Assessing the Role of the ATRX Chromatin Remodeling Protein in Myelination Within the Mouse Central Nervous System

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Assessing the role of the ATRX chromatin remodeling protein in myelination within the mouse central nervous system

Matthew W. Edwards The University of Western Ontario

Supervisor Dr. Nathalie Berube The University of Western Ontario

Graduate Program in Biochemistry A thesis submitted in partial fulfillment of the equirr ements for the degree in Master of Science © Matthew W. Edwards 2016

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Recommended Citation Edwards, Matthew W., "Assessing the role of the ATRX chromatin remodeling protein in myelination within the mouse central nervous system" (2016). Electronic Thesis and Dissertation Repository. 4047. https://ir.lib.uwo.ca/etd/4047

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ATRX is a Rad54 translocase homolog implicated in regulating the epigenetic environment of heterochromatin, mitosis, and gene transcription. Myelin is a specialized membrane encompassing axons of neurons, allowing for efficient propagation of action potentials. CNS myelin arises via oligodendrocyte precursor cell (OPC) differentiation into myelinating oligodendrocytes (OLs), which then extend and wrap axons. Myelination is a tightly regulated process, however the mechanisms involved have yet to fully manifest. Here, we report that ATRX loss results in hypomyelination due to a defect in OPC differentiation and OL maturation, as ATRX loss did not affect OPC production, proliferation, or cholesterol biosynthesis. Additionally, myelin sheath formation was unaffected when ATRX was lost specifically from OLs, suggesting a non cell-autonomous role of ATRX in myelination. Clarifying the role of ATRX in myelination presents a novel mechanism that regulates myelin sheath formation, and findings from this study will benefit future treatments of myelin disorders, including ATR-X.

Keywords

ATRX, ATR-X syndrome, myelination, oligodendrocyte, gene expression, transcriptional regulation, brain development, cholesterol, mouse model

Co-Authorship Statement

I designed and executed all experiments, data analysis, and written material in this thesis with the exception of the following:

Chapter 3.6 – Isolated whole forebrains from P14 control and AtrxFoxG1Cre mice were delivered to Brian Sutherland at the Metabolic Phenotyping Laboratory at Robarts Research Institute (Western University, London ON) for analysis of cholesterol abundance.

Acknowledgments

First and foremost, I would like to thank my parents, as I would not be here without them. I cannot thank you enough for your constant love and support, and always inspiring me to be the best person I can be. Words cannot explain how much the both of you mean to me.

I would like to thank my brother Chris, my sister Callie, and my brother-in-law Jacob for always keeping me grounded and giving me positive advice when I needed it the most.

I would like to give a special thanks to my friends Carlee, Charles, Mike, Piru, Aren, and Ashley. Thank you for all of your support during my time as a graduate student. I deeply cherish all of the coffee breaks and beer nights we had talking not only about science, but life as well.

Thank you to all of the current and past members of the Bérubé lab for all of your support, advice, and friendship over the past several years. Working along side all of you was an absolute pleasure.

Finally, I would also like to thank my supervisor Dr. Nathalie Bérubé. Thank you for giving me the opportunity to pursue my passion in research. During my time in your lab I have gained many attributes that will aid me in becoming a strong and successful scientific researcher.

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Table of Contents

Abstract...... i

Co-Authorship Statement ...... ii

Acknowledgments...... iii

Table of Contents ...... iv

Tables ...... vii

Figures...... viii

Appendices...... xi

Abbreviations...... xi

1 Introduction...... 1

1.1 ATRX ...... 1

1.1.1 The ATRX gene and protein...... 1

1.1.2 Cellular functions of ATRX...... 5

1.1.3 ATR-X syndrome ...... 7

1.1.4 Phenotypic abnormalities associated with ATRX loss...... 10

1.2 Myelination...... 12

1.2.1 The myelin sheath ...... 12

1.2.2 Oligodendrocyte development...... 13

1.2.3 Composition of the myelin sheath...... 21

1.2.3.1 Myelin-specific proteins ...... 22

1.2.3.2 Lipid content of the myelin sheath ...... 24

1.2.4 Pathologies related to myelin sheath dysfunction...... 25

1.3 Hypothesis and thesis objectives...... 28

2 Materials and Methods ...... 31

2.1 Animal husbandry, tissue harvesting, and genotyping ...... 31

2.2 Microarray analysis...... 35

2.3 Quantitative real-time PCR...... 37

2.4 Protein isolation ...... 40

2.5 Western blot analysis...... 41

2.6 Immunofluorescence staining and cell counting...... 44

2.7 Cholesterol biosynthesis...... 48

3 Results ...... 51

3.1 Transcriptional profiling of the postnatal AtrxFoxG1Cre forebrain reveals a decrease in myelin-specific gene expression...... 51

3.2 Loss of ATRX from the mouse forebrain results in hypomyelination...... 56

3.3 Hypomyelination in the AtrxFoxG1Cre mouse forebrain is not due to a reduction in neuronal cell number...... 59

3.4 Oligodendrocyte precursor cell production and morphology is unchanged in AtrxFoxG1Cre mice at birth ...... 62

3.5 Differentiation of oligodendrocyte precursor cells into immature oligodendrocytes is hindered in the absence of ATRX...... 65

3.6 Cholesterol biosynthesis is unaffected in the P14 AtrxFoxG1Cre mouse forebrain...... 68

3.7 Generation of a novel mouse model to study the oligodendrocyte- specific function of ATRX...... 71

3.8 Oligodendrocyte-specific deletion of ATRX does not affect myelin sheath formation in the mouse brain ...... 74

4 Discussion ...... 80

4.1 Summary of results...... 80

4.2 ATRX acts in a non cell-autonomous role to regulate myelin sheath formation...... 84

4.2.1 ATRX may act to control neuronal-specific gene expression to regulate myelin sheath formation...... 84

4.2.2 Astrocyte influence in CNS myelination...... 88

4.2.3 Systemic regulation and growth factor influence of myelin sheath formation in the CNS...... 90

4.3 Future directions...... 92

5 Conclusions...... 98

References ...... 99

Appendices...... 112

Curriculum Vitae ...... 114

Tables

Table 1. Primer sequences for PCR and qRT-PCR...... 38

Table 2. Primary antibodies used for western blot analysis and immunofluorescence staining...... 42

Table 3. Seconardy antibodies used for western blot analysis and immunofluorescence staining...... 43

vii

Figures

Figure 1. The ATRX protein structure and function ...... 2

Figure 2. The oligodendrocyte development pathway ...... 14

Figure 3. Hypothetical mechanism of ATRX in myelination of neurons within the CNS...... 29

Figure 4. Schematic of mice harbouring loxP sites used for Cre-loxP recombination...... 32

Figure 5. Coronal section of the postnatal mouse brain ...... 45

Figure 6. Transcriptional profiling of genes from P17 AtrxFoxG1Cre microarray revealed a significant decrease in expression of myelin-specific genes compared to littermate- matched controls...... 53

Figure 7. Expression of myelin-specific proteins is significantly reduced in P20 AtrxFoxG1Cre mice compared to littermate-matched controls...... 57

Figure 8. Hypomyelination is not due to a reduction in the number of neurons in the mouse brain...... 60

Figure 9. Expression of OPC markers is unchanged in P0.5 AtrxFoxG1Cre mice compared to littermate-matched controls...... 63

Figure 10. The number of APC/OLIG2-postive cells is significantly decreased in P10 AtrxFoxG1Cre mice compared to littermate-matched controls...... 66

Figure 11. Cholesterol levels are unchanged in the absence of ATRX from P14 mice...... 69

Figure 12. ATRX expression is successfully lost from OLs in AtrxCnpCre mice...... 72

Figure 13. Characterization of the P20 AtrxCnpCre and littermate-matched control mouse phenotypes...... 75 viii

Figure 14. Expression of myelin-specific proteins is unchanged between P20 AtrxCnpCre mice compared to littermate-matched controls...... 78

Figure 15. Proposed model of ATRX and its potential mechanism in myelination of neurons in the mouse CNS ...... 96

Appendices

Appendix A. ATRX expression is retained in neurons in P20 AtrxCamkIICre mice, but is lost from neurons in 3-month-old AtrxCamkIICre mice…………………………………..113

Appendix B. Expression of myelin-specific proteins is reduced in 3-month-old AtrxCamkII mice compared to littermate-matched controls……………………………114

Abbreviations

µCi Microcurie

µg Microgram

µL Microlitre

µm Micrometer

µM Micromolar

°C Degree celsius

ADD ATRX-DNMT3-DNMT3L

AKT AKR thymoma

ANOVA Analysis of variance aPAR Ancestral pseudoautosomal region

APC Adenomatous polyposis coli

APS Ammonium persulfate

Ascl1 Achaete-scute homolog 1

ATP Adenosine triphosphate

ATPase Adenosine triphosphatase

ATRX Alpha thalassaemia mental retardation, X-linked

ATRXt Truncated isoform of ATRX

BRG1 BRM/SWI2-related gene 1

BSA Bovine serum albumin

C-terminus Carboxy-terminus

CamkII Calcium/calmodulin-dependent protein kinase II cDNA Complimentary DNA

CNP 2’, 3’-cyclic-nucleotide 3’-phosphodiesterase

CNS Central nervous system

CO2 Carbon dioxide cRNA Complimentary RNA

Cspg4 Chondroitin sulfate proteoglycan 4

Ct Cycle threshold

CTCF CCCTC-binding factor

CTP Cytidine triphosphate

CXCL12 C-X-C motif chemokine 12

DAPI 4’,6-diamidino-2-phenylindole

xii

Daxx Death domain-associated protein

ddH2O Double distilled dihydrogen monoxide

Dlk1 Delta-like 1 homolog

DMNT DNA (cytosine-5) methyltransferase

DNA Deoxyribonucleic acid

Dtx-1 Deltex-1

E Embryonic day

ECL Enhanced chemilluminescence

ECM Extracellular matrix

ErbB4 Receptor tyrosine-protein kinase erbB-4

ERK1 Extracellular signal-regulated kinase 1 f Floxed allele

FACS Fluorescence-activated cell sorting

FGF2 Fibroblast growth factor 2

FGFR1 Fibroblast growth factor receptor 1

FGFR2 Fibroblast growth factor receptor 2

FoxG1 Forkhead box G1

xiii

Fyn Fyn tyrosine kinase protooncogene

GALC Galactocerebrosidase

Gapdh Glyceraldehyde 3-phosphate dehydrogenase

Glast Glial high affinity glutamate transporter

GO Gene ontology

GPCR17 G-protein coupled receptor 17

Gtl2 Gene trap locus 2

H3.3 Histone 3.3

H3K9me3 Histone 3 lysine 9, trimethylation

HbH Hemoglobin H

ID2 Inhibitor of DNA binding 2, HLH protein

ID4 Inhibitor of DNA binding 4, HLH protein

IGF-1 Insulin-like growth factor 1

IGF-1R Insulin-like growth factor receptor 1

IGF-2 Insulin-like growth factor 2

ILK Integrin-linked kinase

INCENP Inner centromere protein

xiv iOL Immature oligodendrocyte kb Kilobase kDA Kilodalton

KD Krabbe disease

LIF Leukemia inhibitory factor

LINGO-1 Leucine-rich immunoglobulin domain-containing, 1

M Molar

MAG Myelin-associated glycoprotein

MBP Myelin basic protein

MeCP2 Methyl-CpG-binding protein 2 mm Millimeter mM Millimolar mL Milliliter

MOBP Myelin-associated oligodendrocyte basic protein

MOG Myelin oligodendrocyte glycoprotein

MRF Myelin regulatory factor

MRI Magnetic resonance imaging

xv mRNA Messenger RNA

MS Multiple sclerosis

N-terminus Amino terminus

N Number of replicates

NaCl Sodium chloride

NaF Sodium fluoride

Na3VO4 Sodium orthovanadate

NCAM Neural cell-adhesion molecule

NeuN Neuronal nuclei antigen

NICD Notch intracellular domain

Nkx2.2 NK2 homeobox protein 2

Nkx6 NK6 homeobox protein nM Nanomolar

NPC Neural progenitor cell

NRG-1 Neuregulin 1

OCT Optical cutting temperature

OL Oligodendrocyte

xvi

OLIG1 Oligodendrocyte lineage transcription factor 1

OLIG2 Oligodendrocyte lineage transcription factor 2

OMIM Online mendelian inheritance of man

OPC Oligodendrocyte precursor cell

P Postnatal day p53 Tumour suppressor protein 53

PBS Phosphate buffered saline

PCR Polymerase chain reaction

PDGFA Platelet derived growth factor, subunit A

PDGFRα Platelet derived growth factor receptor, alpha

PFA Paraformaldehyde

PHD Plant homeodomain

PI3K Phosphatidylinositol-4,5-bisphosphate 3-kinase

PINCH Particularly interesting new cysteine-histidine-rich

protein

PLP Proteolipid protein

PMD Pelizaeus-Merzbacher disease

xvii

PML-NB Promyelocytic leukaemia nuclear bodies

PMSF Phenylmethylsulfonyl fluoride

PNS Peripheral nervous system

PSA-NCAM Polysialylated neural cell adhesion molecule qRT-PCR Quantitative real-time PCR

Rad54 Radiation mutant 54

RIPA Radioimmunoprecipitation assay

RMA Robust multi-array average

RNA Ribonucleic acid

ROS Reactive oxygen species

RPM Revolutions per minute

SAPE Streptavidin-phycoerythrin

SD Standard deviation

SDS Sodium dodecyl sulfate

SDS-PAGE SDS polyacrylamide gel electrophoresis

SEM Standard error of the mean

SHH Sonic hedgehog

xviii

Sip1 Survival of motor neuron protein-interacting protein 1

SMAD Mothers against decapentaplegic

Snf2 Sucrose non-fermenting 2

Sox10 SRY-box 10

SRY Sex-determining region Y

SWI/SNF Switch/sucrose non-fermenting

T3 Triiodothyronine

T4 Thyroxine

TBST Tris buffered saline - Tween 20

TEMED Tetramethylethylenediamine

TNF Tumour necrosis factor

TNFR2 Tumour necrosis factor receptor 2

Tris Tris(hydroxymethyl)aminomethane

TSH Thyroid stimulating hormone

UTP Uridine triphosphate

V Voltage

WAVE2 Veprolin-homologous protein 2

xix

WT Wild-type

ZFP191 Zinc finger protein 191

Zfhx1b Zinc finger homebox 1b

xx 1

1 Introduction

1.1 ATRX

1.1.1 The ATRX gene and protein

The alpha thalassaemia mental retardation, X-linked (ATRX) protein is a chromatin remodeling protein that is a homolog of the radiation mutant 54

(Rad54) translocase (Stayton et al., 1994). The ATRX gene is located on the long-arm of the X-chromosome (Xq13.3), and contains 36 exons spanning over

300 kilobases (kb) of genomic DNA (Gibbons et al., 1995; Picketts et al., 1996a).

Analysis of the ATRX gene revealed that it encodes at least three alternative transcripts. Two of the transcripts are the result of an alternative splicing event of exon 6, resulting in two 10.5 kb mRNA transcripts that are predicted to give rise to slightly different proteins of 265 and 280 (kilodalton) kDa, respectively. The third transcript results from a failure to splice out intron 11 of the ATRX gene, resulting in a truncated isoform of the ATRX protein (ATRXt) (Bérubé et al., 2000;

Garrick et al., 2004). The function of the ATRXt protein has not been fully elucidated, however it has been found to have binding affinity for the full-length

ATRX protein suggesting that it may be required to regulate ATRX activity at heterochromatin (Garrick et al., 2004; Bérubé et al., 2005). The full-length ATRX protein contains two highly conserved domains; a carboxy (C)-terminal helicase/adenosine triphosphatase (ATPase) domain that is a member

Figure 1. The ATRX protein structure and function

A) Representation of the ATRX protein containing two-highly conserved domains (the ADD DNA binding domain and the Snf2 chromatin remodeling domain), as well as the areas of the ATRX protein that interact with Daxx, MeCP2 and PML nuclear bodies. B) Representation of the ATRX protein function at pericentromeric heterochromatin. The ADD domain interacts with DNA through recognition of the H3K9me3 and H3K4me0 epigenetic modifications (Dhayalan et al., 2011; Eustermann et al., 2011; Iwase et al., 2011). ATRX interacts with DAXX to allow incorporation of H3.3 into nucleosomes of chromatin (Drané et al., 2010; Goldberg et al., 2010; Lewis et al., 2010; Wong et al., 2010).

4 of the sucrose non-fermenting 2 (Snf2) family of proteins, as well as an amino

(N)-terminal ATRX-DNMT3-DNMT3L (ADD) domain containing a plant homeodomain (PHD)-like zinc finger, and a C2-C2 GATA-like zinc finger domain

(Figure 1A) (Picketts et al., 1998; Xie et al., 1999; Aapola et al., 2000; Argentaro et al., 2007). On the other hand, the ATRXt protein lacks the C-terminal Snf2

ATPase domain, but retains the N-terminal ADD domain suggesting it may act primarily at pericentromeric heterochromatin (Bérubé, 2011).

ATRX is a member of the Snf2 subgroup of the switch/sucrose non-fermenting

(SWI/SNF) family of proteins as it contains seven highly conserved collinear domains of this family, demonstrating its function in ATPase chromatin remodeling activity (Picketts et al., 1996). SWI/SNF proteins have been shown to play roles in a vast array of cellular functions including transcriptional elongation,

DNA repair, and regulation of mitosis (Carlson and Laurent, 1994; Eisen et al.,

1995; Picketts et al., 1996).

The ATRX protein also contains an ADD chromatin-binding domain. It has been found that ATRX can recognize and bind histone tails through various modifications, including trimethylation on lysine 9 of histone 3 (H3K9me3) and

H3K4me0 (Dhayalan et al., 2011; Eustermann et al., 2011; Iwase et al., 2011). In addition, in vitro studies have also shown ATRX has the ability to bind to lysine 4 of the histone 3 variant, H3.3 (Wong et al., 2010).

1.1.2 Cellular functions of ATRX

ATRX is a nuclear protein crucial for the regulation of the epigenetic environment of heterochromatin, mitosis, and gene transcription. It has been established that

ATRX is required for the proper expression of a subset of genes known as ancestral pseudoautosomal region (aPAR) genes (Levy et al., 2008). It was determined that ATRX promotes the expression of aPAR genes through the incorporation of H3.3 (Levy et al., 2014). ATRX facilitates the incorporation of

H3.3 into nucleosomes of aPAR gene bodies resulting in the relaxation of barrier structures known as G-quadraplexes. Inhibition of G-quadraplex formation allows for elongation of RNA polymerase II along the gene body, thus initiating transcription (Levy et al., 2014). In the absence of ATRX, H3.3 incorporation into nucleosomes of aPAR gene bodies was diminished, thus allowing for the formation of G-quadraplexes. These G-quadraplex structures were found to block, and subsequently stall RNA polymerase II from progressing along the gene body resulting in a significant decrease of aPAR gene expression (Levy et al., 2014). These findings reveal a primary role for ATRX in the regulation of gene expression.

ATRX has been shown to influence gene expression through its interaction with several factors, including methyl-CpG-binding protein 2 (MeCP2), CCCTC- binding factor (CTCF), and cohesin. Studies have shown that ATRX binds with

MeCP2, CTCF, and cohesin to influence the expression of the H19/Igf2 and

Gtl2/Dlk1 imprinted regions in the postnatal brain. It was determined that MeCP2

6 acts as a recruiter for ATRX binding at these sites, as loss of ATRX does not alter the MeCP2 binding efficiency (Kernohan et al., 2010). On the contrary,

ATRX loss causes a decrease in CTCF and cohesin occupancy at these sites, resulting in altered expression of H19, Igf2, and many other imprinted genes

(Kernohan et al., 2010). In addition, ATRX and MeCP2 binding influences gene expression by regulating the chromatin architecture at imprinting control regions.

It was found that binding of ATRX to chromatin through its interaction with

MeCP2, repositions nucleosome placement allowing CTCF binding to these sites

(Kernohan et al., 2014). CTCF binding was found to enable the long-distance recruitment of enhancer elements to imprinting control regions through its interaction with cohesin, resulting in an increase in expression of imprinted genes

(Kernohan et al., 2014).

ATRX has also been found to interact with death domain-associated protein

(Daxx) to influence gene expression (Xue et al., 2003; Tang et al., 2004). It has been shown that the ATRX/Daxx complex exhibits translocase activity, as well as mononucleosome-dependent ATPase activity (Xue et al., 2003; Tang et al.,

2004); however, this activity was found not to reposition nucleosomes to influence gene expression, suggesting that the ATRX/Daxx complex regulates gene expression through an alternative mechanism. Daxx has been discovered to act as a chaperone for the H3.3 histone variant. Recent studies demonstrate that ATRX forms a chromatin remodeling complex with Daxx to deposit H3.3 at both pericentromeric heterochromatin, and telomeres (Figure 1B) (Drané et al.,

2010; Goldberg et al., 2010; Lewis et al., 2010; Wong et al., 2010). It was

7 determined that when ATRX is lost, H3.3 is no longer deposited in nucleosomes at pericentromeric heterochromatin and telomeres, leading to decreased levels of pericentromeric transcripts, and increased levels of telomeric transcripts in embryonic stem cells (Wong et al., 2010).

In addition to influencing gene expression and the epigenetic environment of chromatin, ATRX has also been implicated in the regulation of mitosis. Initially, it was determined that ATRX becomes hyperphosphorylated during mitosis suggesting that it may be required for proper cell-division (Bérubé et al., 2000). In vitro studies have shown that human cells deficient for ATRX, display an irregular assembly of chromosomes at the metaphase plate, disruption of spindle morphology, and a delay in cell-cycle progression resulting in irregular nuclei morphology (Ritchie et al., 2008). In addition, mice lacking ATRX exhibited an increase in the abundance of micronuclei in the ventricular zone of the forebrain coinciding with previous in vitro findings.

These studies show that ATRX is required for multiple cellular functions, and loss of ATRX can lead to severe detriments during development.

1.1.3 ATR-X syndrome

ATR-X syndrome (OMIM#301040) is a neurodevelopmental disorder that is characterized by a wide range of phenotypic aberrations such as severe cognitive deficits, seizures, microcephaly, various developmental abnormalities,

8 and alpha thalassaemia (a decrease of α-globin abundance in the blood)

(Gibbons et al., 1995; Gibbons and Higgs, 2000; Gibbons, 2006). ATR-X syndrome patients have been shown to display intellectual disability associated with a profound mental retardation (95% of patients), genital abnormalities (80%), skeletal abnormalities (90%), microcephaly (reduction in brain size) (75%), seizures (30%), HbH inclusions (87%), and characteristic facial features (94%)

(Gibbons, 2006). Recent magnetic resonance imaging (MRI) studies have also revealed that ATR-X syndrome patients display varying degrees of white matter abnormalities, and delayed myelination (Wada et al., 2013).

ATR-X syndrome results from hypomorphic mutations within the ATRX gene, resulting in protein dysfunction. The majority of mutations in the non-truncated form of ATRX resulting in ATR-X syndrome, occur in the ADD DNA binding domain (49% of cases), and the SWI/SNF2 ATPase chromatin remodeling domain (30% of cases) (Gibbons et al., 2008). It has been postulated that mutations within the remaining domains of the ATRX protein act as neutral polymorphisms rather than lethal mutations (Mitson et al., 2011).

Mutations within the Snf2 ATPase domain were found to result in defective DNA translocase and ATPase activity, as well as abnormal localization of ATRX to promyelocytic leukaemia nuclear bodies (PML-NB) (Bérubé et al., 2008; Mitson et al., 2011). Furthermore, missense mutations within the ADD domain result in a significant decrease of DNA binding ability, and abnormal targeting to pericentromeric heterochromatin (Argentaro et al., 2007). These findings indicate that ATRX has a subnuclear function in maintaining proper brain development.

ATR-X syndrome is an extremely rare disorder, as approximately only 200 people (prevalence of <1-9/1,000,000) have been diagnosed worldwide

(Gibbons, 2006). However, it is estimated that the prevalence of ATR-X may actually be closer to 1/30-40,000 as some patients have yet to be diagnosed

(Wada et al., 2013). ATR-X syndrome primarily affects males, as the ATRX gene is located on the X-chromosome. Females carrying a mutated ATRX allele are phenotyically normal as their cells undergo the phenomenon of non-random X- inactivation (Gibbons et al., 1992).

In addition to ATR-X syndrome, mutations within the ATRX gene have been implicated in several other neurological disorders such as Juberg-Mar-sidi, X- linked mental retardation with spastic paraplegia, Carpenter-Waziri, Holmes-

Gang, Smith-Fineman-Myers, and Chudley-Lowry syndromes (Villard et al.,

1996; Abidi et al., 1999; Lossi et al., 1999; Stevenson et al., 2000; Villard et al.,

2000; Abidi et al., 2005)

ATRX has been implicated to be crucial for proper brain development, as mutations within the ATRX gene result in a severe neurological disorder.

Therefore, it is imperative to study the ATRX protein to further our knowledge of its subcellular function in brain development. By increasing our knowledge of the

ATRX protein, therapies can be devised to combat the neurological disorders that stem from mutations within ATRX.

1.1.4 Phenotypic abnormalities associated with ATRX

loss

As mentioned above, ATR-X syndrome patients display a series of developmental abnormalities, and it has been found that a number of these abnormalities are also illustrated in mice deficient for ATRX. To study the phenotypic changes in mice when ATRX is lost, Bérubé and colleagues developed a mouse model that lacks Atrx expression specifically from the forebrain beginning at embryonic day 8.5 (E8.5) (Bérubé et al., 2005).

Analysis of the prenatal mouse brain revealed that loss of ATRX results in a reduction of overall brain size. This was found to be due to hypocellularity in both the cerebral cortex and hippocampus that results from a dramatic increase in the number of apoptotic cells (Bérubé et al., 2005). It was further determined that the increase in apoptosis is not the only mechanism that contributes to a reduction in brain size when ATRX is lost. Seah and colleagues utilized a mouse model where both ATRX and the tumour suppressing factor p53 were absent (Seah et al., 2008). Analysis of newborn mice lacking both ATRX and p53 expression, displayed decreased brain size compared to littermate matched controls (Seah et al., 2008), suggesting that ATRX functions within multiple mechanisms to regulate proper brain development.

It has recently been reported that ATRX is also required in neural progenitor cell

(NPC) development and maintenance. Loss of ATRX from the mouse forebrain was found to negatively affect cortical layer development in the cerebral cortex.

Layer VI of the cortex was found to be inflated, whereas layers II-IV were found to be reduced with the loss of ATRX (Ritchie et al., 2014). This abnormality in cortical development was determined to be the result of premature cell-cycle exit of NPCs as well as a depletion of apical progenitor cells during the initial stages of neurogenesis (Ritchie et al., 2014). Taken together, these results highlight the importance for proper ATRX function during the early stages of brain development.

In addition to brain development defects, mice lacking ATRX also display additional developmental abnormalities. Mice deficient for Atrx were found to have a shortened lifespan (approximately 23 days on average), and it was resolved that this mortality rate was not due to starvation (Watson et al., 2013).

Total body weight, and body length were significantly reduced in ATRX-null mice when compared to their control counterparts. In addition, these mice also displayed kyphosis (abnormal curvature of the spine), decreased bone mineral density, loss of trabecular bone, and decreased cortical bone thickness (Watson et al., 2013). Watson and colleagues determined that Atrx-deficient mice also display a number of endocrine defects. It was determined that levels of the insulin-like growth factor 1 (IGF-1) as well as circulating thyroxine (T4) levels were significantly reduced in ATRX-null mice (Watson et al., 2013).

1.2 Myelination

1.2.1 The myelin sheath

The myelin sheath is a specialized and modified plasma membrane that encompasses axons of neurons in higher-order vertebrates. Emerging approximately 400 million years ago in cartilaginous fish, the myelin sheath allows for increased propagation of action potentials along nerve fibres, as well as providing trophic support and protection to the axonal cell body (Baumann and

Pham-Dinh, 2001; Zalc et al., 2008). It has been said that myelination of neurons accelerates nerve conduction 20-100 fold then compared to unmyelinated fibres

(Nave and Werner, 2014). Myelinated segments are separated by unmyelinated openings known as the nodes of Ranvier; the location on the axonal membrane where sodium/potassium ion (Na+/K+) exchange occurs. When an electrical current is generated, it travels along the axonal membrane where it maintains its excitatory state due to depolarization at the nodes of Ranvier. Depolarization at these nodes, gives the impression that the electrical signal is jumping from node to node, a sensation known as saltatory conduction. Limiting the abundance of sodium channels as well as extending the distance between channels, significantly reduces the amount of ATP required, thus saving the amount of energy needed to propagate action potentials (Wang et al., 2008). Interestingly, the thickness of the myelin sheath is directly proportional to the thickness of the axon; therefore the thicker the axon, the thicker the myelin sheath needs to be to allow for proper firing of action potentials (Baumann and Pham-Dinh, 2001).

1.2.2 Oligodendrocyte development

In the central nervous system (CNS) the myelin sheath is generated by oligodendrocytes (OLs), whereas the myelin sheath in the peripheral nervous system (PNS) is generated by Schwann cells.

OLs are a subpopulation of glial cell that are responsible for myelinating axons in the CNS. Mature myelinating OLs arise due to the differentiation of OL precursor cells (OPCs) (Figure 2). Multipotential NPCs are directed to become OPCs through several extrinsic factors. Ventrally derived sonic hedgehog (SHH) was found to influence NPC specification by binding to the Notch-1 receptor on the

NPC surface thereby activating the co-receptor Smoothened, which then induces oligodendrocyte transcription factor 2 (Olig2) and NK6 homeobox protein (Nkx6) expression (Pringle et al., 1996; Orentas et al., 1999). Induction of these two factors leads to the first appearance of OPCs in the telencephalon.

In addition to SHH signaling, it has been reported that fibroblast growth factor 2

(FGF2) signaling has an independent role in the transition of NPCs into OPCs

(Chandran et al., 2003; Naruse et al., 2006). It was determined that FGF2 stimulates either FGF receptor 1 or 2 (FGFR1 or FGFR2), which then activates the extracellular signal-regulated kinase 1 and 2 (ERK1 and ERK2) (Chandran et al., 2003; Furusho et al., 2011). Phosphorylation of ERK1 and ERK2 subsequently blocks SMAD signaling, causing the activation of the OL-lineage marker Olig2 (Bilican et al., 2008).

Figure 2. The oligodendrocyte development pathway

Oligodendrcoyte precursor cells (OPCs) arise in the subventricular zone of the mouse brain from the medial ganglionic eminence, lateral ganglionic eminence, and the cerebral cortex beginning at E11.5, E15, and at birth respectively (Kessaris et al., 2006). OPCs then migrate to their targeted axon where they then undergo a wave of differentiation into postmitotic immature oligodendrocytes, which then mature into myelinating oligodendrocytes (Rowitch, 2004).

OPCs initially arise in the subventricular zone of the cortex, where they undergo a wave of proliferation, followed by subsequent migration to their targeted axon

(Kessaris et al., 2006; Richardson et al., 2006). OPCs are identified by the expression of platelet derived growth factor, alpha (PDGFRα), and NG2, a protein encoded by chondroitin sulfate proteoglycan 4 (Cspg4). Both the cell- surface receptor PDGFRα, and the proteoglycan NG2 are important in maintaining the proliferative state of OPCs (Nishiyama et al., 1996; Soriano,

1997). OPC proliferation requires PDGFRα expression, as it is the receptor for the potent PDGFA mitogen (Soriano, 1997). In addition to PDGFA, it has been determined that IGF-1 may also be a factor associated with OPC proliferation.

IGF-1 regulates OPC proliferation by activating the PI3K/Akt pathway via the

IGF-1 receptor (IGF-1R). Activation of the PI3K/Akt pathway promotes the expression and survival of cyclin-D1, a known factor important in cell-cycle progression (Pang et al., 2007; Zeger et al., 2007; Bibollet-Bahena and Almazan,

2009; Romanelli et al., 2009).

Migration of OPCs to their targeted axons relies on various mechanisms including motogenic factors, and cell adhesion molecules. PDGFA and FGF-2 are two of the most important motogenic factors that influence OPC migration.

PDGFA affects OPC migration by activating the phosphorylated version of the

Wasp Veprolin-homologous protein 2 (WAVE2) family of migratory proteins through the Fyn, and cyclin-dependent kinase 5 (Cdk5) pathways (Miyamoto et al., 2008). On the other hand, FGF-2 independently promotes migration through

17 its recognition and stimulation of FGFR1 on OPCs (Bansal et al., 1996;

Osterhout et al., 1997; Bribián et al., 2006).

Cell adhesion molecules also play a vital role in promoting migration of OPCs to their targeted axon. As OPCs migrate throughout the extracellular matrix (ECM), they come into contact with various molecules that further aid their movement. It has been shown that molecules such as laminin, fibronectin, merosin, tenascin-

C, and anosmin-1 are all pro-migratory factors that assist OPC progression throughout the forebrain during development (reviewed in Mitew et al., 2014).

OPCs require several mechanisms acting in unison to properly migrate throughout the forebrain and through the ECM to reach their targeted axon. Upon reaching their final destination, OPCs undergo terminal differentiation into postmitotic immature premyelinating OLs (iOLs) (Rowitch, 2004).

Differentiation of OPCs into iOLs requires several intrinsic and extrinsic factors that act to promote differentiation. It has been shown that OL-specific factors such as achaete-scute homolog 1 (Ascl1) and NK2 homeobox protein 2 (Nkx2.2) are critical for OPC differentiation, as loss of these factors results in a substantial decrease in the number of differentiated iOLs present in the forebrain (Qi et al.,

2001; Sugimori et al., 2008). Furthermore, OLIG2 has been found to activate genes that are required for OPC differentiation. OLIG2 regulates this process by promoting binding of the BRM/SWI2-related gene 1 (BRG1) chromatin remodeling protein to the regulatory regions of genes known to be required for

OPC differentiation including SRY-box 10 (Sox10), and zinc finger homebox 1b

(Zfhx1b) (Yu et al., 2013). Several additional regulators that positively influence

OPC differentiation have been discovered including triiodothyronine (T3). Studies have shown that the abundance of postmitotic iOLs is severely diminished when

T3 levels are reduced, resulting in severe hypomyelination (Mitew et al., 2014).

Although the influence of T3 in the promotion of OPC differentiation is well known, the exact mechanisms influencing differentiation remain unclear.

There are several factors that negatively influence differentiation of OPCs as well. A long known inhibitor of OPC differentiation is the leucine-rich immunoglobulin domain-containing, 1 (LINGO-1) (Mi et al., 2004; 2005). LINGO-

1 negatively regulates OPC differentiation through two independent mechanisms.

First, LINGO-1 negatively regulates the Fyn kinase signaling pathway, which consequently upregulates RhoA signaling which has been found to reduce differentiation capacity (Mi et al., 2005). In addition, LINGO-1 also binds to OPC- specific LINGO-1, which has been shown to negatively regulate OPC differentiation (Jepson et al., 2012). An additional well-known inhibitor of OPC differentiation is the G-protein coupled receptor 17 (GPCR17). It has been shown that overexpression of GPCR17 significantly reduces the differentiation capacity of OPCs by increasing the expression of inhibitor of DNA-binding protein 2 and 4

(ID2 and ID4) proteins (Chen et al., 2009); two known factors in the promotion of cell-cycle progression (Iavarone et al., 1994). Conversely, depletion of GPCR17 has been shown to promote premature differentiation, thus displaying the relevance of GPCR17 function during OL-development (Chen et al., 2009).

Myelination of neurons requires maturation of immature premyelinating OLs into myelinating OLs. Initially, OLs extend membranous processes that contact axonal cell bodies resulting in the initiation of myelination. These membranous processes, also known as filopodia, are abundant in microfilament fibres. Several axonal signaling molecules such as laminin-α2 promote the extension of these processes (Morissette and Carbonetto, 1995; Colognato et al., 2002). Binding of laminin-α2 to the OL-specific β-1 receptor promotes the dephosphorylation of the

Fyn tyrosine kinase leading to the activation of integrin-linked kinase (ILK), particularly interesting new cysteine-histidine-rich protein (PINCH), and α-parvin.

Activation of these factors allows for the initiation of actin polymerization and consequently, filopodia extension (Hu et al., 2009).

Inhibitory factors of myelination initiation have also been well studied.

Polysialylated neural cell adhesion molecule (PSA-NCAM), as well as Jagged-1 signaling through the Notch-1 receptor are well known factors that inhibit OL process extension and contact to the axonal cell body (Wang et al., 1998;

Charles et al., 2000). In addition, LINGO-1 binding to the Nogo receptor on OLs also negatively regulates myelination as it reduces the extension abilities of OL filopodia (Mi et al., 2004; 2005).

After the initial contact of the cytoplasmic membranous processes to the unmyelinated axon, the generation of the growing myelin sheath requires several myelin-specific factors that act to maintain sheath stability, compaction, and thickness. The regulatory mechanisms of myelin-specific gene expression have yet to be completely identified; however, there are several transcription factors

20 that have been found to be important for myelin-specific gene expression and consequently, myelin sheath formation and maintenance.

One such factor that has been found to regulate myelin-specific gene expression is myelin regulatory factor (MRF). MRF was first identified as a transcriptional regulator of myelin-specific gene expression by Emery and colleagues (Emery et al., 2009). MRF is a nuclear protein that contains a DNA binding domain homologous to the Ndt80 yeast transcription factor, and is expressed exclusively in postmitotic OLs (Cahoy et al., 2008; Emery et al., 2009). Emery and colleagues determined that loss of MRF from OLs results in a dramatic decrease in the expression of several myelin-specific proteins including myelin basic protein (MBP), proteolipid protein (PLP), myelin-associated glycoprotein (MAG), and myelin oligodendrocyte glycoprotein (MOG) (Emery et al., 2009). On the other hand, induced expression of MRF in cultured OPCs stimulated the premature expression of these proteins leading to premature myelin formation, suggesting that MRF is directly regulating the expression of these genes (Emery et al., 2009). In addition, MRF has been noted to be important for myelin sheath maintenance, as ablation of MRF from myelinating OLs in 8-week-old mice correlated with severe demyelination (Koenning et al., 2012). Interestingly, Mrf expression was found to be directly regulated by the OL-specific factor SOX10 as it binds to an intronic enhancer in the Mrf transcript (Hornig et al., 2013). MRF has been found to be a potential master regulator of myelin-specific gene expression; however, the exact mechanism of MRF has yet to be elucidated.

In addition to MRF, a transcription factor found to be important in myelin-specific gene expression is the putative transcription factor zinc finger protein 191

(ZFP191). It was determined that loss of ZFP191 results in a severe hypomyelination phenotype in the postnatal day 14 (P14) mouse brain (Howng et al., 2010). Transmission electron microscopy analysis of extracted spinal cord, revealed that ZFP191 mutants displayed a global reduction of myelin correlating with the significant reduction in myelin-specific protein expression (Howng et al.,

2010). Interestingly, it was determined that loss of ZFP191 did not affect OPC production, proliferation, or differentiation into premyelinating OLs suggesting that it is only required for iOL maturation and myelination initiation. Furthermore,

Howng and colleagues determined that ZFP191 also acts in an OL cell- autonomous manner to regulate myelin-specific genes, similar to MRF (Howng et al., 2010). It is strongly suggested that additional regulatory factors are required in the molecular network that controls the expression of myelinating factors; however, the majority of these factors have yet to be identified.

1.2.3 Composition of the myelin sheath

The myelin sheath is a complex, lipid-rich membrane that is assembled and maintained by the intricate interaction of proteins and lipids. It has been established that in the CNS, 15-30% of the dry weight of myelin consists of myelin-specific proteins, and 70-85% consists of lipids (Baumann and Pham-

Dinh, 2001; Quarles, 2005). Interestingly, this proportion of protein to lipids is unique and specific to myelin.

1.2.3.1 Myelin-specific proteins

Two of the most prominent and vital proteins present within the myelin sheath are

MBP and PLP. The MBP protein consists of several different isoforms that are the result of an alternative splicing event of the Mbp gene. Mbp is located on chromosome 18 in both the mouse and human genome, consisting of seven exons and spanning 32 kb and 45 kb respectively (Roach et al., 1983; Saxe et al., 1985; Sparkes et al., 1987; Gogate et al., 1994). Two of the most prevalent isoforms (approximately 95%) of MBP are the 14 and 18.5 kDa isoforms in mice, and the 17.2 and 18.5 kDa isoforms in humans (Staugaitis et al., 1990).

MBP is a crucial factor in myelination as it is required for the formation, stabilization, and maintenance of the multilameller structure of myelin

(Campagnoni and Skoff, 2001). MBP is involved in the tight compaction of the myelin sheath as it found between, and pulls together two negatively charged surfaces of the lipid membrane forming the major dense line of the myelin sheath

(Harauz et al., 2009; Min et al., 2009; Aggarwal et al., 2013). It has been found that mice lacking MBP display significant myelin abnormalities. Mice lacking MBP display severe dysfunction in the compaction of the myelin sheath, demonstrating the necessity of MBP in myelin sheath stability and functionality (Privat et al.,

1979; Roach et al., 1985). MBP is essential in proper brain development as it is a limiting factor in CNS myelination.

In addition to MBP, PLP is also a known factor that is vital to the compaction and stabilization of the myelin sheath. PLP is a 20 kDa protein that is encoded by

Plp1, a small (15 kb) gene located on the X-chromosome. PLP is a tetraspan protein that contains four hydrophobic α-helices that span the lipid bilayer

(Weimbs and Stoffel, 1992). Interestingly, the N-terminus and C-terminus of PLP are found on the cytoplasmic sides of the lipid bilayer, which acts to stabilize the myelin sheath (Weimbs and Stoffel, 1992; Boison et al., 1995). It has been shown that mice lacking PLP, display severe axonal damage due to severe hypomyelination (Quarles and Macklin, 2005). PLP is not only an important factor for the stabilization and maintenance of the myelin sheath, but it is also crucial for providing protection to the axonal cell body. Together, MBP and PLP constitute

80% of the total proteins in myelin sheath composition (Baumann and Pham-

Dinh, 2001). MBP and PLP are the two most prevalent proteins found in the myelin sheath; however there are additional proteins that are vital for proper myelination in the CNS, such as MAG, and MOG.

MAG is a cell-adhesion glycoprotein that is located on the periaxonal glial membrane of the myelin sheath. Biochemical analysis of the MAG protein shows it has high homology to neural cell-adhesion molecule (NCAM) (Lai et al., 1987).

Due to its location as well as its homology to NCAM, it has been postulated that

MAG may be an important factor in myelin sheath initiation. This theory is strengthened as loss of MAG has been shown to disrupt glial-axonal contact in

24 mice (Li et al., 1998). In addition, in vivo mouse studies have shown that CNS myelination is delayed in the absence of MAG (Montag et al., 1994; Bartsch et al., 1997). These studies strongly support the notion that MAG is important in myelin sheath development. Although the exact function of MAG is still being debated, it has been noted to also be important for OL differentiation, as well as myelin sheath maintenance and survival (Quarles, 2007).

MOG is a transmembrane glycoprotein that is found on the outermost surface of the myelin sheath (Pham-Dinh et al., 1993). MOG has been found to be involved in the completion and maintenance of the myelin sheath, as well as being important for cell-cell communication and mediation of homophilic cell-cell adhesion (Roth et al., 1995; Baumann and Pham-Dinh, 2001; Clements et al.,

2003).

1.2.3.2 Lipid content of the myelin sheath

In addition to myelin-specific proteins, the CNS myelin sheath is composed of a high content of lipids. It has been found that the myelin sheath is comprised of cholesterol, phospholipids, galactolipids, and plasmologens at a ratio of 2:2:1:1 respectively (Norton and Poduslo, 1973). Cholesterol is a major constituent of the myelin sheath as it constitutes approximately 26% of the dry weight of myelin

(Norton and Cammer, 1984). Previous studies have found that cholesterol in

CNS myelin is synthesized locally and is independent of cholesterol levels in

25 circulation (Jurevics and Morell, 1995). The myelin sheath in the CNS displays an unusually high content of cholesterol, as it is 2-fold more abundant in CNS myelin compared to any other plasma membrane. The high level of cholesterol affects myelin compartmentalization and intracellular trafficking by influencing membrane thickness and fluidity (Nave and Werner, 2014). Previous in vivo studies have shown that, OLs that cannot synthesize cholesterol display a delay in myelin production, signifying that cholesterol biosynthesis is a rate-limiting step in CNS myelination (Saher et al., 2005).

1.2.4 Pathologies related to myelin sheath dysfunction

There are several known neurodegenerative, neurodevelopmental, and neuropsychological disorders associated with defects in myelination. The most familiar disorder associated with myelin sheath dysfunction is multiple sclerosis

(MS). MS is a neurodegenerative disorder that is the result of demyelination and axonal degeneration. The mechanistic origin of MS remains controversial, as both environmental factors and viral infection have been determined to promote disease onset (Ascherio and Munger, 2007; Compston and Coles, 2008).

Although the origin of MS remains debatable, the disease phenotype is characterized by the formation of sclerotic plaques. Sclerotic plaque formation results in severe inflammation causing demyelination, OL death, astrocytosis, and subsequent axonal degeneration (reviewed in Compston and Coles, 2008).

Although, the myelin sheath is able to partially regenerate, it is unable to maintain

26 its overall integrity, thus leading to permanent demyelination and axonal degeneration. Degradation of axons due to demyelination in MS patients have been found to lead to severe cognitive defects, tremors, reduced visual acuity, spasticity, and over weakness of the body eventually leading to premature death

(reviewed in Compston and Coles, 2008). To date, therapies addressing MS focus on targeting the immune system to suppress the advancement of demyelination of neurons rather than focusing on the mechanisms to induce remyelination (Nave and Werner, 2014). In addition to MS, myelination dysfunction has been characterized in several leukodystrophies, as well as schizophrenia. Leukodystrophies are a family of disorders that arise due to specific genetic mutations resulting in demyelination of CNS axons.

Pelizaeus-Merzbacher disease (PMD) is a member of the leukodystrophy family that results from mutations within the Plp1 gene causing abnormal levels of the

PLP protein (OMIM#312080). Resulting mutations in Plp1 cause severe demyelination in multiple areas of the brain including the cerebral cortex and cerebellum (Seitelberger, 1995). Boys affected with PMD display a severe deterioration in intellectual function, impaired movement, ataxia, and spasticity

(Inoue, 2005). Current therapies for PMD patients target treatment for seizures and spasticity, in addition with physical therapy (Xia and Wang, 2013); however, therapies focusing on targeting the molecular mechanisms of PMD have yet to manifest.

An additional and rare leukodystrophy that is well known is Krabbe disease (KD).

KD is the result of mutations in the gene that codes for the lysosomal enzyme

27 galactocerebrosidase (GALC) (OMIM#245200). Abnormal GALC levels results in a failure to breakdown galactosylceramide and psychosine leading to the degeneration of the myelin sheath (Suzuki, 2003a; 2003b). Current treatments include introduction of chemical chaperones to reactivate defective enzymes

(Berardi et al., 2014); however, no cure has been found.

Recent studies have shown that myelination defects may also be a contributing factor to the disease pathology in individuals with schizophrenia. It was determined that several schizophrenic patients displayed a disruption in both oligodendrocytes and myelination (reviewed in Haroutunian et al., 2014).

Furthermore, expression of genes involved in oligodendrocyte development and myelination was altered in extracted tissue from deceased schizophrenic patients. Genetic alterations resulting in myelination defects, coincide with several neurological and behavioural symptoms seen in individuals with schizophrenia (reviewed in Haroutunian et al., 2014). Experimental procedures performed on animal models of schizophrenia, revealed that moderate changes in OL-associated genes resulted in behaviors similar to that seen in human schizophrenic patients (Gregg et al., 2009), suggesting that abnormal OL development and myelination may be a cause, or result of schizophrenia.

Abnormal OL development and myelination is associated with several neurodevelopmental, neurodegenerative, and neuropsychological disorders; however, the exact mechanisms involved have yet to be fully elucidated, thus restricting the identification of new, improved, and less invasive therapies for these disorders.

1.3 Hypothesis and thesis objectives

It has been recently discovered that in addition to the severe developmental abnormalities, ATR-X syndrome patients also display moderate to severe myelination defects that correlate with diminished cognition (Wada et al., 2013).

As ATRX has been implicated in the regulation of the epigenetic environment of heterochromatin, and gene expression;

I hypothesize that the ATRX chromatin remodeling protein is required for proper myelin formation in the CNS by regulating the expression of myelin- specific genes.

To test this hypothesis, I set out to: (1) determine if myelination is disrupted in the absence of ATRX from the mouse forebrain, (2) determine at what stage in the oligodendrocyte development pathway ATRX is required to initiate myelination, and (3) determine if ATRX has an oligodendrocyte-specific mechanism in CNS myelination.

Figure 3. Hypothetical mechanism of ATRX in myelination of neurons within the CNS

I hypothesize that ATRX directly regulates the expression of myelin-specific genes within oligodendrocytes that code for proteins that are required for oligodendrocyte process extension, myelin sheath initiation, as well as myelin sheath compaction and stabilization

2 Materials and Methods

2.1 Animal husbandry, tissue harvesting, and genotyping

All mice were housed in clear cages at 22±1°C and subjected to a 12/12 hour light/dark cycle. All mice were fed a standard diet (Harlan Laboratories) and a water bottle was placed in each cage so water was available ad libitum. All mice were housed, bred, euthanized, and tissues harvested in a facility designed to maintain mice.

In this study, multiple mouse lines were used. Female mice harbouring loxP sites flanking exon 18 of the Atrx gene (Atrxf/f) were kindly provided by Dr. Douglas

Higgs (Weatherall Institute of Molecular Medicine, John Radcliffe Hospital,

Oxford, United Kingdom). Deletion of exon 18 from the Atrx gene results in a truncated and unstable mRNA transcript, thus becoming vulnerable to degradation and ultimately eradicating the presence of the ATRX protein.

Recombination of exon 18 from the Atrx gene mirrors the effect of an ATRX-null mutation (Bérubé et al., 2005).

Heterozygous female mice containing loxP sites flanking exon 18 of the Atrx gene (Atrxf/wt) were crossed with male mice that have Cre recombinase inserted into the forkhead box G1 (FoxG1) locus, thus allowing for the complete loss of

Atrx expression specifically from the forebrain beginning at E8.5. Male progeny resulting from this cross will be used to assess the role of

Figure 4. Schematic of mice harbouring loxP sites used for Cre-loxP recombination.

The top schematic represents the wild-type allele with normal Atrx expression. The second schematic represents the allele that has loxP sites flanking exon 18 of the Atrx gene. The third schematic represents the result of recombination using Cre-loxP technology resulting in complete loss of exon 18 of the Atrx gene. The white bars represent exons of the Atrx gene, and the black triangles represent loxP sites located in the Atrx gene.

ATRX in the mouse forebrain, as female mice heterozygous for Atrx expression are phenotypically normal. Male mice that have Atrx knocked out from the forebrain (AtrxFoxG1Cre) were used as experimental mice, and male mice that express Cre recombinase but do not contain loxP sites flanking exon 18 of the

Atrx gene were used as control mice.

An additional mouse line was utilized to determine if ATRX has an OL-specific role in myelination development. Male mice that have Cre recombinase inserted into the 2’, 3’-cyclic-nucleotide 3’-phosphodiesterase (Cnp) locus were generated by Dr. Klaus-Armin Nave (Max Planck Institute for Experimental Medicine,

Göttingen, Germany) (Lappe-Siefke et al., 2003), and generously provided by Dr.

Brian Popko (University of Chicago, Chicago, Illinois, U.S.A) with the permission from Dr. Nave. These male mice were mated with heterozygous female mice containing loxP sites flanking exon 18 of the Atrx gene (Atrxf/wt). The resulting male progeny (AtrxCnpCre) from this cross lack ATRX specifically from the OL cell- lineage beginning at E11.5 (Lappe-Siefke et al., 2003).

To determine the genotypes of our mice, genomic DNA was extracted from tail clips/ear notches from each mouse by incubating these samples in a solution of

100µL of DirectPCR and 5µL of proteinase K (ThermoScientific) for 12 hours at

55°C. PCR was then performed using primers designed to amplify the Atrx, Cre, and Sry genes under the following conditions: Step 1. Initial denaturation - 95°C for 3 minutes, Step 2. Denaturation - 95°C for 30 seconds, Step 3. Annealing -

55°C for 30 seconds, Step 4. Elongation - 72°C for 1:30 minutes (Note: Steps 2-4 were performed for 35 cycles), followed by Step 5. 72°C for 5 minutes. Primer

35 designs can be located in Table 1. Following PCR amplification, each sample was then ran on a 1.5% agarose gel for separation, and then imaged using the

BioRad Universal Imaging hood (BioRad).

The mice chosen for either quantitative real-time PCR (qRT-PCR) or western blot analysis were euthanized using carbon dioxide (CO2) inhalation followed by cervical dislocation. The mice chosen for immunohistological analyses were euthanized using CO2 inhalation, and fixed by cardiac perfusion. Cardiac perfusion was performed by first injecting 10mL of 1x phosphate buffered saline

(PBS), followed by injection of a 30mL solution of 4% paraformaldehyde (PFA).

Following cardiac perfusion, whole mouse brains were harvested and placed in a

15mL conical tube containing 10mL of 4% PFA. Post-fixation of whole mouse brains was carried out for 12 hours at 4°C. Once post-fixation was complete, whole mouse brains were then transferred to a solution of 30% sucrose diluted in

1xPBS for cryoprotection. Following cryoprotection, whole mouse brains were embedded in OCT (optical cutting temperature) media and flash frozen using liquid nitrogen. Brain tissues were then stored in a -80°C freezer until further use.

2.2 Microarray analysis

To assess gene expression changes between AtrxFoxG1Cre and control mouse brains at both postnatal day 0.5 (P0.5) and P17, a gene expression microarray was performed by a member of the laboratory, Dr. Michael Levy (unpublished).

RNA was extracted from the forebrain of three pairs of P0.5 AtrxFoxG1Cre and control mice, as well as three pairs of P17 AtrxFoxG1Cre and control mice using the

RNeasy Mini Kit (QIAGEN). Total RNA was then sent to the London Regional

Genomics Centre (LRGC) for microarray analysis. First, cDNA was synthesized using SuperScriptII (Invitrogen) and oligo (dT24) primers. Biotin-labeled cRNA was then prepared by cDNA in vitro transcription using the BioArray High-Yield

RNA Transcript Labeling kit (Enzo Biochem) to incorporate biotinylated UTP and

CTP. Biotinylated labeled cRNA was then hybridized to an Affymetrix Mouse

Genome 430 2.0 array chip for 16 hours at 45°C and 60rpm. One microarray chip per mouse was used. GeneChips were stained with streptavidin-phycoerythrin

(SAPE), followed by an antibody solution and a second SAPE solution, with all liquid handling performed by a GeneChip Fluidics Station 450. GeneChips were scanned with the Affymetrix GeneChip Scanner 3000 7G (Affymetrix) where the

GCOS1.4 program was used to generate probe level (.CEL file) data.

Probe level data was then transferred into the Partek Genomic Suite program

(Partek), and the RMA algorithm (Irizarry et al., 2003) was used to summarize each probe signal, and output the data as probe intensity levels. Summarizing the probe signal utilizing the RMA algorithm consists of, binning the data to reduce the variation within each group, background correction, and normalization of the data from each array chip to each other. Once the probe intensity levels have been determined, a one-way ANOVA was used to calculate the p-values for each probe set. Furthermore, gene expression fold-changes in AtrxFoxG1Cre mouse brains compared to control mouse brains were calculated using the

Fisher’s least significant difference model (Tamhane and Dunlop, 2000). Gene ontology (GO) analysis of specific gene subsets was performed using the

Fisher’s exact test (Rhee et al., 2008) in the Partek Genomic Suite program

(Partek). Heat maps of selected gene targets were generated using respective probe intensity levels from both AtrxFoxG1Cre and control mice.

2.3 Quantitative real-time PCR

Total RNA was isolated from control and knockout mouse forebrains using the

RNeasy Mini Kit (QIAGEN). Quantity and quality of isolated RNA was then measured using the Nanodrop1000 (Thermo Scientific). RNA was then reverse transcribed into cDNA using standard PCR settings (65°C for 10 minutes, followed by 30°C for 10 minutes, then 42°C for 45 minutes). Confirmation of reverse transcription was carried out by PCR using primers designed to the internal control, glyceraldehyde 3-phosphate dehydrogenase (Gapdh) (Table 1).

PCR amplification was performed using standard settings. PCR samples were then loaded onto a 1.5% agarose gel and ran at 95V for 22 minutes, and then imaged. qRT-PCR was carried out to study gene expression changes between

AtrxFoxG1Cre and control mouse forebrains. First, the online Ensembl genome browser 84 database (http://www.ensembl.org/index.html) was used to identify optimal gene

Table 1. Primer sequences for PCR and qRT-PCR Gene Forward primer (5’ to 3’) Reverse primer (5’ to 3’)

Mbp ATCCAAGTACCTGGCCACAG CAGGGAGCCATAATGGGTAG

Mog AAATGGCAAGGACCAAGATG AAGTGCGATGAGAGTCAGCA

Mag GCCTGTACCTGGATCTGGAG ATGACCGTGGCTAGGATCTG

Plp1 GGCCACTGGATTGTGTTTCT GTGGTGTAGAAGCCCTCAGC

Mobp GAGGAGGACTGGATCTGCTG AGTCCTTGAAGGGGACCTTG

Cnp GCTGGTTCCTGACCAAAAAG CACCTGGAGGTCTCTTTCCA

Pdgfrα ACCTCCCACCAGGTCTTTCT GCCAGCTCACTTCACTCTCC

Cspg4 TCCAGATCACTGGGCCTTAC AGGCGTCTGTCTGTGTCTCA

18 R - TGAACCTGGGGACTTCTTTG Atrx AGAACCGTTAGTGCAGGTTCA NeoR - CCACCATGATATTCGGCAAG

Cre TGACCAGAGATCATCCTTAGCG AATGCTTCTGTCCGTTTGCC

Sry GCAGGTGGAAAAGCCTTACA AAGCTTTGCTGGTTTTTGGA

Gapdh CAACGACCCCTTCATTGACCT ATCCACGACCGACACATTGG

locations to design DNA primers for PCR amplification. Using the desired gene locations, the online Primer3 tool (http://bioinfo.ut.ee/primer3-0.4.0/) was used to design optimal DNA primers. Primer sequences for each gene can be found in

Table 1. qRT-PCR was carried out on 96-well PCR plates. Each well on the 96- well plate contained a combined solution of cDNA, H2O, the desired primers, and iQT,MSYBR® Green Mastermix (BioRad). qRT-PCR was carried out using a

Chromo-4 thermocycler (BioRad) under standard conditions [Step 1. Initial denaturation - 95°C for 3 minutes, Step 2. Denaturation - 95°C for 10 seconds,

Step 3. Annealing - 55°C for 20 seconds, Step 4. Elongation - 72°C for 30 seconds (Note: Steps 2-4 were performed for 30-35 cycles), followed by Step 5.

Final elongation - 72°C for 5 minutes]. A final melting curve was constructed in increments of 1°C per plate read. Opticon Monitor 3 sofware (BioRad) was used to calculate Ct values for each sample. Ct values for each well were calculated automatically after resolving the threshold placement on the amplification curve.

Ct values were then transferred into the GeneX (BioRad) program to analyze the gene expression data. Expression of each gene was calculated from two technical replicates and three biological replicates. Gene expression was normalized to Gapdh, and an unpaired students t-test was then used to determine if expression of each gene was significantly different (p<0.05) between control and knockout mice.

2.4 Protein isolation

Total protein was extracted from control and knockout mouse forebrains using

RIPA lysis buffer (150mM NaCl, 1% NP-40, 50mM Tris, pH 8.0, 0.5% deoxycholic acid, 0.1% SDS, 0.2mM PMSF, 0.5mM NaF, 0.1mM Na3VO4, and 1 protease inhibitor cocktail tablet). 200µL of RIPA buffer was added to a 1.5mL

PCR tube containing one half of a mouse forebrain. Forebrain tissue was then homogenized and put on ice for 20 minutes for cell lysis. Total cell extracts were cleared by centrifugation at 10000rpm at 4°C for 20 minutes to pellet cellular debris, and then the supernatant containing the soluble cellular fraction was collected. Protein quantification was then measured by Bradford assay (Bradford,

1976). Bradford assay was carried out by first adding 10µL of each sample into a

96-well plate. 190µL of protein assay dye reagent concentrate (BioRad) was then added to each well. 0.5µg/µL, 1µg/µL, 2 µg/µL, 4µg/µL, 6µg/µL, and 8µg/µL of bovine serum albumin (BSA) was used a controls. The 96-well plate was then loaded into the Wallac Victor2, 1420 multilabel counter machine (Perkin Elmer) where the absorbance of each individual sample was recorded. Each sample was performed in triplicate, and therefore the average absorbance was used to calculate the protein concentration of each sample. An X-Y scatter plot was constructed placing the control BSA concentration values on the Y-axis, and the corresponding wavelength values on the X-axis. A line of best fit was then calculated in excel to determine the equation of the line. The measured wavelength of each sample was inputted into this equation to determine their subsequent protein concentration.

2.5 Western blot analysis

Western blots were carried out using polyacrylamide gels. Each polyacrylamide gel consisted of a 4% stacking gel (780µL ddH2O, 130µL 30% Acrylamide-Bis

(Bio-Rad), 65µL 2M Tris (pH 6.8), 10µL 10% SDS, 8µL 10% APS, 2µL TEMED for a 1mL solution) and a separating gel. In this study, both a 6% separating gel

(5.9mL ddH2O, 2mL 30% Acrylamide-Bis (Bio-Rad), 2mL 2M Tris (pH 8.8),

100µL 10% SDS, 30µL 10% APS, 20µL TEMED for a 10mL solution) and a 12% separating gel (3.9mL ddH2O, 4mL 30% Acrylamide-Bis (Bio-Rad), 2mL 2M Tris

(pH 8.8), 100µL 10% SDS, 30µL 10% APS, 20µL TEMED for a 10mL solution) were used depending on the size of the desired proteins. A total of 8µL of protein ladder (Bio-Rad), and 50ug of protein was loaded into the corresponding wells.

SDS-PAGE gels were run at 90V for 30 minutes, followed by 130V for 2 hours, in

1x running buffer. Protein from the gels was then transferred onto a nitrocellulose membrane (Pall Life Sciences) at 75V for 2 hours in 1x transfer buffer.

Nitrocellulose membranes were then placed into a small plastic dish and blocked with 5% milk-TBST for 1 hour at room temperature. Following this incubation, the desired primary antibody (Table 2) was then diluted in 5mL of 5% milk-TBST and added to the dish containing the membrane, and then incubated overnight at

4°C. The next morning, membranes were place on a rocker where they then underwent 4, 10-minute washes with 1x TBST at room temperature. The corresponding secondary antibody (Table 3) was then diluted in 5mL of 5% milk-

TBST and added on top of the membrane, and then incubated for 1 hour at room temperature. Membranes then underwent another 4, 10-minute washes with

Table 2. Primary antibodies used for Western blot analysis and immunofluorescence staining

Dilution of Dilution of Protein Antibody Description secondary primary antibody antibody

ATRX (D-5) Anti-mouse, polyclonal (Santa Cruz) WB – 1:500 WB – 1:4000

ATRX (H300) Anti-rabbit, polyclonal (Santa Cruz) IF – 1:100 IF – 1:800

IF – 1:200 IF – 1:800 MBP Anti-rat, monoclonal (Abcam) WB – 1:3000 WB – 1:4000

IF – 1:200 IF – 1:800 MOG Anti-mouse, monoclonal (Millipore) WB – 1:3000 WB – 1:3000

MAG Anti-mouse, monoclonal (Abcam) WB – 1:000 WB – 1:4000

CNPase Anti-mouse, monoclonal (Covance) WB – 1:3000 WB – 1:4000

IF – 1:200 IF – 1:800 PDGFRα Anti-rabbit, polyclonal (Abcam) WB – 1:200 WB – 1:4000

APC (CC-1) Anti-mouse, monoclonal (Millipore) IF – 1:100 IF – 1:800

Anti-mouse, monoclonal (Santa OLIG1 IF – 1:100 IF – 1:800 Cruz)

OLIG2 Anti-goat, polyclonal (Santa Cruz) IF – 1:100 IF – 1:800

Actin Anti-rabbit, polyclonal (Sigma) WB – 1:1000 WB – 1:4000

INCENP Anti-rabbit, polyclonal (Sigma) WB – 1:10000 WB – 1:10000

Table 3. Secondary antibodies used for Western blot analysis and immunofluorescence staining

Experimental procedure Secondary antibody and description used

Anti-rat IgG HRP (Santa Cruz) Western blot

Anti-mouse IgG HRP (Santa Cruz) Western blot

Anti-rabbit IgG HRP (Santa Cruz) Western blot

Alexa Fluorophore, goat anti-rabbit IgG 594 nm (Life Technologies) Immunofluorescence

Alexa Fluorophore, goat anti-rat IgG 488 nm (Life Technologies) Immunofluorescence

Alexa Fluorophore, goat anti-mouse IgG 594 nm (Life Technologies) Immunofluorescence

Alexa Fluorophore, donkey anti-goat IgG 594 nm (Life Technologies) Immunofluorescence

1xTBST at room temperature. ECL (enhanced chemilluminescence) was then added to the membrane for approximately 1 minute. The membrane was then exposed onto chemilluminescence film, and then developed with the Konica

Minolta SRV-101A film developer (Konica Minolta). Protein expression was measured by calculating the integrated density of each protein band using

ImageJ software. Protein expression was normalized to the corresponding loading control. INCENP was used as the loading control for proteins that were detected using 6% polyacrylamide gels, and Actin was used as the loading control for proteins that were detected using 12% polyacrylamide gels. Unpaired students t-test was then used to determine if the expression of the protein was significantly different (p<0.05) between control and knockout mice.

2.6 Immunofluorescence staining and cell counting

Embedded whole mouse brains were coronally sectioned (Figure 5) to a thickness of 8µm using the Leica CM100 cryostat (Leica Biosystems) and placed on glass microscope slides. Slides were then placed in a -80°C freezer until further use. 1 hour prior to starting the immunofluorescence protocol, slides containing the frozen brain cryosections were taken out of the freezer to allow for thawing. Once the samples were completely thawed, slides were placed into a

1xPBS solution for 5 minutes to allow for sample rehydration. Slides were then placed into a slide holder containing a pre-warmed solution of 10mM sodium

Figure 5. Coronal section of the postnatal mouse brain

Analysis of the postnatal mouse brain was performed by assessing two areas of the brain that are rich in white matter and are important for proper cognitive function; the corpus callosum and the cerebral cortex (outlined in red boxes).

47 acetate for antigen retrieval. Antigen retrieval was carried out by slightly heating slides in a microwave for 10 minutes. Slides were then taken out, and cooled at room temperature for 20 minutes. Slides were then washed once with

1xPBS, and twice with 1XPBS/0.3% Triton-X. Slides were then placed into a staining chamber. Primary antibodies (Table 2) were diluted in a 1xPBS/0.3%

Triton-X/1% BSA solution, and then 200µL of this solution was added to each slide. Slides were incubated overnight at 4°C. The following morning, slides underwent 3, 5-minute washes with 1xPBS/0.3%Triton-X. Secondary antibodies

(Table 3) were diluted in a 1xPBS/0.3% Triton-X/1% BSA solution, and then

200µL of this solution was added to each slide. Slides were then incubated in the dark at room temperature for 1 hour. Following this incubation, slides underwent

3, 5-minute washes with 1xPBS/0.3%Triton-X. 500µL of DAPI was then added to each slide, and incubated for 5 minutes. Slides were then washed twice with

1xPBS/0.3%Triton-X for 5 minutes each, then washed another two times with

1xPBS for 5 minutes each. Slides were then coverslipped and prepared for microscopy.

Microscopy for immunofluorescence was carried out using the Leica DMI 6000b automated inverted microscope (Leica Biosystems). Images were captured using

Openlab Software (v5.0 Improvision) and then transferred to the Volocity program for further analysis, processing, and cell counting. Using the Volocity software, the correct pixel/µm was set and scale bars were positioned onto the corresponding image. Cell counts were performed on multiple sections from each sample to acquire a global average, and performed on three to four biological

48 replicates. The percentage of cells expressing the protein of interest was determined by comparing the number of cells positive for the protein of interest to the total number of DAPI-positive cells. An unpaired students t-test was then used to determine if the percentage of cells expressing the protein of interest was significantly different (p<0.05) between control and knockout mice.

2.7 Analysis of cholesterol abundance

Cholesterol levels in the forebrains from P14 AtrxFoxG1Cre and control mice were measured by the Metabolic Phenotyping Laboratory at Robarts Research

Institute (Western University, London, Ontario, Canada). Whole mouse-brains were isolated from P14 AtrxFoxG1Cre and control mice as described above, and flash-frozen in dry ice. Brains were then transported to Brian Sutherland at the

Metabolic Phenotyping Laboratory for further processing.

Weights of each brain sample were initially recorded, followed by mincing with a scalpel and transferred to a glass tube. Tissue lipids were extracted using the

Folch extraction method (Folch et al., 1957). First, 2.2mL of methanol was added to each tube containing minced brain samples and incubated for 1 hour at room temperature, followed by the addition of 4.4mL of chloroform.

A radioactive tracer was then incorporated into each sample to allow for lipid recovery analysis. This was achieved by adding 400µL of a 0.1µCi/mL stock of

3H-Cholesterol Oleate to each sample. In parallel, 400µL of this isotope was

49 aliquoted, in duplicate, into scintillation vials to allow for counts added calculations. Counts added allows us to determine the value of 100% recovery.

Samples were then mixed with the isotope, and incubated at 4°C for 24-48 hours to achieve total lipid extraction. The samples were filtered using Whatman paper into a fresh glass tube. The original glass tube was then rinsed with 0.9mL of a

2:1 ratio of chloroform:methanol, and then filtered through Whatman paper into the new glass tubes, followed by the addition of 1.5mL saline and incubated for

1-2 hours at room temperature. The combination of saline and methanol allows for the separation of the chloroform containing the extracted lipids, from the rest of the solution. Using a Pasteur pipette, the chloroform fraction was removed and then placed into a fresh glass tube and made up to a 10mL solution with chloroform.

To calculate the abundance of cholesterol in each sample, two separate assays were performed to assess total cholesterol levels, and free cholesterol levels.

First, a solution containing a combination of Triolein and cholesterol standard, and diluted in isopropanol, was aliquoted in triplicate to calculate a standard curve for each assay. Next, 50-200µL of each sample was aliquoted, in duplicate, into separate glass tubes. Simultaneously, 0.5mL of each sample was aliquoted into scintillation vials containing scintillation fluid, to calculate recovery efficiency.

Each sample and standard were dried under nitrogen to remove the presence of chloroform. 500µL of 1% TritonX-100 was then added to each tube and incubated for 1 hour at room temperature, followed by drying under nitrogen.

50µL deionized water was added to each tube to solubilize the sample, as the

50 assays to assess cholesterol levels cannot be run on chloroform based samples.

Each sample was then placed into a 37°C water bath 15 minutes prior to running each assay.

To calculate the abundance of total cholesterol and free cholesterol, standard kits were used [Wako kits 439-17501, 435-35801 (Wako Diagnostics), respectively].

For each assay, 1mL of colour reagent solution (from the corresponding Wako kit) was added to each sample and cholesterol standard. These solutions were then mixed, and incubated for 15 minutes at 37°C. Follow this incubation, samples were cooled down at room temperature for 5 minutes. The absorbance of each sample and standard was then measured at a wavelength of 600nm using the Beckman DU 640 Spectrophotometer (Beckman Coulter).

An X-Y scatter plot was constructed placing the cholesterol standard values on the Y-axis, and the corresponding absorbance values on the X-axis. A line of best fit was then calculated in excel to determine the equation of the line. The measured absorbance of each sample was inputted into this equation to determine their subsequent concentration. Mean values of total cholesterol and free cholesterol from control and AtrxFoxG1Cre samples were calculated, and the abundance of cholesterol esters is calculated by subtracting free cholesterol from total cholesterol. Unpaired students t-test was then used to determine if cholesterol concentrations between control and AtrxFoxG1Cre samples were significantly different (p<0.05).

3 Results

3.1 Transcriptional profiling of the postnatal AtrxFoxG1Cre

forebrain reveals a decrease in myelin-specific gene

expression

The ATRX chromatin remodeling protein is a known regulator of gene expression; therefore, to examine the gene expression profile in the developing mouse forebrain in the absence of ATRX, a microarray was carried out on P17

AtrxFoxG1Cre and littermate-matched control mouse forebrains. It was determined that gene expression was significantly changed in AtrxFoxG1Cre mice compared to littermate-matched control mice if the fold-change was upregulated, or downregulated by 1.3 fold (p<0.05). Examination of the P17 microarray data revealed that ten of the top fifty-downregulated genes in AtrxFoxG1Cre mice are genes implicated in oligodendrocyte development and CNS myelination (Figure

6A). Furthermore, GO analysis of significantly downregulated genes (-1.5 fold; p<0.05) revealed that genes involved in various aspects of myelination were of the top significantly downregulated genes in the P17 AtrxFoxG1Cre forebrain (Figure

6C). To determine the total number, and identity of the myelin-specific genes that are significantly downregulated in P17 AtrxFoxG1Cre mice, the P17 microarray was compared to an RNA-sequencing transcriptome database of glia, neurons, and vascular cells of the cerebral cortex (Zhang et al., 2014). Using this database, one can determine which genes are expressed, and at what concentration in a

52 specific cell type compared to all other cell types in the cerebral cortex. Here, we sought to determine the identity of genes specifically expressed in mature myelinating OLs, as these cells are responsible for the development of the myelin sheath, and compare this gene set to the significantly downregulated genes from the P17 microarray. First, a list depicting genes enriched in myelinating OLs compared to all other neuronal cells was constructed using the RNA-sequencing database (Zhang et al., 2014). Furthermore, a 4-fold cutoff was set to narrow the set of genes that are truly specific to myelinating OLs. This set of genes was then compared to significantly downregulated genes (-1.3 fold; p<0.05) from the P17 microarray to determine how many of those downregulated genes were in fact myelin-specific genes. It was determined that 76 myelin-specific genes were significantly downregulated in P17 AtrxFoxG1Cre mice compared to their control counterparts (Figure 6B).

Validation of the gene expression changes from the P17 microarray was performed by qRT-PCR on cDNA extracted from P20 AtrxFoxG1Cre and control forebrain samples. It was determined that expression of several myelin-specific genes, was downregulated in P20 AtrxFoxG1Cre mice compared to littermate- matched controls, thereby confirming the P17 microarray data (Figure 6D).

Taken together, this data suggests that ATRX may be an important factor for proper myelin formation in the mouse brain.

Figure 6. Transcriptional profiling of genes from P17 AtrxFoxG1Cre microarray revealed a significant decrease in expression of myelin-specific genes compared to littermate-matched controls.

A) Heat-map displaying the top fifty-downregulated genes in P17 AtrxFoxG1Cre mice compared to controls. B) Heat-map displaying the top downregulated oligodendrocyte-specific genes in P17 AtrxFoxG1Cre mice compared to controls. C) GO-analysis of downregulated genes in P17 AtrxFoxG1Cre mice. D) qRT-PCR of myelin-specific genes from P20 AtrxFoxG1Cre mice. Expression of each gene was normalized to Gapdh. N=2. Error bars represent range of expression for two biological replicates.

3.2 Loss of ATRX from the mouse forebrain results in

hypomyelination

Next, we set out to determine if the decrease in myelin-specific gene expression observed in P20 AtrxFoxG1Cre mice, results in a defect in myelin sheath development. To address this matter, P20 AtrxFoxG1Cre and littermate-matched control mouse brains were processed for western blot analysis and immunofluorescence staining. To determine if loss of ATRX has a deleterious effect on myelin-specific protein expression, western blot analysis of the myelin- specific markers CNP, MOG, MBP, and MAG was carried out using protein that was isolated from P20 mouse forebrains. It was determined that protein expression of CNP, MOG, MBP, and MAG was significantly reduced in P20

AtrxFoxG1Cre forebrains compared to littermate-matched controls (Figure 7A and

7B). Additionally, immunofluorescence co-staining of MBP and MOG was carried out on whole-brain cryosections from P20 AtrxFoxG1Cre and littermate-matched control mice to analyze the abundance and expression pattern of myelin in the mouse forebrain. It was observed that expression of both MBP and MOG was significantly decreased in the corpus callosum (a white-matter-rich area of the brain) of P20 AtrxFoxG1Cre mice compared to their control counterparts (Figure

7C). This finding was also observed in the cerebral cortex, as MBP and MOG expression were both significantly reduced and sparser in P20 AtrxFoxG1Cre mice compared to littermate-matched control mice (Figure 7C). These findings

Figure 7. Expression of myelin-specific proteins is significantly reduced in P20 AtrxFoxG1Cre mice compared to littermate-matched controls.

A) Western blot of ATRX, INCENP, CNP, MBP, MOG, MAG, and Actin from forebrain samples of P20 AtrxFoxG1Cre and control mice. B) Quantitative analysis of western blot. ATRX was normalized to INCENP, whereas CNP, MBP, MOG, and MAG were normalized to Actin. N=4. Error bars represent ± standard error of the mean (±SEM). *p<0.001. C) Immunofluorescence staining of MBP and MOG in P20 AtrxFoxG1Cre and control corpus callosum and cerebral cortex. N=3. Scale bar represents 70µm. DAPI stains the nucleus of the cell.

59 illustrate that loss of the ATRX chromatin remodeling protein results in severe hypomyelination in the mouse forebrain, and therefore we can conclude that

ATRX is required for proper myelination in the CNS.

3.3 Hypomyelination in the AtrxFoxG1Cre mouse forebrain is

not due to a reduction in neuronal cell number

The hypomyelination phenotype seen in the forebrains of P20 AtrxFoxG1Cre mice may be the result of a reduction in the total number of neurons that can be myelinated. To address the possibility that the total number of neurons is decreased in P20 AtrxFoxG1Cre mice, immunofluorescence staining of the neuron- specific marker neuronal nuclei antigen (NeuN) was carried out on whole-brain cryosections of P20 AtrxFoxG1Cre and control mice. Co-staining of P20 AtrxFoxG1Cre and control mouse brain cryosections for DAPI and NeuN allowed us to determine the percentage of neurons in the mouse forebrain. Counts of DAPI- positive cells revealed that the total number of cells in a set area is equal between P20 AtrxFoxG1Cre and control mice (Figure 8A and 8B). Furthermore, it was determined that approximately 76% of DAPI-positive cells are neurons in

P20 control mouse forebrains, and this percentage of neurons was found to be equivalent in the P20 AtrxFoxG1Cre mouse forebrain (Figure 8A and 8C). As neuronal cell number is unchanged between P20 AtrxFoxG1Cre and litter-matched control mice, we can deduce that the hypomyelination phenotype observed

Figure 8. Hypomyelination is not due to a reduction in the number of neurons in the mouse brain.

A) Immunofluorescence staining of NeuN and ATRX in the cerebral cortex of P20 AtrxFoxG1Cre and control mice. N=3. Scale bar represents 70µm. B) Cell counts were performed to analyze the total number of DAPI-positive cells per unit area. C) Cell counts were performed to analyze the percentage of NeuN-positive cells compared to DAPI-positive cells. N=3. Error bars represent ±SEM. p>0.05. DAPI stains the nucleus of the cell.

62 in AtrxFoxG1Cre mice is not due to a reduction in the number of neurons, but due to a defect in the initiation of myelination by OLs.

3.4 Oligodendrocyte precursor cell production is unchanged

in AtrxFoxG1Cre mice at birth

Onset of myelination may be impeded by multiple factors including deficiencies in

OPC production, proliferation, and/or differentiation into postmitotic immature

OLs. We first wanted to assess OPC production and morphology in AtrxFoxG1Cre mice at birth. Microarray analysis performed on P0.5 mouse forebrain samples revealed that expression of two known OPC markers, Pdgfrα and Cspg4 [1.16 fold (p>0.05) and -1.11 fold (p>0.05) respectively], was unchanged in P0.5

AtrxFoxG1Cre mice compared to littermate-matched controls. To confirm that the expression of these OPC markers is unchanged when ATRX is absent, qRT-

PCR was carried out on cDNA from P0.5 AtrxFoxG1Cre and littermate-matched control mouse forebrains. It was determined that expression of both Pdgfrα and

Cspg4 was unchanged in P0.5 AtrxFoxG1Cre mice compared to controls (Figure

9A). To determine if this effect is observed at the protein level, western blot - analysis and immunofluorescence staining of the OPC marker PDGFRα, was performed. Western blot analysis revealed that protein expression of PDGFRα was unchanged between P0.5 AtrxFoxG1Cre and littermate-matched control mice

(Figure 9B and 9C). OPC production was further examined by performing

Figure 9. Expression of OPC markers is unchanged in P0.5 AtrxFoxG1Cre mice compared to littermate-matched controls.

A) qRT-PCR of Atrx, Pdgfrα, and Cspg4 from P0.5 AtrxFoxG1Cre mice. Expression of each gene was normalized to Gapdh. N=3. Error bars represent ±SEM. *p<0.001 for Atrx. p>0.05 for Pdgfrα, and Cspg4. B) Western blot of ATRX, PDGFRα, and INCENP from forebrains of P0.5 AtrxFoxG1Cre and control mice. C) Quantitative analysis of western blot. ATRX, and PDGFRα was normalized to INCENP. N=4. Error bars represent ±SEM. p>0.05. D) Immunofluorescence staining of PDGFRα in the cerebral cortex of P0.5 control and AtrxFoxG1Cre mice. Scale bar represents 70µm. DAPI stains the nucleus of the cell E) Quantitative analysis of the percentage of PDGFRα-positive cells compared to total number of DAPI-positive cells. N=3. Error bars represent ±SEM. p>0.05

65 immunofluorescence staining for PDGFRα on P0.5 whole-brain cryosections. To assess OPC production capability in P0.5 AtrxFoxG1Cre mice compared their control counterparts, cell counts for PDGFRα-positive cells were performed in the cerebral cortex, as the majority of OPCs populating the brain at birth arise from the cerebral cortex (Kessaris et al., 2006). It was determined that the percentage of OPCs in the cerebral cortex was similar between P0.5 AtrxFoxG1Cre and littermate-matched control mice (Figure 9D and 9E). These findings infer that

ATRX loss from the forebrain does not affect OPC production/maintenance, and therefore must have an alternative role in myelin sheath formation.

3.5 Differentiation of oligodendrocyte precursor cells into

immature oligodendrocytes is hindered in the absence

of ATRX

A potential mechanism that ATRX may have in the myelination process is the regulation of OPC differentiation into premyelinating iOLs. To study this possibility, immunofluorescence staining of APC and OLIG2 was carried out on whole-brain cryosections from P10 AtrxFoxG1Cre and littermate-matched control mice. It was determined that the number of APC/OLIG2-positive cells was significantly reduced in the corpus callosum of P10 AtrxFoxG1Cre mice compared to littermate-matched control mice (Figure 10A and 10B). These data

Figure 10. The number of APC/OLIG2-postive cells is significantly decreased in P10 AtrxFoxG1Cre mice compared to littermate-matched controls.

A) Immunofluorescence staining of APC and OLIG2 in the corpus callosum of P10 AtrxFoxG1Cre and control mice. B) Cell counts were performed to determine the percentage of APC/OLIG2-positive cells per DAPI-positive cells. N=3. Scale bar represents 100µm. Error bars represent ±SEM. *p<0.05. DAPI stains the nucleus of the cell.

68 strongly suggests that ATRX is required for OPC differentiation into iOLs.

Although differentiation of OPCs is impeded in the absence of ATRX, the defect in differentiation does not fully explain the severe hypomyelination phenotype seen in P20 AtrxFoxG1Cre mice; therefore, ATRX must also be required during the final stage of myelin sheath development by influencing maturation of iOLs into myelinating OLs.

3.6 Cholesterol biosynthesis is unaffected in the P14

AtrxFoxG1Cre mouse forebrain

Cholesterol biosynthesis is crucial to myelination as 26% of the myelin sheath is composed of cholesterol. Analysis of the P17 microarray revealed that several genes involved in cholesterol biosynthesis were downregulated in the P17

AtrxFoxG1Cre mouse forebrain compared to age-matched controls (data not shown). Therefore, we hypothesized that ATRX was required for cholesterol biosynthesis, as loss of ATRX may cause a decrease in cholesterol levels and consequently cause a delay in myelin formation. To study ATRX and its effects on cholesterol biosynthesis, cholesterol levels were measured in 14 day-old mice

(peak of cholesterol biosynthesis in the mouse brain) in collaboration with Brian

Sutherland at the Robarts Research Institute. This was accomplished by first, isolating lipids from the forebrain, and then using standard kits to quantify total cholesterol as well as free cholesterol levels. Total cholesterol, and free

Figure 11. Cholesterol levels are unchanged in the absence of ATRX from P14 mice.

Total cholesterol, free cholesterol, and cholesterol ester levels. N=4. Error bars represent ±SEM. p>0.05.

71 cholesterol levels were found to be unchanged in P14 AtrxFoxG1Cre mice compared to age-matched controls (Figure 11). Overall, these results suggest that ATRX does not influence myelination by regulating cholesterol biosynthesis.

3.7 Generation of a novel mouse model to study the

oligodendrocyte-specific function of ATRX

Intricate interaction between several key cell types, such as OLs, astrocytes, and neurons is necessary for proper myelin formation; therefore, a disruption in any of these cell types can lead to a hypomyelinated phenotype. Because OLs are the glial cells that myelinate neurons in the CNS, we wanted to determine if

ATRX has an OL-specific effect on myelin sheath formation. To study the function of ATRX within OLs, transgenic male mice that express Cre recombinase under control of the Cnp promoter (Lappe-Siefke et al., 2003) were mated with female mice containing loxP sites flanking exon 18 of the Atrx gene

(Bérubé et al., 2005). This cross results in the deletion of Atrx specifically from

OLs (referred to as AtrxCnpCre) beginning at approximately P5. It was determined that the proportion of oligodendrocytes defined by OLIG1-positive cells, was unchanged between P20 AtrxCnpCre and littermate-matched control mice (Figure

12A and 12B). Immunofluorescence co-staining of OLIG1 and ATRX on cryosections of P20 AtrxCnpCre and littermate-matched control mouse brains

Figure 12. ATRX expression is successfully lost from OLs in AtrxCnpCre mice.

A) Immunofluorescence staining of OLIG1 and ATRX in the corpus callosum of P20 AtrxCnpCre and control mice. N=3. Scale bar represents 70µm. DAPI stains the nucleus of the cell. B) Cells counts that assess the proportion of the number of OLIG1-positive cells per DAPI-positive cells per unit area. C) Cell counts that assess the proportion of the number of ATRX-positive cells per OLIG1-positive cells. Cell counts were performed on an N=2. Errors bars represent the range of two biological replicates.

74 confirmed that Atrx was successfully deleted from OLs in AtrxCnpCre mice (Figure

12A and 12C). These results suggest that this mouse model is a good model to study the OL-specific effects of ATRX in myelination.

3.8 Oligodendrocyte-specific deletion of ATRX does not

affect myelin formation in the mouse brain

As ATRX has been shown to directly regulate gene expression, it was hypothesized that ATRX may influence CNS myelination by regulating the expression of myelin-specific genes in mature myelinating OLs. P20 AtrxCnpCre and littermate-matched control mice were utilized to assess the OL-specific function of ATRX.

Characterization of the mouse phenotype was first performed to assess if loss of

ATRX from OLs results in abnormal physical traits compared to age-matched controls. It was determined that P20 AtrxCnpCre and littermate-matched control mice appeared proportionally similar to one-another in their overall body length

(Figure 13A and 13B)

To determine if loss of ATRX from OLs results in hypomyelination in the mouse brain, we performed western blot analysis of the myelin markers MOG, MBP, and

MAG from protein isolated from whole forebrains of P20 AtrxCnpCre and control mice (Figure 14A). It was found that the expression of MOG and MBP was unchanged in P20 AtrxCnpCre mice compared to littermate-matched controls

Figure 13. Characterization of the P20 AtrxCnpCre and littermate-matched control mouse phenotypes

A) Images of P20 control and AtrxCnpCre mice. B) Measurements of mouse lengths (mm). N=4. Error bars represent ±SEM. p>0.05

(Figure 14B). Additionally, this finding was corroborated by immunofluorescence co-staining of MBP and MOG on brain cryosections. Immunofluorescence staining revealed that expression of MBP and MOG in both the corpus callosum and the cerebral cortex was analogous between P20 AtrxCnpCre and littermate- matched controls (Figure 14C). This finding brings us to the conclusion that the abundance of myelin is unchanged in mice that lack ATRX specifically from OLs.

Taken together, these results strongly suggest that myelin sheath formation does not require ATRX within OL cells and therefore does not directly regulate myelin- specific gene expression. These findings imply that ATRX must be required in a different cell type to allow for proper formation of the myelin sheath in the CNS.

Figure 14. Expression of myelin-specific proteins is unchanged between P20 AtrxCnpCre mice compared to littermate-matched controls.

A) Western blot of MOG, MBP, and Actin from forebrains of P20 AtrxCnpCre and control mice. B) Quantitative analysis of western blot. MBP, and MOG were normalized to Actin. N=3. Error bars represent ±SEM. p>0.05. DAPI stains the nucleus of the cell. C) Immunofluorescence staining of MBP and MOG in the corpus callosum and cerebral cortex of P20 AtrxCnpCre and control mice. N=3. Scale bar represents 70µm.

4 Discussion

4.1 Summary of results

The ATRX chromatin remodeling protein has been found to be an essential factor for proper brain development, as mutations within the human ATRX gene result in ATR-X syndrome. ATR-X is a neurodevelopmental disorder characterized by cognitive defects, seizures, and microcephaly (Gibbons, 2006). Furthermore, recent MRI studies revealed that ATR-X syndrome patients also display varying degrees of white matter abnormalities, and delayed myelination (Wada et al.,

2013). Studies have implicated ATRX in several cellular processes including regulation of the epigenetic environment of heterochromatin, mitosis, and gene expression. The present study identifies not only a novel function of ATRX, but also an additional regulator of myelination within the mouse CNS.

To assess the changes in gene expression when ATRX is lost from P17 mouse forebrains, a microarray was carried out. Transcriptional profiling of the microarray revealed that expression of several myelin-specific genes was significantly downregulated in P17 AtrxFoxG1Cre mouse forebrains compared to age-matched controls. In addition, GO-analysis of significantly downregulated genes in the forebrain of P17 AtrxFoxG1Cre mice revealed that genes involved in several processes of myelination were of the most significantly downregulated genes. To determine if the reduction of myelin-specific gene expression in the

AtrxFoxG1Cre forebrain correlated with a decrease in myelin formation, we analyzed

81 the forebrains of AtrxFoxG1Cre and littermate-matched control mice at postnatal day

20, as the majority of neurons have been fully myelinated by this time. It was determined that loss of ATRX from the P20 forebrain results in significant hypomyelination in both the cerebral cortex and corpus callosum; two areas of the brain that are highly abundant in white matter and crucial for proper cognition.

Therefore, we can conclude that ATRX is in fact required for proper myelination within the CNS.

Hypomyelination in the brain can be the result of numerous consequences, including a reduction in the total number of axons that can potentially be myelinated. It was resolved that neuronal cell numbers were equivalent between

P20 AtrxFoxG1Cre and control samples, implying that the observed hypomyelination phenotype in the P20 AtrxFoxG1Cre forebrain is due to a defect in myelin development and not to due a reduction in the number of myelinated neurons.

I next wanted to determine the mechanism by which ATRX acts to allow for myelin formation to take place. I first analyzed the production and morphology of

OPCs at P0.5, as a decrease in the number of OPCs or malfunction of OPCs would result in a myelination defect. It was established that OPC production and morphology is unaffected in the absence of ATRX, suggesting that ATRX is required further downstream in the OL-development pathway. A possible mechanism of ATRX in myelin sheath formation is its potential requirement for

OPC differentiation into premyelinating iOLs. It was determined that loss of ATRX from the P10 AtrxFoxG1Cre mouse forebrain results in a decrease in the number of premyelinating iOLs. A defect in OPC differentiation can result in a decrease in

82 myelination as it limits the number of OLs that can myelinate axons; however, the decrease in the number of immature OLs when ATRX is absent, does not correlate with the severe hypomyelination phenotype observed in the P20

AtrxFoxG1Cre mouse forebrains. Therefore is it postulated that ATRX is required not only for OPC differentiation into iOLs, but is also required for the maturation of iOLs into myelinating OLs.

The myelin sheath is a modified plasma membrane that is enriched with cholesterol. Cholesterol is a major constituent of the myelin sheath (~26% of the dry weight of myelin), and previous studies have shown that mice with OLs that cannot synthesize cholesterol, display a delay in myelin sheath production

(Norton and Cammer, 1984; Saher et al., 2005). This led us to hypothesize that proper myelin sheath formation may be influenced by ATRX through the regulation of cholesterol synthesis.

Cholesterol levels were assessed at P14 as this time point represents the peak of myelination in mice. It was determined that the abundance of total cholesterol, free cholesterol, and cholesterol esters was analogous between P14 AtrxFoxG1Cre and control mouse forebrains. In addition, it was determined that the abundance of total cholesterol was similar to that of free cholesterol, and the presence of cholesterol esters was negligible. Cholesterol present in the brain is synthesized internally; therefore, the cholesterol found within the brain consists strictly of free cholesterol with no presence cholesterol esters (Baumann and Pham-Dinh, 2001;

Quarles, 2005). These findings allow us to conclude that ATRX does not regulate

CNS myelin sheath formation by influencing cholesterol biosynthesis.

The myelin sheath in the CNS is generated by a subset of glial cells known as

OLs. Regulation of myelin-specific gene expression within OLs is still poorly understood. In recent years, one such factor has emerged as a potential “master regulator” of myelin-specific gene expression. MRF is a transcriptional regulator that has been found to be crucial for myelin sheath formation. Ablation of MRF from OLs impedes myelin sheath formation almost completely; however, induction of MRF in OPCs can promote premature myelination (Emery et al.,

2009).

As ATRX has been implicated in the regulation of gene expression, I hypothesized that ATRX may act in a similar fashion to MRF, and act as a direct transcriptional regulator of several myelin-specific genes that are required for myelin sheath formation. To address this question, I first developed a novel mouse model in which Atrx is completely eradicated specifically from OL-lineage cells (AtrxCnpCre). To our surprise, analysis of both the cerebral cortex and corpus callosum revealed that myelination occurred normally in P20 mice lacking ATRX specifically from OLs; suggesting that ATRX does not directly regulate myelin- specific gene expression in OLs.

Taken together, these results allow for the rejection of my hypothesis. These findings illustrate that ATRX is required for myelin sheath formation in the CNS by not only regulating OPC differentiation, but also by influencing the maturation of iOLs into myelinating OLs, and does so through an OL non cell-autonomous mechanism.

4.2 ATRX acts in a non cell-autonomous role to regulate

myelin sheath formation

Initiation of myelin sheath formation requires numerous intracellular and extracellular mechanisms to act synergistically to allow for proper ensheathment of axons. Myelination in the central nervous system requires precise interaction of several cell types, including OLs, astrocytes, microglia, and neurons. It was hypothesized that ATRX may regulate the expression of myelin-specific genes within OLs, as these cells are the subtype of glial cell that myelinate axons; however, it was determined that myelination occurs normally when ATRX is lost from OLs, suggesting that ATRX must be required within in different cell population to promote myelination.

4.2.1 ATRX may act to control neuronal-specific gene

expression to regulate myelin sheath formation

Myelination is complex and tightly regulated process that is influenced not only by OL-intrinsic factors, but by axonal activity, secreted ligands, and expression of axonal surface ligands as well (Emery, 2010). Preliminary findings from our lab suggest that ATRX may have a neuronal-specific function in the regulation of myelination.

Female mice containing loxP sites flanking exon 18 of the Atrx gene were mated with male transgenic male mice that express Cre recombinase under control of the neuronal-specific marker, calcium/calmodulin-dependent protein kinase II

(CamkII) promoter (Chen et al., 2001). The resulting male progeny (AtrxCamkIICre) from this cross lacks ATRX specifically from postmitotic neurons. Co- immunofluorescence staining for ATRX and NeuN demonstrated that at P20,

ATRX has yet to be lost from postmitotic neurons in AtrxCamkIICre mice (Appendix

A); therefore, these mice cannot be used to study a potential neuronal-specific function of ATRX on myelin sheath formation. Conversely, it was determined that by 8-weeks of age, ATRX has been successfully lost from neurons in AtrxCamkIICre mice (Appendix A), and therefore can be used to study the neuronal-specific effect of ATRX on myelin sheath maintenance at a later age.

Preliminary evidence suggests that myelin sheath maintenance is disrupted in

AtrxCamkIICre mice at 3-months of age, as expression of myelin-specific markers were found to be significantly reduced in AtrxCamkIICre mice compared to controls when analyzed by western blot and immunofluorescence staining (Appendix B).

These preliminary findings suggest that ATRX has a neuronal-specific effect on myelin sheath maintenance, and therefore may have a neuronal-specific effect on myelin sheath formation; however, further experimentation will need to be carried out to resolve this theory.

ATRX may therefore regulate expression of genes that code for secreted factors and/or neuronal-specific factors that function by promoting OPC differentiation into iOLs, as well as maturation of iOLs into myelinating OLs. It has been found

86 that OPC differentiation into premyelinating iOLs relies on signaling molecules from the targeted axonal cell body. One such mechanism is the Notch-1 signaling pathway that is activated through binding of secreted axonal molecule

F3/contactin (Hu et al., 2003). The Notch-1 signaling pathway can both positively and negatively regulate OPC differentiation. Axonally derived F3/contactin binds the Notch-1 receptor on OPCs, and promotes OPC differentiation. This is achieved as F3/contactin binding to Notch-1 promotes the binding of downstream cytoplasmic target Deltex-1 (DTX-1) to the Notch intracellular domain (NICD) protein. This binding causes cleavage of NICD which allows it to translocate into the nucleus, where it then positively regulates the expression of a selection of myelin-specific genes, and thus OPC differentiation into immature premyelinating

OLs (Hu et al., 2003). On the other hand, the Notch-1 signaling pathway can inhibit differentiation, as Jagged-1 binding to Notch-1 inhibits OPC differentiation by activating Hes1 and Hes5 expression (Wang et al., 1998; Kondo and Raff,

2000).

In addition to the promoting the differentiation of OPCs, axonal signaling molecules have also been implicated in initiating the onset of myelination. One such molecule that has been shown to promote myelination is laminin-α2

(Morissette and Carbonetto, 1995; Colognato et al., 2002). Laminin-α2 binds the

OL-specific β-1 receptor, thus promoting the dephosphorylation of the Fyn tyrosine kinase. This dephosphorylation event results in the activation of ILK,

PINCH, and α-parvin leading to the polymerization of actin filaments, resulting in

87 the extension of the OL filopodia and allowing contact initialization with unmyelinated axons (Liang et al., 2004; Bauer et al., 2009; Hu et al., 2009).

A gene expression microarray was performed to assess the expression changes between P0.5 AtrxFoxG1Cre and control mice as well as P17 AtrxFoxG1Cre and control mice. Analysis of these microarrays revealed that expression of contactin-1 (also known as the F3 neural cell surface protein) was slightly decreased in both P0.5 and P17 AtrxFoxG1Cre mice [-1.17 fold (p>0.05) and -1.073 (p>0.05) respectively] compared to controls. It was also revealed that laminin-α2 was also slightly decreased in P17 AtrxFoxG1Cre mice [-1.14 fold (p>0.05)] compared to controls.

Although the expression changes seen are not significant, validation by qRT-

PCR should be performed to reveal the true expression changes of these genes, as microarrays have long been known to have inaccuracies.

A potential mechanism for ATRX is that it may be required to positively regulate the expression of contactin-1 and/or laminin-α2, thus promoting OPC differentiation, and/or the maturation of immature OLs into myelinating OLs.

ATRX may achieve this by directly influencing contactin-1 and laminin-α2 gene expression as described previously (Levy et al., 2014), or through its interaction with MeCP2, CTCF, or Daxx (Drané et al., 2010; Goldberg et al., 2010; Kernohan et al., 2010; Lewis et al., 2010; Wong et al., 2010; Kernohan et al., 2014).

4.2.2 Astrocyte influence in CNS myelination

The majority of studies that examine the mechanisms of CNS myelination have focused primarily on the OL-specific effects, as well as the axonal influences that promote myelin sheath formation. It has been documented that in addition to OLs and neurons, astrocytes are also a key contributor to myelination in the CNS.

Astrocytes are the most abundant and diverse subpopulation of glial cell found in the CNS, and are primarily involved in the organization and maintenance of brain structure and function (Sofroniew and Vinters, 2010; Lundgaard et al., 2014).

Astrocytes have been found to be required for neuronal migration and process extension during brain development, as well as being crucial for neuronal survival

(Powell and Geller, 1999; Wagner et al., 2006). In addition, astrocytes have also been found to be required not only for OL-development, but for myelination initiation and maintenance as well.

Astrocytes have been shown to influence OPC differentiation and myelin sheath formation through its interactions with OLs and neurons. Recently, it was revealed that through two separate mechanisms, OPC differentiation is influenced by the release of the promyelinating cytokine, leukemia inhibitory factor (LIF) from astrocytes (Fischer et al., 2014). OL and astrocyte co-culture experiments determined that the tumor necrosis factor (TNF) derived from OLs, activates TNFR2 on astrocytes. Activation of TNFR2 then results in the PI3K-Akt- dependent secretion of LIF, leading to an increase in expression of myelin- specific factors (Fischer et al., 2014). In addition, Ishibashi and colleagues further

89 determined that during development, neuronal activity simulates the release of

ATP, upon which signals neighboring astrocytes to release LIF (Ishibashi et al.,

2006). As ATRX is a known regulator of gene expression, loss of ATRX specifically from astrocytes may alter the expression pattern of several of these positive and negative regulators, and thereby potentially modifying the myelination phenotype.

It has been shown that loss of ATRX from the mouse forebrain results in an increase in the number of reactive astrocytes (McConkey unpublished). This increase may result in hypomyelination as it has been found that an increase in the number of reactive astrocytes can have detrimental effects in brain development. It has been discussed that an increase in reactive astrocytes can amplify the production of cytokines leading to an exacerbation of inflammation. In addition, reactive astrocytes can increase the release of toxic materials such as glutamate and reactive oxygen species (ROS) (Sofroniew and Vinters, 2010;

Lanciotti et al., 2013). This production and release of toxic substances can result in damage to the myelin sheath leading to increased neurodegeneration.

ATRX has been found to be important for proper NPC proliferation and differentiation. Loss of ATRX was found to result in premature NPC differentiation leading to an increase in the number of outer radial glia, as well as an increase in the generation of postmitotic projection neurons (Ritchie et al., 2014). As radial glial cells are multipotent progenitor cells, they have the capability to differentiate into various cell types including astrocytes (Vaccarino et al., 2007). It is therefore a possibility that premature differentiation of NPCs due to a loss of ATRX, may

90 also result in an increase in the number of reactive astrocytes; however further experimentation is required to assess this hypothesis.

4.2.3 Systemic regulation and growth factor influence of

myelin sheath formation in the CNS

CNS myelination is not only regulated by cell-cell communication between OLs and neurons as well as between OLs and astrocytes, but is also regulated by the

IGF-1 growth factor, and systemically by circulating T4.

T4 and its activated version T3 are tyrosine-based hormones that are produced by the follicular cells of the thyroid gland, and have been implicated to be crucial factors in proper brain development. It has been established that hypothyroidism results in mental retardation and delayed myelination (Dussault and Ruel, 1987); two characteristics observed in ATR-X syndrome patients. Thyroid hormones have been determined to be crucial for both differentiation and maturation of OLs in the CNS. T3 has been found to block OPC proliferation, therefore initiating differentiation of OPCs into immature OLs (Baas et al., 1998; Park et al., 2001).

In addition, hypothyroidism impedes the onset of myelination by delaying the expression of several crucial myelin-associated factors, including fatty acid α- hydroxylase, galactosyltransferase, sialyl-transferase, and sulfotransferase

(Dussault and Ruel, 1987). Furthermore, thyroid hormones have been found to directly regulate the expression of the myelin-specific genes, Mbp and Mag

(Farsetti et al., 1991; Rodriguez-Peña et al., 1993). It was observed that induced hypothyroidism in mice resulted in decreased MBP mRNA expression, and it was further determined that the promoter region of Mbp is susceptible to thyroid hormone binding, suggesting that Mbp expression is directly (at least partially) regulated by thyroid hormones (Farsetti et al., 1991). These studies reveal that thyroid hormones have a fundamental role in the development of the OL pathway and the onset of myelination.

In addition to thyroid hormones, IGF-1 has also been found to be an important regulator of CNS myelination. IGF-1 has been established as a significant factor in the differentiation, maturation, and survival of OLs (Carson et al., 1993;

McMorris et al., 1993). Overexpression of IGF-1 in mice, revealed a substantial increase in both the number of OLs, and the abundance of myelin (Carson et al.,

1993; Ye et al., 1995). On the other hand, mice lacking IGF-1 display a significant reduction in the myelin-specific markers MBP and PLP at 3-weeks of age, a characteristic of delayed myelination (Ye et al., 2002). In addition, it was revealed that mice lacking the IGF1 receptor (IGF1R) from OLs displayed significant reductions in overall brain size, number of OPCs, as well as a reduction in the number of differentiated iOLs leading to a decrease in myelination (Zeger et al., 2007). These findings strongly demonstrate that the

IGF-1/IGF1R signaling pathway is crucial for proper OL and myelin sheath development.

It has been previously shown that loss of ATRX from the P20 mouse forebrain and anterior pituitary, results in decreased circulating T4 and IGF-1 levels

(Watson et al., 2013). In addition, it was also determined that loss of ATRX results in decreased expression of the thyroid stimulating hormone (Tsh)

(Watson et al., 2013). TSH is made by thyrotropes, a population of cells that are located in the anterior pituitary, and who directly influence the regulation of T4.

As these factors are known to be crucial for OL-development and myelination, a potential mechanism of ATRX in myelin sheath formation, may be its requirement for the transcriptional regulation of these factors through previously seen mechanisms (Drané et al., 2010; Goldberg et al., 2010; Lewis et al., 2010; Wong et al., 2010; Kernohan et al., 2014; Levy et al., 2014). It is still unclear whether the loss of ATRX results in decreased levels of T4 and IGF-1 separately, or if the decrease of IGF-1 is the result of T4 dysfunction, as it has been found that T4 can bind to the Igf-1 transcript and thus can regulate its expression (Wang et al.,

2010). Our group is currently addressing the role of ATRX in the regulation of T4 and IGF-1, and findings from this study may further elucidate the role of ATRX in myelin sheath formation.

4.3 Future Directions

To address the hypotheses that ATRX may influence OPC differentiation and maturation of iOLs through neuronal or astrocyte-specific effects, transgenic mouse lines will be utilized to conditionally delete ATRX from postmitotic neurons and from astrocytes in the CNS prior to myelin sheath formation.

To assess the neuronal-specific function of ATRX, female mice containing loxP sites flanking exon 18 of the Atrx gene will be crossed with transgenic male mice containing the Cre recombinase gene downstream of the Nex promoter

(Goebbels et al., 2006). This cross will result in progeny (AtrxNexCre) that will have a deletion of Atrx exclusively from postmitotic neurons beginning at approximately E12.5 (Goebbels et al., 2006).

To assess the astrocyte-specific effect of ATRX, a mouse model to conditionally deplete ATRX from astrocytes that has been previously characterized will be used (McConkey unpublished). Female mice containing loxP sites flanking exon

18 of the Atrx gene were crossed with transgenic male mice containing the Cre recombinase gene downstream of the glial high affinity glutamate transporter

(Glast) promoter. This cross results in progeny (AtrxGlastCreERT) that have a deletion of Atrx exclusively from astrocytes upon administration of tamoxifen.

To address the myelination phenotype when ATRX is lost specifically from neurons and astrocytes, identical experimental procedures as the ones performed in section 3.2 of this thesis, will be carried out. Findings from these experiments will allow us to establish if ATRX has a neuronal-specific and/or an astrocyte-specific effect on myelin sheath formation in the CNS.

To identify extracellular factors that may be required for myelin sheath formation, mass spectrometry will be performed on isolated cell co-culture media from separate OPC/neuronal and OPC/astrocyte cell co-culture experiments, using primary cells extracted from AtrxNexCre and control mice, as well as AtrxGlastCreERT

94 and control mice. Mass spectrometry analysis of cultured medium from these separate cell co-culture experiments will enable us to identify extracellular signaling molecules that are required for myelination, as well as facilitate in the detection of secreted molecules that are differentially expressed when ATRX is lost from neurons and astrocytes.

In addition to secreted ligands, expression of intrinsic neuronal-specific and astrocyte-specific factors that are essential for myelin sheath formation may also be dysregulated in the absence of ATRX. To assess neuronal-specific and astrocyte-specific gene expression changes when ATRX is lost from post-mitotic neurons, RNA-seq will be carried out on RNA isolated from P5, P14, and P20

AtrxNexCre and control cortical neurons, as well as RNA isolated from P5, P14, and

P20 AtrxGlastCreERT and control astrocytes, that have been separated by fluorescence-activated cell sorting (FACS). Cells from P5 mice will be used, as

OPC differentiation is highly active at this time. Cells from P14 mouse forebrains will be used, as this time point represents the peak of myelination, and cells from

P20 mouse forebrains will be used, as the myelin sheath is highly abundant at this time. Identification of dysregulated neuronal-specific and astrocyte-specific genes, that code for factors implicated in myelination is essential as an ATRX- dependent process may regulate the expression of these genes.

Comparative analysis of extracellular signaling ligands by mass spectrometry, and analysis of differential gene expression by RNA-seq will allow us to identify potential neuronal and astrocyte-specific targets whose expression may be

95 regulated by ATRX. This data will allow us to generate additional hypotheses to identify the exact molecular mechanism of ATRX in CNS myelination.

Figure 15. Proposed model of ATRX and its potential mechanism in myelination of neurons in the mouse CNS

ATRX may regulate the expression of neuron-specific genes such as F3/contactin and/or laminin-α2, to influence oligodendrocyte precursor cell differentiation and/or maturation of immature oligodendrocytes into myelinating oligodendrocytes respectively (i). ATRX may also act to regulate astrocyte- specific genes such as leukemia inhibitory factor (LIF) or other astrocyte-specific genes (ASG), which would be required for oligodendrocyte precursor cell differentiation (ii). An additional mechanism of ATRX is that it may act to regulate the expression of thyroid stimulating hormone (Tsh) which then regulates the expression of thyroid hormones (TH) and ultimately oligodendrocyte development and myelination (iii).

5 Conclusions

The present study addresses a novel role of ATRX in the development of OLs and myelin sheath formation in the CNS. It was determined that loss of ATRX from the mouse forebrain results in hypomyelination, and that this phenotype is due to a defect in OPC differentiation into iOLs, and iOL maturation into myelinating OLs. Furthermore, it was resolved that ATRX does not have an OL cell-autonomous role in myelination. It is suggested that ATRX influences myelin sheath formation and maintenance by potentially regulating the expression of neuronal-specific and/or astrocyte-specific factors that are required for myelination, as well as potentially regulating circulating T4 and IGF-1 levels

(Figure 15). Elucidation of the role of ATRX in myelin production would present a novel non cell-autonomous mechanism that regulates myelin formation.

Transcriptional regulation of neuronal-specific and astrocyte-specific genes that code for factors that function by instructing OPC differentiation, and OL maturation have yet to be fully elucidated; therefore, this discovery would be beneficial, as it will further our understanding of the mechanisms involved in myelination in the postnatal mouse forebrain. ATRX causes severe cognitive deficits, seizures, and microcephaly, all of which are associated with loss of white matter. These findings give the prospect of potentially finding a method to alleviate hypomyelination defects in ATR-X patients, which may lead to discoveries of potential therapies for ATR-X patients, as well as other myelin relevant disorders such as multiple sclerosis, various leukodystrophies, and neuropsychiatric disorders.

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Appendices

Appendix A. ATRX expression is retained in neurons in P20 AtrxCamkIICre mice, but is lost from neurons in 3-month-old AtrxCamkIICre mice

Immunofluorescence staining of NeuN and ATRX in the cerebral cortex of P20 and 3-month-old AtrxCamkIICre and control mice. N=1. Scale bar represents 70µm. DAPI stains the nucleus of the cell.

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Appendix B. Expression of myelin-specific proteins is reduced in 3-month- old AtrxCamkII mice compared to littermate-matched controls

A) Western blot of MOG, MBP, and Actin from forebrain samples of 3-month-old AtrxCamkIICre and control mice. B) Quantitative analysis of western blot. MOG and MBP were normalized to Actin. N=2. Error bars represent the range of expression of two biological replicates. C) Immunofluorescence staining of MBP and MOG in P20 AtrxFoxG1Cre and control corpus callosum and cerebral cortex. N=1. Scale bar represents 70µm. DAPI stains the nucleus of the cell.

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Curriculum Vitae

Name: Matthew William Edwards

Post-secondary The University of Western Ontario Education and London, Ontario, Canada Degrees: 2007-2011 B.Sc. in Biology, Honour’s Specialization in Genetics

The University of Western Ontario London, Ontario, Canada 2011-2016 M.Sc. in Biochemistry

Honours and Travel Award, Children’s Health Research Institute, London, Awards: Ontario, Canada 2015

Travel Award, Department of Paediatrics, The University of Western Ontario, London, Ontario, Canada 2015

Best Oral Presentation, Annual Paediatrics Research Day, Children’s Health Research Institute, London, Ontario, Canada 2015

Lawson Health Research Institute Studentship, Lawson Health Research Institute Internal Research Fund, London, Ontario, Canada 2014

Travel Award, Joint meeting of the 20th Biennial Meeting of the International Society for Developmental Neuroscience and the 5th Annual NeuroDevNet Brain Development Conference, Montreal, Quebec, Canada 2014

The Jonathan and Joshua Memorial Graduate Scholarship, Schulich School of Medicine and Dentistry, The University of Western Ontario, London, Ontario, Canada 2014

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Best Poster Presentation, Southern Ontario Neuroscience Association Annual Meeting, London, Ontario, Canada 2014

Best Poster Presentation, Annual Paediatrics Research Day, Children’s Health Research Institute, London, Ontario, Canada 2012

Department of Paediatrics Graduate Studentship, Children’s Health Research Institute, London, Ontario, Canada 2011

Department of Paediatrics Summer Studentship, Children’s Health Research Institute, London, Ontario, Canada 2011

Western Graduate Research Scholarship, The University of Western Ontario, London, Ontario, Canada 2011-2016

Related Work Summer Research Student Experience Children’s Health Research Institute, London, Ontario, Canada 2011

Research Associate, The University of Western Ontario, London, Ontario, Canada 2009-2011

Publications

LA Watson, L Solomon, J Ruizhe, Y Jiang, M Edwards, K Shin-ya, F Beier, NG Bérubé. (2013). Atrx deficiency induces telomere dysfunction, endocrine defects, and reduced life span. J. Clin. Invest. 123(5): p.2049-20634

Poster Presentations

MW Edwards, Y Jiang, NG Bérubé. Regulation of CNS myelination by the ATRX chromatin remodeling protein. The Society for Neuroscience 45th Annual Meeting: Neuroscience 2015, October 2015, Chicago, Illinois, U.S.A

MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. London Health Research Day, March 2015, London, Ontario, Canada

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MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. Joint Meeting of the 20th Biennial Meeting of the International Society for Developmental Neuroscience and the 5th Annual NeuroDevNet Brain Development Conference, July 2014, Montreal, Quebec, Canada

MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. The 2nd Canadian Conference on Epigenetics, June 2014, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. Developmental Biology Research Day, May 2014, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. Annual Paediatrics Research Day, May 2014, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. Southern Ontario Neuroscience Association Annual Meeting, May 2014, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Loss of ATRX causes hypomyelination in the mouse CNS. London Health Research Day, March 2014, London, Ontario, Canada

MW Edwards, Y Jiang, B Sutherland, NG Bérubé. Effects of ATRX deficiency on myelin formation in the CNS. Developmental Biology Research Day, May 2013, London, Ontario, Canada

MW Edwards, Y Jiang, B Sutherland, NG Bérubé. Effects of ATRX deficiency on myelin formation in the CNS. Annual Paediatrics Research Day, May 2013, London, Ontario, Canada

MW Edwards, Y Jiang, B Sutherland, NG Bérubé. Effects of ATRX deficiency on myelin formation in the CNS. London Health Research Day, March 2013, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Effects of the ATR-X syndrome protein on brain myelination. Developmental Biology Research Day, May 2012, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Effects of the ATR-X syndrome protein on brain myelination. Annual Paediatrics Research Day, May 2012, London, Ontario, Canada

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MW Edwards, Y Jiang, NG Bérubé. Effects of the ATR-X syndrome protein on brain myelination. Lawson Research Day, March 2012, London, Ontario, Canada

MW Edwards, AM Laliberté, CML Hutnik, KA Hill. Extracellular matrix remodelling in the retina of the harlequin (hq) mouse as a possible mechanism for human age-related retinal degeneration. Centre for Population, Health and Aging 8th Annual Interdisciplinary Research Event, May 2011, The University of Western Ontario, London, Ontario, Canada

MW Edwards, AM Laliberté, CML Hutnik, KA Hill. Extracellular matrix remodelling in the retina of the harlequin (hq) mouse as a possible mechanism for human age-related retinal degeneration. Lawson Research Day, March 2011, London, Ontario, Canada

Oral Presentations

MW Edwards, Y Jiang, NG Bérubé. Loss of the ATRX chromatin remodeling protein results in hypomyelination in the mouse CNS. Developmental Biology Research Day, June 2015, London, Ontario, Canada

MW Edwards, Y Jiang, NG Bérubé. Loss of the ATRX chromatin remodeling protein results in hypomyelination in the mouse CNS. Annual Paediatrics Research Day, May 2015, London, Ontario, Canada

MW Edwards, AM Laliberté, CML Hutnik, KA Hill. Retinal degeneration in the harlequin (hq) mouse is characterized by early gene expression changes with associated nuclear displacement. Ontario Biology Day, March 2011, Wilfred Laurier University, Waterloo, Ontario, Canada

M Edwards, AM Laliberté, TC MacPherson, CML Hutnik, KA Hill. The power of animal mimics of human eye disease: in vivo imaging, post-mortem histology and molecular biomarkers. Annual Ophthalmology Research Day, November 2010, London, Ontario, Canada