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Investigating the regulation of ARNT2, a neuroprotective protein, in models of multiple sclerosis Rahim, Titissa 2016

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INVESTIGATING THE REGULATION OF ARNT2, A NEUROPROTECTIVE PROTEIN, IN MODELS OF MULTIPLE SCLEROSIS by  Titissa Rahim  B.Sc., The University of British Columbia, 2013  A THESIS SUBMITTED IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF  MASTER OF SCIENCE in THE FACULTY OF GRADUATE AND POSTDOCTORAL STUDIES (Pathology and Laboratory Medicine)  THE UNIVERSITY OF BRITISH COLUMBIA  (Vancouver)  June 2016  © Titissa Rahim, 2016 ii  Abstract  Background: The process of axonal degeneration and neuronal loss has been described as the major cause of irreversible clinical disability in multiple sclerosis (MS). An ideal neuroprotective strategy would be to focus on inhibition of axonal degeneration and on protection against neuronal cell death in addition to immunomodulation. The aryl-hydrocarbon receptor nuclear translocator 2 (ARNT2) is a protein with neuroprotective properties previously described in ischemic insults and oxidative damage. We hypothesize that alterations in ARNT2 expression are associated with changes in cell viability in in vitro and in vivo models of multiple sclerosis. Methods: Following exposure to various compounds mimicking MS disease processes, ARNT2 protein and mRNA levels were observed in primary cortical neuron-enriched cultures using western blotting, quantitative polymerase chain reaction (qPCR) and immunocytochemistry, alongside cytotoxicity measurements, using a lactate dehydrogenase (LDH) release assay/Live/Dead® Viability/Cytotoxicity assay. ARNT2 protein levels were also evaluated in primary cortical astrocytes using immunocytochemistry. Analyses in an animal model of MS, experimental autoimmune encephalomyelitis (EAE) were conducted, with tissue collected at various stages of the disease course, to examine ARNT2 expression patterns in vivo. Results: Examination of individual neurons reveals that most cells demonstrate low-medium ARNT2 expression under steady-state conditions. Exposing cells to both low and higher concentrations of hydrogen peroxide (H2O2) to mimic mild to more severe oxidative stress significantly increases ARNT2 protein levels early, as measured via western blotting and immunocytochemistry. At the mRNA level, oxidative stress fails to drive Arnt2. This increased detection of ARNT2 protein is observed in both neurons and reactive astrocytes specifically iii  within the neuronal-enriched mixed populations. Non-reactive astrocytes also express ARNT2 at baseline conditions. Finally, ARNT2 is differentially expressed in healthy versus EAE tissue at peak disease. Conclusions: This work demonstrates for the first time that ARNT2 can follow altered expression patterns in vitro in neurons depending on the severity/duration of the stimulus involved in MS disease progression. This lays a foundation for understanding the link between ARNT2 expression and neuronal health in vitro.  iv  Preface   Dr. Quandt carried out induction of EAE as described in Chapter 5. Scoring and euthanasia were performed by Katerina Othonos and the author. Once euthanized, brains and spinal cords were collected by Sarah Neil, Katerina Othonos and the author. Andrew Leung, Katerina Othonos and Cheryl Whiting provided assistance with tissue processing and neuronal culture for some of the experiments. Astrocyte cultures were established by Adam Yu. Maintenance of astrocyte cultures was carried out by Cheryl Whiting and the author. Tissue sectioning, immunohistochemistry and tissue data collection was carried out by Adam Yu and analyzed by Dr. Quandt.  All animal work was approved by the UBC Animal Care Committee (certificate #A130281). Cell culture work with biohazardous materials was performed with approval from the UBC Biosafety Committee (certificate #B09-0179).   Otherwise, this thesis comprises original, unpublished work performed by the author, Ms. Titissa Rahim.   v  Table of Contents  Abstract .......................................................................................................................................... ii  Preface ........................................................................................................................................... iv  Table of Contents .......................................................................................................................... v  List of Figures ............................................................................................................................... ix  List of Abbreviations .................................................................................................................... xi  Acknowledgements ..................................................................................................................... xiii  Dedication ................................................................................................................................... xiv  Chapter 1: Introduction ................................................................................................................ 1  1.1	   Multiple Sclerosis ............................................................................................................ 1  1.1.1	   Overview ..................................................................................................................... 1  1.1.1	   Steady state roles of CNS cells ................................................................................... 2  1.1.3	   Pathogenesis ................................................................................................................ 3  1.1.4	   Neurodegeneration and axonal loss ............................................................................. 7  1.1.5	   Experimental autoimmune encephalomyelitis .......................................................... 12  1.1.6	   Current strategies of neuroprotection ........................................................................ 14  1.2	   ARNT2 .......................................................................................................................... 17  1.2.1	   Overview ................................................................................................................... 17  1.2.2	   Expression patterns ................................................................................................... 18  1.2.3	   Functional roles ......................................................................................................... 19  1.2.4	   ARNT2 as a neuronal survival factor ........................................................................ 22  1.2.5	   A role for ARNT2 in inflammatory neurodegeneration ............................................ 23  vi  1.3	   Hypothesis ..................................................................................................................... 26  Chapter 2: Materials and Methods ............................................................................................ 28  2.1	   Primary cortical neuron-enriched cultures .................................................................... 28  2.2	   Primary cortical astrocyte-enriched cultures ................................................................. 28  2.3	   Treatments ..................................................................................................................... 29  2.4	   Cell lysis, protein quantification, SDS-PAGE and western blotting ............................. 29  2.5	   Cell lysis, RNA isolation, reverse transcription and quantitative polymerase chain reaction ...................................................................................................................................... 30  2.6	   Immunocytochemistry ................................................................................................... 32  2.6.1	   Protocol and visualization ................................................................................. 32  2.6.2	   Methods of analysis ........................................................................................... 32  2.7	   Lactate dehydrogenase assay ........................................................................................ 33  2.8	   Live/Dead® viability/cytotoxicity assay ........................................................................ 34  2.9	   Experimental autoimmune encephalomyelitis .............................................................. 35  2.10	   Tissue collection, removal and processing of brain and spinal cord ............................. 36  2.11	   Immunohistochemistry .................................................................................................. 37  2.12	   Statistical Analysis ........................................................................................................ 37  Chapter 3: The effect of excitatory compounds, inflammatory/apoptotic mediators on ARNT2 .......................................................................................................................................... 39  3.1	   Characterization of primary cortical neuron-enriched cultures ..................................... 39  3.2	   The majority of cells in primary cortical neuron-enriched cultures express mild-moderate levels of ARNT2 ....................................................................................................... 39  3.3	   4-aminopyridine and bicuculline have no effect on ARNT2 protein levels ................. 43  3.4	   Potassium chloride has no effect on ARNT2 at low doses, and significantly reduces ARNT2 protein levels at excitatory/cytotoxic doses ................................................................. 44 vii   3.5	   Early exposure to glutamate has no effect on ARNT2 protein levels, while prolonged glutamate exposure results in decreased levels of ARNT2. ...................................................... 47  3.6	   Staurosporine significantly increases ARNT2 protein levels ....................................... 48  3.7	   Hydrogen peroxide significantly alters ARNT2 protein expression ............................. 51  3.8	   Oxidative stress fails to drive Arnt2 mRNA levels ....................................................... 58  3.9	   Oxidative stress significantly alters ARNT2 protein levels in both neuronal and astrocytic populations ................................................................................................................ 60  3.10	   Characterization of primary astrocyte-enriched cultures .............................................. 62  Chapter 4: Optimization of viability/cytotoxicity assays ......................................................... 64  4.1	   Optimizing neuronal treatment protocols to improve sensitivity of LDH detection ..... 64  4.2	   The Live/Dead® viability/cytotoxicity assay is sensitive in distinguishing live and dead populations in primary cortical neuron-enriched cultures ......................................................... 66  Chapter 5: In vivo Results .......................................................................................................... 68  5.1	   EAE results .................................................................................................................... 68  5.2	   Preliminary	  localization of ARNT2 protein in healthy tissues versus tissues undergoing EAE............................................................................................................................................ 69  Chapter 6: Discussion ................................................................................................................. 72  6.1	   Summary and significance ............................................................................................ 72  6.2	   The influence of excitatory compounds on ARNT2 ..................................................... 73  6.3	   ARNT2 at the mRNA versus protein level ................................................................... 74  6.4	   Global versus individual cell-based analysis of ARNT2 expression ............................ 76  6.5	   ARNT2 expression in astrocytes ................................................................................... 77  6.6	   ARNT2 expression and changes in cell viability .......................................................... 80  6.7	   Limitations .................................................................................................................... 83  viii  6.8	   Conclusions and future directions ................................................................................. 85  References .................................................................................................................................... 90  Appendices ................................................................................................................................. 102  Appendix A ............................................................................................................................. 102  A.1	   Antibodies used for western blotting ...................................................................... 102  A.2	   Antibodies used for immunocytochemistry/immunohistochemistry ...................... 102  Appendix B ............................................................................................................................. 103  B.1	   Primers used for qPCR ............................................................................................ 103  B.2	   Run conditions used for qPCR ................................................................................ 103  B.3	   Western blot results for lysates used for qPCR ....................................................... 104  Appendix C ............................................................................................................................. 105  C.1	   Testing of the Live/Dead® viability/cytotoxicity assay with 2µM calcein-AM and EthD-1 ................................................................................................................................. 105          ix  List of Figures  Figure 1.1 Arnt2 and Bdnf spinal cord and Arnt2 brain gene expression over the course of chronic progressive EAE ............................................................................................................................ 25  Figure 1.2 ARNT2 protein detection in the spinal cord of healthy and EAE mice. ...................... 26  Figure 2.1 Chronic progressive EAE disease progression and associated clinical scoring .......... 36 Figure 3.1 Characterization of primary cortical neuron-enriched cultures ................................... 41 Figure 3.2 The majority of cells in primary cortical neuron-enriched cultures express mild-moderate levels of ARNT2 ........................................................................................................... 42  Figure 3.3 4-aminopyridine and bicuculline have no effect on ARNT2 protein levels ................ 44  Figure 3.4 Potassium chloride has no effect on ARNT2 protein levels at doses below and including 10mM ............................................................................................................................ 46  Figure 3.5 Potassium chloride significantly reduces ARNT2 protein levels at excitatory/cytotoxic doses .............................................................................................................................................. 47  Figure 3.6 Early exposure to glutamate has no effect on ARNT2 protein levels ......................... 49  Figure 3.7 Prolonged glutamate exposure results in decreased levels of ARNT2 ........................ 50  Figure 3.8 Staurosporine significantly increases ARNT2 protein levels ...................................... 52  Figure 3.9 Representative phase microscopy images and cytotoxicity measurements showing progression of cell viability changes following H2O2 exposure ................................................... 54  Figure 3.10 Exposure to 25µM H2O2 significantly upregulates ARNT2 protein levels ............... 55  Figure 3.11 Exposure to 50µM H2O2 significantly upregulates ARNT2 protein levels ............... 56  Figure 3.12 Exposure to 100µM H2O2 significantly upregulates ARNT2 protein levels ............. 57  Figure 3.13 Exposure to 300µM H2O2 significantly downregulates ARNT2 protein levels ........ 58  Figure 3.14 Oxidative stress fails to drive Arnt2 mRNA levels .................................................... 59  Figure 3.15 Oxidative stress significantly alters ARNT2 protein levels in both neuronal and astrocytic populations .................................................................................................................... 61 x   Figure 3.16 Characterization of primary astrocyte-enriched cultures ........................................... 63  Figure 4.1 Optimizing neuronal treatment protocols to improve sensitivity of LDH detection ... 65  Figure 4.2 The Live/Dead® viability/cytotoxicity assay is sensitive in distinguishing live and dead populations in primary cortical neuron-enriched cultures .................................................... 67  Figure 5.1 EAE results .................................................................................................................. 69  Figure 5.2 Preliminary localization of ARNT2 in healthy tissues versus tissues undergoing EAE ....................................................................................................................................................... 71    xi   List of Abbreviations  4-AP   4-aminopyridine AhR   Aryl hydrocarbon receptor AMPA   α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid  ANOVA  Analysis of variance ARNT   Aryl hydrocarbon receptor nuclear translocator ARNT2   Aryl hydrocarbon receptor nuclear translocator 2 ARC   Activity-regulated cytoskeletal protein ATP   Adenosine triphosphate BBB   Blood-brain barrier BCA   Bicinchoninic acid BDNF   Brain-derived neurotrophic factor BHLH   Basic helix-loop-helix BRN2   POU domain, class 3, transcription factor 2 CME   Central midline enhancer CNS   Central nervous system CYP1A1  Cytochrome P450, family 1, member A1 DAPI   4',6-diamidino-2-phenylindole DNA   Deoxyribonucleic acid DIV   Days in vitro EAE   Experimental autoimmune encephalomyelitis EthD-1  Ethidium homodimer-1 FOS   FBJ Murine Osteosarcoma Viral Oncogene Homolog GABA   gamma-Aminobutyric acid  GFAP   Glial fibrillary acidic protein GLUT1/3  Glucose transporter 1/3 GST   Glutathione S-transferase H2O2   Hydrogen peroxide HBSS   Hank’s balanced salt solution HIF-1α  Hypoxia inducible factor-1alpha HLA   Human leukocyte antigen HRE   Hypoxia response element IFA   Incomplete Freund’s adjuvant KCL   Potassium chloride LDH   Lactate dehydrogenase MAI   Myelin associated growth-inhibiting factor MAP2   Microtubule-associated protein 2 MHC   Major histocompatibility complex MOG   Myelin oligodendrocyte glycoprotein MS   Multiple Sclerosis NMDA  N-methyl-D-aspartate  NPAS4  Neuronal PAS domain protein 4 xii  PAS   Period circadian protein-aryl hydrocarbon receptor nuclear translocator     protein-single-minded protein (PER-ARNT-SIM) PASRE  PER-ARNT-SIM response element PBS   Phosphate-buffered saline PLP   Proteolipid protein PPMS   Primary progressive multiple sclerosis PRMS   Progressive relapsing multiple sclerosis qPCR   Quantitative polymerase chain reaction RIPA   Radioimmunoprecipitation assay buffer RM   Repeated measures RNA   Ribonucleic acid ROS   Reactive oxygen species RRMS   Relapsing remitting multiple sclerosis SDS   Sodium dodecyl sulfate SDS-PAGE  Sodium dodecyl sulfate polyacrylamide gel electrophoresis SIM1   Single-minded homolog 1 SPMS   Secondary progressive multiple sclerosis TBS   Tris buffered saline TBS-T   Tris buffered saline with Tween 20 TCDD   2,3,7,8-tetrachlorodibenzo-p-dioxin TEMPOL  4-hydroxy-2,2,6,6-tetramethylpiperidine-N-oxyl VEGF   Vascular endothelial growth factor XRE   Xenobiotic response element     xiii  Acknowledgements   I would like to thank my supervisor, Dr. Jacqueline Quandt for her continued support over the duration of my project. I would also like to thank my committee, Dr. Cheryl Wellington, Dr. Neil Cashman and Dr. Marc Horwitz, for their invaluable input. Furthermore, I would like to thank past and present members of the Quandt lab, including Andrew Leung, Sarah Neil, Adam Yu, Katerina Othonos, Negar Farzam-Kia and Cheryl Whiting, and my family for their support.  Many thanks to the Bamji lab for providing us with rat cortices and protocols essential for carrying out my experiments, as well as Izabella Gadawska and the Côté lab, for their help in establishing our qPCR protocol, primers and standards.   Thank you to the Canadian Institute of Health Research, as well as the Multiple Sclerosis Society of Canada for studentship support. As well, thank you to the Department of Pathology and Laboratory Medicine, and the Faculty of Medicine for my scholarships.  Finally, thank you to the endMS Research and Training Network for providing me with opportunities for training and networking, and a travel award. xiv            For Quinn. 1  Chapter 1: Introduction 1.1 Multiple Sclerosis 1.1.1 Overview  Multiple Sclerosis (MS) is the most common neurological disease of young adults in Canada, with prevalence rates ranging from 55 to 240 people per 100,000 [1] [2]. As estimated by the MS Society of Canada three more Canadians are diagnosed with MS every day. First described as a distinct disease by Jean-Martin Charcot in 1868, MS is characterized as a chronic inflammatory and demyelinating neurodegenerative disorder of the central nervous system (CNS), which includes the brain and the spinal cord [2] [3] [4]. Roughly 80% of patients present following the experience of a period of relapsing and remitting disease (RRMS), characterized by acute attacks followed by complete/partial recovery [5] [6]. After 10-20 years, this commonly develops into a secondary progressive disease course (SPMS) with gradual clinical deterioration [5] [6]. About 15% of MS patients have progression from the start of disease, called primary progressive MS (PPMS) [5] [6]. Progressive forms of the disease are characterized by a lack of inflammatory activity despite progression of clinical disability [2]. The rarest type of MS is the progressive relapsing form (PRMS), occurring in 5% of MS patients, characterized by progressive disease with periods of flare-ups [7]. Consequences of MS include loss of balance, cognitive dysfunction, impaired vision/speech, extreme fatigue and paralysis [4] [7].  The etiology of MS is not well understood, but is thought to be multifactorial, including genetic susceptibility, gender, and environmental and viral exposure [8] [9] [10] [11] [12]. First of all, the age of onset of MS is between 20-40 years (with RRMS onset earlier than PPMS/SPMS), which is considerably younger than other neurodegenerative diseases such as Alzheimer’s disease (AD) and Parkinson’s disease (PD) [13] [14]. The strongest genetic 2  association of increased risk of MS development to date is the human leukocyte antigens (HLA)-DRB1*1501, -DRB1*0101, -DQA1*0101, -DQA1*0102, and -DQB1*0602 [15]. The HLA gene family is involved in encoding antigen-presenting proteins and self-antigen recognition [15]. This link is found most strongly in northern European groups [15]. Additionally, family members of MS patients are at a greater risk, with an increased risk of over 20 fold in first-degree relatives of individuals with MS compared to the general population [9]. Furthermore, women are 2-3 times more likely to develop the disease than men [12]. Environmental factors studied include lack of vitamin D exposure, diet, smoking, stress, climate, and infection history [16]. Risk of developing MS has been associated with the human herpes virus 6 and Epstein-Barr virus, the latter of which in one study was present in people with MS at a significantly higher rate than controls (nearly 100% versus 90-95%, respectively) [17].   1.1.2 Steady state roles of CNS cells  The CNS has various resident cells, including neurons and supporting glia, comprised of astrocytes, microglia, and oligodendrocytes. In addition, the blood-brain barrier (BBB) is a structure consisting of astrocytic endfeet, endothelial cells with tight junctions, and functions as a diffusion barrier (in addition to specific transporters) to regulate entry of compounds into the CNS and maintenance of CNS homeostasis [18].   Under normal physiological conditions, astrocytes play many crucial roles in the CNS including metabolic support to neurons, maintenance of the BBB, neurotransmitter uptake and release, and modulation of synaptic transmission, amongst others [19]. Furthermore, subtypes of astrocytes, radial glia and Bergmann glia, play roles in neuron migration and cerebellar development, respectively, and are essential for proper structuring of the developing brain [20]. 3   Under pathological conditions, astrocytes undergo a morphological change to a reactive phenotype, characterized by enhanced intermediate filament expression, further described in section 1.1.3 [20].  Microglia are the resident immune cells of the CNS, and are often likened to macrophages for their roles in inflammation and immune defenses in the CNS through scavenging, phagocytosis, and antigen presentation [21]. Beyond this role in response to insult or injury, microglia play an important role during CNS development in the regulation of apoptosis, neuron/astrocyte proliferation and neuronal differentiation, and synaptic pruning [21] [22].   Oligodendrocytes are the resident myelinating cells of the CNS, maintaining the myelin sheath that covers the axons of many neurons. The myelin sheath plays crucial roles in proper saltatory conduction of action potentials, as well as trophic support for vesicular trafficking along axons [23] [24]. Myelination is only observed in some brain regions at birth, completing at around 25-30 years of age, and continues throughout adulthood [23].  All of these cellular and structural components interact in a complex manner to ensure proper functioning of the CNS, but are also influenced during insult or trauma and contribute to both reparative and pathological processes observed throughout disease onset and progression in neuroinflammation and neurodegeneration.  1.1.3 Pathogenesis  Evidence from an animal model of MS (experimental autoimmune encephalomyelitis, or EAE) has suggested that MS pathogenesis is initiated peripherally (i.e. outside of the CNS) by activated T cells autoreactive to the myelin sheath [3]. In patients activation of autoreactive T cells is thought to occur following an environmental or viral trigger as described in section 1.1.1 4  [3]. While this process of initial activation is not well understood, one potential reason is that myelin and other CNS antigens are expressed in the periphery and are thereby able to activate these autoreactive T cells, which are a normal component of the immune system [25].   Regardless of clinical manifestation, MS pathogenesis is characterized by a series of pathological hallmarks: alterations in blood-brain barrier (BBB) function/increased BBB permeability, inflammatory cell infiltration, demyelination, gliosis, neuronal/axonal damage and loss [3]. These processes affect both the white and grey matter of the CNS, though in a heterogeneous fashion. White matter consists mostly of glia and myelinated axons, while grey matter comprises mostly of neuronal cell bodies/dendrites, some myelinated as well as unmyelinated axons and glia, and together they make up the brain parenchyma [26].   The common relapsing-remitting form of the disease exhibits multifocal inflammation and new lesion formation in the white matter of the brain and spinal cord [27]. White matter lesions associated with the relapsing-remitting stage of the disease are called classical active and chronic active lesions, with profound infiltration of lymphocytes, both T and B, along with significant populations of myelin debris-containing macrophages, encompassing the most pronounced inflammation seen in MS [27]. While grey matter/cortical lesions are fewer at this stage of the disease, they do also exhibit classical active lesions, particularly in the cortical parenchyma [28].  At conversion to the progressive phase, the disease manifests with underlying neuronal and axonal damage and/or loss contributing to the accumulation of disability [29] [30]. Progressive stages of MS are signified by a general lack of inflammatory activity with worsening of clinical symptoms [2]. White matter lesions associated with the progressive stage of the disease include slowly expanding lesions, which are characterized by an inactive lesion core 5  surrounded by microglia/macrophages, with only few containing myelin degradation products, chronic inactive lesions, which have no microglial/macrophage activity around their edges, and remyelinated shadow plaques [27] [31].  Furthermore, the progressive phase is associated with an increase in cortical lesions [32] [33]. There are three types of cortical lesions in MS: cortico-subcortical compound lesions (affecting both grey and white matter), small intra-cortical lesions, and subpial lesions, characterized by superficial demyelination extending into deeper cortical layers [27]. Subpial lesions are the most abundant and are associated with meningeal inflammation, and are unique to MS compared to other neuroinflammatory diseases [34]. White matter damage manifests as motor/movement difficulties, while grey matter damage is correlated with cognitive decline [26][35].    There are three pathways of lymphocyte entry into the CNS: entry from the bloodstream across the choroid plexus, from blood vessels in the subarachnoid space into the cerebrospinal fluid, or directly into the parenchyma under inflammatory conditions [36]. Myelin-specific T cells recruit macrophages and B cells as they enter the CNS [36]. Inflammatory infiltrates mainly consist of T helper cells of the CD8+ subtype (MHC class I restricted), and can cause cell death in two different ways: they can contribute to damage directly via cell to cell death, or indirectly via secretion of pro-inflammatory cytokines, which stimulate B cells, microglia and macrophages to mount an attack against the myelin sheath, via antibodies, free radicals (including superoxide, hydroxyl radicals, hydrogen peroxide and nitric oxide), and proteases, resulting in the generation of focal demyelinating lesions in both white and grey matter [37] [38] [39].   Under pathological conditions, microglia can play paradoxical roles, enhancing disease progression through free radical-mediated mitochondrial damage and axonal degeneration, while 6  also promoting repair and remyelination by cleaning up myelin debris and releasing neurotrophic factors [5] [22].  Furthermore, changes in astrocytic morphology and function are observed. Under inflammatory conditions, astrocytes influence disease progression through hypertrophy, proliferation and conversion to a reactive gliosis state, seen within and at margins of inflammatory demyelinating lesions [40]. In this state, they can also play paradoxical functions; blocking repair and remyelination of axons, while also assisting via glial scar formation, which provides an environment beneficial for repair [40] Astrogliosis may prevail in chronic disease stages, while microglial activity correlates with early plaque phases [40] [6].  Upon demyelination, axonal conduction of action potentials is reduced/blocked, which is followed by compensatory restoration through the redistribution of sodium channels along demyelinated segments, resulting in inefficient non-saltatory conduction along the axolemma (the cell membrane of an axon) [41].   Reduced production of adenosine triphosphate (ATP) due to mitochondrial dysfunction, along with enhanced energy demands for sodium channel reorganization, prevents proper Na+/K+ ATPase activity, which is required in higher amounts to move sodium ions out of the cell, resulting in intracellular sodium ion accumulation [41] [42]. This prompts the sodium-calcium exchanger to operate in reverse, resulting in a rise in intracellular calcium, which activates sodium channels, contributing to further sodium influx [42]. Additionally, increased intracellular sodium results in additional calcium release from intracellular stores, which can damage the mitochondria and activate nitric oxide synthase, proteases and lipases, which lead to neuronal cell death [43]. Increased mitochondrial strain leads to mitochondrial reactive oxygen species (ROS) production, which are highly reactive and damaging compounds to the cell through 7  oxidative stress [41]. This, together with a lack of trophic support from myelin, may lead to impaired axonal mitochondrial function, contributing to axonal degeneration/loss [44].   1.1.4 Neurodegeneration and axonal loss  The two primary components of neurodegeneration are neuronal loss and axonal damage [45]. Neuronal injury and loss in the context of demyelination and astrogliosis contributes to extensive cortical, subcortical and deep grey matter atrophy, while axonal loss in MS is found in white matter lesions (associated with focal inflammation), and also normal appearing white matter [45]. These processes are linked with development of clinical disability [45]. Processes of axonal and neurodegeneration are intertwined, as each one can result in development of the other [45].   Like MS, the final event in Parkinson’s and Alzheimer’s disease (PD and AD) is neurodegeneration [46]. Furthermore, like MS, PD and AD exhibit inflammation, microglial activation and activity, astrogliosis, oxidative damage and mitochondrial dysfunction [46] [47]. AD is also characterized by neuronal loss in the grey matter/cerebral cortex, whereas PD specifically affects dopaminergic neurons of the substantia nigra, resulting in death of these neurons and a lack of innervation to the basal ganglia [47] [48]. Unlike MS, AD and PD both have distinct biomarkers (neurofibrillary tangles and amyloid plaques in AD, Lewy bodies in PD) [47] [48]. In MS, AD, and PD, mitochondrial injury, commonly triggered by free radical-mediated damage, has been suggested as the common pathway of neurodegeneration [47] [48]. Sources of free radicals in all three diseases can include microglia and injured mitochondria [47] [49].  8   Cognitive deficits are more prevalent in the progressive phase than in RRMS, and arise from grey matter neuronal loss, observed as cortical atrophy [26]. However, subpial demyelination and grey matter atrophy are also observed in very early RRMS, and during clinically isolated syndromes (a first episode of neurological symptoms lasting at least 24 hours), suggesting the presence of early neuronal loss [26] [50]. The majority of grey matter lesions are described as “non-inflammatory”, though immune-cell mediated grey matter demyelination has also been described (particularly subpial lesions associated with meningeal inflammation) [26]. Moreover, neuronal loss can occur in both grey matter demyelinated lesions and normal-appearing grey matter, indicating that this damage can occur independently from demyelination [26]. There is evidence for a link between loss in grey matter volume and white matter volume, as well as retrograde neuronal loss through axonal damage [26]. Additionally independent grey matter damage, such as more extensive and independent cortical demyelination compared to white matter demyelination, and mitochondrial injury (induced by microglia/macrophages in both grey and white matter) have been described [26]. Cognitive deficits include reduction in memory and processing speed, attention and concentration, fatigue, and depression/anxiety, amongst others [51].   Axonal degeneration in MS is the main determinant of irreversible clinical disability, and can be divided into two stages, the first representing acute axonal injury in inflammatory MS lesions, associated with primary demyelination, and correlating with lesion activity. The second form is axonal degeneration in non-inflammatory chronic lesions (diffuse axonal injury) (specifically in inactive white matter lesions), associated with inflammation and microglial activity also, but present in whole brain and spinal cord as well as with non-demyelinated neurons [52] [53].  9   Acute axonal injury is characterized by axonal spheroids and end bulbs (representing areas of axonal transection), and can be detected by focal accumulation of proteins (such as amyloid precursor protein, or APP) [54]. This type of injury is mainly seen in actively demyelinating lesions [55]. Transections can lead to Wallerian degeneration (degeneration of the axon distal to the lesion, or anterograde), resulting in focal axonal degeneration [54]. The progressive loss of demyelinated axons that persists after lesion activity is thought to majorly contribute to global axonal loss [55]. Importantly, a fraction of axons in early stages of focal axonal degeneration can recover, and could thus be targeted in neuroprotective approaches for neuroinflammatory disease [56].  Acute axonal loss/axonal degeneration can occur as a result of demyelination resulting in lack of trophic support from oligodendrocytes, or due to release of inflammatory mediators resulting in direct immune cell cytotoxicity, proteolytic enzymes, glutamate excitotoxicity, and oxidative stress, all contributing to mitochondrial dysfunction [42] [57]. In particular, microglia and macrophages closely associate with axonal injury in MS lesions early in disease and are a main source of proteo- and lipolytic enzymes, cytotoxic cytokines, excitotoxins and reactive oxygen or nitric oxide species [57]. Furthermore, in a cyclical fashion, later mitochondrial injury in turn can contribute to demyelination and axonal degeneration, through disrupted ATP production and free radical-mediated damage [42] [57].   In more progressive stages of the disease process, there is a loss of oxidant-antioxidant balance as ROS production overwhelms antioxidant capacity, contributing to cellular injury via free radical-mediated oxidative stress, which causes damage to various cell components including proteins, lipids and DNA [40] [41]. Microglia and macrophages, found in lesion sites, are a major source of early ROS production, along with mitochondria respiratory chain 10  dysfunction [41]. Oxygen and nitric oxide radicals are of particular importance in axonal injury and loss, and demyelinated axons are more vulnerable compared to intact myelinated ones, though injury of myelinated axons is also found in MS [52]. Nitric oxide and superoxide form peroxinitrite, considered the final, common toxic effector molecule mediating damage induced by oxidative stress in MS and other diseases [58]. Peroxynitrite is formed early, correlating with disease activity, and has been consistently shown in acute and chronic active MS lesions, but not in chronic inactive lesions [58]. Like peroxynitrite, hydrogen peroxide is relatively stable and is capable of traveling far distances and generate reactive hydroxyl radicals [41].  As another secondary disease mediator, glutamate excitotoxicity results due to heightened glutamate release [59]. Glutamate is the major excitatory neurotransmitter in the human CNS [59]. Under normal physiological conditions, astrocytes, microglia and oligodendrocytes take up glutamate from the extracellular space via excitatory amino acid transporters, converting it to glutamine through mechanisms that rely on ATP [50]. Glutamine is then taken up by neurons once it is released and converted back to glutamate via the enzyme glutaminase [59]. Under pathological conditions, ATP depletion results in glial cell dysfunction, increased intracellular glutamate, inward transporter reversal and overload of extracellular glutamate [58]. Other sources of extracellular glutamate include release from injured cells, presynaptic vesicular release and release by macrophages/microglia/lymphocytes [58]. Uptake of excess glutamate into a cell results in calcium overload, damage to the mitochondria, activation of proteases and apoptotic pathways, and peroxynitrite production (which can then further induce glutamate excitotoxicity) [58].   Furthermore, in MS a combination of apoptosis and necrosis-mediated neuronal death is observed [34]. While apoptosis is a programmed form of cell death, necrosis is characterized by 11  non-regulated cell swelling and lysis, seen by a loss of membrane integrity [60]. Apoptosis is particularly found in neurons of demyelinated cortical lesions, and is also a mediator of oligodendrocyte cell death [61].   While axonal loss can occur early in the disease course of both MS and EAE, because of compensatory mechanisms within the CNS, clinical consequences of this process are not detected until a threshold level is reached and compensatory resources are depleted [53]. In MS patients, axonal loss can be non-invasively characterized by loss in neuronally enriched N-acetylaspartate and progressive CNS atrophy, and in post-mortem brain tissues by accumulation of APP, indicating axonal transections [53] [55]. Furthermore, in chronic MS lesions, it has been shown that axonal density is reduced by an average 60% [53].  Two models for the relationship between inflammation, demyelination and neurodegeneration with respect to axonal loss has been proposed in which damage can occur either from the outside-in or the inside-out [62]. In the outside-in model, demyelination occurs initially and leads to axonal injury, which can cause more demyelination in turn, forming a cycle of damage, both of which are secondary to inflammatory cells and their mediators [62]. However, this model fails to explain loss of axons with intact myelin, progressive worsening of clinical symptoms after the inflammatory phase has subsided, and neurodegeneration which sometimes precedes inflammatory cell infiltration in EAE [63] [64].   Alternatively, in the inside-out model, axonal damage occurs due to a primary neuronal difficulty, such as energy imbalance, with secondary mitochondrial dysfunction and enhanced production of reactive oxygen species, cell stress and damage, eventually leading to inflammation [62]. Animal models have been developed for the purpose of understanding 12  different components of the underlying disease process and how they interact to induce disease/disability, and some of these will be discussed in the next section.  1.1.5 Experimental autoimmune encephalomyelitis  The most common animal model of MS is EAE, and there are a variety of methods used for its induction, including active immunization and adoptive (passive) transfer [65]. Active immunization involves injection with an encephalitogenic (i.e. able to cause CNS inflammation) myelin protein, an encephalitogenic peptide of a myelin protein, or injection of spinal cord homogenate [65]. Adoptive (passive) transfer of spleen and lymph node cells (purified for myelin-specific CD4+ T helper cells) from an immunized animal into a naïve animal can also be used to induce disease [65] [66].   The clinical course/presentation of EAE that develops depends on the myelin protein/peptide, the mouse strain used, and localization of disease pathology. For example, when myelin oligodendrocyte glycoprotein (MOG) is injected subcutaneously into C57BL/6 mice, it induces chronic progressive EAE, which is characterized by ascending paralysis due to damage localized to the spinal cord, with little pathology seen in the brain [66]. Exposing mice to UV light before subcutaneous injection of MOG peptides in complete Freund adjuvant results in secondary progressive disease [66]. Injecting SJL mice with proteolipid protein (PLP) peptide or adoptive transfer of PLP peptide-specific T cells induce relapsing-remitting disease [66]. Spontaneous relapses and remissions can also be induced in SJL mice injected with mouse spinal cord, pertussis vaccine, and complete Freund adjuvant containing Mycobacterium tuberculosis [66].  13   One of the main challenges in developing animal models is the multifactorial and heterogeneous nature of MS disease progression [67]. Furthermore, aggressive induction in models of EAE is unlike the nature of MS onset in humans, and there are no naturally occurring spontaneous EAE animal models [66]. However, models of spontaneous induction have been developed, and are thought to resemble MS features of paralysis, demyelination and axonal degeneration more closely [66]. These include a C57BL/6 mouse model carrying MOG-specific T cell and B cell receptors, which develop chronic progressive opticospinal EAE [5]. Furthermore, transgenic mice with a MOG-specific T cell receptor, backcrossed to SJL background or C57BL/6 MOG-knockout mice, can also develop spontaneous disease [5]. In particular, T cell receptor transgenic SJL mice can experience damage in the optic nerve and cerebellum, in addition to the spinal cord [5].  Similarities of EAE to MS include early spinal cord neuronal loss, primary initiation by CD4+ T cells, and clonal expansion of CD8+ T cells [52]. Furthermore, MOG-induced EAE has many of the same pathophysiological processes as in MS, including encephalitogenic T cell responses with axonal damage [52]. Both EAE and MS exhibit pronounced axonal injury and loss in lesions, and have similar downstream mechanisms of axonal injury [52]. Furthermore, in both EAE and MS, small-diameter axons are more highly sensitive to axonal degeneration [53]. These similarities allow for EAE, including the C57BL/6 mouse model, to be used to test potential neuroprotective strategies for reducing/preventing axonal damage [52].   Differences between MS and EAE include: in EAE primary demyelination is sparse, axonal destruction with secondary demyelination is profound, and resemblance of pathology in the progressive stage is poor [52]. There is a lack of neurodegeneration in the absence of inflammation, and CNS damage associated with CD8+ T cells and B cells is not as prevalent 14  [67]. For this reason, transgenic and particularly, humanized EAE models may be better suited to address the immunological intricacies seen in MS [67].   1.1.6 Current strategies of neuroprotection  Animal models of MS have been crucial in the development of disease-modifying therapies in MS. Anti-inflammatory, immunosuppressive, immunomodulatory, and immune cell migration-inhibiting strategies are the current standards of disease-modifying therapies, but are only effective in the relapsing-remitting form of the disease, and do not fully prevent progression of disease/disability [45] [67]. The more commonly used therapies can be generally categorized into two groups. The first line therapies, which include interferon- β1a (Avonex®), interferon-β1b (Betaseron®) and glatiramer acetate (Copaxone®), have been shown to provide a reduction in relapse of 29-34% compared to placebo over a two-year period, but exhibit only modest efficacy in slowing progression of disability [68] [69]. Furthermore, phase 3 trials on the follow up of patients on interferon-β therapy have shown a relapse rate of 62 to 75% [70] [71] [72]. The second group includes fingolimod (Gilenya®), natalizumab (Tysabri®) and mitoxantrone (Novantrone®). Natalizumab, when compared to placebo, demonstrated a relapse rate reduction of roughly 65% and reduced risk of progression of disability over two years of about 42% [73]. Fingolimod has been shown to reduce annual relapse rates by more than 50% compared to placebo, and after a two-year follow-up 63-70% of patients were relapse-free [74]. Mitoxantrone treatment prolonged the time to first relapse by roughly 10 months compared to placebo and a significant reduction in progression of disability [75] [76]. While these drugs have higher efficacy than the first line therapies, they are associated with greater health risks, including progressive multifocal leukoencephalopathy (PML) with natalizumab and fingolimod, as well as 15  skin cancer with fingolimod, and cardiac toxicity and acute myelogenous leukemia with mitoxantrone, and are therefore generally reserved for patients with rapidly worsening or progressive relapsing MS [77] [78] [79] [80] [81].  There is a crucial need for therapeutic agents for progressive MS that act in a neuroprotective manner at the site of damage, by allowing remyelination, axonal repair/regeneration and restoration of nerve conduction [82]. An ideal therapeutic strategy in MS would be focusing on inhibition of axonal degeneration and on protection against neuronal cell death whilst promoting repair, in addition to conventional immune modulation [5]. This would include boosting endogenous repair mechanisms, preventing activity of remyelination inhibitors/enhancing oligodendrocyte survival, and blocking excitotoxicity/oxidative stress-induced neuronal injury, amongst others [45] [67].   Neuroprotective agents targeting molecular imbalances in MS include glutamate antagonists (eg. Riluzole), nitric oxide blockers (Furoxan), sodium channel blockers (eg. Flecainide, Phenytoin, Lamotrigine), potassium and calcium channel blockers (eg. Nimopidine, Nifedipine, Ryanodine), cannabinoid receptor antagonists, and erythropoietin/insulin-like growth factor, amongst others [45] [67]. These compounds have all been beneficial in EAE but can have negative effects on the immune system (sodium channel blockers) or the potential for severe toxicity (glutamate antagonists) [45] [67].  Other than modulation of channels involved in MS pathogenesis, remyelination has also been studied as a neuroprotective target. Remyelination of neurons, which protects axon integrity, is limited in the central nervous system for various reasons, and is dependent on oligodendrocyte precursor cells (OPCs) differentiating into mature oligodendrocytes capable of myelination [83]. Upon injury oligodendrocytes either undergo programmed cell death or enter a 16  resting state [83]. Macrophages and microglia are responsible for myelin debris removal, which is crucial for remyelination and neuroaxonal regeneration to occur, as myelin debris contains several inhibitory factors (myelin associated growth-inhibiting factors, or MAIs) [84]. This slow rate of myelin clearance hinders the capacity of regeneration in the CNS [84].  Furthermore, endogenous neuroprotective pathways that could be targeted to enhance remyelination/axonal regeneration include the activity of trophic factors, such as brain-derived neurotrophic factor (BDNF), amongst others [85]. In MS lesions, BDNF is primarily present in T cells, macrophages, microglia and reactive astrocytes, but is also expressed in neurons [85]. Mice lacking immune cell-derived BDNF show enhanced axonal loss and progressive disability in the chronic form of EAE [86]. Mice deficient in astrocytic BDNF exhibit more severe disease and increased axonal loss [86]. BDNF has been shown to be capable of rescuing spinal and bulbar motor neurons following axonal transection and promote axonal preservation and regeneration [86] [87]. Previous studies show that in actively demyelinating lesions, a higher percentage of BDNF+ cells are detected compared to inactive lesions, that immune cells are the major source of BDNF in MS lesions, and that neurons localized to active lesions as well as reactive astrocytes express high levels of one of the isoforms of the BDNF receptor (gp145trkB), which is not found in immune cells [85]. Furthermore, autocrine and paracrine interactions exist between neurons releasing BDNF and expressing gp145trkB receptors [85]. Interestingly, BDNF levels are reduced in patients with MS, and both glatiramer acetate and fingolimod can enhance BDNF levels in vivo [2] [88]. Endogenous neuroprotective factors and their regulators provide a viable target for the development of therapies to prevent damage at the site of disease activity.  17  1.2 ARNT2 1.2.1 Overview  The aryl-hydrocarbon receptor nuclear translocator 2 (ARNT2) is a member of the basic-helix-loop-helix PER-ARNT-SIM (bHLH/PAS) transcription factor family, which generally function by heterodimerizing with other members to direct transcription of target genes in response to various environmental and physiological stimuli [89]. ARNT2 was first described as an isoform of the aryl hydrocarbon receptor nuclear translocator (ARNT), named after its purported role in the nuclear translocation of the aryl hydrocarbon receptor (AhR) [90]. This was refuted later as ARNT and ARNT2 were found to be constitutively expressed in the nucleus and ARNT knockdown did not result in loss of AhR nuclear translocation [91]. ARNT2 shares a >90% amino acid identity with ARNT in the bHLH and >80% in the PAS domain, but while ARNT2 is expressed only in the CNS, kidneys (specifically the tubules), urinary tract, and thymus, ARNT is ubiquitously expressed throughout the body [90] [91] [92] [93]. It has been proposed that ARNT2 developed to partly compensate for some of the functions of ARNT specifically in the CNS [94].  ARNT2 is a member of the class VII helix-loop-helix superfamily of transcriptional regulators, which are classified by a basic region prior to the HLH domain (bHLH), as well as the PAS domain [95]. These are further subclassified into two phylogenetic groups: the AhR group (class I), consisting of proteins AhR, SIM1, SIM2, the HIF-1 family (HIF-1α, HIF-2α, HIF-3α), and HLF, and Arnt group (class II), including proteins ARNT, ARNT2, PER, BMAL1 (ARNT3, MOP3, JAP3, ARNTL1, TIC) and BMAL2 (ARNT4, ARNTL2, MOP9) [95]. Proteins of the first group heterodimerize with members of the second group, though homodimerization can also occur within the class II group [95]. Primary dimerization occurs within the N-terminal 18  bHLH domain, which binds the region within a promoter capable of regulating gene transcription, termed a response element, while the secondary dimerization interface, which defines partner choice/target gene recognition, is the PAS domain [95].   ARNT2 also contains a nuclear localization sequence that interacts with nuclear pore targeting complex proteins for efficient nuclear localization [91]. Deletion of this region results in a cytosolic protein that remains capable of interacting with the AhR [91]. In response to a stimulus, bHLH/PAS transcription factors undergo heterodimerization, bind to a response element within the promoter of the gene of interest via their bHLH domain, and induce transcription of genes required for responding to the stimulus [95].  Loss of ARNT2 has been linked to a lack of hypothalamic neuroendocrine lineage formation, impaired regulation of HIF-1 target genes and thymic defects [96] [97] [98] [99]. ARNT2-null mice and rats die perinatally (within the first 24-48h to 2 weeks after birth), most likely due to improper hypothalamic development and lack of pituitary hormone secretion [97] [98].  1.2.2 Expression patterns  Expression of Arnt2 in the CNS has been described in mouse, rat, human, Xenopus, and zebrafish, mostly via the use of in situ hybridization techniques [92] [95] [100] [101] [102] [103] [104]. In the developing mouse and rat, Arnt2 gene transcripts are highly expressed in the forebrain, neuroepithelium, hypothalamus and habenula (measured at embryonic day 16) [92] [104]. Gene transcripts are moderately expressed in the striatum, and no expression is found in the choroid plexus or the internal capsule [92] [101] [104]. In the adult mouse and rat, Arnt2 gene transcripts are strongly expressed in the cerebral cortex (particularly in the occipital and 19  piriform cortex) and hippocampus (particularly in its ventral part), moderately expressed in the striatum, pons, dorsal midbrain and cerebellum, and not expressed in the olfactory bulb [92] [101] [95]. In the hippocampus Arnt2 is expressed heterogeneously along the Schaeffer collaterals, with highest expression at the CA2 region and gradual reduction in expression towards and including the CA1 region [105] [106]. The thalamus also heterogeneously expresses Arnt2 RNA transcripts, with much higher Arnt2 expression in the posterior region [101] [104]. The expression of ARNT2 in other cells of the CNS (in astrocytes, specifically) has only been described once previously to our knowledge [107].   1.2.3 Functional roles   The bHLH transcription factors control gene expression activity in fundamental biological processes including homeostasis, stress response pathways, cell cycle maintenance and early cell determination and differentiation [89]. While ARNT2 expression is maintained in the nucleus, the majority of its binding partners are kept in the cytoplasm and must relocate to the nucleus for dimerization and functioning, with the exception of neuronal PAS domain protein 4 (NPAS4), which is also nuclear [89] [95].   bHLH transcription factors can regulate themselves by binding the DNA sequence of their own gene to down-regulate/up-regulate production, quickly altering transcription factor levels under various stimuli while also maintaining baseline transcription factor levels [105]. The activity of ARNT2 and its heterodimerization partners can also be negatively regulated [105]. Necdin, a member of the melanoma antigen (MAGE) protein family, thought to be a growth suppressor predominantly expressed in postmitotic neurons, acts as a transcriptional repressor in modulating ARNT2:HIF-1α and ARNT2:SIM1 [105]. Furthermore, a splice variant of ARNT2 20  leads to a truncated protein that can potentially act as a dominant negative factor in the regulation of ARNT2 [108].   The binding partners of ARNT2 fall under the class I bHLH/PAS factors, which include hypoxia inducible factor 1α (HIF-1α), single-minded homolog 1 (SIM1) and the aryl hydrocarbon receptor (AhR) [95].  Under ischemic/hypoxic conditions, the oxygen-regulated HIF-1α becomes activated and travels to the nucleus, where it heterodimerizes with ARNT2 in the CNS [109]. Once heterodimerized, ARNT2/HIF-1α bind to the hypoxia response element (HRE), directing transcription of genes required in cellular metabolism (glycolytic enzymes, phosphoglycerate kinase), glucose transport (GLUT-1 and -3), and angiogenesis (VEGF), combating the hypoxic stimulus [109].   At mouse embryonic day 18.5, ARNT2 and SIM1 guide differentiation of dopaminergic and neuroendocrine neurons within the paraventricular, anterior periventricular and supraoptic nuclei of the hypothalamus, including those responsible for the synthesis and release of corticotrophin-releasing hormone, thyrotropin-releasing hormone, arginine vasopressin, oxytocin and somatostatin [97] [110]. ARNT2-null mice die perinatally as a result of impaired hypothalamic development, phenocopying the defects seen in SIM1-null mice [110]. The differentiation of these cells into cells expressing these hormones is controlled through maintenance of the expression of BRN2 by the SIM1:ARNT2 complex [110].   The Ahr is a receptor for environmental toxins of the dioxin group, as well as endogenous ligands such as kynurenine (which helps regulate processes in inflammation and immune activation) [111]. Following translocation to the nucleus, AhR heterodimerizes with ARNT2, binding the xenobiotic response element (XRE), and inducing transcription of genes for drug 21  metabolizing enzymes, such as cytochrome P450Ia1 (CYP1A1) and glutathione S-transferase (GST), which is also involved in cellular pathways aimed at reduction of oxidative stress [111].  ARNT2 and NPAS4 function together to aid in neuronal survival and cognitive function, through the transcription of BDNF [112]. ARNT2 and NPAS4 have been mostly studied in the hippocampus, where ARNT2 is a central regulator of gene expression, and NPAS4 functions in an activity-dependent negative feedback mechanism to regulate inhibitory synapse development, maintaining excitatory-inhibitory balance [105] [113]. Furthermore, lack of NPAS4 expression has been linked to neurodegenerative diseases and stroke/ischemia models [114] [115] [116]. Under excitatory conditions, a rapid and enhanced influx of calcium into the cell induces NPAS4 transcription, followed by heterodimerization with ARNT2, binding the PER-ARNT-SIM response element (PASRe) and guiding transcription of BDNF at BDNF promoters I and IV of a total of nine promoters [117] [118]. BDNF homozygous null mutants have a shortened life span, and impairments in hippocampal-dependent memory formation, with the latter also seen in heterozygous mutants [119]. Furthermore, decreased BDNF expression has been linked to the development of neurodegenerative diseases, cognitive impairments and affective disorders [119]. The trophic influence of BDNF on neurons is well understood, enhancing neuronal survival by increasing the density of synaptic terminals, promoting growth/regeneration of axons and dendrites, and regulating neurotransmission, amongst others [89]. By regulating these processes BDNF is essential for maintaining proper synaptic function, and has been studied as a potential endogenous therapeutic neuroprotective agent in multiple sclerosis as described in section 1.1.5. [112].   ARNT2 and NPAS4, in their heterodimerized state, also induce the transcription of FBJ Murine Osteosarcoma Viral Oncogene Homolog (FOS), early growth response protein 1 22  (EGR1), activity-regulated cytoskeleton associated protein (ARC), and drebrin [114] [120]. These, like NPAS4 and BDNF, are all immediate early genes, which are activated transiently and rapidly in response to various cellular stimuli [121]. FOS is involved in cell proliferation, differentiation and survival, and in limiting hypoxic damage and promoting angiogenesis [121]. EGR1 may play a role neuronal plasticity (i.e. experience dependent reorganization) and regulation of a protein involved in synaptic exocytosis called synaptobrevin II [121]. Arc mRNA localizes to activated synaptic sites in an NMDA receptor-dependent manner, while its translated protein is thought to play a critical role in learning and memory-related processes [122]. Drebrin is important in synaptic dendritic-cytoskeleton modulation [114].  1.2.4 ARNT2 as a neuronal survival factor              Webb et al. described the case of a family with six children, with a frameshift mutation within Arnt2, resulting in reduced levels of transcript and protein due to the introduction of a premature termination codon, and perinatal death [103]. In these children, congenital kidney and urinary tract defects were found, as well as abnormal hypothalamic and pituitary development [103]. Furthermore, frontal and temporal lobes were hypoplastic, with a thin corpus callosum and a global delay in brain myelination, particularly in the motor and occipital cortices [103]. This microcephaly indicates progressive neurodegeneration, potentially caused by decreased dendritic connection or activity of neurons [103].   Drutel et al. have provided the sole in vitro study to investigate the role of ARNT2 in cell survival in PC12 cells, a cell line derived from a pheochromocytoma of the rat adrenal medulla, with embryonic origin from the neural crest with a mixture of neuroblastic cells and eosinophilic cells [93]. They found that ARNT2 acts as an anti-apoptotic factor in PC12 cells, as 23  downregulation of ARNT2 resulted in apoptosis of these cells. In this study it was also shown that oxidative stress-mediated PC12 cell death was preceded by rapid and strong downregulation of ARNT2 protein, and that cells could be rescued with introduction of Arnt2 cDNA. Furthermore, in a rat model of focal cerebral ischemia, Arnt2 gene transcript levels declined significantly following 2 hours of blood recirculation, preceding neuronal death at 24 hours of blood recirculation [93]. In addition, Arnt2 was found to be differentially expressed in cell cycle regulation of PC12 cells, and increased in expression when the cycle was inhibited at various stages [93]. Knockdown of Arnt2 by treatment with Arnt2 antisense oligonucleotide resulted in an arrest of cell division. Furthermore, previous studies have shown that Arnt2 knockout results in perinatal lethality, further indicating a role in maintaining cell viability/preventing cell death [97] [98].   1.2.5 A role for ARNT2 in inflammatory neurodegeneration   Our lab became interested in examining ARNT2 following a previous study of chronic progressive EAE in animals with antioxidant feed (switched at day 7 post-immunization, when disease is starting) versus control EAE mice. Microarray analysis (Affymetrix Mouse Genome ST 1.0 GeneChip, Santa Clara, CA) of differential gene expression revealed that Npas4 gene expression was lower in TEMPOL-fed versus control EAE (fold change -2.07) at disease pre-onset (day 11) (Quandt lab, unpublished data).  This was followed up by a second EAE study in which tissues were collected at pre-onset (day 7), onset (day 14), peak (day 18), and recovery/stabilization (day 29) stages of the disease and processed for mRNA analysis via qPCR. It was found that in the brain, there was a 24  significant upregulation in both Npas4 and Arnt2 mRNA at the pre-onset stage of the disease in the brain, as well as a significant downregulation in spinal cord Npas4 and Arnt2 mRNA at peak  disease, when/where inflammatory infiltrates and axonal loss are at their highest (Figure 1.1, Quandt lab, unpublished data). This was also concurrent with a reduction in Bdnf mRNA at peak disease in the spinal cord (Figure 1.1, Quandt lab, unpublished data). Furthermore, there was a suggestion of enhanced ARNT2 protein expression in glial populations as measured via immunohistochemistry at peak disease, and was also detected in the infiltrates within the meninges and subarachnoid space of the spinal cord, likely immune cells (Figure 1.2, Quandt lab, unpublished data).   While the importance of ARNT2 in cell survival has been shown in models of ischemic injury, its expression patterns/role in chronic inflammatory and neurodegenerative disease of the CNS had not been characterized [93]. Furthermore, it is likely that ARNT2 may be regulated in response to factors involved in axonal degeneration in MS including oxidative stress and glutamate excitotoxicity, as both of these enhance intracellular Ca+2 levels, which is the upstream regulator in the production of NPAS4 and binding to ARNT2 in the transcription of BDNF [117] [123]. For this reason, we decided to continue our investigation of this protein in both in vitro and in vivo models of MS.       25   Figure 1.1: Arnt2 and Bdnf spinal cord and Arnt2 brain gene expression over the course of chronic progressive EAE. (A) Arnt2 and Bdnf  in the spinal cord follow a similar pattern, with a significant reduction in gene expression levels at peak EAE disease (day 18). (B) in comparison, Arnt2 gene expression in the brain follows a different pattern, with significant upregulation pre-onset of disease (day 7). Healthy animals were immunized with CFA and given pertussis without MOG35-55 to control for immunization/pertussis effects, and littermates (“LM”) received no immunization or pertussis. Bars represent mean±standard deviation, p≤0.05*, Mann Whitney t-test compared to healthy mice. (Quandt et al., unpublished data)  * * !!!!!!!!!!!!!!Arnt2/B-actin ratio x 104  Arnt2/B-actin ratio x 103  A B Spinal cord Brain !!!!!!! Bdnf/B-actin ratio x 104  * 26   Figure 1.2: ARNT2 protein detection in the spinal cord of healthy and EAE mice. (A) staining of healthy spinal cord with hematoxylin, eosin, and anti-ARNT2 antibody (rabbit) shows intense nuclear staining of ARNT2 (pink-red) of the anterior horn cells, with occasional glial cells staining in the white matter, (B) EAE mice at peak disease show neuronal staining as in (A) but also show increased numbers of ARNT2+ cells, possibly glia or immune cells, in regions of inflammatory infiltrates focused in the white matter (black rectangle). (Quandt et al., unpublished data)   1.3 Hypothesis  We hypothesize that alterations in ARNT2 expression are associated with changes in cell viability in in vitro and in vivo models of multiple sclerosis.  Specifically, this work was performed to address the following aims:  1. Study the ability of stressors involved in MS pathogenesis to alter ARNT2 protein  expression in primary cortical neuron-enriched cultures. a. Study the ability of stressors involved in MS pathogenesis to alter ARNT2 protein expression in primary cortical neuron-enriched cultures. b. Examine Arnt2 mRNA levels to determine whether any ARNT2 protein regulation is linked to changes in mRNA levels. A B 27  c. Examine ARNT2 expression within specific cell populations in primary cortical neuron-enriched cultures. d. Examine ARNT2 expression in primary cortical astrocyte-enriched cultures.  2. Optimize viability/cytotoxicity assays for sensitivity and to test potential to  measure ARNT2 intensity in concordance with cell viability changes.  3. Localize and compare ARNT2 expression throughout the initiation and progression  phases of disease in an animal model of MS.      28  Chapter 2: Materials and Methods 2.1  Primary cortical neuron-enriched cultures  Embryonic day 18 (E18) rat cortices, devoid of hippocampus but not striatum, were graciously provided to us by Shernaz Bamji’s research group (University of British Columbia), and were cultured as previously described [124]. Cortices were washed with 37°C Hank’s Balanced Salt Solution (HBSS, Gibco®, Carlsbad, CA), incubated in 0.25% Trypsin-EDTA (Sigma-Aldrich®, St. Louis, MO), DNase (Sigma-Aldrich®) and HBSS for 15 minutes, dissociated in 5ml of HBSS and seeded at 50,000 cells/cm2 for western blotting/qPCR, and 35,000 cells/cm2 for immunocytochemistry on poly-l-lysine coated plates (Corning, Corning, NY), in Neurobasal® media (Thermo Fisher Scientific™, Waltham, MA) supplemented with B-27 antioxidant cocktail (Gibco®). Cells were grown in an incubator (5% CO2, 37°C), and received a full media change at 3 days in vitro (DIV) and a supplementation with half of the media amount at DIV 10. Cells were ready to use between DIV 12-16.  2.2 Primary cortical astrocyte-enriched cultures  The protocol of creating a single-cell suspension was followed as described above [124]. For astrocyte-enriched cultures, cells were instead grown in Dulbecco’s Modified Eagles Medium (DMEM, Thermo Fisher Scientific™) supplemented with 10% horse serum (Gibco®). Media was replaced every 2-3 days for removal of neuronal debris as described previously [125]. Cells were ready to use between DIV 10-14, at which point cells reached roughly 75-95% confluence.  29  2.3 Treatments  Various compounds were used to mimic excitatory and pathological processes. We began our studies by testing doses/time points previously described in neuronal culture experiments in vitro. 4-aminopyridine (4-AP, Tocris, Avonmouth, UK), bicuculline (Bic, Sigma-Aldrich®) and potassium chloride (KCl, Fisher) were tested as excitatory stimuli, and in the case of KCl also tested at neuroprotective and cytotoxic doses [126] [127] [128]. Furthermore, Hydrogen peroxide (H2O2, Fisher, Waltham, US) was used to mimic oxidative damage, glutamate (Sigma-Aldrich®) was used to mimic glutamate excitotoxicity, staurosporine (Sigma-Aldrich®), a non-selective protein kinase inhibitor, was used to induce apoptosis, to mimic various pathological processes involved in axonal degeneration in MS [129] [130] [131].  2.4 Cell lysis, protein quantification, SDS-PAGE and western blotting  All treatments were collected in duplicate per biological replicate. Cells were washed 2x with 37°C PBS, lysed in radioimmunoprecipitation assay buffer (RIPA, consisting of tris-hydrochloric acid (MP Biomedicals, Santa Ana, CA), sodium chloride (Fisher), sodium deoxycholate (Fisher), NP-40 (Thermo Fisher Scientific™), sodium dodecyl sulfate (SDS, MP Biomedicals), and EDTA (MP Biomedicals)) with 5% protease inhibitor (Roche, Basel, CH) on ice, scraped using a cell scraper and centrifuged for 15 minutes at 13,200 rpm, at 4°C, after which the supernatant was collected and stored at -20°C. Protein quantification assays using bicinchoninic acid (Sigma-Aldrich®) and copper (II) sulfate (Sigma-Aldrich®) were performed with 1mg/ml bovine serum albumin (Thermo Fisher Scientific™) for standard curve generation. To the lysate 15% 5x loading buffer and 5% 2M 1,4-Dithiothreitol (DTT, Fisher) were added, and samples were loaded onto a precast 12% tris/glycine gel (Bio-Rad, Hercules, US), with 30  volume loaded adjusted to the same concentration, and run for 45 minutes at 200V. Subsequently, the gels were transferred to a nitrocellulose membrane (Bio-Rad) for 45 minutes at 90V, blocked with 2% skim milk solution containing 10% 10xTBS, and incubated with primary antibody overnight at 4°C (Appendix A.1). Membranes were washed with 0.05% TBST (Tris-buffered saline-Tween 20, (Sigma-Aldrich®)) 3x for 7 minutes, and incubated with secondary antibody at RT for 1hr (Appendix A.1). Membranes were washed again with 0.05% TBST 3x for 7 minutes, and Clarity™ Western ECL Blotting Substrate was added (Bio-Rad). Membranes were imaged using a Bio-Rad ChemiDoc™ MP System, with ARNT2 exposure at 20sec, and tubulin exposure at 10sec. Blots were analyzed using quantitative densitometry on Image Lab™ (Bio-Rad 5.2). ARNT2 levels were normalized to tubulin and untreated levels. The ARNT2 antibody used consistently gave a single band at 79 kDa. Prior to harvest phase microscopy images of cells were obtained, and supernatants were collected from the cells for cytotoxicity measurements described in section 2.7. The linear dynamic range was determined via a titration experiment of ARNT2 protein, where different quantities of cell lysate were loaded to determine the linear range and and set its minimum and maximum detectable concentrations under those assay conditions; all samples were diluted to ensure they would fall within this linear range.  2.5 Cell lysis, RNA isolation, reverse transcription- and quantitative polymerase chain reaction  All treatments were performed in duplicate per biological replicate. Cells were washed 2x with room temperature (RT) PBS, lysed with buffer RLT (Qiagen, Hilden, DE) containing 1% β-2 mercaptoethanol (Thermo Fisher Scientific™), scraped using a cell scraper, pipetted up and down 10x, vortexed for 1 minute and stored at -20°C. RNA was isolated using the Qiagen 31  RNeasy Mini Kit, and protein was precipitated using buffer APP (Qiagen) as per manufacturer’s guidelines. Lysates were transferred to an RNeasy spin column placed in a 1.5ml centrifuge tube, and spun at 11,000 g for 15s. The flow-through from this step was kept for protein precipitation. Columns were then washed with Buffer RW1, and RPE 2x (with a centrifugation step between each). The columns were then washed 2x with 70% ice-cold ethanol made up in ultrapure dH2O, and then eluted in 30ul RNase-free water (Qiagen). RNA quantity and purity (260/280, 260/230) were measured using a NanoDrop™ 2000/2000c spectrophotometer (Wilmington, US). A 260/280 reading of ~1.8 was considered pure, and a 260/230 reading of ~2.0 was considered free of contaminants [132].   The flow-through underwent APP precipitation as described in the All Prep DNA/RNA/Protein Mini Kit (Qiagen). 1 volume of Buffer APP was added to the flow through, mixed and incubated at RT for 10min, then centrifuged at full speed for 10min, after which the supernatant was decanted. 500ul 70% ethanol was added to the protein pellet, centrifuged at full speed for 1min, and supernatant removed. The pellet was dried for 10min at RT, and 100ul 5% sodium dodecyl sulfate was added and vigorously mixed. Protein quantification and western blotting protocols were performed as described in section 2.4.  Reverse transcription was performed using the QuantiTect® Reverse Transcription Kit (Qiagen), yielding a 20ul reaction as per manufacturer’s guidelines. The genomic DNA elimination reaction was prepared with 2ul gDNA Wipeout Buffer (1x concentration final), and template RNA/RNase-free water for a total volume of 14ul. This reaction was incubated for 2min at 42°C using a Thermocycler (PTC-200 Peltier Thermal Cycler, MJ Research, St. Bruno, CAN), and then placed back on ice. This was followed by the reverse-transcription reaction, set up with Quantiscript Reverse Transcriptase at 1ul, Quantiscript RT Buffer at 4ul (1x 32  concentration final), RT Primer Mix at 1ul, and the genomic DNA elimination from the previous step. This was incubated for 15min at 42°C, then for 2min at 95°C to inactivate Quantiscript Reverse Transcriptase.   Fast Start Essential DNA Green Master (Roche) was used for qPCR as per manufacturer’s instructions. 2ul cDNA sample, 5ul SYBR Green and 3ul ultrapure dH2O was combined, for a total 10ul reaction. Samples were loaded onto a 96-well plate (LightCycler®  480 Multiwell plate, Roche) in duplicate, and run on a LightCycler®  480 Instrument II (Roche). qPCR primers and run conditions can be found in Appendices B.1 and B.2, respectively. Data was subsequently analyzed using the LightCycler® 480 software (Version 1.5, Roche).   2.6 Immunocytochemistry 2.6.1 Protocol and visualization  All treatments were performed in duplicate per biological replicate. Cells were washed 2x with 37°C PBS then fixed with 10% phosphate buffered formalin (Fisher) for 20min [133]. Cells were then permeabilized using 0.1% Triton-X 100 (Fisher) for 10min, blocked with 10% normal goat serum (Gibco®) for 30min, then incubated with primary antibody in 2% normal goat serum overnight at 4°C. Cells were then washed 3x for 5min with PBS, and incubated with AlexaFluor® (Thermo Fisher Scientific™) secondary antibody for 1h at RT, and washed again 3x for 5min. Information on primary and secondary antibodies can be found in Appendix A.2. 4',6-diamidino-2-phenylindole (DAPI, a nuclear stain) was added at a 1 in 250 dilution for 5min, and cells were subsequently washed 2x with PBS. Cells were visualized on a Zeiss Axio Vert 200 Inverted Fluorescence Microscope (Oberkochen, DE).   33  2.6.2 Methods of analysis  For analysis of images, Zeiss software Zen (Version 2.0.0.0) was used. 8 images were taken of each well, with 2 technical replicates (i.e. 16 images per treatment total). An analysis protocol was designed to capture either all DAPI-positive nuclei (all cells), or DAPI nuclei surrounded by microtubule associated protein 2 (MAP2)-staining  (indicating neurons), and to quantify ARNT2 expression within that DAPI region. ARNT2 labeled with goat anti-rabbit AlexaFluor® 568 was expressed as mean pixel intensity over total nuclear pixel count (i.e. normalized intensity). For astrocytes, fields were manually scanned for DAPI positive nuclei within a glial fibrillary acidic protein+ (GFAP, an astrocytic marker) cell body. Cells less than 50um2 and greater than 200um2 were excluded from the analysis for their likelihood of being either debris or cell clusters, respectively. An isotype control for each species was used to control for species background fluorescence, and values below the mean of the isotype control plus two standard deviations were considered negligible. Appendix A.2 describes isotype controls used for immunocytochemistry.  To compare ARNT2 expression across treatments, we first determined the distribution of cells using a Shapiro-Wilk normality test. For all of our biological replicates, ARNT2 intensity across cells in our three groups (total DAPI, DAPI within MAP2+ cells, DAPI within GFAP+ cells), were not normally distributed. For this reason, we determined the median of each field examined (i.e. 8 fields per treatment, in duplicate), and took the average of each representative median of each treatment.  34  2.7 Lactate dehydrogenase assay  Cell viability measurements were performed using a lactate dehydrogenase (LDH) cytotoxicity assay kit (Thermo Fisher Scientific™ Pierce™). Supernatants were collected either in combination with a western blotting experiment as described in section 2.4, or treatments were added onto a 96-well plate directly. Supernatants were transferred to 96-well plate, in duplicate, with a substrate mix and assay buffer solution (50ul of each). As controls, supernatants from wells containing dead cells (killed with the lysis buffer provided; maximum LDH release) and wells which had 10% water added to them (for spontaneous release) were included. The plate was incubated at RT away from light for half an hour, after which stop solution was added (50ul), and the plate was measured using a plate reader (Spectra Max, Molecular Devices, Sunnyvale, US) and the fluorescence measured at excitation of 490nm and 680nm (which measures the background signal from the instrument). To determine LDH release, the 680nm values were subtracted from the 480nm values, and the spontaneous release control was subtracted. This value was then expressed as %cytotoxicity by subtracting the spontaneous LDH release value from the LDH release value of the sample, and dividing this number by the maximum LDH release minus the spontaneous LDH release, and multiplying by 100.   2.8 Live/Dead® viability/cytotoxicity assay  A second cell viability measurement was performed using a Live/Dead® viability/cytotoxicity kit (Molecular Probes®, Eugene, US). The protocol was executed as per manufacturer’s instructions, with each treatment in triplicate. Cells were washed once with PBS to remove media, and 100ul PBS was added to each well, followed by addition of two fluorescent dyes: calcein was tested between 1 and 2uM, and ethidium-1 homodimer (EthD-1) at 35  2uM, each made up in PBS (final volume per well: 200ul). Calcein is bound to an acetomethoxy (AM) group, and is able to enter live cells, non-fluorescent in this state [134]. Once inside, the AM group is removed via intracellular esterase activity, causing calcein to get trapped inside, giving off a green fluorescence [134]. EthD-1 can enter dead cells as their membrane permeability is compromised, and binds strongly to nucleic acids, giving off a red fluorescence [135]. A dead control (i.e. positive control for EthD-1) was included by killing cells with 70% methanol, as well as a control with cell-free wells with and without dye added, to determine the background signal. Also, wells with only one of the dyes added were included to control for dye interference. Cells were incubated for 10min at 37°C, then measured on a plate reader (Spectra Max). Calcein and EthD-1 were read at excitation/emission wavelengths of 494/517nm and 528/617nm, respectively.   2.9 EAE induction  8-week old female C57BL/6 mice were immunized for chronic progressive EAE as previously described [136]. Mice received 200ug of MOG35-55  (MEVGWYRSPFSRVVHLYRNGK, 95% pure by high-performance liquid chromatography; Stanford Pan Facility, Stanford, CA) in 4mg/ml Mycobacterium tuberculosis (H37Ra) in incomplete Freund’s adjuvant (IFA) (Difco Laboratories, Detroit, MI). While under anesthesia, 200ul of the emulsion was delivered in total to each mouse via 4 separate subcutaneous injections, 50ul per spot on back and rear flanks. In addition, 200ng pertussis (List Biologicals, Campell, CA) was delivered intraperitoneally. Representative tissues of the controls and chronic disease stage were observed in male mice, induced for disease the same manner at 12-14 weeks of age.  36   Two control groups were included: one sham-immunized with Mycobacterium tuberculosis (H37Ra), IFA and pertussis (to control for differences due to injection/pertussis), and one healthy littermate group. Two days after immunization, the disease and sham-immunized groups were given another dose of pertussis (200ng). In this model, disease is expected 14-15 days post-immunization. Mice were scored daily on a score from 0-5 as seen in Figure 2.1. Mice exhibiting clinical symptoms were given moistened feed on the cage floor to allow access. All animal work was in compliance with the University of British Columbia animal care guidelines.   2.10 Tissue collection, removal and processing of brain and spinal cord  Tissues were collected at pre-onset (days 7, 10), onset (days 14, 15), peak (days 18, 25), recovery and stabilization (days 32, 45) stages of the disease. Mice were euthanized with CO2, and subsequently brains were dissected out of the skull, and spinal cords were flushed with PBS out of the column using a 16g needle attached to a syringe. Both were subsequently chopped up and put directly into RLT buffer containing 1% β-2 mercaptoethanol, disrupted and homogenized, first with a 18g needle 10 times, then with a 20g needle 10 times. Tubes were subsequently spun at 1000rpm for 1.5min, and lysate was collected and aliquoted prior to storing at -80°C.  37   Figure 2.1: Chronic progressive EAE disease progression and associated clinical scoring. (Quandt et al., unpublished data)  2.11 Immunohistochemistry  Tissues collected from mice were fixed in 10% phosphate buffered formalin for 2-3 days, and put into 30% sucrose for stabilization of tissue for 3 days, then washed in PBS. Subsequently, tissues were embedded in OCT (optimal cutting temperature compound), and frozen in a mixture of isopentane and dry ice. Tissues were sectioned using a cryostat at 8µm and collected onto a glass slide and frozen. On each slide 4 representative sections were collected, representing the cervical, thoracic, lumbar and sacral sections of the spinal cord. For staining, slides were thawed at RT for 10-20min. Slides were rehydrated in 1xPBS-Tween 20 wash buffer for 10min, and excess wash buffer was drained. Next, the tissue was surrounded with a hydrophobic barrier using a barrier pen. Tissues were blocked for non-specific staining by incubating with blocking buffer (1% horse serum in PBS) for 30min at RT. Primary antibodies were diluted in incubation buffer and incubated overnight at 4°C. Slides were washed 3x for 15min each in wash buffer, and incubated with secondary antibody diluted in incubation buffer for 60min at RT, and washed again 3x for 15min. Tissues were mounted with cover glass using 0-0.5 0 no disease; 0.5 distal limp tail 1.0 Limp tail 2.0 Weakness in one or 2.5 in both hind limbs/slipping on bars 3.0  Paralysis in one or 3.5 both hind limbs 4.0 (3.0) plus weakness in one or 4.5 in both forelimbs 5.0 Moribund A B Days%post%immuniza.on%EAE%score%38  ProLong® Gold antifade reagent with DAPI (Life Technologies, Carlsbad, US) at 300µl. Appendix A.3 describes primary and secondary antibodies used.  2.12 Statistical analysis  All statistical analysis was performed using SigmaPlot (version 11.0, Systat Software, San Jose, CA) and graphs were prepared using GraphPad Prism (version 5, La Jolla, CA). In all cases, a Shapiro-Wilk’s normality test was performed. If the data was normally distributed, a repeated measures one-way ANOVA was performed, and if the data was not normally distributed, a repeated measures ANOVA on ranks (Friedman test) was performed, when comparing multiple treatments. Both tests were followed up with a Tukey multiple comparisons post test. All data are reported as mean +/- standard deviation unless otherwise indicated. 39  Chapter 3: The effect of excitatory compounds, inflammatory and apoptotic mediators on ARNT2 3.1 Characterization of primary cortical neuron-enriched cultures  Our first aim was to study the ability of stressors involved in MS pathogenesis to alter ARNT2 protein expression in primary cortical neuron-enriched cultures. To culture these cells we used the protocol as described in section 2.1 and performed a purity analysis. We achieved an average 92.6 ±6.9% pure neuronal culture (n=3, Figure 3.1 and 3.2). The remaining cells were GFAP+ (7.4±6.9%) (Figures 3.1 and 3.2). Healthy neurons, characterized by large cell bodies with complex extending axonal and dendritic processes, ranged in nuclear area between 50-200µm2. The astrocytes found in this culture exhibited a reactive morphology, with extensive ramifications in comparison to non-reactive astrocytes, which are flat and polygonal (Figure 3.1B) [125].   3.2 The majority of cells in primary cortical neuron-enriched cultures express mild to moderate levels of ARNT2  Using immunocytochemistry we began our cell population-based analysis by determining the distribution of ARNT2 intensity in our primary neuron-enriched cultures (Figure 3.2). Normalized ARNT2 intensity measurements are described in section 2.6.2, and represent mean pixel count divided by total pixel count in the nucleus to normalize the intensity of the ARNT2 signal for nuclear size. An isotype control was included for each antibody used, and the same program (Zen) was run to determine background normalized ARNT2 intensity levels. Any cell that fell within the range of the mean normalized ARNT2 intensity in the isotype control plus 40  two standard deviations above was considered negligible. Above this negligible range, cells were distributed into four groups depending on the normalized ARNT2 intensity (Figure 3.2), denoted as + (lowest), ++, +++, and ++++ (highest). Cells were assigned into the different groups based on the overall distribution of normalized ARNT2 intensity in the untreated fields, and how these values contributed to visually discernible differences in ARNT2 intensity to allow for categorization (Figure 3.2A). In addition, a cell line lacking in Arnt2 mRNA (bEND.3, a mouse brain endothelial cell line) as previously shown in our lab was used as a negative control. Furthermore, preliminary testing of knockdowns of Arnt2 with three individual constructs revealed a reduction in ARNT2 protein expression by 80-90% both via western blotting and immunocytochemistry, indicating antibody specificity for both of these techniques (Quandt lab, unpublished data).   With measurements in duplicate wells, it was found that 80-95% of all DAPI+ cells and, specifically MAP2+ cells, were positive for ARNT2. Furthermore, for the total DAPI+ population, the majority of cells were in the + to ++ intensity levels within our primary neuron- enriched cultures (encompassing roughly 65% of the population) while +++ encompassed 5-20% and 5-10% for ++++ (n=3, Figure 3.2C). Between 5-20% of cells fell into the negligible range as per our established restrictions (Figure 3.2). This total DAPI+ population is a representative of our total culture, which was used for downstream western blotting and qPCR applications. Overall, it was shown that ARNT2 expression follows a Gaussian-like distribution in primary neuron-enriched cultures, with most cells having low to medium expression under healthy conditions.   It was also found that the astrocytic population in this culture expressed ARNT2 (Figure 3.2B), and that none of the GFAP+ cells fell into the negligible range of detection (i.e. 100% of 41  GFAP+ cells were ARNT2+). This prompted us to also investigate ARNT2 expression in these cells across treatments, and also continue our work on ARNT2 expression in healthy astrocytes, which will be discussed in sections 3.9 and 3.10, respectively.   Figure 3.1: Characterization of primary cortical neuron-enriched cultures. A. Phase microscopy and B. Immunocytochemistry images of rat primary cortical neuron-enriched cultures at DIV 15, both at 10x magnification. A. Healthy adult neurons are characterized by large round cell bodies with extending complex processes. B. Analysis of purity by staining cultures with microtubule-associated protein 2 (MAP2; for neurons) and glial fibrillary acidic protein (GFAP; for astrocytes), revealed a 92.58±6.87% pure neuronal culture (based on three biological replicates). The remaining cells were GFAP-positive (7.42±6.87%).   DAPI%MAP2%GFAP%B A 42   Figure 3.2: The majority of cells in primary cortical neuron-enriched cultures express mild-moderate levels of ARNT2. A. Breakdown of culture purity as well as cell distribution based on normalized ARNT2 intensity in Experiment %neurons  (mean±stdev) %astrocytes (mean±stdev) Isotype control  (mean±stdev, mean pixel/total pixel count) Negligible cut-off (mean pixel/total pixel count) + ++ +++ ++++ 1 85.3±5.1 14.6±4.7 287.2±24.3 335.8 335.8-500   500-600   600-700   700+ 2 93.4±0.2 6.5±1.0 372.4±7.549 387.5 387.5-500 3 89.4±0.02 10.5±1.7 355.0±15.4 385.8 385.8-500 negligible + ++ +++++++0.00.10.20.30.40.50.91.0UnstimulatedArnt2 Intensity Rationegligible + ++ +++++++0.00.10.20.30.40.50.60.91.0UnstimulatedArnt2 Intensity Rationegligible + ++ +++++++0.00.10.20.30.40.50.91.0UnstimulatedArnt2 Intensity RatioExperiment*1*MAP2%GFAP%ARNT2%DAPI%A%B%C% Experiment*2* Experiment*3*MAP2%ARNT2%DAPI%MAP2%GFAP%ARNT2%DAPI%bEnd.3 Astrocyte Negligible + ++ +++ ++++ 43  primary cortical neuron-enriched cultures. B. Representative images of neurons categorized into different normalized ARNT2 intensity groups, as well as a representative astrocyte expressing ARNT2 and a negative control for antibody specificity (bEnd.3, a mouse brain endothelial cell line). C. Individual breakdown of the distribution of normalized ARNT2 intensity in three biological replicates revealed a Gaussian distribution.  3.3 4-aminopyridine and bicuculline have no effect on ARNT2 protein levels  The main binding partner of ARNT2 in the CNS, NPAS4, has been described as an early responder to excitatory stimulation, acting to dampen down the neuronal response, and maintain an excitatory/inhibitory balance through heterodimerization with ARNT2 [105] [113]. However, it has not been tested whether ARNT2 is differentially expressed under these types of stimuli. We tested ARNT2 protein levels with an excitatory dose of 4-aminopyridine, a selective inhibitor of the Kv1 type potassium channel (2.5mM), from 1-5h, to compare to NPAS4 data previously generated at these time points [127] (Quandt lab, unpublished data). There was no significant difference in ARNT2 protein levels when compared to untreated controls (Figure 3.3A), nor were there changes in viability observed in LDH release assays (data not shown) or via phase microscopy (Figure 3.3C).   Bicuculline, a GABAA receptor antagonist, was tested at 50mM for 1 to 5h, to match previous NPAS4 data (Quandt lab, unpublished data) [127]. Similarly to 4-AP, there was no significant change in ARNT2 protein levels or viability with this compound compared to untreated controls (Figure 3.3A/C, n=3, p=0.8691). Overall, there were no changes in ARNT2 with these excitatory compounds at the same doses and time points where one of its CNS binding partners, NPAS4, has been shown to significantly increase [127] (Quandt lab, unpublished data).   44   Figure 3.3: 4-aminopyridine and bicuculline have no effect on ARNT2 protein levels. (A) Points represent three biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots are shown in (B). Furthermore, there is no change in viability as observed via phase microscopy (C) and LDH release (data not shown). Repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  3.4 Potassium chloride has no effect on ARNT2 at low doses, and significantly reduces ARNT2 protein levels at excitatory/cytotoxic doses  Potassium chloride (KCl) has been described as a potential neuroprotective molecule at low doses [126]. For this reason, we tested KCl at 0.01-10mM for up to 8h, to examine the influence on ARNT2 [126]. Overall, we found that KCl did not induce changes ARNT2 protein levels from 1 to 8h exposure at these doses compared to untreated controls (Figure 3.4A).  ARNT2 Tubulin ARNT2 Tubulin unstimulated 4-AP, 5h Bic, 5h C A B 100µm#unstim 1 2 3 4 50.00.51.01.52.0Time (hours)ARNT2/Tubulin Ratio 4-APunstim 1 2 3 4 50.00.51.01.52.0Time (hours)ARNT2/Tubulin Ratio Bic45   KCl was also tested at an excitatory dose (50mM), at which it is known to induce potent Ca+2 influx in primary neuronal cultures [128]. This was followed from 1 to 8h, to compare to previously data showing significant NPAS4 upregulation with a peak at 3-4h exposure (Quandt lab, unpublished data).  We found that ARNT2 expression was significantly lower at 8h exposure compared to untreated control (Figure 3.5A, n=3, p≤0.05), but no significant changes in %cytotoxicity when followed for up to 24h with this dose compared to untreated control (Figure 3.5D). We also tested a cytotoxic dose of KCl (100mM), which showed significantly lower ARNT2 protein levels starting at 4h (Figure 3.5a, n=3, p≤0.01). We also found delayed significantly enhanced LDH release, along with cell swelling and death (18-24h exposure), reaching a maximum of 25% cytotoxicity when compared to untreated control (Figure 3.5D, n=2, p≤0.001) [128]. Overall, potassium chloride does not influence ARNT2 at doses previously shown to be neuroprotective, but does significantly reduce ARNT2 expression with both an excitatory and a cytotoxic dose of KCl, which precedes significant changes in cytotoxicity as measured via LDH release [126] [128]. This excitatory trend does not match NPAS4 expression patterns, indicating that these two proteins are differentially regulated by these compounds (Quandt lab, unpublished data) [113].  46   Figure 3.4: Potassium chloride has no effect on ARNT2 protein levels at doses below and including 10mM. (A) Points represent two biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B) and (C), respectively. Repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.    ARNT2 Tubulin ARNT2 A B Tubulin C unstimulated 100µm#0.01mM, 8h 0.1mM, 8h 1mM, 8h 10mM, 8h unstim0.01 0.1 1 10 0.01 0.1 1 10 0.01 0.1 1 100.00.51.01.52.0Dose of KCl (mM)ARNT2/Tubulin Ratio1h 2h 4hunstim  0.01 0.1 1 10 0.01 0.1 1 100.00.51.01.52.06h 8hDose of KCl (mM)ARNT2/Tubulin Ratio47   Figure 3.5: Potassium chloride significantly reduces ARNT2 protein levels at excitatory/cytotoxic doses. (A) Points represent three biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B). There is no change in %cytotoxicity as measured via LDH release at low doses, but significant increase with 100mM KCl at 18 to 24h (D). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  3.5 Early exposure to glutamate has no effect on ARNT2 protein levels, while prolonged exposure results in decreased levels of ARNT2  Glutamate was used to mimic glutamate excitotoxicity, one of the mediators of axonal damage/degeneration in MS. In primary neuronal cultures, glutamate exposure has been shown to induce acute necrosis in a subpopulation of cells, while remaining neurons die of delayed-onset apoptosis [130].  The influence of glutamate on primary neuron-enriched cultures was tested at 10-500µM up to 24h, to mimic the effect of various doses [130] [128]. Early exposure with this range of 100mM KCl, 4h unstimulated unstim0 6 12 18 24 0 6 12 18 24 0 6 12 18 24 0 6 12 18 24 0 6 12 18 2405101520253035409095100Exposure to KCl (hours)%Cytotoxicity10-2mM 1mM 10mM 50mM 100mMC A B D *** *** ARNT2 Tubulin 100µm#Dose of KCl (hours) %Cytotoxicity unstim 1 2 4 6 8 1 2 4 6 80.00.51.01.550mM 100mMDose of KCl (hours)ARNT2/Tubulin Ratio*******48  glutamate (i.e. up to 7h) had no influence on ARNT2 protein levels; only a consistent decrease with 500µM glutamate at 1, 3, 5, and 7h, though significance was never reached compared to untreated control (Figure 3.6A). Changes in cytotoxicity were observed via LDH release with doses of 50µM and higher at 7h (Figure 3.6A and D, respectively; n=4; p≤0.001 for 50µM, 200µM, and 500µM, p≤0.05 for 100µM). Phase microscopy revealed axonal/dendritic retraction by 1h and the presence of dead/dying cells by 6h exposure to doses of 50µM and above (Figure 3.6C; Figure 3.7C). ARNT2 significantly declined to 50-60% of untreated control at 12 and 24h with 500µM glutamate (Figure 3.7a, n=3, p≤0.05).	  As well, there was a significant increase in %cytotoxicity as measured via LDH release with doses as low as 10uM at 12h exposure, but not earlier. Overall, glutamate has no significant influence on ARNT2 protein levels when followed for up to 7h, and only influenced viability at higher doses at this time point. Also, there is less ARNT2 detected when observed at 12 and 24h with a high dose of glutamate, in concordance with higher changes in %cytotoxicity. At 12h and later significant cytotoxicity changes are also observed with lower doses compared to earlier time points (7h), indicating a potential temporal and dose-dependent response to glutamate.  3.6 Staurosporine significantly increases ARNT2 protein levels  Staurosporine potently induces apoptosis by preventing ATP binding to kinases, preventing protein kinase activity and resulting in growth factor withdrawal [137]. Reduced growth factor activity is linked to caspase-3 and cytochrome c activity, resulting in activation of the apoptotic pathway [138]. Therefore, this compound can be used as a model of apoptosis following blockage of trophic support [138].    49    Figure 3.6: Early exposure to glutamate has no effect on ARNT2 protein levels. (A) Points represent four biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B). Phase microscopy images are shown in (C). There is no change in %cytotoxicity as measured via LDH release until the 7h time point with 50µM-500µM glutamate (D). Repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  ARNT2 A T A T A T unstimulated 50µM, 3h 50µM, 5h 50µM, 7h C A B D 50µM, 1h 1hunstim 5 10 25 50 100 200 5000102030405090100Dose of  glutamate (µM)%Cytotoxicity3hunstim 5 10 25 50 100 200 5000102030405090100Dose of  glutamate (µM)%Cytotoxicity5hunstim 5 10 25 50 100 200 5000102030405090100Dose of  glutamate (µM)%Cytotoxicity7hunstim 5 10 25 50 100 200 5000102030405090100Dose of  glutamate (µM)%Cytotoxicity** ** ***100µm#Tubulin unstim 5 10 25 50 100 200 5000.00.51.01.52.02.5Dose of  glutamate (µM)ARNT2/Tubulin Ratio 1hunstim 5 10 25 50 100 200 5000.00.51.01.52.02.5Dose of  glutamate (µM)ARNT2/Tubulin Ratio 3hunstim 5 10 25 50 100 200 5000.00.51.01.52.02.5Dose of  glutamate (µM)ARNT2/Tubulin Ratio 5hunstim 5 10 25 50 100 200 5000.00.51.01.52.02.5Dose of  glutamate (µM)ARNT2/Tubulin Ratio 7h50   Figure 3.7: Prolonged glutamate exposure results in decreased levels of ARNT2. (A) Points represent three biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B). Phase microscopy images reveal axonal/dendritic retraction and the presence of dead/dying cells by 6h. There is a significant change in %cytotoxicity as measured via LDH release starting at 12h exposure (D). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.   We tested staurosporine at a range of doses from 5 to 1000nM for 24h, where previous studies showed neuronal apoptosis in vitro with 500-1000nM at this time point [137][138].  10nM staurosporine (mimicking a mild apoptotic stimulus) increased ARNT2 levels by 75% compared to unstimulated control at 24h exposure (Figure 3.8A, n=3, p≤0.05). At 500nM to unstimulated 50µM, 6h 50µM, 24h C A B D unstim0 6 12 18 24 0 6 12 18 24 0 6 12 18 24 0 6 12 18 24 0 6 12 18 240102030405090100Exposure to Glutamate (hours)%Cytotoxicity10uM 25uM 50uM 200uM 500uM* *** * * ** *** *** *** *** *** *** ARNT2 Tubulin 100µm#Dose of glutamate (hours) %Cytotoxicity unstim 6 12 24 6 12 24 6 12 24 6 12 240.00.51.01.52.02.510µM 50µM 200µM 500µM* *Dose of  glutamate (hours)ARNT2/Tubulin Ratio51  1000nM, cell death was visually evident, and was associated with ARNT2 levels comparable to untreated controls (Figure 3.8A, n=2/3). Furthermore, while there were no significant increases in %cytotoxicity (1.7% with 100nM, Figure 3.8D, n=2), the presence of dead nuclei was detected via phase microscopy (Figure 3.8C). This discrepancy may be due to the nature of the process of apoptosis versus necrosis [130]. Overall, it was shown that mild inhibition of growth factor activity via the application of staurosporine significantly enhanced ARNT2 expression, while apoptotic levels of staurosporine had no influence on ARNT2 expression despite the clear presence of dead nuclei.   3.7 Hydrogen peroxide significantly alters ARNT2 protein expression   We applied four doses: 25, 50, 100 and 300µM H2O2, to mimic mild to severe oxidative stress in primary cortical neuron-enriched cultures [93]. Representative phase microscopy images and cytotoxicity measurements showed significant changes in cytotoxicity starting at 12h with 50µM H2O2 compared to untreated control (Figure 3.9, two experiments separate from western blotting experiments).   25µM H2O2 induced a significant upregulation of ARNT2 with respect to untreated control from 0.5 to 3h exposure (Figure 3.10A, n=6, p≤0.01). Furthermore, phase microscopy revealed the presence of a healthy culture when followed for up to 24h, and no significant change in LDH release was detected for up to 24h (Figure 3.10C, LDH release data not shown).   52   Figure 3.8: Staurosporine significantly increases ARNT2 protein levels. (A) Points represent three biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B). Phase microscopy images reveal the presence of dead neurons with 500-1000nM. Furthermore, there is no significant change in %cytotoxicity as measured via LDH release (D). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.   Similarly to 25µM, 50µM H2O2 also significantly increased ARNT2 protein levels at 2h exposure (Figure 3.11A, n=6, p≤0.05). In contrast, there was a significant increase in ARNT2 unstim 5 10 25 50 100 200 500 10000.00.51.01.52.02.5Exposure to Staurosporine (24h, nM)ARNT2/Tubulin Ratiounstimulated 10nM 500nM 200nM * unstim 5 10 25 50 100 200 500 100002468109095100Exposure to Staurosporine (24h, nM)%CytotoxicityC A B D Tubulin 12h 24h 10nM 500nM 200nM 100µm###Dose of Staurosporine (nM, 24h) ##Dose of Staurosporine (nM, 24h) ##%Cytotoxicity # #unstim 5 10 25 50 100 200 500 10000.00.51.01.52.02.5Exposure to Staurosporine (24h, nM)ARNT2/Tubulin Ratio##ARNT2/Tubulin Ratio 53  %cytotoxicity as measured via LDH with 10h exposure to 50µM H2O2, with no changes in ARNT2 comparable to untreated controls (Figure 3.11D/A, n=3, p≤0.01).   Exposure to a higher dose of H2O2 (100µM) increased detectable ARNT2 protein at 0.5 and 1h compared to control (Figure 3.12A, n=7, p≤0.05). Morphological changes were not observed until 4h, and LDH release was not significantly upregulated until even later, by 9-10h exposure (Figure 3.12C/D, n=3, p≤0.01 and 0.001, respectively)  Initial analysis revealed that this dose of H2O2 also induces a downregulation of tubulin protein levels as cell viability and thus cytoskeletal integrity starts to be influenced. For this reason, a post-hoc analysis based on changes in cytotoxicity was performed, in which ARNT2 protein levels were normalized to tubulin only up until visually large-scale changes in cytotoxicity occurred (at 6h). Our western blotting observations generally indicated tubulin levels that remained stable after ARNT2 levels declined relative to tubulin. The highest dose of H2O2 tested was 300µM, which induces cell death at 3h exposure, as observed via phase microscopy and significant increases in %cytotoxicity starting at 3h (Figure 3.13C/D, n=2, p≤0.01 at 3h and 0.001 for all subsequent time points compared to unstimulated control). At this dose, there was also a significant downregulation of ARNT2 at 4h exposure compared to control (Figure 3.13A, n=6, p≤0.01; post-hoc analysis shown).  Overall, ARNT2 protein levels appear to follow two patterns when exposed to different levels of oxidative stress, with early up-regulation with 25, 50 and 100µM H2O2, and downregulation when the stimulus is cytotoxic (300µM H2O2). This early upregulation may be suggestive of an early protective response, while the later downregulation with toxic doses may be in concordance with changes in cell viability.  54   Figure 3.9: Representative phase microscopy images and cytotoxicity measurements showing progression of cell viability changes following H2O2 exposure. Phase microscopy images are shown in (A). Representative dead/dying neurons are shown with a black arrow. (B) LDH measurements reveal significant upregulation with 50µM H2O2 and higher. Points represent two biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.      unstim 0 6 12 18 24020406090100* **Time (hours)%CytotoxicityUnstimulated 12.5µM 25µM 50µM 100µM 300µM 100µm#unstim 0 6 12 18 24020406090100Time (hours)%Cytotoxicityunstim 0 6 12 18 24020406090100Time (hours)%Cytotoxicityunstim 0 6 12 18 24020406090100*******Time (hours)%Cytotoxicityunstim 0 6 12 18 24020406090100**********Time (hours)%CytotoxicityA B 6h 12h 18h 24h 55   Figure 3.10: Exposure to 25µM H2O2 significantly upregulates ARNT2 protein levels. (A) Points represent six biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B) and (C), respectively. There is no change in %cytotoxicity as measured via LDH release (data not shown). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.    ARNT2 Tubulin C A B unstimulated 5h 10h 100µm#unstim 0.5 1 2 3 4 5 6 7 8 9 100.00.51.01.52.02.5Time (hours)ARNT2/Tubulin Ratio** ******56   Figure 3.11: Exposure to 50µM H2O2 significantly upregulates ARNT2 protein levels. (A) Points represent six biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B) and (C), respectively. There is a significant change in %cytotoxicity as measured via LDH release at 10h exposure (D). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  ** ARNT2 Tubulin unstimulated 5h 10h 100µm#unstim0.5 1 2 3 4 5 6 7 8 9 100.00.51.01.52.02.53.03.5Time (hours)ARNT2/Tubulin Ratio *A B C D unstim 0.5 1 2 3 4 5 6 7 8 9 1005101520253090100Time (hours)%Cytotoxicity57    Figure 3.12: Exposure to 100µM H2O2 significantly upregulates ARNT2 protein levels. (A) Points represent seven biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B) and (C), respectively. There is a significant change in %cytotoxicity as measured via LDH release starting at 9h exposure (D). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  unstim 0.5 1 2 3 4 5 6 7 8 9 1005101520253090100Time (hours)%Cytotoxicity*** ARNT2 Tubulin unstimulated 5h 10h 100µm#Time (hours) unstim 0.5 1 2 3 4 5 6 7 A B C D ** unstim 0.5 1 2 3 4 5 60.00.51.01.52.02.5Time (hours)ARNT2/Tubulin Ratio **58   Figure 3.13: Exposure to 300µM H2O2 significantly downregulates ARNT2 protein levels. (A) Points represent six biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. Representative blots and images are shown in (B) and (C), respectively. There is a significant change in %cytotoxicity as measured via LDH release starting at 3h exposure (D). p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  3.8 Oxidative stress fails to drive Arnt2 mRNA levels  As part of aim 1, we aimed to examine ARNT2 not only at the protein level, but also at the mRNA level. Using 25, 50 and 100µM H2O2 we observed an early increase in ARNT2 protein levels (Figures 3.10/11/12A). To determine a potential reason for this increase we decided to measure Arnt2 mRNA levels in response to oxidative stress using qPCR. We focused on 25 and 100µM. RNA and protein were isolated from the same lysate for direct comparison as described in section 2.5. Overall, we found that oxidative stress failed to drive early changes in unstim 0.5 1 2 3 4 5 6 7 805101520253090100Time (hours)%CytotoxicityTime (hours) unstim 0.5 1 2 3 4 5 6 ** *** *** *** *** *** ARNT2 Tubulin C unstimulated 5h 10h 100µm#unstim 0.5 1 2 3 40.00.51.01.52.0**Time (hours)ARNT2/Tubulin RatioA B D 59  Arnt2 mRNA levels at 25 or 100µM H2O2, and reduced Arnt2 mRNA levels with 100µM H2O2 from 5-7h exposure compared to untreated control. This was in concordance with a reduction in cell viability as previously determined via LDH release/phase microscopy (Figure 3.14, n=3, 25µM p=0.1636, 100µM p≤0.05 for all significant time points; Figure 3.9). The western blot results, though underpowered, showed the same patterns as our previous western blot results with these doses and time points of H2O2, confirming a divergence between Arnt2 mRNA and protein regulation with H2O2 (Appendix B.2, n=3). Overall, Arnt2 mRNA was not differentially expressed in response to mild to moderate oxidative stress.   Figure 3.14: Oxidative stress fails to drive Arnt2 mRNA levels. Points represent three biological replicates, with two technical replicates each, where lines and whiskers indicate mean and standard deviation. p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  unstim 0.5 1 2 3 4 5 6 70.00.51.01.5Time (hours)Arnt2/B-actin Ratio * * * 25µM H2O2 100µM H2O2 unstim 0.5 1 2 3 4 5 6 7 8 9 100.00.51.01.5Time (hours)Arnt2/B-actin Ratiounstim 0.5 1 2 3 4 5 6 7 8 9 100.00.51.01.5Time (hours)normalized b-actinunstim 0.5 1 2 3 4 5 6 70.00.51.01.5Time (hours)normalized b-actinA B 60  3.9 Oxidative stress significantly alters ARNT2 protein levels in both neuronal and astrocytic populations  For subaim 1c we set out to examine ARNT2 expression in specific cell populations in primary cortical neuron-enriched cultures. We tested the same doses of H2O2 (25µM and 100µM) previously used to investigate changes in ARNT2 expression via western blotting and qPCR, focusing in on earlier time points (0.5, 1, 2, 4 and 6h), to capture the range of early upregulation and later downregulation observed in our global western blotting experiments and to compare this to a more sensitive cell population-based analysis. This analysis was used to discern ARNT2 expression in both neurons and astrocytes separately. We had three separate sets of data: total DAPI+ cells, MAP2+ cells and GFAP+ cells, as described in section 3.2. In one out of three replicates, 25µM H2O2 significantly increased ARNT2 protein levels at 0.5h in all three categories, while in the other two biological replicates, there was no significant upregulation with this dose (data not shown). Therefore, overall we concluded that this dose of H2O2 does not significantly upregulate ARNT2, which is discordant with our western blotting results where a significant upregulation was observed with this dose (Figure 3.10A). With 100µM H2O2, there was a significant increase in ARNT2 protein levels in total DAPI+, MAP2+, and GFAP+ cells at 1h exposure compared to unstimulated control (Figure 3.15B, n=3, p≤0.05 for DAPI+, 0.001 for MAP2+ and 0.01 for GFAP+ cells). The upregulation of ARNT2 at this time point with this dose of H2O2 matches the western blotting results (Figure 3.12). A reduction in ARNT2 protein levels was observed at 4 to 6h in both neuronal and astrocytic populations (i.e. all three data groups, n=3 for each, at 2h: p≤0.01 for GFAP+, at 4h: p≤0.05 for DAPI+, p≤0.01 for GFAP+, at 6h: p≤0.001 for all three) in when visual changes in cell viability were previously observed (Figure 3.9). Therefore, through a sensitive cell-population based analysis, we found a significant early 61  upregulation of ARNT2 with 100µM H2O2 as well as a significant reduction in ARNT2 in both neurons and astrocytes, allowing us to distinguish ARNT2 expression in these two cell populations. Since the astrocytes in our primary cortical neuron-enriched cultures were morphologically reactive, this study prompted us to examine ARNT2 expression in enriched astrocyte cultures under healthy conditions.   Figure 3.15: Oxidative stress significantly alters ARNT2 protein levels in both neuronal and astrocytic populations. A. Representative images of unstimulated cells and cells exposed to 100µM H2O2 at 1h and 6h show death at 6h. B. Breakdown of normalized ARNT2 intensity changes across treatments with 25µM and 100µM H2O2, further categorized into all DAPI+, MAP2+ and GFAP+ cells. Points represent three biological replicates, with eight technical replicates each, where lines and whiskers indicate mean and standard deviation. p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated.  Total DAPI+ cells MAP2+ cells GFAP+ cells Unstimulated MAP2 GFAP ARNT2 DAPI 100uM H2O2, 1h 100uM H2O2, 6h 25µM H2O2 100µM H2O2 unstim 0.5 1 2 4 60300400450500550600Time (hours)ARNT2 Intensityunstim 0.5 1 2 4 60300400450500550600Time (hours)ARNT2 Intensityunstim 0.5 1 2 4 60300400450500550600Time (hours)ARNT2 Intensityunstim 0.5 1 2 4 60300400450500550600Time (hours)ARNT2 Intensityunstim 0.5 1 2 4 60300400500600700800Time (hours)ARNT2 Intensityunstim 0.5 1 2 4 60300400500600700800Time (hours)ARNT2 Intensity** *** * *** *** * *** *** ** A B 62  3.10 Characterization of primary astrocyte-enriched cultures  As our primary cortical neuron-enriched cultures consistently expressed a small population of astrocytes, we decided to examine ARNT2 expression in healthy, astrocyte-enriched cultures compared to reactive astrocytes observed in the neuronally-enriched cultures. In vitro, healthy, non-reactive astrocytes are morphologically flat and polygonal [125]. In two biological replicates, these cultures were 95.4±3.6% GFAP-positive astrocytes, with the remaining cells being MAP2-positive neurons (4.6±3.6%) (Figure 3.16). Furthermore, these cultures were stained for ARNT2 to visually observe the distribution in astrocytes. Overall we found that GFAP+ cells showed variable ARNT2 staining, which requires further investigation (Figure 3.16). Compared to the isotype control, we found that between 80-85% of GFAP+ cells were positive for ARNT2 (i.e. 15-20% were negligible) (n=2, 10 regions of an untreated well examined, in duplicate).   Our lab has started using oxidative stress amongst other compounds previously tested in primary cortical neuron-enriched cultures, to observe the effects of these mediators on astrocytic ARNT2 expression. Currently these results are inconclusive, but will be continued via western blotting and qPCR.     63   Figure 3.16: Characterization of primary astrocyte-enriched cultures.	  Analysis of purity by staining cultures with microtubule-associated protein 2 (MAP2; for neurons) and glial fibrillary acidic protein (GFAP; for astrocytes), revealed a 95.43±3.57% pure astrocytic culture (based on two biological replicates). An inset of the isotype control is included for GFAP (centre) and ARNT2 (right). The remaining cells were MAP2-positive (4.57±3.57%). Protocol adapted from: Aiga M,Levinson JN, Bamji SX, J Biol Chem (2011), and Kaech S, Banker G, Nature (2006).    GFAP MAP2 ARNT2 DAPI ARNT2 64  Chapter 4: Optimization of viability/cytotoxicity assays 4.1 Optimizing neuronal treatment protocols to improve sensitivity of LDH detection  Based on our findings from chapter 3, measuring cell death using the lactate dehydrogenase (LDH) assay showed a delay in significant changes despite apparent cell death in our cultures, with all cytotoxic compounds used. To determine the suitability of the LDH assay for our primary neuron-enriched cultures, we performed a two-fold sensitivity analysis. These optimization tests allowed us to discern the assay sensitivity. For this test, two 96-well plates were seeded with primary neuron-enriched cultures at 50,000 cells/cm2, the same density used for western blotting and qPCR experiments. Cells grew in 150µl neurobasal media as described in section 2.1. For the assay, one plate was adjusted to 100µl, and the second to 70µl. For both plates we tested a range of H2O2 from 12.5µM to 300µM up to 24h, which were all added in 20µl PBS in duplicate. The first plate sat in a final volume of 100µl, and the second in 70µl. At the end of the experiment the supernatants were collected and examined for LDH release as described in section 2.7.   There was a significant increase in %cytotoxicity compared to untreated control with increasing doses of H2O2, starting as early as 6h with as low a dose as 50µM. Furthermore, with this assay modification LDH release matched visual inspection of cells, reaching a maximum of 45% in the 70µl plate and 51% in the 100µl plate with 300µM H2O2 at 24h (Figure 4.1, n=1, overall p≤0.0001 for both 70 and 100µl experiments). Therefore, modifications to the LDH assay may allow for more sensitive cytotoxicity measurements in primary cortical neuron-enriched cultures than our previous approach of removing supernatants directly from wells for western blotting experiments as described in section 2.4. This could be due to a number of reasons, including a higher number of cells per ml in the 96 well plate experiments compared to the 65  western blotting experiments, allowing for concentrating of LDH in the supernatant, and a larger volume of compound added. Furthermore, this test also indicates the suitability of this assay for future applications such as testing potential viability changes in response to knockdown of Arnt2.    Figure 4.1: Optimizing neuronal treatment protocols to improve sensitivity of LDH detection. Treatments were added and supernatants were collected from a 96-well plate at DIV 16. Bars and whiskers represent mean and standard deviation of 1 biological replicate, with two technical replicates. p≤0.05*, p≤0.01**, p≤0.001***, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to unstimulated of each time point.  A B Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 0102030405060901000h 6h 12h 18h 24h70µl********** *******************************Dose of H2O2 (µM)%Cytotoxicity100µlUnstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 Unstim 12.5 25 50 100 300 0102030405060901000h 6h 12h 18h 24h******************* ************************Dose of H2O2 (µM)%Cytotoxicity66  4.2 The Live/Dead® viability/cytotoxicity assay is sensitive in distinguishing live and dead populations in primary cortical neuron-enriched cultures  Suitability of the Live/Dead® viability/cytotoxicity assay was tested using 25, 50 and 100µM H2O2 for up to 24h to test assay sensitivity to a range of mild to more severe oxidative stress. We first tested calcein and EthD-1 at 2µM each. We found that while changes in EthD-1 could be detected as early as 6h with 100µM H2O2, calcein levels remained relatively stable despite visual loss in viable cells, indicating that this dose of calcein was potentially too high for sensitive detection of differences in viability across time points (Appendix D.1).   Therefore, in our next experiment we tested calcein and EthD-1 at 1 and 2µM, respectively, as was also recommended in the protocol. There also was a significant gradual reduction in calcein absorbance, indicating a loss in neuronal function to convert to calcein, starting as early as 6h exposure with 50 and 100µM H2O2, as well as a significant gradual increase in EthD-1 absorbance starting at 8h with 100µM H2O2 (Figure 4.2, n=1). Overall, we found that at these doses, this assay is sensitive in distinguishing changes in cell health and death in primary cortical neuron-enriched cultures, potentially giving us a clearer understanding of the relationship between viability changes in response to oxidative stress and expression patterns of ARNT2. Furthermore, this assay could potentially be suitable for future applications such as testing probable small-scale viability changes in response to knockdown of Arnt2. 67   Figure 4.2: The Live/Dead® Viability/Cytotoxicity Assay is sensitive in distinguishing live and dead populations in primary cortical neuron-enriched cultures. 1uM calcein and 2uM ethidium homodimer-1 (EthD-1) were used. Bars and whiskers represent mean and standard deviation of 3 technical replicates of 1 biological replicate. Calcein is plotted in green on the left y-axis, and EthD-1 is plotted in red on the right y-axis. p≤0.05*/+, p≤0.01**/++, p≤0.001***/+++, repeated measures ANOVA on ranks with Tukey Multiple Comparisons Test to untreated, (* - Calcein, + - EthD-1).010020030040001020304050Calcein EthD-1Time (hours)25µM H2O2Unstim Dead 2 4 6 8 12 18 24************++++++++++++010020030040001020304050CalceinEthD-1Time (hours)50µM H2O2Unstim Dead 2 4 6 8 12 18 24***+++* ********* ***++++++++010020030040001020304050CalceinEthD-1Time (hours)100µM H2O2Unstim Dead 2 4 6 8 12 18 24***+++** ************++++++++ +++68  Chapter 5: In vivo results 5.1 EAE results  In our hands the MOG35-55 EAE model of disease is a reproducible and consistent model of chronic progressive disease. Clinical disease presents consistently in at least 90% of mice with limited variability in the severity of disease. Mice exhibit an acute period of clinical disability and display some degree of recovery. Over time, clinical disability does again worsen, and confirms the model in our hands as that of a chronic and also progressive model of clinical disability. As described in section 1.2.3.6, our preliminary EAE study revealed a significant upregulation of Arnt2 in the brain pre-onset, and a significant downregulation in the spinal cord at peak disease (Quandt lab, unpublished data). Furthermore, at peak disease, ARNT2 expression was observed in regions of inflammatory infiltrates, potentially including immune cells and other glial cells. We set out to repeat this study with higher power, to localize ARNT2 expression throughout initiation and progression phases of disease in progressive EAE. In EAE, the majority of pathological hallmarks are found in the spinal cord, with lesser amounts detected in the brain [52]. Therefore, it appears that the reduction in Arnt2 co-localizes most with the disease stage/region of maximal disease. In the brain, the increase pre-onset may represent a response to a mild stimulus. If so, our in vitro data may potentially mimic this stage of the disease in the early upregulation of ARNT2 at the protein level. To better draw correlations between ARNT2 levels throughout onset, peak, and recovery, this experiment has been repeated with a higher power and additional time points covering healthy littermates and sham-immunized controls. Scoring of clinical disease revealed the typical chronic progressive EAE process (Figure 5.1).   69   Figure 5.1: EAE Results. (A) Points represent mean of 8 individual animals that were followed for the whole study, with standard error (whiskers). (B) Points represent average clinical score in the study on that day (i.e. all animals that were still present in the study, starting with n=65 at day 0, and n=8 at day 45), with mean and standard error  (points and whiskers).   5.2 Preliminary localization of ARNT2 protein in healthy tissues versus tissues undergoing EAE  In a healthy spinal cord, ARNT2 expression is largely limited to the neuronal cell bodies of the tracts and cells of the grey matter with some scattered expression localized to MAP2+ cells in the white matter (Figure 5.2). GFAP expression, indicative of astrocytes, is generally low, mostly found along the meninges of the spinal cord, along the midline to the central canal, and scattered occasionally in the white matter (Figure 5.2). Of these GFAP+ cells, roughly 25-30% are ARNT2+ (Figure 5.2). In contrast, the grey matter of the spinal cord is largely devoid of 7 10 14.5 18 25 32 45012345DayEAE Scoren=65 n=8 A B 5 10 15 20 25 30 35 40 45012345DayEAE Score70  GFAP expressing cells (Figure 5.2). In the spinal cord of EAE mice examined at peak clinical disease, we observed an increase in GFAP+ staining throughout the white matter, as well as the notable appearance of GFAP+ cells throughout the grey matter (Figure 5.2). Notably, GFAP increases in the white and grey matter were observed in sections where inflammatory infiltrates were observed, thus associated with lesion activity (Figure 5.2). In these regions of inflamed/lesioned cord, ARNT2 intensity was also greatly increased (Figure 5.2). Preliminary localization experiments show that ARNT2 most commonly localized within neurons in healthy tissues, but was notably expressed in the majority of GFAP+ cells in lesioned areas, including both regions of white and grey matter (Figure 5.2). The same increases in GFAP were observed in the much later, chronic stages of the disease as well; again, GFAP expression on astrocytes in both white and grey matter regions was typically localized to cells that exhibited intense ARNT2 staining.       71   Figure 5.2: Preliminary localization of ARNT2 in healthy tissues versus tissues undergoing EAE. Immunocytochemistry of moue spinal cord in healthy, acute EAE (day 20) and chronic EAE (day 90). White arrows represent ARNT2+ astrocytes. Lower inserts represent white matter (WM, first panel) and grey matter (GM, second panel) per disease stage, all taken from the top right corner of the whole cord image. Sections represent upper thoracic region of the cord. Images represent a representative of three biological replicates. Age of mice at examination was 24 weeks for healthy and chronic EAE, and 9 weeks for acute EAE.   Healthy Acute EAE Chronic EAE ARNT2 GFAP MAP2 DAPI WM# GM# WM# GM# WM# GM#72  Chapter 6: Discussion 6.1 Summary and significance  Previous work on ARNT2 has elucidated its expression patterns and functional roles in vivo throughout development and under conditions of hypoxia, ischemic injury and neuroprotection, as well as in in vitro models, including primary cells and cell lines. Here we present, for the first time, a description of the effect of various CNS stressors involved in excitatory and neurodegenerative processes on ARNT2 expression in primary cortical neuron-enriched cultures. Specifically, this work has modeled cortical neurons which, in MS, exhibit more neuronal damage/loss in the progressive stage of the disease when compared to those found in white matter, contributing to cognitive deficits observed in MS [32] [33].  In this study we set out to examine the ability of stressors involved in the pathogenesis of MS to alter ARNT2 protein expression in primary cortical neuron-enriched cultures, at the protein and mRNA level, and within different cell populations. We have shown that ARNT2 expression is upregulated under certain stress-initiating stimuli, while declining ARNT2 expression is associated with loss in cell integrity/viability. Furthermore, Arnt2 mRNA is not influenced by oxidative stress in primary cortical neuron-enriched cultures, indicating no change in transcript levels. Cell population-based analysis revealed a homogeneous distribution of ARNT2 protein nuclear intensity, and the presence of strong ARNT2 staining in astrocytes, which are morphologically reactive in these cultures.   We also tested the suitability and optimization of several available viability assays in these cultures as a set up for future functional studies, and found that experimental procedure modifications allowed for more sensitive detection. Finally, we performed an EAE experiment to 73  localize and compare ARNT2 expression throughout various disease stages. Preliminary in vivo results suggest elevated ARNT2 intensity in neurons and astrocytes at peak EAE disease.   The upregulation of ARNT2 protein under conditions of oxidative stress and apoptotic stimuli is a novel finding, and is also suggested in preliminary analysis of EAE at chronic disease stages. With a higher dose of H2O2 a bimodal distribution (significant increase and decrease) of ARNT2 was observed in neurons, with decreases occurring in concordance with a loss in cell health. This adds to the current understanding of ARNT2 regulation in response to stimuli involved in models of neurodegeneration both in vitro and in vivo, indicating an indirect association of ARNT2 with neuronal health.    6.2 The influence of excitatory compounds on ARNT2  Our first investigation was to observe the effects of excitatory compounds on ARNT2, to see if this protein is regulated similarly to NPAS4 in response to these stimuli. Past studies have mostly focused specifically on NPAS4 regulation in vitro and in vivo, but few have looked at how stimuli change ARNT2 levels [114] [118]. Shamloo et al. (2006) found a significant upregulation in Npas4 mRNA in a rat model of ischemic injury [114]. Guidotti et al. (2012) showed a reduction of both Npas4 and Arnt2 mRNA in a rat model of depression [118]. The lack of ARNT2 alteration previously observed under excitatory stimuli matches our observations on 4-AP and Bic [112] (Quandt lab, unpublished data). KCl stimulation, on the other hand, leads to ARNT2 downregulation when NPAS4 levels are still enhanced, which is a novel finding (Quandt lab, unpublished data). This discrepancy may potentially be linked to differential regulation of these binding partners under excitatory processes. While ARNT2 is constitutively expressed in the nucleus, NPAS4, an immediate early gene, is not expressed under baseline conditions and 74  rapidly increases under periods of Ca+2 influx [118]. The importance of NPAS4:ARNT2 dimerization in the transcription of BDNF has been previously reported [89] [118]. This novel description indicates that while ARNT2 and NPAS4 may still be functioning through heterodimerization, following exposure to excitatory compounds for longer periods of time (KCl) shows differential regulation of these proteins. This could potentially result from prolonged exposure to these stimuli or the severity of the stimulus previously untested, or the effect of gradual loss of cell viability on ARNT2 versus NPAS4. Other bHLH factors implicated in maintaining an excitatory-inhibitory balance in the CNS include pancreatic transcription factor 1a (PTF1A) and atonal homolog 1 (ATOH1), which regulate production of inhibitory/excitatory neurons in the cochlear nucleus, and achaete-scute homolog 1 (ASCL1) and PTF1A regulation of GABAergic versus glutamatergic neurons in the dorsal spinal cord, amongst others [139] [140].  6.3 ARNT2 at the mRNA versus protein level  Here we have shown that while ARNT2 at the protein level is significantly upregulated by 25µM (from 0.5-3h), 50µM (at 2h) and 100µM (from 0.5-1h) H2O2, there is no change at the mRNA level in Arnt2 at these doses/time points. This discrepancy indicates that ARNT2 protein upregulation at these doses/time points is not due to enhanced mRNA expression resulting in enhanced translation.   Increased translation of ARNT2 protein by constitutively expressed Arnt2 mRNA might explain this phenomenon. Additionally, ARNT2, like other bHLH transcription factors, is known to regulate its own transcription via a positive feedback loop, thereby ensuring its own constitutive expression [95]. Therefore, one potential mechanism behind this result could be that ARNT2 protein upregulation occurs as a result of translation of constitutively expressed Arnt2 75  mRNA, while ARNT2 protein maintains these mRNA levels. ARNT2 transcriptional activity is also negatively regulated via activity of the necdin protein or an ARNT2 splice variant as described in section 1.2.3, and this could result in a lack of upregulation of Arnt2 mRNA, too [95].  mRNA levels of Arnt2 could also be regulated by microRNA, which has never been described in the regulation of Arnt2, but has been investigated in other bHLH transcription factors, including Hes1 (hairy and enhancer of split-1), which mediates differentiation of neural progenitor cells to astrocytes, the mRNA expression of which is destabilized by microRNA-9 [141]. Furthermore, epigenetic regulation of Arnt2 has previously been described in the context of neuronal differentiation [94]. In this study, P19 cells, which are pluripotent embryonic-derived teratocarcinoma cells in mice and can be treated to differentiate into neuronal cells through retinoic acid application [94]. It was found that the Arnt2 promoter was non-methylated in undifferentiated P19 cells, and contained both a common repressive and a common active marker (H3K27me3 and H3K4me3, marking inactive versus active genes, respectively) [94]. In undifferentiated cells, the Arnt2 promoter contained both markers (termed a “bivalent” marker) [94]. Upon differentiation to neuronal cells, there was a decrease in H3K27me3 at the Arnt2 promoter, which correlated with an increase in ARNT2 protein expression [94]. Thus, in these neuronal cells, ARNT2 protein expression was dependent on the downregulation of gene repression [94]. It would be interesting to study this pattern of epigenetic modulation of the Arnt2 promoter in primary cortical neuron-enriched cultures, and determine whether the lack of Arnt2 mRNA upregulation is due to epigenetic repression of the promoter.  76  6.4 Global versus individual cell-based analysis of ARNT2 expression  Observing overall ARNT2 protein levels through western blotting revealed a significant upregulation under apoptotic conditions and oxidative stress. As oxidative stress is an important mediator of axonal degeneration/neuronal loss through its influence on mitochondrial dysfunction, we decided to continue our investigation of this process using a more sensitive approach [40]. Immunocytochemistry is a sensitive method of detection, and we postulated that it should allow us to observe both constitutive ARNT2 expression and changes in ARNT2 expression in response to oxidative stress at the cellular level. Furthermore, once it became clear that our primary cortical neuron-enriched cultures contain 10.5±4.1% astrocytes (n=3), studying ARNT2 expression at the individual cell-basis became relevant in the context of examining astrocytic ARNT2 expression as well as distinguishing if it is the neuronal, astrocytic, or both populations contributing to the significant upregulation in ARNT2 protein levels via western blotting.   Overall, we found that using both the western blotting and immunocytochemistry, there was a significant upregulation in ARNT2 with 100µM H2O2, but the significant upregulation of ARNT2 with 25µM H2O2 was observed via western blotting was only seen in one of three biological replicates using immunocytochemistry. This discrepancy could be due to different reasons; firstly, the requirement for more power in the immunocytochemistry data. Since the changes in ARNT2 observed are relatively small, we required an n of six for western blotting experiments to rule out a false negative result, but the immunocytochemistry approach was replicated three times. Secondly, cells were seeded at a lower density for immunocytochemistry experiments (35,000 versus 100,000 cells/well for western blotting), and shifts in ARNT2 intensity did not correspond with large-scale changes in the numerical values using this type of 77  analysis. Lastly, for immunocytochemistry 8 regions were examined per well, encompassing about 75% of the total well, whereas quantitative densitometry for western blotting results uses a sum of the ARNT2 intensity in the whole cell lysate loaded, which is mixed prior to loading and thus may represent the whole well.   Using immunocytochemistry we observed a significant downregulation in ARNT2 with 100µM H2O2 in all three experiments. We were unable to achieve this statistically using western blotting as ARNT2 and our loading control, tubulin, were influenced by the loss of cell viability starting 6h of exposure, at which point a majority of cells were dead, losing the biological relevance of our protein quantification.  The positivity of astrocytes for ARNT2 in vitro mimics suggestions in our in vivo observations and will be discussed in the next section.  6.5 ARNT2 expression in astrocytes  As mentioned previously, cortical astrocytic ARNT2 expression has only been described once, in an in vitro model of hypoxia in astrocyte cultures [107]. Wurdak et al. (2010) found that overexpression of Arnt2 in adult hippocampal neural progenitor cells resulted in enhanced neuronal and suppressed astrocytic differentiation [142]. Dela Cruz et al. (2014) found that under normal physiological conditions, GFAP+ cells within the mouse substantia nigra did not express ARNT2 protein [143]. Neuronal restriction of Arnt2 mRNA was also shown in rat brains by Drutel et al. (1999) [93]. We described here, for the first time to our knowledge, ARNT2 expression in both morphologically reactive and non-reactive astrocytes in primary cultures. In our preliminary EAE data, we found a suggestion of an increase in ARNT2+ astrocytic populations at peak disease, and in regions which were not seen in healthy tissue, and need to be 78  investigated further. These astrocytes morphologically resemble reactive astrocytes found in vivo in EAE as well as MS post-mortem tissue, with increased ramifications and high GFAP expression, which play a role in the disease process through astrogliosis, or expansion of astrocyte numbers due to nearby damage, causing scar formation and potential inhibition of axonal regeneration [40]. The expression of ARNT2 in astrocytes in vivo is a novel finding, and could be explained due to differences in regions observed, or previous examination in primarily healthy tissues.  In our primary cortical neuron-enriched cultures, astrocytes also had a reactive phenotype, likely due to a lack of media requirements for their growth in the culture [125]. Furthermore, astrocytic ARNT2 expression was significantly upregulated with 100µM H2O2 at 1h, followed by a significant downregulation, like neuronal populations in this culture. The upregulation of ARNT2 in astrocytes could be due to the direct influence of oxidative stress, or as a bystander result of H2O2 effect on neurons, which could in turn influence astrocytic function and ARNT2 expression [144]. This indicates that the patterns we saw with our global analysis of ARNT2 expression via western blotting result as a combination of astrocytic and neuronal changes in ARNT2, though considerably more highly from the more abundant neuronal population.  Furthermore, it was found that while the MAP2+ population had 5-20% ARNT2 negligible cells, the GFAP+ population in our primary cortical neuron-enriched cultures had no cells that were negligible for ARNT2. Comparison of these reactive astrocytes to our primary cortical astrocyte-enriched cultures indicates a potential higher proportion of ARNT2 positivity within the reactive astrocyte population, as 15-20% of cells within the astrocyte enriched cultures were negligible for ARNT2. Therefore, a shift in elevated ARNT2 expression may be suggestive 79  of an activation of astrocytes to their reactive state. This matches preliminary in vivo data, where reactive astrocytes at peak EAE disease are more apparent with increased GFAP positivity throughout the lesioned areas, which typically associated with ARNT2 expression. Whether or not ARNT2 levels were greater on a per cell basis in astrocytes requires a more quantitative analysis and comparison to ARNT2 expression in astrocytes in healthy tissue, in addition to further in vitro and in vivo investigations being required.    It is known that astrocytes express BDNF, and that this expression is upregulated under conditions of CNS injury [88]. NPAS4 is thought to be a neuron-specific transcription factor, and preliminary results from our laboratory show no NPAS4 expression in astrocytes under baseline or excitatory conditions [118] (Quandt lab, unpublished data). Therefore, ARNT2 may heterodimerize preferentially with a different binding partner in astrocytes. Furthermore, the Bdnf gene has nine promoters, and ARNT2:NPAS4 only act at two of these in neurons [123]. Thus, it is possible that BDNF regulation is not mediated via ARNT2:NPAS4 in astrocytes. For example, endothelin has been shown to increase Bdnf mRNA isoforms in astrocytes (in vitro and in vivo) [145].   While the function of ARNT has been studied in astrocytes in the context of hypoxia, work on astrocytic ARNT2 is lacking [146] [147]. BBB integrity is regulated by astrocytic induction of the hypoxic HIF-1 pathway, in which ARNT2 plays a binding partner role as described in section 1.2.3, switching to the production of factors to promote vessel permeability in an in vitro model of MS [148]. Astrocytic VEGF expression and subsequent angiogenesis is dependent on the HIF-1 pathway in hypoxic astrocytes, too [107] [149]. Activation of ARNT and HIF-1α in astrocytes has been shown to protect astrocytes against oxidative damage and glutamate excitotoxicity [147]. As ARNT2 can often compensate for the role of ARNT, 80  particularly in the CNS, it is possible that ARNT2:HIF-1α heterodimerization also occurs in astrocytes to protect against oxidative stress, though this needs to be further investigated [95].   6.6 ARNT2 expression and changes in cell viability   ARNT2 has previously been implicated in maintenance of cell viability of PC12 cells and in ischemic injury models [93]. In PC12 cells, downregulation of Arnt2 via an antisense oligonucleotide resulted in apoptosis of cells [93]. In a rat model of focal cerebral ischemia, Arnt2 gene transcript levels significantly decreased following 2 hours of recirculation, preceding neuronal death at 24 hours of recirculation [93]. In the chronic progressive model of EAE, we found a significant downregulation of Arnt2 mRNA at peak disease in the spinal cord (Quandt lab, unpublished data). Alterations in ARNT2 expression patterns and loss of cell viability appear to also follow a temporal gradient in our study of the effects of various compounds on neuronal health/viability.   Treatment of neuronally-enriched cultures with KCl significantly downregulated ARNT2 protein levels, which preceded large-scale changes in neuronal viability observed via phase microscopy and LDH release. With glutamate, we did not observe significant changes in ARNT2 protein levels unless cells were treated with a high dose (500µM) for at least 12h, while changes in viability were observed with 10µM glutamate and above at this time point. Staurosporine, which was used to mimic loss of growth factor activity and apoptosis, showed no change in ARNT2 protein levels compared to untreated control at doses which induce apoptosis in our culture, indicating that ARNT2 may be differentially regulated in response to necrotic/apoptotic stimuli and resulting downstream pathways. Cell death via apoptosis is programmed, unlike necrosis, which is spontaneous in nature [130]. The process of apoptosis, unlike necrosis, 81  exhibits maintenance of membrane integrity, and thus likely minimal LDH release compared to necrosis [130]. Another more appropriate measure for apoptosis-mediated cell death is the TUNEL (terminal deoxynucleotidyl transferase dUTP nick end labeling) assay [93]. Furthermore, the effect of growth factor inhibition via the use of staurosporine has also been tested previously in the context of another bHLH factor complex, HIF-1, made up of HIF-1α and ARNT [150]. It was shown that induction of HIF-1 induction reduced staurosporine-induced cell death in A549 cells (lung carcinoma) [150]. Whether induction of ARNT2, which can form the HIF-1 complex in the CNS, can also protect cells from apoptosis warrants investigation. Furthermore, pre-treatment of mouse hippocampal neurons with Bic reduced the percentage of cells undergoing apoptosis following staurosporine treatment, which was abolished with knockdown of NPAS4 [151]. As the primary binding partner of NPAS4 in the CNS is ARNT2, and its previous implication as an anti-apoptotic factor in PC12 cells, their heterodimerization may play a role in protection of cells against apoptotic cell death [89] [93].  Finally, we saw significant ARNT2 reduction with 100µM H2O2 (via immunocytochemistry) and with 300µM H2O2 (via western blotting), which occurred in concordance with cell viability changes (LDH release/phase microscopy). We tested the influence of H2O2 on viability following an optimization of our procedure in obtaining supernatants for LDH release measurements, as well as using a Live/Dead® Viability/Cytotoxicity assay. Optimization of the LDH assay allowed for earlier detection of changes in %cytotoxicity with lower doses (50µM H2O2 for 6h) than the supernatants obtained alongside treatments for western blotting. Furthermore, an earlier upregulation in %cytotoxicity was observed with 100µM H2O2  (6h compared to 9h in the western blotting experiments). Cell death was observed via visual inspection of cells starting at 4-6h with 100µM H2O2. The 82  Live/Dead® Viability/Cytotoxicity assay also showed a significant reduction in calcein absorbance at 6h, indicating a loss in viable cells and a significant upregulation in EthD-1, indicating an increase in dead cells, by 8h. Overall, ARNT2 regulation in response to mediators capable of inducing cell death appears to vary depending on the stimulus used, and appears to be influenced less by apoptotic stimuli compared to when necrosis is primarily occurring. In the case of H2O2, ARNT2 appears to follow a temporal gradient, undergoing a reduction as cell death becomes apparent via visual inspection, LDH release and the Live/Dead® Viability/Cytotoxicity assay.  As these results are correlative in nature, future experiments should include knockdown of Arnt2 and observation of changes in cell viability to deduce a functional role for this protein in maintaining cell health in primary neurons and astrocytes as well as in vivo.  The upregulation of ARNT2 in primary neurons and astrocytes is a novel finding, and has never been described in other cell types in vitro. Previous studies have mostly shown that neuronal ARNT2 levels remain steady, while its binding partners change in localization/expression [112]. We have shown here, for the first time, that ARNT2 expression patterns are indeed enhanced by apoptotic stimuli and oxidative stress in primary cortical neuron-enriched cultures. In vivo data showed an increase in Arnt2 mRNA pre-disease onset concurrent with Bdnf (Quandt lab, unpublished data). It is known that in EAE some neuronal loss is present early in the disease process, and that blocking reactive oxygen species is beneficial in EAE [5] [52]. In the context of neuronal health in response to stressors involved in axonal damage and loss in MS, the upregulation of ARNT2 could potentially result in enhanced BDNF production for the protection of axonal integrity and for axonal regeneration, as described in section 1.2.3 [89]. However, while ARNT2 is known to be responsible for BDNF production 83  through its heterodimerization with NPAS4, their transcriptional activity of this protein in the context of chronic neuroinflammation/neurodegeneration has not yet been studied [112]. In the spinal cord, inflammatory infiltrates are much higher in prevalence than in the brain in EAE, and contribute to neuroaxonal damage and loss via the release of factors including reactive oxygen species [41]. Arnt2 mRNA was significantly reduced at peak disease in our preliminary data in both brain and spinal cord (Quandt lab, unpublished data). The intensity of neuroaxonal damage in the cord at peak disease may therefore correlate with a reduction in Arnt2 mRNA. The reduction in Arnt2 mRNA in the brain could result from spinal cord neuroaxonal damage spreading to the brain, and/or from the few infiltrates in the brain. The significant upregulation of Arnt2 mRNA in the brain pre-onset of the disease is reminiscent of the significant increase in ARNT2 protein with short-term applications of H2O2, and could thus potentially be a response to a mild/early disease stimulus.  6.7 Limitations  There are some weaknesses and limitations associated with this study. The majority of our data comes from an in vitro model of cortical neurons and astrocytes, which, while allowing us to discern changes in ARNT2 expression in response to stimuli in a very specific and controlled manner, is not particularly reflective of the progressive disease. The heterogeneity of MS in terms of disease progression, lesion types and locations affected pose a challenge for modeling multiple aspects of this disease in vitro. For this reason, an in vivo animal model was also used for the purposes of discerning ARNT2 expression patterns. While the EAE model of MS shares many similarities with MS, they also have some differences with respect to disease localization and clinical presentation, as discussed in section 1.1.4 [67]. Additionally, our in vitro 84  work models cortical neurons and astrocytes, while, in EAE, the spinal cord is mostly affected, making it difficult to draw direct correlations between the two models, though allowing us to model various different regions affected in MS [52].   We relied on western blotting for a large amount of this study, which, as a semi-quantitative method, does not allow for optimal sensitivity. For this reason we also used immunocytochemistry, a more sensitive technique. Additionally, there are some limitations associated with using immunohistochemistry techniques. The success of immunohistochemistry relies on proper fixation (type, procedure, duration; capable of influencing antibody affinity and selectivity because of protein conformation alterations and epitope masking), detection method and tissue quality. For this reason, various optimization steps are required, as well as the use of an isotype control, a control using only secondary antibody, and a control staining of tissue devoid of ARNT2. Finally, while our immunohistochemistry was not a quantitative analysis, it was appropriate for our purposes as it allowed for ARNT2 distinction in various cell types, across multiple levels of the spinal cord, and allowed us to observe changes in ARNT2 at specific lesion sites.    Furthermore, the relationship between ARNT2 and changes in cell viability is entirely correlative in this study, and needs to be examined through more functional methodology. Additionally, while the binding partner for ARNT2 in response to stressors involved in MS disease progression could likely be NPAS4, it requires confirmation.  85  6.8 Conclusions and future directions   This study allowed us to observe ARNT2 expression patterns in neurons, and also begin to understand its expression in astrocytes, at the cellular level and in vivo in response to processes that contribute, directly or indirectly, to axonal degeneration and neuronal loss.  The neuroprotective properties of ARNT2 have thus far been studied irrespectively of its ability to produce the neurotrophic protein BDNF [93]. Future experiments should include correlating BDNF levels to ARNT2 levels following exposure to stressors in neuronal cultures, using western blotting or an enzyme-linked immunosorbent assay (ELISA) kit (Promega, Madison, US), as well as in tissue from animal models of MS and post-mortem tissue samples.  In vitro, a luciferase reporter assay at Bdnf promoter IV should allow us to determine whether treatments with oxidative stress/apoptotic stimuli which upregulate ARNT2 enhance Bdnf transcription [152]. For this technique, we would clone this regulatory region of Bdnf upstream of the luciferase gene in an expression vector, introduce the vector into cells and allow them to grow for an optimized period of time. Following collection and lysis of cells, luciferin and co-factors are added to measure luciferase activity via a luminometer. Cross-linked chromatin immunoprecipitation (ChIP), which allows for investigation of protein-DNA interactions, could be used to determine the specific interaction of ARNT2 with the DNA target on the Bdnf promoter in response to these stressors. For this technique, formaldehyde is used to cross-link proteins to DNA, which will need to be optimized, followed by sonication of the lysate, sedimentation of cell debris and immunoprecipitation using ARNT2 antibody. Following protein digestion, the DNA can be purified and identified using various techniques including PCR. 86   To determine whether there is an increase in functional BDNF (i.e. protein) following application of these stressors that increase ARNT2 protein expression, western blots would need to be performed, examining time periods overlapping and extending our ARNT2 investigation. As BDNF can exist in multiple isoforms, the antibody chosen would have to be capable of recognizing a common sequence. In addition, a blocking peptide would be utilized to determine antibody specificity.   ChIP and western blotting could also be used in tissue from mice undergoing EAE (compared to sham immunized and healthy litter mates), using antibodies for ARNT2 and BDNF, to determine the regulatory function of ARNT2, as well as to establish similarities and/or differences in expression patterns of ARNT2 and BDNF at various stages of the disease. Thus far, preliminary data from our lab has shown that Arnt2 and Bdnf mRNA follow the same expression pattern in the spinal cord of EAE mice throughout the disease process.  A co-immunoprecipitation procedure following stimulation with these stressors would clarify the dimerization partner for ARNT2 [89] [90]. Following stimulation, harvest and capture of target, western blots would be run for ARNT2 alongside its known binding partners using specific antibodies. For this, a negative control would include using beads without antibody/with a non-specific antibody, for which we should see no bands via western blotting.   To examine expression patterns of ARNT2 in vivo, they will need to continue to be localized and compared throughout the various stages of the animal model of MS, as well as in post-mortem tissue of individuals with MS, with further refinements based on cell-type (e.g. ARNT2 expression in neurons, astrocytes, infiltrating immune cells, and other glia), with continued use of necessary controls, including an isotype control for each antibody, and staining only with secondary antibody. Findings from this study could then be compared to tissues from 87  Alzheimer’s, Parkinson’s, traumatic brain injury and ischemic tissue. These experiments would further our understanding of the ARNT2 expression patterns in the processes of neuroinflammation and neurodegeneration.   Furthermore, ARNT2 has previously been shown to be an inhibitor of apoptosis in PC12 cells [93]. The role of ARNT2 in cell survival should be further investigated in primary cells. Here we have shown that ARNT2 protein expression in neurons is affected by oxidative stress. To examine the functional relevance of downregulation of ARNT2, knockdowns should be tested in vitro in neuronal/astrocytic cultures, as well as in vivo. In primary cultures, knockdowns could be performed using viral transfection of Arnt2 siRNA. Arnt2 knockdown could be tested at various degrees (e.g. 25%, 50% and 90% knockdown), following optimization of dose delivered, to observe the effects on cell viability. These siRNA would be tagged with fluorescent protein to allow for confirmation of transfection via fluorescence microscopy. To ensure the siRNA will hybridize only to the mRNA of interest, a BLAST search would be performed. These proportional knockdowns would also have to be confirmed using qPCR, compared to untreated (non-transfected) and scrambled siRNA controls. Furthermore, western blots would have to be performed to confirm if RNA knockdown results in downregulation of protein expression. As an antibody control, a blocking peptide would be used to determine antibody specificity. As mRNA/protein expression discordance was observed in our qPCR studies, it is possible that a reduction in Arnt2 mRNA may not immediately result in ARNT2 protein downregulation. If this is the case, the knockdown would be expanded with continued exposure to siRNA, to determine if prolonged and persistent downregulation of Arnt2 mRNA would result in a reduction in protein levels. If no direct loss in cell viability is observed with Arnt2 knockdown, a dose of H2O2 previously shown to not result in cell death in our cultures could be used, to observe 88  whether a reduction in Arnt2 could result in death with this dose. Furthermore, a rescue experiment (introducing Arnt2 cDNA) would be performed. This study would allow us to determine whether ARNT2 may function as a sensitive determinant of neuronal stress, or function as a constitutive influence on neuronal health under both steady state and stress conditions. In vivo, a Cre-Lox recombination method for specific downregulation in neurons versus astrocytes could be used, which allows for specific temporal regulation and could thus allow for Arnt2 downregulation at various stages of the disease. In vivo, knockdowns would have to be established in postnatal mice, as Arnt2 knockdown in embryonic mice results in perinatal death [97] [98]. These mice would be compared to non-transfected controls at each stage of the disease. Additionally, to confirm cell-specific expression, Cre reporter strains designed to express fluorescent protein after Cre recombinase excision of the loxP-flanked stop sequence would be used. Also, to examine the functional relevance of early upregulation of ARNT2 observed in these studies, ARNT2 levels should be increased (via cDNA transfection) to determine whether this rescues cells from a pathway to death under conditions that would normally result in death. In a study by Drutel et al. (1999), oxidative stress induced cell death could be inhibited by introduction of Arnt2 cDNA in PC12 cells, but this has not been studied in neurons or astrocytes to date [94]. Furthermore, Bdnf levels could also be observed following Arnt2 knockdown and rescue using qPCR and western blotting, to further understand the neuroprotective roles of ARNT2 with respect to Bdnf transcription. As an antibody control, tissue could be collected from a region known to have no ARNT2 expression (from lung, for example) as a negative control.   Enhancing our understanding of neuroprotection remains a major goal in identifying novel therapeutics to limit disease severity and progression in MS [82]. 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Neuron. 1998;20(4):709-726. 102  Appendices  Appendix A    A.1 Antibodies used for western blotting    A.2 Antibodies used for immunocytochemistry/immunohistochemistry   Primary Antibody Catalog Number Dilution Species Secondary Antibody Catalog Number Dilution ARNT2 sc-5581 1/2000 rabbit Goat anti-rabbit 111-035-045 1/2000 Tubulin MAB1864 1/1000 rat Donkey anti-rat 712-035-153 1/2000 Primary Antibody Species Catalog Number Dilution ARNT2 Rabbit sc-5581 1:250 GFAP Chicken AB5541 1:2500 (IC) 1:2000 (IH) MAP2 Mouse M9942 1:750 Secondary Antibody Catalog Number Dilution Wavelength (nm) Goat anti-rabbit A11036 1:500 568 Goat anti-chicken A11036  1:1000 (IC) 1:500 (IH) 647 Goat anti-mouse A11029 1:500 488 Normal IgG Species Catalog Number Dilution Rabbit Sc-5581 1:500 Chicken AB5541 1:1000 Mouse M9942 1:160 103  Appendix B   B.1 Primers used for qPCR    B.2 Run conditions used for qPCR          Gene Forward (5’ – 3’) Reverse (5’ – 3’) Species Arnt2 CCA GTC TTG CCA ACA GGA CTC C AGC ATG TCC TGG AAC ACT TCA GTC Mouse, rat, human Actb CCA GCC TTC CTT CCT GGG TAT TGT GTT GGC ATA GAG GTC TTT ACG Rat Program No. Cycles Target Temp (°C) Acquisition Mode Hold (hh:mm:ss) Ramp rate (°C/s) Acquisition (per °C) Sec target (°C) Step Size Step Delay (Cycles) Preincubation 1 95 None 00:10:00 4.4 0 0 0 Amplification 40 95 None 00:00:05 4.4 0 0 0 60 None  00:00:10 2.2 0 0 0 72 Single 00:00:05 4.4 0 0 0 Melting Curves 1 95 None 0:01:00 4.4 45 None 0:00:01 2 95 Continuous 5 Cooling 1 40 None 00:00:01 1.5 0 0 0 104  B.3 Western blot results for lysates used for qPCR experiments  Appendix B.3: Western blots matching the qPCR experiments, though underpowered, exhibit the same patterns as previous western blotting experiments with these doses of H2O2 (Figure 3.10, 3.12; matching qPCR Figure 3.14). Points represent three biological replicates, where each dot represents the average of two technical replicates each, with mean and standard deviation. Repeated measures ANOVA on Ranks with Tukey Multiple Comparisons Test to unstimulated.           unstim 0.5 1 2 3 4 5 6 7 8 9 100.00.51.01.52.02.5Time (hours)ARNT2/Tubulin Ratiounstim 0.5 1 2 3 4 50.00.51.01.52.02.5Time (hours)ARNT2/Tubulin Ratio25µM H2O2 100µM H2O2 A B               Time (hours)                   ARNT2/ Tubulin Ratio               Time (hours)                   ARNT2/ Tubulin Ratio 105  Appendix C   C.1 Testing of the Live/Dead® Viability/Cytotoxicity Assay with 2µM Calcein-AM and EthD-1   Appendix C.1: Testing of the Thermo Fischer Scientific LIVE/DEAD® viability/cytotoxicity assay kit in primary cortical neuron-enriched cultures. 2uM of both calcein and ethidium homodimer-1 (EthD-1) were used. Bars represent mean and standard deviation of three technical replicates. p≤0.05*/+, p≤0.01**/++, p≤0.001***/+++, repeated measures ANOVA on Ranks with Tukey Multiple Comparisons Test to unstimulated, (* - Calcein, + - EthD-1).  02004006008001000020406080100Unstim Dead 2 4 6 8 12 18 24Time  (hours)25µM H2O2***++++ +++++Calcein EthD-102004006008001000020406080100Unstim Dead 2 4 6 8 12 18 24Time (hours)50µM H2O2**++++ +++Calcein EthD-102004006008001000020406080100Unstim Dead 2 4 6 8 12 18 24Time (hours)100µM H2O2***+++ +++Calcein EthD-1+++++++

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