Glucocorticoid Regulation of the G-Protein Coupled Estrogen Receptor (GPER) in Mouse Hippocampal Neurons

Home , Cannabinoid receptor, Cowell (surname), GPER

Kate Colleen Eliza Nicholson

A Thesis presented to The University of Guelph

In partial fulfilment of requirements for the degree of Master of Science in Biomedical Sciences

Guelph, Ontario, Canada

GLUCOCORTICOID REGULATION OF THE G-PROTEIN COUPLED ESTROGEN RECEPTOR (GPER) IN MOUSE HIPPOCAMPAL NEURONS

Kate Colleen Eliza Nicholson Advisor: University of Guelph, 2019 Dr. Neil J. MacLusky

The most prevalent estrogen, 17β-estradiol, binds the non-classical G-protein coupled estrogen receptor (GPER) with high affinity resulting in rapid activation of the c- jun N terminal kinase (JNK) pathway. GPER activation mediates some of the rapid neurotrophic and memory-enhancing effects of 17β-estradiol in the female hippocampus. However, exposure to stressful stimuli may impair these beneficial effects. This thesis characterizes neurosteroid receptor expression in murine-derived mHippoE cell lines that are subsequently used to investigate the glucocorticoid regulation of GPER protein expression and functional activation. This thesis demonstrates that 24-hour treatment with a glucocorticoid receptor agonist reduces

GPER protein expression and activation of JNK in female-derived mHippoE-14s. Using an in vivo model, treatment with glucocorticoids significantly reduces hippocampal activation of JNK in female ovariectomized CD1 mice. Collectively, this thesis uses in vitro and in vivo models to characterize glucocorticoid regulation of GPER expression and signalling in female murine hippocampal neurons.

ACKNOWLEDGEMENTS

To Dr. MacLusky: I would like to thank you for inspiring my passion for science and pursuit of knowledge. Over the past 2 years, you have provided me with countless opportunities to grow as a young researcher and I am tremendously grateful for this. Your ongoing support, guidance, and encouragement has given me the confidence to share my own thoughts, questions, and opinions with other researchers. You have challenged me to improve my scientific communication and critical thinking skills while inspiring enthusiasm for the implications of our research. This opportunity has been uniquely rewarding and I am so very grateful for your dedication as a mentor.

To my advisory committee members, Dr. Bettina Kalisch and Dr. Jon LaMarre: I would like to thank you for your guidance, support, and insight throughout the course of my MSc degree. I feel very fortunate to have been able to approach both of you for friendly advice and opinions in shaping the design and interpretations of this project.

To my MacLusky lab parents, Dr. Ari Mendell and Dr. Carolyn Creighton: I would like to sincerely thank you for your ongoing patience in training me while sharing your wisdom and mentorship throughout graduate school. Carolyn, I am very grateful for your kindness while introducing me to the foundations of lab work and research. Ari, thank you for supporting and challenging me to grow as a graduate student. You have always been the calm in the storm and I am incredibly grateful for your leadership and friendship throughout graduate school. I feel very fortunate to have worked alongside inspiring mentors and friends like you two.

To my MacLusky lab siblings: E-Marti and Lo-dawg, thank you for being my rocks through it all. Thank you for making fun of me when I deserve it and putting up with me when I don’t. Hayl-storm, Dry Rice Lawton, and E-Craig, thank you for all of the lab chats, support, and humour. To all other MacLusky lab members, thank you for making the long lab days seem shorter.

To the faculty and students of the Department of Biomedical Sciences: You have provided me with many rewarding opportunities to grow in and outside of the lab and classrooms. Thank you for making this a welcoming, enriching, and collaborative environment to learn in.

Finally, thank you to my family for all of the love, long phone calls, and support while cheering me on throughout the ups and downs of graduate school. Thank you for always reminding me to look at the bigger picture. To my friends, thank you for keeping me grounded with your love, encouragement, humour, and kindness.

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DECLARATION OF WORK PERFORMED

I declare that I have performed and completed all of the work presented within this Master of Science thesis, with the exception of the following:

All ovariectomy surgical procedures, subcutaneous drug injections, cervical dislocations, and brain extractions in Chapter 5 were performed by Emily Martin (Master of Science candidate supervised by Dr. Neil MacLusky and Dr. Elena Choleris,

Department of Psychology, University of Guelph).

All Sanger sequencing was performed by the University of Guelph’s Advanced

Analysis Centre Genomics facility.

TABLE OF CONTENTS

ABSTRACT ...... ii ACKNOWLEDGEMENTS ...... iii DECLARATION OF WORK PERFORMED ...... iv LIST OF TABLES ...... viii LIST OF FIGURES ...... ix LIST OF ABBREVIATIONS ...... xi LIST OF APPENDICES ...... xiv CHAPTER 1: INTRODUCTION AND REVIEW OF THE LITERATURE ...... 1 Sex Differences in Neuropsychiatric and Neurodegenerative Disorders ...... 2 The hippocampus………………………………………………………………………………..……...4 The Hypothalamic-Pituitary-Gonadal (HPG) Axis ...... 6 Steroid and Estradiol Biosynthesis...... ………………...………………………………………….8 Steroid Receptors ...... 13 Estrogen Receptors in the Hippocampus ...... 14 Functional Effects of Hippocampal Estrogens and GPER Activation ...... 20 Stress and the Hypothalamic-Pituitary-Adrenal (HPA) Axis…………………………………...28 Glucocorticoid Biosynthesis ...... 30 Glucocorticoid Receptor Activation in the Hippocampus ...... 31 Functional Effects of GR Activation in the Hippocampus ...... 31 Stress and Estrogen Interactions in the Hippocampus ...... 34 Rationale ...... 36 Hypothesis and Objectives ...... 37 CHAPTER 2: CHARACTERIZATION OF RECEPTOR MRNA EXPRESSION IN MHIPPOE-14 AND HIPPOE-18 IMMORTALIZED HIPPOCAMPAL NEURONS ...... 39 Introduction ...... 40 Experimental Procedures ...... 47 Cell Culture Conditions ...... 47 RNA Isolation ...... 48 Reverse Transcription and cDNA Synthesis ...... 48 Conventional Polymerase Chain Reactions and Visualization……………………..…………49 Results ...... 53 Discussion ...... 57

Conclusion ...... 63 CHAPTER 3: GLUCOCORTICOID REGULATION OF GPER PROTEIN STEADY STATE LEVELS IN MHIPPOE-14 AND MHIPPOE-18 IMMORTALIZED HIPPOCAMPAL NEURONS 65 Introduction ...... 66 Experimental Procedures ...... 68 Chemical and Stock Solutions ...... 68 Cell Culture Conditions ...... 68 Treatment and Protein Collection ...... 69 Western Blotting...... 70 Statistical Analyses ...... 71 Results ...... 72 GPER Protein Expression is Regulated by Dexamethasone Treatment ...... 72 Discussion ...... 745 Conclusion ...... 78 CHAPTER 4: GLUCOCORTICOID REGULATION OF GPER MEDIATED C-JUN N TERMINAL KINASE (JNK) PHOSPHORYLATION IN IMMORTALIZED HIPPOCAMPAL NEURONS ...... 79 Introduction ...... 80 Experimental Procedures ...... 82 Chemical and Stock Solutions ...... 82 Cell Culture Conditions…………...…………………………………………..…………………..82 G-1 Treatments ...... 83 24 hour Dexamethasone Pre-treatments ...... 83 Protein Collection ...... 83 Western Blotting...... 84 Statistical Analyses: Functional Activation of GPER Using G-1 Treatments ...... 85 Statistical Analyses: Glucocorticoid Regulation of GPER Functional Activation ...... 86 Results ...... 86 mHippoE-14 Cell Lines Express a Functional GPER ...... 86 Glucocorticoids Regulate G-1 Induced Functional Activation of GPER ...... 88 Discussion ...... 90 Conclusion ...... 92 CHAPTER 5: REGULATORY EFFECTS OF GLUCOCORTICOID TREATMENT ON GPER FUNCTIONAL ACTIVATION IN OVARIECTOMIZED FEMALE CD1 MICE ...... 93 Introduction ...... 94 Experimental Procedures ...... 96 Animals ...... 96

Ovariectomy Surgeries ...... 96 Chemical and Stock Solutions ...... 97 Treatment Groups ...... 97 Protein Preparation and Western Blotting ...... 98 Statistical Analyses ...... 100 Results ...... 101 Discussion ...... 104 Conclusion ...... 106 CHAPTER 6: OVERALL CONCLUSIONS AND FUTURE DIRECTIONS ...... 107 REFERENCES ...... 113 APPENDICES...... 137 Appendix 1: Sanger sequencing results for primer pairs that successfully amplified DNA fragments from mHippoE immortalized hippocampal neurons ...... 137 Appendix 2: Antibody information for western blotting experiments ...... 143

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LIST OF TABLES

Table 1: Cycling conditions for initial denaturation, denaturation, primer annealing, extension, and final extension stages of cPCR. ………………………………………...…50

Table 2: Primer sequences used to identify transcription factors and steroid, glutamatergic, and GABAergic mRNA receptors by conventional PCR. ………………..52

Table 3: Antibodies used in western blotting experiments (Appendices) ……………..143

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LIST OF FIGURES

Figure 1.1: Characteristic estradiol, progesterone, LH, FSH, and GnRH levels across the 4-5-day estrous cycle of female mice………………………………………….…………7

Figure 1.2: Steroid biosynthesis pathway showing synthesis and metabolism of progestogens, androgens, glucocorticoids, and estrogens.………………………………12

Figure 1.3: Cellular signaling cascades of membrane bound and intracellular estrogen receptor alpha (ERα), ERβ, and GPER..……………………………………………………20

Figure 1.4: Effects of behavioural testing on estradiol induced increases in spine synapse density in ovariectomized female rats. ………………………………………...…24

Figure 1.5: Dose dependent increase in spine density within the stratum radiatum of the CA1 region within 40 minutes of G-1 treatment in ovariectomized female mice……..…26

Figure 1.6: The hypothalamic-pituitary-adrenal (HPA) axis in response to a stressful stimulus. …………………………………………………………………………………..……29

Figure 1.7: Sexually differentiated effects of stress exposure on conditioned eyeblink responses in male and female rats.……………………………………………………….…33

Figure 2.1: Characterization of receptor mRNA expression for transcription factors and steroid, GABAergic, and glutamatergic receptors in mHippoE-14 and mHippoE-18 immortalized hippocampal neurons.…………………………………………………………55

Figure 2.2: Confirmation of correct amplicon sequencing with forward and reverse GPER primers.…………………………………………………………………………………56

Figure 3.1: Representative western blots depicting GPER protein expression in 10- minute, 1 hour, and 24 hour vehicle and DEX treated mHippoE-14 and mHippoE-18 cell lines, relative to α-tubulin loading control.…………………………………………..………73

Figure 3.2: Representative western blots depicting GPER protein expression in 10 hours and 48-hour vehicle and DEX treated mHippoE-14 and mHippoE-18 cell lines, relative to α-tubulin loading control…………………………………………………..………74

Figure 4.1: Representative western blots depicting JNK phosphorylation in vehicle and 10-minute, 1 hour, and 4 hour G-1 treated samples of a) mHippoE-14 and b) mHippoE- 18 cell lines, relative to β-actin loading control.………………………………………….…87

Figure 4.2: Representative western blots depicting JNK phosphorylation in 24-hour

ix vehicle and DEX pre-treated mHippoE-14 samples.………………………………………89

Figure 5.1: Western blot depicting JNK phosphorylation in ovariectomized female CD1 mice treated with 6 μg/kg of G-1……..……………………………………………..………102

Figure 5.2: JNK phosphorylation in the hippocampus of ovariectomized female CD1 mice treated with G-1 in the presence or absence of CORT………………………...….103

Figure 6.1. Confirmation of correct amplicon sequencing with forward and reverse primers for additional receptor mRNA targets (Appendices)…………….………………142

LIST OF ABBREVIATIONS

5HT1A Serotonin 1A receptor 17β-estradiol Estradiol 17β-HSD 17β-hydroxysteroid dehydrogenase 3β-HSD 3β-hydroxysteroid dehydrogenase 21-OHase 21-hydroxylase 11β-OHase 11β-hydroxylase 17α-OHase 17α-hydroxylase ACh Acetylcholine ACTH Adrenocorticotropic hormone AMPA α-amino-3-hydroxyl-5-methyl-4-ixoxazolepropionic acid AMPA-R2 AMPA receptor type 2 AMPA-R3 AMPA receptor type 3 AMPA-R4 AMPA receptor type 4 ANOVA Analysis of Variance AR Androgen receptor ATP Adenosine triphosphate BDNF Brain-derived neurotrophic factor BLAST Basic Local Alignment Search Tool BSA Bovine serum albumin CA1 Cornu ammonis 1 CA2 Cornu ammonis 2 CA3 Cornu ammonis 3 cAMP Cyclic adenosine monophosphate cDNA Complementary deoxyribonucleic acid CORT Corticosterone cPCR Conventional polymerase chain reaction CREB cAMP response element binding protein CRH Corticotrophin releasing hormone CRHR1 Corticotrophin releasing hormone receptor type 1 CRHR2 Corticotrophin releasing hormone receptor type 2 CS-FBS Charcoal stripped fetal bovine serum DEX Dexamethasone DHEA Dehydroepiandrosterone DHT Dihydrotestosterone DMEM Dulbecco’s modified Eagle medium DMSO Dimethyl Sulfoxide DNA Deoxyribonucleic acid DPN Diarylpropionitrile EDTA Ethylenediaminetetraacetic acid EGF Epidermal growth factor EGFR Epidermal growth factor receptor ER α Estrogen receptor α ER β Estrogen receptor β

ERK Extracellular signal-regulated kinase EB Estradiol Benzoate FBS Fetal bovine serum FSH Follicle stimulating hormone G-1 ±)-1-[(3aR*,4S*,9bS*)-4-(6-Bromo-1,3-benzodioxol-5-yl)- 3a,4,5,9b-tetrahydro-3H-cyclopenta[c]quinolin-8-yl]-ethanone G-15 (3aS*,4R*,9bR*)-4-(6-Bromo-1,3-benzodioxol-5-yl)- 3a,4,5,9b-3H-cyclopenta[c]quinoline GABA - Aminobutyric acid GABA-AR Aminobutyric acid type A receptor GABA-AR α1 Aminobutyric acid type A receptor alpha 1 subunit GABA-AR -α4 Aminobutyric acid type A receptor alpha 4 subunit GABA-AR -α6 Aminobutyric acid type A receptor alpha 6 subunit GABA-AR -β2 Aminobutyric acid type A receptor beta 2 subunit GABA-AR -β3 Aminobutyric acid type A receptor beta 3 subunit GABA-AR -δ Aminobutyric acid type A receptor delta subunit GABA-AR -γ2 Aminobutyric acid type A receptor gamma 2 subunit GHSR Growth hormone secretagogue receptor GnRH Gonadotrophin releasing hormone GPER G -protein coupled estrogen receptor GR Glucocorticoid receptor GRIA2 AMPA selective glutamate receptor 2 GRIN NMDA selective glutamate receptor 1 HBSS Hank’s balanced salt solution HPA Hypothalamic-pituitary-adrenal HPG Hypothalamic-pituitary-gonadal HPRT Hypoxanthine-guanine phosphoribosyl transferase HRP Horse radish peroxidase IR Insulin receptor JNK c-Jun N-terminal kinase kDa Kilodaltons LH Luteinizing hormone Lrf Luman recruitment factor MAP2 Microtubule Associated Protein 2 MAPK Mitogen-activated protein kinase mHippoE-14 Female derived murine embryonic hippocampal neurons mHippoE-18 Male derived murine embryonic hippocampal neurons MR Mineralocorticoid receptor mRNA Messenger ribonucleic acid Na3VO4 Sodium Orthovanadate NMDA N-methyl-D-aspartic acid NMDA-R1 NMDA receptor 1 NPY Neuropeptide Y NSE Neuron specific enolase obRb Leptin receptor OVX Ovariectomized

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P450scc P450 cholesterol side-chain cleavage enzyme PBS Phosphate-buffered saline PCR Polymerase chain reaction PI3K-Akt Phosphoinositide 3-kinase-Protein Kinase B p-JNK Phospho-JNK PKA Protein kinase A PPT 4,4',4''-(4-Propyl-[1H]-pyrazole-1,3,5-triyl)trisphenol RNA Ribonucleic acid SDS-PAGE Sodium dodecyl sulfate polyacrylamide gel electrophoresis SSTY1 Spermiogenesis specific transcript on the Y STAT3 Signal transducer and activator of transcription 3 SV40 Recombinant murine retrovirus harboring simian virus TAE Tris acetic acid EDTA TBS Tris-buffered saline TBS-T Tris-buffered saline with Tween-20 TE Tris-EDTA THRα Thyroid hormone receptor alpha THRβ Thyroid hormone receptor beta TrkA Tropomyosin receptor kinase type 1 TrkB Tropomyosin receptor kinase type 2

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LIST OF APPENDICES

Appendix 1: Sanger sequencing results for primer pairs that successfully amplified DNA fragments from mHippoE immortalized hippocampal neurons

Appendix 2: Antibody information for western blotting experiments

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CHAPTER 1: INTRODUCTION AND REVIEW OF THE LITERATURE

Sex Differences in Neuropsychiatric and Neurodegenerative Disorders Sex differences in structure and function of the brain are observed from the day of birth and persist throughout development and into adulthood (Bao & Swaab, 2002).

Neuropsychiatric, neurodegenerative, and neurological diseases are complex conditions that alter central nervous system structure and function, thereby affecting the overall well-being and cognition of individuals (Hanamsagar & Bilbo 2016). However, there are profound sex differences in the onset, incidence, and severity of many of these conditions (Hanamsagar & Bilbo 2016). For example, women are affected almost twice as often as men by anxiety disorders, post-traumatic stress disorder, and major depressive disorders (Altemus, 2006; Bao & Swaab, 2002; Hanamsagar & Bilbo, 2016) while men are more often affected by severe learning disabilities, schizophrenia, and autism spectrum disorder (Bao & Swaab, 2002). These sex differences are also evident in neurodegenerative conditions such as Parkinson’s disease and amyotrophic lateral sclerosis which more often affects males and multiple sclerosis and Alzheimer’s disease which more often affects females (Baldereschi et al., 2000; Bao & Swaab, 2002; Duarte-

Guterman et al., 2015; Voskuhl & Gold, 2012). While it is difficult to determine the etiology of these sex differences, a substantial amount of work suggests that the interactions between environmental, genetic, and hormonal factors may contribute to their multifactorial development and pathology (Xu et al., 2015).

The innate genetic differences between males and females results in sexually differentiated brain development that is subsequently modulated by sex specific gonadal hormone secretion. These genetic and physiological sex differences are believed to explain the functional basis for the sex differences seen in the neuropsychiatric disorders. Both the primary male and female sex steroids have been shown to mediate

2 neurotrophic and neuroprotective effects, though the timing and extent of these effects may be sexually differentiated (Brann et al., 2007; Garcia-Segura & Balthazart, 2009;

Pike et al., 2009; Pike et al., 2008). It is well-established that varying quantities of these circulating and locally synthesized sex steroids may underly the sexually dimorphic and differentiated development of many neuropsychiatric conditions including anxiety, mood, and learning disorders (Altemus, 2006; Bao & Swaab, 2002). In fact, many studies have demonstrated that females and males innately differ in their behavioural responses to stressful stimuli, implying that there may be mechanistic differences within the brain that result in a sexually dimorphic vulnerability to stressful stimuli (Shors et al., 2001; Wood et al., 2001; Wood & Shors, 1998). For example, exposure to an acute behavioural stressor has been shown to enhance hippocampus dependent associative learning in male rats while impairing cognitive performance in their age-matched female counterparts (Wood & Shors, 1998; Workman et al., 2015). Intriguingly, this female- specific cognitive impairment has been shown to onset within 10-30 minutes after exposure to the stimulus and persist for days after the stressful stimulus has been removed from their environment (Shors et al., 2001; Wood et al., 2001; Wood & Shors,

1998).

The female’s increased vulnerability to mood disorders has been directly correlated with the onset of the cyclic production of the primary female sex steroid, 17β- estradiol (estradiol) (Hayward & Sanborn, 2002; Kessler & Walters, 1998; Lewinsohn et al., 1998) while the loss of circulating estradiol production that occurs with age has also been directly correlated with the onset of many sexually differentiated neurodegenerative conditions (Tang et al., 1996). For example, the rapid and

3 substantial loss of ovarian hormone synthesis that occurs during menopause has been correlated with increased neuronal atrophy and loss of neuroprotection (Tang et al.,

1996). This loss of estrogen mediated neurotrophic and neuroprotective effects could therefore conceivably be contributing to the sexually differentiated atrophy, loss of neuronal connectivity, and subsequently, impaired cognition within aging female populations.

The Hippocampus

Many brain regions are sensitive to the effects of circulating and locally synthesized steroids (McEwen & Milner, 2007; McEwen & Milliken, 1999; Reul & De

Kloet, 1985). For example, in response to a stressful stimulus, the adrenal cortex will synthesize and secrete glucocorticoids into circulation which may exert their effects within the central nervous system by binding to their respective glucocorticoid receptors

(GRs; De Kloet et al., 1988; Reul & de Kloet, 1986; Reul & De Kloet, 1985). These GRs are expressed and become activated within steroid sensitive regions of the brain, resulting in modulation of neuronal structure, function, and subsequently, behaviour

(McEwen, 1982; Reul & De Kloet, 1986).

The hippocampus is a steroid-sensitive limbic structure that plays a profound role in mediating learning and consolidation of short-term and spatial memory through long term potentiation (Day et al., 2003; Riedel et al., 1999; Winters et al., 2004; Winters et al., 2008). Due to its role in learning and memory, the hippocampus is one of the most extensively investigated brain structures as it is often the earliest and most severely affected in a variety of neuropsychiatric and neurodegenerative diseases (Lupien et al.,

1998; McEwen & Milliken, 1997; Snyder et al., 2011). The hippocampus consists of functionally and anatomically distinct ventral and dorsal regions (Witter, 2012). The ventral hippocampus has been shown to be involved in processing and consolidating memories related to anxiety and fear while the dorsal hippocampus plays a larger role in spatial and conceptual memories (Bast et al., 2001; Morris et al., 2006). The hippocampus is organized into distinct regions known as the cornu ammonis (CA) 1,

CA2, CA3, and dentate gyrus that each exhibit distinct morphological features and roles in memory consolidation (Witter, 2012). For example, CA1 is most notable for its role in mediating long term potentiation through strengthening synaptic connectivity and dendritic spine density (Durand et al., 1996; Harris et al., 1984).

The hippocampus is sensitive to a variety of hormones including both sex steroids and stress hormones. As a result, it is a key mediator in sexually differentiated and dimorphic learning, memory, and behavioural responses to stressful stimuli (Handa

& Weiser 2014; Weiser & Handa 2009). For this reason, the hippocampus has been shown to express both intracellular and membrane bound receptors that are bound by glucocorticoids, androgens, and estrogens (González et al., 1999; Ratka et al., 1989;

Simerly et al., 1990; Walf & Frye, 2006). While many sexually differentiated anxiety behaviours have been observed, the cellular mechanisms mediating these sex differences within the hippocampus remain only partially elucidated (Hart et al., 2014;

Kastenberger et al., 2012; Shors et al., 2001; Wood & Shors, 1998). Interactions between sex and stress steroid receptors and cellular signaling mechanisms present one potential avenue for investigating the underlying mechanisms that regulate these behavioural responses.

The Hypothalamic-Pituitary-Gonadal (HPG) Axis

The hypothalamic-pituitary gonadal (HPG) axis is responsible for regulation of sex steroid synthesis through both positive and negative feedback mechanisms (Handa et al., 1994).The pulsatile release of gonadotrophin releasing hormone (GnRH) from the hypothalamus into the hypophyseal portal stimulates secretion of pituitary hormones including follicle stimulating hormone (FSH) and luteinizing hormone (LH) (Viau, 2002;

Viau & Meaney, 1991). Upon release into the female’s circulation, FSH will cause maturation of ovarian follicles that result in the synthesis and secretion of estradiol from the ovaries (Gemzell et al., 1958). From here, circulating estradiol will bind estrogen receptors (ERs) within the hippocampus, hypothalamus, and pituitary glands to exert both negative and positive feedback effects on the secretion of FSH and LH from the anterior pituitary (Brinton et al., 2015; Handa et al., 1994). This negative feedback functions to reduce FSH secretion and return circulating estradiol levels to baseline, while the positive feedback mechanism will enhance production of LH that will cause the ovulatory surge (Handa et al., 1994). Circulating concentrations of estradiol vary according to reproductive stage throughout the course of the human menstrual and rodent estrous cycles (Goldman et al., 2007; Wiele et al., 1970).

In the rodent, the 4-5 day estrous cycle consists of 4 distinct phases including proestrus, estrus, metestrus, and diestrus that can each be characterized by cell type and circulating concentrations of LH, FSH, estradiol, and progesterone (Goldman et al.,

2007). The proestrus phase of the estrous cycle is typically characterized by a rise in estradiol that is accompanied by the LH and FSH ovulatory surge (Caligioni, 2009;

Parkening et al., 1982; Walmer et al., 1992). Following the proestrus ovulatory surge,

6 estradiol levels rapidly decline and this is accompanied by a respective decrease in

FSH and LH levels during the estrus phase of the cycle (Caligioni, 2009). Throughout the following metestrus phase of the cycle, circulating concentrations of estradiol will reach their lowest levels and begin to gradually increase through diestrus in preparation for the proestrus ovulatory surge (Walmer et al., 1992). Circulating levels of LH and

FSH remain steady and relatively low throughout estrus, metestrus, and diestrus phases of the rodent reproductive cycle (Miller & Takahashi, 2013).

Figure 1.1 Characteristic estradiol, progesterone, LH, FSH, and GnRH levels across the 4-5-day estrous cycle of female mice. The peak in estradiol synthesis and circulating concentrations occurs during the proestrus phase of the cycle after which these concentrations rapidly decline into estrus (Miller & Takahashi, 2013). While systemic concentrations of estradiol are regulated by the HPG axis, concentrations of estradiol within the hippocampus have been shown to be significantly higher than those found in the serum (Hojo et al., 2009). As 17β-estradiol, the most potent estrogen, is also synthesized de novo within the brain, local levels can differ considerably from circulating levels. Male hippocampal estradiol concentrations have been shown to reach up to 8.4 nM in comparison to the 0.014 nM seen in their respective circulating concentrations (Hojo et al., 2004). In females, hippocampal

7 estradiol levels during proestrus, estrus, diestrus, and metestrus will range between 0.5-

4.3 nM in comparison to the consistent <0.1 nM circulating levels (Kato et al., 2013).

Steroid and Estradiol Biosynthesis

Within the central nervous system, steroids have been shown to exert both activational and organizational changes. Organizational effects occur during the prenatal/developmental period when exposure to hormones may permanently change the brain’s sensitivity to that same hormone later in life (MacLusky & Naftolin, 1981).

Activational changes occur postnatally following transient hormone exposure that may result in temporary changes in activation and signaling mechanisms in the brain (Arnold,

2009). These steroid hormones exert their effects by binding and activating membrane- bound and/or intracellular receptors (Ramirez & Zheng 1996). These specific steroid receptors were first identified within the brain by their ability to bind circulating steroid hormones with high affinity (McEwen et al., 1986). It was believed that these circulating steroid hormones exerted their effects by crossing the blood brain barrier and directly binding their respective receptors within steroid sensitive regions of the brain (Pardridge et al., 1980). However, it is now understood that steroid hormones may be synthesized de novo by neurons or glia within the central nervous system itself (Baulieu et al.,

2001). Within the brain, these locally synthesized or circulating steroids may also be enzymatically converted into their biologically active metabolites such as testosterone’s conversion to estradiol through the highly expressed aromatase enzyme in the brain

(Baulieu & Robel, 1990). From here, these steroids and their metabolites may activate intracellular receptors or membrane bound receptors that are capable of regulating

8 gene expression either directly through classical mechanisms or indirectly through non- classical activation of signaling cascades, respectively (Carson-Jurica et al., 1990).

Mammalian endocrine glands function to synthesize and secrete four unique classes of hormones, one of which includes steroid hormones (Kim et al., 2016). The regulation of hormone synthesis and secretion is believed to depend on rates of steroid hormone biosynthesis and catabolism (Carson-Jurica et al., 1990). As hydrophobic molecules, steroid hormones are carried in the blood by binding proteins (i.e. sex- steroid binding globulin) and dissociate from these binding proteins within target tissues to exert their effects on membrane-bound and intracellular receptors by diffusing through the lipid bilayer of cells within steroid sensitive organs (Hanukoglu, 1992; Selby,

1990). All steroid hormones share structural similarities in their three six-carbon ring and one conjugated five-carbon ring (Compagnone & Mellon, 2000). These steroid hormones originally derive from the common 27-carbon cholesterol precursor that is processed through a series of enzymatic conversions to produce two functionally distinct steroid groups including the stress response hormones and the sex steroids

(Frick et al., 2015). Through a series of P450 and steroid oxido-reductase enzymatic conversions, the cholesterol molecule is metabolized to produce five major classes of steroid hormones including progestogens, androgens, mineralocorticoids, glucocorticoids, and estrogens (Figure 1.2; Compagnone & Mellon, 2000; Hanukoglu,

1992). The cholesterol precursor is first cleaved to form the 21-carbon progestins including pregnenolone and progesterone through the rate limiting conversion by the P-

450 cholesterol side-chain cleavage enzyme (P450scc) (Lambeth & Stevens 1984).

Identification of P450scc’s wide distribution throughout the brain has expanded our

9 understanding of neurosteroids and neuroendocrinology (Baulieu 1997; Baulieu & Robel

1990; Baulieu et al., 2001). Progesterone may subsequently be converted to produce corticosteroids including corticosterone and cortisol, the most prevalent glucocorticoids in mice and humans, respectively (Mensah-Nyagan et al., 1999). Progesterone may also be metabolized through oxidoreductase reactions to produce 19-carbon androgens such as androstenedione, testosterone, and dihydrotestosterone (Mensah-Nyagan et al., 1999). From here, both androstenedione and testosterone may be cleaved to generate the 18-carbon estrogens through the aromatase enzyme which is highly expressed within the hippocampus (Maclusky et al., 1987; Sasano et al., 1998).

Estrogens are most predominantly synthesized within the female gonadal ovaries but may also be locally synthesized within the brain through de novo synthesis

(MacLusky et al., 1986; MacLusky et al., 1994; Roselli et al., 1985). The class of estrogen steroid molecules include estrone, 17β-estradiol, and estriol all of which share similarities in steroid ring structures that have been commonly derived from the 27 carbon cholesterol precursor (Compagnone & Mellon, 2000; MacLusky et al., 1986).

Estrone and 17β-estradiol are synthesized from androstenedione and testosterone aromatization, respectively (MacLusky et al., 1987; MacLusky et al., 1986). Estrone may be enzymatically converted by 17β-hydroxysteroid to produce the most potent estrogen,

17β-estradiol (Compagnone & Mellon 2000; Murakami et al., 2006). The hippocampus has been shown to express all of the enzymes involved in the biosynthesis of estradiol from the cholesterol precursor, including P450 scc, 17α-hydroxylase/17,20 lyase, 17β- hydroxysteroid dehydrogenase, 3β-hydroxysteroid dehydrogenase, and aromatase

(Compagnone & Mellon, 2000; Hanukoglu, 1992; Hojo et al., 2009; Lambeth & Stevens,

1984; Melcangi et al., 1996).

Both males and females synthesize substantial quantities of progestogens, mineralocorticoids, glucocorticoids, androgens, and estrogens. However, the primary sex difference exists in the quantity of converting enzymes that metabolize the respective steroid molecules (Terasawa & Ojeda, 2009). While catabolism of one steroid leads to the synthesis of another, this makes identifying key sex differences uniquely challenging (Frick et al., 2015).

Figure 1.2 Steroid biosynthesis pathway showing synthesis and metabolism of progestogens, androgens, glucocorticoids, and estrogens. Steroids are all derived from a common cholesterol precursor molecule. Cholesterol is metabolized by P450scc to produce pregnenolone. Pregnenolone may be converted by 17α- hydroxylase into 17α-hydroxy pregnenolone or by 3β-hydroxysteroid dehydrogenase (HSD) into progesterone. Progesterone may go through a series of oxidoreductase reactions to produce corticosterone (the primary glucocorticoid in mice) or cortisol (the primary glucocorticoid in humans). 17α-hydroxy pregnenolone may also be converted by 3β-HSD and two oxidoreductase reactions to result in the synthesis of cortisol. 17α- hydroxy pregnenolone and progesterone may also be converted by 3β-HSD and 17α- hydroxylase (OHase) to produce 17α-hydroxyprogesterone, respectively. 17α-hydroxy pregnenolone may also be metabolized by 17,20-lyase to result in synthesis of dehydroepiandrosterone (DHEA) which may subsequently be converted to androstenediol. Both DHEA and 17α-hydroxyprogesterone may be metabolized to form androstenedione. Androstenedione and androstenediol may both be metabolized to form testosterone. Both androstenedione and testosterone may also undergo

12 aromatization reactions to result in the synthesis of estrone and estradiol, respectively. Estrone may subsequently be metabolized by 17β-HSD to result in further synthesis of estradiol and estriol. Estradiol may be further metabolized to increase estriol synthesis. Figure adapted from Creighton (2017).

Steroid Receptors

Steroid receptors are differentially expressed throughout the male and female brains and may have serious implications for mediating important hormonal feedback and sexually differentiated signaling mechanisms that results in physiological, cognitive, and behavioural differences (Duarte-Guterman et al., 2015; Finley & Kritzer, 1999;

Kritzer, 2006; Milner et al., 2001; Milner et al., 2005). Gonadal and stress steroid hormones exert their effects through binding their respective receptors within brain regions involved in regulating cognition, behavior, mood, learning, and memory. For example, the GR may be bound by endogenous cortisol/corticosterone or exogenous glucocorticoids such as dexamethasone, while the ERs including estrogen receptor α

(ERα), estrogen receptor β (ERβ), and the g-protein coupled estrogen receptor (GPER) bind estrogens including estrone, estriol, and 17β-estradiol. As intracellular receptors,

GR, ERα, and ERβ may activate classical mechanisms that directly regulate the expression of genes that may modulate neuronal structure, function, and subsequently behaviour. Membrane-bound non-classical receptors such as GPER are capable of exerting their rapid and non-genomic effects through activation of intracellular signaling cascades that may indirectly regulate gene expression, neuronal morphology, function, and subsequently behavior. However, these steroid receptors may be regulated and/or modulated by environmental and physiological inputs.

Estrogen Receptors in the Hippocampus

As the most potent estrogen, estradiol is capable of exerting both its short and long term effects through binding and activating both genomic and non-genomic mechanisms mediated by ERα and ERβ which can be found in nuclear, cytoplasmic, and plasma membrane sites or non-classical GPER found within the plasma membrane, endoplasmic reticulum, or Golgi apparatus (Duarte-Guterman et al., 2015;

McEwen et al., 2012; McEwen & Milner 2007; Vasudevan & Pfaff 2008). As a steroid sensitive region of the brain, the hippocampus has been shown to express all three ERs

(ERα, ERβ, and GPER) accounting for its critical role in modulating estrogen sensitivity.

Estradiol has been demonstrated by many to be a potent mediator of neuroprotective and neurotrophic effects within the brain (Frick et al., 2015). Estradiol, a lipophilic steroid molecule, is capable of passing through the lipid bilayer of a plasma membrane to access the intracellular space and activate intracellular classical ER-α and

ER-β (Frick et al., 2015). ERα and ERβ are members of the nuclear receptor superfamily and share 95% similarity in their DNA binding domains and high sequence similarity in their ligand binding domains (Hewitt & Korach 2002; Kuiper et al., 1997).

Upon binding of estradiol and activation of intracellular ERα or ERβ, the hormone receptor complex will undergo conformational change, dimerization, and will translocate to the nucleus where it can bind hormone response elements that initiate transcription of estrogen sensitive genes (Frick et al., 2015). From here, coactivators and corepressors are recruited that allow ERα and ERβ to either facilitate or inhibit transcription of target genes and activate signalling cascades that may differentially modulate dendritic spine plasticity, learning, memory, and behaviour in both males in females (Frick et al., 2015;

Fugger et al., 1998; Sánchez-Andrade & Kendrick 2011). Expression of these classical receptors has been exhibited in many different brain regions responsible for regulating mood, sexual behaviour, and hormone feedback regulation including the prefrontal cortex, entorhinal cortex, perirhinal cortex, hypothalamus, amygdala, cerebellum, thalamus, and lastly the hippocampus (Brinton et al., 2015; Frick et al., 2015; Österlund et al., 1999; Shughrue et al., 1997; Shughrue & Merchenthaler 2001; Simerly et al.,

1990). These classical genomic mechanisms typically require many hours to days to exhibit functional changes and responses within the hippocampus. However, the hippocampus has been previously shown to express relatively few nuclear estrogen receptors (Weiland et al., 1997). Instead, these intracellular receptors may localize within the cytosol and have also been associated with rapidly modulating dendritic spine density, suggesting a role in mediating non-classical cell signalling pathways. For example, systemic estradiol treatment was shown to increase ERβ translocation to the plasma membrane (Sheldahl et al., 2008). Membrane localization of ERα and ERβ has been shown to activate extracellular signal-regulated kinase (ERK) and cyclic adenosine monophosphate (cAMP) response element binding protein (CREB)

(Boulware et al., 2005; Boulware, et al., 2013). In fact, estradiol has been shown to induce ERK-dependent enhancement of memory performance in ovariectomized mice compared to their non-estradiol treated counterparts (Fernandez et al., 2008). This suggests that membrane associated ERs are also capable of facilitating the non- classical effects of estradiol (Micevych & Dominguez 2009).

Within the hippocampus, ERα expression is highest within the CA3 and CA1 regions, while ERβ appears to be more evenly expressed throughout the entire

15 structure of the hippocampus (Mitra et al., 2003). These classical receptors are expressed within both the female and male adult hippocampus within the nucleus, dendrites, dendritic spines, axons, nerve terminals and glia within the hippocampus

(Milner et al., 2001; Milner et al., 2005; Mitra et al., 2003; Mitterling et al., 2010;

Shughrue et al., 1997; Shughrue & Merchenthaler 2001). Within the hippocampus, expression of these receptors may fluctuate according to phase of the cycle, showing highest expression during proestrus when estradiol levels are highest, compared to diestrus females (Frick et al., 2015).

The more recently identified GPER has been shown to bind estradiol with high affinity and mediate rapid non-genomic effects of estradiol through a mechanism distinct from the intracellular classical estrogen receptors (Carmeci et al., 1997; Filardo et al.,

2000; Revankar et al., 2005; Thomas et al., 2005). GPER is a 7 transmembrane domain g-protein coupled receptor that belongs to the class A rhodopsin-like g-protein coupled receptor family (Filardo & Thomas, 2005). GPER has been shown to colocalize with various Gα receptor subunits and a Gβγ dimer to form the GPER complex (Thomas et al., 2005). Structurally and genetically, GPER is unrelated to both ERα and ERβ and has been shown to localize to different subcellular regions depending on the tissue type

(Carmeci et al., 1997). Since its discovery, many studies have demonstrated GPER expression within the cell bodies, dendritic spines, axons, and terminals of neurons within the hypothalamus, cortex, striatum, and hippocampus where it typically associated with postsynaptic scaffolding proteins that are also localized near the plasma membrane (Akama et al., 2013; Waters et al., 2015). GPER mRNA expression has been shown to be significantly high within the steroid-sensitive hippocampus (Gibbs,

2010). Within the hippocampus specifically, GPER has been shown to localize within extranuclear sites of interneurons, glia, and pyramidal neurons (Waters et al., 2015).

The glycosylated form of GPER remains membrane bound and activates signaling cascades through non-classical mechanisms (Pupo et al., 2017) while the non- glycosylated form of GPER may be found intracellularly within the endoplasmic reticulum or Golgi apparatus in the mouse hippocampus (Waters et al., 2015). In the presence of estradiol, GPER is able to translocate from the endoplasmic reticulum and

Golgi apparatus to the plasma membrane where it exerts its rapid non-classical effects through activation of signaling cascades within a rapid minute to hours timescale

(Funakoshi et al., 2006).

Upon activation, GPER undergoes a conformational change that allows guanosine triphosphate to bind to the Gα subunit, causing its dissociation. The Gα subunit will increase cAMP and protein kinase A (PKA) production and activation, respectively (Filardo et al., 2000; Filardo et al., 2007). This estrogen mediated production of cAMP inhibits transactivation of epidermal growth factor receptors

(EGFRs) and inhibition of Raf-1, reducing activation of ERK1 and ERK 2 activity (Filardo et al., 2002). The subunit dissociation allows the Gβγ subunit to stimulate matrix metalloprotease cleavage and induced transactivation of EGFRs that results in phosphorylation and activation of the ERK signaling pathway (Filardo et al., 2000; Qiu et al., 2003). This GPER mediated modulation of ERK signalling has been associated with regulating cellular differentiation through increased c-fos expression (Maggiolini et al.,

2004). GPER activation has previously been shown to modulate proliferative signalling through activation of the phosphoinositide 3-kinase (PI3K)-protein kinase B (Akt)

17 pathways (Gingerich et al., 2010). Following estradiol mediated activation, GPER becomes sequestered within clathrin-coated vesicles and is subsequently transported to the endoplasmic reticulum or Golgi complex to be conserved as opposed to degraded by lysosomes (Cheng et al., 2011).

More recently, hippocampus membrane bound GPER activation has been shown to activate the c-jun N terminal kinase (JNK) signaling pathways that is capable of indirectly regulating gene expression and exerting downstream intracellular effects (Kim et al., 2016). Activation of these kinase pathways has been shown to have profound implications for the rapid modulation of neuronal structure and subsequently, hippocampal mediated learning and memory performance (Gabor et al., 2015; Kim et al., 2016; Lymer et al., 2017). For example, use of the GPER specific synthetic agonist

G-1 in ovariectomized mice has been shown to stimulate JNK phosphorylation that enhances dorsal hippocampus mediated object recognition and spatial memory consolidation that mimics the memory enhancing effects of estradiol (Kim, 2014; Kim et al., 2016). The memory enhancing effects of G-1 have been shown to be independent of ERK phosphorylation but instead dependent on the activation of JNK, establishing

GPER’s unique signaling mechanism distinct from that of ERα and ERβ (Kim et al.,

2016). The mitogen activated protein kinase (MAPK) family includes both the ERK and the JNK family (Peng et al., 2010). JNK1, JNK2, and JNK3 comprise the JNK subfamily with JNK1 and JNK2 playing large developmental, physiological, and pathological roles and JNK3 playing a more significant role in pathological development of conditions like

Alzheimer’s disease (Antoniou & Borsello 2012).

These serine/threonine kinase signaling proteins have been evolutionarily

18 conserved (Antoniou & Borsello, 2012). Upon activation, cytosolic JNK will undergo phosphorylation of the threonine and tyrosine residues and phosphorylated JNK (p-

JNK) may phosphorylate other substrates such as the GR (Qi et al., 2005; Wang et al.,

2005). JNKs have been shown to play important roles in mediating plasticity, learning, and consolidation of memory (Antoniou & Borsello, 2012). Specifically, JNK has been shown to regulate memory formation in hippocampus dependent associative learning using stressful and baseline conditions (Antoniou & Borsello 2012). However, JNK’s role in the consolidation of short and long term memory may differ (Bevilaqua et al. 2003).

Specifically, basal JNK expression is highest within the brain and is often upregulated in response to oxidative stress and DNA damage to regulate anti and pro apoptotic protein synthesis (Antoniou and Borsello 2012). Specifically, primary cultures of hippocampal neurons treated with glucocorticoids exhibit rapid activation of JNK while JNK inhibition has been shown to enhance the expression of the GR in lines of mouse HT22 hippocampal cells (Qi et al., 2005; Wang et al., 2005). Activation and phosphorylation of JNK may regulate transcription, translation, and epigenetic modifications that may subsequently modulate dendritic structure, function, and behavior. Through GPER, the estrogen mediated non-classical activation of JNK occurs rapidly within minutes to hours and has been attributed to enhancements in memory formation and neurotrophic effects within the hippocampus (Gabor et al., 2015; Kim,

2014).

Figure 1.3 Cellular signaling cascades of membrane bound and intracellular estrogen receptors (ERα, ERβ, and GPER). Estradiol mediated activation of classical estrogen receptors may directly regulate gene expression through binding nuclear hormone response elements or activation of intracellular cascades. Activation of the non-classical GPER results in activation of intracellular c-jun N terminal kinase (JNK) cascades that may indirectly regulate gene expression through epigenetic modifications, posttranslational modifications, and modulation or protein synthesis and degradation. This indirect regulation of estrogen sensitive genes may subsequently modulate learning and memory in a rapid minute to hours timescale (Frick et al., 2015).

Functional Effects of Hippocampal Estrogens and GPER Activation The post-synaptic components of excitatory synapses are contained within small dendritic protrusions known as dendritic spines. Since 1990, it has been well established that circulating levels of estrogens can potently modulate hippocampal synaptic plasticity and spine density in female rats (Gould et al., 1990; Woolley et al.,

1990; Woolley & McEwen, 1992). For example, throughout the course of the 4-5 day estrous cycle, the low levels of circulating estradiol during the metestrus and diestrus phases correlate with reduced synapse density within the CA1 hippocampal region while the opposite effects are observed during the high estradiol proestrus and estrus phases of the cycle (Woolley & McEwen, 1992). Additionally, ovariectomized female

20 rats exhibit significantly reduced dendritic spine density in hippocampal pyramidal neurons within the CA1 region (Gould et al., 1990), but these effects are reversed with systemic injections of estradiol (Hao et al., 2003; Tang et al., 2004). These estradiol mediated increases in dendritic spine and synapse density are also mirrored in nonhuman primate hippocampal CA1 and prefrontal cortex regions (Hao et al., 2003;

Leranth, Shanabrough, & Redmond, 2002; Tang et al., 2004). However, the effects of estradiol on hippocampal morphology seem to be limited to dendritic spine and synapse density in female rodents as Sholl analysis reveals that the overall dendritic tree is unaffected ( Mendell et al., 2017). Use of ERα and ERβ specific agonists shows rapid increases in spine density in the mouse hippocampus just 40 minutes after systemic administration while treatment with estradiol has shown increased post-synaptic density within 30 minutes of treatment (MacLusky et al., 2005; Phan et al., 2012; Phan et al.,

2011). Following systemic treatment with estradiol, CA1 dendritic spine density increases in ovariectomized female mice and rats (MacLusky et al., 2005; Phan et al.,

2012) and this has been paralleled in estradiol treated hippocampal slices from female mice (Pozzo-Miller et al., 1999) and male rats (Mukai et al., 2007).

Estradiol has also been shown to increase excitatory synaptic transmission according to various electron microscopy studies (Moult & Harvey, 2008). This is evident through increases in the levels of post synaptic density 95 (PSD-95), synaptophysin, syntaxin, and spinophilin following estradiol exposure in cultured hippocampal neurons and ovariectomized rats (Akama et al., 2013; Lee et al., 2004;

Moult & Harvey, 2008; Murphy & Segal, 1996). Electrophysiological studies have revealed that activation of membranous estrogen receptors in hippocampal slices can

21 evoke an action potential in population or single cell recordings within the CA1 and CA3 regions (Kim et al., 2006; Rudick & Woolley, 2003; Wong & Moss, 1992). These estradiol induced changes in synaptic transmission, coupled with changes in morphology demonstrate estradiol’s ability to modulate long term potentiation (Warren et al., 1995). For example, treatment with estradiol using in vivo and in vitro hippocampal models has been shown to increase the magnitude of long term potentiation while stimulating the upregulation of N-methyl-D-aspartic acid (NMDA) receptor expression and transmission (Moult & Harvey, 2008). Estrogen receptors are often expressed within GABAergic interneurons of the hippocampus and estradiol treatment may partially mediate the transient increase in dendritic spine density through inhibition of GABAergic neurotransmission (Murphy et al., 1998). As an important inhibitory neurotransmitter, GABA may modulate neurogenesis through regulating proliferation of migration of neuroblasts, synapse formation, and synaptic plasticity

(Pallotto & Deprez, 2014).

Neurogenesis within the adult brain requires the processes of cellular proliferation, migration, differentiation into neuron or glia, and survival into the mature phenotype (Duarte-Guterman et al., 2015). Neurogenesis within the dentate gyrus and sub granular zone have been shown to be sensitive to the concentration of circulating estradiol and sex of the animal. While acute estradiol treatment has been shown to enhance neurogenesis and repeated estradiol treatment has been shown to increase cellular proliferation and reduce overall cell death within the dentate gyrus of the female hippocampus (Barker & Galea, 2008), cellular proliferation and apoptosis appear to be insensitive to the effects of estradiol in the male hippocampus (Barker & Galea, 2008).

This clear sex difference is evident by the same enhancements in cell proliferation and reduced apoptosis seen in adult male rats following treatment with testosterone and dihydrotestosterone (Spritzer & Galea, 2007).

Males have been shown to outperform females in spatial learning tasks that include route learning, Morris water maze, and radial arm mazes (Galea & Kimura,

1993; Jonasson, 2005; Voyer et al.,1995). These sex differences are believed to be founded on the males relatively larger contextual fear responses since circulating estradiol levels play a role in reducing the females contextual fear response (Gupta et al., 2001). This sex difference is abolished upon ovariectomy and is re-established when ovariectomized (OVX) females are treated with 10 ug of estradiol (Gupta et al.,

2001). These results, in combination with estradiol mediated enhancements in spine synapse density, long term potentiation, and neurogenesis suggests that estrogens enhance hippocampal plasticity, learning and memory (Foy et al., 1999; Frick et al.,

2004; Tanapat et al., 1999; Warren et al., 1995). However, many findings have demonstrated that spatial reference and working memory, as assessed by the Morris water maze, is more significantly impaired during the high estradiol proestrus and estrus phases of the cycle in comparison to the low estradiol phases (Duarte-Guterman et al.,

2015; Frye, 1995; Galea et al., 1995; Warren & Juraska, 1997). The Morris water maze task measures a rodent’s spatial reference memory in finding a hidden platform submerged in water (Morris, 1984). Although widely accepted as a hippocampus dependent associative learning paradigm, the Morris water maze requires the animal to be submerged in water and therefore, naturally evokes a physiological stress response

(Frick et al., 2004). Additionally, the degree of stress reactivity was previously shown to

23 be higher during the proestrus phase of the cycle when estradiol levels are highest

(Viau & Meaney 1991). Intriguingly, Frick et al., (2004) demonstrated that exposure to an acute behavioural stressor such as the Morris water maze inhibits the effects of estradiol treatment on spine synapse density in the hippocampus of female rats. It is evident that behavioural performance and cognition vary across the estrous cycle, however, the memory enhancing effects appear to be dependent on the degree of stress experienced by the animal as well (Frick et al., 2004).

Figure 1.4 Effects of behavioural testing on estradiol induced increases in spine synapse density in ovariectomized female rats. In the absence of a behavioural stressor (Non WM-tested), treatment with estradiol benzoate (EB) results in a significant increase in spine synapse density in ovariectomized female rats compared to their control treated counterparts. Exposure to the Morris water maze (WM-tested) behavioural stressor eliminates the estradiol induced increase in spine synapse density within the hippocampus of female ovariectomized rats. This study demonstrates that behavioural stressors may eliminate the estradiol induced neurotrophic effects in the CA1 region of the female hippocampus (Frick et al., 2004). The effects of estradiol treatment and natural fluctuation across the estrous cycle indicate that estradiol plays an important role in enhancing dendritic spine and synapse density, plasticity, long-term potentiation, and cognition in the female. Estradiol treatment has been shown to rapidly improve social recognition, object placement, and

24 object recognition in ovariectomized mice within 40 minutes of treatment (Phan et al.,

2011, Phan et al., 2012). Intriguingly, use of the ERα agonist 1,3,5-tris (4- hydroxyphenyl)-4-propyl-1H-pyrazole (PPT) alone improves social recognition and has only a marginal effect on object recognition and object placement performance in female mice while treatment with the ERβ agonist 2,3-bis (4-hydroxyphenyl) propionitrile

(DPN) exerts no effect on object recognition or object placement, but impairs social recognition in female mice (Phan et al., 2011). While these effects do not completely mimic the memory enhancing effects seen with estradiol treatment, GPER has become an alternative candidate of interest for mediating the rapid memory enhancing effects of estradiol.

As a non-classical membrane-bound estrogen receptor, GPER activation has been shown to be a key mediator of these rapid non-genomic effects of estradiol in the hippocampus. GPER’s significant density in synaptic spines suggests that it may play a profound role in regulating plasticity (Gabor et al., 2015). Intriguingly, treatment with the

GPER specific agonist 1-4-(6-bromobenzo [1,3] dioxol-5-yl)-3a,4,5,9b-tetrahydro-3H- cyclopenta[c] quinolin-8-yl)-ethanone (G-1) was shown to rapidly enhance dendritic spine density within CA1 pyramidal neurons from the stratum radiatum of female mice in a dose dependent fashion within 40 minutes of treatment (Gabor et al., 2015). However, other studies have demonstrated that G1 activation of GPER may reduce dentate gyrus

GPER expression and cellular proliferation in female rats (Duarte-Guterman et al.,

2015). Intriguingly, Gingerich et al. (2010) demonstrated that treatment with G-1 at 10 nm and 100 nm doses significantly attenuated glutamate induced neurotoxicity and cell death in immortalized hippocampal neurons (Gingerich et al., 2010).

Figure 1.5 Dose dependent increase in spine density within the stratum radiatum of the CA1 region within 40 minutes of G-1 treatment in ovariectomized female mice. This study demonstrates that GPER’s significant density within dendritic spines may play an important role in the rapid mediation of estradiol’s neurotrophic effects within the female hippocampus(Gabor et al., 2015). GPER expression and activation has been associated with animal models of anxiety and depression. G-1 induced activation of GPER has been shown to evoke anxiogenic effects in OVX females and intact male mice and anxiolytic effects in intact female and gonadectomized male mice (Hart et al., 2014; Kastenberger, et al., 2012). It is speculated that GPER’s association with the serotonin receptor sub-type 5-HT1A may be responsible for 5-HT1A receptor desensitization resulting in GPER’s role in regulating anxiety and depressive like behaviours (McAllister et al., 2012). However,

GPER activation has also been associated with improvement in the Y-maze spatial learning task in OVX female rats when given chronically or acutely (Hammond et al.,

2009, Hawley et al., 2014). Chronic administration of G-1 to OVX rats restores delayed matching to position performance to that of the intact females while G-15 impairs performance in intact females to that of the OVX females (Hammond et al., 2009).

GPER’s high co-localization with cholinergic neurons and G-1’s mediated increase in acetylcholine (ACh) release within the hippocampus suggests that GPER

26 activation may regulate ACh release that may improve acquisition of specific learning tasks including the delayed matching to placement task (Hammond & Gibbs, 2011;

Hammond et al., 2011). Using G-1, Gabor et al. (2015) demonstrated that GPER’s rapid activation improved performance in social, non-social, and spatial recognition tasks in

OVX adult female CD1 mice while other studies demonstrate that GPER rapidly facilitates improvement in social and object recognition, but not object placement learning (Lymer et al., 2017). Using a model of pain-related anxiety Tian et al. (2013) demonstrated that GPER activation within the basolateral amygdala of OVX females maintained an anxiolytic effect through transmission of GABAergic and glutamatergic signaling.

Physiological models of ischemia and hypoxia have demonstrated sexually differentiated effects in neuronal recovery, where treatment with G-1 was particularly beneficial to females and detrimental to intact males (Broughton et al., 2014). These findings indicate that GPER activation may play a sexually differentiated role in response to neurotoxicity and hormonal stress. It has been well-established that activation of the physiological stress response system may negatively regulate HPG axis function through modulation of hormone synthesis, secretion, and mechanism of action. It is therefore plausible that the reciprocal modulation of the hypothalamic- pituitary-adrenal (HPA) and HPG axes may be responsible for the sexually differentiated effects of GPER activation on learning and memory in response to stressful stimuli.

Stress and the Hypothalamic-Pituitary-Adrenal (HPA) Axis

The primary stress response is physiologically regulated by the HPA axis that integrates feedback through the central nervous system and endocrine system

(Altemus, 2006; Smith & Vale, 2006; Viau & Meaney, 1991). In response to environmental, pathological, and physiological stressors, the body is able to regulate its internal environment through this homeostatic control system (Makara, 1985; McEwen,

1998; McEwen & Stellar, 1993). In response to these stressful stimuli, the hypothalamus releases corticotrophin releasing hormone (CRH) into the hypophyseal portal system that subsequently stimulates the anterior pituitary to release adrenocorticotropic hormone (ACTH) into circulation (Smith & Vale, 2006). The circulating ACTH binds to melanocortin receptors within the adrenal glands that stimulate the release of glucocorticoids from the adrenal cortex (Smith & Vale 2006). The secreted glucocorticoids exert their negative feedback effects by binding mineralocorticoid and

GRs within brain regions including the hypothalamus and hippocampus (Dallman et al.,

2002). This negative feedback acts to restore glucocorticoid production from the adrenal cortex to normal levels. However, sustained imbalances or elevations in the stress response may result in high allostatic load that has been associated with increased risk for neurodegenerative conditions such as Alzheimer’s disease (Lupien et al., 1998;

McEwen & Stellar, 1993; McEwen, 1998).

Glucocorticoids are capable of binding and activating both the glucocorticoid and mineralocorticoid receptor in the hippocampus. The hippocampus is a major brain region through which corticosteroids regulate the HPA axis (McEwen et al.,1986). The mineralocorticoid receptor (MR) is highly expressed within the hippocampus and will

28 bind endogenous corticosteroids including glucocorticoids, aldosterone, and mineralocorticoid (McEwen et al., 1968; Reul & De Kloet, 1986). At basal unstressed levels within the hippocampus, the MR will remain stably occupied by circulating corticosteroids (Reul & De Kloet, 1985; Reul et al., 1987). However, upon introduction of a physiological, pathological, or environmental stressor, the elevated glucocorticoid secretion will cause occupation of the glucocorticoid receptor (GR) (De Kloet & Reul,

1987; Reul & De Kloet, 1985; Reul et al., 1987). GRs are widely distributed throughout the entirety of the brain, however, the CA1, CA2, and dentate gyrus of the hippocampus exhibit high expression levels of both the GR and MR (Reul & De Kloet, 1985).

However, it has been shown that GR expression is highest within the CA1 and CA2 pyramidal neurons and within the dentate gyrus at both the mRNA and protein levels

(Van Eekelen et al., 1988).

Figure 1.6 The hypothalamic-pituitary-adrenal (HPA) axis in response to a stressful stimulus. In response to physiological or environmental stressors, corticotrophin release hormone will be secreted into the hypophyseal portal to stimulate the release of adrenocorticotropin (ACTH) from the anterior pituitary gland. ACTH will travel in the blood stream to activate adrenal melanocortin receptors that results in the subsequent release of adrenal glucocorticoids. Glucocorticoids will travel in the blood

29 stream and bind both mineralocorticoid and glucocorticoid receptors (MRs and GRs) in the hippocampus. The hippocampus plays a critical role in regulating negative feedback and initial perception and assessment of the stressors. The hippocampal MR will act to propagate the stress response while the hippocampal GR will act to negatively regulate the stress response (Raabe & Spengler 2013).

Glucocorticoid Biosynthesis The synthesis of endogenous glucocorticoids occurs through a similar series of

P450 and oxido-reductase enzymatic conversions to produce corticosterone/cortisol from the cholesterol precursor (Figure 1.2). These enzymatic conversions result in the synthesis of a 4-ringed, hydrophobic glucocorticoid molecule that contains a C3 keto group (Miller, 1988). As an intracellular steroid receptor, the GR becomes activated by the corticosteroids that have diffused through the lipid membrane. Upon binding and activation, GR forms a homodimer that translocates and binds its respective hormone response element within the nucleus (Beato et al., 1969; Savory et al., 2001). As a result, GR binding will associate with coactivators or co-repressors to regulate transcription of stress sensitive genes. Elevated circulating concentrations of glucocorticoids and prolonged activation of GR binding has long been associated with increased atrophy of the human brain (Lupien et al., 1998; McEwen Harold & Milliken

1997; Watanabe et al., 1992). The hippocampus exhibits high expression of both the

MR and GR and as a result, is a key stress sensitive brain region that regulates the physiological stress response (Ratka et al., 1989). Sustained activation of GR has been associated with atrophy of many brain regions, especially the hippocampal formation and this has become particularly evident in depressive disorders, PTSD, and dementia

(McEwen & Milliken, 1997). The hippocampus also expresses gonadal hormone receptors that play a well-established role in mediating sexually differentiated

30 behavioural responses. While the sexually differentiated behavioral responses in anxiety paradigms have been extensively investigated, the receptor activation, regulation, and cellular signaling mechanisms underlying these sexually dimorphic responses is poorly understood.

Glucocorticoid Receptor Activation in the Hippocampus The GR is an intracellular steroid hormone receptor that is bound by lipophilic glucocorticoids that have passed through the lipid-bilayer of the cell. Unbound, the GR resides within the cellular cytosol (McEwen, 1982). Once bound by cortisol or corticosterone, the GR will homodimerize and translocate to the nucleus where it binds its respective hormone response element to activate or repress transcription of stress sensitive genes (Wranges et al.,1989). However, evidence exists to suggest that GRs may also be membrane bound upon which their activation regulates rapid intracellular signalling cascades (Tasker et al., 2006). Interestingly, GR concentrations within the hippocampus have been shown to be significantly higher than in any other brain region

(McEwen, 1982).

Functional Effects of GR Activation in the Hippocampus

The hippocampus is a limbic structure that plays a critical role in episodic and spatial learning and memory. It is well-established that the hippocampus abundantly expresses both the MR and GR which bind adrenal corticosteroids to regulate neuronal plasticity and neuronal excitability (McEwen & Milliken 1999; McEwen & Milliken, 1997).

By modulating long term potentiation, neurogenesis, and neuronal structure and function, activation of these receptors are subsequently able to modulate cognition, and

31 performance in learning and memory tasks (Schoenfeld & Gould, 2013). For example, both acute and chronic stressful stimuli have been shown to induce atrophy of apical

CA3 dendrites while reducing cell proliferation in the dentate gyrus, dendritic arborization, and spine density (Ferragud et al., 2010; Gould et al., 1992, Gould et al.,

1998; McKittrick et al., 2000; Watanabe et al., 1992).This reduction in cell proliferation is directly correlated with stress and depressive-related behaviours in mouse models

(Mitra et al., 2006; Snyder et al., 2011). This reduction in neurogenesis has been associated with impairments in spatial learning and contextual fear conditioning (Saxe et al., 2006; Shors et al., 2001; Shors et al., 2002; Snyder et al., 2005). Short term glucocorticoid exposure has also been shown to result in reversible cognitive deficits in spatial and episodic memory in both animal models and humans while prolonged stress exposure and elevated glucocorticoids have been shown to have more permanent detrimental effects on neuronal structure, function, and hippocampus mediated cognition (Lupien &, McEwen, 1997; McEwen & Sapolsky, 1995). For example, prolonged stress exposure has been shown to result in CA3 dendritic atrophy and may accelerate the loss of CA1 hippocampal pyramidal neurons (Kerr et al., 1991). However, these findings vary between species, age, and sex (Castilla-Ortega et al., 2011).

Stressful stimuli have been shown to exhibit dramatically different effects in the female and male hippocampus. In response to acute intermittent tail shocks, male rats exhibit enhanced performance in hippocampus dependent associative learning tasks

(Shors et al., 2001; Wood & Shors, 1998). However, when subjected to the same stressful stimuli, age-matched female counterparts exhibit impairments in their respective associative learning (Shors et al., 2001; Wood & Shors, 1998).

Figure 1.7 Sexually differentiated effects of stress exposure on conditioned eyeblink responses in male and female rats. A) 24 hours after exposure to the acute tail shock stressor, male rats exhibit improved acquisition of the conditioned response when compared to their non-stressed counterparts. B) Females exhibit significantly impaired acquisition of the conditioned response 24 hours after exposure to the acute tail shock stressor when compared to their non-stress counterparts (Wood & Shors 1998). Intriguingly, pre-training the females eliminates the male’s advantage in acquisition, suggesting that the stress associated with the novelty of this task may selectively impair females (Perrot-Sinal et al., 1996). This impairment in the female’s performance is variable depending on the presence and concentration of circulating estrogens during various phases of the estrous cycle (Wood et al., 2001; Wood & Shors

1998). As estrogens have been shown to potently modulate spine density, this may mediate the sex differences underlying hippocampal responsiveness and performance in hippocampus dependent paradigms. In fact, males have been shown to exhibit up to

25% increase in apical dendritic spine density 24 hours after exposure to an acute stressful stimulus when compared to their control treated counterparts (Shors et al.,

2001). In contrast, females exposed to the same tail shock stressor during their diestrus stage exhibit an approximate 20% reduction in dendritic spine density 24 hours later during their proestrus (Shors et al., 2001). This indicates that the detrimental effects of

33 stress are most obvious in females when estrogen levels are high. These findings suggest that exposure to an acute behavioural stressor prevents the enhancement of spine density that normally occurs during the high estrogen proestrus phase (Shors et al., 2001). It is evident that the physiological stress response that activates hippocampal

GR may modulate the estrogen mediated neurotrophic effects within the hippocampus and may explain why the female hippocampus is uniquely vulnerable to the detrimental effects of stress.

Stress and Estrogen Interactions in the Hippocampus

The innate mechanistic differences likely contribute to the profound sex differences in the prevalence, onset, and severity of many neuropsychiatric and neurodegenerative disorders. For example, women are more often diagnosed with generalized anxiety, mood, and panic disorders, while males are more likely to be affected by schizophrenia and addictive behaviours (Abel et al., 2010; Barnett et al.,

1987; Bebbington, 1996; Kudielka & Kirschbaum, 2004; Weich, et al., 2001). While these findings may vary across populations and demographics, it is consistently reported that women experience higher prevalence and degrees of anxiety than men

(Altemus, 2006). It has been postulated that this may be associated with sex differences in HPA axis responsivity. Specifically, there are well established interactions between the HPA and HPG axes throughout both development and adulthood.

Developmental interactions between the HPA and HPG axes often alter axis responsiveness later in life (Fernández-Guasti et al., 2012). For example, female offspring have been shown to be significantly more vulnerable to the detrimental effects

34 of prenatal stress showing increased HPA reactivity, anxiety, depression, and loss of hippocampal volume in adulthood (Frye & Wawrzycki, 2003; Koehl et al., 1999;

McCormick et al., 1995; Schmitz et al., 2002; Sternberg, 1999.; Szuran et al., 2000.;

Takahashi et al., 2001; Walf & Frye, 2006; Weinstock et al., 1992). Moreover, female rat offspring that have been exposed to gestational stress exhibit attenuated estradiol mediate antianxiety and anti-depressive behaviour (Frye & Orecki, 2002; Frye & Orecki,

2002; Frye & Wawrzycki, 2003; Walf & Frye, 2006).

Postnatally, androgens have been shown to attenuate HPA axis reactivity while estrogens have been shown to increase HPA axis activity (Burgess & Handa, 1992;

Goel et al., 2014; Peiffer & Barden, 1987). Chronic treatment with estradiol has also been shown to reduce GR expression within the hippocampus of female ovariectomized rats impairing GR’s ability to regulate HPA axis reactivity (Burgess & Handa, 1993). It has also been shown that estradiol treated ovariectomized rats exhibit higher corticosterone concentrations in response to a foot shock stressor and the corticosterone negative feedback is prolonged in the presence of estradiol treatment

(Burgess & Handa, 1992). Intriguingly, estradiol has also been shown to upregulate corticosteroid binding globulin thereby reducing available circulating glucocorticoids

(Brien, 1981; Ycaza Herrera & Mather, 2015). However, these modulatory interactions between the HPA and HPG axes are known to be bi-directional. For example, glucocorticoids have been shown to interfere with normal HPG axis estradiol activity at the level of the hypothalamus, pituitary, and ovaries and this appears to be dependent on whether the stress exposure is acute or chronic (Gore et al.,2006; Tetsuka, 2007;

Tilbrook et al., 2000; Ycaza Herrera & Mather, 2015).

Rationale It is well established the females exhibit an increased prevalence of anxiety and depressive disorders during their reproductive years than their male counterparts (Bao

& Swaab, 2002). Females with major depressive disorder exhibit significantly lower levels of plasma estradiol suggesting an impairment or loss of this estrogen mediated protective mechanism in the hippocampus (Young et al., 2000) and stress responsivity varies greatly depending on the phase of the menstrual cycle. However, with age, plasma cortisol levels steadily increase in both males and females, with females showing higher total plasma concentrations in comparison to their male counterparts

(Kudielka et al., 2004). This age-related rise in circulating stress hormones has been directly attributed to increased atrophy of the hippocampus and it has been demonstrated that females are particularly vulnerable to this stress induced loss of hippocampal volume (Lupien et al., 1998). With menopause, this loss of estrogen mediated neuroprotection may contribute to the female’s susceptibility to neuronal damage and loss of cognition that occurs with age. However, increased exposure to glucocorticoids during the reproductive years may itself impair estrogen mediated signaling and neurotrophic and neuroprotective effects within the hippocampus that may help explain why the female hippocampus is particularly vulnerable to the detrimental effects of stress. For example, mRNA expression of the classical ERα has been shown to be significantly reduced in the amygdala of patients diagnosed with major mental illnesses (Perlman et al., 2004) while glucocorticoid treatment has been shown to cause a significant increase in ERβ protein expression within the hypothalamus of adult female rats (Suzuki & Handa, 2004). Moreover, regulation of ERK and JNK signalling has been shown to be sensitive to glucocorticoid exposure and treatment in many different

36 tissues. For example, glucocorticoid treatment was shown to inhibit both JNK and ERK signalling pathways in human endothelial cells (González et al., 1999). ERK phosphorylation was shown to be significantly increased in the hippocampi of prenatally stressed rats (Cai et al., 2007) while forced swim tests have been shown to result in a much more moderate regulation of JNK in the adult male rat hippocampus (Shen et al.,

2004). However, these findings do not completely explain the sex specific cognitive impairment and loss of estradiol mediated neurotrophic effects in females after exposure to acute stressors. Investigating GPER regulation by glucocorticoids may provide a potential avenue for further elucidating the stress induced loss of functional estradiol signalling in the female hippocampus. To our knowledge, the regulatory effects of glucocorticoid treatment on GPER expression and functional signalling has yet to be investigated in hippocampal neurons.

Hypothesis and Objectives Given the sexually differentiated effects on dendritic morphology, learning, and memory in response to stressful stimuli, it was hypothesized that in vitro treatment with the synthetic GR agonist, DEX, will down-regulate GPER steady state protein levels and functional phosphorylation of JNK in novel in vitro lines of immortalized hippocampal neurons (mHippoE-14 and mHippoE-18). These novel cell lines were originally derived from the hippocampal neurons of a female and male embryonic Swiss Webster mouse, respectively (Gingerich et al., 2010). Since their creation, these cell lines have been used to investigate mechanisms of estradiol and G-1 mediated activation of GPER.

Estradiol was shown to activate the P13/AKT and signal transducer and activator of transcription 3 (STAT3) signaling pathways in both mHippoE-18 and mHippoE-14 cell

37 lines and pre-treatment with G-1 was able to mimic the neuroprotective effects of estradiol (Gingerich et al., 2010). While both of these cell lines have been shown to express GPER, the effects of glucocorticoid treatment on GPER expression and functional signalling have yet to be investigated. To our knowledge, these cells lines have not yet been used to investigate stress and gonadal hormone interactions. As a result, three objectives were developed to test the in vitro hypothesis. Based on the results obtained, a fourth objective was developed to investigate the in vivo effects of glucocorticoid treatment on hippocampal GPER expression.

1) To characterize in vitro neuronal cell models (mHippoE-14s and mHippoE-18s) for

mRNA expression of important neurosteroid receptors, transcription factors, and

neurotransmitter receptors.

2) To determine the effects of glucocorticoid treatment on GPER protein steady state

levels.

3) To determine the effects of glucocorticoid treatment on GPER functional activation of

JNK intracellular signaling cascades in mHippoE-14s.

4) To investigate glucocorticoid induced regulation of hippocampal GPER expression

and functional activation of JNK using an in vivo ovariectomized CD1 mouse model.

CHAPTER 2: CHARACTERIZATION OF RECEPTOR MRNA EXPRESSION IN MHIPPOE-14 AND HIPPOE-18 IMMORTALIZED HIPPOCAMPAL NEURONS

Introduction The hippocampus is a vital component of the learning and memory circuitry in the

central nervous system and, therefore, has been one of the most extensively

investigated brain regions (Frick et al., 2015). Unique neuronal cell types such as

pyramidal neurons, GABAergic interneurons, and granule cells exist within specific sub-

regions of the hippocampus including the cornu ammonis 1 (CA1), CA3, and dentate

gyrus (Andersen et al., 1971). Models of hippocampal neurons have been used to

investigate the underlying mechanisms that regulate neurogenesis, neurodegeneration,

and long term potentiation (Holopainen, 2005; Lopes et al., 2010). To assess relevant

gene expression, protein expression, cellular signaling, and pharmacological effects,

neuronal cell lines are often used for their simple manipulation and ease of control in in

vitro environments (Su et al., 2012). Many in vitro studies rely on proliferative

neuroblastoma and neuron-derived cell lines that lack many of the fundamental

hormone receptor and neurotransmitter system components that are found in vivo

(Gordon et al., 2013; Korecka et al., 2013; Kovalevich & Langford, 2013). While primary

neuron cultures may be generated from an in vivo rodent model, these models lack the

ability to proliferate and, therefore, are short lived in vitro (Gordon et al., 2013).

Although each neuronal culture method offers unique properties, the individual cell lines

are each subject to their own practical and theoretical limitations (Gordon et al., 2013;

Kovalevich & Langford 2013).

Many primary neuron models have been used as a theoretical gold standard to investigate hippocampal cellular mechanisms in vitro ( Hammond et al., 2001; Su et al.,

2012). These primary models are often appealing for their closer resemblance to in vivo neuronal morphology, receptor characteristics, and signaling mechanisms (Gordon et

40 al., 2013). Although the results generated by these models may best recapitulate theoretical generalizability, they are technically challenging to generate and maintain

(Gordon et al.,2013). Typically derived from mammalian embryonic central nervous system tissue, primary neurons can no longer undergo propagation and therefore, are relatively limited in lifespan (Kovalevich & Langford, 2013). Due to the inherent variability in these culture models, consistency between ratios of different astrocytes, oligodendrocytes, and neuronal phenotypes may be inconsistent (Gordon et al., 2013).

Additionally, transfection of primary neurons proves to be much more challenging than transfection in a cell line (Gordon et al., 2013). Practically, these models are subject to significant limitations through their inherent variability and reduced life-span as well as the expertise, time and cost required to generate these cultures on a continual basis

(Gordon et al., 2013).

To overcome the practical and costly limitations of primary neuron models, many researchers rely on cloned neuroblastoma cells that continuously proliferate (Kovalevich

& Langford, 2013). The SH-SY5Y (ATCC CRL-2266 TM) cells are a commonly used model in neurobiology (Kovalevich & Langford, 2013). These cells are a third generation sub-cloned SK-N-SH cells, isolated from a metastatic bone tumor biopsy in a female neuroblastoma patient (Biedler et al., 1973). The original SK-N-SH cells contained both neuroblast-like and epithelial-like cells and, therefore, present with two phenotypically and morphologically distinct cell populations (Ross, Spengler, and Biedler 1983). The neuroblast-like cells usually demonstrate catecholaminergic resemblance through their tyrosine hydroxylase and dopamine-β-hydroxylase expression (Ross et al., 1983).

However, the epithelial-like cells appear to lack these catecholaminergic properties

(Ross et al., 1983). The SH-SY5Y sub-cloned line has been shown to consist only of the homogenous neuroblast-like cells that do express dopamine β-hydroxylase, with negligible choline acetyl-transferase, acetyl cholinesterase and butyryl-cholinesterase activity (Biedler et al., 1973).

In culture, undifferentiated SH-SY5Y cells continuously proliferate in a non- synchronous fluctuating manner, while presenting phenotypically with non-polarized cell bodies and few truncated processes (Påhlman et al., 1981). SH-SY5Y cultures contain both adherent cells and floating cells, with most studies relying only on the adherence cells (Kovalevich & Langford, 2013). In this undifferentiated state, the cells are highly dopaminergic and catecholaminergic and have also been described to express muscarinic and nicotinic acetylcholine receptors (Adem et al., 1987; Kume et al., 2008;

Lopes et al., 2010; Tosetti et al.,1998). However, the SH-SY5Y cells may be differentiated to more closely resemble the morphology of a mature neuron, while maintaining expression of only specific neuronal markers (Påhlman et al., 1981).

Retinoic acid-induced differentiation into a phenotype more closely resembling morphologically mature neurons appears to synchronize, and reduce the proliferative cycles between cells (Ross et al.,1983). Differentiated SH-SY5Y cells exhibit properties of increased excitability and neurite process extensions in addition to increased expression of neuron-specific enzymes and neurotransmitter receptors (Lopes et al.,

2010; Påhlman et al., 1981; Tosetti et al.,1998). Differentiation can be specifically induced to drive SHSY-5Y cells towards cholinergic, adrenergic, or dopaminergic neuronal phenotypes (Kovalevich & Langford, 2013). However, Grimm et al., (2016)

42 demonstrated that the process of differentiation may fundamentally alter the effects of neurosteroids in the SH-SY5Ys.

Following treatment with neurosteroids estradiol, progesterone, estrone, testosterone, and 3α-androstandiol, undifferentiated cells appear to respond positively through increased mitochondrial biosynthesis of ATP (Grimm et al., 2016). However, mitochondrial function in differentiated cells appears relatively unresponsive to all neurosteroids, with the exception of testosterone (Grimm et al., 2016). While SH-SY5Y cells have been shown to express progesterone and classical estrogen receptors, they also appear to be an appropriate model for investigating the effects of androgens (Butler et al., 2001; Grimm et al., 2014; Grimm et al., 2016; Melcangi et al.,1993; Mendell,

2014). Due to their response to androgenic treatment and considerable concentrations of 5α-reductase and 3α-HSD activity, previous studies have used the SH-SY5Ys to characterize the effects of androgenic metabolites (Butler et al., 2001; Melcangi et al.,

1993; Mendell, 2014). SH-SY5Y cell lines have been shown to be relatively sensitive to stress mediating hormones through their expression of corticotrophin releasing hormone receptor 1 (CRHR1) and the glucocorticoid receptor (GR) (Glick et al., 2000; Schoeffter et al., 1999). However, dimethyl sulfoxide (DMSO), a commonly used vehicle for steroid treatment in cell culture, appears to have inherent effects on cell signalling such as the phosphorylation of the extracellular signal-related kinase/mitogen-activated protein kinase (ERK/MAPK) pathway, even at very low concentrations (Mendell, 2014). This may impact the proper interpretation of the effects of treatment seen in SH-SY5Y cells, especially when investigating steroidal effects (Mendell, 2014).

Neurotransmitter synthesis and release is also significantly altered in the process of retinoic acid induced cellular differentiation (Korecka et al., 2013). Specifically, differentiated SH-SY5Ys demonstrate an approximate 70% reduction in key enzymes involved in the neurotransmission of norepinephrine and serotonin (Korecka et al.,

2013). While the dopamine neurotransmitter system appears to be enhanced in the SH-

SY5Ys, the markers and receptors for histamine, gamma-aminobutyric acid

(GABA)ergic, and glutamatergic systems are relatively lacking in both the undifferentiated and differentiated SH-SY5Y models (Korecka et al., 2013). As a result, the relevance of the mechanisms identified in SHSY5Y cells have been disputed due to the relative lack of specific GABAergic and glutamatergic receptor subunits (Andersson et al.,2015; Korecka et al., 2013; Mendell et al., 2018). Specifically, the GABA type A receptor (GABA-AR)-β3, GABA-AR-α1 subunits and the α-amino-3-hydroxyl-5-methyl-4- ixoxazolepropionic acid (AMPA) selective glutamate receptor 2 (GRIA2) and N-methyl-

D-aspartic acid (NMDA) selective glutamate receptor 1 (GRIN1) subunits are expressed, but do not constitute expression of a normal functioning receptor conformation (Korecka et al., 2013; Mendell et al., 2018). Generally, their neuroblast-like morphology, differences in steroid sensitivity, and relative lack of neurotransmitter receptors reduces the generalizability of the findings obtained from studies using these cells (Korecka et al., 2013; Kovalevich & Langford, 2013).

A less commonly used male derived neuroblastoma IMR-32 cell line appears to express multiple cholinergic receptor subunits (α3, α4, α5, α7, β2, and β4), while lacking expression of important sex hormone receptors (Groot Kormelink & Luyten, 1997; Su et al., 2012). In contrast to the SH-SY5Y cells, the IMR-32 line has been well-

44 characterized for its robust GABRA (α 1, α 3, α 4, β 1, β 3, γ2, and δ) subunit expression and functional activation ( Anderson et al.,1993; Mendell et al., 2018;

Meulenberg &, Vijverberg 2003; Noble et al., 2006). The typical functional GABA receptor pentamer usually requires 2α, 2β and 1γ subunits and, therefore, the IMR-32s are frequently used to model synaptic inhibition (Zhou & Smith, 2007).

HT-22s are immortalized murine hippocampal neuronal precursors that have been shown to express functional cholinergic properties and marked insensitivity to glutamate (He et al., 2013; Liu et al.,2009). Therefore, these models are frequently used to model resistance to excitotoxicity (He et al., 2013). The HT-22s lack neuronal morphological properties, neurite outgrowth, and expression of both NMDA and cholinergic receptors (He et al., 2013). Therefore, these cells may represent a poor model of mature hippocampal neuron sensitivity to excitation and long term potentiation, a hallmark of hippocampal function (He et al., 2013). However, immortalized cells undergo rapid proliferation that requires relatively minimal maintenance, in comparison to the primary neuron model (He et al., 2013). Given the limitations of the previously discussed cell models, a non-tumor derived immortalized hippocampal cell line that could be characterized for representative morphology, and neurosteroid, GABAergic, and glutamatergic receptor expression may provide the means to best investigate in vitro hippocampal neurons and mechanisms (Gingerich et al., 2010).

Gingerich et al. (2010) first described the creation of relatively novel immortalized cell lines taken from samples of adult and embryonic mouse hippocampi. As consistent with Belsham et al. (2004) , pregnant female Swiss Webster mice were anaesthetized on embryonic day 18 (Gingerich et al., 2010). Offspring were then sacrificed for the

45 dissection of the fetal hippocampi that were subsequently pooled, triturated, and cultured (Gingerich et al., 2010). These primary cultures were transfected and incubated with the neomycin resistance gene from the pZIPNeo SV(X)1 vector and the recombinant murine retrovirus harboring simian virus (SV40) T-antigen (Gingerich et al.,

2010). Cultures were treated with the aminoglycoside antibiotic geneticin and colonies determined to be resistant were then further expanded and sub-cloned to generate four cell lines: mHippoE-2, -5, -14, -18 (Gingerich et al., 2010).

The mHippoE-14 and mHippoE-18s represent two phenotypically distinct clonal cell lines from the original heterogeneous E-18 hippocampal population, that share similarities in general neuronal and neurite morphology (Gingerich et al., 2010).

Following the generation of both lines, they were individually characterized for receptor mRNA expression profiles (Gingerich et al., 2010). Embryonic mHippoE-14 cell lines were demonstrated to express neuron specific enolase (NSE), estrogen receptor alpha

(ER-α), ER-β, G-protein coupled receptor 30 (GPER), neurotrophic tyrosine kinase receptor type 1 (TrkA), neurotrophic tyrosine kinase receptor type 2 (TrkB), insulin receptor(IR), leptin receptor (obRb), growth hormone secretagogue receptor(GHSR), neuropeptide (NPY), brain derived neurotrophic factor (BDNF), NMDA receptor 1

(NMDA-R1), AMPA receptor 3 (AMPA-R3), AMPA-R4, microtubule-associated protein

2 (MAP2), and androgen receptor (AR) (Gingerich et al., 2010). Embryonic mHippoE-18 cell lines were shown to express: NSE, spermiogenesis specific transcript on the Y

(SSTY1), ER-α, ER-β, GPER, TrkA+B, IR, obRb, GHSR, NPY, BDNF, proglucagon,

NMDA-R1, AMPA-R3, and MAP2 (Gingerich et al., 2010). These cell lines represent phenotypically unique and rapidly proliferating hippocampal neurons that demonstrate

46 potential for their use in models of neurosteroid, GABAergic, and glutamatergic interactions and signaling mechanisms. The lack of additional characterization of mHippoE-14 and mHippoE-18 cell lines may be specific to its use as a model for steroid hormone effects and interactions (Evans et al., 2016; Gingerich et al., 2010). With this in mind, the primary objective of the current research was to further characterize the mHippoE-14 and mHippoE-18 cell lines for mRNA expression of additional neurosteroid,

GABAergic, and glutamatergic receptors. Through multiple conventional polymerase chain reactions (cPCR), the findings of this study allude to the characterization and validation of this model for relevant investigations of neurosteroid interactions in the hippocampus.

Experimental Procedures Cell Culture Conditions

MHippoE-14 and mHippoE-18 aliquots were flash-thawed from storage at -80° C

and carefully transferred to their respected flasks where they were suspended to a total

volume of 10 mL of culture media. Culture media contained Dulbecco’s modified Eagle

medium (DMEM) with 10% fetal bovine serum (FBS) and 1% penicillin-streptomycin

(pen-strep; 100 units penicillin and 100 units streptomycin). MHippoE-14 and mHippoE-

18 cells were both maintained in their own distinct cell culture flasks and housed within

a 37°C/ 5% CO2 incubator. To maintain the rate of cellular proliferation, metabolic

output, and nutrient consumption, mHippoE cell lines were passaged once the culture

flasks reached ~80% density, approximately every 48 hours. Culture media was

aspirated from the flask and 2 mL of 0.25% trypsin 0.53% EDTA solution was added to

lift adherent cells from the flask. Once visibly detached, culture media was added to

suspend the cells. Cellular suspension was then removed from the flask and one

millilitre was plated on 60 mm culture plates, with a record of their respective passage

number. Plates were incubated until 80% confluency was reached.

RNA Isolation

When confluent, media was aspirated from the culture plates and 700µL of

Qiazol lysis buffer (Qiagen-Toronto, ON, Canada) was added. Cellular contents suspended in Qiazol were collected using a cell scraper and pipette and stored in 1.5 mL Eppendorf tubes. The total ribonucleic acid (RNA) from each mHippoE-14 and mHippoE-18 culture lysates was processed for RNA isolation through a series of buffer incubation and elution steps. Isolation steps were consistent with the directions provided in the miRNeasy Mini Kit (Qiagen) and on-column DNase digestion (RNase-free DNase-

Qiagen). The final RNA was collected through 40µL elution volume of RNase-free water. Subsequently, RNA concentrations and qualities were confirmed through µg/µL and 260/280 NanoDrop spectrophotometer analyses (ThermoFisher Scientific,

Burlington, Ontario, Canada). The total RNA was obtained for each mHippoE-14 and mHippoE-18 at multiple passage numbers to ensure consistent gene expression patterns with increasing age of the cells. The remainders of each of the samples were then stored at -80° C before reverse transcription processing.

Reverse Transcription and cDNA Synthesis

After thawing, RNA concentration values were used to aliquot 16µL of RNA combined with RNase free water so that each reaction contained 1000 ng total RNA. An additional 4µL of qScript complementary deoxyribonucleic acid (cDNA) Supermix

(Quanta Biosiences) was added to each diluted sample to total 20 µL reaction volumes.

Following the qScript cDNA Supermix (Quanta Biosciences) protocol, RNA was reverse transcribed to generate cDNA. Subsequently, each sample was further diluted using

10mM Tris, pH 8.8, 0.1 EDTA (TE) buffer to a final 10ng/µL concentration. Samples were stored at -20° Celsius until required for conventional PCR analyses.

Conventional Polymerase Chain Reactions and Visualization

Conventional PCR was performed using a MyCycler Thermal Cycler (Bio-Rad) to determine expression of neurosteroid receptors, transcription factors, and glutamatergic and GABAergic receptors that have been shown to have steroid interactions. PCR reactions were performed using 25µL reaction volumes, made up of 20µL master mix solution that contained 10x PCR buffer, MgCl2, 0.2mM of each dNTP, 0.2 µM of each forward (Fwd) and reverse (Rev) primer (10 μM concentration), 1.25 units AmpliTaq

Gold DNA polymerase (Thermofisher Scientific), in addition to 5 µL of the respective cDNA template (n=5-10 samples per target gene). Reactions were run for 5 minutes at

95°C for initial denaturation, followed by 35 cycles of 30 seconds of 95°C to complete denaturation, annealing for 1 minute at 55°C-57°C (dependent on the primer) for primer annealing and 1 minute at 72 °C for extension, and a final 5 minutes at 72 °C to complete the extension (Table 1). Specifically, PCR reactions were performed for the following targets: hypoxanthine-guanine phosphoribosyl transferase (HPRT), GR, mineralocorticoid receptor (MR), luman, luman recruitment factor (Lrf), CRHR1, CRHR2, thyroid hormone receptor (THR)α, THRβ, AR, GPER, AMPA-R3, AMPA-R2, NMDA-R1,

GABA-AR-A-α1, GABA-AR-α4, GABA-AR-α6, GABA-AR-β2, GABA-AR-β3, GABA-AR-

δ, and GABA-AR-γ2 using the primer pairs listed in table 2.

Table 1. Cycling conditions for initial denaturation, cycling of denaturation, primer annealing and extension, and final extension stages of conventional polymerase chain reactions (cPCR).

Number of Cycles/Time Temperature

Initial Denaturation 5 minutes 95° C

Denaturation 30 seconds 95° C

Primer Annealing 1 minute 55°- 57° C X 35 cycles

Extension 1 minute 72° C

Final Extension 5 minutes 72° C

DNA was purified from one of the PCR samples using the QIAquick PCR

Purification Kit purchased from QIAGEN. Purified DNA samples and respective primers were prepared to 10 ng/uL and 5 μM concentrations, respectively. Samples were sent for amplicon sequencing at the University of Guelph’s Advanced Analysis Centre

Genomics facility. Successful signals were read as Applied Biosystems Sequence

Traces using the SnapGene Viewer 4.3.10 software. FASTA files were saved as for each forward and reverse primer and run through the Basic Local Alignment Search

Tool (BLAST) with the National Center for Biotechnology Information’s nucleotide reference sequence for each respective mus musculus gene (Lipman and Pearson

1985; Pearson and Lipman 1988).

A 2% agarose gel (2.5 mL 50x tris acetic acid EDTA (TAE) buffer, 117.5 mL

MilliQ water, and 1.8 g agarose) containing 4.8 µL of ethidium bromide was used to visualize the final PCR products. Each final PCR sample (20 µL) was combined with

2µL of loading dye and loaded into a well of the agarose gel. DNA ladders were run on each gel to estimate the size of the DNA products. Gels were run in 1x TAE buffer at

90V for 45-60 minutes and subsequently visualized under ultraviolet transillumination for the presence of receptor gene expression. Illumination settings were adjusted for each gel and the Liscap software was used to capture the final images.

Table 2. Primer sequences used to identify transcription factors and steroid, glutamatergic, and GABAergic mRNA receptors by conventional PCR. Primer sequences were generated using Primer3 and National Center for Biotechnology Information’s Primerblast software.

Primers Direction Sequence (5'-3') Primers Direction Sequence (5'-3')

mHPRT Fwd TGACACTGGCAAAACAATGCA mAMPAr3 Fwd AACACGGTACAGGAGCACAG (Gilsbach, 2006)

mHPRT Rev GGTCCTTTTCACCAGCAAGCT mAMPAr3 Rev TCTGGAGAACTGGGAGCAGA

mLuman Fwd AGGTGTATGTCGTGGGGCTG mAMPAr2 Fwd GGGGACAAGGCGTGGAAATA

mLuman Rev CGCTGCTGGTTTTGTTGGCT mAMPAr2 Rev CCAATCTTCCGGGGTCCATT

mLrf Fwd AGCAAGGCCAAGGTCAAGGA mNMDAr1 Fwd CAACGACCACTTCACTCCCA

mLrf Rev TCAGTGTTGAGGCCCCACAA mNMDAr1 Rev GACGCGCATCATCTCAAACC

mCrhr1 Fwd GACCAAACTCCGAGCATCCA mGABA-AR--α1 Fwd GGTTGACCGTGAGAGCTGAA

mCrhr1 Rev GCGGACCTCACTGTTCAGAA mGABA-AR-α1 Rev CTACAACCACTGAACGGGCT

mGR Fwd CAATAGTTCCTGCCGCGCTG mGABA-AR-α4 Fwd GAAGTTCCAGCTGCTCCTGT

mGR Rev GGCGCCCACCTAACATGTTG mGABA-AR-α4 Rev GCTGGTCTTGCTGGAGTGAT

mMR Fwd CAAGGTCACACAGCCCCGTA mGABA-AR-α6 Fwd AAGCGTCTGAATCCCTGCAA

mMR Rev GAGGAGCGCAGAGGGACATT mGABA-AR-α6 Rev GGCGTTCTACTGAGGGCTTT

mAR Fwd GGTTGGCGGTCCTTCACTAA mGABA- AR-β2 Fwd AAGATGCGCCTGGATGTCAA

mAR Rev CATCCTCACACACTGGCTGT mGABA-AR- β2 Rev TCCGTCTAGTTGGGGAGAGG

mGPER Fwd TCTACCTAGGTCCCGTGTGG mGABA-AR-β3 Fwd ACAGGTGCCTATCCTCGACT

mGPER Rev GGTGTAGAGGCAGGAGAGGA mGABA-AR-β3 Rev TGGTGAGCACGGTGGTAATC

mCB1 Fwd TCTGCTTGCGATCATGGTGT mGABA-AR-γ2 Fwd TGGAAGCGCAGTTCTGTTGA

mCB1 Rev GCATGTCTCAGGTCCTTGCT mGABA-AR-γ2 Rev CAGGACAGGACCACGATGAG

mTHRα Fwd AGATCACCCGGAATCAGTGC mGABA-AR-δ Fwd ACGTCTTTGTGTTTGCTGCC

mTHRα Rev ATCTCCTCCTTTCGCCTCCT mGABA-AR-δ Rev CGACGGGAGATAGCCAACTC

mTHRβ Fwd CCACTATCGCTGCATCACCT

mTHRβ Rev CACCAAGTCTGTTGCCATGC

Results Using 20x and 10x magnification, it was shown that both mHippoE-14 female derived and mHippoE-18 male-derived cell lines share similarity in overall neuron and neurite morphology while being morphologically distinct from that seen in typical neuroblastoma derived cell lines (Figure 2.1a). Figure 2.1b depicts representative ethidium bromide-stained gel images following PCR amplification with various primer pairs. Housekeeping HPRT expression appeared visually consistent across multiple passages collected from passage 20 through 30 of both the mHippoE-14 and mHippoE-

18 cells lines. Receptor mRNA characterization in the mHippoE-14 immortalized cells revealed positive expression of GR, MR, Luman, AR, GPER, cannabinoid receptor 1

(CB1), THRα, and THRβ. Glutamatergic and GABAergic receptor expression was limited to the AMPA-R3, GABA-AR-α1, and GABA-AR-β3 subunits. Receptor mRNA characterization in the mHippoE-18 immortalized cells also revealed positive expression of GR, MR, Luman, GPER, CB1, THRα, and THRβ, while AR expression was lacking.

Glutamatergic and GABAergic receptor expression was even more limited in the mHippoE-18 cells to AMPA-R3 and weak GABA-AR-α1subunit expression. Between cell lines, the mHippoE-14s appear to express a greater variety of neurosteroid receptor targets, including the AR and GABA-AR-β3. A summary of these results is shown in

Figure 2.1c.

All DNA fragments detected following PCR amplification were the appropriate

size based on the predicted base-pair length that would be generated from the region of

DNA targeted by the specific primer pairs. To confirm the identity of the PCR products,

the DNA was purified from the PCR reactions and sent for Sanger sequencing. Figure

2.2 shows the confirmation of GPER following Sanger sequencing, with the nucleotide

53 sequence trace shown in Figure 2.2a and the sequence alignment following a BLAST search shown in Figure 2.2b. The sequences for HPRT, MR, GR, AR, GPER, CB1,

THRα, THRβ, AMPA-R3, GABA-AR-α1, and GABA-AR-β3 were confirmed in a similar manner with the results indicated in appendix 1 of this thesis.

Figure 2.1 Characterization of receptor mRNA expression for transcription factors and steroid, GABAergic, and glutamatergic receptors in mHippoE-14 and mHippoE-18 immortalized hippocampal neurons. A) Representative 20x and 10x images of mHippoE-14 and mHippoE-18 morphology. B) Hypoxanthine guanine phosphoribosyl transferase (HPRT) was used as a positive control housekeeping gene for each sample collected from passage 20 (P20) to passage 30 (P30). Representative images of cPCR agarose gel visualization for each specific receptor or transcription factor in the mHippoE-14 and mHippoE-18. Samples were run with cPCR for primers designed to match the glucocorticoid receptor (GR), mineralocorticoid receptor (MR), Luman/Cyclic AMP-responsive element-binding protein3, Luman recruitment factors (Lrf), corticotrophin release hormone receptor (CRHR)-1 and 2, androgen receptor (AR), G-protein coupled estrogen receptor (GPER), cannabinoid receptor type I (CB1), thyroid hormone receptor (THR) α- and β, N-methyl-D-aspartic acid receptor-1 (NMDA-R1), α- amino3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPA-R) subunit 2 and 3, gamma aminobutyric acid A (GABA-AR) receptors α1, α4, α6, β2, β3, δ, and γ subunits in mHippoE-14 and mHippoE-18 lines. C) Respective receptor mRNA expression profile for each mHippoE-14 and mHippoE-18 cell line.

Figure 2.2 Confirmation of correct amplicon sequencing with forward and reverse GPER primers. A) Respective nucleotide applied biosystems sequence trace for the collected purified DNA sample run through Sanger sequencing with the GPER forward primer. B) Output generated by the National Centre for Biotechnology Information’s basic local alignment search tool (BLAST) for mus musculus GPER run with the sample nucleotide FASTA sequence generated by Sanger sequencing using the respective forward and reverse GPER primers. The optimal alignment is indicated by the matching between the query (mus musculus GPER) and the subject (mHippoE sample) nucleotide sequences. Max score indicates total alignment coverage while percent identity indicates degree of alignment between query and subject sequences. Sanger sequencing was used to confirm amplicon sequences for HPRT, MR, GR, AR, GPER, CB1, THRα, THRβ, AMPA-R3, GABRA-α1, and GABRA-β3 (Appendices).

Discussion Current hippocampal cell culture models are relatively limited due to their receptor expression, morphological phenotype, and/or ability to propagate (Gordon et al., 2013). As a result, it is challenging to extrapolate the results obtained from these models to the investigation of regulatory neuroprotective and neurodegenerative mechanisms of disease (Kovalevich & Langford, 2013; Su et al., 2012). The mHippoE-

14 and mHippoE-18 immortalized neurons, two relatively novel cell lines, may be useful for the study of neurotransmission and hormonal interactions due to their receptor expression profile, rapid proliferation, and maintenance (Gingerich et al., 2010). In the present study, both lines, mHippoE-14 and mHippoE-18, expressed GR, MR, Luman,

GPER, CB1, THRα, THRβ, and GABRα1, while the AR and GABRβ3 appear to be expressed in the mHippoE-14s only.

The expression of GR and MR provides a potential avenue for investigating the effects of stress hormone mediated regulation and cellular signaling in both mHippoE-

14s and mHippoE18s. Within the central nervous system, the hippocampus is a primary target for circulating glucocorticoids (Sapolsky, 1987). In vivo, stressful stimuli cause the systemic or de novo endogenous release of glucocorticoids that then bind to their respective high affinity MR and lower affinity GRs within the hippocampus (Kerr et al.,

1992; Reul & De Kloet, 1985; Reul et al.,1987; Reul & De Kloet, 1986). GR and MR activation by glucocorticoids may regulate the transcription of stress-sensitive genes that have been functionally implicated in cognition (Becker et al., 1986; Mifsud & Reul,

2018). While acute glucocorticoid elevation was reported to enhance dendritic morphology within the hippocampus, chronic stimulation of GR and MR may result in significant impairments in neuronal morphology and stress hormone sensitivity (Brown

57 et al., 1999; Sapolsky, 1987). GRs and MRs in the brain have been characterized as transcription factors based on their specific ligand binding properties, molecular structure, and profile of occupancy (Mifsud & Reul, 2018). However, the means by which glucocorticoids may readily access and bind glucocorticoid response elements under baseline conditions and following exposure to stressful stimuli has yet to be comprehensively defined (Mifsud & Reul, 2018).

Luman is an endoplasmic reticulum-bound cellular transcription factor that is highly expressed within the hippocampus (Penney et al., 2017). Luman, itself, is negatively regulated by Lrf (Audas et al., 2008). Martyn et al., (2012) demonstrated that

Lrf functions to downregulate GR activity. Lrf-deficient mice were shown to have significantly reduced circulating corticosterone levels and increased activation of GR, leading to improper attenuation of the stress response (Martyn et al., 2012). Luman deficient mouse models have been shown to exhibit significantly reduced overall stress responses, corticosterone levels, and branching within hippocampal CA3 pyramidal neurons (Penney et al., 2017) These functional effects have been proposed to be the result of an inverse regulatory relationship between Luman and GR expression and activity (Penney et al., 2017). This alludes to the potential role of Luman in regulating hippocampal stress responsiveness and sensitivity through GR modulation (Penney et al., 2017). The mRNA expression of GR, MR, and Luman in both mHippoE-14 and mHippoE-18 cell lines could be used to investigate the regulatory mechanisms between receptor activation and signaling cascades. Intriguingly, for both mHippoE-14s and mHippoE-18s mRNA expression of Lrf was not detected. This lack of Lrf mRNA expression would result in low or no Lrf protein expression in the mHippoE cells which

58 may alter Luman functional activation, providing an interesting caveat of internal control to further delineate the effects of Luman and Lrf activation on regulation of stress hormone receptors and sensitivity in the hippocampus.

These cells have been previously characterized by Gingerich et al. (2010) for the presence and functional activation of both classical ERs, ER-α and ER-β, in addition to the transmembrane GPER (GPER) (Evans et al., 2016; Gingerich et al., 2010). The hippocampus is well-known for its responsiveness and sensitivity to the neurotrophic and neuroprotective effects of classical ER mediated estradiol activation (Frick et al.,

2015). Estradiol mediated ER activation has been shown to play a sexually differentiated role in hippocampal memory function and CA1 dendritic branching

(Fugger et al., 1998; Phan et al., 2011; Phan et al., 2012; Sánchez-Andrade &

Kendrick, 2011). Although GPER is not as well characterized, it has been shown to localize with significant density within dendritic spines of the hippocampus (Akama et al., 2013; Gabor et al.,2015). GPER mediates rapid non-genomic estrogenic effects through activation of the c-Jun N-terminal kinase (JNK) pathways, enhancing the physiological and memory enhancing effects of estrogens (Kim et al., 2016). However, the functional effects and regulatory mechanisms underlying GPER expression and activation remain poorly understood. As the mHippoE-14 and mHippoE-18 cell lines express functional receptors for both the classical ERs and GPER, selective receptor agonists and antagonists could be used to compare respective the effects of activation of the classical ERs with GPER as well as their regulation, and functional signaling under various cellular conditions.

Androgenic regulation of hippocampal structure and function has been a controversial topic for years, with respect to whether these hormones exert limited, suppressive, or beneficial effects (Atwi et al., 2016). Reduced endogenous androgen concentrations have been associated with a predisposition to hippocampal mediated cognitive impairments and neurodegeneration (Drummond et al., 2012). While testosterone has been shown to mediate neuroprotective effects through AR activation, this steroid may also have regulatory effects on separate cellular mechanisms that may indirectly contribute to cognitive impairment (Butchart et al., 2013; Hammond et al.,

2001). Within the hippocampus, testosterone may be readily metabolized to estradiol through the process of aromatization, thereby exerting its effects on relevant estrogen receptors (Baulieu & Mauvais-Jarvis, 1964). Testosterone may also be converted to 5α- reduced metabolites, including the potent androgen dihydrotestosterone, and the

GABA-AR modulator 5α-androstane-3α,17β-diol (Atwi et al., 2016; Melcangi et al.,

1993; Reddy, 2004). Through studies using surgical gonadectomy and aromatase inhibitors, evidence indicates that the androgenic neurotrophic and neuroprotective effects on hippocampal morphology in males is mediated through activation of the AR, rather than the ER (Leranth et al., 2003). As a result, mHippoE-14 expression of AR, in conjunction with relevant estrogen and glucocorticoid target receptors, provides an interesting model for identifying regulatory mechanisms between receptor activation, hormonal sensitivity and cell signalling.

Profound sex differences in hippocampal-mediated learning, memory, and stress responsivity have been observed and, as these mHippoE-14 and mHippoE-18s express relevant androgen and estrogen receptors, this model may provide a dynamic

60 framework to investigate these sex differences (Frick et al., 2015). Specifically, the underlying communication between the hypothalamic-pituitary-adrenal (HPA) and hypothalamic-pituitary-gonadal (HPG) axes are reciprocally modulated (Viau, 2002).

Therefore, the concurrent expression of GR, MR, GPER, and AR may help elucidate regulatory mechanisms and functional cross-talk between sex and stress hormones

(Viau, 2002).

In the context of sexually differentiated stress responses, the concurrent expression of GR, MR, and Luman provides a potential framework to manipulate and determine underlying regulatory functions, signaling interactions, and stress sensitivity in the mHippoE cells. The differential expression of AR between the mHippoE-14 and mHippoE-18s serves as an internal control for comparisons of androgen mediated effects (Gingerich et al., 2010). Therefore, the expression of relevant stress and sex steroid receptors in mHippoE-14 and mHippoE-18 cells provides a valuable model for investigating both stress and sex hormone interactions in the hippocampus.

Endocannabinoids are released post-synaptically to bind presynaptic CB receptors that induce suppression of short or long term neurotransmission (Kawamura et al., 2006). The CB1 and CB2 receptor sub-types, have been determined to play a role within the central nervous system and immune system modulation, respectively

(Howlett et al., 2002). CB1 expression in the central nervous system is heterogeneous, with particularly high expression found in the hippocampus (Davies et al., 2002). CB1 has been shown to localize to presynaptic terminals to inhibit the release of a range of excitatory and inhibitory neurotransmitters (Davies et al., 2002; Kawamura et al., 2006;

Pertwee & Ross, 2002). CB1 activation has been attributed to reducing GABAergic,

61 cholinergic, and noradrenergic neurotransmission (Gifford et al., 2000; Kathmann et al.,

1999; Romero et al., 1997).Within the hippocampus, CB1 activation has also been shown to modulate intrinsic membrane conductance through coupling to calcium channels (Davies et al., 2002; Shen & Thayer, 1998). However, endocannabinoids may play an important role in downregulating high frequency stimulation induced long term potentiation, possibly due to the reduction of excitatory neurotransmission (Misner &

Sullivan, 1999). As the mHippoE-14 and mHippoE-18 cell lines each express CB1, these models could be used to better define the effects of CB1 activation on neurotransmission. Exposure to stressful stimuli has been shown to reduce CB1 expression in both gonadectomized and intact males, while increasing CB1 expression in the dorsal hippocampus of gonadectomized and intact females (Reich et al., 2009).

This sexually differentiated regulation of CB1 could be further explored using the mHippoE-14 and mHippoE-18 cell models that both express relevant stress receptors, since only the mHippoE-14s express AR.

Expression of both THRα and THRβ has been demonstrated in the mHippoE-14 and mHippoE-18s. Functional thyroid hormones are known for their importance in neurological development during the gestational period (Bernal, 2007). Lack of THRβ has been attributed to development of peripheral thyroid hormone resistance and lack of function (Forrest et al., 1996). Lack of THRα has been shown to impair performance in hippocampal dependent anxiety tasks such as the open field test and contextual fear paradigm (Guadaño-Ferraz et al., 2003). Intriguingly, THRα mutant mice have been shown to have fewer GABAergic terminals within CA1 pyramidal neurons and therefore,

THRα may be involved in regulating the structure and neurotransmission of

62 hippocampal neurons (Guadaño-Ferraz et al., 2003). Additionally, circulating thyroid hormones have been shown to mediate an increase in the expression of GR density within the hippocampus, independently of stress hormone mediated effects (Meaney et al., 1987). In conjunction with expressed stress hormone receptors MR, GR, and

Luman, THRα expression may allude to the use of this model to investigate the interactions between glucocorticoids and thyroid hormones (Meaney et al.,1987).

Inconsistent with the findings of the present study, Gingerich et al. (2010) noted expression of the glutamatergic NMDA-R1 subunit in both mHippoE-14 and mHippoE-

18 cell lines. The current study did, however, identify the mRNA expression of AMPA-

R3 in both cell lines, consistent with Gingerich et al. (2010). Although few studies have sought to characterize the receptor expression within these models, in conjunction with the findings of Gingerich et al. (2010), these cells may demonstrate potential resemblance to a glutamatergic neurotransmitter system. GABAergic receptor expression appears to be relatively sparse and, therefore, the mHippoE cells may not be a strong model for investigating inhibitory neurotransmission. However, relatively little characterization has been made to profile respective protein expression and function within these models. With this in mind, the additional mHippoE-14 and mHippoE-18s protein characterization may provide the in vitro framework for investigating the effects of stress, sex, thyroid, and cannabinoid hormonal interactions.

Conclusion Neurobiological investigations have proven challenging to study through tissue culture, as isolated primary neurons are unable to propagate and are, therefore, short lived (Gordon et al.,2013). While many neuroblastoma and neuron-derived cells lines

63 divide at a much higher rate, they often lack many of the fundamental neurotransmitter and hormone receptor systems found in neurons (Kovalevich & Langford, 2013). The present study characterized the presence of many important neurosteroid and related receptors in two novel, immortalized hippocampal neuron lines, mHippoE-14 and mHippoE-18. The receptor expression of MR, GR, Luman, AR, GPER, CB1, THRα and

THRβ provides an important and dynamic framework to investigate hormonal interactions and regulatory effects in a simple and efficient in vitro model.

Unfortunately, the GABAergic receptor expression profile indicates that these lines may be relatively lacking in the expression of many contributors of this neurotransmitter system, while less remains clear about the glutamatergic markers.

However, the neurosteroid receptor expression detected in the respective mHippoE-14 and mHippoE-18 cell lines must be further validated and characterized to confirm that mRNA expression is, in fact, translated into functional protein. While hormonal interactions within the hippocampus have yet to be clearly defined, the current mHippoE-14 and mHippoE-18 may provide an important in vitro model to delineate these effects. Confirming protein expression and receptor activation would additionally validate the use of this model for investigating the regulatory and functional interactions between key neuromodulators.

CHAPTER 3: GLUCOCORTICOID REGULATION OF GPER PROTEIN STEADY STATE LEVELS IN MHIPPOE-14 AND MHIPPOE-18 IMMORTALIZED HIPPOCAMPAL NEURONS

Introduction The G-protein coupled estrogen receptor (GPER), found within the hypothalamus, cortex, striatum, and hippocampus (Gibbs, 2010), mediates some of the rapid non-classical effects of estradiol. GPER contains 7 transmembrane domains and has been shown to be highly expressed within cell bodies, dendritic spines, axons, and terminals of neurons within many brain regions (Akama et al., 2013; Brinton et al., 2015;

Filardo & Thomas, 2005; Waters et al., 2015). Particularly high levels of GPER mRNA expression have been found within the hippocampus itself (Gibbs, 2010) and membrane localization of this receptor is attributed to post-translational modification to its 93 kDa glycosylated form. Upon activation, this membrane localized receptor activates the intracellular c-Jun N-terminal kinase (JNK) signalling cascade (Kim, 2014; Pupo et al.,

2017),which has profound implications for rapidly modulating neuronal structure, function, and, subsequently, hippocampal mediated learning and memory (Gabor et al.,

2015; Kim et al., 2016; Lymer et al., 2017). In fact, the memory enhancing effects of G-1 have been shown to be specific to phosphorylation of JNK and mimic the improvements in object recognition and spatial memory consolidation seen with estradiol treatment

(Kim, 2014; Kim et al.,2016). Specifically, use of G-1 has been shown to enhance dendritic spine density in a dose-dependent fashion within cornu ammonis 1 (CA1) pyramidal neurons of ovariectomized female mice within just 40 minutes (Gabor et al.,

2015). Use of G-1 in immortalized hippocampal neurons has also indicated a role for

GPER activation in attenuating glutamate induced neurotoxicity (Gingerich et al., 2010).

As a result, hippocampal expression and activation of GPER is necessary for mediating both rapid neurotrophic and neuroprotective effects of estradiol.

However, these estradiol-mediated effects of GPER activation are sensitive to modulation by the physiological stress response. Intriguingly, activation of GPER has been shown to elicit sexually differentiated anxiety behaviours that are dependent on circulating gonadal hormones (Hart et al., 2014; Kastenberger et al., 2012). For example, GPER activation in ovariectomized female and intact male mice elicits anxiogenic responses while G-1 treatment results in anxiolytic effects in intact female mice and orchidectomized male mice (Hart et al., 2014; Kastenberger et al., 2012).

Previous studies have also demonstrated that GPER expression and activation in ovariectomized female mice reduces pain-related anxiety and that this may be mediated within the basolateral amygdala through GPER mediated modulation of gamma- aminobutyric acid (GABA)ergic and glutamatergic signalling (Tian et al., 2013). The sexually differentiated effects of GPER activation have also been demonstrated using models of ischemia and hypoxia where G-1 treatment proved beneficial in recovery of ovariectomized females and detrimental to intact male mice (Broughton et al., 2014).

This provides evidence to suggest that there may be an underlying interaction between circulating stress hormones and estradiol mediated activation of GPER. While expression of hippocampal classical estrogen receptor (ER)-α and ER-β have been shown to be regulated by and sensitive to stress, there is little research investigating the effects of stress hormone exposure on GPER protein expression within hippocampal neurons (Perlman et al., 2004; Suzuki & Handa, 2004). As a result, the first objective of this work was to determine the effects of glucocorticoid exposure on GPER protein expression 10 minutes and 1, 10, 24, and 48 hours following treatment. Sex differences

67 in glucocorticoid regulated GPER expression were investigated by using both female derived mHippoE-14 and male derived mHippoE-18 immortalized hippocampal neurons.

Experimental Procedures Chemical and Stock Solutions

Stock solutions of dexamethasone (DEX; 500 μM) were prepared in charcoal- stripped fetal bovine serum (CS-FBS; Invitrogen). CS-FBS was used to reduce confounding effects of steroids and small molecules present within the vehicle

(Gurtovenko & Anwar, 2007; Kalluri & Ticku, 2002). Working stock solutions of 10 μM

DEX were prepared by diluting 500 μM stock solutions of DEX into Dulbecco’s Modified

Eagle Medium (DMEM; Invitrogen) containing 1% CS-FBS and 1% penicillin- streptomycin (pen-strep;100 units penicillin + 100 μg streptomycin; Invitrogen).

Cell Culture Conditions

mHippoE-14 and mHippoE-18 immortalized murine hippocampal neurons

(#CLU198, #CLU199, Cedarlane) were flash-thawed from storage at -80° C. Once thawed, mHippoE-14s and mHippoE-18s were carefully transferred into their own 75 mm cell culture flask and incubated in DMEM that contained 10% FBS and 1% pen- strep. Flasks were cultured at 37°C and 5% CO2 and passaged when culture flasks reached ~80% confluency. Cells were lifted from the flasks with 2 mL of 0.25%/0.53 mM trypsin/EDTA solution (1X Hank’s balanced salt solution (HBSS), 1.2 mg/mL NaHCO3,

0.46 mg/mL EDTA, 2.5 mg/mL trypsin, 0.2μL/mL phenol red; Sigma-Aldrich, Oakville,

Ontario, Canada) and resuspended with 10 mL of DMEM culture media. One mL of the suspension was plated onto each 60 mm culture plate (Corning-Fisher Scientific) to

68 which an additional 3 mL of 10% FBS, 1% pen-strep DMEM was added. Once the plated cells were ~75% confluent the cell culture media was aspirated off and replaced with DMEM containing 1% CS-FBS and 1% pen-strep for 16 hours. After 16 hours, DEX treatment began.

Treatment and Protein Collection

For experiments, 500 μM DEX was diluted in 1 mL of DMEM containing 1% CS-

FBS and 1% pen-strep to a final concentration of 10 μM DEX. From this working stock,

4 μL volumes were injected directly into the 4 mL of media per plate to result in a 10 nM dose of dexamethasone. This dose of DEX was determined based on 10 nM free DEX binding 300 fol/mg over a 330 fol/mg total system binding capacity to allow for an approximate 90% receptor saturation in the rodent hippocampus (Spencer et al., 1990).

However, in an in vitro environment lacking endogenous glucocorticoids, the actual estimated GR occupancy at 10 nM DEX is > 95% (Spencer et al., 1990). Vehicle-treated plates received 4 µL of DMEM containing 1% CS-FBS, 1% pen-strep. Each plate was treated for 10 minutes or 1, 10, 24 or 48 hours before protein collection began. To collect protein samples, culture media was aspirated off and plates were rinsed with 1 mL of 1X ice-cold phosphate-buffered saline (PBS). Subsequently, 200 μL of lysis buffer

(50 mM Tris, 150 mM NaCl, 1% Triton X-100, pH 7.5) containing dissolved protease inhibitors (0.01 mg/mL leupeptin, 0.025 mg/mL aprotinin, 0.010 mg/mL pepstatin A;

Roche, Sigma-Aldrich), 700 units of DNase I (Invitrogen), and 5μM sodium orthovanadate (Na3VO4) (Sigma-Aldrich) was added to each 60 mm plate. Cells were scraped and collected into 1.5 mL Eppendorf tubes, rocked for 10-15 minutes on ice, and subsequently centrifuged for 15 minutes at 17, 530xg and 4°C. After centrifuging,

69 the supernatant containing the protein was collected and the lysate protein concentrations were determined using a Bradford assay (Bradford 1976). After determining concentrations, the protein samples were aliquoted respectively and stored at -20°C for future western blot analyses.

Western Blotting

Collected protein samples were thawed and combined with MilliQ water to prepare western blot samples consisting of 30 μg of protein in a final volume of 30 µL.

Western blot samples were combined with 15 μL of 3x Laemmli buffer (6% SDS, 0.1875

M Tris-HCl pH 6.8, 30% Glycerol, and 0.015% Bromophenol blue) containing 7.5% mercaptoethanol, heated for 4 minutes at 95 °C and vortex mixed. Samples were separated by 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-

PAGE) (10% mini separating gel and 4% mini stacking gel) using the Mini-PROTEAN

Tetra cell system (Bio-Rad) for 2 hours at 100V in running buffer (0.1% SDS, 1.44% glycine, and 0.3% Tris in MilliQ water). Following separation, proteins were subsequently transferred to 0.45 μm nitrocellulose membranes through a semi-dry transfer Trans-Blot Turbo Transfer system at 25 V, 1.5A for 30 minutes (Trans-Blot

Turbo Transfer System, Bio-Rad) in transfer buffer (0.582% Tris, 0.293 % glycine,

0.0107% SDS, and 20% methanol in MilliQ water). Nitrocellulose membranes were then briefly rinsed in 0.1% tris-buffered saline (TBS) containing 0.1% Tween-20 (TBS-T;

2.42% Tris and 8% NaCl in MilliQ water, 0.1% Tween) before blocking in 10% non-fat milk prepared in 0.1% TBS-T for approximately 3 hours. After blocking, membranes were rinsed for 10 minutes with 0.1% TBS-T and incubated at 4°C overnight with the

GPER primary antibody (1:500; Anti-G-protein coupled receptor 30 antibody ab39742;

Abcam) in 0.1 %TBS-T containing 5 % non-fat milk. The next day, nitrocellulose membranes were rinsed with 0.1% TBS-T for 10 minutes and incubated with the horse radish peroxidase (HRP)-conjugated secondary anti-rabbit antibody (1:2500; Cell

Signaling Technology; New England BioLabs, Whitby, Ontario) in 0.1% TBS-T containing 5% milk for 1 hour. Following incubation, membranes were rinsed in 0.1%

TBS-T for 30 minutes and visualized on a ChemiDocTM MP imaging system (Bio-Rad,

Department of Biomedical Sciences) with Luminata Forte Western HRP Substrate

(Millipore). After visualization for 93 kDa GPER bands, nitrocellulose membranes were rinsed in 0.1% TBS-T for 1.5 hours and re-probed for housekeeping protein α-tubulin by incubating with α-tubulin primary antibody (1:500,000; Sigma-Aldrich) in 0.1% TBS-T containing overnight at 4°C. The next day, nitrocellulose membranes were rinsed for 10 minutes and incubated with the anti-mouse secondary antibody (1:2500; Cell Signaling

Technology; New England BioLabs, Whitby, Ontario) and blots were visualized for 50 kDa α-tubulin bands. After visualizing both GPER and α-tubulin loading control blots, semi-quantitative densitometry analyses were performed using Image Lab version 5.0 software (Bio-Rad) to compare the levels of the 93 kDa GPER protein band relative to

α-tubulin loading control between vehicle and 10 nM DEX treated samples isolated after

10-minutes or 1, 10 , 24 or 48 hours of treatment.

Statistical Analyses

After performing inter-blot control corrections, fold changes in GPER expression were compared between the vehicle and DEX treated samples for each 10-minute, 1 hour, 10 hour, 24 hour, and 48 hour time point. Data were tested for normality and homogeneity of variance using Shapiro-Wilk and Bartlett’s tests, respectively. A-priori

71 comparisons within each time point were performed using a two tailed, two-sample unequal variance t-test (heteroscedastic) with Welch’s correction (n=5). Statistical significance was defined by p<0.05. Outliers were removed if values were determined to fall outside of 1.5 times inter-quartile range. All statistical analyses were performed using GraphPad Prism version 7.0 and Microsoft Excel 2016.

Results GPER Protein Expression is Regulated by Dexamethasone Treatment

A representative immunoblot (Figure 3.1a) depicts GPER levels (upper panel) and corresponding α-tubulin levels (lower panel) obtained from control and treated cells.

Both mHippoE-14 and mHippoE-18 cell lines were treated with a single 10 nM dose of the synthetic glucocorticoid receptor agonist, DEX, or a vehicle control for 10 minutes, 1 hour, 10 hours, 24 hours, or 48 hours. When comparing GPER expression fold change in DEX treated samples relative to vehicle controls within each time point, GPER protein expression appeared unaffected at all time points. Densitometric analysis confirmed these observations with the exception of the mHippoE-14 24-hour time point (Figure

3b). A-priori heteroscedastic t-tests revealed a significant downregulation of GPER protein expression after 24-hour treatment with 10 nM DEX in the female derived mHippoE-14s only [t-test; t(7.061)=2.569, p = 0.036805849]. This same downregulation was not observed within any other time points or within the male derived mHippoE-18 cell line (Figure 3c).

Figure 3.1 Effects of dexamethasone on GPER expression in mHippoE cells. (a) Representative western blots depicting GPER protein expression (upper panel) in 10-minute, 1 hour, and 24-hour vehicle and DEX treated mHippoE-14 and mHippoE-18 cell lines, relative to α-tubulin loading control (lower panel). Semi- quantitative densitometric measurements are graphically represented for 93 kDa GPER protein steady state levels in vehicle and DEX treated samples at 10 minutes, 1 hour, and 24 hours in mHippoE-14(b) and mHippoE-18 (c) cells (n=5). A-priori t-testing between control and DEX treated samples within each time point was not statistically significant in mHippoE-14s at 10 minutes [t-test; t(0.3125)=6.471, p = 0.7645] or 1 hour [t-test; t(0.4037)=5.881, p = 0.7007] time points. However, GPER protein steady state levels were significantly different between 24-hour vehicle and DEX treated samples, showing a ~35% reduction in GPER protein levels in the DEX treated samples [t-test; t(7.061)=2.569, p = 0.0368]. A-priori t-testing between control and DEX treated samples within each time point was not statistically significant in mHippoE-18s at 10 minutes [t- test; t(0.602)=8.00, p = 0.5638], 1 hour [t-test; t(0.3892)=6.606, p = 0.7094], or 24 hour [t-test; t(0.1643)=6.963, p = 0.8742] time points. * = p<0.05 determined using a two tailed, two-sample unequal variance t-test (heteroscedastic) to compare differences between vehicle and DEX treated samples. Based on these findings, additional 10 hour and 48 hour time-points were tested for changes in GPER expression following DEX treatment relative to α-tubulin loading control. A representative immunoblot (Figure 3.2a) depicts GPER levels (upper panel) and corresponding α-tubulin levels (lower panel) obtained from control and 10 nM DEX treated cells, respectively. GPER expression fold changes were compared in DEX treated samples relative to vehicle controls within each 10 hour and 48 hour time point.

GPER protein expression was not significantly affected by DEX treatment within either

73 time-point. Densitometric analyses confirmed these observations using a-priori heteroscedastic t-tests in mHippoE-14 10 hour [t-test; t (0.08971)=3.32, p =0.9337 ] or

48 hour [t-test; t(0.2106)=5.354, p =0.8410] time-points and mHippoE-18 10 hour [t-test; t(1.207)=3.948, p =0.2948] or 48 hour [t-test; t(1.027)=2.267, p =0.4015] time points

(Figure 3.2b).

Figure 3.2 Effects of dexamethasone on GPER expression in mHippoE cells. Representative western blots depicting GPER protein expression (upper panel) in 10 hours and 48-hour vehicle and DEX treated mHippoE-14 (a) and mHippoE-18 (b) cell lines, relative to α-tubulin (lower panel) loading control. Semi-quantitative densitometric measurements are graphically represented for 93 kDa GPER protein steady state levels in vehicle and DEX treated samples at 10 and 48 hours (n=4-5). A- priori t-testing between control and DEX treated samples within each time point was not statistically significant in mHippoE-14s (c) at 10 hours [t-test; t (0.08971)=3.32, p =0.9337 ] or 48 hour [t-test; t(0.2106)=5.354, p =0.8410] time points. A-priori t-testing between control and DEX treated samples within each time point was not statistically significant in mHippoE-18s (d) at 10 hour [t-test; t(1.207)=3.948, p =0.2948] or 48 hour [t-test; t(1.027)=2.267, p =0.4015] time points.* = p<0.05 determined using a two tailed, two-sample unequal variance t-test (heteroscedastic) to compare differences between vehicle and DEX treated samples.

Discussion These findings suggest that exposure to a 10 nM dose of DEX results in a significant downregulation of GPER protein steady state levels 24 hours after treatment when compared to vehicle treated controls. This downregulation is moderate, exhibiting an approximate 35 % loss of GPER protein expression in the 24-hour DEX treated mHippoE-14 samples. These findings may have profound implications for stress induced regulation of GPER expression and mediated effects throughout the entire central nervous system and specifically, in the hippocampus.

GPER expression has been shown to be particularly high within extranuclear sites of pyramidal neurons, interneurons, and glial cells of the steroid sensitive hippocampus (Gibbs, 2010; Waters et al., 2015). In the presence of estradiol, GPER has been shown to exert its non-classical effects by translocating from the endoplasmic reticulum and Golgi apparatus to the plasma membrane (Funakoshi et al., 2006). More specifically, 93 kDa plasma membrane GPER expression has been demonstrated within cell bodies, dendritic spines, axons, and nerve terminals within the hippocampus where it has been previously been shown to associate with postsynaptic scaffolding proteins

(Akama et al., 2013; Waters et al., 2015). GPER’s significant density and expression throughout the structure of the neuron suggests that it may play a profound role in regulating estradiol mediated neurotrophic effects and plasticity. For example, GPER activation has been shown to increase dendritic spine density within 40 minutes in CA1 pyramidal neurons of female ovariectomized mice (Gabor et al., 2015). However, it has also been demonstrated that exposure to an acute behavioural stressor may be sufficient to impair the neurotrophic effects of estradiol treatment in the female

75 hippocampus (Frick et al., 2004). As a result, the findings of the current study may provide a potential mechanism to explain why exposure to stressful stimuli may result in reduced estrogen receptor activation due to loss of GPER protein expression.

While the hypothalamic-pituitary-adrenal (HPA) and hypothalamic-pituitary- gonadal (HPG) axes are well established to reciprocally modulate and regulate one another, a majority of studies have focussed only on the behavioural effects and interactions between stress and sex steroids. GPER expression has been associated with the serotonin (5-HT)1A receptor expression and may therefore play a role in estradiol’s desensitization of this receptor (McAllister et al., 2012). This concurrent expression may be involved in GPER’s role in regulating anxiety and depressive like behaviours (McAllister et al., 2012). For example, females with major depressive disorder have been demonstrated to have significantly reduced levels of plasma estradiol and therefore may have reduced estrogen receptor expression and/or activation within their central nervous system (Young et al., 2000). However, most studies have only considered the effects of stress hormone exposure on expression and activation of classical estrogen receptors. For example, amygdala ERα expression is significantly reduced in patients diagnosed with major mental illnesses including major depressive disorder (Perlman et al., 2004). However, ERβ expression within the hypothalamus of female rats is upregulated in response to glucocorticoid treatment

(Suzuki & Handa, 2004). It is therefore evident that classical estrogen receptor expression is sensitive to and may be regulated by the physiological stress response.

However, stress hormone regulation of GPER expression within the hippocampus has been unexplored.

The downregulation of GPER protein expression exhibited 24 hours after DEX treatment was evident only within the female derived mHippoE-14s while the male derived mHippoE-18s remained relatively unaffected by DEX treatment. This observed sex difference is consistent with previous findings demonstrating that the female hippocampus may be particularly vulnerable to the detrimental effects of stress. For example, male rats exposed to an acute tail shock stressor exhibit significant improvements in their hippocampus-dependent associative learning in comparison to their control treated counterparts and this is mimicked through increased hippocampal dendritic spine density ( Shors et al.,2001; Wood et al., 2001; Wood & Shors, 1998).

However, females exposed to the same behavioural stressors exhibit significant cognitive impairments that are mimicked through truncation and loss of dendritic spine density ( Shors et al., 2001; Wood et al., 2001; Wood & Shors, 1998). This sex specific impairment could be explained by glucocorticoid induced loss of GPER expression and subsequent loss of GPER mediated functional effects in the female hippocampus.

This downregulation of GPER protein expression may also have serious implications for the aging brain. With age, there is a consistent and steady rise in plasma cortisol levels that is exhibited in both males and females (Van Cauter et al.,

1996; Lupien et al., 1998). This cumulative exposure to glucocorticoids has been correlated with degree and severity of age-related hippocampal atrophy (Lupien et al.,

1998). The natural loss of circulating ovarian hormones that females experience may predispose all aging females to a higher degree of hippocampal atrophy ( Tang et al.,

1996). However, a sustained exposure to elevated circulating glucocorticoids may help explain why some females are more particularly susceptible to hippocampal atrophy

77 and the onset of sexually differentiated neurodegenerative conditions, such as

Alzheimer’s disease (Tang et al., 1996; Xu et al., 2015; Ycaza-Herrera & Mather, 2015).

Therefore, this glucocorticoid induced loss of GPER protein expression may help explain the vulnerability of the female hippocampus to the detrimental effects of stress.

Conclusion In conclusion, these findings demonstrate that glucocorticoid receptor activation downregulates GPER protein expression in female derived mHippoE-14 hippocampal neurons. These findings may elude to mechanisms that mediate the underlying sex differences observed in hippocampal plasticity and cognition in response to stressful stimuli. Lower of functional GPER expression may account for reduced sensitivity to the neurotrophic and neuroprotective effects of estradiol in the female hippocampus. As a result, downregulated GPER expression may help explain this female sex-specific vulnerability to the detrimental effects of stress.

CHAPTER 4: GLUCOCORTICOID REGULATION OF GPER MEDIATED C-JUN N TERMINAL KINASE (JNK) PHOSPHORYLATION IN IMMORTALIZED HIPPOCAMPAL NEURONS

Introduction The hippocampal G-protein coupled estrogen receptor (GPER) is distinct from classical estrogen receptor (ER)α and ERβ in that it signals through a diverging pathway known as the c-jun N terminal kinase (JNK) cascade (Kim et al.,2016). This non-classical mechanism may indirectly regulate gene expression and activation of other intracellular events that may modulate hippocampal structure, learning, and memory (Gabor et al.,2015; Kim et al., 2016; Lymer et al., 2017). The GPER specific agonist, G-1, has been previously used to demonstrate GPER mediated activation of 54 and 46 kDa isoforms of JNK through phosphorylation of JNK’s threonine 183 and tyrosine 185 residues (Kim et al., 2016; Qi et al., 2005; Wang et al., 2005). This non- classical JNK activation occurs on a rapid timescale within minutes to hours (Gabor et al., 2015). Phosphorylation of JNK has been associated with regulation of hippocampus-mediated associative learning and memory in baseline and stressful conditions (Antoniou & Borsello, 2012). In fact, the memory enhancing effects of GPER activation have been shown to be specific to phospho (p)-JNK and mimic the beneficial effects on memory consolidation that is seen with estradiol treatment (Kim, 2014; Kim et al., 2016). Within 40 minutes of treatment, GPER activation has been shown to significantly increase dendritic spine density within CA1 pyramidal neurons of ovariectomized female mice (Gabor et al., 2015) although, these regulatory mechanisms may differ between short and long term memory consolidation (Bevilaqua et al., 2003).

The female hippocampus is particularly vulnerable to the detrimental effects of stressful stimuli. Previous studies indicate that acute behavioural stressors significantly impair associative learning in females when compared to their control treated

80 counterparts or stress-exposed males who actually exhibit an improvement in performance ( Shors et al.,2001; Wood & Shors, 1998). These findings suggest that there may be an interaction between the physiological rise in glucocorticoids and the neurotrophic and memory enhancing effects of estradiol in the female hippocampus. In fact, Frick et al. (2004) demonstrated that injections with estradiol benzoate resulted in significant increases in spine synapse density within the hippocampus of ovariectomized rats. However, exposing ovariectomized rats treated with estradiol benzoate to the Morris water maze prevented the beneficial and neurotrophic effects of estradiol on spine synapse density (Frick et al., 2004). This would suggest exposure to the Morris water maze may activate the physiological stress response and rise in glucocorticoids that might subsequently impair or prevent functional estrogen receptor activation and/or signalling in the female hippocampus.

GPER is highly expressed within axons, nerve terminals, cell bodies, and dendritic spines and has been largely associated with mediating a rapid increase in dendritic spine density (Akama et al., 2013; Gabor et al., 2015; Waters et al., 2015). As stress has been shown to have such a serious and rapid detrimental effect on neuronal structure in the female hippocampus, it is plausible that stress-induced inhibition of estrogen signalling is mediated through GPER. To investigate the effects of glucocorticoid exposure on GPER activation and signalling, mHippoE-14 and mHippoE-

18 cell lines were both characterized for a functional response to G-1 through JNK phosphorylation. As GPER protein expression was previously shown to be downregulated 24 hours after 10 nM dexamethasone (DEX) treatment, the next objective was

81 to determine whether this downregulation was also mimicked by a loss of functional

GPER mediated activation of JNK.

Experimental Procedures Chemical and Stock Solutions

Stock solutions of G-1 (200 μM) were prepared in 20% (2-Hydroxypropyl)-β- cyclodextrin (cyclodextrin; Sigma-Aldrich) dissolved in sterile filtered MilliQ water.

Working stock solutions of 10 μM G-1 were preparing by diluting 200 μM stock of G-1 into Dulbecco’s modified Eagle medium (DMEM) containing 1% charcoal-stripped fetal bovine serum (CS-FBS; Invitrogen) and 1% penicillin-streptomycin (pen-strep;100 units penicillin + 100 μg streptomycin; Invitrogen). Stock solutions of DEX (500 μM) were prepared in CS-FBS and working stocks were prepared to a final concentration of 10

μM in DMEM containing 1% CS-FBS and pen-strep.

Cell Culture Conditions

mHippoE-14 and mHippoE-18 immortalized murine hippocampal neurons

(#CLU198, #CLU199, Cedarlane) were cultured and incubated in DMEM that contained

10% FBS and 1% pen-strep at 37°C and 5% CO2 and passaged with 2 mL of

0.25%/0.53 mM trypsin/EDTA solution (1X Hank’s balanced salt solution (HBSS), 1.2 mg/mL NaHCO3, 0.46 mg/mL EDTA, 2.5 mg/mL trypsin, 0.2μL/mL phenol red; Sigma-

Aldrich, Oakville, Ontario, Canada) when culture flasks reached ~80% confluency. Lifted cells were resuspended with 10 mL of culture media and 1 mL of the suspension was plated on 60 mm culture plates (Corning-Fisher Scientific) that were brought to a total volume of 4 mL with 10% FBS, 1% pen-strep DMEM culture media. Once plates

82 reached ~75% confluency, culture media was aspirated and replaced with 4 mL of

DMEM containing 1% CS-FBS, 1% pen-strep for 16 hours before experiments began

G-1 Treatments

As a rapidly activated non-classical estrogen receptor, GPER mediated effects occur within minutes to hours (Gabor et al., 2015; Phan et al., 2011; Phan et al., 2015).

To establish functional activation of GPER, mHippoE-14 and mHippoE-18 culture plates were treated with a cyclodextrin vehicle control or a 10 nM dose of G-1 for 10 minutes, 1 hour, or 4 hours. The 10 nM dose of G-1 has been previously demonstrated to sufficiently activate GPER in the mHippoE-14 cell lines (Gingerich et al., 2010).

24 hour Dexamethasone Pre-treatments

To determine glucocorticoid regulated functional activation of GPER, only mHippoE-14 culture plates were used. mHippoE-14s were treated with a vehicle control or 10 nM dose of DEX for 24 hours (pre-treatment). At 24 hours, the same mHippoE-14 culture plates were subsequently treated with a cyclodextrin vehicle control or a 10 nM dose of G-1 for 10 minutes, 1 hour, or 4 hours.

Protein Collection

Protein samples were collected as previously described. Briefly, culture media was removed and culture plates were rinsed with 1 mL of 1X ice-cold phosphate- buffered saline (PBS). Subsequently, 200 μL of Triton-X lysis buffer (50 mM Tris, 150 mM NaCl, 1% Triton C-100, pH 7.5) containing dissolved protease inhibitors (0.01 mg/mL leupeptin, 0.025 mg/mL aprotinin, 0.010 mg/mL pepstatin A; Roche, Sigma-

Aldrich), 700 units of DNase I (Invitrogen), and 5 μM sodium orthovanadate (Na3VO4;

Sigma-Aldrich) was added to each 60 mm plate. Cells were collected into 1.5 mL

Eppendorf tubes, rocked for 10-15 minutes on ice, and subsequently centrifuged for 15 minutes at 17, 530 g and 4°C. After centrifuging, the supernatant containing protein was collected, quantified using a Bradford assay (Bradford 1976), and aliquoted respectively for storage at -20°C.

Western Blotting

Collected protein samples were thawed and prepared with MilliQ water to total 30

μg of protein in a final volume of 30 μL. Prepared samples were combined with 15 μL of

3x Laemmli buffer (6% SDS, 0.1875 M Tris-HCl pH 6.8, 30% Glycerol, and 0.015%

Bromophenol blue) containing 7.5% mercaptoethanol, heated for 4 minutes at 95 °C and vortex mixed. separated through 10% sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) (10% mini separating gel and 4% mini stacking gel) using the Mini-PROTEAN Tetra cell system (Bio-Rad) for 2 hours at 100V in running buffer

(0.1% SDS, 1.44% glycine, and 0.3% Tris in MilliQ water). Protein samples were subsequently transferred to 0.45 μm nitrocellulose membranes through a semi-dry transfer Trans-Blot Turbo Transfer system at 25 V, 1.5A for 30 minutes (Trans-Blot

Turbo Transfer System, Bio-Rad) in transfer buffer (0.582% Tris, 0.293% glycine,

0.0107% SDS, and 20% methanol in MilliQ water) and subsequently blocked in 10% bovine serum albumin (BSA) prepared in tris-buffered saline (TBS) containing 0.1% tween-20 (TBS-T; TBS-T; 2.42% Tris and 8% NaCl in MilliQ water, 0.1% Tween) for approximately 1.5 hours. After blocking, membranes were rinsed with 0.1% TBS-T and incubated at 4°C overnight with antibodies to detect total JNK (1:500; JNK antibody D-

2:sc-7345, Santa Cruz Biotechnology) or p-JNK (1:2000; phosphor-SAPK/JNK

[Thr183/Tyr185] antibody #9251, Cell Signaling Technology; New England BioLabs,

Whitby, Ontario). The next day, nitrocellulose membranes were rinsed with 0.1% TBS-T and incubated with the horse radish peroxidase-conjugated secondary anti-rabbit or anti-mouse antibody (1:2500; Cell Signaling Technology; New England BioLabs,

Whitby, Ontario) for 1 hour. Following incubation, membranes were rinsed in 0.1% TBS-

T and visualized using the ChemiDocTM MP imaging system (Bio-Rad, Department of

Biomedical Sciences) with Luminata Forte Western HRP Substrate (Millipore). After visualization, nitrocellulose membranes were rinsed in 0.1% TBS-T for 1.5 hours and re- probed for housekeeping protein β-actin by incubating with the β-actin primary antibody

(1:5000; Santa Cruz Biotechnology Inc., Dallas, Texas, USA) overnight at 4°C. The next day, nitrocellulose membranes were rinsed and incubated with the anti-mouse secondary antibody (1:2500; Cell Signaling Technology; New England BioLabs, Whitby,

Ontario, Canada) and blots were visualized as described. After capturing images of p-

JNK, JNK, and β-actin blots, semi-quantitative densitometric analyses were performed using Image Lab version 5.0 software (Bio-Rad) to compare 54 and 46 kDa p-JNK expression relative to total JNK expression, normalized to β-actin loading controls.

Statistical Analyses: Functional Activation of GPER Using G-1 Treatments

After performing inter-blot control corrections, fold changes in p-JNK relative to total JNK expression were compared between the cyclodextrin vehicle and G-1 treated

10 minute, 1 hour, and 4 hour samples. A two-way analysis of variance (ANOVA) was used to assess overall effects of treatment and time. Linear contrast tests were subsequently performed to assess for overall effects of G-1 treatment in comparison to vehicle treated controls (n=6). Statistical significance was defined by p<0.05. Outliers

85 were removed if values were determined to fall outside of 1.5 times inter-quartile range.

All statistical analyses were performed using GraphPad Prism version 7.0 and Microsoft

Excel 2016.

Statistical Analyses: Glucocorticoid Regulation of GPER Functional Activation

Inter-blot corrections were performed for vehicle treated samples. Subsequently, p-JNK expression was quantified as a fold change value relative to total JNK expression values for both 54 and 46 kDa isoforms. Overall effects of DEX pre-treatment between control and G-1 treated samples were analyzed using a two-way ANOVA (n=7) followed by a Tukey’s post hoc-test for multiple comparisons. Data were tested for normality and homogeneity of variance using Shapiro-Wilk and Bartlett’s tests, respectively. Statistical significance was defined by p<0.05 and all statistical analyzes were performed using

Microsoft Excel 2016 and GraphPad Prism version 7.0.

Results mHippoE-14 Cell Lines Express a Functional GPER

Both mHippoE-14 and mHippoE-18 cell lines were treated with a single 10 nM dose of the GPER specific agonist, G-1 (Gingerich et al., 2010). Representative immunoblots indicate respective JNK phosphorylation in vehicle and G-1 treated mHippoE-14s (Figure 4.1a) and mHippoE-18s (Figure 4.1b) at 10 minute, 1 hour, and 4 hour time points. Western blot visualization revealed steady increases in JNK phosphorylation within mHippoE-14 female derived cell lines while JNK phosphorylation appeared much more variable in G-1 treated male derived mHippoE-18s. Semi- quantitative densitometric analyses were used to calculate levels of p-JNK expression

86 relative to total JNK expression as a fold change value that was normalized to β-actin expression for 54 and 46 kDa isoforms in both mHippoE-14 (Figure 4.1a) and mHippoE-

18 (Figure 4.1b) cell lines. While correcting for inhomogeneity of variance, linear contrast tests were used. The p-JNK fold change values in the 10-minute, 1 hour, and 4 hour treated samples were significantly increased in comparison to the cyclodextrin vehicle samples in both 54 (F=16.670, p = 0.002) and 46 (F=12.135, p = 0.003) kDa isoforms in the female derived mHippoE-14s (Figure 4.1 a). The male derived mHippoE-

18 cell line exhibited similar trends in p-JNK activation at 10 minutes, 1 hour, and 4 hour time points, however, no statistically significant differences in the 46 kDa (F=4.659, p =

0.053) or 54 kDa (F=1.421, p = 0.0256) isoforms of JNK were identified (Figure 4.1 b).

Figure 4.1 Effects of GPER agonist G1 on JNK signaling in mHippoE cells. Representative western blots depicting JNK phosphorylation (upper panel), total JNK (middle panel) and actin (bottom panel) in vehicle and 10-minute, 1 hour, and 4 hour G-1 treated samples of a) mHippoE-14 and b) mHippoE-18 cell lines. Semi- quantitative densitometric measurements are graphically represented for 54 and 46 kDa isoforms of p-JNK relative to total JNK and β-actin loading controls. Using linear contrast, p-JNK levels are significantly increased in mHippoE-14 10-minute, 1 hour, and 4-hour G-1 treated samples when compared to cyclodextrin vehicle control for both 54 (F=16.670, p = 0.002) and 46 (F=12.135, p = 0.003) kDa isoforms of JNK (**=p<0.01). Expression of p-JNK is not significantly increased in mHippoE-18 10-minute, 1 hour, and 4-hour G-1 treated samples for both 54 (F=1.421, p = 0.256) and 46 (F=4.659, p = 0.053) kDa isoforms when compared to cyclodextrin vehicle controls. The 46 kDa isoform of p-JNK is trending towards significance in mHippoE-18s (t = p < 0.1).

Glucocorticoids Regulate G-1 Induced Functional Activation of GPER

After characterizing a functional response to G-1 in mHippoE-14 female derived hippocampal cell lines, effects of glucocorticoid treatment on GPER functional activation were investigated. mHippoE-14s were exposed to vehicle or DEX pre-treatment 24 hours prior to treatment with a cyclodextrin vehicle control or G-1 treatment for 10 minutes, 1 hour, or 4 hours. Representative immunoblots for vehicle pre-treated mHippoE-14s indicate a steady increase in p-JNK expression in G-1 treated 10 minute,

1 hour, and 4 hour samples when compared to the cyclodextrin vehicle control (Figure

4.2 upper panel). Representative immunoblots for DEX pre-treated mHippoE-14s show a lack of JNK phosphorylation in G-1 treated 10 minute, 1 hour, and 4 hour samples when compared to the cyclodextrin vehicle control (Figure 4.2 upper panel). After performing interblot corrections, semi-quantitative analyses revealed that fold change levels of p-JNK relative to total JNK were significantly decreased in the 24 hour DEX pre-treated samples in comparison to the vehicle pretreated samples after 10 minutes, 1 hour, and 4 hour treatments with G-1 (Figure 4.2 lower panel). For the 54 kDa isoform, a significant effect of DEX pre-treatment [F(1,48)=23.83, p = < 0.0001] and a significant interaction effect [F(3,48)=3.187, p=0.0319] were detected. Similar differences were also observed with the 46 kDa isoform including a significant effect of DEX pre- treatment [F(1,47)=31.66, p = < 0.0001] and a significant interaction effect [F(3,47)=

3.659, p = 0.0189] (Figure 4.2). Tukey post hoc tests revealed significantly reduced p-

JNK in 24 hour DEX pre-treated samples at 1 hour (p = 0.0035) and 4 hour (p=0.0337), but not 10 minutes (p=0.3029) following G-1 treatment when compared to vehicle pre- treated samples in the 54 kDa isoform of JNK. However, Tukey post hoc tests revealed

88 that the 46 kDa isoform of p-JNK was significantly reduced in 24-hour DEX pre-treated samples following 10 minute (p=0.0319), 1 hour (p = 0.0106), and 4 hour (p = 0.0022) treatments with G-1 when compared to the vehicle pre-treated samples.

Figure 4.2 Representative western blots depicting JNK phosphorylation in 24- hour vehicle and DEX pre-treated mHippoE-14 samples. After 24-hour pre- treatment, cells were treated with a vehicle control or G-1 for 10 minutes, 1 hour, or 24 hours. Semi-quantitative densitometric measurements are graphically represented for fold changes of 54 and 46 kDa isoforms of p-JNK relative to total JNK and β-actin loading controls. Using a 2-way ANOVA, p-JNK levels were significantly decreased in the 24 hour DEX pre-treated mHippoE-14 10-minute, 1 hour, and 4-hour G-1 treated samples when compared to 24 hour vehicle pre-treated samples for both 54 kDa isoform [F(1,48)=23.83, p = < 0.0001] and 46 kDa isoform [F(1,47)=31.66, p = ***< 0.0001]. A significant interaction effect was observed for the 54 kDa isoform [F(3,48)=3.187, p=0.0319] and 46 kDa isoform [F(3,47)= 3.659, p = 0.0189]. Tukey post-hoc tests revealed significant downregulation of 54 kDa p-JNK at 1 hr G-1 (p = 0.0035) and 4 hour G-1 (p = 0.0337) in DEX pre-treated samples when compared to vehicle pre-treated samples. Tukey post-hoc tests revealed significant downregulation at 10 minute G-1 (p = 0.0319), 1 hour G-1 (p = 0.0106), and 4 hour G-1 (p=0.0022) DEX pre-treated samples when compared to vehicle pre-treated samples.

Discussion These findings indicate that G-1 induced activation of GPER, as measured by

JNK phosphorylation, is significantly inhibited by glucocorticoid pre-treatment when compared to vehicle pre-treated controls in the female derived mHippoE-14s. This overall effect of DEX pre-treatment is significant for both 54 and 46 kDa isoforms of

JNK. The glucocorticoid induced loss of GPER functional activation may help explain the mechanism underlying the stress induced loss of functional estradiol signalling and neurotrophic effects in the female hippocampus.

As a non-classical estrogen receptor, GPER activation results in rapid phosphorylation of the JNK intracellular signalling cascade (Kim, Szinte, Boulware, &

Frick, 2016). As a member of the mitogen-activated protein kinase family, JNK has been shown to play an important role in the modulation of synaptic plasticity, neuronal differentiation, and regulation of apoptosis (Farías et al., 2009; Seo et al., 2012).

Inhibition of hippocampal JNK has been shown to result in significantly impaired induction of long-term potential within murine hippocampal CA1-CA3 synapses (Seo et al., 2012). Additionally, JNK activation has been shown to enhance recruitment of clusters of post-synaptic density-95 scaffolding within mammalian hippocampal neurons

(Farías et al., 2009).

Many protein kinase pathways such as JNK are sensitive to stressful stimuli and glucocorticoid treatment. For example, glucocorticoid exposure has been shown to downregulate JNK signalling in human endothelial cells (González et al., 1999).

Hippocampal extracellular signal-related kinase (ERK) phosphorylation is well established to increase in response to stressful stimuli, while JNK regulation has been shown to be more moderately regulated within the male rat hippocampus (Cai et al.,

2007; Shen et al., 2004). However, other studies have demonstrated a rapid activation of JNK following behavioural stressors or glucocorticoid treatment in hippocampal neurons (Qi et al., 2005). While JNK’s molecular role in response to glucocorticoid receptor activation remains unclear, JNK has been shown to regulate hippocampus- mediated memory formation under baseline and stressful conditions (Antoniou &

Borsello, 2012).

As a result, GPER mediated JNK activation may play an important role in the underlying sexually differentiated response to stressful stimuli. Within the hippocampus, there are evident sex differences in behavioural and morphological responses to stress.

Exposure to a 30 minute intermittent tail shock stressor has been shown to have sexually differentiated effects on behaviour and apical spine density in the hippocampus

(Shors et al., 2001). For example, stressed males exhibit improved cognition with a 30% increase in apical spine density, while females exhibit cognitive impairments and up to a

20% reduction in dendritic spine density 24 hours after exposure to the behavioural stressor (Shors et al., 2001). Moreover, exposure to a behavioural stressor has been shown to sufficiently impair the rapid neurotrophic effects of estradiol treatment in the female hippocampus (Frick et al., 2004). However, GPER expression is similar in CA1,

CA3, and dentate gyrus dendrites, spines, and axons in both male and female mice

(Waters et al., 2015). Therefore, the current findings demonstrating the stress induced downregulation of GPER expression and functional signalling in female murine hippocampal neurons may help explain the underlying sex differences in hippocampal vulnerability to stress. Using the female derived mHippoE-14 neuronal cell lines, we have demonstrated that GPER mediated phosphorylation of JNK is significantly

91 impaired following exposure to glucocorticoids.

Loss of functional GPER activation and JNK signalling may significantly impair estradiol mediated neurotrophic and neuroprotective effects within the female hippocampus. This may have profound implications for the aging brain and the onset of many sexually differentiated neurodegenerative conditions. The natural loss of estrogen synthesis that occurs during menopause may predispose the female brain to an age- related loss of estradiol’s neurotrophic and neuroprotective effects. Given these current findings, women who experience sustained elevations in glucocorticoids may experience significantly reduced GPER expression and functional activation in their hippocampi. In comparison to their male counterparts, women exposed to stressful stimuli may be more vulnerable to hippocampal atrophy and the onset of sexually differentiated neurodegenerative conditions, such as Alzheimer’s disease (Pike et al.,

2009).

Conclusion In conclusion, these findings indicate that exposure to glucocorticoids significantly impair functional activation of hippocampal GPER in mHippoE-14 female derived hippocampal neurons. These results may provide an explanation for the mechanisms underlying sex differences in hippocampal morphology and cognition in response to stress stimuli. Glucocorticoid induced downregulation of GPER mediated

JNK signalling may explain the stress induced loss of estradiol’s neurotrophic and neuroprotective effects within the female hippocampus and may help explain why the female hippocampus is more vulnerable to the detrimental effects of stress.

CHAPTER 5: REGULATORY EFFECTS OF GLUCOCORTICOID TREATMENT ON GPER FUNCTIONAL ACTIVATION IN OVARIECTOMIZED FEMALE CD1 MICE

Introduction The behavioural and morphological responses to stressful stimuli are well established to be sexually differentiated in both male and female rodent models (Shors et al., 2001; Wood & Shors, 1998). These findings indicate a role for underlying interactions between stress and gonadal hormones. Specifically, these findings elude to a female sex-specific vulnerability to the detrimental effects of stress. However, the underlying cellular and molecular mechanisms mediating these sexually differentiated responses has yet to be fully understood. The in vitro findings presented in chapters 1-4 of this thesis indicate an interaction between glucocorticoid exposure and GPER expression and functional activation in female derived immortalized mHippoE-14 neurons. The downregulation of GPER protein expression coupled with the glucocorticoid induced loss of G1-mediated JNK activation may provide a potential explanation for why the female hippocampus is particularly vulnerable to the detrimental effects of stress.

These findings may have profound implications for understanding glucocorticoid regulated mechanisms in the female hippocampus. In vitro models provide us with the ability to easily control and manipulate physiological systems that allow us to more clearly identify direct cellular mechanisms. However, the application and generalizability of these findings is significantly limited as the physiological variables, feedback mechanisms, and systemic responses present in an in vivo model cannot be accounted for. As a result, the next objective of this work was to determine whether these in vitro mHippoE-14 findings are similar to what occurs in an in vivo mouse model. Previous studies have investigated the effects of stress and estradiol interactions within the hippocampus through use of behavioural stressors such as acute tail shocks, the Morris

94 water maze, or restraint stress (Anderson et al.2013; Frick et al., 2004; Frye & Orecki,

2002; Wood & Shors, 1998). However, it remains difficult to determine the underlying mechanisms of action as the physiological stress results in activation of multiple stress receptors including, glucocorticoid, mineralocorticoid, and corticotrophin-releasing hormone receptors. Although, in vivo treatment with dexamethasone (DEX) would solely result in activation of glucocorticoid receptors (GRs) and elucidate a potential mechanism, this may poorly represent the effects of stress exposure. Previous studies have investigated the effects of subcutaneous corticosterone (CORT) treatment on stress response physiology (Choleris et al., 2013). As CORT results in both mineralocorticoid receptor (MR) and GR activation, this may provide the framework to properly investigate a more refined mechanism within the physiological stress response.

While many studies have determined the in vivo morphological and behavioural effects of G-1 treatment, few have considered interactions between GPER activation and the physiological stress response. Previous findings indicate that GPER activation may result in sexually differentiated anxiolytic and anxiogenic effects in male and female rodents (Hart et al., 2014; Kastenberger et al., 2012). However, the reciprocal stress hormone regulation of GPER expression or activation has yet to be investigated in vivo.

While previous literature indicates that behavioural stressors sufficiently impair estradiol mediated neurotrophic and neuroprotective effects in the female hippocampus (Frick et al., 2004), GPER has not yet been investigated as a potential mediator of these effects.

The current in vitro findings suggest that glucocorticoid exposure may downregulate

GPER protein expression and functional activation within the female hippocampus and this may help explain why the female hippocampus is particularly vulnerable to the

95 detrimental effects of stress. To best understand the generalizability of these findings, the last objective was to investigate these mechanisms using an in vivo model.

Experimental Procedures Animals

Naive (2-3 month) female CD1 mice (Mus musculus) were purchased from

Charles River Laboratories, St. Constance, QC. The animal utilization protocols for procedures involving the use of the mice were consistent with the guidelines of the

Canadian Council of Animal Care and were approved by the University of Guelph’s

Institutional Animal Care and Use Committee. After arrival to the University of Guelph’s

Central Animal Facility, mice were triple-housed in an enriched polyethylene cage (16 cm x 12 cm x 26 cm) containing a paper cup, paper pad, and cotton nesting. Mice were maintained on a reversed 12-hour light/dark cycle (lights off at 8:00 am) at 21 ± 1° with ad libitum access to Teklad global 14% protein rodent diet (Envigo) and tap water. After arrival, mice were given 1 week to habituate to their environment.

Ovariectomy Surgeries

All mice received a subcutaneous injection of 50 mg/kg of the veterinary anti- inflammatory drug, carprofen (Pfizer Canada Inc, Kirkland, Qc, Canada) at least one hour before beginning surgeries. All mice were placed in an induction chamber and anesthetized with 5% isoflurane gas (Benson Medical Industries, Markham, ON).

Subsequently, mice were placed in a nose cone administering 2% isoflurane gas and the lumbar region of their back was cleaned and shaved before topical administration of

0.02 mL of 0.67% lidocaine (Alveda Pharmaceuticals, Toronto, ON, Canada) and 0.17%

96 bupivacaine (Hospira Inc., Montreal, Qc, Canada) before making a ~1 cm subcutaneous incision to reduce post-operative pain. After the subcutaneous incision was made, mice received a 0.5 mL sub-cutaneous injection of warm saline to maintain hydration. After making a midline dorsal incision, bilateral incisions were made through the underlying connective tissue and muscle above each ovary using the blunt edges of surgical scissors. The fat pads containing the ovaries were drawn out from the abdominal cavity, the oviduct was clamped, and the fat pad containing the ovary was removed by cutting above the clamp. After sufficient clotting, the fallopian tubes were tucked back into the abdominal cavity and the subcutaneous incision in the skin was sealed with wound clips

(9mm wound clips, MikRon Precision Inc., Gardena, CA). Following surgery, the mice were single housed and monitored on a daily basis to avoid re-opening of the surgical incision for at least one week following surgery.

Chemical and Stock Solutions

Stock solutions of CORT (C2505; Sigma-Aldrich, Oakville, ON) were prepared by dissolving CORT in a 0.1% ethanol and 0.9% physiological saline solution. Stock solutions of G-1 (Tocris Biosciences, UK) were prepared in sesame oil.

Treatment Groups

Mice were randomly allocated to four separate treatment groups (n=7 per group): vehicle-injection control, G-1 without CORT injection, G-1 with CORT injection, or

CORT without G-1 injection. After weighing each mouse to determine appropriate volumes for injection, CORT and G-1 were retracted in the same syringe to reduce the number of subcutaneous injections in each mouse. Treatments suspended in sesame oil were retracted first followed by treatments suspended in saline were retracted. Mice

97 were subcutaneously injected with 5 mg/kg of CORT (Choleris et al., 2013) with or without 6 μg/kg of G-1 (Gabor et al., 2015) and the vehicle control animals received 5 mg/kg saline and 6 μg/kg of sesame oil. The 5 mg/kg dose of CORT was chosen based on previous studies indicating a dose dependent increase in plasma CORT that subsequently decreased 40 minutes following injection (Choleris et al., 2013). The 6

μg/kg dose of G-1 was chosen according to a previous study indicating G-1 mediated increases in hippocampal dendritic spine density through subcutaneous injection (Gabor et al., 2015). Injection sites were sealed using liquid bandage to prevent subcutaneous leakage. The results of a pilot study conducted within our lab indicate increased JNK phosphorylation within 15 minutes of G-1 treatment. As a result, mice for the current study were sacrificed by cervical dislocation 15 minutes after injection and the hippocampus was bilaterally extracted from each animal’s brain within 2-3 minutes.

Hippocampi were flash frozen in liquid nitrogen and transferred to -80 °Celsius for storage.

Protein Preparation and Western Blotting

Following removal from -80°Celsius, tissue samples were weighed and homogenized in lysis buffer (50mM Tris,150mM NaCl,1% Triton X-100, pH 7.5) containing dissolved protease inhibitors tablets (Sigma-Aldrich, Oakville, ON), 5 μM sodium orthovanadate, and 700U/ml DNase. Low frequency sonication was used briefly and each sample was rocked on ice for 15-20 minutes. Samples were subsequently centrifuged twice at 17, 530xg for 15 minutes at 4°Celsius. After centrifugation, each supernatant was transferred into an Eppendorf tube and the remaining cell waste pellets were discarded. Using a Bradford assay (Bradford, 1976), protein concentrations were

98 determined and samples were aliquoted respectively for storage at -20° Celsius.

Hippocampal lysates were combined with MilliQ water to contain 30 ug of total protein in a final volume of 30 μL Samples were mixed with 15 μL of 3x Laemmli buffer

(6% SDS, 0.1875 M Tris-HCl pH 6.8, 30% Glycerol, and 0.015% Bromophenol blue) containing 7.5% mercaptoethanol, heated for 4 minutes at 95 °C and vortex mixed.

Samples were subsequently added into wells of western blot gels (10% mini separating gel and 4% mini stacking gel). Samples were subjected to sodium dodecyl sulfate polyacrylamide gel electrophoresis (SDS-PAGE) using the Mini-PROTEAN Tetra cell system (Bio-Rad) for 2 hours at 100 V in running buffer (0.1% SDS, 1.44% glycine, and

0.3% Tris in MilliQ water). Proteins were subsequently transferred to nitrocellulose membranes through a semi-dry transfer Trans-Blot Turbo Transfer system at 25 V, 1.5

A, for 30 minutes (Trans-Blot Turbo Transfer System, Bio-Rad) in transfer buffer

(0.582% Tris, 0.293 % glycine, 0.0107% SDS, and 20% methanol in MilliQ water).Blots were subsequently rinsed in tris buffered saline (TBS) containing 0.1% tween-20 (TBS-

T; 2.42% Tris and 8% NaCl in MilliQ water, 0.1% Tween), and blocked at room temperature in 5% bovine serum albumin (BSA) dissolved in 0.1% TBS-T for 1.5 hours on a rocker. After blocking, membranes were rinsed with TBS-T and incubated at 4°C overnight with JNK (1:500; JNK antibody D-2:sc-7345, Santa Cruz Biotechnology) or p-

JNK (1:2000; phosphor-SAPK/JNK [Thr183/Tyr185] antibody #9251, Cell Signaling

Technology; New England BioLabs, Whitby, Ontario) primary antibodies. The next day, nitrocellulose membranes were rinsed with 0.1% TBS-T and incubated on a rocker with their respective goat anti-rabbit or goat anti-mouse secondary antibodies (1:2500; Cell

Signaling Technology; New England BioLabs, Whitby, Ontario) in 0.1 % TBS-T

99 containing 5% BSA for 1 hour at room temperature. After 4 rinses with 0.1 % TBS-T for a total of 30 minutes, western blots were visualized on a ChemiDoc TM MP imaging system (Bio-Rad, Department of Biomedical Sciences) with Luminata Forte Western

HRP Substrate (Millipore, Oakville, ON). After visualization for p-JNK or JNK densitometry, nitrocellulose membranes were rinsed in TBS-T for 1.5 hours and re- probed for β-Actin by incubating with the β-Actin primary antibody (1:5000; Cell

Signaling Technology; New England BioLabs, Whitby, Ontario) overnight at 4°C. The next day, nitrocellulose membranes were rinsed again with 0.1% TBS-T and incubated with the goat anti-rabbit secondary antibody (1:2500; Cell Signaling Technology; New

England BioLabs, Whitby, Ontario) and blots were visualized for β-Actin bands. Semi- quantitative densitometry analyses were performed using Image Lab version 5.0 software (Bio-Rad) to compare p-JNK and JNK (54 and 46 kDa) expression normalized to a house-keeping (β-Actin) protein from the same blot. Analyses were used to compare p-JNK fold changes between treatment groups.

Statistical Analyses

Following semi-quantitative densitometric analyses and inter-blot control corrections, fold changes in p-JNK relative to JNK levels for the 54 and 46 kDa isoforms were compared between the vehicle, G-1, CORT, and G-1 & CORT groups (n=5-6 per group). Natural log transformations were used to account for inhomogeneity of variance between groups. Subsequently, a one-way analysis of variance (ANOVA) was used to assess for overall effects of treatment within each JNK isoform followed by Tukey’s post hoc-test for multiple comparisons. Statistical significance was defined by p < 0.05 and outliers were removed if values were determined to fall outside of ± 2 standard

100 deviations from the mean. All statistical analyses were performed using GraphPad

Prism version 7.0 and Microsoft Excel 2016.

Results Based on previous studies indicating that a 6 μg/kg subcutaneous injection of G-1 induced dendritic spine formation within 40 minutes of treatment (Gabor et al., 2015), a brief pilot study was performed to determine a potential time point for G-1 induced JNK phosphorylation in the hippocampus following subcutaneous injection. Using OVX female CD1 mice, animals were treated with a 6 μg/kg subcutaneous injection of G-1 and subsequently sacrificed at 15 minute time intervals up to 1 hour following injection (pilot n=1 per group).

Although no statistical analyses were performed, the representative immunoblot and semi- quantitative densitometric analyses were used to quantify fold changes in p-JNK expression over total JNK expression, all relative to the β-actin loading control. Semi-quantitative densitometric analyses revealed stable 54 kDa p-JNK expression and an approximate 33% increase in 46 kDa p-JNK expression at the 15 minute time point, relative to the vehicle control.

The 30 minute time point showed fairly stable 54 kDa and 46 kDa p-JNK expression relative to the vehicle control, while the 45 minute and 1 hour time points exhibit a gradual loss of 54 kDa and 46 kDa p-JNK expression relative to the vehicle control.

Based on the very preliminary results of this pilot study, the 15 minute time point was chosen as the most likely to exhibit a G-1 induced JNK phosphorylation. Female CD1 mice

(n=24) were bilaterally ovariectomized and left to recuperate from surgical stress for approximately 15 days. Mice were randomly allocated (n=6 per group) to receive subcutaneous injections of sesame oil and saline vehicle control, G-1 and saline, CORT and sesame oil, or a combination of G-1 & CORT. The representative immunoblot (Figure 5.2 upper panel) shows respective p-JNK expression relative to total JNK expression and protein loading control β-actin levels. The representative western blot suggests a lack of G-1 induced JNK phosphorylation

101 when comparing the vehicle vs G-1 treated groups (Figure 5.2 upper panel). However, the representative western blot also suggests that CORT and G1 & CORT samples exhibit reduced p-JNK expression relative to the vehicle and G-1 treated groups. Semi-quantitative densitometric analyses were performed to determine fold changes in the phosphorylation of the

54 and 46 kDa isoforms of JNK relative to total JNK, normalized to β-actin loading controls.

Following natural log transformation, a one-way ANOVA revealed a significant effect of treatment for both 54 kDa isoform [F(3,19)=7.492, p = 0.0017] and 46 kDa isoform

[F(3,20)=7.116, p =0.0019] (Figure 5.2). Tukey Kramer post-hoc multiple comparison tests revealed significant decrease in 54 kDa JNK phosphorylation in the G-1 + CORT group compared to the vehicle (p = 0.0054) and G-1 (p = 0.0062) groups. Tukey Kramer post-hoc multiple comparison tests revealed a significant reduction in the phosphorylation of 46 kDa JNK in the CORT group when compared to either the G-1 (p=0.0336) or vehicle (p=0.0339) groups and in the G-1 + CORT groups when compared to treatment with G-1 alone (p=0.0109) or the vehicle (p=0.0111) group.

Figure 5.1 Western blot depicting JNK phosphorylation in ovariectomized female CD1 mice treated with 6 μg/kg of G-1. To pilot the amount of time required for subcutaneous G-1 treatment to reach the hippocampus and increase p-JNK levels, ovariectomized female CD1 mice were treated with 6 μg/kg of G-1 (Gabor et al., 2015) and sacrificed at 15 minutes, 30 minutes, 45 minutes, or 1 hour following treatment. Relative to total JNK and β-actin levels, p-JNK expression seems to rapidly increase within 15 minutes of G-1 injection and gradually decline over the remaining 45 minutes.

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Figure 5.2 JNK phosphorylation in the hippocampus of ovariectomized female CD1 mice treated with G-1 in the presence or absence of CORT. Ovariectomized female CD1 mice were subcutaneously injected with a vehicle control, G-1, CORT, or a combination of G-1 & CORT and sacrificed 15 minutes later. (a) Representative immunoblots obtained from hippocampal lysates of control and treated mice separated by SDS-PAGE and transferred to nitrocellulose membranes. The top panel represents phosphorylated 46 and 52 kDa JNK expression, the middle panel depicts total JNK levels and the bottom panel shows β-actin levels from the same blot. Semi-quantitative densitometric measurements are graphically represented for fold changes of 54 (b) and 46 kDa (c) isoforms of p-JNK relative to total JNK and β-actin loading controls. Following natural log transformation to account for inhomogeneity of variance, a one- way ANOVA was used to assess for overall effects of treatment within each isoform. The ANOVA analyses reveal a significant effect of for both 54 kDa isoform [F(3,19)=7.492, p = 0.0017] and 46 kDa isoform [F(3,20)=7.116, p =0.0019]. Tukey Kramer post-hoc multiple comparison tests revealed significant downregulation of 54 kDa p-JNK in the G-1 & CORT group compared to the vehicle (p = 0.0054) and G-1 (p = 0.0062) groups. Tukey Kramer post-hoc multiple comparison tests revealed significant downregulation of 46 kDa p-JNK in the CORT group when compared to the G-1 (p=0.0336) and Vehicle (p=0.0339) groups and in the G-1 & CORT groups when compared to the G-1 (p=0.0109) and Vehicle (p=0.0111) groups (p=**<0.001, p=*<0.05).

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Discussion These findings suggest that physiological circulating concentrations of glucocorticoids may prevent phosphorylation of JNK in the female hippocampus. As a subcutaneous injection with 5 mg/kg of CORT has been shown to rapidly increase plasma glucocorticoids levels within 10 minutes of subcutaneous injection (Choleris et al., 2013), these findings may provide important insight into the rapid and detrimental effects of stressful stimuli within the female hippocampus. Specifically, JNK activation and phosphorylation through GPER has been associated with rapid enhancements in dendritic spine density and cognition (Gabor et al., 2015, Kim et al., 2016). While these findings may elude to a potential interaction between glucocorticoid regulation of GPER activation, the induction of JNK phosphorylation by G-1 alone was not significantly differently from than that seen in hippocampal lysates obtained from vehicle-treated mice. As a result, it is difficult to discern whether or not this loss of p-JNK is being mediated through inhibition of GPER activation or through alternative mechanisms that may regulate JNK signalling. A considerable limitation of this study was the anticipated timing of G-1 induced JNK activation within the hippocampus. While there is considerable in vivo variability in physiological responses, feedback, and timing, a more thorough pilot study should be performed to determine the optimal peak in JNK phosphorylation before the onset of dendritic spine changes that occurs within 40 minutes of injection (Gabor et al., 2015). Higher doses of G-1 should also be investigated to best determine if this loss of JNK phosphorylation is mediated through

GPER inhibition. Despite a lack of induction of p-JNK by G-1, these results provide support for the previous in vitro findings that indicate a reduction in p-JNK following exposure to glucocorticoids. While male rats exhibit significant increases in dendritic

104 spine density and cognitive performance after exposure to a behavioural stressor, their age matched female counterparts trend in the opposite direction, exhibiting loss of dendritic spines, learning and memory (Shors et al., 2001; Wood & Shors, 1998). While the JNK cascade has been shown to play an important role in rapid modulation of synaptic plasticity, apoptosis, and neuronal differentiation (Farías et al., 2009; Seo et al., 2012), this glucocorticoid induced loss of JNK phosphorylation may impair long term potentiation, learning, and memory formation in the female hippocampus (Seo et al.,

2012). As the neuroprotective and neurotrophic effects of estradiol treatment may be inhibited by exposure to a stressful stimulus (Frick et al., 2004), these findings may provide support for an underlying signalling mechanism that may mediate these behavioural and morphological impairments. The evident morphological and behavioural sex differences seen following stress in the male and female hippocampus may be at least partially accounted for by the glucocorticoid induced loss of JNK phosphorylation that may be mediaed by GPER activation. If this downregulation is indeed mediated through inhibition of GPER, these findings may help explain why the female hippocampus is more vulnerable to the detrimental effects of stress.

These in vivo findings may have further implications for the developing and aging female brain. While activation of sex steroid receptors during development play an important role in sexual differentiation of the brain (MacLusky & Naftolin, 1981; Arnold,

2009), exposure to elevated stress hormones may have serious implications for regulation of estrogen sensitivity in the fetal hippocampus. Additionally, these findings may also have profound implications for the aging female brain that is predisposed to sexually differentiated neurodegenerative conditions such as Alzheimer’s disease. The

105 natural loss of estrogen signalling that occurs in the aging female may be further confounded by glucocorticoid induced downregulation of GPER or functional JNK signalling that may be ultimately detrimental to the female hippocampus.

Conclusion In conclusion, the findings of the current study provide evidence to support glucocorticoid induced downregulation of JNK activation. While the G-1 induction of p-

JNK expression was not significantly different than that of the vehicle, the loss of basal p-JNK expression may still have serious implications for impaired neurotrophic and neuroprotective effects in the female hippocampus. These findings may elude to an underlying mechanism that may help explain the sexually differentiated behavioural and morphological effects exhibited following exposure to a stressful stimulus.

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CHAPTER 6: OVERALL CONCLUSIONS AND FUTURE DIRECTIONS

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Neuronal tissue culture models have proven challenging to study cellular and molecular mechanisms in vitro as they are unable to propagate and are, therefore, short lived and time-consuming to generate (Gordon et al.,2013). Many neuroblastoma and neuron-derived cells lines divide but are limited in their expression of specific neurotransmitter markers and steroid hormone receptors (Kovalevich & Langford,

2013). As a result, Chapter 1 of the present study sought to characterize novel lines of female and male-derived immortalized murine hippocampal neurons. The rapid division of both mHippoE-14 and mHippoE-18 cell lines makes this model very efficient and easy to work with. However, as first described by Gingerich et al. (2010), these cell lines had yet to be characterized for important transcription factors as well as steroid and neurotransmitter receptors. The present study characterized the mRNA expression of receptors involved in modulating the stress response, gonadal axis, metabolism, and neurotransmission. The identified receptor expression of Mineralocorticoid receptor

(MR), glucocorticoid receptor (GR), Luman, androgen receptor (AR), G-protein coupled estrogen receptor (GPER), cannabinoid receptor type I (CB1), Thyroid hormone receptor-alpha (THR-α) and THR-β provides a dynamic framework to investigate hormonal interactions and regulatory effects within hippocampal neurons in a simple and efficient in vitro model. However, the lack of GABAergic and glutamatergic receptor mRNA expression in both mHippoE-14 and mHippoE-18 cell lines indicates that these cell lines may not be the best model for investigating steroid interactions with mechanisms of neurotransmission. These current findings suggest that the mHippoE-14 and mHippoE-18 cell lines may provide an important in vitro model that may be easily manipulated to identify steroid interactions within the hippocampus. Future studies

108 should confirm respective protein expression to further validate the use of these models for investigating the regulatory and functional interactions between key cellular constituents.

Based on the concurrent expression of GR and GPER identified in both

mHippoE-14 and mHippoE-18s, the findings of the present study indicate that

glucocorticoid receptor activation may downregulate GPER protein expression in female

derived mHippoE-14 hippocampal neurons only. This reduced protein expression of

GPER may help explain the loss of estradiol sensitivity exhibited within the female

hippocampus after exposure to a stressful stimulus. This stress hormone induced loss

of GPER expression may have serious implications for sex differences observed in

cognition and hippocampal plasticity with the aging brain. As a result, the next objective

was to determine whether or not glucocorticoid treatment would similarly regulate GPER

mediated signalling mechanisms. It was subsequently determined that 24 hour in vitro

glucocorticoid pre-treatment inhibited G-1 induced GPER activation in the female

derived mHippoE-14s as is evident by the significant loss of JNK phosphorylation.

Given these in vitro findings, we sought to determine whether or not these regulatory

effects of glucocorticoid treatment are also exhibited in vivo. Findings from the in vivo

experiment suggest that 6 μg/kg of G-1 may not have been sufficient to significantly

induce JNK phosphorylation in female CD1 mice within 15 minutes. While previous

studies have indicated that subcutaneous injections of G-1 induce dendritic spine

formation within 40 minutes in the female hippocampus (Gabor et al., 2015), the timing

of GPER induced JNK phosphorylation is relatively unknown. Previous studies have

used intrahippocampal infusions to demonstrate activation of GPER induced JNK

109 signalling within 5 minutes of G-1 treatment infusion (Kim et al., 2016). However, this significant increase in JNK phosphorylation has been shown to return quickly to baseline within 15 minutes of dorsal intrahippocampal infusion with 4 ng/hemisphere of

G-1 (Kim et al., 2016). As a result, timing of G-1 induced GPER activation via subcutaneous injection is difficult to discern given time spent in circulation and crossing the blood brain barrier to reach the hippocampus. However, the findings of the in vivo study do suggest that treatment with subcutaneous CORT injections may significantly downregulate phosphorylation of JNK. Subcutaneous injections of 5 mg/kg of CORT have been previously shown to rapidly increase circulating levels of glucocorticoids within 10 minutes of injection (Choleris et al., 2013). As a result, the timing of CORT elevation may also confound the timing of G-1 induced JNK phosphorylation in the G-1

& CORT group. Further studies must include a more extensive pilot investigation to best elucidate the optimal dosage and timing of subcutaneous G-1 induced JNK phosphorylation in the hippocampus. This would allow us to further investigate these regulatory mechanisms and interactions between stress and GPER activation in the hippocampus.

While the in vitro findings indicated an approximate 35% reduced expression of

GPER protein following 24 hour treatment with dexamethasone, the in vitro 24 hour dexamethasone pre-treatment much more significantly reduced expression of p-JNK.

While some of this reduced p-JNK may be attributable to loss of GPER expression, it is likely that this reduction in GPER expression does not account for the entirety of glucocorticoid regulated JNK phosphorylation. This suggests that glucocorticoids may act through an alternative mechanism such as inhibiting synthesis of GPER agonists,

110 antagonizing hippocampal GPER, or inhibiting JNK phosphorylation downstream of

GPER activation. The findings from the in vivo study suggest that glucocorticoids may regulate JNK phosphorylation independently of changes in GPER expression as it would be unlikely that protein expression would significantly change within 15 minutes of treatment. However, the findings from the in vivo study also indicate that CORT may be capable of regulating JNK phosphorylation independently of GPER activation as 6

μg/kg of G-1 failed to significantly induce JNK phosphorylation at 15 minutes. While the use of CORT is physiologically more meaningful, it differs from the experimental conditions used in vitro that selectively activated the glucocorticoid receptor and this may also contribute to differences seen between these studies.

Collectively, the findings from the in vitro work clearly indicate that there may be an interaction between glucocorticoid receptor activation and GPER expression and functional signalling in female hippocampal neurons. Given the nature of in vitro work, these findings may be the result of a controlled environment that lacks a physiological stress axis feedback mechanism. While the in vivo study suggests that an elevation in circulating glucocorticoids (CORT) may have similar inhibitory effects on JNK phosphorylation in the female hippocampus, it has yet to be determined if this mechanism is mediated through inhibition of GPER in the intact hippocampus.

GPER activation has been shown to play a profound role in mediating some of estradiol’s rapid neurotrophic and neuroprotective effects in the female hippocampus

(Gabor et al., 2015; Gingerich et al., 2010). The stress hormone-induced loss of GPER signalling may provide further support for a mechanism that may underly the sex differences observed in hippocampal morphology, cognition, and behaviour following

111 stress exposure (Shors et al., 2001; Wood & Shors, 1998). The loss of GPER mediated

JNK activation may have profound implications for the aging brain. While cortisol levels continuously rise in both aging males and females, these findings may help explain the sexually differentiated hippocampal atrophy and cognitive impairments that occur in the female hippocampus (Lupien et al., 1998). This stress induced loss of functional GPER activation may help explain why the female hippocampus is particularly vulnerable to the detrimental effects of stress.

These regulatory effects may have serious implications for hippocampal morphology and cognitive function. As a result, future studies should seek to determine the importance of this stress induced loss of GPER activation by investigating these mechanisms in vivo. As GPER activation plays a profound role in rapidly modulating dendritic spine density, future studies should investigate whether or not stress hormones prevent these G-1 mediated neurotrophic effects at both short- and long-term time points to most accurately mimic the design of this in vitro work. Additionally, studies should consider the behavioural implications of this loss as GPER has been shown to facilitate object and social recognition performance. It may also be beneficial to consider the influence of naturally circulating ovarian hormones through investigating intact females treated with letrozole or G-15 to determine the effects of inhibited estradiol synthesis and GPER activation. Collectively, these studies could further elucidate the role of the stress response in GPER regulation as it pertains to sexually differentiated impairments in hippocampal learning and memory.

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APPENDICES Appendix 1: Sanger sequencing results for primer pairs that successfully amplified DNA fragments from mHippoE immortalized hippocampal neurons

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Figure 6.1. Confirmation of correct amplicon sequencing with forward and reverse primers. Output generated by the National Centre for Biotechnology Information’s basic local alignment search tool (BLAST) for mus musculus HPRT, MR, GR, AR, GPER, CB1, THRα, THRβ, AMPA-R3, GABRA-α1, and GABRA-β3 run with the sample nucleotide FASTA sequence generated by Sanger sequencing using the respective forward and reverse primers. The optimal alignment is indicated by the matching between the query (mus musculus GPER) and the subject (mHippoE sample) nucleotide sequences. Max score indicates total alignment coverage while percent identity indicates degree of alignment between query and subject sequences.

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Appendix 2: Antibody information for western blotting experiments

Table 3. Antibodies used in western blotting experiments

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