Scaffolds with Oriented Microchannels for Angioneural Regenerative Engineering By

Scaffolds with Oriented Microchannels for

Angioneural Regenerative Engineering

A Thesis

Submitted to the Faculty

Drexel University

Aybike Saglam in partial fulfillment of the requirements for the degree

Master of Science in Biomedical Engineering

December 2010

To my father…

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 ACKNOWLEDGMENTS

It is very difficult to express how grateful I am to the people who contributed to this project.

First of all, I want to thank to my committee members: Dr. Peter I. Lelkes, Dr.

Philip Lazarovici, Dr. Anat Perets Katsir and Dr. Fred Allen. This thesis would have been impossible without all of you. Dr. Lelkes was a great advisor and mentor who was a big influence in my academic life. Dr. Perets was an amazing advisor and friend with constant optimism and encouragement. Dr. Lazarovici was an inspiration for good academic character and integrity and I hope to be as good an academian some day. Dr. Allen was a particularly positive source of encouragement in the beginning of my degree. Special thanks to Dr. Onaral, Dean of Biomedical

Engineering, Science, and Health Systems, for serving as such a positive and tenacious role model for me throughout my degree.

Thanks to many individuals and groups who have helped me considerably along the way: Dr. Itzhak Fisher,Dr. Gianluca Gallo, and to their laboratory members for their help on primary neuron extraction and culture, particularly Dr. Katherine

Kollins; Dr. J. Yasha Kresh and Anant for letting me use the rheometer even for extended hours; Dr. Yvonne Mueller for her help on Fluorescence Activated Cell

Sorting technique; and Dr. Qingwei Zhang and Gozde Senel for their help on

Scanning Electron Microscopy.

Thanks, too, to my friends and labmates: Mie was a great friend, advisor, mentor, roommate, classmate, and labmate; Greg, Diane and Leko who were never too busy

to help me; Adam who was personally involved in some of my experiments and

designed the controlled freeze-drying machine with Ho-Lung Li according to Dr.

Peret’s directives; my “sisters,” Gozde, Seda and particularly Asli made me feel at

home; our program manager, Susan; our program coordinator, Tantri; Dr. Har-el was

a source of advice and support; labmates Mike, Collin, Patrick, Gaurav, Sean, Jingjia,

Tracy, Jessica, and Anna; and finally, Paul for fixing the lyophilization machine.

Thanks to many institutions and programs on campus for their logistical support: the Biomed office, specifically Aylin, Natalia, Danielle and Lisa; and the ISSS office, especially to Adrienne.

Finally, but not least, to my family and friends to whom I am grateful for their support.

 TABLE OF CONTENTS  ACKNOWLEDGMENTS ...... iv  TABLE OF CONTENTS ...... vi  List of Figures ...... vii  ABSTRACT ...... viii  INTRODUCTION ...... 1 .1 Overall goal & hypothesis ...... 2 .2 Aims of the thesis ...... 3 .3 Definitions ...... 4 .4 Rationale ...... 4  BACKGROUND ...... 8 .1 Past & current tissue engineering strategies for SCI ...... 8 .2 Spinal cord organization ...... 10 .3 Spinal cord injury ...... 11 .4 Overview on Scaffolds for SCI ...... 12 .5 Hydrogels for engineered scaffolds ...... 16 .6 Freezedrying techniques ...... 16 .7 The effect of pore diameter on axon extension and ECM secretion ...... 19 .8 Angiogenesis & vasculogenesis: ...... 20 .9 Neuritogenesis ...... 22 .10 Neurovascular crosstalk ...... 24 .10.1 Interactions between nervous & vascular systems: ...... 24 .11 Crosstalk between neurons and ECs ...... 25 .11.1 Angiogenic factors’ effects on the nervous system: ...... 27 .11.2 Neurogenic factors’ effects on the vascular system ...... 28  MATERIALS AND METHODS ...... 30 .1 Preparation of genipin crosslinked gelatin polymer (GCG) ...... 30 .2 Scaffold production by regular and controlled freezedrying ...... 30 .3 Measurements of mechanical properties of the scaffold ...... 33 .4 Invitro degradation of the GCGCFDSs ...... 33 .5 Cells ...... 34 .6 Cell proliferation ...... 37 .7 Fluorescence activated cell sorting (FACS) ...... 37 .8 Endothelial cell labeling ...... 38 .9 Populating the scaffold with cells and treatment with angioneurins ...... 39 .10 Confocal Microscopy ...... 39 .11 Scanning Electron Microscopy ...... 40 .12 Statistics ...... 40  RESULTS & DISCUSSION ...... 41 .1 The morphology of the scaffolds ...... 41 .2 The mechanical properties of the scaffolds ...... 43 .3 The degradability of the GCGCFDS in the serum ...... 44 .4 Biocompatibility of the GCGCFDS for homogeneous cell populations ...... 45 .5 Biocompatibility of the GCGCFDS for heterogeneous cell populations ...... 49  SUMMARY ...... 53  CONCLUSION& FUTURE WORK...... 56  LIST of REFERENCES ...... 60  APPENDIX ...... 67 .1 Evaluate the effects of NGF & FGF2 on ECs & PC12 cells ...... 67 .2 Evaluate the coculture proliferation dynamics for both endothelial and neuronal cells ...... 68 .3 Protocols ...... 70

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 List of Figures

Figure 1 ...... 18 Figure 2 ...... 27 Figure 3 ...... 32 Figure 4 ...... 35 Figure 5 ...... 36 Figure 6 ...... 38 Figure 7 ...... 42 Figure 8 ...... 43 Figure 9 ...... 43 Figure 10 ...... 45 Figure 11 ...... 46 Figure 12 ...... 47 Figure 13 ...... 49 Figure 14 ...... 50 Figure 15 ...... 51 Figure 16 ...... 52 Figure 17 ...... 57 Figure 18 ...... 67 Figure 19 ...... 69

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Scaffolds with Oriented Microchannels for Angioneural Regenerative Engineering by

Aybike Saglam

Submitted to the Department of Biomedical Engineering on December 30, 2010 in partial fulfillment of the requirements for the degree of Master of Science in Biomedical Engineering

 ABSTRACT

Spinal cord injury (SCI) affects over 12,000 people annually and, due to limited therapeutic options, results in an estimated 1.3 million people living with chronic motor dysfunction or paralysis in the U.S. The medical costs associated with SCI can pose significant financial burden to patients (estimated at 2 million dollars in a lifetime-NSCICS). Finding a therapeutic strategy for repair of SCI would dramatically improve quality of life for these patients as well as ease the costs associated with chronic care. Prior studies have shown that on non-structured substrates, oriented axonal growth is very poor; axons tend to grow erratically and axon connection pathways cannot find and connect to their target neurons. Additionally spinal cord development, and presumably spinal cord regeneration requires angiogenesis for nutrient delivery as well as regulation of neuronal growth through paracrine signaling. This is supported by previous studies that showed increased micro-vessel density correlates with regeneration and functional recovery in spinal cord tissue. Based on these finding this study focuses on combining structural support for directed axonal growth with angiogenic cues for the purpose of enhancing SCI repair. It hypothesizes that the crosstalk between the vascular and the nervous systems, i.e., between cues that lead to angiogenesis and neuritogenesis is crucial for axonal regeneration across a spinal cord lesion. To test this hypothesis, a scaffold is evaluated for promoting neuritogenesis in vitro by seeding its longitudinally oriented 3D pores with endothelial cells prior to seeding with neural-crest derived PC12 cells.

This study designed and tested a biodegradable, genipin cross-linked gelatin scaffold by using a controlled uniaxial freezing technique followed by lyophilization to improve mechanical properties of the scaffold and to generate longitudinal microporous channels. The scaffold had mechanical properties similar to those of the rat spinal cord with elastic moduli of (~51kPa), and it supported growth and differentiation of endothelial cells and PC12 cells. An “angioneural tissue” was generated by first seeding endothelial cells in the scaffold to generate a monolayer lining the channel walls, followed by seeding with neuronal PC12 cells a week later. In support of the hypothesis, the presence of endothelial cells facilitated the differentiation of nerve growth factor (NGF)-induced PC12 cells significantly leading to enhanced PC12 cell survival and neuritogenesis. Our results are promising for the promotion of neuritogenesis through endothelial cell-derived cues (angioneural crosstalk, i.e., concomitantly promoting angiogenesis and neuritogenesis), which may facilitate tissue regeneration and recovery after spinal cord injury.

Thesis Supervisors:

Dr. Peter I. Lelkes1

Dr. Philip Lazarovici1,2

Dr. Anat Perets Katsir1

Dr. Fred Allen1

1School of Biomedical Engineering, Drexel University, Philadelphia, PA, USA

2School of Pharmacy Institute for Drug Research, Faculty of Medicine, The Hebrew

University of Jerusalem, Jerusalem, Israel

 INTRODUCTION

Injury to the nervous system is a major health problem worldwide. The limited regenerative potential of the adult nervous system after injury often causes chronic sensory or motor function failures as well as neuropathic pains. In the peripheral nervous system (PNS), surgical end-to-end reconnections, nerve autographs, allografts and xenografts are common strategies to improve regeneration after nerve transection injuries (Trumble et al., 2000; Evans et al., 1994; Platt et al., 1990). However autographs can be effective only for small gaps, often require multiple surgeries and can cause function loss at donor sites. On the other hand allografts and xenografts

(donor tissue from cadavers and animals, respectively) can cause disease transfer and immunological rejections (Cao et al., 2009). Moreover, none of these methods are effective enough to help regeneration in the central nervous system (CNS) (Fawcett et al., 1999).

The regeneration in the CNS, including brain and spinal cord, is very limited and complicated as innate factors exist within the body that inhibit repair; therefore it is a great challenge to researchers studying CNS therapies. Millions of people suffer from

CNS injury annually and as a result, most of them endure irreversible disabilities.

Among these, spinal cord injury (SCI) affects over 12,000 people annually (National

Spinal Cord Injury Statistical Center- NSCICS) and due to limited therapeutic options, results in an estimated 1.3 million people living with chronic motor dysfunction or paralysis in the U.S, (report available at www.christopherreeve.org), making the search for novel avenues of SCI repair a critical area of research. Moreover the medical costs associated with SCI can pose significant financial burden to patients

(estimated at $2 million dollars in a lifetime-NSCICS). Therefore finding a therapeutic strategy for repair of SCI, a focus point of this project, would dramatically improve the quality of life for patients as well as ease the costs associated with chronic SCI care.

.1 Overall goal & hypothesis

Unfortunately, to date, complete recovery after spinal cord injury has not been achieved. As stated above, most of the regenerative strategies for the PNS are focused on alternatives to the nerve graft. Comparatively, strategies for SCI require more complex designs that necessitate a permissive environment for regeneration (Schmidt et al., 2003). Therefore, to achieve a successful neural regeneration, specifically for

CNS, there remains a need for a biodegradable and biocompatible tissue engineered scaffold, which mimics the natural tissue’s architecture as well as biochemical and mechanical properties (Taipale et al., 1997; Berthiaume et al., 1996).

This project’s approach on engineering a scaffold for future neuronal tissue repair relies on combining structural guidance with biocompatibility for concomitant growth of two different populations of cells: neurons and endothelial cells (ECs). A degradable, genipin cross-linked gelatin scaffold was designed by using a controlled freeze-drying technique in an effort to improve the mechanical properties of the scaffold and to give it a longitudinal structure that would guide axonal extensions throughout the scaffold in the presence of ECs. The hypothesis is that the ECs will induce neuronal cell survival and differentiation. We propose to exhibit neuronal differentiation using PC12 cells and FBNs in an endothelial cell permissive environment and to test the ability of this novel approach as a promising starting point

3 for future spinal cord injury repair studies.

.2 Aims of the thesis

Specific Aim 1: To characterize properties of genipin-crosslinked-gelatin scaffolds

for various methods of fabrication.

 Specific Aim 1.1: Rheologically characterize the viscoelastic properties of

the scaffolds.

 Specific Aim 1.2: Measure the pore-size of the scaffolds for different

concentrations of genipin crosslinker.

 Specific Aim 1.3: Evaluate the in vitro degradation of the scaffolds.

Specific Aim 2: To show the effects of angiogenesis on neuronal growth in a model

system.

 Specific Aim 2.1: Evaluate the cell biocompability of the optimal scaffold

for neurons and endothelial cells individually.

 Specific Aim 2.2: Establish angioneural CFDSs by seeding ECs and

neuronal cells in combination.

Specific aim 2.2.1: Evaluate the optimum cell seeding density.

 Specific Aim 2.3: Quantify and compare neuritogenesis in mono-cultures

and co-cultures.

.3 Definitions

PC12- pheochromocytoma cells; GFP-PC12- green fluorescence protein transduced PC12 cells; EAhy926- a hybrid cell line obtained by fusion of human umbilical vein endothelial cell line and lung carcinoma cell line, FBN- chicken forebrain primary neurons; SEM- scanning electron microscope; GCG- genipin crosslinked gelatin; DMEM- Dubelco’s modified eagle medium, Medium A- DMEM supplemented with 7.5% fetal bovine serum FBS, 7.5% horse serum; Medium B-

Dulbecco’s modified Eagle’s medium supplemented with 10% FBS; E- a parameter describing the degree of neurite extension; N- a parameter describing percentage of cells with neurites, FBS- fetal bovine serum; CFDS- controlled freeze dried scaffold;

RFDS-regular freeze dried scaffold; 2D- two dimensional; 3D- three dimensional;

ECM- extracellular matrix; CNS- central nervous system; PNS-peripheral nervous system; GFs- growth factors; BSCB- blood spinal cord barrier; NSCISC- National

Spinal Cord Injury Statistical Center; BDNF- brain-derived neurotrophic factor;

GDNF- Glial cell-derived neurotrophic factor; VEGF- vascular endothelial growth factor

.4 Rationale

An ideal scaffold for neuronal tissue engineering needs to mimic endogenous neural tissue according to structural geometry, mechanical properties and chemical composition. Numerous degradable natural and synthetic polymers of different composition such as collagen, gelatin, fibronectin, alginate, and poly-glycolic acid/ploy-lactic acid (PGA/PLA) are considered candidates for nerve repair strategies

(Nomura et al., 2006). Gelatin was the first biodegradable material to be tested as a

5 nerve guide (IJkema-Paassen et al., 2004) and was considered a good candidate matrix for production of present controlled-freeze dried scaffolds (CFDSs). Gelatin is produced by thermal denaturation or chemical degradation of collagen, which is the major component of the extracellular matrix (ECM), and is advantageous since it is a biodegradable polymer with adequate biocompatibility, plasticity and adhesiveness and does not express antigenicity. The mechanical and chemical properties of gelatin scaffolds can be optimized by crosslinking with genipin. Genipin is a natural compound derived from geniposide (Bigi et al., 2002), which has been demonstrated to exhibit neurite outgrowth inducing activity in PC12 cells (Liu et al., 2006), chicken dorsal root ganglia cultures (Sundararaghavan et al., 2009) and confers neuroprotection in hippocampal neurons (Lee et al., 2006). In view of these reports, it was considered as the cross linker for preparation of present CFDSs.

One of the strategies for the repair of neural tissue lesions, specifically for SCI, considers the use of porous scaffolds prepared by freeze-drying and then populating this scaffold with neuronal cells (Rauch et al., 2009). This type of scaffold, upon implantation, should provide tropic support and guidance to the injured neuronal tissue and allow regeneration (Mollers et al., 2009). In addition, a porous scaffold will provide a sufficient surface for cell seeding, a place for their proliferation and enable

ECM deposition (Thomson et al., 2000; Patrick et al., 1998). However, the development of clinically relevant, porous scaffolds for neuronal tissue engineering remains a tremendous challenge.

Studies show that on non-structured substrates, longitudinal axonal regrowth is very poor; axons tend to grow erratically and axon connection pathways cannot find

6 and connect to their target neurons. Smeal et al., (2005) mentions that, at early developmental stages of embryogenesis, a natural mechanism might be formed in order to guide axons. In the initial stages of development, axons are guided by chemotaxis over short distances to their targets. At later stages of development, axons traverse longer distances and trace these cylindrical substrates, called growth pathways. It is currently uncertain how these structures provide guidance (Smeal et al.,

2005). But this knowledge gives us the insight that an optimum scaffold for the injured neural tissue requires a guiding structure, which allows biocompatible cell seeding.

Recently a few groups reported an effective, novel technology to generate porous materials by using a directional freeze-drying procedure (Thomson et al., 2000), which contains pores in the form of longitudinal channels (Stokols et al., 2004; Bozkurt et al., 2007; Mollers et al., 2009; Wu et al., 2010). In contrast to porous non-structured scaffolds, where the neuronal cells are restricted in their ability to generate organized pathways, channeled micro-structured scaffolds allow the neurons to develop long axonal pathways enabling synapse connections (Bozkurt et al., 2009). The overarching purpose of this project was to further improve this approach and to generate a freeze- dried scaffold with longitudinal channels by using an in-house designed device allowing controlled, gradient temperature decrease during the freezing process. Our goal is to achieve a constant cooling rate since there is a relationship between cooling rate and unidirectional (unbranched) channel formation as well as pore diameter.

Another critical process important for the integration of the implanted scaffold into the neuronal tissue relates to angiogenesis. Angiogenesis is the formation of blood vessels, either from ECs populating the scaffold (Rauch et al., 2009) or endothelial

7 stem cells penetrating into the scaffold from the peripheral blood upon implantation

(Allen et al., 2010). It has previously been shown that increased micro-vessel density correlates with regeneration and functional recovery in spinal cord tissue (Glaser et al.,

2006; Kaneko et al., 2006). More importantly, regenerating axons grow along blood vessels in the spinal cord (Bearden et al., 2005), which indicate that restoration of angiogenesis would help neuronal cell guidance. These results coupled with the similarity in action of angiogenic and neurotrophic factors along with the presence of their receptors in vascular ECs and neuronal cells (Lazarovici et al., 2006) exemplify the crosstalk between angiogenesis and neurogenesis. These factors emphasize the important role of including ECs in generating scaffolds for neural engineering.

Therefore the new generation of scaffolds needs to be biocompatible for both ECs and neuronal cells. Furthermore, treatment with angioneurins (Zacchigna et al., 2008) such as fibroblast growth factor-type 2 (FGF-2) and nerve growth factor (NGF) has been shown to be obligatory for the survival and generation of both neurons (Moore et al., 2007) and capillary-like structures in the scaffold (Perets et al., 2003).

Cumulatively, these considerations call for proof of concept studies in the preparation of novel complex scaffolds with populations of both endothelial and neuronal cells.

 BACKGROUND .1 Past & current tissue engineering strategies for SCI

Axonal regeneration in the nervous system is subjected to many cellular and environmental difficulties. Injured nerve cells are incapable of regenerating spontaneously, particularly those in the CNS, because of specific inhibitory factors

(explained in the SCI section) released post-injury. In the 1990’s, researchers thought

CNS regeneration to be impossible (Feigin et al., 1951). However, over the past two decades, studies have shown that neurons in the CNS can regenerate axons after trauma, although limited, (Aguayo et al., 1981; Bray et al., 1987; Fawcett et al., 1998) when treated with neurotrophic factors (Richardson et al., 1991), appropriate grafts

(Bregman et al., 2002) or tissue-engineered scaffolds (Hynes et al., 1992; McCaig et al., 1991).

The concept of nerve suture was first described in ancient times by Paul of Aegina

(625-690 ac) (Chang et al., 2010) and the first documented surgery on humans to reconnect severed nerve tissue was performed in 1864 by Laugier and Nelaton

(Holmes et al., 1951). The potential strategies for CNS regeneration include the use of peripheral nerve grafts, embryonic spinal cord grafts (Richardson et al., 1980;

Bregman et al., 1986; Horvat et al., 1991; Asada et al., 1998; Itoh et al., 1999;

Duchossoy et al., 2001), cultured Schwann cells (Paino et al., 1991; Xu et al., 1999;

Pinzon et al., 2001) and the development of cell-seeded or cell-free bridges made from biocompatible materials. Additionally, neural stem cells, embryonic neural progenitor cells (Mc Donald et al., 1999; Ogawa et al., 2002) and Schwann cell–loaded biodegradable scaffolds (Olson et al., 2009) have been recently used in the generation

9 of functional artificial neural tissue (Jurga et al., 2009), and for spinal cord tissue restoration (Nishio et al., 2006). While embryonic spinal cord grafts were shown to increase neuronal survival and support new axonal growth in a topographical manner

(Mori et al., 1997; Bernstein et al., 1997), peripheral nerve grafts seem to only supply a permissive substrate for the regenerating axons (Vidal-Sanz et al., 1997).

Today, it is commonly accepted that an ideal scaffold for efficient neural tissue engineering should physically guide regenerating axons throughout the transected gap

(Schmidt et al., 2003). Therefore, recent studies in this field focused on designing longitudinally micro-structured scaffolds for the control of oriented axon regeneration

(Stang et al., 2005; Keilhoff et al., 2003; Lietz et al., 2006; Sinis et al., 2005; Bozkurt et al. 2007).

While some of the approaches for the repair of SCI focuses on longitudinally micro-structured scaffolds and how axons elongate and migrate in response to different chemical and mechanical substrate materials, other studies focus on signaling molecules that are attractive or repulsive to the axons. Many of these studies hold promising results for therapeutic reconstruction of neural connections, however it is important to note that the main obstacles remain: remyelination of axons and the integration of the regenerated neurons with the functional synaptic pathways.

Moreover even if the axonal connections throughout the transected gap could be achieved, it cannot be foreseen what type of functional recovery could be achieved.

Future SC tissue engineering strategies should combine guidance and biomolecular related therapies for optimization of a permissive scaffold through an understanding of the signaling mechanisms and effects of the scaffold characteristics on axonal

10 regeneration (Norman et al., 2009).

.2 Spinal cord organization

The function of the spinal cord is to receive and integrate information between the brain and the peripheral regions of the body. The gross anatomy of the spinal cord is divided into four major regions (sacral, lumbar, thoracic, cervical) along a rostral- caudal axis respectively, each responsible in providing somatic and visceral motor innervations for the body.

In the cervical-lumbar regions, somatic motor neurons; similarly at the thoracic- sacral regions sympathetic and parasympathetic preganglionic neurons provide motor innervations to the musculature. The medial motor column is present at all levels. At cervical and lumbar levels of the spinal cord, the lateral motor column supplies motor nerves to the limb musculature. Sympathetic preganglionic neurons reside in the intermediolateral cell column at thoracic levels whereas parasympathetic preganglionic neurons present at sacral levels (Carpenter et al., 2002).

The spinal cord is composed of two cell types, neurons and glial cells (astrocytes and oligodendrocytes). While neurons (which consist of dendrites, axons, and a cell body) are the basic structural and functional elements of the nervous system, glial cells give support to the function of neurons. Glial cells are present at higher numbers and can undergo mitosis, whereas neurons cannot divide but can regenerate or sprout new processes under certain conditions. One type of glial cell, the astrocyte, contributes to the blood-spinal cord barrier. Oligodendrocytes, on the other hand, serve to myelinate the axons in the CNS to increase the speed of the nerve impulse; just like the sheath of

Schwann cells in the PNS (Schmidt et al., 2003).

In the center of the spinal cord, the butterfly shaped gray matter contains neuronal cell bodies, blood vessels and glial cells, whereas the surrounding white matter is comprised of axons and oligodendrocytes along with blood vessels, microglia

(immune cells) and astrocytes (McDonald et al., 1999; Becker et al., 2007). The ascending and the descending tracts of axons are located in the dorsal and the ventral regions of the white matter, respectively. Ascending tracts of axons send sensory information received from the body to the brain and the descending tracts of axons send motor signals from the brain to the spinal cord. After the spinal cord receives signals from the brain, it sends these signals to the muscles to carry out functions

(Silverthorn et al., 2004).

.3 Spinal cord injury

SCI can result from direct damage to the tissue or trauma to surrounding tissues.

After injury to the spinal cord, glial scar tissue forms which is mainly composed of myelin and cellular debris as well as astrocytes, microglia and oligodendrocytes and is inhibitory to regenerating neurons. Other possible cells that may comprise the glial scar are fibroblasts, monocytes, and macrophages (Schmidt et al., 2003). The hypertrophic astrocytes present in the glial scar not only act as a physical barrier to regenerating axons by expressing intermediate filamement proteins (vimentin, glial fibrillary acidic protein, etc.), but also secrete nerve growth-inhibitory chondroitin sulfate proteoglycans (neurocan, phosphacan, versican, etc) (McKeon et al., 1991;

Schmidt et al., 2003; Rhodes et al., 2004). Another cause for the limited regeneration capability after CNS injury is the slow infiltration of the machrophages to the injury site, which causes delayed myelin removal (Avellino et al., 1995). The main reason

12 behind this delay in the CNS is the blood-spinal cord barrier (BSCB), which limits macrophage entry into the tissue.

Surprisingly, it has been found that reactive astroglia also express some growth- promoting ECM molecules as well, but the growth-inhibitory molecule secretion significantly dominates (McKean et al., 1991). In addition to astrocytes and macrophages, it has been shown that after extreme injuries, in which meninges get damaged, connective tissue cells such as fibroblasts may also invade in to the lesion

(Preston et al., 2001). Previously, NG2 expressing cells have been thought to represent oligodendrocyte precursor cells (Levine et al., 2005) and their role after injury was questioned (Jones et al., 2003). However, recent studies show that NG2 cells generate oligodendrocytes, protoplasmic astrocytes and in some instances neurons in vivo

(Trotter et al., 2010). Moreover Busch et al., (2010) suggest that adult NG2 expressing cells are permissive to neurite outgrowth and increase neuronal survival after SCI.

The scar tissue formation begins right after injury (within hours) and is most significant in the areas where BSCB breakdown occurs (Fitch et al., 1999). Despite inhibitory effects of the gliosis (the increased production of astrocytes), studies show that the glial scar helps to protect the surrounding regions from secondary injury

(Busch et al., 2007), also prevents an increase in lesion size, cell death, immune response or motor deficits (Faulkner et al., 2004).

.4 Overview on Scaffolds for SCI

Recent strategies for neural regeneration rely on the replacement of the inhibitory environment with artificial scaffolds, which is a very promising and important topic in biomedical research. The composite biomaterials of these scaffolds are produced in a

13 liquid solvent and then polymerized into macromers in desired shapes by a thermal, chemical or photo-crosslinking reaction. To create a porous architecture, polymer solution may be mixed with porogens such as sodium chloride crystals or gas bubbles that are produced by peroxides or air-foaming. In the next step, the porogens are removed, leaving grooves behind. By adjusting the variables, such as the crystal size or the water content, the pore size of the scaffold can be controlled (Madigan et al.,

2009).

As mentioned before, physical guidance of a scaffold is one of the important criteria for successful axonal regeneration. For that aim, hollow tubes and porous foam rods are commonly used techniques due to their simple fabrication methods.

Experiments show that host tissue fills the implanted hollow tube with an oriented fibrin matrix, which guides axonal regeneration in vivo. The axonal alignment in amorphous foam rods, however, is open to discussion. Recent studies have concentrated on the fabrication of scaffolds that has longitudinally aligned fibers or channels, which require more advanced processing techniques. Some of these advanced techniques are magnetic polymer fiber alignment, injection molding, phase separation, solid free-form fabrication, and ink-jet polymer printing (Schmidt et al.,

2003).

Injection molding is one of the most commonly used techniques for axonal regeneration strategies (Madigan et al., 2009). Teng et al., (2002) produced a highly porous, two-phase poly (lactic-co-glycolic acid) scaffold for spinal cord repair by using injection molding technique and phase separation (explained in more detail in

“Freeze-drying techniques” section) technique. The polymer was dissolved in dioxane,

14 which was crystallized in an oriented manner through a thermal gradient. After the molding process, they induced the solid-liquid phase separation thermally and produced longitudinal channels in the scaffold (Teng et al., 2002). In another study,

Moore et al., (2006) fabricated spinal cord scaffolds from PLGA, by using injection molding to give it a channeled structure and seeded it with Schwann cells to show axonal growth is supported.

It has been shown that the scaffolds, which have magnetically aligned protein polymer matrices, support axonal regeneration both in vitro and in vivo when compared to random matrices (Ceballos et al., 1999; Dubey et al., 2001). Axons also show alignment on micro and nanofibres (30µm-50nm) of polymers that generated by electrospinning (Madigan et al., 2009), a technique that uses electric charge to produce very fine fibers from a polymer solution. As the pump moves the polymer (collagen,

PLGA, PCL) solution out of the syringe, the polymer droplet goes into a high voltage and the body of the liquid becomes charged and evaporates, leaving behind polymer filaments which is laid down on an grounded target (Yang et al., 2005). Yoshii and colleagues used this technique to generate 20 µm-scale uniform collagen fibres and showed improved axonal guidance in 2D. However results from the same laboratory showed these scaffolds have high cytotoxicity in vivo (Yoshii et al., 2004; Yoshii et al., 2009). The uses of spun or extruded collagen fibers for axonal regeneration were also examined. Tong et al., (1994), investigated the regeneration capability of the transected axons when treated with laminin and fibronectin coated collagen fibers.

Likewise, it has been shown that, regeneration was permitted over 15mm gaps by longitudinally oriented polyamide filaments placed into silicone tubes (Lundborg et

15 al., 1997), and over 80 mm gaps by collagen fibers placed into poly(glycolic acid)- collagen tubes in vivo (Matsumoto et al., 2000). However this enhanced regeneration capability was not significantly different than tubes filled with collagen sponge, which can be fabricated more easily compared to fibers (Toba et al., 2001). As a conclusion, methods that are developed to generate longitudinal porous internal structure and that are also not too difficult to fabricate would be preferential for axonal regeneration.

One of the other advanced scaffold fabrication techniques, the solid freeform fabrication (SFF) technique, is designed to produce complex features from the models in the computer. This technique can be used to fabricate the mold or the polymer scaffold itself. To fabricate 3D structures, three-dimensional printing (3DP), which is a type of SFF technique, can be used. These 3D scaffolds can mimic the architecture of the native tissue such as key spinal cord tracts (Friedman et al., 2002). In spite of these advantages, this technique is very expensive and some processing conditions are not convenient for biological components to be used. An alternative technique that can be used to fabricate 3D structures with desired thickness and dimensions is ink-jet printing. Moreover, this technique is available for biomolecule and neurotrophic factor incorporation. Researchers in MicroFab Inc., in Plano, Texas, have generated bifurcated degradable polymer tubes as well as non-bifurcated ridged tubes that is filled with fluorescent dye (Schmidt et al., 2003). Even though these advanced fabrication techniques provide powerful advantages for spinal cord regeneration, today we are still the earliest stages of technology for axon repair, and there is much to be learned.

.5 Hydrogels for engineered scaffolds

Hydrogels hold great potential for engineered tissue scaffolds due to their native

ECM-like characteristics and biocompatibility (Annabi et al., 2010). ECM plays an important role in regulating cell adhesion, proliferation, migration and differentiation, therefore Liang and colleagues (2007) suggest that a close imitation of the ECM may provide a more favorable environment for such cellular behaviors. Another critical feature for an ideal scaffold is porosity, which provides sufficient surface for cell seeding and a place for their proliferation (Thomson et al., 2000; Patrick et al., 1998).

Today, advanced techniques enable researchers to give hydrogels a controlled porous structure, generating engineered scaffolds with similar functions to native tissues

(Annabi et al., 2010). An artificial scaffold that combines these properties can be beneficial to guide axonal connections in CNS tissue engineering.

.6 Freezedrying techniques

Freeze-drying is a well-known method that is used widely to manufacture porous hydrogels for tissue engineering purposes. The logic behind this technique lies in phase separation, which means separating distinct phases that are existing within the same system. Phase separation can be achieved by thermodynamic instability in a rapidly cooled sample. Then, by removing the ice crystals (porogens) via sublimation under vacuum conditions, the sample get a porous structure as removed ice crystals leave voids in the scaffold. Such scaffolds are favorable since they allow metabolite and nutrient exchange, support revascularization and are biodegradable (Annabi et al.,

2010).

Regardless of different designs in terms of chemical organization and properties,

17 researchers agree that an ideal scaffold for efficient neural tissue engineering should not only support axonal regeneration but also guide it (Schmidt et al., 2003; Stang et al., 2005; Keilhoff et al., 2003; Lietz et al., 2006; Sinis et al., 2005; Bozkurt et al.

2007). In 2004, Stokols and colleagues modified the freeze-drying technique

(explained in more detail in the “Materials and methods” section) to fabricate agarose hydrogels with linear pores. These channeled scaffolds, which later were used for other tissue engineering strategies such as cartilage (Wu et al., 2009) and muscle

(Chen et al. 2007), served as a starting point for novel nerve tissue regeneration strategies. In a spinal cord injury model, one month after the surgery, it was shown that scaffolds fabricated with this method were integrated within host tissue, cells were able to penetrate individual channels, and the grown axons were linear (Stokols et al.,

2006).

These results were confirmed by a few other studies. At 2007, Bozkurt and colleagues used a different technique to produce similar inner microstructured collagen scaffolds by creating a temperature gradient with two sinks placed above and below the polymer solution. They loaded it with dorsal root ganglion (DRG) cells and showed that neurofilament positive axons migrated into the channels of the scaffold.

In a different study, the same group also showed the cytocompability of these scaffolds for human neuroblastoma cell line as well as olfactory nerve ensheathing cells and astrocytes (Bozkurt et al., 2009). Similarly, Mollers et al., (2009) showed that collagen scaffolds with similar inner microstructures support glial cell attachment, maintain viability, proliferation, and migration, and support axonal growth from differentiated human neuroblastoma cells. Tang et al., (2010) evaluated the

18 compatibility of collagen-heparan sulfate scaffolds with olfactory ensheathed cells in vitro. Hu et al., (2009) examined such scaffolds in bridging peripheral nerve gaps in vivo. They manufactured a collagen-chitosan scaffold to bridge a 15mm long sciatic nerve defect in rats and achieved regeneration and functional recovery equivalent to that of an autograft, without the exogenous delivery of regenerative agents or cell transplantation.

A B

Figure 1: SEM pictures of longitudinal section (A) and cross section (B) of the GCG-CFDSs.

In a recent study, Zhang et al., (2010) published SEM photographs of an acellularized xenogeneic nerve matrix which was porous and contained longitudinal grooves. By immunostaining, they have further proved that the main components of natural nerve matrix were collagen I and III. With all of this knowledge, we designed and manufactured similar microstructured gelatin (an essentially denatured collagen) scaffolds in our laboratory (Figure 1- A, B) by using our in-house designed controlled freeze-drying device. By controlling the temperature decrease with a PID controller, we obtained more uniform, unbranched micro-channels (Figure 1- A). Moreover we rheologicaly characterized our scaffolds to closely mimic the natural tissue, which has

19 not been done in previous studies. When compared to acelularized nerve matrix

(Zhang et al., 2010), it is evident that our scaffolds resemble the typical architectural characteristics of the natural tissue.

.7 The effect of pore diameter on axon extension and ECM secretion

Unlike most cells, neurons can generate axons that may extend for meters through the ECM and other tissue-specific cells by secreting proteases from growth cones to degrade the surroundings (McFarlane et al. 2003). During embryogenesis, the growth cones of pioneer neurons migrate through the embryonic tissue, which is still short and relatively uncomplicated, and reach to their target tissue. Then, these pioneer neurons, which may die later in development, are followed by other neurons (Klose and Bentley et al., 1989). If these pioneer neurons would not guide to other neurons, the neuronal connections would not be able to be established which further proves that an optimum scaffold for the injured neural tissue requires a guiding structure in which neurons can regenerate and axonal outgrowths can connect to their target neurons.

Francisco et al., (2007) proves that 3-D constraints can be used to guide and control the length of axons. While the percentage of neurons extending axons in fully enclosed chambers were decreased as a function of chamber width, the percentage of neurons extending axons in corridors were similar to those on 2-D. In other words, the axon extension was not affected by the confinement of long corridors, when the remaining two directions do not have constraints. There was a tendency toward more neurons having grown axons in corridors between 20 and 50µm however no statistical differences were obtained comparing the data (Francisco et al., 2007). In a different

20 study, it has also been shown that 100µm width channels in poly(lactide-co-glycolide) scaffolds were more effective than larger channels for neuronal guidance and resulted in longer neurite extension, as well as less secondary branching (Houchin et al. 2007).

This result confirms previous studies where axons exhibit decreased extension as a function of increased matrix porosity (Bellamkonda et al. 1995, Dillon et al. 1998).

The pore size of the scaffolds also significantly affects ECM secretion, cell growth and penetration in 3-D (Annabi et al., 2010). Lien et al., (2009) claims that the chondrocytes increase their ECM secretion directly to the pore size. It was found that in genipin crosslinked gelatin hydrogels, cells tend to proliferate rather than secreting

ECM proteins in small pores, and they became overconfluent during differentiation.

Accordingly the extent of ECM secretion decreased compared to that of scaffolds with larger pores (Lien et al., 2009).

Collectively, these data suggests that the optimum channel diameter for neural tissue engineering should be big enough for sufficient oxygen and nutrient transport within the deeper compartments of the biomaterial but small enough to direct axon elongation and ECM deposition. With the previously described, sophisticated preparation techniques, the enhanced ability to control the architecture, degradation rate, pore size and mechanical properties of scaffolds will drive neural tissue engineering research towards regeneration.

.8 Angiogenesis & vasculogenesis:

During embryogenesis, vascularization occurs either by angiogenesis or by vasculogenesis in the tissues. There is a controversy about how to distinguish these two terms, either by the origin of endothelial cell or by the steps of maturation of the

21 vascular network. Pudliszewski et al., (2005) used initial definitions and referred vasculogenesis as the development of a vascular network from intrinsic ECs, while angiogenesis was described as blood vessel formation by extrinsic ECs from a pre- existing network.

An experiment on mouse embryos showed that vascularization in a majority of vessels in the limbs was by angiogenesis, whereas in visceral organs it was by vasculogenesis (Pudliszewski et al., 2005). This data corresponds to previous research that had been done in 1989 by Pardanaud and colleagues, which shows that when an ectoderm and somatopleural mesoderm originating from one species is grafted into a host from other species, it is colonized by host ECs, meaning it is vascularized by angiogenesis (Pardanaud et al., 1989).

Brain-derived neurotrophic factor (BDNF) is one of the neuroprotective factors that have been known to stimulate angiogenesis (Raab & Plate et al., 2007), and its administration increases the vascular density after SCI, which shows a relationship with recovery (Yoshihara et al., 2007). Bone marrow-derived mononuclear cells also improve vessel densities and recovery after SCI (Yoshihara et al., 2007). Another neuroprotective and angiogenic factor is vascular endothelial growth factor (VEGF), but it can be harmful in large amounts due to increasing vessel permeability (Benton &

Whittemore et al., 2003).

Capillaries in the CNS support the tissue with energy and nutrient transfer (Raab &

Plate et al., 2007) therefore vessel density enhancement and the restoration of microvessels in the injured tissue is a promising research area for spinal cord repair

22 and recovery (Rauch et al., 2009).

.9 Neuritogenesis

Neuritogenesis is accepted to be the first step of neuronal differentiation in the developing nervous system and characterized by emerging neurites from the neuronal soma (Da Silva et al., 2003). The key mechanism underlying this morphological change, the reorganization of the neuron cytoskeleton, is still not understood completely but known to be stimulated by a number of microtubule-associated proteins (MAPs) (Bouquet et al., 2004; Dehmelt and Halpain et al., 2004; Dent and

Gertler et al., 2003), and Rho-GTPases (Govek et al., 2005; Li et al., 2006; Luo et al.,

2000), and the surface receptors that bind ephrins, netrins, semaphorins and other major ligands (Gallard et al., 2005; Huber et al., 2003). The distal tips of the neurites have locomotor organelles called growth cones, which was discovered by Ramon and

Cahal over a century ago (1890). These structures can respond to the same types of signals that migrating cells can sense and secrete proteases to degrade surroundings, therefore enabling neurons to generate axons that may extend for meters through the

ECM and other tissue-specific cells (McFarlane et al., 2003). Experiments on growth cones may help us to understand the environmental change-related axonal responses following CNS trauma, and may direct future therapeutic approaches (Hou et al.,

2008). The growth cone consists of two domains, a microtubule-rich, cytoplasmic central (C-) domain; and a surrounding peripheral (P-) domain from which filopodia and lamellipodia extend (Gordon-Weeks et al., 2004). Microtubules are polarized (fast growing plus and slow growing minus ends) (Heidemann et al., 1981) and grow slowly due to hydrolysis of GTP bound to the filaments, by reorganizing and directing

23 growth cone shape continuously (Gordon-Weeks et al., 2004). Additionally, microtubule-binding proteins enable turning movements of the growth cone (Dent and

Gertler et al., 2003). Lamellipodia and filopodia formations rely on F-actin filament polymerization and represent antenna like sensors, which investigates the radius-wide environment (Davenport et al., 1993). Following polymerization F-actin is transported in a retrograde fashion to the center of the growth cone and filaments are depolymerized to their actin subunits, which later may be used again for F-actin polymerization (Gallo and Letourneau et al., 2004).

Both inhibitory molecules (ephrins, semaphorns, etc) and guidance signals influence the growth cone behavior. A variety of environmental signals modulate these molecules, such as growth factors (GFs) (neurotrophins), ECM components, cell adhesion molecules (CAMs), neurotransmitters, electrical stimulation, calcium signaling, etc. (Hou et al., 2008).

A recent study by Blackmore and Letourneau et al. (2006) showed that intrinsic changes developing in the old neurons limit their regeneration capability. They compared embryonic day 15 (E15) primary chick neurons with embryonic day 9 (E9) neurons in a permissive environment. E15 neurons demonstrated a much slower axonal extension rate and axon regeneration was 90% less than E9 neurons. A better understanding of the signaling mechanisms of the intrinsic and extrinsic factors that effect neuronal regeneration capability may enable researchers to control them and reactivate positive signals in the adult neural tissue (Hou et al., 2008).

.10 Neurovascular crosstalk

It has been shown that some angiogenic factors also have effects on neural cells

(Carmeliet et al., 2003). The fact that some of these angiogenic factors have originated from the nervous system drove the researchers to search whether neurotrophic factors also have angiogenic properties. Zacchigna et al., (2008), termed molecules that effect both neural and vascular cells as angioneurins. It is also of interest by which mechanisms these factors carry out their activities and whether other cell types are functional in the mechanism that angioneurins affect vascular and neural cells

(Zacchigna et al., 2008). Zacchigna et al., (2008) summarized the effects of angioneurins as the regulation of angiogenesis, blood–brain barrier (BBB) integrity, vascular perfusion, neuroprotection, neuroregeneration and synaptic plasticity. The effects of the GFs that have such neurovascular properties will be categorized and described in more detail below.

.10.1 Interactions between nervous & vascular systems:

The first tissue systems that appear in embryogenesis are the nervous and vascular systems. In addition, during neurogenesis, neural cells and endothelial cells are coordinated closely, and they construct a neurovascular niche (Park et al., 2003).

Similarity in development of these systems, common organizational features [similar signaling cues for new cell formation, cell migration, cell fate, formation of boundaries

(Lok et al., 2007)] and responsiveness to similar GFs by similar receptors show that they have evolved coordinately (Park et al., 2003; Nicole et al., 2004).

The morphological developments of ECs and neural cells and most importantly their guidance within target tissues are also parallel (Park et al., 2003; Lok et al.,

2007). Moreover, since their anatomic patterning is similar, in many cases they share the same physical space and they mutually regulate each other: glucose, oxygen and hormone delivery is provided by the vascular network as well as the regulation of nerve response to the local environment, while the neural network regulates the location and pattern of blood vessels (Park et al., 2003). This knowledge suggests that angiogenesis stimulation may also induce neurogenesis (Lok et al., 2007).

Understanding the details of this cross-talk between neurogenesis and angiogenesis may guide the researchers to design improved strategies for recovery following injury.

.11 Crosstalk between neurons and ECs

As mentioned before, blood vessels and nerves are located closely together and they communicate with and regulate each other by intercellular interactions. In more detail, neuronal cells and ECs answer to the changing conditions in the environment by producing GFs and their receptors; which in turn effect them in a paracrine and/or autocrine way and induce their differentiation, migration and proliferation (Park et al.,

2003). Park and colleagues, (2003) suggest that these similar receptors and factors are likely to have common signaling pathways.

During hippocampal neurogenesis in the adult, neural and endothelial precursors proliferate together in the subgranule zone (SGZ). SGZ is where new neurons being generated lifelong and is also responsible from the recovery of the damaged neurons.

It has been shown that the neuronal precursors from SGZ encourage the EC proliferation to the region, which will in turn secrete neuroangiogenic factors to support the migration and maturation of neurons (Park et al., 2003). Leventhal and colleagues, (1999) made experiments on monolayers of astrocytes, fibroblasts and ECs

26 to examine neural outgrowth and they showed that ECs were the only cell type that supports neural outgrowth derived from adult SGZ. This finding proves that ECs are not only functional as a channel for the blood supply, but also are active in cell signaling (Lok et al., 2007). In other words, neurogenesis and angiogenesis are intricately linked (Lok et al., 2007).

Furthermore it has also shown that endothelial signals can affect neural cell fate determination and neuronal cells can induce endothelial cell differentiation and network patterning (Park et al., 2003). TrkB is a high affinity catalytic receptor for several neurotrophins whereas TrkB-Fc inhibits BDNF-induced proliferation of TrkB transfected cells. BDNF blockage by TrkB-Fc chimera interrupted the endothelial cell effects on neurons and the addition of exogenous BDNF into the co-culture replicated the effects (Lok et al., 2007). Therefore, it is proven that ECs secrete the BDNF, which supports neurogenesis (Figure 2).

ECs also express KDR/Flk-1, Flt-1 and NRP-1 (neuropilin-1), which are the receptors of one of the most important growth factors in angiogenesis and neurogenesis; VEGF (Figure 2). This growth factor is known to have fundamental effects in endothelial cell survival, migration, proliferation, also in stimulation of axon growth, neurotrophy and neuroprotection (Park et al., 2003); and is known to be expressed by neurons.

Figure 2. Illustration of systemic coordination between angiogenesis and neurogenesis by confocal pictures. Neuronal cells express VEGF that induces proliferation of ECs through its receptors, VEGFR. Conversely, endothelial cells produce BDNF, which binds to the Trk receptor on nerve cells, and stimulates neurite outgrowth.

Numerous neuroangiogenic factors and receptors are expressed by neuronal and endothelial cells. These factors have many effects during neuroangiogenesis, such as proliferation, migration, and differentiation of endothelial and neuronal cells. These mechanisms are not generated from a single cell, but from the coordinated interaction of cell-cell units (Park et al., 2003).

.11.1 Angiogenic factors’ effects on the nervous system:

VEGF, one of the most important growth factors in angiogenesis and neurogenesis, has been shown to have supporting effects on EC survival, migration, proliferation, and also in stimulation of axon growth, neurotrophy and neuroprotection (Park et al.,

2003). VEGF is expressed in ECs, neurons and astrocytes and binds to the neuropilin receptor NRP-1, a receptor that is expressed both in neural and endothelial cells (Lok et al., 2007). NRP-1 knockout mice phenotypically expressed failure in the maturation

28 and remodeling of the neurovascular network, suggesting necessity of NRP-1 for neuroangiogenesis.

After SCI, it has been found that VEGF and NRP-1 are upregulated. This knowledge is important for both neuronal guidance and EC survival, since besides

VEGF, semaphorins also bind to NRP-1, which in turn results in the repelling of growing axons. Therefore there is a complex coordination between the neurons and

ECs, via VEGF. Insulin-like growth factor-1 (IGF-1), TGF-β and EGF were also shown to control vascular and neuronal proliferation. Moreover, IGF-1 has a protective effect on neurons (Park et al., 2003).

Fibroblast growth factor-2 (bFGF or FGF-2) is also an angiogenic factor, which is upregulated in neuropathologic conditions after spinal cord injuries (Park et al., 2003).

In vitro, FGF-2 has been shown to be mitogenic for neuronal stem cells (Ciccolini et al., 1998; Palmer et al. 1997). In addition to FGF, epidermal growth factor (EGF) has been also shown to regulate the survival and motility of neural cells in invertebrates

(Chao et al., 2000).

As a conclusion, the cross-talk between cells and interactions between signaling pathways seem to have positive and crucial effects on both neurogenesis and angiogenesis. For example FGF-2 stimulates both neurogenesis and angiogenesis by stimulating VEGF and VEGFR expression in ECs (Lok et al. 2007).

.11.2 Neurogenic factors’ effects on the vascular system

The coordinative effects of VEGF and semaphorins have already been mentioned above. In addition, the other 3 major families of neuronal guidance factors (ephrins, slits and netrins) have also been proven to stimulate angiogenesis (Carmeliet et al.,

2005). This knowledge takes attention to the evolutionary link between neurogenesis and angiogenesis once again.

Wilson and colleagues, (2006) showed that netrin-1,-2, and -4 stimulate migration, proliferation, and tube formation in ECs. In addition, in diabetic experiment animals it was shown that netrins could stimulate neoangiogenesis by enhancing capillary density and restoring nerve conduction velocity (Lok et al. 2007).

ECs sense the changes in the environment and express necessary neutrophic factors: NGF, BDNF, NT-3, and NT-4/5. These neurotrophins bind to Trks (a kind of receptor tyrosine kinase) to regulate neuronal survival, plasticity, and differentiation.

Of these, NGF and glial cell-derived neurotrophic factor (GNDF) were found to be most powerful for motor neurons, whereas GDNF and BDNF were most effective for sensory neurons (Lok et al. 2007).

As a conclusion, studies show that these neuroangiogenic factors have autocrine and paracrine effects on both neuronal and endothelial cells (Park et al., 2003;

Carmeliet et al., 2005; Zacchigna et al., 2008).

 MATERIALS AND METHODS

.1 Preparation of genipin crosslinked gelatin polymer (GCG)

Individual (aqueous) solutions of 5% gelatin (American Master*Tech Scientific

Inc., CA) and 2% genipin (HPLC grade, Wako, VA) were prepared in distillated water, degassed and filtered through 0.2 μm syringe filters. Thereafter, 6 ml of each solution were mixed (1:1 v/v) to generate a working solution containing 1% genipin

(cross-liker) and 2.5% gelatin solution. This solution was further processed for scaffold production (see below) or left at room temperature for 2 hr followed by cooling at 4ºC for 14 hr to generate a crosslinked hydrogel gel.

.2 Scaffold production by regular and controlled freezedrying

To generate a conventional random porous regular-freeze dried scaffold (RFDS),

48ml GCG was poured into 24 well-plate immediately after preparation and left at -

20ºC for 2 days. The gels were dried for 48 hr by lyophilization in a freeze-dryer

(Christ model, Alpha 1-4, Gefriertrocknungsanlagen, Osterode am Harz, Germany) at

-50ºC, and200 millitor vacuum.

Unidirectional controlled freeze dried scaffolds (CFDSs) were prepared essentially as previously described (Wegst et al., 2010). Controlled freezing was carried out in a homemade freezing apparatus, which was designed and built following published literature (Wegst et al., 2010). In brief: 12 ml freshly prepared GCG solution was poured into a specialized sealed cup (Figure 3-A4, B4). The bottom of the cup is made of a thermally conductive copper disc (Figure 3-B4); the sides are made of thermally insulating polytetrafluoroethylene (PTFE). This setup allows for unidirectional heat

31 transfer, i.e. temperature changes will affect the base of the sample without affecting the sides and top, thus creating a unidirectional longitudinal temperature gradient within the polymer solution sample (Wegst et al., 2010). The copper-PTFE cup

(Figure 3-A4, B4) was placed atop a copper rod (Figure4-A,B2), whose bottom end was submerged into a container of liquid nitrogen (Figure 3-A1, B1). A heating element (Figure 3-A3, B3) was wrapped around the copper rod just below where the copper-PTFE cup rests. The heating element is controlled by a PID controller (not shown) based on the input of a thermocouple (Figure 3-A5) placed at the base of the copper-PTFE cup. The heating element maintains the top of the copper rod at RT and then, by lowering the current voltage of the heating element, it maintains a controlled

(3oC/min) decrease in temperature for approximately 120 minutes until the polymer solution is frozen.

In normal freezing processes, a freezing equilibrium is quickly reached through the sample all at once. In this controlled freezing process, a temperature gradient throughout the sample is achieved, such that the bottom is colder than the top (Stokols et al., 2006; Bozkurt et al., 2007; Wegst et al., 2010). The PTFE sides do not conduct temperature well, which makes all heat transfer to go through the copper on the bottom of the cup.

A unidirectional temperature gradient upward through the sample is maintained. A freezing front, which is the interface between solid and liquid phases of the sample, moves slowly through the sample along the temperature gradient. Different portions of the sample will freeze at different times based on the controlled freezing front velocity.

The bottom of the aqueous solution becomes frozen first, creating ice crystals among

32 the cross-linked polymer scaffold. As the freezing front moves upward through the sample, the ice crystals continue to form as columns through the polymer. Once the entire sample is frozen, the ice columns sublime during the two

Figure 3: The controlled freeze-drying device, a macro view of the scaffold and microscopical image of channels. A, a photograph of the device; B, a scheme of the controlled freezing device; C, a typical cylindrical scaffold; D, the light microscopic view of the typical longitudinal channels. 1 = Dewar flask for liquid nitrogen, 2 = copper rod, 3 = heating element, 4 = copper-PTFE cup, 5 = thermocouple, 6 = polymer solution.

33 days lyophilization procedure, which generates the hollow channels within the scaffold

(Figure 3-D). The scaffolds (Figure 3-C) are kept dry in a desicator until use; microscopical examinations of the scaffold indicate their integrity up to at least 6 months storage.

For cell culture experiments, the scaffold was cut (into 1x1x2mm-sized pieces), which were then exposed to UV for 30 min for sterilization before seeding with cells.

Excess genipin, which not cross-linked to gelatin, was removed, by washing the scaffolds 5 times with complete serum-containing cell culture medium.

.3 Measurements of mechanical properties of the scaffold

The mechanical properties of the scaffolds were determined using a Bohlin

Rhometer (Malvern Instruments Ltd, Worcestershire, United Kingdom). Freeze-dried scaffolds samples were hydrated and cut into cylindrical pieces (8mm diameter,

1.5mm height) large enough to cover completely the upper plate of the instrument.

Three samples were prepared for each concentration of scaffold and measurements of each sample were performed in triplicates.

The conversion equation for shear modulus (G) and elastic modulus (E) is:

E=2(1+v) G where E and G represents elastic and shear moduli respectively, and v represents the Poisson's ratio. For perfectly elastic materials, the Poisson's ratio is given as 0.5, therefore the equation becomes E=3G; when v = 0.5.

.4 Invitro degradation of the GCGCFDSs

In this experiment, the dry weight (W0) of two different genipin concentrations of

GCG-CFDSs was measured before they were placed into cell culture medium (10%

FBS) and kept at 370C up to 3 weeks. Every week, a sample of GCG-CFDS from each concentration was washed, freeze-dried and weighed (Wt). The weight loss percentage, which represents the degradation rate (D) of the sample, was calculated by the equation: D = (W0 – Wt)/W0 x 100%.

.5 Cells

Rat adrenal medullary PC12 rat pheochromocytoma neuronal cells are neural crest derived cells of sympathoadrenal lineage and an established model for neuronal differentiation (Fujita et al., 1989). PC12 cells were grown in DMEM (Cellgro,

Mediatech Inc., VA) supplemented with 7.5% fetal bovine serum (FBS), 7.5% horse serum (ES) and 0.5% penicillin streptomycin (medium A), in T 75 cm flasks (BD

Falcan, CA) that were maintained at 37°C in a 5% CO2 incubator. Cells were split at

50% confluence by gently mechanically detaching them from the flask and propagated at a split ratio 1:7. For the majority of the experimants we used a neuritogenic GFP- labeled PC12 subclone (GFP-PC12), which has been characterized before (Arien-

Zakay et al., 2009). Scaffolds were seeded at a density of 50.000 cells/scaffold, which in preliminary experiments was found to be the optimal seeding density.

Primary forebrain neurons from chick embryo were isolated at the day of the experiment and purified according to an established procedure (Heidemann et al.,

2003; Francisco et al., 2009). After isolation, the cells, containing 90% neurons and

10-20% glia and other cells were transferred to fresh M199 (containing 10% FBS) medium (Francisco et al., 2009), and counted in a hemocytometer. Scaffolds were seeded with 50.000 primary cells.

Figure 4: Day 8 chicken embryo forebrain neurons after extraction on 2D polylisine coated surfaces (Green: tubulin immunostaining with DMIA + FIT-C. Red: actin staining with rhodamine phalloidin).

Immortalized EA.hy926 endothelial cells, derived from the fusion of human umbilical vein endothelial cells (HUVECs) with A549 lung carcinoma cells (Edgell et al., 1983), were grown in T 75 flasks in Dulbecco’s modified Eagle’s medium

(DMEM) supplemented with 10% fetal bovine serum (FBS) and 0.5% penicillin streptomycin (medium B). The medium was changed every other day and cells were split at 80% confluence at a ratio of 1:6.

For co‐culture with neural cells we needed to unify the diverse culture media.

Since EA.hy926 cells easily grow in a variety of media, we opted for adapting the

ECs to grow in the respective neural cell medium. For this, ECs were gradually

36 transfered from endothelial medium B to medium A in the following order: from

25% A + 75% B to 50% A + 50% B to 75% A + 25% B and ending with 100% A.

Figure 5: Cell proliferation of adapted cells (pink) compared with EA.hy926 cells (blue) in regular medium (graphs represent different initial seeding concentrations: 1, 20, 50, 100 and 200K form top left to bottom right respectively).

The proliferation of adapted EA.hy926 cells and EA.hy926 cells in regular medium was analyzed every second day over a period of up to 6 days using the

Alamar Blue staining method (Figure 5). The cells were incubated for 3 hours with the

Alamar Blue reagent and then, supernatants were collected and their fluorescence intensity was evaluated in an ELISA reader (excitation: 560 nm & emission: 595 nm).

The difference between cell proliferation of adapted cells compared to EA.hy926 cells in regular medium, decreased and disappeared as the cell concentration increases. This result points out the positive effect of the cell-cell communication and the secreted

GFs.

.6 Cell proliferation

Cell proliferation was analyzed every second day, over a period of up to 6 days using the Alamar Blue staining method (Nikolaychik et al., 1996; Lelkes et al., 1997).

Briefly, the cells washed once in PBS and then incubated for 3 hours with the their respective growth medium supplemented with 5% Alamar Blue reagent (Abd Serotec,

Raleigh, NC). Supernatants were collected and their fluorescence intensity was evaluated by excitation at 560 nm and emission at 595 nm using an ELISA Synergy 4 reader (BioTek, Winooski, VT). The data, expressed as mean ± SD, is presented in arbitrary fluorescence units.

.7 Fluorescence activated cell sorting (FACS)

Sorting of the GFP/lentivirus-transduced PC12 4F cloned cells (Arien-Zakay et al.,

2009) was performed using a fluorescence-activated cell sorter (FACS Aria, BD

Biosciences, Franklin Lakes, NJ) with fluorescence excitation at 488 nm using the

FITC channel. In brief, the GFP-labeled PC12 cells (GFP-PC12) were mechanically dislodged from the dish and resuspended in Hank’s Buffered Saline Solution (HBSS) containing 3% BSA.

By pipetting up and down, the cell aggregates were carefully separated into individual cells followed up by visual inspection evaluation. The final suspension contained 2×107 cells/ml was filtered through a 35μm filter directly before the sorting.

The population of cells exhibiting the highest fluorescence intensity (0.9% of all cells) were separated, collected and cultured in medium A, till experimental use.

Figure 6: Fluorescence activated cell sorting of GFp-PC12 cells.

.8 Endothelial cell labeling

To easily track the fate of EA.hy926 endothelial cells and to distinguish them from the neuronal cells in the heterogeneous co-cultures, ECs were labeled with a red fluorescent cell tracker dye Cm-Dil derivative (C7001, Molecular Probes, Carlsbad,

CA), essentially as previously described (Kruyt et al., 2003). In brief: A 1mg/mL stock solution was diluted 1:250 in warmed HBSS. ECs were resuspended in this medium at a final concentration of 2x106/cells ml and maintained for 3 minutes in the incubator

(Kruyt et al., 2003) followed by an additional 15 min at 40C to allow the dye to label the membrane and to slow down the endocytotic process, according to manufacturer instructions. Cells were pelleted by gentle centrifugation (800 rpm) and then resuspended in medium A. in select cases we also used fluorescent phallotoxins

(TRITC-phalloidin) to visualize cells in scaffolds seeded with homogeneous cell- populations using previously described protocols (Wulf et al., 1979; Lelkes et al.,

1986).

.9 Populating the scaffold with cells and treatment with

angioneurins

In preliminary studies we established the optimal volume (10μl), and the optimal cell number (50,000 cells) for seeding our scaffolds (1x1x2 mm). Unless mentioned otherwise, the scaffolds were always seeded under these conditions. In the first step each scaffold was mounted vertically (in an upright position with the longitudinal pores facing upwards) in a single well of 48 well plate. A single drop of the cell suspension (50,000 cells in 10μl) was added on top of each scaffold, and incubated for

30 minutes at 5% CO2, 37°C to allow distribution and attachment into the longitudinal channels. Thereafter, using a fine forceps, the scaffolds were turned horizontally, immersed; in 200μl medium A and the plates were returned to the incubator. The efficacy of population of the scaffold was found to be 85%, and determined by counting the cells not attached to the scaffold one day after seeding and subtracting this value from the total number of cells at the beginning of the experiment. To promote neuronal differentiation of the GFP-PC12 cells (Arien Zakay et al., 2009) and endothelial cells angiogenesis (Lecht et al., 2010 A; Lecht et al., 2010 B), the scaffolds were treated with a mixture of 50 ng/ml NGF and 10 ng/ml FGF-2.

.10 Confocal Microscopy

The scaffolds were fixed (in 3.7% formaldehyde for one hour), and the nuclei were

40 stained with 4', 6-diamidino-2-phenylindole (DAPI). The scaffolds were examined in a confocal laser scanning fluorescence microscope (Olympus Fluoview 1000), using excitation and emission wavelengths of 488 (for GFP+ PC12 cells), 543 (for ECs) and

405nm (for nuclei) at different magnifications. Photographs were acquired and 3-D z- projections were generated by using the FluoView confocal software.

.11 Scanning Electron Microscopy

The scaffolds were coated with carbon for 30s at a pressure of 0.025mbar using a carbon rod, thermal evaporative coater (Cressington 208 High Vacuum Turbo Carbon

Coater) and their structure was evaluated by scanning electron microscopy at an acceleration voltage of 5 kV using either an FEI XL30 Environmental FESEM

(Hillsboro, OR) or a Zeiss Supra 50VP FESEM (Thornwood, NY). Photographs were taken at a magnification of 500-1500 X.

.12 Statistics

Where appropriate, all data are expressed as mean ± SEM, evaluating at least 3 samples, from 3 to 6 independent experiments. Comparisons between groups were analyzed by Student’s t-test or one-way analysis of variance (Microsoft Office Excel,

2003) to determine significant differences. P values less than 0.05 were considered statistically significant.

 RESULTS & DISCUSSION

.1 The morphology of the scaffolds

To evaluate the microscopic structure of the scaffolds, we characterized longitudinal and transverse sections of dry scaffolds by SEM. Figure 7-B-a,b,c and d presents a typical organization of the GSG-CFDSs indicating the presence of continuous longitudinal channels of different diameter in accordance to genipin crosslinker concentration.

At 0.005% genipin and a constant 2.5% gelatin the typical diameter measured was around 20 µm (Figure 7-B-a,b) while at 1% genipin it was around 50 µm (Figure 7-B- c,d). Both values were independent of scaffold hydration conditions (Figure 7-A) and being in the right range for a successful neuron orientation (Francisco et al., 2007).

These findings indicate a 2.7 fold increase in the diameter upon of the increase in the concentration of the cross linker.

Figure 7-A-insert also presents a direct relationship between regular freeze-dried scaffold pore size and genipin concentration. However these RFDSs lacked channels and their pore sizes were very large which would be suitable for other tissue engineering purposes (Figure 7-B-e,f). The scaffolds were characterized by a strong autofluorescence due to the genipin crosslinker, under confocal microscope (Figure 8).

Morphological analysis of dry (Figure 7- B) or wet scaffolds (Figure 8) after six months up to one year indicated preservation of the longitudinal channels organization, continuity and similar pore size compared to freshly prepared scaffolds

(data not shown). Therefore the present CFDSs enabled the production of a scaffold with continuous, stable, longitudinal pores at an optimal diameter size of 20-50µm.

Figure 7: Longitudinal channel diameter measurements (A) and SEM micrographs (B) of GCG-CFDSs and RFDSs. A. White represent dry and black represents wet GCG-CFDSs. The insert represents pore size measurements of RFDSs with different concentrations of genipin while the gelatin concentration kept constant at 2.5%. B. The SEM micrographs of the GCG-CFDSs (a,b,c and d) and RFDS (e and f). a and b represents longitudinal and transverse sections of 1% genipin- 2.5% gelatin respectively, while c and d represents longitudinal and transverse sections 0.005% genipin- 2.5% gelatin respectively. e and f represents 0.005% and 1% genipin- crosslinked RFDSs respectively. The scale bars represent 30μm for a and b, 50μm for b and d, and 200μm for e and f. The data represent average ±SD of three groups, each one n=10 each symbol p<0.005.

Figure 8: The confocal photographs of the autofluorescent 0.005% genipin (right) and 1% genipin (left) scaffolds in medium (scale bar represents 100µm).

.2 The mechanical properties of the scaffolds

To characterize the mechanical properties of GCG-CFDS and to compare them to that of conventional RFDS or GCG gels we prepared samples at different genipin

Figure 9: Mechanical properties of the GCG-CFDS (triangle), RFDS (square) and gel scaffolds (diamond). Groups of three wet scaffolds (n=3) were submitted for rheological measurements. The data represent average SD and * p<0.05 compared to RFDS; ** p<0.05 compared to the gels.

44 concentration, while keeping the gelatin concentration constant at 2.5% and measured their complex modulus by rheology. The data presented in Figure 9 showed that the elastic modulus for CFDLS, RFDS and GCG ranged between 4.2, 6.6 and 1.5 kPa to

51, 20.7 and 5.4 kPa, respectively, depending on genipin concentration. Figure 9 clearly demonstrated the increase in the elastic moduli of RFDS and GCG gels from

0.04 (0.005%) to 44.2 mM (1%) genipin.

To date, little information has been published about the mechanical properties of micro-channeled scaffolds; however, our measurements of the conventionally available collagen scaffolds from Bozkurt and colleagues showed that the mechanical properties of these collagen scaffolds are 8.31 kpa which might be too soft for spinal cord repair. In another study by Lin et al., (2003) scaffolds prepared by using a solution coating and porogen decomposition method, found to have a compressive modulus between 43.5–168.3 MPa, which might be extremely high. Therefore our results demonstrated in this current study represent the most appropriate range of mechanical properties for spinal cord injury repair strategies.

.3 The degradability of the GCGCFDS in the serum

To characterize the stability of GCG-CFDS in cell culture medium containing 10%

FBS, similar pieces of scaffolds of about 6 mg weight, of two different genipin concentrations were incubated up to a month and weekly washed, dried and weight measured. The data presented in Figure 10 clearly indicates that 0.005% GCG scaffold degrades by 85% in the first week while 1% GCG scaffold was very stable, losing

25% weight after 3 weeks.

Figure 10: Time course of GCG-CFDSs degradation. Groups of 3 similar weight scaffols (n=3) were incubated in medium supplemented 10% serum at 370C for three weeks. The scaffolds were weekly dried and their weight measured. The data represent average ± SD. Black and white represent GCG- CFDSs obtained by crosslinking 2.5% gelatin with 44.2mM (1%) and 0.2mM (0.005%) genipin respectively.

.4 Biocompatibility of the GCGCFDS for homogeneous cell

populations

Considering the long-term goal of use of GCG-CFDS for neuronal engineering, for biocompability purposes, we investigated the ability of PC12 sympathetic neuron,

FBN and ECs to grow for two weeks in these 3D scaffold conditions. Figure 11 presents typical growth curves of cell growth in the scaffold. Initial doubling time of

PC12 cells was about 2 days similar to regular 2-D conditions; thereafter, 6.5 days were required for their doubling time. The doubling time of EC cells in the scaffold was around 4.5 days compared to 2 days in regular 2-D conditions. These findings

46 indicated a retardation effect of the scaffold conditions on both cell types. However,

FBN cells did not proliferate in the scaffolds as expected from post mitotic primary neurons.

Figure 11: The proliferation of homogenous cell cultures in the GCG-CFDSs. Scaffolds were populated with PC12 (circle) cells, ECs (square) and FBN (triangle) and cell proliferation was estimated by the Alamar Blue Method.

Confocal microscope analyses of individual cultures of ECs, FBN and PC12 cells grown separately for 7 days in the GCG-CFDS are presented in Figure 12 and Figure

14 A and B. The scaffolds were characterized by a strong autofluorescence, which assisted in identification of the longitudinal channels (Figure 12, arrows). The presence of ECs was visualized using red CellTracker (CmDIL). It was evident that the cells were attached to the longitudinal channels surface and that they did not block the continuity of the channels (Figure 12 A and B).

Figure 12: Confocal microscopy images of monoculture in the scaffold after 7 days. A and B represents ECs (red), C and D represents FBNs (green). Nuclei were counterstained with DAPI (blue). The scale bar represents 100μm for A and B and 20μm for B and D.

After 7 days in the scaffold, increased adhesion and cell flattening were observed

(Figure 12 A and B), generating an efficient coverage of the longitudinal channels with ECs, resembling a capillary like organization. FBNs were prepared freshly and seeded onto the scaffolds. After 7 days the presence of FBNs were visualized by measuring the expression of neuronal specific marker TUJ-1 (Karkkainen et al., 2009),

(Figure 12 C and D, green).

It was evident that GCG-CFDS promoted adhesion and long neurite outgrowths directed longitudinally to the scaffold axis (Figure 12 C, D) as also observed on 2-D culture of FBN on gelatin-coated surfaces (Shimazaki et al., 2001; He et al., 2005). To further evaluate neuronal biocompatibility of the scaffolds, GFP-PC12 cells were seeded in GCG-CFDS scaffolds. After 7 days, we found that the cells were populated within the scaffold channels (Figure 14 A) but axonal outgrowth was poor (Figure 14

B, Figure 15 A and B) in contrast to strong differentiation process of GFP-PC12 cells on 2-D (Arien-Zakay et al., 2009). To further show the discrepancy between the FBN and PC12 cell differentiation in the scaffolds we quantified the time course of neurite outgrowth of both cell types in the scaffold up to seven days. Figure 15 A and B clearly demonstrates a decrease in neurite expression when GFP-PC12 cells were grown alone in the GCG-CFDS.

PC12 cells are well-characterized neuron-like cells, which stop dividing upon treatment with NGF and produce axonal outgrowths (Fujita et al., 1989). Before the first report on PC12 cells appeared, researchers were struggling to understand how growth factors, specially NGF, works on neurons because primary neuron cultures are both difficult to harvest and culture, and moreover they are dependent on growth factors for survival. Thus it is difficult to evaluate the effects of growth factors on primary neurons. Today, PC12 cells are widely used to study the actions of the growth factors, since their survival is growth factor independent (Fujita et al., 1989).

Therefore the additional factors that are present in the FBN medium (M199) might cause the difference of axonal growth capability between FBNs and PC12, in monocultures.

.5 Biocompatibility of the GCGCFDS for heterogeneous cell

populations

Figure 13:The proliferation of homogenous (A) and heterogeneous (B) cell cultures in the GCG- CFDSs. Scaffolds were populated with either monoculture or heterogeneous cultures and cell proliferation was projected by the Alamar Blue Method. A. PC12 (circle), ECs (square) and FBN (triangle). B. ECs with PC12 cells (square), FBNs with ECs (diamond) in the GCG-CFDSs.

In the next step we evaluated the proliferation for two weeks of PC12 and FBN cells in GCG-CFDS upon co-seeding with the same number of ECs adjusted for growth in neuronal medium conditions (Figure 13). It was found that upon an initial phase of proliferation of 2 - 4 days, the cultures stopped proliferating reaching a constant density. We propose that the proliferation is representing the PC12 and EC cultures while FBN didn’t proliferate. Whatever the identity of the proliferating cell type, these experiments together with the confocal images of the cultures presented in

Figure 14, indicate survival and biocompatibility of the scaffolds for 7-13 days. The survival of these heterogeneous cultures in GCG-CFDS was further maintained up to four weeks (data not shown).

Figure 14: Differentiation of homogeneous and heterogeneous cultures of neurons in GCG- CFDSs. Confocal microscopy images of homogeneous PC12 cells after 7 days (A and B); the heterogeneous culture of the ECs with PC12 cells (C and D); the heterogeneous culture of the ECs with FBNs (E and F). The scale bar represents 100μm for A, C and E and 50μm for B, D and F.

Figure 15: Quantification of the confocal images according to two parameters, elongation of neurite outgrowths (A) and percentage of cells with neurites (B) in the GCG-CFDSs. A: The elongation ratio of the neurites of heterogeneous cultures of neurons with ECs (gray bars) compared to homogenous single neuronal cell cultures (white bars) B: The percentage of differentiated neurons in the heterogeneous cultures (gray bars) compared to homogeneous cultures of neurons (black bars).

Upon comparing the differentiation of PC12 cells under two different conditions, it was evident that their differentiation in the presence of ECs in the scaffold compared to their single growth was higher (N) by 35% and the length of their neurite outgrowths (E) significantly increased from a value of 4.1 to 6.8 (Figure 15 A, B and

Figure 14-C, D compared to A, B) In contrast, the differentiation of FBN in the heterogeneous scaffold containing ECs was not significantly different than the homogeneous cultures grown alone in the scaffolds (Figure 15 A, B and Figure 14 E, F compared to Figure 12 C, D).

Figure 16: The PC12 cell differentiation in the previously EC populated scaffolds at day 1, 3 and 7 according to two parameters: elongation ratio of the neurites (stripped) and the percantage of differentiated neurons (dotted).

 SUMMARY

The present study improves upon recently reported technologies for preparing longitudinally structured scaffolds (Stokols et al., 2006; Bozkurt et al., 2007; Mollers et al., 2009; Wu et al., 2010) by implementing a decreasing gradient temperature methodology to achieve controlled freeze-dried scaffolds (CFDS). This method is very convenient for the generation of unbranched, yet aligned channels, which may be advantageous in the design of neuronal pathways. The scaffold used for our in vivo experiments was composed of 2.5% gelatin cross-linked with 1% genipin (GCG) which exhibited longitudinal channels of ~50μm diameter and had an elastic modulus of ~51kPa. In addition, the scaffold was found to be biocompatible when applied to an in vitro angioneural cell interaction model by displaying cell-cell cross-talk (via

BDNF, VEGF, other common GFs and receptors) between EAhy926 endothelial cells

(ECs) and either PC12 or forebrain neuronal cells (FBN). Therefore, we surmise that the scaffold is appropriate for future testing in animal models with spinal cord injury.

The biodegradability of a scaffold is an important characteristic for future in vivo experiments since a nondegradable scaffold can cause inflammatory side effects related to the its continuous presence within neuronal tissues (Schmidt et al., 2003).

Biodegradable scaffolds also present a smaller risk of nerve compression since they degrade as the neuronal tissue regenerates, reducing volumetric pressures. Our present scaffolds have been found to be biodegradable as GCG-CFDS cross-linked with

44.2mM (1%) genipin degraded by 25% within 3 weeks while scaffolds cross-linked with 0.2mM (0.005%) genipin degraded by 80% within three weeks. Due to the fact that the optimum degradation rate of a scaffolds used for CNS injury has yet to be

54 determined (Hejcl et al., 2008), our GCG-CFDS are appropriate for implantation as their biodegradability rate, stiffness and other mechanical properties can be adjusted by cross-linker concentration.

The studies on the mechanical properties of the central nervous system structures varies according to whether the area measured is brain or spinal cord whether the measurements are in gray or white matter, and by the presence of pia matter and other technical variants (Bilston et al., 1996). On one extreme, Bilston and Thibault et al.,

(1996) as well as Ichihara et al., (2001) reported that the elastic moduli of human and bovine spinal cord tissues ranged between 940-1230kPa for white matter and

~1660kPa for gray matter. On the other extreme, Ozawa et al., (2001) reported that the spinal cord elastic modulus values were between 3 to 4 kPa. The mean shear stiffness of cerebral white matter was 13.6 kPa while that of gray matter was lower at around

5.2 kPa (Kruse et al., 2008). Chang et al., (1981) found that the elastic modulus of the intact spinal cord was 40 kPa while Mazuchowski et al., (2003) reported 89 kPa in the absence of pia mater. The GCG-CFDS developed in the present study are characterized by mean elastic moduli between 4.2-51 kPa, therefore suitable for implantation in brain and spinal cord but not appropriate for peripheral nerve regenerative purposes. This is due to the fact that peripheral nerve trunks were characterized as having very high elastic moduli in the range of 10-100MPa (Abrams et al., 1998; Rydevik et al., 1990). However, studies from our laboratory show that a different crosslinker or a higher concentration of gelatin (data not shown) could be used to obtain higher stiffness’s and there mimic PNS tissue.

We assume that matching the mechanical properties of GCG-CFDS to the

55 surrounding neuronal tissue in future in vivo experiments will more closely mimic the endogenous CNS tissue, as well as reduce inflammation and facilitate cell migration across the tissue–implant boundary. Indeed, preliminary experiments in our laboratory indicate that the GCG-CFDSs implanted within rats with spinal cord injuries integrated into the tissue and did not cause inflammation (data not shown).

Efforts were invested to characterize the biocompatibility of the GCG-CFDS towards angioneural cell models. In the presence of EAhy926 ECs, neuronal PC12 cells survived and proliferated up to 15 days in the scaffolds while FBN expressed long neurites as early as 3 days after populating of the scaffold (data not shown).

Present findings are in line with other studies indicating that micro-structured porous scaffolds are biocompatible for population with neurons supporting attachment, proliferation and elongation of aligned neurites (Stokols et al., 2006; Bozkurt et al.,

2007; Mollers et al., 2009; Wu et al., 2010). The advantage of the present GCG-CFDS relies in its ability to promote not only single cell populations but also proliferation and differentiation of heterogeneous cultures of endothelial and neuronal cells.

Therefore GCG-CFDSs may be used in future studies to generate blood capillary as well as neuronal pathways.

Results clearly indicate that the presence of ECs is beneficial for the differentiation of the neuronal cells. While primary neuronal FBN alone could differentiate within the scaffold; PC12 neuronal cell monocultures poorly differentiated within the same scaffolds. However, scaffolds primed with ECs for two weeks facilitated PC12 differentiation and suggest that ECs might release some angioneurins that accumulate in the scaffold and promote neurite outgrowth elongation. Another possibility to be

56 considered to explain the poor differentiation of monoculture of PC12 cells in the scaffold relates to the dimensional and mechanical properties of the scaffold.

Differentiation of PC12 in 3-D conditions was found to be delayed compared to 2-D conditions (Arien-Zakay et al., 2009). Also in PEG 3-D scaffolds, PC12 neurite outgrowth length was found to decrease as the stiffness of the scaffold increased (Scott et al., 2010). With this background and considering that PC12 cells were derived from adrenal medulla tumor tissue (which is a soft tissue characterized by a stiffness in the range of 5.9-6.3 kpa (Egorov et al., 2008)) we assume that 51kpa GCG-CFDSs used in these experiments possibly was too high for the PC12 cells therefore contributing to the inhibitory effect on PC12 cell differentiation.

 CONCLUSION& FUTURE WORK

Tissue engineering holds great promise for neural tissue repair and recent experimental work has made great strides within this field. Until now, engineers and scientists focused on mechanical, biological, chemical or electrical stimuli to enhance nerve repair. However, nerve regeneration is a very complex process and unfortunately none of these approaches could achieve an effective clinical therapy yet (Schmidt et al., 2003). Today, researchers are focusing on multiple stimuli (Miller et al., 2002;

Rauch et al., 2009) to better mimic the complex nature of the neural tissue. Studies further show that a coordinated regeneration approach is more successful than single stimuli strategies in promoting directional outgrowth of neurites (Miller et al., 2002).

For that reason, in this study, we designed a coordinated regeneration model to achieve a more successful neural regeneration by combining physical guidance and biological

57 cues.

Different studies show that on non-structured substrates, longitudinal axonal regrowth was very poor; axons tended to grow erratically and connection pathways could not meet with their target neurons (Davies et al., 1999; Smith et al., 2010).

Another critical process in CNS development is angiogenesis, or the formation of blood vessels, which is required for nutrient delivery as well as regulation of neuronal growth through paracrine signaling (Rauch et al., 2009). It has previously been shown that increased micro-vessel density correlates with regeneration and functional recovery of spinal cord tissue (Glaser et al., 2006; Kaneko et al., 2006). More importantly, regenerating axons grew along blood vessels in the spinal cord suggesting that the restoration of vascular networks would also help neuronal cell guidance

(Bearden et al., 2005). Thus, crosstalk between angiogenesis and neurogenesis are crucial for guidance of axonal growth across a spinal cord lesion.

Figure 17: The confocal microscopy image of the coculture of endothelial and PC12 cells in the longitudinal channels of the CFDSs (red represents CM-DIL labeled endothelial cells whereas green represents TUJ-1 labeled PC12 cells and blue represents the nuclei).

In this project, we designed a coordinated regeneration model in which our micro-

58 channeled scaffold provides physical guidance while endothelial cells provide physical and chemical signaling cues. In addition, we investigated the biologic response (such as neuritogenesis) of neuronal cells when implemented within this model. Ultimately, a degradable, genipin cross-linked gelatin scaffold was designed to restore spinal cord neuronal growth post-injury by using a controlled freeze-drying technique to improve both the mechanical properties of the scaffold as well as to give a longitudinal structure that can guide axonal connections throughout the injured tissue. To the best of our knowledge, the current study represents the first co-culture in micro-structured scaffolds and has the most appropriate range of mechanical properties for spinal cord injury repair. Moreover the degradation rate of these scaffolds was found to be similar to previous studies (Chang et al., 2009), which has been shown to be successful for peripheral nerve regeneration.

There are many studies in which PC12 cells were used as a neuronal cell model and differentiated in 3-D scaffolds. However in these studies, differentiation of PC12 in 3-D conditions was found to have delays when compared to 2-D conditions (Arien-

Zakay et al., 2009). Also in PEG 3-D scaffolds, PC12 neurite outgrowth length was found to decrease as the stiffness of the scaffold increased (Scott et al., 2010). The reason behind this delayed PC12 differentiation (when cultured as mono-cultures) may be because of the limited communication between cells, when compared to the 2-D conditions. Therefore we aimed to provide an ECM-like platform for cellular crosstalk within neurogenic niches. Our results showed that in the presence of ECs, both the axonal outgrowth percentage and the axonal outgrowth length were increased suggesting that the ECs may release some angioneurins or secrete extracellular matrix

59 components into the microenvironment that could promote neurite differentiation. This methodology is a promising approach for future neural tissue engineering and future studies are being developed with this methodology

In conclusion, many studies have promising results for therapeutic reconstruction of neural connections, however even if the axonal connections throughout the transected gap could be achieved, it cannot be foreseen what type of functional recovery can be achieved. Future spinal cord tissue engineering strategies should combine guidance as well as biomolecular related therapies for optimization of a permissive scaffold thorough an understanding of the signaling mechanisms and effects of the scaffold characteristics on axonal regeneration. Therefore GCG-CFDSs may be used in future studies to generate blood capillary as well as neuronal pathways.

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 APPENDIX

.1 Evaluate the effects of NGF & FGF2 on ECs & PC12 cells

The effects of NGF and FGF2 and a combination of these growth factors were evaluated and compared to the control (which is cultured without any additional GFs) on monocultures of ECs and PC12 cells for 7 days (Figure 18).

Figure 18: The effects of NGF (top right) and FGF2(top left) and a combination of these( bottom left) compared to the control (bottom right) on monocultures of ECs at day 7 (scale bar represents 100µm).

While there was no difference observed for PC12 cells (the PC12 cells in

monocultures remained round shaped even with additional growth factors- data not

shown), ECs seemed to have higher proliferation rate in the presence of additional

GFs (Figure 18). FGF-2 seemed to have a spreading effect on ECs (Figure 18 top

left), whereas NGF seemed to cause a flattening effect on the cells (Figure 18 top right). The best morphology was observed under the combined effects of FGF-2 and

NGF (Figure 18 bottom left). However none of these assumptions were quantified or

measured by any other technique.

.2 Evaluate the coculture proliferation dynamics for both endothelial and neuronal cells

It is a known fact that ECs are stimulated by shear stress both in vivo and in vitro.

They respond to shear stress with altered gene expression and morphological

reorganization (Dahl et al., 2010). Researchers claimed that oscillatory flow shear

stress stimulates adhesion molecule expression (Chappell et al., 1998) and promotes

endothelial cell migration and proliferation (De Paola et al., 1992) in vitro. In another

study, while laminar shear stress was suggested to induce cell alignment in the

direction of flow; turbulent shear stress was shown to stimulate endothelial DNA

synthesis, but in that case no cell alignment was observed (Davies et al., 1986).

However some of more recent studies show that non-uniform shear stress may cause

endothelial cell dysfunction (Cicha et al., 2008) and impair vascular functions

(Thacher et al., 2007). Hahn et al., (2009) summarizes the effects of shear stress, as

“Atheroprotective effects of high laminar shear are lost, whereas low or disturbed flow activates pro-inflammatory pathways”.

Figure 19: The Alamar Blue Test results of endothelial cells, PC12 cells and the co-culture of these cell lines under dynamic versus static conditions.

To better understand the effects of shear stress for seeded cells in the present

CFDSs, the ECs, PC12 cells and the co-culture of these were cultured under dynamic conditions by using a belly dancer and compared to the proliferation rate of static conditions (Figure 19). Results from our laboratory showed that the proliferation rate was higher under static conditions, when compared to dynamic conditions (Figure

19). Therefore we used static conditions for the rest of our experiments.

.3 Protocols

Cell preparation protocol for GFP-PC12 cell sorting

1. Prepare 1% BSA with HBSS (weight 60mg and dissolve it in 6ml HBSS) 2. Prepare 6 ml 1% BSA in HBSS to block tubes. (tubes are 5ml each so 6ml should be enough to block 6 tubes and to leave 200 μl in each of them, leave one tube for GFP-PC12 cells that are going to be sorted, two tubes for setting up the sorter with untransfected and transfected cells, and 3 tubes for collecting.) 3. Fill a collection tube (BD Falcon 5 ml polystyrene round bottom tube #352054) with 1%BSA in HBSS, wait 2 seconds and empty the tube to the second tube and so on. Prepare 6 tubes as total (we do not have this tube in our lab, so I have asked Dr. Mueller and she said I can take a few FACS tubes from them). 4. Leave around 200 μl of 1%BSA in HBSS in the collection tube for the cells to be sorted in. 5. Prepare a media with 3% BSA in HBSS (since HBSS has 0.50 g Glucose in it, it would be better for cells than PBS). 6. Detach cells from the flask by gently hitting to the bottom of the flask. Check for cell detachment under the microscope. 7. After cells detached, add 5ml of pre-warmed medium (3% BSA in HBSS) and collect the cells, pipette up and down (with a 5ml pipette) a few times by washing the bottom of the flask. 8. Then pipette up the suspension, press the tip of the pipette against the bottom wall of the flask and slowly spray the suspension while pipetting down, to separate cells. Repeat this step 5-6 times. 9. Check if the solution is in single cells under microscopy. If not, repeat the previous step till you get a single cell suspension. 10. Collect the entire cell solution from 4 different flasks, spin down and add 1ml fresh medium (HBSS with 3% BSA). 11. Prepare a suspension with 20x106 cells/ml 12. Also prepare  One tube with 2 million transfected cells in to FACS tubes (BD Falcon 5 ml polystyrene round bottom tube #352054) (The cells for setting up the sorter will be each in 300μl of sorting media.)  One tube with 2 million transfected cells in 300 μl of sorting media 13. Keep the cells on ice. 14. Filter the cells with previously autoclaved 35 μm filter directly before the sort that will be done in Flow Cytometry Core Facility, at Queen Lane Campus. (collect them into the FACS tubes that are blocked with 1% BSA in HBSS) 15. Collect the cells into the previously prepared FACS tubes. 16. Label T-25 flasks for different clones of GFP-PC12 cells. 17. Add 5 ml warm medium (7.5%FBS, 7.5% Equine Serum, 0.5% Pen.Strip in high glucose DMEM) to each T25 flask and then seed cells to the flasks.

Forebrain Dissection (general protocol) – (Note: prepare the scaffolds and seed them with ECs one week before if you aim to have co-cultures)

1. Arrange one large glass dish and two smaller glass dishes, and forceps. 2. Turn on UV light for 15 minutes prior to dissection. 3. Use (1-4) E8 chicks for most experiments. For all steps in which culture tubes are incubated in water bath, ethanol sterilize these tubes prior to opening them in the hood. 4. Wash the egg with ethanol over waste can, then use handle of large forceps to gently tap around top of eggshell and remove without puncturing embryo. 5. Break membrane and remove embryo with large forceps, placing on large glass tissue culture dish. Sever head from spinal column, and position in F12 medium with forebrain facing up. 6. Moisten fine forceps with F12 (F12HS10) media prior to removal of cerebral hemispheres to minimize tissue sticking. 7. Use fine forceps to peel away skull and meninges, first clipping a slit in the medial aspect of the skull and then clipping around the orbits. Sever the forebrain from the midbrain and gently scoop the forebrain into a small volume of F12 medium. 8. Position each hemisphere with medial side facing up, and gently clip away the upper portion of the forebrain (thin flap…trim off any thick portions). Mince the tissue with forceps (should be ~ 100 mm chunks). 9. Add 4 mL of CMF-PBS (in “clean” TC fridge) to a sterile 15 mL centrifuge tube and transfer the minced tissue into this tube (should have a final total volume of 5 mL). Vortex gently for 15 sec. (five 3 sec. pulses with vortex). 10. Incubate neurons in 37o C water bath for 10 minutes, during which time the trypsin can be removed from the freezer (with 1 min left) and allowed to thaw. 11. Spin cells for 4 minutes. 12. Remove CMF-PBS and add Trypsin solution to the cell pellet (Trypsin is diluted 1:1 with CMF/PBS at a total volume of 5 mL). Gently roll tube to mix solution. 13. Incubate in 37o C water bath for 5 minutes, vortexing once per minute (6 second vortex intervals) 14. Spin cells for 4 minutes. 15. Remove Trypsin solution and add back 2 mL of F12HS10, triturate 30X with a 1 mL pipette tip, and then add 3 mL F12HS10. 16. Spin cells for 4 minutes. 17. Remove F12HS10 and add back 2 mL M199 Medium, then resuspend by triturating 15X as before. 18. Use hemocytometer (use ~ 10 uL) to determine the cell density and calculate the volume needed (use 50 x 103 cells to seed one scaffold in M199 Medium.