Citation
Identification and Characterization of Neural Stem and Progenitor Cells in Vivo and in Vitro

Material Information

Title:
Identification and Characterization of Neural Stem and Progenitor Cells in Vivo and in Vitro
Creator:
WALTON, NOAH MATTHEW ( Author, Primary )
Copyright Date:
2008

Subjects

Subjects / Keywords:
Astrocytes ( jstor )
Cell growth ( jstor )
Cells ( jstor )
Cultured cells ( jstor )
In vitro fertilization ( jstor )
Microglia ( jstor )
Neuroblasts ( jstor )
Neurogenesis ( jstor )
Neurons ( jstor )
Progenitor cells ( jstor )

Record Information

Source Institution:
University of Florida
Holding Location:
University of Florida
Rights Management:
Copyright Noah Matthew Walton. Permission granted to the University of Florida to digitize, archive and distribute this item for non-profit research and educational purposes. Any reuse of this item in excess of fair use or other copyright exemptions requires permission of the copyright holder.
Embargo Date:
12/31/2016

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Full Text

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IDENTIFICATION AND CHARACTERIZATION OF NEURAL STEM AND PROGENITOR CELLS IN VIVO AND IN VITRO By NOAH MATTHEW WALTON A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 2006

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Copyright 2006 by Noah Matthew Walton

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This work is dedicated to my parents, who supported me both before and long after I should have become self-sufficient.

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iv ACKNOWLEDGMENTS Many people have had influence on the work contained within the pages of this document, including contributions scientific and otherwise: most notably, my mentor Dennis Steindler, who oversaw my training and development as not only a scientist but a human being; Bjorn Scheffler, whose time and effort proved to be the gold standard for what a scientist is; my committee members, who generously provided time, guidance, and reagents for my betterment; and my labmates, who toiled with me. I would also like to thank the members of my soccer team Dynamo, who provided a much-needed venue for me to display my meager nonscientific talents.

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v TABLE OF CONTENTS page ACKNOWLEDGMENTS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . iv LIST OF FIGURES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . ix ABBREVIATIONS...........................................................................................................xii ABSTRACT . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . xiii CHAPTER 1 INTRODUCTION TO ADULT NEURAL STEM AND PROGENITOR CELLS . . . . . 1 Overview of Neural Stem Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1 Adult Neural Stem/Progenitor Cells as a Therapeutic Cell Source . . . . . . . . . . . . . . . . . . . . . . . . . . . . 3 2 MATERIALS AND METHODS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Supplier Information . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Strains of Mice . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Cell Lines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 7 Reagents . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Aphidicolin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Artificial CSF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 Avertin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 bFGF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 8 BDNF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 BrDU . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Cyclosporin A . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Ara C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 DAPI . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 9 Dibutyl Cyclic AMP . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 EGF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 EGCG . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 FGF-8 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Hoechst 33528 . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Immunocytochemistry Hybridization Buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 IBMX . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 10 Laminin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Mitomycin C . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Mitosox Dye . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 NSC Differentiation Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 NSC Proliferation Culture Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 11 Nerve Growth Factor . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Neurosphere Proliferation Media . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12

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vi 4% Paraformaldehyde . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Picrotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Pleiotrophin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Poly-L-Lysine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Polyornithine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 12 Propidium Iodide . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Retinoic Acid . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 RIPA Buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Senescence-Associated -Galactosidase Solution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Sonic Hedgehog . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 TEA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Tetrodotoxin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 13 Trypsin . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Western Blot Hybridization Buffer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Xanthosine . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Antibody List . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Primary Antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Immunocytochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 14 Western Blot . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16 Immunoprecipitation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16 Secondary Antibodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16 Immunocytochemistry . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 16 Western Blot . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 Methods . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 Isolation, Derivation, and Expansion of Rodent NSCs In Vitro . . . . . . . . . . . . . . . . . 17 Regional microdissection of neuropoietic cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 17 Isolation and expansiton of primary subventricular dissociates . . . . . . . . . . 18 Neurosphere generation, differentiation, and quantification . . . . . . . . . . . . . . . 19 Inducible differentiation of rodent SVZ-derived cells . . . . . . . . . . . . . . . . . . . . . . . . 20 Tissue sectioning and storage . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 Antibody application . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 21 Nuclear lableling and lipophilic dye application . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 Electrophysiology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 22 Documentation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23 Live-cell microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24 Unbiased cell counting . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 24 Electron Microscopy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 Western blotting and semi-quantitative protein analysis . . . . . . . . . . . . . . . . . . . . . . . . . . . . 25 FACS studies on cell cycle/ploidy . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 Clonal seeding experiments in adherent conditions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 26 In vivo BrDU administration . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 Cell growth experiments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 Apoptosis/necrosis detection . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 27 MitoSox dye addition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28 Senescence-associated -galactosidase stain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 28

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vii Rescue of endogenous neurogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 Microglial depletion . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 29 Immunoprecipitation studies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 Human neural tissue collection and culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 30 Cellular irradiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 Karyotyping . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 Generation of human neuronal cell types . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 32 Transplantation of human progenitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33 ELISA . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34 3 PHENOTYPIC AND FUNCTIONAL CHARACTERIZATION OF ADULT BRAIN NEUROPOIESIS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35 Recapitulation of SVZ Neurogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36 Development of Neuroblasts into Interneurons . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38 Dynamic Characterization and In Vitro Correlation of Neurogenesis . . . . . . . . . . . . . . . . . . . . . . 39 SVZ Astrocytes Mature into Mature Interneuronal Phenotypes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 40 4 TUJ1+ CELLS IDENTIFY NEUROGENIC CELLS OF THE SUBVENTRICULAR ZONE AND CHARACTERIZE THE ASTROCYTE-TO-NEURON GESTALT . . . 49 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 49 Tuj1 Defines a Subset of Immature Glial Cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 52 Tuj1 Expression Characterizes Cells Undergoing Astrocyte-to-Neuron Differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53 Developmentally Intermediate Cells Possess Enhanced Multipotentiality and Proliferative Potential . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 57 5 BROMODEOXYURIDINE HAS CYTOTOXIC EFFECTS ON NEURAL STEM CELLS AND IMMORTAL CELL LINES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 67 BrDU Attenuates the Growth of Primary Neurogenic Cells and Cell Lines . . . . . . . . . . . . . 68 BrDU Affects Neurosphere Formation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 69 BrDU Induces Senescence In Vivo and In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 70 6 NEURAL STEM CELLS ARE DEPENDENT ON MICROGLIAL SIGNALING FOR NEUROGENESIS AND NEURONAL DEVELOPMENT BUT NOT SELFRENEWAL OR MULTIPOTENTIALITY . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 77 Loss of Neurogenesis Accompanies Continued NSC Proliferation . . . . . . . . . . . . . . . . . . . . . . . . . . 81 Loss of Inducible Neurogenesis is not Linked to NSC Depletion . . . . . . . . . . . . . . . . . . . . . . . . . . . . 83 Microglial Depletion Correlates with Loss of Inducible Neurogenesis . . . . . . . . . . . . . . . . . . . . 85 Microglia Rescue Inducible Neurogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

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viii 7 DERIVATION AND LARGE-SCALE EXPANSION OF MULTIPOTENT ASTROGLIAL PROGENITORS FROM ADULT HUMAN BRAIN . . . . . . . . . . . . . . . . . . . 102 Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 102 Characterization and Expansion of Primary Cells as AHNPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 105 AHNPs Maintain Growth Sensitivity and Avoid Immortalization . . . . . . . . . . . . . . . . . . . . . . . . . 109 Expanded AHNPs Function as a Transplantable, Modifiable Cell Source . . . . . . . . . . . . . 112 8 DISCUSSION AND CONCLUSIONS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 133 Isolation, Establishment, and Characterization of SVZ Neurogenesis In Vitro . . . . . . 133 Tuj1 Characterizes the Astrocyte-to-Neuron Metamorphosis of SVZ Neurogenesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 136 Bromodeoxyuridine Alters NSC Function and Induces Senescence In Vivo and In Vitro . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 139 Microglia Undergo Age-Dependent Neurogenic and Neuronizing Support Roles . 141 Derivation, Expansion, and Potential Uses of Human Progenitor Cells . . . . . . . . . . . . . . . . . 145 LIST OF REFERENCES . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 150 BIOGRAPHICAL SKETCH . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 162

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ix LIST OF FIGURES Figure page 1-1 Rostral migratory stream of the adult rodent CNS . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 6 3-1 Monolayers of SVZ cells can be inducibly differentiated into newborn neuroblasts . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 42 3-2 Generation of cellular heterogeneity accompanies differentiation. . . . . . . . . . . . . . . . . . . . . . . . . 43 3-3 SVZ-generated neuroblasts develop into interneuron phenotypes . . . . . . . . . . . . . . . . . . . . . . . . . 44 3-4 A mitotically active, multipotent cell emerges transiently during early stages of in vitro neuropoiesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45 3-5 Differentiating cells recapitulate membrane development consistent with a glialto-neuron transition . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 46 3-6 Immunophenotypic alteration accompanying neurogenesis and maturation of neuronal progeny following inducible differentiation in culture . . . . . . . . . . . . . . . . . . . . . . . . . . . . 48 4-1 Tuj1 labels putative NSC/progenitor populations . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 59 4-2 Tuj1 is coexpressed in immature and astrocytic cells derived from both adherent monolayers and neurospheres . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60 4-3 Tuj1 + cells characterize the astrocyte-to-neuron transition of rodent SVZ cells . . . . . 61 4-4 Immature markers are lost from Tuj1 + cells upon induction of differentiation . . . . . . . 62 4-5 Tuj1 + cells drive neurogenesis through characteristic mitotic dynamics . . . . . . . . . . . . . . . . 63 4-6 Intermediate progenitors retain multipotentiality and self-renewal . . . . . . . . . . . . . . . . . . . . . . . . 65 5-1 BrDU abrogates cell growth in cell lines and neurogenic astrocytes . . . . . . . . . . . . . . . . . . . . . 71 5-2 BrDU addition affects NS number and size in NS derived from young and adult animals . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 72 5-3 Systemic BrDU administration results in increases in senescent cell types . . . . . . . . . . . . 73 5-4 Addition of BrDU results in morphological alteration of both subventricular astrocytes and cell lines . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 74 5-5 BrDU does not abrogate growth through necrosis or apoptosis, but does induce mitochondrial stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 75

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x 6-1 An adherent culture model allows cultivation and inducible neurogenesis in an SVZ-derived astrocyte monolayer . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90 6-2 Neuropoietic tissues are indefinitely expandable as astrocyte monolayers, but lose mitogen-withdrawal prompted neurogenic potential . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 91 6-3 Expanded cultures maintain neurosphere-forming cells. PD3 and PD300 SVZ dissociates contain similar numbers of neurosphere-forming cells that do not differ in number, appearance, or size. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 92 6-4 Neurospheres derived from PD3 and PD300 SVZ cells are multipotent . . . . . . . . . . . . . . . . 93 6-5 Non-neurogenic cells promote neuroblast generation in antiadhesive culture systems . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 94 6-6 Microglia depletion parallels loss of inducible neurogenesis in proliferating cultures . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 95 6-7 XS has negligible effects on proliferative rate and NSC frequency . . . . . . . . . . . . . . . . . . . . . . . 96 6-8 Levels of apoptosis and necrosis in microglia remain constant throughout differentiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 97 6-9 Microglial cells modulate inducible neurogenesis through a soluble factor(s) . . . . . . . 98 6-10 CD11b/MAC1-conjugated (MAC1-SAP) and unconjugated ( uncon-SAP) saporin toxin application to proliferating and differentiating cultures does not affect microglial depletion or proliferation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 99 6-11 Microglial undergo age-dependent alterations in neurogenic support and neuronal development . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 100 7-1 Surgical specimens employed for derivation of AHNP cell strains . . . . . . . . . . . . . . . . . . . . . 117 7-2 Culture paradigm for the generation of AHNPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 118 7-3 Expansion of primary neural cells as a homogenous population of AHNPs . . . . . . . . 119 7-4 AHNPs avoid immortalizing mutations, and exhibit mitogenand telomerasedependent growth . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 120 7-5 AHNPs maintain viability and assume glial phenotypes upon ventricular transplantation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 122 7-6 Optimized lentiviral infection of AHNPs . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 123 7-7 Transfection of human progenitors with GDNF . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 124

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xi 7-8 Genetically modified human progenitors integrate and express transgenes in vivo . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 126 7-9 Cortically-implanted AHNPs adopt predominantly neuronal fates . . . . . . . . . . . . . . . . . . . . . . 128 7-10 Distribution and phenotype of transplanted progenitors . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 130 7-11 Differentiation of AHNPs into neuronal cell types . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 131 7-12 Generation of mature and tyrpsine-hydroxylase-expressing neurons . . . . . . . . . . . . . . . . . . . 132

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xii ABBREVIATIONS AHNP, adult human neural progenitor; AraC, cytosine D -Arabinofuranoside; BDNF, brain-derived neurotrophic factor; BrDU, 5’bromo-2-deoxyuridine; BSA, bovine serum albumin; CNS, central nervous system; CSF, cerebrospinal fluid; DAPI, 4’,6-diamidino2-phenylindole dihydrochloride; DF, dulbecco’s modified eagle medium with F12 supplements; DIV, days in vitro; DMSO, dimethyl sulfoxide; DTT, dithiothreonine; ECL, enhanced chemiluminescence system; EGCG, epigallocatechin-3-gallate; EGF, epidermal growth factor; eGFP, enhanced green fluorescent protein; ELISA, enzyme-linked immunosorbent sandwich assay; ES, embryonic stem; FACS, fluorescent activated cell sorting; FCS, fetal calf serum; bFGF, basic fibroblast growth factor; GABA, gamma aminobutyric acid; GAD, glutamic acid decarboxylase; GDNF, glial-derived neurotrophic factor; GFAP, glial fibrillary acidic protein; HNA, human ribonuclear protein; hTERT, human telomerase, IBMX, 3-isobutyl-1-methylxanthine; LGE, lateral ganglionic eminence; LPO, laminin-polyornithine; NGF, nerve growth factor; NS, neurosphere; NSC, neural stem cell; PBS, phosphate buffered saline; PD, population doubling; PDGF, platelet-derived growth factor; PNS, peripheral nervous system; RMS, rostral migratory stream; SA--Gal, senescence-associated beta galactosidase; SSC, sodium chloride/sodium citrate; SVZ, subventricular zone; TBST, tris buffered saline containing tween-20; TEA, tetraethylammonium; TTX, tetrodotoxin; TUNEL, TdTmediated dUTP nick-end labeling; XS, xanthosine.

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xiii Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy IDENTIFICATION AND CHARACTERIZATION OF NEURAL STEM AND PROGENITOR CELLS IN VIVO AND IN VITRO By Noah Matthew Walton December 2006 Chair: Dennis Steindler Major Department: Medical SciencesNeuroscience Neural stem cells (NSCs) of the postnatal mammalian forebrain are maintained throughout life in tissue surrounding the paired lateral ventricles. These cells give rise to neurons and glia throughout life, and have generated recent interest as a donor source of replacement material for damaged or injured neural tissue. Fundamental challenges to establishing therapeutically useful populations include isolating and maintaining NSCs, manipulation and characterization of differentiation, and assessing development of their committed progeny. Neither in vivo or in vitro approaches to these issues are alone capable of prospectively identifying NSCs, effectively evaluating dynamics of differentiation, quantification of cellular kinetics, identification of intermediate cell types during differentiation, and/or the identification of neurogenic correlates in vivo. As such, current methods of investigation do not allow for effective, direct characterization of many of the aspects involved in identification of NSCs or of factors pertinent to the development of their progeny. To address these questions, we have created an experimental paradigm based on a novel two-dimensional in vitro model capable of sustaining subventricular NSCs in conditions conducive to continual observation and

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xiv characterization. Inducing differentiation in this system leads to the rapid generation of neurons and glia in a process that faithfully and accurately recapitulates the developmental progression of in vivo subventricular zone (SVZ) neurogenesis. Using this model, we have identified several novel cell types involved in neurogenesis, including a unique population of Tuj1 + astrocytes, which characterize the intermediate cells involved in the astrocyte-to-neuron transition, and microglia, which facilitate neurogenesis through a soluble factor contribution. As rodent SVZ was expandable in our culture system without loss of attendant NSCs, we applied growth conditions favoring the maintenance and expansion of NSCs to human neural tissue, resulting in the identification and expansion of a uniform astrotypic progenitor population that retains the potential to differentiate to neurons in vivo and in vitro. Through the use of a reductionist culture model, it is possible to further characterize the behaviors and cells of postnatal neurogenesis, allowing unprecedented observations to be made on an increasingly complex biological process.

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1 CHAPTER 1 INTRODUCTION TO ADULT NEURAL STEM AND PROGENITOR CELLS Overview of Neural Stem Cells In higher vertebrates, neural stem cells (NSCs) are restricted to discrete neuropoietic zones, where lifelong neurogenesis occurs (reviewed in Alvarez-Buylla and Lim, 2004). The largest consensus neuropoietic zone is located in the mammalian forebrain, within the subventricular zone (SVZ), which harbors NSCs that are maintained as a 2-3 cell thick layer of subependymal cells which emerge between the medial and lateral ganglionic eminences (Doetsch et al. 1997). Throughout life, these cells constitutively generate populations of migrating neuroblasts, which transit the rostral migratory stream (RMS, Fig. 1-1) to the olfactory bulb and develop into mature interneurons (Lois and Alvarez-Buylla, 1994). Correlate neurogenic regions in the human SVZ have been shown to exist, but do not appear to maintain an analogous RMS (Sanai et al. 2004). Establishing consensus markers for identification of NSCs within various neuropoietic niches has proven difficult (reviewed in Morshead and van der Kooy, 2004) however multiple studies have indicated that the stem cells of the rodent SVZ are glial in nature (Doetsch et al. 1999; Laywell et al. 2000; Imura et al. 2003). In animals in which the RMS is temporarily ablated through addition of antimitotic compounds, populations of mitotically quiescent subependymal astrocytes were sufficient to regenerate the RMS (Doetsch et al. 1999 a, b), suggesting that a population of mitotically quiescent astrocytes acts as a founding population for subsequent neurogenic events. In a complementary study, it was found that depletion of the RMS was a

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2 sufficient signal to induce NSCs to actively divide ( Morshead et al. 1994), indicating the SVZ remains responsive to exogenous cues. Astrotypic NSCs (often referred to as ‘B’ cells) are believed to generate committed progeny via transit-amplifying cells (‘C’ cells), a rapidly dividing intermediate that becomes progressively restricted in developmental potential before terminally dividing into new neuroblasts (‘A’ cells, reviewed in Doetsch, 2003). Multiple identifying markers for this cell intermediate type exist (Doetsch et al. 1999 b; Aguirre et al. 2004). In practice however, separating NSCs and their progeny by morphometric, immunological, or functional methodologies in vivo has proven difficult. In vitro studies are frequently used to definitively identify and quantitate NSCs from the adult CNS. Isolated NSCs from the postnatal SVZ can be maintained in vitro in media containing epidermal and basic fibroblast growth factor (EGF and bFGF) (Gritti et al. 1996), and have the capability to self-renew and demonstrate multilineage potential through the formation of neurospheres: single cell-derived aggregations which self-renew and contain multiple cells types (Reynolds and Weiss, 1992). Furthermore, a number of protocols exist for inducibly differentiating NSCs exist for a variety of culture conditions (Lim et al. 1999; Skogh et al. 2001; Gritti et al. 2002). Using primarily in vitro approaches, limited observations on the dynamics of differentiation have been made. Phenotypic differentiation in vitro is accompanied by extensive expression changes in multiple genes (Gurok et al. 2004), and several markers have been identified to prospectively identify NSCs (Lendahl et al. 1990; Roy et al. 2000; Mignone et al. 2004; Imura et al. 2003), and have been used to monitor stem cell development in the CNS in vivo and in vitro.

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3 A recurrent theme in the investigation of postnatal NSCs and neurogenesis is the failure to correlate in vitro and in vivo findings. Most notably, no study has demonstrated a rigorous recapitulation of prescribed SVZ neurogenesis in vitro. As such, the dynamic transition between glial progenitors and mature neurons and/or glial cell types has not been directly observed. As the identity and developmental state of neurosphere-forming cells remains unclear, such a study would give new insights on the interpretation of existing in vitro investigation techniques, as current approaches cannot answer this question. Transit-amplifying ‘C’ cells have been described as being able to self-renew in vitro while maintaining intermediate cell mitotic kinetics (Doetsch et al. 2002). As subependymal astrocytes are reported to have insufficient mitotic kinetics to form neurospheres (Morshead et al. 1998; Doetsch et al. 2002), the identity of neurosphere-forming cells is indeed questionable. Adult Neural Stem/Progenitor Cells as a Therapeutic Cell Source Recent advances in developmentally regulating embryonic stem cells to a neuronal fate has prompted the attempt to utilize cultured embryonic stem (ES)-derived neural progenitors as a potential source for replacement tissue (Benninger et al. 2003), with the transplantation of ES-derived neuronal progenitors successfully treating animal models of neural disease (Kim et al. 2002). Though technically challenging, derivation of analogous donor populations from adult sources provides an equally attractive avenue for cellular replacement, and may subvert existing ethical and/or immunological concerns regarding ES cells. Existing protocols have been established for obtaining an enriched fraction of neuroblasts from cultured adult SVZ NSCs (Lim et al. 1999). Both in vivo transplantation and organotypic slice culture models allow for observation of integration,

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4 migration, and functional characterization of transplanted cells, and have been demonstrated for use in the engraftment of embryonic glial and neuronal progenitors in the hippocampus (Scheffler et al. 2003; Benninger et al. 2003). The maintenance of multipotent, self-renewing NSCs is a key prerequisite for derivation and growth of neural tissues for replacement strategies, as well as maintenance and regeneration of endogenous stem cell populations. Both human and rodent somatic cells are subjected to a maximal lifespan in culture. Human somatic cells are subjected to the telomere-imposed limit of approximately 50-60 population doublings (PDs) in culture (Hayflick, 1965). Cellular lifespans of rodent cells are regulated similarly by nontelomeric mechanisms. In vivo and in vitro, stem cells and cancerous cells exist as notable exceptions to conventional proliferative limits (reviewed in Wright and Shay, 2000). Indeed, rodent NSCs have been proliferated beyond conventional limits in vitro (Gobbel et al. 2003). Recent reports of culture conditions which allow expansion of nonpluripotent cells beyond traditional limits of expansion without immortalization suggest such upper barriers of proliferation may be more flexible than previously imagined (Mathon et al. 2001; Tang et al. 2001). It has been suggested that postnatal stem cells are maintained in vivo through asymmetric division involving nonrandom segregation of chromosomes, frequently suggested as a mechanism for preventing replication-associated damage (Potten et al. 2002). Cell cycle control genes have been implicated in the control of asymmetric division (Sherley et al. 1995), but the mechanism by which this nonrandom division is accomplished is unclear. Nonrandom segregation of genomic DNA has been implicated in NSC maintenance (reviewed in Sommer and Rao, 2002), providing two potential insights on the identification and apparent loss of NSCs

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5 throughout life. First, asymmetric division may lead to unequal labeling of dividing cells by thymidine analogs, a potentially problematic issue for in vivo identification of stem cells in the brain (Gould and Gross, 2002). Second, a failure to self-renew via symmetric division may account for the observed progressive reduction in NSCs throughout life (Tropepe et al. 1997). Populations of NSCs have been maintained in vitro without the predictable depletion of stem cells resulting from extensive passaging (Gobbel et al. 2003), but it remains unclear whether this phenomenon is the result of a reversion to symmetrical division or another undescribed mechanism. Finally, the effect of support cells on NSCs is a considerable factor to their function. Cell contacts have been implicated as being essentially for maintenance of the SVZ as a neuropoietic niche and as possibly supporting or directing differentiation (Doetsch et al. 1997; 1999 a, b). Nonneural cell types have also been reported in self-renewal of NSCs (Shen et al. 2004), transdifferentiation (Wurmser et al, 2004), and in cell fusion paradigms (Weimann et al. 2003). Within neuropoietic regions, astrocytes have been suggested as being essential in hippocampal (Song et al. 2002) and SVZ (Lim et al. 1999) neurogenesis, a curious observation in light of the fact neurons are formed before astrocytes in the developing brain. Based on these observations, it is increasingly clear that the cells and processes of postnatal neurogenesis are increasingly complex, and require innovative approach in their study.

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6 Fig. 1-1. Rostral migratory stream of the adult rodent CNS. Migratory PSA-NCAM + neuroblasts (stained for in lower left reconstruction) originating from subventricular astrocytes adjacent to the paired lateral ventricles migrate anteriorally to the olfactory bulb, where they tangentially migrate into the mitral cell layer (see diagram) and integrate as periglomerular or interneurons. Scale bar 500 m.

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7 CHAPTER 2 MATERIALS AND METHODS Supplier Information Abcam (Cambridge, MA), Acris (Hiddenhaus, Germany), Advanced Targeting Systems (San Diego, CA), Altamonte Labs (Jerusalem, Israel), Amersham (Piscataway, NJ), Axon Instruments (Union City, CA), Becton-Dickinson (San Jose, CA), Bio-Rad (Hercules, CA), Carl Zeiss Microimaging Inc (Thornwood, NY), Chemicon (Temecula, CA), Corning Inc (Corning, NY), DAKO (Carpintera, CA), Electron Microscopy Sciences (Hatfield, PA), Gibco (Grand Island, NY), Hyclone (Logan, UT), Invitrogen (Carlsbad, CA), Jackson Labs (West Grove, PA), Leica (Bannockburn, IL), Millipore (Billerica, MA), Molecular Devices (Sunnyvale, CA), Molecular Probes (Carlsbad, CA), Promega (Madison, WI), Qiagen (Valencia, CA), R&D Systems (Minneapolis, MN), Roche Diagnostics (Indianapolis, IN), Rockland (Gilbertsville, PA), Santa Cruz Biotechnologies (Santa Cruz, CA), Sigma Aldrich (St. Louis, MO), Sutter Instruments (Novato, CA), Upstate (Waltham, MA), Vector Labs (Burlingame, CA), Willco Wells BV (Amsterdam, The Netherlands). Strains of Mice The following strains of mice were used in this study: C57/B6 (The Jackson Laboratory, Bar Harbor, ME) -Actin-GFP (The Jackson Laboratory) NestineGFP (provided by Grigori Enikolopov, Cold Spring Harbor Laboratory, NY) Fox-Chase immunodeficient (NOD-SCID, Charles River Labs, Charles River, MA) Cell Lines These studies employ NIH derived and maintained H1299, RG2, and NCI HE57 and HUVEC cells.

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8 Reagent Preparation Aphidicolin -(Sigma) Stock solution was prepared in water at 10 mg/ml and stored at -20C for up to one year. Aphidicolin was applied at a final concentration of 1 ug/ml. Artificial cerebrospinal fluid (CSF) (all reagents obtained from Sigma) Artificial cerebrospinal fluid was composed of, in mM: 125 NaCl, 3 KCl, 26 NaHCO 3 , 1.25 NaH 2 PO 4 , 20 glucose, 1 MgCl 2 , and 2 CaCl 2 , prepared freshly for and maintained at 35 C during experiments. Avertin -(Sigma) Stock solution of Avertin (2-2-2 Tribromoethanol) was prepared in tert-amyl alcohol, sterilized using a 50 mL “Steriflip” disposable vacuum filtration system (0.22 ! m pore, Millipore), and stored in a 50 mL sterile polystyrene conical tube. Working solution was freshly prepared for each use at 20 mg/ml by diluting 0.6 ml of stock solution with 39.4 ml of phosphate buffered saline. NOTE: As avertin is toxic if allowed to photo-oxidize, the stock bottle was kept tightly capped and wrapped in aluminum foil. Stable at room temperature for well over one year. bFGF -(R&D Systems) Recombinant human basic fibroblast growth factor (bFGF, 25 ! g, R&D systems) was suspended in ice-cold 3 ml phosphate buffered saline (PBS) containing 0.003 g Fraction V heat shock Bovine Serum Albumin (BSA, Roche Diagnostics) and 3 ! l of 1M dithioerythritol (DTT, Sigma). Working solution was sterile-filtered using a 5 cc syringe w/ regular luer tip (Becton Dickinson) and sterile syringe filter (0.45 ! m pore size, Corning Inc). Aliquots of 100 ! L were prepared and stored at -80C. Final concentration 10 ng/ ! l, and used at 1:1,000.

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9 Brain Derived Neurotrophic Factor (BDNF) -(Invitrogen) Stock solution was prepared in water at a concentration of 1.0 mg/ml, and stored at C for a maximum of 2 weeks. Working dilution was 20 ng/ml. 5’Bromo-2-deoxyuridine (BrDU) –(Sigma) Stock solution was prepared by dissolving BrDU in Dulbecco’s Modified Eagle Medium with F12 supplements (DF) in a laminar flow hood as a sterile solution at a concentration of 2 mM. A working solution BrDU was diluted to 500 ! M in culture media, and pH was adjusted to 7.4. Solution was stored stably for 1 year at C. Working dilution ranged from 10 to 100 ! M. (Sigma, 10 M for cellular labeling not related to toxicity studies and 1-100 ! M for toxicity studies). Cyclosporin A -(Sigma) Stock solution was prepared at 50 mg/ml in dimethylsulfoxide (DMSO), and stored at C for 1 year. Working dilution was 1 ! g/ml. Cytosine ! D -Arabinofuranoside ( Ara C) -(Sigma) Stock solution was prepared in water at a concentration of 50 mg/ml and stored at 4C for a maximum of 1 month. Working dilution was 0.1 ! m. 4',6-Diamidino-2-phenylindole dihydrochloride (DAPI) -(Sigma) Stock solution was created at 0.5 g/ml in glycerol, and applied during the mounting process of sections or slides.

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10 Dibutyl cyclic AMP -(Sigma) Stock solution was prepared in ddH 2 O at a concentration of 100 mg/ml. Stable at 4C for 1 month. Working dilution was 0.5 mM. EGF -(R & D Systems) Stock solution of recombinant human epidermal growth factor (EGF, 200 ! g) was dissolved in 30 ml of ddH 2 O containing 17.24 ! l of glacial acetic acid and a final concentration of 0.01% of fraction V heat shock BSA. Solution was sterile-filtered and stored in 500 ! l aliquots at C. Final concentration was 20 ng/ ! l, diluting stock solution 1:1,000 in culture media. Epigallocatechin-3-gallate (EGCG) -(Sigma) Stock solution was prepared in water at a concentration of 500 ! M. Working dilution was 10 ! M. FGF-8 -(Sigma) Stock solution was prepared in PBS containing 0.1% fraction V heat shock BSA at a concentration of 50 ! g/ml and stored at C for a maximum of 1 month. Working dilution was 1 ng/ml. Hoechst 33528(Sigma) Stock solution was prepared in PBS at 10 mg/ml. Working dilution was 1 mg/ml. Immunocytochemistry Hybridization Buffer -Sterile working solution was composed of PBS containing 10% fetal calf serum (FCS, Hyclone), 5% normal goat serum (NGS, Hyclone), and 0.1% triton X-100 (Sigma). 50 ml aliquots were prepared and stored at 4C. Stable for 1 week. 3-Isobutyl-1-methylxanthine ( IBMX) -(Sigma) Stock solutions were dissolved in 100% methanol at a final concentration of 50 mg/ml, and were stable at 4C for 3 months. Working concentration was 0.05 mM.

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11 Laminin -(Sigma) Stock solution was created in water at a concentration of 5 mg/ml and stored at C for up to 6 months. Working dilution was 1 g/ml. Mitomycin C -(Sigma) Stock solution was prepared in water and was stored at 4C for a maximum of 1 month. Mitomycin C was applied at a final concentration of 10 ! g/ml. MitoSox Superoxide Indicator Dye -(Molecular Probes) Stock solution was dissolved in DMSO and prepared at a concentration of 5 mM. Stock solution was stored for up to one week. Working dilution was 5 M in Hank’s Balanced Salt Solution (Sigma). NSC Differentiation Media (Inducible Differentiation Media) -Differentiation media was composed of DF (Invitrogen) containing standard N2 supplements (sodium selenite, putrescine, insulin, progesterone, transferrin), 35 g/ml bovine pituitary extract, and antibiotic/antimycotic cocktail (1 ml per 100 ml total media volume, Sigma). NSC Differentiation media was stored at 4C for up to 2 weeks. NSC Proliferation Culture Media (Defined Proliferative Media) -Defined proliferative media was composed of (DF, Invitrogen) containing N2 supplements (sodium selenite, putrescine, insulin, progesterone, transferrin), 35 g/ml bovine pituitary extract, antibiotic/antibiotic cocktail (1 ml per 100 ml total media volume, Invitrogen), 5% FCS (HyClone), and 40 ng/ml EGF) and bFGF (R&D Systems). NSC proliferative media was stored at 4C for up to 2 weeks.

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12 Nerve Growth Factor (NGF) -(Sigma) NGF (2.5S isoform derived from submaxillary gland) was prepared in phosphate buffered saline and stored at 10 mg/ml. Working dilution was 0.5 mM. Neurosphere Proliferation Media -Composed of defined proliferative media containing 1% methylcellulose (Sigma). Media was stored at 4C for up to 2 weeks. 4% Paraformaldehyde -(Sigma) Paraformaldehyde (4% w/v, Sigma) was mixed with 60 ml of distilled water heated to approximately 55C in a glass beaker and covered with aluminum foil while mixing for 10 minutes. Two drops of 1N NaOH was added to the solution followed by an additional 5 minutes of mixing, after which the solution became semi-clear. 30 mL of 3x PBS was added to the solution and the pH adjusted to 7.2 using 1N HCl and 1N NaOH. The solution was then brought to a final volume of 100mL with ddH 2 0, filtered through Whatman paper into clean glass bottle and stored at 4C. Solution is stable for approximately 1 week. Picrotoxin -(Sigma) Picrotoxin was added to 0.9% NaCl in ddH 2 O for each experiment. W orking concentration was 50 M. Pleiotrophin -(Sigma) Stock solution was prepared in phosphate buffered saline at a concentration of 1 mg/ml, and stored at C for up to one month. Working dilution was 5 ! g/ml. Poly-L-Lysine -(Sigma) Stock solution was prepared in a laminar flow hood in PBS at a concentration of 1 mg/ml and stored at C for up to 6 months. Polyornithine -(Sigma) Stock solution was prepared in ddH 2 O at a concentration of 1 mg/ml. Stock solution was diluted in ddH 2 O to form a working solution with a

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13 dilution of 10 g/ml. Propidium Iodide -(Sigma) Working solution was prepared in PBS at a dilution of 50 ! g/ml. Retinoic Acid -(Sigma) Stock solution was prepared in ethanol at a concentration of 2.7 mg/ml, and was stored at C for a maximum of 2 weeks. Working dilution was 0.5 ! M. RIPA Buffer -RIPA buffer contains (in mM): 150 NaCl, 50 EDTA (pH 7.5), 50 sodium ! -glycerophosphate, 50 NaF, 5 sodium pyrophosphate, 2 EDTA, 2 EGTA, 1 DTT, 1 phenylmethylsulfonyl fluoride, 1 sodium orthovanadate with 1% Triton X-100, 10 ! g/ml leupeptin, and 10 ! g/ml aprotinin (Sigma) in ddH 2 O. Senescence-Associated Beta-Galactosidase (SA--Gal) Solution-SA! -Gal solution contains 1 mg/ml 5-bromo-4-chloro-3-indolyl ! -DGalactosidase (X-Gal, Sigma), 20 g/ml dimethylforamide (Sigma), and (in mM) 150 NaCl, 40 citric acid/sodium phosphate (pH 6.0), 5 potassium ferrocyanide, 2 MgCl 2 (Sigma) in ddH 2 O. Sonic Hedgehog -(Sigma) Stock solution was prepared in PBS containing 0.1% BSA at a concentration of 1 mg/ml and stored at C for a maximum of 1 month. Working dilution was 5 ! g/ml. Tetraethylammonium (TEA) -(Sigma) Stock solution was prepared in ddH 2 O at a concentration of 50 mg/ml and was stored at C for a maximum of 1 month. Working concentration was 200 mM. Tetrodotoxin (TTX) -(Alomone Labs) Working concentration (400 nM) added from powder to PBS and used immediately.

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14 Trypsin -(Sigma) Trypsin was reconstituted Versene and TD (2:1 ratio) from powdered form to a final concentration of 0.25% (vol/vol), and established at pH 7.3. Trypsin was stored at 4C and was stable for 1 month. Western Blot Hybridizing Buffer -Hybridizing buffer was composed of trisbuffered saline (TBST) [20 m M Tris-HCl (pH 7.5), 500 m M sodium chloride, and 0.05% Tween-20, obtained from Sigma] containing 5% nonfat dried milk (Invitrogen), and was stored at room temperature for up to 7 days. Xanthosine (XS) -(Sigma) Stock solution was reconstituted in sterile phosphate buffered saline at a concentration of 100 mg/ml and stored at 4C for up to three months. Working dilution was 10 ! M. Antibody List Primary Antibodies Immunocytochemistry A2B5 (recombinant A2B5-105, 1:500, Chemicon) ! -III-tubulin/Tuj1 (mouse monoclonal, 1:300, Promega, Madison, WI; rabbit polyclonal, 1:500, Covance, Denver, PA) BrDU (mouse monoclonal, 1:50, BD Biosciences, San Jose, CA) CD11b (rat monoclonal, BD Biosciences, 1:500) CD15 (mouse monoclonal, 1:300, Abcam) CD45 (mouse monoclonal, 1:400, Chemicon) CD133/Prominin-1 (rabbit polyclonal, 1:600, Abcam)

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15 CNPase (mouse monoclonal, 1:250, Chemicon, Temecula, CA) dlx-2 (goat polyclonal, Santa Cruz Biotechnology, 1:50) doublecortin (goat polyclonal, 1:200, Chemicon) eGFP (rabbit polyclonal, Chemicon, 1:500) EGFR (sheep polyclonal, 1:500, Upstate) FGFR1 (rabbit polyclonal, 1:500, GeneTex) Glutamic acid decarboxylase (GAD) 65/67 (rabbit polyclonal, Santa Cruz Biotechnology, 1:125) Glial-derived neurotrophic factor (GDNF) (rabbit polyclonal, 1:500, Chemicon) Glial fibrillary acidic protein (GFAP) (rabbit polyclonal, 1:600, DAKO, Carpinteria, CA) glutamine synthetase (rabbit polyclonal, 1:100, Abcam, Cambridge, MA) human ribonuclear protein (HNA, mouse monoclonal, 1:300, Acris, Hiddenhausen, Germany) Ki-67 (mouse monoclonal, 1:300, BD Biosciences) map2a-c (chicken polyclonal, 1:30,000, gift from Dr. Gerry Shaw) nestin (mouse monoclonal, 1:50, Chemicon) nestin (human specific, mouse monoclonal, 1:300, Chemicon) NeuN (mouse monoclonal, 1:500, Chemicon) NG2 (rabbit polyclonal, 1:1,000, Chemicon) neurofilament L (mouse monoclonal, 1:300, Chemicon) neurofilament M (mouse monoclonal, 1:500, gift from Dr. Gerry Shaw) O4 (mouse monoclonal IgM, 1:150, Chemicon) PDGFR (mouse monoclonal IgG, Abcam, 1:600) PSA-NCAM (mouse monoclonal IgM, Chemicon, 1:400)

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16 S100! (rabbit polyclonal, 1:100, Swant, Bellinzona, Switzerland) Sox1 (chicken polyclonal, 1:300, Chemicon) Sox2 (goat polyclonal, 1:500, R&D Systems) Sox3 (rabbit polyclonal, 1:300, Chemicon) hTERT (rabbit polyclonal, 1:200, Santa Cruz Biotechnology, Santa Cruz, CA) tyrosine hydroxylase (rabbit polyclonal, 1:1,000, Sigma) vimentin (mouse monoclonal, 1:500, Chemicon) Western Blot cyclin A (rabbit " human, 1:200, Santa Cruz) cyclin D1 (mouse " human, 1:2000, Santa Cruz) cyclin E (rabbit " human, 1:200, Santa Cruz) p16 (rabbit " human, 1:200, Santa Cruz) p21 (rabbit " human, 1:200, Santa Cruz) p53 (mouse " human, 1:500, Santa Cruz) hTERT (1:200, rabbit " human, Santa Cruz) Immunoprecipitation Cystatin C (rabbit polyclonal, 0.2 g/ml, Upstate) Secondary Antibodies Immunocytochemistry Alexa-555 goat anti chicken (1:300, Molecular Probes, Carlsbad, CA) AMCA goat " rabbit IgG (Jackson Labs, 1:50) Cy3 goat anti mouse IgG (1:300, Jackson Labs, West Grove, PA),

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17 Cy3 goat anti mouse IgM (1:600, Jackson Labs) Oregon Green donkey anti goat (Molecular Probes, 1:200) Oregon Green goat anti mouse (Molecular Probes, 1:1000) Oregon Green goat anti rabbit (1:600, Molecular Probes) Western Blot Donkey anti rabbit (1:10,000, Amersham) Donkey anti mouse (1:5000, Amersham) Methods Isolation, Derivation, and Expansion of Rodent NSCs In Vitro Regional microdissection of neuropoietic cells To obtain SVZ cells, 8-day-old and adult C57/B6, -actin-eGFP, and nestin-eGFP mice were deeply perfused with avertin and decapitated. Brains were removed and placed in ice-cold DF containing 20 mg/ml penicillin, 20 mg/ml streptomycin, and 25 ng/ml amphotericin B (collectively abx), and the paired lateral ventricles were exposed via coronal sectioning. Periventricular tissue extending 1 mm laterally from the ventricular wall was microdissected under a surgical microscope. Comparable volumes of parietal neocortex and cerebellar tissue were similarly gathered as non-neuropoietic control tissues.

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18 Isolation and expansion of primary subventricular dissociates Under aseptic conditions, ageand sex-matched tissue ( # 5 animals, for C57/B6 mice) was pooled, placed in PBS, and manually dissociated into 1 mm 3 pieces and differentiated as described (Scheffler et al, 2005). Briefly, tissue was enzymatically digested in 0.25% trypsin (15 min, 37 C, pH 7.3) and cultured overnight in uncoated T75 plastic tissue culture dishes (Costar) in defined proliferative media (described in Reagents). Unattached cells were collected and triturated using bore-restricted pipetting, replated onto uncoated plastic dishes (TPP) and proliferated to confluency in defined proliferative media supplemented bidaily with 20 ng of EGF and FGF. Cells were maintained in these conditions and split 1:2 when confluent. Live cells were counted in triplicate using trypan dye exclusion. Under these conditions, SVZ-derived monolayers from C57/B6 mice and nestin-eGFP mice were maintained >330 population doublings (PDs). Confluent monolayers were frozen in aliquots of 1 million cells and maintained in liquid nitrogen for future use. To induce differentiation in primary dissociates, cells were plated on glass coverslips coated with laminin (Sigma, 5 g/ml) and polyornithine (Sigma, 10 g/ml) (LPO) at a minimum density of 20,000 cells/cm 2 . Cells were induced to differentiate by removal of EGF, bFGF and serum from culture media. Proliferating cells were labelled with BrDU (Sigma, 10 M) administered before and after differentiation. Xanthosine (XS, 10 M) was added to identically cultured proliferating cultures immediately following initial plating in proliferative medium, and was supplemented bidaily. Aphidicolin (1 g/ml) was added to proliferating cell cultures at 5 and 300 PDs. Nestin-eGFP ( Mignone et al. 2004) and ! -actin-eGFP transgenic animals were decapitated, their brains placed overnight in ice-cold DF containing antibiotics, and

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19 processed identically. All procedures were performed in accordance with institutional IACUC guidelines. Soluble factor and coculture experiments for rescue of neurogenesis were performed on C57/B6 SVZ cells that had undergone > 300 PDs. Cells were plated onto LPO-coated coverslips at a density of 2x10 5 cells/cm 2 in a 1:1 ratio with identically isolated cerebellar cells from 3 PD -actin-eGFP-expressing mice. Cells were allowed to attach for 48 hours in defined proliferative media, then were differentiated as described. Cells were evaluated for neuroblast production 3, 7, and 10 days later. Cells were plated on LPO-coverslips in defined proliferative medium supplemented with 50% (v/v) conditioned medium from passage 3 ( $ 3 PDs) SVZ or cerebellar-derived astrocytes from 8-day-old -actin-eGFP mice. Cellular contamination (GFP + cells) was evaluated 3d following differentiation. Cells were proliferated to confluency (3 days) and differentiated by removing conditioned medium, serum, and growth factors. Primary and expanded cells were immunophenotyped while proliferating, and 3 and 7 days following induction of differentiation. For experimental comparison of adult and early postnatal animals, postnatal day 90 -actin-eGFP mice (n=4) were sacrificed, and their SVZ and cerebellum processed identically. Neurosphere (NS) generation, differentiation, and quantification Passage 5 ( " 5 PDs) cells from P8 and adult SVZ were trypsinized, counted, and re-suspended in nonadhesive 6-well plates (Corning, NY) in 2 ml/well of N5 media containing 1% methylcellulose as described (Kukekov et al. 1999; Laywell et al. 2000). To verify clonality of NS generated under these conditions, serial dilution of SVZ cells was performed from 0.6-20 x10 3 cells/cm 2 . A linear seeding/NS relationship was

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20 observed between densities of 2.5-20 x10 3 cells/cm 2 (data not shown). All NS experiments were performed at a seeding density of 10 4 cells/cm 2 . Polyclonal aggregation spheres were generated at a density of 1 x10 7 cells/cm 2 in otherwise identical conditions in defined proliferative medium lacking methylcellulose. To quantify presence of sphere-forming cells, total numbers of primary NS were counted per well using bright field microscopy (n=3 for each, adult and P8 cultures, derived from 2 independent experiments). Secondary NS were derived from dissociated primary NS and evaluated similarly (n=2 for each, adult and P8 cultures, from 2 independent experiments). NS were evaluated at 14-21 days after cell seeding. The level of statistical significance was set at p <0.05, and was calculated using the student’s t -test or one-way ANOVA. NS were plated on LPO-coated glass coverslips overnight in defined proliferative media lacking EGF and bFGF. Cells migrating out of NS were allowed to differentiate for 7 days after removing growth factors and serum from culture media. Individual primary and secondary NS were assessed for multipotentiality by doubleimmunofluorescence analysis with neuronal ( ! -IIItubulin, Map2, NeuN) and glial (CNPase, GFAP, O4) marker proteins. Neurosphere diameter was measured in a minimum of 25 randomly selected spheres per experimental condition using Spot Axiovision software. All values expressed as mean + S.E.M. Inducible differentiation of rodent SVZ-derived cells To induce differentiation, cells were plated on glass coverslips coated with LPO or poly-L-lysine at densities of approximately 2x10 5 cells/cm 2 . Cells were proliferated to 90-100% confluency and were induced to differentiate by removing growth factors and

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21 serum from culture media. Differentiating progeny were labelled with BrDU (Sigma, 10 M) administered 0-72 hours following differentiation. Tissue sectioning and storage Animals were deeply anesthetized with avertin, sacrificed by cutting the descending aorta, and were perfused with 20 ml of ice-cold 4% paraformaldehyde. Perfused brains were removed and placed in 2% paraformaldehyde containing sucrose (30% vol/vol) overnight at 4C. Preserved brains were mounted and sectioned into 15 ! m sections in a coronal or sagittal plane on a freezing microtome. Sections were serially stored at C. Antibody application Cells plated on coverslips coated with poly-L-lysine or laminin and polyornithine (LPO) were fixed with 4% paraformaldehyde (15 min, 25 C, Sigma). After washing with PBS, cells were blocked 20 min (attached cells) or 2 hours (tissue sections) in PBS containing 10% FCS (Hyclone), 5% NGS (Sigma), and 0.1% Triton X-100 (Sigma). Primary antibodies were applied for either 1 hour at 25 C or overnight at 4 C in PBS containing 10% FCS and 0.1% Triton X-100. Secondary antibodies were applied for 1 hr at 25 C in PBS containing 10% FCS and 0.1% Triton X-100. For BrDU imaging, cells were incubated in sodium chloride/sodium citrate (SSC)-formamide (1:1, 37 C, 2 hr), washed 3 x 10 min in SSC, incubation in 2N HCl (37 C, 30 min), and washed with 0.1 M borate buffer (25 C, 10 min). For immunoperoxidase detection of antigen, tissue sections

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22 were pretreated with 1% H 2 O 2 in 70% methanol (15 min, 25 C) and visualized using an ABC Elite detection kit (Vector Labs, Burlingame, CA). Nuclear labeling and lipophilic dye application Nuclei were stained by application of either DAPI (1 g/ml, 25 C, 10 min, Sigma) or propidium iodide (50 g/ml, 25 C, 10 min, Sigma) prior to mounting. Lipophilic dye addition was performed with FM-143 (Molecular Probes). Adherent cells were placed in ice-cold Hank’s Balanced Salt Solution containing 5 g/ml lipophilic dye for 1 minute. Dyed cells were mounted and immediately examined. Electrophysiology Adherent cells were placed in a holding chamber continuously perfused with oxygenated artificial cerebrospinal fluid containing, in mM: 125 NaCl, 3 KCl, 26 NaHCO 3 , 1.25 NaH 2 PO 4 , 20 glucose, 1 MgCl 2 , and 2 CaCl 2 and maintained at 35 C during experiments. The intracellular pipette solution comprised of, in mM: 145 Kgluconate, 10 HEPES, 10 EGTA, and 5 MgATP (pH 7.2, osmolarity 290). Cells were visualized using Axioskop-FS DIC microscope ( Zeiss, Thornwood, NY). Patch electrodes were pulled from borosilicate capillary glass using a Flaming-Brown P-87 microelectrode puller (Sutter Instruments, Novato, CA) and had a resistance of 4-6 M % when filled with internal solution comprising of (in mM): 130 Kgluconate, 10 HEPES, 0.2 EGTA, 2 ATP and 0.3 GTP (pH 7.2, osmolarity 290). For experiments in which post-synaptic currents were recorded, 145 K-gluconate was replaced with 125 KCl and 20 K-gluconate. Recordings were performed with an Axopatch-1D (Axon Instruments,

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23 Union City, CA) and filtered at 5 kHz. Clampex 8.2 (Axon Instruments) was used to deliver command potentials and for data collection. Series resistances were < 20 M % and checked frequently to ensure that they did not deviate. For voltage-clamp experiments, a step protocol was applied which held the membrane at potentials between mV and +60 mV for 50 ms after a pre-pulse period of 200 ms at mV. During current-clamp experiments a step protocol was utilized in which currents between 10-100 pA were applied per step. Clampfit 8.2 (Axon Instruments) was used to analyze voltage and current traces. For recordings of human progenitors and their progeny, series resistances were 10-20 M % and recordings were discarded if a change of series resistances was > 10%. Cells were held at -65 mV. The values of capacitance and input resistance were obtained by applying 10 mV voltage pulse to cells. Picrotoxin (PIC) was applied at a concentration of 50 M, TEA at 20mM, and TTX at 400nM (Alomone Labs). Chemicals and reagents were obtained from Sigma unless otherwise noted. Data are expressed as mean S.E.M. Documentation In vitro images of cultured cells were captured using a Nikon Eclipse TS-100 bright field microscope and a Spot 3.1 digital camera (Diagnostic Instruments, Sterling Heights, MI). Fluorescence microscopy was performed on a Leica DMLB upright microscope (Leica) and images were captured with a Spot RT color CCD camera (Diagnostic Instruments). Confocal microscopy was performed on an Olympus IX-70 microscope (Melville, NY) using Confocal 1024 ES software (BioRad, Hercules, CA).

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24 Live-cell microscopy Passage 3 SVZ cells were grown to confluency in N5 media on LPO-coated 3 cm glass coverslip dishes ( Willco Wells BV, Amsterdam, The Netherlands). Cells were induced to differentiate as described, and were monitored under standard culture conditions (37 C, 5% humidified CO 2 ) on a Zeiss Cell Observer system (Carl Zeiss Microimaging Inc., Thornwood, NY). Five randomized visual fields (20X) were selected for analysis 24 hours following induction of differentiation. Phase images were taken every five minutes for up to 72 hours. Images were compiled into movies using Axiovision software (Zeiss). Unbiased cell counting For quantification of adherent cells, a minimum of twelve randomized fields were selected at 40X magnification and counted per experimental condition. For quantification in vivo, serial coronal or saggital sections were created, and every sixth section quantified for a minimum of 12 randomized fields at 40X magnification per section per experimental condition. For evaluation of SA--Gal, each subventricular region (lateral periventricular tissue extending 100 ! m lateral to the lateral wall of the paired ventricle) was evaluated in each section examined (a minimum of 12 sections per condition, for a minimum of 2 independent experiments). For sections containing hippocampus, senescent cells within the cytoarchitecture were quantified into the following regions: dorsal and ventral blades (constrained to the subgranular zone on both defined blades) and hilus (defined by the subgranular blades of the dorsal and ventral

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25 blades and laterally defined by the incoming projection of CA3). A minimum of 12 hippocampi from 2 independent experiments were evaluated per condition. Electron microscopy Passage 3 SVZ cells were grown in N5 media on LPO-coated aclar coverslips. Fixation and processing was standard. All reagents were obtained from Electron Microscopy Sciences. Samples were visualized on a Leica EM10A transmission electron microscope at magnifications between 1-16,000X. Images were captured using a CCD digital camera (Finger Lakes Instrumentation, Lima, NY). Western blotting and semi-quantitative protein analysis Cells were lysed in a modified RIPA buffer containing (in mM): 150 NaCl, 50 EDTA (pH 7.5), 50 sodium ! -glycerophosphate, 50 NaF, 5 sodium pyrophosphate, 2 EDTA, 2 EGTA, 1 DTT, 1 phenylmethylsulfonyl fluoride, 1 sodium orthovanadate with 1% Triton X-100, 10 g/ml leupeptin, and 10 g/ml aprotinin (Sigma). Equal amounts of lysates were resolved on a 12% SDS-polyacrylamide gel and transferred to a nitrocellulose membrane. The membrane was blocked in TBST [20 m M Tris-HCl (pH 7.5), 500 m M sodium chloride, and 0.05% Tween-20] containing 5% nonfat dry milk for 2 hours and then incubated with primary antibodies in TBST containing 1% BSA at room temperature for 2 hours. Horseradish peroxidase-labeled secondary antibodies were applied in TBST containing 5% nonfat dry milk for 2 hours. Protein was visualized by using an enhanced chemiluminescence (ECL) detection system (Amersham).

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26 Semiquantitative was performed using fixed-area denitometry comparison using ImageJ (NIH). FACS studies on cell cycle/ploidy Fluorescent-activated cells sorting (FACS) analysis was performed on both proliferating human progenitor populations and neuropoietic and non-neuropoietic rodent neural tissues under conditions favoring differentiation and proliferation. Adherent monolayers were trypsinized as described, resuspended in 5 ml of 0.9% NaCl, and fixed with the addition of 5 ml of ice-cold 90% ethanol. Cells were collected via centrifugation at 400 xg, and stored as pellets at 4C for up to 1 week. For experiments evaluating cell cycle and ploidy analysis, pelleted cells were resuspended in 50 mg/ml propidium iodide containing 1 mg/ml RNAse I for 30 minutes. Cells were gated for forward and side scatter, and were evaluated for overall DNA content and cell cycle analysis. A minimum of 10 5 cells from > 3 independently derived samples were analyzed per experimental condition. For these studies, a FACScan flow cytometer (BD Biosciences, San Jose, CA) was employed. Analysis of data was performed using Cell Quest data analysis software (BD Biosciences). Clonal seeding experiments in adherent conditions Adherent monolayers were dissociated with trypsin as described, and attached overnight to LPO-coated coverslips in defined proliferative medium at a density of 12.5

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27 cells/cm 2 . Cells were differentiated as described, and evaluated for proliferative activity and generation of committed cell types 3 and 7 days later. In vivo BrDU administration Young (postnatal day 8) C57/B6 mice housed under identical conditions were anesthetized with halothane, restrained, and injected intraperitoneally with saline or 250 mg/kg BrDU. BrDU was administered every other day for 6 days. Groups received 0, 1, or 3 injections of BrDU (n=3 per group), with animals receiving one injection receiving 2 saline injections. 24 hours following the final injection, animals were sacrificed and transcardially perfused, and their brains were removed and processed as described. Cell growth experiments Primary SVZ astrocytes and RG2 cells were plated at a density of 2x10 4 cells/cm 2 and proliferated in defined proliferative media (astrocytes) or DF containing 10% FCS. 0, 1, 10, 50, or 100 ! M BrDU was added to proliferating cultures, either constitutively, or in 24-hour intervals. Confluent cells layers were split 1:2 or 1:4. Cell number was quantified as described, and expressed as doublings/day. Statistical significance was determined using a student’s t test or one-way ANOVA with Bonferroni post hoc analysis. Apoptosis/necrosis detection Apoptotic cells were fluorescently labeled using TdT-mediated dUTP nick-end labeling (TUNEL) using a protocol adapted from the DeadEnd Fluorometric TUNEL

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28 System (Promega). Briefly, adherent cells were fixed using 4% paraformaldehyde (15 min, room temp). Sections were washed 3 x 5 minutes with PBS containing 0.1% triton X-100. Cells were suspended in equilibration buffer for 10 minutes, and treated with rTdT incubation buffer (equilibration buffer: nucleotide mix: rTdT enzyme=45:5:1) for 60 min at 37C. Negative controls were prepared without rTdT enzyme. Reactions were quenched in excess 2X SSC for 15 min and coverslipped with DAPI. TUNEL + apoptotic cells were expressed as a percentage= TUNEL + /DAPI + . Necrotic cells were identified using propidium dye exclusion criteria. Cultured cells were placed in 50 mg/ml propidium iodide in Hank’s Balanced Salt Solution (37C, 10 minutes). Cells were mounted in warm mounting media containing DAPI and examined within 30 minutes. Necrotic cells taking up propidium iodide were expressed as a percentage= PI + /DAPI + . MitoSox dye addition Mitosox dye was applied to proliferating cells cultured for 24 hours in the presence of a 0-100 M BrDU dose supplemented to culture medium. Proliferating cells were removed from culture media and placed in 1 ml of 5 M MitoSox dye in Hank’s Balanced Salt Solution for 10 minutes. Cells were washed three times with culture media, mounted, and examined within 30 minutes. Senescence-associated -galactosidase stain SA! -Gal expression was assessed 7 days after addition of growth arrestors as described (Dimri et al. 1995). Briefly, cells were fixed in PBS containing 2%

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29 formaldehyde and 0.2% glutaraldehyde (25 C, 15 min). Following wash in PBS, cells were incubated (37 C, 12 hr) with fresh SA! -Gal solution. Rescue of endogenous neurogenesis Soluble factor analysis was performed on C57/B6 SVZ cells that had undergone > 300 PDs. Cells were plated on LPO-coverslips in defined proliferative medium supplemented with 50% (v/v) conditioned medium from passage 3 ( $ 3 PDs) SVZ or cerebellar-derived astrocytes from 8-day-old actin-eGFP mice. Cellular contamination (GFP + cells) was evaluated 3 days following differentiation. Cells were proliferated to confluency (3 days) and differentiated by removing conditioned medium, serum, and growth factors. Primary and expanded cells were immunophenotyped while proliferating, and 3 and 7 days following induction of differentiation. Microglial depletion Saporin-conjugated CD11b/MAC1 antibody or unconjugated saporin (10 pg/ml, Advanced Targeting Systems) was added to matched cultures of PD300 SVZ dissociates for 4, 18, and 24 hours (37C, 5% humidified CO 2 ) in defined proliferative media. Cells were washed with sterile PBS and either fixed with 4% paraformaldehyde (15 min, 25C) or induced to differentiate by removal of growth factors and serum and evaluated four days later.

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30 Immunoprecipitation studies 3 and 300 PD cells were grown for 3 days in 15 ml of defined proliferative medium. Conditioned medium was removed and centrifuged at 1,000xg for 5 min. 150 l protein A slurry beads were added, and cells were incubated 30 min. at 4 C. Cells were centrifuged at 1,000xg for 15 minutes, and 5 g anti-cystatin C antibody (rabbit polyclonal, Abcam) was added (90 minutes, 25 C), followed by 150 l protein A slurry (1 hour, 4 C). Protein A beads were collected by centrigugation, washed twice with PBS, and combined with an equal volume of Laemmli buffer. Samples were heated to 90 degrees for 5 min, centrifuged at 10,000xg (5 min). Equal amounts of supernatant were resolved on a 12% SDS-polyacrylamide gel and transferred to a nitrocellulose membrane. The membrane was blocked in TBST [20 m M Tris-HCl (pH 7.5), 500 m M sodium chloride, and 0.05% Tween-20] containing 5% nonfat dry milk for 2 hours and then incubated with anti-cystatin C (rabbit polyclonal, 1:1,000, Abcam) in TBST containing 1% BSA at room temperature for 2 hours. Horseradish peroxidase-labeled secondary antibodies (donkey " rabbit, 1:10,000, Amersham) were applied in TBST containing 5% nonfat dry milk for 2 hours. Protein was visualized by using an ECL detection system (Amersham). Human neural tissue collection and culture Primary tissue was gathered from individuals undergoing surgery related to medically intractable temporal lobe epilepsy. Primary tissue was removed and stored overnight in ice-cold DF (Gibco) medium containing 20 mg/ml penicillin, 20 mg/ml streptomycin, and 25 ng/ml amphotericin B (collectively abx, Invitrogen). Hippocampus

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31 (containing hilus) and temporal cortex (not containing subventricular zone and substantia nigra) were microdissected from biopsied tissue samples (n=5, data presented from 17 year-old female and 34 and 78 year-old males). Neural cell collection from patients undergoing deep brain stimulation was performed in accordance with University of Florida Institutional Review Board, as specified in protocol 17-2006. All procedures were performed with informed consent and were performed in accordance with human tissue handling and use guidelines. Dissected tissues were placed in 1X PBS (pH 7.3) lacking CaCl 2 or MgCl 2 , and were manually dissociated into 1mm 3 pieces under sterile conditions. Tissues were collected and resuspended in 0.25% trypsin (15 min, 37 C, pH 7.3, Sigma), and were further triturated using restricted bore pipetting. Cells were collected, resuspended in proliferative media, and seeded onto uncoated T75 culture flasks overnight (12 hr, 37 C, 5% humidified CO 2 ). Unattached cells were collected and seeded onto uncoated 60 mm plastic dishes in proliferative media. Proliferative media was comprised of DF containing N2 supplements, 35 g/ml bovine pituitary extract (Sigma), abx, 5% FCS ( Hyclone), and 40 ng/ml of EGF and bFGF (R&D Systems). 20 ng of EGF and FGF were supplemented bidaily. When necessary, media was changed every fourth day. A total of 7 lines from 5 patients were gathered. Cells were frozen in aliquots of 1 million cells in DF containing 10% FCS and 20% dimethyl sulfoxide (v/v, Sigma). Cells were passaged 1:2 when confluent. Cells were dissociated with 0.25% trypsin, counted (using trypan dye exclusion as viability criteria), and were replated onto uncoated 60 mm plastic dishes (Sigma). For growth monitoring experiments, 1 g/ml aphidicolin or 20 M EGCG (Sigma) were added to culture media 1 hour after plating. Irradiated cells were treated with a single 3 Gy dose of X-irradiation. Following period

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32 of application, cells were washed in 1X PBS and were resuspended in proliferative media or fixed. To assess viability of cells in various growth conditions, equal numbers of culture-matched cells were seeded into triplicate wells of various proliferative conditions, and counted 7, 14, and 21 days later. For growth factor analysis, EGF and/or bFGF were removed for seven days, and surviving cells were returned to proliferative media. Significance (p<0.05) was calculated using a student’s t -test. Cellular irradiation Cultured cells were trypsinized in 0.25% trypsin and suspended in PBS. Cells were exposed to 3 Gy total dose of X-irradiation. Karyotyping All cytogenetic analysis was done by the University of Florida core facility for cytogenetics. Briefly, confluent cell layers were incubated with 300 L Karyomax (Gibco), dissociated with 0.25% trypsin and resuspended in 75 mM KCl for 6 minutes. Cells were collected and resuspended in 3:1 (vol/vol) ethanol:acetic acid. Cells were visualized on a coverslip using light microscopy. Generation of human neuronal cell types To differentiate adherent cells serum, EGF and bFGF were removed from the culture media and supplemented with 0.5 mM 3-isobutyl-1-methylxanthine (IBMX), 0.5 mM 1-dibutyryl cAMP, and 25 ng/ml NGF ( Ronnett et al. 1990). Media supplements were replaced every third day. FGF-8 (100 ng/ml), sonic hedgehog peptide (500 ng/ml),

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33 pleiotrophin (100 ng/ml), and retinoic acid (0.5 M) were purchased from Sigma. Matched differentiating cells were cocultured with 10 m BrDU for 2 days following factor addition. Cells were immunocytochemically evaluated 2, 3, 5 and 7 days later, and were electrophysiologically evaluated 7 days following differentiation. To dedifferentiate adherent cells serum, EGF and bFGF were removed from the culture media and a subset supplemented with 0.5 mM IBMX, 0.5 mM 1-dibutyryl cAMP, and 25 ng/ml NGF (Ronnett et al., 1990) . BrDU (10 M) was added to age-, region-, and culture-matched cells for 2 days following factor addition. Cells were immunocytochemically evaluated for fate choice 2, 3, 5 and 7 days later, and were electrophysiologically evaluated 7 days following differentiation. Transplantation of human progenitors Cultured progenitors (30 PDs) were trypsinzed and resuspended at a density of 10 5 cells in 2 ! l PBS. Cells were injected into the lateral ventricle or cortex of anesthetized postnatal day 3 C57/B6 mice (n=6) or immunocompromised adult NODSCID mice (n=3, Taconic, Hudson, NY) at the following stereotactic coordinates: bregma, .06 mm; interaural +2.74 mm; 1 mm left of midline to a depth of 1 (cortical, NOD-SCID) or 2 (ventricular, C57/B6) mm. Transplantation coordinates for human progenitor injection into the substantia nigra were 4 mm anterior to the lambdoid suture and 2 mm lateral to the midline at a depth of 8.5 mm. Animals were sacrificed and perfused with 4% paraformaldehyde 7 (C57/B6) or 30 (NOD-SCID) days later. Brains were removed and placed in 2% paraformaldehyde containing 30% sucrose (v/v) overnight, and were sectioned into 20 m sagittal and coronal sections on a freezing

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34 microtome. Sections were stored at C in cryoprotectant solution until analyzed. Immunosuppressed animals (n=3) were injected with 10 mg/kg cyclosporin A (Sigma) immediately prior to transplantation and bidaily thereafter. Transplanted cells were identified by size (>20 ! m cell body diameter) and immunoreactivity for HNA, and were immunophenotyped by evaluating glial and neuronal marker (GFAP, O4, -III-tubulin, and NeuN) expression. ELISA Enzyme-linked immunosorbent sandwich assay (ELISA) was performed for glialderived neurotrophic factor using a protocol and kit adapted from a standard ELISA assay (Santa Cruz). Briefly, whole cell lysates of 10 5 primary human progenitors were created using RIPA buffer as previously described for Western blot applications. 50 ! l of protein (2 ! g sample protein + 48 ! l carbonate buffer) were added to carbonate-coated 96-well plates (pH 9.0) and incubated overnight at 4C. Carbonate solution was removed, and wells were blocked with PBS containing 1% BSA for 1 hour at room temp. Blocking buffer was removed, and GDNF antibody was added (1:500) for 1 hour at room temp. Wells were washed 3 x 5 min with PBS containing 0.05% Tween-20. Wells were washed once with diethanolamine buffer (10 mM diethanolamine, 0.5 mM MgCl 2 , pH 9.5), GDNF was added at 1 mg/ml for 10 minutes, and reaction was stopped with 0.1 M EDTA (pH 7.5). Plates were read on a spectrophotometer at OD 490 nm.

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35 CHAPTER 3 PHENOTYPIC AND FUNCTIONAL CHARACTERIZATION OF ADULT BRAIN NEUROPOIESIS Introduction 1 Attempts to trace the cellular source of ongoing neurogenesis in the adult CNS have recently led to the surprising conclusion that dedicated glial cells give rise to new neurons throughout life (Chiasson et al. 1999; Doetsch et al. 1999; Laywell et al. 2000; Goldman, 2003; Steindler and Laywell, 2003). Even though neurons and glia are both derived from the embryonic neuroepithelium, sharing common signaling pathways and downstream transcription factors during development (Rowitch, 2004), it seems rather difficult to imagine how one major cell class in the adult brain can transpose into the other. Postnatal neurogenesis in the SVZ of rodents proceeds as a characteristic series of events, where multipotent glial cells (referred to as type-B cells) are capable of dividing to form colonies of neuroblasts (type-A cells) through a transit-amplifying cell population (type-C cells) (Doetsch, 2003). Newborn neuroblasts migrate from the SVZ through the rostral migratory stream and mature to ! aminobutyric acid (GABA)-ergic granule cells and periglomerular cells, which integrate as inhibitory interneurons into the olfactory bulb of rodents 3-4 weeks after generation (Lois and Alvarez-Buylla, 1994; Belluzzi et al. 2003; Carleton et al. 2003). Specific features such as nestinand GFAP-expression are ascribed to the founder cells of postnatal neurogenesis ( Lendahl et al. 1990; Morshead et 1 This chapter contains figures and text excerpted from Scheffler et al, 2005.

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36 al. 1992; Gates et al. 1995; Laywell et al. 2000), but a distinctive antigenic and functional profile remains elusive. Traditional approaches for the isolation and characterization of persistent neurogenesis have relied on the in vitro neurosphere (NS) assay (Reynolds and Weiss, 1992; Kukekov et al. 1999) or on post hoc identification depending upon the incorporation of BrdU and/or retroviral constructs to label precursors during cell division. However, neither of these methods affords the recognition of the dynamic processes of maturation of individual cells en route from stem cell to fully differentiated neural phenotypes. Here, we present an alternative culture model that closely recapitulates in vivo postnatal/adult SVZ neurogenesis, allowing us to monitor the entire sequence of hierarchical events from glial-like stem cell to functionally mature (inter)neurons, and to expose phenotypical and electrophysiological properties of developmentally intermediate cell types. Recapitulation of SVZ Neurogenesis To expand a putative SVZ stem cell population, and to remove cells of limited proliferative capacity, single cell suspensions of microdissected SVZ tissue were proliferated as adherent monolayers >5 PDs (at least 3 passages) in defined proliferative media supplemented with EGF and FGF (Fig. 3-1 a). Cells expressing mature neuronal (NeuN, Map2) or oligodendrocyte (CNPase, O4) markers were not detected in expanded proliferating cultures at this point. Instead, a mixture of mature astrocytes expressing glial fibrillary acidic protein (GFAP), and immature nestin + glial phenotypes was present (Fig. 3-1 b). Ultrastructural analysis of proliferating cells exposed abundant protoplasmic

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37 astrotype morphologies with extensive bundled intermediate filaments (Fig. 3-1 c and d). SVZ-derived monolayers could be proliferated for more than 75 PDs without senescence, and when transferred to non-adhesive conditions multipotent primary and secondary neurospheres formed at passage 5 (3.2.06% and 0.83.25% respectively) and at passage 25 (4.12 0.79% and 0.81 0.13%) (data not shown). These observations indicate the maintenance of multipotent self-renewing cells in adherent conditions. Further examination of proliferating SVZ cultures revealed a nestin + /A2B5 + population of cells, among which 26.6 6.3% co-expressed low levels of GFAP. These cells showed compact morphologies and frequently signs of mitotic activity (Fig. 3-1 e and inset 3-1 e). Removal of growth factors and serum induced simultaneous differentiation of proliferating cultures, and resulted in the sequential reduction of the number of A2B5/GFAP low -expressing cells (Fig. 3-1 f). At the same time, rapid generation of committed neuronal progeny was observed. At 24 hours, glial cells expressing A2B5 and nestin (not shown) transiently co-express ! -III-tubulin (Fig. 3-1 g and h), a polycomb gene family member expressed during neuronal fate choice (Cai et al. 2000; Dennis et al. 2002). These morphologically unique cells are frequently devoid of GFAP (not shown), and contain large quantities of mitochondria and ribosomes (Fig. 3-1 i and j). Compacted cell colonies appear 3-4 days following growth factor withdrawal. Individual cells were small and round, and ubiquitously expressed ! -IIItubulin, nestin and A2B5 (not shown) as well as PSA-NCAM and low levels of Dlx2 (Fig. 3-1 k and l). Phenotypic and ultrastructural hallmarks correspond to mixtures of A and C cells described in vivo (Doetsch et al. 1997) (Fig. 3-1 m). This phenomenon was specific to cultures from the SVZ, as characteristic A and C cells could not be derived from cell dissociations of

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38 identically prepared parietal neocortex. To evaluate the potential for large-scale production of neuroblasts, confluent monolayers of passage 5 proliferating cells (n=9, at 80,000 cells/cm 2 ) were differentiated as described, yielding 39,545 2,691 neuroblasts ( ! -III-tubulin + cells)/cm 2 four days after differentiation. These cells are stably maintained in colonies on underlying glia, and were easily isolatable for subculture by detachment from underlying cells. Clonally plated cells were able to generate neuroblasts, albeit far less efficiently than adherent monolayers. Proliferating monolayers contained extremely low numbers of CD15 + cells (Fig. 3-2 a), which have been reported as being involved in postnatal neurogenesis. Examination of differentiating clusters of cells shows irregular distribution of PDGF and FGF receptor (Fig. 3-2 b,c), which may account for the concomitant production of CNPase + oligodendrocyte progenitors produced following differentiation (Fig. 3-2 e). III-tubulin + neuroblasts were largely positive for neurofilament L (Fig. 3-5 f), suggesting that the majority of the cells generated through induction of differentiation are dedicated neuroblasts. Development of Neuroblasts into Interneurons To examine the feasibility of developing glial-derived neuroblasts into mature cell types we induced terminal differentiation using retinoic acid and longitudinally monitored isolated colonies of newborn cells. Following their generation, immature A2B5 + /nestin + / ! -IIItubulin + neuroblasts (Fig. 3-3 a) lose nestin antigens and begin to extend bipolar processes (Fig. 3-3 b). With time, A2B5 expression is successively downregulated in developing clusters of neuroblasts, neuritic arborisation becomes increasingly complex, and more mature neuron markers are expressed (Fig. 3-3 c; NeuN

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39 not shown). Three to four weeks after growth factor withdrawal, characteristic mature interneuronal phenotypes are observed (Fig. 3-3 d and e). These neurons almost exclusively express GAD 65/67, a marker for GABAergic interneurons (Fig. 3-3 e). Taken together, these results suggest it is possible to recapitulate and longitudinally track a heteromorphic differentiation process in vitro that temporally and phenotypically follows SVZ neurogenesis in vivo. Dynamic Characterization and In Vitro Correlation of Neurogenesis Real-time microscopy was employed to capture and characterize the immediate dynamics of localized neurogenic events (Fig. 3-4 a). After induction of differentiation, cells maintain a flat, amorphous-glial appearance for one day. Following this 24 hr ‘silent’ period, a sudden and widespread series of rapid cell divisions leads to the initial appearance of neuroblasts. The observed rapid proliferation during this period appears to confirm the approximately 12.7-hour cell cycle time reported for the transit-amplifying SVZ population in vivo (Morshead et al. 2002). The timing of appreciated mitotic activity is consistent with in vivo observations where ! 50% of putative stem cells are mitotically active 48 hours after depleting constitutively proliferating SVZ cells (Morshead et al. 1994). Similar to in vivo studies (Doetsch et al. 1999), thymidine analog labeling using BrdU indicates the majority of neuroblasts are born 48-72 hours following induction of differentiation (Fig. 3-4 b). Our observations of immediate neurogenic events also correlated with the ability of differentiating cells to form clonally derived, multipotent NS (Fig. 3-3 c). Adherent cells were trypsinized during proliferation and after 24 and after 96 hr of growth factor withdrawal, counted, and plated at clonal

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40 density in a conventional NS-forming assay (see Methods). In accordance with findings of decreased neurogenesis in adult brain (Kukekov et al. 1999), total numbers (and diameters) of NS formed in this assay varied with age of SVZ tissue. However, a significant and comparably transient increase of the frequency of NS-forming cells was observed regardless of donor brain age at 24 hours after induced differentiation (1.86 and 1.6 times in P8 and adult-derived SVZ cultures, respectively). For both age groups, dissociated primary NS yielded clonally derived, multipotent secondary NS (Fig. 3-4 d and e), additionally demonstrating the self-renewing potential of rapidly amplifying cells in our culture system. SVZ Astrocytes Differentiate Into Mature Interneuronal Phenotypes Whole-cell patch clamp recordings were used to monitor the longitudinal transition of proliferating SVZ glial cells into mature neuronal phenotypes in vitro. Proliferating cells (Fig. 3-5 a) displayed a predominance of A-type K + currents (IK A ) with hyperpolarized resting membrane potentials. One day after induction of differentiation, rapidly amplifying cells exhibit low potassium conductances and a predominance of delayed-rectifying K + currents (IK dr ). At four days after induction, membrane properties of clustered newborn cells resemble those of SVZ-born neuroblasts previously characterized in vivo (Wang et al. 2004). Neuroblasts present characteristic IK dr , with no significant sodium channel contribution. Significant changes in passive membrane properties accompany the dramatic morphological transition observed during the first four days of differentiation (Fig. 3-5 b). Following the initial period of rapid change, a second, protracted period of functional neuronal maturation is entered, which

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41 mimics SVZ neurogenesis in vivo (Belluzzi et al. 2003; Carleton et al. 2003). This includes gradual repolarization, an increasing membrane capacitance, and a slowly decreasing membrane resistance (Fig. 3-5 b). Similar to both in vitro (Stewart et al. 1999) and in vivo (Belluzzi et al. 2003) recordings, immature neurons display TEA sensitive IK dr at 9 days after induction (Fig. 3-5 c). In some bipolar cells, application of TEA exposed underlying IK A , a typical potassium current of proliferating immature/glial cells (Fig. 3-5 a and c). Mature glial-derived neurons (28 4 days) could be elicited to fire series of TTX-sensitive action potentials (Fig. 3-5 d). Surprisingly, spontaneous synaptic activity consisted almost exclusively of inhibitory events (6/6 recorded cells), as demonstrated by application of picrotoxin, an inhibitor of GABAmediated synaptic transmission (Fig. 3-5 d). This suggests that, similar to SVZ neurogenesis in vivo, the fate choice of glial-derived neurons in vitro is largely restricted to inhibitory phenotypes. A broad range of immunophenotypic markers are associated with the generation of new neurons, many of which are altered in the glia-to-neuron transition, we examined the global changes in immunophenotype accompanying organotypic neurogenesis. Matched cultures were induced to differentiate and multiple markers were tracked at throughout generation of mature neurons (Fig. 3-6). These findings closely match the characteristic patterning of a rapid generation of mitotically active intermediate cells that form neuroblasts which undergo a protracted period of neuronal development culminating with the generation of mature neuronal cell types.

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42 Fig. 3-1. Monolayers of SVZ cells can be inducibly differentiated into newborn neuroblasts. (a) Phase contrast appearance of a proliferating passage 5 SVZ culture. (b) 3-dimensional reconstruction. Large GFAP + cells overly compact nestin-expressing cells. (c) Electron micrographs show cells with astrotypic morphology, containing abundant intermediate filaments (d). (e) Low levels of GFAP are found in a subpopulation of underlying nestin + (not shown)/A2B5 + cells. Arrow points to a GFAP low+ /A2B5 + cell. The inset shows one of these cells undergoing division. (f) Proliferating SVZ cultures contain 57.3 4.3% GFAP + and 42.3 1.5% A2B5 + cells. Following induction of differentiation, GFAP-expression among A2B5 + cells (26.6 6.3% during proliferation) decreases sequentially to 15.7 1.5 and 2.9 1.5% at 1 and 4 days after growth factor withdrawal, respectively. (g-h) 24 hours following growth factor removal, differentiating cells display heterogeneous nuclear compaction and transiently co-express A2B5 and ! -III-tubulin (h, inset, mitotic cell identified by asterisk in g). (i)

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43 These cells maintain an intermediate morphology, including cell size and nuclear diameter, and display extremely abundant mitochondria and free ribosomes (j). (k) Defined neuroblasts become abundant 4 days following initiation of differentiation, and are morphologically distinctive as round, phase dark cells with compacted nuclei (inset). (l) Cells are PSA-NCAM + , weakly express Dlx-2, and are ultrastructurally similar to mixtures of type A and C cells in the SVZ (m). Scale bars ( m): a=200; b=50; e ,h,l=30; c=10; i,m=5; d,j=2. Fig. 3-2. Generation of cellular heterogeneity accompanies differentiation. (a) One day following inducible differentiation, CD15 + cells are expressed at extremely low frequency (<0.01%). (b) Primordial clusters of cells exhibit alteration in receptor distribution, as evidenced by disparate distribution of platelet-derived growth factor (PDGF) and FGF-1 receptors (b and c) in the same visual field. (d) PSA-NCAM + early neuroblasts are closely associated with cells expressing Sox1, but do not express the marker when evaluated three days after the induction of differentiation. (e) CNPase + oligodendrocyte progenitor cells are generated near clusters of Tuj1 + new neurons, but do not coexpress Tuj1. (f) Neurofilament L (NF-L) is frequently coexpressed in Tuj1 + clusters of neuroblasts. Scale bars 100 m (a), 50 m (b-f).

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44 Fig. 3-3. SVZ-generated neuroblasts develop into interneuron phenotypes. (a) Upon appearance, neuroblasts co-express nestin and ! -III-tubulin. (b) Soon after, nestin expression is lost and bipolar processes are extended. (c) Expression of more mature neuronal markers corresponds to increasingly complex neuronal arborization. At 28 days, abundant interneuron phenotypes resemble granule cell (d) and periglomerular

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45 morphologies (e), expressing GAD 65/67. Scale bars ( m): a-c=25; d ,e=30; days=time after induction of differentiation. Fig. 3-4. A mitotically active, multipotent cell emerges transiently during early stages of in vitro neuropoiesis. (a) Time lapse microscopy of growth factor withdrawn SVZ cultures reveals a transient period characterized by rapid cell divisions leading to the initial appearance of neuroblasts within 27 hours. (b) BrDU applied at 48-72 hours following initiation of differentiation labels >95% of all generated neuroblasts. (c) Clonal NS were derived from cultured postnatal day 8 (P8) and adult SVZ cells (left panel for morphological comparison). Total numbers and relative frequencies (right panel) of NS generated from 100,000 cells per condition increase significant but

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46 transiently at 24 hours after WD (*p<0.01 for adult and P8 compared to nonWD and 4dWD. (d) Primary and secondary P8 and adult (shown here) neurospheres yield neurons and glia (e). Scale bars ( m): a=15; b=60; c=200; d,e=20. WD, refers to time after growth factor withdrawal. Fig. 3-5. Differentiating cells recapitulate membrane development consistent with a glialto-neuron transition. (a) Proliferating (prolif; n= 45), rapidly dividing (1 day; n=8), and newborn neuroblast (n=20) cell populations were identified in culture using infrared differential interference contrast (DIC) microscopy. Patch-clamp studies (voltage clamp, center; current clamp, right) reveal characteristic changes of functional phenotypes. Cells were incapable of generating action potentials during this period. A small percentage (6/45) of proliferating cells possessed TTX-sensitive voltage-gated sodium channels. (b) Passive membrane properties (membrane potential, V m ; capacitance, C m ; and input resistance, R m ) during development of SVZ glial cells into functionally mature neuronal

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47 phenotypes. (c) Voltage clamp characterization of IK dr 9 days after growth factor withdrawal (n=20). In some cells (cell 1), TEA application exposes underlying IK A . (d) Glial-derived mature neurons (n=14) fire TTX-sensitive action potentials. Spontaneous synaptic activity can be recorded and entirely blocked by application of picrotoxin (lower traces).

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48 Type Marker Proliferative State Undifferentiated 1D Diff 3D Diff 7-28D Diff Neuronal Doublecortin + +++ +/PSA-NCAM + +++ + NeuN ++ Map2a-c + ++ Tyrosine Hydroxylase +/+/GAD 65/67 ++ Dlx-2 ++ +++ + -III-Tubulin/ Tuj1 + ++ +++ +++ Progenitor/ Cell Cycle A2B5 +++ ++ + Nestin +++ +++ ++ + Sox1 ++ ++ + Sox2 +++ +++ ++ + Sox3 ++ + + + CD133 + +/FGFR1 +++ +++ +++ + EGFR +++ +++ +++ + CD15 + Ki-67 ++ +++ +++ +/Glial GFAP ++ ++ + + NG2 + + + Vimentin + + + + S100 + + + +/Glutamine Synthetase + ++ + +/GalC/O4/ CNPase +/+ + Lineage CD11b + + + + CD45 + Fig. 3-6. Immunophenotypic alteration accompanying neurogenesis and maturation of neuronal progeny following inducible differentiation in culture. Semiquantitative analysis was used as a marker-specific comparison in the following expression ratios: ()=0-0.1%, (+/-)=0.1-1%, (+)=1-25%, (++)=25-50%, (+++)= 50-100%.

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49 CHAPTER 4 TUJ1+ CELLS IDENTIFY NEUROGENIC CELLS OF THE SUBVENTRICULAR ZONE AND CHARACTERIZE THE ASTROCYTE-TO-NEURON GESTALT Introduction Prospective identification of primordial cells in the postnatal brain has become a source of continuing interest, both in identifying cells for repair and replacement in the central nervous system (CNS) and as a means to characterize endogenous adult neurogenesis ( Morshead and van der Kooy, 2004). Neurogenesis in the subventricular zone (SVZ), the largest postnatal neurogerminal region, is accomplished by an astrotypic NSC (‘B cell’) which gives rise to committed neuroblasts (A cells) via a rapidly dividing intermediate cell (Doetsch et al. 1999) . While significant evidence exists describing NSCs as glial in nature (Doetsch 2003), the inaccessible nature of the ventricular niche makes prospective identification of NSCs and their behaviors difficult. In vitro efforts to characterize NSCs generally rely on identification of clonally derived, proliferative-active cells that form multicellular aggregates containing multilineage neural cell types (Reynolds and Weiss, 1992). Such assays require post hoc NSC identification, and it remains unclear whether committed cell types generated in this system reflects the process described in vivo. As such, existing in vivo and in vitro methods of NSC study are not well-suited to (a) prospectively identify NSCs and (b) track the NSC ! progenitor ! neuroblast transition during the generation of progeny.

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50 To better identify and characterize primordial NSC populations, we previously created a novel adherent culture condition containing primary dissociates from neuropoietic SVZ, which retain astrotypic cells with NSC characteristics in an adherent system (Scheffler et al. 2005). When induced to terminally differentiate via mitogenic withdrawal, these cells synchronously undergo a concerted terminal differentiation which characteristically recapitulates temporal and phenotypic generation postnatal neurogenesis appreciated in vivo. Using this system, it is possible to employ parallel avenues of investigation to isolate, track, and analyze the developmental dynamics and cells comprising the transition of astrocyte to neuron. An analysis of undifferentiated NSCs using a variety of immunological markers reveals heterologous class III -tubulin (Tuj1) expression in a subpopulation of SVZderived protoplasmic astrocyte-like cells. This is interesting, as Tuj1 represents a polycomb gene member involved in fate choice and proliferative potential ( Katsetos et al. 2003). Although Tuj1 + cells comprise a relatively small portion of cultured cells, a high percentage of these cells colocalize with putative NSC and progenitor markers. To examine the role of Tuj1-expressing cells in organotypic postnatal neurogenesis, we longitudinally monitored Tuj1 + cells in conditions favoring NSC differentiation. Immediately following induction of differentiation, Tuj1 positive cells assume either morphologically intermediate phenotypes or (more commonly) a compacted morphology. As differentiation proceeds, only Tuj1 + cells divide to form multicellular aggregations of increasing size, which bridge the astrocyte-to-neuron transition and resolve into discrete clusters of neuroblasts within 96 hours of induction of differentiation.

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51 Ultrastructural examination of differentiating clusters of cells reveals dramatic changes in cellular morphology accompanied by cellular remodeling. Transient increases in free ribosomes, mitochonria, and increased endoplasmic reticulum, golgi bodies, and nuclear pore complex formation are also appreciated, indicating increases in transcription, translation, and cellular energy consumption. To further characterize Tuj1 + cells as essential for organotypic differentiation, we examined the role of Tuj1 + in the proliferative dynamics of differentiation. Alterations in proliferative dynamics have been reported in the generation of progeny from differentiating SVZ NSCs (Doetsch et al. 2002) . Unlike astrocytes derived from nonneurogenic regions, cells from neurogenic regions respond to mitogenic induction of differentiation with an initial period of slowed growth, followed by a period of increased mitotic activity culminating with the appearance of mature neuroblasts. Analysis of this delayed proliferative response reveals a lagging upregulation of multiple cyclin levels, the activation of which correlates to increases in expression of Ki-67 in Tuj1 + cells, suggesting that these cells underly the proliferative response to induction of differentiation. To correlate the ‘organotypic’ generation of neuronal progeny to current in vitro standards, neurospheres were generated from differentiating cells at various time points. Cells derived from periods of differentiation corresponding to an enrichment for transitamplifying ‘C cells’ produced significantly higher numbers of neurospheres than other developmental timepoints. Furthermore, only cells for this developmental period remained capable of generating equivalent numbers of secondary neurospheres,

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52 suggesting the developmental intermediates retain characteristics traditionally ascribed to quiescent NSCs. To further assign proliferative potential and assess fate choice of cells in various stages of neurogenesis, single cells from various stages of differentiation were randomly selected, plated and assessed for differentiation and proliferative scope under conditions favoring differentiation. Cells isolated from transit-amplifying enriched phases of development display the greatest proliferative potential and were preferentially enriched for neuroblast generation. By applying this model to characterize the relatively uncharacterized process of postnatal neurogenesis, a more detailed study becomes possible, allowing identification of novel cell types and processes governing postnatal neurogenesis. Additionally, employing this model to characterize NSCs and their behaviors may act as a basis for translating findings between in vivo and in vitro studies. Tuj1 Defines a Subset of Immature Glial Cells To characterize the transformation of gliotypic stem cells into committed neuronal cell types, we cultured whole-dissociate SVZ tissue in a monolayer culture as previously described (Scheffler et al. 2005). Initial characterization of passage 1 proliferating cultures indicated a heterogenous composition, including PSA-NCAM + and NeuN + neuronal cell types, CNPase + oligodendroglial cells, GFAP + astroglial cells, and CD11b + microglial cells (data not shown). Following culture in defined proliferative medium for 3 population doublings (PDs, approximately 7-10 days), neuronal and oligodendroglial cell types (as distinguished by morphology and immunocytochemistry) were not appreciated. The primary proliferating cultures appear to be a heterogeneous mixture of

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53 cells displaying multiple immature markers, including nestin and A2B5, and glial markers, including GFAP and vimentin (Scheffler et al. 2005). Microglial cell types were still detectable as a minor (<2% of total cell number) population, and remain relatively constant throughout culture. A minority (22.2+/-8.4%) of undifferentiated SVZ cells express the intermediate filament Tuj1 (class III tubulin), often in cells displaying a protoplasmic astrocyte morphology. To determine whether Tuj1 was a potential NSC and/or progenitor marker, we examined Tuj1 + cells for coexpression with phenotypic markers associated with neural progenitor and stem cells. Tuj1 + cells frequently coexpress A2B5 (Fig. 4-1 a), FGF receptor 1 (Fig. 4-1 b), CD133 (Fig. 4-1 c), and multiple sox-family proteins (Fig. 4-1 e-g). Nestin and GFAP are also frequently expressed (Fig. 4-1 d and 4-3 a). Furthermore, Tuj1 is frequently expressed adjacent to the lateral ventricle in vivo (Fig. 1h). Tuj1 was expressed in similar ratios in nonneuropoietic tissue. An A2B5 + /GFAP + /nestin + progenitor cell has been implicated as a founder cell for generation of progeny (see chapter 3). To evaluate whether Tuj1 is expressed in cells displaying both progenitor and astrocytic markers, we analyzed coexpression of Tuj1, GFAP, and A2B5 and/or nestin (Fig. 4-2). Tuj1 was heterologously expressed in cells expressing GFAP (Fig. 4-2 a), nestin (Fig. 4-2 b), and A2B5 (Fig. 4-2 c). These findings further suggest that Tuj1 may label a population of primordial astrotypic cells, and may represent a potential NSC marker. Tuj1 expression characterizes cells undergoing astrocyte-to-neuron differentiation The predominant stem cell of the adult brain is described as astrotypic (Doetsch 2003). To further verify that Tuj1 expression occurs in astrocytes, we examined the

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54 morphology of undifferentiated Tuj1 + cells. Tuj1 is largely expressed in cells retaining protoplasmic morphology (with or without GFAP expression, data not shown), which typically extend one or more processes to contact adjacent cell bodies (Fig. 4-3 a). Ultrastructural examination of undifferentiated cells confirms the presence of intermediate tubules in otherwise unremarkable astrotypic cells (Fig. 4-3 b, c). To determine whether Tuj1 labels cells involved in the generation of neuronal cell types, passage 3 ageand density-matched cultures were induced to differentiate and longitudinally examined at 12-hour intervals until the appearance of defined clusters of PSA-NCAM + neuroblasts 72-96 hours later. Two distinct populations of Tuj1 + cells were present in culture 12 hours after induction of differentiation: a less prevalent hybrid cell type (Fig. 4-3 d) which are morphologically similar to previously described intermediate cells with morphology intermediate to astrocyte and neuron (Laywell et al. 2005), and (predominantly) in single cells containing scant cytoplasm and a compacted morphology (Fig. 4-3 e). GFAP is very rarely expressed in either cell type. Following their appearance in culture, single Tuj1 + cells rapidly expand in number, forming densely packed clusters. Cells within these characteristic formations exhibit increases in free ribosomes, endoplasmic reticulum, and nuclear pore complexes (Fig. 4-3 f-h), suggesting according increases in protein synthesis and cell cycle. Tuj1 + cells within these clusters express EGF receptor (Fig. 4-3 i, inset), matching a report for the expression of this receptor in developmentally intermediate cells (Sun et al. 2005). Defined clusters of Tuj1 + cells increase in size following their initial appearance (Fig. 4-3 i-k). Cells with apparently contiguous cytoplasmic elements are occasionally present within these clusters, but are eventually supplanted by individual discrete cell bodies (indicated by

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55 live cell labeling with lipophilic fluorophore conjugated dye, data not shown). Increases in both mitochondria and lysosomes on cells within these clusters is appreciated (Fig. 4-3 l,m). A progressive reduction in average nuclear size also accompanies differentiation within these clusters (9.6+/-0.2 vs 5.1+/-0.17 m 72 hours following differentiation). Discrete clusters of newly-generated neuroblasts, which are Tuj1 + , GFAP , and nestin + are present 60-72 hours after induction of differentiation (Fig. 2n,o). These cells remain mitotically active (indicated by continued Ki-67 expression, BrDU uptake, and nuclear cleavages, data not shown and Fig. 4-3 p), display scant cytoplasm, reduced numbers of mitochondria, and appear in closely associated clusters. Following their appearance in culture, individual neuroblasts dissociate as maturation of neurons occurs, including process extension and upregulation of mature markers (Fig. 4-3 q). To ensure Tuj1 + acts to label maturing populations of cells instead of discrete progenitor (i.e., as a polycomb gene marker) and mature populations (i.e., as a functional marker), Tuj1 expression was measured in cells induced to differentiate and compared to progenitor marker expression. Tuj1 cells assuming characteristic intermediate morphologies cease to express both A2B5 (Fig. 4-4 a) and Sox2 (Fig. 4-4 b), suggesting these cells mature upon induction of differentiation. To further examine the role of Tuj1 + cells in the generation of neuroblasts, ageand density-matched passage 3 SVZ and cerebellar astrocytes were induced to differentiate and were analyzed for total cell number, thymidine analog incorporation, cyclin expression, ploidy, and cell cycle throughout differentiation. Total cell number measurements revealed that, while non-neuropoietic tissue fails to proliferate following the induction of differentiation via mitogen withdrawal (see methods), SVZ tissue enters

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56 a period of relative quiescence lasting approximately 24 hours, followed by a rapid increase in cell number (Fig. 4-5 a). To confirm this pattern of proliferation, and to further evaluate the proliferative dynamics throughout differentiation, BrDU was added in 24-hour intervals to differentiating SVZ and cerebellar tissues. While nonneuropoietic cells remain mitotically inactive, a significant increase in BrDU incorporation is appreciated 24-72 hours following induction of differentiation (Fig. 4-5 b). Protein expression comparison in differentiating neuropoietic astrocytes reveals transient increases in multiple cyclin levels 12-24 hours following differentation (Fig. 4-5 c), which may drive the appreciated proliferative activity. To determine whether Tuj1 + cells are involved in the observed proliferation accompanying differentiation, Ki-67 expression in Tuj1 + cells was measured for neuropoietic and non-neuropoietic tissue. Increases in Ki-67 expression were appreciated in only SVZ-derived tissue following differentiation (Fig. 4-5 d), including frequent expression within Tuj1 + cells comprising characteristic clusters bridging the astrocyte to neuron transition (Fig. 4-5 d, inset). Increases in BrDU incorporation were also appreciated in SVZ-derived Tuj1+ cells 24-72 hours following differentiation (data not shown). As alteration of proliferative dynamics have been reported in the generation of progeny (Doetsch et al. 2002), we FACsorted differentiating SVZ astrocytes for ploidy and cell cycle analysis throughout differentiation. As indicated by previous analysis of total cell number, a relatively small initial population of cells begin differentiation in S phase, but dramatically increase following 60 hours in conditions favoring differentiation (Fig. 4-5 e). The period initially following differentiation is characterized by an increased fraction of cells entering G1/2 (Fig. 4-5 f). This period of hypertrophy

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57 corresponds to a period of increased aneuploidy, which is significantly reduced as terminal differentiation occurs. Forward scatter analysis reveals cells from periods enriched with G1/2 cycling cells to be largely hyperploidic (data not shown). Developmentally intermediate cells possess enhanced multipotentiality and proliferative potential NSCs of the ventricular neuraxis are classically defined in vitro as having the capacity to form self-renewing and multilineage neurospheres from clonally cultured cells (Reynolds and Weiss 1992). To evaluate the ability of cells undergoing ‘organotypic’ differentiation to self-renew and generate multipotent progeny, randomly selected SVZ-derived dissociates were seeded into antiadhesive conditions at clonal density while proliferating and 1, 4, and 7 days after induction of differentiation and assessed for neurosphere formation. Cells from all developmental states were able to generate neurospheres in varying proportions, with cells derived from states of advanced differentiation progressively less able to generate primary neurospheres (Fig. 4-6 a). To assess whether the generation of neurospheres is transient or sustainable, primary neurospheres were dissociated into secondary neurospheres. While sphere-forming cells derived from other developmental time points generate secondary neurospheres at lower rates, sphere-forming cells derived from an intermediate point 24 hours following differentiation are able to generate equivalent numbers of secondary neurospheres (Fig. 4-6 a). Primary neurospheres derived from cells spending 7 days in conditions favoring differentiation are unable to generate secondary neurospheres. Secondary spheres generate equivalent numbers of tertiary neurospheres, irrespective of initial developmental state or proliferative frequency (Fig. 4-6 b).

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58 Finally, the intrinsic capabilities of Tuj1 + cells throughout differentiation was examined. To examine the proliferative and multilineage capacity of isolated individual cells gathered from dissociates undergoing progressive ‘organotypic’ differentiation, randomly selected cells were selected from differentiating SVZ astrocytes and re-plated into conditions favoring their differentiation at clonal density, and evaluated forproduction of committed cell types 3 days later. While cells derived from all stages of differentiation were able to generate all cell types, cells isolated from conditions favoring the increased presence of intermediate progenitors (‘C cells’, following 24 hours of differentiation) generate significantly larger numbers of neuroblasts than undifferentiated or more differentiated isolates (Fig. 4-6 c, d). Isolated cells were also evaluated for proliferative ability following re-plating in differentiation to evaluate the proliferative ability of cells from various states of differentiation. Cells isolated from progenitorenriched stages of differentiation are enriched for cells that exhibit extensive proliferative activity (Fig. 4-6 d, e). Cells derived from undifferentiated or more differentiated states display reduced proliferative indexes, suggesting reduced proliferative capacity to generate progeny (Fig. 4-6 e).

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59 Fig. 4-1. Tuj1 labels putative NSC/progenitor populations. Expression of Tuj1 in cells coexpress A2B5 (a, arrows), FGFR1 (b), and CD133/Prominin-1 (c) in undifferentiated passage 3 SVZ dissociates. Tuj1 is expressed in the mitotic spindle of dividing cells (insets in a and c). (d) Tuj1 is expressed in a substantial fraction of cells expressing NSC and progenitor markers. CD15 and NG2 were examined and were not expressed in undifferentiated cultures. (e-g) Tuj1 is expressed in multiple members of the Sox protein

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60 family. Coronal section of the SVZ reveals Tuj1 + cells are present in the subependymal layer adjacent to the lateral ventricle (LV, arrows). Scale bars 50 m (a, c, e-g), 25 m (b), 100 m (h). Fig. 4-2. Tuj1 is coexpressed in immature and astrocytic cells derived from both adherent monolayers and neurospheres. (a) Monolayer-derived astrocytes plated in defined proliferative media exhibit intermittent coexpression of Tuj1 and GFAP. Monoclonal neurosphere-derived cells also contain Tuj1 and GFAP coexpressing cells, which heterologously express nestin-eGFP (nes-eGFP). Scale bar 500 ! m (a-c).

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61 Fig. 4-3. Tuj1 + cells characterize the astrocyte-to-neuron transition of rodent SVZ cells. (a) Tuj1 + cells are initially appreciated in undifferentiated cultures in cells possessing a protoplasmic astrocyte morphology (Tuj1, red; GFAP, blue; nestin-eGFP, green). Intermediate 25 nm tubules label astrotypic cells with uncompacted nuclei and broad cytoplasm (b, arrows in c). (d,e) Twelve hours following induction of differentiation, both intermediate cell types and compact single Tuj1 + cells are present, which proliferate

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62 to form clusters (f) characterized by increases in ribosomes (g), mitochondria (g), and rough endoplasmic reticulum (h). (i,j) Increasingly large groupings of intermediate cells remain ubiquitously positive for Tuj1 (red), and do not express GFAP (green). Cells from this developmental stage express EGF receptor (red, inset i; Tuj1, yellow). (k) Average Tuj1 + cellular aggregation following differentiation. (l,m) 48 hours after induction of differentiation, an increase in mitochondrial density is accompanied by increased in lysosomal remodeling. (n) 72 hours after induction of differentiation, discrete clusters of Tuj1 + neuroblasts are formed (red, Tuj1; green, nestin-eGFP; blue, GFAP). These cells remain in clusters (o) but remain mitotically active (nuclear cleavage in p) and mature to express mature marker map2a-c (p). Images counterstained with DAPI. *p<0.05, **p<0.01, one way ANOVA. Fig. 4-4. Immature markers are lost from Tuj1 + cells upon induction of differentiation. (a) One day after inducible differentiation of monolayer, A2B5 is not expressed in differentiation-transiting Tuj1 + intermediate cells. (b) Tuj1 + cells are also negative for the transcription factor Sox2, which exhibits continued expression in underlying cells. Scale bar 250 ! m (a ,b).

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63 Fig. 4-5. Tuj1 + cells drive neurogenesis through characteristic mitotic dynamics. (a) SVZ-derived cells proliferate in response to removal of mitogenic stimuli, following a lag period of 24 hours, which is not reflected in cerebellar (nSVZ) tissue. (b) 24-hour incremental BrDU labeling of differentiating cultures reveals increases in thymidine analog incorporation 24-72 hours following induction of differentiation. (c) 12 hr semiquantitative measurement of cyclin levels in proliferating and differentiating cells. 42

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64 KDa control band (-actin) shown. (d) Ki-67 expression is increased in Tuj1 + cells in neurogenic, but not non-neurogenic, tissue following induction of differentiation (Inset: Ki-67 expression (red) in Tuj1 + cells (green)). (e, f) SVZ cells induced to differentiate initially pass through a 24-hour period of minimal proliferation, increased G1/2, and increased aneuploidy before resolving into a conventional proliferative population. Image counterstained with DAPI. *p<0.05, one-way ANOVA.

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65 Fig. 4-6. Intermediate progenitors retain multipotentiality and self-renewal. (a) Primary and secondary neurosphere production from undifferentiated proliferating (prol) and differentiating (1, 4, or 7 days, respectively) SVZ cells. (b) Tertiary sphere formation from secondary spheres generated in (a). NS generated from cells differentiating 7d did not form 2 NS. (c) Progeny analysis of individual cells expanded as adherent clones for 3 days. (c) Cells derived from SVZ cells placed in differentiating conditions for one day

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66 display Tuj1 + neuroblasts (red), nestin-eGFP (green), and GFAP (blue). (e) Proliferative scope of individual differentiating SVZ cells placed in conditions favoring differentiation for 3 days. *p<0.05, one way ANOVA.

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67 CHAPTER 5 BROMODEOXYURIDINE HAS CYTOTOXIC EFFECTS ON NEURAL STEM CELLS AND IMMORTAL CELL LINES Introduction Examination of dividing cells is frequently performed by pulsed addition of thymidine analog. BrDU is widely employed as one such analog, and has been shown to incorporate into cells during S phase (Nowakowski et al. 1989), and has been used extensively to characterize the proliferative dynamics of and prospectively identify NSCs in vivo (Gould and Gross, 2002). In addition to functioning as a labeling agent, BrDU incorporation may result in developmental defects in persistently dividing cells during development. BrDU toxicity has been suggested during both embryonic and neonatal development (Garner, 1974; Agnish & Kochhar, 1976 a, b; Pollard et al. 1976; Dribin & Jacobson, 1978; Bannigan & Cottell, 1991; Nagao et al. 1997; Kolb et al. 1999). Specifically, BrDU administration to gestating mothers has led to developmental malformations in rodents in a timeand dose-dependent. BrDU has also been reported as a mutating and photosensitizing agent (reviewed in Wilt & Anderson, 1972). Previous studies have observed a correlation between the application of BrDU with doesdependent reduction in the generation of committed cell types in embryonic rodent cells, which was attributed to classic cell death mechanisms (Caldwell et al. 2005).

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68 Several reports exist suggesting stem cells divide via an asymmetric mechanism, with preferential sequestration of newly synthesized DNA into daughter cells acting as a means to prevent DNA damage (Sherley et al. 1995; Potten et al. 2002). This phenomenon has been observed in the developing CNS (Sun et al. 2005). The relation between asymmetric cell division and the effects of BrDU incorporation is unclear. To investigate this phenomenon in persisting postnatal neural stem cells, we applied BrDU constitutively to both primary ( neurogenic) subventricular astrocytes and NCI/HE57 tumor cell line, resulting in the abrogation of growth and the failure of subventricular astrocytes to generate neuroblasts when induced to differentiate. Both neurosphere number and size were reduced in an age-dependent manner when generated in the presence of BrDU, and increased numbers of senescent cells were appreciated in the neurogerminal zones of adult rodents injected with BrDU. Interestingly, both SVZ astrocytes and RG2 glioma cells exhibit a senescent phenotype in the presence of BrDU, with unremarkable levels of apoptosis or necrosis. However, mitochondrial stress appears to be significantly increased in both primary and glioma cells, suggesting BrDU may have specific functions as an inducer of cellular senescence in proliferative-active cells of the CNS. BrDU Attenuates the Growth of Primary Neurogenic Cells and Cell Lines To evaluate the overall effect of BrDU on proliferative potential, growth rates of NCI/HE57 tumor cells and PD3 SVZ astrocytes cultured in proliferative media supplemented with 10 ! M BrDU were measured. Both immortalized and primary cell’s growth rates were reduced significantly with the application of thymidine analog (Fig. 51 a,b). To evaluate whether BrDU application was systemic (i.e., conventionally toxic)

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69 or ongoing (i.e., a result of incorporation), a single, 24-hour dose of 10 ! M BrDU was applied to primary and immortalized cells and their growth rates were recorded. Interestingly, a progressive decrease in growth rates was appreciated in both cell types, suggesting a single dose may have long-lasting effects in proliferative potential (Fig 5-1 c). Inhibition of mitosis through application of reversible growth inhibitors (1 ! g/ml aphidicolin 24 hours prior to and during BrDU application) was capable of preventing this reduction in growth and restoring normal rates of cell division following washout (data not shown). Similar to previous reports (Caldwell et al, 2005), a single dose of 10 ! M BrDU 24 hours prior to induction of differentiation was sufficient to result in almost complete abrogation of neuroblast production (Fig. 5-1 d). BrDU Affects Neurosphere Formation The effect of BrDU on the self-renewal on NSCs in postnatal animals was evaluated by neurosphere formation in cells derived from young and adult mice grown in the presence of BrDU. Primary neurospheres derived from young animals were equivalent in number when cultured in the presence of BrDU (Fig. 5-2 a); however, adult-derived neurospheres exhibited a dose-dependent reduction in neurosphere number (Fig. 5-2 b). Self-renewal of neurospheres is also affected by BrDU addition, as both young and adult-derived neurospheres were incapable of generating secondary neurospheres, regardless of the presence of BrDU (data not shown). Interestingly, neurosphere size is affected in young animals, resulting in a reduction in average neurosphere diameter with increasing BrDU dose (Fig. 5-2 c). Neurospheres remain morphologically normal in the presence of BrDU (Fig. 5-2 d).

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70 BrDU Induces Senescence In Vivo and In Vitro In vivo effects of BrDU on neurogenesis were measured by unbiased cell counting of SA--Gal in consensus regions of ongoing postnatal neurogenesis. repeated injections of BrDU resulted in an increase in SA--Gal + cells in both lateral SVZ and hippocampus (Fig. 5-3 a). 250 mg/kg injections of BrDU (equivalent to an intracellular concentration of " 10 ! M) BrDU demonstrated increased expression within the lateral subventricular wall (Fig. 5-3 b) and the hippocampus (Fig. 5-3 c), most notably in the hilus and dorsal blade (Fig. 5-3 d). To determine if BrDU was capable of inducing a senescent phenotype in cultured cells, primary SVZ astrocytes and RG2 glioma cells were cultured in the presence of BrDU and evaluated for phenotype 3 days later. Primary SVZ astrocytes exhibited a flattened, dystrophic morphology (Fig. 5-4), often characterized by vacuole formation (Fig. 5-4, lower left panel). RG2 cells display a compacted morphology in the absence of BrDU (Fig. 5-4 upper right panel), but become increasingly large, frequently displaying giant cells characterizing a senescent state (Fig. 5-4, lower right panel). SA--Gal analysis of these cells revealed increases in senescent markers after a minimum of three days in the presence of 10 ! M BrDU (data not shown). Oxidative stress has been a frequently attributed mechanism for aging on a cellular and molecular level (Sohal et al, 2002). To examine whether BrDU promotes cell death, RG2 cells were grown in the presence of 10 ! M BrDU and examined for necrotic (propidium iodide + ) and apoptotic (TUNEL + ) cells. Propidium iodide was expressed in vital cells at low levels, regardless of BrDU dosage (Fig. 5-5 a,c). TUNEL + cells were also expressed at low frequency in culture in a BrDU-independent manner

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71 (Fig. 5-5 b). To determine if mitochondrial-driven oxidative stress is upregulated in the presence of BrDU, mitochondrial oxidative stress was measured through addition of mitoSox dye (Molecular Probes) to vital cells. RG2 cells exhibited mitochondria-specific oxidative product production in a dose-dependent manner (Fig. 5-5, d), suggesting a potential mechanism underlying senescence. These findings suggest that BrDU achieves irreversible, incorporation-dependent growth inhibition and induction of senescence in postnatal NSCs. BrDU’s effects are apparently nontoxic, and appears to trigger an upregulation in intracellular oxidative stress. Fig. 5-1. BrDU abrogates cell growth in cell lines and neurogenic astrocytes. NCI/HE57 teratoma cells (a) and passage 3 subventricular astrocytes (b, derived from pooled postnatal day 8 C57/B6 periventricular tissue as described) grown in culture conditions containing continuous 10 ! M BrDU. (c) A single 24 hour application of BrDU (10 ! M) is sufficient to create a progressive reduction in the growth rate of both passage 2 subventricular astrocytes (SVZ) or the RG2 glioma cell line. Single 24 hour application of 10 ! M BrDU accompanying induction of differentiation effectively

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72 abrogates neuroblast generation in passage 3 SVZ astrocytes when examined 3 and 7 days later (3 days shown). All growth rates calculated by either total estimated cell number (a,b) or relative growth rate (c) based on per passage cell counting. *p<0.05 (a,b,c) or <0.0001 (d). Fig. 5-2. BrDU addition affects NS number and size in NS derived from young and adult animals. (a) Young (postnatal day 8 animals) exhibit insignificant differences in numbers of NS generated when grown in the constitutive presence of 10 ! M BrDU. However, NS derived from adult animals (postnatal day 90, b) exhibit a dose-dependent reduction in NS number. NS derived from young animals exhibit a dose-dependent decrease in NS size when cultured in constitutively applied 0-100 ! M doses of BrDU (c, 0 vs 50 ! M BrDU for young animal-derived NS grown for 14 days). *p<0.05, one way ANOVA. Scale bar 25 ! m (d). Scale bar 25 ! m (d).

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73 Fig. 5-3. Systemic BrDU administration results in increases in senescent cell types. (a) Unbiased stereological quantification of subventricular and hippocampal tissues from mice injected with either saline or 1 or 3 injections of 250 mg/kg BrDU every other day for 6 days reveals increases SA--Gal + cells in the SVZ, hilus, dorsal and total granular cell layer (GCL), but not ventral blade. (b) Lateral ventricle examination of injected rodents reveals an increase in blue SA--Gal cells in periventricular niche, as compared to saline-injected control. (c) Hippocampal SA--Gal measurement reveals increased expression of senescence markers (magnification of boxed area shown in d). *p<0.05, one-way ANOVA. Scale bars 1 mm (b,c); 50 m (d, magnified ventricular wall in b).

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74 Fig. 5-4. Addition of BrDU results in morphological alteration of both subventricular astrocytes and cell lines. Passage 3 SVZ astrocytes gathered from periventricular tissue and rat glioma (RG2) cells were cultured in the presence of 10 M BrDU for 3 days. Cells grown in the presence of BrDU exhibit morphological alterations, including a flattened protoplasmic morphology and multinucleated cell types (arrow in lower left panel) and/or presence of cytoplasmic vacuoles (arrow in lower right panel).

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75 Fig. 5-5. BrDU does not abrogate growth through necrosis or apoptosis, but does induce mitochondrial stress. (a) Live-cell analysis for propidium iodide (PI) uptake in RG2 cells treated for 48 hours with 10 M BrDU reveals a low, consistent level of cell death in BrDU-treated cultures. Similarly, the frequency of apoptotic cells (measured by TUNEL in b) remains low in similarly cultured RG2 cells. (c) PI expression in cells cultured with 50 M BrDU reveals infrequently occurring dead or dying cells. (d) Upon addition of BrDU, increases in mitochondrial oxidative stress (as measured by mitochondrial superoxide indicator dye MitoSox) are appreciated within 48 hours of BrDU application.

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76 Superoxide stress is localized to mitochondria (punctate reactivity in arrow-labeled cells in inset). Scale bar 50 m (c ,d).

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77 CHAPTER 6 NEURAL STEM CELLS ARE DEPENDENT ON MICROGLIAL SIGNALING FOR NEUROGENESIS AND NEURONAL DEVELOPMENT BUT NOT SELF-RENEWAL OR MULTIPOTENTIALITY Introduction 1 Persistent neurogenesis within the postnatal brain remains a source of ongoing investigation with respect to both restorative biological processes in normal adult brain, and as a potential source of replacement cells for the treatment of injury or disease (Emsley et al. 2005) . While neurogenesis is an accepted phenomenon in both the hippocampus and the SVZ, the SVZ alone is thought to contain true NSCs ( Seaberg and van der Kooy, 2002; Seaberg and van der Kooy, 2003). In vivo investigations of the SVZ (Doetsch et al. 1999 a, b; Doetsch et al. 1997; Doetsch et al. 2002; Imura et al. 2003) suggest that neurogenesis is recapitulated postnatally via the asymmetric division of quiescent subependymal astrocytic NSCs (‘B cells’) which generate mitotically active, developmentally intermediate cells (‘C cells’) that transit the rostral migratory stream while dividing further to yield committed neuroblasts (‘A cells’) that integrate as granule or periglomerular cell interneurons (Luskin, 1993) . NSCs in the SVZ are maintained and actively divide throughout life (Gates et al. 1995; Thomas et al. 1996; Tropepe et al. 1997), generating large numbers of progeny capable of long-term morphological and functional integration (Belluzzi et al. 2003). Despite the lifelong presence of NSCs in the 1 1 This chapter contains directly excerpted text and figures from Walton et al, 2006b.

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78 ventricular neuraxis, a progressive reduction in neurogenesis accompanies aging (Tropepe et al. 1997), suggesting underlying changes in number or function of NSCs or supportive cells essential to neurogenesis. Elucidation of the mechanisms governing NSC behaviors is hampered by several factors, notably the lack of an appropriate approach allowing for detailed concurrent evaluation of NSC maintenance and generation of committed progeny. While supplying a native environment for studying multifactorial contributions to NSC maintenance and differentiation, the inaccessible nature of the neuropoietic niches makes it difficult to identify, separate, and quantify NSCs, their intermediates, and their progeny, severely limiting the identification and quantification of factors contributing to the regulation of postnatal neurogenesis (Morshead and van der Kooy, 2004; Seaberg and van der Kooy, 2003). In vitro analyses of NSC populations frequently rely on post hoc identification of NSCs from clonally-derived multipotent neurospheres (Reynolds and Weiss, 1992). While allowing both quantifiable analysis of NSC number and activity, several relevant aspects to the study of native NSCs are lost using this approach, including multicellular environment (including attendant cell-cell contacts, paracrine/endocrine factor secretion and interaction) and the ability to evaluate non-NSC contribution in NSC maintenance and differentiation ( Morshead and van der Kooy, 2004). Finally, the inherent differences between the in vitro and in vivo paradigms of studying endogenous NSCs create difficulties in identifying relevant aspects to NSC function (Seaberg and van der Kooy, 2003). As a result of such technical difficulties, several pertinent questions remain regarding postnatal factors involved in NSC function. A central question in the study of stem cells remains whether constitutively dividing NSCs are bounded by the proliferative

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79 limits imposed on normal somatic cells (Hayflick, 1965) . Siginificant evidence demonstrates that embryonic and certain adult stem cells are not subject to a maximal number of divisions (Rubin, 2002), and there is increasing evidence that adult stem cells within their niche may not demonstrate the growth inhibition observed in vitro (Rubin, 2002). Invitro investigations have documented increased proliferative abilities in the SVZ (Gobbel et al. 2003) and hippocampus (Palmer et al. 1995) in rodents. However, the extent to which endogenous NSCs are dependent on mitogens and support cells is unclear. While in vitro investigations provide a straightforward approach for identification of NSCs, current in vitro assays for NSCs are unable to identify non-NSC contribution due to the clonality requirement of the assay (Reynolds and Weiss, 1992) . To address these questions, we have employed a previously-characterized adherent culture system that allows for the establishment of monolayers of multipotent astrocytes which may be induced to generate neuroblasts analogous to that observed in vivo (Scheffler et al. 2005). In this culture system, non-neurogerminal tissues enter a senescencent state following ! 15-20 PDs. However, NSC-containing tissue derived from SVZ was expandable as a gliotypic monolayer for >300 PDs with no signs of senescence or hyperplasic transformation. In spite of this expandability, SVZ-derived astrocytes exhibit a progressive attenuation in neuroblast production when induced to differentiate. This failure to generate committed progeny is not the result of depletion of endogenous NSC populations, as neurosphere-forming cells are longitudinally retained throughout culture, and produce multipotent, self-renewing neurospheres (NS). Previous studies have identified mechanisms for NSC-independent control of neurogenesis, including nonneural cell contribution regulation of neurogenesis (Shen et al. 2004; Weimann et al.

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80 2003; Wurmser et al. 2004). Application of a reductionist culture system allows measurement of persistent neural and non-neural cell types, resulting in the identification of a population of CD11b + microglial cells which are lost in a manner paralleling the appreciated loss of inducible neurogenesis. Microglial cells represent a major hematopoietic contribution to the CNS (Eglitis et al. 1997), and have previously been demonstrated as acting in hematopoiesis (Fisher et al, 1998), normal development (Scott et al. 1994), and in states of CNS health and disease (Streit, 2002). Addition of Xanthosine (XS), a nucleotide analog shown to circumvent p53-mediated cell death (Sherley, 1991) associated with the limited proliferative limits of primary neural cells (Evans et al. 2003), results in a significant extension of both microglial lifespan and neurogenic potential in SVZ-derived cells. Addition of microglia-enriched cells or microglia-conditioned medium restored neurogenesis in highly expanded SVZ-derived cultures, suggesting a soluble contribution as a mechanism of action. As microglia have been implicated in a variety of mechanisms of neuronal development and function (Polazzi and Contestabile, 2002), we compared microglial contributions of young and adult animals. Adult-derived conditioned media is less effective in reconstituting inducible neurogenesis in SVZ-derived highly expanded cultures, but is markedly more effective in promoting the rapid morphological development of axonal processes in generated neuroblasts. These findings suggest that microglia are essential to neurogenesis, and may undergo age-related alteration in function, from one facilitating neurogenesis to one promoting the development of individual neurons.

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81 Loss of Neurogenesis Accompanies Continued NSC Proliferation To investigate the effect of continued proliferation on the composition, selfrenewal, and generation of progeny in neuropoietic (periventricular tissue) and nonneuropoietic (cerebellum) brain regions, primary dissociates from 8 day old mice were established and maintained as adherent monolayers as described ( Scheffler et al., 2005). Under the influence of defined mitogenic stimuli, cells re-enter the cell cycle and are expandable as a monolayer culture (Fig. 6-1 a). When growth factors are removed from proliferating cells, subventricular, but not cerebellar dissociates differentiate into mature cell types (Fig. 6-1 a). While initial cultures contain neurons, astrocytes, and oligodendrocytes, proliferating SVZ cells (at PD3) contain primarily astrocytic immunophenotypes, including GFAP + cells containing a subset of vimentin + and nestin + cells (Fig. 6-1 b, c). A2B5 + cells are also present in culture, either with or without GFAP coexpression (Fig. 6-1 d). Proliferating cultures also express FGF receptor 1 and putative stem cell markers Prominin-1 and Sox-1, 2, and 3 (data not shown). Bromodeoxyuridine (BrDU) labeling in proliferating cultures demonstrates that no neuronal or oligodendroglial cell types are generated or proliferative-active at PD3 (data not shown). Transit-amplifying ‘C cells’, distinguishable by a Dlx2 + /PSA-NCAM antigen profile are present 1-2 days following induction of differentiation (Fig. 6-1 e-g). These cells give rise to oligodendrocytes and PSA-NCAM + /TuJ1 + neuroblasts within 3 days of differentiation (Fig. 6-1 e, i). Neuroblasts generated in this manner exhibit increasingly elaborate neuritic arborization (Fig. 6-1 i) and express interneuron markers and

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82 morphology by 14 days following differentiation (Fig. 6-1 j). To determine whether this differentiation was a terminal event, SVZ dissociates induced to differentiate for 7 days were placed in proliferative medium for 3 days, then induced to differentiate as described. Proliferative media did not restore previous growth rates in SVZ-derived cells, and was unsuccessful in priming cells for generation of additional committed phenotypes (as defined by BrDU labeling 3d following subsequent growth factor withdrawal) following differentiation. Parietal neocortex from age-matched animals fails to yield significant numbers of committed cell types when cultured and induced to differentiate under identical conditions. We next investigated the ability of endogenous NSCs to self-renew, retain NSC characteristics, and recapitulate neurogenesis as reported in vivo. Matched cultures of SVZ and parietal neocortex dissociates were expanded 1:2 in defined proliferative medium in monolayer cultures, and parallel cultures were differentiated as described following each PD. Multiple lines of SVZ-derived primary tissue display continual logarithmic expansion kinetics with an average doubling time of 3.72+0.69 days, and were expanded >330 PDs without detection of a maximum number of cell divisions (Fig. 6-2 a). However, identically cultured cerebellar-derived cultures remain mitotically limited, typically achieving a maximum of 20 PDs before cessation of growth (Fig. 6-2 b) accompanied by the expression of senescence-associated " -galactosidase ( Dimri et al, 1995). SVZ dissociates cultured >300 PDs maintained a stable morphology and a constant, contact-inhibited growth rate over time (Fig. 6-2 c), and ceased division following addition of the antimitotic agent aphidicolin to culture media (data not shown). Giant and/or multinucleated cells, characteristic of transformed cells, were rarely detected

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83 in culture. When growth factors are removed from extensively passaged proliferating cells, cells immediately enter a period of growth arrest, and become unviable (Fig. 6-2 d). SVZ dissociates were induced to differentiate as described (Scheffler et al., 2005) each passage, and evaluated for neuroblast production 3 days later. Quantification of neuroblast production demonstrates a progressive attenuation in Tuj1 + neuroblast production in proliferative-active cultures (Fig. 6-2 e), eventually resulting in a complete abrogation in neuroblast production by 10 PDs (Fig. 6-2 f). Oligodendrocytes are generated at an extremely low frequency and do not increase with time in culture. These findings suggest SVZ-derived neuropoietic cells retain a unique ability to continue proliferation, but lose the ability to generate committed progeny with continued expansion in culture. Loss of Inducible Neurogenesis is not Linked to NSC Depletion To evaluate whether multipotent, self-renewing NSCs are maintained in proliferating primary cultures, SVZ astrocytes at 3 and 300 PDs in defined proliferative conditions were challenged in the neurosphere assay. Both PD3 and 300 astrocytes formed primary and secondary neurospheres in similar ratios (Fig. 6-3 a, b). Primary and secondary sphere diameters did not significantly differ between neurospheres derived from primary or extensively expanded astrocytes (Fig. 6-3 b, c), suggesting uniform proliferative potential. NSC proliferation has been reported to be dependent on the production of a glycosylased form of cystatin C (Taupin et al. 2000). Immunoprecipitation of PD3 and 300 SVZ-derived cells express equivalent levels

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84 cystatin C (Fig. 6-3 d). Nestin has also been reported as a key factor in the generation of primary and secondary NS formation ( Mignone et al. 2004). Using a transgenic mouse expressing enhanced green fluorescent protein (eGFP) under the nestin promoter (Mignone et al. 2004), nestin was found to be ubiquitously present in PD3-derived neurospheres (212/212 primary NS examined) and PD300-derived neurospheres (288/288 primary NS examined, Fig. 6-3 e). To assess multipotency in neurospheres formed from PD3 and 300 SVZ-derived cells, neurospheres were attached to coverslips and differentiated as described. Adherent neurospheres from both populations initially contain a mixture of GFAP + and/or TuJ1 + cells (Fig. 6-4 a, b). When induced to differentiate, NS from both 3 and 300 PDs age groups yield PSA-NCAM + /TuJ1 + neuroblasts and CNPase + progenitor cells (Fig. 6-4 c-h). These findings indicate multipotent, self-renewing cells matching the classical definition of NSCs are longitudinally maintained in culture. However, the ability of these cells to recapitulate organotypic neurogenesis is gradually lost with continued proliferation in culture, and the generation of multiple lineages may not reflect true niche-specific neurogenesis. To examine the effects of non-neural cell types on neurogenesis in an analogous antiadhesive culture conditions, we compared the generation of progeny from clonally isolated neurospheres and multiclonal aggregation neurospheres, which contain variegated, niche-specific cell types (including cells of hematopoietic origin). While both monoclonal and polyclonal neurospheres generate neuronal cell types, only polyclonal spheres yielded characteristic A cells (Fig. 6-5), suggesting included non-neural cell types were essential for organotypic neurogenesis.

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85 Microglia Depletion Correlates with Loss of Inducible Neurogenesis Loss of a non-NSC cell type(s) from culture over time may be an underlying factor in the appreciated loss of neurogenesis accompanying continued proliferation. To identify such populations, we immunophenotyped 3 and 300 PD SVZ cells for a variety of markers correlating to neural and non-neural phenotypic markers (Fig. 6-6 a). Highly expanded populations display nonsignificant reductions in GFAP, nestin, and A2B5 expression over time (Fig. 6-6 a). Both nestin + and A2B5 + /nestin + cells, which have been described as potential NSCs (Lendahl et al. 1990; Scheffler et al. 2005), are retained in culture in ratios similar to the percentage of NS-forming cells in highly expanded cultures. GFAP + cells have been reported as being the NSC of the adult mammalian brain (Doetsch et al. 1999; Imura et al. 2003; Laywell et al. 2000), and were retained in culture in frequencies approximating those of primary neurosphere-forming cells (5.7+3.7 vs 6.63 + 0.64% respectively). Putative NSC markers Sox2 and Prominin-1 were examined, and were present in both PD3 and 300 cultures (data not shown). CD15 and NG2 expression in proliferating cells was also examined, and are expressed in frequencies insufficient (<0.01%) to account for the appreciated production of neurospheres. Cells of nonneural origin, including endothelial cells, have been implicated as having the capacity to influence the proliferation, neurogenicity, and fate choice of neural stem cells through soluble factor influence (Shen et al. 2004) , direct contact (Wurmser et al., 2004), or fusion (Weimann et al. 2003) . To examine contributions from cells of hematopoietic origin, we examined cultures for

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86 panhematopoietic lineage marker CD45 and microglial marker CD11b. While CD45 + /CD11b cells are initially present at very low frequency (<0.01%), CD11b + cells are initially expressed in a subset of cultured cells, but are not present in extensively expanded cells (Fig. 6-6 a). To examine whether microglial loss mirrors a loss of neurogenic potential in SVZ-derived cultured cells, we measured the prevalence of microglia thoughout culture. In proliferative cultures, microglia are lost from culture at a rate that closely parallels the observed loss of neurogenic potential in proliferative-active SVZ cultures (Fig. 6-6 b,c). Microglia Rescue Inducible Neurogenesis To calculate the presence of microglia to the neurogenic potential of NSCs in vitro, XS was added to proliferating SVZ dissociates. Addition of XS to proliferating cultures was able to extend the duration of microglial lifespan in culture conditions (Fig. 6-6 b), with a concomitant extension of neurogenic lifespan in vitro. However, microglial cell types were permanently eliminated from culture following 20 PDs regardless of supplementation. XS treatment has no effect on the proliferation rate of either neuropoietic (Fig. 6-7 a) or non-neuropoietic (cerebellar, data not shown) tissues, nor is it effective in reconstituting neuroblast generation in cells expanded beyond 20 PDs in defined proliferative media. XS treatment does not significantly alter the frequency of primary or secondary NS formation in primary or extensively expanded tissues (Fig. 6-7 b,c), suggesting XS does not act upon NSC self-renewal.

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87 To examine the conditions underlying the depletion of microglial cells, CD11b + cells were measured for markers of apoptosis and necrosis throughout the initial period of expansion. Microglia display consistent, infrequent apoptotic and necrotic cells, \which do not exceed 3.5% of total number of microglial cell number (Fig. 6-8 a). As XS application was successful in extending microglial lifespan, it is likely a proliferationassociated mechanism ultimately limits the proliferative abilities of microglia. Examination of the proliferative marker Ki-67 in passage 3 cultures reveals a reduced expression in microglial cells (Fig. 6-8 b-d), suggesting a reduced rate of proliferation underlies microglial depletion. To analyze the means by which microglia effect a contribution to neurogenesis, the distribution and number of microglia were examined throughout differentiation in 3 PD SVZ cultures. Microglia exhibit a uniform distribution, both in vivo (Cuadros and Navascues, 2001) and in vitro, and do not appear to be closely associated with either astrocytes (Fig. 6-9 a) or clusters of developing neuroblasts (Fig. 6-9 b), suggesting a cell contact-independent mechanism of action. Microglial prevalence and morphology do not undergo significant alteration in response to mitogenic withdrawal-initiated terminal differentiation, suggesting they are relatively insensitive to mitogenic alteration of culture conditions necessary for NSC differentiation (Fig. 6-10 a). Hematopoietic cells have been implicated in cell fusion, which have been questioned as a potential mechanism in the generation of committed cell types (Terada et al. 2002), alteration of cellular plasticity (Ying et al. 2002), and as a means of non-neural cellular contribution to neurogenic cells (Weimann et al. 2003). To determine if microglial presence has an effect on the neurogenesis of subventricular dissociates,

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88 eGFP-expressing cells from PD3 cerebellar dissociates were added to C57/B6 300 PD tissues. CD11b-enriched populations were able to rescue neurogenesis in CD11bdepleted subventricular tissues (Fig. 6-9 c,d). No neuronal phenotypes were generated that expressed eGFP, suggesting that fusion does not occur in cocultures (Fig. 6-9 e). Based on the apparent lack of fusion and microglial contact before and during differentiation, it appears microglia may act through a soluble factor. Conditioned medium from eGFP-expressing PD3 cells from SVZ or cerebellum was added to extensively expanded SVZ cells. Conditioned medium from both PD3 SVZ and cerebellar dissociates was effective in restoring neurogenesis in adherent cultures (Fig. 69 f). Interestingly, conditioned medium derived from cerebellar tissue (which contained larger numbers of microglia compared to ageand culture-matched SVZ, data not shown) is markedly more effective in eliciting neurogenesis (Fig. 6-9 f). To establish that microglia, and not another unspecified cell type are essential to neurogenesis, microglia were selectively ablated from PD3 SVZ dissociates using a saprotoxin-conjugated antibody to CD11b (Fig. 6-9 g). Significant microglial depletion was appreciated following a minimum of 18 hours of conjugated antibody application (Fig. 6-9 g), which was not observed in untreated or unconjugated saporin application. Proliferating cultures, which maintain Ki-67 expression throughout targeted toxin application (Fig. 6-10 b), and microglia do not undergo significant proliferation in response to selective ablation or saporin presence (Fig. 6-10 a). Following incubation with targeted toxin, SVZ dissociates were induced to differentiate as described (Fig. 6-9 h), which results in significant declines in neuroblast production in microglial-depleted cultures but not untreated or unconjugated antibody culture.

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89 In vivo neurogenesis occurs throughout life, albeit at reduced rates with increasing age (Tropepe et al. 1997). As microglia have been implicated in a variety of mechanisms associated with aging and senescence, we examined the changes in microglial abundance in early postnatal (P8) and adult (P90) mice. Early postnatal and adult animals contain equivalent numbers of microglial cells (Fig. 6-11 a). However, conditioned medium from adult animals is unable to rescue neurogenesis to the extent observed from that of early postnatal animals (Fig. 6-11 b). In addition to putatively facilitating neurogenesis, microglial cell types are implicated in a variety of paradigms for ongoing neuronal maintenance (i.e., axon sprouting, survival). To determine if agedependent differences in microglia-driven neuronal development existed, we tracked alterations in neuronal morphology in developing neuroblasts following their coculture with microglial-enriched eGFP cells from early postnatal and adult animals. Regardless of age of neuroblast-generating tissues, neuroblasts generated far longer processes than those maintained in conditioned medium from younger animals (Fig. 6-11, middle panel). To examine whether this alteration in microglial support of neurogenesis was purely agespecific or was affected by regional specification, we evaluated both SVZ and cerebellar conditioned media to identify differences in neuronal development (Fig. 6-11, middle panel). No differences were appreciated between regions employed, suggesting global changes occur in microglia to alter their function from neurogenesis-enabling to neuronodevelopmental.

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90 Fig. 6-1. An adherent culture model allows cultivation and inducible neurogenesis in an SVZ-derived astrocyte monolayer. (a) Dissociated periventricular tissue (red and blue area) was expanded in culture. Removal of growth factors (EGF, bFGF) and serum from culture media induces differentiation, yielding defined clusters of neuronal progenitors. (b-d) Undifferentiated (PD3) SVZ tissues contain predominantly astrocytic (GFAP + , vimentin + ) cells, with subsets displaying immature markers (nestin and/or A2B5). (e-g) Differentiating SVZ dissociates phenotypically and temporally recapitulate SVZ neurogenesis, displaying Dlx-2 + ‘C Cells’ (arrowhead) and Dlx-2 + /PSANCAM + ‘A Cells’ 24-48 hrs later. Following differentiation, mature cell types emerge, including oligodendrocytes (h), astrocytes (i), and neurons (i) that mature into GAD65/67 + interneurons (j). Scale bars 50 m (a, b-d, e-g, h, i), 25 m (j).

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91 Fig. 6-2. Neuropoietic tissues are indefinitely expandable as astrocyte monolayers, but lose mitogen-withdrawal prompted neurogenic potential. (a) SVZ-derived dissociates exhibit long-term logarithmic expansion, which is not matched by cerebellar cultures (b). (c) mitotic rate (average relative increase in cell #/day) for SVZ cells between 1-10 and 300-310 PDs (photo inset). (d) SVZ cells (>300 PDs) remain growth dependent on N2 proliferative media (prol) supplemented with EGF/FGF (EF), and/or serum (ser). (e) A progressive reduction in mitogen withdrawal-induced neuroblasts (small, phase dark cells) accompanies continued proliferation. (f) SVZ dissociates fail to generate neuroblasts by 10 PDs in defined proliferative conditions. Scale bar 75 m (e).

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92 Fig. 6-3. Expanded cultures maintain neurosphere-forming cells. PD3 and PD300 SVZ dissociates contain similar numbers of neurosphere-forming cells that do not differ in number (a), appearance (b), or size (relative measurement of sphere diameter, c). (d) 3 and 300 PD dissociates from neuropoietic zones express equivalent amounts of both glycosylated (21 kD) and non-glycosylated (17 kD) cystatin C. (e) Highly expanded dissociates continue to ubiquitously express nestin in clonally-derived NS. Scale bars 100 m (b), 75 m (d).

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93 Fig. 6-4. Neurospheres derived from PD3 and PD300 SVZ cells are multipotent. Neurospheres from PD3 (a) and PD300 (b) populations were composed of phenotypically intermediate cells that display astrocytic and neuronal markers. (c) Upon mitogenic withdrawal, Tuj1 + /nestin + neuroblasts are generated. (d) maturing neurons (boxed area from c) develop an elaborate neuritic processes and lose nestin expression following generation. Neurospheres from 3 (e) and 300 (f) PD SVZ are capable of generating multilineage progeny, including PSA-NCAM + neurons (g), GFAP + astrocytes (g, h), and CNPase + oligodendrocyte progenitors (h, 300 PD shown). Scale bars 50 m (d, g), 100 m (a-c, e-f, h).

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94 Fig. 6-5. Non-neurogenic cells promote neuroblast generation in antiadhesive culture systems. Comparison of monoclonal (single cell-derived, left column) neurospheres and polyclonal (high density-aggregation, right column) neurospheres. Monoclonal and polyclonal neuropheres were formed from primary tissue at low and high density respectively (see materials and methods) under otherwise identical culture conditions. When induced to differentiate, only polyclonal neurospheres produce morphologically distinctive A cells (highlighted area, magnified in inset).

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95 Fig. 6-6. Microglia depletion parallels loss of inducible neurogenesis in proliferating cultures. (a) The frequency of expression of a variety of markers associated with endogenous neural stem cells, pan-hematopoietic markers, and monocyte/macrophage lineages reveals a complete depletion of microglial (CD11b + ) cells in PD300 astrocyte monolayers. (b) The majority of microglial cell loss occurs within 10 PDs in culture. XS supplementation extends microglial lifespan (b) and neurogenic potential (c) in vitro. Microglial loss mirrors reduction in neuroblast generation from matched cultures. *p<0.05, one-way ANOVA, for PD 9-21 (c).

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96 Fig. 6-7. XS has negligible effects on proliferative rate and NSC frequency. (a) total estimated number (circles) and total population doublings (triangles) for culture-matched SVZ dissociates with (orange) or without (blue) XS supplementation. Relative frequency of primary (b) and secondary (c) NS generation in 3 and 300 PD SVZ expanded with or without XS supplementation.

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97 Fig. 6-8. (a) Levels of apoptosis and necrosis in microglia remain constant throughout differentiation. (b) PD3 CD11b + cells display reduced Ki-67 expression when compared to CD11b cells. (c) Microglial cells proliferate in culture (Ki-67 expression, arrow), but are eventually lost from cultures (d). Scale bar 50 m (c,d). *p<0.05, one-way ANOVA.

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98 Fig. 6-9. Microglial cells modulate inducible neurogenesis through a soluble factor(s). Microglia present prior to (a) and 3d following differentiation (b) in PD3 SVZ monolayers appear to assume reactive morphologies and do not form close associations with proliferative-active astrocytes (a) or neuroblasts (b). (c) Coculture of eGFP

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99 expressing PD3 cerebellar cells (eGFP-CB) with C57/B6 PD300 SVZ cells rescues inducible neurogenesis to the astrocyte ! neuroblast ! neuron transition appreciated in vivo. No eGFP neuroblasts (d) or neurons (e) are generated, suggesting no microglia fusion-mediated cellular recombination occurs. (f) Addition of conditioned medium from SVZ (+ SVZ CM) or cerebellum (+CB CM) is effective in restoring inducible neurogenesis in microglia-depleted (300 PD) SVZ dissociates. (g) Microglia are selectively ablated by saporin-conjugated CD11b/MAC1 antibody (MAC1-SAP) but not unconjugated saporin (uncon-SAP). (h) Inducible neurogenesis is reduced in immunotoxin treated cultures, as evidenced by Tuj1 + neuroblast production 3d following differentiation of microglia-depleted cultures. Scale bars 50 m (a-b, e), 25 m (d). *p<0.05, one-way ANOVA. . Fig. 6-10. (a) CD11b/MAC1-conjugated (MAC1-SAP) and unconjugated (uncon-SAP) saporin toxin application to proliferating and differentiating cultures does not affect microglial depletion or proliferation (measured by % cells CD11b + ). (b) Addition of conjugated or unconjugated toxins to proliferating cultures for 24 hours does not affect overall proliferation rates (% total cells Ki-67 + ).

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100 Fig. 6-11. Microglial undergo age-dependent alterations in neurogenic support and neuronal development. (a) PD3 cerebellar dissociates from postnatal day 8 and 90 mice contain equivalent numbers of microglia. (b) Conditioned cerebellar medium from adult animals was less able to rescue neurogenesis to the extent observed in early postnatal animals. (c) 7 days after differentiation, Tuj1 + neurons derived from PD300 SVZ

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101 dissociates with conditioned medium from adult (c, left panel) and early postnatal (c, right panel) animals. Representative camera lucida neuron etchings from PD3 young and adult animals demonstrate regionally-insensitive, age-dependent effect on axonal outgrowth (c, central panel). Scale bar 50 ! m (c). *p<0.05, one-way ANOVA.

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102 CHAPTER 7 DERIVATION AND LARGE-SCALE EXPANSION OF MULTIPOTENT ASTROGLIAL NEURAL PROGENITORS FROM ADULT HUMAN BRAIN Introduction 1 Significant attention has been focused on the development of primary human neural tissue sources for multiple applications in the central and peripheral nervous system (CNS, PNS). Several techniques exist for generating such cell types, including cultivation of multipotent neurogenic progenitors from developing brain (Carpenter et al. 1999; Vescovi et al. 1999; Ostenfeld et al. 2000; Zhang et al. 2001). Despite their promise in generating variegated cell types for therapeutic application, pluripotent cell sources are beset by ethical concerns surrounding their use, as well as concerns over potential tumor formation (Odorico et al. 2001) and immunorejection (Martin et al. 2005). Additionally, primary multipotent cells from either embryonic or postnatal sources produce primarily heterogeneous mixtures of committed cells, providing potential concerns over the generation of pure populations of cells for type-specific transplant or bioassay applications ( Schuldiner et al. 2000; Shamblott et al. 2001) . This is particularly relevant, as many investigations focus on the derivation of committed neuronal cell types, and do not address the concomitant need for committed glial cells. Primary cells derived from adult brain may provide committed cell types which are developmentally and immunologically matched for transplantation or other biological 1 This chapter contains figures and text directly excerpted from Walton et al, 2006a.

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103 assays when expanded as proliferating precursor populations in vitro. However, a significant barrier to the use of normal somatic cells is an intrinsic lack of sustainable ex vivo mitosis in culture (Kiyono et al. 1998; Evans et al. 2003) . Characterization of clonally expanded multipotent human progenitors reveals similar proliferative limits to those appreciated in somatic neural cell types (Nunes et al. 2003). Despite these observations, several recent findings suggest that barriers to expansion of postnatal progenitor populations may be more flexible than previously believed. The appreciated lack of expandability of primary human cells has been linked to the cell cycle arrest and entry into senescence via activation of cyclin-dependent kinase inhibitor p21 WAF1 (and subsequent activation of the p16 INK4A ) pathway, which has been reported to initially arrest growth of cultured astrocytes after ! 20 population doublings ( PDs) (Evans et al. 2003) . Recent reports describing culture of normal rodent glia indicate that the static upper boundary for maximal cell divisions is more flexible than previously imagined (Mathon et al. 2001; Tang et al. 2001), and may be circumventable using appropriate culture conditions. A growing body of evidence indicates a disparity in replicative competency of cells in vivo compared to in vitro , particularly for stem cells of high turnover organ systems (Rubin, 2002) , theoretically allowing the extensive expansion of endogenous progenitor populations upon the application of correct growth criteria. The lack of telomerase expression in non-neurogenic regions of human brain may also be a limiting factor in the long-term expansion of widely distributed progenitor populations. Catalytic telomerase (hTERT) is believed to play a critical role in maintaining telomere length, and has been related to lifespan in a variety of human tissues. Though hTERT expression has been reported in the neuropoietic regions of adult rodents (Caporaso et al. 2003), only

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104 low levels of telomerase have been reported in ex vivo cultured human cells (Ostenfeld et al. 2000). However, telomerase has been detected in germ cells (Kim et al. 1994) and hematopoietic cells in vivo (Broccoli et al. 1995; Counter et al. 1995; Hiyama et al. 1995), and can be attenuated by physiological alterations endometrium cells (Kyo et al. 1997), suggesting it may possible to induce and/or longitudinally maintain telomerase expression in human cell populations under appropriate conditions. Glial cells comprise the majority of CNS cell types, and are increasingly recognized for their role in injury (i.e., glial scar formation (Silver and Miller, 2004)) and as a potential tool for treatment of neurological disease (i.e., secretion of neuroprotective factors ( Kordower, 2003; Tai YT, 2004)). Although some glial cells have been implicated to represent endogenous NSCs in rodents (Doetsch et al. 1999; Laywell et al. 2000; Doetsch, 2003; Scheffler et al. 2005), there exists limited characterization of neurogenic cells in the adult human brain (Nunes et al. 2003) . To examine the feasibility of deriving and expanding endogenous adult human neural progenitors, we applied growth conditions favoring the propagation of gliotypic rodent neural progenitor/stem cells in dissociated monolayer culture ( Scheffler et al. 2005) to both neurogenic and nonneurogenic regions of postnatal human brain. Using these conditions, a single population of highly expandable neural progenitors from multiple forebrain regions was isolated and maintained as a homogenous population in vitro. These cells retain morphologies consistent with type I astrocytes in culture and display multiple progenitor markers. Furthermore, these cells are highly expandable, and can be maintained for over 300 days and >60 PDs with minimal signs of senescence or immortalizing mutations. Interestingly, progenitors derived from neurogenic and non-neurogenic regions express

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105 telomerase, the continued expression of which appears to be linked to a synergistic mitogenic effect and is directly coupled to the continued growth of adult human neural progenitors (AHNPs). To test the ability of ex vivo gliotypic cells to incorporate into the CNS, expanded cells were engrafted in the ventricle and cortex of early postnatal and adult rodents and examined longitudinally for survival, integration, distribution, and fate choice. Implanted cells effectively incorporate in a variety of host brain regions and adopt both neuronal and glial phenotypes. Stable and long-term genetic modification of AHNPs was achieved using both transient and long-term transfection approaches. Finally, it was possible to preferentially differentiate AHNPs to rapidly generate neuronal cell types in vitro. These findings suggest a means for rapid expansion of uniform, transplantable, and genetically modifiable multipotent human neural progenitor populations, allowing for new applications of readily obtainable postmortem or autologous cell sources. Characterization and Expansion of Primary Cells as AHNPs Human brain tissue (anterolateral temporal lobe) was derived from patients undergoing resection associated with medically intractable epilepsy. Tissue was microdissected into regions containing hippocampus, SVZ or temporal cortex gray matter (Fig. 7-1). Isolated dissociates were maintained as a monolayer on uncoated plastic dishes throughout culture in defined proliferative media (described in Fig. 7-2), modified from a standard protocol for the culture of NSCs (Scheffler et al. 2005). To identify cultured cell types, primary cells were examined for expression of phenotypic markers. Following dissociation, immunocytochemistry on primary cells 3

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106 days in vitro (DIV) revealed a heterogeneous population containing predominantly astrotypic (GFAP + ) cells, but included neuronal (NeuN + , PSA-NCAM + ) and oligodendrocyte (CNPase + , O4 + ) phenotypes. Nestin + cells, generally displaying a protoplasmic morphology were initially present in early cultures. Following expansion in defined growth medium, a population of progenitors is established as the sole proliferating population by 14 DIV. These cells are defined by the conserved expression of nestin (Fig. 7-3 a), with the retention of the morphological and antigenic properties ascribed to type I protoplasmic astrocytes (Norenberg and Martinez-Hernandez, 1979; Raju et al. 1980; Cammer and Tansey, 1988). Cells present 14 DIV frequently coexpress both immature and astrotypic markers, including A2B5 (97.2 +/1.3%), nestin (99.8 +/0.1), NG2 (96.7 +/2.2%), GFAP (95.4 +/3.2), S100 " (89.8 +/4.1), and glutamine synthetase (90.4 +/4.4) (% positive +/S.E.M.) (Fig. 7-3 a-d,g). To further characterize these cells, we performed single cell patch clamp recordings for highly expanded (30 PDs) cells (n=4). Recorded cells exhibited ubiquitous gliotypic membrane potentials (Sontheimer, 1994), with an RMP of .3 +/4.2 mV, C m of 277.2 +/189.7 pF, R m of 214.5 +/156.1 M # , and R a of 14.9 +/3.1 M # . Recorded cells did not fire action potentials, but displayed prominent Na + channel activity and K + channel activity (Fig. 7-3 e). To determine the composition and dynamics of proliferating populations, cells undergoing 10, 20, and 30 PDs were cultured in the presence of the thymidine analog BrDU. In our culture conditions, only nestin + cells appear to re-enter the cell cycle as shown by their rapid increase in prevalence (55.2+/-17.2% at 3 days in culture vs 99.7+/0.2% at 30 days in culture) and near-ubiquitous incorporation of BrDU throughout culture (average 98.9+/-0.8% (nestin + /BrDU + )/BrDU + for 10, 20, and 30 PD populations

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107 following 48 hour BrDU administration). This finding was confirmed through appreciation of extensive intermediate filaments in cells subjected to ultrastructural examination (data not shown). Furthermore, the rate at which BrDU increases in culture is contiguous with the known doubling rate of expandable populations derived from multiple forebrain regions (Fig. 7-3 h). In the absence of defined growth factors, BrDU incorporation is rapidly attenuated in nestin + cells, which accompanies a cessation of growth (Fig. 7-3 h). Neurons (PSA-NCAM + ) and oligodendrocytes ( CNPase + ) were not appreciated in proliferating culture conditions after 14 DIV. Cells displaying a stellate or reactive morphology were rarely detected in culture. Microglia (CD11b + ) were present initially and did not significantly decline upon continued culture. Selecting for and proliferating unattached cells 12 hrs after initial plating decreased microglial presence in culture to nearly undetectable levels. FACScan analysis of 30 PD cell cycle revealed a single proliferating population with minimal side scatter (data not shown). Immature astrocyte-like cells have been implicated as “immortal” neural stemlike cells maintained throughout life in the hippocampus and subventricular zone (Laywell et al. 2000; Seri et al. 2001; Sanai et al. 2004). In rodents and humans, these cells have been described as existing throughout life (Tropepe et al. 1997; Sanai et al. 2004) and cells cultured from these regions may represent a system-specific stem cell population which can be expanded beyond the limits ascribed to somatic cells (Potten and Morris, 1988). To detect potential NSCs, proliferating temporal cortex and hippocampal astrocytes were clonally seeded and assayed for neurosphere formation as described (Kukekov et al. 1999) every fifth passage (n=3 wells/assay). Cultured cells fail to generate multipotent neurospheres at clonal seeding densities at any point, suggesting

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108 they were a separate population from neurosphere-forming cells. Despite a lack of proliferation in neurosphere-forming conditions, clonally seeded cells remained viable for up to 14 days, and were expandable as adherent cultures following substrate reattachment. Cells isolated in this manner were expandable as individual clones (153/182 clones examined for a minimum of 5 PDs). To assess whether clonally isolated cells recapitulate the progenitor populations isolated from initial derivation, low density adherent cultures (500 cells/cm 2 from primary and 10 PD temporal cortex) were created from both adherent cells and cells surviving 14 days under clonal antiadhesive conditions. Cells derived from both culture conditions ubiquitously express nestin (>99.9%), with a large subset (similar to previously described for initial derivation) expressing GFAP (Fig. 7-3 g). Similarly, addition of BrDU immediately following attachment resulted in the progressive increase in BrDU incorporation to >99% within 72 hours (Fig. 7-3 h), suggesting virtually all cells maintain a constitutively proliferative state. In both initial and clonal derivations, BrDU incorporation was limited to nestin + cells indicating a homogenous population of proliferating progenitors with frequent astrotypic immunophenotypes compose the proliferating culture, rather than a heterogeneous culture composed of proliferating lineage-negative proliferating progenitors which generate postmitotic, but immature astroglial cells. Removal of mitogenic stimuli results in a progressive attenuation of BrDU incorporation and subsequent failure to thrive (Fig. 7-3 h). However, nestin expression remains in >99% of cells regardless of mitogenic presence, suggesting a single, mitogen-dependent progenitor population comprises expanding cultures. Thymidine analog incorporation in GFAP + cells does not significantly differ from that of nestin + cells, suggesting no additional

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109 proliferative ability of GFAP + /nestin cells. Based on these observations it appears we have established conditions for the isolation of a unique population of widely distributed, highly expandable forebrain cells. To determine the proliferative limits for purified gliotypic cells in these conditions, cells were grown in vitro and their expansion quantified via cell counting. To ensure a homogenous population for measurement of proliferation, neural cell dissociates were grown continuously for 30 days prior to quantification of growth ( ! 10 PDs) to remove postmitotic cells. Following this initial culture period, 10 6 cells from hippocampus and temporal cortex were plated in defined proliferative media and supplemented with EGF and bFGF bidaily. Proliferating cultures derived from multiple regions maintain a constant contact-inhibited growth rate for both hippocampaland temporal cortex-derived cells (0.34 +/0.04 and 0.35 +/0.04 doublings/day respectively) (Fig. 7-3 i), while retaining constant morphology and size throughout culture (Fig. 7-3 j). Upon reaching confluency, cultured cells were passaged 1:2 and total cell number counted. Both temporal cortex, subventricular, and hippocampal astrocytes exhibited logarithmic growth expansion in defined growth medium for over 300 DIV, with a maximal expansion of > 60 PDs (Fig. 7-3 k), equivalent to one cell giving rise to >10 16 cells. AHNPs Maintain Growth Sensitivity and Avoid Immortalization Purified expanding cell populations may undergo growth-specific genetic modification(s) resulting in circumvention of cell cycle regulatory mechanisms and manifesting in an immortalized phenotype, allowing for extensive clonal expansion

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110 similar to that observed. Immortalized cells frequently contain accumulated neoplastic mutations in genes linked to cell cycle control, apoptosis, and survival, and may be characterized by a lack of response to physiological or chemical arrestors of the cell cycle. Furthermore, transformed cells often exhibit irregular or hyperplasic growth rates, and can be tumorogenic when transplanted. To determine whether such immortalizing mutations were present in AHNPs, we examined the molecular and cytogenetic profiles of expanded populations. Immortalization of human cells is frequently marked by the aberrant expression of key regulatory proteins. To determine the activation status of cell cycle proteins in expanding cells, protein expression levels for major cell cycle regulatory proteins were measured throughout the culture period (Fig. 7-4 a). AHNPs longitudinally express major cell cycle checkpoints, including p53, a key initiator of cellular senescence. Expanding AHNPs also express p16, the deletion of which is reported to be essential for immortalization in both epithelial cells (Kiyono et al., 1998) and astrocytes (Evans et al. 2003) in humans (Fig. 7-4 a). Though p53 remains constant throughout culture, other cyclin-dependent kinase inhibitors (i.e., p21) and cyclins (i.e., cyclin E) increased throughout the culture period. This observation agrees with noted increases in both promitotic and inhibitory proteins during the extended culture of glial progenitors in rodents (Mathon et al. 2001; Tang et al. 2001). Interestingly, robust hTERT expression was appreciated in cultured cells initially, matching a report of initial expression of telomerase in cultured fetal human brain tissue (Ostenfeld et al. 2000). hTERT is expressed at progressively lower levels during expansion in defined proliferative conditions. None of the populations examined (n=6 from 4 individuals) was capable of

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111 indefinite growth (avg 62.1+/-2.31 PDs, range 59-65). Karyotypic analysis of metaphasic high passage cells (n=7) revealed no gross cytogenetic abnormalities in highly expanded cells (Fig. 7-4 b). To determine whether AHNPs remain sensitive to chemical and physiological regulators of the cell cycle, highly expanded AHNPs were treated with the DNA synthesis inhibitor aphidicolin or X-irradiation and were assessed for the senescent cell marker SA" -gal (Dimri et al., 1995) (Fig. 7-4 d,e), 7 days later. Treated cells expressed significantly higher levels of SA" -gal than age-matched controls, suggesting cell cycle checkpoint mechanisms remain sensitive throughout culture period. Telomerase, the holoenzyme responsible for telomeric extension, is longitudinally expressed during the observed period of growth. This is unique, as telomerase is rarely reported in adult neural tissues, and its expression and loss may be related to the continuing expandability of cells in culture. To investigate the relationship between telomerase expression and continued growth in culture, the telomerase inhibitor epigallocatechin-3-gallate (Naasani et al., 1998) was added to highly expanded cells, and telomerase expression and growth rate were measured 7 days later (Fig. 7-4 c-e). Despite significant reduction in the rate of cellular proliferation, hTERT expression remained ubiquitous in expanding AHNPs (Fig. 7-4 c). To further examine the potential relationship between growth conditions, telomerase expression, and expandability in AHNPs EGF, bFGF, and serum were selectively removed from culture medium of highly expanded cells. Removal of EGF and/of bFGF resulted in the loss of telomerase expression within 7 days accompanied by a failure to continue to proliferate (Fig. 7-4 d, e). Interestingly, AHNPs treated with EGCG or aphidicolin returned to normal growth

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112 rates within 7 days following replating in proliferative media (Fig. 7-4 e), while cells deprived of growth factor failed to regain previous proliferative levels and subsequently became unviable (Fig. 7-4 f). Less than 4% of AHNPs exhibited SA" -gal in defined proliferative conditions, suggesting cells continue to be mitotically active and are expandable for over 60 PDs (Fig. 7-4 d). Multinucleated and/or giant cells, characteristic of senescent cells, were rarely observed at any point throughout culture period. These findings suggest cultured cells do not spontaneously immortalize, while remaining highly expandable, mitogen dependent, and telomerase-positive. Expanded AHNPs Function as a Transplantable, Modifiable Cell Source To assess the ability of AHNPs to survive, integrate, and assume a committed phenotype in vivo, 10 5 AHNPs (20-30 PDs) were injected into the right lateral ventricle of early postnatal (P3) C57/B6 mice. Engrafted cells were assessed for patterns of incorporation and immunophenotype 7 days later using human ribonuclear protein (HNA) to identify engrafted cells. Moderate reactive gliosis was appreciated in transplanted animals, which may affect overall survival of transplanted cells. Immunosupression of young animals with cyclosporin A substantially reduced reactive gliosis and increased survival and engrafted cell distribution substantially. Engrafted cells were primarily detected within the ependymal wall of the injected ventricle, with increasingly frequent distribution immediately adjacent to the injection site (Fig. 7-5 a, b). HNA + cells were also frequently detected in the choroid plexus, adjacent to the third ventricle, cerebral aqueduct, and (rarely) in the cerebellum. Immunocytochemistry

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113 revealed integrating cells frequently adopt mature neuronal morphologies. Though many ventricularly-engrafting cells did not express mature phenotype markers, engrafted cells were frequently found to coexpress GFAP (Fig. 7-5 c-f). Surviving ventricularlyengrafted cells did not coexpress neuronal markers, and rarely expressed nestin, suggesting they mature to largely postmitotic astrotypic cell types upon integration. The highly expandable nature of AHNPs suggests a potential role as a substrate for a transplantable cell source, frequently envisioned for use with single or multiple gene products in drug delivery and neuroprotective paradigms (Kordower, 2003; Tai YT, 2004). To concurrently examine the amenability of AHNPs to ex vivo genetic modification and examine the feasibility of single-gene alteration in influencing fate choice toward a neuronal phenotype, AHNPs were transfected with a plasmid containing a 2 kb gene encoding the neural patterning gene Pax6 and eGFP using both activated dendrimer transfection and non-liposomal lipid transfection. Stably transfected cells were detected at low frequency 3 days following transfection (Fig. 7-5 g,h), but Pax6 expression did not result in phenotypic alteration to neuronal fates using this approach. We also sought to evaluate the long-term transfection efficiency of viral vector transformation on expanded AHNPs. Using a lentiviral vector expressing eGFP under the SV40 promoter, we were able to identify optimal transfection conditions for expression of lentiviral vector expression in expanded populations (Fig. 7-5 i and 7-6 ac). The ability to insert therapeutically relevant genes was examined by creating a lentiviral vector expressing glial-derived neurotrophic factor (GDNF, Fig. 7-7 a-f). Human progenitors infected with this construct express detectable levels of GDNF (Fig.

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114 7-7 e) while maintaining an astrocytic identity, and exhibit increased GDNF production (Fig. 7-7 h). Human progenitors infected with a bicistronic construct expressing eGFP under the GDNF promoter resulted in detectable eGFP expression 7 days following transfection (Fig. 7-7 g). These results indicate it is possible to viably manipulate human progenitor cells with single gene products. To examine the ability of GDNF-infected human progenitor cells to effectively integrate and act as cellular dug delivery vehicles, human progenitors expressing GFP and GDNF were injected into the striatum of immunosupressed rodents (Fig. 7-8 a), and evaluated 2 and 10 days later. Both GDNF (Fig. 7-7 b, and lower panel) and GFP (Fig. 7-8 b and lower panel) expressing cells were detected following transplantation in the targeted region, suggesting these cells may be suitable as a transplantable population for drug delivery. To examine survival and integration of AHNPs in the adult CNS, AHNPs (30 PDs) were transplanted into the cortex of adult (P90) NOD-SCID immunocompromised mice. Transplanted AHNPs (HNA + ) were detectable following a 30-day engraftment period. In contrast to ventricularly engrafted cells, cortically implanted cells were found to express -III-tubulin and adopt neuronal morphologies with significant process extension (Fig. 7-9 a,c). Astrocytic (GFAP + , Fig. 7-9 b) or oligodendrocytic (CNPase + , data not shown) phenotypes were rarely detected in cortically-integrating HNA + cells. AHNPs that displayed non-neuronal immunophenotypes were infrequently detected. Transplanted AHNPs were largely concentrated around the injection site, with limited migration along the dorsal-ventral axis. Serial analysis of adjacent sections revealed modest migration of up to 250 m laterally and 400 m radially. Very infrequently,

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115 HNA + / -III-tubulin + cells were present in the CA1 and CA3 regions, which adopt morphologies and extend process characteristic of endogenous pyramidal neurons (Fig. 7-9 d,e). Engrafted cells very rarely coexpressed the proliferative cell cycle marker Ki-67 and were nestin (data not shown), suggesting they were largely postmitotic following transplantation. Transplanted cells adopt mature neuronal phenotypes, as demonstrated by the expression of NeuN in cortically transplanted HNA + cells 30 days following engraftment (Fig. 7-9 f,g). Transplanted cells exhibit modest medial-lateral migration and limited anterior-posterior migration (Fig. 7-10). Phenotypic identity appeared heavily reliant on site-specific integration, with cortically-engrafting cells tending toward a neuronal identity and ventricularly-engrafting cells adopting a more glial fate (Fig. 710). Controlled alteration and in vitro manipulation of cellular phenotype are increasingly envisaged in tissue culture paradigms. Attempts to induce in vitro differentiation in adherent cells as previously described for attached human neurospheres (Ostenfeld and Svendsen, 2004) and adherent rodent NSCs (Scheffler et al. 2005) were unsuccessful in producing multiple differentiated cell types. To further test the potential for alteration of phenotype in AHNPs, expanded cells (>20 PDs) were subjected to multiple combinations of culture supplements, including FGF-8, retinoic acid, sonic hedgehog, dibutyl cAMP, nerve growth factor (NGF), 1-isobutyl-3-methylxanthine (IBMX), retinoic acid, and serum. Application of dibutyl cAMP, NGF, and IBMX, combined with the removal of serum and growth factors, was found to induce a rapid phenotypic alteration in proliferating cells (Fig. 7-11 a) that yielded morphologically and electrophysiologically characteristic immature neurons within 7 days. When examined 3

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116 days following this induction of phenotypic alteration, a subset of cells displayed a hybridized somato-dendritic morphology intermediate to astrocyte and neuron, and displayed both neuronal and astrocytic markers (Fig. 7-11 b). These intermediate cells progressively disappear, and neuronal cells expressing a characteristic morphology and characteristic immunophenotypic markers of newborn neurons become increasingly prevalent (Fig. 7-11 c,d). Immature neurons produced 5 days after induction of differentiation (n=4) display electrophysiological properties reminiscent of immature neuronal characteristics: RMP of +/20.8 mV, C m of 32.6 +/1.3 pF, R m of 1.3 +/0.3 G # , and R a of 16.7 +/5.5 M # , with prominent Na + and K + channels (Fig. 7-11 e). Immature neurons fire single evoked action potentials (Fig. 7-10 e). Oligodendrocytes were not detected following induction of neurogenesis. Defined cells displaying an exclusively neuronal phenotype universally incorporate BrDU (Fig. 7-11 f) and express Ki-67 (data not shown), suggesting they are the product of subsequent divisions of cells following differentiation. New neurons continue to express immature neuronal markers (PSA-NCAM, Fig. 7-11 g) and maturing neuronal markers (Fig. 7-11 h). Addition of serum to AHNPs induced to differentiate largely abrogates the production of neuronal cell types. To determine if region-specific neuronal cell types could be generated from correlate tissue, substantia nigra was dissociated and cultured as described, and induced to differentiate. Both mature (Fig. 7-12 a) and tyrosine hydroxylase + neurons (Fig. 7-12 b-f) were occasionally generated in culture, which assume variegated morphologies (Fig. 7-12 b, d). This finding suggests endogenous astrocytes may have intrinsic abilities to regenerate specific cell loss.

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117 Sample ID Age/Sex Region Long term culture established Max # PDs Neuron generation 1 27/F 1 Ctx GM, WM Ctx, GM 63, 60 Y 2 29/M 1 Ctx GM, WM, H, ASVZ Ctx GM, ASVZ 59 ND 3 17/F 1 Ctx WM, GM, A-SVZ WM 62 Y 4 5/M 1 Ctx GM, WM, H, A, M, P-SVZ H, P-SVZ 65, 64 Y 5 78/M 2 H, SN, A, M, P-SVZ ND ND Y Fig. 7-1. Surgical specimens employed for derivation of AHNP cell strains. Ctx: WM, White matter cortical tissue; GM, grey matter cortex; H, Hippocampus; SN, substantia nigra; SVZ, subventricular tissue: A, anterior; M, medial; P, posterior. 1 Surgical specimen derived from resection associated with medically intractable epilepsy. 2 Pathological specimen (36 hour postmortem interval) derived from a patient suffering from Parkinson’s disease.

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118 Fig. 7-2. Culture paradigm for the generation of AHNPs. Progenitors were isolated from a broad range of forebrain locations (green, representative biopsy sites shown for established cell lines) and cultured as mitogen-dependent adherent cultures. Previously described neurosphere-forming cells were isolated from periventricular (red) and hippocampal tissue, and were clonally expanded in nonadhesive culture. Differentiation of neurospheres yields all CNS cell types, while application of ‘neuronization’ conditions induces the formation of neurons (but not oligodendrocytes) in adherent progenitors.

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119 Fig. 7-3. Expansion of primary neural cells as a homogenous population of AHNPs. (a) Highly expanded (> 60 PDs) cells ubiquitously express nestin (red), with a large subset of GFAP + cells (green). (b) AHNPs express widespread immature neuronal and glial markers, including A2B5 (red) and NG2 (green). (c,d) AHNPs (nestin + , green) express astrotypic markers in a large subset of cells, including S100 (c, red) and glutamine synthetase (d, green). (e) Voltage-clamp profile of these cells reveals prominent Na + and

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120 K + channel activity. Data shown for temporal cortex-derived cells. (f) Nestin + (green) AHNPs proliferated in the presence of BrDU (red) uniformly incorporate thymidine analog. (g) Stereological evaluation of proliferating AHNPs reveals a uniform nestin + population which frequently coexpress glial cell markers (GFAP shown). Maintaining these cells in growth medium supplemented with BrDU results in label saturation in AHNPs (BrDU + Nestin + cells) at a rate of incorporation analagous to previously characterized proliferative dynamics (h). Removal of mitogenic stimuli (GF=EGF+bFGF) results in failure of AHNPs to divide (See Fig. 7-3 f). (i,j) Both hippocampal and temporal cortex-derived AHNPs maintain comparable, stable doubling rates and uniform, protoplasmic morphologies throughout culture. (k) AHNPs derived from temporal cortex and hippocampus reveals continuous logarithmic expansion throughout culture. Scale bars: 25 m (a,b,f,j), 50 m (c), 75 m (d). Images counterstained with DAPI. Fig. 7-4. AHNPs avoid immortalizing mutations, and exhibit mitogenand telomerasedependent growth. (a) Cultured AHNPs express major growth regulatory proteins longitudinally throughout culture. (b) Karyotyped AHNPs display normal ploidy and

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121 have no gross cytogenetic malformations. (c) Following growth arrest by an exogenous hTERT inhibitor (EGCG) or growth factor withdrawal cultured cells express SA" -Gal. However, only mitogen-withdrawn (-bFGF) cells lose hTERT expression when evaluated 7 days later. (d) Physiological (3 Gyx-irradiation) or chemical inhibitors (apidicolin, EGCG) consistently increase the fraction of cells expressing SA" -Gal. (e) Application of reversible growth inhibitors yields a significant reduction in growth rate. AHNPs revert to previous proliferative levels following arrestor washout. Age-matched AHNPs placed in either basic media (N2) or media containing EGF or bFGF only (N2E, N2F) enter irreversible growth arrest compared to defined proliferative conditions (N2EF) and subsequently become unviable. Data shown for temporal cortex derived cells. *p<0.05, student’s t-test. Scale bar: 75 m (c).

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122 Fig. 7-5. AHNPs maintain viability and assume glial phenotypes upon ventricular transplantation. (a) AHNPs cells injected into the right lateral ventricle of postnatal day 3 mice were detectable with HNA in periventricular tissue adjacent to injection site (*). (b) HNA + cells (boxed in a) were primarily located within 100 m of the ventricular wall in the ependymal and subependymal cell layer. (c-e) HNA + cells (red) integrating into the LV wall display conserved morphology of astrotypic cells, and frequently coexpress

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123 GFAP (green). (f) Confocal micrograph shows GFAP + process extension from an AHNP into the subependymal zone. (g, h) Activated dendrimer transfection of Pax6-eGFP of 30 PDs AHNPs 3d post-transfection. (i) Lentiviral-eGFP transfection of 30 PD AHNPs (20 moi). Scale bars: 40 m (b), 25 m (c-e), 8 m (f), 20 m (g-i). Images (c-f, h) counterstained with DAPI. Fig. 7-6. Optimized lentiviral infection of AHNPs. (a) 7d postinfection AHNPs were sorted for eGFP expression against untreated cells using standard fluorescence gating. (b) Following FACS analysis, plated cells remain viable and demonstrate continual eGFP expression. (c) Gated threshold measurements of eGFP expression at 5-20 moi.

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124 Fig. 7-7. Transfection of human progenitors with GDNF. Passage 10 (15 PDs) adult human hippocampal astrocytes were infected with either 5 moi/24 hr or 10 moi/12 hr lentivirus expressing a GDNF-eGFP coding region cloned from existing adeno-associated virus. Plated cells were examined for GDNF and eGFP expression 7 days after infection. (a-f) GFAP + astrocytes were detectable in culture in both WT and GDNF-eGFP infected

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125 cells 7 days following plating. Infected cells frequently express GDNF, which is not appreciated in uninfected cells. (g) eGFP expression (arrow) was detected in a nominal frequency in attached cells 7 days following infection. (h) ELISA evaluation of GDNF expression in human cells 7 days following infection. 5 moi/24 hrs and 10 moi/12 hrs compared to wild type. Significant (*p<0.05, student’s t-test) increase in GDNF expression was appreciated in 10 moi/ 24 hr infection conditions. GDNF expression in infected cells compared relative to control expression. Scale bars: 50 m (a, c, e, g), 25 m (b, d, f).

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126 Fig. 7-8. Genetically modified human progenitors integrate and express transgenes in vivo. Human progenitor cells (30 population doublings in culture) were co-infected with lentivirus containing recombinant green fluorescent protein (GFP) and glial-derived neurotrophic factor (GDNF) under the SV40 promoter. 5x10 4 progenitors were stereotactically injected into the striatum of immunocompromised Fox-SCID mice (a). Transplanted mice were sacrificed, and their brains fixed and processed into 20 m serial

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127 sections on a freezing microtome. Progenitor integration was evaluated 2 and 10 days after transplantation. Ten days following transplantation, progenitor cells were visible in the striatum (b) and co-express HNA and GFP (inset, b). GFP and GDNF expression was detected using serial section immunocytochemistry. Both GFP (c, magnified in d) and GDNF (e, magnified in f) are expressed in adjacent sections. Scale bars: 0.5 mm (b, c, e), 50 m (b, inset), 100 m (d, f).

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128 Fig. 7-9. Cortically-implanted AHNPs adopt predominantly neuronal fates. (a) Coronal section of engrafted left hemisphere shows " -III-tubulin + /HNA + donor cells adjacent to engraftment site. Schematic representation includes two-dimensional proximal-distal and lateral distribution of the majority of AHNPs and ectopically migrating cells in two transplanted animals (blue and yellow). (b) Fate analysis indicates few cells adopt an astroglial identity. (c) Integrating AHNPs within the primary engraftment site adopt

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129 neuronal morphologies and immunophenotypes. (d) " -III-tubulin + cells present within the hippocampus of engrafted animals occasionally displayed HNA (e, from boxed area in d) in CA1 and CA3, where they adopted apparent pyramidal neuron morphologies. (f,g) Single plane confocal image of cortically-implanted AHNPs. HNA + cells form mature neuronal (NeuN + ) cell types, which co-exist with endogenous neurons (arrowheads). Scale bars: 200 (a,b,d), 50 (c,f,g), 100 (e).

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130 Fig. 7-10. Distribution and phenotype of transplanted progenitors. Distribution of SVZ and cortically implanted AHNPs 7 days after transplantation. Both SVZ and cortically implanted AHNPs exhibit modest migration on the anterior-posterior axis. While SVZ implanted AHNPs exhibit limited migration from the ventricular surface into the cortex, AHNPs transplanted into the cortex exhibit strong bidirectional migration on the mediallateral axis. Immunophenotyping of AHNPs on the primary engrafting section

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131 identifying astrocytes (GFAP + ), neurons (-III-tubulin + ), or undifferentiated cells (GFAP , -III-tubulin ). Data presented as average percentage (n=3 animals per experimental condition). Fig. 7-11. Differentiation of AHNPs into neuronal cell types. (a) Proliferating cells (30 PDs) assume a compacted morphology immediately following removal of mitogens and addition of dibutyl cAMP, IBMX, and NGF. (b) 3 days following induction of differentiation, intermediate cells displaying a developmentally intermediate phenotype are appreciated. (c) Maturing cells concurrently lose GFAP and continue to strongly express " -III-tubulin. (d) Newly generated neurons in vitro frequently coexpress immature neuron markers, and assume typical bipolar morphologies. (e) Current and voltage clamp analysis of 7 day old neurons. New neurons exhibit prominent Na + and K + channels, and were able to fire elicited action potentials when polarized to mV. (f) " III-tubulin neurons generated in the presence of thymidine analog universally incorporate BrDU. Cells generated in this manner display additional type-specific neuronal markers, including PSA-NCAM (g) and neurofilament M (NF-M, h). Scale bars: 75 m (a), 25 m (b,h), 100 m (c,g). Cells counterstained with DAPI.

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132 Fig. 7-12. Generation of mature neurons and tyrosine hydroxylase-expressing neurons from human astrotypic progenitors derived from the substantia nigra of a deceased 78 y/o patient suffering from Parkinson’s disease. (a) Newly-generated neurons (3 days following inducible generation) express Tuj1 and NeuN. (b) 7 days following their appearance in culture, neurons occasionally express tyrosine hydroxylase in Tuj1 + cells, which retain morphologies similar to interneurons (b) or projection neurons (d-f).

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133 CHAPTER 8 DISCUSSION AND CONCLUSIONS Isolation, Establishment, and Characterization of SVZ Neurogenesis In Vitro 1 Using a novel culture paradigm, we have created a two-dimensional model recreating the subventricular niche (including neuropoietic and non-neuropoietic cells), which can be isolated and inducibly differentiated to form committed cell types. These observations not only expand the previous observations of both ourselves and others that describe how stem cell-derived neurons develop basic biophysical characteristics (Strubing et al. 1995; Finley et al. 1996; Song et al. 2002; Benninger et al. 2003), but describe the manner by which we can in longitudinally track the birth and maturation of precursors specific to the postnatal/adult SVZ in vitro, observing similar timing of developmental events and lineage restrictions as they occur in vivo. Moreover, our model offers the advantage of inducible neurogenesis, thereby generating populations of synchronously differentiating SVZ stem cells which are in monolayers readily available for single cell analysis in real time throughout early and protracted stages of differentiation. Use of this culture system also enabled us to focus on previously uncharacterized events during the first 96 hours of SVZ stem cell development. GFAP low /A2B5 + /nestin + / ! IIItubulin /Dlx-2 /PSA-NCAM proliferating glial cells develop 1 This chapter contains text excerpted from Scheffler et al, 2005, Walton et al, 2006a, and Walton et al, 2006b.

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134 within 24 hours into a multipotent, transit-amplifying GFAP /A2B5 + /nestin + / ! IIItubulin + /Dlx-2 -/+ /PSA-NCAM population, which give rise to GFAP /A2B5 + /nestin + / ! IIItubulin + /Dlx-2 + /PSA-NCAM + precursors specific to the SVZ. Closely resembling the glial-to-neuron transition of type B, C, and A cells in vivo (Doetsch, 2003), we present, for the first time, their comprehensive electrophysiological characterization. Significant changes of biophysical properties parallel stage-specific morphological and ultrastructural characteristics of differentiating cells. Following induced differentiation, morphologically unique A2B5 cells seem to first lose GFAP and then nestin before converting into functionally mature A2B5 GABAergic interneurons. Thus the A2B5 antigen, a developmentally regulated tetrasialoganglioside (Eisenbarth et al. 1979), appears to denote the sequential differentiation of a neural stem/progenitor cell en route to becoming an interneuron. Furthermore, we propose that a subpopulation of A2B5 + cells that co-expresses nestin and low levels of GFAP could represent the founders for SVZ-specific generation of interneurons. Even though the monoclonal antibody A2B5 is not cell-type specific as it detects different precursor cells in a variety of developing and adult CNS structures (Raff et al. 1983; Rao et al. 1998; Mi et al. 1999; Noble et al. 2003; Nunes et al. 2003), we found it intriguing that this antigen is also expressed by a subpopulation of radial glial cells in the lateral ganglionic eminence (LGE) between E13.5 and E15.5 (Li et al. 2004). The LGE supplies interneurons to the olfactory bulb during brain development (Wichterle et al. 1999), and is an ancestor of the postnatal/adult striatal SVZ. In separate pilot studies we performed immunolabeling on sections of postnatal rat and mouse forebrain (data not shown) and found A2B5 + cells spatially restricted in the dorsolateral ( striatal) aspect of the lateral ventricles; this in part

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135 could explain the origin of postnatal interneuron-genesis as we now have completely recapitulated in vitro. The use of reductionist culture models remains crucial for uncovering functional aspects of neural and non-neural stem cell biology (Colter et al. 2001; Song et al. 2002; Shen et al. 2004). Yet, recent studies have indicated that growth factors can significantly change the transcriptional profile of neural precursor cells (Gabay et al. 2003; Hack et al. 2004). It is therefore a major concern that cellular ‘reprogramming’ induced by EGF, bFGF, serum or other factors influences the behaviors of isolated cells in culture. Of course, cells in vitro might not always exactly behave as their in vivo counterparts; however, we demonstrate here, in spite of a disruption of a system that is clearly based on a large degree of inter-cellular communication, all characteristic neurogenic events unique to the rodent subventricular zone can be recapitulated in dissociated tissue culture. It can therefore be concluded that ‘organotypic’ postnatal neurogenesis does not necessarily require a total recapitulation of the environmental cues of the host brain. Alternatively, and on a broader perspective, the pioneering work of Dexter et al. (1977) showed how cells from bone marrow establish themselves in adhesive culture conditions that mimic specific environmental cues for proliferation and differentiation of hematopoietic cells. Our model could hence provide all essential environmental contributions of the postnatal SVZ neuropoietic niche, and results presented here would further support our originally touted similarities between hematopoiesis and neuropoiesis (Scheffler et al. 1999 ). Uncovering the precise morphogenic cadence during early stages of neuropoiesis, including transitional states and sequential differentiation as shown here, thus provides insights as well as a protocol for targeted isolation and generation of distinct populations

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136 from the spectrum of stem cells to fully differentiated neurons and glia in the adult brain. The SVZ in vivo and in vitro obviously has a propensity to generate GABAergic interneurons. Such a fate restriction associated with the known establishment of a forebrain adult neuronal lineage challenges future research aimed at manipulating neuropoiesis for desired neuronogenesis. Tuj1 Characterizes the Astrocyte-to-Neuron Metamorphosis of SVZ Neurogenesis Using an adherent model of inducible neurogenesis intermediate to conventional in vivo and in vitro approaches, we have defined a unique approach for the the identification of novel markers for neurogenic cells, as well as defining their role in the astrocyte-to-neuron transition of subventricular neurogenesis. Employing this approach, we have identified a novel marker, Tuj1, as a conserved marker that (a) frequently coexpresses markers associated with astrotypic NSCs and neural progenitors, (b) is expressed in the developmentally intermediate structures that bridge the astrocyte-toneuron transition, and (c) is present in newly generated committed neuronal cell types. The identification of such a marker is thus unique as it, labels primordial, differentiating, and newly generated cells comprising subventricular neurogenesis, unlike existing astrocytic (i.e., GFAP), NSC (i.e., nestin, Sox2) or intermediate/transit-amplifying (i.e., dlx2) markers. Utilizing Tuj1 as a marker for differentiating subventricular astrocytes in an inducible model of subventricular neurogenesis enabled us to characterize the mechanism by which subventricular neurogenesis occurs. Though Tuj1 is expressed in both neuropoietic and non-neuropoietic tissue, Tuj1 + cells from neuropoietic regions uniquely

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137 respond to prompts to differentiate via rapid alterations in morphology to assume a compacted cell body, which undergo further cell divisions to yield clusters of committed neuroblasts. Differentiating cells display concomitant intracellular remodeling, including reductions in cytoplasmic volume and nuclear size, and transient increases in cell cycle upregualtion and cellular machinery required to perform extensive intermediate division. Tuj1 appears to label the dominant cell comprising the astrocyte-to-neuron transition. Differentiating cells from neuropoietic regions exhibit extensive upregulation of cell cycle regulatory protein Ki-67, particularly within Tuj1 + characteristic intermediate cell formations present prior to the generation of defined neuroblasts. Alteration in the frequency of cells in growth vs. mitotic phase, as well as the prevalence of aneuploid cells in differentiating cultures closely match the cell division dynamics appreciated in Tuj1 + intermediate formations, further underscoring their role as the primary cell type comprising astrocyte-to-neuron transition. To correlate cells undergoing ‘organotypic’ differentiation with classical neurosphere formation, we compared the ability of undifferentiated (‘B cell’ enriched), 24 hour differentiated (‘C cell’ enriched) and 4 day differentiated (‘A cell’ enriched) to generate neurospheres. Our findings indicate cells isolated from progenitor-rich developmental states are enriched for neurosphere-forming cells, agreeing with evidence suggesting neurospheres are largely derived from the transit amplifying cells of the CNS, which are induced to form neurospheres when exposed to EGF in clonal conditions (Doetsch et al. 2002) . However, the appreciated period in which neurosphere-forming cells are enriched occurs 24 hours following the removal of EGF (albeit followed by growth in clonal conditions containing EGF and bFGF), suggesting a previously

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138 unreported intrinsic potential of transit-amplifying cells to form neurospheres in vitro. Neurosphere-forming cells derived from transit-amplifying cells possess an enhanced ability to self-renew, as they generate higher proportions of secondary neurospheres than ‘B’ or ‘A cell’ enriched developmental states. Interestingly, secondary neurospheres derived from all developmental states form equivalent numbers of tertiary neurospheres, regardless of the origin of the initial neurosphere-forming population. From a practical standpoint, this may indicate a necessity to generate secondary neurospheres for the accurate estimation of neurosphere-forming populations. To further evaluate the developmental potential of cells from various developmental states, similar populations of differentiating neuropoietic cells were isolated and fully differentiated in adherent conditions. Progenitor rich populations again displayed the highest proliferative potential and propensity for generation of neuroblasts, further suggesting they are an enriched population for “classic” NSC characteristics. Based on the ubiquitous nature of Tuj1 expression in cells displaying primordial markers, involved in differentiation, and as an established marker of newly generated neuroblasts, it appears likely that Tuj1 may be useful as a pan-developmental marker for NSC identification. By employing this knowledge in a controlled model of differentiation, we are able to demonstrate unprecedented ability of intermediate progenitor cells, previously thought to be limited in scope of differentiation and proliferation. These findings provide grounding for prospective avenues of investigation into further definition of NSCs and their differentiating progeny.

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139 Bromodeoyuridine Alters NSC Function and Induces Senescence In Vivo and In Vitro Using pulsed labeling and intraperitoneal injections, dissociated subventricular tissue, immortalized glioma and tumor cell lines, and early postnatal rodents were exposed to BrDU for both abbreviated and extended time periods. In all cases, BrDU addition was accompanied by deleterious on proliferation in vitro. To determine if BrDU possessed endogenous toxicity independent of incorporation, reversible mitotic inhibitors were applied to cells treated with BrDU. As cells proliferated under these conditions were able to resume normal rates of division, it appears that BrDU achieves an effect through incorporation into newly synthesized DNA. This observation is underscored by two additional expereimental observations: first, a single dose of BrDU has increasing effect on the reduction in growth rate per round of replication in expanding cells. Second, in cultured primary astrocytes from the SVZ, administration of BrDU prior to induction of differentiation results in the prevention of subsequent neuroblast production. As the production of neuroblasts in the postnatal mammalian brain is closely linked to cellular proliferation (Doetsch et al. 1999), it appears the use of BrDU as a labeling agent may have potentially confounding effectsin the investigation of NSCs and their function. Evaluation of asymmetric division, both in undifferentiated and differentiating NSCs was performed in separate pilot experiments. Over 1,500 dividing SVZ-derived cells were examined for asymmetric division did not detect asymmetric distribution of genomic DNA following mitosis (data not shown). This suggests that NSCs are likely incorporating BrDU. Experiments involving the delayed administration of BrDU (shown in Fig. 3-4) are successful in labeling

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140 neuroblasts, presumably due to the few rounds of replication remaining in the astrocyteto-neuron transition implicit in subventricular neurogenesis. While not definitively inhibiting the identification of clonally isolated NCSs, the presence of BrDU clearly affects neurosphere production. Despite the failure to significantly affect the total number of primary neurospheres generated in younger animals, reduced neurosphere size (combined with the failure to continue to form secondary neurospheres when further dissociated) suggests an analogous reduction in proliferative potential. Systemic application of BrDU to juvenile mice resulted in a dosedependent increase in the total levels of senescent cells within the two major defined neurogenic niches in the CNS. This is particularly interesting, as SA--Gal expression in non-neurogenic zones (i.e., cortical layers IV-VI) is not significantly different in BrDUtreated animals, and suggests that BrDU may be incorporation-dependent in vivo as well as in vitro. To identify a mechanism by which BrDU addition abrogates growth, primary astrocytes and glioma cells were examined (work presented and unpublished observations by Lindsay Levkoff, personal communication). Interestingly, BrDU does not appear to have a direct cytotoxic effect, as suggested by in vivo observations, but appears to abrogate growth through a progressive increase in senescent cell fraction. Additionally, it appears mitochondrial stress (measured by the increased production of superoxide products) is substantially increased by BrDU. It is unclear as to whether this oxidative stress is a secondary result of primary DNA damage or cell cycle alteration, or is a primary consequence of mitochondrial damage or alteration. However,

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141 mitochondrial disruption has been implicated in multiple disorders associated with aging and senescence, and disruption of mitochondrial function may ultimately underlie the appreciated consequences of BrDU administration. As such, BrDU application likely affects the fidelous recapitulation of neurogenesis in vivo and in vitro through alteration in growth rate and level of intracellular stress. Microglia Undergo Age-Dependent Neurogenic and Neuronizing Support Roles Using an adherent model for expanding and inducibly differentiating populations of constituent neuropoietic and non-neuropoietic niches provides conditions for recapitulation of ‘organotypic’ neurogenesis, allowing direct evaluation of factors relevant to maintenance and function of NSCs of the SVZ. By selectively depleting cell types through continued proliferation, we have identified a concomitant decline in microglial cells with the neurogenic potential of undifferentiated SVZ astrocytes. The essential requirement of microglial cells for neurogenesis was confirmed by selective depletion of microglial cells using a targeted toxin. Microglial contact is not required for facilitation of neurogenesis, as conditioned medium from multiple microglia-containing regions is able to restore neurogenesis, nor is it essential for maintenance of neurosphereforming cells. As such, microglial input does not appear to affect NSC self-renewal or multipotentiality, but appears to be a critical regulator of postnatal neurogenesis. NSCs are retained and expanded in culture, as evidence by the extensive selfrenewal in adherent culture, conservation of NSC marker expression, and continued generation of multipotent neurospheres. When induced to differentiate, a progressive

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142 reduction in neuroblast formation is appreciated in subventricular tissue, resulting in a complete loss of neurogenesis by ! 10 PDs. Addition of the nucleoside analog XS, a compound reported to function both as a suppressor of asymmetric cellular kinetics (Sherley et al. 1995) and as a suppressor of p53-mediated cell death/senescence (Sherley 1991), resulted in significant extension of the neurogenic window for undifferentiated SVZ dissociates. Using this approach, we were able to correlate microglial lifespan to neurogenic competency of undifferentiated neuropoietic tissue. NSC prevalence and self-renewal (evidenced by neurosphere formation and overall proliferative rate) are unaffected by microglial depletion, suggesting microglial contributions (a) do not affect basal NSC self-renewal, inherent multipotentiality, or proliferative index and (b) are constrained to the facilitation of neurogenesis. As conditioned medium from microglia is able to reconstitute neurogenesis in highly expanded dissociates, it is unlikely that microglia act to keep neurogenic cells alive, but rather function as external regulators of neurogenesis. This may reflect a commonly found developmental motif of a separate regulatory cell controlling a population of dedicated primordial cells, such as stromal cell control over proliferation and differentiation of primordial HSC progenitors in (Verfaillie 1993), and as a developmental mechanism for the restoration of radial glial cells (Gierdalski et al. 2005). However, this concept remains relatively undescribed in the postnatal brain, and may accordingly affect the prevailing view of neural stem cells as autonomous entities. On a related note, the process by which NSCs generate committed cell types via the neurosphere assay bears few similarities to the developmental continuum described in vivo for the production of committed cell types. For example, neurosphere-derived progeny do not appear to recapitulate the astrocyte-to-neuron

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143 transition appreciated in vivo, and are frequently comprised of cells morphologically intermediate to astrocyte and neuron (Laywell et al. 2005). As these types of cells are not fully described in vivo, these cells may not represent a developmental fidelity to the generation of progeny described in vivo or in primary dissociated cells described here. To further examine this phenomenon, we compared the generation of defined neuroblasts from monoclonal neurospheres to aggregation spheres (described in Laywell et al. 2005) containing non-neurogenic cells. Unlike monoclonal spheres, these polyclonal aggregation spheres generate A cells in a manner temporally and phenotypically consistent with that observed in vivo and in vitro. This observation further underscores the potential importance of non-NSC cells in recapitulation of prescribed organotypic neurogenesis. Finally, due to the lack of evidence presented for hyperplasic or transformed phenotype, it is unlikely that neurosphere-forming cells are retained through spontaneous immortalization. Microglia are hematopoietic in origin (Eglitis and Mezey, 1997), and begin CNS infiltration at birth which continues throughout adulthood (Cuadros and Navascues, 2001). As such, the regulatory mechanisms of neurogenesis originating from microglia are (a) non-neural in origin, (b) inherently postnatal, and (c) potentially capable of agerelated alteration. To further explore the latter possibility, we supplemented and subsequently differentiated highly expanded SVZ astrocytes using conditioned medium derived from adult animals. Despite containing a comparable number of microglia, adultderived conditioned medium were less effective in promoting neurogenesis. However, neurons generated using adult-derived conditioned medium developed into mature neuronal phenotypes more rapidly than controls using conditioned medium from young

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144 animals. This preliminary finding suggests a possible ongoing role of microglia may alter with age, from that of neurogenesis during the early postnatal period to neuronodevelopmental in the established adult brain, which bears further study. Microglia appear to be regionally-insensitive in their ability to affect neurogenesis, as cells and conditioned medium from variegated brain regions were successful in reconstituting neurogenesis. Conditioned medium derived from tumorogenic and HUVEC cell lines, as well as primary fibroblasts, were not capable of reconstituting neurogenesis. The soluble mechanism by which microglia affect neurogenesis remains currently unknown. Exposure of conditioned medium to heat (65C) and proteolytic digestion indicate the signal is highly labile. To attempt identification of the soluble factor(s) enabling neurogenesis, gel filtration chromatography was used to fractionate media from early postnatal dissociates (3-150 kDa). Fractions depleted of proteins <3 kDa were successful in inducing neurogenesis, however, individual fractions were unable to reconstitute neurogenesis, and were only successful when recombined. This observation suggests a pleitrophic, protein-driven interaction (possibly between multiple cell types), and further underscores the possibility of organotypic postnatal neurogenesis being multifactorial in origin. Astroglia have been shown to promote neurogenesis in hippocampal progenitors through a contact-mediated mechanism (Song et al. 2002), suggesting the existence of potential mechanistic and cellular differences underlying the regulation of SVZ and hippocampal neurogenesis. Our findings establish a more complex framework for the developmental regulation of NSCs than previously imagined, and may have far-reaching consequences

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145 in understanding and defining the basal regulatory elements underlying neurogenesis into adulthood. Derivation, Expansion, and Potential Uses of Human Progenitor Cells We demonstrate a method for isolating multipotent astrotypic progenitors from primary neural tissue and describe culture conditions necessary for their extensive expansion as a homogenous population. AHNPs maintain a stable doubling rate throughout culture, and do not exhibit characteristics of transformed cells, including loss of key cell cycle checkpoint proteins, loss of sensitivity to arrestors of the cell cycle, and cell contact inhibition of growth. Our results suggest it is possible to isolate and expand astroglial cell lines from epileptic temporal lobe resections with characteristics of relatively uncommitted progenitor cells without senescence or cellular transformation. Similar to all embryonic, germ, and certain adult stem cells, telomerase is ubiquitously expressed in cultured cells, initially at high levels and at subsequently lower levels. Unlike immortalized cells, telomerase expression is not coupled to a loss of key regulatory proteins required for immortalization (Kiyono et al. 1998; Evans et al. 2003) . Thus, the observed growth phenotype is a hybrid one, allowing expansion past proliferative limits while avoiding immortalizing mutations. Cultured cells display conserved primordial markers, with a large subset displaying characteristics of type I protoplasmic astrocytes. Thymidine analog incorporation reveals a uniform proliferative capacity among AHNPs, suggesting we have identified dedicated culture conditions for the isolation and expansion of a homogenous progenitor population rather than conditions which favor or maintain a

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146 subset of primordial cells that constitutively generate progressively more differentiated/postmitotic cell types. Similar to previously described neurogenic cells (Nunes et al. 2003), AHNPs maintain similar immunological expression and mitogenic dependency throughout culture. However, our cells were expandable as an adherent monolayer, were derived from multiple regions, and display a uniform potential for generating neuronal cell types. Comparison of AHNPs derived from the hippocampus and temporal cortex reveals no difference in growth rates, cellular composition, or significant physiological factors. This is interesting, as the hippocampus is believed to contain astrocyte-like NSCs that have documented self-renewal and multipotentiality in vitro and in vivo (Eriksson et al. 1998). Thus, our findings present data suggesting the possibility of a broadly distributed population of neurogenic progenitor cells. These cells are not inherently multipotent or self-renewing in clonal conditions by virtue of their failure to produce multipotent neurospheres, indicating they do not meet the existing criteria for neural stem cells. Despite a lack of inherent multipotentiality, AHNPs can be induced to form neuronal cell types in vitro and in vivo. Utilizing a paradigm adapted from the generation of neuronal cell types from cortical neuronal cell lines (Ronnett et al. 1990), it was possible to generate immature neurons in a similar manner. Similar to studies of postnatal neurogenesis in rodents and humans (Laywell et al. 2000; Seri et al. 2001; Sanai et al. 2004; Scheffler et al. 2005), neurogenesis is accompanied by cell division, as evidenced by ubiquitous BrDU incorporation and cell cycle marker expression characteristic of dividing cells. Neuronizing AHNPs frequently display an “asteron”

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147 hybrid phenotype, similar to recent findings in cultured rodent neural cells (OkanoUchida et al. 2004; Laywell et al. 2005). Whereas the previous studies report a neuronto-glia transdifferentiation, the transition reported here represents a glia-to-neuron commitment, suggesting the isolation of conditions (Ronnett et al. 1990) promoting the phenotypic alteration of widely distributed endogenous AHNPs may be possible. Similarly, neurons derived in this manner possess electrophysiological profiles occasionally appreciated in astrocytic cells derived from astrocytic tumors (Bordey and Sontheimer, 1998). However, when transplanted to the cerebral cortex, neurons assumed almost exclusively neuronal phenotypes, suggesting the appreciated electrophysiological development is at least partially a product of in vitro culture. It was not possible to produce oligodendrocytes using this approach, suggesting that that the observed generation of neuronal phenotypes may be direct result of signal-driven developmental cues, rather than intrinsic differentiation potential. Taken together, these finding indicate a cell lacking traditional NSC characteristics may be isolated and maintained as a cellular substrate for the inducible generation of neurons. AHNPs remain sensitive to exogenous cell cycle inhibitors (aphidicolin, EGCG), but continue to express telomerase and return to previous levels of proliferation upon removal of exogenous growth inhibitors. EGCG is able to rapidly inhibit proliferation of primary cells in this culture system, in contrast to previous work done in immortal cell lines (Naasani et al., 1998), but does not abrogate hTERT expression. However, upon mitogenic withdrawal, telomerase expression is promptly lost and continued proliferation ceases, neither of which is restored upon reversion to defined proliferative media. This anecdotal coupling of hTERT expression to cellular proliferation and mitogenic

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148 dependence provides for several interesting possibilities. First, these results suggest a system whereby environmental mitogens (provided in constant supply) provide a condition-specific synergistic growth effect, allowing for both hTERT expression and continued expansion. A loss of environmental support factors may trigger a demonstrably irreversible loss of hTERT expression, which may effectively mortalize cells or possibly, in the case of AHNPs, trigger their immediate and irreversible entry into a state of replicative senescence. The appreciation of rapid senescence with the loss of hTERT expression in aged cells suggests that both telomere length and telomerase expression may be critical to the maintenance of continued cell division in populations of proliferatively active cells, such as neural progenitor populations. This agrees with a number of reported examples of poor correlation between telomere length and replicative senescence, including one example in which cells rescued from replicative senescence by viral transfection of telomerase maintained shorter telomere length than replication incompetent counterparts (Yang et al. 1999; Zhu et al. 1999). The ability to massively expand progenitor cell populations possesses implications for diagnostic neurobiology as well as for therapeutic approaches involving tissue replacement. By extensively expanding primary cells from various brain regions, it is possible to create a substrate for neural cell bioassays (i.e., primary cell drug testing) without relying on clonally derived cell lines that contain potentially masking genotoxic mutations or inaccurately reflect the homeostenosis of target cells. Recent efforts for cell replacement therapies in the brain have prompted a focus on transplantation biology, including the use of glia genetically modified to express neurotrophins (Kordower, 2003; Tai YT, 2004). Employing a logarithmic ex vivo expansion of endogenous cells allows

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149 heretofore unprecedented applications in neurotransplantation and neural cell bioassays. This fact, combined with the demonstrated amenability to alteration of gene expression and fate choice in these cells provides an exciting substrate for further investigations addressing disorders and repair of the human CNS.

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162 BIOGRAPHICAL SKETCH Noah Walton was born in Charlotte, North Carolina, on March 28, 1980. Upon graduation from Jacksonville Senior High School in 1998, he attended Duke University, graduating in 2001 with a major in biology and minors in chemistry and sociology. Following graduation, Noah worked as a research technician in the laboratories of Drs. Donald Schmechel and Patrick Sullivan at Duke University until enrolling in the University of Florida Interdisciplinary Program in Biomedical Science in 2002. Noah joined the laboratory of Dr. Dennis Steindler in 2003, and began study of the biology underlying the maintenance and differentiation of neural stem cells. Noah has, to date, published multiple first author papers, and has presented over a dozen posters and oral presentations at multiple international meetings, including the annual meetings of the Society for Neuroscience and the International Society for Stem Cell Research, and has received a predoctoral research fellowship from the NIH. While in Gainesville, Noah has also run several marathons and iron distance triathlons.