Medicine, Faculty of
Pathology and Laboratory Medicine, Department of
DSpace
UBCV
Abdul-Rahman, Raneen
Master of Science - MSc
University of British Columbia
Multiple sclerosis (MS) is characterized by inflammatory demyelination and axonal loss, which contribute to irreversible clinical disability. Both hallmarks affect calcium homeostasis, partly due to mediators released by inflammatory cells, like glutamate and hydrogen peroxide (H₂O₂). Nuclear calcium transients can activate neuroprotective factors, including neuronal PAS-domain containing protein 4 (NPAS4), a neuronal activity-dependent gene. NPAS4 and aryl hydrocarbon receptor nuclear translocator 2 (ARNT2) work independently or in combination to regulate genes responsible for neuronal survival. We sought to determine the effects of soluble inflammatory mediators and inflammatory cells relevant to MS pathogenesis on NPAS4 and ARNT2 expression. Murine neuron-enriched cortical cultures were treated with increasing concentrations of glutamate (1-30µM) and H₂O₂ (12.5-100µM) or co-cultured in pilot experiments with resting or activated CD4⁺ T-cells. NPAS4 and ARNT2 protein expression were characterized via immunocytochemistry by examining the neuronal nuclear mean fluorescent intensity (MFI), the percentage of positive cells, and staining intensity changes using fluorescence microscopy across time and concentrations. In untreated conditions, NPAS4 was detected in less than 10% of neurons, while greater than 70% of neurons expressed low-medium ARNT2 levels. Glutamate increased the NPAS4 MFI and percentage of NPAS4-positive neurons, with increased staining intensity observed early at all doses. ARNT2 staining levels were similar across the glutamate doses and times. Prolonged glutamate exposure (10-30µM) was associated with morphological changes and cell loss. H₂O₂ (50-100µM) increased the NPAS4 MFI and percentage of NPAS4-expressing cells at later time points (5hrs and greater), with a simultaneous decrease and loss of ARNT2 expression. Exposure to H₂O₂ at 50-100µM tended to impact neuronal cell viability with significant cell loss by 9 hours compared to controls. Lastly, the addition of resting or activated CD4⁺ T-cells, at a 10- to 20-fold higher ratio to neurons, tended to increase the NPAS4 MFI and rapidly increased the percentage of high-intensely expressing cells without affecting ARNT2 expression. This study shows that MS-related inflammatory mediators and CD4⁺ lymphocytes modulate in vitro neuronal NPAS4 and ARNT2 protein expression. Further research is required to elucidate the relationship between these proteins and how changes in their expression might impact neuronal health in inflammatory degenerative settings.
Characterizing the Neuronal Expression of Neuronal PAS-Domain Containing Protein 4 and Aryl Hydrocarbon Receptor Nuclear Translocator 2 in Multiple Sclerosis Related Inflammatory Settings In Vitro by Raneen Abdul-Rahman 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) November 2024 © Raneen Abdul-Rahman, 2024 ii The following individuals certify that they have read, and recommend to the Faculty of Graduate and Postdoctoral Studies for acceptance, the thesis entitled: Characterizing the Neuronal Expression of Neuronal PAS-Domain Containing Protein 4 and Aryl Hydrocarbon Receptor Nuclear Translocator 2 in Multiple Sclerosis Related Inflammatory Settings In Vitro Submitted by Raneen Abdul-Rahman in partial fulfilment of the requirements for the degree of Master of Science in Pathology and Laboratory Medicine Examining Committee: Jacqueline Quandt, Assistant Professor, Pathology and Laboratory Medicine, UBC Supervisor Wayne Moore, Clinical Professor, Pathology and Laboratory Medicine, UBC Supervisory Committee Member Kevin Bennewith, Professor, Pathology and Laboratory Medicine, UBC Additional Examiner Clare Beasley, Associate Professor, Psychiatry, UBC Additional Examiner Additional Supervisory Committee Members: Shernaz Bamji, Professor, Cellular and Physiological Sciences, UBC Supervisory Committee Member Yu Tian Wang, Professor, Medicine, UBC Supervisory Committee Member iii Abstract Multiple sclerosis (MS) is characterized by inflammatory demyelination and axonal loss, which contribute to irreversible clinical disability. Both hallmarks affect calcium homeostasis, partly due to mediators released by inflammatory cells, like glutamate and hydrogen peroxide (H2O2). Nuclear calcium transients can activate neuroprotective factors, including neuronal PAS-domain containing protein 4 (NPAS4), a neuronal activity-dependent gene. NPAS4 and aryl hydrocarbon receptor nuclear translocator 2 (ARNT2) work independently or in combination to regulate genes responsible for neuronal survival. We sought to determine the effects of soluble inflammatory mediators and inflammatory cells relevant to MS pathogenesis on NPAS4 and ARNT2 expression. Murine neuron-enriched cortical cultures were treated with increasing concentrations of glutamate (1-30µM) and H2O2 (12.5-100µM) or co-cultured in pilot experiments with resting or activated CD4+ T-cells. NPAS4 and ARNT2 protein expression were characterized via immunocytochemistry by examining the neuronal nuclear mean fluorescent intensity (MFI), the percentage of positive cells, and staining intensity changes using fluorescence microscopy across time and concentrations. In untreated conditions, NPAS4 was detected in less than 10% of neurons, while greater than 70% of neurons expressed low-medium ARNT2 levels. Glutamate increased the NPAS4 MFI and percentage of NPAS4-positive neurons, with increased staining intensity observed early at all doses. ARNT2 staining levels were similar across the glutamate doses and times. Prolonged glutamate exposure (10-30µM) was associated with morphological changes and cell loss. H2O2 (50-100µM) increased the NPAS4 MFI and percentage of NPAS4-expressing cells at later time points (5hrs and greater), with a simultaneous decrease and loss of ARNT2 expression. Exposure iv to H2O2 at 50-100µM tended to impact neuronal cell viability with significant cell loss by 9 hours compared to controls. Lastly, the addition of resting or activated CD4+ T-cells, at a 10- to 20-fold higher ratio to neurons, tended to increase the NPAS4 MFI and rapidly increased the percentage of high-intensely expressing cells without affecting ARNT2 expression. This study shows that MS-related inflammatory mediators and CD4+ lymphocytes modulate in vitro neuronal NPAS4 and ARNT2 protein expression. Further research is required to elucidate the relationship between these proteins and how changes in their expression might impact neuronal health in inflammatory degenerative settings. v Lay Summary Brain cell loss is a major factor driving disease progression in multiple sclerosis (MS), making neuronal protection a critical focus in MS research. This study aims to characterize how neuronal PAS-domain containing protein 4 (NPAS4) and aryl hydrocarbon receptor nuclear translocator 2 (ARNT2), two proteins thought to protect neurons, are changed when neurons are exposed to inflammatory factors and cells associated with MS. Exposure to both glutamate and hydrogen peroxide (H2O2) increased NPAS4 expression, with glutamate exerting more rapid effects, while high doses increased cell death and tended to decrease ARNT2 levels. Initial tests with immune cells increased NPAS4 expression but had little effect on ARNT2. These results suggest that NPAS4 and ARNT2 may play a role in neuronal damage in MS stages where these factors are present. More research is needed to understand the impact of these proteins on neuron health in MS-like inflammation. vi Preface All experiments were conducted at the University of British Columbia (UBC) to fulfill the requirements for the Master of Science degree in Pathology and Laboratory Medicine. Animal studies were conducted in accordance with Protocol A18-0361 and A22-0283, both approved by the UBC Animal Care Committee. Figures 1.1 and 2.1 were created in BioRender.com. Figure 1.2 was created by Dr. Jacqueline Quandt and Quandt laboratory members and includes unpublished data as well as previously published data from “Expression of the Neuroprotective Protein Aryl Hydrocarbon Receptor Nuclear Translocator 2 Correlates with Neuronal Stress and Disability in Models of Multiple Sclerosis” by Rahim et al. (2018)1. I prepared all remaining figures using GraphPad Prism 10 and the Zen Blue 3 Software. Emily Kamma assisted with the isolation and purification of CD4+ T-cells. The results presented in Chapter 3 were from experiments I conducted with guidance on experimental planning from my supervisor, Dr. Jacqueline Quandt, and recommendations provided by my supervisory committee. Otherwise, all remaining work comprises my original, unpublished work. vii Table of Contents Abstract ......................................................................................................................................... iii Lay Summary ................................................................................................................................ v Preface ........................................................................................................................................... vi Table of Contents ........................................................................................................................ vii List of Figures ............................................................................................................................... ix List of Abbreviations ................................................................................................................... xi Acknowledgements ..................................................................................................................... xv Chapter 1: Introduction ............................................................................................................... 1 1.1 Multiple Sclerosis ....................................................................................................................... 1 1.1.1 Overview ................................................................................................................................................ 1 1.1.2 Pathogenesis ........................................................................................................................................... 2 1.1.3 Inflammation and Neurodegeneration.................................................................................................... 6 1.1.4 Current Therapeutic Strategies ............................................................................................................... 9 1.2 Neuronal PAS-Domain Containing Protein 4 (NPAS4)........................................................ 13 1.2.1 Overview .............................................................................................................................................. 13 1.2.2 Expression Patterns .............................................................................................................................. 15 1.2.3 Functional Roles................................................................................................................................... 16 1.2.4 Neuroprotective Roles .......................................................................................................................... 18 1.2.5 ARNT2: NPAS4’s Primary Binding Partner ....................................................................................... 22 1.2.6 NPAS4/ARNT2: A Role in Chronic Inflammatory Neurodegenerative Settings Relevant to MS...... 22 1.3 Thesis Objectives and Hypothesis ........................................................................................... 25 1.3.1 Aims ..................................................................................................................................................... 25 1.3.2 Clinical Significance ............................................................................................................................ 25 Chapter 2: Materials and Methods ........................................................................................... 27 2.1 Mice ........................................................................................................................................... 27 2.2 In Vitro Cultures ...................................................................................................................... 27 2.2.1 Murine Primary Neuron-Enriched Cortical Cultures ........................................................................... 27 2.2.2 CD4+ T-Cell Isolation .......................................................................................................................... 28 2.2.3 Murine Immortalized Brain Endothelial Cell Line (bEnd.3) ............................................................... 29 2.2.4 Soluble Inflammatory Mediators ......................................................................................................... 29 2.3 Immunocytochemistry ............................................................................................................. 29 2.3.1 Staining Protocol .................................................................................................................................. 29 2.3.2 Image Acquisition and Immunocytochemistry Analysis ..................................................................... 30 2.4 Statistical Analysis.................................................................................................................... 33 Chapter 3: Effects of Inflammatory Mediators and Cells on NPAS4 and ARNT2 Expression .................................................................................................................................... 34 3.1 Characterization of Primary Cortical Neuron-Enriched Cultures ..................................... 34 3.2 Effects of Inflammatory Mediators on NPAS4 Expression .................................................. 36 3.2.1 Glutamate Rapidly Alters NPAS4 Expression ..................................................................................... 36 3.2.2 High Doses of H2O2 Induce Late NPAS4 Expression ......................................................................... 46 viii 3.3 Effects of Inflammatory Mediators on ARNT2 Expression ................................................. 53 3.3.1 High Doses of Glutamate Tended to Reduce ARNT2 Levels ............................................................. 53 3.3.2 Prolonged Exposure to High Doses of H2O2 Reduced ARNT2 Levels ............................................... 59 3.4 Linking Effects of Inflammatory Mediators on ARNT2 and NPAS4 Intensity Levels to Viability Across Select Focus Treatment Groups .............................................................................. 64 3.4.1 Categorical Changes in NPAS4 and ARNT2 Expression During Glutamate and H2O2 Exposure...... 64 3.5 Effect of Inflammatory Cells on NPAS4 and ARNT2 Expression....................................... 70 3.5.1 The Number of Inflammatory T-Cells Rather than Activation Status Seems to Induce Early NPAS4 Expression .......................................................................................................................................................... 70 3.5.2 CD4+ T-lymphocytes Do Not Alter ARNT2 Expression ..................................................................... 73 Chapter 4: Discussion ................................................................................................................. 76 4.1 Summary of Key Findings ....................................................................................................... 76 4.2 NPAS4 and ARNT2 in the Cortex .......................................................................................... 77 4.3 The Influence of Glutamate on NPAS4 and ARNT2 Expression ........................................ 81 4.4 Effects of H2O2 on NPAS4 and ARNT2 Expression .............................................................. 85 4.5 NPAS4 and ARNT2 as Potential Mediators of Cell Viability .............................................. 89 4.6 Effects of Inflammatory Cells on NPAS4 and ARNT2 Expression ..................................... 91 4.7 Global Assessment of Combined Changes in NPAS4 and ARNT2 ..................................... 92 4.8 Limitations ................................................................................................................................ 94 4.9 Future Directions and Conclusions ........................................................................................ 95 References .................................................................................................................................. 100 Appendix: List of Specific and Isotype/Irrelevant Antibodies Used for Immunocytochemistry .............................................................................................................. 119 ix List of Figures Figure 1.1 ARNT’s role in activity-dependent upregulation of genes in the CNS.. .............. 22 Figure 1.2 Expression changes in Arnt2, Npas4 and Bdnf RNA throughout the course of active EAE. .................................................................................................................................. 24 Figure 2.1 Flowchart depicting experimental protocol for culturing primary cortical cells, application of inflammatory mediators and quantification of NPAS4 and ARNT2 protein expression..................................................................................................................................... 32 Figure 3.1 Characterization of primary neuron-enriched cortical cultures.. ....................... 35 Figure 3.2 NPAS4 expression in response to excitatory stimuli. ............................................ 37 Figure 3.3 Glutamate rapidly induced NPAS4 expression...................................................... 41 Figure 3.4 Early changes in NPAS4 intensity at all doses of glutamate................................. 43 Figure 3.5 Decreased cell viability following exposure to higher doses of glutamate. .......... 45 Figure 3.6 Prolonged exposure to high doses of H2O2 induced NPAS4 expression. ............. 49 Figure 3.7 Changes in NPAS4 intensity observed as early as 3 hours post-exposure to high doses of H2O2. .............................................................................................................................. 50 Figure 3.8 Decreased cell viability at high doses of H2O2. ....................................................... 52 Figure 3.9 ARNT2 is expressed constitutively in DIV10 cortical neurons. ........................... 54 Figure 3.10 High doses of glutamate tended to reduce ARNT2 levels. .................................. 57 Figure 3.11 High doses of glutamate tended to reduce ARNT2 intensity levels. .................. 59 Figure 3.12 Prolonged exposure to high doses of H2O2 reduces ARNT2 expression levels. 62 Figure 3.13 Changes in ARNT2 intensity levels observed at prolonged exposure to H2O2. 64 x Figure 3.14 High dose glutamate rapidly upregulates NPAS4 while downregulating ARNT2.. ....................................................................................................................................... 66 Figure 3.15 Prolonged exposure to 100µM H2O2 increases NPAS4 expression while simultaneously decreasing ARNT2 levels. ................................................................................ 69 Figure 3.16 Number but not activation status of CD4+ T cells drives NPAS4 expression. . 73 Figure 3.17 ARNT2 expression was unchanged by neuronal cell incubation with resting or activated CD4+ T-cells. ............................................................................................................... 75 xi List of Abbreviations AAV Adeno-associated virus ACC Animal care committee AD Alzheimer’s Disease AMPA α-amino-3-hydroxy-5-methylisoxazole-4-propionate ANOVA Analysis of variance APC Antigen-presenting cell APP Amyloid precursor protein ARNT2 Aryl hydrocarbon nuclear translocator 2 ATP Adenosine triphosphate AUC Area under the curve BBB Blood-brain barrier BDNF Brain-derived neurotrophic factor bEnd.3 Murine immortalized brain endothelial cell bHLH-PAS Basic-helix-loop-helix PER ARNT SIM Ca2+ Calcium CNS Central nervous system CO2 DIV DMEM DMT DNA Carbon dioxide Day in vitro Dulbecco’s modified eagle medium Disease-modifying therapy Deoxyribonucleic acid xii DPBS Dulbecco’s phosphate-buffered saline EAE Experimental autoimmune encephalomyelitis EBV Epstein Barr virus EDTA Ethylenediaminetetraacetic acid EGTA Ethylene glycol tetraacetic acid ELISA Enzyme-linked immunosorbent assay FAD Focal axonal degeneration FBS Fetal bovine serum FDA Food and drug administration GFAP Glial fibrillary acidic protein GLAST Glutamate/aspartate transporter H2O2 Hydrogen peroxide HBSS Hank’s balanced salt solution HEPES 2-(4-(2-hydroxyethyl)-1-piperazinyl)-ethanesulfonic acid HIF-1α Hypoxia-inducible factor 1α HLA Human leukocyte antigen IEG Immediate early gene IFN-γ Interferon-gamma IL Interleukin IQR Interquartile range KCl Potassium chloride KO Knockout xiii MAP2 Microtubule-associated protein 2 MFI Mean fluorescent intensity MHC Major histocompatibility complex MRI Magnetic resonance imaging mRNA Messenger ribonucleic acid MS Multiple sclerosis NBM Neurobasal medium NCoR2 Nuclear receptor co-repressor 2 NEAA Non-essential amino acids NeuN Neuronal nuclear protein NFH Neurofilament heavy chain NMDA NMM N-methyl-D-aspartate Neuronal maintenance media NPAS4 Neuronal PAS-domain containing protein 4 OPC Oligodendrocyte precursor cell PBS Phosphate buffered saline PDC Preclinical Discovery Center PPMS Primary progressive multiple sclerosis qPCR Quantitative polymerase chain reaction RCT Randomized controlled trial RNA Ribonucleic acid ROS Reactive oxygen species xiv RPMI RRMS Roswell Park Memorial Institute medium Relapsing-remitting multiple sclerosis SIM1 SPMS Single-minded homolog 1 Secondary progressive multiple sclerosis TBS Tris-buffered saline TBST TCM TCR TEMPOL TNF-α UBC VGCC Tris-buffered saline with tween T-cell media T-cell receptor 4-hydroxy-2,2,6,6-tetramethylpiperidine-N-oxyl Tumour necrosis factor alpha University of British Columbia Voltage-gated calcium channel xv Acknowledgements Firstly, I would like to thank my supervisor, Dr. Jacqueline Quandt, for her unwavering support, insightful guidance and expertise, which have been invaluable throughout this journey. I am grateful for your mentorship. Thank you for continuously challenging me to think critically and improve my work, and in doing so, enabling my personal and professional growth. I would also like to express my deepest gratitude to my committee members, Dr. Helene Cote, Dr. Shernaz Bamji, Dr. Wayne Moore, Dr. Yu Tian Wang, and Dr. Dana Devine, for their time, guidance, and invaluable input during committee meetings. I would also like to extend a heartfelt thanks to past and present members of the Quandt lab for their assistance and support. I am forever grateful for the friendships made with such wonderful people. I would especially like to thank Pierre Becquart who took the time to train me and teach me numerous experimental techniques throughout the first two years of this degree, as well as Curtis Wong for his guidance and expertise. I would also like to thank Emily Kamma for her guidance and assistance in T-cell isolation experiments. I would like to give a special thanks to Katie Mayne for her encouragement and support. I would also like to express my appreciation to the staff at the Preclinical Discovery Center for their help and guidance involving breeding and timed pregnancies. To my friends and family, your unwavering support and belief in me have been my anchor. Thank you for always being there to listen, offer advice, and keep me motivated. Your love, patience, and constant encouragement have carried me throughout this journey, and I am deeply grateful for all you have done to help me reach this milestone. I am truly grateful to have such amazing people in my life, and I would not be the person I am today without your steadfast support. 1 Chapter 1: Introduction 1.1 Multiple Sclerosis 1.1.1 Overview Multiple sclerosis (MS) is a chronic, inflammatory, and neurodegenerative disorder of the central nervous system (CNS), comprising the brain and spinal cord2. Affecting over 100,000 Canadians and more than 2.8 million people globally, MS is the most common cause of non-traumatic neurological disability amongst young adults, with evidence indicating a rising prevalence3-5. It consists of different subtypes, each of which is important for prognosis and treatment decisions6. The majority of individuals (~80%) present with relapsing-remitting MS (RRMS), characterized by episodes of acute attacks followed by periods of remission. In RRMS, an inflammatory response leads to activated immune cells attacking myelin and nerve fibers. This attack results in inflammatory demyelinating lesions, axonal loss and ultimately, the common symptoms associated with MS, such as visual impairments, paresis, fatigue and spasticity7. Over time, approximately 65% of RRMS patients transition to secondary progressive MS (SPMS), which is marked by a gradual decline in neurological function independent of relapses8. A smaller proportion of individuals (~15%) present with primary progressive MS (PPMS), where disability accumulates progressively from onset without distinct relapses or remissions8,9. While axonal loss begins early in the disease course, as confirmed by in vivo magnetic resonance imaging (MRI) studies, it is the irreversible axonal loss and sustained neurodegeneration that is strongly associated with chronic disability in MS10,11. MS is a multifaceted disease, and while the etiology of the disease remains largely unknown, numerous environmental and genetic factors contribute to its development and progression6. Most individuals develop MS between 20 and 40 years of age, with RRMS onset 2 typically occurring 10 years earlier than progressive MS (i.e., PPMS or SPMS)12,13. Numerous environmental factors, including exposure to viral agents (notably Epstein Barr virus [EBV] and human herpes virus type 6)14,15, smoking16, vitamin D deficiency17, diet18,19 and ultraviolet radiation exposure20 have all been associated with an increased risk of developing MS. A longitudinal study following a cohort of EBV-negative individuals at baseline recently showed more than a 30-fold increase in MS risk following EBV infection15,21, which the authors suggested was data in support of a causal link between EBV and MS. Genetic predisposition also contributes to MS risk, as indicated by familial studies where MS risk increased based on the amount of shared genetic information22-24. Studies examining the familial recurrence rate of MS showed that in the cases where one twin is affected, the risk rate of the other twin developing MS is 25% for monozygotic twins (100% genetic similarity) and 2-5% for dizygotic twins (50% genetic similarity)22-24. These genetic predisposition studies also pointed to the human leukocyte antigen (HLA) of chromosome 6 as a significant genetic factor, increasing MS risk by up to threefold25. Additionally, MS is more prevalent in females, with three times as many females diagnosed as males5,26-28. Therefore, while the exact cause of MS remains unknown, both genetic and environmental factors contribute to the likelihood of developing MS. 1.1.2 Pathogenesis MS pathogenesis is characterized by several pathological hallmarks, including vascular alterations (i.e., BBB dysfunction and increased permeability), inflammatory cell infiltration, demyelination, gliosis, oligodendrocyte loss and, ultimately, axonal damage/loss and neurodegeneration2,29. Historically, MS has been regarded as a white matter disease due to the characteristic demyelination of myelinated axonal tracts30. White matter, which also contains oligodendrocytes responsible for myelination, lies beneath the CNS gray matter to enable 3 cortical and cortical to subcortical connections as well as connecting the brain to the spinal cord. In contrast, the gray matter, composed primarily of neuronal cell bodies, dendrites and some myelinated and unmyelinated axons, is located on the brain’s surface and deeper subcortical structures31. It plays a central role in information processing and higher cognitive functions. Despite their differences, the CNS white and gray matter work together to coordinate a broad spectrum of cognitive, sensory and motor functions. However, despite the historical belief that MS predominantly affects the white matter, recent histological and imaging findings suggest that the pathological hallmarks of MS differentially affect both the CNS white and gray matter32-35. White matter lesions evolve differently based on the disease stage and phase29. In individuals living with RRMS, focal new and classical active white matter lesions are most common. These lesions are marked by hypercellular demyelinated plaques heavily infiltrated by immune cells, both B- and T-lymphocytes, and myelin-laden macrophages29,36. In the progressive stages of MS, the most common plaques include slowly expanding active lesions with an inactive demyelinated center surrounded by a rim of activated myelin-laden microglia and macrophages29,36. The most prevalent lesions in the MS brain are inactive lesions, characterized by demyelination, partial axonal preservation, and reactive gliosis in the absence of active inflammation29,36. Although cortical demyelination has been observed in the early stages of MS, it significantly increases in the progressive phase36-38 and is a key factor driving cognitive decline in MS39. The CNS was traditionally considered an immune-privileged organ due to a physical BBB formed by tight junctions between endothelial cells, which restricts the passage of serum factors from the periphery to the CNS and limits pathogen and immune cell entry40-43. Recent research has revealed that the CNS is not as isolated from the immune system as previously 4 thought. While the brain parenchyma relies on surveillance from microglia, the CNS-resident immune cells44, evidence has shown the presence of circulating peripheral immune cells (i.e., CD4+ and CD8+ T-cells) and lymphatic vessels45,46. Therefore, the CNS undergoes constant immune surveillance by both resident and peripheral immune cells, working together to detect and respond to threats while minimizing damage. However, under inflammatory conditions such as MS, this homeostasis between the immune system and the CNS is disrupted40,47. Activated lymphocytes can enter through one of three pathways, either the choroid plexus or the subarachnoid space from the bloodstream. Activated T-cells can also pass directly into the brain parenchyma through the BBB due to the increased expression of cell adhesion molecules in response to inflammation, which interact with integrin receptors on activated T-cells to allow their migration across the BBB40,47,48. These inflammatory infiltrates are primarily composed of CD8+ T-lymphocytes, which greatly outnumber CD4+ T-cells49-52. All T-lymphocytes express a T-cell receptor (TCR) complex consisting of a variable antigen binding TCR and CD3 signaling proteins. The TCR complex enables CD4+ and CD8+ T-cells to recognize antigens on major histocompatibility complex (MHC) class II and I molecules, respectively53. To activate and differentiate naïve T-lymphocytes, two signals presented by APCs are required: the engagement of the TCR with the peptide-MHC complex and the interaction of the coactivating receptor (CD28) with a co-activating ligand53. Activated CD4+ and CD8+ T-lymphocytes can both contribute directly to CNS damage via cell-to-cell death, i.e., the engagement of cell-death receptors of the Fas-FasL or TNF receptor pathway or release of lytic granules containing factors such as granzyme B and perforin53,54. These T-lymphocytes also contribute indirectly to CNS damage via the release of pro-inflammatory cytokines (for example, interferon-gamma (IFN-γ) and tumour necrosis factor-5 alpha (TNF-α)) amongst others, that further exacerbate inflammation by recruiting and activating B-cells, microglia, and macrophages resulting in a cascade of events such as the production of antibodies, reactive oxygen species (ROS; i.e., superoxide, hydroxide radicals, hydrogen peroxide & nitric oxide) and proteases. These events act in combination to create a highly toxic environment that further amplifies the damage, ultimately leading to neuronal/axonal loss and the formation of focal demyelinating lesions in the CNS white and grey matter55-57. Two main theories have developed over time to explain MS pathogenesis. In the “outside-in” hypothesis, external triggers, including environmental pathogens (bacterial or viral), are believed to activate immune cells in the periphery58. While the exact trigger is unknown, one proposed mechanism is that exposure to a myelin or neuronal protein peptide mimic epitope expressed on pathogens – bacteria or viruses - could prime pathogenic T-cells58. Other self-epitopes generated by damage, infection or cell apoptosis, such as fragments cleaved from the cytoskeleton, may also be cross-presented by APCs59. The activated cross-reactive and now auto-reactive myelin-specific T-cells can then cross the blood-brain barrier (BBB). Once in the CNS, these T-cells encounter self/myelin antigens presented on APCs in the CNS, such as microglia or memory B-cells, causing an immune cascade that leads to further myelin destruction, either through direct cell-cell death or through the release of inflammatory mediators60,61. Memory B-cells recruited in the CNS can also interact with T-cells in follicle-like structures within the CNS to cause clonal T-cell expansion6258,63,64. Support for this hypothesis comes from the experimental autoimmune encephalomyelitis (EAE) animal model, the most used preclinical model of MS63. EAE is induced by generating pathogenic autoreactive T-cell responses to myelin antigens, either by immunizing animals with myelin proteins with an immune adjuvant (active EAE) or by transferring primed/activated myelin-specific T-cells (passive EAE)55,65,66. 6 The competing hypothesis, the “inside-out” model of MS, suggests that the primary pathogenesis begins within the CNS, attributable to internal factors or insults such as oligodendrocyte injury or axonal damage that occur as primary cytodegenerative processes60,61. Some insults cause the release of cellular materials or neuroaxonal components, which can themselves alarm inflammatory responses against myelin debris or other CNS elements. This may also be accompanied by the release of antigens to the periphery, which would otherwise not be present. The presentation of these antigens by APCs to T-cells adds the additional element of autoreactive T-cells to the proposed pathogenesis, similar to that proposed for molecular mimicry. This hypothesis suggests that damage characterized by cytotoxicity, axonal loss and neurodegeneration starts early and is maintained/amplified throughout the disease60. 1.1.3 Inflammation and Neurodegeneration MS has previously been viewed as a “two-stage disease” consisting of an initial focal inflammatory phase followed by a secondary neurodegeneration phase50,67. A significant emphasis was placed on inflammation as it was believed to be the primary driver of accumulating disability and neurodegeneration in the progressive stages of MS68. However, emerging imaging and neuropathological evidence revealed that neurodegeneration and axonal injury are present at disease onset10,69. Axonal injury has been shown to correlate with the level of inflammation, and both inflammatory and neurodegenerative phases are observed throughout all stages of MS10,38,50. Additionally, clinical observations indicate that disease progression can occur independent of inflammation, suggesting a more prominent role for neurodegeneration in disease progression70. Proper axonal function depends on oligodendrocyte support for proper myelin sheath formation; therefore, myelin loss renders axons and neurons susceptible to damage/loss71,72. 7 Axonal or neuronal malfunction, driven by demyelination and inflammation, can then activate a myriad of self-destructive processes, including apoptosis (programmed cell death), autophagia and synaptic pruning via microglial activation73-75. Axonal spheroids, a common hallmark seen in early axonal damage, disrupt protein and organelle transport along the axon and are found mainly in actively demyelinating lesions76,77. These can be detected immunohistochemically by identifying amyloid precursor protein (APP) accumulations, as APP can no longer be transported and will accumulate in localized regions11,78-80. Additionally, the non-phosphorylated form of neurofilament heavy chain (NFH), which is typically phosphorylated, serves as a marker for axonal damage81. Damaged axons may then trigger one of two active axonal degeneration processes: anterograde (Wallerian) or retrograde degeneration, where axonal degeneration occurs distal or proximal to the site of injury, respectively, resulting in axonal dissolution82. Axonal damage can also occur independently of inflammation, as axonal abnormalities were observed in gray matter regions lacking inflammatory cell infiltrates83. Focal axonal degeneration (FAD), a multi-step process involving the formation of semi-stable axonal swellings in inflammatory environments, has also been observed early in MS and EAE84. It is characterized by subcellular changes, including swollen/shortened mitochondria with decreased membrane potential, lowered ATP levels, increased ROS generation, impaired calcium (Ca2+) buffering and the release of pro-apoptotic signals84,85. While some axons in FAD progress to axon fragmentation, others experience a full recovery, with axonal loss or survival correlating with high and low intra-axonal Ca2+ levels, respectively84,85. Ultimately, these results suggest that Ca2+ levels may predict neurodegeneration. Several key factors are believed to drive neurodegeneration in MS, including glutamate excitotoxicity, which has been suggested to link inflammatory and neurodegenerative processes 8 in MS86. As the CNS’ major excitatory and most abundant neurotransmitter, glutamate regulates critical functions, including memory, cognition, and mood86,87. Astrocytes, microglia and oligodendrocytes tightly regulate extracellular/synaptic levels of glutamate, as excess levels result in cellular toxicity/death87. These glial cells remove glutamate from the extracellular space via excitatory amino acid transporters, converting it to glutamine and transporting it to presynaptic neurons86,87. In MS, glial cell dysfunction, glutamate release caused by demyelination or from damaged cells or activated immune cells (such as CD4+ T-cells), and oxidative stress that impairs glutamate clearance mechanisms all contribute to glutamate accumulation in the extracellular space86-89. Glutamate accumulation triggers N-methyl-D-aspartate (NMDA) receptor activation, resulting in calcium influx into the cell90. Elevated calcium levels then activate secondary pathways contributing to neurodegeneration, including mitochondrial depolarization, caspase activation, ROS production and cellular toxicity91,92. Oxidative stress is also a major factor thought to promote MS-related neurodegeneration and can result from intrinsic or extrinsic sources, those being within cells or produced by cells and released into the local environment, respectively. Oxidative stress can occur due to numerous factors, including energy imbalance within neurons or oligodendrocytes, mitochondrial damage, microglia/macrophage and astrocyte activation, iron accumulation due to aging and meningeal inflammation37,93-96. ROS, such as superoxide radicals and hydrogen peroxide, are primarily produced by the mitochondria during oxidative phosphorylation for adenosine triphosphate (ATP) production97 and during cellular metabolism98,99. Resident microglia and recruited macrophages in the CNS express the necessary enzymes for ROS production (i.e., NADPH oxidase); however, under homeostatic conditions, ROS production is tightly regulated to prevent excess accumulation97. While ROS are typically minimized by a 9 balanced oxidant-antioxidant system100, inflammation and neurodegeneration disrupt this balance, leading to free radical formation and cellular damage97. In response to inflammation or infection, the microglia/macrophage populations in active MS lesions produce ROS, contributing to demyelination and free-radical-associated tissue injury101. Mitochondrial dysfunction in MS also causes excessive ROS production. This dysfunction leads to axonal damage as axons are highly susceptible to mitochondrial dynamics due to the inefficient distribution of mitochondria in axons94,102. Lastly, iron accumulation in the aging brain or from oligodendrocyte damage amplifies oxidative injury by increasing ROS release in microglia94. Importantly, oxidative stress also contributes to neurodegeneration by directly altering the function of voltage-gated Ca2+ channels, promoting the release of Ca2+ from the endoplasmic reticulum103. Histopathological evidence of oxidative injury in both actively enhancing lesions (RRMS) and non-active lesions (progressive MS) has suggested its presence throughout the disease course104,105. While axonal injury and degeneration occur early in the disease course, the brain’s plasticity and compensatory mechanisms buffer the onset of permanent clinical disability93,106,107. Over time, accumulating CNS damage exceeds these mechanisms, leading to brain atrophy and physical and cognitive impairments108. In imaging studies, damage is apparent as changes in brain parenchymal fraction and brain volume over time109. Therefore, while inflammation and neurodegeneration play key roles in disease onset and progression, their interaction and impact on clinical symptoms remain unclear, highlighting the importance of further understanding this relationship for therapeutic strategies67. 1.1.4 Current Therapeutic Strategies Health Canada has approved 19 disease-modifying therapies (DMTs) for MS110; however, none are curative. These treatments are primarily anti-inflammatory and 10 immunomodulatory, reducing inflammation and relapse rates111. As a result, most approved DMTs are effective only in RRMS and afford little benefit in the later progressive stages of MS111. DMTs are categorized into first- and second-line treatments. First-line DMTs in Canada include injectable options, such as glatiramer acetate (Copaxone®), interferon-β-1a (Avonex®) and interferon-β-1b (Betaseron®), as well as oral treatments such as teriflunomide (Aubagio®) and dimethyl fumarate (Tecfidera®)112. Phase-3 randomized controlled trials (RCT) have shown these first-line DMTs moderately reduced relapses (by 29-50%) and new lesions over two years compared to placebos, however, they were less effective at preventing disability progression113-116. Second-line DMTs, or higher efficacy treatments, also include a mixture of oral treatments such as fingolimod (Gilenya®) and cladribine (Mavenclad®), and monoclonal antibodies such as natalizumab (Tysabri®), ocrelizumab (Ocrevus®), and alemtuzumab (Lemtrada®)112. These have demonstrated significant efficacy in phase-3 RCTs. Both cladribine and fingolimod not only resulted in a higher relapse-free rate but also reduced the risk of disability progression over either a 3- or 24-month period and the number of brain lesion counts as detected by MRI when compared to placebos117,118. Ocrelizumab, in particular, has shown superior effectiveness compared to interferon β-1a in reducing disease activity rates and progression over 96 weeks119. However, while second-line DMTs are more effective than first-line treatments, they pose more significant health risks. They are typically reserved for individuals who do not respond adequately to first-line treatments or are prescribed early for those with highly active forms of MS characterized by rapid progression, frequent relapses and severe disability accumulation over a short period of time120. 11 Ocrelizumab is currently the only DMT approved by the Food and Drug Administration (FDA) and Health Canada for progressive MS112,121. The FDA has also approved four additional drugs for SPMS: cladribine, mitoxantrone, siponomod, and interferon β-1b122. However, these treatments are primarily recommended for individuals with “active” SPMS, characterized by ongoing relapses or new MRI lesions, and offer limited benefit for those with inactive SPMS122. Additionally, optimal results from these treatments are seen when administered within five years of the first demyelinating episode, highlighting the importance of early intervention in individuals with RRMS123,124. Given the evidence that axonal loss/injury are key drivers of clinical disability and disease progression in MS and that neurodegenerative mechanisms amplify immune-mediated damage, it has become evident that immunomodulatory drugs alone are insufficient to prevent or minimize CNS damage37,82,85,93-95. The inability of these DMTs to prevent neurodegeneration underscores the need for neuroprotective therapies that not only limit inflammation but also prevent axonal loss/injury and subsequent disease progression. Current research is exploring various neuroprotective strategies, including CNS-specific, regenerative biological processes (trophic factor support, axonal preservation and remyelination), prevention of oxidative stress/excitotoxicity induced axonal damage/loss, energy depletion/mitochondrial dysfunction, and apoptosis82. Both oxidative stress and glutamate excitotoxicity are at the root of neurodegenerative-associated damage in MS, prompting the investigation of antioxidants and glutamate receptor inhibitors as neuro or cytoprotectants82. Despite these therapies showing promising results in animal models of MS82,125, clinical trials in individuals with SPMS have yielded contradictory results, with some individuals experiencing transient symptom worsening or no significant improvements82,126-129. The selected drugs targeted axonal pathobiological features that play a 12 role in neurodegeneration in MS, including glutamate excitotoxicity, calcium overload and providing sufficient energy for neurons. However, these findings suggest that solely targeting these aspects of axonal pathobiology is insufficient to prevent axonal loss and neurodegeneration, as inflammation and remyelination are also implicated in disease progression and must also be targeted. Further research is required to determine their effectiveness when combined with existing immunomodulators or other neuroprotective treatments. Given the central role of demyelination in MS, promoting remyelination has become a key focus for enhancing axonal and neuronal survival. Remyelination, where a new myelin sheath forms around a recently demyelinated axon, is essential for maintaining axonal integrity, restoring cellular function, and reducing the risk of axonal and neuronal degeneration130. This process depends on several factors, including the activation and recruitment of oligodendrocyte precursor cells (OPCs) to the damaged site, followed by their differentiation into mature oligodendrocytes capable of myelinating axons130-132. Additionally, microglia and macrophages play an important role in clearing inhibitory myelin debris that can impede OPC progression and remyelination133-135. However, remyelinated axons typically have thinner myelin sheaths with shorter internodal distances compared to the original axon136,137. Given the complexity of the remyelination process, even minor disruptions in OPC recruitment, differentiation or debris clearance could lead to remyelination failure. For example, post-mortem forebrain tissue histopathological assessments revealed that two-thirds of individuals remyelinated less than a quarter of their lesions138. Although several drugs to enhance endogenous remyelination are being tested in clinical trials with some efficacy, a deeper understanding of remyelination in humans is essential for optimizing treatment success82,139. 13 In addition to remyelination, trophic factor signaling offers a promising neuroprotective strategy by supporting neuronal health and survival128,140,141. Brain-derived neurotrophic factor (BDNF), a member of the neurotrophin family, has been implicated in MS-related neuroinflammation and neuroprotection, increasing oligodendrocyte lineage cells and myelin proteins, rescuing injured neurons, and promoting axonal outgrowth142-144. While neurons in the CNS are a major source of Bdnf145, it is also expressed by immune cells, microglia, and astrocytes146,147. The release of BDNF by activated lymphocytes, monocytes, and astrocytes in inflammatory lesions supports its role in myelin repair146. Actively demyelinating areas had a higher number of BDNF+ immune cells compared to inactive regions within the same lesion148, suggesting a link between BDNF and active inflammation. Further support for this link between BDNF and myelin repair comes from studies on Bdnf-knockout mice, where deficits in myelin proteins and OPC proliferation are observed149,150. Deficiencies in immune-cell Bdnf resulted in enhanced axonal loss and disability progression in chronic EAE, whereas direct supplementation of exogenous Bdnf to the lesion site rescued this effect151. Individuals living with RRMS152,153 and SPMS154 displayed reduced serum BDNF levels compared to their healthy counterparts152-154. Lastly, two currently approved DMTs, glatiramer acetate and fingolimod, have been associated with increased levels of BDNF in humans and rodents, respectively152,155. Ultimately, trophic factors such as BDNF remain valuable therapeutic targets as they are key modulators of cellular processes promoting neuronal survival. 1.2 Neuronal PAS-Domain Containing Protein 4 (NPAS4) 1.2.1 Overview Synaptic inputs can drive gene expression changes via transduction molecules that 14 activate specific transcription factors156. Among these are the basic-helix-loop-helix PER-ARNT-SIM (bHLH-PAS) proteins, a family of transcriptional regulators that play vital roles in numerous physiological and developmental events, making up the second most populated transcription factor family in the human genome157,158. These transcription factors possess several conserved functional domains: an N-terminal DNA-binding motif, a dimerization motif known as the PAS-homology domain, and a C-terminal domain for activating or repressing target gene transcription159,160. These three motifs led to the identification of neuronal PAS-domain-containing protein 4 (Npas4), initially discovered while searching for relatives of a bHLH-PAS transcriptional repressor (Sim2)159. Initially thought to be localized exclusively to neuronal cells, NPAS4 was suggested to have a role in the transcriptional regulation of neural genes159. Further characterization of Npas4 highlighted its function as an inducible immediate early gene (IEG) activated within minutes of stimulation to regulate the excitatory/inhibitory balance in the CNS156,159,161. The bHLH-PAS proteins are further divided into two distinct classes: class I, which includes specialized transcription factors activated in response to specific stimuli, and class II proteins, which are ubiquitously expressed160. Like other class I members, NPAS4 functions through heterodimerization with class II members, notably aryl hydrocarbon receptor nuclear translocator 1 and 2 (ARNT or ARNT2), to form a functional transcriptional complex that enables spatial and temporal gene activation or repression160,162. The two ARNT paralogues share similarities in their highly conserved protein sequences, with over 80% amino acid sequence similarities in the bHLH and PAS domains163. While ARNT is universally expressed throughout the body, ARNT2 is localized to the brain and kidneys159,163. Given the localized expression of ARNT2 and NPAS4 in the brain, ARNT2 is believed to be NPAS4’s major 15 binding partner159,163. However, recent evidence in hippocampal neurons has suggested that although NPAS4 has a stronger affinity for ARNT2, both can form functional stimulus-dependent heterodimers, leading to differential DNA binding and gene regulation162,164. 1.2.2 Expression Patterns NPAS4 expression has primarily been localized to CNS neuronal cells. Conserved brain-specific expression has been observed in several mammals, including rodents (i.e., mice and rats)165-167 and humans159, as well as in lower vertebrates, including zebrafish and Xenopus168. Unlike ubiquitously expressed class II bHLH-PAS proteins, basal levels of NPAS4 in the rodent brain are minimal. Enriched expression has been observed in the limbic system165, a group of subcortical cultures (i.e., hippocampus, amygdala, fornix and the cingulate gyrus) involved in learning/memory, emotional regulation, and olfaction169-171. Expression is highest in the hippocampus, specifically in the pyramidal cell layers of the CA1, 2 and 3 regions and the granule cell layer of the dentate gyrus165,166,172,173. NPAS4 has also been observed in the basal ganglia (i.e., striatum), olfactory bulb, cerebellum, hypothalamus and the cerebral cortex, with the highest expression observed in cortical layers III and V165,166,172,174-176. Additionally, while Npas4 RNA transcripts have been identified outside of the CNS, including endocrine organs and tissues such as the pancreas and adrenal glands, and the thymus177, protein expression outside of the CNS has been described only in the adrenal glands, endothelial cells, and pancreatic beta cells178. As mentioned previously, Npas4 is nuclear-localized, however, immunofluorescence studies in rat primary cortical neurons have detected NPAS4 protein in the soma, neurites and synapses166, suggesting a potential role for NPAS4 in neuronal activity regulation and synaptic plasticity. 16 At the cellular level, NPAS4 is mainly expressed by excitatory neurons161 but has also been observed in rodent inhibitory neurons174,179,180, with distinct functions observed in each subtype. Dissociated primary cultures of mouse cortical neurons showed negligible Npas4 RNA or protein, but each was upregulated following KCl administration161,181. Similar to other IEGs, Npas4 is rapidly and transiently activated, with maximal mRNA and protein expression occurring within 0.5-1 hour and 1-2 hours, respectively, in depolarized cultured neurons161.181 However, unlike other IEGs, Npas4 is selectively induced by neuronal activity (via Ca2+-dependent activation) and not by growth factors, neurotrophins or kinase pathways161,181. In rat cortical cultures, Npas4 transcript and protein levels were negligible1, while application of transcription and translation inhibitors also maintained Npas4 mRNA and protein at negligible following exposure to an excitatory stimulus (Quandt lab, unpublished), suggesting that de novo transcription and translation may be required for NPAS4 expression in these cells. In vivo, Npas4 induction occurs more rapidly, with mRNA and protein levels increasing within 5- and 30-minutes post-stimulation, respectively161,181. 1.2.3 Functional Roles Due to its activity-dependent nature, NPAS4 has an extensive influence on brain function and in maintaining homeostasis. Many studies have aimed to characterize its major functional roles that link neuronal activity to adaptive changes, including activity-dependent gene expression, maintenance of the excitation-inhibition balance, behavioural and cognitive functions underlying synaptic plasticity, and the brain’s response to stressful environments. One essential function of NPAS4 is the regulation of genes that control inhibitory synapse development, thereby maintaining the excitatory-inhibitory (EI) balance in the CNS161,182. While NPAS4 is expressed in both inhibitory and excitatory neurons, it activates a 17 distinct set of genes in each type179. Npas4-RNAi knockdown in cultured hippocampal neurons decreased the number of inhibitory synapses on neurons, while overexpression had the opposite effect161. NPAS4’s role in the activity-dependent upregulation of inhibitory synapses was also shown in vivo183. Therefore, NPAS4 modulates inhibition levels by increasing inhibition on excitatory neurons while increasing excitation on inhibitory neurons, maintaining a homeostatic response by balancing activity across hippocampal, cortical, limbic and thalamocortical circuits179. Given that NPAS4 can transduce synaptic changes into longer-lasting changes (i.e., gene transcription)184, it also plays a key role in learning, memory, and synaptic plasticity156. For example, following sensory experience, Npas4 gene activation inhibited Mdm2, facilitating dendritic spine formation in olfactory bulb granule cells180. During contextual memory formation, Npas4 was shown to reduce the number of synaptic connections between hippocampal mossy fibers and CA3 pyramidal neurons185. Deletion of Npas4 not only prevented contextual memory formation but also prevented the synaptic changes previously observed185. Furthermore, Npas4 plays an important role in long-term memory formation in the hippocampus and amygdala, as its deletion in the CA3 region prevented long-term contextual memory formation, with restoration upon Npas4 re-expression181. These findings emphasize NPAS4’s role as a key regulator of experience-dependent synaptic plasticity. Lastly, NPAS4 is a potential therapeutic target of interest for CNS diseases due to its role in stress protection. Rodents exposed to stressors such as conditioned fear186,187, sleep deprivation188, and anxiogenic environments189 displayed reduced NPAS4 levels. Altered NPAS4 expression was also evident in impaired neurogenesis and hippocampus-dependent fear memory in mice190, contextual fear exposure in rats191 and early maternal separation leading to 18 impaired social behavior in adult mice192. Lastly, prenatal stress in Npas4 knockout (KO) mice resulted in impaired social recognition193. Overall, NPAS4 plays a key role in long-term synaptic plasticity across multiple brain circuits involved in learning and memory, sensory processing, cognitive functions, emotional regulation, and behavioural responses. Its ability to confer protection against numerous CNS insults via modulation of the EI balance makes it a key neuroprotective target. 1.2.4 Neuroprotective Roles Given the above-listed functional roles, NPAS4 has been shown to contribute to neuronal health and survival. Due to its role in modulating the EI balance, Npas4 was shown to provide neuroprotection in disorders with disrupted EI balance, as evidenced by its upregulation following focal cerebral ischemia166,174, kainate-induced hippocampal injury159, and epileptic seizures194. Following seizure activity, Npas4 mRNA and protein rapidly increased in the hippocampus within 0.5-1 hour and 2 hours, respectively, returning to baseline within 2 and 4 hours, respectively194. Npas4-deficient mice (Npas4 KO) experienced accelerated seizure initiation, suggesting that NPAS4 functions as a switch to help suppress seizure initiation and recurrence194. Furthermore, adeno-associated virus (AAV)-mediated delivery of Npas4 to the dorsal hippocampus helped protect against seizure-induced cell death, with over 92% of cells surviving compared to the control contralateral hemisphere195. Conversely, Npas4 KO mice were more susceptible to hippocampal neuronal damage following kainite-induced seizure activity, which resulted in death more than one-third of the time159. These mice also displayed larger lesion sizes as well as signs of increased neurodegeneration, such as gliosis, increased proinflammatory cytokine levels (i.e., interleukin (IL)-6 and tumour necrosis alpha (TNFα)) and a switch from apoptotic to necrotic cell death196. Interestingly, even in the absence of seizure 19 activity, Npas4 KO mice exhibited hippocampal and cortical neurodegeneration after three months of age, with over 50% mortality by 16 months of age172,197. NPAS4’s neuroprotective functions have also been explored in in vitro studies. Neuronal Npas4 expression prior to potassium cyanide treatment167, growth factor withdrawal or treatment with an apoptotic inducer (staurosporine)195 prevented cell death. This neuroprotective effect was reversed following RNAi-mediated knockdown of Npas4167. Similarly, Npas4 knockdown increased cell death susceptibility in mouse cortical neurons exposed to ischemic-like conditions, such as hypoxia and hypoglycemia196. Both in vivo and in vitro findings highlight NPAS4’s role in neuroprotection, prompting research into its involvement in several neurodegenerative diseases. In Alzheimer’s disease (AD), Npas4 expression is reduced as the disease progresses, notably during lesion development in the transentorhinal/entorhinal cortex, which is associated with cognitive and memory-related impairments198. Furthermore, APP-deficient neurons displayed reduced Npas4 expression and increased GABA production199, suggesting a link between NPAS4 and AD, as NPAS4 may be a relevant downstream target in the synaptic functions regulated by APP199. Overall, NPAS4’s neuroprotective functions are important for maintaining neuronal health and functions, specifically in the case of increased activity, stress, or potential injury/insult. Although its neuroprotective functions are well-documented in contexts of altered EI balance and cellular stress or injury, any role or function in chronic inflammatory neurodegenerative diseases of the CNS remains unknown. 1.2.5 ARNT2: NPAS4’s Primary Binding Partner As NPAS4 relies on heterodimerization with class II bHLH-PAS proteins to regulate 20 target gene expression, its binding partners had to be identified. Through an examination of protein-protein interactions in neuroblastoma cells, NPAS4 was shown to preferentially bind with the ARNT subclass of bHLH-PAS proteins, specifically ARNT2159,172,200 (Figure 1.1). ARNT2 is highly conserved across mammals, with expression observed in rodents, humans, Xenopus and zebrafish201-207. While predominantly localized to the CNS, Arnt2 is also expressed in the kidneys, urinary tract, and thymus163,201,208,209. Within the CNS, Arnt2 gene transcripts are highly and preferentially expressed within the hippocampus210, with an expression gradient seen from the CA2 region (highest) to the CA1 region (lower)210,211. Expression within the CNS has also been observed in the cerebral cortex, the striatum, pons, dorsal midbrain, thalamus and cerebellum201,202,204,207. However, unlike Npas4, Arnt2 mRNA and protein expression have also been observed in glial cells, including astrocytes and oligodendrocytes1,212. Given its ubiquitous expression, ARNT2 has also been implicated in cell health and survival. ARNT2 was shown to play an important role in overcoming hypoxia by binding to hypoxia-inducible factor 1α (HIF-1α) and regulating genes involved in cellular metabolism, glucose transport and angiogenesis213. Furthermore, lack of Arnt2 expression in mice results in perinatal death due to improper hypothalamic development214. Additionally, a major decline in Arnt2 expression preceding cell death was observed due to ischemia or H2O2-mediated oxidative stress in neuroblastoma cells209. In the absence of neuronal activity, ARNT2 is thought to play a role in the activity-dependent upregulation of genes by binding to nuclear receptor co-repressor 2 (NCoR2), which results in the repression of inducible genes such as Npas4215 (Figure 1.1A). However, in response to excitatory stimuli, rapid calcium influx drives Npas4 transcription, which Arnt2 then recruits to the proper activity-dependent regulatory elements to regulate gene transcription and promote 21 cell health and survival (Figure 1.1B). To form a stable heterodimer, NPAS4 and ARNT2 interact through their PAS domains, and the bHLH DNA binding motif allows the complex to bind to the promoter regions of target genes164,216. Once bound, the complex recruits co-activators and transcriptional machinery components to initiate the transcription of numerous genes involved in neuronal activity, health, and survival. Most importantly, this complex regulates the expression of several downstream genes, including activity-dependent Bdnf mRNA levels. BDNF is an important neurotrophic factor in the CNS, with roles in neuronal differentiation217, maturation218, and survival and growth219, significantly impacting synaptic plasticity underlying learning, memory and neurogenesis220,221. Furthermore, BDNF provides neuroprotection under ischemic, hypoglycemic, neurotoxic or excitotoxic conditions222,223, and its deficiency in mice resulted in a shortened life span224. The link between NPAS4 and BDNF was demonstrated via RNAi interference of Npas4 in cultures, which led to a 50% decrease in depolarization-induced Bdnf expression compared to control cultures161, while overexpression of Npas4 increased Bdnf expression225. These results suggest that NPAS4 mediates neuronal activity-dependent upregulation of BDNF. As mentioned in section 1.1.4, BDNF has become a target for enhanced neuroprotection in MS and many other neurodegenerative diseases, including Parkinson’s, Alzheimer’s and Huntington’s disease. 22 Figure 1.1 ARNT2’s role in activity-dependent upregulation of genes in the CNS. (A) In the absence of neuronal activity, ARNT2 recruits and binds nuclear receptor co-repressor 2 (NCoR2) to repress the transcription of inducible genes such as Npas4, helping to maintain low basal levels of these inducible genes215. (B) Following neuronal activity, calcium (Ca2+) influx into the cell induces neuronal PAS-domain containing protein 4 (Npas4) transcription. NPAS4 is then recruited by aryl hydrocarbon receptor nuclear translocator 2 (ARNT2) to activity-dependent regulatory elements (PasRE) of genes to upregulate transcription of key neuroprotective factors, including brain-derived neurotrophic factor (Bdnf). Infiltrating immune cells and oxidative stress may be an additional driver of Ca2+ influx into the cell that induces NPAS4 expression. Created in Biorender.com. 1.2.6 NPAS4/ARNT2: A Role in Chronic Inflammatory Neurodegenerative Settings Relevant to MS Given the growing evidence linking NPAS4 and ARNT2 to neuronal health and survival, our lab sought to characterize their expression in chronic inflammatory conditions modelling those in MS. This interest stemmed from earlier findings that examined TEMPOL, an antioxidant with anti-inflammatory and neuroprotective properties226,227, in mice with chronic EAE. Gene expression microarray analyses showed that before the expected onset of disease symptoms, TEMPOL-fed mice with lower disease incidence and severity had two-fold lower A B 23 Npas4 expression in the brain and spinal cord compared to non-TEMPOL-fed mice (Quandt lab, unpublished data). They next explored Npas4 and Arnt2 expression levels in the CNS over the course of EAE. qPCR mRNA analysis of tissues obtained at pre-onset (day 7), onset (day 14), peak (day 18), and recovery (day 29) showed upregulation of both Npas4 and Arnt2 mRNA in the brain (Quandt lab, unpublished) and the spinal cord1 pre-onset when immune cell infiltration is believed to be in its earliest stages228 (Figure 1.1B and C). This was followed by a decrease in Npas4 and Arnt2 levels during onset and peak disease in the spinal cord, with levels rising again during recovery1. Spinal cord Bdnf levels mirrored these expression patterns (Figure 1.2C). EAE mice also experienced a 20-50% reduction in NeuN+ARNT2+ cells compared to healthy littermates, with levels inversely correlating to the degree of immune infiltration and disability1. Interestingly, ARNT2 expression was also reported in GFAP+ astrocytes and Olig2+ oligodendrocytes, with a marked decrease of ARNT2+ oligodendrocytes at peak EAE212. The changes in CNS Npas4 and Arnt2 levels throughout EAE at points associated with the presence of immune cells in the CNS suggest that immune cells and inflammatory mediators may be responsible for changes in NPAS4 and ARNT2 levels. To examine this, rat cortical neuronal cultures were exposed to H2O2 as a proxy for oxidative stress. At lower concentrations, neuronal ARNT2 levels increased and returned to baseline over time1. In contrast, higher concentrations resulted in decreased ARNT2 levels that preceded cell death1. Overall, these findings suggest potential roles for both NPAS4 and ARNT2 in the pathobiology of MS and its animal model, as both may be regulated by factors involved in neuroaxonal degeneration, such as oxidative stress. However, while much is known about the independent and combined protective roles of NPAS4 and ARNT2 in acute experimental or developmental settings, their expression patterns and functional roles in CNS chronic 24 inflammatory neurodegenerative disease remain to be fully characterized, warranting further investigation. Figure 2.2 Expression changes in Arnt2, Npas4 and Bdnf RNA throughout the course of active EAE. (A) Brain and spinal cord tissues were collected from 5-8 mice at preclinical, onset, peak and recovery times to be analyzed by qPCR. Arrows represent time points at which spinal cord samples were taken. (B) qPCR analysis of brain tissues showing Npas4 and Arnt2 message levels normalized to β-actin (Quandt lab, unpublished data). The dotted red line represents sham-immunized (CFA) baseline levels. Mann-Whitney comparison to healthy/sham-immunized mice, *p<0.05. (C) qPCR analysis of spinal cord tissues showing Npas4, Arnt2 and Bdnf message levels normalized to β-actin. One-way ANOVA followed by Dunnett’s multiple comparison test against CFA/sham immunized mice, **p<0.01, ***p < 0.001, ****p < 0.0001 (Adapted from Rahim et al., 2018). Each point represents values obtained from one mouse, with bars representing mean +/- SD. 25 1.3 Thesis Objectives and Hypothesis 1.3.1 Aims Given the neuroprotective roles of NPAS4 and ARNT2 and previous findings linking these factors to chronic inflammation in MS animal models, I hypothesize that inflammatory mediators and cells associated with pathological processes in a preclinical model of MS may alter NPAS4 and ARNT2 protein expression. To better understand the role of NPAS4 and ARNT2 in chronic inflammatory neurodegenerative environments relevant to MS pathogenesis, this work was structured to address the following aims: Aim 1: Characterize NPAS4 and ARNT2 protein expression in resting or steady-state conditions in primary neuron-enriched cortical cultures. Aim 2: Examine the effects of soluble inflammatory mediators and inflammatory cells implicated in MS or EAE pathogenesis on NPAS4 and ARNT2 protein expression in primary neuron-enriched cortical cultures. 1.3.2 Clinical Significance Despite significant advances in successful treatments for individuals living with MS, the majority of treatments are most effective for individuals living with RRMS and offer limited benefit in the later progressive stages marked by smouldering inflammation and ongoing neurodegeneration and axonal loss. This highlights the dire need for approaches that protect neuro-axonal components to promote recovery while limiting damage, one of the primary goals in MS research. This project aims to characterize the role of NPAS4 and ARNT2, two neuroprotective molecules associated with neuronal and axonal survival via the activation of downstream neuroprotective factors, in inflammatory degenerative settings relevant to MS pathogenesis. Ultimately, understanding how neuroinflammation affects NPAS4 and ARNT2 26 expression levels and mechanisms associated with neuroprotection relevant to MS may identify these two proteins as targets with the potential to limit neuronal damage in inflammatory settings. 27 Chapter 2: Materials and Methods 2.1 Mice Male and female C57BL/6NCrl mice (strain code 027) were purchased from Charles River (Wilmington, USA) and housed and bred at the Preclinical Discovery Center (PDC) at the Djavad Mowafaghian Center for Brain Health at the University of British Columbia (UBC) (Vancouver, Canada). Mice were exposed to a 12h:12h light cycle at a constant room temperature with free access to food and water. All animals were bred in accordance with protocols A19-0220 and A23-0173, while all experimental procedures were carried out in accordance with protocols A18-0361 and A22-0283, approved by the Animal Care Committee (ACC) at UBC. 2.2 In Vitro Cultures 2.2.1 Murine Primary Neuron-Enriched Cortical Cultures To increase the chances of successful timed pregnancies, estrus was induced in female C57BL/6NCrl mice by exposing them to male bedding for two days. One or two sexually mature females (8 weeks or older) were paired with a sexually mature male (8 weeks or older) for 48 hours and, followed for euthanasia and isolation of embryonic day 15.5 to 17.5 (E15.5-17.5) embryos. As previously described, cortices from E15.5-E17.5 male and female embryos were pooled and used to establish primary neuron-enriched cortical cultures229 (Figure 2.1). Cortices (freed of meninges, hippocampus and the striatum) were washed with Hank’s Balanced salt solution (HBSS) buffer, composed of HBSS (Gibco, Waltham, US, cat #25200056) supplemented with 0.01M HEPES (Gibco, cat #1563080), 6mg/ml glucose (Sigma-Aldrich, St-Louis, US, cat #G7528) in Dulbecco’s phosphate buffered saline (DPBS; Gibco, cat #14190144) and 1% Penicillin-Streptomycin (100 units/ml penicillin/100µg/ml streptomycin; Gibco, cat 28 #15140-122). Cortices were incubated in 0.25% Trypsin-EDTA (Gibco, cat #25200056) and 0.2mg/ml DNAse (Sigma-Aldrich, cat #DN25) for 15 minutes at 37°C, and washed again with 37°C HBSS buffer. The cortices were dissociated into a single cell suspension in 5ml HBSS buffer and seeded onto glass bottom 96-well plates (Cellvis, Mountain View, US, cat #P96-1.5H-N) coated with 50µg/ml poly-d-lysine (Gibco, cat #A38904-01). Cells were plated at 50,000 cells/cm2 in neuronal plating media (DMEM (Invitrogen, Waltham, US, cat #11430-030) supplemented with 10% fetal bovine serum (FBS; PAA Laboratories, Zandt, DE, cat #A15-701), 4.5mg/ml glucose, 1mM sodium pyruvate (Invitrogen, cat #11360-070), 2mM glutaMAX (Invitrogen, cat #35050-161), and 1% Penicillin-Streptomycin). After 4 hours, the media was replaced with neuronal maintenance media (NMM) (Neurobasal medium (NBM, Invitrogen, cat #21103-049) supplemented with 2mM glutaMAX, 1% Penicillin-Streptomycin and 2% B-27 supplement (Invitrogen, cat #17504-044)). Cells were incubated at 37°C with 5% CO2, with 50% media changes every 2-3 days until days in vitro (DIV) 9-11 (Figure 2.1). 2.2.2 CD4+ T-Cell Isolation Spleens from 12-16-week-old C57BL/6NCrl male mice were aseptically isolated, manually dissociated in HBSS and passed through a 40µm filter to create a single-cell suspension. Red blood cells were lysed using ACK Lysing Buffer (Gibco, cat #A1049201), followed by two washes in HBSS. Naïve CD4+ T-cells were then purified via negative selection using the Naïve Mouse CD4+ T-cell Isolation Kit (Miltenyi Biotec, Bergisch Gladbach, DE, cat #130-104-453) with MACS LS columns (Miltenyi Biotec, cat #130-042-401). Cells were either used immediately as resting or following activation by seeding for 20 hours at 250,000 cells/well on 96-well flat bottom plates (Corning, Corning, US, cat #CLS3585) with 4µg/ml plate-bound anti-CD3 (Biocon, Bengaluru, IN, clone 145-2c11) and 2µg/ml soluble anti-CD28 (clone 37.51; 29 eBioscience, San Diego, US, cat #16-0281-85) in T-cell media (TCM: RPMI (Gibco, cat #21870-076) supplemented with 5% FBS, 1% Penicillin-Streptomycin, 2mM L-glutamine (Sigma-Aldrich, cat #G7513), and 0.1mM non-essential amino acids (NEAA; Gibco, cat #11140050)) at 37°C in a humidified incubator supplemented with 5% CO2. Cells were harvested and washed twice in DPBS. 2.2.3 Murine Immortalized Brain Endothelial Cell Line (bEnd.3) Murine immortalized brain endothelial cells (bEnd.3, ATCC, Manassas, US, CRL-2299) were plated at 15,000 cells/cm2 in DMEM supplemented with 10% FBS, 1% Penicillin-Streptomycin, 1mM Sodium Pyruvate, 0.1mM MEM NEAA and 4mM L-glutamine. Cells were incubated for 2 days at 37°C with 5% CO2 on glass bottom plates (Cellvis) to serve as an immunocytochemical staining negative control. 2.2.4 Soluble Inflammatory Mediators Freshly prepared hydrogen peroxide (H2O2, Fisher Scientific, Waltham, US, cat #H325-500) in DPBS was added to neuronal cultures in neuronal maintenance media at final concentrations of 12.5, 25, 50 and 100 M and glutamate (Sigma-Aldrich, cat #G5667) was applied in DPBS to 1, 3, 10 and 30 M final concentration for 1, 3, 5, 7 and 9 hours (Figure 2.1). Potassium chloride (25mM KCl, Thermo Fisher Scientific, cat #A11662) in DPBS was used as an excitatory stimulus and positive control for NPAS4 protein induction and antibody staining. 2.3 Immunocytochemistry 2.3.1 Staining Protocol Following treatments, media was aspirated, and cells were fixed with 10% neutral buffered formalin (Fisher Scientific, cat #SF100-4) for 20 minutes at room temperature. Cells 30 were then washed twice with DPBS and stored at 4°C until stained. All staining and wash steps were performed on an orbital shaker (Model 13687700, Fisher Scientific) at 100 rpm. Cells were washed for 5 minutes with Tris-Buffered Saline with 0.5% Tween (TBST; Tris-Buffered saline (TBS): 200mM trizma base (Fisher Scientific, cat #BP152-500) pH adjusted to 7.6 and Tween-20: Fisher Scientific, cat #AC233360010), permeabilized for 20 minutes at room temperature with 0.1% Triton-X-100 (Fisher Scientific, cat #BP151100) in TBST and blocked for 30 minutes at room temperature with 10% normal donkey serum (Sigma Aldrich, cat #D9663) in TBST. Primary antibodies (Appendix) were applied in TBST with 2% normal donkey serum and incubated overnight at 4C. Cells were then washed three times for 5 minutes with TBST followed by a 1-hour secondary antibody (Appendix) incubation at 4C. After two additional 5-minute TBST washes, cells were incubated with 2µg/ml Hoechst-33342 (Invitrogen, cat #H3570) in TBST for 10 minutes for nuclear staining before a final wash in TBS. 2.3.2 Image Acquisition and Immunocytochemistry Analysis All cell visualization and analysis steps were performed using the Zeiss software Zen Blue (Version 3.4, Oberkochen, DE). Cells were imaged with a Zeiss Axio Vert 200 Inverted Fluorescence Microscope (Oberkochen, DE) using two filters: the 90 HE DAPI/GFP/Cy3/Cy5 filter for the 350, 488 and 647 channels, and the 31 Alexa Fluor 568 filter for the 568 channel. All images were captured at 20x magnification (0.8 numerical aperture) with an Axiocam 506 monochrome camera, with similar light intensity and exposure times applied to images from the same experimental set. Four 3x3 images were taken for each well, with 2-3 technical replicates across all experiments (i.e., 8-12 3x3 images total per treatment), typically capturing 800-1000 cells per well. To minimize variations in background fluorescence, a background subtraction on 31 all fluorescence channels was applied using a “rolling ball” algorithm at a radius of 220 across all fields of view. Neuronal ARNT2 and NPAS4 expression were examined by first selecting all Hoechst-positive nuclei. A NeuN threshold was then applied by averaging the nuclear NeuN mean fluorescent intensity (MFI) of our negative staining control population, bEnd.3 cells, plus two standard deviations to select for Hoechst+NeuN+ (neuronal) cells. A nuclear area cutoff was also applied, and only cells with nuclei between 50 and 200µm2 were chosen for analysis due to the likelihood of cells below and above this range being pyknotic nuclei/cell debris or cell clusters, respectively. ARNT2 and NPAS4 MFI were calculated as the average intensity per pixel over Hoechst+NeuN+ neuronal cells (Figure 2.1). MFI values of all cells were then averaged per well, and the 2-3 technical replicates were averaged to provide one value per treatment per experiment. These values were then normalized to the control treatment by dividing the average MFI per treatment by the average MFI of the PBS-treated wells (i.e., 0µM) at each time point. To quantify the percentage of ARNT2- and NPAS4-expressing cells, bEnd.3 cells (confirmed by qPCR as ARNT2 and NPAS4 negative) were used to set a positive ARNT2 and NPAS4 threshold (the average ARNT2 and NPAS4 MFI in bEnd.3 cells plus 1 and 3 standard deviations, respectively) (Figure 2.1). Cells with nuclear MFI below this threshold were considered negative and cells above were considered positive. ARNT2 and NPAS4 expression were further characterized into a categorical distribution of NPAS4 and ARNT2 staining intensity in Hoechst+NeuN+ neurons (Figure 2.1). Each cell’s MFI was divided by the bEnd.3-derived positivity threshold value to obtain a fold change from the threshold. Cells with a normalized value below 1, indicating an MFI value below the threshold, were classified as negative (Neg). Cells with a normalized value above 1 were further 32 classified into four other categories based on a fold-change from the threshold: values 1 but 2 (Low), 2 but 3 (Med), 3 but 4 (High) or 4 (Very high), with the highest category determined using the positive staining and treatment control, KCl. To determine culture purity, the percentage of Hoechst+NeuN- cells was calculated by dividing the number of Hoechst+NeuN- cells by the total Hoechst+ cells. The identity of some of the Hoechst+NeuN- cells was determined using a purity stain with primary antibodies to examine the presence of GFAP+ astrocytes and IBA-1+ microglia (Appendix). As a proxy for changes in cell viability, changes in the number of neuronal cells were examined by calculating the average Hoechst+NeuN+ cells from each well and averaging across 2-3 technical replicates to obtain one value per treatment per experiment. Values were then normalized by dividing them by the average Hoechst+NeuN+ cells across all PBS (“0 µM”) time points. Figure 2.1 Flowchart depicting experimental protocol for culturing primary cortical cells, application of inflammatory mediators and quantification of NPAS4 and ARNT2 protein expression. Representative images showing NPAS4 expression in NeuN+ neuronal cells depicting the different methods of quantifying NPAS4 and ARNT2 protein expression. The dotted line represents the nucleus as delineated by the Hoechst staining, while the arrows represent NPAS4-positive cells. Created in Biorender.com 33 2.4 Statistical Analysis All statistical analyses and graph visualizations were performed using GraphPad Prism (version 10.3.0, La Jolla, CA). A Shapiro-Wilk test was performed on all data sets to assess normality. A paired-ratio t-test with a Bonferroni correction was conducted on select doses and time points with the greatest effect to evaluate differences between dose and time curves. An area under the curve (AUC) analysis was also conducted, and data were compared using a Kruskal-Wallis analysis of variance (ANOVA) on ranks test with a Dunn’s multiple comparisons post-hoc test either to the 0µM dose or across all time points. Follow-up Mann-Whitney Rank T-test compared to the 0µM dose was carried out on doses with the greatest effect. To evaluate changes in NeuN cell counts, a repeated measures two-way ANOVA with a Dunnet’s multiple comparisons post-hoc test was conducted comparing NeuN cell counts at each treatment dose to the PBS treated control at the same exposure time. Data are reported as median +/- range or interquartile range (IQR) or mean +/- standard deviation (SD). 34 Chapter 3: Effects of Inflammatory Mediators and Cells on NPAS4 and ARNT2 Expression 3.1 Characterization of Primary Cortical Neuron-Enriched Cultures We used primary neuron-enriched cortical cultures to examine the effects of inflammatory mediators on NPAS4 expression in an in vitro setting. Over time, neurons became progressively complex with increased length of the microtubule-associated protein 2 (MAP2)-staining axodendritic processes and number of cell-cell connections (i.e., neuron-neuron and neuron-glia) (Figure 3.1A). By DIV10, the majority of cells exhibited the characteristic morphological states of neurons and were characterized by round, oval or pyramidal soma (Figure 3.1B and C). Most neurons had formed a network of connections with neighbouring neurons, with complex axonal or dendritic processes extending from the soma of one cell to another, enabling cell-cell communication (Figure 3.1C). An examination of all Hoechst+ nuclei revealed that 69  6% of all nuclei were NeuN+ neuronal cells (n=9). The remaining NeuN- cells were typically GFAP+ astrocytes, star-shaped cells characterized by ovoid nuclei and long processes extending towards neuronal cells, as shown by our purity stains (Figure 3.1D and E). The remaining NeuN-GFAP- cells were not characterized and could have been occasional oligodendrocytes or microglia (data not shown). 35 Figure 3.1 Characterization of primary neuron-enriched cortical cultures. Immunocytochemistry (A, B and D) and phase (C and E) images of mouse primary cortical neuron-enriched cultures. (A) Increasing complexity of cultures over time as shown by microtubule-associated protein 2 (MAP2) staining, with increases in the length and number of connections. (B and C) DIV10 healthy adult neurons were characterized by large pyramidal or round cell bodies with numerous extending processes and increasing complexity. (D and E) Purity analysis revealed 69±6% of Hoechst+ nuclei were NeuN+ neuronal cells (nine biological replicates). Some remaining cells were GFAP+ astrocytes (arrows), characterized by ovoid nuclei. Hoechst NeuN GFAP 50µm 50µm 50µm Hoechst NeuN B C D E 20µm DIV3 DIV 7 DIV 10 DIV 14 HoechstMAP2 A 50µm * 36 3.2 Effects of Inflammatory Mediators on NPAS4 Expression 3.2.1 Glutamate Rapidly Alters NPAS4 Expression As activated immune cells can release glutamate in MS and EAE lesions89,230, we were interested in examining the effects of glutamate on NPAS4 expression in an in vitro setting. PBS or glutamate in PBS was added to neuronally enriched cultures for up to 9 hours to a final concentration of 1-30µM, a range which includes the physiologically relevant in vivo concentrations of extracellular glutamate (2-4 µM)231 as well as higher doses meant to induce glutamate excitotoxicity232. In steady-state untreated cells at or around DIV10, NPAS4 protein expression in neurons was largely negligible, where roughly 95% of PBS-treated NeuN+ cells were NPAS4-negative (Figure 3.2 A and B). Untreated neurons that expressed NPAS4 had low levels of expression, and very few medium, high or very high NPAS4+ cells were detected. Conversely, neurons treated with an excitatory stimulus of KCl at 25mM demonstrated high levels of NPAS4 expression, with more than 60% in the highest staining intensity category and less than 10% in the negligible staining category (Figure 3.2A, B and C). Across all treatments, NPAS4 staining at all intensity categories was localized to the nucleus (Figure 3.2C). 37 Figure 3.2 NPAS4 expression in response to excitatory stimuli. (A and B) Breakdown of the percentage of cells in each of the different NPAS4 staining intensity categories in Hoechst+NeuN+ cells for PBS- or KCl-treated (25mM) wells (n=7, mean % in each category, error bars represent SD). (C) Immunocytochemistry images showing representative NPAS4 intensity categories where negative was most common in untreated NeuN+ cells and very high was most representative of KCl-treated cultures. bEnd.3 cells served as a negative control for NPAS4. We first focused on global changes in expression by comparing changes in the mean fluorescent intensity (MFI) and the percentage of NPAS4-expressing or NeuN+NPAS4+ cells. Increasing glutamate concentrations rapidly increased the NPAS4 MFI 4-6-fold by 1 and 3 hours (Figure 3.3A) compared to no glutamate. While both 10 and 30µM tended to alter the MFI between 1-3 hours, considerable variability in the treatment effect meant only 10µM at 3 hours differed from the untreated control (Figure 3.3A, p=0.0125). However, when examining changes in MFI expression over 9 hours, the area under the curve (AUC) analysis showed significant 38 differences in the MFI at 10 and 30µM compared to no glutamate (Figure 3.3B, p=0.05 and 0.02, respectively). The percent-positivity seemed to be a more discriminatory readout for NPAS4 induction as glutamate tended to increase the percentage of NeuN+NPAS4+ cells at a dose as low as 1µM (Figure 3.3C). Despite the tendency for glutamate to induce rapid changes in the percent positivity at all doses, variability in the treatment effect meant only the 10µM at 3 (p=0.0016) and 7 hours (p=0.0155) were significantly different compared to no glutamate (Figure 3.3C). The AUC analysis also suggested that the percentage of NPAS4-expressing cells was a more sensitive measure, and NPAS4 induction was observed as low at 1µM in some experiments and at 3µM in all cultures (Figure 3.3D). However, only 10 and 30µM showed a significant increase in the percentage of NPAS4-expressing cells compared to the PBS control over 9 hours in all experiments (Figure 3.3D, p=0.02 and 0.04, respectively). NPAS4 expression changed rapidly in the presence of glutamate, with increases in the MFI observed as early as 1 hour, with values remaining high at 3 hours (Figure 3.3E). A more robust increase was observed at higher doses (Figure 3.3E). The AUC analysis showed the greatest increase in the MFI occurred across all doses at 3 hours; MFI values dropped thereafter (Figure 3.3F). Changes in the percent of NPAS4-expressing cells were also observed as early as 1 hour (Figure 3.3G). However, the AUC analysis showed that maximal expression in the percentage of NPAS4-expressing cells occurred earlier than changes in the MFI expression. NPAS4 percent positivity was highest at 1 hour, after which the proportion declined (Figure 3.3H). Evaluation of NPAS4 induction at the single cell level explains why changes in the percent positivity appeared earlier than changes in the MFI expression: MFI expression peaked later, given the lag in sufficient cells with an increased NPAS4 intensity to increase the population’s MFI. Despite the high NPAS4 expression observed at 1 or 3 hours, similar trends at 39 both time points meant there was no significant difference between expression patterns in either the MFI or the percentage of NPAS4-expressing cells at the two earliest time points tested. Ultimately, the increases in NPAS4 MFI and the percentage of NPAS4-expressing cells suggest that expression is increased early and remains elevated for 3-5 hours with fewer NPAS4-positive cells detected at later time points. 40 0 1 3 10 30010203040Glutamate Dose (μM)AUC0.050.021 3 5 7 9050100150200Time (h)AUC0.031 3 5 7 90500100015002000Time (h)AUCA B C D E F G H 0 5 10 15 20 25 30024681012Dose (μM)Normalized NPAS4 MFI 1h3h5h7h9h0 2 4 6 8 10024681012Time (h)Normalized NPAS4 MFI 0μM1μM3μM10μM30μM* 0 2 4 6 8 10020406080100Time (h)NPAS4 % positivity0μM1μM3μM10μM30μM** 0 5 10 15 20 25 30020406080100Dose (μM)NPAS4 % positivity1h3h5h7h9h0 1 3 10 300200400600Glutamate Dose (μM)AUC0.020.04* 41 Figure 3.3 Glutamate rapidly induced NPAS4 expression. Normalized NPAS4 MFI (A and E) and percent of NPAS4-expressing cells (C and G) at different times or doses of glutamate exposure. Each point represents the average of 3 biological replicates, with each replicate being the average of 2-3 technical replicates. Ratio-paired t-test with Bonferroni correction comparing 10 or 30µM to 0µM at 1-5 hours for the MFI (A) and 1-7 hours for the percent of NPAS4-expressing cells (C), or comparisons between 1 and 3 hours, or 1, 3 or 5 hours compared to 9 hours at 3, 10 and 30µM (E and G); * to the left of the data point indicate p<0.05, **p<0.01. All points represent the median with error bars and shaded regions representing the range. Associated area under the curve (AUC) analysis of the MFI (B) and percent positivity (D) across five different time points or the MFI (F) and percent positivity (H) across five different glutamate doses. Each point represents the average of 2-3 technical replicates for that biological replicate. Kruskal-Wallis test with Dunn’s multiple comparisons test to 0µM (B and D) or across all time points (F, H). All bars represent median +/- IQR. While the MFI and the percentage of positive cells are useful parameters, the former is less discriminatory, as there would need to be either a change in several neurons or a more considerable change in fewer neurons to cause an appreciable change in the MFI. At the same time, the latter is binary and doesn’t provide an accurate readout of the ranges of intensity observed for a protein, and it is unknown how the amount of protein expressed may influence biology within the cell. Therefore, we applied an alternate readout that would reflect both intensity and proportion across the NeuN+ cells in culture by examining the proportion of neurons with specific staining intensity ranges. The categories were determined as described in section 2.3.2, and individual cells were allocated into five categories: negative, low, medium, high and very high expression. The proportions of NPAS4-negative and the remaining NPAS4-low cells in untreated neuronal cultures were similar over the 9 hours (Figure 3.4A). Similar to the percent of NPAS4+ cells, the percent-based intensity category readout reflected changes in NPAS4 expression as low as 1µM and as early as 1 hour of glutamate exposure. At this dose, we saw a decrease in the percentage of NPAS4-negative cells and an increase in the number of cells in the low and medium expression categories compared to no glutamate. By 5 hours, a decrease in the 42 proportion of higher intensity category NPAS4-staining cells was observed and closely mirrored the control treatment’s values. Doses of 3µM and higher showed increasing percentages of higher intensity NPAS4 staining cells as early as 1 hour (Figure 3.4A and B). However, despite the rapid decline in the percentage of NPAS4-negative cells at 3µM (1-3 hours), values eventually returned to steady-state levels by 7-9 hours, with minimal NPAS4+ cells present (Figure 3.4B). Rapid shifts by 1-3 hours in NPAS4+ categories were also observed at the two highest tested doses of glutamate, with the greatest change occurring at 10µM (Figure 3.4A and B). At 10 or 30µM glutamate, NPAS4 expression seemed to be sustained: by 5-9 hours, roughly 25% of cells still expressed NPAS4, with over 10% of cells in the medium to very high NPAS4 expression categories. This was unlike lower doses where NPAS4 was typically in the low or negligible categories by 5 hours (Figure 3.4A and B). 43 Figure 3.4 Early changes in NPAS4 intensity at all doses of glutamate. (A) Breakdown of the percentage of NeuN+ cells in each of the NPAS4 staining intensity categories following glutamate exposure at different doses and exposure times. Each bar represents the average of 3 biological replicates, with each replicate being the average of 2-3 technical replicates. Bars represent mean +/- SD. (B) Representative immunocytochemistry images showing NPAS4 expression intensities after shorter (3 hour) and longer (7 hour) exposure to lower (3µM) versus higher doses (10µM) of glutamate. In some glutamate treatments, we observed an apparent decrease in NeuN+ cells, which may represent decreased cell viability or cell death. We examined changes in cell numbers across treatments by normalizing the number of NeuN+ cells in the treatment well to the number of 3h 7h 0µM 3µM 10µM Hoechst NeuN NPAS4 B Hoechst NeuN NPAS4 50µm 50µm 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+cellsNeg Low Med High Very High—— 0μM —— —— 3μM ———— 1μM —— — 10μM — — 30μM —A 44 NeuN+ cells in the control 0µM treatment well (Figure 3.5A). At 1 and 3µM glutamate, we saw no significant reduction in the number of NeuN+ cells per well over the entire 9 hours, except at 3µM at 9 hours (Figure 3.5A, p=0.013). At 10 and 30µM, we detected a significant reduction in NeuN+ cells by 9 (p=0.002) and 7 hours (p=0.018), respectively; this represented as much as a 50% loss in NeuN+ cells/region of interest compared to the 0µM or untreated dose (Figure 3.5A). Visual changes in cell morphology could also be observed as early as 5 hours of treatment with 30µM glutamate, evidenced by ruptured plasma membranes and the presence of pyknotic nuclei that were still detectable for up to 9 hours (Figure 3.5B). By 7-9 hours, complete axonal dissolution was further evidence of cell degeneration. The decrease in NPAS4 at prolonged exposure to 30µM glutamate may be associated with reduced cell viability as the average NeuN counts tended to decrease in each biological replicate when examined over the 9-hour assay. Despite this trend of decreasing cell counts, the presence of ~25-30% NPAS4-expressing cells at 7-9 hours for 10 and 30µM showed that some of the viable cells are NPAS4+ (Figure 3.4A). 45 Figure 3.5 Decreased cell viability following exposure to higher doses of glutamate. (A) Average Hoechst+NeuN+ counts per similar unit area across different doses and time points of glutamate exposure. Each shape represents a biological replicate (n=3), with each point being the mean of 2-3 technical replicates. Repeated measures two-way ANOVA with Dunnet’s multiple comparisons post-hoc test comparing each dose to 0µM at the same time point, *p<0.05, **p<0.01. Bars represent median +/- IQR. (B) Representative phase and immunocytochemistry images showing changes in cell morphology/viability over time, including condensed or pyknotic nuclei (*) or axonal dissolution (arrowhead) during exposure to 30µM glutamate. 0μM 1μM 3μM 10μM 30μM KCl0.00.51.01.52.0TreatmentsAverage NeuN Counts 1h3h5h7h9hKCl0µM 30µM 1h 30µM 3h 30µM 5h 30µM 9h 20µm A B 20µm 30µM 7h Hoechst NeuN Hoechst NeuN Hoechst NeuN 20µm 20µm 20µm 20µm * * * ** * 46 3.2.2 High Doses of H2O2 Induce Late NPAS4 Expression As a proxy for mild to moderate oxidative stress, we exposed primary neuronal cortical cultures to H2O2 in PBS at a final concentration of 12.5, 25, 50 and 100µM. Doses of 50µM and greater tended to increase the NPAS4 MFI as early as 3 hours after addition (Figure 3.6A) but did not reach significance compared to 0µM at any of the assessed time points (3-7 hours). A higher dose of 100µM did not increase the MFI further, as MFI values remained comparable until 9 hours, when greater experimental variability was observed (Figure 3.6A). The AUC analysis showed that increases in the NPAS4 MFI at 50 and 100µM over 9 hours were comparable and significantly different to the control dose (Figure 3.6B, p=0.03 and p=0.04, respectively). As a more discriminatory measure, the percentage of NPAS4+ cells allowed us to discern differences at 50µM and above. Secondary analyses showed that the percentage of NPAS4-expressing cells was significantly increased with 100µM H2O2 at 3 (p=0.02) and 5 hours (p=0.02) compared to the untreated controls (Figure 3.6C). The AUC analysis over 9 hours showed the percentage of NPAS4-expressing cells at 100µM tended to be greater than the percentage seen in cells treated with 50µM (Figure 3.6D), but the large variability at these higher doses meant no dose reached significance (Figure 3.6D). A secondary analysis comparing the AUC of 50 and 100µM doses to the control showed that the 50µM treatment elevated NPAS4 percent positivity over the 9 hours compared to the control treatment (Figure 3.6D, p=0.0286). Notably, no differences between NPAS4 expression (neither MFI nor percent-positivity) were observed between controls and 12.5 or 25µM (Figure 3.6A and C). Irrespective of dose, the earliest differences in NPAS4 expression measured by MFI tended to appear as early as 3 hours (Figure 3.6E). Maximal NPAS4 MFI expression was observed at 5 hours of treatment, where a secondary analysis showed that 50µM at 5 hours was significantly elevated over the 1-hour time point (Figure 3.6E, p=0.028). The AUC analysis 47 showed that H2O2 tended to increase the MFI as early as 3 hours, however, levels were highest at 5 hours across all doses (Figure 3.6F, p=0.007). Increases in the percentage of NPAS4-expressing cells followed a similar trend to those observed for the MFI, however, changes were observed as early as 3 hours (Figure 3.6G). No changes were observed at any dose following one hour of exposure, which was subsequently used as a baseline comparator. Maximal increases in the percentage of NPAS4-expressing cells were observed at 5 hours, where 100µM was the greatest driver compared to 1 hour (Figure 3.6G, p=0.047). The AUC analysis suggested that the percent positivity tended to increase across all examined doses as early as 3 hours but did not reach significance with the high variability between biological replicates (Figure 3.6H). 48 49 Figure 3.6 Prolonged exposure to high doses of H2O2 induced NPAS4 expression. Normalized NPAS4 MFI (A and E) and percent of NPAS4-expressing cells (C and G) at different times or doses of H2O2 exposure. Each point represents the average of 4 biological replicates, with each replicate being the average of 2-3 technical replicates. Ratio-paired t-test with Bonferroni correction comparing 50 and 100 µM to 0µM at 3-7 hours (A and C) or comparing 3-7 hours to 1 hour at 50 and 100µM (E and G), * to the left of a data point is p<0.05. Bars represent the median, and whiskers or shaded regions represent the IQR. Associated area under the curve (AUC) analysis of the MFI (B) and percent positivity (D) across five different time points or the MFI (F) and percent positivity (H) across five different doses of H2O2 exposure. Each point represents the average of 2-3 technical replicates. Kruskal-Wallis test with Dunn’s multiple comparisons test to 0µM (B and D) or across all time points (F, H) with secondary Mann-Whitney t-test comparing 50 and 100µM to 0µM (D). All bars represent median +/- IQR. We examined the % of cells staining with different NPAS4 intensity categories and saw no differences after treatment with 12.5 or 25µM compared to controls; 80-90% of cells were NPAS4-negative, while the rest were low NPAS4-expressing (Figure 3.7A and B). This breakdown of expression intensity was comparable to that observed in untreated cells. At 50 and 100µM, the distribution of the staining intensities over time was quite similar (Figure 3.7A). By 3 hours, there was a decrease in the number of NPAS4-negative cells with a corresponding increase in cells in the low category and the emergence of cells in the medium to very high categories compared to 1 hour (Figure 3.7A). By 5 hours, we observed the greatest changes at 50 and 100µM, where approximately 50% of cells were NPAS4+, while 15-20% were very high NPAS4-expressing cells (Figure 3.7A and B); these distributions were largely maintained until 9 hours when we ended the experiment. 50 Figure 3.7 Changes in NPAS4 intensity observed as early as 3 hours post-exposure to high doses of H2O2. (A) Breakdown of the selected NPAS4 staining intensity categories at different doses and time points of H2O2 exposure. Each bar represents the average of 4 biological replicates, with each replicate being the average of 2-3 technical replicates. Bars represent mean +/- SD. (B) Representative immunocytochemistry images showing changes in NPAS4 expression following exposure to low (12.5µM) or high (50 µM) doses of H2O2 at shorter (1 hour) or longer (5 hour) intervals. Similar to glutamate, prolonged exposure to high doses of H2O2 also impacted neuronal morphology and viability, impacting NeuN+ cell counts (Figure 3.8A). The changes in NPAS4 expression (MFI, percent positivity and the different intensity categories) observed at 50µM (3-7 1h 5h 0µM 12.5µM 50µM Hoechst NeuN NPAS4 B 50µm Hoechst NeuN NPAS4 50µm 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High—— 0μM —— — 12.5μM — — 50μM —— 25μM — — 100μM —A 51 hours) and 100µM (3 hours) appeared to precede cell death, as the number of NeuN+ cells tended to decrease over time (Figure 3.8A). However, at 50µM (9 hours) and 100µM (5-7 hours), we observed altered morphology and a decreasing trend in cell loss, as shown by a 1.5-3-fold decrease in the NeuN+ counts compared to the PBS-treated wells (Figure 3.8A). By 9 hours of 100µM exposure, a significant decrease in the NeuN+ counts was observed compared to PBS-treated wells (Figure 3.8A, p=0.0130). Following prolonged exposure to 50 (7-9 hours) or 100µM H2O2 (5-9 hours), visual changes emerged, with evidence of cell swelling, ruptured cell membranes, and pyknotic nuclei (Figure 3.8B). By 9 hours, most regions had complete dendritic and/or axonal dissolution and multiple pyknotic nuclei that suggested ongoing cellular loss from 5 to 9 hours. Given this observed decrease in cell viability at 100µM from 7-9 hours, no secondary comparisons for the NPAS4 MFI or percentage of NPAS4-expressing cells were performed at these time points despite the increased NPAS4 expression observed. Despite the apparent NeuN+ cell loss over the 7-9 and 5-9 hours of exposure to 50 and 100µM, respectively, 25-40% of cells remained NPAS4+ in each of these analyses. These findings suggest the NeuN+ cells still present following 100µM for 7-9 hours of exposure were positive for NPAS4 (Figure 3.7A). 52 Figure 3.8 Decreased cell viability at high doses of H2O2. (A) Average Hoechst+NeuN+ counts across different doses and time points of H2O2 exposure. Each shape represents a biological replicate (n=4), and each point is the average of 2-3 technical replicates. Repeated measures two-way ANOVA with Dunnet’s multiple comparisons post-hoc test comparing each dose to 0µM at the same time point, *p<0.05. Bars represent median +/- IQR. (B) Representative phase and immunocytochemistry images showing changes in cell morphology suggestive of altered viability over time following exposure to 100µM H2O2. By 5-7 hours of exposure to 100µM H2O2, some cells showed swelling and occasional pyknotic nuclei (*) with loss of cytoplasmic processes and condensed nuclei and nuclear shrinkage (arrowheads) in the majority of cells by 7-9 hours. 0μM 12.5μM 25μM 50μM 100μM KCl0.00.51.01.52.0TreatmentsAverage NeuN Counts1h3h5h7h9hKCl20µm A 0µM 100µM 1h 100µM 3h 100µM 5h 100µM 9h B 100µM 7h Hoechst NeuN Hoechst NeuN Hoechst NeuN * * * * * 20µm 20µm 20µm 20µm 20µm 53 3.3 Effects of Inflammatory Mediators on ARNT2 Expression 3.3.1 High Doses of Glutamate Tended to Reduce ARNT2 Levels As the major primary binding partner of NPAS4 in the CNS, we also examined the effects of inflammatory mediators on ARNT2 expression. Unlike NPAS4, which requires a stimulus to be induced, ARNT2 was constitutively expressed, but levels were heterogeneous across all NeuN+ cells. In untreated conditions, over 97% of Hoechst+NeuN+ cells at DIV10 expressed ARNT2 (n=7) (Figure 3.9A and B), where the majority (~70%) of cells displayed low- to medium-intensity expression. The remaining positive cells (~28%) fell in the high to very high intensity staining category (Figure 3.9C). 54 Figure 3.9 ARNT2 is expressed constitutively in DIV10 cortical neurons. (A and B) Breakdown of the percentage of cells in each of the different ARNT2 staining intensity categories in Hoechst+NeuN+ cells for PBS-treated wells (n=7, mean % in each category, error bars represent SD). (C) Representative immunocytochemistry images showing the ARNT2 intensity categories, where low and medium cells comprised nearly 75% of untreated NeuN+ cells. bEnd.3 cells served as a negative control for ARNT2. 55 Exposure to glutamate tended to alter ARNT2 expression levels only at 30µM and as early as 1 hour post-exposure (Figure 3.10A). ARNT2 MFI levels tended to be lower at most of the time points examined, as represented by the AUC analysis of ARNT2 MFI at 30µM (Figure 3.10B). Similarly, the percentage of ARNT2-expressing cells tended to drop at 30µM, as early as 1 hour, with more robust changes by 3-5 hours (Figure 3.10C). Similar to the MFI, the percentage of ARNT2-expressing cells also tended to be lower at all time points examined, as represented by the AUC analysis of the percentage of ARNT2-expressing cells (Figure 3.10D). The exposure time did not appear to affect ARNT2 levels at any dose, as no differences were observed in either the ARNT2 MFI or the percentage of ARNT2-expressing cells (Figure 3.10E and G). This was also reflected in the AUC analysis for both the MFI and the percent positivity, where both remained comparable across all time points regardless of the dose (Figure 3.10F and H). 56 A B C D E F G H 0 2 4 6 8 10050100Time (h)ARNT2 %Positivity0μM1μM3μM10μM30μM0 1 3 10 300246810Dose (μM)AUC0 1 3 10 3002004006008001000Dose (μM)AUC0 5 10 15 20 25 300.00.51.01.5Dose (μM)Normalized ARNT2 MFI 1h3h5h7h9h0 5 10 15 20 25 30050100Dose (μM)ARNT2 %Positivity1h3h5h7h9h1 3 5 7 9010203040Time (h)AUC1 3 5 7 90100020003000Time (h)AUC0 2 4 6 8 100.00.51.01.5Time (h)Normalized ARNT2 MFI 0μM1μM3μM10μM30μM57 Figure 3.10 High doses of glutamate tended to reduce ARNT2 levels. Normalized ARNT2 MFI (A and E) and percent of ARNT2-expressing cells (C and G) at different times or doses of glutamate exposure. Each point represents the average of 3 biological replicates, each the average of 2-3 technical replicates. Bars represent the median, and whiskers, dotted lines, or shaded regions represent the range. Associated area under the curve (AUC) analysis of the MFI (B) and percent positivity (D) across five different time points or the MFI (F) and percent positivity (H) across five different doses of glutamate exposure. Each point is a biological replicate derived from 2-3 technical replicates. Kruskal-Wallis test with Dunn’s multiple comparisons test to 0µM (B and D) or across all time points (F and H). All bars represent median +/- IQR. We examined the breakdown of ARNT2 staining intensity categories and found the majority of untreated or low dose (1-3µM) glutamate-treated cells expressed low to medium intensity ARNT2 (Figure 3.11A and B). At each of these doses and across all time points, the percentage of NeuN+ cells in each of the five categories remained comparable. Reductions in the percentage of cells in each intensity category were first apparent at 10µM, with a noticeable increase in the percentage of ARNT2-negative cells seen as early as 3 hours (Figure 3.11A). At 30µM, there tended to be more ARNT2-negative cells present by 1 hour of exposure (~20%) (Figure 3.11A and B). By 3 hours and longer, negative cells represented 40-50% of NeuN+ cells (Figure 3.11A and B), although the variability between biological replicates was high. The variability in ARNT2 expression following glutamate exposure made it difficult to associate expression with viability. However, the 2-fold reduction in NeuN+ cell counts at 7-9 hours of 30µM glutamate suggests that reductions in ARNT2 expressing cells as early as 1 and 3 hours preceded changes in morphology or cell viability. 58 Hoechst NeuN ARNT2 -------- 1h -------- ARNT2 0µM 3µM 30µM B Hoechst NeuN ARNT2 ARNT2 50µm 50µm 50µm 50µm -------- 9h -------- 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High—— 0μM —— —— 1μM —— —— 3μM —— — 10μM — — 30μM —A 59 Figure 3.11 High doses of glutamate tended to reduce ARNT2 intensity levels. (A) Breakdown of the selected ARNT2 staining intensity categories at different doses and time points of glutamate exposure. Each bar represents the average of 3 biological replicates, with each replicate being the average of 2-3 technical replicates. Bars represent mean +/- SD. (B) Representative immunocytochemistry images showing reductions in ARNT2-expressing NeuN+ cells following high-dose glutamate exposure, which was not detected at 3uM. 3.3.2 Prolonged Exposure to High Doses of H2O2 Reduced ARNT2 Levels Reductions in the ARNT2 MFI expression were observed as low as 50µM H2O2, and as early as 3 hours, reaching significance compared to controls at 7 hours (Figure 3.12A, p=0.0192). The highest dose (100µM) caused significant reductions in the ARNT2 MFI from 5-9 hours (Figure 3.12A, p=0.0081, 0.0061, 0.0169, respectively), although reductions were also apparent as early as 3 hours. The AUC analysis spanning the full 9 hours of testing showed the AUC MFI tended to drop with 50µM, yet only the 100µM dose differed from the control dose (Figure 3.12B, p=0.03). Changes in the percentage of ARNT2-expressing cells occurred later than changes in the MFI, reflecting the high frequency of ARNT2+ cells in these cultures. The percentage of ARNT2-expressing cells dropped minimally by 7 hours of exposure to 50µM H2O2 but tended to be more robust in response to 100µM H2O2 at 7 and 9 hours (Figure 3.12C). Changes in the AUC were comparable, as 100µM significantly reduced ARNT2 MFI over the 9 hours compared to the control dose (Figure 3.12D, p=0.03). No changes in ARNT2 MFI across all NeuN+ cells or the percentage of NeuN+ARNT2+ cells were observed at 12.5 and 25µM (Figure 3.12A and C) at any timepoint compared to PBS-treated controls (Figure 3.12B and D). The duration of H2O2 exposure did affect neuronal ARNT2 levels. At the two highest tested doses, the ARNT2 MFI tended to be reduced at 3 hours (Figure 3.12E), yet significant differences in the MFI were only observed at 100µM at 5, 7, and 9 hours compared to 1 hour (Figure 3.12E, p=0.0069, 0.0045, 0.0097, respectively). The AUC analysis also tended to show a 60 reduction at 3 to 5 hours, reaching significance at 7 hours (Figure 3.12F, p=0.05). Overall, longer exposure tended to be associated with a greater reduction in the percentage of ARNT2-expressing cells (Figure 3.12G); this was also reflected in the AUC analysis, where slight reductions in the AUC for the percent of ARNT2-expressing cells gradually decreased over the longer exposure times tested (Figure 3.12H). The earlier reduction in ARNT2 MFI compared to the percent positivity across 50 and 100µM suggests a decrease in intensity precedes the complete loss in expression. 61 A B C D E F G H 0 2 4 6 8 100.00.51.01.5Time (h)Normalized ARNT2 MFI 0μM12.5μM25μM50μM100μM0 2 4 6 8 10050100Time (h)ARNT2 %Positivity0μM12.5μM25μM50μM100μM0 12.5 25 50 1000246810Dose (μM)AUC0.020 12.5 25 50 10002004006008001000Dose (μM)AUC0.030 50 1000.00.51.01.5Dose (μM)Normalized ARNT2 MFI 1h3h5h7h9h0 50 100050100Dose (μM)ARNT2 %Positivity1h3h5h7h9h1 3 5 7 9050001000015000Time (h)AUC1 3 5 7 9050100150Time (h)AUC0.05** ** * * ** ** ** 62 Figure 3.12 Prolonged exposure to high doses of H2O2 reduces ARNT2 expression levels. Normalized ARNT2 MFI (A and E) and percentage of ARNT2-expressing cells (C and G) at different times and doses of H2O2 exposure. Each point represents the average of 4 biological replicates, with each biological replicate being the average of 2-3 technical replicates. Ratio-paired t-test to 0µM at 5-9 hours for 50 (A) and 100µM (A and C) or comparisons between 1 hour and 5-9 hours for 50 and 100µM (E and G), * to the left of the datapoint is p<0.05, **p<0.01. Bars represent the median, and whiskers or the shaded region represent the IQR. Associated area under the curve (AUC) analysis of the MFI (B) and percent positivity (D) across five different time points or the MFI (F) and percent positivity (H) across five different doses of H2O2 exposure. Each point represents the average of 4 biological replicates, each derived from the average of 2-3 technical replicates. Kruskal-Wallis test with Dunn’s multiple comparisons test to 0µM (B and D) or across all time points (F, H). All bars represent median +/- IQR. The proportions of cells in each ARNT2 staining category were largely unchanged over the 9 hours of exposure to 12.5 or 25µM H2O2 compared to controls (Figure 3.13A and B). Regardless of the dose, there was no change in the ARNT2 expression 1-hour post-exposure (Figure 3.13A and B). At 50 and 100µM, altered ARNT2 expression was observed at 5-9 hours, suggesting that H2O2 requires time to affect ARNT2 expression. Reductions in the percentage of ARNT2-expressing cells and MFI intensity at 100µM seemed to coincide with time points at which there was also a reduction in the number of NeuN+ cells (Figure 3.8A). At 5 hours of 100µM H2O2 exposure, roughly 50% of NeuN+ cells had been lost, therefore, changes in ARNT2 expression must be interpreted with caution because of the extensive NeuN+ cell drop-off. However, the observed reduction in ARNT2 MFI and the slight reduction in the percentage of ARNT2-expressing cells seen at 50µM appeared to be independent of changes in the number of neuronal cells (Figure 3.8A). The reduction in the ARNT2 MFI appearing as early as 3 hours for 100µM also appeared to precede cell death and seemed to be linked to a decrease in cell viability at the later time points. Ultimately, these results suggest prolonged exposure to high doses of H2O2 gives way to reduced ARNT2 staining and, ultimately, a total loss of ARNT2 expression in some cells. 63 0µM 12.5µM 50µM Hoechst NeuN ARNT2 ARNT2 B Hoechst NeuN ARNT2 ARNT2 50µm 50µm 50µm 50µm -------- 9h -------- -------- 1h -------- 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+cellsNeg Low Med High Very High—— 0μM —— — 12.5μM — — 25μM — — 50μM — — 100μM —A 64 Figure 3.13 Changes in ARNT2 intensity levels observed at prolonged exposure to H2O2. (A) Breakdown of the selected ARNT2 intensity categories at different doses and time points of H2O2 exposure. Each bar represents the average of 4 biological replicates, with each replicate being the average of 2-3 technical replicates. Bars represent mean +/- SD. (B) Representative immunocytochemistry images showing changes in ARNT2 MFI in NeuN+ cells and the reduction in NeuN+ARNT2+ cells observed at prolonged exposure to high doses of H2O2 (50µM). 3.4 Linking Effects of Inflammatory Mediators on ARNT2 and NPAS4 Intensity Levels to Viability Across Select Focus Treatment Groups 3.4.1 Categorical Changes in NPAS4 and ARNT2 Expression During Glutamate and H2O2 Exposure Given the relationship between NPAS4 and ARNT2 and their ability to heterodimerize in the CNS to upregulate transcription of key neuroprotective factors, we were also interested in comparing the general trends in ARNT2 and NPAS4 intensity levels. We focused on 30µM glutamate as this dose tended to show the most differences in ARNT2 expression levels. Compared to the control treatment, 30µM glutamate rapidly upregulated NPAS4 expression with roughly 50% of cells expressing NPAS4 across 1-5 hours post-exposure (Figure 3.14A). This change in intensity was primarily due to an increase to about 25-30% in the number of medium to very high expressing cells. By 7 and 9 hours, the proportion of cells staining in the medium to very high category was greatly decreased, with only 5-10% of cells expressing medium to very high NPAS4. By 1 hour of 30µM glutamate exposure, there was already an increase in the number of ARNT2-negative cells (Figure 3.14B). By 3 hours, roughly 40-50% of cells were ARNT2-negative (Figure 3.14B), whereas approximately 50% were NPAS4-negative at this time point (Figure 3.14A). Across the population, glutamate exposure drove NPAS4 but also led to reductions in ARNT2 (Figure 3.14C). This trend might suggest a reduced likelihood of these two factors interacting in inflammatory settings but remains to be examined at the individual cell level. A qualitative attempt to examine colocalization revealed that many of the NPAS4 65 moderate or bright expressing NeuN+ cells tended to be low or negative for ARNT2 (Figure 3.14C, 30µM at 3 and 5 hours). The NeuN+ staining on many of these cells was bright, and cell bodies and cytoplasmic processes appeared typical. However, the decrease in ARNT2 levels suggests that NPAS4 may be less likely to interact with ARNT2 following exposure to high doses of glutamate. 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High-------------------- 0μM -------------------- ------------------- 30μM -------------------1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High-------------------- 0μM -------------------- ------------------- 30μM -------------------NPAS4 ARNT2 A B 66 Figure 3.14 High dose glutamate rapidly upregulates NPAS4 while downregulating ARNT2. Breakdown of the selected NPAS4 (A) and ARNT2 (B) intensity categories in untreated conditions or following exposure to 30µM glutamate (the same samples/an excerpt of 0 and 30µM treatments from Figures 3.4A and 3.11A). Each bar is the average of 3 biological replicates, with each being the average of 2-3 technical replicates. All bars represent mean +/- SD. (C) Representative phase and immunocytochemistry images showing changes in neuronal morphology and NeuN, ARNT2 and NPAS4 staining over time following exposure to 30µM glutamate. Many NPAS4 moderate or bright expressing NeuN+ cells tended to be low or negative for ARNT2 (30µM at 3 and 5 hours; cells with asterisks, dotted circles in NPAS4 and ARNT2 panels, respectively). The NeuN+ staining on many of these cells was bright, and cell bodies and cytoplasmic processes (arrowheads) were largely typical. 67 Similarly, we focused on H2O2 at 100µM exposure over time. NPAS4 increases were not observed until 3 hours of exposure (Figure 3.15A). At this time point, there was an emergence of cells in the medium to very high intensity categories. By 5 hours, roughly 20-25% of cells expressed medium to very high NPAS4 levels, and overall, 40-50% of NeuN+ cells were NPAS4+. By 3 hours of treatment, we also saw increases in the percentage of ARNT2-negative cells (Figure 3.15B). The proportions of ARNT2 and NPAS4 cells in each staining category following 5, 7 and 9 hours of 100µM treatment were noticeably different compared to the control cells at those time points. At these time points, opposite changes in NPAS4 and ARNT2 expression were observed, as 40-50% of NeuN+ cells had become NPAS4+ while 40-60% of NeuN+ cells had become ARNT2-negative (Figure 3.15C). Strikingly, the NPAS4+ cells at 5, 7 or 9 hours of 100µM H2O2 exposure seemed to be predominantly ARNT2-low or, more often, ARNT2-negative. 68 1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High-------------------- 0μM -------------------- ------------------ 100μM ------------------1h 3h 5h 7h 9h 1h 3h 5h 7h 9h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High-------------------- 0μM -------------------- ------------------ 100μM ------------------NPAS4 ARNT2 A B 69 Figure 3.15 Prolonged exposure to 100µM H2O2 increases NPAS4 expression while simultaneously decreasing ARNT2 levels. Breakdown of the selected NPAS4 (A) and ARNT2 (B) intensity categories in steady-state untreated conditions or following exposure to 100µM H2O2. Each bar is the average of 4 biological replicates, with each being the average of 2-3 technical replicates. All bars represent mean +/- SD. (C) Representative phase and immunocytochemistry images showing changes in neuronal morphology and NeuN, ARNT2 and NPAS4 staining over time following exposure to 100µM H2O2. Many of the NPAS4 moderate or bright expressing NeuN+ cells tended to be low or negative for ARNT2 (100µM at 5, 7 or 9 hours; cells with asterisks, dotted circles in NPAS4 and ARNT2 panels, respectively). The NeuN+ staining on many of these cells was bright, and cell bodies appeared typical with several cytoplasmic processes (arrowheads). 70 3.5 Effect of Inflammatory Cells on NPAS4 and ARNT2 Expression 3.5.1 The Number of Inflammatory T-Cells Rather than Activation Status Seems to Induce Early NPAS4 Expression We also examined the effects of resting and activated murine CD4+ T-cells on neuronal NPAS4 expression in co-culture conditions. In a pilot study, freshly isolated naïve or activated (anti-CD3 and anti-CD28) CD4+ T-cells were added to E15.5-17.5 murine primary neuron-enriched cortical cultures for 1-6 hours. Cells were initially co-cultured at a ratio of five T-cells to 1 neuron in each well (i.e. 75,000 T-cells:15,000 neurons) (Figure 3.16A), a dose which previously gave robust NPAS4 changes in western blot analyses (Quandt lab, unpublished data). At this dose, extended incubation with resting CD4+ T-cells tended to upregulate NPAS4 MFI from 2-6 hours in two of the three experiments, while no increase in the NPAS4 MFI was observed following exposure to activated T-cells (Figure 3.16A). However, an increase in the ratio of CD4+ T-cells to neurons tended to increase the NPAS4 MFI (Figure 3.16B). Both a 10:1 (150,000 T-cells) and 20:1 (300,000 T-cells) ratio of resting or activated T-cells to neurons increased the NPAS4 MFI (Figure 3.16B). This increase was observed as early as 1 hour, with a 1.5-1.75-fold increase in the MFI over the controls. By 2 hours, the NPAS4 MFI with 150,000 and 300,000 resting and activated T-cells yielded a 2-3-fold increase in NPAS4 MFI compared to control media (Figure 3.16B). We also examined the different staining intensities of NPAS4-expressing cells at a 5:1 ratio (Figure 3.16C) and a 10:1 and 20:1 ratio of T-cells to neurons (Figure 3.16D). We saw that short exposure to the control treatment (neuronal maintenance media – “NMM”) led to an increase in the proportion of low (~20%) or medium (~10%) NPAS4-expressing neurons (Figure 3.16C). However, by 4-6 hours, all NeuN+ cells were again either NPAS4-negative or low. Slight differences in NPAS4 expression tended to occur at 1 or 2 hours for resting or activated T-cells, 71 respectively, as a few NPAS4 high and very high cells could be detected (Figure 3.16C). An increase in the ratio of resting or activated T-cells to neurons tended to increase the proportion of NPAS4+ cells, with a higher percentage of cells in the medium to very high expression category (Figure 3.16D). After 2 hours of co-culture in the presence of 150,000 and 300,000 T-cells, roughly 15-20% and 25-40% of neurons were in the very high NPAS4 expression category, respectively. This increase in the percentage of very high NPAS4 cells matched a decrease in the proportion of NPAS4-negative cells, with roughly 20-25% of cells staining as NPAS4-negative following exposure to 300,000 T-cells for 2 hours (Figure 3.16D). Notably, the trends were similar for both resting and the activated T-cells across time points and similar T-cell amounts. These results suggest that the number, rather than the activation status of T-cells, is a greater driver of NPAS4 expression at these time points. However, given the low sample size (n=1-3) and the lack of data examining the activation status of the CD4+ T-cells, further replicates are required to examine the effects of these cells on NPAS4 expression. 72 1h 2h 3h 4h 6h 1h 2h 3h 4h 6h 1h 2h 3h 4h 6h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High———— NMM ———— ———— Rest ———— ———— Act ————1h 2h 3h 4h 5h 6h 1h 2h 3h 4h 5h 6h 2h01234568TreatmentNormalized NPAS4 MFI—— Rest —— —— Act —— KCl75k150k300k 75k150k300k 75k150k300k 75k150k300k0123456Normalized NPAS4 MFI—Rest— —Act— —Rest— —Act————— 1h ———— ———— 2h ————TreatmentA B C 75k 150k 300k 75k 150k 300k 75k 150k 300k 75k 150k 300k0255075100Treatment% of Hoechst+NeuN+ cellsNeg——— Rest ———Low Med High Very High——— Act ——— ——— Rest ——— ——— Act ———-------------------- 1h -------------------- -------------------- 2h --------------------D 73 Figure 3.16 Number but not activation status of CD4+ T cells drives NPAS4 expression. Normalized NPAS4 MFI at different incubation times with 75,000 resting or activated (anti-CD3 and anti-CD28) CD4+ T-cells (A) or following 1- or 2-hour incubation with 75, 150, or 300k resting or activated T-cells (B). Each symbol represents a different biological replicate (n=1-3). Breakdown of the NPAS4 staining categories after different incubation times with added neuronal maintenance media (NMM), 75,000 resting or activated T-cells (C), or 1- or 2-hour incubation with 75, 150, or 300k resting or activated T-cells (D). Bars represent the average of 1-3 biological replicates per experiment, and error bars represent median +/- IQR (A and B) or mean +/- SD (C and D); some incubations could only be performed with one dose of T cells or fewer time points. 3.5.2 CD4+ T-lymphocytes Do Not Alter ARNT2 Expression At all tested time points, the ARNT2 MFI values remained similar regardless of exposure to resting or activated CD4+ T-cells at low or higher ratios to neurons (Figure 3.17A and B). ARNT2 was detected at a level similar to neuronal maintenance media-treated wells, which notably did not change with the addition of NMM as observed for NPAS4 expression (Figure 3.17A). An increase in the ratio of T-cells to neurons did not alter ARNT2 expression measured by MFI compared to the control treatment (Figure 3.17B). ARNT2 intensity category proportions also remained similar. Increasing the ratio of T-cells to neurons had no impact on the ARNT2 staining category distribution, regardless of cell activation status (Figure 3.17D). More experiments are required to examine whether longer CD4 T cell coincubation may alter the percentage of cells in each ARNT2 staining category. 74 1h 2h 3h 4h 6h 1h 2h 3h 4h 6h 1h 2h 3h 4h 6h0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High---------- NMM ---------- ---------- Rest ---------- ---------- Act ----------1h 2h 3h 5h 4h 6h 1h 2h 3h 4h 5h 6h 2h0123TreatmentNormalized ARNT2 MFI——Rest—— ——Act—— KCl75k150k300k 75k150k300k 75k150k300k 75k150k300k0123TreatmentNormalized ARNT2 MFI—Rest— —Act— —Rest— —Act————— 1h ———— ———— 2h ————A B C 75k 150k 300k 75k 150k 300k 75k 150k 300k 75k 150k 300k0255075100Treatment% of Hoechst+NeuN+ cellsNeg Low Med High Very High------ Rest ------ ------ Act ------ ------ Rest ------ ------ Act -------------------------- 1h -------------------- -------------------- 2h --------------------D 75 Figure 3.17 ARNT2 expression was unchanged by neuronal cell incubation with resting or activated CD4+ T-cells. Normalized ARNT2 MFI at different times of exposure to 75,000 resting or activated (anti-CD3 and anti-CD28) CD4+ T-cells (A) or to 1- or 2-hour exposure to 75k, 150k, or 300k resting or activated T-cells (B). Each symbol represents a different biological replicate (n=1-3). Breakdown of the selected ARNT2 intensities at different times of exposure to neurobasal media (NBM) or 75,000 resting or activated T-Cells (C) or to 1- or 2-hour exposure to 75k, 150k, or 300k resting or activated T-cells (D). Bars represent the average of 1-3 biological replicates per experiment, and error bars represent median +/- IQR (A and B) or mean +/- SD (C and D); some incubations were only performed with one dose of T cells or fewer time points due to limited T cell numbers. 76 Chapter 4: Discussion 4.1 Summary of Key Findings The independent functional roles of NPAS4 and ARNT2 in hypoxic196,213, ischemic165,174,196 and neurodegenerative settings have been studied extensively. Recent findings have also suggested a role for these two proteins in in vivo inflammatory degenerative settings relevant to CNS chronic inflammatory conditions1. While their expression patterns and functional roles have been well-documented, much of the published literature has explored the expression and function of these neuroprotective factors in anatomical locations associated with learning and memory165,174, sensory processing180 and synaptic plasticity, such as the hippocampus, limbic system, and olfactory bulb. Only recently have studies begun to examine the functional relevance of NPAS4 in the cortex, specifically following ischemic stroke. Npas4 was shown to promote neuroprotection following ischemia, as Npas4 knockdown was associated with decreased cell viability196. While investigations of cortical NPAS4 expression are emerging, very few studies have examined ARNT2 expression in the cortex. Investigating the cortex is relevant to MS as early axonal damage and neuronal loss are observed in this region9,10. Cortical involvement has also been associated with disease progression as cortical damage worsens in the progressive stages of MS and contributes to the cognitive deficits observed in individuals living with MS231. Therefore, we sought to characterize the in vitro expression patterns of NPAS4 and ARNT2 by examining the effects of different inflammatory mediators and cells relevant to MS pathogenesis on ARNT2 and NPAS4 protein expression. Using primary murine neuron-enriched cortical cultures, we saw minimal NPAS4 expression in untreated cultures, with very low NPAS4 MFI and percentage of NPAS4-expressing cells following PBS treatment. Glutamate exposure rapidly increased both the percentage of NPAS4-expressing cells and NPAS4 MFI at 77 doses as low as 1 and 3µM, respectively. NPAS4 was also upregulated following prolonged H2O2 exposure, as shown by increases in the MFI and percent of positive cells, but only at the two highest tested doses. At these two highest doses of glutamate, a decrease in the NeuN+ cell counts was also observed over time, which may have indicated a reduction in cell viability and increased toxicity. Conversely, steady state conditions showed that over 95% of cells were ARNT2-expressing, with the large majority (~70%) being low- or medium-ARNT2 expressing. While glutamate exposure did not affect ARNT2 levels, the highest tested dose tended to decrease ARNT2 MFI and percentage of expressing cells throughout the 9 hours. These results may also have been associated with the changes in cell viability observed over time, as shown by a reduction in NeuN+ cell counts. H2O2 exposure, however, did decrease ARNT2 expression levels with changes in intensity preceding the complete loss of ARNT2 expression, with the latter coinciding with time points at which a decrease in NeuN+ cells was observed. The changes observed in cell morphology and cell counts may have also been an indicator of increased toxicity at higher doses of H2O2 and changes in ARNT2 results must be interpreted with caution. Lastly, resting or activated CD4+ T-cells were insufficient to alter NPAS4 levels unless present at a minimum of a tenfold higher ratio than neurons. Despite a trend towards increased NPAS4 expression, no changes in ARNT2 expression were observed in the presence of CD4+ T-cells. Ultimately, these results suggest inflammatory mediators such as glutamate and H2O2 can alter the steady-state levels of NPAS4 and ARNT2 expression in primary murine cortical cells. 4.2 NPAS4 and ARNT2 in the Cortex Previous studies examining NPAS4 have largely focused on the hippocampus due to its well-characterized nature and NPAS4’s functional roles in learning and memory. This study aimed to examine expression patterns of NPAS4 in an in vitro cortical cell model. While this 78 study adds to the recently growing literature examining NPAS4 in the cortex, it is one of the first to examine the effects of inflammatory-like settings on cortical NPAS4 expression. We focused on the cortex as it is particularly impacted in individuals living with MS. Cortical involvement has been detected early in the disease course, and subsequent cortical damage has been associated with disease progression. Therefore, while previous studies mostly took a more focused approach by examining a specific anatomical region or cell type (such as the striatum, hippocampus, or even a cell line), we used a broader approach by using a mixed cortical culture to evaluate different subtypes of cortical cells in an attempt to model the heterogeneity observed in the cortex. Our mixed cortical cultures included a mixture of neurons carrying different neurotransmitters, including glutamatergic, GABAergic, cholinergic, catecholaminergic and dopaminergic neurons233. They also may have included different subtypes of excitatory (i.e., L2/3 IT, L4/5 IT, L5 IT, L6 IT) and inhibitory neurons (i.e., parvalbumin, somatostatin and vasoactive intestinal peptide)234. We felt cell lines represented a less physiologically relevant/often immortalized or oncogenic neuronal population. In particular, our in vitro model is able to capture the diversity in the cortex of neuronal populations specifically for evaluation by immunocytochemistry at the cellular level. At DIV 10, our cortical cells showed minimal NPAS4 expression in untreated conditions, with less than 10% positive cells. This observation was similar to previous investigations in murine hippocampal neurons, in which NPAS4 expression was not detectable in DIV 7 neurons161. In preliminary studies examining DIV 14 cortical cultures (data not shown), we also found no changes in NPAS4 expression; this was unlike DIV14 hippocampal cultures in which increased NPAS4 expression was observed and believed to be associated with synapse formation and maturation161. One potential explanation for this difference is that synapse formation and 79 maturation occur at different rates in these two cell types. Another explanation for this difference is that increased stressors in the culture media due to longer culture times and less frequent media changes may differentially impact hippocampal and cortical cells since most cell culture media contain antioxidants and nutrients that minimize stress in neuronal cultures. These results are also consistent with in vivo observations, where Npas4 mRNA levels were found to be low in cortical regions of healthy mice, including the prefrontal235, cingulate and piriform cortex236. Those mice also had negligible basal levels of Npas4 in the dorsal and ventral hippocampus, as well as the amygdala235. Protein levels were consistent with mRNA expression, with low protein expression observed in the frontal and piriform cortex of healthy rats174. In contrast to the limbic regions (i.e. nucleus accumbens and ventral pallidum), where NPAS4-positive cells were absent, the cortical regions exhibited a sparse distribution of these positive cells, with a slightly higher prevalence in the frontal cortex174. The presence of a small population of positive cells is similar to what we observed in our cultures, where approximately 10% of cells were NPAS4-positive. The cortex is composed of different regions with specialized functions and specific neuronal populations to help support these functions, thus NPAS4 expression is not uniform throughout the cortex. In our collection of cortical neurons, it is possible that different neuronal subtypes are more prone to expressing NPAS4 either due to differences in their basal neuronal activity, spontaneous synaptic activity, ongoing development and maturation or specialized roles. The low NPAS4 levels we observed were expected since NPAS4 is an IEG selectively induced by Ca2+ influx predominantly in neurons161. In the absence of a stimulus (i.e., neuronal activity), de novo transcription of Npas4 does not occur, and basal levels remain very low. Therefore, while these results are not novel, the similarities in basal NPAS4 expression in our murine primary neuron-enriched cortical cultures and previous in vivo 80 and in vitro results suggest these cultures are a suitable model to examine mediators that may alter NPAS4 in inflammatory degenerative settings relevant to MS pathogenesis. ARNT2 expression studies have primarily focused on its expression in the hypothalamus, given its role in hypothalamic development and function. Despite ARNT2 being ubiquitously expressed in neurons throughout the CNS, few studies have examined ARNT2 in the cortex. At DIV10, steady-state untreated conditions resulted in less than 5-10% of cells being ARNT2-negative, with roughly 70% of cells expressing low- to medium-ARNT2. This constitutive expression of ARNT2 has previously been reported in rat cortical neurons, where over 15% of cortical cells were found to be ARNT2-negative, and ~60% were found to be low- or medium-ARNT2-expressing1. In vitro, ARNT2 expression has also been previously reported in non-neuronal cells, specifically astrocytes1,237 and oligodendrocytes212. While we did not specifically assess oligodendrocytes, we did examine brightly expressing NeuN-ARNT2+ cells, several of which were confirmed to be astrocytes (data not shown). In vivo, investigations of ARNT2 in healthy mice provide further evidence of its constitutive expression. Arnt2 mRNA was found to be ubiquitously expressed in the large pyramidal neurons of the cerebral cortex211. Constitutive Arnt2 mRNA expression was also observed in the Purkinje cell layer of the cerebellum211 as well as the dentate gyrus and CA1-3 regions of the hippocampus211,238. Similar to Npas4, Arnt2 mRNA was not uniformly expressed throughout the cortex or the hippocampus, with variations in intensity levels observed, further matching the staining intensity variations observed in our cultures. Once again, these results are not novel and match what is previously known about ARNT2 expression patterns. ARNT2 is a member of the class II bHLH-PAS family, meaning that it is ubiquitously expressed throughout the CNS160. However, while the large majority of 81 cells express ARNT2, the heterogeneous level of protein observed in our cultures is consistent with that described both in in vitro and in vivo studies. 4.3 The Influence of Glutamate on NPAS4 and ARNT2 Expression Following exposure to glutamate, changes in the percentage of NPAS4-expressing cells (either distinguished as positive or as different intensity categories) appeared to precede changes in the MFI. The MFI examines the bulk population and, therefore, requires a small increase in many or large changes in fewer cells to result in a significant difference. Examining the percentage of expressing cells can better discern differences in a heterogeneous population of cortical cells. Given the expected diversity of neuronal subtypes in our culture (inhibitory, excitatory), we would similarly expect that cells would have varying responses to glutamate and thus capacities to upregulate NPAS4. In contrast, KCl induced NPAS4 expression in over 95% of NeuN+ cells. KCl is often used to drive global depolarization of all neurons, acting directly on voltage-gated Ca2+ channels (VGCC) to enable Ca2+ influx into the cell239. This influx then triggers secondary signaling pathways, including the induction of NPAS4 in various cells. Conversely, glutamate is an excitatory neurotransmitter that is tightly regulated in the nervous system due to its importance for cellular function and its excitotoxic nature. In vivo, findings showed that extracellular glutamate basal levels are tightly regulated and maintained between 2-4µM231. Glutamate also exerts its effects by triggering an influx of Ca2+ into the cell, however, glutamate does so by binding to receptors on the postsynaptic neuron. Ca2+ influx primarily occurs through NMDA receptors and, to a lesser extent, through α-amino-3-hydroxy-5-methylisoxazole-4-propionate (AMPA) or kainite receptors240. While NMDA receptors are widespread throughout the cerebral cortex, not all neurons express these receptors241. The type and density of glutamate receptors are 82 also neuron-specific, with some neurons expressing more AMPA or kainite receptors based on their function242. Therefore, KCl is capable of causing generalized depolarization in any cell with VGCCs, leading to a broader induction of NPAS4 in most cells. Glutamate’s effects are more selective due to its reliance on the presence and activation of specific glutamate receptors, which are not uniformly expressed in all cells. Changes at all tested doses of glutamate occurred rapidly within 1-hour of exposure. This rapid upregulation in NPAS4 expression may potentially be explained by fluctuations in intracellular free Ca2+ levels. Previous experiments have shown that primary neuronal cultures exposed to 250µM glutamate for 50 minutes experienced an initial rapid increase in intracellular free Ca2+ concentration ([Ca2+]), observed 5-10 minutes post-exposure90, followed subsequently by a more considerable secondary increase in [Ca2+]243-248. While we did not assess doses as high as 250µM, other studies have shown that doses as low as 5µM were also able to increase intracellular [Ca2+] in the soma and dendritic processes of rat neocortical cells249. Previous studies have also shown that calcium chelation (using EGTA) prevents NPAS4 induction following exposure to KCl161 or 30µM glutamate (Quandt lab, unpublished data). The prevention of glutamate-induced NPAS4 expression following calcium chelation suggests that glutamate may be driving NPAS4 upregulation through its direct effect on glutamate receptors that trigger a Ca2+ influx, which may be occurring at all tested doses of glutamate. Glutamate was previously shown to increase Ca2+ levels in a dose-dependent manner until 10µM, at which there was no further change in the Ca2+ levels249. It is plausible the direct effects of glutamate on its receptors on certain cortical cells may have resulted in Ca2+ influx into the cell, which could explain the increase in NPAS4 protein observed at all doses of glutamate; knowing definitively would require the specific measurement of intracellular Ca2+ levels. 83 At low doses of glutamate, we found NPAS4 levels decreased over time, with expression returning to baseline by 7-9 hours. This change over time observed at low doses could be due to the rapid and transient nature of NPAS4 expression161, which is only induced for short periods following Ca2+ influx to maintain circuit homeostasis. This transient upregulation matches previous findings that examined the short half-life of Npas4 mRNA178. Following a 2-hour KCl stimulation, NPAS4 protein levels were reduced by 86% after only the first hour post-KCl treatment. This short half-life allows cells to quickly adapt to changes and leads to a more transient and tightly regulated protein expression. The rapid degradation of Npas4 mRNA was shown to occur via posttranscriptional modifications250, regulated by microRNAs that target the 3’ untranslated region to negatively regulate Npas4 mRNA stability251. Additionally, mechanisms are in place to ensure that glutamate does not remain in the synapse for extended periods due to its excitotoxic nature. It is possible that at these low doses of glutamate, glutamate transporters present in glial cells and the presynaptic terminals of neurons, such as glutamate/aspartate transporter (GLAST) in mice, uptake the glutamate and convert it to glutamine252. Removal of glutamate from the synaptic space prevents it from being able to bind to glutamate receptors to further excite neurons. In this manner, diminished levels of glutamate or its removal from the extracellular synapse (which would increase transcription and, therefore, RNA levels), as well as degradation of Npas4 mRNA, may each explain the reduced levels of RNA and protein. While NPAS4 levels did decrease over time at the two highest tested doses, there was a distinct population of cells that were NPAS4+ several hours later, even with a decrease in cell counts. At high doses of glutamate, glutamate clearance was possibly impaired due to transporter saturation and could have resulted in glutamate receptor activation and continued intracellular 84 Ca2+ influx253. Another potential explanation could be linked to the “delayed calcium overload” or the secondary larger increase in intracellular Ca2+ levels following glutamate exposure243. Previous reports found that this “delayed calcium overload”243 occurred 30 minutes before cell death and was thought to be associated with the irreversible damage caused by glutamate excitotoxicity, leading to a cascade of effects (i.e., ROS production, mitochondrial stress) that result in neuronal death254-256. It is possible that a secondary influx in Ca2+ levels may be restimulating de novo Npas4 transcription, which could be why some viable cells remain NPAS4+, but this would require assessment with the introduction of calcium chelators or by measuring intracellular Ca2+ levels at different time points of glutamate exposure. Furthermore, the decrease in cell viability observed at the highest doses of glutamate may also potentially explain the inability of the cells to return to baseline NPAS4 levels. The doses typically used to induce rapid excitotoxicity in in vitro cultures tend to be greater than 100µM257, however, extended exposure to lower doses also resulted in excitotoxicity258 and may explain the observed cell death. Furthermore, while there are several pathways to death following glutamate excitotoxicity (i.e., apoptosis, necrosis, etc.)259,260, the observed visual changes where cells were lost suggest a rupturing of the cell membrane. Following the loss of cell membrane integrity, the intracellular contents of the neurons would be released to trigger an inflammatory response in the remaining viable cells. These could include soluble mediators such as ATP, protons, ROS or more glutamate that could potentially increase intracellular Ca2+ levels and upregulate NPAS4 expression. We found glutamate exposure did not significantly affect ARNT2 expression, however, expression tended to be lower with 30µM glutamate treatment at all time points. While the trends in the MFI and the percentage of ARNT2-expressing cells were similar, the percentage 85 of ARNT2-expressing cells showed a larger reduction in ARNT2 expression compared to the MFI at all time points. Changes were also observed at 10µM, however, this was only evident when examining the breakdown of the different ARNT2 intensity levels. One potential explanation for why this change was not apparent for the MFI or the percent positivity is that only a few cells may have undergone changes, with only these few switching between “low” to negligible. Given the cell loss attributed to reduced viability observed at 30µM glutamate, it is difficult to fully discern whether the decreasing trend in ARNT2 is due to the loss of cells or some other mechanism associated with glutamate excitotoxicity. It may also point to various regulatory mechanisms that contribute to the stability of Arnt2 mRNA or posttranslational modifications that impact the ARNT2 protein. Regulation of Arnt by microRNAs has been described, pointing to the possibility that Arnt2 may also be regulated in a similar manner. These mechanisms may contribute to a decrease in the stability of Arnt2 mRNA following glutamate exposure, however, this requires further investigation. 4.4 Effects of H2O2 on NPAS4 and ARNT2 Expression Unlike glutamate, we found that NPAS4 expression was upregulated in response to H2O2 exposure but only following exposure to 50 and 100µM. However, while the same MFI expression levels were seen for the 50 and 100µM, the percent positivity could better discern expression differences between these two doses. The percent positivity seems better able to detect subtle changes in expression within small or possibly specific subpopulations. It is also better at detecting low-level expression within a small number of cells, which the average MFI values would otherwise dilute. It is possible that at 50 and 100µM, the overall expression levels 86 were similar, but 100uM may have caused a minor change in a larger number of cells, which would have been easily detected by changes in the percent of positive cells. Changes in NPAS4 expression following H2O2 exposure occur later than changes following glutamate exposure. This may be explained by the delayed Ca2+ influx observed following treatment with different concentrations of H2O2. In a previous report, exposure to 30µM H2O2 was insufficient to cause changes in intracellular Ca2+ levels after 3 hours of exposure, yet 100µM H2O2 resulted in a noticeable increase between 2-3 hours of exposure261. This timeline aligns well with that observed in our study of changes in NPAS4 expression at both 50 and 100µM with a shift in the number of medium to high expressing cells by 3 hours. Increases in intracellular Ca2+ levels following H2O2 exposure have also been observed in multiple other cell types, including HeLa cells262, pancreatic acinar cells263 , and myocytes264. Therefore, it is possible that the delayed response following exposure to H2O2 may be due to a delay in intracellular Ca2+ influx. This agrees with unpublished data from our lab showing that the H2O2-induced NPAS4 upregulation observed at 4 hours in rat neuronal-enriched cultures could be prevented using calcium chelators such as EGTA. However, further assessments are required to examine changes in intracellular Ca2+ over time following H2O2 exposure to be able to correlate them with changes in NPAS4 expression. Alongside this delayed effect, H2O2 exposure did not upregulate NPAS4 expression to the same extent as glutamate. Following treatment with H2O2, fewer cells became NPAS4+, and not as many were medium to very-high NPAS4-expressing cells. While glutamate is a tightly regulated neurotransmitter that directly acts on receptors to allow Ca2+ entry into the cell, Ca2+ influx caused by H2O2 exposure is mainly secondary to the activation of oxidative stress pathways. One study found that H2O2 promotes Ca2+ release in rat hepatocytes by opening ATP-87 sensitive, non-specific cation channels as a result of mitochondrial stress and reduced intracellular ATP levels265. Micromolar concentrations of H2O2 mainly resulted in agonist-sensitive calcium release via activation of receptors on the endoplasmic reticulum, while millimolar concentrations triggered mitochondrial calcium release263. Therefore, H2O2 may not drive NPAS4 to the same degree as glutamate because of their distinct roles in cellular stress responses. H2O2 is a signaling molecule that modulates stress responses, apoptosis and antioxidant defense, while glutamate is involved in the direct modulation of synaptic activity. Glutamate’s direct effects on calcium influx make it a more potent inducer of NPAS4. Fewer cells respond to H2O2 because it may not robustly activate these calcium-dependent pathways in most cells, particularly when culture media may have high levels of antioxidants to combat oxidants or different cell types may have different antioxidant capacity levels. This resistance to oxidative stress may explain why more than 50% of cells remained NPAS4-negative. Additionally, the delayed effects of H2O2 exposure on increased NPAS4 expression may also point to an indirect manner of NPAS4 induction. Visual assessments of cell cultures over time suggested that high doses of H2O2 impacted glial cells, and these cells were often lost by 3 hours of treatment. The death of glial cells may have resulted in the release of glutamate, ATP, or other mediators that could be driving NPAS4 expression in neuronal cells. While H2O2 exposure (i.e., 50 and 100µM) upregulated NPAS4 as early as 3 hours post-exposure, expression was still observed in ~25-40% of the population from 5-9 hours, with no major fluctuations in the amount of very high-expressing cells. Unlike glutamate, which must be removed from the extracellular space, H2O2 undergoes natural decomposition, which may occur slowly over time in the absence of an enzyme266. Therefore, the presence of H2O2 could be continuing to activate oxidative stress pathways, potentially maintaining high intracellular levels 88 of Ca2+. Furthermore, 100µM H2O2 was previously shown to result in apoptosis-induced cell death 3 hours post-exposure261. H2O2 has also been shown to cause cell death through autophagy and necrosis, with the resultant cell death pathway highly dependent on the concentration of H2O2267. This also matches our results where membrane swelling and blebbing were observed visually at 100µM from 5 hours onwards. While the specific cell death pathway was not examined, rupturing of the cell membrane may indicate the release of intracellular contents that might also increase intracellular Ca2+ levels independently of H2O2. When examining the effects of H2O2 on ARNT2 expression, we found exposure to 100µM significantly decreased ARTN2 expression, with lesser effects seen in response to 50µM. At both concentrations, the decrease in ARNT2 MFI was observed a few hours before the decrease in the percentage of ARNT2-expressing cells. Our results suggest that following exposure to oxidative stress, cells experience a gradual decrease in ARNT2, where ARNT2 levels might be slowly declining before losing ARNT2 expression completely. Previous investigations showed that short exposure to 25, 50 and 100µM H2O2 upregulated ARNT2 protein expression in rat cortical neurons, as shown by western blots, up until 2-3 hours post-exposure1. These results were also observed immunocytochemically, in which an increase in ARNT2 protein expression was observed in DAPI+MAP2+ neuronal cells following 1-hour exposure to 100µM H2O21. In contrast, cells in this study showed no upregulation in ARNT2 protein expression at any assessed time points. One potential explanation for this difference is the protein detection method. While western blots can quantify protein levels within a sample, they do not provide valuable information about cell-specific protein expression. Therefore, this analysis would not have excluded ARNT2 from contaminating glial cells such as astrocytes1 or oligodendrocytes212. A cell-specific analysis is 89 invaluable to distinguish cell types, and flow cytometry was considered. Flow cytometry, however, inevitably means some damage to adherent cells as they are harvested by trypsin or otherwise. Our observations suggest that even the slightest damage to cells may directly or indirectly induce the expression of NPAS4 while also not allowing for the distinction between nuclear or cytoplasmic localization. Our immunocytochemistry analysis, however, enabled a focused evaluation of NeuN+ neuronal cells. 4.5 NPAS4 and ARNT2 as Potential Mediators of Cell Viability Treatment with high doses of glutamate and H2O2 both tended to reduce the number of neuronal cells in these cultures. Despite this decrease, there was a noticeable number of viable NPAS4-expressing cells that remained following prolonged exposure to high doses of glutamate and H2O2. While this may be a result of a delayed Ca2+ influx caused by several factors, as mentioned previously, it may also suggest a potential link between NPAS4 and neuroprotection. One explanation is the relationship between NPAS4 and BDNF expression, the latter of which is a key neurotrophin involved in various aspects of neuronal survival, development and plasticity (as mentioned in section 1.1.4). NPAS4 was previously shown to upregulate BDNF expression in both cortical161 and hippocampal CA3 pyramidal neurons183. Additionally, changes in Bdnf RNA levels were found to mimic changes in Npas4 RNA levels in chronic active EAE mice1 (Figure 1.2C). Downstream genes other than Bdnf may also be mediating this neuroprotective effect. One previous study found that NPAS4-induced Gem expression following ischemia was able to promote neuroprotection via its action on L-type VGCCs268. This prevented excessive Ca2+ influx into the cell, promoting neuroprotection in excitotoxic conditions. Therefore, it is possible that NPAS4 upregulation in some cells helps to protect against cell death via the activation of downstream neuroprotective genes such as Bdnf. This link must be further assessed by 90 examining associated changes in BDNF protein expression when exposed to these inflammatory mediators. Given the loss of high to very high NPAS4-expressing cells and the increase in NPAS4-negative cells over time, this may also indicate the preferential loss of high NPAS4-expressing cells, pointing to potential detrimental effects of sustained NPAS4 expression. While NPAS4 expression has typically been associated with neuroprotection, several studies have also examined its detrimental effects. NPAS4 overexpression in the basal ganglia following intracerebral hemorrhage (ICH) not only promoted neuronal cell death but was also found to increase concentrations of proteins that promote cell pyroptosis, including the NLRP6 inflammasome, cleaved caspase-1 and cleaved caspase-11269. NPAS4 overexpression also resulted in cell death in both HeLa cells and cultured cerebellar granular neurons, in which the number of viable cells declined over 3 or 5 days of Npas4 overexpression, respectively167. Therefore, NPAS4 overexpression may also be linked to changes in cell viability as the potential sustained NPAS4 induction could be identifying those cells about to die. Determining the likelihood of NPAS4 being neuroprotective or detrimental relies largely on determining whether or not NPAS4 induction precedes cell death at the individual cell level and will involve an assessment of changes in NPAS4 over time within individual live cells to be able to examine whether the “ever or still” NPAS4-expressing or “always negative” negative population is being lost. ARNT2 has been linked to cellular health and viability in studies where siRNA-mediated Arnt2 knockdown drove apoptosis in PC-12 cells209. Furthermore, treatment with 50µM H2O2 reduced ARNT2 expression levels in rat cortical neurons, which preceded increases in cleaved caspase-3 staining1. Apoptosis may also be occurring in our cultures, where a reduction in 91 ARNT2 protein expression was observed prior to cell death following exposure to high doses of glutamate and H2O2. At high doses of glutamate or H2O2, we observed the presence of numerous ARNT2-negative cells. Although we were unable to pinpoint the exact timing of ARNT2 protein expression loss in these cells, it is possible that the complete reduction of ARNT2 could serve as a prognostic indicator of impending cell death. These observations align with previous studies that demonstrated complete ARNT2 downregulation preceded ischemia-induced cell death and oxidative stress-induced cell loss in PC-12 cells209. Similar to examining roles for NPAS4 in cell survival, an examination of temporal changes in ARNT2 or targeted induction or knockdown with an examination of associated changes in cell viability are essential to further investigate this question. 4.6 Effects of Inflammatory Cells on NPAS4 and ARNT2 Expression Because Npas4 and Arnt2 were both increased prior to the onset of clinical symptoms in the CNS, we postulated that inflammatory cells might be responsible for those changes. While changes in NPAS4 expression were subtle in the presence of a 5:1 ratio of T-cells to neurons, increasing the number of resting and activated T-cells was enough to increase NPAS4 expression by 3-4-fold at 2 hours post-incubation. CD4+ T-cells play a key role in MS pathogenesis by releasing inflammatory soluble mediators, such as proinflammatory cytokines, including IFN-, interleukin (IL)-6, IL-17A and IL-17F270. Several proinflammatory cytokines produced by T helper-1 or Th1 cells, such as interferon-gamma (IFN-γ) or TNFα, were found to elevate calcium levels in rodent islet cells in a dose-dependent manner271,272. Another T helper subtype, Th17 cells that produce significant amounts of IL-17, were found to increase intra-axonal Ca2+ levels through direct cell-cell contact with neurons and the release of glutamate into the extracellular 92 space, resulting in glutamate toxicity89. As naïve cells also resulted in NPAS4 expression, it is unlikely that they would rapidly produce these cytokines in abundance, and cytokines are not the likely mediators but would have to be tested. Previous studies have also shown that CD8+ T-cells can alter the electrical excitability of cells via the release of perforin273, which results in the formation of voltage-independent polyperforin channels. These channels enable non-selective ion fluxes into the cell and could potentially cause membrane depolarization. Therefore, based on previous reports demonstrating the effects of different immune cells on ion flux into the cell, the observed changes may not be specific to CD4+ and may also result from CD8+ T-cell interactions. Given the similar effects of increased resting and activated T-cell amounts on NPAS4 expression, it is also possible that plating any type of immune cell at a high ratio compared to neurons could potentially alter ion homeostasis and cause changes in NPAS4. No noticeable changes were observed in ARNT2 expression even when increasing amounts of resting and activated CD4+ T-cells were added to the neurons. ARNT2, like other bHLH-PAS transcription factors, has been shown to autoregulate its transcription202. Via a positive feedback loop, ARNT2 constantly works to make sure that a baseline level of expression is maintained. Therefore, the number of inflammatory cells or the amount of soluble mediators being released may be insufficient to overcome this autoregulation and disrupt expression patterns. The steady ARNT2 expression may also explain why no visual changes in cell viability were observed following neuronal exposure to resting or activated CD4+ T-cells. 4.7 Global Assessment of Combined Changes in NPAS4 and ARNT2 Given that ARNT2 is thought to be NPAS4’s primary binding partner in the CNS, we might expect to see similar trends for these two proteins in the presence of inflammatory mediators and cells. However, a cursory qualitative review showed this was not generally the 93 case. Despite early NPAS4 increases following glutamate exposure at all doses, a decreasing trend in ARNT2 expression was only observed at 30µM. While neurons exposed to 50 and 100µM H2O2 showed increased NPAS4 from 3 hours onwards, ARNT2 expression was reduced at these same time points. These opposing trends might suggest that the likelihood of these two proteins engaging in inflammatory environments is low and was supported by a number of NPAS4+ cells after several hours of H2O2 exposure, which showed no ARNT2 expression. The two factors may also exert functions independently of each other. NPAS4 has been shown to form functional heterodimers with ARNT164 in the CNS and may partner with ARNT instead of ARNT2 in these settings. Furthermore, ARNT2 also has other functional binding partners, such as hypoxia-inducible factor 1α (HIF-1α)213 and single-minded homolog 1 (SIM1)274. ARNT2 is also capable of homodimerizing to exert its effects. Therefore, it is possible that ARNT2 also relies on other binding partners or on itself in inflammatory settings. However, more work is required to assess the functional binding partners of NPAS4 and ARNT2 in inflammatory settings. Recent studies elucidated a role for ARNT2 in suppressing activity-dependent gene transcription, therefore maintaining low basal levels of inducible genes215. It was found that in the absence of neuronal activity, ARNT2 recruits nuclear receptor co-repressor (NCoR2), a transcriptional coregulatory protein that enables the recruitment of histone deacetylases275, to activity-dependent regulatory elements in much the same way it recruits NPAS4. The ARNT2/NCoR2 complex then works to recruit other co-repressors to restrict inhibitory synaptic plasticity and prevent NPAS4 induction. This process helps to maintain circuit homeostasis by preventing the aberrant induction of genes and may relate to our findings at low doses of H2O2 (i.e., 12.5 and 25µM) or to prolonged exposure to low doses of glutamate (i.e., 1 and 3µM). In 94 these settings, ARNT2 expression may play a role in maintaining low basal levels of NPAS4 via the ARNT2/NCoR2 complex. Alternately, in the presence of neuronal activity or stimulation, ARNT2 recruits NPAS4 to induce transcription, resulting in increased somatic inhibition215. Following stimulation, NCoR2 is also recruited to NPAS4-bound enhancers; however, it works to increase activity-dependent gene expression and may explain our findings at low doses of glutamate in which rapid NPAS4 upregulation at 1 and 3µM was observed alongside ARNT2 expression. Whether similar pathways are functioning at high doses of glutamate and H2O2 would require further investigation. 4.8 Limitations In this study, we utilized primary murine neuron-enriched cortical cultures, which represent a heterogeneous population of more than one neuronal cell type. To minimize any variability associated with differentially matured or proportions of specific neuronal populations, we used embryos at similar developmental stages and carried out experiments on cells at consistent DIV. However, the presence of glial cells and other non-neuronal contaminating cells posed a limitation in this study. Unlike neurons, these cells can proliferate and migrate to different regions of the well, particularly if recruited by soluble mediators. Glia can uptake glutamate to prevent excitotoxic effects and may have attenuated the observed effects. Glia may also respond to these inflammatory mediators to differing degrees and in different ways, potentially releasing their own BDNF or other protective factors to impact the neuronal milieu. Glia are responsive to glutamate and also H2O2 and may release proinflammatory cytokines or become less viable and release ATP that could drive neuronal NPAS4 or alter ARNT2. Despite these limitations, this served as an appropriate and physiologically relevant model of the cortex as it mimicked the in vivo environment in which complex cellular interactions occur between 95 glial and neuronal cells. If glia impacts are to be characterized, those cells could be isolated from the culture with measures to limit their growth. These cultures represent a simplified environment that may not accurately model the complex immune responses, signaling pathways and other cellular interactions that occur in vivo during MS. The inflammatory environment was also created using exogenous factors, such as soluble inflammatory mediators, resulting in an oversimplified representation of an MS-like inflammatory setting. While this model was useful for this study, it is also important to examine an in vivo model, such as an EAE mouse model, to examine NPAS4 and ARNT2 expression patterns and colocalization in healthy and model disease conditions. This study relied mainly on immunocytochemistry techniques performed on fixed cells, which capture only a snapshot in time. Although we assessed a range of doses and time points to gain a better understanding of the temporal and dose-dependent changes in NPAS4 and ARNT2 expression, we were unable to examine changes in live cells over time to give us a better idea of the kinetics of NPAS4 and ARNT2 protein expression within individual cells. Our approach and initial studies enabled us to detect only four proteins reliably due to the number of fluorophores and filters available on this microscope; we focused on NPAS4 and ARNT2 in nuclei of neuronal cells. The signal limitation meant we could not distinguish specific neuronal subtypes (i.e., inhibitory versus excitatory) or examine changes in viability with fluorophores, and it would be an important line of investigation in future studies. 4.9 Future Directions and Conclusions Overall, this study examined NPAS4 and ARNT2 protein expression in primary neuron-enriched cortical cultures and has provided further insight into the different inflammatory mediators that may influence the expression of these proteins associated with 96 neuronal function and protection. As hypothesized, we found that glutamate and H2O2, two inflammatory mediators known to cause damage in MS, altered in vitro NPAS4 and ARNT2 protein expression. While the CD4+ T cells we studied agreed with the hypothesis and increased NPAS4 levels, changes in ARNT2 expression were not altered by those immune cells at the doses studied. To fully characterize the effects of these two proteins in chronic inflammatory degenerative settings, additional questions and approaches to address them should be explored. Inflammatory cells relevant to MS pathogenesis can exert their effects by releasing soluble inflammatory mediators, including proinflammatory cytokines and glutamate, the latter of which we have already interrogated directly. These primary neuron-enriched cortical cultures can be utilized to test the direct effects of cytokines typically released by resident and infiltrating cells on NPAS4 and ARNT2 expression. This would include an assessment of proinflammatory cytokines implicated in MS and EAE pathogenesis, including IL-1, IL-17A, IFN-, TNF- and granulocyte-macrophage colony-stimulating factor (GM-CSF) at different doses and time points to examine changes in RNA and/or protein expression via qPCR and immunocytochemical methods. Additionally, given the desire to characterize the neuroprotective properties of NPAS4 and ARNT2, the association between expression of these factors and cell viability and function should further be examined. Supernatants were collected from all neuronal cultures co-incubated with glutamate and H2O2 with the goal of examining lactate dehydrogenase (LDH) release as a signal of cell death in association with NPAS4 or ARNT2 expression. While this provides a global assessment of cell viability from one treatment to another, it is not a cell-specific assay, and outcomes associated with expression of either factor specifically would not be apparent. Therefore, viability can also be examined using Calcein and propidium iodide to differentiate 97 between live and dead cells, respectively. Calcein-AM is a non-fluorescent membrane permeable compound, which, upon entry into live cells, gets converted by esterase activity into Calcein, a fluorescent dye that remains trapped within the cytoplasm of live cells276. Propidium iodide is membrane-impermeable, meaning it can only penetrate cells with compromised membranes (i.e., dead or dying cells) and binds to DNA, producing autofluorescence277. This method, in conjunction with immunocytochemical techniques, can allow for staining of neuronal cells and ARNT2 and/or NPAS4 expression and may provide an indication of the association between NPAS4 and ARNT2 expression to cell viability. NPAS4 and ARNT2 expression were examined in response to inflammatory mediators and cells, but the levels of each have not yet been determined at the individual cell level. Colocalization of the two proteins following different stimuli would inform the likelihood that these proteins are acting in combination, independently, or with alternate partners. A co-localization analysis to examine NPAS4 and ARNT2 intensity levels within the same cell is the next analysis to undertake. One potential method is to use Pearson’s correlation coefficient to measure the linear correlation between the intensity distributions of NPAS4 and ARNT2. NPAS4 and ARNT2 will also be broken down into different intensity categories, as previously done, to examine the combined changes in the same cell. One of the major downstream targets of NPAS4 and ARNT2 heterodimerization is BDNF. To better elucidate the neuroprotective properties of both NPAS4 and ARNT2, BDNF production should also be examined. The neuronal cell culture supernatants collected from the glutamate and H2O2 experiments can be examined using a BDNF enzyme-linked immunosorbent assay (ELISA). BDNF protein levels will then be correlated to NPAS4 and ARNT2 levels to examine how levels have changed in response to different soluble inflammatory mediators. 98 However, BDNF can also be produced by glial cells, therefore, it would be hard to distinguish the true source of BDNF expression using the ELISA. Future experiments could also include an immunocytochemical examination of BDNF, NPAS4 and ARNT2 protein levels that would enable analysis at the cellular level. This would provide an indication of how these three proteins are altered within the same cell and can allow us to differentiate between neuronal and glial BDNF production. The above studies are largely associative and do little to characterize the functional roles and mechanistic pathways impacted by loss or overexpression of NPAS4 and/or ARNT2. The impact of Npas4 and/or Arnt2 depletion on cell viability in the inflammatory settings we studied is best examined using murine primary neuron-enriched cortical cultures derived from Npas4 knockout mice (KO) or via siRNA-mediated knockdown of Arnt2, as we described previously212. These cells will be exposed to glutamate, H2O2 and inflammatory cells to monitor changes in neuronal morphology and viability as described above (i.e., LDH or Calcein/propidium-iodide staining). RNAseq can identify the gene pathways disrupted or enhanced by either of these factors and serve to identify downstream targets that might be considered for novel therapeutic intervention strategies. Similarly, the impact of NPAS4 overexpression should also be examined to determine whether prolonged expression is harmful to cells and the potential pathways involved. This would best be performed using lipid-based approaches or through the use of AAV-viral vectors. 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J Microbiol Methods 37, 77-86 (1999). https://doi.org/10.1016/s0167-7012(99)00048-2 119 Appendix: List of Specific and Isotype/Irrelevant Antibodies Used for Immunocytochemistry Primary Antibody Species Company City, Country Catalog Number Final concentration (µg/ml) NeuN Chicken Polyclonal (IgY) Millipore Burlington, US ABN91 1.0 ARNT2 Mouse Monoclonal (IgG2b) Santa Cruz Biotechnology Dallas, US sc-393683 0.4 NPAS4 Rabbit Monoclonal (IgG) Activity Signaling San Diego, US NP41-2 1.0 GFAP Chicken Polyclonal (IgY) Millipore Burlington, US AB5541 0.67 IBA-1 Rabbit Polyclonal (IgG) Wako Richmond, US 019-19741 0.5 NeuN Mouse Monoclonal (IgG1) Millipore Burlington, US MAB377 0.37 MAP2 Chicken Polyclonal (IgY) Abcam Cambridge, UK AB5392 1.27 Irrelevant antibodies/isotype controls (listed in order for primaries Species Company City, Country Catalog Number Final concentration (µg/ml) Chicken IgY Chicken R&D Systems Minneapolis, US AB-101-c 1.0 Mouse IgG2b Mouse Santa Cruz Biotechnology Dallas, US sc-3879 0.4 Rabbit IgG Rabbit R&D Systems Minneapolis, US MAB1050-500 1.0 Chicken IgY Chicken R&D Systems Minneapolis, US AB-101-c 0.67 Rabbit IgG Rabbit R&D Systems Minneapolis, US MAB1050-500 0.5 Mouse IgG1 Mouse Santa Cruz Biotechnology Dallas, US sc-3877 0.37 Chicken IgY Chicken R&D Systems Minneapolis, US AB-101-c 1.27 120 Secondary Antibody (listed in order as used for primaries above Company City, Country Catalog Number Wavelength Final concentration (µg/ml) Donkey anti-chicken IgY Jackson ImmunoResearch West Grove, US 703-545-155 488 3.0 Donkey anti-mouse IgG2 Invitrogen Waltham, US A10037 568 4.0 Donkey anti-rabbit IgG Invitrogen Waltham, US A32795 647 4.0 Donkey anti-chicken IgY Jackson ImmunoResearch West Grove, US 703-545-155 488 3.0 Donkey anti-mouse IgG2 Invitrogen Waltham, US A31571 647 4.0 Donkey anti-chicken IgY Jackson ImmunoResearch West Grove, US 703-545-155 488 3.0
Thesis/Dissertation
eng
Pathology and Laboratory Medicine
Vancouver : University of British Columbia Library
Attribution-NonCommercial-NoDerivatives 4.0 International
Graduate
Quandt, Jacqueline
Characterizing the neuronal expression of neuronal PAS-domain containing protein 4 and aryl hydrocarbon receptor nuclear translocator 2 in multiple sclerosis related inflammatory settings in vitro
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