The Molecular Characterization and Role of Teneurin C-Terminal Associated Peptide (TCAP)-1 in the Regulation of Neuronal Cytoskeletal Dynamics and Male Reproduction

Home , Arrestin, Teneurin

The Molecular Characterization and Role of Teneurin C-terminal Associated Peptide (TCAP)-1 in the Regulation of Neuronal Cytoskeletal Dynamics and Male Reproduction

Dhan Sidhartha Chand

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Cell and Systems Biology University of Toronto

The Molecular Characterization and Role of Teneurin C-terminal Associated Peptide (TCAP)-1 in the Regulation of Neuronal Cytoskeletal Dynamics and Male Reproduction

Dhan Sidhartha Chand

Doctor of Philosophy

Cell and Systems Biology University of Toronto

2013

Abstract

Teneurin C-terminal associated peptides (TCAPs) are a novel family of peptides encoded on the last exon of the teneurin genes. The predicted peptide sequences are highly conserved across metazoans and possess the structural hallmarks of a cleavable bioactive peptide that are 40 or 41 amino acid residues. One of the peptides in the family, TCAP-1, is a potent regulator of neurite outgrowth and dendritic spine density in the hippocampus and inhibits corticotropin-releasing factor (CRF)-associated stress-induced and cocaine-seeking behaviours. The effects of TCAP-1 are long lasting, suggesting that TCAP-1 plays a significant role in the regulation of cell-to-cell communication and cellular plasticity. Moreover, TCAP-1 regulates cellular energy, metabolism and cell survival and may, therefore, possess functional attributes outside of the CNS. However, the molecular mechanisms associated with TCAP-1-mediated trophic effects are not known. My research was aimed to 1) determine whether TCAP-1 exerts its effects as part of a direct teneurin-1 function, whereby TCAP-1 represents a functional region of the large teneurin-1 protein, or if it has an independent role, either as a splice variant or post-translational proteolytic cleavage product of teneurin-1; 2) map the distribution of TCAP-1-immunoreactivity and TCAP-

1 binding sites in mouse; 3) elucidate the molecular mechanism by which TCAP-1 regulates cytoskeletal dynamics; and 4) investigate a role for TCAP-1 in male reproduction. My research establishes that the C-terminal region of teneurin-1, corresponding to TCAP-1, can be both structurally and functionally independent from teneurin-1 in both the brain and testis of the adult mouse. Furthermore, I provide novel evidence that functionally links the teneurin-TCAP-1 system with the dystroglycan complex and provide new insight into the molecular and signaling mechanisms by which TCAP-1 regulates cytoskeletal dynamics. These studies implicate the teneurins in a broader range of neuroendocrine and trophic functions than previously thought and furthers our understanding of the mechanisms associated with TCAP-1-mediated function in the body.

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Acknowledgments

My doctoral studies have been an enriching, enjoyable and fulfilling experience. I am forever grateful to the many people who have helped me along the arduous road to success. First, I thank my supervisor and mentor Dr. David Lovejoy. David, thank you for facilitating my dreams of being a scientist, the opportunity to purse many exciting projects, your unwavering support and belief in my abilities, and providing me with all the opportunities and tools needed to succeed as a scientist. It has been a great privilege and honour to be your student. I consider myself extremely fortunate to have had you as a supervisor. Thank you for the incredible journey.

Over the course of my graduate studies, I had the pleasure of working with many brilliant scientists and colleagues. I acknowledge Dr. Arij Al Chawaf, who provided me with tremendous support and guidance during my first research project; Dr. Dalia Barsyte-Lovejoy, from whom I learnt many technical and problem-solving skills early on in my graduate studies; Dr. Cláudio

Casatti, whose dedication and patience inspired me to persevere during difficult experiments;

Lifang Song, for her technical assistance with many experiments and insightful discussions; and to Laura Tan, Tanya Nock, Tiffany Ng, Reuben DeAlmeida, Mei Xu and Louise deLannoy, for their friendship, support in the lab and numerous adventurous conference trips. I also acknowledge, Michael Colacci and Autumn Otchengco, two stellar undergraduates whom I had the privilege of mentoring.

A special thank you to my collaborators at the Samuel Lunenfeld Research Institute at Mount

Sinai Hospital, Dr. Theodore Brown, for providing me with great advice, insight and guidance on many scientific matters; and Dr. Alexandra Kollara, who has been a great friend, mentor and teacher throughout my graduate studies.

My scientific achievements would not have been possible without the love and support of my family and friends outside of the laboratory. First and foremost, I thank my parents, Pamela

Chand and Rabindra Chand for their sacrifices, encouragement and never-ending support. The sacrifices you have made for me always inspire me to overcome whatever challenge I face. You are my tower of strength. Thank you for challenging me to be best that I can be.

I also thank my ‘second-parents’, John and Anne-Marie Webster for their unwavering support, love and confidence in me during this journey. Thank you for providing me with a ‘home away from home’ and a refuge from the rigours of university life. To my sisters, Thalia, Tejika and

Ryhan Chand, and Alexandria and Angelique Webster, thank you for always being there for me and filling my life with so much joy, laughter and love when I needed it the most.

I express a special thank you to my partner, Melissa Orilall, for her steadfast support, understanding and motivation during this journey. Thank you for uplifting my spirits and always encouraging me to never give up. It is impossible for me to express in words how much I appreciated your love and support throughout these years. Also, I express sincere gratitude to

Mohan and Paulette Orilall for their support.

I am eternally grateful and blessed to have you all in my life. I am reminded that “we are all connected to each other, biologically; to the earth, chemically; and to the rest of the universe, atomically” – Neil DeGrasse Tyson.

Table of Contents

Abstract ...... ii

Acknowledgments ...... iv

List of Figures ...... xiii

List of Tables ...... xv

List of Abbreviations ...... xvii

1 Chapter One: Introduction: Integrating developmental proteins, trophic factors and the cytoskeleton in the regulation of brain and testis function...... 1

1.1 Abstract ...... 1

1.2 The Teneurin family of proteins ...... 2

1.2.1 Teneurin Structure...... 4

1.2.2 Processing of the teneurin proteins ...... 6

1.2.3 Teneurin Expression ...... 8

1.2.4 Teneurin Function ...... 10

1.2.5 Teneurin interacting proteins and cell signalling ...... 12

1.3 Dystroglycans ...... 16

1.3.1 Dystroglycan Structure ...... 18

1.3.2 Dystroglycan interacting proteins and cell signalling ...... 19

1.4 Developmental proteins in the mammalian testes ...... 22

1.5 Regulation of Cytoskeletal dynamics ...... 26

1.5.1 Actin cytoskeleton...... 26

1.5.2 Microtubule cytoskeleton ...... 28

1.5.3 Cytoskeletal dependent cellular protrusions: Lamellipodia and filopodia formation ...... 31

1.6 Teneurin C-terminal Associated Peptide ...... 33

1.7 Objectives and Hypothesis ...... 37

1.8 References ...... 39

2 Chapter Two: Processing, localization and uptake of the C-terminal region of teneurin-1 ...... 52

2.1 Abstract ...... 52

2.2 Introduction ...... 53

2.3 Materials and Methods ...... 55

2.3.1 Animals ...... 55

2.3.2 Cell Culture ...... 56

2.3.3 Northern blot analysis ...... 56

2.3.4 Synthesis of TCAP Peptides ...... 57

2.3.5 Production of TCAP-1 Antisera ...... 57

2.3.6 Western Blot Analysis ...... 59

2.3.7 Immunofluorescent and immunohistochemical staining ...... 60

2.3.8 Preparation of Fluoresceinisothiocyanate (FITC)-labeled TCAP-1 ...... 62

2.3.9 FITC-TCAP-1 uptake and binding studies ...... 62

2.3.10 Image Analysis ...... 63

2.4.0 Results ...... 63

2.4.1 Transcription and translation of TCAP-1 ...... 63

2.4.2 Localization of Teneurin-1 and TCAP-1 to hippocampal cells and tissues ...... 66

2.4.3 Caveoli-dependent endocytosis of TCAP-1 in hippocampal cells ...... 69

2.5 Discussion ...... 72

2.6 References ...... 81

3 Chapter Three: Localization of TCAP-1-immunoreactivity in the adult mouse brain ...... 86

3.1 Abstract ...... 86

3.2 Introduction ...... 87

3.3 Materials and Methods ...... 89

3.3.1 Animals ...... 89

3.3.2 Synthesis of TCAP Peptides ...... 89

3.3.3 Antibody Characterization and Controls ...... 90

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3.3.4 Immunofluorescent and immunohistochemical staining ...... 91

3.3.5 Imaging ...... 92

3.4 Results ...... 93

3.4.1 Localization of TCAP-1-immunoreactivity in the mouse brain ...... 93

3.5 Discussion ...... 104

3.5.1 Methodological considerations ...... 105

3.5.2 Comparison of TCAP-1 and teneurin localization in the vertebrate brain ...... 106

3.5.3 Cerebral Cortex ...... 106

3.5.4 Hindbrain: Cerebellum, Medulla and Pons ...... 107

3.5.5 Midbrain ...... 109

3.5.6 Thalamus and Hypothalamus ...... 110

3.5.7 Hippocampus ...... 112

3.5.8 Limbic Regions ...... 112

3.5.9 Integrating TCAP-1-regulated psychiatric disorders and sensorimotor function ...... 113

3.6 References ...... 117

4 Chapter Four: Regulation of cytoskeletal dynamics by TCAP-1 ...... 122

4.1 Abstract ...... 122

4.2 Introduction ...... 123

4.3 Materials and Methods ...... 125

4.3.1 Cell Culture ...... 125

4.3.2 Antibodies and Reagents ...... 125

4.3.3 Synthesis of TCAP Peptides ...... 126

4.3.4 Preparation of Fluoresceinisothiocyanate (FITC)-labeled TCAP-1 ...... 127

4.3.5 FITC-TCAP-1 binding studies ...... 127

4.3.6 Immunofluorescent staining ...... 128

4.3.7 Cytoskeletal and Morphological analysis ...... 128

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4.3.8 Western Blot Analysis ...... 130

4.3.9 Image Analysis ...... 131

4.3.10 Statistical Analyses ...... 132

4.4 Results ...... 132

4.4.1 Binding of TCAP-1 to hippocampal cells and tissues...... 132

4.4.2 TCAP-1 co-localization with the Dystroglycan complex ...... 134

4.4.3 TCAP-1 increases actin polymerization and tubulin cytoskeletal elements ...... 136

4.4.4 TCAP-1 induces filopodia formation and elongation ...... 139

4.4.5 TCAP-1 induces ERK-dependent phosphorylation of Stathmin ...... 141

4.4.6 TCAP-1 induces phosphorylation of p90RSK and filamin A ...... 144

4.5 Discussion ...... 146

4.6 References ...... 152

5 Chapter Five: In vivo effects of TCAP-1 in the testes and epididymis of the adult mouse ...... 157

5.1 Abstract ...... 157

5.2 Introduction ...... 158

5.3 Materials and Methods ...... 161

5.3.1 Animals ...... 161

5.3.2 Antibodies ...... 161

5.3.3 Synthesis of TCAP-1 Peptides ...... 162

5.3.4 TCAP-1 administration and tissue collection ...... 162

5.3.5 Fecal collection ...... 162

5.3.6 Preparation of Fluoresceinisothiocyanate (FITC)-labeled TCAP-1 ...... 163

5.3.7 Histology and Immunofluorescent labeling ...... 163

5.3.8 FITC-TCAP-1 binding studies ...... 164

5.3.9 Image and morphological analysis ...... 165

5.3.10 Serum and fecal analysis ...... 166

5.3.11 Statistical Analyses ...... 167

5.4 Results ...... 167

5.4.1 Teneurin-1 and TCAP-1 localization in the mouse testis...... 167

5.4.2 Teneurin-1 and TCAP-1 co-localizes with dystroglycan in the testis and epididymis ..... 169

5.4.3 TCAP-1 binding in the mouse testis and epididymis ...... 172

5.4.4 Localization of TCAP-1 and TCAP-1 binding sites in the seminal vesicles ...... 174

5.4.5 TCAP-1-immunoreactivity in the epididymis of TCAP-1-treated mice ...... 174

5.4.6 TCAP-1 increases testis size ...... 176

5.4.7 TCAP-1 regulates seminiferous tubule size ...... 178

5.4.8 TCAP-1 increases the size of the caput and corpa epididymis ...... 181

5.4.9 TCAP-1 increases circulating levels of testosterone ...... 183

5.5 Discussion ...... 185

5.6 References ...... 194

6 Chapter Six: Significance of findings, future studies and conclusions ...... 200

6.1 Abstract ...... 200

6.2 Teneurin-TCAP relationship: An evolutionary conserved system ...... 201

6.2.1 The teneurin-dystroglycan complex and the putative TCAP-1 receptor ...... 205

6.2.2 Further interactions at the plasma membrane: TCAP-1 co-localizes with 17 β- hydroxysteroid dehydrogenase 10 (HSD17β10) ...... 208

6.2.3 Identifying TCAP-1 binding proteins ...... 211

6.3 TCAP-1 signal transduction: A novel mechanism in the regulation of cytoskeletal dynamics and cell survival...... 215

6.3.1 A role for TCAP-1-mediated cytoskeletal regulation during cell division ...... 219

6.4 Regulating CRF-mediated and stress-induced physiology and behaviours ...... 221

6.5 A broader role for TCAP-1 in reproduction ...... 227

6.5.1 Furthering the case for TCAP-1 in male reproduction ...... 227

6.5.2 A role for TCAP-1 in female reproduction ...... 231

6.5.3 Enhancing fertility with TCAP-1 ...... 236

6.6 A role for TCAP-1 in cancer ...... 237

6.7 Conclusions ...... 241

6.8 References ...... 243

List of Figures

Figure 1.1. Proposed structural properties of the mouse teneurins ...... 5

Figure 1.2. Domain organization of vertebrate teneurins ...... 7

Figure 1.3. Current teneurin signalling model ...... 15

Figure 1.4. A diagrammatic representation of α- and β-dystroglycan at the cell surface ...... 18

Figure 1.5. Current model of the the laminin-dystroglycan-dystrophin signaling cascade ...... 21

Figure 1.6. Cytoskeletal elements and the formation of lamellipodia and filopodia ...... 32

Figure 2.1. Expression and processing of the TCAP portion of mouse teneurin-1 ...... 65

Figure 2.2. TCAP-1 and teneurin-1-immunoreactivity in mouse E14 hippocampal cells ...... 67

Figure 2.3. TCAP-1-immunoreactivity in the neuronal layers of the mouse hippocampal formation ...... 68

Figure 2.4. TCAP-1 uptake in mouse E14 hippocampal cells ...... 70

Figure 2.5. Caveolin-dependent endocytosis of FITC-[K8]-TCAP-1 in mouse E14 hippocampal cells .... 71

Figure 2.6. Proposed mechanism for TCAP-1 processing, release and uptake in hippocampal cells ...... 80

Figure 3.1. Distribution of TCAP-1-immunoreactivity in the cerebral cortex ...... 94

Figure 3.2. Localization of immunoreactive-TCAP-1 in the cerebellum ...... 95

Figure 3.3. Distribution of immunoreactive-TCAP-1 in the pons ...... 96

Figure 3.4. Distribution of immunoreactive-TCAP-1 in the midbrain ...... 97

Figure 3.5. Distribution of immunoreactive-TCAP-1 in the medulla ...... 98

Figure 3.6. Localization of immunoreactive-TCAP-1 in the thalamus ...... 99

Figure 3.7. Localization of immunoreactive-TCAP-1 in the hypothalamus ...... 100

Figure 3.8. Localization of immunoreactive-TCAP-1 in the hippocampus ...... 101

Figure 3.9. Localization of immunoreactive-TCAP-1 in the bed nucleus of the stria terminalis ...... 102

Figure 3.10. Distribution of immunoreactive-TCAP-1 in the limbic regions ...... 103

Figure 3.11. Diagrammatic representation of the distribution of immunoreactive-TCAP-1 in the adult mouse brain ...... 116

Figure 4.1. FITC-[K8]-TCAP-1 binding in mouse hippocampal formation ...... 133

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Figure 4.2. TCAP-1 binding in immortalized mouse E14 hippocampal cells ...... 134

Figure 4.3. TCAP-1 co-localization with the dystroglycan complex in immortalized mouse E14 hippocampal cells ...... 135

Figure 4.4. TCAP-1 treatment increases actin polymerization in immortalized E14 hippocampal cells . 137

Figure 4.5. TCAP-1 treatment increase tubulin immunoreactivity ...... 138

Figure 4.6. TCAP-1 treatment increases filopodia formation and mean filopodia length in immortalized E14 hippocampal cells ...... 140

Figure 4.7. TCAP-1 activates the Mitogen-activated Protein Kinase (MAPK/ERK) pathway and induces ERK-dependent stathmin phosphorylation at serine-25 in E14 hippocampal cells ...... 143

Figure 4.8. TCAP-1 induces ERK-dependent p90RSK phosphorylation and filamin A phosphorylation in E14 hippocampal cells ...... 145

Figure 4.9. Proposed mechanism of TCAP-1 effects on the cytoskeleton in the mouse hippocampus .... 147

Figure 5.1. Teneurin-1 and TCAP-1-immunoreactivity in the seminiferous tubule of the mouse testis .. 168

Figure 5.2. Teneurin-1 and TCAP-1 co-localizes with the Dystroglycan complex in the seminiferous tubule of the mouse testis ...... 170

Figure 5.3. Teneurin-1 and TCAP-1-immunoreactivity and co-localization with the dystroglycan complex in the mouse epididymis ...... 171

Figure 5.4. FITC-[K8]-TCAP-1 binding in the seminiferous tubule of the mouse testis ...... 172

Figure 5.5. FITC-[K8]-TCAP-1 binding in the mouse epididymis ...... 173

Figure 5.6. Localization of TCAP-1-immunoreactivity and FITC-[K8]-TCAP-1 binding sites in the mouse seminal vesicles ...... 174

Figure 5.7. TCAP-1-immunoreactivity in the epididymides of TCAP-1-treated mice ...... 175

Figure 5.8. TCAP-1 treatment increases testicular size ...... 177

Figure 5.9. TCAP-1 treatment regulates seminiferous tubule diameter ...... 179

Figure 5.10. TCAP-1 treatment increases the size of the caput and corpa epididymis ...... 182

Figure 5.11. Effect of TCAP-1 treatment on fecal testosterone and corticosterone ...... 183

Figure 6.1. TCAP-immunoreactivity in Ciona intestinalis ...... 204

Figure 6.2. MALDI quadrupole time-of-flight analysis of synthetic TCAP-1 ...... 205

Figure 6.3. Internalization of teneurin-1 by TCAP-1 ...... 207

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Figure 6.4. FITC-labelled TCAP-1 co-localizes with mitochondrial marker hydroxysteroid (17-β) dehydrogenase 10 (HSD17β10) ...... 209

Figure 6.5. Characterizing TCAP-1 binding proteins ...... 213

Figure 6.6. FITC-labeled TCAP-1 binding in fixed mouse E14 hippocampal cells ...... 214

Figure 6.7. TCAP-1 binding sites in the nucleus ...... 215

Figure 6.8. TCAP-1 dephosphorylates stathmin at serine-16 and -63 and induces phosphorylation BAD in E14 hippocampal cells ...... 217

Figure 6.9. Proposed TCAP-1 signal transduction...... 218

Figure 6.10. TCAP-1 co-localizes with tubulin during cell division in mouse E14 hippocampal cells ... 220

Figure 6.11. TCAP-1-immunoreactivity in adult mouse female reproductive tract ...... 232

Figure 6.12. FITC-[K8]-TCAP-1 binding in the adult mouse female reproductive tissue ...... 234

Figure 6.13. Western blot analysis on cancer cell lines using TCAP-1 antiserum ...... 238

Figure 6.14. TCAP-1 expression in human testes with mixed germ cell tumor ...... 239

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List of Tables

Table 1.1: Teneurin expression in mouse ...... 10

Table 5.1: Mean serum testosterone, progesterone, prolactin and growth hormone levels (ng/ml) from saline-treated, 25pmol and 250pmol TCAP-1-treated mice after 9 days of treatment ...... 184

Table 6.1: Comparing the molecular characteristics of CRF-associated pathologies and TCAP-1 effects in the central nervous system (CNS) ...... 226

List of Abbreviations

α-DG alpha dystroglycan β-DG beta dystroglycan γ-TuRC γ-tubulin ring complex 7n facial nerve 7N facial nucleus aca anterior part of the anterior commissure AcbSh nucleus accumbens shell AchC nucleus accumbens core aci intrabulbar part of the anterior commissure ADP Adenosine diphosphate Amg amygdaloid nucleus ANOVA analysis of variance AOB accessory olfactory bulb AOD dorsal part of the anterior olfactory nucleus AOM medial anterior olfactory nucleus AOP posterior part of the anterior olfactory nucleus AOV ventral anterior olfactory nucleus APP amyloid precursor protein Arp2/3 actin-related proteins 2 and 3 ATP Adenosine-5'-triphosphate Aβ β-amyloid BAD Bcl-2-associated death promoter BCA bicinchoninic acid Bcl-2 B-cell lymphoma 2 BDNF brain-derived neurotrophic factor BLA basolateral nucleus of the amygdala BSA bovine serum albumin BST bed nucleus of the stria terminalis BSTMA anterior part of the medial division of the BST

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BSTMP medial division of the BST C/EBP CCAAT/enhancer binding protein CA Cornu Ammonis CAP/ponsin c-Cb1 associated protein Cb cerebellum cc corpus callosum Cdc42 cell division control protein 42 homolog Cdk1 cyclin-dependent kinase 1 CHOP C/EBP homology protein CNS central nervous system CNTF ciliary neurotrophic factor cp corpus luteum CPu caudate putamen CPZ chlorpromazine CREB cyclic adenosine 3',5'-monophosphate response element binding protein CRF corticotropin-releasing factor CRFR CRF receptor Cu/Zn Copper/Zinc DABCO 1,4-diazabicyclo[2.2.2]octane DAG1 dystroglycan gene (mouse) DAPI 4',6-diamidino-2-phenylindole DG dentate gyrus dgn-1 dystroglycan gene (Caenorhabditis elegans) DHT dihydrotestosterone DIC Differential interference contrast microscopy DMEM Dulbecco’s Modified Eagle Medium DMSO dimethyl sulfoxide DOC downstream of CHOP ECD extracellular domain ECM extracellular matrix EF-1α elongation factor 1α

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EGF epidermal growth factor Egr1 early growth response 1 ELISA enzyme-linked immunosorbent assay Ena/VASP vasodilator-stimulated phosphoprotein epi-1 laminin α-chain gene (Caenorhabditis elegans) EPM elevated plus maze ERK extracellular signal-regulated kinase FAK focal adhesion kinase FBS fetal bovine serum FCA Freund’s complete adjuvant FGF fibroblast growth factor fi fimbria of the hippocampus FITC fluoresceinisothiocyanate FSH Follicle stimulating hormone GDP Guanosine diphosphate GEF Guanine nucleotide exchange factors GFAP glial fibrillary acidic protein GLUT glucose family of transporters GPCR G-protein coupled receptor Grb2 growth factor receptor-bound protein 2 GRK G protein-coupled receptor kinase GTP Guanosine-5'-triphosphate H&E Hematoxylin and Eosin HPA hypothalamic-pituitary-adrenal axis HPG hypothalamic-pituitary gonadal axis HPLC high performance liquid chromatography HSD17β10 17 beta-Hydroxysteroid dehydrogenase 10 IC inferior colliculus ICD intracellular domain ICV intracerebroventricular IGF-1 insulin-like growth factor 1

xviii ina-1 α-integrin gene (Caenorhabditis elegans) IntP interposed cerebellar nucleus IO inferior olive ir immunoreactive IV intravenous IVF in vitro fertilization JNK c-Jun N-terminal kinase LC locus coeruleus LD laterodorsal thalamic nucleus LDH-A lactate dehydrogenase A lfp longitudinal fasciculus of the pons LG laminin G LH lateral hypothalamic area ll lateral lemniscus of the pons LPH1 latrophilin 1 LPMR mediorostral part of the lateral posterior thalamic nucleus LS lateral septal nucleus m/z mass to charge ratio M1 primary motor cortex M2 secondary motor cortex MAP mitogen-activated protein MAP2 microtubule-associated protein 2 MAPK mitogen-activated protein kinase MAPs microtubule-associated proteins MBD1 methyl-CpG-binding protein 1 MAPK mitogen-activated protein kinase MEK mitogen-activated protein kinase kinase ml medial lemniscus MPA medial preoptic area MPta medial parietal association cortex MTOC Microtubule Organizing Center

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NGF nerve growth factor NGS Normal Goat Serum NLS nuclear localization signal OF open field tests PBS phosphate buffer saline PBS-T PBS Tween-20 Pef perifornical nucleus PFA paraformaldehyde Pir piriform cortex PKA protein kinase A PML promyelocytic leukemia protein Pn pontine nuclei PnO oral pontine reticular nucleus Py pyramidal layer RGD arginine-glycine-aspartic acid RIP regulated intramembrane proteolysis RIPA radioimmunoprecipitation assay ROS reactive oxygen species RSA retrosplenial agranular cortex RSG retrosplenial granular cortex RSK ribosomal S6 kinase; RT room temperature SC superior colliculus SC-TCAP-1 Scrambled Teneurin C-terminal Associated Peptide SH2 Src homology 2 SH3 Src homology 3 SL stratum lucidum SNC substantia nigra pars compacta SNR substantia nigra pars reticulata SO stratum oriens SOD superoxide dismutase

SpVe spinal vestibular nucleus SR stratum radiatum STMN stathmin TCAP Teneurin C-terminal Associated Peptide ten-1 teneurin-1 gene (Caenorhabditis elegans) ten-a tenascin-like molecule accessory ten-m tenascin-like molecule major TFA trifluoroacetic acid TM transmembrane domain TNFα tumor necrosis factor-α V2ML mediolateral area of the secondary visual cortex V2MM mediomedial area of the secondary visual cortex VMH ventromedial hypothalamic nucleus VO ventral orbital cortex VP ventral pallidum VPL ventral posterolateral thalamic nucleus VPM ventral posteromedial thalamic nucleus VTA ventral tegmental area WASp Wiskott-Aldrich syndrome protein YD tyrosine-aspartic acid zic-1 zinc finger protein

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1 Chapter One: Introduction: Integrating developmental proteins, trophic factors and the cytoskeleton in the regulation of brain and testis function.

1.1 Abstract

The regulation of neuronal plasticity and tissue morphology is highly dependent on a combination of several key events, such as cell-cell and cell-extracellular matrix (ECM) interactions, trophic factors, receptor-ligand binding, and the subsequent recruitment of multiple signalling pathways. Central to all of these processes is the complex and dynamic regulation of cytoskeletal elements that not only aids in the organization of the molecular machinery but also represents the foundation that underlies cell shape and cell movement. Despite the enormous complexity of metazoan species, the repertoire of known signalling systems that coordinate these developmental and cytoskeletal-dependent processes are limited and relatively similar between neural and non-neural tissues. Therefore, they are likely to share a common set of evolutionary conserved substrates and mechanisms. Thus the elucidation of such integrative systems is of considerable interest. One such emerging candidate system is the recently described teneurin family of proteins. These proteins are conserved across vertebrate and invertebrate species and play a fundamental role in development both in neural and non-neural tissue. At the carboxyl terminal tip of all teneurins are a novel family of peptides termed the teneurin C-terminal associated peptides (TCAPs). The TCAP sequence is 40-41 amino acids long and possesses the characteristics of a bioactive peptide. One member of the family, TCAP-1, has shown both in vitro and in vivo effects, including the modulation of neuronal morphology, conferring neuroprotection under stress conditions, and inhibiting corticotropin-releasing factor (CRF)-

1 induced stress-associated behaviours. TCAP’s precise function, however, has not yet been elucidated, but current studies point to a complex signalling system involving the recruitment of kinases and cytoskeletal proteins. Therefore, the objective of this thesis is to characterize the teneurin-TCAP-1 system and elucidate a mechanism by which TCAP-1 exerts its effects in the mouse brain and testis.

1.2 The Teneurin family of proteins

The teneurins comprise a family of four glycosylated type II transmembrane proteins that were originally discovered in Drosophila as tenascin-like molecule accessory (ten-a;

Baumgartner and Chiquet-Ehrismann, 1993), tenascin-like molecule major (ten-m; Baumgartner et al., 1994) and odd oz (odz; Levine et al., 1994), by two independent groups in a search intended to identify orthologues of the vertebrate tenascins (Baumgartner et al., 1994) and tyrosine phosphorylated proteins (Levine et al., 1994). However, the teneurins turned out to be structurally and functionally distinct from the tenascins despite the high degree of conservation of their epidermal growth factor (EGF)-like repeats (Tucker et al., 2006).

Subsequently, the vertebrate orthologues of ten-a and ten-m were discovered. Mouse

DOC4, the first vertebrate member of the teneurin family, was identified in a screen for proteins that were expressed in response to perturbation of protein folding in the endoplasmic reticulum

(Wang et al., 1998) and since then, several laboratories independently described the ten-a and ten-m/odz homologs in zebrafish (Mieda et al., 1999), chicken (Minet et al., 1999; Rubin et al.,

1999), mouse (Oohashi et al., 1999), rat (Otaki and Firestein, 1999), human (Minet et al., 1999;

Minet and Chiquet-Ehrismann, 2000) and most recently in Caenorhabditis elegans (Drabikowski et al., 2005). In most invertebrates only one teneurin copy has been identified, with the exception

2 of insects where two paralogues have been discovered. However, unlike invertebrates, four teneurin paralogues have been reported in vertebrates.

It has been postulated that the teneurins arose from a single ancestral gene, as comparison of the gene organization among human ten-1, Drosophila ten-a and ten-m and the

Caenorhabditis elegans ten-1 reveals the presence of several conserved intron locations (Minet and Chiquet-Ehrismann, 2000; Tucker et al., 2012). Sequence comparisons of teneurins show that it is not possible to class any of the vertebrate teneurins specifically with either Drosophila ten-a or ten-m. Furthermore, phylogenetic analysis suggests that insects and vertebrates separated before the teneurin gene started to duplicate in each. The Drosophila teneurin ancestor gene therefore, duplicated once to allow for two teneurin paralogues, ten-a and ten-m, whereas the duplication of the teneurins in the vertebrates occurred before the vertebrates radiated into fish, birds and mammals.

Given the independent discoveries of the teneurins, the nomenclature of these homologous proteins in different species is not standardized. The family members were called teneurin in chicken (Minet et al., 1999; Rubin et al., 1999), teneurin or Odz in humans (Minet and Chiquet-Ehrismann, 2000; Minet et al., 1999), neurestin in rat (Otaki and Firestein, 1999), ten-m in zebrafish (Mieda et al., 1999) and ten-m1-4 or odz-1-4 in mouse (Oohashi et al., 1999).

Murine ten-m4 is identical to DOC4 protein, a downstream target of the transcription factor

CHOP (C/EBP homology protein) (Wang et al., 1998), whereas neurestin is most similar to teneurin-2 (Otaki and Firestein, 1999). Taken together, the name teneurin reflects a fusion of the original name, ten-a, and the observation that the common site of expression of the teneurin molecules in vertebrates and invertebrates is in developing neurons. Therefore, given that the

3 studies described in this thesis are restricted to that in the mouse, I will use the term ‘‘teneurins’’ to describe the rodent homologues.

1.2.1 Teneurin Structure

The four teneurin genes encode large proteins that are composed of about 2800 amino acids and possess a distinct domain architecture. The teneurins are type II transmembrane proteins that contain an N-terminal intracellular domain (ICD), a single span transmembrane domain (TM) and a large highly conserved C-terminal extracellular domain (ECD) (Fig. 1.1;

Rubin et al., 1999; Tucker and Chiquet-Ehrismann, 2006). The ICD contains two EF-hand-like motifs, two polyproline motifs and several conserved tyrosines which are predicted to be phosphorylated (Tucker et al., 2007). On the highly conserved extracellular side, there are 8 tenascin-type EGF-like repeats, a region of conserved cysteine residues, and a unique stretch of

26 tyrosine-aspartate (YD)-repeats (Fig. 1.2; Minet and Chiquet-Ehrismann, 2000; Young and

Leamey, 2009).

The large highly conserved 2400 amino acid C-terminal extracellular domain has the tendency to form homodimers or heterodimers with other teneurin family members (Fig. 1.1;

Feng et al., 2002; Oohashi et al., 1999). The second and fifth EGF-like repeat have an odd number of cysteines and it was proposed that the unpaired cysteines may form disulfide bridges with an adjacent teneurin molecule leading to homo- or heterodimer formation (Oohashi et al.,

1999). Following the EGF-like repeats is a region containing 17 cysteine residues that are conserved throughout family members in all species and may be required for correct protein folding (Tucker et al., 2007).

Among eukaryotic proteins, the 26 YD repeats occur only in teneurins. Electron microscopy revealed that this C-terminal half of the protein forms large globular domains that are rich in N-linked glycosylation and are connected to the rod like EGF-like repeats (Feng et al.,

2002). Furthermore, they may also be involved in homophilic and/or heterophilic juxtacrine interactions between teneurins on opposing cells (Oohashi et al., 1999; Rubin et al., 2002). The

YD-repeats are most similar to the repeats encoded by the core of the rearrangement hot spot

(rhs) elements of Escherichia coli, suggesting that the teneurin ancestor is a candidate gene for the source of the rhs core acquired by horizontal gene transfer (Minet and Chiquet-Ehrismann,

2000). Studies have shown that the heavily glycosylated YD-repeats of teneurin-1 can bind to heparin (Minet et al., 1999) further suggesting that the YD-repeats of teneurin-1 may be important for carbohydrate binding.

Figure 1.1. Proposed structural properties of the mouse teneurins. Teneurins are type II transmembrane proteins that can dimerize via the second and fifth EGF repeats. The C-terminus has a globular shape which can be intensively glycosylated. [Adapted from Feng et al., (2002); Permission obtained from American Society for Biochemistry and Molecular Biology].

1.2.2 Processing of the teneurin proteins

Alternative splicing of teneurin transcripts (Tucker et al., 2001) and the presence of several conserved protein cleavage sites (Minet and Chiquet-Ehrismann, 2000) could generate a large variety of different molecular combinations which would significantly enhance the functional complexity of the teneurin family of proteins. Chicken teneurin-2 contains at least three splice variants; a short splice variant composed only of the intracellular domain, transmembrane domain, and the first seven EGF-like repeats, and two long variants that differ from each other only in the presence or absence of eight amino acids between the 7th and 8th

EGF-like repeats (Tucker et al., 2001). In Caenorhabditis elegans, the two promoters driving expression of the sole teneurin gene, ten-1, leads to the formation of two proteins that have distinct expression patterns. The long form, ten-1L, corresponds to the full length protein and is found mainly in neurons, somatic gonad and muscle, whereas the short form ten1-S is mainly expressed in a subset of neurons (Drabikowski et al., 2005).

Teneurins also possess several cleavage sites that are conserved across the metazoans

(Fig. 1.2; Minet and Chiquet-Ehrismann, 2000). Several lines of evidence indicate that teneurins may undergo regulated intramembrane proteolysis (RIP), a lesser known form of proteolytic cleavage whose substrates are transmembrane (Lemberg, 2011). The translocation of the N- terminal domain of teneurin-1 and teneurin-2 to the nucleus suggests that teneurins may be a candidate for RIP (Tucker et al., 2001), similar to that of many other RIP-regulated transmembrane proteins such as Notch (Chan and Jan, 1998) and amyloid precursor protein

(APP) (Haass and Strooper, 1999; Ebinu and Yankner, 2002). The protease responsible for the liberation of the teneurin intracellular domain remains to be identified. However, there is only a limited set of known intramembrane proteases, all of which exhibit specificity for either type-I or

6 type-II transmembrane proteins. Given that teneurins are oriented as type-II transmembrane proteins, possible candidates include site-2 protease or signal peptide peptidase (Weihofen et al.,

2002).

Figure 1.2. Domain organization of vertebrate teneurins. The intracellular domain contains nuclear localization signal (NLS), EF-hand like motifs (EF), prolin-rich stretches (PP) and putative tyrosine phosphorylation sites (Y). Teneurins have a single trans-membrane domain (TM) and a large extracellular domain (ECD). The ECD contains eight EGF-like repeats (green), a region with conserved cysteine residues (yellow) and condensed and YD repeats (blue). Three proteolytic cleavage sites are indicated by arrows. Arrow 1 indicates a cleavage site in or near the transmembrane domain resulting in the release of the intracellular domain. Arrow 2 and 3 mark furin cleavage sites that results in the release of the extracellular domain. [Adapted from Tucker and Chiquet-Ehrismann, (2006); Permission obtained from Elsevier].

In addition to the processing of the intracellular domain, several postulated dibasic cleavage sites are present at the extracellular domain of teneurins. Firstly, a putative furin cleavage site present in all mouse teneurins, both Drosophila ten-m and ten-a, as well as

Caenorhabditis elegans ten-1 is located between the transmembrane domain and the EGF-like repeats (Fig. 1.2; Oohashi et al., 1999; Drabikowski et al., 2005; Tucker and Chiquet-Ehrismann,

2006). This suggests that ectodomain shedding is important for the function of teneurins. Furin is a member of the subtilisin-like proprotein convertase family and cleaves proteins with the consensus dibasic amino acid target sequences, Arg-X-(Arg/Lys)-Arg (Rholam and Fahy, 2009).

Processing of the teneurins at this furin cleavage site has been shown to be functional in vivo

(Rubin et al., 1999) and leads to the release of the extracellular domain from the cell surface.

Interestingly, RIP requires an initial cleavage of the extracellular domain prior to cleavage within the transmembrane region (Young and Leamy, 2009).

Moreover, all teneurins possess a conserved furin cleavage site 100 amino acids from the

C-terminal end (Fig. 1.2; Tucker and Chiquet-Ehrismann, 2006) that could liberate a 40 to 41 amino acid bioactive peptide termed the ‘teneurin C-terminal associated peptide’ (TCAP) (Qian et al., 2004; Wang et al., 2005). Many functionally important cellular peptides, including hormones, neuropeptides, and growth factors, are synthesized as inactive precursor polypeptides, and require post-translational proteolytic processing to become biologically active. Sequence analysis of the last exon of all teneurins, which encodes the TCAP peptides, reveals the presence of numerous conserved potential basic/dibasic cleavage motifs suggesting that release of the

TCAP peptide can be achieved by the action of a relatively small number of proteases that belong to a family of pro-protein convertases.

1.2.3 Teneurin Expression

The main site of teneurin expression, as the name implies, is the developing and adult nervous system and has been studied extensively across the metazoans (Mieda et al., 1999;

Oohashi et al., 1999; Otaki and Firestein, 1999; Tucker et al., 2000 Ben-Zur et al., 2000; Rubin et al., 2002; Zhou et al., 2003). In chicken embryo, teneurin expression in the central nervous system (CNS) is mainly expressed in complementary, non-overlapping populations of neurons that are associated with specific pathways of the visual and olfactory systems (Kenzelmann et al.,

2007; Kenzelmann et al., 2008). In the mouse brain, all four teneurins are mainly expressed in the cortex, thalamus, hippocampus and cerebellum (Zhou et al., 2003). However, although

8 different teneurins are expressed in distinct populations of neurons within these areas, there is some overlapping expression during development and adulthood (Zhou et al., 2003). When examined by in situ hybridization, neocortical expression of the teneurin paralogues is graded rostrocaudally (Li et al., 2006). In all studies investigating the expression of vertebrate teneurins, mRNA expression was observed both in cell bodies and in axons (Tucker et al., 2007), whereas antiserum directed to the C-terminal region of the teneurin protein showed distinct membrane staining (Rubin et al., 1999).

Much of the characterization of teneurin expression has focussed on the CNS, and little is known about the expression and localization of teneurins outside of the CNS. A few studies have shown teneurin expression in non-neuronal tissues, mainly at sites of pattern formation and cell migration (Oohashi et al., 1999; Zhou et al., 2003; Drabikowski et al., 2005; Tucker et al., 2007) further highlighting the importance of teneurins during development. For example, Drosophila ten-m was found in the tracheal system, muscle attachment sites and cardiac cells (Baumgartner and Chiquet-Ehrismann, 1993; Baumgartner et al., 1994). In Caenorhabditis elegans, expression of teneurin-1 outside of the CNS was detected in the somatic gonad precursor cells, pharynx, intestine and sex muscles (Drabikowski et al., 2005). Similarly, mouse teneurin-1 has been observed in the adult testes, smooth muscle cells in lungs and kidney glomeruli (Oohashi et al.,

1999), whereas teneurins-3 and -4 have been detected in the developing limb and adipose tissue

(Ben-Zur et al., 2000). A summary of the reported invertebrate and vertebrate teneurin expression can be found in Table 1.1.

Table 1.1: Teneurin expression in mouse

Caenorhabditis teneurin-1 Somatic gonad, intestines, pharynx, vulva, subset of neurons, elegans gut, some hypodermal and muscle cells1

teneurin-1 developing and adult CNS6,10,14, visual system6,10, smooth

muscle cells in lungs10, kidney glomeruli10, adult testes10

teneurin-2 developing and adult CNS14, visual system6

teneurin-3 developing and adult CNS2,3,4,6,13,14 visual system2,6, spinal

Mouse cord2,14, notochord14, craniofacial mesenchyme2, tongue2,

dermis2, saccule2, developing limb2, periosteum2

teneurin-4 developing and adult CNS2,6,7,14, visual system6, somites14,

spinal cord2, trachea2, nasal epithelium2, saccule2, joints2,

adipose tissue2, tail bud and limbs7

1Drabikowski et al., 2005, 2Ben-Zur et al., 2000, 3Kenzelmann et al., 2008, 4Leamey et al., 2007, 5Leamey et al., 2008, 6Li et al., 2006, 7Lossie et al., 2005, 8Mieda et al., 1999, 9Minet et al., 1999, 10Oohashi et al., 1999, 11Rubin et al., 1999, 12Tucker et al., 2000, 13Tucker et al., 2001, 14Zhou et al., 2003

1.2.4 Teneurin Function

When teneurins were first discovered, their close structural relationship to the tenascins suggested they may play a role in wound healing, nerve regeneration and tumourigenesis

(Chiquet-Ehrismann, 2004; Copertino et al., 1997). However, despite extensive genetic studies and determining the sites of expression, the fundamental role of teneurins in vertebrates remains elusive, partly due to the genetic redundancy that could result in compensation by other family members, and possible splice variants increasing the number of potential protein versions.

However, in Caenorhabditis elegans, the single teneurin-1 gene is required for several aspects of cell migration and morphogenesis. Gene knock-down studies using RNA interference results in a pleiotropic phenotype, including ectopic germline formation, gonad disorganization, germ cell leakage, distal tip cell migration, axonal guidance defects and nerve cord defasciculation (Drabikowski et al., 2005; Trzebiatowska et al., 2008). Studies in Drosophila indicated that ten-m/odz appeared to be the only pair-rule gene that was not a transcription factor

(Baumgartner et al., 1994; Levine et al., 1994). In Drosophila, mutations of the ten-m gene indicate that teneurins are required for proliferation, survival and cellular specification (Kinel-

Tahan et al., 2007). Drosophila ten-m mutants are embryonic lethal due to the fusion of adjacent denticle belts (Baumgartner et al., 1994; Levine et al., 1994). Moreover, late ten-m mutants show defects in ventral nerve cord development, cardiac cells and eye patterning (Levine et al., 1994;

Kinel-Tahan et al., 2007). Similar defects in cuticle formation and eye development have been described for the second Drosophila gene, ten-a (Rakovitsky et al., 2007).

In vertebrates, transfection studies, both in vitro and in vivo, have provided evidence that teneurins can promote neurite outgrowth (Rubin et al., 1999; Leamey et al., 2008), cell adhesion

(Rubin et al., 2002; Leamey et al., 2008) and neuronal pathfinding (Leamey et al., 2007;

Kenzelmann-Broz et al., 2010). In neuroblastoma (Nb2A) cells, overexpression of teneurin-2 led to enhanced neurite elongation, enlarged growth cones, increased filopodia formation and co- localization with actin-containing filopodia (Rubin et al., 1999; Rubin et al., 2002). Similar effects were also observed in chick dorsal root ganglia explants plated on recombinant teneurin-1

YD-repeats (Minet et al., 1999). In mice, a missense mutation that causes an alanine to be replaced with a proline at position 2642 near the globular C-terminus region of teunerin-4 results in delayed gastrulation and neural tube defects prior to embryonic lethality (Lossie et al., 2005).

Knock-out mice lacking teneurin-3 show abnormalities in mapping of ipsilateral projections, and exhibit deficits when performing visually mediated behavioural tasks (Leamey et al., 2007).

However, these deficits were described as mild, suggesting functional redundancy with other teneurins. Interestingly, the intracellular N-terminal domain, once cleaved, can translocate to the nucleus and regulate zic-1 transcriptional activity (Bagutti et al., 2003; Kenzelmann et al., 2008).

However, the events triggering the release of the N-terminal fragment are not known. Moreover, the mechanisms by which these developmental and trophic functions occur remain uncharacterized, but point to a recruitment and re-organization of cytoskeletal elements and cytoskeletal regulatory proteins.

1.2.5 Teneurin interacting proteins and cell signalling

Extracellular matrix proteins, such as the tenascins are typically composed of multiple protein domains, with each individual domain having distinct cellular functions (Orend and

Chiquet-Ehrismann, 2006). Similarly, the complex and unique domain architecture of the teneurins, the ability to form dimers and the evidence of proteolytic processing suggest that different domains of the teneurins could act separately to mediate distinct teneurin functions. The teneurins, along with other extracellular matrix proteins often act as the first sensors of various stimuli from their environment. This initiates a stimuli-dependent transmembrane signalling cascade that is relayed by several effector proteins by mechanisms that include protein modifications, phosphorylation or non-covalent interactions. However, despite the complexity and diversity of multicellular organisms only a limited number of conserved signalling pathways are used repetitively throughout evolution to regulate growth, morphology and development.

In vertebrates, teneurins have a highly conserved and unique intracellular domain with proline-rich stretches, which are characteristic of SH3-binding sites, and two EF-handlike putative Ca2+ binding sites and a number of conserved putative phosphorylation sites. A yeast two-hybrid screen, using part of the teneurin-1 intracellular domain as bait, identified two proteins that could interact with the proline-rich domain (Nunes et al., 2005). One of them, c-

Cb1 associated protein (CAP/ponsin) is a cytoskeleton adapter protein that plays an important role in cell adhesion (Zhang et al., 2006), insulin signalling (Ribon et al., 1998), transcriptional activation and glucose transport (Kioka et al., 2002). This interaction could potentially link the intracellular domain of teneurins to the actin-based cytoskeleton through CAP/ponsin’s interactions with vinculin (Fig. 1.3; Nunes et al., 2005).

The second teneurin-1 N-terminal binding protein identified was the methyl-CpG-binding protein 1 (MBD1), a known transcriptional repressor (Fujita et al., 1999; Wade, 2001).

Interestingly, MBD1 mutant mice display deficiencies in neurogenesis and impaired spatial learning (Zhao et al., 2003). The biological function of teneurin-1 interacting with CAP/ponsin and MBD1 has not been fully resolved but suggests that teneurin-1 may be required for linking the actin cytoskeleton to transcriptional machinery. Similarly, the intracellular domain of teneurin-2 could be detected in the nuclei of fibrosarcoma cell line HT1080 cells in discrete spots that often co-localize with the endogenous tumor suppressor transcription factor promyelocytic leukemia protein (PML) (Bagutti et al., 2003). The teneurin-2 intracellular domain has a nuclear function in repressing odd-paired-a (opa) activity in Drosophila (Baumgartner et al., 1994) and the corresponding vertebrate homologue zinc finger protein (zic-1) in a similar signalling cascade (Bagutti et al., 2003). Zic-1 and opa are both implicated in mediating neuronal differentiation, cellular morphology and development (Benedyk et al., 1994; Aruga et al., 1996).

RIP has been suggested as the mechanism by which the intracellular domain is released to influence transcription.

The domain architecture of the teneurin extracellular domain and the high number of conserved dibasic cleavage motifs suggests that the extracellular fragment could also act as a ligand. Previous studies have postulated functional interaction with the extracellular matrix

(Tucker et al., 2001; Trzebiatowska et al., 2008). Immunolabelling with an antiserum against the extracellular domain of teneurin-2 co-localizes with the laminin staining in certain chicken basement membranes (Tucker et al., 2001) suggesting that the shed extracellular domain may bind to the surrounding extracellular matrix. Moreover, recent studies have demonstrated a genetic interaction between Caenorhabditis elegans ten-1 and ina-1 (α-integrin), dgn-1

(dystroglycan) and epi-1 (laminin α-chain), as well as functional redundancy at the basement membrane in gonadal development and maintenance of basement membrane integrity

(Drabikowski et al., 2005; Trzebiatowska et al., 2008). Many extracellular matrix proteins at the basement membrane are heavily glycosylated, and the YD-repeats on the teneurin extracellular domain are also predicted to be glycosylated. Furthermore, recombinantly expressed YD-repeats have been shown to bind the glycosaminoglycan heparin (Minet et al., 1999), suggesting that the teneurins could interact with the carbohydrate moieties in the extracellular matrix. Such a signaling mechanism would allow the teneurins to act as extracellular adhesion molecules and anchor proteins, but also as signaling molecules themselves.

Some studies have also implicated a functional relationship between the teneurins and the integrins. Integrins act to add mechanical stability to a cell, and like the teneurins, have diverse roles in cell adhesion, migration, cell signaling and differentiation (Wickström and Fässler,

2011). Integrins can bind to a number of ligands that possess an arginine-glycine-aspartic acid

14 motif (RGD), such as fibronectin and vitronectin (Ruoslahti, 1996). In Drosophila, the teneurins possess an RGD motif, 72 amino acids from the C-terminus, just upstream from the putative

Drosophila TCAP peptide (Graner et al., 1998). However this site is not conserved among vertebrate teneurins and is not found in any of the mouse teneurin sequences (Lovejoy et al.,

2006) suggesting that some functional differences exist between Drosophila and vertebrate teneurins.

Figure 1.3. Current teneurin signalling model. Teneurin is depicted as a dimer linked through the EGF-like repeats two and five. The possible release of a C-terminal neuropeptide (TCAP) is indicated. In the cell on the left teneurin is anchored to the cytoskeleton through its interaction with CAP/ponsin, which in turn is linked to actin through vinculin. Upon homophilic binding to teneurin on the cell to the right, the intracellular domain is cleaved and translocates to the nucleus where it may interact with transcriptional regulators such as MBD1 or zic [Adapted from Tucker and Chiquet-Ehrismann, (2006); Permission obtained from Elsevier].

Although there is considerable evidence of teneurin dimerization, extensive processing and release of the extracellular domain and the presence of a cleavable bioactive peptide TCAP, at the tip of the C-terminus, little is known about the receptor or ligands that they interact with or

15 the corresponding signal transduction mechanisms that they may induce. In fact, the current model offers little explanation for the number of biological functions attributed to the teneurins

(Fig. 1.3; Tucker and Chiquet-Ehrismann, 2006). Nevertheless, the unique domain architecture of the teneurin proteins and the combination of alternative splicing of teneurin transcripts

(Tucker et al., 2001) allows for multiple binding partners and a large variety of different molecular combinations that would significantly enhance the functional capability and complexity of the teneurins.

1.3 Dystroglycans

The current body of evidence regarding the teneurins points to a fundamental role in mediating cytoskeletal dynamics through a functional interaction with the extracellular matrix and basement membrane proteins (Tucker et al., 2001; Drabikowski et al., 2005; Trzebiatowska et al., 2008). The interaction between a cell and its surroundings is regulated by adhesion molecules at the plasma membrane which allow the extracellular matrix to communicate with the intracellular cytoskeleton. One such adhesion molecule that is of considerable interest because of its functional redundancy to the Caenorhabditis elegans teneurin-1 is dystroglycan

(Trzebiatowska et al., 2008). The Caenorhabditis elegans genome contains three dystroglycan related genes: dgn-1, dgn-2 and dgn-3 (Johnson et al., 2006), whereas vertebrates possess a single dystroglycan (Holt et al., 2000).

Dystroglycan was first discovered in brain tissue as a laminin-binding membrane protein termed cranin (Smalheiser and Schwartz, 1987). It was later discovered to be identical to the dystroglycans identified in skeletal muscles and further characterized as a component of a large multimeric protein assembly, the dystrophin–glycoprotein complex (Smalheiser and Schwartz,

1987; Ervasti and Campbell, 1991; Smalheiser and Kim, 1995). The distribution of dystroglycan in adult mouse brain appears to be restricted to neurons of the cerebral cortex, hippocampus, olfactory bulb, basal ganglia, thalamus, hypothalamus, brainstem and cerebellum (Zaccaria et al.,

2001). However, dystroglycan is also distributed throughout the basement membranes of the whole body and was found in most tissues and organs examined in rodents (Durbeej et al., 1998), primates (Royuela et al., 2003) and teleost fish (Guyon et al., 2003). Although the dystroglycans were first discovered in the brain, most research so far has focused on the functions of dystroglycan in muscle and neuromuscular junctions, while its role in the brain and other organs, such as the testis has largely been neglected.

The protein matrices that form the extracellular matrix and basement membranes are critical for cell and tissue viability and the dystroglycans provide mechanical stability to cells by linking the extracellular matrix on the extracellular surface with the actin cytoskeleton on the intracellular side (Chen et al., 2003). Apart from its central involvement in a variety of muscular dystrophies, recent findings suggest that dystroglycan is important in brain development, synapse formation, neuronal plasticity, nerve-glia interactions and maintenance of the blood-brain barrier and basement membrane integrity (Peng et al., 1999; Jacobson et al., 2001; Zaccaria et al., 2001;

Saito et al., 2003; Nico et al., 2003). Moreover, when dystroglycan is overexpressed in fibroblasts, it results in an increase in actin protrusions such as microvilli and filopodia

(Higginson and Winder, 2005). It is therefore, likely, that dystroglycan simply acts as part of the foundation or scaffold necessary for the organization of other adhesion proteins and signalling molecules, both inside and outside of the cell.

1.3.1 Dystroglycan Structure

The dystroglycan gene, DAG1, encodes a single 97-kDa polypeptide that is post- translationally cleaved at serine-654 to yield two glycoproteins, α-dystroglycan and β- dystroglycan (Ibraghimov-Beskrovnaya et al., 1993; Holt et al., 2000; Jayasinha et al., 2003).

Figure 1.4. A diagrammatic representation of α- and β-dystroglycan at the cell surface. The extracellular α-dystroglycan (α) and transmembrane β-dystroglycan (β) are shown in blue. Agrin, perlecan and laminin isoforms bind to the carbohydrate moieties of the α-dystroglycan mucin- like region via their laminin G domain (LG) modules. Rapsyn associates with membrane- proximal residues in the β-dystroglycan cytoplasmic tail, and GRB2 binds via a SH3-dependent interaction to one of the several potential SH3 binding motifs in the β-dystroglycan C-terminus. Dystrophin or utrophin bind the C-terminal of β-dystroglycan via the WW domain motif PPxY; the utrophin and dystrophin N-termini bind to F-actin. [Adapted from Winder et al., (2001); Permission obtained from Elsevier].

The α- and β-subunits are extensively glycosylated and are non-covalently bound to form a type-

1 heterodimeric transmembrane protein (Fig. 1.4; Winder et al., 2001). α-Dystroglycan is a

18 dumb-bell-shaped peripheral membrane protein that contains three N-linked glycosylation sites and several O-linked glycosylation sites (Winder, 2001). Depending on the extent of glycosylation and glycosaminoglycans, the molecular mass of dystroglycan can range from around 156 kDa in skeletal muscle to 140 kDa in cardiac muscle and 120 kDa in brain and peripheral nerve tissue (Muntoni et al., 2004).

The C-terminal globular domain of α-dystroglycan is non-covalently associated with the

N-terminus of β-dystroglycan (β-DG) (Ervasti and Campbell 1993; Jung et al., 1995). The 43kD

β-DG subunit contains a single N-linked glycosylation site, a single-pass transmembrane domain and a proline rich cytoplasmic tail (Ibraghimov-Beskrovnaya et al., 1993). The dystroglycan complex is, therefore, anchored in the cell membrane by the membrane-spanning β-dystroglycan subunit that functionally links the intracellular region of the cell to the extracellular partners via

α-dystroglycan. The addition of carbohydrate moieties and complex sugar modifications are implicated in regulating dystroglycan binding partners, signal transduction and function.

1.3.2 Dystroglycan interacting proteins and cell signalling

Dystroglycan contains binding sites for several known extracellular proteins and cytoplasmic proteins. The α-subunit of dystroglycan is a receptor for laminin and other proteins with laminin G-like domains such as perlecan (Peng et al., 1998), agrin (Gee et al., 1994), and neurexin (Sugita et al., 2001) (Fig. 1.4). These interactions further emphasize a role for dystroglycan in development and maintenance of basement membrane integrity in cells and tissue. Laminin and perlecan are extracellular matrix proteins that are ubiquitously expressed in basal lamina, structures that provide support for cell attachment, differentiation, migration, and survival (Colognato and Yurchenco, 2000). The accumulation and assembly of laminin and

19 perlecan on the cell surface and basement membranes is a dynamic process that is mediated by dystroglycan (Henry and Campbell, 1998; Henry et al., 2001). Agrin is an extracellular matrix protein that mediates the clustering of acetylcholine receptor (AChR) at the neuromuscular junction (Grady et al., 2000), whereas neurexins are neuronal proteins which are concentrated at the postsynaptic densities of excitatory synapses and function in cell-cell interaction (Sugita et al., 2001).

The long β-dystroglycan C-terminal cytoplasmic tail binds to cytoskeletal-associated proteins utrophin and dystrophin in the cytoplasm (Matsumura et al., 1992) and even directly to actin (Fig. 1.5; Chen et al., 2003). However, the ability of dystroglycan to spatially organize other molecules might also reflect a signal transduction capacity suggesting that dystroglycan has a more active function in the transduction of a variety of signalling pathways through its β- dystroglycan subunit. The cytoplasmic tail of β-dystroglycan contains several proline-rich regions and tyrosine phosphorylation consensus sequences (Ibraghimov-Beskrovnaya et al.,

1992) similar to that reported in teneurins and which can interact with the Src homology 2 (SH2) and Src homology 3 (SH3) domains of signaling proteins. One such example is the interaction of the β-dystroglycan cytoplasmic region with the SH3 domains of the growth factor receptor- bound protein 2 (Grb2), an adaptor protein involved in signal transduction and cytoskeletal organization (Fig. 1.5; Yang et al., 1995). Via Grb2, dystroglycan can interact with the C– terminal domain of focal adhesion kinase (FAK) (Cavaldesi et al., 1999). Although the dystroglycans functionally interact with the integrins through binding to laminin, the N–terminal domain of FAK also binds to intracellular portions of integrins, which could further enhance the functional connection between dystroglycan and the integrin system. In addition, β-Dystroglycan serves as a scaffold for the mitogen-activated protein (MAP) kinase 2 (MEK2) and extracellular

20 signal-regulated kinase (ERK) signaling cascade (Fig. 1.5; Spence et al., 2004). The MEK and

ERK kinases form part of an important signalling cascade, the receptor tyrosine kinase/Ras/Raf/MAP kinase pathway that are involved in a vast number of cellular processes, including growth, morphology and synaptic plasticity. Furthermore, β-dystroglycan binds the cytoskeletal adaptor protein ezrin, and modulates actin reorganization in fibroblasts via cell division control protein 42 homolog (Cdc42), a small GTPase of the Rho-subfamily (Spence et al., 2004).

Figure 1.5. Current model of the the laminin-dystroglycan-dystrophin signaling cascade. The β-dystroglycan tail serves as a scaffold for a number of kinases including MEK1/2, ERK1/2, GRB2 and JNK. In addition, β-dystroglycan binds to utrophin to mediate actin polymerization.

Adding further to the complexity of dystroglycan binding proteins and signalling pathways is the interaction of dynamin (Zhan et al., 2005) and caveolin-3 (Sotgia et al., 2000) to the cytoplasmic C–terminal tail of β-dystroglycan. Dynamin is a GTPase involved in

21 endocytosis, recycling of presynaptic vesicles and receptor internalization in all cell types including neurons. On the other hand, caveolin-3, which acts as scaffolding proteins in the formation of caveolae during caveolae-mediated endocytosis, competes with dystrophin for binding to the β-dystroglycan tail (Sotgia et al., 2000).

The complex structural arrangement of dystroglycan and its association with numerous binding proteins and signaling molecules on both the extracellular and the intracellular side confirms its versatile role in many cellular functions. It also suggests that dystroglycan may be a receptor for a multitude of ligands that could recruit and induce a unique combination of the dystroglycan-associated signalling pathways. The characterization of these proteins interacting with dystroglycan, as well as the elucidation of potential ligands will give insight into the different and distinct functions attributed to the dystroglycan complex.

1.4 Developmental proteins in the mammalian testes

In most cells and tissues, developmental and extracellular matrix proteins provide structural support and interact with cytoskeletal proteins to maintain normal epithelial morphology, organization and function. The role of these proteins, therefore, has been of considerable interest in mediating testicular function (Siu and Cheng, 2004; Lie et al., 2010). In the testis, the seminiferous tubule is the functional unit that carries out spermatogenesis (de

Kretser and Kerr, 1988), whereas steroidogenesis occurs at the Leydig cells (Steinberger, 1971).

Spermatogenesis is a continuous, energy-demanding and precisely regulated process by which stem cells called spermatogonia divide and differentiate via mitosis and meiosis into spermatocytes and spermatids during the seminiferous epithelial cycle (Lie et al., 2010). During spermatogenesis, the germ cells migrate towards the tubule lumen. Spermatids, which are located

22 closer to the tubule lumen, undergo extensive morphological changes that include chromosomal condensation, formation of the acrosome, tail and residual body in a process defined as spermiogenesis (Lie et al., 2010). At the completion of spermiogenesis, elongated spermatids line the tubule lumen, awaiting release from the seminiferous epithelium at spermiation. These processes are not only mediated by neuroendocrine signaling (Steinberger, 1971; McLachlan et al., 1995), but are highly dependent on the precise control of testicular cell-cell interactions, growth factors and the integration of autocrine and paracrine signaling pathways in the seminiferous tubules (Siu and Cheng, 2004). The role of the cytoskeleton in mediating germ cell shape, size, differentiation and migration during spermatogenesis has been well characterized

(Russell et al., 1989; Lie et al., 2010) but less is known about the role of the extracellular matrix proteins that lie upstream of the cytoskeleton.

Each seminiferous tubule is about 1 m in length and 0.5 mm in diameter and undergoes extensive restructuring and changes in size and morphology during spermatogenesis (Cheng and

Mruk, 2002) so as to accommodate the multitude of cellular events occurring within the seminiferous tubule (Wing and Christensen, 1982). These changes are largely integrated at the basement membrane of the tunica propria, a modified form of extracellular matrix that is constituted largely by type-IV collagen, laminin, integrin and heparan sulfate proteoglycan (Dym

1994; Siu and Cheng, 2008). The basement membrane separates the interstitial cells from the germ cells and Sertoli cells within the seminiferous tubule, and is thus in a unique position to mediate communication between the interstitial and seminiferous tubule compartments. Sertoli cells and spermatogonia are in close physical contact with the basement membrane and rely on it for structural and hormonal support throughout the different stages of spermatogenesis (Mruk and Cheng, 2004; Siu and Cheng, 2008). In fact, infertile patients with aspermatogenesis were

23 shown to have abnormal basement membrane structures (Lustig et al., 2000), thus highlighting the importance of maintaining basement membrane integrity. Of particular interest is the integrin–laminin complex that functions in concert with other extracellular matrix proteins to facilitate germ cell movement and maintain structural integrity of the basement membrane in the seminiferous tubule (Siu and Cheng, 2004b; Siu and Cheng, 2008). In addition, studies by

Trzebiatowska et al. (2008) found that teneurin-1 and dystroglycan function redundantly in maintaining basement membrane integrity in the gonad of Caenorhabditis elegans. It is likely that teneurin-1 and dystroglycan may also possess a functional role in the mammalian testis.

The extracellular matrix at the basement membrane also provides a functional link between the interstitium and cells within the seminiferous tubule (Dym, 1994; Siu and Cheng,

2004), suggesting that they may also play a role in mediating steroidogenesis. Spermatogenesis and male fertility are heavily dependent upon the presence of testosterone that is secreted by luteinizing hormone-stimulated Leydig cells in the interstitium (McLachlan et al., 1995). The absence of the androgen receptor that it targets in the Sertoli cells, results in spermatogenic arrest at the meiosis stage (Haywood et al., 2003; Chang et al., 2004). Sertoli cells are the major cellular target for the testosterone signaling that is required to support male germ cell development and survival (Griswold, 1998). When testosterone diffuses across the plasma membrane and binds to the androgen receptor, the androgen receptor undergoes a conformational change that allows it to be translocated to the nucleus where is binds to specific DNA sequences called androgen response elements (Walker, 2011). This facilitates the recruitment of co- regulator proteins and regulates transcription of target genes related to tubular restructuring, cell junction dynamics and the cytoskeleton (Verhoeven et al., 2010). This main pathway by which androgens exert their activities is referred to as the classical genomic pathway.

However, non-classical signalling pathways have been identified by which androgens such as testosterone may affect the function of target cells without a need for direct effects on gene expression (Silva et al., 2002). Two such pathways have been so far identified, testosterone- mediated [Ca2+] influx pathway (Gorczynska and Handelsman, 1995) and the activation of the

MAP kinase cascade (Cheng et al., 2007). The recruitment of the MAPK-ERK pathway has been shown to induce phosphorylation of the ribosomoal S6 kinase (p90RSK) which, in turn, phosphorylates cyclic adenosine 3',5'-monophosphate response element binding protein (CREB) at serine-133 (Cheng et al., 2007) and result in activation of CREB-regulated genes such as lactate dehydrogenase A (LDH-A) and early growth response 1 (Egr1) (Fix et al., 2004). In addition, peritubular myoid cells, situated near the basement membrane, also express androgen receptors and could mediate androgen-dependent functioning in Sertoli cells and spermatogonia which are in close physical contact to the basement membrane (Walker, 2011). Interestingly, in vitro studies indicate that testosterone can act via the activation of MEK-ERK kinases to facilitate Sertoli-germ cell attachment (Cheng et al., 2007) and is thus likely to have a similar function in vivo at the basement membrane.

Cytokines and growth factors such as tumor necrosis factor-α (TNFα; Siu et al., 2003) and insulin-like growth factor 1 (IGF-1; Saez et al., 1989) also affect both spermatogenesis and steroidogenesis and are likely to functionally interact with the extracellular matrix at the basement membrane. The relationship of these extracellular matrix proteins as well as the ligands they associate with are, therefore, pertinent to understanding how developmental proteins, trophic factors and sex steroids integrate to mediate testicular function.

1.5 Regulation of Cytoskeletal dynamics

The cellular cytoskeleton is a vital and dynamic scaffolding component of all eukaryotic cells that is responsible for a wide range of functions including cell division, cellular motility, organelle transport, membrane trafficking, maintenance of cellular integrity and shape, establishing cell polarity and regulating cellular extensions. The cytoskeleton is composed of three distinct polymers: actin, microtubules and intermediate filaments. These cytoskeletal elements are distinguished by the architecture and function of the networks they form, their mechanical stiffness and the binding proteins with which they associate. However, despite their structural differences, the three cytoskeletal elements often have overlapping functions in controlling the shape and mechanics of eukaryotic cells. Cytoskeletal polymer dynamics, particularly for actin and tubulin, are energetically expensive yet evolutionarily conserved, suggesting important biological roles.

1.5.1 Actin cytoskeleton

Actin is the most abundant cellular protein found in eukaryotes and is comprised by a highly conserved family of proteins; α-actin, β-actin and γ-actin. In non-muscle cells such as neurons, actin plays a variety of myosin-independent roles such as maintaining cellular architecture and mediating receptor-mediated responses of the cell to external signals. In fact, the actin cytoskeleton is mainly controlled by the Rho family of small GTP-binding proteins and is continually assembled and disassembled in response to the local activity of signalling systems.

The actin monomer is a 43 kDa protein with a single nucleotide binding site for either ATP or

ADP (Steinmetz et al., 1997). In its monomeric form actin is referred to as globular actin or G- actin, whereas in its polymerized form, actin is referred to as filamentous actin or F-actin.

The conversion of soluble G-actin into F-actin is a highly dynamic and reversible process regulated by ATP binding and hydrolysis and actin-binding proteins (Kabsch and

Vandekerckhove, 1992). Polymerization is first initiated by a process referred to as nucleation and is often induced by the actin-related proteins 2 and 3 (Arp2/3) complex (Wear et al., 2000;

Rácz and Weinberg, 2008). Upon activation, Arp2/3 binds existing actin, nucleating them and stimulating actin polymerization into a branched network of actin ﬁlaments (Goley and Welch,

2006). Actin filaments have an inherent polarity, with one end referred to as the plus or ‘barbed’ end and the other end as the minus or ‘pointed’ end. ATP-bound G-actin has a higher affinity than ADP-bound G-actin for the barbed ends of actin filaments, and thus nucleotide exchange of

ADP for ATP on G-actin promotes the monomeric addition to the barbed end (Wear et al., 2000;

Steinmetz et al., 1997). Actin monomers within the actin filament have an intrinsic ATPase activity that slowly hydrolyzes ATP-bound actin to ADP-bound actin. ADP-bound actin monomers depolymerize or dissociate from the actin filament, from the pointed end. This dynamic process of polymerization and depolymerization is referred to as treadmilling (Kabsch and Vandekerckhove, 1992).

Remodeling of the cytoskeleton is central to the modulation of cell shape and migration and the dynamic regulation and stabilization of elaborated actin-based cytoskeletal structures is regulated by a multitude of actin-binding proteins such as cofilin, α-actinin, ezrin, elongation factor-1, dystrophin, utrophin and filamin A (dos Remedios et al., 2003). The bundles and networks of actin filaments created by these cross-linking proteins are highly stable and resistant to rapid polymerization and depolymerization (Puius et al., 1998). Of particular interest is filamin A, a 280-kD non-muscle actin-binding phosphoprotein that cross-links actin filaments into orthogonal networks. Filamin A is phosphorylated at multiple sites by several protein

27 kinases, such as protein kinase A (PKA; Jay et al., 2004), cyclin-dependent kinase 1 (Cdk1;

Cukier et al., 2007) and p90RSK (Woo et al., 2004) and plays a key role in inducing filopodia formation (Ohta et al., 1999). In addition to cross-linking F-actin, filamin A can interact with integrins, transmembrane receptor complexes, and second messengers through its C-terminal region and recruit signaling proteins to the vicinity of sites of actin polymerization and remodeling (Stossel et al., 2001; Nakamura et al., 2011). Moreover, the association of filamin with the Rho-GEF (Guanine nucleotide exchange factors) proteins (Bellanger et al., 2000) and the RSK family (Woo et al., 2004) supports an involvement of filamin A as a docking site for signaling molecules This suggests that filamin A could serve a potential link between transmembrane proteins and the actin cytoskeleton.

1.5.2 Microtubule cytoskeleton

Microtubules are the stiffest of the three polymers and serve as the main structural components within cells (Fletcher and Mullins, 2010). Microtubules important for a wide array of functions such as neurite outgrowth, growth cone stabilization, cell migration, vesicular trafficking, cell polarity, flagella movement, and cell division. The microtubule networks are assembled from α-tubulin and β-tubulin heterodimers and have a more complex assembly and disassembly dynamics as compared to the actin cytoskeleton. However, similar to actin polymerization, microtubule assembly occurs in two phases, nucleation and elongation.

Microtubules are nucleated at the Microtubule Organizing Center (MTOC), which is often located in the perinuclear regions of the cell (Desai and Mitchison, 1997). Contained within the

MTOC is another type of tubulin, γ-tubulin, which is distinct from the α-tubulin and β-tubulin.

The γ-tubulin combines with several other associated proteins to form a circular structure known

28 as the "γ-tubulin ring complex" (γ-TuRC) which acts as a scaffold for α- and β-tubulin dimers to begin polymerization (Nogales, 1999).

Polymerization dynamics allow microtubules to adopt spatial arrangements that can change rapidly in response to cellular needs. Tubulin heterodimers polymerize end-to-end to form linear protofilaments that then bundle into hollow cylindrical microtubule filaments. The protofilaments arrange themselves in an imperfect helix with one turn of the helix containing 13 tubulin dimers each from a different protofilament (Desai and Mitchison, 1997). The protofilaments are also polarized into a plus and minus end. The minus end is anchored at the

MTOC and is considered relatively static as compared to the rapidly growing or shrinking plus end, which frequently projects to the cell's periphery. Within the microtubule, the tubulin heterodimers are oriented in such a way that the microtubule plus end is capped by β-tubulin subunits, while the minus end is capped by α-tubulin (Desai and Mitchison, 1997). Like actin, tubulin polymerization is also dependent on nucleotide hydrolysis. GTP-bound tubulin adds to the MTOC, and during polymerization, the GTP bound to β-tubulin is hydrolyzed to GDP and does not exchange while β-tubulin remains in the polymer (David-Pfeuty et al., 1977, MacNeal and Purich, 1978). However, GTP bound to α-tubulin remains stable and is not hydrolyzed during polymerization (Spiegelman et al., 1977). When hydrolysis catches up to the entire tip of the microtubule, a rapid depolymerization and shrinkage of the microtubules occurs, in a process known as ‘catastrophe’. Upon depolymerisation, the released β-tubulin subunits can exchange

GDP for GTP and undergo another round of polymerization (Mitchison, 1993). Since the GTP- bound tubulin lattice is more stable, capping proteins are recruited to shield the protofilaments from hydrolysis and subsequent catastrophe (Cassimeris and Spittle, 2001).

The highly dynamic nature of microtubules allows them to mediate an array of cellular functions that require the cytoskeleton to have complex high order architectures. This dynamic nature is controlled by numerous microtubule-associated proteins (MAPs). One of the major

MAPs that regulate microtubule dynamics is an 18kDa ubiquitously expressed cytosolic phosphoprotein known as stathmin or oncoprotein-18 (Hailat et al., 1990; Cassimeris, 2002).

When unphosphorylated, stathmin promotes microtubule catastrophe in two ways. First, stathmin can reduce the microtubule polymer mass through direct binding and sequestration of soluble tubulin in a ratio of 1 stathmin to 2 tubulin dimers, also called a T2S complex (Howell et al.,

1999). Alternatively, stathmin can increase the catastrophe frequency by binding directly to the ends of polymerized microtubules (Belmont and Mitchison, 1996). Interestingly, the catastrophe frequency is considerably greater at minus ends than at plus ends (Manna et al., 2006).

Stathmin microtubule destabilizing activity is attenuated through serine phosphorylation at four regulatory sites, serine-16, -25, -38 and -63, and is correlated with the action of multiple extracellular stimuli including trophic factors (Curmi et al., 1999). Stathmin phosphorylation can be regulated by several classes of kinases, such as the MAP kinase/ERK family, cAMP- dependent protein kinase, cyclin-dependent kinase 1 (cdk1) and c-Jun N-terminal kinase (JNK) that target specific serine residues (Sobel, 1991; Manna et. al., 2009; Ng et al., 2010). Hyper- phosphorylation of stathmin at all four regulatory sites has shown to drive cell division (Larsson et al., 1997). On the other hand, phosphorylation at serine-38 is typically associated with stress- induced JNK activation, whereas phosphorylation at serine-25 is mainly associated with ERK- dependent growth and differentiation (Doye et al., 1990; Beretta et al., 1993; Di Paolo et al.,

1996).

1.5.3 Cytoskeletal dependent cellular protrusions: Lamellipodia and filopodia formation

The interaction between the actin and microtubule cytoskeletons has emerged as a fundamental process required for spatial regulation of cell protrusion, retraction, growth cone guidance and dendritic remodelling activities (Rodriguez et al., 2003). The mechanisms governing these interactions are not well known but have been shown to be mediated by common cytoskeletal binding proteins and signal transduction pathways (Condeelis et al., 1995;

Chesarone et al., 2010). Two major types of structures dominate actin formations in neurons and cells at the leading edge, branched networks of unbundled filaments that form lamellipodia and unbranched filaments that are bundled together to form filopodia. The formation, movement and integrity of these structures are also dependent on microtubule dynamics that act, in part, to re- organize and anchor the actin cytoskeleton in these structures (Fig. 1.6; Schober et al., 2007;

Lowery and Vactor, 2009).

Lamellipodia are actin protrusions with branched, sheet-like structures located at the leading edge of the cell (Small et al., 2002). The branched actin structures of lamellipodia are the result of actin nucleation at the plasma membrane in response to the Arp2/3 complex, which binds pre-existing actin filaments and creates nucleation cores. This facilitates the polymerization of new actin filaments, which is then followed by barbed-end capping (Welch et al., 1997; Mejillano et al., 2004). The protrusive action of lamellipodia is generated by actin treadmilling (Wang, 1985) which is enhanced by actin-binding proteins such as cofilin and profilin (Didry et al., 1998). These processes are mediated by the activation of the Rho-family

GTPases, particularly, Rac and Cdc42 (Small et al., 2002). Activation of Cdc42 recruits the

Wiskott-Aldrich syndrome protein (WASp) family receptors and in conjuction with Rac1 signalling, activates the Arp2/3 complex (Small et al., 2002).

Protruding from the lamellipodia actin network, are finger-like F-actin-rich protrusive structures, known as filpodia. When filopodia were first discovered they were described as

“extremely fine filamentous pseudopodia” at surface of neuronal growth cones (Harrison, 1910).

However, they have since been observed on all types of migratory cells. The rodlike characteristic of filopodia extensions are approximately 0.1–0.2μm in diameter and are composed of unbranched bundled actin filaments oriented with their fast-growing filament “plus ends” toward the tip (Small et al., 1978; Nemethova et al., 2008).

Figure 1.6. Cytoskeletal elements and the formation of lamellipodia and filopodia. The leading edge of neurons or cells consists of dynamic, finger-like filopodia that are separated by sheets of membrane between the filopodia called lamellipodia-like veils. Long bundled actin filaments (F-actin bundles) form the filopodia whereas mesh-like branched F-actin networks give structure to lamellipodia. Additionally, individual dynamic microtubules explore the region along F-actin and along with the stable microtubule bundles help to determine the shape and movement of the neuron or cell. [Adapted from Lowery and Vactor, (2009); Permission obtained from Nature Publishing Group].

Filopodia and growth cones can serve as sensory structures and are important for correct navigation of outgrowing axons during development (Tessier-Lavigne and Goodman, 1996).

Certain filaments within the branched network generated by the Arp2/3 complex are selected for elongation due to the binding of a ‘tip complex’ (Johnston et al., 2008). The Ena/VASP

(vasodilator-stimulated phosphoprotein) proteins are found at the leading edge of lamellipodia and at the tips of filopodia and promote the spatially regulated actin polymerization. These actin bundles are then grouped together by actin cross-linking proteins such as filamin A and along with the microtubule cytoskeleton, help to support filopodia formation and architecture (Ohta et al., 1999; Schober et al., 2007; Lowery and Vactor, 2009).

The induction of filopodia formation through a recruitment of the Arp2/3 complex and the Ena/VASP proteins is also preceded by the activation of the Rho family of small Ras-related

GTPases (Ohta et al., 1999). The activation of Rac and Cdc42 can be mediated by stimulation of both growth factor (Hall, 1998) and integrin (Price et al., 1998) receptors and requires the GDP–

GTP exchange factors, GEFs (Small et al., 2002). In addition, the activation of Rac1, Cdc42, and

RhoA can lead to the activation of the MEK-ERK signalling pathway via Ras (Brakebusch et al.,

2002) and subsequent recruitment of ERK-mediated cytoskeletal regulatory proteins. Together, this suggests that the filopodia are intimately associated with lamellipodia. However, recent evidence has shown that filopodia formation can occur independently of lamellipodia and the

Arp2/3 complex in mammalian cells (Steffen et al., 2006). However, the mechanism by which this occurs remains unresolved.

1.6 Teneurin C-terminal Associated Peptide

The TCAP family of peptides was first discovered in a search for novel corticotropin- releasing factor (CRF) homologues (Qian et al., 2004). Since then, four TCAPs have been identified in vertebrates, annotated as TCAP-1,-2,-3 and 4, based on their location on the

33 terminal exon of the four teneurin genes. The four TCAPs are the same size as CRF but possess less than 20% sequence similarity to the CRF family of peptides (Lovejoy et al., 2006) and differ markedly in structure (Tan et al., 2011b). However, the TCAP family is highly conserved across the metazoans. The TCAP sequences possess 73–88% sequence identity among its four human paralogues and 71–87% among mouse paralogues relative to TCAP-1. Mouse and rat TCAPs are identical whereas the identity between human and mouse orthologues is 95–100%. There is only one amino acid difference between mouse and human TCAP-1. At position 5, a glycine residue in mouse TCAP-1 has been substituted with a serine in human TCAP-1. Human TCAP-2 and

TCAP-3 are identical to mouse TCAP-2 and TCAP-3 respectively. For TCAP-4, there are two amino acid differences. In mouse TCAP-4, asparagine in position 5 and threonine in position 18 has been replaced with serine and isoleucine respectively in human TCAP-4. Furthermore, the

TCAP family of peptides are amidated (GKR) at the C-terminal end, possess a pyroglutamyl residue in position 1 and cleavage sites, all of which are structural hallmarks of a cleavable bioactive peptide (Qian et. al., 2004). Although the evolutionary origin of the TCAP region of the teneurins has not been established, the TCAP family of peptides are phylogenetically old, and such high level of conservation may be indicative of functional constraints, suggesting that they are important in the development and survivability of a variety of species (Lovejoy et al., 2006).

In the genome, TCAPs are annotated as being part of the extracellular domain of teneurin but previous studies indicate that some members of the family may possess functions that are independent of the larger teneurins. In the rodent brain, teneurin-1 (Zhou et al., 2003) and

TCAP-1 (Wang et al., 2005) mRNA expression appear to be distinct in some regions such as in the limbic areas, but overlap in others areas such as the olfactory bulb and cerebellum, suggesting that parts of the teneurin gene could be differentially regulated. In fact, northern blot

34 studies in the adult brain and embryonic hypothalamic cell culture, demonstrated that only

TCAP-1 and -3 can be independently synthesized from the larger teneurins, whereas TCAP-2 and -4 are synthesized as part of the full-length teneurin (Al Chawaf, 2008). It is likely that the four TCAPs are regulated differently during development as is the case for the teneurins (Li et al., 2006). This would indicate that in some cases TCAP-1 may be functionally independently from teneurin-1.

A number of biological functions both in vitro and in vivo have been attributed to the

TCAP family of peptides. The first studies conducted by Qian et al., (2004) found that in immortalized Gn11 mouse neurons, TCAP-3 treatment had a dose-dependent effect on cAMP levels, teneurin-1 gene expression and cell proliferation as measured by an MTT assay.

Subsequent studies by Wang et al., (2005) and Al Chawaf et al., (2007a) observed similar in vitro effects of TCAP-1 in immortalized N38 hypothalamic mouse cells. Further studies with

TCAP-1 showed numerous in vitro effects in neuronal cell lines. TCAP-1 was neuroprotective in hypothalamic neurons subjected to alkalotic and oxygen free radical stress by inhibiting caspase-

3 cleavage and necrotic cell pathways, increasing superoxide dismutase (SOD)-1, SOD-1 corresponding copper (Cu2+) chaperone and catalase (Trubiani et al., 2007). In unstressed hypothalamic cells and primary hippocampal neurons, TCAP-1 treatment increases expression of

α-actinin-4, β-actin and β-tubulin, induced neurite outgrowth, dendritic arborization, and axon fasciculation (Al Chawaf et al., 2007a). TCAP-1 also inhibits brain-derived neurotrophic factor

(BDNF) expression and translation in hypothalamic neurons (Ng et al., 2012).

In vivo, TCAP-1 has emerged as a novel candidate for the integration and modulation of a number of psychiatric disorders including stress, anxiety, and addiction (Rotzinger et al., 2010;

Tan et al., 2011b). Acute administration of TCAP-1 into the basolateral nucleus of the amygdala

(BLA) in rats modulated acoustic startle behaviour by increasing the startle-response in the low- startle group and decreasing startle response in the high-startle group (Wang et al., 2005).

Further anxiety studies, using an elevated plus maze (EPM) and open field tests (OF), showed that TCAP-1 modulates CRF-regulated behaviours. Repeated intravenous (IV) administration of

TCAP-1 had an anxiolytic effect on CRF-induced responses in both the EPM and OF tests (Al

Chawaf et al., 2007b). However, intracerebroventricular (ICV) administration of TCAP-1 yielded differential behavioural responses based on the presence or absence of a stress challenge

(Tan et al., 2008). In the absence of a CRF-mediated stressor, TCAP-1 had mild anxiolytic effects in the EPM and OF tests of behaviour; however, in the presence of a stressor TCAP-1 appeared to have anxiogenic effects in the EPM and OF tests (Tan et al., 2008). Interesting, the

TCAP-1 induced effects on behaviour are long-lasting, with effects persisting 21 days after

TCAP-1 injection (Wang et al., 2005), suggesting that TCAP-1 may regulate neuronal plasticity in the brain.

Further in vivo studies strengthened the notion of a functional link between the TCAP-1 and CRF systems. TCAP-1 blocked CRF-mediated c-fos synthesis in the hippocampus and amygdala of the adult rat (Tan et al., 2009) and increased spine density in the CA1 and CA3 neurons of the rodent hippocampus but not in the amygdala (Tan et al., 2011). Moreover, TCAP-

1 was shown to completely ablate CRF-induced cocaine seeking reinstatement in rats

(Kupferschmidt et al., 2011). However, despite the similarity between the CRF and TCAP family of peptides, the expression of TCAP-1 mRNA in CRF-responsive regions in the brain (Wang et al., 2005), and the ability for TCAP-1 to cross the blood brain barrier (Al Chawaf et al., 2007), binding studies indicate that TCAP-1 does not bind to CRF receptor 1 (CRFR1) or CRFR2

(Nock, 2009) or the CRF binding protein (A.F Seasholtz and D.A Lovejoy, unpublished

36 findings). These findings suggest that TCAP-1 could be acting through its own receptor signalling system to mediate neuroplastic changes and inhibit CRF-driven systems.

1.7 Objectives and Hypothesis

The teneurin-TCAP system represents a highly evolutionary conserved system across the metazoans and play fundamental roles in development and homeostasis. I hypothesize that the

TCAP-1 region of teneurin-1 acts independently of the larger teneurin-1 to regulate cytoskeletal elements as a prelude to the changes associated with TCAP-mediated neuroplastic changes and inhibition of CRF-driven systems. TCAP-1’s role in remodelling the neural cytoskeleton could explain the long-term modifications in brain plasticity and modulation of CRF-mediated behaviours. In addition, TCAP-1 unique set of functional attributes suggests that it may possess biological activity outside of the CNS.

The biological effects of TCAP-1 in the brain are well documented (Tan et al., 2011b) but raise a number of questions. First, it is not known if TCAP-1 exerts its effects as part of a direct teneurin function, whereby TCAP represents a functional region of the larger teneurin protein, or if it has an independent role, either as a splice variant or post-translational proteolytic cleavage product of teneurin. Secondly, if TCAP-1 is structurally distinct from the teneurin-1, and possesses its own complement of functional attributes, it would be expected to have distinct sites of immunoreactivity and action on cells in the brain. Much has been done to resolve the distribution of the teneurins in the CNS, but little is known of the distribution of the TCAP-1 region of teneurin-1 in the mammalian brain and whether or not it corroborates with the stress- regulating brain areas that are responsive to TCAP-1 (Tan et al., 2009). A detailed description of the TCAP-1 network in the mouse brain would provide an integrative framework that would aid

37 in the interpretation of many of the TCAP-1-regulated psychiatric disorders. Thirdly, a multitude of biological effects are attributed to TCAP-1, but the receptor-signalling system and mechanisms that are recruited by TCAP-1 remain unresolved. Fourthly, less is known about the localization and role of the vertebrate teneurin-TCAP system outside the CNS. The majority of studies on the teneurin-TCAP system have centered on resolving its function in the brain, with a few studies pointing to a teneurin function in the gonads of invertebrates (Drabikowski et al.,

2005; Trzebiatowska et al., 2008). Given that TCAP-1 is most efficacious at regulating homeostasis under stress conditions and is associated with metabolically demanding functions, it is likely that TCAP-1 may also be bioactive in other metabolically active tissues outside the

CNS, such as the gonads.

The studies described in this thesis were therefore aimed at addressing the following objectives:

1.) To determine the expression and processing of the TCAP-1 peptide encoded on the

terminal exon of the teneurin-1 gene.

2.) To characterize TCAP-1 binding sites, uptake and the cellular and tissue localization

of immunoreactive teneurin-1 and TCAP-1 in the mouse.

3.) To identify a possible receptor candidate and elucidate the molecular mechanism by

which TCAP-1 could regulate cytoskeletal dynamics and neuronal plasticity in the

mouse hippocampus.

4.) To investigate TCAP-1 localization, binding and in vivo functions outside the CNS,

particularly in the adult mouse testis.

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2 Chapter Two: Processing, localization and uptake of the C- terminal region of teneurin-1

This chapter has been published in Molecular and Cellular Neuroscience:

Chand D*, Casatti CA, de Lannoy L, Song L, Kollara A, Brown TJ, Lovejoy DA (2012) C- terminal processing of the teneurin proteins: Independent actions of a teneurin C-terminal associated peptide in hippocampal cells. Mol Cell Neurosci (In Press). (Authorization for reproduction obtained from Elsevier).

2.1 Abstract

The teneurins are a recently described family of proteins that play a significant role in visual and auditory development. Encoded on the terminal exon of the teneurin genes are a new family of bioactive peptides, termed teneurin C-terminal associated peptides (TCAPs) that regulate mood- disorder associated behaviours. Thus, the teneurin-TCAP system could represent a novel neurological system underlying the origins of a number of complex neuropsychiatric conditions.

However, it is not known if TCAP-1 exerts its effects as part of a direct teneurin function, whereby TCAP represents a functional region of the larger teneurin protein, or if it has an independent role, either as a splice variant or post-translational proteolytic cleavage product of teneurin. In this study, I show that TCAP-1 can be transcribed as a smaller mRNA transcript of the same size as the terminal exon of the teneurin-1 gene. After translation, further processing yields a smaller 15 kDa protein containing the TCAP-1 region. In the mouse hippocampus, immunoreactive (ir) TCAP-1 is exclusively localized to the pyramidal layers of the CA1, CA2 and CA3 regions. Although the localization of TCAP and teneurin in hippocampal regions is similar, they are distinct within the cell as most ir-teneurin is found at the plasma membrane, whereas ir-TCAP-1 is predominantly found in the cytosol. Moreover, in mouse embryonic hippocampal cell culture, FITC-labeled TCAP-1 binds to the plasma membrane and is taken up

52 into the cytosol via dynamin-dependent caveolae-mediated endocytosis. These data provide novel evidence that TCAP-1 is structurally and functionally distinct from the larger teneurins.

2.2 Introduction

The teneurins are a family of four type-II transmembrane proteins predominantly expressed in the central nervous system (CNS) (Levine et al., 1994; Tucker et al., 2007; Ben-Zur et al., 2000) and are implicated in visual, auditory and olfactory integration in the CNS (Rubin et al., 2002; Wang et al., 2005; Kenzelmann et al., 2007; 2008; Leamey et al., 2007; 2008; Young and Leamey, 2009). They were originally discovered in Drosophila as odd oz (odz; Levine et al.,

1994) on the basis of its expression pattern and tenascin-major (ten-m; Baumgartner et al., 1994), because of their structural homology to the tenascin proteins. The teneurins are highly conserved in metazoans indicating that they may play a fundamental role in the evolution of sensorimotor integration (Tucker et al., 2012).

Teneurin homologs are found in all metazoans examined thus far, and are functionally implicated in axon guidance (Young and Leamey, 2009), neurite outgrowth (Rubin et al., 1999;

Leamey et al., 2008), transcriptional regulation (Kenzelmann et al., 2008; Bagutti et al., 2003;

Nunes et al., 2005), cell proliferation (Kinel-Tahan et al., 2007) and adhesion (Rubin et al., 2002;

Leamey et al., 2008). Its long evolutionary history is reflected by the highly conserved domain heterogeneity of the teneurin structure. The teneurins consist of about 2800 amino acids and possess a complex set of functional domains (Minet et al., 1999; Feng et al., 2002; Leamey et al.,

2007). The short amino-terminus intracellular domain of vertebrate teneurins contains two EF- hand-like calcium-binding motifs and two polyproline domains characteristic of SH3-binding sites and which can interact with the actin-cytoskeleton via CAP/ponsin and vinculin (Nunes et

53 al., 2005). The extracellularly oriented, carboxyl-terminus domain is represented by about 2400 amino acid residues and is highly conserved across vertebrates. The carboxyl-terminus contains eight epidermal growth factor (EGF)-like repeats that interact via cysteine linkages with other teneurin paralogs to create homo- and heterodimers (Feng et al., 2002; Oohashi et al., 1999) and possesses 26 tyrosine-aspartic acid (YD) repeats, a distinct feature unique to this class of proteins in the eukaryotic proteomes (Kenzelmann et al., 2007).

Given the complexity of the teneurin structure, it is not clear how all of these functions are regulated in both the developing and adult CNS. However, studies indicate that some family members also exist as a number of splice variants (Lossie et al., 2005) or as smaller soluble proteins derived from further proteolytic cleavage of the carboxy terminal regions (Kenzelmann et al., 2008). One such region is a 40- to 41-amino acid sequence at the tip of the carboxy terminus of the teneurin proteins that possess characteristics of a bioactive peptide and is encoded by the teneurin 3’ terminal exon (Qian et al., 2004; Wang et al, 2005). This sequence is termed the ‘teneurin C-terminal associated peptide’ (TCAP).

Structurally, the TCAP family of peptides possess an amidation motif at the carboxyl- terminus and a pyroglutamic acid residue at the amino-terminus (Wang et al., 2005), features that are characteristic of bioactive peptides. Recent evidence indicates that the TCAP-1 region of the teneurin-1 may act as a novel neuroplastic regulatory factor in the rodent brain (Tan et al.,

2011b). In vitro, TCAP-1 increases cytoskeletal proteins including β-actin and β-tubulin, modulates neurite outgrowth in cultured hippocampal neurons (Al Chawaf et al., 2007a) and inhibits brain derived neurotrophic factor (BDNF) expression in hypothalamic cells (Ng et al.,

2012). In vivo, synthetic TCAP-1 regulates stress-induced behaviour (Wang et al., 2005; Al

Chawaf et al., 2007b; Tan et al., 2008), increases spine density in the CA1 and CA3 neurons of

54 the rodent hippocampus (Tan et al., 2011) and blocks CRF-mediated c-fos synthesis in the hippocampus and amygdala of the adult rat (Tan et al., 2009). Furthermore, TCAP-1 ablates cocaine-seeking reinstatement in adult rats (Kupferschmidt et al., 2011). These studies suggest that TCAP-1 may have applications in the treatment of stress-, anxiety- and addiction-like disorders and, together with the teneurins, may serve to link some pathophysiological elements of these conditions.

However, it is not known if TCAP-1 exerts its effects as part of a direct teneurin function, whereby TCAP-1 represents a functional region of the large teneurin-1 protein, or if it has an independent role, either as a splice variant or post-translational proteolytic cleavage product of teneurin. If TCAP-1 is structurally distinct from the teneurin-1, and possesses its own complement of functional attributes, it would be expected to have distinct binding sites and action on cells of the hippocampus. In this study, I investigated the expression and processing of the TCAP-1 peptide encoded on the terminal exon of teneurin-1 and further characterized the cellular and tissue localization, and uptake mechanism in the mouse hippocampus.

2.3 Materials and Methods

2.3.1 Animals. Adult male BALB/c mice obtained from Charles River Laboratories (Montreal,

Quebec, Canada) were housed in a controlled environment with a 12 h light/dark cycle at a constant temperature of 21C. For studies of adult brain, BALB/c mice aged 6-8 weeks were killed by cervical dislocation and brain tissues were dissected and snap-frozen in liquid nitrogen or fixed overnight in 4% paraformaldehyde (PFA; Sigma-Aldrich Canada, Oakville, ON,

Canada), pH 7.4, and paraffin-embedded. For immunization, adult male New Zealand white

55 rabbits were obtained from Charles River Laboratories. All procedures were approved by the local animal care committee and were in accordance with the Canadian Council on Animal Care.

2.3.2 Cell Culture. Immortalized mouse embryonic E14 hippocampal cells (Gingerich et al.,

2010; provided by Dr. D. Belsham, University of Toronto) were cultured in Dulbecco’s Modified

Eagle Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 100

U/ml penicillin and 100 µg/ml streptomycin (Invitrogen, Burlington, ON, Canada). Cells were maintained at 60-70% confluency at 37C in a humidified C02 incubator.

2.3.3 Northern blot analysis. Probes to mouse TCAP-1, teneurin-1 and β-actin were prepared by

RT-PCR using the following primers: mouse TCAP-1, forward 5’-ttcatttccttggatcagcttcctatg-3’, reverse 5’-aagctgctgcttttctccctctgtcca-3’ (585 bases); teneurin-1, forward 5’- gtgtcacctgatggcaccctctat-3’, reverse 5’-tcctgggtatgtcatcaaggccaa-3’ (402 bases); β-actin, forward

5’-caggtcatcactattggcaacgag-3’ reverse 5’-ctcatcgtactcctgcttgctgat-3’ (357 bases). The amplified sequences were purified using a Purelink Quick Gel DNA Extraction Kit (Invitrogen) and subcloned to pCR2.1-TOPO plasmids TOPO TA Cloning kit (Invitrogen). Plasmids with the insert were purified using the QIAprep Spin Mini Kit (Qiagen, Mississauga, ON, Canada) and sequence verified (ACGT, Toronto, ON, Canada). The northern blot was performed using the

NorthernMax kit (Ambion, Austin, TX, USA). Five micrograms (5 μg) of the total RNA from whole mouse brain were denatured for 15 min at 65ºC and separated in denaturing 2% formaldehyde-agarose gel. The RNA was UV-crosslinked to a positively-charged BrightStar-

Plus nylon membrane (Ambion; Stratalinker UV Crosslinker 1800, Stratagene), prehybridized with Ultrahyb overnight at 42ºC followed by hybridization with Ultrahyb and 1-5 ng/ml of

56 denatured biotinylated probe for 24-48 h at 42ºC. The membrane was washed for 2-3 h at 46-

59ºC according to manufacturer’s protocol (Ambion). The hybridization was visualized using a chemiluminescent nucleic acid detection system (Pierce Biotechnology, Rockford, IL, USA) followed by exposure to Kodak BIOMAX-MR film (Perkin Elmer, Waltham, MA, USA).

2.3.4 Synthesis of TCAP Peptides. Mouse TCAP-1, TCAP-2, and the fragments TCAP-1(1-8),

TCAP-1(9-41) and TCAP-1(1-37) were synthesized at 95% purity using f-moc-based solid phase synthesis (American Peptide Company, Sunnyvale, CA, USA). TCAP-1 variants with lysine substituted for arginine at positions 8 or 37 (K8 and K37) were synthesized as previously described (Wang et al., 2005). Two 25-amino acid peptides spanning either the mouse C-terminal sequence TCAP-1 (C-TCAP-1) or the N-terminal TCAP-1 (N-TCAP-1), were synthesized and conjugated with keyhole limpet hemocyanin (KLH) in a 1:1 ratio (w/w) by the Dalton Chemical

Company (Toronto, ON, Canada). The peptides were solubilized by exposure to ammonium hydroxide vapors for 2 min before dilution in phosphate-buffered saline (PBS) pH 7.4 with 10 mM sodium phosphate.

2.3.5 Production of TCAP-1 Antisera. The sequences used as haptens were KLH- pEQLLGTGRVQGYDGYFVLSVEQYLE-OH (pE: pyroglutamic acid) and KLH-

57 immunization. The conjugated peptides were emulsified with Freund’s complete adjuvant (FCA) in 1:1 ratio (V/V) at a final peptide concentration of 100 μg/ml. Four weeks after the injection, rabbits received a booster injection of peptides with Freund’s incomplete adjuvant. Periodic blood samples were collected to assess antibody titer. Booster injections were repeated until antibody titers were sufficient as determined by dot and western blotting.

The binding specificity of the antisera was determined by an enzyme-linked immunosorbent assay (ELISA). Nunc Maxisorp flat-bottom plates were coated with goat anti- rabbit IgG Fc fragment (Pierce Biotechnology) at 10 μg/ml overnight at 4°C. The remaining binding sites were blocked with 1% bovine serum albumin (BSA) in PBS 0.05% Tween-20

(PBS-T) for 2-16h and the wells then washed with PBS-T. TCAP-specific antisera, at 1:1000 dilution in PBS-T with 1% BSA, were bound to the goat anti-rabbit IgG Fc fragment-coated plates for 2h. The wells were washed 3x with PBS-T. [K8]- and [K37]-TCAP-1 variants were labeled with biotin using EZ-Link Sulfo-NHS-LC-Biotinylation Kit (Pierce Biotechnology).

Serial dilutions of unlabelled TCAP and 10 ng/ml biotinylated TCAP in PBS-T 1% BSA were incubated for 4h at room temperature (RT). The wells were washed 4x with PBS-T and incubated with streptavidin-HRP (Pierce Boitechnology) at 0.1 µg/ml in PBS-T 1% BSA for 25 min. After 3 washes with PBS-T, the substrate Super Signal TMB (Pierce Biotechnology) was added for 10-30 min, the reaction was stopped with sulfuric acid, and HRP activity measured at

450 nm. The competition curves and 50% binding values were calculated with GraphPad Prism 4

(San Diego, CA) software using variable slope sigmoidal regression and F test to compare the

50% binding values.

2.3.6 Western Blot Analysis. Whole brain tissue specimens were homogenized in cold lysis buffer (0.1 M PBS, pH 7.4, 1% Triton X-100, 1 mM PMSF and 1% protease inhibitor cocktail;

Invitrogen) using a Polytron tissue homogenizer. The nuclei and cell debris were removed by centrifugation at 15,000 g for 20 min at 4ºC. The supernatant was then collected and stored at -

80ºC to be used for western blotting experiments. Cultured mouse embryonic E14 hippocampal cells were lysed in ice-cold radioimmunoprecipitation assay (RIPA) lysis buffer (50 mM TRIS-

HCl, 150 mM NaCl, 1% Triton X-100, 0.1% SDS, 1% sodium deoxycholate, 1 mM EDTA, 1 mM PMSF, 1 mM sodium orthovanadate, 1 mM sodium fluoride and 1 μg/ml each of pepstatin, aprotinin and leupeptin) and placed on ice for 15 min. Cell debris was removed by centrifugation at 20,000g for 15 min at 4C. The supernatant was collected for western blot analysis. The protein concentration was determined using the bicinchoninic acid (BCA) kit as per instructions

(Pierce Biotechnology).

For Western blot analysis, homogenized adult mouse brain tissue and E14 RIPA cell lysate (50 μg/lane) were resolved by SDS-PAGE (BioRad mini-protean III cell) and transferred onto a nitrocellulose membrane (GE Healthcare, Piscataway, NJ, USA). After blocking with 5%

BSA in PBS-T for 1 h, the membrane was incubated with rabbit polyclonal affinity purified N- terminally directed TCAP-1 (TNR308) antiserum (1:1000), pre-immunized antiserum (1:1000) or mouse monoclonal teneurin-1 antiserum (1:1500; ABNOVA, Taipei City, Taiwan) overnight at 4°C. After washing with PBS-T, the membrane was incubated with HRP-labeled goat anti- rabbit IgG (1:5000; New England Biolabs, Pickering, ON, Canada) for 1 h at RT. The membrane was analyzed using the ECL western blotting detection and analysis kit (GE Healthcare).

For peptide extraction, whole brain tissues were weighed and homogenized in cold lysis buffer (5 ml/gram of tissue) as described above. The supernatant was collected, mixed with an

59 equal volume of chloroform and centrifuged at 15,000 g for 20 min at 4°C to remove lipids from the homogenate. After centrifugation, the supernatant was diluted with 0.1% trifluoroacetic acid

(TFA) in water at a ratio of 1:3 and passed through a Water Oasis™ HLB Plus extraction cartridge (SPE; Waters Limited, Mississauga, ON, Canada). The cartridge was washed with 5 ml of 0.1% TFA and 3 ml of 20% acetonitrile/80% water containing 0.1% TFA. Bound material was eluted with 3 ml of 60% acetonitrile/40% water containing 0.1% TFA and then concentrated using a vacuum centrifuge (Eppendorf Canada, Mississauga, ON, Canada). Western blot analysis, as described above, using mouse monoclonal anti-teneurin-1 (1:1500), TNR308 and affinity purified C-terminally direct TCAP-1 (TCR108) antisera at 1:1000 dilution were used to analyze the eluted fractions. The specificity of the affinity purified rabbit anti-mouse-TCAP-1

TNR308 and TCR108 antisera were tested by pre-adsorption with a 5-fold excess of synthetic mouse TCAP-1 peptide.

2.3.7 Immunofluorescent and immunohistochemical staining. Immortalized mouse E14 hippocampal cells were plated on poly-D-lysine-coated cover-slips (BD Biosciences,

Mississauga, ON, Canada) and cultured to 60% confluency in DMEM with 10% FBS as described above. The cells were fixed in 4% PFA (Sigma-Aldrich Canada) for 20 min and permeabilized for 10 min with 0.3% Triton X-100 (Fisher Scientific, Ottawa, ON, Canada). After blocking with 10% normal goat serum (NGS; Vector Laboratories, Burlington, ON, Canada) for

1 h, the cells were incubated with affinity-purified TNR308 TCAP-1 (1:1000), mouse monoclonal teneurin-1 (1:1000; ABNOVA), chicken anti-cow-microtubule-associated protein 2

(MAP2, 1:8000; Abcam, Cambridge, MA, USA) or chicken anti-cow-glial fibrillary acidic protein (GFAP, 1:2000; Abcam) primary antibodies, overnight at 4°C in PBS and 1% NGS. The

60 cells were washed in PBS and incubated with Alexa 594 goat anti-rabbit (1:400; Invitrogen),

Alexa 488 donkey anti-mouse (1:400; Invitrogen), FITC goat anti-chicken (1:250; Abcam) or

Texas-Red goat anti-chicken (1:200; Abcam) secondary antibodies in 0.01 M PBS for 1 h. Cells were counterstained with 4',6-diamidino-2-phenylindole (DAPI; Sigma-Aldrich Canada) for 1 min and mounted with 1,4-diazabicyclo[2.2.2]octane (DABCO; Sigma-Aldrich Canada). The specificity of the teneurin-1 antiserum was tested by pre-adsorption with a 5-fold excess of synthetic mouse recombinant teneurin-1 protein fragment (ABNOVA). For controls, cells were incubated with diluted pre-immune serum, primary antibody pre-adsorbed with their respective immunizing peptides, primary antibody alone, secondary antibody alone or, neither primary nor secondary antibodies.

For immunohistochemical analysis, paraformaldehyde-fixed paraffin-embedded sections

(5 µm) of adult mice brains were deparaffinized in xylene and rehydrated. The sections were rinsed in double-distilled water and placed in 10mM citrate buffer for antigen retrieval by heating in a 750W microwave oven 4 x 5 min with gentle agitation between each period.

Sections were permeabilized with 0.3% Triton X-100 (Fisher Scientific) for 10 min. The tissue sections were blocked with 10% NGS (Vector Laboratories) in PBS for 1 h and incubated overnight at 4˚C with affinity-purified TNR308 TCAP-1 antiserum (1:1000). After washing with

PBS, sections were incubated with Alexa 594 goat anti-rabbit IgG secondary antibody (1:400;

Invitrogen) in 0.01 M PBS for 1 h at RT. The sections were counterstained with DAPI (Sigma-

Aldrich Canada) for 1 min and mounted with DABCO (Sigma-Aldrich Canada). Controls were performed on adjacent sections in the manner described above.

2.3.8 Preparation of Fluoresceinisothiocyanate (FITC)-labeled TCAP-1. [K8] or [K37]-TCAP-1 was labeled with FITC according to the EZ-Label FITC Protein labeling kit (Pierce

Biotechnology) and as previously described (Al Chawaf et al., 2007b). The size of the FITC-

TCAP-1 conjugate was confirmed by non-reducing SDS-PAGE on a 10–20% Tris-Tricine gel

(BioRad, Hercules, CA, USA). The amount of FITC required to be used as a control was determined by titrating PBS-diluted preparations of FITC on a nitrocellulose membrane.

Fluorescence was measured using an Epi Chemi II Darkroom system (UVP) with the sequential integration function of LabWorks Image acquisition and analysis software (V4.0.0.8, Upland,

CA, USA).

2.3.9 FITC-TCAP-1 uptake and binding studies. Immortalized mouse E14 hippocampal cells were plated on poly-D-lysine-coated cover-slips as described above. The cells were rinsed with

PBS and incubated with conjugated mouse FITC-TCAP-1 diluted at 1:400 in DMEM with 10%

FBS for 60 min at 37°C and 5% CO2. The cells were washed with PBS, fixed in 4% PFA

(Sigma-Aldrich Canada) in PBS and counterstained with DAPI as described above. To determine the mechanism by which TCAP-1 is internalized, cells were pretreated with endocytic inhibitor,

Dynasore (100µM; Sigma-Aldrich Canada), caveolin inhibitor, Filipin (1µg/ml; Sigma-Aldrich

Canada) or clathrin inhibitor, chlorpromazine (CPZ, 40µM; Sigma-Aldrich Canada) for 1 h at

37°C and 5% CO2 before the addition of FITC-labeled TCAP-1. The cells were washed, fixed and counterstained with DAPI as described above. For controls, cells were incubated with unconjugated FITC (1:400) and treated with a corresponding dose of dimethyl sulfoxide

(DMSO). To further confirm the uptake mechanism of FITC-labeled TCAP-1, immunofluorescent co-localization with rabbit anti-caveolin-1 (New England Biolabs) was

62 performed on mouse E14 hippocampal cells that were pretreated with FITC-labeled TCAP-1 for

30 min as described above. Controls were performed as described above.

2.3.10 Image Analysis. Cells were analyzed using a Zeiss LSM 510 confocal laser-scanning microscope (Carl Zeiss MicroImaging, Göttingen, Germany) with a Q imaging Retiga EXi camera (Q Imaging, Surrey, BC, Canada). Tissue sections were analyzed using a Zeiss Axio

Imager Z1 Deacon microscope (Carl Zeiss MicroImaging) with a Q imaging Retiga EXi camera

(Q Imaging). The resulting images were adjusted for brightness and contrast using Volocity 5.2

(Improvision, Waltham, MA, USA) and Adobe Photoshop CS4 Extended (San Jose, CA, USA).

2.4.0 Results

2.4.1 Transcription and translation of TCAP-1

A northern blot was performed on mRNA extracted from whole mouse brain to verify the size of the TCAP-1 transcript. A cDNA probe prepared against exon 31, the terminal exon encoding the TCAP region, showed the presence of two transcripts, approximately 700 bp and

7000-8000 bp, similar to the expected size of the terminal exon (832 bp) and full-length mRNA(8341 bp; NCBI Accession No: NM_011855.3) of the teneurin-1 gene, respectively (Fig.

2.1A). In contrast, a cDNA probe directed to exon 25, upstream of the TCAP region, showed only the presence of the full-length mRNA transcript of the teneurin-1 gene (Fig. 2.1A).

Sequence analysis of the last exon of teneurin-1, of which the short transcript encodes, revealed the presence of numerous potential basic/dibasic cleavage motifs (Fig. 2.1B) suggesting that cleavage of this region could liberate a smaller protein possessing the TCAP region. Five potential cleavage sites are indicated as CL1-CL5, however, only three of the basic/dibasic

63 cleavage motifs (CL2, CL3 and CL5) are conserved amongst the terminal exons of all teneurins.

The paired basic amino acid sequence (RR) indicated by CL2 yields a fragment of 130-133 amino acid residues or a peptide around 14.1–14.8 kDa. The cleavage motif CL3 (RxRR) predicts a fragment 107-110 amino acids in length, corresponding to size of 11.6 – 12.3 kDa.

Cleavage motif CL5 would result in the liberation of the 41-residues for TCAP-1.

To verify if shorter proteins, immunoreactive (ir) for the TCAP-1 portion, could be observed in cell extracts, I examined the size of ir-TCAP-1 proteins by western blot using

TCAP-1 specific antisera. To do this, antisera were generated to the C- and N-terminal regions spanning the 41 residues of the TCAP-1 sequence (Fig. 2.1B). Two antisera, TNR308 (TCAP amino terminus directed) and TCR108 (TCAP carboxyl terminus directed) possessed non- overlapping epitopes with the desired binding characteristics and were specific only to TCAP-1

(Fig. 2.1B,C). TCAP-1 TCR108 and TNR308 antisera both bound strongly to the full-length

TCAP-1 peptide (Fig. 2.1C). However, the truncated TCAP-19-41 fragment was also recognized by TCR108 antiserum (B50 =11.7nM; Fig. 2.1C). TCAP-1 TNR308 antiserum showed weak binding affinity to the TCAP-19-41 (B50 >200nM) and 1-8 (B50 >200nM) TCAP-1 fragments (Fig.

2.1C). TCAP-2 was not recognized by TNR308 or TCR108 antisera (B50 >200nM; Fig. 2.1C) suggesting that both antisera are specific to TCAP-1. However, TNR308 antiserum was used for all immunofluorescence studies as it showed the stronger binding affinity (B50 =10.7nM) and higher specificity for the TCAP-1 peptide relative to TCR108 (B50 =22nM; Fig. 2.1C).

Western blot analysis with TCAP-1 TNR308 antiserum, indicated that the TCAP region of teneurins was primarily present as two smaller protein fragments suggesting cleavage events at the carboxy terminus of the teneurin protein. TCAP-1 TNR308 and TCR108 antisera both

Figure 2.1. Expression and processing of the TCAP portion of mouse teneurin-1. A. Northern blot of whole mouse brain. The size of the transcripts (kb) is indicated by the scale on the left. B. Translated portion of the terminal exon of mouse teneurin-1. Basic residues are shown in bold. The putative translation site (TS) and amidation motifs are boxed. Five potential cleavage spots are indicated as CL1- CL5. The epitopes of the three antisera are shown as light gray boxes. C. Binding specificity of the TCAP antisera used in the study: TNR308 (N- terminally directed) and TCR108 (C-terminally directed). D. Western blot of mouse brain extracts (Br), E14 RIPA cell lysate and mouse brain peptide extracts (SPE) using TCAP-1 (TNR308 and TCR108) antisera and teneurin-1(Ten-1/Odz-1) antiserum.

detected immunoreactivity of the putative 15 kDa ir-TCAP fragment whereas the much larger 70 kDa ir-TCAP protein was more strongly detected with the C-terminal directed TCR108 TCAP-1 antiserum (Fig. 2.1D; SPE). A band of about 15 kDa could correspond to the cleavage position at

CL2 or CL3 described above. Although I could not find evidence of a 4.7 kDa peptide corresponding to the terminal 41 residues, the majority of ir-TCAP was clearly present as smaller

65 protein fragments. Immunoreactivity greater than 250 kDa, possibly that of a full-length teneurin was also observed in hippocampal cell lysate.

Western blot analysis with teneurin-1 antiserum, whose epitope is directed to a region upstream of the TCAP-1 sequence but within the amino acid sequence encoded by the terminal teneurin-1 exon (Fig. 2.1B), also detected immunoreactivity of the putative 15 kDa band in whole brain peptide extracts. A smaller 10 kDa ir-teneurin-1 fragment (Fig. 2.1D; SPE) was also detected, as would be expected if the 4.7 kDa TCAP-1 peptide was cleaved at position CL5 as described above. Strong ir-teneurin at 70 kDa, similar to that observed with TCAP-1 TNR308 and TCR108 antisera was also detected in mouse brain peptide extracts (Fig. 2.1D; SPE).

Western blot analysis of mouse E14 hippocampal cell lysate probed with teneurin-1 antiserum detected a band greater than 250 kDa, similar to that detected with TCAP-1 TNR308 antiserum

(Fig. 2.1D). In addition, teneurin-1 immunoreactivity was also detected at approximately 200 kDa and 50 kDa in E14 hippocampal cell lysate (Fig. 2.1D).

2.4.2 Localization of Teneurin-1 and TCAP-1 to hippocampal cells and tissues

The localization of TCAP-1 using TCAP-1 TNR308 antiserum was examined and compared to that obtained for teneurin-1-immunoreactivity to determine whether TCAP-1 and teneurin-1 labeling was similarly distinct. In immortalized mouse E14 hippocampal cells, immunofluorescent labeling with teneurin-1 antiserum localized the larger type-II transmembrane teneurin-1 protein to the plasma membrane (Fig. 2.2A,B,G,H). However, fluorescent labeling with TCAP-1 TNR308 antiserum showed intense punctate-like immunoreactivity of TCAP-1 in the cytosol (Fig. 2.2C,D,I,J). Teneurin-1 and TCAP-1 co- localization studies showed that ir-TCAP-1 did not co-localize with ir-teneurin-1 in the cytosol

Figure 2.2. TCAP-1 and teneurin-1 immunoreactivity in mouse E14 hippocampal cells. Immunofluorescent labeling with monoclonal teneurin-1 antiserum (red) on E14 hippocampal cells distinctly localized teneurin-1 to the plasma membrane (B and H). However, affinity purified TNR308 antiserum (green) showed strong punctate-like TCAP-1 immunoreactivity in the cytosol (D and I). Weak teneurin-1 and TCAP-1 co-localization (yellow; arrow) was observed near the plasma membrane (J). Immunoreactivity was not observed in sections incubated with pre-adsorbed ODZ-1 antiserum (E) or pre-adsorbed TNR308 antiserum (F). For each fluorescence image, the corresponding DIC image is represented on the left (A, C, and G). Only neuronal cells were present in cell culture as indicated by the presence of MAP2 (K) and absence of GFAP (L). All sections were counterstained with DAPI to highlight cellular DNA. Magnification, 630X (A-F) and 200X (K and L). Scale bars, 25μm. MAP2, microtubule- associated protein 2; GFAP, glial fibrillary acidic protein.

of E14 hippocampal cells, but weak co-localization of both teneurin-1 and TCAP-1 was observed near the plasma membrane (Fig. 2.2G,J; white arrow). TCAP-1 (Fig. 2.2E) and teneurin-1 (Fig.

2.2F) immunoreactivity was completely abolished with the respective pre-adsorbed antisera.

Only neuronal cells were present in the cell culture as indicated by the presence of

67 immunoreactive microtubule-associated protein 2 (MAP2; Fig. 2.2K) and absence glial-specific marker glial fibrillary acidic protein (GFAP; Fig. 2.2L).

Figure 2.3. TCAP-1 immunoreactivity in the neuronal layers of the mouse hippocampal formation. Sections labeled with TCAP-1 TNR308 antiserum (red) showed strong TCAP-1 immunoreactivity in the cytosol of the neuronal layers of the hippocampus (B, E, H and K). TCAP immunoreactivity was strongest in the pyramidal layer (Py) of the CA2 (E) and CA3 regions (H) but weak in the dentate gyrus (K). TCAP-1 immunoreactivity was also observed in the stratum oriens (SO) of the CA1 (B) and stratum lucidum (SL) and stratum radiatum (SR) of the CA2 (E) and CA3 (H) regions of the mouse hippocampus. Immunoreactivity was not observed in sections incubated with pre-adsorbed TNR308 antiserum (C, F, I and L). For each fluorescence image, the corresponding DIC image is represented on the left (A, D, G and J). All sections were counterstained with DAPI. Magnification, 200X. Scale bars, 50μm.

To determine the distribution of ir-TCAP-1 in the mouse hippocampus, I used fluorescent labeling with TCAP-1 TNR308 antiserum on coronal sections of the mouse brain. TCAP-1- labeling was strongly localized to the two main neuronal cellular layers of the hippocampus, the granular layer of the dentate gyrus and the pyramidal layer (Py) of the CA1, CA2 and CA3 (Fig.

2.3). TCAP-1 immunoreactivity was strong in the CA1 (Fig. 2.3A,B), CA2 (Fig. 2.3D,E) and

CA3 (Fig. 2.3G,H) pyramidal layer and weak in the granular layer of the dentate gyrus (Fig.

2.3J,K). TCAP-1 was also localized to the stratum oriens (SO) of the CA1 (Fig. 2.3A,B) and stratum lucidum (SL) and stratum radiatum (SR) of the CA2 (Fig. 2.3D,E) and CA3 (Fig.

2.3G,H) regions of the mouse hippocampus. Immunoreactivity was absent in sections incubated with pre-adsorbed TCAP-1 TNR308 antiserum (Fig. 2.3C,F,I,L).

2.4.3 Caveoli-dependent endocytosis of TCAP-1 in hippocampal cells

The cytoplasmic binding of FITC-TCAP-1 was suggestive of an active cellular transport mechanism. Therefore, TCAP-1-binding sites were further examined by incubating cultured mouse E14 hippocampal cells with FITC-conjugated TCAP-1. Some cells incubated with FITC-

[K8]-TCAP-1 for 60 min, showed strong binding on or near the plasma membrane (Fig. 2.4A), and in other cells appeared to be internalized as a large bolus (Fig. 2.4B) or distributed in a punctuate-like pattern throughout the cytosol (Fig. 2.4C). As further confirmation, fluorescence was not observed in any of the cells incubated with the un-conjugated FITC control (data not shown). To elucidate the uptake mechanism of TCAP-1 in cell culture, immortalized mouse E14 hippocampal cells were pretreated with various inhibitors and incubated with FITC-[K8]-TCAP-

1 for 60 min. In cells pretreated with dynamin inhibitor, Dynasore, FITC-TCAP-1 remained bound at or near the plasma membrane (Fig. 2.5C,D). A complete lack of FITC-labeled TCAP-1

69 uptake was similarly observed in hippocampal cells that were pretreated with caveoli inhibitor, filipin (Fig. 2.5E,F). This is in contrast to cells that were pretreated with clathrin inhibitor, CPZ, which showed strong uptake of FITC-labeled TCAP-1 to the cytosol (Fig. 2.5G,H), suggesting that the endocytosis of TCAP-1 is independent of clathrin-mediated endocytosis. The internalization of FITC-labeled TCAP-1 to the cytosol was unaffected by DMSO (Fig. 2.5A,B).

Figure 2.4. TCAP-1 uptake in mouse E14 hippocampal cells. Cultured E14 hippocampal cells incubated with FITC-[K8]-TCAP-1 for 60 min showed binding to the plasma membrane (A), internalization (B) and strong punctate-like labeling in the cytosol (C). All sections were counterstained with DAPI. Magnification, 630X. Scale bars, 10μm.

To further confirm the uptake mechanism of FITC-labeled TCAP-1, immunofluorescent co-localization with rabbit anti-caveolin-1 was performed on mouse E14 hippocampal cells that were pretreated with FITC-labeled TCAP-1. Cells incubated with FITC-labeled TCAP-1 for 30 min showed strong FITC-labeled TCAP-1 binding (Fig. 2.5I,K) and corresponding caveolin-1 immunoreactivity (Fig. 2.5I,J) at the plasma membrane. Immunoreactive caveolin-1 was also distributed throughout the cytosol (Fig. 2.5I,J). Strong co-localization between FITC-labeled

TCAP-1 and caveolin-1 was observed near the plasma membrane of E14 hippocampal cells (Fig.

2.5I,L; yellow, white arrow), providing further confirmation that the TCAP-1-receptor internalization at the plasma membrane is dependent on caveolae-mediated endocytosis.

Figure 2.5. Caveolin-dependent endocytosis of FITC-[K8]-TCAP-1 in mouse E14 hippocampal cells. Cultured E14 hippocampal cells incubated with FITC-[K8]-TCAP-1 for 60 min showed strong uptake and labeling in the cytosol (A). Cells pretreated with dynamin inhibitor Dynasore (100µM) strongly inhibited FITC-labeled TCAP-1 uptake (D). Pretreatment with caveolin inhibitor filipin (1μg/ml) completely inhibited TCAP-1 uptake (F). However, TCAP-1 uptake was not affected by pretreatment with clathrin inhibitor chlorpromazine (CPZ, 40μM; H). Cells incubated with FITC-labeled TCAP-1 for 30 min showed strong FITC-labeled TCAP-1 binding at the plasma membrane (I and K). Caveolin-1 immunoreactivity was detected both in the cytosol and at the plasma membrane (I and J). Strong co-localization between FITC- labeled TCAP-1 and caveolin-1 was observed near the plasma membrane (I and L; yellow, white arrow). Corresponding DIC images are represented on the left (A, C, E, G and I). All sections were counterstained with DAPI. Magnification, 630X. Scale bars, 20μm (A-H) and 15μm (I-L).

2.5 Discussion

The data described in this study strongly support the hypothesis that under some situations, TCAP-1 is functionally distinct from teneurin-1 in the mouse hippocampus. I show that TCAP-1 can be liberated both by cleavage from the full-length teneurin and by subsequent transcription, translation, and processing of a smaller TCAP-1 specific transcript. Several lines of evidence support this. First, cDNA probes to TCAP-1 indicate it can be transcribed as mRNA corresponding to the full-length teneurin gene and a shorter TCAP-1-encoding transcript about the same size as the terminal exon. Second, after translation, either the short form or the full- length form may be processed into a 15 kDa TCAP form. Third, specific antisera to the TCAP-1 region predominantly labels an epitope in the cytosol, whereas antiserum directed upstream from the TCAP-1 region primarily labels plasma membrane regions. Finally, the 41-residue form binds to and is internalized by the cell corroborating previous functional studies of a direct action on cells. Therefore, I postulate that TCAP-1 can exert its actions either as a cleaved or tethered peptide associated with the teneurins and also as an independently transcribed gene or splice variant (Fig. 2.6).

A cDNA probe directed to the 3’terminal exon that encodes the TCAP-1 region showed the presence of about a 700-bp and full-length transcripts as determined by Northern blot. The short transcript size is consistent with the postulated size of the 3’ terminal exon if it was independently transcribed either as a separate gene or as a teneurin-derived splice variant.

Alternative splice variants for the teneurin proteins have been described (Lossie et al., 2005;

Kenzelmann et al., 2008); however, splice variants encompassing primarily the final exon have not yet been described. Thus my finding of the short transcript may be specific to particular brain regions, as whole mouse brain extracts were used in the northern blot analysis. Interestingly, the

72 sequence encoded by the terminal exon lacks a signal peptide, thus TCAP-1 could be synthesized by free ribosomes as part of the non-secretory pathway. Proteins destined for cytosolic locations lack a signal peptide and remain essentially where they are after being translated. This is also consistent with my finding of extensive cytosolic labeling of ir-TCAP-1 and the existence an ir- teneurin-1 and -TCAP-1 15 kDa protein that may represent part of the teneurin-TCAP-1 propeptide. It is notable that the independent transcription, translation and processing of the smaller

TCAP-1-containing transcript, is similar to that utilized by other cytosol-soluble peptides such as ciliary neurotrophic factor (CNTF; Stockli et al., 1989; Stockli et al., 1991) and fibroblast growth factor 1 (FGF-1; Jackson et al., 1992). However, consistent with previous studies (Oohashi et al.,

1999), I also detected a full-length transcript using the terminal exon cDNA probe. Thus this suggests that the TCAP-1 region of the teneurin gene can also be transcribed as part of the final full-length teneurin protein.

Generally, soluble proteins are synthesized on free ribosomes whereas secretory and integral proteins are synthesized by the ribosomes on the endoplasmic reticulum (ER). The latter mechanism limits the ER-synthesized proteins to those that possess hydrophobic sequences as part of the signal peptide or other domains, such as membrane spanning regions (Stephens and

Nicchitta, 2008; Lerner et al., 2009). Teneurins, with their stretch of hydrophobic residues in the membrane spanning region, are therefore sorted via this latter route, where they become incorporated into the plasma membrane upon fusion of the secretory vesicles with the plasma membrane. My studies show the existence of the full-length transcription of teneurin-1 that includes the TCAP-1-containing exon, the existence of a 250-300 kDa protein on both the

TCAP-1 antiserum- and teneurin-1 antiserum-probed western blots and weak co-localization between teneurin-1 and TCAP-1 at the plasma membrane, suggesting that in some cases, the

TCAP-1 sequence may indeed be translated as part of the larger teneurin-1 protein. It may not have been possible to observe strong immunoreactive TCAP-1 signal on the plasma membrane because the epitope may have been obscured by the globular arrangement of the extracellular domain of the teneurins (Feng et al., 2002). Assuming this to be case, then TCAP-1 may act as a functional unit of teneurin by remaining tethered to it, in a similar manner to the notch signaling of serrate and delta ligands (Artavanis-Tsakonas et al., 1999; Hicks et al., 2002). Given this scenario, TCAP can be liberated by proteolytic cleavage by vesicle or plasma membrane bound peptidases, and then secreted into the extracellular matrix.

Assuming then that TCAP-1 may be included as either the full-length plasma membrane teneurin protein, or as a smaller soluble protein resident in the cytosol, then it could be liberated from its precursor by a common mechanism. Removal of the ‘‘prepro’’ sequence during progression through the secretory pathway is an essential step in the processing of secreted proteins to their mature form (Brechler et al., 1996) and is often achieved by cleavage at dibasic residues (Lim et al. 2007) and consensus sequences by prohormone convertases such as furin

(Seidah et al., 1996; Seidah, 2011). The carboxyl-terminus of teneurin-1 is characterized by a highly conserved furin-like cleavage site located about 135 amino acids from the C-terminus

(Rubin et al., 1999; Tucker and Chiquet-Ehrismann, 2006). Cleavage of the precursor at this motif would liberate a protein of about 14.8 kDa, similar with my findings of a 15 kDa immunoreactive TCAP-1 in cell extracts western blots. A similar mechanism could occur on the plasma membrane form of TCAP-1 as well. The expected TCAP-processing mechanism at the plasma membrane may, therefore, resemble the ectodomain-shedding mechanism of tumor necrosis factor (TNF) (Garton et al., 2001) or Apo-2 ligand (Pitti et al., 1996). These peptides are also type-II transmembrane proteins whose extracellular C-terminus can be cleaved via a

74 membrane-bound member of the prohormone convertase family of processing enzymes. In support of this, and similar to that detected with TCAP-1 antiserum, previous teneurin investigations have reported the existence of ir-teneurin fragments at 250 and 150 kDa, which are the result of ectodomain shedding of the large C-terminal fragment and/or regulated intramembrane proteolysis of teneurin-1 (Kenzelmann et al., 2008). Further proteolytic processing of the teneurin C-terminus can occur to liberate several smaller fragments or pro- peptides. Our detection of ir-tenurin-1 and -TCAP-1 at approximately 70 kDa, which is much larger than the expected size of the translated portion of the entire terminal exon but smaller than the full length teneurin-1 is consistent with further proteolytic cleavage of the C-terminus of teneurin and is well supported by previous teneurin studies that have also reported teneurin fragments of similar size in mouse neuronal cells (Rubin et al., 1999).

Given the strength and the consistency of my observations of the 15 kDa immunoreactive

TCAP-1 form, it is likely that this form is relatively stable in the cytosol. However, a strong consensus peptide cleavage site separates the TCAP-1 region from the remainder of the elongated 15 kDa form. Although numerous previous studies have verified the biological activity of synthetic TCAP-1 (Tan et al., 2011b), I could not consistently detect the 41-mer TCAP-1 in purified hippocampal cell extracts and brain tissue homogenates under a variety of conditions.

One possibility is that this 41-mer does not exist in vivo, but rather represents the functional region of the 15 kDa form. However, the uptake of FITC-labeled TCAP-1, an R/K-Xn-R/K prohormone convertase cleavage motif (Seidah and Chretien, 1999) adjacent to the mature peptide and bioactivity of synthetic TCAP-1 both in vitro and in vivo (Tan et al., 2011b), suggests that cleavage of the 15 kDa pro-peptide could occur to liberate TCAP-1. TCAP-1 may, therefore, be released as the 15 kDa form, but cleaved in the extracellular region immediately

75 before binding and uptake by surrounding cells. Thus, similar to nerve growth factor (NGF),

TCAP-1, in its precursor form, may be released in an activity-dependent manner, where it is quickly processed by the coordinated release and action of proenzymes and enzyme regulators

(Bruno and Cuello, 2006; Lim et al., 2007) before binding and internalization by surrounding cells.

Thus, both the previous transcription and translation studies support an independent role for TCAP-1 under some circumstances. Moreover, the immunoreactive labeling of TCAP-1 and teneurins showed two distinctive patterns. My studies show that TCAP-1 immunoreactivity corresponds to TCAP-1 mRNA (Wang et al., 2005) and teneurin-1 mRNA expression (Zhou et al., 2003) in the hippocampus, but differs markedly from that of teneurin-1 immunoreactivity both in vitro and in vivo (Baumgartner et al., 1994; Zhou et al., 2003). This further supports a mechanism by which TCAP-1 is independently processed from teneurin-1 and/or cleaved from the C-terminal region by proteolytic cleavage (Rubin et al., 1999). As observed in the mouse hippocampus, the TCAP-1 portion of teneurin-1 remains distributed in the cytosol, whereas the remaining teneurin-1 protein is deposited in the plasma membrane or to axonally targeted regions distant from the cell body (Zhou et al., 2003; Kenzelmann et al., 2008).

However, the role of TCAP-1 could also to be closely associated with that of teneurin-1 because TCAP-1 is immunoreactive in all cell groups of the mouse hippocampus that show both teneurin-1 (Zhou et al., 2003) and TCAP-1 mRNA expression (Wang et al., 2005). Interestingly, the expression pattern of teneurin-1 mRNA does not coincide with the localization pattern of the protein in the mouse hippocampus (Zhou et al., 2003). Teneurin-1 immunoreactivity was mainly localized within the molecular layers with weak immunoreactivity in the stratum pyramidale, whereas the mRNAs were specifically confined to the cell somata of the stratum pyramidale and

76 the granular layer of the dentate gyrus (Zhou et al., 2003). TCAP-1, on the other hand, was strongly localized to the stratum pyramidale, with weak immunoreactivity in the stratum oriens, stratum lucidum and stratum radiatum of the mouse hippocampus. Likewise, the expression pattern of Drosophila teneurin mRNA does not coincide with the protein (Baumgartner et al.,

1994) similar to that of other neural cell adhesion molecules, such as L1 (Jucker et al., 1996).

While axonal transport has been postulated for the discrepancy between the mRNA and protein pattern (Zhou et al., 2003), the differential mRNA and protein pattern of teneurin-1 could also be accounted for by the fact that teneurin-1 is a secreted protein. Thus, the localization pattern of the teneurin-1 protein would be ultimately determined by the localization of the postulated receptor.

Recently, the C-terminal region of teneurin-2 has been shown to be a novel juxtacrine-signaling ligand for the neuronal G protein–coupled receptor latrophilin 1 (Silva et al., 2011). I have recently established that TCAP-1 may be novel ligand for the dystroglycans in hippocampal neurons (Chand et al., 2012; Chapter 4). Unlike the dystroglycans, latrophilin 1 does not appear to be internalized or associated with endocytotic-machinery.

Regardless of the processing mechanism, the 41-mer TCAP-1 binds to, and is internalized by hippocampal neurons. I have previously demonstrated a functional interaction between TCAP-1 and the cytoskeletal-regulating laminin-binding protein, dystroglycan, at the plasma membrane of hippocampal neurons and a corresponding induction of ERK-dependent phosphorylation of cytoskeletal-binding proteins, stathmin and filamin A (Chand et al., 2012;

Chapter 4). Here, I show that TCAP-1 not only binds to the plasma membrane but is internalized via dynamin-dependent caveolae-mediated endocytosis. This is further supported by previous studies that have shown that dynamin, a GTPase involved in endocytosis, recycling of presynaptic vesicles and receptor internalization in most cells including neurons, has a high

77 affinity for dystroglycan (Zhan et al., 2005). Interestingly, the dystroglycans have also been previously shown to be functionally related to teneurin-1 (Trzebiatowska et al., 2008).

Furthermore, caveolin-3, which along with other caveolins, acts as a scaffolding protein in the formation of caveolae to form large invaginations at the plasma membrane, can bind to the C– terminus of β-dystroglycan (Sotgia et al., 2000). Like TCAP-1, FGF-1 and FGF-2 are also internalized by receptor-mediated endocytosis to reach cytosolic compartments (Wiedlocha and

Sorensen, 2004). However, the exact cytosolic target remains unresolved, but it has been previously established that that TCAP-1 binds to at least two cytosolically-expressed proteins

(Ng et al., 2012) and shows strong and specific binding to the somata of the stratum pyramidale of the mouse hippocampus (Chand et al., 2012; Chapter 4). Moreover, TCAP-1 stimulates transcription of cytoskeletal proteins β-actin and β-tubulin (Al Chawaf et al., 2007a) and inhibits transcription of brain derived neurotrophic factor (BDNF; Ng et al., 2012).

Central to the interpretation of these studies, was the concept that TCAP-1 can be released from its cytosolic-stores to function as an autocrine or paracrine signaling molecule.

Like FGF-1 (Jackson et al., 1992; Shin et al., 1996), interleukin 1b (Rubartelli et al., 1990), galectin-1 (Cooper and Barondes 1990) and other cytosol-soluble proteins, TCAP-1 lacks a signal sequence for secretion, thus its release mechanism is likely to follow the non-classical protein export or ER/Golgi-independent secretion of peptides. The molecular mechanisms and components that mediate ER/Golgi-independent secretion of peptides are unknown but is thought to be achieved through cell injury/death, cell leakage (D’Amore, 1990), the formation of labile structures such as exosomes in a process known as membrane blebbing (Nickel, 2003), or a novel and distinct secretion pathway. However, the release of TCAP-1 may be stimulus dependent. TCAP-1 can block CRF-related stress-induced behaviours (Wang et al., 2005; Al

Chawaf et al., 2007b; Tan et al., 2008), addiction behaviours (Kupferschmidt et al., 2011) and regulate corresponding morphological changes (Tan et al., 2011) in the hippocampus. TCAP-1 may, therefore, be released under stress conditions similar to that of interleukin 1b (Rubartelli et al., 1990) and FGF-1 (Jackson et al., 1992; Shin et al., 1996).

Although, the teneurins have been established as type-II transmembrane proteins

(Oohashi et al., 1999) with evidence of proteolytic cleavage in the mouse, rat and chicken (Rubin et al., 1999), a majority of the investigations have focused on the release and translocation of the

N-terminal fragment (Kenzelmann et al., 2008). Little is known about the processing, function and localization of the cleaved C-terminal fragment of teneurin-1. My study shows for the first time that the C-terminal region of teneurin-1, corresponding to TCAP-1, can be processed and localized independently from the much larger teneurin-1 (Fig. 2.6). This study also raises several interesting questions. Are the other three TCAP homologs structurally and functionally independent from their teneurin precursors? Is TCAP-1 independently processed from teneurin-1 in other brain regions? This study focusses primarily on the mouse hippocampus, and thus the teneurin-TCAP-1 relationship in this region of the brain may not be representative of teneurin-

TCAP-1 processing in the whole brain. In the mouse hippocampus, TCAP-1 is mainly localized to cytosol and pyramidal neurons of the hippocampus, in contrast to teneurin-1, which is primarily localized to the plasma membrane and molecular areas. Although further studies are needed to locate the promoter region corresponding to the smaller TCAP-1 transcript and positively identify the smaller teneurin-TCAP-1 fragments, my study adds further confirmation to the growing body of evidence of teneurin-1 processing in vertebrates. Moreover, I provide the first evidence that the cleaved TCAP-1 peptide is internalized via dynamin-dependent caveolae- mediated endocytosis in hippocampal neurons. Therefore, in conjunction with previous

79 investigations, this study strongly supports the hypothesis that TCAP-1 can function independently from teneurin-1 as either a paracrine or autocrine signaling molecule in the mouse hippocampus. The teneurin-TCAP system may, therefore, be part of a previously unknown neurological signaling system that may regulate neuronal function and some neuropathologies.

Figure 2.6. Proposed mechanism for TCAP-1 processing, release and uptake in hippocampal cells. TCAP is transcribed as part of the full-length teneurin mRNA or part of a shorter splice variant (1). The full-length mRNA is translated as part of the secretory pathway (2) and the TCAP portion may be cleaved in secretory vesicles or on the plasma membrane (3). The shorter TCAP mRNA is translated by free ribosomes and remains in the cytosol as part of the soluble protein translation ER/Golgi-independent pathway (4). TCAP is released from the cytosol during rupture of the plasma membrane and may undergo further proteolytic cleavage in the extracellular milieu (5). Once released, TCAP binds to its receptor and is taken up by neighboring cells via dynamin-dependent caveolae-mediated endocytosis (6).

2.6 References

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Tan, L.A., Xu, K., Vaccarino, F.J., Lovejoy, D.A., Rotzinger, S., 2009. Teneurin C-terminal associated peptide (TCAP)-1 attenuates corticotropin-releasing factor (CRF)-induced c-Fos expression in the limbic system and modulates anxiety behavior in male Wistar rats. Behav. Brain Res. 201, 198-206. Trzebiatowska, A., Topf, U., Sauder, U., Drabikowski, K., Chiquet-Ehrismann, R., 2008. Caenorhabditis elegans teneurin, ten-1, is required for gonadal and pharyngeal basement membrane integrity and acts redundantly with integrin ina-1 and dystroglycan dgn-1. Mol. Biol. Cell 19, 3898-3908. Tucker, R.P., Beckmann, J., Leachman, N.T., Scholer, J., Chiquet-Ehrismann, R., 2012. Phylogenetic analysis of the teneurins: conserved features and premetazoan ancestry. Mol. Biol. Evol. 29, 1019-1029. Tucker, R.P., Chiquet-Ehrismann, R., 2006. Teneurins: a conserved family of transmembrane proteins involved in intercellular signaling during development. Dev. Biol. 290, 237-245. Tucker, R.P., Kenzelmann, D., Trzebiatowska, A., Chiquet-Ehrismann, R., 2007. Teneurins: transmembrane proteins with fundamental roles in development. Int. J. Biochem. Cell. Biol. 39, 292-297. Wang, L., Rotzinger, S., Al Chawaf, A., Elias, C.F., Barsyte-Lovejoy, D., Qian, X., Wang, N.C., De Cristofaro, A., Belsham, D., Bittencourt, J.C., Vaccarino, F., Lovejoy, D.A., 2005. Teneurin proteins possess a carboxy terminal sequence with neuromodulatory activity. Brain Res. Mol. Brain Res. 133, 253-265. Wiedlocha, A., Sorensen, V., 2004. Signaling, internalization, and intracellular activity of fibroblast growth factor. Curr. Top. Microbiol. Immunol. 286, 45-79. Young, T.R., Leamey, C.A., 2009. Teneurins: important regulators of neural circuitry. Int. J. Biochem. Cell. Biol. 41, 990-993. Zhan, Y., Tremblay, M.R., Melian, N., Carbonetto, S., 2005. Evidence that dystroglycan is associated with dynamin and regulates endocytosis. J. Biol. Chem. 280, 18015-18024. Zhou, X.H., Brandau, O., Feng, K., Oohashi, T., Ninomiya, Y., Rauch, U., Fassler, R., 2003. The murine Ten-m/Odz genes show distinct but overlapping expression patterns during development and in adult brain. Gene Expr. Patterns 3, 397-405.

3 Chapter Three: Localization of TCAP-1-immunoreactivity in the adult mouse brain

This chapter is in submission for publication in Journal of Comparative Neurology

Chand D*, Bittencourt JC, Kollara A, Brown TJ, Lovejoy DA (2012) Localization of the Teneurin C-terminal Associated Peptide (TCAP-1) in the adult mouse (mus musculus) brain.

3.1 Abstract

Recently, the teneurins have emerged as an interneuronal and extracellular matrix signaling system that plays a significant role in brain development and neuronal communication. Encoded on the last exons of the teneurin genes are a new family of four bioactive peptides termed the

‘Teneurin C-terminal associated peptides’ (TCAPs). Previous in vivo studies indicate that TCAP-

1 blocks corticotropin-releasing factor (CRF)-related stress-induced behaviours, addiction behaviours and regulate corresponding neuronal morphological changes. Although TCAPs are highly conserved across the metazoan phylogeny, little is known about the distribution of TCAP-

1 in the mammalian brain. In this study I characterized the organization of immunoreactive-

TCAP-1-expressing cells in the mouse brain using immunohistochemical-staining and fluorescent-labelling with antiserum raised against mouse TCAP-1. Throughout the brain,

TCAP-1-immunoreactivity (TCAP-1-ir) was restricted to the cell soma and neuronal projections.

Strong TCAP-1-ir was detected in the external and internal granular layers of the cerebral cortex, purkinjie cells of the cerebellum and pyramidal neurons of the hippocampus. TCAP-1-ir was particularly strong in regions of the medulla, pons, midbrain and hypothalamus, but weak in the thalamus. Of particular interest was strong TCAP-1-labelling in limbic regions associated with stress, anxiety and addiction, such as the nucleus accumbens shell, piriform cortex, amygdala, bed nucleus of the stria terminalis and accessory olfactory regions. These results reveal

86 numerous shared features with the teneurins, but also important differences. A number of neuropsychiatric conditions have a common set of neurological substrates associated with the integration of sensorimotor processing and my results support the hypothesis that the teneurin-

TCAP-1 system might participate in the control and integration of these functions.

3.2 Introduction

The teneurins are a family of four type-II transmembrane proteins predominantly expressed in the central nervous system (CNS) (Tucker et al., 2007; Ben-Zur et al., 2000). They were originally discovered in Drosophila as odd oz (odz; Levine et al., 1994) on the basis of its expression pattern and tenascin-major (ten-m; Baumgartner et al., 1994), because of its structural homology to the tenascin proteins. Teneurin homologues have been found in all metazoans examined thus far, and have been functionally implicated in axon guidance (Young and Leamey,

2009), neurite outgrowth (Rubin et al., 1999; Leamey et al., 2008), transcriptional regulation

(Kenzelmann et al., 2008; Bagutti et al., 2003; Nunes et al., 2005), cell proliferation (Kinel-

Tahan et al., 2007) and adhesion (Rubin et al., 2002; Leamey et al., 2008).

The teneurins are highly conserved in metazoans and have been implicated in visual, auditory and olfactory integration in the CNS (Rubin et al., 2002; Wang et al., 2005;

Kenzelmann et al., 2007; 2008; Leamey et al., 2007; 2008; Young and Leamey, 2009), indicating that they may play a fundamental role in the evolution of sensorimotor integration (Tucker et al.,

2012). Given the complexity and highly conserved domain heterogeneity of the teneurin structure, it is not clear how all of these functions are regulated in both the developing and adult

CNS. Studies suggests that some family members may also exist as a number of splice variants

(Lossie et al., 2005) or as smaller soluble proteins derived from further proteolytic cleavage of

87 the carboxy terminal regions (Kenzelmann et al., 2008). One such region is a 40- to 41-amino acid sequence at the tip of the carboxy terminus of the teneurin proteins that possess characteristics of a cleavable bioactive peptide (Qian et al., 2004; Wang et al., 2005) and shares structural and physiological properties with the corticotropin-releasing factor (CRF) family of peptides (Lovejoy et al., 2006). This sequence is referred to as the ‘teneurin C-terminal associated peptide’ (TCAP).

TCAP-1 mRNA, is highly expressed in regions of the forebrain and limbic system, including the hippocampus, amygdala, cerebellum, hypothalamus, and cortex (Wang et al., 2005) and has recently emerged as a novel candidate in the integration of a number of psychiatric disorders (Rotzinger et al., 2010) such as CRF-mediated stress-induced behaviours (Al Chawaf et al., 2007b), anxiety (Tan et al., 2008) and addiction (Kupferschmidt et al., 2011). Psychiatric disorders encompass a broad suite of symptoms but common to these conditions is the aberrant processing of sensorimotor integration (Bolmont et al., 2002; Jovanvic et al., 2009; Canbeyli,

2010; Epstein et al., 2011). The elucidation of a common neurological system to link these disorders is of considerable interest and has classically focused on neurotransmitter systems

(Millan, 2006; Detera-Wadleigh and Akula, 2001; Drago et al., 2011). However, during the last decade the role of adhesion and juxtacrine signaling systems has garnered much attention (Costa et al., 2001; Bauer et al., 2002; Miguel-Hidalgo et al., 2011). Recent evidence indicates that the

TCAP-1 region of the teneurin-1 may act as a novel neuroplastic regulatory factor in the rodent brain (Tan et al., 2011b), through a dystroglycan-associated signaling mechanism to regulate cytoskeletal elements as a prelude to the changes associated with TCAP-mediated neuronal morphology and behaviours (Chand et al., 2012).

While TCAP-1 has been to shown to possess functions that are independent of the larger teneurins (Tan et al., 2011b), little is known about the expression of TCAP-1 in the CNS. If

TCAP-1 is structurally distinct from the teneurin-1, and possesses its own complement of functional attributes, it would be expected to have distinct sites of immunoreactivity and action on cells in the brain. In this study, I used antiserum specific for the TCAP-1 region of teneurin-1 and immunohistochemical techniques to characterize the localization of TCAP-1- immunoreactivity (TCAP-1-ir) in the adult mouse brain. A detailed description of the mouse- brain TCAP-1 network should provide an integrative framework that would aid in the interpretation of many of the TCAP-1-regulated psychiatric disorders.

3.3 Materials and Methods

3.3.1 Animals. Adult male BALB/c mice obtained from Charles River Laboratories (Montreal,

Quebec, Canada) were housed in a controlled environment with a 12 h light/dark cycle at a constant temperature of 21C. For immunehistochemical studies, BALB/c mice aged 6-8 weeks were killed by cervical dislocation and brain tissues were dissected and snap-frozen in liquid nitrogen or fixed overnight in 4% paraformaldehyde (PFA; Sigma-Aldrich Canada, Oakville,

ON, Canada), pH 7.4, and paraffin-embedded. All procedures were approved by the local animal care committee and were in accordance with the Canadian Council on Animal Care.

3.3.2 Synthesis of TCAP Peptides. Mouse TCAP-1, TCAP-2, and the fragments TCAP-1(1-8),

89 described (Wang et al., 2005). Two 25-amino acid peptides spanning either the mouse C- terminal sequence TCAP-1 (C-TCAP-1) or the N-terminal TCAP-1 (N-TCAP-1), were synthesized and conjugated with keyhole limpet hemocyanin (KLH) in a 1:1 ratio (w/w) by the

Dalton Chemical Company (Toronto, ON, Canada). The peptides were solubilized by exposure to ammonium hydroxide vapors for 2 min before dilution in phosphate-buffered saline (PBS) pH

7.4 with 10 mM sodium phosphate.

3.3.3 Antibody Characterization and Controls. The sequences used as haptens were KLH- pEQLLGTGRVQGYDGYFVLSVEQYLE-OH (pE: pyroglutamic acid) and KLH-

VLSVEQYLELSDSANNIHFMRQSEI- NH2. The purity of fragments was measured by high performance liquid chromatography (HPLC) at over 80% using a Vydac C18 reverse-phase column. Two un-conjugated peptide sequences of N-TCAP-1 and C-TCAP-1 were used for antisera specificity studies. Two rabbits were each immunized with one of the conjugated peptides. The pre-immune sera were collected to assess the background immunoreactivity before immunization. The conjugated peptides were emulsified with Freund’s complete adjuvant (FCA) in 1:1 ratio (V/V) at a final peptide concentration of 100 μg/ml. Four weeks after the injection, rabbits received a booster injection of peptides with Freund’s incomplete adjuvant. Periodic blood samples were collected to assess antibody titer. Booster injections were repeated until antibody titers were sufficient as determined by dot and western blotting.

450 nm. The competition curves and 50% binding values were calculated with GraphPad Prism 4

(San Diego, CA, USA) software using variable slope sigmoidal regression and F test to compare the 50% binding values.

Competition studies were carried out involving a comparison of staining patterns and intensities achieved using TCAP-1 TNR308 antiserum that was pre-incubated (overnight at 4°C) with 0–250 µM synthetic TCAP-1 peptide. Another method used to search for specificity of the antiserum was titering the antibody to the point at which it just barely stains the brain tissue (as described below) and then blocking with 50 µM/ml of TCAP-1 peptide (in the diluted antiserum), using a 1:200 – 1:2000 dilution. The pre-adsorbed antisera were used in all experiments as controls.

3.3.4 Immunofluorescent and immunohistochemical staining. For immunohistochemical analysis, paraformaldehyde-fixed paraffin-embedded sagittal and coronal sections (5 µm) of adult mice brains were deparaffinized in xylene and rehydrated. The sections were rinsed in double-distilled

91 water and placed in 10mM citrate buffer for antigen retrieval by heating in a 750W microwave oven 4 x 5 min with gentle agitation between each period. Sections were permeabilized with

0.3% Triton X-100 (Fisher Scientific) in PBS for 10 min and endogenous peroxidase was inactivated by incubation in 0.3% H2O2 in methanol for 20 min. The tissue sections were blocked with 10% NGS (Vector Laboratories) in PBS for 1 h and incubated overnight at 4˚C with affinity-purified TNR308 TCAP-1 antiserum (1:1000) in PBS containing 1% NGS. After washing in PBS, sections were incubated with biotinylated goat anti-rabbit immunoglobulin

(IgG) antibody (1:200; Vector Laboratories) in PBS containing 1% NGS for 1 hr, followed by horseradish peroxidase-streptavidin conjugate (1:400; Vector Laboratories) in PBS for 30 min.

To visualize staining, the sections were incubated with 0.07% 3,3-diaminobenzidine (Sigma; St.

Louis, MO) and 0.03% H2O2 in PBS for 12 min and counterstained with hematoxylin (Sigma).

For immunofluorescent analysis, sections were incubated instead with Alexa 594 goat anti-rabbit

IgG secondary antibody (1:400; Invitrogen) in 0.01 M PBS for 1 h at RT and counterstained with

DAPI (Sigma-Aldrich Canada) for 1 min. Sections were mounted with DABCO (Sigma-Aldrich

Canada). I have followed the parcellation as described by Paxinos and Franklin in “The Mouse

Brain in Stereotaxic Coordinates, Second Edition” (2001) to aid in the histological analysis.

3.3.5 Imaging. Diaminobenzidine-stained sections were analyzed using an Aperio ScanScope XT

(Vista, CA, USA) equipped with an Olympus 40x/0.75NA UPlanFL N objective (Shinjuku,

Tokyo, Japan). Sections were scanned to a resolution of 0.25 microns/pixel, and images visualized using Aperio ImageScope Version 10. Fluorescently-labelled tissue sections were analyzed using a Leica DMRXE fluorescent microscope (Leica Microsystems, Wetzlar,

Germany) with a Sony DXC-970MD 3CCD camera (Minato, Tokyo, JapanJapan). The resulting

92 images were adjusted for brightness and contrast using Eclipse V8 (Empix Imaging, Inc,

Mississauga, ON, Canada) and Adobe Photoshop CS4 Extended (San Jose, CA, USA).

3.4 Results

3.4.1 Localization of TCAP-1-immunoreactivity in the mouse brain

The distribution of TCAP-1-ir in the adult mouse brain was widespread but mainly restricted to cytosolic compartments and neuronal projections. TCAP-1-ir was detected in the cortex (Fig. 3.1), cerebellum (Fig. 3.2), pons (Fig. 3.3), midbrain (Fig. 3.4), medulla (Fig. 3.5), thalamus (Fig. 3.6), hypothalamus (Fig. 3.7), hippocampus (Fig. 3.8) and limbic regions (Fig. 3.9 and 3.10).

3.4.1.1 Cortex. In the cortex, strong TCAP-1-ir was detected in the ventral orbital cortex (VO;

Fig. 3.1A) and mediomedial area of the secondary visual cortex (V2MM; Fig. 3.1K), whereas, moderate immunoreactivity was observed in the mediolateral area of the secondary visual cortex

(V2ML; Fig. 3.1I), medial parietal association cortex (MPtA; Fig. 3.1G) and primary motor cortex (M1; Fig. 3.1E). TCAP-ir was comparatively weaker in the secondary motor cortex (M2;

Fig. 3.1C). Within the cortical layers, TCAP-1-ir was absent in the molecular layer (layer 1) but strongest in the external granular layer (layer 2) and internal granular layer (layer 4) of the MPtA

(Fig. 3.1G) and V2ML (Fig. 3.1I). In comparison, moderate TCAP-1-ir was detected in the external pyramidal layer (layer 3), internal pyramidal layer (layer 5) and multiform layer (layer

6) of the cerebral cortex (Fig. 3.1). Preadsorption with synthetic TCAP-1 significantly diminished staining of TCAP-1-ir in cells of the cortex (Fig. 3.1B,D,F,H,J,L) as compared with the control.

Figure 3.1. Distribution of TCAP-1-immunoreactivity in the cerebral cortex. TCAP-1-ir was detected in the ventral orbital cortex (VO) (A), secondary motor cortex (M2) (C), primary motor cortex (M1) (E), medial parietal association cortex (MPtA) (G), mediolateral area of the secondary visual cortex (V2ML) (I) and mediomedial area of the secondary visual cortex (V2MM) (K). TCAP-1-ir was absent in the molecular layer (layer 1) but strongest in the external granular layer (layer 2) and internal granular layer (layer 4) of the MPtA and V2ML. Moderate TCAP-1-ir was detected in the external pyramidal layer (layer 3), internal pyramidal layer (layer 5) and multiform layer (layer 6) of the cerebral cortex. Immunoreactivity was not observed in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D, F, H, J and L). Scale bars, 100μm.

3.4.1.2 Cerebellum. Strong TCAP-1-ir was detected in the purkinjie neurons of the cerebellum

(Fig. 3.2A) with sparse immunoreactivity in the granular layer and molecular layer of the cerebellum (Fig. 3.2A). Immunoreactive-TCAP-1 was strongest in the cell soma of the purkinjie neurons, with moderate immunoreactivity in the axon hillock (Fig. 3.2A). Similarly, TCAP-ir was exclusively localized to the soma of the neuronal cells in the interposed cerebellar nucleus

(Fig. 3.2C). Preadsorption with synthetic TCAP-1 significantly diminished staining of TCAP-1- ir in the purkinjie neurons (Fig. 3.2B) and cells of interposed cerebellar nucleus (Fig. 3.2D).

Figure 3.2. Localization of immunoreactive-TCAP-1 in the cerebellum. TCAP-1-ir was detected in the soma and axonal projections of the purkinjie neurons in the cerebellum (A). Weak TCAP-1-immunoreactivity was detected in the granular layer and molecular layer of the cerebellum (A). Strong TCAP-ir was also localized to the soma of the neuronal cells in the interposed cerebellar nucleus (C). TCAP-1-ir was significantly diminished in sections incubated with pre-adsorbed TCAP-1 antiserum (B and D). Scale bars, 50μm.

3.4.1.3 Pons. In the pons, strong TCAP-1-ir was observed in the soma and neuronal projections of cells of the pontine nuclei (Pn; Fig. 3.3A), oral pontine reticular nucleus (PnO; Fig. 3.3B) and the locus coeruleus (LC; Fig. 3.3C). However, immunoreactive-TCAP-1 was not detected in the

95 longitudinal fasciculus of the pons (lfp) nor the lateral lemniscus (ll) of the pons (Fig. 3.3A).

Preadsorption with synthetic TCAP-1 significantly diminished staining of TCAP-1-ir in cells of the Pn (Fig. 3.3B), PnO (Fig. 3.3D) and LC (Fig. 3.3F) as compared with the control.

Figure 3.3. Distribution of immunoreactive-TCAP-1 in the pons. Strong TCAP-1-ir was localized to soma of cells in the pontine nuclei (Pn) but was not detected in the longitudinal fasciculus of the pons (lfp) or the lateral lemniscus (ll) of the pons (A). Immunoreactive-TCAP-1 was observed in the soma and neuronal projections of cells of the oral pontine reticular nucleus (PnO) (B) and the locus coeruleus (LC) (C). Immunoreactivity was significantly diminished in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D and F). Scale bars, 150μm (A and B); 50μm (C-F).

3.4.1.4 Mid Brain. TCAP-1-ir in the mid brain was mainly localized to the substantia nigra, ventral tegmental area (VTA) and inferior colliculus (IC). In the substantia nigra pars reticulata

(SNR; Fig. 3.4A) TCAP-1-ir was sparse in comparison to the dense immunoreactive-TCAP-1- staining observed in the substantia nigra pars compacta (SNC; Fig. 3.4C).

Figure 3.4. Distribution of immunoreactive-TCAP-1 in the midbrain. In the substantia nigra, sparse TCAP-1-ir was detected in the substantia nigra pars reticulata (SNR) (A) whereas strong dense immunoreactive-TCAP-1-staining was observed in the substantia nigra pars compacta (SNC) (C). Strong TCAP-1-ir was detected in the cell soma and neuronal projections of cells in the ventral tegmental area (VTA) (E) and inferior colliculus (IC) (G). Immunoreactivity was significantly diminished in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D, F and H). Scale bars, 100μm.

Strong TCAP-1-ir was also detected in the cell soma and neuronal projections of cells in the

VTA (Fig. 3.4E) and IC (Fig. 3.4G). Preadsorption with synthetic TCAP-1 significantly diminished immunoreactive-TCAP-1 staining in the SNC (Fig. 3.4D) and VTA (Fig. 3.4F) and completely ablated TCAP-1-ir in the SNR (Fig. 3.4B) and IC (Fig. 3.4H) as compared with the control.

3.4.1.5 Medulla. In the medulla, strong TCAP-1-ir was detected in the facial nucleus (7N), with intense staining localized to both the cell soma and neuronal projections (Fig. 3.5A). Similar

TCAP-1-staining was also observed in cells of the corresponding facial nerve (7n) (data not shown). Moderate TCAP-1-ir was observed in cells of the inferior olive (IO; Fig. 3.5C), and comparatively less dense staining observed in cells of the spinal vestibular nucleus (SpVe; Fig.

3.5E). Preadsorption with synthetic TCAP-1 significantly diminished staining of TCAP-1-ir in cells of the 7N (Fig. 3.5B), IO (Fig. 3.5D) and SpVe (Fig. 3.5F) as compared to the control.

Figure 3.5. Distribution of immunoreactive-TCAP-1 in the medulla. Strong immunoreactive- TCAP-1-staining was detected in the soma and neuronal projections of cells in the facial nucleus (7N) (A). Strong TCAP-1-ir was also observed in the soma of cells of the inferior olive (IO) (C) and spinal vestibular nucleus (SpVe) (E). Immunoreactivity was not observed in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D and F). Scale bars, 100μm.

3.4.1.6 Thalamus and Hypothalamus. In the thalamus, weak TCAP-1-ir was detected in the cytosol of cells in the mediorostral part of the lateral posterior thalamic nucleus (LPMR; Fig.

3.6A), laterodorsal thalamic nucleus (LD; Fig. 3.6C) and ventral posterolateral thalamic nucleus

(VPL; Fig. 3.6E), whereas, moderate TCAP-1-ir was observed in cells of the ventral posteromedial thalamic nucleus (VPM; Fig. 3.6E). In contrast to the thalamus, strong dense

TCAP-1-ir was observed in the cell soma and neuronal projections of cells in the hypothalamic regions, such as the ventromedial hypothalamic nucleus (VMH), lateral hypothalamic area (LH;

Fig. 3.7A), perifornical nucleus (PeF; Fig. 3.7C) and medial preoptic area (MPA; Fig. 3.7E).

Preadsorption with synthetic TCAP-1 significantly diminished staining of TCAP-1-ir in both the thalamic (Fig. 3.6B,D,F) and hypothalamic (Fig. 3.7B,D,F) regions as compared with the control.

Figure 3.6. Localization of immunoreactive-TCAP-1 in the thalamus. Weak TCAP-1-ir was observed in the soma of cells in the mediorostral part of the lateral posterior thalamic nucleus (LPMR) (A), laterodorsal thalamic nucleus (LD) (C) and ventral posterolateral thalamic nucleus (VPL) (E). Moderate immunoreactive-TCAP-1-staining was detected in cells of the ventral posteromedial thalamic nucleus (VPM) (E). Immunoreactivity was not observed in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D and F). Scale bars, 100μm.

Figure 3.7. Localization of immunoreactive-TCAP-1 in the hypothalamus. Strong TCAP-1-ir was observed in the cell soma and neuronal projections of cells in the lateral hypothalamus (LH) (A), perifornical nucleus (PeF) (B) and medial preoptic area (MPA) (C). Immunoreactivity was not observed in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D and F). Scale bars, 100μm (A, B, E and F); 50μm (C and D).

3.4.1.7 Hippocampus. TCAP-1-ir was localized to the two main neuronal cellular layers of the hippocampus, the granular layer of the dentate gyrus and the pyramidal layer of the CA1, CA2 and CA3 (Fig. 3.8). The distribution of TCAP-1-ir was strong in the pyramidal layer (Py) of the

CA1 (Fig. 3.8A), CA2 (Fig. 3.8C) and CA3 (Fig. 3.8E) and weak in the stratum lucidum (SL).

Weak TCAP-1-ir was also detected in the granular layer of the dentate gyrus (Fig. 3.8G).

Immunoreactivity was absent in sections incubated with pre-adsorbed TCAP-1 antiserum (Fig.

3.8B,D,F,H).

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Figure 3.8. Localization of immunoreactive-TCAP-1 in the hippocampus. Strong TCAP-1-ir was detected in the cytosol of the neuronal layers of the hippocampus (A, C, E and G). TCAP-1- ir was strongest in the pyramidal layer of the CA2 (C) and CA3 regions (E), moderate in the CA1 region (A) and weak in the dentate gyrus (G). TCAP-1-ir was also observed in the stratum lucidum of the CA1 (A) and CA2 (C) regions of the mouse hippocampus. Immunoreactivity was significantly diminished in sections incubated with pre-adsorbed TCAP-1 antiserum (B, D, F and H). Scale bars, 25μm.

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3.4.1.8 Limbic Regions. In addition to TCAP-1-ir in the hippocampus, TCAP-1 was also localized to various regions that comprise the limbic system (Fig. 3.9 and 3.10). In the bed nucleus of the stria terminalis (BST), immunoreactive-TCAP-1 staining was weak in the anterior part of the medial division of the BST (BSTMA; Fig. 3.9A). However, strong TCAP-1-ir was localized to the soma of cells in the posterior part of the medial division of the BST (BSTMP;

Fig. 3.9B).

Figure 3.9. Localization of immunoreactive-TCAP-1 in the bed nucleus of the stria terminalis. Weak TCAP-1-ir was detected in the anterior part of the medial division of the bed nucleus of the stria terminalis (BSTMA) (A), whereas strong TCAP-1-ir was localized to the soma of cells in the posterior part of the medial division of the bed nucleus of the stria terminalis (BSTMP) (B). Immunoreactivity not observed in sections incubated with pre-adsorbed TCAP-1 antiserum (B and D). Scale bars, 100μm.

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Figure 3.10. Distribution of immunoreactive-TCAP-1 in the limbic regions. Sections fluorescently-labelled with TCAP-1 antiserum (red) showed strong cytosolic TCAP-1-ir in cells of the the nucleus accumbens shell (AcbSh) (A) but weak and sparse labelling in the the nucleus accumbens core (AchC) (B). Strong cytosolic labelling was also detected in piriform cortex (Pir) (C), ventral anterior olfactory nucleus (AOV) (E), lateral anterior olfactory nucleus (AOL) (G) and posterolateral cortical amygdaloid nucleus (PLCo) (I). Moderate TCAP-1-ir was detected in cells of the medial anterior olfactory nucleus (AOM) (H) and accessory olfactory bulb (AOB) (F) with sparse TCAP-1-labelling in the lateral septal nucleus (LS) (D). Scale, 50 μm (A-G); 150 μm (H and I).

In addition, fluorescent-labelling with TCAP-1 TNR308 antiserum on coronal sections of the mouse brain showed strong TCAP-1-ir in cells of the nucleus accumbens shell (AcbSh; Fig.

3.10A), piriform cortex (Pir; Fig. 3.10C), ventral anterior olfactory nucleus (AOV; Fig. 3.10E), lateral anterior olfactory nucleus (AOL; Fig. 3.10G) and posterolateral cortical amygdaloid nucleus (PLCo; Fig. 3.10I). In the AOL, TCAP-1-ir was observed to be strongest in cells that are

103 located adjacent to the intrabulbar part of the anterior commissure (aci; Fig. 3.10G). Moderate

TCAP-1-ir was detected in cells of the medial anterior olfactory nucleus (AOM; Fig. 3.10H) and accessory olfactory bulb (AOB; Fig. 3.10F) with comparatively less labelling in the lateral septal nucleus (LS; Fig. 3.10D). In the nucleus accumbens core (AchC) TCAP-1-ir was sparse, with only few cells showing weak TCAP-1-labelling (Fig. 3.10B). Although TCAP-1-ir was localized to some regions that are also associated with the basal ganglia, TCAP-1-ir was not found in the caudate putamen (CPu) or the ventral pallidum (VP) (data not shown). Preadsorption with synthetic TCAP-1 completely ablated TCAP-1 staining in the BST (Fig. 3.9B,D) and fluorescently-labelled sections (data not shown).

3.5 Discussion

When the TCAP sequences were first discovered, they were annotated as being part of the last exon in the teneurin genes. Structurally, the TCAP family of peptides possess an amidation motif at the carboxyl-terminus and a pyroglutamic acid residue at the amino-terminus

(Wang et al., 2005), features that are characteristic of bioactive peptides. The expression of the region of the teneurin-1 gene that encodes for TCAP-1 and the characterization of TCAP-1-ir shows that the distribution of teneurin-1 in the vertebrate brain is far more widespread than previously reported. The majority of immunoreactive-teneurin-1 in the brain is localized to the cell membrane and neuronal processes (Zhou et al., 2003; Li et al., 2006; Kenzelmann et al.,

2008) whereas the majority of TCAP-1-ir was found to be cytosolic. The characterization of

TCAP-1 in the mouse brain adds to the growing body of evidence that the TCAP-1 region of teneurin-1 can be both structurally and functionally independent of the larger teneurin-1 (Chapter

2; Tan et al., 2011b; Chand et al., 2012b). This is thought to be due in part to region specific

104 processing of the C-terminal region of teneurin-1 (Kenzelmann et al., 2008). Notably, TCAP-1-ir appeared to be strongly immunoreactive in regions associated with the control and integration of sensorimotor function (Fig. 3.11).

3.5.1 Methodological considerations

The TCAP-1 antiserum was generated in rabbit against a mouse antigen. The adsorption tests and binding specificity of the TCAP-1 antibody presented a reliable outcome showing that the antiserum recognized the antigen specifically. In preadsorption experiments of TCAP-1 antiserum with the corresponding protein, immunoreactivity was completely ablated in most brain regions. In some regions low TCAP-1 immunostaining was still present but was significantly diminished as compared to control. This residual labelling might be due to incomplete blocking of the TCAP-1 antibody after preadsorption since these regions showed intense immunoreaction in sections treated without antibody preadsorption. Alternatively, residual TCAP-1-staining may be due to cross-reactivity with the other three TCAP homologs.

However, this is less likely, as the antibody showed strong binding affinity to TCAP-1 as opposed to TCAP-2. Although all teneurins have a potential dibasic cleavage motif near the C- terminus that would generate a TCAP peptide, only TCAP-1 mRNA is independently expressed in the adult brain (Chapter 2; Chand et al., 2012b), whereas TCAP-2, TCAP-3 and TCAP-4 are all transcribed as part of the larger teneurin genes (C.A Cassati and D.A Lovejoy, unpublished findings).

Additionally, the antiserum labeled the pyramidal neurons of the hippocampus strongly, the exact same region where TCAP-1 binding sites have been identified (Chapter 4; Chand et al.,

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2012) and where TCAP-1 is biologically active (Tan et al., 2009). Unfortunately, no comparative studies of immunoreactive TCAP-labelling have been reported to date.

3.5.2 Comparison of TCAP-1 and teneurin localization in the vertebrate brain

Teneurins are a novel family of transmembrane proteins implicated in neuronal plasticity, pattern formation and morphogenesis (Young and Leamey, 2009). The expression and function of teneurins has been extensively studied in the CNS of several species and these studies suggest that each of the genes has its own distinct developmental roles that are evolutionarily conserved throughout the metazoans (Mieda et al., 1999; Tucker and Chiquet-Ehrismann, 2006; Tucker et al., 2007; Kenzelmann et al., 2008; Young and Leamey, 2009; Tucker et al., 2012).

3.5.3 Cerebral Cortex

The cerebral cortex is composed of areas that are anatomically and functionally distinct, recent findings suggest an important role for the teneurin family of proteins in establishing cortical arealization and patterning in the developing brain (Beckman et al., 2011). All four teneurins are each expressed in distinctive rostro-caudal gradients within the cortical plate of the cerebral cortex. The cortical expression gradient of teneurin-1 is rostral high and caudal low, while cortical expression gradients of the other teneurin members are rostral low and caudal high

(Li et al., 2006). In the developing mouse brain, teneurin-1 was mainly expressed in layer IV, teneurin-2 in layer V, teneurin-3 in layers IV and V and teneurin-4 in layers II–V with the most extensive expression in layer V (Li et al., 2006).

Similarly, all teneurin mRNAs were found to be expressed throughout the cerebral cortex of the adult mouse brain with strong signals between the layer II and layer VI, while no signals

106 were reported in layer I (Zhou et al., 2003). Consistent with the expression of teneurin mRNAs, immunoreactive TCAP-1-staining was heterogeneously distributed throughout the cerebral cortex, with strong staining in layer II and layer IV, comparatively weaker staining in the layer

III, layer V and layer VI and a lack of staining in layer I. While the teneurin-1 proteins have been reported in neurons of the adult mouse cerebral cortex (Oohashi et al., 1999), the precise distribution within the cortical layers has not been clearly established. However, given the strong correlation between teneurin-1 expression and TCAP-1 immunoreactivity in the cortex, TCAP-1 may be acting as part of a direct teneurin-1 function to mediate neuronal plasticity and synapse formation (Al Chawaf et al., 2007a).

3.5.4 Hindbrain: Cerebellum, Medulla and Pons

Similar to TCAP-1 mRNA expression (Wang et al., 2005), TCAP-1-ir was strongly localized to the soma and neuronal projections of the Purkinje cell layer in the cerebellum.

Interestingly, teneurin-1 mRNA expression was absent from Purkinje cells but present exclusively in the granular layer (Zhou et al., 2003), where only weak TCAP-1-staining was observed. This result suggests, that TCAP-1 may be transcribed independently of the teneurin-1 gene in the mouse cerebellum. This is further supported by the lack of teneurin-1- immunoreactivity in the Purkinje cell layer, intense teneurin-1-staining in the molecular layer and weaker staining around cells in the granular layer (Oohashi et al., 1999; Zhou et al., 2003). I also observed strong TCAP-1-ir in the interposed cerebellar nucleus, but only teneurin-1 has been reported to be present in the cerebellar nuclei (Oohashi et al., 1999) and has not been fully characterized.

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In comparison to the other teneurin homologs, teneurin-2, -3 and -4 mRNA transcripts were all detected in the Purkinje cell layer, in addition to teneurin-2 expression in the molecular layer, and teneurins-3 and -4 expression in the white matter of the mouse cerebellum (Zhou et al., 2003). However, despite mRNA expression in the Purkinje cell layer, all teneurin proteins were found to be mainly localized to the molecular layer and granular layer, with the addition of weak teneurin-3-labelling in the Purkinje cell layer and white matter of the mouse cerebellum

(Zhou et al., 2003). TCAP-1-ir was not detected in the white matter of the mouse cerebellum. My results show important differences in cerebellar distribution of TCAP-1 and the teneurins, further supporting the hypothesis that TCAP-1 can, in some cases, be both structurally and functionally independent of the teneurins.

In the medulla oblongata, strong TCAP-1-ir was detected in the cell soma and neuronal projections of the facial nucleus, inferior olive spinal vestibular nucleus. Teneurin-1 expression has not been determined in the medulla oblongata, but teneurin-2 (Zhou et al., 2003) and teneurin-3 (Ben-zur et al., 2000) mRNAs have been reported in the rostral portion of the medulla oblongata in the developing brain. Interestingly, teneurin-4 is not expressed in the medulla oblongata (Ben-zur et al., 2000). The detection of strong TCAP-1-ir in the medulla oblongata may be accounted for by the fact that teneurin-1 is only expressed later in brain development and most prominently in adult. Alternatively, TCAP-1 may be expressed independently of teneurin-1, similar to that observed in other brain regions, such as the hippocampus where TCAP-1 has already been shown to possess functions that are independent of the larger teneurins (Wang et al., 2005; Tan et al., 2011; Ng et al., 2012).

Only teneurin-3 and teneurin-4 has been reported to be expressed in the pons (Ben-zur et al., 2000). This study shows for the first time that the C-terminal region of teneurin-1 is strongly

108 localized in the pontine nuclei, oral pontine reticular nucleus and locus coeruleus of the pons. My finding of strong TCAP-1-ir in the pons is consistent with the localization of TCAP-1 in integrative and relay centers of the brain. The pontine nuclei projects to the cerebellar cortex and cerebellar nuclei, receives direct input from the cerebral cortex and forms the key relay center in the cerebropontocerebellar pathway mediating cerebrocerebellar communication (Allen and

Tsukahara 1974; Mihailoff et al., 1978). A key finding of strong TCAP-1-ir in the locus coeruleus, an important homeostatic control center involved in mediating stress (Benarroch,

2009) and addiction (Devenyi et al., 1982) behaviours is also consistent with previous investigations that showed TCAP-1 blocks CRF-mediated stress-induced behaviours (Al Chawaf et al., 2007b), anxiety (Tan et al., 2008) and cocaine-reinstatement (Kupferschmidt et al., 2011) in rats.

3.5.5 Midbrain

In the midbrain, TCAP-1-ir was particularly strong and dense in the inferior colliculus, substantia nigra pars compacta and ventral tegmental area. The midbrain is involved in the integration and relay of multi-modal sensory perception, and TCAP-1 has been shown to regulate the acoustic startle response in rats, which is integrated in particular by the inferior colliculus

(Wang et al., 2005). The distribution of teneurin proteins in the midbrain remains unresolved, but the expression of teneurin mRNAs has been reported in the developing midbrain of zebrafish

(Mieda et al., 1999), chicken (Rubin et al., 2002) and mouse (Wang et al., 1998; Zhou et al.,

2003).

In the midbrain, teneurin mRNAs are strongly expressed in a complimentary, non- overlapping subset of neurons or sub-regions similar to that observed for TCAP-1 staining. In

109 both mouse and zebrafish, teneurin-3 is expressed throughout the entire midbrain, except for the most caudal part, whereas, teneurin-4 is strongly expressed at the most caudal part of the midbrain, marking the midbrain-hindbrain boundary (Mieda et al., 1999; Wang et al., 1998;

Zhou et al., 2003). In mouse, teneurin-2 was mainly expressed at the roof the midbrain (Zhou et al., 2003), analogous to the strong expression of chicken teneurin-1 (Kenzlemann et al., 2008) and teneurin-2 (Rubin et al., 2002) in the optic tectum. In contrast to the other teneurins, teneurin-1 mRNA is only detected after embryonic day 15.5 in mouse, and is predominantly expressed in the dorsal part of the midbrain, with moderate expression extending to the caudal regions (Zhou et al., 2003), consistent with that observed for TCAP-1-staining. All four teneurins are known to form homophilic and heterophilic dimers at the C-terminal regions (Oohashi et al.,

1999; Feng et al., 2002; Bagutti et al., 2003). Thus, the complimentary teneurin-TCAP-1 expression patterns maybe part of a network that mediates synapse formation and neuronal communication between subsets of neurons or sub-regions within the midbrain.

3.5.6 Thalamus and Hypothalamus

TCAP-1-ir was weak in the thalamus as compared to other regions of the mouse brain.

Moderate immunoreactivity was detected in the ventral posteromedial thalamic nucleus but

TCAP-1-staining was weakest in the dorsal thalamic nuclei of the thalamus. Little is known about the distribution of the teneurin proteins in the thalamus, but all four teneurin mRNAs are highly expressed in the thalamus of the developing mouse brain (Li et al., 2006). However, only tenueurin-1 and teneurin-4 mRNAs are expressed in the thalamus of the adult mouse (Zhou et al., 2003). At embryonic day 15.5, teneurin-1 is the only teneurin gene expressed in the ventroposterior nucleus of the dorsal thalamus (Li et al., 2006), an area that shows weak TCAP-

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1-ir. In contrast, teneurin-2, teneurin-3 and teneurin-4 are mainly expressed in the ventral subdivision of medial geniculate nucleus and posterior complex of the dorsal thalamus (Li et al.,

2006).

Interestingly, the expression and localization of the teneurins may be age-dependent. At postnatal day 2, teneurin-1 is only weakly expressed in the ventroposterior nucleus of the dorsal thalamus but relatively stronger expression is detected in the ventral posterior thalamic nuclei (Li et al., 2006), similar to that observed with immunoreactive-TCAP-1-staining in the thalamus. In contrast to teneurin-1 mRNA expression, teneurin-2, teneurin-3 and teneurin-4, are all heavily expressed in both dorsal lateral geniculate nucleus and posterior complex of the dorsal thalamus

(Li et al., 2006). These results suggest that, unlike in the cerebellum, TCAP-1 may be expressed as part of the larger teneurin-1 in the thalamus.

TCAP was first discovered in the Onchorynchus mykiss hypothalamus (Qian et al.,

2004), and TCAP-1 mRNA has been reported in the ventromedial nucleus of the hypothalamus and subthalamic nucleus in the rat (Wang et al., 2005). This is consistent with my findings of strong TCAP-1-ir in the lateral hypothalamic area, perifornical nucleus and medial preoptic area.

Teneurin proteins have not been reported in the mouse hypothalamus, and teneurin-1 expression in the hypothalamus was not detected in either embryonic or postnatal mice (Li et al., 2006) whereas, only sub-regions of hypothalamus are stained positive for teneurin-2, teneurin-3, and teneurin-4 (Ben-zur et al., 2000; Li et al., 2006). The lack of teneurin-1 mRNA but strong

TCAP-1 mRNA and TCAP-1-ir in the hypothalamus supports previously reported observations that TCAP-1 can exist and possess functions that are independent of the larger teneurins (Wang et al., 2005; Tan et al., 2011; Ng et al., 2012).

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3.5.7 Hippocampus

TCAP-1 is immunoreactive in all cell groups of the mouse hippocampus that show both teneurin-1 (Zhou et al., 2003) and TCAP-1 mRNA expression (Wang et al., 2005). However, the expression pattern of teneurin-1 mRNA does not coincide with the localization pattern of the protein in the mouse hippocampus (Zhou et al., 2003). Teneurin-1 proteins were mainly localized within the molecular layers (Oohashi et al., 1999) with weak immunoreactivity in the stratum pyramidale, whereas the mRNAs were specifically confined to the cell somata of the stratum pyramidale and the granular layer of the dentate gyrus (Zhou et al., 2003). TCAP-1 on the other hand was strongly localized to stratum pyramidale, with weak immunoreactivity in the stratum lucidum of the mouse hippocampus. Likewise, it has been shown that the expression pattern of

Drosophila teneurin mRNA does not coincide with the protein (Baumgartner et al., 1994) similar to that of other neural cell adhesion molecules, such as L1 (Jucker et al., 1996).

These studies show that TCAP-1-immunoreactivity corresponds to TCAP-1 mRNA

(Wang et al., 2005) and teneurin-1 mRNA expression (Zhou et al., 2003) in the hippocampus, but differs markedly from that of teneurin-1-immunoreactivity both in vitro and in vivo

(Baumgartner et al., 1994; Oohashi et al., 1999; Zhou et al., 2003). This further supports a mechanism by which TCAP-1 is independently processed from teneurin-1 and/or cleaved from the C-terminal region by proteolytic cleavage (Rubin et al., 1999).

3.5.8 Limbic Regions

In addition to TCAP-1-ir (Chapter 2; Chand et al., 2012b) and TCAP-1 binding (Chapter

4; Chand et al., 2012) in the pyramidal neurons of the hippocampus, strong TCAP-1-ir was also detected in other brain regions associated with the limbic system (Fig. 3.11). The localization of

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TCAP-1-ir to the the bed nucleus of the stria terminalis, nucleus accumbens, piriform cortex, olfactory nuclei, amygdala, accessory olfactory bulb and lateral septal nucleus is consistent with

TCAP-1 (Wang et al., 2005) and teneurin-1 (Kenzelmann et al., 2008) mRNA expression in these regions. Furthermore, this study also supports previous in vivo studies that show TCAP-1 is highly active in regulating CRF-mediated stress-induced behaviours (Al Chawaf et al., 2007b;

Tan et al., 2009) and addiction-like behaviours (Kupferschmidt et al., 2011) that are associated with these brain regions. CRF can dose-dependently increase c-Fos in the brain (Bittencourt and

Sawchenko, 2000) and TCAP-1 administration has been shown to attenuate CRF-induced c-Fos expression in the hippocampus, amygdala, medial prefrontal cortex, lateral and medial septum, and dorsal raphe nucleus in the adult male rat brain (Tan et al., 2009).

The limbic system is well characterized as an interconnected region associated with integrating and regulating sensory information related to stress, emotionality, fear, reward and addiction (Herman et al., 2005). While the teneurins, particularly teneurin-1 has been implicated in mediating neuronal connectivity in these areas (Kenzelmann et al., 2008), this study supports the hypothesis that TCAP-1 and the teneurins may play a fundamental role in the regulation of stress-related behaviours, anxiety and addiction (Rotzinger et al., 2010; Tan et al., 2011b).

3.5.9 Integrating TCAP-1-regulated psychiatric disorders and sensorimotor function

Teneurin-1 and TCAP-1 share many similar functions in regulating neuronal development and plasticity, and this study shows that they are also distributed in many of the same regions in the mouse brain. In this case, TCAP-1 may function as part of the larger teneurin-1 protein. However, in some regions like the cerebellum and hippocampus, the TCAP-1

113 region of teneurin-1 can also exist independently from the larger 2800 amino acid transmembrane protein.

One of the most important findings in this study was the strong localization of TCAP-1 in areas associated with the integration and regulation of sensorimotor processing and behaviours, such as the cerebral cortex, cerebellum, hippocampus, locus coeruleus, hypothalamus and limbic regions. TCAP-1 effects on CRF-associated behaviours are long-lasting, with effects persisting

21 days after TCAP-1 injection (Wang et al., 2005). Moreover, acutely-administered TCAP-1 attenuates CRF-induced c-Fos accumulation in the limbic system (Tan et al., 2009). Although this study shows the localization of TCAP-1 in areas that also express CRF receptors

(Bittencourt and Sawchenko, 2000; Chalmers et al., 1995; van Pett et al., 2000), TCAP-1 does not appear to regulate the hypothalamic-pituatary adrenal axis through CRF receptor signaling

(Wang et al., 2005). Instead, TCAP-1 regulates cytoskeletal elements (Al Chawaf et al., 2007a;

Chand et al., 2012) and modulates dendritic arborization (Tan et al., 2011) suggesting that the

TCAP-mediated regulation of addiction, stress and anxiety-like behaviours may be associated with the remodeling of neuronal networks and dendritic morphology. This is consistent with role of the teneurins in modulating neuronal plasticity (Tucker et al., 2007).

The brain regions that show strong TCAP-1-ir are also responsive in vivo to TCAP-1 treatment (Tan et al., 2009). More importantly, these areas such as hippocampus, inferior colliculus, medial prefrontal cortex, locus coeruleus and hypothalamus have also been characterized as areas of high neuronal activity and correspondingly high metabolic activity

(Sokoloff, 1981), thus making them susceptible to metabolic insults and homeostatic challenges.

It has been previously established that TCAP-1 is neuroprotective against alkalotic stress

(Trubiani et al., 2007), inhibits brain-derived neurotrophic factor (BDNF) expression and

114 translation in hypothalamic neurons (Ng et al., 2012) and increases actin stress-fibers in hippocampal cells (Chapter 4; Chand et al., 2012). Therefore, the widespread distribution of

TCAP-1 in these highly active areas and other brain regions maybe part of a unique family of peptides that serve a broader function in maintaining homeostasis, neuroprotection and resistance to metabolic insults.

This study provides novel information on the distribution of the TCAP-1 region of teneurin-1 in the adult mouse brain (Fig. 3.11). My study not only adds to a growing body of evidence of dynamic teneurin-1 expression in the vertebrate brain but also implicates the teneurins in a broader range of neuroendocrine and sensorimotor functions. Moreover, the characterization of TCAP-1 distribution in the brain helps to provide a better understanding of the mechanisms involved in TCAP-1-regulated psychiatric disorders (Wang et al., 2005; Al

Chawaf et al., 2007b; Tan et al., 2009; Rotzinger et al., 2010; Tan et al., 2011; Kupferschmidt et al., 2011). A number of neuropsychiatric conditions have a common set of neurological substrates associated with the integration of sensorimotor processing and my results support the hypothesis that the teneurin-TCAP-1 system might participate in the control and integration of these functions.

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Figure 3.11. Diagrammatic representation of the distribution of immunoreactive-TCAP-1 in the adult mouse brain. Immunoreactive-TCAP-1 was localized to the cell soma and neuronal projections in various regions throughout the adult mouse brain. In the cerebral cortex, TCAP-1 was localized to the ventral orbital cortex (VO), secondary motor cortex (M2), primary motor cortex (M1), medial parietal association cortex (MPta), mediolateral area of the secondary visual cortex (V2ML), mediomedial area of the secondary visual cortex (V2MM), retrosplenial agranular cortex (RSA) and retrosplenial granular cortex (RSG). In the cerebellum (Cb), TCAP- 1-ir was mainly localized to the purkinjie neurons and interposed cerebellar nucleus (IntP). In the medulla, strong TCAP-1-ir was localized to cells within the spinal vestibular nucleus (SpVe), facial nucleus (7N), facial nerve (7n) and inferior olive (IO). TCAP-1 was not localized in the cuneate nucleus (Cu). In the pons strong TCAP-1-ir was localized to cells of the pontine nuclei (Pn), oral pontine reticular nucleus (PnO) and locus coeruleus (LC). However, TCAP-1-ir was not detected in the longitudinal fasciculus of the pons (lfp) nor the lateral lemniscus of the pons (ll). In the midbrain, TCAP-1-ir was strongly localized to cells of the inferior colliculus (IC), ventral tegmental area (VTA) and substantia nigra pars compacta (SNC). Weak TCAP-1-ir was detected in the superior colliculus (SC) and substantia nigra pars reticulata (SNR). In the hippocampus, TCAP-1-ir was strongly localized to the pyramidal neurons of the CA1, CA2 and CA3 regions, whereas weak TCAP-1-ir was detected in the granular layer of the dentate gyrus (DG). In the thalamus, weak TCAP-1-ir was localized to cells of the mediorostral part of the lateral posterior thalamic nucleus (LPMR), laterodorsal thalamic nucleus (LD), ventral posteromedial thalamic nucleus (VPM), and ventral posterolateral thalamic nucleus (VPL). TCAP-1-ir was not detected in the medial lemniscus (ml). However, in the hypothalamus, strong TCAP-1-ir was detected in the soma and neuronal projections of cells in the perifornical nucleus (Pef), lateral hypothalamic area (LH), and ventromedial hypothalamic nucleus (VMH) and medial preoptic area (MPA). In the limbic regions, TCAP-1-ir was localized to cells within the bed nucleus of the stria terminalis (BST), amygdaloid nucleus (Amg), lateral septal nucleus (LS), nucleus accumbens shell (AcbSh), nucleus accumbens core (AchC), piriform cortex (Pir), ventral anterior olfactory nucleus (AOV), accessory olfactory bulb (AOB), medial anterior olfactory nucleus (AOM), dorsal part of the anterior olfactory nucleus (AOD) and posterior part of the anterior olfactory nucleus (AOP). TCAP-1-ir was not found in the caudate putamen (CPu), ventral pallidum (VP), intrabulbar part of the anterior commissure (aci), anterior part of the anterior commissure (aca), fimbria of the hippocampus (fi) or corpus callosum (cc).

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3.6 References

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Kinel-Tahan Y, Weiss H, Dgany O, Levine A, Wides, R (2007) Drosophila odz gene is required for multiple cell types in the compound retina. Dev. Dyn. 236:2541-2554. Kupferschmidt DA, Lovejoy DA, Rotzinger S, Erb S (2011) Teneurin C-terminal associated peptide-1 blocks the effects of corticotropin-releasing factor on reinstatement of cocaine seeking and on cocaine-induced behavioural sensitization. Br. J. Pharmacol. 162:574-583. Leamey CA, Glendining KA, Kreiman G, Kang ND, Wang KH, Fassler R, Sawatari A, Tonegawa S, Sur M (2008) Differential gene expression between sensory neocortical areas: potential roles for Ten_m3 and Bcl6 in patterning visual and somatosensory pathways. Cereb. Cortex 18:53-66. Leamey CA, Merlin S, Lattouf P, Sawatari A, Zhou X, Demel N, Glendining KA, Oohashi T, Sur M, Fassler R (2007) Ten_m3 regulates eye-specific patterning in the mammalian visual pathway and is required for binocular vision. PLoS Biol. 5:e241. Levine A, Bashan-Ahrend A, Budai-Hadrian O, Gartenberg D, Menasherow S, Wides R (1994) Odd Oz: a novel Drosophila pair rule gene. Cell 77:587-598. Li H, Bishop KM, O'Leary DD (2006) Potential target genes of EMX2 include Odz/Ten-M and other gene families with implications for cortical patterning. Mol Cell Neurosci 33(2):136-149. Lossie AC, Nakamura H, Thomas SE, Justice MJ (2005) Mutation of l7Rn3 shows that Odz4 is required for mouse gastrulation. Genetics 169:285-299. Lovejoy DA, Al Chawaf A, Cadinouche MZ (2006) Teneurin C-terminal associated peptides: an enigmatic family of neuropeptides with structural similarity to the corticotropin-releasing factor and calcitonin families of peptides. Gen Comp Endocrinol 148(3):299-305. Mieda M, Kikuchi Y, Hirate Y, Aoki M, Okamoto H (1999) Compartmentalized expression of zebrafish ten-m3 and ten-m4, homologues of the Drosophila ten(m)/odd Oz gene, in the central nervous system. Mech Dev 87:223-227. Miguel-Hidalgo JJ, Overholser JC, Jurjus GJ, Meltzer HY, Dieter L, Konick L, Stockmeier CA, Rajkowska G (2011) Vascular and extravascular immunoreactivity for intercellular adhesion molecule 1 in the orbitofrontal cortex of subjects with major depression: age-dependent changes. J Affect. Disord 132:422-431. Mihailoff GA, Burne RA, Woodward DJ (1978) Projections of the sensorimotor cortex to the basilar pontine nuclei in the rat: an autoradiographic study. Brain Res 145(2):347-354. Millan MJ (2006) Multi-target strategies for the improved treatment of depressive states: Conceptual foundations and neuronal substrates, drug discovery and therapeutic application. Pharmacol Ther 110:135-370. Ng T, Chand D, Song L, Al Chawaf A, Watson JD, Boutros PC, Belsham DD, Lovejoy DA (2012) Identification of a novel brain derived neurotrophic factor (BDNF)-inhibitory factor: regulation of BDNF by teneurin C-terminal associated peptide (TCAP)-1 in immortalized embryonic mouse hypothalamic cells. Regul Pept 174:79-89.

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Nunes SM, Ferralli J, Choi K, Brown-Luedi M, Minet AD, Chiquet-Ehrismann R (2005) The intracellular domain of teneurin-1 interacts with MBD1 and CAP/ponsin resulting in subcellular codistribution and translocation to the nuclear matrix. Exp Cell Res 305:122-132. Oohashi T, Zhou XH, Feng K, Richter B, Morgelin M, Perez MT, Su WD, Chiquet-Ehrismann R, Rauch U, Fassler R (1999) Mouse ten-m/Odz is a new family of dimeric type II transmembrane proteins expressed in many tissues. J Cell Biol 145:563-577. Qian X, Barsyte-Lovejoy D, Wang L, Chewpoy B, Gautam N, Al Chawaf A, Lovejoy DA (2004) Cloning and characterization of teneurin C-terminus associated peptide (TCAP)-3 from the hypothalamus of an adult rainbow trout (Oncorhynchus mykiss). Gen Comp Endocrinol 137:205-216. Rotzinger S, Lovejoy DA, Tan LA (2010) Behavioral effects of neuropeptides in rodent models of depression and anxiety. Peptides 31(4): 736-756. Rubin BP, Tucker RP, Brown-Luedi M, Martin D, Chiquet-Ehrismann R (2002) Teneurin 2 is expressed by the neurons of the thalamofugal visual system in situ and promotes homophilic cell-cell adhesion in vitro. Development 129:4697-4705. Rubin BP, Tucker RP, Martin D, Chiquet-Ehrismann R (1999) Teneurins: a novel family of neuronal cell surface proteins in vertebrates, homologous to the Drosophila pair-rule gene product Ten-m. Dev Biol 216:195-209. Sokoloff L (1981) Localization of functional activity in the central nervous system by measurement of glucose utilization with radioactive deoxyglucose. J Cereb Blood Flow Metab. 1(1):7-36. Tan LA, Al Chawaf A, Vaccarino FJ, Boutros PC, Lovejoy DA (2011) Teneurin C-terminal associated peptide (TCAP)-1 modulates dendritic morphology in hippocampal neurons and decreases anxiety-like behaviors in rats. Physiol Behav 104:199-204. Tan LA, Chand, D, De Almeida, R, Xu M, De Lannoy L, Lovejoy DA (2011b) Modulation of neuroplastic changes and corticotropin-releasing factor-associated behavior by a phylogenetically ancient and conserved peptide family. Gen. Comp Endocrinol 176:309-313. Tan LA, Xu K, Vaccarino FJ, Lovejoy DA, Rotzinger S (2009) Teneurin C-terminal associated peptide (TCAP)-1 attenuates corticotropin-releasing factor (CRF)-induced c-Fos expression in the limbic system and modulates anxiety behavior in male Wistar rats. Behav Brain Res 201:198-206. Tan LA, Xu K, Vaccarino FJ, Lovejoy DA, Rotzinger S. (2008) Repeated intracerebral teneurin C-terminal associated peptide (TCAP)-1 injections produce enduring changes in behavioral responses to corticotropin-releasing factor (CRF) in rat models of anxiety. Behav Brain Res 188(1): 195-200. Trubiani G, Al Chawaf A, Belsham DD, Barsyte-Lovejoy D, Lovejoy DA (2007) Teneurin carboxy (C)-terminal associated peptide-1 inhibits alkalosis-associated necrotic neuronal death

120 by stimulating superoxide dismutase and catalase activity in immortalized mouse hypothalamic cells. Brain Res 1176:27–36. Tucker RP, Beckmann J, Leachman NT, Scholer J, Chiquet-Ehrismann R (2012) Phylogenetic analysis of the teneurins: conserved features and premetazoan ancestry. Mol Biol Evol 29:1019- 1029. Tucker RP, Kenzelmann D, Trzebiatowska A, Chiquet-Ehrismann R (2007) Teneurins: transmembrane proteins with fundamental roles in development. Int J Biochem Cell Biol 39:292- 297. Van Pett K, Viau V, Bittencourt JC, Chan RK, Li HY, Arias C, Prins GS, Perrin M, Vale W, Sawchenko PE (2000) Distribution of mRNAs encoding CRF receptors in brain and pituitary of rat and mouse. J Comp Neurol 428(2):191-212. Wang L, Rotzinger S, Barsyte-Lovejoy D, Qian X, Elias CF, Bittencourt JC, De Cristofaro A, Wang NC, Belsham D, Vaccarino F, Lovejoy DA (2005) Teneurin proteins possess a carboxy terminal corticotropin-releasing factor-like sequence that modulates emotionality and neuronal growth. Mol Brain Res 133:253-265. Wang XZ, Kuroda M, Sok J, Batchvarova N, Kimmel R, Chung P, Zinszner H, Ron D (1998) Identification of novel stress-induced genes downstream of chop. EMBO J. 17:3619–3630. Young TR, Leamey CA (2009) Teneurins: important regulators of neural circuitry. Int J Biochem Cell Biol 41:990-993. Zhou XH, Brandau O, Feng K, Oohashi T, Ninomiya Y, Rauch U, Fässler R (2003) The murine Ten-m/Odz genes show distinct but overlapping expression patterns during development and in adult brain. Gene Expr Patterns 3:397-405.

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4 Chapter Four: Regulation of cytoskeletal dynamics by TCAP-1

This chapter have been published in Neuroscience:

Chand D*, Song L, deLannoy L, Barsyte-Lovejoy D, Ackloo S, Boutros PC, Evans K, Belsham DD, Lovejoy DA (2012) C-terminal region of teneurin-1 co-localizes with dystroglycan and modulates hippocampal neurite outgrowth through ERK-dependent stathmin and filamin A modulation of the cytoskeleton. Neuroscience 219:255-270. (Authorization for reproduction obtained from Elsevier)

4.1 Abstract

The pyramidal neurons in the hippocampus are extremely neuroplastic, and the complexity of dendritic branches can be dynamically altered in response to a variety of stimuli, including learning and stress. Recently, the teneurin family of proteins have emerged as an interneuronal and extracellular matrix signaling system that play a significant role in brain development and neuronal communication. Encoded on the last exon of the teneurin genes are a new family of bioactive peptides termed the teneurin C-terminal associated peptides (TCAPs). Previous studies indicate that TCAP-1 regulates axon fasciculation and dendritic morphology in the hippocampus.

This study was aimed at understanding the molecular mechanisms by which TCAP-1 regulates these changes in the mouse hippocampus. FITC-labeled TCAP-1 binds to the pyramidal neurons of the CA2 and CA3, and dentate gyrus in the hippocampus of the mouse brain. Moreover,

FITC-TCAP-1 co-localizes with β-dystroglycan upon binding to the plasma membrane of cultured immortalized mouse E14 hippocampal cells. In culture, TCAP-1 stimulates ERK1/2- dependent phosphorylation of the cytoskeletal regulatory proteins, stathmin at serine-25 and filamin A at serine-2152. In addition, TCAP-1 induces actin polymerization, increases immunoreactivity of tubulin-based cytoskeletal elements and causes a corresponding increase in filopodia formation and mean filopodia length in cultured hippocampal cells. I postulate that the

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TCAP-1 region of teneurin-1 has a direct action on the cytoskeletal reorganization that precedes neurite and process development in hippocampal neurons. My data provides novel evidence that functionally links the teneurin and dystroglycan systems and provides new insight into the molecular mechanisms by which TCAP-1 regulates cytoskeletal dynamics in hippocampal neurons. The TCAP-dystroglycan system may represent a novel mechanism associated with the regulation of hippocampal-function.

4.2 Introduction

In the developing central nervous system (CNS), the regulation of neurite outgrowth and retraction are fundamental for the precise formation of synapses (O'Leary et al., 1986). The mechanism by which this occurs remains unresolved, but is thought to involve a reorganization of actin- and tubulin- based cytoskeletal elements (Tada and Sheng, 2006; Gu et al., 2008). In the brain, rapid reorganization of microtubule cytoskeleton is mainly regulated by an 18 kDa ubiquitously expressed phosphoprotein known as stathmin that destabilizes microtubulin when unphosphorylated (Cassimeris, 2002). On the other hand, the dynamic regulation of actin cytoskeleton could be mediated by the actin cross-linking protein filamin A (Stossel et al., 2001) or the actin-binding protein dystroglycan (Chen et al., 2003). Together, these systems may have an important role in establishing a cytoskeletal topography that is required for the regulation of neuronal plasticity, synapse formation and brain development.

The teneurins are a family of four transmembrane proteins predominantly expressed in the CNS (Tucker et al., 2007; Ben-Zur et al., 2000) and play a significant role in brain development and neuronal connectivity (Kenzelmann et al., 2007). Originally discovered in

Drosophila as odd oz (odz; Levine et al., 1994) and tenascin-major (ten-m; Baumgartner et al.,

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1994), the teneurins consist of about 2800 residues and possess a complex set of functional domains (Minet et al., 1999; Feng et al., 2002; Leamey et al., 2007). However, some family members may also exist as a number of splice variants (Lossie et al., 2005) or as smaller soluble proteins derived from further proteolytic cleavage of the carboxy terminal regions (Kenzelmann et al., 2008). One such region is a 40- to 41-amino acid sequence at the tip of the carboxy terminus of the teneurin proteins that possess characteristics of a bioactive peptide (Qian et al.,

2004; Wang et al, 2005). This sequence was termed the ‘teneurin C-terminal associated peptide’

(TCAP).

One of these peptides, TCAP-1, is highly expressed in regions of the forebrain and limbic system, including the hippocampus, amygdala, cerebellum, hypothalamus, and cortex (Wang et al., 2005). Synthetic versions of TCAP-1 exert numerous biological effects associated with the limbic system. In vitro, TCAP-1 increases cytoskeletal proteins including β-actin and β-tubulin and modulates neurite outgrowth in cultured primary hippocampal neurons (Al Chawaf et al.,

2007a). In vivo, synthetic TCAP-1 regulates stress-induced behaviour (Wang et al., 2005; Al

Chawaf et al., 2007b; Tan et al., 2008) and blocks the stress-associated neurohormone corticotropin-releasing factor (CRF)-mediated c-fos synthesis in the hippocampus and amygdala of the adult rat (Tan et al., 2009). These effects on behaviour are long-lasting, with effects persisting 21 days after TCAP-1 injection (Wang et al., 2005). Recently, TCAP-1 was shown to completely ablate CRF-induced cocaine seeking reinstatement in rats, suggesting that the peptide may have applications for the treatment of mood and addictive disorders (Kupferschmidt et al.,

2011). Furthermore, TCAP-1 increases spine density in the CA1 and CA3 neurons of the rodent hippocampus but not in the amygdala (Tan et al., 2011) suggesting that the inhibition of stress

124 and anxiety-like behaviour by TCAP-1 may be associated with the remodeling of neuronal networks and dendritic morphology in the hippocampus.

Thus, the TCAP region of the teneurins may act as a novel neuroplastic regulatory factor in the hippocampus. I hypothesize that TCAP-1 regulates cytoskeletal elements as a prelude to the changes associated with TCAP-mediated neuronal morphology and behaviours within the hippocampus. In this study, I investigate the molecular mechanism by which TCAP-1 regulates cytoskeletal dynamics in the mouse hippocampus. The TCAP-1 signaling mechanism may, therefore, represent a novel system involved in the regulation of cytoskeletal dynamics and neuronal plasticity in hippocampal neurons.

4.3 Materials and Methods

4.3.1 Cell Culture. Immortalized mouse embryonic E14 hippocampal cells (Gingerich et al.,

2010; provided by Dr. D. Belsham, University of Toronto) were cultured in Dulbecco’s Modified

Eagle Medium (DMEM) supplemented with 10% heat-inactivated fetal bovine serum (FBS), 100

U/ml penicillin and 100 µg/ml streptomycin (Invitrogen, Burlington, ON, Canada). Cells were maintained at 60-70% confluency at 37C in a humidified CO2 incubator.

4.3.2 Antibodies and Reagents. Chicken anti-cow-microtubule-associated protein 2 (MAP2;

1:8000) and chicken anti-cow-glial fibrillary acidic protein (GFAP; 1:2000) were obtained from

Abcam (Cambridge, MA, USA). Mouse monoclonal anti--tubulin (1:200) and anti--tubulin

(1:200) were obtained from Sigma-Aldrich Canada (Oakville, ON, Canada). Rabbit anti-human-

-actin and anti-human-phospho-FAK (Tyr925), -MEK1/2 (Ser217/221), -ERK1/2

(Thr202/Tyr204), -Stathmin (Ser38), -p90RSK (Ser380) and –filamin A (Ser 2152) were all used

125 at a dilution of 1:1000 and were obtained from New England Biolabs (Pickering, ON, Canada).

Rabbit anti-human-phospho-Stathmin (Ser25; 1:500) and rabbit anti-human-β-Dystroglycan (β-

DG, 1:500) were obtained from Santa Cruz Biotechnology (Santa Cruz, CA, USA). The corresponding pan antibodies rabbit anti-human-MEK1/2, -Stathmin (STMN), -Filamin A, - ribosomal S6 kinases (RSK1/2/3) and -rat-ERK1/2 were all used at a dilution of 1:1000 and were obtained from New England Biolabs. Secondary antibodies HRP-labeled goat anti-rabbit IgG

(1:5000) was obtained from New England Biolabs, Alexa 594 goat anti-rabbit (1:400), Alexa

594 donkey anti-mouse (1:400) and Alexa 488 donkey anti-mouse (1:400) were obtained from

Invitrogen and FITC goat anti-chicken (1:250) and Texas-Red goat anti-chicken (1:200) were obtained from Abcam. Alexa Fluor Phalloidin-594 was obtained from Invitrogen and used according to the manufacturer’s instructions. MEK inhibitor U0126 (New England Biolabs) was dissolved in dimethyl sulfoxide (DMSO) leading to a 50 mM U0126 stock solution.

4.3.3 Synthesis of TCAP Peptides. Mouse TCAP-1 and scrambled TCAP-1 (SC-TCAP-1) were synthesized at 95% purity using f-moc-based solid phase synthesis (American Peptide Company,

Sunnyvale, CA, USA). TCAP-1 variants with lysine substituted for arginine at positions 8 or 37

(K8 and K37) were synthesized as previously described (Wang et al., 2005). The peptides were solubilized by exposure to ammonium hydroxide vapors for 2 min before dilution in phosphate- buffered saline (PBS) pH 7.4 with 10 mM sodium phosphate. Concentrations of 1 and 100 nM

TCAP-1 were used throughout the study as it has been previously established that this concentration appears to be most efficacious (Qian et al., 2004; Wang et al., 2005; Al Chawaf et al., 2007a).

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4.3.4 Preparation of Fluoresceinisothiocyanate (FITC)-labeled TCAP-1. [K8] or [K37]-TCAP-1 and –SC-TCAP-1 were labeled with FITC according to the EZ-Label FITC Protein labeling kit (Pierce Biotechnology, Rockford, IL, USA) and as previously described (Al Chawaf et al.,

2007b). The size of the FITC-TCAP-1 and FITC-SC-TCAP-1 conjugates were confirmed by non-reducing SDS-PAGE on a 10–20% Tris-Tricine gel (BioRad, Hercules, CA, USA). The amount of FITC required to be used as a control was determined by titrating PBS-diluted preparations of FITC on a nitrocellulose membrane. Fluorescence was measured using an Epi

Chemi II Darkroom system (UVP) with the sequential integration function of LabWorks Image acquisition and analysis software (V4.0.0.8, Upland, CA, USA).

4.3.5 FITC-TCAP-1 binding studies. Paraformaldehyde-fixed paraffin-embedded sections (5 µm) of adult mice brains were deparaffinized in xylene and rehydrated. Sections were permeabilized with 0.3% Triton X-100 (Fisher Scientific, Ottawa, ON, Canada) for 10 min. The tissue sections were blocked with 10% normal goat serum (NGS, Vector Laboratories, Burlington, ON, Canada) in PBS for 1 h and incubated overnight at 4˚C with FITC-TCAP-1 or FITC-SC-TCAP-1 at 1:400 dilution in 0.1 M PBS containing 1% NGS. Sections were washed with PBS, counterstained with

4',6-diamidino-2-phenylindole (DAPI; Sigma-Aldrich Canada) for 1 min and mounted with 1,4- diazabicyclo[2.2.2]octane (DABCO; Sigma-Aldrich Canada).

Immortalized mouse E14 hippocampal cells were plated on poly-D-lysine-coated cover- slips (BD Biosciences, Mississauga, ON, Canada) and cultured to 60% confluency in DMEM with 10% FBS as described above. The cells were rinsed with PBS and incubated with conjugated mouse FITC-TCAP-1 or FITC-SC-TCAP-1 diluted at 1:400 in DMEM with 10%

FBS for 15 min at 37°C and 5% CO2. The cells were washed with PBS, fixed in 4%

127 paraformaldehyde (PFA, Sigma-Aldrich Canada) in PBS, counterstained with DAPI and mounted with DABCO as described above.

For live imaging, E14 hippocampal cells were cultured to 60% confluency in DMEM with 10% FBS in an 8-well glass slide chamber (Fisher Scientific) and incubated with FITC- labeled TCAP-1 for 10 min as described above. The cells were then washed twice with fresh

DMEM and incubated with DMEM with 10% FBS. Live confocal microscopy and time lapse imaging was performed at 37°C and 5% CO2 for a minimum duration of 5 min. For controls, cells were incubated with unconjugated FITC (1:400) or FITC-SC-TCAP-1 (1:400) as described above.

4.3.6 Immunofluorescent staining. Mouse E14 hippocampal cells were plated on poly-D-lysine- coated cover-slips and cultured overnight to 60% confluency in DMEM with 10% FBS. The cells were pretreated with FITC-labeled TCAP-1 for 30 min as described above, fixed in 4%

PFA for 20 min and permeabilized for 10 min with 0.3% Triton X-100. After blocking with 10%

NGS for 1 h, the cells were incubated with rabbit anti--DG overnight at 4°C in PBS and 1%

NGS. The cells were washed in PBS and incubated with Alexa 594 goat anti-rabbit secondary antibody in 0.01 M PBS for 1 h. Cells were counterstained with DAPI and mounted with

DABCO as described above. For controls, cells were incubated with, primary antibody alone, secondary antibody alone, or neither primary nor secondary antibodies.

4.3.7 Cytoskeletal and Morphological analysis. Mouse E14 hippocampal cells were plated on poly-D-lysine-coated cover-slips in triplicate as described above. At 60% confluency, the cells were synchronized in serum-free DMEM overnight. The cells were washed twice with PBS and

128 then treated with water, 1nM TCAP-1, 100nM TCAP-1 or 100nM SC-TCAP-1 dissolved in fresh

DMEM, 1% FBS for 1 h at 37°C and 5% CO2. Immunofluorescent labeling with the indicated antibodies was performed as described above to determine the effect of TCAP-1 treatment on the cytoskeleton. For quantitative analysis of immunoreactive tubulin, Image J software (NIH,

Bethesda, MD, USA) was used to assess the whole cell fluorescence intensity as previously described by Al Chawaf et al., (2007a) and Gavet and Pines (2010). A minimum of 100 cells per treatment were analyzed by observers blinded to the study design.

The relative proportions of filamentous F-actin and G-actin across treatments were analyzed using the G-Actin/F-actin In Vivo Assay Biochem Kit and quantitated according to the manufacturer’s instructions (Cytoskeleton Inc, Denver, CO, USA). Briefly, mouse E14 hippocampal cells were treated with water, 100nM TCAP-1 or 100nM SC-TCAP-1 for 60 min as described as above. Cells were homogenized in 100 µl of F-actin stabilization and lysis buffer supplemented with 1mM ATP and 1% protease inhibitor cocktail (Cytoskeleton Inc.). Cell lysate was centrifuged at 100,000g, for 1 h at 37°C. Supernatants containing G-actin were removed and stored at -80°C and the remaining F-actin pellets were re-suspended in 100 µl of F-actin depolymerization buffer (Cytoskeleton Inc.) and incubated on ice for 1 h to allow for actin depolymerization to occur. Four microliters of supernatant (G-actin) and pellet (F-actin) fractions were subjected to western blot analysis using an anti-actin rabbit polyclonal antibody

(Cytoskeleton Inc). The amounts of F-actin to G-actin were determined from a standard curve for actin protein ranging from 10 – 50 ng by densitometry using an Epi Chemi II Darkroom and

Laboratory works V4.0.0.8 (UVP) as per the manufacturer’s instructions (Cytoskeleton Inc.) and as described by Zhang et al. (2005). The G-actin and F-actin assays were performed in quadruplicate.

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To determine whether TCAP-1 treatments affected cell morphology, a minimum of 100 cells were analyzed per treatment using NIS-Elements Basic Research V2.30 (Nikon,

Mississauga, ON, Canada) and scored for number of filopodia per cell and filopodia length by observers blinded to the study design. Cell size was determined using the Image J software

(NIH) by observers blinded to the study design. Filopodia were identified according to the morphological criteria described by Grutzendler et al. (2002) and Matsumoto-Miyai et al. (2009).

Briefly, serial images of E14 hippocampal cells treated with water, 1nM TCAP-1, 100nM

TCAP-1 or 100nM SC-TCAP-1 and labeled for F-actin (Phalloidin-594; Invitrogen), were collected at z-steps of 0.10 µm using a 40X and 63X oil-immersion objective (Leica

Microsystems, Wetzlar, Germany). Filopodia were identified as long thin structures with a ratio of head to neck diameter smaller than 1.2:1, and a ratio of length to neck diameter larger than 3:1

(Grutzendler et al., 2002). Untreated cells arbitrarily represented the 100% values.

4.3.8 Western Blot Analysis. For Western blot analysis of TCAP-1 signaling pathway and cytoskeletal regulating proteins, mouse E14 hippocampal cells were cultured in triplicate as described above. At 60% confluency, cells were synchronized in serum-free DMEM overnight.

The cells were washed twice with PBS and then treated for 0, 1, 5, 10, 30 or 60 min with 100nM

TCAP-1, 100nM scrambled TCAP-1 (SC-TCAP-1) or vehicle (water) in DMEM, 1% FBS. To inhibit mitogen-activated protein kinase (MAPK) kinase (MEK) activity, MEK inhibitor, U0126

(10µM), was added 30 min before the addition of TCAP-1. In parallel, control cells were treated with DMSO alone. The cells were washed once with PBS, lysed in ice-cold radioimmunoprecipitation assay (RIPA) lysis buffer (50 mM TRIS-HCl, 150 mM NaCl, 1%

Triton X-100, 0.1% SDS, 1% sodium deoxycholate, 1 mM EDTA, 1 mM PMSF, 1 mM sodium

130 orthovanadate, 1 mM sodium fluoride and 1 μg/ml each of pepstatin, aprotinin and leupeptin) and placed on ice for 15 min. Cell debris was removed by centrifugation at 20,000g for 20 min at

4C. The supernatant was collected for Western blot analysis. The protein concentration was determined using the bicinchoninic acid (BCA) kit as per instructions (Pierce Biotechnology).

E14 protein cell lysates (50 μg/lane) were resolved by SDS-PAGE using a Mini-Protean 3

Cell (BioRad) electrophoresis unit and transferred onto a nitrocellulose membrane (GE Health

Care, Piscataway, NJ, USA). After blocking with 5% bovine serum albumin (BSA, Sigma-

Aldrich Canada) in PBS-T for 1 h, the membrane was incubated with the indicated antibodies overnight at 4°C. After washing with PBS-T, the membrane was incubated with HRP-labeled goat anti-rabbit IgG for 1 h at room temperature. The membrane was analyzed using the ECL western blotting detection and analysis kit (GE Health Care). Experiments were repeated three times. Blots were scanned and optical density was determined using an Epi Chemi II Darkroom and Laboratory works V4.0.0.8 (UVP). All assays were performed in triplicate and the band intensity for each immunoreactive protein was compared with baseline lane intensity according to the quantification procedure utilized for this software (Qian et al., 2004).

4.3.9 Image Analysis. Cells were analyzed using a Zeiss LSM 510 confocal laser-scanning microscope (Carl Zeiss MicroImaging, Göttingen, Germany) with a Q imaging Retiga EXi camera (Q Imaging, Surrey, BC, Canada). Tissue sections were analyzed using a Zeiss Axio

Imager Z1 Deacon microscope (Carl Zeiss MicroImaging) with a Q imaging Retiga EXi camera

(Q Imaging). The resulting images were adjusted for brightness and contrast using Volocity 5.2

(Improvision, Waltham, MA, USA) and Adobe Photoshop CS4 Extended (San Jose, CA, USA).

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4.3.10 Statistical Analyses. Data were analyzed by one-way or two-way ANOVA followed by

Bonferroni’s post hoc test using GraphPad Prism Ver. 4. Mean values were obtained from a minimum of three independent experiments and data were considered statistically significant when ***P<0.001, **P<0.01 or *P<0.05.

4.4 Results

4.4.1 Binding of TCAP-1 to hippocampal cells and tissues

Coronal sections of the mouse brain were incubated with FITC-labeled TCAP-1 in order to identify TCAP-1 binding and uptake sites in the mouse hippocampus. FITC-labeled TCAP-1 bound exclusively to the pyramidal neurons in the CA1, CA2, CA3 and granular layer of the dentate gyrus of the mouse hippocampus (Fig. 4.1). FITC-conjugated TCAP-1 binding was not observed in the stratum oriens, stratum lucidum, stratum radiatum or stratum lacunosum- moleculare (Fig. 4.1). TCAP-1 binding was strong in the CA2 (Fig. 4.1D,E), CA3 (Fig. 4.1G,H), and dentate gyrus (Fig. 4.1J,K) regions and weak in the CA1 region (Fig. 4.1A,B) of the mouse hippocampus. Fluorescence was not observed in any of the controls sections incubated with the

FITC-labeled scrambled TCAP-1 control (FITC-SC-TCAP-1; Fig. 4.1C,F,I,L).

To corroborate the FITC-TCAP-1 binding in the brain, TCAP-1-binding sites were further examined by incubating cultured immortalized mouse E14 hippocampal cells with FITC- conjugated TCAP-1. Cells incubated with FITC-[K8]-TCAP-1 for 10 min and analyzed by rapid time-lapse confocal microscopy for 5 min showed strong TCAP-1 binding on or near the plasma membrane (Fig. 4.2A). FITC-TCAP-1 remained bound at or near the plasma membrane after 12 min (Fig. 4.2B) and 15 min (Fig. 4.2C) from the time of initial treatment. As further confirmation, cell incubated with FITC-conjugated TCAP-1 for 15 min, fixed and analyzed by

132 confocal microscopy also showed strong binding at the plasma membrane (Fig. 4.2D,E) indicating a ligand-plasma membrane interaction. Fluorescence was not observed in any of the cells incubated with the FITC-SC-TCAP-1 control (Fig. 4.2F).

Figure 4.1. FITC-[K8]-TCAP-1 binding in mouse hippocampal formation. Sections incubated with FITC-labeled TCAP-1 showed strong binding in the cytosol and plasma membrane of the neuronal layers of the hippocampus (B, E, H and K). FITC-TCAP-1 binding was strongest in the pyramidal layer (Py) of the CA2 (E) and CA3 regions (H) but weak in the CA1 (B) region of the hippocampus. Moderate binding was observed in the dentate gyrus (K). FITC-labeled TCAP-1 binding was not observed in the stratum lucidum (SL), stratum radiatum (SR), stratum lacunosum-moleculare (SLM) or stratum oriens (SO). Immunofluorescence was not observed in sections incubated with FITC-labeled scrambled TCAP-1 (FITC-SC-TCAP-1) (C, F, I and L). For each fluorescence image, the corresponding DIC image is represented on the left (A, D, G and J). All sections were counterstained with DAPI. Magnification, 200X. Scale bars, 50μm.

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Only neuronal cells were present in the cell culture as indicated by the presence of immunoreactive microtubule-associated protein 2 (MAP2; Fig. 4.2G) and absence of glial- specific marker glial fibrillary acidic protein (GFAP; Fig. 4.2H).

Figure 4.2. TCAP-1 binding in immortalized mouse E14 hippocampal cells. Cultured E14 hippocampal cells incubated with FITC-[K8]-TCAP-1 for 10 min and analyzed by time-lapse confocal microscopy showed strong FITC-TCAP-1 binding to the plasma membrane (A). FITC- conjugated TCAP-1 remained bound 12 min (B) and 15 min (C) after the initial incubation. Cells incubated with FITC-[K8]-TCAP-1 for 15 min also showed binding to the plasma membrane (E). Corresponding DIC image is represented on the left (D). Fluorescence was not observed in cells incubated with FITC-SC-TCAP-1 (F). Only neuronal cells were present in cell culture as indicated by the presence of MAP2 (G) and absence of GFAP (H). Sections were counterstained with DAPI to highlight cellular DNA (E-G). Magnification, 400X (A-C), 630X (D and E) 200X (F and G). Scale bars, 20μm (A-C), 15μm (D and E) and 50 μm (F and G). FITC-SC-TCAP-1, FITC-labeled scrambled TCAP-1; GFAP, glial fibrillary acidic protein; MAP2, microtubule- associated protein 2.

4.4.2 TCAP-1 co-localization with the Dystroglycan complex

Previous gene array studies indicated that TCAP-1 upregulated cytoskeletal regulating proteins such as the dystroglycan system (D.A. Lovejoy, D.D. Belsham and M.J. Brownstein, unpublished observations). To determine whether TCAP-1 functionally interacts with the

134 dystroglycan complex, cultured immortalized mouse E14 hippocampal cells were incubated with

FITC-[K8]-TCAP-1 for 30 min and co-labeled with β-dystroglycan (β-DG) antiserum. Cells treated with FITC-labeled TCAP-1 showed strong binding to the plasma membrane (Fig.

4.3A,B) in a pattern similar to that observed for immunoreactive β-DG (Fig. 4.3A,C). Strong co- localization between FITC-[K8]-TCAP-1 and β-DG was observed at the plasma membrane (Fig.

4.3A,D).

Figure 4.3. TCAP-1 co-localization with the dystroglycan complex in immortalized mouse E14 hippocampal cells. Cultured E14 hippocampal cells incubated with FITC-[K8]- TCAP-1 for 30 min showed strong binding to the plasma membrane and punctate- like labeling in the cytosol (B). β-Dystroglycan (β-DG) immunoreactivity was mostly observed near the plasma membrane with weak immunoreactivity in the cytosol (C). Strong co-localization (yellow; white arrows) between FITC-[K8]-TCAP-1 and β-Dystroglycan was observed at the plasma membrane (D). The corresponding DIC image is represented on the left (A). All sections were counterstained with DAPI. Magnification, 630X. Scale bars, 15μm.

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4.4.3 TCAP-1 increases actin polymerization and tubulin cytoskeletal elements

TCAP-1 co-localization with the dystroglycan complex was consistent with previous observations that TCAP-1 could regulate cytoskeletal elements (Al Chawaf et al., 2007a).

Therefore, the effect of TCAP-1 on the cellular localization of β-actin, F-actin, α-tubulin and β- tubulin in E14 hippocampal cells were analyzed by immunofluorescent confocal microscopy after 1 h of treatment. TCAP-1-treated hippocampal cells showed an increase in long β-actin strands and a much greater prevalence of actin cross-linking and polymerization (Fig. 4.4B). This observation is in contrast to untreated (Fig. 4.4A) and 100nM scrambled TCAP-1-treated (Fig.

4.4C) hippocampal cells, where immunoreactive β-actin was present mainly as monomeric actin subunits. Similarly, TCAP-1-treated E14 hippocampal cells labeled with phalloidin showed an increase in long filamentous F-actin (Fig. 4.4E) as compared to untreated (Fig. 4.4D) and scrambled TCAP-1-treated (Fig. 4.4F) hippocampal cells, indicating a direct action on F-actin polymerization.

To confirm the immunofluorescence-labeling, I used western blot analysis to compare the proportions of G-actin and F-actin in E14 hippocampal cells treated with water (untreated),

100nM TCAP-1 or 100nM scrambled TCAP-1 (Fig. 4.4G,H). In untreated hippocampal cells, G- actin content was 76.44 ± 1.53% of total cellular actin and F-actin content was 23.56 ± 1.54% of total cellular actin (Fig. 4.4H). However TCAP-1 significantly decreased G-actin to 46.20 ±

1.56% and significantly increased F-actin to 53.80 ± 1.56% of total cellular actin (P<0.001, one- way ANOVA with Bonferroni’s post hoc test, n=4) as compared to untreated and scrambled

TCAP-1-treated hippocampal cells (Fig. 4.4H). Cells treated with scrambled TCAP-1 showed no significant change in G-actin (78.87 ± 2.41% of total cellular actin) or F-actin (21.13 ± 2.41% of total cellular actin) as compared to untreated hippocampal cells (Fig. 4.4H).

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Figure 4.4. TCAP-1 treatment increases actin polymerization in immortalized E14 hippocampal cells. Cultured E14 hippocampal cells treated with 100nM TCAP-1 for 60 min showed increase labeling of long β-actin polymers (B) as compared to untreated (water) (A) and 100nM SC-TCAP-1-treated (C) cells where β-actin was mostly present as individual subunits. Phalloidin-labeling showed an increase in F-actin immunoreactivity and actin polymerization in TCAP-1 treated cells (E) as compared to untreated (D) and 100nM SC-TCAP-1-treated (F) hippocampal cells. Western blot analysis of G- and F-actin in hippocampal cells treated with water (Untreated), 100nM TCAP-1 and 100nM SC-TCAP-1 (G). Densitometric analysis showed a significant decrease of G-actin and a corresponding significant increase of F-actin in hippocampal cells treated with 100nM TCAP-1 for 60 min as compared to untreated or 100nM SC-TCAP-1-treated hippocampal cells (n=4, two-way ANOVA with a Bonferroni’s post hoc test, ***P<0.001, compared to untreated controls). Values are the mean ± SEM and expressed as a percent of total actin (H). Magnification, 630X. Scale bars, 15μm (A-F). SC-TCAP-1, scrambled TCAP-1.

TCAP-1 treatment also induced changes in the microtubulin-based cytoskeleton (Fig.

4.5). E14 hippocampal cells treated with 100nM TCAP-1 showed a 2.41 ± 0.15 fold increase in

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α-tubulin (Fig. 4.5B,G) and 2.16 ± 0.11 fold increase in β-tubulin (Fig. 4.5E, H) immunofluorescence-labeling as compared to untreated hippocampal cells (Fig. 4.5A,G, α- tubulin; Fig. 4.5D,H, β-tubulin) (P<0.001, one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group; n=3). Cell treated with 100nM SC-TCAP-1 showed no significant change in α-tubulin (Fig. 4.5C,G) or β-tubulin (Fig. 4.5F,H) immunofluorescence-labeling as compared to untreated hippocampal cells (Fig. 4.5A,G, α-tubulin; Fig. 4.5D,H, β-tubulin).

Figure 4.5. TCAP-1 treatment increase tubulin immunoreactivity. Cultured E14 hippocampal cells treated with 100nM TCAP-1 for 60 min showed increase labeling of α-tubulin (B) and β-tubulin (E) as compared to water- (A, α-tubulin and D, β-tubulin; untreated) and 100nM SC-TCAP-1-treated (C, α-tubulin and F, β-tubulin) hippocampal cells. Strong tubulin immunoreactivity was detected in areas of cellular protrusions and outgrowth in TCAP-1-treated cells (B and E). Quantification of whole cell fluorescence, showed a significant increase in α- tubulin (G) and β-tubulin (H) immunofluorescence in TCAP-1 treated cells as compared to untreated and SC-TCAP-1-treated cells (one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group (n=3); ** p<0.01, *** p<0.001, compared to untreated controls). Magnification, 630X. Scale bars, 15μm (A-F). SC-TCAP-1, scrambled TCAP-1.

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4.4.4 TCAP-1 induces filopodia formation and elongation

The increase in actin polymerization (Fig. 4.4) and tubulin immunoreactivity (Fig. 4.5),

β-actin expression (Al Chawaf et al., 2007a) and dendritic spine density (Tan et al, 2011) in the mouse hippocampus indicated that TCAP-1 may regulate filopodia formation and elongation.

Cellular morphology was observed in cells immunolabeled for β-tubulin (A-H) or phalloidin- labeled F-actin (Fig. 4.6 I-L). Filopodia were analyzed from cells stained for F-actin (Fig. 4.6I-L) and according to the morphological criteria described by Grutzendler et al. (2002) and

Matsumoto-Miyai et al. (2009). In E14 hippocampal cells, TCAP-1 induced a 46.9 ± 10.2% and

56.4 ± 10.7% increase (P<0.001, one-way ANOVA with Bonferroni’s post hoc test, minimum

100 cells per group; n=3) in filopodia per cell at 1 and 100 nM TCAP-1 respectively, as compared to untreated (water-treated) cells (Fig. 4.6N).

Apart from increased filopodia formation, TCAP-1 also induced a significant increase in mean filopodia length, in E14 hippocampal cells treated with 1 and 100nM TCAP-1 (P<0.001, one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group; n=3) as compared to untreated cells (Fig. 4.6O). Hippocampal cells treated with 1 and 100nM TCAP-1 had a mean filopodia length of 10.07 ± 0.42µm and 11.44 ± 0.47µm respectively. In comparison, the mean filopodia lengths in untreated and SC-TCAP-1-treated hippocampal cells were 7.69 ±

0.32µm and 7.44 ± 0.46µm respectively. There was no significant change in filopodia formation per cell (Fig. 4.6N) or filopodia length (Fig. 4.6O) between 1 nM and 100 nM TCAP-1-treated cells (p>0.05, one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group; n=3).

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Figure 4.6. TCAP-1 treatment increases filopodia formation and mean filopodia length in immortalized E14 hippocampal cells. Representative images of cultured E14 hippocampal cells, immunolabeled for β-tubulin (A-D; red, E-H; grayscale) and phalloidin-stained F-actin (I- L; grayscale), showing increase cellular protrusions and filopodia formation (white arrows) after 1 h of 1nM TCAP-1 (B,F,J) an 100nM TCAP-1 (C,G,K) treatment as compared to water (A,E,I; untreated) and 100nM scrambled-TCAP-1 (D,H,L; SC-TCAP-1) treated cells. TCAP-1 induced a significant increase in the number of tubulin-labeled cellular protrusions (M), filopodia (N) and mean filopodia length (O) at 1nM and 100nM (one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group (n=4); *** p<0.001, compared to untreated controls). TCAP-1 did not affect overall cell size (P). Values are the mean ± SEM are indicated (M,N,O,P) and are expressed as a percentage increase over untreated cells (M,N). Magnification, 200X (A-D), 630X (E-L). Scale bars, 50µm (A-D), 15µm (E-L).

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In addition to filopodia formation, I also observed a significant increase in tubulin- labeled cellular protrusions in 1nM (Fig. 4.6B,F,M) and 100nM (Fig. 4.6C,G,M) TCAP-1-treated cells (P<0.001, one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group; n=3) as compared to untreated hippocampal cells (Fig. 4.6A,E,M). Despite the changes in filopodia formation and length, TCAP-1 treatment at 1 and 100nM did not significantly affect overall cell size (Fig. 4.6P). In addition, cells treated with 100nM SC-TCAP-1 showed no significant change in tubulin-labeled cellular protrusions (Fig. 4.6M), filopodia per cell (Fig.

4.6N), mean filopodia length (Fig. 4.6O) or cell size (Fig. 4.6P) as compared to untreated hippocampal cells (p>0.05, one-way ANOVA with Bonferroni’s post hoc test, minimum 100 cells per group; n=3).

4.4.5 TCAP-1 induces ERK-dependent phosphorylation of Stathmin

The dystroglycan complex is a multifunctional scaffold capable of interacting with components of the mitogen-activated protein (MAP) kinase kinase 2 (MEK2) and extracellular signal-regulated kinase (ERK) cascade (Spence et al., 2004). To elucidate whether TCAP-1 activates the MEK-ERK1/2 pathway, E14 hippocampal cells were treated with 100nM TCAP-1 and immunoblotted for levels of phosphorylated MEK and ERK. A TCAP-1 concentration of

100nM was used for these studies, as this concentration provided the most efficacious and acute action of the peptide. TCAP-1-treated hippocampal cells induced a significant activation of phosphorylated MEK after 1 min, with the strongest increase of MEK phosphorylation occurring after 5 min of TCAP-1 treatment (Fig. 4.7A,D). This corresponded with a similar activation of phosphorylated ERK1/2, the downstream kinase of MEK, with peak phosphorylation occurring from 5 to 30 min (Fig. 4.7B,E). There was no significant change in levels of phosphorylated

141 focal adhesion kinase (FAK), in TCAP-1 treated hippocampal cells (Fig. 4.7A,D), indicating that

TCAP-1 is not interacting through this pathway.

The activity of stathmin is dynamically regulated by serine phosphorylation on four highly conserved serine residues (serine-16, -25, -38, and -63) and is a major target of several classes of kinases, such as the mitogen-activated protein kinase (MAPK/ERK) family (Manna et. al., 2009). Mouse E14 hippocampal cells treated with 100nM TCAP-1 demonstrated a robust increase in phosphorylation at serine-25 after 5 min, with peak phosphorylation occurring from

10 to 60 min after TCAP-1 treatment (Fig. 4.7B,F).

However, there was no significant induction of phosphorylation of stathmin at serine-38 over the course of TCAP-1 treatment (Fig. 4.7B,F). In order to determine whether the phosphorylation of stathmin is mediated by the MEK-ERK1/2 signaling cascade induced by

TCAP-1, E14 hippocampal cells were pretreated with the MEK inhibitor, U0126, prior to incubation with TCAP-1. The inhibition of the MEK-ERK1/2 signaling pathway effectively blocked TCAP-1-induced phosphorylation of ERK1/2 and stathmin at serine-25 over the course of TCAP-1 treatment (Fig. 4.7G). Cells treated with a scrambled version of TCAP-1 peptide

(SC-TCAP-1) did not induce phosphorylation of MEK1/2, ERK1/2 nor stathmin at serine-25

(Fig. 4.7C). Immunoblot analysis with pan-MEK, -ERK and -stathmin antibodies indicated that

MEK, ERK and stathmin levels were not significantly changed during TCAP-1 treatment (Fig.

4.7).

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Figure 4.7. TCAP-1 activates the Mitogen-activated Protein Kinase (MAPK/ERK) pathway and induces ERK-dependent stathmin phosphorylation at serine-25 in E14 hippocampal cells. Cultured E14 hippocampal cells treated with water (untreated), 100nM TCAP-1 or scrambled-TCAP-1 for 0, 1, 5, 10, 30 and 60 min were analyzed by immunoblotting. A. Lysates from E14 cells treated with 100nM TCAP-1 over a time course were immunoblotted for levels of phosphorylated MEK1/2 and FAK. Immunoblotting with pan-MEK showed equivalent total MEK levels. B. Immunoblotting analysis with site specific phosphoserine stathmin (STMN) antibodies and phospho-ERK antibody. Total protein levels were also revealed by immunoblotting with pan-ERK1/2 and pan-STMN antibodies. C. Lysates from E14 cells treated with 100nM scrambled-TCAP (SC-TCAP) over a time course were immunoblotted for levels of phosphorylation of MEK1/2, ERK1/2 and STMN Ser-25 and total MEK, ERK and STMN. Phosphorylated FAK and MEK1/2 (D), phosphorylated ERK1/2 (E) and phosphorylated STMN Ser-25 and -38 (F) were quantitated by densitometric analysis and expressed as a fold increase over untreated controls. Values are the mean ± SEM (n=3, one-way ANOVA with a Bonferroni’s post hoc test, *p<0.05, **p<0.01, ***P<0.001, compared to untreated controls). G. Lysates from E14 hippocampal cells treated with TCAP-1 (100nM) and cells pretreated with U0126 (10µM, 30min) before TCAP-1 treatment (100nM) or left untreated were immunoblotted for levels of phosphorylated ERK1/2 and STMN Ser-25 as well total ERK1/2 and STMN. DMSO (0.1% (v/v)) was used as a vehicle control.

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4.4.6 TCAP-1 induces phosphorylation of p90RSK and filamin A

The actin-cross-linking protein filamin A, is a membrane-associated p90RSK substrate, which together can be mediated by the MEK-ERK1/2 signaling cascade (Woo et al., 2004). My finding of increased actin cross-linking and the recruitment of the MEK-ERK1/2 signaling cascade in TCAP-1-treated hippocampal cells raised the question of whether the p90RSK- filamin A pathway is also activated in response to TCAP-1 treatment. Immunoblot analysis in

E14 hippocampal cells treated with 100nM TCAP-1 showed a significant increase in the levels of phosphorylation of p90RSK at serine-380 after 5 min of TCAP-1 treatment (Fig. 4.8A,C).

Furthermore, TCAP-1 also induced serine phosphorylation of filamin A at serine-2152 after 1 min, with peak phosphorylation occurring from 30 to 60 min after TCAP-1 treatment (Fig.

4.8A,D). Incubation with 100nM SC-TCAP-1 did not induce serine phosphorylation of p90RSK nor filamin A (Fig. 4.8B).

To determine if the recruitment of the p90RSK-filamin A pathway was mediated by the

MEK-ERK1/2 signaling pathway, E14 hippocampal cells were pretreated with the MEK inhibitor, U0126, prior to incubation with TCAP-1. U0126 blocked TCAP-1 induced phosphorylation of p90RSK (Fig. 4.8E) throughout the course of treatment and significantly abolished serine phosphorylation of filamin A from 10 to 60 min (Fig. 4.8E,F). Immunoblot analysis with pan-RSK1/2/3 and –filamin A antibodies indicated that RSK and filamin A levels were not significantly changed during TCAP-1 treatment (Fig. 4.8).

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Figure 4.8. TCAP-1 induces ERK-dependent p90RSK phosphorylation and filamin A phosphorylation in E14 hippocampal cells. Cultured E14 hippocampal cells treated with water (untreated), 100nM TCAP-1 or scrambled-TCAP-1 for 0, 1, 5, 10, 30 and 60 min were analyzed by immunoblotting. A. Lysates from E14 cells treated with 100nM TCAP-1 over a time course were immunoblotted for levels of phosphorylated p90-RSK Ser-380 and filamin A Ser-2152 and total RSK1/RSK2/RSK3 and filamin A. B. Lysates from E14 cells treated with 100nM scrambled-TCAP (SC-TCAP) over a time course were immunoblotted for levels of phosphorylation of p90RSK Ser-380 and filamin A Ser-2152. Total protein levels were also determined by immunoblotting with RSK1/2/3 and pan-filamin A antibodies. Phosphorylated p90RSK (C) and phosphorylated filamin A (D) were quantitated by densitometric analysis and expressed as a fold increase over untreated controls. Values are the mean ± SEM (n=3, one-way ANOVA with a Bonferroni’s post hoc test, *p<0.05, **p<0.01, ***P<0.001, compared to untreated controls). E. Lysates from E14 hippocampal cells treated with TCAP-1 (100nM) and cells pretreated with U0126 (10µM, 30min) before TCAP-1 treatment (100nM) or left untreated were immunoblotted for levels of phosphorylated p90-RSK and filamin A as well total RSK1/RSK2/RSK3 and total filamin A. DMSO (0.1% (v/v)) was used as a vehicle control. F. Levels of phosphorylated p90-RSK and filamin A before and after U0126 and TCAP-1 treatment over time were quantified by densitometric analysis and expressed as a fold increase over untreated controls (vehicle). Values are the mean ± SEM (n=3, one-way ANOVA with a Bonferroni’s post hoc test, **p<0.01, ***P<0.001, compared to vehicle).

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4.5 Discussion

TCAP-1 is a member of a novel family of neuropeptides that has been highly conserved throughout evolution and is strongly expressed in the hippocampus and other nuclei in the limbic system (Wang et al., 2005). The neuromodulatory in vivo and in vitro effects of synthetic TCAP-

1 suggested a role for TCAP-1 in regulating cytoskeletal dynamics. The data described in this study strongly support the hypothesis that TCAP-1 can mediate a rapid reorganization of actin- and tubulin-based cytoskeletal elements and are associated with an increase in filopodia formation and elongation in the hippocampal cell. This study provides a molecular mechanism underlying the TCAP-1-mediated dynamic regulation of the cytoskeleton and subsequent filopodia outgrowth. I postulate that this mechanism is a dystroglycan-associated, MEK-

ERK1/2-mediated integration of stathmin and p90RSK-filamin A systems (Fig. 4.9).

In culture, FITC-labeled TCAP-1 co-localizes with β-dystroglycan at the plasma membrane. Dystroglycan is an important laminin-binding membrane receptor protein linking the actin cytoskeleton, via utrophin and dystrophin to the extracellular matrix (Winder, 2001) and is functionally related to the teneurins (Trzebiatowska et al., 2008). Although it has primarily been implicated in muscle development, the dystroglycans are also found in brain tissue (Smalheiser and Schwartz, 1987), but its role in the brain is not well understood. However, in the mouse brain, dystroglycan is predominantly localized in the neurons of the hippocampus, cerebellum, cerebral cortex, olfactory bulb and hypothalamus (Zaccaria et al., 2001). Specifically, Zaccaria et al., (2001) demonstrated that the dystroglycans were mainly localized to the somata and apical dendrites of pyramidal cells in the CA1, CA2 and CA3 regions of the mouse hippocampus, similar to that observed with teneurin-1 expression (Zhou et al., 2003; Kenzelmann et al., 2008) and TCAP-1 binding. Moreover, TCAP-1 induces changes in dendritic arborization in the CA1

146 and CA3 neurons and attenuates anxiety-like behaviour in the elevated plus maze when administered in vivo (Tan et al., 2011).

Figure 4.9. Proposed mechanism of TCAP-1 effects on the cytoskeleton in the mouse hippocampus. TCAP-1 released from neighboring cells or cleaved from the teneurins first interacts with the –dystroglycan (-DG) subunit of the dystroglycan complex. TCAP-1 binding to -DG activates the mitogen-activated protein (MAP) kinase kinase 1/2 (MEK1/2) and its downstream kinase extracellular signal-regulated kinase 1/2 (ERK1/2). Activation of ERK1/2 leads to the phosphorylation of stathmin at serine-25 and p90-Ribosomal S6 Kinase (p90-RSK) at serine-380. TCAP-1 dependent phosphorylation of stathmin in turn contributes to the regulation microtubule dynamics and organization. Phosphorylation of p90RSK, acts in part, to phosphorylate filamin A at serine-2152 which results in the regulation of actin polymerization. TCAP-1 does not activate focal adhesion kinase (FAK), suggesting that signaling through the MEK-ERK pathway is integrin-independent. The regulation of cytoskeletal dynamics by TCAP- 1 results in an increase in filopodia formation and elongation in mouse hippocampal cells.

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Dystroglycans can serve as a scaffold for the MEK-ERK1/2 signaling cascade (Spence et al., 2004). In fact, a yeast two-hybrid screen identified mitogen-activated protein MEK2 and

ERK1/2 as a β-dystroglycan interactor (Spence et al., 2004). Moreover, dystroglycan MAPK- signaling is also thought to be achieved indirectly through focal adhesion kinase (FAK) and the integrin system (Cavaldesi et al., 1999). Integrin-mediated activation of FAK regulates microtubule stabilization (Palazzo et al., 2004); however, TCAP-1 does not stimulate phosphorylation of FAK. Given that TCAP-1 co-localizes with β-dystroglycan in hippocampal cell culture and stimulates the activation of MEK1/2 and its downstream kinase ERK1/2 but not

FAK, suggests that TCAP-1 may be a novel ligand for dystroglycan signaling and regulation of cytoskeletal elements in neuronal cells (Fig. 4.9). This is further supported by the observation that TCAP-1 remains bound to the plasma membrane after prolonged incubation with FITC- conjugated-TCAP-1 suggesting that the TCAP-1 target that lies upstream of this activation is at the plasma membrane and not localized intracellularly at the focal adhesion complex.

TCAP-1 increases β-actin and β-tubulin expression in neuronal culture (Al Chawaf et al.,

2007a) and modulate dendritic arborization and spine density in the hippocampus in vivo (Tan et al., 2011). Neurite outgrowth, dendritic remodeling and the formation of spines depends on the rapid reorganization of actin filaments and dynamic regulation of tubulin cytoskeleton

(Hotulainen and Hoogenraad, 2010). I have shown that TCAP-1 increases actin polymerization and α- and β-tubulin immunofluorescent-labeling within hippocampal neurons. Although the regulation of actin cytoskeleton may be occurring through direct interaction between dystroglycan and actin, my data suggest that the rapid reorganization of α- and β-tubulin may be primarily mediated by the tubulin-destabilizing phosphoprotein, stathmin. Stathmin, also known as oncoprotein-18 (Hailat et al., 1990) is a cytosolic protein whose microtubule destabilizing

148 activity is attenuated through serine phosphorylation at four regulatory sites (serine-16,-25,-38 and -63) and is correlated with the action of multiple extracellular stimuli (Curmi et al., 1999).

When unphosphorylated, stathmin destabilizes microtubules by reducing the microtubule polymer mass through sequestration of soluble tubulin and by increasing the catastrophe frequency by binding directly to polymerized microtubules (Cassimeris, 2002). Stathmin phosphorylation can be regulated by several classes of kinases, such as the MAP kinase/ERK family, cAMP-dependent protein kinase, cyclin-dependent kinase 1 (Manna et. al., 2009) and c-

Jun N-terminal kinase (JNK; Ng et al., 2010) that target specific serine residues. Like nerve growth factor (NGF; Di Paolo et al., 1996), TCAP-1 specifically induces ERK1/2-dependent phosphorylation of stathmin at serine-25 but not serine-38, thus only partially inhibiting its tubulin-destabilizing activity and, therefore, facilitating the rapid reorganization of α- and β- tubulin to areas of cellular protrusions in hippocampal neurons. This is consistent with previous studies showing that serine-25, which is the preferential site for MAP kinase (Beretta et al.,

1993; Leighton et al., 1993), is also the major target of many neurotrophic factors such as NGF

(Di Paolo et al., 1996), epidermal growth factor and fibroblast growth factor (Doye et al., 1990).

Furthermore, stathmin mRNA is increased in the hippocampus following an acute dose of methamphetamine, a treatment which increases dendritic spines (Ujike et al., 2002). Recently, stathmin phosphorylation at serine-25 and -38 by MAP-kinase and JNK, respectively, has been implicated as a novel cytoprotective response to stress and toxic stimuli that results in cytoskeletal reorganization (Ng et al., 2010). TCAP-1 does not induce phosphorylation of stathmin at serine-38, suggesting that TCAP-1 signaling to stathmin is analogous to other neurotrophic factors such as NGF and fibroblast growth factor.

149

My observations that TCAP-1 treatment induced an apparent increase in actin polymerization and cross-linking suggested that besides dystroglycans, actin cross-linking proteins are involved in the TCAP-1 signaling pathway. Filamin A is the most widely distributed isoform of a family dimeric actin cross-linking proteins that has been well characterized (Stossel et al. 2001). Moreover, filamin A interacts with membrane receptors, small GTPases, and protein kinases (Stossel et al. 2001). Recently, filamin A associated actin cross-linking activity was shown to be highly regulated by ribosomal S6 kinase (p90RSK), a key player in the MEK-

ERK1/2 pathway (Woo et al., 2004). I have shown that TCAP-1 stimulates ERK-dependent phosphorylation of p90RSK at serine-380, which results in subsequent phosphorylation of filamin A at serine-2152. In conjunction with previously reported studies, my data suggest that

TCAP-1 stimulation of the MEK-ERK signaling cascade may control actin cytoskeletal changes via p90RSK phosphorylation of filamin A.

The role of TCAP-1 in regulating cytoskeletal dynamics and neuronal morphology may be closely associated with that of teneurin-1 because TCAP-1 binds to all cell groups of the hippocampus that show teneurin expression (Zhou et al., 2003). Moreover, my observations of

TCAP-1 induced cellular protrusions, filopodia formation and increased filopodia length, are consistent with previous studies that show that the teneurins play a key role in the formation of filopodia and neurite outgrowth (Rubin et al., 1999). Filopodia are highly dynamic actin rich structures that are used as both sensory and mechanical structures to explore the extracellular matrix and surfaces of other cells and mediate appropriate cell matrix adhesion (Wood and

Martin, 2002). In its capacity as sensory structures, filopodia are extremely sensitive and react to subtle changes in soluble or substrate-bound signaling molecules (Kater and Rehder, 1995;

Wood and Martin, 2002) as observed by the significant increase in filopodia and filopodia length

150 even with a 1nM concentration of TCAP-1. Actin and its associated proteins such as the Arp2/3 complex (Johnston et al., 2008), vinculin (Sydor et al., 1996), and myosin (Lin et al., 1996; Bohil et al., 2006) play a fundamental role in the formation, elongation and retraction of filopodia.

However, it remains to be seen whether the TCAP-1-induced increase in filopodia formation is due to an increase in the frequency of initiation and/or a reduction in the frequency of retraction.

Interestingly, the teneurins have been shown to co-localize with filamentous actin at cell-cell contact sites (Rubin et al., 2002) and the interaction between the intracellular domain of teneurin-

1 and the adaptor protein c-Cbl-associated protein CAP/Ponsin represents a possible link between these transmembrane proteins and the actin cytoskeleton (Nunes et al., 2005).

Therefore, in addition to dystroglycan and filamin A, it is possible that TCAP-1 may functionally recruit other actin-binding proteins to influence a reorganization of the actin cytoskeleton and dynamically regulate filopodia formation.

These studies delineate a novel mechanism of cytoskeletal regulation and provide new insight into the molecular mechanisms by which TCAP-1 exerts its effect in the hippocampus.

My data indicate that TCAP-1 may be a novel ligand for the dystroglycan signaling pathway, and functionally links the teneurin and dystroglycan systems. However, it remains to be seen whether

TCAP-1 functionally interacts with the dystroglycan complex and mediates cytoskeletal dynamics as part of a direct teneurin-1 juxtacrine signaling mechanism, or, if it has an independent role, either as a splice variant or post-translational proteolytic cleavage product of teneurin. Nevertheless, the TCAP-1-stimulated MEK-ERK1/2-dependent phosphorylation of stathmin and p90RSK-filamin A system suggests a mechanism by which TCAP-1 induces a rapid reorganization of both actin- and tubulin-based cytoskeletal elements which correlates with an increase in filopodia formation and elongation. I postulate that these changes in the

151 cytoskeleton are a prelude to, and are associated with changes in neuronal morphology that may underlie the mechanism that ultimately modulates TCAP-induced dendritic spine formation, inhibition of CRF-associated stress behaviours and neuronal plasticity.

4.6 References

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5 Chapter Five: In vivo effects of TCAP-1 in the testes and epididymis of the adult mouse

This chapter is in submission for publication in Journal of Histochemistry and Cell Biology

Chand D*, Colacci M, Kollara A, Brown TJ, Lovejoy DA (2012) C-terminal region of teneurin-1 co-localizes with the dystroglycan complex in the seminiferous tubules and epididymis of the adult mouse testes and regulates testicular size and testosterone production.

5.1 Abstract

Testicular size is directly proportional to fertility potential and is dependent on the integration of developmental proteins, trophic factors and sex steroids. The teneurins are a family of transmembrane glycoproteins that function as signaling molecules and cell adhesion proteins in the establishment and maintenance of the somatic gonad, gametogenesis and basement membrane integrity. Moreover, the teneurins are thought to function redundantly to the extracellular matrix protein, dystroglycan. Encoded on the last exon of the teneurin genes are a family of bioactive peptides termed the teneurin C-terminal associated peptides (TCAPs).

Previous studies indicated that TCAP-1 functions as a neuromodulatory peptide with trophic characteristics independent from the teneurins. However, little is known about the localization and relationship between the teneurin-TCAP-1 system and the dystroglycans in the male gonad.

In the adult mouse testis, immunoreactive TCAP-1 was localized to the spermatogonia and early spermatocytes, co-localizing with β-dystroglycan. In contrast, teneurin-1 was localized exclusively to the basement membranes of the seminiferous tubule and epididymis, co-localizing with α-dystroglycan. TCAP-1 binding sites were identified in the germ cell layers and spermatid regions of the seminiferous tubules, and epithelial cells of the epididymis. In vivo, TCAP-1 administration significantly increased testicular size, seminiferous tubule short-diameter, epididymis short-diameter, and testosterone levels. TCAP-1-treated mice also showed increased

157

TCAP-1 immunoreactivity in the epithelial cell layer of the caput and corpa epididymis. My data provides novel evidence of TCAP-1 localization in the testes that is distinct from the teneurins, but integrated through an association with the dystroglycan complex.

5.2 Introduction

Male fertility is highly dependent on the precisely regulated and highly specialized process of spermatogenesis in the seminiferous tubule of the testes as well as storage and maturation of spermatozoa in the epididymis. Central to the process of spermatogenesis is the migration and maturation of germ cells, which is associated with extensive restructuring of the seminiferous epithelium and dynamic changes in seminiferous tubule morphology (Cheng and

Mruk, 2002; Mruk and Cheng, 2004). This process is not only mediated by autocrine and paracrine signaling pathways but is highly dependent on the integration and control of testicular cell-cell interactions (Siu et al., 2003b; Siu and Cheng 2004).

In the mammalian testes, these cell-cell interactions are integrated at the basement membrane of the seminiferous epithelium, a modified form of the extracellular matrix constituted largely by type IV collagen and laminins (Dym 1994). Throughout the different stages of the seminiferous tubule epithelial cycle, Sertoli cells and germ cells, particularly spermatogonia are in close physical contact with the basement membrane and rely on it for structural and hormonal support (Mruk and Cheng 2004). In addition, the basement membrane, along with the rest of the extracellular matrix that forms the tunica propria, mediates intracellular communication between the seminiferous tubules and the steroidogenic interstitial cells

(Reventos et al., 1989; Saez et al., 1989). Such is the importance of the basement membrane that infertile patients with aspermatogenesis were shown to have abnormal basement membrane

158 structures (Salomon et al., 1982; Lehmann et al., 1987). Maintenance of basement membrane integrity is, therefore, crucial to spermatogenesis.

Recently, the teneurins have emerged as a novel extracellular matrix signaling system that play a significant role in development, cell-cell interaction, cell adhesion and cell migration

(Tucker and Chiquet-Ehrismann, 2006; Kenzelmann et al., 2007; Young and Leamey, 2009).

The teneurins are a family of four highly conserved, type-II transmembrane proteins that are predominantly expressed in the central nervous system (CNS) (Mieda et al., 1999; Oohashi et al.,

1999; Ben-Zur et al., 2000; Rubin et al., 2002; Zhou et al., 2003). However, recent studies have shown that the Caenorhabditis elegans teneurin ortholog, ten-1, is prominently expressed in the gonadal somatic precursor cells and plays an important role in gonad epithelialization and maintenance of basement membrane integrity (Drabikowski et al., 2005; Trzebiatowska et al.

2008). In fact, deletion of the teneurin-1 gene, ten-1, leads to gonad disorganization, germ cell leakage due to gonad rupture and sterility, similar to phenotypes reported for other basement membrane mutants such as integrin α-ina-1 (Baum and Garriga, 1997), laminin αB epi-1 (Huang et al., 2003), and dystroglycan dgn-1 (Johnson et al., 2006). The heterodimeric dystroglycan transmembrane protein binds directly to laminin (Smalheiser and Schwartz, 1987), which in turn forms a functional complex with integrin (Ido et al., 2004), and together links the extracellular matrix to the intracellular cell cytoskeleton. However, little is known about the relationship between the teneurin-1 protein and the dystroglycan complex in the mammalian testes. Thus, the relative position of these extracellular matrix proteins in the testis is of considerable interest.

The teneurins possess a complex set of functional domains (Minet et al., 1999; Feng et al., 2002; Leamey et al., 2007) and some family members may also exist as a number of splice variants (Lossie et al., 2005), or as smaller soluble proteins derived from further proteolytic

159 cleavage of the carboxy terminal regions (Kenzelmann et al., 2008). One such region is a 40- to

41-amino acid sequence encoded by the teneurin 3’ terminal exon and possesses characteristics of a cleavable bioactive peptide (Qian et al., 2004; Wang et al, 2005). This sequence was termed the ‘teneurin C-terminal associated peptide’ (TCAP). Recent evidence indicates that the TCAP region of the teneurin-1 protein acts as a novel neurotrophic factor in the rodent brain (Tan et al.,

2011b), through a dystroglycan-associated and extracellular-signal-regulated kinase (ERK1/2)- dependent signaling mechanism to regulate cytoskeletal elements as a prelude to the TCAP- mediated physiological changes (Chand et al., 2012). TCAP-1 is highly expressed in the CNS

(Wang et al., 2005), and in addition to regulating cytoskeletal elements and cellular morphology

(Al Chawaf et al., 2007; Chand et al., 2012), has also been reported to inhibit stress-induced cell death (Trubiani et al., 2007). However, the localization and action of TCAP-1 outside the brain is unresolved.

The hormonal control of spermatogenesis has been well documented (McLachlan et al.,

1995) but the factors mediating the corresponding changes in seminiferous tubule and epididymis size are less understood. Given that teneurin-1 (Oohashi et al., 1999) and the dystroglycan complex (Durbeej et al., 1998) are expressed outside of the CNS, and have related and partly redundant functions in Caenorhabditis elegans gonadal development (Trzebiatowska et al. 2008), I hypothesize that TCAP-1 may also serve a broader role in regulating gonadal morphology and function in the adult mouse. However, it is not known if TCAP-1 exerts its effects as part of a direct teneurin function, whereby TCAP-1 represents a functional region of the large teneurin-1 protein, or if it has an independent role, similar to that reported in the mouse brain (Chand et al., 2012b; Tan et al., 2011b). If TCAP-1 is structurally distinct from teneurin-1, and possesses its own complement of functional attributes, it would be expected to have distinct

160 binding sites and action in the testes. Therefore, these studies were aimed at determining the localization and action of the TCAP-1 region of teneurin-1 in the adult mouse testes, and further characterize the distribution and relationship between the C-terminal regions of teneurin-1 and the dystroglycan complex.

5.3 Materials and Methods

5.3.1 Animals. All procedures were approved by the local animal care committee and were in accordance with the Canadian Council on Animal Care. Adult male BALB/c mice (n=31,

Charles River Laboratories, Montreal, Quebec, Canada) were housed in groups of two to three in shoebox cages in a controlled environment with a 12 h light/dark cycle at a constant temperature of 21ºC. Mice were provided with standard mouse chow and water ad libitum and allowed to acclimatize to laboratory conditions for two weeks before injection.

5.3.2 Antibodies. Rabbit anti-mouse affinity-purified TCAP-1 antiserum previously characterized by (Chand et al., 2012b) was used at a dilution of 1:1000. Mouse monoclonal TCAP-1 antiserum was used at a dilution of 1:1000. Mouse monoclonal teneurin-1 antiserum (1:1000) was obtained from ABNOVA (Taipei City, Taiwan). Rabbit anti-human-α-dystroglycan antiserum (α-DG,

1:500) and rabbit anti-human-β-dystroglycan (β-DG, 1:500) were obtained from Santa Cruz

Biotechnology (Santa Cruz, CA, USA). Secondary antibodies Alexa 594 goat anti-rabbit (1:400),

Alexa 594 donkey anti-mouse (1:400) and Alexa 488 donkey anti-mouse (1:400) were obtained from Invitrogen (Burlington, ON, Canada).

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5.3.3 Synthesis of TCAP-1 Peptides. Mouse TCAP-1 and scrambled TCAP-1 (SC-TCAP-1), were synthesized using f-moc-based solid phase synthesis and purified to 95% purity (American

Peptide Company, Sunnyvale, CA, USA). The SC-TCAP-1 is a randomly varied amino acid sequence based on the amino acid composition of the mouse TCAP-1 sequence. TCAP-1 variants with lysine substituted for arginine at positions 8 or 37 (K8 and K37) were synthesized at the University of Toronto (Toronto, ON, Canada) as previously described (Wang et al., 2005).

5.3.4 TCAP-1 administration and tissue collection. Lyophilized synthetic TCAP-1 peptide was solubilized by exposure to ammonium hydroxide vapor for 2 min before dilution in sterile saline to a stock concentration of 10-4 M as previously described (Al Chawaf et al., 2007b). Injections were administered subcutaneously between 1000 h and 1200 h once daily for nine consecutive days using a syringe with a 30-gauge stainless steel needle. Mice were injected with 300 μl of

25pmol TCAP-1 (n=11), 250pmol TCAP-1 (n=10) or saline (control; n=10). On the tenth day, animals were anesthetized with 3% isoflurane, and blood was collected by cardiac puncture.

Mice were then decapitated and the epididymides and testes immediately removed. The testes were weighed and the long- and short-diameter of each testis was measured using a digital vernier caliper. The extracted epididymides and testes were then fixed overnight in 4% paraformaldehyde (PFA; Sigma-Aldrich Canada, Oakville, ON, Canada), pH 7.4, and paraffin- embedded.

5.3.5 Fecal collection. For fecal collection, animals were placed in a sterile cage, and approximately 0.5g of fresh fecal matter was collected twice daily, for 10 days using sterile forceps. Fecal boli was collected both before and 4 h after saline or TCAP-1 injection and stored

162 at -20ºC, without preservatives, until assay for steroid content. Fecal boli contaminated with urine were not collected for steroid assay.

5.3.6 Preparation of Fluoresceinisothiocyanate (FITC)-labeled TCAP-1. [K8] or [K37]-TCAP-1 and –SC-TCAP-1 were labeled with FITC according to the EZ-Label FITC Protein labeling kit (Pierce Biotechnology, Rockford, IL, USA) and as previously described (Al Chawaf et al.,

Chemi II Darkroom system (UVP) with the sequential integration function of LabWorks Image acquisition and analysis software (V4.0.0.8, Upland, CA, USA).

5.3.7 Histology and Immunofluorescent labeling. For histological and morphological analysis, paraformaldehyde-fixed paraffin-embedded sections (5 µm) of adult mice testes and epididymides across treatments were deparaffinized in xylene, rehydrated and counterstained with hematoxylin (Vector Laboratories, Burlington, ON, Canada) and eosin (Sigma-Aldrich

Canada). The sections were dehydrated and mounted with VectaMount mounting media (Vector

Laboratories).

For immunohistochemical analyses, paraformaldehyde-fixed paraffin-embedded sections

(5 µm) of adult mice testes, epididymides and seminal vesicles were deparaffinized in xylene and rehydrated. The sections were rinsed in double-distilled water and placed in 10mM citrate buffer for antigen retrieval by heating in a 750W microwave oven 4 x 5 min with gentle agitation

163 between each period. Sections were permeabilized with 0.3% Triton X-100 (Fisher Scientific,

Ottawa, ON, Canada) for 10 min. The tissue sections were blocked with 10% normal goat serum

(NGS; Vector Laboratories) in PBS for 1 h and incubated overnight at 4˚C with the indicated antibodies. After washing with PBS, sections were incubated with the appropriate secondary antibody in 0.01 M PBS for 1 h at RT. The sections were counterstained with 4',6-diamidino-2- phenylindole (DAPI; Sigma-Aldrich Canada) for 1 min and mounted with 1,4- diazabicyclo[2.2.2]octane (DABCO; Sigma-Aldrich Canada).

The specificity of the affinity purified rabbit anti-mouse-TCAP-1 TNR308 and mouse monoclonal TCAP-1 antisera were tested by pre-adsorption with a 5-fold excess of synthetic mouse TCAP-1 peptide as previously described (Chand et al., 2012b). The specificity of the teneurin-1 antiserum was tested by pre-adsorption with a 5-fold excess of synthetic mouse recombinant teneurin-1 protein fragment (ABNOVA) (Chand et al., 2012b). For controls, sections were incubated with diluted pre-immune serum, primary antibody pre-adsorbed with their respective immunizing peptides, primary antibody alone, secondary antibody alone or, neither primary nor secondary antibodies.

5.3.8 FITC-TCAP-1 binding studies. To identify TCAP-1 binding sites, paraformaldehyde-fixed paraffin-embedded sections (5 µm) of adult mice testes, epididymides and seminal vesicles from non-treated mice were deparaffinized in xylene and rehydrated. Sections were permeabilized with 0.3% Triton X-100 (Fisher Scientific) for 10 min, blocked with 10% NGS (Vector

Laboratories) in PBS for 1 h and incubated with FITC-TCAP-1 or FITC-labeled scrambled

TCAP-1 at 1:400 dilution in 0.1 M PBS containing 1% NGS for 1 h at room temperature.

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Sections were then washed with PBS, counterstained with DAPI and mounted with DABCO as described above.

5.3.9 Image and morphological analysis. Immunofluorescent-labeled tissue sections were analyzed using a Zeiss LSM 510 confocal laser-scanning microscope (Carl Zeiss MicroImaging,

Göttingen, Germany) with a Q imaging Retiga EXi camera (Q Imaging, Surrey, BC, Canada).

The resulting images were adjusted for brightness and contrast using Volocity 5.2 (Improvision,

Waltham, MA, USA) and Adobe Photoshop CS4 Extended (San Jose, CA, USA).

Hematoxylin and Eosin (H&E)-stained testes sections were analyzed using a Nikon

Optiphot microscope (Nikon Canada, Mississauga, ON). The number of open and closed lumen and the diameters of the seminiferous tubule were measured across the minor axis of their profiles (longest short-diameter) from H&E-stained testes sections from all 12 stages of the seminiferous tubule cycle using NIS Elements Software Basic v2.30 (Nikon, Mississauga, ON) by three observers blinded to the study design and as described by Wing and Christensen (1982).

All the measurements were performed in 10 randomly selected H&E-stained sections of each testis and from each mouse across all treatments. Every complete seminiferous tubule profile in a section (usually 5-8) was included in the analyses. The mean of 50 measurements for each testis and from each mouse were included in the analyses.

H&E-stained epididymides were analyzed using an Aperio ScanScope XT (Vista, CA,

USA) equipped with an Olympus 40x/0.75NA UPlanFL N objective (Shinjuku, Tokyo, Japan).

Sections were scanned to a resolution of 0.25 µm/pixel, and images visualized using Aperio

ImageScope Version 10. The short-diameter and lumen diameter were measured across the minor axis of their profiles using Aperio ImageScope Version 10 by three observers blinded to

165 the study design. All the measurements were performed in 10 randomly selected H&E-stained epididymis sections from each mouse across all treatments. The left and right epididymides were pooled as one group and only complete epididymal profiles in a section were included in the analyses. The mean of 20 measurements from the caput-corpa epididymis and the mean of 10 measurements from the cauda epididymis, from each section and from each mouse were included in the analyses.

5.3.10 Serum and fecal analysis. Blood samples obtained by cardiac puncture were allowed to coagulate and centrifuged at 2000 g for 30 min at room temperature. Serum was removed and stored at -80ºC for further analysis. Serum samples were subsequently assayed for testosterone, progesterone, prolactin and growth hormone by a Clinical Laboratory Improvement

Amendments (CLIA)-certified biomarker testing laboratory using an enzyme-linked immunosorbent assay (ELISA) (Myriad RBM, Inc., Austin, TX, USA) and as described by Lee et al., (2010).

Testosterone and corticosterone were extracted from fecal samples as previously described by Muir et al. (2001). Briefly, 0.5g of fecal boli were lyophilized, mixed with 5.0 ml of

80% methanol in distilled water and incubated overnight at 4ºC on a rotary shaker. Samples were then centrifuged at 2500 RPM for 15 min and the supernatant collected and stored at -20°C.

Supernatants were subsequently assayed for testosterone and corticosterone by ELISA at the

Center for Biological Timing and Cognition research institute (CBTC; Toronto, ON, Canada) as described by Munro and Stabenfeldt (1984).

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5.3.11 Statistical Analyses. Data were analyzed by one-way or two-way ANOVA followed by

Bonferroni’s post hoc test using GraphPad Prism Ver. 4. Mean values were obtained from two independent in vivo experiments and expressed as mean ± standard error of the mean (SEM).

Serum hormone data were analyzed by Kruskal-Wallis test with Dunn’s post hoc test using

GraphPad Prism Ver. 4. Data were considered statistically significant when ***P<0.001,

**P<0.01 or *P<0.05.

5.4 Results

5.4.1 Teneurin-1 and TCAP-1 localization in the mouse testis.

The localization of TCAP-1 using rabbit-anti-mouse TCAP-1 antiserum was examined to determine the distribution of TCAP-1 in the adult mouse testis, and compared to that of immunoreactive-teneurin-1. In adult mouse testis, immunofluorescent labeling with teneurin-1 antiserum localized the larger type-II transmembrane teneurin-1 protein exclusively to the basement membrane of the seminiferous tubule and surrounding interstitial cells (Fig.

5.1A,B,C,J; white arrows). Immunoreactive-teneurin-1 was detected over the entire cell surface of the seminiferous epithelium that is enclosed by the basement membrane.

However, fluorescent labeling with TCAP-1 antiserum showed intense TCAP-1- immunoreactivity specifically in germ cells (Fig. 5.1E,F,G,K). TCAP-1-immunoreactivity was strongest in spermatogonia and early spermatocytes and was exclusively localized and uniformly distributed in the cytoplasm of these spermatogenic cells (Fig. 5.1E,F,G,K). Some punctate-like

TCAP-1-immunoreactivity was also observed near the spematid regions (Fig. 5.1E,F,G).

Teneurin-1 and TCAP-1 co-localization studies showed that immunoreactive-TCAP-1 did not

167 co-localize with immunoreactive-teneurin-1 (Fig. 5.1I-L) indicating that in the mouse seminiferous tubule, TCAP-1 localization is distinct from the larger teneurins. In the seminiferous tubules, immunoreactive-TCAP-1 (Fig. 5.1H) and -teneurin-1 (Fig. 5.1D) were completely abolished with the respective pre-adsorbed antisera.

Figure 5.1. Teneurin-1 and TCAP-1-immunoreactivity in the seminiferous tubule of the mouse testis. Immunofluorescent-labeling with monoclonal teneurin-1 antiserum (green) on adult mouse testis distinctly localized teneurin-1 to the basement membrane of the seminiferous tubule and interstitial cells (B, C and J; white arrows). However, sections labeled with TCAP-1 TNR308 antiserum (red) showed strong TCAP-1-immunoreactivity in the cytosol of spermatogonia and early spermatocytes (F, G and K). Teneurin-1 and TCAP-1 did not co- localize in sections co-labeled with teneurin-1 and TCAP-1 antisera (L). Immunoreactivity was not observed in sections incubated with pre-adsorbed teneurin-1 antiserum (D) or pre-adsorbed TCAP-1 antiserum (H). For each fluorescence image, the corresponding DIC image is represented on the left (A, E, and I). Magnification, 200X. Scale bars, 50μm.

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5.4.2 Teneurin-1 and TCAP-1 co-localizes with dystroglycan in the testis and epididymis.

To characterize the localization of the dystroglycan complex and relationship with the C- terminal region of teneurin-1 in the mouse gonad, testis and epididymis sections were immunolabeled for either α-dystroglycan or β-dystroglycan and co-labeled for immunoreactive- teneurin-1 or -TCAP-1. In the testis, immunoreactive-teneurin-1 (Fig. 5.2A,B,E,F) and -α- dystroglycan (Fig. 5.2A,C,E,G; α-DG) were strongly localized to the basement membrane of the seminiferous tubules and surrounding interstitial cells. Moreover, α-dystroglycan- immunoreactivity was noted over the entire cell surface of the seminiferous epithelium that is enclosed by the basement membrane, similar to that observed for teneurin-1. Strong co- localization between teneurin-1 and α-dystroglycan was observed at the basement membrane and along the interstitial cells (Fig. 5.2A,D,E,H; white arrow).

In the seminiferous tubules, β-dystroglycan was also strongly localized to the spermatogonia and early spermatocytes, with weak immunoreactivity at the basement membrane

(Fig. 5.2I,K; β-DG; white arrows). Similarly, TCAP-1 was strongly localized in the cytosol of spermatogonia and early spermatocytes, with punctate-like labeling near the spermatid regions

(Fig. 5.2I,J), similar to that observed with the rabbit anti-mouse TCAP-1 antiserum (Fig. 5.1).

Strong co-localization between TCAP-1 and β-dystroglycan was observed in spermatogonia and early spermatocytes near the basement membrane of the seminiferous tubules (Fig. 5.2I,L; white arrows). However, co-localization was not observed at the basement membrane or near the spermatid regions.

In the epididymis, immunoreactive-teneurin-1 (Fig. 5.3A,B,E,F,I,J; white arrows), α- dystroglycan (Fig. 5.3E,G) and β-dystroglycan (Fig. 5.3I,K) were strongly localized to the basement membrane and smooth muscle layer of the epididymis, similar to that observed in the

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Figure 5.2. Teneurin-1 and TCAP-1 co-localizes with the Dystroglycan complex in the seminiferous tubule of the mouse testis. Teneurin-1-immunoreactivity was observed at the basement membrane of the seminiferous tubule and interstitial cells in the adult mouse testis (B and F). Similarly, testis sections labeled with α-dystroglycan (α-DG) antiserum also showed strong α-DG-immunoreactivity at the basement membrane of the seminiferous tubule and interstitial cells (C and G). Strong teneurin-1 and α-DG co-localization (yellow) was observed at the basement membrane of the seminiferous tubule and interstitial cells (D and H; white arrows). Immunofluorescent-labeling with mouse monoclonal TCAP-1 antiserum localized TCAP-1 to the cytosol of the germ cells, with strong TCAP-1-immunoreactivity in spermatogonia and punctate-like labeling near the spermatid regions (J). Immunoreactive-β-dystroglycan (β-DG) was strongly localized to the cytosol of spermatogonia and early spermatocytes, with weak immunolabeling at the basement membrane (K; white arrow). Strong TCAP-1 and β-DG co- localization (yellow) was observed in spermatogonia and early spermatocytes near the basement membrane (L; white arrow). For each fluorescence image, the corresponding DIC image is represented on the left (A, E, and I). Magnification, 200X (A-D, I-L); 630X (E-H). Scale bars, 50μm (A-D, I-L); 15μm (E-H).

170 seminiferous tubules (Fig. 5.1A,B,). In addition, punctate-like teneurin-1-immunoreactivity was also detected near the lumen of the epididymis (Fig. 5.3A,B,E,F,I,J). However, TCAP-1- immunoreactivity was not detected in the epididymis (Fig. 5.3A,C). At the basement membrane

Figure 5.3. Teneurin-1 and TCAP-1-immunoreactivity and co-localization with the dystroglycan complex in the mouse epididymis. Teneurin-1-immunoreactivity was detected along the basement membrane of the epididymis with punctate-like immunoreactivity near the lumen (B, F and J; white arrows). However, TCAP-1-immunoreactivity was not detected in the epididymis (C). Alpha-dystroglycan (G; α-DG) and β-dystroglycan (K; β-DG)-immunoreactivity was detected along the basement membrane of the epididymis. Strong co-localization (yellow) between teneurin-1 and α-DG was observed at the basement membrane (H). Teneurin-1 did not co-localize with β-DG, but appeared juxtaposed to immunoreactive-β-DG (L). Immunoreactivity was not observed in sections incubated with pre-adsorbed teneurin-1 antiserum (D). For each fluorescence image, the corresponding brightfield image is represented on the left (A, E, and I). Magnification, 200X (A-D); 630X (E-L). Scale bars, 50μm (A-D); 15μm (E-L).

171 of the epididymis, there was strong co-localization between teneurin-1 and α-dystroglycan (Fig.

5.3E,H; white arrow) but not between teneurin-1 and β-dystroglycan (Fig. 5.3I,L). In fact, teneurin-1 was localized juxtaposed to β-dystroglycan at the basement membrane of the epididymis. Preadsorption with synthetic teneurin-1 completely abolished teneurin-1- immunoreactivity in the epididymis (Fig. 5.3D).

5.4.3 TCAP-1 binding in the mouse testis and epididymis.

Mouse testes cross-sections were incubated with FITC-labeled TCAP-1 in order to identify TCAP-1 binding sites in the mouse testis. Similar to the localization of immunoreactive-

TCAP-1, FITC-labeled TCAP-1 bound strongly to the spermatogonia and early spearmatocyte germ cell layer along the basement membrane (Fig. 5.4A,B). However, unlike TCAP-1- immunoreactivity, FITC-labeled TCAP-1-binding was mainly associated within the nucleus of

Figure 5.4. FITC-[K8]-TCAP-1 binding in the seminiferous tubule of the mouse testis. Sections incubated with FITC-labeled TCAP-1 showed strong TCAP-1 binding to the spermatogonia and early spermatocyte germ cell layer (A and B). FITC-TCAP-1 binding was mainly associated within the nucleus of spermatogonia and spermatocytes (B). FITC-TCAP-1 also labeled large irregularly shaped granules in the spermatid region (C). Immunofluorescence was not observed in sections incubated with FITC-labeled scrambled TCAP-1 (FITC-SC-TCAP- 1) (D). Magnification, 200X (D); 400X (A and B) and 630X (C). Scale bars, 25μm (A and B); 15μm (C) and 50μm (D). spermatogonia and early spermatocytes (Fig. 5.4A,B). In addition, TCAP-1 binding sites were also detected near the lumen of the seminiferous tubule (Fig. 5.4C). FITC-TCAP-1 strongly labeled large irregularly shaped granules near the spermatid regions of the seminiferous tubule

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(Fig. 5.4C). FITC-labeled TCAP-1 did not label the basement membrane, sertoli cells, spermatozoa or interstitial cells. Fluorescence was not observed in testes sections incubated with

FITC-labeled scrambled TCAP-1 control (Fig. 5.4D).

In the epididymis, FITC-labeled TCAP-1 bound exclusively to the principal epithelial cells and smooth muscle cells of the caput (Fig. 5.5A,B,C), corpa (Fig. 5.5,D,E,F) and cauda

(Fig. 5G,H,I) epididymis. FITC-labeled TCAP-1 binding appeared strongest in the caput and

Figure 5.5. FITC-[K8]-TCAP-1 binding in the mouse epididymis. Sections incubated with FITC-labeled TCAP-1 showed strong TCAP-1 binding to the principal epithelial cells and smooth muscle cells of the caput epididymis (B and C). Moderate FITC-TCAP-1 binding was observed in the principal epithelial cells and smooth muscle cells of the corpa epididymis (E and F), whereas weak FITC-TCAP-1 binding was observed in the cauda epididymis (H and I). For each fluorescence image, the corresponding DIC image is represented on the left (A, D, and G). Magnification, 400X. Scale bars, 25μm.

corpa epididymis, with moderate binding in the cauda epididymis. FITC-labeled TCAP-1 did not bind to the stereocilia or spermatozoa in the epididymis (Fig. 5.5). Fluorescence was not

173 observed in epididymis sections incubated with FITC-labeled scrambled TCAP-1 control (data not shown).

5.4.4 Localization of TCAP-1 and TCAP-1 binding sites in the seminal vesicles.

Immunofluorescent-labeling with TNR308 TCAP-1 antiserum showed that TCAP-1- immunoreactivity was not present in the seminal vesicles (Fig. 5.6A,B). Similarly, incubation with FITC-labeled TCAP-1 showed a lack of TCAP-1 binding sites in the seminal vesicles (Fig.

5.6C,D).

Figure 5.6. Localization of TCAP-1-immunoreactivity and FITC-[K8]-TCAP-1 binding sites in the mouse seminal vesicles. Sections immunolabeled with TNR308 TCAP-1 antiserum showed no TCAP-1-immunoreactivity in the seminal vesicles (B). Sections incubated with FITC-labeled TCAP-1 also showed a lack of TCAP-1 binding sites in the seminal vesicles (D). For each fluorescence image, the corresponding brightfield image is represented on the left (A, and C). Magnification, 200X. Scale bars, 50μm.

5.4.5 TCAP-1-immunoreactivity in the epididymis of TCAP-1-treated mice.

TCAP-1-immunoreactivity was undetectable in the caput (Fig. 5.7A), corpa (Fig. 5.7D) and cauda (Fig. 5.7G) epididymides from saline-treated mice. However, mice treated with

25pmol (Fig. 5.7B,E,H) or 250pmol (Fig. 5.7C,F,I) TCAP-1 showed increased TCAP-1- immunoreactivity in the cytosol of the principal cells of the caput (Fig. 5.7B,C) and corpa (Fig.

5.7E,F) epididymis as compared to saline-treated mice (Fig. 5.7A, caput; 5.7D, corpa). The increased TCAP-1-immunoreactivity in the caput and corpa epididymis appeared to be greatest

174 in mice treated with 250pmol TCAP-1 (Fig. 5.7C,F) as compared to the 25pmol TCAP-1-treated mice (Fig. 5.7B,E) and saline-treated mice (Fig. 5.7A,D). There was no apparent change in

TCAP-1-immunoreactivity in the cauda epididymis among saline-treated (Fig. 5.7G), 25pmol

Figure 5.7. TCAP-1-immunoreactivity in the epididymides of TCAP-1-treated mice. Immunofluorescent-labeling with TNR308 TCAP-1 antiserum (red) on epididymides from saline-treated mice showed no TCAP-1-immunoreactivity in the caput (A), corpa (D) or cauda (G) epididymis. However, TCAP-1-immunoreactivity was detected in the cytosol of the principal cells in the caput (B and C) and corpa (E and F) epididymis of TCAP-1-treated mice. TCAP-1- immunoreactivity was strongest in the caput epididymis from 25pmol TCAP-1-treated (B) and 250pmol TCAP-1-treated (C). In the corpa epididymis, moderate TCAP-1-immunoreactivity was detected in 250pmol TCAP-1-treated mice (F), whereas comparatively weaker TCAP-1- immunoreactivity was observed in corpa epididymis from 25pmol TCAP-1-treated mice (E). The increased TCAP-1-immunoreactivity in the caput and corpa epididymis appeared to be greatest in 250pmol TCAP-1-treated mice (C and F) as compared to the 25pmol TCAP-1-treated (B and E) and saline-treated (A and D) mice. There was no apparent change in TCAP-1- immunoreactivity in the cauda epididymis among saline-treated (G), 25pmol TCAP-1-treated (H) or 250pmol TCAP-1-treated (I) mice. Magnification, 200X. Scale bars, 50μm.

175

TCAP-1-treated (Fig. 5.7H) or 250pmol TCAP-1-treated (Fig. 5.7I) mice. In the epididymis of

TCAP-1-treated mice, the increased TCAP-1-immunoreactivity was strongest in the caput epididymis (Fig. 5.7B,C) as compared to the corpa (Fig. 5.7E,F) and cauda epididymis (Fig.

5.7H,I). In the corpa epididymis, moderate TCAP-1-immunoreactivity was detected in 250pmol

TCAP-1-treated mice (Fig. 5.7F), whereas comparatively weaker TCAP-1-immunoreactivity was observed in corpa epididymis from 25pmol TCAP-1-treated mice (Fig. 5.7E).

5.4.6 TCAP-1 increases testis size.

Adult male mice were injected with 25pmol TCAP-1, 250pmol TCAP-1 or saline

(control) daily for 9 days to elucidate a possible role for TCAP-1 in the gonads. TCAP-1-treated mice showed increased testicular protrusion at both 25pmol TCAP-1 (Fig. 5.8B) and 250pmol

TCAP-1 (Fig. 5.8C) treatment as compared to saline-treated mice (Fig. 5.8A). To determine whether TCAP-1 affected testicular size, both the long- and short-diameter of the extracted testes were measured across treatments.

In mice treated with 25pmol of TCAP-1, there was no significant increase in the long- diameter of the left testis (7.304 ± 0.068mm) or right testis (7.183 ± 0.100mm) as compared to saline-treated mice (7.245 ± 0.094mm, left testis; 6.946 ± 0.189mm, right testis) (Fig. 5.8F).

However, treatment with 250pmol of TCAP-1 significantly increased the long-diameter of both the left testis (7.693 ± 0.133mm, p<0.05, two-way ANOVA with a Bonferroni’s post hoc test) and right testis (7.626 ± 0.173mm; p<0.01, two-way ANOVA with a Bonferroni’s post hoc test) as compared to saline-treated mice (Fig. 5.8F). Interestingly, in the left testes of mice treated with 250pmol TCAP-1, the increased long-diameter was not significantly greater than that measured in mice treated with 25pmol TCAP-1 (p>0.05, two-way ANOVA with a Bonferroni’s

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Figure 5.8. TCAP-1 treatment increases testicular size. Representative images of mouse testes from saline-treated (A), 25pmol TCAP-1-treated (B) and 250pmol TCAP-1-treated mice (C). TCAP-1 treatment had no significant effect on overall body mass (D; one-way ANOVA with a Bonferroni’s post hoc test; p>0.05, compared to saline-treated mice) or testes mass (E; two-way ANOVA with a Bonferroni’s post hoc test; p>0.05, compared to saline-treated mice). There was no significant change in the long-diameter of left or right testis from 25pmol TCAP-1-treated mice as compared to saline-treated mice (F). However, treatment with 250pmol of TCAP-1 significantly increased the long-diameter of both the left testis and right testis (two-way ANOVA with a Bonferroni’s post hoc test, *p<0.05, **p<0.01, compared to saline-treated mice) (F). In the left testes of mice treated with 250pmol TCAP-1, the increased long-diameter was not significantly greater than that measured in mice treated with 25pmol TCAP-1 (p>0.05, two-way ANOVA with a Bonferroni’s post hoc test). Only in the right testis was the increased long- diameter in 250pmol TCAP-1-treated mice significantly greater than that measured in 25pmol TCAP-1-treated mice (p<0.05, two-way ANOVA with a Bonferroni’s post hoc test) (F). TCAP-1 treatment had no significant effect on the mean short-diameter of the mouse testes (G; two-way ANOVA with a Bonferroni’s post hoc test; p>0.05, compared to saline-treated mice). All values are the mean ± SEM obtained from two independent experiments. Number of mice from each experiment; saline (n=10), 25pmol TCAP-1 (n=11), 250pmol TCAP-1 (n=10).

177 post hoc test). Only in the right testis was the increased long-diameter in 250pmol TCAP-1- treated mice significantly greater than that measured in 25pmol TCAP-1-treated mice (p<0.05, two-way ANOVA with a Bonferroni’s post hoc test; Fig. 5.8F). There was no significant change in the mean short-diameter of testes from 25pmol (4.994 ± 0.095mm, left testis; 4.839 ±

0.076mm, right testis) or 250pmol (4.982 ± 0.142mm, left testis; 5.095 ± 0.126mm, right testis)

TCAP-1-treated mice as compared to the short-diameter of testes measured from saline-treated mice (4.824 ± 0.005mm, left testis; 4.959 ± 0.004mm, right testis) (Fig. 5.8G).

Despite the TCAP-1-induced increase in testicular size, there was no significant change in the mean testis mass of TCAP-1-treated mice at 25pmol TCAP-1 (0.090 ± 0.002g, left testes;

0.093 ± 0.003g, right testes) or 250pmol TCAP-1 (0.092 ± 0.003g, left testes; 0.096 ± 0.004g, right testes) treatment as compared to saline-treated mice (0.092 ± 0.002g, left testes; 0.090 ±

0.002g, right testes; p>0.05, two-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.8E). In addition, there was no significant change in the mean body mass across all three treatments

(p>0.05, one-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.8D).

5.4.7 TCAP-1 regulates seminiferous tubule size.

The increase in testicular size suggested that TCAP-1 may affect seminiferous tubule diameter. To characterize changes in seminiferous tubule size, the longest short-diameter of

H&E stained seminiferous tubule cross-sections representing all 12 stages of the seminiferous tubule cycle, were measured from saline-treated (Fig. 5.9A), 25pmol TCAP-1-treated (Fig. 5.9B) and 250pmol TCAP-1-treated (Fig. 5.9C) mice.

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Figure 5.9. TCAP-1 treatment regulates seminiferous tubule diameter. Representative images of hematoxylin and eosin (H&E) stained testis cross-sections from saline-treated (A), 25pmol TCAP-1-treated (B) and 250pmol TCAP-1-treated mice (C). The mean seminiferous tubule short-diameter significantly decreased in mice treated with 25pmol of TCAP-1 as compared to saline-treated mice (D). The significant decrease in short-diameter was observed in both the left (E) and right testis (F). However, mice treated with the 250pmol of TCAP-1 showed a significant increase in the mean seminiferous tubule short-diameter as compared to saline- treated mice (D, E and F) All values are the mean ± SEM, one-way ANOVA with a Bonferroni’s post hoc test, ***P<0.001, compared to saline-treated mice (D, E and F). H&E stained seminiferous tubule sections scored for the number of open and closed lumen showed no significant change between the percentage of open and closed lumen in saline-treated mice (G). However, TCAP-1 induced a significant increase in the percentage of open lumen to closed lumen in 25pmol TCAP-1-treated mice and 250pmol TCAP-1-treated mice (G). In left testis, only the 250pmol TCAP-1-treated mice showed a significant increase in the percentage of open to closed lumen (H). There was no significant change observed in the 25pmol TCAP-1-treated or saline-treated mice (H). In the right testis, the percentage of open lumen to closed lumen was significantly greater in both the 25pmol TCAP-1-treated and 250pmol TCAP-1-treated mice (I). All values are the mean ± SEM expressed as a percentage of total lumen per treatment, *p<0.05, **p<0.01, ***p<0.001, two-way ANOVA with a Bonferroni’s post hoc test (G, H and I). All values are from two independent experiments. The mean of 50 measurements for each testis and from each mouse were included in the analyses. Number of mice from each experiment; saline (n=10), 25pmol TCAP-1 (n=11), 250pmol TCAP-1 (n=10). Magnification, 100X. Scale bars, 100μm.

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The mean seminiferous tubule short-diameter significantly decreased in mice treated with

25pmol TCAP-1 (139.1 ± 1.1µm) but significantly increased in 250pmol TCAP-1-treated mice

(157.4 ± 1.4µm) as compared to saline-treated mice (145.8 ± 1.4µm; p<0.001, one-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.9D). In the left testis, the mean seminiferous tubule short-diameter was smaller in 25pmol TCAP-1-treated mice (138.5 ± 1.4µm) but significantly larger in 250pmol TCAP-1-treated mice (157.1 ± 1.7µm) as compared to saline-treated mice

(142.6 ± 1.5µm, p<0.001, one-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.9E).

Similar significant changes in seminiferous tubule short-diameter were observed in the right testis (Fig. 5.9F). The mean seminiferous tubule short-diameter of the right testis was smaller in

25pmol TCAP-1-treated mice (136.6 ± 1.5µm) but significantly larger in 250pmol TCAP-1- treated mice (154.9 ± 1.4µm) as compared to saline-treated mice (143.6 ± 1.8µm, p<0.001, one- way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.9F).

H&E stained seminiferous tubules cross-sections, were also scored for the number of open and closed lumen, to determine whether the change in seminiferous tubule diameter also correlated with changes at the lumen. In saline-treated mice, there was no significant difference between the percentage of open (47.29 ± 3.43%) and closed lumen (52.71 ± 3.43%) (Fig. 5.9G).

However, TCAP-1 induced a significant increase in the percentage of open lumen to closed lumen in 25pmol TCAP-1-treated mice (54.89 ± 1.87%, open; 45.11 ± 1.87%, closed; p<0.05, two-way ANOVA with a Bonferroni’s post hoc test) and 250pmol TCAP-1-treated mice (66.70

± 1.25%, open; 33.30 ± 1.25%, closed; p<0.001, two-way ANOVA with a Bonferroni’s post hoc test). (Fig. 5.9G). Interestingly, in the left testis, only the 250pmol TCAP-1-treated mice showed a significant increase in the percentage of open to closed lumen (p<0.001, two-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.9H). There was no significant change in the 25pmol

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TCAP-1-treated or saline-treated mice (Fig. 5.9H). However, in the right testis, the percentage of open lumen to closed lumen was significantly greater in both the 25pmol TCAP-1-treated

(p<0.01, two-way ANOVA with a Bonferroni’s post hoc test) and 250pmol TCAP-1-treated

(p<0.001, two-way ANOVA with a Bonferroni’s post hoc test) mice (Fig. 5.9I).

5.4.8 TCAP-1 increases the size of the caput and corpa epididymis.

The presence of TCAP-1-binding sites (Fig. 5.5) and increased TCAP-1- immunoreactivity in the epididymis (Fig. 5.7) of TCAP-1-treated mice, suggest that TCAP-1 treatment may affect the size of the epididymis. Mice treated with 250pmol TCAP-1 showed a significant increase in the mean short-diameter of the caput and corpa epididymis (120.3µm ±

2.2µm) as compared to saline-treated mice (109.5 ± 2.9µm; p<0.05, one-way ANOVA with a

Bonferroni’s post hoc test) and 25pmol TCAP-1-treated mice (101.0 ± 3.3µm; p<0.001, one-way

ANOVA with a Bonferroni’s post hoc test) (Fig. 5.10A). Treatment with 25pmol TCAP-1 had no significant effect on the mean short-diameter of the caput and corpa epididymis as compared to saline-treated mice (Fig. 5.10A).

Despite significant changes in the mean short diameter of the caput and corpa epididymis,

TCAP-1 treatment did not significantly affect the short-diameter of the lumen of the caput and corpa epididymis (Fig. 5.10B). There was no significant change in the mean short-diameter of the lumen from mice treated with 25pmol TCAP-1 (49.1 ± 1.6µm) or 250pmol TCAP-1 (53.4 ±

1.2µm) as compared to saline-treated mice (52.5 ± 1.2µm) (Fig. 5.10B). However, mice treated with 250pmol TCAP-1 showed a significant increase in the basement membrane-epithelium diameter of the caput and corpa epididymis (65.5µm ± 1.8µm) as compared to saline-treated mice (57.5 ± 2.3µm; p<0.05, one-way ANOVA with a Bonferroni’s post hoc test) and 25pmol

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TCAP-1-treated mice (51.9 ± 2.4µm; p<0.001, one-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.10C). Treatment with 25pmol TCAP-1 had no significant effect on the basement membrane-epithelium diameter of the caput and corpa epididymis as compared to saline-treated mice (Fig. 5.10C).

Figure 5.10. TCAP-1 treatment increases the size of the caput and corpa epididymis. Mice treated with 250pmol TCAP-1 showed a significant increase in the mean short-diameter of the caput and corpa epididymis as compared to saline-treated mice and 25pmol TCAP-1-treated mice (A). Treatment with 25pmol TCAP-1 had no significant effect on the mean short-diameter of the caput and corpa epididymis as compared to saline-treated mice (A). TCAP-1 treatment had no significant effect on the short-diameter of the lumen of the caput and corpa epididymis (B). However, mice treated with 250pmol TCAP-1 showed a significant increase in the basement membrane-epithelium diameter of the caput and corpa epididymis as compared to saline-treated and 25pmol TCAP-1-treated mice (C). Treatment with 25pmol TCAP-1 had no significant effect on the basement membrane-epithelium diameter of the caput and corpa epididymis as compared to saline-treated mice (C). TCAP-1 treatment had no significant effect on the mean short- diameter of the cauda epididymis (D). All values are the mean ± SEM, *p<0.05, ***p<0.001, one-way ANOVA with a Bonferroni’s post hoc test, compared to saline-treated mice. Number of mice; saline (n=10), 25pmol TCAP-1 (n=11), 250pmol TCAP-1 (n=10).

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In addition, TCAP-1 treatment had no effect on the size of the cauda epididymis (Fig.

5.10D). There was no significant change in the mean short-diameter of the cauda epididymis from 25pmol TCAP-1-treated (139.1 ± 5.0µm) or 250pmol TCAP-1-treated (137.2 ± 4.2µm) mice as compared to saline-treated mice (137.0 ± 3.0µm) (Fig. 5.10D).

5.4.9 TCAP-1 increases circulating levels of testosterone.

Strong TCAP-1-immunoreactivity and binding in the seminiferous tubules and increase in testicular size in TCAP-1-treated mice suggests that TCAP-1 may also mediate steroidogenesis. Fecal samples collected daily from TCAP-1-treated mice showed strong and significant increases in testosterone as compared to saline-treated mice (Fig. 5.11A). Mice

Figure 5.11. Effect of TCAP-1 treatment on fecal testosterone and corticosterone. TCAP-1- treatment significantly increased testosterone levels as measured from fecal samples collected daily for ten days and as compared to saline-treated mice (A). Mice treated with 25pmol TCAP- 1, had significantly higher levels of fecal testosterone on day 5, day 6, day 9 and day 10 (**p<0.01, ***p<0.001, two-way ANOVA with a Bonferroni’s post hoc test, compared to saline-treated mice) (A). Mice treated with 250pmol TCAP-1 had significantly lower levels of fecal testosterone on day 3 but significantly higher levels of fecal testosterone at day 5, day 8 and day 10 (**p<0.01, ***p<0.001, two-way ANOVA with a Bonferroni’s post hoc test, compared to saline-treated mice) (A). There were no significant changes in fecal testosterone from saline- treated mice during the nine days of treatment or on the tenth day (p>0.05; one-way ANOVA with a Bonferroni’s post hoc test, compared to day 1 (A). TCAP-1 treatment had no significant effect on fecal corticosterone levels (p>0.05; two-way ANOVA with a Bonferroni’s post hoc test, compared to saline-treated mice) (B). All values are the mean ± SEM, expressed as ng/g of feces. Number of mice; saline (n=10), 25pmol TCAP-1 (n=11), 250pmol TCAP-1 (n=10).

183 treated with 25pmol TCAP-1, had significantly higher levels of fecal testosterone on day 5

(89.11 ± 5.39ng/g of feces; p<0.01), day 6, (98.20 ± 6.33ng/g of feces; p<0.001) day 9 (93.25 ±

3.79ng/g of feces; p<0.001) and day 10 (77.30 ± 2.39ng/g of feces; p<0.01) as compared to saline-treated mice (two-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.11A).

However, mice treated with 250pmol TCAP-1 had significantly lower levels of fecal testosterone on day 3 (36.60 ± 1.78ng/g of feces; p<0.01) but significantly higher levels of fecal testosterone at day 5 (113.12 ± 9.95ng/g of feces; p<0.001), day 8 (94.01 ± 9.76ng/g of feces; p<0.001) and day 10 (81.23 ± 2.77ng/g of feces; p<0.01) as compared to saline-treated mice

(two-way ANOVA with a Bonferroni’s post hoc test) (Fig. 5.11A). There were no significant changes in fecal testosterone from saline-treated mice during the nine days of treatment or on the tenth day (p>0.05; one-way ANOVA with a Bonferroni’s post hoc test, compared to day 1) (Fig.

5.11A). To corroborate my observations of TCAP-1-mediated increases in fecal testosterone, serum testosterone was also analyzed from mice after nine days of saline or TCAP-1 treatment.

Mice treated with 25pmol and 250pmol TCAP-1-treated mice had significantly higher levels of serum testosterone (p<0.05; Kruskal-Wallis test with a Dunn’s post hoc test) as compared to saline-treated mice (Table 5.1).

Table 5.1: Mean serum testosterone, progesterone, prolactin and growth hormone levels (ng/ml) from saline-treated, 25pmol and 250pmol TCAP-1-treated mice after 9 days of treatment.

All values are the mean ± SEM, expressed as ng/ml of serum; * p<0.05, Kruskal-Wallis test with a Dunn’s post hoc test. Number of mice; saline (n=10), 25pmol TCAP-1 (n=11), 250pmol TCAP-1 (n=10).

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TCAP-1 treatment had no significant effect on fecal corticosterone levels as compared to saline-treated mice (p>0.05, two-way ANOVA with a Bonferroni’s post hoc test) during the nine days of treatment (Fig. 5.11B). In addition, TCAP-1 treatment did not significantly affect levels of serum progesterone, prolactin, or growth hormone as compared to saline-treated mice (Table

5.1; p>0.05, Kruskal-Wallis test with a Dunn’s post hoc test).

5.5 Discussion

Previous studies have implicated a functional relationship between the teneurins and dystroglycan in the gonads of Caenorhabditis elegans (Drabikowski et al., 2005; Trzebiatowska et al. 2008). Furthermore, recent evidence indicates that the bioactive signaling peptide encoded on the last exon of the teneurin-1 gene, TCAP-1, acts independently from the larger teneurin-1 as trophic factor through a dystroglycan-associated ERK1/2-dependent signaling mechanism in the rodent brain (Chapter 4; Chand et al., 2012). However, less is known about the localization and functional relationship between the teneurin-TCAP-1 and dystroglycan systems in the mammalian gonad. The data described in this study provide the first evidence of TCAP-1- immunoreactivity and action outside the central nervous system and strongly support the hypothesis of a functional relationship between the C-terminal regions of teneurin-1 and the dystroglycans in the adult mouse. First, immunohistochemical studies show that teneurin-1- immunoreactivity is restricted to the basement membranes of the testis and epididymis, whereas the TCAP-1 region of teneurin-1 is independently localized to the germs cells of the seminiferous tubules. Second, I show for the first time the relative positions of the α- and β- dystroglycan subunits in the seminiferous tubule and epididymis and further characterized the relationship between the teneurin-TCAP-1 system and the dystroglycan complex. Teneurin-1 co-

185 localizes with α-dystroglycan, whereas TCAP-1 co-localizes with the β-dystroglycan subunit.

Finally, I provide novel evidence of a direct action of TCAP-1 in the adult testis and epididymis that is independent from the full length teneurin-1 protein. TCAP-1 administration modulates testicular and epididymal size and increases circulating levels of testosterone. Therefore, I postulate that the action of TCAP-1 in the adult mouse testis is closely associated with the teneurin-1-dystroglycan complex at the basement membrane, corroborating previous in vitro and in vivo studies of a functional relationship between the teneurins and dystroglycans (Drabikowski et al., 2005; Trzebiatowska et al. 2008; Chapter 4; Chand et al., 2012).

The localization of teneurin-1 to the basement membrane of the testis and epididymis is consistent with teneurin-1 playing a crucial role in maintaining gonadal basement membrane integrity (Drabikowski et al., 2005; Trzebiatowska et al., 2008) and corroborates previous studies that show the localization of the teneurin-2 extracellular domain at the cell surface and the basement membrane of the avian diencephalon and optic stalk (Tucker et al., 2001). However, my findings of strong teneurin-1-immunoreactivity at the basement membrane of the mouse seminiferous tubule is in contrast to previous studies by Oohashi et al., (1999) that showed teneurin-1-immunoreactivity near the spermatid region. This discrepancy may be due to the fact that the antiserum used recognized different epitopes (Oohashi et al., 1999) and may have been obscured by the globular arrangement of the extracellular domain of the teneurins (Feng et al.,

2002). Moreover, teneurins possess a complex set of functional domains (Minet et al., 1999;

Feng et al., 2002; Leamey et al., 2007) and some teneurin family members may also exist as a number of splice variants (Lossie et al., 2005) or as smaller soluble proteins derived from further proteolytic cleavage of the carboxy terminal regions (Kenzelmann et al., 2008). In fact, antiserum specific to the TCAP-1 region of teneurin-1, specifically localized TCAP-1 to the

186 germ cells and showed no apparent co-localization with teneurin-1 in the seminiferous tubule.

This is consistent with my findings in the mouse brain where the TCAP-1 region of the teneurin-

1 can be expressed, processed and localized independently form the full length teneurin-1

(Chand et al., 2012b).

Although the C-terminal region of teneurin-2, termed ‘Lasso’ acts as a ligand for the G- protein coupled receptor latrophilin (LPH1) in hippocampal neurons (Silva et al., 2011), the C- terminal region of teneurin-1 appears to be more closely associated with the dystroglycans

(Trzebiatowska et al., 2008; Chand et al., 2012). Dystroglycan is a transmembrane heterodimeric protein complex that consists of α- and β-dystroglycan (Ibraghimov-Beskrovnaya et al., 1992) and links the extracellular matrix protein laminin and the laminin-binding protein integrin, to the intracellular cytoskeleton and signaling proteins (Winder 2001). The dystroglycan complex is required for basement membrane assembly, adhesion and signal transduction (Higginson and

Winder, 2005). Immunohistochemical studies by Durbeej et al., (1998) using an antiserum that detected both the α- and β-dystroglycan subunits showed that dystroglycan complex was primarily confined to the basement membrane region of the seminiferous tubules where it co- localizes with laminin α1- and α2-chains and weakly labeled the most primitive spermatogenic cells, the spermatogonia (Durbeej et al., 1998). My study shows for the first time the relative positions of the α- and β-dystroglycan subunits in the seminiferous tubule and epididymis. In the seminiferous tubule, α-dystroglycan-immunoreactivity was restricted to the basement membrane, whereas the β-dystroglycan subunit was localized juxtaposed to the α-dystroglycan subunit, primarily to the spermatogonia, with weak labeling at the basement membrane region. In the epididymis, the α- and β-dystroglycan subunits were also localized juxtaposed to each other at the basement membrane.

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Despite being independently localized in the testis, the role of TCAP-1 may be closely associated with that of teneurin-1 because both TCAP-1 and teneurin-1 functionally interacts with the dystroglycan complex. Furthermore, the genetic interactions between ten-1, ina-1, dgn-1 and epi-1, suggest that teneurin, integrin, and dystroglycan have related and partly redundant functions in Caenorhabditis elegans gonadal development and maintenance of basement membrane integrity (Drabikowski et al., 2005; Trzebiatowska et al. 2008). The strong co- localization between teneurin-1 and α-dystroglycan at the basement membrane of the seminiferous tubule and epididymis, suggests that teneurin-1 may be functioning as part of the larger dystroglycan-laminin-integrin extracellular matrix complex in maintaining basement membrane integrity. The C-terminal region of teneurin-1 harbors 26 repetitive sequence motifs termed tyrosine-aspartic acid (YD)-repeats, that are strongly involved in binding carbohydrates

(Minet et al., 1999). These YD-repeats may, therefore, interact with the carbohydrate moieties on the α-dystroglycan subunit that are important for interacting with extracellular matrix proteins

(Ervasti and Campbell, 1993; Pall et al., 1996; Andac et al., 1999; Michele et al., 2002). This is further supported by the co-localization of the teneurin-2 extracellular domain with laminin in the basement membrane of some tissues (Tucker et al., 2001) and the lack of co-localization between teneurin-1 and β-dystroglycan. Alpha-dystroglycan binds to laminin (Ibraghimov-

Beskrovnaya et al., 1992), whereas β-dystroglycan serves as a scaffold for intracellular signal transduction kinases and cytoskeletal regulatory proteins (Cavaldesi et al., 1999; Chen et al.,

2003; Spence et al., 2004). The strong co-localization between TCAP-1 and β-dystroglycan in the germ cell layer of the seminiferous tubule is consistent with my previous observations that show, in vitro, TCAP-1 functionally interacting with β-dystroglycan and activating an ERK1/2- signal transduction cascade to target cytoskeletal regulatory proteins (Chapter 4; Chand et al.

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2012). Together, these studies provide evidence that the teneurin-TCAP-1 system may serve as a novel ligand for the dystroglycan complex.

The association of TCAP-1 with the dystroglycan complex, and the localization of immunoreactive-TCAP-1 and TCAP-1-binding sites to the early spermatocyte stages as well as near the spermatid region of the seminiferous tubule suggest that TCAP-1 treatment may have trophic and morphological effects in the testis during spermatogenesis. Spermatogenesis is a continuous and precisely regulated process associated with extensive changes in cell shape, size movement and differentiation. During spermatogenesis, the seminiferous epithelium undergoes extensive restructuring and is coupled with dynamic changes in seminiferous tubule size to facilitate germ cell migration and differentiation throughout the 12 stages of the seminiferous epithelial cycle (Oakberg 1956; Wing and Christensen 1982; Cheng and Mruk 2002; Mruk and

Cheng 2004). Dynamic regulation of the underlying cytoskeleton (Lie et al., 2010) and recruitment of signal transduction kinases such as focal adhesion kinase (FAK) and ERK1/2 (Siu and Cheng 2004; Xia and Cheng, 2005) are of paramount importance to the restructuring and morphological changes occurring in the seminiferous tubule. These events occur primarily at the level of the basement membrane (Siu and Cheng 2004), where actin-rich, smooth muscle-like cells reside (Clermont 1958; Baillie 1962; Ross 1967) and is consistent with my findings of a teneurin-TCAP-1-dystroglycan system at the basement membrane region.

Morphometric studies on the mammalian seminiferous epithelium by Wing and

Christensen (1982), showed a direct and functional correlation between seminiferous tubule diameter, luminal volume and stage of the seminiferous epithelial cycle. The seminiferous tubule diameter increases significantly from stages I-VIII and correlates with an increase in luminal volume and germinal epithelium (Wing and Christensen 1982). This is similar to that observed in

189 mice treated with 250pmol of TCAP-1, where increases in seminiferous tubule diameter corresponded with in an increase in lumen opening, suggesting that TCAP-1 may function to increase the size of the germinal epithelium to accommodate germ cell differentiation and migration. This is further supported by my observations in the epididymis. In the caput and corpa epididymis, TCAP-1 treatment did not affect the size of lumen but rather increased the basement membrane-principal cell width, an area that showed strong TCAP-1-binding and increased

TCAP-1-immunoreactivity after TCAP-1 treatment. The increase in TCAP-1-immunoreactivity in the caput and corpa epididymis and uptake in the seminiferous tubules could be due to a dystroglycan-associated caveolae-mediated uptake mechanism as previously reported in the brain (Chand et al., 2012b). Interestingly, TCAP-1 did not affect the size of the cauda epididymis, consistent with a lack of TCAP-1-immunoreacitvity and weak dystroglycan and teneurin-1-labeling in this region (data not shown). This suggests that TCAP-1 may play a more important role in spermatogenesis rather than the storage of spermatozoa.

Wing and Christensen (1982) also reported a significant decrease in seminiferous tubule diameter and luminal volume between stages VIII and IX. This prominent decrease in the diameter of the seminiferous tubule and luminal volume is caused by the depletion of the luminal contents due to spermiation, rather than a loss of the germinal epithelium, since the latter remains relatively constant over the duration of the decline in the cycle (Wing and Christensen 1982).

TCAP-1 binding sites have been identified near the lumen of the seminiferous tubule and mice treated with 25pmol dose of TCAP-1 showed a significant decrease in the seminiferous tubule size and lumen opening, suggesting a role for TCAP-1 in spermiation, a process that is underscored by a complex pattern of actin- and tubulin-dependent cell movements, tubulobulbar complex rearrangements, cytoplasmic elimination and myoid cell contractility necessary to effect

190 sperm release (Fawcett and Phillips 1969; Russell et al., 1989). The physiological significance of the TCAP-1-mediated changes in seminiferous tubule size remains unknown but may be intimately linked to its ability to regulate cytoskeletal elements through a dystroglycan- associated ERK1/2-dependent mechanism (Chand et al., 2012). Furthermore, interactions between the intracellular domain of teneurin-1 and the adaptor protein c-Cbl-associated protein

CAP/Ponsin represent a possible link between these transmembrane proteins and the actin cytoskeleton (Nunes et al., 2005). These studies also suggest that TCAP-1 treatment could have a dose-dependent effect similar to that observed in the rodent brain (Tan et al., 2011).

Apart from regulating seminiferous tubule and epididymal size, TCAP-1 also increased circulating levels of testosterone. Spermatogenesis and male fertility are dependent upon the presence of testosterone in the testis (Walker 2011). Steroidogenesis occurs in Leydig cells found in the interstitium, whereas spermatogenesis takes place in the seminiferous tubule (Steinberger

1971; Walker 2011) which is separated from the interstitium by the extracellular matrix- associated blood-testis barrier (Reventos et al., 1989). Communication between the cells in the seminiferous tubule and the interstitium is mediated not only by testosterone from Leydig cells, but also by trophic factors secreted from the seminiferous tubule (Saez et al., 1989) and direct cell-cell contact via extracellular matrix proteins anchored at the basement membrane (Siu and

Cheng, 2004). Together these systems act to increase the steroidogenic responsiveness and activity of the steroidogenic pathway in Leydig cells (Reventos et al., 1989).

TCAP-1 is unlikely to have any acute steroidogenic effects on Leydig cells as these cells were devoid of TCAP-1-immunoreactivity and TCAP-1-binding sites. Alternatively, TCAP-1 may be acting to increase testosterone production through a functional interaction with the extracellular matrix at the basement membrane, similar to many other testosterone stimulating

191 cytokines and growth factors expressed in the seminiferous tubule such as tumor necrosis factor-

α (TNFα; Siu et al., 2003) and insulin-like growth factor 1 (IGF-1; Saez et al., 1989). My study shows strong teneurin-1 and α-dystroglycan-labeling and co-localization in the instertitium and at the basement membrane of the seminiferous tubule. Teneurins are known to interact in a homophilic manner (Oohashi et al., 1999; Rubin et al., 2002; Bagutti et al., 2003; Leamey et al.,

2008) and to date, no ligand has been identified. These studies suggest that teneurin-1-associated communication between the seminiferous tubule and interstitium could occur either by homophilic interaction or by juxatcarine-signaling with α-dystroglycan in a similar manner to the notch signaling of serrate and delta ligands (Artavanis-Tsakonas et al., 1999; Hicks et al., 2002).

TCAP-1 co-localizes with β-dystroglycan and could potentially stimulate Leydig cell testosterone production via a functional interaction with dystroglycan complex at the basement membrane.

It remains to be seen whether the TCAP-1-mediated increase in testicular and epididymal size is due to a direct TCAP-1 trophic effect, and/or indirectly via an increase in testosterone production; both of which could be mediated via teneurin-1-dystroglycan-associated and ERK- dependent mechanisms (Chand et al., 2012; Robaire and Hamzeh, 2011). Studies have shown that high levels of testosterone in the testis stimulate an increase in seminiferous tubule lumen development (opening) through their actions on Sertoli cells, but have little effect on seminiferous tubule diameter (Bressler and Lustbader, 1978). The lack of growth hormone production by TCAP-1 suggests that the TCAP-1-mediated trophic effects are independent of growth hormone, which is known to increase testicular size and body weight (Laron et al., 1983).

Majority of the teneurin investigations have focused on the expression and function of the teneurins in the CNS (Kenzelmann et al., 2007; Young and Leamey, 2009) and little is known

192 about the distribution of teneurin-1 or the localization and action of the cleaved C-terminal fragment of teneurin-1 outside the CNS. My study shows for the first time that the teneurin-1 transmembrane protein and the C-terminal region of teneurin-1 corresponding to TCAP-1, is highly localized in the mammalian testes. In the testis, TCAP-1 is distributed in the germ cells and is structurally independently from the much larger teneurin-1 which was exclusively localized to the basement membrane. Furthermore, I highlighted the relative positions of the α- dystroglycan and β-dystroglycan subunits at the basement membrane in the seminiferous tubule and epididymis of the adult mouse and provided further evidence that functionally links the teneurin and dystroglycan systems. Moreover, this study shows that TCAP-1 may be part of a unique family of trophic factors expressed in the mammalian testes that regulate testicular size and functionally interact with the extracellular matrix at the basement membrane. Like TCAP-1,

TNFα is also expressed in the germ cells of the seminiferous tubules and plays a crucial role in regulating basement membrane dynamics (Siu et al., 2003), germ cell apoptosis (Pentikäinen et al., 2001) and Leydig cell steroidogenesis (Hong et al., 2004). The teneurin-TCAP-dystroglycan system may therefore, be part of the family of systems involved the regulation of gonadal function through the integration of developmental proteins, trophic factors and sex steroids.

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Tan LA, Al Chawaf A, Vaccarino FJ, Boutros PC, Lovejoy DA (2011) Teneurin C-terminal associated peptide (TCAP)-1 modulates dendritic morphology in hippocampal neurons and decreases anxiety-like behaviors in rats. Physiol Behav 104:199-204. Tan LA, Chand, D, De Almeida, R, Xu M, De Lannoy L, Lovejoy DA (2011b) Modulation of neuroplastic changes and corticotropin-releasing factor-associated behavior by a phylogenetically ancient and conserved peptide family. Gen. Comp Endocrinol 176:309-313. Tan LA, Xu K, Vaccarino FJ, Lovejoy DA, Rotzinger S (2009) Teneurin C-terminal associated peptide (TCAP)-1 attenuates corticotropin-releasing factor (CRF)-induced c-Fos expression in the limbic system and modulates anxiety behavior in male Wistar rats. Behav Brain Res 201:198-206. Trubiani G, Al Chawaf A, Belsham DD, Barsyte-Lovejoy D, Lovejoy DA (2007) Teneurin carboxy (C)-terminal associated peptide-1 inhibits alkalosis-associated necrotic neuronal death by stimulating superoxide dismutase and catalase activity in immortalized mouse hypothalamic cells. Brain Res 1176:27–36. Trzebiatowska A, Topf U, Sauder U, Drabikowski K, Chiquet-Ehrismann R (2008) Caenorhabditis elegans teneurin, ten-1, is required for gonadal and pharyngeal basement membrane integrity and acts redundantly with integrin ina-1 and dystroglycan dgn-1. Mol Biol Cell 19:3898-3908. Tucker RP, Chiquet-Ehrismann R (2006) Teneurins: a conserved family of transmembrane proteins involved in intercellular signaling during development. Dev Biol 290:237-245. Walker WH (2011) Testosterone signaling and the regulation of spermatogenesis. Spermatogenesis 1:116-120. Wang L, Rotzinger S, Barsyte-Lovejoy D, Qian X, Elias CF, Bittencourt JC, De Cristofaro A, Wang NC, Belsham D, Vaccarino F, Lovejoy DA (2005) Teneurin proteins possess a carboxy terminal corticotropin-releasing factor-like sequence that modulates emotionality and neuronal growth. Mol Brain Res 133:253-265. Winder SJ (2001) The complexities of dystroglycan. Trends Biochem Sci 26:118-124. Wing TY, Christensen AK (1982) Morphometric studies on rat seminiferous tubules. Am J Anat. 165(1):13-25. Xia W, Cheng CY (2005) TGF-beta3 regulates anchoring junction dynamics in the seminiferous epithelium of the rat testis via the Ras/ERK signaling pathway: An in vivo study. Dev Biol 280:321-343. Young TR, Leamey CA (2009) Teneurins: important regulators of neural circuitry. Int J Biochem Cell Biol 41:990-993. Zhou XH, Brandau O, Feng K, Oohashi T, Ninomiya Y, Rauch U, Fässler R (2003) The murine Ten-m/Odz genes show distinct but overlapping expression patterns during development and in adult brain. Gene Expr Patterns 3:397-405.

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6 Chapter Six: Significance of findings, future studies and conclusions

6.1 Abstract

The studies completed in this thesis have provided novel insight on the processing and distribution of TCAP-1 in the mouse and have elucidated a TCAP-1-mediated signalling mechanism that provides new insight into the molecular mechanisms by which TCAP-1 exerts its biological functions in the CNS. Furthermore, I also showed for the first time, that the TCAP-

1 region of teneurin-1 has potent biological effects outside the CNS, particularly in the adult male testis of the mouse, further expanding on the functions attributed to the teneurin-TCAP-1 system. When the TCAP sequences were first discovered they were annotated as being encoded by the last exon of the teneurin genes. My studies show that the C-terminal region of teneurin-1 corresponding to the TCAP-1 sequence can be transcribed independently from the full-length teneurin-1. This was further supported by the independent localization of the TCAP-1 region to the cytosol and the teneurin-1 protein to the plasma membrane in both cell culture and tissues, suggesting that TCAP-1 may be processed through the non-classical secretory pathway. Using an embryonic hippocampal cell line model, I found that TCAP-1 associates with the dystroglycan complex, activates a MEK-ERK1/2-signalling cascade and is subsequently internalized by caveolae-mediated endocytosis. The activation of the dystroglycan-associated, MEK-ERK1/2 signalling cascade results in the phosphorylation and partial inhibition of a tubulin-destabilizing binding protein known as stathmin, and the activation and recruitment of the actin cross-linking p90RSK-filamin A system. This corresponded with TCAP-1-induced actin polymerization and tubulin reorganization in immortalized hippocampal cells and correlated with increases in cell

200 protrusions and filopodia formation and outgrowth. These changes in the cytoskeleton could be a prelude to, and associated with changes in neuronal morphology that may underlie the mechanism that ultimately modulates TCAP-1-induced dendritic spine formation, inhibition of

CRF-associated stress behaviours and neuronal plasticity. One of the most significant findings outlined in this thesis is the evidence of teneurin-1 and TCAP-1 immunoreactivity, TCAP-1 binding sites and in vivo functions in the adult male testis and epididymis. This represents the first evidence of TCAP-1 outside the CNS. Consistent with my studies in the mouse hippocampus, TCAP-1 in the testis also associated with the β-dystroglycan subunit of the dystroglycan complex and modulated testis and epididymal size. Interestingly, I found that teneurin-1 co-localizes with the α-dystroglycan subunit, further confirming a functional link between the C-terminal region of teneurin-1 and the dystroglycan complex in the gonads.

However, the results of this thesis, also raises a number of interesting questions about its function and clinical applicability in development, metabolism, stress and reproduction.

6.2 Teneurin-TCAP relationship: An evolutionary conserved system.

The evolutionary origin of the TCAP region at the carboxy terminus of the teneurin proteins has not been fully established, but was initially suggested to be the result of an ancestral gene duplication that became associated with the carboxy terminus of the teneurins (Qian et al.,

2004; Lovejoy et al., 2006). Gene duplication drives the evolution and functional expansion of protein families in eukaryotes. However, it has been recently postulated that the teneurins are a complex hybrid of fusion proteins that evolved in the choanoflagellate Monosiga brevicollis, the group of organisms most related to metazoans via horizontal gene transfer from both prokaryotic and diatom or algal genes (Tucker et al., 2012). The cysteine rich region of the teneurins may

201 have been acquired from diatoms, whereas the NHL domain and YD-repeats may have been acquired from prokaryotes (Tucker et al., 2012). If TCAP is present in prokaryotes then it too may have been acquired via horizontal gene transfer, perhaps in an effort to improve the integration of growth and homeostatic mechanisms. The Choanoflagellate, Monosiga brevicollis, possesses a single teneurin gene, much like that of Caenorhabditis elegans and Ciona intestinalis

(Drabikowski et al., 2005; Tucker et al., 2012). Therefore, the evolution of the teneurin-TCAP families may be a combination of horizontal gene transfer in prokaryotes, followed by gene duplication in eukaryotes.

The objectives completed in this thesis, support an independent role for TCAP-1. TCAP-

1 is structurally distinct from the teneurin-1 protein, possesses distinct binding sites and its own complement of functional attributes. Moreover, all vertebrate teneurins possess a conserved furin cleavage site 100 amino acids from the C-terminal end (Tucker and Chiquet-Ehrismann, 2006) that could liberate the TCAP sequence and allow it to function as an independent signalling molecule (Qian et al., 2004; Wang et al., 2005). However, the association of TCAP to the teneurins may have added to its complement of functional attributes. In some cases, as observed in the testis and epididymis, TCAP-1 appears to be closely associated with that of the teneurins as the C-terminal region of teneurin-1 and TCAP-1 are intimately linked to the dystroglycan complex. The existence of the full-length transcription of teneurin-1 that includes the TCAP-1- containing exon, the existence of a 250-300 kDa immunoreactive forms of TCAP-1 and weak co-localization between teneurin-1 and TCAP-1 at the plasma membrane, suggests that in some cases, the TCAP-1 sequence may, indeed, be translated and function as part of the larger teneurin-1 protein. In addition, TCAP-1 shows some overlapping and complimentary expression

202 patterns with that of the teneruins in some regions of the rodent brain (Wang et al., 2005; Zhou et al., 2003; Li et al., 2006).

In the protochordate Ciona intesinalis, the single teneurin gene encodes a teneurin sequence that is structurally similar to mouse teneurin-1 (Tucker et al., 2012). However, the

Ciona intesinalis teneurin sequence possesses fewer NHL repeats and a third furin cleavage site located between the first and second NHL repeat, suggesting that the orientation and arrangement of the extracellular domain is less complex as compared to vertebrates. In fact, my preliminary immunofluorescent studies in Ciona intesinalis tissue sections with anti-mouse

TCAP-1 antiserum localized the TCAP region to areas that are known to strongly express the teneurin protein in vertebrates (Fig. 6.1). TCAP was localized to the neuromuscular junction of the intestinal tract (Fig. 6.1B,C), an area in metazoans that is rich in dystroglycan (Bogdanik et al., 2008) and whose function and structural integrity is dependent on teneurin signalling (Mosca et al., 2012). In the Ciona intesinalis testis, TCAP-immunoreactivity was localized to the basement membrane and spermatid regions (Fig. 6.1E,F). Similarly, the Ciona intesinalis sperm duct, an area that is analogous to the mouse epididymis, localized TCAP-1 to the basement membrane (Fig. 6.1H,I). In the ovarian follicle, TCAP-labelling was strongest in the oocyte (Fig.

6.1K, L). It may not have been possible to observe a strong immunoreactive-TCAP-1 signal on the plasma membrane in cell culture or at the basement membrane in mouse tissue, because the epitope may have been obscured by the globular arrangement of the extracellular domain of the teneurins (Feng et al., 2002). Assuming this to be case, then TCAP-1 may act as a functional unit of teneurin by remaining tethered to it, in a manner similar to the notch signaling of serrate and delta ligands (Artavanis-Tsakonas et al., 1999; Hicks et al., 2002). Nevertheless, TCAP region of

203 the teneurins are highly conserved and are important in the development and survivability of a variety of species (Lovejoy et al., 2006).

Figure 6.1. TCAP-immunoreactivity in Ciona intestinalis. TCAP was localized to the neuromuscular junction in the intestines (B, C), basement membrane and spermatid region of the testis (E, F), basement membrane of the sperm duct (H, I) and in the oocyte of the ovarian follicle (K, L). Corresponding DIC images are represented on the left (A, D, G and J). Corresponding images with DAPI overlay (B, E, H and K). Magnification, 200X. Scale bars, 50μm. (Chand, Colacci, Sephton and Lovejoy, manuscript in preparation).

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6.2.1 The teneurin-dystroglycan complex and the putative TCAP-1 receptor. All four teneurins have the ability to homo- and hetero-dimerize, a characteristic that may be related to a receptor-like function (Feng et al., 2002). Western blot analysis suggested that the

TCAP-1 peptide may form polymers, particularly when in high concentrations. Protein aggregation is known to confer stability and it is also a prerequisite in the mechanism for storage in secretory granules for many peptides such as prolactin and growth hormone (Dainnies, 2002).

A strong tendency to form aggregates may also account for the larger immunoreactive-TCAP-1- related peptides observed on the western blot and an inability to detect the free peptide.

Figure 6.2. MALDI quadrupole time-of-flight analysis of synthetic TCAP-1. MALDI quadrupole time-of-flight analysis of synthetic TCAP-1 revealed that TCAP-1 was mainly present in its monomeric with a mass to charge ratio (m/z) of 4688. TCAP-1 was also detected at m/z 9381, m/z 14069 and m/z 18752. (Nock, Chand, Ackloo and Lovejoy, manuscript in preparation).

A MALDI quadrupole time-of-flight analysis performed on 5 ng/mL, 1 mg/mL and 5 mg/mL of synthetic TCAP-1 confirmed the ability of TCAP-1 to bind to itself and form polymers (Fig. 6.2). The 5 ng/mL sample showed two signals, one at mass to charge ratio (m/z)

4688 and one at m/z 4742. The 1 mg/mL sample showed four major signals that were clearly multiples of m/z 4688. They were m/z 9381, m/z 14069, m/z 18752 and m/z 23444 (Fig. 6.2). As

205 expected, the highest concentration showed a similar pattern to the mid-range sample but with higher intensities.

The ability for TCAP-1 to bind to itself, also suggests that TCAP-1 could bind to the teneurins at the plasma membrane as a prerequisite to internalization and/or the activation of downstream signaling pathways. This is supported by several pieces of evidence. First, current binding studies suggest that TCAP-1 can bind to the C-terminal region of teneurin-1 (Song,

Chand and Lovejoy, unpublished findings). Second, I have established that the C-terminal region of teneurin-1 functionally interacts with the dystroglycan complex. Teneurin-1 co-localizes with the laminin-binding α-dystroglycan subunit, whereas TCAP-1 functionally interacts with β- dystroglycan, the cytoskeletal interacting subunit of the dystroglycan complex and intracellular scaffold for the MEK-ERK1/2 signalling cascade of which TCAP-1 activates. In addition, previous studies have shown that teneurin-1 and dystroglycan have related and partly redundant functions in the maintenance of basement membrane integrity and cell adhesion (Drabikowski et al., 2005; Trzebiatowska et al. 2008). Third, the overexpression of teneurin or TCAP results in similar phenotypes such as increased filopodia formation, cellular adhesion and neurite outgrowth (Rubin et al., 1999; Al Chawaf et al., 2007a; Tan et al., 2012). Lastly, mouse E14 hippocampal cells treated with 1nM or 100nM TCAP-1 for 1 h, results in the internalization of both the teneurin-1 (Fig. 6.3) and β-dystroglycan proteins (data not shown) from the plasma membrane to the cytosol.

I have shown that the internalization of TCAP-1 occurs via a dynamin-dependent caveolae-mediated endocytosis. Interestingly, both dynamin (Zhan et al., 2005) and caveolin-3

(Sotgia et al., 2000) bind directly to the C–terminal cytoplasmic tail of β-dystroglycan suggesting that the internalization of the teneurin is intimately linked to the dystroglycans. Recent evidence

206 has shown that the C-terminal region of teneurin-2, termed ‘Lasso’ acts as a novel ligand to the

G-protein coupled receptor (GPCR), latrophilin 1 (LPH1) in hippocampal neurons (Silva et al.,

2011). However, TCAP-1 does not appear to bind latrophilin (Song, Chand and Lovejoy, unpublished findings) nor does latrophilin associate with dynamin or caveolin-associated endocytotic machinery.

Figure 6.3. Internalization of teneurin-1 by TCAP-1. In untreated mouse E14 hippocampal cells, teneurin-1 was mainly localized to the plasma membrane (B). Hippocampal cells treated with 1nM (D) or 100nM (F) TCAP-1 showed strong punctate-like teneurin-1-immunoreactivity in the cytosol. In 100nM TCAP-1-treated cell, teneurin-1-immunreacitivity was primarily cytosolic (F). Corresponding DIC images are represented on the left (A, C, E). Magnification, 630X. Scale bars, 20μm. (Chand and Lovejoy, unpublished data).

The intimate relationship between teneurin-1 and the dystroglycan complex may represent part of the putative TCAP-1-receptor complex. If that is the case, then TCAP-1 may signal via autocrine, paracrine or juxtacrine manners. Each of the signalling possibilities could result in its own unique biological outcome or degree of TCAP-1 bioactivity, thus adding another level of biological control. The studies presented in this thesis used models that already expressed teneurin-1 and dystroglycan. The extent to which TCAP-1 relies on teneurin-1 for any

207 of its biological effects can be determined in future studies using teneurin knockdown models and subsequent treatment with synthetic TCAP-1.

6.2.2 Further interactions at the plasma membrane: TCAP-1 co-localizes with 17 β-

hydroxysteroid dehydrogenase 10 (HSD17β10).

In addition to strong FITC-labelled TCAP-1 co-localization with β-dystroglycan and caveolin, FITC-labelled TCAP-1 also showed strong co-localization with the mitochondrial enzyme 17 β-hydroxysteroid dehydrogenase 10 (HSD17β10) at the plasma membrane (Fig. 6.4).

HSD17β10, also known as β-amyloid (Aβ) binding alcohol dehydrogenase (ABAD) is a member of the short chain dehydrogenase-reductase family (Yang et al., 2005). Unlike other 17β-HSD members, HSD17β10 seems to be concentrated in mitochondria of both neurons and cells and is an important contributor to the regulation of metabolic homeostasis (He et al., 2001; Yang et al.,

2005). The HSD17β10 gene encodes a 27 kDa protein, which in vivo forms 108 kDa homotetramers (He et al., 2001). The strong co-localization between TCAP-1 and HSD17β10 at the plasma membrane suggests that HSD17β10 may be part of the putative teneurin-dystroglycan receptor complex and as such an important part of the TCAP-1-signalling mechanism.

TCAP-1 may be involved in the recruitment and enrichment of mitochondria to the plasma membrane, where it can regulate calcium homeostasis and provide the energy needed to facilitate filopodia elongation, neuronal development, synaptic function and dendritic spine formation and morphogenesis (Lin et al., 2007; Sung et al., 2008). The molecular mechanisms that mediate mitochondrial distribution in neurons are poorly understood, but have been shown to involve interactions between mitochondria and the microtubules and F-actin cytoskeleton

(Morris and Hollenbeck, 1995). The action of TCAP-1 on the cytoskeleton is well established,

208 and the recruitment of HSD17β10 may be part of the TCAP-1-induced signalling mechanism involved in the regulation of filopodia formation, neuronal morphology and dendritic spine density (Al Chawaf et al., 2007a; Tan et al., 2011). Moreover, recent evidence has shown that

TCAP-1 can mediate cellular metabolism and energy levels (Xu and Lovejoy, unpublished findings) as well as calcium signalling (Chand and Lovejoy, data not shown) in neuronal cell culture, further implicating a strong functional relationship between TCAP-1 and cellular mitochondria.

Figure 6.4. FITC-labelled TCAP-1 co-localizes with mitochondrial marker hydroxysteroid (17-β) dehydrogenase 10 (HSD17β10). Cultured E14 hippocampal cells incubated with FITC- [K8]-TCAP-1 for 30 min showed strong binding to the plasma membrane and punctate-like labeling in the cytosol (B). Mitochondrial marker HSD17β10/ERAB immunoreactivity was mostly observed near the plasma membrane with weak labelling in the cytosol (C). Strong co- localization (yellow) between FITC-[K8]-TCAP-1 and HSD17β10 was observed at the plasma membrane (D and E). The corresponding DIC image is represented on the left (A). Magnification, 630X. Scale bars, 20μm. (Chand and Lovejoy, unpublished data).

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One of the major functions of HSD17β10 is the regulation of sex steroid metabolism

(Yang et al., 2005). In rodents, HSD17β10 is highly expressed in Leydig cells, and preferentially convert 3α-androstanediol into the more potent dihydrotestosterone (DHT) and estradiol to the less potent estrone (He et al., 2000; Ivell et al., 2003). The HSD17β10 enzyme is therefore essential for maintaining appropriate levels of male and female sex hormones. Although TCAP-1 can stimulate testosterone synthesis, the lack of TCAP-1-binding in Leydig cells suggests that

TCAP-1 does not directly interact with HSD17β10 in the testes to regulate steroidogenesis.

However, if HSD17β10 recruitment to or near the plasma membrane by TCAP-1 is part of a functional association with the teneurin-dystroglycan receptor complex, then it is possible that

TCAP-1 could indirectly influence HSD17β10 activity through a functional interaction with the teneurin-dystroglycan complex at the basement membrane, which facilitates cross-talk between cells of the seminiferous tubules and Leydig cells within the interstitium (Siu and Cheng, 2004).

In addition to regulating mitochondrial function and sex steroid metabolism, HSD17β10 has a strong affinity for β-amyloid peptide, and is thus linked to the mitochondrial dysfunction seen in Alzheimer's disease. When cell cultures are exposed to β-amyloid, HSD17β10 inside the cell is rapidly redistributed to the plasma membrane (Yan et al., 1997). Over-expression of β- amyloid and binding to HSD17β10 results in neuronal stress, apoptosis, free-radical generation, and impairment of learning and memory (Yan et al., 1997; Lustbader et al., 2004). The elucidation of peptides that can inhibit HSD17β10-β-amyloid interaction is of considerable interest and has become a therapeutic target in β-amyloid–associated pathologies such as

Alzheimer's disease (Yan et al., 1997; Lustbader et al., 2004; Yao et al., 2011). The strong co- localization between TCAP-1 and HSD17β10 suggests that TCAP-1 may have therapeutic benefits for neurodegenerative diseases such as Alzheimer’s disease. However, it remains to be

210 seen whether the association between TCAP-1 and HSD17β10 at the plasma membrane is indeed a physiological and functional interaction that could inhibit HSD17β10-β-amyloid interaction and the corresponding β-amyloid toxicity.

Interestingly, there has been considerable evidence linking chronic stress to Alzheimer’s disease. Prolonged stress and chronic activation of the CRF1 receptor, hyper-phosphorylates tau proteins and in turn leads to the formation of β-amyloid plaques that are prevalent in Alzheimer’s disease pathology (Rissman et al., 2007; Carroll et al., 2011). Although, TCAP-1 is highly expressed (Wang et al., 2005) and strongly immunoreactive in the pyramidal neurons of the hippocampus, little is known about whether TCAP-1 plays a role in learning, and memory.

However, TCAP-1 is potent at inhibiting CRF-mediated stress-induced changes in hippocampal neuronal morphology and behaviours (Al Chawaf et al., 2007b; Tan et al., 2008; Tan et al., 2009;

Kupferschmidt et al., 2011; Tan et al., 2011) and, thus, could be a useful therapeutic strategy in the pharmacological manipulation of CRF-mediated and stress-associated pathophysiology of

Alzheimer’s disease.

6.2.3 Identifying TCAP-1 binding proteins.

The studies presented in this thesis have elucidated much about the proteins that functionally interact with and are recruited by the TCAP-1 peptide. However, little is known about the proteins that TCAP-1 interacts with. The ability for TCAP-1 to cross the blood-brain barrier (Al Chawaf et al., 2007b) and blood-testes barrier, the punctate-labelling of immunoreactive-TCAP-1 in the cytosol and uptake of FITC-TCAP-1 to reach cytosolic and nuclear targets all point to the existence of TCAP-1 receptors and/or binding proteins (Ng et al.,

2012), that could aid in TCAP-1 transport and protect it from degradation by peptidases.

211

Affinity chromatography and TCAP-1-binding studies have revealed the presence of several TCAP-1-interactors. Resolution of the eluted nuclear material using SDS-PAGE showed the clear presence of a 50 kDa protein with lower intensity bands occurring at about 30, 40 and

100 kDa (Fig. 6.4A). The cytosolic extract showed the presence of 40, 50 and 100 kDa bands as well as a 60 kDa band but not the 30 kDa band (Fig. 6.5A). In addition, synthetic TCAP-1 appeared to associate with a protein of about 50 kDa in mouse F9 testicular carcinoma protein extracts consistent with the dominant form found in the affinity column elutants (Fig. 6.5B). To further confirm the presence of TCAP-binding proteins, fixed E14 hippocampal cells treated with triton-X100 to expose potential TCAP-1 binding sites showed strong web-like FITC-TCAP-

1 labelling in the cytosol (Fig. 6.6) and the existence of TCAP-1 binding sites in the nucleus

(Fig. 6.7). Preliminary mass spectrometry analysis of these TCAP-1-associated interactors has revealed the presence of cytoskeletal-modulating proteins and mitochondrial-associated proteins, which are consistent with known TCAP-1 functions. A positive identification on all the bands is yet to be determined but are similar in size to proteins that TCAP-1 appears to functionally interact with, such as the 43 kDa β-dystroglycan (Winder, 2001) and the 108 kDa HSD17β10 homotetramer (He et al., 2001).

Of particular interest is the identity of the 50 kDa protein, which has tentatively been identified as elongation factor-1α (EF-1α). EF-1α is highly expressed in the pyramidal cell bodies and dendrites of the hippocampus (Huang et al., 2005), an area that shows strong TCAP-

1-immunreactivity and TCAP-1-binding. In the hippocampus, EF-1α facilitates the integration and regulation of both actin- and tubulin-based cytoskeletal elements (Moore et al., 1998; Liu et al., 2002; Huang et al., 2005), and plays a crucial role in stabilizing dendritic spines. The formation, elimination, and stability of dendritic spines in the hippocampus are highly dependent

212 on the actin cytoskeleton and the regulation of microtubules, where new spines anchor their actin filaments (Hotulainen and Hoogenraad, 2010). This interaction between TCAP-1 and EF-1α is, therefore, consistent with TCAP-1 regulating both actin- and tubulin-based cytoskeletal elements and modulating dendritic spine density (Al Chawaf et al., 2007a; Tan et al., 2011).

Figure 6.5. Characterizing TCAP-1 binding proteins. A. Coomassie blue-stained PAGE resulting from elutants obtained from cytosolic and nuclear extracts of mouse N38 cells passed through a TCAP-1 affinity column. B. Binding of synthetic TCAP-1 to a 50kDA TCAP-1-related protein in mouse testicular carcinoma F9 extracts. (Chand and Lovejoy, unpublished data).

Moreover, EF-lα also acts to promote cell growth, confer resistance to stress-induced apoptosis and maintain neuronal morphology by stabilizing microtubules through a calcium/calmodulin-dependent manner and binding directly to actin to create actin-EF-1α bundles that are resistant to depolymerization (Murray et al., 1996; Moore et al., 1998; Talapatra et al., 2002). TCAP-1 also promotes neurite outgrowth, inhibits stress-induced cell death and maintains neuronal morphology during stress (Al Chawaf et al., 2007a, Trubiani et al., 2007; Ng,

2010). In addition, knockdown of all three TCAPs present in N38 cells using siRNA results in a

213 decrease in neurite number and abnormal neuronal morphology (Trubiani, 2008) suggesting that the function of TCAP-1 in regulating cytoskeletal dynamics and promoting active mechanisms that maintain cell morphology and integrity may be intimately linked to EF-1α.

Figure 6.6. FITC-labeled TCAP-1 binding in fixed mouse E14 hippocampal cells. FITC- TCAP-1 binding in cultured E14 hippocampal cells, fixed in 4% PFA and permeabilized with 0.3% Triton X-100 showed strong binding and web-like cytoplasmic labelling (B). The corresponding DIC image is represented on the left (A). Magnification, 630X. Scale bars, 20μm. (Chand and Lovejoy, unpublished data).

Current studies show that TCAP-1 does not significantly affect the synthesis of EF-1α in hippocampal cell culture, but rather regulates the distribution within the cytosol, specifically, to areas that correlate with greater actin polymerization (DeLannoy, Chand and Lovejoy, unpublished findings). The distribution of EF-1α directly correlates with the distribution of β- actin mRNA and is due primarily to the strong binding of β-actin mRNA to EF-1α-F-actin bundles (Liu et al., 2002). It has been previously established that TCAP-1 increases β-actin mRNA in neuronal cell culture (Al Chawaf et al., 2007a). Thus, given that the conventional role of EF-1α is protein synthesis, it is conceivable that both EF-1α and β-actin mRNA are localized to sites where translation would supply β-actin protein near sites of actin polymerization,

214 consistent with the observations of TCAP-1-induced actin polymerization and re-distribution of

EF-1α to areas of actin polymerization.

Figure 6.7. TCAP-1 binding sites in the nucleus. Distinct TCAP-1 binding sites were observed within the nucleus of E14 hippocampal cells incubated with FITC-[K8]-TCAP-1 (B) and also with TCAP-1 TNR308 antiserum (C). Strong co-localization (yellow) with affinity purified TNR308 confirmed the strong uptake to and binding of FITC-[K8]-TCAP-1 in the nucleus (D). Corresponding DIC image is represented on the left (A). Magnification, 630X. Scale bars, 10μm. (Chand and Lovejoy, unpublished data).

6.3 TCAP-1 signal transduction: A novel mechanism in the regulation of cytoskeletal

dynamics and cell survival.

The cytoskeleton is extremely sensitive to a variety of stimuli, serves an important structural and architectural scaffold in the cell, is extremely versatile and is critical in the adaption response to stress and homeostatic imbalance. I have shown that the TCAP-1-mediated regulation of cytoskeletal elements is preceded by the activation and recruitment of a dystroglycan-associated, MEK-ERK1/2-signal transduction pathway that targets cytoskeletal regulatory proteins, stathmin and filamin A. This offers a potential mechanism by which TCAP-

1 mediates neuronal plasticity (Al Chawaf et al., 2007a; Tan et al., 2011) and induces long-term adaptive behavioural responses to CRF-mediated stress, anxiety and addiction (Wang et al.,

2005; Tan et al., 2009; Kupferschmidt et al., 2011). However, TCAP-1 is also neuroprotective

215 under alkalotic and oxidative stress conditions (Trubiani et al 2007; Ng, 2010) and can mediate cellular metabolism and energy levels (Xu and Lovejoy, unpublished findings). The molecular mechanisms by which these effects occur have only partially been described, and points to the recruitment of other proteins in the TCAP-1-induced signal transduction cascade.

One such protein that I have already identified is the microtubule-destabilizing protein stathmin, that may play a central role in the TCAP-1 response to stress. Phosphorylation of stathmin inhibits its destabilizing activity and promotes the polymerization of microtubules.

Microtubules are the stiffest of the three cytoskeletal polymers and serve as the main structural components within cells (Fletcher and Mullins, 2010). In neuronal cells, the phosphorylation of stathmin at serine-25 and serine-38 by ERK1/2 and c-Jun N-terminal kinase (JNK) confers neuroprotection against a range of cellular stressors (Ng et al., 2010). However, at 100nM,

TCAP-1-treatment results in the ERK1/2-mediated phosphorylation of stathmin at serine-25 and causes no change in phosphorylation at serine-38. Interestingly, TCAP-1 (100nM) also induces a corresponding dephosphorylation of stathmin at serine-16 and serine-63 (Fig. 6.8), thus, enhancing its tubulin destabilizing activity. Therefore, instead of promoting the building of a strong tubulin network and cellular architecture to withstand stressful stimuli, TCAP-1 could be acting to decouple the tubulin network and facilitate cellular migration away from the stressor.

This is consistent with my findings of tubulin re-organization towards the cell periphery, the formation of actin-rich filopodia and the internalization of teneurin-1 (Fig. 6.3) and β- dystroglycan (data not shown) that would result in a disassociation from the extracellular matrix.

This response may be indicative of an extreme stress-event, such as nearby cell death due to trauma or injury that would result in release of the cytosolic contents and, thus, a large amount of the TCAP-1 peptide. TCAP-1 has been shown to have a dose-dependent effect (Wang et al.,

216

2005) and cells treated with only 1nM of TCAP-1 show less internalization of teneurin-1 (Fig.

6.3C,D) and fewer filopodia.

Figure 6.8. TCAP-1 dephosphorylates stathmin at serine-16 and -63 and induces phosphorylation BAD in E14 hippocampal cells. Cultured E14 hippocampal cells treated with water (untreated) or 100nM TCAP-1 for 0, 1, 5, 10, 30 and 60 min were analyzed by immunoblotting. A. Immunoblotting analysis with site specific phosphoserine stathmin (STMN) antibodies showed a dephosphorylation of STMN at serine-16 and serine-63 after 5 min of TCAP-1 treatment. B. TCAP-1 induced phosphorylation of p90RSK at serine-380 and BAD at serine-112 in E14 hippocampal cells after 5 mins of TCAP-1 treatment. (Chand and Lovejoy, unpublished data).

The TCAP-1 signalling pathway that I elucidated extends beyond the targeting of and recruitment of cytoskeletal regulating proteins, and suggests a mechanism that could mediate the pro-survival effects attributed to TCAP-1 (Fig. 6.9). Mouse E14 hippocampal cells treated with

100nM TCAP-1 showed a strong induction of phosphorylation of B-cell lymphoma 2 (Bcl-2)- associated death promoter (BAD) at serine-112 (Fig. 6.8B) similar to that detected for TCAP-1- induced ERK1/2 and p90RSK activation. Studies by Bonni et al., (1999) showed that the

MAPK-signaling pathway can mediate growth factor-dependent cell survival through the recruitment of p90RSK and subsequent phosphorylation of the pro-apoptotic protein BAD at serine-112. The phosphorylation of BAD at serine-112 promotes binding of BAD to 14-3-3 proteins and prevents it from inhibiting pro-survival factors, Bcl-2 and Bcl-xl (Zha et al., 1996).

However, it is yet to be determined whether the neuroprotective effects of TCAP-1 are, indeed, mediated by the recruitment of the ERK1/2-dependent p90RSK-BAD system (Fig. 6.9).

217

Figure 6.9. Proposed TCAP-1 signal transduction. TCAP-1 released from neighbouring cells or cleaved from teneurin-1 first interacts with the β–dystroglycan (β-DG) subunit of the dystroglycan complex and is internalized by caveolae-mediated endocytosis. TCAP-1 binding to β-DG activates the mitogen-activated protein (MAP) kinase kinase 1/2 (MEK1/2) and its downstream kinase extracellular signal-regulated kinase 1/2 (ERK1/2). Activation of ERK1/2 leads to the phosphorylation of stathmin at serine-25 and p90-Ribosomal S6 Kinase (p90-RSK) at serine-380. Phosphorylation of p90RSK induces phosphorylation of filamin A at serine-2152, and BAD at serine 133. TCAP-1 does not activate focal adhesion kinase (FAK), suggesting that signaling through the MEK-ERK pathway is integrin-independent.

218

6.3.1 A role for TCAP-1-mediated cytoskeletal regulation during cell division.

The effect of TCAP-1 on cell proliferation (Al Chawaf et al., 2007), tubulin dynamics and modulation of stathmin phosphorylation raised the question as to whether TCAP-1 may also play a role in cell division. Mouse E14 hippocampal cells during anaphase show strong co- localization between α-tubulin and endogenous TCAP-1, particularly near kinetochore microtubules and aster formations. Co-localization between TCAP-1 and α-tubulin was also detected during telophase (Fig. 6.10). However, TCAP-1 does not co-localize with the α-tubulin during interphase.

The dynamics of microtubule polymerization/depolymerization during the different phases of the cell cycle are regulated by a balance between microtubule-stabilizing and microtubule-destabilizing factors. During interphase, microtubules are stable primarily because of a low frequency of catastrophes. However, during mitosis, microtubules become short and dynamic because of a roughly ten-fold increase in the catastrophe frequency compared with that in interphase (Andersen, 2000). Stathmin plays an important role in the regulation of microtubule dynamics during cell cycle progression (Iancu et al., 2001). When cells enter the mitotic phase of the cell cycle, stathmin is phosphorylated at all four regulatory sites, thus, allowing tubulin to polymerize and form a mitotic spindle (Larsson et al., 1997). However, when unphosphorylated, stathmin promotes the depolymerization of the microtubules that make up the mitotic spindle

(Marklund et al., 1996; Iancu et al., 2001). TCAP-1 does not hyper-phosphorylate stathmin, but rather causes a corresponding dephosphorylation of stathmin at serine-16 and -63 (Fig. 6.9A), two regulatory residues that must be phosphorylated for cell division to proceed (Larsson et al.

1997). Although TCAP-1 does affect cell proliferation, it does not appear to do so by inducing

219 mitosis. The TCAP-1-mediated dephosphorylation of stathmin may be necessary for the disassembly of the mitotic spindle, exit from mitosis and entry into a new cell cycle.

Figure 6.10. TCAP-1 co-localizes with tubulin during cell division in mouse E14 hippocampal cells. Immunofluorescent co-labeling with TCAP-1 antiserum (green) and α- tubulin (red) in E14 hippocampal cells showed strong co-localization of TCAP-1 and α-tubulin (yellow) during anaphase near aster formations and kinetochore microtubules. Weak co- localization was observed during telophase at or near kinetochore microtubules. All sections were counterstained with DAPI to highlight cellular DNA. Magnification, 630X. Scale bars, 20μm. (Chand and Lovejoy, unpublished data).

220

The role of TCAP-1 on mediating cytoskeletal dynamics has focussed mainly on action of exogenous TCAP-1. Although little is known about the role of endogenous TCAP-1, it is likely to possess similar functions in regulating cytoskeletal elements, as observed in TCAP-1 knockdown studies (Trubiani, 2008) and strong co-localization with α-tubulin near kinetochore microtubules and aster formations. Interestingly, EF-1α, the putative 50 kDa TCAP-1-interacting protein, is also part of the mitotic apparatus and has been shown to promote aster formation and mitotic spindles in vitro (Toriyama et al., 1988; Moore et al., 1998). The relationship between

TCAP-1 and EF-1α during mitosis remains unresolved. However, current studies suggest that

TCAP-1 regulates the distribution of EF-1α in mouse hippocampal cells during interphase

(DeLannoy, Chand and Lovejoy, unpublished observations). Endogenous TCAP-1 may be functionally interacting with EF-1α or EF-1α-related proteins to help regulate tubulin dynamics at the microtubule organisation complex (MTOC) during cell division. The elucidation of TCAP-

1-EF-1α relationship could therefore serve as a strong platform for future studies on elucidating the role of endogenous TCAP-1 in the cell.

6.4 Regulating CRF-mediated and stress-induced physiology and behaviours.

The TCAP family of peptides was first discovered in a search for novel corticotropin- releasing factor (CRF) homologues (Qian et al., 2004). Since then, the action of the TCAP peptides on CRF-mediated systems has been of considerable interest. One member of the family,

TCAP-1, has emerged as a novel candidate in the modulation of a number of CRF-associated disorders including stress, anxiety and addiction (Wang et al., 2005; Al Chawaf et al., 2007b;

Tan et al., 2009; Rotzinger et al., 2010; Kupferschmidt et al., 2011; Tan et al., 2011). In addition,

I have shown that TCAP-1 is mainly localized to areas that are associated with the regulation of

221 the behavioural stress response, particularly the pyramidal layer of the hippocampus and in neurons of the basolateral nucleus of the amygdala (BLA). The hippocampus and amygdala are important substrates for TCAP-1 action (Tan et al., 2009). However, the mechanism by which

TCAP-1 regulates CRF-mediated behaviours remains unresolved. TCAP-1 does not bind to the

CRF receptors (Nock, 2009; Wang et al., 2005), nor does it affect hypothalamic-pituitary-adrenal axis (HPA) activation (Al Chawaf et al., 2007b). One possibility is that TCAP-1, acting on a receptor system distinct from the CRF receptors, could modulate either signal transduction or transcriptional elements downstream of CRF receptor activation.

This could occur at several different levels. First, TCAP-1 could promote the internalization and/or desensitization of the CRF receptors via the recruitment of G protein- coupled receptor kinase (GRK) and β-Arrestin mechanisms (Hauger et al., 2009). β-arrestins are important regulators of G-protein coupled receptors, and influence regional regulation of CRF1 and CRF2 receptor signaling to prevent deleterious effects of unrestrained and excessive signal transduction (Hauger et al., 2009). Under this scenario TCAP-1 would reduce the cell’s responsivity to CRF-induced signalling and stress. This is supported by previous studies that show TCAP-1 blocks CRF-induced c-Fos expression in key stress regions such as the hippocampus, amygdala, medial prefrontal cortex, lateral and medial septum, and dorsal raphe nucleus in the adult male rat brain (Tan et al., 2009). Moreover, TCAP-1 completely ablates

CRF-mediated stress-induced behaviours (Al Chawaf et al., 2007b), anxiety (Tan et al., 2008) and cocaine-reinstatement (Kupferschmidt et al., 2011) in rats.

Alternatively, cross-talk between the TCAP-1 and CRF-signalling pathways could lead to the inhibition of CRF-mediated stress-induced physiology and behaviours. Chronic stress- induced CRF-mediated action in the hippocampus involves destabilization of spine F-actin and

222 destruction of thin dendritic spines (Chen et al., 2008; Chen et al., 2012). Dendritic spine dynamics and remodeling of the F-actin that provides the bulk of the cytoskeleton within the spine, are regulated by a family of Rho GTPases, including RhoA, Rac1, and cdc42 (Hall, 1998;

Ethell and Pasquale, 2005). However, prolonged and intense stimulation of RhoA, by chronic application of CRF, such as in severe stress, leads to the rapid dephosphorylation (activation) of the F-actin-destabilizing protein, cofilin and subsequent spine disintegration and impaired neuronal plasticity in adult hippocampus (Chen et al., 2008; Chen et al., 2012). TCAP-1 on the other hand activates a dystroglycan-associated MEK-ERK1/2 signalling cascade that targets cytoskeletal regulatory proteins that promotes actin polymerization, cytoskeletal stabilization

(Fig. 6.9) and increased dendritic branching and spine density in the hippocampus (Al Chawaf et al., 2007a; Tan et al., 2011). During times of stress, TCAP-1 administration blocks the effect of

CRF in the hippocampus and alters anxiety-like behaviour, in part, via modification of dendritic spines (Tan et al., 2011).

TCAP-1 may, therefore, be acting antagonistically to CRF by suppressing RhoA activity.

One way, is the suppression of RhoA activation by protein kinase A (PKA) (Qiao et al., 2003).

Although the dystroglycans have no intrinsic ability to activate PKA, α-dystroglycan is intimately linked to the integrins which act as a scaffold for PKA-signalling (Lim et al., 2008). In addition, the C-terminal region of teneurin-1 is also functionally linked to α-dystroglycan and integrin at the plasma membrane (Trzebiatowska et al., 2008). TCAP-1 binding to the teneurin-

1-dystroglycan complex could indirectly induce an integrin-mediated protein kinase A activation at the leading edge of cells (Lim et al., 2008) to inhibit CRF-driven RhoA activation and thus promote dendritic branching and neuronal connectivity (Chen et al., 2012). This is further supported from studies by Noren et al., (2001; 2003) who showed that the induction of cell-cell

223 contacts and cadherin-associated cell adhesion activates Rac1 and Cdc42 and strongly inhibits

RhoA activation. Teneurin-1 and dystroglycan are both known to promote cell adhesion, and cadherin-like domains have been identified in α-dystroglycan (Dickens et al., 2002). This suggests that a functional relationship between the teneurin-TCAP-1-dystroglycan complex and the cadherins could exist at the plasma membrane and in turn form an integrative system that could inhibit CRF-driven RhoA activity and prevent spine disintegration.

TCAP-1 could also act antagonistically to CRF at the level of cytoskeletal organization as means of preparing cells for an anticipated stress event. TCAP-1 stabilizes the cytoskeleton by inducing actin polymerization and cross-linking, re-organizing microtubulin from a perinuclear- cytosolic scaffold to the periphery of the cell, recruits cytoskeletal stabilizing proteins and maintains cellular integrity in stress conditions (Fig. 6.9; Tan et al., 2012). This indicates that

TCAP-1 could protect dendritic spines from CRF-induced destruction by promoting active cytoskeletal mechanisms that maintain and strengthen cellular morphology and network integrity. This scenario would require the release and action of TCAP-1 prior to the anticipated stress.

TCAP-1 lacks a signal sequence for secretion and is likely to leave the cell either through cell injury/death, cell leakage (D’Amore, 1990), the formation of labile structures such as exosomes in a process known as membrane blebbing (Nickel, 2003), or a novel and distinct secretion pathway. Regardless, the release of small amounts of TCAP-1 and action on neighbouring neurons could act as a warning mechanism for an incoming stressor. This would then put the neurons in a “prepared state” by upregulating cytoskeletal elements for maintaining and strengthening cellular integrity, and inducing filopodia formation and elongation for enhanced sensitivity to environmental cues. Although a small amount of TCAP-1 release in the

224 brain may signal mild brain distress or acute injury, the effects of repeated TCAP-1 would communicate a message of distress and as such would warrant a more severe response, such as detachment from the extracellular matrix as observed by internalization of teneurin-1 (Fig. 6.3) and β-dystroglycan and/or the promotion of cellular migration as observed by the increase in

TCAP-1-induced lamellipodia and filopodia formation (Fig. 6.9). This type of signalling coupled with the ability to anticipate danger and recognize hostile cues would represent one of the simplest, oldest and most beneficial forms of communication between cells that would have existed before the evolution of multicellular organisms.

A similar effect has been observed in vivo. Behavioural studies show TCAP-1 is mildly anxiolytic in ‘unstressed’ rats as measured on the elevated plus maze (EPM) and open field (OF) tests (Tan et al., 2008; Tan et al., 2011). However, in the presence of a stressor, repeated administration of TCAP-1 results in an anxiogenic response in the EPM and OF tests (Tan et al.,

2008; Tan et al., 2009). This phenomena has been postulated by Tan et al., (2012) to be a coping mechanism that makes the organism more aware of injurious cues, whereby TCAP-1 promotes different coping mechanisms by preparing the brain for stressors by increasing the exploratory drive in a novel environment (active coping) and increasing protective and defensive behaviours

(passive coping) in a harmful environment.

Many neuropsychiatric conditions have a common set of neurological substrates associated with the integration of sensorimotor processing. Common to these conditions is the aberrant processing of sensorimotor integration (Bolmont et al., 2002; Jovanovic et al., 2009;

Canbeyli, 2010; Epstein et al., 2011). The teneurin-TCAP family of proteins has recently emerged as a novel candidate system to integrate the origin of these disorders (Rotzinger et al.,

2009). The ten-m gene appears to be upregulated by stressors (Wang et al., 1998) and TCAP-1 is

225 efficacious at regulating CRF-mediated and stress-induced physiology and behaviours (Lovejoy et al., 2006; Rotzinger et al., 2010; Tan et al., 2012). However, the exact mechanisms linking the teneurin-TCAP-1 and CRF systems require further investigation. The studies presented in this thesis have shown that TCAP-1 is uniquely positioned to illicit a number a functional responses and offers novel insight into how TCAP-1 could regulate growth, morphology and stress. At the heart of the TCAP-1-mediated mechanism is the association with extracellular matrix proteins and the regulation of cytoskeletal dynamics, that could represent a novel neurological system underlying the origins of a number of complex neuropsychiatric conditions. TCAP-1 could, therefore, be part of a promising therapeutic strategy that targets the cytoskeleton to provide preventative and therapeutic benefit for the treatment of a wide range of stress-related disorders

(Table 6.1).

Table 6.1: Comparing the molecular characteristics of CRF-associated pathologies and TCAP-1 effects in the central nervous system (CNS)

Stress-related abnormalities TCAP-1-treatment in the CNS in the CNS 1 Spine loss and dendritic abnormality Increased spine density and dendritic (Chen et al., 2012) morphology (Tan et al., 2011) 2 Disruption of F-actin and tubulin Increased actin-polymerization and cross- dynamics (Chen et al., 2008) linking. Regulation of tubulin-organization. 3 Excess BDNF and intracellular Decreased BDNF expression (Ng, 2010; BDNF trafficking (Ma et al., 1999) Ng et al., 2012) 4 Impaired Filamin A Function (Sheen Induces Filamin A activity et al., 2001) 5 Hyper-phosphorylation of stathmin Dephosphorylation of stathmin at serine-16 and stabilization of microtubules and serine-63, no change at serine-38 and (Nakamura et al., 2006) phosphorylation at serine-25

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6.5 A broader role for TCAP-1 in reproduction.

The studies presented in this thesis have expanded our knowledge of the teneurin-TCAP-

1 system beyond the CNS. I have presented the first detailed study of the localization of teneurin-1 and TCAP-1 in the adult mammalian testis, identified the dystroglycans a possible receptor-candidate for the teneurin-TCAP-1 system, clarified the molecular organization of the dystroglycan complex at the basement membrane of the testis and epididymis, and showed that

TCAP-1 is potent regulator of testis and epididymis size. Together, these studies provide a strong framework for future investigations regarding TCAP-1 as a novel regulator of mammalian reproduction.

6.5.1 Furthering the case for TCAP-1 in male reproduction.

The studies characterizing the localization and action of TCAP-1 in the adult male testis and epididymis highlights a broader role for the teneurin-TCAP system in the gonads. TCAP-1- treated mice showed increased testicular size, epididymal size and testosterone levels, suggesting that TCAP-1 may be part of the family of trophic factors expressed in the mammalian testes that can regulate testicular function. However, the precise function of TCAP in the gonads is not yet fully characterized. Like many other growth factors and cytokines that are expressed in the mammalian testes, TCAP-1 may also possess a multitude of roles in the testes. This is supported by several pieces of evidence.

First, TCAP-1-immunoreactivity and TCAP-1-binding sites in spermatogonia suggest a broader role in mediating the continuous energy-demanding and highly metabolic process of spermatogenesis. Metabolic stress and energy unbalance are frequently coupled to disturbed reproductive maturation and infertility (Herrera et al., 2000). Much of the current research on

227 how energy availability and metabolism influences the development and function of the reproductive axis has focussed mainly at the level of the hypothalamic centers governing reproduction. Metabolic cues and a number of neuropeptide signalling molecules, such as the kisspeptin family, have been identified as key players in the metabolic regulation of fertility

(Meczekalski et al., 2011). However, the highly active seminiferous tubules of the testes also possess a system deeply involved in the metabolic regulation of spermatogenesis (Boussouar and

Benahmed, 2004). This energy-dependent growth and differentiation of the germ cells in the seminiferous tubule is tightly controlled by Sertoli cells (Jegou, 1993). Meiotic and post-meiotic spermatogenic cells rely on oxidative ATP production for their energy metabolism (Reyes et al.,

1990). Follicle stimulating hormone (FSH) induces glucose uptake and its conversion to lactate in Sertoli cells. Lactate is then transported to the meiotic and post-meiotic spermatogenic cells, providing them with the primary metabolic fuel necessary for oxidative metabolism (Hall and

Mita, 1984). The mobilization of glucose and lactate in spermatogonia and other germs cells seminiferous tubule is dependent on hexose transporters (Angulo et al., 1998) and the glucose family of transporters (GLUT) (Boussouar and Benahmed, 2004). However, little is known about the signalling molecules regulating this system within the testicular germ cells. Current research shows that TCAP-1 modulates glucose metabolism and increases cellular ATP-levels in neuronal cell culture (Xu and Lovejoy, personal communication) and may, therefore, possess a similar a role in the testis. Given that TCAP-1 associates with the dystroglycan complex in the seminiferous tubule, it could modulate cytoskeletal elements similar to that in the hippocampus as a prelude to the mobilization of hexose and GLUT transporters, ultimately regulating ATP- levels within the germ cells.

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Secondly, TCAP-1 may confer cellular protection in the characteristically oxygen-poor environment of the seminiferous tubule similar to that observed in neuronal cell culture

(Trubinani et al., 2007; Ng, 2010). The high metabolic activity within the testis predisposes it to metabolic toxicity that could compromise the integrity of germ cell DNA. During spermatogenesis, reactive oxygen species (ROS) are generated as a by-product of normal cellular metabolism. ROS expression at appropriate levels has physiological roles in cellular differentiation (Sohal et al., 1986) and sperm capacitation (de Lamirande et al., 1997). However, oxygen-free radicals at concentrations beyond physiological limits result in oxidative stress, which can damage DNA, proteins, and lipids (Halliwell, 1996), ultimately leading to apoptosis or necrosis. Although germ cells are susceptible detrimentally to exogenous and endogenous reactive oxygen species (ROS), spermatogonia which show strong TCAP-1-immunoreactivity and TCAP-1-binding are highly tolerant to ROS attack, whereas advanced-stage germ cells, such as spermatozoa which lack TCAP-1 and TCAP-1-binding sites, are much more susceptible. This is due, in part, to the presence of high levels of copper/zinc (Cu/Zn) superoxide dismutase (SOD) that render spermatogonia resistant to ROS, and consequently protected from oxidative stress thus ensuring the fidelity and integrity of germ cell DNA (Celino et al. 2011). It has been previously shown, in vitro, that TCAP-1 inhibits necrotic cell death and is protective against oxygen free radical stress via an increase of SOD expression, SOD Cu2+ chaperone expression and catalase expression and activity (Trubiani et al., 2007). Spermatogonia have evolved with a greater need for elevated levels of protective factors against DNA damage and TCAP-1 may, therefore, serve a broader role as part of the family of trophic factors that maintain homeostasis within the seminiferous tubule during spermatogenesis.

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Thirdly, apart from spermatogenesis, TCAP-1 also appears to play a role in steroidogenesis. However, the lack of TCAP-1-immunoreactivity and TCAP-1-binding sites in the Leydig cells suggests that TCAP-1 is unlikely to have direct steroidogenic effects on Leydig cells. Moreover, TCAP-1 lacks a signal sequence for secretion and is, therefore, unlikely to be secreted from the seminiferous tubules. Alternatively, TCAP-1 may act to increase testosterone production through a functional interaction with the extracellular matrix at the basement membrane, similar to many other testosterone stimulating cytokines and growth factors expressed in the seminiferous tubule such as tumor necrosis factor-α (TNFα; Siu et al., 2003) and insulin-like growth factor 1 (IGF-1; Saez et al., 1989). Communication between the cells in the seminiferous tubule and the interstitium is mediated not only by testosterone from Leydig cells, but also by trophic factors secreted from the seminiferous tubule (Saez et al., 1989) and direct cell-cell contact via extracellular matrix proteins anchored at the basement membrane (Siu and

Cheng, 2004) that increases the steroidogenic responsiveness and activity of the steroidogenic pathway in Leydig cells (Reventos et al., 1989). My study shows strong teneurin-1 and α- dystroglycan co-localization in the interstitium and at the basement membrane of the seminiferous tubule. Teneurins are known to interact in a homophilic manner (Oohashi et al.,

1999; Rubin et al., 2002; Bagutti et al., 2003; Leamey et al., 2008) and to date, no ligand has been identified. These studies suggest that teneurin-1-associated communication between the seminiferous tubule and interstitium could occur either by homophilic interaction or by juxtacrine-signaling with α-dystroglycan in a similar manner to the notch signaling of serrate and delta ligands (Artavanis-Tsakonas et al., 1999; Hicks et al., 2002). Given that TCAP-1 co- localizes with β-dystroglycan, TCAP-1 could potentially regulate Leydig cell testosterone production via a functional interaction with the dystroglycan complex at the basement membrane

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The presence of TCAP-1 binding sites in the seminiferous tubule, uptake of iodinated

TCAP-1 in the rodent testis (Tan, 2011) and trophic effects on the testes and epididymis suggests that TCAP-1 can cross the tightly-regulated blood-testes-barrier (Mruk and Cheng, 2010; Su et al., 2011). It has already been demonstrated that TCAP-1 can cross the blood-brain barrier (Al

Chawaf et al., 2007), however, the exact transport and uptake mechanism remains to be elucidated. The ability for TCAP-1 to traverse these highly-regulated and restrictive passages further highlights the importance and fundamental role that this highly conserved peptide plays in mediating physiological function.

6.5.2 A role for TCAP-1 in female reproduction.

Gametogenesis in both males and females share similar constraints and challenges pertaining to morphology, migration, differentiation and metabolism. The strong localization and biological functions of TCAP-1 within the male reproductive system raises the question of whether TCAP-1 may also play a role in female reproduction. In the female reproductive tract, immunoreactive-TCAP-1 was detected in the ovarian follicle, corpus luteum and fallopian tube

(Fig. 6.11A). TCAP-1-immunoreactivity in the female reproductive tissues was restricted to the cytosol (Fig. 6.11) similar to that observed in the brain and testes. TCAP-1-immunoreactivity was strongly localized to the cytosol of the granulosa cells and oocyte of the ovarian follicle

(Fig. 6.11A, B). Immunoreactive-TCAP-1 was not detected in the contractile cells of the theca externa or steroidogenic theca interna of the ovarian follicle. In the corpus luteum, immunoreactive-TCAP-1 was also localized to the granulosa cells but was comparatively weaker as compared to the TCAP-1-stained granulosa cells of the ovarian follicle (Fig. 6.11A). In addition, strong TCAP-1-immunoreactivity was also detected at the ovarian surface epithelium

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(Fig. 6.11F). In the fallopian tube, TCAP-1-immunoreactivity was restricted to the columnar epithelial cells (Fig. 6.11D).

Figure 6.11. TCAP-1-immunoreactivity in adult mouse female reproductive tract. TCAP-1- immunoreactivity was detected in cytosol of the granulosa cells and oocyte of the ovarian follicle (A, B), the granulosa cells of the corpus luteum (cp) (A), columnar epithelial cells of the fallopian tube (A, D) and the ovarian surface epithelium (F). Sections preadsorbed with synthetic TCAP-1 completely ablated TCAP-1 staining (C,E). (Chand, Kollara, Brown and Lovejoy, manuscript in preparation).

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TCAP-1-binding sites were also detected in the same regions of the female reproductive tract that were immunoreactive for TCAP-1. In the ovarian follicle, TCAP-1 binding sites were localized specifically to cytosolic regions of the granulosa cells (Fig. 6.12A-C). TCAP-1-binding appeared stronger in the cumulus granulosa cells that surround the oocyte. Unlike the strong

TCAP-1-immunreactivity observed in the oocyte, FITC-conjugated TCAP-1 did not label the oocyte. TCAP-1-binding was also observed in granulosa lutein cells of the corpus luteum (Fig.

6.12D-F) and cells within the endometrium of the uterus (Fig. 6.12G-I). In addition, FITC-

TCAP-1 binding was not observed in theca cells of the ovarian follicle or the theca lutein cells of the corpus luteum (Fig. 6.12B,C,E and F), consistent with the lack of TCAP-1-immunreactivity in these areas (Fig. 6.11). In the fallopian tube, FITC-conjugated TCAP-1 exclusively labeled the cytosol and nuclei of the mucosal cells (Fig. 6.12J-L).

TCAP-1 appears to have a paracrine action as FITC-TCAP-1-binding appears in cells adjacent to those that are immunoreactive for TCAP-1 (Fig. 6.11 and Fig. 6.12). Moreover,

TCAP-1 immunostaining in the ovarian follicle suggests cytoplasmic streaming may be occurring between the oocyte and granulosa cells. The granulosa cells surrounding the oocyte, an area that shows strong TCAP-1 binding, are connected with the oocyte by way of gap junctions and cytoplasmic bridges (Gutzeit, 1986). These bridges facilitate communication between the growing oocyte and the granulosa cells that surrounds it and are heavily dependent on extensive modulation of both actin and tubulin-based cytoskeletal elements (Gutzeit, 1986; Bohrmann and

Biber, 1994). This is consistent with previous reports that have shown the oocyte to be an important paracrine regulator of granulosa cell proliferation and differentiation (Vanderhyden et al., 1990; Gilchrist et. al., 2006). TCAP-1 may, therefore, be part of the family of signalling molecules secreted by the oocyte to mediate granulosa cell function.

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Figure 6.12. FITC-[K8]-TCAP-1 binding in the adult mouse female reproductive tissues. Sections incubated with FITC-labeled TCAP-1 showed strong binding in the cytosol of granulosa cells of the ovarian follicle (B and C). FITC-TCAP-1 binding was also observed in the granulosa lutein cells of the corpus luteum (E and F) and cells within the endometrium of the uterus (H and I). In the fallopian tube, FITC-TCAP-1 labeled the cytosol and nuclei of the mucosal cells (K and L). Corresponding DIC images are represented on the left (A, D, G and J). Magnification, 400X (A-I); 630X (J-L). Scale bars, 25μm (A-I); 15μm (J-L). (Chand, Kollara, Brown and Lovejoy, manuscript in preparation).

Similar to the testes, the ovaries are also a site of massive cellular proliferation, differentiation, migration and high metabolic activity (Rábiee et al., 1997). Consequently, the

234 female reproductive tract is also vulnerable to oxidative stress. ROS affect multiple physiological processes from oocyte maturation to fertilization (Agarwal et. al., 2005). In the female reproductive tract, TCAP-1 expression appears to be primarily localized to the proliferative epithelial-type cells and may be involved in energy metabolism, homeostatic and protective mechanism similar to that postulated in the testis and reported in cell culture (Trubiani et al.,

2007; Ng, 2010).

Alternatively, TCAP-1 may also function to modulate the extensive morphological changes in the ovaries (Pask, 2012) and the fallopian tube (Crow et al., 1994) during the female reproductive cycle, similar to that observed in the adult mouse testis and epididymis. Much is known about the role of the hypothalamic-pituitary gonadal (HPG) axis and the hormones produced from each level of the axis in the regulation of the female reproductive cycle (Maffucci and Gore, 2009). However, recent studies have identified a crucial role for extracellular matrix proteins such as the integrins in regulation of ovarian morphology and function (Berkholtz et al.,

2006; Monniaux et al., 2006). In the mammalian ovary, extracellular matrix proteins are involved in the recruitment of multiple signalling and scaffolding proteins, including ERK1/2- mediated modulation of cytoskeletal elements to regulate ovarian morphology, follicle maturation, cellular proliferation, apoptosis, differentiation, implantation and migration (Reszka et al., 1997; Berkholtz et al., 2006; Monniaux et al., 2006). Although it is yet to be determined, it is likely that the teneurin-1 and the dystroglycan complex are also expressed in the female reproductive tract, particularly at the basement membrane and oocyte similar to that reported for the integrins and laminins (Berkholtz et al., 2006; Monniaux et al., 2006) and may also functionally interact with TCAP-1 as observed in the mouse testis.

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6.5.3 Enhancing fertility with TCAP-1.

The role for TCAP in reproduction appears to be evolutionary conserved from tunicates

(Fig. 6.1) to mice. These studies raise the possibility that TCAP-1 could be a useful therapeutic agent for fertility disorders related to impaired spermatogenesis or poor oocyte quality. In the adult mouse, TCAP-1 increased testicular size, epididymal size and testosterone levels. Future studies should be aimed at determining whether these TCAP-1-mediated in vivo effects correlate with an increase in spermatozoa and/or improved sperm quality.

In addition, TCAP-1 could also enhance the current standard of in vitro fertilization

(IVF). Oocyte quality has long been considered the main limiting factor for IVF and is directly linked to ATP levels within the oocyte (Wang et al., 2009). Mitochondria are the primary source of cellular energy and provide ATP to the oocytes to support the rapid cell division for fertilization and preimplantation embryo development (Torner et al., 2004). High quality oocytes are rich in mitochondria and ATP. Mitochondrial dysfunction, therefore, leads to suboptimal

ATP levels and consequently poor quality oocytes. Poor oocyte quality is often characterized by increased mitochondrial DNA damage, chromosomal aneuploidies and apoptosis, all of which leads to a high level of developmental retardation and arrest of preimplantation embryos (Wang et al., 2009). Apart from increasing maternal age, oxidative damage from excessive ROS has also been implicated as the leading causes of mitochondrial dysfunction and poor oocyte quality

(Tarín, 1996; Trifunovic et al., 2005). The ability for TCAP-1 to modulate glucose metabolism and increase cellular ATP-levels (Xu and Lovejoy, personal communication), as well as protect against ROS (Trubiani et al., 2007) makes it a novel therapeutic candidate for improving female fertility and current IVF standards.

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6.6 A role for TCAP-1 in cancer.

Cancer is characterized by disordered and deregulated cellular proliferation, reduced cell death and the ability to survive under stresses such as nutrient and growth factor deprivation, hypoxia, and loss of cell-to-cell contacts. TCAP-1 has significant effects on cellular morphology, survival and proliferation (Lovejoy et al., 2006; Tan et al., 2012) and is mapped to areas of massive proliferation and high metabolic activity in reproductive tissues. In the female reproductive tract, most ovarian cancers appear at the ovarian surface epithelium, whereas in the testis, germ cell tumours are the most common testicular tumours. I have shown that both of these areas, the ovarian surface epithelium in mouse ovaries and the germ line of the mouse testes are highly immunoreactive for TCAP-1. This raises the question as to whether TCAP-1 may also have a functional role in cancer biology.

Given that TCAP-1 is localized mainly to epithelial type cells I hypothesized that TCAP-

1 would also be expressed or even up-regulated in most epithelial-type cancers. I found that

TCAP-1 was highly immunoreactive in cell lysates from mouse testicular F9 carcinoma, ovarian carcinoma OVCAR-3 and SK-OV-3 cells, breast carcinoma, lung carcinoma, lung fibroblasts and glioblastoma (Fig. 6.13). In these cells, TCAP-1-immunoreactivity was mainly detected as a

50 kDa band. In addition, a larger 250 kDa band was also detected in the OVAR-3 and SK-OV-3 cell lysate (Fig. 6.13) and may be a result of ectodomain shedding of the large C-terminal fragment of teneurin-1 (Kenzelmann et al., 2008).These immunoreactive TCAP-1 bands are consistent with those observed in normal tissues and cell lines. Interestingly, HTB and NTERA-2 testicular carcinoma cells and ES-2 and HEY ovarian carcinoma cells showed little to no TCAP-

1-immunoreactivity (Fig. 6.13). In addition, immunofluorescent-labelling with TCAP-1 antiserum on human testis with mixed germ cell tumour (5% Seminona, 85% Embryonal

237 carcinoma and 10% mature teratoma) found intense TCAP-1-immunoreativity specifically localized to the germ cells (Fig. 6.14), similar to that observed in normal mouse testis (Chapter

5).

Figure 6.13. Western blot analysis on cancer cell lines using TCAP-1 antiserum. Strong expression of a 50kDa TCAP-related protein was detected in F9, OVAR-3, SK-OV-3, breast carcinoma, lung carcinoma, glioblastoma and lung fibroblasts cell lines. A 250kDa TCAP- fragment was detected in the OVCAR-3 and SK-OV-3 cell lines. (Chand and Lovejoy, unpublished data).

The role of TCAP-1 in cancer may lie in its ability to regulate cytoskeletal elements and apoptotic machinery. Many cancer cells are particularly hardy because they have switched off their apoptotic machinery, thus protecting them from the suicide process that their aberrant behaviour would otherwise trigger, even in stressful environments. TCAP-1 has been shown to prevent apoptotic cell death by increasing the synthesis of proteins associated with oxidative stress (Trubiani et al., 2007) and pro-apoptotic protein BAD, activates pro-survival factors such as p90RSK (Fig. 6.8; Fig. 6.9) and confers cellular protection under hypoxic conditions (Ng,

2010).

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Moreover, two biological features that characterize cancer, deregulated cell proliferation and invasive behaviour are typical the result of aberrant cytoskeletal regulation. Of particular interest are three TCAP-1-targeted cytoskeletal regulating proteins, stathmin, also known as oncoprotein-18 (Sherbet and Cajone, 2005), EF-1α (Edmonds et al., 1996) and dystroglycan

(Sgambato and Brancaccio, 2005). Stathmin is expressed at high levels in a wide variety of cancers, particularly ovarian cancer (Iancu et al., 2001) and the inhibition of stathmin expression in malignant cells interferes with their orderly progression through the cell cycle (Mistry and

Atweh, 2002). I have shown that TCAP-1 can regulate the phosphorylation of stathmin and consequently the microtubule cytoskeleton. In addition to inducing phosphorylation of stathmin at serine-25 in mouse hippocampal cells, TCAP-1 also causes a corresponding dephosphorylation of stathmin at serine-16 and -63 (Fig. 6.8), two regulatory residues that are strongly implicated in cell division (Larsson et al. 1997) and tumour growth (Sherbet and

Cajone, 2005).

Figure 6.14. TCAP-1 expression in human testes with mixed germ cell tumor. Immunofluorescence labeling with TCAP-1 antiserum (Red) showed distinct TCAP-1 expression in the germ cells of testes. Sertoli and Leydig cells were not immunoreactive for TCAP-1. Magnification, 200X. Scale bars, 50μm. (Chand and Lovejoy, unpublished data).

In metastasis, actin-binding and microtubule-associating protein EF-1α is overexpressed in metastatic cells and whole tumours and is highly localized to cytoskeletal structures thought to

239 be important for supporting the cellular motility required for metastasis (Edmonds et al., 1996).

In non-metastatic cells, EF-lα stabilizes microtubules through a calcium/calmodulin-dependent manner (Moore et al., 1998) and alters the rates of actin polymerization (Murray et al., 1996) by reducing the critical concentration of actin and binding directly to actin to create actin-EF-1α bundles that are resistant to depolymerization (Murray et al., 1996). Current studies show that

TCAP-1 does not significantly affect the synthesis of EF-1α in hippocampal cell culture, but rather regulates the distribution within the cytosol, specifically, to areas that correlate with greater actin polymerization (DeLannoy, Chand and Lovejoy, unpublished findings).

Abnormalities in the expression of the dystroglycan complex have also been implicated in tumour invasion and dissemination (Sgambato and Brancaccio, 2005). Dystroglycan provides a molecular bridge between the extracellular matrix and the intracellular cytoskeleton (Winder,

2001) and cytosolic components involved in signal transduction (Cavaldesi et al., 1999; Russo et al., 2000). Functionally, dystrolgycans are involved in cellular adhesion, epithelial cell development, formation of the basement membrane and maintenance of tissue integrity. This makes the dystroglycan complex a prime candidate for the study of metastatic tumour cells. The behaviour of metastatic tumour cells is intimately linked to interactions between tumour cells and adhesion molecules and extracellular matrix proteins such as the dystroglycans (Kawaguchi,

2005). In certain tumours and cancers, mutations in the dystroglycan complex can lead to the loss of cell-cell contacts, aberrant cell signalling and/or dysregulation of cytoskeletal elements

(Sgambato and Brancaccio, 2005).

I have shown that α-dystroglycan co-localizes with teneurin-1, and previous studies have demonstrated that teneurin-1 and dystroglycan have related and partly redundant functions in the maintenance of basement membrane integrity and cell adhesion (Drabikowski et al., 2005;

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Trzebiatowska et al. 2008), suggesting that the role of dystroglycan in cancer biology may also be closely linked to teneurin-1 function. Furthermore, my studies have uncovered an intimate relationship between TCAP-1 and the actin-binding and cell signaling β-dystrgoycan subunit. To determine whether TCAP-1 plays a role in cancer biology, future studies should be aimed at elucidating the teneurin-TCAP-1-dystroglycan relationship and role in cancer cells. Future studies could also take advantage of the testicular and ovarian carcinomas that show high TCAP-

1-immunoreactivity (F9, OVAR-3 and SK-OV-3) and those that show little to no immunoreactive-TCAP-1 (ES-2 and HEY) to investigate the effect of TCAP-1 on stathmin phosphorylation and cytoskeletal organization in cancer cells.

6.7 Conclusions

The studies presented in this thesis provide novel information on the molecular characterization and localization of TCAP-1 in the mouse and furthers our understanding of the mechanisms associated with TCAP-1-mediated function in the body. TCAP-1 can be both structurally and functionally independent from the much larger teneurin-1 and is localized to areas in the brain that are associated with the integration of sensorimotor processing and stress systems. My studies suggest that TCAP-1 may be a novel ligand for the dystroglycan complex, where it activates a novel and integrative mechanism that regulates cytoskeletal elements and cell survival systems. Furthermore, TCAP-1 can be internalized to reach cytosolic and nuclear targets. The dynamic changes in the cytoskeleton provide new insight into the molecular mechanisms that are likely to underlie a number of TCAP-1-induced effects in the hippocampus such as dendritic spine formation, inhibition of CRF-associated stress behaviours and neuronal plasticity.

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The function of TCAP-1 extends beyond the CNS and appears to play an evolutionary conserved role in reproduction. TCAP-1 is bioactive at mediating testicular size, epididymal size and steroidogenesis in the adult mouse. In the testis and epididymis, my studies further support a functional interaction between the teneurins and the dystroglycans, which may form part of the putative TCAP-1-receptor complex. Although many questions remain, these studies implicate the teneurin-TCAP system in a broader range of neuroendocrine and trophic functions than previously thought. Together, the data presented here has furthered our understanding of the enigmatic nature of TCAP-1 and lays a foundation of work that may lead to the use of TCAP-1 as a potential therapeutic agent in the treatment of a number of pathologies associated with development, metabolism, stress and reproduction.

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