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Tracking neuronal development in the adult brain Thèse Karen Bakhshetyan Doctorat en biophotonique Philosophiæ doctor (Ph. D.) Québec, Canada © Karen Bakhshetyan, 2017

Tracking neuronal development in the adult brain€¦ · neuronal migration and is defined by transmembrane Cl-gradient. This, in turn is controlled by the Cl-extruding co-transporter

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Page 1: Tracking neuronal development in the adult brain€¦ · neuronal migration and is defined by transmembrane Cl-gradient. This, in turn is controlled by the Cl-extruding co-transporter

Tracking neuronal development in the adult brain

Thèse

Karen Bakhshetyan

Doctorat en biophotonique

Philosophiæ doctor (Ph. D.)

Québec, Canada

© Karen Bakhshetyan, 2017

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Tracking neuronal development in the adult brain

Thèse

Karen Bakhshetyan

Sous la direction de:

Armen Saghatelyan, directeur de recherche

Tigran Galstian, codirecteur de recherche

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Résumé

Les connaissances des voies moléculaires et cellulaires régulant le développement neuronal

dans le cerveau adulte peuvent être utilisées pour mettre au point des stratégies efficaces de

thérapie de remplacement cellulaire. L’étude de la dynamique et des mécanismes

nécessaires à la neurogenèse adulte, requiert des techniques d’imagerie en temps réel. En

outre, il est important de développer des méthodes d'imagerie sans marqueurs.

Mon travail vise, en partie, à relever ces défis. Les néo-neurones générés chez l’adulte

migrent densément le long des vaisseaux sanguins et des tubes gliaux dans le courant de

migration rostral (CMR). Cet alignement peut créer une anisotropie qui peut être détectée

en lumière polarisée. J'ai d'abord essayé cette technique pour la détection sans marqueurs

des cellules migratrices dans le CMR. Bien que l’imagerie avec la lumière polarisée suscite

certaines espérances, elle a toutefois fait apparaître que l'anisotropie des cellules

migratrices est très faible et que sa détection est entravée par des signaux de fortes

intensités provenant des axones myélinisés se trouvant à proximité.

Ensuite, j'ai étudié la migration des neuroblastes marqués viralement pour élucider certains

mécanismes nécessaires à leur migration. La signalisation GABAergique joue un rôle

important dans la migration neuronale, déterminée par le gradient chlorique trans-

membranaire. Ce dernier est, à son tour, contrôlé par KCC2, un co-transporteur responsable

de l'extrusion de Cl-. Il est connu que KCC2 est exprimé dans les stades de développement

plus avancés. Toutefois, le rôle de KCC2 dans la migration neuronale est inconnu et mes

expériences suggèrent que ce co-transporteur est impliqué dans la migration radiale, mais

pas tangentielle, de neuroblastes.

Enfin, j'ai exploré in vivo comment la plasticité structurelle des néo-neurones générés chez

l’adulte dans le bulbe olfactif (BO) est modulée par les odeurs. On ne sait pas comment le

fonctionnement du BO s’ajuste à l’environnement olfactif de facon rapide lorsque de

nouvelles synapses de néo-neurones ne sont pas encore formées. Mes données in vivo

d'imagerie à deux photons complètent les travaux antérieurs de notre laboratoire, révélant

une nouvelle forme de plasticité structurale dans le cerveau adulte.

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Ainsi, en utilisant diverses méthodes d'imagerie j'ai essayé de mieux comprendre la

migration et la plasticité des nouveaux neurones dans le cerveau adulte.

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Abstract

The knowledge about molecular and cellular pathways orchestrating neuronal development

in the adult brain can be used to build up efficient strategies for cell replacement therapies.

Adult neurogenesis is a very dynamic process, and it is crucial to monitor it directly to

decipher mechanisms required for neuronal development. Furthermore, it is important to

develop label-free imaging methods.

My work is, in part, aimed at addressing these challenges. Adult-born neurons migrate

densely along blood vessels and glial tubes in the rostral migratory stream (RMS). This

alignment may create anisotropy which can be detected in polarized light. I first tried this

technique for label-free detection of migratory cells in the RMS. While this imaging may

have some promises, it showed that anisotropy in migrating cells is quite low and its

detection is hampered by large signals deriving from nearby myelinated axons.

I further studied the migration of virally labeled neuroblasts to elucidate some of the

mechanisms required for their migration. GABAergic signaling plays an important role in

neuronal migration and is defined by transmembrane Cl- gradient. This, in turn is controlled

by the Cl- extruding co-transporter KCC2, known to have a late developmental expression.

The role of KCC2 in neuronal migration is unknown and my experiments suggest that this

co-transporter is involved in the radial, but not tangential migration of neuroblasts.

Finally, I explored in vivo the odor-related structural plasticity of adult-born neurons in the

olfactory bulb (OB). It remains unknown how OB functioning is adjusted to rapidly

changing odor environment when new synapses of adult-born neurons have not yet been

formed. My in vivo two-photon imaging data complements the previous work in our lab,

revealing altogether a new form of structural plasticity in the adult OB.

Thus, using diverse imaging methods I tried to better understand the migration and

plasticity of new neurons in the adult brain.

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Table of Contents

Résumé .................................................................................................................................. iii

Abstract ................................................................................................................................... v

Table of Contents .................................................................................................................. vi

List of tables ........................................................................................................................... x

List of figures ........................................................................................................................ xi

List of abbreviations ............................................................................................................ xiv

Acknowledgements ............................................................................................................ xvii

Foreword ............................................................................................................................ xviii

1. Introduction ..................................................................................................................... 1

1.1. Adult Neurogenesis .................................................................................................. 1

1.1.1. Neuronal development in adult rodent brain ..................................................... 1

1.1.2. The olfactory system ......................................................................................... 4

1.1.3. Proliferation of neuronal precursors in the SVZ ............................................... 9

1.1.4. Migration of neuroblasts in the adult brain ..................................................... 10

1.1.5. Maturation and integration of adult-born interneurons in the OB .................. 12

1.1.6. Synaptic development of adult-born GCs ....................................................... 15

1.1.7. Adult neurogenesis as a form of structural plasticity to adjust OB

network to changing environmental conditions .............................................. 17

1.2. Overview of GABAergic signaling in the brain ..................................................... 19

1.2.1. The role of GABA in neuronal development during embryogenesis and

in adulthood ..................................................................................................... 20

1.2.2. Cation-chloride cotransporters ........................................................................ 23

1.2.3. The role of KCC2 in brain development ......................................................... 24

1.2.4. The role of KCC2 and Cl- gradient in neuronal migration .............................. 27

1.3. Tools for imaging neuronal development .............................................................. 29

1.3.1. Fundamentals of optical microscopy............................................................... 29

1.3.2. Wide-field and two-photon imaging ............................................................... 33

1.3.3. Light polarization ............................................................................................ 36

1.3.4. Label-free imaging using polarized light ........................................................ 38

1.3.5. Polarized light imaging in biomedical applications ........................................ 40

1.4. Neurodegenerative disorders. Parkinson’s disease. ............................................... 41

2. Thesis objectives ........................................................................................................... 43

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3. Results - Comparative study of myelinated fiber bundles with polarized light

imaging under normal and pathological conditions ...................................................... 45

3.1. Résumé ................................................................................................................... 46

3.2. Abstract .................................................................................................................. 47

3.3. Introduction ............................................................................................................ 48

3.4. Materials and methods............................................................................................ 53

3.4.1. Animals ........................................................................................................... 53

3.4.2. Preparation of human tissue ............................................................................ 53

3.4.3. Myelin staining ................................................................................................ 54

3.4.4. Immunohistochemistry .................................................................................... 54

3.4.5. Image acquisition and processing ................................................................... 54

3.4.6. Data analysis ................................................................................................... 55

3.5. Results .................................................................................................................... 56

3.6. Discussion .............................................................................................................. 58

3.7. References .............................................................................................................. 60

3.8. Figures .................................................................................................................... 81

4. Results - Activation of KCC2 affects radial but not tangential migration of

neuronal precursors in the adult brain ........................................................................... 87

4.1. Résumé ................................................................................................................... 88

4.2. Abstract .................................................................................................................. 89

4.3. Introduction ............................................................................................................ 90

4.4. Materials and Methods ........................................................................................... 91

4.4.1. Animals ........................................................................................................... 92

4.4.2. Stereotaxic injections ...................................................................................... 92

4.4.3. Immunohistochemistry .................................................................................... 92

4.4.4. Slice preparation and time-lapse video-imaging ............................................. 93

4.4.5. Statistical analysis ........................................................................................... 93

4.5. Results .................................................................................................................... 94

4.5.1. KCC2 expression in the SVZ-OB pathway ..................................................... 94

4.5.2. KCC2 activation does not affect tangential migration in the RMS ................. 94

4.5.3. KCC2 activation fosters radial migration of neuroblasts in the OB ............... 95

4.6. Discussion .............................................................................................................. 96

4.7. References .............................................................................................................. 99

4.8. Figures .................................................................................................................. 101

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5. Results - Principal cell activity induces spine relocation of adult-born interneurons

in the olfactory bulb .................................................................................................... 107

5.1. Résumé ................................................................................................................. 109

5.2. Abstract ................................................................................................................ 110

5.3. Introduction .......................................................................................................... 111

5.4. Results .................................................................................................................. 112

5.4.1. Dendritic spines of adult-born GC relocate in the OB network .................... 112

5.4.2. Spine relocation is preceded by spine head filopodia growth ....................... 113

5.4.3. Relocated spines are maintained in the OB network ..................................... 115

5.4.4. MC activity induces SHF directional growth and spine relocation .............. 115

5.4.5. Glutamate released from MC controls the motility of SHF .......................... 117

5.4.6. MC-derived BDNF induces the spine relocation of adult-born GC .............. 118

5.4.7. Spines with SHF are maintained after sensory deprivation .......................... 119

5.4.8. Spine relocation is involved in odor information processing ........................ 120

5.5. Discussion ............................................................................................................ 121

5.6. Methods ................................................................................................................ 123

5.6.1. Animals ......................................................................................................... 123

5.6.2. Stereotaxic injection ...................................................................................... 124

5.6.3. Time-lapse two-photon imaging in vivo ........................................................ 124

5.6.4. Time-lapse two-photon imaging in vitro ....................................................... 125

5.6.5. Image analysis ............................................................................................... 126

5.6.6. Stimulation of mitral cells ............................................................................. 128

5.6.7. Iontophoresis and puff application ................................................................ 129

5.6.8. Unilateral nostril occlusion ........................................................................... 129

5.6.9. Immunohistochemistry .................................................................................. 130

5.6.10. In situ hybridization ...................................................................................... 131

5.6.11. OB network model ........................................................................................ 131

5.6.12. Statistical analysis ......................................................................................... 132

5.7. References ............................................................................................................ 132

5.8. Figures .................................................................................................................. 136

5.9. Authors’ contributions .......................................................................................... 162

6. General conclusions .................................................................................................... 163

References .......................................................................................................................... 165

ANNEX A - Tracking neuronal migration in adult brain slices ......................................... 185

A1. Résumé .................................................................................................................... 186

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A2. Abstract .................................................................................................................... 187

A3. Introduction ............................................................................................................. 188

A4. Basic Protocol .......................................................................................................... 189

A4.1. Time-lapse video-imaging of neuronal migration in adult acute brain slices .. 189

A4.2. Materials ........................................................................................................... 189

A4.3. Protocol steps .................................................................................................... 190

A5. Support Protocol ...................................................................................................... 192

A5.1. Stereotaxic injection of viral vectors into the SVZ of the adult mouse ............ 192

A5.2. Materials ........................................................................................................... 193

A5.3. Protocol steps .................................................................................................... 193

A5.4. Reagents and Solutions ..................................................................................... 194

A6. Commentary ............................................................................................................ 194

A6.1 Background Information .................................................................................... 194

A6.2. Critical Parameters and Troubleshooting ......................................................... 196

A6.3. Anticipated Results ........................................................................................... 197

A6.4. Time Considerations ......................................................................................... 197

A7. References................................................................................................................ 198

A8. Figures ..................................................................................................................... 201

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List of tables

Chapter 3

Figure 1. Clinical characteristics of human subjects

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List of figures

Chapter 1

Figure 1. First histological evidence of adult neurogenesis in rodents.

Figure 2. A schematic drawing of the adult mouse forebrain and cells in the subventricular

zone (SVZ), rostral migratory stream (RMS), and olfactory bulb (OB).

Figure 3. Diagram of olfactory system pathways.

Figure 4. Diagram of olfactory bulb neuronal elements, grouped into categories of (A)

afferent fibers, (B) principal cells and (C) local interneurons.

Figure 5. Connectivity of olfactory bulb showing basic molecular, cellular and functional

organization.

Figure 6. Diagram of morphological classes of adult-born granule cells and their positions

in the olfactory bulb.

Figure 7. Electron micrographs of synapses in the EPL.

Figure 8. Regulation of KCC2 functions by transcriptional control, subcellular targeting

and protein phosphorylation.

Figure 9. Basic configurations of a modern compound microscope with an infinity-

corrected objective lens.

Figure 10. Light path through a compound microscope with transmission geometry.

Figure 11. Schematic diagram of an epifluorescence microscope.

Figure 12. Simplified Jablonski diagram.

Figure 13. Optical layout of fluorescence microscopy techniques.

Figure 14. Two-photon excitation microscopy.

Figure 15. Different states of light polarization.

Figure 16. Polarized light imaging geometry.

Chapter 3

Figure 1. Schematic diagram of polarized light imaging.

Figure 2. Mouse brain dataset in polarized light, sagittal sections.

Figure 3. Fiber orientation information obtained with polarized light.

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Figure 4. Comparison of polarized light imaging and TH immunostaining.

Figure 5. Comparative assessment of fiber intensity in polarized light between control and

PD in internal capsule.

Figure 6. Comparison of PLI and Luxol Fast Blue staining for myelin, anterior limb of

internal capsule.

Chapter 4

Figure 1. Tracking tangential and radial migration in adult brain slices.

Figure 2. Comparison of KCC2 immunohistochemical staining in RMS and OB.

Figure 3. Tangential migration parameters of adult-born neuroblasts in the RMS in

response to KCC2 activation.

Figure 4. Tangential migration parameters of adult-born neuroblasts in the RMS-OB in

response to KCC2 activation.

Figure 5. Radial migration parameters of adult-born neuroblasts in the RMS-OB in

response to KCC2 activation.

Figure 6. Radial migration parameters of adult-born neuroblasts in the GCL in response to

KCC2 activation.

Chapter 5

Figure 1. Relocation of mature spines of adult-born GC in the OB.

Figure 2. SHF determine the relocation of the spines of adult-born but not early-born GC.

Figure 3. Relocated spines are stabilized in the OB network and are part of the

dendrodendritic synapses.

Figure 4. Olfactory sensory activity stabilizes SHF and promotes spine relocation.

Figure 5. Glutamate released from MC stabilizes spine head filopodia.

Figure 6. BDNF application promotes spine relocation.

Figure 7. The activity of BDNF-lacking MC does not induce spine relocation.

Figure 8. Spines with SHF are selectively maintained after sensory deprivation.

Figure 9. Spine relocation promotes fast synchronization of MC with functional

consequences for odor information processing.

Supplementary Figure 1. Direction vector amplitudes of GC spines and dendrites.

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Supplementary Figure 2. SHF dynamic at different maturational stages.

Supplementary Figure 3. Random stimulation pattern of MC does not induce GC spine

relocation.

Supplementary Figure 4. Activation of AMPARs is required for SHF motility.

ANNEX A

Figure 1. Tissue preparation.

Figure 2. Tracking tangential and radial migration in adult brain slices.

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List of abbreviations

AD Alzheimer’s disease

AMPA α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid

ANOVA Analysis of variance

AP Anterio-posterior

α-SN α-synuclein

BDNF Brain derived neurotrophic factor

BL Baseline

BrdU 5-bromo-2-deoxyuridine

BV blood vessel

CaMK Ca2+/calmodulin-dependent protein kinase

CC Corpus callosum

CCC Cation-chloride cotransporter

CCD Charge-coupled-device

CNS Central nervous system

CTD C-terminal domain

DAB Diaminobenzidine

DAPI 4,6-Diamidino-2-phenylindole

DCX Doublecortin

DG Dentate Gyrus

DIC Differential interference contrast

DNA Deoxyribonucleic acid

DPI Days post-injection

DTI Diffusion tensor imaging

DTT Diffusion tensor tractography

DV Dorso-ventral

EGF Epidermal growth factor

EPL External plexiform layer

EPSC Excitatory post synaptic current

GABA Gamma aminobutyric acid

GAD Glutamic acid decarboxylase

GC Granule cell

GCL Granule cell layer

GDNF Glial cell line-derived neurotrophic factor

GFAP Glial fibrillary acidic protein

GFP Green fluorescent protein

GL Glomerular layer

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HD Huntington’s disease

IPSC Inhibitory post synaptic current

IR Infrared

KCC K+/Cl- cotransporter

KO knockout

LB Lewy body

LN Lewy neurite

LOT Lateral olfactory tract

LTP Long-term potentiation

LV Lateral ventricle

MC Mitral cell

MCL Mitral cell layer

mEP Miniature excitatory post synaptic current

mGluR2 metabotropic glutamate receptor 2

MIA Melanoma inhibitory activity

mIPS Miniature inhibitory post synaptic current

ML Medio-lateral

MRI Magnetic resonance imaging

NA Nucleus accumbens

NCAM Neural cell adhesion molecule

NeuN Neuronal nuclei

NIR Near-infrared

NKCC Na+/K+/Cl- cotransporter

NMDA N-Methyl-D-Aspartate

NSC Neural stem cells

NTD N-terminal domain

OB Olfactory bulb

OGB Oregon Green BAPTA

ONL Olfactory nerve layer

OSN Olfactory sensory neurons

Pax6 Paired box gene 6

PBS Phosphate-buffered saline

PD Parkinson's disease

PET Positron emission tomography

PFA Paraformaldehyde

PG Periglomerular cell

PGL Periglomerular cell layer

PLI Polarized light imaging

PSA-NCAM Polysialylated neuronal cell adhesion molecule

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PSD95 Postsynaptic density protein 95

RI Refractive index

RMS Rostral migratory stream

SD Standard deviation

SEM Standard error of the mean

SGZ Subgranular zone

SHF Spine head filopodia

SLC Solute carriers

SN Substantia nigra

SPECT Single photon emission computerized tomography

STIM Stimulation

SVZ Subventricular zone

TCS Transcranial sonography

TH Tyrosine hydroxylase

TrkB Tropomyosin related kinase B

TTX Tetradotoxin

VEGF Vascular endothelial growth factor

VBM Voxel based morphometry

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Acknowledgements

Firstly, I want to express my deep gratitude and appreciation to my Director of Research,

Dr. Armen Saghatelyan, for his tremendous support and for teaching me a host of

techniques that will help me throughout my career. I am also grateful to Dr. Tigran Galstian

for his guidance and for sharing his insights in physics and optics. Thanks to Dr. Marina

Snapyan for her professionalism and for always being there to support and encourage. I

also thank Dr. Daniel Côté who helped me a lot with the different imaging techniques. I am

grateful to Dr. Gurgen Melkonyan, for developing the first prototypes of label-free imaging

and for his sincere support that helped starting my project. I thank the members of Armen’s

laboratory, past and present: Vincent, Archana, Delphine, Sarah, Arthur, Cedric, Marcos,

Helia, Rodrigo, Lynda, Lusine, Jivan, Qian and Caroline, as well the members of the

laboratories of Drs. Daniel Côté and Martin Parent and all my colleagues CRIUSMQ who

made this journey more enjoyable.

Thank you all of my friends who are so close no matter how far. Thank you, Alik and

Tsovik - you and I can share the silence, finding comfort together - the way old friends do.

Thank you, Armina - with you to live the dreams we always had. Thank you, my darlings,

Maya and Hayk, thanks for all the joy you're bringing. And last but not least, thank you, my

mother and father for bringing me into this world and for all your love and care that

surrounds me every single day of my life. With you forever, nothing really matters to me.

Any way the wind blows…

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Foreword

During the first part of my PhD work I focused on label-free imaging methods for tracking

the development of migrating cells in the migratory stream of forebrain. I also studied the

mechanisms controlling migration and the implication of mature adult-born neurons in the

functioning of olfactory system. My thesis consists of one published paper where I am a

coauthor, one paper that is in preparation and the third work which should be explored

further.

First part of my results represents the paper which is preparation, where we did

comparative study of myelinated fibers in human brain sections under normal and

pathological conditions (Parkinson’s disease). We demonstrate significant differences of

fiber intensities imaged in polarized light between these two groups. Our results highlight

the importance of further multimodal imaging studies in the areas of the brain which are

generally not known to be associated with neurodegenerative diseases. I am the main

contributor in this paper; I co-developed the polarimetric imaging technique, conducted all

experiments and data analysis. Dr. Gurgen Melkonyan has designed the first prototype of

polarimeter and assisted in some experiments. Dr. Martin Parent has provided with human

brain samples from healthy subjects and patients with Parkinson’s disease. Dr. Armen

Saghatelyan and Dr. Tigran Galstian are my director and co-director respectively and they

designed and supervised this project.

The second part of my results aims to unveil the role of KCC2 in different types of

neuronal migration. Our preliminary results demonstrate that KCC2 activation fosters radial

migration of neuroblasts in the rostral migratory stream and in the olfactory bulb. This

work needs to be continued to reveal the exact mechanisms of KCC2 action on radially

migrating neuroblasts. I had main contribution in this paper by conducting all experiments.

Dr. Armen Saghatelyan has designed and supervised this project.

In third part of my thesis I present a research article published in Nature Communications

in 2016 and titled “Principal cell activity induces spine relocation of adult-born

interneurons in the olfactory bulb”. Here we reveal a new form of structural remodeling

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where mature spines of adult-born but not early-born neurons relocate in an activity-

dependent manner. The publication of this paper was done thanks to important contribution

from several co-authors. Vincent Breton-Provencher is the first author of the paper; he

performed most of the experiments, analyzed the data, co-designed the study and co-wrote

the paper with Armen Saghatelyan. I performed the majority of in vivo imaging

experiments.

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1. Introduction

1.1. Adult Neurogenesis

1.1.1. Neuronal development in adult rodent brain

In contrast to some lower vertebrates, known to regenerate whole regions in the central

nervous system (CNS), traditionally it was assumed that all neurons in mammals are being

produced only in embryonic and early postnatal stages of development. This dogma has

been, however, challenged by discovery of persistent proliferation, migration and

integration of neurons in the adult mammalian brain. First evidence of adult neurogenesis

dates half a century ago, when Altman and Das have shown the presence of

deoxyribonucleic acid (DNA) synthesis marker, 3H-thymidine in hippocampal neurons of

adult rodent brains (Altman and Das, 1965). They further discovered another even larger

neurogenic niche in the postnatal brain giving rise to granule and periglomerular cells of the

olfactory bulb (OB) (Fig. 1; Altman, 1969). Despite continuous research in the field during

next two decades, the results didn’t get proper acknowledgment by scientific community

mainly due to the lack of evidence of adult neurogenesis in the primates at that time (Rakic,

1985).

The situation has changed only in 1990’s when adult-born cell proliferation in

subventricular zone (SVZ) and further migration via rostral migratory stream (RMS) have

been discovered (Lois and Alvarez-Buylla, 1993; Reynolds and Weiss, 1992). These

important studies were followed by a series of publications revealing a physiological

importance of adult neurogenesis by showing that environmental enrichment and learning

modulates the level of production of new neurons in the adult rodent brain (Gould et al.,

1999; Kempermann et al., 1997). Adult neurogenesis was also discovered in the dentate

gyrus (DG) of hippocampus of primates and humans (Eriksson et al., 1998), further

reinforcing the notion that specific regions of the adult brain produce new neurons

throughout the life. Nowadays adult neurogenesis is a widely accepted fact, shown on

practically all mammal species studied and the current research topics range from the

molecular basis and functional implications of adult neurogenesis to its role in

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neurodegenerative disorders and brain injury recovery. In mammals adult neurogenesis

under normal, uninjured conditions is confined to two main regions. One is the subgranular

zone (SGZ) of DG which supplies the hippocampus with new granule cells, and second one

Figure 1. First histological evidence of adult neurogenesis in rodents (adapted from

Altman, 1969).

A) Drawing of sagittal sections of the brain from rats of different ages showing the RMS (black). CC, corpus

callosum; CO, cerebral cortex; CP, caudate-putamen; HI, hippocampus; LV, inferior horn of lateral ventricle;

NA, nucleus accumbens; OB, olfactory bulb; SE, septum. B) Photomicrograph of the subependymal layer of

the inferior horn of the lateral ventricle (LV) and the RMS (arrow) in sagittal section in a 21 day old rat.

Cresyl violet, X16. C) The RMS at higher magnification.

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is the SVZ bordering the lateral ventricles (LV) from where the new-born cells migrate via

RMS to the OB to become interneurons in granule cell and periglomerular cell layers (GCL

and PGL, respectively). The neurogenesis in the OB of rodents is much more pronounced

with much higher cell turnover (~30 000 cells produced per day) and longer migration

distances (several millimeters) (Lois and Alvarez-Buylla, 1994; Winner et al., 2002). The

cells proliferate in SVZ, where a GFAP-positive subpopulation of stem cells gives rise to

proliferating transit amplifying cells that, in turn, differentiate into migrating immature

neurons (neuroblasts) (Fig. 2; Doetsch et al., 1999; Ming and Song, 2005).

These newly formed neuroblasts migrate tangentially in chains towards the OB via RMS in

a tubular structure formed by glial cells (Lois et al., 1996). They also use blood vessels as a

scaffold for migration and source of molecular factors (Snapyan et al., 2009). The

migration of neuroblasts from the SVZ to the OB is orchestrated by various mechanisms.

Figure 2. A schematic drawing of the adult mouse forebrain and cells in the subventricular

zone (SVZ), rostral migratory stream (RMS), and olfactory bulb (OB) (adapted from

Gengatharan et al., 2016).

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The chain migration of neuroblasts highly depends on polysialylated-neural cell adhesion

molecule (PSA-NCAM) proteins on the surface of the migrating neuroblasts. Several other

factors are also known to coordinate the motility of neuroblast migration in chains, such as

netrin, ephrin-B2, integrin and γ-aminobutyric acid A receptor (GABAAR) activity (Bolteus

and Bordey, 2004; Conover et al., 2000; Hu et al., 1996; Murase and Horwitz, 2002).

Netrin is also involved in the directionality of chain migration, as is the Slit/Robo signaling

(Murase and Horwitz, 2002; Nguyen-Ba-Charvet et al., 2004; Wu et al., 1999).

Having reached the core of OB, the neuroblasts drastically change the mode and

directionality of migration. They detach from the chains and undertake individual radial

migration into GCL and PGL. The process of detachment from chains is initiated by reelin

(Hack et al., 2002) and tenascin-R, which then control the radial migration to their final

destination in the OB (Saghatelyan et al., 2004). Whether the cells will stop in GCL or

continue their radial migration to reach PGL, is influenced by transcription factor Pax6

(Hack et al., 2005; Kohwi et al., 2005). Once reached their target area, the cells get mature

and integrate in the olfactory network as functional granule cells (GC) and periglomerular

cells (PG) (Lledo and Saghatelyan, 2005). Both cell types are axonless interneurons making

reciprocal dendrodendritic synapses with mitral or tufted cells (Shepherd, 2004).

1.1.2. The olfactory system

The olfactory system is essential for the survival of many animal species, providing vital

information about food location and influencing social and sexual behaviours. In mammals,

odor information is conveyed by olfactory sensory neurons (OSN) located in the olfactory

epithelium into the OB where it is processed by bulbar principal cells and interneurons

(Shepherd, 2004). From the OB, the information is conveyed to the piriform cortex,

amygdala, hippocampus, and entorhinal cortex (Davis, 2004; Shepherd, 2004). Olfactory

system is the only sensory system where information processing occurs without a thalamic

relay. It is assumed that the OB “substitutes” thalamus in terms of processing odor

information and relaying it to higher cortical areas (Kay and Sherman, 2007).

The OB consists of multiple layers each containing morphologically distinct cells (Fig. 3).

The neurons in the OB are classified according to the layers in which their cell bodies

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reside, such as PG in the glomerular layer (PGL), tufted cells in the external plexiform

layer (EPL), mitral cells (MC) in the mitral cell layer (MCL), and GCs in the GCL (Fig. 4).

The OB receives the sensory information from OSNs in the olfactory epithelium (Shepherd,

2004). The mammals have ~1000 different functional odorant receptors and, interestingly,

each OSN expresses only a single odorant receptor. The axons from different OSNs

expressing the same receptor come together into a spherical structure in the OB, called

glomerulus. Therefore, the glomeruli are considered to be the functional units in the OB

and together they represent spatial map of the different olfactory receptors located in the

olfactory epithelium. The unique affinity profiles of odorant receptors together with

specific arrangement of terminals in glomeruli gives rise to significant combinatorial

complexity in odor processing, which allows to discriminate more than 1.7 trillion of

different odors (Bushdid et al., 2014). While these data were nominated for “Breakthrough

of the Year 2014” and heavily promoted by the popular press, they raised a lot of

skepticism in the scientific community (Gerkin and Castro, 2015; Meister, 2015). In

Figure 3. Diagram of olfactory system pathways (Shepherd, 2004).

Here some of the major connections in the mammalian olfactory bulb are shown. The olfactory sensory

neurons in the epithelium send the input signal to olfactory bulb, which in turn projects it to the olfactory

cortex. Olfactory epithelium populated with overlapping types of olfactory sensory neurons project to

individual glomeruli. c.f., centrifugal fiber; G, granule cell; M, mitral cell; OSN, olfactory sensory neuron; P,

pyramidal cell; PG, periglomerular cell; r.c., recurrent axon collateral; T, tufted cell.

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particular, the authors used mathematical modeling of experimental data showing that

humans can successfully discriminate 148 pairs of odors (Bushdid et al., 2014). However,

several flaws in the analytical model were identified (Gerkin and Castro, 2015; Meister,

2015). Moreover, if the same method would have been applied to human color vision, one

would estimate that humans can distinguish at least 1027 colors, which is in dramatic

conflict with experimental evidence (Meister, 2015). Therefore, the claim that humans can

discriminate 1.7 trillions of odors is unjustified (Gerkin and Castro, 2015; Meister, 2015)

and further studies are required to solve this issue.

In glomeruli, the terminals of OSN axons synapse with the primary dendrites of the

principal neurons (mitral or tufted cells) that convey the sensory information from the

periphery to the second-order neurons, forming thereby the basis of an odor processing

column (Mori and Sakano, 2011; Ressler et al., 1994). The PG and short axon neurons in

the GCL play an important modulatory role via inter- and intraglomerular excitatory and

inhibitory connections to the principal neurons (Aungst et al., 2003). The PGs can be

distinguished based on the expression of calretinin, calbindin, parvalbumin, glutamate and

γ-aminobutyric acid (GABA), with some of the latter co-expressing tyrosine hydroxylase

(TH), an enzyme involved in dopamine synthesis (Bagley et al., 2007; Brill et al., 2009;

Kosaka et al., 1995; Whitman and Greer, 2007a). This abundance of heterogeneous

subpopulations of cells is not fully clear yet, but is apparently related to complexity of the

functions in the first stages of odor processing.

Beneath the GL is the EPL where the lateral dendrites of principal output neurons make

dendrodendritic synapses with the GCs. The EPL is also sparsely populated by somas of

tufted cells, astrocytes and parvalbumin-expressing interneurons that make synapses to

output neurons or other interneurons (Hamilton et al., 2005). Next follows a thin layer of

MC somas from which this layer has got its name. The long lateral dendrites of mitral cells

and tufted cells (further jointly referred as principal cells) reach the EPL where they form

glutamatergic synaptic connections with the dendrites of interneurons (Fig. 4). The

principal cells transfer the olfactory information with their axons to the olfactory cortex via

the lateral olfactory tract (LOT), which, in turn, conveys the signals further to piriform

cortex, amygdala, hippocampus (Davis, 2004; Igarashi et al., 2012; Shepherd, 2004).

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Figure 4. Diagram of olfactory bulb neuronal elements, grouped into categories of (A)

afferent fibers, (B) principal cells and (C) local interneurons (adapted from Shepherd,

2004).

ONL, olfactory nerve layer; GL, glomerular layer; EPL, external plexiform layer; MCL, mitral cell layer; IPL,

internal plexiform layer; GRL, granule cell layer. In A: ON, olfactory nerve fibers. Centrifugal afferents are

from the contralateral anterior olfactory nucleus (cAON), ipsilateral anterior olfactory nucleus (iAON), tenia

tecta (TT), olfactory cortex (OC), horizontal limb of the diagonal band (HDB), locus coeruleus (LC), and

raphe nucleus (Ra). pE, pars externa of the AON; pM, pars medialis of the AON. In B: dendrites and axon

collaterals of a mitral cell (M), and internal tufted cell (iT, or a displaced mitral cell, dM), a middle tufted cell

(mT), and an external tufted cell (eT) are illustrated, a, axon; d, dendrite. In C: GI, GII, and GIII, three types

of granule cells; PG, periglomerular cell; SA(S), superficial short-axon cell; SA(B), Blanes cell.

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Deeper from MCL is the GCL consisting mostly of the cell bodies of GCs, as well as a

sparse amount of short-axon cells. The GCs are notable for their lack of axon and they have

characteristically large dendritic spines. The primary dendrite of each GC extends radially

toward the outer layers of OB, giving rise to secondary branches (distal dendrites) and

terminating in the EPL (Fig. 4c). The rest of GC processes are short basal dendrites serving

Figure 5. Connectivity of olfactory bulb showing basic molecular, cellular and functional

organization (adapted from Shepherd, 2004).

Molecular components (left): OR - olfactory receptor, ON - olfactory nerve, AMPA - 2-amino-5-

phosphonovaleric acid, NMDA - N-methyl-D-aspartate, M/T - mitral/tufted cell, PG - periglomerular cell,

GluR - ionotropic glutamate receptor, GABA R - GABA receptor, DAR - dopamine receptor, NE -

norepinephrine, aAR - alpha adrenoreceptor, mGluR2 - metabotropic glutamate receptor, GR - granule cell,

Synaptic circuit (middle): ORN - olfactory receptor neuron, J, K - ORN subsets, e - excitatory, i - inhibitory.

Structural-functional relations (right): top - overlapping responses of ORNs to a range of odors (1..n); middle

- ORN subsets connectivity to the glomeruli; bottom - due to lateral inhibition, the response spectra of M/T

cells have less overlap

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as inputs from other brain regions. The GCL is comparatively wide and depending where

the GC somas reside, their morphology and functional role might be different. The

dendritic ramifications of superficial GCs reach EPL where they mainly contact the tufted

cell dendrites, while the dendrites of deep GCs synapse to the MCs in deeper layers of EPL

(Greer, 1987; Mori et al., 1983; Orona et al., 1983). Less heterogeneous than PGs, the GCs

are nevertheless shown to have small subpopulations expressing calretinin, glycoprotein

5T4, metabotropic glutamate receptor 2 (mGluR2), Neurogranin and Ca2+/calmodulin-

dependent protein kinases IIα (CaMKIIα) and IV (CaMKIV) (Batista-Brito et al., 2008;

Gribaudo et al., 2009; Imamura et al., 2006; Jacobowitz and Winsky, 1991; Liu, 2000;

Néant-Fery et al., 2012; Zou et al., 2002).

The GCs provide recurrent and lateral inhibition to lateral dendrites of principal neurons

(Arevian et al., 2008; Isaacson and Strowbridge, 1998; Schoppa and Urban, 2003; Tan et

al., 2010; Urban, 2002). This inhibition is mediated through reciprocal synapses, where

glutamate is released from the lateral dendrites of the mitral or tufted cells onto the spine of

a GC, which in turn induces the release of GABA back onto the principal cell dendrites

(Isaacson and Strowbridge, 1998; Price and Powell, 1970; Shepherd, 2004). The GCs and

PGs play an important role in the rhythmic activity of the OB. PGs coordinate theta activity

by regulating baseline and odor-evoked inhibition, whereas GCs are involved in the

synchronization of MC activity and generation of gamma rhythm (Arevian et al., 2008;

Fukunaga et al., 2014; Urban, 2002; Yokoi et al., 1995). An overview of synaptic

connectivity of OB is presented in Fig. 5. A striking feature of adult OB is its ability to

produce new GCs and PGs throughout the life of an animal. In the next parts, I will discuss

different processes leading to the generation, migration, maturation and functional

integration of adult-born interneurons and will discuss the current understanding of the role

played by these cells in olfactory system.

1.1.3. Proliferation of neuronal precursors in the SVZ

Adult-born neurons in the OB are derived from stem cells located in the SVZ bordering

lateral ventricles. From the SVZ neuronal precursors migrate several mm to reach the OB

and integrate into its neuronal network (Fig. 2). The SVZ neurogenic niche contains

various cell types, each having an important contribution in neurogenic activity: neural

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stem cells (NSC), transit-amplifying cells, neuroblasts, astrocytes, endothelial and

ependymal cells (Ming and Song, 2011). Adult neurogenesis comprises a complicated and

interrelated chain of events which are orchestrated by timely involvement of molecular

cues. The first step of these events is considered to be a division of slow dividing NSCs in

the SVZ - the Type B cells, which are derived from radial glial cells producing the neurons

during embryonic life (Fuentealba et al., 2015; Merkle et al., 2014; Young et al., 2007). The

type B cells are astrocyte-like NSCs giving rise to transit-amplifying cells (Type C cells),

which in turn divide rapidly to give rise to neuronal precursors (Type A cells) (Doetsch et

al., 1999). NSCs are located in the specialized microenvironment where they need to

integrate multiple signals to maintain their quiescence or get activated to generate neurons.

They contact blood vessels via their basal processes and are exposed to a variety of factors

from the cerebrospinal fluid via small apical process bearing a non-motile primary cilium

(Fig. 2; Doetsch et al., 1999). The non-neuronal cells in SVZ such as the ependymal and

endothelial cells modulate adult neurogenesis by providing molecular factors.

1.1.4. Migration of neuroblasts in the adult brain

The migration of neuroblasts in the SVZ-OB pathway consists of succession of migratory

and stationary phases (Snapyan et al., 2009). Typically the initial periods of extension of

leading process and exploration are followed by the displacement of the cell body towards

the place of the leading process (Wichterle et al., 1997). These processes are regulated by

interplay of different intracellular mechanisms (Lalli, 2014). For instance, the stabilization

of the leading process is attributed to doublecortin (DCX), a microtubule-associated protein

(Belvindrah et al., 2011; Koizumi et al., 2006), while the movement of nucleus in the

cytoplasm is supported by Lis1/Ndel1 complex (Hippenmeyer et al., 2010).

In the SVZ, neuroblasts assemble together in tightly packed chains and begin to migrate

towards the OB through a special environment in the RMS along topographically aligned

blood vessels (Saghatelyan, 2009; Snapyan et al., 2009; Whitman and Greer, 2009). It has

been proposed that cilia beating of ependymal cells creates a gradient of chemorepellent

molecules secreted from the septum and choroid plexus, such as Slit (Sawamoto et al.,

2006; Wu et al., 1999), that pushes away neuroblasts from the SVZ (Sawamoto et al.,

2006). While this gradient and ependymal cells cilia beating play an important role in the

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neuroblasts migration in the SVZ, it is difficult to imagine how the unidirectional vectorial

gradient of a single molecule might explain the complex-shaped curvature migration of

newborn cells in the RMS. It would be intuitive to suggest that the further migration of

neuroblasts is somehow controlled by attractive factors coming from OB. However surgical

removal of the OB did not affect neuroblasts migration in the RMS (Jankovski et al., 1998;

Kirschenbaum et al., 1999), questioning the role of the OB in the directional guidance of

neuronal precursors. The work from our lab has revealed that blood vessels play an

important role in the neuroblasts migration (Snapyan et al., 2009). They topographically

outline the migratory stream and neuroblasts use these long, parallel running blood vessels

as a substrate for their migration. In the context of vasculature-associated neuronal

migration model, no attractive molecules secreted from the OB are needed and a single

repellent molecule might be sufficient to push cells away from the posterior parts of the

brain (SVZ), with vasculature providing local cues to keep migrating precursors in the

RMS and guide them towards the OB (Snapyan et al., 2009). Therefore, the migration of

neuroblasts in the RMS would be controlled by the local factors and in line with this,

myriads of local factors derived either from neuroblasts, or astrocytes and endothelial cells

have been shown to regulate neuronal migration.

Astrocytes ensheath the chains of neuroblasts and their processes undergo rapid structural

changes in response to neuroblasts-secreted repulsive protein Slit1 (Kaneko et al., 2010).

Slit1 repels astrocytic processes via Robo receptor and leads to the formation of structurally

permissive glial tunnels enabling neuronal migration (Kaneko et al., 2010). Astrocytes may

also release soluble factors that modulate neuroblasts migration and/or survival. For

example, astrocytes may release glutamate or melanoma inhibitory activity (MIA) protein,

which are required for neuroblasts migration (Mason et al., 2001; Platel et al., 2010).

Astrocytes also express high affinity GABA transporters and inhibition of GABA uptake

reduces neuroblasts migration (Bolteus and Bordey, 2004).

Brain derived neurotrophic factor (BDNF) released from the endothelial cells promotes

migration via low affinity BDNF receptor, p75NTR present on neuroblasts (Snapyan et al.,

2009). Interestingly, migrating neuroblasts control their own migration by regulating the

amount of free extracellular BDNF. GABA released from neuronal precursors induces

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Ca2+-dependent insertion of high-affinity TrkB receptors into the plasma membrane of

astrocytes that ensheath migrating neuroblasts and contact blood vessels. This leads to

trapping of extracellular BDNF and, therefore, to the entry of migrating cells to the

stationary phase. Through this mechanism, neuronal precursors regulate availability of

BDNF required for their own migration (Snapyan et al., 2009).

A number of other molecular factors also affect neuronal migration in the adult brain.

These include ephrin-B (Conover et al., 2000), and integrin families (Belvindrah et al.,

2007; Murase and Horwitz, 2002); the ErbB4 (Anton et al., 2004) and prokineticin 2 (Ng et

al., 2005) receptors; as well as various growth factors such as glial cell line-derived

neurotrophic factor (GDNF; Paratcha et al., 2006), and vascular endothelial growth factor

(VEGF; Bozoyan et al., 2012; Wittko et al., 2009).

Tangential chain-arranged migration continues until the neuroblasts reach the OB. Here

they detach from chains and start individual radial migrations along blood vessels to outer

layers of OB (Bovetti et al., 2007; Lledo and Saghatelyan, 2005; Snapyan et al., 2009).

While many migration related mechanisms here are similar to the ones operating during

tangential migration, there are however some specific differences. The processes of

separation of neuroblasts from chains are associated with the expression of extracellular

matrix glycoprotein tenascin-R in the RMS-OB (David et al., 2013; Saghatelyan et al.,

2004). Also the glycoprotein reelin released by the principal cells of the OB has a role in

the transition to radial migration and in the final placement of adult-born neuroblasts in the

OB (Hack et al., 2002; Kim et al., 2002).

1.1.5. Maturation and integration of adult-born interneurons in the OB

When the precursor cells find their final position in the OB, they differentiate and mature to

become fully functional OB interneurons. The neuroblasts differentiate into two types of

interneurons - the GCs and the PGs and while doing so their morphology evolves from

bipolar cells with short processes into a fully developed neurons with dendritic branches

extending hundreds of microns (Carleton et al., 2003; Lledo et al., 2006; Lledo and

Saghatelyan, 2005; Petreanu and Alvarez-Buylla, 2002). The maturation stages of GCs are

divided into 5 classes (Fig. 6; Carleton et al., 2003; Petreanu and Alvarez-Buylla, 2002).

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According to this classification, the class 1 cells are the tangentially migrating neuroblasts

of RMS-OB. It takes 5 days for the neuronal precursors to get there from the SVZ. The

neuroblasts detaching from the chains and starting to migrate radially are considered class 2

cells. Then within 9 to 15 days from their birth, the growing primary dendrites of class 3

cells start to reach the EPL, while not yet contacting the MC. Starting from approximately

11 days from their generation, yet immature GCs of class 4 begin to develop their

secondary dendrites that are spineless at this stage. Eventually, near the 20th-30th day, the

GCs reach the morphological class 5 by developing multiple dendritic spines and become

synaptically integrated in OB.

Class 5 adult-born GCs are morphologically similar to the mature GCs produced at

embryonic and neonatal age (thereafter called early-born neurons), however they tend to

Figure 6. Diagram of morphological classes of adult-born granule cells and their positions

in the olfactory bulb (adapted from Petreanu and Alvarez-Buylla, 2002).

EPL - External plexiform layer; GL - glomeruli layer; MCL - mitral cell layer; GCL - granule cell layer; RMS

- rostral migratory stream.

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occupy deeper areas in the GCL (Imayoshi et al., 2008), while early-born GCs are more

concentrated in the superficial parts of the GCL (Lemasson et al., 2005). There is a

correlation between the position of GC in GCL and the area of EPL where their dendrites

reach, meaning that adult-born and early-born GCs may preferentially target the dendrites

of mitral or tufted cell, respectively (Greer, 1987; Shepherd et al., 2004). This evidence

hints on the differences in functional roles of adult-born and early-born GC populations.

There are less studies of adult-born PGs and classification of morphological changes of

adult born PGs has not been done (Belluzzi et al., 2003; Mizrahi, 2007). The migration of

PG neuroblasts seems to be faster than for GCs, despite the longer distance (Hack et al.,

2005), but dendritogenesis and synaptogenesis take longer time as compared to GC

development (Mizrahi, 2007; Whitman and Greer, 2007a).

Apart from differences in morphology, the evidence of electrophysiological distinctions

(such as spiking activity and sodium conductance) between early-born and adult-born GCs

further supports the idea of distinct functional role of these two subpopulations (Belluzzi et

al., 2003; Carleton et al., 2003). Both tangentially and radially migrating neuroblasts have

no synaptic currents, but their ionotropic AMPA and GABA receptors are already

functional (Carleton et al., 2003) The radially migrating cells express NMDA receptors,

however their presence in earlier stages is a matter of controversy (Carleton et al., 2003;

Platel et al., 2010).

The post-synaptic currents start to appear together with the growth of the primary dendrite

of adult-born GCs. The inhibitory synaptic activity prevails at this stage, probably due to

presence of GABA (Carleton et al., 2003; Panzanelli et al., 2009). With the development of

secondary dendrites and spines, excitatory post-synaptic activity appears (Carleton et al.,

2003) and coincidentally the cells begin to give rise to sodium currents (Carleton et al.,

2003). Na+ current become more prominent with further growth of GC distal dendrites in

the EPL (Carleton et al., 2003). The sodium currents and action potentials of adult-born GC

interneurons develop at a later maturational phase than those of early-born GCs (Belluzzi et

al., 2003; Carleton et al., 2003; Kelsch et al., 2008). Interestingly, compared to their

neonatal counterparts, the synaptically integrated adult-born GCs and PGs have higher

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sodium currents, their conductance-voltage ratio is more steep and the activation threshold

is more negative (Belluzzi et al., 2003; Carleton et al., 2003; Saghatelyan et al., 2005).

1.1.6. Synaptic development of adult-born GCs

GCs are axonless neurons that have basal and apical dendrites. The apical dendrite is

composed of an unbranched proximal segment and highly branched distal segment with

numerous spines. The spines of GCs are constituents of dendrodendritic synapses that these

interneurons form with the lateral dendrites of MCs. In dendrodendritic reciprocal synapses

glutamate is released from the lateral dendrites of the MC onto the spine of a GC, which in

turn induces the release of GABA back onto the principal cell dendrites (Isaacson and

Strowbridge, 1998). The dendrodendritic synapses are the only output synapses of GCs in

the OB (Shepherd, 2004) and are responsible for recurrent and lateral inhibition of principal

neurons (Isaacson and Strowbridge, 1998; Schoppa and Urban, 2003).

During the development of adult-born GCs, they first receive input synapses on the basal

dendrites and on the proximal part of the primary dendrite, before forming the output ones

on the distal dendrites (Kelsch et al., 2008; Pallotto et al., 2012; Panzanelli et al., 2009;

Whitman and Greer, 2007b). This contrasts with development of early-born interneurons

that receive input and output synapses simultaneously (Kelsch et al., 2008). Early in the

development of the adult-born GCs, there are more GABAergic than glutamatergic

synapses, which is assumed to be an important factor for the proper formation of dendrites

and spines in the EPL (Pallotto et al., 2012; Panzanelli et al., 2009). Soon after, the output

dendrodendritic synapses between distal dendrites of adult-born cells and lateral dendrite of

MC or tufted cells begin to develop (Kelsch et al., 2008; Panzanelli et al., 2009). Electronic

microscopy study confirmed the presence of dendrodendritic synapses between adult-born

GCs and MC lateral dendrites (Fig. 7; Whitman and Greer, 2007b).

It is thought that adult-born neurons by first receiving input synapses before forming the

output ones, silently integrate into the OB without disturbing already functional neuronal

network (Kelsch et al., 2008; Lledo and Saghatelyan, 2005). Optogenetic targeted-

stimulation of adult-born cells has demonstrated their GABAergic outputs onto the MCs

(Alonso et al., 2012; Bardy et al., 2010; Valley et al., 2013). While no glutamatergic inputs

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has been yet recorded from the adult-born GC following optogenetic or electrical

stimulation of MC, the synapses between GC and MC have clear dendrodendritic

architecture (Fig. 7), high presence of PSD95 puncta in the spine of GCs (Panzanelli et al.,

2009; Whitman and Greer, 2007b) and express AMPA and NMDA receptors (Breton-

Provencher et al., 2014). All this suggest that adult-born GC spines receive glutamatergic

input from bulbar principal neurons. The details and the mechanism of reciprocal synapse

development on the distal dendrites of adult-born GCs are not yet fully known. What we

know so far is that initially when the cells integrate in the OB, their spine density increases

and later with maturation it decreases again (Pallotto et al., 2012; Whitman and Greer,

Figure 7. Electron micrographs of synapses in the EPL ( from Whitman and Greer, 2007b)

A - Dendrodendritic synapse in the EPL between a mitral cell dendrite and a granule cell spine, unlabeled. B,

C - Examples of mitral to granule excitatory synapses on labeled spines at 42 dpi. Virally labeled cells were

marked by GFP immunohistochemistry and DAB; the spines of new granule cells are darkly stained. D - An

example of a bidirectional dendrodendritic synapse on the same spine head. Md - Mitral cell dendrite; Gr -

granule cell spine.

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2007b). This can be explained by increased search of synaptic partners during early

maturational periods, followed by synaptic pruning that may lead to the decreased spine

density at later maturational stages (Panzanelli et al., 2009; Whitman and Greer, 2007b).

The work from our laboratory has shown that MC activity orchestrates the initial

spinogenesis and formation of contact between GC and MC dendrites (Breton-Provencher

et al., 2014). Using time-lapse two-photon imaging of adult-born GCs at their different

maturational stages in acute slices of the OB, our lab has demonstrated that principal cell

activity regulates the filopodia dynamic during the early but not the late maturational stages

through the activation of NMDA receptors (NMDARs) on adult-born GC dendrites

(Breton-Provencher et al., 2014). Recent in vivo data suggested that GCs have high synaptic

turnover even when they are fully mature (Sailor et al., 2016). The mechanisms leading to

stabilization of some spines and disappearance of others are not clear, but it is plausible that

these mechanisms are activity-dependent.

1.1.7. Adult neurogenesis as a form of structural plasticity to adjust OB network

to changing environmental conditions

It is estimated that tens of thousands new neurons are added to the OB every day. This cell

addition is accompanied by apoptosis of GCs in order to maintain OB volume.

Interestingly, the largest amount of dying cells belongs to immature population of adult-

born neurons (Kuhn et al., 2005; Petreanu and Alvarez-Buylla, 2002). In fact, since the

sites of progenitor generation (SVZ) and adult-born cells integration (OB) are located

several mm apart, it is though that adult brain produces an excess of new neurons with

more than half destined for programmed cell death. However, this overproduction allows

for having large number of immature cells in the OB at any moment during the life of

animal which leads to up- or downregulation of the number of integrating adult-born

neurons in response to changing odor environment. Indeed, the number of adult-born cells

can be modulated by odor enrichment (Rochefort et al., 2002), odor deprivation (Petreanu

and Alvarez-Buylla, 2002; Saghatelyan et al., 2005; Yamaguchi and Mori, 2005),

pheromones (Mak et al., 2007; Shingo et al., 2003), and associative tasks based on olfaction

(Mouret et al., 2008; Sultan et al., 2010). In contrast, early-born neurons persist over time

in the OB (Lemasson et al., 2005).

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Several studies have shown that when adult-born neurons integrate into functional bulbar

network, they are more active as compared to their early-born counterparts. For example,

odor stimulation induces higher activation of adult-born as compared to early-born

interneurons (Carlén et al., 2002) and blocking adult neurogenesis only for 1 month results

in about 50% reduction in the inhibitory activity received by principal cells (Breton-

Provencher et al., 2009; Saghatelyan et al., 2005). Furthermore, the constant arrival of

adult-born interneurons and, consequently, the persistent formation and elimination of

dendrodendritic synapses formed by these cells with the principal neurons allows the OB

circuitry to maintain and modulate the spatio-temporal coding of sensory information and

facilitate odor learning and memory (Alonso et al., 2012; Breton-Provencher et al., 2009;

Mak et al., 2007; Sultan et al., 2010).

Why OB needs such an elaborated process for promoting plasticity? The main reason for

this could be because of the nature of output synapse in the OB. The main output synapse is

dendrodendritic reciprocal synapse between interneurons and bulbar principal cells. It is

therefore difficult to imagine other forms of plasticity in these synapses, such as long-term

potentiation (LTP) or long-term depression that induce changes the efficacy of the synapse

and allow adjustment of neuronal network in other brain regions. Because of the reciprocal

nature of dendrodendritic synapse, any changes in the efficacy of glutamatergic part will be

immediately accompanied by counterbalancing changes in the inhibitory part of the same

synapse. In line with this, no LTP was found in dendrodendritic OB synapses (Dietz and

Murthy, 2005; Gao and Strowbridge, 2009) and therefore other forms of structural

plasticity should exist to adjust bulbar network functioning.

The continuous supply of new neurons provides the OB with a reservoir of plastic cells and

it is thought to constantly sculpt the bulbar network in response to changing environmental

conditions (Sailor et al., 2016). The plasticity brought about by adult-born neurons depends

on the formation, retraction and/or stabilization of new synaptic contacts (Breton-

Provencher et al., 2014; Livneh and Mizrahi, 2012; Mizrahi, 2007) and growing evidence

suggests that new adult-born neurons in the OB are involved in the different types of

behavior, including short- (Breton-Provencher et al., 2009; Rochefort et al., 2002) and

long-term odor memory (Alonso et al., 2012; Sultan et al., 2010), odor discrimination

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(Enwere et al., 2004), parental (Mak and Weiss, 2010) and social (Mak et al., 2007)

behavior. However, there is an important conceptual problem since environmental changes

can be very rapid, whereas the synaptogenesis of adult-born neurons occurs over a longer

time scale. It remains thus completely unknown how bulbar network functions when rapid

and persistent changes in environmental conditions occur, even though new synapses have

not yet been formed.

In the third objective of my thesis, I undertook in vivo imaging of adult-born neurons with a

relatively rapid acquisition rate (once every 5 min), to understand how these cells react to

rapid changes in the odor environment and if there are other forms of structural plasticity in

the bulbar network that adapt its functioning to rapid and persistent changes in the

environmental conditions.

1.2. Overview of GABAergic signaling in the brain

During the formation of the nervous system, the cells undergo several crucial

developmental changes which are regulated by many molecular cues. Being the main

inhibitory neurotransmitter in the adult CNS (Watanabe et al., 2002), GABA is known to

play an excitatory role during early neuronal development (Rivera et al., 1999).

GABAergic synapses develop before the appearance of glutamatergic ones (Chen et al.,

1995; Del Rio et al., 1992) and the shift from the excitatory to inhibitory actions of GABA

is regulated by neuronal activity and expression of cation-chloride cotransporters (Ben-Ari

et al., 1997; 2002).

GABA binds to three types of neurotransmitter receptors: ligand-gated ion-channels such as

GABAA and GABAC (also called GABAA-rho) receptors; and G protein-coupled

metabotropic GABAB receptors. Both GABAA and GABAC are ligand-gated Cl- channels,

with GABAC being bicuculline insensitive and entirely composed of rho (ρ) subunits

(Johnston, 2013). GABAC has slow onset and offset kinetics (Johnston, 2013) and is

predominantly expressed in the retina (Qian and Ripps, 2001). GABAB receptors are

metabotropic G protein-coupled receptors that modulate neuronal functions via their

coupling to K+ and Ca2+ channels. Activation of K+ channels, and particularly Kir3.1

channels, leads to the generation of slow postsynaptic inhibitory potentials that

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hyperpolarize the membrane and inhibit neuronal activity by shunting excitatory

transmission (Lüscher and Slesinger, 2010). Pre- and postsynaptic GABAB receptors also

modulate voltage-gated Ca2+ channels, which affects synaptic transmission and plasticity

(Chalifoux and Carter, 2011; Pérez-Garci et al., 2006).

GABAA receptors are responsible for the most of the physiological actions of GABA in

CNS. They are pentameric receptors composed from different subunits surrounding a

central chloride ion-selective channel (Sigel and Steinmann, 2012). The differences in

subunit composition determine the functional characteristics of GABAA receptors (Farrant

and Nusser, 2005). In the mature brain GABAA receptors located in the postsynaptic

membrane mediate rapid (millisecond range) neuronal inhibition, whereas those located

extrasynaptically respond to ambient GABA and induce longer inhibition (Sigel and

Steinmann, 2012). In the developing brain, GABAA-mediated depolarization activates

voltage-gated Ca2+ channels that releases Mg2+ block from NMDA channels and induces

further increase in the Ca2+ entry (Ben-Ari, 2002). This synergistic action of GABAA and

NMDA receptors in immature neurons underlies so called Giant Depolarizing Potentials

(GDP), a pattern of synchronized large oscillations in the intracellular Ca2+, which is

required for the activity-dependent maturation and establishment of neuronal network (Ben-

Ari, 2002; Wu and Sun, 2015).

Not only GABA regulates maturation of neuronal networks, but it also modulates

proliferation of neuronal precursors, their migration, differentiation, axonal and dendritic

growth and synaptogenesis. Below, I briefly discuss the role of GABA in all these

developmental processes.

1.2.1. The role of GABA in neuronal development during embryogenesis and in

adulthood

1.2.1.1. Neuronal proliferation

GABAergic neurons first appeared at embryonic day 12 (E12) are localized in the

subventricular and ventricular zones, the areas of extensive neuronal proliferation (Del Rio

et al., 1992). At this stage, GABA plays an excitatory role and it has been shown that it

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inhibits DNA synthesis and thus proliferation of cortical progenitors (Jovanovic and

Thomson, 2011; Wang and Kriegstein, 2009). From mechanistic point of view, this

inhibition is likely due to depolarization-induced Ca2+ fluxes that affect the cell cycle

length (LoTurco et al., 1995). Interestingly, the effect of GABA on cell proliferation is cell

type- and context-specific. For example, GABA increases the proliferation of neuronal

progenitors in the embryonic ventricular zone, but conversely decreases proliferation of

progenitors located in the SVZ (Haydar et al., 2000). While the reasons for such differential

effects are still not clear, the findings indicate that the depolarizing effect of GABA on

neuronal proliferation is cell type specific.

GABA also affects proliferation of neuronal progenitors located in the adult SVZ and SGZ

of DG, two neurogenic regions in the adult brain. In the DG, GABA is released from

parvalbumin-expressing interneurons and inhibits proliferation of stem cells-like radial glia

cells through activation of the γ2-subunit-containing GABAA receptor (Song et al., 2012).

In the SVZ, GABA also depolarizes neuronal progenitors, but the mechanism of GABA

release is different (Liu et al., 2005; Wang et al., 2003). Migrating neuroblasts are

GABAergic cells (Wang et al., 2003; Snapyan et al., 2009) and release GABA in a non-

synaptic, non-vesicular fashion (Liu et al., 2005). This induces tonic activation of GABAA

receptors in the neuronal progenitor cells and decreases their proliferation (Fernando et al.,

2011; Liu et al., 2005; Nguyen et al., 2003). Both in the embryonic and adult brains

GABAA receptor activation induces phosphorylation of the histone proteins, likely via Ca2+

signaling, which in turn mediates the inhibitory effect of GABA on the cell cycle (Fernando

et al., 2011).

1.2.1.2. Neuronal migration

GABA also affects migration of neuronal precursors in the embryonic and adult brain. Both

GABAA and GABAB receptors may be involved in the migratory effects observed after

GABA application (Behar et al., 1998; López-Bendito et al., 2003; Snapyan et al., 2009;

Wang et al., 2003). Intriguingly, at higher concentration GABA promotes migration of

interneuronal precursors via activation of GABAB receptors and G-protein coupled

mechanisms, whereas at low concentration GABA increases migration of progenitors for

excitatory neurons via GABAA-mediated mechanism (Behar et al., 1998). Blocking the

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depolarizing action of GABA affects neuronal migration and circuit formation (Cancedda

et al., 2007; Manent et al., 2005; Wang and Kriegstein, 2011). It has been also shown that

GABAA receptors may have a dual role on radially migrating neurons by acting as a

migration-promoting signal in lower layers and as a STOP signal in upper cortical layer,

cortical plate (Luhmann et al., 2015).

In the adult, GABAA receptors are expressed by the neuronal progenitors in both

neurogenic regions: SGZ of the DG and SVZ-OB pathway. The role of GABAA receptors

on the migration of neuronal precursors in the SGZ has not been yet explored, mainly

because of very short migration distance that these cells propagate radially, along the

processes of radial glia (Bond et al., 2015; Bordey, 2007). Interestingly, it has been recently

revealed that these cells migrate much larger distances tangentially, along the blood vessels

(Sun et al., 2015). Since GABAergic signaling plays an important role in the vasculature-

mediated migration of neuronal precursors in the SVZ-OB pathway (Snapyan et al., 2009),

it is conceivable that GABA may also modulate tangential migration of neuronal precursors

in the adult SGZ. In the adult SVZ-OB pathway, activation of GABAA receptors on the

migrating neuroblasts reduces the speed of cell migration (Bolteus and Bordey, 2004;

Snapyan et al., 2009). GABAA receptors are also expressed by the astrocytes that ensheath

the chains of migrating neuroblasts (Lois et al., 1996; Luskin, 1998). Activation of GABAA

receptors on astrocytes induces insertion of TrkB, a high affinity BDNF receptors, on the

plasma membrane which traps vasculature-derived BDNF and leads to the entry of

migratory cells into the stationary phase (Snapyan et al., 2009).

1.2.1.3. Neuronal differentiation and maturation

GABAergic signaling controls cell differentiation by regulating expression of several key

proneuronal genes (Bertrand et al., 2002; Hevner et al., 2006). For example, application of

GABA increases expression of NeuroD (a basic helix-loop-helix transcriptional factor)

which is a downstream regulator of neuronal differentiation (Schwab et al., 2000). In

addition to promoting differentiation towards neuronal phenotype, GABAergic signaling,

which operates before the glutamatergic one, regulates neurite outgrowth and the level of

dendritic arborization (Khazipov et al., 2001; Tyzio et al., 1999). In adults, GABA also

promotes dendritic development and synaptic integration of adult-born neuronal

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progenitors (Gascon et al., 2006; Ge et al., 2006). Switching GABA action from

depolarizing to hyperpolarizing, delayed neuronal maturation (Ge et al., 2006).

Depolarizing GABA leads to the Ca2+ entry, which in turn induces activation of second

messengers and a wide range of trophic actions (Garaschuk et al., 2000; Leinekugel et al.,

1997; Yuste and Denk, 1995). It has been shown that GABA-induced Ca2+ entry promotes

microtubules stability (Gascon et al., 2006), which is known to play an important role in

dendritic stability (Bray et al., 1978; Dehmelt et al., 2003; Henley and Poo, 2004).

Altogether, these data suggest that GABA via GABAA receptors plays a pivotal role in the

development of nervous system via its depolarizing action. At the end of neuronal

development, GABA becomes hyperpolarizing and exerts its inhibitory role in neuronal

networks. How this shift from depolarizing to hyperpolarizing action is orchestrated?

Among the plethora of channels and transporters that regulate intracellular Cl-

concentration, two have emerged as being major players: the chloride importer and the

chloride exporter. In the next chapter, I will briefly describe the regulation of Cl-

homeostasis in the cells and will focus on the role of potassium-chloride transporter

member KCC2 in neuronal development, which is one of the objectives of my thesis.

1.2.2. Cation-chloride cotransporters

Maintaining a proper concentration of intracellular chloride is crucial for controlling cell

volume and pH and it also plays a role in the neuronal response to GABA and glycine

(Misgeld et al., 1986). This is done by electroneutral cation-chloride cotransporter (CCC)

family, which in turn are part of the bigger family of solute carriers (SLC) of around 300

transporter proteins (Hediger et al., 2004). SLC transporters are present in the plasma

membrane and in the intracellular membranes of nearly all cells and organelles, where their

main function is the intake/efflux of sugars, aminoacids, nucleotides, ions.

The CCC family regulates ion homeostasis via influx or outflux of ions, depending on

electrochemical gradients provided by active transporters. Basing on their ion transport

features and peptide sequences, the CCCs are classified into two groups: the Na+-K+-

dependent Cl- importers and the K+-dependent Cl- extruders (Gagnon and Delpire, 2013).

Using the K+ gradient, the K+/Cl- cotransporters (KCC) extrude Cl- out of the cell, whereas

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the Na+ gradient is used by Na+/K+/Cl- cotransporters (NKCC) to collect Cl- into the cell.

To date, four K+/Cl- cotransporters (KCC1 to KCC4) and two Na+/K+/Cl- cotransporters

(NKCC1, NKCC2) are known. NKCC1 and KCC1 have been found in all types of tissues,

and NKCC2 is expressed only in kidney. KCC2 is specific to neural cells only, while the

expression KCC3 and KCC4 has been shown both in CNS as well as in other tissues

(Delpire and Mount, 2002; Payne et al., 2003).

All CCCs transport equal amounts of cations and anions each cycle, so the net result of

charge transfer through the plasma membrane is eventually electroneutral. This is

particularly important for excitable cells, including muscle and nerve cells, because

electroneutral activity of CCCs allows for regulation of intracellular ion concentrations

while maintaining membrane potential at the same level (Payne, 2012).

Present in all types of tissues, the CCCs are involved in wide range of physiological

processes, such as regulation of the cell volume and the blood pressure, transport of solute

and water (Arroyo et al., 2013; Blaesse et al., 2009; Gagnon and Delpire, 2013; Gamba,

2005; Hebert et al., 2004). Dysfunction of CCCs has been shown to have a relation to

various heterogeneous diseases, including arterial hypertension, osteoporosis, cancer, and

epilepsy (Gamba, 2005).

1.2.3. The role of KCC2 in brain development

The neuron-specific KCC2 is a glycoprotein consisting from 12 transmembrane domains,

an N-glycosylated extracellular domain between the 5th and the 6th transmembrane

domains, and two intracellular domains: the N-terminal domain (NTD) and C-terminal

domain (CTD), adjacent to the transmembrane domains (Fig. 8; Gerelsaikhan et al., 2006;

Kahle et al., 2013; Kyte and Doolittle, 1982). CTD is also where the majority of

phosphorylation occurs (Chamma et al., 2012; Song et al., 2002), with some of these sites

within CTD known to be implicated in regulatory phosphorylation of KCC2 during

development and in response to neuronal activity (Chamma et al., 2012; Kahle et al., 2013).

In mammals, the first exon of KCC2 can be differentially spliced, giving rise to two

isoforms with similar co-transport properties, the KCC2a and KCC2b (Uvarov et al., 2007).

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KCC2a isoform is expressed at comparatively low levels throughout pre- and postnatal

development, whereas the KCC2b undergoes significant up-regulation postnatally (Stein et

al., 2004; Uvarov et al., 2007, 2009). It is thought that KCC2 is responsible for the

Figure 8. Regulation of KCC2 functions by transcriptional control, subcellular targeting

and protein phosphorylation (adapted from Blaesse et al., 2009; Kahle et al., 2013).

Upper left: Several regulatory mechanisms of transcriptional control of KCC2. Upper right: Important

regulatory phosphoresidues of KCC2. Orange dots indicate the positions of phosphoresidues in the

cytoplasmic C terminus of the transporter that are critical for functional regulation of KCC2, including

tyrosine 903 (Y903), threonine 906 (T906), serine 940 (S940), threonine 1007 (T1007), and tyrosine 1087

(Y1087). The pink region denotes the KCC2 'ISO' domain, required for hyperpolarizing GABAergic

transmission. Lower left: Protein 4.1N anchors the KCC2 expressed in spine to the cytoskeleton, however it is

unknown if it does also to KCC2 of dendritic shafts and somata (question mark). Lower right: Kinases and

phosphatases acting on KCC2 can influence endo- and exocytosis rates, thus modifying the KCC2 function. It

is not known if the intrinsic rate of ion transport of KCC2 is directly controlled (question mark).

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“developmental shift” and changing the GABA action from depolarizing to

hyperpolarizing (Blaesse et al., 2009).

The reversal potential for GABA responses (EGABA) is established by the electrochemical

gradient of Cl-. As mentioned above, in the immature neurons there is high amount of

intracellular Cl-. This results in EGABA shift to the values higher than the resting membrane

potential, which in turn is causing a depolarization when GABAA receptors are activated.

With the developmental upregulation of KCC2, the intracellular Cl- decreases because of

extrusion of Cl- ions from the cell and EGABA settles at levels more negative than the resting

membrane potential. This creates the hyperpolarizing response to GABA in mature neurons

(Rivera et al., 1999).

In vitro and in vivo models with diminished or overexpressed KCC2 have shown a

functional link of KCC2 with the development and function of GABAergic and

glutamatergic synapses (Blaesse et al., 2009; Chamma et al., 2012). First to demonstrate the

causal role for KCC2 in creating the driving force for hyperpolarizing effects of GABA was

a study where KCC2 was knocked down using antisense oligonucleotide (Rivera et al.,

1999).

Apart from well-known physiological function in establishing the response of neurons to

activation of GABAA and glycine receptors, KCC2 was recently also found to play an ion-

transport independent role in the structural maintenance of glutamatergic synapses in

studies using transport-inactive mutants of KCC2 (Fiumelli et al., 2013; Horn et al., 2010;

Li et al., 2007). Fiumelli and colleagues used in utero electroporation with post hoc

iontophoretic injection of Lucifer Yellow in layer 2/3 pyramidal neurons of somatosensory

cortex. The results indicate that premature up-regulation of KCC2 leads to significant

increase in dendritic spine density of pyramidal neurons, resulting in elevated spontaneous

excitation in KCC2-transfected neurons. The increase in spine density was observed also in

chloride transport deficient KCC2-mutant cells with deleted N-terminal of KCC2. On the

other hand, the cells with transfected C-terminal domain of KCC2, which is related to

development of dendritic cytoskeleton, also had increased spine density (Fiumelli et al.,

2013). Overexpression of KCC2 was shown to affect the proper development in mouse

embryo areas related to the neural tube and neural crest. In transgenic embryos of E9.5-11.5

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age the neuronal differentiation has been diminished with thin neural tube and anomalies in

neural crest and embryo body. Later, at E11.5-15.5 they had notably smaller brain

structures and a characteristic cleft palate. Using transport-lacking variant of KCC2 with

deleted N-terminal resulted in similar results, suggesting that they are not dependent on

KCC2 Cl- transporter function (Horn et al., 2010). To examine the role of Cl- gradient from

the earliest embryonic stages, Reynolds and colleagues overexpressed KCC2 in zebrafish

embryos, thereby inverting Cl- gradient. As a result, glycine by hyperpolarizing all neurons

of CNS induced reduction in number of motoneurons and interneurons, resulting in smaller

brain and axonal tracts, as well as disrupted motor functions. These findings advocate for a

critical role of chloride-mediated excitation in neurogenesis from the onset of embryonic

development (Reynolds et al., 2008). Therefore, when studying the implication of KCC2 in

diseased or injured brain, it is important to consider both the ion transport role as wells as

other, ion transport unrelated functions of this multifunctional protein.

1.2.4. The role of KCC2 and Cl- gradient in neuronal migration

Proper neuronal proliferation, migration and synaptic integration depend on timely actions

of various mechanisms. The migration of neuroblasts in the RMS is regulated by molecules

of extracellular matrix, cell adhesion molecules, chemoattractors, chemorepellents and

neurotransmitters. The tonic application of GABA activating the GABAA receptors was

demonstrated to depolarize the neuroblasts leading to decrease of their migration (Bolteus

and Bordey, 2004; Wang et al., 2003). Eventually it is the concentration of intracellular Cl-

and the transmembrane Cl- gradient which define if the effect imposed by GABAA receptor

activity on the cell membrane potential will be depolarizing or hyperpolarizing (Ben-Ari et

al., 1989; Misgeld et al., 1986; Mueller et al., 1984). The concentration of intracellular Cl-,

in turn, depends on the balance of activity of NKCC1 Cl- importer and KCC2 Cl- extruder.

GABA neurotransmitter signalling via GABAA receptors, is a major signalling function

present in all stages of neuronal development (Henschel et al., 2008; Young and Bordey,

2009). A series of in vitro and in vivo experiments in embryonic, young and adult animals

where GABAA receptor signalling was pharmacologically impaired, has demonstrated its

regulatory role in proliferation of neurons (Cesetti et al., 2011; Liu et al., 2006) and in their

ability to develop dendritic branches (Ge et al., 2006; Wang and Kriegstein, 2008). GABAA

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activity is also reported to control dendritic growth by stabilizing lamellipodia in migratory

neuroblasts of the OB in acute slices (Gascon et al., 2006).

In immature neurons, due to high expression of NKCC1 Cl- importer and lack of KCC2 Cl-

extruder, the activation of GABAA receptor leads to depolarization (Bordey, 2006; Owens

and Kriegstein, 2002; Platel et al., 2010). Recently, in vivo function of GABAA signaling

on neural stem cell proliferation and dendrite development has been studied by using

electroporation and short-hairpin (sh) RNA against the NKCC1 cotransporter (shNKCC1)

(Young et al., 2012). Premature reduction of GABAA-induced depolarization in neuronal

progenitors and neuroblasts by shNKCC1 in NPCs of mouse SVZ has led to reduction in

proliferation and density of newborn neurons. The density of neurons is recovered when

using an inducible Cre-encoding plasmid to target GABAA activity of immature neurons

only. Loss of GABAA depolarization also negatively impacts the dendritogenesis in critical

stages of integration and plasticity of newborn cells (Nissant et al., 2009). Early in

development, a depolarizing GABA signaling arises in some regions of CNS which is

thought to regulate the excitability of neurons leading to their proper maturation (Gascon et

al., 2006; Ge et al., 2006). Interestingly, NKCC1 is also often expressed in developing

brain, and it is shown to act in synchrony with depolarizing GABAA activation (Delpy et

al., 2008; Ge et al., 2006; Plotkin et al., 1997; Wang and Kriegstein, 2008; Yamada et al.,

2004). The role of NKCC1 in shaping the GABA activity during migration and maturation

of neuronal precursors has been explored in another recent study (Mejia-Gervacio et al.,

2011). Manipulations of NKCC1 activity revealed a strong dependence of GABAA-induced

depolarization in RMS neuroblasts from this chloride transporter. In more mature GCs

though, the driving force of depolarizing GABA is much weaker and interestingly appears

to be unaffected by manipulations of NKCC1 activity (Mejia-Gervacio et al., 2011).

It has been reported that KCC2 mRNA is not expressed in neuronal progenitors, whereas it

is abundant in mature neurons (Blaesse et al., 2009; Rivera et al., 1999). After the second

postnatal week, coincidentally with the appearance of KCC2 expression, it was

demonstrated that GCs begin to be hyperpolarized by GABA (Wang et al., 2005). With

maturation, NKCC1 becomes functionally down-regulated, leading to decrease in [Cl-]i in

GCs, and hyperpolarization of EGABA. Later, the up-regulating KCC2 gets involved in

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synaptic connectivity of GCs, similar to other systems (Cancedda et al., 2007; Li et al.,

2007).

1.3. Tools for imaging neuronal development

1.3.1. Fundamentals of optical microscopy

Optical microscopy is basically the visual observation of how light interacts with a sample.

For centuries microscopy has been used to investigate small structures that are not

distinguished with the naked eye. In optical microscopy, the object is illuminated with light

and the signal, which may be transmitted, reflected, or emitted by the object, is analyzed.

The invention of the microscope is attributed to father and son Hans and Zacharias Jansen

of Middleburg, Holland in the end of XVIth century. The optical construction made of two

lenses and diaphragms mounted inside moving tubes, allowed to see cells with up to nine

times magnification. The first documented experiments using a microscope were done by

Robert Hooke in 1665. He actually introduced the term "cell" for the small structural units

observed in the thin slices of cork (Croft, 2006). A decade later, van Leeuwenhoek with his

improved microscope was the first to observe and describe single cellular organisms, as

well as muscle fibers, bacteria, spermatozoa and blood flow in blood vessels.

In the optical microscope, the magnified image of the specimen is projected onto the image

Figure 9. Basic configurations of a modern compound microscope with an infinity-

corrected objective lens (Hornak, 2002).

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detector which can be a camera or an eye. Fig. 9 illustrates the schematic layout of a

modern microscope with an infinity-corrected objective lens. The objective lens collimates

the light from a point object in the object plane. A tube lens is generally used to focus the

collimated light onto the intermediate image plane located at its back focal plane. The

microscope generally consists of components including light source, illumination system,

condenser, diaphragms, the stage, specimen in air or immersion fluid, microscope

Figure 10. Light path through a compound microscope with transmission geometry

(Hornak, 2002).

The condenser forms a well-defined light cone that is concentrated onto the specimen. Then light passes

through the specimen into the objective, which then projects a real, inverted, and magnified image of the

specimen to a fixed plane within the microscope. The distance between the backfocal plane of the objective

and the intermediate image is called the optical tube length. The lens of the eyepiece further magnifies the real

image projected by the objective. The eye of the observer sees this secondarily magnified virtual image as if it

were at a distance of 25 cm from the eye.

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objective, tube, tube lens, eyepiece, filters, polarizers, and other optical elements. In

transmission optical microscope, such as in Fig. 10, the light source is on one side of the

sample and the light detection part is on the other side. For thick or opaque specimens epi-

illumination configurations are employed, where illumination light and the light from the

specimen are on the same side (Fig. 11). An important progress in biological imaging has

been made with the invention of fluorescence microscopy (Fig. 11). Fluorescence is a

quantum mechanical process of relaxation of a molecule from an excited state (higher

energy) to its ground (lower energy) state, which is visible as optical radiation by the

emission of a photon (Fig. 12).

Being mostly conjugated hydrocarbons, fluorescent molecules (also called fluorophores)

are typically targeted to attach to other complex molecules, like proteins and antibodies.

Figure 11. Schematic diagram of an epifluorescence microscope

Fluorescence microscope typically includes an excitation filter, an emission filter, and a dichroic beamsplitter.

The white light from arc-discharge lamp or other source is first filtered by excitation filter (blue) and directed

onto the specimen through the microscope objective, and then using that same objective the emitted

fluorescence is selected by emission filter (red) and is captured by camera or other light detector.

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Fluorophores in a biological system are commonly categorised as intrinsic or extrinsic. The

intrinsic fluorophores are the proteins or small molecules in cells that are naturally

fluorescent, such as the green fluorescent protein (GFP). Alternatively, using an extrinsic

fluorophore, one can label proteins, nucleic acids, lipids, or small molecules. The most

commonly used fluorophore is GFP, which is a naturally inherent protein in the

luminescent jellyfish Aequorea victoria (Shimomura et al., 1962). The GFP fluorescence

occurs when the protein is illuminated with 488 nm light, leading to emission around 520

nm. The emitted photons can be detected using fluorescence microscopy techniques,

allowing for precise localization of the GFP molecule (Saggau, 2006). The GFP family

proteins are relatively compact, and chemically inert. Among many uses of GFP in life

sciences are labeling of proteins and subcellular compartments in live cells, indicator for

gene activity, tracking of GFP-labeled cells in tissues.

Currently, many other types of fluorescent proteins of different colors have been discovered

and are used as fluorescent tags for imaging in biology (Stepanenko et al., 2011). Excitation

wavelengths of currently used fluorophores range from ultraviolet through the visible light

spectrum, with the emission spectra ranging from visible light into the near infrared region.

Figure 12. Simplified Jablonski diagram (Jabłoński, 1933):

A photon of higher energy (low wavelength) is absorbed by electron; then a lower energy (high wavelength)

photon is emitted.

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1.3.2. Wide-field and two-photon imaging

In a conventional wide-field fluorescence microscope (Figs. 11, 13a), large volume of the

specimen is simultaneously illuminated by a light source, which is generally a mercury or

xenon lamp. The light beam passes through optical filter (excitation filter) which selects the

wavelength of excitation light. The excitation light is projected to the sample via a dichroic

mirror, designed to be transparent to specific wavelength range, but to reflect all the others.

Then the light emitted by fluorophores of the sample passes thought the emission filter,

which rejects all the wavelengths except that of fluorophore and the final filtered signal is

detected by image sensor (Fig. 13a).

A limiting factor for applications of wide-field epifluorescence microscopy is the sample

thickness. Since the whole field of view of the objective is being illuminated, the out-of-

Figure 13. Optical layout of fluorescence microscopy techniques (adapted from Saggau,

2006):

a) epifluorescence microscopy; b) confocal microscopy; c) two-photon excitation microscopy

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focus signal intermingles with the useful one coming from the areas in focus, leading to

low-contrast images. To obtain high contrast images, optical sectioning techniques, such as

confocal and multi-photon can be implemented.

The confocal microscope eliminates the out-of-focus light with a special pinhole located at

a point conjugated with the illumination point. This design allows rejecting the photons

coming from the areas out of focus, while passing only those originating from the in-focus

regions of the sample (Fig. 13b). To acquire the image from the whole area of interest, the

illumination point is scanned across the sample, thus allowing acquisition of high resolution

three-dimensional (3D) images.

However, while confocal microscopy allows collecting photons only from focalized areas,

it still has disadvantage, as the wide-field microscopy, and requires illumination of the

entire sample area, leading to high photobleaching. This problem is resolved in the two-

photon microscope (Fig. 13c). The non-linear effect of two-photon occurs only in special

high irradiance conditions, when an electron absorbs two photons of the same wavelength

and emits a single photon with nearly half the wavelength of original photons (Fig. 14;

Helmchen and Denk, 2005; Svoboda and Yasuda, 2006). This effect occurs only at high

photon flux in the focal point of objective, where the photon density gets sufficient to allow

for the two-photon excitation.

To achieve this high light intensity, a pulsed laser source is used usually with pulse width

around 100 fs and 100 MHz repetition rate. Most widely used is the Ti:Sapphire laser with

tunable 700 – 1100 nm output range. Since only in-focus regions are being excited, this

allows for inherent optical sectioning. Another important advantage of two-photon

microscopy is the usage of excitation at infrared (IR) and near-infrared (NIR) wavelengths.

Being more transparent for these wavelengths, the biological tissue scatters and diffuses the

IR light much less than the visible light, thus allowing for high contrast imaging with

deeper penetration into the tissue (Ntziachristos, 2010).

Traditionally, many studies of adult neurogenesis are based on histological sections and

acute slices that extrapolate the acquired data about the morphology and behavior of adult-

born cells in the OB to real in vivo conditions. The direct in vivo observations of neuronal

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maturation and integration in adulthood were hampered by limited light penetration of

conventional microscopy techniques. Thanks to its high optical resolution and limited

photobleaching and photodamage, two-photon microscopy has facilitated imaging and

long-term tracking of fluorescent neuron dynamics in the tissues up to 500 µm deep.

This technique currently addresses a variety of neurobiological in vivo questions in

neurobiology, such as imaging neural activity and morphological stability and plasticity. By

combining two-photon microscopy with fluorescent calcium indicators, it has become

Figure 14. Two-Photon Excitation Microscopy (adapted from Svoboda and Yasuda, 2006)

a) Simplified Jablonski diagram of the two-photon excitation effect. b) Small volume of excitation localized

in sample (black). The objective focuses the excitation light (red) into a diffraction-limited area, exciting

green fluorescence only in that part of a dendritic branch. c) Fluorescence collection in a sample. The

excitation volume emits fluorescent photons. Even scattered fluorescence photons carry useful signal, since

all of them come only from the area in focus. d) Schematic diagram of a two-photon microscope with

epifluorescence and transmission fluorescence detection.

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possible to stain entire local populations of neurons and to image their activity patterns in

vivo on time ranges from milliseconds (Helmchen et al., 1999; Svoboda et al., 1997) to

months (Grutzendler et al., 2002; Holtmaat et al., 2005; Trachtenberg et al., 2002; Zuo et

al., 2005). For the third objective of my thesis I used in vivo two-photon imaging of adult-

born neurons in the OB to characterize the morphological plasticity of these cells under

baseline conditions and in response to the odors.

1.3.3. Light polarization

Polarization is one of the fundamental features of light, as are the intensity, wavelength and

coherence. An essential component of the light is its electric field vector E(r; t) oscillating

in the plane perpendicular to the direction of light propagation direction. In wave theory,

polarization is associated to the pattern of oscillation by the electric field vector E(r; t) as a

function of time t at a given location r. Light is said to be linearly polarized when its

electric field vector oscillates in one direction, whereas when the rotation of the vector

describes an ellipse, then we talk about elliptically polarized light. Circular polarization is a

Figure 15. Different states of light polarization.

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special case of elliptic polarization, when electric field vector rotates in circle. Finally, if

the rotation of the electric field is irregular with its vector randomly changing the amplitude

and orientation, then the light is said to be unpolarized (Fig. 15).

To describe the light polarization and its interaction with the medium, Jones and Stokes-

Mueller formalisms are generally used (Hecht, 2002; Jones, 1941). Jones formalism is well

adapted to totally polarized states; however when depolarization is to be taken into account,

such as for light propagation in scattering medium, the Stokes-Mueller formalism is

needed. In Stokes-Mueller formalism the polarization of light is represented as a four-

element vector S (Stokes vector):

The first parameter, S0 (also called I) represents the total light intensity. The parameter S1

(or Q) is the amount of linear horizontal (x) or vertical (y) polarization, while S2 (or U)

shows the amount of linear polarization along the +45° or -45° azimuths. Finally, the

parameter S3 (or V) quantifies the amount of circular polarization with its left and right

components.

The degree of polarization is defined as:

The degree of polarization can take any values between 0 (totally unpolarized light, with Q,

U and V all zero) to 1 in totally polarized states.

From the total degree of polarization shown above, we can derive the degree of linear

polarization:

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and the degree of circular polarization:

The resulting effect of a medium on the incident light can be described by the Mueller

matrix equation:

This 4 x 4 transformation matrix M of a medium named after its inventor, Hans Mueller,

transforms the incident Stokes vector Si into the corresponding output Stokes vector So, i.e.

So = M*Si. The importance of Stokes-Mueller calculus is underlined by the fact that the

output light Stokes vector carries information about the Stokes vector of the incident light,

whereas the Mueller matrix is a function of the medium only. With the help of 16 Mij real

and measurable coefficients of Mueller matrix, it is possible to fully characterize the optical

properties of the medium (Bickel and Bailey, 1985).

1.3.4. Label-free imaging using polarized light

Due to its complexity the brain remains the least explored organ in human body. Current

imaging technologies together with recent advances in genetics have given a high impetus

in deeper understanding of development and functioning of CNS. Most of these methods

being exogenous by nature require introduction of foreign DNAs, fluorophores and

immunohistochemical labeling. In some cases, though, these exogenous techniques are

impossible or undesirable to implement, for example in human brain studies. Polarized

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Light Imaging (PLI) is a method that allows for label-free analysis of brain tissue (Axer et

al., 2000a; M. Axer et al., 2011a; Fraher and MacConaill, 1970; Larsen et al., 2007a).

In terms of light geometry there are two distinct methods of imaging with polarized light:

transmission and reflection (or backscattering) polarized light imaging (Fig. 16). In

thebasic setup of polarizer microscope with transmission geometry, the linearly polarized

light from the first polarizer passes through the histological brain section to the second

polarizer (the analyzer) which is aligned perpendicularly to the first one (Fig. 16a). The

birefringent tissue will cause the linearly polarized beam to become elliptically polarized,

and a fraction of it will then be able to pass through the analyzer and will be registered by

the camera.

Figure 16. Polarized light imaging geometry

a) transmission polarized light imaging; b) reflection polarization Imaging

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1.3.5. Polarized light imaging in biomedical applications

The tissue can cause light depolarization, the degree of which is a useful parameter for

determining tissue pathologies when imaging with polarized light (Jacques et al., 2002).

The deeper the photons penetrate into tissue, the more scattering events they experience,

hence they lose their initial polarization state and get randomly polarized (depolarized).

Therefore, polarization can be used as light "gating" method to discriminate two distinct

types of light-tissue interactions: a) superficially scattered or ballistically transmitted light

(initial polarization is preserved) and b) diffusely scattered or transmitted (depolarized)

light (Demos and Alfano, 1996; Schmitt et al., 1992). Spectroscopic polarization

measurements of superficially scattered light were used to study the changes in morphology

of cells related to pre-cancerous states (Backman et al., 1999; Gurjar et al., 2001).

The tissues with structural anisotropy have birefringent properties, making them an

interesting probe to examine their structural conditions. The birefringence is a typical

feature of many fibrous tissues such as muscle, collagen, cartilage and skin, so their

condition in pathology or treatment can be assessed by birefringence measurements.

Presence of optically active (chiral) molecules in the tissue leads to rotation of linear

polarization. Being an optically active molecule, glucose has been in the center of attention

of several polarimetric studies, as there is a potential for non-invasive measurements of

glucose concentration in blood of diabetic individuals. Polarimetry measurements have also

been used to monitor the glucose in the aqueous humor of the eye (Baba et al., 2002; Coté

et al., 1992). Being optically transparent, the eye benefits from lack of light depolarization,

a problem which hampers the development of non-invasive polarimetric measurements of

glucose in other tissues.

Polarization microscopy allows visualizing the bundles of myelin-covered fibers in

histological brain sections with high detail. The myelin sheaths are multilayer structures

composed of radially ordered lipids and proteins that act as insulators and significantly

increase the conduction speed of the action potential (Martenson, 1992; Norton and

Cammer, 1984). Owing to this highly organized anisotropic structure, myelinated fibers

exhibit optical birefringence (de Campos Vidal et al., 1980a), a type of anisotropy where

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the refractive index (RI) depends on the polarization and the propagation direction of light

(Hecht, 2002). This feature of myelinated fibers provides additional contrast when imaging

with polarized light as compared to traditional intensity imaging (Schmitt and Bear, 1937a;

Setterfield and Weaver, 1937).The latest developments in PLI demonstrated fiber tract 3D

reconstruction with quantitative estimates of the directions of principal fiber bundles in

histological sections of human brain (Axer et al., 2000a; Larsen et al., 2007a). Thanks to its

high resolution histological PLI method can be used to validate MRI data and to study fiber

architecture in more detail (M. Axer et al., 2011a).

Various neurodegenerative diseases are characterized by the damage and loss of specific

subpopulations of neurons. PLI may be used to perform label-free assessment of these

neurological changes. In the first objective of my thesis, I have used PLI to assess changes

in the myelination following one of such neurodegenerative disorder, Parkinson’s disease

(PD).

1.4. Neurodegenerative disorders. Parkinson’s disease.

Many neurodegenerative disorders, such as Parkisnon's disease (PD), Huntington’s disease

(HD) and Alzheimer’s disease (AD) are associated with a gradual loss of specific neuronal

populations.

Being the most wide-spread movement disorder in the world, the PD is characterized by

progressive degeneration of neurons of the nigrostriatal system, which is preceded synaptic

dysfunction and damage of axons (Galvin et al., 1999; Hashimoto et al., 2003; Maguire-

Zeiss and Federoff, 2003). The degeneration affects mostly the dopaminergic cells, but

other cell types are also reported to be damaged, namely the cholinergic cells in the nucleus

basalis, adrenergic neurons in the locus ceruleus, autonomic ganglia, amygdala,

hippocampus, OB (Braak et al., 2003, 2004; Braak and Braak, 2000; Jellinger, 1991;

Jellinger and Attems, 2006). A common neuropathological attribute in all forms of PD is

the generation of Lewy bodies (Shults, 2006; Spillantini et al., 1997; Takeda et al., 1998;

Trojanowski and Lee, 1998; Wakabayashi et al., 1997). These formations are composed of

α-synuclein, ubiquitin and elements of cytoskeleton.

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The main mechanism causing the neuronal degeneration in PD is supposed to be the

abnormal mutations, such as oligomerization and aggregation of α-synuclein in the cell

synapses and neurites (Hashimoto et al., 2004), hence a potential therapy should be aimed

to diminish these mutations of α-synuclein. Current treatments though use symptomatic

approaches, such as the efforts of increasing the level of dopamine in the nigrostriatal

system. Cell replacement therapies appear to be promising, for instance dopaminergic cell

transplants from fetus are shown to survive and develop in the brain of PD patient (Hagell

and Brundin, 2001).

However, the lack of donor tissue, variability in the outcome, and adverse side effects

(graft-related dyskinesias) in some patients are currently slowing down the progress of cell

replacement therapies in PD (Björklund et al., 2003). The neurogenic niche of adult

mammalian forebrain, on the other hand, provides an interesting and accessible pathway for

providing cellular replacement.

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2. Thesis objectives

Adult neurogenesis is regulated by diverse cellular and molecular mechanisms. It is

traditionally studied by immunohistochemical, viral and molecular biology tools in the

fixed tissues. However, adult neurogenesis is very dynamic process, encompassing among

other processes, the migration and maturation of neuronal progenitors. It is therefore crucial

to monitor these processes directly to depict their dynamic and pin-point specific

mechanisms required for their development.

In addition, tools to study adult neurogenesis are all exogenous in nature and require

introduction of viral vectors or fluorophores into the cells or analysis of fixed samples. This

may lead to some undesired effects, and it is important to develop label-free methods to

study different processes underlying adult neurogenesis.

I therefore started my PhD thesis by addressing these two challenging topics in the field.

Neuronal precursors migrate in the specialized structure called RMS, where thousands of

these cells are aligned along each other, and along parallelly aligned blood vessels and glial

tubes. This alignment may create non-uniform, anisotropic structures which can be detected

by polarized light imaging. I hypothesized that PLI in both transmission and reflection

geometry might be used for label-free detection of migratory cells in the RMS.

I started my PhD by testing this hypothesis and establishing PLI in transmission and

reflection geometry in brain slices. Unfortunately, label-free detection of migrating cells

appeared to be extremely challenging and not reproducible. In addition, low signal from

migrating cells was hidden by very large signal from myelinated axons. I therefore decided

to adapt my first PhD project and use PLI for label-free imaging of myelinated fibers under

normal and disease conditions. I was particularly interested in Parkinson disease (PD) since

little is known about changes in myelination in PD. Although PLI does not provide direct

information about fiber number or fiber density in white matter, nevertheless the reliable

fiber directions datasets may help in indirect comparative estimates for these parameters

between groups of normal and diseased brains. In the first objective of my thesis, after

using PLI in mouse brain sections, I employed this imaging method to characterize changes

in the myelination following PD.

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In the second objective of my thesis, I perform time-lapse imaging of virally labeled

neuroblasts in the adult brain slices to study the dynamics of their migration and investigate

some of the mechanisms required for their displacement from the SVZ into the OB.

GABAergic signaling plays a major role in the migration of neuroblasts and its action is set

by the gradient of Cl- along the membrane. KCC2 is the main co-transporter that extrudes

Cl- ions from the cell and allows lowering intracellular Cl- concentration. The role of KCC2

in neuronal migration remains unknown. We hypothesize that because of late

developmental expression profile of KCC2, this co-transporter plays a role in the radial

but not tangential migration of neuroblasts. I used recently developed KCC2 activator to

address this question.

Finally, in the third objective of my thesis I performed in vivo two-photon imaging of adult-

born neurons in the OB to explore their structural modification under baseline conditions

and following odor presentation. It is well accepted that continuous supply of new neurons

constantly sculpts the bulbar network in response to changing environmental conditions.

However, there is an important conceptual problem since environmental changes can be

very rapid, whereas the synaptogenesis of adult-born neurons occurs over a longer time

scale. How bulbar network functions when rapid and persistent changes in

environmental conditions occur, even though new synapses have not yet been formed? I

performed in vivo two-photon imaging at relatively rapid timescale to address this question.

My data complement the previous work in the lab and altogether reveal a new form of

structural plasticity in the adult OB.

Thus by bringing together different imaging approaches I have tried to better understand

many aspects of adult neurogenesis, and in particular neuronal migration and plasticity of

adult-born neurons once they are fully integrated into the bulbar operational network.

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3. Results - Comparative study of myelinated fiber

bundles with polarized light imaging under normal

and pathological conditions

Karen Bakhshetyan1, Gurgen Melkonyan2, Martin Parent1, Tigran Galstian2 and Armen

Saghatelyan1

CRIUSMQ1, COPL2, Université Laval, Québec

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3.1. Résumé

Grâce à l’imagerie sans marquage utilisant la lumière polarisée, nous avons analysé des

sections sagittales, coronales et horizontales de cerveau de souris, réalisant ainsi une carte

des fibres cérébrales en lumière transmise à polarisation croisée. Via la détection de la

biréfringence des gaines de myéline, cette méthode permet la transmission de la lumière

polarisée dans le but d’évaluer quantitativement l’orientation des fibres au sein des tranches

imagées. Nous avons en outre mis en œuvre cette technique dans le but de réaliser une

étude comparative des faisceaux de fibres de la capsule interne sur des coupes histologiques

de cerveau humain en conditions normales ou pathologiques (maladie de Parkinson).

L'analyse des données obtenues a révélé des différences significatives entre ces deux

groupes concernant l’intensité des fibres imagées en lumière polarisée. A l’heure actuelle,

aucune précédente étude n’a rapporté d’altérations des fibres de myéline au niveau de la

capsule interne des patients Parkinsoniens. Nos résultats mettent ainsi en évidence

l'importance de nouvelles études d'imagerie multimodale au niveau de zones du cerveau qui

ne sont généralement pas connues comme étant associées aux maladies neurodégénératives.

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3.2. Abstract

By using label-free imaging with polarized light, we made an analysis of mouse brain

histological sections in sagittal, coronal and horizontal cuts, creating hereby a map of

mouse brain fibers in cross-polarizer transmission light. By detecting the birefringence of

the myelin sheaths, this method uses the transmission of polarized light to quantitatively

estimate the fiber orientation in the imaged section. We further implemented this technique

for comparative study of fiber bundles of internal capsule in the histological sections of

human brain under normal and pathological conditions (Parkinson’s disease). The analysis

of obtained data has shown significant differences of fiber intensities imaged in polarized

light between these two groups. There are no previous reports of myelin damage in the

fibers of internal capsule of Parkinson patients and as such our results highlight the

importance of further multimodal imaging studies in the areas of the brain which are

generally not known to be associated with the neurodegenerative diseases.

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3.3. Introduction

Various neurodegenerative diseases are associated by the damage and loss of selective

subpopulations of neurons leading to progressive dysfunction of specific brain systems. For

example, the pathology of PD is characterized by progressive accumulation of

intraneuronal inclusion bodies, namely the different types of so-called Lewy neurites (LN)

within neuronal processes and Lewy bodies (LB) in the perikarya of nerve cells (Gibb and

Lees, 1991; Pollanen et al., 1993). The formation of these inclusion bodies is associated

with aggregations of a misfolded protein, α-synuclein (α-SN) with subsequent damage of

dopaminergic nerve cells (Damier et al., 1999). These cells have axonal projections mostly

to dorsal putamen and caudate nucleus, known to be most severely dopamine-depleted

regions of the striatum in the PD (Damier et al., 1999).

The dopaminergic dysfunction in PD has been successfully studied by functional imaging

approaches, such as positron emission tomography (PET) and single photon emission

computerized tomography (SPECT), at clinical (Marek et al., 2001; Morrish et al., 1996;

Vingerhoets et al., 1994) and preclinical stages (Hilker et al., 2002; Piccini et al., 1999;

Ponsen et al., 2004), as well as to study the effects of therapies with neuroprotective agents

(Parkinson Study Group, 2002; Rakshi et al., 2002; Rascol et al., 2000; Whone et al.,

2003).

For studying the structural changes in PD, imaging approaches such as transcranial

sonography (TCS) and magnetic resonance imaging (MRI) are used. Reports with TCS in

PD patients demonstrate increased echogenicity from the lateral midbrain, hinting on

increased amounts of iron deposition in substantia nigra (SN) (Becker et al., 1995; Berg et

al., 2001; Walter et al., 2003). Spatial resolution is a limiting factor of this technique;

therefore it requires histological verification to localize the precise position and to confirm

disturbances of iron metabolism. MRI methodologies, on the other hand, allow pinpointing

the area of abnormal iron deposition with sub-centimeter precision (Hu et al., 2001;

Hutchinson and Raff, 2000; Michaeli et al., 2007).

A recent development of MRI technique, voxel based morphometry (VBM) [72] localizes

significant changes in grey matter density related to disease. VBM studies analyzing

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patterns of brain atrophy in PD have revealed progressive cortical atrophy in PD patients

with dementia (Burton et al., 2004; Nagano-Saito et al., 2005; Ramírez-Ruiz et al., 2005).

Diffusion tensor imaging (DTI) and diffusion tensor tractography (DTT) are other new

MRI techniques that have increasingly been employed to detect and provide differential

diagnosis of parkinsonian syndromes. DTI analyzes the directionality and magnitude of

water diffusion in each voxel, thus allowing evaluation of fiber tract integrity in white

matter. In healthy brain the water diffusion is anisotropic, as it is constrained along nerve

fibers. Degeneration of tracts leads to loss of directional diffusivity of water, which can be

detected by DTI (Chan et al., 2007; Matsui et al., 2007; Worker et al., 2014; Yoshikawa et

al., 2004). DTT, on the other hand, is an analytical methodology for reconstruction of major

fiber bundles in the brain based on the anisotropy of water movement in myelinated axons.

The projection axons that are predisposed to develop the pathology are long, thin and

unmyelinated or poorly myelinated, while sturdily myelinated axons are resistant to

formation of LNs/LBs (Braak and Braak, 2000; Morrison et al., 1998). Compared to

myelinated axons, the maintenance of unmyelinated axons requires significantly higher

energy (Nieuwenhuys, 1998), possibly resulting in continuous oxidative stress, which is

considered an important factor in the pathogenesis of PD (Dias et al., 2013).

The myelination of axons takes place in the final stages of cortical maturation, as a result

the areas that myelinate last (or do not myelinate at all) are the most prone to develop the

lesions. Interestingly though, some recent DTI and VBM studies hint on degeneration of

myelinated regions, such as olfactory tract (Rolheiser et al., 2011; Scherfler et al., 2006)

and brainstem (Jubault et al., 2009) in early PD patients. The loss or decrease of anisotropy

detected by MRI reflects water diffusion changes in oriented structures such as the

membrane, myelin and elements of cytoskeleton (Le Bihan, 2003). However, interpretation

of DTI data is not always easy; for example the measurement results may reflect axonal

damage (increased axial diffusivity; Aung et al., 2013; Budde et al., 2009) or myelin

damage (increased radial diffusivity; Aung et al., 2013; Klawiter et al., 2011). Despite

continuous efforts of improving the characterization of brain changes (Alexander, 2008;

Assaf et al., 2008; Jensen et al., 2005), the specificity of the diffusion parameters in MRI

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(Foerster et al., 2013; Surova et al., 2016) as well as sonography results in TCS (Berg et al.,

2001) often remain suboptimal.

The spatial resolution of current diffusion MRI tools usually doesn’t exceed the millimeter

scale (Gaggl et al., 2014), which, although better than in TCS, is still not sufficient to detect

and more accurately portray the minute changes in early stages of neurodegeneration.

Therefore, the validation of MRI results is often carried out by high-resolution microscopes

using traditional histological staining or fluorescent markers. However the applications of

these methods are quite limited for in vivo studies of human brain and, on the other hand,

meaningful analysis of unlabeled brain tissue with traditional wide-field microscopy is

rather challenging due strong scattering of visible light by such non-uniform medium

(Tuchin, 2015). Therefore, there is a growing demand in high-resolution imaging methods

that can use intrinsic optical properties of brain tissue. One promising label-free imaging

method is based on using polarized light to visualize highly anisotropic myelinated fibers

(Axer et al., 2000b; M. Axer et al., 2011b; Göthlin, 1913; Larsen et al., 2007b). Having

spatial resolution in micrometer range and being label-free imaging method, polarized light

imaging (PLI) has been shown as a suitable tool to complement and verify MRI data (H.

Axer et al., 2011). Although this technique has been mostly used for post-mortem analysis

of histological brain sections in transmission geometry, it can potentially be extended to in

vivo studies in reflection geometry, during a surgery or medical diagnostic work.

Observations of nerve fibers with polarized light began more than a century ago

(Brodmann, 1903; Göthlin, 1913; Schmitt and Bear, 1937b; Setterfield and Weaver, 1937).

The anisotropy of fibers is attributed to myelin sheaths covering the axons. The myelin

sheaths are multilayer structures that act as insulators and significantly increase the

conduction speed of the electrical signals in the axons (Martenson, 1992; Morell, 1984).

They are composed of radially ordered lipids and proteins, glycoprotein, and myelin basic

protein arranged tangentially to the axon. Owing to this highly organized structure, the

myelin sheaths exhibit optical birefringence (de Campos Vidal et al., 1980b), a type of

anisotropy where the refractive index (RI) depends on the polarization and the propagation

direction of light (Hecht, 2002). Therefore the birefringence of a myelinated fiber is a

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function of two principal refractive indices, parallel (npar) and orthogonal (nort) to the

physical fiber axis:

∆n ≈ (npar – nort)cos2α (1)

where α is the inclination of the fiber is inclined by the angle α with respect to the front of

light wave (Larsen et al., 2007b).

A linearly polarized light beam passing through a birefringent sample such as a thin section

of white matter, splits into two perpendicular components, the ordinary and the

extraordinary ray and becomes elliptically polarized. Since the propagation speeds of these

two rays are not the same, this causes a phase shift δ and a difference in amplitude of rays

depending on fiber orientation. The induced phase shift depends on light wavelength λ, the

thickness of section d, and the optical birefringence ∆n (Hecht, 2002):

δ = 2πd∆n/λ (2)

These birefringent effects can be detected when using an imaging technique based on use of

linear polarizers. In the basic polarized light imaging (PLI) system, the histological brain

sections are put between two crossed linear polarizers and an unpolarized light is passed

through the system (Fig. 1). The first polarizer filters cuts down the light to a linearly

polarized beam, which then passes through the section. If the section is not birefringent

then there will be no change in the polarization state of the beam and it will be totally

absorbed by the second polarizer. If the section is birefringent it will cause the linearly

polarized beam to become elliptically polarized, and a fraction of it will then be able to pass

through the second polarizer (also called analyzer) and will be recorded by the camera. As

the results obtained by this imaging system depend on the orientation of fibers in the

imaging plane, therefore the imaging of brain section should be done at several different

rotation angles (ρ) of polarizer-analyzer assembly, while always keeping their mutual

orientation unchanged.

Due to the birefringence of the fiber, the measured light transmittance I varies in a

sinusoidal manner with respect to the rotation angles ρ, depending on fiber orientation in x-

y imaging plane (ϕ), and its longitudinal inclination in z-axis (α). PLI method uses linear

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optical elements and the light reaching the sample from first polarizer is fully polarized, so

the light tramsittance by system can be described by Jones calculus (Jones, 1941; Larsen et

al., 2007b):

I = I0 sin2(2ρ − 2ϕ) sin2(δ/2) (3)

where I0 dentotes the intensity of the incident light, ρ is the angle of the transmission axis of

the first polarizer, ϕ is the local in-plane fiber direction relative to the zero position of the

polarimeter (ρ = 0˚), and δ is the phase shift (Eq. 2).

By measuring the intensity of light transmittance per image pixel at discrete rotation angles

ρ and by further merging together the data obtained ad different rotations, average intensity

maps of myelinated fibers can be obtained. Previous research (Axer et al., 2000b; H. Axer

et al., 2011) makes comparison between brain section images obtained with PLI and myelin

staining and the results show validity of PLI method for visualizing myelinated axonal

fibers.

In the present study, using PLI technique, we first made an analysis of mouse brain

histological sections in sagittal, coronal and horizontal cuts, creating hereby a map of

mouse brain fibers in cross-polarizer transmission light. Then we examined the histological

brain tissues from patients with PD and from healthy subjects and made a comparative

assessment of myelinated fiber bundles in the anterior limb of internal capsule between

these two groups. The internal capsule passes between caudate and putamen, both

containing the same types of neurons and circuits that are involved in parkinsonian

neurodegeneration. As, to our knowledge there are no reports about the damage of

myelinated fibers of internal capsule in PD, we were interested to check the potential

changes in this region. Our results show decreased intensity of myelinated fibers of internal

capsule observed in polarized light in PD patient tissues as compared to healthy subjects.

This data highlights the potential of high-resolution label-free imaging methods, which can

complement and validate the established neuroimaging techniques.

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3.4. Materials and methods

3.4.1. Animals

The experiments were performed in adult 4-month-old C57BL/J6 male mice (Charles

River). All the animal experiments were approved by the animal protection committee of

Université Laval. The mice were deeply anesthetized and transcardially perfused with 0.9%

NaCl followed by 4% paraformaldehyde (PFA). The brains were postfixed in 4% PFA

overnight at 4°C and then embedded in 4% agar and cut into 100μm-thick free-floating

vibratome sections (Leica TV1000S).

3.4.2. Preparation of human tissue

The postmortem human brain tissue samples were obtained from Human Brain Bank

facility at Centre de recherche de l'Institut universitaire en santé mentale de Québec

(CRIUSMQ), Université Laval. Individuals used as controls had no clinical or pathological

signs of neurological or psychiatric diseases. The Ethics Committee at Université Laval

approved the brain collecting procedures, as well as the storage and handling of post-

mortem human brain tissues. The postmortem delay for all control and PD subjects was

within 24 h period. Four male patients with PD aged 68-82 years (mean ± SD: 74.8 ± 6.42

years) and four male control subjects aged 51-82 years (mean ± SD: 70.4 ± 12.84 years)

were examined. Table 1 summarizes the clinical characteristics for each subject.

Donor

Age

Sex

PM delay,

hours

Weight, gr

State

Fixation

H41 73 M 20 1320 Normal PFA

H32 51 M 21 1595 Normal PFA

H23 82 M 18 1250 Normal PFA

H47 65 M 24 1550 Normal PFA

C0002 78 M 2.5 1170 Parkinson PFA

H4 82 M 3 1350 Parkinson PFA

H1 78 M 10 1270 Parkinson PFA

C0016 68 M 17 1250 Parkinson PFA

Table 1. Clinical characteristics of human subjects

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3.4.3. Myelin staining

For the staining of myelin sheaths we based on the staining protocol (Kluver and Barrera,

1953) using luxol fast blue solution (Sigma). The sections were mounted on slides and

dried overnight. Then the luxol fast blue solution was heated in oven maintaining at 56°C

and the slides were immersed into solution for 12h. After staining in the Luxol solution the

sections were first immersed into 95% ethanol and then distilled water, during 10 to 30 sec,

respectively. The sections were then differentiated in 0.05% lithium carbonate for 30sec

and further in 70% ethanol for 30 sec and then rinsed in distilled water and phosphate-

buffered saline (PBS). The results were checked with microscope and if the myelinated

fibers weren’t sufficiently revealed, the differentiation steps were repeated. The sections

were coverslipped in resinous mounting medium (Depex, VWR Internation ltd).

3.4.4. Immunohistochemistry

To reveal the unmyelinated axons, both human and mouse brain sections were

immunostained with Tyrosine Hydroxylase (TH). The sections were incubate with primary

antibody anti-mouse IgG TH, 1:500 (Immunostar, Hudson) overnight at 4°C (antibody

diluted in Triton 0.5% + BSA). The secondary antibody incubation was done using goat

anti-mouse IgG Alexa 488, 1:500 (Life Technologies) for 3h at room temperature. The

sections were coverslipped in Daco fluorescent mounting medium.

3.4.5. Image acquisition and processing

The images were acquired with bright-field microscope Olympus BX5 additionally

equipped with a pair of film linear polarizers (Edmund Optics #A86-188). In transmission

light configuration, the light from halogen white lamp (OSRAM BC1469) passes through

the system from below and the section image is captured by a CCD camera (cooled color

12-bit, QICAM Fast1394). In this configuration the sample position is fixed, while the

polarizer and analyzer are rotated to achieve imaging different fiber orientations while

keeping the relative orientation of polarizers unchanged. To reveal the anisotropic

structures, such as fibers covered with myelin sheaths, we imaged using polarizer and

analyzer set with their transmission axes set perpendicularly.

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The image tiles (1388x1036 pixels each) were obtained using the motorized stage Prior

H101A of the microscope. For acquisition of the images, Image-Pro Plus 6.0 software with

Oasis-4I Turboscan stage controller module has been used. These individual tiles of images

were later stitched together with ImageJ 1.46r software using stitching plugin described

elsewhere (Preibisch et al., 2009). For imaging the human sections, Olympus 10x 0.3 NA

objective has been used yielding 2.18μm × 2.18μm pixel resolution, whereas the mouse

brain sections were imaged using Olympus 20x 0.75NA yielding 4.29μm × 4.29μm pixel

resolution.

To obtain detailed map of mouse brain in polarized light, three brain hemispheres were

serially cut into 100μm-thick coronal, sagittal and horizontal sections. 121 coronal, 44

sagittal and 47 horizontal sections were serially collected, mounted with Daco mounting

medium, and coverslipped without staining.

For each section 3 sets of mosaic images were obtained with crossed polarizers at 0º, 30º

and 60º orientation angle from X axis of the stage. These three mosaic images were stacked

together to make a composite average intensity image which was further used for

computational analysis. Imaging of human brain samples was performed at anterior (pre-

commissural) level of the human striatum according to human brain atlas (Mai et al., 2008).

Coronal, 50μm-thick sections were imaged with crossed polarizers at 0º, 30º and 60º

rotation angles and processed using same method described above. For fluorescence

imaging of TH stained samples, the same microscope has been used in epifluorescence

mode, without polarizer assembly.

Prior to each imaging session the system critical parameters (incident light intensity,

polarizer alignment, camera white balance, shading control) were calibrated to minimize

processing errors.

3.4.6. Data analysis

For comparative analysis, the human brain tissue samples from four male patients with

Parkinson disease and four male control subjects were used. The region of interest was the

anterior limb of the internal capsule.

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For every tissue sample up to 100 sub-regions with densely packed myelinated fibers were

selected in ImageJ software and for each regions the mean pixel intensities and standard

error of mean (SEM) were calculated. Similarly we selected background regions in

surrounding putamen and caudate areas, which due to lack of myelin are almost completely

dark in cross-polarizer images and calculated their respective mean pixel intensities and

SEM. We used up to 3 sections per subject and for final analysis of the mean and SEM

values for each subject’s “signal” (myelinated) and “background” (non-myelinated) sub-

region values were used.

Results are expressed as means ±SEM. Statistical significance was determined by an

ANOVA for multiple groups and a Student’s t test to compare two groups (*p 0.05, **p

0.001, and ***p 0.0001).

3.5. Results

For initial verification of PLI method, histological sections from perfused mouse brains

were used. 3 hemispheres were cut serially into sagittal, coronal and horizontal sections of

100 µm thickness and were imaged with cross-polarizer setup (Fig. 2). By rotating

polarizer-analyzer assembly around the stationary brain slice, we imaged the principal axes

of birefringent structures (fiber axes) by a CCD camera at discrete rotation angles ρ (0º, 30º

and 60º from x axis of the stage). Hereby we obtained a dataset of birefringent myelinated

fibers of whole brain comprised of 44 sagittal, 121 coronal and 47 horizontal sections.

Using this data we created a false color-coded fiber orientation map under polarized light

(Fig. 3). By using image alignment and registration algorithms this 3-plane imaging dataset

may be converted to a full 3D atlas of mouse fiber orientation maps, which though was not

in the scope of this study.

After verification of the imaging principle on mouse brain, we have extended its application

to study of human brain histological sections. We examined the samples with PD which is

characterized by damage of dopaminergic neurons in substantia nigra and loss of their

projections to the dorsal putamen in striatum. The axons of dopaminergic neurons are not

covered with myelin sheaths and therefore cannot be examined in polarized light. To reveal

the dopaminergic neurons, human Parkinson and control brain sections of internal capsule,

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as well as whole-mount mouse brain sections were immunostained with dopaminergic

neuron marker Tyrosine Hydroxylase (TH). The same sections were then imaged with PLI

method and the resulting images were merged for analysis (Fig. 4). As expected, both for

mouse and human samples there is no visible colocalization of TH-stained axons and

myelinated fibers.

It was important to see if we can find out a measurable change in degree of myelination in

Parkinson's versus the control samples in the regions adjacent to dopamine-depleted

putamen. For fiber assessment experiments we produced totally 17 high-resolution PLI

datasets of brain tissue samples from four PD patients and four control subjects, up to 3

sections each. In the quantifications we considered the closely packed myelinated axonal

fibers in anterior limb of internal capsule as “Signal”, while the “Background” values were

selected the unmyelinated putamen and caudate nucleus areas. Using ImageJ, we have

selected 31 to 117 signal regions of interest (ROI) and 9 to 23 background ROIs of

rectangular shape in 17 sections derived from totally 8 subjects under study. The areas of

ROIs ranged from 0.0023 mm2 to 31.4 mm2 (mean 1.01 mm2). For intensity comparisons

we used the signal to background ratio (SBR) defined as ratio of relative mean intensity of

myelinated area (IS - “Signal”) to mean intensity of adjacent non-myelinated area (IB -

“Background”):

𝑆𝐵𝑅 =𝐼𝑆 − 𝐼𝐵

𝐼𝐵

The intensity values IS and IB represent myelination-dependent light tramsittance through

the tissue put between two crossed polarizers. The values of IB are very low as they

represent non-birefringent background lacking myelin. IS, on the other hand, is a function of

fiber birefringence, which, when measured with technique, depends on locally prevailing

spatial orientation of fibers in the voxel.

As shown in equation 3 [I = I0 sin2(2ρ − 2ϕ) sin2(δ/2)] , the measured intensity is a function

of the orientation of polarizer ρ and are the orientation of axis of the fiber ϕ, as well as the

phase difference δ caused by birefringence. In particular cases when ρ = ϕ or δ = 2kπ the

absence of light does not necessarily mean a lack of birefringence but instead it can be

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caused by orientation of fibers or thickness of the slice. To minimize the chance alignment

where axes of fiber and the polarizer coincide (ρ = ϕ), we used the average of images at 3

orientations of the pair of polarizers (0º, 30º and 60º), while keeping their transmission axes

perpendicular. Also, by using white light illumination, we reduced the risk of phase shift

being equal to integer number of wavelengths (δ = 2kπ).

Quantification of SBR revealed a statistically significant reduction of fiber intensity of PD

compared with the control subjects (2.801 ± 0.303 and 3.724 ± 0.07 respectively; *p <

0.05), (Fig. 5). To additionally verify the relevance of selecting the myelinated and

unmyelinated ROIs basing on PLI data, we stained the sections with traditional myelin

staining using luxol fast blue solution. The staining confirmed good correlation between

two methods (Fig. 6).

3.6. Discussion

Over the two past decades, imaging approaches have evolved considerably thus allowing

for better understanding of the degenerative process in Parkinson's disease (PD) and other

neurodegenerative disorders. Functional imaging with PET and SPECT not only detects

dopaminergic dysfunction in PD, but also allows following the progression of disease as

manifested by dopamine transporter alterations (Marek et al., 2001; Morrish et al., 1996;

Pirker, 2003; Winogrodzka et al., 2003). On the other hand, the structural information

obtained by MRI shows evidence of cortical thinning in PD patients (Lee et al., 2013;

Vaillancourt et al., 2009; Zarei et al., 2013) and increase in iron levels of SN in PD patients

(Hutchinson and Raff, 2000; Martin et al., 2008; Michaeli et al., 2007).

Typically manifested by degeneration of non-myelinated dopaminergic axons, PD even at

its early stages has surprisingly been shown to affect some myelin-rich white matter areas,

particularly the olfactory tract (Rolheiser et al., 2011; Scherfler et al., 2006) and brainstem

(Jubault et al., 2009). These studies were done using recently developed diffusion MRI

techniques, which measure the anisotropy and diffusion of water molecules in the brain and

thus provide important clues of the structure and geometric organization of tracts in white

matter (Chan et al., 2007; Yoshikawa et al., 2004)..

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The techniques described above are rapidly developing but generally remain bulky, very

complex and expensive. There is a growing need for simpler methods for label-free

verification of histological sections at high spatial resolution. The polarized light imaging

method we used corresponds to these requirements and only requires a slightly modified

regular wide-field microscope. The method is based on the birefringence of the myelin

sheaths covering axons, and enables the high-resolution analysis of fiber tracts. PLI has

been recently demonstrated to complement and cross-validate the tractography results

derived from DTT for complex white matter bundles (H. Axer et al., 2011). The major

benefit of polarized light microscopy for the biology and medicine sciences lies in the

label-free imaging of structural parameters with the potential to follow the cellular

development and the tissues under physiological conditions.

Typically, the voxel size in MRI is in the mm range and one such voxel can contain tens of

thousands of fibers passing in ambiguous directions (Jones, 2010). Due to this restriction in

resolution, it is difficult to explore the details of complex fiber systems andd small fiber

tracts. With PLI, in contrast, it is possible to acheve nearly 1000-fold higher resolution (H.

Axer et al., 2011), hereby providing much higher detail about the directions of fibers.

It is important to note that, although the intensity signal of PLI is sensitive to myelination,

it carries information only about neuronal fiber orientation and it does not provide direct

evaluation of the number, density or thickness of fibers. The estimates of fiber orientation

from PLI measurements may have significant level of ambiguity, because light intensity

profile depends on several factors, such as fiber inclination, section thickness and light

wavelength. While the wavelength and thickness are controllable parameters, however the

the inclination of fibers in z direction can have any values and it can’t be derived with the

current PLI method. The fibers going along z axis, can’t be detected or will have very low

intensity profile, so they can be erroneously attributed to lack of myelin, therefore a priori

knowledge about fiber orientations can be very useful. However, with careful consideration

and selecting the slices where the fibers have minimal longitudinal inclination angles but

are instead aligned in imaging plane it is nevertheless possible to indirectly estimate the

myelination degree of PLI data. This will reduce the ambiguity in the interpretation of the

intensity profile measurements.

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With the simple two-polarizer approach that we used, the orientation of fibers in x-y is

obtained by rotating the crossed-polarizers, however since the intensity varies sinusoidally,

the orientation can only be resolved with 90º ambiguity (see eq. 3). A more advanced

approach (M. Axer et al., 2011a) helps to elimintate the orientation by the addition of a

quarterwave plate between two poarziers. In the scope of our study though detecting the

myelin-related intensity was more important that resolving the orientation of the fibers,

therefore we used only crossed-polarizers which also allowed to use less rotations of

polarizer (in range of 0º to 90º with cross-polarizers, instead of 0º to 180º when

quarterwave plate is to be added).

Therefore, for better quantification of fiber density and axon structure details, PLI needs to

be complemented with classical histological staining and imaged with high resolution

imaging modalities, such as the confocal microscope which would allow to resolve in high

detail the orientations and amount of fibers.

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3.8. Figures

Figure 1. Schematic diagram of polarized light imaging

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Figure 2. Mouse brain dataset in polarized light, sagittal sections

ML – medial-lateral distance from the midline of the brain

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Figure 3. Fiber orientation information obtained with polarized light

A) Mouse sagittal, coronal and horizontal sections, imaged in 3 angles of polarizers

B) Pseudo-color coded merge showing the directions of fibers.

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Figure 4. Comparison of polarized light imaging and TH immunostaining

A) Mouse sagittal sections

B) Human sections, internal capsule

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Figure 5. Comparative assessment of fiber intensity in polarized light between control and

PD in internal capsule.

Control subjects section in brightfield microscopy (A) and PLI (B).

PD section in brightfield microscopy (C) and PLI (D).

s and b denote “signal” and “background” ROIs respectively.

Cumulative frequency distribution and mean fiber pixel intensities in PLI (E).

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Figure 6. Comparison of PLI and Luxol Fast Blue staining for myelin, anterior limb of

internal capsule.

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4. Results - Activation of KCC2 affects radial but not

tangential migration of neuronal precursors in the

adult brain

Karen Bakshetyan1, Yves de Koninck1,2, and Armen Saghatelyan1,2

Affiliations

1Cellular Neurobiology Unit, Centre de recherche de l'Institut Universitaire en Santé

Mentale de Québec, Québec City, QC, Canada G1J 2G3

2Department of Psychiatry and Neuroscience, Université Laval, Quebec City, QC, Canada

G1K 7P4

Acknowledgements

We thank Dr. Annie Castonguay for support and valuable advices related to using the

KCC2 activator drug, Karine Bachand and Modesto Peralta for testing the drug metabolism

in mice, Dr. Louis-Étienne Lorenzo for support in immunohistochemical staining of KCC2

and Dr. Marina Snapyan for conducting the confocal imaging. A.S. holds a Canada

Research Chair in postnatal neurogenesis.

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4.1. Résumé

L’alteration de l’homéostasie chlorique dans les neurones semble responsable de plusieurs

troubles neurodéveloppementaux et sa régulation stricte est nécessaire pour le

développement normal du cerveau. Le gradient de Cl- au niveau de la membrane neuronale

est régulée par deux principaux co-transporteurs NKCC1 et KCC2 qui font rentrer et sortir

les ions chloriques, respectivement. Le modèle de la neurogenèse adulte dans le bulbe

olfactif permet d’étudier le développement neuronal et des études antérieures ont mis en

évidence le rôle de NKCC1 dans la migration neuronale. Cependant, le rôle de KCC2 dans

le processus de migration reste méconnu. Dans nos travaux actuels, nous avons utilisé un

activateur très puissant et sélectif de KCC2, CLP257 afin de faire de l’imagerie à temps réel

de neuroblastes viralement marqués. Ainsi nous avons étudié le rôle de KCC2 dans les trois

types de migrations différentes dans le bulbe olfactif adulte. Nous avons démontré que

l'activation de KCC2 n'affecte pas la migration tangentielle des précurseurs neuronaux de la

SVZ où se trouvent les cellules souches vers le bulbe olfactif. En revanche, l'application de

CLP257 a favorisé la migration radiale des neuroblastes dans le courant migratoire rostrale

(CRM) entrant dans le bulbe olfactif où les cellules migratoires changent leur mode de

migration tangentielle à radiale ainsi que la migration radiale dans le bulbe olfactive. Ces

données corroborent avec nos résultats montrant l'absence de neuroblastes KCC2

immunopositives migrant tangentiellement et la présence de KCC2 immunopositives dans

les neuroblastes qui migrent radialement. Bien que les mécanismes exacts de l'action de

KCC2 sur la migration radiale des neuroblastes restent à étudier, nos données suggèrent que

l'activation de ce transporteur favorise la migration neuronale.

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4.2. Abstract

Impaired Cl- homeostasis in neurons is thought to underlie several neurodevelopmental

disorders and its tight regulation is required for normal brain development. The Cl- gradient

across the neuronal membrane is regulated by two main co-transporters NKCC1 and KCC2

that import and export Cl- ions, respectively. Using adult OB neurogenesis as a model

system to study neuronal development, previous studies have revealed the role of NKCC1

in neuronal migration. Little is known, however, about the role of KCC2 in this process. In

the current study we used recently developed highly potent and selective KCC2 activator,

CLP257, and time-lapse video-imaging of virally labeled neuroblasts to study the role of

KCC2 in three different types of migration in the adult OB. We demonstrated that KCC2

activation does not affect tangential migration of neuronal precursors from the SVZ where

stem cells are located into the OB. By contrast, CLP257 application fostered radial

migration of neuroblasts in the rostral migratory stream (RMS) entering OB where

migrating cells switch their mode of migration from tangential to radial ones and in the OB.

These data are consistent with our results showing absence of KCC2 immunopositive

signal in the tangentially migrating neuroblasts and its appearance in the radially migrating

cells. While the exact mechanisms of KCC2 action on radially migrating neuroblasts

remain to be studied, our data suggest that activation of this transporter promotes neuronal

migration.

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4.3. Introduction

Neuronal migration is essentials for proper assembly and function of nervous system. The

neuronal migration is particularly prominent in the embryonic and prenatal brains with

thousands of neurons migrating to their final destinations and establishing synaptic

connections with their targets (Evsyukova et al., 2013). In the adult brain, the neuronal

migration is largely ceased, except few regions that retain a remarkable capacity to produce

new neurons throughout the life-span of animals. The mammalian forebrain is the largest

neurogenic region in the adult brain and neuronal precursors, born in the subventricular

zone (SVZ) bordering lateral ventricle, migrate along the RMS into the OB where they

mature and integrate into the neuronal network (Breton-Provencher and Saghatelyan,

2012). There are about 30000-40000 neuronal progenitors that arrive every day to the OB

(Alvarez-Buylla and Garcia-Verdugo, 2002; Lledo and Saghatelyan, 2005), and this

massive arrival and integration of new neurons represents an ideal model to study neuronal

migration in the adult fully-formed brain tissue.

The neuronal precursors born in the SVZ migrate first tangentially, in chains, along the

RMS and once in the OB, turn to migrate radially and individually out of the RMS into the

bulbar layers. In the adult RMS, neuroblasts travel in chains ensheathed by astrocytic

processes (Kaneko et al., 2011; Lois and Alvarez-Buylla, 1994) and their migration is under

the control of various signaling molecules, including those derived from the astrocytes

(Gengatharan et al., 2016; Saghatelyan, 2009). Cellular interactions between neuroblasts

and blood vessels are also required for faithful migration toward the OB. We and others

have demonstrated that neuronal precursors use for their migration blood vessels that

topographically outline the RMS (Snapyan et al., 2009; Whitman et al., 2009).

Furthermore, endothelial cells release trophic factors that foster neuronal migration

(Snapyan et al., 2009).

In addition to factors derived from astrocytes and endothelial cells that modulate neuronal

migration, factors released by neuroblasts themselves may affect their own migration. For

example, autocrine action of GABA on neuronal migration through GABAA receptors

present of neuroblasts has been documented (Bolteus and Bordey, 2004; Platel et al., 2008;

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Snapyan et al., 2009). Signaling through GABAA receptors depends on the intracellular Cl-

concentration and its gradient across the membrane. During neuronal development

immature cells have higher intracellular Cl- concentration and the action of inhibitory

neurotransmitters such as GABA and glycine is depolarizing. Neuronal maturation is

associated with the decrease in the intracellular Cl- concentration leading to hyperpolarizing

action of inhibitory neurotransmitters (Ben-Ari et al., 2012; Kaila et al., 2014). The chloride

gradient across the membrane is regulated by two main co-transporters Na+-K+-Cl-

cotransporter (NKCC1) and K+-Cl- cotransporter (KCC2) that import and export Cl- ions,

respectively (Kaila et al., 2014). Previous work has demonstrated that genetic or

pharmacological alteration of NKCC1 expression and/or function decreased tangential

migration of neuroblasts in the RMS (Mejia-Gervacio et al., 2011; Young et al., 2012). It

remains, however, unknown how radial migration of neuroblasts is affected following

dysregulation of Cl- gradient and if KCC2 activation modulates tangential or radial

migrations.

The role of KCC2 in neuronal development is particularly important since several

neurodevelopmental disorders, such as autism spectrum disorders are associated with the

loss of KCC2 activity, leading to the maintenance of high intracellular Cl- concentration in

the developing neurons and altered response to GABA (Tyzio et al., 2014). KCC2 is

particularly promising therapeutic target for CNS disorders, since in contrast to NKCC1

and other members of cation chloride co-transporters, its expression is specifically

restricted to the CNS (Kaila et al., 2014). In the current study, we used recently developed

selective KCC2 activator, CLP257, and time-lapse video-imaging of virally labelled

neuroblasts, to examine their tangential and radial migrations and impact of KCC2

activation on different parameters of cell migration. Our results indicate that CLP257 does

not affect the tangential migration of neuronal precursors, but fosters radial migration of

neuroblasts in the OB.

4.4. Materials and Methods

This section provides a brief description of the methods used in the current study. The

detailed description is presented in the paper published in the Current Protocols in

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Neuroscience (Bakhshetyan and Saghatelyan, 2015) which is included in the thesis as the

Annex A. In that paper we described in great detail the entire procedure for stereotaxic

injection, preparation of acute slices for tangential and radial migration, time-lapse video-

imaging of cell migration and analysis. We also discuss the troubleshooting of time-lapse

video-imaging of cell migration.

4.4.1. Animals

Adult (> 2 months old) male C57BL/6 mice (Charles River) were used for all the

experiments, which were performed in accordance with the Canadian Guide for the

Care and Use of Laboratory Animals and were approved by the Université Laval

Animal Protection Committee. The mice were kept on a 12 h light/dark cycle at a constant

temperature (22°C) with food and water ad libitum.

4.4.2. Stereotaxic injections

A GFP-encoding retrovirus was stereotaxically injected into the SVZ of both brain

hemispheres (Fig. 1a). The retrovirus was purchased from the Platform for Cellular

Imaging of the Centre de Recherche de l'Institut Universitaire en Santé Mentale de Québec.

The following coordinates were used for the stereotaxic injections (with respect to the

bregma): anterior-posterior: 0.7 mm; medial-lateral: ± 1.2 mm; and dorsal-ventral: 1.9 mm.

After the injection, the mice were returned to their cages for 5-7 days post-injection (dpi)

for analysis of tangential migration and 10-14 dpi for analysis of radial migration (Fig. 1b).

4.4.3. Immunohistochemistry

Immunohistochemistry was performed as described previously (David et al., 2013; Snapyan

et al., 2009). Briefly, the mice previously injected with GFP retroviruses were deeply

anesthetized and transcardially perfused with 0.9% NaCl followed by 4%

paraformaldehyde (PFA). The brains were post-fixed in 4% PFA overnight at 4°C, and 40-

µm-thick free-floating vibratome (VT 1000S, Leica) sections were incubated with a mouse

anti-KCC2 (1:500; Upstate) antibody and then with an Alexa Fluor conjugated anti-mouse

564 secondary antibody (1:1000; Life Technologies).

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4.4.4. Slice preparation and time-lapse video-imaging

Acute horizontal 250-µm-thick slices were prepared for time-lapse video-imaging of cell

migration in the RMS and OB. The mice were first perfused transcardially with ice-cold

sucrose-based artificial cerebrospinal fluid (ACSF) containing (in mM) 250 sucrose, 3 KCl,

0.5 CaCl2, 3 MgCl2, 25 NaHCO3, 1.25 NaHPO4, and 10 glucose. The brains were rapidly

removed and immersed in the solution used for the transcardial perfusion. Sagittal or

horizontal slices of the forebrain were obtained using a vibratome for video-imaging of

tangential and radial migrations, respectively. After a 30-min recovery period at 32°C, the

slices were incubated in CLP257 (100 µM diluted in ACSF and DMSO) or control vehicle

(ASCF with DMSO) for 2 h. There were then placed in the recording chamber and were

continuously perfused with oxygenated ACSF containing (in mM) 124 NaCl, 3 KCl, 2

CaCl2, 1.3 MgCl2, 25 NaHCO3, 1.25 NaHPO4, and 10 glucose at a rate of 1 ml/min

(bubbled with 95% O2/5% CO2; pH ≈ 7.4). The slices were perfused with or without

CLP257 depending on the pre-incubation conditions. For time-lapse video-imaging of cell

migration, multiple z-stacks images (at least 6-10 z-sections at 3 µm intervals) were

acquired every 30 s for at least 1 h for tangential migration and 2 h for radial migration.

The image acquisition was performed using a BX61WI up-right microscope (Olympus)

equipped with CCD camera (CoolSnap HQ2). Cell migration was analyzed using Imaris

software. The total displacement during 1 h of cell migration, the speed of migration, and

the percentage of the stationary phase with respect to the total migration time were

calculated. For analysis of the stationary periods, only those phases that were intercepted by

two migratory periods were used. Time-lapse video-images of the RMS, RMS of the OB

(RMSOB) and granule cell layer (GCL) were acquired to track tangential migration of

neuroblasts in the RMS and RMSOB and radial migration in the RMSOB and GCL (Fig.

1c,d).

4.4.5. Statistical analysis

Results are expressed as means ± SEM. Statistical significance was determined using

Student's t-test (*p < 0.05 and **p < 0.01).

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4.5. Results

4.5.1. KCC2 expression in the SVZ-OB pathway

To establish the expression profile of KCC2 in the SVZ-OB pathway, we retrovirally

labeled neuronal progenitors in the SVZ and carried out immunohistochemical analysis of

GFP-infected neuroblasts in the RMS and GCL. KCC2 immunofluorescence performed on

sagittal forebrain sections containing the SVZ and the RMS of adult 2-3-months-old mice

revealed that KCC2 expression is not detectable in the SVZ and posterior RMS (Fig. 2a).

In contrast, our results have demonstrated that KCC2 was expressed homogenously by all

the granule cells in the GCL (Fig. 2b). These data suggest that KCC2 expression is

upregulated in neuroblasts following their arrival into the OB.

4.5.2. KCC2 activation does not affect tangential migration in the RMS

Despite the fact that KCC2 is not detected in tangentially migrating neuroblasts in the

RMS, it is still possible that low level of this transporter is present in migrating cells and

plays a functional role. We therefore used recently developed KCC2 activator, CLP257,

(Gagnon et al., 2013) and time-lapse video-imaging of virally labeled neuroblasts

(Bakhshetyan and Saghatelyan, 2015) to directly address this question. 5-7 days after the

viral labeling, sagittal forebrain sections were prepared and pre-incubated for 2h either in

CLP257 (100 µM) or control vehicle (Fig. 1a,b). Time-lapse video-imaging of neuroblasts

for an additional hour either in the presence or absence of CLP257 did not reveal any

differences in the tangential migration of neuroblasts in the RMS (Fig. 3). The total

distance of migration (109.4 ± 4.8 µm for control and 123.9 ± 5.9 µm for CLP257 during

1h of imaging; 108 and 96 cells from 7 and 6 mice, respectively; Fig. 3a), track straightness

(0.47 ± 0.03 µm for control and 0.39 ± 0.03 µm; Fig. 3b), the speed of migration (178.2 ±

4.2 µm/h for control and 170.6 ± 3.1 µm/h for CLP257;Fig. 3c), and the duration of

migratory phases (31.7 ± 3.0 % for control and 37.7 ± 3.2 % for CLP257;Fig. 3d) were not

affected by KCC2 activation. These data are consistent with our immunohistochemical

results and suggest that KCC2 does play a role in the tangential migration of neuroblasts in

the RMS.

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4.5.3. KCC2 activation fosters radial migration of neuroblasts in the OB

Following their arrival into the OB, neuroblasts detach from each other, turn by 90 degree

and start to migrate radially in the RMSOB and then in the GCL. Since KCC2 is expressed

during late developmental stages of neurons (Ben-Ari et al., 2012), we thus asked if KCC2

activity is important for migration of more developed neuronal precursors, undertaking

their tangential migration in the RMSOB, radial migration in the RMSOB or radial migration

in the GCL. We first studied tangential migration of neuroblasts 10-14 days after viral

labeling of neuronal progenitors in the SVZ (Fig. 4). We hypothesize that if KCC2 activity

is required for stopping tangential migration and initiation of radial one, than

pharmacological activation of this transporter should reduce tangential migration. Our

experiments revealed, however, that CLP257 application does not affect the migration

distance (74.4 ± 2.9 µm for control and 77.5 ± 2.6 µm for CLP257 during 1h of imaging;

200 and 216 cells from 21 mice, respectively; Fig. 4a), track straightness (0.41 ± 0.02 for

control and 0.36 ± 0.02; Fig. 4b), the speed of migration (119.5 ± 2.6 µm/h for control and

118.2 ± 2.6 µm/h for CLP257; Fig. 4c), and the duration of migratory phases (27.1 ± 1.8 %

for control and 28.8 ± 1.6 % for CLP257; Fig. 4d). These data suggest that KCC2 activity

does not contribute for a “STOP” signal for tangentially migrating neuroblasts in the

RMSOB.

We next studied the radial migration of neuroblasts in the RMSOB. Since radially migrating

neuroblasts have slower dynamic, we performed time-lapse video-imaging for 2h,

following 2h pre-incubation of acute slices either in CLP257 or control vehicle (Fig. 5).

Interestingly, while total displacement (63.0 ± 2.5 µm for control and 69.4 ± 3.6 µm for

CLP257 during 1h of imaging; 56 and 34 cells from 17 and 15 mice, respectively; Fig. 5a),

migration speed (98.0±5.1 µm/h for control and 98.6 ± 5.8 µm/h for CLP257; Fig. 5c) and

duration of migratory phases (26.8 ± 2.4 % for control and 28.7 ± 2.2 % for CLP257; Fig.

5d) were not affected by CLP257 application, our data revealed that activation of KCC2

transporter affects the straightness of migration (0.39 ± 0.02 for control and 0.33 ± 0.02 for

CLP257; Fig. 5b). The straightness of migration is the ratio of the vector of migration to

overall path that migratory cell propagate and changes in this parameter suggest that KCC2

activity may regulate the exit of radially migrating cells from the RMSOB into the GCL.

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In the GCL, the neuroblasts continue to migrate to take their final position in the different

bulbar layers. We thus asked if KCC2 activation affect radially migrating neuroblasts in the

GCL, as well. We used the same pre-incubation and imaging approach as for radially

migrating neuroblasts in the RMSOB and observed that CLP257 application fosters radial

migration of neuroblasts in the GCL (Fig. 6). In particular, up-regulation of KCC2 activity

increased the total distance of migration (58.8 ± 3.4 µm for control and 79.7 ± 8.5 µm for

CLP257 during 1h of imaging; p=0.03 with Student t-test; 24 and 21 cells from 12 and 11

mice, respectively; Fig. 6a). This increased migration distance is likely due to the combined

effect of CLP257 on the speed of migration (80.8 ± 6.8 µm/h for control and 92.1 ± 6.0

µm/h for CLP257; Fig. 6c) and duration of migratory phases (28.3 ± 2.4 % for control and

31.3 ± 3.9 µm for CLP257; Fig. 6d). While none of these two parameters reached

significance after CLP257 application, there was a tendency for higher values in both of

these parameters after KCC2 activation. Interestingly, the straightness of migration in the

GCL had also showed some tendency towards lower values after CLP257 application (0.4 ±

0.04 for control and 0.34 ± 0.03 for CLP257; Fig. 6b). Altogether, our data indicates that

KCC2 activity is required for radial migration of neuroblasts in the RMSOB and OB, while

it has no role in tangentially migrating neuroblasts.

4.6. Discussion

The results presented in the current study needs to be elaborated further and several

questions remain unanswered. Meanwhile, our data provide the first indication that

activation of KCC2 transporter may affect radial, but not tangential, migration of neuronal

precursors in the adult mammalian forebrain. Our immunohistochemical data demonstrate

that KCC2 is not expressed in tangentially migrating cells and starts to be upregulated at

later maturational stages, when neuronal precursors reach OB. This expression profile is

consistent with the one observed during embryonic and early postnatal development, with

KCC2 being expressed late during neuronal development (Ben-Ari et al., 2012; Kaila et al.,

2014). KCC2 upregulation during development leads to the inversion of Cl- gradient and

changes the action of inhibitory neurotransmitters such as GABA and glycine from

excitatory depolarizing to inhibitory hyperpolarizing (Ben-Ari et al., 2012; Kaila et al.,

2014). In the adult RMS, the action of GABA is also depolarizing (Mejia-Gervacio et al.,

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2011) and several studies have shown the crucial role of GABA in modulating tangential

migration of neuroblasts (Bolteus and Bordey, 2004; Platel et al., 2008; Snapyan et al.,

2009). Interestingly, genetic downregulation of NKCC1, another Cl- transporter that import

Cl- ions, in tangentially migrating neuroblasts produces significant hyperpolarization of

EGABA from about -30 mV to -50 mV (Mejia-Gervacio et al., 2011). This, in turn, decreases

GABA-depolarizing responses in neuroblasts and their tangential migration (Mejia-

Gervacio et al., 2011) and reduces density of newborn cells in the OB (Young et al., 2012).

It is remains unclear how downregulation of NKCC1 affects radial migration of neuroblasts

in the RMSOB and OB and at which exact maturational stage the inversion of Cl- gradient

occurs. Our results suggest that KCC2 start to be expressed in neuroblasts reaching the

RMSOB and previous work has revealed that downregulation of NKCC1 in mature newborn

neurons does not affect EGABA (Mejia-Gervacio et al., 2011). It is therefore likely that shift

in Cl- gradient and GABA-induced responses occurs in the RMSOB when cells change their

mode of migration from tangential to radial one. In line with this are our results that

increasing KCC2 activity does not influence tangential migration of neuroblasts in the

RMSOB, but affect the radial migration in the RMSOB and GCL.

It is peculiar, however, that downregulation of NKCC1 in tangentially migrating cells and

increasing KCC2 activity in radially migrating cells induced opposite effects on cells

migration. While both of these manipulations should lead to lowering intracellular Cl-

concentration and more hyperpolarizing (or less depolarizing) GABA responses, tangential

migration was decreased (Mejia-Gervacio et al., 2011) whereas radial migration was

increased. These apparently contradictory data may be reconciled by several different

mechanisms. First, it is possible that intracellular Cl- concentration have different impact of

two different migratory modes. Second, it should be noted that KCC2 is a multifunctional

protein and, in addition of lowering intracellular Cl- concentration, it has also an ion-

transport independent role (Fiumelli and Woodin, 2007; Horn et al., 2010; Llano et al.,

2015). It is thus conceivable that that tangential migration of neuroblasts is regulated by

NKCC1 function via chloride-transport-dependent manner, whereas radial migration of

neuroblasts is regulated by KCC2 via chloride-transport-independent manner. It has been

recently shown that precocious expression of KCC2 leads to the increased spine density of

pyramidal neurons in the somatosensory neurons and this effect does not vanish when N-

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terminal deleted form of KCC2, which lacks the chloride transporter function, is expressed

(Fiumelli and Woodin, 2007). Furthermore, it has been recently shown that KCC2 may

directly affect actin dynamic during synaptogenesis (Llano et al., 2015). Downregulation of

KCC2 leads to increased stability of actin filaments in dendritic spines which can be

restored by expression of chloride-transport-deficient mutants of KCC2 (Llano et al., 2015).

These data suggest that KCC2 regulates actin dynamics in a chloride-transport-independent

manner during synaptogenesis. Therefore, it is possible that also during radial migration,

KCC2 affect actin dynamic which leads to the increased migration of these cells in the

RMSOB and GCL. Thus, according to this scenario, tangential migration of neuroblasts is

regulated by NKCC1 function via chloride-transport-dependent manner, whereas radial

migration of neuroblasts would be regulated by KCC2 via chloride-transport-independent

manner. If this is the case, then this implies that CLP257 affects chloride-transport-

independent functions of KCC2 which still needs to be investigated.

Finally, it is possible that application of CLP257 affects cell migration in non-cell-

autonomous manner. Neuroblasts migrating radially in the RMSOB and GCL are surrounded

by thousands of mature neurons that strongly express KCC2. Application of CLP257 will

not only increase activity of KCC2 in the migrating neurons, but also in the mature

neurons. This, in turn, may affect OB network functioning and lead indirectly to changes in

the radial migration. In line with this are previous reports showing that sensory deprivation,

which reduces activity of principal cells in the OB, leads to the accumulation of neuroblasts

in the RMSOB and affected radial migration (Saghatelyan et al., 2004). These effects were

attributed to the activity-dependent expression of tenascin-R (Saghatelyan et al., 2004). It

should be also noted that principal cells express Reelin which also affects radial migration

of neuroblasts (Hack et al., 2002). Thus, changes in network activity following CLP257

application may lead to the modulation of radial migration via these or other molecular cues

in the OB.

Altogether, our results highlighted the role of KCC2 in the radial migration of neuroblasts.

While this topic requires further investigation, our data indicate that KCC2 activity may be

required in cell-autonomous or non-autonomous manner for specific neurodevelopmental

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processes. This is particularly interesting since KCC2 is promising therapeutic target for

neurodevelopmental disorders.

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4.8. Figures

Figure 1. Tracking tangential and radial migration in adult brain slices.

a) Schematic diagram of adult-born cell labeling by stereotaxic injection of GFP-encoding viral particles into

the SVZ.

b) Tangential migration is monitored in the RMS with slices cut sagitally, while radial migration is recorded

in the horizontal sections of OB and the RMS of the OB.

c,d) Time-lapse imaging of GFP-labeled neuroblasts in acute RMS and OB slices respectively. The arrows

and numbers indicate the soma of different migratory cells.

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Figure 2. Comparison of KCC2 immunohistochemical staining in RMS and OB.

Expression of immunohistochemically labeled KCC2 (red) in RMS (a) and in GCL (b). In green are shown

the GFP-labeled adult born neuroblasts. Scale bar, 10µm.

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Figure 3. Tangential migration parameters of adult-born neuroblasts in the RMS in

response to KCC2 activation.

Track length (a), track straightness (b), speed of migration (c), and the ratio of migratory periods of cells (d)

measured in the presence of KCC2 activator, CLP257 versus the control vehicle analyzed upon 1 hour time-

lapse video-imaging.

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Figure 4. Tangential migration parameters of adult-born neuroblasts in the RMS-OB in

response to KCC2 activation.

Track length (a), track straightness (b), speed of migration (c), and the ratio of migratory periods of cells (d)

measured in the presence of KCC2 activator, CLP257 versus the control vehicle analyzed upon 1 hour time-

lapse video-imaging.

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Figure 5. Radial migration parameters of adult-born neuroblasts in the RMS-OB in

response to KCC2 activation.

Track length (a), track straightness (b), speed of migration (c), and the ratio of migratory periods of cells (d)

measured in the presence of KCC2 activator, CLP257 versus the control vehicle analyzed upon 1 hour time-

lapse video-imaging.

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Figure 6. Radial migration parameters of adult-born neuroblasts in the GCL in response to

KCC2 activation.

Track length (a), track straightness (b), speed of migration (c), and the ratio of migratory periods of cells (d)

measured in the presence of KCC2 activator, CLP257 versus the control vehicle analyzed upon 1 hour time-

lapse video-imaging.

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5. Results - Principal cell activity induces spine

relocation of adult-born interneurons in the

olfactory bulb

Vincent Breton-Provencher1, Karen Bakhshetyan1,#, Delphine Hardy1,#, Rodrigo Roberto

Bammann1, Francesco Cavarretta2,3, Marina Snapyan1, Daniel Côté1,4, Michele

Migliore2,3,5, and Armen Saghatelyan1,6,*

Affiliations

1Cellular Neurobiology Unit, Institut Universitaire en santé mentale de Québec, Québec

City, QC, Canada G1J 2G3

2Department of Mathematics, University of Milan, 20133 Milan, Italy

3Department of Neurobiology, Yale University School of Medicine, New Haven, CT

06520, USA

4Centre d’optique, photonique et laser (COPL), Université Laval, Québec City, QC, Canada

G1V 0A6

5Institute of Biophysics, National Research Council, 90146 Palermo, Italy

6Department of Psychiatry and Neuroscience, Université Laval, Québec City, QC, Canada

G1V 0A6

#These authors contributed equally to the work.

Acknowledgements

We thank Drs Kenneth Campbell (Cincinnati Children’s Hospital Medical Center) for

providing the CAG-CAT-EGFP reporter mice, E. Castren (the University of Helsinki,

Finland) for providing the BDNF plasmid, and Gordon M. Shepherd for useful discussions.

We also thank Mireille Massouh for preparing illustration for the cover art. This work was

funded by an operating grant from the National Science and Engineering Research Council

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of Canada (NSERC) and a grant from the Canadian Institutes of Health Research (CIHR) to

A.S. V.B.-P. was supported by a PhD fellowship from FRSQ and a training grant in

neurophotonics from CIHR. M.M. and F.C. are also grateful for support of the SenseLab

project by grant 01 DC 00997701-06 from the National Institute of Deafness and Other

Communication Disorders, the CINECA consortium (Bologna, Italy) and the PRACE

association (Partnership for Advanced Computing in Europe) for granting access to the

IBM BlueGene/Q FERMI system. D.C. holds a Canada Research Chair in biophotonics,

and A.S. holds a Canada Research Chair in postnatal neurogenesis.

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5.1. Résumé

Les néo-neurones générés chez l’adulte régulent le fonctionnement du réseau du bulbe

olfactif en réponse aux changements environnementaux par la formation, la rétraction et/ou

la stabilisation de nouveaux contacts synaptiques. Tandis que certains changements

olfactifs environnementaux sont rapides, la synaptogénèse des néo-neurones générés chez

l’adulte se produit sur une échelle de temps plus longue. On ne sait donc pas comment le

système bulbaire fonctionne lorsque des changements rapides et persistants dans les

conditions environnementales se produisent. Ici, nous révélons une nouvelle forme de

remodelage structurel où les épines matures des néo-neurones produits chez l’adulte, mais

pas celles des cellules pré-existantes, se déplacent de façon dépendante de l’activité. Ce

déplacement est précédé par la croissance de filopodes à l’extrémité des épines et est

contrôlé par l’activité des cellules principales. Le glutamate et le BDNF, dérivé de cellules

principales, règulent, respectivement, la motilité des filopodes et la direction de

relocalisation des épines; de plus, les épines avec filopodes sont sélectivement conservées

après une privation sensorielle. Notre modèle 3D a montré que le déplacement des épines

permet une réorganisation rapide du réseau bulbaire avec des conséquences fonctionnelles

sur le traitement de l’information olfactive.

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5.2. Abstract

Adult-born neurons adjust olfactory bulb (OB) network functioning in response to changing

environmental conditions by the formation, retraction, and/or stabilization of new synaptic

contacts. While some changes in the odor environment are rapid, the synaptogenesis of

adult-born neurons occurs over a longer time scale. It is thus not known how the bulbar

network functions when rapid and persistent changes in environmental conditions occur.

Here, we reveal a new form of structural remodeling where mature spines of adult-born but

not early-born neurons relocate in an activity-dependent manner. This relocation is

preceded by the growth of spine head filopodia (SHF) and is driven by the activity of

principal cells. Principal cell-derived glutamate and BDNF regulate SHF motility and

directional spine relocation, respectively; and spines with SHF are selectively preserved

following sensory deprivation. Our 3D model showed that spine relocation allows fast

reorganization of OB network with functional consequences for odor information

processing.

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5.3. Introduction

The olfactory bulb (OB) is one of the few regions in the adult brain that displays a high

level of structural plasticity due to a constant supply of adult-born periglomerular and

granule cells (GC)1,2. GC form dendrodendritic reciprocal synapses with the lateral

dendrites of principal cells (mitral cells; MC). The inhibition provided by these synapses

synchronizes the activity of MC, allowing for fine spatio-temporal tuning of their responses

to odors3. Adult-born GC play an important role in this process4, and several studies have

shown that they are involved in short-4 and long-term odor memory5, odor discrimination6,

and social behavior7. The central role played by adult-born neurons in different odor-

dependent tasks is likely due to their increased responsiveness to odor stimulation8 as well

as to transient experience-dependent synaptic modifications at proximal glutamatergic

fiber-GC synapses9.

The continuous supply of new neurons provides the OB with a reservoir of plastic cells that

enables synaptic remodeling due to the continuous formation, elimination, and/or

stabilization of new spines of immature10-14 and mature adult-born neurons15-17. Adult-born

GCs thus constantly sculpt the bulbar network in response to changing environmental

conditions. However, environmental changes can be rapid, whereas the synaptogenesis of

adult-born neurons occurs over a longer time scale. It is thus conceivable that the OB

network requires considerably quicker structural modifications to existing dendrodendritic

synapses in order to adapt the functioning of the OB network to changing environmental

conditions.

In the present study, we used in vivo and in vitro two-photon time-lapse imaging, with a

relatively rapid acquisition rate (once every 5 min), to show that synaptic remodeling in the

OB network also relies on the relocation of the spines of mature adult-born neurons.

Chronic in vivo imaging revealed that relocated spines are stabilized in the bulbar network

and are directly opposed to synaptophysin-labeled puncta. We linked spine relocation to the

growth of thin filopodia-like protrusions from the spine head (spine head filopodia; SHF)

that depended on the level of odor-induced activity. Stimulation of a single MC induced

directional growth of SHF toward activated MC dendrites that in turn promoted spine

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relocation. Glutamate released from MC induced the AMPA receptor- (AMPAR)

dependent motility of SHF, whereas the activity-dependent release of BDNF by MC drove

directional growth of SHF and spine displacement. On the other hand, there were fewer

SHF on the spines of early-born GC, and they were unaffected by odor stimulation. Our

findings suggested that the spines of adult-born neurons can relocate from inactive to active

principal cell dendrites, highlighting a new form of structural plasticity in the constantly

remodeling OB network. Furthermore, our computational modeling experiments showed

that the relocation of mature spines allows rapid adjustment of the OB network to odor-

induced activity that can have functional consequences for odor information processing.

5.4. Results

5.4.1. Dendritic spines of adult-born GC relocate in the OB network

To study the structural remodeling of adult-born neurons over relatively short time

intervals, we performed in vivo two-photon time-lapse imaging in the adult OB. Neuronal

precursors were labeled by the stereotaxic injection of a GFP-encoding lentivirus into the

adult RMS (Fig. 1a) and time-lapse imaging of adult-born GC in the OB was performed

every 5 min for 60-240 min (Fig. 1b,c). Our experiments revealed that adult-born GCs

undergo a novel form of structural remodeling via the relocation of some spines on the

apical part of the dendrites (Fig. 1c,d). In some cases, the spines relocated by at least 2-4

μm (Fig. 1d,h; Supplementary Movie 1). The relocation of the spines was observed at 30-

50 and 120-150 days post-injection (dpi) of the viral vector into the RMS, indicating that

the relocating spines were located on fully integrated and mature adult-born GC. A

comparison of relocating and non-relocating spines showed that the relocation was not

associated with changes in spine volume (Supplementary Fig. 1d). In addition, spine

relocation was observed in acute OB sections at 14, 28, 42, and more than 77 dpi (163 dpi

being the longest time-point studied) (Fig. 1e-h; Supplementary Fig. 1a; Supplementary

Movie 2), confirming our results in more stable imaging conditions than in vivo. To

quantify the percentage of relocating spines and the extent of their relocation, we tracked

514 randomly selected spines at different dpi and identified relocating spines based on the

z-score of their direction vector amplitude, which is a measure of the directionality of spine

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relocation (Supplementary Fig. 1b,c). Under baseline conditions, 4.5% of the spines

relocated above the threshold (Fig. 1i). This percentage was similar at different dpi (range

3-8% for 14, 28, 42, and 77 dpi in vitro and 30-60 dpi in vivo). The mean ratios of spine

relocation and of the amplitudes of the direction vectors were 1.2±0.2 and 1.26±0.09 μm,

respectively (n= 21 spines from 18 cells, 16 mice) for spines that significantly relocated

(above z-score) (Fig. 1g,h). These findings suggested that the structural plasticity in the OB

network depends not only on the continuous arrival of new neurons but also on the

relocation of mature spines of adult-born GC.

5.4.2. Spine relocation is preceded by spine head filopodia growth

Our in vivo and in vitro two-photon time-lapse imaging revealed that spine relocation is

often preceded by the growth of thin filopodia-like protrusion from the spine head,

hereafter called spine head filopodia (SHF; Fig. 1f). The SHF were on average 2.1 ± 0.1

µm in length (n = 97 SHF, 3 cells, 3 mice). Based on 60 min of baseline recordings,

approximately 45% of the spines both in vivo and in vitro displayed SHF, several of which

arose from the same spine and grew in different directions (Fig. 1f and Fig. 2a,b). SHF

were also observed on adult-born GC spines in fixed tissues (Fig. 2c). The percentage of

spines with SHF was similar in fixed tissues and in snapshot images of time-lapse movies

(7.7 ± 0.9% of the spines with SHF in fixed tissues (n = 28 cells,3 mice) and 7.6 ± 1.2% in

acute OB slices (n = 16 cells, 10 mice). The total distance of spine relocation as well as the

directional vector were higher for the spines with SHF compared to the spines devoid of

SHF (Fig. 2d,e), and 95.3% of the spines relocating above the threshold had SHF (20 of 21

spines, n = 16 mice; red dots on Fig. 2d). These results indicated that SHF are required for

spine relocation. To verify whether SHF are sufficient for spine relocation, we examined

the relocation of spines with and without SHF. Of the 344 spines with at least one SHF (n

=57 cells, 42 mice), 6.1% relocated above threshold (range 4.5-13.5% for 14, 28, 42, and

77 dpi in vitro and 30-60 dpi in vivo), whereas only one of 160 spines devoid of SHF

(0.6%) relocated above the threshold (n = 57 cells, 42 mice). To determine whether the

direction of SHF growth determined the direction of spine relocation, we calculated the

average vector of SHF growth (Fig. 2f,g; black arrow) for relocating spines (red dots on

Fig. 2d). Our analysis suggested that spines relocate toward the position where SHF had

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grown (Fig. 2g,h; compare red and black solid arrows). These results indicated that the

presence of SHF is required but not sufficient for spine relocation and that the direction of

SHF growth determines the direction of spine relocation.

We then looked at whether spine relocation occurs on the GC dendritic segments showing

increased structural dynamics. We examined the dynamics of SHF on ~100-µm-long

dendritic segments centered around relocating (above a z-score) and stable (without SHF)

spines. Our analyses revealed higher structural dynamics on dendritic segments close to

relocating spines (< 15 µm) than those at remote locations (> 15 µm) of the same dendrite

as well as on dendrites with non-relocating spines (Supplementary Fig. 2).

Since the relocation of spines on adult-born GC was maintained at all maturational stages

studied, we then determined the dynamics and lifetimes of SHF at 14, 28, 42, and more

than 77 dpi. We observed no differences in SHF lifetimes and the number of appearing and

retracting SHF (Fig. 2i and Supplementary Fig. 2), suggesting that SHF on adult-born

spines appear at 14 dpi and remain beyond 77 dpi. To show that adult-born GC spines with

SHF are mature, we determined whether markers of mature spines, including PSD95 and

synaptoporin, the latter being expressed by GC but not MC 14,18. Our findings showed that

these spines did indeed express PSD95 (93.2 ± 2.3%) and synaptoporin (85.2 ± 7.4%) (n =

26 cells, 3 mice). SHF were devoid of these markers, which is consistent with their rapid

formation and retraction. To provide further evidence that spines with SHF are fully

functional, we performed Ca2+ imaging of spines with and without SHF following the

induction of back-propagating action potential. We first filled td-tomato virus-labeled

adult-born GC with Oregon Green BAPTA, a Ca2+ indicator, using a patch pipette (Fig. 2j).

We then performed line scan imaging of spines with and without SHF following back-

propagating action potentials induced by GC depolarization via a patch pipette (Fig. 2k).

Spines with SHF produced robust transient Ca2+ signals. These transients were slightly

higher, albeit not significantly, than those produced by spines without SHF, indicating that

spines with SHF are fully functional (Fig. 2l-m).

We then determined whether spine relocation is specific to adult-born GC or whether it can

also be observed in early-born interneurons. We thus labeled early-born GC by stereotaxic

injection of a GFP-encoding lentivirus into the RMS of P10 pups and performed in vivo

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two-photon imaging in the OB 45-60 days later. No spine relocation of early-born GC was

observed. Moreover, early-born GC had a lower percentage of spines with SHF, a lower

SHF dynamics, and a tendency for higher lifetime than their adult-born counterparts (Fig.

2n-p). Altogether, these results suggested that the SHF growth vector determines the

position of spine relocation and that this new form of structural plasticity of mature, fully

functional spines in the OB network is mediated by adult-born but not by early-born GC.

5.4.3. Relocated spines are maintained in the OB network

To determine whether relocating spines are maintained in the OB network, we performed

chronic in vivo two-photon imaging. We first imaged spines of adult-born 30-60 dpi GC to

identify relocating spines and retrieved them 24-48 h later during a second imaging session

(Fig. 3a,b). The mean direction vector amplitude of relocated spines during the first

imaging session was 0.96 ± 0.2 µm (n = 15 spines from 11 mice). Interestingly, 93.3% of

these spines were found on the second imaging day, indicating that relocating spines are

stabilized and maintained in the OB network (Fig. 3c,d). In comparison, 90.7% of non-

relocating spines were found on the second imaging day (n=43 spines from 11 mice). Spine

relocation was observed on the apical dendrites of adult-born GC in the external plexiform

layer where they receive exclusive input from bulbar principal neurons19. We then

investigated whether the relocated spines were part of dendrodendritic synapses with MC

dendrites. To address this issue, we identified relocated spines by in vivo two-photon

imaging and then performed immunolabeling in the fixed tissue to detect synaptophysin,

which is expressed by both MC and GC spines18. Our analysis revealed that relocating

spines express synaptophysin and are directly opposed to synaptophysin-positive puncta

(Fig. 3e). This result was consistent with the dendrodendritic nature of GC apical dendrite

synapses and suggested that relocated spines are part of synaptic contacts with MC.

5.4.4. MC activity induces SHF directional growth and spine relocation

Since several SHF may emerge from the same spine and grow in different directions under

baseline conditions, we hypothesized that SHF on adult-born GC spines may be important

in sensing the OB microenvironment and that their dynamics might depend on odor

stimulation. To test this hypothesis, we performed in vivo two-photon imaging of adult-

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born and early-born GC during odor stimulation (Fig. 4a,b). We presented, via an

olfactometer, a mixture of butyraldehyde and methylbenzoate, which is known to activate

the medio-lateral region of the dorsal surface of the OB20. Our experiments revealed that

odor stimulation stabilizes the SHF of adult-born GC at both 30-50 and 120-150 dpi (Fig.

4c). Interestingly, however, the odor stimulation had no effect on the SHF dynamics of

early-born GC (Fig. 4c), which again implies that adult-born GC make a specific

contribution to the structural remodeling of the bulbar network over a relatively rapid time

frame.

We then investigated whether SHF are preferentially guided toward activated MC dendrites

that in turn induce spine relocation. To directly address this question, we selectively

stimulated single MC in acute OB slices, avoiding the effect of broad activation of the

bulbar network induced by odor stimulation. We filled the MC with Alexa594, and imaged

SHF dynamics on GFP-labeled adult-born GC spines positioned close to (5 μm) and far

away (> 10 μm) from the Alexa594-filled MC dendrite (Fig. 4d,e). The MC was stimulated

with two different patterns, one that mimicked the activity of these cells induced by odor

presentation (hereafter called physiological pattern stimulation; Fig. 4f,g)21 and one that

consisted of the same number of spikes given in a random order (hereafter called random

stimulation, Supplementary Fig. 3). The images under control conditions were taken over

a 30-min period, followed by 30 min of imaging during the stimulation of the MC (Fig.

4g). Consistent with our in vivo results following odor stimulation, the SHF lifetimes of

spines located close to the stimulated MC dendrite were significantly longer than those

located far away after physiological pattern MC stimulation (Stim) compared to the

baseline condition (BL) (Fig. 4h,i). Interestingly, the tracking of each SHF with respect to

the MC dendrite location and the calculation of the cosines of the angle between them

showed that SHF growth became highly directional toward the stimulated MC dendrite

(Fig. 4j-l). We then determined whether the directional growth of SHF toward stimulated

MC dendrites is followed by spine relocation. Our analysis revealed a significant increase

in the ratio of spine displacement (Fig. 4m), and the amplitude of the direction vector (Fig.

4n,p) after physiological MC stimulation. Moreover, stimulating the MC with a

physiological pattern increased the percentage of spines with SHF that relocated above the

threshold from 3.7 % to 25.9 % (30 min imaging under baseline and stim. conditions, n =

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27 spines, 11 MC-GC pairs, 9 mice). In contrast, the stimulated MC dendrite was stable and

did not show any displacement (0.08 ± 0.01 µm, n = 7 MC dendrites, Supplementary Fig.

3). In addition, no differences were observed in the lifetime, directional growth, and spine

relocation of the SHF when the MC was stimulated with a random pattern (Fig. 4o;

Supplementary Fig. 3). These results indicated that spine relocation does not occur

passively over time and has to be induced by a physiological pattern of stimulation and not

the overall activity of the MC.

5.4.5. Glutamate released from MC controls the motility of SHF

Since MC are glutamatergic neurons22, adult-born GC express both AMPARs and

NMDARs23-25 and since stimulating MC affects the initial stages of GC integration10, we

next investigated whether SHF dynamics and spine relocation depend on MC-derived

glutamate and the activation of AMPARs and/or NMDARs on adult-born GC. To address

this issue, we extracellularly stimulated the lateral olfactory tract (LOT) where all the axons

of MC converge and applied a pattern that mimicked MC responses to odors (Fig. 5a). We

used this approach, rather than single MC stimulation, since it allowed us to acquire

prolonged time-lapse images of SHF under baseline (BL) conditions for 45 min followed

by LOT stimulation for 45 min and then to apply NMDAR (APV, 50 µM) or AMPAR

(NBQX, 25 µM) antagonists while stimulating the MC for an additional 45 min (Fig. 5c).

To ascertain that the imaged adult-born cell was located in the region activated by the

electrical stimulation, we also recorded local field potentials in that area. Our results

showed that MC stimulation stabilizes SHF by increasing their lifetime and reducing their

motility (Fig. 5b and Supplementary Fig. 4a-c). These effects depended on the activation

of AMPARs, but not NMDARs, since the application of NBQX, but not APV, completely

blocked the changes in SHF dynamics and lifetime induced by MC stimulation (Fig. 5b;

Supplementary Fig. 4d). No changes were observed under control conditions when we

imaged spines of adult-born GC for the same period of time without MC stimulation

(Supplementary Fig. 4b, left panel).

While these experiments revealed that AMPARs play a role in the motility of SHF, the

extracellular stimulation of thousands of MC cannot be used to assess the directionality of

SHF growth or spine relocation. To this end, we applied AMPA locally by iontophoresis

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and monitored SHF dynamics and spine relocation. We first recorded baseline dynamics for

45 min and then positioned a pipette containing 10 mM AMPA and 10 µM Alexa594 (to

visualize the position of the pipette) approximately 5 µm from the spine of an adult-born

GC. A brief pulse of negative current ranging from 150 to 250 nA via the pipette was

applied to release the AMPA (Fig. 5d). AMPA iontophoresis increased the lifetime of SHF

(Fig. 5e,f) and decreased the dynamics (Supplementary Fig. 4e), which is in agreement

with the LOT stimulation (Fig. 5a-c) and odor administration (Fig. 4a-c) results.

Interestingly, however, an analysis of the cosines of the angles between the SHF and the

position of the iontophoresis pipette did not reveal any directionality effect by AMPA (Fig.

5g). In addition, we observed no spine displacement following AMPA iontophoresis (Fig.

5h).

5.4.6. MC-derived BDNF induces the spine relocation of adult-born GC

While our results indicated that glutamate controls the motility of SHF via AMPAR

activation, the directional growth of SHF and spine relocation should be regulated by other

factors released in an activity-dependent manner. It is well established that target cell-

derived trophic factors may be required for activity-dependent reorganization of synaptic

contacts26,27 and that MC express BDNF, which affects the spine density of adult-born GC

in a TrkB-dependent manner28. We thus examined the involvement of MC-derived BDNF

in the directional growth of SHF and in spine displacement. We first performed in situ

BDNF hybridization and observed high levels of BDNF mRNA in the MC and glomerular

(GL) layers, but not in the GC layer (Fig. 6a), as previously reported28. BDNF fluorescent

in situ hybridization combined with MC marker PGP9.5 immunolabeling confirmed that

MC expressed BDNF in the OB (Fig. 6b). To address the role of BDNF signaling in SHF

dynamics and spine relocation, we pressure-applied BDNF (10 ng/ml) and monitored SHF

dynamics on the spines of adult-born GC. Interestingly, puff application of BDNF did not

affect the lifetimes of SHF (Fig. 6d,e) but induced directional growth of SHF toward the

BDNF-containing pipette (Fig. 6d,f). The directional growth of SHF following the puff

application of BDNF was accompanied by the relocation of adult-born GC spines (Fig.

6g,h). Moreover, while we did not observed any relocation of spines with SHF under 45

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min of baseline imaging, BDNF puff induced relocation of 38.1 % of spines with SHF (n =

21 spines, 12 cells from 4 mice).

We then produced a BDNF-knockout subpopulation of MC in the adult OB by injecting a

Cre-mCherry viral construct above the MC layer of adult BDNFfl/fl mice (Fig. 7a). To

ensure the efficacy of the Cre-dependent recombination, we first injected the Cre-mCherry

viral construct into GFP reporter mice. We observed multiple GFP+ MC as determined by

co-labeling of mCherry with PGP9.5 (Fig. 7b,c). We then patched the BDNF-lacking Cre-

mCherry+ MC in BDNFfl/fl mice, filled it with Alexa594, and monitored the SHF

dynamics and spine relocation of GFP+ adult-born GC following a physiological pattern of

MC stimulation (Fig. 7d-f). The stimulation of the BDNF-lacking MC still increased the

lifetime of SHF (Fig. 7g), which is consistent with a role for MC-derived glutamate in the

motility of SHF (Fig. 5). However, no directional growth of SHF (Fig. 7h) and, as such, no

spine displacement of adult-born GC (Fig. 7f,i) toward the simulated BDNF-lacking MC

was observed. These results suggested that the activity-dependent release of BDNF from

MC is required for the directional growth of SHF and the spine relocation of adult-born

GC.

5.4.7. Spines with SHF are maintained after sensory deprivation

Our results showed that SHF are required for spine relocation and are guided toward MC

dendrites by BDNF released in an activity-dependent manner. This structural plasticity may

be instrumental in the maintenance of some adult-born GC spines in the constantly

remodeling OB network. We thus investigated the SHF dynamics and spine maintenance of

adult-born GC in the sensory-deprived OB of BDNFfl/fl mice injected with the Cre-

mCherry viral construct (Fig. 8a). We first verified the efficiency of the sensory

deprivation by confirming the decrease in TH expression in the GL layer 14 days following

nostril occlusion (Fig. 8b)29,30. We then assessed the spine density of adult-born GC in

regions containing a high density of Cre-mCherry+ versus Cre-mCherry- MC and

compared it to the spine density in sensory-deprived ipsilateral versus control contralateral

OBs (Fig. 8c). Sensory deprivation decreased the overall spine density of adult-born GC

regardless of the expression of BDNF in MC (Fig. 8d), which is in agreement with

previous reports31,32. However, an increased percentage of GC spines with SHF in the

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regions of Cre- MC was observed after sensory deprivation (Fig. 8e). This effect was seen

in vivo as well as in vitro in acute OB slices derived from sensory-deprived mice (Fig. 8g).

These results suggested that sensory deprivation does not affect spines with SHF and leads

to the elimination of spines without SHF. Specific maintenance of spines with SHF in the

sensory-deprived OB depended on the expression of BDNF by MC since the knockout of

BDNF expression by the Cre-mCherry viral construct completely abolished the increase in

the percentage of spines with SHF (Fig. 8e, Cre-mCherry+ and Cre-mCherry- regions in

the ipsilateral OB). We next determined whether the presence of spines with SHF in the

sensory-deprived OB is due to the increased motility of SHF. We acquired time-lapse two-

photon images of adult-born GC dendrites from control and odor-deprived OB to measure

the lifetime of SHF. We observed a decrease in the lifetime of SHF and an increase in their

dynamics (Fig. 8f). These effects were, however, independent of BDNF expression (Fig.

8f), which is consistent with our observations that MC-derived BDNF affects SHF

directional growth and spine relocation but not SHF dynamics (Figs. 6-7). These results

suggested that a reduction in odor-induced activity triggers SHF dynamics in a BDNF-

independent manner followed by BDNF-dependent maintenance of spines with SHF.

5.4.8. Spine relocation is involved in odor information processing

Previous studies have suggested that inhibition promotes MC synchronization33 and that a

significant part of this inhibition is conveyed by the GC organized in sparse, segregated,

and distributed columns34. Given these observations, we investigated how and to what

extent mature spine relocation affects the synchronization between MC belonging to

different glomeruli and how this structural plasticity impacts odor information processing.

To address this issue, we used a 3D OB network model35 that represents the natural

arrangement of MC and GC and provides a realistic representation of their overlapping

dendrites. We started from a network configuration representing three neighboring

glomerular (GL) units after a learning session that resulted in well-formed columns of

potentiated GC synapses below each GL (Fig. 9a). At the end of the learning period there

were 3269 GC synapses that were fully potentiated (>95% of the peak conductance value).

Based on our in vivo experimental data showing that 3-8% of GC spines relocate (Fig. 1),

we ran a simulation in which 3% or 6% of the total GC spines that initially connected GL37

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with GL86 were switched to connect GL37 with GL123. In general, the number of

synapses connecting any two GL units is intrinsically determined by the spatial distribution

of the MC and GC dendrites. The synchronization between two GL units was measured as

the cross-correlation between the post-stimulus time histograms (PSTHs) of GL37 and of

either GL86 or GL123 obtained from 14 sniffs (Fig. 9b, raster plots) using a 20-ms time

bin. While a visual inspection of both the raster plots and PSTHs did not reveal any clear

difference, the cross-correlation between the involved GL units gradually changed with the

proportion of relocated spines from GL86 to GL123 (Fig. 9c). These results suggested that

the relocation of relatively few spines in response to a new sensory input can be an

effective mechanism for quickly changing the set of synchronized MC, which in turn

affects odor information processing.

5.5. Discussion

We discovered a new form of structural plasticity in the OB that relies on the relocation of

the dendritic spines of mature adult-born but not early-born, GC. This relocation is driven

by MC activity and is preceded by the growth of SHF from the spine heads. Principal cell-

derived glutamate controls the motility of SHF via the activation of AMPARs, whereas

activity-dependent BDNF release from MC dendrites drives the directional growth of SHF

and spine displacement. Spines with SHF were maintained in sensory-deprived OB, and

this maintenance depended upon expression of BDNF by MC. Our modeling studies also

showed that spine relocation contributes to odor information processing and allows for the

fast synchronization of MC in olfactory learning paradigms.

The filopodia-like structures emerging from spines have been observed in primary36,37 and

organotypic38,39 hippocampal cultures, and in acute slices40, as well as in vivo in the visual

cortex during the critical period41. The length of these structures ranges from short 167-

880-nm spinules37,42 to longer structures called spine head protrusions38,39,41. It has been

previously suggested that spinules and spine head protrusions may be distinct structures

based on differences in microtubule transendocytosis36. The filopodia-like structures

observed on GC in the OB are long (> 2 µm) and resemble, in terms of their length,

lifetime, and dynamics, the spine head protrusions observed in hippocampal organotypic

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cultures38,39 and, in vivo, the mouse visual cortex during the period of synaptic plasticity41.

While both spinules and spine head protrusions are associated with synaptic

plasticity36,37,42, their function is unknown. The results of the present study revealed that

SHF are required for a new form of structural plasticity where mature functional spines

relocate in an activity-dependent manner. We propose that these SHF may also be required

for the structural remodeling of spines in other sensory systems during periods of massive

synaptic competition and activity-dependent refinement of neuronal networks.

The release of glutamate and BDNF from MC dendrites has two distinct functions in order

to ensure spine stabilization and spine motility. The release of glutamate from MC may

activate the AMPA receptors on adult-born GC spines, reinforcing the structures that are

synaptically active and stopping the formation of SHF in the OB network. It has been

suggested that NMDA receptors trigger the formation of new spines43,44, whereas AMPA

receptors play a role in synapse maintenance by stabilizing actin-based motility45. We

previously showed that NMDA receptor activity is important for the initial step of spine

formation on adult-born GC10. We now propose that AMPA receptors play a role in the

stabilization of the spines of adult-born mature interneurons. On the other hand, local

application of BDNF revealed a trophic mechanism for inducing SHF directional growth

and spine relocation. We hypothesize that, when a period of lower synaptic activity occurs,

SHF protrude and grow toward trophic factors such as BDNF released from active MC.

This in turn promotes spine relocation and the synaptic maintenance of adult-born

interneurons. The role of BDNF in synaptic maintenance and competition has been studied

in visual cortex where mosaic depletion of BDNF alters spine density46. Furthermore, the

deletion of TrkB, a high-affinity BDNF receptor, results in fewer spines on adult-born

neurons in OB28. We propose that, in the adult OB, the concurrent actions of MC-derived

glutamate and BDNF regulate the structural modifications of mature adult-born GC.

Several lines of evidence suggested that SHF guide the relocation of spines from inactive

toward active MC dendrites. First, spine relocation was directional and occurred over the

space of several micrometers. The MC dendrites were very stable and did not show any

displacement, suggesting that GC spines relocate to another stable MC dendrite. Second,

the relocating spines were stable and were found at the same position 24-48 h later. Third,

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the relocated spines expressed pre- and postsynaptic markers and were directly opposed to

presynaptic structures, indicating that they are part of dendrodendritic synapses. These

results suggested that relocating spines rapidly form new functional units with their target

cells in an activity-dependent manner.

GC-to-MC synapses are involved in the generation of fast evoked oscillations by

synchronizing the activity of MC in the OB3. Adult-born GC play an important role in this

process by providing approximately 45% of the inhibition received by MC4,32. These adult-

born GC are highly sensitive to sensory activity8,47, and odor-induced activity regulates the

integration of these neurons into the OB network31,32. These reports suggest that the

continuous formation, stabilization, and/or elimination of new synapses by adult-born GC

regulate the functioning of the OB network, which in turn affects some, but not all,

olfactory behavioral tasks48. The synaptogenesis of adult-born neurons occurs over time

scales that are longer than the rapid and persistent changes in environmental conditions, and

the OB would require much faster structural plasticity to react to environmental changes. In

our experiments, we did not observe the formation of new spines during 4-h imaging

periods of adult-born GC dendrites. This contrasts with spine relocation, which can occur

within a few minutes. In this respect, the relocation of mature spines of adult-born GC from

inactive toward active MC dendrites may allow a much faster adaptation of the OB network

to rapidly changing environmental conditions. Indeed, our computational modeling

experiments revealed that the relocation of a few spines may be sufficient to synchronize a

new subset of principal cells during new sensory input processing. Our study revealed a

new form of activity-dependent structural plasticity that allows for the rapid adaptation of

the OB network to new sensory inputs.

5.6. Methods

5.6.1. Animals

Adult (>2-month-old) male C57BL/6 mice (Charles River), postnatal day 10 (P10) C57Bl/6

and CD1 mice (Charles River) adult male and female BDNF fl/fl mice (Jackson

Laboratories), and CAG-CAT-EGFP reporter mice (kindly provided by Dr. Kenneth Campbell,

Cincinnati Children's Hospital Medical Center) were used. The experiments were performed in

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accordance with Canadian Guide for the Care and Use of Laboratory Animals

guidelines. All the animal procedures were approved by the Université Laval animal

protection committee. The investigator was not blinded to the experimental group. One to

four mice per cage were kept on a 12-h light/dark cycle at a constant temperature (22°C)

with food and water ad libitum.

5.6.2. Stereotaxic injection

A GFP-encoding lentivirus (100-300 nl; 1x109iu/ml, UNC Vector Core) was injected into

the RMS of both brain hemispheres of the mice to label adult-born GC. The following

coordinates were used for 20-25 g mice (from bregma): anterior-posterior: 2.55 mm;

medial-lateral: ± 0.82 mm; and dorsal-ventral: 3.15 mm. After injection, the mice were

returned to their cages and were kept for different periods of time (14, 28, 42, and > 77

days post-injection (dpi)). The > 77 dpi group included animals kept for up to 163 dpi. We

detected few GFP+ cells in the RMS of the OB, indicating that the viral construct did not

infect stem cells10. With our injection method, we thus obtained an accurate assessment of

the age of the adult-born neurons. In some experiments we infected neuronal progenitors in

the RMS with td-tomato-encoding lentivirus particles (100-300 nl; 1.5x1010iu/ml, UNC

Vector Core) to perform Ca2+ imaging of adult-born GCs spines with Oregon Green

BAPTA. To label early-born neurons, we injected a GFP-encoding lentivirus into the RMS

of P10 mice at the following coordinates (from bregma): anterior-posterior: 2.05 mm;

medial-lateral: ± 0.65 mm; and dorsal-ventral: 2.7 mm. To selectively knockout BDNF

expression in the OB, an AAV Cre-mCherry (2x1012 iu/ml, UNC Vector Core) viral

construct was injected into the OB of BDNF fl/fl mice. The following coordinates were

used: anterior-posterior: 5.00 mm; medial-lateral: ± 1.50 mm; and dorsal-ventral: 1.38 mm.

5.6.3. Time-lapse two-photon imaging in vivo

Adult mice injected with a GFP-encoding lentivirus in the RMS either at P10 or P60 were

used for in vivo two-photon imaging at 45-60 dpi (for P10 mice) or 30-50 and 120-150 dpi

(for P60 mice). The mice were anesthetized with 2-3% isoflurane during cranial window

implantation. The temperature was maintained at 37.5˚C during the entire procedure using

an infrared blanket (Kent Scientific). After removing the scalp, we drilled a circular

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craniotomy centered over the OB hemispheres. We removed the bones protecting the OB,

leaving the dura intact. Bleeding was controlled with gel foam pieces. A 3-mm-diameter

coverslip was centered on top of the OB, and Kwik-seal (World Precision Instruments) and

dental cement was used to maintain the coverslip on the surface of the OB. A small head

plate glued to the skull was used to keep the head from moving during the imaging

procedure. Once the surgery was completed, the amount of isoflurane was gradually

decreased, and the mouse was injected with ketamine-xylazine to maintain anesthesia

during the imaging period.

GC structural dynamics were imaged using a custom-made video-rate two-photon

microscope49. Thirty to 60 min after surgery, we positioned the mouse using the head plate

on a custom-made stereotaxic frame controlled by a micromanipulator (MPC 200; Sutter).

A 20x water immersion objective (XLUMPlanFl 20X/0.95; Olympus) was used to locate

the region of interest for dendritic imaging. For some isolated GFP+ GC, we used a simple

neurite tracer plugin to reconstruct the morphology of the neuron (ImageJ). Structural

dynamics were imaged using a 60x water immersion objective (XLUMPlanFl/IR 60X/0.90;

Olympus). 70 to 100-µm image stacks (2 µm steps) were acquired in the EPL at a rate of

one image stack every 5 min. Before each session, GC were identified by locating their

soma, which were 200-500 µm down from the brain surface. For chronic experiments, GC

spines were first imaged on day 1 for 2-4 h and were re-imaged 24-48 h later.

To study the effects of sensory activity on the structural dynamics of GC, we delivered

odors with an olfactometer (Knosis) at the beginning of each stack acquisition (30 s of odor

delivery every 5 min). We used a mixture of butyraldehyde and methylbenzoate at a

concentration of 10-3 that has been shown to activate the mediolateral region of the dorsal

surface of the OB20. The effect of the sensory activation was compared to the results of a 60

min baseline period in the absence of odor.

5.6.4. Time-lapse two-photon imaging in vitro

We acquired time-lapse two-photon images of adult-born GC as previously described10.

Briefly, deeply anesthetized mice were transcardially perfused with ice-cold oxygenated

artificial cerebrospinal fluid (aCSF-sucrose) containing the following (in mM): 250

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sucrose, 3 KCl, 0.5 CaCl2, 3 MgCl2, 25 NaHCO3, 1.25 NaH2PO4, and 10 glucose.

Horizontal slices (250-300-µm-thick) of the OB were obtained using a vibrating blade

microtome (HM 650V; Thermo Scientific). The slices were transferred into oxygenated

aCSF maintained at 32°C that contained the following (in mM): 124 NaCl, 3 KCl, 2 CaCl2,

1.3 MgCl2, 25 NaHCO3, 1.25 NaH2PO4, and 10 glucose. For the time-lapse imaging

experiments, we placed the acute slices in a perfusion chamber equipped with a temperature

controller (Warner Instruments). We acquired images of the spines of apical dendrites of

GFP-expressing GC located at least 40-µm-deep in the slices using a custom-made two-

photon microscope (modified from an Olympus FV 300 system) equipped with a 60x

water-immersion objective optimized for infrared light imaging (LUMPlanFl/IR 60x NA:

0.90; Olympus) and a Ti:Sapphire laser (Coherent). To assess spine dynamics, time-lapse z-

stack images were acquired every 5 min using Fluoview 5.0 software (Olympus). Two

detection channels allowed us to simultaneously image the GFP and Alexa594 signals for

the MC/GC and puff/iontophoresis experiments. GFP and Alexa594 were excited at 900-

940 nm. The same imaging system was used for Ca2+ imaging in adult-born GC spines. For

these experiments, adult-born GC were labeled with td-tomato-expressing lentiviral

particles and were filled with Oregon Green BAPTA (OGB) organic Ca2+ indicator (100

µM) at 21-28 dpi via a patch electrode. The Td-tomato-expressing GC were excited with a

1040 wavelength laser (Spectra-Physics). Ten to 15 min after filling the GC with OGB, line

scan imaging was performed on spines with and without SHF following a single

depolarizing pulse (200 ms) applied using a patch electrode to induce action potential.

5.6.5. Image analysis

The time-lapse images were analyzed using custom-made MATLAB programs (version

R2010a; Mathworks). We extracted maximum projection 512 x 512 pixel images of each

time point. The slice drift between each time point was corrected using a cross-correlation-

based algorithm. The original images were cropped to facilitate the visualization and

analysis of structural dynamics. For some analyses, we also false-colored every time point

(previous stack: red; current stack: green) to detect appearing and retracting SHF as well as

spine displacement.

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Several criteria must be met to detect SHF. First, they have to clearly protrude out of the

spine head by at least 1 μm. Second, they need to be easily identifiable with a high signal-

to-noise ratio. Third, they must protrude from a spine identified as stubby or mushroom,

i.e., the spine head volume has to be considerable. Lastly, they have to be transient. SHF

that were present during the whole time-course of a recording were discarded. We used

these criteria to quantify SHF dynamics on adult-born GC by calculating the number of

SHF forming/retracting on spines for a period of 1 h per µm of dendrite. The number of

SHF was obtained by manually counting the number of SHF forming or retracting on adult-

born GC spines. The SHF lifetimes were calculated by tracking each SHF manually using

the MTrackJ plugin (developed by Dr. Erik Meijering, Biomedical Imaging Group,

University Medical Center Rotterdam, Rotterdam, The Netherlands) for ImageJ. The

resolution of the lifetime measurements was limited by the sampling rate (5 min) and the

length of the time-lapse acquisition (30-240 min, depending on the experiment). SHF

tracking also made it possible to measure the direction of SHF growth. The angle of SHF

growth was measured based on the position of either the MC lateral dendrite or the location

of the puff/iontophoresis pipette (see illustration in Fig. 4j). We averaged the direction

angle for each time point during the SHF observation period and calculated the cosine of

this angle to obtain its direction. The direction measurement ranged from +1 (an SHF

pointed directly at the target) to –1 (an SHF pointed away from the target).

We assessed spine motility and volume using a custom-made program to automatically

detect the contour of the spine head. Spine relocation was measured based on the center of

the spine contour for each time point. These values were normalized to those of the region

of the dendrite from which the spine emerged. The spine displacement values were

determined for 30 to 90 min, depending on the experiment. We also calculated the direction

vector in order to assess the directionality of spine relocation. The direction vector was

calculated by averaging the vector between the origin (t = 0 min) and each subsequent time

point. For analyzing spine head relocation, we used spines that had a good GFP signal-to-

noise ratio throughout the whole imaging session and avoided spines that were clustered

together to facilitate tracking. We also investigated the spatial distribution of spine

relocation and SHF plasticity by measuring the relationship between the number of SHF

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per spine and the distance of each spine from either relocating or non-relocating spines. The

distance was measured by drawing a straight line between the two spines.

5.6.6. Stimulation of mitral cells

To assess the role of MC activity on the motility of spines of adult-born GC, we stimulated

the LOT where all the axons of MC of the OB converge10. Briefly, the LOT was electrically

stimulated using 0.1-ms pulses at ∼100 μA using an A360 stimulus isolator (World

Precision Instruments) triggered by a Digidata 1440A data acquisition system (Molecular

Devices). The pattern of electrical stimulations (five 0.1-ms pulses at 25 Hz repeated 60

times every 500 ms) was designed to reproduce the typical response of MC to odors21. We

assessed the specificity of the specific stimulation pattern by comparing the results to a

random stimulation pattern in which the same number of spines were evoked after random

delays. We recorded the local field potential responses in the EPL using a Multiclamp 700B

amplifier (Molecular Devices). We positioned the stimulating pipette as far as possible

from the imaging region to avoid direct stimulation of the adult-born GC. To investigate the

role of NMDA and AMPA receptors in the SHF dynamics of adult-born GC, we bath-

applied 50 μM APV (NMDA receptor antagonist) or 25 μM NBQX (AMPA receptor

antagonist) during the LOT stimulation protocol.

To assess the role of a single MC stimulation on the spine dynamics of adult-born GC, we

performed whole-cell patch-clamp recordings of MC. Electrophysiological recordings and

MC stimulations were performed with a Multiclamp 700B amplifier (Molecular Device).

Patch electrodes (ranging from 2.5 to 4 MΩ) were filled with an intracellular solution

containing (in mM): 122.5 K-methylsulfate, 10 KCl, 10 HEPES, 0.2 EGTA, 2 ATP, 0.3

GTP, and 10 glucose. We added 10 µM of Alexa594 fluorescent dye (Life Technologies) to

the intracellular solution to visualize MC lateral dendrites. The MC was maintained in

current-clamp mode during the recording. We recorded time-lapse sequences of adult-born

GC spines located within 5 μm of the lateral dendrite of the MC. A baseline period of 30

min in which no current was applied to the MC was followed by a stimulation period in

which the current was applied to the MC during each acquisition. The pattern of the

injected current was as follow: 150 ms pulses of ~100 pA repeated every 500 ms 60 times.

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MC with a resting potential above –50 mV were discarded. We used spines located at least

10 µm away from the stimulated MC dendrite as controls. In the experiments investigating

the role of MC-derived BDNF in spine motility, we took recordings from MC infected with

the Cre-mCherry viral construct. The construct was injected 7 or 14 days before the

experiment. We clearly identified the Cre-mCherry+ cells by bright field and fluorescence

signals before approaching the patch pipette. Cre-mCherry- cells located far from the

injection site were used as controls.

5.6.7. Iontophoresis and puff application

We used iontophoresis to locally apply AMPA to the spines of adult-born GC distal

dendrites and to assess changes in SHF dynamics and spine displacement. After recording

baseline dynamics for 45 min, we placed a pipette filled with AMPA (10 mM diluted in

aCSF) 5 µm from the spine of interest. We applied one 2-ms pulse of negative current

(ranging from 150 to 250 nA) at the beginning of the second period of time-lapse

acquisition using an ION-100 apparatus (Dagan). To assess the trophic role of BDNF in

SHF formation on spines, we pressure applied BDNF (10 ng/ml) for 10 ms at 10 psi at

every time-lapse acquisition using a PV930 Pneumatic PicoPump (World Precision

Instruments). In both the puff and iontophoresis experiments, the position of the pipette was

visualized by adding Alexa594 (10 μM) to the solution. SHF dynamics were calculated for

a 20 × 20 μm region centered on the pipette. For the iontophoresis application experiments,

we also calculated SHF dynamics at remote positions on dendritic segments of adult-born

GC located >10 μm from the pipette. We used the tip of the pipette in the puff and

iontophoresis application experiments as the reference for measuring the direction angle of

SHF. The pressure application of ACSF was used as a control.

5.6.8. Unilateral nostril occlusion

The mice were anesthetized with an injection of ketamine-xylazine. Their reflexes were

monitored to assess the depth of anesthesia before beginning the surgery. Occlusion plugs

were fabricated using polyethylene tubing (PE50, I.D. 0.58 mm, O.D. 0.965 mm; Becton

Dickinson), with the center blocked using a tight fitting knot made from Vicryl sutures (3-

0; Johnson & Johnson). The ~5mm-long petroleum jelly-coated plugs were inserted in the

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left nostrils of the mice (n = 12). We verified that the tip of the plug was fully inserted in

the nostril to avoid removal by the mouse. The nostrils were occluded before stereotaxically

injecting the GFP lentivirus in the RMS and the AAV Cre-mCherry viral construct in the

OB. We assessed the success of the occlusion 14 days later by comparing the TH-labeling

intensities of the ipsilateral versus the contralateral bulb. Only mice showing a significant

decrease in TH-labeling intensity were used in the study.

5.6.9. Immunohistochemistry

The mice were given an overdose of pentobarbital and were perfused transcardially with

0.9% NaCl followed by 4% PFA. The OB were resected and post-fixed in 4% PFA at 4°C.

Horizontal sections (50-µm-thick) were cut using a vibratome and were incubated with the

following primary antibodies: mouse anti-TH (24 h, 1:1000, catalog number 22941;

ImmunoStar), rabbit anti-PSD 95 (48 h, 1:1000, catalog number 51-6900; Invitrogen),

rabbit anti-synaptoporin (48 h, 1:500, catalog number 102 002, Synaptic System), rabbit

anti-GFP (24 h, 1:1000, catalog number A11122; Molecular Probes), rabbit anti-PGP 9.5

(24 h, 1:1000, catalog number 31A; Ultra Clone), mouse anti-Cre(24 h, 1:1000, catalog

number MAB3120; Millipore), and anti-mCherry(24 h, 1:1000, catalog number 5411-100;

Biovision). The corresponding secondary antibodies were used. The immunohistochemistry

for PSD95 and synaptoporin was performed on 4 mice, for GFP, PGP9.5, and Cre on 4

mice, for TH on 12 mice, and for mCherry on 3 mice, respectively. Six animals used for in

vivo imaging were processed for anti-synaptophysin labeling. Anti-synaptophysin antibody

was coupled to a fluorescent antibody using Mix-n-Stain CF568 antibody labeling kits

(Biotium). Anti-synaptophysin immunohistochemistry was performed on 100-µm-thick OB

sections. The sections were incubated for 72 h at 4ºC. To find the same GC dendrites in the

OB slices, the mice used for the in vivo imaging were perfused. The post-fixed brains were

placed in 4% agar and were positioned on the set-up used for the in vivo imaging. The same

dendrites were found, and a high power Ti:Sapphire laser beam was directed at the same

depth as the imaged spines but 300-500 µm apart in the y or x-axis to burn small regions.

The OB was then cut into 100-µm-thick sections. The sections containing the burned region

as well as two adjacent sections were kept for immunohistochemistry. Fluorescent images

of fixed tissues were obtained using a confocal microscope (FV 1000; Olympus) with 100x

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(UPlanSApoN 100x/NA 1.40; Olympus) and 40x (UPlanSApoN 40x/NA 0.90; Olympus)

oil immersion objectives.

5.6.10. In situ hybridization

Antisense RNA probes were labeled using DIG RNA labeling kits (Roche Diagnostics) and

were purified on ProbeQuant G-50 columns (GE Healthcare). In situ hybridization was

performed on 50-μm-thick vibratome sections from 5 mice. The signals were revealed with

nitroblue-tetrazolium-chloride/5-bromo-4-chlor-indolyl-phosphate (Promega). The

antisense probes were obtained from plasmid-containing mouse BDNF (kindly provided by

Dr. Castren, University of Helsinki, Finland). BDNF mRNA images were obtained using a

bright-field microscope (BX51; Olympus) equipped with a 20x objective (UPlanSApo 20x;

Olympus). Fluorescent in situ hybridization was performed as previously described50.

5.6.11. OB network model

We used a recent novel 3D model of the OB35 that represents the natural 3D arrangement of

MC and GC, including their overlapping dendrites, for all the simulations. Briefly, the

model implemented the reported spatial distribution of 128 glomeruli distributed over ≈2

mm2 of the dorsal area and activated by natural odors. The model was composed of 635

MC (5 for each glomerulus) and 97,017 GC, which is consistent with the commonly

accepted estimate of 1:20 for the MC/GC ratio. We selected a subset of 15 MC projecting

to three different glomeruli (GL37, GL86, and GL123) and the relative ensemble of GC

(n=14652) connected to them through dendrodendritic synapses. Unless noted otherwise,

we will refer to the ensemble of MC projecting to a given glomerulus and the set of GC

with synaptic connections to them as a glomerular unit.

The odor input and synaptic plasticity rule were identical to those used in Migliore et al.

(2015)35. Briefly, all the synaptic weights started at zero and, in response to an odor input,

each component (inhibitory or excitatory) of each dendrodendritic synapse was

independently modified according to the local spiking activity in the lateral dendrite of the

MC or in the GC synapse. The characteristics of the synaptic clusters predicted by this

model were consistent with those observed experimentally, and their formation was an

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extremely robust process. Unless noted otherwise, a learning session consisted of a 7-s odor

stimulation, which was sufficient to achieve a stable configuration of the synaptic weights

under all conditions.

All simulations were carried out in a fully integrated NEURON+Python parallel

environment (NEURON v7.3) (Hines and Carnevale, 1997) on a BlueGene/Q IBM

supercomputer (CINECA, Bologna, Italy). The model and simulation files used for the

present work are available for public download in the ModelDB section of the Senselab

database suite (http://senselab.med.yale.edu, acc.n. 186771).

5.6.12. Statistical analysis

The data are expressed as means ± SEM for bar graphs, individual values for scatter plots,

and individual values and means for population comparisons. The normality of the samples

was assessed using the Lilliefors test. Statistical significance was tested using a paired or

unpaired two-sided Student's t-test, depending on the experiment. Equality of variance for

unpaired t-test was verified using F-test. One-way ANOVA and Tukey post hoc tests were

used to compare the groups at different times after injection. The levels of significance

were set as follows: *: p < 0.05, **: p < 0.01 and ***: p < 0.001. The investigator was not

blinded to the experimental condition.

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5.8. Figures

Figure 1

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Figure 1. Relocation of mature spines of adult-born GC in the OB

a, Protocols used for labeling adult-born GC and for in vivo two-photon imaging. Neuronal

precursor cells were infected with GFP-expressing lentiviral particles in the RMS. Thirty to

150 days post-injection (dpi), the spines of GFP+ cells were imaged in vivo by two-photon

microscopy through a cranial window. b, Low magnification projection image of a

population of adult-born GC and periglomerular cells taken in vivo. c, Top – Projection

image of an isolated adult-born GC. Bottom – Lateral view (left) and top view (right) of the

reconstructed neuron shown above. d, Example from two adult-born GC dendrites imaged

in vivo at 60-70 dpi showing spine head relocation. The white arrows indicate spine head

filopodia (SHF). The first and last z-stacks have been false-colored in red and green to

highlight the extent of spine relocation. e, Example of an adult-born GC imaged in an acute

slice. f, Sequences of time-lapse two-photon images showing spine relocation on 14 dpi

(upper panels) and 42 dpi (lower panels) adult-born GC distal dendrites. g,h, Ratio of total

spine displacement and direction vector amplitude that shows the directionality of spine

relocation over 95 min for spines monitored at different maturational stages (n = 68, 124,

93, 103, and 126 spines from 7, 7, 7, 5, and 16 mice for the 14, 28, 42, 77 dpi, and in vivo

conditions, respectively). i, Histogram showing the direction vector amplitudes for 514

randomly selected spines. Spines with direction vector amplitudes above a z-score value are

shown in red. Scale bars for b, c, e are 100, 20, and 50 µm, respectively. The scale bar for

d, f is 2 µm.

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Figure 2

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Figure 2. SHF determine the relocation of the spines of adult-born but not early-born GC

a, Low magnification image of an adult-born GC used for in vivo two-photon imaging. b,

Image sequence of the boxed region in a. The arrow indicates the formation and retraction

of an SHF. c, GFP+ adult-born GC from fixed OB sections showing the presence of SHF.

d, Relationship between the number of SHF and the amplitude of the spine relocation

direction vector (n = 505 spines). e, Normalized spine displacement and amplitude of

direction vector for spines with and without SHF (n = 161 and 344 spines, respectively).

f,g, Measurement of the correlation between the direction of SHF protrusion and the

direction of spine relocation. Each dashed black line represents individual SHF growing

from the same spine. The solid black and red arrows represent the average direction of SHF

protrusion and the direction of spine relocation, respectively. h, The difference between the

average angle of SHF protrusion and the spine displacement vector was calculated for

spines showing relocation above the z-score (n = 21 spines, highlighted in red in e; ***p =

2.88x10-5 using a one-sample t-test). i, SHF lifetime values for different maturational stages

of adult-born GC (n = 12, 14, 15, and 8 cells from 10, 10, 9, and 5 mice for the 14, 28, 42,

and 77 dpi condition, respectively). j, Example of an Oregon Green BAPTA (OGB)-filled

td-tomato expressing adult-born GC at 30 dpi. k, Line scan Ca2+ imaging of a GC spine

with SHF (lower panel) following single action potentials induced by the depolarization of

the GC (upper panel). l, m Individual traces and quantification of Ca2+ transients in spines

with (green) and without (black) SHF (n = 12 and 13 spines with and without SHF, 8 cells,

8 mice). n-p, Quantification of the percentage of spines with SHF and SHF dynamics and

lifetimes for a period of 1 h of in vivo two-photon imaging for early-born at 45-60 dpi and

adult-born GC at 30-50 and 120-150 dpi (n = 7, 16, and 10 cells from 6, 6, and 4 mice,

respectively; *: p < 0.05 using one-way ANOVA). Scale bars for a, b, c, and j is 20, 5, 2,

and 20 µm, respectively.

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Figure 3

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Figure 3. Relocated spines are stabilized in the OB network and are part of the

dendrodendritic synapses

a, Experimental procedure for chronic in vivo two-photon imaging. b, Example of GC

dendrite images acquired on two consecutive days. c, Higher magnification of the boxed

area in b. Time-lapse two-photon imaging of relocating spine during the first imaging day

(upper panel). Note that the same spine is stabilized at the relocated position and that no

displacement is observed during the second imaging day. The first and last z-stacks have

been false-colored in red and green to compare the first and last images of the time-lapse

sequence, respectively. d, Two other examples of relocating spines. e, In vivo image

showing the spine relocation of an adult-born GC (lower panels). At the end of the in vivo

imaging, the animal was perfused, and OB sections were prepared for anti-synaptophysin

immunohistochemistry. Confocal image showing the dendrite of the imaged GC (upper

middle panel) and the anti-synaptophysin labeling (upper right panel). Note that spine

relocating in vivo is directly opposed to synaptophysin puncta. Scale bars: 20 µm for b, 2

µm for c, d; 5µm for e (center panel) and 2 µ for e right panels.

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Figure 4

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Figure 4. Olfactory sensory activity stabilizes SHF and promotes spine relocation

a-b, Experimental procedure and time-lapse imaging during sensory stimulation in vivo.

Scale bar: 5 µm. c, Quantification of effects of sensory stimulation on the SHF lifetime in

vivo for early-born GC at 45-60 dpi and adult-born GC at 30-50 and 120-150 dpi (n = 7, 8

and 7 cells from 6, 6, and 4 mice, respectively). d, Example of Alexa594-filled MC (red)

and adult-born GC (green) dendrites. Scale bar: 20 µm. e, Spines of an adult-born GC

located within 5 µm of MC dendrite. Scale bar: 5 µm. f-g, Stimulation pattern used to

reproduce MC activity in response to an odor, and the experimental design. The top traces

in f show an example of an MC recorded during stimulation. h, Image sequence of the

spine highlighted in e, before (baseline) and during (Stim) the MC stimulation. Scale bar: 2

µm. i, Comparison of the effect of MC stimulation on the SHF lifetimes of spines located

within 5 μm and more than 10 μm away from the stimulated lateral dendrites. j, Illustration

of the method used to calculate the SHF direction based on the position of the MC lateral

dendrite. k, Graph showing the direction of the growth and lifetimes of the SHF. The

arrows indicate the orientation of the SHF with respect to the dendrite of the MC. The

lengths of the arrows represent the SHF lifetimes (n = 43 and 32 SHF for baseline and

stimulation conditions, respectively). l, Quantification of the cosine of the angle between

the SHF and the MC lateral dendrite for each spine analyzed before and during MC

stimulation. i-l: n = 11 MC-GC pairs, 9 mice. m-n, Quantification of the normalized spine

displacement and direction of amplitude vector at baseline and during MC stimulation (n =

27 spines,11 MC-GC pairs, 9 mice). o, Random stimulation pattern delivered to a single

MC did not induce spine relocation (n = 18 spines from 9 MC-GC pairs, 9 mice). p,

Comparison of the effect of MC stimulation on spine relocation for spines close to or far

from MC dendrite. m-n, *: p<0.05 with paired Student’s t-test.

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Figure 5

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Figure 5. Glutamate released from MC stabilizes spine head filopodia

a, Illustration of the protocol used to apply a physiological pattern of activity to MC in the

OB while imaging adult-born GC. A stimulating pipette was inserted in the lateral olfactory

tract (LOT) while an extracellular pipette recorded the local field potential in the external

plexiform layer. A distal dendrite of an adult-born GC was then imaged in the region where

MC activity was induced. b, Experimental design of the LOT stimulation experiment. After

recording the baseline spine dynamics of an adult-born GC, the dynamics under LOT

stimulation were recorded. A glutamate receptor antagonist was applied while maintaining

the LOT stimulation. c, Effect of bath application of an AMPA and an NMDA receptor

antagonist (NBQX and APV, respectively) on the stabilization of SHF observed after LOT

stimulation (n = 11 and 12 cells from 7 and 5 mice for NBQX and APV, respectively; *p

<0 .05 and **p < 0.01 using a paired t-test). d, Illustration of the protocol used for the local

application of AMPA. The angle of the SHF was calculated based on the position of the tip

of the iontophoresis pipette. e, Example of an image sequence showing the effect of AMPA

iontophoresis on SHF dynamics. Scale bar: 10 µm. f, Quantification of the lifetime of the

SHF at baseline (45 min) and following AMPA iontophoresis. g, Quantification of the

angle of the SHF with respect to the tip of iontophoresis pipette at baseline and following

the application of the AMPA. h, Effect of AMPA iontophoresis on adult-born GC spine

displacement over 45 min (n = 20 spines). f-h, Data from 7 cells from 10 mice. p values

were calculated using the paired Student’s t-test.

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Figure 6

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Figure 6. BDNF application promotes spine relocation

a, In situ hybridization of BDNF in the OB. Note that BDNF is selectively expressed in the

MC and GL layers. EPL - External plexiform layer; GCL - granule cell layer. b, Confocal

images of BDNF expression by MC. Green - immunohistochemistry of PGP9.5, a marker

for MC. Red - fluorescent in situ hybridization of BDNF mRNA. c, Two-photon image of

the distal dendrite of an adult-born GC and the location of the BDNF puff pipette. d, Time-

lapse acquisition of the spine highlighted by an arrow in c at baseline followed by the

application of BDNF (10 ng/ml) at the beginning of the second imaging session. e,

Quantification of the lifetime of the SHF following a BDNF puff. f, Quantification of the

directional growth of the SHF toward the BDNF puff. g-h, BDNF-triggered spine

relocation compared to baseline. Spine relocation was analyzed for 45 min under both

baseline and BDNF puff conditions (n = 21 spines). Normalized spine displacement (g) and

direction vector amplitude (h) are shown. e-h, n = 12 cells from 4 mice. p values are

calculated using the paired Student’s t-test. Scale bars: 200, 20, 10, and 2 µm for a, b, c and

d, respectively.

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Figure 7

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Figure 7. The activity of BDNF-lacking MC does not induce spine relocation

a, Illustration of the method used to knock out BDNF from a fraction of MC in the adult

OB. The RMS of adult-born GC were first infected with GFP-expressing lentiviral

particles. Seven days later, an AAV viral vector expressing Cre-mCherry was injected in

the MC layer of the OB of BDNF fl/fl mice. The effect of knocking out BDNF was

assessed 7 days layer (14 dpi for the adult-born GC). b, Confocal images of the GFP signal

in the MC layer of the GFP reporter line injected with the Cre-mCherry virus. c, Confocal

images of the PGP9.5 protein signal in the MC layer of wild-type mice injected with the

Cre-mCherry virus. d, Reconstruction of the lateral dendrites of an MC injected with the

Cre-mCherry viral construct and filled with Alexa594 using a patch pipette. e, Enlarged

view of the boxed area in d. f, Image sequence of a spine illustrating the effect of an

electrical stimulation of a BDNF-deficient MC. g, Effect of the activity of a Cre-mCherry+

MC on the lifetime of the SHF. These results were obtained from time-lapse images

acquired over 30 min of baseline recordings followed by a further 30 min of MC

stimulation. h, Comparison of the direction of an SHF forming in baseline conditions and

following the stimulation of a BDNF-deficient Cre-mCherry+ MC. i, Effect of the

stimulation of a BDNF-deficient Cre-mCherry+ MC on spine displacement over 30 min (n

= 20 spines). g-i, n = 7 cells from 6 mice. p values were calculated using the paired

Student’s t-test. Scale bars: 500 and 2 µm for a and f, respectively; and 50 µm for b-d.

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Figure 8

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Figure 8. Spines with SHF are selectively maintained after sensory deprivation

a, Experimental design of occlusion and simultaneous injections of GFP and Cre-mCherry

viruses. The mice were sacrificed 14 days after the surgery. b, Tyrosine hydroxylase (TH)

expression in the glomeruli layer demonstrating the efficiency of unilateral nostril

occlusion. Scale bar: 100 µm. c, Representative images of the distal dendrites of adult-born

GC in the contralateral and ipsilateral bulbs in injected and non-injected regions of BDNF

fl/fl mice. Scale bar: 5 µm. d, Quantification of spine density on adult-born GC in different

regions of the OB following sensory deprivation and BDNF knockout. e, Quantification of

the percentage of spines with SHF in fixed tissues (n = 45 cells from 3 mice in each

condition; *: p < 0.05, **: p < 0.01, ***: p <0.001 – One way ANOVA with Tukey post-

hoc test for panels d and e). f, Effect of sensory deprivation on the lifetime of SHF in the

control and sensory-deprived bulbs of wild-type mice and in the sensory-deprived bulb of

BDNF KO mice. The lifetime of the SHF was measured over a period of 60 min in time-

lapse experiments using acute OB slices. g, Percentage of spines with SHF. f-g n = 12 and

9 cells from the contralateral and ipsilateral bulbs of 10 and 6 mice, respectively, and n = 4

from 1 BDNF KO mouse. *: p<0.05; **: p<0.01 using the unpaired Student’s t-test.

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Figure 9

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Figure 9. Spine relocation promotes fast synchronization of MC with functional

consequences for odor information processing

a, 3D OB network model with glomerular units used in all simulations. Input was delivered

to form individual glomerular units. MC dendritic segments are color coded based on the

normalized peak inhibitory conductance they received from the GC. The green colored

points below the mitral cells represent the somas of GC in which at least one synapse was

strongly potentiated (more than 95% of its peak value). b, Post-stimulus time histograms

and the relative raster plot for MC from the different glomeruli under control conditions

(left plots) and after spine relocation (right plots). Test simulations were performed by

stimulating two glomeruli at a time (GL37 and GL123, red plots) or (GL37 and GL86,

black plots). Each simulation lasted for 14 s, with 1 sniff/s. c, Results of spike cross-

correlations between glomeruli as a function of the proportion of relocated spines.

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Supplementary Figure 1

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Supplementary Figure 1. Direction vector amplitudes of GC spines and dendrites

a, Location of spines shown in Fig. 1f. Scale bars: 10 µm for 1st and 3rd panels, and 5 µm

for 2nd and 4th panels. b, Illustration of the methodology for calculating spine relocation and

the average direction vector. The spine head location was calculated for each time-point for

30 to 90 min. Spine relocation was the sum of the distance between positions at each time

point normalized to that of the dendrite from which the spine emerged. The direction vector

was calculated by averaging the vector between the origin (t = 0 min) and each time point.

The direction vector gave a direct measurement of the directionality of the spine relocation.

The greater the amplitude of spine direction, the more the spine moved in a specific

direction. Scale bar: 2 µm. c, Quantification of direction vector amplitudes for GC

dendrites versus spines (n = 25, 34, 31, 28, and 38 dendritic segments and n = 68, 124, 93,

103, and 126 spines from 7, 7, 7, 5, and 16 mice for the 14, 28, 42, 77 dpi, and in vivo

conditions, respectively). d, Quantification of volume changes normalized on a local

portion of a dendrite for relocating (direction vector > 0.86 µm) and non-relocating

(direction vector < 0.4 µm) spines. n = 13 and 17 dendrites from 10 and 13 cells from 8 and

11 mice for relocating and non-relocating spines.

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156

Supplementary Figure 2

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157

Supplementary Figure 2. SHF dynamic at different maturational stages

a, Quantification of SHF dynamics for different maturational stages of adult-born GC. b-c,

SHF dynamics considered separately for appearing and retracting SHF at different

maturational stages. No significant differences between groups were observed with a one-

way ANOVA with Tukey post-hoc test (n = 16, 17, 15 and 10 cells from 10, 10, 9, and 5

mice for the 14, 28, 42, and 77 dpi conditions, respectively). d-f, Analysis of structural

dynamic on the spines located close to and far away from the relocating (d,f) and non-

relocating (e) spines. Scale bar: 10 μm.

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158

Supplementary Figure 3

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159

Supplementary Figure 3. Random stimulation pattern of MC does not induce GC spine

relocation

a-c, Comparison of physiological (left panel in a-c) and random (right panel in a-c) patterns

of stimulation delivered to a single MC. The random stimulation pattern did not cause an

increase in the lifetime of SHF (a, right panel), spine direction ratio (b, right panel), or

direction of SHF growth (c, right panel). n = 18 spines from 9 MC-GC pairs from 9 mice

and n = 27 spines from 11 MC-GC pairs from 9 mice for the random and physiological

stimulations, respectively. p values were calculated using the paired Student’s t-test. d,

Time-lapse imaging of Alexa594-filled MC dendrite showing its stability over the time.

Scale bar: 10 µm.

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160

Supplementary Figure 4

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161

Supplementary Figure 4. Activation of AMPARs is required for SHF motility.

a, Sequence of time-lapse two-photon images of a distal dendrite of an adult-born GC taken

at baseline and during lateral olfactory tract (LOT) stimulation. Arrow 1 indicates a spine

that did not display SHF dynamics after LOT stimulation. Arrow 2 indicates a SHF that

stabilized after LOT stimulation. Scale bar: 5 μm. b, Quantification of SHF dynamics of the

controls (left panel; n = 30 cells from 12 mice) and during LOT stimulation (right panel; n

= 35 cells from 18 mice). c, Effect of LOT stimulation on the lifetime of SHF (n = 33 cells

from 18 mice). d, Effect of the bath application of the AMPA and NMDA receptor

antagonists, NBQX and APV respectively, on the decrease in SHF dynamics observed

following MC stimulation. The SHF dynamics were normalized to the baseline value (n =

11 and 12 cells from 7 and 5 mice for NBQX and APV, respectively; *p < 0.05 using the

Student’s paired t-test). e, Effect of the local application of AMPA on the SHF dynamics of

spines located within 10 μm of the iontophoresis pipette (n = 7 cells from 3 mice).

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162

5.9. Authors’ contributions

A.S. and V.B.P. designed the study. V.B.P performed most of the experiments and

analyzed the data. K.B., V.B.P., and A.S. performed the in vivo imaging experiments. D.H.

examined the spine dynamics of adult-born neurons in the control and odor-deprived

olfactory bulb and the random stimulation experiments. R.R.B performed the Ca2+

imaging experiments. M.S. performed the immunohistochemistry and in situ hybridization.

D.C. provided the video-rate two-photon system for the in vivo imaging. M.M. and F.C.

performed the modeling studies. M.M compiled and drafted the modeling results. A.S.

supervised the project and obtained the funding. V.B.P. and A.S. wrote the paper.

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163

6. General conclusions

In my work, I bring together several questions related to particular processes governing the

migration of neuroblasts and their functional role in the olfactory system later in the

maturation. Using neurogenesis of adult OB as a model system to study neuronal

development, I investigated the role of KCC2 Cl- transporter in neuronal migration via

time-lapse imaging of neuroblasts with pharmacological activation of KCC2. My results

suggest that KCC2 fosters radial migration of neuroblasts in the OB and in the RMS, while

it doesn’t play role during tangential migration of cells. Further studies are required to

discover the precise mechanisms of KCC2 actions on neuroblast migration.

I studied then the functional role of these adult-born in their later stage of development

when they get mature and fully integrate in the OB. Our research was aimed at elucidating

the precise mechanisms that adult-born cells use to adjust the bulbar network functions in

the presence of rapid and persistent changes in odor environmental conditions occur. Our in

vivo data, reinforced with in vitro experiments, reveal a new form of structural remodeling

where mature dendritic spines of adult-born but not early-born granule cells relocate in an

activity-dependent manner. This relocation is preceded by the growth of spine head

filopodia (SHF) and is driven by the activity of principal cells. We also demonstrate that the

relocation of spines facilitates fast reorganization of OB network with functional

consequences for odor information processing.

Overall, studying the SVZ-OB model of adult neurogenesis helps us better comprehend the

myriad of complex processes responsible for proper development of neuronal network and

brings us closer to applying the knowledge about these mechanisms for cell replacement

therapies. To this end, however, it’s needed also to develop imaging strategies that will

allow avoiding utilization of exogenous labeling, thus extending their application to human

studies. In my research, I did imaging with polarized light which takes advantage of

intrinsic anisotropic properties of white matter consisting of dense bundles of myelinated

axons. My data suggests potential degradation of myelinated fibers observed with polarized

light in tissue samples from Parkinson patients as compared to those of healthy subjects.

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164

This highlights the potential of high-resolution label-free imaging methods, which can

complement and validate the established neuroimaging techniques.

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165

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ANNEX A - Tracking neuronal migration in adult

brain slices

Karen Bakhshetyan1 and Armen Saghatelyan1,2,*

Affiliations

1Cellular Neurobiology Unit, Institut Universitaire en santé mentale de Québec, Québec

City, QC, Canada G1J 2G3

2Department of Psychiatry and Neuroscience, Université Laval, Québec City, QC Canada

G1K 7P4

Acknowledgements

This work was supported by an operating grant from the Canadian Institutes of Health

Research (CIHR) to A.S. K.B. was partially supported by a training grant in neurophotonics

from CIHR. A.S. holds a Canada Research Chair in postnatal neurogenesis.

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A1. Résumé

La migration neuronale est l'un des processus fondamentaux sous-tendant le bon

assemblage et fonctionnement des circuits neuronaux. La majorité des précurseurs

neuronaux sont générés loin de leur site d'intégration, nécessitant ainsi une migration sur de

longues distances afin d’atteindre leur destination finale. Ce phénomène de migration

neuronale se produit non seulement au niveau du cerveau embryonnaire, mais aussi au sein

de quelques régions du cerveau adulte telles que le bulbe olfactif et l’hippocampe.

Cependant, les mécanismes sous-tendant la migration des cellules dans le cerveau adulte

sont peu connus, malgré leur pertinence clinique. Nous décrivons ici une méthode

d’imagerie en temps réel de la migration cellulaire sur des tranches de cerveau aiguës. Ce

procédé, combiné à des manipulations génétiques et / ou pharmacologiques de différents

mécanismes moléculaires, permet de déterminer la dynamique et les mécanismes

moléculaires de la migration des cellules dans le cerveau adulte. En outre, l'imagerie en

temps réel permet de suivre les mouvements des cellules dans un microenvironnement

similaire aux conditions in vivo, et d'étudier le déplacement des neuroblastes le long

d'autres composants cellulaires tels que les astrocytes et les vaisseaux sanguins.

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A2. Abstract

Neuronal migration is one of the fundamental processes underlying the proper assembly

and function of neural circuitry. The majority of neuronal precursors are generated far away

from their sites of integration and need to migrate substantial distances to reach their final

destination. Neuronal migration occurs not only in the embryonic brain but also in a few

regions of the adult brain such as the olfactory bulb and hippocampus. The mechanisms

orchestrating cell migration in the adult brain are, however, poorly understood, despite their

clinical relevance. Here we describe a method for time-lapse imaging of cell migration in

acute brain slices. This method, combined with genetic and/or pharmacological

manipulations of different molecular pathways, makes it possible to determine the

dynamics and molecular mechanisms of cell migration in the adult brain. In addition, time-

lapse imaging in acute brain slices makes it possible to monitor cell movement in a

microenvironment that closely resembles in vivo conditions and to study neuroblast

displacement along other cellular elements such as astrocytes and blood vessels.

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A3. Introduction

Neuronal precursors born in the adult subventricular zone (SVZ) retain the remarkable

capacity to migrate long distances along the rostral migratory stream (RMS) into the

olfactory bulb (OB) (Alvarez-Buylla and Garcia-Verdugo, 2002; Marin and Rubenstein,

2003). Neuronal precursors first migrate tangentially in chains along the RMS and then,

once in the OB, turn and migrate radially and individually out of the RMS into the bulbar

layers (Alvarez-Buylla and Garcia-Verdugo, 2002; Lois and Alvarez-Buylla, 1994;

Saghatelyan, 2009). A number of molecular cues affect neuronal migration in the adult

brain. These include polysialated neural cell adhesion molecule (PSA-NCAM) (Cremer et

al., 1994; Hu et al., 1996; Ono et al., 1994); members of the Slit (Kaneko et al., 2011),

ephrin-B (Conover et al., 2000), and integrin families (Belvindrah et al., 2007; Murase and

Horwitz, 2002); astrocyte-derived factor of unknown identity (Mason et al., 2001); the

ErbB4 (Anton et al., 2004) and prokineticin 2 (Ng et al., 2005) receptors; neuroblast-

derived GABA (Bolteus and Bordey, 2004); as well as various growth factors such as

GDNF (glial cell line-derived neurotrophic factor) (Paratcha et al., 2006), BDNF (brain-

derived neurotrophic factor) (Chiaramello et al., 2007; Snapyan et al., 2009), and VEGF

(vascular endothelial growth factor) (Bozoyan et al., 2012; Wittko et al., 2009). Reelin

and tenascin-R, two extracellular matrix molecules, also contribute to the radial migration

of neuronal precursors (David et al., 2013; Hack et al., 2002; Saghatelyan et al., 2004). In

addition to these molecular cues, cellular interactions between neuroblasts, astrocytes, and

blood vessels (BVs) are required for faithful migration toward the OB. Adult neuronal

precursors use BVs that topographically outline the RMS for their migration (Snapyan et

al., 2009; Whitman et al., 2009) and create a migratory path by repelling astrocytic

processes (Kaneko et al., 2011).

Understanding the cellular and molecular mechanisms orchestrating neuronal migration in

the adult brain may be important for the development of new strategies aimed at the

recruitment of endogenous and/or grafted neuronal precursors into damaged/diseased brain

areas. Indeed, the possibility of manipulating these mechanisms in order to increase the

number and dispersal of endogenous and/or grafted neuronal precursors in diseased areas

may open novel avenues for the treatment of devastating neurodegenerative diseases and

brain trauma. Interestingly, a massive migration of endogenous neuronal precursors from

their generation site (SVZ) to damaged areas of the brain has been observed following

stroke and in some neurodegenerative conditions such as Parkinson’s disease (Arvidsson et

al., 2002; Falk and Frisen, 2005). These observations suggest that the injured/diseased brain

may re-route the migration of neuronal precursors as a repair/compensation mechanism. If

so, this also suggests that intervening in this process might be an effective therapy for

treating degenerative diseases and brain injuries.

In this unit we will describe a method for monitoring and studying the migration of adult

neuronal precursors in acute brain slices in a microenvironment that closely resembles in

vivo conditions. We will describe stereotaxic injections of viral particles to infect neuronal

precursors in vivo, the labeling of BVs, the preparation of acute adult brain slices, time-

lapse video-imaging of cell migration, and analysis. While in this unit we describe neuronal

migration along BVs in the adult RMS. we have successfully applied the same procedure

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for monitoring neuronal migration along BVs in the ischemic striatum (Grade et al., 2013)

and along astrocytes in the developing and adult RMS (Bozoyan et al., 2012). A video

showing some of the procedures in this unit can be viewed at

http://www.jove.com/video/4061/ (Khlghatyan and Saghatelyan, 2012).

NOTE: All protocols using live animals must be reviewed and approved by institutional

animal care and use committees and must comply with applicable government legislation

and regulations on the humane care and use of laboratory animals.

A4. Basic Protocol

A4.1. Time-lapse video-imaging of neuronal migration in adult acute brain

slices

This protocol describes the basic steps for visualizing and studying neuronal migration in

acute adult brain slices. The use of acute slices makes it possible to assess cell migration in

a microenvironment that closely resembles in vivo conditions and in brain regions that are

difficult to access by in vivo imaging. While neuronal migration can be also visualized

using DIC optics (Snapyan et al., 2009), we recommend using time-lapse video-imaging of

fluorescently labeled neuronal precursors, which makes it possible to reliably monitor the

tangential and radial migration of individual cells and to follow the dynamics of leading

processes of neuroblasts (Bozoyan et al., 2012; David et al., 2013; Grade et al., 2013;

Khlghatyan and Saghatelyan, 2012; Snapyan et al., 2009). In the Support Protocol, we

describe the procedure to stereotaxically inject viral particles into the SVZ in order to infect

neuronal precursors.

A4.2. Materials

Acute brain slices prepared from adult mice infected with viral particles 3-7 days

(for monitoring tangential migration in the RMS) or 6-12 days (for monitoring radial

migration in the OB and the RMS of the OB) before the preparation of the slices (see

Support Protocol)

Surgical instruments used for extracting the brain (scissors, scalpel, forceps, etc.)

Artificial cerebrospinal fluid (ACSF) and cutting solutions (see recipe)

Vibratome (Microm HM 650V; Thermo Scientific)

Fluorescence wide-field upright microscope (BX61WI; Olympus) with a motorized z-

drive

482/35nm and 536/40nm excitation and emission filters for imaging GFP-labeled

cells

14-bit cooled CCD camera (CoolSnap HQ2; Photometrics) with 1392 x 1040 imaging

pixels

40x water immersion objective lens with a 0.8 numerical aperture or higher

(Olympus)

Illumination system (Lambda DG-4; Sutter Instruments) equipped with a 175 W

xenon lamp (30-100 ms excitation time per z plane)

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Imaging chamber (PH1 Series 20; Harvard Apparatus) mounted on a microscope

stage

Automatic heating system (TC-344B; Harvard Apparatus) for maintaining the

temperature in the imaging chamber at ~32°C

Multidimensional time-lapse data acquisition software (MetaMorph; Molecular

Devices).

Software for Z-stack image acquisition (usually 5–10 z-planes at 3-µm intervals)

every 15 s for 1 h (MetaMorph; Molecular Devices)

System for the continuous perfusion of slices with oxygenated ACSF at a 1-2 ml/min

flow rate

Slice fixation mesh (nylon with 0.12-mm-diameter and ~1-mm2 holes)

NOTE: The imaging and acquisition systems described in this unit are used in the authors’

laboratory and serve only as examples of the equipment required to perform time-lapse

imaging in acute slices. Other commercial and custom-made systems can be used.

However, we strongly recommend using imaging systems that can acquire time-lapse

images at least every 15 s. This is important to ensure the unambiguous identification of the

migratory and stationary phases of neuronal precursors, to better understand the dynamics

of cell migration.

A4.3. Protocol steps

A4.3.1. Preparation of acute slices and imaging

1. Inject 200 μl of Dextran Texas Red (10 mg/ml) into the tail vein and wait 15-30 min

before decapitating the mouse. Alternatively, inject 200 μl of Dextran Texas Red into

the left ventricle of the heart of a deeply anesthetized mouse and wait 2-3 min before

decapitating the animal.

2. Anesthetize the mouse in which neuronal precursors are labeled with GFP-encoding

viral vectors (see Support Protocol) and BVs are labeled with Dextran Texas Red.

3. Decapitate the mouse and rapidly immerse the head in ice-cold cutting solution. Use

scissors to remove the scalp and make an incision in the skull over the cerebellum.

4. Using forceps, gently remove the cranial bones to expose the lateral and dorsal sides

of the brain (Fig. 1A, B).

5. Without removing the brain, cut out the caudal part of the brain using a scalpel.

Sagittally remove approximately 2mm of brain tissue from the most lateral part of

each hemisphere (dotted lines in Fig. 1B). Cut the brain sagittally in the midline to

separate the two hemispheres from each other (Fig. 1C).

6. Gently remove both hemispheres from the skull using a spatula and place in ice-cold

cutting solution.

7. Place the two hemispheres side by side on a 4% agar block with the dorsal side

touching the agar (Fig. 1D). Then glue the agar block with both hemispheres to the

platform of a vibratome with the laterally cut sides facing to the platform, keeping the

medial side face up (Fig. 1E).

8. Place the platform in the chamber of the vibratome and fill the platform with cutting

solution. Keep the solution oxygenated throughout the slice preparation procedure by

bubbling with 95% O2/5% CO2.

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9. Prepare 250-μm-thick slices using the vibratome. Cut slices at low speed and with

high-frequency vibration to obtain high-quality slices.

10. Gently remove the slices from the cutting solution and place them in an incubation

chamber filled with oxygenated ACSF maintained at 32oC in a water bath.

NOTE: This procedure should be performed as quickly as possible to ensure high-quality

slices.

11. Start the perfusion of the imaging chamber mounted on a fluorescent upright

microscope (BX61WI; Olympus) with ACSF (1-2 ml/min). Make sure the

temperature remains stable at approximately 32°C.

12. Carefully transfer the slice into the imaging chamber of the microscope. To avoid

slice drift during imaging, gently cover the slice with a nylon mesh stretched on a

silver frame.

13. Engage a 10x objective to find the region of interest and make sure the nylon mesh

does not obstruct the field of view. Slightly reposition the mesh if it obstructs the

field of view and gently press down on the silver frame on which the nylon mesh is

stretched.

14. Engage a 40x objective and select the region for imaging.

15. Set the time-lapse acquisition parameters in MetaMorph (exposure time, the number

and distance of z-planes, time-lapse intervals, and total imaging duration). The

acquired time-lapse data is automatically saved by MetaMorph and the z-stack of

each time point is stored as a multipage Tiff file.

16. To avoid changes in the focal plane during imaging, continuously monitor the amount

and temperature of the ACSF in the imaging chamber and make sure the perfusion of

the ACSF remains constant.

17. We recommend occasionally verifying the data acquired during the imaging by

selecting Review Multidimensional Data in Metamorph. If there are significant drifts

or changes in focus that could affect the analysis, stop, discard the data, resolve the

issue, and restart the time-lapse recording.

18. Perform the imaging for at least 1 h, or more, to ensure that both the migratory and

stationary phases of cell migration are recorded. Avoid imaging cells that are too

close to the surface of the slice. Image at a depth of 20 to 100 μm. For reliable cell

tracking, we recommend short acquisition intervals (15-30 s).

19. To study particular molecular factors that might play role in neuroblast migration, the

ACSF can be supplemented with pharmacological agents such as

agonists/antagonists, growth factors/blockers, etc. Alternatively the expression of

particular genes of interest can be affected using viral vectors (gain-of-function or

loss-of-function approaches using overexpression constructs, siRNA, miRNA, the

injection of Cre viruses into animals where the gene of interest is flanked by two loxp

sites, etc.).

A4.3.2. Analysis of cell migration

Use Imaris software (Bitplane) for the 3D analysis of migrating cells.

20. The acquired time-lapse movie is loaded by dragging and dropping the first time

point Tiff file onto the Imaris program icon.

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21. When the movie is loaded in Imaris, adjust the brightness in the Display adjustment

window so that the cells are clearly distinguished from the background. Set the voxel

size based on objective calibration data and define the time-lapse acquisition interval

in the Image Properties and Set Equidistant Time Points windows, respectively.

22. To initiate cell tracking, run the Spots function in the Surpass menu and then follow

the wizard-style steps. In Source Channel, set the spot diameter and then define the

threshold for cell detection. Adjust the threshold if not all the migrating cells were

tracked. Next, set the maximal distance between the spot positions of consecutive

time points as well as the maximal time gap in cell tracking.

23. After creating the tracks, it is possible to make corrections to tracks whenever

necessary. For example, different tracks that are actually the same cell but were not

recognized as such by Imaris can be connected together. The parts of tracks that on

careful examination belong to different cells but were misinterpreted as one cell can

also be disconnected.

24. Delete all irrelevant tracks such as those that were tracked for too short a time. Also

delete the tracks that remained stationary throughout the imaging period since these

cells may have been damaged during the cutting procedure or may be non-migratory

cells.

25. Once all the corrections have been made, export the cell tracking data to an Excel file

by clicking on the Export All Statistics to File icon. The exported file contains per

time point information on the number of tracks and cell speed and displacement, as

well as per cell information on displacement length, track duration, and track length.

26. Load the data from Imaris into a custom-made Excel template to extract the final

migration data and to identify the migratory and stationary phases of each cell. The

stationary phases are defined by a speed threshold of 0.03 µm/s. This corresponds to

the means (± SD) of cell displacement per second for all cells multiplied by 1.5.

27. The final cell tracking statistics data include following parameters: total migration

distance per cell, mean speed per cell, percentage of migratory phases per cell, and

percentage of migrating cells per time point.

A5. Support Protocol

A5.1. Stereotaxic injection of viral vectors into the SVZ of the adult mouse

While neuroblast migration in acute brain slices can be also monitored using DIC optics

(Snapyan et al., 2009), we recommend imaging fluorescently labeled neuronal precursors

(Bozoyan et al., 2012; David et al., 2013; Grade et al., 2013; Khlghatyan and Saghatelyan,

2012; Snapyan et al., 2009). This makes it possible to reliably track cell migration during

prolonged periods, which is quite difficult using DIC optics. In addition, cells can be

tracked deeper in the tissue, and the dynamics of leading process of neuroblasts can be

visualized (Grade et al., 2013). In this Support Protocol, we describe the basic steps for

labeling neuroblasts with GFP-encoding viral vectors. This procedure can also be used to

overexpress or knock-out and/or knock-down the protein of interest and study its role in

neuronal precursor migration. We use AAV, retroviruses, and/or lentiviruses to label

neuronal precursors in the SVZ using the following coordinates: anterio-posterior (AP)

0.70; medio-lateral (ML) 1.20, and dorso-ventral (DV) 1.90.

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A5.2. Materials

Micropipette puller

Stereotaxic injection apparatus equipped with a digital stereotaxic coordinate read-out

system (WPI)

Mouse adaptor mounted on a stereotaxic apparatus (WPI)

Sterile surgical instruments such as scissors, scalpels, forceps, etc.

Microdrill system

Heating pad

Nanoliter injector (WPI) and nanoliter injector controller

Binocular microscope

Proviodine or 70% ethanol

A5.3. Protocol steps

1. Sterilize all the instruments before starting the surgery using a bead sterilizer or an

autoclave.

2. Pull very thin-tipped (1-2 μm) micropipettes using a micropipette puller. We use

glass capillaries suitable for the nanoliter injector.

3. Backfill the pipettes with paraffin oil until half-full and insert the plunger of a

nanoliter injector.

4. Using a nanoliter injector controller, force the paraffin oil downwards with the

plunger until a small drop extrudes from the tip. Lower the pipette tip into a small

drop (0.5-1 μl) of solution containing the viral particles.

5. Use the withdraw function of the nanoliter injector to backfill the pipette.

6. Anesthetize 22-24-g adult C57Bl/6 mice using an IP injection of ketamine/xylazine

(10 mg/1 mg per 10 g of body weight) or by isoflurane inhalation.

7. Shave the head of the mouse and use wet gauze to thoroughly clean the surgical site.

8. Inject lidocaine under the skin at the surgical site and administer an analgesic IP

(anafen 10 mg/kg).

9. Place the mouse in the stereotaxic frame on the heating pad and fix the head in place

using ear and tooth bars.

10. Use gauze soaked with Proviodine or 70% ethanol to disinfect the surgical site.

11. Cut the skin at the surgical site and expose the underlying skull.

12. Use the bregma and midline as the zero coordinates to set the anterio-posterior (AP)

and medio-lateral (ML) stereotaxic coordinates, respectively.

13. Drill a small hole in the skull over each hemisphere at the desired coordinates. Avoid

damaging the underlying brain tissue.

14. Carefully remove the dura using an angled needle or forceps.

15. Set the zero for the dorso-ventral (DV) stereotaxic coordinates on the surface of the

brain.

16. Slowly insert the glass micropipette tip into the brain using the desired coordinates

and slowly inject a small amount (we inject 100-500 nl at 5 nl/s) of the solution

containing the viral particles.

17. Wait 1-2 min and then slowly withdraw the glass micropipette.

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18. Suture the skin over the surgical site. Wait until the animal recovers, and return it to

its cage.

A5.4. Reagents and Solutions

Use deionized, distilled water for all solutions. Maintain solutions at a pH of 7.3-7.4, and

continually oxygenate by bubbling with 95% O2/5% CO2. Chill the cutting solution until

ice cold using liquid nitrogen.

Cutting solution

210.3 mM sucrose

3 mM KCl

3 mM MgCl2•6H2O

0.5 mM CaCl2•2H2O

26 mM NaHCO3

1.25 mM NaH2PO4

20 mM glucose

Artificial cerebrospinal fluid (ACSF)

125 mM NaCl

3 mM KCl

1.3 mM MgCl2•6H2O

2 mM CaCl2•2H2O

26 mM NaHCO3

1.25 mM NaH2PO4

20 glucose

A6. Commentary

A6.1 Background Information

Neuronal migration is one of the fundamental processes underlying proper brain

development. In the adult brain, neuronal migration has largely ceased and the SVZ-OB

pathway is probably the only region where massive neuronal migration still occurs in

normal conditions. It should be noted, however, that neuronal migration in the adult brain

can be induced by brain damage and/or neurodegenerative diseases (Arvidsson et al., 2002;

Grade et al., 2013; Lindvall et al., 2004; Ohab et al., 2006). Interestingly, induced neuronal

migration toward damaged/diseased brain areas shares at least some features of constitutive

neuronal migration in the SVZ-OB pathway. For example, neuronal precursors use BVs as

a physical scaffold and a source of molecular cues in both the RMS (Snapyan et al., 2009;

Whitman et al., 2009) and the ischemic striatum (Grade et al., 2013; Ohab et al., 2006).

Astrocytes, which play an important role in neuroblast migration in the RMS (Kaneko et

al., 2011; Lois and Alvarez-Buylla, 1994; Snapyan et al., 2009), are also involved in

neuronal migration in ischemic brain regions (Imitola et al., 2004; Yan et al., 2007).

Studying the cellular and molecular mechanisms of neuronal migration in the SVZ-OB

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pathway is thus not only important from a fundamental point of view but may also have

clinical relevance. It is conceivable that a better understanding of the mechanisms

controlling neuronal migration in the undamaged adult brain may be of use in the

development of new strategies to re-route neuronal precursors from their natural migratory

pathway and/or to increase the dispersal of grafted neuronal progenitors in

diseased/damaged brain areas (Falk and Frisen, 2005; Lindvall et al., 2004; Massouh and

Saghatelyan, 2010). Since ischemic damage and neurodegeneration mostly occurs in the

adult/aged brain, studying the mechanisms of neuronal migration in the adult SVZ-OB

where massive neuronal migration is observed (Lois and Alvarez-Buylla, 1994; Marin and

Rubenstein, 2003) may be particularly important for the development of new cell

replacement strategies.

There are several approaches for studying neuroblast migration. These include 2D cultures

of SVZ progenitor cells (Lois et al., 1996), 3D cultures of SVZ explants on Matrigel

(Courtes et al., 2011; Lois et al., 1996; Saghatelyan et al., 2004), organotypic cultures

(Mejia-Gervacio et al., 2012; Mejia-Gervacio et al., 2011; Murase and Horwitz, 2002), and

acute brain slices (Bolteus and Bordey, 2004; Bozoyan et al., 2012; Comte et al., 2011;

David et al., 2013; Grade et al., 2013; Platel et al., 2008; Snapyan et al., 2009). Each of

these methods has advantages and limitations. Time-lapse imaging of adult acute slices

allows the monitoring of neuroblast migration in a microenvironment that closely

resembles in vivo conditions. In addition, complex cellular and molecular interactions with

other cellular elements such as astrocytes and BVs can be studied (Bolteus and Bordey,

2004; Bozoyan et al., 2012; Comte et al., 2011; Platel et al., 2008; Snapyan et al., 2009).

The main drawback of this method is that cutting-induced damage may affect neuronal

migration.

If the role of a particular molecular pathway in neuroblasts needs to be studied without

interference from astrocytes and/or endothelial cells, 2D and 3D SVZ cultures may be used

(Courtes et al., 2011; Saghatelyan et al., 2004). In addition, co-cultures of neuroblasts and

astrocytes (Bozoyan et al., 2012; Garcia-Marques et al., 2010) or endothelial cells (Snapyan

et al., 2009) may be used to study the role of these cells in neuronal migration. It should be

noted, however, that neuroblasts migrating out of SVZ explants are in a microenvironment

that is completely different from in vivo conditions, which may affect the dynamics and

mechanisms of cell migration. Organotypic cultures have also been used to study neuronal

migration, especially in the developing RMS (Mejia-Gervacio et al., 2012; Mejia-Gervacio

et al., 2011; Murase and Horwitz, 2002). This method provides the advantage of studying

neuroblast migration over long periods of time (up to several days). It should be kept in

mind, however, that cellular organization (Bozoyan et al., 2012; Law et al., 1999; Peretto et

al., 2005) and neuronal migration (Bozoyan et al., 2012; David et al., 2013) are quite

different in the developing and the adult RMS. Moreover, long-term culturing may also

alter the migratory properties of the cells. Regardless of the method used to study

neuroblast migration, we recommend using short intervals (15-30 s) between consecutive

acquisitions in order to reliably identify the migratory and stationary phases and to quantify

of the speed of migration solely during migratory phases. Moreover, an assessment of the

periodicity of the migratory and stationary phases as well as the duration of the migratory

phases provide additional information on the dynamics of cell movement and the impact of

particular treatments on cell migration (Bortone and Polleux, 2009; Snapyan et al., 2009).

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A6.2. Critical Parameters and Troubleshooting

Preparation of acute brain slices. The quality of the acute brain slices is particularly

important for the successful implementation of time-lapse video-imaging of cell migration.

It is thus important to carefully inspect the quality of the slices using DIC optics before

starting the time-lapse video-imaging. Acute slices from the adult mouse brain (2-3 months

old) are particularly challenging to prepare. A number of modifications in the ionic

composition of the cutting solution can improve cell viability and decrease cellular swelling

and damage. For example, increasing Mg2+ and decreasing Ca2+ concentrations to dampen

synaptic transmission as well as replacing Na+ by sucrose to dampen neuronal excitability

(see cutting solution recipe) during the cutting procedure improve the quality of the slices.

It is also important to prepare the slices as quickly as possible. The brain should be rapidly

extracted and immersed in ice-cold cutting solution. The vibratome chamber should be also

filled with oxygenated ice-cold solution, and the entire slicing procedure should be

performed when the cutting solution is still ice cold. The solution should be oxygenated

throughout the slice preparation, and the brain should be cut at low speed and high

frequency vibration to obtain high-quality slices.

If BVs do not need to be labeled, we recommend that the mouse be transcardially perfused

with ice-cold cutting solution before removing the brain. In our hands, this approach

markedly improves the quality of the slices.

Stable conditions during the time-lapse video-imaging. An irregular flow of ACSF,

unstable temperature, and slice drift may all lead to changes in the focal plane and

unreliable imaging of cell migration. Make sure that all these parameters are stable before

transferring the slice into the imaging chamber and starting the time-lapse video-imaging.

ACSF perfusion can be pump-driven or gravity-fed. In both cases, irregular flows of ACSF

may be resolved by adjusting the rates of inflow and outflow. We use a gravity-fed

perfusion system and an ultra-quiet imaging chamber (RC-27D; Warner Instruments) that

eliminates vibrations and provides a more constant flow of ACSF.

Temperature fluctuation during imaging is another critical parameter that must be

controlled. In fact, one of the most effective ways of blocking cell migration is to perform

imaging at room temperature. We usually image cell migration at about 32˚C. In our hands,

increasing the temperature above this value leads to unstable imaging of cell migration.

This is mainly due to the formation of numerous small bubbles at the interface between the

heated ACSF and the air in the open-bath chamber. The temperature should not fluctuate

more than ±1˚C to ensure stable imaging. Pre-heated ACSF can be used to avoid

overheating the imaging chamber.

Slice drift is another parameter that may affect the time-lapse video-imaging of cell

migration. This can be controlled using slice anchors. We use custom slice anchors made

from nylon mesh that is tightly stretched over and glued to a silver frame. We usually

prepare new slice anchors every month or as soon as we notice significant slice drift.

Selecting the region for imaging. It is important to carefully inspect the slice and select the

appropriate region for imaging. The RMS in the field of view should be continuous and

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197

uninterrupted. Do not image cells in the region where the RMS was cut or is spanned with

axonal bundles. Avoid imaging cells that are too close to the slice surface. Rather, image

cells at a depth of 20 to 100 μm. For reliable cell tracking we recommend short acquisition

intervals (15-30 s). Perform the imaging for at least 1 h, or more, so that both the migratory

and stationary phases of cell migration are recorded.

A6.3. Anticipated Results

Time-lapse video-imaging of cell migration in acute adult brain slices improves our

understanding of the dynamics and cellular and molecular mechanisms of cell movement in

a microenvironment that closely resembles in vivo conditions. Typical real-time video

images of neuroblasts migrating tangentially and radially are shown in Fig. 2. Neuronal

precursors labeled by the stereotaxic injection of GFP-encoding retroviruses into the SVZ

are shown in Fig. 2A, B. Images of tangential migration in the RMS in acute slices

prepared 5 days after the viral vector injections are shown in Fig. 2C. Images of radial

migration in the OB in acute slices prepared 9 days after the injection of the viral vector

into the SVZ are shown in Fig. 2D. Regions of interest containing GFP-labeled neuroblasts

were selected based on the parameters described in the Critical Parameters and

Troubleshooting section. Time-lapse video-imaging of cell migration was performed for 1 h

with 15 s intervals between consecutive acquisitions. The exposure time was 30 ms, and

seven z-planes with 3-μm intervals were used to acquire time-lapse movies. Time-lapse

imaging snapshots containing several migrating cells in the RMS (numbered arrows; each

number reflect different cell) are shown in Fig. 2C. Migration in the RMS was relatively

rapid and ranged from 120 to 180 μm/h. The mean distance that neuroblasts migrate during

1 h is about 60 μm, and cells spend approximately 40% in the migratory phase.

When neuroblasts arrive in the OB, they change their mode of migration from tangential to

radial (Fig. 2D). Several modes of migration can be studied in the OB. Tangential

migration in the RMS of the OB can be studied to determine whether particular molecular

cues play a role in stopping tangential migration (David et al., 2013). Radial migration in

the RMS of the OB and the granule cell layer (GCL) can be also studied. For example,

time-lapse imaging of neuroblasts in the OB showed that extracellular matrix glycoprotein

does not affect tangential migration in the RMS of the OB but sustained radial migration in

the RMS of the OB and the GCL (David et al., 2013). The distance covered by tangentially

and radially migrating neuroblasts in the OB and the RMS of the OB is shorter than that of

tangentially migrating cells in the RMS. This is mainly due to lower speed of migration

(David et al., 2013). Radially migrating neuroblasts in the OB migrated at about 80-120

μm/h, had roughly 40% of migratory phases, and propagated an average of 35±5 μm during

1 h of imaging.

A6.4. Time Considerations

It takes an average of 30-40 min to perform the stereotaxic injections of the viral vectors

into the SVZ. The survival time after injection can vary and depends on the type of

migration to be studied. If tangential migration of neuroblasts is to be studied, the

preparation of the acute slices and time-lapse imaging may be performed 3-7 days after the

viral vector injection. If radial migration of neuroblasts in the OB and the RMS of the OB

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is to be studied, the time-lapse imaging of acute slices may be performed 7-12 days after

the injection of the viral vector into the SVZ. The acute slices should be prepared as

quickly as possible. Acute slices can be kept for up to 6-8 h.

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A8. Figures

Figure 1. Tissue preparation

A: Dorsal view of the mouse head with removed scalp. Dotted line represents the line of incision for removal

of cranial bones.

B: Dorsal view of the brain with removed cranial bones.

C: Dorsal view of the brain with removed caudal and lateral parts.

D: Separated hemispheres placed on agar block.

E: The agar block with hemispheres glued on vibratome platform.

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Figure 2. Tracking tangential and radial migration in adult brain slices

A. Schematic diagram of adult-born cell labeling by stereotaxic injection of GFP-encoding viral particles into

the SVZ. The adult forebrain slices are prepared 3-7 days or 7-12 days after the injection of the viral

vectors for monitoring tangential and radial migration, respectively.

B: Tangential migration is monitored in the RMS with slices cut sagittally, while radial migration is recorded

in the horizontal sections of OB and the RMS of the OB.

C: Time-lapse wide-field imaging of GFP-labeled neuronal precursors in acute slices of the adult mouse

forebrain. GFP-expressing retrovirus was injected into the SVZ 4 days before time-lapse imaging in the

RMS. The arrows and numbers indicate the soma of the different migratory cells. Time is indicated in

minutes in the upper right corner of each photograph.

D: Time-lapse imaging of GFP-labeled neuroblasts in acute OB slices. GFP-expressing retrovirus was

injected into the SVZ 8 days before time-lapse imaging. The arrows and numbers indicate the soma of

different migratory cells. Time is indicated in minutes in the upper right corner of each photograph.