REGENERATIVE MEDICINE APPROACHES TO SPINAL CORD INJURY

A Dissertation

Presented to

The Graduate Faculty of The University of Akron

In Partial Fulfillment

of the Requirements for the Degree

Doctor of Philosophy

Ashley Elizabeth Mohrman

March 2017

i

ABSTRACT

Hundreds of thousands of people suffer from spinal cord injuries in the U.S.A. alone, with very few patients ever experiencing complete recovery. Complexity of the tissue and inflammatory response contribute to this lack of recovery, as the proper

function of the central nervous system relies on its highly specific structural and spatial

organization. The overall goal of this dissertation project is to study the central nervous

system in the healthy and injured state so as to devise appropriate strategies to recover

tissue homeostasis, and ultimately function, from an injured state. A specific spinal cord

injury model, syringomyelia, was studied; this condition presents as a fluid filled cyst

within the spinal cord. Molecular evaluation at three and six weeks post-injury revealed a

large inflammatory response including leukocyte invasion, losses in neuronal

transmission and signaling, and upregulation in important osmoregulators. These

included osmotic stress regulating metabolites betaine and taurine, as well as the

betaine/GABA transporter (BGT-1), potassium chloride transporter (KCC4), and water

transporter aquaporin 1 (AQP1). To study cellular behavior in native tissue, adult neural

stem cells from the subventricular niche were differentiated in vitro. These cells were

tested under various culture conditions for cell phenotype preferences. A mostly pure

(>80%) population of neural stem cells could be specified using soft, hydrogel substrates

with a laminin coating and interferon-γ supplementation. To guide and possibly recruit

native stem cells, as well as reduce injury in the spinal cord, an injectable delivery

iii strategy is necessary. An in situ cross-linking hydrogel could increase latency and localization of treatments. In this project, a chitosan/PEG based hydrogel was tailored for

CNS tissues with low swelling post-gelation, a low elastic modulus (0.37 kPa), and very low cytotoxicity. When injected into the spinal cord parenchyma, the hydrogel elicited close to the same response as the saline injected surgical sham group. Overall, these platforms can be used to manufacture future strategies to locally deliver therapeutics that combat spinal cord injury.

iv

ACKNOWLEDGEMENTS

Graduate school has been a long and bumpy road. I definitely would not have made it to the end in one piece without many of wonderful people encouraging, helping, and sometimes down right pushing me along the way.

I would like to thank Dr. Leipzig, for taking me into his lab and teaching me to do research when I had no real experience. I’m sure there were some frustrating times along the way (writing the mini-book comes to mind!), but you stuck with me and made me into a stronger person. I appreciate all the times you came into the lab to teach us techniques and you open door policy. Getting so much face time with your advisor can be a rare thing, and I am happy to have received direction “from the source.” Thank you for encouraging a sense of family in the lab, from semi-annual lab outings to sharing your garden produce and dahlia tubers.

A big thank you to my committee, Drs. Chase, Cheng, Joy, Willits, and the newly joined Dr. Monty! Your comments, critiques, and questions have helped to mold this project. I am grateful that I got to learn from you in and out of the classroom, you all greatly inspire me. A special thank you to Dr. Monty and Dr. Willits for talks and advice,

I strive to someday be the role-model to future graduate students the way you were to me.

To all the students I have had the pleasure of working with, graduate and undergraduate, I am extremely grateful for research discussions, non-research talks, pick- me-ups after failed experiments, and all around brightening my day. I believe that great research comes from a collaborative effort, where minds from different backgrounds and

v areas of expertise really meld to make something special. Thank you to Andrew, Hannah,

Mahmoud, Pritam, Shahrzad, and Trevor for answering my incessant questions, being excellent sounding-boards, suggesting solutions to problems big and small, and just for being amazing scientists and surrounding me with positivity.

The early (read: glory) years would not have been the same without my previous labmates, who are now very dear friends. Hang, Aleesha, Natalie, and Liza: I miss you every day and wish each of you were still in my daily life! I must also thank and spread the appreciation to friends and sources of knowledge outside of my own lab. Frank, you helped me immensely in and out of the machine shop! Not only did you make amazing things that greatly enhanced my research, but you kept me sane with all the trips to the rec and introduced me to new friends through crazy walleyball games. Mary Beth, I definitely would not have finished everything up without our running (a.k.a. therapy) sessions towards the end of my project. Thank you for being a great friend and a kick- butt colleague. You all were so supportive and integral to my success as a graduate student. I look up to each one of you, and I hope we continue to stay in contact and remain friends.

Saving the best for last, I must acknowledge my family, without whom I certainly would not have achieved so much in my life. I am eternally grateful that I get to call you all family (be it by blood or by choice), you are my rock, my lifeline, my cheerleaders, my angry mob, my everything. Also, thank you for keeping me grounded in the fact that there is life going on OUTSIDE of a laboratory. I cannot name everyone here, I do like trees, but need to say a few special thanks to Anthony, Caitlin, and Kent, and to Leah and

Amanda, my oldest friends. I must officially thank my husband, Brian, you should be

vi nominated for saint-hood. I cannot wait to see what else life has in store for us (should be easy compared to these last couple years)! I would not have even started this journey if not for the constant love, support, and encouragement of my parents and my brother, all of whom have helped make me the person I am today. Thank you for teaching me by example, the person that I want to be.

vii

TABLE OF CONTENTS

Page

LIST OF FIGURES ...... xiv

CHAPTER

I. INTRODUCTION ...... 1

II. CENTRAL NERVOUS SYSTEM GROSS ANATOMY, CELL TYPES, AND

MATERIAL BACKGROUND ...... 7

Introduction ...... 7

Anatomy of the CNS & Progress of Neurological Damage ...... 9

Anatomy & Physiology of the CNS...... 9

Loss of Neural Function...... 18

Neurodegenerative Diseases ...... 25

Role of Glia in Degeneration & Regeneration of CNS Axons ...... 26

Biomaterials for Scaffold Preparation ...... 28

Definition of Biomaterial & Requirements for Neural TE Scaffolds ...... 28

Biodegradable Scaffolds ...... 29

Sample of current Biomaterials in CNS TE ...... 33

Cell Sources for CNS TE ...... 38

viii

Primary Cell Treatment of CNS Injury: Glial Cells ...... 40

Pluripotent Stem Cells: Embryonic Stem Cells ...... 42

Induced Stem Cells ...... 46

Adult Stem Cells: Endogenous Stem Cells in the Brain & Spinal Cord ...... 48

Mesenchymal Stem Cells ...... 52

Stimulation & Guidance ...... 55

Physical Cues ...... 56

Chemical Cues ...... 63

Electrical Stimulation...... 73

Concluding Remarks ...... 75

III. MOLECULAR LEVEL INVESTIGATION OF SPINAL CORD INJURY MODEL

...... 77

Summary ...... 77

Introduction ...... 78

Materials and Methods ...... 82

Surgical protocol ...... 82

Animal perfusion and tissue harvest ...... 83

Histological Staining ...... 84

RNA sequencing and transcript processing ...... 85

Metabolite processing and identification ...... 86

ix

Network generation and mapping ...... 88

Results ...... 89

Excitotoxic injury has a unique molecular signature associate with tissue

pathology...... 89

Inflammation and immune response over six weeks ...... 92

Neuronal-related loss and functional testing ...... 95

Fluid and solute transport perturbation ...... 97

Discussion ...... 100

Conclusions ...... 110

Acknowledgments...... 110

Supplemental Information ...... 111

IV. INVESTIGATE AND MANIPULATE NATIVE STEM CELLS OF THE CNS ... 112

Summary ...... 112

Introduction ...... 113

Materials and Methods ...... 115

Substrate Preparation ...... 115

Adhesive Protein Immobilization ...... 116

Surface Protein Verification ...... 117

NSC Isolation and Expansion ...... 118

NSC Differentiation ...... 118

x

Immunocytohistochemistry...... 119

Cell Counting ...... 120

Statistical Analysis ...... 120

Results ...... 121

Substrate Analysis ...... 121

Cell Density of NSCs ...... 121

NSC Differentiation Cell Types ...... 122

Discussion ...... 125

Coupled compliance and chemistry ...... 125

Cell viability and proliferation ...... 126

NSC differentiation and morphology...... 130

Conclusions ...... 132

Acknowledgments...... 134

Chapter 5 ...... 135

V. IN SITU GELLING CHITOSAN-PEG COPOLYMER EVALUATION FOR USE IN

THE SPINAL CORD ...... 135

Introduction ...... 135

Materials and Methods ...... 138

Materials and equipment used ...... 138

Hydrogel precursors and gel formation ...... 138

xi

Thiol confirmation and content ...... 139

Gelation and mechanical properties ...... 140

Hydrogel swelling and degradation ...... 140

In vitro toxicity ...... 141

In vivo material injection ...... 141

Temperature sensitivity testing ...... 142

Tissue sectioning and staining ...... 143

Statistical analyses ...... 143

Results ...... 144

Chitosan gel properties ...... 144

Cytotoxicity of fibroblasts and spinal cord cells...... 145

Host response safety assessment ...... 147

Discussion ...... 149

Conclusion ...... 155

VI. CONCLUSIONS ...... 156

REFERENCES ...... 161

APPENDICES ...... 193

APPENDIX A. SUPPORTING INFORMATION FOR CHAPTER 3 ...... 194

xii

APPENDIX B. SHORT DURATION ELECTRICAL STIMULATION TO

ENHANCE NEURITE OUTGROWTH AND MATURATION OF ADULT NEURAL

STEM PROGENITOR CELLS ...... 220

xiii

LIST OF FIGURES

Figure Page

Figure 2.1. General tissue engineering strategy. 9

Figure 2.2. Spinal cord anatomy 10

Figure 2.3. Major cell types of the CNS 14

Figure 2.4. CNS injury response 22

Figure 2.5. Diagram of neuronal development from the blastula in Xenopus 44

Figure 2.6. Diagram of differentiation paths for ES and iPS cells 47

Figure 2.7. Illustration of topographies. 60

Figure 2.8. Attraction and repulsion cues. 64

Figure 2.9. Common methods of protein surface attachment. 69

Figure 3.1. Syringomyelia study design. 81

Figure 3.2. Analytical overview of data. 90

Figure 3.3. Immune response for EXC versus CTL groups. 94

Figure 3.4. CNS parenchymal response. 97

Figure 3.5. Syrinx propagation following EXC injury. 99

Figure 4.1: Illustration of coverslip preparation showing silanation and immobilization of proteins 116

Figure 4.2: FTIR absorbance scans of immobilized of proteins on chitosan 122

Figure 4.3: Cellular density of NSCs on treated coverslips after 7 d of culture with

IFN-γ. 123

xiv

Figure 4.4: Cell fate of NSCs after 7 d in culture with IFN-γ on modified surfaces. 125

Figure 4.5: IHC images of NSCs exposed to laminin functionalized surfaces after 7 d in

the presence of IFN-γ showing neuronal markers βIII tubulin and synapsin. 128

Figure 4.6: Immunostaining for glial markers in NSCs exposed to laminin functionalized surfaces after 7 d in culture with IFN-γ. 129

Figure 5.1: Schematic of chitosan thiolation and gelation by mixing with

PEG copolymer. 138

Figure 5.2: Mechanical characterization of chitosan/PEG hydrogel 145

Figure 5.3: Cytotoxicity measurements and cell images. 146

Figure 5.4: Temperature sensitivy testing and immunostaining from spinal cord injection

subjects 148

Figure 5.5: Metrics from histological images were measured on three rodents per group at

week one. 151

xv

CHAPTER 1

INTRODUCTION

The brain and spinal cord, making up the central nervous system (CNS), are the information processing center and highway for the whole body. Due to this, injuries to the CNS are usually devastating to correct. Holistic function can be affected, including proper speech, interpretation, memory, sensation, movement, and bladder control, to name a few. The physical, emotional, and monetary strain on the patient and caretakers of

CNS deficit are immense, and any regain in functional control is a massive achievement.

In fact, the National Spinal Cord Injury (SCI) Statistical Center estimates that of the estimated 280k people with a SCI, less than 1% experience complete neurological recovery; the majority (45%) are within the incomplete tetraplegia category. At the cellular and molecular level, damage in the spinal cord tissue continues long after the initiating injury. A cascade of secondary events leads to further damage to the tissue and eventual scar formation which can block afferent and efferent signal transduction through the spinal cord. To combat loss of spinal cord tissue, information must be gathered on the pathways and mechanisms responsible for tissue and functional loss. The study of spinal cord disease and injury is a critical step in formulating a recovery strategy. Once a proper target is sighted, incorporating essential elements to reduce further deterioration and increase regeneration is important.

1

One specific spinal cord disorder is syringomyelia, or the formation of a fluid filled cyst (syrinx) within the spinal cord. Syrinxes are not well understood as they can have very strange and different presentations; cases in humans have described as asymptomatic to causing pain, tingling, or numbness. The National Institute of

Neurological Disorders and Stroke estimate at least 40,000 people in the US alone are affected, with symptoms beginning most commonly during young adulthood. For this reason, a better knowledge of the initiation and volumetric growth of syrinxes is necessary.

Native adult neural stem cells (NSCs) persist within the subventricular zone, hippocampus, and spinal cord central canal; the in vitro and in vivo behavior of these stem cells has been studied for regenerative strategies in CNS tissue engineering. These cells can be manipulated by physical and chemical cues to differentiate into neurons, oligodendrocytes and astrocytes, the three main cells of the CNS. NSCs possess the potential to replace particular cell types, specifically neurons, often lost in injury and disease.

Currently, there are no encompassing solutions to address brain and spinal cord injuries. Healing the CNS after injury is extremely complex due to toxic or inhibitory signals. To combat the negative environment post-injury, tissue engineering and regenerative medicine strategies are being developed that attack the problem from multiple angles: survival, clearing of inhibition, and regeneration of lost cells and architecture. These schemes often involve the use of scaffolds containing cells and/or therapeutic agents to augment endogenous regeneration. The overall goal of this dissertation is to discern important molecular level aspects of the syrinx environment and

2

combine them with a biomaterial therapy system aimed at steering the local tissue

towards regeneration.

Specific Aim 1: Closely examine syringomyelia at the molecular level and uncover

important pathways to target in subsequent regeneration strategies.

A detailed understanding of the injury environment is crucial to identifying targets for

tissue engineering. Spinal cord syrinx formation and progression will be studied by two

parallel, highly sensitive techniques. Next generation RNA (ribonucleic acid) sequencing

will allow the identification of major genes dysregulated in the sub-acute injury stage.

Transcripts are identified from the mRNA (messenger RNA) stage in the cell, and the techniques now employed are quantitative allowing the discernment of the abundance of particular sequences. As a precursor to protein, mRNA can relate to the amount of proteins cells are producing or the RNA can be a transcription factor in the cell, influencing cellular pathways. In parallel, the small molecules, or metabolites, will be quantified and identified using mass spectrometry. The transcript data will be linked with metabolomics data to hone in on important biological pathways disrupted during syringomyelia in the rat and confirmed with some immunostaining of the tissue. The driving hypothesis is that the aquaporin family of water transporters will be highly dysregulated along with one or two other pathways also involved in fluid flow and pressure build up in the spinal cord.

Specific Aim 2: Investigate and manipulate native stem cells of the CNS in vitro to

gain knowledge that can be incorporated into future tissue engineering schemes toward guiding NSC behaviors.

3

Adult NSCs derived from the lateral ventricles of the brain could serve as an

endogenous source for neuron and oligodendrocyte regeneration in the CNS. This

population of stem cells in particular allows for high expansion in culture and flexiblility

in expressing all three parenchymal phenotypes. These cells will be studied in the

laboratory in response to stimulation by chemical and physical cues. Particularly, hard

and soft substrates coated with different native cell matrix components will be compared.

Outcomes of these studies will be used downstream to direct cell behavior in the injured

spinal cord. The guiding hypothesis is that NSCs can be directed towards close to 100% neuronal population through culture on soft substrates with immobilized laminin.

Specific Aim 3: Develop a tissue engineered construct with the aim of diminishing

syrinx volume and damage in the spinal cord and possibly achieving regeneration of

lost tissue using knowledge of the injury environment and native cell behavior.

An injectable hydrogel will be developed to deliver several chemical cues that combat

the injury environment and promote native healing. The construct will mimic native

tissue stiffness and gel in a sufficient time to localize treatment. Material characterization

and release of regenerative molecules will be studied in vitro. A brief in vivo

investigation will ensure safety of the material for spinal cord injection by evaluating host

response using immunostaining techniques. The overall hypothesis is that a chitosan-

based material can be modified to gel in under 5 minutes with a biomimetic stiffness.

Further, that this material being based in chitosan will not exhibit cellular toxicity in

culture or in situ.

A detailed background is included in Chapter 2. Introduction to the CNS and

native parenchymal cells gives some background on the endogenous, healthy

4

environment of the brain and spinal cord. A brief background on neuronal degeneration

gives some information on the cascade of cell death that can result from injury or disease.

Characteristics of some common biomaterials used in the CNS are described here.

Chitosan in particular is discussed in several applications. Finally, stem cell sources for

CNS tissue engineering applications are reviewed.

Following the background of CNS anatomy and injury process, Chapter 3 details

a molecular investigation into syringomyelia in a rat model. Previous comprehensive

investigation of the movement of CSF in syringomyelia and initial investigations into

water channels done by the Stoodley group prompted this work. The goal was to

characterize molecular players in the initiation and propagation of the syrinx by utilizing

advances in nucleic acid sequencing alongside the newer parallel technique of

fingerprinting small molecules, or more commonly, metabolomics.

Chapter 4 details in vitro manipulation of NSCs to guide neuronal differentiation

sans serum. Native neural stem cells from the subventricular zone were cultured under

specific substrate conditions to elucidate their phenotypic responses. Learning to control

these stem cells could lead to better cell impregnated scaffold design or incorporating of

proper growth conditions to induce infiltration of native cells.

In Chapter 5, the characterization of an injectable biomaterial system is described and the host response from injection in situ. Attempts to modify chitosan for gelation

without radical mediated cross-linking culminated in a thiol-conjugated material that was

able to react with a co-polymer for rapid gelation, or auto-gelation over the course of

days. Chemical, mechanical, and toxicological characterizations were then performed to

ensure the material is appropriate for CNS application. The highlight of this chapter

5 includes preliminary evaluation of the host responses in the spinal cord following material injection compared to a surgical sham.

Concluding remarks in Chapter 6 summarize the findings of the entire project, from injury-to cell-to material analyses. These final remarks aim to put all the pieces together with lessons learned to offer up suggestions for future directions.

6

CHAPTER 2

CENTRAL NERVOUS SYSTEM GROSS ANATOMY, CELL TYPES, AND

MATERIAL BACKGROUND1

Introduction

The brain and spinal cord compose the Central Nervous System (CNS), which is

the control center of the body. Inputs from muscles, involuntary organs, and senses travel

through the nerves of the Peripheral Nervous System (PNS) into the CNS where they are

interpreted. Signals may travel within the brain to separate functional areas. Instructions

are then sent outward again for voluntary movement and involuntary regulation to

complete the endless loop of the nervous system circuitry. The CNS is critical to function

of the entire body, which is why incurred injury and disease cripple one’s quality of life.

In the United States alone approximately 265,000 people are estimated to have spinal

cord injuries (SCIs), with over 10,000 new injuries occurring each year.[1] Patients with

SCI experience decreased lifespan in addition to life-costs from one to four million

dollars, depending on the extent of injury, which is especially disturbing considering the

fact that the average age of a spinal cord injured person is 31.[1] Moreover, traumatic

1 Chapter 2 contains material published as a review article as Wilkinson, A.E., McCormick, A.M., Leipzig, N.D. Central Nervous System Tissue Engineering: Current Considerations and Strategies. Synthesis Lectures on Tissue Engineering. Athanasiou, K.A. and Leach, J.K. Eds, 2011. Morgan & Claypool Publishers, p. 1-120.

7 brain injuries (TBIs) occur to over 1.7 million people each year.[2] The devastating physical and psychological effects of CNS damage are felt by both patients and their families. Of SCI individuals experiencing paraplegia or tetraplegia, less than 1% achieve full neurological recovery post treatment.[1] Solutions to recover neurological function are desperately needed for all CNS injuries.

Tissue engineering (TE) in the CNS is extremely difficult because of the intrinsic restrictions and complexity of native CNS tissue. Generally, multi-component approaches are used in attempt to restore natural function to the brain or spinal cord. First and foremost, an understanding of tissue formation and function as well as tissue responses to damage is needed in order to formulate treatments to correct injury and disease in the

CNS. Knowledge of native tissue and pathological development will foster improvement of strategies for overcoming damage to the CNS. For the most severe CNS disorders, a complex TE construct involving multiple cues is most likely needed to combat the physical and chemical obstacles of the CNS; the general building blocks of these constructs are physical scaffolding from biomaterials, endogenous or exogenous cells, and stimulatory cues from chemical, mechanical and electrical signals within the construct or on its surface (Fig. 2.1). Within this review, scaffold formation techniques and common biomaterials in CNS TE will be discussed followed by potential cell sources. Subsequent sections will discuss the myriads of stimulatory and guidance techniques currently being employed in CNS strategies. The hope of this review is to give the reader the basic tools for designing or understanding strategies aimed at regenerating the CNS and also for exposure to current approaches. PNS regenerative strategies are

8

often discussed to augment basic understanding and many of these techniques do

translate to the CNS. In depth review articles and books are suggested throughout for

further reading on particular topics.

Figure 2.1. General tissue engineering strategy: knowledge of native cell environment is used to combine cell sources with mechanical, chemical, and electrical cues into a tailored tissue engineering construct. This assembly is grown under proper stimulation for translation to host tissue for regeneration.

Anatomy of the CNS & Progress of Neurological Damage

Anatomy & Physiology of the CNS

Neurons throughout the body are organized into three main structures: ganglia, nuclei, and lamina.[3] Outside of the CNS, neuronal bodies are grouped together into ganglia; an example of this formation is the dorsal root ganglia (DRG) of sensory cell

9

bodies that form just outside of the spinal column in the PNS. Within the brain, neuronal

bodies with a common function are grouped together into nuclei. The bulk of the brain is

organized into a layered cortex, including most of the cerebrum and cerebellum. The

cerebrum, diencephalon, cerebellum, and brain stem make up the parts of the brain and

are all housed within the skull (Fig. 2.2).

Figure 2.2. (A) Illustration of the CNS displaying the brain and spinal cord. (B) Cross- section of the spinal column showing the spinal cord protected within the vertebrae. (C) The spinal cord is segregated with white matter surrounding gray matter. Spinal roots exit on the ventral side and enter on the dorsal side to and from the PNS. Figure reprinted from.[4]

10

The cerebral cortex is extremely complex and has a number of functions including, but

not limited to, systemic sensory and motor control, speech, recognition and

understanding.[5] The spinal cord runs inferior to the brain stem in a columnar form, protected by the vertebrae of the spine. The inner core of the spinal column, with a butterfly like shape, contains the gray matter while the surrounding axons are white matter. Dorsal horn (sensory), ventral horn (motor), intermediate zone and commissural region comprises the gray matter.[3] White matter is made up of the anterior, posterior

and lateral columns.[3]. Gray matter is largely unmyelinated while myelination provides

white matter with its name and color. Nerve fibers enter and exit the spine at each

vertebra though holes called Foramen, allowing information to pass to and from the PNS.

There are four main groups of spinal nerves that exit at different levels of the spinal cord.

Named in descending order down the vertebral column these are: cervical (neck),

thoracic (upper back), lumbar (lower back) and sacral (base) nerves. While the PNS is

made up of groups of axons termed nerves, in the CNS axons run in groups called tracts

that are bound together by the processes of astrocytes, often called ‘end-feet.’[3]

Descending (efferent) pathways include the pyramidal and extrapyramidal tracts, and

ascending (afferent) pathways include the spinothalamic and spinocerebellar tracts as

well as the gracile and cuneate fasciculi. Proper bodily function depends on these paths to transmit information between the brain and periphery.

The extracellular matrix (ECM) is an important component of the CNS, and it accounts for around 20% of the adult brain [6]. More thorough reviews of CNS ECM are described elsewhere, only a few of the common ECM molecules are discussed here.[6, 7]

11

The brain and spinal cord are primarily made up of proteins and proteoglycans -

macromolecules with a protein core and glycosaminoglycan (GAG) side chains. GAGs

are linear, negatively charged polymers of repeating disaccharide units also present in the

extracellular space that help to properly hydrate tissues. Not only does the ECM provide the natural scaffolding for tissue, but it plays an active role in the regulation of diffusion of soluble proteins and in localizing membrane proteins to functional domains. Collagens and laminins are the primary ECM proteins of the CNS, which mainly make-up the basal lamina, and contain specific amino acid sites that interact with cell receptors.[6, 8]

Several types of collagens are found in the brain and spinal cord (I, II, IV, XVII, XIX) however, they are not as abundant in the CNS as they are in most other tissues.[9]

Integrins are the receptors that cells use to interact with the ECM. These transmembrane glycoproteins are made up of an alpha and a beta subunit that complex to activate a host of signaling pathways within the cell. Integrins provide a link from the ECM to the cytoskeleton. In the CNS, β1 integrins are most relevant, binding to ligand sites on laminins, and are critical for proper neuronal migration.[7, 8] A major GAG found in the

CNS and important constituent of the brain ECM is hyaluronic acid (HA).[6] HA is implicated in many cellular functions, including regulating the diffusion of synaptic elements. Specifically, HA rich ECM in the synaptic region of the brain serves to restrict the escape of neurotransmitters and provides a physical barrier preventing the diffusion of the post-synaptic α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid (AMPA) receptor to other areas of the plasma membrane.[6] AMPA receptors are cell ion channels that allow current to pass when activated by bound glutamate and are involved in fast

12

synaptic transmission in the CNS.[10] In this way, HA contributes to maintaining proper synaptic signaling. Chondroitin sulfate proteoglycans (CSPGs) are the most common proteoglycans in the CNS, and include aggrecan, brevican, neurocan, and versican.[11]

These CSPGs are termed lecticans due to their lectin-like domain, or a sugar binding domain.[12] Along with heparan sulfate proteoglycans (HSPGs), CSPGs are known to inhibit axon regeneration, but have been implicated in growth factor retention and presentation in healthy CNS tissue.[6, 7]

Neurons are the fundamental units of the nervous system that process and transmit information by chemical and electrical signaling. Neurons are regionalized specifically to carry out signaling and can either be efferent, sending information away from the brain, afferent, sending information toward the brain, or interneurons, that send information between functional groups of neurons. The dendrites and soma compile and interpret cues from other cells and the surrounding environment (Fig. 2.3A). The soma serves as the

trophic center of the neuron, regulating and producing proteins to be sent to various parts

of the neuron.[3] Located here are most of the organelles common to all eukaryotic cells,

including the nucleus and endoplasmic reticulum. Most cytoplasmic components are

made in the soma and transported to the processes directly or in vesicles. Dendrites

sprouting from the soma mainly function to provide space for synapse, or connection to

other neurons, and interpret the summation of excitatory and inhibitory signals from other

neurons or extracellular space. They take on different formations depending on the type

of neuron, but dendrites are thicker than axons and generally have many branches,

sometimes even containing protruding spines to enhance synaptic reception area.[5]

13

Although synapses may form on the soma or axon, the majority of synapses are located in the dendrites of a neuron.[3]

Figure 2.3. Major cell types of the CNS. (A) Different neurons have varied dendrite and axon conformations depending on their specific roles. The typical neuron has a large number of dendrites to gather information and a long axon to transmit the signal to its target. Insets show the terminus of the axon: Left: the growth cone is present during development and regeneration. Microtubules make up the core of the growth cone, while the peripheral lamellipodia and filopodia are comprised of depolymerized and F-actin. Right: once the axon finds its target it forms a synapse full of neurotransmitter vesicles and mitochondria. The neurotransmitters affect target receptors across the synaptic cleft. Illustration of an astrocyte in red (B) and oligodendrocyte in blue (C).Morphologically each cell follows its name; astrocytes are generally star shaped while oligodendrocytes have many branches, allowing them to myelinate more than one neuron at a time.

The axons are the fiber-optic pipeline of the neuron, passing information at high speeds. Electrical signals, or action potentials, travel away from the soma down the axon in an all-or-nothing manner. The cytoskeleton of the axon is made up of intermediate 14 filaments aligned along the axon and some microtubules constructed from tubulin, although not as many as are found in the dendritic processes. Tubulin is a polymer of repeating α and β subunits whose stability is promoted by microtubule-associated proteins (MAPs) and specific nucleotides.[5] Actin microfilaments are found throughout the axon, but are particularly important at the terminal end of the axon, or the growth cone, and will be discussed in greater detail later. The axon can be shorter, as in a satellite neuron, or very long to carry signals great distances. For example, adult human spinal cord axons can reach a length of several feet.[13] Whatever the length, vesicles and proteins need to be transported quickly along the axon to and from the soma. This is accomplished by motor proteins that use ATP hydrolysis to travel along microtubules; kinesin is responsible for soma to terminus (anterograde transport), and dynein carries organelles or vesicles from the axon to the soma (retrograde transport).[5] Axon transport is extremely important for delivering neurotrophins and neurotransmitters to the terminus, as well as bringing proteins back to the soma for reprocessing. Depending on the molecule or vesicle being relocated, rate of transport in the axon can range from around 1 to 300 mm/d.[3]

Axons have developed an evolved way of traveling to their target cell; the terminus of the axon, or the growth cone, interprets growth and directional signaling molecules and guides the axon along its path. Growth cones contain microtubules in their core and actin in their periphery (Fig. 2.3A). Long filaments of actin (F-actin) are found in the spikes protruding from the growth cone (filopodia) and disassembled actin is found in the web like lamellipodia that are just proximal to the filopodia.[5, 14] Actin is

15

extremely important to the dynamic movement of the growth cone. Permissive substrates

facilitate focal adhesion attachments of the growth cone to the substrate and stabilization of F-actin keeps the filopodium from being retracted. The lamellipodia follows in tow, advancing the growth cone.[14, 15] New cytoskeleton and membrane must be added to the axon as the growth cone hastens forth; constant transport of membrane components to the terminus as well as protein synthesis within the growth cone itself allows significant advancement even if the cell machinery is distant.[5, 16]

Once the growth cone has arrived at its innervating target, a synapse is formed, either chemical or electrical. The chemical synapse includes the pre-synaptic terminal of the axon, the post-synaptic area of the cell being acted upon, and the very small gap (30-

40 nm) between the two termed the synaptic cleft.[3] The pre-synaptic area of the neuron

is filled with mitochondria and synaptic vesicles containing neurotransmitters (Fig.

2.3A). When an action potential is generated along the axon it travels all the way to the synapse, leading to an increase in calcium levels that causes neurotransmitters to be

exocytosed into the synaptic cleft. These neurotransmitters travel to receptors in the post-

synaptic region, diffuse into surrounding tissue, or are endocytosed by nearby

astrocytes.[3, 5, 6] Receptors activated by neurotransmitters change the permeability of the membrane to specific ions, which will lead to either a depolarization (excitatory response) or hyperpolarization (inhibitory response) of the target cell. Synapses in the dendritic region tend to be excitatory, while those on the soma are usually inhibitory.[3]

In electrical synapses, neurons are connected by gap junctions that allow ions to pass

directly from one cell to the next without mediation by chemical transmitters. In contrast

16

to chemical synapses, electrical signaling has no delay, but is only excitatory, is not

amplified, and may be bidirectional.[5]

The main role of astrocytes (Fig. 2.3B) in the CNS is supporting neuronal function by creating and protecting the neuronal microenvironment. There are over one hundred times more astrocytes in the CNS than neurons, and they maintain and mimic neurons directly by absorbing and releasing neurotransmitters into the synapse.[3]

Astrocytes protect the CNS by forming the outer glial layer of the brain and enwrapping

vasculature to create the blood-brain barrier that is infamous for its extreme

selectivity.[3] The structure of the CNS is largely due to the action of astrocytes; they form the outer and inner glial membrane and isolate axons throughout the brain and spinal cord. In the event of injury, astrocytes proliferate and become a glial scar, which is a major blockade for neural TE and will be discussed in greater detail shortly.

Oligodendrocytes are the myelin forming cells of the CNS, serving to surround

axons in an electron-dense myelin sheath. Processes of oligodendrocytes wrap around a

neuron’s axons many times, squeezing out most of the cytoplasm. Layers of lipid rich

plasma membrane are left, tightly encircling and insulating the axon.[5]

Oligodendrocytes have many processes (Fig. 3C) and a single oligodendrocyte can

myelinate 30-60 axons at once. Myelin is an electrical insulator, whose main purpose is

to increase the speed of electrical conduction through the axon while preventing signal

loss. Electrical current cannot conduct through the myelinated portions of the axonal

membrane, it only occurs at small gaps between myelin, termed nodes of Ranvier, which

are several micrometers in length.[3, 5] Not all axons are myelinated, but the ones that

17

are have much faster signal transmission times due to the saltatory conduction of current

“jumping” from node to node along the axon. Injury to oligodendrocytes, and subsequent

demyelination of axons, has been shown to lead to nervous system degeneration.[17, 18]

Similarly, demyelination of axons is the causative factor for the symptoms of multiple

sclerosis (MS).

Microglia are the immune cells of the CNS. Microglia originate from monocytes

that have been trapped in the CNS during development and evolve into a less active state,

or resting state.[3] At any sign of injury or disease these resting microglia proliferate and

may become active, expressing class I major histocompatibility complex (MHC). Due to

their reactivity, microglia are used in research to gauge the extent of an insult to the brain

or spinal cord by detecting the amount of activated microglia in the area.[19-21]

Ependymal cells serve to line the ventricles and central canal of the CNS. These cells, in conjunction with blood vessels in the brain, secrete cerebrospinal fluid (CSF).[3]

Cilia on the surface of ependymal cells circulate CSF in the ventricles. Recently, they have been found to possess plasticity, and have the ability to differentiate into glia of the

CNS in response to specific stimuli.[22-24]

Loss of Neural Function

Injury response and subsequent nerve regeneration is very different in the CNS and PNS, resulting in contrasting outcomes. Due to differences in the two healing environments, PNS axons are able to reinnervate their targets while CNS axons are inhibited by physical and chemical blockades.

18

Throughout the course of this review, different models of evaluation for TE

strategies will be discussed. For understanding the implications of particular situations, a

basic knowledge of injury models and evaluation techniques is necessary. Simulating

injury in the brain and spinal cord has been standardized to some degree to enable global

comparisons of different treatments across research labs. Cut or crush injuries are

generally discussed for the spinal cord. Cut injuries are mimicked in the laboratory by

surgical removal of all or a columnar portion of the spinal cord, termed complete

transection or hemisection, for half of the spinal cord. To simulate a crush injury, the

spinal cord is usually exposed and an electromagnetic or weight drop device is used to

contuse the tissue. Animal disease models also exist where a key feature of a disease (e.g.

demyelination) is simulated genetically for comparison of therapeutic methods.

Once an injury or disease model is created for study, specific methods for gauging deterioration or recovery are used to gather results. “Functional recovery” is a very subjective term and is applied mainly to in vivo work but sometimes it can be applied to

in vitro models as well. Ex vivo cell and tissue are analyzed in a number of ways

including, examination of the cell and tissue anatomy (through histology,

immunohistochemistry (IHC), and microscopy) as well as particular DNA, RNA, protein,

or receptor expression (via polymerase chain reaction (PCR), microarrays, enzyme-linked

immunosorbent assay (ELISA) and patch clamping). In animal models, functional

improvement is usually estimated using behavioral observations and a rating score. For

motor function, exercise tests and open field walking tests are often used. A popular and

systematically defined scoring system is the Basso, Beattie and Bresnahan (BBB)

19

locomotor rating scale.[25] The designers of this scoring system observed rats walking in

an open field and assigned points for movements in paw joints, plantar steps,

coordination, and limb alignment. In addition, exercise tests are sometimes used to assess

motor recovery including swimming, rotor clinging, and narrowing track tests.[26] For sensory evaluation, reflex tests are used; reaction time to a toe pinch or heat application correlate to scoring of the animal’s sensory recovery.

There are two modes through which axons remodel in the body, retraction (small scale pruning of axons and dendrites) and degeneration (large scale elimination of large portion of the primary axon or collateral branches). Retraction and Wallerian degeneration will be covered as well as possible mechanisms for these types of neuronal preservation models. For a more in depth understanding of these occurrences please see.[17]

Retraction typically concerns small scale maintenance of the nervous system where multiple target innervations are eliminated by local pruning of axonal and dendritic branches. The mechanisms of retraction are currently poorly understood; as such, the basic definition of retraction is a change in cell shape, manipulation of the cytoskeleton, signaling pathways, or environmental cues. As described above, cytoskeletal components such as microtubules and actin are involved in growth cone migration. Axon retraction is a dynamic process involving interaction between these factors. Retraction of neurons results when microtubule polymerization is inhibited; whereas, retraction does not occur with blocked reduction in ATP microtubule assembly.[27] Inhibition of dynein on intact microtubules can lead to axon retraction; however, this does not occur when

20

microfilaments are depleted.[28] Therefore, during retraction it is believed that motor

molecules counterbalance changes in microfilaments. Cytoskeletal regulation and

changes result from intercellular signaling cascades. Inhibition of rho-associated protein

kinase (ROCK) blocks Ras homolog gene family, member A (RhoA) downstream

signaling activation on axonal and dendritic retraction in hippocampal neurons.[29]

Inhibitory guidance molecules located in the extracellular environment initiate RhoA

signaling cascades, and are therefore thought to be key players in axon retraction.[30, 31]

Even though many of the causative initiators of axon retraction remain unclear, it plays a

significant role in neuronal development and the establishment of functional connections.

Uncovering and fully understanding these mechanisms more clearly could result in better

models and functional recovery outcomes.

For Wallerian degeneration following injury, a latent period precedes rapid progression into active cytoskeletal breakdown, membrane blebbing, and axon fragmentation (Fig. 2.4).[17, 32-35] Upon injury, degeneration occurs in 3-4 d, however, in lower vertebrates and invertebrates the axon segment can last for a period of more than ten times longer before degeneration proceeds.[36] Although Wallerian degeneration

occurs after a cut or crush injury, it is also similar to axon degeneration observed in the

later stages of some neurodegenerative diseases.[17, 32] Study of this response is

invaluable for understanding axonal degeneration due to this commonality. PNS, CNS,

and explanted nerves exhibit Wallerian degeneration, allowing for controllable Wallerian

degeneration initiation in vitro and in vivo and thus the creation of valuable research

models.

21

Figure 2.4. (A) CNS injury response is extremely restrictive. Damage incurred in the spinal cord is followed by fragmentation of axons and myelin, with limited clearance. Reactive astrocytes form a glial scar, preventing the axons from resynapsis. (B) Axon degeneration pathways showing the scope and complexity of Wallerian degeneration. AAD, acute axon degeneration; EAE, experimental autoimmune encephalomyelitis; ERK-1, extracellular signal regulated kinase 1; NO, nitric oxide; SIRT 1, silent information regulator; VCP, valosin-containing protein; WLDS, slow Wallerian degeneration protein. Images reprinted from [37] and [32].

The discovery of the WldS mutant mouse by Lunn et al. in 1850 accelerated the study of Wallerian degeneration mechanisms and allowed for better understanding of some nervous system diseases.[34, 35] Lunn and associates originally observed that a

22

special strain of mice was able to propagate action potentials for over two weeks

following sciatic nerve transection, whereas, after injury, wild-type mice only carry

action potentials for 1.5 d.[38] This delay was correlated to axon degeneration and facilitated further study of physical and molecular events that occur following nerve injury. Subsequent studies revealed that WldS axons degrade in a more gradual atrophic

process whereas wild-type axon degeneration is self-regulated by pre-existing machinery in the axon, similar in manner but not in mechanism to apoptosis.[17, 33, 39, 40] The wide-held belief is that WldS mice have the ability to either defer or completely suppress

Wallerian degeneration since injured axons die by different mechanisms in these mutants.

Obstructed axonal degradation of WldS mice is dependent and a sole property of neurons

themselves, requiring zero assistance from the neural system or glia of the mutant

strain.[41, 42] Found mainly in the nucleus, WldS is a fusion protein containing nicotinamide mononucleotide adenylyltransferase 1 (Nmnat1), a specific segment of ubiquitin conjugation factor E4 B (Ube4b), and a unique 18 amino acid sequence joining the two.[34, 43] The question remains as to whether WldS protein acts from within the

nucleus by recruiting and directing other pathways or if low concentrations within the

axon are sufficient to interfere with degeneration. There is controversy over whether

slowing degeneration is beneficial to injury recovery. Several studies have shown that in

cases where degeneration was slowed using WldS mice; regeneration was also delayed

and often muted after axotomy.[44, 45] While attempting to compare developmental

pruning to axonal injury, Martin et al. found that regenerating axons in developing

23

zebrafish avoided persistent fragments from induced axotomy.[46] The group speculated

that slow clearance of degenerating axons may be detrimental to innervation.

The complete molecular pathway for Wallerian degeneration is still unknown,

despite the recent identification of several key players within the axon that are associated

with the process (Fig. 2.4B). Axonal transport failure and microtubule dissociation are

believed to be one of the earliest initiators of Wallerian degeneration.[17, 32, 47] Once

the axon is lesioned, transport is limited or completely cut-off. A lag phase follows in which the distal axon is separated from the proximal portion, yet is still capable of conducting action potentials. Once the latent period is over, a catastrophic and active breakdown of the distal segment takes place via innate machinery within the axon. One culprit implicated in axonal degradation is the ubiquitin-proteasome system (UPS).[17,

32, 46, 47] Ubiquitin acts to tag proteins for destruction via proteasomes, enzymes that

denature proteins by peptide bond scission. This system is present in most cells and is

important to many biological processes. Recent studies have shown that pharmacological

and genetic inhibition of the UPS can significantly increase the lag time after axotomy,

but only when administered before a lesion is made.[46, 47] The need for priming of UPS

inhibition suggests its involvement in the early events of Wallerian degeneration.

Additionally, a rise in intracellular calcium ion (Ca2+) concentration is necessary for the

progression of Wallerian degeneration and subsequent regeneration. Increased Ca2+

promotes cyclic adenosine monophosphate (cAMP) activity and neurofilament

breakdown by calpain, a calcium dependent protease.[47, 48] Axon degeneration is often compared to apoptosis, or programmed cell death, because of similarities in the way each

24

works on the cell including targeted ubiquitination.[32-34] Since axon degradation is performed by activation of similar cell proteins to apoptosis, studies have attempted to uncover similar pathways for the two processes. So far, research has revealed that the mechanisms of each are independent; moreover, when distal portions of injured axons were subject to nerve growth factor (NGF) deprivation, UPS inhibition resulted in delayed axon degeneration and only inhibition of apoptosis saved both the soma and axon of deprived neurons.[47]

Neurodegenerative Diseases

Wallerian-like degeneration has been observed in many neurodegenerative diseases, including Charcot-Marie-Tooth, MS, amyotrophic lateral sclerosis (ALS),

Alzheimer’s, Parkinson’s, Huntington’s, and prion diseases, such that axonal transport is disrupted without transection or crush injury.[17, 32-34, 49, 50] The base cause of these

diseases often occurs because of improper or missing axonal transport protein function,

frequently implicating either microtubules or antero and retrograde motor proteins.

Proteins and organelles become trapped in varicosities, or the minor axonal swellings,

leading them to become spheroid formations (major swelling).[32] Aberrant swelling and

spheroid formation within the axon from loss of transport has been linked to initiation of

Wallerian degeneration in disease models.[32, 51, 52] After spheroid formation, the axon

degrades in a strikingly similar fashion to traditional Wallerian degeneration.[51, 52]

Much of the focus of therapeutic Wallerian degeneration in neurodegenerative diseases

has shifted to axonal survival (or sparing) rather than strictly neuronal survival and the

25

prevention of apoptosis. Administration of WldS protein, or portions of it, could

potentially act as a therapeutic agent to delay degeneration as shown in models of

Parkinson’s and motor neuron disease.[17, 49, 53-55] In the case of diseases, slowing

degeneration may have significant benefits as opposed to injury where it could inhibit

regeneration.

Role of Glia in Degeneration & Regeneration of CNS Axons

Owing to different extracellular milieu, CNS neurons have a more difficult time

regenerating than in the PNS. Schwann cells (SCs) are an important component of PNS

regeneration, and act with macrophages to breakdown myelin and form new sheaths to

guide the axon back to its target.[56] SC sheaths, termed bands of Büngner, are important

for isolating the axon and growth cone from the damaged environment. Response to disease and injury in the CNS is quite different, since the guiding glial tubes that protect axons from surrounding environment are not present. Oligodendrocytes do not clear inhibitory myelin debris in the CNS and astrocytes form a glial scar that permanently blocks passage of regenerating growth cones (Fig. 2.4A).[37, 56] Microglia in the CNS

are not nearly as efficient at clearing axon fragments as SCs and macrophages in the

PNS, which can be detrimental to the nerve stump attempting to regenerate.

Some membrane proteins, as well as fragmented and intact myelin from

oligodendrocytes, are capable of inhibiting neurite outgrowth of CNS neurons. The Nogo

family of membrane proteins is known to inhibit axon growth, and two specific inhibitory

domains have been identified in Nogo-A that can both be detected on the extracellular

26 surface of oligodendrocytes.[57] A recent study used gene silencing to knockdown Nogo receptors in transplanted neural stem cells (NSCs), which led to increased functional recovery in rats with TBI over cells transplanted without the receptor silencing.[58] The

Nogo family of proteins exhibits complex interactions with axons.[15, 59] Myelin- associated glycoprotein (MAG) has been shown to not only inhibit neurite outgrowth, but to initiate growth cone collapse.[60] Although MAG has so far proved detrimental to axon regeneration, it is known to encourage embryonic neurite outgrowth and has been implicated in maintaining and encouraging healthy, myelinated axons.[61]

Membrane proteins from oligodendrocytes and myelin are not the only source of inhibition at CNS injury sites. Microglia and astrocytes are recruited to damaged CNS areas, and while some astrocytes may support axons, injury stimulates a reactive phenotype characterized by cell hypertrophy. In response to injury, engorged astrocytes upregulate glial fibrillary acidic protein (GFAP) and vimentin expression in response to cytokines and growth factors released by microglia and other immune cells.[62] Vimentin and GFAP have both been shown to negatively affect axon regeneration.[63] Reactive astrocytes also upregulate CSPG expression, which in turn inhibits regeneration at high concentrations. At the site of injury the concentration of CSPGs is very high but decreases as distance from the center of the insult increases.[64] The mechanism of inhibition from CSPGs is still not completely clear; some evidence suggests the GAG side-chains are to blame while other research points to the protein core. Treatment of injured sites with chondroitinase ABC (enzyme that cleaves GAGs from proteoglycans) decreases inhibition and could thus be used therapeutically to aid regeneration.[37] In the

27

event of injury, the permeability of the blood-brain barrier and blood-spinal cord barrier

is affected, resulting in infiltration of macrophages and cytokines that induce an

inflammatory response.[65-67] Ultimately, astrocytes play a protective role in reestablishing these barriers and preventing the spread of injury; however, the effect is a highly inhibitory environment, caused by glial scar formation and release of neurodestructive molecules, resulting in failed neuron resynapsis and thus permanent loss of CNS function.[68]

Biomaterials for Scaffold Preparation

Definition of Biomaterial & Requirements for Neural TE Scaffolds

Most TE approaches (Fig. 2.1) begin with a biomaterial scaffold of natural or synthetic origin to provide structure for cells while preventing cavitation caused by massive tissue loss. Although the exact make-up of an ideal CNS TE construct is rarely agreed upon, some general requirements are widely accepted.[69, 70] A desirable scaffold is:

• biodegradable with the ability to release therapeutic agents if necessary

• mechanically similar to target host tissue

• easy to manufacture and process

• adhesive for cells or can easily be modified to be cell adhesive

• biocompatible or elicits an appropriate host response minimizing inflammatory

and immune reactions

28

The ECM is extremely important for normal cell behavior and tissue function.[71]

A common approach in TE scaffold design is to mimic the natural environment to facilitate full regeneration of damaged tissue. Often this includes using different materials to recreate the ECM. Surface interactions of biomaterials with both endogenous and exogenous tissue are extremely important due to the dependence of cell adhesion and migration to cellular functions. Chemical or physical surface modifications can be used as a strategy to remedy the unfavorable tissue interactions of some materials. An introduction to materials and common modifications will accompany specific material examples and subsequent sections will carry these topics forward. In depth examination of biomaterials discussed here and others can be found in review papers dedicated to biomaterials.[69, 72-75]

Biodegradable Scaffolds

Non-degradable biomaterials are used in neural TE; however, utilizing a material that can degrade over time and be completely replaced by natural tissue is typically the preferred approach. Most non-degradable materials offer control of synthesis and less complex design considerations, but at the cost of becoming a permanent fixture in the body.[74] Degradable scaffolds eradicate the necessity for surgical removal and allow for complete reinstallation of host function. In addition, when materials are designed to be degraded and eradicated from the body, they do not induce lasting immune or inflammatory responses. In the body, implanted materials can chemically degrade by enzymatic mechanisms or by hydrolysis. Cell interactions with the surrounding matrix

29

are important for homeostasis, allowing for endogenous or exogenous cell migration

throughout the area during regeneration.[71] The polymer on its own must be harmless to

the body; in addition, its monomers should be nontoxic during the length of degradation

time without eliciting any response preventing it from fulfilling its purpose. This often

requires preventing significant alteration to the local cellular environment (e.g., pH,

osmolarity, microglial or astrocytic activation). For example, poly(lactic-co-glycolic acid) (PLGA) degrades into lactic acid and glycolic acid, which can decrease the local tissue pH, encouraging inflammatory responses.[72] Slowed degradation could circumvent pH changes, as long as degradation products are removed from the local environment quickly.

Most naturally derived biomaterials, such as proteins or polysaccharides, can be degraded by enzymes, and do not have toxic byproducts since specific elimination mechanisms exist in the body. In particular, proteolytic enzymes are integral to the processes of tissue remodeling and formation where migrating cells require active control of the ECM. Matrix metalloproteinases (MMPs) have been identified as important proteases in cell migration and ECM remodeling.[76, 77] The study of MMP proteolysis has lead to the identification of specific peptide sequences or substrates for each

MMP.[78-80] These peptide substrates are short (< 7 amino acids) and have been incorporated into polymeric cross-linkers in biomaterial scaffolds.[76, 77, 81, 82] The

MMP-1, or collagenase, cleavable sequence Ala-Pro-Gly-↓-Leu (↓ for cleavage site) has been incorporated into the backbone of photopolymerizable poly(ethylene glycol) (PEG) allowing MMP-1 mediated cell migration through the hydrogel scaffold.[76, 77] The

30

MMP-2, or gelatinase, peptide substrate Pro-Val-Gly-↓-Leu-Ile-Gly (PVGLIG) has been used as a linker for dextran and methotrexate to enable the creation of MMP-2 activated drug delivery microparticles.[82] The PVGLIG peptide has also been included in larger self assembling peptides for the creation of MMP-2 sensitive self-assembled hydrogels.

The gelatinases (MMP-2 and 9) are important in neural development and differentiation.[83] MMP-2 has been shown to be particularly important in postnatal development and in migration [84] and axon outgrowth.[85] In the native CNS environment, MMPs facilitate migration and differentiation, thus, CNS tailored biomaterial scaffolds should allow for material remodeling to promote differentiation, migration and cell process extension.

In contrast to enzymatic degradation, hydrolytic degradation occurs at hydrolytically labile bonds, such as esters, orthoesters, and anhydrides. Due to the patient-to-patient variation in autogenous enzymes, hydrolytic degradation of materials is more predictable over time and location, while holding standard throughout different populations of patients, as compared to enzymatic degradation.[86, 87] Polymers degrade via bulk or surface erosion mechanisms. Bulk deterioration of hydrolytically degraded polymers occurs over the whole volume due to faster water penetration as compared to hydrolytic cleavage of bonds at the surface.[69] Surface erosion of a polymer occurs when water does not penetrate easily and bonds are cleaved before water can reach the inner mass of the material. This type of degradation is characterized by mass loss at the surface that penetrates inward over time. Most often, surface erosion leads to a more linear release of any encapsulated agents included in the scaffold.[69, 88] The mechanism

31

and rate of degradation are typically selected depending on the application.

Enzymatically degradable materials are incorporated more often into TE constructs and

hydrolytically cleavable materials are typically used to achieve stable release profiles in

drug delivery applications. An example of a scaffold with both types of degradation is

examined later, where an enzyme susceptible scaffold is impregnated with a nanoparticle

delivery system with the potential to release therapeutic drugs or growth factors.[89]

Hydrogels are a very popular choice for neural TE scaffolds due to their characteristic similarity to CNS tissue. Hydrogels are polymer networks typically composed of 1-5 wt% polymers that swell with water and can have a low mechanical stiffness, similar to native soft tissues.[69, 70] Mechanical, as well as surface, properties have been shown to be important for neural cell adhesion, migration and survival, as well as the differentiation of stem cells.[90-93] To control the degree of cross-linking, and indirectly the mechanical properties, the polymer can be chemically cross-linked, photo- polymerized, or irradiated either in a dry state or in solution.[69, 73] Photo-initiation of cross-linking in gels is the preferred approach to fill lesioned cavities of irregular shape and size, especially in neural TE where reduced chemical linking is required for soft gel formation. Alternatively, preformed gels can use any cross-linking method and be shaped accordingly. Swelling behavior of a hydrogel influences properties such as nutrient/waste diffusion, surface properties, optical properties, and mechanical properties.[69] Diffusion of nutrients and waste products throughout the scaffold is essential for cell maintenance within the scaffold and for encouraging the ingrowth of homologous cells and tissues when the construct has been incorporated into the body. One way to control swelling

32 behavior is through environmentally sensitive hydrogels. The most common environmental cues are pH and temperature.[69] Researchers have utilized environmentally sensitive hydrogels that exhibit complexing in the presence of a stimulant to initiate physical cross-linking from inter or intrapolymer interactions. For this reason, thermo-gelling polymers have become popular because they can be injected, and cross-link into place at body temperature, obviating the need for invasive surgical implantation.[89, 94-97] Overall, hydrogels are the most common form of CNS scaffolds, and have been widely employed to fill nerve guidance tubes for PNS constructs.[98, 99]

Sample of current Biomaterials in CNS TE

There are many different biomaterials that have been used in CNS regeneration, and those discussed here are not an exhaustive list. The following discussion should serve as an introduction that is designed to provide the reader with a basic understanding of common neural TE biomaterials, with specific examples of past applications and unique features of each material. Reiterated throughout the literature is the commonality that a successful TE construct requires a composite of many stimuli built into the scaffold.

Many of the materials discussed here are typically used as copolymers, blends, or in conjunction with other molecules to encourage the appropriate tissue responses both in vitro and in vivo.

Many researchers have used the body’s most abundant ECM component, collagen, to form constructs for neural TE. Collagen is composed of chains of amino acids linked by peptide bonds, or the covalent bond between the amino group of one

33

amino acid and the carboxylic acid of another amino acid. These chains form secondary

structures and tertiary structures; combinations of multiple tertiary structures form higher

order structures and finally collagen fibrils.[72] Over 20 collagens have been identified,

however, collagens I-IV are the most abundant in the body and only not all collagen types are found in the brain.[6, 72] Since collagen is native to the body it is recognized by cells, promoting adhesion, however, with the potential for immune response. For this reason, allograft or xenograft collagens are especially avoided due to their immunogenicity and possibility of disease transfer.[72, 100] Enzymatic degradation of collagen is carried out by MMPs at specific cleavage sites. As previously noted, collagen I can be cleaved by

MMP-1 to allow for matrix remodeling and cell migration. MMP-2 and -9, both

gelatinases, can degrade collagen IV, a common basement membrane protein found in the

CNS.[83] As an insoluble protein, collagen is most often dissolved in a cold, acidic

solution and can be made into a hydrogel by adjusting the pH and temperature to physiological conditions.[74] In this way, collagen can be used as an injectable hydrogel

for TE applications. In contrast to physically formed gels, chemical cross-linking of

collagen (by aldehydes, carbodiimides, polyepoxides, hexamethylene-diisocyanate, etc.)

is often used to control scaffold properties such as the mechanical properties, degradation

time, and immune reactivity.[72, 100] Concentration and the degree of cross-linking can

be used to adjust the stiffness of collagen gels, as discussed earlier; collagen can also be

blended with other materials to increase mechanical compliance or stiffness, depending

on the additive. As a native ECM component, collagen has integrin binding sites, and can

be further enhanced for neuronal regeneration by incorporating growth factors such as

34

neurotrophins. Neurotrophin-3 (NT-3) was recombinantly functionalized with a collagen binding domain (CBD) allowing it to complex to collagen I scaffolds; these scaffolds were subsequently implanted into the transected spinal cord of rats with positive functional results.[101] The collagen-NT-3 combination yielded significant locomotor improvement over collagen scaffolds with soluble NT-3. NGF was also functionalized with a CBD and used as an injectable scaffold for sciatic nerve regeneration.[102]

Gelatin, a hydrolyzed derivative of collagen, can also be used in neural TE to make gel scaffolds.[103, 104] Collagen can be processed into gels, meshes, and even a powdered form to suit a variety of needs. In addition, collagen can be electrospun into a fibrous mesh and cross-linked for stability, leading to excellent neural outgrowth responses in vitro.[105, 106]

Since it is a regular component of the ECM, HA (hyaluronic acid) is being studied as a biomaterial for the CNS.[69, 73] HA is very viscous due to its high molecular weight and can form soft gels via chemical cross-linkers, such as carbodiimide, or modified for photocross-linking.[74, 107] Natural degradation of HA occurs in the body because of interaction with free radicals, MMPs, and hyaluronidases allowing for native remodeling of these scaffolds.[72] Increased spreading and attachment of hippocampal neurons on

HA gels as compared to unmodified HA gels were achieved with immobilized Nogo receptor antibodies (antiNgR).[90, 108] Further, HA-antiNgR gels were able to support

NSC survival and differentiation.[90] HA on its own has poor mechanical properties, thus it is often blended with polymers and stiffening agents to increase rigidity. HA has been used with collagen and methylcellulose to decrease their compressive elastic moduli

35 closer to ~1 kPa, which is similar to brain tissue.[97, 107] HA may be preformed or injected to form brain and spinal TE constructs.

Methylcellulose (MC) is an inverse thermogelling polymer that is derived from cellulose (a polysaccharide found in plant cell walls), and has been shown to invoke a low inflammatory response in vivo.[70, 73] Due to its hydrophilicity, MC has low cellular adhesion unless modified, but shows biocompatibility in vitro and in vivo.[109, 110] An injectable hydrogel composite of HA and methylcellulose (HAMC) incorporating synthetic polymer nanoparticles showed very little microglial response when injected into the intrathecal space of rat spinal columns.[89] The HAMC combination allowed for fast gelling at physiological temperature.[97] Blank nanoparticles were included as a test for a drug delivery vehicle that could release therapeutic agents in future studies.[89]

Chitosan is derived from chitin, a naturally abundant polysaccharide found in the shells of crabs and other shellfish. Considered biocompatible by most researchers, chitosan may illicit an inflammatory response through macrophage activation with low deacetylation, therefore, it is typically used at deacetylation percentages above 80%.[111]

Chitosan is recognized to have antibacterial properties, which are a useful property for biomaterial scaffolds.[112] Chitosan can be covalently bonded or thermogelled in the presence of glycerophosphate salt or blending with a thermogelling polymer.

Thermogelled chitosan was able to support comparable mouse cortical neuron survival and neurite outgrowth in vitro to cells grown on poly(D-lysine), a common adhesion enhancing agent.[113] Alternatively, a methacrylated form of chitosan has been synthesized that can be formed into hydrogels via exposure to UV light in the presence of

36 a photoinitiator.[114] This methacrylated chitosan can support the survival and differentiation of cultured NSCs and allows for the easy tuning of substrate stiffness via photoinitiator concentration or exposure time.[92, 114] Chitosan can be formed into a cryogel, either alone or blended with other polysaccharides, similar to alginate and agarose.[115-118] In addition, chitosan blends have the capacity to be electrospun, and these scaffolds have been used in PNS TE.[119, 120]

PEG is a hydrophilic polymer that is biocompatible and typically nonfouling and therefore often requires modifications to increase cellular adhesion.[73, 74] One way to adjust the surface interactions of PEG or its degradation characteristics is through polymer blends or composites. Bjugstad et al. recently performed a comprehensive assessment of the biocompatibility of PEG hydrogels in the brain. In the study,

PEG/lactic acid (PEG/LA) gels were injected into rat cortexes, with the slow degrading

(less LA) and non-degradable (no LA) PEG gels demonstrating the lowest microglial and astrocytic response.[121] The non-degradable and slower degrading gels showed less glial activation in a 50-200 µm region surrounding the implant than the sham. While LA is useful for making a PEG based scaffold degradable, it likely leads to observed glial activation by increasing the acidity in the local environment. In other studies, photopolymerized LA/PEG hydrogels were utilized as a delivery system for the neurotrophic agent NT-3.[122] When the scaffolds were implanted into hemisectioned rat spinal cords, improved movement and coordination, axon ingrowth, and host NT-3 concentrations were observed.

37

Neural TE scaffolds exhibit strict demands on their bulk biomaterial properties as well as surface characteristics. The materials discussed here all possess strengths and flaws, which should be weighed against another depending on the specific application. In many studies, natural and synthetic materials can be blended or conjugated together for improved properties and to gain additional functionality. For CNS TE, it is important to remember the restrictive environment that can be created by microglial and astrocytic activation, and to make sure any material used in a TE scaffold does not illicit any undesirable response. Subsequent discussions will also provide insight into further surface modifications, physical and chemical, that can mask bulk material properties and further enhance cellular responses of the scaffold material.

Cell Sources for CNS TE

When designing CNS tissue constructs, inclusion of encapsulated and surrounding cells are of great importance to achieve tissue regeneration (Fig. 2.1). Biomaterial scaffolds discussed previously are commonly impregnated with cells to expedite recovery by replacing lost tissue or by decreasing the migrational distance of natural cells in the surrounding area. In this section, a brief overview of commonly used and promising cell sources follows, and is meant to give the reader an introduction to each one.

TE design criteria for cells are similar to those for biomaterials, in that any elicited response that impedes regeneration is unacceptable. For this reason, autologous cells are the gold standard for TE since they rarely evoke an undesired response (there are exceptions, discussed later). However, the price of using the patient’s own cells is paid by

38

donor site morbidity, as well as additional time spent on surgical isolation and culture. In

addition, care must be taken to correctly choose cells for different injury and disease

states. For demyelinating wounds and diseases (e.g. MS), reestablishment of the myelin

sheath by glial cells is of utmost importance. In contrast, when neurons or both neurons

and glia are lost, typically strategies shift focus to the neuron. Subsequent discussion of

specific cell types will show that opinions differ on the optimal therapy in these cases.

Some researchers attempt neuronal implantation while others incorporate glia or stem

cells to encourage native neurons to regrow through the lesioned area. An overall strategy

for cell incorporation or stimulation is not agreed upon, and is usually formulated for the

specific application.

Somatic cells of the CNS are rarely used as a cell source for CNS TE. Issues of

secondary injury sites, surgical accessibility, and poor mitotic ability restrict the use of

primary cells. As an alternative, the use of the peripheral myelinating cells have become

popular, this includes protective ensheathing cells within the nasal cavity. During

homeostasis and development, glia serve as the supporting cells of the neuron. Most CNS

TE strategies incorporating glia focus on maintaining unaffected surrounding tissue while remyelinating damaged tissue.

Stem cells offer an attractive alternative to somatic cells, and provide increased

cell expansion and decreased donor site morbidity. Stem cells can be derived from a

number of sources, and depending on their location and age they possess different

variations of two intrinsic stem cell characteristics: differentiation (or ability to form

multiple cell types) and proliferation (or ability to self-renew). Since they can be readily

39

expanded, less stem cells are needed initially to achieve high cell numbers compared to

somatic cells. Totipotent stem cells have the ability to become any cell type of the body,

as well as expand indefinitely; from here differentiation potential or plasticity decreases

to pluripotent, to multipotent and finally to progenitor stem cells. Pluripotency is possessed by cells very early on in development, namely embryonic cells, but also has been induced in somatic cells by genetic alteration. Multipotent and progenitor stem cells

persist throughout the body in stem cell niches into adulthood, and several populations of

adult NSCs reside within the CNS.

Primary Cell Treatment of CNS Injury: Glial Cells

As mentioned previously, SCs (Schwann Cells) are the myelinating cells of the

PNS. SCs wrap around the axon many times allowing multiple SCs to myelinate a single axon. In contrast to oligodendrocytes in the CNS, SCs resorb fragmented myelin and the distal axon from an injured site, recruit macrophages, and then work in synergy to clear debris and encourage axon resynapsis.[123, 124] SCs play an encouraging role in PNS regeneration and serve to augment axon synapse reformation. They do this by transforming to a non-myelinating phenotype, increasing secretion of axon recruitment

factors brain derived neurotrophic factor (BDNF, sometimes referred to as BDGF),

ciliary neurotrophic factor (CNTF), and NGF, and forming a that tube guides the growth

cone of the proximal axon to its destination.[125, 126] Though not typically found in the

CNS, SCs are capable of myelinating CNS neurons; and endogenous SCs have been

found to migrate into the injured spinal cord and myelinate CNS axons.[127-129] The

40

mechanism of SC migration into the CNS is still unknown, as is the extent of their

participation in CNS nerve regeneration, though it is known that their contribution is

minimal.[130, 131] Encouragement of SC migration into the injured spinal cord has not been well researched; rather, the majority of studies to date have injected or transplanted

SCs into the injured area in an attempt to encourage regeneration. SC myelination of axons is limited to regions with low astrocyte numbers. In vivo and in vitro, astrocytes and SCs are known to inhabit mutually exclusive areas. Early studies by Blakemore et al.

created areas of demyelination in the cortex or spinal cord with ethidium bromide, which

resulted in low or compromised astrocyte populations.[132, 133] SCs were able to

myelinate endogenous axons in the areas with decreased astrocytic inhabitance.

Modifications to SCs using exogenous proteins or transcription factors increase SC interactions with astrocytes and improve their migration and integration into the CNS.

Recent work has focused on altering SC neural cell adhesion molecules (NCAMs) by inducing SC expression of polysialic acid (PSA) through viral delivery of sialyltransferase X (STX).[134, 135] PSA associates with the fourth domain

(extracellular portion) of NCAM and decreases adhesiveness so that the cells can more

easily separate. Expression of PSA-NCAM is found on oligodendrocyte precursors during development and regeneration, and has been found to increase SC migration into astrocyte territory without adversely affecting their myelination capabilities.[134] Adult

primate SCs were transplanted near experimentally demyelinated areas of the spinal

column and demonstrated faster and more efficient migration when transfected with STX

viral vectors.[135] In this study, remyelination was enhanced in transfected SCs;

41

however, no functional analyses were conducted. One significant advantage of SCs is their ease of isolation and expansion capabilities. Obtaining SCs by biopsy, expanding them in culture with mitogenic agents, and purifying them from fibroblasts provides a

source for autologous cells.[136-138] Glial growth factor (GGF) and the cAMP activator

forskolin are two popular mitogenic agents, but as with any mitotic factor, their use must

be closely monitored and inhibited before implantation to avoid tumorigenesis.[136, 137,

139, 140]

Pluripotent Stem Cells: Embryonic Stem Cells

Pluripotent stem cells, namely embryonic stem (ES) cells, have the ability to become neurons or glia for use in CNS TE when provided the correct cues. ES cells have also been proposed as an abundant source for NSCs, not only for use in cell based therapies but for screening assays and disease progression studies. It has even been argued that NSCs derived from ES cells retain better differentiation capabilities or

‘developmental competence.’[141]

One attractive property of ES cells for neural TE, other than their excellent proliferative capabilities, is their predictable behavior in response to developmental cues.

Embryonic cells are derived from the inner cell mass of the blastula, and neuronal cell induction begins once the blastula enters gastrulation. Development of the neural system begins with induction of neural precursors which are differentiated to neurons and glia, proceeded by axon journey and preliminary synapse formation to their targets. Neural functionality is made possible by final remodeling of the neural network. Neural cells

42

arise from the ectoderm, or the outermost layer of the gastrula, on the dorsal side where

the inhibitory bone morphogenetic proteins (BMPs) are suppressed by the organizer

molecules noggin, chordin, follistatin, cerberus and Xenopus nodal-related 3 (XNr3)

protein (Fig. 2.5).[5, 142] After the inhibition of BMPs, ectodermal cells are permitted to

become neural precursor cells and are directed further by neural promoting molecules.

Once the neural tube is formed in the embryo, spatial patterning occurs to direct cell fate.

Caudal formation is led by Wnt-8, basic fibroblast growth factor (bFGF), and retinoic acid (RA).[143, 144] The spinal cord is patterned ventrally by sonic hedgehog (Shh)

protein and RA, and dorsally mainly by BMPs.[145, 146] The PNS is derived from

neural crest cells that migrate from the neural tube; neural crest cells may become PNS

neurons and glia, smooth muscle cells, or pigment cells.[147-149] The cell type of neural

crest cells is determined by migration route and environmental cues encountered. Events

directing development have been well studied and are often mimicked by applying

certain factors in a sequential manner to ES cells in order to derive NSCs and even specific somatic cell types.[150-152]

43

Figure 2.5. Diagram of neuronal development from the blastula in Xenopus (A) and chick (B), illustrat-ing the roles of BMP, FGF and Wnt. For both species, FGF promotes neuronal induction by blocking BMP activity. Chd, chordin; Nog, noggin; NXR, nodal- related factors. Image reprinted from.[494].

Defined differentiation protocols eliminating the use of serum are in development to improve the clinical relevance of ES cells. In many cases, neural differentiation is initiated by culturing ES cells on feeder cells (typically embryonic mouse fibroblasts), followed by culture as suspended embryoid bodies with RA. This culture regime has been found to promote a neuronal phenotype and suppress mesodermal cell types.[153, 154]

Embryoid bodies are subsequently dissociated and plated on laminin coated surfaces with

N2, a serum free neuronal supplement, in media for the neuronal phenotype.[154] Motor

neuron specification has been achieved by activating the Shh pathway subsequent to RA;

RA is necessary for early neural induction but should be followed by other factors if

further differentiation is desired.[150-152] This differentiation regime mimics

development closely, as ES cells are given caudalization signaling from RA and then

sequential ventralization signaling from Shh. Testing of in vitro differentiated ES cells by

44

most groups involves immunostaining or electrophysiology testing, however, Wichterle

et al. alternatively implanted differentiated motor neurons into developing chick embryos

to test functionality.[152] This group observed that ES derived cells localized into the

correct area of the spinal cord and projected axons to the appropriate peripheral regions.

Directing ES cell differentiation has not been limited to development-linked factors and

molecules. A recent study found that soluble amyloid precursor protein (APP) not only

induced neural progenitor differentiation but also led to ES cell expression of βIII

tubulin, a common neuronal marker, in only 5 d.[155] APP, a protein thought to be a synapse regulator, is more well known for its transformation into amyloid plaques in

Alzheimer’s disease.[51, 52]

Since success has been achieved in directing ES cell fate into neural cell types, focus has shifted on incorporating ES cells or ES-derived cells into functional neural TE constructs. In an attempt to enhance serotonergic fibers two weeks after a hemisectioned

SCI, embryonic neural precursor cells (NPCs) from the neural tube were injected caudal to the injury with generally positive results.[156] Even though the NPCs showed superior

cell morphology and axon extension in tissue sections, functional improvement of

coordination and maneuverability were comparable to injected mesenchymal stem cells

(MSCs) and both showed significant improvement over control mice. Similarly,

successful treatment of a simulated crush injury demonstrated the regenerative efficacy of

human NPCs transfected to express Neurogenin 2 (Ngn2), a proneural factor that inhibits

astrocytic differentiation, injected a week post-injury in adult rats.[157] NPCs grafted

without Ngn2 did not result in any functional improvement; however, NPCs that

45 expressed Ngn2 resulted in significantly higher scores in functional motor skill tests. This study by Perrin et al. showed the utility of human embryonic NPCs in a more realistic model where induced cells were administered a week after injury. Outside of the research lab, injury to CNS tissue is not treated immediately following insult due to delays in diagnosis, treatment availability, not to mention many other obstacles. The ability of human cells to perform well in animal models coupled with a more realistic delay of treatment makes this work very exciting for future CNS regeneration treatments.

Induced Stem Cells

Ethical considerations surrounding ES cells and the associated controversy have dampened the enthusiasm for incorporating them as a cell source into TE strategies. In recent years, an alternative source of pluripotent stem cells is under active development.

In 2006, Shinya Yamanaka’s group demonstrated the creation of pluripotent stem cells from adult fibroblasts using gene therapy to introduce Oct3/4, Sox2, c-Myc, and Klf4 expression.[158] Yamanaka’s group termed these cells, ‘induced pluripotent stem (iPS) cells’, and observed multiple lineage formation after injection into adult nude mice or blastocysts. Development of two factor iPS production (only Oct4 and Klf4) from mouse

NSCs may offer better clinical relevance since it eliminates the c-Myc gene that is a well know oncogene implicated in tumorigenicity.[159] iPS cells behave similarly to ES cells and can be induced to form neurons in a similar manner (Fig. 2.6).

46

Figure 2.6. Diagram of differentiation paths for ES and iPS cells into specific populations of neurons. hESCs, human embryonic stem cells; hips, human induced pluripotent stem cells; hNPs, human neu-ral precursor cells; DA, dopaminergic neurons; MN motor neurons; PNS, peripheral nervous system derivatives; EB, embryoid body. Image reprinted from [201].

Despite the attractiveness of using the patient’s own cells and avoiding embryonic cell use, iPS cells still have many problems. Transfection of genes has inherent difficulties. Viruses are typically used to delivery genetic material to generate iPS cells since they are equipped with natural mechanisms that can penetrate the cell wall and release nucleotides into the tight security of the cell nucleus. The downside is that viral gene delivery by retroviral and even lentiviral vectors causes random insertion into the host genome and could lead to unwanted mutations.[160] Adenovirus vectors, as well as non-viral methods, do not carry the same threat of random genome insertion but are often less efficient.[160] Alternative methods to viral gene delivery have been developed that

47

are more effective, involving nanoparticle delivery or the use of commercial transfection

kits.[161-163] Electroporation using commercially available systems allows increased

permeability of the cell and nuclear membrane and has been used to deliver a single

plasmid containing all four genes for induction of iPS cells.[161] Other commonly used

methods include complexation with a cationic polymer or lipid to stabilize DNA and

cross the cell membrane.[164] In depth discussion of other nonviral systems and

subsequent uptake into the cell and nucleus can be found in other reviews.[165, 166]

Although iPS cells are still far from clinical studies in CNS TE, they are being

actively developed for diagnostic uses. It is proposed that patient specific cells could be

used to screen drugs and therapeutic treatments, or conversely to study genetic and

contracted diseases using patient iPS derived neurons or glia.[167] Disease specific iPS

cells are currently being used to investigate neurodegenerative disorders including ALS,

Parkinson’s, and Huntington’s disease, among others.[168-170] Continued work with iPS

cells may prove them valuable in CNS TE approaches in the future.

Adult Stem Cells: Endogenous Stem Cells in the Brain & Spinal Cord

Stem cells niches throughout the postnatal body retain populations of multipotent

cells into adulthood. In contrast to ES or iPS cells, NSCs within the brain and spinal cord

closely interact with their surrounding cells and are committed to the neural lineage.

Adult NSCs may also offer a more popularly acceptable source of stem cells because they lack the stigma that surrounds embryonic research and possess decreased tumorigenicity.

Defined populations of multipotent adult NSCs have been identified in the subventricular

48

zone (SVZ) of the lateral ventricles, the dentate gyrus (DG) of the hippocampus, and

around the central canal of the spinal cord.[171]

SVZ and DG cells can differentiate into neurons, oligodendrocytes, or

astrocytes.[172, 173] In vitro, these two stem cell populations behave very similarly.

Both areas in the brain produce neurons under non-pathological conditions. Physiological neurogenesis in the DG takes place in the innermost region of the subgranular zone of the hippocampus, quickly becoming neurons with mature phenotype that incorporate into the granule layer.[174-177] Cells from the SVZ naturally migrate along a track lined by astrocytes to the olfactory bulb where they differentiate into granule cells.[178-181] The

rostral migratory pathway is specific for these migratory cells, and transplantation of

other neurons into the SVZ does not result in migration to the olfactory bulb.[178] On the contrary, NSCs from the DG of the adult hippocampus can be transplanted into the SVZ migration pathway and will differentiate into neurons typical of the olfactory bulb.[182]

Adult NSCs are of interest for endogenous activation or exogenous transplantation for

TBI or brain diseases owing to their multipotency and natural brain origin. Their

implantation has been shown to decrease recovery time in SCI and enhance tissue

bridging when used in conjunction with a nerve guidance conduit (NGC).[183]

Due to the delicacy of brain surgery and implantation, endogenous activation of

the SVZ or DG populations is a motivating interest for many researchers. Multiple injury

models have been used to stimulate cell activity in the NSC niches of the brain. In a study

by Rice et al., TBI was induced in adult rats and proliferating cells were labeled and

counted in tissue sections, which revealed increased proliferation and migration of cells

49

from both the SVZ and DG post injury.[184] Interestingly, results from this study showed

significantly timed ’waves’ of increased mitotic activity from the DG. Similarly, after

traumatic axonal injury, proliferation and migration of SVZ and DG cells, as well as an

increased gliogenesis, persisted even eight weeks post-injury.[185] After inducing demyelinated lesions in the mouse brain, SVZ cells were observed to expand and migrate towards the lesion.[186] Observations show that NSCs in the adult brain respond differently to different injury models, presenting astrocytic preference in response to TBI and oligodendrocyte preference after demyelination.[187] Alternatively to injury-induced activation, injections of different neurotrophic agents have been studied for stimulation of growth, migration, and differentiation of NSCs in the brain. System perfusion of insulin- like growth factor I (IGF-I) in rats produced marked proliferation and neuronal differentiation in the DG without any increase in astrocyte production.[188] In the rat

SVZ and olfactory bulb, proliferation and neuronal differentiation was induced by viral delivery of BDNF.[189] New neurons persisted 5-8 wks after the adenoviral injection into the lateral ventricles. It is clear that NSCs in the brain will respond to injury and a local or systemic delivery of neurotrophic agents. A properly designed CNS TE system has the potential to augment the stimulation of large populations of new neurons and glia from the SVZ and DG.

In vitro, SVZ derived adult NSCs can be induced to differentiate into neurons and glia by a variety of factors including IGF-I, NGF, BDNF, angiopoietin-1, cAMP, BMP-2, platelet derived growth factor (PDGF-AA), and the cytokine interferon gamma (IFN-γ), among others.[190-194] IGF-I has similar action in vitro to in vivo, and is mitogenic and

50

neurogenic on NSCs.[195, 196] The combinatorial administration of IFN-γ in

combination with an analog of cAMP to SVZ NSCs leads to a significantly higher

population of neurons than soluble administration of neurotrophins, NGF, and BDNF, or

tumor necrosis factor alpha (TNF-α).[193, 194, 197] Even though biochemical treatments have been found to induce neuronal and glial differentiation for transplantation into the injured CNS, the question still remains whether these cells will integrate into the existing network of tissue, resulting in functional recovery.

Ependymal cells located near the central canal of the spinal cord have been identified that possess high proliferation rates and multipotency under specific conditions.[23, 24, 171, 198-201] Similar to NSC responses in the brain, injury of the spinal cord results in a protective response of the spinal cord ependymal cells.[198, 199,

202] Additionally, introduction of EGF and bFGF into the spinal column leads to increased proliferation of ependymal cells.[203] These results are expected, since EGF and bFGF are mitogenic agents, well known for their use in the in vitro expansion of

NSCs. Ependymal NSCs were less effective than SVZ NSCs as a direct cell treatment for

SCI in rats.[183]

Obtaining adult NSCs from the brain and spinal cord is difficult and risky to the patient, which is the biggest drawback of these cells. However, isolating these cells has been shown possible and further advances in surgical techniques will increase their possibility for clinical use.[204] Another drawback of adult NSCs, in contrast to pluripotent cells such as iPS and ES cells, is that these cells show decreased proliferation with age.[205] Research with NSCs is still extremely useful as it could be applied

51 directly to NSCs derived from ES or iPS cells. TE strategies that target endogenous activation of stem cell populations in the brain and spinal cord would be useful cues to include in CNS constructs. In this respect, incorporation of these signals into a degradable scaffold could allow for a long release time to facilitate long-term regeneration by native tissue stimulation.

Mesenchymal Stem Cells

MSCs have also been utilized for the purpose of CNS therapies. These multipotent stem cells are derived from a number of locations in the adult body and have the capability to differentiate into many different types of tissues with varying efficiency

[206]. One of the main sources of MSCs is the bone marrow; these cells can be differentiated into smooth muscle cells, osteoblasts, chondrocytes, cardiomyocytes, liver cells, SCs, and to some degree, neurons.[207-212] In addition, MSCs isolated from the umbilical cord have similar characteristics and can express neuronal phenotypes following neural induction.[213-215]

MSCs are able to promote regeneration in neural TE; however, experts are unsure of the therapeutic mechanism by which they accomplish this. MSCs may transdifferentiate into neurons and glia to augment tissue regeneration; alternately, inflammatory and immunological agents may be recruited to the damaged site by way of

MSCs.[216-218] A secondary protection mode of MSCs is their secretion of neuroprotective factors that provide neural sparing and encourage endogenous axon regeneration. Crigler et al. studied MSCs for gene coding and expression of neurotrophic

52

factors and found cultured MSCs could express BDNF and NGF.[219] Further testing

showed that neuronal blastoma cells as well as explanted DRG both co-cultured with

MSCs demonstrated significantly increased neurogenesis and neurite outgrowth. In these

studies, inhibition of BDNF activity resulted in small decreases in outgrowth and

proliferation, but the effects of MSCs were not completely negated, suggesting that other

neurogenic factors are also secreted by MSCs.

Transplantation of undifferentiated MSCs into SCIs has shown positive

therapeutic effects. Adult bone marrow MSCs transplanted two days after a spinal

contusion injury showed neuroprotective effects that led to increased myelination at the

injury site over the control group.[220] Histological analysis revealed increased laminin

expression with MSCs along with neurite extension and alignment with spinal cord

direction, but functional analysis using BBB scoring showed no significant difference

between MSC treated injuries and the controls. Results suggested that MSC expression of

laminin was responsible for aiding neurite alignment. As mentioned above, MSCs are

also known to differentiate into myelinating cells. Three days after a focal demyelinated

lesion was made in adult rats, undifferentiated MSCs delivered to the site helped to

improve electrical conduction velocity across the lesion.[221] Spinal cord axons treated with MSCs contained myelination typical of the PNS, suggesting MSC differentiation and myelination within the wound. Direct injection and intravenous administration of

MSCs in response to a demyelinated lesion in the rat spinal cord revealed that remyelination occurred in a dose-dependent fashion, as shown by histological examination.[222] This work also demonstrated the effectiveness of intravenous delivery

53

of MSCs; the treatment is less invasive, although, almost 100 times more cells had to be

used. MSCs also have aided healing in brain injury models. In an in situ environment, co-

transplantation of MSCs with NSCs into hippocampal slice cultures led to the majority of

NSCs differentiating into oligodendrocytes.[223] In contrast, when the NSCs were

transplanted in the tissue slices alone, the majority became astrocytes. This effect

translates to in vivo studies as well. A recent study involving MRI tracking of tissue and

cortical blood flow (CBF) showed that administration of MSCs in the brain significantly

reduced ventricle expansion and helped maintain CBF in regions adjacent to injury.[224]

Compared to saline injections, the MSC-treated rats demonstrated significant functional improvements as assessed by neurological severity score and the Morris water maze test.

From a TE standpoint, MSCs could provide an efficient and elegant way of administering therapeutic agents in multi-cued constructs for CNS injury and disease. They provide neuroprotection, can be manipulated to secrete many growth factors, and can differentiate into myelinating cells. Ongoing research suggests MSCs could be used as a source of growth factors as well as for inflammatory and immune agents. Thus, incorporation of

MSCs may address several issues at once. Isolation of cells from a non-invasive, although painful, bone marrow biopsy procedure is also very attractive to limit extra surgical procedures outside of treatment. In depth investigation of the long term effects of

MSC transplantation are underway, and based on the above findings, MSCs present a promising choice for cell therapy for CNS regeneration applications.

One flaw that is redundant throughout adult cell therapy techniques is the decreased plasticity and activity of aged cells. A comparison of SKPs derived from

54

different regions of skin from patients 8 months to 85 years old revealed that proliferation

and differentiation capabilities drastically declined in cells from the elderly.[225] SKPs

still have many advantages and continue to be developed for additional CNS TE

applications.

Many cell sources exist to choose from for treatment of CNS injuries and

diseases. The specific needs of TBI, SCI, and neurological diseases call for tailored

approaches when developing and optimizing neural regenerative methods. TE constructs

that incorporate cells and/or encourage ingrowth have the capability to augment and

accelerate the restoration of normal CNS function, improving quality of life.

Stimulation & Guidance

Multi-cued CNS TE constructs are made possible through incorporation of

surface and soluble prompts combined with scaffolding and cell components. Cell

migration in combination with process extension are essential early steps in nervous

system development, thus developmental signals are often the source for stimulation

schemes aimed at controlling cell behavior and reinnervation after injury. Immature cells are partitioned and guided to the appropriate targets by a complex mixture of chemical and physical cues. During nervous system development, neurons rely on the surrounding environment for guidance in order to innervate their targets and create a functional neural network system. Initially, neurites extend from the soma in all directions, and some

gradually unite to become the primary axon while remaining neurites form

dendrites.[226, 227] The growth cone, located at the tip of the axon (Fig. 2.3A), serves to

55 decipher chemical and physical cues, both in space and time, determining the axon’s trajectory.[228, 229] As it senses these signals, the growth cone pauses and enlarges as the cytoskeleton reorganizes, preparing itself for the next move.[230] As discussed earlier in this review, in the periphery of the growth cone are located lamellipodia that contain a web of actin. From this region, finger-like projections called filopodia protrude and sense the environment. Microtubules are responsible for transmitting this information from the growth cone to the soma and back to the tip. Axon branching can occur where the growth cone will split or interstitial branches will form from actin remnants as a result of cytoskeletal changes.[230] Guidance of the growth cone is vital to properly guide axons to their targets. Several strategies for encouraging and directing growth cones to specific targets will be introduced including physical, chemical, and electrical applications. Most can be used in conjunction with each other for complex control over neuronal behavior.

Physical Cues

Guidance and stimulation of cells can result from physical connection with the environment around them. Cells are sensitive to changes in substrate stiffness, mechanical stresses as well as topographical changes. Analyzing how physical cues modulate cellular adhesion, differentiation and guidance will provide a framework for developing optimal biomaterials and bioreactors to optimize translation of neural TE constructs in vivo (Fig. 2.1). Therefore, material stiffness, physical elongation and topographical guidance methods used for neural regenerative strategies will be outlined here.

56

Many biomaterials used in TE have tunable mechanical properties that can be

adjusted by material blending and degree of cross-linking. This is important for

biological applications, because cells tend to favor their native tissue moduli.

Additionally, during development and in certain disorders, changes as well as gradients in

ECM stiffness are important for proper development and healing responses. Cell

interpretation of mechanical stimuli and the resulting response, including the activation of

downstream pathways, is known as mechanotransduction. Vascular tissue and cell

migration mechanotransduction is better understood compared to responses in the

nervous system; thus, there are still many questions concerning the pathways activated in

response to mechanical forces in the nervous system.[231] Cells adhere to surfaces and exert contractile forces in order to migrate to different areas. In CNS and PNS TE, low substrate elastic modulus (E) is important for neuronal outgrowth.[232, 233] Several studies using DRG have shown that gels with lower stiffness result in more branching and longer neurites than stiffer gels.[234-236] In a study by Flanagan et al., mouse spinal

cord neurons and glia were grown on polyacrylamide gels with elastic moduli ranging

from 50 to 550 Pa.[232] This group found that on the softer acrylamide, neuron

branching was three times higher and glial survival decreased. Gradient stiffness gels can

induce neuronal durotaxis, as neurites have grown significantly longer down a decreasing

stiffness gradient.[235] Interestingly, a threshold effect has also been observed when

stiffnesses were higher than a shear modulus (G) of 100 Pa (E ≅ 600 Pa); neurite outgrowth was still present at higher moduli but was not as pronounced as in the range of

G = 10-100 Pa (E ≅ 60-600 Pa).[237] Differentiation into a specific lineage or cell type

57

can be specified by culture substrate compliance. Leipzig et al. found that extremely soft

gels with stiffness, E around 800 Pa, will elicit neuronal differentiation from NSCs, while

slightly stiffer gels with an elastic modulus around 7,000 Pa yields an oligodendrocyte

phenotype.[92]

The neuronal growth cone is indeed an important way that axons extend towards a

target, but it is by no means the only method of elongation in the axon. During

development, growth of the organism continues after synapses are formed. To maintain

the neuronal network, the axon must continue to grow in response to the continued

tension placed on them. This process has been exploited in a number of experimental

settings to elongate axons to great distances.[13, 238-240]

Axons can be elongated by micropipette towing of the terminus.[241-243] This

pioneering method of axon stretching has been used to investigate mechanical properties

of axons. Alternatively, the Smith lab has been very active in extreme axon

elongation.[13, 238-240, 244-246] They have developed a device that can stretch millions

of axons at a time and accelerate axonal stretch rates up to 1 cm/d without breakage or

thinning. Embryonic rat DRG cells were seeded across two substrates to allow neural cell

bodies to adhere to each; the substrates are then separated in a step-rest pattern causing the axons between the two populations of cell bodies to be elongated. Stretched constructs have been shown to retain the same electrophysiological competence as control cultured neurons by displaying similar voltage channel density and patch clamping.[244] When stretch-generated constructs were implanted into rat sciatic nerve lesions, they showed promising integration into host tissue and axons within the tubular

58

graft displayed signs of host myelination.[247] The promise of tension-induced axonal

elongation for use in spinal injuries is enticing. Damaged tissue could be excised and

bridged by pre-grown neurons stretched to the correct length, cutting down on healing

time since large gaps would already be filled with existing axons. Iwata et al. teamed up

with Smith to create stretch grown constructs and to test their ability to repair a rat

hemisection SCI.[248] Embryonic DRG were elongated to 10 mm using tensile elongation, encapsulated in collagen and implanted 10 days after injury. Tissue bridging was observed after four weeks, but the functional benefits were not significant over collagen alone. Thus, proper interfacing and reconnection with host tissue is still a concern, especially in the spinal cord where there are complex organizations of axon tracts.

Physical cues can be used to directionally stimulate cells for guidance strategies.

Advancements in microfabrication techniques have allowed for new methods of surface

patterning to be generated with enhanced resolution. A few common surface

manufacturing approaches will be discussed briefly followed by specific patterns

attempted both in vitro and in vivo for neural guidance (Fig. 2.7). Although the specific cellular effects and changes in signaling due to topography have yet to be elucidated, some theories on cytoskeletal and functional mechanisms will be mentioned. Further review of topographical and surface guidance can be found in the following papers.[249-

251]

59

Figure 2.7. Illustration of topographies that can be used in cell guidance, on the nano or micro scale. Image reprinted from [294].

One of the most popular anisotropic patterns for cellular alignment is grooved or ridged substrates. These can be made in a large range of feature sizes by adjusting the height, width, and spacing of the grooves in the nano and micro scale. Micron scale

grooves have been used with different cell types to induce specific guidance, but with

mixed results. With neurons, smaller grooves (<10-20 µm) tend to cause a higher

occurrence of perpendicular alignment (across grooves), but also include parallel

alignment of neurites to grooves.[252-254] Recently, a phenomenon of cell bridging

across micron sized grooves was observed with DRG neurons, hippocampal neurons,

SCs, and neuroblastoma cells (B104).[255]. Each cell type was seen to extend processes 60

across adjacent plateaus (spaced 30-100 µm), especially with increasing cell number and plateau width (30-100 µm width). In addition to directional orientation, microgrooves have been shown to induce neuronal polarization (axon establishment). Embryonic hippocampal neurons on 1 to 2 µm wide grooves were shown to have significantly higher occurrence of polarization than on flat PDMS, especially when groove depth was increased from 400 µm to 800 µm.[252] From a TE standpoint, this work could be useful

in inducing polarization of differentiating stem cells for CNS work. It is also important to

note that the current trend is toward the development of degradable, as well as, surface

patterned materials. Traditionally, most topographical studies have been performed on

PDMS or other non-degradable materials due to their ease of fabrication with current

techniques. In order to transition to implantable materials that are biodegradable, some

researchers have begun utilizing PLA for grooved substrates.[254, 256]

Another common physical pattern, especially for neural TE, is channels. Large

tubular NGCs (nerve guidance conduits) and smaller channels imitate white matter tracts

found in the CNS and are therefore often used to lead spinal cord axons parallel to the

spinal column during regeneration. NGCs are the current preferred bioengineering

strategy for regenerating the PNS. Yu and Shoichet synthesized longitudinally oriented

multichannel NGC that increased DRG adhesion and outgrowth especially in peptide

modified channels.[257] However NGC applications can be utilized in the CNS, as

mentioned previously in the materials section, where agarose channels were implanted

into a rat spinal cord.[258] Also, NGCs incorporating additional components such as

neurotrophins or supporting cells were able to generate enhanced axon growth,

61 suggesting that NGCs could be useful for SCIs.[259, 260] Recently, one study synthesized semiconductor nanotubes (1 to 10 µm in diameter) out of silicon and germanium and formed them into arrays that encourages single axon penetration and outgrowth into the nanostructure.[261] This model electrically and physically mimicked the native myelin and therefore could be utilized for future neural network applications.

The ECM-like morphology of fibers with small geometries has been used to stimulate and guide cells. At a cell and structural level, the ECM ranges in size from 50 to 500 nm.[262, 263] Nanofibrous scaffolds have been created using three main techniques: self-assembly, phase separation, and electrospinning (see review [264]).

Aligned fibers have been used in neural cell culture to induce neurite sprouting and guidance.[265, 266] Fiber diameter is also an important factor to consider in optimizing neural outgrowth and guidance.[267, 268] Although not a true fiber scaffold, the nano surface topography of nerve basal lamina was recently replicated using a PDMS stamp on a polystyrene substrate.[269] The resulting topography resembled nanofibrous laminin structures, and supported the growth and alignment of DRG neurons. This study illustrates that cell reaction to fibrous scaffolds is very much tied to the surface structure and the feature size must be small enough for cells to sense and interact with it.

As a substitute to using actual cell topographies in co-culture, biomimetic surfaces aimed at reproducing these structural cues have been fabricated. Microcontact printing has been used to align SCs, which in turn were used for neural guidance.[270, 271] In a creative study by Kofron et al., PDMS was used to make both aligned cell topography

(taken from live cultures), and mimetic cell topography from a computer assisted design

62

(CAD) program.[272] The CAD generated topographies were tested against live cultures

of astrocytes, endothelial cells, and SCs for DRG neurite alignment and length and

displayed comparable or improved results in the CAD topographies as compared to the

live topographies. Kofron and Hoffman-Kim also optimized and quantitatively analyzed

cellular monolayers of astrocytes, endothelial cells and SCs using Design of Experiment

(DOS) and Response Surface Methodology (RSM).[273] The statistical experimental

design of this study afforded insights into the micropatterned feature sizes that affect

cellular adhesion, alignment and confluence. These powerful tools, along with other

modeling programs, take into account the multiple and complex biological variables and

are meant to alleviate time and resources spent on bench top experiments.

Chemical Cues

Several biochemical guidance signals have been identified including:

chemoattractive factors such as neurotrophins and netrins, chemorepulsive agents like

semaphorins and slits, or contact-mediated molecules such as ephrins and those located in the ECM (Fig. 2.8). Deciphering biomolecular guidance activity during nervous system development as well as injury is key to generating new techniques and tactics for improving and restoring function to the nervous system after injury. For this reason, general immobilization techniques and specific PNS and CNS chemical guidance strategies will be preceded by an overview of promising molecules commonly used in these studies.

63

Figure 2.8. The extracellular matrix (ECM) alongside a combination of contact-mediated and soluble factors guide the axons of maturing neurons to their innervating target during development. While the attractive cues pull on axons, the repulsive cues push axons to the path towards their proper targets. Images reprinted from [6].

Proteins and polysaccharides primarily make up the complex structural

framework of the ECM, which is secreted and organized into specific tissues by cellular

architects. In turn, the assembled ECM plays a vital role in regulating cell behavior

throughout development and adulthood, providing anchorage, mechanical buffering,

segregating different tissues and aiding cell-cell communication. The ECM is paramount to progenitor cell migration and differentiation, as well as to axonal genesis and extension.

Throughout the body, laminins serve as a key component of the ECM, providing binding sites for self-polymerization, other ECM macromolecules and cells. They are key contact mediated cell adhesion promoters for neural cells. To date, fifteen laminins have been characterized and all share a similar trimer structure that is generated from five α, four β and three γ chains.[274] Within the nervous system, laminins make up basement

64

membranes that are essential to interactions between neurons and glia. Research has

demonstrated that laminin is required for neuronal migration in the developing

cerebellum.[275, 276] Laminin contains several binding motifs that interact predominantly with cell-surface integrins (primarily α1β1 and α6β1) and secondarily with

α-distroglycan.[277, 278] Interactions between integrin receptors to their ligands contained in ECM proteins are central for anchoring and directing the growth cone in order to initiate guidance.[279, 280] More detailed information regarding integrin interaction and signaling is reviewed elsewhere.[281-283] Jacques et al. have showed that neural precursor migration on laminin can be significantly halted by inhibition of

α6β1 integrin subunits.[284] Developmental studies have observed that knocking-out β1 integrin expression blocks basement membrane formation and the expression of laminin protein.[285] Laminin provides essential attachment points enabling axons to extend and exert forces on the ECM.[286-288] Collagens and fibronectin are also important in the nervous system and are generally supportive to neuronal cell migration and neurite extension.[289, 290] Fibronectin, laminins, and collagens activate similar cell integrin receptors, thus all three are important in cell substrate adhesion.

Proteoglycans are important ECM molecules that have been implicated in neuronal and axonal guidance; these highly negatively charged molecules consist of a core protein with numerous GAG side chains and are grouped into two major classes:

HSPGs (heparan sulfate) and CSPGs (chondroitin sulfate). Several studies have demonstrated that either exogenous addition of HSPG or enzymatic removal of HSPG leads to axonal guidance defects during development.[291-293] This has led to findings

65

that demonstrate a functional association between HSPGs and several secreted and

transmembrane proteins. As a result, the role of HSPGs in guidance appears to be tied to

sequestering slit, netrin and semaphorin proteins (thoroughly reviewed in [294]). The

influence of CSPG on guidance is not as well understood as that of HSPG; however, it is

clear that CSPG also has a potent inhibitory effect on neuron guidance. Studies have

demonstrated that CSPG makes up part of the glial scar that effectively halts CNS axonal

regeneration, and leads to the generation of abnormal axonal growth cones.[295] Work

has shown that, similar to HSPGs, CSPGs also functionally associate with semaphorins

and inhibit axonal extension.[296] Li et al. utilized microfluidic techniques to create

parallel and opposing gradients of laminin and CSPG.[297] They found that cultured

DRG neurons show preference for higher laminin and lower CSPG in opposing gradients

by exhibiting strong axon directionality.

Tenascins are ECM glycoproteins primarily involved in development as well as wound healing. The tenascin (TN) subtypes, TN-C and TN-R, are two of the four TN family members found in the nervous system and are generally repulsive to axons.

However, TN-C and TN-R contain multiple domains that can be either repulsive or attractive depending on presentation, and may provide more precise control of neurite extension, migration and guidance.[298] The inhibitory effects of TN-R on neurite

outgrowth can be overcome by laminin and fibronectin used in combination, as shown in

a RGC outgrowth assay.[299]

For brevity, this review has focused only on a few important neurotrophins in cell

control and guidance since these have the highest inclusion in TE scaffolds to date. A

66

better understanding of these molecules (and many others not included here) and their

interactions will help in the future formulation of approaches for enhanced control of

axonal guidance, branching, pruning and synapse formation for the purposes of

regenerative medicine and TE. The following discussion is by no means exhaustive,

especially in regard to developmental findings and signaling pathways. For more

complete reviews please see.[15, 300-302]

The neurotrophin family is composed of secreted chemoattractant proteins that are integral to nervous system development and maintenance.[303] NGF, NT-3, BDNF, and

neurotrophin-4/5 (NT-4/5) are the four major neurotrophins and are similar in structure

and sequence.[304] Tropomyosin receptor kinases (Trks) and p75 neurotrophin receptors

(p75NTRs), located primarily on the terminal end of axons, are vital for the initiation of neurotrophin signaling.[303, 305] Three Trk receptors have been identified to date. NGF

binds specifically to TrkA,[306-308] whereas, BDNF and NT-4/5 prefer TrkB receptors.[309, 310] NT-3 complexes with TrkC with high affinity but can also interact

with the other Trk receptors.[311, 312] P75NTR shows high affinity for NGF and serves

as a low-affinity receptor for all the neurotrophins.[313]

The first neurotrophin to be discovered was NGF, which was shown to encourage

neurite extension from sensory ganglia in vitro.[305] Subsequent work found that NGF

exists at significant concentrations in adult tissue showing neuronal specificity, however,

NGF has been shown to also elicit responses from other tissues and cell types.[305] The

existence of other neurotrophic factors was postulated early on, since neurons depend on

specific targeting for proper innervation during development. BDNF was the next

67

neurotrophin identified from mammalian brain isolates and has been shown to

significantly act on CNS and directly associated neuronal populations.[314, 315] BDNF has been shown to play an important role in the regulation of synapse structure and function, especially in glutamatergic synapses.[316] NT-3 shares significant homology to

NGF and BDNF, however, NT-3’s primary role occurs during the development of many tissues as indicated by the broad distribution of its messenger RNA.[317-319] Both

BDNF and NT-3 have been shown to significantly influence axon path-finding, as well as aiding axonal regeneration in rats following SCI.[320, 321] Moreover, during development both neurotrophins direct the path-finding of maturing axons to their targets by a combination of long range attractive and repulsive cues.[322] NT-4/5, also known as

NT-4 and NT-5, influences the survival and outgrowth of sensory and sympathetic neurons.[309, 323, 324] The importance of these neurotrophins in the nervous system have been reviewed in [325]. New experimental evidence has determined that bFGF and

GDNF may also be important for nerve regeneration in the PNS and CNS. Both growth factors have been shown to influence neurons, SCs, and oligodendrocytes towards axonal growth and remyelination following injury.[326, 327] Finally, CNTF has been shown to act primarily on neurons as a survival factor following injury in the PNS.[328] To date,

CNTF has not demonstrated any functional or regenerative benefits for nerve repair;[329] however, it may show synergistic effects in the presence of other neurotrophins.[330]

68

Figure 2.9. Common methods of protein surface attachment. Physisorption is the physical nonspecific adsorption of proteins to the surface. Chemisorption utilizes covalent bonding between protein functional groups and a modified substrate. This figure depicts thiol- maleimide interaction. Bioaffinity depicts the strong noncovalent affinity steptavidin (tetrameric protein) has with biotin (small molecule on the protein). This type of immobilization shows specific linkage of a protein to modified surface. Figured produced by Aleesha M. McCormick.

Protein patterning has gained significant attention with advances in techniques such as lithography, microfluidics, microcontact printing, and biosensors. Protein patterning has enabled high-throughput measurements of biological responses using micro and nano scale protein assays. There are three primary methods used to immobilize protein: physisorption, chemisorption and bioaffinity (Fig. 2.9). Physisorption is the physical adsorption of proteins to the surface via intermolecular forces such as hydrophobic and polar interactions. This type of immobilization is nonspecific, such that proteins are adsorbed heterogeneously across the surface and are oriented to minimize repulsive forces with other molecules and the substrate. Adsorption is cheaper and easier than chemical means; however; the binding affinities are low and proteins can desorb easily, quickly leaving the original surface exposed. Chemisorption, on the other hand, is

69

a type of immobilization that results in covalent bonding between exposed side-chain

functional groups and modified surfaces. N-Hydroxysuccinimide (NHS)-amine, carboxyl,

thiol-maleimide, epoxy, and photoactive chemistry are all common strategies used for

covalent attachment. Often with these techniques, nonspecific chemisorption takes place

as exterior residues containing the reactive groups on the protein attach to the surface.

Nonspecific attachment could block active regions of the protein of interest or inhibit

important conformational changes. Therefore, site-specific protein immobilization is

desired to reduce unwanted attachments. This requires the insertion of functional moieties

into proteins specifically for immobilization to surfaces with the corresponding coupling

molecule. The biotin-avidin system is a well-known biochemical immobilization strategy

15 - largely because it exhibits one of the strongest non-covalent bonds known (Kd=10 M

1).[331] Streptavidin is a tetrameric protein that has a comparable affinity to biotin because it is structurally similar to avidin. Alternatively, histidine/nickel interaction, that allow tagged histidine regions on proteins to bind to nickel chelated complexes such as

Ni-nitiloacetic acid (NTA), and DNA-directed systems, such as DNA microarray technology and DNA-protein bioconjugation, are two other types of bioaffinity immobilization approaches commonly used for protein immobilization.[332-334] These methods of immobilization are attractive due to the specificity and homogeneity of the oriented molecules. For more information on immobilization strategies, please see.[335,

336]

Recent work has begun to reveal how tethering or immobilization of growth factors and guidance molecules modifies cell and stem cell function.[337-346]

70

Immobilization of bioactive factors to biomaterial substrates allows for not only

migrational stimulation, but spatial control of differentiation with sustained dosing, which is not possible with soluble factors.[338, 342] At the same time, cytokine immobilization

allows for the study of the dynamics and the necessity of cellular internalization for

activation of signal transduction.[345] Research has recently shown that IFN-γ as well as

PDGF-AA can be immobilized to hydrogel scaffolds to spatially guide the differentiation of NSCs.[337, 343, 346] Immobilization offers a number of advantages over soluble dosing. Immobilized molecules do not diffuse away from a scaffold like they would in soluble form and require smaller amounts of bioactive molecules over the course of treatment maintaining local concentrations, potentially maximizing cell interactions while reducing cost. Immobilization also allows for the creation of permanent gradients and this strategy has been utilized for small adhesion peptides [347-350] and more recently for growth factors.[351, 352] Consequently, it has become a preferred strategy employed to modify cell guidance and migration.[347-350] In achieving these effects, the presentation of proteins in gradients and/or 3D patterns can be highly beneficial since it more closely mimics how cells experience many proteins in vivo. Control of spatial distributions of proteins is key to achieving greater control of cell and tissue functions.

Attachment peptides derived from ECM proteins have been immobilized to culture substrates for neural guidance to mimic ligand presentation during development and injury. A RGDC peptide (found in laminin, collagen and fibronectin) and axonin-1, a cell adhesion protein, were patterned to a culture surface using photolithography techniques; this allowed in vitro neurite extension and network formation.[353] Similar

71

studies by Luo et al. demonstrated that 2D photo-immobilized GRGDS enhanced local

neurite density and elongation, as well as DRG extension, into immobilized GRGDS in

3D.[354] Photolithography paired with the development of multiphoton scanning

microscopes,[355-357] allows for even tighter control of immobilized factor

concentration and spatial location, potentially providing axon guidance with submicron

precision. Recent work from Yu et al. has utilized two-photon confocal patterning to

tether NGF to chitosan surfaces.[351, 352] They demonstrated that immobilized NGF does encourage DRG axon extension in vitro and that axons can be guided by gradients of immobilized NGF.

Recombinant fusion proteins of growth factors and binding domains have been created previously to control immobilization to specific ECMs, biomaterials and cells.

Fibronectin cell-binding domains have been incorporated in fusion proteins along with bFGF and EGF to stimulate vascularization and wound healing.[358] Collagen binding

domains have also been incorporated into fusion proteins of bFGF,[359] EGF,[360, 361]

PDGF,[362] hepatocyte growth factor (HGF),[363] and NGF for targeted wound

regeneration.[102, 364] Recently Sun et al. compared collagen binding NGF (CBD-NGF)

to native NGF (NAT-NGF) in a crushed rat sciatic nerve model.[102] They found that

CBD-NGF injection treatment enhanced remyelination of axons after crush injury. CBD-

NGF resulted in better myelinated axons when compared to NAT-NGF and PBS sham

treatments at 8 and 12 wks. Dodla and Ballamkonda created nerve guidance scaffolds

containing gradients of immobilized laminin and NGF in agarose hydrogels and showed

that axons were able to bridge a 20 mm nerve gap after sciatic nerve injury using

72

scaffolds with a continuous gradient of laminin and a step gradient of NGF in the same

direction.[98]

Electrical Stimulation

Natural electrical activity of the nervous system has led to investigation on the

effects of electrical stimulation, especially on neurons and subcellular behavior. The

application of electric stimulants to neural cells is not a new idea by any means, and has

been studied for over 30 years.[365] Despite its long history, mixed effects of applied electric fields still leave questions about their control over cell behavior. Neural stimulation in vitro could be used to promote desired cell behaviors such as alignment and outgrowth of processes. In vivo, electrical activity could be promoted by incorporating a conductive polymer, such as PPy or poly(aniline).

Electric fields have been used to stimulate neurite outgrowth and have been found to align cells, giving them polarity. The presence of a direct current (DC) electric field causes growing axons in vitro to align, extend, and accelerate toward the cathode and to

increase branching.[366-370] DRG in particular have been used extensively in studies of

applied electric fields to study neurite sprouting, length, and alignment.[371-373]

Neurites from DRG were observed to align in the direction of the field and extend

significantly farther than control treatments, but these results were governed by surface

properties.[373] Laminin coated surfaces induced the significant neurite lengthening,

while collagen surfaces did not. In addition, substrate surfaces have shown variable

results in neurite alignment and directionality in applied electric fields.[251, 374]

73

Electrical stimulation has been investigated as a differentiation cue in mammalian neural stem cells. Ariza et al. recently found that DC electric fields of around 500 mV/mm applied across NSCs from the DG (dentate gyrus) induced significantly higher neuronal differentiation than alternating current (AC) electric fields or no electric field application.[375] In the experiments, DC voltage was applied in vitro for the first three days and the last day of culture but AC was applied the full 6 days. DC voltage did appear to cause a lower cell density and some cell death. Results suggest that DC electric fields might be selective for the neuronal phenotype and cause alignment of neurons from

NSCs of the DG.[375] NSC migration in the presence of electric fields is of interest in inducing or guiding endogenous stem cells to injury sites. Adult rat NSCs did have directional migration toward the cathode over cells on which no electric field was applied

[376].

Animal studies have shown that brief electric field stimulation (1 h/d) works as well or better than long stimulation (4 h/d to continuous).[377, 378] Electric field stimulation has been shown to greatly enhance nerve regeneration in animal models, resulting in significant decreases in regeneration time. One hour a day of electrical stimulation has reduced healing time from 9 wk to only 3 wk in rat models.[377]

Electric fields have shown considerable promise for enhanced neuronal development and regeneration; however, further work is needed to understand the optimal way to utilize electric fields for TE approaches, especially for methodologies involving stem cells.

74

Concluding Remarks

Tissue engineering is poised to generate new techniques to replace or restore tissue damaged from injury or disease. The concept itself is not new, as it was contemplated by several visionary scientists speculating on where basic biological knowledge gained decades ago would take future researchers.[379] Although significant progress has been made toward neural TE, especially in the PNS, there is still a long road ahead to formulate clinically relevant CNS solutions that achieve significant long-term functional benefits. Major issues in the area remain unresolved including global injury and disease models and functional assessment, complete eradication unwanted of immune response, an absolute cell source, and concrete mechanisms and techniques for physical, chemical, and electrical cues.

The continuing advancement of new technology and techniques coupled with the uncovering of basic knowledge brightens the CNS TE outlook and the creation of restorative therapies for devastating injuries such as SCI or TBI. Design of biomaterials from the ground up, including proper cellular and tissue functionalities, will allow for the creation of ideal brain and spinal cord regenerative constructs. In any TE strategy it is important to incorporate cues (chemical, physical, electrical, etc.) derived from native cellular microenvironment (Fig. 2.1) to instruct cells and tissues to predictably regenerate. The incorporation of multiple cues experienced during development and regeneration into CNS treatments has propelled neural regeneration and TE into new exciting frontiers. Neural TE researchers are progressing towards the hallmark milestones

75 of enabling the restoration of speech after a TBI and empowering a paralyzed SCI patient to walk again.

76

CHAPTER 3

MOLECULAR LEVEL INVESTIGATION OF SPINAL CORD INJURY MODEL

Summary

Syringomyelia is a condition of the spinal cord in which a syrinx, or fluid filled cavity, forms from trauma, malformation or general disorder. Previous work has shown that in noncanalicular syringomyelia irregular flow and pressure conditions enhance the volumetric growth of syrinxes. A better understanding of the underlying molecular pathways associated with syrinx formation will unveil targets for treatments and possibly prevention of syringomyelia in the future. In this study, we performed an established surgical induction of a syrinx using quisqualic acid and kaolin injections in rats to characterize the injury at the molecular level by RNA sequencing and metabolomics techniques at three and six weeks post injury. Syrinxes nearly averaging nearly 10 mm in length formed in the rats’ spinal cords; however, smaller syrinxes were also detected in saline injected surgical shams, complicating interpretation of results. Our current results indicate a robust immune response coupled with overall decreases in neuronal signal transmission of syrinx containing animals compared to controls. Although transcriptional changes indicated gliosis and loss of neurons, no neuropathic pain was detected by von

Frey allodynia testing. Unique transporters were revealed to be highly dysregulated,

77

including significant increases in betaine/glycine transporter (BGT-1), K+/Cl-

cotransporter (KCC4), and aquaporin 1 (AQP1), along with the upregulation of small

molecule osmolytes taurine and betaine. The identified metabolites are of particular interest due to their involvement in osmotic homeostasis and need to be investigated further for their specific involvement in trauma induced syrinxes.

Introduction

Syringomyelia results from the formation of a fluid filled syrinx in the spinal cord, and can be associated with several central nervous system (CNS) disorders including Chiari Type-I malformation, hydrocephalus, trauma, cancer, or spinal cord infection.[380-383] The occurrence of syringomyelia changes across different ethnicities, and its detection rate is increasing due to higher prevalence of magnetic resonance imaging, improvements in diagnostic imaging, or both.[384] Syrinxes are dynamic and

sometimes asymptomatic, but large syrinxes often cause significant damage to the spinal

cord, are often difficult to treat and are debilitating to patients who experience pain,

numbness, stiffness, or weakening of the limbs.[385] Reviews on syringomyelia,

symptoms, and treatment options offer more in depth discussion.[380] Currently,

syringomyelia is mainly addressed by treating the associated condition to halt formation

or progression of a spinal cord cyst.[382] Treating Chiari or arachnoiditis often involves

decompression of the skull or spine with or without expansive duraplasty, essentially

creating space in which the CSF can properly flow. Idiopathic syringomyelia is most

commonly addressed by treating symptoms or, in the worst cases, by shunting the syrinx.

78

Leading researchers in the field have recently highlighted the need for basic research and

mechanistic insights into syringomyelia to improve both identification and

treatments.[386]

A clear grasp of the mechanisms responsible for syrinx initiation remains

particularly elusive, even as the knowledge of hydrodynamic factors responsible for

enlarging syrinxes has grown. Excitotoxic injury is part of the initiation process in post-

traumatic syringomyelia and involves energy depletion and subsequent destruction of

neurons.[387] Studies have shown the importance of subarachnoid blockage in significantly expanding syrinx size and increasing damage to the syrinx parenchyma as compared to syrinx size without arachnoiditis.[388-390] Preventing the formation of a syrinx within the spinal cord is paramount; however, many cases are asymptomatic with reversible pathology until the syrinx is of considerable size, so understanding how to halt syrinx enlargement is also important. Additional information on the cells and molecules that induce tissue dysfunction is needed before new syrinx treatments are formulated.

Irregular flow and pressure conditions drive the growth of the syrinx; these hydrodynamic forces have been shown to increase flow in the perivascular spaces that moves toward the central canal in healthy animals, whereas in surgically induced syringomyelia the fluid moves toward expanding syrinxes as well as the central canal.[390-393] Tracing fluid flow has also shown that arterial pulsation exacerbates syrinx growth in the spinal cord,[394] again highlighting the importance of hydrodynamic homeostasis in the CNS and how slight perturbations can have catastrophic results. The specific enzymes, ion channels, or neurotransmitters that initiate

79

fluid accumulation within the spinal cord and their cell localization are largely unknown

to date within the spinal cord. Previous reports have largely focused on the water channel

aquaporin 4 (AQP4) and syrinx formation.[395, 396]

In this study, the primary aim was to utilize a surgically induced syringomyelia rat

model to reveal the molecular events in the spinal cord during syrinx formation and

expansion. The goal was to determine several molecular markers specific to the injury

environment that may serve as molecular candidates for future treatments. Based on the

importance of hydrodynamic equilibrium in the brain and spinal cord, we hypothesized

that one or two particular fluid transporters, including AQP4, would show highly

irregular expression in syrinx containing animals. Additionally, we hypothesized that

several small molecule metabolites would be increased in syrinx containing animals over

healthy animals that could be used to fingerprint the disease state. The overall

experimental design and associated analyses are presented in Figure 3.1. Specifically, we

used a previously reported surgical procedure for excitotoxic induction of a syrinx in the

spinal cord that incorporates an intraparenchymal injection of quisqualic acid and a

subarachnoid injection of kaolin.[389-392, 397] The excitotoxic quisqualic acid begins killing local neurons to initiate a cavity in the spinal cord itself while the kaolin induces neighboring arachnoiditis to allow the irregular flow of CSF expand the syrinx size.[389]

In this study, at week three and six, animals were designated for fixation/histology or molecular assays. We chose to focus on revealing new information at the molecular level by isolating RNA and small molecule metabolites to create a multidimensional model of gene and metabolite expression for therapeutic target identification. RNA sequencing

80 data provided information about transcript levels that complimented metabolite data from tandem mass spectrometry analysis. The transcript and metabolite data was related using protein-small molecule interactions including enzyme reactions and transporter reactions, as well as any regulatory information available from the literature to provide a physiologic characterization of syringomyelia in the employed model.

Figure 3.1. Three experimental groups were analyzed: healthy control (CTL) animals; animals receiving sham surgical treatment (SHAM) where both injections were saline; and the excitotoxic injury group (EXC) that included animals receiving surgical treatment and injections of kaolin and quisqualic acid (QA). Some of the samples were chemically fixed for histology and imaging, while in the opposing set small molecules were isolated from tissue for transcriptomics or metabolomics. Datasets were integrated and interpreted for biological results.

81

Materials and Methods

Surgical protocol

All animal manipulations were done in accordance with the regulations of the

University of Akron Institutional Animal Care and Use Committee (IACUC). The groups included 1. healthy animals with no surgical treatment (control group), 2. animals receiving full surgical treatment with saline injections (1% Evans blue (Sigma-Aldrich,

St. Louis, MO, USA)in saline) into spinal cord and 3. subarachnoid space (sham group), and animals that received full surgical treatment with an intraspinal injection of quisqualic acid (QA; Enzo Life Sciences, Farmingdale, NY, USA) /1% Evans blue and a subarachnoid injection of kaolin (Avantor, Center Valley, PA, USA)) (excitotoxic injury group) beginning with 10 wk male Wistar rats. Animal numbers included: 8 healthy controls for molecular analyses; 12 sham animals, 8 for molecular analyses and 4 for histology; 32 excitotoxic injury animals, 16 for molecular analyses and 16 for histology.

All surgical procedures were performed under aseptic conditions. After induction of anesthesia, animals were maintained on isoflurane for the duration of the procedure. The surgery included a midline incision through the skin along the direction of the spine followed by exposure of the spinal column by blunt dissection of the muscles. A C7-T1 laminectomy was performed and a bent, sharp 30 gauge needle (Hamilton Syringes,

Franklin, MA, USA) was used to inject 5 µl of 250 mg/ml kaolin and 1% Evans blue into the subarachnoid space followed by a 2 µl injection of 24 mg/ml QA and 1% Evans blue

(in saline) using a blunt 30 gauge needle that was advanced 0.5 mm into the spinal cord.

82

Specifically, the injection was made in right dorsal quadrant of the cord, between spinal

nerves C7 and C8. The muscle was sutured together with 4-0 Vicryl and the midline incision was closed with Michel clips. For the sham group, the same surgical procedure was performed; however, saline plus 1% Evans blue was injected into the right dorsal quadrant of the cord and in the subarachnoid space. One animal had to be euthanized prior to the established endpoint due to extreme weight loss and three were lost during surgery, which is believed to be extreme sensitivity of the animals to anesthetics.

Animal perfusion and tissue harvest

At week three or six all animals were perfused with 3.7% formaldehyde (Sigma-

Aldrich) fixative (histology samples) or phosphate buffered saline (PBS) (for metabolite and RNAseq samples). PBS perfusion was used to displace blood before tissue dissection. Rats were administered an overdose of ketamine/acepromazine/xylazine intramuscularly; once the animals were sufficiently anesthetized and the heart had slowed, the ribs were opened and the heart exposed. An 18 gauge needle was inserted through the left ventricle into the ascending aorta. The needle was kept visible inside the ascending aorta and was clamped in the ascending aorta to prevent slipping. The right atrium was incised to allow fluid to flow out. A peristaltic pump was used at 15 ml/min to perfuse the animals with first heparin (Sigma-Aldrich), which was transferred to either fixative or PBS for perfusion. The cords were immediately isolated from the animal for post-processing and analysis. Fixed cords were left overnight in fresh 3.7% formaldehyde before rinsing in PBS. The unfixed animals provided samples for 1. RNA sequencing that

83 were stored in RNAlater (Life Technologies, Carlsbad, CA, USA) according to manufacturer’s instructions; and 2. the metabolite tissue samples that were immediately frozen at -80oC until use.

Histological Staining

Formaldehyde fixed and dissected cervical and thoracic spinal columns were sectioned at 40 μm using a microtome (Leica SM2000R, Leica Microsystems, Buffalo

Grove, IL, USA) with frozen stage. Sections were stained for astrocyte marker glial fibrillary associated protein (GFAP, Millipore, Billerica, MA, USA); oligodendrocyte marker RIP (Developmental Studies Hybridoma Bank, Iowa City, IA, USA); neuronal cytoskeletal protein βIII tubulin (TUJ1, Covance, Princeton, NJ, USA); macrophage marker ED1 (Millipore); cytoskeletal marker found predominantly in ependymal cells

βIV tubulin (Sigma-Aldrich); macrophage maker for M1 phenotype CD86 (AbD Serotec,

Raleigh, NC, USA); macrophage marker for M2 phenotype CD163 (Bio-Rad, Hercules,

CA, USA). Slides were blocked and permeabilized in normal serum (goat or donkey serum, Life Technologies) in 0.1% Triton-X100 for 1 h before incubation with the primary antibody overnight. Washes in phosphate or tris buffered saline preceded incubation with the secondary antibody overnight (either goat-anti-mouse-IgG-

Alexafluor546 or donkey-anti-rabbit-IgG-AlexaFluor488, Life Technologies). Long washes in PBS or TBS were followed by mounting in Prolong Gold (Life Technologies).

The channels BGT-1 (SLC6a12, betaine/glycine transporter, Millipore), KCC4 (K+/Cl- cotransporter, Bioss, Woburn, MA, USA), and Aquaporin 1 (AQP1, Millipore) were co-

84

stained to determine localization; this consisted of the procedure listed above for mouse

primary antibody and goat-anti-mouse-IgG followed by the repeat of the protocol with

rabbit primary antibody and donkey-anti-rabbit-IgG. Hoechst 33442 (Life Technologies) was used last to counterstain the nuclei. Sections were imaged with an inverted epifluorescence microscope (IX81, Olympus, Center Valley, PA, USA) and an Olympus confocal microscope (Fluoview FV1000). Images were processed for publication using

Adobe Photoshop (Adobe, San Jose, CA, USA) for color correction and proper contrast.

Syrinx dimensions in week six animals were calculated from sections; syrinx diameter was measured in MetaMorph (Molecular Devices, Sunnyvale, CA, USA) using the caliper tool across the largest distance on the cross sections. Length was found by adding consecutive sections containing a syrinx and multiplying by section thickness of 40 μm.

RNA sequencing and transcript processing

RNA was isolated from the tissue using TRIzol (Ambion, Foster City, CA, USA), following the manufacturer’s protocol. Briefly, tissue samples were homogenized by mortar and pestle in TRIzol then left at RT to incubate for 5 min. Chloroform (molecular grade, Sigma-Aldrich) was used to extract the RNA and spun down at 12,000 × g for 15 minutes at 4°C. The top aqueous phase was precipitated with isopropanol (molecular grade, Sigma-Aldrich) for 10 min and centrifuged at 12,000 × g for 10 min at 4°C.

Ethanol (molecular grade, Sigma-Aldrich) at 75% was used to wash the RNA twice, centrifuging at 7500 × g for 5 minutes at 4°C in between. Pellets were air dried ~20 min and resuspended in Tris-EDTA buffer (Fluka, Buchs, Switzerland). RNeasy MiniElute

85

Cleanup Kit (Qiagen, Germantown, MD, USA) was used on samples if quality or concentration was too low. The RNA samples were sent to Case Western Reserve

Genomics Core for library preparation (TruSeq stranded total RNA library prep kit,

Illumina, San Diego, CA, USA) and sequencing using Illumina HiSeq 2500 (2x 100bp run, 500-60M read output per flow cell, pooled samples). Sequence quality control was performed using FastQC and low quality neucliotides at the end of reads were removed using TrimGalore! (http://www.bioinformatics.babraham.ac.uk/projects/trim_galore/).

Tophat was used to align sequences to the rat reference genome rn6;[398] Gene expression per transcript (fragments per kilobase of transcript per million mapped reads,

FPKM) was obtained using Cufflinks and differentially expressed genes (rat gene annotation rn6) were identified with the CuffDiff method from Cufflinks package as described in Trapnell et al.[399] using a p < 0.01 to select statistically significant transcripts.[399, 400]

Metabolite processing and identification

Metabolites were isolated from spinal cord tissue by using a modified Bligh and

Dyer Extraction method to obtain both hydrophilic metabolites and lipids.[401] Tissue was lysed by the addition of 180 µl HPLC grade water and 20 µl methanol then subjected to a three cycles of freezing in liquid nitrogen, followed by thawing, and sonication. 750

µl 1:2 (v: v) CHCl3: MeOH and 125 µL CHCl3 were added to each sample. The samples were vortexed and an additional 250 µl of water was added. After incubation at -20°C for one h, samples were then centrifuged at 1000 × g for 10 min at 4°C. Hydrophilic

86

metabolites were collected from the aqueous phase and lipid from the bottom, organic

phase. All the extracted samples were dried in a CentriVap Concentrator (LABCONCO,

Kansas, MO, USA) at room temperature, and then preserved at -80°C until resuspension

and analysis. Hydrophilic interaction liquid chromatography was performed with a Luna

3μ NH2 100Å, 150mm×1.0mm, column(Phenomenex, Torrance, CA, USA) on an

Micro200LC system (Eksigent, Redwood, CA, USA). Polar metabolites were re- suspended in 200 μl of 35:65 (v:v) acetonitrile: water solution and 5 μl of each sample was analyzed. Mobile phase A of water and a mobile phase B of acetonitrile, each with the addition of 5 mM ammonium acetate and 5 mM ammonium hydroxide were used for the chromatography. The flow rate was 30 μl/ min. The gradient consisted of the following linear changes in mobile phase B over time: 0 min 98%, 0.5 min 98%, 1 min

95%, 5 min 80%, 6 min 46%, 13 min 14.7%, 17 min 0%, 17.1 min 100%, 23 min 100%.

Samples were analyzed on a 5600+ TripleToF Mass Spectrometer (AB SCIEX,

Framingham, MA, USA) in positive mode and were processed with Information

Dependent Acquisition (IDA) under the following conditions: the ion source nebulizer gas (GS1),15 psi, heater gas (GS2) was 20 psi, and the curtain gas (CUR) was 25 psi. A survey scan was performed over the mass range of 60-1,000 Da with a 250 ms accumulation time. The ionspray voltage was set to +5000 V ionspray voltage with a

+100 V declustering potential. Fragmentation data was subsequently collected using a collision energy spread of + (25-40) V. The HILIC-MS data were processed using

MarkerView (version 1.2.1.1). Isotopic ion peaks were excluded from monoisotopic ion peaks before analysis. Student’s T-tests (unequal sizes, equal variance) or Mann Whitney

87

U-tests were used to determine features that were significantly changed (p < 0.05, and fold change > 2) between experimental and control groups. Principal component analysis

(PCA) was performed with the Pareto Scaling method to compare groups. Putative metabolite/pathway identification was done using Metabolizer (ChemAxon, Budapest,

Hungary) and association with transcripts (especially enzymes and transporters) was performed with the KEGG database (Kyoto Encyclopedia of Genes and Genomes) and

SMPDB (Small Molecule Pathway Database) pathway information. [402, 403]

Compounds were initially confirmed through matching accurate mass and fragmentation

to the Metlin database and HMDB (Human Metabolome Database) and authentic

standards.[404, 405]

Network generation and mapping

Using transcript expression measures obtained using Cufflinks, FPKM values

were input into the Ingenuity Pathway Analysis software (IPA, Qiagen) in order to

calculate transcript expression fold changes of surgically manipulated animals as

compared to healthy animals. Significant fold change values were analyzed for gene set

enrichment in canonical pathways and upstream regulator analysis with no fold change

cut off implemented. Pathways and networks from IPA and from KEGG database were

further edited in Cytoscape,[406] where transcript and metabolite fold change values

were mapped to node color. All graphical displays and tables show only molecules with a

significant p-value and absolute fold change above 2 unless otherwise specified.

88

Results

Excitotoxic injury has a unique molecular signature associated with tissue pathology

Histological and molecular analyses revealed striking differences between healthy

animals and injured groups, including both the SHAM (saline injections) and excitotoxic

injury (EXC; QA and kaolin injections) surgical groups (Fig. 3.2). Although increased

GFAP expression is seen in affected areas of the dorsal region of the spinal cord, at week

three it is difficult to determine any cavitation in the tissue histologically (Fig. 3.2A).

Syrinxes at week six are large (0.56 ± 0.052 mm) and span a noteworthy distance of 8.8 ±

2.4 mm (excitotoxic group); this time point was determined to be the peak syrinx volume

by Tu et al.[407] Both the EXC and SHAM groups showed increased staining of GFAP

at week six, and transcript data reflects this increase with GFAP expression of over 2-fold in both EXC (log2(FC) = 1.1) and SHAM (log2(FC) = 1.0) groups (as compared to

healthy control) by week six (p<0.01). Somewhat surprisingly at week six we observed

syrinxes in the EXC as well as smaller but distinct syrinxes in the SHAM group; this was

not anticipated based on previously reported findings.[390] Unfortunately, there were not

enough samples designated for histology in the SHAM group to confirm that 100% of

animals would develop syringomyelia and compare syrinx size to EXC groups

statistically.

89

Figure 3.2. Analytical overview of data collected. (A) Histology images GFAP; week three and six EXC and SHAM groups where syrinx is denoted by star (scale bar represents 500 μm). (B) Table shows differential analyses of the different groups at week three and six. The numbers represent significant transcript/metabolites as determined by t-test comparisons that also have fold change greater than 2 or less than 0.5. Significance for metabolites p<0.05, significance for transcripts p<0.01; n=4 for CTL, n=4 for SHAM,

90

n=8 for EXC. (C) The categorical breakdown of perturbed transcripts and metabolites are shown at week three and six.

Differential comparisons of the groups in the molecular analyses revealed

thousands of significantly different transcript expression levels (p<0.01), and hundreds of

significant metabolite concentrations between the EXC and CTL groups (summarized in

Fig. 3.2B). Suppementary Tables 3.1-4 can be found in Appendix B and include the first

100 dysregulated transcripts or metabolites (tables truncated for inclusion). Between the

SHAM and EXC groups there was still a large, significant difference in transcript expression and metabolism (Supp. Tables 3.5-6). We believe this demonstrates the importance of the mechanical factors in the spinal cord that lead to injury and development of a syrinx as well as the chemical upheaval from necrotic cell debris and secondary injury cascade. Although the sham comparison to the excitotoxic injury group is important and intriguing, in this study we will focus mainly on the comparison of the

EXC group to the CTL group in a subsequent, more detailed analyses. In the future we hope to delve deeper into the differences between the SHAM and CTL, and SHAM and

EXC groups.

From Ingenuity Pathway Analysis (IPA) ranking of top biological functions (Fig.

3.2C), the transcriptional data revealed the main disturbances at week three are in cell death/survival and neurological function (deficits). At week six, perturbed pathways were involved in cellular movement, cell-to-cell signaling, and vascular related functions.

When the differentially expressed transcripts are compared categorically the fraction of significant changes increases from week three to week six in the immune and inflammatory response as well as cell movement. Conversely, the percentages of

91 transcript expression changes in cellular activities such as protein synthesis and modification, small molecule metabolism, and molecular transport decreases from week three to six.

Metabolyzer analysis of the metabolite profiling data allowed identification of significant peaks using the accurate mass, and the function of these molecules was inferred from KEGG database pathway involvement. KEGG categorization showed similar changes in pathways involved in signaling, amino acid metabolism, and co-factor metabolism (Fig. 3.2C).

Inflammation and immune response over six weeks

A major subset of differential transcript expression in the EXC groups suggest mobilization of phagocytes and granulocytes at week three, with recruitment fading by week six (Fig. 3.3). Even though data shows immune recruitment is tapering off, response to the injury is still observable at week six in the high levels of GFAP protein staining that are maintained by reactive astrocytes (Fig. 3.2A) and a large portion of disrupted transcripts that are involved in inflammation response and cell mediated immunity (Fig. 3.2C). The high levels of reactive microglia/ infiltrating macrophages, as shown by transcript levels and histological staining in Figure 3.3B, are expected, especially when considering the large syrinxes observed in the spinal cords. It is unclear whether the ED1+, CD163+, or CD86+ cells originated within or outside of the CNS from the likely compromised spinal cord-blood barrier.

92

Macrophage polarization was also analyzed using transcriptome data (Fig. 3.3B), but is difficult to interpret, as transcripts from EXC at week three have a larger magnitude of differences from CTL levels than at week six, in general. Important cellular markers of each M1, M2a, M2b, and M2c polarization phase were selected based on commonality in the literature and detection in our dataset (macrophage polarization reviewed in [408-

410]). When each time point is compared separately, markers of both M1 (pro- inflammatory) and M2 (anti-inflammatory) are upregulated suggesting a mix of macrophage types. A very low expression level of arginase 1 at week three, that then recovers at week six suggests a transition from a population with more M1 phenotypes to a population with more M2 phenotypes by week six in the EXC group. Tissue sections from the injury site revealed increased expression of ED1 and GFAP around the lesion formed in the dorsal spine (Fig. 3.3B), but mixed staining from the M1 marker CD86 and the M2 marker CD163. The increase in CD86 or CD163 was not as pronounced as ED1 staining in the cord, and actual protein expression may not reflect transcript level increases detected in RNA sequencing.

93

Figure 3.3. Immune response for EXC versus CTL groups. (A) Network displaying perturbation in transcript levels involved in immune cell movement. Week three data is represented by the inner node color and 6 week data is displayed as the outer node color. Gray nodes are not significant as determined by p-value (p < 0.01; n=4 for CTL, n=8 for EXC), and white nodes have a log2(fold change) value between -1 and 1 (and thus may not be significant biologically). (B) Macrophage polarization/ subtype dominance at week three and six from transcript data. The colored nodes represent the log2(fold change) value of the transcript levels in EXC as compared to CTL levels. p<0.01; n=4 for CTL, n=8 for EXC. Week six immunostaining in EXC animals for ED1, M1 and M2 macrophages is on the right side, confirming transcript data. The antibody used to identify M1 polarization is CD86, while CD163 shows M2 macrophages. Scale bar represents 100 μm.

94

Neuronal-related loss and functional testing

Transcriptomics revealed a major down regulation of transcripts related to

neuronal functions in the syrinx containing animals (Fig. 3.4). Due to the high cellular

density of CNS tissue, spinal cord cavitation likely leads to severe axonal losses, with

accompanying death of neuronal and glial cell bodies. RNA processing allowed the

compensation of lost cells by normalizing the sample to sample total RNA variation, but

reactive gliosis and invading immune cells undoubtedly skewed syrinx containing

samples to have a decreased fraction of neurons compared to healthy tissue.[389, 390]

From the data we attempted to interpret neuronal loss from decreased/increased

expression of the remaining neurons. A large subset of down-regulated mRNA was detected in transcripts involved in neurotransmission, as well as sensation and release of synaptic vesicles (Fig. 3.4A). Axonal guidance signaling was one of the most significant and highly dysregulated canonical pathways in the week six dataset. Detection of increased transcript levels involved in neurite growth and branching, displayed in the network of Figure 3.4A, is exciting considering the loss of neuronal cells. The activation of neurite growth pathways at three weeks is not sustained, as by six weeks these transcripts are upregulated, but with smaller fold changes.

Animals were tested for mechanical allodynia using von Frey filaments (Fig.

3.4B); however, no increased sensitivity was seen when testing the left and right hind paws (p > 0.05). Putative metabolites involved or associated with neuropathic pain show mainly decreased levels at week three as compared to healthy animals (Fig. 3.4C).

Probing the RNA sequencing data showed that there are conflicting transcript levels in

95 the neuropathic pain pathway, which may partially explain why no outward sign of sensitivity was noted in the animals (Fig. 3.4D).

96

Figure 3.4. CNS parenchymal response. (A) Categorical network showing transcript log2(fold change) values indicating neuronal damage and their relationship to specific functions including synaptic vesicle release, neurite outgrowth and branching, neural signal transmission, and sensation (includes pain), p<0.01; n=4 for CTL, n=8 for EXC. Gray indicates the change was not statistically significant while white denotes values between -1 and 1. (B) Animal response to allodynia testing revealed no significant changes in any of the groups across time. The average response of each animal was rated 0 to 4 for each von Frey Hair used to test the left and right hindpaw. Data depicted is the response to a 60 g von Frey Hair where a rating of 0 is no reaction, 1 is paw lift, 2 is quick withdraw, 3 is vocalization, and 4 represents licking of the paw, p<0.05. (C) Table depicting putative metabolites involved in neuropathic pain signaling. All are statistically significant but the fold change may not be biologically significant, p<0.05; n=4 for CTL, n=8 for EXC. (D) Canonical pathway of neuropathic pain in the dorsal horn, p<0.01; n=4 for CTL, n=8 for EXC. Nodes represent log2(fold change) values of EXC/CTL (center=3 w/ border=6 w) where gray indicates the change was not statistically significant while white denotes values between -1 and 1.

Fluid and solute transport perturbation

The expansion of the syrinx has been attributed to irregular CSF flow and pressure conditions in the spinal cord.[390-392, 397] Figure 3.5 shows key transcripts

and metabolites involved in transport and osmotic control in the CNS. Severely reduced expression of ion channels KCC2, and Na+/K+ pump in the EXC group compared to healthy CTL group was indicated by the transcript data and likely due to the loss of neurons/axons at week three and six of syrinx development. A sharp increase in AQP1 and KCC4 transcript levels was detected at week three and six when comparing EXC group to both the CTL group and SHAM group (Fig. 3.5C). No significant change was detected in AQP4 channel expression between any groups; APQ4 is known to be preferentially located near the astrocytic end-feet on the blood-brain barrier.[411]

Similarly, metabolites involved in maintaining osmotic balance in the CNS are shown in

Figure 3.5B. Metabolite IDs were confirmed by comparing fragmentation data obtained

97

from MS2 experiments to authentic standards (Supp. Fig. 3.1). The representative stained

sections shown in Figure 3.5D demonstrate the localization of BGT-1, KCC4, and AQP1 in the injured spinal cord. Perhaps one of the most interesting increases is the rise in

BGT-1 mRNA levels at week six (Fig. 3.5C). Studies indicate low levels of the transporter BGT-1 in the CNS and that these channels localize in the leptomeninges.[412,

413] We observed BGT-1 in the peripheral areas of the spinal cord cross sections. Due to the large increase in KCC4 transcript levels, co-staining was performed to confirm a dominant cell type expressing this ion channel. Figure 3.5D displays representative images with dual staining of KCC4 with TUJ1 and minimal GFAP co-localization, and was largely absent when dual stained with RIP, an alternative oligodendrocyte marker.

These images are taken mainly from the white matter tracts in the spinal cord, towards the dorsal side of the cross-section, where the most staining was seen. Increased intensity and frequency of KCC4 staining was observed rostral to the tissue surrounding the syrinx. AQP1 transcripts are highly abundant at week three and six; within the tissue increased intensity of the water channel was seen in the dorsal region of cord cross sections, and in what resemble sulci traveling from the edge of the spinal cord inward.

98

Figure 3.5. Syrinx propagation following EXC injury. (A) Network of transcripts showing progression of water and ion transporters responsible for osmotic control where the node center displays week three log2(fold change)data and the border week six log2(fold change) data. (EXC vs CTL shown here, EXC vs SHAM is shown in Supp. Fig. 3.2). Gray color indicates no statistical significance (p<0.01). (B) Metabolites important for osmotic protection (e.g., taurine and betaine) that are present in the tissue at high concentrations. Most are shown to be upregulated after injury and could be there to provide a protective effect. (C) Specific channels are highlighted that interact with increased metabolites (BGT-1) or were highly dysregulated (KCC4 and AQP1). On the left are box-and-whisker plots of the transcript levels of the channels while (D) the right side shows localization in the tissue at week six of EXC group tissue samples. Arrowheads indicate regions of dual staining, as seen by yellow overlap of fluorescent signals. Scale bar represents 50 μm. Statistical significance delineated by: * p<0.05; ** p<0.01, *** p<0.001; n=4 for CTL, n=8 for EXC.

99

Discussion

The syringomyelia model we chose to employ is an excitotoxic injury model

following on the work by Dr. Marcus Stoodley’s group who extensively studied the

progression of syringomyelia from this surgical procedure.[389-392, 395-397, 407, 414,

415] Typically an injury is created in the cervical or thoracic vertebral regions of the

spinal cord via injection of an excitotoxic agent (QA) that initiates the death of neurons

leading to syrinx formation. The severity is enhanced by following the initial injury with

a localized blockage in the subarachnoid space (similar to arachnoiditis) that disrupts

normal CSF flow (Fig. 3.1). Increased intraspinal injury has been shown to be associated

with SCI models that simulate arachnoiditis, classically consisting of an intrathecal injection of kaolin.[388] Past work in the field has characterized some of the effects of the surgically induced syrinx, including syrinx size progression, increased GFAP and

ED1 presence near the lesion, and cerebrospinal fluid (CSF) flow into the perivascular space around syrinxes.[389, 392, 397] The surgical method was shown in several studies

to reliably produce large syrinxes in the majority of, if not all, experimental animals when

the two injections were combined.[389, 390] Before beginning this study, we postulated that syrinxes from the excitotoxic injury (EXC) group would mimic and give the best initial insight into syrinxes formed as a result of trauma to the spinal cord. In addition, a surgical sham group (SHAM) receiving saline injections was included which could allow us to narrow down the molecular changes that are more strongly associated with the excitotoxic α-amino-3-hydroxy-5-methyl-4-isoxazolepropionic acid receptor (AMPAR)

100

agonist QA and the kaolin blockage in the subarachnoid space. It is known from other

surgical models of syringomyelia that a syrinx, albeit a smaller one, will form without the

excitatory amino acid-mimic, such as clip compression and arachnoid kaolin

injections.[416-419] Even so it was unexpected to see distinct cavitation in our SHAM

group as this is not reported as often. Because we did not have a high enough sample

number for histology to determine incidence of SHAM animal syrinx development, it is

unclear whether some of the trends we have uncovered are related specifically to syringomyelia or are a result of spinal cord trauma (via the fluid injection) in general.

Future studies will elucidate these associations.

Previous transcriptome wide studies on general SCI have revealed some master

regulator genes for neuronal growth following injury, especially by comparing

regeneration that does occur in the peripheral system to the stagnant behavior in the adult

CNS (review in [420, 421]). Recent work has identified gene networks occurring in

peripheral nerve injuries and used the preserved regulators from multiple injury types to

screen for drugs that alter gene expression profiles of DRG to improve neurite

outgrowth.[422] Exciting work such as this could potentially transition to improving CNS

outcomes by altering a host of transcription factors and major regulators, casting a wider

net as compared to many strategies that only target one or two genes at a time.

Investigations into spinal cord injuries have revealed coincident transcript pathways most

altered in the CNS and some master regulatory factors. A transcriptome wide weighted

gene network analysis was performed on a full transection SCI model by Duan et al.[423]

in which modules of genes were determined and monitored over time. Unlike our

101 findings, the rats that underwent a full transection retained upregulation of immune and inflammatory response over 90 days.[423] From the literature, the most conserved regulators reported are STAT3[422, 424] and ATF3[422, 425]. We saw significant increase in our excitotoxic and sham dataset compared to healthy controls for these transcripts, especially ATF3. However, because of the low abundance usually found in the CNS, this increase was likely due to invading leukocytes.

Inflammation in the vicinity of the injury was expected, especially as studies have shown the vasculature is compromised in syringomyelia, which leads to increased extravasation of immune cells.[426] Indeed inflammation was seen in both SHAM and

EXC groups in response to trauma caused by the surgical procedure. A breach in vasculature and increased extravasation of leukocytes can be seen in the increased mobilization of immune cells shown in IHC for ED1, CD86 and CD163 (Fig. 3.3B) and transcript levels in the network of Figure 3.3A. RNA levels are on average much higher at week three while ED1 staining was more intense and covered a larger area in the vicinity of the syrinx at week six. We believe the magnitude of leukocyte, specifically macrophage, recruitment and mobilization may be transcriptionally increased at week three, but the accumulation of immune cells over time leads to increased histological detection at week six. The large number of overexpressed migration markers for phagocytes, granulocytes, and lymphocytes suggest that there is a high possibility the vasculature was compromised or that a high degree of extravasation occurred near the injured area. Compromised vasculature in the presence of a syrinx may also contribute to

102

enhanced mobilization of immune cells, as tracing studies have revealed using

horseradish peroxidase to uncover leakage in the injured cord.[426]

Determination of the polarization of the accumulated macrophages and reactive

microglia surrounding the syrinx was unclear. Some studies suggest a spectrum of

macrophages rather than clear polarization of M1 vs M2.[427] Ebb in M1 markers was present in transcriptional data from week three to week six; however, distinct changes in

CD86 and CD163 at the protein level was not as apparent (Fig. 3.3B). A transition towards an M2, or regenerative state, in the injured tissue from week three to week six would agree with studies from the Stoodley lab that have investigated syrinxes over time, showing that the size does not progress after six weeks.[407]

Of note is the lack of detection of any neuropathic pain in the rats even though syrinxes were detected in the histology. Small decreases in creatine and choline (Fig.

3.4C) may be attributed to neuronal loss, but could also mean oligodendrocyte death as the two molecules are typically seen more in glial cells than neurons.[428, 429]

Protective effects of phosphocreatine and carnitine have been studied in hypoxia,[430] although how these protective effects extend to the week three and six time points is unclear. The convoluted milieu of conflicting transcriptional changes (Fig. 3.4D) could explain the lack of detectable allodynia or any kind of sensitivity in the animals.

Transmission and sensation was mildly interrupted, according to RNA sequencing (Fig.

3.4A). Decreased signaling may be due to neuronal death from the syrinx formation, and increased signaling along the parallel route within the pain pathway may show compensation by remaining neurons near the lesion. In humans, symptoms of

103

syringomyelia are widespread and inconsistent from patient to patient. Studies have

described dysesthesia as a common symptom, alongside hypoesthesia and hyperesthesia,

often occurring in the same patient on different regions of the body.[431] It is very

possible that we did not test the correct area for allodynia on the rats, or that they actually had desensitized paws due to the loss of neurons in the dorsal horn area of the spinal cord. Further, conflicting expression data for neurite growth and branching was revealed;

there were losses in neurite growth/branching, but the majority of expressional changes

are increased fold changes suggesting neuronal attempts to reconnect across the injury or

form new connections to recover loss in signal transmission. The increased fold change

of transcripts in growth/branching of neurons and even a few in neurotransmission and

sensation, shown as red nodes in Figure 3.4A, are highly biologically significant when

considering the loss of neurons and increased fractions of glia in RNA samples from

syrinx containing animals. In total, these data suggest the injured tissue is moving

towards regaining homeostasis and guiding neurites to reconnect lost neural circuits.

A case study from Lohle et al.[432] detected very high increases in the amount of

proteins in syrinx fluid (as compared to regular CSF) in an epedymoma associated case of

syringomyelia. This previous finding correlates well with the detected increases in our

study of metabolites and many mRNA transcripts in the EXC group compared to healthy

animals. In general, the greatest magnitude of increases in mRNA and metabolite

expression occurred at week three and became largely attenuated at week six. Further, in

the Lohle et al. study the authors postulated that the compromised vasculature likely

resulted in increased migration of serum proteins into the syrinx (82% of detected

104 proteins were serum proteins), leading to a further hypothesis that increased osmotic pressure in the syrinx from the high colloidal concentration caused lower absorption of syrinx fluid. This preferentially directed fluid travel implicates extravasation as the main contributor to syrinx growth in the spinal cord. Indeed, work from Brodbelt et al. [391,

397] has shown via horseradish peroxidase tracing that fluid flows preferentially into the syrinx and to the central canal. Their work revealed that most of the fluid travelled through the perivascular spaces although fluid travel did still occur in the parenchyma.

One of the most interesting changes that occurred in the EXC group compared to

CTL and SHAM animals was the sharp increase in important metabolites, specifically betaine, carnitine, and taurine and the expression of corresponding transporter channels

(Fig. 3.5). These osmolytes are vital to maintaining osmotic homeostasis to preserve proper cell volume. High concentrations could be due to solute release from necrotic cells or possibly released from surviving cells in an attempt to regulate the tissue and restore hydrodynamic balance; increases of the osmolytes at week three will have to be considered carefully as they can be part of inflammation due to the surgical procedure. It is not certain whether increases in osmolytes and channel overexpression of BGT-1,

SLC6a6, and SLC22a5 are contributing to increased cyst size, a direct result of syrinx formation, or a result of surgical trauma. The mRNA expression patterns suggest a correlation between syrinx size and the levels of betaine channel and carnitine channel

(BGT-1 and SLC22a5, respectively), because their expression levels do not change significantly until six weeks after the initial injury (Fig. 3.5A). More time points evaluating protein and metabolite levels against syrinx size would likely yield additional

105

information as to kinetics and whether these rises in betaine, carnitine and their respective

channels are, in fact, causative. The increase in BGT-1 expression was significant statistically but not “biologically significant” according to the applied cut-off for fold change values; however, we believe this change is indeed highly biologically relevant to syrinx development due to the extremely low expression levels of BGT-1 typically found

in the CNS and because it does not occur until 6 weeks after the surgery suggesting it is

not tied to the procedure itself. Conflicting evidence exists in the literature as to cellular

presentation of the betaine/GABA transporter BGT-1; however, more recent work using multiple antibodies for the channel only detected BGT-1 in the leptomeninges of the

brain.[413] Our findings see this continuing into the spinal cord, with almost all BGT-1

observed around the periphery of the spinal cord and some processes extending toward

the center (Fig. 3.5D); there appears to be partial colocalization with GFAP, but not with

βIII tubulin (TUJ1, neurons) or βIV tubulin (ependyma). In addition to solute

transporters, ion transport can critically impact the movement of fluid and osmotic

balance in the CNS. Study of localization of KCC4 in the CNS has shown that it is

present at higher levels in the spinal cord than brain, where it is mainly located along the

apical membrane of the choroid plexus epithelial cells, as well as in cranial nerves and

nuclei of the brainstem.[433] Karadshesh et al. also found that in the spinal cord KCC4

co-stained with only CNPase positive cells (oligodendrocytes) while in the peripheral

nerves it co-stained with MAP2 positive cells (neurons). The observed differences seen in

our syrinx containing sections versus healthy sections from Karadshesh et al. are most

likely a result of the tissue response to syrinx formation and attempt to osmoregulate in

106

the face of syrinx expansion. Alterations in the expression of water/solute/ion channels

(Fig. 3.5A), especially a large decrease in the critical ion transporter Na+/K+ pump,

could also lead to irregular water movement in the syrinx animals and may be an

important contributor to syrinx propagation throughout the six weeks. Na+/K+ pump

(ATP1) is an important ion channel and one of the main channels implicated in cell osmoregulation. In addition to solute movement, there can be secondary active transport of water molecules, even against osmotic gradients, by solute transporters such as KCC,

NKCC1, GABA, etc.[434, 435] where water is co-transported with the substrate, and not in a passive manner or in response to an osmotic gradient. The alternative culprits responsible for water movement are the aquaporins, water channels responsible for exchanging water in cells between different compartments such as the blood/brain or blood/spinal cord barriers. The choroid plexus is the main location for AQP1 in the CNS, where it aids water transport across the apical membrane.[435] Staining for AQP1 is

shown in Figure 3.5D, and located near the syrinx including positive staining examples

near capillaries running through the spinal cord. Compared to similar work from Hemley

et al.[395] studying AQP4 expression in syringomyelia, we see different spatial patterns

of AQP1 staining. Hemley et al. used the same excitotoxic injection surgery applied in

our study including subarachnoid blockage with kaolin with the only obvious difference

being rat strain of Sprague Dawley versus Wistar.[395] In their results, AQP4 was

predominantly found around the circumference of the cord, surrounding blood vessels

and around the central canal expressed in ependymal cells; in our sections, AQP1 was

found near the edge of the spinal cord especially near the dorsal horn with very little of

107 the channel detectable in the white matter. Figure 3.5D shows AQP1 in the dorsal region of the spinal cord, near to but not colocalizing with GFAP; in addition, any significant staining in the white matter of the spinal cord is in what appears to be sulci running from the perimeter of the cord in towards the center with some co-expression with βIV tubulin, indicating meningeal localization. What is interesting is that in our RNA sequencing dataset levels of AQP4 remained stable, while huge increases in AQP1 transcription were seen at both week three and six (Fig. 3.5 A,C).

One unique aspect of this work included identification of perturbed metabolites, and we were able to identify severely dysregulated osmolytes that are known to be important in osmoregulation. Betaine and taurine are two of the most highly recognized osmolytes in the CNS.[434] We are particularly interested in the possible application of these two metabolite markers as potential diagnostic aids that could predate syrinx detection by MRI. Betaine and taurine are specific, as their imbalance is not typically linked to general SCI; the magnitude of the change in concentration suggests that they could be easily detected after lumbar puncture with low probability of false positives.

Additionally, both taurine and betaine are highly upregulated at week three even though there is small histological evidence of syrinx formation in our sections and they stay elevated at week six as inflammation abates. The imbalance of taurine and betaine should be studied in future work to determine if these two molecules indeed precede any physical sign of syringomyelia. Additionally, it would be valuable to determine if betaine or taurine levels in CSF samples can correlate to injury progression in syrinx containing rats, and could thus be studied over the duration of syringomyelia. Insertion and sampling

108 via an intrathecal catheter has been successful in rats, and is translatable to patients who could receive an epidural catheter or have CSF sampling done during a scheduled CNS surgery.[436-440]

A difficulty of this study was evaluating all the statistically identified features from the metabolomics dataset, as some of these were not able to be confirmed on current databases. Unfortunately this problem is common to the field of metabolomics as there is currently no comparable and agreed upon open source repository for metabolites, like has existed for decades for genomic, RNA and protein data via the National Center for

Biotechnology Information. There is fear of missing pieces of the puzzle of syrinx enlargement due to still unidentified metabolites. Of the identified metabolites in this study, we believe several are very important not only to spinal cord injuries but to syringomyelia in particular. The integration of metabolite and transcript data was key in establishing importance and likely implications of the specific changes in the spinal cord metabolome discussed.

The data presented here represent an important first step towards understanding the molecular mechanisms involved specifically in syringomyelia. We are particularly interested in connection with the hydrodynamic aspects of the injury as these are largely responsible for syrinx enlargement. Several transport channels for fluids and solutes were implicated here by the metabolite and transcript data that should be further studied and monitored before and during syrinx formation to confirm their association with the syrinx due to the confusion of SHAM animals containing small cavities. We plan to delve into

109

the temporal profiles of BGT-1, KCC4, and AQP1 as well as betaine and taurine in future

work to possibly confirm causal relationships with the cyst formation and size.

Conclusions

The results of this study expand upon previously implicated water channels, namely aquaporins, in syrinx exacerbation and identify other key players that may be important in syringomyelia, e.g., ion channels KCC4, Na+/K+ pump and solute channel

BGT-1. Syrinxes were successfully initiated and studied at three and six weeks using surgical induction via QA and kaolin, but some cavitation was seen in saline injected

SHAM animals that complicates confirmation of our interpretations. Significantly dysregulated transcript levels showed a marked immune response and a decreased population of neurons leading to transmission loss. Of note, the broad view of sequenced transcript levels and identified metabolites allowed the pinpointing of several key species including the transporters BGT-1, KCC4, and AQP1 along with highly upregulated metabolites betaine and taurine. These solutes and channels are suspected to be important in the preferential flow of fluid into the syrinx from surrounding spaces and parenchyma because of their known involvement in osmotic balance and fluid movement.

Acknowledgments

We would like to thank Conquer Chiari for funding this work, as well as the

University of Akron and Choose Ohio First for partial funding of personnel involved in the studies. We would also like to acknowledge support from AB SCIEX Young

110

Investigator Award, to LPS. Assistance provided by Bethany Noble and Parag Joshi was greatly appreciated. Finally, special thanks to Andrew Brodbelt, Patricia Sloan, and Kim

Stakleff who were imperative to initial surgical protocol details.

Supplemental Information

Information supporting this study can be found in Appendix B.

111

CHAPTER 4

INVESTIGATE AND MANIPULATE NATIVE STEM CELLS OF THE CNS

Summary

New developments in growth factor and pharmaceutical control of cells for neural regenerative strategies have built a need for high throughput assays to conveniently screen these treatments in vitro before moving to more complex experiments. Towards this application, we have studied an easy and highly reproducible culture regime with a renewable cell source for central nervous system (CNS) strategies. Adult neural stem cells (NSCs) derived from the CNS are attractive for use in screening assay because they are easily expanded and can differentiate into all major CNS cell types. NSCs were cultured on glass alone or methacrylamide chitosan (MAC) hydrogel coated glass substrates, immobilized with extracellular matrix (ECM) proteins collagen and laminin at varying densities. Proteins were also adsorbed on the surfaces as a control. We found that adsorbed protein on hydrogel coated glass resulted in the highest cell densities after 8 d, over twice the density of immobilized groups or adsorbed protein on glass. No significant differences were observed between collagen, laminin, or both proteins together regarding

cell differentiation (p > 0.05); however, the morphological spreading and branching of

differentiated NSC processes was enhanced on MAC substrates with covalently

112

immobilized laminin protein. The results of this study suggest that a soft MAC hydrogel

surface with adsorbed protein would be most desirable for specifying neuronal

differentiation of large numbers of stem cells due to the high cellularities they supported.

Introduction

There is an urgent need for the development of regenerative strategies aimed at

central nervous system (CNS) recovery from injury or disease due to the poor innate

ability of the brain and spinal cord to heal unattended.[37, 441-443] In their attempt to

protect the CNS and prevent further damage, astrocytes wall off the site of injury and

prevent axonal reconnection. The lack of treatments offering complete functional

recovery results in consequences such as a massive drop in patient quality of life and

reduced life-span. As a first step toward the development of implantable tissue

engineering strategies, biomimetic assays are required to study cellular processes and

responses to varied physical, electrical, and chemical stimuli.[254, 374, 444-448] In

particular, the design of simple, successful, and repeatable assays is necessary to gather

meaningful data on protein and/or drug analyses for guiding desired cellular responses.

Cell-based assays call for a reliable cell source and a simple and highly reproducible cell

culture method. The neural regeneration community is specifically interested in how such

assays can be used to stimulate cellular processes such as axonal sparing, extension, and

neurogenesis.[446-451]

Adult neural stem cells (NSCs) from the subventricular zone present an appropriate cell source for neural regeneration that are capable of becoming all major cell 113

types of the CNS (neurons, astrocytes, and oligodendrocytes) and are easily expanded in

culture.[172, 452, 453] These cells are derived from the CNS itself, and naturally reside

in the walls of the lateral ventricles. Many recent strategies have proposed using brain

derived NSCs for tissue engineering constructs aimed at regenerating the CNS.[58, 109,

187, 191, 454, 455] Therefore, it is important to establish methods for controlling the

proliferation and differentiation of these cells so that they can be easily incorporated into

implantable scaffolds. At the current time homologous NSCs are an unlikely cell source

for regenerative medicine due to the invasive procedures required to isolate them from

the patient.[204] However, donor matched NSCs offer intriguing possibilities while

serving as a template for the “CNS restricted” multipotentiality that should be achieved

with other stem cell sources.

Within the body cells receive a multitude of chemical and physical cues from

their environment on a regular basis. In the CNS stimuli originate from surrounding cells

sending paracrine factors or electrical signals, extracellular matrix (ECM), and the flow

of cerebrospinal fluid around and through the brain and spinal cord.[456-459] It stands to

reason that in order to effectively and repeatedly control cell behavior in vitro, a multi-

cued scheme is a necessity. In this study we have combined surface chemistry, concentration, and stiffness to reveal the effects of the culture substrate on NSC differentiation. In order to better understand and control NSC behavior, we chose native adhesive proteins preferred by neural tissue, laminin and collagen I.[236, 460-462]

Common signaling proteins/growth factors used to promote neuronal differentiation of

NSCs include nerve growth factor, brain derived neurotrophic factor, angiopoietin I, and

114

the cytokine interferon-γ (IFN-γ) used by us and others to yield a high percentage of

neurons.[190, 192-194, 197, 337, 463] In a simple yet effective scheme using silanated

coverslips, we demonstrate in this study the covalent immobilization of proteins in

increasing concentration allowing us to reveal the most effective substrates that combine

with IFN-γ to increase neuronal differentiation and subdue glial cell lineage specification.

We hypothesized that a combinatorial approach using a low density of laminin on a soft

substrate would encourage stem cell differentiation into neuronal over glial lineages.

Materials and Methods

Substrate Preparation

Glass coverslips (12 mm dia., VWR, Bridgeport, NJ, USA) were first cleaned

repeatedly in deionized (DI) water and 1 M NaOH for 10 min and 30 min, respectively.

The coverslips were then functionalized with amine groups (3-

Aminopropyltriethoxysilane, Chem-Impex, Wood Dale, IL, USA) or methacryl groups

(3-(Trimethoxysilyl)propyl, Sigma-Aldrich, St Louis, MO, USA) for protein immobilization, as illustrated in Figure 4.1. Briefly, the coverslips were covered in ethanol and bubbled with N2. Each siloxane was reacted separately in ethanol at 10% v/v,

and allowed to react for 2 h with 1 min N2 bubbling every 30 min. The coverslips were rinsed in DI water and placed in 70% ethanol for 20 min to sterilize. The methacryl coverslips were further modified with a methacrylamide chitosan (MAC) hydrogel layer.

MAC was synthesized as reported previously,[92, 114, 463] by the reaction of

115 methacrylic anhydride with chitosan. MAC was dissolved at 2 wt% in DI water and sterilized by autoclave on liquids cycle. A photoinitiator (1-hydroxycyclohexyl phenyl ketone, Sigma-Aldrich), dissolved in 1-vinyl-2-pyrrolidinone (Sigma-Aldrich), was added at 0.6 mg/g of MAC and mixed thoroughly at 1500 rpm for 1 min (SpeedMixer

DAC 150 FVZ, Hauschild Engineering, Hamm, Germany). A thin layer of

MAC/photoinitiator was polymerized onto the methacryl coverslips by pipetting 40 µl of the polymer onto sterile Teflon and placing the coverslip overtop; 2 min of UV exposure

(365 nm) created gel coatings with a Young’s elastic modulus of 0.5-0.7 kPa as we have characterized previously.[92, 463] The gels were rinsed thoroughly in phosphate buffered saline (1xPBS).

Figure 4.1: Illustration of coverslip preparation showing silanation and immobilization of proteins including a surface of MAC hydrogel (A) or glass only (B). Notice that MAC coverslips had an extra preparation step where the hydrogel layer was polymerized onto the methacrylated slips (A).

Adhesive Protein Immobilization

116

Mouse laminin (Life Technologies, Carlsbad, CA, USA), rat tail collagen I

(isolated from rat tail tendons), used together and separately, were immobilized to the

amine coverslips or MAC coverslips using 1-ethyl-(3-dimethylaminopropyl)carbodiimide hydrochloride (EDC, Chem-Impex) and N-hydroxysulfosuccinimide sodium salt (sulfo-

NHS, Chem-Impex).[464] The proteins were first diluted to 1 ng/ml, 10 ng/ml, and 500 ng/ml, in 1xPBS. EDC was added to the protein solutions at 50 mM and mixed well, and then sulfo-NHS was added to the solutions at 5 mM. The protein solutions were immediately added to the surfaces and allowed to react for 1 h at RT. After 1 h, the surfaces were rinsed thoroughly in 1xPBS. In addition, separate coverslips were made using standard techniques for cell culture where each protein was adsorbed on either glass or MAC coverslips at 5 µg/ml for 2 h after the adsorption of poly-D-lysine (Sigma-

Aldrich) at 50 µg/ml overnight.

Surface Protein Verification

Coverslips from each group (12 in total) as well as plain MAC coated coverslips, non-reacted amine functionalized coverslips, and non-reacted methacryl functionalized coverslips were analyzed by Fourier Transform Infrared (FTIR) spectroscopy for validation of attachment and comparison of protein concentrations. All coverslips were lyophilized prior to FTIR readings to remove water. Absorbance scans were read by a

Nicolet-6700 (Thermo Scientific) with 10 co-added scans at resolution of 4 cm-1.

117

NSC Isolation and Expansion

All protocols involving the use of animals were performed with prior approval

from the University of Akron Institutional Animal Care and Use Committee (IACUC).

NSCs were harvested from the subventricular zone of 6-8 wk old female Wistar rats

purchased from Charles River Labs (Wilmington, MA, USA) as described

previously.[463] Briefly, one week after arrival rats were sacrificed using CO2 and the

lining of the lateral ventricles was excised and digested using a papain dissociation kit

(Worthington Biochemical Corporation, Lakewood, NJ). NSCs were expanded as

neurospheres in a suspension culture of growth medium containing neurobasal (NB, Life

Technologies, Grand Island, NY, USA), B27 supplement (Life Technologies), 100 µg/ml

penicillin-streptomycin (PS, Life Technologies), 2 mM L-glutamine (L-glut, Life

Technologies), 2 µg/ml heparin (Sigma-Aldrich), 20 ng/ml epidermal growth factor

(Sigma-Aldrich), and 20 ng/ml basic fibroblast growth factor (PeproTech, Rocky Hill,

NJ) and incubated at 37°C and 5% CO2. Cells were expanded by passaging weekly until

use.

NSC Differentiation

NSCs from passages 3-8 were plated on coverslips at an initial seeding density of

40,000 cells/cm2 in growth medium overnight. A full media change was made 18 h later and differentiation medium was used containing NB, B27, 100 µg/ml PS, 2mM L-glut,

and 150 ng/ml IFN-γ (PeproTech). NSCs were cultured in differentiation medium for an

118

additional 7 d with a half media change at 3 d. At 8 d, the cells were rinsed with 37 oC

1xPBS and fixed with 3.7% paraformaldehyde for 7 min.

Immunocytohistochemistry

To visualize cell type, the antibodies βIII tubulin (neuronal), synapsin I (mature neuronal), nestin (stem cell), and NG2 (oligodendroglial precursor) were used. Washing with 1xPBS was performed in-between each of the following steps. Cells were first permeabilized with 0.1% Triton-X (Sigma-Aldrich) in 1xPBS for 5 min, followed by 1 h

incubation of coverslips in 2% goat serum (Life Technologies) in 1xPBS. Next, the

primary antibody monoclonal mouse anti-β-III tubulin (1:500, Covance, Princeton, NJ,

USA) or mouse monoclonal anti-nestin (1:1000, BD Biosciences, San Jose, CA, USA) in

1xPBS plus 1.5% BSA was left on the slips for at least 1 h, followed by incubation of the

secondary antibody goat anti-mouse IgG AlexaFluor 546 (1:400, Life Technologies) in

1xPBS plus 1.5% BSA for over 1 h. Coverslips were then blocked for a second time with

2% donkey serum (Life Technologies) in 1xPBS for 1 h. βIII tubulin stained coverslips

were also incubated with rabbit monoclonal anti-synapsin I (1:500, Abcam, Cambridge,

MA, USA), while nestin coverslips were also incubated with rabbit polyclonal anti-NG2

(1:500, Abcam) for over 1 h. Donkey anti-rabbit IgG AlexaFluor 488 (1:400, Life

Technologies) was applied to all coverslips for at least 1 h. For visualization of nuclei, cells were treated with 1 µM Hoechst 33342 (Life Technologies) for 10 min. Finally,

119

coverslips were mounted on glass slides with Prolong Gold hard mount (Life

Technologies).

Cell Counting

NSCs were imaged at 6 fields of view (FOV) per coverslip (n=4 coverslips for each group) with a 20x objective on a fluorescent microscope (Olympus IX81, Tokyo,

Japan). Cell density was calculated by manually counting all nuclei and normalizing by the area. Positive staining for each antibody marker was counted in each of the overlay images of nuclei and cell marker and normalized by total cell number. Positive staining percentages were then averaged for each antibody over all 4 coverslips per group.

Statistical Analysis

Each cell marker data and cell density data were analyzed by a 3 factor ANOVA using SAS 9.2 statistical software (Cary, NC, USA). The factors and levels analyzed are listed in Table 1 and include: surface type (glass or MAC), protein (collagen, laminin or both), and concentration (adsorbed, high, medium, or low). Significant differences between interaction terms (cross of 2 or more factors) is denoted if present; if no interaction terms were significant, differences between factors were displayed. A level of

α = 0.05 was used to denote significance. Values are reported as mean ± SD, and if relevant, interaction means are used.

120

Results

Substrate Analysis

The IR absorption spectra of all the groups are shown in Figure 4.2. MAC groups

were all normalized by a plain MAC hydrogel coated coverslip, while the glass groups were normalized by an un-reacted amine functionalized coverslip. Differences were seen between the amine silanated glass and hydrogel coated glass because of the characteristic

bands in the range of 3600-2800 cm-1, 1700-1200 cm-1, and 1200-800 cm-1 from the

chitosan (base material of MAC).[465, 466] Amide bands can be seen at 1600–1670 cm-1,

1510–1560 cm-1, and 1220–1450 cm-1,[467] which is where we expect to see a change in

peak sizes since surface amines form an amide bond when a carboxylic acid on proteins

are covalently attached to the polymer via carbodiimide chemistry, as used in this study.

-1 In addition, the 3600-2800 cm band represents NH2 and NH3 vibrations.[465]

Cell Density of NSCs

Cell surface density showed little variation between the different proteins but was

very different among the increasing concentrations and surface type, as shown in Figure

4.3. Statistical analysis revealed significance (p < 0.0001) between the interaction of

surface typse (MAC or glass) and concentration (adsorbed, low, medium, or high). NSCs

grown on adsorbed proteins with a MAC coating maintained over double the number of

cells than all other surfaces (83,900 ± 27,800 cells/cm2). This was true regardless of the

protein adsorbed. In general, all soft MAC surfaces had visibly higher cell numbers than 121

corresponding glass surfaces; however, the difference was not significant due to the

interaction term between surface type and concentration. The lowest cell density of NSCs

occurred on the glass + medium concentration groups, with an average of only 9,010 ±

2 3,640 cells/cm .

Figure 4.2: FTIR absorbance scans of lyophilized surfaces were used to confirm immobilization of proteins. Adsorbed controls show different IR spectrums from immobilized groups as seen by comparing characteristic amide peaks at 1600-1670 cm-1, 1510-1560 cm-1, and 1220-1450 cm-1.[467] Characteristic bands can also be seen for chitosan in the range of 3600-2800 cm-1, 1700-1200 cm-1, and 1200-800 cm-1.[465, 466]

NSC Differentiation Cell Types

To determine cell fate of NSCs after 8 d of culture, the coverslips were stained for

the stem cell marker nestin, the glial progenitor marker NG2, or neuronal markers βIII

tubulin and synapsin I. The percentages of cell staining for each marker are displayed in

Figure 4.4. The factors of substrate type, protein type, and protein concentration were

examined and the interaction term between surface type and concentration was

significant for all markers except NG2 (p < 0.0001).

122

Figure 4.3: Cellular density of NSCs on treated coverslips after 7 d of culture with IFN-γ, with an initial seeding density of 40,000 cells/cm2. Results are separated by protein treatment: collagen alone (A), laminin alone (B) and collagen and laminin in combination (C). Significantly different groups are indicated by letter using multi-factor ANOVA, where “A” represents the highest mean (p<0.05). Mean ± SD, n=4.

The two groups with the greatest βIII tubulin and synapsin expression (Fig.

4.4A,B) were adsorbed protein on MAC (79 ± 6.0% βIII tubulin, 42 ± 15% synapsin), as well as low concentration of protein attached to glass coverslips (72 ± 8.0% βIII tubulin,

52 ± 11% synapsin); adsorbed glass had the lowest neuronal positive staining percentage with an average of 44 ± 21% βIII tubulin and 16 ± 8.0% synapsin. For nestin staining 123

(Fig. 4.4C), most groups exhibited similar expression, with the exception of proteins

adsorbed on glass, which had an average of 12 ± 5.0% nestin positive cells.

Contrastingly, for NG2 expression significant differences were seen between each

the substrate type factor (p < 0.0001) and protein concentration factor (p < 0.0001), but not between the interaction terms (Fig. 4.4D). There was no significant difference between the proteins themselves for NG2 expression either (p = 0.4267). Glass had higher positive staining for NG2 at 75 ± 9.0% whereas MAC surfaces only averaged 56 ±

12%; low protein concentration had the highest number of NG2 positive cells (74 ± 11%) followed by adsorbed (67 ± 14%), high (63 ± 15%), and medium concentrations (57 ±

12%), in descending order.

There were noticeable differences in cell morphologies between the groups.

Figure 4.5 shows representative images of each group cultured on laminin coverslips stained with βIII tubulin and synapsin. Morphologically the cells on adsorbed MAC coverslips had a higher occurrence of cell processes and branching than low concentration on glass. Although glass surfaces supported lower cell numbers, compared to MAC cultured NSCs on glass still presented similar neuronal staining but decreased neurite extension. To reveal the glial lineage specification of surface treatments, Figure

4.6 displays NSCs immunostained with nestin and NG2. NSCs cultured on MAC had more cell processes and arborization. Glass adsorbed protein also elicited some glial progenitor process extension (Fig. 4.6). All groups had relatively high positive staining of

NG2, as seen by the prominence of green in Figure 4.6; only the adsorbed protein groups suppressed high levels of nestin.

124

Figure 4.4: Cell fate of NSCs after 7 d in culture with IFN-γ on modified surfaces. Antibodies were used against the specific cell markers to determine cell type: βIII tubulin for neurons (A), synapsin I for mature neurons (B), nestin for neural stem cells (C), NG2 for glial precursor cells (D). Significance is denoted by letter using multi-factor ANOVA (p<0.05). Mean ± SD, n=4.

Discussion

Coupled compliance and chemistry

Our methods, outlined in Figure 4.1, resulted in successfully conjugated proteins

to modified coverslips as confirmed by IR spectroscopy. Figure 4.2 shows the subtraction

of base surfaces from post modified surface so that the peaks shown indicate the presence

of protein on all surfaces, with slight changes in peaks when comparing immobilized to

adsorbed groups. Differences in the FTIR absorption spectra can be seen in the 125

characteristic amide bands including 1600–1670 cm-1, 1510–1560 cm-1, and 1220–1450

cm-1. We also created different substrate types by gel coating glass coverslips with our

MAC hydrogel, which has a low elastic modulus that has been shown conducive to

neuronal differentiation.[92, 463] By combining substrate stiffness and surface chemistry

we successfully created varying cell culture environments to specify NSC behavior. This

multi-cued approach is meant to mimic the native environment while revealing the

factors, alone and in combination, that signal NSCs to become terminal cell types in vivo.

In this work we were able to use two known stimuli in conjunction with each other and

common techniques that can be applied in most laboratories. Our results should be

valuable to future studies including assays or tissue engineering constructs that require

control of NSC behavior.

Cell viability and proliferation

A trend in cell densities at the end of culture can be seen between the soft MAC surfaces and the hard glass (Fig. 4.3); however, the statistical significance can be seen between the interaction of surface type and protein concentration (p < 0.0001). We hypothesize that confounding factors such as possible protein desorption and/or length of culture could lead to these statistical results. Methacrylamide chitosan, our hydrogel material, has a slight positive charge from the amine side groups that could lead to a more sustained protein adsorption than negatively charge glass (even with adsorbed poly-d- lysine). A different length of culture in future studies may reveal a more pronounced effect of substrate stiffness and immobilized protein density. The visual difference in cell 126

number can be seen in the representative immunohistochemistry images (Figs. 4.5 and

4.6), especially the adsorbed protein on MAC group, where over twice as many nuclei

can be observed than other groups. The initial seeding density for all surfaces was 40,000

cells/cm2; thus MAC coverslips resulted in enhanced NSC viability and adhesion, and

proliferation in the case of the adsorbed MAC groups when compared to glass surfaces.

This has been shown in a number of studies investigating hard versus soft culture

substrates for NSCs.[92, 468] We and others believe that the preference for softer

substrates is because the native NSC environment (niche) is very soft compared to many

other tissues (elastic modulus 0.5-1kPa).

Because of the importance of laminin in the NSC niche,[456, 469] in these studies

we expected to see differences in proliferation in response to either collagen or laminin.

Recent work by Hall et al. resulted in human NSC growth with the addition of soluble laminin in culture;[470] however, we did not see any difference in total cell number

between collagen, laminin, or both proteins together in our work with rat NSCs (Fig. 4.3).

In a study of embryonic mouse NSCs, Flanagan et al. found that laminin coated surfaces

resulted in over twice the cell density than matrigel coated surfaces after 3 days in

culture.[471] Because matrigel is a combination of many ECM proteins and growth

factors, including both laminin and collagen, we would expect it to be somewhat similar

to our collagen and laminin group. Imaging of NSCs (Figs. 4.5 and 4.6) revealed visual

differences between cell shape on coverslips with laminin and those without; however,

we did not observe the sharp increase in cell number as Flanagan et al. This suggests to

127 us that laminin could play a role in the morphology of these cells but not enough to affect proliferation.

Figure 4.5: IHC images of NSCs exposed to laminin functionalized surfaces after 7 d in the presence of IFN-γ showing neuronal markers βIII tubulin (red) and synapsin (green). Cell nuclei are shown in blue. The differences between groups in terms of cell number 128 and cell spreading can be observed. In general, MAC surfaces had brighter neuronal staining and more cell processes (indicated by arrowheads). Scale bar is 100 µm in all images.

Figure 4.6: IHC images of NSCs exposed to laminin functionalized surfaces after 7 d in culture with IFN-γ showing neural stem cell marker nestin (red) and the glial marker NG2 (green). Interestingly, groups with a higher concentration of βIII tubulin also 129

generally had a high concentration of NG2. Arrowheads indicated stem cell or glial processes. Scale bar is 100 µm in all images.

NSC differentiation and morphology

Our results show that substrate type and immobilized protein density were coupled in their statistical significance in terms of NSC differentiation (Fig. 4.4). Despite similar immunostaining quantification results (Fig. 4.4), morphologically, NSCs grew and spread very differently on glass and MAC substrates. We are not surprised that the combination of stiffness and protein concentration significantly affects cell morphology, as this corroborates findings from Engler et al. where they found that gel stiffness and ligand density coupled together affected smooth muscle cell (SMC) shape and cytoskeletal expression.[472] Although NSCs and SMCs are vastly different cells, the coupling effect of mechanical and chemical cues is common among both, and could be due to focal adhesion and myosin interactions on different substrates.[473-475] Future studies will further probe the link between mechanical and chemical cell sensing by looking into receptors and transduction pathways in NSCs.

Protein type did not significantly affect neuronal differentiation in our studies

(Fig. 4.4 A,B); however, it has been seen to help in neuronal differentiation in embryonic derived NSCs.[476, 477] Mouse embryonic NSCs had over twice as many neuronal cells than fibronectin or collagen after 2 or 20 days in culture.[471, 476] There is not any information on whether or not NSCs lose their sensitivity to these ECM proteins over time, but possibly the integrin expression changes with development. Figures 4.5 and 4.6

display images from NSCs cultured on laminin and represent the most spread and

130 branched cells of the other two protein groups. As described above, laminin is known be a key component of the neural stem cell niche,[456, 469] and is generally preferred in neurite outgrowth and guidance studies for NSC or primary neurons.[98, 234, 257, 447,

461, 478, 479] In this study, differentiated NSCs developed longer and more branched processes on laminin, followed by collagen/laminin, and collagen coated surfaces.

Studies of human embryonic neural stem cells in vivo and in vitro have revealed that laminin does cause more cell migration, neurite extension, and neurite branching compared to other ECM proteins including fibronectin.[471, 480] When assayed for cell motility and neurite length, the NSCs behaved similarly on matrigel and laminin, both of which had more motility and longer neurites than fibronectin, suggesting that perhaps collagen does not have a pronounced effect on NSC morphology in the presence of laminin.[471]

With respect to low, medium, and high concentrations of immobilized protein, there is a trend for βIII tubulin, nestin, and NG2 expression (Fig. 4.4 A, C, and D). It is clear that low concentration has the higher expression levels of all proteins, and there appears to be a negative correlation between protein concentration and protein expression levels. Interestingly, the low concentration of protein has the highest expression after adsorbed groups for positive neuronal staining which could be due to the larger spacing in between protein molecules. Some NSCs did retain expression of the stem cell marker nestin at day 8 (Figs. 4.4 and 4.6). In contrast to our hypothesis, a large percentage of

NSCs expressed the glial precursor indicator NG2, even those groups with a high percentage of neuronal staining. We hypothesize that the cells were not fully committed

131 to the neuronal cell type and thus still exhibited some features of glial precursor cells.

This may also be the reason for the overall low staining percentages of synapsin, a more mature neuronal marker.

In response to covalently attached proteins or surfaces with adsorbed proteins, we saw that the adsorbed protein had far higher neuronal staining than any of the immobilized groups. We had hypothesized that, at the correct concentration, immobilized protein would exceed or match adsorbed protein in differentiation capacity. In literature, the most common attachment of laminin to NSC culture substrates are by physical adsorption, and we confirm that adsorbed laminin provokes good cell attachment and neuronal differentiation under the right media conditions.[471, 480] One possible reconciliation of our findings is that ECM molecules or fragments need to be taken up by

NSCs to have a more pronounced effect. Internalization could explain laminin caused cell proliferation in embryonic mouse NSCs in recent work by Hall et al. when they cultured the NSCs in suspension with soluble laminin.[470] Because of the ease and commonality of physically adsorbing laminin to culture substrates, in addition to it eliciting the most desired response from adult NSCs, we have chosen it as the best culture condition for future work.

Conclusions

In this study we report the successful immobilization of cell attachment proteins to hard and soft surfaces to control adult NSC differentiation as compared to the

132 commonly used adsorption method. The ECM molecules laminin, collagen I, and a combination of both proteins were conjugated and analyzed on either glass coverslips or hydrogel coated glass coverslips for their ability to maintain cell viability and specify neuronal differentiation. Methods used to manufacture and apply these substrates could be easily replicated in most laboratories, making them suitable for application to high- throughput assays for drug and growth factor screening.

The results of this work support that a MAC surface with adsorbed protein would be most desirable for specifying neuronal differentiation of large numbers of stem cells due to the high cellularities they supported. Additionally, MAC surfaces resulted in NSCs with more cell processes, arborization, and with decreased glial tendency, which was especially true on laminin surfaces. Results of this work will be valuable in future studies to determine favored culture conditions for NSC and other adult stem cell proliferation and differentiation into both neurons and glia. In fact, a follow up study was completed by our lab comparing DC electric field stimulation on NSCs. The manuscript, authored by Ms. Liza J. Kobelt, is included in Appendix B. The data show that electric field stimulation can vastly enhance neurite outgrowth. Generating significant numbers of these CNS phenotypes currently represents a major hurdle toward CNS tissue engineering. Combinatorial approaches using chemical, mechanical, and electrical stimulation seek to recapitulate native environments and overcome obstacles to regeneration.

133

Acknowledgments

We are grateful for funding from the University of Akron that supported this work. We would like to acknowledge Dr. Steven S. Chuang and his students Seyed Ali

Modjtahedi and Matthew T. Isenberg for assistance with FTIR.

134

CHAPTER 5

IN SITU GELLING CHITOSAN-PEG COPOLYMER EVALUATION FOR USE IN

THE SPINAL CORD

Introduction

Endogenous wound repair in any location is a complex process. Enhancing healing in the spinal cord is made more complex by the innate but elaborate neural circuitry and natural barriers to the healing process. Important events, key cellular players, and secondary injury cascade are reviewed in great detail in several review articles.[409, 481, 482] While scar tissue is never an ideal replacement for natural tissue, the signal processing and transferring functions of spinal cord make it even less appropriate because scar formation can block communication. Spinal cord injury, or damage by degenerative diseases, can lead to inflammation and much more detrimental secondary injury processes. Syringomyelia, or the formation of a fluid filled cavity in the spinal cord, can develop after injuries of this type.[381, 483] The syrinx offers the additional challenge of not being as easily accessible as a partial or complete transection;

however, if the cyst resolves the tissue can recover function.

To combat spinal cord cavitation, especially syringomyelia, therapies involving

pharmaceutical delivery are being developed. An ideal treatment would involve a

135 biomaterial vehicle delivering a drug or protein aimed at combating specific cell processes to alleviate a disease or treat symptoms. Injectable treatments for the spinal cord are reviewed in greater detail elsewhere.[484, 485] The ideal material delivery system should gel in the native environment and remain in place, induce minimal inflammation due to the gel or any byproducts, match the tissue mechanical properties to avoid stress to surrounding tissue, release its treatment in an appropriate time-frame, and resorb over time so that native tissue can replace the cavity. While this “wish list” has been difficult to attain, significant progress has been made towards finding materials compatible for the spinal cord. Hydrogels are a prime focus based on their high water content and generally low mechanical stiffness that is similar to soft tissues.

Polysaccharide based hydrogels in particular have become very popular CNS biomaterials because of their similarities to native brain or spinal cord extracellular matrix (ECM). There are many naturally sourced polymers capable of producing soft hydrogels being used in spinal cord engineering, with the most popular being made of alginate [486], chitosan [423, 487, 488], hyaluronic acid [489, 490], methylcellulose

[110, 491], or combinations [492, 493] of these.

Chitosan, a deacetylated form of chitin, is chemically similar to the native CNS

ECM component hyaluronic acid. The free amine group on chitosan makes it simple to modify with different functional moieties for cross-linking and specific immobilization.

A thiolated chitosan (thiomer) material has been described thoroughly by Andreas

Bernkop-Schnurch in several papers as used for a mucoadhesive and drug delivery agent,[494, 495] even being tested as a tissue scaffold.[496] Here we have combined

136

thiolated chitosan with a multi-arm polyethylene glycol (PEG) cross-linker terminated in

maleimides that allows for almost instantaneous gelation once the two components are

mixed. The thiol-maleimide click reaction provides a quick gelation at physiological

conditions sans the use of harmful initiators or chemical byproducts (reviewed in [497]).

Similar gels using this reaction have seen long lasting hydrogels and no cytotoxicity.[498,

499]

The goal of this work was to determine a suitable, injectable hydrogel material for the spinal cord that can cross-link in situ and does not cause unacceptable host responses

following mild injury. Towards this end, we first grafted a thiol functional group onto

chitosan to allow rapid gelation via the addition of a maleimide terminated multi-arm

PEG cross-linker. Several chitosan precursors were synthesized, gelled and tested for

swelling properties. The most appropriate thiolated chitosan for this application was fully

characterized in vitro. Finally, an in vivo safety assessment was performed (Figure 5.1) to

ensure no undue host reaction occurred when the gel was injected into the spinal cord.

The extent of immune infiltration and inflammatory response was characterized and

interpreted based on similar studies.

137

Figure 5.1: The reaction of chitosan with Traut’s reagent leaves the polysaccharide with a thiol functional group for the center gelation scheme. The thiolated chitosan and maleimide terminated PEG precursors are mixed immediately prior to injection, allowing them to gel in place in the spinal cord.

Materials and Methods

Materials and equipment used

Chemical components including buffer salts and reagents were purchased from

Sigma Aldrich (St. Louis, MO, USA) unless specified otherwise. Biological reagents

including basal media and supplements as well as antibodies were purchased from

Thermo Fisher Scientific (Waltham, MA, USA) unless stated otherwise. Animals used in

experiments were used in accordance with protocols approved by the University of Akron

IACUC.

Hydrogel precursors and gel formation

Three thiolated chitosan variations (CS-SH) were made for initial testing in vitro,

all were synthesized from PROTASAN UP B 80/20 (Novamatrix) with a reported

deacetylation of 80-89% and an average MW of 200 kDa. CS-T is chitosan reacted with

Traut’s reagent (2-iminothiolane, ChemImpex); CS-TGA5 is thiolgycolic acid (TGA) conjugated chitosan that was reacted at a pH of 5; lastly, CS-TGA6 describes chitosan reacted with TGA at pH 6.

CS-T reaction consisted of 0.25 g of chitosan in 175 mL of 1% acetic acid solution to which 0.1 g of Traut’s reagent was added. The pH was adjusted to 6 using 5

138

M NaOH and reaction left at RT for 24 h. CS-TGA was synthesized by dissolving 0.25 g chitosan in 2 mL of 1 M HCl and adding DI water to make a 1 wt% solution [496]. For 3 h chitosan is reacted with 0.25 g TGA and 0.6 g of 125 mM 1-ethyl-3-(3- dimethylaminopropyl)carbodiimide (EDC, ChemImpex, Wood Dale, IL, USA). The pH was adjusted to 5 or 6 (as specified in the name). All three CS-SH materials were dialyzed the same way: 2 changes of 5 mM HCl buffer, 2 changes of 5 mM HCl + 1%

NaCl, and 2 changes of 1 mM HCl buffer. They were each then frozen, lyophilized, and stored at -20°C until further use. For use the material was dissolved in PBS to 3 wt% with the help of tris-(carboxyethyl)phosphine hydrochloride (TCEP; 5mM for CS-T and 20 mM for CS-TGA). In the resulting solution pH was adjusted to ~6-6.5 by adding 0.1 mM

NaOH until the concentration reached 2.5 wt% and finally autoclaved for sterility.

The 4 arm polyethylene glycol terminated in maleimides (PEG-4mal, 20 kDa

MW, JenKem Technology, Plano, TX, USA) was diluted to 1.5 wt% in phosphate buffered saline (PBS) before use and sterilized by 0.2 μm filtration. The two hydrogel precursors were mixed vigorously using connected syringes and then injected into an appropriate vessel, and allowed to gel at 37°C for subsequent studies unless otherwise specified.

Thiol confirmation and content

Unmodified chitosan and CS-T were dissolved overnight in 1 mM HCl at 60°C at

2 wt%. Some of the material was made into films that were imaged with electron microscopy and EDS. Some of the dissolved chitosan and CS-T was diluted in 1X PBS

139

and 1 mM EDTA (pH adjusted to 7-8). These samples were tested for thiol content using

Ellman’s assay by mixing the reagent with the sample for 15 min then reading the

absorbance at 412 nm. A standard curve of Traut’s reagent was used to determine thiol

concentration in the samples. Unmodified chitosan was used as a blank.

Gelation and mechanical properties

An ARES II rheometer (TA Instruments, Newcastle, DE, USA) was used to collect data on 6 different samples cut to 8 mm in diameter using a metal punch. The hydrogel precursors (CS-SH and PEG-4mal) were mixed as previously described and rinsed in PBS to establish final mechanical properties once the gel has reached equilibrium. The cut out gels were measured on parallel plate geometries using a frequency sweep (strain controlled) and strain sweep to establish the linear range. The storage (G’), loss (G”), and complex (G*) moduli were recorded at 0.1-80% strain and 1-

100 rad/s.

Hydrogel swelling and degradation

Hydrogels were formed as described previously and their initial masses recorded.

Pre-weighed vessels were used to manipulate the gels during measurements and carefully

blotted to remove excess fluid. Gels were placed in ~20% excess of fluid. For post-

gelation swelling ratio measurements, gels were formed, transferred to PBS containing wells, and stored at 37°C and between measurements. They were weighed one day later

140

so that gels could reach equilibrium without any degradation (no appreciable degradation

observed in 1 w). Post-gelation swelling ratio was calculated as (swollen mass – initial

mass)/initial mass. The swelling ratio, Q, was calculated by taking the swollen gels and

freeze drying them to obtain hydrogel dry mass. Q was calculated as (swollen mass/dry mass). For degradation, gels were formed as described, freeze dried, and covered with either PBS or lysis buffer at 37ºC for 1 or 2 wks. Gels were then frozen at the appropriate time-point, dried, and massed to compare final dry weights to the initial dry weight.

In vitro toxicity

For cytotoxicity experiments, human dermal fibroblasts from neonatal foreskins

were seeded and given time to become confluent. The next day, the hydrogels were pre-

formed and transferred to transwell inserts (VWR, Radnor, PA, USA) and this was

recorded as day 0. The media was aspirated 24 h and PrestoBlue in PBS was incubated on the wells for 30 min. The solution was read for fluorescence intensity at 555 nm excitation and 590 nm emission according to manufacturer’s directions. Next, cells were washed with PBS and fixed using 4% formaldehyde. Cell nuclei were stained with

Hoechst and cytoskeleton with phalloidin conjugated to AlexaFluor488 dye. Wells were imaged on a fluorescent microscope. Cell numbers per image were counted using ImageJ and normalized by area of the image.

In vivo material injection

141

The hydrogel precursors were injected into rat spinal cords to assess gelation in situ and determine host response. Due to the rapid gelation of CS-T and PEG-4mal, they were injected using separate Hamilton syringes, mixed in a y-connector, and injected through a 27 gauge needle. All surgical procedures were performed aseptically. Rats were sedated using isoflurane and a local anesthetic was used as the skin and muscles were pulled back to reveal the spinal column. A laminectomy was performed on C7 to T1 and a 5 μL injection of sterile saline or sterile gel precursors was injected. Injections were 0.5 mm deep in the right dorsal region between C7 and C8 rootlets. The incision was closed and animals received analgesic for at least 24 h following surgery. After 7 or 14 d, animals were sacrificed via administration of intraperitoneal injection of ketamine/xylazine/acepromazine followed by transcardial perfusion of PBS. Tissue was dissected and post fixed.

Temperature sensitivity testing

Weekly functional assessments were made to detect any allodynia occurring in rats. Prior to surgery, rats were acclimated to hot/cold plate and enclosure at room temperature and subsequently tested at 48°C and 5°C. Time was recorded and animals were removed from hot/cold plate at rapid foot weight change (also any sign of distress such as vocalization, jumping, or paw licking). More moderate temperatures were tested first, if distress seen immediately time of 0 s was recorded for both the temperature tested and the more extreme corresponding temperature. After 7 and 14 d rats were tested again to determine latency times after incurring injury.

142

Tissue sectioning and staining

Spinal cord tissue containing gel injection site was post-fixed in 3.7%

paraformaldehyde for 48 h at 4°C then rinsed in PBS. Tissue samples were equilibrated in OCT medium at 4°C 18-24 h to preserve the tissue before sectioning. Sections were taken in 14-20 μm slices to see gross and fine architecture within the tissue. Thicker sections were necessary in some samples with larger cavities in the spinal cord. Select sections were stained overnight using the primary antibodies mouse-anti-GFAP

(Millipore, San Diego, CA, USA) or mouse-anti-ED1 (Bioss, Woburn, MA, USA) after blocking with 5% goat serum. The secondary antibody goat-anti-mouse IgG

AlexaFluor546 was incubated next on the samples and Hoechst was applied for nuclear staining. Alternate slides were stained with Hematoxylin and Eosin. Tissue section semi- quantitative analysis included calibrated measurements of stained sections under the microscope in ImageJ software.

Statistical analyses

Most analyses were run using analysis of variance (ANOVA), either one or two factor using JMP software. For two factor analysis, if the cross between the two factors is significant, a post-hoc Tukey’s test is used on the cross group averages. If there is no significance in the cross of the factors or if run as a single factor ANOVA, any significance in a single factor is investigated with Tukey’s post-hoc test. Results reported

143

in paper and used in graphs are mean ± standard deviation unless stated. Fold changes are

ratios of an experimental group mean to a “control” group mean and the standard

deviation is calculated as:

( / ) = ( / ) ( )/ + ( )/ 2 ( / )/( ) . 2 2 𝑆𝑆𝑆𝑆𝑆𝑆𝑆𝑆𝑆𝑆 𝑥𝑥 𝑦𝑦 𝑥𝑥̅ 𝑦𝑦� ∗ ��𝑉𝑉𝑉𝑉𝑉𝑉 𝑥𝑥 𝑥𝑥̅ 𝑉𝑉𝑉𝑉𝑉𝑉 𝑦𝑦 𝑦𝑦� − ∗ 𝐶𝐶𝐶𝐶𝐶𝐶 𝑥𝑥 𝑦𝑦 𝑥𝑥̅ ∗ 𝑦𝑦� �

Results

Chitosan gel properties

The three thiolated precursors were tested for swelling behavior immediately after

gelation with PEG-4mal, by immersing them in PBS for one day. Figure 5.2A displays

the results for each modification, showing that CS-T (Traut’s modification) along with

CS-TGA5 (thioglycolic acid modification, pH 5) experienced slight shrinking while CS-

TGA6 had very mild swelling. All three materials had similar ratios of swollen gel to dry weight (Fig. 5.2B). CS-T seemed the safest choice to avoid undue swelling and damage in vivo. We also observed the most consistency with the reaction and gelation of CS-T in

vitro and chose to move forward with this material for further testing. Thiol addition to

chitosan material using Traut’s reagent was confirmed using Ellman’s assay to be 255

µmol of thiol per g of polymer. Degradation studies show that the hydrogel loses roughly

30% of its dry mass (Fig. 5.2C) over two weeks, with no appreciable difference between

lysozyme or PBS buffers. Mechanical testing is displayed in Figure 5.2D; the complex

shear modulus of CS-T at 5% strain and 10 rad/s is 126 Pa. The elastic modulus (E) can

144 be determined by assuming a poisson’s ratio (ν) of 0.5 and using the expression

E=2G*(1+ν), yielding an elastic modulus of 378 Pa for CS-T/PEG-4mal hydrogels.

Figure 5.2: Mass change from hydrogel swelling is shown for post-gelation swelling (A) and the more traditional swelling ratio, Q (B). Post-gelation swelling tracks the mass change from an immediately mixed and formed hydrogel to 24 h in PBS, whereas the Q is the ratio of swollen gel mass to the dry mass. (C) Degradation profile of CS-T hydrogels in PBS and lysis buffer. (D) Rheometry graphs showing gel equilibrium mechanical properties of CS-T/PEG-4mal hydrogels. All shear moduli G’, G”, and G* are shown for frequency and strain sweeps. Data is displayed as mean ± SD; letters denote p<0.001; * = p<0.05, ** = p<0.001; n=4 for A-C; n=6 for D.

Cytotoxicity of fibroblasts and spinal cord cells

Cell cytotoxicity analysis after 24 h of exposure to chitosan/PEG gels revealed that none of the synthesized materials or their respective gels elicited an adverse reaction from fibroblasts. Figure 5.3A shows significant drops in cell activity as measured by

145

PrestoBlue in the CS-TGA5 and CS-TGA6 treatments; however, the activity level was

still above 70% of the healthy control cell level, which is the acceptable cutoff from ISO

10993-5 for in vitro cytotoxicity testing. This drop in cell activity was not reflected in cell

count data where there was no observable difference between groups (Fig. 5.3B). Figure

5.3C shows qualitatively that gels made from CS-T precursor closely resemble healthy

control cells, whereas the CS-TGA5 and CS-TGA6 gels have cells with irregular morphology including small, rounded cells. These data also lead to the choice of CS-T as the superior material for in vivo testing.

Figure 5.3: (A) Cell activity as measured by PrestoBlue; statistical significance was found between healthy control / CS-T wells and CS-TGA wells (p< 0.001). (B) Cell 146 number shown by nuclear staining and counting comparing control samples to gel containing samples. (C) Images of fibroblasts during test, comparing control wells with wells containing gels. Data is displayed as mean ± SD; n=4.

Host response safety assessment

The injected rodents showed no overt signs of functional deficit after initial recovery from surgery (~1 d). Sensitivity to heat or cold was tested using a Peltier block; data for responses at 48°C and 5°C are shown in Figure 5.4A. Latency before surgery was subtracted from latency measured at the endpoint of the study; thus, a negative change reflects a decrease in latency after spinal cord injection and a positive change indicates a longer post-treatment latency at the respective temperature. We expected latencies to decrease for all temperatures after surgery, more so at the acute 1 wk time- point. Decrease in latency suggests an increase in temperature sensitivity; however, no real differences or trends emerged from the data. Due to the large variability and tendency of rats to attempt jumping out during the second round of testing, no statistical analysis was performed in this data. Spinal cord response to surgical injection fell into two general categories: cavitation or hematoma. In some animals, as pictured in Figure

5.4, an open cavity was seen. Usually this area was surrounded by an area of high ED1 and GFAP staining, indicating reactive gliosis and potentially infiltration of macrophages. In other animals, a dark spot showing the affected area could be seen as the tissue was being sectioned. This area stained in H&E for red blood cells, thus we assumed it is a hematoma-like formation within the cord. In both saline and CS-T injected groups, at least one rodent experienced a cavitation or a hematoma-like response.

147

To compare the host response to injection of CS-T to the surgical sham animals

(injected with saline) the affected areas were imaged and measured for three animals in

the one week time-point. The affected shapes were very irregular, thus area measurements were used for comparison. Figure 5.5 shows bright areas of antibody recognition for reactive glia. In H&E sections, affected areas were denoted as irregular dark sections with too many cells, red staining of red blood cells, or cell lined cavities.

No significant difference could be detected between groups, but there was a high level of variability. In future studies, a larger sample size would give more power to analyses.

Figure 5.4: (A) Change in latency time from before and after surgical treatment for 48°C and 5°C. Testing was performed on n=4 animals per group, except 1 wk saline which had n=6. IHC images from CS-T injected spinal cord (B) and saline injected spinal cord (C) 148

showing the two typical responses; animals had cavitation in some cases (B) or red blood cell filled hematoma-like area (C). Inflammation can be seen in the bright areas in each image showing both reactive astrocytes (GFAP) and reactive microglia or invading macrophages (ED1). Scale bar 500 micron; WM: white matter; GM: grey matter.

Discussion

Successful thiolation of chitosan and gelation via the addition of a 4 armed PEG

terminated in maleimide groups was confirmed in the chemical and mechanical testing

data shown in Figure 5.2. Ellman’s assay demonstrated that the functionalized chitosan

contains about 255 µmol of thiol per g of polymer, which is consistent with previous

literature.[500] In addition, observations of modified chitosan confirmed thiolation was

successful, as the material was more soluble in water and would autogel in several days if

left unperturbed (data not shown). To minimize any further tissue damage after needle

puncture and material injection, a material that remains the same volumetric size is

desirable. Severe swelling is to be avoided, as outward pressure can damage neurons and

surrounding tissue, impeding the signaling function of the spinal cord. Here, we chose to

go forward with Traut’s modified chitosan (CS-T) because it shows slight shrinking in

post-gelation swelling mass change data (Fig. 5.2A). This approach was used to

specifically observe immediate swelling once the chitosan and PEG precursors were

mixed and the gel formed. Material swelling is a critical consideration in evaluating

safety; if the gel swells during formation it could apply high pressure to surrounding

tissue. The swelling ratio, Q (Fig. 5.2B), revealed that the three chitosan precursors are most likely very similar in cross-linking density and mechanical properties, as they had no difference or trend in swelling ratio. Our ratio (swollen mass/ dry mass ~60) is in the

149

range that observed for chitosan,[463] HA,[501] or combinations with PEG, [502, 503]

which range from 10-80. Further degradation testing showed that a small mass loss

occurred over 2 wk (Fig. 5.2C), but no difference in mass was seen when the buffer

contained lysozyme, a hydrolytic enzyme known to degrade chitosan. Mass loss may not

have been due to degradation in this time period, but perhaps unreacted material slowly

diffusing out of the scaffold over time. When handling the CS-T/PEG-4mal gel it was extremely deformable, sticky, and resistant to cutting. Rheometry data shows the high elasticity of the material, as it has a much lower viscous component (Fig. 5.2D). The complex shear modulus is 126 Pa and the elastic modulus is 378 Pa, or ~0.4 kPa, well within the range of reported mechanical properties of CNS tissue (50 Pa to 15 kPa).[504]

The resulting stiffness we have measured is within the range of a recent study using AFM to characterize Young’s modulus from both indentation and creep studies of the spinal cord in which researchers measured elastic moduli from about 50-125 Pa across white and grey matter.[505] As there are many methods and test parameters with which CNS tissue has been measured, the 0.1 kPa hydrogels reported here should be within the accepted range of mechanical stiffness.

150

Figure 5.5: Metrics from histological images were measured on three rodents per group at week 1 including: cavitation (A), affected area from H&E as well as macrophage (ED1) and reactive glia (GFAP) immunostaining (B). The ratio of affected area intensity was normalized to surrounding tissue (matching white/gray matter) and compared (C). A sample image shows an example of how the irregular areas were measured (D). Data in A are represented as individual data points to show variability in cavitation response; all other data shown as mean ± SD.

To assess the cellular response to thiolated chitosan-PEG gels, cultured fibroblasts

were used in a standard ISO 10993-5 test. Hydrogels made from CS-T, CS-TGA5, or CS-

TGA6 mixed with PEG-4mal were formed in transwell inserts and placed in wells of

confluent fibroblasts for 24 h. Figure 5.3A displays measured cell activity level in live

cells and although there was a significantly lower level in the gels made from CS-TGA, all cell activity levels were above 70% of the healthy controls. Seventy percent is the

cutoff suggested by ISO10993-5. Overall cell number was determined from immunostaining cells and there was no statistical difference between any groups (Fig.

5.3B). Morphologically, however, the CS-T/PEG-4mal gels showed the healthiest looking cells (Fig. 5.3C). Fibroblasts in the CS-TGA/PEG4mal treated wells showed many rounded, small cells with disrupted actin cytoskeletons. The favorable response, 151 along with swelling data, provided justification to continue on with Traut’s modified chitosan to in vivo studies.

The spinal cord, and CNS in general, is a complicated tissue with generally poor healing properties. In order to adequately evaluate the safety of the thiolated chitosan, it was intentionally injected directly into the cord. Direct injection into a three dimensional structure invariably stressed the surrounding tissue as room was made for the injected material. The same can be said for the saline injected animals, which served as a surgical sham. The initial injection site was assumed to create a small, focal injury in order to test the material under inflammatory conditions. Subsequent analysis aimed to evaluate tissue response beyond the mechanical insult of surgery and volume displacement. Grossly, there was no detectable difference between saline and material injected animals in their behavior, ambulation, or sensitivity to temperature (Fig. 5.4A). Behavioral testing in animals can be difficult as each has their own “personality” which affects their response to testing. Although rodents were acclimated to the testing apparatus, they did not enjoy being tested and thus some would attempt to jump out even though the temperature of the plate was not unpleasant. Overall, no response to treatment was seen in the animals other than brief surgical recovery.

One week after surgical treatment, vascular invasion via disrupted blood vessels, signaling and recruiting of native and non-native cells, and reactive gliosis in the lesion perimeter should be at or close to the maximum for an acute to sub-acute response.[506]

Acute events such as inflammation and parenchymal cell death will be visible at this time, if present. Qualitatively, the animals responded in two ways, which are exemplified

152

in Figure 5.4. In a few cords, large, dark hematoma-like areas appeared with little to no cavitation. While sectioning, a dark spot was visible in the nearly white tissue; staining with H&E revealed red blood cells in the affected area that were dark pink or red color surrounded by increased ED1 and GFAP staining. In other samples, large cavitation occurred with some increased reactive gliosis nearby. One sample in both the CS-T

injected group and saline injected group resulted in a large hematoma-like structure. At

least one saline injected rat also showed large cavitation as well. Thus responses were

mixed and likely due to the nature of the surgical procedure. A large inflammatory

response was not anticipated, considering the similarity in structure of chitosan and the

native ECM component hyaluronic acid (HA). HA and chitosan have been used

previously, and successfully, in CNS applications.[183, 487, 507] The placement of

injection in this study may yield higher inflammation from glia as it is disrupting tissue

within the cord from the initial injection where in many studies a material is placed

intrathecally, or somehow on the periphery of the native tissue. In the material injected

animals, no hydrogel was confirmed during staining, as material was probably lost during

the extensive washing and the slightly positive charge of the material repelling the

positively charged slides. Positive reactive glia immunostaining and cavitation did appear

more extensive overall in the material injected animals than in the surgical shams.

Variation in the CS-T injected group could also be due to irregular mixing and injection

of the co-polymers. Extensive material testing and larger group sizes in future studies will

help to confirm the pilot data in this chapter.

153

Semi-quantitative measures help describe the overall group effect of the

treatment. The surgical procedure essentially caused an acute, focal insult in the spinal

cord (regardless of material injected); however, injected CS-T persists in the space while

saline can diffuse away in a relatively short amount of time. Figure 5.5 describes the

surgical groups as a whole, through measurements of the area and intensity of

inflammatory markers and cavitation. Determining the correct area can be subjective but

bias was avoided whenever possible during area measurements of H&E and

immunostaining, however, it was not able to be blinded. Comparison with other studies

observing inflammation due to biomaterial implantation is difficult, as experimental

conditions are extremely important. Important considerations are that this is an acute

time-frame (versus long-term implantation), injection placement was within the spinal

cord as opposed to a peripheral area, no injury was created before injection, and no anti-

inflammatory therapeutic was included. As expected, affected areas measured (~1.5 mm2)

are comparable to most reported values. In a study by Kang et. Al., delivery of a hyaluronic acid/ methylcellulose composite gel releasing the neuroprotective agent erythropoietin caused an affected area of 6-10 x 105 µm2 (by ED1 signal) six weeks after

a spinal cord contusion injury.[508] With the intraspinal injection used in this study,

variation within groups made differences between groups undetectable in a statistical

analysis. Pilot data reported here shows that the chitosan/PEG material works well as a

bulk injectable material for the spinal cord parenchyma, and has potential in future

delivery applications because it does not show extreme signs of toxicity.

154

Conclusion

An in situ cross-linking hydrogel was successfully synthesized and characterized in this study for intended use in regenerative CNS strategies. Chitosan, an abundant natural polymer similar to ECM components, was functionalized with Traut’s reagent to add a reactive thiol groups. When combined with maleimide terminated PEG, the precursors formed a covalently cross-linked hydrogel with appropriate mechanical properties for the CNS including a low complex modulus with high elasticity, minor shrinking in PBS, and low susceptibility to lysozyme degradation. The material was non- toxic to fibroblasts in vitro, and showed a similar muted host response to a saline sham group when injected directly in the spinal cord of rats. Taken together, we believe the chitosan-PEG copolymer to be an appropriate base biomaterial for future spinal cord delivery strategies.

155

CHAPTER 6

CONCLUSIONS

The guiding objective of this thesis was to combine aspects of spinal cord injury

with stem cell guidance techniques into a biomaterial therapeutic system to promote

regeneration. The devastating effects of spinal cord injury are felt in the healthcare

system and more importantly they decrease patient quality of life. The National SCI

Statistical Center estimates there are 17,000 new cases of SCI each year in the USA alone. Average age at injury is 42 years old, and for para- or tetraplegia patients overall life expectancy can decrease by 10-20 years. Important groundwork for a therapeutic strategy targeting the CNS, and spinal cord in particular, has been laid down in Chapters

3-5. This work provides an in depth look at injury in the spinal cord and makes steps toward combating further damage and tissue loss. Specifically, investigation of syrinx progression helped to better characterize the injury environment and establish molecular targets. Following this, two major approaches were developed to aid in future regeneration strategies. First, control of native NSC differentiation using soluble and surface cues yielded a high percentage of cells maturing into a neuronal phenotype.

Second, an injectable hydrogel was created and initial testing was performed in order to determine future utility in spinal cord delivery schemes.

156

Spinal cord cavitation was described biochemically in Chapter 3, with an

overview on transcriptional change and a deeper query into critical aspects of fluid movement. Validation of the injury was shown through massive changes in inflammatory markers and losses in neuronal transmission. A dramatic increase in migratory transcripts for white blood cells suggested a compromised vasculature in syrinx containing samples, disrupting the delicate balance normally maintained in the CNS through the blood-brain

barrier. Loss of neuronal transmission was seen through loss of signaling transcripts and a

decrease in myelin membrane components. Indeed this loss of neurons and neuronal

transmission manifested in the animals, where no increased sensitivity was seen in

excitotoxic animals compared to healthy controls. It was hypothesized that the aquaporin

family would be highly dysregulated and that it and a few other pathways would stand

out in the analysis as being primarily responsible for syrinx enlargement. In reality,

culprits responsible for syrinx enlargement were confounded by the massive

inflammatory markers and changes in neuronal signaling molecules. Led by extensive

literature describing the fluid movement out of syrinx as it expands and suspected

pressure and flow differentials responsible for cavitation, osmolytes and their respective

transporters as well as water channels were scrutinized. Osmolytes betaine, taurine, and

carnitine drastically increased by 3 wks after injury, and the betaine/GABA (BGT-1) and

carnitine (SLC22A5) transporters saw a corresponding increase at 6 wks. Meanwhile, the

potassium channel KCC4 and water channel AQP1 both increased over 3 and 6 wks.

Although it was difficult to determine cell type(s) responsible for the increase in channel

expression, we believe these to be important in fluid movement out of the syrinx. This

157

investigation set the stage for strategies of combating SCI, saving and replacing lost

tissue, and overall diminishing injury progression. Carrying on this work in the Leipzig

Lab, there is current investigation into antagonizing BGT-1 and aquaporin channels.

Looking at the injury at more time-points would also be an interesting addition; perhaps a

time-course investigation could shed more light on which molecules are responsible and

which are results of cavitation.

In Chapter 4, control of native stem cell behavior was explored. NSC phenotype

could be significantly changed by chemical and physical cues, demonstrating the

importance of culture substrate properties. This is especially true with sensitive cells such

as stem cells, which readily respond to the surrounding environment. Overall the cells

preferred the softer, chitosan based substrate regardless of cell adhesion protein; all MAC

coated groups had a higher cell number than the corresponding glass groups. Determining

the optimal culture surface to specify cell phenotype was more confounded. Laminin

coated MAC groups showed the most consistent neuronal population (with high βIII

tubulin and synapsin staining) while expressing lower trends of glial markers.

Surprisingly, observations of NSCs in adsorbed laminin surfaces showed more desirable

morphology and trended toward more neuronal phenotype than covalently immobilized

protein. This data disproved part of the guiding hypothesis that immobilized laminin on

hydrogel substrates would yield the most neuronal differentiation. A nonspecific

immobilization strategy was used to covalently attach proteins laminin and collagen,

however, enzyme immobilization strategies have shown that site specific immobilization leads to increased activity over random attachment.[509] Results from Chapter 4 could

158

also suggest an internalization of the basement membrane protein is important in its

signaling cascade. Additionally, soluble IFN- γ was present to help guide NSC behavior.

The final results were a largely pure population of cells maturing into a neuronal

phenotype. This work and the electrical stimulation data from Appendix B shows that

stem cell fate can be directed with the proper cues. Specifically, the electrical stimulation

data showed that precise levels of DC electrical field yielded more neurite branching and

length; too much current resulted in cell death. It also became apparent that media

supplementation greatly affected neurite extension in the electric field. Future studies

involving cell migration would add nicely to this work. Additionally, a conductive

substrate in the presence of an electric field could enhance cell maturation and neurite

elongation on a chitosan based substrate.

Findings described here have set up future studies aiming to devise specific

combative strategies to SCI, especially syringomyelia. The chitosan/PEG hydrogel was

shown to be non-toxic both in vitro and in vivo, and displayed excellent mechanical

properties in vitro suggesting it would make an excellent delivery vehicle in the spinal

cord for cells, pharmaceuticals, guidance cues, and/or trophic factors. Post gelation

swelling was actually negative, indicating slight shrinking, which can be safer for the

surrounding tissue when the material is injected in situ. Future development on this

material could involve design of a syringe device that enhances homogenous mixing for consistent delivery, incorporation of an enzymatically labile linker to speed degradation, or addition of pharmaceuticals and/or proteins in particles for sustained release. In delivering specific factors, Chapter 3 provides probable molecular targets involved in

159

fluid or solute transport. To combat further cavitation, a pharmaceutical or antagonist to osmolyte transporters (BGT-1 or carnitine transporter), ion channels (KCC4), or aquaporins could be delivered. These agent(s) would promote survival of tissue and aim to prevent any further damage. To repair damaged or lost parenchyma, native NSCs from the brain or central canal of the spinal cord could be recruited and guided into specific phenotypes. An interesting future direction of this work would include probing the recruitment of NSCs, and subsequently directing their differentiation into a specified phenotype. This work shows that release or specific immobilization of laminin and IFN-γ

should lead to a large population of neurons. In conclusion, these neural regenerative approaches can span the gap to create multi-faceted tissue constructs or devices that are injected into various injury sites for tissue sparing and regeneration.

160

REFERENCES

1. Spinal cord injury facts and figures at a glance. 2010, National Spinal Cord Injury Statistical Center: Birmingham, AL. p. 2. 2. Faul M, X.L., Wald MM, Coronado VG, Traumatic brain injury in the United States: emergency department visits, hospitalizations, and deaths. 2010, Centers for Disease Control and Prevention, National Center for Injury Prevention and Control: Atlanta, GA. 3. Jacobson, S. and E.M. Marcus, Neuroanatomy for the neuroscientist. 2008, Springer: Boston, MA. 4. Schmidt, C.E. and J.B. Leach, Neural tissue engineering: Strategies for repair and regeneration. Annual Review of Biomedical Engineering, 2003. 5: p. 293- 347. 5. Levitan, I.B. and L.K. Kaczmarek, THE NEURON CELL AND MOLECULAR BIOLOGY. Levitan, I. B. and L. K. Kaczmarek. the Neuron: Cell and Molecular Biology. Xii+450p. Oxford University Press, Inc.: New York, New York, USA; Oxford, England, Uk. Illus, 1991: p. XII+450P. 6. Dityatev, A., C.I. Seidenbecher, and M. Schachner, Compartmentalization from the outside: the extracellular matrix and functional microdomains in the brain. Trends in Neurosciences, 2010. 33(11): p. 503-512. 7. Yu, L.M.Y., N.D. Leipzig, and M.S. Shoichet, Promoting neuron adhesion and growth. Materials Today, 2008. 11(5): p. 36-43. 8. Milner, R. and I.L. Campbell, The integrin family of cell adhesion molecules has multiple functions within the CNS. Journal of Neuroscience Research, 2002. 69(3): p. 286-291. 9. Hubert, T., et al., Collagens in the developing and diseased nervous system. Cellular and Molecular Life Sciences, 2009. 66(7): p. 1223-1238. 10. Suzuki, T., et al., Recent advances in the study of AMPA receptors. Nihon yakurigaku zasshi. Folia pharmacologica Japonica, 2003. 122(6): p. 515-26. 11. Costa, C., et al., Mapping of aggrecan, hyaluronic acid, heparan sulphate proteoglycans and aquaporin 4 in the central nervous system of the mouse. Journal of Chemical Neuroanatomy, 2007. 33(3): p. 111-123. 12. Ruoslahti, E., Brain extracellular matrix. Glycobiology, 1996. 6(5): p. 489-492. 13. Smith, D.H., Stretch growth of integrated axon tracts: Extremes and exploitations. Progress in Neurobiology, 2009. 89(3): p. 231-239. 14. Mallavarapu, A. and T. Mitchison, Regulated actin cytoskeleton assembly at filopodium tips controls their extension and retraction. Journal of Cell Biology, 1999. 146(5): p. 1097-1106.

161

15. Huber, A.B., et al., Signaling at the growth cone: Ligand-receptor complexes and the control of axon growth and guidance. Annual Review of Neuroscience, 2003. 26: p. 509-563. 16. Davis, L., et al., Protein-synthesis within neuronal growth cones. Journal of Neuroscience, 1992. 12(12): p. 4867-4877. 17. Luo, L. and D. O'Leary, Axon retraction and degeneration in development and disease. Annual Review of Neuroscience, 2005: p. 127-156. 18. Edgar, J.M. and K.A. Nave, The role of CNS glia in preserving axon function. Current Opinion in Neurobiology, 2009. 19(5): p. 498-504. 19. Ebneter, A., et al., Microglial Activation in the Visual Pathway in Experimental Glaucoma: Spatiotemporal Characterization and Correlation with Axonal Injury. Investigative Ophthalmology & Visual Science, 2010. 51(12): p. 6448-6460. 20. Prewitt, C.M.F., et al., Activated macrophage/microglial cells can promote the regeneration of sensory axons into the injured spinal cord. Experimental Neurology, 1997. 148(2): p. 433-443. 21. Thored, P., et al., Long-Term Accumulation of Microglia with Proneurogenic Phenotype Concomitant with Persistent Neurogenesis in Adult Subventricular Zone After Stroke. Glia, 2009. 57(8): p. 835-849. 22. Cizkova, D., et al., Response of Ependymal Progenitors to Spinal Cord Injury or Enhanced Physical Activity in Adult Rat. Cellular and Molecular Neurobiology, 2009. 29(6-7): p. 999-1013. 23. Hamilton, L.K., et al., Cellular organization of the central canal ependymal zone, a niche of latent neural stem cells in the adult mammalian spinal cord. Neuroscience, 2009. 164(3): p. 1044-1056. 24. Hugnot, J.P. and R. Franzen, The spinal cord ependymal region: A stem cell niche in the caudal central nervous system. Frontiers in Bioscience-Landmark, 2011. 16: p. 1044-1059. 25. Basso, D.M., M.S. Beattie, and J.C. Bresnahan, A SENSITIVE AND RELIABLE LOCOMOTOR RATING-SCALE FOR OPEN-FIELD TESTING IN RATS. Journal of Neurotrauma, 1995. 12(1): p. 1-21. 26. Lebedev, S.V., et al., Exercise Tests and BBB Method for Evaluation of Motor Disorders in Rats after Contusion Spinal Injury. Bulletin of Experimental Biology and Medicine, 2008. 146(4): p. 489-494. 27. Solomon, F. and M. Magendantz, Cytochalasin separates microtubule disassembly from loss of asymmetric morphology. J Cell Biol, 1981. 89(1): p. 157-61. 28. Ahmad, F.J., et al., Motor proteins regulate force interactions between microtubules and microfilaments in the axon. Nat Cell Biol, 2000. 2(5): p. 276- 80. 29. Nakayama, A.Y., M.B. Harms, and L. Luo, Small GTPases Rac and Rho in the maintenance of dendritic spines and branches in hippocampal pyramidal neurons. J Neurosci, 2000. 20(14): p. 5329-38. 30. Guan, K.L. and Y. Rao, Signalling mechanisms mediating neuronal responses to guidance cues. Nat Rev Neurosci, 2003. 4(12): p. 941-56.

162

31. He, Z. and V. Koprivica, The Nogo signaling pathway for regeneration block. Annu Rev Neurosci, 2004. 27: p. 341-68. 32. Coleman, M., Axon degeneration mechanisms: Commonality amid diversity. Nature Reviews Neuroscience, 2005: p. 889-898. 33. Saxena, S. and P. Caroni, Mechanisms of axon degeneration: From development to disease. Progress in Neurobiology, 2007: p. 174-191. 34. Coleman, M.P. and M.R. Freeman, Wallerian Degeneration, Wld(S), and Nmnat. Annual Review of Neuroscience, Vol 33, 2010. 33: p. 245-267. 35. Waller, A., Experiments on the sections of glossopharyngeal and hypoglossal nerves of the frog and observations of the alterations produced thereby in the structures of their primitive fibers. Philosophical Transactions of the Royal Society London, 1850. 140. 36. Benbassat, D. and M.E. Spira, The survival of transected axonal segments of cultured aplysia neurons is prolonged by contact with intact nerve-cells. European Journal of Neuroscience, 1994. 6(10): p. 1605-1614. 37. Yiu, G. and Z.G. He, Glial inhibition of CNS axon regeneration. Nature Reviews Neuroscience, 2006. 7(8): p. 617-627. 38. LUNN, E., et al., ABSENCE OF WALLERIAN DEGENERATION DOES NOT HINDER REGENERATION IN PERIPHERAL-NERVE. European Journal of Neuroscience, 1989: p. 27-33. 39. Beirowski, B., et al., The progressive nature of Wallerian degeneration in wild- type and slow Wallerian degeneration (WldS) nerves. BMC Neuroscience, 2005. 6(February 1). 40. Raff, M.C., A.V. Whitmore, and J.T. Finn, Neuroscience - Axonal self-destruction and neurodegeneration. Science, 2002. 296(5569): p. 868-871. 41. Glass, J.D., et al., PROLONGED SURVIVAL OF TRANSECTED NERVE-FIBERS IN C57BL/OLA MICE IS AN INTRINSIC CHARACTERISTIC OF THE AXON. Journal of Neurocytology, 1993. 22(5): p. 311-321. 42. Wright, A.K., et al., Synaptic Protection in the Brain of Wld(S) Mice Occurs Independently of Age but Is Sensitive to Gene-Dose. Plos One, 2010. 5(11): p. 9. 43. Mack, T.G.A., et al., Wallerian degeneration of injured axons and synapses is delayed by a Ube4b/Nmnat chimeric gene. Nature Neuroscience, 2001. 4(12): p. 1199-1206. 44. Bisby, M.A. and S. Chen, DELAYED WALLERIAN DEGENERATION IN SCIATIC-NERVES OF C57BL/OLA MICE IS ASSOCIATED WITH IMPAIRED REGENERATION OF SENSORY AXONS. Brain Research, 1990. 530(1): p. 117- 120. 45. Brown, M.C., E.R. Lunn, and V.H. Perry, CONSEQUENCES OF SLOW WALLERIAN DEGENERATION FOR REGENERATING MOTOR AND SENSORY AXONS. Journal of Neurobiology, 1992. 23(5): p. 521-536. 46. Martin, S.M., et al., Wallerian degeneration of zebrafish trigeminal axons in the skin is required for regeneration and developmental pruning. Development, 2010. 137(23): p. 3985-3994.

163

47. Zhai, Q.W., et al., Involvement of the ubiquitin-proteasome system in the early stages of Wallerian degeneration. Neuron, 2003. 39(2): p. 217-225. 48. Ghosh-Roy, A., et al., Calcium and Cyclic AMP Promote Axonal Regeneration in Caenorhabditis elegans and Require DLK-1 Kinase. Journal of Neuroscience, 2010. 30(9): p. 3175-3183. 49. Ferri, A., et al., Inhibiting axon degeneration and synapse loss attenuates apoptosis and disease progression in a mouse model of motoneuron disease. Current Biology, 2003. 13(8): p. 669-673. 50. Horste, G.M.Z., et al., The Wlds transgene reduces axon loss in a Charcot-Marie- Tooth disease 1A rat model and nicotinamide delays post-traumatic axonal degeneration. Neurobiology of Disease, 2011. 42(1): p. 1-8. 51. Beirowski, B., et al., Mechanisms of Axonal Spheroid Formation in Central Nervous System Wallerian Degeneration. Journal of Neuropathology and Experimental Neurology, 2010: p. 455-472. 52. Mi, W.Q., et al., The slow Wallerian degeneration gene, Wld(S), inhibits axonal spheroid pathology in gracile axonal dystrophy mice. Brain, 2005. 128: p. 405- 416. 53. Sajadi, A., B.L. Schneider, and P. Aebischer, Wld(s)-mediated protection of dopaminergic fibers in an animal model of Parkinson disease. Current Biology, 2004. 14(4): p. 326-330. 54. Samsam, M., et al., The Wld(s) mutation delays robust loss of motor and sensory axons in a genetic model for myelin-related axonopathy. Journal of Neuroscience, 2003. 23(7): p. 2833-2839. 55. Wang, M.S., et al., The Wld(S) protein protects against axonal degeneration: A model of gene therapy for peripheral neuropathy. Annals of Neurology, 2001. 50(6): p. 773-779. 56. Vargas, M.E. and B.A. Barres, Why is Wallerian degeneration in the CNS so slow? Annual Review of Neuroscience, 2007. 30: p. 153-179. 57. Huebner, E.A., et al., A Multi-domain Fragment of Nogo-A Protein Is a Potent Inhibitor of Cortical Axon Regeneration via Nogo Receptor 1. Journal of Biological Chemistry, 2011. 286(20): p. 18026-18036. 58. Wang, D., et al., Neural stem cell transplantation with Nogo-66 receptor gene silencing to treat severe traumatic brain injury. Neural Regeneration Research, 2011. 6(10): p. 725-731. 59. Gonzenbach, R.R. and M.E. Schwab, Disinhibition of neurite growth to repair the injured adult CNS: Focusing on Nogo. Cellular and Molecular Life Sciences, 2008. 65(1): p. 161-176. 60. Li, M., et al., Myelin-associated glycoprotein inhibits neurite/axon growth and causes growth cone collapse. Journal of Neuroscience Research, 1996. 46(4): p. 404-414. 61. Quarles, R.H., A Hypothesis About the Relationship of Myelin-Associated Glycoprotein's Function in Myelinated Axons to its Capacity to Inhibit Neurite Outgrowth. Neurochemical Research, 2009. 34(1): p. 79-86.

164

62. Ridet, J.L., et al., Reactive astrocytes: cellular and molecular cues to biological function. Trends in Neurosciences, 1997. 20(12): p. 570-577. 63. Menet, V., et al., Axonal plasticity and functional recovery after spinal cord injury in mice deficient in both glial fibrillary acidic protein and vimentin genes. Proceedings of the National Academy of Sciences of the United States of America, 2003. 100(15): p. 8999-9004. 64. Niederost, B.P., et al., Bovine CNS myelin contains neurite growth-inhibitory activity associated with chondroitin sulfate proteoglycans. Journal of Neuroscience, 1999. 19(20): p. 8979-8989. 65. Bartanusz, V., et al., The blood-spinal cord barrier: morphology and clinical implications. Ann Neurol, 2011. 70(2): p. 194-206. 66. Sharma, H.S., Pathophysiology of blood-spinal cord barrier in traumatic injury and repair. Curr Pharm Des, 2005. 11(11): p. 1353-89. 67. Pedersen, M.O., et al., Cell death in the injured brain: roles of metallothioneins. Prog Histochem Cytochem, 2009. 44(1): p. 1-27. 68. Bonhomme, V., et al., NEURON-SPECIFIC ENOLASE AS A MARKER OF INVITRO NEURONAL DAMAGE .2. INVESTIGATION OF THE ASTROCYTE PROTECTIVE EFFECT AGAINST KAINATE-INDUCED NEUROTOXICITY. Journal of Neurosurgical Anesthesiology, 1993. 5(2): p. 117-120. 69. Biomaterials science: An introduction to materials in medicine. Biomaterials science: An introduction to materials in medicine, ed. B.D. Ratner, et al. 1996: Academic Press, Inc.; Academic Press Ltd. xi+484p. 70. Zhong, Y. and R.V. Bellamkonda, Biomaterials for the central nervous system. Journal of the Royal Society Interface, 2008. 5(26): p. 957-975. 71. Peyton, S.R., et al., The emergence of ECM mechanics and cytoskeletal tension as important regulators of cell function. Cell Biochemistry and Biophysics, 2007. 47(2): p. 300-320. 72. Nair, L.S. and C.T. Laurencin, Biodegradable polymers as biomaterials. Progress in Polymer Science, 2007. 32(8-9): p. 762-798. 73. Nisbet, D.R., et al., Neural tissue engineering of the CNS using hydrogels. Journal of Biomedical Materials Research Part B-Applied Biomaterials, 2008. 87B(1): p. 251-263. 74. Straley, K.S., C.W.P. Foo, and S.C. Heilshorn, Biomaterial Design Strategies for the Treatment of Spinal Cord Injuries. Journal of Neurotrauma, 2010. 27(1): p. 1- 19. 75. Lee, K.Y. and D.J. Mooney, Hydrogels for tissue engineering. Chemical Reviews, 2001. 101(7): p. 1869-1879. 76. West, J.L. and J.A. Hubbell, Polymeric biomaterials with degradation sites for proteases involved in cell migration. Macromolecules, 1999. 32(1): p. 241-244. 77. Lee, S.H., et al., Poly(ethylene glycol) hydrogels conjugated with a collagenase- sensitive fluorogenic substrate to visualize collagenase activity during three- dimensional cell migration. Biomaterials, 2007. 28(20): p. 3163-3170. 78. Kridel, S.J., et al., Substrate hydrolysis by matrix metalloproteinase-9. Journal of Biological Chemistry, 2001. 276(23): p. 20572-20578.

165

79. Sottrup-Jensen, L. and H. Birkedal-Hansen, Human fibroblast collagenase-alpha- macroglobulin interactions. Localization of cleavage sites in the bait regions of five mammalian alpha-macroglobulins. J Biol Chem, 1989. 264(1): p. 393-401. 80. Turk, B.E., et al., Determination of protease cleavage site motifs using mixture- based oriented peptide libraries. Nature Biotechnology, 2001. 19(7): p. 661-667. 81. Chau, Y., et al., Incorporation of a matrix metalloproteinase-sensitive substrate into self-assembling peptides - A model for biofunctional scaffolds. Biomaterials, 2008. 29(11): p. 1713-1719. 82. Chau, Y., F.E. Tan, and R. Langer, Synthesis and characterization of dextran- peptide-methotrexate conjugates for tumor targeting via mediation by matrix metalloproteinase II and matrix metalloproteinase IX. Bioconjugate Chemistry, 2004. 15(4): p. 931-941. 83. Tonti, G.A., et al., Neural stem cells at the crossroads: MMPs may tell the way. Int J Dev Biol, 2009. 53(1): p. 1-17. 84. Ogier, C., et al., Matrix metalloproteinase-2 (MMP-2) regulates astrocyte motility in connection with the actin cytoskeleton and integrins. Glia, 2006. 54(4): p. 272- 84. 85. Zuo, J., et al., Neuronal matrix metalloproteinase-2 degrades and inactivates a neurite-inhibiting chondroitin sulfate proteoglycan. J Neurosci, 1998. 18(14): p. 5203-11. 86. Tayebjee, M.H., et al., Effects of age, gender, ethnicity, diurnal variation and exercise on circulating levels of matrix metalloproteinases (MMP)-2 and-9, and their inhibitors, tissue inhibitors of matrix metalloproteinases (TIMP)-1 and-2. Thrombosis Research, 2005. 115(3): p. 205-210. 87. Katti, D.S., et al., Toxicity, biodegradation and elimination of polyanhydrides. Advanced Drug Delivery Reviews, 2002. 54(7): p. 933-961. 88. Leong, K.W., B.C. Brott, and R. Langer, BIOERODIBLE POLYANHYDRIDES AS DRUG-CARRIER MATRICES .1. CHARACTERIZATION, DEGRADATION, AND RELEASE CHARACTERISTICS. Journal of Biomedical Materials Research, 1985. 19(8): p. 941-955. 89. Baumann, M.D., et al., Intrathecal delivery of a polymeric nanocomposite hydrogel after spinal cord injury. Biomaterials, 2010. 31(30): p. 7631-7639. 90. Pan, L., et al., Viability and Differentiation of Neural Precursors on Hyaluronic Acid Hydrogel Scaffold. Journal of Neuroscience Research, 2009. 87(14): p. 3207-3220. 91. Orive, G., et al., Biomaterials for promoting brain protection, repair and regeneration. Nature Reviews Neuroscience, 2009. 10(9): p. 682-U47. 92. Leipzig, N.D. and M.S. Shoichet, The effect of substrate stiffness on adult neural stem cell behavior. Biomaterials, 2009. 30(36): p. 6867-6878. 93. Saha, K., et al., Substrate Modulus Directs Neural Stem Cell Behavior. Biophysical Journal, 2008. 95(9): p. 4426-4438. 94. Cong Truc, H., N. Minh Khanh, and D.S. Lee, Injectable Block Copolymer Hydrogels: Achievements and Future Challenges for Biomedical Applications. Macromolecules, 2011. 44(17): p. 6629-6636.

166

95. Jain, A., et al., In situ gelling hydrogels for conformal repair of spinal cord defects, and local delivery of BDNF after spinal cord injury. Biomaterials, 2006. 27(3): p. 497-504. 96. Nguyen, M.K. and D.S. Lee, Injectable biodegradable hydrogels. Macromolecular bioscience, 2010. 10(6): p. 563-79. 97. Gupta, D., C.H. Tator, and M.S. Shoichet, Fast-gelling injectable blend of hyaluronan and methylcellulose for intrathecal, localized delivery to the injured spinal cord. Biomaterials, 2006. 27(11): p. 2370-2379. 98. Dodla, M.C. and R.V. Bellamkonda, Differences between the effect of anisotropic and isotropic laminin and nerve growth factor presenting scaffolds on nerve regeneration across long peripheral nerve gaps. Biomaterials, 2008. 29(1): p. 33- 46. 99. Pfister, L.A., et al., Nerve conduits and growth factor delivery in peripheral nerve repair. Journal of the Peripheral Nervous System, 2007. 12(2): p. 65-82. 100. Lynn, A.K., I.V. Yannas, and W. Bonfield, Antigenicity and immunogenicity of collagen. Journal of Biomedical Materials Research Part B-Applied Biomaterials, 2004. 71B(2): p. 343-354. 101. Fan, J., et al., Linear Ordered Collagen Scaffolds Loaded with Collagen-Binding Neurotrophin-3 Promote Axonal Regeneration and Partial Functional Recovery after Complete Spinal Cord Transection. Journal of Neurotrauma, 2010. 27(9): p. 1671-1683. 102. Sun, W., et al., The effect of collagen-binding NGF-beta on the promotion of sciatic nerve regeneration in a rat sciatic nerve crush injury model. Biomaterials, 2009. 30(27): p. 4649-56. 103. Zhang, T., et al., Three-dimensional gelatin and gelatin/hyaluronan hydrogel structures for traumatic brain injury. Journal of Bioactive and Compatible Polymers, 2007. 22(1): p. 19-29. 104. Zhang, H.Z., et al., Gelatin-siloxane hybrid scaffolds with vascular endothelial growth factor induces brain tissue regeneration. Current Neurovascular Research, 2008. 5(2): p. 112-117. 105. Liu, T., et al., Photochemical crosslinked electrospun collagen nanofibers: Synthesis, characterization and neural stem cell interactions. Journal of Biomedical Materials Research Part A, 2010. 95A(1): p. 276-282. 106. Timnak, A., et al., Fabrication of nano-structured electrospun collagen scaffold intended for nerve tissue engineering. Journal of Materials Science-Materials in Medicine, 2011. 22(6): p. 1555-1567. 107. Wang, T.-W. and M. Spector, Development of hyaluronic acid-based scaffolds for brain tissue engineering. Acta Biomaterialia, 2009. 5(7): p. 2371-2384. 108. Wei, Y.-T., et al., Hyaluronic Acid Hydrogel Modified with Nogo-66 Receptor Antibody and Poly(L-Lysine) Enhancement of Adherence and Survival of Primary Hippocampal Neurons. Journal of Bioactive and Compatible Polymers, 2009. 24(3): p. 205-219.

167

109. Cullen, D.K., et al., In vitro neural injury model for optimization of tissue- engineered constructs. Journal of Neuroscience Research, 2007. 85(16): p. 3642- 3651. 110. Tate, M.C., et al., Biocompatibility of methylcellulose-based constructs designed for intracerebral gelation following experimental traumatic brain injury. Biomaterials, 2001. 22(10): p. 1113-1123. 111. Crompton, K.E., et al., Inflammatory response on injection of chitosan/GP to the brain. Journal of Materials Science-Materials in Medicine, 2006. 17(7): p. 633- 639. 112. Sudarshan, N.R., D.G. Hoover, and D. Knorr, Antibacterial action of chitosan. Food Biotechnology, 1992. 6(3): p. 257-272. 113. Crompton, K.E., et al., Polylysine-functionalised thermoresponsive chitosan hydrogel for neural tissue engineering. Biomaterials, 2007. 28(3): p. 441-449. 114. Yu, L.M.Y., K. Kazazian, and M.S. Shoichet, Peptide surface modification of methacrylamide chitosan for neural tissue engineering applications. Journal of Biomedical Materials Research Part A, 2007. 82A(1): p. 243-255. 115. Cao, Z., R.J. Gilbert, and W. He, Simple Agarose-Chitosan Gel Composite System for Enhanced Neuronal Growth in Three Dimensions. Biomacromolecules, 2009. 10(10): p. 2954-2959. 116. Dainiak, M.B., et al., Gelatin-fibrinogen cryogel dermal matrices for wound repair: Preparation, optimisation and in vitro study. Biomaterials, 2010. 31(1): p. 67-76. 117. Nikonorov, V.V., et al., Synthesis and Characteristics of Cryogels of Chitosan Crosslinked by Glutaric Aldehyde. Polymer Science Series A, 2010. 52(8): p. 828-834. 118. Orrego, C.E. and J.S. Valencia, Preparation and characterization of chitosan membranes by using a combined freeze gelation and mild crosslinking method. Bioprocess and Biosystems Engineering, 2009. 32(2): p. 197-206. 119. Jeong, S.I., et al., Electrospun Chitosan-Alginate Nanofibers with In Situ Polyelectrolyte Complexation for Use as Tissue Engineering Scaffolds. Tissue Engineering Part A, 2011. 17(1-2): p. 59-70. 120. Prabhakaran, M.P., et al., Electrospun Biocomposite Nanofibrous Scaffolds for Neural Tissue Engineering. Tissue Engineering Part A, 2008. 14(11): p. 1787- 1797. 121. Bjugstad, K.B., et al., Biocompatibility of poly(ethylene glycol)-based hydrogels in the brain: An analysis of the glial response across space and time. Journal of Biomedical Materials Research Part A, 2010. 95A(1): p. 79-91. 122. Piantino, J., et al., An injectable, biodegradable hydrogel for trophic factor delivery enhances axonal rewiring and improves performance after spinal cord injury. Experimental Neurology, 2006. 201(2): p. 359-367. 123. Baichwal, R.R., J.W. Bigbee, and G.H. Devries, MACROPHAGE-MEDIATED MYELIN-RELATED MITOGENIC FACTOR FOR CULTURED SCHWANN- CELLS. Proceedings of the National Academy of Sciences of the United States of America, 1988. 85(5): p. 1701-1705.

168

124. Han, M.H., et al., THE ROLE OF SCHWANN-CELLS AND MACROPHAGES IN THE REMOVAL OF MYELIN DURING WALLERIAN DEGENERATION. Acta Histochemica Et Cytochemica, 1989. 22(2): p. 161-172. 125. Heumann, R., et al., DIFFERENTIAL REGULATION OF MESSENGER-RNA ENCODING NERVE GROWTH-FACTOR AND ITS RECEPTOR IN RAT SCIATIC-NERVE DURING DEVELOPMENT, DEGENERATION, AND REGENERATION - ROLE OF MACROPHAGES. Proceedings of the National Academy of Sciences of the United States of America, 1987. 84(23): p. 8735- 8739. 126. Meyer, M., et al., Enhanced synthesis of brain-derived neurotrophic factor in the lesioned peripheral-nerve - Different mechanisms are responsible for the regulation of BDNF and NGF messenger-RNA. Journal of Cell Biology, 1992. 119(1): p. 45-54. 127. Gilmore, S.A. and D. Duncan, ON PRESENCE OF PERIPHERAL-LIKE NERVOUS AND CONNECTIVE TISSUE WITHIN IRRADIATED SPINAL CORD. Anatomical Record, 1968. 160(4): p. 675-&. 128. Hirano, A., Zimmerma.Hm, and S. Levine, ELECTRON MICROSCOPIC OBSERVATIONS OF PERIPHERAL MYELIN IN A CENTRAL NERVOUS SYSTEM LESION. Acta Neuropathologica, 1969. 12(4): p. 348-&. 129. Raine, C.S., OCCURRENCE OF SCHWANN-CELLS WITHIN NORMAL CENTRAL NERVOUS-SYSTEM. Journal of Neurocytology, 1976. 5(3): p. 371- 380. 130. Iwashita, Y. and W.F. Blakemore, Areas of demyelination do not attract significant numbers of Schwann cells transplanted into normal white matter. Glia, 2000. 31(3): p. 232-240. 131. Iwashita, Y., et al., Schwann cells transplanted into normal and x-irradiated adult white matter do not migrate extensively and show poor long-term survival. Experimental Neurology, 2000. 164(2): p. 292-302. 132. Blakemore, W.F., LIMITED REMYELINATION OF CNS AXONS BY SCHWANN- CELLS TRANSPLANTED INTO THE SUB-ARACHNOID SPACE. Journal of the Neurological Sciences, 1984. 64(3): p. 265-276. 133. Blakemore, W.F., A.J. Crang, and R. Curtis, THE INTERACTION OF SCHWANN-CELLS WITH CNS AXONS IN REGIONS CONTAINING NORMAL ASTROCYTES. Acta Neuropathologica, 1986. 71(3-4): p. 295-300. 134. Bachelin, C., et al., Ectopic expression of polysialylated neural cell adhesion molecule in adult macaque Schwann cells promotes their migration and remyelination potential in the central nervous system. Brain, 2010. 133: p. 406- 420. 135. Lavdas, A.A., et al., Schwann cells genetically engineered to express PSA show enhanced migratory potential without impairment of their myelinating ability in vitro. Glia, 2006. 53(8): p. 868-878. 136. Eccleston, P.A., K.R. Jessen, and R. Mirsky, CONTROL OF PERIPHERAL GLIAL-CELL PROLIFERATION - A COMPARISON OF THE DIVISION RATES

169

OF ENTERIC GLIA AND SCHWANN-CELLS AND THEIR RESPONSE TO MITOGENS. Developmental Biology, 1987. 124(2): p. 409-417. 137. Jin, Y.-Q., et al., Efficient Schwann cell purification by differential cell detachment using multiplex collagenase treatment. Journal of Neuroscience Methods, 2008. 170(1): p. 140-148. 138. Singh, A.K., et al., Schwann cell culture from the adult animal sciatic nerve: Technique and review. Journal of Clinical Neuroscience, 1996. 3(1): p. 69-74. 139. Funk, D., C. Fricke, and B. Schlosshauer, Aging Schwann cells in vitro. European Journal of Cell Biology, 2007. 86(4): p. 207-219. 140. Langford, L.A., S. Porter, and R.P. Bunge, IMMORTALIZED RAT SCHWANN- CELLS PRODUCE TUMORS INVIVO. Journal of Neurocytology, 1988. 17(4): p. 521-529. 141. Joannides, A.J. and S. Chandran, Human embryonic stem cells: An experimental and therapeutic resource for neurological disease. Journal of the Neurological Sciences, 2008. 265(1-2): p. 84-88. 142. Rogers, C.D., S.A. Moody, and E.S. Casey, Neural Induction and Factors That Stabilize a Neural Fate. Birth Defects Research Part C-Embryo Today-Reviews, 2009. 87(3): p. 249-262. 143. Maden, M., Retinoic acid in the development, regeneration and maintenance of the nervous system. Nature Reviews Neuroscience, 2007. 8(10): p. 755-765. 144. Villanueva, S., et al., Posteriorization by FGF, Wnt, and retinoic acid is required for neural crest induction. Developmental Biology, 2002. 241(2): p. 289-301. 145. Jessell, T.M., Neuronal specification in the spinal cord: Inductive signals and transcriptional codes. Nature Reviews Genetics, 2000. 1(1): p. 20-29. 146. Liem, K.F., T.M. Jessell, and J. Briscoe, Regulation of the neural patterning activity of sonic hedgehog by secreted BMP inhibitors expressed by notochord and somites. Development, 2000. 127(22): p. 4855-4866. 147. Delfino-Machin, M., et al., The proliferating field of neural crest stem cells. Developmental Dynamics, 2007. 236(12): p. 3242-3254. 148. Le Douarin, N.M., G.W. Calloni, and E. Dupin, The stem cells of the neural crest. Cell Cycle, 2008. 7(8): p. 1013-1019. 149. Nagoshi, N., et al., Neural Crest-Derived Stem Cells Display a Wide Variety of Characteristics. Journal of Cellular Biochemistry, 2009. 107(6): p. 1046-1052. 150. Li, X.J., et al., Specification of motoneurons from human embryonic stem cells. Nature Biotechnology, 2005. 23(2): p. 215-221. 151. Li, X.J., et al., Directed differentiation of ventral spinal progenitors and motor neurons from human embryonic stem cells by small molecules. Stem Cells, 2008. 26(4): p. 886-893. 152. Wichterle, H., et al., Directed differentiation of embryonic stem cells into motor neurons. Cell, 2002. 110(3): p. 385-397. 153. Bain, G., et al., Retinoic acid promotes neural and represses mesodermal gene expression in mouse embryonic stem cells in culture. Biochemical and Biophysical Research Communications, 1996. 223(3): p. 691-694.

170

154. Bibel, M., et al., Differentiation of mouse embryonic stem cells into a defined neuronal lineage. Nature Neuroscience, 2004. 7(9): p. 1003-1009. 155. Freude, K.K., et al., Soluble Amyloid Precursor Protein Induces Rapid Neural Differentiation of Human Embryonic Stem Cells. Journal of Biological Chemistry, 2011. 286(27): p. 24264-24274. 156. Boido, M., et al., Embryonic and adult stem cells promote raphespinal axon outgrowth and improve functional outcome following spinal hemisection in mice. European Journal of Neuroscience, 2009. 30(5): p. 833-846. 157. Perrin, F.E., et al., Grafted Human Embryonic Progenitors Expressing Neurogenin-2 Stimulate Axonal Sprouting and Improve Motor Recovery after Severe Spinal Cord Injury. Plos One, 2010. 5(12): p. 7. 158. Takahashi, K. and S. Yamanaka, Induction of pluripotent stem cells from mouse embryonic and adult fibroblast cultures by defined factors. Cell, 2006. 126(4): p. 663-676. 159. Kim, J.B., et al., Pluripotent stem cells induced from adult neural stem cells by reprogramming with two factors. Nature, 2008. 454(7204): p. 646-U54. 160. Schatzlein, A.G., Non-viral vectors in cancer gene therapy: principles and progress. Anti-Cancer Drugs, 2001. 12(4): p. 275-304. 161. Kaji, K., et al., Virus-free induction of pluripotency and subsequent excision of reprogramming factors. Nature, 2009. 458(7239): p. 771-U112. 162. Lee, C.H., et al., The generation of iPS cells using non-viral magnetic nanoparticlebased transfection. Biomaterials, 2011. 32(28): p. 6683-91. 163. Montserrat, N., et al., Simple Generation of Human Induced Pluripotent Stem Cells Using Poly-beta-amino Esters As the Non-viral Gene Delivery System. Journal of Biological Chemistry, 2011. 286(14). 164. Okita, K., et al., Generation of Mouse Induced Pluripotent Stem Cells Without Viral Vectors. Science, 2008. 322(5903): p. 949-953. 165. Khalil, I.A., et al., Uptake pathways and subsequent intracellular trafficking in nonviral gene delivery. Pharmacological Reviews, 2006. 58(1): p. 32-45. 166. Ma, H. and S.L. Diamond, Nonviral gene therapy and its delivery systems. Current Pharmaceutical Biotechnology, 2001. 2(1): p. 1-17. 167. Malgrange, B., et al., Using human pluripotent stem cells to untangle neurodegenerative disease mechanisms. Cellular and Molecular Life Sciences, 2011. 68(4): p. 635-649. 168. Dimos, J.T., et al., Induced pluripotent stem cells generated from patients with ALS can be differentiated into motor neurons. Science, 2008. 321(5893): p. 1218- 1221. 169. Park, I.H., et al., Disease-specific induced pluripotent stem cells. Cell, 2008. 134(5): p. 877-886. 170. Soldner, F., et al., Parkinson's Disease Patient-Derived Induced Pluripotent Stem Cells Free of Viral Reprogramming Factors. Cell, 2009. 136(5): p. 964-977. 171. Weiss, S., et al., Multipotent CNS stem cells are present in the adult mammalian spinal cord and ventricular neuroaxis. Journal of Neuroscience, 1996. 16(23): p. 7599-7609.

171

172. Lois, C. and A. Alvarezbuylla, Proliferating subventricular zone cells in the adult mammalian forebrain can differentiate into neurons and glia. Proceedings of the National Academy of Sciences of the United States of America, 1993. 90(5): p. 2074-2077. 173. Gage, F.H., et al., Multipotent progenitor cells in the adult dentate gyrus. Journal of Neurobiology, 1998. 36(2): p. 249-266. 174. Altman, J. and G.D. Das, AUTORADIOGRAPHIC AND HISTOLOGICAL EVIDENCE OF POSTNATAL HIPPOCAMPAL NEUROGENESIS IN RATS. Journal of Comparative Neurology, 1965. 124(3): p. 319-&. 175. Eriksson, P.S., et al., Neurogenesis in the adult human hippocampus. Nature Medicine, 1998. 4(11): p. 1313-1317. 176. Gould, E., et al., Hippocampal neurogenesis in adult Old World primates. Proceedings of the National Academy of Sciences of the United States of America, 1999. 96(9): p. 5263-5267. 177. Kornack, D.R. and P. Rakic, Continuation of neurogenesis in the hippocampus of the adult macaque monkey. Proceedings of the National Academy of Sciences of the United States of America, 1999. 96(10): p. 5768-5773. 178. Jankovski, A. and C. Sotelo, Subventricular zone olfactory bulb migratory pathway in the adult mouse: Cellular composition and specificity as determined by heterochronic and heterotopic transplantation. Journal of Comparative Neurology, 1996. 371(3): p. 376-396. 179. Nissant, A. and M. Pallotto, Integration and maturation of newborn neurons in the adult olfactory bulb - from synapses to function. European Journal of Neuroscience, 2011. 33(6): p. 1069-1077. 180. Pignatelli, A., C. Gambardella, and O. Belluzzi, Neurogenesis in the adult olfactory bulb. Neural Regeneration Research, 2011. 6(8): p. 575-600. 181. Morshead, C.M. and D. Vanderkooy, Postmitotic death is the fate of constitutively proliferating cells in the subependymal layer of the adult-mouse brain. Journal of Neuroscience, 1992. 12(1): p. 249-256. 182. Suhonen, J.O., et al., Differentiation of adult hippocampus-derived progenitors into olfactory neurons in vivo. Nature, 1996. 383(6601): p. 624-627. 183. Nomura, H., et al., Extramedullary chitosan channels promote survival of transplanted neural stem and progenitor cells and create a tissue bridge after complete spinal cord transection. Tissue Engineering Part A, 2008. 14(5): p. 649- 665. 184. Rice, A.C., et al., Proliferation and neuronal differentiation of mitotically active cells following traumatic brain injury. Experimental Neurology, 2003. 183(2): p. 406-417. 185. Bye, N., et al., Neurogenesis and Glial Proliferation Are Stimulated Following Diffuse Traumatic Brain Injury in Adult Rats. Journal of Neuroscience Research, 2011. 89(7): p. 986-1000. 186. Nait-Oumesmar, B., et al., Progenitor cells of the adult mouse subventricular zone proliferate, migrate and differentiate into oligodendrocytes after demyelination. European Journal of Neuroscience, 1999. 11(12): p. 4357-4366.

172

187. Kulbatski, I., et al., Endogenous and exogenous CNS derived stem/progenitor cell approaches for neurotrauma. Current Drug Targets, 2005. 6(1): p. 111-126. 188. Aberg, M.A.I., et al., Peripheral infusion of IGF-I selectively induces neurogenesis in the adult rat hippocampus. Journal of Neuroscience, 2000. 20(8): p. 2896-2903. 189. Benraiss, A., et al., Adenoviral brain-derived neurotrophic factor induces both neostriatal and olfactory neuronal recruitment from endogenous progenitor cells in the adult forebrain. Journal of Neuroscience, 2001. 21(17): p. 6718-6731. 190. Bai, Y., et al., Ectopic expression of angiopoietin-1 promotes neuronal differentiation in neural progenitor cells through the Akt pathway. Biochemical and Biophysical Research Communications, 2009. 378(2): p. 296-301. 191. Bath, K.G. and F.S. Lee, Neurotrophic Factor Control of Adult SVZ Neurogenesis. Developmental Neurobiology, 2010. 70(5): p. 339-349. 192. Rosa, A.I., et al., The Angiogenic Factor Angiopoietin-1 Is a Proneurogenic Peptide on Subventricular Zone Stem/Progenitor Cells. Journal of Neuroscience, 2010. 30(13): p. 4573-4584. 193. Wong, G., Y. Goldshmit, and A.M. Turnley, Interferon-gamma but not TNF alpha promotes neuronal differentiation and neurite outgrowth of murine adult neural stem cells. Experimental Neurology, 2004. 187(1): p. 171-177. 194. Zahir, T., et al., Neural Stem/Progenitor Cells Differentiate In Vitro to Neurons by the Combined Action of Dibutyryl cAMP and Interferon-gamma. Stem Cells and Development, 2009. 18(10): p. 1423-1432. 195. Arsenijevic, Y. and S. Weiss, Insulin-like growth factor-I is a differentiation factor for postmitotic CNS stem cell-derived neuronal precursors: Distinct actions from those of brain-derived neurotrophic factor. Journal of Neuroscience, 1998. 18(6): p. 2118-2128. 196. Arsenijevic, Y., et al., Insulin-like growth factor-1 is necessary for neural stem cell proliferation and demonstrates distinct actions of epidermal growth factor and fibroblast growth factor-2. Journal of Neuroscience, 2001. 21(18): p. 7194- 7202. 197. Kim, S.J., et al., Interferon-gamma promotes differentiation of neural progenitor cells via the JNK pathway. Neurochemical Research, 2007. 32(8): p. 1399-1406. 198. Adrian, E.K., Jr. and B.E. Walker, Incorporation of thymidine-H3 by cells in normal and injured mouse spinal cord. Journal of neuropathology and experimental neurology, 1962. 21: p. 597-609. 199. Matthews, M.A., M.F. Stonge, and C.L. Faciane, ELECTRON-MICROSCOPIC ANALYSIS OF ABNORMAL EPENDYMAL CELL-PROLIFERATION AND ENVELOPMENT OF SPROUTING AXONS FOLLOWING SPINAL-CORD TRANSECTION IN THE RAT. Acta Neuropathologica, 1979. 45(1): p. 27-36. 200. Meletis, K., et al., Spinal cord injury reveals multilineage differentiation of ependymal cells. Plos Biology, 2008. 6(7): p. 1494-1507. 201. Rakic, P. and R.L. Sidman, SUBCOMMISSURAL ORGAN AND ADJACENT EPENDYMA - AUTORADIOGRAPHIC STUDY OF THEIR ORIGIN IN MOUSE BRAIN. American Journal of Anatomy, 1968. 122(2): p. 317-&.

173

202. Attar, A., et al., Electron microscopic study of the progeny of ependymal stem cells in the normal and injured spinal cord. Surgical Neurology, 2005. 64: p. 28- 32. 203. Kojima, A. and C.H. Tator, Intrathecal administration of epidermal growth factor and fibroblast growth factor 2 promotes ependymal proliferation and functional recovery after spinal cord injury in adult rats. Journal of Neurotrauma, 2002. 19(2): p. 223-238. 204. Ayuso-Sacido, A., et al., Long-term expansion of adult human brain subventricular zone precursors. Neurosurgery, 2008. 62(1): p. 223-229. 205. Kuhn, H.G., H. DickinsonAnson, and F.H. Gage, Neurogenesis in the dentate gyrus of the adult rat: Age-related decrease of neuronal progenitor proliferation. Journal of Neuroscience, 1996. 16(6): p. 2027-2033. 206. Barry, F.P. and J.M. Murphy, Mesenchymal stem cells: clinical applications and biological characterization. International Journal of Biochemistry & Cell Biology, 2004. 36(4): p. 568-584. 207. Alexanian, A.R., An efficient method for generation of neural-like cells from adult human bone marrow-derived mesenchymal stem cells. Regenerative Medicine, 2010. 5(6): p. 891-900. 208. Galmiche, M.C., et al., STROMAL CELLS FROM HUMAN LONG-TERM MARROW CULTURES ARE MESENCHYMAL CELLS THAT DIFFERENTIATE FOLLOWING A VASCULAR SMOOTH-MUSCLE DIFFERENTIATION PATHWAY. Blood, 1993. 82(1): p. 66-76. 209. Jackson, K.A., et al., Regeneration of ischemic cardiac muscle and vascular endothelium by adult stem cells. Journal of Clinical Investigation, 2001. 107(11): p. 1395-1402. 210. Theise, N.D., et al., Derivation of hepatocytes from bone marrow cells in mice after radiation-induced myeloablation. Hepatology, 2000. 31(1): p. 235-240. 211. Zimmet, J.M. and J.M. Hare, Emerging role for bone marrow derived mesenchymal stem cells in myocardial regenerative therapy. Basic Research in Cardiology, 2005. 100(6): p. 471-481. 212. Kashani, I.R., et al., Schwann-like cell differentiation from rat bone marrow stem cells. Archives of Medical Science, 2011. 7(1): p. 45-52. 213. Battula, V.L., et al., Human placenta and bone marrow derived MSC cultured in serum-free, b-FGF-containing medium express cell surface frizzled-9 and SSEA-4 and give rise to multilinelage differentiation. Differentiation, 2007. 75(4): p. 279- 291. 214. Datta, I., et al., Neuronal plasticity of human Wharton's jelly mesenchymal stromal cells to the dopaminergic cell type compared with human bone marrow mesenchymal stromal cells. Cytotherapy, 2011. 13(8): p. 918-32. 215. Lu, L.-L., et al., Isolation and characterization of human umbilical cord mesenchymal stem cells with hematopoiesis-supportive function and other potentials. Haematologica-the Hematology Journal, 2006. 91(8): p. 1017-1026.

174

216. Walker, P.A., et al., Progenitor cell therapy for the treatment of central nervous system injury: a review of the state of current clinical trials. Stem cells international, 2010. 2010: p. 369578. 217. Joyce, N., et al., Mesenchymal stem cells for the treatment of neurodegenerative disease. Regenerative Medicine, 2010. 5(6): p. 933-946. 218. Vaquero, J.V.J. and M. Zurita, Functional recovery after severe CNS trauma: Current perspectives for cell therapy with bone marrow stromal cells. Progress in Neurobiology, 2011. 93(3): p. 341-349. 219. Crigler, L., et al., Human mesenchymal stem cell subpopulations express a variety of neuro-regulatory molecules and promote neuronal cell survival and neuritogenesis. Experimental Neurology, 2006. 198(1): p. 54-64. 220. Ankeny, D.P., D.M. McTigue, and L.B. Jakeman, Bone marrow transplants provide tissue protection and directional guidance for axons after contusive spinal cord injury in rats. Experimental Neurology, 2004. 190(1): p. 17-31. 221. Akiyama, Y., C. Radtke, and J.D. Kocsis, Remyelination of the rat spinal cord by transplantation of identified bone marrow stromal cells. Journal of Neuroscience, 2002. 22(15): p. 6623-6630. 222. Inoue, M., et al., Comparative analysis of remyelinating potential of focal and intravenous administration of autologous bone marrow cells into the rat demyelinated spinal cord. Glia, 2003. 44(2): p. 111-118. 223. Rivera, F.J., et al., Mesenchymal Stem Cells Promote Oligodendroglial Differentiation in Hippocampal Slice Cultures. Cellular Physiology and Biochemistry, 2009. 24(3-4): p. 317-324. 224. Li, L.A.L.L.A., et al., Transplantation of Marrow Stromal Cells Restores Cerebral Blood Flow and Reduces Cerebral Atrophy in Rats with Traumatic Brain Injury: In vivo MRI Study. Journal of Neurotrauma, 2011. 28(4): p. 535- 545. 225. Gago, N., et al., Age-Dependent Depletion of Human Skin-Derived Progenitor Cells. Stem Cells, 2009. 27(5): p. 1164-1172. 226. Szebenyi, G., et al., Interstitial branches develop from active regions of the axon demarcated by the primary growth cone during pausing behaviors. J Neurosci, 1998. 18(19): p. 7930-40. 227. Dotti, C.G., C.A. Sullivan, and G.A. Banker, The establishment of polarity by hippocampal neurons in culture. J Neurosci, 1988. 8(4): p. 1454-68. 228. Wen, Z. and J.Q. Zheng, Directional guidance of nerve growth cones. Curr Opin Neurobiol, 2006. 16(1): p. 52-8. 229. Farrar, N.R. and G.E. Spencer, Pursuing a 'turning point' in growth cone research. Dev Biol, 2008. 318(1): p. 102-11. 230. Kalil, K., G. Szebenyi, and E.W. Dent, Common mechanisms underlying growth cone guidance and axon branching. J Neurobiol, 2000. 44(2): p. 145-58. 231. Suter, D.M. and K.E. Miller, The emerging role of forces in axonal elongation. Progress in Neurobiology, 2011. 94(2): p. 91-101. 232. Flanagan, L.A., et al., Neurite branching on deformable substrates. Neuroreport, 2002. 13(18): p. 2411-2415.

175

233. Gunn, J.W., S.D. Turner, and B.K. Mann, Adhesive and mechanical properties of hydrogels influence neurite extension. Journal of Biomedical Materials Research Part A, 2005. 72A(1): p. 91-97. 234. Marquardt, L. and R.K. Willits, Neurite growth in PEG gels: Effect of mechanical stiffness and laminin concentration. Journal of Biomedical Materials Research Part A, 2011. 98A(1): p. 1-6. 235. Sundararaghavan, H.G., et al., Neurite Growth in 3D Collagen Gels With Gradients of Mechanical Properties. Biotechnology and Bioengineering, 2009. 102(2): p. 632-643. 236. Willits, R.K. and S.L. Skornia, Effect of collagen gel stiffness on neurite extension. Journal of Biomaterials Science-Polymer Edition, 2004. 15(12): p. 1521-1531. 237. Leach, J.B., et al., Neurite outgrowth and branching of PC12 cells on very soft substrates sharply decreases below a threshold of substrate rigidity. Journal of Neural Engineering, 2007. 4(2): p. 26-34. 238. Pfister, B.J., et al., Extreme stretch growth of integrated axons. Journal of Neuroscience, 2004. 24(36): p. 7978-7983. 239. Pfister, B.J., et al., Development of transplantable nervous tissue constructs comprised of stretch-grown axons. Journal of Neuroscience Methods, 2006. 153(1): p. 95-103. 240. Smith, D.H., J.A. Wolf, and D.F. Meaney, A new strategy to produce sustained growth of central nervous system axons: Continuous mechanical tension. Tissue Engineering, 2001. 7(2): p. 131-139. 241. Bray, D., AXONAL GROWTH IN RESPONSE TO EXPERIMENTALLY APPLIED MECHANICAL TENSION. Developmental Biology, 1984. 102(2): p. 379-389. 242. Bueno, F.R. and S.B. Shah, Implications of tensile loading for the tissue engineering of nerves. Tissue Engineering Part B-Reviews, 2008. 14(3): p. 219- 233. 243. Lamoureux, P., et al., Growth and Elongation Within and Along the Axon. Developmental Neurobiology, 2010. 70(3): p. 135-149. 244. Pfister, B.J., et al., Stretch-grown axons retain the ability to transmit active electrical signals. Febs Letters, 2006. 580(14): p. 3525-3531. 245. Pfister, B.J., et al., Neural engineering to produce in vitro nerve constructs and neurointerface. Neurosurgery, 2007. 60(1): p. 137-141. 246. Pfister, B.P., et al., Engineering nerve constructs for clinical application. Journal of Neurotrauma, 2004. 21(9): p. P98. 247. Huang, J.H., et al., Long-Term Survival and Integration of Transplanted Engineered Nervous Tissue Constructs Promotes Peripheral Nerve Regeneration. Tissue Engineering Part A, 2009. 15(7): p. 1677-1685. 248. Iwata, A., et al., Long-term survival and outgrowth of mechanically engineered nervous tissue constructs implanted into spinal cord lesions. Tissue Engineering, 2006. 12(1): p. 101-110.

176

249. Hoffman-Kim, D., J.A. Mitchel, and R.V. Bellamkonda, Topography, Cell Response, and Nerve Regeneration, in Annual Review of Biomedical Engineering, Vol 12, M.L.D.J.S.G.M.L. Yarmush, Editor. 2010. p. 203-231. 250. Khan, S. and G. Newaz, A comprehensive review of surface modification for neural cell adhesion and patterning. Journal of Biomedical Materials Research Part A, 2010. 93A(3): p. 1209-1224. 251. Roach, P., et al., Surface strategies for control of neuronal cell adhesion: A review. Surface Science Reports, 2010. 65(6): p. 145-173. 252. Gomez, N., et al., Immobilized nerve growth factor and microtopography have distinct effects on polarization versus axon elongation in hippocampal cells in culture. Biomaterials, 2007. 28(2): p. 271-284. 253. Jang, M.J., et al., Directional neurite growth using carbon nanotube patterned substrates as a biomimetic cue. Nanotechnology, 2010. 21(23). 254. Li, J.M., H. McNally, and R. Shi, Enhanced neurite alignment on micro-patterned poly-L-lactic acid films. Journal of Biomedical Materials Research Part A, 2008. 87A(2): p. 392-404. 255. Goldner, J.S., et al., Neurite bridging across micropatterned grooves. Biomaterials, 2006. 27(3): p. 460-472. 256. Miller, C., S. Jeftinija, and S. Mallapragada, Synergistic effects of physical and chemical guidance cues on neurite alignment and outgrowth on biodegradable polymer substrates. Tissue Engineering, 2002. 8(3): p. 367-378. 257. Yu, T.T. and M.S. Shoichet, Guided cell adhesion and outgrowth in peptide- modified channels for neural tissue engineering. Biomaterials, 2005. 26(13): p. 1507-1514. 258. Gros, T., et al., Regeneration of long-tract axons through sites of spinal cord injury using templated agarose scaffolds. Biomaterials, 2010. 31(26): p. 6719- 6729. 259. Midha, R., et al., Growth factor enhancement of peripheral nerve regeneration through a novel synthetic hydrogel tube. J Neurosurg, 2003. 99(3): p. 555-65. 260. Xu, X.M., et al., Bridging Schwann cell transplants promote axonal regeneration from both the rostral and caudal stumps of transected adult rat spinal cord. J Neurocytol, 1997. 26(1): p. 1-16. 261. Yu, M., et al., Semiconductor nanomembrane tubes: three-dimensional confinement for controlled neurite outgrowth. ACS Nano, 2011. 5(4): p. 2447-57. 262. Elsdale, T. and J. Bard, Collagen substrata for studies on cell behavior. J Cell Biol, 1972. 54(3): p. 626-37. 263. Barnes, C.P., et al., Nanofiber technology: designing the next generation of tissue engineering scaffolds. Adv Drug Deliv Rev, 2007. 59(14): p. 1413-33. 264. Smith, L.A. and P.X. Ma, Nano-fibrous scaffolds for tissue engineering. Colloids Surf B Biointerfaces, 2004. 39(3): p. 125-31. 265. Yang, F., et al., Electrospinning of nano/micro scale poly(L-lactic acid) aligned fibers and their potential in neural tissue engineering. Biomaterials, 2005. 26(15): p. 2603-10.

177

266. Corey, J.M., et al., Aligned electrospun nanofibers specify the direction of dorsal root ganglia neurite growth. J Biomed Mater Res A, 2007. 83(3): p. 636-45. 267. Shaw, D. and M.S. Shoichet, Toward spinal cord injury repair strategies: peptide surface modification of expanded poly(tetrafluoroethylene) fibers for guided neurite outgrowth in vitro. J Craniofac Surg, 2003. 14(3): p. 308-16. 268. Wen, X. and P.A. Tresco, Effect of filament diameter and extracellular matrix molecule precoating on neurite outgrowth and Schwann cell behavior on multifilament entubulation bridging device in vitro. J Biomed Mater Res A, 2006. 76(3): p. 626-37. 269. Karlsson, M., F. Johansson, and M. Kanje, Polystyrene replicas of neuronal basal lamina act as excellent guides for regenerating neurites. Acta Biomaterialia, 2011. 7(7): p. 2910-2918. 270. Wang, D.Y., et al., Microcontact printing of laminin on oxygen plasma activated substrates for the alignment and growth of Schwann cells. J Biomed Mater Res B Appl Biomater, 2007. 80(2): p. 447-53. 271. Schmalenberg, K.E. and K.E. Uhrich, Micropatterned polymer substrates control alignment of proliferating Schwann cells to direct neuronal regeneration. Biomaterials, 2005. 26(12): p. 1423-30. 272. Kofron, C.M., et al., Neurite Outgrowth at the Biomimetic Interface. Annals of Biomedical Engineering, 2010. 38(6): p. 2210-2225. 273. Kofron, C.M. and D. Hoffman-Kim, Optimization by Response Surface Methodology of Confluent and Aligned Cellular Monolayers for Nerve Guidance. Cell Mol Bioeng, 2009. 2(4): p. 554-572. 274. Miner, J.H. and P.D. Yurchenco, Laminin functions in tissue morphogenesis. Annu Rev Cell Dev Biol, 2004. 20: p. 255-84. 275. Selak, I., J.M. Foidart, and G. Moonen, Laminin promotes cerebellar granule cells migration in vitro and is synthesized by cultured astrocytes. Dev Neurosci, 1985. 7(5-6): p. 278-85. 276. Liesi, P., et al., Domain-specific antibodies against the B2 chain of laminin inhibit neuronal migration in the neonatal rat cerebellum. J Neurosci Res, 1995. 40(2): p. 199-206. 277. Smalheiser, N.R. and N.B. Schwartz, Cranin: a laminin-binding protein of cell membranes. Proc Natl Acad Sci U S A, 1987. 84(18): p. 6457-61. 278. Smalheiser, N.R., Cranin interacts specifically with the sulfatide-binding domain of laminin. J Neurosci Res, 1993. 36(5): p. 528-38. 279. Goh, E.L., et al., beta1-integrin mediates myelin-associated glycoprotein signaling in neuronal growth cones. Mol Brain, 2008. 1: p. 10. 280. Plantman, S., et al., Integrin-laminin interactions controlling neurite outgrowth from adult DRG neurons in vitro. Mol Cell Neurosci, 2008. 39(1): p. 50-62. 281. Kim, S.H., J. Turnbull, and S. Guimond, Extracellular matrix and cell signalling: the dynamic cooperation of integrin, proteoglycan and growth factor receptor. J Endocrinol, 2011. 209(2): p. 139-51.

178

282. Wojcik-Stanaszek, L., A. Gregor, and T. Zalewska, Regulation of neurogenesis by extracellular matrix and integrins. Acta Neurobiol Exp (Wars), 2011. 71(1): p. 103-12. 283. Luo, B.H., C.V. Carman, and T.A. Springer, Structural basis of integrin regulation and signaling. Annu Rev Immunol, 2007. 25: p. 619-47. 284. Jacques, T.S., et al., Neural precursor cell chain migration and division are regulated through different beta1 integrins. Development, 1998. 125(16): p. 3167-77. 285. Li, S., et al., Matrix assembly, regulation, and survival functions of laminin and its receptors in embryonic stem cell differentiation. J Cell Biol, 2002. 157(7): p. 1279-90. 286. Varnum-Finney, B. and L.F. Reichardt, Vinculin-deficient PC12 cell lines extend unstable lamellipodia and filopodia and have a reduced rate of neurite outgrowth. J Cell Biol, 1994. 127(4): p. 1071-84. 287. Jay, D.G., A Src-astic response to mounting tension. J Cell Biol, 2001. 155(3): p. 327-30. 288. Woo, S. and T.M. Gomez, Rac1 and RhoA promote neurite outgrowth through formation and stabilization of growth cone point contacts. J Neurosci, 2006. 26(5): p. 1418-28. 289. Letourneau, P.C., M.L. Condic, and D.M. Snow, Interactions of developing neurons with the extracellular matrix. J Neurosci, 1994. 14(3 Pt 1): p. 915-28. 290. Reichardt, L.F. and K.J. Tomaselli, Extracellular matrix molecules and their receptors: functions in neural development. Annu Rev Neurosci, 1991. 14: p. 531-70. 291. Wang, L. and J.L. Denburg, A role for proteoglycans in the guidance of a subset of pioneer axons in cultured embryos of the cockroach. Neuron, 1992. 8(4): p. 701-14. 292. Walz, A., et al., Essential role of heparan sulfates in axon navigation and targeting in the developing visual system. Development, 1997. 124(12): p. 2421- 30. 293. Irie, A., et al., Specific heparan sulfate structures involved in retinal axon targeting. Development, 2002. 129(1): p. 61-70. 294. de Wit, J. and J. Verhaagen, Proteoglycans as modulators of axon guidance cue function. Semiaphorins: Receptor and Intracellular Signaling Mechanisms, 2007. 600: p. 73-89. 295. Tom, V.J., et al., Studies on the development and behavior of the dystrophic growth cone, the hallmark of regeneration failure, in an in vitro model of the glial scar and after spinal cord injury. J Neurosci, 2004. 24(29): p. 6531-9. 296. Kantor, D.B., et al., Semaphorin 5A is a bifunctional axon guidance cue regulated by heparan and chondroitin sulfate proteoglycans. Neuron, 2004. 44(6): p. 961- 75. 297. Li, G.N., J. Liu, and D. Hoffman-Kim, Multi-molecular gradients of permissive and inhibitory cues direct neurite outgrowth. Annals of Biomedical Engineering, 2008. 36(6): p. 889-904.

179

298. Jones, F.S. and P.L. Jones, The tenascin family of ECM glycoproteins: structure, function, and regulation during embryonic development and tissue remodeling. Dev Dyn, 2000. 218(2): p. 235-59. 299. Lang, D.M., et al., Tenascin-R and axon growth-promoting molecules are up- regulated in the regenerating visual pathway of the lizard (Gallotia galloti). Dev Neurobiol, 2008. 68(7): p. 899-916. 300. Lykissas, M.G., et al., The role of neurotrophins in axonal growth, guidance, and regeneration. Current Neurovascular Research, 2007. 4(2): p. 143-151. 301. TessierLavigne, M. and C.S. Goodman, The molecular biology of axon guidance. Science, 1996. 274(5290): p. 1123-1133. 302. Chilton, J.K., Molecular mechanisms of axon guidance. Dev Biol, 2006. 292(1): p. 13-24. 303. Huang, E.J. and L.F. Reichardt, Neurotrophins: roles in neuronal development and function. Annu Rev Neurosci, 2001. 24: p. 677-736. 304. Hallbook, F., Evolution of the vertebrate neurotrophin and Trk receptor gene families. Curr Opin Neurobiol, 1999. 9(5): p. 616-21. 305. Levi-Montalcini, R., The nerve growth factor: thirty-five years later. Biosci Rep, 1987. 7(9): p. 681-99. 306. Cordon-Cardo, C., et al., The trk tyrosine protein kinase mediates the mitogenic properties of nerve growth factor and neurotrophin-3. Cell, 1991. 66(1): p. 173- 83. 307. Kaplan, D.R., et al., The trk proto-oncogene product: a signal transducing receptor for nerve growth factor. Science, 1991. 252(5005): p. 554-8. 308. Klein, R., et al., The trk proto-oncogene encodes a receptor for nerve growth factor. Cell, 1991. 65(1): p. 189-97. 309. Berkemeier, L.R., et al., Neurotrophin-5: a novel neurotrophic factor that activates trk and trkB. Neuron, 1991. 7(5): p. 857-66. 310. Ip, N.Y., et al., Mammalian neurotrophin-4: structure, chromosomal localization, tissue distribution, and receptor specificity. Proc Natl Acad Sci U S A, 1992. 89(7): p. 3060-4. 311. Lamballe, F., R. Klein, and M. Barbacid, trkC, a new member of the trk family of tyrosine protein kinases, is a receptor for neurotrophin-3. Cell, 1991. 66(5): p. 967-79. 312. Huang, E.J., et al., Expression of Trk receptors in the developing mouse trigeminal ganglion: in vivo evidence for NT-3 activation of TrkA and TrkB in addition to TrkC. Development, 1999. 126(10): p. 2191-203. 313. Rodriguez-Tebar, A., G. Dechant, and Y.A. Barde, Neurotrophins: structural relatedness and receptor interactions. Philos Trans R Soc Lond B Biol Sci, 1991. 331(1261): p. 255-8. 314. Barde, Y.A., D. Edgar, and H. Thoenen, Purification of a new neurotrophic factor from mammalian brain. EMBO J, 1982. 1(5): p. 549-53. 315. Leibrock, J., et al., Molecular cloning and expression of brain-derived neurotrophic factor. Nature, 1989. 341(6238): p. 149-52.

180

316. Carvalho, A.L., et al., Role of the brain-derived neurotrophic factor at glutamatergic synapses. Br J Pharmacol, 2007. 317. Hohn, A., et al., Identification and characterization of a novel member of the nerve growth factor/brain-derived neurotrophic factor family. Nature, 1990. 344(6264): p. 339-41. 318. Maisonpierre, P.C., et al., Neurotrophin-3: a neurotrophic factor related to NGF and BDNF. Science, 1990. 247(4949 Pt 1): p. 1446-51. 319. Rosenthal, A., et al., Primary structure and biological activity of a novel human neurotrophic factor. Neuron, 1990. 4(5): p. 767-73. 320. Tobias, C.A., et al., Delayed grafting of BDNF and NT-3 producing fibroblasts into the injured spinal cord stimulates sprouting, partially rescues axotomized red nucleus neurons from loss and atrophy, and provides limited regeneration. Exp Neurol, 2003. 184(1): p. 97-113. 321. Yuan, X.B., et al., Signalling and crosstalk of Rho GTPases in mediating axon guidance. Nat Cell Biol, 2003. 5(1): p. 38-45. 322. Tessier-Lavigne, M. and C.S. Goodman, The molecular biology of axon guidance. Science, 1996. 274(5290): p. 1123-33. 323. Hallbook, F., C.F. Ibanez, and H. Persson, Evolutionary studies of the nerve growth factor family reveal a novel member abundantly expressed in Xenopus ovary. Neuron, 1991. 6(5): p. 845-58. 324. Lewin, G.R. and Y.A. Barde, Physiology of the neurotrophins. Annu Rev Neurosci, 1996. 19: p. 289-317. 325. Lykissas, M.G., et al., The role of neurotrophins in axonal growth, guidance, and regeneration. Curr Neurovasc Res, 2007. 4(2): p. 143-51. 326. Blesch, A. and M.H. Tuszynski, Cellular GDNF delivery promotes growth of motor and dorsal column sensory axons after partial and complete spinal cord transections and induces remyelination. J Comp Neurol, 2003. 467(3): p. 403-17. 327. Jungnickel, J., et al., Faster nerve regeneration after sciatic nerve injury in mice over-expressing basic fibroblast growth factor. J Neurobiol, 2006. 66(9): p. 940- 8. 328. Sendtner, M., et al., Endogenous ciliary neurotrophic factor is a lesion factor for axotomized motoneurons in adult mice. J Neurosci, 1997. 17(18): p. 6999-7006. 329. Kelleher, M.O., et al., The use of ciliary neurotrophic factor to promote recovery after peripheral nerve injury by delivering it at the site of the cell body. Acta Neurochir (Wien), 2006. 148(1): p. 55-60; discussion 60-1. 330. McCallister, W.V., et al., Regeneration along intact nerves using nerve growth factor and ciliary neurotrophic factor. J Reconstr Microsurg, 2004. 20(6): p. 473- 81. 331. Green, N.M., et al., Advances in Protein Chemistry, 1975: p. 85-133. 332. Wegner, G.J., et al., Fabrication of histidine-tagged fusion protein arrays for surface plasmon resonance imaging studies of protein-protein and protein-DNA interactions. Anal Chem, 2003. 75(18): p. 4740-6. 333. Niemeyer, C.M., The developments of semisynthetic DNA-protein conjugates. Trends Biotechnol, 2002. 20(9): p. 395-401.

181

334. Hendrickson, E.R., et al., High sensitivity multianalyte immunoassay using covalent DNA-labeled antibodies and polymerase chain reaction. Nucleic Acids Res, 1995. 23(3): p. 522-9. 335. Rusmini, F., Z. Zhong, and J. Feijen, Protein immobilization strategies for protein biochips. Biomacromolecules, 2007. 8(6): p. 1775-89. 336. Ganesan, R.e.a., Multicomponent protein patterning of material surfaces. Journal of Materials Chemistry, 2010. 20: p. 7322-7331. 337. Leipzig, N.D., et al., Differentiation of neural stem cells in three-dimensional growth factor-immobilized chitosan hydrogel scaffolds. Biomaterials, 2011. 32(1): p. 57-64. 338. Kuhl, P.R. and L.G. Griffith-Cima, Tethered epidermal growth factor as a paradigm for growth factor-induced stimulation from the solid phase. Nat Med, 1996. 2(9): p. 1022-7. 339. Kapur, T.A. and M.S. Shoichet, Chemically-bound nerve growth factor for neural tissue engineering applications. J Biomater Sci Polym Ed, 2003. 14(4): p. 383-94. 340. Fan, V.H., et al., Tethered epidermal growth factor provides a survival advantage to mesenchymal stem cells. Stem Cells, 2007. 25(5): p. 1241-51. 341. Nakajima, M., et al., Combinatorial protein display for the cell-based screening of biomaterials that direct neural stem cell differentiation. Biomaterials, 2007. 28(6): p. 1048-60. 342. Alberti, K., et al., Functional immobilization of signaling proteins enables control of stem cell fate. Nat Methods, 2008. 5(7): p. 645-50. 343. Aizawa, Y., et al., The effect of immobilized platelet derived growth factor AA on neural stem/progenitor cell differentiation on cell-adhesive hydrogels. Biomaterials, 2008. 29(35): p. 4676-83. 344. Shen, Y.H., M.S. Shoichet, and M. Radisic, Vascular endothelial growth factor immobilized in collagen scaffold promotes penetration and proliferation of endothelial cells. Acta Biomater, 2008. 4(3): p. 477-89. 345. Wall, S.T., et al., Multivalency of Sonic hedgehog conjugated to linear polymer chains modulates protein potency. Bioconjug Chem, 2008. 19(4): p. 806-12. 346. Leipzig, N.D., et al., Functional immobilization of interferon-gamma induces neuronal differentiation of neural stem cells. J Biomed Mater Res A, 2010. 93(2): p. 625-33. 347. Kang, C.E., E.J. Gemeinhart, and R.A. Gemeinhart, Cellular alignment by grafted adhesion peptide surface density gradients. J Biomed Mater Res A, 2004. 71(3): p. 403-11. 348. Adams, D.N., et al., Growth cones turn and migrate up an immobilized gradient of the laminin IKVAV peptide. J Neurobiol, 2005. 62(1): p. 134-47. 349. Guarnieri, D., et al., Covalently immobilized RGD gradient on PEG hydrogel scaffold influences cell migration parameters. Acta Biomater, 2010. 6(7): p. 2532-9. 350. Bhangale, S.M., et al., Biologically active protein gradients via microstamping. Advanced Materials, 2005. 17(7): p. 809-+.

182

351. Yu, L.M., J.H. Wosnick, and M.S. Shoichet, Miniaturized system of neurotrophin patterning for guided regeneration. J Neurosci Methods, 2008. 171(2): p. 253-63. 352. Yu, L.M., F.D. Miller, and M.S. Shoichet, The use of immobilized neurotrophins to support neuron survival and guide nerve fiber growth in compartmentalized chambers. Biomaterials, 2010. 31(27): p. 6987-99. 353. Sorribas, H., C. Padeste, and L. Tiefenauer, Photolithographic generation of protein micropatterns for neuron culture applications. Biomaterials, 2002. 23(3): p. 893-900. 354. Luo, Y. and M.S. Shoichet, A photolabile hydrogel for guided three-dimensional cell growth and migration. Nat Mater, 2004. 3(4): p. 249-53. 355. Hoffmann, J.C. and J.L. West, Three-dimensional photolithographic patterning of multiple bioactive ligands in poly(ethylene glycol) hydrogels. Soft Matter, 2010. 6(20): p. 5056-5063. 356. Jeon, H., et al., Chemical Patterning of Ultrathin Polymer Films by Direct-Write Multiphoton Lithography. Journal of the American Chemical Society, 2011. 133(16): p. 6138-6141. 357. Wylie, R.G., et al., Spatially controlled simultaneous patterning of multiple growth factors in three-dimensional hydrogels. Nature Materials, 2011. 10(10): p. 799-806. 358. Hashi, H., et al., Angiogenic activity of a fusion protein of the cell-binding domain of fibronectin and basic fibroblast growth factor. Cell Struct Funct, 1994. 19(1): p. 37-47. 359. Andrades, J.A., et al., Engineering, expression, and renaturation of a collagen- targeted human bFGF fusion protein. Growth Factors, 2001. 18(4): p. 261-75. 360. Nishi, N., et al., Collagen-binding growth factors: Production and characterization of functional fusion proteins having a collagen-binding domain. Proceedings of the National Academy of Sciences of the United States of America, 1998. 95(12): p. 7018-7023. 361. Hayashi, M., M. Tomita, and K. Yoshizato, Production of EGF-collagen chimeric protein which shows the mitogenic activity. Biochimica Et Biophysica Acta- General Subjects, 2001. 1528(2-3): p. 187-195. 362. Lin, H., et al., The effect of collagen-targeting platelet-derived growth factor on cellularization and vascularization of collagen scaffolds. Biomaterials, 2006. 27(33): p. 5708-5714. 363. Ota, T., et al., A fusion protein of hepatocyte growth factor enhances reconstruction of myocardium in a cardiac patch derived from porcine urinary bladder matrix. Journal of Thoracic and Cardiovascular Surgery, 2008. 136(5): p. 1309-1317. 364. Sun, W., et al., Promotion of peripheral nerve growth by collagen scaffolds loaded with collagen-targeting human nerve growth factor-beta. J Biomed Mater Res A, 2007. 83(4): p. 1054-61. 365. McCaig, C.D., NERVE GUIDANCE - A ROLE FOR BIO-ELECTRIC FIELDS. Progress in Neurobiology, 1988. 30(6): p. 449-468.

183

366. Jaffe, L.F. and M.M. Poo, Neurites grow faster towards the cathode than the anode in a steady field. J Exp Zool, 1979. 209(1): p. 115-28. 367. Erskine, L. and C.D. McCaig, Growth cone neurotransmitter receptor activation modulates electric field-guided nerve growth. Dev Biol, 1995. 171(2): p. 330-9. 368. McCaig, C.D., Nerve branching is induced and oriented by a small applied electric field. J Cell Sci, 1990. 95 ( Pt 4): p. 605-15. 369. McCaig, C.D., L. Sangster, and R. Stewart, Neurotrophins enhance electric field- directed growth cone guidance and directed nerve branching. Dev Dyn, 2000. 217(3): p. 299-308. 370. Patel, N. and M.M. Poo, Orientation of neurite growth by extracellular electric fields. J Neurosci, 1982. 2(4): p. 483-96. 371. Wan, L.D., R. Xia, and W.L. Ding, Low-frequency electrical stimulation improves neurite outgrowth of dorsal root ganglion neurons in vitro via upregulating Ca(2+)-mediated brain-derived neurotrophic factor expression. Neural Regeneration Research, 2010. 5(16): p. 1256-1260. 372. Wood, M. and R.K. Willits, Short-duration, DC electrical stimulation increases chick embryo DRG neurite outgrowth. Bioelectromagnetics, 2006. 27(4): p. 328- 331. 373. Wood, M.D. and R.K. Willits, Applied electric field enhances DRG neurite growth: influence of stimulation media, surface coating and growth supplements. Journal of Neural Engineering, 2009. 6(4). 374. Rajnicek, A.M., K.R. Robinson, and C.D. McCaig, The direction of neurite growth in a weak DC electric field depends on the substratum: Contributions of adhesivity and net surface charge. Developmental Biology, 1998. 203(2): p. 412- 423. 375. Ariza, C.A., et al., The Influence of Electric Fields on Hippocampal Neural Progenitor Cells. Stem Cell Reviews and Reports, 2010. 6(4): p. 585-600. 376. Arocena, M., et al., A Time-Lapse and Quantitative Modelling Analysis of Neural Stem Cell Motion in the Absence of Directional Cues and in Electric Fields. Journal of Neuroscience Research, 2010. 88(15): p. 3267-3274. 377. Al-Majed, A.A., et al., Brief electrical stimulation promotes the speed and accuracy of motor axonal regeneration. J Neurosci, 2000. 20(7): p. 2602-8. 378. Sisken, B.F., et al., Stimulation of rat sciatic nerve regeneration with pulsed electromagnetic fields. Brain Res, 1989. 485(2): p. 309-16. 379. Langer, R. and J.P. Vacanti, TISSUE ENGINEERING. Science, 1993. 260(5110): p. 920-926. 380. Brodbelt, A.R. and M.A. Stoodley, Post-traumatic syringomyelia: a review. Journal of Clinical Neuroscience, 2003. 10(4): p. 401-408. 381. Stoodley, M.A., Pathophysiology of syringomyelia. Journal of Neurosurgery, 2000. 92(6): p. 1069-1070. 382. National Institutes of Health (U.S.) and National Institute of Neurological Disorders and Stroke (U.S.). Office of Communications and Public Liaison., Syringomyelia fact sheet, in NIH publication. National Institute of Neurological Disorders and Stroke, National Institutes of Health.: Bethesda, MD.

184

383. Milhorat, T.H., et al., Anatomical basis of syringomyelia occurring with hindbrain lesions. Neurosurgery, 1993. 32(5): p. 748-754. 384. Brickell, K.L., et al., Ethnic differences in syringomyelia in New Zealand. J Neurol Neurosurg Psychiatry, 2006. 77(8): p. 989–991 385. Sixt, C., et al., Evaluation of quality of life parameters in patients who have syringomyelia. Journal of Clinical Neuroscience, 2009. 16(12): p. 1599-1603. 386. Fehlings, M.G. and J.W. Austin, Posttraumatic syringomyelia. Journal of Neurosurgery-Spine, 2011. 14(5): p. 570-572. 387. Hu, Z. and J. Tu, The roads to mitochondrial dysfunction in a rat model of posttraumatic syringomyelia. BioMed research international, 2015. 2015: p. 831490-831490. 388. Austin, J.W., M. Afshar, and M.G. Fehlings, The Relationship between Localized Subarachnoid Inflammation and Parenchymal Pathophysiology after Spinal Cord Injury. Journal of Neurotrauma, 2012. 29(10): p. 1838-1849. 389. Brodbelt, A.R., et al., The role of excitotoxic injury in post-traumatic syringomyelia. Journal of Neurotrauma, 2003. 20(9): p. 883-893. 390. Yang, L.Q., et al., Excitotoxic model of post-traumatic syringomyelia in the rat. Spine, 2001. 26(17): p. 1842-1849. 391. Brodbelt, A.R., et al., Fluid flow in an animal model of post-traumatic syringomyelia. European Spine Journal, 2003. 12(3): p. 300-306. 392. Stoodley, M.A., et al., Mechanisms underlying the formation and enlargement of noncommunicating syringomyelia: experimental studies. Neurosurgical focus, 2000. 8(3): p. E2-E2. 393. Stoodley, M.A., et al., Arterial pulsation-dependent perivascular cerebrospinal fluid flow into the central canal in the sheep spinal cord. Journal of Neurosurgery, 1997. 86(4): p. 686-693. 394. Bilston, L.E., M.A. Stoodley, and D.F. Fletcher, The influence of the relative timing of arterial and subarachnoid space pulse waves on spinal perivascular cerebrospinal fluid flow as a possible factor in syrinx development Laboratory investigation. Journal of Neurosurgery, 2010. 112(4): p. 808-813. 395. Hemley, S.J., et al., Aquaporin-4 Expression in Post-Traumatic Syringomyelia. Journal of Neurotrauma, 2013. 30(16): p. 1457-1467. 396. Hemley, S.J., et al., Aquaporin-4 expression and blood spinal cord barrier permeability in canalicular syringomyelia Laboratory investigation. Journal of Neurosurgery-Spine, 2012. 17(6): p. 602-612. 397. Brodbelt, A.R., et al., Altered subarachnoid space compliance and fluid flow in an animal model of posttraumatic syringomyelia. Spine, 2003. 28(20): p. E413-E419. 398. Trapnell, C., L. Pachter, and S.L. Salzberg, TopHat: discovering splice junctions with RNA-Seq. Bioinformatics, 2009. 25(9): p. 1105-1111. 399. Trapnell, C., et al., Differential gene and transcript expression analysis of RNA- seq experiments with TopHat and Cufflinks. Nature Protocols, 2012. 7(3): p. 562- 578.

185

400. Trapnell, C., et al., Transcript assembly and quantification by RNA-Seq reveals unannotated transcripts and isoform switching during cell differentiation. Nature Biotechnology, 2010. 28(5): p. 511-U174. 401. Bligh, E.G. and W.J. Dyer, A rapid method of total lipid extraction and purification. Canadian journal of biochemistry and physiology, 1959. 37(8): p. 911-7. 402. Ogata, H., et al., KEGG: Kyoto Encyclopedia of Genes and Genomes. Nucleic Acids Research, 1999. 27(1): p. 29-34. 403. Frolkis, A., et al., SMPDB: The Small Molecule Pathway Database. Nucleic Acids Research, 2010. 38: p. D480-D487. 404. Smith, C.A., et al., METLIN - A metabolite mass spectral database. Therapeutic Drug Monitoring, 2005. 27(6): p. 747-751. 405. Wishart, D.S., et al., HMDB: the human metabolome database. Nucleic Acids Research, 2007. 35: p. D521-D526. 406. Shannon, P., et al., Cytoscape: A software environment for integrated models of biomolecular interaction networks. Genome Research, 2003. 13(11): p. 2498- 2504. 407. Tu, J., et al., Reaction of endogenous progenitor cells in a rat model of posttraumatic syringomyelia. Journal of Neurosurgery-Spine, 2011. 14(5): p. 573- 582. 408. Hu, X., et al., Microglial and macrophage polarization -new prospects for brain repair. Nature Reviews Neurology, 2015. 11(1): p. 56-64. 409. David, S. and A. Kroner, Repertoire of microglial and macrophage responses after spinal cord injury. Nature Reviews Neuroscience, 2011. 12(7): p. 388-399. 410. Novak, M.L. and T.J. Koh, Macrophage phenotypes during tissue repair. Journal of Leukocyte Biology, 2013. 93(6): p. 875-881. 411. Nielsen, S., et al., Specialized membrane domains for water transport in glial cells: High-resolution immunogold cytochemistry of aquaporin-4 in rat brain. Journal of Neuroscience, 1997. 17(1): p. 171-180. 412. Oenarto, J., et al., Expression of organic osmolyte transporters in cultured rat astrocytes and rat and human cerebral cortex. Archives of Biochemistry and Biophysics, 2014. 560: p. 59-72. 413. Zhou, Y., et al., The betaine-GABA transporter (BGT1, slc6a12) is predominantly expressed in the liver and at lower levels in the kidneys and at the brain surface. American Journal of Physiology-Renal Physiology, 2012. 302(3): p. F316-F328. 414. Tu, J., et al., Differentiation of Endogenous Progenitors in an Animal Model of Post-Traumatic Syringomyelia. Spine, 2010. 35(11): p. 1116-1121. 415. Wong, J., et al., Fluid Outflow in a Large-Animal Model of Posttraumatic Syringomyelia. Neurosurgery, 2012. 71(2): p. 474-480. 416. Seki, T. and M.G. Fehlings, Mechanistic insights into posttraumatic syringomyelia based on a novel in vivo animal model. Journal of Neurosurgery- Spine, 2008. 8(4): p. 365-375. 417. Kobayashi, S., et al., Experimental Syringohydromyelia Induced by Adhesive Arachnoiditis in the Rabbit: Changes in the Blood-Spinal Cord Barrier,

186

Neuroinflammatory Foci, and Syrinx Formation. Journal of Neurotrauma, 2012. 29(9): p. 1803-1816. 418. Takahashi, A., et al., Effect of cerebrospinal fluid shunting on experimental syringomyelia: Magnetic resonance imaging and histological findings. Neurologia Medico-Chirurgica, 1999. 39(9): p. 668-675. 419. Chakrabortty, S., et al., Experimental syringomyelia: Late ultrastructural changes of spinal cord tissue and magnetic resonance imaging evaluation. Surgical Neurology, 1997. 48(3): p. 246-254. 420. Dulin, J.N., et al., Transcriptomic Approaches to Neural Repair. Journal of Neuroscience, 2015. 35(41): p. 13860-13867. 421. Munro, K.M. and V.M. Perreau, Current and Future Applications of Transcriptomics for Discovery in CNS Disease and Injury. Neurosignals, 2009. 17(4): p. 311-327. 422. Chandran, V., et al., A Systems-Level Analysis of the Peripheral Nerve Intrinsic Axonal Growth Program. Neuron, 2016. 89(5): p. 956-970. 423. Duan, H.M., et al., Transcriptome analyses reveal molecular mechanisms underlying functional recovery after spinal cord injury. Proceedings of the National Academy of Sciences of the United States of America, 2015. 112(43): p. 13360-13365. 424. Smith, R., et al., Transcriptional profiling of intrinsic PNS factors in the postnatal mouse. Molecular and Cellular Neuroscience, 2011. 46(1): p. 32-44. 425. Stam, F.J., et al., Identification of candidate transcriptional modulators involved in successful regeneration after nerve injury. European Journal of Neuroscience, 2007. 25(12): p. 3629-3637. 426. Hemley, S.J., et al., Role of the blood-spinal cord-barrier in posttraumatic syringomyelia Laboratory investigation. Journal of Neurosurgery-Spine, 2009. 11(6): p. 696-704. 427. Mosser, D.M. and J.P. Edwards, Exploring the full spectrum of macrophage activation. Nature Reviews Immunology, 2008. 8(12): p. 958-969. 428. Gill, S.S., et al., Brain metabolites as 1H NMR markers of neuronal and glial disorders. NMR in biomedicine, 1989. 2(5-6): p. 196-200. 429. Govind, V., K. Young, and A.A. Maudsley, Proton NMR chemical shifts and coupling constants for brain metabolites (vol 13, pg 129, 2000). Nmr in Biomedicine, 2015. 28(7): p. 923-924. 430. Rauchova, H., et al., Hypoxia-induced lipid peroxidation in rat brain and protective effect of carnitine and phosphocreatine. Neurochemical Research, 2002. 27(9): p. 899-904. 431. Cohodarevic, T., A. Mailis, and W. Montanera, Syringomyelia: Pain, sensory abnormalities, and neuroimaging. Journal of Pain, 2000. 1(1): p. 54-66. 432. Lohle, P.N.M., et al., The pathogenesis of syringomyelia in spinal-cord ependymoma. Clinical Neurology and Neurosurgery, 1994. 96(4): p. 323-326. 433. Karadsheh, M.F., et al., Localization of the KCC4 potassium-chloride cotransporter in the nervous system. Neuroscience, 2004. 123(2): p. 381-391.

187

434. Pasantes-Morales, H. and S. Cruz-Rangel, Brain volume regulation: osmolytes and aquaporin perspectives. Neuroscience, 2010. 168(4): p. 871-884. 435. MacAulay, N. and T. Zeuthen, Water transport between CNS compartments: Contributions of aquaporins and cotransporters. Neuroscience, 2010. 168(4): p. 941-956. 436. den Daas, I., et al., Serial CSF sampling over a period of 30 h via an indwelling spinal catheter in healthy volunteers: headache, back pain, tolerability and measured acetylcholine profile. European Journal of Clinical Pharmacology, 2013. 69(5): p. 1083-1090. 437. Xu, F., T.Z. Li, and B.X. Zhang, An improved method for protecting and fixing the lumbar catheters placed in the spinal subarachnoid space of rats. Journal of Neuroscience Methods, 2009. 183(2): p. 114-118. 438. Jasmin, L. and P.T. Ohara, Long-term intrathecal catheterization in the rat. Journal of Neuroscience Methods, 2001. 110(1-2): p. 81-89. 439. Milligan, E.D., et al., A method for increasing the viability of the external portion of lumbar catheters placed in the spinal subarachnoid space of rats. Journal of Neuroscience Methods, 1999. 90(1): p. 81-86. 440. Storkson, R.V., et al., Lumbar catheterization of the spinal subarachnoid space in the rat. Journal of Neuroscience Methods, 1996. 65(2): p. 167-172. 441. Fawcett, J.W. and R.A. Asher, The glial scar and central nervous system repair. Brain Research Bulletin, 1999. 49(6): p. 377-391. 442. Fitch, M.T., et al., Cellular and molecular mechanisms of glial scarring and progressive cavitation: In vivo and in vitro analysis of inflammation-induced secondary injury after CNS trauma. Journal of Neuroscience, 1999. 19(19): p. 8182-8198. 443. Bovolenta, P., F. Wandosell, and M. Nietosampedro, CNS glial scar tissue- A source of molecules which inhibit central neurite outgrowth. Neuronal-Astrocytic Interactions: Implications for Normal and Pathological Cns Function, 1992. 94: p. 367-379. 444. Lamoureux, P., et al., A cytomechanical investigation of neurite growth on different culture surfaces. Journal of Cell Biology, 1992. 118(3): p. 655-661. 445. Kapur, T.A. and M.S. Shoichet, Immobilized concentration gradients of nerve growth factor guide neurite outgrowth. Journal of Biomedical Materials Research Part A, 2004. 68A(2): p. 235-243. 446. Xie, J.W., et al., Neurite Outgrowth on Nanofiber Scaffolds with Different Orders, Structures, and Surface Properties. Acs Nano, 2009. 3(5): p. 1151-1159. 447. Fereol, S., et al., Micropatterned ECM Substrates Reveal Complementary Contribution of Low and High Affinity Ligands to Neurite Outgrowth. Cytoskeleton, 2011. 68(7): p. 373-388. 448. Mukhatyar, V.J., et al., Role of fibronectin in topographical guidance of neurite extension on electrospun fibers. Biomaterials, 2011. 32(16): p. 3958-3968. 449. Wilkinson, A.E., A.M. McCormick, and N.D. Leipzig, Central Nervous System Tissue Engineering: Current Considerations and Strategies, in Central Nervous

188

System Tissue Engineering: Current Considerations and Strategies, K.A.a.J. Leach, Editor. 2011, Morgan & Claypool Publishers. p. 1-120. 450. McCormick, A.M. and N.D. Leipzig, Neural Regenerative Strategies Incorporating Biomolecular Axon Guidance Signals. Annals of Biomedical Engineering, 2012. 40(3): p. 578-597. 451. Ferrari, A., et al., Neuronal polarity selection by topography-induced focal adhesion control. Biomaterials, 2010. 31(17): p. 4682-4694. 452. Whittemore, S.R., et al., Mitogen and substrate differentially affect the lineage restriction of adult rat subventricular zone neural precursor cell populations. Experimental Cell Research, 1999. 252(1): p. 75-95. 453. Johansson, C.B., et al., Identification of a neural stem cell in the adult mammalian central nervous system. Cell, 1999. 96(1): p. 25-34. 454. Horne, M.K., et al., Three-Dimensional Nanofibrous Scaffolds Incorporating Immobilized BDNF Promote Proliferation and Differentiation of Cortical Neural Stem Cells. Stem Cells and Development, 2010. 19(6): p. 843-852. 455. Seidenfaden, R., et al., Glial conversion of SVZ-derived committed neuronal precursors after ectopic grafting into the adult brain. Molecular and Cellular Neuroscience, 2006. 32(1-2): p. 187-198. 456. Campos, L.S., et al., beta 1 integrins activate a MAPK signalling pathway in neural stem cells that contributes to their maintenance. Development, 2004. 131(14): p. 3433-3444. 457. Scadden, D.T., The stem-cell niche as an entity of action. Nature, 2006. 441(7097): p. 1075-1079. 458. Pierret, C., et al., Developmental cues and persistent neurogenic potential within an in vitro neural niche. Bmc Developmental Biology, 2010. 10. 459. Reilly, G.C. and A.J. Engler, Intrinsic extracellular matrix properties regulate stem cell differentiation. Journal of Biomechanics, 2010. 43(1): p. 55-62. 460. Stabenfeldt, S.E., et al., Biomimetic Microenvironment Modulates Neural Stem Cell Survival, Migration, and Differentiation. Tissue Engineering Part A, 2010. 16(12): p. 3747-3758. 461. Deister, C., S. Aljabari, and C.E. Schmidt, Effects of collagen 1, fibronectin, laminin and hyaluronic acid concentration in multi-component gels on neurite extension. Journal of Biomaterials Science-Polymer Edition, 2007. 18(8): p. 983- 997. 462. Hiraoka, M., et al., Enhanced Survival of Neural Cells Embedded in Hydrogels Composed of Collagen and Laminin-Derived Cell Adhesive Peptide. Bioconjugate Chemistry, 2009. 20(5): p. 976-983. 463. Li, H., A. Wijekoon, and N.D. Leipzig, 3D Differentiation of Neural Stem Cells in Macroporous Photopolymerizable Hydrogel Scaffolds. Plos One, 2012. 7(11): p. 11. 464. Hermanson, G.T., Bioconjugate techniques. Bioconjugate techniques, 1996: p. xxvii+785p.

189

465. Begum, A.A., R. Radhakrishnan, and K.P. Nazeer, Study of Structure-Property Relationship on Sulfuric Acid Crosslinked Chitosan Membranes. Malaysian Polymer Journal, 2011. 6(1): p. 27-38. 466. Radhakumary, C., et al., Biopolymer composite of Chitosan and methyl methacrylate for medical applications. Trends in Biomaterials and Artificial Organs, 2005. 18(2): p. 117-124. 467. Krishnamoorthy, G., et al., Development of D-lysine-assisted 1-ethyl-3-(3- dimethylaminopropyl)-carbodiimide/N-hydroxysuccinimide-init iated cross linking of collagen matrix for design of scaffold. Journal of Biomedical Materials Research Part A, 2013. 101A(4): p. 1173-1183. 468. Saha, K., et al., Substrate Modulus Directs Neural Stem Cell Behavior. Biophysical Journal, 2008. 95(9): p. 4426-4438. 469. Georges-Labouesse, E., et al., Essential role of alpha 6 integrins in cortical and retinal lamination. Current Biology, 1998. 8(17): p. 983-986. 470. Hall, P.E., et al., Laminin enhances the growth of human neural stem cells in defined culture media. Bmc Neuroscience, 2008. 9: p. 10. 471. Flanagan, L.A., et al., Regulation of human neural precursor cells by laminin and integrins. Journal of Neuroscience Research, 2006. 83(5): p. 845-856. 472. Engler, A., et al., Substrate compliance versus ligand density in cell on gel responses. Biophysical Journal, 2004. 86(1): p. 617-628. 473. Bach, C.T.T., et al., Tropomyosin isoform modulation of focal adhesion structure and cell migration. Cell Adhesion & Migration, 2010. 4(2): p. 226-234. 474. Vicente-Manzanares, M., et al., Non-muscle myosin II takes centre stage in cell adhesion and migration. Nature Reviews Molecular Cell Biology, 2009. 10(11): p. 778-790. 475. Wylie, S.R. and P.D. Chantler, Separate but linked functions of conventional myosins modulate adhesion and neurite outgrowth. Nature Cell Biology, 2001. 3(1): p. 88-92. 476. Ma, W., et al., Cell-extracellular matrix interactions regulate neural differentiation of human embryonic stem cells. Bmc Developmental Biology, 2008. 8: p. 13. 477. Goetz, A.K., et al., Temporally restricted substrate interactions direct fate and specification of neural precursors derived from embryonic stem cells. Proceedings of the National Academy of Sciences of the United States of America, 2006. 103(29): p. 11063-11068. 478. Balgude, A.P., et al., Agarose gel stiffness determines rate of DRG neurite extension in 3D cultures. Biomaterials, 2001. 22(10): p. 1077-1084. 479. Curley, J.L. and M.J. Moore, Facile micropatterning of dual hydrogel systems for 3D models of neurite outgrowth. Journal of Biomedical Materials Research Part A, 2011. 99A(4): p. 532-543. 480. Tate, M.C., et al., Specific beta(1) integrins mediate adhesion, migration, and differentiation of neural progenitors derived from the embryonic striatum. Molecular and Cellular Neuroscience, 2004. 27(1): p. 22-31.

190

481. Zhang, N., et al., Inflammation & apoptosis in spinal cord injury. Indian Journal of Medical Research, 2012. 135(3): p. 287-296. 482. Hausmann, O., Post-traumatic inflammation following spinal cord injury. Spinal Cord, 2003. 41(7): p. 369-378. 483. Klekamp, J., The pathophysiology of syringomyelia - Historical overview and current concept. Acta Neurochirurgica, 2002. 144(7): p. 649-664. 484. Macaya, D. and M. Spector, Injectable hydrogel materials for spinal cord regeneration: a review. Biomedical Materials, 2012. 7(1). 485. Pakulska, M., B. Ballios, and M. Shoichet, Injectable hydrogels for central nervous system therapy. Biomedical Materials, 2012. 7(2). 486. des Rieux, A., et al., Vascular endothelial growth factor-loaded injectable hydrogel enhances plasticity in the injured spinal cord. Journal of Biomedical Materials Research Part A, 2014. 102(7): p. 2345-2355. 487. Li, H., et al., A Hydrogel Bridge Incorporating Immobilized Growth Factors and Neural Stem/Progenitor Cells to Treat Spinal Cord Injury. Adv Healthc Mater, 2016. 5(7): p. 802-12. 488. Li, H., et al., In vivo assessment of guided neural stem cell differentiation in growth factor immobilized chitosan-based hydrogel scaffolds. Biomaterials, 2014. 35(33): p. 9049-9057. 489. Park, J., et al., Nerve regeneration following spinal cord injury using matrix metalloproteinase-sensitive, hyaluronic acid-based biomimetic hydrogel scaffold containing brain-derived neurotrophic factor. Journal of Biomedical Materials Research Part A, 2010. 93A(3): p. 1091-1099. 490. Fuhrmann, T., et al., Click-crosslinked injectable hyaluronic acid hydrogel is safe and biocompatible in the intrathecal space for ultimate use in regenerative strategies of the injured spinal cord. Methods, 2015. 84: p. 60-69. 491. Stabenfeldt, S.E. and M.C. LaPlaca, Variations in rigidity and ligand density influence neuronal response in methylcellulose-laminin hydrogels. Acta Biomaterialia, 2011. 7(12): p. 4102-4108. 492. Austin, J.W., et al., The effects of intrathecal injection of a hyaluronan-based hydrogel on inflammation, scarring and neurobehavioural outcomes in a rat model of severe spinal cord injury associated with arachnoiditis. Biomaterials, 2012. 33(18): p. 4555-4564. 493. Elliott Donaghue, I., C.H. Tator, and M.S. Shoichet, Sustained delivery of bioactive neurotrophin-3 to the injured spinal cord. Biomaterials Science, 2015. 3(1): p. 65-72. 494. Kast, C.E. and A. Bernkop-Schnurch, Thiolated polymers - thiomers: development and in vitro evaluation of chitosan-thioglycolic acid conjugates. Biomaterials, 2001. 22(17): p. 2345-2352. 495. Hornof, M.D., C.E. Kast, and A. Bernkop-Schnurch, In vitro evaluation of the viscoelastic properties of chitosan-thioglycolic acid conjugates. European Journal of Pharmaceutics and Biopharmaceutics, 2003. 55(2): p. 185-190.

191

496. Kast, C., et al., Chitosan-thioglycolic acid conjugate: a new scaffold material for tissue engineering? International Journal of Pharmaceutics, 2003. 256(1-2): p. 183-189. 497. Nair, D., et al., The Thiol-Michael Addition Click Reaction: A Powerful and Widely Used Tool in Materials Chemistry. Chemistry of Materials, 2014. 26(1): p. 724-744. 498. Phelps, E.A., et al., Maleimide Cross-Linked Bioactive PEG Hydrogel Exhibits Improved Reaction Kinetics and Cross-Linking for Cell Encapsulation and In Situ Delivery. Advanced Materials, 2012. 24(1): p. 64-+. 499. Darling, N., et al., Controlling the kinetics of thiol-maleimide Michael-type addition gelation kinetics for the generation of homogenous poly(ethylene glycol) hydrogels. Biomaterials, 2016. 101: p. 199-206. 500. Bernkop-Schnurch, A., M. Hornof, and T. Zoidl, Thiolated polymers-thiomers: synthesis and in vitro evaluation of chitosan-2-iminothiolane conjugates. International Journal of Pharmaceutics, 2003. 260(2): p. 229-237. 501. Lee, F., J.E. Chung, and M. Kurisawa, An injectable hyaluronic acid-tyramine hydrogel system for protein delivery. Journal of Controlled Release, 2009. 134(3): p. 186-193. 502. Fan, M., et al., Cytocompatible in situ forming chitosan/hyaluronan hydrogels via a metal-free click chemistry for soft tissue engineering. Acta Biomaterialia, 2015. 20: p. 60-68. 503. Truong, V.X., et al., In situ-forming robust chitosan-poly(ethylene glycol) hydrogels prepared by copper-free azide-alkyne click reaction for tissue engineering. Biomaterials Science, 2014. 2(2): p. 167-175. 504. Cheng, S., E. Clarke, and L. Bilston, Rheological properties of the tissues of the central nervous system: A review. Medical Engineering & Physics, 2008. 30(10): p. 1318-1337. 505. Koser, D., et al., CNS Cell Distribution and Axon Orientation Determine Local Spinal Cord Mechanical Properties. Biophysical Journal, 2015. 108(9): p. 2137- 2147. 506. Burda, J.E. and M.V. Sofroniew, Reactive Gliosis and the Multicellular Response to CNS Damage and Disease. Neuron, 2014. 81(2): p. 229-248. 507. Che, Y.J., et al., In Situ Gel Delivery System of Methylprednisolone for Post Traumatic Spinal Injuries. Journal of Biomaterials and Tissue Engineering, 2015. 5(7): p. 552-556. 508. Kang, C.E., et al., A New Paradigm for Local and Sustained Release of Therapeutic Molecules to the Injured Spinal Cord for Neuroprotection and Tissue Repair. Tissue Engineering Part A, 2009. 15(3): p. 595-604. 509. Holland-Nell, K., et al., Specifically immobilised aldo/keto reductase AKR1A1 shows a dramatic increase in activity relative to the randomly immobilised enzyme. Chembiochem, 2007. 8(9): p. 1071-1076.

192

APPENDICES

193

APPENDIX A

Supporting Information for Chapter 3

Supplementary Figure 3.1. Chemical standards for several important metabolites were

purchased (if available) and examined by mass spectrometry. Each graph shows the MS2

data above from samples with the standard fragmentation extending below the x-axis in gray for comparison of identifying peaks.

194

Supplementary Figure 3.2. Network displaying log2(fold change) values of EXC group vs

SHAM group at week three (center) and six (border). Gray color indicates no statistical significance between groups (p<0.01).

195

Supplemental Table 3.1

Week 3 CTL vs EXC Transcripts Gene ID Gene Symbol Gene Name Log2(fold p-value change) NM_001025276 RGD1305807 hypothetical LOC298077 inf 5.00E- 05 NM_001002280 Mrgprb1 MAS-related GPR, member inf 5.00E- X2 05 NM_030584 Sost sclerosteosis inf 5.00E- 05 NM_053365 Fabp4 fatty acid binding protein 4, 7.1626 0.0076 adipocyte 5 NM_001107702 Msr2 Fc receptor-like S, scavenger 6.76429 5.00E- receptor 05 NM_001013894 Gp49b leukocyte immunoglobulin- 6.64026 0.0002 like receptor, subfamily B, member 4; similar to GP49B1 NM_022179 Hk3 hexokinase 3 (white cell) 6.35621 0.0003 NM_133298 Gpnmb glycoprotein (transmembrane) 6.27534 5.00E- nmb 05 NM_198746 Klra5 killer cell lectin-like receptor, 5.90322 0.0037 subfamily A, member 5 5 NM_173136 Akr1b8 aldo-keto reductase family 1, 5.70272 0.0044 member B8 NM_001168285 RGD1559482 similar to immunoglobulin 5.38082 5.00E- superfamily, member 7 05 NM_001031638 Cd68 Cd68 molecule 5.31024 5.00E- 05 NM_001106314 Ifitm1 interferon induced 5.22687 5.00E- transmembrane protein 1 05 NM_031538 Cd8a CD8a molecule 5.21901 5.00E- 05 NM_001107503 Cd22 CD22 molecule 5.15674 5.00E- 05 NM_001173386 Clec7a C-type lectin domain family 7, 4.98871 5.00E- member A 05 NM_001007729 Cxcl4 platelet factor 4 4.92852 5.00E- 05 NM_001128494 Lyc2 lysozyme 2; lysozyme C type 4.56569 5.00E- 2 05 NM_198748 Scin scinderin 4.45596 0.0001 5 196

NM_019325 Myh4 myosin, heavy chain 4, 4.36608 5.00E- skeletal muscle 05 NM_001025115 RGD1311584 signal transducing adaptor 4.35246 5.00E- family member 1 05 NM_013085 Plau plasminogen activator, 4.32848 5.00E- urokinase 05 NM_022379 Tfec transcription factor EC 4.28581 5.00E- 05 NM_001013927 RGD1308734 phospholipase B domain 4.2262 5.00E- containing 1 05 NR_027324 H19 H19 fetal liver mRNA 4.15076 5.00E- 05 NM_013025 Ccl3 chemokine (C-C motif) ligand 4.12364 5.00E- 3 05 NM_031713 Pirb leukocyte immunoglobulin- 4.11551 5.00E- like receptor, subfamily B 05 (with TM and ITIM domains), member 3; similar to paired- Ig-like receptor A11; leukocyte immunoglobulin- like receptor, subfamily B (with TM and ITIM domains), member 3-like; paired-Ig-like receptor A2 NM_012907 Apobec1 apolipoprotein B mRNA 4.00678 5.00E- editing enzyme, catalytic 05 polypeptide 1 NM_001109439 LOC681325 hypothetical protein 3.98563 0.0043 LOC681325 NM_001107777 Siglec1 sialic acid binding Ig-like 3.96423 5.00E- lectin 1, sialoadhesin 05 NM_001106283 Folr2 folate receptor 2 (fetal) 3.85912 5.00E- 05 NM_001008518 MGC105649 hypothetical LOC302884 3.85641 5.00E- 05 NM_022849 Dmbt1 deleted in malignant brain 3.74382 0.0003 tumors 1 NM_031530 Ccl2 chemokine (C-C motif) ligand 3.72447 5.00E- 2 05 NM_001191939 Msr1 macrophage scavenger 3.71724 5.00E- receptor 1 05 NM_031598 Pla2g2a phospholipase A2, group IIA 3.71664 0.0001 (platelets, synovial fluid) 197

NM_001013433 Arl11 ADP-ribosylation factor-like 3.69556 5.00E- 11 05 NM_022221 Mmp8 matrix metallopeptidase 8 3.6556 5.00E- 05 NM_001202463 Cd300le Cd300 molecule-like family 3.60416 5.00E- member E 05 NM_001270701 Kcnn4 potassium intermediate/small 3.58598 5.00E- conductance calcium-activated 05 channel, subfamily N, member 4 NM_001168284 RGD1561778 similar to dendritic cell- 3.58422 0.0005 derived immunoglobulin(Ig)- 5 like receptor 1, DIgR1 - mouse NM_133306 Oldlr1 oxidized low density 3.5599 5.00E- lipoprotein (lectin-like) 05 receptor 1 NM_020104 Myl1 myosin, light polypeptide 1 3.53797 0.0003 NM_001007612 Ccl7 chemokine (C-C motif) ligand 3.53751 0.0075 7 5 NM_053734 Ncf1 neutrophil cytosolic factor 1 3.53037 5.00E- 05 NM_001107336 Dok3 docking protein 3 3.51196 5.00E- 05 NM_017143 F10 coagulation factor X 3.48524 5.00E- 05 NM_001105971 Slamf9 SLAM family member 9 3.48407 5.00E- 05 NM_001205348 Cd300a Cd300a molecule 3.48054 5.00E- 05 NM_001011987 Glipr1 GLI pathogenesis-related 1 3.46454 5.00E- 05 NM_023953 Mox2r CD200 receptor 1 3.46333 5.00E- 05 NM_001013147 Axl Axl receptor tyrosine kinase 3.4306 5.00E- 05 NM_001107491 RGD1304717 chemokine (C-X-C motif) 3.42815 0.0026 ligand 17 5 NM_053372 Slpi secretory leukocyte peptidase 3.40279 0.0043 inhibitor 5 NM_001106748 Batf basic leucine zipper 3.39281 0.0024 transcription factor, ATF-like 5 NM_019316 Mafb v-maf musculoaponeurotic 3.37971 5.00E- fibrosarcoma oncogene 05

198

homolog B (avian) NM_001191994 Slc37a2 solute carrier family 37 3.36868 5.00E- (glucose-6-phosphate 05 transporter), member 2 NM_001107980 Rspo1 R-spondin homolog (Xenopus 3.3378 5.00E- laevis) 05 NM_001170481 Pi16 peptidase inhibitor 16 3.31968 5.00E- 05 NM_012771 Lyz lysozyme 2; lysozyme C type 3.30125 5.00E- 2 05 NM_012530 Ckm creatine kinase, muscle 3.27475 5.00E- 05 NM_031832 Lgals3 lectin, galactoside-binding, 3.27282 5.00E- soluble, 3 05 NM_133416 Bcl2a1 B-cell leukemia/lymphoma 2 3.24853 5.00E- related protein A1d 05 NM_031012 Anpep alanyl (membrane) 3.24744 5.00E- aminopeptidase 05 NM_012912 Atf3 activating transcription factor 3.22 5.00E- 3 05 NM_053565 Socs3 suppressor of cytokine 3.19816 5.00E- signaling 3 05 NM_001134716 Clec12a C-type lectin domain family 3.19775 5.00E- 12, member A 05 NM_001106338 Ms4a7 membrane-spanning 4- 3.18561 0.0019 domains, subfamily A, 5 member 7 NM_001017496 Cxcl13 chemokine (C-X-C motif) 3.18497 5.00E- ligand 13 05 NM_019144 Acp5 acid phosphatase 5, tartrate 3.17689 0.0001 resistant NM_030848 Bst1 bone marrow stromal cell 3.16583 0.0001 antigen 1 5 NM_013154 Cebpd CCAAT/enhancer binding 3.16583 5.00E- protein (C/EBP), delta 05 NM_133530 Mmp13 matrix metallopeptidase 13 3.16068 5.00E- 05 NM_138850 Fap fibroblast activation protein, 3.15002 5.00E- alpha 05 NM_001135767 Gngt2 similar to guanine nucleotide 3.14706 5.00E- binding protein (G protein), 05 gamma transducing activity polypeptide 2; guanine nucleotide binding protein (G 199

protein), gamma transducing activity polypeptide 2 NM_053635 St14 suppression of tumorigenicity 3.13688 5.00E- 14 (colon carcinoma) 05 NM_031644 Ptgds2 prostaglandin D2 synthase 2, 3.11911 5.00E- hematopoietic 05 NM_012687 Tbxas1 thromboxane A synthase 1, 3.07026 5.00E- platelet 05 NM_023965 Cybb cytochrome b-245, beta 3.03161 5.00E- polypeptide 05 NM_133555 Csf2rb1 colony stimulating factor 2 3.01428 5.00E- receptor, beta, low-affinity 05 (granulocyte-macrophage) NM_001100984 Ncf2 neutrophil cytosolic factor 2 3.00396 5.00E- 05 NM_198049 Soat solute carrier family 10 3.00291 5.00E- (sodium/bile acid 05 cotransporter family), member 6 NM_001009681 Oasl1 2'-5'-oligoadenylate 2.95747 5.00E- synthetase-like 05 NM_001013428 Pla2g2d phospholipase A2, group IID 2.94995 0.0044 5 NM_001101009 Tlr8 toll-like receptor 8 2.94843 5.00E- 05 NM_176074 C6 complement component 6 2.9437 5.00E- 05 NM_019285 Adcy4 adenylate cyclase 4 2.93836 5.00E- 05 NM_053544 Sfrp4 secreted frizzled-related 2.90956 5.00E- protein 4 05 NM_031115 Sctr secretin receptor 2.90627 0.0001 5 NM_001107742 Slc43a1 solute carrier family 43, 2.90526 0.0003 member 1 NM_001008839 RT1-Aw2 RT1 class I, CE14; RT1 class 2.89927 5.00E- I, CE16; RT1 class Ia, locus 05 A2; RT1 class Ib, locus Cl; RT1 class Ia, locus A1; RT1 class I, A3 NM_001031656 Serinc2 serine incorporator 2 2.89096 0.0001 NM_199082 Sectm1b secreted and transmembrane 2.88578 0.0068 1B NM_017185 Tnni2 troponin I type 2 (skeletal, 2.88087 0.0023 200

fast) 5 NM_031560 Ctsk cathepsin K 2.85889 5.00E- 05 NM_022617 Mpeg1 macrophage expressed gene 1 2.85559 5.00E- 05 NM_001191940 Aoah acyloxyacyl hydrolase 2.85004 5.00E- (neutrophil) 05 NM_172222 C2 complement component 2 2.83532 5.00E- 05 NM_053843 Fcgr3 Fc fragment of IgG, low 2.8321 5.00E- affinity IIb, receptor (CD32); 05 Fc fragment of IgG, low affinity IIa, receptor (CD32) NM_001037660 Csf2ra granulocyte-macrophage 2.83194 5.00E- colony stimulating receptor 05 alpha

201

Supplemental Table 3.2

Week 6 CTL vs EXC Transcripts Gene ID Gene Symbol Gene Name Log2(fold p-value change) NM_012666 Tac1 tachykinin 1 -1.74482 0.0007 NM_176861 Kcnmb2 potassium large conductance -1.62067 0.0015 calcium-activated channel, 5 subfamily M, beta member 2 NM_00110608 Tex15 similar to testis expressed gene -1.60752 5.00E- 7 15; testis expressed 15 05 NM_080587 Gabra4 gamma-aminobutyric acid -1.43469 5.00E- (GABA) A receptor, alpha 4 05 NM_052805 Chrna3 cholinergic receptor, nicotinic, -1.42403 0.0004 alpha 3 NM_012869 Npy5r neuropeptide Y receptor Y5 -1.41599 5.00E- 05 NM_00110954 Nkain3 Na+/K+ transporting ATPase -1.39927 0.0007 0 interacting 3 5 NM_00119184 Atp6ap1l ATPase, H+ transporting, -1.37169 0.0028 3 lysosomal accessory protein 1- 5 like NM_00102424 Gpr139 G protein-coupled receptor 139 -1.36808 0.0066 1 5 NM_00100202 Slco4c1 solute carrier organic anion -1.35636 0.0033 4 transporter family, member 4C1 NM_012568 Glra2 glycine receptor, alpha 2 -1.34906 5.00E- 05 NM_133570 Grp gastrin releasing peptide -1.33337 0.0018 NM_032065 Slc1a6 solute carrier family 1 (high -1.32376 0.0018 affinity aspartate/glutamate transporter), member 6 NM_00110923 RGD156279 neurogenic differentiation 6 -1.30854 0.0001 7 3 5 NM_00111318 Gria4 glutamate receptor, ionotropic, -1.28724 5.00E- 5 AMPA4 05 NM_053724 Glra3 glycine receptor, alpha 3 -1.2819 5.00E- 05 NM_00113448 Plcxd2 phosphatidylinositol-specific -1.27931 5.00E- 1 phospholipase C, X domain 05 containing 2; CD96 molecule NM_012686 Vsnl1 visinin-like 1 -1.2684 5.00E- 05

202

NM_022302 Efcbp1 N-terminal EF-hand calcium -1.26821 5.00E- binding protein 1 05 NM_012971 Kcna4 potassium voltage-gated -1.26734 5.00E- channel, shaker-related 05 subfamily, member 4 NM_053937 Kcnh6 potassium voltage-gated -1.26663 0.0003 channel, subfamily H (eag- 5 related), member 6 NM_023968 Npy2r neuropeptide Y receptor Y2 -1.2665 0.0085 NM_021680 Nxph4 neurexophilin 4 -1.26408 5.00E- 05 NM_017053 Tacr3 tachykinin receptor 3 -1.24953 0.0001 NM_00110930 RGD156550 similar to PSST739 protein -1.2469 0.0001 8 1 (von Willebrand factor C domain-containing protein 2- like) NM_031736 Slc27a2 solute carrier family 27 (fatty -1.24541 5.00E- acid transporter), member 2 05 NM_00110604 Dleu7 deleted in lymphocytic -1.24444 5.00E- 3 leukemia, 7 05 NM_080691 Cacng3 calcium channel, voltage- -1.23361 0.0002 dependent, gamma subunit 3 5 NM_023974 Synpr synaptoporin -1.23282 5.00E- 05 NM_183326 Gabra1 gamma-aminobutyric acid -1.23279 5.00E- (GABA) A receptor, alpha 1 05 NM_053594 Ptprr protein tyrosine phosphatase, -1.22631 0.0002 receptor type, R 5 NM_00116293 Ctxn2 cortexin 2 -1.22374 5.00E- 5 05 NM_00101274 Cbln2 cerebellin 2 precursor -1.22081 5.00E- 0 05 NM_198786 Mal2 mal, T-cell differentiation -1.21868 5.00E- protein 2 05 NM_022280 Lrat lecithin-retinol acyltransferase -1.21743 5.00E- (phosphatidylcholine-retinol-O- 05 acyltransferase) NM_00102567 Rassf6 Ras association (RalGDS/AF-6) -1.21517 0.0003 1 domain family member 6 NM_012727 Camk4 calcium/calmodulin-dependent -1.21253 5.00E- protein kinase IV 05 NM_012765 Htr2c 5-hydroxytryptamine -1.20722 5.00E- (serotonin) receptor 2C 05

203

NM_00113581 Mgat4c mannosyl (alpha-1,3-)- -1.20004 5.00E- 4 glycoprotein beta-1,4-N- 05 acetylglucosaminyltransferase, isozyme C (putative) NM_00110088 St6gal2 ST6 beta-galactosamide alpha- -1.19991 0.0003 8 2,6-sialyltranferase 2 5 NM_013165 Cckbr cholecystokinin B receptor -1.1977 0.0002 5 NM_013099 Mc4r melanocortin 4 receptor -1.196 0.0001 5 NM_012958 Galr1 galanin receptor 1 -1.19381 5.00E- 05 NM_053427 Slc17a6 solute carrier family 17 (Na- -1.19112 5.00E- dependent inorganic phosphate 05 cotransporter), member 6 NM_00127057 Snap25 synaptosomal-associated -1.19079 5.00E- 5 protein 25, transcript variant 1 05 NM_00102497 RGD130584 hypothetical LOC294883 -1.18907 5.00E- 8 4 05 NM_00101220 Dpp10 dipeptidylpeptidase 10 -1.18737 5.00E- 5 05 NM_019348 Sstr2 somatostatin receptor 2 -1.18643 0.0002 5 NM_053393 Cdh8 cadherin 8 -1.17739 5.00E- 05 NM_00112818 Myadml2 myeloid-associated -1.17442 0.0015 5 differentiation marker-like 2 5 NM_00104787 RGD156519 contactin associated protein-like -1.1736 5.00E- 3 4 5B 05 NM_012563 Gad2 glutamate decarboxylase 2 -1.1727 5.00E- 05 NM_019344 Rgs8 regulator of G-protein signaling -1.17032 5.00E- 8 05 NM_170666 Rims4 regulating synaptic membrane -1.16923 0.0004 exocytosis 4 NM_00110639 Hs3st5 heparan sulfate (glucosamine) -1.16698 5.00E- 2 3-O-sulfotransferase 5 05 NM_053428 Fgf13 fibroblast growth factor 13 -1.16675 5.00E- 05 NM_00111336 connector enhancer of kinase -1.16567 5.00E- 6 suppressor of Ras 2 05 NM_00110899 Trhde thyrotropin-releasing hormone -1.16285 5.00E- 1 degrading enzyme 05

204

NM_024000 Camkv CaM kinase-like vesicle- -1.16147 5.00E- associated 05 NM_053346 Nrn1 neuritin 1 -1.15862 5.00E- 05 NM_00110956 LOC689978 neurensin 2 -1.15736 5.00E- 1 05 NM_031044 Hnmt histamine N-methyltransferase -1.1538 0.0003 5 NM_013148 Htr5a 5-hydroxytryptamine -1.15142 0.0022 (serotonin) receptor 5A NM_170668 Slc13a5 solute carrier family 13 (Na- -1.14979 5.00E- dependent citrate transporter), 05 member 5 NM_145777 Olfm3 olfactomedin 3 -1.14917 5.00E- 05 NM_012920 Camk2a calcium/calmodulin-dependent -1.14657 5.00E- protein kinase II alpha 05 NM_00119196 -1.14345 5.00E- 6 05 NM_017122 Hpca hippocalcin -1.14242 5.00E- 05 NM_031758 Mchr1 melanin-concentrating hormone -1.14145 0.002 receptor 1 NM_00110605 Klhl1 kelch-like 1 (Drosophila) -1.13931 5.00E- 4 05 NM_00110745 Rgs17 regulator of G-protein signaling -1.13919 5.00E- 9 17 05 NM_013002 Pcp4 Purkinje cell protein 4 -1.1383 5.00E- 05 NM_019162 Tac2 tachykinin 2 -1.13752 5.00E- 05 NM_053859 Slc17a7 solute carrier family 17 (Na- -1.13609 0.0033 dependent inorganic phosphate 5 cotransporter), member 7 NM_00110802 Lrfn5 leucine rich repeat and -1.136 5.00E- 4 fibronectin type III domain 05 containing 5 NM_00110846 Usp29 ubiquitin specific peptidase 29 -1.13033 5.00E- 5 05 NM_00110888 RGD156617 kelch-like 14 (Drosophila) -1.13021 0.0006 5 8 5 NM_031778 Kcns3 potassium voltage-gated -1.12948 5.00E- channel, delayed-rectifier, 05

205

subfamily S, member 3 NM_017007 Gad1 glutamate decarboxylase 1 -1.12732 5.00E- 05 NM_024394 Htr3a 5-hydroxytryptamine -1.12682 0.0021 (serotonin) receptor 3a NM_173146 Unc13c unc-13 homolog C (C. elegans) -1.12444 5.00E- 05 NM_00110932 RGD156514 melanoma associated antigen -1.12306 5.00E- 1 8 (mutated) 1-like 1 05 NM_00113474 zinc finger, matrin type 4 -1.12077 5.00E- 7 05 NM_00101427 LOC367515 similar to RIKEN cDNA -1.11915 0.0013 1 1700081O22 NM_012799 Nmbr neuromedin B receptor -1.11868 0.0038 NM_00110929 RGD156208 similar to expressed sequence -1.11744 0.0004 4 4 AI118078 NM_053996 Slc6a7 solute carrier family 6 -1.11616 5.00E- (neurotransmitter transporter, L- 05 proline), member 7 NM_00119172 -1.11524 5.00E- 1 05 NM_00101399 LOC305691 similar to hypothetical protein -1.11216 5.00E- 2 FLJ22419 05 NM_00101418 RGD130699 similar to Protein C20orf103 -1.11141 5.00E- 3 1 precursor 05 NM_00119172 -1.11074 5.00E- 6 05 NM_012853 Htr4 5-hydroxytryptamine -1.10862 0.0004 (serotonin) receptor 4 5 NM_053991 vasoactive intestinal peptide -1.10601 0.0002 NM_012585 Htr1a 5-hydroxytryptamine -1.10542 0.0026 (serotonin) receptor 1A 5 NM_00104786 RGD156634 contactin associated protein-like -1.1051 5.00E- 6 3 5A; similar to contactin 05 associated protein-like 5 isoform 1 NM_00113056 metallophosphoesterase domain -1.10402 5.00E- 9 containing 1 05 NM_00110238 Nts neurotensin -1.10364 5.00E- 1 05 NM_017295 Gabra5 gamma-aminobutyric acid -1.10258 5.00E- (GABA) A receptor, alpha 5 05

206

NM_00110917 LOC499506 ankyrin repeat domain 34B -1.10112 5.00E- 4 05 NM_019189 Hapln1 hyaluronan and proteoglycan -1.09539 5.00E- link protein 1 05

207

Supplemental Table 3.3

Week 3 CTL vs EXC Metabolites Mass-to-charge ratio Retention Time (m/z) 68 16.24 69.0236 10.36 86.0448 10.23 87.0296 10.23 103.7636 8.52 104.0521 10.27 104.0889 21.13 105.0914 9.02 110.0512 10.94 118.0645 9.41 125.9987 12.98 132.013 9.73 136.0363 9.76 140.0425 9.38 144.0751 9.38 161.0997 10.62 162.0832 9.82 163.0857 9.82 168.1167 10.91 169.9584 12.72 171.1171 4.38 176.8417 20.34 182.133 12.92 184.0407 15.28 188.984 10.63 192.0912 11.32 200.0545 9.67 204.0855 9.36 205.0869 9.36 208.0935 9.1 210.5985 11.89 214.14 13.42 220.0973 20.75 225.6164 10.56 227.0717 11.62 208

229.1561 9.26 229.6033 9.07 232.113 9.07 232.6112 12.67 233.6151 9.13 233.6226 9.46 234.0345 13.12 238.1127 13.42 239.1204 10.57 244.0472 9.36 246.1208 9.04 246.1248 8.19 246.1275 9.04 248.1037 9.36 248.1242 14.42 249.0312 17.58 249.1143 9.54 249.1223 9.28 249.1429 9.91 254.0791 9.35 254.1471 9.07 254.6404 21.03 257.0946 13.01 263.0628 10.94 263.0984 9.74 265.0159 21.31 267.09 13.45 268.0539 9.13 269.0582 9.13 273.1373 13.01 277.0386 9.48 282.1493 9.2 284.0468 10.54 285.0424 10.94 288.9296 12.89 290.0248 9.58 291.1827 9.28 292.144 9.41 293.0218 12.42

209

294.9406 12.73 297.0929 9.44 299.1234 13.97 306.0268 10.57 306.619 9.44 307.029 10.57 310.6192 9.69 315.6317 9.83 316.1903 4.31 316.9238 12.7 322.1455 14.85 325.1024 9.04 331.0546 12.79 332.6375 9.76 333.6349 12.57 335.4901 9.55 335.6282 13.08 336.6462 12.26 339.8357 9.18 341.1427 13.67 342.2005 4.31 344.2157 4.38 345.0632 12.7 345.2197 4.22 346.6256 12.4 347.2088 12.62

210

Supplemental Table 3.4

Week 6 CTL vs EXC Metabolites Mass-to-charge ratio Retention Time (m/z) 70.0543 20.3 72.0683 10.32 78.9717 20.18 86.0806 9.86 87.076 20.3 111.0735 20.36 113.0868 20.3 118.0645 9.41 124.0359 6.05 124.0359 4.54 125.9987 12.98 126.0785 6.97 130.1097 20.91 130.1346 4.36 131.9929 9.77 134.978 12.12 136.0363 9.76 136.5493 20.51 140.0425 9.38 141.0953 9.16 144.0752 10.71 144.0782 11.36 147.0653 6.69 147.0653 6.21 147.9755 12.7 148.0666 6.21 149.9857 11.78 156.1215 20.36 156.5672 9.04 157.1154 9.12 158.0672 20.3 159.0338 9.35 159.0474 10.87 162.0832 9.82 163.0857 9.82 211

166.0796 9.3 166.0807 20.36 169.9584 12.72 171.0173 20.6 171.1171 4.38 176.0398 10.61 176.8417 20.34 176.8665 19.42 179.0868 20.3 184.0394 16.81 187.5994 20.3 188.0913 20.36 189.0848 20.3 189.0993 9.04 190.1667 9.65 190.1674 20.82 192.0935 14.57 193.5769 9.26 196.1122 11.1 196.6107 20.64 197.0948 20.36 197.4426 9.58 198.0529 10.26 200.0855 20.36 200.5869 20.36 200.6037 20.39 201.102 20.3 202.053 20.36 202.0596 9.26 202.0841 20.36 203.0151 11.99 203.0156 9.56 203.0449 20.3 203.0536 13.64 203.1098 9.06 204.0166 9.9 204.0271 11.88 204.0855 9.36 204.1316 20.27

212

205.0869 9.36 205.6052 20.27 206.0178 16.81 206.1074 20.27 209.1148 20.39 209.6052 12.44 210.098 20.36 210.5995 20.3 211.6055 9.32 212.1473 9.34 214.0192 10.87 214.0841 20.45 214.1176 20.36 214.2138 4.45 216.1029 20.3 216.5949 20.39 217.0891 20.36 217.6053 13.33 218.0957 9.23 218.6124 20.27 218.8955 11 219.1129 20.39 219.7571 20.66 220.0676 20.3 220.1169 20.39 220.1171 13.02

213

Supplemental Table 3.5

Week 3 SHAM vs EXC Metabolites Mass-to-charge ratio Retention Time (m/z) 118.0645 9.41 125.9987 12.98 133.603 9.69 140.0425 9.38 144.0751 9.38 162.0832 9.82 163.0857 9.82 204.0855 9.36 271.6403 20.58 315.9497 20.36 316.1903 4.31 339.8357 9.18 341.2422 4.22 342.2005 4.31 342.2456 4.71 344.2157 4.38 344.2172 8.28 345.2197 4.22 365.8019 8.95 370.2266 8.19 370.2285 4.22 372.2425 8.28 381.2289 4.48 381.23 4.22 388.2328 8.73 398.2052 4.54 407.8144 20.58 417.7806 4.47 418.3861 4.51 418.4843 4.42 418.8193 4.42 419.506 4.45 419.8078 4.51 437.5005 9.06 463.8944 12.7 214

466.9849 11.78 489.9195 10.18 489.9228 9.55 494.5156 9.06 494.8533 9.04 499.5223 9.06 570.2913 9.38 620.6716 9.76 640.5348 11.61 653.8999 9.42 672.1864 4.5 697.2837 9.13 730.0997 9.88 741.2763 9.03 741.7845 9 742.2716 9 742.7736 9.06 748.2773 9.06 748.7793 9.06 749.2815 9.06 749.7839 9.06 766.6058 9.78 940.8192 9.75 948.2773 9.71

215

Supplemental Table 3.6

Week 6 SHAM vs EXC Metabolites Mass-to-charge ratio Retention Time (m/z) 60.0367 3.1 72.0683 10.32 118.0645 9.41 134.978 12.12 140.0425 9.38 147.0636 4.36 147.0653 6.21 147.0653 6.69 147.9755 12.7 148.0666 6.21 148.0666 4.36 162.0832 9.82 163.0857 9.82 169.9584 12.72 197.5947 12.13 200.1999 6.67 200.2024 7.77 204.0855 9.36 205.0869 9.36 234.0369 9.49 246.1248 8.19 254.7903 20.45 269.1972 10.49 289.2174 11.07 294.7877 20.64 316.9238 12.7 324.9446 12.65 331.0546 12.79 335.0684 9.08 402.1836 12.6 422.0251 12.57 422.8659 12.52 426.9171 11.97 441.9136 12.64 460.0742 9.79 216

463.8944 12.7 471.7151 20.54 494.5156 9.06 494.8533 9.04 495.5165 9 499.5223 9.06 499.8599 9.06 503.878 12.61 505.1677 4.35 506.5116 9.1 507.9642 10.66 508.4276 9.55 508.429 8.97 508.6759 9.55 508.6801 9 516.9142 10.66 517.6627 9.17 517.9203 9.03 549.1893 4.35 553.2628 9.38 566.9099 12.7 575.1957 4.35 612.8887 9.06 614.1708 9.68 617.903 9.06 653.2379 9 653.5662 9 653.8999 9.42 653.9019 9 668.9638 9.1 675.0761 8.97 676.8982 9.02 676.8999 9.55 676.9035 10 677.2339 9.42 677.238 8.97 677.5621 10.66 677.5646 9.42 677.5677 21.04 677.5686 8.97 217

677.9001 10.75 677.9067 8.97 678.5684 8.97 682.5747 9.06 682.903 9.06 684.2279 11.32 684.2285 10.59 684.2314 9.06 684.5537 9.01 684.8861 9.01 684.8898 10.57 685.5512 9.01 687.8917 9.06 688.2437 8.98 689.3223 20.36 689.5475 9.06 689.8806 9.52 689.8811 9.06 690.2121 9.01 690.5524 9.06 690.8887 9.09 705.9061 9.02 706.576 8.97 707.7317 9.13 708.4757 9.15 717.1999 9.19 741.2763 9.03 741.7845 9 742.2716 9 742.7736 9.06 748.2773 9.06 748.7793 9.06 749.2815 9.06 749.7839 9.06 752.7632 9.04 766.6058 9.78 767.2665 10.39 767.285 9 767.5313 9 767.7855 9 218

805.5016 9.07 808.7455 9.13 835.5954 9 844.5332 9 848.2784 9.01 848.5289 9.13 891.2784 9.15 891.6089 9.2 943.6455 9.18 943.96 9.15 944.6374 9.15

219

APPENDIX B

SHORT DURATION ELECTRICAL STIMULATION TO ENHANCE NEURITE

OUTGROWTH AND MATURATION OF ADULT NEURAL STEM PROGENITOR

CELLS2

Summary

New therapies are desperately needed for human central nervous system (CNS)

regeneration to circumvent the lack of innate regenerative ability following traumatic

injuries. Previously attempted therapies have been stymied by barriers to CNS

regeneration largely because of protective mechanisms such as the blood brain barrier,

inhibitory molecules, and glial scar formation. The application of electric fields (EFs)

have shown promise for enhancing peripheral nervous system regeneration, but to date

have not been as thoroughly studied for CNS regeneration. The objective of this study is

to better understand how short duration EFs can be harnessed to direct adult neural stem

progenitor cell (NSPC) neurogenesis, neurite extension, and maturation. Herein, NSPCs

were exposed to physiological electrical fields of 25 and 50 V/m of direct current (DC)

for 10 min/d for 2 d with a total differentiation time of 3 d. Culturing conditions consisted

22 The main author of this work is Liza J. Kobelt. This manuscript has been published as Kobelt, L. J., Wilkinson, A.E., McCormick, A.M., Willits, R.K., Leipzig, N.D. Short Duration Electrical Stimulation to Enhance Neurite Outgrowth and Maturation of Adult Neural Stem Progenitor Cells. Annals of Biomedical Engineering. (2014) 42(10): 2164-2176. 220

of either mitogenic growth factors or the neuronal differentiation factor interferon-γ (IFN-

γ). Stimulated NSPCs showed lengths that were over five times longer than control cells

(0 V/m; 112.0 ± 88.8 μm at 25 V/m vs. 21.3 ± 8.5 μm for 0 V/m with IFN-γ) with the longest neurites reaching up to 600 µm. Additionally, EF stimulation resulted in mature neuronal morphologies and signs of differentiation through positive βIII tubulin, neuronal nuclei (NeuN), and better organized filamentous-actin (f-actin) staining with growth cone formation. Additionally, the neurites and soma of stimulated NSPCs showed increases in intracellular Ca2+ during EF stimulation, also signifying the presence of functional

neurons capable of electrical conductance and communication with other cells. Our study

demonstrates that short stimulation times (10 min/d) result in significant neurite

extension of stem cells in a quick time frame (3 d). This electrical stimulation modality is

potentially advantageous for promoting axon re-growth at an injury site using delivered adult stem cells; however, significant work still remains to understand both the delivery approach of cells as well as EF application in vivo.

Introduction

Traumatic brain and spinal cord injuries affect over 2 million people in the US annually and are generally irreversible (1). Failure to heal central nervous system (CNS) injuries is largely due to intrinsic inhibition of both axonal re-growth and interconnection past the site of injury (6, 11, 14, 51, 48). Many approaches are being tested to heal CNS injuries; however, a completely successful methodology generating functional restoration has yet to be discovered. Electric fields (EFs) occur endogenously and have long been used for both healing and pain management in medical practice (50). The endogenous 221

EFs that exist in the brain play an important role in injury, development, and

galvanotaxis, defined by the directional movement of a biological entity in response to

EFs (38). EF stimulation is also well established for its role in enhancing peripheral nerve regeneration (39, 37, 48).

For the most part, the molecular mechanisms affected by EFs are not fully

understood, especially within the CNS; however, it is believed that they play an

important role in homeostasis due to the inherent electrical nature of the tissue (15). In

addition, EFs have been shown to have multiple effects on various cell types outside the

nervous system, affecting aspects of development, wound healing, and regeneration (35,

48). For example, direct current (DC) EFs are enhanced physiologically at injury sites and are thought to be partially responsible for healing tissues immediately around the wound (50, 40).

EFs are a key aspect of normal brain function in that they aid in the transmission

of cell signaling. Electrical signals in the brain are due to spatial changes in the function

of ion pumps and small ion secretions from individual cells. These small changes drive

extracellular flow of ions and establish voltage gradients in tissues (35). During

development, and to a limited degree in adulthood, cells migrate in the brain from their

origin to their place of maturation in other areas, which could be due in part to EFs (31,

1). Growth during development has been shown to be dramatically affected by endogenous EFs; they promote cell migration, cell death, limb growth, and are vital during initial developmental stages (30, 19, 40). Without normal EFs, developmental abnormalities are more likely to form (30, 45).

222

NSPCs are an attractive cell source for tissue regenerative purposes because of

their ability to differentiate into the three primary cell types of the CNS: astrocytes,

oligodendrocytes and neurons (32, 47, 27). In addition, cytokines such as interferon

gamma (IFN-γ), epidermal growth factor (EGF), and fibroblast growth factor (FGF) play an important role in innate immunity and, therefore, in preservation and regeneration of the brain. Previous studies have revealed that IFN-γ can be used as a differentiation factor for NSPCs and functions to significantly enrich neuronal populations over oligodendrocytes or astrocytes (25-27). EGF and FGF are commonly used to expand

NSPCs in number due to their mitogenic capacity (10, 23). During homeostasis, EFs exist

in the CNS and the resulting currents play a key role in cell function and likely affect

stem cell responses. However, it is difficult to pinpoint an exact range that should be

applied to facilitate neuronal maturation and axon extension after CNS trauma. In the

native injury response, endogenous levels of 42-150 V/m have been recorded (28).

Based on the literature, these applied EFs can range from 33 V/m to 437 V/m depending

on the animal model chosen and specific cell type of interest (34, 38, 3, 15). The combined treatment of NSPCs with EFs and known differentiation molecules has not been thoroughly studied. By utilizing both mitogenic and differentiation cues, we aim to determine if DC EFs can override and/or provide synergistic responses by directing both neurite extension and the cell fate of NSPCs in culture. Therefore, the primary objective of this study was to determine the ability of DC EF stimulation to enhance neurite extension and guidance of NSPCs and their differentiation into neurons with the addition of IFN-γ or in the presence of EGF and FGF (25-27). This regime can potentially be used

223

in the future in combination with other current tissue engineered therapies. Based on

previous work with both primary neurons and stem cells, we hypothesized that short DC

EF stimulation would enhance outgrowth of NSPC neurites, cause alignment, and

increase maturation as evidenced by mature axon/growth cone formation and expression

of neuronal proteins.

Materials and Methods

Cell Culture and Isolation

NSPCs were isolated from the subventricular zone (SVZ) of 6 wk old adult

female Wistar rat (Charles River, Wilmington, MA, USA) forebrains and expanded as

neurospheres in growth medium containing neurobasal (NBM; Life Technologies, Grand

Island, NY, USA), 100 μg/mL penicillin streptomycin (PS; Life Technologies), 2 mM L-

glutamine (L-glut; Life Technologies), B-27 (Life Technologies), 0.02 μg/mL heparin

(Sigma-Aldrich, St. Louis, MI, USA), 20 ng/mL of EGF (Sigma-Aldrich) and 20 ng/mL

FGF (Peprotech, Rocky Hill, NJ, USA) as described previously (52).

NSPC Stimulation Regime

Electrical stimulation chambers consisted of a 63 mm long non-conductive open

silicone rectangular box with a 75 x 25 mm glass slide bottom. Platinum electrodes were

adhered to the silicone frame and connected to a DC power source (MPJA, West Palm

Beach, FL, USA). The platinum electrodes had a width 7.0 ± 1.0 mm and height of 11 ±

0.48 mm with a separation of 54 ± 0.88 mm between each electrode. Glass coverslips in

24-well culture plates were used as controls. Before experimentation, control coverslips 224

and stimulation chambers were coated with 50 µg/mL poly-d-lysine (PDL; BD

Biosciences, Bedford, MA, USA) and 5 µg/mL laminin (Life Technologies, Carlsbad,

CA). Cells were seeded at 40,000 cells/cm2 in growth medium overnight and cultured in

either differentiation or growth medium the following day. Differentiation media

contained NBM, 150 ng/mL IFN-γ for neuronal specification (Peprotech), 100 μg/mL PS,

2 mM L-glut and B-27. Stimulation was performed in a copper-lined humidified CO2

incubator (HERAcell 150i; Thermo Scientific, Sunnyvale, CA, USA) at 37°C and 5%

CO2 providing effective shielding from any external interference. After about 10 h in differentiation medium, cells were stimulated in PBS (pH=7.4) at 25 or 50 V/m for 10 min/d for 2 d (3 d total differentiation), followed by replacement of the proper medium.

The voltages we selected for this study we arrived upon after preliminary cell studies from 0-28 V/m (Suppl. Figs. 1-2), as well as the fact that an electrochemical reaction

occurred above 50 V/m. Overall current at each voltage was determined by measuring the

resistance of each chamber using a multimeter (DM-40; Greenlee, Lake in the Hills, IL,

USA). Current calculations at 25 V/m were determined to be 1.42 μA and 28.3 μA at 50

V/m. Stimulation above 50 V/m resulted in significant electrochemical reactions.

225

Supplemental Figure 1. Average neurite lengths after 3 d of culture (with 10 min/d EF stimulation on day 1 and 2). Untransformed values are included for each group. Stimulated cells (16, 25, 28 V/m) showed significance from control groups (*, p <0.0001) by single factor ANOVA. Mean ± SD.

Supplemental Figure 2. Polar plots from neurite lengths of initial experiments with NSPCs stimulated at 16, 25, 28 V/m or no stimulation with addition of (A) IFN-γ or (B) EGF/FGF. No directional bias was detected using radial statistical analyses (p > 0.05).

226

NSPCs Length, Polarity and Differentiation Analysis

Live phase contrast images were obtained 24 h after the final stimulation (at 3

days) to allow quantification of neurite outgrowth and polarity. Images were obtained at

random areas throughout the culture area, and approximately 6-10 images were taken per

group. All primary neurites of cells in each image taken were included in analysis.

Images were analyzed with ImageJ software. The length of each neurite and angle was

determined by measuring from the soma to the end of each process, or measured to the

termination of a neurite at a neighboring cell. After imaging, cells were fixed with 3.7%

paraformaldehyde for 5 min, and then permeabilized with 0.1% Triton X-100 (Sigma-

Aldrich) in PBS (pH=7.4) for 5 min for immunohistochemistry. In PBS, a 10% goat serum (Sigma) block was applied for 1 h at room temperature (RT). Samples were again washed with PBS and primary antibody monoclonal mouse anti-βIII tubulin (1:500;

Covance, Princeton, NJ, USA) or mouse anti-nestin (1:10; Developmental Studies

Hybridoma Bank, Iowa City, IA, USA) was left on overnight at 4°C. Cells were washed three times in PBS for 15 min each wash, then the secondary antibody goat anti-mouse

IgG Alexa-Fluor 546 (1:400; Life Technologies) was incubated for at least 1 h at RT.

Samples were washed for 15 min in PBS three times then blocked again with 10% donkey serum (Sigma) in PBS for 1 h at RT. After rinsing samples with PBS, primary antibody rabbit anti-neuronal nuclei (NeuN; 1:800; Millipore, Billerica, MA, USA) was incubated with anti-βIII tubulin samples and Alexa-Fluor 488 phallodin (for filamentous- actin (f-actin); 1:200; Life Technologies) was co-stained with anti-nestin samples overnight at 4°C. After 15 min PBS washes, secondary antibody donkey anti-rabbit IgG

227

Alexa-Fluor 488 (1:400; Life Technologies) was added to anti-NeuN samples for at least

1 h at RT. Following three more 15 min PBS washes, all nuclei were stained with

Hoechst 33342 (10 μM; Life Technologies) for 10 min. Coverslips were mounted with prolong gold (Life Technologies) and imaged with an Olympus IX81 florescent microscope (Tokyo, Japan).

Live Time-lapse Imaging

Additional live brightfield imaging was performed using AxioVision software with an incubated Zeiss microscope (San Diego, CA, USA) held at 37°C at time points before, during, and after stimulation. Chambers of control and stimulated groups were coated with PDL and laminin, and NSPCs were seeded at 40,000 cells/cm2 on day 0.

Cells were stimulated in PBS at 25 V/m for 10 min on day 1 and 2 and imaged at time

points before, during, and after stimulation on day 2.

NSPC Metabolic Activity Assay

PrestoBlue reagent (Life Technologies) was used to assess cell activity before and

after stimulation according to the manufacturer’s instructions. The assay measures the

reducing capacity of cells, which is tied to metabolic functions. Cells were seeded

overnight at 40,000 cells/cm2 at day 0; before the first medium change (day 1) and after

the final EF stimulation (day 3), PrestoBlue reagent (diluted in the appropriate media)

was incubated with the same NSPCs at 37°C for 45 min. Fluorescence intensity (I) was

read (Ex/Em: 555 nm/590 nm) at 45 min for cells and controls using an Infinite M200

spectrophotometer (Tecan, Männedorf, Switzerland). Since PrestoBlue measures the

228

reducing environment of the cell, dithiothreitol (DTT, Chem-Impex International, Wood

Dale, IL, USA), at concentrations of 50, 100, and 1000 µM, was used as a positive control to normalize the data. The fold change in activity was calculated as Iday0/Ida3.

After the incubation period, cells were washed with PBS and either stimulated with an EF

or fixed for IHC analysis, depending on the time point.

Supplemental Figure 3. Metabolic activity of NSPCs assayed using PrestoBlue reagent presented as fold change from day 0 to day 3. All IFN- γ groups decreased in viability while all EGF/FGF cultured cells increased in metabolic activity, resulting in a significant difference between these growth factor treatments as revealed by ANOVA (p < 0.0001).

Calcium Flux Experiments

NSPCs were seeded at 40,000 cells/cm2 in silicone isolators (JTR13R-2.0-Press-

To-Seal Silicone Isolator; Grace Bio-Labs, Bend, OR, USA) inside of 60 mm petri dishes between platinum wires spaced 45 mm apart. PDL and laminin were adsorbed to the petri dish prior to cell seeding as described previously. After overnight culture in growth

229

medium cells were incubated in differentiation media containing 150 ng/mL IFN-γ. No

EF stimulation was performed until calcium fluctuation measurements on day 2.

Prior to analysis on day 2 silicone isolators were removed and Fluo-4, AM (Life

Technologies) molecular probe was prepared at 3 mM in dimethyl sulfoxide (DMSO;

BDH, West Chester, PA USA) and was mixed with an equal amount of 20 w/v % of

Pluronic F-127 (Life Technologies) also dissolved in DMSO. This solution was added to

Hank’s balanced salt solution (HBSS; Life Technologies) to ensure a final concentration of 3 μM for Fluo-4, AM and 2 w/v % for Pluronic F-127. NSPCs were incubated with this mixture for 1 h and then washed in HBSS for at least 1 h. During stimulation and imaging Leibovitz media (L-15; Life Technologies) replaced HBSS. Calcium flux was observed using a 40 X objective on an upright Olympus FV1000 microscope with

Olympus Fluoview software that included settings for Fluo-4 acquisition (494 nm excitation, 506 nm emission). The observed calcium fluctuations are a result of changes due to either extracellular calcium influx or the release of calcium stores from within the cell. About 10 cells per group were examined and a representative video and images were obtained for each group. Each interval lasted 5 min and 10 s, with initially no stimulation for 10 s, followed by 3 min of stimulation at 25 and 50 V/m in separate experiments, then

2 min of no stimulation. Additionally, sequential increases from 25 V/m to 50 V/m to 100

V/m for approximately 1 min each were performed in place of the 3 min stimulation step.

In this experimental set-up and during the short stimulation and culturing time, 100 V/m did not cause an electrochemical reaction at the electrodes, Fluorescent intensity ratios were determined and analyzed using the Olympus Fluoview software. Briefly, straight-

230

line regions of interest (ROI) were drawn along selected neurites as well as across the

soma, and fluorescent intensity versus time data was collected (18). The intensity values

were normalized by the starting intensity values from the same ROI, and then plotted.

Statistical Analysis

Neurite length was tested for normal distribution using the Shapiro-Wilk test. It

was determined that the data was non-normal, therefore, neurite lengths were transformed

using a log transformation as follows: transformed length = log10(length+1). Factors included stimulation levels of (i) 25 V/m and (ii) 50 V/m or (iii) 0 V/m (no stimulation), as well as growth conditions, either under (i) EGF/FGF or (ii) IFN-γ. JMP 10.0 software was utilized to run a two-factor ANOVA model assessing the interaction between these stimulation levels and growth conditions. An α level of 0.05 was used to determine significance. Data is shown as a mean ± standard deviation (SD). Statistical analysis of neurite directionality was performed in Excel using the Rayleigh Z Test. Angular data was doubled since it was bimodal, and the test was performed also using an α level of

0.05 to establish significance. Metabolic activity fold change data was analyzed in JMP

10.0 using a two-factor ANOVA model (factors and levels listed above) and an α level of

0.05.

Results

NSPCs Length, Polarity and Differentiation Analysis

Neurite length data was not normally distributed; therefore, a log transformation was used to fit the data to a proper distribution for statistical analysis as described above. 231

A two-way ANOVA on the transformed lengths showed the interaction between voltage and media conditions to be significantly different (p = 0.0071). Therefore a post hoc,

Tukey’s HSD (honestly significant difference) test was used to analyze significance between groups. Both 25 V/m and 50 V/m groups containing IFN-γ had significantly longer extensions (112.0 ± 88.8 μm, 105.4 ± 58.0 μm), compared to the control groups

(p-value <0.0001) (0 V/m) (Figs. 1-2). The maximum single neurite length seen in stimulated samples containing IFN-γ was 580.0 μm at 25 V/m, while a maximum length of 280.5 μm was measured at 50 V/m for the EGF/FGF stimulated group. The length of each neurite and angle was determined by measuring from the soma to the end of each process, or measured to the termination of a neurite at a neighboring cell. Control groups, receiving no stimulation, averaged lengths of 21.2 ± 8.3 μm and 36.2 ± 32.7 μm with the addition of INF-γ and EGF/FGF, respectively. The maximum single neurite length seen in all controls was 141.9 μm with the addition of IFN-γ. In preliminary experiments a select number of neurites reached lengths up to 600 μm with addition of IFN-γ (Fig. 2C).

We originally hypothesized that EFs would affect polarity of neurite length, but statistical analysis rejected this hypothesis revealing that neurites did not extend significantly in a specific orientation or alignment to the field (p > 0.05).

232

Figure 1. Average neurite lengths after 3 d of culture (with 10 min/d EF stimulation on day 1 and 2) transformed by log10(x+1) for linear statitical analysis. Untransformed values are included for each group inside each bar (μm). Both media conditions ( IFN-γ or EGF/FGF) and varying EF stimulation resulted in a significant difference of NSPC length between groups (p < 0.0001). Letters denote significance by two-factor ANOVA with post hoc Tukey HSD analyses. Transformed and untransformed values presented as mean ± SD, with n ranging from 50-150 neurites.

Qualitatively studying the maturity of NPSCs with EF stimulation in both

medium conditions demonstrated that groups stimulated at 25 V/m and 50 V/m had

superior positive neuronal staining compared to controls (Fig. 3-4). More specifically, the

neuronal stains, β-III tubulin and NeuN, indicated differentiation and maturation of

NSPCs and was apparent at both 25 V/m and 50 V/m (Figs. 3-4 E,I,H,L). However,

nestin indicated that NSPCs did not completely differentiate after 3 d in culture (Figs. 3-4

C,G,K,O). The presence of F-actin was more visible and better organized in the 25 V/m

and 50 V/m stimulated groups compared to controls (Figs. 3-4 F,J). Phase-contrast

images of NSPCs at 25 V/m and 50 V/m also reveal morphological differences and

extensive lengths as compared to controls (Fig. 5). Additionally, mature cytoskeletal

growth cone structures with actin filopodia-like projections were seen in these same two

stimulated groups as well (Fig. 6).

233

Figure 2. (A, B) Polar plots presenting neurite lengths of NSPCs stimulated in vitro at 0 (control), 25 and 50 V/m, with addition of (A) IFN-γ or (B) EGF/FGF. No directional bias was detected using radial statistical analyses (p > 0.05). Angles are in degrees and lengths are shown in μm. (C) Long neurites were especially noticeable in EF stimulated NSPC treatments. Scale bar = 200 μm.

Live Time-lapse Imaging

Live imaging of NSPCs showed that short time (< 2 h) responses were varied. We observed a response showing initial retraction upon the application of an electric field followed by elongation of neurites post stimulation (Suppl. Vid. 1 available online).

Supplemental video 2 (available online) shows that stimulation can result in an almost 234

immediate response and can sometimes result in neurite retraction. Overall the NSPC

neurites seemed to be very active during and directly after stimulation especially

compared to controls (Suppl. Vid. 3 available online).

Figure 3. NSPCs after 3 d in IFN-γ differentiation medium. (A-D) Control (0 V/m) NSPCs display neuronal and cytoskeletal structures but not as prominent as 25 V/m (E- H) and 50 V/m (I-L), which show longer interconnected neurites and more mature neuronal staining (NeuN). All nuclei are stained with Hoechst 33342. Scale bar = 100 µm.

Cell Metabolism Assay

NSPC metabolic activity was tested before and after stimulation using the

PrestoBlue assay so that the fold change in cell activity could be determined; these data are shown in Supplemental Figure 3. Results revealed that cells cultured in IFN-γ are less metabolically active after the 3 d culture period whether or not they were stimulated; while NSPCs cultured in EGF/FGF were significantly more active than the IFN-γ groups

(p < 0.0001). The largest decrease in metabolic activity occurred in the 25 V/m group 235

cultured in IFN-γ with a decrease of 0.48 ± 0.07 fold compared to day 0 (before differentiation factor treatment and electrical stimulation), followed by the 50 V/m group

that decreased by 0.51 ± 0.03 fold compared to day 0.

Figure 4. NSPCs after 3 d in EGF/FGF growth medium. (A-D) NSPCs that did not undergo stimulation have very few neurites and little NeuN staining. (E-H) Cells stimulated at 25 V/m show increased neurite outgrowth and mature neuronal staining (E and H). (I-L) The 50 V/m group showed very high number of neurites with mature looking cell processes. All nuclei are stained with Hoechst 33342. Scale bar = 100 µm.

Calcium Flux Experiments

Calcium flux experiments show that free calcium inside the soma and the neurite

can increase during stimulation at 25 V/m as compared to periods where no EF is present

(Fig. 7, Suppl. Vid. 4 available online). In a representative stimulated neurite at 25 V/m,

the maximum normalized Ca2+ intensity was seen 68 s after the voltage was applied, and then began to wane. Turning off the voltage at 180 s did not appreciably change the rate of decrease as it proceeded back to initial levels. In the soma a different response was

236

observed, and the normalized Ca2+ intensity reached a maximum at 110 s and did not decrease. This observation is partially due to the fact that more calcium was present in the soma to begin with and the laser setting was selected such that we could observe flux in the neurites. In separate experiments EF stimulation was also increased in a step-wise fashion from 0 to 100 V/m (Fig. 7, Suppl. Vid. 5 available online). In the soma of NSPCs stimulated with increasing voltage, there was no measureable increase of Ca2+ intensity at

10 s with 25 V/m, but calcium did increase at 110 s with 50 V/m. Most noticeably, we found that when voltage was increased to 100 V/m at 210 s, there was a sharp decline in

Ca2+. A similar response in the neurite of the same cell was also seen, but to a lesser extent. No electrochemical response was noted for the short interval of stimulation at 100

V/m. This drastic change is believed to be directly tied to the relatively high current (57.5

μA) that was present at this time point or could be a result of depletion of calcium stores.

Discussion

With our experimental set-up, the stimulation of NPSCs with DC EFs of 25 or 50

V/m produced mature neurons with extensions almost 3 times greater than controls after a period of only days. Maximum neurite lengths for stimulated cells were up to 600 μm

(for 25 V/m) with maximum control lengths of no greater than 200 μm (Figs. 1-2). Other studies involving neural stem cells have not observed neurite lengths or extension such as ours. Ariza et al. (3) treated hippocampal-derived NSPCs with DC EFs of 437 V/m for longer stimulation periods, while resulting in a lower cell viability. Additionally, electrical stimulation did not significantly affect cell activity in our study (p > 0.05,

237

Suppl. Fig. 3). It is important to note that only comparing EF strengths (V/m) between studies proves to be difficult as both the current and the resistance are important and these numbers were not reported in the Ariza et al. study (3).

Figure 5. Phase-contrast images of NSPCs stimulated at 0 (control), 25, and 50 V/m. NSPCs stimulated with addition of IFN-γ (left column) and EGF/ FGF (right column). Control neurites are shorter and show less mature characteristics as compared to stimulated groups. Scale bars = 50 μm.

In an attempt to gauge cell viability, metabolic activity of NSPCs was assayed before and after both stimulations for all treatment groups (Suppl. Fig. 3). Groups cultured with EGF/FGF resulted in a significant increase in metabolic activity, as compared to IFN- γ treated groups, which demonstrated comparably decreased metabolic activity (p < 0.0001). These results suggest that total cell metabolic activity was similar in 238

all treatments by day 3, but more focused on neuronal differentiation and neurite

extension in groups cultured in IFN-γ. It would be necessary to test NSPCs over a longer

time period to see if the viability gap between these groups becomes larger or significant

between electrically stimulated cells.

Figure 6. Representative 60X images of NSPC growth cones at day 3. (A) After no stimulation, tubulin structure is most noticeable. (B-C) After DC electrical stimulation regime at 25 V/m and 50 V/m, respectively, f-actin is prominently displayed as the finger-like projections that are characteristic of growth cones as tubulin is present in the shaft of the axon and center of the growth cone. Scale bar = 5 µm.

Other groups have shown that short duration DC EFs (10-60 min/day) result in

longer outgrowth in different cell types, such as chick embryo dorsal root ganglia,

possibly due to changes in regulation of growth factors (49, 46). Combining our results

with those of short duration stimulation regimes, it appears that short duration EF

stimulation in the range of 25-50 V/m (1.42 - 28.3 μA), approximately 10-60 min/day, is

all that is required to provide significant benefit in terms of neurite extension as well as neuronal differentiation and maturation of NSPCs. Furthermore, stimulating for too long or at higher voltages may result in negative effects such as a reduction in neurite length or unorganized morphology.

In this study, one of our goals was to determine if applied DC EFs act as a differentiation factor, thus we compared EF stimulation of NSPCs exposed to a known 239

neuronal differentiation factor (IFN-γ) to cells maintained in expansion medium

(EGF/FGF). Without EF stimulation, we have previously shown that IFN-γ stimulation of

SVZ-derived NSPCs generated a population containing greater than 60% neurons, whereas culture in EGF/FGF yielded a low percentage of neurons after 8 days (27). The experiments in this study showed that EF stimulation encouraged mature neuronal staining in both differentiation and growth media, particularly at 25 V/m and 50 V/m

(Fig. 3-4). These results suggested that EFs combined with appropriate growth factors can be used to both enhance and accelerate the actions of known neuronal differentiation factors, such as IFN-γ. Furthermore, EFs alone may provide a non-chemical means of inducing differentiation as shown by others (33, 7). Our finding of upregulated positive neuronal staining in EGF/FGF media plus 25 and 50 V/m show that DC EF stimulation can help to override mitogen signaling (Fig. 4). This is especially interesting when contrasting our findings with those of a recent study where mouse NSPCs in mitogens were stimulated with a salt-bridge at 250 V/m for 8 h. Babona-Pilipos et al. (4) found that NSPCs maintained progenitor markers, and preferred to migrate toward the cathode.

The mechanisms involved in EF cell differentiation is uncertain and could involve signaling pathways that regulate calcium flux or modification to proteins (12, 43). One of the most striking responses we observed in these experiments was the morphological difference between control groups compared to 25 V/m and 50 V/m groups (Figs. 3-6).

The emergence of prominent growth cones in our studies (Fig. 6) after only 2 d of stimulation (3 d total) may be indicative of more mature cells generated from adult

NSPCs in a relatively short period of time. Growth cones play an important role in the

240

function of the neuron by guiding it to its synaptic target. Since we know that EFs occur

endogenously within the body (38), it is quite possible that this may serve as an initiation cue for the growth cone.

Also of interest to us in this study was if DC EFs would encourage alignment of

NSPC neurites. Many cell types are known to preferentially migrate towards the cathode during long duration EF stimulation such as in human keratinocytes (44), HUVEC cells

(53), bovine aortic endothelial cells (29), and hippocampal-derived NSPCs (3).

Subependymal NSPCs and human embryonic derived NPSCs both exhibited directed migration toward the cathode when DC EFs were applied after 2.5 and 1 hr, respectively, and galvanotaxis immediately ceased when stimulation was removed (4, 13). Others have shown alignment where more neurites were directed toward the anode, although less often, such as seen with human granulocytes (41) and PC-12 cells (9). Our results (Fig. 2) agreed with similar studies where DRG do not show any conclusive evidence of alignment (49, 3, 22). This lack of alignment for NSPCs is most likely due to the short duration of stimulation, 10 min/d that we chose to employ. Further, reported differences among alignment preference between cell types could indicate that different pathways responsible for galvanotaxis vary according to the cells under consideration.

To better understand the mechanisms by which cells use endogenous EFs in development and injury models, the intervals at which retraction and growth occur in neurites must be revealed in future studies to formulate optimal timing of EF stimulation.

We have found that within the first minutes of stimulation, cells exposed to DC EFs can

undergo retraction in response to the EF; however, post stimulation cells extend neurites

241

rapidly (Suppl. Vid. 2). This result has been confirmed in studies with developing

amphibians where neural crest cells show retraction within seconds of stimulation

followed by accelerated growth (38). In future live imaging EF stimulation experiments it

would be instructional to monitor the entire culture period to better understand the

dynamics and timing of cellular responses. Overall, the observed response to EF

stimulation from 25-50 V/m is positive based on the extreme lengths and mature neuronal

phenotypes that this DC EF stimulation encourages after 3 d (Fig. 1-7).

Figure 7. Ca2+ flux experiments in DC electrically stimulated NSPCs. (A) Graphs and images of Ca2+ flux recordings in the soma and a neurite of a single selected NSPC (cultured in IFN-γ for 3 d) stimulated at 25 V/m. (B) Graphs and images of Ca2+ flux recordings from soma and neurite of a selected NSPC cultured in IFN-γ for 3 d stimulated at 25, 50, and 100 V/m in a ramping step-wise fashion. Videos of each experiment (A, B) are included as supplemental data (Suppl. Vids. 4, 5).

To date, we are unaware of the exact cause of why EFs are beneficial to neurons, but there are many suggested mechanisms by which extension of neurites occur in response to EFs. More specifically, mobility of the growth cone and extension/retraction of neurites is strongly linked to the influx of calcium into the cell, caused by 242

depolarization (8). Increased depolarization by DC EFs increased intracellular calcium, as

we show in our system (Fig. 7). Even though calcium changes were variable and

sometimes small in magnitude in some parts of the cell, these variation are still important

as calcium fluctuations are known to activate several important downstream cellular

mechanisms including neuron extension, differentiation and plasticity (2, 16). For

example, calcium is known to be partially responsible for cadherin based cell attachment,

transport of organelles, and vesicle fusion, which all can dramatically affect the

outgrowth of neurites and formation of growth cones (24).

In our experiments, we were interested to find if differentiating NSPCs would exhibit calcium signaling in response to DC stimulation as would be expected from primary neurons. Overall the observed calcium fluctuations of stimulated cells demonstrated electrical functionality (Fig. 7, Suppl. Vids. 4-5). Generation of an action potential in cells has been shown to be created by an applied extracellular current

(20).We saw that NSPCs demonstrated immediate calcium flux increases when DC EFs were applied (Fig. 7), with the largest change in intensity occurring in the neurite at 25

V/m. The changes in calcium levels are similar in both the soma and the neurite when the

EF was increased from 25 V/m to 100 V/m. Previous work has revealed that neuronal somas contain higher concentrations of calcium channels relative to concentrations found in axons (5). In our ramped stimulation experiment, increasing the applied voltage from 0 to 100 V/m (at 25 V/m each min) resulted in increasing calcium flux until reaching 100

V/m, where an almost immediate drop of flux was noted (Fig. 7). In compendium, the application of 100 V/m DC resulted in decreased f-actin and tubulin organization, shorter

243

neurite lengths, and decreased calcium flux, suggesting that 100 V/m results in cellular

damage and interference with the cell’s ability to properly regulate calcium flux, which is vital to many neuronal signal transduction pathways and processes (5, 20). As is explained by Kater and Mills, there is suggested to be an optimal level of intracellular calcium for growth cone function and promotion of neurite extension (21). We believe that exogenous EFs play a vital role in tuning intracellular calcium levels (Fig. 7). From our studies 25 V/m and 50 V/m short duration EF results in significantly longer neurites compared to controls (Fig. 1). In addition, in other work the filopodia guiding growth cone movements have been shown to retract initially from an increase in intracellular calcium, followed by a recovery period where normalization of calcium levels by cellular mechanisms occurs (42). This agrees with what we have seen with initial periods of growth cone retraction (Suppl. Vid. 2, available online) followed by neurite extension

(Suppl. Vid. 1, available online). With the addition of an outside stimulus, i.e., EFs of 25 and 50 V/m, it is possible that intracellular calcium was increased to more preferred levels for neurite extension. When EFs of 100 V/m were applied in short-term experiments, available calcium was above the permissive range, potentially halting neurite extension.

There is great debate over the effectiveness of EFs as a regenerative strategy, largely due to a lack of agreed upon stimulation type (DC/AC) and regime (exposure time, voltage, etc.). Thus significant work remains to be completed to better understand the phenomena to translate these findings. In peripheral nerves of the adult guinea-pig, it

shown there is no correlation between the use of a DC EF (20 μA) and regeneration (36).

244

Again, in the rat sciatic nerve encased in silicon tubes, weak EFs of 10 μA applied in vivo failed to not only heal the injury but to actually inhibit regrowth (17). Conversely, studies have found that DC EF stimulation (1 μA) has been effective in healing and improving recovery in the rat sciatic nerve (37). In our study, we have found that DC EFs are beneficial to NSPCs in vitro as seen by an increase in neurite extension and better neuronal differentiation, which could be valuable for nervous system tissue engineering type approaches incorporating stem cells that require axonal extension.

Conclusions

The effects of short duration DC EF stimulation of NSPCs in vitro in combination with EGF/FGF or IFN-γ produced morphologically mature neurons with longer neurite lengths in a relatively short time period (<10 min/d for 2 d) compared to no stimulation.

We have found that DC stimulation of neuronally-differentiated NSPCs caused an elevated influx of calcium into the cell within seconds, and that there appears to be a current/voltage threshold for which these cells can endure. Live imaging during stimulation revealed that DC EFs can cause retraction before elongation, but cells remain viable and active during this stimulation regime. This result along with the appearance of mature markers, growth-cone characteristics, and rapid extension of NSPCs suggests that using EFs could provide a potential new solution to CNS regeneration. We hope to apply these results to neural regenerative strategies in the near future.

245

Acknowledgements

We would like to thank The University of Akron for the funding that supported

this work.

Supporting Information

Supplemental videos are described below; links to these videos are available online at https://link.springer.com/article/10.1007%2Fs10439-014-1058-9.

Supplemental Video 1. Live imaging videos of cells stimulated at 25 V/m with addition of IFN-γ (2 d culture). Video shows initial retraction seen in some cells at start of stimulation. Video time is 30 min, with voltage turned on at 10 min and turned off at 20 min.

Supplemental Video 2. Live imaging videos of cells stimulated at 25 V/m under addition of IFN-γ. Video shows elongation of neurite following stimulation for 10 min. Video time is 30 min, with voltage turned on at 10 min and turned off at 20 min.

Supplemental Video 3. Live imaging videos of cells with no stimulation with addition of

IFN-γ (2 d culture). Video time is 30 min, with voltage turned on at 10 min and turned off at 20 min.

Supplemental Video 4. Calcium flux videos of cells stimulated at 25 V/m as summarized in Fig. 7. Video shows increase in calcium permeability during stimulation and decreasing after saturation and cessation of stimulation.

Supplemental Video 5. Calcium flux videos of cells stimulated at with voltage increased step-wise ramp from 25 to 100 V/m as summarized in Fig. 7. Videos show stimulation

246

increasing calcium permeability during stimulation then sharply decreasing shortly after

100 V/m is initiated.

References

1. 2011. Spinal Cord Injury Facts at a Glance. National Spinal Cord Injury Statistical Center, 2 pp. July, 2013. https://www.nscisc.uab.edu. 2. Anglister, L., Farber, I. C., Shahar, A. and Grinvald, A."Localization of Voltage- Sensitive Calcium Channels Along Developing Neurites: Their Possible Role in Regulating Neurite Elongation." Developmental Biology 94(2): 351-365, 1982. 3. Ariza, C. A., Fleury, A. T., Tormos, C. J., Petruk, V., Chawla, S., Oh, J., Sakaguchi, D. S. and Mallapragada, S. K."The Influence of Electric Fields on Hippocamapal Neural Progenitor Cells." Stem Cell Review and Repair 6(4): 585-600, 2010. 4. Babona-Pilipos, R., Droujinine, I. A., Popovic, M. R. and Morshead, C. M."Adult Subependymal Neural Precursors, but Not Differentiated Cells, Undergo Rapid Cathodal Migration in the Presence of Direct Current Electric Fields." PLoS ONE 6(8), 2011. 5. Bolsover, S. and Spector, I."Measurements of Calcium Transients in the Soma, Neurite, and Growth Cone of Single Cultured Neurons " Journal of Neuroscience 6(7): 1934-1940, 1986. 6. Bovolenta, P., Wandosell, F. and Nieto-Sampedro, M."CNS Glial Scar Tissue- A Source of Molecules Which Inhibit Central Neurite Outgrowth." Progress in Brain Research 94: 367-379, 1992. 7. Chang, K. A., Kim, J. W., Kim, J. A., Lee, S. E., Kim, S., Suh, W. H., Kim, H. S., Kwon, S., Kim, S. J. and Suh, Y. H."Biphasic Electrical Currents Stimulation Promotes Both Proliferation and Differentiation of Fetal Neural Stem Cells." PLoS One 6(4): e18738, 2011. 8. Connor, J. A."Digital Imaging of Free Calcium Changes and of Spatial Gradients in Growing Processes in Single, Mammalian Central Nervous System Cells." Proc. Natl. Acad. Sci 83(16): 6179-6183, 1986. 9. Cork, R. J., McGinnis, M. E., Tsai, J. and Robinson, K. R."The Growth of PC-12 Neurites Is Biased Towards the Anode of an Applied Electrical Field." Developmental Neurobiology 25(12): 1509-1516, 1994. 10. Craig, C., Tropepe, V., Morshead, C., Reynolds, B., Weiss, S. and Kooy, D. v. d."In Vivo Growth Factor Expansion of Endogenous Subependymal Neural Precursor Cell Populations in the Adult Mouse Brain." Journal of Neuroscience 16(8): 2649-2658, 1996. 11. Fawcett, J. and Asher, R."The Glial Scar and Central Nervous System Repair." Brain Research Bulletin 49(6): 377-391, 1999. 12. Felaco, M., Reale, M., Grilli, A., De Lutiis, M. A., Barbacane, R. C., Di Luzio, S. and Conti, P."Impact of Extremely Low Frequency Electromagnetic Fields on CD4 Expression in Peripheral Blood Mononuclear Cells." Mol Cell Biochem 201(1-2): 49-55, 1999. 247

13. Feng, J.-F., Liu, J., Zhang, X.-Z., Zhang, L., Jiang, J.-Y., Nolta, J. and Zhao, M."Brief Report: Guided Migration of Neural Stem Cells Derived from Human Embryonic Stem Cells by an Electric Field." Stem Cells 30(2): 349-355, 2012. 14. Fitch, M., Doller, C., Combos, C., Landreth, G. and Silver, J."Cellular and Molecular Mechanisms of Glial Scarring and Progressive Cavitation: In Vivo and in Vitro Analysis of Inflammation-Induced Secondary Injury After CNS Trauma." Journal of Neuroscience 19(19), 1999. 15. Frohlich, F. and McCormick, D. A."Endogenous Electric Fields May Guide Neocortical Network Activity." Neuron 67(1): 129-143, 2010. 16. Ghosh, A. and Greenberg, M. E."Calcium Signaling in Neurons: Molecular Mechanisms and Cellular Consequences." Science 268, 1995. 17. Hanson, S. M. and McGinnis, M. E."Regeneration of Rat Sciatic Nerves in Silicone Tubes: Characterization of the Response to Low Intensity D.C. Stimulation." Neuroscience 58(2): 411-421, 1994. 18. Haugland, R. P. “The Handbook: A Guide to Fluorescent Probes and Labeling Technologies.” Invitrogen Corp, 2005, 1126 pp. Edited by Michelle T. Z. Spence. 19. Hotary, K. B. and Robinson, K. R."Endogenous Electrical Currents and the Resultant Voltage Gradients in the Chick Embryo." Developmental Biology 140(1): 149- 160, 1990. 20. Jimbo, Y., Robinson, H. P. C. and Kawana, A."Simultaneous Measurement of Intracellular Calcium and Electrical Activity from Patterned Neural Networks in Culture." IEEE Transactions on Biomedical Engineering 40(8): 804-810, 1993. 21. Kater, S. B. and Mills, L. R."Regulation of Growth Cone Behavior by Calcium." The Journal of Neuroscience 11(4): 891-899, 1991. 22. Koppes, A. N., Seggio, A. M. and Thompson, D. M."Neurite Outgrowth Is Significantly Increased by the Simultaneous Presentation of Schwann Cells and Moderate Exogneous Electric Fields." Journal of Neural Engineering 8(4), 2011. 23. Kuhn, H. G., Winkler, J., Kempermann, G., Thai, L. J. and Gage, F. H."Epidermal Growth Factor and Fibroblast Growth Factor-2 Have Different Effects on Neural Progenitors in the Adult Rat Brain." The Journal of Neuroscience 17(15): 5820-5829, 1997. 24. Lankford, K. L. and Letourneau, P. C."Evidence That Calcium May Control Neurite Outgrowth by Regulating the Stability of Actin Filaments." The Journal of Cell Biology 109(3): 1229-1243, 1969. 25. Leipzig, N. D., Xu, C., Zahir, T. and Shoichet, M. S."Functional Immobilization of Interferon-Gamma Induces Neuronal Differentiation of Neural Stem Cells." Journal of Biomedical Materials Research Part A 93A(2): 625-633, 2010. 26. Leipzig, N. D., XU, C., Zahir, T. and Shoichet, M. S."Differentiation of Neural Stem Cells in Three-Dimensional Growth Factor-Immobilized Chitosan Hydrogel Scaffolds." Biomaterials 32(1): 57-64, 2011. 27. Li, H., Wijekoon, A. and Leipzig, N. D."3d Differentiation of Neural Stem Cells in Macroporous Photopolymerizable Hydrogel Scaffolds." PLoS ONE 7(11): e48824, 2012.

248

28. Li, L. and Jiang, J."Stem Cell Niches and Endogenous Electric Fields in Tissue Repair." Frontiers of Medicine 5(1): 40-44, 2011. 29. Li, X. and Kolega, J."Effects of Direct Current Electric Fields on Cell Migration and Actin Filament Distribution in Bovine Vascular Endothelial Cells." Journal of Vascular Research 39(5): 391-404, 2001. 30. Lipton, S. A."Blockade of Electrical Activity Promotes the Death of Mammalian Retinal Ganglion Cells in Culture." Neurobiology 83(24): 9774-9778, 1986. 31. Lois, C. and Alvarez-Buylla, A."Long-Distance Neuronal Migration in the Adult Mammalian Brain." Science 264(5162): 1145-1148, 1994. 32. Lois, C. and Alvarezbuylla, A."Proliferating Subventricular Zone Cells in the Adult Mammalian Forebrain Can Differentiate into Neurons and Glia." Proceedings of the National Academy of Sciences of the United States of America 90(5): 2074-2077, 1993. 33. Matos, M. A. and Cicerone, M. T."Alternating Current Electric Field Effects on Neural Stem Cell Viability and Differentiation." Biotechnol Prog 26(3): 664-670, 2010. 34. McCaig, C. D. and Rajnicek, A. M."Electrical Fields, Nerve Growth and Nerve Regeneration " Experimental Physiology 76(4): 473-494, 1991. 35. McCaig, C. D., Song, B. and Rajicek, A. M."Electrical Dimensions in Cell Science." Journal of Cell Science 122: 4267-4276, 2009. 36. McGinnis, M. E. and Murphy, D. J."The Lack of an Effect of Applied D.C. Electric Fields on Peripheral Nerve Regeneration in the Guinea Pig." Neuroscience 51(1): 231-244, 1992. 37. Mendonca, A. C., Barbieri, C. H. and Mazzer, N."Directly Applied Low Intensity Direct Electric Current Enhances Peripheral Nerve Regeneration in Rats." Journal of Neuroscience Methods 129(2): 183-190, 2003. 38. Mycielska, M. E. and Djamgoz, M. B. A."Cellular Mechanisms of Direct-Current Electric Field Effects: Galvanotaxis and Metastatic Disease." Journal of Cell Science 117(Pt 9): 1631-1639, 2004. 39. Politis, M. J., Zanakis, M. F. and Albala, B. J."Facilitated Regeneraton in the Rat Peripheral Nervous System Using Applied Electric Fields." Journal of Trauma-Injury Infecion & Critical Care 28(9), 1988. 40. Pullar, C. E. The Physiology of Bioelectricity in Development, Tissue Regeneration, and Cancer, 2011, 342 pp. Boca Raton, FL: abCRC Press, 41. Rapp, B., A., d. B.-C. and Gruler, H."Galvanotaxis of Human Granulocytes." European Biophysics Journal 16(5): 313-319, 1988. 42. Rehder, V. and Kater, S. B."Regulation of Neuronal Growth Cone Filopodia by Intracellular Calcium." The Journal of Neuroscience 12(8): 3175-3186, 1992. 43. Rollwitz, J., Lupke, M. and Simko, M."Fifty-Hertz Magnetic Fields Induce Free Radical Formation in Mouse Bone Marrow-Derived Promonocytes and Macrophages." Biochim Biophys Acta 1674(3): 231-238, 2004. 44. Sheridan, D. M., Isseroff, R. R. and Nuccitelli, R."Imposition of a Physiologic Dc Electric Field Alters the Migratory Response of Human Keratinocytes on Extracellular Matrix Molecules." Journal of Investigative Dermatology 106(4): 642-646, 1996.

249

45. Shi, R. and Borgens, R. B."Three-Dimensional Gradients of Voltage During Development of the Nervous System as Invisible Coordinates for the Establishment of Embryonic Pattern." Developmental Dynamics 202: 101-114, 1995. 46. Wan, L., Xia, R. and Ding, W."Low-Frequency Electrical Stimulation Improves Neurite Outgrowth of Dorsal Root Ganglion Neurons in Vitro Via Upregulating Ca2+- Mediated Brain-Derived Neurotrophic Factor Expression." Neural Regeneration Research 5(16): 1256-1260, 2010. 47. Whittemore, S. R., Morassutti, D. J., Walters, W. M., Liu, R. H. and Magnuson, D. S. K."Mitogen and Substrate Differentially Affect the Lineage Restriction of Adult Rat Subventricular Zone Neural Precursor Cell Populations." Experimental Cell Research 252(1): 75-95, 1999. 48. Wilkinson, A. E., McCormick, A. M. and Leipzig, N. D."Central Nervous System Tissue Engineering: Current Considerations and Strategies." Synthesis Lectures on Tissue Engineering 8: 1-112, 2011. 49. Wood, M. and Kuntz Willits, R."Short-Duration, DC Electrical Stimulation Increases Chick Embryo Drg Neurite Outgrowth." Bioelectromagnetics 27: 328-331, 2006. 50. Yang, G., Long, H., Wu, J. and Huang, H. (2008). A Novel Electrical Field Bioreactor for Wound Healing Study. International Conference on Biomedical Engineering and Informatics. 51. Yiu, G. and He, Z."Glial Inhibition of CNS Axon Regeneration." Nature Reviews Neuroscience 7(8): 617-627, 2006. 52. Zahir, T., Nomura, H., Guo, X., Kim, H., Tator, C., Morsehead, C. and Shoichet, M."Bioengineering Neural Stem/Progenitor Cell-Coated Tubes for Spinal Cord Injury and Repair." Cell Transplant 17(3): 245-254, 2008. 53. Zhao, M., Bai, H., Wang, E., Forrester, J. V. and McCaig, C. D."Electrical Stimulation Directly Induces Pre-Angiogenic Responses in Vascular Endothelial Cells by Signaling through VEGF Receptors." Journal of Cell Science 117(Pt. 3): 397-405, 2003.

250