Cellular and Molecular Mechanisms of Transient Receptor Potential Melastatin 7 (TRPM7) Channel in Neuronal Development and Injury

Home , Naltriben

Ekaterina Turlova

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy Department of Physiology University of Toronto

Cellular and Molecular Mechanisms of Transient Receptor Potential Melastatin 7 (TRPM7) Channel in Neuronal Development and Injury

Ekaterina Turlova

Doctor of Philosophy

Department of Physiology University of Toronto

2018 Abstract

Transient Receptor Potential Melastatin 7 (TRPM7) is a ubiquitously expressed, calcium- permeable ion channel that regulates intracellular levels of divalent cations, cytoskeletal dynamics, cell adhesion and other processes. As these processes are critical for neurite elongation during development, this thesis investigated the role of TRPM7 in neurite outgrowth and development using a pharmacological approach. It was found that blocking TRPM7 activity preferentially enhanced axonal outgrowth of cultured hippocampal neurons at least in part through TRPM7 interaction with cytoskeletal proteins, actin and α-actinin. TRPM7 activity is potentiated under the conditions of stress that commonly occur in injury or disease, such as increased extracellular acidity and production of reactive oxygen species. Therefore, the effect of hypoxia, a common cause of damage to the developing brain, on TRPM7 activity and TRPM7- mediated neurite outgrowth was investigated next. It was found that prolonged hypoxia caused axonal retraction, while blocking TRPM7 activity attenuated this effect. Finally, the therapeutic potential of TRPM7 blocker has been evaluated in a mouse model of neonatal hypoxic-ischemic brain injury, and blocking TRPM7 pharmacologically had neuroprotective and therapeutic effects (up to 1 hour after injury). Further short and long-term evaluation demonstrated ii preservation of brain morphology and functional motor and memory outcomes. Proteomic analysis revealed that TRPM7 may mediate neuronal injury at least in part through calcium- dependent cytoskeletal regulation by calcium/calmodulin-dependent kinase and phosphatase.

Overall, this thesis establishes the role of TRPM7 in neuronal outgrowth and development under normal and stress conditions. Moreover, it evaluates a specific pharmacological inhibitor of

TRPM7 as a candidate for further drug development for hypoxic-ischemic injury to the developing brain.

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Acknowledgements

First, I would like to thank Dr. Hong-Shuo Sun for his supervision and numerous opportunities that allowed me to develop professionally over the years. He showed me that hard work and perseverance always pay off and provided a great example of what a dedicated scientist should be. I truly could not have asked for a better mentor, and I am very grateful for his guidance. I would also like to thank Dr. Zhong-Ping Feng for her care, support and advice. Together, they created an intellectually and scientifically stimulating environment that helped me develop into a well-rounded individual and provided me with opportunities for learning and conducting high- quality research. I would also like to thank my committee members, Dr. Cindi Morshead, Dr. Philippe Monnier and Dr. Jeffrey Henderson, for giving me advice and guidance and for challenging me to think critically and creatively about my work. I am also thankful for the help and technical support from our collaborators – Dr. Andrea Fleig and Dr. F. David Horgen.

I would like to extend special thanks to the members and alumni of Sun and Feng labs – Dr. Andrew Barszczyk, Marielle Deurloo, Raymond Wong, Vivian Szeto, Sammen Huang, Feiya Li, Dr. Wenliang Chen, Dr. Baofeng Xu, Qing Li, Ji-Sun Kim and many others – for their help, support and friendship throughout my program.

Finally, I would not be where I am today without the care, love and support of my family and my best friend and partner, Steven. Thank you for believing in me and always encouraging me to do and be my best.

Table of Contents

Acknowledgements ...... iv

Table of Contents ...... v

List of Tables ...... xiv

List of Figures ...... xv

List of Abbreviations ...... xix

Chapter 1 Introduction ...... 1

1 Mechanisms of neurite outgrowth ...... 2

1.1 Current model of neurite outgrowth ...... 2

1.2 The cytoskeletal structure of neuronal growth cone ...... 3

1.3 Cellular events in neuronal polarization ...... 5

1.3.1 Directed membrane and protein trafficking and cytoplasmic flow ...... 5

1.3.2 Cytoskeletal dynamics ...... 6

1.3.3 Extracellular and substrate cues ...... 7

1.3.4 Current model of axon specification in neuronal polarization...... 8

1.4 Cellular mechanisms of neurite outgrowth ...... 8

1.4.1 The role of Ca2+ in neurite outgrowth ...... 8

1.4.2 Ca2+-conducting membrane channels in neurite outgrowth ...... 9

1.4.3 Ca2+-dependent modulators of neurite outgrowth ...... 10

1.5 The effects of hypoxia on the developing brain ...... 15

1.5.1 In vitro studies of effects of hypoxia on neurite outgrowth ...... 15

1.5.2 The effects of hypoxia on neuronal cytoskeleton ...... 15

1.6 In vivo models of hypoxic-ischemic injury to the developing brain ...... 16

1.6.1 Hypoxic-ischemic encephalopathy ...... 16

1.6.2 Current treatments and interventions ...... 17

1.6.3 Current animal models of neonatal HIE ...... 17

1.6.4 Selective regional and cellular vulnerability ...... 19

1.6.5 Evidence of axonal damage in rodents and humans ...... 20

1.7 Molecular mechanisms of neonatal hypoxic-ischemic brain injury ...... 20

1.7.1 Overview of pathogenesis of neonatal HIE ...... 20

1.7.2 Excitotoxic cascade ...... 21

1.7.3 Inflammation ...... 21

1.7.4 Mitochondrial damage and dysfunction...... 23

1.7.5 Cell death mechanisms ...... 23

1.7.6 Glutamate-independent mechanisms ...... 25

1.8 Transient Receptor Potential (TRP) Channel Superfamily ...... 27

1.8.1 TRP superfamily classification ...... 27

1.8.2 TRPM family ...... 28

1.9 TRPM7 ...... 29

1.9.1 Gene and protein structure ...... 29

1.9.2 Biophysical characteristics and ion permeation ...... 30

1.9.3 Channel gating and regulation ...... 31

1.9.4 Current pharmacological modulators ...... 32

1.9.5 Tissue distribution ...... 35

1.9.6 TRPM7’s physiological and pathophysiological roles ...... 35

Chapter 2 Rationale, Hypothesis and Aims ...... 42

2 TRPM7 regulates neuronal development and injury through regulation of cytoskeleton ...... 42

2.1 Aim 1 ...... 42

2.2 Aim 2 ...... 43 vi

2.3 Aim 3 ...... 43

2.4 Aim 4 ...... 43

Chapter 3 Matrials and Methods ...... 45

3 Materials and methods ...... 45

3.1 Cell culture ...... 45

3.1.1 Dissociated culture of primary embryonic hippocampal neurons ...... 45

3.1.2 Drug treatment of hippocampal neurons...... 45

3.1.3 Hypoxic treatment of hippocampal neurons ...... 46

3.1.4 Flag-murine TRPM7/pCDNA4 HEK-293 cells...... 46

3.2 Functional recordings...... 47

3.2.1 Calcium imaging ...... 47

3.2.2 Electrophysiological recordings...... 47

3.3 Immunolabelling ...... 48

3.3.1 Immunocytochemistry ...... 48

3.3.2 Immunohistochemistry ...... 48

3.3.3 Antibodies used for immunolabelling ...... 48

3.4 Confocal microscopy and image analysis ...... 50

3.4.1 Image acquisition for neurite length analysis ...... 50

3.4.2 Neurite length analysis ...... 50

3.4.3 Protein Colocalization Analysis ...... 50

3.5 Identification of protein-protein interactions ...... 51

3.5.1 Co-immunoprecipitation ...... 51

3.5.2 Protein identification by mass spectrometry ...... 51

3.6 Mouse model of neonatal hypoxia-ischemia ...... 51

3.6.1 Animals ...... 51

3.6.2 Neonatal hypoxic-ischemic injury model ...... 52 vii

3.6.3 Drug administration ...... 52

3.7 Histological assessments ...... 52

3.7.1 Infarct volume measurement...... 53

3.7.2 Nissl staining ...... 53

3.8 Behavioural assessments ...... 53

3.8.1 Short-term neurobehavioural assessments ...... 53

3.8.2 Accelerated Rotarod Test ...... 54

3.8.3 Passive avoidance test ...... 54

3.8.4 Novel object recognition ...... 54

3.9 Proteomic analysis ...... 55

3.9.1 Visualization of proteomic analysis ...... 55

3.10 Western blotting ...... 55

3.11 Statistical Analysis ...... 57

Chapter 4 Results ...... 58

4 TRPM7 regulates axonal outgrowth and maturation of primary hippocampal neurons...... 58

4.1 TRPM7 block with waixenicin A enhances neurite outgrowth in developing hippocampal neurons ...... 59

4.1.1 TRPM7 protein is expressed in neurites and growth cones of developing hippocampal neurons ...... 59

4.1.2 Waixeinicin A reduces TRPM7-like current in hippocampal neurons in a dose-dependent manner ...... 61

4.1.3 TRPM7 block by waixenicin A decreases calcium influx into hippocampal neurons ...... 62

4.1.4 Waixenicin A enhances neurite outgrowth of cultured hippocampal neurons...... 63

4.1.5 Waixencin A preferentially enhances axonal growth of cultured hippocampal neurons ...... 67

4.1.6 Waixenicin A accelerates progression of hippocampal neurons through developmental stages ...... 69

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4.2 TRPM7 activation by naltriben mesylate induces neurite retraction in developing hippocampal neurons ...... 73

4.2.1 Naltriben activates TRPM7 currents in HEK-293 cells and primary hippocampal neurons ...... 73

4.2.2 Naltriben induces calcium influx under magnesium-free and basal conditions in hippocampal neurons ...... 75

4.2.3 Naltriben induces axon retraction and loss of processes in hippocampal neurons in a dose-dependent manner ...... 77

4.2.4 Waixenicin A attenuates the effect of naltriben on axonal lengths of hippocampal neurons ...... 79

4.3 TRPM7 mediates neurite outgrowth through cytoskeleton interaction ...... 80

4.3.1 A-actinin-1 and F-actin are identified as potential binding partners of TRPM7 by ESI-TRAP MS and confirmed by co-immunoprecipitation ...... 80

4.3.2 TRPM7 co-localizes with actin and α-actinin-1 at the neuronal growth cone ...... 81

4.4 Summary ...... 85

5 Characterizing the effects of hypoxia on TRPM7-mediated neurite outgrowth ...... 86

5.1 Short and long-term hypoxia differentially affects axonal outgrowth through regulating TRPM7 activity ...... 87

5.1.1 Short-term hypoxia enhances axonal outgrowth ...... 88

5.1.2 Short-term hypoxia causes downregulation of TRPM7 protein levels and activation of MEK/ERK and PI3K/Akt signaling pathways ...... 90

5.1.3 Long-term hypoxia causes axonal and dendritic retraction ...... 92

5.1.4 Naltriben exacerbates the effect of long-term hypoxia on axonal outgrowth ...... 93

5.1.5 Short-term hypoxia attenuates and long-term hypoxia potentiates TRPM7-like current in hippocampal neurons ...... 93

5.1.6 Waixenicin A attenuates hypoxia-induced axonal retraction when administered immediately after hypoxia ...... 95

5.2 Presence of astrocytes in culture exacerbates the effects of hypoxia on neurons ...... 97

5.2.1 Presence of astrocytes in culture exacerbates hypoxia-induced neurite retraction ...... 97

5.2.2 The effect of astrocytes on TRPM7-like current in hippocampal neurons ...... 99

5.2.3 The effect of soluble factors on axonal retraction ...... 101

5.3 Summary ...... 103

6 Neuroprotective and therapeutic effects of waixenicin A in mouse model of neonatal hypoxic-ischemic (HI) brain injury ...... 104

6.1 Model development and experimental timeline ...... 105

6.2 Waixenicin A has neuroprotective and therapeutic effects in neonal model of HI ...... 106

6.2.1 Waixenicin A pre-treatment reduces brain injury 24 hours after HI ...... 106

6.2.2 Waixenicin A reduces brain injury 24 hours after HI when administered up to 1 hour after the onset of injury ...... 107

6.2.3 Naltriben exacerbates brain injury 24 hours after HI when administered as a pre-treatment ...... 109

6.2.4 Waixenicin A pre-treatment reduces apoptotic brain death, reactive astrocyte activation and TRPM7 upregulation in ischemic penumbra...... 109

6.3 Waixenicin A preserves short-term function and overall brain morphology 7 days after HI ...... 112

6.3.1 Waixenicin A preserves brain morphology 7 days after HI injury ...... 112

6.3.2 Waixenicin A preserves short-term function following HI injury ...... 114

6.4 Waixenicin A preserves long-term function and overall brain morphology up to 32 days after HI ...... 116

6.4.1 Waixenicin A pre-treatment preserves Accelerated Rotarod outcomes ...... 116

6.4.2 Waixenicin A pre-treatment preserves memory test outcomes ...... 117

6.4.3 Waixenicin A pre-treatment preserves overall brain morphology ...... 120

6.5 Summary ...... 122

7 TRPM7-dependent molecular mechanisms in neonatal HI ...... 123

7.1 Characterization of protein changes 6 hours after HI ...... 124

7.1.1 Proteins with calcium-binding and cytoskeletal regulation functions ...... 126

7.2 TRPM7 signaling through CaMKII and regulation of cytoskeleton ...... 126

7.2.1 Proteomic changes of CaMKII, calcineurin and calmodulin ...... 127

7.2.2 Downstream signaling and cytoskeleton regulation from CaMKII, calcineurin and calmodulin ...... 127

7.3 Summary ...... 130

Chapter 5 Discussion and Conclusions ...... 131

8 Discussion ...... 132

8.1 Summary of findings...... 132

8.2 TRPM7 as a negative regulator of axonal outgrowth during development ...... 133

8.2.1 TRPM7 expression in hippocampal neurons suggests a developmental function ...... 134

8.2.2 TRPM7 maintains proper axonal development by inhibiting excessive growth and premature axonal specification ...... 134

8.2.3 TRPM7 regulates axonal outgrowth through calcium influx ...... 135

8.2.4 TRPM7 may serve as mechanosensor during neuronal development ...... 135

8.2.5 A model for TRPM7-mediated axonal outgrowth via interactions with actin and actinin ...... 136

8.3 TRPM7’s role in axonal retraction under pathophysiological conditions...... 137

8.3.1 TRPM7 and cytoskeleton in axonal retraction...... 138

8.3.2 Axonal injury in hypoxia-ischemia ...... 139

8.3.3 TRPM7 downregulation by short-term hypoxia ...... 140

8.3.4 The role of astrocytes in axonal retraction under hypoxic conditions...... 141

8.4 TRPM7 in neonatal hypoxic-ischemic brain injury...... 142

8.4.1 Suitability of the neonatal HI model ...... 143

8.4.2 Long-term effects of waixenicin A treatment ...... 144

8.4.3 Waixenicin A is a promising therapeutic target for drug development for neonatal HI ...... 145

8.5 TRPM7-mediated molecular mechanisms in HI brain injury ...... 146

8.5.1 Proteomic analysis of HI brain injury ...... 146

8.5.2 TRPM7-mediated regulation of cytoskeleton in HI ...... 147

8.6 Limitations of the current work ...... 149

8.6.1 The role of magnesium in neurite outgrowth ...... 149

8.6.2 PKC-dependent signaling in TRPM7-mediated neurite outgrowth ...... 150

8.6.3 Naltriben as TRPM7 activator and its potential off-target effects ...... 150

8.6.4 Systemic administration of waixenicin A in vivo ...... 151

8.6.5 The effect of waixenicin A on other cell types in the brain ...... 152

8.7 Future directions ...... 153

8.7.1 Other proteins at the neuronal growth cone ...... 154

8.7.2 The role of factors released by glia ...... 155

8.7.3 TRPM7 in other neurodegenerative conditions ...... 156

8.8 Significance...... 158

Appendices ...... 159

9 Appendix ...... 159

9.1 Appendix 1: Waixenicin A extraction method ...... 159

9.2 Appendix 2: Electrophysiological recordgings ...... 159

9.3 Appendix 3: Proteomic analysis ...... 160

9.3.1 TMT Q-Exactive samples preparation and analysis ...... 160

9.3.2 Database searching...... 160

9.3.3 Criteria for protein identification ...... 161

9.3.4 Quantitative data analysis ...... 161

9.4 Appendix 4: Confirmation of TRPM7 expression in HEK-293 cells...... 162

9.5 Appendix 5: Data sumary of axonal and dendritic lengths under normoxic and hypoxic conditions...... 163

9.6 Appendix 6: Data summary of axonal lengths under normoxic and hypoxic conditions with and without naltriben treatment...... 164

9.7 Appendix 7: Data summary of axonal and dendritic lengths in neuron-astrocyte co- cultures...... 164

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9.8 Appendix 8: Data summary of axonal lengths of neurons that received conditioned media from hypoxic neurons or hypoxic co-cultures ...... 165

9.9 Appendix 9: Proteins identified by LC-MS/MS ...... 165

9.10 Appendix 10: PKC interacts with TRPM7 C-terminus...... 169

9.11 Appendix 11: Neuron densities of neuronal cultures and neuron-astrocyte co-cultures. 171

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List of Tables

Table 1: Ca2+-dependent cytoskeletal regulators involved in neurite outgrowth...... 13

Table 2: Media/Solution Composition for Primary Hippocampal Neuronal Culture ...... 45

Table 3: Drug concentrations for treatment of hippocampal cultures...... 46

Table 4: Media/Solution Composition for HEK-293 cell culture ...... 47

Table 5: Antibodies used for immunolabelling...... 49

Table 6: Antibodies used for Western blot...... 56

Table 7: Summary of the co-localization analysis...... 84

Table 8: Data summary for Figure 25...... 163

Table 9: Data summary for Figure 26...... 164

Table 10: Data summary for Figure 29...... 164

Table 11: Data summary for Figure 31...... 165

Table 12: List of 95 differentially expressed proteins identified by LC-MS/MS...... 165

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List of Figures

Figure 1: The process of neuronal development of hippocampal neurons in culture...... 3

Figure 2: Cytoskeletal structure of neuronal growth cone ...... 4

Figure 3: Ion channels involved in hypoxic-ischemic neuronal death...... 27

Figure 4: Transient Receptor Potential (TRP) channel superfamily ...... 28

Figure 5: TRPM7 structural and biophysical characteristics...... 34

Figure 6: TRPM7 expression pattern in DIV2 hippocampal neurons ...... 60

Figure 7: Waixenicin A inhibits TRPM7 activity in a dose-dependent manner ...... 61

Figure 8: Waixenicin A reduces calcium influx in hippocampal neurons ...... 62

Figure 9: Waixenicin A enhances neurite outgrowth through neurite length and number of neurites 6 hours after treatment...... 64

Figure 10: Waixenicin A enhances neurite outgrowth and number of neurites 24 and 96 hours after treatment...... 66

Figure 11: Waixenicin A treatment preferentially enhances axonal outgrowth and branching ... 68

Figure 12. Treatment with Waixenicin A promotes maturation of hippocampal neurons...... 70

Figure 13: Quantitative analysis of neuronal transition between second and third developmental stages...... 72

Figure 14: Effects of waixenicin A and naltriben on TRPM7-like currents in TRPM7- overexpressing tetracycline-inducible HEK293 cells and dissociated hippocampal neurons...... 74

Figure 15: Effects of naltriben on TRPM7 calcium influx in dissociated hippocampal neurons . 76

Figure 16: Activation of TRPM7 by naltriben induces axonal retraction...... 78

Figure 17: Waixenicin A prevents axonal retraction caused by naltriben ...... 79 xv

Figure 18: TRPM7 coprecipitates with actin and α-actinin-1 ...... 80

Figure 19: TRPM7 colocalizes with actin and α-actinin-1...... 82

Figure 20: Changes in co-localization between TRPM7 and cytoskeletal proteins after treatment with waixenicin A...... 83

Figure 21: Western blot analysis of alpha-actinin-1 protein levels in control, vehicle and waixenicin A-treated neurons...... 85

Figure 22: The experimental timeline of hypoxia treatment...... 88

Figure 23: Short-term hypoxia enhances axonal outgrowth...... 89

Figure 24: Short-term hypoxia causes downtregulation of TRPM7 and activation of ERK and Akt singalling pathways...... 91

Figure 25: Long-term hypoxia causes axonal and dendritic retraction and TRPM7 block attenuates this retraction...... 92

Figure 26: Naltriben excerbates axonal retraction under hypoxic conditions...... 93

Figure 27: TRPM7 activity was reduced with short-term hypoxia, and enhanced with long-term hypoxia...... 94

Figure 28: TRPM7 block attenuates axonal retraction when administered immediately after hypoxia...... 96

Figure 29: Presence of astrocytes in culture exacerbates axonal retraction caused by hypoxia. .. 98

Figure 30: TRPM7 activity of neurons co-cultured with glia under short-term and long-term hypoxia...... 100

Figure 31: Axonal lengths following a media switch...... 102

Figure 32: Timeline of treatments, histological, morphological and neurobehavioural experiments...... 106

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Figure 33: Morphological assessment of brain injury in pre-treatment paradigm 24 hours after hypoxic ischemic insult...... 107

Figure 34: Morphological assessment of brain injury in post-treatment paradigm 24 hours after hypoxic ischemic insult...... 108

Figure 35: Morphological assessment of brain injury in pre-treatment paradigm with naltriben treatment 24 hours after hypoxic ischemic insult...... 109

Figure 36: Biochemical assessment of brain injury following hypoxic-ischemic insult on neonatal brain in a pre-treatment paradigm...... 110

Figure 37: Immunohistochemical assessment of brain injury following hypoxic-ischemic insult on neonatal brain in a pre-treatment paradigm...... 111

Figure 38: Morphological assessment of brain injury 7 days after hypoxic-ischemic insult on neonatal brain ...... 113

Figure 39: Assessment of motor and vestibular function up to 7 days after HI...... 115

Figure 40: Accelerated Rotarod Test administered 28 days after HI...... 116

Figure 41: Novel object recognition test 4 weeks after HI injury ...... 118

Figure 42: Passive avoidance test 4 week after HI...... 120

Figure 43: Brain morphology, brain weight and body weight 32 days after HI...... 121

Figure 44: Overview of proteomic changes between sham, vehicle and waixenicin A-treated groups after HI...... 125

Figure 45: Calcium-binding cytoskeletal regulators that were indentified by LC-MS/MS...... 126

Figure 46: Changes in CaMKII, calcineurin and calmodulin protein levels...... 127

Figure 47: Biochemical assessment of signalling pathways affected by hypoxic-ischemic insult on neonatal brain in a pre-treatment paradigm...... 129

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Figure 48: A model of TRPM7-mediated neurite outgrowth under normal and adverse conditions...... 137

Figure 49: Proposed TRPM7-mediated mechanism of neuronal cell death in neonatal HI...... 149

Figure 50: Confirmation of TRPM7 expression in TET-inducible HEK-293 cells...... 162

Figure 51: PKC interacts with TRPM7 C terminus ...... 162

Figure 52: Hypoxia does not have an effect on neuronal density ...... 162

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List of Abbreviations

AD Alzheimer's disease

ADF Actin-depolymerizing factor

ADP Adenosine diphosphate aEEG Amplitude-integrated electroencephalogram

ALS Amyotrophic lateral sclerosis

AMPA α-amino-3-hydroxy-5-methyl-4-isoxazole

APP β-amyloid precursor protein

ATP Adenosine triphosphate

BBB Blood-brain barrier

BDNF Brain-derived neurotrophic factor

BFA Brefeldin A

CaM Calmodulin

CaMKII Ca2+/calmodulin-dependent protein kinase II

CNS Central Nervous System

CRAC Calcium release activated channel

DIV Day in vitro

DMSO Dimethyl sulfoxide

DNA Deoxyribonucleic acid

EPSP Excitatory postsynaptic potential

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ERK Extracellular signal-regulated kinase

GAP-43 Growth associated protein 43

GFAP Glial fibrillary acidic protein

GFP Green Fluorescent Protein

GLAST Glutamate aspartate transporter

GLT-1 Glutamate transporter-1

GSK3-β Glycogen synthase kinase 3 beta

GTP Guanosine triphosphate

HEK Human embryonic kidney

HI Hypoxia-ischemia

HIE Hypoxic-ischemic encephalopathy

HUVEC Human umbilical vein endothelial cell

IgG Immunoglobulin G

IL Interleukin

JNK c-Jun N-terminal kinase

LC-MS/MS Liquid chromatography tandem-mass spectrometry

MAP2 Microtubule-associated protein 2

MAPK Mitogen-activated protein kinase

MARCKS Myristoylated alanine-rich C kinase substrate

MCAO Middle cerebral artery occlusion

MPO Multiparameter optimization

NCAM Neural cell adhesion molecule

NeuN Neuronal nuclear antigen

NgCAM Neuron-glia cell adhesion molecule

NGF Nerve growth factor

NMDA N-methyl-D-aspartate

NO Nitric oxide

NOS Nitric oxide synthase

PD Parkinson's disease

PI3K Phosphoinositide-3 kinase

PIP2 Phosphatidylinositol-4,5-bisphosphate

PKA Protein kinase A

PKC Protein kinase C

PLC Phospholipase C

PVL Periventricular white matter

RNA Ribonucleic acid

ROS Reactive oxygen species

SAC Stretch-activated channel

Sem3A Class 3 semaphorin siRNA Small interfering RNA

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SOCE Store-operated calcium entry

TGF-beta Transforming growth factor beta

TLR Toll-like receptor

TNF Tumor necrosis factor

TRAIL TNF-related apoptosis induced-ligand

TRP Transient Receptor Potential

TRPC Transient Receptor Potential Canonical

TRPM Transient Receptor Potential Melastatin

TRPV Transient Receptor Potential Vanilloid

TTC Triphenyl tetrazolium chloride

TTX Tetrodotoxin

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Chapter 1 Introduction

The functions of central nervous system (CNS) are dependent on the formation of highly complex neuronal networks. These networks develop through directed and tightly regulated outgrowth of axons and dendrites, collectively known as neurites. Several factors and signalling cascades have been implicated in regulation of neurite outgrowth and many are still being elucidated. Understanding the mechanisms of proper neuronal development may provide us with the knowledge needed to develop therapeutic interventions that can overcome the challenges of regeneration in the CNS during injury or disease.

Transient Receptor Potential Melastatin 7 (TRPM7), a member of the melastatin subfamily of the Transient Receptor Potential (TRP) channel superfamily, is a ubiquitously expressed cation channel that conducts Ca2+, Mg2+ and trace metal ions. TRPM7 has been shown to play a role in a wide range of cellular processes, such as ion homeostasis, cell survival and proliferation, neurotransmitter release, and cytoskeletal regulation. For many of these processes, the role of TRPM7 was tightly linked to its condunctance of divalent cations, such as Ca2+ and Mg2+. Based on the current knowledge of the mechanisms involved in neurite outgrowth, such as Ca2+- dependent regulation of the cytoskeleton, we hypothesized that TRPM7 might be involved in neurite elongation. Although TRPM7 is constitutively active, various factors such as pH and changes in extracellular divalent cations can modulate its activity. Often, these factors occur under the conditions of stress, and TRPM7 has previously been implicated in neuronal death under hypoxic/ischemic conditions in vitro and in vivo. The effect of hypoxia on neurite outgrowth has been controversial. Therefore, we also proposed to characterize the effect of hypoxia on neurite elongation/retraction and determine if TRPM7 regulates this process under the conditions of stress. Finally, we proposed to determine the therapeutic potential of a novel TRPM7 blocker waixenicin A and investigate TRPM7-dependent molecular mechanisms in an animal model of neonatal hypoxic-ischemic brain injury, a developmental condition caused by reduced oxygen supply to the neonatal brain due to trauma or disease.

Overall, this thesis will contribute knowledge on the regulatory mechanisms of neurite outgrowth by determining the role of TRPM7 in this process under normal and pathophysiological conditions in vitro. As well, it will determine the therapeutic potential of a novel TRPM7 blocker

1 2 in hypoxic-ischemic injury to the developing brain, thus proposing a potential treatment for this condition in the future.

1 Mechanisms of neurite outgrowth 1.1 Current model of neurite outgrowth

The directional outgrowth of neuronal projections, also known as neurites, is the fundamental process that occurs at every stage of neuronal development, leading to formation of two distinct compartments – a single axon and several dendrites – that differ in morphology, membrane composition, synaptic polarity and molecular constituents. Our current understanding of neurite outgrowth during neuronal development comes from a model described by Dotti and colleagues in 1988 (Dotti et al., 1988). In this model, the process of neuronal development has been characterized using embryonic hippocampal neurons and divided into five highly stereotyped and morphologically distinct stages (Dotti et al., 1988).

The first stage is characterized by formation of lamellipodia around the cell periphery and occurs shortly after the cells attach to substrate, usually within the first 6 hours in culture. The lamellipodia are thought to serve as sensors of the extracellular environment and contribute to neuronal motile machinery. The second stage is characterized by emergence of several motile processes that are similar in size, growth behaviour and molecular constituents. These processes develop within the first 24 hours in culture from the discrete patches of lamellipodia at intervals along the cell periphery and extend to a length of 10-15 μm (Dotti et al., 1988). The transition between this and the third stage marks the development of neuronal polarity (Craig and Banker, 1994). Several hours after the processes first appear, one of them begins to elongate at rate much faster than the others. This process acquires distinct growth behavior, morphological and molecular characteristics of an axon and can be clearly identified as soon as its growth rate intensifies. At this point, the neuronal symmetry is broken, and the rest of the processes become dendrites. The fourth stage is characterized by dendritic growth and development and takes place following approximately four days in culture. The final stage, known as maturation stage, is characterized by further development of axonal and dendritic arbors and formation of synaptic connections. The current model of neurite outgrowth is depicted in Figure 1.

Figure 1: The process of neuronal development of hippocampal neurons in culture. This image is adapted from Dotti et al., 1988, and depicts the transition of cultured hippocampal neurons through 5 morphologically distinct developmental stages. Lamellipodia, minor processes, axon and dendrites are marked by coloured arrows. 1.2 The cytoskeletal structure of neuronal growth cone

Growth cones, highly dynamic structures at the tips of elongating neurites, act as “vehicles” which move the protruding neurites towards their ultimate targets (Lowery and Van, 2009). Contact-mediated molecules including cell adhesion molecules (CAMs) and extracellular matrix proteins, and soluble cues such as neurotransmitters, growth factors and neurotrophic factors (Tessier-Lavigne and Goodman, 1996; Kiryushko et al., 2004; Lowery and Van, 2009), direct the growth cone behaviour and movement by triggering intracellular signaling cascades that lead to cytoskeletal rearrangement. A growth cone is separated into three regions based on its cytoskeletal composition as outlined in Figure 2 (Stiess and Bradke, 2011). The most distal region, also known as peripheral domain or P domain, contains mostly actin bundles (F-actin) that form filopodia and meshwork of actin filaments that form lamellipodia. The central domain of the growth cone or C domain contains stable microtubules, vesicles, and organelles that are transported along the microtubule bundles. Between the P and C domains there is a transition zone or T zone where microtubules and actin bundles interact (Lowery and Van, 2009; Stiess and Bradke, 2011). This highly specialized structure of the growth cone allows for fast cytoskeletal rearrangement in response to environmental guidance cues encountered by the growth cone.

According to the current model of neurite outgrowth first proposed by Mitchison and Kirschner in 1988, the growth cone of an elongating neurite repetitively progresses though three morphologically distinct stages. First, the encounter between the growth cone and adhesive substrate triggers a formation of “molecular clutch”, which is a physical coupling between the growth cone actin structures and the substrate. The formation of the “clutch” favours polymerization of actin filaments at the front and depolymerisation of actin filaments behind the clutch, leading to the protrusion of filopodia, also known as protrusion stage. Second, as actin behind the clutch undergoes depolymerisation, the microtubules invade the space previously occupied by actin filaments while bringing the organelles and vesicles along. This stage is known as engorgement. During the third stage, known as consolidation, actin filaments in the proximal end of the growth cone disappear, the microtubules become tightly bundled, and the membrane is reduced. What is left behind is a newly added segment of the neurite shaft (Kalil, 1996; Lowery and Van, 2009; Stiess and Bradke, 2011).

Figure 2: Cytoskeletal structure of neuronal growth cone. This image is adapted from Stiess and Bradke, 2011, and depicts the cytoskeletal elements that comprise the neuronal growthcone as well as the peripheral and central domains and transition zone.

1.3 Cellular events in neuronal polarization

Neurons are highly specialized cells with two structurally, molecularly and functionally distinct compartments, axons and dendrites. This structural asymmetry allows for a unidirectional flow of information within the neuronal network, with multiple dendrites receiving inputs from their surroundings and an axon integrating and transmitting these signals onto the next cell. Therefore, establishing and maintaining neuronal polarity is an important step towards a neuron becoming a functional cell.

The morphological changes that occur during neuronal polarization have been previously described by Dotti and colleagues (Dotti et al., 1988). The polarity is established once the neuron transitions from stage 2 to stage 3, normally within 24 hours in culture. At this point, one of the neurites begins to elongate rapidly, while the others exhibit little net elongation. The elongating neurite becomes and axon and acquires specific morphological and molecular characteristics (Dotti et al., 1988). Interestingly, once established the polarity is rather flexible and can be reversed under specific conditions. For example, when an axon of a hippocampal neuron was cut next to the cell body, one of the other processes, that would have become a dendrite initially, acquired axonal characteristics instead (Dotti and Banker, 1987). The distinguishing characteristic that may be the key to axonal development is the neurite length, as it has been shown that if the cut axon remains longer than the rest of the neurites, it still becomes an axon (Goslin and Banker, 1989). Moreover, the reversibility of polarity persists even in mature neurons. One study showed that a dendrite can be transformed into an axon even at a later stage of neuronal development (Bradke and Dotti, 2000). The evidence from these studies suggests that any neurite possesses the capacity to become an axon.

1.3.1 Directed membrane and protein trafficking and cytoplasmic flow

Establishment of an axon and its subsequent elongation require a large and coordinated increase in membrane surface area, cytoplasmic volume and axon-specific proteins. Therefore, it is possible that the transport of these materials is coordinated prior to axonal specification events that take place during stage 3. An observation of living neurons by Bradke and colleagues revealed that the polarized transport of lipids, organelles, vesicles and cytoplasmic proteins into the future axon exists before the neuron becomes polarized (Bradke and Dotti, 1997). Another study demonstrated that inhibition of vesicular traffic from the Golgi network by brefeldin A

(BFA) prevented axonal formation in stage 2 unpolarized neurons and inhibited axonal growth in stage 3 polarized neurons (Jareb and Banker, 1997).

It has been suggested that there are at least two separate mechanisms that contribute to polarized protein sorting during neuronal polarization. By using GFP-tagged axon and dendrite-specific proteins and observing their distribution at the cell surface, Burack and colleagues determined that some proteins are directly transported to the appropriate membrane, and some proteins are delivered non-specifically within the cells, where their insertion into the membrane is later determined by a post-transport mechanism (Burack et al., 2000; Silverman et al., 2001). These mechanisms have been found to be present in neurons as early as stage 3 (Silverman et al., 2001). Therefore, they could account at least in part to the formation of single axon.

1.3.2 Cytoskeletal dynamics

In addition to the cytoplasmic flow and protein sorting, the cytoskeletal dynamics at the growth cone of the future axon contribute to its specification. Local instability of actin, which is found within the peripheral region of the growth cone and composes filopodia and lamellipodia, has been shown to determine the initial neuronal polarization (Bradke and Dotti, 1999). Local application of cytochalasin D, a potent inhibitor actin polymerization, to one growth cone of a stage 2 neuron was sufficient to initiate axon formation, while a bath application resulted in a formation of multiple axons (Bradke and Dotti, 1999). Therefore, the most dynamic growth cone with greatest number of filopodia and lamellipodia in the stage 2 neuron is more likely to develop into an axon than the rest of the growth cones (Bradke and Dotti, 1999). Moreover, it’s been suggested that increased actin dynamics allow for enhanced polymerization of microtubules at the growth cone, leading to enhanced neurite elongation that follows axon specification (Forscher and Smith, 1988). Several studies consistently demonstrated that actin cytoskeleton regulates the microtubule cytoskeleton at the growth cone. For example, one study demonstrated the extension of microtubules from the central domain of the growth cone into the periphery, minutes after the application of cytochalasin D to cultured Aplysia neurons (Forscher and Smith, 1988).

While enhanced actin dynamics are widely accepted as an important phenomenon that contributes to axon specification, the enhanced stability of growth cone microtubules was also shown to contribute to neuronal polarization. As opposed to actin instability, microtubule

7 stability seems to correlate with axon formation (Witte et al., 2008). A study by Witte and colleagues demonstrated that during initial neuronal development and before the polarization, microtubule stability is increased in one of the neurites (Witte et al., 2008). Moreover, local microtubule stabilization by focal application of microtubule-stabilizing agent taxol, strongly predisposes the application site to axonal formation (Witte et al., 2008). Additionally, the stability of the microtubules seems to also influence actin dynamics at the growth cone. A recent study demonstrated that increased microtubule stability increases actin dynamics at the growth cone with the help of actin-binding protein Drebrin (Zhao et al., 2017), that’s been shown to facilitate microtubule entry into F-actin-rich structures such as growth cones (Worth et al., 2013).

Overall, this evidence suggests that the growth cone of the neuron that is destined to become an axon exhibits greater actin dynamics and greater microtubule stability.

1.3.3 Extracellular and substrate cues

Aside from the intrinsic cues such as polarized cytoplasmic flow and cytoskeleton dynamics, several lines of evidence implicate extracellular cues, both in vitro and in vivo, as regulators of neuronal polarization. Several studies demonstrated the role of different adhesion molecules and other extracellular cues in neuronal polarization with the use of striped substrates. One study demonstrated that in neurons plated on stripes of poly-L-lysine and either laminin or the neuron- glia cell adhesion molecule (NgCAM), the axons preferentially formed from neurites that came in contact with laminin or NgCAM (Esch et al., 1999). A different study, using a similar approach, showed that neurites of immature hippocampal neurons that came in contact with brain-derived neurotrophic factor (BDNF) coating reliably underwent axonal formation (Shelly et al., 2007).

A study by Polleux and colleagues demonstrated that in vivo axon emergence was regulated at least in part by secreted class 3 semaphorin, Sem3A, that repulses axonal initiation ventrally towards the ventricle (Polleux et al., 1998). Another diffusible cue that originates at the ventricular zone and induces axon formation in cortical neurons is transforming growth factor beta (TGF-beta). Elimination of TGF-beta results neurons lacking axons, while local application of TGF-beta in vitro is sufficient to induce axonal specification (Yi et al., 2010).

1.3.4 Current model of axon specification in neuronal polarization

According to an axon specification model proposed by Arimura and Kaibuchi (Arimura and Kaibuchi, 2007), the neurites of an unpolarized immature neuron extend and retract in a random fashion. Neurite extension is driven by increase in plasma membrane due to vesicle recruitment and fusion, local concentration and activation of growth-promoting signaling molecules, and enhanced actin dynamics and microtubule stabilization. Inhibitory growth signals sent by neighbouring neurites counteract positive regulation, induce microtubule destabilization, decrease actin dynamics and inhibit vesicle fusion to plasma membrane. Therefore, each neurite maintains its overall length though internal balance of inhibitory and growth-promoting events. When this balance is upset due to extracellular cues, contact with adhesion molecules or recruitment of specific signaling molecules, one of the neurites begins to elongate faster than others and the neuron transitions from stage 2 to stage 3. Continuous elongation of this neurite is supported by the positive feedback loop of increased membrane recruitment, protein transport, enhanced actin dynamics and microtubule assembly, while the progressively generated inhibitory signals interfere with axon specification of other neurites. Consequently, the growing neurite becomes an axon while the rest of the neurites acquire dendritic characteristics.

1.4 Cellular mechanisms of neurite outgrowth

1.4.1 The role of Ca2+ in neurite outgrowth

While many regulators of neuronal outgrowth have been described in the literature, there are numerous intracellular pathways that remain to be elucidated. It has been established that Ca2+ plays a major role in neurite outgrowth and motility, and intracellular calcium concentrations are stringently regulated (Kater and Mills, 1991). Brundage and colleagues showed that in migrating cells there was an asymmetric Ca2+ gradient with Ca2+ concentration being the lowest at the leading edge of the cell (Brundage et al., 1991). Within the neuronal growth cone, spontaneous Ca2+ “sparks” were shown to cause filopodial retraction and slow neurite outgrowth (Robles et al., 2003). Experimental evidence strongly supports the Ca2+ set-point hypothesis, first proposed by Kater and Mills, which states that normal growth cone motility depends upon an optimal range of intracellular Ca2+ concentrations (Kater and Mills, 1991). Ca2+ concentrations above the optimal range cause microtubule catastrophe, actin depolymerisation and growth cone collapse, while Ca2+ concentrations below the optimal range stabilize cytoskeletal structures and inhibit

9 growth cone motility (Kater and Mills, 1991). However, there is an additional complexity to Ca2+ signals within the neuron. Intracellular Ca2+ transients differing in amplitude, frequency and entry route were shown to have a range of effects on neurite elongation and motility (Kater and Mills, 1991; Mattson and Kater, 1987; Henley and Poo, 2004; Gomez and Zheng, 2006). For 2+ example, IP3-mediated release of Ca from internal stores promotes outgrowth (Takei et al., 1998), while Ca2+ spikes within the growth cones inhibit neurite outgrowth (Gomez and Spitzer, 1999; Tang et al., 2003) .Channels that are involved in generating these signals are often regulated by means other than voltage changes or neurotransmitter release (Shim et al., 2009; Kerstein et al., 2013) and in some cases are not yet identified (Gomez et al., 2001).

1.4.2 Ca2+-conducting membrane channels in neurite outgrowth

Ca2+ influx through plasma membrane channels was shown to have a diverse range of effects on growth cone behavior and neurite outgrowth. Several Ca2+-conducting channels have been shown to mediate neurite outgrowth and retraction. In rodent neonatal dorsal root ganglion neurons, the depolarization caused by high K+, veratidine or bradykinin, inhibited neurite outgrowth by 60% (Robson and Burgoyne, 1989). However, this inhibition was abolished by nifedipine application, suggesting that L-type Ca2+ channels may negatively regulate neurite outgrowth (Robson and Burgoyne, 1989). Similarly, mammalian cortical neurons showed enhanced axonal outgrowth in the presence of L-type Ca2+ channel blockers (Tang et al., 2003).

Another type of channels that has been studied in this context is plasma membrane stretch- activated channels (SACs), which are permeable to Ca2+ and monovalent cations. SACs have been shown to be densely distributed at the growth cone membrane of leech neurons, and inhibition of these ion channels with gentamicin significantly enhanced total length of the axon (Calabrese et al., 1999). Another study showed that in Xenopus spinal cord neurons Ca2+ influx through SACs negatively regulates neurite outgrowth, while Ca2+ influx through TRPC channels and from intracellular stores through IP3 and ryanodine receptors promotes neurite outgrowth (Jacques-Fricke et al., 2006).

The roles of Transient Receptor Potential (TRP) channels in neurite outgrowth have also been extensively studied due to their mechanosensing properties. Membrane stretch-dependent activation of TRPV2 in mouse dorsal root ganglia neurons enhanced neurite outgrowth (Shibasaki et al., 2010), while Ca2+ influx through TRPC1 channel in Xenopus spinal cord

10 neurons had an inhibitory effect on neurite outgrowth (Kerstein et al., 2013). TRPC5 channel has also been shown to regulate neurite outgrowth, however, there has been some controversy around it specific effects. In rat hippocampal neurons TRPC5 has been identified as a negative regulator of neurite extension (Greka et al., 2003), while in NG108-15 cells and primary rodent cerebellar granule neurons Ca2+ influx through TRPC5 was shown to initiate neurite outgrowth (Wu et al., 2007). TRPM2 channel activity, on the other hand, was shown to negatively regulate neurite outgrowth. Neurons isolated from TRPM2-deficient mice showed significantly longer neurites compared to wild type neurons, while activation of TRPM2 by ADP ribose in PC12 cells was associated with neurite retraction (Jang et al., 2014).

Extensive evidence demonstrates that Ca2+ signaling within the neuron and neuronal growth cone is complex and results in a wide range of behaviours which depend on the amplitude, frequency and entry route of the Ca2+ signals.

1.4.3 Ca2+-dependent modulators of neurite outgrowth

How could Ca2+ influx through different types of ion channels exert such diverse effects on neurite outgrowth? Jacques-Fricke and colleagues suggested that different channel types may function within specific and distinct microdomains (Jacques-Fricke et al., 2006). Ca2+ microdomains (clusters of ion channels) or Ca2+ nanodomains (single ion channel) are thought to be highly localized sites where Ca2+-conducting ion channels are functionally linked to specific Ca2+-sensitive effectors that exert a variety of downstream effects on the growth cone cytoskeleton leading to differential effects on neurite outgrowth (Jacques-Fricke et al., 2006).

Among the best described Ca2+-dependent effectors of neurite elongation are protein kinase C (PKC) and Ca2+/calmodulin-dependent protein kinase II (CaMKII)/calcineurin signaling cascades. Activation of these and other Ca2+-dependent effectors leads to phosphorylation of a number of proteins that interact with actin and microtubule cytoskeleton and modulation of the rate of neurite outgrowth/retraction. Ca2+-dependent effector proteins, their downstream targets and effects on neurite cytoskeleton and outgrowth are discussed below and summarized in Table 1.

1.4.3.1 Protein Kinase C (PKC)

Members of Ca2+ and phospholipid-dependent PKC family have been shown to be enriched at the neuronal growth cone (Cheng et al., 2000). Sustained activation of PKC has been shown to be necessary for filopodial protrusion of the growth cone in human SH-SY5Y neuroblastoma cells (Fagerstrom et al., 1996), while pharmacological inhibition of PKC in rat embryonic spinal cord cultures caused a significant retraction of neurites (Yang et al., 2010). Another study showed that neural cell adhesion molecule (NCAM)-stimulated neuritogenesis and neurite elongation in PC12-E2 cells is highly dependent upon activation of both PKC and MAPK/ERK signaling pathways (Kolkova et al., 2000). At the growth cone, activated PKC has also been shown to phosphorylate Growth Associated Protein 43/Neuromodulin (GAP-43), which leads to stabilization and formation of long actin filaments and growth cone protrusion (Liu and Storm, 1990). Another major substrate of PKC at the growth cone is myristoylated alanine-rich C kinase substrate (MARCKS), an actin cross-linking protein (Mikule et al., 2003a). MARCKS has been shown to localize to the growth cone membrane and stabilize integrin-mediated adhesions in its non-phosphorylated form. However, MARCKS phosphorylation by PKC causes it to translocate to the cytosol, resulting in localized growth cone detachment and collapse (Gatlin et al., 2006b). Additionally, MARCKS actin cross-linking activity has been shown to be inhibited by binding to Ca2+-calmodulin (CaM), suggesting that MARCKS is a potential conversion point of PKC and CaM signaling in regulation of actin cytoskeleton (Hartwig et al., 1992b).

1.4.3.2 Calmodulin, CaM-Kinase II and Calcineurin

Calmodulin (CaM), one of the key Ca2+-binding proteins that is abundant at the growth cone, binds Ca2+ at concentrations slightly above basal (Zucker, 1999). Ca2+/CaM modulate activity of variety downstream proteins that regulate growth cone motility. Several members of myosin family of motor proteins, which are well-known regulators of actin-based growth cone motility, have been shown to be regulated by Ca2+/CaM (Dent and Gertler, 2003b). In chick dorsal root neurons, myosin 1c and myosin II were shown to play opposing roles in growth cone protrusion (Diefenbach et al., 2002a). Ca2+/CaM was also shown to bind two structurally related proteins CAP23 and GAP43 that regulate neurite outgrowth through actin cytoskeleton rearrangement (Frey et al., 2000b). Several actin-binding proteins that regulate growth cone motility and protrusion, such as MARCKS, α-actinin, spectrin, MAP2 and tau, have also been shown to interact with and bind to Ca2+/CaM (Sobue, 1993b; Dehmelt and Halpain, 2004b).

Binding of Ca2+ to CaM leads to activation of CaMKII α and β isoforms, which are highly enriched in neurons and phosphorylate a wide range of substrates. CaMKII and ERK1/2 have been implicated in neuritogenesis and neurite outgrowth of cerebellar granule cells (Borodinsky et al., 2003), while CaMKII isoform β was shown to regulate neurite outgrowth in cultured hippocampal neurons (Fink et al., 2003). Cytosolic Ca2+ elevation caused by gangliosides was shown to lead to CaMKII activation and subsequent actin polymerization and formation of new filopodia in NG108-15 cells ad hippocampal neurons (Chen et al., 2003). In rat embryonic hippocampal neurons, overexpression of human recombinant S100A12 protein induced significant neurite outgrowth through activation of PKC, CaMKII, MEK/ERK and phospholipase C (PLC) pathways (Mikkelsen et al., 2001), suggesting that many different upstream regulators converge on these pathways to modulate neurite outgrowth.

Binding of Ca2+ to CaM can also activate phosphatase calcineurin that is also enriched at the growth cone and was shown to participate in neurite outgrowth regulation. Pharmacological inhibition of calcineurin has been reported to have conflicting effects on neurite outgrowth depending on the culture system. In Xenopus spinal neurons, pharmacological inhibition of calcineurin by cyclosporin A, deltamethrin and peptide inhibitors blocked Ca2+-dependent reduction of neurite outgrowth potentially through regulation of GAP-43 activity and actin cytoskeleton (Lautermilch and Spitzer, 2000b). In cultured chick dorsal root ganglia neurons, chromophore-assisted laser inactivation of calcineurin in regions of growth cone lead to filopodial and lamellipodial retraction (Chang et al., 1995a). Calcineurin was reported to regulate the activity of ADF/cofilin, an actin depolymerizing protein involved in regulation of neurite outgrowth, through its dephosphorylation (Meberg et al., 1998).

Table 1: Ca2+-dependent cytoskeletal regulators involved in neurite outgrowth.

Protein Regulation Function in Function in References cytoskeleton neurite regulation outgrowth

ADF/cofilin Activated by Depolymerizes Inhibits or (Meberg et al., calcineurin; F-actin enhances neurite 1998; Chang et inactivated by outgrowth al., 1995b; Wen LIM kinases depending on the et al., 2004; system Lautermilch and Spitzer, 2000a; Zhao et al., 2012)

Α-actinin Ca2+/CaM Actin-cross Involved in (Sobue, 1993c; binding linking protein; filopodial Sobue and anchors actin to protrusion, Kanda, 1989) membrane and motility, neurite cytoskeletal outgrowth and structures growth cone adhesion

Calpain Ca2+-dependent Protease; cleaves Activation slows (Robles et al., activation enzymes, neurite 2003; Glading et cytoskeletal, outgrowth, al., 2002) membrane and reduces receptor proteins lamellipodial protrusion, stabilizes filopodia

CAP23 CaM binding Promotes F-actin Induces neurite (Frey et al., accumulation at outgrowth and 2000a) the membrane branching

GAP-43 CaM binding; Binds to and Overexpression (Frey et al., PKC stabilizes long enhances neurite 2000c; phosphorylation actin filaments outgrowth Hocquemiller et al., 2010; Korshunova and Mosevitsky, 2010)

Gelsolin Ca2+-dependent F-actin severing Initiates (Lu et al., 1997) activation protein filopodial retraction

MAP2 CaM binding; Inhibits Enhances neurite (Dehmelt and microtubule outgrowth Halpain, 2004c; CaMKII assembly when Caceres et al., phosphorylation phosphorylated 1992; Poplawski by CaMKII; actin et al., 2012) cross-linking in unphosphorylated form

MARCKS Ca2+/CaM F-actin cross- Phosphorylation (Hartwig et al., binding; PKC linking protein leads to growth 1992a; Mikule et phosphorylation cone collapse al., 2003b; Gatlin et al., 2006a)

Myosin (I, II, V) Regulated by Drives retrograde Growth cone (Dent and Ca2+/CaM F-actin flow at extension; Gertler, 2003a; the growth cone regulation of Diefenbach et growth cone al., 2002b; Wang shape et al., 1996)

Spectrin Ca2+/CaM Cross-links actin Regulates growth (Sobue, 1993a) binding to plasma cone adhesion membrane adhesion

Synaptotagmin I Ca2+-dependent Exocytosis of Enhances neurite (Madgavkar et regulation secretory vesicles outgrowth al., 1977)

Tau CaM binding; Inhibits Regulates growth (Dehmelt and CaMKII microtubule cone elongation Halpain, 2004a) phosphorylation assembly when and motility phosphorylated

1.5 The effects of hypoxia on the developing brain

Hypoxia is a common cause of damage to the developing brain (Vannucci and Perlman, 1997; Vannucci, 2000; Perlman, 2006). The consequences of this injury are devastating, and survivors of perinatal hypoxia often face life-long disabilities, including cerebral palsy, neurodevelopmental delays, visual and learning deficits and other permanent disabilities. The intrinsic vulnerability of the neonatal brain to the hypoxic injury differs from the adult, as mechanisms such as glucose uptake and vasoregulation are largely underdeveloped in the neonate (Cimino et al., 2005). In the animal models, hypoxic injury to the developing brain causes enhanced activation of calpains, caspase-3 and mitochondrial pro-apoptotic signaling pathways compared to adult brain, leading to significant cell loss (Blomgren et al., 2001). Therefore, the pathophysiology of cerebral ischemia in neonate and adult differs at the molecular level. While understanding of the pathogenesis of the hypoxic brain injury in the fetus and neonate has been increasing, there are still many mechanisms that remain to be understood.

1.5.1 In vitro studies of effects of hypoxia on neurite outgrowth

To date, the effects of hypoxic conditions on neurite outgrowth are still under debate. It has been shown that exposure of rat cortical neurons in culture to hypoxic conditions caused neurite retraction with complete loss of processes after 24 hours of 5% hypoxia (Glass et al., 2002). However, it was also shown that severe (0.5%) hypoxia initiated neurite outgrowth in PC12 cells without addition of NGF for the first two days in culture, and caused neurite retraction on day in vitro 3 (DIV3) and cell death on DIV4 (O'Driscoll and Gorman, 2005). Another study on PC12 cells showed that exposure to 5% hypoxia for 6 hours caused a significant retraction of BDNF- and NGF-induced neurites (Woronowicz et al., 2007). While these discrepancies may arise from the differences in cell types, the clear effects of prolonged hypoxia on dendritic and axonal outgrowth of neurons and underlying molecular mechanisms have not been demonstrated.

1.5.2 The effects of hypoxia on neuronal cytoskeleton

Cytoskeletal structure and dynamics are compromised under hypoxic conditions making developing neurons especially sensitive to hypoxic insult. In Aplysia bag cell neurons, F-actin structure and dynamics, such as actin assembly at the leading edge, retrograde actin flow and filopodial protrusion at the growth cone were shown to be controlled by ROS (Munnamalai and

Suter, 2009). In rat retinas, hypoxic conditions were shown to lead to activation of calpains, subsequent degradation of cytoskeletal proteins and neuronal cell death (Tamada et al., 2005). Cytoskeletal actin rearrangements were also shown to take place in cortical neurons under hypoxic conditions, and inhibition of these changes by sequestering monomeric actin by latrunculin-A provided neuroprotection (Khan et al., 2008). Infusion of excitotoxin kainic acid into the brains of rats caused neuronal loss and cytoskeletal abnormalities in the hippocampus (Stein-Behrens et al., 1994). However, concrete mechanisms of cytoskeleton regulation by hypoxia and their effects on neuronal development and outgrowth are yet to be established.

1.6 In vivo models of hypoxic-ischemic injury to the developing brain

1.6.1 Hypoxic-ischemic encephalopathy

Hypoxic-ischemic encephalopathy (HIE) due to perinatal asphyxia, or impaired respiratory gas exchange accompanied by acidosis, is a major cause of neurological disability in term neonates (Johnston et al., 2011). It has an incidence of 1-8 per 1000 live births in developed countries and as high as 26 per 1000 live births in underdeveloped countries (Kurinczuk et al., 2010) and causes as many as a million neonatal deaths worldwide every year (Lawn et al., 2005). Several causes for neonatal HIE have been described, including antepartum factors such as pre- eclampsia, maternal thyroid disease and intrauterine growth retardation, as well as acute intrapartum obstetrical events, have been associated with this condition (Johnston et al., 2011). The outcomes of neonatal HIE are highly variable, and depend on a variety of factors such as duration and severity of the insult, number of repetitions of the insult and the maturity and general condition of the fetus (Johnston et al., 2011).

Some of the more severe outcomes of neonatal HIE include mental retardation, cerebral palsy, epilepsy and seizure disorders, motor impairments and visual and auditory impairments (Perez et al., 2013; Vannucci, 2000; Perlman, 2006). The risk of schizophrenia and other nonaffective psychoses is also elevated following hypoxia-ischemia-related neonatal complications (Zornberg et al., 2000).

1.6.2 Current treatments and interventions

Several clinical trials for treatments for neonatal HIE, including whole-body hypothermia (Shankaran et al., 2005), selective head cooling (Gluckman et al., 2005) and erythropoietin (Fauchere et al., 2008; Juul et al., 2008; McPherson and Juul, 2010), have been published. A randomized, controlled trial was conducted where whole-body cooling to esophageal temperature of 33.5 ºC was shown to reduce the rate of death or moderate/severe disability in term infants with HIE when administered within 6 hours after birth and continued for 72 hours (Shankaran et al., 2005). While the trial showed reduction in death rates from 62% (control group) to 44% (hypothermia) and cerebral palsy rates from 30% (control) to 19% (hypothermia), it was designed to include infants with high probability of death or disability and excluded preterm infants, those with congenital abnormalities and those that were unable to enroll in the trial within 6 hours of age (Shankaran et al., 2005).

The Cool-Cap Trial, a large, randomized controlled trial of delayed and selective head cooling with systemic hypothermia (34-35 ºC rectal temperature) only showed improved survival without severe disability in term infants (born at 36 weeks or longer gestation) with less severe abnormalities on amplitude-integrated electroencephalogram (aEEG) (Gluckman et al., 2005).

Erythropoietin (Epo) has been evaluated for neuroprotection in both term (37 weeks or later) infants (Zhu et al., 2009) and very low birth weight infants (Fauchere et al., 2008; Juul et al., 2008) with moderate success and few reported side effects. Neuroprotective effects of Epo were shown to be mediated potentially through PI3K/Akt and GSK3-β signaling (Zhang et al., 2006a; Byts et al., 2008; Shang et al., 2007).

Despite the recent advances in clinical and basic experimental research, the clinical prognosis for HIE remains poor suggesting that there is a need for novel therapies (Johnston et al., 2011). Therefore, animal models that closely resemble the events that take place in humans are of the utmost importance for moving the novel pre-clinical research towards drug and treatment development.

1.6.3 Current animal models of neonatal HIE

There is a wide variety of animal models used to study the pathogenesis of neonatal HIE. Until recently, umbilical cord occlusion (Gonzalez et al., 2005; Lotgering et al., 2004) or exposure to

18 maternal hypoxemia (Gleason et al., 1990; Harris et al., 2001) in fetal sheep were the most widely utilized models in this field. However, due to large costs associated with this model, as well as the lack of clinical evidence of brain injury in surviving lambs, other animal models had to be developed (Yager, 2004). Clancy and colleagues have previously established neurodevelopmental parallels, such as neurogenesis and axonal outgrowth, between different species, including rodents, cats, monkeys and humans (Clancy et al., 2001). These findings helped with the development of more accessible rodent models of neonatal HIE. The most commonly used model, 7-day-old rat or mouse, has been shown to have the brain maturity that corresponds to the early third trimester human fetus.

While many models utilizing different surgical and injection approaches are currently being used (reviewed in (Yager, 2004; Northington, 2006)), one of the most robust models of neonatal brain injury has been developed by Rice and colleagues (Rice, III et al., 1981). The investigators in this study adapted the Levine preparation, which involved unilateral ligation of the common carotid artery followed by hypoxia and was previously used on adult animals, to 7-day-postnatal rats (Rice, III et al., 1981). Since then, this procedure has been also adapted to a variety of mouse strains, thus providing the avenue for valuable pre-clinical molecular and biochemical investigations and testing of potential treatments and therapies.

Other, less widely used models involve induction of hypoxia without a preceding ischemic procedure. These models have been used to induce seizures in neonatal rodents (Rodriguez- Alvarez et al., 2015; Sampath et al., 2015), and manifest several behavioural phenotypes, such as hyperactivity, aggression, sleep disturbances and memory impairments (Decker et al., 2005; Mikati et al., 2005; Venerosi et al., 2006). While these hypoxia-only models have the potential to induce brain injury without unphysiological occlusion of the common carotid artery, there are currently several major disadvantages that need to be addressed. Currently, there is very little consensus in methodology, including age or strains of animals and experimental conditions such as severity and duration of hypoxia (Millar et al., 2017). Additionally, very few studies used standard behavioural testing, making it difficult to compare to other animal models (Millar et al., 2017). Finally, gene transcription analysis of rat pups that have undergone this model failed to show any difference in stress marker expression compared to controls, suggesting that this model does not generate adequate injury (Boss et al., 2005). Overall, far more careful considerations and design are needed, before these models can be used reliably.

Several other models that use inflammatory agents to induce cerebral inflammation, as seen in human patients, have been described. Administration of live E.coli into the uterus of pregnant rodents and intracervical injections of cell wall lipopolysaccharide (LPS) and simulation of viral infection by synthetic viral RNA have been associated with elevated cytokines, white matter injury and behavioural phenotypes in surviving pups (Bergeron et al., 2013; Wang et al., 2007; Rousset et al., 2013; Richetto et al., 2013). One of the strengths of this model is that it mimics the risk factors for severity of neonatal HI, such as maternal infection (Neufeld et al., 2005). However, the connection between maternal infection and fetal brain damage is currently under debate (Leviton et al., 1999). Moreover, there is a high variability in reported results, which makes this model difficult to compare to other models.

1.6.4 Selective regional and cellular vulnerability

HI causes selective damage to different brain structures that is determined by severity and duration of the insult and developmental stage of the brain at the time of the insult (Johnston et al., 2001; McQuillen et al., 2003; Schmidt-Kastner, 2015). Clinical evidence suggests that HI preferentially affects structures that control movement and tone. In pre-term infants (<32 weeks of gestation), periventricular white matter (PVL) is particularly vulnerable, resulting in motor, cognitive and sensory deficits (Johnston et al., 2002). In term infants, HI causes selective damage to sensorimotor cortex, thalamus, basal ganglia and brain stem leading to motor disability with impairment of upper limbs and speech difficulties (Johnston et al., 2001; Martin et al., 1997; Hoon, Jr. et al., 1997; Menkes and Curran, 1994).

The Rice-Vannucci procedure, which involves the unilateral carotid artery ligation combined with exposure to moderate hypoxia, causes ipsilateral injury of several brain regions in neonatal mice and rats. Numerous studies have investigated short and long-term functional and behavioural outcomes, demonstrating motor and cognitive impairments and severe cerebral abnormalities (Skoff et al., 2007; Chen et al., 2015a; Huang et al., 2017c; Huang et al., 2017b; Sun et al., 2015; Xiao et al., 2015; Xu et al., 2015). Additionally, it’s been shown that HI injury evolves over time, causing prolonged tissue damage (Geddes et al., 2001). Clinical and pre- clinical evidence suggests that this regional vulnerability is primarily due to excessive activity of excitatory synapses (Pu et al., 2000). The selectively vulnerable regions, such as somatosensory cortex, basal ganglia and thalamus, have been shown to have high glucose metabolic rate

(Blennow et al., 1995). Functional studies found that increased glucose metabolism strongly correlates with enhanced synaptic activity (Sokoloff, 1999; Sibson et al., 1998; Pfund et al., 2000). Moreover, these brains regions have been shown to be interconnected by active glutamatergic neurons, suggesting that regional differences in brain injury after HI depend on the position of the region within excitatory circuits (Johnston and Hoon, Jr., 2000).

1.6.5 Evidence of axonal damage in rodents and humans

Axonal injury is an important, but poorly understood event that contributes to brain injury following HI in adults (Wang et al., 2016; Hinman, 2014), and preterm and term infants with mild and severe forms of HIE (Govaert et al., 2008; Mathur et al., 2010; Martinez-Biarge et al., 2012). Moderate to severe white matter/axonal abnormality has been reported in approximately 60% of term infants (gestational age of 36-43 weeks) with HIE. These axonal abnormalities were also found to correlate with outcomes such as global developmental delay, visual, behavioural and communication abnormalities, and seizures (Martinez-Biarge et al., 2012).

Rodent studies have reported that developing axons are particularly vulnerable to ischemic injury during the period of myelination (Fern et al., 1998), and that HI causes significant axonal damage, as suggested by increased accumulation of β-amyloid precursor protein (APP) (Lin et al., 2004).

1.7 Molecular mechanisms of neonatal hypoxic-ischemic brain injury

1.7.1 Overview of pathogenesis of neonatal HIE

HI brain injury is an evolving process that develops over the period of days or even weeks (Sarnat and Sarnat, 1976). During normal conditions, the human brain has a high requirement for oxygen and glucose that are used to produce an adenosine triphosphate (ATP) via the process of oxidative phosphorylation. HI conditions rapidly inhibit this process, resulting in a primary energy failure, leading to massive releases of glutamate into the extracellular space. At this point, the severity and duration of HI insult will dictate the pattern and severity of the subsequent brain injury. During this time, the fetus is able to maintain some degree of homeostasis by reducing nonobligatory energy consumption by the organs and maintaining anaerobic respiration (Vannucci, 1990; Jensen and Berger, 1991; Jensen et al., 1999). However, these conditions will

21 rapidly lead to accumulation of lactic acid and severe metabolic acidosis (Graham et al., 2008). If the initial insult is prolonged and severe, a second wave of energy failure starts as early as 6 hours after initial injury, and is accompanied by production of reactive oxygen species (ROS), increases in intracellular Ca2+ concentrations, inflammation and mitochondrial dysfunction (Sanders et al., 2010). During this phase, majority of the cell death occurs. A tertiary phase may also occur within days after initial injury and may continue for months afterwards. This phase involves late cell death, astrogliosis, remodeling and repair (Sanders et al., 2010).

1.7.2 Excitotoxic cascade

One of the earliest events that occur during HI injury is an excito-oxidative biochemical cascade that results from a shift from high-energy phosphate metabolism towards anaerobic respiration due to insufficient oxygen (Jensen and Berger, 1991). The failure of the cell membrane ion transport leads to subsequent activation of intracellular cascades that result in cell death. As the Na+/K+ pumps on the cell membrane stop functioning due to ATP shortage, the change in membrane voltage leads to depolarization and release of pre-synaptic glutamate (Jensen et al., 2003; Johnston et al., 2001). Since glucose availability is severely limited, reuptake of glutamate by glia is also reduced, resulting in overactivation of glutamate receptors. Neurons express several glutamate receptors, including ligand-gated N-methyl-D-aspartate (NMDA) receptors, α- amino-3-hydroxy-5-methyl-4-isoxazole (AMPA) receptors, and kainate receptors. Overstimulation of glutamate ionotropic receptors leads to increased Ca2+ influx into the neurons, causing activation of lipases, proteases, and endonucleases and cytoskeletal degradation (Johnston et al., 2002; Choi, 1988). Several rodent studies demonstrated that cytoplasmic accumulation of cations causes severe cell swelling and results in necrotic cell death (Carloni et al., 2007; Chavez-Valdez et al., 2012). Excitotoxicity affects multiple highly metabolic brain regions, such as cerebral cortex, thalamus, and putamen (Sie et al., 2000; Johnston et al., 2001; Johnston et al., 2002).

1.7.3 Inflammation

HI brain injury initiates a rapid inflammatory response that involves both intrinsic and infiltrating immune cells and lasts for days and even weeks after the initial insult (Hedtjarn et al., 2004; Winerdal et al., 2012). The role of the initial inflammation is targeted at removing damaged cells and debris and limiting infections at the site of injury. Following the early pro-

22 inflammatory phase of the response, the immune system initiates anti-inflammatory and reparative phases. The initial inflammatory response is characterized by rapid activation of microglia and mast cells, the resident immune cells of the brain (McRae et al., 1995; Jin et al., 2009). This innate immune response has toxic influences on the neurons, oligodendrocyte precursor cells and the vasculature. This leads to increased permeability of the blood-brain barrier (BBB) which allows the peripheral macrophages, monocytes and neutrophils to infiltrate the injury site (Ek et al., 2015). Both intrinsic and infiltrating cells produce cytokines (e.g. IL-1α, IL-18, TNF-α) (Jin et al., 2009; Bona et al., 1999; Palmer et al., 2004), chemokines, ROS, excitatory amino acid agonists, and tumor necrosis factors (e.g. FasL, TRAIL, TWEAK, TNF-α, TNF-β) (Kichev et al., 2014) which could contribute to cell death.

Microglial activation and subsequent release of pro-inflammatory cytokines, as early as 2 hours after the injury, was shown to contribute to axonal damage (Deng et al., 2008; Deng et al., 2010) and play an important role in initiating secondary energy failure (Baburamani et al., 2014). However, pharmacological depletion of microglia prior to injury exacerbates outcome and enhances the release of pro-inflammatory cytokines, suggesting that there is a subpopulation of microglia that have beneficial effects (Faustino et al., 2011). Microglial phagocytosis has been shown to be important in tissue recovery during the delayed recovery phase (Woo et al., 2012). It’s been suggested that there are different functional phenotypes of microglia, with some participating in acute inflammatory phase, while others contribute to anti-inflammatory recovery phase (Faustino et al., 2011).

Astrocytes, the major type of glia in the brain, play a critical role in glutamate uptake, formation of glial scar that surrounds the injury site, and comprise some of the BBB (Koizumi et al., 2018; Li et al., 2008; Sen and Levison, 2006). Some evidence also implicates reactive astrocytes in contributing to the injury development by releasing pro-inflammatory cytokines TNF-α and IL-6 (Deng et al., 2010). Aquaporin-4 (AQP4) channels expressed on astrocytes have been shown to contribute to brain edema during HI, thus potentially exacerbating the extent of the injury (Fu et al., 2007).

Other immune cells have also been shown to participate in the inflammatory processes following HI injury. Neutrophils accumulate in neonatal brain in much smaller amounts than in adult brain, and have been suggested to contribute to early injury progression (Hudome et al., 1997; Palmer

23 et al., 2004). Mast cells, other immune cells present in the brain, have been implicated as early contributors to the disruption of the blood-brain barrier (BBB) and formation of edema (Lindsberg et al., 2010).

1.7.4 Mitochondrial damage and dysfunction

While primary energy failure can be rapidly compensated with redistribution of blood flow by cerebroprotective mechanisms (Jensen and Berger, 1991; Jensen et al., 1999), the secondary energy failure that occurs with prolonged hypoxia or during brain reperfusion causes a substantial exhaustion of cellular energy reserves (ATP) and leads to cell death (Blumberg et al., 1997). This secondary energy depletion is associated with increased production of lactate, pH fluctuation and increased production of oxidative particles (Hope et al., 1984; Penrice et al., 1997). As we know, brain has a high requirement for aerobic respiration which translates into higher rate of mitochondrial respiratory activity. During HI, mitochondrial electron transport chain is disrupted, leading to a dissipation of the proton gradient in the inner membrane. This in turn, leads to accumulation of Ca2+ in the inner membrane and mitochondrial depolarization (Hagberg et al., 2014). Excessive Ca2+ accumulation leads to activation of nitric oxide synthase (NOS), which in turn starts to produce nitric oxide (NO). Inhibition of NOS prior to insult and deletion of neuronal NOS (nNOS) has been shown to be highly protective in rodent models of neonatal HI (Hamada et al., 1994; Ferriero et al., 1996; van den Tweel et al., 2002). NO combines with superoxide radicals to produce peroxynitrite, which then quickly decomposes into NO2+ (nitrogen dioxide) and hydroxyl radicals (Beckman, 1991). This causes mitochondrial permeabilization and loss of function. Accumulation of oxidative particles also increases trafficking of cytochrome C and apoptosis-inducing factor (AIF) from the outer mitochondrial membrane into the cytoplasm, where it triggers activation of caspase-3. This in turn initiates intrinsic pathway-mediated apoptosis and DNA fragmentation (Robertson et al., 2009; Hagberg et al., 2009). In neonatal brain, the inhibition of Bax, a protein that initiates the permeabilization of the outer mitochondrial membrane and causes the release of cytochrome C into the cytoplasm, has been shown to significantly reduce brain injury (Wang et al., 2010; Han et al., 2011).

1.7.5 Cell death mechanisms

Neuronal cell death is the hallmark of HI brain injury, and leads to irreparable damage to the brain and life-long disability in 20-40% of HIE patients (Johnston et al., 2011). Several types of

24 cell death that take place in the neonatal brain following HI have been broadly classified as apoptotic and non-apoptotic. Apoptosis is thought to account for a significant portion of neuronal death following HI, and occurs during early reperfusion and in the following hours and days (Wang et al., 2001). Oxidative stress and mitochondrial dysfunction are central to initiating apoptotic cell death (Wang et al., 2001). Histologically, it can be characterized by cell rounding, membrane blebbing, cytoplasmic condensation, chromatin condensation (pyknosis) and fragmentation, and formation of membrane-bound apoptotic bodies (Elmore, 2007). On the molecular level, the key proteins involved in apoptosis are apoptotic peptidase-activating factor 1 (APAF1), Bax, cytochrome C and executioner caspase-3 (Cheng et al., 1998; Wang et al., 2001; Ota et al., 2002; Wang et al., 2010). It is important to mention that apoptosis is critical in normal brain development and regulation of the size and shape of the central nervous system (CNS) (Raff et al., 1993; Kuan et al., 2000). In some brain regions, more than half of the initially formed neurons undergo apoptosis (Raff et al., 1993), and mice lacking caspase-3 exhibit hyperplasic and disorganized brains (Kuida et al., 1996).

While the canonical apoptotic pathway is one of the most studied cell death mechanisms, the majority of cell death in neonatal brain occurs by necrosis (Northington et al., 2011b). Histologically, it has been characterized by cytoplasmic swelling, nuclear dissolution and membrane rupture that leads to an inflammatory response, and begins early during the development of HI injury (Laster et al., 1988). A highly regulated programmed form of necrosis, known as necroptosis (Galluzzi et al., 2011), was shown to occur under the conditions of extreme ATP depletion (Leist et al., 1997) or during caspase inhibition (Vercammen et al., 1998), and recognized as an important mechanism of cell death in immature brain after injury (Northington et al., 2011a; You et al., 2008). Necroptosis is commonly induced by death receptor ligands (TNF-α, FasL, TRAIL) or by Toll-like receptors (TLR) 3 and 4 (Vanlangenakker et al., 2012). This initiates the recruitment of receptor-interacting kinase 1 (RIP1) which leads to recruitment and interaction with RIP3 and formation of necrosome. RIP3 then recruits its substrate, pseudokinase mixed lineage kinase domain-like (MLKL), into the necrosome and initiates its phosphorylation and oligomerization. In this form the necrosome is “activated” and can facilitate the permeabilization of cell and organelle membranes leading to cell lysis observed in necrosis (Galluzzi et al., 2011). Caspase-8 has been shown to actively inhibit necroptosis through

25 degradation of RIP1 and RIP3 and initiate apoptosis instead (Lin et al., 1999; Oberst et al., 2011).

Another distinct type of cell death that’s been described in neonatal HI brain injury is autophagy, a degradative pathway that involves the delivery of cytoplasmic cargo to the lysosome (Galluzzi et al., 2016). The major molecular characteristic of autophagy is sequestration of damaged proteins and organelles into a double-membrane autophagosome which is marked by LC3, the autophagy marker, that later fuses with the lysosome for protein degradation (Choi et al., 2013). There is accumulating evidence showing that autophagy plays a significant role in neuronal cell death in neonatal HI brain injury. Mice deficient in autophagy-related protein 7 (Atg7) exhibited remarkable neuroprotection (Xie et al., 2016), while in neonatal rats most of the neuronal cell death in CA3 region appeared to be by autophagy, as characterized by high number of autophagosomes, empty vacuoles and ballooning of the perinuclear space (Koike et al., 2008). Moreover, the autophagy marker LC3 has been detected in the brains of human neonates who died after severe perinatal asphyxia (Koike et al., 2008).

1.7.6 Glutamate-independent mechanisms

A previous clinical strategy was to pharmacologically target activation of NMDA receptors after the excessive glutamate release following HI injury, thus preventing excitotoxicity from occurring. Compounds such as dezocilipine maleate (MK-801) (Buchan et al., 1992; Gerriets et al., 2003), aptiganel hydrochloride (Cerestat) (Lees, 1997), dexthrometorphan (DMX) (van Rijen et al., 1991) and CGS 19755(Selfotel) (Grotta et al., 1990a; Grotta et al., 1990b) were tested in rodent models with promising results. However, these findings did not translate to humans and by 2001 all clinical trials using NMDA receptor antagonists for HI injury were deemed unsuccessful (Albers et al., 2001; Morris et al., 1999; Albers et al., 1999; Davis et al., 1997; Davis et al., 2000; Sacco et al., 2001). Therefore the clinical and pre-clinical efforts were re- focused on finding novel non-glutamate therapeutic targets for HI brain injury, and several candidates were identified as potential therapeutic targets.

Acid sensing ion channels (ASICs) have been identified as glutamate-independent contributors to HI brain injury. Specifically, the activation of ASIC1a, an ion channel that is widely expressed in the brain, has been shown to contribute to neuronal cell death in both in vitro and in vivo models (Xiong et al., 2004; Yang et al., 2011b; Yang et al., 2007). Anaerobic respiration during

HI injury leads to production of lactic acid, causing the cerebral pH in the neonate to drop as low as 6.7 (Yang et al., 2011b). This state of acidosis was shown to induce Ca2+ influx through ASIC1a channels, thus implicating them in injury development.

ATP-sensitive potassium (KATP) channels are outwardly rectifying channels that act as metabolic sensors in pancreatic β cells and are involved in insulin secretion. In the neonatal brain, these channels have been implicated in ischemic tolerance and neuroprotection by hypoxic preconditioning following HI (Sun et al., 2015). The increase in ADP/ATP ratio following HI injury activates KATP channels causing hyperpolarization and decrease in neuronal excitability (Munoz et al., 2003). Therefore, a potential strategy would be to potentiate the activity of these channels to increase ischemic tolerance in the brain.

Volume-regulated anion channels (VRACs) regulate cell volume via efflux of ions that are followed by water, thus contributing to cerebral edema following HI (Pedersen et al., 2015). VRACs expressed in astrocytes have also been implicated in excitotoxicity, by conducting excitatory amino acids (EAAs), aspartate and glutamate, following HI injury (Alibrahim et al., 2013; Zhang et al., 2008; Feustel et al., 2004).

Connexin hemichannels, proteins that form the pore of the gap junction between neighbouring cells, have also been implicated in neuronal death in HI injury (Thompson et al., 2006). It’s been shown that they contribute to spread of anoxic depolarization into the penumbra region, and allow for efflux of vital nutrients from neurons, thus worsening the effects of energy failure caused by low oxygen conditions (Kozoriz et al., 2010; Davidson et al., 2012; Davidson et al., 2014).

Transient Receptor Potential Melastatin (TRPM) channels are non-selective, ubiquitously expressed cation channels (Fleig and Penner, 2004; Harteneck, 2005). Two closely related members of TRPM family, TRPM7 and TRPM2, have recently emerged as major contributors to non-glutamate cell death following HI (Bae and Sun, 2013; Aarts and Tymianski, 2005; Zhan et al., 2016). In vitro and in vivo rodent studies showed contribution of TRPM7 to neuronal death in following HI (Aarts et al., 2003; Sun et al., 2009). A closely related TRPM family member, TRPM2, has also been investigated in its contribution to HI injury in the neonatal brain (Huang et al., 2017c; Huang et al., 2017b).

Figure 3: Ion channels involved in hypoxic-ischemic neuronal death.

1.8 Transient Receptor Potential (TRP) Channel Superfamily

1.8.1 TRP superfamily classification

Transient Receptor Potential (TRP) superfamily is a group of unique ion channels that are conserved across vertebrates and invertebrates. These channels are widely expressed in numerous cell types and serve as cellular sensors for a wide variety of physical and chemical stimuli. TRP channels were shown to be involved in physiological processes such as sight, hearing, touch, smell, smell and temperature and pain sensation (Ramsey et al., 2006).

The first TRP channel was discovered in a mutant strain of Drosophila melanogaster lacking a functional copy of trp gene (Cosens and Manning, 1969). Unlike the wild-type fruit flies which exhibited a plateau-like receptor potential to continuous light, this mutant showed an abnormal transient receptor potential electroretinogram response (Cosens and Manning, 1969). The

28 predicted trp gene product was postulated to be an ion channel (Montell and Rubin, 1989), and was later confirmed to be a Ca2+-permeable ion channel that was activated by light in photoreceptors (Hardie and Minke, 1992).

There are currently 28 identified mammalian TRP channels, all of which contain 6 transmembrane domains and are permeable to cations causing cell depolarization once activated. They are grouped by sequence homology into 6 different subfamilies: 1) TRPC (canonical, TRPC1-7), 2) TRPM (Melastatin, TRPM1-8), 3) TRPV (vanilloid, TRPV1-6), 4) TRPA (ankyrin, TRPA1), 5) TRPML (mucolipin, TRPML1-3) and 6) TRPP (polycystin, TRPP1-3) (Ramsey et al., 2006). Since classification of TRP channels into subfamilies is based on sequence similarity rather than functional features, the members of the same subfamilies exhibit great functional diversity.

Figure 4: Transient Receptor Potential (TRP) channel superfamily. TRP superfamily is divided into 6 subfamilies based on amino acid sequence: TRPC, TRPM, TRPV, TRPA, TRPML and TRPP. TRPM (melastatin) subfamily has 8 family members grouped together based on amino acid sequence.

1.8.2 TRPM family

The mammalian TRPM subfamily contains 8 members, TRPM1-8. They are further divided into three separate subgroups based on amino acid sequence similarity: TRPM1/TRPM3,

TRPM4/TRPM5 and TRPM6/TRPM7, with TRPM2 and TRPM8 are being separate from the rest of the family (Zheng, 2013).

All TRPM members contain a very long (~700 amino acids) TRPM homology region located in the N-terminus. TRPM4 and TRPM5 are unique, due to their selectivity for monovalent cations, while TRPM2, TRPM6 and TRPM7 contain enzymatically active C-terminal domains (Zheng, 2013).

1.9 TRPM7

1.9.1 Gene and protein structure

In the human genome, the TRPM7 gene, also known as CHAK1, TRP-PLIK and LTRPC7, is located on chromosome 15q21.2, spans 127 kb of DNA sequence and encodes 39 exones (NCBI Gene ID 54822) (Runnels et al., 2001; Nadler et al., 2001) (Figure 5A). The human TRPM7 gene encodes for a protein that is 1,865 amino acid residues long with a predicted molecular weight of 212 kDa (Schmitz et al., 2005), and TRPM7 orthologs with 94.3 – 99.89% human amino acid similarity have been identified in chimpanzee, mouse, rat, dog and cow (Fleig and Chubanov, 2014).

Similarly to other TRPM family members, TRPM7 monomers are through to form tetrameric units (Nadler et al., 2001; Schnitzler et al., 2008). The TRPM7 monomer can be subdivided into several distinct domains. The N-terminus contains hydrophobic region (H1) and TRPM homology domain (MHD), both with poorly understood functions. Next, there are 6 putative transmembrane domains with a pore forming loop between segments 5 and 6 (Nadler et al., 2001). C-terminal to S6, there is a highly conserved 24 amino acid “TRP box” which is unique to all identified TRP channels, and is thought to be an interaction site with phosphatidylinositol-

4,5-bisphosphate (PIP2). The “TRP box” is followed by a cytoplasmic coiled-coil (CC) domain that is thought to mediate channel assembly and trafficking (Fujiwara and Minor, Jr., 2008). Distal to CC domain there is a functional serine/threonine kinase domain that is homologous to atypical α-kinase family. This domain is unique to TRPM7 and its paralog TRPM6 (Nadler et al., 2001; Runnels et al., 2001). Structural evidence shows that the kinase domain contains an ATP- binding site and Zn2+ and Mg2+ binding sites (Yamaguchi et al., 2001) (Figure 5B).

Mass spectrometry studies on TRPM7 identified 47 autophosphorylation sites mostly located in the serine/threonine-rich region of the kinase domain, which is thought to be important for kinase substrate binding (Clark et al., 2008c). Other key phosphorylation sites have been identified on the cytoplasmic C-terminus, with 3 located in CC domain and 7 located in the serine/threonine- rich region and 2 other sites located distal to the kinase region (Kim et al., 2012; Matsushita et al., 2005).

The functional roles of distinct TRPM7 domains have been identified with the use of various TRPM7 mutants. Mutations S138L and P1040R correspond to mutations in human TRPM6 that are associated with an inherited disease of hypomagnesemia with secondary hypocalcemia (Schlingmann et al., 2002; Walder et al., 2002). In TRPM7, P1040R results in a dominant- negative channel (Chubanov et al., 2007), while S138L interferes with channel assembly and membrane trafficking (Chubanov et al., 2004). The E1047Q mutation in ion selectivity filter of the pore-forming segment results in an active channel, permeable to monovalent, but not divalent cations (Schnitzler et al., 2008; Li et al., 2007). The S1107E mutation produces a constitutively active channel that is insensitive to intracellular Mg2+ concentrations (Hofmann et al., 2014). The double mutation of K1112Q and R1115Q disrupts channel function with no PIP2 sensitivity (Xie et al., 2011). The K1646R mutation produces a “kinase-dead” channel without any phosphotransferase activity (Kaitsuka et al., 2014).

1.9.2 Biophysical characteristics and ion permeation

TRPM7 is a constitutively active outward-rectifying cation channel with single channel conductance of ~40 pS and a reversal potential of ~0 mV (Runnels et al., 2001; Penner and Fleig, 2007; Wu et al., 2010). TRPM7 exhibits selectivity for divalent metal ions at hyperpolarized potentials and carries a small inward current. At depolarized potentials, TRPM7 carries a strong outward current as intracellular cations are forced to exit the cell (Nadler et al., 2001) (Figure 6C). This strong outwardly rectifying current-voltage (I/V) relationship is due to voltage- dependent permeation block of monovalent cations by Ca2+ and Mg2+. If divalent cations are removed, TRPM7 shows no intrinsic voltage dependence and exhibits a quasi-linear I/V relationship (Nadler et al., 2001). The level of TRPM7 activity is regulated by a wide range of intracellular and extracellular factors.

TRPM7 permeation profile to divalent metal ions is as follows: Zn2+ = Ni2+ > Ba2+ > Co2+ > Mg2+ > Mn2+ > Sr2+ > Cd2+ > Ca2+. As opposed to monovalent cations, the conductance of these ions is not affected by the permeation block (Monteilh-Zoller et al., 2003). Several amino acid residues located in the TRPM7 pore have been identified as important for channel’s Ca2+ and Mg2+ permeability. Substitution of glutamic acid with neutral glutamine in mouse and human at residues 1047 and 1052 strongly reduces the channel’s affinity for Ca2+ and Mg2+, respectively (Li et al., 2007; Schnitzler et al., 2008). Additionally, the permeation block by divalent cations was shown to be regulated by aspartic acid residues at 1054 and 1059 (Numata and Okada, 2008).

1.9.3 Channel gating and regulation

One of the most important regulators of TRPM7 activity is intracellular free Mg2+ (Nadler et al., 2001), and its effect can be mimicked by Ba2+, Sr2+, Zn2+ and Mn2+ (Kozak and Cahalan, 2003). Mg2+ -mediated inhibition involves two separate binding sites, one located within the kinase domain and another on the channel itself (Demeuse et al., 2006; Chokshi et al., 2012a). TRPM7 activity has been shown to increase when intracellular Mg2+ and Mg2+-bound nucleotides were chelated or omitted from solution altogether (Runnels et al., 2001; Kozak and Cahalan, 2003; Demeuse et al., 2006). Single channel analysis of TRPM7 conductance revealed that the primary effect of Mg2+ inhibition is due to reduction of the number of conducting channels and a modest reduction (~20%) in unitary channel conductance (Chokshi et al., 2012b). Mg2+-bound nucleotides (Mg-ATP and Mg-ADP) have been shown to regulate channel activity synergistically with Mg2+ and are postulated to have binding sites independent from Mg2+ (Demeuse et al., 2006).

Several studies have reported regulation of TRPM7 activity through various receptor-mediated pathways. In HEK293 cells TRPM7 inactivation was induced by carbachol stimulation of heterologously co-expressed muscarinic 1 receptors and subsequent depletion of PIP2 by phospholipase C (PLC) in the plasma membrane (Runnels et al., 2002). In rat CA1 hippocampal neurons, TRPM7-like currents were inhibited in a dose- and time-dependent manner through NGF-mediated activation of TrkA receptor and PLC (Tian et al., 2007). In pig isolated ventricular myocytes nonhydrolysable GTP analogues inhibited TRPM7-like current, potentially through G-protein coupled receptor and PLC activity (Macianskiene et al., 2008). Stimulation of

32 beta-adrenergic receptors in TRPM7-overexpressing HEK293 cells was shown to enhance TRPM7 activity through protein kinase A (PKA) activity and required an intact TRPM7 kinase domain (Takezawa et al., 2004).

Cell volume and mechanical stress were also shown to modulate TRPM7 activity. In HeLa cells, endogenous TRPM7 channels were directly activated by stretch or increased cell volume (Numata et al., 2007), and in TRPM7-overexpressing HEK293 cells osmotically induced changes in cell volume were shown to modulate channel activity primarily through changes in cytosolic free Mg2+ (Bessac and Fleig, 2007). In vascular smooth muscle cells sheer stress induced by laminar flow caused TRPM7 translocation to the plasma membrane, suggesting that TRPM7 may be involved in mechano-sensing (Oancea et al., 2006).

Extracellular acidity (~ pH 6) also greatly potentiates TRPM7 current (Monteilh-Zoller et al., 2003) via direct competition of protons with divalent cations for specific binding sites in the channel pore (Jiang et al., 2005). This competition leads to change in TRPM7 selectivity and increased permeation by monovalent cations at negative membrane potentials (Jiang et al., 2005). Specifically, TRPM7’s pH sensitivity was shown to be governed by glutamic acid residues 1047 and 1052 (Li et al., 2007). This biophysical characteristic of TRPM7 becomes relevant under pathophysiological conditions, when extracellular environment becomes acidic.

1.9.4 Current pharmacological modulators

Several pharmacological inhibitors of TRPM7 have been described in the literature, although most of them lack specificity, potency or both. Some of the non-specific TRPM7 blockers are: 3+ 3+ trivalent ions (i.e. Gd , IC50 ~1.4-2.5 μM; La , IC50 ~17 μM) (Aarts et al., 2003), spermine (IC50

~2.3 μM) (Kozak et al., 2002), 2-aminoethyl diphenylborinate (2-APB, IC50 ~50-174 μM)

(Prakriya and Lewis, 2002; Li et al., 2006), SKF-96365 (IC50 ~20 μM) (Kozak et al., 2002), carvacrol (IC50 ~307 μM) (Parnas et al., 2009), serine protease inhibitor and coagulant nafamostat mesylate (IC50 ~617 μM) (Chen et al., 2010b) and several 5-lipoxygenase inhibitors

(NDGA, IC50 ~6.3 μM; AA861, IC50 ~6.0 μM; MK886, IC50 ~8.6 μM) (Chen et al., 2010a). Several other compounds, such as ginsenoside Rg3 (Kim et al., 2011), ginsenoside-Rd (Kim, 2013), aripiprazole (Sato-Kasai et al., 2016), midazolam (Chen et al., 2016) and coomassie brilliant blue G-250 (Norenberg et al., 2016), were also reported as TRPM7 inhibitors in the high μM range, but their exact effects need further investigation.

Sphingosine (IC50 ~600 nM), a component of the plasma membrane spingolipids, and its structural analogue fingolimod (IC50 ~720 nM), an FDA-approved treatment for multiple sclerosis, inhibit TRPM7 activity by reducing the open probability of the channel (Qin et al., 2013). Another study, using aequorin bioluminescence-based assay, identified several Ca2+- activated K+ channel inhibitors to act on TRPM7: quinine, CyPPA, dequalinium, NS8593, SKA31, and UCL1684 (Chubanov et al., 2012). Among those, the most potent TRPM7 inhibitor 2+ NS8593 (IC50 ~1.6 μM) was thought to interfere with Mg -dependent regulation of TRPM7 (Chubanov et al., 2012).

The first specific and potent inhibitor of TRPM7 waixeinicin A, a natural terpenoid of Sarcothelia edmondsoni, was identified by Zeirler and colleagues with the use of high- throughput drug-screening bioassay using fluorescent-based Mn2+ quench in TRPM7- overexpressing HEK293 cells (Zierler et al., 2011). Waixenicin A was shown to block TRPM7 currents in a magnesium-dependent manner (IC50 ~16 nM), and had no effects on the other members of TRPM family (TRPM2, TRPM4 and TRPM6) or Ca2+ release-activated Ca2+ (CRAC) channels (Zierler et al., 2011). This study provided the scientific community with a novel pharmacological tool to further develop in-depth understanding of physiological and pathophysiological functions of TRPM7.

Small molecular activators of TRPM7 have also been recently investigated. Among 20 drug-like compounds, naltriben, δ opioid antagonist, showed selectivity for TRPM7 and was able to 2+ activate the current without prior depletion of intracellular Mg and under low PIP2 conditions (Hofmann et al., 2014). Two benzimidazole relatives of TRPM7 inhibitor NS8593, mibefradil and NNC 50-0396, have also been recently identified as specific Mg2+-sensitive agonists of TRPM7 (Schafer et al., 2016).

Pharmacological tools for manipulation of TRPM7 kinase activity in living cells are also being developed. A study by Davis and colleagues screened 72 kinase inhibitors, and found that TG100-115, a potent inhibitor of phosphoinositide 3-kinase-γ (PI3Kγ), was able to inhibit purified TRPM6 kinase domain with IC50 of 8 nM (Davis et al., 2011). A later study found that

TG100-115 also inhibited TRPM7 kinase domain with IC50 of ~2 μM (Song et al., 2017).

In summary, a set of potent and specific pharmacological tools has been identified for further investigation of TRPM7 function, as well as future development of high-affinity drugs that target the channel.

Figure 5: TRPM7 structural and biophysical characteristics. A) The TRPM7 gene is located on chromosome 15 and contains 39 exons. B) The structure of TRPM7 subunit and pharmacological modulators. C) Heterologously expressed TRPM7 current.

1.9.5 Tissue distribution

TRPM7 is a ubiquitously expressed ion channel (Nadler et al., 2001; Runnels et al., 2001). As shown by quantitative real-time reverse transcriptase polymerase chain reaction (qRT-PCR) analysis of human tissue, TRPM7 was expressed in the following organs and tissues: brain, pituitary, heart, lung, liver, fetal liver, kidney, skeletal muscle, stomach, intestine, spleen, peripheral blood mononuclear cells, macrophages, adipose, pancreas, prostate, placenta, bone, cartilage and bone marrow. The highest expression of TRPM7 was found in heart, pituitary, bone and adipose (Fonfria et al., 2006). Compared to other members of TRPM family, TRPM7 showed a consistently higher expression levels across majority of tissues (Fonfria et al., 2006).

Studies have also investigated TRPM7 expression in mouse organ systems. While TRPM7 levels vary across different mouse strains, TRPM7 was found to be the most abundantly expressed out of all TRP channels, with the highest expression levels in brain, lung, kidneys, intestine and testis in the adult mouse (Kunert-Keil et al., 2006). During mouse development, TRPM7 gene expression seems to follow a bimodal expression pattern, first peaking at embryonic day 18 (E18) and then again after postnatal day 4 (P4) and maintaining stable levels thereafter (Staaf et al., 2010).

TRPM7 endogenous currents have also been confirmed in a wide range of commonly used cell lines, such as embryonic kidney (HEK293), mast cells (rat RBL-2H3), lymphocytes (human Jurkat T) (Nadler et al., 2001), human breast cancer (MDA-MD-231) (Guilbert et al., 2013) and others.

1.9.6 TRPM7’s physiological and pathophysiological roles

TPM7 has been implicated in a wide variety of physiological processes, including ion homeostasis, cytoskeleton regulation, cell survival, proliferation and migration and organ development. Disruption of normal TRPM7 function has been associated with neurodegeneration under the conditions of stress, progression of cancer and cardiovascular disease.

1.9.6.1 TRPM7 in ion homeostasis

TRPM7 was shown to regulate intracellular levels of divalent cations such as Mg2+, Ca2+ and heavy metal ions. Targeted TRPM7 deletion from chicken B lymphocytes led to reduced

36 intracellular Mg2+ levels, cell proliferation arrest and cell death which was rescued by supplementing with high extracellular Mg2+ (Schmitz et al., 2003). Embryonic stem cells derived from heterozygous TRPM7(kinase) mice displayed a proliferation arrest phenotype that can be rescued by Mg2+ supplementation. Additionally, these mice developed hypomagnesaemia due to a defect in intestinal Mg2+ absorption (Ryazanova et al., 2010). However, tissue-specific deletion of TRPM7 in mouse T lymphocytes did not alter total Mg2+ content of these cells nor systemic Mg2+ levels in these mice (Jin et al., 2008). This was likely due to compensation by other Mg2+ transporters present in those cells, such as MagT1 (Li et al., 2011).

TRPM7 was also shown to be important for maintaining cytosolic Ca2+ concentrations during several physiological processes. TRPM7 kinase-dead K1646R knock-in mice exhibited splenomegaly and impaired blastogenesis (Beesetty et al., 2018). This was due to reduction in store-operated calcium entry (SOCE) pathway in these cells, which required functional TRPM7 kinase (Beesetty et al., 2018). Mice with a loss-of-function mutation in TRPM7 kinase domain showed altered platelet responses, such as decreased granule secretion and aggregation, also due to defective SOCE mechanism (Gotru et al., 2018).

Moreover, TRPM7 was also shown to be involved in heavy metal ion homeostasis. In osteoblasts, TRPM7 was implicated in cadmium (Cd2+) uptake and cytotoxicity (Martineau et al., 2010; Levesque et al., 2008). In mouse embryonic stem cells, TRPM7 was shown to be located on intracellular vesicles that were distinct from other organelles and sequestered Zn2+ during cytosolic overload. It has been hypothesized that TRPM7-mediated Zn2+ release from these vesicles regulated ROS signaling during embryonic development (Abiria et al., 2017).

1.9.6.2 TRPM7 in cell survival, proliferation and differentiation

Several lines of evidence show that genetic or pharmacological inhibition of TRPM7 activity in proliferating tissues arrests G0/G1 transition of the cell cycle. In rat mast cells RBL-2H3, TRPM7-like current was significantly enhanced during the G0/G1 transition, and TRPM7 suppression by siRNA increases the rate of apoptosis in these cells (Ng et al., 2012; Tani et al., 2007). Similarly, majority of TRPM7-deficient DT-40 B lymphocytes was shown to be arrested in quiescence/G0 phase, and this state can be reversed by supplemental Mg2+ or induction of TRPM7 expression (Sahni et al., 2010). In human malignant glioma MGR2 cells, midazolam, a commonly used clinical anesthetic, induced G0/G1 cell cycle arrest through inhibition of

TRPM7 activity (Chen et al., 2016). Pharmacological inhibition of TRPM7 in hepatic stellate cells led to apoptosis induced by TNF-related apoptosis induced-ligand (TRAIL), suggesting that TRPM7 activity is necessary for survival of these cells (Liu et al., 2012). Similarly, in primary human lung mast cells (HLMCs) and human mast cell lines, LAD2 and HMC-1, TRPM7 knockdown by shRNA induced cell death that was not rescued by Mg2+ supplementation (Wykes et al., 2007).

The role of TRPM7 in proliferation has been confirmed in numerous cell lines. In rat liver cells, TRPM7 channels were shown to be differentially expressed during different developmental stages, with higher expression found in proliferating cells compared to terminally differentiated and non-dividing cells (Lam et al., 2012). TRPM7 knockdown in human breast cancer lines (MCF-7 and hBCE) (Guilbert et al., 2009), head and neck carcinoma lines (FaDu, SCC25) (Jiang et al., 2007), glioblastoma lines (U87, U251) (Chen et al., 2015b), retinoblastoma cells (Hanano et al., 2004), human osteoblast-like cells (Abed and Moreau, 2007), rat aortic vascular smooth muscle cells (Yang et al., 2017), human colorectal adenocarcinoma (HT-29) (Huang et al., 2017a), mouse cortical astrocytes (Zeng et al., 2015) and macrophages (Schilling et al., 2014) led to suppression of proliferation of these cell types.

1.9.6.3 TRPM7 in regulation of cytoskeleton

TRPM7 has been implicated in cell adhesion, migration, guidance and overall cell morphology through regulation of cytoskeletal structures. TRPM7 overexpression in HEK293 cells led to cell rounding and detachment from substrate (Nadler et al., 2001; Su et al., 2006) through stress- dependent activation of p38 mitogen-activated (MAP) and c-Jun N-terminal (JNK) kinases and stimulation of m-caplain activity (Su et al., 2010). The activation of these kinases was thought to be dependent on TRPM7 activity, and inhibitors of these kinases abolished TRPM7-induced cell rounding and detachment (Su et al., 2010). Interestingly, this effect was not dependent on the nature of the divalent cation conducted by TRPM7, as alteration of external Ca2+ and Mg2+ concentrations to favour the permeation of one cation over the other produced similar rounding and m-caplain activity levels (Su et al., 2010).

TRPM7 kinase domain was also implicated in cell adhesion. In N1E-115 neuroblastoma cells, TRPM7 activation by bradikinin was shown to cause TRPM7 kinase activation, interaction with actomyosin cytoskeleton and phosphorylation of myosin IIA heavy chain (Clark et al., 2006). On

38 the morphological level, TRPM7 overexpression led to cell spreading and reorganization of cytoskeleton to facilitate podosome formation. These effects were kinase-dependent, as cells expressing the kinase-dead TRPM7-D1775A mutant did not show similar responses to bradikinin stimulation (Clark et al., 2006). TRPM7 was also shown to physically interact with myosin IIA and β-actin, suggesting that it may exert its effects on actomyosin function through direct interaction with cytoskeletal proteins (Clark et al., 2006; Clark et al., 2008b). Additionally, it was shown that TRPM7 and TRPM6 also phosphorylate the assembly domain of myosin IIB and IIC, and that the interaction of TRPM7 with its substrates may be dependent on the massive autophosphorylation of Ser/Thr-rich region (Clark et al., 2008a).

TRPM7 was also shown to regulate cell migration, a complex process that requires coordinated rearrangement of the cytoskeletal structures. In human T lymphocytes, TRPM7, together with KCa3.1 channel, was shown to regulate cell migration through localizing to the uropod of the cell and regulating T cell migration machinery (Kuras et al., 2012). In human, embryonic lung fibroblasts (WI-38), genetic suppression of TRPM7 reduced the number of high-calcium microdomains or “calcium flickers”, membrane tension and traction force at the leading-edge of the migrating cells (Wei et al., 2009). In Swiss 3T3 fibroblasts, TRPM7 depletion by RNA interference altered cell morphology, disrupted the cytoskeleton and reduced the ability of cells to form lamellipodia to carry out cell movements through inhibition of Rac and Cdc42. However, the expression of Mg2+ transporter SLC41A2 reversed these phenotypic changes, suggesting that Mg2+ concentrations may control TRPM7-dependent cytoskeleton regulation (Su et al., 2011).

More recently, TRPM7 was studied in a variety of cancer cell lines as a regulator of invasiveness of these cells. TRPM7 knockdown or pharmacological inhibition of TRPM7 activity was shown to suppresses the migration and invasion in bladder cancer (Gao et al., 2017), prostate cancer (Chen et al., 2017), glioma (Chen et al., 2016), pancreatic ductal adenocarcinoma (Rybarczyk et al., 2017), breast cancer (Guilbert et al., 2013), human glioblastoma (Chen et al., 2015b), neuroblastoma (Middelbeek et al., 2016), nasopharyngeal carcinoma (Qin et al., 2016), ovarian carcinoma (Wang et al., 2014), retinoblastoma (Hanano et al., 2004) and leukemia (Zierler et al., 2011). TRPM7 regulation of cytoskeleton is thought to be at least in part responsible for its oncogenic properties.

1.9.6.4 TRPM7 in neurotransmitter release

TRPM7 was also implicated in vesicle trafficking and neurotransmitter release (Krapivinsky et al., 2006). TRPM7 was found to localize to the membrane of synaptic vesicles of rat presynaptic sympathetic neurons and interact with synaptic vesicle proteins synapsin I, synaptotagmin and snapin. It was hypothesized that TRPM7 Ca2+ conductance is crucial to neurotransmitter release, and the changes in TRPM7 protein levels correlated with changes in post-synaptic EPSP amplitudes and kinetics (Krapivinsky et al., 2006). In a subsequent study, TRPM7 was also found in acetylcholine-secreting small synaptic-like vesicles in PC12 cells. TRPM7 knockdown or expression of dominant-negative TRPM7 altered the neurotransmitter quanta and release probability (Brauchi et al., 2008). This study postulated that due to TRPM7 pH sensitivity and

PIP2 gating mechanism, the channel acts as a “coincidence detector” and opens only when vesicle pH is low and PIP2 in the plasma membrane binds the cytoplasmic domain of TRPM7. These events occur just prior to fusion, and enhanced TRPM7 activity at this point would facilitate vesicle fusion with the plasma membrane (Brauchi et al., 2008).

1.9.6.5 TRPM7 in organ development

The role of TRPM7 in embryonic development and organogenesis has been studied with the use of several genetically-modifiable vertebrate and invertebrate models. Several mouse lines that carry mutant alleles in Trpm7 gene have been described (Jin et al., 2008; Ryazanova et al., 2010). Trpm7 null mutant mice and mutants that lack exons encoding the kinase domain were shown to be embryonic lethal on day 6.5-7.5 (Jin et al., 2008). Conditional mutagenesis using Cre/loxP recombination, revealed that TRPM7 is indispensible before and during organogenesis, as disruption of Trpm7 after e14.5 did not have an effect on embryo development, and Trpm7 inactivation in adult mice produced no distinguishable phenotype (Jin et al., 2008).

Several transgenic mouse lines with tissue-specific Cre activity were used to study the involvement of Trpm7 in organ development. Trpm7 deletion in the mouse T cell lineage led to disruption of thymopoiesis and developmental block of thymocytes without altering Mg2+ homeostasis (Jin et al., 2008). Moreover, knockout thymocytes exhibited disrupted production of growth factors needed for maintenance of medullary epithelial cells (Jin et al., 2008). In kidney, Trpm7 disruption in metanephric mesenchyme resulted in abnormal nephrogenesis with decreased number of glomeruli, dilation of renal tubules and cyst formation (Jin et al., 2012).

Disruption of Trpm7 at e10.5 in neural crest cells resulted in the loss of skin pigmented cells and dorsal root ganglion sensory neurons, leading to discoloration of lower trunk fur and hind leg paralysis (Jin et al., 2012). Early (before e9) cardiac deletion of Trpm7 resulted in congestive heart failure and death by e11.5, and Trpm7 deletion between e.9 and e.13 produced 50% rate of cardiomyopathy, impaired repolarization and ventricular arrhythmias. Trpm7 deletion after e.13 did not affect ventricular size or function (Sah et al., 2013).

Zebrafish (D. rerio) and frog (Xenopus laevis) have also been used as models to study Trpm7 function. Trpm7-deficient zebrafish undergo normal early morphogenesis, but later develop defective melanin synthesis, death of melanophores, kidney stones, loss of touch responsiveness, reduced proliferation of epithelial cells in the exocrine pancreas and lethality in later larval life (Elizondo et al., 2005; Elizondo et al., 2010; Low et al., 2011; McNeill et al., 2007; Yee et al., 2011). Partial rescue of melanophores and epithelial cells of the exocrine pancreas could be accomplished with Mg2+ supplementation (Yee et al., 2011; Elizondo et al., 2010). In the frog embryo, Trpm7 was shown to regulate cell polarity and convergent extension movements during gastrulation leading to an abnormal phenotype. This phenotype could be reversed with Mg2+ supplementation or expression of Mg2+ transporter SLC41A2 (Liu et al., 2011).

Trpm7 function was also studied in two invertebrate models – fruit fly (D. melanogaster) and roundworm (C. elegans). In the fruit fly, a single Trpm gene lacking the kinase domain is highly expressed in the Malpighian tubules that function in removal of electrolytes and toxic compounds, and Trpm-null mutants exhibited pupal lethality (Hofmann et al., 2010). In C.elegans, three TRPM channels GON-2, GTL-1 and GTL-2 also lack kinase domains and regulate Mg2+ uptake at the intestine and Mg2+ excretion by excretory cells (Teramoto et al., 2005; Teramoto et al., 2010).

In summary, several studies using different animal models demonstrated TRPM7 role in embryonic development, organogenesis and ion homeostasis.

1.9.6.6 TRPM7 in cerebral ischemia

In the brain, the role of TRPM7 is most often studied under the conditions of stress. Several lines of evidence implicate TRPM7 in contributing to glutamate-independent Ca2+ overload under ischemic conditions. A number of in vitro studies demonstrate the contribution of TRPM7 to

41 neuronal death under oxygen-glucose deprivation conditions (Aarts et al., 2003; Zhang et al., 2011). Conditions that occur in the brain under hypoxic stress, such as low extracellular pH (Jiang et al., 2005), ROS production (Aarts et al., 2003; Coombes et al., 2011), changes in divalent cations (Wei et al., 2007) and intracellular Zn2+ accumulation (Inoue et al., 2010) also potentiate TRPM7 activity and lead to cell death. Consistent with in vitro data, small hairpin RNA-mediated suppression of TRPM7 in the CA1 hippocampal region of adult rats, protected the neurons from cell death and preserved normal neuronal morphology after transient global ischemia (Sun et al., 2009). Functionally, these animals also demonstrated better outcomes on fear-associated and spatial navigation memory tests compared to controls (Sun et al., 2009). Moreover, TRPM7 expression was shown to be upregulated following middle cerebral artery occlusion in the hippocampus region, suggesting increased vulnerability of these neurons to ischemic injury (Jiang et al., 2008). Recently, TRPM7 currents in primary cortical neurons were shown to be regulated by the pro-inflammatory cytokine IL-6, which contributes to pathogenesis of cerebral ischemia (Liu et al., 2016). Another study demonstrated that increased Ca2+ influx through TRPM7 increased the levels of autophagy through CaMKKβ-dependent pathway, a process that is associated with cell death under ischemic conditions (Oh et al., 2015). Additionally, TRPM7-like currents have also been identified in rat brain microglia (Jiang et al., 2003) and mouse cortical astrocytes (Zeng et al., 2015), suggesting that TRPM7 activation under adverse conditions may contribute to inflammatory responses mediated by these cells.

A current model of TRPM7 in cerebral ischemia proposes that “Ca2+ paradox” or initial reductions in extracellular divalent cations and concomitant decrease in extracellular pH potentiate TRPM7 activity. Further progression of ischemic condition leads to ROS production, further potentiating TRPM7 currents. This positive feedback mechanism may prolong neuronal Ca2+ overload, thus contributing to neuronal cell death even after inactivation of NMDARs. However, recent evidence suggests that TRPM7 contribution to cerebral ischemia may be more complex, and there are additional molecular mechanisms that remain to be identified. While increasing evidence strongly implicates TRPM7 in neuronal cell death in cerebral ischemia, the channel’s role remains largely phenomenal. Concrete molecular mechanisms need to be identified to better understand TRPM7 contribution to pathogenesis of ischemic injury and develop targeted therapies for this condition.

Chapter 2 Rationale, Hypothesis and Aims 2 TRPM7 regulates neuronal development and injury through regulation of cytoskeleton

Rationale: Based on the current evidence from the literature, TRPM7 plays a role in a variety of physiological processes, such as actin cytoskeleton regulation, ion homeostasis, adhesion and migration. TRPM7 was also shown to modulate and directly interact with cytoskeletal proteins. As these processes are crucial in neuronal development and maturation, TRPM7 channels may play a role in regulation of neurite outgrowth during development of mammalian neurons. Moreover, TRPM7 biophysical characteristics, such as sensitivity to changes in pH, divalent cation concentrations and ROS, demonstrate that TPRM7 has an important role under pathophysiological conditions.

In my graduate program, I proposed to investigate the molecular mechanisms that are regulated by TRPM7 activity during normal neuronal development and under hypoxia-induced stress in vitro. I also proposed to determine the TRPM7-mediated molecular mechanisms of brain injury in an in vivo model of neonatal hypoxia-ischemia and test a novel specific TRPM7 blocker for therapeutic potential.

I hypothesized that TRPM7 activity mediates neurite outgrowth under normal conditions via actin cytoskeleton regulation, and that hypoxia causes neurite retraction through potentiation of TRPM7 activity in vitro. I also hypothesized that inhibition of TRPM7 activity preserves behavioural outcomes and reduces brain injury through cytoskeletal regulation in a mouse model of neonatal hypoxic-ischemic brain injury.

I proposed to test this hypothesis by carrying out the following aims.

2.1 Aim 1: To determine the role of TRPM7 activity in neurite outgrowth of hippocampal neurons in vitro

Hypothesis: TRPM7 activity regulates neurite outgrowth of cultured hippocampal neurons through regulation of actin cytoskeleton.

42 43

I proposed to test this hypothesis by answering the following questions:

Q1: What is the role of TRPM7 activity in neurite outgrowth during development in vitro?

Q2: What is the TRPM7-dependent molecular mechanism of actin regulation in cultured neurons in vitro?

2.2 Aim 2: To characterize the effect of hypoxia on TRPM7- mediated neurite outgrowth of cultured neurons in vitro.

Hypothesis: Hypoxia induces neurite retraction through potentiating TRPM7 activity in vitro.

I proposed to test this hypothesis by answering the following questions:

Q1: How does hypoxia regulate TRPM7 activity in vitro?

Q2: How does hypoxia affect TRPM7-mediated neurite outgrowth in vitro?

2.3 Aim 3: To determine the effects of TRPM7 block with waixenicin A on brain injury and behavioural outcomes in vivo using a neonatal hypoxia-ischemia model.

Hypothesis: Inhibition of TRPM7 activity with waixenicin A reduces brain injury and preserves behavioural outcomes following hypoxia-ischemia in vivo.

I proposed to test this hypothesis by answering the following questions:

Q1: What are short and long-term effects of hypoxic-ischemic injury on the developing brain?

Q2: What are the effects of waixenicin A on behavioural outcomes following HI-induced brain injury?

2.4 Aim 4. To determine TRPM7-dependent molecular mechanisms in vivo using a neonatal hypoxic-ischemic injury model.

Hypothesis: TRPM7 activity mediates neuronal injury in vivo via calcium-dependent cytoskeletal regulation.

I proposed to test this hypothesis by answering the following questions:

Q1: What are the proteomic changes that take place after HI injury to the developing brain?

Q2: What is the potential molecular mechanism mediated by TRPM7 during neonatal HI brain injury?

Chapter 3 Matrials and Methods 3 Materials and methods 3.1 Cell culture

3.1.1 Dissociated culture of primary embryonic hippocampal neurons

All procedures were performed in accordance with animal welfare guidelines at the University of Toronto and were approved by the institutional animal care and use committee. Embryonic hippocampal cultures were prepared from E16.5 CD1 mice as described previously (Fath et al., 2009). Dissected hippocampi were digested with 0.025% Trypsin/EDTA at 37°C for 15 min. Cell density was determined using an Improved Neubauer hemocytometer and 1.0x104 cells were plated on poly-D-lysine (Sigma)-coated glass coverslips (12mm #1 German Glass, Bellco cat #1943-10012). The cells were kept in 5% CO2 at 37°C in culture medium (Table 2).

Table 2: Media/Solution Composition for Primary Hippocampal Neuronal Culture Solution Ingredients Concentration

Coating solution Poly-D-Lysine in sterile H2O 0.1 mg/mL Culture Medium Neurobasal Medium (Neuronal B-27 supplement 1.8% (vol/vol) cultures) GlutaMAX supplement 0.25% (vol/vol) Penicillin-Streptomycin 1% (vol/vol) Culture Medium Neurobasal Medium (Co-cultures) B-27 supplement 1.8% (vol/vol) GlutaMAX supplement 0.25% (vol/vol) Penicillin-Streptomycin 1% (vol/vol) Fetal Bovine Serum 10% (vol/vol)

3.1.2 Drug treatment of hippocampal neurons

Waixenicin A was obtained through collaboration with Dr. Andrea Fleig (University of Hawaii Cancer Center) and Dr. F. David Horgen (Hawaii Pacific University). Detailed extraction methods are described in Appendix 1.

The stock of waixenicin A was prepared by dissolving the compound in 100% methanol and adding PBS to dilute methanol to 2.5% concentration. Waixenicin A was added to cell cultures on day in vitro (DIV) 2, 24 hours post-plating. Naltriben mesylate (Tocris, #0892) was dissolved in DMSO and added to cultures on DIV4. Control neurons were either left untreated or treated with appropriate vehicle at the concentration shown below.

Table 3: Drug concentrations for treatment of hippocampal cultures.

Compound Final concentration in culture

Waixenicin A (drug) 10 nM, 30 nM, 50 nM, 100 nM, 300 nM, 500 nM, 800 nM

Methanol (vehicle) 0.0025% (vol/vol)

Naltiben mesylate (drug) 500 nM, 1 μM, 10 μM, 25 μM, 50 μM

DMSO (vehicle) 0.001% (vol/vol)

3.1.3 Hypoxic treatment of hippocampal neurons

Neuronal cultures and neuron-astrocyte co-cultures were grown under normal conditions (37C,

5% CO2) until DIV4. On DIV4, the cultures were transferred into an airtight Hypoxia Incubator

Chamber, which was flused with hypoxic gas (5% O2) for 5 minutes as described previously (Glass et al., 2002). The Hypoxia Incubator Chamber was then transferred into an incubator (37C) and kept there for up to 10 hours. The chamber was flushed with hypoxic gas every 2 hours. Upon completion of hypoxic treatment, cells were either immediately fixed with 4% PFA and prepared for immunolabelling, or transferred to an incubator with normal conditions.

3.1.4 Flag-murine TRPM7/pCDNA4 HEK-293 cells

HEK- 293 cells with stable expression of Flag-murine TRPM7/ pCDNA4 (gift from Dr. Sharenberg, Seattle, WA) were cultured in 10 cm2 tissue culture dish and grown in growth medium until 85% confluency as described previously (Chen et al., 2015b). 1-2 to two days prior to the tetracycline induction, Flag-TRPM7 HEK293 cells were plated on 12mm poly-D-lysine- coated coverslips at 20% confluency with growth media, and incubated at 37°C with 5% CO2.

TRPM7 expression was induced by adding 1 μg/mL tetracycline to the growth medium for 20 hours.

Table 4: Media/Solution Composition for HEK-293 cell culture Solution Ingredients Concentration

Coating solution Poly-D-Lysine in sterile H2O 0.1 mg/mL Growth Medium Minimum Essential Medium (MEM) Fetal Bovine Serum 10% (vol/vol) GlutaMAX supplement 1% (vol/vol) Zeocin 0.2% (vol/vol) Blasticidin 0.1% (vol/vol) Penicillin-Streptomycin 1% (vol/vol) Induction Tetracycline 1 mg/mL

3.2 Functional recordings

3.2.1 Calcium imaging

Intracellular Ca2+ concentration was measured using a Fura-2 ratiometric Ca2+ imaging system as described previously (Chen et al., 2010b). Neurons were pre-loaded with Fura-2 AM (5 μM) in dark for 40 min at room temperature. Fura-2 calcium signal was acquired at alternate excitation wavelengthes of 340 and 380 nm by a Deltaram V single monochromator controlled by EasyRatioPro (PTI), while neurons were perfused with control solution containing (mM) 129

NaCl, 2 CaCl2, 1 MgCl2, 25 HEPES, 30 glucose, and 5 KCl (with pH of 7.3-7.4 and osmolality ranging from 320-330 mOsm, containing 500 nM tetrodotoxin (TTX), 25 μM D-(−)-2-amino-5- phos-phonopentanoate (APV, Abcam), 40 μM 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX, 2+ Tocris), 5 μM nimodipine (Sigma)), or low Mg /Ca2+ solution (0 MgCl2, 0.5 CaCl2) with or without 500 nM waixenicin A or 50 μM naltriben mesylate. Signals were digitized by an intensified charged-coupled device (ICCD) camera (PTI) and fluorescence intensity (Poenie- Tsien) ratios of images were calculated by using EasyRatioPro.

3.2.2 Electrophysiological recordings

For methods on electrophysiological recordgings refer to Appendix 2.

3.3 Immunolabelling

3.3.1 Immunocytochemistry

Immunocytochemistry was performed as described previously (Nejatbakhsh et al., 2011). Neurons were fixed with pre-warmed 4% paraformaldehyde/4% sucrose in PBS for 15 min, permeabilized with 0.1% Triton X-100 in PBS for 10 min, and blocked with a blocker solution (2% bovine serum albumin, 2% fetal bovine serum, and 0.2% fish gelatin in PBS) for 1 hour at room temperature. Neurons were then incubated the primary antibodies for 2 hours at room temperature or at 4C overnight. Next, neurons were incubated with a secondary antibody for 1 hour at room temperature. Following incubation, were washed with 1xPBS 3 times and mounted on slides with ProLong Gold Antifade reagent.

3.3.2 Immunohistochemistry

3.3.2.1 Brain slice preparation

Brains were collected from pups in all treatment groups 3 days after HI at P10 and fixed in 4% paraformaldehyde/ 30% sucrose at 4ºC overnight. Brains were then embedded into cryomatrix embedding resin (#6769006, Thermo Fisher Scientific, Ottawa, ON, Canada), sectioned coronally into 80 μm slices with Leica CM3050 S cryostat (Leica, Concord, ON, Canada) and placed in 4% paraformaldehyde/ 30% sucrose at 4ºC overnight. Slices were then washed with 1xPBS for 15 minutes and probed with primary antibodies at 4C overnight. Subsequently, slices were washed with 1xPBS three times and probed with secondary antibodies for 1 hour at room temperature, washed with 1xPBS and mounted on glass coverslips with ProLong Gold antifade reagent (P36930, Thermo Fisher Scientific, Ottawa, ON, Canada).

3.3.2.2 Confocal microscopy of brain slices

Slices were imaged with Zeiss LSM 700 confocal laser scanning microscope (Zeiss, Germany). Three coronal slices per brain and three brains per treatment group were used. Quantitative analysis was performed by counting cells in 5-7 fields with 20x objective in the ipsilateral (penumbra) and contralateral hemispheres.

3.3.3 Antibodies used for immunolabelling

The antibodies in a table below were used for immunolabelling.

Table 5: Antibodies used for immunolabelling.

Antibody Dilution Primary/Secondary Cells/Tissue Vendor

B-tubulin, 1:1000 Primary Cells Sigma, T-0198 mouse

TRPM7, goat 1:200 Primary Cells Abcam, Ab729

Α-actinin-1, 1:500 Primary Cells Abcam, Ab18061 mouse

Tau, mouse 1:500 Primary Cells Millipore, MAB3420

MAP2, 1:500 Primary Cells Millipore, chicken AB15452

NeuN, mouse 1:200 Primary Tissue Millipore, MAB377

GFAP, rat 1:1000 Primary Tissue Abcam, Ab7260

Caspase-3, 1:500 Primary Tissue Cell Signalling rabbit Technology, #9661

TRPM7, 1:500 Primary Tissue Abcam, Ab85016 mouse

Alexa Fluor 1:500 Secondary Cells/Tissue Thermo Fisher, 488, mouse A11001

Alexa Fluor 1:500 Secondary Tissue Thermo Fisher, 405, rabbit A31556

Alexa Fluor 1:500 Secondary Tissue/Cells Thermo Fisher, 568, rat and A11077 chicken

3.4 Confocal microscopy and image analysis

3.4.1 Image acquisition for neurite length analysis

Images of neurons were captured with either a Leica TCS LS confocal laser-scanning microscope (Heidelberg, Germany; Leica confocal software, version 2.5; build 1347; Leica Microsystems) with either 40x (NA 1.25) or 63x (NA 1.32) oil-immersion lenses, and 488, 543 and 633 nm lasers, or a Carl Zeiss Confocal Laser Scanning Microscope LSM700 with either 63x DIC (NA 1.40) or 40x DIC (NA 1.3) oil immersion lenses, and 405, 488, and 543 nm lasers (Sun et al., 2009). Each Z-plane was 0.3 μm. All cells were imaged using the same magnification and laser settings. All cells were imaged at a resolution of 1024x1024 pixels using the same magnification and laser settings.

3.4.2 Neurite length analysis

Neurite lengths were analyzed with SynD, a semi-automated image analysis routine (Schmitz et al., 2011). For neurons (DIV>6) that require a larger field of view to capture a single neuron, multiple images were taken under the 40x lens first and mosaics of overlapping images were assembled in ImageJ (NIH, http://rsb.info.nih.gov/ij) using MosaicJ plugin before neurite length analysis.

3.4.3 Protein Colocalization Analysis Confocal images of cells that were triple-stained with antibodies targeting the proteins of interest were scanned at a resolution of 2048 x 2048 pixels using a 63x (NA 1.40) oil-immersion lens (Zeiss LSM700). Each fluorescence signal was imaged separately with corresponding channel, without pixel saturation. Protein colocalization was estimated by quantifying the degree of overlap between the fluorescence intensities in the each pixel located in the region of interest (ROI). For each image, an ROI was selected in the cell body, neurite, growth cone, and a background area (outside of the cell) to be used as the control. A Pearson’s correlation coefficient (Rr) was measured using an ImageJ plugin called the intensity correlation analysis, as described previously (Nejatbakhsh et al., 2011). The Rr value of 1.0 indicates close colocalization, and 0 indicates low probability for colocalization.

3.5 Identification of protein-protein interactions

3.5.1 Co-immunoprecipitation Protein isolated from MDA-MB-231 cells (250 µg) (Sigma-Aldritch) in RIPA lysis buffer (Santa Cruz Biotechnology, sc-24948) was incubated with 2 µg of anti-TRPM7 antibody (Abcam, ab85016) for 90 min at 4ºC and 10 min at room temperature (RT). Then 20 µl of pre-washed Protein A/G PLUS-Agarose beads (Santa Cruz Biotechnology, sc-2003) were added, and the mixture was incubated and shaking at 4ºC overnight. The protein mixture was vortexed, heated from 5 min and centrifuged for 10 min at 3000 rpm. The supernatant was collected and run SDS- PAGE.

3.5.2 Protein identification by mass spectrometry

LC-MS/MS and protein identification were performed as described previously (Silverman- Gavrila et al., 2009). In-gel trypsin digestion of proteins was performed using an In-gel Tryptic Digestion Kit (Pierce, Thermo Fisher Scientific Inc, Rockford, IL, USA). Peptides were sent to the Ontario Cancer Biomarker Network Facility Centre for protein identification by nano LC/MS/MS HPLC chromatographic separation. Mascot Generic and Mascot Daemon were used to combine and analyze data. After the MS/MS Ions Search against UniProt Universal Protein Resources knowledge base (http://www.uniprot.org/) a protein was accepted as identified if the total Mascot score was greater than the significance threshold.

3.6 Mouse model of neonatal hypoxia-ischemia

3.6.1 Animals

All procedures were performed in accordance with animal welfare guidelines at the University of Toronto and were approved by the institutional animal care and use committee. Timed pregnant CD1 mice were obtained from Charles River laboratories (Sherbrooke, Quebec, Canada) and housed at an ambient temperature of 20 ± 1°C and a 12-hour light/dark cycle with food and water fed ad libitum. Postnatal day 7 (P7) pups were used in all experiments. All procedures and protocols were carried out in accordance with Canadian Council on Animal Care guidelines and approved by University of Toronto Animal Care Committee.

3.6.2 Neonatal hypoxic-ischemic injury model

A modified method of Rice-Vannucci procedure (Rice, III et al., 1981) of neonatal hypoxic- ischemic (HI) brain injury was carried out as described previously (Huang et al., 2017c; Xiao et al., 2015; Sun et al., 2015; Xu et al., 2015; Huang et al., 2017b; Chen et al., 2015a). Briefly, P7 pups of either gender were anesthetized with isoflurane (3.0% for induction and 1.5% for maintenance). The right common carotid artery (CCA) was exposed and ligated using bipolar electrical coagulation (Vetroson). The incision was closed with tissue adhesive (3M Vetbond). Mice were then returned to their dam and allowed to recover for 90 minutes. Subsequently, pups were placed in an airtight, transparent chamber (A-Chamber A-15274 with ProOx 110 Oxygen Controller/E-720 Sensor, Biospherix, NY, USA) perfused with humidified gas mixture containing 7.5% oxygen balanced with 92.5% nitrogen at 37°C for 60 minutes and returned to the dam afterwards. Chamber temperature was monitored using a homoeothermic blanket control unit (K-017484 Harvard Apparatus, Massachusetts, USA). Sham controls were subjected to anesthesia and the common carotid artery was exposed without ligation and subsequent exposure to hypoxia.

3.6.3 Drug administration

Waixenicin A: P7 pups of either gender were randomly assigned to either vehicle or waixenicin A groups. The compound was administered by intraperitoneal injection as a single dose of 37 ng/g body weight in 100 μl of saline solution with 0.0037% methanol. For sham and vehicle controls an equal volume of vehicle solution (saline with 0.0037% methanol) was administered in an identical manner.

Naltriben: Naltriben mesylate (Tocris, #0892) was administered before ischemia by interaperitoneal injection as a single dose of 42 μg/g of body weight in 100 μl of saline solution (0.004% DMSO). For sham and vehicle controls an equal volume of saline (0.004% DMSO) was administered in an identical manner.

3.7 Histological assessments

Histological assessments were carried out as described previously (Huang et al., 2017c; Xiao et al., 2015; Sun et al., 2015; Xu et al., 2015; Huang et al., 2017b; Chen et al., 2015a).

3.7.1 Infarct volume measurement

Twenty-four hours after HI injury, whole brains were removed from P8 pups and sectioned coronally into approximately 1mm slices. Slices were stained with 1% 2,3,5-triphenyl-2H- tetrazolium chloride (TTC) in saline and placed in a dark incubator maintained at 37C for 20 minutes. The corrected infarct volumes were calculated as follows: corrected infarct volume (%) = ((contralateral hemisphere volume – ipsilateral hemisphere + infarct volume) /contralateral hemisphere volume) x 100%.

3.7.2 Nissl staining

Seven days after HI injury, whole brains were removed, fixed, imaged, sectioned coronally into approximately 100 µm slices and stained with Nissl (0.1% cresyl violet). Thirty two days after HI injury, whole brains were removed, fixed and imaged and hemispheric liquefaction volumes were calculated. The hemispheric liquefaction volumes and infarct volumes were measured using the ImageJ software (National institute of Health, Bethesda, MD, USA).

3.8 Behavioural assessments

3.8.1 Short-term neurobehavioural assessments

Short-term neurobehavioural assessments were carried out as described previously (Huang et al., 2017c; Xiao et al., 2015; Sun et al., 2015; Xu et al., 2015; Huang et al., 2017b; Chen et al., 2015a).

Pups in all treatment groups were subjected to the following highly reproducible strain- and gender-independent neurobehavioural assessments 1,3 and 7 days after HI: 1) righting reflex, where animals are placed in supine position and the time needed to get to prone position is recorded; 2) cliff avoidance reflex, where animals were placed with forepaws over the edge and the time required to turn at least 90º was recorded; 3) geotaxis reflex, where animals were placed on an inclined board (45º) facing downwards and the time required to turn around is recorded; 4) grip strength test, where animals were allowed to hang on a horizontal wire by forelimbs and the time to release the wire was recorded. All tests were repeated for a total of three trials, and average scores were used for analysis (Feather-Schussler and Ferguson, 2016c).

3.8.2 Accelerated Rotarod Test

Four weeks after the HI procedure, mice in all three groups underwent Accelerated Rotarod Test (LE8205, Panlab, Harvard Apparatus, Barcelona, Spain) to test for balance and motor function. The speed rotation of the drum accelerated from 4 to 40 rpm within 120 sec, and the latency and velocity at which each mouse fell off the drum was recorded. The test was repeated for three consecutive trials and the results were averaged.

3.8.3 Passive avoidance test

Four weeks after the HI procedure, mice in all three groups underwent Passive Avoidance Test to test for the ability to inhibit innate preference for a dark confined space after a single association with an aversive stimulus, as described previously (Haelewyn et al., 2007). The training protocol consisted of three sessions performed on separate days: habituation, acquisition and retention. The apparatus consisted of a large (250(W) x 250(D) x 240 (H) mm) illuminated compartment and a small (195 (W) x 108 (D) x 120 (H) mm) dark compartment with electrified grid floor separated by a guillotine gate (LE872, Panlab, Harvard Apparatus, Barcelona, Spain). During habituation session the mouse was placed into the illuminated compartment and allowed to explore for 1 minute. After 1 minute, the door to the dark compartment was opened, latency to enter the dark room was recorded (step-through maximal latency: 300 sec) and the door was closed for 30 sec before the mouse was returned to its home cage. During acquisition session, the protocol was the same as habituation session, but the mouse received an inescapable foot shock (0.4 mA, 2 sec) upon entering the dark compartment. The retention session took place 48 hours after acquisition session. The mouse was placed into the illuminated compartment, the door to the dark compartment was opened after 5 sec and latency to enter the dark compartment was recorded (step-through maximal latency: 300 sec). The latency to enter dark compartment during the retention session was taken as an index of memory performance.

3.8.4 Novel object recognition

Four weeks after HI procedure, the mice in all three groups underwent Novel Object Recognition test to evaluate spontaneous exploratory behaviour triggered by novelty, as described previously (Leger et al., 2013; Sik et al., 2003). The objects used for this test were similar in size and odour, but different in shape and texture. The test was carried out in two sessions separated by an intersession interval of 4 hours. During familiarization session, the mouse was placed in an open-

55 field arena (25 cm x 25 cm x 24 cm) facing away from two identical objects placed 5 cm apart and was allowed to explore both objects freely for 3 minutes of session duration. During the test session, one of the familiar objects was replaced by an identical copy to avoid olfactory cues, and another familiar object was replaced by a novel object. The mouse was placed in an open-field arena facing away from the objects and allowed to explore the objects freely for 3 min of session duration. The exploration time of each object was recorded and preference for novel object was calculated as a percentage of total exploration time.

3.9 Proteomic analysis

Sample preparation, database searching, protein identification and quantitative data analysis were performed by SickKids SPARC (Proteomics, Analytics, Robotics and Chemical Biology) BioCentre. Detailed methods are described in Appendix 3.

3.9.1 Visualization of proteomic analysis

Protein network analysis was performed using STRING 10.5 (https://string-db.org/). Heatmap of the proteins was created using the heatmap3 package in RStudio.

3.10 Western blotting

The protein was collected from dissociated mouse hippocampal neuronal cells, MDA-MB-213 cells or ipsilateral hemispheres of pups 6 or 24 hours after HI in RIPA buffer (Santa Cruz Biotechnology, sc-24948) with protease inhibitor mixture (Santa Cruz Biotechnology, sc-29131). The protein samples (20 μg/well) were separated on a 10% SDS-PAGE gel and proteins were then transferred to a nitrocellulose membrane (200 mA per gel, 60 min). Membranes were incubated with specific primary antibodies summarized in Table 4 at 4ºC overnight. Horseradish peroxidase–conjugated anti-mouse (1:7500; Chemicon AP130P), anti-rabbit (1:7500; Chemicon AP132S) and anti-goat (1:7500; Chemicon AP180P) IgG antibodies were used as secondary antibodies and were detected with the ECL system (PerkinElmer, Inc, USA). Images were analyzed using an image-analysis system (NIH Image J 1.47v).

Table 6: Antibodies used for Western blot.

Antibody Dilution Cells/Tissue Vendor

Α-actinin-1 1:1000, 1:2000 Cells, tissue Abcam, Ab18061

GAPDH 1:7500, 1:10000 Cells, tissue Cell Signalling Technology, 2118S

CaMKII, p-CaMKII 1:1000, 1:3000 Tissue Santa Cruz Biotechnology, sc- 5306, sc-12886

Calcineurin B 1:3000 Tissue Abcam, Ab154650

Calmodulin 1:2000 Tissue Abcam, Ab107977

P38, p-p38 1:1000, 1:2000 Tissue Cell Signalling Technology, 9212S, 4511S

Cleaved caspase-3 1:1000 Tissue Cell Signaling Technology, 9664

Cleaved caspase-9 1:1000 Tissue Cell Signaling Technology, 9504S

Bax 1:1000 Tissue Cell Signaling Technology, 2772

Bcl-2 1:1000 Tissue Cell Signaling Technology, 2870

Cofilin, p-cofilin 1:1000, 1:2000 Tissue Abcam, Ab134963, Ab12866

Β-actin 1:3000 Tissue Sigma

Flag 1:10000 Cells Sigma

3.11 Statistical Analysis

Data are presented as the mean ± SEM. Statistical analysis was performed using GraphPad Prism (GraphPad Software, San Diego, CA). The multiple comparisons of means between experimental and control conditions were performed with Bonferroni post hoc test following the one-way ANOVA. Values of p < 0.05 are taken as statistically significant.

Chapter 4 Results 4 TRPM7 regulates axonal outgrowth and maturation of primary hippocampal neurons.

Contributions

Part of this work was completed in collaboration with Christine YouJin Bae, Marielle Deurloo, Dr. Wenliang Chen, Dr. Andrew Barszczyk, Dr. David F. Horgen, Dr. Andrea Fleig, Dr. Zhong- Ping Feng and Dr. Hong-Shuo Sun. I hold first authorship on this manuscript, and I performed all of the cell culture studies and analysis, including neurite outgrowth studies and colocalization studies, and wrote the manuscript. Christine YouJin Bae performed TRPM7 shRNA knockdown experiments (not included in this thesis) and assisted with mass spectrometry and co- immunoprecipitation experiments. Marielle Deurloo and I performed calcium imaging experiments and analysis. Dr. Wenliang Chen performed electrophysiology experiments and analysis. Dr. Andrew Barszczyk contributed to the manuscript preparation. Drs. David F. Horgen and Andrea Fleig supplied waixenicin A. Drs. Zhong-Ping Feng and Hong-Shuo Sun contributed to study design and manuscript preparation.

Sections 4.1 and 4.3 of this chapter contain a re-creation of the published material with the following citation:

Turlova et al., TRPM7 regulates axonal outgrowth and maturation of primary hippocampal neurons. Mol. Neurobiol. 2016. 53(1): 595-610.

Publication was recreated with permission and copyright license.

Section 4.2 was completed in collaboration with Raymond Wong (PhD student in Sun lab). Raymond Wong performed electrophysiological recordings. I performed cell culture experiments and calcium imaging experiments.

4.1 TRPM7 block with waixenicin A enhances neurite outgrowth in developing hippocampal neurons

4.1.1 TRPM7 protein is expressed in neurites and growth cones of developing hippocampal neurons

Earlier studies have shown that TRPM7 protein is extensively expressed in hippocampal neurons in both rat brain slices (Sun et al., 2009) and rat or mouse primary cell cultures (Wei et al., 2007). Here we first investigated the distribution of TRPM7 in the growth cone region of the cultured hippocampal neurons dissociated from E16 embryonic CD1 mice. Confocal immunofluorescence imaging revealed that TRPM7 channels are present in growth cones of DIV2 hippocampal neurons (Figure 6B), in addition to the somata and processes (Figure 6A).

To further characterize the distribution of TRPM7 relative to cytoskeletal structures, hippocampal neurons were triple-labeled with anti-TRPM7, anti-β-tubulin and rhodamine phalloidin for filamentous actin (F-actin). High magnification confocal images showed the relation between the spatial distribution of TRPM7 to two cytoskeletal structures, β-tubulin and F-actin (Figure 6A). In the central microtubule-containing domain of the growth cone, TRPM7 expression was low, as evident by the lower fluorescent intensity. In the peripheral domain containing filopodia and lamellipodia, TRPM7 channels are highly expressed along the actin bundles (Figure 6B-C). Further quantification of TRPM7 fluorescence intensity showed higher expression of TRPM7 in the periphery of the growth cone, where filopodia are located (Figure 6C-E). As actin dynamics determine growth cone motility and are necessary for directed axonal outgrowth (Bentley and Toroian-Raymond, 1986), our results suggest that TRPM7 may mediate axonal outgrowth through actin regulation.

Figure 6: TRPM7 expression pattern in DIV2 hippocampal neurons. A) DIV2 hippocampal neuron triple-stained with antibodies recognizing α-tubulin (green) and TRPM7 (blue) and rhodamine phalloidin for F-actin (red). 2-image and 3-image overlays were included to show the relative distribution of TRPM7 with respect to cytoskeletal structures. B) and C) Distribution of TRPM7 and cytoskeletal proteins in two separate growth cones. D) Schematic of the domains within the growth cone. E) Ratio of TRPM7 intensities in central and peripheral domains of the growth cone to TRPM7 intensity in the neurite; N=5 neurons, n=9 growth cones. Scale bars are 20 μm.

4.1.2 Waixeinicin A reduces TRPM7-like current in hippocampal neurons in a dose-dependent manner

Waixenicin A is a newly identified specific TRPM7 inhibitor and blocks the activity of either recombinant channel or native channel (Zierler et al., 2011). We investigated whether waixenicin A blocks ion conductance of TRPM7 and affects axonal outgrowth in hippocampal culture. Our whole-cell patch-clamp recording showed that waixenicin A decreased the TRPM7-like current in DIV3-DIV7 hippocampal neurons in a dose-dependent manner (Figure 7A and B), with an

IC50 value of 362.7±1.5 nM (Figure 7C). The TRPM7-like current was recorded with a ramp protocol in the presence of tetrodotoxin (TTX), D-(-)-2-amino-5-phos-phonopentanoate (APV), 6-cyano-7-nitroquinoxaline-2,3-dione (CNQX) and nimodipine in the bath solution to block the activation of voltage-gated Na+, NMDA, non-NMDA glutamate-activated and L-type Ca2+ voltage-gated channels, respectively (Sun et al., 2009).

Figure 7: Waixenicin A inhibits TRPM7 activity in a dose-dependent manner. A) Mean currents of TRPM7 in DIV3-DIV7 hippocampal neurons without or with different concentration waixenicin A (Waix A) (50nM, n=6; 125 nM, n=7; 250 nM, n=13; 500 nM, n=11; 1000 nM, n=8; 10000 nM, n=3) incubation. B) Mean outward current measured at 100 mV on control (0 nM Waix A, n=27) and different concentrations of waix A treatments (30 min incubation time on hippocampal neurons) (50nM, n=6; 125 nM, n=7; 250 nM, n=13; 500 nM, n=11; 1000 nM, n=8; 10000 nM, n=3), respectively (* vs. control, p<0.05, One-way ANOVA). C) Nolinear curve fitting analysis for the inhibition rate of Waix A on TRPM7 currents at 100 mV. It indicated that the IC50 of Waix A for TRPM7 current in hippocampal neurons is 362.7±1.5 nM.

The data in this figure was contributed by Dr. Wenliang Chen.

4.1.3 TRPM7 block by waixenicin A decreases calcium influx into hippocampal neurons

Lowering extracellular concentrations of divalent cations, such as Mg2+, potentiates Ca2+ influx through TRPM7 channels (Sun et al., 2009). To test if TRPM7 block by waixenicin A reduces the influx of calcium in hippocampal neurons, we carried out ratiometric fura-2 Ca2+ imaging at various extracellular Mg2+ concentrations. Fura-2 Ca2+ signals were monitored under the same Mg2+ -free and low Ca2+ condition in the absence or presence of 500 nM waixenicin A. In the presence of waixenicin A, the net increase of the Ca2+ signal was 0.03±0.004 (n=10, p<0.05), which was significantly lower than that recorded in the absence of waixenicin A (0.09±0.008, n=10, p<0.05) (Figure 8B, left panel). The decrease was 58%±8% (n=10) from control value (Figure 8B, right panel). The KCl-evoked Ca2+ signal was not significantly different between the two groups. Representative images are shown in Figure 8A. Two consecutive applications of Mg2+-free and low Ca2+ solution did not produce a desensitization of the calcium signal (Figure 8C). Our findings showed that waixenicin A suppressed the free Mg2+-induced Ca2+ influx through TRPM7 channels in hippocampal neurons.

Figure 8: Waixenicin A reduces calcium influx in hippocampal neurons. A) Representative images of control and waixenicin A- treated neurons. B) Representative 340/380 ratio trace of neurons perfused with Mg2+-free solution followed by Mg2+-free solution together with 500 nM waixenicin A (left panel); average peak 340/380 ratio values of TRPM7-induced calcium influx (right panel). EC Representative 340/380 ratio trace of two consecutive applications of Mg2+-free solution to hippocampal neurons (left panel) and average peak 340/380 ratio values of TRPM7-induced calcium influx (right panel). For waixenicin A- treated neurons n=10, control neurons n=19. Data is presented as mean±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc (*p<0.05).

62 63

4.1.4 Waixenicin A enhances neurite outgrowth of cultured hippocampal neurons.

We next examined whether pharmacological inhibition of TRPM7 activity was sufficient to affect neurite outgrowth. This approach allowed us to examine the morphological changes in the early stages in culture. Various concentrations of waixenicin A (10, 30, 50, 100, 300, 500 and 800 nM) and 0.0025% of methanol (vehicle control) were added to hippocampal culture on DIV2, 24 hours after plating. Neurons were stained for two major cytoskeletal components, β- tubulin and F-actin, 6 hours post-treatment to visualize the morphology of the cells (Figure 9A). Compared to control neurons, neurons treated with waixenicin A exhibited significant increases in total neurite length (Figure 9B: control: 70.4 ± 4μm, n = 81; methanol (0.0025%): 68.3 ± 3.7 μm, n = 78; waixenicin A 10 nM: 117 ± 6.3 μm, n = 62; waixenicin A 30 nM: 108 ± 6 μm, n = 76; waixenicin A 50 nM: 141.7 ± 5.4 μm, n = 84; waixenicin A 100 nM: 143.5 ± 9.9 μm, n = 98; waixenicin A 300 nM: 152.2 ± 7 μm, n = 84; waixenicin A 500 nM: 183 ± 8 μm, n = 95; waixenicin A 800 nM: 158 ± 8.5 μm, n = 74), average length of longest neurite (Figure 9C: control: 20.9 ± 1.2 μm, n = 81; methanol (0.0025%): 22.2 ± 1.4 μm, n = 78; waixenicin A 10 nM: 36.4 ± 2.4 μm, n = 62; waixenicin A 30 nM: 29.1 ± 1.8 μm, n = 76; waixenicin A 50 nM: 42.2 ± 2.4 μm, n = 84; waixenicin A 100 nM: 39.7 ± 1.9 μm, n = 98; waixenicin A 300 nM: 39.6 ± 2.2 μm, n = 84; waixenicin A 500 nM: 55 ± 2.8 μm, n = 95; waixenicin A 800 nM: 36.1 ± 2.2 μm, n = 74) and average number of neurites per neuron (Figure 9D: control: 6.2 ± 0.2, n = 81; methanol (0.0025%): 6 ± 0.2, n = 78; waixenicin A 10 nM: 8.3 ± 0.4, n = 62; waixenicin A 30 nM: 8.5 ± 0.4, n = 76; waixenicin A 50 nM: 8.4 ± 0.3, n = 84; waixenicin A 100 nM: 9.7 ± 0.5, n = 98; waixenicin A 300 nM: 10.5 ± 0.5, n = 84; waixenicin A 500 nM: 12 ± 0.6, n = 95; waixenicin A 800 nM: 9.4 ± 0.4, n = 74).

We found that TRPM7 block with waixenicin A increased neurite outgrowth both through enhanced length of neurites and the increased number of neurites, and 500 nM was the most effective dose.

Figure 9: Waixenicin A enhances neurite outgrowth through neurite length and number of neurites 6 hours after treatment. A) Representative images of DIV2 control, vehicle control neurons, and neurons treated with increasing doses of Waixenicin A stained for β-tubulin (green) and F-actin (red). Scale bars are 10 μm. B) Total neurite length, C) average length of longest neurite and D) average number of neurites per neuron of DIV2 neurons (control n =81, vehicle n=78, waixenicin A 10 nM n=62, 30 nM n=76, 50 nM n=84, 100 nM n=98, 300 nM n=84, 500 nM n=96, 800 nM n=74).

Moreover, neurons treated with 500 nM of waixenicin A 24 hours after plating exhibited substantial increases in outgrowth as compared to the untreated control or vehicle control neurons 24 and 96 hours post-treatment, on DIV3 and DIV6. Representative images of DIV3 and DIV6 neurons are presented in Figures 10A and B, respectively. The total neurite lengths per neuron enhanced significantly in waixenicin A group on both DIV3 (control: 479.2±13.6 μm, n=104; vehicle: 470.1±16.4 μm, n=99; waixenicin A: 624.8±18.6 μm, n=96; p < 0.01; Figure 10C) and DIV6 (Control: 1641.8±53.8 μm, n=87; vehicle: 1676.2±94.6 μm, n=71; waixenicin A: 2272.5±78.9 μm, n=86; p < 0.01; Figure 10D). We also found that total number of neurites was significantly higher in waixenicin A-treated groups at both time points (Figure 10E DIV3, control: 15 ± 0.8, n = 104; vehicle: 16 ± 0.5, n = 99; waixenicin A: 21 ± 1, n = 96; Figure 10F DIV6, control: 25 ± 2, n = 87; vehicle: 24 ± 1.3, n = 71; waixenicin A: 28.4 ± 1.4, n = 86).

These data demonstrated that blocking TRPM7 activity with waixenicin A enhanced neurite outgrowth at multiple time points in culture.

Figure 10: Waixenicin A enhances neurite outgrowth and number of neurites 24 and 96 hours after treatment. Representative images of hippocampal neurons on A) DIV3 and B) DIV6. C) Total neurite length of DIV3 neurons (control n=104, vehicle n=99, waixenicin A n=96). D) Total neurite length of DIV6 neurons (control n=87, vehicle n=71, waixenicin A n=76). For DIV3 and DIV6 neurons. Total number of neurites of E) DIV3 and F) DIV6 neurons. Waixenicin A dose was 500 nM. Data is shown as mean±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc. *p<0.05, **p<0.01, ***p<0.001. All scale bars are 20 μm.

4.1.5 Waixencin A preferentially enhances axonal growth of cultured hippocampal neurons

As the increase in total neurite outgrowth could be contributed by increases in axonal length, dendritic length or both, we next evaluated the effects of TRPM7 channel blockade on axons and dendrites. Neurons were stained for axonal (tau-1) and dendritic (MAP2) markers, and representative images of neurons on DIV4, DIV6 and DIV8 are shown in Figure 11A, B and C, respectively. At DIV4, neurons treated with the TRPM7 blocker showed a 71% increase in axonal length and 33% increase in dendritic length compared to controls. At DIV6, axonal and dendritic lengths increased by 48% and 25% respectively, while at DIV8 axonal length increased by 53%, and dendritic length increased only by 14% (Figure 11D shows axonal lengths and 11E shows dendritic lengths). Sholl analysis revealed a consistent increase in branching of axons at all times points while dendrites only showed greater branching on DIV4 and DIV6, but not DIV8. Sholl analysis of axons on DIV4 is shown in Figure 11F and dendrites in Figure 11G. Therefore, we concluded that under normal conditions TRPM7 is a potential negative regulator of axonal growth.

Figure 11: Waixenicin A treatment preferentially enhances axonal outgrowth and branching. A-C) Representative images of neurons in different treatment groups on DIV4, DIV6 and DIV8. D) Average axonal and E) dendritic lengths of DIV4 (control n=56, vehicle n=83, waixenicin A n=59), DIV6 (control n=50, vehicle n=46, waixenicin A n=49) and DIV8 (control n=50, vehicle n=49, waixenicin A n=32) neurons. F) Sholl analysis of axons of DIV8 neurons indicates increased branching and length of branches with Waixenicin A treatment (control n=50, vehicle n=49, waixenicin A n=32). G) Sholl analysis of dendrites of DIV8 neurons showed increased number of branches only (n=31). Data is shown as mean±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc. **p<0.01, ***p<0.001.

4.1.6 Waixenicin A accelerates progression of hippocampal neurons through developmental stages

Dotti et al. described that hippocampal neurons go through five stages of development in culture (Dotti et al., 1988). Here, we show that our E16 hippocampal cultures closely follow these developmental stages (Figure 12A). At stage 1, neurons protrude veil-like lamellipodia that explore extracellular environment within ~6 hours in culture. The second stage is characterized by appearance of several neurites of similar length (~24 hours in culture). Around 36-48 hours in culture, neurons move into the third stage, also known as neuronal polarization. At this stage, one of the neurites begins elongating faster than the rest. Molecular marker expression of that neuron also changes, as axon starts to preferentially express tau-1, and dendrites to express MAP2 (Figure 12B). Stage 4 is characterized by increased branching and development of axons and dendrites (>4 days in culture), and neurons at stage 5 (>14 days in culture) show further maturation of axonal and dendritic arbors and formation of synapses. We outlined the neuronal transition through these stages in Figure 12A. We focused on the transition between stage 2 and stage 3, as it is crucial for breaking neuronal symmetry and progression towards functionality.

We asked whether TRPM7 channel activity is required for neuronal development. Since the developmental transitional stages take place within the first three days in culture, we examined the effects of waixenicin A on the developmental stages from the neurons in DIV2 and DIV3 cultures. As shown in Figure 12C, blocking TRPM7 activity by waixenicin A shifted the developmental transitional stage towards an early time in culture. At DIV2, untreated and vehicle-treated neuron cultures had 14% and 15% of neurons in stage 3 (axon distinguished by ICC with anti-tau-1 antibody; dendrites with anti-MAP2 antibody), while neurons treated with 500 nM waixenicin A had 30% of neurons in stage 3 (Figure 12C, left). In DIV3 cultures, untreated and vehicle-treated groups showed that 32% and 31% of neurons progressed into stage 3, while neurons treated with waixenicin A had 54% neurons in stage 3 (Figure 12C, right).

Figure 12. Treatment with Waixenicin A promotes maturation of hippocampal neurons. A) Representative images of developmental stages of hippocampal neurons according to Dotti et al. Top panel: stage 1 (left), stage 2 (center) and stage 3 (right). Bottom panel: stage 4 (left) and stage 5 (center and right). B) Representative images of neurons in stage 2 (top panel) and stage 3 (bottom panel) in different treatment groups. Scale bars are 20 μm. C) Proportion of neurons in stage 2 and 3 in different treatment groups. Left panel: DIV2 cultures (6 hours post-treatment; n=137); right panel: DIV3 cultures (24 hours post treatment; n=144). Chi-square: DIV2 untreated control vs. Waixenicin A P=0.006, DIV3 untreated control vs. Waixenicin A P=0.04.

We have also addressed the progression of neurons from stage 2 into stage 3 quantitatively in a different population of neurons. As stage 3, or neuronal polarization, starts when one of the neurites begins to elongate faster than others, we calculated the difference between the length of the longest neurite and the average length of the remaining neurites (Figure 13A). We found an overall increase in length difference in both DIV2 and DIV3 hippocampal neurons treated with waixenicin A, when compared to control groups, suggesting an increase in the rate of axonal growth, an increase in the number of stage 3 neurons or both (Figure 13B). Upon further investigation, we found that the length difference of neurons that were morphologically defined as stage 2 was significantly lower than that of neurons morphologically defined as stage 3, yet remained consistent between different groups (Figure 13C, DIV2 stage 2: untreated control 0.44±0.02; vehicle control 0.5±0.02; waixenicin A 0.05±0.02; DIV3 stage 2: untreated control 0.52±0.02; vehicle control 0.54±0.02; waixenicin A 0.58±0.02; DIV2 stage 3: untreated control 0.77±0.02; vehicle control 0.74±0.03; waixenicin A 0.84±0.02; DIV3 stage 3: untreated control 0.81±0.02; vehicle control 0.86±0.02; waixenicin A 0.87±0.02), suggesting that there are more stage 3 neurons in waixenicin A-treated group, while the rate of axonal elongation does not change between the groups. Based on these data, we defined a neuron to be in stage 3 if the difference between the length of its longest neurite and the average length of the remaining neurites is greater or equal to 0.67. Based on this definition, we have found that untreated control, vehicle control and waixenicin A-treated neurons had 15%, 15% and 33% of neurons in stage 3 at DIV2 and 35%, 32% and 53% at DIV3, respectively, and that these data correspond well with our morphological data (Figure 13D). These findings suggest that TRPM7 plays a role in inhibiting premature transition between the developmental stages under normal conditions, and thus regulates neuronal maturation.

Figure 13: Quantitative analysis of neuronal transition between second and third developmental stages. A) Formula for calculating the difference between the length of longest neurite and the average length of the remaining neurites. B) Average length difference of DIV2 and DIV3 neurons in different treatment groups. C) Average length difference of stage 2 and stage 3 neurons in different treatment groups at DIV2 and DIV3. Data is presented as mean±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc (*p<0.05; ***p<0.001). D) Distribution of neurons in stage 2 and stage 3 in different treatment groups at DIV2 and DIV3, based on the assumption that the length difference for stage 3 is ≥ 0.67. Chi-square: DIV2 untreated control vs. waixenicin A P=0.0015; DIV3 untreated control vs. waixenicin A P= 0.0046. For groups at DIV2: untreated control n=73; vehicle control n=68; waixenicin A n=69. For groups at DIV3: untreated control n=63; vehicle control n=62; waixenicin A n=62.

4.2 TRPM7 activation by naltriben mesylate induces neurite retraction in developing hippocampal neurons

Next, to confirm the involvement of TRPM7 activity in axonal outgrowth, we used a specific TRPM7 activator, naltriben mesylate (Hofmann et al., 2014). We observed the effects of TRPM7 activation by naltriben on TRPM7 currents, calcium dynamics and axonal and dendritic outgrowth.

4.2.1 Naltriben activates TRPM7 currents in HEK-293 cells and primary hippocampal neurons

To confirm the stimulatory effect of naltriben on TRPM7 current, we first carried out whole-cell patch-clamp recording on HEK-293 cells overexpressing recombinant TRPM7 channels. Tetracycline was used to induce TRPM7 overexpression in HEK-293 cells. Confirmation of TRPM7 expression in HEK-293 cells is described in Appendix 4. Figure 14B shows that the TRPM7 currents in HEK-293 cells treated with tetracycline were large and outwardly rectifying. The TRPM7 inhibitor, waixenicin A (500 nM), significantly reduced TRPM7 current density at +100 mV by ~60% (from 30 ± 4 pA/pF to 13 ± 3 pA/pF) in the HEK-293 cells induced with tetracycline (Figure 14B, n=8, p<0.01). On the contrary, the TRPM7 agonist, naltriben (10 μM), potentiated the TRPM7 current density by ~70% (from 31 ± 4 pA/pF to 51 ± 7 pA/pF; Figure 14D, n=7, p<0.001), and was also sensitive to block by waixenicin A (the naltriben-potentiated current was reduced to 19 ± 2 pA/pF; Figure 14D, n=7, p<0.0001). In HEK-293 cells without induction by tetracycline, neither waixenicin A (Figure 14A, n=4) nor naltriben (Figure 14C, n=5) had any effect on the endogenous current.

We next tested whether naltriben can activate TRPM7-like currents in DIV3-7 primary mouse hippocampal neurons, and if so, whether waixenicin A can reduce this current. As shown in Figure 14E (middle), the current density of the basal outwardly rectifying current at +100 mV was 11 ± 2 pA/pF. Perfusion of naltriben potentiated TRPM7-like currents by approximately three folds, and the current density became 34 ± 2 pA/pF (p<0.0001, n=6). Subsequent waixenicin A perfusion significantly reduced the naltriben-potentiated current by ~50% to 17 ± 1 pA/pF (p<0.001, n=6). Additionally, naltriben also potentiated the inward current at -100 mV by

~65% (from -1.1 ± 0.3 pA/pF to -1.9 ± 0.4 pA/pF; p<0.001, n=6), and subsequently reversed by waixenicin A (to -1.2 ± 0.3; p<0.001, Figure 14E (right); n=6).

Our results suggest that TRPM7-like currents in primary mouse hippocampal neurons can be potentiated by naltriben, and this enhancement of current is sensitive to inhibition by waixenicin A.

Figure 14: Effects of waixenicin A and naltriben on TRPM7-like currents in TRPM7-overexpressing tetracycline-inducible HEK293 cells and dissociated hippocampal neurons. A) Representative current-voltage (IV) trace of TRPM7 current (1 = bath; 2 = waixenicin A and TRPM7 outward current in non-induced HEK293 cells (p>0.05; n=4). B) Representative IV trace of TRPM7 current (1 = bath; 2 = waixenicin A) and TRPM7 outward current in TRPM7-overexpressed HEK293 cells (p<0.001; n=8). C) Representative IV trace of TRPM7 current (1 = bath; 2 = naltriben) and TRPM7 outward current in non-induced HEK293 cells (p>0.05; n=5). D) Representative IV trace of TRPM7 current (1 = bath; 2 = naltriben; 3 = waixenicin A + naltriben) and outward current in TRPM7-overexpressed HEK293 cells (** p<0.01; *** p<0.001; #### p<0.0001 (n=8)). E) Representative IV trace of TRPM7 current (1 = bath; 2 = naltriben; 3 = waixenicin A + naltriben) and TRPM7 outward (left) and inward (right) currents in primary hippocampal neurons (** p<0.01; ***/### p<0.001; **** p<0.0001 (n=6)).

The data in this figure were contributed by Raymond Wong.

4.2.2 Naltriben induces calcium influx under magnesium-free and basal conditions in hippocampal neurons

To determine the effect of naltriben on TRPM7 Ca2+ dynamics, we performed fura-2 ratiometric Ca2+ imaging on DIV4-5 mouse hippocampal neurons. In the basal solution the perfusion of neurons with 50 μM naltriben resulted in increase of the ratio of emission signal at 510 nm following excitation at 340 nm/380 nm (340/380 ratio) by 0.4 ± 0.02 (Figure 15A, n=34) from the baseline level while subsequent perfusion of naltriben together with 500 nM waixenicin A increased the ratio by a lesser degree of 0.1 ± 0.02 (Figure 15A, n=34) from the baseline level, a 63% reduction in naltriben-induced calcium signal. A representative 340/380 ratio trace for this experiment is shown in Figure 15A.

Next, we investigated the effect of naltriben on Mg2+-free TRPM7 calcium influx in mouse hippocampal neurons. We found that perfusing neurons with Mg2+-free solution increased the net Ca2+ signal by 0.2 ± 0.02 from the basal level (n=19, Figure 15B). Perfusion of 50 μM naltriben in Mg2+-free solution resulted in the net increase in Ca2+ signal by 0.4 ± 0.02 (n=19, Figure 15B), a 72% increase from Mg2+-free Ca2+ influx. Perfusion of 50 μM naltriben together with 500 nM waixenicin A under Mg2+ -free conditions resulted in a smaller increase in Ca2+ signal by 0.14 ± 0.01 (n=19, Figure 15D), a 58% reduction in naltriben-induced Ca2+ signal and a 28% reduction in Mg2+-free Ca2+ influx. These waixenicin A findings were consistent with our previous findings. A representative 340/380 ratio trace for this experiment is shown in Figure 15B.

These data confirm that naltriben can modulate TRPM7-mediated Ca2+ signal in hippocampal neurons under basal and Mg2+-free conditions.

Figure 15: Effects of naltriben on TRPM7 calcium influx in dissociated hippocampal neurons. A) Representative 340/380 fura-2 ratio trace recorded from DIV4 hippocampal neurons and TRPM7 calcium influx under basal conditions (*p<0.05, n=34). B) Representative 340/380 fura-2 ratio trace recorded from DIV4 hippocampal neurons and TRPM7 calcium influx under low- calcium and magnesium-free conditions (*p<0.05, n=19). All data presented as mean±SEM. Statistical analysis was done by student t-test or one-way ANOVA with Bonferroni post-hoc (*p<0.05).

4.2.3 Naltriben induces axon retraction and loss of processes in hippocampal neurons in a dose-dependent manner

We next used naltriben to confirm the role of TRPM7 in neurite outgrowth of cultured hippocampal neurons. DIV4 hippocampal neurons were exposed to several doses of naltriben for 4 hours. The representative images of neurons treated with naltriben are shown in Figure 16A. Naltriben treatment caused a dose-dependent axonal retraction (untreated: 239.3 ± 14.8 μm, n=68; vehicle (0.001% DMSO): 218 ± 19.80 μm, n=56; 500 nM naltriben: 239 ± 13.2 μm, n=71; 1 μM naltriben: 183.2 ± 10.5 μm, n=63; 10 μM naltriben: 162.7 ± 9 μm, n=71; 25 μM naltriben: 129 ± 7.5 μm, n=51; 50 μM naltriben: 114.5 ± 9.5 μm, n=34) (Figure 16B). The dendritic lengths were shown to significantly decrease at high naltriben concentrations (untreated: 155.4 ± 9.5 μm, n=68; vehicle (0.001% DMSO): 132.5 ± 7.6 μm, n=56; 500 nM naltriben: 134.4 ± 10.5 μm, n=71; 1 μM naltriben: 147 ± 8.2 μm, n=63; 10 μM naltriben: 136.2 ± 10.2 μm, n=71; 25 μM naltriben: 98.5 ± 8 μm, n=51; 50 μM naltriben: 43 ± 5.1 μm, n=34) (Figure 16C). Total number of neurites per neuron showed a decreasing pattern, similar to axons (untreated: 31.4 ± 2.1 μm, n=68; vehicle (0.001% DMSO): 35 ± 2.3 μm, n=56; 500 nM naltriben: 20.2 ± 1.2 μm, n=71; 1 μM naltriben: 20 ± 1 μm, n=63; 10 μM naltriben: 13.7 ± 1 μm, n=71; 25 μM naltriben: 12.2 ± 0.6 μm, n=51; 50 μM naltriben: 6.1 ± 0.7 μm, n=34) (Figure 16D).

Figure 16: Activation of TRPM7 by naltriben induces axonal retraction. A) Representative images of DIV4 cultured hippocampal neurons treated with various concentrations of naltriben for 4 hours. B) Mean axonal and C) dendritic lengths measured immediately after 4 hour exposure to naltriben. D) Total number of neurites measured immediately after 4 hour exposure to naltriben. Data is presented as mean±SEM, *p<0.05 compared to untreated control, #p<0.05. Statistical significance: ANOVA with Bonferroni post-hoc.

4.2.4 Waixenicin A attenuates the effect of naltriben on axonal lengths of hippocampal neurons

Next, we examined the effect that naltriben and waixenicin A would have on axonal outgrowth when applied together. We found that when naltriben was administered together with waixenicin A, naltriben was able to inhibit waixenicin A-induced axonal outgrowth (untreated: 232.5 ± 14 μm, n = 62; vehicle (0.001% DMSO): 180.7 ± 14.2 μm, n = 42; vehicle (0.025% methanol): 229.4 ± 18 μm, n = 53; waix A 500 nM: 282.8 ± 23 μm, n = 43; naltriben 1 μM: 153.1 ± 11 μm, n = 37; naltriben 1 μM + waix A: 227.1 ± 13.1 μm, n = 39; naltriben 10 μM: 125.6 ± 9.6 μm, n = 23; naltriben 10 μM + waix A: 182.1 ± 9.3 μm, n = 38) (Figure 17A). The effect on the dendrite lengths was similar to axons, but less pronounced (untreated: 122.8 ± 10.2 μm, n = 62; vehicle (0.001%DMSO): 97.7 ± 8.2 μm, n = 42; vehicle (0.025% methanol): 109.3 ± 7.6 μm, n = 53; waix A 500 nM: 112.5 ± 11 μm, n = 43; naltriben 1 μM: 86 ± 10 μm, n = 37; naltriben 1 μM + waix A: 92.8 ± 8.4 μm, n = 39; naltriben 10 μM: 66.7 ± 4.8 μm, n = 23; naltriben 10 μM + waix A: 90.1 ± 9.4 μm, n = 38) (Figure 17B).

Overall, in this section we demonstrated that activation of TRPM7 by naltriben negatively regulates axonal outgrowth of hippocampal neurons.

Figure 17: Waixenicin A prevents axonal retraction caused by naltriben. A) Mean axonal and B) dendritic lengths measured immediately after 4 hour exposure to naltriben, waixenicin A or both. Data is presented as mean±SEM, *p<0.05, #p<0.05 compared to untreated control. Statistical significance: one-way ANOVA with Bonferroni post-hoc.

4.3 TRPM7 mediates neurite outgrowth through cytoskeleton interaction

4.3.1 A-actinin-1 and F-actin are identified as potential binding partners of TRPM7 by ESI-TRAP MS and confirmed by co- immunoprecipitation

To explore the mechanisms underlying the effect of TRPM7 activity on neuronal growth, we carried out affinity pull-down assay using TRPM7 channel protein antibody in hippocampal proteins followed by 1D SDS-PAGE and mass spectrometry to search for potential binding partners of TPRM7 channel protein. The potential binding partners were identified by ESI-TRAP MS in combination with database analysis. We found both α-actinin-1 and F-actin in the TRPM7-binding protein complex (Figure 18A). TRPM7-to-actin or TRPM7-to-actinin binding was further confirmed by co-immunoprecipitation (co-IP) assays with proteins isolated from MDA-MB-231 cells, a breast cancer cell line with high TRPM7 expression (Middelbeek et al., 2012). As shown in Figure 18B, both α-actinin-1 and F-actin were found in the TRPM7 binding protein complex.

Figure 18: TRPM7 coprecipitates with actin and α-actinin-1. A) Mass spectrometry analysis of proteins co-immunoprecipitating with anti-TRPM7 antibod from mouse hippocampal tissue. B) β-actin and α-actinin-1 coimmunoprecipitate with TRPM7 in protein isolated from MDA-MB-231 cells.

4.3.2 TRPM7 co-localizes with actin and α-actinin-1 at the neuronal growth cone

We then conducted the quantitative colocalization analysis to determine the degree of overlap between TRPM7 and α-actinin-1 or F-actin at the same pixel location on the multi-channel fluorescence images. Colocalization of fluorescent signals suggests a high probability of two antigens co-occurring in close proximity, indicated by Pearson correlation coefficient (Rr; one signifies a high degree of colocalization, and zero indicates low likelihood of localization) (Nejatbakhsh et al., 2011). The colocalization of TRPM7 and α-actinin-1 or F-actin was focused within the lamellipodia (mesh-like structures) and filopodia (filamentous structures) of the neuronal growth cones. The Rr for TRPM7 and α-actinin was 0.59±0.02 in filopodia (n=22) and 0.43±0.04 in lamellipodia (n=22) in the normal culture condition. Rr values for TRPM7 and F- actin were 0.82±0.02 in filopodia (n=22) and 0.74±0.04 in lamellipodia (n=22). Surprisingly, colocalization between F-actin and α-actinin was not as strong as expected, with Rr of .0.43±0.03 (n=22) in filopodia and 0.29±0.03 in lamellipodia (n=22) (Figure 19).

Figure 19: TRPM7 colocalizes with actin and α-actinin-1. Protein colocalization analysis using fluorescence correlation analysis indicates strong colocalization between TRPM7 and F-actin and moderate colocalization between TRPM7 and α-actinin-1 and between F-actin and α-actinin-1. Rr is Pearson correlation coefficient. Scale bar is 20 μm.

We then studied whether the colocalization requires TRPM7 activity. Interestingly, the Rr values of F-actin and α-actinin-1 increased to 0.52±0.03 in filopodia and 0.35±0.03 in lamellipodia in neurons treated with 500 nM waixenicin A for 24 hours (Figure 20A), and the Rr between TRPM7 and α-actinin decreased to 0.52±0.03 in filopodia and 0.32±0.04 in lamellipodia (Figure 20B). Colocalization between TRPM7 and F-actin remained the same as in control neurons (Figure 20C). These findings suggest that TRPM7 is more strongly colocalized with F-actin than with α-actinin in the growth cone. Blocking TRPM7 by waixenicin A reduces TRPM7-to-actinin colocalization, while enhances (α-actinin)-to-(F-actin) colocalization. All of the colocalization data are summarized in Table 7.

Figure 20: Changes in co-localization between TRPM7 and cytoskeletal proteins after treatment with waixenicin A. Co- localization changes in A) F-actin vs α-actinin-1, B) TRPM7 vs α-actinin-1 and C)TRPM7 vs F-actin 24 hours after waixenicin A treatment. Control neurons n = 22, waixenicin A-treated neurons n = 15. Data is presented as mean±SEM, *p<0.05. Statistical significance: t-test.

Table 7: Summary of the co-localization analysis.

To determine whether chronic treatment of waixenicin A affects expression levels of the proteins, we measured α-actinin level in the untreated, vehicle-treated and waixenicin A-treated neurons (24 hour-treatment) and found no significant differences in protein levels (Figure 21). Thus, it is likely that the changes in colocalization between TRPM7 and actinin were due to spatial rearrangement of α-actinin. Taken together, our observations suggest that TRPM7 forms a potential complex with F-actin and α-actinin-1 that could be involved in regulation of neurite growth. Blocking TRPM7 activity brings F-actin and α-actinin-1 into closer proximity, which allows for stronger anchoring of F-actin to the membrane and enhanced neurite outgrowth.

Figure 21: Western blot analysis of alpha-actinin-1 protein levels in control, vehicle and waixenicin A-treated neurons. Western blot analysis indicates that protein levels of α-actinin-1 do not significantly differ between control and Waixenicin A-treated groups (n=4 for each group, one-way ANOVA followed by Bonferroni post-hoc). Data is shown as mean±SEM. 4.4 Summary

In this chapter, it was demonstrated that:

1. In cultured hippocampal neurons, TRPM7 ion channel is expressed at neuronal growth cones, and its expression pattern is similar to expression pattern of F-actin.

2. A novel TRPM7 blocker, waixenicin A, reduces TRPM7-like currents and TRPM7- mediated calcium influx in cultured hippocampal neurons.

3. TRPM7 block by waixenicin A enhances neurite outgrowth at several points in culture, with a preference for axonal outgrowth.

4. TRPM7 block accelerates the progression of cultured neurons through developmental stages.

5. Activation of TRPM7 by naltriben causes neurite retraction in a dose-dependent manner which can be prevented by waixenicin A.

6. TRPM7 may regulate neurite outgrowth through interaction with two cytoskeletal proteins, F-actin and α-actinin-1, at the growth cone.

Overall, it was shown that TRPM7 is a negative regulator of axonal outgrowth and development in hippocampal neurons under normal conditions in vitro.

5 Characterizing the effects of hypoxia on TRPM7- mediated neurite outgrowth

Contributions

The work in this chapter was performed in collaboration with Raymond Wong (PhD student in Sun lab). Raymond Wong performed electrophysiological recordings. I performed cell culture experiments.

5.1 Short and long-term hypoxia differentially affects axonal outgrowth through regulating TRPM7 activity

While hypoxic brain injury is often the consequence of ischemia, or reduced blood supply to the brain, hypoxic injury can often occur without ischemia, when arterial oxygen levels are reduced. In adults and children this type of hypoxia can occur during severe anemia, drowning, cyanide poisoning or at high altitudes, and prolonged hypoxia can be damaging to the brain, causing severe and irreversible neurocognitive deficits (Peterson, 1977; Craft et al., 1993; Fayed et al., 2006; Suominen and Vahatalo, 2012). In a fetus, intrauterine hypoxia can occur due to a variety of maternal, placental and fetal conditions, some of which are a hypoxic environment (high altitudes), maternal infections, hematological conditions such as thalassemia (reduced hemoglobin production due to genetic mutation) or iron deficiency anemia, chronic inflammation and smoking (Hutter et al., 2010). All of these conditions limit maternal oxygen uptake, thus reducing oxygen supply to the fetus in the presence of adequate blood flow (Hutter et al., 2010).

The effects of reduced oxygen conditions on neurite outgrowth and development are still under debate, and different studies report conflicting findings (O'Driscoll and Gorman, 2005; Glass et al., 2002; Woronowicz et al., 2007). TRPM7 activity has been shown to be potentiated by a variety of extracellular conditions that may occur due to hypoxia, such as changes in pH (Jiang et al., 2005), production of ROS (Coombes et al., 2011) and changes in extracellular divalent cations (Wei et al., 2007). This evidence, taken together with our findings that TRPM7 activity may negatively regulate axonal outgrowth and development, suggest that TRPM7 activity may also play a role in axonal outgrowth under hypoxic conditions. The aim of this chapter is to characterize the effects of hypoxia on axonal outgrowth and TRPM7 activity.

To investigate the effects of hypoxia on axonal outgrowth, E16 hippocampal neuron cultures were used as a model. Briefly, cultures were grown until DIV4 and then separated into three treatment groups (untreated, vehicle-treated and waixenicin A-treated) and subjected to hypoxia

(5% O2) as described previously (Glass et al., 2002). In the pre-treatment paradigm waixenicin A application took place immediately before the onset of hypoxia and neurons were exposed to hypoxic conditions for 1, 2, 4, 6, 8 and 10 hours. Pre-hypoxia control neurons were fixed on DIV4 immediately before the onset of hypoxia and were used as control. The timeline of the experiment is shown in Figure 22. Neurons were fixed immediately after the hypoxia treatment,

88 and axonal lengths and dendritic lengths were analyzed using ICC with confocal microscopy and SynD analysis routine.

Figure 22: The experimental timeline of hypoxia treatment.

5.1.1 Short-term hypoxia enhances axonal outgrowth

It was found that one hour-long hypoxia treatment significantly enhanced axonal lengths in all treatment groups compared to neurons kept under normoxic conditions (pre-hypoxia control: 253 ± 19 μm, n = 43; normoxia untreated: 241±22 um, n=43; normoxia vehicle: 216 ± 13 μm, n=68; normoxia waixenicin A: 335 ± 20 μm, n=63; hypoxia untreated: 304 ± 26 μm, n=32; hypoxia vehicle: 300 ± 21 μm, n=50; hypoxia waixenicin A: 338 ± 20 μm, n=53) (Figure 23B). Dendritic analysis revealed a significant reduction in length during hypoxia treatment, which could not be prevented by waixenicin A treatment (pre-hypoxia control: 200 ± 17 μm, n = 41; normoxia untreated: 162 ± 17 μm, n=43; normoxia vehicle: 150 ± 12 μm, n=68; normoxia waixenicin A: 171 ± 11 μm, n=63; hypoxia untreated: 120 ± 13 μm, n=32; hypoxia vehicle: 126 ± 8 μm, n=50; hypoxia waixenicin A: 145 ± 10 μm, n=53) (Figure 23C). Representative images of the neurons in different treatment groups are shown in Figure 23A.

Figure 23: Short-term hypoxia enhances axonal outgrowth. A) Representative images of neurons in different treatments under normoxic and hypoxic conditions. B) Mean axonal and C) mean dendritic lengths of neurons in different treatment conditions. All data is presented as mean ±SEM, *p<0.05 (one-way ANOVA with Bonferroni post-hoc).

5.1.2 Short-term hypoxia causes downregulation of TRPM7 protein levels and activation of MEK/ERK and PI3K/Akt signaling pathways

Under hypoxic conditions, phospho-activation of MAPK/ERK kinase MEK1/2 and its downstream target ERK1/2 increased cell survival in mouse cultured cortical neurons through phospho-inactivation of pro-apoptotic Bcl2 family protein Bad (Jin et al., 2002). In neonatal brain, activation of ERK1/2 through BDNF signaling pathway provided neuroprotection against hypoxic-ischemic injury (Han and Holtzman, 2000). Interestingly, a study on human umbilical vein endothelial cells (HUVECs) showed that both pharmacological inhibition of TRPM7 and downregulation of TRPM7 by siRNA enhanced cell growth and proliferation via activation of ERK signaling pathway (Inoue and Xiong, 2009). Moreover, TRPM7 was also shown to regulate proliferation of hepatic stellate cells via ERK and PI3K signaling pathways (Fang et al., 2013). We have also recently demonstrated that ERK/MEK and PI3K/Akt signaling cascades are involved in TRPM7-mediated cytoskeletal regulation and migration of neuroblastoma cells (Chen et al., 2015b; Wong et al., 2017). These signaling cascades have also been shown to mediate neurite outgrowth. Hence, we investigated activation of these signaling pathways Western blot results indicated a downregulation of TRPM7 expression together with activation of ERK and Akt signaling pathways (Figure 24). These data suggest that short-term hypoxia causes an increase in axonal outgrowth through concurrent activation of pro-growth ERK and Akt signaling and downregulation of TRPM7 protein levels.

Figure 24: Short-term hypoxia causes downtregulation of TRPM7 and activation of ERK and Akt singalling pathways. Representative western blot of protein extracted from DIV4 hippocampal neurons immediately following 1 hour hypoxia and quantification of TRPM7, p-Akt/Akt and p-ERK/ERK protein levels in hippocampal neurons immediately following 1 hour hypoxia (N=3). All data is presented as mean ±SEM, *p<0.05 (one-way ANOVA with Bonferroni post-hoc).

5.1.3 Long-term hypoxia causes axonal and dendritic retraction

Next, we characterized the effect of long-term hypoxia on axonal and dendritic outgrowth. Prolonged (2, 4, 6, 8 and 10 hours) hypoxic treatment caused a significant reduction of both axonal (Figure 25A) and dendritic lengths (Figure 25B), compared to normoxic neurons. However, this retraction was attenuated in groups treated with waixenicin A, suggesting an involvement of TRPM7 in later stages of hypoxic response. The summary of all the measurements is presented in Appendix 5.

Figure 25: Long-term hypoxia causes axonal and dendritic retraction and TRPM7 block attenuates this retraction. A) Axonal and B) dendritic lengths under normoxic and hypoxic conditions. All data is presented as mean ±SEM, *p<0.05 (ANOVA with Bonferroni post-hoc).

5.1.4 Naltriben exacerbates the effect of long-term hypoxia on axonal outgrowth

To confirm the involvement of TRPM7 activity in long-term hypoxia-induced axonal retraction, axonal lengths were measured in neurons subjected to hypoxia for 1,2 and 4 hours with and without 1μM or 10 μM naltriben. Neurons kept in normoxia under the same conditions were used as controls. As seen in Figure 26, naltriben causes axonal retraction in normoxic neurons and significantly exacerbates axonal retraction in hypoxic neurons after 2 and 4 hours of hypoxia. Detailed data are presented in Appendix 6.

Figure 26: Naltriben excerbates axonal retraction under hypoxic conditions. Axonalof DIV4 neurons lengths under normoxic and hypoxic conditions with and without addition of naltriben. All data is presented as mean ±SEM, *p<0.05 (ANOVA with Bonferroni post-hoc).

5.1.5 Short-term hypoxia attenuates and long-term hypoxia potentiates TRPM7-like current in hippocampal neurons

To examine the effects that short and long-term hypoxia had on TRPM7 activity, whole-cell patch-clamp recordings were carried out on DIV4-5 primary hippocampal neurons in normoxic conditions (control), after short- (~1 h) or long-term (>2 h) hypoxia. As a proof-of-principle assessment of whether waixenicin A can inhibit TRPM7 activity in primary hippocampal neurons under hypoxic conditions, we used the waixenicin A-sensitive current component for all analyses. Note that since a pan-inhibitor cocktail (500 nM TTX, 25 μM APV, 40 μM CNQX, and 5 μM nimodipine) was not added, current contamination from voltage-gated sodium and calcium channels and AMPA receptors cannot be ruled out.

Representative traces of TRPM7-like current in normoxic neurons, neurons after short-term hypoxia and neurons after long-term hypoxia with and without application of waixenicin A are shown in Figure 27A, B and C, respectively. As shown in Figure 27D, we found that short-term hypoxia reduced the TRPM7 current density from 10.51±1.16 pA/pF to 6.55±0.83 pA/pF (p<0.05; n=4/group). On the contrary, long-term hypoxia enhanced the TRPM7 current density to 15.41±0.61 pA/pF (p<0.05; n=4/group) (Figure 27D). These results confirm our previous finding that on the functional level TRPM7 activity is affected by the duration of hypoxic exposure. Overall, the findings in this chapter suggest that short-term hypoxia enhances axonal outgrowth through downregulation of TRPM7 activity, while long-term hypoxia leads to axonal retraction at least in part due to potentiation of TRPM7 activity. This axonal retraction can be prevented by application of waixenicin A prior to onset of hypoxia.

Figure 27: TRPM7 activity was reduced with short-term hypoxia, and enhanced with long-term hypoxia. Representative traces of TRPM7 current density before and after application of 500 nM waixenicin A in A) control (normoxic) neurons; B) short-term hypoxic (~1 hr in hypoxic chamber) neurons; and C) long-term hypoxic (>2 h in hypoxic chamber) neurons. D) Comparison of TRPM7 current density (i.e. waixenicin A-sensitive inhibition component) in control (normoxic), short-term hypoxic, and long- term hypoxic neurons. * represents p<0.05, comparison to control; ‡‡‡ p<0.001 (ANOVA; n=4/group). Bars represent SEM.

The data in this figure were contributed by Raymond Wong.

5.1.6 Waixenicin A attenuates hypoxia-induced axonal retraction when administered immediately after hypoxia

To determine if waixenicin A application can rescue axonal retraction induced by long-term hypoxia, neurons were subjected to post-treatment paradigm. Briefly, neurons were grown for 4 days under normal conditions. On DIV4, neurons were exposed to 2-hour hypoxia. This treatment length was chosen because this is the shortest length of hypoxia that was shown to induce axonal retraction and potentiate TRPM7 current. Immediately after hypoxic treatment, neurons were treated with 500 nM waixenicin A, vehicle or left unteated and were transferred to the incubator with normal conditions. After 24 hours of recovery, neurons were fixed and axonal and dendritic lengths were measured. Neurons that were kept under normoxic conditions were used as control. It was found that during the recovery period following 2-hour hypoxia, neurons underwent further axonal retraction. This axonal retraction was rescued by waixenicin A application, suggesting that potentiation of TRPM7 activity by hypoxia could also persist during the recovery process and cause further axonal retraction under normal conditions (neurons immediately after hypoxia, untreated: 184.7 ± 11 μm, n = 52; neurons 24 hours after hypoxia, untreated: 140.2 ± 11 μm, n = 51; vehicle: 130.4 ± 8.7 μm, 41; waixenicin A: 188.7 ± 9 μm, n = 53) (Figure 28A). Dendritic lengths and total number of neurites increased 24 hours after hypoxia (Figures 28B and C).

These findings demonstrated that axons continue to retract even during recovery after hypoxia and TRPM7 block by waixenicin A can attenuate this retraction. However, TRPM7 block does not promote further outgrowth in these neurons.

Figure 28: TRPM7 block attenuates axonal retraction when administered immediately after hypoxia. A) Axonal lengths, B) dendritic lengths and C) total number of neurites of neurons before (normoxia), immediately after and 24 hours after hypoxia. All data is presented as mean ±SEM, *p<0.05, #p<0.05 compared to immediately after hypoxia (one-way ANOVA with Bonferroni post-hoc).

5.2 Presence of astrocytes in culture exacerbates the effects of hypoxia on neurons

To investigate how the presence of astrocytes in culture affects hypoxia-mediated axonal outgrowth, retraction and TRPM7-like current in neurons, the experiments described previsouly were performed on neuron-astrocyte co-cultures.

5.2.1 Presence of astrocytes in culture exacerbates hypoxia-induced neurite retraction

Next we examined how the presence of astrocytes in the culture affected hypoxia-induced neurite retraction and TRPM7-like currents in neurons. Neuron-astrocyte co-cultures were grown for 4 days. On DIV4, the cultures were separated into three treatment groups: utreated, vehicle- treated and waixenicin A-treated. Immediately after the treatment, cultures were subjected to hypoxia for 1, 2, 4 and 6 hours and fixed immediately after. Axonal and dendritic lengths were analyzed and compared to neurons in the same treatment group that were kept under normoxic conditions. As seen in Figure 29A, presence of astrocytes in the culture abolished an increase in axonal outgrowth caused by short-term hypoxia in neuronal cultures. At the same time, the retraction of axons caused by long-term hypoxia was much faster in co-cultures when compared to neuronal cultures (neuronal cultures: 27% after 2 hours, 51% after 4 hours, 54% after 6 hours; neuron-astrocyte co-cultures: 53% after 2 hours, 76% after 4 hours, 83% after 6 hours). TRPM7 block by waixenicin A prevented this retraction for up to 4 hours of hypoxia, suggesting that TRPM7 activity plays a role in axonal retraction in this culture system. Dendrites showed retraction pattern similar to axons (Figure 29B). Summary of the data is presented in Appendix 7.

Figure 29: Presence of astrocytes in culture exacerbates axonal retraction caused by hypoxia. A) Axonal and B) dendritic lengths of DIV4 neurons in neuron-astrocyte co-culture under normoxic and hypoxic conditions. All data is presented as mean ±SEM, *p<0.05 (ANOVA with Bonferroni post-hoc).

5.2.2 The effect of astrocytes on TRPM7-like current in hippocampal neurons

Next, the effect of astrocytes on TRPM7-like currents in neurons was investigated. As a proof- of-principle assessment of whether waixenicin A can inhibit TRPM7 activity in primary hippocampal neurons under hypoxic conditions, we used the waixenicin A-sensitive current component for all analyses. Note that since a pan-inhibitor cocktail (500 nM TTX, 25 μM APV, 40 μM CNQX, and 5 μM nimodipine) was not added, current contamination from voltage-gated sodium and calcium channels and AMPA receptors cannot be ruled out.

Representative traces of TRPM7-like current in normoxic neurons, neurons after short-term hypoxia and neurons after long-term hypoxia with and without application of waixenicin A are shown in Figure 30A, B and C, respectively. In neurons that were co-cultured with glial cells, short-term hypoxia had no significant effect on neuronal TRPM7 activity (i.e. 8.59±0.81 pA/pF and 9.23±1.61 pA/pF under normoxic control and short-term hypoxic conditions, respectively Figure 30D). Nevertheless, long-term hypoxia increased neuronal TRPM7 activity to 12.84±0.80 pA/pF (Figure 30D) (p<0.05 compared to normoxic control; not significant compared to short- term hypoxia group). These results demonstrate that in this culture system, TRPM7 was also regulated by hypoxic conditions.

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Figure 30: TRPM7 activity of neurons co-cultured with glia under short-term and long-term hypoxia. Representative traces of TRPM7 current density before and after application of 500 nM waixenicin A in A) control (normoxic) neurons; B) short-term hypoxic (~1 hr in hypoxic chamber) neurons; and C) long-term hypoxic (>2 h in hypoxic chamber) neurons. D) Comparison of TRPM7 current density (i.e. waixenicin A-sensitive inhibition component) in control (normoxic), short-term hypoxic, and long- term hypoxic neurons. * represents p<0.05, comparison to control (One-way ANOVA with Bonferonni multiple comparison tests; n=3/group). Bars represent SEM.

The data in this figure were contributed by Raymond Wong.

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5.2.3 The effect of soluble factors on axonal retraction

Finally, we investigated whether the effect of astrocytes on axonal retraction was due to soluble factors released by these cells, cell-cell interactions or both. We performed a set of media switch experiments outline below.

Neuronal cultures or neuron-astrocyte co-cultures were grown for 4 days under normal conditions. On DIV4, these cultures were subjected to hypoxia for 1, 2, 4 and 6 hours. Immediately after, the media from these cultures were collected and placed on DIV4 neuronal cultures under normal conditions for 2 hours. After 2 hours, neuronal cultures that received conditioned media were fixed with 4% PFA and prepared for immunolabelling. Control cultures were kept under normal conditions and fixed at 1, 2, 4 and 6 hour timepoints for comparison. At the time of the media switch, neurons receiving conditioned media were also treated with vehicle, waixenicin A or left untreated.

Overall, axonal lengths followed the patterns described in previous sections, suggesting that the soluble factors released under hypoxic conditions may play an important role in initiating axonal retraction. The media collected from hypoxic neuronal cultures resulted in an increase in axonal outgrowth after short-term hypoxia and subsequent trend towards axonal retraction (Figure 31A). The media collected from hypoxic neuron-asotrcyte co-cultures resulted in progressive axonal retraction as hypoxia exposure increased (Figure 31B). The data are summarized in Appendix 8.

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Figure 31: Axonal lengths following a media switch. Axonal lengths of neurons that received conditioned media from A) hypoxic neuronal cultures or B) hypoxic neuron-astrocyte co-cultures. The numbers on X axes represent duration of hypoxia in hours. All data is presented as mean ±SEM, *p<0.05 (ANOVA with Bonferroni post-hoc).

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5.3 Summary

In this chapter, it was demonstrated that:

1. Short-term hypoxia (1 hour) enhanced axonal outgrowth of neuronal cultures.

2. Short-term hypoxia activated MEK/ERK and PI3K/Akt pathways.

3. Long-term hypoxia ( > 2.0 hours) caused a progressive retraction of axons and dendrites and waixenicin A attenuated this retraction.

4. Short-term hypoxia reduced TRPM7 activity, while long-term hypoxia potentiated TRPM7 activity in cultured hippocampal neurons.

5. Presence of astrocytes in cultures enhanced axonal retraction caused by hypoxia.

6. In the presence of astrocytes, short-term hypoxia had no effect on TRPM7 activity, while long-term hypoxia potentiated TRPM7 activity.

7. Neurons subjected to media conditioned on hypoxic neuronal cultures or neuron- astrocyte co-cultures exhibited similar patterns of axonal outgrowth/retraction as cultures subjected to hypoxia, suggesting that soluble factors present in media may play a role in axonal retraction. Overall, this chapter characterizes the effect of hypoxia on TRPM7 activity and axonal outgrowth/retraction with and without astrocytes present in culture.

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6 Neuroprotective and therapeutic effects of waixenicin A in mouse model of neonatal hypoxic-ischemic (HI) brain injury

Contributions

This work was completed in collaboration with Dr. Baofeng Xu, Dr. Wengliang Chen, Sammen Huang, Feiya Li, Dr. David F. Horgen, Dr. Andrea Fleig, Dr. Zhong-Ping Feng and Dr. Hong- Shuo Sun. Dr. Wenliang Chen provided assistance with model development and Western blot experiments. Dr. Baofeng Xu, Sammen Huang, Feiya Li and myself performed animal surgery, histological and short-term behavior assessments. I performed immunohistochemistry experiments and long-term behavior experiments. Drs. David F. Horgen and Andrea Fleig supplied waixenicin A. Drs. Zhong-Ping Feng and Hong-Shuo Sun contributed to study design and manuscript preparation.

This chapter contains a recreation of the manuscript in preparation for submission with the following citation:

Turlova et al., TRPM7 mediates cell death through cytoskeletal regulation via calcium/calmodulin-dependent protein kinase II and calcineurin in neonatal hypoxic-ischemic brain injury. 2018. In preparation.

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6.1 Model development and experimental timeline

As it was previously shown, TRPM7 plays an important role in hypoxic-ischemic brain injury in adult rodents in vivo (Sun et al., 2009). It is clinically important to evaluate the neuroprotective and therapeutic potential of the novel TRPM7 blocker waixenicin A in an in vivo model of hypoxic-ischemic brain injury. Due to differences in responses of neonatal and adult brains to hypoxia-ischemia, the response of neonatal brain to TRPM7 block by waixenicin A may also differ from that of adult, and thus needs to be determined. Additionally, there is an urgent need for novel treatments for neonatal hypoxic-ischemic brain injury, due to permanent and debilitating neurocognitive impairments. Therefore, an in vivo model of neonatal hypoxic- ischemic brain injury will be used to evaluate therapeutic potential of waixenicin A.

Postnatal day 7 (P7) pups were used in these experiments, as it was shown that brain development of P7 pups corresponds closely to brain development of a full-term baby (Clancy et al., 2001). The injury was induced at P7 by permanent ligation of the right carotid artery following the subjection of pups to mild (7.5% O2) hypoxia for 60 minutes, a well-described method that results in an infarct lesion in the hemisphere ipsilateral to the carotid occlusion (Rice, III et al., 1981). Experimental timeline: Experimental timeline of drug administration and experimental procedures are outlined in Figure 32. The following timelines for drug administration were used: Pre-treatment or pre-ischemia: waixenicin A or vehicle solution were administered 30 minutes prior to surgical occlusion of common carotid artery. Post-treatment 1 or post-ischemia: waixenicin A or vehicle solution were administered 30 minutes prior to onset of hypoxia. Post-treatment 2 or immediately post-hypoxia: waixenicin A or vehicle solution were administered immediately after hypoxic treatment. Post-treatment 3 or 1 hour post-hypoxia: waixenicin A or vehicle solution were administered 1 hour after hypoxic treatment.

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Figure 32: Timeline of treatments, histological, morphological and neurobehavioural experiments.

6.2 Waixenicin A has neuroprotective and therapeutic effects in neonal model of HI

To determine the effect of waixenicin A on the infarct volume following HI injury, waixenicin A (37 ng/g) or vehicle (saline with 0.0037% methanol) were administered as a single intraperitoneal injection to postnatal P7 pups according to the timeline in Figure 32.

6.2.1 Waixenicin A pre-treatment reduces brain injury 24 hours after HI

The pups in “pre-treatment” group were injected 30 minutes before induction of HI. The brains were harvested and stained with triphenyl tetrazolium chloride (TTC) 24 hours after injury. We found that compared to the vehicle-treated group, the waixenicin A-treated group had significantly reduced corrected infarct volumes of the ipsilateral hemispheres (vehicle: 48 ± 7 %, n=11; waixenicin A: 25 ± 7 %, n=10), as illustrated in Figure 33B. Figure 33A shows representative TTC stained images of vehicle and waixenicin A-treated brains 24 hours following the injury.

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Figure 33: Morphological assessment of brain injury in pre-treatment paradigm 24 hours after hypoxic ischemic insult. A) Representative TTC staining and B) corrected infarct volume of vehicle and waixenicin A groups. All data presented as mean±SEM. Statistical analysis was done by student t-test (*p<0.05).

6.2.2 Waixenicin A reduces brain injury 24 hours after HI when administered up to 1 hour after the onset of injury

The “post-treatment 1” group received an injection of vehicle or waixenicin A 30 minutes before the onset of hypoxia. When compared to the vehicle-treated group, the waixenicin A-treated group also had significantly smaller infarct volumes 24 hours following the injury in this treatment paradigm (vehicle: 51 ± 4 %, n=11; waixenicin A: 26 ± 3 %, n=15). Figure 34A (left) shows representative TTC stainings, while Figure 34A (right) shows corrected infarct volumes of the ipsilateral hemisphere.

The “post-treatment 2” group received an injection of vehicle or waixenicin A immediately following hypoxia exposure. Figure 34B (left) shows representative TTC staining, while Figure 34B (right) shows a significant reduction in corrected infarct volumes in the waixenicin A- treated group, compared to the vehicle-treated group (vehicle: 63 ± 12 %, n=11; waixenicin A: 34 ± 7 %, n=14).

The “post-treatment 3” group received an injection of vehicle or waixenicin A 1 hour after the hypoxia event. Representative TTC staining of brains treated with vehicle and waixenicin A are shown in Figure 34C (left), while Figure 34C (right) shows a significant reduction in corrected

108 infarct volumes in the waixenicin A-treated group compared to the vehicle-treated group (vehicle: 55 ± 5 %, n=17; waixenicin A: 34 ± 5 %, n=10) 24 hours following the injury.

These data show that application of waixenicin A reduces HI-induced brain damage when administered both before and after HI injury.

Figure 34: Morphological assessment of brain injury in post-treatment paradigm 24 hours after hypoxic ischemic insult. A) Representative TTC staining and corrected infarctvolume of vehicle and waixenicin A groups treated according to post-treatment 1 paradigm. B) Representative TTC staining and corrected infarct volume of vehicle and waixenicin A groups treated according to post-treatment 2 paradigm. C) Representative TTC staining and corrected infarct volume of vehicle and waixenicin A groups treated according to post-treatment 3 paradigm. All data presented as mean±SEM. Statistical analysis was done by student t-test (*p<0.05).

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6.2.3 Naltriben exacerbates brain injury 24 hours after HI when administered as a pre-treatment

Next, we examined the effect of TRPM7 activator naltriben (Hofmann et al., 2014) on the extent of brain injury following HI induction. Naltriben was injected intraperitoneally as a single dose of 42 μg/g of body weight as a pre-treatment (30 minutes before the onset of ischemia). The brains were harvested and stained with triphenyl tetrazolium chloride (TTC) 24 hours following the injury. We found that naltriben application significantly increased the corrected infarct volumes as shown in Figure 35 (vehicle (0.001% DMSO): 53 ± 2 %, n=22; naltriben: 71 ± 4 %, n=15). These data demonstrate that application of the TRPM7 activator naltriben exacerbates brain damage after neonatal HI.

Figure 35: Morphological assessment of brain injury in pre-treatment paradigm with naltriben treatment 24 hours after hypoxic ischemic insult. Representative TTC staining and corrected infarct volume of vehicle and naltriben groups treated according to pre-treatment paradigm. All data presented as mean±SEM. Statistical analysis was done by student t-test (***p<0.001).

6.2.4 Waixenicin A pre-treatment reduces apoptotic brain death, reactive astrocyte activation and TRPM7 upregulation in ischemic penumbra

Next, we assessed the effects of waixenicin A on apoptotic signalling, neuronal survival and TRPM7 expression by performing biochemical analysis of sham, vehicle-treated and waixenicin A-treated brains 24 hours after HI injury. We found that HI injury upregulated cleaved caspase-3 and caspase-9 levels and significantly reduced Bcl-2/Bax protein ratio, and waixenicin A pre-

110 treatment restored the levels of these proteins (Figure 36B: caspase-9/β-actin normalized to sham group expression: vehicle-treated: 3.6 ± 0.6, n=8; waixeincin A-treated: 1.2 ± 0.4, n=8; Figure 36C: caspase-3/β-actin normalized to sham group expression: vehicle-treated: 17 ± 1.2, n=8; waixenicin A-treated: 8.9 ± 0.8, n=8; Figure 36D: Bcl-2/Bax normalized to sham group expression: vehicle-treated: 0.6 ± 0.02, n=8; waixenicin A-treated: 1 ± 0.02, n=8).

Figure 36: Biochemical assessment of brain injury following hypoxic-ischemic insult on neonatal brain in a pre-treatment paradigm. A) Apoptotic signalling is reduced in waixenicin A pre-treated group 24 hours after HI injury as seen by B) reduction in caspase-9 and C) cleaved caspase-3 expression and D) restoration of Bcl-2/Bax ratio. The data is presented as mean±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc, *p<0.05.

We also performed immunohistochemical staining and analysis of the penumbra area of the brain slices in sham, vehicle- and waixenicin A-treated groups three days after HI injury. Representative confocal images are shown in Figure 37A. As seen in Figure 37B, waixenicin A pre-treatment significantly reduced the loss of NeuN-positive cells when compared to vehicle- treated group (as a part of contralateral hemisphere: sham: 1 ± 0.04, n=9; vehicle: 0.5 ± 0.04 n=9; waixenicin A: 0.7 ± 0.03 n=9). Moreover, we also observed an upregulation of GFAP expression in astrocytes which represents astroglial activation and reactive gliosis during trauma or neurodegeneration in vehicle-treated group which was significantly attenuated in waixenicin A-treated group. These data are shown in Figure 37C and represent GFAP-positive cells per 20X field (sham: 13 ± 6, n=6; vehicle: 245 ± 61, n=6; waixenicin A: 41 ± 18, n=6). We also found a significant upregulation of cleaved caspase-3 and TRPM7 protein levels in vehicle-treated groups with attenuation of both proteins in waixenicin A-treated group. This is shown in Figure 37D as the number of cleaved caspase-3 positive cells per 20X field (sham: 7 ± 1, n=5; vehicle: 100 ± 11, n=5; waixenicin A: 38 ± 9, n=5) and Figure 37E as relative TRPM7 levels compared to

111 contralateral hemisphere (sham: 1 ± 0.06, n=10; vehicle: 2 ± 0.2, n=11; waixenicin A: 1.3 ± 0.2, n=8 of contralateral hemisphere).

These data demonstrate that waixenicin A pre-treatment reduces apoptotic and delayed neuronal cell death and attenuates glial cell activation in ischemic penumbra.

Figure 37: Immunohistochemical assessment of brain injury following hypoxic-ischemic insult on neonatal brain in a pre- treatment paradigm. A) Representative images of sham, vehicle and waixenicin A-treated coronal brain slices stained for NeuN (neuronal marker), GFAP (astrocyte marker), cleaved caspase-3 (apoptotic number) and TRPM7 three days following injury. B) Ratio of ipsilateral/contralateral NeuN-positive cells is restored in waixenicin A-treated group. C) GFAP-positive cells per 20X field. D) Cleaved caspase-3 positive cells per 20X field. E) TRPM7 expression is upregulated three days after HI injury and is attenuated by waixenicin A administration. The data is presented as mean±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc, *p<0.05, **p<0.01, ***p<0.001, #p<0.05 as compared to sham.

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6.3 Waixenicin A preserves short-term function and overall brain morphology 7 days after HI

6.3.1 Waixenicin A preserves brain morphology 7 days after HI injury

To determine the effects of waixenicin A treatment on preserving brain morphology following HI injury, we evaluated corrected ipsilateral liquefaction volumes and brain weight in different treatment groups 7 days after HI injury. The overall brain morphology was preserved as shown in Figure 38A and B (pre-treatment) and Figure 38E and F (post-treatment 2). Waixenicin A- treated groups showed significantly reduced corrected ipsilateral liquefaction volumes in pre- treatment as shown in Figure 38C (sham: 0, n=3; vehicle: 46 ± 3 %, n=13; waixenicin A: 27 ± 3 %, n=12) and post-treatment, as shown in Figure 38G (sham: 0, n=5; vehicle: 55 ± 6 %, n=12; waixenicin A: 34 ± 5 %, n=11) . Brain weights were also significantly greater in both waixenicin A pre-treatment and post-treatment groups, as seen in Figure 38D (vehicle: 0.33 ± 0.02 g, n=14; waixenicin A: 0.39 ± 0.01 g, n=18) and Figure 38H (vehicle: 0.32 ± 0.01 g, n=13; waixenicin A: 0.38 ± 0.01 g, n=11), respectively.

These data show that waixenicin A prevents brain mass loss and preserves brain morphology in the short term after HI injury.

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Figure 38: Morphological assessment of brain injury 7 days after hypoxic-ischemic insult on neonatal brain. A) Overall brain morphology is preserved in the pre-treatment paradigm 7 days after HI injury. B) Nissl staining shows reduced liquefaction volume in waixenicin A-treated group in pre-treatment paradigm. C) Brain weight is significantly higher in waixenicin A-treated group in pre-treatment paradigm. D) Corrected ipsilateral liquefaction volume is significantly reduced in waixenicin A-treated group in pre-treatment paradigm. E) Overall brain morphology is preserved in the post-treatment 2 paradigm 7 days after HI injury. F) Nissl staining shows reduced liquefaction volume in waixenicin A-treated group in post-treatment 2 paradigm. G) Brain weight is significantly higher in waixenicin A-treated group in post-treatment 2 paradigm. H) Corrected ipsilateral liquefaction volume is significantly reduced in waixenicin A-treated group in post-treatment 2 paradigm. All data presented as mean±SEM. Statistical analysis was done by student t-test or one-way ANOVA with Bonferroni post-hoc (*p<0.05).

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6.3.2 Waixenicin A preserves short-term function following HI injury

To determine the effects of waixenicin A on function after HI injury, we subjected the pups to multiple tests 1, 3 and 7 days following HI to assess vestibular, proprioceptive and motor functions as well as strength and fatigability of muscles. We found that pre-treatment (before ischemia) administration of waixenicin A significantly improved outcomes of righting reflex, geotaxis reflex, cliff avoidance reflex and grip strength test.

Righting reflex is a motor ability of the pup to flip back to its feet from a supine position, and appears in rodents on P5 to P10. This is a reflex, and therefore it has no learning component (Feather-Schussler and Ferguson, 2016b). We found that it significantly improved in waixenicin A-treated group compared to vehicle-treated group 1 day (sham: 0.8 ± 0.09 sec, n=14; vehicle: 2.7 ± 0.6 sec, n =13; waixenicin A: 1.5 ± 0.3 sec, n = 12)) after HI (Figure 39A).

Negative geotaxis or geotaxis reflex is an innate behavior that is used to assess motor coordination in response to a vestibular cue. Pups were placed facing down a 45ºC slope and the time to turn around and face up the slope was recorded. This reflex appears in rodents on P7 to P15 (Feather-Schussler and Ferguson, 2016a). Geotaxis reflex performance was also significantly improved in waixenicin A-treated group compared to vehicle-treated group on day 1 (sham: 4.4 ± 0.4 sec, n=14; vehicle: 8.3 ± 1.3 sec, n=13; waixenicin A: 5.3 ± 0.4 sec, n=12) and day 3 (sham: 2.9 ± 0.3 sec, n=14; vehicle: 5.7 ± 0.7 sec, n=13; waixenicin A: 3.1 ± 0.3 sec, n=12) post-HI (Figure 39B).

Cliff avoidance reflex appears in rodents on P1 to P14, and is used to test for vestibular function and motor coordination. Pups were placed with the digits of their front paws and snout over a flat elevated edge, and the time it took the pup to move its paws and snout away from the edge was recorded (Feather-Schussler and Ferguson, 2016e). Cliff avoidance reflex showed significant improvement in waixenicin A-treated group compared to vehicle-treated group on day 1 (sham: 2.2 ± 0.2 sec, n=14; vehicle: 6.7 ± 1.5 sec, n=13; waixenicin A: 3.2 ± 0.4 sec, n=12) and day 3 (sham: 1.7 ± 0.1 sec, n=14; vehicle: 4.5 ± 0.7 sec, n=13; waixenicin A: 2.9 ± 0.3 sec, n=12) post- HI (Figure 39C).

Grip strength test is used to assess the paw strength of front paws. The pups were allowed to hold on to the horizontal wire and the time it took the pup to fall down was recorded (Feather-

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Schussler and Ferguson, 2016d). Grip strength was significantly improved with waixenicin A treatment on day 7 post-HI (sham: 22 ± 2.6, n=14; vehicle: 9.4 ± 1 sec, n=13; waixenicin A: 20 ± 4.3 sec, n=12) (Figure 39D).

These data demonstrate that waixenicin A pre-treatment significantly improves functional recovery after HI in the short-term, as evident by improved performance on a battery of neurobehavioural tests.

Figure 39: Assessment of motor and vestibular function up to 7 days after HI. A) Righting reflex 1 and 3 days after HI, B) geotaxis reflex 1,3 and 7 days after HI, C) cliff avoidance reflex 1,3 and 7 days after HI and D) grip strength 1, 3 and 7 days after HI. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, *p<0.05.

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6.4 Waixenicin A preserves long-term function and overall brain morphology up to 32 days after HI

To determine whether waixenicin A treatment would also improve functional recovery in the long-term, we subjected mice in the pre-treatment paradigm to a battery of motor and memory tests 4 weeks after HI.

6.4.1 Waixenicin A pre-treatment preserves Accelerated Rotarod outcomes

To test for motor and balance, we subjected the mice to Accelrated Rotarod Test. We found that mice in waixenicin A-treated groups had a significantly improved motor function, compared to vehicle-treated mice as was evident by longer latency (Figure 40A: sham: 61 ± 3 sec, n = 24; vehicle: 27 ± 3 sec, n=15; waixenicin A: 51 ± 4 sec, n=16) and higher speed (Figure 40B: sham: 22 ± 1 rpm, n=24; vehicle: 13 ± 1 rpm, n=15; waixenicin A: 19 ± 1 rpm, n=16) at which waixenicin A-treated mice fell off the drum.

Figure 40: Accelerated Rotarod Test administered 28 days after HI. In the Accelerated Rotarod Test, mice in waixenicin A- treated groups had a significantly improved motor and balance function as was evident by A) longer latency and B) higher speed at which waixenicin A-treated mice fell off the drum. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, **p<0.01, ***p<0.001.

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6.4.2 Waixenicin A pre-treatment preserves memory test outcomes

To determine whether waixenicin A treatment attenuates memory impairment after HI (Blasi et al., 2014; Hattori et al., 2000), we subjected mice in pre-treatment paradigm to two different memory tests 4 weeks after HI injury.

First, the mice were subjected to novel object recognition test to evaluate spontaneous exploratory behavior triggered by novelty (Leger et al., 2013). The objects used for this test were similar in size and odour, but different in shape and texture. The test was carried out in two sessions separated by an intersession interval of 4 hours (Sik et al., 2003) (Figure 41A). During familiarization session, the mouse was placed in an open-field arena facing away from two identical objects placed 5 cm apart and was allowed to explore both objects freely for 3 minutes of session duration (Leger et al., 2013). During the test session, one of the familiar objects was replaced by an identical copy to avoid olfactory cues (Leger et al., 2013), and another familiar object was replaced by a novel object. The mouse was placed in an open-field arena facing away from the objects and allowed to explore the objects freely for 3 min of session duration. The exploration time of each object was recorded and preference for novel object was calculated as a percentage of total exploration time. We found that mice in waixenicin A-treated groups had a significantly greater novel object preference, compared to vehicle-treated mice (Figure 41B: % of time spent exploring both objects: sham: 65 ± 2 %, n = 12; vehicle: 48 ± 3%, n=8; waixenicin A: 64 ± 2%, n=11).

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Figure 41: Novel object recognition test 4 weeks after HI injury. A) The schematic of the experimental setup. B) Mice in waixenicin A-treated groups had a significantly greater novel object preference compared to vehicle-treated mice. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, *p<0.05.

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To test for the ability to inhibit innate preference for a dark confined space after a single association with an aversive stimulus the mice were subjected to passive avoidance test, as described previously (Haelewyn et al., 2007). The training protocol consisted of three sessions performed on separate days: habituation, acquisition and retention (Figure 42A). Briefly, during habituation session the mouse was placed into the illuminated compartment and allowed to explore for 1 minute. After 1 minute, the door to the dark compartment was opened and latency to enter the dark room was recorded. During acquisition session, the protocol was the same as habituation session, but the mouse received an inescapable foot shock (0.4 mA, 2 sec) upon entering the dark compartment. The retention session took place 48 hours after acquisition session. The mouse was placed into the illuminated compartment, the door to the dark compartment was opened after 5 sec and latency to enter the dark compartment was recorded. The latency to enter dark compartment during the retention session was taken as an index of memory performance. Waixenicin A-treated mice also showed better memory function compared to vehicle-treated mice, as was evident from significantly longer latency to enter the dark room 48 hours after the foot shock (Figure 42B: sham: 116 ± 31 sec, n=12; vehicle: 25 ± 9 sec, n = 8; waixenicin A: 87 ± 17 sec, n=11).

Together, these tests indicate that mice in waixenicin A-treated groups perform better on motor and memory functions than vehicle-treated mice as late as 4 weeks after HI injury.

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Figure 42: Passive avoidance test 4 week after HI. A) A schematic diagram of the experimental setup. B) Waixenicin A-treated mice showed better memory function as was evident from significantly longer latency to enter the dark room 48 hours after the foot shock. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, *p<0.05.

6.4.3 Waixenicin A pre-treatment preserves overall brain morphology

Next, brain morphology of mice in different treatment groups was examined. Liquefaction brain volume of waixenicin A-treated mice was significantly smaller than of vehicle-treated mice (Figure 43B, % of ipsilateral hemisphere: sham: 0, n=12; vehicle: 47 ± 10%, n=8; waixenicin A: 19 ± 6%, n=14). Representative images of brains extracted 32 days after HI procedure are shown in Figure 43A. Waixenicin A-treated mice also had significantly greater brain weight (Figure 43C: sham: 0.5±0.01 g, n=18; vehicle: 0.37 ± 0.02 g, n=8; waixenicin A: 0.44 ± 0.01 g, n=14) and body weight (Figure 43D: sham: 26 ± 1 g, n=18; vehicle: 18 ± 0.9 g, n=8; waixenicin A: 23 ± 1 g, n=14), indicating an enhanced general recovery in the long-term.

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Figure 43: Brain morphology, brain weight and body weight 32 days after HI. A) Overall brain morphology is preserved in waixenicin A-treated group, and B) liquefaction brain volume of waixenicin A-treated mice was significantly smaller 32 days after HI. Waixenicin A-treated mice had significantly greater C) brain and D) body weights, indicating an enhanced general recovery in the long-term. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, *p<0.05, **p<0.01. ***p<0.001.

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6.5 Summary

In this chapter, it was demonstrated that:

1. Waixenicin A has neuroprotective and therapeutic properties in an animal model of neonatal HI.

2. Administration of waixenicin A prior to onset of injury reduces brain damage and preserves brain morphology.

3. Administration of waixenicin A prior to onset of injury reduces apoptotic cell death after HI.

4. Administration of waixenicin A prior to onset of injury improves short-term motor and proprioceptive functions.

5. Administration of waixenicin A prior to onset of injury preserves long-term motor and memory functions and brain morphology.

Overall, in this chapter it was shown that TRPM7 may be a potential therapeutic target for neonatal HI brain injury, and that waixenicin A is a promosing candidate for future testing and development.

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7 TRPM7-dependent molecular mechanisms in neonatal HI

Contributions

This chapter contains a re-creation of the manuscript in preparation with the following citation:

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7.1 Characterization of protein changes 6 hours after HI

So far, it’s been shown that TRPM7 regulates axonal outgrowth and retraction in vitro, and blocking TRPM7 with waixenicin A has neuroprotective and therapeutic outcomes in an in vivo model of neonatal HIE. Next, TRPM7-dependent molecular mechanisms must be investigated.

A mouse neonatal HI model was also used in this chapter, and the changes in protein levels between sham, vehicle and waixenicin A-treated groups were determined by liquid chromatography tandem-mass spectrometry (LC-MS/MS) 6 hours after HI.

Samples from sham, vehicle-treated and waixenicin A-treated groups were collected 6 hours after HI and subjected to liquid chromatography tandem-mass spectrometry (LC-MS/MS) using Orbitrap analyzer. We found 95 proteins that were differentially expressed, displaying more than 1.5-fold change, between sham and vehicle groups and were also identified in waixenicin A group (Figure 44A). The full list of the proteins with Uniprot ID and fold change can be found in Appendix 9. The proteins were uploaded into the STRING 10.5 database (Szklarczyk et al., 2017) to identify protein interactions based on co-expression, information from curated databases, experimentally determined interactions and text mining (Figure 44B).

Classification of proteins by molecular function reveled that majority of the proteins were involved in enzyme activity, nucleotide and nucleic acid binding, cytoskeletal protein binding and ion binding (Figure 44C) (enzyme activity 21%, nucleotide binding 13%, cytoskeletal protein binding 12%, RNA binding 10%, metal ion binding 9%, receptor activity 7%, transmembrane transport activity 7%, DNA binding 6%, kinase activity 6%, calcium ion binding 5%, lipid binding 5%). Classification of proteins by cellular compartments showed that majority of proteins were present in the cytoplasm, nucleus, plasma membrane or associated with the cytoskeleton (Figure 44D) (cytoplasm 28%, nucleus 21%, cell membrane 13%, cytoskeleton 11%, mitochondrion 7%, synapse 6%, secreted 5%, golgi apparatus 5%, endoplasmic reticulum 4%).

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Figure 44: Overview of proteomic changes between sham, vehicle and waixenicin A-treated groups after HI. A) A heatmap of 95 differentially expressed proteins. B) STRING interaction network of these proteins. C) Molecular function classification and D) cellular component categories of the proteins.

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7.1.1 Proteins with calcium-binding and cytoskeletal regulation functions

To build on the previously discovered TRPM7-mediated molecular mechanism, we focused on the proteins with molecular function of calcium binding and regulation of the cytoskeleton. The list of the proteins and their STRING network are presented in Figure 45.

These data indicate that waixenicin A attenuates HI brain injury potentially through calcium- dependent signalling cascade.

Figure 45: Calcium-binding cytoskeletal regulators that were indentified by LC-MS/MS.

7.2 TRPM7 signaling through CaMKII and regulation of cytoskeleton

We focused on the analysis of three identified proteins – calcium/calmodulin-dependent protein kinase II, calmodulin and calcineurin – and their downstream signaling cascades. We chose these proteins because they were shown to be present at the neuronal growth cone and affect neurite outgrowth through upstream regulation of actin-interacting proteins, such as α-actinin, which we previously identified as potential TRPM7 binding partner (Henley and Poo, 2004).

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7.2.1 Proteomic changes of CaMKII, calcineurin and calmodulin

We found that protein levels of Ca2+/calmodulin-dependent protein kinase II (CaMKII) (Uniprot ID P11798), calcineurin (regulatory subunit B) (Uniprot ID Q63810) and calmodulin (Uniprot ID P0DP2) were significantly different between the treatment groups (Figure 46A, fold change normalized to sham group expression: CaMKII, vehicle-treated group: 0.66±0.07, n=3, waixenicin A-treated group: 1.00±0.03, n=4; Figure 46B: calcineurin regulatory subunit B, vehicle-treated group: 1.5±0.3, n=3, waixenicin A-treated group: 0.99±0.15, n=4; Figure 46C: calmodulin, vehicle-treated group: 2.06±0.34, n=3, waixenicin A-treated group: 1.03±0.2, n=4).

Figure 46: Changes in CaMKII, calcineurin and calmodulin protein levels. Changes in A) CaMKII, B) calcineurin (regulatory subunit B) and C) calmodulin levels, as detected by LC-MS/MS 6 hours after HI. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, *p<0.05.

7.2.2 Downstream signaling and cytoskeleton regulation from CaMKII, calcineurin and calmodulin

Next, we examined the changes in CaMKII, calmodulin and calcineurin levels as well as downstream proteins in sham, vehicle-treated and Waixenicin A-treated groups 6 and 24 hours after HI. We found that HI injury significantly reduced CaMKII protein levels, CaMKII and p38 phosporylation levels and increased calcineurin B protein levels in the ipsilateral hemisphere, and waixenicin A pre-treatment restored the protein levels of these proteins 6 (Figure 47A) and 24 hours (Figure 47B) after HI (Figure 47A: p-p38/p38 normalized to sham group expression: vehicle-treated: 0.62 ± 0.03, n=4, waixenicin A-treated: 0.86 ± 0.08, n=4; CaMKII/β-actin normalized to sham group expression: vehicle-treated: 0.69±0.07, n=4, waixenicin A-treated: 0.92±0.06, n=4; p-CaMKII/CaMKII normalized to sham group expression: vehicle-treated: 0.75 ± 0.04, n=4; waixenicin A-treated: 0.94 ± 0.06, n=4; calcineurin B normalized to sham group

128 expression: vehicle-treated: 1.26±0.05, n=4; waixenicin A-treated: 1.05±0.06, n=4; Figure 47B: p-p38/p38 normalized to sham group expression: vehicle-treated: 0.5 ± 0.07, n=8; waixenicin A- treated: 0.7 ± 0.06, n=8; CaMKII/β-actin normalized to sham group expression: vehicle-treated: 0.53±0.11, n=4, waixenicin A-treated: 0.95±0.23, n=4; p-CaMKII/CaMKII normalized to sham group expression: vehicle-treated: 0.7 ± 0.13, n=4; waixenicin A-treated: 0.9 ± 0.14, n=4; calcineurin B normalized to sham group expression: vehicle-treated: 1.24±0.04, n=4; waixenicin A-treated: 1.11±0.02, n=4). Calmodulin protein levels were significantly upregulated in the ipsilateral hemisphere in HI group 6 hours after injury (Figure 47A), but not 24 hours after injury (Figure 47B) (Figure 47A: calmodulin normalized to sham group expression: vehicle-treated: 1.32±0.08, n=4; waixenicin A-treated: 1.08±0.06, n=4).

Modification of actin cytoskeleton have also been linked to cell death under hypoxic conditions (Posmantur et al., 1996). Since we have previously shown that TRPM7 interacts with α-actinin-1 (Turlova et al., 2014), we evaluated the effect of HI injury and waixenicin A treatment on the expression levels of two cytoskeleton-modulating proteins, α-actinin-1 and cofilin. We found that HI injury significantly upregulated α-actinin-1 expression levels and significantly reduced cofilin phosphorylation levels, and waixenicin A pre-treatment restored the levels of these proteins 24 hours after HI (Figure 47B), while there was a similar trend in changes in these proteins 6 hours after HI (Figure 47A) (Figure 47A: α-actinin-1/β-actin normalized to sham group expression: vehicle-treated: 1.08 ± 0.08, n=4, waixenicin A-treated: 1.05 ± 0.09, n=4, p- cofilin/cofilin normalized to sham group expression: vehicle-treated: 0.84 ± 0.07, n=4, waixenicin A-treated: 0.96 ± 0.07, n=4; Figure 47B: α-actinin-1/β-actin normalized to sham group expression: vehicle-treated: 1.8 ± 0.24, n=8,waixenicin A-treated: 1.4 ± 0.13, n=8, p- cofilin/cofilin normalized to sham group expression: vehicle-treated: 0.6 ± 0.07, n=4, waixenicin A-treated: 0.9 ± 0.07, n=4).

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Figure 47: Biochemical assessment of signalling pathways affected by hypoxic-ischemic insult on neonatal brain in a pre- treatment paradigm. A) Changes in downstream signalling of CaMKII and calcineurin as detected by Western blot 6 hours after HI. Significant changes were detected in CaMKII, calmodulin, calcineurin B, p-CaMKII/CaMKII ratio and p-p38/p38 ratio. Alpha-actinin-1 levels showed an increasing trend, while p-cofilin/cofilin ratio showed a decreasing trend. B) Changes in downstream signalling of CaMKII and calcineurin as detected by Western blot 24 hours after HI Significant changes were detected in CaMKII, calcineurin B, p-CaMKII/CaMKII, p-p38/p38 ratio, alpha-actinin-1 levels and p-cofilin/cofilin ratio. Calmodulin levels showed an increasing trend. All data presented as mean±SEM. Statistical analysis: one-way ANOVA with Boferroni post-hoc, *p<0.05.

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7.3 Summary

Overall, in this chapter it’s been demonstrated that:

1. Neonatal HI causes proteomic changes in the ipsilateral hemisphere 6 hours after injury, and waixenicin A administration prior to injury attenuates these changes.

2. Significant changes in several proteins with calcium-binding and cytoskelon regulation functions were detected, including CaMKII, calcineurin and calmodulin.

3. TRPM7 may mediate cell death in HI injury through several calcium-dependent signaling pathways, including CaMKII/calcineurin-dependent regulation of cytoskeleton.

Overall, it’s been shown that waixenicin A may act through several signalling cascades to attenuate the HI brain injury to the neonatal brain. The proposed TRPM7-mediated signaling mechanism is discussed in detail in the next chapter.

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Chapter 5 Discussion and Conclusions

Contributions

Section 8.2 of this chapter contains a re-creation of the published material with the following citation:

Turlova et al., TRPM7 regulates axonal outgrowth and maturation of primary hippocampal neurons. Mol. Neurobiol. 2016. 53(1): 595-610.

Publication was recreated with permission and copyright license.

Section 8.5 of this chapter contains a re-creation of the manuscript in preparation for submission to Journal of Experimental Medicine with the following citation:

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8 Discussion 8.1 Summary of findings

This work provides insight into the role of TRPM7 activity in hippocampal neurons under normal and adverse conditions in vitro, and evaluates TRPM7 blocker waixenicin A for therapeutic potential in a model of neonatal hypoxic-ischemic injury. First, we determined the role of TRPM7 activity in neurite outgrowth and development in cultured hippocampal neurons under normal conditions. It was first confirmed that waixenicin A inhibited TRPM7-like currents in cultured hippocampal neurons with IC50 concentration in the nanomolar range. This result demonstrated the potency of waixenicin A in primary neurons. Since Ca2+ is a major regulator of neurite outgrowth, the effect of waixenicin A on Ca2+ influx through TRPM7 under Mg2+-free conditions was also investigated. It was also found that at the early stages of neuronal development (DIV2 – 3), blocking TRPM7 resulted in accelerated neuronal polarization and axonal specification. At the later stages in culture (DIV4 – 8), after the axon has been specified, blocking TRPM7 resulted in preferential enhancement of axonal outgrowth. Since both of these events are dependent on cytoskeletal dynamics at the neuronal growth cone, the interaction between TRPM7 and cytoskeletal proteins was examined. It was found that TRPM7 physically interacts with F-actin and α-actinin-1 and co-localizes with these proteins at the neuronal growth cone. Therefore, based on these findings, a TRPM7-mediated neurite outgrowth model has been proposed.

TRPM7’s biophysical characteristics, such as pH and reactive oxygen species sensitivity, make it a compelling target for investigation under the conditions of stress, such that occur in a variety of neurodegenerative disorders. Given our findings of TRPM7 activity’s role in neuronal development, we next characterized the effect of hypoxia on TRPM7 activity and neurite outgrowth with and without astrocytes present in culture. It was found that short-term hypoxia (~1 hour) downregulated TRPM7 activity and expression. Moreover, short-term hypoxia significantly enhanced neurite outgrowth at least in part through activation of ERK and Akt signaling pathways. Long-term hypoxia (2 hours and longer) caused a potentiation of TRPM7 activity as well as significant retraction of axons that was attenuated by TRPM7 block. Moreover, it was also shown that pharmacological activation of TRPM7 under hypoxic conditions exacerbated axonal retraction. The neurons grown in co-culture with astrocytes

133 exhibited more pronounced axonal retraction. Moreover, the effects of short-term hypoxia on axonal outgrowth and TRPM7 current were abolished under these conditions.

Finally, we evaluated a novel and specific blocker of TRPM7 channel waixenicin A for neuroprotective and therapeutic properties in a neonatal hypoxic-ischemic brain injury mouse model. The findings collectively demonstrated preventative and therapeutic effects of waixenicin A in a mouse model of neonatal HI. It was found that administration of waixenicin A as a pre- treatment and up to 1 hour after the hypoxia exposure, as a post-treatment, significantly reduced brain infarction volume compared to vehicle controls 24 hours after HI. On the contrary, it was found that administration of TRPM7 activator naltriben exacerbated the extent of brain injury 24 hours following HI. As well, waixenicin A administration reduced the long-term brain mass loss and preserved overall brain morphology up to 32 days following HI injury when compared to the vehicle group. Moreover, preservation of short-term and long-term function was shown in waixenicin A-treated group by improved performance in a battery of neurobehavioural tests involving motor, proprioceptive and memory functions when compared to vehicle-treated group. Overall, these findings showed that waixenicin A has preventative and therapeutic properties in a neonatal HI brain injury model in mice. Additionally proteomic and western blot analysis revealed that waixenicin A may attenuate brain injury after HI at least in part through Ca2+/calmodulin – dependent kinase (CaMKII) and phosphatase (calcineurin) regulation of cytoskeleton. Based on these findings, TRPM7-mediated mechanism of cell injury in HI was proposed and is discussed later in this chapter.

8.2 TRPM7 as a negative regulator of axonal outgrowth during development

This work showed for the first time that TRPM7, a calcium-permeable non-selective cation channel, participates in regulating neurite outgrowth and maturation, with preferential effect on axonal growth, through cytoskeletal regulation at the growth cone. We showed that block by waixenicin A promotes neurite outgrowth in embryonic hippocampal neuronal culture. We also demonstrated TRPM7 expression pattern in neuronal growth cones and propose a model for TRPM7-mediated axonal outgrowth thorough cytoskeleton regulation.

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8.2.1 TRPM7 expression in hippocampal neurons suggests a developmental function

Several studies demonstrated TRPM7 expression patterns in different types of mammalian neurons in vitro. In mouse and rat hippocampal neurons, TRPM7 showed a diffused expression pattern in the cell bodies and processes (Wei et al., 2007; Sun et al., 2009). In hippocampal neurons, broader TRPM7 expression correlates well with its role in divalent ion homeostasis (Wei et al., 2007) and cell survival (Sun et al., 2009). In this study, we showed that TRPM7 is expressed in the growth cones of mouse hippocampal neurons. Since the growth cone is a key structure responsible for axonal navigation in developing nervous system, TRPM7 expression patterns shown in this study suggests a specific developmental role.

8.2.2 TRPM7 maintains proper axonal development by inhibiting excessive growth and premature axonal specification

Neuronal differentiation is essential to nervous system development and function. Development of cultured hippocampal neurons has been characterized and separated into several stages, starting from neuronal sprouting followed by axonal specification and elongation (Dotti et al., 1988). Growth cones, highly dynamic structures at the tips of elongating neurites, act as “vehicles” which move the protruding neurites towards their ultimate targets (Lowery and Van, 2009).

Here we utilized a pharmacological approach to study the role of TRPM7 channels in neuronal development. We report that blocking TRPM7 conductance by waixenicin A promoted axonal outgrowth and enhanced axonal specification by enhancing neuronal polarization. We also report that blockade of TRPM7 enhances axonal branching. Our data describe that under physiological conditions TRPM7 channel functions as an inhibitor of axonal outgrowth and maturation. As these processes are crucial for proper target finding and formation of functional neuronal networks, they must be tightly regulated to avoid overgrowth and connection with unwanted targets. Indeed, postmortem analysis of brain tissue from individuals with autism showed excessive axonal branching in several prefrontal areas associated with attention, emotions and social interactions (Zikopoulos and Barbas, 2010). In fact, the maintenance of axonal morphology is an active process that requires constant suppression of sprouting through limiting actin polymerization along the neurite shaft by calpain proteases (Mingorance-Le and O'Connor,

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2009). Thus, TRPM7 proteins represent one of the potential mechanisms for regulation of proper axonal outgrowth.

8.2.3 TRPM7 regulates axonal outgrowth through calcium influx In this study we also showed that TRPM7 block by waixenicin A causes a significant decrease in calcium influx (58 ± 8% decrease). Taken together with our findings on the effect of waixenicin A on neurite outgrowth, these results suggest that calcium influx through TRPM7 plays an important role in neurite outgrowth regulation, as cytoskeletal dynamics at the growth cone are tightly controlled by local intracellular calcium levels (Montell, 2005; Lohmann et al., 2005; Gomez and Spitzer, 2000), which in turn regulates neurite elongation and motility (Mattson and Kater, 1987; Henley and Poo, 2004). Nevertheless, the role of TRPM7 as a scaffolding protein that brings together cytoskeletal proteins and the effector proteins that regulate neurite outgrowth cannot be ruled out in this study.

Moreover, previous study has also shown that magnesium influx through TRPM7 channels can modulate cell proliferation in DT40 B cells (Nadler et al., 2001). As magnesium is an important regulator of the cytoskeleton (Prescott et al., 1988), it is possible that magnesium influx through TRPM7 contributes to TRPM7-mediated neurite outgrowth. Under low magnesium conditions, TRPM7 via its C-terminal kinase domain (Krapivinsky et al., 2014) enhances phosphorylation of eukaryotic elongation factor 2 kinase (eEF2K) (Perraud et al., 2011). Phosphorylation of eEF2 inhibits protein synthesis at developing synapse (Scheetz et al., 2000). The increase in axonal outgrowth by TRPM7 inhibition could be through suppression of eEF2K phosphorylation. Our findings support the notion of TRPM7 functions as a negative regulator to mediate optimal axonal outgrowth. However, a significant cytoskeletal rearrangement at the growth cone or change in expression of cytoskeleton-mediating proteins due to inhibitionTRPM7 of activity cannot be ruled out as potential reasons for observed axonal outgrowth. Therefore, these mechanisms need to be investigated in future studies.

8.2.4 TRPM7 may serve as mechanosensor during neuronal development

The regulatory role of TRPM7 in neurite elongation can be explained by mechanosensitive nature of the channel (Wei et al., 2009). Stretch-activated channels (SACs) were long thought to be candidates for regulating growth cone motility in neurons (Sigurdson and Morris, 1989;

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Jacques-Fricke et al., 2006). It was proposed that SACs regulate neurite outgrowth by mediating local calcium dynamics within specific growth cone microdomains (Jacques-Fricke et al., 2006). These microdomains are functionally linked to specific calcium-sensitive effectors that regulate cytoskeletal rearrangement at the growth cone (Jacques-Fricke et al., 2006). Blocking these stretch-activated channels accelerates axonal outgrowth, however, their identity remains unknown (Jacques-Fricke et al., 2006). Previous study with migrating embryonic lung fibroblasts has shown that TRPM7 transduces the local mechanical stress into an intracellular calcium signals in the form of calcium flickers at the leading end of migrating cells (Wei et al., 2009). Our results further support the hypothesis of TRPM7 channel function as a mechanosensitive regulator of neuronal cytoskeleton. Therefore, it is conceivable that the extent of growth cone protrusion and neurite elongation would depend on a ratio of open to closed TRPM7 channels.

8.2.5 A model for TRPM7-mediated axonal outgrowth via interactions with actin and actinin

Actin dynamics at the elongating growth cone are locally and tightly regulated by actin- mediating proteins in response to guidance cues, adhesive/repulsive substrate encounters and calcium fluxes (Gomez and Letourneau, 2014). This regulation of actin assembly, turnover and interactions with the membrane and cytoskeleton are critical in generating the traction and pulling force needed to protrude the elongating neurite towards its target (Gomez and Letourneau, 2014). Here, we showed that TRPM7 interacts and colocalizes with the cytoskeletal protein actin, and actin-binding protein α-actinin-1. Our data support the notion that TRPM7 mediates axonal outgrowth via actin and α-actinin-1 protein complex, or a TRPM7-mediated protein microdomain in the growth cone. This microdomain regulates actin anchoring by α- actinin-1 and calcium influx through TRPM7. Actin anchoring affects actin polymerization, which is crucial in filopodial protrusion and growth cone motility (Forscher and Smith, 1988). Suppressing TRPM7 activity by waixenicin A would allow for greater interaction between F- actin and α-actinin-1, thus enhancing growth cone filopodial protrusion and leading to increased neurite outgrowth, branch formation and axonal differentiation. On the other hand, potentiation of TRPM7 current would cause dissociation of F-actin from the membrane leading to axonal degeneration and growth cone collapse (Figure 48).

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Figure 48: A model of TRPM7-mediated neurite outgrowth under normal and adverse conditions.

8.3 TRPM7’s role in axonal retraction under pathophysiological conditions.

Several lines of evidence indicate that TRPM7 activity becomes potentiated under the conditions of stress. More specifically, conditions such as low extracellular pH (Jiang et al., 2005), ROS production (Aarts et al., 2003; Coombes et al., 2011), changes in divalent cations (Wei et al., 2007) and intracellular Zn2+ accumulation (Inoue et al., 2010) have all been shown to enhance TRPM7 activity. Given our findings that TRPM7 may act as a negative regulator of axonal extension and development, it is reasonable to hypothesize that TRPM7 may contribute to axonal retraction and degeneration under the conditions of stress. Axons are particularly vulnerable to injury and disease, and axonal damage is a hallmark of many neurodegenerative conditions. Many neurological disorders, such as Alzheimer’s disease, Parkinson’s disease, multiple sclerosis, stroke and traumatic brain injury exhibit axonal degeneration (Lingor et al., 2012). Compared with axonal outgrowth and development, relatively little is known about the molecular mechanisms of axonal retraction. Since axonal retraction is a change in cell shape, it is reasonable to assume that it is based on modifications of cytoskeleton.

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8.3.1 TRPM7 and cytoskeleton in axonal retraction

Several studies implicated the integrity of the cytoskeleton in maintaining axonal stability. More specifically, abnormalities in F-actin dynamics have been shown to contribute to axonal dysfunction. Multiple actin regulators, including Rho-associated kinases (ROCKs) (Saal et al., 2015), actin-depolymerizing factor/cofilin (Heredia et al., 2006), and others have been reported to play a role in a range of axonal retraction/degeneration conditions. Early modulation of F- actin was observed in transected axons of primary hippocampal neurons and phosphorylated cofilin was shown to be enriched in their soluble axoplasm (Garland et al., 2012). Moreover, destabilizing actin cytoskeleton with cytochalasin D in the axonal compartment is sufficient to generate the same response as axonal injury in mouse dorsal root ganglion neurons (Valakh et al., 2015). The high ATP demand for the maintenance of dynamic F-actin organization in axons makes it possible for ischemic/hypoxic conditions to disrupt actin organization with subsequent impairment in axonal integrity, ion channel function, organization of neurotransmitter vesicles and synaptic communication.

The potential role of TRPM7 in axonal retraction is twofold. First, Ca2+ influx through TRPM7 into the axon under the conditions of stress may trigger several processes leading to axonal retraction. First, Ca2+ ions decrease interaction between F-actin and α-actinin-1 (Burridge and Feramisco, 1981) which, according to our findings, may be closely associated with TRPM7 in neuronal growth cones (Turlova et al., 2014). Decreased cross-linking of actin by α-actinin could in turn lead to instability of F-actin, leading to growth cone collapse.

Second, TRPM7 has been shown to undergo caspase-mediated cleavage that results in a functional membrane channel and releases the kinase domain to translocate to the nucleus and participate in chromatin remodeling under the conditions of stress (Krapivinsky et al., 2014). Moreover, this cleavage was shown to potentiate TRPM7 current (Desai et al., 2012), which can further contribute to disruption of F-actin organization. Additionally, TRPM7 kinase domain was previously linked to regulation of actomyosin contractility and cell adhesion (Clark et al., 2006). In neuroblastoma cell line, overexpression of wild-type TRPM7, but not kinase-dead TRPM7, leads to phosphorylation and inhibition of myosin IIA (Clark et al., 2006). During axonal retraction, myosin II contractility plays a major role in shortening of the axon. Actomyosin tension that is generated by active myosin II leads to rearrangement of F-actin bundles into intra-

139 axonal F-actin bundles and loss of F-actin in growth cone lamellipodia and filopodia. This allows for coordinated and effective shortening of the axon (Gallo, 2006). Since TRPM7 kinase has been shown to inhibit actomyosin contractility, cleavage of TRPM7 following the onset of the stress conditions may lead to decrease in myosin II phosphorylation, thus enhancing tensile force generated by actomyosin and resulting in axonal shortening. Overall, it’s possible that TRPM7 cation conductance and kinase activity both contribute to axonal retraction/degeneration under injurious conditions.

8.3.2 Axonal injury in hypoxia-ischemia

Hypoxic-ischemic brain injury causes a rapid and significant loss of axons that continues long after the onset of injury. It has been estimated that for every minute that ischemic injury to the brain goes untreated, approximately 7 miles of axons are lost (Saver, 2006). It has been proposed that there are three distinct stages of axonal degeneration following HI injury: primary acute phase, secondary progressive degeneration phase and third degenerative/regenerative phase (Hinman, 2014).The first phase of axonal injury affects the axonal tracts in the ischemic core, where they undergo catastrophic loss of energy and intra-axonal Ca2+ influx. This ionic influx destabilizes axolemma, activates proteases and leads to breakdown of structural components (Stys and Steffensen, 1996).

Following this acute phase, the axons located in ischemic penumbra undergo continued retraction/degeneration over the following days. This is known as second progressive phase of axonal injury (Zhang et al., 2012). The molecular mechanisms that drive this degeneration are still being investigated. Few studies investigated the effect of HI on axonal organization and integrity. In a model of mild cerebral hypoperfusion, structural integrity of axons was compromised resulting in axonal damage (Reimer et al., 2011).

From our findings, it’s possible that TRPM7 participates and facilitates the second progressive degeneration phase that occurs in peri-infarct region. The surviving neurons in the ischemic penumbra experience progressive energy loss, and there is an accumulation of extracellular protons and ROS due to hypoperfusion of the region. These conditions potentiate TRPM7 activity and facilitate the retraction of the axons though cytoskeletal mechanisms described previously. We demonstrated that neurons exhibit progressive retraction of axons when exposed to hypoxic conditions for longer than 2 hours. Moreover, we also found that neurons exposed to

140 prolonged hypoxia exhibit reduced axonal lengths 24 hours after being returned to normal conditions, suggesting that axonal retraction may persist even during the recovery phase. TRPM7 inhibition by waixenicin A attenuated axonal retraction caused by prolonged hypoxia for up to 6 hours, suggesting that axonal degeneration may be delayed by TRPM7 block. Additionally, when administered after prolonged hypoxia and immediately before return to normal conditions, TRPM7 block was able to prevent further axonal retraction. These findings suggest that TRPM7 may be one of the regulators of axonal stability following HI and contribute to the observed progressive degeneration of axons.

8.3.3 TRPM7 downregulation by short-term hypoxia

An interesting observation that we came across during characterization of the effect of hypoxia on neurite outgrowth and TRPM7 activity is that short-term hypoxia (no longer than 1 hour) caused a decrease in TRPM7 activity and protein level. Moreover, as consistent with our previous findings of regulation of axonal outgrowth by TRPM7, short-term hypoxia enhanced axonal outgrowth.

TRPM7 protein level downregulation under short-term hypoxic conditions can be explained by activation of TrkA pathway by nerve growth factor (NGF). Constitutive production of NGF has been previously demonstrated in hippocampal neuronal cultures (Di et al., 2000). When these cultures were exposed to mild hypoxia together with a neutralizing antibody against interleukin- 1β (IL-1β), NGF production was shown to be upregulated compared to normoxic neurons (Di et al., 2000). Activation of TrkA by NGF is widely accepted to have neuroprotective properties in cerebral ischemia (Luk et al., 2004; Yang et al., 2011a). Once activated, TrkA activates three main pathways: PI3K/Akt, MEK/ERK and PLC-γ (Kaplan and Miller, 1997). It has been shown that NGF can stabilize neuronal calcium homeostasis and protect neurons against insults. It was demonstrated that in the model of middle cerebral artery occlusion (MCAO), TRPM7 expression was significantly upregulated in the ipsilateral hippocampus, and high levels of TRPM7 were maintained for up to 30 hours after injury. Intracerebroventricular administration of NGF significantly reduced TRPM7 expression (Jiang et al., 2008). These findings were confirmed in hippocampal neurons in vitro that were subjected to oxygen-glucose deprivation. Interestingly, co-administration of wortmannin, PI3K inhibitor, abolished NGF effect on TRPM7 expression, suggesting that NGF potentially regulates TRPM7 expression through PI3K pathway (Jiang et

141 al., 2008). In our results, we observed upregulation of PI3K/Akt and MEK/ERK signaling pathways concomitant with downregulation of TRPM7 expression, which is in agreement with previous findings of NGF-mediated TRPM7 downregulation through TrkA activation.

Further studies will be needed in the future to confirm the involvement of NGF in TRPM7 expression under hypoxic conditions.

8.3.4 The role of astrocytes in axonal retraction under hypoxic conditions.

The functions of astrocytes in neuronal survival during hypoxia/ischemia are still under debate, and astrocytes have been shown to serve both protective and damaging roles. In our study, the presence of astrocytes in the culture accelerated axonal retraction/degeneration, suggesting a detrimental effect. One of the critical roles of astrocytes in the brain is their ability to uptake excess glutamate from extracellular space. It has been postulated that the developing brain is more vulnerable to excitotoxicity than the adult brain because of the reduced glutamate uptake capacity that may be at least in part due to low expression of astrocytic glutamate transporters (Morken et al., 2014), glutamate aspartate transporter (GLAST) and glutamate transporter-1 (GLT-1) (Furuta et al., 1997). As our co-cultures were derived from embryonic mice, it is conceivable that the astrocytes in these cultures have a limited capacity of glutamate uptake. In fact it has been shown that in astrocytes co-cultured with neurons, the expression of GLAST and GLT-1 was low at early stages in culture and only began to increase after 10 days of co-culture (Perego et al., 2000). Under pathophysiological conditions astrocyte-expressed transporters have been shown to operate in reverse, thus contributing to excessive glutamate and other excitatory neurotransmitters in the extracellular space (Malarkey and Parpura, 2008). Therefore, it is possible that the extracellular glutamate concentration in co-cultures is higher than in neuronal cultures, since glutamate is released from both types of cells. Additionally, excessive glutamate has been demonstrated to induce rapid axonal degeneration in cortical neuronal cultures (Chung et al., 2005). Therefore, it would be important to investigate and compare the extracellular concentrations of excitatory neurotransmitters in neuronal cultures and co-cultures.

Additionally, hypoxia was shown to induce transcriptional activation of IL-1β in human and mouse astrocytes (Zhang et al., 2006b). IL-1β production may contribute to the detrimental effects of hypoxia that we observed in co-cultures. A previous study also linked IL-1β to

142 activation of intraneuronal inflammosome and axonal degeneration in rodent and human CNS neurons (Kaushal et al., 2015). Interestingly, a recent study demonstrated that IL-6, another cytokine produced under hypoxic-ischemic conditions in the brain, negatively regulated TRPM7 activity in cultured cortical neurons (Liu et al., 2016). This finding suggested that IL-6 may promote cell survival during HI injury by inhibiting TRPM7 activity (Liu et al., 2016). Another study demonstrated that in cultured hypoxic neurons immunoneutralization of IL-1β resulted in upregulation of IL-6 production, suggesting that IL-1β may negatively regulate IL-6 production (Di et al., 2003). Therefore, it is conceivable that TRPM7 activity may be regulated by the levels of IL-1β and IL-6 present in the culture. Enhanced production of IL-1β by astrocytes in co- cultures may lead to decrease in IL-6 production, which usually negatively regulates TRPM7 activity, thus leading to potentiation of TRPM7 activity, as we observed in our study.

Together, the factors discussed in this section could lead to exacerbation of axonal retraction, as observed in our study. However, future in-depth studies are necessary to better understand the interplay between factors produced by astrocytes, their effect on TRPM7 activity and axonal retraction.

8.4 TRPM7 in neonatal hypoxic-ischemic brain injury.

Here, we evaluated a novel and specific blocker of TRPM7 channel waixenicin A as a potential treatment in a neonatal HI brain injury mouse model. Our data collectively demonstrate strong preventative and therapeutic effects of waixenicin A in our model of neonatal HI. We found that administration of waixenicin A as a pre-treatment and up to 1 hour after the hypoxia exposure, as a post-treatment, significantly reduced brain infarction volumes compared to vehicle controls 24 hours after HI. On the contrary, we found that administration of TRPM7 activator naltriben exacerbated the extent of brain injury 24 hours following HI. As well, we found that waixenicin A administration reduced the long-term brain mass loss and preserved overall brain morphology up to 32 days following HI injury as compared to the vehicle group, as well as preserved short- term and long-term function. These data show that waixenicin A has neuroprotective and therapeutic properties in a neonatal hypoxic-ischemic brain injury model in mice.

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8.4.1 Suitability of the neonatal HI model

Neonatal HI is the most common cause of death and disability in human neonates. It accounts for 23% of infant mortality worldwide and affects over 1 million infants annually (Lawn et al., 2005). Neonatal HI remains a significant cause of mortality and disability in the developed countries where the incidence of the condition has not decreased in the past two decades (Himmelmann et al., 2005; Vincer et al., 2006). Clinically, neonatal HI injury is defined as asphyxia of the umbilical blood supply to the human fetus which occurs at 36 gestational weeks or later (Perlman, 1997; Volpe, 2012; Shah et al., 2006). The rates of disability remain high throughout life for survivors of neonatal HI, with 5-10% of infants exhibiting motor deficits and 20-50% displaying sensory and cognitive abnormalities (Hack et al., 1992; Volpe, 2012; Lee et al., 2013). Other disorders, including seizures, hearing and vision loss, language abnormalities, microcephaly and other abnormal neurological signs have been reported (Robertson and Finer, 1985; Shankaran et al., 1991; Shankaran et al., 2012).

Models that intend to replicate clinical symptoms of neonatal HI have been extensively described in rodents (mice and rats) (Hagberg et al., 2002; Dean et al., 2015), with few researchers studying larger animals such as sheep, pigs and primates (Roohey et al., 1997; Dean et al., 2015; Volpe, 2012). The Rice-Vannucci model, one of the most published models of neonatal HI, was used in our study (Rice, III et al., 1981). The wide spread usage of the model allows for comparisons with many published studies. It has been chosen due to several distinct advantages, which are described below. First, this model parallels the anatomical damage seen in human neonates. The gray matter injury has been observed in the cortical regions, hippocampus, thalamus and basal ganglia (Rice, III et al., 1981; Andine et al., 1990; Vannucci et al., 1999). White matter injury has also been reported, with the severity of the injury correlating with the duration of hypoxia (Liu et al., 2002). Metabolically, decreased cerebral blood flow and glucose uptake, brain acidosis and inflammation have been demonstrated (Vannucci et al., 1988; Yager et al., 1991; Vannucci et al., 1989; Bona et al., 1999).

Second, this model gives rise to well-documented behavioural phenotypes, including impaired motor and balance functions, sensory abnormalities, and cognitive abnormalities such as memory and attention deficits (Balduini et al., 2000; Balduini et al., 2001; Park et al., 2015; Buwalda et

144 al., 1995). Long-term behavioural characterization also mimics the outcomes of the neonatal HI in humans (Lubics et al., 2005).

However, several drawbacks of the model must also be mentioned. First, the invasiveness of the procedure (severing of the common carotid artery) does not replicate the nature of the injury that occurs in human neonates (Hill, 1991). Second, there is high variability in size and severity of injury due to differences in experimenters, conditions and genetic background of animals, making a direct comparison more difficult (Vannucci and Hagberg, 2004). However, the advantages of this model outweigh the drawbacks, thus making it a suitable model for our study.

8.4.2 Long-term effects of waixenicin A treatment

Several studies have shown that ischemic injury is an evolving process with neuronal loss being detected as late as 14 days after the onset of injury (Li et al., 1995; Du et al., 1996). Indeed, the post-injury mild hypothermia in a rodent model of neonatal HI was shown to reduce brain injury when evaluated several days post-HI, but did not show protection when the evaluation was performed one month post-HI, suggesting that the onset of injury was delayed rather than prevented (Trescher et al., 1997). Therefore, the long-term neurological outcome is one of the most important assessments of a neuroprotective method. The extent of neonatal HI injury in rodents has been reported to span cortical (Zhao et al., 2007) and hippocampal (Zhao et al., 2007) areas, perirhinal cortex (Zhao et al., 2007) and striatum (Bouet et al., 2007). Here we show that neonatal HI causes long-term functional deficits four weeks after injury and that waixenicin A alleviates these deficits. Waixenicin A reduces damage to higher cortical areas including parietal cortex (Bouet et al., 2007) as evident from improved motor and coordination performances of mice on Accelerated Rotarod Test. A better performance on novel object recognition task in waixenicin A-treated group suggests reduced damage to hippocampal and parahippocampal brain regions, such as perirhinal cortex (Hammond et al., 2004; Antunes and Biala, 2012). Passive avoidance behaviour has been shown to require the function of perirhinal (Burwell et al., 2004), parietal (Hogg et al., 1998) and frontal (Ostrovskaya et al., 1999) cortices, as well as striatum (Sandberg et al., 1984) and basal forebrain (Torres et al., 1994) and was significantly improved in waixenicin A-treated mice in our study. Our data collectively suggest that waixenicin A alleviates injury at least in part to the affected brain regions.

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8.4.3 Waixenicin A is a promising therapeutic target for drug development for neonatal HI

Despite the efforts in scientific and clinical research, the development of therapeutic strategies for neonatal HI faces numerous challenges, including drug delivery properties, dosage and use in neonates. Waixenicin A is a worthwhile target for further investigation and potential drug development for this condition for several reasons discussed below.

8.4.3.1 Limitations to CNS drug delivery

One of the challenges faced by CNS drug development is the ability of the compound to cross the blood brain barrier (BBB) (Alavijeh et al., 2005). Therefore, CNS drugs are limited to lipid- soluble and low molecular weight (below 400 kDa) compounds, which hinders the process of developing effective CNS therapeutics (Pardridge, 2005). According to a novel central nervous system multiparameter optimization (CNS MPO) algorithm analysis done by our collaborators, waixenicin A achieved the score of 4.5 (unpublished data not shown in this thesis), which highly predicts BBB penetration (MPO desirability score >4, scale 0-6). This analysis was created as a tool for prospective drug design and focuses on physicochemical properties used by medicinal chemists, such as lipophilicity, molecular weight, polar surface area and others (Wager et al., 2010). These findings suggest that waixenicin A is a desirable candidate for further testing for neonatal HI and other conditions of the CNS.

8.4.3.2 Limitations to drug dosage for neonates

In the case of pediatric patients, clinical translation for compounds is more complicated when compared to adult patients, and most compounds are approved only for adult use. The pediatric patient population is highly variable, and therefore extrapolating dosages and predicting adverse reactions can be extremely difficult (Turner, 2011; Dabliz and Levine, 2012). The advantage of waixenicin A, as presented in the current thesis work, is the low dose that is required to achieve significant reduction of brain damage (nanomolar range). Therefore, taken together with desirable BBB permeability, it is more likely that an effective and low-range dose can be found for pediatric patients in the future.

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8.4.3.3 Limitations to timing of drug delivery

Although hypothermia has been established as the standard of care for neonatal HI, the protection it provides is incomplete. Therefore, there is an urgent need for adjuvant neuroprotective therapies (Johnston et al., 2011). An advantage of waixenicin A that has been observed in the current study is that it provides protection for up to one hour following HI in our model. While this effect has to be confirmed in different animal models of neonatal HI, it could potentially provide some flexibility in terms of the timing of treatment administration. Drug administration before or during hypothermia may have adverse effects on liver and kidney clearance of the drug, or even initiate degeneration in the brain, thus counteracting the neuroprotective effects of hypothermia (Cornette, 2012).Therefore, future studies are needed to examine the best combination of hypothermia and waixenicin A administration to provide long- lasting synergistic protection and achieve the best possible outcome for the patients.

8.5 TRPM7-mediated molecular mechanisms in HI brain injury

8.5.1 Proteomic analysis of HI brain injury

We have investigated proteomic changes in the brain following HI injury in vehicle-treated and waixenicin A-treated groups. Proteomic analyses following HI injury have been previously performed in several rodent models, and our findings were generally in agreement with published literature (Rosenkranz et al., 2012; Shao et al., 2017; Hu et al., 2006; Yang et al., 2014).

Several studies identified groups of proteins involved in ion homeostasis, energy metabolism, enzyme regulation and cytoskeletal and structural components (Yang et al., 2014; Shao et al., 2017; Hu et al., 2006). Generally, our findings are in agreement with these studies, as we found groups of proteins with similar functions differentially expressed in our model as well.

A study by Rosenkranz and colleagues identified several proteins that were significantly affected following neonatal HI in Wistar rats. One of those proteins was Coronin-A, an actin-biding protein that was also identified as an immunohistochemical marker for microglial cells. Upregulation of this protein in the lesioned hemisphere was reflective of an inflammatory response. Glial fibrillary acidic protein (GFAP) was another highly upregulated protein identified in this study. Since GFAP is present predominantly in astrocytes, the authors postulated it to be

147 reflective of astrogliosis following HI. Finally, calcineurin, a Ca2+/calmodulin-dependent phosphatase was also upregulated in the injured hemisphere (Rosenkranz et al., 2012). This particular result is in agreement with our data, and will be discussed in detail in the next section.

8.5.2 TRPM7-mediated regulation of cytoskeleton in HI

In this study, we focused on three proteins, calmodulin, calcineurin and CaMKII that were differentially expressed in the vehicle-treated group after HI. Their differential expression was also attenuated in waixenicin A-treated groups. We focused on these proteins because they were shown to be highly expressed at the growth cone and regulate neurite outgrowth through upstream regulation of actin cytoskeleton. To build upon our previously identified TRPM7- cytoskeleton molecular complex, we aimed to investigate if changes in actin cytoskeleton regulation were present in our model. Further analysis of the downstream signaling of these proteins revealed that waixenicin A acts potentially on several actin cytoskeleton-regulating proteins, including p38, cofilin and α-actinin-1. Actin cytoskeleton has been reported to play a role in cell death under hypoxic conditions (Posmantur et al., 1996; Posmantur et al., 1997; Kampfl et al., 1996; Endres et al., 1999).

CaMKII, one of the most abundant protein kinases in the nervous system, plays a key role in synaptogenesis during development (Rongo and Kaplan, 1999). Our findings show a drastic depletion of CaMKII following HI, and a restoration of its expression following waixenicin A treatment, as supported by a previous a report (Tang et al., 2004). Here we also reported that TRPM7 modulates actin cytoskeleton through interaction with α-actinin-1. Ca2+ influx via N- methyl-D-aspartate (NMDA)-type glutamate receptors has long been considered as a hallmark of excitotoxic neuronal injury during ischemic insults (Sattler and Tymianski, 2001; Chen et al., 2008). Binding of α-actinin to NMDA was shown to enhance activity of the receptor, while CAMKII binding attenuated its activity (Leonard et al., 2002). Moreover, actinin has been shown to play a role in transport of -amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA) receptors, well-known contributors to neuronal injury during stroke (Soundarapandian et al., 2005), to dendritic spines (Schulz et al., 2004). Additionally, CaMKII was shown to inhibit TRPM7 currents in hepatocytes (Mishra et al., 2009); however, this mechanism is yet to be confirmed in neurons. Together, these findings suggest that upregulation of actinin and downregulation of CaMKII may contribute to neuronal death and potential increase in TRPM7

148 activity and leave neurons vulnerable to further injurious stimuli. Waixenicin A treatment reduced actinin upregulation and restored CaMKII levels thus conferring neuroprotection.

Calcineurin, Ca2+/calmodulin-regulated phosphatase, has also been implicated in neuronal cell death under oxidative stress conditions (Ankarcrona et al., 1996; See and Loeffler, 2001; Asai et al., 1999). In our study, we observed upregulation of calcineurin B (regulatory subunit) and calmodulin, suggesting an increase in calcineurin activity following HI. Waixenicin A treatment restored the levels of both proteins to sham levels. Similarly, activation of cofilin or actin depolymerizing factor (ADF) via dephosphorylation (Suzuki et al., 1995) has been linked to neuronal death under oxidative stress conditions, as well as blood-brain barrier disruption (Alhadidi et al., 2016; Alhadidi et al., 2017). It has been previously shown that CaMKII and calcineurin control cofilin activity via phosphorylation and dephosphorylation, respectively, through Ca2+-dependent regulation of LIM-kinase1and phosphatase Slingshot-1L (Zhao et al., 2012). In our study, we observed a significant decrease in p-cofilin following HI injury, suggesting its upstream activation by calcineurin, while waixenicin A restored p-cofilin levels, potentially through restoring CaMKII expression and activity.

The role of p38 mitogen-activated protein kinase (p38 MAPK) in HI brain injury has been controversial. While it has been reported that inhibition of p38 MAPK is neuroprotective against HI injury (Hee et al., 2002), another report suggested that activation of p38 MAPK reduced apoptotic cell death under HI conditions (Pfeilschifter et al., 2010). While it is possible that conflicting reports are due to differences in animal models and experimental conditions, in our model of neonatal HI we observed a drastic decrease in p38 MAPK activation following HI, which was restored by waixenicin A treatment. Interestingly, p38 activation was previously implicated in downstream reorganization of actin cytoskeleton in endothelial cells (Nguyen et al., 2004) and neurons (Correa and Eales, 2012), and CaMKII was shown to be a potential activator of p38 (Nguyen et al., 2004). A proposed signaling cascade of TRPM7-mediated neuronal cell death in neonatal HI is illustrated in Figure 49. Together, our findings show that waixenicin A could reduce brain injury through upstream regulation of actin cytoskeleton, potentially affecting both neuronal and non-neuronal cells.

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Figure 49: Proposed TRPM7-mediated mechanism of neuronal cell death in neonatal HI.

8.6 Limitations of the current work

8.6.1 The role of magnesium in neurite outgrowth

This thesis work shows that ion conductance through TRPM7 is sufficient to affect axonal outgrowth process. While Ca2+ is a well known regulator of axonal outgrowth, this is not the only ion that is conducted by TRPM7. Therefore, the role of Mg2+ influx through TRPM7 in axonal outgrowth and cytoskeletal regulation cannot be ruled out. Mg2+ has been shown to be important for cytoskeleton integrity, and elevated intercellular Mg2+ concentrations were shown to lead to disassembly of microtubules (Prescott et al., 1988). It has also been shown that neurite formation and extension in response to NGF treatment in PC12 cells is Mg2+-dependent (Koike, 1983). Similarly, a strong substrate attachment of these cells and subsequent neurite extension were shown to be heavily dependent on the presence of Mg2+ ions in the media (Turner et al., 1987). Since TRPM7 has been shown to be essential for control of cellular Mg2+ homeostasis (Ryazanova et al., 2010), further investigation of TRPM7 role in axonal outgrowth under varying Mg2+ concentration should be carried out in the future.

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8.6.2 PKC-dependent signaling in TRPM7-mediated neurite outgrowth

Protein kinase C (PKC) is a well-established regulator of neurite outgrowth (Qian et al., 1994; Kolkova et al., 2000; Fagerstrom et al., 1996; Brodie et al., 1999). PKC has a broad range of targets, including several ion channels (Shearman et al., 1989) and regulates neurite outgrowth through PKC-dependent (Brodie et al., 1999; Fagerstrom et al., 1996; Kolkova et al., 2000; Miloso et al., 2004a; Tsuneishi, 1992; Auer et al., 2012), Raf/MEK/ERK (Miloso et al., 2004b; Kolkova et al., 2000), PI3K/Akt (Auer et al., 2012) and other mechanisms. TRPM7 C terminus contains several potential phosphorylation sites (Kim et al., 2009). We found that GST-fusion protein of TRPM7 C-terminus co-immunoprecipitates with PKC in mouse hippocampal tissue. Moreover, we found that broad PKC inhibitors abolished the increase in axonal outgrowth when applied together with waixenicin A. These findings are shown in Appendix 10. Therefore, it cannot be ruled out that TRPM7 may also affect axonal outgrowth through a PKC-dependent signaling mechanism or that under certain conditions TRPM7 activity may be affected by PKC phosphorylation. The role of TRPM7-PKC complex in axonal outgrowth must be investigated in depth in the future.

8.6.3 Naltriben as TRPM7 activator and its potential off-target effects

In our study, we used naltriben mesylate to confirm the role of TRPM7 activity in axonal outgrowth in vitro and show that administration of naltriben in vivo exacerbated brain damage after HI. Naltriben was previously characterized as a moderately potent (EC50 of 20 μM) TRPM7 agonist with no effects on other related TRP channels (TRPM2, TRPM3 and TRPM8) at the concentrations used in our study (up to 50 μM) (Hofmann et al., 2014). Additionally, naltriben displayed other favourable characteristics, such as being completely washed out, reversible and able to induce endogenous TRPM7 currents in RBL-1 cells (Hofmann et al., 2014). In our hands, naltriben was also able to activate endogenous TRPM7-like currents in cultured hippocampal neurons.

Naltriben was also shown to be a potent δ, μ and ĸ-opioid receptor antagonist with IC50 in the nanomolar range, with in vivo selectivity for δ2-receptor (Miyamoto et al., 1993; Portoghese et al., 1991; Thorat and Hammond, 1997). Both μ and δ opioid receptors have been shown to be expressed in rodent cultured hippocampal neurons (Guo et al., 2015). Activation of μ-opioid receptors was shown to inhibit conotoxin-sensitive and conotoxin-insensitive low-threshold (T-

151 type) high-threshold Ca2+ currents in cultured neurons (Schroeder et al., 1991). While activation of these receptors in hippocampal cultures under normal conditions has not been explicitly demonstrated, endogenous opioid receptor agonists, such as dynorphin and enkephalin, were shown to be produced in hippocampal cultures in vitro (He et al., 1992). Therefore, it is possible that using naltriben in culture would inhibit the receptors present on these neurons, thus dis- inhibiting Ca2+ transients and contributing to Ca2+ influx observed in our experiments. Therefore, experiments to determine the relative contribution of TRPM7 activation and opioid-receptor inhibition to Ca2+ influx in hippocampal neurons need to be conducted in the future.

Additionally, opioid receptor activation has been shown to be involved in neuroprotection mediated by TRPV1 activation in the rat retina against excitotoxicity (Sakamoto et al., 2017). Similarly, activation of δ-opioid receptors in cultured cortical neurons significantly reduced cell injury when neurons were exposed to 1% hypoxia (Zhang et al., 2002). Moreover, δ-opioid receptor activation attenuated brain injury in a model of cerebral ischemia with middle cerebral artery occlusion in a rat (Yang et al., 2009). Therefore, the exacerbation of brain injury by naltriben in our model can be at least in part through inhibition of opioid receptors. The relative contributions of TRPM7 activation and opioid receptor inhibition by naltriben to hypoxic- ischemic brain injury in our model need to be confirmed in the future.

8.6.4 Systemic administration of waixenicin A in vivo

In the present work, waixenicin A was administered via an intraperitoneal injection as opposed to local injection into a specific brain region. However, this type of systemic administration may result in suboptimal drug concentration being achieved in the CNS, thus leading to incorrect conclusions about drug efficacy. While this type of drug administration in the case of waixenicin A has proven to be effective in reducing brain infarction, we could not reliably estimate the concentration of compound in the brain. The current study serves as the proof of principle of waixenicin A’s neuroprotective and therapeutic effects in the mouse model of neonatal HI. However, two things remain to be clarified during the further testing of the compound. First, since TRPM7 is a ubiquitously expressed ion channel, it would be necessary to determine the uptake of the drug by other organs following systemic administration. This could be achieved by radiolabelling of waixenicin A and measuring its concentration in various organ systems. Any adverse effects of waixenicin A on different organs should also be investigated during this

152 testing. Second, a precise measurement of waixenicin A concentration in cerebrospinal fluid following systemic administration will provide us with a better understanding of BBB permeability to this compound and help us refine the dose necessary to achieve the desirable effect.

8.6.5 The effect of waixenicin A on other cell types in the brain

Here we show that waixenicin A administration reduces infarct volume and preserves the number of NeuN-positive cells potentially through actin cytoskeleton regulatory mechanism. However, the effect of waixenicin A on other cell types present in the brain has not been investigated in our study.

TRPM7 expression has been demonstrated in vascular endothelial cells derived from the umbilical vein (HUVEC), and silencing of TRPM7 expression with siRNA promoted proliferation in these cells via activation of ERK signaling pathway (Inoue and Xiong, 2009). Additionally, oxidative stress and free radicals were shown to cause an upregulation of TRPM7 protein in human endothelial cells (Baldoli et al., 2013). However, in human capillary endothelial cells (HMEC), silencing TRPM7 had a growth-inhibitory effect (Baldoli and Maier, 2012). While these results are conflicting, possibly due to differences in the cell lines, they suggest that TRPM7 could be a possible contributor to angiogenesis. The ischemic penumbra becomes an active site of remodeling and angiogenesis and begins as early as 12-24 hours after ischemia in rodents (Hayashi et al., 2003; Marti et al., 2000). Therefore, the effects of inhibition of TRPM7 activity by waixenicin A on angiogenesis following HI in our model must be considered in future studies.

TRPM7 expression has also been confirmed in mouse cortical astrocytes. Silencing and pharmacological inhibition of TRPM7 in these cells impaired their proliferation and migration through via ERK and c-Jun N-terminal kinase (JNK) signaling, but not through p38 MAPK or Akt (Zeng et al., 2015). Astrocytes participate in several processes after hypoxic-ischemic injury in the brain. Emergence of reactive astrocytes after stroke in peri-infarct area contributes to formation of “glial scar”, which has been previously shown to inhibit axonal regeneration (Ridet et al., 1997). However, astrocytes have also been implicated in preserving neuronal tissues during early phase of the brain injury, maintaining and repairing the blood-brain barrier and decrease immune cell infiltration (Bush et al., 1999; Rossi et al., 2007; Okada et al., 2006).

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Knockout mice lacking glial fibrillary acidic protein (GFAP) and vimentin, the two proteins that are commonly expressed by astrocytes, showed a reduction in reactive astrocytes and glutamate transport, as well as increased infarct volumes after stroke (Li et al., 2008). Therefore, astrocytes could play both detrimental and protective roles after stroke, and inhibition of TRPM7 activity in these cells could interfere with the critical functions of these cells.

Microglial cells are another type of cell that may be affected by waixenicin A blocking of TRPM7 activity. Microglial cells become activated in the CNS after injury such as hypoxia- ischemia and can exist in different activation states that can be either pro- or anti-inflammatory. Stimulation of microglia with either IL-4 or IL-10 cytokines was shown to facilitate brain repair by reducing acute inflammation by regulating anti-inflammatory mediators and neurotrophic factors and facilitating removal of debris (Colton, 2009). Inhibition of TRPM7 activity in these anti-inflammatory microglia was shown to reduce the migration and invasiveness of these cells, which could potentially have a detrimental effect to brain injury (Siddiqui et al., 2014). Similarly, TRPM7 currents have been demonstrated in anti-inflammatory microphages, and inhibition of these channels abolished proliferation and polarization of these cells (Schilling et al., 2014). While the role of TRPM7 activity in microglial cells is still being studied, it’s possible that inhibition of TRPM7 activity in these cells may actually have a detrimental impact on brain injury.

Overall, inhibition of TRPM7 activity with waixenicin A in our model was beneficial and resulted in reduced infarct volume, preservation of neuronal numbers, reduction in apoptotic cell death and preservation of short and long-term function. However, an in-depth understanding of the effects of TRPM7 inhibition on different cell types and their contribution to evolution of hypoxic-ischemic brain injury in vivo is required for waixenicin A to be successfully developed into a potential therapeutic tool.

8.7 Future directions

This thesis work implicates TRPM7 activity in axonal outgrowth regulation through cytoskeletal dynamics under normal and pathophysiological conditions. Moreover, it demonstrates neuroprotective and therapeutic properties of TRPM7 inhibitor waixenicin A in a mouse model of neonatal HI brain injury, thus creating the basic foundation for future investigation of this compound as a drug development candidate for this condition. Finally, it investigates several

154 molecular mechanisms downstream of TRPM7 that contribute to neuronal injury and cell death in HI. Below I propose several new directions for investigation that can contribute to further understanding of TRPM7 in neuronal development and injury.

8.7.1 Other proteins at the neuronal growth cone

Proteomic analysis of HI injury identified several differentially expressed proteins that are usually found at the neuronal growth cone. In our study, administration of waixenicin A prior to onset of HI attenuated the changes in these proteins. Therefore, it would be interesting to investigate the relationship between TRPM7 and these proteins in the context of axonal outgrowth under normal and adverse conditions.

Cyclin-dependent kinase 5 (Cdk5) is a serine/threonine kinase expressed in CNS that plays a key role in axonal outgrowth and is present in the growth cones of extending neurites (Fu et al., 2002). We found this kinase to be significantly upregulated in the lesioned hemisphere, and waixenicin A administration reduced Cdk5 expression to normal levels. Inhibition of Cdk5 activity was shown to reduce infarct volumes and promote functional recovery in a rat neonatal HI model (Tan et al., 2015), which is in accordance with our proteomic data. Cdk5 null mice display defects in axonal elongation, and in cortical neurons inhibition of Cdk5 activity attenuates NGF-mediated neurite outgrowth (Nikolic et al., 1996; Paglini and Caceres, 2001). Cdk5 phosphorylation targets include several Ca2+-conducting ion channels, including TRPA1 channel (Sulak et al., 2018; Furusawa et al., 2014). Given the localization of Cdk5 to the neuronal growth cone, it would be interesting to investigate a potential interaction between TRPM7 and Cdk5, and if TRPM7 kinase domain plays a role in Cdk5 activation and vice versa.

Dynamin 2 (Dyn2), a GTPase that associates with actin-binding protein contractin in the lamellipodia in epithelial cells, was significantly downregulated in our data. This downregulation was abolished by waixenicin A. A study by Kurklinsky and colleagues demonstrated that Dyn2 localizes to the growth cones of developing hippocampal neurons and forms a physical complex with contractin and α-actinin-1. Overexpression of Dyn2 reduced actin dynamics and resulted in large, flat and static growth cones (Kurklinsky et al., 2011). Given our previous findings of TRPM7-α-actinin-1 interactions, it would be interesting to investigate the interactions between TRPM7 and Dyn2 and their effects on neurite outgrowth.

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Finally, given our findings and previous literature, CaMKII and calcineurin interactions with TRPM7 at neuronal growth cone should be investigated. CaMKII and calcineurin were previously shown to act as a switch for cytoskeletal dynamics at the growth cone, and were shown to be controlled by local Ca2+ levels (Wen et al., 2004). Since our findings suggest that TRPM7 may regulate neurite elongation at least in part through Ca2+ influx, it would be worthwhile to investigate if TRPM7 may be one of the upstream regulators of CaMKII and calcineurin during neuronal development.

Overall, several Ca2+-dependent actin-modulating proteins should be investigated in the context of TRPM7-mediated neurite outgrowth and linked to the signaling pathways investigated in vivo.

8.7.2 The role of factors released by glia

Our findings suggest that factors released by glial cells present under hypoxic conditions may accelerate axonal retraction, and this retraction also coincides with potentiation of TRPM7 activity. Therefore, an in-depth investigation of the factors present in these cultures and their effects on TRPM7 expression and activity should be undertaken. A large body of literature suggests that the most prominent factors released by glial cells in co-cultures under adverse conditions are tumour necrosis factor-α (TNF-α), interleukin-6 (IL-6) and interleukin-1β (IL-1β) (Andersson et al., 2005; Minogue et al., 2012; Machacek et al., 2016; Garwood et al., 2011). Moreover, IL-1β, IL-6 and TNF-α were found to be upregulated in cerebrospinal fluid of neonates diagnosed with HIE, and IL-1β was shown to correlate with the severity of brain injury (Aly et al., 2006).

Several members of TNF superfamily were shown to be capable of regulating neurite outgrowth in the developing nervous system. TNF-α was shown to inhibit neurite outgrowth and branching of hippocampal neurons co-cultured with astrocytes. This effect was reversed in cultures derived from TNF-receptor deficient mice or in wild-type cultures where TNF receptor was blocked with a soluble TNF receptor IgG fusion protein (Neumann et al., 2002). Interestingly, TNF-α was also shown to modulate expression and activity of several different ion channels in vitro and in vivo (Vicente et al., 2004; Czeschik et al., 2008). Involvement of this cytokine in TRPM7-mediated axonal outgrowth/retraction and TRPM7 activity can be studied in culture with the use of combination of TRPM7 modulators (waixenicin A and naltriben) and factors that stimulate the

156 release of TNF-α from glial cells (IL-1β and IFN-γ). Moreover, TNF-α deficient mice have also been characterized and are commercially available.

Il-6 is another obvious target for further investigation. As mentioned previously, IL-6 was shown to functionally inhibit TRPM7 currents in a dose-dependent manner in primary cultured neurons (Liu et al., 2016). Interestingly, administration if IL-6 after spinal cord injury enhanced outgrowth and regeneration of the damaged axons (Yang et al., 2012). Additionally, IL-6 was shown to promote enhanced outgrowth and sprouting of axons in injured organotypic hippocampal slice cultures (Hakkoum et al., 2007). Moreover, IL-6 has been shown to play a role in neuronal differentiation and neurogenesis in the developing brain. Given the previously published inhibitory effects of IL-6 on TRPM7 activity and commercial availability of IL-6 deficient mice, it would be a reasonable next step to investigate in-depth regulation of TRPM7 by IL-6 in the context of neuronal development under normal and adverse conditions.

IL-1β has been shown to contribute to pathogenesis of hypoxic-ischemic brain injury, and IL-1β down-regulation by siRNA in vitro and lentivirus in vivo significantly attenuated brain edema, astrocyte dysfunction and neurological deficit in rat neonatal HI model (Liu et al., 2015). This effect was postulated to be due to IL-1β regulation of IL-6. More specifically, downregulation of IL-1β caused an upregulation of IL-6 production (Liu et al., 2015). Therefore, it is conceivable that IL-1β may indirectly regulate TRPM7 activity through IL-6 expression.

Overall, in-depth investigations of the effects of these factors on TRPM7 activity, expression and neurite outgrowth are needed and will contribute to further understanding of TRPM7 role in neuronal development and injury.

8.7.3 TRPM7 in other neurodegenerative conditions

While TRPM7 has been extensively studied under conditions of cerebral ischemia, its biophysical characteristics may also implicate it in other neurodegenerative conditions. Particularly, more attention should be dedicated to studying TRPM7 in Alzheimer’s disease (AD), Parkinson’s disease (PD) and amyotrophic lateral sclerosis (ALS).

AD is a devastating and progressive neurodegenerative condition leading to complete loss of cognitive abilities. A key pathogenic factor of AD is accumulation of amyloid-β (Aβ), a peptide derived from amyloid precursor protein (APP), in the extracellular space. TRPM2, a close

157 relative of TRPM7, has been implicated in cerebrovascular dysfunction caused by activation of TRPM2, possibly via production of cyclic ADP-ribose by poly (ADP-ribose) polymerase (Park et al., 2014). Overproduction of ROS also contributes to pathogenesis of AD, leading to deregulation of intracellular Ca2+ concentrations (Ermak and Davies, 2002). As previously described, TRPM7 activity is potentiated under oxidative conditions, thus making it a potential player in neuronal injury in AD. Moreover, mutations in presenilin genes (PS1 and PS2), which are responsible for Familial Alzheimer’s disease (FAD), were also shown to upregulate PIP2 levels, thus enhancing potentially detrimental Ca2+ influx through TRPM7 (Oh et al., 2012). Therefore, waixenicin A therapeutic properties should also be investigated in vitro and in vivo models of AD.

PD is another progressive neurodegenerative disorder that is characterized by the loss of dopaminergic neurons in the substantia nigra brain region, which leads to motor disabilities. A study using TRPM7 mutant zebrafish showed deficits in production and release of dopamine and hypomotile phenotype (Decker et al., 2014). Additionally, overexpression of kinase-dead TRPM7 in SH-SY5Y cells, which are dopaminergic, caused cell death (Decker et al., 2014). As well, carvacrol was recently shown to provide neuroprotection in a rodent model of PD that used unilateral injection of 6-hydroxy-dopamine into the striatum, potentially at least in part through TRPM7 inhibition (Dati et al., 2017). Therefore, further studies establishing the role of TRPM7 in pathogenesis of PD are necessary.

ALS is a neurodegenerative disease, characterized by loss of motor neurons (Shaw, 2005). Several studies suggest that ALS and Parkinsonism Dementia, a disorder related to ALS, are connected to deregulation of intracellular levels of Ca2+ and Mg2+ (Hermosura and Garruto, 2007). A missense mutation of TRPM7 was also found in some of ALS/PD patients in Guam, but similar findings were not observed in the Kii peninsula of Japan, suggesting that TRPM7 contributes to the pathogenesis of the disease at least in a subset of patients (Hara et al., 2010; Hermosura and Garruto, 2007). The role of TRPM7 in neurotransmitter release in sympathetic neurons (Krapivinsky et al., 2006) may also be important in ALS, as one of the earliest pathogenic events of this condition is the loss of neuromuscular synapses (Frey et al., 2000d).

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Overall, this thesis work validated waixenicin A as a potent pharmacological tool for TRPM7 activity modulation, thus allowing for further investigation of TRPM7’s roles in a variety of neurodegenerative conditions of the CNS.

8.8 Significance

Overall, this thesis work contributes to understanding of TRPM7 role in neuronal development under normal and pathophysiological conditions. It validates waixenicin A as a pharmacological tool for TRPM7 activity modulation and tests neuroprotective and therapeutic properties of this compound in an animal model of neonatal brain injury. Finally, it investigates the molecular mechanisms of TRPM7 in cellular injury and death.

Appendices 9 Appendix 9.1 Appendix 1: Waixenicin A extraction method

Waixenicin A was obtained from freeze-dried polyps of the soft coral Sarcothelia edmondsoni, which was collected in Kailua Bay (Oahu, Hawaii), as described previously (Zierler et al., 2011). The biological material was ground and exhaustively extracted with hexane. The solvent was removed under vacuum to give a crude extract residue. The residue was first fractionated by silica high performance liquid chromatography (HPLC) to give semi-purified waixenicin A. Final purification was accomplished by reversed phase (C18) HPLC, and compound identity and purity was established by nuclear magnetic resonance spectroscopy (NMR, d6-benzene) in comparison to in-house reference data (Zierler et al., 2011). Aliquots of the compound were stored as a dried film in a dessicator until use.

9.2 Appendix 2: Electrophysiological recordgings

Whole-cell TRPM7-like currents were recorded from DIV3 to DIV7 cultured mouse hippocampal neurons and non-induced and induced HEK-293 cells, using an Axopatch 700B (Axon Instruments, Inc.) as described previously (Chen et al., 2015a). Currents were recorded using a 400 ms voltage ramp protocol (-100 to +100 mV) at a holding potential of -70 mV with an interval of 5s at 2 kHz and digitized at 5 kHz. pClamp 9.2 software was used for data acquisition and Clampfit 9.2 was used for data analysis. All experiments were carried out at room temperature. Patch pipette resistance was between 5-9 megaohms after filling with pipette solution containing (in mM): 145 cesium methanesulfonate, 8 NaCl, 10 EGTA, and 10 HEPES, pH adjusted to 7.2 with CsOH. The bath solution contained (in mM) 140 NaCl, 5 KCl, 2 CaCl2, 20 HEPES, and 10 glucose (pH adjusted to 7.4 with NaOH). The cells were perfused with 500 nM waixeincin A (Hawaii Pacific University) or 50 μM naltriben mesylate.

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9.3 Appendix 3: Proteomic analysis

9.3.1 TMT Q-Exactive samples preparation and analysis

Samples were reduced, alkylated, digested, and TMT labelled according to manufacturer’s directions (Thermo Fisher TMT 10 Plex, Product 90110). Labelled peptides from all samples were combined and lyophilized. Peptides were then fractionated into 3 fractions using the Pierce High pH Reversed-Phase Peptide Fractionation Kit (Pierce Cat 84868) as per manufacturer’s directions (fractions 3, 5, X). Samples were analyzed on a Orbitrap analyzer (Q-Exactive, ThermoFisher, San Jose, CA) outfitted with a nanospray source and EASY-nLC nano-LC system (ThermoFisher, San Jose, CA). Lyophilized peptide mixtures were dissolved in 0.1% formic acid and loaded onto a 75μm x 50 cm PepMax RSLC EASY-Spray column filled with 2μM C18 beads (ThermoFisher San, Jose CA) at a pressure of 800 Bar and a temperature of 60C.. Peptides were eluted over 180 min at a rate of 250nl/min using a gradient set up as 0%-40% gradient of Buffer A (0.1% Formic acid; and Buffer B, 0.1% Formic Acid in 80% acetonitrile). Peptides were introduced by nano-electrospray into the Q-Exactive mass spectrometer (Thermo-Fisher). The instrument method consisted of one MS full scan (525–1800 m/z) in the Orbitrap mass analyzer with an automatic gain control (AGC) target of 5e5, maximum ion injection time of 100 ms and a resolution of 35 000 followed by 15 data-dependent MS/MS scans with a resolution of 35,000, an AGC target of 5e5, maximum ion time of 100ms, and one microscan. The intensity threshold to trigger a MS/MS scan was set to an underfill ratio of 1.0%. Fragmentation occurred in the HCD trap with normalized collision energy set to 28. The dynamic exclusion was applied using a setting of 35 seconds.

9.3.2 Database searching

Tandem mass spectra were extracted, charge state deconvoluted and deisotoped by Xcalibur version 2.2. All MS/MS samples were analyzed using Sequest (Thermo Fisher Scientific, San Jose, CA, USA; version 1.4.1.14) and X! Tandem (The GPM, thegpm.org; version CYCLONE (2010.12.01.1)). Both search engines were set up to search Uniprot-Mus Musculus-Oct- 172016_reviewed.fasta (Download Oct 17 2016, 16818 entries) assuming the digestion enzyme trypsin. Sequest and X! Tandem were searched with a fragment ion mass tolerance of 0.020 Da and a parent ion tolerance of 20 PPM. Carbamidomethyl of cysteine and TMT6plex of lysine and the n-terminus were specified in Sequest and X! Tandem as fixed modifications. Deamidated of

161 asparagine and glutamine and oxidation of methionine were specified in Sequest as variable modifications. Glu->pyro-Glu of the n-terminus, ammonia-loss of the n-terminus, gln->pyro-Glu of the n-terminus, deamidated of asparagine and glutamine and oxidation of methionine were specified in X! Tandem as variable modifications.

9.3.3 Criteria for protein identification

Scaffold (version Scaffold_4.8.4, Proteome Software Inc., Portland, OR) was used to validate MS/MS based peptide and protein identifications. Peptide identifications were accepted if they could be established at greater than 95.0% probability by the Scaffold Local FDR algorithm. Protein identifications were accepted if they could be established at greater than 95.0% probability and contained at least 2 identified peptides. Protein probabilities were assigned by the Protein Prophet algorithm (Nesvizhskii et al., 2003). Proteins that contained similar peptides and could not be differentiated based on MS/MS analysis alone were grouped to satisfy the principles of parsimony. Proteins were annotated with GO terms from GOA mouse.gaf (downloaded Nov 11, 2016) (Ashburner et al., 2000).

9.3.4 Quantitative data analysis

Scaffold Q+ (version Scaffold_4.8.4, Proteome Software Inc., Portland, OR) was used to quantitate Label Based Quantitation (iTRAQ, TMT, SILAC, etc.) peptide and protein identifications. Peptide identifications were accepted if they could be established at greater than 95.0% probability by the Scaffold Local FDR algorithm. Protein identifications were accepted if they could be established at greater than 95.0% probability and contained at least 2 identified peptides. Protein probabilities were assigned by the Protein Prophet algorithm (Nesvizhskii et al., 2003). Proteins that contained similar peptides and could not be differentiated based on MS/MS analysis alone were grouped to satisfy the principles of parsimony. Normalization was performed iteratively (across samples and spectra) on intensities, as described in Statistical Analysis of Relative Labeled Mass Spectrometry Data from Complex Samples Using ANOVA (Oberg et al., 2008). Means were used for averaging. Spectra data were log-transformed, pruned of those matched to multiple proteins and those missing a reference value, and weighted by an adaptive intensity weighting algorithm. Of 207066 spectra in the experiment at the given thresholds, 189703 (92%) were included in quantitation.

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9.4 Appendix 4: Confirmation of TRPM7 expression in HEK-293 cells.

HEK-293 cells were grown to 85% confluency and the media was replaced with media containing tetracycline (1 μg/ml)(Chen et al., 2015b). After 20 hours the cells were imaged and proteins were extracted for Western blot. Figure 50A shows the control (-TET) and tetracycline- induced (+TET) cells that exhibited characteristic cell rounding (Runnels et al., 2002). Figure 50B shows TRPM7 and flag expression in (-TET) and (+ TET) HEK-293 cells.

Figure 50: Confirmation of TRPM7 expression in TET-inducible HEK-293 cells. A. +TET cells exhibit characteristic cell rounding after 20 hours of treatment. B.Representative western blot showing TRPM7 and flag expression in +TET cells after 20 hours of treatment.

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9.5 Appendix 5: Data sumary of axonal and dendritic lengths under normoxic and hypoxic conditions.

Table 8: Data summary for Figure 25.

Normoxic neurons - Axonal lengths (μm) Hypoxic neurons - Axonal lengths (μm) 500 nM 500 nM Hours Untreated Vehicle Waixenicin A Untreated Vehicle Waixenicin A Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N

0 248.7 11.2 137 248.7 11.2 137 248.7 11.2 137 248.7 11.2 137 248.7 11.2 137 248.7 11.2 137 1 241 21.7 43 227.6 12.9 63 317 22.9 73 303.9 25.9 32 300 21.1 50 338.1 20.2 53 2 279.3 14.9 58 287.6 16.4 53 334.5 19.7 63 180.2 10.5 46 197.5 10.7 65 265.6 16.9 43 4 317.9 28.4 32 316.9 23.6 39 397.6 29.3 40 120.6 16.2 32 126.8 19 43 296.9 21.3 43 6 365.8 14 57 325.9 16.2 32 416.3 12.4 42 114.1 14.6 46 100.6 13.7 54 320.4 20.9 50 8 398.3 17.1 59 399 16.5 41 466.2 18.5 44 49.4 27.6 57 53.64 17.7 49 178.3 15.9 45 10 421 20.6 42 430.7 22.4 44 635.7 27.1 63 23.96 8.96 51 15.58 5.58 40 79.81 12.3 42 Normoxic neurons - Dendritic lengths (μm) Hypoxic neurons - Dendritic lengths (μm) 500 nM 500 nM Hours Untreated Vehicle Waixenicin A Untreated Vehicle Waixenicin A Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N

0 188.1 7.95 160 188.1 7.95 160 188.1 7.95 160 188.1 7.95 160 188.1 7.95 160 188.1 7.95 160 1 190.1 7.35 43 199.7 12.2 65 191.1 11.3 63 120.7 13.3 32 126.1 7.88 50 145.1 9.93 53 2 202.3 9.42 59 207.9 8.97 58 198.5 12.1 76 105 9.8 46 111 8.3 61 172.9 14.5 44 4 223.5 10.8 33 223.1 8.61 39 221.2 8.41 39 92.03 14.5 32 90.43 12.1 43 148.4 12.9 43 6 227.7 10.9 55 243.3 15.2 32 222.2 13.6 42 66.54 7.63 45 66.56 7.55 52 109.7 10.3 50 8 291.4 16.9 59 268.9 17.8 41 304.2 18.6 44 41.42 5.62 57 53.56 8.43 49 53.93 8.13 45 10 326.1 27.9 42 320.3 23 44 330.6 24.4 63 6.62 1.62 51 6.487 2.49 40 7.975 4.98 42

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9.6 Appendix 6: Data summary of axonal lengths under normoxic and hypoxic conditions with and without naltriben treatment.

Table 9: Data summary for Figure 26. Normoxia Untreated Normoxia Normoxia Hours (μm) Normoxia Vehicle (μm) Naltriben 1 uM (μm) Naltriben 10 uM (μm) Mean SEM N Mean SEM N Mean SEM N Mean SEM N

0 263.4 25.0 50 263.4 25.0 50 263.4 25.0 50 263.4 25.0 50 1 268.7 25.4 54 275.8 35.0 48 244.9 29.1 44 227.3 18.0 51 2 275.0 38.0 41 264.8 31.9 38 204.1 21.7 58 183.6 15.3 46 4 301.9 44.6 42 284.9 36.7 38 184.4 13.6 46 142.7 8.4 52 Hypoxia Hypoxia Hours Hypoxia Untreated (μm) Hypoxia Vehicle (μm) Naltriben 1 uM (μm) Naltriben 10 uM (μm) Mean SEM N Mean SEM N Mean SEM N Mean SEM N

0 263.4 25.0 50 263.4 25.0 50 263.4 25.0 50 263.4 25.0 50 1 305.7 50.1 32 324.9 45.1 34 244.5 35.0 40 186.8 18.7 45 2 202.9 30.5 35 215.8 36.4 36 123.7 29.1 44 107.7 29.9 18 4 145.6 13.3 37 175.4 11.8 40 98.5 14.0 37 53.9 15.3 18

9.7 Appendix 7: Data summary of axonal and dendritic lengths in neuron-astrocyte co-cultures.

Table 10: Data summary for Figure 29.

Normoxic neurons - Axonal lengths (μm) Hypoxic neurons - Axonal lengths (μm)

Hours Untreated Vehicle Waix A Untreated Vehicle Waix A Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N

0 750 42 50 750 42 50 750 42 50 750 42 50 750 42 50 750 42 50 1 779 48 53 744 43 50 805 49 48 502 38 43 402 26 51 658 43 44 2 832 59 38 884 59 38 897 53 41 350 23 46 366 15 47 687 49 43 4 833 44 56 862 43 61 954 70 40 182 22 54 219 21 52 743 32 43 6 905 43 52 909 36 44 1110 50 41 125 18 34 95 19 38 459 36 41 Normoxic neurons - Dendritic lengths (μm) Hypoxic neurons - Dendritic lengths (μm)

Hours Untreated Vehicle Waix A Untreated Vehicle Waix A Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N Mean SEM N

0 168 11 50 168 11 50 168 11 50 168 11 50 168 11 50 168 11 50 1 178 12 53 178 14 50 184 11 48 94 6 43 88 6 51 141 9 44 2 179 18 38 179 16 38 207 16 41 102 9 46 96 9 47 147 12 43 4 185 11 56 193 11 61 203 20 40 127 10 54 110 9 52 160 15 43 6 181 8 52 190 9 44 213 11 41 30 5 34 36 4 38 103 11 41

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9.8 Appendix 8: Data summary of axonal lengths of neurons that received conditioned media from hypoxic neurons or hypoxic co-cultures

Table 11: Data summary for Figure 31.

Control neurons Switch from neuronal cultures - Axonal lengths (um) Hours Control Untreated Vehicle Waix A Mean SEM N Mean SEM N Mean SEM N Mean SEM N 0 231 42 50 231 42 50 231 42 50 231 42 50 1 237 20 41 301 19 35 293 23 38 314 34 34 2 255 19 35 279 17 31 255 19 37 308 29 30 4 262 17 39 227 23 39 222 27 37 299 31 27 6 264 17 39 198 20 31 181 14 31 254 20 33 Control neurons Switch from neuron-astrocyte co-cultures - Axonal lengths (um) Hours Control Untreated Vehicle Waix A Mean SEM N Mean SEM N Mean SEM N Mean SEM N 0 231 42 50 231 42 50 231 42 50 231 42 50 1 237 20 41 201 18 34 192 23 38 224 25 30 2 255 19 35 173 17 38 159 19 42 219 20 29 4 262 17 39 144 23 30 141 22 34 185 29 36 6 264 17 39 118 20 31 124 19 31 174 26 31

9.9 Appendix 9: Proteins identified by LC-MS/MS

Table 12: List of 95 differentially expressed proteins identified by LC-MS/MS.

Protein Uniprot ID Size Vehicle Waix A

Protein dpy-30 homolog Q99LT0 11 kDa 3.16 1.29

Bromodomain-containing protein 8 Q8R3B7 103 kDa 2.60 1.12

Calmodulin P0DP26 17 kDa 2.06 1.03

Cyclin-dependent kinase-like 5 Q3UTQ8 105 kDa 1.97 1.04

RNA-binding protein FUS P56959 53 kDa 1.97 1.15

FACT complex subunit SSRP1 Q08943 81 kDa 1.97 1.18

Lactoylglutathione lyase Q9CPU0 21 kDa 1.92 1.14

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Serine/threonine-protein kinase 26 Q99JT2 47 kDa 1.89 1.03

Proteasome subunit beta type-7 P70195 30 kDa 1.85 1.14

Transcription factor 20 Q9EPQ8 216 kDa 1.83 1.08

SPATS2-like protein Q91WJ7 62 kDa 1.82 1.31

Cellular nucleic acid-binding protein P53996 20 kDa 1.79 1.07

Complexin-2 P84086 15 kDa 1.77 0.96

Coagulation factor XIII A chain Q8BH61 83 kDa 1.74 1.34

Small integral membrane protein 13 E9Q942 10 kDa 1.74 1.11

Protein Shroom3 Q9QXN0 215 kDa 1.74 1.01

Beta-adrenergic receptor kinase 1 Q99MK8 80 kDa 1.73 1.12

Peroxiredoxin-4 O08807 31 kDa 1.72 1.08

Tubulin polymerization-promoting protein family member 3 Q9CRB6 19 kDa 1.71 0.91

Hypoxanthine-guanine phosphoribosyltransferase P00493 25 kDa 1.71 1.00

10 kDa heat shock protein, mitochondrial Q64433 11 kDa 1.70 0.91

BRISC and BRCA1-A complex member 1 Q3UI43 37 kDa 1.69 1.10

Protein MB21D2 Q8C525 48 kDa 1.68 1.23

Basigin P18572 42 kDa 1.63 1.04

Uncharacterized protein KIAA1211 Q5PR69 132 kDa 1.63 1.04

Proteasome subunit beta type-6 Q60692 25 kDa 1.62 1.06

Proteasome subunit alpha type-4 Q9R1P0 29 kDa 1.61 1.03

Plasminogen activator inhibitor 1 RNA-binding protein Q9CY58 45 kDa 1.61 1.14

Small nuclear ribonucleoprotein Sm D3 P62320 14 kDa 1.60 0.97

Delta-aminolevulinic acid dehydratase P10518 36 kDa 1.60 0.98

Acidic leucine-rich nuclear phosphoprotein 32 family member A O35381 29 kDa 1.60 1.06

Sentrin-specific protease 7 Q8BUH8 116 kDa 1.58 1.35

Microtubule-associated protein tau P10637 76 kDa 1.58 1.03

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Stathmin P54227 17 kDa 1.58 1.04

Ran-specific GTPase-activating protein P34022 24 kDa 1.57 1.02

Proteasome subunit alpha type-2 P49722 26 kDa 1.57 1.05

Vacuolar protein sorting-associated protein 45 P97390 65 kDa 1.57 1.04

Nucleobindin-1 Q02819 53 kDa 1.56 1.06

Myristoylated alanine-rich C-kinase substrate P26645 30 kDa 1.55 0.92

Myotrophin P62774 13 kDa 1.55 1.21

Fumarylacetoacetase P35505 46 kDa 1.55 1.10

Proteasome subunit beta type-2 Q9R1P3 23 kDa 1.55 0.99

MARCKS-related protein P28667 20 kDa 1.55 1.00

Reticulocalbin-2 Q8BP92 37 kDa 1.54 0.98

Small nuclear ribonucleoprotein E P62305 11 kDa 1.54 1.05

Calbindin P12658 30 kDa 1.53 0.94

SPARC-like protein 1 P70663 72 kDa 1.52 0.99

Protein SET Q9EQU5 33 kDa 1.52 0.99

Cytochrome c, somatic P62897 12 kDa 1.52 1.03

Protein phosphatase 1 regulatory subunit 14B Q62084 16 kDa 1.51 1.04

Visinin-like protein 1 P62761 22 kDa 1.51 1.00

Proteasome subunit alpha type-6 Q9QUM9 27 kDa 1.51 1.05

Copper transport protein ATOX1 O08997 7 kDa 1.49 1.05

Calcineurin subunit B type 1 Q63810 19 kDa 1.49 0.99

Small nuclear ribonucleoprotein-associated protein B P27048 24 kDa 1.48 1.10

Alpha-2-HS-glycoprotein P29699 37 kDa 1.48 1.09

Ubiquitin-60S ribosomal protein L40 P62984 15 kDa 1.48 1.18

Acidic leucine-rich nuclear phosphoprotein 32 family member E P97822 30 kDa 1.48 1.04

28S ribosomal protein S36, mitochondrial Q9CQX8 11 kDa 1.47 1.02

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Neuronal calcium sensor 1 Q8BNY6 22 kDa 1.47 1.04

Prosaposin Q61207 61 kDa 1.47 1.00

Heterogeneous nuclear ribonucleoprotein A/B Q99020 31 kDa 1.47 1.09

Protein piccolo Q9QYX7 551 kDa 1.47 0.99

Neuron-specific calcium-binding protein hippocalcin P84075 22 kDa 1.46 1.05

Clathrin light chain A O08585 26 kDa 1.46 1.12

Centrin-2 Q9R1K9 20 kDa 1.45 1.00

Translation machinery-associated protein 7 Q8K003 7 kDa 1.45 0.98

ATP synthase subunit delta, mitochondrial Q9D3D9 18 kDa 1.45 1.11

Thioredoxin P10639 12 kDa 1.45 1.05

Neurocalcin-delta Q91X97 22 kDa 1.44 0.97

Regulator of G-protein signaling 7 O54829 55 kDa 0.74 1.09

Coatomer subunit epsilon O89079 35 kDa 0.73 0.98

Long-chain fatty acid transport protein 4 Q91VE0 72 kDa 0.73 0.99

L-lactate dehydrogenase B chain P16125 37 kDa 0.71 0.91

Neurofilament light polypeptide P08551 62 kDa 0.71 0.95

Twinfilin-2 Q9Z0P5 39 kDa 0.71 1.02

Protein YIPF5 Q9EQQ2 28 kDa 0.69 1.05

Beta-centractin Q8R5C5 42 kDa 0.69 0.92

NADH dehydrogenase ubiquinone 1 beta subcomplex subunit 4 Q9CQC7 15 kDa 0.68 1.05

Set1/Ash2 histone methyltransferase complex subunit ASH2 Q91X20 68 kDa 0.67 0.91

Filamin-C Q8VHX6 291 kDa 0.67 1.02

Coatomer subunit gamma-2 Q9QXK3 98 kDa 0.67 1.04

PDZ domain-containing protein GIPC2 Q9Z2H7 34 kDa 0.65 0.96

Mitochondrial dicarboxylate carrier Q9QZD8 32 kDa 0.65 1.05

169

Calpain-1 catalytic subunit O35350 82 kDa 0.65 0.96

Exocyst complex component 6B A6H5Z3 94 kDa 0.65 1.06

Calcium/calmodulin-dependent protein kinase type 1B Q9QYK9 39 kDa 0.63 0.99

General transcription factor 3C polypeptide 2 Q8BL74 100 kDa 0.62 0.69

MTSS1-like protein Q6P9S0 77 kDa 0.62 1.05

Death-inducer obliterator 1 Q8C9B9 247 kDa 0.62 0.90

Prolyl 3-hydroxylase OGFOD1 Q3U0K8 63 kDa 0.59 0.83

Sorting nexin-17 Q8BVL3 53 kDa 0.58 0.96

Calcium/calmodulin-dependent protein kinase type II subunit alpha P11798 54 kDa 0.58 0.98

Hemoglobin subunit beta-2 P02089 16 kDa 0.48 0.70

Eukaryotic translation initiation factor 4E-binding protein 2 P70445 13 kDa 0.38 0.61

9.10 Appendix 10: PKC interacts with TRPM7 C-terminus.

Co-immunoprecipitation assay with GST-fusion protein of TRPM7 C-terminus (nucleotides 4018-5090) and mouse hippocampal tissue was carried out to identify potential interactions of TRPM7 with PKC. Following GST-fusion protein binding to mouse hippocampal proteins, PKC was identified as one of the binding partners of TRPM7.

To investigate the involvement of specific PKC isoforms in TRPM7-mediated axonal outgrowth, several PKC inhibitors have been tested. E16 cultured hippocampal neurons have been treated with various inhibitors on DIV2, 24 hours after plating, and axonal and dendritic lengths have been analyzed on DIV4 to ensure that most neurons in culture have developed distinct axons and dendrites. Axonal and dendritic lengths have been analyzed separately with the use of ICC (anti- MAP2 for dendritic marker; anti-tau1 for axonal marker) and confocal microscopy, and SynD analysis routine. Broad PKC inhibitors such as staurosporine (IC50 = 0.7nM) and Go 6983 (IC50

170

= 7-10 nM) and GF109203X (IC50 = 20 nM), an inhibitor selective for α and β1 isoforms, abolished the enhancement in axonal outgrowth when applied together with waixenicin A. (Figure 50).

Figure 51: PKC interacts with TRPM7 C terminus. PKC interacts with TRPM7 C terminus. A) TRPM7 with phosphorylation cites (image adapted from Kim et al., 2012) and co-immunoprecipiation between GST-fustion TRPM7 and PKC. B) Mean axonal lengths of DIV4 neurons treated with different PKC inhibitors with and without waixenicin A. All data is presented as mean ±SEM. Statistical analysis: one-way ANOVA with Bonferroni post-hoc. *p<0.05.

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9.11 Appendix 11: Neuron densities of neuronal cultures and neuron-astrocyte co-cultures.

To determine the effect of hypoxia on neuronal survival, neuronal density was measured at different time points of hypoxia. Neurons were stained with neuronal marker NeuN and cell nuclei were stained with DAPI. The numer of NeuN and DAPI positive cells was measured in neuronal cultures and neuron-astrocyte co-cultures. No significant differences were detected between the groups in the same treatment groups or between treatment groups, suggesting that this hypoxic treatment does not cause immediate cell death.

Figure 52: Hypoxia does not have an effect on neuronal density. Neuronal density was measured in neuronal cultures and neuron- astrocyte co-cultures at different times of hypoxia duration. Data is presented as mean ± SEM. Statistical analysis: ANOVA with Bonferroni post-hoc.

The following citation has been re-created in this thesis with copyright permission and lisence:

Turlova et al., TRPM7 regulates axonal outgrowth and maturation of primary hippocampal neurons. Mol. Neurobiol. 2016. 53(1): 595-610.

Journal: Molecular Neurobiology

Permission type: Republish or display content

Type of use: Thesis/dissertation

Order Lisence ID: 4292190440589

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