasing the propensity for energy to be stored as fat throughout the body.

1.4 Choline and betaine protect against fatty liver and insulin resistance

Non-alcoholic fatty liver disease (NAFLD) has become the most prominent chronic disease in humans and its prevalence has risen concomitant with insulin resistance, type 2 diabetes (T2D) and obesity over the past 30 years (48). NAFLD comprises a wide spectrum of pathologies, ranging from lipid droplets and simple steatosis to hepatitis to fibrosis and cirrhosis (49). Dietary factors such as high caloric intake, high fat intake (50), high fructose intake, refined grain and processed meat consumption have been positively correlated with NAFLD development (51). Though extensively

studied, the mechanism responsible for the pathogenesis of NAFLD remains poorly understood.

Dietary choline is an important nutrient for maintaining optimal hepatic function. Choline deficiency has widespread effects on the one carbon metabolic system and lipid synthesis in the liver. In choline deficient mice, SAM concentrations were decreased by

50% after the consumption of a choline deficient diet for 2 weeks (52). Due to this decrease in SAM concentration, there is a concomitant decrease in PEMT activity, and therefore PE derived PC phospholipid content (53). The PEMT enzyme is important for the synthesis of PC used in very low-density lipoproteins (VLDL), which is critical for

TAG export from the liver (54). PC is a critical component of VLDL particles and a diminished capacity for the liver to synthesize PC via the PEMT pathway results in decreased lipoprotein secretion and TAG accumulation in hepatocytes (55). Hepatic steatosis resulting from choline deficiency is a hallmark feature of VLDL deficiency in species such as humans and cats (52, 56). Choline supplementation improves hepatic function in mice by normalizing the expression of genes involved in lipogenesis

(SREBP1, FAS, SCD1) and lipolysis (ATGL, HSL, LPL), as well as FA oxidation

(PPARα) and mitochondrial biogenesis (PGC-1α) (40). These changes result in a decrease in hepatic TAG (40, 57) and plasma acylcarnitines, which is an indication of improvements in mitochondrial fatty acid metabolism (40, 58).

Betaine has been shown to be hepatoprotective in mice with respect to numerous toxic substances such as ethanol, lipopolysaccharide and dimethylnitrosamine (59, 60, 61).

The liver injury resulting from these toxicants is largely due to altered sulfur amino acid

metabolism, and betaine has been shown to be beneficial in this regard (62, 63).

Hepatocyte cell volume in mammalian systems is often altered during oxidative stress and these changes in cell volume activate signal transduction cascades in an attempt to allow the cell to respond to stress more effectively (64, 65). BHMT, CHDH and PEMT expression is increased in hypotonic conditions, indicating a coordinated response to increase cell volume and regenerate methionine from Hcy for eventual methyl group donation (66). Additionally, betaine is a lipotropic compound and also helps to increase the levels of SAM and GSH in the liver which are critical for maintaining proper methylation and redox states (67).

Betaine has been shown in mice to reduce body weight induced by HFD consumption as well as hepatic and visceral fat mass by increasing AMPK activation (68). AMPK is a positive regulator of fatty acid oxidation and decreases the expression of genes such as

SREBP-1c, ACC and FAS which promote lipogenesis (69). Furthermore, AMPK can directly phosphorylate PGC-1α to stimulate mitochondrial biogenesis (70). Betaine has also been shown to reverse increased serum insulin levels and improve glucose tolerance (71) while also reducing serum TAG and cholesterol levels. These findings are supplemented by the activation of insulin signaling in the liver as seen by increasing phosphorylation of IRS1 and activation of Akt (72).

In mice, plasma acyl-carnitine levels are lowered substantially with choline and betaine supplementation, which is beneficial as an abundance of acyl-carnitines in the plasma is indicative of inefficient mitochondrial FA oxidation, a metabolic issue that often affects obese individuals (40, 57). As a result, acyl-carnitine levels can provide an indication of

the ability of the cell to oxidize FA (73) and to maintain optimal mitochondrial function

(74). In mice, long chain acyl-CoA species are critical regulators of metabolism as they inhibit mitochondrial adenine nucleotide translocase (ANT), which regulates ATP/ADP exchange across the inner mitochondrial membrane (75). Additionally, malonyl-CoA is an allosteric inhibitor of CPT1, the rate limiting enzyme in the oxidation of long chain fatty acyl-CoA by facilitating its entry into the mitochondrial matrix. Moreover, choline supplementation, decreased expression of genes which are critical for fatty acid synthesis (FAS and ACC) while the expression of genes involved with FA oxidation was increased (PPARα and PGC-1α) (76).

Supplementation of choline and betaine in vivo has been shown to ameliorate perturbations with lipid metabolism that arise in obesity. Choline and betaine supplementation in mice decreased liver and adipose tissue weight while also reducing lipid droplet size which is largely due to decreased FA incorporation into TAG (40). In addition, betaine supplementation significantly decreased plasma TAG content while choline supplementation reduced collagen deposition, indicating decreased inflammation and fibrosis throughout the liver (40). Membrane composition was also remodeled with choline supplementation as the ratio of important lipid raft components

FC:SM was increased (76). The optimal balance between FA oxidation and lipogenesis was restored with choline supplementation by activating the main regulator of skeletal muscle FA oxidation AMPK (76, 77). Subsequently, mTORC1 activation was decreased which in turn facilitate a decrease in SREBP1c nuclear translocation for lipogenic gene transcription (76). With choline supplementation, glycogen content, which is often

depleted in insulin resistant skeletal muscle, was also restored, indicating an enhanced ability to store glucose taken up from the blood.

Therefore, choline and betaine are critical for mitigating the development of NAFLD.

Choline and betaine both decrease lipid accumulation in the liver while also decreasing hepatic fibrosis. Metabolism is impacted by stimulating TAG degradation by lipolysis to generate a fasted energy deficient state. Choline is the predominant phospholipid component of VLDL which is important for exporting TAG from the liver. Improving choline and betaine levels promote fatty acid oxidation by increasing AMPK activation and improving insulin signaling.

1.5 Phospholipids in skeletal muscle and mitochondrial function

Skeletal muscle is a major site for energy expenditure and fatty acid oxidation and muscle mass is inversely correlated with health ramifications such as falls, fractures and poor quality of life (78, 79, 80). Miyake et al. (2019) concluded that the preservation of skeletal muscle mass is negatively correlated with all-cause mortality in individuals suffering from type 2 diabetes mellitus (81). Additionally, Hou et al. (2019) found that protein supplementation helped mitigate losses in maintain muscle mass in elderly individuals (82). Mutations which is affect lipid oxidation and mitochondrial respiration are detrimental to skeletal muscle function, leading to severe muscle pathologies (83).

In particular, loss of function mutations in genes that encode enzymes that help maintain phospholipid homeostasis can promote rhabdomyolysis, or rapid skeletal

muscle breakdown. An example is LPIN1, which encodes for a phosphatidate phosphatase to generate DAG required phospholipid synthesis. Additionally, Lipin1 can serve as a transcriptional co-activator of PPARα and PGC-1α (84), which are key for fatty acid oxidation and mitochondrial biogenesis. The link between mutations in LPIN1 and rhabdomyolysis in children has not been elucidated but likely involves an overaccumulation of intramuscular phospholipids and sarcoplasmic reticulum stress

(85).

Skeletal muscle is an important site for mitochondrial fatty acid oxidation, and phospholipids are also integral to optimal mitochondrial function (86, 87, 88).

Mitochondria have an atypical phospholipid composition whereby PE and cardiolipin

(CL) comprise roughly 50% of the phospholipid content (89). The major reasons for this enrichment of PE and CL in mitochondria are two-fold: these lipids induce negative membrane curvature to give cristae their tubular shape (90) and to help anchor the complexes of the electron transport chain in the inner mitochondrial membrane (91).

Additionally, fatty acid composition within these phospholipids plays an important role in mitochondrial function (92). Approximately 90% of mitochondrial cardiolipin species in skeletal muscle exist as tetralinoleoylcardiolipin (93), and alterations in cardiolipin configuration are often indicative of oxidative stress (94). Diminished PUFA content in mitochondrial phospholipids has been correlated with longevity (95), potentially due to a greater likelihood of these molecules to be peroxidized by reactive oxygen species (96).

On the other hand, As stated in section 1.2, PSD is an enzyme specific to mitochondria which is important as PE created via CDP-ethanolamine does not substantially

contribute to mitochondrial PE content (97). Moreover, mitochondria from fibroblasts lacking the PSD enzyme appear swollen and fragmented and consequently and this is linked to the depression of electron transport chain function and ATP synthesis (97).

1.6 The role of choline and epigenetics in pregnancy and gestation

During pregnancy, maternal choline affects metabolic and physiological functions of the offspring through numerous inter-related mechanisms. Maternal choline is vital for optimal placental development and maintaining optimal fetal growth (98, 99). The increased requirement for choline during gestation is largely due to an increased use of betaine as a methyl group donor (100) and a higher demand for choline utilization for

PC synthesis (101). This could lead to substantial depletion of choline derived methyl donors such as betaine and SAM in pregnant women (102, 103). A reduction in choline and betaine leads to elevated homocysteine and decreased SAM levels, thereby decreasing methyltransferase activity as SAM is a positive regulator of MAT (104).

Genetic variants that increase choline requirements can leave an individual susceptible to choline inadequacy (105), and this effect can be magnified during reproduction (106).

Some of these SNPs are located in genes such as PEMT (rs12325817), choline dehydrogenase (CHDH) (rs12676) and MTHFR (rs1801133) which are critical for proper functioning of the one carbon metabolic system (107, 108). In addition, estimations from the NHANES study estimated that only 1 out of 10 pregnant women in the US are

consuming adequate amounts of choline (109), which is a troubling statistic as choline is a vital nutrient for fetal development.

The prenatal period is vital for the establishment and maintenance of the epigenome

(110). The DNA methylation patterns of gametes are mostly abolished after conception, therefore de novo methylation is crucial for gene silencing (111). DNA methylation is closely linked to histone modification in order to facilitate time-sensitive alterations throughout fetal development (112). In addition, the fetal epigenome can be affected by numerous maternal environmental factors such as nutrition (113). During pregnancy, it has been determined that maternal choline supply has dramatic effects on the fetal epigenome (114). The fetuses of choline deficient mothers exhibited hypermethylation of the insulin growth factor 2 (IGF2) gene (115), which is critical for embryonic development (116). Additionally, choline deficient mothers exhibited hypomethylation of

DNMT1 and this was correlated with the epigenetic and expression changes of IGF2, suggesting that maternal choline deficiency exerted its effects on IGF2 via DNMT1 hypomethylation (115).

Maternal choline intake during pregnancy is critical for optimal cognitive function for a variety of reasons (116, 117).There is substantial evidence that adequate maternal choline levels during pregnancy are required for optimal hippocampal function, and therefore maintaining cognitive abilities (118, 119, 120). Choline deficiency during pregnancy has been shown to decrease hippocampal methylation of the cyclin dependent kinase inhibitor 3 (CDKN3) gene and increase the expression of kinase associated phosphatase (Kap), a known inhibitor of cell proliferation (121). Maternal PC

levels during pregnancy alter hippocampal cholinergic function of the offspring, which is associated with depression and anxiety later in life (122). This is because PC is important for ensuring adequate neuron density (123), propagation of intracellular signals (124) and optimal membrane configuration (125). PC can also be used for acetylcholine synthesis, and therefore cholinergic transmission (126). Choline intake is an important component of neurological development (127) as it has a large influence on structural integrity, function and cell signaling within the brain (128, 129).

Additionally, choline supplementation in pregnant women has been shown to increase placental FATP4 content, which transports DHA to the fetus for neurological development (130). Moreover, maternal choline supplementation increases the levels of

DHA-enriched PC (PC-DHA), which is mainly synthesized by the PEMT pathway (33).

The PEMT pathway is especially important for fortifying phospholipids with DHA as this enzyme prefers PE-DHA as a substrate (131). This is in contrast to the CDP-choline pathway which predominantly synthesizes PC containing saturated medium length FA

(120). With this in mind, maternal choline supplementation can serve as a dietary approach to supply the developing fetus with DHA. Umbilical cord choline content can be up to 5-fold higher than what is observed in maternal blood, further emphasizing the importance of choline in fetal development (96). Furthermore, the PEMT gene has an estrogen response element within its promoter region, meaning that its expression can be induced when estrogen is present (132). Estrogen is typically abundantly during pregnancy, and is therefore important for the fortification of the developing fetus with

PC-DHA (132). In fact, women with SNPs in within the estrogen response element are

much more susceptible to developing choline deficiency (133). Prenatal omega-3 fatty acid supplementation in pregnant dams that consuming a diet low in folate and vitamin

B12 normalizes global DNA methylation levels in the placenta and the brain (134). This implies that omega-3 fatty acids are involved in modifying methylation patterns (135).

Taken together, choline and PC have critical roles with respect to pregnancy and fetal development. Choline is a critical molecule which can ultimately serve as a methyl donor within the one carbon metabolic system. As a result, choline deficiency has a negative impact on fetal development due to perturbed epigenetic regulation. In addition, choline deficiency has detrimental effects on hippocampal development, and can lead to cognitive deficiencies. Lastly, PC is an important molecule for fetal DHA enrichment and hormones (ie. estrogen) and enzymes (PEMT) help facilitate this process.

1.7 Neuroprotective roles of choline

Choline levels have been shown to be integral to maintaining optimal neurological function over time. As the number of older individuals in our population increases, the impact of choline deficiency on the health of these populations becomes more important. With aging, cerebral function becomes impaired through myelin degradation, decreased synaptic function and dysregulation of DNA methylation. Due to these pathologies, older adults experience cognitive decline and are increasingly affected by neurological disorders like Alzheimer’s disease (AD). Studies have established a

positive correlation between individuals carrying mutations which alter MTHFR activity and AD, linking choline metabolism and neurological function. Choline can act as a reserve methyl donor when MTHFR function is diminished but this will deplete choline pools that are required for various neurological functions. These functions include the integration of choline into the neurotransmitter acetylcholine and lipids like PC, which are associated with hallmark neurological perturbations of AD such as memory impairment and anxiety (136). Moreover, the quantity of lysoPC species, which are pro- inflammatory lipid mediators, as well as sphingolipid breakdown products have been found to be markedly increased in AD patients (137, 138).

The focus on nutrition has become increasingly important with regards to neurological disorders, largely because synaptic membranes require numerous nutrients to be synthesized. van Wijk et al. (139) report that cognitively impaired subjects had lower circulating levels of choline and folate, both of which are fundamental to membrane phospholipid synthesis. Mellott et al. (140) have shown that neonatal choline consumption can mitigate AD related cognitive decline later in life by attenuating amyloid plaque formation. As in other models, in utero choline supplementation has been shown to prevent a decline in hippocampal neurogenesis in adulthood while also rescuing cholinergic function, which are hallmarks of AD (140, 141). In mice, maternal choline supplementation has therapeutic potential by normalizing the expression of genes involved with synaptic plasticity in offspring. This is key for protecting basal forebrain cholinergic neurons mitigating decline in spatial cognition (142, 143). Cytidine

5’-Diphosphocholine (citicoline), a PC precursor, has also been shown to improve

cognitive performance in AD patients as a safe and effective agent to increase PC levels in the brain (144, 145).

1.8 Interplay between fatty acids and phospholipids in modulating membrane biophysics

It is widely conceived that fatty acid esterification in critical for efficiently incorporating n-

3 fatty acids into the brain (146, 147), and that phospholipids are important carriers of n-

3 fatty acids (148). Esterified fatty acids are more readily absorbed into the body relative to fatty acids in TAG or unesterified fatty acids (149). Additionally, n-3 fatty acids are more resistant to oxidation when incorporated into phospholipids relative to TAG (150).

Moreover, fatty acids esterified in phospholipids relative to TAG are more bioavailable

(151), which is especially important in brain development (152, 153).

An important role of n-3 fatty acids is their role in modulating biophysical properties within the cell membrane which play a role in signal transduction cascades (154, 155,

156). For instance, Li et al. (2007) demonstrated that plasma membrane DHA content is integral for elevating eNOS activity by altering the lipid:protein interactions in caveolar microdomains, thereby facilitating the translocation of eNOS from the plasma membrane (157). Moreover, glucose transport has been shown to be modulated by the effects of n-3 fatty acids on its configuration within the membrane (158, 159, 160).

Imbalances in the activity of enzymes that maintain phospholipid homeostasis can facilitate pathological conditions resulting from membrane perturbations (161).

2+ Mutations in the PLA2G6 gene which encodes Ca dependent phospholipase A2

(iPLA2) have been linked to infantile neuroaxonal dystrophy which is characterized by motor and sensory impairment (162). iPLA2 works to breakdown PC, generating free fatty acids and lysoPC and serves as an antagonist to Pcyt1, which is a key enzyme in

PC production (163). Imbalances between the activity of iPLA2 and Pcyt1 can lead to lipid imbalances which are detrimental for membrane integrity (164). One of the lipids that is most often liberated by iPLA2 is arachidonic acid (165), an eicosanoid precursor which can facilitate PLD activation leading to the activation of subsequent signaling cascades (166). Moreover, excessive iPLA2 mediated arachidonic acid release can lead to 4-hydroxy-2-nonenal (4-HNE) production (167). 4-HNE is a peroxidized product of arachidonic acid which can have a significant impact on cell functions by forming protein adducts and neuroaxonal dystrophy (168, 169). Additionally, a subset of individuals with mutations in the PLA2G6 gene develop brain iron deposits and abnormal EMG readings likely resulting from axonal swelling and deterioration (170, 171). The mechanism for brain iron accumulation is unclear but it is likely due to many factors from the dysfunction of proteins involved with iron transport and storage (172) and the ability for iron to participate in the Fenton reaction to generate free radicals from products of mitochondrial respiration (173).

CHAPTER 2: HYPOTHESES, RATIONALE AND OBJECTIVES

The overall goal of this thesis was to shed light on PC and PE metabolism by utilizing unique genetic models linked to obesity and neurodegenerative disease. Dietary choline intake is critical for regulating metabolism and this thesis examines the role of choline in skeletal muscle lipid metabolism as this has not been extensively investigated. As choline is important for regulating metabolism, it was necessary to understand how choline is transported into the cell to be utilized. SLC44A1 deficient cells were utilized to further our understanding of cellular choline uptake. Additionally, choline and ethanolamine are similar in structure and are both used as substrates for de novo phospholipid biogenesis. However, the mechanism of ethanolamine uptake in humans has not been elucidated which prompted us to address this question.

It was hypothesized that:

I: Choline supplementation in insulin resistant Pcyt2+/- mice improves muscle function

The development of insulin resistance, metabolic syndrome and type 2 diabetes is strongly correlated with increased DAG and TAG accumulation in skeletal muscle.

Choline deficiency compromises muscle cell function by decreasing mitochondrial abundance, increasing TAG accumulation and perturbing phospholipid and FA metabolism. We postulated that choline supplementation will be important for mitigating

Pcyt2+/- muscle pathologies by restoring FA and energy homeostasis to improve insulin signaling in muscle. The primary objective was to analyze the effects of choline supplementation TAG accumulation and markers of energy metabolism in skeletal muscle of Pcyt2+/- mice exhibiting an obese phenotype.

II: Dysfunctional CTL1/SLC44A1 is implicated in widespread disturbances in membrane composition

We characterized two adolescent patients with a neurodegenerative disease caused by different homozygous frameshift mutations in the SLC44A1 gene. Our group postulated that these mutations have a severe effect on choline uptake, leading to downstream disturbances in phospholipid homeostasis. We analyzed the molecular mechanisms for this hypothesis in fibroblasts isolated from skin biopsies from the two patients. The primary objective was to establish how the SLC44A1 mutations lead to the widespread disturbances in membrane composition.

III: CTL1 functions as an ethanolamine transporter

PE is an important component of cellular membranes, yet little is known about how one of its constituents, ethanolamine, is transported into the cell. Through past experiments, we have seen that choline and ethanolamine seem to compete with each other for uptake into the cell. Moreover, we have seen that blocking the CTL1 transporter with an

antibody leads to diminished PE production. The primary objective was to determine if

CTL1 has ethanolamine transport capacity, and if so to characterize uptake kinetics using a variety of cell lines.

CHAPTER 3 RATIONALE

It is widely known that dietary choline consumption is strongly correlated with optimal health. Studies have shown that a diet deficient in choline is implicated in fatty liver (10) and cognitive impairment (15). However, little is known about choline supplementation on skeletal muscle lipid metabolism within the context of obesity in a mouse model. We utilized our Pcyt2+/- mouse model which has diminished substrate flow through the CDP- ethanolamine branch of the Kennedy pathway. This was an adequate model to utilize because these mice gradually develop insulin resistance and obesity as well as DAG and TAG accumulation in skeletal muscle (40).

CHAPTER 3: ADAPTATIONS TO EXCESS CHOLINE IN INSULIN RESISTANT AND PCYT2 DEFICIENT SKELETAL MUSCLE

3.1 Abstract

It was hypothesized that choline supplementation in insulin resistant (IR)

CTP:phosphoethanolamine cytidylyltransferase deficient (Pcyt2+/-) mice would ameliorate muscle function by reducing TAG accumulation and altering the expression of key players in energy metabolism. Pcyt2+/- mice either received no treatment or were allowed access to 2 mg/ml choline in drinking water for 4 weeks. Skeletal muscle from legs was harvested from choline treated and untreated mice. Lipid analysis and metabolic gene expression and signaling pathways were compared between untreated

Pcyt2+/- mice, treated Pcyt2+/- mice and Pcyt2+/+ mice. The major positive effect of choline supplementation in muscle was the reduction of TAG accumulation and increased muscle glycogen storage. Choline reduced the expression of genes devoted to FA and TAG formation (Scd1, Fas, Srebp1c, Dgat1/2), upregulated the expression of genes devoted to FA oxidation (Cpt1, Pparα, Pgc1α) and had minor effects on genes devoted to phospholipid synthesis and lipolysis. Pcyt2+/- muscle had reduced protein content of key players involved with insulin signaling (IRS1), autophagy (LC3) and choline transport (CTL1) proteins that were restored by choline treatment. Additionally,

AMPK and Akt phosphorylation were increased while mTORC1 phosphorylation was decreased. These data suggest that choline supplementation can play a role in improving muscle glucose metabolism by reducing lipogenesis and improving

mitochondrial and intracellular signaling for protein and energy metabolism in Pcyt2 deficient mice.

3.2 Introduction

The nutrient choline is required for the formation of membrane phospholipids PC and sphingomyelin, the neurotransmitter acetylcholine (ACh) (174) and the methyl group donor betaine (175). Choline is a vital component of skeletal muscle function and choline deficiency is directly implicated in the development of muscular dystrophy (176).

Choline specific transporter CTL1/SLC44A1 is abundantly present in skeletal muscle

(177) and upregulated in numerous mitochondrial and muscular myopathies (178).

Inherited mutations in choline kinase beta result in the development of muscular dystrophy in both mouse and humans (176, 177). ACh is essential for muscle contraction (178) and both neuron-specific (179) and ubiquitous choline transporters

(180) supply choline for muscle ACh synthesis (181). Choline deficiency compromises muscle cell function by decreasing mitochondrial abundance, increasing TAG accumulation and perturbing phospholipid and FA metabolism (182).

The ATTICA study (183) as well as the Nurses’ Health Study (184) demonstrated that diets rich in choline and betaine could reduce inflammatory biomarkers, including C- reactive protein, interleukin-6 and tumor necrosis factor alpha. Adequate intakes of choline and betaine have been frequently associated with lower plasma Hcy, an independent risk factor for cardiovascular disease. In both animal models and humans,

choline deficiency results in the formation of non-alcoholic fatty liver and an increased sensitivity to inflammation (185). Choline and betaine are lipotropic dietary agents as they can reduce the accumulation of TAG in the liver. Betaine has been shown to curb liver fat accumulation and it has been broadly recognized as a safe and effective therapy for alcoholic fatty liver disease (186). The beneficial role of choline and betaine was also clearly observed with respect to non-alcoholic steatohepatitis (which develops in obesity) when oral supplements improved homocysteine levels, lipid profile

(attenuation of fatty liver and proper export and formation of triglycerides and VLDL), steatosis, inflammation and fibrosis (71). Therefore, it is clear that a diet enriched in choline is beneficial in the prevention and treatment of obesity related disorders.

However, it is not known if dietary choline could regulate muscle function and mitigate lipotoxicity.

It has becoming increasingly evident that skeletal muscle phospholipids are important mediators of insulin sensitivity and muscle contraction. Muscle-specific knockout models for CTP:phosphoethanolamine cytidylyltransferase/Pcyt2 (187) and choline/ethanolamine phosphotransferase 1/CEPT1 (188) have been recently developed. These enzymes catalyze the rate-limiting step and the final step respectively in the CDP-ethanolamine (Kennedy) pathway for PE biosynthesis. The two knockout models established that impaired PE biosynthesis has a direct impact on muscle lipid metabolism and function (187, 188). The muscle-specific knockout (187) accumulated

TAG and DAG but did not develop IR, probably because of upregulated mitochondrial

FA oxidation. On the other hand, CEPT1-deleted muscle (188) was protected from high-

fat diet induced IR but was exercise intolerant, which has been attributed to PE dependent regulation of endoplasmic reticulum calcium handling. We have generated a heterozygous Pcyt2 mouse (Pcyt2+/-) which has systemically reduced PE biosynthesis

(189). Young Pcyt2+/- mice are asymptomatic but begin to become more insulin resistant and gain weight more readily than their wildtype counterparts after 20 weeks of age.

Moreover, TAG and DAG accumulate in the skeletal muscle of these older Pcyt2+/- mice

(40, 190). Choline treatment reduced Pcyt2+/- mediated weight gain and TAG accumulation in the liver (191). Here, we hypothesize that choline will also aid in mitigating Pcyt2+/- muscle pathologies by reducing TAG accumulation, which is likely beneficial in improving the expression of key players in insulin signaling and mitochondrial function in muscle.

3.3 Methods

3.3.1 Animals and choline treatments

Generation of Pcyt2 deficient (Pcyt2+/-) mice and genotyping were described previously

(189). All procedures were approved by the University of Guelph's Animal Care

Committee and were in accordance with guidelines of the Canadian Council on Animal

Care. Mice were exposed to a 12-h light/12-h dark cycle beginning with light at 7:00 a.m. Mice had ad libitum access to a standardized chow diet (Harlan Teklad S-2335,

50% calories from carbohydrate, 30% calories from fat, 20% calories from protein) and water. Pcyt2+/- mice progressively develop gain weight and develop insulin resistance

starting at 20 weeks of age (190). In the present study, 8-10-month old mice were divided into three groups (n = 6 - 10 in each group): Pcyt2+/- treated (Pcyt2+/- CHO), and untreated (Pcyt2+/-) with choline, and wildtype littermate controls (Pcyt2+/+). Treated mice were kept in separate cages and had free access to water containing 2 mg/ml choline (Sigma) for 4 weeks. Choline supplementation did not influence food and water intake, as determined on a daily basis. The choline intake from drinking water was ~240

μg/g/day as determined from the choline concentration (2 mg/ml), water intake (5 ml/day/mouse), and mouse weight (191). Thus, in addition to 17 mg/day obtained from diet, the Pcyt2+/- CHO group obtained 10 mg supplemental choline/day (or 37% of total choline ingested) in water. Mice were sacrificed by CO2 asphyxiation and total leg skeletal muscle was harvested for future analyses.

3.3.2 Analysis of phospholipids and cholesterol

The analysis of muscle phospholipids and cholesterol was performed by HPLC (Agilent

Technologies, Santa Clara, CA, USA) as previously described (40). Briefly, 1 mg of skeletal muscle homogenates from Pcyt2+/- CHO, Pcyt2+/- and Pcyt2+/+ mice (n = 4) was extracted in the presence of an internal standard (50 mg dipalmitoyl-phosphatidyl dimethyl ethanolamine) using the method of Folch et al. (192). The extracted lipids were dried, resuspended in chloroform:isooctane (1:1) and separated by HPLC using a 3- solvent gradient and a normal phase column (Onyxmonolithic silica; Phenomenex

Incorporated, Torrance, CA, USA). The amounts of PC, PE, phosphatidylinositol (PI),

sphingomyelin (SM), cholesterol (FC), cholesterol ester (CE) and free fatty acids (FFA) were determined using appropriate standards and expressed in mg of lipid/mg of protein.

3.3.3 Analysis of DAG and TAG

Muscle samples (50 mg, n = 4 per group) were homogenized in 200 μl 5% PBS-T, the homogenates were twice heated to 100°C, cooled to room temperature and centrifuged to remove protein debris. Total TAG content (μg TAG/mg of muscle) was determined using the Wako Diagnostics Kit (Mountain View, CA, USA). In addition, 50 mg muscle samples (n = 3 per group) were extracted by the method of Bligh and Dyer (193). The lower organic phase was dried and resuspended in 100 μl of chloroform. DAG and TAG were resolved by TLC using a solvent system of heptane/isopropyl ether/acetic acid

(60:40:3). Authentic standards were spotted in parallel to the samples, the plates were dried and exposed to iodine vapour for visualization. DAG and TAG spots were scanned and quantified with densitometry (190).

3.3.4 Analysis of muscle glycogen

Skeletal muscle samples (50 mg; n = 4 per group) were hydrolyzed in 500 μl 30% KOH.

Homogenates were heated to 100°C for 30 min with frequent vortexing. Glycogen was precipitated with 600 μl of 95% ethanol, kept on ice for 45 min and centrifuged at 840 x

g for 30 min. The glycogen pellet was dissolved in 1 ml of water by vortexing. The glycogen solution was mixed with 1 ml of 5% phenol and 5 ml of 98% sulfuric acid was then added and kept at room temperature for 10 min. Samples were cooled on ice for

30 min and developed colour which was determined spectrophotometrically at 490 nm.

A standard curve was generated using pure glycogen (Roche) (194).

3.3.5 Immunoblotting

Muscle samples (50 mg; n = 3 per group) were homogenized in cold lysis buffer (10 mM

Tris-HCl (pH 7.4), 1 mM EDTA and 10 mM NaF) containing protease (1/10) and phosphatase (1/100) inhibitor cocktails (Sigma). The lysates were centrifuged for 20 minutes at 10,000 x g at 4oC to remove cell debris and 50 µg of lysate was used for analysis. For CTL1 protein detection, the lysates were mixed with a non-denaturing loading buffer (62 mM Tris-HCl and 0.01% bromophenol blue in 10% glycerol) and resolved with an 8% native gel at 90 V for 2 h. All other proteins were analyzed under denaturing conditions with a loading buffer comprised of 400 mM Tris-HCl, 50% glycerol, 10% SDS, 0.25% (w/v) bromophenol blue and 3% (w/v) DTT. AMPKα, pAMPKα, ACC, pACC, PKCα, PI3K, IRS-1, Akt, p308Akt, p473Akt, mTORC1 and p2443mTORC1 were resolved with a 10% denaturing gel. LC3 I and LC3 II were resolved with a 15% denaturing gel. All proteins were transferred to PVDF membranes (VWR).

Membranes were blocked with 5% skim milk in PBS-T and incubated with the C- terminal CTL1 antibody (ENS-627; 1:500). Membranes were also incubated with

AMPKα, pAMPKα, PI3K, Akt, p308Akt, p473Akt, mTORC1 and p2443mTORC1 (Cell

Signaling), LC3 I/II (Abcam), IRS-1 (Millipore) antibodies as a 1:1,000 dilution with 5%

BSA in PBS-T overnight at 4oC. Membranes were washed 2 times with PBS-T and incubated with horseradish peroxidase-conjugated anti-rabbit IgG secondary antibody

(NEB) (1:5,000) in 5% skim milk in PBS-T for 1 h. After washing with PBS-T, the proteins were visualized using a chemiluminescent substrate (Sigma). β-tubulin (Sigma) was used as a loading control.

3.3.6 Analysis of gene expression mRNA was extracted from 50 mg of Pcyt2+/- CHO, Pcyt2+/- and Pcyt2+/+ skeletal muscle

(n = 5-8 per group) using TRIzol reagent (Invitrogen). cDNA was prepared from 2 µg of total mRNA using a 3-AP primer and SuperScript II reverse transcriptase (Invitrogen).

For the phospholipid biogenesis pathways, the expression of the choline transporter like protein 1 (Ctl1), CTP:phosphocholine cytidylyltransferase (Pcyt1), phosphatidylserine synthase 1 and 2 (Pss1 and Pss2) and phosphatidylserine decarboxylase (Psd) was quantified. The expression of genes involved in FA and TAG synthesis including sterol regulatory binding protein 1c (Srebp1c), FA synthase (Fas), stearoyl-CoA desaturase 1

(Scd1), and diacylglycerol acyltransferase 1 and 2 (Dgat1 and Dgat2) was analyzed.

The expression of genes devoted to lipid degradation such as lipoprotein lipase (Lpl), hormone-sensitive lipase (Hsl) and adipocyte triglyceride lipase (Atgl) was also analyzed. The expression of the mitochondrial regulatory genes peroxisomal

proliferation activation receptor α (Pparα), Ppar gamma coactivator 1α (Pgc1α), and the

FA genes acyl CoA carboxylase (Acc) and carnitine palmitoyltransferase 1 (Cpt1) was analyzed. Glyceraldehyde-3-phosphate dehydrogenase (Gapdh) mRNA was used as a control. The primers and PCR conditions are in Table 3.1 and as described (40, 189,

191).

Table 3.1: Primer sequences and melting temperatures (TM) for polymerase chain reaction (PCR) using mouse RNA

O Gene Forward Primer 5’-3’ Reverse Primer 5’-3’ TM ( C) Size (bp)

Gapdh ACCACAGTCCATGCCATCAC TCCACCACCCTGTTGCTGTA 57 452

Pss2 GAGTGGCTGTCCCTGAAGAC TCGTAGATCTCACGCATGGC 59 305

Atgl CAACGCCACTCACATCTACGG GGACACCTCAATAATGTTGGCAC 57 106

Hsl ACGCTACACAAAGGCTGCTT TCGTTGCGTTTGTAGTGCTC 59 125

Lpl GCTCGCACGAGCGCTCCATT CCTCGGGCAGGGTGAAGGGAA 59 351

Pgc1α TTGACTGGCGTCATTCGGG GAAGGACTGGCCTCGTTGTC 59.5 396

Fas AGATGGAAGGCTGGGCTCTA GAAGCGTCTCGGGATCTCTG 59 268

Ctl1 GAACGCTCTGCGAGTGGCTGC CGGCTTTAGCTCTCGGGCGT 49 376

Pcyt1 ATGCACAGAGAGTTCAGCTAAAG GGGCTTACTAAAGTCAACTTCAA 50 170

Pss1 CTGTTGTGCAATGGTGGTGG GGCTGGCTTGGAACACAAAG 59.5 283

Psd GTTTGCTGTCACGTGCCTGTG CAGTGCAAGCCACATACGGG 59.5 205

Dgat2 GGCGCTACTTCCGAGACTAC TCTTTAGGGTGACTGCGTTC 58.5 255

Ssd1 CGCATCTCTATGGATATCGCCCC CTCAGCTACTCTTGTGACTCCCG 57 279

Dgat1 ATCCAGACAACCTGACCTACCG GACCGCCAGCTTTAAGAGACGC 57 257

Cpt1 CCCATGTGCTCCTACCAGAT GGTCTCATCGTCAGGGTTGT 57 396

Pparα CGCATGTGAAGGCTGTAAGGGC GGCTTCGTGGATTCTCTTGCCC 57 289

Srebp1c TCACAGGTCCAGCAGGTCCC GGTACTGTGGCCAAGATGGTCC 57 438

Acc CGCATCTCTATGGATATCGCCCC CTCAGCTACTCTTGTGACTCCCG 57 295

Scd1 CGCATCTCTATGGATATCGCCCC CTCAGCTACTCTTGTGACTCCCG 57 279

3.3.7 Statistical analysis

Data are represented as mean ± SEM. One-way ANOVA with post-hoc Tukey’s test was performed to compare means between WT, Pcyt2+/- and Pcyt2+/- CHO groups. A difference at 95% confidence interval (p ≤ 0.05) was considered as significant. All statistical tests were performed with GraphPad Prism 4 software.

3.4 Results

3.4.1 Choline modifies membrane composition and reduces TAG accumulation in

Pcyt2+/- muscle

The analysis of the membrane lipids in choline treated and control muscle was shown in

Figure 3.1. The content of the major phospholipids PE and PC in the Pcyt2+/- muscle was similar to the content in the Pcyt2+/+ muscle. The Pcyt2+/- muscle however had reduced levels of phosphatidylinositol (PI) and the ‘lipid raft’ free cholesterol (FC), which are important molecules for signal transduction. Choline treatments did not affect PI content but significantly increased PC, SM and FC to levels similar to control muscle.

Additionally, free FA content was dramatically reduced (60%) by choline treatment

(Figure 3.1). Overall, choline treatment increased the membrane phospholipid ratio

(PC:PE, Figure 3.1) but did not alter the ‘lipid rafts’ ratio (SM:FC, Figure 3.1).

The levels of Pcyt2+/- muscle TAG were elevated relative to control muscle but were reduced by choline treatment. Pcyt2+/- muscle had 60% (Figure 3.2) more TAG than the

control muscle and choline reduced muscle TAG by 40%. A similar trend was observed in TAG and DAG relative ratios (Figure 3.2); Pcyt2+/- muscle DAG was elevated by 30% relative to control Pcyt2+/+ muscle and choline (Pcyt2+/- CHO) returned the elevated

Pcyt2+/- DAG to the control levels. Taken together, the lipid analysis demonstrated that a choline enriched diet can help increase choline containing membrane phospholipids

(PC and SM) while reducing DAG and TAG content in Pcyt2+/- muscle.

While not presented in this particular study, the Pcyt2+/- mice exhibited a reduction in whole body weight gain, adipose weight and adipose cell size and liver weight with choline treatment. Moreover, liver TAG and plasma TAG content were diminished in

Pcyt2+/- mice supplemented with choline. Choline treatment also had a beneficial impact on PC remodeling and plasma acylcarnitine levels in Pcyt2+/- mice (191 Laila liver paper).

Figure 3.1: Choline modifies muscle glycerolipid content. (A) Choline supplementation modified choline-containing phospholipid and cholesterol content. (B) The relative phospholipid ratio of PC:PE increased with choline supplementation, and (C) the relative SM:FC ratio was not affected with choline treatment. *p < 0.05 compared to Pcyt2+/+ and #p < 0.05 compared to Pcyt2+/-.

Figure 3.2: Choline modifies muscle TAG and DAG content and glycogen storage. (A, B) Choline reduced muscle TAG and DAG. (C) Choline increased muscle glycogen content. *p < 0.05 compared to

Pcyt2+/+ and #p < 0.05 compared to Pcyt2+/-.

3.4.2 Choline helps restore Pcyt2+/- muscle glycogen

Choline treatment helped increase Pcyt2+/- muscle glycogen levels (Figure 3.2).

Glycogen was reduced in Pcyt2+/- muscle relative to Pcyt2+/+ wildtype muscle by 50%.

One month of dietary choline supplementation was able to increase Pcyt2+/- muscle glycogen levels by 60% (p < 0.05). Taken together, the data in Figure 3.2 demonstrated that choline can influence the remodeling of Pcyt2+/- muscle TAG and glycogen content to be more similar to the lean phenotype of Pcyt2+/+ muscle.

3.4.3 Choline does not modify phospholipid gene expression

Choline is incorporated into PC via the CDP-choline (Kennedy) pathway. The mRNA expression of the choline transporter (Ctl1/Slc44a1) and the main regulatory gene for

PC synthesis (Pcyt1) did not differ among the three groups. The expression of Pss1, which catalyzes the degradation of PC and the formation of PS from PC, was decreased by 15% (p < 0.05) with choline treatment. A 20% reduction was also observed with the expression of Pss2, which catalyzes the formation of PS from PE (p <

0.05). The expression of Psd, which catalyzes the formation of PE from PS in mitochondria, was also not altered with choline treatments and it was similar across the three groups (Figure 3.3). Taken together, these data demonstrated that dietary choline did not significantly modify the expression of genes devoted to phospholipid synthesis in muscle.

3.4.4 Choline downregulates Pcyt2+/- muscle lipogenesis

Since choline treatment reduced FA, DAG and TAG levels in muscle of obese mice

(Figure 3.3), we then investigated the effect of choline on the expression of genes involved in FA and TAG synthesis (Figure 3.3). The expression of major regulators of lipogenesis, Srebp1c, and its target genes Fas and Scd1 was elevated in Pcyt2+/- relative to Pcyt2+/+ control muscle and they were reduced by choline treatment (25%,

15% and 15% respectively, p < 0.05), to become similar to the expression observed in the wildtype muscle. The expression of Dgat1 and Dgat2, which catalyze the formation of TAG from FA and DAG, were elevated in Pcyt2+/- relative to Pcyt2+/+ muscle and both decreased with choline treatment (20% and 35% respectively, p < 0.05). Reduced muscle lipolysis is linked to IR and obesity (195). Choline supplementation modestly increased the expression of Lpl (10%, p < 0.05). Hsl and adipose triglyceride lipase

(Atgl) were only slightly different from wildtype muscle and unaffected by choline treatments. Taken together, the gene expression data and the metabolic analysis in

Figures 3.1 and 3.2 implicate that in Pcyt2+/- muscle, choline supplementation primarily reduced the mRNA expression of key players in de novo FA synthesis and lipogenesis while having a minor effect on the key players that facilitate DAG/TAG degradation by lipolysis.

Figure 3.3: Choline positively affects the expression of genes involved in lipogenesis and lipolysis. (A) mRNA expression of genes devoted to PC synthesis and PC breakdown. (B) mRNA expression of key players in lipogenesis. (C) mRNA expression of key players in lipolysis. *p < 0.05 compared to Pcyt2+/+ and #p < 0.05 compared to Pcyt2+/-.

3.4.5 Choline improves AMPK activity and FA oxidation in Pcyt2+/- muscle

As shown in Figure 3.4, AMPK activity (pAMPK:AMPK) was dramatically reduced in

Pcyt2+/- muscle relative to the wildtype muscle (3-fold, p < 0.05). AMPK activity was significantly increased (2-fold) by choline treatment, indicating improvements in protein signaling with respect to mitochondrial energy production in Pcyt2+/- mice. Additionally,

ACC phosphorylation (pACC:ACC), which was diminished in Pcyt2+/- mice (2-fold, p <

0.05), was increased with choline supplementation (2-fold, p < 0.05), indicating that choline helped inhibit the first committed step in de novo FA synthesis (Figure 3.4). In addition to decreasing the expression of key players in FA synthesis and increasing pAMPK activity, there was an increase in the expression of genes devoted to FA oxidation with choline treatment. The expression of genes involved in FA oxidation -

Pparα, Cpt1 and Pgc1α (Figure 3.4) were reduced 20-35% in Pcyt2+/- muscle relative to

Pcyt2+/+ control muscle. Choline (Pcyt2+/- CHO) increased their expression by 20-25%, back to the levels observed in the control muscle. Acc mRNA expression was similar between the groups, however the inhibition of ACC activity through increased phosphorylation (AMPK activation) was described in Figure 3.4.

Figure 3.4: Choline has a positive effect on the expression of key players in fatty acid oxidation.

(A, B) Increased AMPK phosphorylation and (B) ACC phosphorylation with choline treatment. (C)

Representative immunoblots of AMPK and ACC. (D) mRNA expression of key players in fatty acid oxidation and mitochondrial biogenesis. *p < 0.05 compared to Pcyt2+/+ and #p < 0.05 compared to

Pcyt2+/-.

3.4.6 Choline improves nutrient signaling in Pcyt2+/- muscle

To support alterations in gene expression with regards to FA formation (Figure 3.3) and mitochondrial oxidation (Figure 3.4), we then assessed the effect of choline on key components of insulin signaling. As shown in Figure 3.5, the most dramatic effect of

choline was the restoration of IRS1 protein content and Akt phosphorylation (Figure

3.5). The amount of IRS1 in Pcyt2+/- muscle was diminished by 80% and choline recovered IRS1 protein to the control levels (Figure 3.5). Downstream from IRS1, the most affected was the phosphorylation of Akt at Ser473 (p473Akt:Akt). Pcyt2+/- p473Akt was reduced 50% and choline increased p473Akt levels 3-fold (Figure 3.5E).

Interestingly, Akt phosphorylation at Thr308 (p308Akt:Akt, Figure 3.5D) was not modified by Pcyt2 deficiency or affected by choline treatment. Finally, choline dramatically reduced (50%) PKCα content (Figure 3.5), a well-established inhibitor of muscle insulin signaling (196).

As choline improved Pcyt2+/- muscle AMPK activity and insulin signaling, we also probed the regulation of mTORC1 (protein synthesis) and LC3 (autophagy), downstream of both AMPK and insulin signaling (Figure 3.6). Indeed, there was a 60% increase in mTORC1 phosphorylation at Ser2448 in Pcyt2+/- muscle with respect to wildtype muscle, and choline largely mitigated this increase in phosphorylation

(p2448mTORC1:mTORC1 in Figure 3.6). Since mTORC1 is a negative regulator of autophagy, we also probed the level of lipidation of the autophagy marker LC3 (LC3

II:LC3 I in Figure 3.6). The lipidation of LC3 was reduced in Pcyt2+/- muscle and choline increased LC3 lipidation to the level present in the control muscle. As there are alterations in autophagy occurring with choline treatment, it was also of interest to determine the protein content of the choline transporter CTL1. It has previously shown to be degraded by the autophagosomal-lysosomal pathway in palmitate treated muscle cells (198). CTL1 protein content as well as the total LC3 protein content were however

dramatically reduced (by 70%) in Pcyt2+/- muscle and almost completely recovered by choline treatment, implying elevated proteosomal degradation in Pcyt2+/- muscle (Figure

3.6). Taken together, the above data implicate a beneficial role for choline in regulating insulin signaling as well as protein synthesis and turnover in Pcyt2+/- muscle.

Figure 3.5: Choline reduces PKCα and restores muscle insulin signaling. (A - F) Representative immunoblots showing differences in expression of key players in insulin signaling. Each experiment was repeated 3 times with n = 3 per group; one-way ANOVA: *p < 0.05 compared to Pcyt2+/+ and #p < 0.05 compared to Pcyt2+/-.

Figure 3.6: Choline reduces mTORC1 activation and increases expression of key players in autophagy. (A) Representative immunoblots for mTORC1, p2448mTORC1, LC3 I, LC3 II, CTL1, and β tubulin in choline treated and untreated groups. (B) Choline treatment reduces muscle mTORC1 protein phosphorylation. (C) Choline increases expression of key players in autophagy. (D) Choline restores

CTL1 protein content. Each experiment was repeated 3 times (n = 3 per group); one-way ANOVA: *p <

0.05 compared to Pcyt2+/+ and #p < 0.05 compared to Pcyt2+/-.

3.5 Discussion

The effects of dietary choline on Pcyt2+/- muscle were assessed in this study. Choline decreased muscle TAG accumulation largely by reducing the expression of FA and

TAG biosynthetic genes while increasing the expression of genes devoted to energy utilization. Choline helped facilitate increased muscle glycogen levels and restored the activity of the metabolic regulators AMPK, Akt and mTORC1. Overall, choline improved glucose and lipid balance and insulin signaling in Pcyt2+/- muscle.

One month of choline supplementation in Pcyt2+/- mice increased the content of choline- containing phospholipids (PC and SM) and cholesterol (FC). This increased the membrane phospholipid ratio but did not modify the lipid raft ratio in skeletal muscle.

The major effect of choline supplementation was the reduction in FA, DAG and TAG content that were otherwise elevated in the Pcyt2+/- muscle. Based on previous studies

(197), one might have anticipated that choline facilitated TAG degradation by lipolysis but this was not the case. Choline mediated effects were primarily by reducing the expression of genes devoted to FA synthesis via lipogenesis and FA esterification/TAG formation regulated by Srebp1c and Dgat1/2 respectively. The expression of Srebp1 and its target genes Fas and Scd1 as well as Dgat1/2 were significantly reduced while the lipolytic genes Lpl and Atgl were not affected by choline treatment. Srebp1 was positively regulated by insulin signaling and downstream mTORC1 but it is not known how it becomes upregulated in IR states to cause elevated lipogenesis and an accumulation of TAG in the liver and perhaps other tissues. On the other hand, Liu et al.

(2012) showed that AMPK mediated mTORC1 inhibition decreases the ability for

SREBP1c to localize to the nucleus in IR skeletal muscle (198). Here we established a relationship with impaired insulin signaling whereby mTORC1 was upregulated and

AMPK was down regulated in Pcyt2+/- muscle. Choline supplementation improved insulin signaling, upregulated AMPK activation and downregulated mTORC1 activation, consequently reducing the expression of Srebp1c and lipogenesis.

We previously established through metabolomic studies that the majority of choline supplied to Pcyt2+/- mice becomes oxidized to betaine, which enters the one carbon cycle and stimulates mitochondrial oxidation. The increased flux through the one-carbon cycle prevents the accumulation of choline and betaine in the circulation (40). Plasma acyl-carnitine composition after choline supplementation to Pcyt2+/- mice further supports this possibility (40). Therefore, the main positive effect of choline on Pcyt2+/- muscle can likely be attributed to its action as a methyl group donor, and a potential stimulation of mitochondrial FA oxidation. High dietary choline and betaine intakes are significantly associated with favorable body composition in humans (199, 200, 201), demonstrating that choline supplementation lowers oxidative stress during exercise. In young adult male athletes, 600 mg per day of supplemental alpha glycerylphosphorylcholine has recently been demonstrated to improve strength (202). In addition, betaine has been used as an ergogenic agent to improve muscle performance and to increase lean body mass in humans (203), possibly by increasing circulating IGF levels leading to increased PI3K/Akt signaling (204). More recently, Ejaz et al. (2016) has demonstrated in mice that betaine supplementation reduces obesity and systemic

IR by upregulation of Fgf21 (205).

In addition to the effects on muscle FA oxidation through the betaine/one carbon cycle, choline could have influenced acetylcholine signaling, which is essential for muscle contraction. Reduction in acetylcholine production has a marked influence on muscle performance and most sport nutrition products are supplemented with choline to maintain acetylcholine levels (192). In mice, the mechanism of muscle contraction after acetylcholine stimulation primarily depends on AMPK activation and is independent of insulin (206). IR is associated with muscle weakness and exercise ameliorates IR via

AMPK activation and Ca2+ signaling (207, 208). Therefore, the activation of AMPK (208) by choline supplementation likely improved the metabolic condition of Pcyt2+/- muscle.

The question remains if muscle Ca2+ signaling is also activated by choline supplementation and will require further investigation.

The general role of AMPK is to stimulate FA oxidation and to diminish TAG synthesis by lipogenesis, and these processes are likely both in effect in choline supplemented

Pcyt2+/- muscle. Deactivation of ACC with choline treatment can be beneficial because

ACC catalyzes the formation of malonyl-CoA, the rate limiting step in FA biosynthesis.

Additionally, choline increased the mRNA expression of the mitochondrial regulators

Pparα, Pgc1α and Cpt1. Pparα stimulates FA oxidation and Pgc1α regulates mitochondrial biogenesis, while Cpt1 is the rate limiting step in FA transport in the mitochondria.

Choline supplemented Pcyt2+/- muscle demonstrated improvements in insulin signaling.

There was an increase in IRS1, an increase in p473Akt (mTORC2 site) and increased muscle glycogen content with choline treatment. Glycogen synthesis and content are

typically decreased in IR. Liu et al. (2015) have shown that this can occur because elevated GSK3 phosphorylation leads to the inhibition of glycogen synthase activity

(209). Furthermore, Stretton et al. (2015) have shown that GSK phosphorylates Raptor, an mTORC1 associated scaffold protein, which is important in mTORC1 activation

(210). mTORC1 was hyperactivated in Pcyt2+/- muscle and there was a decrease in activation with choline treatment. mTORC1 is a regulator of protein synthesis (211), and the activation of this protein was normalized with choline treatment. AMPK activation has been shown by Hong-Brown et al. (2010) to inhibit the activation of mTORC1 (212) and is likely most responsible for the normalization of protein turnover in choline supplemented Pcyt2+/- muscle. Why might mTORC1 have been more active in Pcyt2+/- muscle? Um et al. (2004) demonstrated that chronic mTORC1 activation can promote insulin resistance via S6K-dependent phosphorylation of IRS1 leading to its degradation and the inhibition of the insulin signaling pathway (213). Typically, mTORC1 is activated via Akt phosphorylation but this is not the case in IR muscle. mTORC1 hyperactivation was likely a consequence of increased nutrient availability/elevated proteolysis in

Pcyt2+/- muscle, and choline supplementation helped to restore this balance (214).

Choline supplementation has shown a capacity to affect muscle metabolism and improve insulin signaling. The mechanisms that link choline with improvements in muscle insulin signaling and energy metabolism involve multiple parameters. These include the normalization of gene expression to favour less FA formation and more FA oxidation. In turn, this can be linked with a depression of lipogenesis, increases in

muscle glycogen and activation of AMPK. These data suggest that a choline enriched diet can help normalize muscle FA and TAG homeostasis.

Mouse husbandry and study design: L.C. Schenkel, M. Bakovic, experimental work: A.

Taylor, M. Yokich, manuscript preparation: A. Taylor, M. Bakovic. Thanks to Audric

Moses for assistance with phospholipid and cholesterol analysis.

CHAPTER 4 RATIONALE

With additional knowledge regarding how choline modulates metabolism from Chapter

3, it also became important to shed light on how choline is transported into the cell to be utilized. In Chapter 4, we wanted to analyze how mutations within the coding region of

SLC44A1 in human skin fibroblasts affect phospholipid and fatty acid homeostasis. In addition, the individuals with mutations in SLC44A1 exhibit a wide array of motor and cognitive difficulties with brain iron accumulation, which potentially implicates SLC44A1 in human disease.

CHAPTER 4: THE CHOLINE TRANSPORTER LIKE PROTEIN 1 IS INTEGRAL FOR MAINTAINING PHOSPHOLIPID AND FATTY ACID HOMEOSTASIS

4.1 Abstract

Cerebral choline metabolism is crucial for normal brain function, and its homoeostasis depends on carrier mediated transport. Here, we report on two adolescent individuals with neurodegenerative disease and homozygous mutations, Asp517Metfs*19 and

Ser126Metfs*8 respectively, in the SLC44A1 gene encoding the choline transporter-like protein 1, CTL1. The progressive symptoms were ataxia, tremor, cognitive decline, dysphagia, opticus atrophy, poor speech and dysarthria. Cerebral MRI showed changes compatible with Neurodegeneration with Brain Iron Accumulation, NBIA. Cultured fibroblasts demonstrated ultrastructural changes of plasma membrane, increased number of vacuoles, and abnormal mitochondria. Iron uptake in mutated fibroblasts was increased, and iron accumulated in mutated cells indicating that CTL1 deficiency is indeed a subtype of NBIA. Functional studies of skin fibroblasts showed diminished choline transport yet the membrane PC content remained unchanged. As part of the mechanism to preserve choline and phosphatidylcholine the SLC44A1 mutations cause impairments to very long chain fatty acid and membrane phospholipid homeostasis. In conclusion, we outlined a new type of NBIA highlighted by CTL1 deficiency with autosomal recessive inheritance.

4.2 Introduction

The choline transporter-like protein 1 (CTL1) is a highly conserved Na+-independent, intermediate-affinity transporter of choline (215, 216, 217). It is present as a major 3.5 kb transcript which is broadly expressed in different regions of the brain including the striatum, cerebral cortex, hippocampus, hypothalamus, olfactory bulbs and spinal cord

(218, 219). CTL1 is considered an important mediator of choline transport across both the plasma membrane and the mitochondrial membranes (216). The CTL1 protein is encoded by the SLC44A1 gene (215, 216, 217); no association with human disease has been reported so far.

Choline is vital for protecting the brain from neuropathological changes associated with

Alzheimer's disease as well as neurological damage associated with epilepsy, fetal alcohol syndrome and inherited conditions such as Down and Rett syndromes (220).

Choline is a precursor of membrane phospholipid PC (221) and the neurotransmitter acetylcholine (222). The oxidation of choline to betaine is important for the production of the methyl group donor S-adenosylmethionine (223). As choline is a cationic species, its uptake is dependent on carrier mediated transport. Choline transporters are categorized into three transporter families according to their affinity for choline (215, 224): 1) The high affinity choline transporter CHT1, encoded by the SLC5A7 gene, is unique for cholinergic neurons and has been implicated in autosomal dominant distal hereditary motor neuropathy associated with vocal cord paralysis (DHMN7A). This is in addition to autosomal recessive inherited diseases in a spectrum of severe muscle weakness ranging from a lethal antenatal form of arthrogryposis and severe hypotonia to a

neonatal form of congenital myasthenic syndrome with episodic apnea and/or early infantile lethality (225, 226); 2) three different organic cation carriers are known (OCT1-

3, encoded by the SLC22A1-3 genes) recognizing a multitude of exogenic organic cations. OCT1 and OCT2 accept choline as a substrate with a relatively low affinity, while OCT3 does not recognize choline as a substrate (215); 3) the CTL transporters include CTL1-5 (encoded by SLC44A1-5) and have an intermediate affinity for choline.

CTL2 is present in two isoforms of which CTL2-P2 exhibits detectable transport activity of choline in humans (227). CTL2 has been associated with an increased susceptibility to venous thromboembolism (228) and has been linked to progressive sensorineural hearing loss in mice (229). The functions of CTL3-CTL5 at the molecular level are still unclear (224) however, CTL4 has been suggested to be a locus conferring increased susceptibility to age-related macular degeneration (230). So far, CTL1 has not been associated with human disease, but CTL1 has been implicated in the formation and/or maintenance of the myelin sheath. It has also been suggested that malfunction of this transporter could have important implications in nervous system development and repair following injury and in neurodegenerative disease (27).

NBIA is a group of brain iron deposition diseases that lead to mixed extrapyramidal features and progressive dementia. Today, ten genes are thought to be implicated in

NBIA: PANK2, PLA2G6, C19orf12, COASY, FA2H, ATP13A2, WDR45, FTL, CP,

DCAF17, SCP2 and GTPBP2 (231), but a significant proportion of patients remain without genetic diagnosis (232). Some of these genes are directly involved in iron metabolism, while the rest are involved in a variety of other functions including CoA

synthesis, lipid metabolism, autophagy and membrane remodeling (232, 233). Some subgroups of NBIA highlight a link between neurodegeneration and alteration of sphingolipid metabolism, mitochondrial morphology and membrane remodeling (233).

We outlined two adolescent individuals with a neurodegenerative disease caused by homozygous mutations in the SLC44A1 gene. We demonstrated that this disease is a new subtype of NBIA and described the consequences of the genetic defect on a cellular level.

4.3 Methods

4.3.1 Whole exome sequencing and Sanger sequencing

Whole exome sequencing, WES, for individual A-II-2 was performed on data delivered from Dep. Genetics & Pharmacogenomics, Merck Research Labs, Boston. Average sequencing depth was 105x, 94 % of the exome was covered at least 10x, and 92 % was covered at least 20x. Raw reads were processed using the Burrows-Wheeler

Alignment tool (BWA-MEM) v. 0.7.12 (Li, 2013) and the GATK Best Practice pipeline v.

3.3-0 was used for variant calling (234). Variant annotation and filtering were performed using VarSeq (Golden Helix). As autosomal recessive inherited disease was suspected due to parental consanguinity, only rare homozygous variants in the protein coding region were considered (minor allele frequency <1%, ExAc). In silico functional prediction algorithms (Polyphen2, SIFT and MutationTaster) were applied to evaluate the significance of identified missense variants. Sanger sequencing of members of

family A on SLC44A1, exon 13 (RefSeq: NM_080546.4) was analyzed by bidirectional sequencing using the BigDye Terminator v. 3.1 cycle sequencing kit and an ABI3730XL capillary sequencer (Applied Biosystems). Primers used were SLC44A1e13F:

AGGGTGCCATGTGTTTGACT and SLC44A1e13R:

AAAATACTTGGTTTTACCTTGCCA.

WES of individual B-II-1 was performed on an Illumina HiSeq 2500 (Illumina, San

Diego, CA, USA) after enrichment with the SureSelect Human All Exome 50 Mb Kit following standard manufacturer protocol (Agilent, Santa Clara, Ca, USA) and as described previously (235). After calling of variants and calculating coverage parameters, filtering of candidate variants consisted of three steps: (i) common (minor allele frequency > 1%) genetic variants (by public databases (ExAc, ESP, 1000G) and in-house database of more than 1000 exomes) were excluded, (ii) under the assumption of an autosomal recessive phenotype due identical by descent homozygous variants, only these were selected, and (iii) variants were then checked for quality

(manual checking of the aligned reads, and Sanger sequencing where needed), pathogenicity (based on minor allele frequencies in different populations, on in silico predictions and conservation) and causality (gene functions and disease associations).

PCR and Sanger sequencing were used to exclude technical artefacts and validate segregation. Although the parents are related, and the probability of an autosomal recessive phenotype is high, we also have considered that the inheritance may be of different modus since there is only one affected boy in the family. Thus, we have repeated the sequencing of the index and of his parents in the sense of a trio analysis.

Using the available data, we (a) repeated the analysis for homozygous variants (and achieved similar results as in the first experiment), (b) performed analysis for autosomal recessive compound heterozygous variants based on the same three steps of filtering,

(c) performed analysis for X chromosomal recessive inheritance based on the same three steps of filtering and (d) performed analysis regarding a de novo mutation using the same three steps of filtering with modified MAF of a maximum of one heterozygous case in each of the available databases.

4.3.2 Isolation and treatment of skin fibroblasts

Skin biopsies from individual A-II-2 (M1 patient) and individual B-II-1 (M2 patient) were minced into small pieces and grown in minimum essential medium (MEM, Fisher

Scientific), supplemented with 20% fetal bovine serum and 1% penicillin/streptomycin.

o Flasks were kept in a humidified atmosphere at 37 C and 5% CO2. After 3 weeks, fibroblast cells began to grow out from the skin pieces, and once they reached confluency, cells were trypsinized and transferred to a new flask. Cells were kept in culture for 10 days before passage and media was changed every 3 days. An aliquot of cells was kept frozen in liquid nitrogen. Control skin fibroblasts (ie. no mutations in

SLC44A1) from a healthy teenage female were maintained under the same conditions.

For treatments with choline, fibroblasts were grown for 21 days with 2.5 and 5 mg/L choline chloride (Sigma Aldrich, Oakville, ON, Canada) added to the media and

compared with fibroblasts grown without choline. During the 21-day treatment, the cells were passaged every 3-4 days at 3.0 x 105 cells/ml. No differences in cell number or growth rate were observed. After treatment, cells were washed with PBS and used for future analyses.

4.3.3 RNA Isolation and RT-PCR

Total RNA from choline treated M1, M2 and control fibroblasts and untreated M1, M2 and control fibroblasts was isolated with TRIzol (Invitrogen, Life Technologies

Incorporated, Burlington, ON, Canada). DNase I was used to eliminate genomic DNA and cDNA was then synthesized from 2 μg RNA using a 3-AP primer and SuperScript III reverse transcriptase (Invitrogen, Life Technologies Incorporated). Expression of

SLC44A1, SLC44A2, SLC44A3, SLC44A4, SLC44A5, SLC22A1, SLC22A2, SLC22A3,

SLC22A4, SLC22A5, CHT1, Pcyt1, Pcyt2, PLD1, PLD2, CK, EK, PSS1, PSS2, PSD,

PEMT, DGAT1, DGAT2, LPIN1, FAS, SREBP-1c, PGC-1α, PPARα, CPT1, ELOVL1,

ELOVL2, ELOVL3, ELOVL4, ELOVL5, ELOVL6 and ELOVL7 was determined by PCR using the primers and conditions outlined in Table 4.1. Reactions were standardized by amplifying glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and relative band intensity was quantified using ImageJ software (NIH, Bethesda, MD, USA).

Table 4.1: Primer sequences and melting temperatures (TM) for polymerase chain reaction (PCR) using human fibroblast RNA

Gene Forward Primer (5’-3’) Reverse Primer (5’-3’) TM Size (oC) (bp) GAPDH ACCACAGTCCATGCCATCAC TCCACCACCCTGTTGCTGTA 58 452 SLC44A1 5’ probe ACGTAGCTGGCAAGGTCTTC GATCCAAATCAGGCCCACCA 55 221 SLC44A1 3’ probe (M1) TCAACAGCACCAACTTCTGC ACAGGAAGCAATGAGCGACT 55 246 SLC44A1 5’ probe (M2) CTCTTCTGCATTGGGATGGGA CTTGGACACGCTGCTACACA 57 248 PLD1 CGTCCAGTGAGTCTGAGCAA GGCGTGGAGTACCTGTCAA 56 198 PLD2 CATCTGCGGGCTTCGTACAC TCCGCAGACTCAAGGCAAAC 57 248 Pcyt1 CCCTCTTTCCGATGGCCCTT TGTTTGGGTTCCCCCAGTCC 53 267 Pcyt2 ACATCATCGCGGGCTTACAC TGACACACCAGGTCCACCTT 53 190 CK GAGTTCCACATCAGTGTCATCAGA ATGGCCTCAGCCCCTTGAAAT 57 203 EK AATGGTGCCGGGAGTACTTG CCCTCTGGGAAGACTCCGTA 57 274 PSS1 CGTTCACTCGACCTCATCCA GATTGTCCAGCAGAGACCGT 56 302 PSS2 ATGCACCGCACACCCTTTA TCTTGGTGTTGTAGGCCGTG 57 457 PSD TTAGGTGGGGAGTAGCCACA GCTTCCGTAAGGGCTGTAGG 57 273 DGAT1 ACTACCGTGGCATCCTGAAC GCTGGGAAACACAGAATGGT 56 289 DGAT2 AGTGGGTCCTGTCCTTCCTT TCACTTCTGTGGCCTCTGTG 55 321 LPIN1 TAGCCGGCCAGGTGTTTGTC TGTCAACCACTTTCTCTCGGG 55 180 SREBP-1c TTTCCGAGGAACTTTTCGCC AAAGTGCAATCCATGGCTCC 55 343 PGC-1α TGCCCAAGTCAGGACGAAC GCGGCTGTTACTCTCTCTCC 56 495 PPARα TCATCACGGACACGCTTTCA CATCCCGACAGAAAGGCACT 56 299 FAS CAGAGCAGCCATGGAGGAG TCCTCGGAGTGAATCTGGGT 56 316 CPT1 GTTTCTGGCTCGTTGGATGC TGACCACGTTCTTCGTCTGG 56 428 PEMT AGGTAACGAACAGCTCGGTG GGGTCTTGTGTTCCCATCGT 56 201 SLC44A2 CCACGCCTACAAGGGTGTCC CCATGGCAATGACCAGGCCT 58 203 SLC44A3 TTCTTCTGGGTCCTCTGGGT GGTACCTGGACAATGCACCA 56 397 SLC44A4 ATCACTCCGGAGACTGAGCCA CCCTGGCAGACAAAAGTTCCT 58 473 SLC44A5 TTGCAGTGATTCAGGCCTACTT AGGTTTAGCAACACGGAGGG 56 372 SLC22A1 CTGCCCACCTTCCTCTTCC GTGCGGAACAGGTCTGCAA 55 212 SLC22A2 TCTGCCCTGGTTGAATTCCC GAGCCGGTAGACCAGGAATG 55 318 SLC22A3 TTTTTGCTGCTGTGCCTGAC GTTCAGGATGGCTTGGGTGA 56 423 SLC22A4 CTCCCTGTTCTTCGTAGGCG AGTGGCAGCAGCATATAGCC 55 306 SLC22A5 AGGATGGGAAAGAAGCCTGC GAGATCTGGCCCATGCCTAC 58 272 SLC5A7 GCAGACGACTTGACTTGGGA AGATCCTTCCACAGAGGGCT 55 383

4.3.4 Immunoblotting

All fibroblasts were washed 3 times with 1X PBS (Fisher Scientific) and subjected to a lysis buffer (25 mM Tris, 15% glycerol, 1% Triton X-100, 8 mM MgCl2, 1 mM DTT, protease inhibitor cocktail and phosphatase inhibitor cocktail) at 4oC for 30 minutes and scraped into 1.5 ml tubes. Cell debris was removed by centrifugation at 13,500 g for 20 minutes at 4oC. Protein concentration was determined with bicinchoninic acid (BCA;

Pierce, Rockford, IL, USA). The LV58 (N-terminus) and ENS627 (C-terminus) antibodies were previously developed by our laboratory and were shown to detect the

72 kDa SLC44A1 protein under native (non-denaturing) conditions. The LC3 I/II antibody (Abcam) detects proteins at 14 and 16 kDa under denaturing conditions.

Samples were mixed with loading buffer (62 mM Tris-HCl, 0.01% bromophenol blue and

10% glycerol) and separated by PAGE at 100 V for 1.5 h. CTL1 was resolved on an 8% native gel and LC3 I/II were resolved on a 15% denaturing gel. Proteins were transferred onto PVDF membranes (Pall Canada, Mississauga, ON, Canada) by a semi-dry transfer system and stained with Ponceau S. Membranes were blocked in 5% skim milk in Tris Buffered Saline-Tween 20 (TBS-T) solution and then incubated with either one of the SLC44A1 antibodies (1:500 in 5% skim milk in TBS-T) or the LC3 I/II antibody (1:1,000 in 5% skim milk in TBS-T) overnight at 4oC. Membranes were washed with TBS-T and then incubated with an anti-rabbit horseradish peroxidase conjugated secondary antibody (New England Biolabs, 1:5.000 in 5% skim milk in TBS-T) for 2 h.

Membranes were washed in TBS-T and proteins were visualized using a chemiluminescent substrate (Sigma Aldrich, Oakville, ON, Canada).

4.3.5 Choline uptake

Choline uptake was measured according to our standardized protocols (216). Briefly, cells were washed with KRH buffer (130 mM NaCl, 1.3 mM KCl, 2.2 mM CaCl2, 1.2 mM

MgSO4, 1.2 mM KH2PO4, 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid

(HEPES) (pH 7.4) and 10 mM glucose and incubated with 0.5 μCi/ml [3H]-choline

(American Radiolabeled Chemicals, St. Louis, MO, USA) with or without 100 μM hemicholinium-3 (HC-3) for 15 min at room temperature. Cells were then washed in ice cold KRH buffer containing 1 mM ‘cold’ choline to stop the uptake and remove the radiolabeled choline from the cell surface. Cells were then lysed in 500 μl ice cold lysis buffer (10 mM Tris-HCl, 1 mM EDTA and 10 mM NaF), and [3H]-choline was analyzed by liquid scintillation counting. The rate of choline uptake was calculated as DPM/mg at

15 min. Protein was quantified using the BCA protein assay (Thermo Fisher,

Mississauga, ON, Canada).

4.3.6 Electron microscopy

Cultured cells were fixed in 2% glutaraldehyde in 0.04M phosphate buffer, washed in buffer and centrifuged to form pellets. The pellets were further post fixed in osmium tetroxide for 1 hr. The specimens were dehydrated in graded alcohols and propylene oxide and infiltrated with epoxy resin. The epoxy was polymerized at 60°C. Ultrathin sections, 60 μm, were cut and the sections were stained on the grids with uranyl acetate and lead citrate. The sections were analyzed with a JEOL2100 electron microscope.

4.3.7 Iron determination with ferrozine

Total iron content was determined as described (236, 237). Briefly, fibroblasts were cultured 24 h in serum free DMEM in the presence and absence of 100 μM ferric ammonium citrate (FAC) and lysed with 50 mM NaOH for 2 h with rocking. A standard curve was generated using 0-50 μM FAC in 10 mM HCl, 50 mM NaOH, iron releasing agent and iron detecting agent. Iron-releasing agent: 1.4 M HCl:4.5% (w/v) KMnO4 in

H2O. Iron-detecting agent: 6.5 mM ferrozine, 6.5 mM neocuproine, 2.5 M ammonium acetate, 1 M ascorbic acid. Iron-releasing agent was added to lysate and the mixture was incubated for 2 h at 60oC. After cooling to room temperature, 60 μL of the iron- detecting reagent was added to lysate. After 30 minutes, the lysate was transferred into

96 well plate and the absorbance measured at 550 nm. The iron concentration was normalized against the protein content of the sample.

4.3.8 Pulse and pulse-chase radiolabeling with [3H]-choline and [14C]- ethanolamine

For pulse experiments, M1, M2 and control cells were incubated for 1-3 h with 0.5 μCi

[3H]-choline chloride or [14C]-ethanolamine (ARC, St. Louis, MO), washed with PBS and then chased 1-3 h with an excess of unlabeled choline or ethanolamine. At each time point, cells were washed 2X with PBS and total lipids were extracted with the method of

Bligh and Dyer (194). The water soluble CDP-choline metabolites (choline, P-choline

(P-Cho) and CDP-choline) were separated in ethanol:water:ammonia (48:95:7) and

visualized with 0.5% phosphomolybdic acid in ethanol:HCl (19:1). The radiolabeled PC was isolated from the lipid phase after TLC separation with chloroform:methanol:acetic acid:water (40:12:2:0.75) and staining with 15% sulfuric acid and 0.5% K2CrO7. Specific bands were scraped from plates and radioactivity was determined with LSC. The water soluble CDP-ethanolamine metabolites (ethanolamine-Etn), P-ethanolamine (P-Etn) and CDP-Ethanolamine (CDP-Etn) were separated in methanol:0.5% NaCl:ammonia

(50:50:5) and visualized with 0.8% ninhydrin in 95% ethanol. Specific bands were scraped from plates and radioactivity was determined with LSC.

3 14 4.3.9 Equilibrium radiolabeling with [ H]-glycerol and [ C-(CH3)]-methionine

To establish PC, PE and PS equilibrium pools, cells with and without supplemental choline were incubated for 24 h with 0.5 μCi [3H]-glycerol (ARC, St. Louis, MO).

Additionally, to measure the PC pool produced from PE methylation, M1, M2 and control cells with and without supplemental choline were incubated for 24 h with 0.5 μCi

14 [ C-(CH3)]-methionine (ARC, St. Louis, MO). In both scenarios, cells were washed 2X with PBS and total lipids were extracted with the method of Bligh and Dyer (194). An aliquot of cells was used to measure the protein concentration. The radiolabeled PC, PE and PS were isolated from the lipid phase by TLC separation with chloroform:methanol:acetic acid water (40:12:2:0.75) and staining with 15% sulfuric acid and 0.5% K2CrO7. Specific bands were scraped from plates and radioactivity was determined with LSC.

4.3.10 Phospholipase D activity assay

This assay was conducted as previously described (40). In brief, cells were labeled for

24 h with 5 μCi/ml [3H]-myristic acid and then incubated with 0.5% ethanol for 30 mins at 37oC. Cells were solubilized in 0.2% SDS and 5 mM EDTA and protein concentration was determined with the BCA assay. Lipids were extracted with chloroform:methanol:lysate:acetic acid (1:2:0.8:0.08 v/v). The radiolabeled phosphatidic acid (PA) and phosphatidylethanol were separated by TLC using ethyl acetate:chloroform:acetic acid:water (65:15:10:50 v/v) and visualized in iodine vapor.

Bands were scraped from plates and radioactivity was determined with LSC.

4.3.11 Analysis of membrane phospholipids and fatty acids

The lipid analysis was conducted as previously described (40, 189). Prior to experiment, samples were spiked with internal standards 17:0 – 14:1 PS, 12:0 – 13:0 PE and 17:0 –

14:1 PC. Volume was adjusted to 1 ml and lipids were extracted by the method of Bligh and Dyer (194). Organic phases were evaporated under nitrogen gas and reconstituted in a methanol:chloroform mixture. Mass spectrometry was performed by direct infusion into an AB SCIEX QTRAP 5500 (Framingham, MA) in both positive and negative ESI modes. Samples were infused at a constant rate of 5 µl/min and data were acquired by precursor ion and neutral loss scans. Results were based on the peak area and expressed as frequency (%) of each species in the total lipid identified.

Lipidomic clustering of individual phospholipid species was done to partition the data into K=3 groups such that the sum of squares from points to the assigned cluster centers was minimized. Using “clusplot” function, a bivariate cluster plot was made with default method where all the observations were represented by points in the plot, using the principal components. Then, the hierarchical cluster analysis was performed using

“hclust” function where distance was calculated using “Pearson” method and linkage method used was “Complete”. “Heatmap.2” function was used to make the heatmaps.

This analysis was performed for all the three clusters generated by k-means analysis.

4.3.12 Data analysis

All measurements are expressed as means from quadruplets ± SEM. Statistical analysis was performed using GraphPad Prism 4.0 software (GraphPad, Inc.). Data were subjected to students T-test and one way-ANOVA tests. Differences were considered statistically significant at *p < 0.05.

4.4 Results

4.4.1 Clinical description

The family of the M1 patient is Danish but of Turkish origin. The parents are healthy non-consanguineous parents from the same small village in Turkey. The M1 patient is

an 18-year-old girl. She was born at term with birth weight 3522 g (median), birth length

53 cm (+1.5 SD) and head circumference 37 cm (+2 SD). At 2 months, she was hospitalized with liver affection of unknown cause, it resolved spontaneously on prednisolone and nutritional supplements. Language development was delayed, and she did not speak sentences before age 3-4 years. At 7.5 years of age, she was diagnosed with tremor, hypertonia and ataxia, retrospectively gross motor problems and hand tremor was present since 5 years of age. She had a broad forehead and macrocephaly (head circumference 54.5 cm, > 97th percentile). EEG showed an epileptiform pattern, but no seizures was present. At 15 years of age, a PEG-tube was inserted because of increasing problems with swallowing. She had then almost stopped speaking, was increasingly hyperactive, and had problems with sleeping. Melatonin was introduced with some effect. Growth was initially normal with height following -1 SD but decreasing to 152 cm (< -2 SD) at age 15 years of age. Ophthalmologic investigation at

8.5 years of age showed hyperopia +3.5 diopter, visual acuity 0.5 bilaterally, full-field

ERG was normal, but pattern and flash visual evoked potentials was with delayed latency. At 14.5 years of age visual acuity had decreased to 0.1 in both eyes, optic nerves were atrophic and optical coherence tomography (OCT) showed thinning of retinal nerve fiber layers secondary to optic atrophy. Eye movements, both saccades and smooth pursuits were progressively affected. She had a reduced blinking rate causing dry eyes. A cognitive decline was evident as demonstrated by performance tests: WISC-III at 8 years of age showed an overall IQ score 40 (verbal comprehension score: 58, performance IQ:45); RIAS test at 13 years of age showed an overall IQ score

of 36 (verbal intelligence index 42, nonverbal intelligence index 32); and a WPPSI-R test at 15 years of age showed that she performed as 3 year old child. At her current age, she has ataxia, tremor, dystonic movements, and reduced facial mimicry. She gets water through gastric tube but eats most food by herself; she has a tendency of drooling. She has dysarthria and can say single words and short sentences. She has urinary incontinence (since 13.5 years of age). She can walk short distances without support. Through time, a wide range of other biochemical, metabolic, and genetic investigations (Array-CGH Agilent 400K, FXN-gene, GMA2-gene) has shown normal results. At 3 months of age, a small needle liver biopsy revealed complete loss of normal liver architecture, presence of swollen hepatocytes and a prominent macrophage response in connective tissue. Moreover, fibrosis and inflammation with lymphocytes, plasma cells and granulocytes were present. There was no sign of hemochromatosis or antitrypsin deficiency. Muscle biopsies at the age of 7 and 16 years showed unspecific myopathic changes including type 1 predominance with only

25% type 2 fibers and type 2 fiber atrophy. Activity of respiratory chain enzymes and pyruvate dehydrogenase measured on a muscle biopsy was normal at 15 years of age.

The suggested diagnosis was NBIA, Neurodegeneration with Brain Iron Accumulation.

The family of the M2 patient is German but of Turkish origin. Parents are consanguineous. The M2 patient was, after an uneventful pregnancy, born at term by an uncomplicated caesarean section, weight 3990 g (+1 SD), length 56 cm (+3 SD), and head circumference 38 cm (+3 SD). He is the parents’ only child. Development within the first 15 months appeared to be normal. However, motor developmental

milestones were reached relatively late (walking without support around the age of 19 months, ability to climb stairs around the age of 3 years). According to his parents, speech development started rather early (around the age of 10 months) but didn’t progress adequately and he had ongoing difficulties clearly expressing sounds or words. Starting from the age of 2 years, swallowing difficulties were noted. A first brain

MRI was performed at the age of 4 years but didn’t show clear abnormalities.

Laboratory work-up, metabolic studies as well as human genetic investigations

(exclusion of Simpson-Golabi-Behmel syndrome, fragile X syndrome, DiGeorge syndrome, etc.) were without specific findings. Starting from the age of 7 years, a decline in motor skills was noted. His gait became broad-based and unsecure, dysarthria worsened. A neurodegenerative disorder was suspected and further investigations were performed (muscle biopsy for mitochondrial respiratory chain analysis, exclusion of ataxia-telangiectasia, etc.). At the age of 10 years, bilateral optic atrophy became evident. Urinary incontinence presented at the age of 11 years. At the current age of 16 years, the boy suffers from severe ataxia, dysarthria, dysphagia, dystonia and reduced facial expression. He is wheelchair bound but can stand upright and walk short distances with assistance. Measurements at 13 years were length 160 cm (31st percentile), weight 68 kg (89th percentile) and head circumference 54 cm

(20th percentile).

4.4.2 Cerebral MRI scans

Common features for both patients were bilateral hyperintensities affecting periventricular and cerebellar white matter, low signal intensity in the globus pallidus with hyperintense streaking and low signal intensity in substantia nigra.

4.4.3 Genetic results

A homozygous SLC44A1 variant (M1), (NM_080546.4):c.1549del,p.(Asp517Metfs*19) was identified in individual A-II-2 by investigation of exome data. Sanger sequencing confirmed that the parents were heterozygous carriers, and that the younger brother (A-

II-3) carried the homozygous mutation. A homozygous variant (M2),

(NM_080546.4):c.377_380del,p.(Ser126Metfs*8) was identified in individual B-II-1 by investigation of exome data (235).

Figure 4.1: Photos of the individuals, pedigrees and brain MRI studies. Photos of individuals A-II-2

(M1 patient) and B-II-1 (M2 patient) are shown with the permission of the parents. The pedigrees are shown with indication of the genetic defect in the SLC44A1 gene in individuals, WT=wildtype. Individual

A-II-3 was still an infant at the time of writing and has not developed neurodegenerative disease. Cerebral

Brain MRI studies: A-D) M1 patient, age 13 years; E-H) M1 patient, age 15 years. A-B) and E-F) shows

T2-weighted images, axial view demonstrating bilateral hyperintensities affecting periventricular and cerebellar white matter (white arrows). C-D) and G-H) shows susceptibility weighted imaging (SWI), axial view showing low signal intensities in the globus pallidus and substantia nigra respectively (white arrows) suggesting deposition of paramagnetic material.

4.4.4 Histological investigations

Electron microscopy of skin fibroblasts from both affected individuals demonstrated coarse folding of the plasma membrane, increased presence of secondary lysosomes/endosomes, increased number of vacuoles, and abnormal mitochondria

(Figure 4.2). Additionally, the fibroblasts of the affected individuals displayed an accumulation of iron as observed with the ferric ammonium citrate (FAC) assay (Figure

4.2). Following a 24 h incubation period with ferric ammonium citrate (FAC), all fibroblasts from affected individuals exhibited a profound accumulation of intracellular iron (2-3-fold change) while control fibroblasts showed a 2-fold increase (Figure 4.2).

Immunoblot analysis of FAC treated fibroblasts also demonstrated an increased content of autophagy marker LC3 II in affected individuals (Figure 4.2).

Figure 4.2: Ultrastructure and iron load of cultured skin fibroblasts. (A) Electron microscopy of

SLC44A1 mutated cells (M1, top left and M2, middle left) reveal coarse and rounded protrusions in the plasma membrane compared to the slender projection of the control cells (bottom left) (E). There are multiple small structures containing degenerated material, secondary lysosomes/endosomes, and increased number of vacuoles some of which appears to be degenerated mitochondria with remnants of cristae in M1 (top right) and M2 (middle right). Cytoplasm of normal fibroblasts was seen in bottom right image. Scale bar: 2 μm. (B) Colorimetric ferrozine iron quantification assay. (C) Mutated fibroblasts had increased lipidation of the autophagy marker LC3 (LC3 II/LC3 I ratio) when exposed to high amounts of iron. * denotes significant difference (p < 0.05).

4.4.5 M1 mutant produces a truncated CTL1 protein while M2 mutant results in no protein being produced

The SLC44A1 gene encodes a 657 amino acid protein containing 9 transmembrane domains, with internal N-terminus and external C-terminus (216) (Figure 4.3). The frameshift mutation in M1 introduces a premature stop codon, which predicts a truncated 536 amino acids CTL1 protein with 7 transmembrane domains (Figure 4.3).

The frameshift mutation in M2 introduces a premature stop codon at amino acid 132

(Figure 4.3). To see if the mutants are expressed, the SLC44A1 mRNA was probed in

M1 and M2 cells with primers within 5’- and 3’-end SLC44A1 regions. The amplification of the 5’-region showed that they are expressed but in SLC44A1 mRNA content was

40% lower in M1 and SLC44A1 mRNA content was 75% lower in M2 than in Control cells. Because of the frameshift mutations and the prediction of the premature stop codons, the SLC44A1 mRNA content was not detected in M1 and M2 cells with 3'-end

SLC44A1 primers (Figure 4.3). The N-terminus CTL1 antibody further confirmed that the CTL1 protein was a truncated 59 kDa protein in M1 cells, as opposed to the full length 72 kDa CTL1 protein. The amount of CTL1 protein in M1 was also decreased by

50% with respect to the native CTL1 protein (Figure 4.3). As expected, the C-terminus

CTL1 antibody detected the full length CTL1 protein but did not detect the protein in M1

(Figure 4.3). CTL1 protein was not detected by either CTL1 antibody in M2 (Figure 4.3).

4.4.6 Choline uptake in mutant fibroblasts was reduced but improves with choline treatment

The uptake of choline was reduced by 70-80% in M1 and M2 fibroblasts and the CTL1 specific inhibitor hemicholinium (HC-3) additionally reduced the choline uptake in the mutant fibroblasts (Figure 4.3). Choline supplementation improved the uptake in a concentration dependent manner and the uptake in M1 cells at 5 mg/ml choline was

50% of basal control levels (Figure 4.4). Under the same conditions the CTL1 protein content was increased by in M1 cells 30% at 2.5 mg/ml choline and 70% at 5 mg/ml choline (Figure 4.4). The supplemental choline was also able to increase SLC44A1 mRNA content up to 60% in M1 (Figure 4.4).

Figure 4.3: Topology and transport function of the native and mutated CTL1 proteins. (A, B, C)

Schematics of SLC44A1 topology in Control, M1 and M2 cells. (D) mRNA expression of SLC44A1 in

Control, M1 and in (E) M2 cells. (F) SLC44A1 protein content in Control, M1 and (G) M2 cells. (H) Choline uptake in M1 and (I) M2 cells after HC-3 inhibition. * denotes significant difference (p < 0.05).

Figure 4.4: Choline stimulates CTL1 transport and protein abundance in M1 cells. (A) Choline uptake in Control and M1 cells supplemented with choline. (B, C) CTL1 protein content in Control and M1 cells supplemented with choline. (D) mRNA expression of SLC44A1 after choline supplementation in

Control and M2 cells. * denotes statistically significant difference (p < 0.05) relative to 0 mg/L choline treatment, # denotes statistically significant difference (p < 0.05) relative to 2.5 mg/L choline treatment.

4.4.7 Altered expression of other choline transporters in mutant fibroblasts

The mRNA expression of various families of transporters which have demonstrated the ability to transport choline is stated in Figure 4.5. The expression of organic cation transporter 1, SLC22A1 (OCT1), was reduced 40% in mutant fibroblasts. The main role of OCT1 is to transport cationic drugs (238) but it also has a low binding affinity for choline (239). Interestingly, carnitine transporter 2, SLC22A5 (OCTN2), which also has a low binding affinity for choline (240) was highly upregulated in mutant fibroblasts however completely absent in the control fibroblasts. The expression of SLC44A2

(CTL2) and SLC22A4 (OCTN1) was not altered between mutant and control cells.

Lastly, SLC44A3 (CTL3), SLC44A4 (CTL4), SLC44A5 (CTL5), SLC22A2 (OCT2),

SLC22A3 (OCT3) and the neuron-specific choline transporter SLC5A7 (CHT1) (179), were not detected in any of the cells (Figure 4.5).

Figure 4.5: mRNA expression of choline related transporters is dysregulated in mutant fibroblasts.

Among organic cation transporters, SLC22A1 (OCT1 organic cation transporter) mRNA was decreased by 40% while SLC44A4 (CTL4 choline transporter) mRNA was increased 25% in M1 and M2 fibroblasts.

Interestingly, SLC22A5 (OCTN2 carnitine transporter associated with primary carnitine deficiency) was abundantly detected in mutant fibroblasts but not in control fibroblasts. Lastly, the transcripts for SLC44A2

(CTL2 choline transporter) and SLC22A4 (OCTN1 carnitine transporter) were not affected, and SLC44A3

(CTL3), SLC44A5 (CTL5), SLC22A2 (OCT2), SLC22A3 (OCT3) and SLC5A7 (CHT1 neuron specific choline transporter) were not present in Control and mutant fibroblasts. * p < 0.05 relative to Control cells.

4.4.8 Changes in phospholipid gene expression as a likely measure to preserve membrane PC

Due to the diminished ability to transport choline, it was not surprising that the expression of the genes involved in PC synthesis and degradation were altered in the mutant cells (Figure 4.5). The rate-limiting enzyme for PC synthesis via the CDP-choline pathway, Pcyt1, was decreased by 50-70% and choline kinase (CK) and ethanolamine kinase (EK) were decreased by 20-30%. The mRNA expression of Pcyt2, the rate- limiting enzyme for PE synthesis via the CDP-ethanolamine pathway and the PS synthesis (PSS1, PSS2) and decarboxylation (PSD) genes were not altered in the mutant cells. To generate additional PC and to counteract the diminished ability to use the extracellular choline, the expression of PE methyltransferase, PEMT, that facilitates

PC production via PE methylation was increased by 25%. Phospholipase D (PLD1 and

PLD2) that facilitates the release of choline from PC was also increased in M1 by 20-

25% (Figure 4.6).

Figure 4.6: mRNA expression of phospholipid genes is dysregulated in mutant fibroblasts. Among phospholipid genes de novo PC synthesis genes PCYT1 (CTP: phosphocholine cytidylyltransferase) and

CK (choline kinase) were 50% and 20% downregulated in mutant cells. In the de novo PE synthesis pathway, EK (ethanolamine kinase) expression was decreased by 30%. The genes for PS synthesis

(PSS1, PSS2) and PE synthesis PSD (PS decarboxylase) were not affected in mutant cells. The expression of PEMT (PE methyltransferase) that produces PC from PE was increased by 25% in mutant cells. Finally, the expression of phospholipase D (PLD1 and PLD2) which degrades PC, was 20-25% higher in mutant cells relative to control cells. * p < 0.05 relative to Control cells.

4.4.9 Increased expression of lipogenic genes and reduced expression of fatty acid oxidation genes as a response to deficiency in choline transport

The expression of a group of genes involved in de novo FA and triglyceride synthesis by lipogenesis (SREBP1c, FAS, DGAT1 and DGAT2) were modestly but significantly upregulated in mutant cells (Figure 4.5). Conversely, the expression of genes involved in mitochondrial fatty acid transport and oxidation (PGC1α, PPARα and CPT1) were downregulated 35%-60% in mutant cells (Figure 4.7). This pattern where the expression of genes devoted to lipogenesis are increased while the expression of genes devoted to

FA oxidation are decreased is well documented in scenarios where choline uptake and substrate flow through the CDP-choline pathway are suboptimal (241).

Figure 4.7: mRNA expression of fatty acid genes is dysregulated in mutant fibroblasts. Among fatty acid genes (D), the lipogenic genes for fatty acid and triglyceride synthesis (FAS, SREBP1 and DGAT1/2) were upregulated 20-30% while the genes responsible for the mitochondrial fatty acid transport and oxidation (CPT1, PGC1α, PPARα) were significantly downregulated (35-60%) in mutant cells. * p < 0.05 relative to Control cells.

4.4.10 Preservation of mutant cell PC results in deficiency in other membrane phospholipids

The effect of diminished choline transport on de novo PC synthesis was investigated by monitoring choline flux through the CDP-choline (Kennedy) pathway (216). The reduced rate of [3H]-choline incorporation into P-Cho, CDP-Cho and PC in mutant cells is an indication of reduced de novo PC synthesis (Figure 4.8). On the other hand, the PC produced de novo from choline was more readily degraded in the mutant cells (Figure

4.8), likely to be used as a source of additional choline. The substrate flux through the

CDP-ethanolamine branch of the Kennedy pathway was similarly reduced and the 14C-

PE formed from 14C-ethanolamine was more readily degraded in the mutant cells than in the Control cells (Figure 4.8). Surprisingly, the equilibrium radiolabeling of phospholipids with [3H]-glycerol revealed that the total PC pool was not reduced in M1 and M2 cells; however the PE and PS pools were largely reduced in the mutant cells

(Figure 4.8). The reduced PE and PS pools can imply that those phospholipids were used to maintain membrane PC under conditions of reduced choline transport. Indeed, choline supplementation to M1 and M2 cells did not change the PC pool but significantly improved the PE and PS pools in mutant cells (Figure 4.8). Since the expression of PE methyltransferase (PEMT) was increased in mutant cells (Figure 4.8), the PE degradation to generate PC by the PE methylation pathway was monitored using [14C-

(CH3)]-methionine (Figure 4.8). The results showed that the mutant cells could produce

PC from PE and that choline addition could provide a small reduction in PE methylation.

Additionally, the increased PC degradation (Figure 4.8) and elevated PLD expression

(Figure 4.8) to produce more choline also took place in mutant cells. This was further supported with elevated PLD activity in the mutant cells that was reversed by choline treatment (Figure 4.8).

Figure 4.8: PC and PE de novo biosynthesis and turnover. (A) [3H]-choline pulse experiments of

CDP-choline pathway metabolites. (B) [3H]-choline pulse and pulse-chase experiments of PC production and degradation. (C) Pulse and pulse-chase experiments with [14C-Etn] of Etn and PE degradation and

PE production.

Figure 4.9: Analysis of equilibrium phospholipid levels. (A) Equilibrium levels of PC, PE and PS determined with [3H]-glycerol radiolabeling. (B) PE methyltransferase activity quantification with the [14C-

(CH3)]-methionine radiolabel. (C) Phospholipase D enzyme activity analysis. * denotes statistically significant difference (p < 0.05) relative to control cells, # denotes statistically significant difference (p <

0.05) relative to untreated mutant cells.

4.4.11 Membrane phospholipid profiles are greatly modified in mutant cells

In agreement with the metabolic fluxes and the phospholipid pool sizes in Figure 4.9, the M1 cells had unaltered total PC but were deficient in two other membrane phospholipids PE and PS (Figure 4.9). The M1 cells also had an increased % contribution of PI, PA and PG, likely indicating further changes in the CDP-DAG pathway for PI and CL synthesis (Figure 4.9).

Predominant ether phospholipids in Control fibroblasts are ether-PC (43%) and ether-

PE (38.8 %) and their levels decreased 6-11% in mutant cells. At the same time, ether-

PI (12.5%) and ether-PA (4.3 %) increased 8-9% (Figure 4.9). Ether phospholipids are known to be enriched in polyunsaturated fatty acids (PUFA) (37, 242). The ether lipid profiling showed that C18:1 n-9 (oleic acid), C20:4 n-6 (arachidonic acid, AA) and C20:5 n-3 (EPA, eicosapentaenoic acid) were more abundant in M1 cells while these same cells were depleted of very-long chain fatty acids C22:n=3, 4, 5 or 6 (VLCFA). A similar deficiency in VLCFA was present in PC (Figure 4.9) and in all other membrane lipids, thus representing a general feature of the mutant cells. There was also an enrichment in

C20:4 n-6 (arachidonic acid, AA) in the main phospholipids which produced an altered ratio of AA (n-6) and DHA (n-3) in the mutant cells. The increased abundance of AA at the expense of DHA is frequently seen in plasmalogen deficiency disorders (37) and likely plays a pathogenic role in the two individuals with CTL1 mutations. Although

VLCFA were decreased, the expression of multiple elongation genes was elevated

(ELOVL3, 5, 6, 7 > 1.5-fold) (Figure 4.9) apparently as compensation for the primary defects in PUFA elongation and desaturation pathways. Finally, the lipidomic analysis

showed that M1 cells generally had shorter carbon chains in PE and PS and longer carbon chains in PG and PI. Additionally, the main phospholipids (PC, PE and PS) were generally more saturated whereas the metabolic products PA and PG were enriched in n-6 PUFA, apparently because of generally increased fatty acid remodeling and drastic

PE and PS depletion in the mutant cells.

Figure 4.10. Glycerolipid subclasses and fatty acid composition are modified in mutant cells. (A)

Display of phospholipid composition and (B) ether lipid composition in Control and M1 cells. (C) Cluster analysis demonstrating the differences in fatty composition in PC between Control and M1 cells. (D) mRNA expression of ELOV genes in Control, M1 and M2 cells. * p < 0.05 relative to Control cells.

4.5 Discussion

We have presented two unrelated adolescent individuals with homozygous frameshift mutations in the SLC44A1 gene encoding the choline transporter 1 protein (CTL1). The molecular defect resulted in an expressed but truncated CTL1 protein in one individual, while CTL1 was completely absent in the other. The clinical features were emblematic of a neurodegenerative disorder with progressive ataxia, tremor, dystonia, cognitive decline, optic nerve atrophy with normal ERG and urinary incontinence. Cerebral MRI scans showed hypointensity of globus pallidus with a hyperintensive streak, hypointensity of substantia nigra, white matter disturbances and atrophy. The clinical features together with the MRI changes raised suspicion of an NBIA.

We have demonstrated that CTL1 deficiency results in abnormal cell membranes with increased accumulation of membrane materials in cells. The increased number of lysosomes and vacuoles in mutant cells most likely reflects disturbances in the regulation of vesicular trafficking, in which CTL1 plays an important role (11, 243), a feature which was recently established as a common defect in NBIA (236). We report that the mutant fibroblasts retain abnormally high levels of iron, a hallmark characteristic of NBIA. Moreover, as in other NBIA cases (236), LC3 II content was increased in mutant cells, which was indicative of an induction of autophagosome formation (244) and intracellular iron accumulation (236).

These studies implicate CTL1 deficiency in a new subtype of NBIA. NBIA is a group of neurodegenerative diseases caused by accumulation of iron in basal ganglia with

characteristic clinical features and MRI scans. Clinical core symptoms comprise a combination of early onset dystonia, pyramidal and extrapyramidal signs with ataxia, cognitive declines, behavioral abnormalities and retinal and axonal neuropathy variably accompanying these core features. Increased non-age-associated brain iron, most pronounced in the basal ganglia, is considered a unifying characteristic of these disorders (245). Subgroups of NBIA demonstrate a link between neurodegeneration and alteration of phospholipid metabolism, mitochondrial morphology and membrane remodeling (233). Comparing the features of our patients with different subgroups of

NBIA, we find most similarities to MPAN (Mitochondrial Membrane Protein-Associated

Neurodegeneration). Therefore, the characteristics in common with MPAN patients are ataxia and Parkinsonian features (tremor and reduced facial mimicry), optic nerve atrophy combined with a normal ERG, which is considered rather distinct for MPAN

(246, 247), progressive cognitive decline (246), a characteristic pattern of hyperintense streaking that highlights the entire length of the medullary lamina between the internal and external segments of the globus pallidus (248), hypointensity of substantia nigra and white matter defects around lateral ventricles and in the cerebellum (249). The causative gene for MPAN is C19orf12, whose function is still unknown. CTL1 mediated choline transport is directly functionally coupled with the CDP-choline (Kennedy) pathway for PC synthesis (215, 216, 217) and not surprisingly, the substrate flux through this pathway was downregulated in mutant fibroblasts. The content of PC was however unchanged showing that the primary adaptation to CTL1 deficiency was to preserve PC as an additional source of choline. The constant PC levels were however

maintained at the expense of other membrane phospholipids (250), predominantly PE, which was reduced in CTL1 mutant cells. PC could be made from PE via PE methylation (237) and this pathway was modestly upregulated in mutant cells. Deficient cells also had reduced membrane PS. PS is formed from PE and PS, and its low levels are likely caused by a combination of reduced synthesis and/or increased degradation to PE.

Efforts to produce choline intracellularly were further evident from the increased phospholipase D activity and excessive abundance of phosphatidic acid product in

CTL1 mutant cells. Phosphatidic acid is an important signaling molecule which was likely redirected from PC and PE synthesis towards PI and PG synthesis upon Kennedy pathway impairment. This occurs via the CDP-DAG pathway, as manifested by the accumulation of PI and PG in the mutant cells. The ether (alkyl acyl; plasmalogen) phospholipid deficiencies are strongly linked to peroxisomal disorders and more common neurological disorders like Alzheimer’s disease and Parkinson’s disease (137).

Although their biological function is not clear, the ether lipids are abundant in the brain, particularly in PE (251). There was a deficiency in ether-PE in mutant cells that could be contributing to the observed pathologies by the inhibition of membrane fusion and ion transport (251, 252). As a further consequence of adaptation to CTL1 transport deficiency (40, 191), the mutant cells had reduced expression of genes devoted to mitochondrial β-oxidation. There was a general deficiency in very long chain fatty acids

(C22 n=3, 4, 5 and 6) and an accumulation of arachidonic acid (C20:4 n-6) in membrane phospholipids. How CTL1 deficiency affects the very long chain fatty acids

elongation at the level of endoplasmic reticulum and the β-oxidation system in the mitochondria is not clear but a link likely exists between very long chain fatty acid content and CTL1 function.

Functional studies showed an improved uptake of choline in patient cells when choline was added to the medium. This finding provokes the thought of choline supplementation as a possible treatment for this disease. Since the genetic defect was discovered, patients have been treated with supplemental choline and later, GPC. Treatment of patients with choline has not resulted in an improvement of symptoms, but the condition of the M1 patient has stabilized during treatment. The symptoms of the M2 patient progressed during treatment, and the GPC supplementation was discontinued after a treatment course of 1 year. The symptoms in both adolescent patients had been progressing steadily before initiation of treatment, and the fact that the symptoms of the

M1 patient have been unchanged during treatment might indicate a positive effect of the treatment. Importantly, the M1 patient has a residual truncated CTL1 function, which might explain the difference in treatment response to the M2 patient. Since deficiency in

PE, PS, ether lipids, and very long chain fatty acids represents the major abnormalities and are known contributing factors in multiple neuronal pathologies (251, 252), future therapies may focus on phospholipid supplementation, particularly n-3 enriched phospholipids from krill oil. The observed deficiency in ether phospholipids could be reduced by supplementation of alkylglycerol precursors, enriched in shark liver oil, a well-known natural source of alkylglycerols. The M2 patient, which did not respond well to choline therapy, may particularly benefit from those phospholipid therapies.

We have shown that biallelic pathogenic variants in the SLC44A1 gene encoding the

CTL1 protein are implicated in a new subtype of NBIA. This new subtype of NBIA resembles MPAN in clinical features as well as cerebral MRI changes. The primary outcome of CTL1 deficiency was severely diminished choline transport, and membrane

PC was subsequently utilized as an intracellular source of choline. The homeostatic response to this essential need for PC (and choline) results in the loss of inner membrane phospholipids PE and PS and deficiency in very long chain fatty acids.

These features together represent the most critical causes of defective plasma membrane and organelle function in CTL1 deficiency.

Study design: C.R. Fagerberg, F. Distelmaier, M. Bakovic, A. Taylor, Experimental work: A. Taylor, C.R. Fagerberg, F. Distelmaier, Manuscript preparation: C.R.

Fagerberg, F. Distelmaier, M. Bakovic, A. Taylor, M. Kibaek, D. Wieczorek, H.D.

Schroder, M.J. Larsen, R.A. Jamra, E. Gade, L. Markovic, D. Klee, L. Brady, M.

Tarnopolsky, P. Agarwal, V. Dolinsky. Special thanks to Heiko Runz for exome sequencing assistance and Michael Leadley for lipidomics assistance.

CHAPTER 5 RATIONALE

In Chapter 4, we further elucidated the role of SLC44A1 in choline uptake. However, the mechanism of ethanolamine uptake has yet to be elucidated in humans. Furthermore, in the literature there are data which suggest that choline and ethanolamine may be taken up into the cell by the same transporter. This is largely thought to be because choline and ethanolamine share similar molecular structures which are believed to be critical for protein mediated uptake. To elucidate the mechanism of ethanolamine uptake, we utilized the human skin fibroblasts from Chapter 4 to study the implications of SLC44A1 in ethanolamine uptake.

CHAPTER 5: THE NOVEL ROLE OF SLC44A1/CTL1 IN ETHANOLAMINE TRANSPORT

5.1 Abstract

PC and PE comprise the bulk of cell membrane phospholipids. While it is established that CTL1 mediates cellular choline transport, an ethanolamine transporter has yet to be identified. In this study, we examine the function of CTL1 as an ethanolamine transporter in skin fibroblasts in addition to its known role in the transport of choline.

Ethanolamine uptake was mediated by a pH dependent system which was also Na+ independent. Control cells have two distinct ethanolamine transport systems as demonstrated by two different Km values (66.5 ± 8.5 μM (Km1) and 299.0 ± 13.1 μM

(Km2). CTL1 overexpression enhanced ethanolamine uptake in M2 cells, which are devoid of CTL1 function. Furthermore, we demonstrated a further decrease in ethanolamine uptake when CTL2 expression was knocked down in M2 cells, further supporting the roles for both CTL1 and CTL2 in ethanolamine uptake. Moreover, both

CTL1 and CTL2 appear to facilitate mitochondrial ethanolamine uptake, with CTL1 being the predominant transport system. Furthermore, choline and ethanolamine treatment in tandem reduced the uptake of both choline and ethanolamine. This was an indication that these two substrates compete for the same transport system. Lastly, treatment with HC-3 and CTL1 antibodies reduced PE synthesis. Together, these data demonstrate the novel role for CTL1 in the transport of ethanolamine.

5.2 Introduction

PC and PE are major components of cellular membranes and they are also involved with essential cellular processes (25, 26). The major pathway for PC and PE production is via de novo synthesis through the CDP-choline and CDP-ethanolamine branches of the Kennedy pathway, respectively, of which choline and ethanolamine are initial substrates. Three choline transport systems have previously been established: low- affinity diffusion: intermediate-affinity, Na+-independent transport; and high-affinity, Na+- dependent transport (253). The intermediate-affinity choline transporter-like protein 1

(CTL1/SLC44A1) is ubiquitously expressed in mouse, rat and human tissues and was originally discovered as a choline transporter for PC synthesis (27). More recently,

CTL1 was shown to mediate the transport of choline across mitochondrial membranes

(216). Choline transport has been extensively studied however, there has been less focus on elucidating the mechanisms of ethanolamine transport, despite its biological importance.

The supply of ethanolamine as a substrate for the initial step in the CDP-ethanolamine pathway is essential for PE synthesis (254). PE is the most abundant lipid on the cytoplasmic leaflet of cellular membranes and plays important roles in cell division, membrane fusion, autophagy and apoptosis (255). Moreover, PE is important for the production of other phospholipids, such as PC via PE N-methyltransferase (PEMT) and

PS via PS synthase 2 when free ethanolamine is released. PE lipolysis to generate

DAG and fatty acids by phospholipases also releases ethanolamine or phosphoethanolamine that needs to be transported in and out of cells or reincorporated

into PE. In addition, the inner mitochondrial membrane is enriched with PE, meaning that mitochondrial ethanolamine transport is likely critical for mitochondrial biogenesis

(256).

The supply of ethanolamine as a substrate for the initial step in the CDP-ethanolamine pathway is essential for de novo PE synthesis. PE is the most abundant lipid on the cytoplasmic leaflet of cellular membranes and plays important roles in cell division, membrane fusion, autophagy and apoptosis. Moreover, PE is important for the production of other phospholipids, such as PC via PE methyltransferase and to phosphatidylserine (PS) by phosphatidylserine synthase 2 when free ethanolamine is released. PE lipolysis to DAG and fatty acids by phospholipases also releases ethanolamine or phosphoethanolamine that needs to be transported in and out of the cells or reincorporated into PE.

While the importance of ethanolamine for many biological processes is clear, an ethanolamine uptake system has yet to be characterized in humans. Ethanolamine is not endogenously produced and therefore needs to be transported into the cell (255).

While ethanolamine transport has not been characterized in humans, previous investigators have identified examples of ethanolamine uptake systems in other organisms (16, 257, 258). In bacteria found in the human gastrointestinal tract, the EutH protein facilitates the uptake of ethanolamine to be utilized as an intracellular source of carbon and nitrogen (259, 260). Kinetic studies in bovine endothelial cells, human Y79 retinoblastoma cells, chick embryo neurons and glial cells. Additionally, ethanolamine transport likely shares biophysical properties with choline as the hydroxyl groups of both

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compounds are thought to help facilitate hydrogen bonds with the transport protein

(261).

Here, we demonstrated that CTL1 and CTL2 are capable of facilitating ethanolamine transport, in addition to their role in choline transport. We characterized ethanolamine transport in cells with errors in CTL1 transport due to inherited SLC44A1 gene mutations and in cells overexpressing SLC44A1/CTL1. Additionally, we examined the effects of choline and ethanolamine competition on ethanolamine and choline uptake.

Finally, the effects of pharmacological and antibody induced CTL1 inhibition on PC and

PE synthesis were evaluated to separate the contributions of CTL1 and CTL2 in ethanolamine metabolism and PE synthesis. This study was the first to our knowledge to demonstrate the activity of SLC44A1/CTL1 and SLC44A2/CTL2 as ethanolamine transporters.

5.3 Methods

5.3.1 Maintenance of cell lines

MCF-7 human breast cancer cells, MCF-10 human mammary epithelial cells and COS-

7 fibroblast-like cells were maintained in DMEM (Fisher Scientific) supplemented with

10% fetal bovine serum and 2% penicillin/streptomycin; Control, M1 and M2 skin fibroblasts were maintained in MEM (Fisher Scientific) supplemented with 20% fetal

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bovine serum and 2% penicillin/streptomycin. Cells were kept in a humidified

o atmosphere at 37 C and 5% CO2.

5.3.2 Drug treatments in cancer cells

Approximately 8.0 x 104 cells/well were seeded in 6-well plates and grown for 24 h.

After 24 h of growth, drugs or antibodies were added to cells and incubated for 24 h.

Working concentrations of HC-3 and CTL1 antibody were specific to each cell line representing the IC50 concentrations.

5.3.3 Radiolabeling and PC and PE determination

Cells were passaged and grown for 24 h. After 24 h of growth, media was removed and replaced with media containing 0.2 μCi [3H]-choline or [14C]-ethanolamine (ARC St.

Louis, MO) for 24 h. Media was then removed and cells were washed 2x with 1x PBS.

Lipids were then extracted with the method of Bligh and Dyer (194). PE and PC content were determined by LSC.

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5.3.4 Kinetics of ethanolamine transport

Kinetic analyses were performed as described (239). Cells were incubated (20 min) with increasing concentrations of unlabeled ethanolamine (0-1000 µM) before being treated with 0.2 µCi [14C]-ethanolamine for 20 minutes.

5.3.5 Choline and ethanolamine uptake

Choline and ethanolamine uptake were measured according to our previously standardized protocols (216). Briefly, cells were washed with KRH buffer (130 mM

NaCl, 1.3 mM KCl, 2.2 mM CaCl2, 1.2 mM MgSO4, 1.2 mM KH2PO4, 10 mM 4-(2- hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) (pH 7.4) and 10 mM glucose and incubated with 0.2 μCi [3H]-choline or [14C]-ethanolamine with the compound of interest for 20 minutes at room temperature. Cells were then washed in ice cold KRH buffer containing 500 μM ‘cold’ choline or ethanolamine to stop the uptake reaction.

Cells were then lysed in 500 μl ice cold lysis buffer (10 mM Tris-HCl, 1 mM EDTA and

10 mM NaF) and choline and ethanolamine uptake were analyzed by LSC.

For the choline/ethanolamine uptake competition assay, control cells were incubated with 200 µM choline, 200 µM ethanolamine or 200 µM choline + 200 µM ethanolamine for 20 minutes. To study the effects of cationic compounds (choline, ethanolamine, HC-

3, betaine, verapamil, quinine and nifedipine; all purchased from Sigma Aldrich) on ethanolamine uptake, control and M2 cells were incubated with the compounds for 20

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minutes. To assess the effects of Na+ concentration on ethanolamine uptake, control and M2 cells were subjected to either standard KRH buffer or KRH buffer with LiCl instead of NaCl. To study the effect of pH on ethanolamine uptake, control and M2 cells were treated with KRH buffers of varying pH (pH 5.5, 6, 6.5, 7, 7.5, 8 and 8.5). Buffers were prepared by mixing 10 mM MES (pH 5.5) and 10 mM bicine (pH 8.5).

5.3.6 RNA extraction and RT-PCR

Total RNA from Control and M2 cells was isolated with TRIzol (Invitrogen, Life

Technologies Incorporated, Burlington, ON, Canada). DNase I was used to eliminate genomic DNA and cDNA was synthesized from 2 μg RNA using a 3-AP primer and RNA

SuperScript III reverse transcriptase (Invitrogen, Life Technologies Incorporated).

Expression of CTL1, CTL2, CTL3, CTL4, CTL5, CK, EK, Pcyt1, Pcyt2, PSD, PSS1 and

PSS2 was determined by PCR using the primers and conditions outlined in Table 2.

Reactions were standardized by amplifying glyceraldehyde 3-phosphate dehydrogenase (GAPDH) and relative band intensity was quantified using ImageJ software (NIH, Bethesda, MD, USA).

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Table 5.1: Primer sequences and melting temperatures (TM) for polymerase chain reaction (PCR) using human fibroblasts RNA

Gene Forward Primer (5’-3’) Reverse Primer (5’-3’) TM Size (oC) (bp) GAPDH ACCACAGTCCATGCCATCAC TCCACCACCCTGTTGCTGTA 58 452 SLC44A1 CTCTTCTGCATTGGGATGGGA CTTGGACACGCTGCTACACA 57 248 Pcyt1 CCCTCTTTCCGATGGCCCTT TGTTTGGGTTCCCCCAGTCC 53 267 Pcyt2 ACATCATCGCGGGCTTACAC TGACACACCAGGTCCACCTT 53 190 CK GAGTTCCACATCAGTGTCATCAGA ATGGCCTCAGCCCCTTGAAAT 57 203 EK AATGGTGCCGGGAGTACTTG CCCTCTGGGAAGACTCCGTA 57 274 PSS1 CGTTCACTCGACCTCATCCA GATTGTCCAGCAGAGACCGT 56 302 PSS2 ATGCACCGCACACCCTTTA TCTTGGTGTTGTAGGCCGTG 57 457 PSD TTAGGTGGGGAGTAGCCACA GCTTCCGTAAGGGCTGTAGG 57 273 SLC44A2 CCACGCCTACAAGGGTGTCC CCATGGCAATGACCAGGCCT 58 203 SLC44A3 TTCTTCTGGGTCCTCTGGGT GGTACCTGGACAATGCACCA 56 397 SLC44A4 ATCACTCCGGAGACTGAGCCA CCCTGGCAGACAAAAGTTCCT 58 473 SLC44A5 TTGCAGTGATTCAGGCCTACTT AGGTTTAGCAACACGGAGGG 56 372

5.3.7. Expression of Myc-tagged rSLC44A1 cDNA and SLC44A2 siRNA in fibroblasts

Confluent control and M2 cells were transfected with 5 µg of pCMV3-ORF-C-Myc-

SLC44A1 cDNA (Sino Biologics) or empty vector (pcDNA4His-Max B) using

Lipofectamine 2000 (Invitrogen). Control and M2 cells were transfected with 30 nM

CTL2 siRNA (Santa Cruz Biotechnology; sc-62163) using siPORT lipid transfection agent.

5.3.8 Immunoblotting

All fibroblasts were washed 3x with 1x PBS and subjected to a lysis buffer (25 mM Tris,

15% glycerol, 1% Triton X-100, 8 mM MgCl2, 1 mM DTT, protease inhibitor cocktail and

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phosphatase inhibitor cocktail) at 4oC for 30 minutes and scraped into 1.5 ml tubes.

Protein concentration was determined with the bicinchoninic (BCA) assay (Pierce,

Rockford, IL, USA). The LV58 (N-terminus) and ENS-627 (C-terminus) antibodies (both

1:500 in skim milk in TBS-T) detect the 72 kDa CTL1 protein under native (non- denaturing) conditions. Samples were mixed with loading buffer (62 mM Tris-HCl,

0.01% bromophenol blue and 10% glycerol) and separated by PAGE at 120 V for 1.5 h.

CTL1 was resolved on an 8% native gel and proteins were transferred onto PVDF membranes (Pall Canada, Mississauga, ON, Canada) by a semi-dry transfer system and stained with Ponceau S. Membranes were blocked in 5% skim milk in Tris Buffered

Saline-Tween 20 (TBS-T) solution and then incubated with either one of the CTL1 antibodies (1:500 in 5% skim milk in TBS-T) overnight at 4oC. Membranes were washed with TBS-T and then incubated with an anti-rabbit horseradish peroxidase conjugated secondary antibody (New England Biolabs, 1:10,000 in 5% skim milk in TBS-T) for 2 h.

Membranes were washed in TBS-T and proteins were visualized using a chemiluminescent substrate (Sigma Aldrich, Oakville, ON, Canada).

5.3.9 Mitochondrial isolation

Mitochondria were isolated from control and M2 cells as described (20). In brief, cells were incubated for 20 minutes in ice-cold RSB swelling buffer and homogenized; 19 ml

MS buffer was added and the cell homogenate was centrifuged at 2500 rpm for 5 minutes. This step was repeated twice and the final supernatant was centrifuged at

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12,500 rpm. The resulting pellet was resuspended in MS buffer and mitochondrial ethanolamine uptake was conducted as previously described.

5.3.10 Lipid analysis

Phospholipid (PC, PS and PE) and neutral lipid (DAG and TAG) pools were determined as previously described (190). The lipids from both classes were extracted by the method of Bligh and Dyer (194) and separated by TLC.

5.3.11 Pcyt2 Activity Assay

The assay was conducted as described (262). In brief, Control and M2 cells were cultured for 24 h and 50 µg protein was dissolved in Pcyt2 reaction mixture. The mixture was treated for 15 minutes with 0.2 µCi [14C]-P-ethanolamine and the reaction was terminated by boiling for 2 minutes. The radiolabeled Pcyt2 reaction product [14C]-CDP- ethanolamine was isolated by TLC as previously described (262). Pcyt2 activity was expressed as nmol/min/mg protein.

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5.3.12 Statistical analysis

All measurements are expressed as means from quadruplets ± SEM. Statistical analysis was performed using GraphPad Prism 4.0 software (GraphPad, Inc.). Data were subjected to Student’s T-test. Differences were considered statistically significant at *p < 0.05.

5.4 Results

5.4.1 CTL1 inhibition with HC-3 and CTL1 antibody on PC and PE synthesis

MCF-7 and MCF-10 cells were treated with 50 μM HC-3 and 0.5 μg of CTL1 antibodies to assess the magnitude by which CTL1 affects PC and PE production. CTL1 inhibition by HC-3 reduced PC production in MCF-7 and MCF-10 by 50% and 75% whereas PE production was reduced by 50% in MCF-7 and MCF-10 cell lines (Figure 5.1). CTL1 antibody treatment reduced PC content by 50% and 60% in the MCF-10 and MCF-7 cell lines respectively. PE content was decreased by 50% in both cell lines after CTL1 antibody treatment. This suggests CTL1 may mediate the transport of ethanolamine, in addition to choline (Figure 5.1).

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Figure 5.1: CTL1 inhibition with HC-3 and CTL1 antibody on PC and PE synthesis. PC and PE determination in MCF-10 and MCF-7 cells after HC-3 and CTL1 antibody treatment in (A) MCF-10 and

(B) MCF-7 cells. ** p < 0.01, *** p < 0.001.

5.4.2 Kinetics of ethanolamine uptake and effects of extracellular pH and Na+ on ethanolamine uptake

To analyze the kinetics of ethanolamine uptake, cells were treated for 20 minutes with

[14C]-ethanolamine at concentrations ranging from 0 to 1000 μM. Our kinetic analysis

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yielded Michaelis-Menten (Km) constants of 66.5 ± 8.5 μM (Km1) and 299.0 ± 13.1 μM

(Km2) in Control cells, 55.6 ± 14.8 μM (Km1) and 277.3 ± 7.9 μM (Km2) in COS-7 cells,

279.6 ± 7.1 μM (Km1) in M1 cells and 275.4 ± 3.9 μM (Km1) in M2 cells. Vmax values were 26.9, 26.3, 21.2 and 20.6 nmol/mg protein/min for Control, COS-7, M1 and M2 cells respectively. The Eadie-Hofstee plots were non-linear in Control and COS-7 cells but linear in M1 and M2 cells. This suggests that ethanolamine uptake was mediated by two transport systems in Control and COS-7 cells but only one transport system in M1 and M2 cells.

We analyzed ethanolamine uptake with and without the presence of 10 μM extracellular

Na+ in both Control and M2 cells (Figure 5.2). Ethanolamine uptake was modestly decreased when NaCl was replaced by LiCl in the uptake buffer. Additionally, we analyzed the effect of extracellular pH on ethanolamine uptake in Control and M2 cells by changing the pH of the uptake buffer (Figure 5.2). Ethanolamine uptake was decreased when extracellular pH was 5.5 and 6 but was slightly increased when pH was 8.5. These data indicate that ethanolamine uptake in all cells was pH dependent.

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Figure 5.2: Kinetics of ethanolamine uptake. (A) Ethanolamine uptake curves (0-1000 μM) of Control,

COS-7, M1 and M2 cells. The Eadie-Hofstee plots demonstrate that two systems facilitated ethanolamine uptake in Control and COS-7 cells and one system facilitated ethanolamine uptake in M1 and M2 cells.

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Figure 5.3: Effects of extracellular pH and Na+ on ethanolamine uptake. (A) Effects of extracellular pH on [14C]-ethanolamine uptake in Control and M2 cells. The uptake was measured for 20 minutes under different pH conditions. * denotes statistically significant difference (p < 0.05) relative to pH 7.5. (B)

Time course of 10 μM [14C]-ethanolamine uptake in the presence and absence of extracellular Na+ in

Control and M2 cells for 60 minutes. Na+ free buffer was prepared by replacing NaCl with an equimolar concentration of LiCl.

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5.4.3 Effects of various compounds on CTL1 mediated uptake

In Control cells, we conducted a competition assay to elucidate how choline and ethanolamine influence the uptake of each other (Figure 5.3). Choline uptake was diminished by 40%, 70% and 80% with 200 µM ethanolamine, 200 µM choline and 200

µM ethanolamine + 200 µM choline treatments respectively (Figure 5.3). Ethanolamine uptake was diminished by 50%, 40% and 70% with 200 µM ethanolamine, 200 µM choline and 200 µM ethanolamine + 200 µM choline respectively (Figure 5.3). The inhibition of choline and ethanolamine uptake by a high concentration of the opposite substrate demonstrates that both substrates are competing for the same transport system.

It is known that OCTs and CTL1 recognize numerous organic cations as substrates and display an overlap in substrate specificity. We assessed the effects of numerous organic cations on ethanolamine uptake in Control and M2 cells. Numerous organic cations inhibited ethanolamine uptake in a concentration dependent manner. Ki values were calculated from inhibition curves and are displayed in Table 5.2. Organic compounds including HC-3 (prevalent inhibitor of CTL1), unlabeled choline, unlabeled ethanolamine, betaine, verapamil, quinine and nifedipine affected ethanolamine uptake in both Control and M2 cells.

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Figure 5.4: Effects of various compounds on CTL1 mediated uptake. (A) Control cells were pre- incubated with ethanolamine, choline or both compounds for 30 minutes and the uptake of [14C]- ethanolamine or [3H]-choline was measured for 20 minutes. ** p < 0.01, *** p < 0.001. (B) Control and M2 cells were pre-incubated with test compounds for 30 minutes and the uptake of 10 μM [14C]-ethanolamine was measured for 20 minutes.

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5.4.4 mRNA expression of CTL transporters and Kennedy pathway genes

As CTL1 mediated ethanolamine capacity was diminished in M2 cells, it made sense that the expression of genes that are dedicated to PE synthesis and degradation were altered in M2 cells. We analyzed the mRNA expression of CTL1-5 (Figure 5.4) and numerous genes in the Kennedy pathway. CTL1 mRNA expression was reduced by

80% in M2 cells while CTL2 expression was unchanged. CTL3, CTL4 and CTL5 mRNA content was not detected. The mRNA expression of the rate limiting enzyme for CDP- choline pathway mediated PC synthesis, Pcyt1, was decreased by 70% in M2 cells.

Moreover, the expression of Pcyt2, the rate limiting enzyme for CDP-ethanolamine pathway mediated PE synthesis, was decreased by 20% in M2 cells. The expression of choline kinase and ethanolamine kinase were decreased by 25% in M2 cells but the expression of all other Kennedy pathway genes was not affected (Figure 5.4).

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Figure 5.5: mRNA expression of CTL transporters and Kennedy pathway genes and Pcyt2 activity assay. (A) mRNA expression of CTL genes. The transcripts of CTL3, CTL4 and CTL5 were not detected.

(B) mRNA expression of genes devoted to phospholipid metabolism. (C) Pcyt2 enzyme activity assay. * <

0.05 relative to control cells.

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5.4.5 Effects of CTL1 overexpression on ethanolamine uptake

Control and M2 cells were transfected with a Myc-tagged SLC44A1 cDNA to further characterize the role of CTL1 in ethanolamine uptake. In Control cells, ethanolamine uptake increased by 25% 24 h after transfection (Figure 5.5) whereas ethanolamine uptake was increased in M2 cells by 50% 24 h after transfection (Figure 5.5). These data demonstrate that CTL1 overexpression significantly improves ethanolamine uptake in both Control and cells with CTL1 defects. In addition, M2 cells exhibit minimal CTL1 uptake capacity. Therefore, rescuing ethanolamine uptake in M2 cells provides conclusive evidence that CTL1 functions as an ethanolamine transporter.

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Figure 5.6: Effects of CTL1 overexpression on ethanolamine uptake. (A) CTL1 Western blot in

Control and M2 cells demonstrating the presence of the Myc tag. (B) Role of CTL1 overexpression on ethanolamine uptake in M2 cells. (C) Role of CTL1 overexpression on choline uptake in Control cells. (D)

Role of CTL1 overexpression on ethanolamine uptake in Control cells. * p < 0.05, ** p < 0.01, *** p <

0.001 relative to vehicle.

5.4.6 Effects of CTL2 knockdown on ethanolamine uptake

We measured uptake over a 120-minute period in Control and M2 cells to understand the dynamics of mitochondrial ethanolamine transport (Figure 5.6). The addition of HC-3

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blunted ethanolamine uptake in a time dependent manner in both cell lines. As M2 cells have negligible CTL1 uptake capacity, we knocked down CTL2 expression to further analyze ethanolamine uptake. Ethanolamine uptake was decreased by 50% after CTL2 knockdown, demonstrating the role of CTL2 in mitochondrial ethanolamine uptake.

Furthermore, we treated Control cells with 0-1000 μM ethanolamine after knocking down CTL2. Ethanolamine uptake was decreased when CTL2 was knocked down. In addition, the Eadie-Hofstee plots demonstrate that Control cells have 2 ethanolamine uptake mechanisms but the cells with CTL2 knocked down only had one ethanolamine uptake system in play.

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Figure 5.7: Characteristics of mitochondrial ethanolamine transport. (A) Western blots and PCR demonstrating adequate mitochondrial purity and CTL2 knockdown. (B) Effects of CTL2 knockdown on mitochondrial ethanolamine uptake. (C) Time course analysis of mitochondrial ethanolamine uptake in

Control cells treated with 20 µM HC-3. (D) Time course analysis of whole cell ethanolamine uptake in

Control cells treated with 20 µM HC-3. (E) Time course analysis of mitochondrial ethanolamine uptake in

M2 cells treated with 20 µM HC-3. (F) Time course analysis of whole cell ethanolamine uptake in Control cells treated with 20 µM HC-3. * p < 0.05, ** p < 0.01, *** p < 0.001.

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5.4.7 Lipid analysis in Control and M2 cells

To understand the influence on CTL1 on lipid content, we quantified membrane phospholipids (Figure 5.7) and neutral lipids (Figure 5.7). Membrane content of PC was not significantly different between Control and M2 cells. However, PE and PS content was decreased in M2 cells relative to Control. TAG content was increased whereas

DAG content was decreased in M2 relative to Control.

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Figure 5.8: Lipid analysis in Control and M2 cells. (A) PC levels were unchanged but PE and PS content was decreased in M2 cells relative to Control cells. (B) TAG content was increased whereas DAG content was decreased. * p < 0.05 relative to Control cells.

5.5. Discussion

Ethanolamine is vital for cellular function as it is an important component of the membrane phospholipid PE. Deficiencies in PE synthesis can lead to a lack of

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membrane integrity and subsequent metabolic disorders (255). As the mechanism of ethanolamine uptake had not been elucidated, it was of great importance to determine the main molecular system responsible for ethanolamine uptake.

In our study, we determined that CTL1 was largely responsible for ethanolamine uptake in human skin fibroblasts was pH dependent and partially Na+ dependent. Ethanolamine uptake was diminished by acidification of extracellular media, an indication that ethanolamine was likely transported by a choline/H+ antiporter system such as CTL1.

Additionally, there are two predominant ethanolamine uptake systems at play, as characterized by the Km values and Eadie-Hofstee plots of COS-7 and Control cells. We also showed that CTL1 mediated uptake was also sensitive to numerous organic cations, which has been demonstrated by numerous groups (152, 154, 263). Therefore,

CTL1 may be involved in the transport of these drugs but further studies would be needed to understand the role of CTL1 in drug transport.

With the CTL1 deficient M2 cells, we demonstrated that ethanolamine uptake can be rescued by overexpressing CTL1. This helps to clarify the role of CTL1 as an ethanolamine transporter. It has been suggested that choline and ethanolamine are transported via different systems. However, we show that choline and ethanolamine compete with one another for uptake. Moreover, choline and ethanolamine treatment in tandem further decreased the uptake of both substrates. While we used high concentrations of both substrates, the ability for ethanolamine to inhibit choline uptake has also been documented at physiological concentrations (28, 29).

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There was also the question of how the CTL1 deficient M2 cells take up ethanolamine.

By knocking down CTL2 expression in M2 cells, we observed that ethanolamine uptake was reduced by 50%, highlighting the importance of CTL2 in ethanolamine transport.

Moreover, CTL2 knockdown led to a 50% decrease in ethanolamine uptake in mitochondria. This is important to note as CTL2, not CTL1, seems to be the more prominent CTL protein present in mitochondria (253, 263). Ethanolamine plays a very important structural role in mitochondria, largely by serving as the headgroup of PE

(264, 265). The mitochondrion is the major organelle devoted to ATP production and they include structures with tight folds called cristae. The main goal of cristae is to maximize the amount of surface area on the inner mitochondrial membrane for ATP production to be as robust as possible. To maintain these folds, phospholipids which induce negative curvature, such as PE, need to be abundant. Moreover, PE has demonstrated to be important for stabilizing proteins of the electron transport chain which drive ATP synthesis (18, 19, 42).

We found that HC-3 and CTL1 antibody treatment led to widespread decreases in PC and PE production in MCF-7 and MCF-10 cells. This was intriguing because CTL1 has been traditionally thought to facilitate choline uptake for subsequent flow through the

CDP-choline branch of the Kennedy pathway.

To our knowledge, this was the first report delineating the roles of CTL1 and CTL2 in ethanolamine uptake. Therefore, it was important to verify the function of CTL1 and

CTL2 in ethanolamine uptake by overexpressing CTL1 and knocking down CTL2 expression.

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In conclusion, we investigated ethanolamine transports systems in human skin fibroblasts. Our studies demonstrate that ethanolamine uptake occurs through the CTL1 and CTL2 transporters in a pH and Na+ dependent manner. Moreover, we demonstrated that CTL2 facilitates ethanolamine uptake in mitochondria. Together, our data have highlighted the uptake mechanisms of ethanolamine which had not been clearly elucidated in human systems.

Study design: A. Taylor, M. Bakovic, Experimental work: A. Taylor, S. Grapentine, J.

Ichhpuniani, Manuscript preparation: A. Taylor, S. Grapentine, M. Bakovic.

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CHAPTER 6: INTEGRATIVE DISCUSSION

The importance of choline in skeletal muscle lipid homeostasis and neurodegenerative disease have not been extensively investigated to date. Additionally, the transport mechanism of ethanolamine in humans has not been elucidated so far. This thesis examined the relationships between choline, ethanolamine, CTL1/SLC44A1 and lipid metabolism in various states of choline deficiency and metabolic diseases. First of all, choline uptake and the CDP-choline pathway were analyzed in Pcyt2 deficient mice

(Chapter 3). Secondly, choline metabolism and lipid homeostasis were characterized in skin fibroblasts of two patients with frameshift mutations in SLC44A1 that also suffer from Neurodegeneration with Brain Iron Accumulation (NBIA) (Chapter 4). Third,

CTL1/SLC44A1 mediated ethanolamine uptake was characterized in skin fibroblasts

(Chapter 5). These studies demonstrated the benefits of choline supplementation on improving skeletal muscle lipid metabolism (Chapter 3), NBIA (Chapter 4) and the role of CTL1/SLC44A1 in ethanolamine uptake (Chapter 5).

6.1 Choline treatment in regulating lipid metabolism

As demonstrated in Chapters 3 and 4, choline supplementation can influence energy homeostasis as well as glycerolipid and fatty acid metabolism. We showed the effects of choline supplementation in two different studies: the CTL1 deficient M1 and M2

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fibroblasts which have perturbed choline uptake and the Pcyt2+/- mice which have diminished substrate flow through the ethanolamine branch of the Kennedy pathway.

Choline supplementation of M1 and M2 fibroblasts stimulated the increase of choline uptake in M1 fibroblasts by elevating CTL1/SLC44A1 expression. 28 days of choline treatment also ameliorated the obese phenotype of the Pcyt2+/- mice (76). In both studies, choline supplementation decreased the expression of genes involved in lipogenesis while increasing the expression of genes devoted to fatty acid oxidation. In

M1 and M2 fibroblasts, choline treatment diminished the sparing effect of PE for PC production. Choline supplementation improved lipid metabolism by altering the expression of genes involved with lipogenesis and fatty acid oxidation, which was similar to the findings of others (57, 58, 76, 80). The mechanisms by which choline treatment improves lipid homeostasis are likely interrelated. The expression and activation of key regulatory factors like SREBP1c, AMPK, PPARα and PGC-1α was modified by choline treatment and restored the expression of key lipogenic and lipolytic genes (57, 58). Moreover, choline is important for regulating epigenetic changes which can also play a role in modifying the expression of genes involved in obesity (91, 134,

175). During conditions such as obesity in Pcyt2+/- mice or choline uptake deficiency, supplemental choline has a protective effect by diminishing excessive TAG accumulation and preventing the overload of toxic intermediates like DAG.

There were a number of limitations with the study in Chapter 3. First off, choline treatment in Pcyt2+/+ mice was not examined as the goal of this study was to study the beneficial effects of a dietary intervention on a diseased condition rather than study the

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effects of a dietary intervention within a healthy population. That being said, a lack of study on choline supplementation in Pcyt2+/- mice fails to provide a complete picture of the impact of choline on physiological function. Moreover, there was a lack of functional analyses conducted in relation to the choline intervention. As an example, ITT and GTT analyses were not conducted which would have been a good complement to the quantification of key molecular markers involved with insulin signaling. Additionally, a key component of this study’s narrative was TAG accumulation within skeletal muscle but there were no histological analyses performed for a visual representation of skeletal muscle in these mice. Lastly, there was no specification between muscle fiber types (ie. slow twitch vs. fast twitch) which should be considered for future studies as oxidative function greatly differs (266).

6.2 Consequences of CTL1/SLC44A1 deficiency

We demonstrated that insufficient choline levels have widespread effects on cellular function. Chapter 3 described how diminished CTL1 content negatively impacted TAG metabolism and insulin signaling. Chapter 4 outlined the M1 and M2 fibroblasts who have unaffected PC levels but diminished CTL1 content and cellular choline uptake capacity. This was an extremely rare case whereby choline uptake deficiency was caused by a marked reduction and function of CTL1. However, the M1 and M2 fibroblasts were not deficient in PC but had reduced PE and PS relative to control fibroblasts. TAG content was also increased in M1 and M2 cells, which was a similar

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result to what was observed in Pcyt2+/- mice from Chapter 3. In addition, previous studies have demonstrated that both reduced substrate flow through the Kennedy pathway and diminished choline uptake are linked to an accumulation of TAG.

CTL1 deficiency has detrimental effects on cell membrane and cell organelle configuration. In Chapter 4, we noted that the M1 and M2 fibroblasts had an accumulation of abnormal materials, leading to an increased number of lysosomes and vacuoles. The function of these organelles was likely disrupted, and this has previously been associated with CTL1 function (11, 243). Moreover, these patients have are believed to have NBIA, which is characterized by neurodegeneration and cognitive decline. NBIA has been associated with perturbations in phospholipid metabolism and compromised CTL1 function likely also plays a role.

With regards to the study in Chapter 4, there were a number of parameters that could have been addressed. First off, there were no visual localization experiments about

CTL1 localization within the plasma membrane, which would have been beneficial in a study concerning CTL1 sufficiency. Moreover, part of the narrative of this chapter was intracellular iron accumulation but there were no analyses of proteins dedicated to iron uptake or iron storage (ie. ferritin, IRP2, TfR1) included in this work. Additionally, while the EM data can provide insight about the intracellular organization within Control, M1 and M2 cells, there were no images to document how chronic choline treatment helped improve intracellular configuration. Also, neurological cells such as iPSC derived cortical neurons were not utilized in this study, which would likely have been a relevant cell type to study as the patients have iron accumulation in the brain. Conducting studies with a

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type of neurological cells would have generated data that is more relevant to cerebral cell physiology. This is largely because there are likely more drastic cell specific differences that need to be accounted for when comparing skin fibroblasts to neurological tissue. Lastly, it would have been beneficial to utilize a more sensitive technique for detecting trace metals such as IPC-MS as opposed to the ferrozine assay

(267).

6.3 Regulation of ethanolamine transport by CTL1/SLC44A1

In addition to choline, we demonstrated that CTL1 was important for ethanolamine uptake. The CTL1 uptake deficits in M1 and M2 fibroblasts highlighted in Chapters 3 and 4 helped show that ethanolamine uptake was also compromised. Through uptake kinetics analyses, ethanolamine uptake in M1 and M2 fibroblasts occurred via one transporter whereas two transport systems were responsible for ethanolamine in control and COS-7 cells. As a result, CTL1 was likely heavily involved in ethanolamine transport because the M1 and M2 fibroblasts have deficiencies in CTL1. Moreover, ethanolamine uptake was more strongly affected in control fibroblasts treated with HC-3 relative to M2 cells. This is evidence of CTL1 being implicated in ethanolamine uptake as HC-3 has a strong capacity to inhibit CTL1 function. We also demonstrated that

CTL1 overexpression improved both choline and ethanolamine uptake in control and

M2 cells, further evidence of CTL1’s role in ethanolamine uptake. Overall, CTL1 demonstrates ethanolamine uptake capacity, which was an important finding as

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ethanolamine is implicated in numerous biological processes from mitochondrial biogenesis to cell division and apoptosis.

There were a number of additional analyses that could have been conducted to fortify the study in Chapter 5. To begin, a dose-response analysis of ethanolamine relative to anti-CTL1 treatment would have been beneficial to further solidify the role of CTL1 in ethanolamine uptake. Furthermore, pulse and pulse-chase analyses could have been conducted in conjunction with the transfection experiments. By studying metabolic flux through the CDP-ethanolamine branch of the Kennedy pathway, this would have strengthened the narrative of CTL1 facilitating ethanolamine uptake for the purposes of ethanolamine synthesis. Lastly, the rationale for the increased inhibitory effect of HC-3 on mitochondrial ethanolamine uptake relative to whole cell ethanolamine uptake was not explored. Studies to examine the molecular species (ie. fatty acids, proteins) within the plasma membrane and the mitochondrial membranes that help CTL1 and CTL2 maintain optimal topological configuration can provide insight about the key players that allow the transporters to function.

6.4 Integration of CDP-choline and CDP-ethanolamine pathways

We demonstrated that the CDP-choline and the CDP-ethanolamine pathways are important for regulating lipid glycerolipid homeostasis and preventing lipid accumulation.

Moreover, we showed how the CDP-choline pathway functions when substrate flow through the CDP-ethanolamine pathway was hindered using Pcyt2+/- mice (Chapter 3).

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Phospholipid levels (PC, PE, PS) in Pcyt2+/- mice do not significantly change because of reduced production and degradation of PE. These Pcyt2+/- mice accumulate TAG throughout the body because there was a decreased ability for DAG to be utilized for

PE synthesis and is more readily esterified to TAG (76, 191). We have demonstrated that supplemental choline treatment increases substrate flow through the choline branch of the Kennedy pathway. As a result, fatty acid synthesis and TAG accumulation were reduced because of increased DAG incorporation into the Kennedy pathway. The stimulation of the choline branch of the Kennedy pathway during ethanolamine branch impairment demonstrates the importance of both Kennedy pathway branches in reducing TAG accumulation.

In M1 and M2 fibroblasts, which have severe disruptions with CTL1 mediated choline uptake, we observed a sparing effect whereby PE was utilized as a substrate for PC synthesis. To our surprise, we observed PEMT enzyme activity whereby PC was

14 synthesized by S-adenosylmethionine methylation of PE. With [ C-CH3]-methionine labeling, we saw increased PC and PE content in M1 and M2 fibroblasts but this was decreased with choline supplementation. In addition, we saw that these cells have reduced substrate flow through the choline branch of the Kennedy pathway. As a result, there was increased expression of genes devoted to lipogenesis with decreased expression of fatty acid oxidation genes. This was likely because of decreased DAG integration into the Kennedy pathway due to diminished substrate flow through the choline branch.

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6.5 Conclusion and future directions

The findings presented in this thesis provide knowledge to better understand the role of choline in muscle lipid homeostasis and neurodegenerative disorders while also providing insight on the ethanolamine uptake mechanism in humans. Overall, phospholipid homeostasis and glycerolipid homeostasis were affected when Kennedy pathway substrate flow and CTL1 mediated substrate uptake are disrupted. However, supplemental choline modified both TAG and phospholipid homeostasis, demonstrated that additional choline can be beneficial in ameliorating the adverse effects of obesity and neurodegenerative disease (76). Additionally, this thesis also demonstrated the interplay between the CDP-choline and CDP-ethanolamine branches of the Kennedy pathway in maintaining phospholipid homeostasis.

In addition, the implication of CTL1 in both choline and ethanolamine uptake was also established. Ethanolamine uptake was severely impacted by the abolishment of CTL1, as demonstrated in M2 fibroblasts. Moreover, PE appeared to have a sparing effect on

PC in fibroblasts deficient in CTL1 activity, likely meaning that the causes of PE deficiency were two-fold: PE methylation to generate PC and decreased ethanolamine being taken up to be utilized for PE synthesis. Furthermore, the individuals with diminished CTL1 activity demonstrated perturbations in organelle architecture and function. As a result, it would be interesting to see the influence of choline and ethanolamine treatment on neurological disorders characterized by organelle dysfunction.

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There are numerous directions that be explored based on this thesis. The study in

Chapter 3 concerns choline supplementation with respect to metabolic function and composition of skeletal muscle. Moreover, there have been numerous observational studies and clinical trials conducted in humans which dictate that plasma choline levels are correlated with better body composition (199) and fitness level (268). However, choline is implicated in athletic performance (269) and contractile function (270) within skeletal muscle but there is a lack of studies which focus on dietary choline relative to these functions.

Another research area to explore would be the involvement of creatine in high-energy phosphate dependent substrate cycling during thermogenesis. Kazak et al. (2015) (271) conducted a mitochondrial proteomics analysis and determined that phosphatase activity is likely elevated for purposes of creatine cycling when ADP is limiting.

Moreover, PHOSPHO1 (a phosphatase with high activity towards phosphocholine and phosphoethanolamine) was identified as a candidate for involvement in creatine substrate cycling. As a result, it would be interesting to explore the influence of phosphocholine and phosphoethanolamime supplementation on thermogenesis as thermogenic activity has been shown to combat obesity (272, 273, 274).

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REFERENCES

(1) Awwad, H.M., Ohlmann, C.H., Stoeckle, M., Aziz, R., Geisel, J. and Obeid, R. 2016. Choline-phospholipids inter-conversion is altered in elderly patients with prostate cancer. Biochimie, 126, pp. 108-114.

(2) Cho, E., Zeisel, S.H., Jacques, P., Selhub, J., Dougherty, L., Colditz, G.A. and Willett, W.C. Dietary choline and betaine assessed by food-frequency questionnaire in relation to plasma total homocysteine concentration in the Framingham Offspring Study. 2006. American Journal of Clinical Nutrition. 83(4), pp. 905-911.

(3) Bidulescu, A., Chambliss, L.E., Siega-Ruiz, A.M., Zeisel, S.H., Heiss, G. 2007. Usual choline and betaine dietary intake and incident coronary heart disease: the Atherosclerosis Risk in Communities (ARIC) study. BMC Cardiovascular Disorders. 7(20), pp. 1-8.

(4) Bidulescu, A., Chambliss, L.E., Siega-Ruiz, A.M., Zeisel, S.H., Heiss, G. 2009. Repeatability and measurement error in the assessment of choline and betaine dietary intake: the Atherosclerosis Risk in Communities (ARIC) study. Nutrition Journal. 8(14), pp. 1-6.

(5) Cho, E., Willett, W.C., Colditz, G.A., Fuchs, C.S., Wu, K., Chan, A.T., Zeisel, S.H. and Giovannucci, E.L. 2007. Dietary choline and betaine and the risk of distal colorectal adenoma in women. Journal of the National Cancer Institute. 99(16), pp. 1224-1231.

(6) Richard, C., Lewis, E.D., Zhao, Y.Y., Asomaning, J., Jacobs, R.L., Field, C.J. and Curtis, J.M. 2016. Measurement of the total choline content in 48 commercial dairy products or dairy alternatives. Journal of Food Composition and Analysis. 45, pp. 1-8.

(7) Asomaning, J., Zhao, Y.Y., Lewis, E.D., Wu, J., Jacobs, R.L., Field, C.J. and Curtis, J.M. 2017. The development of a choline rich cereal based functional food: Effect of processing and storage. LWT – Food Science and Technology. 75, pp. 447-425.

(8) de Veth, M.J., Artegoitia, V.M., Campagna, S.R., Lapierre, H., Harte, F. and Girard, C.L. 2016. Choline absorption and evolution of bioavailability markers when supplementing choline to lactating dairy cows. Journal of Dairy Science. 99(12), pp. 9732-9744.

(9) Savoini, G., Agazzi, A., Invernizzi, G., Cattaneo, D., Pinotti, L. and Baldi, A. 2010. Polyunsaturated fatty acids and choline in dairy goats nutrition: Production and health benefits. Small Ruminant Research. 88, pp. 135-144.

(10) Mehta, R., Birendinc, A. and Younossi, Z.M. 2014. Host Genetic Variants in Obesity-Related Nonalcoholic Fatty Liver Disease. Clinics in Liver Disease. 18(1), pp. 249-267.

134

135

(11) Gao, X., Wang, Y. and Sun, G. 2017. High dietary choline and betaine intake is associated with low insulin resistance in the Newfoundland population. Nutrition. 33, pp. 28-34.

(12) Michel, V., Singh, R.K. and Bakovic, M. 2011. The impact of choline availability on muscle lipid metabolism. Food & Function. 2, pp. 53-62.

(13) Rinella, M.E. and Green, R.M. 2004. The methionine-choline deficient dietary model of steatohepatitis does not exhibit insulin resistance. Journal of Hepatology. 40(1), pp. 47-51.

(14) Payne, F., Lim, K., Girousse, A., Brown, R.J., Kory, N., Robbins, A., Xue, Y., Sleigh, A., Cochran, E., Adams, C., Borman, A.D., Russel-Jones, D., Gorden, P., Semple, R.K., Saudek, V., O'Rahilly, S., Walther, T.C., Barroso, I. and Savage, D.B. 2014. Mutations disrupting the Kennedy phosphatidylcholine pathway in humans with congenital lipodystrophy and fatty liver disease. Proceedings of the National Academy of Sciences of the United States of America. 111(24), pp. 8901-8906.

(15) Velazquez, R., Ash, J.A., Powers, B.E., Kelley, C.M., Strawderman, M., Luscher, Z.I., Ginsberg, S.D., Mufson, E.J. and Strupp, B.J. 2013. Maternal choline supplementation improves spatial learning and adult hippocampal neurogenesis in the Ts65Dn mouse model of Down syndrome. Neurobiology of Disease. 58, pp. 92-101.

(16) Lipton, BA., Yorek, MA. and Ginsberg, BH. 1988. Ethanolamine and choline transport in cultured bovine aortic endothelial cells. Journal of Cell Physiology. 137(3), pp. 571-576.

(17) Wishart, DS., Tzur, D., Knox, C., Eisner, R., Guo, AC., Young, N., Cheng, D., Jewell, K., Arndt, D., Sawhney, S., Fung, C., Nikolai, L., Lewis, M., Coutouly, MA., Forsythe, I., Tang, P., Shrivastava, S., Jeroncic, K., Stothard, P., Amegbey, G., Block, D., Hau, DD., Wagner, J., Miniaci, J., Clements, M., Gebremedhin, M., Guo, N., Zhang, Y., Duggan, GE., MacInnis GD., Weljie, AM., Dowlatabadi, R., Bamforth, F., Clive, D. Greiner, R., Li, L., Marrie, T., Sykes, BD., Vogel, HJ. and Querengesser, L. 2007. HMDB: the Human Metabolome Database. Nucleic Acids Research. 35, D521-D526.

(18) Heden, TD., Neufer, PD. and Funai, K. 2016. Looking Beyond Structure: Membrane Phospholipids of Skeletal Muscle Mitochondria. Trends in Endocrinology & Metabolism. 27(8), pp. 553-562.

(19) Ikon, N. and Ryan, RO. 2017. Cardiolipin and mitochondrial cristae organization. Biochimica et Biophysica Acta. 1859(6), pp. 1156-1163.

(20) Kano-Sueoka, T., Watanabe, T., Miya, T. and Kasai, H. 1991. Analysis of Cytosolic Phosphoethanolamine and Ethanolamine and Their Correlation with Prognostic Factors in Breast Cancer. Japanese Journal of Cancer Research. 82(7), pp. 829-834.

135

136

(21) Modica-Napolitano, JS. and Renshaw, PF. 2004. Ethanolamine and phosphoethanolamine inhibit mitochondrial function in vitro: implications for mitochondrial dysfunction hypothesis in depression and bipolar disorder. Biological Psychiatry. 55(3), pp. 273-277.

(22) Sasaki, H., Kume, H., Nemoto, A., Narisawa, S. and Takahashi, N. 1997. Ethanolamine modulates the rate of rat hepatocyte proliferation in vitro and in vivo. Proceedings of the National Academy of Sciences of the United States of America. 94(14), pp. 7320-7325.

(23) Garsin, DA. 2010. Ethanolamine utilization in bacterial pathogens: roles and regulation. Nature Reviews Microbiology. 8(4), pp. 290-295.

(24) Shin, KO. and Lee, YM. 2016. Simultaneous analysis of mono-, di-, and tri- ethanolamine in cosmetic products using liquid chromatography coupled tandem mass spectrometry. Archives of Pharmacal Research. 39(1), pp. 66-72

(25) Niculescu, M.D. and Zeisel, S.H. 2002. Diet, Methyl Donors and DNA Methylation: Interactions between Dietary Folate, Methionine and Choline. The Journal of Nutrition. 132, pp. 2333S-2335S.

(26) Vance, J.E. and Vance, D.E. 1986. Specific Pools of Phospholipids Are Used for Lipoprotein Secretion by Cultured Rat Hepatocytes. The Journal of Biological Chemistry. 261(10), pp. 4486-4491.

(27) Machova, E., O’Regan, S., Newcombe, J., Meunier, FM., Prentice, J., Dove, R., Lisa, V. and Dolezal, V. 2009. Detection of choline transporter-like 1 protein CTL1 in neuroblastoma x glioma cells and in the CNS, and its role in choline uptake. Journal of Neurochemistry. 110(4), pp. 1297-1309.

(28) Zelinski, TA. and Choy, PC. 1984. Ethanolamine inhibits choline uptake in the isolated hamster heart. Biochimica et Biophysica Acta. 794(2), pp. 326-332.

(29) Zha, X., Jay FT. and Choy, PC. 1992. Effects of amino acids and ethanolamine on choline uptake and phosphatidylcholine biosynthesis in baby hamster kidney-21 cells. Biochemistry and Cell Biology. 70(12), pp. 1319-1324.

(30) Ridgway, N.D. and Vance, D.E. 1987. Purification of Phosphatidylethanolamine N- methyltransferase from Rat Liver. The Journal of Biological Chemistry. 262(35), pp. 17231-17239.

(31) Vance, D.E. 2014. Phospholipid methylation in mammals: from biochemistry to physiological function. Biochimica et Biophysica Acta. 1838(6), pp. 1477-87.

(32) Nishimaki-Mogami, T., Yao, Z. and Fujimori, K. 2002. Inhibition of phosphatidylcholine synthesis via the phosphatidylethanolamine methylation pathway 136

137

impairs incorporation of bulk lipids into VLDL in cultured rat hepatocytes. Journal of Lipid Research. 43, pp. 1035-1045.

(33) Jiang, X., West, A.A. and Caudill, M. 2014. Maternal choline supplementation: a nutritional approach for improving offspring health? Trends in Endocrinology and Metabolism. 25(5), pp. 263-273.

(34) Watkins, S.M., Zhu, X. and Zeisel, S.H. 2003. Phosphatidylethanolamine-N- methyltransferase activity and dietary choline regulate liver-plasma lipid flux and essential fatty acid metabolism in mice. Journal of Nutrition. 133(11), pp. 3386-3391.

(35) Lo Van, A., Sakayori, N., Hachem, M., Belkouch, M., Picq, M., Lagarde, M., Osumi, N. and Bernoud-Hubac, N. 2016. Mechanisms of DHA transport to the brain and potential therapy to neurodegenerative diseases. Biochimie. 130, pp. 163-167.

(36) Thies, F., Pillon, C., Moliere, P., Lagarde, M. and Lecerf, J. 1994. Preferential incorporation of sn-2 lysoPC DHA over unesterified DHA in the young rat brain. American Journal of Physiology. 267, pp. 1273-1279.

(37) Dorninger, F., Broode, A., Braverman, N.E., Moser, A.B., Just, W.W., Forss-Petter, S., Brugger, B. and Berger, J. 2015. Homeostasis of phospholipids – The level of phosphatidylethanolamine tightly adapts to changes in ethanolamine plasmalogens. Biochimica et Biophysica Acta. 1851(2), pp. 117-128.

(38) Braverman, N.E. and Moser, A.B. Functions of plasmalogen lipids in health and science. 2012. Biochimica et Biophysica Acta. 1822(9), pp. 1442-1452.

(39) Foster, D.A. 2013. Phosphatidic acid and lipid-sensing by mTOR. Trends in Endocrinology and Metabolism. 24(6), pp. 272-278.

(40) Schenkel, L.C., Singh, R.K., Michel, V., Zeisel, S.H., da Costa, K.A., Johnson, A.R., Mudd, H.S. and Bakovic, M. 2015. Mechanism of choline deficiency and membrane alteration in postural orthostatic tachycardia syndrome primary skin fibroblasts. The Journal of the Federation of American Societies for Experimental Biology. 29(5), pp. 1663-1675.

(41) Vance, J.E. and Tasseva, G. 2013. Formation and function of phosphatidylserine and phosphatidylethanolamine in mammalian cells. Biochimica et Biophysica Acta. 1831(3), pp. 543-554.

(42) Martensson, C.U., Doan, K.M. and Becker, T. 2016. Effects of lipids on mitochondrial functions. Biochimica et Biophysica Acta. 1862(1), pp. 102-113.

(43) Vance, J.E. 2014. MAM (mitochondria-associated membranes) in mammalian cells: Lipids and beyond. Biochimica et Biophysica Acta. 1841(4), pp. 595-609.

137

138

(44) Zhou, Y., Holmseth, S., Hua, R., Lehre, A.C., Olofsson, A.M., Poblete-Naredo, I., Kempson, S.A. and Danbolt, N.C. 2012. The betaine-GABA transporter (BGT1, slc6a12) is predominantly expressed in the liver and at lower levels in the kidneys and at the brain surface. American Journal of Physiology. 302(3), pp. 316-328.

(45) Anwar, S.L. and Lehmann, U. 2014. DNA methylation, microRNAs, and their crosstalk as potential biomarkers in hepatocellular carcinoma. World Journal of Gastroenterology. 20(24), pp. 7894-7913.

(46) Teng, Y.W., Mehedint, M., Garrow, T. and Zeisel, S. 2011. Deletion of betaine- homocysteine S-methyltransferase in mice perturbs choline and 1-carbon metabolism, resulting in fatty liver and hepatocellular carcinomas. The Journal of Biological Chemistry. 286(42), pp. 36258-36267.

(47) Anstee, Q.M. and Day, C.P. 2012. S-adenosylmethionine (SAMe) therapy in liver disease: A review of current evidence and clinical utility. Journal of Hepatology. 57(5), pp. 1097-1109.

(48) Noureddin, M., Mato, J.M. and Lu, S.C. 2015. Nonalcoholic fatty liver disease: Update on pathogenesis, diagnosis, treatment and the role of S-adenosylmethionine. Experimental Biology and Medicine. 240(6), pp. 809-820.

(49) Seko, Y., Sumida, Y., Tanaka, S., Mori, K., Taketani, H., Ishiba, H., Hara, T., Okajima, A., Umemara, A., Nishikawa, T., Tamaguchi, K., Moriguchi, M., Kanemasa, K., Yasui, K., Imai, S., Shimada, K. and Itoh, Y. 2018. Insulin resistance increases the risk of incident type 2 diabetes mellitus in patients with nonalcoholic fatty liver. Hepatology Research. 48(3), pp. 42-51.

(50) Toohey, J.I. 2014. Sulfur Amino Acids in Diet-induced Fatty Liver: A New Perspective Based on Recent Findings. Molecules. 19(6), pp. 8334-8349.

(51) Shim, P., Choi, D. and Park, Y. 2017. Association of Blood Fatty Acid Composition and Dietary Pattern with the Risk of Non-Alcoholic Fatty Liver Disease in Patients Who Underwent Cholecystectomy. Annals of Nutrition and Metabolism. 70(4), pp. 303-311.

(52) Zeisel, S.H. and Blusztajn, J.K. 1994. Choline and Human Nutrition. Annual Review of Nutrition. 14, pp. 269-296.

(53) Obeid, R. 2013. The Metabolic Burden of Methyl Donor Deficiency with Focus on the Betaine Homocysteine Methyltransferase Pathway. Nutrients. 5(9), pp. 3481-3495.

(54) Khaire, A., Rathod, R., Kale, A. and Joshi, S. 2015. Vitamin B12 and omega-3 fatty acids together regulate lipid metabolism in Wistar rats. Prostaglandins, Leukotrienes and Essential Fatty Acids. 99, pp. 7-17.

138

139

(55) Begley, H.N., Wang, Y., Campbell, M.S., Yu, X., Lane, R.H. and Joss-Moore, L.A. 2013. Maternal docosahexaenoic acid increases adiponectin and normalizes IUGR- induced changes in rat adipose deposition. Journal of Obesity. [Online] DOI: 10.1155/2013/312153

(56) Verbrugghe, A. and Bakovic, M. 2013. Peculiarities of One-Carbon Metabolism in the Strict Carnivorous Cat and the Role in Feline Hepatic Lipidosis. Nutrients. 5(7), pp. 2811-2835.

(57) Pooya, S., Blaise, S., Garcia, M.M., Giudicelli, J., Alberto, J.M., Gueant-Rodriguez, R.M., Jeannesson, E., Gueguen, N., Bressenot, A., Nicolas, B., Malthiery, Y., Daval, J.L., Peyrin-Biroulet, L., Bronowicki, J.P. and Gueant, J.L. 2012. deficiency impairs fatty acid oxidation through PGC-1α hypomethylation and decreased ER-α, ERR-α, and HNF-4α in the rat liver. Journal of Hepatology. 57, pp. 344-351.

(58) Dahlhoff, C., Worsch, S., Saller, M., Hummel, B.A., Fiamonici, J., Uebel, K., Obeid, R., Scherling, C., Geisel, J., Bader, B.L. and Daniel, H. 2014. Methyl-donor supplementation in obese mice prevents the progression of NAFLD, activates AMPK and decreases acyl-carnitine levels. Molecular Metabolism. 3(5), pp. 565-580.

(59) Schofield, Z., Reed, M.A.C., Newsome, P.N., Adams, D.H., Gunther, U.L. and Lalor, P.F. 2017. Changes in human hepatic metabolism in steatosis and cirrhosis. World Journal of Gastroenterology. 23(15), pp. 2685-2695.

(60) Auta, J., Zhang, H., Pandey, S.C. and Guidotti, A. 2017. Chronic Alcohol Exposure Differentially Alters One-Carbon Metabolism in Rat Liver and Brain. Alcoholism: Clinical and Experimental Research. 41(6), pp. 1105-1111.

(61) Liu, H.S., Lu, P., Guo, Y., Farrell, E., Zhang, X., Zheng, M., Bosano, B., Zhang, Z., Allard, J., Liao, G., Fu, S., Chen, J., Dolim, K., Kuroda, A., Usaka, J., Cheng, J., Tao, W., Welch, K., Liu, Y., Pease, J., de Keczer, S.A., Masjedizadeh, M., Hu, J.S., Weller, P., Garrow, T. and Peltz, G. 2010. An integrative genomic analysis identifies Bhmt2 as a diet-dependent genetic factor protecting against acetaminophen-induced liver toxicity. Genome Research. 20(1), pp. 28-35.

(62) Robinson, J.L. and Bertolo, R.F. 2016. The Pediatric Methionine Requirement Should Incorporate Remethylation Potential and Transmethylation Demands. Advances in Nutrition. 7(3), pp. 523-534.

(63) Cote-Robitaille, M.E., Girard, C.L., Guay, F. and Matte, J.J. 2015. Oral supplementations of betaine, choline, creatine and vitamin B6 and their influence on the development of homocysteinaemia in neonatal piglets. Journal of Nutritional Science. 4(31), pp. 1-7.

139

140

(64) Miettinen, T.P., Pessa, H.K.J., Caldez, M.J., Fuhrer, T., Diril, M.K., Sauer, U., Kaldis, P. and Bjorklund, M. 2014. Identification of Transcriptional and Metabolic Programs Related to Mammalian Cell Size. Current Biology. 24(6), pp. 598-608.

(65) Ginzberg, M.B., Kafri, R. and Kirschner, M. 2015. On being the right (cell) size. Science. 348(6236), pp. 1-19.

(66) Hoffmann, L., Brauers, G., Gehrmann, T., Haussinger, D., Mayatepek, E., Schless, F. and Schwahn, B. 2013. Osmotic regulation of hepatic betaine metabolism. Gastroenterology and Liver Physiology. 304(9), pp. G835-G846.

(67) Ahn, C.W., Choi, Y.J., Hong, S.H., Jun, D.S., Na, J.D., Choi, Y.J. and Kim, Y.C. 2015. Involvement of multiple pathways in the protection of liver against high-fat diet- induced steatosis by betaine. Journal of Functional Foods. 17, pp. 66-72.

(68) Than, N.N. and Newsome, P.N. 2015. A concise review of non-alcoholic fatty liver disease. Atherosclerosis. 239(1), pp. 192-202.

(69) Kathirvel, E., Morgan, K., Nandgiri, G., Sandoval, B.C., Caudill, M.A., Bottiglieri, T., French, S.W. and Morgan, T.R. 2010. Betaine improves nonalcoholic fatty liver and associated hepatic insulin resistance: a potential mechanism for hepatoprotection by betaine. American Journal of Physiology, Endocrinology Metabolism and Gastrointestinal Physiology. 299(5), pp. 1068-1077.

(70) Jager, S., Handschin, C., St-Pierre, J. and Spiegelman, B.M. 2007. AMP-activated protein kinase (AMPK) action in skeletal muscle via direct phosphorylation of PGC-1α. Proceedings of the National Academy of Sciences of the United States of America. 104(29), pp. 12017-12022.

(71) Buzzetti, E., Pinzani, M. and Tsochatzis, E.A. 2016. The multiple-hit pathogenesis of non-alcoholic fatty liver disease (NAFLD). Metabolism. 65(8), pp. 1038-1048.

(72) Hirsch, E., Costa, C. and Ciraolo, E. 2007. Phosphoinositide 3-kinases as a common platform for multi-hormone signaling. Journal of Endocrinology. 194(2), pp. 243-256.

(73) Longo, N., Frigeni, M. and Pasquali, M. 2016. Carnitine transport and fatty acid oxidation. Biochimica et Biophysica Acta. 1863(10), pp. 2422-2435.

(74) ter Veld, F., Primassin, S., Hoffmann, L., Mayatepek, E. and Spiecherkoetter, U. 2009. Corresponding increase in long-chain acyl-CoA and acylcarnitine after exercise in muscle from VLCAD mice. Journal of Lipid Research. 50(8), pp. 1556-1562.

(75) Neess, D., Bek, S., Engelsby, H., Gallego, S.F. and Faergeman, N.J. 2015. Long- chain acyl-CoA esters in metabolism and signaling: Role of acyl-CoA binding proteins. Progress in Lipid Research. 59, pp. 1-25. 140

141

(76) Taylor, A., Schenkel, L.C., Yokich, M. and Bakovic, M. 2016. Adaptations to excess choline in insulin resistant and ,deficient skeletal muscle. Biochemistry and Cell Biology. 95(2), pp. 223-231.

(77) Jiao, F., Yan, X., Yu, Y., Zhu, X., Ma, Y., Yue, Z., Ou, H. and Yan, Z. 2016. Protective Effects of Maternal Methyl Donor Supplementation on Adult Offspring of High-fat Diet-fed Dams. The Journal of Nutritional Biochemistry. 34, pp. 42-51.

(78) Thomas, DT., Schnell, DM., Redzic, M., Zhao, M., Abraha, H., Jones, D., Brim, H., Yu, G., 2019. Local In Vivo Measures of Muscle Lipid and Oxygen Consumption Change in Response to Combined Vitamin D Repletion and Aerobic Training in Older Adults. Nutrients. 11(4): pii: E930.

(79) Davis, HM., Essex, AL., Valdez, S., Deosthale, PJ., Aref, MW., Allen, MR., Bonetto, A., Plotkin, LI., 2019. Short-term pharmacologic RAGE inhibition differentially affects bone and skeletal muscle in middle-aged mice. Bone. 124: 89-102.

(80) Adelnia, F., Cameron, D., Bergeron, CM., Fishbein, KW., Spencer, RG., Reiter, DA., Ferrucci, L., 2019. The Role of Muscle Perfusion in the Age-Associated Decline of Mitochondrial Function in Healthy Individuals. Front Physiol. 10:427.

(81) Miyake, H., Kanazawa, I., Tanaka, KI., Sugimoto, T., 2019. Low skeletal muscle mass is associated with the risk of all-cause mortality in patients with type 2 diabetes mellitus. Ther Adv Endocrinol Metab. 10:2042018819842971.

(82) Hou, L., Lei, Y., Li, X., Huo, C., Jia, X., Yang, J., Xu, R., Wang, X., 2019. Effect of Protein Supplementation Combined with Resistance Training on Muscle Mass, Strength and Function in the Elderly: A Systematic Review and Meta-Analysis. J Nutr Health Aging. 23(5): 451-458.

(83) Chan, EK., Kornberg, AJ., Ryan, MM. 2015. A diagnostic approach to recurrent myalgia and rhabdomyolysis in children. Arch Dis Child. 100(8): 793-797.

(84) Ren, H., Federico, L., Huang, H., Sunkara, M., Drennan, T., Frohman, MA., Smyth, SS., Morris, AJ., 2010. A phosphatidic acid binding/nuclear localization motif determines lipin1 function in lipid metabolism and adipogenesis. Mol Biol Cell. 21(18): 3171-3181.

(85) Rashid, T., Nemazanyy, I., Paolini, C., Tatsuta, T., Crespin, P., de Villeneuve, D., Brodesser, S., Benit, P., Rustin, P., Baraibar, MA., Agbulut, O., Olivier, A., Protasi, F., Langer, T., Chrast, R., de Lonlay, P., de Foucoauld, H., Blaauw, B., Pende, M. 2019. Lipin deficiency causes sarcoplasmic reticulum stress and chaperone-responsive myopathy. 38(1): pii: e99576.

(86) Gerling, CJ., Mukai, K., Chabowski, A., Heigenhauser, GJF., Holloway, GP., Spriet, LL., Jannas-Vela, S., 2019. Incorporation of Omega-3 Fatty Acids Into Human Skeletal

141

142

Muscle Sarcolemmal and Mitochondrial Membranes Following 12 Weeks of Fish Oil Supplementation. Front Physiol. 10:348.

(87) Slodki, S., Bogucka, J., 2019. Mitochondrial Theory of Skeletal Muscle Ageing – New Facts, New Doubts. J Vet Res. 63(1): 149-160.

(88) Abrigo, J., Simon, F., Cabrera, D., Vilos, C., Cabello-Verrugio, C., 2019. Mitochondrial dysfunction in skeletal muscle pathologies. Curr Protein Pept Sci. doi: 10.2174/1389203720666190402100902.

(89) Pennington, ER., Funai, K., Brown, DA., Shaikh, SR., 2019. The role of cardiolipin concentration and acyl chain composition on mitochondrial inner membrane molecular organization and function. Biochim Biophys Acta Mol Cell Biol Lipids. 1864(7): 1039- 1052.

(90) Ren, M., Phoon, CK., Schlame, M., 2014. Metabolism and function of mitochondrial cardiolipin. Prog Lipid Res. 55: 1-16.

(91) Duncan, AL., Ruprecht, JJ., Kunji, ERS., Robinson, AJ., 2018. Cardiolipin dynamics and binding to conserved residues in the mitochondrial ADP/ATP carrier. Biochim Biophys Acta Biomembr. 1860(5): 1035-1045.

(92) Mejia, EM., Hatch, GM., 2016. Mitochondria phospholipids: role in mitochondrial function. J Bioenerg Biomembr. 48(2): 99-112.

(93) Raja, V., Reynolds, CA., Greenberg, ML. 2017. Barth syndrome: A life-threatening disorder caused by abnormal cardiolipin remodeling. J Rare Dis Res Treat. 2(2): 58-62.

(94) Lores-Arnaiz, S., Lombardi, P., Karadayian, AG., Cutrera, R., Bustamante, J., 2019. Changes in motor function and brain cortex mitochondrial active oxygen species production in aged mice. Exp Gerontol. 118: 88-98.

(95) Sullivan, EM., Pennington, ER., Sparagna, GC., Torres, MJ., Neufer, PD., Harris, M., Washington, J., Anderson, EJ., Zeczycki, TN., Brown, DA., Shaikh, SR. 2018. Docosahexaenoic acid lowers cardiac mitochondrial enzyme activity by replacing linoleic acid in the phospholipidome. J Biol Chem. 293(2): 466-483.

(96) Gonzalez-Friere, M., de Cabo, R., Bernier, M., Sollott, SJ., Fabbri, E., Navas, P., Ferrucci, L. 2015. Reconsidering the Role of Mitochondria in Aging. J Gerontol A Biol Sci Med Sci. 70(11): 1334-1342.

(97) Shiyo, YJ., Lupo, G., Vance, JE., 1995. Evidence that phosphatidylserine is imported into mitochondria via a mitochondria-associated membrane and that the majority of mitochondrial phosphatidylethanolamine is derived from decarboxylation of phosphatidylserine. J Biol Chem. 270(19): 11190-11198.

142

143

(98) King, J.H., Kwan, S.T., Yan, J., Klatt, K.C., Jiang, X., Roberson, M.S. and Caudill, M.A. 2017. Maternal Choline Supplementation Alters Fetal Growth Patterns in a Mouse Model of Placental Insufficiency. Nutrients. 9, pp. 1-16.

(99) Nam, J., Greenwald, E., Jack-Roberts, C., Ajeeb, T.T., Malysheva, O.V., Caudill, M.A., Axen, K., Saxena, A., Semernina, E., Nanobashvili, K. and Jiang, X. 2017. Choline prevents fetal overgrowth and normalizes placental fatty acid and glucose metabolism in a mouse model of maternal obesity. Journal of Nutritional Biochemistry. 49, pp. 80-88.

(100) Yan, J., Jiang, X., West, A.A., Perry, C.A., Malysheva, O.V., Devapatla, S., Pressman, E., Vermeylen, F., Stabler, S.P., Allen, R.H. and Caudill, M.A. 2012. Maternal choline intake modulates maternal and fetal biomarkers of choline metabolism in humans. The American Journal of Clinical Nutrition. 95(5), pp. 1060-1071.

(101) Yan, J., Jiang, X., West, A.A., Perry, C.A., Malysheva, O.V., Brenna, J.T., Stabler, S.P., Allen, R.H., Gregory, J.F. and Caudill, M.A. 2013. Pregnancy alters choline dynamics: results of a randomized trial using stable isotope methodology in pregnant and nonpregnant women. The American Journal of Clinical Nutrition. 98(6), pp. 1459- 1467.

(102) Molloy, A.M., Mills, J.L., Cox, C., Daly, S.F., Conley, M., Brody, L.C., Kirke, P.N., Scott, J.M. and Ueland, P.M. 2005. Choline and homocysteine interrelations in umbilical cord and maternal plasma at delivery. American Journal of Clinical Nutrition. 82(4), pp. 836-842.

(103) Velzing-Aarts, F.V., Holm, P.I., Fokkema, M.R., Van Der Dijs, F.P., Ueland, P.M. and Muskiet, F.A. 2005. Plasma choline and betaine and their relation to plasma homocysteine in normal pregnancy. American Journal of Clinical Nutrition. 81(6), pp. 1383-1389.

(104) Brosnan, J.T. and Brosnan, M.E. 2006. The Sulfur-Containing Amino Acids: An Overview. American Society for Nutrition. 136, pp. 1636-1640.

(105) Ganz, A.B., Cohen, V.V., Swersky, C.C., Stover, J., Vitiello, G.A., Lovesky, J., Chuang, J.C., Shields, K., Fomin, V.G., Lopez, Y.S., Mohan, S., Ganti, A., Carrier, B., Malysheva, O.V. and Caudill, M.A. 2017. Genetic Variation in Choline-Metabolizing Enzymes Alters Choline Metabolism in Young Women Consuming Choline Intakes Meeting Current Recommendations. International Journal of Molecular Sciences. 18, pp. 1-23.

(106) Jadavji, N.M., Deng, L., Malysheva, O., Caudill, M.A. and Rozen, R. 2015. MTHFR deficiency or reduced intake of folate or choline in pregnant mice results in

143

144

impaired short-term memory and increased apoptosis in the hippocampus of wild-type offspring. Neuroscience. 300, pp. 1-9.

(107) da Costa, K.A., Kozyreva, O.G., Song, J., Golanko, J.A., Fischer, L.M. and Zeisel, S.H. 2006. Common genetic polymorphisms affect the human requirement for the nutrient choline. The Journal of the Federation of American Societies for Experimental Biology. 20(9), pp. 1336-1344.

(108) Kohlmeier, M., da Costa, K.A., Fischer, L.M. and Zeisel, S.H. 2005. Genetic variation of folate-mediated one-carbon transfer pathway predicts susceptibility to choline deficiency in humans. Proceedings of the National Academy of Sciences. 102(44), pp. 16025-16030.

(109) Jensen, H.H., Batres-Marquez, S.P., Carriquiry, A. and Schalinske, K.L. 2007. Choline in the diets of the U.S. population: NHANES, 2003-2004. The Journal of the Federation of American Societies for Experimental Biology. 21, pp. 219.

(110) Provencal, N. and Binder, E.B. 2015. The effects of early life stress on the epigenome: From the womb to adulthood and even before. Experimental Neurology. 268, pp. 10-20.

(111) Smith, Z.D. and Meissner, A. 2013. DNA methylation: roles in mammalian development. Nature Reviews Genetics. 14(3), pp. 204-220.

(112) Schneider, E., Dittrich, M., Bock, J., Nanda, I., Muller, T., Seidmann, L., Tralau, T., Galetzka, D., El Hajj, N. and Haaf, T. 2016. CpG sites with continuously increasing or decreasing methylation from early to late human fetal brain development. Gene. 592(1), pp. 110-118.

(113) Hala, D., Huggett, D.B. and Burggren, W.W. 2014. Environmental stressors and the epigenome. Drug Discovery Today: Technologies. 12, pp. 3-8.

(114) Haggarty, P. 2012. Nutrition and the Epigenome. Progress in Molecular Biology and Translational Science. 108, pp. 427-446.

(115) Kovacheva, V.P., Mellott, T.J., Davison, J.M., Wagner, N., Lopez-Coviella, I., Schnitzler, A.C. and Blusztajn, J.K. 2007. Gestational Choline Deficiency Causes Global and Igf2 Gene DNA Hypermethylation by Up-regulation of Dnmt1 Expression. The Journal of Biological Chemistry. 282(43), pp. 31777-31888.

(116) Agaoglu, O.K., Agaoglu, A.R., Guzeloglu, A., Aslan, S., Kurar, E., Kayis, S.A. and Schafer-Somi, S. 2016. Gene expression profiles of some cytokines, growth factors, receptors, and enzymes (GM-CSF, IFNγ, MMP-2, IGF-II, EGF, TGF-β, IGF-IIR) during pregnancy in the cat uterus. Theriogenology. 85(4), pp. 638-644.

144

145

(117) Borodovitsyna, O., Flamini, M. and Chandler, D. 2017. Noradrenergic Modulation of Cognition in Health and Disease. Neural Plasticity. [Online]. [29 March 2018]. Available from: https://www.ncbi.nlm.nih.gov/pmc/articles/PMC5450174/pdf/NP2017- 6031478.pdf

(118) da Costa, K.A., Rai, K.S., Craciunescu, C.N., Parikh, K., Mehedint, M.G., Sanders, L.M., McLean-Pottinger, A. and Zeisel, S.H. 2009. Dietary Docosahexaenoic Acid Supplementation Modulates Hippocampal Development in the Pemt-/- Mouse. The Journal of Biological Chemistry. 285(2), pp. 1008-1015.

(119) da Costa, K.A., Sanders, L.M., Fischer, L.M. and Zeisel, S.H. 2011. Docosahexaenoic acid in plasma phosphatidylcholine may be a potential marker for in vivo phosphatidylethanolamine N-methyltransferase activity in humans. The American Journal of Clinical Nutrition. 93(5), pp. 968-974.

(120) Nyaradi, A., Li, J., Hickling, S., Foster, J. and Oddy, W.H. 2013. The role of nutrition in children's neurocognitive development, from pregnancy through childhood. Frontiers in Human Neuroscience. 7(97), pp. 1-16.

(121) Niculescu, M.D., Craciunescu, C.N. and Zeisel, S.H. 2006. Dietary choline deficiency alter global and gene-specific DNA methylation in the developing hippocampus of mouse fetal brains. The Journal of the Federation of American Societies for Experimental Biology. 20(1), pp. 43-49.

(122) Schulz, K.M., Pearson, J.N., Gasparrini, M.E., Brooks, K.F., Drake-Frazier, C., Zajkowski, M.E., Kreisler, A.D., Adams, C.E., Leonard, S. and Stevens, K.E. 2014. Dietary choline supplementation to dams during pregnancy and lactation mitigates the effects of in utero stress exposure on adult anxiety-related behaviors. Behavioural Brain Research. 268, pp. 104-110.

(123) Hancock, S.E., Freidrich, M.G., Mitchell, T.W., Truscott, R.J.W. and Else, P.L. 2017. The phospholipid composition of the human entorhinal cortex remains relatively stable over 80 years of adult aging. GeroScience. 39(1), pp. 73-82.

(124) Boughter, C.T., Monje-Galvan, V., Im, W. and Klauda, J.B. 2016. Influence of Cholesterol on Phospholipid Bilayer Structure and Dynamics. The Journal of Physical Chemistry. 120(45), pp. 11761-11772.

(125) Kolisnyk, B., Al-Onaizi, M.A., Hirata, P.H., Guzman, M.S., Nikolova, S., Barbash, S., Soreg, H., Bartha, R., Prado, M.A. and Prado, V.F. 2013. Forebrain deletion of the vesicular acetylcholine transporter results in deficits in executive function, metabolic, and RNA splicing abnormalities in the prefrontal cortex. The Journal of Neuroscience. 33(37), pp. 14908-14920.

145

146

(126) Wurtman, R.J. 2015. How Anticholinergic Drugs Might Promote Alzheimer's Disease: More Amyloid-β and Less Phosphatidylcholine. Journal of Alzheimer's Disease. 46(4), pp. 983-987.

(127) Caudill, M.A., Strupp, B.J., Muscalu, L., Nevins, J.E.H. and Canfield, R.L. 2018. Maternal choline supplementation during the third trimester of pregnancy improves infant information processing speed: a randomized, double-blind, controlled feeding study. The Journal of the Federation of American Societies for Experimental Biology. 32(4), pp. 2172-2180.

(128) Zhu, C., Wu, T., Jin, Y., Huang, B., Zhou, R., Wang, Y., Luo, X. and Zhu, H. 2016. Prenatal choline supplementation attenuates spatial learning deficits of offspring rats exposed to low-protein diet during fetal period. Journal of Nutritional Biochemistry. 32, pp. 163-170.

(129) Craciunescu, C.N., Albright, C.D., Mar, M.H., Song, J. and Zeisel, S.H. 2003. Choline Availability During Embryonic Development Alters Progenitor Cell Mitosis in Developing Mouse Hippocampus. Journal of Nutrition. 133(11), pp. 3614-3618.

(130) Kwan S.T., King, J.H., Yan, J., Wang, Z., Jiang, X., Hutzler, J.S., Klein, H.R., Brenna, J.T., Roberson, R.S. and Caudill, M.A. 2017. Maternal Choline Supplementation Modulates Placental Nutrient Transport and Metabolism in Late Gestation of Mouse Pregnancy. Journal of Nutrition. 147(11), pp. 2083-2092.

(131) DeLong, C.J., Shen, Y.J., Thomas, M.J. and Cui, Z. 1999. Molecular Distinction of Phosphatidylcholine Synthesis between the CDP-Choline Pathway and Phosphatidylethanolamine Methylation Pathway. Journal of Nutritional Biochemistry. 274(42), pp. 29683-29688.

(132) Resseguie, M., Song, J., Niculescu, M.D., da Costa, K.A., Randall, T.A. and Zeisel, S.H. 2007. Phosphatidylethanolamine N-methyltransferase (PEMT) gene expression is induced by estrogen in human and mouse primary hepatocytes. The Journal of the Federation of American Societies for Experimental Biology. 21(10), pp. 2622-2632.

(133) Fischer, L.M., da Costa, K.A., Kwock, L., Stewart, P.W., Lu, T.S., Stabler, S.P., Allen, R.H. and Zeisel, S.H. 2017. Sex and menopausal status influence human dietary requirements for the nutrient choline. The American Journal of Clinical Nutrition. 85(5), pp. 1275-1285.

(134) Sundrani, D., Khot, V. and Joshi, S. 2016. DNA Methylation for Prediction of Adverse Pregnancy Outcomes. In: Epigenetic Biomarkers and Diagnostics. San Diego, California, United States: Academic Press. pp. 351-376.

146

147

(135) Kulharni, A., Dangat, K., Kale, A., Sable, P., Chavan-Gautam, P. and Joshi, S. 2011. Effects of Altered Maternal Folic Acid, Vitamin B12 and Docosahexaenoic Acid on Placental Global DNA Methylation Patterns in Wistar Rats. PLoS ONE. 6, e17706.

(136) Bahous, RH., Cosin-Tomas, M., Deng, L., Leclerc, D., Malysheva, O., Ho, MK., Pallas, M., Kaliman, P., Bedell, BJ., Caudill, MA. And Rozen, R. 2018. Early Manifestations of Brain Aging in Mice Due to Low Dietary Folate and Mild MTHFR Deficiency. Molecular Neurobiology. [Epub ahead of print].

(137) Dorninger, F., Moser, AB., Kou, J., Weisinger, C., Forss-Petter, S., Gleiss, A., Hinterberger, M., Jungwirth, S., Fischer, P. and Berger, J. 2018. Alterations in the Plasma Levels of Specific Choline Phospholipids in Alzheimer’s Disease Mimic Accelerated Aging. Journal of Alzheimer’s Disease. 62(2), pp. 841-854.

(138) Lin, W., Zhang, J., Liu, Y., Wu, R., Yang, H., Hu, X. and Ling, X. 2017. Studies on diagnostic biomarkers and therapeutic mechanism of Alzheimer’s disease through metabolomics and hippocampal proteomics. European Journal of Pharmaceutical Sciences. 15(105), pp. 119-126.

(139) van Wijk, N., Slot, RER., Duits, FH., Strik, M., Biesheuvel, E., Sijben, JWC., Blankenstein, MA., Bierau, J., van der Flier, WM., Scheltens, P. and Teunissen, CE. 2017. Nutrients required for phospholipid synthesis are lower in blood and cerebrospinal fluid in mild cognitive impairment and Alzheimer's disease dementia. Alzheimer’s and Dementia.16(8), pp. 139-146.

(140) Mellott, TJ., Huleatt, OM., Shade, BN., Pender, SM., Liu, YB., Slack, BE. and Blusztajn, JK. 2017. Perinatal Choline Supplementation Reduces Amyloidosis and Increases Choline Acetyltransferase Expression in the Hippocampus of the APPswePS1dE9 Alzheimer's Disease Model Mice. PLoS One. 12(1), e0170450.

(141) Nickerson, CA., Brown, AL., Yu, W., Chun, Y. and Glenn, MJ. 2017. Prenatal choline supplementation attenuates MK-801-induced deficits in memory, motor function, and hippocampal plasticity in adult male rats. Neuroscience. 361, pp.116-128.

(142) Alldred, MJ., Chao, HM., Lee, SH., Beilin J., Powers, BE., Petkova, E., Strupp, BJ. and Ginsberg, SD. 2018. CA1 pyramidal neuron gene expression mosaics in the Ts65Dn murine model of Down syndrome and Alzheimer's disease following maternal choline supplementation. Hippocampus. 28(4), pp. 251-268.

(143) Powers, BE., Kelley, CM., Velazquez, R., Ash, JA., Strawderman, MS., Alldred, MJ., Ginsberg, SD., Mufson, EJ. and Strupp, BJ. 2017. Maternal choline supplementation in a mouse model of Down syndrome: Effects on attention and nucleus basalis/substantia innominata neuron morphology in adult offspring. Neuroscience. 340, pp. 501-514.

147

148

(144) Iulia, C., Ruxandra, T., Costin, LB. and Liliana-Mary, V. 2017. Citicoline - a neuroprotector with proven effects on glaucomatous disease. Romanian Journal of Ophthalmology. 61(3), pp. 152-158.

(145) Roberti, G., Tanga, L., Michelessi, M., Quaranta, L., Parisi, V., Manni, G. and Oddone, F. 2015. Cytidine 5'-Diphosphocholine (Citicoline) in Glaucoma: Rationale of Its Use, Current Evidence and Future Perspectives. International Journal of Molecular Sciences. 16(12), pp. 28401-28417.

(146) Thies, F., Delachambre, MC., Bentejac, M., Lagarde, M., Lecerf, J., 1992. Unsaturated Fatty Acids Esterified in 2-Acyl-1-Lysophosphatifylcholine Bound to Albumin Are More Efficiency Taken up by the Young Rat Brain than the Unesterifed Form. J Neurochem. 59(3): 1110-1116.

(147) Lagarde, M., Hachem, M., Bernoud-Hubac, N., Picq, M., Vericel, E., Guichardant, M. 2015. Biological properties of a DHA-containing structured phospholipid (AceDoPC) to target the brain. Prostaglandins Leukot Essent Fatty Acids. 92: 63-65.

(148) Lo Van, A., Sakayori, N., Hachem, M., Belkouch, M., Picq, M., Lagarde, M., Osumi, N., Bernoud-Hubac, N., 2016. Mechanisms of DHA transport to the brain and potential therapy to neurodegenerative diseases. Biochimie. 130: 163-167.

(149) Tamura, A., Tanaka, T., Yamane, T., Nasu, R., Fujii, T., 1985. Quantitative studies on translocation and metabolic conversion of lysophosphatidylcholine incorporated into the membrane of intact human erythrocytes from the medium. J Biochem. 97(1): 353-9.

(150) Maligan, JM., Estiasih, T., Kusnadi, J., 2012. Structured Phospholipids from Commercial Soybean Lecithin Containing Omega-3 Fatty Acids Reduces Atheroslcerosis Risk in Male Sprague Dawley Rats Which Fed with an Atherogenic Diet. WASET. 6(9): 731-737.

(151) Murru, E., Banni, S., Carta, G., 2013. Nutritional properties of dietary omega-3- enriched phospholipids. Biomed Res Int. doi: 10.1155/2013/965417.

(152) Fernandes, MF., Mutch, DM., Leri, F., 2017. The Relationship between Fatty Acids and Different Depression-Related Brain Regions, and Their Potential Role as Biomarkers of Response to Antidepressants. Nutrients. 9(3): pii: E298.

(153) Schonfeld, P., Reiser, G., 2017. Brain energy metabolism spurns fatty acids as fuel due to their inherent mitotoxicity and potential capacity to unleash neurodegeneration. Neurochem Int. 109: 68-77.

(154) Schuschardt, JP., Huss, M., Stauss-Grabo, M., Hahn, A., 2010. Significance of long-chain polyunsaturated fatty acids (PUFAs) for the development and behavious of children. Eur J Pediatr. 169(2): 149-164. 148

149

(155) Calder, PC., 2009. Polyunsaturated fatty acids and inflammatory processes: New twists in an old tale. Biochimie. 91(6): 791-795.

(156) Bazan, NG., Molina, MF., Gordon, WC., 2011. Docosahexaenoic acid signalolipidomics in nutrition: significance in aging, neuroinflammation, macular degeneration, Alzheimer’s, and other neurodegenerative diseases. Annu Rev Nutr. 31: 321-351.

(157) Li, Q., Zhang, Q., Wang, M., Liu, F., Zhao, S., Ma, J., Luo, N., Li, N., Li, Y., Xu, G., Li, J., 2007. Docosahexaenoic acid affects endothelial nitric oxide synthase in caveolae. Arch Biochem Biophys. 466(2): 250-259.

(158) Ma, DW., 2007. Lipid mediators in membranes rafts are important determinants of human health and disease. Appl Physiol Nutr Metab. 32(3): 341-350.

(159) Pifferi, F., Jouin, M., Alessandri, JM., Haedke, U., Roux, F., Perriere, N., Denis, I., Lavialle, M., Guesnet, P., 2007. n-3 Fatty acids modulate brain glucose transport in endothelial cells of the blood-brain barrier. Prostaglandins Leukot Essent Fatty Acids. 77(5-6): 279-286.

(160) Pifferi, F., Jouin, M., Alessandri, JM., Roux, F., Perriere, N., Langelier, B., Lavialle, M., Cunnane, S., Guesnet, P., 2010. n-3 long-chain fatty acids and regulation of glucose transport in two models of rat brain endothelial cells. Neurochem Int. 56(5): 703-710.

(161) Murakami, M., Taketomi, Y., Miki, Y., Sato, H., Hirabayashi, T., Yamamoto, K., 2011. Recent progress in phospholipase A2 research: from cells to animals to humans. Prog Lipid Res. 50(2): 152-192.

(162) Strokin, M., Reiser, G., 2016. Mitochondria from a mouse model of the human infantile neuroaxonal dystrophy (INAD) with genetic defects in VIA iPLA2 have disturbed Ca(2+) regulation with reduction in Ca(2+) capacity. Neurochem Int. 99: 187-193.

(163) Morgan, NV., Westaway, SK., Morton, JE., Gregory, A., Gissen, P., Sonek, S., Cangul, H., Coryell, J., Canham, N., Nardocci, N., Zorzi, G., Pasha, S., Rodriguez, D., Desguerre, I., Mubaidin, A., Bertini, E., Trembath, RC., Simonati, A., Schanen, C., Johnsonm CA., Levinson, B., Woods, CG., Wilmot, B., Kramer, P., Gitschier, J., Maher, ER., Hayflick, SJ., 2006. PLA2G6, encoding a phospholipase A2, mutated in neurodegenerative disorders with high brain iron. Nat Genet. 38(7): 752-754.

(164) Baburina, I., Jackowski, S., 1999. Cellular responses to excess phospholipid. J Biol Chem. 274(14): 9400-9408.

(165) Farooqui, AA., Ong, WY., Horrocks, LA., Farooqui, T., 2000. Brain Cytosolic Phospholipase A2: Localization, Role, and Involvement in Neurological Diseases. Neuroscientist. 6(3): 169-180. 149

150

(166) Klein, J., Chalifa, V., Liscovitsch, M., Loffelholz, K., 1995. Role of phospholipase D activation in nervous system physiology and pathophysiology. 65(4): 1445-1455.

(167) Tamango, E., Robino, G., Obbili, A., Bardini, P., Aragno, M., Parola, M., Danni, O., 2003. H2O2 and 4-hydroxynonenal mediate amyloid beta-induced neuronal apoptosis by activating JNKs and p38MAPK. Exp Neurol. 180(2): 144-155.

(168) Gegotek, A., Skrzydlewska, E., 2019. Biological effect of protein modifications by lipid peroxidation products. Chem Phys Lipids. 221: 46-52.

(169) Beck, G., Sugiura, Y., Shinzawa, K., Kato, S., Setou, M., Tsujimoto, Y., Sakoda, S., Sumi-Akamuru, H., 2011. Neuroaxonal dystrophy in calcium-independent phospholipase A2β deficiency results from insufficient remodeling and degeneration of mitochondrial and presynaptic membranes. J Neurosci. 31(31): 11411-11420.

(170) Carrilho, I., Santos, M., Guimaraes, A., Teixeira, J., Chorao, R., Martins, M., Dias, C., Gregory, A., Westaway, S., Nguyen, T., Hayflick, S., Barbot, C., 2008. Infantile neuroaxonal dystrophy: what’s most important for the diagnosis? Eur J Paediatr Neurol. 12(6): 491-500.

(171) Colombelli, C., Aoun, M., Tiranti, V., 2015. Defective lipid metabolism in neurodegeneration with brain iron accumulation (NBIA) syndromes: not only a matter of iron. J Inherit Metab Dis. 38(1): 123-136.

(172) Dusek, P., Schneider, SA., Aaseth, J., 2016. Iron chelation in the treatment of neurodegenerative diseases. J Trace Elem Med Biol. 38: 81-92.

(173) Lv, H., Shang, P. 2018. The significance, trafficking and determination of labile iron in cytosol, mitochondria and lysosomes. Metallomics. 10(7): 899-916.

(174) Niculescu MD. 2013. Choline and Phosphatidylcholine. Encyclopedia of Human Nutrition. 1:346-351.

(175) Zeisel SH. 2012. Dietary choline deficiency causes DNA strand breaks and alters marks on DNA and histones. Mutat Res. 733: 34-8.

(176) Sher RB, Aoyama C, Huebsch KA, Ji S, Kerner J, Yang Y, Frankel WN, Hoppel CL, Wood PA, Vance DE, Cox GA. 2006. A rostrocaudal muscular dystrophy caused by a defect in choline kinase beta, the first enzyme in phosphatidylcholine biosynthesis. J. Biol. Chem. 281: 4938-48.

(177) Yuan Z, Wagner L, Poloumienko A, Bakovic M. 2004. Identification and expression of a mouse muscle-specific CTL1 gene. Gene. 341: 305-12.

150

151

(178) Uno, J., Obara, K., Suzuki, H., Miyatani, S., Chino, D., Yoshio, T., Tanaka, Y., 2017. Inhibitory Effects of Antidepressants on Acetylcholine-Induced Contractions in Isolated Guinea Pig Urinary Bladder Smooth Muscle. Pharmacology. 99(1-2): 89-98.

(179) Haga T. 1971. Synthesis and release of [14C]-acetylcholine in synaptosomes. J Neurochem. 18:781-98.

(180) Michel V, Bakovic M. 2012. The Ubiquitous Choline Transporter SLC44A1. Cent Nerv Syst Agents Med Chem. 12:70-81.

(181) Nassenstein C, Wiegand S, Lips KS, Li G, Klein J, Kummer W. 2015. Cholinergic activation of the murine trachealis muscle via non-vesicular acetylcholine release involving low-affinity choline transporters. Int Immunopharmacol. 29(1):173-80.

(182) Michel V, Singh RK, Bakovic M. 2011. The impact of choline availability on muscle lipid metabolism. Food Funct. 2:53-62.

(183) Detopoulou P, Panagiotakos DB, Antonopoulou S, Pitsavos C, Stefanadis C. 2008. Dietary choline and betaine intakes in relation to concentrations of inflammatory markers in healthy adults: the ATTICA study. Am J Clin Nutr. 424-30.

(184) Zeisel SH, da Costa KA. 2009. Choline: an essential nutrient for public health. Nutr Rev. 67:615-23.

(185) Tryndyak VP, Marrone AK, Latendresse JR, Muskhelishvili L, Beland FA, Pogribny IP. 2016. MicroRNA changes, activation of progenitor cells and severity of liver injury in mice induced by choline and folate deficiency. J Nutr Biochem. 28:83-90.

(186) Purohit V, Abdelmalek MF, Barve S, Benevenga NJ, Halsted CH, Kaplowitz N, Kharbanda KK, Liu QY, Lu SC, McClain CJ, Swanson C, Zakhari S. 2007. Role of S- adenosylmethionine, folate, and betaine in the treatment of alcoholic liver disease: summary of a symposium. Am J Clin Nutr. 86:14-24.

(187) Selathurai A, Kowalski GM, Burch ML, Sepulveda P, Risis S, Lee-Young RS, Lamon S, Meikle PJ, Genders AJ, McGee SL, Watt MJ, Russell AP, Frank M, Jackowski S, Febbraio MA, Bruce CR. 2015. The CDP-Ethanolamine Pathway Regulates Skeletal Muscle Diacylglycerol Content and Mitochondrial Biogenesis without Altering Insulin Sensitivity. Cell Metab. 21:718-30.

(188) Funai K, Lodhi IJ, Spears LD, Yin L, Song H, Klein S, Semenkovich CF. 2016. Skeletal Muscle Phospholipid Metabolism Regulates Insulin Sensitivity and Contractile Function. Diabetes. 65:358-70.

(189) Basu P, Alibhai FJ, Tsimakouridze EV, Singh RK, Paglialunga S, Holloway GP, Martino TA, Bakovic M. 2015. Male-Specific Cardiac Dysfunction in

151

152

CTP:Phosphoethanolamine Cytidylyltransferase (Pcyt2)-Deficient Mice. Mol Cell Biol. 35:2641-57.

(190) Fullerton MD, Hakimuddin F, Bakovic M. 2007. Developmental and Metabolic Effects of Disruption of the Mouse CTP:Phosphoethanolamine Cytidylyltransferase Gene (Pcyt2). Mol Cell Biol. 27:3327-36.

(191) Schenkel LC, Sivanesan S, Zhang J, Wuyts B, Taylor A, Verbrugghe A, Bakovic M. 2015. Choline supplementation restores substrate balance and alleviates complications of Pcyt2 deficiency. J Nutr Biochem. 26(11): 1221-1234

(192) Folch J, Lees M, Sloane-Stanley GH. 1957. A simple method for the isolation and purification of total lipids from animal tissues. J Biol Chem. 226:497-509.

(193) Bligh EG, Dyer WJ. 1959. A rapid method of total lipid extraction and purification. Can J Biochem Physiol. 37:911-7.

(194) Lo S, Russell JC, Taylor AW. 1970. Determination of glycogen in small tissue samples. J Appl Physiol. 28:234-6.

(195) Mellor H. and Parker PJ. 1998. The extended protein kinase C superfamily. Biochem J. 332: 281-292.

(196) Schmitz-Peiffer C. 2002. Protein kinase C and lipid-induced insulin resistance in skeletal muscle. Ann N Y Acad Sci. 967:146-57.

(197) Kase ET, Feng YZ, Badin PM, Bakke SS, Laurens C, Coue M, Langin D, Gaster M, Thoresen GH, Rustan AC, Moro C. 2015. Primary defects in lipolysis and insulin action in skeletal muscle cells from type 2 diabetic individuals. Biochim Biophys Acta. 1851:1194-1201.

(198) Liu X, Yuan Y, Niu Y, Niu W, Fu L. 2012. The role of AMPK/mTOR/S6K1 signaling axis in mediating the physiological process of exercise-induced insulin sensitization in skeletal muscle of C57BL/6 mice. Biochim Biophys Acta. 1822:1716-26.

(199) Gao X, Wang Y, Randell E, Pedram P, Yi Y, Gulliver W, Sun G. 2016. Higher Dietary Choline and Betaine Intakes Are Associated with Better Body Composition in the Adult Population of Newfoundland, Canada. PLoS One 11(5):e015540.

(201) Sachan DS , Hongu N, Johnsen M. 2005. Decreasing oxidative stress with choline and carnitine in women J Am Coll Nutr. 24(3):172-6.

(202) Bellar D, LeBlanc NR, Campbell B. 2015. The effect of 6 days of alpha glycerylphosphorylcholine on isometric strength. J Int Soc Sports Nutr. 17;12:42.

152

153

(203) Cholewa JM, Guimaraes-Ferreira L, Zanchi NE. 2014. Effects of betaine on performance and body composition: a review of recent finding and potential mechanisms. Amino Acids. 46:1785-93

(204) Senesi P, Luzi L, Montesano A, Mazzocchi N, Terruzzi I. 2013. Betaine supplement enhances skeletal muscle differentiation in murine myoblasts via IGF-1 signaling activation. J Transl Med. 11:1-12

(205) Ejaz A, Martinez-Guino L, Goldfine AB, Ribas-Aulinas F, De Nigris V, Ribo S, Gonzalez-Franquesa A, Garcia-Roves PM, Li E, Dreyfuss JM, Gall W, Kim JK, Bottiglieri T, Villarroya F, Gerszten RE, Patti ME, Lerin C. 2016. Dietary Betaine Supplementation Increases Fgf21 Levels to Improve Glucose Homeostasis and Reduce Hepatic Lipid Accumulation in Mice. Diabetes 65:902-12.

(206) Koh HJ, Brandauer J, Goodyear LJ. 2008. LKB1 and AMPK and the regulation of skeletal muscle metabolism. Curr Opin Clin Nutr Metab Care. 11:227-32.

(207) Park DR, Park KH, Kim BJ, Yoon CS, Kim UH. 2015. Exercise ameliorates insulin resistance via Ca2+ signals distinct from those of insulin for GLUT4 translocation in skeletal muscles. Diabetes. 64:1224-34.

(208) Merlin J, Evans BA, Csikasz RI, Bengtsson T, Summers RJ, Hutchinson DS. 2010. The M3-muscarinic acetylcholine receptor stimulates glucose uptake in L6 skeletal muscle cells by a CaMKK–AMPK–dependent mechanism. Cell Signal. 22:1104-13.

(209) Liu TY, Shi CX, Gao R, Sun HJ, Xiong XQ, Ding L, Chen Q, Li YH, Wang JJ, Kang YM, Zhu GQ. 2015. Irisin inhibits hepatic gluconeogenesis and increases glycogen synthesis via the PI3K/Akt pathway in type 2 diabetic mice and hepatocytes. Clin Sci. 129:839-50.

(210) Stretton C, Hoffmann TM, Munson MJ, Prescott A, Taylor PM, Ganley IG, Hundal HS. 2015. GSK3-mediated raptor phosphorylation supports amino-acid dependent mTORC1-directed signalling. Biochem J. 470:207-21.

(211) Wullschleger S, Loewith R, Hall MN. 2006. TOR signaling in growth and metabolism Cell. 124:471-484.

(212) Hong-Brown LQ, Brown CR, Kazi AA, Huber DS, Pruznak AM, Lang CH. 2010. Alcohol and PRAS40 knockdown decrease mTOR activity and protein synthesis via AMPK signaling and changes in mTORC1 interaction. J Cell Biochem. 109:1172-1184.

(213) Um SH, Frigerio F, Watanabe M, Picard F, Joaquin M, Sticker M, Fumigalli S, Allegrini PR, Kozma SC, Auwerx J, Thomas G. 2004. Absence of S6K1 protects against age- and diet-induced obesity while enhancing insulin sensitivity. Nature. 431:200-5.

153

154

(214) Bond P. 2016. Regulation of mTORC1 by growth factors, energy status, amino acids and mechanical stimuli at a glance. J Int Soc Sports Nutr. 13:8.

(215) Michel, V., et al., 2006. Choline transport for phospholipid synthesis. Exp Biol Med (Maywood). 231, 490-504.

(216) Michel, V., Bakovic, M., 2009. The solute carrier 44A1 is a mitochondrial protein and mediates choline transport. Faseb j. 23, 2749-58.

(217) Michel, V., Bakovic, M., 2012. The ubiquitous choline transporter SLC44A1. Cent Nerv Syst Agents Med Chem. 12, 70-81.

(218) O'Regan, S., et al., 2000. An electric lobe suppressor for a yeast choline transport mutation belongs to a new family of transporter-like proteins. Proc Natl Acad Sci U S A. 97, 1835-40.

(219) Traiffort, E., et al., 2005. Molecular characterization of the family of choline transporter-like proteins and their splice variants. J Neurochem. 92, 1116-25.

(220) Blusztajn, J.K., Slack, B.E., Mellott, T.J., 2017. Neuroprotective Actions of Dietary Choline. Nutrients. 9.

(221) Zeisel, S.H., Klatt, K.C., Caudill, M.A., 2018. Choline. Adv Nutr. 9, 58-60.

(222) Ennis, E.A., Blakely, R.D., 2016. Choline on the Move: Perspectives on the Molecular Physiology and Pharmacology of the Presynaptic Choline Transporter. Adv Pharmacol. 76, 175-213.

(223) Ueland, P.M., 2011. Choline and betaine in health and disease. J Inherit Metab Dis. 34, 3-15.

(224) Inazu, M., 2014. Choline transporter-like proteins CTLs/SLC44 family as a novel molecular target for cancer therapy. Biopharm Drug Dispos. 35, 431-49.

(225) Bauche, S., et al., 2016. Impaired Presynaptic High-Affinity Choline Transporter Causes a Congenital Myasthenic Syndrome with Episodic Apnea. Am J Hum Genet. 99, 753-761.

(226) Wang, H., et al., 2017a. Choline transporter mutations in severe congenital myasthenic syndrome disrupt transporter localization. Brain. 140, 2838-2850.

(227) Kommareddi, P.K., et al., 2010. Isoforms, expression, glycosylation, and tissue distribution of CTL2/SLC44A2. Protein J. 29, 417-26.

154

155

(228) Germain, M., et al., 2015. Meta-analysis of 65,734 individuals identifies TSPAN15 and SLC44A2 as two susceptibility loci for venous thromboembolism. Am J Hum Genet. 96, 532-42.

(229) Kommareddi, P., et al., 2015. Hair Cell Loss, Spiral Ganglion Degeneration, and Progressive Sensorineural Hearing Loss in Mice with Targeted Deletion of Slc44a2/Ctl2. J Assoc Res Otolaryngol. 16, 695-712.

(230) Cheng, C.Y., et al., 2015. New loci and coding variants confer risk for age-related macular degeneration in East Asians. Nat Commun. 6, 6063.

(231) Di Meo, I., and Tiranti, V. 2018. Classification and molecular pathogenesis of NBIA syndromes. Eur J Paediatr Neurol. 22, 272-284.

(232) Tello, C., et al., 2017. On the complexity of clinical and molecular bases of neurodegeneration with brain iron accumulation. Clin Genet.

(233) Colombelli, C., Aoun, M., Tiranti, V., 2015. Defective lipid metabolism in neurodegeneration with brain iron accumulation (NBIA) syndromes: not only a matter of iron. J Inherit Metab Dis. 38, 123-36.

(234) Van der Auwera, G.A., et al., 2013. From FastQ data to high confidence variant calls: the Genome Analysis Toolkit best practices pipeline. Curr Protoc Bioinformatics. 43, 11.10.1-33.

(235) Reuter, M.S., et al., 2017. Diagnostic Yield and Novel Candidate Genes by Exome Sequencing in 152 Consanguineous Families With Neurodevelopmental Disorders. JAMA Psychiatry. 74, 293-299.

(236) Drecourt, A., et al., 2018. Impaired Transferrin Receptor Palmitoylation and Recycling in Neurodegeneration with Brain Iron Accumulation. Am J Hum Genet. 102, 266-277.

(237) Riemer, J., et al., 2004. Colorimetric ferrozine-based assay for the quantitation of iron in cultured cells. Anal Biochem. 331, 370-5.

(238) Boxberger, K.H., Hagenbuch, B., Lampe, J.N., 2014. Common drugs inhibit human organic cation transporter 1 (OCT1)-mediated neurotransmitter uptake. Drug Metab Dispos. 42, 990-5.

(239) Yara, M., et al., 2015. Molecular and functional characterization of choline transporter in the human trophoblastic cell line JEG-3 cells. Placenta. 36, 631-7.

(240) Wang, T., et al., 2007. Choline transporters in human lung adenocarcinoma: expression and functional implications. Acta Biochim Biophys Sin (Shanghai). 39, 668- 74. 155

156

(241) Michel, V., Singh, R.K., Bakovic, M., 2011. The impact of choline availability on muscle lipid metabolism. Food Funct. 2, 53-62.

(242) Saitoh, M., et al., 2009. Phosphatidyl ethanolamine with increased polyunsaturated fatty acids in compensation for plasmalogen defect in the Zellweger syndrome brain. Neurosci Lett. 449, 164-7.

(243) Wang, Y., et al., 2017b. Arabidopsis choline transporter-like 1 (CTL1) regulates secretory trafficking of auxin transporters to control seedling growth. PLoS Biol. 15, e2004310.

(244) Saitsu, H., et al., 2013. De novo mutations in the autophagy gene WDR45 cause static encephalopathy of childhood with neurodegeneration in adulthood. Nat Genet. 45, 445-449.

(245) Wiethoff, S., Houlden, H., 2017. Neurodegeneration with brain iron accumulation. Handb Clin Neurol. 145, 157-166.

(246) Gregory, A., Hayflick, S., 1993. Neurodegeneration with Brain Iron Accumulation Disorders Overview. In: GeneReviews(R). Vol., M.P. Adam, H.H. Ardinger, R.A. Pagon, S.E. Wallace, L.J.H. Bean, H.C. Mefford, K. Stephens, A. Amemiya, N. Ledbetter, ed.^eds. University of Washington, Seattle University of Washington, Seattle. GeneReviews is a registered trademark of the University of Washington, Seattle. All rights reserved., Seattle (WA).

(247) Langwinska-Wosko, E., et al., 2016. Retinal and optic nerve abnormalities in neurodegeneration associated with mutations in C19orf12 (MPAN). J Neurol Sci. 370, 237-240.

(248) Amaral, L.L., et al., 2015. Neurodegeneration with Brain Iron Accumulation: Clinicoradiological Approach to Diagnosis. J Neuroimaging. 25, 539-51.

(249) Skowronska, M., et al., 2017. Evolution and novel radiological changes of neurodegeneration associated with mutations in C19orf12. Parkinsonism Relat Disord.

(250) Al-Saeedi, F., Smith, T., Welch, A., 2007. [Methyl-3H]-choline incorporation into MCF-7 cells: correlation with proliferation, choline kinase and phospholipase D assay. Anticancer Res. 27, 901-6.

(251) Farooqui, A.A., Horrocks, L.A., 2001. Plasmalogens: workhorse lipids of membranes in normal and injured neurons and glia. Neuroscientist. 7, 232-45.

(252) Farooqui, A.A., Horrocks, L.A., Farooqui, T., 2000. Glycerophospholipids in brain: their metabolism, incorporation into membranes, functions, and involvement in neurological disorders. Chem Phys Lipids. 106, 1-29.

156

157

(253) Traiffort, E., O’Regan, S., Ruat, M., 2013. The choline transporter-like family SLC44: properties and roles in human diseases. Mol Aspects Med. 34, 646-654.

(254) McMaster, CR., Tardi, PG., Choy, PC., 1992. Modulation of phosphatidylethanolamine biosynthesis by exogenous ethanolamine and analogues in the hamster heart. Mol Cell Biochem. 116, 69-73.

(255) Patel, D., Witt, SN., 2017. Ethanolamine and Phosphatidylethanolamine: Partners in Health and Disease. Oxid Med Cell Longev. Epub 2017 Jul 12.

(256) van der Veen, JN., Kennelly, JP., Wan, S., Vance, JE., Vance, DE., Jacobs, RL., 2017. The critical role of phosphatidylcholine and phosphatidylethanolamine metabolism in health and disease. Biochem Biophys Acta Biomembr. Epub 2017 Apr 11.

(257) Yorek, MA., Dunlap, JA., Spector, AA., Ginsberg, BH., 1986. Effect of ethanolamine on choline uptake and incorporation into phosphatidylcholine in human Y79 retinoblastoma cells. J Lipid Res. 11, 1205-1213.

(258) Massarelli, AC., Dainous, F., Hoffmann, D., Mykita, S., Freysz, L., Dreyfus, H., Massarelli, R., 1986. Uptake of ethanolamine in neuronal and glial cell cultures. Neurochem Res. 11, 29-36.

(259) Anderson, CJ., Satkovich, J., Koseoglu, VK., Agaisse, H., Kendall, MM., 2018. The Ethanolamine Premease EutH Promotes Vacuole Adaptation of Salmonella enterica and Listeria monocytogenes during Macrophage Infection. Infect Immun. 86, pii:e00172-18.

(260) Stojiljokovic, I., Baumler, AJ., Heffron, F., 1995. Ethanolamine utilization in Salmonella typhimurium: nucleotide sequence, protein expression, and mutational analysis of the cchA cchB eutE eutJ eutG eutH gene cluster. J Bacteriol. 177, 1357- 1366.

(261) Rifkin, MR., Strobos, CA., Fairlamb, AH., 1995. Specificity of ethanolamine transport and its further metabolism in Trypanosoma brucei. J Biol Chem. 270, 16160- 16166.

(262) Poloumienko, A., Cote, A., Quee, AT., Zhu, L., Bakovic, M. 2004. Genomic organization and differential splicing of the mouse and human Pcyt2 genes. Gene. 325, 145-155.

(263) Iwao, B., Yara, M., Hara, N., Kawai, Y., Yamanaka, T., Nishihara, H., Inoue, T., Inazu, M., 2016. Functional expression of choline transporter like-protein 1 (CTL1) and CTL2 in human brain microvascular endothelial cells. Neurochem Int. 93, 40-50.

157

158

(264) Basu Ball, W., Baker, CD., Neff, JK., Apfel, GL., Lagerborg, KA., Zun, G., Petrovic, U., Jain, M., Gohil, V., 2018. Ethanolamine ameliorates mitochondrial dysfunction in cardiolipin-deficient yeast cells. J Biol Chem. 293, 10870-10883.

(265) Yang, H., Xiong, X., Li, T., Yin, Y., 2016. Ethanolamine enhances the proliferation of intestinal epithelial cells via the mTOR signaling pathway and mitochondrial function. In Vitro Cell Dev Biol Anim. 52, 562-567.

(266) Bonora, M., Wieckowsk, MR., Chinopoulos, C., Kepp, O., Kroemer, G., Galluzzi, L., Pinton, P., 2015. Molecular mechanisms of cell death: central implication of ATP synthase in mitochondrial permeability transition. Oncogene. 34(12): 1608.

(267) Michalke, B., Willkommen, D., Venkataramani, V., 2019. Iron Redox Speciation Analysis Using Capillary Electrophoresis Coupled to Inductively Coupled Plasma Mass Spectrometry (CE-ICP-MS). Front Chem. 7:136.

(268) Gonzales, MM., Tarumi, T., Kaur, S., Nualnim, N., Fallow, BA., Pyron, M., Tanaka, H., Haley, AP., 2013. Aerobic fitness and the brain: increased N-acetyl- aspartate and choline concentrations in endurance-trained middle-aged adults. Brain Topogr. 26(1): 125-134.

(269) von Allworden, HN., Horn, S., Kahl, J., Feldheim, W., 1993. The influence of lecithin on plasma choline concentration in triathletes and adolescent runners during exercise. Eur J Appl Physiol Occup Physiol. 67(1): 87-91.

(270) Lagace, TA., Ridgway, ND., 2013. The role of phospholipids in the biological activity and structure of the endoplasmic reticulum. Biochim Biophys Acta. 1833(11): 2499-2510.

(271) Kazak, L., Chouchani, ET., Jedrychowski, MP., Erickson, BK., Shinoda, K., Cohen, P., Vetrivelan, R., Lu, GZ., Laznik-Bogoslavski, D., Hasenfuss, SC., Kajimura, S., Gygi, SP, Spiegelman, BM., 2015. A creatine-driven substrate cycle enhances energy expenditure and thermogenesis in beige fat. Cell. 163(3): 643-655.

(272) Oliveira, TE., Castro, E., Belchior, T., Andrade, ML., Chaves-Filho, AB., Peixoto, AS., Moreno, MF., Ortiz-Silva, M., Moreira, RJ., Inague, A., Yoshinaga, MY., Miyamoto, S., Moustaid-Moussa, N., Festuccia, WT., 2019. Fish Oil Protects Wild Type and Uncoupling Protein 1-Deficient Mice from Obesity and Glucose Intolerance by Increasing Energy Expenditure. Mol Nutr Food Res. 63(7): e1800813.

(273) Fuller-Jackson, JP., Henry, BA., 2018. Adipose and skeletal muscle thermogenesis: studies from large animals. J Endocrinol. 237(3): R99-R115.

(274) Betz, MJ., Enerback, S., 2018. Targeting thermogenesis in brown fat and muscle to treat obesity and metabolic disease. Nat Rev Endocrinol. 14(2): 77-87.

158

159