Metformin Pretreatment Rescues Subependymal Zone Neurogenesis and Long-Term Olfactory Memory in a Juvenile Model of Cranial Irradiation

Daniel Derkach

A thesis submitted in conformity with the requirements for the degree of Master of Science Institute of Medical Sciences University of Toronto

© Copyright by Daniel Derkach 2019 Metformin Pretreatment Rescues Subependymal Zone Neurogenesis and Long-Term Olfactory Memory in a Juvenile Model of Cranial Irradiation Daniel Derkach Master of Science Institute of Medical Sciences University of Toronto 2019 Abstract

Neural stem cells (NCSs) in the mammalian brain contribute to neurogenesis throughout life and in response to brain injuries to promote neurorepair. Cranial irradiation (IR), used as an adjuvant therapy in the treatment of childhood brain tumors, results in cognitive deficits associated with long-term impairments to neurogenesis in rodent models. Metformin is an antidiabetic drug that enhances functional neurogenesis under physiological conditions and in response to brain injuries. Herein, we investigated metformin’s potential to rescue deficits to neurogenesis and long-term olfactory memory (LTOM) following cranial IR. Juvenile mice displayed acute and persistent deficits in periventricular neurogenesis following 8 Gy cranial IR.

We show that metformin pretreatment was sufficient to enhance the recovery of proliferating neuroblasts and completely restore LTOM. This study is the first to report that metformin pretreatment promotes neurogenesis and functional recovery following juvenile cranial IR and supports its consideration as a therapeutic intervention to enhance neurorepair.

ii Acknowledgements This Master’s thesis is the product of three years of research and support from my mentors, colleagues, friends, and family. This experience has greatly contributed to my self- development as a scientist, student, and educator, and has also provided me with a sense of both humility and reward. I am extremely grateful to my mentors Dr. Cindi Morshead, Dr. Rebecca Ruddy, and Master’s student Ashkan Azimi for the training, support, and critical feedback. Your guidance and insights expedited the learning process and helped me improve my critical thinking and reasoning skills. Your enthusiasm in helping me succeed has motivated me to work with passion and instilled a desire to pay it forward through dedicated mentoring of other undergraduate students. You have imparted with me the notion that success is not only found in personal accomplishment, but through the accomplishments of those who have learned from or have been influenced by you. I would like to acknowledge all my colleagues who have shared with me the experiences of wonder, confusion, frustration, perseverance, failure, and success. To Dr. Rebecca Ruddy, Dr. Emily Gilbert, Dr. Jessica Livingston-Thomas, Ashkan Azimi, and Kelsey Adams, your technical expertise and extensive understanding of neural stem cell biology has been immensely helpful in developing relevant and meaningful questions, as well as the experiments to address these questions. You’ve encouraged me to constantly learn and ask questions until there are no remaining answers. To Stasja Drecun, Nancy Liu, and Fil Stojic, your roles as devil’s advocates and source of philosophical discussion have continuously challenged me to think critically and through multiple perspectives, and to argue effectively and with humility. To Elana Sefton, Clara Bourget, Nishanth Lakshman, Rehnuma Islam, Stephanie Iwasa, Ilan Vonderwalde, Tom Enbar, and Dr. Erica Scott, thank you for your enduring friendship and for promoting a sense of family within the lab. To the undergraduate students that I have had the pleasure of mentoring, you’ve taught me just as much as I have hopefully taught you. Most importantly, I am eternally thankful for the unconditional support, sacrifice, and love of my family and my partner Katie. You’ve been my greatest source of inspiration and emotional support, and have instilled values within me that have guided every aspect of my life.

iii Statement of Contributions

Artificial cerebrospinal fluid (aCSF) and serum-free media (SFM) used for tissue culturing were prepared by lab manager Venkateswaran Subramaniam. Assistance with transcardial perfusions was provided by lab technician Ricky Siu and Master’s student Ashkan Azimi. Assistance with genotyping GFAP::GFP mice was provided by lab technician Ricky Siu.

Eight GFAP::GFP mice that were used to establish a colony were provided by Dr. Maryam Faiz’s lab. The fluorescence-activated cell sorting (FACS) protocol was provided by Dr. Fiona Doetsch’s lab. Doctoral student Kelsey Adams and Master’s student Elana Sefton assisted with optimizing the FACS protocol for the Morshead lab.

Undergraduate student Mohsen Heidari assisted in sectioning brains. Undergraduate student Tarlan Kehtari assisted in the execution and analysis of conditioned media experiments. Undergraduate student Matthew Renaud assisted in sectioning brains, food restriction of mice, and the execution of the long-term olfactory memory (LTOM) task.

Training with the neurosphere assay and on the cesium-137 gamma irradiator (Best Theratronics, Gamacell 40 Exactor) was provided by doctoral student, and more recently, Dr. Rebecca Ruddy.

Principal Investigator Dr. Cindi Morshead provided thesis manuscript feedback. Program Advisory Committee (PAC) members Dr. Rebecca Laposa, Dr. Freda Miller, and Dr. Cindi Morshead advised on project viability and experimental design.

This research was supported by funding from the Canadian Institutes of Health Research (CIHR), Brain Canada (via the Hospital for Sick Children, Toronto, Canada), the Stem Cell Network (SCN), and the Carlton and Marguerite Smith Medical Research Fellowship (Division of Anatomy within the Department of Surgery, University of Toronto).

iv Table of Contents Acknowledgements...... iii

Statement of Contributions ...... iv

Table of Contents ...... v

List of Abbreviations ...... ix

List of Figures ...... xv

List of Appendices ...... xvi

1. Literature Review ...... 1

1.1. Regenerative Medicine ...... 1

1.2. Neural Stem and Progenitor Cells ...... 1

1.2.1. Initial characterization of neural stem cells ...... 1

1.2.2. Existence of NSCs in the postnatal human and rodent brain ...... 2

1.2.3. Neural stem cells during development ...... 3

1.2.4. Neural stem cells during adulthood...... 5

1.3. The role of adult SEZ NSCs under baseline conditions ...... 7

1.3.1. Neural stem and progenitor pool maintenance ...... 8

1.3.2. In vivo NSC lineage and olfactory bulb neurogenesis ...... 10

1.3.3. The role of neurogenesis in olfactory behaviour ...... 12

1.3.4. Neurogenesis vs. gliogenesis ...... 14

1.4. The SEZ NSC Niche ...... 15

1.4.1. Cytoarchitecture ...... 15

1.4.2. Ependymal Cells and Cerebrospinal Fluid ...... 17

1.4.3. Microglia ...... 19

1.4.4. Vasculature ...... 21

v 1.4.5. Extracellular matrix (ECM) ...... 22

1.4.6. Neurotransmitters ...... 23

1.5. Postnatal NSC-niche heterogeneity and endogenous response to injury ...... 24

1.5.1. Regional heterogeneity within the SEZ ...... 24

1.5.2. Temporal heterogeneity within the SEZ ...... 26

1.5.3. Changes to NSC and niche dynamics following injury ...... 29

1.6. Assaying for NSCs ...... 32

1.6.1. Assaying for NSCs in vitro...... 32

1.7. Cranial Irradiation ...... 34

1.7.1. IR-induced apoptosis and senescence ...... 35

1.7.2. IR-induced NPC dysfunction and deficits to neurogenesis and oligodendrogenesis...... 35

1.7.3. Niche-mediated effects of cranial IR ...... 38

1.8. Endogenous NPC activation using metformin ...... 41

1.8.1. Pharmacology & toxicology ...... 41

1.8.2. Metformin in the brain ...... 43

1.8.3. Metformin’s pro-neurogenic effects via direct action on NPCs ...... 44

1.8.4. Metformin’s protective effects in the neurogenic niche ...... 46

2. Research Aims and Hypothesis ...... 50

2.1. Hypothesis...... 50

2.2. Objectives...... 50

3. Methods ...... 53

3.1. Animals ...... 53

3.2. Metformin administration ...... 53

vi 3.3. Cranial irradiation ...... 54

3.4. Neurosphere assay...... 54

3.4.1. Tissue dissection ...... 54

3.4.2. Cell culture ...... 55

3.4.3. Conditioned media ...... 55

3.4.4. Quantification ...... 56

3.5. Fluorescence-activated cell sorting (FACS) ...... 56

3.5.1. Reagents and solutions ...... 56

3.5.2. Tissue dissection ...... 56

3.5.3. Cell suspension and staining ...... 57

3.5.4. Sorting ...... 59

3.6. Immunohistochemistry and immunofluorescence ...... 59

3.6.1. Perfusions and tissue preparation ...... 59

3.6.2. Antibody labeling ...... 60

3.6.3. Microscopy and quantification ...... 60

3.7. Long-term olfactory memory task ...... 61

3.7.1. Food restriction, acclimation, and materials ...... 61

3.7.2. Pre-training and training ...... 62

3.7.3. Testing ...... 63

3.8. Statistical Analysis ...... 64

4. Results ...... 65

4.1. Effects of metformin on aNSC pool size ...... 65

4.1.1. Metformin expands the neonatal, but not the juvenile NSC pool in the SEZ...... 65

4.1.2. Metformin induces a short-term expansion of the aNSC pool in the SEZ...... 67

vii 4.2. Effects of metformin pretreatment on aNSC pool size following juvenile cranial IR ... 68

4.2.1. Juvenile cranial IR causes an acute depletion of aNSCs that recover spontaneously over time...... 68

4.2.2. Metformin pretreatment does not prevent the acute depletion of aNSCs following juvenile cranial IR...... 69

4.2.3. Metformin pretreatment does not affect aNSC pool recovery following juvenile cranial IR...... 70

4.3. Effects of metformin pretreatment on NSC activation state in the SEZ following juvenile cranial IR ...... 72

4.4. Factors in the irradiated SEZ niche reduce aNSC pool size in vitro...... 74

4.5. Effects of metformin pretreatment on SEZ neuroblasts following juvenile cranial IR . 76

4.6. Metformin pretreatment rescues deficits to long-term olfactory memory following juvenile cranial IR...... 78

5. Discussion...... 80

6. Conclusions ...... 92

7. Future Directions ...... 94

References ...... 98

Appendices ...... 130

viii List of Abbreviations AC – adenylate cyclase AICAR – 5-aminoimidazole-4-carboxamide ribonucleotide Akt (PKB) – protein kinase B AMP – adenosine monophosphate AMPK – AMP-activated protein kinase aNSC – activated neural stem cell APC/CC1 – adenomatous polyposis coli clone CC1 aPKC – atypical protein kinase C AQP – aquaporin Ara-C – cytosine--D- arabinofuranoside ATP – adenosine triphosphate BBB – blood-brain barrier BDNF – brain-derived neurotrophic factor BEC – brain endothelial cell BTC – betacellulin CBP – CREB-binding protein CCL2/MCP-1 – chemokine ligand 2 (also known as monocyte chemoattractant protein-1) CCR2 – CC chemokine receptor type 2 CD133 – cluster of differentiation 133 (also known as prominin-1) CD31 (PECAM-1) – cluster of differentiation 31 (platelet endothelial cell adhesion molecule 1) CD68/ED1 – cluster of differentiation 68 Cdc42 – cell division control protein 42 Cdk – cyclin dependent cyclase CM – conditioned media CNS – central nervous system CP – cortical plate CREB – cAMP response element-binding protein CSF – cerebrospinal fluid (aCSF = artificial CSF)

ix Ctrl – control CX3CR1 – CX3C chemokine (fractalkine) receptor 1 CXCR4/CD184 – CXC chemokine receptor type 4 (also known as fusin) D2L – D2-like dopamine receptor DA – dopamine DAPI – 4’,6-diamidino-2-phenylindole DCC – deleted in colorectal carcinoma DCX – doublecortin DG – dentate gyrus DLL1/3 – Delta-like canonical Notch ligand 1/3 Dlx2 – distal-less homeobox 2 DMEM/F12 – Dulbecco’s modified eagle medium/Ham’s F12 DNA DSB – DNA double-strand break ECM – extracellular matrix EdU - 5-ethynyl-2’-deoxyuridine EGF – epidermal growth factor EGFR – epidermal growth factor receptor EL – ependymal layer Emx1 – empty spiracles homeobox 1 ERK – extracellular signal-regulated kinase FACS – fluorescence-activated cell sorting FBPase – fructose-1,6-bisphosphatase FBS – fetal bovine serum FGF – fibroblast growth factor FMO – fluorescence minus one FoxJ1 – forkhead box protein J1 FOXO3 – forkhead box O3 FUCCI – fluorescence ubiquitination cell cycle indicator GABA – -aminobutyric acid

x GCL – granule cell layer GCV – ganciclovir GFAP – glial fibrillary acidic protein GFP – green fluorescent protein GL – glomerular layer GLAST – glutamate aspartate transporter Gsh2 – glutathione synthetase 2 GSK3 - glycogen synthase 3 H/I – hypoxic-ischemic HPC – hippocampus HSPG – heparan sulphate proteoglycan IAA – isoamyl acetate Iba1 – ionized calcium-binding adapter molecule 1 ICAM-1 – intercellular adhesion molecule-1 IFN - interferon- IGF – insulin-like growth factor IHC – immunohistochemistry IL – interleukin IP – intraperitoneal IR – irradiation IZ – intermediate zone L-DOPA – levodopa LeX – Sialyl-Lewis X [(also known as CD15 (cluster of differentiation 15) or SSEA-1 (stage-specific embryonic antigen 1)] LGE – lateral ganglionic eminence LSP – lipopolysaccharide LTOM – long-term olfactory memory LV – lateral ventricle LVCPsec – lateral ventricle choroid plexus conditioned media

xi MAP2 – microtubule associated protein-2 MAPK – mitogen-activated protein kinase Mash1 (or Ascl1) – mammalian achaete-scute homolog 1 MBP – myelin basic protein Mcm2 – minichromosome maintenance complex component 2 MEK (MAPKK/MAP2K) – mitogen-activated protein kinase kinase MPTP – 1-methyl-4-phenyl-1,2,3,6-tetrahydropyridine MT5-MMP – membrane-type 5 matrix metalloproteinase mTOR(C) – mammalian/mechanistic target of rapamycin (complex) MZ – marginal zone NeuN – neuronal nuclear antigen NF-κB – nuclear factor κ-light-chain-enhancer of activated B cells NG2 – neuron-glial antigen 2 nIPC – neurogenic intermediate progenitor cell NO – nitric oxide NOAEL – no observable adverse effect level NOS – nitric oxide synthase (iNOS = inducible NOS; eNOS = endothelial NOS) NPC – neural precursor cell NSAID – nonsteroidal anti-inflammatory drug NSC – neural stem cell NT3 – neurotrophin 3 OB – olfactory bulb OCT – organic cation transporter oIPC – oligodendrocytic intermediate progenitor cell Olig2 – oligodendrocyte transcription factor 2 OPC – oligodendrocyte progenitor cell p75NTR – p75 neurotrophin receptor Pax6 – paired box 6 PBS – phosphate-buffered saline

xii PCNA – proliferating cell nuclear antigen PD – Parkinson’s disease PEDF – pigment epithelium-derived factor PFA – paraformaldehyde PFK – phosphofructokinase PGL – periglomerular layer PI3K – phosphatidylinositol 3-kinase PIPES – 1,4-piperazinediethanesulfuronic acid PKA – protein kinase A PlGF – placental growth factor PSA-NCAM – polysialylated-neural cell adhesion molecule qNSC – quiescent neural stem cell qRT-PCR – quantitative reverse transcription polymerase chain reaction RG – radial glia RMS – rostral migratory stream RNS/ROS – reactive nitrogen/oxygen species ROI – region of interest S100B – S100 calcium-binding protein B SDF1 – stromal-derived factor 1 SEZ – subependymal zone SFM – serum-free media SFM+EFH – SFM + epidermal growth factor + basic fibroblast growth factor + heparin SFRP5 – secreted frizzled-related protein 5 SGZ – subgranular zone SLC – solute carrier Sox – sex determining region Y-box STAT6 – signal transducer and activator of transcription 6 STOM – short-term olfactory memory SVZ – subventricular zone

xiii TAP – transit amplifying progenitor TAp73 – transcriptionally active p73 Tbr2 – T-box brain protein 2 TEM – transmission electron microscopy TGF - transforming growth factor- TMZ – temozolomide TNF - tumour necrosis factor- TREM2 – triggered receptor expressed on myeloid cells 2 TrkB – tyrosine receptor kinase B Tuj1 – III-tubulin VCAM1 – vascular cell adhesion molecule 1 VZ – ventricular zone WT – wild type 5HT – 5-hydroxytryptamine (serotonin) 6-OHDA – 6-hydroxydopamine

xiv List of Figures Figure 1.1: Embryonic neurogenesis. Image from Kriegstein and Alvarez-Buylla, 2009 ……………. 5 Figure 1.2: SEZ-OB neurogenic axis. Image from Fiorelli et al., 2015 ……………………………………..... 8 Figure 1.3: Cytoarchitecture of SEZ. Image from Ihrie and Álvarez-Buylla, 2011 ……………………. 17 Figure 1.4: Cranial IR-induced cellular effects on NSC activation and signaling. Image from Chaker et al., 2016 …………………………………………………………………………………………………….. 38 Figure 1.5: Metformin-mediated aPKC-CBP pathway. Image from Potts and Lim, 2012 ………... 46 Figure 3.1. Schematic of long-term olfactory memory task ...... ………. 64 Figure 4.1: Metformin expands the neonatal, but not the juvenile aNSC pool in the SEZ ...... 66

Figure 4.2: Metformin expands the male and female neonatal aNSC pool ...... 67 Figure 4.3: Metformin induces a short-term expansion of the aNSC pool in the SEZ ...... 68 Figure 4.4: Spontaneous recovery of aNSCs occurs within 30 days post-IR ...... 69 Figure 4.5: Metformin pretreatment does not prevent the acute depletion of aNSCs 2 days post- IR …………………………………………………………………...... …………………………...... 70 Figure 4.6: Metformin pretreatment does not affect aNSC pool recovery following juvenile cranial IR ………………………………………………………………………...... …………………………...... 71 Figure 4.7: Neither metformin pretreatment nor juvenile cranial IR resulted in an altered ratio of aNSCs to qNSCs ………………………………………………………………...... ……………...... 73

Figure 4.8: Factors in the irradiated SEZ niche reduce aNSC pool size in vitro ...... 75 Figure 4.9: Metformin enhances the recovery of neuroblast proliferation in the SEZ following

juvenile cranial IR …………………………………………………………………...... …………….…...... 77 Figure 4.10: Metformin pretreatment rescues deficits to LTOM following juvenile cranial IR .. 79

xv List of Appendices

Appendix 1: Artificial cerebrospinal fluid (aCSF) ...... 127

Appendix 2: Serum-free media (SFM) ...... 128 Appendix 3: Reagents and solutions for FACS ...... 129

xvi 1

1. Literature Review 1.1. Regenerative Medicine The ability of organisms to regenerate tissues in response to damage is widespread and diverse, differing from species to species and from tissue to tissue. The regenerative capacity of mammals is relatively limited, and this is particularly highlighted in the central nervous system (CNS). However, recent advances in stem cell research and regenerative medicine have led to the emergence of tools and techniques that facilitate the regeneration of human cells, tissues, and organs. One of the most promising strategies for neural repair involves the activation of endogenous stem cells, with high likelihood for clinical success, given its potential to be minimally invasive relative to other regenerative strategies such as cell transplantation.

1.2. Neural Stem and Progenitor Cells 1.2.1. Initial characterization of neural stem cells For decades, it was the prevailing belief that neurogenesis (the generation of new neurons) halted after fetal development and did not persist in the postnatal and adult mammalian brain. This belief was first contradicted in the 1960s, when seminal work by Joseph Altman highlighted the existence of proliferating cells in the hippocampus (HPC) and surrounding the lateral ventricles (LVs) that were suggested to give rise to neurons in the dentate gyrus (DG) and olfactory bulb (OB), respectively (Altman, 1963; Altman and Das, 1965; Altman 1969). These cells would eventually become classified as neural stem and progenitor cells, together termed neural precursor cells (NPCs). The neural stem cell (NSC) is a multipotent stem cell and gives rise to all cell types of the nervous system, including neurons, oligodendrocytes, and astrocytes (Reynolds and Weiss, 1996). Given their role in neurogenesis, NSCs have been studied extensively since their initial identification and isolation from the postnatal mammalian brain, with the goals of understanding their behaviour under baseline and pathological conditions, and how they can be harnessed to repair the CNS. Despite Altman’s seminal work, it was not until the mid-1990s when the scientific community took seriously the notion of adult mammalian neurogenesis,

2 thanks to the in vitro isolation of NSCs by Brent Reynolds and Samuel Weiss in 1992. In these experiments, cells were isolated from the adult mouse brain, dissociated into single cells, and plated in the presence of epidermal growth factor (EGF) to form free-floating colonies termed neurospheres. Neurospheres could be passaged to exhibit self-renewal capacity, as well as differentiated into both astrocytes and neurons, displaying their multipotentiality (Reynolds and Weiss, 1992).

1.2.2. Existence of NSCs in the postnatal human and rodent brain In the postnatal mammalian brain and extending into adulthood, NSCs comprise rare populations within two neurogenic regions: the subgranular zone (SGZ) of the hippocampal dentate gyrus (DG) and the subependymal zone (SEZ) lining the lateral ventricles (LVs) (reviewed in Doetsch, 2003). The neurogenic niches have been well characterized in rodents throughout the lifespan, although their persistence in the postnatal human brain remains controversial. Neurogenesis in the adult human brain was first reported in a postmortem study involving brain tissue from cancer patients treated with the thymidine analog bromodeoxyuridine (BrdU), which labels DNA during S phase. Co-labeling of BrdU with neuronal nuclear antigen (NeuN) in the DG suggested the persistence of neurogenesis late into adulthood (Eriksson et al., 1998). Other studies have since corroborated this finding through a variety of approaches, including the use of proliferating cell nuclear antigen (PCNA) and G1 cell cycle marker Ki67 to label proliferating cells in the SEZ (Quiñones-Hinojosa et al., 2006), or polysialylated-neural cell adhesion molecule (PSA-NCAM) and the microtubule associated protein doublecortin (DCX) to label migrating neuronal progenitors or immature neurons termed neuroblasts from the adult SEZ (Curtis et al., 2007; Wang et al., 2011; Ernst et al., 2014) and DG (Knoth et al., 2010). Importantly, this phenomenon has been observed in humans as a response to ischemic stroke, where postmortem analyses found an increase in Ki67+ cells and neuroblasts in the SEZ ipsilateral to the infarct (Marti-Fabregas et al., 2010). Similarly, significant increases in cell proliferation have been observed in the postmortem SEZ of patients with multiple sclerosis (Nait-Oumesmar et al., 2007).

However, some studies suggest that postnatal neurogenesis sharply declines by 18 months of age (Sanai et al., 2011) and continues to decline throughout childhood (Dennis et al., 2016), although it is unclear and extremely challenging to investigate how functionally relevant these dynamics are in humans. Some of the most recently conducted human studies continue to draw conflicting conclusions, which demonstrates the enduring controversy surrounding this question and the need for further investigation (Sorrells et al., 2018; Boldrini et al., 2018). However, Moreno-Jiménez et al. attribute failure to identify human adult neurogenesis to inconsistent tissue fixation conditions and histological pretreatment of brain tissue and observed persistent DG neurogenesis throughout the human lifespan (Moreno-Jiménez et al., 2019). In order to specifically address the aims of this thesis, this literature review will focus only on SEZ neurogenesis in the mouse.

1.2.3. Neural stem cells during development During embryogenesis, neuroepithelial cells in the periventricular region, referred to as the ventricular zone (VZ), give rise to radial glia (RG) which are the NSCs of the developing fetal brain (reviewed in Kriegstein and Alvarez-Buylla, 2009). RG have apical-basal polarity with a single primary cilium on the apical process contacting the ventricle and a long basal process that contacts blood vessels and the meninges (pia) during development (Takahashi et al., 1990; reviewed in Kriegstein and Alvarez-Buylla, 2009). The majority of RG are produced during mid- fetal development, which persist as a relatively quiescent population of NSCs in the postnatal brain and into adulthood (Merkle et al., 2004; Fuentealba et al., 2015). In the developing human fetus, RG are identifiable by their expression of the intermediate filament proteins vimentin and glial fibrillary acidic protein (GFAP), as well as the glia-specific glutamate aspartate transporter (GLAST) (Antanitus et al., 1976; Choi and Lapham, 1978; Sasaki et al., 1988; Stagaard and Møllgård, 1989; Gould et al., 1990; Honig et al., 1996; Zecevic, 2004). Conversely, GFAP is not expressed in rodent RG (Bignami and Dahl, 1974; Sancho-Tello et al., 1995). Instead, rodent RG express the intermediate filament protein vimentin, GLAST, nestin, vascular cell adhesion molecule 1 (VCAM1), brain-lipid binding protein (BLBP), and the extracellular matrix (ECM) glycoprotein Tenascin-C (Dahl et al., 1981; Schnitzer

4 et al., 1981; Feng et al., 1994; Sancho-Tello et al., 1995; Götz et al., 1998; Hartfuss et al., 2001; Park et al., 2009), as well as the transcription factors sex determining region Y-box 2 (Sox2) and paired box 6 (Pax6) (Götz et al., 1998; Yuzwa et al., 2017). In mice, cortical neurogenesis occurs from approximately embryonic day 11 (E11) to E18, when RG divide asymmetrically to directly produce neurons or intermediate basal progenitors that go on to generate neurons (Noctor et al., 2001, 2004; Haubensak et al., 2004; reviewed in Malatesta et al., 2008). Cortical neurons are generated in an inside-out gradient, with deep layers being generated first, followed by outer layers. Earlier-born neurons migrate short distances to form the inner cortical layers (V-VI) whereas later-born neurons use the basal processes of RG as scaffolds to migrate radially towards the pial surface and form the outer cortical layers (II-IV) (reviewed in Nadarajah and Parnavelas, 2002). By E17.5, the majority of RG are no longer in cell cycle, although RG continue to make up many of the VZ cells at the time of birth. However, by P7, the proportion of RG is significantly reduced until their complete depletion by P15, as RG transition to postmitotic multi-ciliated FoxJ1+VCAM1- ependymal cells from ~E17.5 to P5 and astrocytes (Tramontin et al., 2003; Spassky et al., 2005; Hu et al., 2017; Yuzwa et al., 2017; Redmond et al., 2019). The RG-derived astrocytes lose vimentin expression during this time (P0-15), while becoming GFAP+ (Pixley and Vellis, 1984). These astrocytes maintain a VCAM1+ apical process with a single short cilium extending into the LV, as well as a basal process contacting blood vessels, and constitute the NSC pool that persists into adulthood (Doetsch et al., 1999a,b; Mirzadeh et al., 2008; Hu et al., 2017). The astrocytic cell bodies are separated from the LV lumen by a layer of ciliated ependymal cells, thereby creating two distinct layers — a single-cell thick ependymal layer (EL) and a subependymal zone (SEZ), often referred to as the subventricular zone (SVZ). The periventricular region (EL and SEZ) shrinks during the first two weeks of postnatal development, resembling the adult EL and SEZ by P15 (Tramontin et al., 2003).

Figure 1.1. Schematic of neurogenesis in the embryonic ventricular zone (VZ), which gives rise to the EL immediately adjacent to the LV and the SEZ (SVZ) which lies adjacent to the EL (Kriegstein and Alvarez-Buylla, 2009). CP = cortical plate, IZ = intermediate zone, MZ = marginal zone, nIPC = neurogenic intermediate progenitor cell, oIPC = oligodendrocytic intermediate progenitor cell, SVZ = subventricular zone, VZ = ventricular zone.

1.2.4. Neural stem cells during adulthood In the adult SEZ, which is the layer of cells separated from the LVs by the single-cell thick EL, NSCs persist as a relatively quiescent population of cells (Morshead et al., 1994) and are commonly referred to as type B cells. On the ventricular surface, NSCs extend an apical process that forms the centre of a pinwheel-like structure, surrounded by multiciliated (E1) and biciliated (E2) ependymal cells (Mirzadeh et al., 2008). Adult NSCs exist in either quiescent (non- proliferative; qNSCs) or activated (proliferative; aNSCs) states, with distinct protein expression signatures. Both qNSCs and aNSCs express GLAST, GFAP, Lewis X (LeX, also known as CD15 or SSEA-1), and prominin-1 (CD133), while only aNSCs express nestin, epidermal growth factor receptor (EGFR), and BLBP (Doetsch et al., 1997; Capela and Temple, 2002; Morshead et al., 2003; Mori et al., 2006; Pastrana et al., 2009; Platel et al., 2009; Beckervordersandforth et al., 2010; Daynac et al., 2013; Codega et al., 2014; Giachino et al., 2014; Morizur et al., 2018). In vitro, qNSCs and aNSCs are both multipotent, although qNSCs display low protein synthesis and rarely and slowly give rise to colonies, while aNSCs significantly upregulate global

6 protein synthesis and form colonies rapidly (Codega et al., 2014; reviewed in Baser et al., 2017).

While an initial estimate of NSC cell cycle time (TC) was ~15 days, this estimate pre-dated the discovery of qNSC and aNSC subpopulations and was likely limited to qNSCs that rarely divided

(Morshead et al., 1998). More recently, the cell cycle time (TC) of aNSCs has been reported to be ~17-18 hours, which includes an S phase length (TS) of 4 hours (Ponti et al., 2013). Since the discovery of these two cell states, the question of whether qNSCs and aNSCs comprise two independent cell populations has been addressed through flow cytometry and passaging experiments, suggesting that qNSCs and aNSCs can interconvert between states (Codega et al., 2014). Furthermore, single-cell transcriptomics have suggested that these cell states exist on a continuum, and that an intermediate “primed” state may exist, where NSCs upregulate genes associated with ribosomal biogenesis and increase protein synthesis prior to becoming proliferative and expressing the G1 cell cycle marker Ki67 (Llorens-Bobadilla et al., 2015; Shin et al., 2015; Dulken et al., 2017). Several factors contribute to the regulation of quiescence and activation of NSCs, including intrinsic transcriptional programs, vasculature, cell-cell and ECM communication, and diffusible signals, which can be modulated by various injuries and drugs (Hitoshi et al., 2002; Kazanis et al., 2010; Le Belle et al., 2011; Kokovay et al., 2012; Daynac et al., 2013; López-Juárez et al., 2013; Ottone et al., 2014; Chavali et al., 2018). Importantly, it is apparently necessary for a qNSC pool to be actively maintained, as unregulated NSC activation results in the depletion of NSCs (Imayoshi et al., 2010; Encinas et al., 2011). Certain cell cycle inhibitors, such as p21, are necessary to maintain NSC quiescence through the regulation of Sox2. The loss of p21 has been reported to deplete the NSC pool in vivo via replicative stress and reduce in vitro self-renewal capacity (Marqués-Torrejón et al., 2013). Certain molecules involved in cell-cell contact are also required to maintain a qNSC pool, such as VCAM1 and N-cadherin (Kokovay et al., 2012; Porlan et al., 2014). Furthermore, NSCs are suggested to self-regulate their activation through negative feedback, in which aNSCs expressing the Notch ligands Delta-like 1 (Dll1) and Dll3 directly contact qNSCs that highly express the Notch2 receptor to promote Notch-Hes signaling and maintenance of quiescence.

Similarly, direct inhibition of Notch increases NSC activation (Imayoshi et al., 2010; Kawaguchi et al., 2013; Llorens-Bobadilla et al., 2015). Similarly, the transition from quiescence to activation relies on numerous cell processes. N-cadherin-mediated adhesion of NSCs to ependymal cells contributes to quiescence, and the cleavage of N-cadherin by the metalloproteinase MT5-MMP is necessary for activation of NSCs in both physiological and regenerative conditions (Porlan et al., 2014). It has also been demonstrated that Wnt signaling can regulate NSC kinetics. Non-canonical Wnt signaling through Wnt5A increases N-cadherin expression in NSCs and activates the Rho-GTPase Cdc42, resulting in the maintenance of NSC polarity and adhesion, and therefore quiescence (Chavali et al., 2018). Using a BrdU label-retaining experiment to identify qNSCs, the direct inhibition of Cdc42 reduced qNSC pool size and increased proliferation in the SEZ, suggesting a disruption to quiescence, and the activation of NSCs (Chavali et al., 2018). Therefore, NSC quiescence is unlikely a passive cell state, and the active maintenance of qNSCs may be necessary to sustain adult neurogenesis throughout the mammalian lifespan.

1.3. The role of adult SEZ NSCs under baseline conditions NSCs in the SEZ give rise to constitutively proliferating progenitors known as transit- amplifying progenitors (TAPs) or C cells (Morshead and van der Kooy, 1992; Doetsch et al., 1999a,b). Single cell clonal analysis suggested that NSCs divide asymmetrically to self-renew and generate TAPs and downstream progeny of which >60% undergo cell death (Morshead et al., 1998). Under healthy conditions, the surviving TAPs give rise to neuroblasts (also known as A cells) that migrate 3-8mm along the rostral migratory stream (RMS) and differentiate into interneurons within the granule cell layer (GCL) and periglomerular layer (PGL) of the OB (Fig. 1.2) (Luskin, 1993; Lois and Alvarez-Buylla, 1994; Doetsch and Alvarez-Buylla, 1996; Lois et al., 1996; Craig et al., 1999; Fuentealba et al., 2015). Functionally, these newborn neurons in the OB are reported to underlie olfactory learning and memory (reviewed in Whitman and Greer, 2009; reviewed in Lazarini and Lledo, 2011).

Figure 1.2. SEZ-OB neurogenic axis (Fiorelli et al., 2015). (A-B) 3D representation of mouse brain with ventricles highlighted in grey and NPCs (including their progeny) in orange. (C) Diagrams of coronal sections highlighted in (B) with NPCs and their progeny in orange. GCL = granule cell layer, GL = glomerular layer, LV = lateral ventricle, OB = olfactory bulb, RMS = rostral migratory stream, SVZ = subventricular zone (SEZ).

1.3.1. Neural stem and progenitor pool maintenance Morshead et al. used high doses of 3H-thymidine to kill actively dividing NPCs in the SEZ and showed that NSCs are recruited to repopulate the NPC pool to baseline levels within 8 days, and that most NSCs reverted to quiescence within 4 days (Morshead et al., 1994). This was one of the first indicators of both a quiescent and an activated NSC state, although the identification of unique expression profiles and isolation of qNSCs and aNSCs were not accomplished for another 15 years (Pastrana et al., 2009; Codega et al., 2014). A number of studies have since recognized the regenerative potential of the SEZ in vivo using a variety of techniques. Doetsch and colleagues used the antimitotic agent cytosine--D- arabinofuranoside (Ara-C) to eliminate progenitors, and showed that NSCs spontaneously regenerate the SEZ-RMS network of progenitors and neuroblasts within 10 days, thereby highlighting the plasticity and resilience of the SEZ neurogenic niche (Doetsch at al., 1999a,b). To identify the source of regeneration in the SEZ, Codega and colleagues confirmed that Ara-C

9 infusion resulted in complete ablation of nestin+GFAP+ aNSCs, and that nestin- qNSCs were responsible for the regeneration of both aNSCs and neuroblasts (Codega et al., 2014). Therefore, the maintenance and recruitment of endogenous SEZ qNSCs is a promising therapeutic strategy, which has the potential to replace lost cells following certain brain injuries. The mechanism underlying the generation of TAPs involves the symmetry of NSC division. It has long been accepted that aNSCs can divide symmetrically in vitro to give rise to identical daughter cells, thereby expanding the NSC pool (Reynolds and Weiss, 1992; Gritti et al., 1996). Conversely, NSCs predominantly give rise to TAPs in vivo, which suggests the absence of a positive signal in the SEZ that induces symmetric self-renewal (Morshead et al., 1998; Obernier et al., 2018). For several years, it has been reported that NSCs divide asymmetrically in vivo to self- renew and generate a progenitor cell, which would subsequently divide symmetrically to maintain the progenitor pool or eventually migrate to the OB via the RMS (Morshead et al., 1998; Calzolari et al., 2015). This seems highly plausible, given the robust capacity of NSCs to regenerate downstream progenitor and neuroblast pools without becoming depleted (Morshead et al., 1994; Doetsch et al., 1999a,b). The asymmetric mode of division has recently been challenged by Obernier et al., who reported that NSCs rarely or never undergo asymmetric division, but instead divide symmetrically to self-renew (20-30% occurrence), or undergo a consuming symmetrical division to generate a pair of TAPs (70-80% occurrence) at the expense of the parent NSC (Obernier et al., 2018). However, this study relied on ex vivo time-lapse imaging from juvenile mice, in which environmental factors that may influence the mode of NSC division were potentially absent. In contrast, the majority of literature pertaining to the symmetry of NSC division continues to support the existence of asymmetric divisions (Karpowicz et al., 2005; Bonaguidi et al., 2011; Costa et al., 2011; Calzolari et al., 2015). Costa and colleagues propose a lineage in which relatively quiescent NSCs with an in vitro cell TC of ~53 hours symmetrically divide to generate 2-4 quickly dividing NSCs, that divide either symmetrically to generate 2 TAPs, or asymmetrically to generate 1 NSC and 1 TAP (Costa et al., 2011).

Much of the conflict surrounding NSC division symmetry may be a product of inconsistent use of in vitro and in vivo models and the methods used to label and track dividing NSCs. Given that the SEZ neurogenic niche changes dramatically with age, it is also possible that the mode of NSC division changes with age and environment (Ruddy and Morshead, 2018). Despite uncertainties regarding the symmetry of NSC division under physiological conditions, it is possible to modulate NSC division symmetry. Enhancing Wnt signaling expands the SEZ NSC pool size by promoting self-renewing symmetric divisions, without affecting cell survival or TAPs (Piccin and Morshead, 2011). Therefore, targeting the symmetry of NSC division may be a promising approach to improving the regenerative capacity of the mammalian brain.

1.3.2. In vivo NSC lineage and olfactory bulb neurogenesis Similar to aNSCs, TAPs continue to express nestin and EGFR, although GFAP expression is lost (Karpowicz et al., 2005; Codega et al., 2014; Calzolari et al., 2015). TAPs are also characterized by the expression of mammalian achaete scute homolog (Mash1/Ascl1) (Parras et al., 2004). TAPs were initially estimated to have a TC of 12.7 hours via quantification of cumulative BrdU incorporation over time within coronal brain sections, although the TC of TAPs has since been re-evaluated using whole mounts and 3 cell cycle analyses to be 18-25 hours with a long TS of 14-17 hours (Morshead and van der Kooy, 1992; Ponti et al., 2013). Following the generation of TAPs from NSCs, double thymidine analog and percentage of labeled mitoses methods have been used to demonstrate that TAPs undergo 3-4 rounds of proliferative symmetric divisions prior to a consuming symmetric division generating DCX+ neuroblasts at the expense of the TAP (Costa et al., 2011; Ponti et al., 2013). Neuroblasts are characterized by an elongated cell body with radial morphology, as well as DCX, PSA-NCAM, TuJ1 (III-tubulin), and CD24 expression (Lois and Alvarez-Buylla, 1993;

Calaora et al., 1996; Doetsch et al., 1997). Under physiological conditions, neuroblasts have a TC of ~18 hours, and divide 1-2 times before migrating away from the SEZ (Ponti et al., 2013). Interestingly, neuroblasts in the SEZ act as a major source of gamma-aminobutyric acid (GABA) which activates GABAA receptor (GABAAR) signaling in NSCs to maintain quiescence in a

11 feedback system that likely prevents qNSC pool depletion under physiological conditions (Liu et al., 2005; Daynac et al., 2013). Neuroblasts migrate 3-8mm rostrally in chains of 30-40 cells parallel to the lateral wall of the SEZ, which form the RMS that underlies the corpus callosum and extends into the OB. Along the RMS, neuroblasts are elongated, interconnected, and ensheathed in a glial tube of astrocytes (Rousselot et al., 1995; Lois et al., 1996; Doetsch et al., 1997). This chain structure of migrating neuroblasts is necessary for directional migration to the OB and is PSA-NCAM- dependent (Battista and Rutishauser, 2010). Certain chemoattractant factors also support this directional migration, such as the expression of insulin-like growth factor-1 (IGF-1) in the OB (Hurtado-Chong et al., 2009), as well as secretion of netrin by mitral cells of the OB and interaction with its receptor Deleted in Colorectal Carcinoma (DCC) on neuroblasts (Murase and Horwitz, 2002). A rostral to caudal gradient of the chemorepulsive factor Slit2 also supports the migration of neuroblasts from the SEZ into the RMS (Nguyen-Ba-Charvet et al., 2004). From their origin in the SEZ, neuroblasts migrate at ~26-30m/hour along the RMS, reaching the OB after 5-7 days (Lois and Alvarez- Buylla, 1994; Luskin and Boone, 1994; Craig et al., 1999). Within the GCL of the OB, the olfactory activity-dependent expression of the ECM glycoprotein Tenascin-R causes the detachment of neuroblasts from the RMS and concurrent radial migration along vessels in the OB, where neuroblasts terminally differentiate into predominantly GABAergic interneurons within the GCL and to a lesser extent, the PGL (Rousselot et al., 1995; Betarbet et al., 1996; Lois et al., 1996; Zigova et al., 1996; Carleton et al., 2003; Saghatelyan et al., 2004; Bovetti et al., 2007). Newborn OB granule neurons receive GABAergic inputs within 3 days following their arrival in the OB and functionally integrate within 2-4 weeks (Petreanu and Alvarez-Buylla, 2002; Carlén et al., 2002; Carleton et al., 2003; Panzanelli et al., 2009). Upon maturation, neurons lose DCX expression and express microtubule associated protein-2 (MAP2), as well as neuronal nuclear marker NeuN, which is observed in the GCL within two weeks of neuroblast formation in the SEZ (Mullen et al., 1992; reviewed in Johnson and Jope, 1992; Zigova et al., 1998; Magavi et al., 2005; Imayoshi et al., 2008). The survival of adult-born mature OB granule neurons is activity dependent and

12 sustained for at least 4 months, even in aged mice (9 months old), despite the arrest of OB growth at 3-4 months of age (Pomeroy et al., 1990; Corotto et al., 1993; Petreanu and Alvarez- Buylla, 2002). Approximately 30-70% of newborn OB neurons undergo apoptosis during synaptogenesis, although increased OB activity via odorant exposure and olfactory learning increase survival and integration rates (Rochefort et al., 2002; Yamaguchi and Mori, 2005; Lemasson et al., 2005; Alonso et al., 2006; Kim et al., 2007; Mouret et al., 2008). Interestingly, the investigation of anosmic mice (unable to smell) revealed normal generation and migration of adult-born neuroblasts; however, olfactory input was necessary for the survival of GCL neurons during synaptic integration and dendritic spine formation (Corotto et al., 1994; Petreanu and Alvarez-Buylla, 2002). Therefore, adult OB neurogenesis may respond and contribute to changes in olfactory experiences. In the rodent OB, it has been proposed that adult neurogenesis is required to continuously replace old GCL neurons, as the OB’s size remains relatively stable throughout adulthood (Kaplan et al., 1985). Indeed, by permanently labeling all nestin+ NSC-derived cells in young adult mice, Imayoshi et al. reported that nearly all deep GCL neurons are replaced, and that adult-born neurons comprise ~50% of superficial GCL neurons after 10-12 months (Imayoshi et al., 2008). Hence, OB neurons generated during P3-7 predominantly settle in the superficial GCL and survive significantly longer than young adult-born neurons (Lemasson et al., 2005). Taken together, these studies suggest that adult neurogenesis contributes to cell addition in the superficial GCL and cell replacement in the deep GCL under physiological conditions throughout life.

1.3.3. The role of neurogenesis in olfactory behaviour The functions of OB neurogenesis in rodents are controversial and variably considered to be involved in different forms of olfactory discrimination, learning, and memory (reviewed in Whitman and Greer, 2009; reviewed in Lazarini and Lledo, 2011). A general lack of standardized and reproducible olfactory behaviour tasks has resulted in the use of different experimental paradigms which make it challenging to assess similar behaviours. For example, Ghuesi et al.

13 disrupted neuroblast migration via NCAM deficiency (Cremer et al., 1994), which resulted in GCL volume reduction and impaired olfactory discrimination between familiar and novel odors, with odor detection thresholds and short-term olfactory memory (STOM) being unaffected (Ghuesi et al., 2000). In contrast, Pan et al. deleted the extracellular signal-regulated kinase 5 (ERK5) mitogen-activated protein kinase (MAPK) in nestin+ cells within both neurogenic niches of the brain to disrupt neurogenesis, and this resulted in deficits to STOM, odor and pheromone detection thresholds, and odor-cued associative learning, while olfactory discrimination was unaffected (Pan et al., 2012a,b; Zou et al., 2012). The use of intraventricular Ara-C infusion to ablate neurogenesis has also demonstrated a necessity of OB neurogenesis for STOM and long- term olfactory memory (LTOM) (Breton-Provencher et al., 2009; Sultan et al., 2010). Other approaches to identify the functions of adult OB neurogenesis typically rely on gain-of-function manipulations via olfactory enrichment experiments designed to target postnatally-derived GCL neurons. Young adult mice housed in an odor-enriched environment for 40 days exhibited enhanced survival of newborn GCL neurons which correlated with extended STOM relative to control mice (Rochefort et al., 2002). Similarly, by rearing neonatal (P0-9) mice in a citral-scented environment, mice acquired a specific preference for citral that was sustained after 2 months and was associated with increased neonatal, but not adult GCL neurogenesis. This finding suggests that neurogenesis coincides with the acquisition of LTOM (Lemasson et al., 2005). A study by Moreno et al. reported that improved olfactory discrimination following repeated odorant exposure was associated with increased activity and survival of newborn GCL GABAergic interneurons, which was inhibited by blocking neurogenesis via Ara-C infusion before and during odorant exposure, thereby suggesting the necessity of neurogenesis for perceptual olfactory memory (Moreno et al., 2009). In rodents, neurogenesis-dependent olfactory perceptual learning may be crucial throughout adulthood for reproductive success, as it is reportedly necessary for female preference of dominant-male pheromones, recognition of mates by females, and paternal recognition of offspring (Mak et al., 2007; Mak and Weiss, 2010; Oboti et al., 2011).

Despite some discrepancies amongst studies, the majority of findings suggest functionally significant roles for turnover of OB GABAergic interneurons throughout adulthood and that newly generated adult-born neurons in the OB undergo distinct modifications in response to olfactory experiences (Lazarini and Lledo, 2011). These features provide a strong basis for the likelihood that adult OB neurogenesis underlies olfactory discrimination, learning, and memory in various contexts. Therefore, the SEZ-OB neurogenic axis and its putative behavioural implications provide a robust experimental model to investigate the potential to manipulate neurogenesis and the brain’s capacity for repair.

1.3.4. Neurogenesis vs. gliogenesis Although the majority of adult-born SEZ-derived cells become GABAergic interneurons in the OB, a small proportion of newborn SEZ-derived cells generate oligodendrocytes in the corpus callosum, striatum, and fornix, thereby making the SEZ a potential target for enhancing repair in demyelinating diseases (Hack et al., 2005; Marshall et al., 2005; Menn et al., 2006; Xing et al., 2014). In NPCs within the SEZ, the transcription factors Pax6 and Olig2 are major determinants of whether a TAP gives rise to neuroblasts or oligodendrocyte progenitor cells (OPCs) (Hack et al., 2005). Pax6 overexpression in NPCs increases neuroblast formation in vitro and in vivo, while its conditional deletion or expression of its dominant-negative form reduces neuroblast formation (Hack et al., 2004; Hack et al., 2005). In contrast, Olig2 promotes TAP-derived generation and migration of Sox10+ OPCs into the corpus callosum, where they differentiate into adenomatous polyposis coli (APC) clone CC1+ (also known simply as CC1+) oligodendrocytes at the expense of neuroblast formation and OB neurogenesis. Similarly, the expression of loss- of-function or dominant-negative (Olig2VP16) forms of Olig2 increase neuroblast formation and completely deplete oligodendrogenesis in the corpus callosum (Hack et al., 2005). Distal-less homeobox 2 (Dlx2) is another transcription factor involved in neurogenesis, which is necessary for the specification of TAPs to a neuronal fate. Interestingly, Dlx2 also interacts with Pax6 in RMS neuroblasts to shift adult-born OB neurons to a dopaminergic fate in the PGL at the expense of GABAergic GCL interneurons (Hack et al., 2005; Brill et al., 2008;

Bonzano et al., 2016). Other pathways implicated in neuronal fate decision include bone morphogenic protein (BMP)-mediated signaling in NSCs via Smad4, which is required for neurogenesis and suppression of oligodendrogenesis (Colak et al., 2008), as well as Wnt/- catenin and fibroblast growth factor-2 (FGF2) signaling, which increase OPC generation, survival, and differentiation from the SEZ (Azim and Butt, 2011; Azim et al., 2012b; Azim et al., 2014b). Spontaneous remyelination has been observed in multiple sclerosis patients and rodent models of demyelinating injury, but parenchymal OPCs (pOPCs) have historically been considered the source of adult oligodendrogenesis (Prineas and Connell, 1979; Gensert and Goldman, 1997). However, it is now recognized that the SEZ contributes to remyelination and that chronic demyelination can deplete SEZ NCSs and reduce TAP proliferation, inciting the need to better understand the mechanisms underlying the balance between SEZ oligodendrogenesis and neurogenesis in the SEZ (Nait-Oumesmar et al., 2007; Brousse et al., 2015).

1.4. The SEZ NSC Niche Within the SEZ, the maintenance and behaviour of NSCs and their progeny rely on interactions with the ECM, cerebrospinal fluid (CSF) in the LVs, factors released by the intraventricular choroid plexus, vasculature, microglia, and ependymal cells, as well as secreted factors from neighbouring and distant cells (reviewed in Doetsch, 2003; Riquelme et al., 2008; Adams and Morshead, 2018; Ruddy and Morshead, 2018). This neurogenic niche undergoes changes throughout development and aging, as well as in response to injuries, thereby influencing cell kinetics and neurogenesis. In order to preserve or manipulate neurogenesis, an understanding of niche dynamics under physiological and pathological conditions is required.

1.4.1. Cytoarchitecture The postnatal SEZ is 3-4 cells thick and is separated from the lumen of the LVs by an epithelial monolayer of ependymal cells, which are predominantly multiciliated and organized in pinwheel structures around the apical process of NSCs (Doetsch et al., 1997; Doetsch, 2003;

Sawamoto et al., 2006; Mirzadeh et al., 2008). The LVs contain the choroid plexus that produces CSF, which flows in a rostral direction. Furthermore, an extensive network of serotonergic axons coats the apical surface of the LV walls (Sawamoto et al., 2006; Tong et al., 2014). SEZ astrocytes can be divided into two types based on morphology, position, and function. Type B1 astrocytes are NSCs that have radial morphology, with an apical process punctuating the centre of ependymal cell pinwheels and extending a short primary cilium into the LV (Mirzadeh et al., 2008). A basal process extends and terminates in an endfoot contacting the endothelial cells of a planar vascular plexus lying parallel to the LV wall (Shen et al., 2008; Tavazoie et al., 2008). Type B2 astrocytes do not extend an apical process into the LV and are located closer to the striatum and vasculature (Doetsch et al., 1997). NSCs give rise to non- radial, rounder TAPs that also reside in close proximity to blood vessels within the SEZ. In turn, TAPs give rise to neuroblasts that arrange in long migratory chains ensheathed by type B1 (NSCs) and B2 astrocytes, forming the RMS that snakes along the vasculature towards the OB (Doetsch et al., 1997; Ihrie and Alvarez-Buylla, 2011). The SEZ also houses microglia, the resident immune cell of the brain that are ramified under physiological conditions (Walton et al., 2006; reviewed in Ekdahl et al., 2009; reviewed in Su et al., 2014). All cell types within the SEZ contact the ECM, which anchors NPCs in place and controls growth factor availability to modulate proliferation kinetics (Mercier et al., 2002; Kerever et al., 2007; Shen et al., 2008). The cellular and extracellular organization supporting NPC survival, proliferation, and differentiation is dynamic and heterogeneous, providing several potential targets for therapeutic manipulation.

Figure 1.3. Cytoarchitecture of SEZ (Ihrie and Álvarez-Buylla, 2011). A = neuroblast, B1 = neural stem cell, B2 = SEZ astrocyte, Bv = blood vessel, C = transit amplifying progenitor, E = ependymal cell, Mg = microglia.

1.4.2. Ependymal Cells and Cerebrospinal Fluid The majority of ependymal cells are generated from E14-16 by RG and become multiciliated from P0-4, both occurring in a caudorostral progression. Although they arise from the same RG that generate the postnatal NSC pool, ependymal cells are postmitotic (Spassky et al., 2005; Muthusamy et al., 2018; Shah et al., 2018; Redmond et al., 2019). During maturation, ependymal cells expand their apical surface and organize in pinwheel structures in a caudorostral progression from P5-10, which are maintained in adulthood (Mirzadeh et al., 2008; Redmond et al., 2019). Ependymal cells express CD133, FoxJ1, CD24 (also expressed by neuroblasts), nestin, and the calcium-binding protein S100B, which is also expressed by mature astrocytes, but not NSCs (Didier et al., 1986; Calaora et al., 1996; Doetsch et al., 1997; Chiasson

18 et al., 1999; Lim et al., 2000; Raponi et al., 2007; Jacquet et al., 2009; Muthusamy et al., 2018; Shah et al., 2018). Ependymal cells serve as a barrier between the CSF-filled ventricles and also to maintain homeostasis within the neurogenic niche through cell-cell contact with NSCs and indirect CSF- mediated effects. NSCs express VCAM1 on their apical endfeet, which is required to maintain adhesion to ependymal cells. Loss of VCAM1 results in NSC detachment from ependymal cells, disrupted pinwheel architecture, and subsequent activation and depletion of NSCs (Kokovay et al., 2012). Ependymal cells produce Noggin, a BMP antagonist (McMahon et al., 1998). NSCs express BMPs and their receptors (BMPRs), permitting BMP-Smad signaling which reduces in vitro and in vivo neurogenesis, and instead promotes glial differentiation; however, this is rescued in the presence of Noggin (Lim et al., 2000). In contrast, it has been reported that Smad4 deletion in SEZ NSCs significantly increases Olig2+ OPC production and corpus callosum oligodendrogenesis at the expense of neuroblasts, suggesting that tight regulation of BMP- Smad signaling in NSCs is necessary for normal levels of SEZ neurogenesis, since its aberrant upregulation or downregulation both reduce neurogenesis (Colak et al., 2008). Therefore, Noggin secretion by ependymal cells may serve to stabilize BMP-regulated neurogenesis. Planar polarity of ependymal cells established during early postnatal ependymal cell maturation is required for proper orientation of cilia and CSF flow, as well as prevention of hydrocephalus (Ohata et al., 2014; Ohata et al., 2015). The synchronized beating of ependymal cilia is required for the normal caudal to rostral flow of CSF, which supports a rostral to caudal gradient of the chemorepulsive factor Slit2 in the LV and dorsal SEZ. This gradient is necessary for proper migration of neuroblasts from the SEZ into the RMS (Nguyen-Ba-Charvet et al., 2004; Sawamoto et al., 2006). Despite initial reports characterizing ependymal cells as NSCs that contribute to continuous baseline neurogenesis or become proliferative and neurogenic in response to injuries (Johansson et al., 1999; Coskun et al., 2008; Carlén et al., 2009), it has since been convincingly demonstrated that ependymal cells do not possess NSC qualities in vitro or in vivo (Doetsch et al., 1999a; Muthusamy et al., 2018; Shah et al., 2018). Earlier studies often relied on

19 putative ependymal cell markers that were also expressed in some NSCs (CD133, FoxJ1), or labeled all cells contacting the CSF and failed to account for the LV-contacting apical process of NSCs, thereby mischaracterizing some bona fide NSCs as ependymal cells with NSC characteristics (Jacquet et al., 2009; Beckervordersandforth et al., 2010; Luo et al., 2015; Shah et al., 2018). Taken together, ependymal cells and CSF flow regulate the maintenance, proliferation, differentiation, and migration of NPCs. Although the specific roles of CSF components in regulating neurogenesis are less clear, CSF-borne insulin-like growth factors (IGFs) and inflammatory cytokines of the interleukin (IL) family have been proposed as potential regulators of NPC survival and proliferation (Lehtinen et al., 2011; reviewed in Singhal et al., 2014).

1.4.3. Microglia Microglia are the resident macrophages of the brain, comprising ~10% of CNS cells and are highly ramified, but rarely overlap with each other. Their motile ramifications survey their environment to detect and respond to extracellular signals to maintain homeostasis (Salter and Stevens, 2017). Although microglia have historically been considered to strictly be involved in CNS inflammation and relatively silent in the healthy brain, recent studies have revealed a variety of microglial states and responses distinct from inflammation (reviewed in Salter and Beggs, 2014). RNA sequencing and epigenetic analyses of microglia throughout development suggest three developmental stages of microglia associated with neuronal development: early microglia (E10.5-14) which shift to pre-microglia (from E14 to a few weeks after birth) implicated in synaptic pruning and neuronal maturation, and adult microglia (4 weeks and older) important for tissue maintenance and homeostasis (Matcovitch-Natan et al., 2016). Pre-microglia during early postnatal development also secrete the cytokines IL-1, IL-6, tumor necrosis factor- (TNF), interferon- (IFN), and IGF-1, which play differing roles in promoting early postnatal neurogenesis, oligodendrogenesis, and neuroblast migration (Hurtado-Chong et al., 2009; reviewed in Gonzalez-Perez et al., 2012; Shigemoto-Mogami et al., 2014).

Microglial activation occurs via 2 pathways: the classical M1 inflammatory and often cytotoxic pathway, or the alternative anti-inflammatory M2 pathway involved in cleanup, regeneration, and potentially neuroprotection (reviewed in Franco and Fernández-Suárez, 2015). One role of microglia is the phagocytosis of dead and dying neurons, although recent studies have demonstrated that microglia may also release neurotoxic factors to induce apoptosis and phagocytosis of stressed-but-viable neurons – a process termed phagoptosis (reviewed in Marín-Teva et al., 2012; Brown and Neher, 2014). Regarding the direct effects of microglia on the neurogenic niches, most studies have been restricted to hippocampal neurogenesis, which suggest a prominent regulatory role of microglia via apoptosis of NPCs (Sierra et al., 2010; Bachstetter et al., 2011; Gemma and Bachstetter, 2013). However, the SEZ and RMS are reported to contain a unique population of microglia with enlarged cell bodies and fewer, thicker ramifications that express the chemokine receptor CX3CR1, and express lower levels of typical markers of microglia (Iba1) and microglial activation (TREM2, CD68/ED1) in addition to being ~5 times more proliferative in the SEZ/RMS than in the striatum or corpus callosum (Jung et al., 2000; Goings et al., 2006; Ribeiro Xavier et al., 2015). SEZ microglia are positioned in close contact with NSCs, TAPs, neuroblasts, and vasculature (Mosher et al., 2012; reviewed in Matarredona et al., 2018). Studies have demonstrated that microglia in the neurogenic niche are necessary for NPC survival and migration. Microglial ablation results in a buildup of apoptotic neuroblasts and impaired migration along the RMS, and subsequent reduction in OB neurogenesis (Ribeiro Xavier et al., 2015). Neuroblast phagocytosis is rare in the RMS, suggesting that microglia in the SEZ and RMS provide trophic support to induce survival. This is supported by their expression of phosphorylated STAT6 (pSTAT6), IL-4, and IL-10 which are typical of M2 microglia. In contrast, microglial activation (amoeboid morphology, TREM2+) and phagocytosis are robust in the OB where interneuron turnover and apoptosis during maturation of newborn neurons occur (Ribeiro Xavier et al., 2015). The apparent supportive roles of microglia in the SEZ and RMS are also observed in vitro, where repeated passaging of NPCs reduces neuronal differentiation capacity, which is rescued by coculture with microglia (Walton et al., 2006). Together, these studies reveal an important role of microglia in regulating NPC behaviours.

1.4.4. Vasculature The SEZ contains a planar vascular plexus with a modified blood-brain barrier (BBB) that periodically lacks NG2+ pericyte and aquaporin 4-expressing (AQP4+) astrocytic endfeet coverage, allowing for direct contact of CD31+ vascular endothelial cells with NSCs and TAPs (Shen et al., 2008; Tavazoie et al., 2008). Following Ara-C-mediated ablation of proliferating cells, the majority of newly generated NPCs directly contacted endothelial cells, suggesting that the vasculature serves as a scaffold for regeneration in the SEZ (Tavazoie et al., 2008). The vasculature may serve not only to deliver blood-borne nutrients, hormones, or growth factors to cells within the SEZ, but the endothelial cells themselves directly regulate NSC behaviour through secreted factors and cell-cell contact or juxtacrine signaling (Shen et al., 2004; Shen et al., 2008; Snapyan et al., 2009; Delgado et al., 2014; Crouch et al., 2015; Ottone and Parrinello, 2015). Endothelial cells were first shown to modify NSC behaviour when Shen et al. cocultured them with NSCs, resulting in enhanced NSC symmetric self-renewal capacity through Notch-Hes1 signaling (Shen et al., 2004). This was later proposed to be caused by pigment epithelium-derived factor (PEDF) in vivo, which is secreted by endothelial cells (Ramírez-Castillejo et al., 2006). Numerous other endothelial cell-derived factors have been reported to support NSC maintenance and progenitor proliferation, such as neurotrophin 3 (NT3) which promotes quiescence and placental growth factor 2 (PlGF2) which stimulates aNSC and TAP proliferation (Delgado et al., 2014; Crouch et al., 2015). Shen et al. demonstrated that the ECM protein laminin is highly expressed by vascular cells in the SEZ, and that NSC expression of laminin receptor 61 integrin is necessary for their adhesion to endothelial cells on cultured monolayers and in vivo. Infusion of an antibody blocking 61 integrin into the LVs caused a significant increase in proliferating cells in the dorsal SEZ (Shen et al., 2008). Mechanistically, this effect is mediated by endothelial membrane-bound ligands ephrinB2 and Jagged1, which suppress MAPK-mediated cell-cycle entry and promote transcription of qNSC-associated genes in NSCs, respectively (Ottone et al., 2014). Together, these findings suggest that endothelial cell adhesion is required for proper NSC pool maintenance.

Furthermore, endothelial cells and ependymal cells both secrete the chemokine stromal-derived factor 1 (SDF1), which binds its receptor CXCR4 on NPCs, specifically upregulating EGFR and 6 integrin in aNSCs and TAPs, as well as increasing neuroblast motility in vitro (Kokovay et al., 2010). Endothelial cells also secrete betacellulin (BTC), which activates EGFR and ErbB4 in TAPs and neuroblasts, respectively, thereby increasing TAP proliferation via the MEK/ERK pathway and neuroblast proliferation via Akt signaling (Gómez-Gaviro et al., 2012). Accordingly, vasculature is longitudinally aligned with the RMS and endothelial cells have been reported to directly contact both neuroblasts and the astrocytic arborizations of the glial tube (Whitman et al., 2009). The endothelial cells along the RMS secrete brain-derived neurotrophic factor (BDNF) which binds the tyrosine receptor kinase B (TrkB) on astrocytes and the p75 neurotrophin receptor (p75NTR) on neuroblasts to maintain their chain-like migratory phenotype towards the OB (Snapyan et al., 2009). BDNF has also been demonstrated to enhance neuroblast survival in vitro and increase neuroblast arrival in the OB following infusing in the LV (Kirschenbaum and Goldman, 1995; Zigova et al., 1998). Together, these findings reveal critical roles of vascular endothelial cells in both SEZ NSC pool maintenance and OB neurogenesis.

1.4.5. Extracellular matrix (ECM) In addition to the cell-cell signaling-mediated regulation of SEZ-OB neurogenesis, the ECM also plays a significant role in the behaviour of NSCs and their progeny (Gattazzo et al., 2014). One of the best-documented features of the SEZ ECM are fractones, which are extensions of the vascular basement membrane comprised of laminin, collagen IV, nidogen, and heparan sulphate proteoglycan (HSPG) (Mercier et al., 2002; Kerever et al., 2007). Interestingly, HSPG-containing fractones entrap the mitogen FGF2 and are necessary for FGF2- mediated proliferation in the adult SEZ (Douet et al., 2013). As previously mentioned, laminin is abundant near endothelial cells in the SEZ and promotes 61 integrin-mediated adhesion of NSCs to the vasculature for proper NSC pool maintenance (Shen et al., 2008). In addition to NSC pool maintenance, ECM components such as the glycoprotein Reelin, play a significant role in neuronal migration and positioning during

23 development (Frotscher, 2010). Deletion of Reelin has been reported to disrupt NSC-derived neuroblast generation and chain migration of NPCs in vitro, and significantly reduces neuroblast migration through the RMS (Won et al., 2006; Massalini et al., 2009). The ECM glycoprotein Tenascin-R, which mediates neuroblast detachment and migration away from the RMS in the OB, is expressed in an olfactory activity-dependent manner, and its ectopic expression is sufficient to reroute neuroblasts (Saghatelyan et al., 2004). Furthermore, ECM stiffness alters NPC differentiation (Saha et al., 2008; Keung et al., 2011), suggesting that ECM components influence NSC pool maintenance, as well as the proliferation and differentiation of their progeny.

1.4.6. Neurotransmitters Neurotransmitter signaling in the adult brain has long been associated primarily with communication between differentiated neurons, although its role in neurogenesis has become more prominent in recent years. Diverse neurotransmitter signaling exists in both neurogenic niches, although the types of signaling and subsequent effects vary significantly between the SEZ and DG (reviewed in Platel et al., 2010; Berg et al., 2013). D2-like (D2L) dopamine (DA) receptors are expressed by TAPs and neuroblasts in the SEZ, the former receiving DA afferents from the substantia nigra. Using the selective DA neurotoxins MPTP or 6-hydroxydopamine (6-OHDA), ablation of DA neurons reduced proliferation in the SEZ, which was rescued with 3 weeks of treatment with the DA precursor levodopa (L-DOPA) or a single injection of the D2L agonist ropinirole (Höglinger et al., 2004). Several studies have reported that presence of DA or D2L agonists also increases baseline SEZ proliferation, as well as in vitro TAP proliferation, which is blocked by D2L antagonists, suggesting that DA signaling directly enhances NPC proliferation (Baker et al., 2004; Höglinger et al., 2004; O’Keeffe et al., 2009; Winner et al., 2009; Kim et al., 2010). In addition to DA, serotonergic signaling plays an important role in the SEZ. This was first demonstrated when inhibition of serotonin (5-hydroxytryptamine (5HT)) production or lesions to 5HT neurons in the raphe nuclei were reported to reduce SEZ proliferation (BrdU+ labeling) by 50% (Brezun and Daszuta, 1999). Both acute and chronic administration of 5HT1A or 5HT2C

24 receptor agonists increase SEZ proliferation and subsequent OB neurogenesis, without affecting differentiation profiles, suggesting a proliferative effect of 5HT in the SEZ (Banasr et al., 2004). More recently, the use of a retrograde tracer in the lumen of the LVs allowed Tong et al. to identify an extensive network of 5HT axons lying on the apical surface of the LV walls originating from the raphe nuclei of the midbrain. NSCs were observed via transmission electron microscopy (TEM) to apically extend microvilli that ensnared 5HT axonal varicosities. Despite the absence of synaptic communication, NSCs express the 5HT2C receptor which positively regulates proliferation in the SEZ (Tong et al., 2014). Similar to 5HT signaling, GABA is released and perceived within the SEZ in a non-synaptic fashion (Wang et al., 2003; Liu et al., 2005). GABA is released from neuroblasts via spontaneous depolarizations, inducing GABAAR signaling and inhibition of cell cycle progression in qNSCs, which has been proposed as a feedback system to control NSC pool depletion (Liu et al., 2005;

Daynac et al., 2013). Mechanistically, GABAAR activation using the GABAAR agonist muscimol activates the ATM/ATR PI3K-related kinases, resulting in phosphorylation of histone H2AX, an epigenetic modification that reduces NPC proliferation and OB neurogenesis (BrdU+NeuN+ cells). However, this can be reversed with the GABAAR antagonist bicuculline in vitro and in vivo, resulting in long-lasting increases to OB neurogenesis (Fernando et al., 2011). Taken together, these findings suggest critical roles for neurotransmitter signaling in the SEZ neurogenic niche, whereby their disruption can induce long-lasting impairments to OB neurogenesis. However, it is still unclear whether neurotransmitter signaling plays a prominent role in NPC survival, or primarily in proliferation, as most studies to date have focused on mechanisms involved in proliferation, while studying survival in vivo has been challenging (Berg et al., 2013).

1.5. Postnatal NSC-niche heterogeneity and endogenous response to injury 1.5.1. Regional heterogeneity within the SEZ Heterogeneity exists amongst NSCs and their progeny within the SEZ, with regionally distinct subpopulations of NPCs biased towards specific lineages and neuronal subtypes

(reviewed in Kriegstein and Alvarez-Buylla, 2009). This parcellation occurs during embryonic development of the telencephalon via restricted expression of transcription factors, which are partially conserved in the adult SEZ (reviewed in Wilson and Rubenstein, 2000; Young et al., 2007; Delgado and Lim, 2015). Using Cre recombinase driven by transcription factor promoters specific to different regions of the developing VZ and adult SEZ, Young et al. determined the embryonic origins of various NPC populations in the adult SEZ and their postmitotic fate. These lineage tracing experiments found that Gsh2+ precursors in the lateral ganglionic eminence (LGE) predominantly gave rise to NPCs in the dorsolateral corner and lateral wall of the adult SEZ, while Emx1+ precursors in the dorsal VZ gave rise to NPCs in the dorsolateral corner and dorsal wall of the adult SEZ. Adult-born calbindin+ and DA neurons in the PGL were primarily derived from lateral Gsh2+ precursors, while calretinin+ interneurons were derived from dorsal and dorsolateral Emx1+ precursors (Young et al., 2007). Similar work by Merkle et al. confirmed these findings, and also demonstrated that this regional heterogeneity is cell-intrinsic, since regionally distinct SEZ NSCs maintain their post-mitotic fate even after being cultured and differentiated in vitro, or when heterotopically transplanted into donor mice (Merkle et al., 2007). The proliferation of NPCs in different regions of the SEZ and their relative contributions to neurogenesis are also heterogeneous. Using Cre recombinase lineage tracing and BrdU labeling, Young et al. reported that the majority of adult-born OB neurons were generated from the lateral SEZ; however, it was not determined if this was due to laterally enriched NSC or TAP proliferation (Young et al., 2007). Using the G1 phase marker Mcm2 and absence of the thymidine analog (S phase marker) 5-ethynyl-2’-deoxyuridine (EdU) following 3 days of EdU administration, relatively quiescent (Mcm2+/EdU-) NSCs were shown to be homogeneously distributed along the lateral wall, with some located in the lateral region of the dorsal wall. A similar distribution was also observed for (Mash1+) TAPs and proliferating (Ki67+) cells, which were slightly rostrally enriched (Azim et al., 2012a). Likewise, the transcription factor Gsx2 is predominantly expressed in the dorsolateral SEZ, where it is associated with NSC activation and control of lineage progression to TAPs (López-Juárez et al., 2013).

Similar to NSCs, neuroblasts and OPCs also exhibit regional heterogeneity. The transcription factor Tbr2+ labels neuroblasts fated to become glutamatergic neurons (Brill et al., 2009), which are scarce and restricted to the lateral region of the dorsal SEZ, while Dlx2+ GABAergic neuroblasts are ~40 times more prevalent and enriched in the dorsolateral SEZ (Azim et al., 2012a). Furthermore, postnatal OPC production is restricted to the dorsal SEZ due to dorsally enriched canonical Wnt3/-catenin signaling, which suppresses glycogen synthase 3 (GSK3)-mediated regulation of oligodendrogenesis (Azim et al., 2014a,b).

1.5.2. Temporal heterogeneity within the SEZ At birth, the periventricular region is much larger (~300µm thick) than in adults (<100µm) and contains many RG. By P7, RG-derived immature ependymal cells comprise ~30% of the cells in the periventricular region, and by P15, the periventricular region resembles that of the young adult where RG are completely absent, having given rise to mature ependymal cells and NSCs (Tramontin et al., 2003; Merkle et al., 2004; reviewed in Fiorelli et al., 2015; Redmond et al., 2019). Although the overall structure of the SEZ is generally conserved into adulthood, several changes in the composition of cell types, signaling pathways, and cell fate are observed throughout postnatal life (Capilla-Gonzalez et al., 2014a; Fiorelli et al., 2015; reviewed in Adams and Morshead, 2018; Ruddy and Morshead, 2018). These age-related changes significantly reduce proliferation and neurogenesis in the SEZ due to both intrinsic changes in NPCs and extrinsic niche-mediated changes, although it is controversial whether reduced neurogenic activity is due to reduced NPC pool size, mitotic activity, differentiation, cell survival, or a combination of these effects (Luo et al., 2006; Bouab et al., 2011; reviewed in Conover and Shook, 2011; Katsimpardi et al., 2014; Piccin et al., 2014; reviewed in Chaker et al., 2016). Studies pertaining to the effects of aging on neurogenesis typically compare 2-3 age groups: young adult (2 to 5-months old), middle-aged (8 to 12 months old), and old/aged (18-24 months old). Initial studies typically focused on quantification of NSCs and their progeny, with some reports of ~50-75% reductions in proliferative cells and putative NSCs of old mice relative to young adults (Tropepe et al., 1997; Enwere et al., 2004; Maslov et al., 2004; Ahlenius et al.,

2009; Capilla-Gonzalez et al., 2014). Similar reductions in ependymal cells, TAPs, neuroblasts (including the fraction of proliferating neuroblasts), and OPCs have also been reported using combinations of immunohistochemistry (IHC) and TEM (Enwere et al., 2004; Ahlenius et al., 2009; Capilla-Gonzalez et al., 2014). In concurrence with reduced numbers of proliferative cells within the aged SEZ, some studies report increased frequency of division and reduced survival amongst remaining proliferative cells in vitro and in vivo, although these studies do not differentiate between NSCs and progenitors (Stoll et al., 2011; Shook et al., 2012). Similarly, while some studies report dramatic (75%) reductions in total numbers of NSCs (Shook et al., 2012; Capilla-Gonzalez et al., 2014), many of these studies rely on markers that exclusively quantify proliferative cells or aNSCs and TAPs, such as Nestin, Mcm2, and thymidine analogs (S-phase markers) with short chase periods, ultimately precluding qNSCs. In contrast, other groups have demonstrated the loss of TAPs and neuroblasts, but report sustained NSC pools and instead attribute progenitor reductions to increased NSC cell cycle time or quiescence in middle-aged and old mice (Tropepe et al., 1997; Luo et al., 2006; Bouab et al., 2011; Giachino et al., 2014). Increased neuroblast apoptosis has also been reported, thereby implicating both cell survival and proliferation in age- associated neurogenic decline (Luo et al., 2006). In accordance with diminished TAP and neuroblast populations in the SEZ, dramatic reductions in neuroblast proliferation in the rostral RMS (~50%) and newborn neurons in the OB (~75% reduction) have also been reported in both middle-aged and old mice (Bouab et al., 2011; Shook et al., 2012). Mechanistically, reduced proliferation in the aged SEZ has been attributed to reduced expression of proliferation-related proteins such as EGFR, which is associated with reduced OB neurogenesis and significant deficits in discriminating between similar odors (Enwere et al., 2004). In 12-month-old mice, telomerase expression is reduced, which is associated with telomere shortening and increased p53 activity in neurosphere-forming cells and reduced OB neurogenesis (d’Adda di Fagagna et al., 2003; Ferrón et al., 2009). Indeed, reduced in vitro or in vivo NPC proliferation caused by telomerase deficiency was rescued in p53-null mice, suggesting intrinsic cell deficits in aged NPCs (Ferrón et al., 2009). Certain genes in the aging brain have also been proposed to control intrinsic NPC behaviour such as the transcription

28 factor FOXO3, which is expressed in NPCs and maintains quiescence (via cell cycle regulators CDKN1B/p27KIP1 and cyclin G2) to prevent NPC pool exhaustion in middle-aged mice (Webb et al., 2013). Similarly, expression of the cyclin dependent kinase (Cdk) inhibitor p16INK4a is dramatically increased in the SEZ of the aged brain, which inhibits cell cycle re-entry upon sensing DNA damage. Aged p16INK4a-deficient mice display alleviated declines in SEZ proliferation, OB neurogenesis, and in vitro NPC proliferation, while p16INK4a deficiency has no effect in young mice (Molofsky et al., 2006). However, age-related changes to NPC behaviour have also been attributed to alterations in the SEZ niche. A downregulation of fibronectin and integrins is observed, resulting in altered architecture, reduced cell density, and poor diffusion of macromolecules (Wang et al., 2011). In the aged niche, larger and more penetrative fractones extend from the vasculature and ependymal cells appear flattened with tangled cilia, resulting in cilia-devoid patches. The aged SEZ also exhibits increased numbers microglia with reduced ramifications, suggestive of activation, which contribute to reduced neurogenesis (Capilla-Gonzalez et al., 2014; Solano Fonseca et al., 2016). Indeed, it has recently been reported that NSC maintenance throughout aging is due to increased NSC quiescence that is maintained by increased inflammation and SFRP5-mediated inhibition of non-canonical Wnt activity. The NSC transcriptome changes minimally with age and NSCs behave similarly in young and old mice once activated (Giachino et al., 2014; Kalamakis et al., 2019). NSC colony formation is reduced when old SEZ tissue is cultured in vitro, although this reduction is lost when old NSCs are cultured as a pure population, suggesting that intrinsic NSC self-renewal capacity is maintained with age (Ahlenius et al., 2009). Heterochronic transplantation of SEZ-derived NPCs demonstrates that NPC migratory behaviour is entirely dependent on the age of the host niche, and that factors in conditioned media from the young niche enhance in vitro NSC colony formation and in vivo NPC proliferation in old mice (Piccin et al., 2014). Likewise, heterochronic parabiosis of 2 and 15-month-old mice caused significant increases in NPCs and proliferating cells in the SEZ of old mice relative to isochronic old controls and also rescued olfactory sensitivity, suggesting that circulating factors from young mice are sufficient to enhance functional neurogenesis in old mice. This was associated with a decline in

29 blood vessel volume in aged mice that was reversed by factors in young blood (Katsimpardi et al., 2014). The LV choroid plexus secretome contains factors that promote NSC and TAP proliferation, which changes with age. Silva-Vargas et al. demonstrated that young aNSCs cultured in choroid plexus conditioned media (LVCPsec) from old mice reduced in vitro clone formation by ~50% relative to young aNSCs cultured in LVCPsec from young mice. Likewise, infusion of LVCPsec from old mice into the LVs of young mice significantly reduced NSC proliferation while LVCPsec from young mice increased NSC proliferation in old mice (Silva- Vargas et al., 2016). Specific factors from young LVCPsec that promoted clone formation include IGF-1, IGF-2, and BMP5, while IL-1 and NT3 in old LVCPsec reduced clone formation (Lehtinen et al., 2011; Silva-Vargas et al., 2016). Hence, a multitude of age-related alterations to the SEZ niche underlie the reduced neurogenic activity observed in middle-aged and old mice. Although various intrinsic mechanisms have been proposed, several studies have demonstrated that old NSCs behave like young NSCs when removed from their niche or supplemented by factors from young mice.

1.5.3. Changes to NSC and niche dynamics following injury CNS injuries and diseases can induce robust responses from NPCs and the SEZ niche. Not surprisingly, these responses vary depending on age and type of injury/disease. The ability for NPCs to respond to CNS injuries has led to significant interest in harnessing these cells for regenerative therapies and has highlighted the importance of protecting these cells from environmental factors or injuries that may affect their neurogenic capacity throughout the lifespan. Stroke/ischemia has been a prominent rodent injury model to study the endogenous response of SEZ NPCs to injury, as well as therapies that may enhance their response to promote functional recovery. Following stroke, NSCs transiently proliferate and generate neuroblasts, both of which migrate through the striatum towards striatal and/or cortical lesions where they contribute to neurogenesis and glial scar formation (Arvidsson et al., 2002; Zhang et al., 2004a; Faiz et al., 2015; reviewed in Chaker et al., 2016).

Similar to neuroblast migration along the RMS, injury-induced migration through the striatum relies on chemoattraction and can occur via individual or chain migration at similar rates as RMS migration and in close association with astrocytes and vasculature (Yamashita et al., 2006; Kimura et al., 2008; Zhang et al., 2009). Newly generated GABAergic and cholinergic neurons following stroke have been shown to receive synaptic inputs and fire action potentials, suggesting functional integration (Hou et al., 2008). However, this endogenous response is insufficient to restore stroke-induced loss of neurons since >80% of stroke-generated neurons are estimated to die within 2 weeks after differentiation, ultimately replacing <1% of lost neurons (Arvidsson et al., 2002). In order to improve cellular recovery and promote functional recovery following such injuries, an understanding of the mechanisms underlying injury-induced neurogenesis is required. Injury-induced NPC proliferation is necessary for repair, as Ara-C treatment inhibits SEZ-derived neurogenesis in the striatum following stroke (Arvidsson et al., 2002). Altering the symmetry of NSC division may be required, as suggested by the endogenous transient shift to symmetric division also observed after stroke (Zhang et al., 2004b). Interestingly, the recruitment of qNSCs to the aNSC pool has also been suggested, since the non-canonical Wnt signaling target Cdc42 that maintains NSC quiescence is downregulated following demyelination injury, thereby initiating NSC lineage progression and tissue repair (Chavali et al., 2018). Expression of transcription factor Gsx2, which is normally restricted to the dorsolateral corner of the SEZ, is upregulated and ectopically expressed throughout the SEZ following either hypoxia-ischemia or excitotoxicity-induced lesions, which is necessary to increase TAP and neuroblast production (López-Juárez et al., 2013). Therefore, the activation of numerous pathways involved in the proliferation of NSCs and their progeny contribute to injury-induced neurogenesis. In addition to the cellular responses of NSCs and their progeny, it is crucial to understand the injury-induced alterations to the niche, including secondary damage that may impede recovery, since these effects may reveal other potential targets for therapeutic intervention. For example, cortical stroke in young adult, but not old mice induced an expansion of the SEZ aNSC pool size associated with upregulation of Wnt signaling in young

31 mice only, suggestive of a niche-mediated effect since both young and old NSCs behave similarly in the absence of their niche (Piccin et al., 2014). Growth factor signaling is also implicated in stroke-induced neurogenesis, evidenced by upregulated IGF-1 expression in lesion sites, which is necessary for the proliferative response of NPCs (Yan et al., 2006). Accordingly, intraventricular infusion of EGF and erythropoietin enhanced the SEZ neurogenic response and promoted cellular and functional motor recovery, even when treatment was delayed 7 days post-stroke (Kolb et al., 2007). Growth factor signaling may also enhance injury-induced neuroblast migration, since transient striatal angiogenesis that guides neuroblasts to lesion sites is observed post-stroke, and is enhanced by IGF-1 overexpression leading to motor functional recovery (Thored et al., 2007; Zhu et al., 2008). In addition to stroke, several brain injuries modify microglial activation leading to distinct NPC responses depending on the type and location of injury, as well as the duration or extent of inflammation (Ekdahl et al., 2009). Bacterial lipopolysaccharide (LPS) has commonly been administered to experimentally elicit acute and intense inflammatory microglial activation, which is cytotoxic to NPCs and reduces both neurogenesis and oligodendrogenesis (Butovsky et al., 2006; Cacci et al., 2008). In line with the detrimental effects of acute microglial activation, administration of the anti-inflammatory drug indomethacin improved neuroblast survival in lesion sites in the first month following stroke (Hoehn et al., 2005). However, SEZ microglia have previously been identified as unique from microglia of other brain regions, predominantly acting in an anti-inflammatory supportive role (Ribeiro Xavier et al., 2015). Accordingly, while peri-infarct microglia exhibited an inflammatory phenotype, SEZ microglia exhibited an anti-inflammatory phenotype and expressed IGF-1 for 4 months following stroke, likely reducing apoptosis while enhancing NSC proliferation and neuronal differentiation (Ekdahl et al., 2009; Thored et al., 2009; Deierborg et al., 2010). In contrast to stroke-induced neurogenesis and anti-inflammatory microglial activation in the SEZ, other injuries have been shown to promote inflammatory microglial activation and inhibit neurogenesis (Su et al., 2014). In humans, Parkinson’s Disease (PD) depletes EGFR+ cells in the SEZ and the 6-OHDA rodent model of PD similarly exhibits reduced EGF production, TAP proliferation, and OB neurogenesis, which is rescued by L-DOPA or D2L agonists (Baker et al.,

2004; O’Keeffe et al., 2009; Winner et al., 2009). Interestingly, the MPTP model of PD causes inflammatory microglial activation, resulting in reactive oxygen and nitrogen species (ROS and RNS) production, which activates GSK3-mediated -catenin degradation. Ultimately, this deficit in Wnt/-catenin signaling causes reduced NPC proliferation and neuroblast formation, but is rescued by GSK3 or HCT1026 administration, a derivative of the nitric oxide (NO)-donating nonsteroidal anti-inflammatory drug (NSAID) flurbiprofen (L’Episcopo et al., 2010; L’Episcopo et al., 2012). Taken together, several signaling pathways including neurotransmitters, microglial activation, and growth factors contribute to both direct and niche-mediated neurogenic responses to certain injuries, highlighting the need for multifaceted or pleiotropic therapeutic approaches.

1.6. Assaying for NSCs 1.6.1. Assaying for NSCs in vitro Due to a lack of definitive markers and unique morphology of adult NSCs, researchers have heavily relied on the use of functional criteria for their post-hoc identification, namely self- renewal and multipotentiality (Morshead et al., 1994). The predominant technique for the in vitro identification and analysis of NSCs is the neurosphere assay, which was established by Reynolds and Weiss in 1992 and continues to be the gold standard for in vitro NSC assays (Reynolds and Weiss, 1992; Walker and Kempermann, 2014). The neurosphere assay consists of the dissection of primary tissue, dissociation into a single cell suspension, and subsequent culturing in the presence of EGF, FGF2, and heparin (Coles-Takabe et al., 2008; Azari et al., 2010; Walker and Kempermann, 2014). These conditions support the survival and proliferation of NSCs and their progeny, resulting in the formation of free-floating spherical cell clusters termed neurospheres, typically comprised of hundreds of cells. Given the single cell origin of neurospheres, the size of the NSC pool is reflected in the number of neurospheres formed when cells are plated at clonal density (Coles-Takabe et al., 2008). The majority of the cells comprising a neurosphere are neural progenitors and to a lesser extent, differentiated cells, which thereby allows for the estimation of progenitor pool size via

33 the quantification of neurosphere size (Walker and Kempermann, 2014; reviewed in Adams and Morshead, 2018). Neurospheres can be passaged several times in order to assess self-renewal capacity (Reynolds and Weiss, 1996), and can subsequently be differentiated on adherent substrates via the removal of EGF and addition of fetal bovine serum (FBS), which supports the differentiation of NPCs into neurons, oligodendrocytes, and astrocytes, thereby exhibiting the multipotentiality of single neurospheres (Ahmed et al., 1995; Reynolds and Weiss, 1996). Using these outcome measures, the behaviour of NSCs and their progeny can be assessed in response to extrinsic cues via the addition of such factors to the neurosphere growth or differentiation media such as in conditioned media and co-culture experiments, or by growing neurospheres from injured or drug-treated animals (reviewed in Adams and Morshead, 2018). When using the neurosphere assay, it must be recognized that niche factors may persist in primary cultures. In order to explicitly characterize cell autonomous (intrinsic) NSC properties, passaged neurospheres can instead be assessed. Another important consideration when using the neurospheres assay to quantify NSC pool size is the fact that it is a retroactive assay, reliant on the expression of EGFR by NSCs in order to respond to culture conditions and proliferate to form a sphere (reviewed in Pastrana et al., 2011). Given more recent work examining the expression profiles and behaviour of neurosphere forming cells, one of the hallmarks of an aNSC is their ability to form neurospheres when cultured for 7-10 days. Conversely, qNSCs require 10-12 days in culture before acquiring an activated state conducive to proliferation and neurosphere formation due to their minimal/lack of EGFR expression when plated from primary dissections of the SEZ (Codega et al., 2014). NSCs can also be assayed in vitro using adherent monolayer cultures, which may improve the homogeneity of cells due to reduced cell-cell communication and cell density- related issues that exist within neurospheres (Walker and Kempermann, 2014). Adherent NSC cultures may also enhance the purity of NSCs and undifferentiated progenitors relative to neurosphere cultures by reducing spontaneous differentiation, while maintaining their multipotentiality (Conti et al., 2005; Pollard et al., 2006; Conti and Cattaneo, 2010). However, a major limitation of adherent monolayer cultures relative to the neurosphere assay is the

34 inability to quantify and isolate individual single cell-derived clones, making the neurosphere assay more favorable for quantifying NSC pool size and progenitor pool size (Walker and Kempermann, 2014; reviewed in Adams and Morshead, 2018).

1.7. Cranial Irradiation Since certain forms of brain injury, including stroke, rely on recruiting endogenous NPCs for cell replacement and functional recovery, it is vital to protect postnatal NPCs and their niche from injuries and diseases that may affect their survival, proliferation, differentiation, and migration. One form of injury that impairs these behaviours is cranial irradiation (IR), which is a prominent adjuvant therapy in the treatment of childhood primary brain tumours and acute lymphoblastic leukemia (Packer et al., 1999; reviewed in Duffner, 2004; Redmond et al., 2013). As long-term survival rates have increased with advancements in treatment, several late cognitive impairments have been identified amongst surviving patients. Gradual reductions in IQ and academic performance arise over several years (Mabbott et al., 2005; reviewed in Palmer et al., 2007), and are thought to be caused by earlier deficits to processing speed, working memory, broad attention, and executive function (Schatz et al., 2000; Palmer et al., 2013). Interestingly, decline in intellectual function was more rapid and severe in patients who received cranial IR at younger ages (early childhood) (Chin and Maruyama, 1984; Spiegler et al., 2004; Tonning Olsson et al., 2014). In rodent models of cranial IR, cognitive deficits are associated with long lasting reductions in neurogenesis in both the SEZ and DG (Hellström et al., 2009), which is not surprising given the relatively high levels of mitotic activity in NPCs which make them particularly sensitive to IR-mediated ablation (Shinohara et al., 1997). Since NPCs are required for lifelong functional neurogenesis and endogenous repair following certain brain injuries, it is crucial to identify the mechanisms underlying IR-induced cellular and behavioural deficits that may serve as targets for therapeutic intervention.

1.7.1. IR-induced apoptosis and senescence It is widely accepted that IR induces cell death through DNA damage and chromosomal dysfunction leading to apoptosis. Although many forms of DNA damage are repairable by cellular repair mechanisms, DNA double stranded breaks (DSBs) are proposed to be more difficult to repair due to the lack of an intact template strand to guide DNA repair machinery, resulting in chromosomal aberrations, mutagenesis, and cell death (reviewed in Iliakis, 1991; Goodhead, 1994; Little, 2000; Vignard et al., 2013). Furthermore, IR-induced NPC death occurs through caspase 3 and p53-mediated apoptosis in the young adult rodent SEZ at doses as low as 2 Gy, with apoptosis peaking 4-6 hours post-IR and subsiding by 48 hours (Bellinzona et al., 1996; Shinohara et al., 1997; Chow et al., 2000). However, deficits to neurogenesis cannot be attributed solely to acute cell loss caused by apoptosis, since deficits to normal NPC behaviour persist several weeks to months after apoptosis subsides (Fukuda et al., 2005). In addition to apoptosis, cranial IR induces transcript and protein expression profiles suggestive of cellular senescence (Cheng et al., 2017). In the DG, various senescence-associated proteins are upregulated 2 months post-IR, including p16INK4a, p19ARF, p53, p21, phosphorylated p38 (pp38), 4-hydroxy-2-neonal, IL-6, and -H2AX (Cheng et al., 2017). Similar to the upregulation of p16INK4a in NPCs of the aging SEZ, the upregulation of these factors is associated with reduced proliferative capacity and terminal differentiation into S100B+ astrocytes (Molofsky et al., 2006; Zou et al., 2012).

1.7.2. IR-induced NPC dysfunction and deficits to neurogenesis and oligodendrogenesis In young adult rodents, depletion of proliferating cells is observed in the SEZ 1 day following cranial IR with doses as low as 2 Gy. Recovery occurs over 7-14 days in a dose- dependent manner and subsequently decreases again, resulting in >50% reductions after 6 months for mice receiving 7.5 Gy IR or greater, suggesting deficits to long-term NPC pool maintenance (Tada et al., 1999). Following 25 Gy cranial IR in adults, or 15 Gy in neonates, proliferating cells in the SEZ were immediately depleted and remained depleted 3-15 months

36 later, suggesting that high doses of cranial IR cause a permanent reduction in the NPC pool (Panagiotakos et al., 2007; McGinn et al., 2008). Interestingly, nestin expression is significantly upregulated in the adult SEZ 1-4 weeks following 20 Gy cranial IR, although proliferation remains absent, suggesting that higher doses of IR may recruit nestin- qNSCs to the nestin+ aNSC pool to replace lost cells, but proliferation remains impeded (Fig. 1.4) (Shi et al., 2002). Using a lower dose of 4 Gy cranial IR in young adult mice, Daynac et al. showed that p53-dependent apoptosis still occurs in the SEZ during the first 24 hours post-IR and that LeX+EGFR- qNSCs become activated 2 days later, as visualized using transgenic Fluorescence Ubiquitination Cell Cycle Indicator (FUCCI) green mice, in which cells fluoresce green during S and G2 phases (Sakaue-Sawano et al., 2008; Daynac et al., 2013).

Activation and proliferation of qNSCs was linked to reduced NSC GABAAR signaling as a result of IR-induced neuroblast depletion, a major source of GABA in the SEZ (Liu et al., 2005; Fernando et al., 2011). Short-term administration of the GABAAR agonist muscimol prevented the repopulation of aNSCs and TAPs 96 hours post-IR, despite spontaneous recovery of non-treated mice at this time, confirming the role of GABA signaling in NSC response to IR (Daynac et al., 2013). The age at which cranial IR is administered also affects neurogenic outcomes. Following 6 Gy cranial IR at P9 or P23, a significantly increased number of apoptotic (cleaved caspase 3+/p53+) cells was observed in the P9 SEZ compared to the P23 SEZ, which correlates with increased Nestin+ NPCs and DCX expression under baseline conditions at P9 compared to P23 (Fukuda et al., 2005). Although the deficits were greater at P9 than at P23, Nestin+ NPCs in the SEZ were significantly reduced in both age groups 6 hours post-IR, followed by a partial recovery over 7 days, and subsequent depletion 10 weeks later. Taken together, this suggests a transient activation of NSCs to compensate for lost cells, which is insufficient to restore the NPC pool (Fukuda et al., 2005). Given the deficits to NPC proliferation and differentiation in the SEZ, it is not surprising that a near-complete reduction in newborn neurons is observed in the OB 3 months after 15 Gy focal IR to the SEZ. Concomitant with the IR-induced deficits to OB neurogenesis, irradiated mice exhibited a reduced odor-cued, but not audio-cued fear response in a fear conditioning

37 task at both 6 and 26 weeks post-IR (Valley et al., 2009). Similarly, 15 Gy focal IR (over 3 fractions) to the SEZ causes significant SEZ and OB neuroblast reduction 7 months post-IR, as well as reduced newborn OB neuron production resulting in deficits to LTOM (Lazarini et al., 2009). To confirm whether these deficits were caused by dysfunction to NSCs or their progeny, Achanta et al. used TEM and IHC to identify the specific cell types involved. TAPs and neuroblasts remained almost completely depleted 1 month after 10 Gy focal IR to the SEZ, despite NSCs being present at ~50% of control levels, which suggests that NSCs retain regenerative capacity and that downstream NPC behaviours are perturbed (Achanta et al., 2012). When the anterior SEZ and RMS were focally irradiated, neuroblasts were spared in the posterior SEZ, but would not migrate rostrally through the irradiated tissue, suggesting that IR also disrupts the microenvironment and prevents neuroblast migration to the OB (Achanta et al., 2012). In addition to neurogenic deficits, OPCs in the corpus callosum are immediately and permanently depleted following 25 Gy cranial IR, which has been shown via magnetic resonance imaging (MRI) and ultrastructural analyses to reduce white matter volume and cause progressive demyelination 9-12 months post-IR (Panagiotakos et al., 2007). Similarly, when 8 Gy cranial IR is delivered at P8 in rats, an important time for OPC production, myelin basic protein (MBP) expression is significantly reduced in both the striatum and corpus callosum 1 week later during the window of peak myelination (Fukuda et al., 2004). This has important implications for injury-induced NPC activation. For example, when a moderate dose of 10 Gy is focally applied to the SEZ via computed tomography (CT)-guided IR, a demyelinating injury to the striatum is able to elicit a response from the SEZ 1 month post-IR, but to a lesser extent than in non-irradiated mice (Capilla-Gonzalez et al., 2014b). Therefore, cranial IR not only causes deficits to baseline neurogenesis, oligodendrogenesis, and olfactory behaviour, but it also impedes the ability of the SEZ to respond to injuries.

Figure 1.4. Cranial IR-induced cellular effects on NSC activation and signaling (Chaker et al., 2016).

1.7.3. Niche-mediated effects of cranial IR While IR directly kills NPCs and disrupts the function of surviving NPCs, IR also induces secondary damage to the neurogenic niches that results in indirect suppression of neurogenesis. This is suggested by regional differences in spontaneous recovery between the DG and SEZ. While the neonatal rodent DG and SEZ both show ~90% reductions in proliferating cells 1 day after 6 Gy cranial IR, there is partial recovery of proliferating cells (~50%) and newborn neurons (~40% of controls) 9 weeks later in the SEZ and OB, but not the DG (~10% & <5% recovery, respectively). Interestingly, GFAP+nestin+ putative NSCs surpassed control levels in the SEZ 9 weeks post-IR but remained depleted (~15% recovery) in the DG (Hellström et al., 2009), suggesting that the NSC microenvironment plays a significant role in mediating the recovery of NSCs and their progeny following IR. Based on reports that have suggested a lack of bona fide NSCs in the adult DG (Seaberg and van der Kooy, 2002; Clarke and van der Kooy, 2011), it can be argued that regional differences in recovery are caused by the activation of surviving qNSCs in the SEZ (Daynac et al.,

2013), which may be absent in the DG. However, when healthy NPCs are grafted into the irradiated DG 1 month post-IR, an ~80% reduction in newborn neurons is observed compared to grafted NPCs into the non-irradiated DG (Monje et al., 2002). Furthermore, when NPCs are grafted into the 15 Gy-irradiated or naïve SEZ, their survival and neuroblast formation are comparable, although proliferation is only observed in the non-irradiated host (Pineda et al., 2013). These findings suggest that persistent IR-induced deficits to proliferation and differentiation are mediated by the niche and not simply due to intrinsic NPC deficits. Indeed, there is strong evidence suggesting that cranial IR induces inflammation, which may be a primary mediator of IR-induced deficits to neurogenesis (Hong et al., 1995; reviewed in Wong and Van der Kogel, 2004). Following 15 Gy cranial IR, Pineda et al. observed an increase in transforming growth factor- (TGF-) production by endothelial cells in the SEZ that specifically activated TGF-/Smad3 signaling in NSCs and TAPs (Pineda et al., 2013). Interestingly, TGF- has been suggested to be anti-inflammatory in certain settings (Ledeboer et al., 2000), although TGF-/Smad signaling has also been shown to have inflammatory and apoptotic roles in other forms of brain injury (Patel et al., 2017). Administration of a TGF- neutralizing antibody or TGF- receptor-I (TRI) inhibitor 1 month post-IR reduced apoptosis in the SEZ and RMS immediately following treatment, and increased neuroblast production and arrival in the OB 3 weeks following treatment. Furthermore, co-culture of irradiated brain endothelial cells (BECs) with neurosphere-derived Mash1+ TAPs caused apoptosis of TAPs, which was blocked in the presence of an anti-TGF- neutralizing antibody, confirming that the irradiated niche contributes to cell death (Pineda et al., 2013). Although the irradiated niche appears to reduce TAP survival, proliferation, and differentiation, the niche’s effects on NSC behaviours are unclear. While the majority of studies pertaining to IR-induced neuroinflammation focus on the DG as opposed to the SEZ, several parallels are found (Hellström et al., 2009; Kalm et al., 2009). Seminal studies reported increases in activated pro-inflammatory (CD68+) microglia and BrdU+CD11b+NG2+ cells considered to be proliferating macrophages recruited from the periphery in the DG 2 months post-IR (Monje et al., 2002; Mizumatsu et al., 2003; Monje et al., 2003). Furthermore, treatment with the nonsteroidal anti-inflammatory drug indomethacin

40 starting 2 days prior to 10 Gy cranial IR until 2 months post-IR attenuated this inflammatory response and promoted partial (~25%) recovery of newborn DG neurons (Monje et al., 2003). However, while an increase in CD68+ microglia in both neurogenic niches is consistently observed in the first 24 hours post-IR (Hellström et al., 2009), others report that this response subsides as early as 1 week later and that macrophage infiltration does not occur (Han et al., 2016). Oddly, IR-induced acute upregulation of the proinflammatory cytokines IL-1 and IL-1, as well as chemokine ligand 2 (CCL2/MCP-1) and its receptor CCR2 are consistently reported in both neurogenic niches of juvenile and adult rodents, although conflicting studies suggest they either return to baseline expression levels within 1 week (Kalm et al., 2009), or remain elevated 1 month later (Han et al., 2016). Although CCL2 is not linked to microglial activation but is instead considered to promote proliferation of activated microglia and recruit peripheral macrophages following brain injury, these aforementioned studies reveal conflicting hypotheses of IR-induced local vs. hematogenous inflammation, and between acute vs. chronic inflammation (Schilling et al., 2009; Hinojosa et al., 2011). Despite these dichotomies, it is irrefutable that cranial IR induces some form of inflammation that perturbs neurogenesis. To address this issue, Lee et al. demonstrated that 10 Gy cranial IR in young adult mice transiently upregulated CCL2 during the first 12 hours post-IR, resulting in no recruitment of peripheral macrophages, but increased BrdU+Iba1+ and CD68+ microglia in the DG 1 month post-IR. This was rescued in CCL2-null mice, suggesting that IR-induced acute upregulation of CCL2 specifically promotes proliferation of activated microglia that persist for at least 1 month post-IR (Lee et al., 2013). Accordingly, administration of the pro-inflammatory cytokine inhibitor MW-151 immediately following IR did not prevent reduced proliferation in the DG, but rescued neuroblasts 2 months post-IR, which led to functional recovery in HPC-dependent memory (Jenrow et al., 2013). Similarly, 10 Gy cranial IR impaired Morris water maze performance 2 months post-IR, which was prevented in CCR2-null mice, suggesting that this inflammatory pathway significantly contributes to impaired functional neurogenesis (Belarbi et al., 2013).

1.8. Endogenous NPC activation using metformin While stem cell transplantation has been a popular approach to replace lost or dying tissues and cells in virtually every organ system, transplantation is associated with several caveats, including the challenge of identifying appropriate stem cell sources, the potential requirement of immunosuppression to avoid host-graft rejection, and the reliance on engineered scaffolds or delivery vehicles to ensure cell survival and integration following grafting (reviewed in Mahla, 2016). Thus, the field of regenerative medicine has invested significant effort into endogenous stem cell activation strategies, whereby resident stem and progenitor cell populations are recruited to respond to injuries, or their endogenous responses are enhanced (reviewed in Miller and Kaplan, 2012). Since the discovery of NSCs in the postnatal mammalian brain, enhancing neurogenesis has become an attractive strategy for neurorepair following brain injury, in part due to the observation that NPCs are endogenously recruited from the SEZ following a variety of injuries (Arvidsson et al., 2002; Zhang et al., 2004a; Yamshita et al., 2006; L’Episcopo et al., 2012; Faiz et al., 2015). Experimental strategies to enhance this endogenous response often include growth factor or hormone administration such as EGF or erythropoietin (Kolb et al., 2007; Thored et al., 2007; Zhu et al., 2008), and more recently, researchers have turned to the highly prescribed and well-tolerated drug metformin (reviewed in Potts and Lim, 2012).

1.8.1. Pharmacology & toxicology Metformin (1,1-dimethylbiguanide) is an oral antihyperglycaemic (antidiabetic) drug that reduces gluconeogenesis and improves insulin sensitivity in the liver without causing hypoglycemia or hyperinsulinemia (reviewed in Dunn and Peters, 1995; Hundal et al., 2000; reviewed in Gong et al., 2012; reviewed in Pernicova and Korbonits, 2014). Metformin is a polar cationic molecule, requiring membrane-bound transporters such as organic cation transporters (OCTs) or solute carriers (SLCs) for cell entry and exit (reviewed in Graham et al., 2011; Pernicova and Korbonits, 2014). Metformin is largely absorbed in the small intestine, does not bind plasma proteins, is not metabolized by the liver, and is excreted in the urine unchanged (Dunn and Peters, 1995; Graham et al., 2011; Gong et al., 2012). Following oral administration,

OCT1 (SLC22A1) and OCT3 (SLC22A3) mediate intestinal transport of metformin, as well as hepatocyte entry. OCT1 and OCT2 (SLC22A2) mediate renal epithelial cell uptake, while multidrug and toxin extrusion 1 (MATE1/SLC47A1) and MATE2-K (SLC47A2) mediate renal excretion of metformin (Gong et al., 2012). Plasma elimination half-life is dependent on route of administration but not dose, and is consistently reported as shorter when administered intravenously (IV) (Dunn and Peters, 1995; Graham et al., 2011). Typical doses in human patients range from 500-2,550 mg/day. The plasma elimination half-life is 1.5-4.5 hours following IV administration with >80% excretion in the urine only, or 2.0-8.7 hours following oral administration with similar excretion occurring through both urine and feces (Tucker et al., 1981; reviewed in Dunn and Peters, 1995; reviewed in Scheen, 1996). In rodents, the no observable adverse effect level (NOAEL) has been reported to be 200 mg/kg/day following 13 weeks of oral administration, with doses ≥600 mg/kg/day resulting in weight loss, and doses ≥900 mg/kg/day resulting in mortality (Quaile et al., 2010). Similarly, plasma elimination half-life in rodents is 2-2.5 hours or 3-7 hours following IV or oral administration, respectively (Choi et al., 2006). Furthermore, metformin is not detectable or nearly completely absent in plasma 24 hours following administration in both rodents and humans (Wilcock and Bailey, 1994; Robert et al., 2003). Once inside the cell, metformin accumulates in mitochondria and transiently inhibits the mitochondrial respiratory chain complex I, resulting in increased cellular concentration of adenosine monophosphate (AMP), which binds and inhibits adenylate cyclase (AC) (El-Mir et al., 2000; Owen et al., 2000; reviewed in Viollet et al., 2012). In hepatocytes, this AMP accumulation inhibits AC-mediated conversion of ATP to cyclic AMP (cAMP), which reduces protein kinase A (PKA) activity and its downstream effects on PFK/FBPase 1-mediated gluconeogenesis, as well as CREB signaling involved in gluconeogenic gene expression (reviewed in Pernicova and Korbonits, 2014; Saisho, 2015). Metformin-mediated AMP upregulation also causes activation of AMP-activated protein kinase (AMPK), which promotes the phosphorylation of the histone acetyltransferase CREB- binding protein (CBP) at serine (S) 436 by atypical protein kinase C (aPKC), thereby causing dissociation of the CREB-CBP-TORC2 transcription complex necessary for gluconeogenic gene

43 expression (Zhou et al., 2001; Zhou et al., 2004; He et al., 2009; reviewd in Viollet et al., 2012). In addition to being involved in lipid and glucose metabolism, AMPK also inhibits the inflammatory nuclear factor κB (NF-κB)-IL-6 pathway and mammalian target of rapamycin (mTOR) signaling in several cell types, which are thought to contribute to metformin’s reported lifespan improvement and anti-tumour properties (reviewed in Giovannucci et al., 2010; Gong et al., 2012; Martin-Montalvo et al., 2013; Pernicova and Korbonits, 2014).

1.8.2. Metformin in the brain Following oral administration, metformin rapidly crosses the BBB and is found in the mouse brain within 30 minutes, reaching peak concentration after 4-6 hours (Wilcock and Bailey, 1994). Following either acute (single dose of 150 mg/kg) or chronic (150mg/kg, twice daily for 3 weeks) administration, metformin is most concentrated within the CSF (~37-44µM) and is also found in the OB (~10-11µM), HPC (~5-7µM), and striatum (~2µM). Interestingly, when LPS is co-administered with metformin to induce an inflammatory response, striatal metformin concentration increases 3 to 5-fold (Łabuzek et al., 2010b). In rodents, OCTN1 (SLC22A4) is the predominant cation transporter expressed in Nestin+ NPCs and OCTN2 (SLC22A5) is expressed in astrocytes, confirming that metformin can directly exert its effects within NPCs and glial cells, although it is unclear whether Nestin- qNSCs also express these cation transporters (Inazu et al., 2006; Ishimoto et al., 2014). Similar to hepatocytes, metformin stimulates AMPK activity in NPCs, which activates the aPKC-CBP pathway (Wang et al., 2010; Potts and Lim, 2012; Wang et al., 2012; Fatt et al., 2015). Within several cell types, metformin also enhances the activity of various proteins involved in NPC survival and proliferation, including FOXO3 and TAp73, which are both expressed in NPCs (Renault et al., 2009; Hou et al., 2010; Sato et al., 2012; Fatt et al., 2015; Kim et al., 2018). Furthermore, metformin exhibits anti-inflammatory and angiogenic effects in microglial and vascular endothelial cells, both of which are abundant within the neurogenic niches of the brain, suggesting that metformin may be neuroprotective following cranial IR through multiple direct and indirect effects on NPCs (Hattori et al., 2006; Łabuzek et al., 2010a; Jin et al., 2014; Liu et al., 2014).

1.8.3. Metformin’s pro-neurogenic effects via direct action on NPCs Investigation into metformin’s effects on neurogenesis were spurred by a study in which Wang et al. demonstrated that CBP-mediated histone acetylation of neural gene promoters supports neuronal and glial cell differentiation during development in the embryonic mouse cortex and is dependent on CBP S436 phosphorylation by aPKC (Wang et al., 2010). Wang et al. later demonstrated that metformin activates the aPKC-CBP pathway in embryonic mouse NPCs and human embryonic stem cell-derived forebrain NPCs (Fig. 1.5). In this study, the presence of 500µM metformin increased production of III-tubulin+ neurons at the expense of Pax6+ and Sox2+ embryonic precursors in vitro in an aPKC-CBP-dependent manner (Wang et al., 2012). Furthermore, metformin exposure also increased the production of astrocytes and oligodendrocytes, suggesting that metformin activates the aPKC-CBP pathway to promote general differentiation of NPCs (Wang et al., 2012; Dadwal et al., 2015). Similar to metformin, the AMPK activator 5-aminoimidazole-4-carboxamide ribonucleotide (AICAR) increases neuronal differentiation from adult SEZ-derived neurospheres in an aPKC-CBP dependent manner, suggesting that metformin’s effects on NPC differentiation are indeed mediated by the AMPK-aPKC-CBP pathway (Fatt et al., 2015). In addition to promoting the differentiation of NPCs in vitro, multiple studies have reported that metformin enhances in vitro and in vivo NPC proliferation and self-renewal. Using the neurosphere assay, 4-7 days of in vivo metformin treatment prior to dissection, or 1 week of in vitro metformin exposure increases neonatal and adult SEZ-derived neurosphere number and size (Dadwal et al., 2015; Fatt et al., 2015). In vivo administration of metformin (200mg/kg/day) for 7 days increases SEZ and DG cell proliferation in adult mice, while BrdU injections followed by 21 days of metformin treatment result in increased newborn neurons in the OB. Metformin treatment also increases the number of DCX+ neuroblasts and newborn neurons in the DG in a CBP-dependent manner (Wang et al., 2012; Fatt et al., 2015). Importantly, 5 weeks of metformin treatment resulted in increased DG neurogenesis that coincided with improved spatial memory flexibility (improved performance in a Morris water maze reversal probe). When the antimitotic drug temozolomide (TMZ) was co-administered

45 with metformin, the metformin-induced DG cell proliferation and enhanced spatial memory were lost, suggesting a causal link between increased neurogenesis and spatial memory (Wang et al., 2012). To assess whether metformin could enhance neurogenesis to promote functional recovery following injury, Dadwal et al. administered metformin for 1 week following a hypoxic- ischemic (H/I) insult to P8 mice, rescuing motor deficits in the cylinder test 2 weeks post-H/I in an aPKC-CBP-dependent manner. Using lineage tracking of nestin+ cells and their progeny, metformin-mediated functional recovery was associated with increased SEZ-derived neurogenesis in the striatum and oligodendrogenesis in the striatum, motor cortex, and corpus callosum 2 weeks post-H/I (Dadwal et al., 2015). Together, these results suggest that metformin may be an effective therapeutic agent by activating endogenous NPCs to promote neurogenesis and oligodendrogenesis following brain injury. While metformin has been shown to promote both NPC proliferation and differentiation, the mechanisms underlying these effects differ. Fatt et al. demonstrated that the presence of the PKC inhibitor chelerythrine or mutation of CBP at its aPKC phosphorylation site did not prevent the metformin-mediated increase in neurosphere formation, although CBP mutation prevented the metformin mediated increase in neuronal differentiation, suggesting that the aPKC-CBP pathway is required for metformin’s effects on NPC differentiation, but not proliferation and self-renewal (Fatt et al., 2015). Previous studies have shown that tumor suppressor TAp73-deficient mice exhibit ~2- fold reductions in adult OB neurogenesis and give rise to significantly fewer SEZ-derived neurospheres, which become depleted upon passaging, suggesting that TAp73 is required for NSC self-renewal and adult neurogenesis (Fujitani et al., 2010). Interestingly, metformin has been shown to increase TAp73 expression by inhibiting mTOR in cancer cells (Rosenbluth et al., 2008). Similarly, quantitative reverse transcription polymerase chain reaction (qRT-PCR) showed that metformin increased TAp73 expression in adult SEZ-derived neurospheres, which coincides with the finding that metformin enhances cell proliferation, as well as primary and secondary neurosphere formation from the SEZ of WT, but not TAp73-deficient mice (Fatt et al., 2015). Therefore, metformin increases NPC proliferation and differentiation to enhance

46 postnatal neurogenesis through multiple pathways and mobilizes NPCs in the injured brain to promote functional recovery, making it a strong candidate for further investigation as a therapeutic agent for neurorepair.

Figure 1.5. Schematic of metformin-mediated aPKC-CBP pathway in a (A) hepatocyte and (B) embryonic neural precursor cell (Potts and Lim, 2012).

1.8.4. Metformin’s protective effects in the neurogenic niche While the stimulation of NPCs to proliferative and differentiate may enhance neurogenesis, injuries or interventions that disrupt the neurogenic niche such as cranial IR have made it apparent that directly targeting NPCs may be insufficient to maintain or recover normal levels of neurogenesis. However, as an AMPK activator, metformin confers several therapeutically relevant effects on multiple cell types within the neurogenic niches. One potential mechanism by which this occurs is by regulating the activity of FOXO3, a transcription factor that promotes oxidative stress resistance, which is phosphorylated and activated by AMPK in human cells (Greer et al., 2007). Metformin activates FOXO3 via AMPK in human vascular endothelial cells to reduce intracellular ROS levels induced by fatty acids and in glioma- initiating cells to inhibit glioblastoma formation by inducing their differentiation to nontumorigenic cells (Hou et al., 2010; Sato et al., 2012). Interestingly, FOXO3 is expressed in Sox2+ NPCs in the SEZ and in nestin+ cells from SEZ-derived neurospheres, where it is required for proper NSC pool maintenance and self-renewal throughout life, since FOXO3-deficient mice give rise to reduced SEZ-derived primary and secondary neurospheres (Renault et al., 2009).

Inflammatory microglia, which are induced by cranial IR, produce ROS and RNS that impair neurogenesis following injuries to the SEZ, suggesting that reducing oxidative stress may enhance recovery of NPCs (L’Episcopo et al., 2012). In several cell types, metformin is proposed to have anti-inflammatory effects through AMPK-dependent and independent inhibition of NF-κB, as well as anti-inflammatory (M2) microglial activation (Łabuzek et al., 2010a; reviewed in Saisho, 2015). In vitro exposure to 2mM metformin increases microglial production of the anti-inflammatory cytokines IL-10 and TGF- under baseline conditions and enhances this upregulation following LPS-induced activation of microglia (Łabuzek et al., 2010a). Although TGF-/Smad signaling has been shown to have inflammatory and apoptotic roles in mild traumatic brain injury (TBI) (Patel et al., 2017), TGF- and IL-10 suppress microglial NO synthesis, while IL-10 also suppresses LPS-induced production of inflammatory cytokines IL-6, TNFα, and IL-1 (Ledeboer et al., 2000). Accordingly, metformin reduces LPS-induced microglial ROS production and NO synthesis with concurrent downregulation of inducible NO synthase (iNOS) and upregulation of arginase I (Łabuzek et al., 2010a). In the irradiated brain, TNFα is upregulated, which stimulates acute NF-κB signaling to promote CCL2 production in astrocytes, microglia, and endothelial cells (Hayashi et al., 1995; Thibeault et al., 2001; Lee et al., 2013). However, metformin has been shown to inhibit TNFα- induced NF-κB activation in human vascular endothelial cells in vitro, thereby decreasing the NF-κB-dependent transcription of proinflammatory genes, such as CCL2, intercellular adhesion molecule-1 (ICAM-1), VCAM-1, and E-selectin, which are involved in monocyte extravasation in regions of inflammation (Hattori et al., 2006). Furthermore, AMPK has also been shown to regulate CCR2, which is the major receptor for CCL2 that directs macrophage migration to sites of injury during the inflammatory response. Both AMPKα1 activation and NF-κB inhibition decreased LPS-induced CCR2 expression in macrophages in vitro (Kumase et al, 2016). Interestingly, CCR2 has also been implicated as a mediator of cognitive impairments following cranial IR. In a comparison of wild-type and CCR2-null mice, CCR2 deficiency attenuated deficits to HPC-dependent spatial learning following 10 Gy cranial IR, suggesting that metformin’s anti- inflammatory effects may promote recovery following cranial IR (Belarbi et al. 2013).

While metformin reduces the inflammatory phenotype of activated microglia and subsequent ROS/RNS production in vitro, chronic metformin treatment has been shown to promote AMPK-dependent alternative (M2) microglial activation following stroke, resulting in functional recovery associated with increased angiogenesis (increased density of CD31+ endothelial cells) in the ischemic striatum and SEZ neurogenesis (BrdU+DCX+ cells) 2 weeks post- stroke (Jin et al., 2014). In contrast to microglia, metformin-mediated AMPK activation in vascular endothelial cells has been shown to activate endothelial NOS (eNOS), which is required for normal SEZ-derived neurosphere formation and in vitro NPC migration (Chen et al., 2005; Calvert et al., 2008). Following stroke, eNOS activation contributes to BDNF-mediated NPC proliferation and migration from the SEZ, as well as angiogenesis, which are enhanced with metformin treatment (Chen et al., 2005; Liu et al., 2014). Therefore, metformin’s various effects in microglia and endothelial cells further support its potential role as a neuroprotective agent following injuries such as cranial IR, where preservation of the niche is crucial. Metformin may also exert neuroprotective effects through regulation of mTOR, which is a serine/threonine kinase found in 2 complexes: mTORC1 and mTORC2, each regulated by and involved in a variety of distinct signaling pathways in the brain (reviewed in Bockaert and Marin, 2015). In addition to activating NF-κB signaling, TNF increases mTORC1 activity in cancer cells, although it is unclear whether this mechanism persists in NPCs post-IR (Lee et al., 2007). However, mTORC1 activation in NPCs occurs within 30 minutes of 1 Gy IR in vitro and contributes to NPC apoptosis, since inhibition of mTORC1 prevents IR-induced caspase 3/7 activation (Sharlow et al., 2016). Similarly, in various non-neural cell types, mTORC1 inhibition reduces IR-mediated DNA damage and cellular senescence (Iglesias-Bartolome et al., 2012; Zheng et al., 2016). Furthermore, increased mTORC1 activity in adult-born DG neurons results in spatial memory deficits in Disrupted-in-schizophrenia 1 (Disc1) mutant mice, which is rescued by rapamycin-mediated mTORC1 inhibition (Zhou et al., 2013). While the direct effects of metformin on mTOR activity in NPCs are not yet characterized, metformin inhibits mTORC1 via AMPK-dependent and AMPK-independent pathways in several non-neural cell types, providing a basis for its potential as a protective agent in the brain’s neurogenic niches (Inoki et al., 2003; Gwinn et al., 2008; Sahra et al., 2011).

In summary, metformin promotes angiogenesis and neurogenesis in the postnatal brain through multiple pathways involved in NPC proliferation and differentiation, which improve cellular and functional outcomes following neonatal and adult stroke (Wang et al., 2012; Dadwal et al., 2015; Jin et al., 2014; Fatt et al., 2015). While some of these outcomes are mediated by direct signaling in NPCs, metformin also evokes responses from microglia and endothelial cells to promote angiogenesis and reduce neuroinflammation and oxidative stress within the SEZ (Łabuzek et al., 2010a; Liu et al., 2014). Given these pleiotropic effects, there is a strong basis for investigating metformin’s ability to promote recovery of neurogenesis following cranial IR.

2. Research Aims and Hypothesis

Cranial IR is a prominent adjuvant therapy used in the treatment of adolescent brain tumours. Despite improving survival rates, cranial IR induces delayed cognitive deficits. In rodent models, IR-induced cognitive deficits are associated with immediate and sustained reductions in neurogenesis. The SEZ of the lateral ventricles contains NSCs and their highly proliferative progeny that are necessary for lifelong OB neurogenesis and olfactory learning, and contribute to endogenous neurorepair following certain brain injuries. Since these cells are highly sensitive to IR and damage to their niche, therapies that protect their niche and enhance their recovery are necessary. Metformin, a drug commonly used to treat type 2 diabetes, has recently shown anti-inflammatory efficacy and enhances neurogenesis to promote functional recovery following neonatal stroke. Since the effects of cranial IR have predominantly been studied in adult rodents when neurogenesis is already on the decline, this thesis will investigate the impact of juvenile cranial IR. Given the clinical potential to deliver a prospectively neuroprotective drug prior to cranial IR, I will investigate the effects of metformin pretreatment on SEZ-derived neurogenesis following cranial IR in juvenile mice.

2.1. Hypothesis In a clinically relevant rodent model of juvenile cranial irradiation, metformin treatment will enhance subependymal zone neurogenesis and promote long-term olfactory memory recovery.

2.2. Objectives Objective 1: Investigate the effects of metformin on SEZ-derived aNSC pool size following juvenile cranial IR. Metformin will be administered for varying durations in neonatal and juvenile mice, since the SEZ continues to undergo architectural and compositional changes during the first 2 weeks of postnatal life (Tramontin et al., 2003). This developmental period may therefore exhibit differences in response to metformin depending on timing and duration of

51 administration and will provide insights into how the developing SEZ niche responds to metformin. NSCs become increasingly quiescent with age and have been shown to completely regenerate the NPC pool within 10 days following Ara-C-mediated ablation of proliferating cells in the adult SEZ (Doetsch et al., 1999a). However, the early postnatal SEZ contains a greater proportion of aNSCs and proliferating cells, resulting in increased oxidative stress and apoptosis, and a greater loss of nestin+ cells and DCX+ neuroblasts following IR-mediated ablation at P9, which is sustained in rats 10 weeks post-IR (Fukuda et al., 2005). However, nestin labels both aNSCs and TAPs, and therefore does not quantify aNSC pool size. Therefore, the neurosphere assay will be used to quantify aNSCs. a) Metformin will be administered once daily for 4 or 7 days beginning on P9 (neonatal) or P18 (juvenile), and the neurosphere assay will subsequently be employed within 24 hours of the final metformin injection to estimate aNSC pool size. b) After confirming the optimal timing of metformin administration, the neurosphere assay will be employed to estimate the size of the aNSC pool at varying times (ranging from 1 day to 1 month after terminating treatment) to identify whether metformin’s effects on the aNSC pool are maintained over time. c) The neurosphere assay will be employed at varying times following cranial IR to characterize the acute and chronic effects of juvenile cranial IR on the aNSC pool. d) The neurosphere assay will be employed at varying times following metformin pretreatment and cranial IR to assess whether metformin prevents NSC pool depletion or enhances the recovery of aNSCs following juvenile cranial IR.

Objective 2: Investigate potential mechanisms involved in metformin-induced and cranial IR- induced neurogenic responses in the SEZ. Long-term cellular deficits implicated in reduced neurogenesis have typically been characterized in vivo via IHC-based quantification of label-retaining cells and proliferative NPCs, lineage tracing of newborn OB neurons, and more rarely the expression of various receptors involved in growth factor signaling (Tada et al., 1999). However, the mechanisms underlying

52 injury-induced reductions in neurogenesis are often unclear, including whether NSC activation is correlated with downstream cellular outcomes. It is also unclear whether injury-induced reductions in SEZ neurogenesis are intrinsically regulated (e.g., due to altered transcriptomes) or mediated by the niche, which is a question that has recently become more prevalent (Su et al., 2014; Tong et al., 2014; Chaker et al., 2016). Recent findings have implicated the neurogenic niche as the dominant mediator of quiescence in the aged SEZ, making it plausible that the niche also regulates neurogenesis in response to injury (Piccin et al., 2014; Silva-Vargas et al., 2016; Kalamakis et al., 2019). a) The ratio of qNSCs to aNSCs within the SEZ will be quantified via FACS 2 days following metformin pretreatment and cranial IR to assess whether changes to NSC activation are correlated with downstream cellular outcomes (Objectives 1c-d, 2c). b) Conditioned media (CM) will be generated from SEZ tissue of mice 1 day after receiving cranial IR (IR-CM) or sham-IR (Ctrl-CM). The neurosphere assay will be used to culture SEZ-derived cells from naïve mice in normal growth media, Ctrl-CM, or IR-CM to assess whether secreted factors from the irradiated niche are sufficient to alter NSC pool size. c) DCX+ and Ki67+ cells within the SEZ will be quantified in vivo via IHC at varying times following metformin pretreatment and cranial IR to assess recovery of proliferating cells and neuroblasts.

Objective 3: Investigate long-term olfactory memory following metformin treatment and juvenile cranial IR. Postnatal SEZ-OB neurogenesis plays a crucial role in the retention of long-term olfactory memory (LTOM) (Lemasson et al., 2005; Sultan et al., 2010). However, deficits to neurogenesis caused by cranial IR result in deficits to LTOM that do not spontaneously recover (Lazarini et al., 2009). Therefore, the effects of metformin pretreatment on LTOM will be assessed. a) Mice will be subjected to a LTOM task in which they will be trained to associate an odor with a reward 1 month post-IR and will subsequently be tested 1 month after training to assess whether metformin pretreatment and juvenile cranial IR affect LTOM.

3. Methods 3.1. Animals Young adult (6-week-old) C57Bl/6 mice were purchased from Charles River (#027; Montréal, Canada) for use as breeders. For flow cytometry experiments, transgenic hGFAP::GFP mice expressing green fluorescent protein (GFP) under the control of the human GFAP promoter were purchased from The Jackson Laboratory (#003257; Ellsworth, Maine, USA) and used as breeders (Zhuo et al., 1997). Mice of both sexes were used for all experiments. All mice were maintained on a 12-hour day/night light cycle and housed in cages containing a nestlet and shelter. Food and water were available ad libitum, except for mice undergoing the long- term olfactory memory (LTOM) task, in which food was restricted for 10 days prior to and during training, as well as 3 days prior to testing in order to maintain a body weight of 85-90% of baseline levels. Mice were euthanized on P13, P16, P18, P19, P22, P25, P47, or P84 according to experiment, via an overdose of the anaesthetic avertin, which was delivered by intraperitoneal (IP) injection. Avertin was prepared by dissolving 0.25g 2,2,2-tribromoethanol (Sigma-Aldrich, T48402) and 0.5mL 2-methyl-2-butanol (Sigma-Aldrich, 152463) in 20mL of sterile double-distilled water. Following avertin overdose, mice underwent cervical dislocation and decapitation for neurosphere assay experiments, or transcardial perfusion with 4% paraformaldehyde (PFA). All animal work was performed in accordance with institutional guidelines and approved by the University of Toronto Animal Care Committee (animal use protocol 20011476).

3.2. Metformin administration For in vivo metformin treatment, metformin (1,1-dimethylbiguanide hydrochloride; Sigma-Aldrich, D150959) was dissolved in sterile phosphate-buffered saline (PBS) at a concentration of 20mg/mL and administered once daily via subcutaneous injection at 200mg/kg body weight (10µL/g body weight), a dose that has been reported to be safe for long-term rodent administration (Quaile et al., 2010). PBS was administered (10µL/g body weight) as a vehicle control. For all metformin and PBS injections, a 25G needle was used.

3.3. Cranial irradiation For cranial IR experiments, mice were anaesthetized on P17 via IP injection of avertin at a dose of 20µL/g body weight (250mg/kg body weight 2,2,2-tribromoethanol). Mice were secured inside a lead shield with the head exposed by a circular opening in the top of the shield, and subsequently placed in a cesium-137 gamma irradiator (Best Theratronics, Gamacell 40 Exactor). Mice received 8 grays (Gy) of whole brain cranial IR at a rate of ~1 Gy/minute. Following cranial IR, mice were returned to their home cage where they recovered under a heat lamp until regaining consciousness.

3.4. Neurosphere assay 3.4.1. Tissue dissection Mice were euthanized via avertin overdose (IP injection) and cervical dislocation. Heads and necks were doused with 70% ethanol to matte down fur prior to decapitation with Mayo scissors. An incision was made through the skin from the base of the skull to between the eyes in order to peel away the skin. The skull was cut along the sagittal suture from the base of the skull towards the coronal suture and up through the midline of the nasal bones. The nasal, frontal, and parietal bones were peeled away using forceps to expose the brain. Brains were removed from their skull using a thin tipped spatula and transferred into separate 60x15mm polystyrene petri dishes (Fisher Scientific, FB0875713A) pre-filled with cold artificial cerebrospinal fluid (aCSF; Appendix 1A) on ice. Brains were cut in half along the longitudinal fissure using a straight razor to separate hemispheres. With the medial aspect of the hemisphere facing up and using microdissection scissors, an incision was made from the dorsal surface of the cortex to the posterior aspect of the corpus callosum at the midline, allowing for the cortex to be peeled back and cut away to expose the lateral ventricles and striatum. With the dorsal aspect of the brain facing up and using fine, curved microdissection scissors, incisions were made on either side of the medial and lateral walls of the lateral ventricles to remove the medial and lateral SEZ. SEZ tissue from each sample was transferred to individual Falcon 15mL polystyrene conical tubes (Fisher

Scientific, 0552790) and suspended in 0.25mL cold hi/lo aCSF (Appendix 1B) on ice until processing.

3.4.2. Cell culture SEZ tissue was enzymatically digested in 6mL/sample enzyme solution consisting of 1.33mg/mL trypsin (Sigma-Aldrich, T1005), 1245 units/mL (~0.76mg/mL) hyaluronidase (Sigma- Aldrich, H6254), and 0.13mg/mL kynurenic acid (Sigma-Aldrich, K3375) in hi/lo aCSF. During enzymatic digestion, samples were incubated on a rocker at 37oC for 25 minutes, and then centrifuged for 5 minutes at 1500rpm. The supernatant was decanted and replaced with 2mL/sample trypsin inhibitor solution consisting of 0.67mg/mL ovomucoid trypsin inhibitor (Worthington, LS003086) in serum-free media (SFM; Appendix 2). Samples were gently triturated 30x with a P1000 micropipette (set to 1000µL) to dissociate tissue into single cells and then centrifuged for 5 minutes at 1500rpm. To wash the samples, the supernatant was decanted and replaced with 2mL/sample SFM supplemented with mitogens (SFM+EFH), and samples were gently triturated 10x with a P1000 micropipette. SFM+EFH consisted of 20ng/mL epidermal growth factor (EGF; Gibco, PMG8041), 10ng/mL basic fibroblast growth factor (bFGF; Gibco, PHG0266), and 2µg/mL heparin (Sigma-Aldrich, H3149) in SFM. Samples were then centrifuged once more for 3 minutes at 1500rpm. After decanting the supernatant, cells were suspended in 1mL SFM+EFH and gently triturated 5x with a P1000 micropipette. Live cell density was quantified using a haemocytometer. Cells were subsequently plated at 10 cells/µL (Coles-Takabe et al., 2008) in 500µL SFM+EFH/well in Nunc Cell Culture Treated 24-well plates (Thermo Fisher Scientific, 142475). Cultures were incubated at 37oC with

5% carbon dioxide (CO2) and neurospheres were counted after 7 days of undisturbed incubation.

3.4.3. Conditioned media For conditioned media experiments, mice were euthanized on P18, and brains were isolated and dissected as described in section 3.4.1. The SEZ tissue from 3 brains per group (Control vs. IR) was pooled with tissue from the same group. SEZ tissue was then digested and

56 suspended as single cells in SFM+EFH as described in section 3.4.2. Live cell density was quantified using a haemocytometer. To generate conditioned media, cells were plated at 40 cells/µL in SFM+EFH in Falcon Tissue Culture Treated 50mL flasks (Fisher Scientific, 101269).

o Cultures were incubated at 37 C with 5% CO2 for 24 hours, whereupon samples were transferred to 15mL polystyrene conical tubes and centrifuged for 5 minutes at 1500rpm. The supernatant was collected and filtered through a 0.22µm Millex-GP sterile syringe filter (Fisher Scientific, SLGP033RS) for use as conditioned media. SEZ tissue was isolated from P19 control mice (sham-irradiated on P17) and processed as described in section 3.4.2. Cells were then plated in 500µL SFM+EFH alone, control conditioned media (from sham-irradiated mice), or IR conditioned media (from P17-irradiated mice) at 10 cells/µL in Nunc Cell Culture Treated 24-well plates. Cultures were incubated at

o 37 C with 5% CO2 and neurospheres were counted after 7 days of undisturbed incubation.

3.4.4. Quantification For all neurosphere assays, including conditioned media experiments, cultures were incubated for 7 days prior to quantification. Neurospheres were characterized as round clusters of cells having a diameter ≥80µm. Smaller clusters were excluded from analyses.

3.5. Fluorescence-activated cell sorting (FACS) 3.5.1. Reagents and solutions FACS of qNSCs and aNSCs was performed as described by Codega et al. (2014) with various modifications described below. All stock and working solutions were prepared in sterile conditions and stored according to Appendix 3.

3.5.2. Tissue dissection Mice were euthanized on P19, and brains were isolated as described in section 3.4.1. Brains were removed from their skulls and transferred to 60x15mm polystyrene petri dishes (Fisher Scientific, FB0875713A) pre-filled with cold PBS on ice. SEZ tissue was isolated as described in section 3.4.1 and was transferred to 35x10mm polystyrene petri dishes (Fisher

Scientific, 08757100A) pre-filled with freshly made cold 1,4-Piperazinediethanesulfuronic acid (PIPES) solution (Appendix 3B) on ice. Given the relative scarcity of NSCs compared to other cells in the postnatal brain, SEZ tissue from 5 GFAP::GFP mice per group was pooled for each FACS experiment. For single-colour controls used for setting gates and adjusting channel compensation, the SEZ tissue from 2 wild type (WT) C57Bl/6 mice was pooled for each FACS experiment. Once in PIPES solution, the SEZ tissue was finely minced into small pieces using microdissection scissors.

3.5.3. Cell suspension and staining Immediately prior to mincing the SEZ tissue into small pieces, papain (Worthington, LS003127) was activated in papain activating solution (Appendix 3B) at 45 units/mL (2.25mg/mL). Papain was activated for 20-30 minutes at room temperature before use. After mincing, pooled SEZ tissue from each group (5 GFAP::GFP brains/group, 2 WT brains) was transferred to Falcon 15mL polystyrene conical tubes (Fisher Scientific, 0552790). The volume of PIPES solution was adjusted to 8mL/sample (4mL for WT control tissue). 2mL activated papain was then added to each sample (1mL for WT control tissue) and was mixed with tissue by inverting the samples several times. Samples were incubated on a rocker at 37oC for 10 minutes and were subsequently centrifuged for 5 minutes at 1300rpm at 4oC. The supernatant was decanted and replaced with 1mL Dulbecco’s Modified Eagle Medium/Ham’s F12 (DMEM/F12; DMEM from Gibco, 12100046; F12 from Gibco, 21700075), which was supplemented with 100µL ovomucoid solution (7mg/mL; Appendix 3A) to inhibit papain activity and 33.4µL DNase solution (15mg/mL; Appendix 3A) to reduce viscosity caused by genomic DNA. Samples were gently triturated 20x with a P1000 micropipette (set to 1000µL) and then gently triturated 50x with a P200 micropipette (set to 150µL) to dissociate tissue into single cells. The cell suspension from each tube was divided into 2 new conical tubes pre-filled with 4mL cold 22% Percoll (Appendix 3B) by gently dripping half of each cell suspension on top of each Percoll gradient solution without disturbing the layering. Samples were then centrifuged for 10 minutes at 1800rpm at 4oC. A P1000 micropipette was used to remove myelin residuals

58 and pink media from the top of the Percoll gradient solution, while the remaining Percoll was removed by gently decanting. 2mL of staining solution (Appendix 3B) was added to each tube and samples were gently triturated 10x with a P1000 micropipette. Samples that were divided for the Percoll gradient were re-combined (totalling 4mL staining solution) and centrifuged for 5 minutes at 1300rpm at 4oC. The supernatant was decanted and replaced with 720µL staining solution for GFAP::GFP samples, while the WT sample was topped up to 180µL staining solution. The following 5 single colour controls were prepared for adjusting compensation parameters: Unstained, DAPI, PE-Cy7, A647, GFP. The following 3 fluorescence minus one (FMO) controls were prepared for setting gates: no GFAP::GFP FMO (fully stained WT), no CD133-PE-Cy7 FMO, no EGF-A647 FMO. All single colour and FMO controls were prepared by first adding 30µL of GFAP::GFP sample (GFP, CD133 FMO, A647 FMO) or WT sample (unstained, DAPI, PE-Cy7, A647, GFP FMO) to 1.5mL polypropylene microcentrifuge tubes (Corning Life Sciences, MCT-150-C), while the remaining 600µL GFAP::GFP samples were used for sorting. CD133 (prominin-1) was labeled using an anti-CD133-biotin antibody (eBioscience, 13-1331-82) at a concentration of 1:300 in staining solution. After adding anti-CD133-biotin to the appropriate tubes, controls and sort samples were incubated on ice for 15 minutes, washed with 5mL staining solution, and centrifuged for 5 minutes at 1300rpm at 4oC. The supernatant was decanted, and sort samples were topped up to 600µL, while single colour and FMO controls were topped up to 200µL staining solution. Streptavidin-PE-Cy7 (eBioscience, 25-4317- 82) at 1:1000 was added to bind anti-CD133-biotin, while EGFR was labelled by EGF conjugated to Alexa Fluor 647 (EGF-A647; Invitrogen, E35351) at 1:300. Upon adding streptavidin-PE-Cy7 and EGF-A647 to the appropriate tubes, samples were again incubated on ice for 15 minutes, washed with 5mL staining solution, and centrifuged for 5 minutes at 1300rpm at 4oC. Supernatant was decanted, and sort samples were topped up to 600µL, while single colour and FMO controls were topped up to 200µL staining solution. 4′,6-diamidino-2-phenylindole (DAPI; Invitrogen, D1306) was then added to the appropriate tubes at 1:10,000 for exclusion of dead cells during sorting. Samples were strained through sterile 40µm nylon cell strainers (Fisher Scientific, 22-363-547) into Falcon 5mL round-bottom polypropylene tubes (Corning Life

Sciences, C352063) to eliminate clumps of cells that may clog FACS machinery. Cells were then immediately analyzed via FACS.

3.5.4. Sorting All cell sorting was conducted using a BD Influx (BD Biosciences, California, USA) or BD FACS Aria IIIu (BD Biosciences, California, USA) with a sheath pressure of 13psi and 100µm nozzle aperture. FMO controls were used to set gates and single colour controls were used to adjust compensation parameters. BD FACSDiva software (BD Biosciences, California, USA) was used for equipment setup and data acquisition, while FlowJo 9.3 (FlowJo LLC, Oregon, USA) was used for data analysis. Cells were first gated using forward scatter area and side scatter area to exclude debris, followed by forward scatter area and forward scatter height to exclude doublets. DAPI exclusion was used to gate for live cells. The GFAP::GFP FMO, CD133-Pe-Cy7 FMO, and EGF-A647 FMO controls were used to set the gates for GFAP::GFP+, CD133+, and EGFR+ populations, respectively. These gates were subsequently used to sort for GFAP::GFP+CD133+EGFR- qNSCs and GFAP::GFP+CD133+EGFR+ aNSCs, and data was presented using biexponential scaling (Fig. 4.8B-C). The relative proportions of qNSCs and aNSCs were compared (Fig. 4.8D).

3.6. Immunohistochemistry and immunofluorescence 3.6.1. Perfusions and tissue preparation Mice were euthanized on P19, P47, or P84 via avertin overdose. An incision was made from the midline of the abdomen and up through the ribs and diaphragm to expose the heart. Mice were transcardially perfused with ice cold PBS, followed by transcardial perfusion with ice cold 4% PFA (7.2-7.3 pH) in PBS. Brains were removed from skulls as described in section 3.4.1 and post-fixed in 4% PFA at 4oC for 4-5 hours. Brains were then cryoprotected in 30% sucrose solution at 4oC until sectioned on an HM525 NX Cryostat (Thermo Scientific, 956640). Brains were mounted in Shandon Cryomatrix embedding resin (Thermo Scientific, 6769006) and frozen at -24oC within the cryostat. Coronal sections (20µm thick) were then mounted on

Superfrost Plus microscope slides (Fisher Scientific, 12-550-15) for antibody labeling and imaging.

3.6.2. Antibody labeling Coronal sections were first rehydrated in PBS for 5 minutes at room temperature before being submerged in citrate buffer (pH 6.0), comprised of 1.92g/L citric acid (Sigma-Aldrich,

240621) and 0.05% Tween 20 (Sigma-Aldrich, P1379) in double distilled water (ddH2O). Sections in citrate buffer underwent heat-induced epitope retrieval in a pressure cooker (Nesco, PC-6- 25-30TPR) on high heat setting for 15 minutes. Slides were allowed to cool at room temperature for 10 minutes and were washed 3x in PBS for 5 minutes per wash at room temperature. A blocking solution was applied for 1 hour at room temperature and was comprised of 5% normal donkey serum (Sigma-Aldrich, D9663), 10mg/mL bovine serum albumin (Sigma-Aldrich, A9647), and 0.3% Triton X-100 (Sigma-Aldrich, T9284) in PBS. Primary antibodies were prepared in blocking solution and applied to slides for overnight (16 hours) incubation at 4oC. Neuroblasts were labeled using a mouse anti-doublecortin (DCX) antibody (Santa Cruz Biotechnology, sc-271390) at 1:400 and proliferating cells were labeled using a rabbit anti-Ki67 antibody (Abcam, ab15580) at 1:500. After overnight incubation with primary antibodies, slides were washed 3x in 0.2% Tween 20 in PBS for 5 minutes per wash at room temperature. Secondary antibodies and DAPI (Invitrogen, D1306) were prepared in PBS and applied to slides for 90 minutes at room temperature. DCX was labeled by donkey anti-mouse Alexa Fluor 568 (Invitrogen, A10037) at 1:750 and Ki67 was labeled by donkey anti-rabbit Alexa Fluor 488 (Invitrogen, A-21206) at 1:750. DAPI was used to label nuclei at a concentration of 1:10,000. Slides were then washed 3x in PBS for 5 minutes per wash at room temperature, followed by application of mounting medium (Dako, S302380-2) and coverslips. Coverslips were allowed to seal at room temperature overnight and slides were subsequently stored at -20oC.

3.6.3. Microscopy and quantification Imaging was performed on an Axio Observer Z1 inverted motorized microscope (Zeiss, Germany) with a high-speed spinning disk CSU-X1 confocal scanner unit (Yokogawa, Texas, USA)

61 and an Axiocam 506 high resolution camera (Zeiss, Germany). Zen (Zeiss, Germany) and Fiji (Schindelin et al., 2012) software were used for image acquisition and processing. Three sections per animal selected based on their rostral-caudal location. An anterior section was selected where the forceps minor of the corpus callosum met at the midline, a posterior section was selected where the anterior commissure met at the midline, and a middle section was selected halfway between these two points. The LV of each hemisphere was imaged at 20x magnification and tiles were stitched together to display the entire LV and SEZ, totaling 6 tiled images per mouse. Three regions of interest (ROIs) in the SEZ were analyzed and pooled from each LV. The dorsolateral corner, the midpoint of the lateral wall, and the ventral corner were analyzed in 150x50µm ROIs. The total number of DCX+Ki67+ cells in each ROI was counted and expressed as a proportion of DAPI+ cells. All cells were manually counted using the Fiji Cell Counter plugin.

3.7. Long-term olfactory memory task 3.7.1. Food restriction, acclimation, and materials A sand digging task was utilized to measure long-term olfactory memory, in which an odor cue is paired with a food reward (Zou et al., 2015). In order to motivate mice to search for an odor-cued food reward, mice were food restricted prior to and during behavioural training and testing. Food and mice were weighed daily from P32-37 to determine average ad libitum food consumption and body weight. Mice were food restricted to 50-80% of daily ad libitum food consumption starting 10 days prior to training until the end of training (from P37-54) to maintain 85-90% of baseline body weight (Fig. 3.1A). Prior to training and test sessions, mice were allowed to acclimate in a clean, empty cage with bedding for 30 minutes, which was used as the training and testing arena. From P41- 43, mice underwent daily acclimation sessions to sugar reward pellets (Bio-Serv, F05550) and the reward/sand delivery apparatus, in which 3 sugar pellets were placed in the base of a 35x10mm polystyrene petri dish (Fisher Scientific, 08757100A) atop the reward/sand delivery apparatus. The reward/sand delivery apparatus was constructed by gluing two 48-well culture plate lids (Fisher Scientific, 12-565-322) perpendicular to each other at their edges to form the

62 platform base and handle. An additional lid cut to 4 cm tall was used as a divider and glued between the base and the handle to separate left and right sides of the platform. The lid from a 35x10mm polystyrene petri dish was glued upside-down on either side of the divider to hold a reward/sand-filled 35x10mm polystyrene petri dish (Fig. 3.1B, left image). Dishes were filled with autoclaved sand (Canadian Tire, 059-4557-4) for all trials after P41-43 acclimation. During training trials, sand was scented with 100 µL 10mM isoamyl acetate (IAA; Sigma, W205508) in the reward dish and 100 µL 10mM Citralva (3,7-dimethyl-2,6- octadienenitrile; Sigma, W512508) in the non-reward dish.

3.7.2. Pre-training and training From P44-46, mice underwent daily pre-training sessions (2 blocks of 4 trials, 1 minute inter-trial interval, 4 hour inter-block interval) to learn to dig and retrieve sugar pellets from the sand, during which only one of the two dishes were filled with sand and baited with a sugar pellet. The left or right location of the sand-filled dish with a sugar pellet was randomized among the trials. On the first day of pre-training (P44), the sugar pellet was placed on top of the sand. On the second day of pre-training (P45), the sugar pellet was partially buried (half exposed) for the first block of trials, and completely buried just beneath the surface of the sand for the second block of trials. On the third day of pre-training (P46), the sugar pellet was completely buried at the bottom of the dish. Following pre-training, mice were trained to associate the reward (sugar pellet) with IAA, while Citralva was used to scent the non-reward dish. Training took place for 8 days (P47- 54) with 2 blocks of 4 trials per day (1 minute inter-trial interval, 4 hour inter-block interval). On the reward/sand delivery apparatus, both dishes were filled with sand. The dish containing the reward was scented with 100 µL 10mM IAA and the non-reward dish with 100 µL 10mM Citralva. The location of the reward dish containing a buried sugar pellet was randomized between trials (Fig. 3.1B). Upon placing the loaded sand/reward delivery apparatus in the cage, mice were allowed to dig in one dish. If a mouse dug in the reward dish, the trial was recorded as correct. If a mouse dug in the non-reward dish, the trial was recorded as incorrect. The apparatus was removed immediately after a mouse had correctly or incorrectly dug for the

63 reward (mice were not permitted to dig in both dishes). Scores were quantified as a percentage of correct trials relative to total training trials on each day.

3.7.3. Testing Mice that correctly chose the reward-associated dish at an average rate of ≥80% over the final two training days (P53-54) passed the training phase and were tested 1 month after the final training day (P84). Food and mice were weighed daily from P76-81 to determine average ad libitum food consumption and body weight, and mice were food restricted to 50% of daily ad libitum consumption levels from P81-84. Testing on P84 involved a single block of 4 trials (1 minute inter-trial interval) with no reward under either of the scented dishes. The location of the 10mM IAA- and 10mM Citralva-scented dishes was randomized between trials. If a mouse dug in the IAA-scented (reward-associated) dish, the trial was scored as correct, while digging in the Citralva-scented (non-reward-associated) dish, the trial was scored as incorrect. As in training, mice were only permitted to dig in one dish. Scores were quantified as a percentage of correct trials relative to total testing trials.

Figure 3.1. Schematic of long-term olfactory memory task. (A) Experimental timeline. (B) Schematic of training/testing arena with reward/sand delivery apparatus (image from Zou et al., 2015) and training/testing paradigm.

3.8. Statistical Analysis All raw data was recorded and processed using Microsoft Excel (Microsoft, Washington, USA). Statistical analyses and graphing were performed using GraphPad Prism 6 (GraphPad Software, California, USA). All quantified values were presented as group mean ± standard error of the mean (SEM). Column analyses between 2 groups were performed using unpaired t tests, or multiple unpaired t tests when comparing 2 groups over multiple time points. Column analyses between 3 groups were performed using one-way ANOVA with Tukey’s test for multiple comparisons. Grouped analyses were performed using two-way ANOVA with Tukey’s test for multiple comparisons. Statistical significance was characterized as p<0.05.

4. Results 4.1. Effects of metformin on aNSC pool size 4.1.1. Metformin expands the neonatal, but not the juvenile NSC pool in the SEZ. One of our primary objectives was to examine the potential for metformin to promote neurogenesis in a model of juvenile cranial IR. Previous work by our lab demonstrated that metformin-induced functional recovery in a neonatal model of hypoxia-ischemia was correlated with expansion of the SEZ NSC pool (Dadwal et al., 2015). To determine an optimal time for metformin treatment, and building on previous studies that suggest there are age-dependent effects of metformin on the SEZ NSC pool (Ruddy et al., 2019a), metformin was administered via subcutaneous injection once daily (200mg/kg/day) for either 4 or 7 days, beginning on P9 (neonatal) or P18 (juvenile). The neurosphere assay was performed 1 day after the final metformin injection and neurospheres were counted to assess the size of the aNSC pool. Both 4 and 7 days of metformin administration in neonates significantly increased the number of SEZ- derived neurospheres formed relative to PBS-treated controls (Fig. 4.1A-B). However, 4 or 7 days of metformin administration in juvenile mice did not result in significant changes to neurosphere numbers (Fig. 4.1C-D). Hence, with the goal of expanding the aNSC pool to promote neural repair, we used the 7 day treatment paradigm (from P9-15) for subsequent experiments. Since previous studies have demonstrated sex-dependent effects of metformin on NPCs (Ruddy et al., 2019a), we next compared the NSC expansion in males and females from the P16 SEZ following 1 week (P9-15) metformin treatment (Fig. 4.2). As predicted, metformin administration resulted in a similar expansion in the SEZ pool of both sexes (1.49±0.19-fold versus 1.42±0.20-fold increase, males and females, respectively), likely due to the prepubescent age at which metformin was administered. Interestingly, our lab has recently identified that when metformin is administered to neonatal mice at a lower dose of 20mg/kg/day, an equivalent neurosphere expansion is observed in both sexes (~1.4-fold increase) (Ruddy et al., 2019a). Therefore, both sexes were used for all subsequent experiments. Despite the potential

66 for metformin to expand the NSC pool at a lower dose, 200mg/kg/day was chosen for subsequent experiments since it has not yet been determined whether lower doses are sufficient for metformin to exert its pleiotropic effects on other cell types in the CNS (e.g., neuroblasts and microglia).

Figure 4.1. Metformin expands the neonatal, but not the juvenile aNSC pool in the SEZ. (A) Neurosphere formation after 4 days of neonatal metformin treatment (***p<0.001; n≥15 animals per group). (B) Neurosphere formation after 1 week of neonatal metformin treatment (**p<0.01; n≥15 animals per group). (C) Neurosphere formation after 4 days of juvenile metformin treatment (n≥10 animals per group). (D) Neurosphere formation after 1 week of juvenile metformin treatment (n≥8 animals per group). All comparisons were made using unpaired t tests. Data is presented as mean ± SEM.

Figure 4.2. Metformin expands the male and female neonatal aNSC pool. Data represents neurosphere formation after 1 week of neonatal metformin treatment (*p<0.05 relative to same sex control, unpaired t tests; n≥7 animals per group). Two-way ANOVA revealed a significant effect of metformin (p<0.01), but not sex (p=0.81). Data is presented as mean ± SEM.

4.1.2. Metformin induces a short-term expansion of the aNSC pool in the SEZ. We have recently established a juvenile model of cranial IR in which mice receive 8 Gy IR to the entire brain on P17 (Ruddy et al., 2019b, submitted to Brain Research). Briefly, mice are anesthetized with an IP injection of Avertin (tribromoethanol, 250mg/kg) and placed in a lead shield to exclusively expose the head to IR. Mice are placed in a Cesium-137 irradiator and receive 8 Gy cranial IR (~1 Gy/min) prior to being returned to their home cage under a heat lamp until recovery from anesthesia. In a first series of experiments, we sought to determine how long the metformin-induced expansion of the neonatal aNSC pool is sustained relative to the time when cranial IR would be employed. Mice received metformin treatment from P9-15 and were sacrificed on P16, P19, P25, or P47 for the neurosphere assay (Fig. 4.3A). We observed significantly greater numbers of neurospheres at P16 and P19 relative to PBS-treated controls. By P25, the metformin-induced increase in neurosphere formation was no longer statistically significant (Fig. 4.3B). These findings suggest that in vivo metformin treatment promotes a transient expansion of the aNSC pool, since cessation of treatment leads to the

68 aNSC pool returning to control levels. Importantly, this reveals that metformin-induced expansion of the aNSC pool does not cause persistent growth or NSC pool depletion.

Figure 4.3. Metformin induces a short-term expansion of the aNSC pool in the SEZ. (A) Experimental timeline, in which the neurosphere assay was conducted on P16, P19, P25, and P47 following 1 week (P9-15) of metformin treatment. (B) Neurosphere formation at the ages specified in (A) after 1 week of neonatal metformin treatment (*p<0.05, **p<0.01, multiple unpaired t tests; n≥7 animals per group). Data is presented as mean ± SEM.

4.2. Effects of metformin pretreatment on aNSC pool size following juvenile cranial IR 4.2.1. Juvenile cranial IR causes an acute depletion of aNSCs that recover spontaneously over time. Prior to assessing whether metformin pretreatment could prevent or reduce cranial IR- medicated deficits to aNSC pool size, we first characterized the acute and chronic effects of juvenile cranial IR on SEZ aNSC pool size by employing the neurosphere assay at 2, 8, and 30

69 days post-IR (Fig. 4.4A). Similar to previous reports of spontaneous NSC recovery at 9 weeks post-IR (Hellström et al., 2009), we found that neurosphere formation partially recovered 8 days post-IR and had recovered to non-irradiated control levels by 30 days post-IR (Fig. 4.4B).

Figure 4.4. Spontaneous recovery of aNSCs occurs within 30 days post-IR. (A) Experimental timeline. (B) Neurosphere formation relative to age-matched non-irradiated controls at the ages specified in (A) following cranial IR at P17 (*p<0.05, ****p<0.0001, multiple unpaired t tests; n≥7 animals per group). Data is presented as mean ± SEM.

4.2.2. Metformin pretreatment does not prevent the acute depletion of aNSCs following juvenile cranial IR. Given that neonatal metformin treatment (from P9-15) expanded the SEZ aNSC pool until P19 (Fig. 4.3B), we predicted that this expansion may result in attenuated deficits to aNSC pool size caused by cranial IR. Therefore, we used the neurosphere assay at P19 to investigate whether metformin pretreatment protects NSCs against cranial IR on P17 (Fig. 4.5A). Mice that received 8 Gy cranial IR on P17 had a >50% reduction in neurospheres at P19 relative to non-

70 irradiated controls. However, metformin pretreatment did not prevent this deficit to neurosphere formation (Fig. 4.5B).

Figure 4.5. Metformin pretreatment does not prevent the acute depletion of aNSCs 2 days post- IR. (A) Experimental timeline. (B) Neurosphere formation relative to non-irradiated controls at P19 following metformin pretreatment and cranial IR (***p<0.001, ****p<0.0001, one-way ANOVA with Tukey’s test for multiple comparisons; n≥11 animals per group). Data is presented as mean ± SEM.

4.2.3. Metformin pretreatment does not affect aNSC pool recovery following juvenile cranial IR. Metformin pretreatment did not prevent the acute IR-induced depletion of the SEZ aNSC pool at P19 so we next asked if metformin pretreatment could enhance recovery of aNSCs, potentially due to metformin’s reported anti-inflammatory and/or pro-survival effects in the SEZ niche (Hattori et al., 2006; Hou et al., 2010; Łabuzek et al., 2010a; Jin et al., 2014). The finding that spontaneous recovery of the SEZ aNSC pool occurs following juvenile cranial IR (Fig. 4.4B) supports a possible compensatory mechanism of NSC activation to mitigate the persistent deficits to cell proliferation and neurogenesis in the SEZ and OB, respectively

(Hellström et al., 2009). Thus, we investigated whether metformin pretreatment was sufficient to accelerate or enhance aNSC pool recovery at 8 and 30 days post-IR (Fig. 4.6A). As shown in Figure 4.6B, metformin pretreatment did not alter the recovery of neurosphere formation following cranial IR.

Figure 4.6. Metformin pretreatment does not affect aNSC pool recovery following juvenile cranial IR. (A) Experimental timeline. (B-C) Neurosphere formation relative to non-irradiated controls at (B) P25 (n≥13 animals per group) and at (C) P47 (n≥7 animals per group). At both ages, no significant differences were found between groups using one-way ANOVA with Tukey’s test for multiple comparisons. Data is presented as mean ± SEM.

4.3. Effects of metformin pretreatment on NSC activation state in the SEZ following juvenile cranial IR Repopulation of the NSC pool has been observed in a number of injury models that deplete the proliferating progenitors in the SEZ (Morshead et al. 1994; Doetsch et al., 1999a). More recently, Daynac et al. showed that a low dose (4 Gy) of cranial IR in young adult mice resulted in a depletion of aNSCs at 2 days post-IR and subsequent activation of qNSCs to repopulate the aNSC pool at 3 days post-IR (Daynac et al., 2013). We hypothesized that the return of aNSCs to control levels after cranial IR and metformin pretreatment was due to activation of qNSCs. To test this hypothesis, we used GFAP::GFP mice to isolate NSCs based on their activation state with FACS (Codega et al., 2014). We performed FACS at 2 days post-IR from control, metformin-treated, cranial IR, and metformin pretreatment + cranial IR mice (Fig. 4.7A). The SEZ of 5 mice per treatment group were pooled and stained for CD133 and EGFR as previously described (Codega et al., 2014), in order to quantify GFAP::GFP+CD133+EGFR- qNSCs and GFAP::GFP+CD133+EGFR+ aNSCs (Fig. 4.7B-C). We compared the relative percentages of activated and quiescent NSCs from all groups. Approximately 60% of NSCs were activated in control and metformin-treated mice, suggesting that metformin did not expand the size of the NSC pool by pulling NSCs out of the quiescent state. While not significantly different (via two- way ANOVA), a trend was observed in cranially irradiated mice, which skewed the NSC pool towards quiescence (Fig. 4.7D). These findings therefore suggest that juvenile cranial IR does not result in a reduced proportion of aNSCs relative to qNSCs at 2 days post-IR.

Figure 4.7. Neither metformin pretreatment nor juvenile cranial IR resulted in an altered ratio of aNSCs to qNSCs. (A) Experimental timeline. (B-C) Representative FACS plots showing gating strategy, in which (B) GFAP::GFP+ cells are first selected and then (C) gated for CD133-PECy7 and EGF-A647. (D) Quantification of GFAP::GFP+CD133+EGFR- qNSCs and GFAP::GFP+CD133+EGFR+ aNSCs via FACS 2 days post-IR (n≥3 sorts per group). Two-way ANOVA found no significant effects on aNSC% by cranial IR or metformin and no significant differences were found between groups with Tukey’s test for multiple comparisons. Data is presented as mean ± SEM.

4.4. Factors in the irradiated SEZ niche reduce aNSC pool size in vitro. Cranial IR-induced inflammation has been shown to contribute to reduced DG neurogenesis and subsequent impairments to hippocampal-dependent spatial memory, which can be partially rescued with anti-inflammatory drugs or deficiency of pro-inflammatory cell signaling components (Monje et al., 2003; Belarbi et al., 2013; Lee et al., 2013; Han et al. 2016). These studies highlight the significant impact of the niche on regulating NSC behaviour and neurogenesis. Inflammation has also been observed in the SEZ following cranial IR, although it is not known whether the IR-induced inflammation is sufficient to reduce the size of the NSC pool (Kalm et al., 2009). Notably, SEZ neural precursors (NSCs and progenitor cells) cultured in the presence of irradiated brain endothelial cells or transplanted into the irradiated SEZ undergo apoptosis and reduced proliferation, respectively (Pineda et al., 2013). Herein, we sought to determine whether factors from the irradiated SEZ can affect NSC behaviour from non-irradiated mice. The neurosphere assay was employed using cells from the SEZ of non-irradiated P19 mice cultured in conditioned media (CM) derived from the P18 SEZ of control (Ctrl CM) or cranially irradiated (IR CM) mice (Fig. 4.8A). As predicted, IR CM resulted in a reduction in the numbers of neurospheres formed relative to Ctrl CM (Fig. 4.8B), suggesting that the irradiated niche contains factors that reduce aNSC pool size. Importantly, Ctrl CM did not significantly affect neurosphere formation relative to normal neurosphere growth conditions [serum free media supplemented with EGF, FGF, and heparin (SFM+EFH)] alone, suggesting that these effects are specific to niche factors present in the irradiated SEZ.

Figure 4.8. Factors in the irradiated SEZ niche reduce aNSC pool size in vitro. (A) Schematic outlining the generation of conditioned media (CM) from non-irradiated (Ctrl CM) or irradiated (IR CM) mice used for culturing neurospheres. (B) Neurosphere formation of cells derived from the non-irradiated SEZ in the presence of Ctrl or IR CM relative to cells cultured in SFM+EFH (**p<0.01, repeated measures one-way ANOVA with Tukey’s test for multiple comparisons; n=5 experiments). Data is presented as mean ± SEM.

4.5. Effects of metformin pretreatment on SEZ neuroblasts following juvenile cranial IR While spontaneous recovery of NSCs occurs in the adult SEZ following cranial IR, long- term deficits to proliferation and neurogenesis persist in the SEZ and OB, respectively (Hellström et al., 2009; Lazarini et al., 2009; Achanta et al., 2012; Pineda et al., 2013). Given the finding that factors from the irradiated SEZ niche are sufficient to reduce neurosphere formation, it is likely that the irradiated SEZ niche may also perturb the behaviour of the progeny of NSCs, namely neuroblasts. We therefore asked if metformin pretreatment prevents or rescues IR-induced deficits to SEZ-derived neurogenesis in vivo. Mice were sacrificed at 2 or 30 days post-IR and the numbers of DCX+ neuroblasts and DCX+Ki67+ proliferating neuroblasts were assessed in coronal sections of the SEZ (Fig. 4.9A). We found a >95% depletion of DCX+ and DCX+Ki67+ neuroblasts at 2 days post-IR, which was not prevented by metformin pretreatment (Fig. 4.9B-D). However, by 30 days post-IR, the total number of DCX+Ki67+ proliferative neuroblasts was rescued with metformin pretreatment (Fig. 4.9E-G). While the total number of DCX+ neuroblasts remained reduced by >25% in irradiated mice 30 days post-IR, this trend was not statistically significant (p=0.17) by one-way ANOVA and Tukey’s test for multiple comparisons. Hence, this suggests that metformin pretreatment specifically enhances the recovery of neuroblast proliferation in the SEZ following cranial IR.

Figure 4.9. Metformin enhances the recovery of neuroblast proliferation in the SEZ following juvenile cranial IR. (A) Experimental timeline. (B, E) Representative images of the dorsolateral corner of the SEZ at (B) 2 and (E) 30 days post- IR (cyan, DAPI; green, Ki67; red, DCX). Scale bar = 100µm. (C-D) Quantification of (C) DCX+ and (D) DCX+Ki67+ neuroblasts in the SEZ at 2 days post-IR (**p<0.01, ****p<0.0001, one-way ANOVA with Tukey’s test for multiple comparisons; n≥3 animals per group). (F-G) Quantification of (F) DCX+ and (G) DCX+Ki67+ neuroblasts in the SEZ at 30 days post-IR (*p<0.05, one-way ANOVA with Tukey’s test for multiple comparisons; n=4 animals per group). Data is presented as mean ± SEM.

4.6. Metformin pretreatment rescues deficits to long-term olfactory memory following juvenile cranial IR. The most clinically relevant outcome of neural repair strategies is functional recovery. Herein we used a model of olfactory memory to determine whether the metformin-induced recovery of proliferative neuroblasts at 30 days post-IR was functionally relevant (Fig. 4.9E-G). We made the strong prediction that metformin pretreatment would improve long-term olfactory memory (LTOM), which has been shown to decline following adult cranial IR (Lazarini et al., 2009). To assess LTOM at 30 days post-metformin pretreatment and cranial IR, mice were trained to associate the odorant isoamyl acetate (IAA) with a sugar reward, and citralva with no reward, and were tested 1 month later to interrogate LTOM retention (Fig. 4.10A). Briefly, mice were acclimated to the reward and training apparatus from P41-43 prior to undergoing pre- training from P44-46 to learn to dig for the reward in a sand-filled dish. Training took place for 8 days x 8 trials/day from P47-54, where mice were presented with 2 sand-filled dishes scented with IAA (containing reward) or citralva (no reward) in a randomized location (left vs. right) and were allowed to dig in one dish per trial. Mice that correctly chose the reward-associated dish at an average rate of ≥80% over the final 2 training days passed the training phase and were tested 1 month later (P84), which involved a single block of 4 trials with no reward under either of the scented dishes (Fig. 410A). No significant differences between groups were observed during the final 2 training days, although cranially irradiated mice displayed deficits on the 6th day of training, potentially suggesting a delayed acquisition of the odor-reward association relative to non-irradiated controls (Fig. 410B). As predicted, juvenile cranial IR caused significant deficits to LTOM 1 month post-training, which was completely rescued by metformin pretreatment (Fig. 410C). This finding indicates that metformin pretreatment is able to promote functional SEZ-OB neurogenesis associated with LTOM following cranial IR.

Figure 4.10. Metformin pretreatment rescues deficits to LTOM following juvenile cranial IR. (A) Experimental timeline. (B) Quantification of % correct choice (characterized by digging in the IAA-scented dish) during the training phase in which mice learned to associate the odorant IAA with a sugar reward and citralva with no reward (*p<0.05, two-way ANOVA with Tukey’s test for multiple comparisons; n≥10 animals per group). Data is presented as mean ± SEM. (C) Quantification of % correct choice (characterized by digging in the IAA-scented dish) 1 month post-training with no reward in either dish (**p<0.01, two-way ANOVA with Tukey’s test for multiple comparisons; n≥6 animals per group). Data is presented as mean ± SEM.

5. Discussion

This study is the first to identify a prophylactic therapy that rescues deficits to neuroblast behaviour and promotes functional recovery following cranial IR. The findings presented here provide strong support for the consideration of metformin as a pro-neurogenic and neuroprotective drug. Neurogenesis involves several distinct cellular behaviours, including NSC activation, self-renewal, progenitor production and proliferation, differentiation, and migration. Furthermore, these processes are influenced and manipulated by numerous cell intrinsic and extrinsic processes, resulting in both spatial and temporal NPC heterogeneity (Chaker et al., 2016; Adams and Morshead, 2018). Fortunately, the neurosphere assay serves as a robust tool to assess these behaviours in vitro, and thus, was used extensively to assess the effects of metformin and cranial IR on NSCs. Despite the strengths of the neurosphere assay, it is important to consider the limitations of the assay (reviewed in Pastrana et al., 2011; Gil-Perotín et al., 2013). One such limitation is the potential persistence of niche factors in primary cultures, which may confound the ability to study cell autonomous (intrinsic) properties of NSCs. However, the neurosphere assay was used in this study to assess NSC pool size, irrespective of intrinsic vs. extrinsic effects, making this aspect of the neurosphere assay inconsequential. Although the passaging of neurospheres may eliminate niche components from primary cultures, assessing passaged neurospheres may obscure the effects of in vivo injuries on NSC pool size by selecting for viable or healthier cells upon passaging. Furthermore, neurospheres may also establish their own niche where the centre of a neurosphere receives fewer inputs from diffusible factors in media, producing heterogeneity within and between neurospheres (Bez et al., 2003). While these potentially confounding aspects of the neurosphere assay may impede its ability to reveal mechanisms involved in NPC pool expansion and intrinsic vs. extrinsic signaling, it remains an effective tool for measuring relative changes in NPC behaviours when NPCs are grown at clonal density (Coles-Takabe et al., 2008). Herein we used the neurosphere assay following metformin treatment in neonatal and juvenile mice to quantify aNSC pool size. Notably, qNSCs are excluded from this quantification, as they express low to negligible levels of EGFR and therefore fail to proliferate in the presence

81 of EGF, FGF, and heparin (collectively termed EFH), which are mitogens commonly used in neurosphere growth media (Coles-Takabe et al., 2008; Azari et al., 2010; Codega et al., 2014). Interestingly, we found that metformin expanded the aNSC pool of the SEZ in neonatal, but not juvenile mice (Fig. 4.1), which was initially concerning, given that NPCs from both neonatal and adult mice generate greater numbers of neurospheres when grown in the presence of metformin, suggesting that metformin directly acts on NSCs to enhance their self- renewal and proliferation (Dadwal et al., 2015; Fatt et al., 2015). However, we administered metformin in vivo prior to growing neurospheres in the absence of metformin in order to assess metformin’s in vivo effects on aNSC pool size, which has not previously been reported. While neurospheres grown in the presence of metformin in vitro allow for the elucidation of metformin’s direct effects on NSCs, this model is unable to account for metformin’s niche-mediated effects on NSCs, as well as normal physiological cues that may alter the response of NSCs to metformin. For example, spontaneous neuroblast depolarization in the SEZ results in GABA release that tonically activates GABAAR signaling in NSCs to maintain their quiescence (Liu et al., 2005). Therefore, if metformin alters neuroblast-mediated GABA release or GABAAR expression in NSCs in vivo, neurosphere formation following in vivo metformin administration will likely differ from neurosphere formation in the presence of metformin in vitro. Interestingly, it was recently reported that metformin activates the AMPK-

FOXO3 axis in hippocampal neurons to upregulate membrane insertion of GABAAR via increased transcription of GABAAR-associated protein (GABARAP) (Fan et al., 2019). Although it is unclear whether metformin induces this response in NSCs, FOXO3 is expressed in NSCs of both neurogenic niches and is required for the maintenance of quiescence and prevention of NPC depletion (Renault et al., 2009). Therefore, while NSCs from mice of varying ages respond to metformin in vitro to increase neurosphere formation, age-related changes to the neurogenic niche may modify or counteract the ability of in vivo metformin treatment to expand the aNSC pool, which may underlie the differences between neonatal and juvenile neurosphere formation in Figure 4.1. Alternatively, it has also been reported that EGF-responsive TAPs generate neurospheres, which if true, would signify that the quantification of neurospheres is

82 inappropriate for comparing relative aNSC pool size, since aNSC- and TAP-derived neurospheres may not be distinguishable (Doetsch et al., 2002). However, this finding has been refuted using transgenic GFAP-tk mice expressing thymidine kinase under the control of the GFAP promoter, which is expressed in NSCs and astrocytes, but not TAPs (Doetsch et al., 1999b; Doetsch et al., 2002). When ganciclovir (GCV) is administered to GFAP-tk mice, proliferating GFAP+ cells (i.e., NSCs) are selectively ablated, and 3 days of GCV administration is sufficient to reduce neurosphere formation by >95%, suggesting that the neurosphere assay provides a reliable estimate of aNSC pool size (Morshead et al., 2003). After selecting a 1-week window for metformin pretreatment (P9-15), we asked whether this metformin treatment causes a sustained expansion of the SEZ aNSC pool. Based on the finding that transient metformin exposure in primary neurospheres enhanced neurosphere formation upon multiple passages in the absence of metformin (Fatt et al., 2015), we predicted that 1 week of neonatal in vivo metformin treatment would also result in a sustained increase in aNSC pool size. Despite a slight trend of increased aNSC pool size 1 month after metformin treatment, the significant increase was lost within 10 days after treatment, suggesting that neonatal metformin treatment results in transient expansion of the aNSC pool that is lost in juvenile mice (Fig. 4.3). Again, this apparent discrepancy is likely due to our use of in vivo metformin treatment as opposed to in vitro metformin exposure, suggesting that factors in the SEZ niche either induce aNSCs to revert to quiescence, undergo apoptosis, or undergo consuming divisions to generate TAPs following metformin treatment (Obernier et al., 2018). However, apoptosis is unlikely given that metformin acts directly on NPCs to reduce apoptosis, as well as on other targets such as microglia and endothelial cells to reduce inflammation and oxidative stress (Hattori et al., 2006; Łabuzek et al., 2010a; Jin et al., 2014; Fatt et al. 2015). Furthermore, the absence of sustained aNSC pool expansion suggests that metformin’s effects are not neoplastic/tumorigenic. This is a crucial safety concern in the field of regenerative medicine, specifically in strategies aimed at promoting cell proliferation. Previous work has revealed that neurosphere formation no longer depends on the presence of mitogens following the fourth passage, and that repeated (>10 times) passaging results in significantly increased number and size of neurospheres (Morshead et al., 2002). The altered proliferation

83 kinetics of passaged NPCs therefore suggests the potential for neoplasm formation. Indeed, embryonic rat-derived NSCs transplanted into the transected thoracic spinal cord of adult rats have been reported to migrate and form ectopic colonies at long distances from their transplant site (Steward et al., 2014). Multifocal brain tumour formation in humans following human fetal NSC transplantation has also been reported, highlighting the risks of uncontrolled NSC proliferation in neurorepair strategies (Amariglio et al., 2009). However, the absence of aNSC pool expansion 10 days after metformin treatment (Fig. 4.3B) suggests that metformin’s in vivo effects are non-tumorigenic. Since the mechanisms regulating the metformin-induced neonatal aNSC pool expansion and its return to baseline in the juvenile mouse could not be elucidated using the neurosphere assay, multiple interpretations are possible. Specifically, it is unclear whether metformin recruits qNSCs to the aNSC pool or enhances the symmetrical self-renewal of previously activated NSCs. If metformin acts on aNSCs to enhance their symmetrical self-renewal, the inability of metformin to expand the juvenile aNSC pool may be due to an endogenous reduction in aNSCs from the neonatal to juvenile age. Indeed, a ~50% reduction of nestin+ cells in the SEZ of P23 juvenile rats relative to P9 neonatal rats has been reported (Fukuda et al., 2005). However, since nestin is also expressed in TAPs, it is unclear whether a reduction in aNSCs or TAPs underlies this age-associated reduction (Doetsch et al., 1997). Longitudinal studies of the ratio of qNSC to aNSCs throughout neonatal and juvenile development are therefore necessary to address this question. Given the finding that qNSCs fail to activate after 6 days in culture (Codega et al., 2014), FACS-mediated isolation of qNSCs would allow for their growth in the presence or absence of metformin, allowing for the elucidation of metformin’s mechanism of action on NSCs. It was recently proposed that metformin requires TAp73 to promote self-renewal and proliferation, measured via neurosphere number and size, respectively (Fatt et al., 2015). While this finding contributes to the understanding of cell signaling pathways in NSCs, it does not explicitly identify whether TAp73 is expressed in qNSCs, aNSCs, or both. Genetic ablation of TAp73 (TAp73-/-) results in reduced primary and passaged neurosphere formation from the adult SEZ, even when cultured in the presence of metformin. However, the finding that

84 metformin increases passaged neurosphere formation in WT mice suggests that metformin acts specifically on aNSCs, given the assumption that passaging neurospheres selects for aNSCs and their progeny, but not qNSCs (Codega et al., 2014; Fatt et al., 2015). Alongside our observation that metformin was insufficient to increase the ratio of aNSCs to qNSCs (Fig. 4.7D), these data suggest that metformin does not directly promote the activation of qNSCs, but instead promotes the symmetrical self-renewal of aNSCs. As previously described, FOXO3 is also required for long-term NPC maintenance since FOXO3-/- mice give rise to reduced primary and passaged neurospheres, suggesting that both TAp73 and FOXO3 are required for the maintenance of NSC self-renewal capacity (Renault et al., 2009). However, inducing self-renewal is distinct from maintaining self-renewal capacity, which may rely on the capacity of the NSC pool to maintain a reserve populations of qNSCs. Therefore, it is difficult to conclude that metformin specifically acts on TAp73 to enhance adult NPC self-renewal and proliferation, since it is plausible that the NPC pool is depleted or debilitated in TAp73-/- mice prior to adulthood (i.e., during development), resulting in an inability of TAp73-/- adult-derived NPCs to respond to metformin in vitro. This same logic applies to FOXO3-/- mice, although metformin’s capacity to enhance neurosphere formation from FOXO3-/- mice has not yet been established. Furthermore, TAp73 is required for long-term NPC maintenance via regulation of the transcription factor Hey2, which prevents premature differentiation (Fujitani et al., 2010). Therefore, it is possible that the premature lineage progression or differentiation of NPCs results in NSC pool depletion, which underlies the inability of metformin to enhance neurosphere formation in TAp73-/- mice. Conditional knockout or artificial upregulation of TAp73 or FOXO3 during metformin exposure may therefore provide greater insight into whether metformin acts on either of these proteins to promote self-renewal and proliferation. To identify whether metformin affects the ratio of aNSCs to qNSCs in vivo, we used FACS to quantify the percentage of NSCs that were activated (GFAP::GFP+CD133+EGFR+) or quiescent (GFAP::GFP+CD133+EGFR-) on P19, following P9-15 metformin pretreatment and cranial IR on P17 (Fig. 4.7). In control mice receiving no metformin or cranial IR, ~60% of SEZ-derived NSCs were activated, which was greater than the previously reported level of activation (~40%) in

85 adults (Codega et al., 2014; Llorens-Bobadilla et al., 2015). This aligns with our predictions, given that NSCs gradually become more quiescent in vivo with age (Luo et al., 2006; Bouab et al., 2011; Giachino et al., 2014; Piccin et al., 2014; Silva-Vargas et al., 2016; Kalamakis et al., 2019). The finding that P9-15 metformin treatment did not significantly alter the ratio of qNSCs to aNSCs in vivo (Fig. 4.7D) was initially surprising and seemingly conflicts with our observation that P9-15 metformin treatment increases neurosphere formation at P19 (Fig. 4.3B). However, this can be reconciled by previous reports that qNSCs enter a primed quiescent state characterized by reduced GLAST expression and increased protein synthesis prior to becoming activated and upregulating EGFR (Llorens-Bobadilla et al., 2015). Therefore, it is possible that metformin primed qNSCs for activation, which would not be detectable using our labeling strategy for FACS (Fig. 4.7), although these qNSCs may be more likely to activate and form neurospheres in the presence of mitogens (as in Fig. 4.3B). Alternatively, it is possible that an increased proportion of aNSCs relative to qNSCs may induce a negative feedback mechanism in which aNSCs spontaneously revert to quiescence in order to conserve energy or prevent pool depletion. It is also possible that the expansion of aNSCs results in premature lineage progression, which may be detectable as an increase in TAP or neuroblast formation. Furthermore, we predicted an overwhelming increase in the proportion of qNSCs following IR, since cranial IR has previously been shown to deplete proliferating cells in the SEZ, which are predominantly comprised of aNSCs, TAPs, and neuroblasts (Fukuda et al., 2005; Daynac et al., 2013). Despite observing a slight trend suggestive of a shift to quiescence, this was not significantly different than non-irradiated controls 2 days after cranial IR (~50%) using FACS (Fig. 4.7D), which may conflict with our finding that cranial IR significantly reduces neurosphere formation 2 days post-IR (Fig. 4.4B, 4.5B). However, the recovery or maintenance of normal qNSC to aNSC ratio following cranial IR does not ensure the survival or normal proliferation/self-renewal of aNSCs, given the persistent inflammatory environment of the SEZ and our finding that factors in the irradiated niche contribute to reduced neurosphere formation (Fig. 4.8B). Therefore, our observation of reduced primary neurosphere formation 2 days post-IR is compatible with a consistent ratio of aNSCs to qNSCs, given that the niche

86 factors present during primary neurosphere growth may influence NSC survival and neurosphere formation without reducing the ratio of aNSCs to qNSCs. In contrast to our finding of unaltered qNSC to aNSC ratio 2 days post-IR, Daynac et al. used FACS to identify LeX+EGFR- qNSCs and LeX+EGFR+ aNSCs, and found that qNSCs were upregulated and that aNSCs were reduced by >50% in the adult SEZ 2 days after 4 Gy IR, while aNSCs recovered 3 days post-IR and significantly surpassed control levels thereafter (Daynac et al., 2013). This qNSC-mediated recovery of aNSCs was partially attributed to the loss of neuroblasts following cranial IR, which are normally a source of GABA signaling-mediated quiescence (Daynac et al., 2013). However, the dynamics of NSC activation following cranial IR or other CNS injuries in the juvenile mouse likely differ from adults and have yet to be investigated. The lack of aNSC depletion 2 days post-IR (Fig. 4.7) in the juvenile mouse may be due to an accelerated compensatory response. In accordance with neuroblast-mediated control of NSC quiescence (Liu et al., 2005; Daynac et al., 2013), we observed a >95% depletion of neuroblasts 2 days post-IR (Fig. 4.9C), which may underlie the lack of qNSC expansion relative to aNSCs, due qNSC-derived repopulation of aNSCs. Given that neurogenesis declines with age (Capilla- Gonzalez et al., 2014), it is also possible that the juvenile SEZ promotes enhanced or accelerated conversion of qNSCs to aNSCs relative to the adult SEZ. However, it is unclear if such a heightened response is beneficial, as the over-activation of NSCs may eventually lead to the premature depletion of the NSC pool (Imayoshi et al., 2010; Encinas et al., 2011). Therefore, it will be valuable to investigate the activation kinetics of NSCs at earlier times following cranial IR (6, 12, and 24 hours post-IR), and whether repeated doses of cranial IR (i.e., fractionated doses) in the juvenile mouse will cause repeated ablation of aNSCs and activation of qNSCs, resulting in depletion of the NSC pool. It is also necessary to note that not all NSCs express CD133 (used in the present study) or LeX (used in Daynac et al., 2013) in vivo, and their colocalization has not been reported, suggesting that they label distinct subpopulations of NSCs (Coskun et al., 2008; Mirzadeh et al., 2008; Sun et al., 2009). Furthermore, the specificity of LeX has been challenged in several studies, which report LeX expression by hippocampal neurons, astrocytes, OPCs, cortical

87 neurons, and cells in the corpus callosum and striatum (reviewed in Hennen and Faissner, 2012). Accordingly, in cells isolated from the SEZ via microdissection and dissociation, acute staining revealed that as low as 18% of LeX+ cells were also GFAP+, suggesting that LeX is not a specific marker of NSCs (Capela and Temple, 2002). Therefore, the relative IR-induced increase in the proportion of qNSCs may have been overestimated in previous reports (Daynac et al., 2013). Similarly, we may be selecting for a specific subpopulation of NSCs by using CD133, resulting in potential discrepancies between studies, and between assays used to quantify aNSCs, such as the neurosphere assay. Therefore, a combination of techniques and labeling strategies at multiple time points may be necessary to accurately identify NSC activation dynamics following injury or metformin treatment. Since metformin pretreatment did not prevent the acute depletion of neurosphere- forming aNSCs 2 days post-IR (Fig. 4.5B), it can also be interpreted that NSCs reverted to quiescence following metformin treatment and cranial IR, or that IR-mediated apoptosis was not prevented by metformin. The latter is more likely given the proliferative nature of aNSCs, resulting in their increased susceptibility to IR-induced DNA damage and apoptosis (Bellinzona et al., 1996; Shinohara et al., 1997; Chow et al., 2000), as well as our observation that P9-15 metformin treatment did indeed expand the neurosphere-forming aNSC pool at P19 (Fig. 4.3B). Conversely, since the metformin-mediated expansion of neurosphere-forming aNSCs was maintained until P19, it can be reasoned that this increase in proliferation may actually increase DNA damage-mediated apoptosis following P17 cranial IR, since cells in S-phase are more likely to experience DNA DSBs (Vignard et al., 2013). Notwithstanding the inability of metformin pretreatment to prevent the acute IR-induced depletion of the aNSC pool, metformin pretreatment may still protect the niche against secondary damage induced by acute and chronic inflammation (Hattori et al., 2006; Łabuzek et al., 2010a; Jin et al., 2014; reviewed in Saisho, 2015). Despite an initial depletion of neurosphere-forming aNSCs, the neurosphere-forming aNSC pool partially recovers 8 days post-IR and fully recovers within 1 month following 8 Gy cranial IR at P17 (Fig. 4.4), which is consistent with previous reports of NSC recovery typically occurring weeks to months post-IR, with variability attributable to IR dose and age at IR

(Hellström et al., 2009; Achanta et al., 2012; Daynac et al., 2013). However, deficits to proliferation in the SEZ persist, even when NSCs recover to control levels (Hellström et al., 2009; Pineda et al., 2013), warranting an investigation of downstream neurogenic processes. Despite our observation that the SEZ aNSC pool spontaneously recovers within 1 month post-IR (Fig.4.5), others have reported that the irradiated NSC pool surpasses control levels 9 weeks post-IR, which may signify a compensatory expansion to offset downstream deficits to proliferation and OB neurogenesis (Hellström et al., 2009). Although P9-15 metformin treatment alone did not result in an expanded aNSC pool at P25 or P47 (Fig. 4.3), we considered the possibility that metformin pretreatment would enhance spontaneous recovery following cranial IR by minimizing perturbations to the niche. Metformin’s anti-inflammatory, pro- neurogenic, and angiogenic effects may prevent damage to the niche caused by activated microglia (Łabuzek et al., 2010a; Jin et al., 2014) and secreted factors by endothelial cells (Hattori et al., 2006). Therefore, we predicted that metformin would prime the niche to support accelerated recovery of neurosphere-forming aNSCs following cranial IR, since NSC activation is strongly mediated by extrinsic factors (reviewed in Adams and Morshead, 2018; Ruddy and Morshead, 2018). However, recovery of the neurosphere-forming aNSC pool was not significantly enhanced by metformin at 8 (P25) or 30 (P47) days post-IR (Fig. 4.6B-C), which may signify that metformin pretreatment is insufficient to enhance the intrinsic or extrinsic mechanisms promoting NSC self-renewal. However, we did not assess the aNSC pool size at time points later than 1 month post-IR, so we cannot rule out the possibility that the NSC pool continues to expand at later times, as reported 9 weeks post-IR (Hellström et al., 2009). Furthermore, long-term over-expansion of the NSC pool may not necessary be beneficial, given that OB neurogenesis remains impaired in other models, concomitant with long-term deficits to neurogenesis-dependent olfactory behaviour (Hellström et al., 2009; Lazarini et al., 2009). Instead, over-expansion of the NSC pool may signify deficits to either intrinsic NSC maintenance (Renault et al., 2009; Fujitani et al., 2010; Webb et al., 2013) or extrinsic feedback mechanisms (Liu et al., 2005; Daynac et al., 2013; Ramírez-Castillejo et al., 2006) that prevent long-term NSC exhaustion. Indeed, other studies have reported NSC over- expansion concurrent with deficits to neuroblasts following cranial IR (Daynac et al., 2013;

Pineda et al., 2013). This may result in aberrant NSC activation due to a lack of neuroblast- mediated GABAAR signaling in NSCs that normally promotes quiescence to avoid exhaustion of the NSC pool (Liu et al., 2005). Interestingly, when using a relatively low 4 Gy dose of cranial IR in adult mice, FACS revealed aNSC over-expansion and neuroblast depletion 4 days post-IR, with both cell types returning to control levels 10 days post-IR (Daynac et al., 2013). However, when P9 rats received a greater dose of 6 Gy cranial IR, aNSC pool size was >1.5-fold larger 9 weeks post-IR (Hellström et al., 2009), while 8 Gy cranial IR at P9 resulted in significant reductions in neuroblasts 10 weeks post-IR (Fukuda et al., 2005), suggesting a strong correlation between neuroblast depletion and aNSC overexpansion that is potentially exacerbated with younger age and increased dose of IR. Indeed, a more pronounced apoptotic response and long-term reduction of neuroblasts was reported when cranial IR was delivered at P9 than at P23 (Fukuda et al., 2005). Interestingly, the severity and onset of cognitive deficits in human patients is negatively correlated with the age at which they receive cranial IR (Chin and Maruyama, 1984; Spiegler et al., 2004; Tonning Olsson et al., 2014). Therefore, an absence of aNSC over-expansion following cranial IR may in fact be a sign of enhanced regulation of NSC kinetics and intact homeostasis in the niche. In accordance with this assumption, metformin pretreatment rescued neuroblast proliferation 1 month post-IR (Fig. 4.9), indicating that metformin may have protective effects on the neurogenic niche, thereby supporting normal neuroblast function. Importantly, our finding that metformin rescues neuroblast proliferation following cranial IR is strongly supported by other studies reporting similar findings. Two weeks of metformin administration in young adult Zucker diabetic fatty rats rescued deficits to proliferating (Ki67+) cells and DCX+ neuroblasts in the SGZ (Hwang et al., 2010), which is hypothesized to be BDNF-dependent (Yoo et al., 2011). More recently, 1 week of metformin administration immediately following stroke in the rat brain was also shown to promote increased neuroblast proliferation in the SGZ (Yuan et al., 2019), highlighting metformin as an effective pharmacological agent to rescue neurogenesis following various insults to the CNS. However, it will also be important to investigate the downstream

90 effects of enhanced neuroblast proliferation, such as neuroblast migration and neuronal survival and maturation. Although neuroblast migration was not assessed in the present study, metformin’s anti- inflammatory and angiogenic effects support the prediction that metformin pretreatment would enhance neuroblast migration through the RMS. When the RMS is focally irradiated, leaving the SEZ intact, neuroblasts are unperturbed in the SEZ, although they fail to migrate through the RMS, supporting the hypothesis that the irradiated niche impairs normal neuroblast behaviour (Achanta et al., 2012). Metformin reduces inflammatory microglial activation and ROS/RNS production, as well as endothelial NF-κB-dependent transcription of pro-inflammatory genes (Hattori et al., 2006; Łabuzek et al., 2010a). Following stroke, metformin treatment also promotes anti-inflammatory microglial activation and eNOS production, resulting in angiogenesis, BDNF-mediated NPC proliferation and migration, and functional recovery (Chen et al., 2005; Jin et al., 2014; Liu et al., 2014). Interestingly, the pharmacological ablation of microglia following 9 Gy cranial IR in adult mice has been shown to improve cognitive function related to medial prefrontal cortex and hippocampal learning and memory 5-6 weeks post-IR, suggesting a link between IR-induced inflammation and cognitive deficits (Acharya et al., 2016). Possibly our most compelling evidence for metformin’s neuroprotective effects against cranial IR was the observation that metformin pretreatment completely rescued deficits to LTOM. Deficits to LTOM can either be attributed to an inability to consolidate a new memory or recall an established memory. It has been reported that adult-born neuron maturation and survival in the OB is positively correlated with repeated activation of newborn neurons, which occurs during the acquisition of olfactory memory (Lemasson et al., 2005; Magavi et al., 2005; Moreno et al., 2009). Therefore, it is probable that OB neurogenesis is required for memory consolidation as opposed to recall. Since we did not investigate cellular recovery after P47 (the time at which LTOM training began), it cannot be concluded whether deficits to neurogenesis persisted until the LTOM retention test at P84 (1 month after the final LTOM training day). Therefore, it cannot be confirmed whether IR-induced deficits to neurogenesis contribute to

91 impaired consolidation or recall. This can, however, be determined by delivering cranial IR after acquiring the odor-reward association, leaving neurogenesis intact during consolidation.

6. Conclusions

Using the neurosphere assay, we determined that in vivo metformin administration (200mg/kg/day) expands the aNSC pool in the SEZ of neonatal, but not juvenile mice, which is consistent with our lab’s work demonstrating age-specific effects of metformin on NPCs (Ruddy et al., 2019a). This expansion was not sex-dependent, which strengthens metformin’s potential for clinical use. Furthermore, we found that 1 week of neonatal metformin administrated promoted a transient expansion of the aNSC pool that returned to baseline levels after 10 days, indicating that metformin’s in vivo effects are non-tumorigenic in the neonatal/juvenile mouse brain. Taken together, these findings support metformin’s candidacy as a clinically relevant drug for promoting neurogenesis. We next sought to investigate whether metformin’s pro-neurogenic effects were sufficient to prevent deficits to aNSCs following cranial IR. Similar to previous studies in adult rodents (Achanta et al., 2012; Pineda et al., 2013), we found that a clinically relevant dose (8 Gy) of cranial IR in juvenile P17 mice induces an acute depletion aNSCs and neuroblasts in the SEZ, measured via the in vitro neurosphere assay and in vivo labeling of neuroblasts, respectively. While the aNSC pool spontaneously recovers, deficits to neuroblast proliferation persist 1 month post-IR. This indicates that the downstream progeny of NSCs fail to regain normal function in the irradiated niche despite recovery of NSCs, highlighting the necessity to protect or enrich the neurogenic niche in order to restore neurogenesis. Metformin pretreatment from P9-15 does not significantly modify the spontaneous recovery of aNSCs following cranial IR, but promotes a complete rescue of neuroblast proliferation. This novel finding suggests that metformin pretreatment protects the SEZ niche from cranial IR in order to rescue deficits to neurogenesis. However, further investigation is necessary to elucidate the effects of metformin pretreatment on the SEZ niche following cranial IR. We were unable to conclude whether metformin recruits qNSCs to the activated pool for multiple reasons. Firstly, we used FACS 4 days following metformin treatment, which may have resulted in transient metformin-induced activation and subsequent reversion to quiescence, or lineage progression of NSCs. Secondly, our labeling strategy may have identified a subpopulation of NSCs, resulting in the possibility of sampling error. Lastly, we did not assess

93 the potential for metformin to induce a primed quiescent state, thereby precluding our ability to conclude whether metformin alters the activation state of NSCs. Cranial IR did not significantly alter NSC activation state 2 days post-IR, although we cannot exclude the possibility that aNSC depletion and subsequent activation of qNSCs occurs within this 2 day window without assessing NSC activation at earlier times post-IR. Due to previous reports of increased inflammation in the neurogenic niches following cranial IR (Monje et al., 2002; Kalm et al., 2009; Han et al., 2016), we assessed whether the irradiated niche was sufficient to induce deficits to aNSCs. Using conditioned media from the irradiated SEZ, we found that factors in the irradiated niche were sufficient to reduce neurosphere formation. While previous reports have identified that the irradiated neurogenic niche inhibits the survival, proliferation, migration, and differentiation of healthy NPCs (Monje et al., 2002; Achanta et al., 2012; Pineda et al., 2013), this study is the first to identify that the secreted factors in the irradiated SEZ induce deficits to aNSC self-renewal. Lastly, we confirmed that juvenile cranial IR induces deficits to LTOM, as it does in adult models of cranial IR (Lazarini et al., 2009). While cranially irradiated mice are able to establish an odor-reward association 30 days post-IR, they are unable to retain LTOM 1 month later. However, metformin pretreatment completely rescues the cranial IR-induced deficit to LTOM. These findings are the first to provide strong evidence in support of the hypothesis that metformin pretreatment enhances SEZ neurogenesis and promotes functional recovery in a clinically relevant rodent model of juvenile cranial IR. This study primarily identifies metformin as a potential prophylactic agent against cranial IR, resulting in cellular and functional recovery that may translate to attenuated cognitive deficits in human patients. However, the significance of this study extends beyond the rescue of long-term deficits induced by cranial IR, since intact neurogenesis is required for endogenous repair following certain brain injuries. For example, stroke induces NPC activation, where SEZ-derived cells contribute to neurogenesis and glial scar formation (reviewed in Chaker et al., 2016). Therefore, protecting the SEZ niche and rescuing neurogenesis following cranial IR may not only ameliorate IR-induced cognitive deficits, but may also contribute to the life-long preservation of SEZ-mediated neurorepair capacity.

7. Future Directions

While we were able to show that metformin pretreatment promotes functional SEZ neurogenesis following cranial IR, several questions remain pertaining to metformin’s mechanisms of action. While previous reports suggest that metformin acts on TAp73 to induce self-renewal and proliferation of adult-derived NPCs, it is unclear whether TAp73 deficiency specifically prevents metformin from expanding the NSC pool, or if TAp73 deficiency causes NPC deficits during development that thereby negate metformin’s effects on adult NSCs (Fujitani et al., 2010; Fatt et al., 2015). Since the effects of TAp73 deficiency on self-renewal capacity mirror those of FOXO3 deficiency (Renault et al., 2009), it will be valuable to transiently inhibit or upregulate TAp73 or FOXO3 in the presence of metformin to identify whether these proteins are involved in metformin-mediated expansion of aNSCs. Furthermore, it remains unknown whether metformin recruits qNSCs to the activated pool, or if metformin enhances the symmetrical self-renewal of already activated aNSCs. Therefore, FACS-purified qNSCs or aNSCs should be cultured in the presence of metformin to identify if metformin can promote activation of qNSCs, enhanced symmetrical self-renewal of aNSCs, or both. This study supports the hypothesis that the irradiated SEZ niche contributes to long- term deficits to neurogenesis. However, the specific factors in the niche that contribute to reduced aNSC pool size are unknown, as previous studies have not established any causal relationships or have failed to distinguish between NSCs and TAPs. To identify secreted factors implicated in reduced neurogenesis, secretome analysis of the irradiated SEZ with or without metformin pretreatment will contribute to the identification of specific factors that may be down- or upregulated following metformin pretreatment and cranial IR. Following this analysis, the addition of specific IR-induced secreted factors to neurosphere growth media will allow for a causal link to be drawn between IR-induced niche perturbation and reduced neurogenesis. Understanding which factors play a significant role in damage or recovery will allow for the development of more targeted therapeutic interventions, while minimizing any potential side- effects. It is also unclear whether metformin-induced neurogenesis and functional recovery are mediated by changes to microglial activation or proliferation. It has previously been shown that

95 inflammatory microglia in other brain injury models produce ROS and RNS that impair neurogenesis following injuries to the SEZ (L’Episcopo et al., 2012). Since cranial IR induces microglial proliferation and persistent inflammation (Hattori et al., 2006; Han et al., 2016), quantification of inflammatory and proliferative microglia, as well as microglial ROS/RNS production following metformin pretreatment and cranial IR will reveal whether metformin’s therapeutic effects depend on its ability to reduce inflammation. Despite our finding that metformin restores neuroblast proliferation following cranial IR, we did not identify whether OB neurogenesis was also restored. Based on previous findings of persistent deficits to OB neurogenesis following adult cranial IR (Hellström et al., 2009; Lazarini et al., 2009), we would predict to observe reduced OB neurogenesis following juvenile cranial IR. However, given that metformin pretreatment rescued neuroblast proliferation and LTOM, we would also predict that this pretreatment paradigm would rescue deficits to OB neurogenesis. This could be determined via quantification of EdU+NeuN+ newborn neurons in the OB following EdU injections. If a correlation between rescued OB neurogenesis and LTOM is found, we could subsequently ablate neurogenesis using GFAP-tk or DCX-tk mice and GCV administration prior to and during LTOM training in order to identify a causal relationship between rescued neurogenesis and functional recovery. While we and others have shown neurogenesis-associated deficits to LTOM following cranial IR (Lazarini et al., 2009), it is still unclear whether these deficits pertain to memory consolidation or recall. To test whether IR-induced deficits to neurogenesis cause deficits to LTOM consolidation or recall, cranial IR should be delivered after acquiring the odor-reward association, prior to testing LTOM retention, which would thereby leave neurogenesis intact during consolidation. We would predict that delivery of cranial IR after LTOM consolidation would not induce deficits to LTOM, since previous studies have reported that increased neurogenesis at the time of olfactory training coincides with olfactory discrimination and memory formation (Lemasson et al., 2005; Moreno et al., 2015). While most studies identify acute deficits to neurogenesis following cranial IR, the persistence and severity of deficits varies significantly between studies. While the age at which mice receive cranial IR and the dose of IR have previously been identified as factors mediating

96 the severity of neurogenic deficits, it has yet to be identified whether a single dose or fractionated doses of cranial IR result in significantly different outcomes. Repeated IR may result in exacerbated deficits to both NSCs and their niche, due to the potential for IR to repeatedly deplete proliferating cells. If a single dose of cranial IR depletes proliferating cells, which are repopulated by qNSCs, multiple doses of cranial IR may result in repeated cycles of activation and depletion. Indeed, studies involving 3 fractions of 5 Gy IR separated by 48 hour intervals (totaling 15 Gy) typically report permanent deficits to neurogenesis and LTOM (Lazarini et al., 2009; Pineda et al., 2013). However, it is unclear whether the severity of these deficits is associated with an increased total dose of IR, or rather the repeated depletion of proliferating cells. Since this IR paradigm creates the potential for accelerated depletion of the postnatal NSC pool, it will valuable to compare the effects of single dose and fractionated cranial IR, with total dose remaining equivalent between irradiated groups. Metformin treatment prior to and during the course of fractionated cranial IR should be investigated to assess if this treatment paradigm is also sufficient to prevent neurogenesis-associated cellular and cognitive deficits. It has previously been reported that SEZ NSCs and proliferating cells fully and partially recover, respectively, while DG NSCs and proliferating cells remain depleted 9 weeks post-IR (Hellström et al., 2009). While the present study replicates this finding in the juvenile SEZ (full recovery of aNSCs, but not proliferating neuroblasts) and also identifies metformin as sufficient to rescue proliferating neuroblasts, it will be interesting and clinically relevant to repeat this investigation in the DG. Since spontaneous recovery of NSC pool size is not a guarantee of rescued neurogenesis, DG neuroblasts and hippocampal neurogenesis-dependent behaviours should be assessed following metformin pretreatment and cranial IR. However, based on our findings of metformin-induced recovery in the SEZ, as well as the finding that metformin enhances hippocampal-dependent spatial memory (Wang et al., 2012), we would predict that metformin pretreatment would rescue long-term IR-induced deficits to neuroblasts in the DG and hippocampal-dependent spatial memory. Lastly, it will be valuable to assess the effects of combined metformin pre- and post- treatment in relation to cranial IR, since NPCs may be more responsive to metformin’s pro-

97 neurogenic effects following cranial IR if pretreatment protects the niche and supports normal NPC function. Combined metformin pre- and post-treatment may have additive effects on niche protection and cellular recovery, which would provide additional proof of principle that the SEZ response to neurogenic stimuli is maintained. This would further advance metformin’s candidacy as a clinically relevant neuroprotective drug since certain brain injuries including stroke rely on SEZ NPCs for endogenous repair responses (Arvidsson et al., 2002; Zhang et al., 2004a; Faiz et al., 2015; Chaker et al., 2016). Therefore, brain injuries that typically induce a neurogenic response such as stroke should be delivered in adult mice that received metformin treatment and juvenile cranial IR. This will allow us to investigate whether long-term neurogenic deficits induced by cranial IR impair the SEZ neurogenic response to other injuries later in life, and whether metformin treatment protects or rescues the integrity of the endogenous neurogenic response.

References

Achanta, P., Capilla-Gonzalez, V., Purger, D., Reyes, J., Sailor, K., Song, H., Garcia-Verdugo, J.M., Gonzalez-Perez, O., Ford, E., and Quinones-Hinojosa, A. (2012). Subventricular Zone Localized Irradiation Affects the Generation of Proliferating Neural Precursor Cells and the Migration of Neuroblasts. STEM CELLS 30, 2548–2560. Acharya, M.M., Green, K.N., Allen, B.D., Najafi, A.R., Syage, A., Minasyan, H., Le, M.T., Kawashita, T., Giedzinski, E., Parihar, V.K., et al. (2016). Elimination of microglia improves cognitive function following cranial irradiation. Scientific Reports 6, 31545. Adams, K.V., and Morshead, C.M. (2018). Neural stem cell heterogeneity in the mammalian forebrain. Progress in Neurobiology 170, 2–36. Ahlenius, H., Visan, V., Kokaia, M., Lindvall, O., and Kokaia, Z. (2009). Neural stem and progenitor cells retain their potential for proliferation and differentiation into functional neurons despite lower number in aged brain. J. Neurosci. 29, 4408–4419. Ahmed, S., Reynolds, B.A., and Weiss, S. (1995). BDNF enhances the differentiation but not the survival of CNS stem cell- derived neuronal precursors. J. Neurosci. 15, 5765–5778. Alonso, M., Viollet, C., Gabellec, M.-M., Meas-Yedid, V., Olivo-Marin, J.-C., and Lledo, P.-M. (2006). Olfactory discrimination learning increases the survival of adult-born neurons in the olfactory bulb. J. Neurosci. 26, 10508–10513. Altman, J. (1963). Autoradiographic investigation of cell proliferation in the brains of rats and cats. The Anatomical Record 145, 573–591. Altman, J. (1969). Autoradiographic and histological studies of postnatal neurogenesis. IV. Cell proliferation and migration in the anterior forebrain, with special reference to persisting neurogenesis in the olfactory bulb. Journal of Comparative Neurology 137, 433–457. Altman, J., and Das, G.D. (1965). Autoradiographic and histological evidence of postnatal hippocampal neurogenesis in rats. Journal of Comparative Neurology 124, 319–335. Amariglio, N., Hirshberg, A., Scheithauer, B.W., Cohen, Y., Loewenthal, R., Trakhtenbrot, L., Paz, N., Koren-Michowitz, M., Waldman, D., Leider-Trejo, L., et al. (2009). Donor-derived brain tumor following neural stem cell transplantation in an ataxia telangiectasia patient. PLoS Medicine 6, e1000029. Antanitus, D.S., Choi, B.H., and Lapham, L.W. (1976). The demonstration of glial fibrillary acidic protein in the cerebrum of the human fetus by indirect immunofluorescence. Brain Research 103, 613–616. Arvidsson, A., Collin, T., Kirik, D., Kokaia, Z., and Lindvall, O. (2002). Neuronal replacement from endogenous precursors in the adult brain after stroke. Nature Medicine 8, 963–970. Azari, H., Rahman, M., Sharififar, S., and Reynolds, B.A. (2010). Isolation and expansion of the adult mouse neural stem cells using the neurosphere assay. J Vis Exp.

Azim, K., and Butt, A.M. (2011). GSK3β negatively regulates oligodendrocyte differentiation and myelination in vivo. Glia 59, 540–553. Azim, K., Fiorelli, R., Zweifel, S., Hurtado-Chong, A., Yoshikawa, K., Slomianka, L., and Raineteau, O. (2012a). 3-dimensional examination of the adult mouse subventricular zone reveals lineage-specific microdomains. PLoS ONE 7, e49087. Azim, K., Fischer, B., Hurtado‐Chong, A., Draganova, K., Cantù, C., Zemke, M., Sommer, L., Butt, A., and Raineteau, O. (2014a). Persistent Wnt/β-Catenin Signaling Determines Dorsalization of the Postnatal Subventricular Zone and Neural Stem Cell Specification into Oligodendrocytes and Glutamatergic Neurons. STEM CELLS 32, 1301–1312. Azim, K., Raineteau, O., and Butt, A.M. (2012b). Intraventricular injection of FGF‐2 promotes generation of oligodendrocyte‐lineage cells in the postnatal and adult forebrain. Glia 60, 1977–1990. Azim, K., Rivera, A., Raineteau, O., and Butt, A.M. (2014b). GSK3β regulates oligodendrogenesis in the dorsal microdomain of the subventricular zone via Wnt‐β‐catenin signaling. Glia 62, 778–789. Bachstetter, A.D., Morganti, J.M., Jernberg, J., Schlunk, A., Mitchell, S.H., Brewster, K.W., Hudson, C.E., Cole, M.J., Harrison, J.K., Bickford, P.C., et al. (2011). Fractalkine and CX3CR1 regulate hippocampal neurogenesis in adult and aged rats. Neurobiology of Aging 32, 2030–2044. Baker, S.A., Baker, K.A., and Hagg, T. (2004). Dopaminergic nigrostriatal projections regulate neural precursor proliferation in the adult mouse subventricular zone. European Journal of Neuroscience 20, 575–579. Banasr, M., Hery, M., Printemps, R., and Daszuta, A. (2004). Serotonin-Induced Increases in Adult Cell Proliferation and Neurogenesis are Mediated Through Different and Common 5-HT Receptor Subtypes in the Dentate Gyrus and the Subventricular Zone. Neuropsychopharmacology 29, 450–460. Baser, A., Skabkin, M., and Martin-Villalba, A. (2017). Neural Stem Cell Activation and the Role of Protein Synthesis. Brain Plast 3, 27–41. Battista, D., and Rutishauser, U. (2010). Removal of Polysialic Acid Triggers Dispersion of Subventricularly Derived Neuroblasts into Surrounding CNS Tissues. J. Neurosci. 30, 3995–4003. Beckervordersandforth, R., Tripathi, P., Ninkovic, J., Bayam, E., Lepier, A., Stempfhuber, B., Kirchhoff, F., Hirrlinger, J., Haslinger, A., Lie, D.C., et al. (2010). In Vivo Fate Mapping and Expression Analysis Reveals Molecular Hallmarks of Prospectively Isolated Adult Neural Stem Cells. Cell Stem Cell 7, 744–758. Belarbi, K., Jopson, T., Arellano, C., Fike, J.R., and Rosi, S. (2013). CCR2 deficiency prevents neuronal dysfunction and cognitive impairments induced by cranial irradiation. Cancer Res 73, 1201–1210.

100

Bellinzona, M., Gobbel, G.T., Shinohara, C., and Fike, J.R. (1996). Apoptosis is induced in the subependyma of young adult rats by ionizing irradiation. Neurosci. Lett. 208, 163–166. Berg, D.A., Belnoue, L., Song, H., and Simon, A. (2013). Neurotransmitter-mediated control of neurogenesis in the adult vertebrate brain. Development 140, 2548–2561. Betarbet, R., Zigova, T., Bakay, R.A., and Luskin, M.B. (1996). Dopaminergic and GABAergic interneurons of the olfactory bulb are derived from the neonatal subventricular zone. Int. J. Dev. Neurosci. 14, 921–930. Bez, A., Corsini, E., Curti, D., Biggiogera, M., Colombo, A., Nicosia, R.F., Pagano, S.F., and Parati, E.A. (2003). Neurosphere and neurosphere-forming cells: morphological and ultrastructural characterization. Brain Res. 993, 18–29. Bignami, A., and Dahl, D. (1974). Astrocyte-specific protein and radial glia in the cerebral cortex of newborn rat. Nature 252, 55. Bockaert, J., and Marin, P. (2015). mTOR in Brain Physiology and Pathologies. Physiological Reviews 95, 1157–1187. Boldrini, M., Fulmore, C.A., Tartt, A.N., Simeon, L.R., Pavlova, I., Poposka, V., Rosoklija, G.B., Stankov, A., Arango, V., Dwork, A.J., et al. (2018). Human Hippocampal Neurogenesis Persists throughout Aging. Cell Stem Cell 22, 589-599.e5. Bonaguidi, M.A., Wheeler, M.A., Shapiro, J.S., Stadel, R.P., Sun, G.J., Ming, G., and Song, H. (2011). In vivo clonal analysis reveals self-renewing and multipotent adult neural stem cell characteristics. Cell 145, 1142–1155. Bonzano, S., Bovetti, S., Gendusa, C., Peretto, P., and De Marchis, S. (2016). Adult Born Olfactory Bulb Dopaminergic Interneurons: Molecular Determinants and Experience- Dependent Plasticity. Front. Neurosci. 10. Bouab, M., Paliouras, G.N., Aumont, A., Forest-Bérard, K., and Fernandes, K.J.L. (2011). Aging of the subventricular zone neural stem cell niche: evidence for quiescence-associated changes between early and mid-adulthood. Neuroscience 173, 135–149. Bovetti, S., Hsieh, Y.-C., Bovolin, P., Perroteau, I., Kazunori, T., and Puche, A.C. (2007). Blood Vessels Form a Scaffold for Neuroblast Migration in the Adult Olfactory Bulb. J. Neurosci. 27, 5976–5980. Breton-Provencher, V., Lemasson, M., Peralta, M.R., and Saghatelyan, A. (2009). Interneurons Produced in Adulthood Are Required for the Normal Functioning of the Olfactory Bulb Network and for the Execution of Selected Olfactory Behaviors. J. Neurosci. 29, 15245– 15257. Brezun, J.M., and Daszuta, A. (1999). Depletion in serotonin decreases neurogenesis in the dentate gyrus and the subventricular zone of adult rats. Neuroscience 89, 999–1002. Brill, M.S., Ninkovic, J., Winpenny, E., Hodge, R.D., Ozen, I., Yang, R., Lepier, A., Gascón, S., Erdelyi, F., Szabo, G., et al. (2009). Adult generation of glutamatergic olfactory bulb interneurons. Nat. Neurosci. 12, 1524–1533.

101

Brill, M.S., Snapyan, M., Wohlfrom, H., Ninkovic, J., Jawerka, M., Mastick, G.S., Ashery-Padan, R., Saghatelyan, A., Berninger, B., and Götz, M. (2008). A Dlx2- and Pax6-Dependent Transcriptional Code for Periglomerular Neuron Specification in the Adult Olfactory Bulb. J Neurosci 28, 6439–6452. Brousse, B., Magalon, K., Durbec, P., and Cayre, M. (2015). Region and dynamic specificities of adult neural stem cells and oligodendrocyte precursors in myelin regeneration in the mouse brain. Biology Open 4, 980–992. Brown, G.C., and Neher, J.J. (2014). Microglial phagocytosis of live neurons. Nature Reviews Neuroscience 15, 209–216. Butovsky, O., Ziv, Y., Schwartz, A., Landa, G., Talpalar, A.E., Pluchino, S., Martino, G., and Schwartz, M. (2006). Microglia activated by IL-4 or IFN-gamma differentially induce neurogenesis and oligodendrogenesis from adult stem/progenitor cells. Mol. Cell. Neurosci. 31, 149–160. Cacci, E., Ajmone-Cat, M.A., Anelli, T., Biagioni, S., and Minghetti, L. (2008). In vitro neuronal and glial differentiation from embryonic or adult neural precursor cells are differently affected by chronic or acute activation of microglia. Glia 56, 412–425. Calaora, V., Chazal, G., Nielsen, P.J., Rougon, G., and moreau, H. (1996). mCD24 expression in the developing mouse brain and in zones of secondary neurogenesis in the adult. Neuroscience 73, 581–594. Calvert, J.W., Gundewar, S., Jha, S., Greer, J.J.M., Bestermann, W.H., Tian, R., and Lefer, D.J. (2008). Acute Metformin Therapy Confers Cardioprotection Against Myocardial Infarction Via AMPK-eNOS–Mediated Signaling. Diabetes 57, 696–705. Calzolari, F., Michel, J., Baumgart, E.V., Theis, F., Götz, M., and Ninkovic, J. (2015). Fast clonal expansion and limited neural stem cell self-renewal in the adult subependymal zone. Nat. Neurosci. 18, 490–492. Capela, A., and Temple, S. (2002). LeX/ssea-1 Is Expressed by Adult Mouse CNS Stem Cells, Identifying Them as Nonependymal. Neuron 35, 865–875. Capilla-Gonzalez, V., Cebrian-Silla, A., Guerrero-Cazares, H., Garcia-Verdugo, J.M., and Quiñones-Hinojosa, A. (2014a). Age-related changes in astrocytic and ependymal cells of the subventricular zone. Glia 62, 790–803. Capilla-Gonzalez, V., Guerrero-Cazares, H., Bonsu, J.M., Gonzalez-Perez, O., Achanta, P., Wong, J., Garcia-Verdugo, J.M., and Quiñones-Hinojosa, A. (2014b). The Subventricular Zone Is Able to Respond to a Demyelinating Lesion After Localized Radiation. Stem Cells 32, 59– 69. Carlén, M., Cassidy, R.M., Brismar, H., Smith, G.A., Enquist, L.W., and Frisén, J. (2002). Functional Integration of Adult-Born Neurons. Current Biology 12, 606–608. Carlén, M., Meletis, K., Göritz, C., Darsalia, V., Evergren, E., Tanigaki, K., Amendola, M., Barnabé- Heider, F., Yeung, M.S.Y., Naldini, L., et al. (2009). Forebrain ependymal cells are Notch-

102

dependent and generate neuroblasts and astrocytes after stroke. Nature Neuroscience 12, 259–267. Carleton, A., Petreanu, L.T., Lansford, R., Alvarez-Buylla, A., and Lledo, P.-M. (2003). Becoming a new neuron in the adult olfactory bulb. Nat. Neurosci. 6, 507–518. Chaker, Z., Codega, P., and Doetsch, F. (2016). A mosaic world: puzzles revealed by adult neural stem cell heterogeneity. Wiley Interdisciplinary Reviews: Developmental Biology 5, 640– 658. Chavali, M., Klingener, M., Kokkosis, A.G., Garkun, Y., Felong, S., Maffei, A., and Aguirre, A. (2018). Non-canonical Wnt signaling regulates neural stem cell quiescence during homeostasis and after demyelination. Nature Communications 9, 36. Chen, J., Zacharek, A., Zhang, C., Jiang, H., Li, Y., Roberts, C., Lu, M., Kapke, A., and Chopp, M. (2005). Endothelial Nitric Oxide Synthase Regulates Brain-Derived Neurotrophic Factor Expression and Neurogenesis after Stroke in Mice. J. Neurosci. 25, 2366–2375. Cheng, Z., Zheng, Y.Z., Li, Y.-Q., and Wong, C.S. (2017). Cellular Senescence in Mouse Hippocampus After Irradiation and the Role of p53 and p21. J Neuropathol Exp Neurol 76, 260–269. Chiasson, B.J., Tropepe, V., Morshead, C.M., and van der Kooy, D. (1999). Adult mammalian forebrain ependymal and subependymal cells demonstrate proliferative potential, but only subependymal cells have neural stem cell characteristics. J. Neurosci. 19, 4462– 4471. Chin, H.W., and Maruyama, Y. (1984). Age at treatment and long-term performance results in medulloblastoma. Cancer 53, 1952–1958. Choi, B.H., and Lapham, L.W. (1978). Radial glia in the human fetal cerebrum: A combined golgi, immunofluorescent and electron microscopic study. Brain Research 148, 295–311. Choi, Y.H., Kim, S.G., and Lee, M.G. (2006). Dose‐independent pharmacokinetics of metformin in rats: Hepatic and gastrointestinal first‐pass effects. Journal of Pharmaceutical Sciences 95, 2543–2552. Chow, B.M., Li, Y.-Q., and Wong, C.S. (2000). Radiation-induced apoptosis in the adult central nervous system is p53-dependent. Cell Death and Differentiation 7, 712. Clarke, L., and van der Kooy, D. (2011). The adult mouse dentate gyrus contains populations of committed progenitor cells that are distinct from subependymal zone neural stem cells. Stem Cells 29, 1448–1458. Codega, P., Silva-Vargas, V., Paul, A., Maldonado-Soto, A.R., DeLeo, A.M., Pastrana, E., and Doetsch, F. (2014). Prospective Identification and Purification of Quiescent Adult Neural Stem Cells from Their In Vivo Niche. Neuron 82, 545–559. Colak, D., Mori, T., Brill, M.S., Pfeifer, A., Falk, S., Deng, C., Monteiro, R., Mummery, C., Sommer, L., and Götz, M. (2008). Adult Neurogenesis Requires Smad4-Mediated Bone Morphogenic Protein Signaling in Stem Cells. J. Neurosci. 28, 434–446.

103

Coles‐Takabe, B.L.K., Brain, I., Purpura, K.A., Karpowicz, P., Zandstra, P.W., Morshead, C.M., and Kooy, D. van der (2008). Don’t Look: Growing Clonal Versus Nonclonal Neural Stem Cell Colonies. STEM CELLS 26, 2938–2944. Conover, J.C., and Shook, B.A. (2011). Aging of the Subventricular Zone Neural Stem Cell Niche. Aging Dis 2, 49–63. Conti, L., and Cattaneo, E. (2010). Neural stem cell systems: physiological players or in vitro entities? Nature Reviews Neuroscience 11, 176–187. Conti, L., Pollard, S.M., Gorba, T., Reitano, E., Toselli, M., Biella, G., Sun, Y., Sanzone, S., Ying, Q.- L., Cattaneo, E., et al. (2005). Niche-independent symmetrical self-renewal of a mammalian tissue stem cell. PLoS Biol. 3, e283. Corotto, F.S., Henegar, J.R., and Maruniak, J.A. (1994). Odor deprivation leads to reduced neurogenesis and reduced neuronal survival in the olfactory bulb of the adult mouse. Neuroscience 61, 739–744. Coskun, V., Wu, H., Blanchi, B., Tsao, S., Kim, K., Zhao, J., Biancotti, J.C., Hutnick, L., Krueger, R.C., Fan, G., et al. (2008). CD133+ neural stem cells in the ependyma of mammalian postnatal forebrain. PNAS 105, 1026–1031. Costa, M.R., Ortega, F., Brill, M.S., Beckervordersandforth, R., Petrone, C., Schroeder, T., Götz, M., and Berninger, B. (2011). Continuous live imaging of adult neural stem cell division and lineage progression in vitro. Development 138, 1057–1068. Craig, C.G., D’sa, R., Morshead, C.M., Roach, A., and van der Kooy, D. (1999). Migrational analysis of the constitutively proliferating subependyma population in adult mouse forebrain. Neuroscience 93, 1197–1206. Cremer, H., Lange, R., Christoph, A., Plomann, M., Vopper, G., Roes, J., Brown, R., Baldwin, S., Kraemer, P., and Scheff, S. (1994). Inactivation of the N-CAM gene in mice results in size reduction of the olfactory bulb and deficits in spatial learning. Nature 367, 455–459. Crouch, E.E., Liu, C., Silva-Vargas, V., and Doetsch, F. (2015). Regional and stage-specific effects of prospectively purified vascular cells on the adult V-SVZ neural stem cell lineage. J. Neurosci. 35, 4528–4539. Curtis, M.A., Kam, M., Nannmark, U., Anderson, M.F., Axell, M.Z., Wikkelso, C., Holtås, S., Roon- Mom, W.M.C. van, Björk-Eriksson, T., Nordborg, C., et al. (2007). Human Neuroblasts Migrate to the Olfactory Bulb via a Lateral Ventricular Extension. Science 315, 1243– 1249. d’Adda di Fagagna, F., Reaper, P.M., Clay-Farrace, L., Fiegler, H., Carr, P., Zglinicki, T. von, Saretzki, G., Carter, N.P., and Jackson, S.P. (2003). A DNA damage checkpoint response in telomere-initiated senescence. Nature 426, 194. Dadwal, P., Mahmud, N., Sinai, L., Azimi, A., Fatt, M., Wondisford, F.E., Miller, F.D., and Morshead, C.M. (2015). Activating endogenous neural precursor cells using metformin leads to neural repair and functional recovery in a model of childhood brain injury. Stem Cell Reports 5, 166–173.

104

Dahl, D., Rueger, D.C., Bignami, A., Weber, K., and Osborn, M. (1981). Vimentin, the 57 000 molecular weight protein of fibroblast filaments, is the major cytoskeletal component in immature glia. Eur. J. Cell Biol. 24, 191–196. Daynac, M., Chicheportiche, A., Pineda, J.R., Gauthier, L.R., Boussin, F.D., and Mouthon, M.-A. (2013). Quiescent neural stem cells exit dormancy upon alteration of GABAAR signaling following radiation damage. Stem Cell Research 11, 516–528. Deierborg, T., Roybon, L., Inacio, A.R., Pesic, J., and Brundin, P. (2010). Brain injury activates microglia that induce neural stem cell proliferation ex vivo and promote differentiation of neurosphere-derived cells into neurons and oligodendrocytes. Neuroscience 171, 1386–1396. Delgado, A.C., Ferrón, S.R., Vicente, D., Porlan, E., Perez-Villalba, A., Trujillo, C.M., D’Ocón, P., and Fariñas, I. (2014). Endothelial NT-3 delivered by vasculature and CSF promotes quiescence of subependymal neural stem cells through nitric oxide induction. Neuron 83, 572–585. Delgado, R.N., and Lim, D.A. (2015). Embryonic Nkx2.1-expressing neural precursor cells contribute to the regional heterogeneity of adult V–SVZ neural stem cells. Developmental Biology 407, 265–274. Dennis, C.V., Suh, L.S., Rodriguez, M.L., Kril, J.J., and Sutherland, G.T. (2016). Human adult neurogenesis across the ages: An immunohistochemical study. Neuropathology and Applied Neurobiology 42, 621–638. Didier, M., Harandi, M., Aguera, M., Bancel, B., Tardy, M., Fages, C., Calas, A., Stagaard, M., Møllgård, K., and Belin, M. (1986). Differential immunocytochemical staining for glial fibrillary acidic (GFA) protein, S-100 protein and glutamine synthetase in the rat subcommissural organ, nonspecialized ventricular ependyma and adjacent neuropil. Cell and Tissue Research 245, 343–351. Doetsch, F. (2003). A niche for adult neural stem cells. Current Opinion in Genetics & Development 13, 543–550. Doetsch, F., and Alvarez-Buylla, A. (1996). Network of tangential pathways for neuronal migration in adult mammalian brain. PNAS 93, 14895–14900. Doetsch, F., Caillé, I., Lim, D.A., García-Verdugo, J.M., and Alvarez-Buylla, A. (1999a). Subventricular zone astrocytes are neural stem cells in the adult mammalian brain. Cell 97, 703–716. Doetsch, F., García-Verdugo, J.M., and Alvarez-Buylla, A. (1997). Cellular composition and three- dimensional organization of the subventricular germinal zone in the adult mammalian brain. J. Neurosci. 17, 5046–5061. Doetsch, F., García-Verdugo, J.M., and Alvarez-Buylla, A. (1999b). Regeneration of a germinal layer in the adult mammalian brain. Proc. Natl. Acad. Sci. U.S.A. 96, 11619–11624.

105

Doetsch, F., Petreanu, L., Caille, I., Garcia-Verdugo, J.M., and Alvarez-Buylla, A. (2002). EGF converts transit-amplifying neurogenic precursors in the adult brain into multipotent stem cells. Neuron 36, 1021–1034. Douet, V., Kerever, A., Arikawa‐Hirasawa, E., and Mercier, F. (2013). Fractone-heparan sulphates mediate FGF-2 stimulation of cell proliferation in the adult subventricular zone. Cell Proliferation 46, 137–145. Duffner, P.K. (2004). Long-Term Effects of Radiation Therapy on Cognitive and Endocrine Function in Children With Leukemia and Brain Tumors: The Neurologist 10, 293–310. Dulken, B.W., Leeman, D.S., Boutet, S.C., Hebestreit, K., and Brunet, A. (2017). Single-Cell Transcriptomic Analysis Defines Heterogeneity and Transcriptional Dynamics in the Adult Neural Stem Cell Lineage. Cell Reports 18, 777–790. Dunn, C.J., and Peters, D.H. (1995). Metformin. Drugs 49, 721–749. Ekdahl, C.T., Kokaia, Z., and Lindvall, O. (2009). Brain inflammation and adult neurogenesis: The dual role of microglia. Neuroscience 158, 1021–1029. El-Mir, M.Y., Nogueira, V., Fontaine, E., Avéret, N., Rigoulet, M., and Leverve, X. (2000). Dimethylbiguanide inhibits cell respiration via an indirect effect targeted on the respiratory chain complex I. J. Biol. Chem. 275, 223–228. Encinas, J.M., Michurina, T.V., Peunova, N., Park, J.-H., Tordo, J., Peterson, D.A., Fishell, G., Koulakov, A., and Enikolopov, G. (2011). Division-Coupled Astrocytic Differentiation and Age-Related Depletion of Neural Stem Cells in the Adult Hippocampus. Cell Stem Cell 8, 566–579. Enwere, E., Shingo, T., Gregg, C., Fujikawa, H., Ohta, S., and Weiss, S. (2004). Aging Results in Reduced Epidermal Growth Factor Receptor Signaling, Diminished Olfactory Neurogenesis, and Deficits in Fine Olfactory Discrimination. J. Neurosci. 24, 8354–8365. Eriksson, P.S., Perfilieva, E., Björk-Eriksson, T., Alborn, A.-M., Nordborg, C., Peterson, D.A., and Gage, F.H. (1998). Neurogenesis in the adult human hippocampus. Nature Medicine 4, 1313–1317. Ernst, A., Alkass, K., Bernard, S., Salehpour, M., Perl, S., Tisdale, J., Possnert, G., Druid, H., and Frisén, J. (2014). Neurogenesis in the Striatum of the Adult Human Brain. Cell 156, 1072– 1083. Faiz, M., Sachewsky, N., Gascón, S., Bang, K.W.A., Morshead, C.M., and Nagy, A. (2015). Adult Neural Stem Cells from the Subventricular Zone Give Rise to Reactive Astrocytes in the Cortex after Stroke. Cell Stem Cell 17, 624–634. Fan, J., Li, D., Chen, H.-S., Huang, J.-G., Xu, J.-F., Zhu, W.-W., Chen, J.-G., and Wang, F. (2019). Metformin produces anxiolytic-like effects in rats by facilitating GABAA receptor trafficking to membrane. British Journal of Pharmacology 176, 297–316.

106

Fatt, M., Hsu, K., He, L., Wondisford, F., Miller, F.D., Kaplan, D.R., and Wang, J. (2015). Metformin acts on two different molecular pathways to enhance adult neural precursor proliferation/self-renewal and differentiation. Stem Cell Reports 5, 988–995. Feng, L., Hatten, M.E., and Heintz, N. (1994). Brain lipid-binding protein (BLBP): A novel signaling system in the developing mammalian CNS. Neuron 12, 895–908. Fernando, R.N., Eleuteri, B., Abdelhady, S., Nussenzweig, A., Andäng, M., and Ernfors, P. (2011). Cell cycle restriction by histone H2AX limits proliferation of adult neural stem cells. PNAS 108, 5837–5842. Ferrón, S.R., Marqués-Torrejón, M.A., Mira, H., Flores, I., Taylor, K., Blasco, M.A., and Fariñas, I. (2009). Telomere shortening in neural stem cells disrupts neuronal differentiation and neuritogenesis. J. Neurosci. 29, 14394–14407. Fiorelli, R., Azim, K., Fischer, B., and Raineteau, O. (2015). Adding a spatial dimension to postnatal ventricular-subventricular zone neurogenesis. Development 142, 2109–2120. Franco, R., and Fernández-Suárez, D. (2015). Alternatively activated microglia and macrophages in the central nervous system. Prog. Neurobiol. 131, 65–86. Frotscher, M. (2010). Role for Reelin in stabilizing cortical architecture. Trends Neurosci. 33, 407–414. Fuentealba, L.C., Rompani, S.B., Parraguez, J.I., Obernier, K., Romero, R., Cepko, C.L., and Alvarez-Buylla, A. (2015). Embryonic origin of postnatal neural stem cells. Cell 161, 1644–1655. Fujitani, M., Cancino, G.I., Dugani, C.B., Weaver, I.C.G., Gauthier-Fisher, A., Paquin, A., Mak, T.W., Wojtowicz, M.J., Miller, F.D., and Kaplan, D.R. (2010). TAp73 Acts via the bHLH Hey2 to Promote Long-Term Maintenance of Neural Precursors. Current Biology 20, 2058–2065. Fukuda, A., Fukuda, H., Swanpalmer, J., Hertzman, S., Lannering, B., Marky, I., Bjork-Eriksson, T., and Blomgren, K. (2005). Age-dependent sensitivity of the developing brain to irradiation is correlated with the number and vulnerability of progenitor cells. Journal of Neurochemistry 92, 569–584. Fukuda, H., Fukuda, A., Zhu, C., Korhonen, L., Swanpalmer, J., Hertzman, S., Leist, M., Lannering, B., Lindholm, D., Björk-Eriksson, T., et al. (2004). Irradiation-induced progenitor cell death in the developing brain is resistant to erythropoietin treatment and caspase inhibition. Cell Death and Differentiation 11, 1166–1178. Gattazzo, F., Urciuolo, A., and Bonaldo, P. (2014). Extracellular matrix: a dynamic microenvironment for stem cell niche. Biochim Biophys Acta 1840, 2506–2519. Gemma, C., and Bachstetter, A.D. (2013). The role of microglia in adult hippocampal neurogenesis. Front. Cell. Neurosci. 7. Gensert, J.M., and Goldman, J.E. (1997). Endogenous Progenitors Remyelinate Demyelinated Axons in the Adult CNS. Neuron 19, 197–203.

107

Gheusi, G., Cremer, H., McLean, H., Chazal, G., Vincent, J.-D., and Lledo, P.-M. (2000). Importance of newly generated neurons in the adult olfactory bulb for odor discrimination. PNAS 97, 1823–1828. Giachino, C., Basak, O., Lugert, S., Knuckles, P., Obernier, K., Fiorelli, R., Frank, S., Raineteau, O., Alvarez‐Buylla, A., and Taylor, V. (2014). Molecular Diversity Subdivides the Adult Forebrain Neural Stem Cell Population. STEM CELLS 32, 70–84. Gil‐Perotín, S., Duran‐Moreno, M., Cebrián‐Silla, A., Ramírez, M., García‐Belda, P., and García‐ Verdugo, J.M. (2013). Adult neural stem cells from the subventricular zone: a review of the neurosphere assay. The Anatomical Record 296, 1435–1452. Giovannucci, E., Harlan, D.M., Archer, M.C., Bergenstal, R.M., Gapstur, S.M., Habel, L.A., Pollak, M., Regensteiner, J.G., and Yee, D. (2010). Diabetes and cancer: a consensus report. Diabetes Care 33, 1674–1685. Goings, G.E., Kozlowski, D.A., and Szele, F.G. (2006). Differential activation of microglia in neurogenic versus non-neurogenic regions of the forebrain. Glia 54, 329–342. Gómez-Gaviro, M.V., Scott, C.E., Sesay, A.K., Matheu, A., Booth, S., Galichet, C., and Lovell- Badge, R. (2012). Betacellulin promotes cell proliferation in the neural stem cell niche and stimulates neurogenesis. Proc. Natl. Acad. Sci. U.S.A. 109, 1317–1322. Gong, L., Goswami, S., Giacomini, K.M., Altman, R.B., and Klein, T.E. (2012). Metformin pathways: pharmacokinetics and pharmacodynamics. Pharmacogenet Genomics 22, 820–827. Gonzalez-Perez, O., Gutierrez-Fernandez, F., Lopez-Virgen, V., Collas-Aguilar, J., Quinones- Hinojosa, A., and Garcia-Verdugo, J.M. (2012). Immunological regulation of neurogenic niches in the adult brain. Neuroscience 226, 270–281. Goodhead, D.T. (1994). Initial events in the cellular effects of ionizing radiations: clustered damage in DNA. Int. J. Radiat. Biol. 65, 7–17. Götz, M., Stoykova, A., and Gruss, P. (1998). Pax6 Controls Radial Glia Differentiation in the Cerebral Cortex. Neuron 21, 1031–1044. Gould, S.J., Howard, S., and Papadaki, L. (1990). The development of ependyma in the human fetal brain: an immunohistological and electron microscopic study. Developmental Brain Research 55, 255–267. Graham, G.G., Punt, J., Arora, M., Day, R.O., Doogue, M.P., Duong, J.K., Furlong, T.J., Greenfield, J.R., Greenup, L.C., Kirkpatrick, C.M., et al. (2011). Clinical pharmacokinetics of metformin. Clin Pharmacokinet 50, 81–98. Greer, E.L., Oskoui, P.R., Banko, M.R., Maniar, J.M., Gygi, M.P., Gygi, S.P., and Brunet, A. (2007). The Energy Sensor AMP-activated Protein Kinase Directly Regulates the Mammalian FOXO3 Transcription Factor. J. Biol. Chem. 282, 30107–30119. Gritti, A., Parati, E.A., Cova, L., Frolichsthal, P., Galli, R., Wanke, E., Faravelli, L., Morassutti, D.J., Roisen, F., Nickel, D.D., et al. (1996). Multipotential stem cells from the adult mouse

108

brain proliferate and self-renew in response to basic fibroblast growth factor. J. Neurosci. 16, 1091–1100. Gwinn, D.M., Shackelford, D.B., Egan, D.F., Mihaylova, M.M., Mery, A., Vasquez, D.S., Turk, B.E., and Shaw, R.J. (2008). AMPK Phosphorylation of Raptor Mediates a Metabolic Checkpoint. Molecular Cell 30, 214–226. Hack, M.A., Saghatelyan, A., Chevigny, A. de, Pfeifer, A., Ashery-Padan, R., Lledo, P.-M., and Götz, M. (2005). Neuronal fate determinants of adult olfactory bulb neurogenesis. Nature Neuroscience 8, 865. Hack, M.A., Sugimori, M., Lundberg, C., Nakafuku, M., and Götz, M. (2004). Regionalization and fate specification in neurospheres: the role of Olig2 and Pax6. Molecular and Cellular Neuroscience 25, 664–678. Han, W., Umekawa, T., Zhou, K., Zhang, X.-M., Ohshima, M., Dominguez, C.A., Harris, R.A., Zhu, C., and Blomgren, K. (2016). Cranial irradiation induces transient microglia accumulation, followed by long-lasting inflammation and loss of microglia. Oncotarget 7, 82305–82323. Hartfuss, E., Galli, R., Heins, N., and Götz, M. (2001). Characterization of CNS Precursor Subtypes and Radial Glia. Developmental Biology 229, 15–30. Hattori, Y., Suzuki, K., Hattori, S., and Kasai, K. (2006). Metformin inhibits cytokine-induced nuclear factor κB activation via AMP-activated protein kinase activation in vascular endothelial cells. Hypertension 47, 1183–1188. Haubensak, W., Attardo, A., Denk, W., and Huttner, W.B. (2004). Neurons arise in the basal neuroepithelium of the early mammalian telencephalon: a major site of neurogenesis. Proc. Natl. Acad. Sci. U.S.A. 101, 3196–3201. Hayashi, M., Luo, Y., Laning, J., Strieter, R.M., and Dorf, M.E. (1995). Production and function of monocyte chemoattractant protein-1 and other beta-chemokines in murine glial cells. J. Neuroimmunol. 60, 143–150. He, L., Sabet, A., Djedjos, S., Miller, R., Sun, X., Hussain, M.A., Radovick, S., and Wondisford, F.E. (2009). Metformin and insulin suppress hepatic gluconeogenesis through phosphorylation of CREB binding protein. Cell 137, 635–646. Hellström, N.A.K., Björk-Eriksson, T., Blomgren, K., and Kuhn, H.G. (2009). Differential Recovery of Neural Stem Cells in the Subventricular Zone and Dentate Gyrus After Ionizing Radiation. Stem Cells 27, 634–641. Hennen, E., and Faissner, A. (2012). LewisX: A neural stem cell specific glycan? The International Journal of Biochemistry & Cell Biology 44, 830–833. Hinojosa, A.E., Garcia-Bueno, B., Leza, J.C., and Madrigal, J.L. (2011). CCL2/MCP-1 modulation of microglial activation and proliferation. J Neuroinflammation 8, 77. Hitoshi, S., Alexson, T., Tropepe, V., Donoviel, D., Elia, A.J., Nye, J.S., Conlon, R.A., Mak, T.W., Bernstein, A., and Kooy, D. van der (2002). Notch pathway molecules are essential for

109

the maintenance, but not the generation, of mammalian neural stem cells. Genes Dev. 16, 846–858. Hoehn, B.D., Palmer, T.D., and Steinberg, G.K. (2005). Neurogenesis in Rats After Focal Cerebral Ischemia is Enhanced by Indomethacin. Stroke 36, 2718–2724. Höglinger, G.U., Rizk, P., Muriel, M.P., Duyckaerts, C., Oertel, W.H., Caille, I., and Hirsch, E.C. (2004). Dopamine depletion impairs precursor cell proliferation in Parkinson disease. Nature Neuroscience 7, 726. Hong, J.-H., Chiang, C.-S., Campbell, I.L., Sun, J.-R., Withers, H.R., and McBride, W.H. (1995). Induction of acute phase gene expression by brain irradiation. International Journal of Radiation Oncology*Biology*Physics 33, 619–626. Honig, L.S., Herrmann, K., and Shatz, C.J. (1996). Developmental Changes Revealed by Immunohistochemical Markers in Human Cerebral Cortex. Cereb Cortex 6, 794–806. Hou, S.-W., Wang, Y.-Q., Xu, M., Shen, D.-H., Wang, J.-J., Huang, F., Yu, Z., and Sun, F.-Y. (2008). Functional Integration of Newly Generated Neurons Into Striatum After Cerebral Ischemia in the Adult Rat Brain. Stroke 39, 2837–2844. Hou, X., Song, J., Li, X.-N., Zhang, L., Wang, X., Chen, L., and Shen, Y.H. (2010). Metformin reduces intracellular reactive oxygen species levels by upregulating expression of the antioxidant thioredoxin via the AMPK-FOXO3 pathway. Biochemical and Biophysical Research Communications 396, 199–205. Hu, X.-L., Chen, G., Zhang, S., Zheng, J., Wu, J., Bai, Q.-R., Wang, Y., Li, J., Wang, H., Feng, H., et al. (2017). Persistent Expression of VCAM1 in Radial Glial Cells Is Required for the Embryonic Origin of Postnatal Neural Stem Cells. Neuron 95, 309-325.e6. Hundal, R.S., Krssak, M., Dufour, S., Laurent, D., Lebon, V., Chandramouli, V., Inzucchi, S.E., Schumann, W.C., Petersen, K.F., Landau, B.R., et al. (2000). Mechanism by which metformin reduces glucose production in type 2 diabetes. Diabetes 49, 2063–2069. Hurtado-Chong, A., Yusta-Boyo, M.J., Vergaño-Vera, E., Bulfone, A., de Pablo, F., and Vicario- Abejón, C. (2009). IGF-I promotes neuronal migration and positioning in the olfactory bulb and the exit of neuroblasts from the subventricular zone. Eur. J. Neurosci. 30, 742– 755. Hwang, I.K., Kim, I.Y., Joo, E.J., Shin, J.H., Choi, J.W., Won, M.-H., Yoon, Y.S., and Seong, J.K. (2010). Metformin normalizes type 2 diabetes-induced decrease in cell proliferation and neuroblast differentiation in the rat dentate gyrus. Neurochem. Res. 35, 645–650. Iglesias-Bartolome, R., Patel, V., Cotrim, A., Leelahavanichkul, K., Molinolo, A.A., Mitchell, J.B., and Gutkind, J.S. (2012). mTOR Inhibition Prevents Epithelial Stem Cell Senescence and Protects from Radiation-Induced Mucositis. Cell Stem Cell 11, 401–414. Ihrie, R.A., and Álvarez-Buylla, A. (2011). Lake-Front Property: A Unique Germinal Niche by the Lateral Ventricles of the Adult Brain. Neuron 70, 674–686.

110

Iliakis, G. (1991). The role of DNA double strand breaks in ionizing radiation‐induced killing of eukaryotic cells. BioEssays 13, 641–648. Imayoshi, I., Sakamoto, M., Ohtsuka, T., Takao, K., Miyakawa, T., Yamaguchi, M., Mori, K., Ikeda, T., Itohara, S., and Kageyama, R. (2008). Roles of continuous neurogenesis in the structural and functional integrity of the adult forebrain. Nat. Neurosci. 11, 1153–1161. Imayoshi, I., Sakamoto, M., Yamaguchi, M., Mori, K., and Kageyama, R. (2010). Essential Roles of Notch Signaling in Maintenance of Neural Stem Cells in Developing and Adult Brains. J. Neurosci. 30, 3489–3498. Inazu, M., Takeda, H., Maehara, K., Miyashita, K., Tomoda, A., and Matsumiya, T. (2006). Functional expression of the organic cation/carnitine transporter 2 in rat astrocytes. Journal of Neurochemistry 97, 424–434. Inoki, K., Zhu, T., and Guan, K.-L. (2003). TSC2 Mediates Cellular Energy Response to Control Cell Growth and Survival. Cell 115, 577–590. Ishimoto, T., Nakamichi, N., Hosotani, H., Masuo, Y., Sugiura, T., and Kato, Y. (2014). Organic cation transporter-mediated ergothioneine uptake in mouse neural progenitor cells suppresses proliferation and promotes differentiation into neurons. PLoS ONE 9, e89434. Jacquet, B.V., Salinas-Mondragon, R., Liang, H., Therit, B., Buie, J.D., Dykstra, M., Campbell, K., Ostrowski, L.E., Brody, S.L., and Ghashghaei, H.T. (2009). FoxJ1-dependent gene expression is required for differentiation of radial glia into ependymal cells and a subset of astrocytes in the postnatal brain. Development 136, 4021–4031. Jenrow, K.A., Brown, S.L., Lapanowski, K., Naei, H., Kolozsvary, A., and Kim, J.H. (2013). Selective inhibition of microglia-mediated neuroinflammation mitigates radiation-induced cognitive impairment. Radiat. Res. 179, 549–556. Jin, Q., Cheng, J., Liu, Y., Wu, J., Wang, X., Wei, S., Zhou, X., Qin, Z., Jia, J., and Zhen, X. (2014). Improvement of functional recovery by chronic metformin treatment is associated with enhanced alternative activation of microglia/macrophages and increased angiogenesis and neurogenesis following experimental stroke. Brain, Behavior, and Immunity 40, 131–142. Johansson, C.B., Momma, S., Clarke, D.L., Risling, M., Lendahl, U., and Frisén, J. (1999). Identification of a Neural Stem Cell in the Adult Mammalian Central Nervous System. Cell 96, 25–34. Johnson, G.V., and Jope, R.S. (1992). The role of microtubule-associated protein 2 (MAP-2) in neuronal growth, plasticity, and degeneration. J. Neurosci. Res. 33, 505–512. Jung, S., Aliberti, J., Graemmel, P., Sunshine, M.J., Kreutzberg, G.W., Sher, A., and Littman, D.R. (2000). Analysis of fractalkine receptor CX(3)CR1 function by targeted deletion and green fluorescent protein reporter gene insertion. Mol. Cell. Biol. 20, 4106–4114.

111

Kalamakis, G., Brüne, D., Ravichandran, S., Bolz, J., Fan, W., Ziebell, F., Stiehl, T., Catalá- Martinez, F., Kupke, J., Zhao, S., et al. (2019). Quiescence Modulates Stem Cell Maintenance and Regenerative Capacity in the Aging Brain. Cell 176, 1407-1419.e14. Kalm, M., Fukuda, A., Fukuda, H., Öhrfelt, A., Lannering, B., Björk-Eriksson, T., Blennow, K., Márky, I., and Blomgren, K. (2009). Transient Inflammation in Neurogenic Regions after Irradiation of the Developing Brain. Rare 171, 66–76. Kaplan, M.S., McNelly, N.A., and Hinds, J.W. (1985). Population dynamics of adult-formed granule neurons of the rat olfactory bulb. J. Comp. Neurol. 239, 117–125. Karpowicz, P., Morshead, C., Kam, A., Jervis, E., Ramunas, J., Cheng, V., and Kooy, D. van der (2005). Support for the immortal strand hypothesis: neural stem cells partition DNA asymmetrically in vitro. J Cell Biol 170, 721–732. Katsimpardi, L., Litterman, N.K., Schein, P.A., Miller, C.M., Loffredo, F.S., Wojtkiewicz, G.R., Chen, J.W., Lee, R.T., Wagers, A.J., and Rubin, L.L. (2014). Vascular and Neurogenic Rejuvenation of the Aging Mouse Brain by Young Systemic Factors. Science 344, 630– 634. Kawaguchi, D., Furutachi, S., Kawai, H., Hozumi, K., and Gotoh, Y. (2013). Dll1 maintains quiescence of adult neural stem cells and segregates asymmetrically during mitosis. Nature Communications 4, 1880. Kazanis, I., Lathia, J.D., Vadakkan, T.J., Raborn, E., Wan, R., Mughal, M.R., Eckley, D.M., Sasaki, T., Patton, B., Mattson, M.P., et al. (2010). Quiescence and Activation of Stem and Precursor Cell Populations in the Subependymal Zone of the Mammalian Brain Are Associated with Distinct Cellular and Extracellular Matrix Signals. J. Neurosci. 30, 9771– 9781. Kerever, A., Schnack, J., Vellinga, D., Ichikawa, N., Moon, C., Arikawa‐Hirasawa, E., Efird, J.T., and Mercier, F. (2007). Novel Extracellular Matrix Structures in the Neural Stem Cell Niche Capture the Neurogenic Factor Fibroblast Growth Factor 2 from the Extracellular Milieu. Stem Cells 25, 2146–2157. Keung, A.J., de Juan-Pardo, E.M., Schaffer, D.V., and Kumar, S. (2011). Rho GTPases Mediate the Mechanosensitive Lineage Commitment of Neural Stem Cells. Stem Cells 29, 1886–1897. Kim, J.-M., Yoo, H., Kim, J.-Y., Oh, S.H., Kang, J.W., Yoo, B.R., Han, S.Y., Kim, C.S., Choi, W.H., Lee, E.-J., et al. (2018). Metformin Alleviates Radiation-Induced Skin Fibrosis via the Downregulation of FOXO3. Cell. Physiol. Biochem. 48, 959–970. Kim, W.R., Kim, Y., Eun, B., Park, O., Kim, H., Kim, K., Park, C.-H., Vinsant, S., Oppenheim, R.W., and Sun, W. (2007). Impaired migration in the rostral migratory stream but spared olfactory function after the elimination of programmed cell death in Bax knock-out mice. J. Neurosci. 27, 14392–14403. Kim, Y., Wang, W.-Z., Comte, I., Pastrana, E., Tran, P.B., Brown, J., Miller, R.J., Doetsch, F., Molnár, Z., and Szele, F.G. (2010). Dopamine stimulation of postnatal murine

112

subventricular zone neurogenesis via the D3 receptor. Journal of Neurochemistry 114, 750–760. Kimura, A., Ohmori, T., Kashiwakura, Y., Ohkawa, R., Madoiwa, S., Mimuro, J., Shimazaki, K., Hoshino, Y., Yatomi, Y., and Sakata, Y. (2008). Antagonism of Sphingosine 1-Phosphate Receptor-2 Enhances Migration of Neural Progenitor Cells Toward an Area of Brain Infarction. Stroke 39, 3411–3417. Kippin, T.E., Kapur, S., and Kooy, D. van der (2005). Dopamine Specifically Inhibits Forebrain Neural Stem Cell Proliferation, Suggesting a Novel Effect of Antipsychotic Drugs. J. Neurosci. 25, 5815–5823. Kirschenbaum, B., and Goldman, S.A. (1995). Brain-derived neurotrophic factor promotes the survival of neurons arising from the adult rat forebrain subependymal zone. PNAS 92, 210–214. Knoth, R., Singec, I., Ditter, M., Pantazis, G., Capetian, P., Meyer, R.P., Horvat, V., Volk, B., and Kempermann, G. (2010). Murine features of neurogenesis in the human hippocampus across the lifespan from 0 to 100 years. PLoS ONE 5, e8809. Kokovay, E., Goderie, S., Wang, Y., Lotz, S., Lin, G., Sun, Y., Roysam, B., Shen, Q., and Temple, S. (2010). Adult SVZ lineage cells home to and leave the vascular niche via differential responses to SDF1/CXCR4 signaling. Cell Stem Cell 7, 163–173. Kokovay, E., Wang, Y., Kusek, G., Wurster, R., Lederman, P., Lowry, N., Shen, Q., and Temple, S. (2012). VCAM1 Is Essential to Maintain the Structure of the SVZ Niche and Acts as an Environmental Sensor to Regulate SVZ Lineage Progression. Cell Stem Cell 11, 220–230. Kolb, B., Morshead, C., Gonzalez, C., Kim, M., Gregg, C., Shingo, T., and Weiss, S. (2007). Growth Factor-Stimulated Generation of New Cortical Tissue and Functional Recovery after Stroke Damage to the Motor Cortex of Rats. J Cereb Blood Flow Metab 27, 983–997. Kriegstein, A., and Alvarez-Buylla, A. (2009). The Glial Nature of Embryonic and Adult Neural Stem Cells. Annual Review of Neuroscience 32, 149–184. Kumase, F., Takeuchi, K., Morizane, Y., Suzuki, J., Matsumoto, H., Kataoka, K., Al-Moujahed, A., Maidana, D.E., Miller, J.W., and Vavvas, D.G. (2016). AMPK-activated protein kinase suppresses Ccr2 expression by inhibiting the NF-[kappa]B pathway in RAW264.7 macrophages. PLoS ONE 11, e0147279. L’Episcopo, F., Tirolo, C., Caniglia, S., Testa, N., Serra, P.A., Impagnatiello, F., Morale, M.C., and Marchetti, B. (2010). Combining nitric oxide release with anti-inflammatory activity preserves nigrostriatal dopaminergic innervation and prevents motor impairment in a 1- methyl-4-phenyl-1,2,3,6-tetrahydropyridine model of Parkinson’s disease. Journal of Neuroinflammation 7, 83. L’Episcopo, F., Tirolo, C., Testa, N., Caniglia, S., Morale, M.C., Deleidi, M., Serapide, M.F., Pluchino, S., and Marchetti, B. (2012). Plasticity of Subventricular Zone Neuroprogenitors in MPTP (1-Methyl-4-Phenyl-1,2,3,6-Tetrahydropyridine) Mouse Model of Parkinson’s Disease Involves Cross Talk between Inflammatory and Wnt/β-

113

Catenin Signaling Pathways: Functional Consequences for Neuroprotection and Repair. J. Neurosci. 32, 2062–2085. Lazarini, F., and Lledo, P.-M. (2011). Is adult neurogenesis essential for olfaction? Trends Neurosci. 34, 20–30. Lazarini, F., Mouthon, M.-A., Gheusi, G., Chaumont, F. de, Olivo-Marin, J.-C., Lamarque, S., Abrous, D.N., Boussin, F.D., and Lledo, P.-M. (2009). Cellular and behavioral effects of cranial irradiation of the subventricular zone in adult mice. PLOS ONE 4, e7017. Łabuzek, K., Liber, S., Gabryel, B., and Okopień, B. (2010a). Metformin has adenosine- monophosphate activated protein kinase (AMPK)-independent effects on LPS- stimulated rat primary microglial cultures. Pharmacological Reports 62, 827–848. Łabuzek, K., Suchy, D., Gabryel, B., Bielecka, A., Liber, S., and Okopień, B. (2010b). Quantification of metformin by the HPLC method in brain regions, cerebrospinal fluid and plasma of rats treated with lipopolysaccharide. Pharmacological Reports 62, 956– 965. Le Belle, J.E., Orozco, N.M., Paucar, A.A., Saxe, J.P., Mottahedeh, J., Pyle, A.D., Wu, H., and Kornblum, H.I. (2011). Proliferative Neural Stem Cells Have High Endogenous ROS Levels that Regulate Self-Renewal and Neurogenesis in a PI3K/Akt-Dependant Manner. Cell Stem Cell 8, 59–71. Ledeboer, A., Brevé, J.J.P., Poole, S., Tilders, F.J.H., and Dam, A.-M.V. (2000). Interleukin-10, interleukin-4, and transforming growth factor-β differentially regulate lipopolysaccharide-induced production of pro-inflammatory cytokines and nitric oxide in co-cultures of rat astroglial and microglial cells. Glia 30, 134–142. Lee, D.-F., Kuo, H.-P., Chen, C.-T., Hsu, J.-M., Chou, C.-K., Wei, Y., Sun, H.-L., Li, L.-Y., Ping, B., Huang, W.-C., et al. (2007). IKK beta suppression of TSC1 links inflammation and tumor angiogenesis via the mTOR pathway. Cell 130, 440–455. Lee, S.W., Haditsch, U., Cord, B.J., Guzman, R., Kim, S.J., Boettcher, C., Priller, J., Ormerod, B.K., and Palmer, T.D. (2013). Absence of CCL2 is sufficient to restore hippocampal neurogenesis following cranial irradiation. Brain Behavior and Immunity 30, 33–44. Lehtinen, M.K., Zappaterra, M.W., Chen, X., Yang, Y.J., Hill, A.D., Lun, M., Maynard, T., Gonzalez, D., Kim, S., Ye, P., et al. (2011). The Cerebrospinal Fluid Provides a Proliferative Niche for Neural Progenitor Cells. Neuron 69, 893–905. Lemasson, M., Saghatelyan, A., Olivo-Marin, J.-C., and Lledo, P.-M. (2005). Neonatal and adult neurogenesis provide two distinct populations of newborn neurons to the mouse olfactory bulb. J. Neurosci. 25, 6816–6825. Lim, D.A., Tramontin, A.D., Trevejo, J.M., Herrera, D.G., García-Verdugo, J.M., and Alvarez- Buylla, A. (2000). Noggin Antagonizes BMP Signaling to Create a Niche for Adult Neurogenesis. Neuron 28, 713–726. Little, J.B. (2000). Radiation carcinogenesis. Carcinogenesis 21, 397–404.

114

Liu, X., Wang, Q., Haydar, T.F., and Bordey, A. (2005). Nonsynaptic GABA signaling in postnatal subventricular zone controls proliferation of GFAP-expressing progenitors. Nature Neuroscience 8, 1179. Liu, Y., Tang, G., Zhang, Z., Wang, Y., and Yang, G.-Y. (2014). Metformin promotes focal angiogenesis and neurogenesis in mice following middle cerebral artery occlusion. Neuroscience Letters 579, 46–51. Llorens-Bobadilla, E., Zhao, S., Baser, A., Saiz-Castro, G., Zwadlo, K., and Martin-Villalba, A. (2015). Single-Cell Transcriptomics Reveals a Population of Dormant Neural Stem Cells that Become Activated upon Brain Injury. Cell Stem Cell 17, 329–340. Lois, C., and Alvarez-Buylla, A. (1993). Proliferating subventricular zone cells in the adult mammalian forebrain can differentiate into neurons and glia. PNAS 90, 2074–2077. Lois, C., and Alvarez-Buylla, A. (1994). Long-distance neuronal migration in the adult mammalian brain. Science 264, 1145–1148. Lois, C., García-Verdugo, J.-M., and Alvarez-Buylla, A. (1996). Chain Migration of Neuronal Precursors. Science 271, 978–981. López-Juárez, A., Howard, J., Ullom, K., Howard, L., Grande, A., Pardo, A., Waclaw, R., Sun, Y.-Y., Yang, D., Kuan, C.-Y., et al. (2013). Gsx2 controls region-specific activation of neural stem cells and injury-induced neurogenesis in the adult subventricular zone. Genes Dev. 27, 1272–1287. Luo, J., Daniels, S.B., Lennington, J.B., Notti, R.Q., and Conover, J.C. (2006). The aging neurogenic subventricular zone. Aging Cell 5, 139–152. Luo, Y., Coskun, V., Liang, A., Yu, J., Cheng, L., Ge, W., Shi, Z., Zhang, K., Li, C., Cui, Y., et al. (2015). Single-Cell Transcriptome Analyses Reveal Signals to Activate Dormant Neural Stem Cells. Cell 161, 1175–1186. Luskin, M.B. (1993). Restricted proliferation and migration of postnatally generated neurons derived from the forebrain subventricular zone. Neuron 11, 173–189. Luskin, M.B., and Boone, M.S. (1994). Rate and pattern of migration of lineally-related olfactory bulb interneurons generated postnatally in the subventricular zone of the rat. Chem. Senses 19, 695–714. Mabbott, D.J., Spiegler, B.J., Greenberg, M.L., Rutka, J.T., Hyder, D.J., and Bouffet, E. (2005). Serial evaluation of academic and behavioral outcome after treatment with cranial radiation in childhood. Journal of Clinical Oncology 23, 2256–2263. Magavi, S.S.P., Mitchell, B.D., Szentirmai, O., Carter, B.S., and Macklis, J.D. (2005). Adult-Born and Preexisting Olfactory Granule Neurons Undergo Distinct Experience-Dependent Modifications of their Olfactory Responses In Vivo. J. Neurosci. 25, 10729–10739. Mahla, R.S. (2016). Stem Cells Applications in Regenerative Medicine and Disease Therapeutics. Int J Cell Biol 2016.

115

Mak, G.K., Enwere, E.K., Gregg, C., Pakarainen, T., Poutanen, M., Huhtaniemi, I., and Weiss, S. (2007). Male pheromone–stimulated neurogenesis in the adult female brain: possible role in mating behavior. Nature Neuroscience 10, 1003–1011. Mak, G.K., and Weiss, S. (2010). Paternal recognition of adult offspring mediated by newly generated CNS neurons. Nature Neuroscience 13, 753–758. Malatesta, P., Appolloni, I., and Calzolari, F. (2008). Radial glia and neural stem cells. Cell Tissue Res. 331, 165–178. Marín-Teva, J.L., Cuadros, M.A., Martín-Oliva, D., and Navascués, J. (2011). Microglia and neuronal cell death. Neuron Glia Biology 7, 25–40. Marqués-Torrejón, M.Á., Porlan, E., Banito, A., Gómez-Ibarlucea, E., Lopez-Contreras, A.J., Fernández-Capetillo, Ó., Vidal, A., Gil, J., Torres, J., and Fariñas, I. (2013). Cyclin- Dependent Kinase Inhibitor p21 Controls Adult Neural Stem Cell Expansion by Regulating Sox2 Gene Expression. Cell Stem Cell 12, 88–100. Marshall, C.A.G., Novitch, B.G., and Goldman, J.E. (2005). Olig2 directs astrocyte and oligodendrocyte formation in postnatal subventricular zone cells. J. Neurosci. 25, 7289– 7298. Marti-Fabregas, J., Romaguera-Ros, M., Gomez-Pinedo, U., Martinez-Ramirez, S., Jimenez- Xarrie, E., Marin, R., Marti-Vilalta, J.-L., and Garcia-Verdugo, J.-M. (2010). Proliferation in the human ipsilateral subventricular zone after ischemic stroke. Neurology 74, 357–365. Martin-Montalvo, A., Mercken, E.M., Mitchell, S.J., Palacios, H.H., Mote, P.L., Scheibye- Knudsen, M., Gomes, A.P., Ward, T.M., Minor, R.K., Blouin, M.-J., et al. (2013). Metformin improves healthspan and lifespan in mice. Nature Communications 4, 2192. Maslov, A.Y., Barone, T.A., Plunkett, R.J., and Pruitt, S.C. (2004). Neural Stem Cell Detection, Characterization, and Age-Related Changes in the Subventricular Zone of Mice. J. Neurosci. 24, 1726–1733. Massalini, S., Pellegatta, S., Pisati, F., Finocchiaro, G., Farace, M.G., and Ciafrè, S.A. (2009). Reelin affects chain-migration and differentiation of neural precursor cells. Mol. Cell. Neurosci. 42, 341–349. Matarredona, E.R., Talaverón, R., and Pastor, A.M. (2018). Interactions Between Neural Progenitor Cells and Microglia in the Subventricular Zone: Physiological Implications in the Neurogenic Niche and After Implantation in the Injured Brain. Front Cell Neurosci 12. Matcovitch-Natan, O., Winter, D.R., Giladi, A., Vargas Aguilar, S., Spinrad, A., Sarrazin, S., Ben- Yehuda, H., David, E., Zelada González, F., Perrin, P., et al. (2016). Microglia development follows a stepwise program to regulate brain homeostasis. Science 353, aad8670. McGinn, M.J., Sun, D., and Colello, R.J. (2008). Utilizing X-irradiation to selectively eliminate neural stem/progenitor cells from neurogenic regions of the mammalian brain. Journal of Neuroscience Methods 170, 9–15.

116

McMahon, J.A., Takada, S., Zimmerman, L.B., Fan, C.-M., Harland, R.M., and McMahon, A.P. (1998). Noggin-mediated antagonism of BMP signaling is required for growth and patterning of the neural tube and somite. Genes Dev 12, 1438–1452. Menn, B., Garcia-Verdugo, J.M., Yaschine, C., Gonzalez-Perez, O., Rowitch, D., and Alvarez- Buylla, A. (2006). Origin of Oligodendrocytes in the Subventricular Zone of the Adult Brain. J. Neurosci. 26, 7907–7918. Mercier, F., Kitasako, J.T., and Hatton, G.I. (2002). Anatomy of the brain neurogenic zones revisited: Fractones and the fibroblast/macrophage network. Journal of Comparative Neurology 451, 170–188. Merkle, F.T., Mirzadeh, Z., and Alvarez-Buylla, A. (2007). Mosaic Organization of Neural Stem Cells in the Adult Brain. Science 317, 381–384. Merkle, F.T., Tramontin, A.D., García-Verdugo, J.M., and Alvarez-Buylla, A. (2004). Radial glia give rise to adult neural stem cells in the subventricular zone. PNAS 101, 17528–17532. Miller, F.D., and Kaplan, D.R. (2012). Mobilizing Endogenous Stem Cells for Repair and Regeneration: Are We There Yet? Cell Stem Cell 10, 650–652. Mirzadeh, Z., Merkle, F.T., Soriano-Navarro, M., Garcia-Verdugo, J.M., and Alvarez-Buylla, A. (2008). Neural stem cells confer unique pinwheel architecture to the ventricular surface in neurogenic regions of the adult brain. Cell Stem Cell 3, 265–278. Mizumatsu, S., Monje, M.L., Morhardt, D.R., Rola, R., Palmer, T.D., and Fike, J.R. (2003). Extreme Sensitivity of Adult Neurogenesis to Low Doses of X-Irradiation. Cancer Res 63, 4021–4027. Molofsky, A.V., Slutsky, S.G., Joseph, N.M., He, S., Pardal, R., Krishnamurthy, J., Sharpless, N.E., and Morrison, S.J. (2006). Increasing p16INK4a expression decreases forebrain progenitors and neurogenesis during ageing. Nature 443, 448–452. Monje, M.L., Mizumatsu, S., Fike, J.R., and Palmer, T.D. (2002). Irradiation induces neural precursor-cell dysfunction. Nature Medicine 8, 955–962. Monje, M.L., Toda, H., and Palmer, T.D. (2003). Inflammatory blockade restores adult hippocampal neurogenesis. Science 302, 1760–1765. Moreno, M.M., Linster, C., Escanilla, O., Sacquet, J., Didier, A., and Mandairon, N. (2009). Olfactory perceptual learning requires adult neurogenesis. PNAS 106, 17980–17985. Moreno-Jiménez, E.P., Flor-García, M., Terreros-Roncal, J., Rábano, A., Cafini, F., Pallas-Bazarra, N., Ávila, J., and Llorens-Martín, M. (2019). Adult hippocampal neurogenesis is abundant in neurologically healthy subjects and drops sharply in patients with Alzheimer’s disease. Nature Medicine 1. Mori, T., Tanaka, K., Buffo, A., Wurst, W., Kühn, R., and Götz, M. (2006). Inducible gene deletion in astroglia and radial glia—A valuable tool for functional and lineage analysis. Glia 54, 21–34.

117

Morizur, L., Chicheportiche, A., Gauthier, L.R., Daynac, M., Boussin, F.D., and Mouthon, M.-A. (2018). Distinct Molecular Signatures of Quiescent and Activated Adult Neural Stem Cells Reveal Specific Interactions with Their Microenvironment. Stem Cell Reports 11, 565–577. Morshead, C.M., Benveniste, P., Iscove, N.N., and Kooy, D. van der (2002). Hematopoietic competence is a rare property of neural stem cells that may depend on genetic and epigenetic alterations. Nature Medicine 8, 268. Morshead, C.M., Craig, C.G., and van der Kooy, D. (1998). In vivo clonal analyses reveal the properties of endogenous neural stem cell proliferation in the adult mammalian forebrain. Development 125, 2251–2261. Morshead, C.M., Garcia, A.D., Sofroniew, M.V., and Kooy, D.V.D. (2003). The ablation of glial fibrillary acidic protein-positive cells from the adult central nervous system results in the loss of forebrain neural stem cells but not retinal stem cells. European Journal of Neuroscience 18, 76–84. Morshead, C.M., Reynolds, B.A., Craig, C.G., McBurney, M.W., Staines, W.A., Morassutti, D., Weiss, S., and van der Kooy, D. (1994). Neural stem cells in the adult mammalian forebrain: a relatively quiescent subpopulation of subependymal cells. Neuron 13, 1071–1082. Morshead, C.M., and van der Kooy, D. (1992). Postmitotic death is the fate of constitutively proliferating cells in the subependymal layer of the adult mouse brain. J. Neurosci. 12, 249–256. Mosher, K.I., Andres, R.H., Fukuhara, T., Bieri, G., Hasegawa-Moriyama, M., He, Y., Guzman, R., and Wyss-Coray, T. (2012). Neural progenitor cells regulate microglia functions and activity. Nat. Neurosci. 15, 1485–1487. Mouret, A., Gheusi, G., Gabellec, M.-M., de Chaumont, F., Olivo-Marin, J.-C., and Lledo, P.-M. (2008). Learning and survival of newly generated neurons: when time matters. J. Neurosci. 28, 11511–11516. Mullen, R.J., Buck, C.R., and Smith, A.M. (1992). NeuN, a neuronal specific nuclear protein in vertebrates. Development 116, 201–211. Murase, S., and Horwitz, A.F. (2002). Deleted in colorectal carcinoma and differentially expressed integrins mediate the directional migration of neural precursors in the rostral migratory stream. J. Neurosci. 22, 3568–3579. Muthusamy, N., Brumm, A., Zhang, X., Carmichael, S.T., and Ghashghaei, H.T. (2018). Foxj1 expressing ependymal cells do not contribute new cells to sites of injury or stroke in the mouse forebrain. Scientific Reports 8, 1–9. Nadarajah, B., and Parnavelas, J.G. (2002). Modes of neuronal migration in the developing cerebral cortex. Nature Reviews Neuroscience 3, 423–432. Nait-Oumesmar, B., Picard-Riera, N., Kerninon, C., Decker, L., Seilhean, D., Höglinger, G.U., Hirsch, E.C., Reynolds, R., and Evercooren, A.B.-V. (2007). Activation of the

118

subventricular zone in multiple sclerosis: Evidence for early glial progenitors. PNAS 104, 4694–4699. Nguyen-Ba-Charvet, K.T., Picard-Riera, N., Tessier-Lavigne, M., Evercooren, A.B.-V., Sotelo, C., and Chédotal, A. (2004). Multiple Roles for Slits in the Control of Cell Migration in the Rostral Migratory Stream. J. Neurosci. 24, 1497–1506. Noctor, S.C., Flint, A.C., Weissman, T.A., Dammerman, R.S., and Kriegstein, A.R. (2001). Neurons derived from radial glial cells establish radial units in neocortex. Nature 409, 714–720. Noctor, S.C., Martínez-Cerdeño, V., Ivic, L., and Kriegstein, A.R. (2004). Cortical neurons arise in symmetric and asymmetric division zones and migrate through specific phases. Nature Neuroscience 7, 136–144. O’Keeffe, G.C., Tyers, P., Aarsland, D., Dalley, J.W., Barker, R.A., and Caldwell, M.A. (2009). Dopamine-induced proliferation of adult neural precursor cells in the mammalian subventricular zone is mediated through EGF. PNAS 106, 8754–8759. Obernier, K., Cebrian-Silla, A., Thomson, M., Parraguez, J.I., Anderson, R., Guinto, C., Rodas Rodriguez, J., Garcia-Verdugo, J.-M., and Alvarez-Buylla, A. (2018). Adult Neurogenesis Is Sustained by Symmetric Self-Renewal and Differentiation. Cell Stem Cell 22, 221-234.e8. Oboti, L., Schellino, R., Giachino, C., Chamero, P., Pyrski, M., Leinders-Zufall, T., Zufall, F., Fasolo, A., and Peretto, P. (2011). Newborn Interneurons in the Accessory Olfactory Bulb Promote Mate Recognition in Female Mice. Front. Neurosci. 5. Ohata, S., Herranz-Pérez, V., Nakatani, J., Boletta, A., García-Verdugo, J.M., and Álvarez-Buylla, A. (2015). Mechanosensory Genes Pkd1 and Pkd2 Contribute to the Planar Polarization of Brain Ventricular Epithelium. J. Neurosci. 35, 11153–11168. Ohata, S., Nakatani, J., Herranz-Pérez, V., Cheng, J., Belinson, H., Inubushi, T., Snider, W.D., García-Verdugo, J.M., Wynshaw-Boris, A., and Álvarez-Buylla, A. (2014). Loss of Dishevelleds Disrupts Planar Polarity in Ependymal Motile Cilia and Results in Hydrocephalus. Neuron 83, 558–571. Ottone, C., Krusche, B., Whitby, A., Clements, M., Quadrato, G., Pitulescu, M.E., Adams, R.H., and Parrinello, S. (2014). Direct cell–cell contact with the vascular niche maintains quiescent neural stem cells. Nature Cell Biology 16, 1045–1056. Ottone, C., and Parrinello, S. (2015). Multifaceted control of adult SVZ neurogenesis by the vascular niche. Cell Cycle 14, 2222–2225. Owen, M.R., Doran, E., and Halestrap, A.P. (2000). Evidence that metformin exerts its antidiabetic effects through inhibition of complex 1 of the mitochondrial respiratory chain. Biochemical Journal 348, 607–614. Packer, R.J., Goldwein, J., Nicholson, H.S., Vezina, L.G., Allen, J.C., Ris, M.D., Muraszko, K., Rorke, L.B., Wara, W.M., Cohen, B.H., et al. (1999). Treatment of Children With Medulloblastomas With Reduced-Dose Craniospinal Radiation Therapy and Adjuvant Chemotherapy: A Children’s Cancer Group Study. Journal of Clinical Oncology 17, 2127– 2127.

119

Palmer, S.L., Armstrong, C., Onar-Thomas, A., Wu, S., Wallace, D., Bonner, M.J., Schreiber, J., Swain, M., Chapieski, L., Mabbott, D., et al. (2013). Processing speed, attention, and working memory after treatment for medulloblastoma: an international, prospective, and longitudinal study. J. Clin. Oncol. 31, 3494–3500. Palmer, S.L., Reddick, W.E., and Gajjar, A. (2007). Understanding the Cognitive Impact on Children Who are Treated for Medulloblastoma. J Pediatr Psychol 32, 1040–1049. Pan, Y.-W., Chan, G.C.K., Kuo, C.T., Storm, D.R., and Xia, Z. (2012a). Inhibition of adult neurogenesis by inducible and targeted deletion of ERK5 mitogen-activated protein kinase specifically in adult neurogenic regions impairs contextual fear extinction and remote fear memory. J. Neurosci. 32, 6444–6455. Pan, Y.-W., Kuo, C.T., Storm, D.R., and Xia, Z. (2012b). Inducible and Targeted Deletion of the ERK5 MAP Kinase in Adult Neurogenic Regions Impairs Adult Neurogenesis in the Olfactory Bulb and Several Forms of Olfactory Behavior. PLoS One 7. Panagiotakos, G., Alshamy, G., Chan, B., Abrams, R., Greenberg, E., Saxena, A., Bradbury, M., Edgar, M., Gutin, P., and Tabar, V. (2007). Long-Term Impact of Radiation on the Stem Cell and Oligodendrocyte Precursors in the Brain. PLOS ONE 2, e588. Panzanelli, P., Bardy, C., Nissant, A., Pallotto, M., Sassoè-Pognetto, M., Lledo, P.-M., and Fritschy, J.-M. (2009). Early synapse formation in developing interneurons of the adult olfactory bulb. J. Neurosci. 29, 15039–15052. Park, D., Xiang, A.P., Zhang, L., Mao, F.F., Walton, N.M., Choi, S.S., and Lahn, B.T. (2009). The radial glia antibody RC2 recognizes a protein encoded by Nestin. Biochem. Biophys. Res. Commun. 382, 588–592. Parras, C.M., Galli, R., Britz, O., Soares, S., Galichet, C., Battiste, J., Johnson, J.E., Nakafuku, M., Vescovi, A., and Guillemot, F. (2004). Mash1 specifies neurons and oligodendrocytes in the postnatal brain. EMBO J 23, 4495–4505. Pastrana, E., Cheng, L.-C., and Doetsch, F. (2009). Simultaneous prospective purification of adult subventricular zone neural stem cells and their progeny. Proc. Natl. Acad. Sci. U.S.A. 106, 6387–6392. Pastrana, E., Silva-Vargas, V., and Doetsch, F. (2011). Eyes Wide Open: A Critical Review of Sphere-Formation as an Assay for Stem Cells. Cell Stem Cell 8, 486–498. Patel, R.K., Prasad, N., Kuwar, R., Haldar, D., and Abdul-Muneer, P.M. (2017). Transforming growth factor-beta 1 signaling regulates neuroinflammation and apoptosis in mild traumatic brain injury. Brain Behav. Immun. 64, 244–258. Pernicova, I., and Korbonits, M. (2014). Metformin--mode of action and clinical implications for diabetes and cancer. Nat Rev Endocrinol 10, 143–156. Petreanu, L., and Alvarez-Buylla, A. (2002). Maturation and death of adult-born olfactory bulb granule neurons: role of olfaction. J. Neurosci. 22, 6106–6113.

120

Piccin, D., and Morshead, C.M. (2011). Wnt signaling regulates symmetry of division of neural stem cells in the adult brain and in response to injury. Stem Cells 29, 528–538. Piccin, D., Tufford, A., and Morshead, C.M. (2014). Neural stem and progenitor cells in the aged subependyma are activated by the young niche. Neurobiology of Aging 35, 1669–1679. Pineda, J.R., Daynac, M., Chicheportiche, A., Cebrian‐Silla, A., Felice, K.S., Garcia‐Verdugo, J.M., Boussin, F.D., and Mouthon, M.-A. (2013). Vascular‐derived TGF‐β increases in the stem cell niche and perturbs neurogenesis during aging and following irradiation in the adult mouse brain. EMBO Molecular Medicine 5, 548–562. Pixley, S.K.R., and de Vellis, J. (1984). Transition between immature radial glia and mature astrocytes studied with a monoclonal antibody to vimentin. Developmental Brain Research 15, 201–209. Platel, J.-C., Gordon, V., Heintz, T., and Bordey, A. (2009). GFAP-GFP neural progenitors are antigenically homogeneous and anchored in their enclosed mosaic niche. Glia 57, 66– 78. Platel, J.-C., Stamboulian, S., Nguyen, I., and Bordey, A. (2010). Neurotransmitter signaling in postnatal neurogenesis: The first leg. Brain Research Reviews 63, 60–71. Pollard, S.M., Conti, L., Sun, Y., Goffredo, D., and Smith, A. (2006). Adherent Neural Stem (NS) Cells from Fetal and Adult Forebrain. Cereb Cortex 16, i112–i120. Pomeroy, S.L., LaMantia, A.S., and Purves, D. (1990). Postnatal construction of neural circuitry in the mouse olfactory bulb. J. Neurosci. 10, 1952–1966. Ponti, G., Obernier, K., Guinto, C., Jose, L., Bonfanti, L., and Alvarez-Buylla, A. (2013). Cell cycle and lineage progression of neural progenitors in the ventricular-subventricular zones of adult mice. PNAS 110, E1045–E1054. Porlan, E., Martí-Prado, B., Morante-Redolat, J.M., Consiglio, A., Delgado, A.C., Kypta, R., López- Otín, C., Kirstein, M., and Fariñas, I. (2014). MT5-MMP regulates adult neural stem cell functional quiescence through the cleavage of N-cadherin. Nature Cell Biology 16, 629– 638. Potts, M.B., and Lim, D.A. (2012). An old drug for new ideas: metformin promotes adult neurogenesis and spatial memory formation. Cell Stem Cell 11, 5–6. Prineas, J.W., and Connell, F. (1979). Remyelination in multiple sclerosis. Annals of Neurology 5, 22–31. Quaile, M.P., Melich, D.H., Jordan, H.L., Nold, J.B., Chism, J.P., Polli, J.W., Smith, G.A., and Rhodes, M.C. (2010). Toxicity and toxicokinetics of metformin in rats. Toxicology and Applied Pharmacology 243, 340–347. Quiñones-Hinojosa, A., Sanai, N., Soriano-Navarro, M., Gonzalez-Perez, O., Mirzadeh, Z., Gil- Perotin, S., Romero-Rodriguez, R., Berger, M.S., Garcia-Verdugo, J.M., and Alvarez- Buylla, A. (2006). Cellular composition and cytoarchitecture of the adult human

121

subventricular zone: A niche of neural stem cells. The Journal of Comparative Neurology 494, 415–434. Ramírez-Castillejo, C., Sánchez-Sánchez, F., Andreu-Agulló, C., Ferrón, S.R., Aroca-Aguilar, J.D., Sánchez, P., Mira, H., Escribano, J., and Fariñas, I. (2006). Pigment epithelium-derived factor is a niche signal for neural stem cell renewal. Nat. Neurosci. 9, 331–339. Raponi, E., Agenes, F., Delphin, C., Assard, N., Baudier, J., Legraverend, C., and Deloulme, J.-C. (2007). S100B expression defines a state in which GFAP-expressing cells lose their neural stem cell potential and acquire a more mature developmental stage. Glia 55, 165–177. Redmond, S.A., Figueres-Oñate, M., Obernier, K., Nascimento, M.A., Parraguez, J.I., López- Mascaraque, L., Fuentealba, L.C., and Alvarez-Buylla, A. (2019). Development of Ependymal and Postnatal Neural Stem Cells and Their Origin from a Common Embryonic Progenitor. Cell Reports 27, 429-441.e3. Redmond, K.J., Mahone, E.M., Terezakis, S., Ishaq, O., Ford, E., McNutt, T., Kleinberg, L., Cohen, K.J., Wharam, M., and Horska, A. (2013). Association between radiation dose to neuronal progenitor cell niches and temporal lobes and performance on neuropsychological testing in children: a prospective study. Neuro-Oncology 15, 360– 369. Renault, V.M., Rafalski, V.A., Morgan, A.A., Salih, D.A.M., Brett, J.O., Webb, A.E., Villeda, S.A., Thekkat, P.U., Guillerey, C., Denko, N.C., et al. (2009). FoxO3 regulates neural stem cell homeostasis. Cell Stem Cell 5, 527–539. Reynolds, B.A., and Weiss, S. (1992). Generation of neurons and astrocytes from isolated cells of the adult mammalian central nervous system. Science 255, 1707–1711. Reynolds, B.A., and Weiss, S. (1996). Clonal and Population Analyses Demonstrate That an EGF- Responsive Mammalian Embryonic CNS Precursor Is a Stem Cell. Developmental Biology 175, 1–13. Ribeiro Xavier, A.L., Kress, B.T., Goldman, S.A., Lacerda de Menezes, J.R., and Nedergaard, M. (2015). A Distinct Population of Microglia Supports Adult Neurogenesis in the Subventricular Zone. J. Neurosci. 35, 11848–11861. Riquelme, P.A., Drapeau, E., and Doetsch, F. (2008). Brain Micro-Ecologies: Neural Stem Cell Niches in the Adult Mammalian Brain. Philosophical Transactions: Biological Sciences 363, 123–137. Robert, F., Fendri, S., Hary, L., Lacroix, C., Andréjak, M., and Lalau, J. (2003). Kinetics of plasma and erythrocyte metformin after acute administration in healthy subjects. Diabetes & Metabolism 29, 279–283. Rochefort, C., Gheusi, G., Vincent, J.-D., and Lledo, P.-M. (2002). Enriched odor exposure increases the number of newborn neurons in the adult olfactory bulb and improves odor memory. J. Neurosci. 22, 2679–2689.

122

Rosenbluth, J.M., Mays, D.J., Pino, M.F., Tang, L.J., and Pietenpol, J.A. (2008). A Gene Signature- Based Approach Identifies mTOR as a Regulator of p73. Molecular and Cellular Biology 28, 5951–5964. Rousselot, P., Lois, C., and Alvarez‐Buylla, A. (1995). Embryonic (PSA) N-CAM reveals chains of migrationg neuroblasts between the lateral ventricle and the olfactory bulb of adult mice. Journal of Comparative Neurology 351, 51–61. Ruddy, R.M., Adams, K.V., and Morshead, C.M. (2019a). Age- and sex-dependent effects of metformin on neural precursor cells and cognitive recovery in a model of neonatal stroke. Science Advances 5, eaax1912. Ruddy, R.M., Derkach, D., Dadwal, P., and Morshead, C.M. (2019b) Cranial irradiation in juvenile mice leads to early and sustained deficits in the stem and progenitor cell pools and late cognitive impairments. Brain Research, submitted. Ruddy, R.M., and Morshead, C.M. (2018). Home sweet home: the neural stem cell niche throughout development and after injury. Cell Tissue Res 371, 125–141. Saghatelyan, A., de Chevigny, A., Schachner, M., and Lledo, P.-M. (2004). Tenascin-R mediates activity-dependent recruitment of neuroblasts in the adult mouse forebrain. Nat. Neurosci. 7, 347–356. Saha, K., Keung, A.J., Irwin, E.F., Li, Y., Little, L., Schaffer, D.V., and Healy, K.E. (2008). Substrate Modulus Directs Neural Stem Cell Behavior. Biophys J 95, 4426–4438. Sahra, I.B., Regazzetti, C., Robert, G., Laurent, K., Marchand-Brustel, Y.L., Auberger, P., Tanti, J.- F., Giorgetti-Peraldi, S., and Bost, F. (2011). Metformin, Independent of AMPK, Induces mTOR Inhibition and Cell-Cycle Arrest through REDD1. Cancer Res 71, 4366–4372. Saisho, Y. (2015). Metformin and Inflammation: Its Potential Beyond Glucose-lowering Effect. Endocr Metab Immune Disord Drug Targets 15, 196–205. Sakaue-Sawano, A., Kurokawa, H., Morimura, T., Hanyu, A., Hama, H., Osawa, H., Kashiwagi, S., Fukami, K., Miyata, T., Miyoshi, H., et al. (2008). Visualizing Spatiotemporal Dynamics of Multicellular Cell-Cycle Progression. Cell 132, 487–498. Salter, M.W., and Beggs, S. (2014). Sublime microglia: expanding roles for the guardians of the CNS. Cell 158, 15–24. Salter, M.W., and Stevens, B. (2017). Microglia emerge as central players in brain disease. Nature Medicine 23, 1018–1027. Sanai, N., Nguyen, T., Ihrie, R.A., Mirzadeh, Z., Tsai, H.-H., Wong, M., Gupta, N., Berger, M.S., Huang, E., Garcia-Verdugo, J.-M., et al. (2011). Corridors of migrating neurons in the human brain and their decline during infancy. Nature 478, 382–386. Sancho‐Tello, M., Vallés, S., Montoliu, C., Renau‐Piqueras, J., and Guerri, C. (1995). Developmental pattern of GFAP and vimentin gene expression in rat brain and in radial glial cultures. Glia 15, 157–166.

123

Sasaki, A., Hirato, J., Nakazato, Y., and Ishida, Y. (1988). Immunohistochemical study of the early human fetal brain. Acta Neuropathol 76, 128–134. Sato, A., Sunayama, J., Okada, M., Watanabe, E., Seino, S., Shibuya, K., Suzuki, K., Narita, Y., Shibui, S., Kayama, T., et al. (2012). Glioma-Initiating Cell Elimination by Metformin Activation of FOXO3 via AMPK. Stem Cells Transl Med 1, 811–824. Sawamoto, K., Wichterle, H., Gonzalez-Perez, O., Cholfin, J.A., Yamada, M., Spassky, N., Murcia, N.S., Garcia-Verdugo, J.M., Marin, O., Rubenstein, J.L.R., et al. (2006). New neurons follow the flow of cerebrospinal fluid in the adult brain. Science 311, 629–632. Schatz, J., Kramer, J.H., Ablin, A., and Matthay, K.K. (2000). Processing Speed, Working Memory, and IQ: A Developmental Model of Cognitive Deficits Following Cranial Radiation Therapy. Neuropsychology 14, 189–200. Scheen, A.J. (1996). Clinical Pharmacokinetics of Metformin. Clinical Pharmacokinetics 30, 359– 371. Schilling, M., Strecker, J.-K., Ringelstein, E.B., Schäbitz, W.-R., and Kiefer, R. (2009). The role of CC chemokine receptor 2 on microglia activation and blood-borne cell recruitment after transient focal cerebral ischemia in mice. Brain Res. 1289, 79–84. Schindelin, J., Arganda-Carreras, I., Frise, E., Kaynig, V., Longair, M., Pietzsch, T., Preibisch, S., Rueden, C., Saalfeld, S., Schmid, B., et al. (2012). Fiji: an open-source platform for biological-image analysis. Nat. Methods 9, 676–682. Schnitzer, J., Franke, W.W., and Schachner, M. (1981). Immunocytochemical demonstration of vimentin in astrocytes and ependymal cells of developing and adult mouse nervous system. The Journal of Cell Biology 90, 435–447. Seaberg, R.M., and van der Kooy, D. (2002). Adult rodent neurogenic regions: the ventricular subependyma contains neural stem cells, but the dentate gyrus contains restricted progenitors. J. Neurosci. 22, 1784–1793. Shah, P.T., Stratton, J.A., Stykel, M.G., Abbasi, S., Sharma, S., Mayr, K.A., Koblinger, K., Whelan, P.J., and Biernaskie, J. (2018). Single-Cell Transcriptomics and Fate Mapping of Ependymal Cells Reveals an Absence of Neural Stem Cell Function. Cell 173, 1045- 1057.e9. Sharlow, E.R., Leimgruber, S., Lira, A., McConnell, M.J., Norambuena, A., Bloom, G.S., Epperly, M.W., Greenberger, J.S., and Lazo, J.S. (2016). A Small Molecule Screen Exposes mTOR Signaling Pathway Involvement in Radiation-Induced Apoptosis. ACS Chemical Biology 11, 1428–1437. Shen, Q., Goderie, S.K., Jin, L., Karanth, N., Sun, Y., Abramova, N., Vincent, P., Pumiglia, K., and Temple, S. (2004). Endothelial cells stimulate self-renewal and expand neurogenesis of neural stem cells. Science 304, 1338–1340. Shen, Q., Wang, Y., Kokovay, E., Lin, G., Chuang, S.-M., Goderie, S.K., Roysam, B., and Temple, S. (2008). Adult SVZ Stem Cells Lie in a Vascular Niche: A Quantitative Analysis of Niche Cell-Cell Interactions. Cell Stem Cell 3, 289–300.

124

Shi, M., Wei, L.-C., Cao, R., and Chen, L.-W. (2002). Enhancement of nestin protein- immunoreactivity induced by ionizing radiation in the forebrain ependymal regions of rats. Neuroscience Research 44, 475–481. Shigemoto-Mogami, Y., Hoshikawa, K., Goldman, J.E., Sekino, Y., and Sato, K. (2014). Microglia enhance neurogenesis and oligodendrogenesis in the early postnatal subventricular zone. J. Neurosci. 34, 2231–2243. Shin, J., Berg, D.A., Zhu, Y., Shin, J.Y., Song, J., Bonaguidi, M.A., Enikolopov, G., Nauen, D.W., Christian, K.M., Ming, G., et al. (2015). Single-Cell RNA-Seq with Waterfall Reveals Molecular Cascades underlying Adult Neurogenesis. Cell Stem Cell 17, 360–372. Shinohara, C., Gobbel, G.T., Lamborn, K.R., Tada, E., and Fike, J.R. (1997). Apoptosis in the subependyma of young adult rats after single and fractionated doses of X-rays. Cancer Res. 57, 2694–2702. Shook, B.A., Manz, D.H., Peters, J.J., Kang, S., and Conover, J.C. (2012). Spatiotemporal Changes to the Subventricular Zone Stem Cell Pool through Aging. J. Neurosci. 32, 6947–6956. Sierra, A., Encinas, J.M., Deudero, J.J.P., Chancey, J.H., Enikolopov, G., Overstreet-Wadiche, L.S., Tsirka, S.E., and Maletic-Savatic, M. (2010). Microglia Shape Adult Hippocampal Neurogenesis through Apoptosis-Coupled Phagocytosis. Cell Stem Cell 7, 483–495. Silva-Vargas, V., Maldonado-Soto, A.R., Mizrak, D., Codega, P., and Doetsch, F. (2016). Age- Dependent Niche Signals from the Choroid Plexus Regulate Adult Neural Stem Cells. Cell Stem Cell 19, 643–652. Singhal, G., Jaehne, E.J., Corrigan, F., Toben, C., and Baune, B.T. (2014). Inflammasomes in neuroinflammation and changes in brain function: a focused review. Front. Neurosci. 8. Snapyan, M., Lemasson, M., Brill, M.S., Blais, M., Massouh, M., Ninkovic, J., Gravel, C., Berthod, F., Götz, M., Barker, P.A., et al. (2009). Vasculature guides migrating neuronal precursors in the adult mammalian forebrain via brain-derived neurotrophic factor signaling. J. Neurosci. 29, 4172–4188. Solano Fonseca, R., Mahesula, S., Apple, D.M., Raghunathan, R., Dugan, A., Cardona, A., O’Connor, J., and Kokovay, E. (2016). Neurogenic Niche Microglia Undergo Positional Remodeling and Progressive Activation Contributing to Age-Associated Reductions in Neurogenesis. Stem Cells and Development 25, 542–555. Sorrells, S.F., Paredes, M.F., Cebrian-Silla, A., Sandoval, K., Qi, D., Kelley, K.W., James, D., Mayer, S., Chang, J., Auguste, K.I., et al. (2018). Human hippocampal neurogenesis drops sharply in children to undetectable levels in adults. Nature 555, 377–381. Spassky, N., Merkle, F.T., Flames, N., Tramontin, A.D., García-Verdugo, J.M., and Alvarez-Buylla, A. (2005). Adult Ependymal Cells Are Postmitotic and Are Derived from Radial Glial Cells during Embryogenesis. J. Neurosci. 25, 10–18. Spiegler, B.J., Bouffet, E., Greenberg, M.L., Rutka, J.T., and Mabbott, D.J. (2004). Change in neurocognitive functioning after treatment with cranial radiation in childhood. J. Clin. Oncol. 22, 706–713.

125

Stagaard, M., and Møllgård, K. (1989). The developing neuroepithelium in human embryonic and fetal brain studied with vimentin-immunocytochemistry. Anat Embryol 180, 17–28. Steward, O., Sharp, K.G., and Matsudaira Yee, K. (2014). Long-Distance Migration and Colonization of Transplanted Neural Stem Cells. Cell 156, 385–387. Stoll, E.A., Habibi, B.A., Mikheev, A.M., Lasiene, J., Massey, S.C., Swanson, K.R., Rostomily, R.C., and Horner, P.J. (2011). Increased Re-Entry into Cell Cycle Mitigates Age-Related Neurogenic Decline in the Murine Subventricular Zone. STEM CELLS 29, 2005–2017. Su, P., Zhang, J., Zhao, F., Aschner, M., Chen, J., and Luo, W. (2014). The interaction between microglia and neural stem/precursor cells. Brain Research Bulletin 109, 32–38. Sultan, S., Mandairon, N., Kermen, F., Garcia, S., Sacquet, J., and Didier, A. (2010). Learning- dependent neurogenesis in the olfactory bulb determines long-term olfactory memory. The FASEB Journal 24, 2355–2363. Sun, Y., Kong, W., Falk, A., Hu, J., Zhou, L., Pollard, S., and Smith, A. (2009). CD133 (Prominin) negative human neural stem cells are clonogenic and tripotent. PLoS ONE 4, e5498. Tada, E., Yang, C., Gobbel, G.T., Lamborn, K.R., and Fike, J.R. (1999). Long-term impairment of subependymal repopulation following damage by ionizing irradiation. Exp. Neurol. 160, 66–77. Takahashi, T., Misson, J.P., and Caviness, V.S. (1990). Glial process elongation and branching in the developing murine neocortex: a qualitative and quantitative immunohistochemical analysis. J. Comp. Neurol. 302, 15–28. Tavazoie, M., Van der Veken, L., Silva-Vargas, V., Louissaint, M., Colonna, L., Zaidi, B., Garcia- Verdugo, J.M., and Doetsch, F. (2008). A Specialized Vascular Niche for Adult Neural Stem Cells. Cell Stem Cell 3, 279–288. Thibeault, I., Laflamme, N., and Rivest, S. (2001). Regulation of the gene encoding the monocyte chemoattractant protein 1 (MCP-1) in the mouse and rat brain in response to circulating LPS and proinflammatory cytokines. J. Comp. Neurol. 434, 461–477. Thored, P., Heldmann, U., Gomes‐Leal, W., Gisler, R., Darsalia, V., Taneera, J., Nygren, J.M., Jacobsen, S.-E.W., Ekdahl, C.T., Kokaia, Z., et al. (2009). Long-term accumulation of microglia with proneurogenic phenotype concomitant with persistent neurogenesis in adult subventricular zone after stroke. Glia 57, 835–849. Thored, P., Wood, J., Arvidsson, A., Cammenga, J., Kokaia, Z., and Lindvall, O. (2007). Long-term neuroblast migration along blood vessels in an area with transient angiogenesis and increased vascularization after stroke. Stroke 38, 3032–3039. Tong, C.K., Chen, J., Cebrián-Silla, A., Mirzadeh, Z., Obernier, K., Guinto, C.D., Tecott, L.H., García-Verdugo, J.M., Kriegstein, A., and Alvarez-Buylla, A. (2014). Axonal Control of the Adult Neural Stem Cell Niche. Cell Stem Cell 14, 500–511.

126

Tonning Olsson, I., Perrin, S., Lundgren, J., Hjorth, L., and Johanson, A. (2014). Long-term cognitive sequelae after pediatric brain tumor related to medical risk factors, age, and sex. Pediatr. Neurol. 51, 515–521. Tramontin, A.D., García-Verdugo, J.M., Lim, D.A., and Alvarez-Buylla, A. (2003). Postnatal Development of Radial Glia and the Ventricular Zone (VZ): a Continuum of the Neural Stem Cell Compartment. Cereb Cortex 13, 580–587. Tropepe, V., Craig, C.G., Morshead, C.M., and van der Kooy, D. (1997). Transforming growth factor-alpha null and senescent mice show decreased neural progenitor cell proliferation in the forebrain subependyma. J. Neurosci. 17, 7850–7859. Tucker, G.T., Casey, C., Phillips, P.J., Connor, H., Ward, J.D., and Woods, H.F. (1981). Metformin kinetics in healthy subjects and in patients with diabetes mellitus. Br J Clin Pharmacol 12, 235–246. Valley, M., Mullen, T.R., Schultz, L., Sagdullaev, B.T., and Firestein, S. (2009). Ablation of mouse adult neurogenesis alters olfactory bulb structure and olfactory fear conditioning. Front. Neurosci. 3. Vignard, J., Mirey, G., and Salles, B. (2013). Ionizing-radiation induced DNA double-strand breaks: A direct and indirect lighting up. Radiotherapy and Oncology 108, 362–369. Viollet, B., Guigas, B., Sanz Garcia, N., Leclerc, J., Foretz, M., and Andreelli, F. (2012). Cellular and molecular mechanisms of metformin: an overview. Clin. Sci. 122, 253–270. Walker, T.L., and Kempermann, G. (2014). One Mouse, Two Cultures: Isolation and Culture of Adult Neural Stem Cells from the Two Neurogenic Zones of Individual Mice. J Vis Exp. Walton, N.M., Sutter, B.M., Laywell, E.D., Levkoff, L.H., Kearns, S.M., Marshall, G.P., Scheffler, B., and Steindler, D.A. (2006). Microglia instruct subventricular zone neurogenesis. Glia 54, 815–825. Wang, C., Liu, F., Liu, Y.-Y., Zhao, C.-H., You, Y., Wang, L., Zhang, J., Wei, B., Ma, T., Zhang, Q., et al. (2011). Identification and characterization of neuroblasts in the subventricular zone and rostral migratory stream of the adult human brain. Cell Research 21, 1534–1550. Wang, D.D., Krueger, D.D., and Bordey, A. (2003). GABA Depolarizes Neuronal Progenitors of the Postnatal Subventricular Zone Via GABAA Receptor Activation. The Journal of Physiology 550, 785–800. Wang, J., Gallagher, D., DeVito, L.M., Cancino, G.I., Tsui, D., He, L., Keller, G.M., Frankland, P.W., Kaplan, D.R., and Miller, F.D. (2012). Metformin activates an atypical PKC-CBP pathway to promote neurogenesis and enhance spatial memory formation. Cell Stem Cell 11, 23– 35. Wang, J., Weaver, I.C.G., Gauthier-Fisher, A., Wang, H., He, L., Yeomans, J., Wondisford, F.E., Kaplan, D.R., and Miller, F.D. (2010). CBP histone acetyltransferase activity regulates embryonic neural differentiation in the normal and Rubinstein-Taybi syndrome brain. Developmental Cell 18, 114–125.

127

Wang, J., Yin, L., and Chen, Z. (2011). New insights into the altered fibronectin matrix and extrasynaptic transmission in the aging brain. Journal of Clinical Gerontology and Geriatrics 2, 35–41. Webb, A.E., Pollina, E.A., Vierbuchen, T., Urbán, N., Ucar, D., Leeman, D.S., Martynoga, B., Sewak, M., Rando, T.A., Guillemot, F., et al. (2013). FOXO3 Shares Common Targets with ASCL1 Genome-wide and Inhibits ASCL1-Dependent Neurogenesis. Cell Reports 4, 477– 491. Whitman, M.C., Fan, W., Rela, L., Rodriguez-Gil, D.J., and Greer, C.A. (2009). Blood vessels form a migratory scaffold in the rostral migratory stream. J. Comp. Neurol. 516, 94–104. Whitman, M.C., and Greer, C.A. (2009). Adult Neurogenesis and the Olfactory System. Prog Neurobiol 89, 162–175. Wilcock, C., and Bailey, C.J. (1994). Accumulation of metformin by tissues of the normal and diabetic mouse. Xenobiotica 24, 49–57. Wilson, S.W., and Rubenstein, J.L.R. (2000). Induction and Dorsoventral Patterning of the Telencephalon. Neuron 28, 641–651. Winner, B., Desplats, P., Hagl, C., Klucken, J., Aigner, R., Ploetz, S., Laemke, J., Karl, A., Aigner, L., Masliah, E., et al. (2009). Dopamine receptor activation promotes adult neurogenesis in an acute Parkinson model. Experimental Neurology 219, 543–552. Won, S.J., Kim, S.H., Xie, L., Wang, Y., Mao, X.O., Jin, K., and Greenberg, D.A. (2006). Reelin- deficient mice show impaired neurogenesis and increased stroke size. Experimental Neurology 198, 250–259. Wong, C.S., and Van der Kogel, A.J. (2004). Mechanisms of radiation injury to the central nervous system: implications for neuroprotection. Molecular Interventions 4, 273–284. Xing, Y.L., Röth, P.T., Stratton, J.A.S., Chuang, B.H.A., Danne, J., Ellis, S.L., Ng, S.W., Kilpatrick, T.J., and Merson, T.D. (2014). Adult Neural Precursor Cells from the Subventricular Zone Contribute Significantly to Oligodendrocyte Regeneration and Remyelination. J. Neurosci. 34, 14128–14146. Yamaguchi, M., and Mori, K. (2005). Critical period for sensory experience-dependent survival of newly generated granule cells in the adult mouse olfactory bulb. Proc. Natl. Acad. Sci. U.S.A. 102, 9697–9702. Yamashita, T., Ninomiya, M., Acosta, P.H., García-Verdugo, J.M., Sunabori, T., Sakaguchi, M., Adachi, K., Kojima, T., Hirota, Y., Kawase, T., et al. (2006). Subventricular Zone-Derived Neuroblasts Migrate and Differentiate into Mature Neurons in the Post-Stroke Adult Striatum. J. Neurosci. 26, 6627–6636. Yan, Y.-P., Sailor, K.A., Vemuganti, R., and Dempsey, R.J. (2006). Insulin-like growth factor-1 is an endogenous mediator of focal ischemia-induced neural progenitor proliferation. Eur. J. Neurosci. 24, 45–54.

128

Yoo, D.Y., Kim, W., Nam, S.M., Yoo, K.-Y., Lee, C.H., Choi, J.H., Won, M.-H., Hwang, I.K., and Yoon, Y.S. (2011). Reduced cell proliferation and neuroblast differentiation in the dentate gyrus of high fat diet-fed mice are ameliorated by metformin and glimepiride treatment. Neurochem. Res. 36, 2401–2408. Young, K.M., Fogarty, M., Kessaris, N., and Richardson, W.D. (2007). Subventricular Zone Stem Cells Are Heterogeneous with Respect to Their Embryonic Origins and Neurogenic Fates in the Adult Olfactory Bulb. J. Neurosci. 27, 8286–8296. Yuan, R., Wang, Y., Li, Q., Zhen, F., Li, X., Lai, Q., Hu, P., Wang, X., Zhu, Y., Fan, H., et al. (2019). Metformin reduces neuronal damage and promotes neuroblast proliferation and differentiation in a cerebral ischemia/reperfusion rat model: NeuroReport 30, 232–240. Yuzwa, S.A., Borrett, M.J., Innes, B.T., Voronova, A., Ketela, T., Kaplan, D.R., Bader, G.D., and Miller, F.D. (2017). Developmental Emergence of Adult Neural Stem Cells as Revealed by Single-Cell Transcriptional Profiling. Cell Reports 21, 3970–3986. Zecevic, N. (2004). Specific characteristic of radial glia in the human fetal telencephalon. Glia 48, 27–35. Zhang, R., Chopp, M., Gregg, S.R., Toh, Y., Roberts, C., LeTourneau, Y., Buller, B., Jia, L., Davarani, S.P.N., and Zhang, Z.G. (2009). Patterns and dynamics of subventricular zone neuroblast migration in the ischemic striatum of the adult mouse. J Cereb Blood Flow Metab 29, 1240–1250. Zhang, R., Zhang, Z., Wang, L., Wang, Y., Gousev, A., Zhang, L., Ho, K.-L., Morshead, C., and Chopp, M. (2004a). Activated Neural Stem Cells Contribute to Stroke-Induced Neurogenesis and Neuroblast Migration toward the Infarct Boundary in Adult Rats. J Cereb Blood Flow Metab 24, 441–448. Zhang, R., Zhang, Z., Zhang, C., Zhang, L., Robin, A., Wang, Y., Lu, M., and Chopp, M. (2004b). Stroke Transiently Increases Subventricular Zone Cell Division from Asymmetric to Symmetric and Increases Neuronal Differentiation in the Adult Rat. J. Neurosci. 24, 5810–5815. Zheng, H., Wang, M., Wu, J., Wang, Z.-M., Nan, H.-J., and Sun, H. (2016). Inhibition of mTOR enhances radiosensitivity of lung cancer cells and protects normal lung cells against radiation. Biochem. Cell Biol. 94, 213–220. Zhou, G., Myers, R., Li, Y., Chen, Y., Shen, X., Fenyk-Melody, J., Wu, M., Ventre, J., Doebber, T., Fujii, N., et al. (2001). Role of AMP-activated protein kinase in mechanism of metformin action. J Clin Invest 108, 1167–1174. Zhou, M., Li, W., Huang, S., Song, J., Kim, J.Y., Tian, X., Kang, E., Sano, Y., Liu, C., Balaji, J., et al. (2013). mTOR Inhibition Ameliorates Cognitive and Affective Deficits Caused by Disc1 Knockdown in Adult-Born Dentate Granule Neurons. Neuron 77, 647–654. Zhou, X.Y., Shibusawa, N., Naik, K., Porras, D., Temple, K., Ou, H., Kaihara, K., Roe, M.W., Brady, M.J., and Wondisford, F.E. (2004). Insulin regulation of hepatic gluconeogenesis through phosphorylation of CREB-binding protein. Nature Medicine 10, 633.

129

Zhu, W., Fan, Y., Frenzel, T., Gasmi, M., Bartus, R.T., Young, W.L., Yang, G.-Y., and Chen, Y. (2008). Insulin Growth Factor-1 Gene Transfer Enhances Neurovascular Remodeling and Improves Long-Term Stroke Outcome in Mice. Stroke 39, 1254–1261. Zhuo, L., Sun, B., Zhang, C.L., Fine, A., Chiu, S.Y., and Messing, A. (1997). Live astrocytes visualized by green fluorescent protein in transgenic mice. Dev. Biol. 187, 36–42. Zigova, T., Betarbet, R., Soteres, B.J., Brock, S., Bakay, R.A., and Luskin, M.B. (1996). A comparison of the patterns of migration and the destinations of homotopically transplanted neonatal subventricular zone cells and heterotopically transplanted telencephalic ventricular zone cells. Dev. Biol. 173, 459–474. Zigova, T., Pencea, V., Wiegand, S.J., and Luskin, M.B. (1998). Intraventricular Administration of BDNF Increases the Number of Newly Generated Neurons in the Adult Olfactory Bulb. Molecular and Cellular Neuroscience 11, 234–245. Zou, J., Pan, Y.-W., Wang, Z., Chang, S.-Y., Wang, W., Wang, X., Tournier, C., Storm, D.R., and Xia, Z. (2012). Targeted Deletion of ERK5 MAP Kinase in the Developing Nervous System Impairs Development of GABAergic Interneurons in the Main Olfactory Bulb and Behavioral Discrimination between Structurally Similar Odorants. J Neurosci 32, 4118– 4132. Zou, J., Wang, W., Pan, Y.-W., Lu, S., and Xia, Z. (2015). Methods to measure olfactory behavior in mice. Curr Protoc Toxicol 63, 11.18.1-11.18.21.

130

Appendices Appendix 1: Artificial cerebrospinal fluid (aCSF)

Appendix 1A: Regular aCSF

For 100mL regular aCSF:

− 72.4mL ddH2O − 6.2mL 2M NaCl (Sigma-Aldrich, S5886) − 0.5mL 1M KCl

− 0.13mL 1M MgCl2•6H2O (Sigma-Aldrich, M2393)

− 16.9mL 155mM NaHCO3 (Gibco, 25080094) − 1mL 1M glucose (Sigma-Aldrich, G6152)

− 1.852mL 108mM CaCl2 − 1mL (5,000 units) penicillin-streptomycin (Gibco, 15070063)

Appendix 1B: Hi/lo aCSF

For 100mL Hi/Lo aCSF:

− 74mL ddH2O − 6.2mL 2M NaCl (Sigma-Aldrich, S5886) − 0.5mL 1M KCl

− 0.32mL 1M MgCl2•6H2O (Sigma-Aldrich, M2393)

− 16.9mL 155mM NaHCO3 (Gibco, 25080094) − 1mL 1M glucose (Sigma-Aldrich, G6152)

− 0.09256mL 108mM CaCl2 − 1mL (5,000 units) penicillin-streptomycin (Gibco, 15070063)

131

Appendix 2: Serum-free media (SFM)

For 100mL SFM:

− 75mL ddH2O − 10mL 10x DMEM/F12 (DMEM from Gibco, 12100046; F12 from Gibco, 21700075) − 2mL 30% glucose (Sigma-Aldrich, G6152)

− 1.5mL 7.5% NaHCO3 (Gibco, 25080094) − 0.5mL 1M HEPES (Gibco, 15630080) − 1mL 200mM L-glutamine (Gibco, 25030081) − 1mL (5,000 units) penicillin-streptomycin (Gibco, 15070063) − 10mL hormone mix − To make 1L hormone mix stock solution:

▪ 750mL ddH2O ▪ 100mL 10x DMEM/F12 (DMEM from Gibco, 12100046; F12 from Gibco, 21700075) ▪ 20mL 30% glucose (Sigma-Aldrich, G6152)

▪ 15mL 7.5% NaHCO3 (Gibco, 25080094) ▪ 5mL 1M HEPES (Gibco, 15630080) ▪ 1g transferrin (Sigma-Aldrich, T2252) and 96.1mg putrescine dissolved in

40mL ddH2O ▪ 250mg insulin (Wisent Bioproducts, 511-016-CM) dissolved in 5mL 0.1N

HCl (BioShop, HCL666), added to 35mL ddH2O ▪ 100µL selenium (Sigma-Aldrich, S9133) ▪ 100µL progesterone (Sigma-Aldrich, P6149)

132

Appendix 3: Reagents and solutions for FACS

Appendix 3A: Stock solutions

For 50mL 10x Salt Solution (stored at room temperature):

− 50mL ddH2O − 3.5g NaCl (Sigma-Aldrich, S5886) − 0.186g KCl

For 50mL 1M PIPES Stock Solution (pH adjusted to 7.5; stored at room temperature):

− 50mL ddH2O − 15.1g PIPES (Sigma-Aldrich, P1851)

For 50mL 30% Glucose Solution (stored at 4oC):

− 50mL ddH2O − 15g glucose (Sigma-Aldrich, G6152)

For 5mL Ovomucoid Solution (7mg/mL; stored at -20oC):

− 5mL ddH2O − 35mg ovomucoid (Worthington, LS003086)

For 5mL DNase Solution (15mg/mL; stored at -20oC):

− 5mL ddH2O − 75mg DNase (Worthington, LK003172)

Appendix 3B: Working solutions

For 50mL Papain Activating Solution (stored at room temperature):

− 50mL ddH2O − 43.3mg cysteine HCl − 50µL 0.5M EDTA (Invitrogen, AM9260G)

133

For 500mL Staining Solution (1% BSA, 0.1% glucose; stored at 4oC):

− 484mL HBSS (Gibco, 14185052) − 1.666mL 30% Glucose Solution − 14.33mL 35% bovine serum albumin (BSA; Sigma-Aldrich, A7979)

For 50mL 22% Percoll Solution (stored at 4oC):

− 34mL ddH2O − 11mL Percoll (Sigma-Aldrich, P4937) − 5mL 10x PBS

For 50mL PIPES Solution (stored on ice):

− 42mL ddH2O − 1mL 1M PIPES Stock Solution − 5mL 10x Salt Solution − 750µL 30% Glucose Solution − 500µL antibiotic-antimycotic (Sigma-Aldrich, A5955) − 50µL phenol red (pH adjusted until solution is orange-pink colour; pH 7.2-7.5)

134

Figure 1.1: Reprinted from Annual Review of Neuroscience, Vol. 32, A. Kriegstein and A. Álvarez-Buylla, The glial nature of embryonic and adult neural stem cells, pp. 149-184, Copyright 2009, with permission from Annual Reviews and Copyright Clearance Center. License ID: 4644911431098

Figure 1.2: Reprinted from Development, Vol. 142, R. Fiorelli, K. Azim, B. Fischer, and O. Raineteau, Adding a spatial dimension to postnatal ventricular-subventricular zone neurogenesis, pp. 2109-2120, Copyright 2015, with permission from Company of Biologists and Copyright Clearance Center. License ID: 4644931010525

Figure 1.3: Reprinted from Neuron, Vol. 70, R. A. Ihrie and A. Álvarez-Buylla, Lake-front property: a unique germinal niche by the lateral ventricles of the adult brain, pp. 674-686, Copyright 2011, with permission from Elsevier and Copyright Clearance Center. License ID: 4644931181466

Figure 1.4: Copyright 2016 Z. Chaker, P. Codega, and F. Doetsch. This is an open access article available under the terms of the Creative Commons Attribution Non-Commercial License CC BY- NC and permits non-commercial use, distribution, and reproduction in any medium, provided the original work is properly cited.

Figure 1.5: Reprinted from Cell Stem Cell, Vol. 11, M. B. Potts and D. A. Lim, An old drug for new ideas: metformin promotes adult neurogenesis and spatial memory formation, pp. 5-6, Copyright 2012, with permission from Elsevier and Copyright Clearance Center. License ID: 4644940497038