THE CHARACTERIZATION of HYALURONIC ACID and POLYETHYLENE GLYCOL HYDROGELS for NEURAL TISSUE ENGINEERING by EMILY ROSE AURAND

THE CHARACTERIZATION OF HYALURONIC ACID AND POLYETHYLENE GLYCOL

HYDROGELS FOR NEURAL TISSUE ENGINEERING

EMILY ROSE AURAND

B.S., Colorado State University, 2009

A thesis submitted to the

Faculty of the Graduate School of the

University of Colorado in partial fulfillment

of the requirements for the degree of

Doctor of Philosophy

Neuroscience Program

2014

EMILY ROSE AURAND

This thesis for the Doctor of Philosophy degree by

Emily Rose Aurand

has been approved for the

Neuroscience Program

John Caldwell, Chair

Kimberly Bjugstad, Advisor

Bruce Appel

Curt Freed

Robin Shandas

John Sladek

Date 03/10/2014

Aurand, Emily Rose (Ph.D., Neuroscience)

The Characterization of Hyaluronic Acid and Polyethylene Glycol Hydrogels for

Neural Tissue Engineering

Thesis directed by Assistant Professor Kimberly Bjugstad.

ABSTRACT

Neural tissue engineering through the use of biomaterials holds great promise for treating a wide variety of neurological disorders. The customizable nature of hydrogels provides the opportunity to mimic the brain’s unique extracellular matrix (ECM). Hydrogels can be used to recreate this ECM environment to support neural cells in vitro, through 3D culturing, or during transplantation procedures. To be effective, hydrogels must be characterized chemically, physically, and mechanically, and the biocompatibility of these materials with neural cells and brain tissue must be defined. Twenty-five hydrogels were created from ratios of hyaluronic acid (HA) and poly(ethylene glycol) (PEG). Hydrogels were assessed for the properties of polymerization, degradation, and compressive modulus, and the cytocompatibility with encapsulated neural progenitor cells (NPC) from fetal and adult sources. The physical and mechanical properties of the hydrogels were found to be dependent on the polymer concentration. Additionally, the compressive moduli of the hydrogels were comparable to rodent brain tissue, indicating that the hydrogel formulations developed were physiologically relevant. Subsequently, NPC derived from fetal and adult rats (fNPC and aNPC, respectively) were

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encapsulated within the hydrogels. Twenty-four hour cell survival was highest at lower concentrations of HA and PEG. Three-week fNPC and aNPC differentiation was demonstrated to be influenced by mechanical properties. Fetal-NPC generally produced greater numbers of astrocytes in stiffer hydrogels, while increased numbers of neurons were observed in softer hydrogels. Greater numbers of aNPC became neuronal, regardless of stiffness. When two chosen hydrogels were used to implant NPC into the brain, the results suggested that encapsulated NPC survived at up to 50% two months post-implantation,

indicating good cytocompatibility. Further, the implanted cells were able to

migrate from the hydrogel into the surrounding brain tissue farther than

unencapsulated cells. Immunolabeling for glial cells demonstrated that the

hydrogels elicited a similar immune response as control treatments, establishing

the histocompatibility with brain tissue. Based on these studies, HA-PEG

hydrogels were biocompatible and could be used therapeutically in the brain.

Further modifications and specializations of these hydrogels, such as the

inclusion of growth factors or attachment factors, may provide specific

therapeutic support for encapsulated cells and/or neurodegenerative disorders.

The form and content of this abstract are approved. I recommend its

publication.

Approved: Kimberly Bjugstad

I dedicate this work to my parents for their ever present encouragement and

enthusiasm; and to Sammie and Lucy.

ACKNOWLEDGMENTS

Thank you to my mentor and advisor, Dr. Kim Bjugstad for her guidance, enthusiasm, and creativity. I also thank my advisory committee: Drs. Bruce

Appel, Curt Freed, Robin Shandas, John Sladek, and my Chair, Dr. John

Caldwell, for their advice and direction.

I would like to thank those individuals whose assistance along the way contributed to the accomplishment of this work, in particular: Dr. Angela

Rachubinski, Dr. Kyle Lampe, Connie Brindley, Jennifer Wagner; and the members of the Neuroscience program, including my peer group: Pam Lopert,

Pirooz Parsa, and Josh St.Clair.

I also acknowledge the support of the Department of Pediatrics and the

Dean’s Academic Enrichment Fund for the majority of my financial support.

I must also thank my early science mentors, including members of the

Tobet Lab, Dr. Joan King, and especially Dr. Stu Tobet for recognizing the value

of undergraduates in scientific research. I am also greatly appreciative of the late

Mr. Dan Preble, who, albeit unknowingly, allowed me to discover my passion for

neuroscience.

TABLE OF CONTENTS

CHAPTER

I. INTRODUCTION ...... 1

II. LITERATURE REVIEW ...... 5

Introduction ...... 5

Polymers and Hydrogels ...... 9

Polymerization...... 12

Degradation ...... 15

Mechanical Properties and Physical Architecture ...... 19

Drug Delivery ...... 30

Biocompatibility ...... 36

CNS Environment and Immune Response of the Brain ...... 36

Contributions of Hydrogel Properties to Biocompatibility ...... 42

Summary ...... 49

Acknowledgements ...... 50

III. HYDROGEL FORMULATION DETERMINES CELL FATE OF FETAL AND ADULT NEURAL PROGENITOR CELLS ...... 51

Introduction ...... 51

Methods ...... 54

Hydrogel Formulations and Polymerization ...... 54

Hydrogel Degradation ...... 56

Mechanical Testing ...... 57

Fetal NPC Derivation ...... 58

Adult NPC Derivation ...... 59

NPC Encapsulation ...... 61 vii

NPC 24-hour Survival ...... 61

Cell Numbers and Differentiation at Three Weeks ...... 63

Immunocytochemistry and Microscopy ...... 64

Data Analysis and Statistics ...... 65

Results ...... 66

Polymerization...... 66

Degradation ...... 67

Compressive Modulus ...... 68

Fetal NPC Survival and Differentiation ...... 69

Adult NPC Survival and Differentiation ...... 74

Discussion ...... 78

IV. IN VIVO BIOCOMPATIBILITY OF HYALURONIC ACID AND POLY(ETHYLENE GLYCOL) HYDROGELS ...... 87

Introduction ...... 87

Methods ...... 91

Hydrogel and NPC Preparation ...... 91

Implantation Surgery ...... 94

Tissue Preparation and Immunohistochemistry ...... 95

Microscopy and Statistics ...... 97

Results ...... 99

Cytocompatibility ...... 99

Histocompatibility ...... 102

Discussion ...... 105

V. SUMMARY AND CONCLUDING REMARKS ...... 114

Hydrogel Properties ...... 114

viii

In Vitro Biocompatibility ...... 118

In Vivo Biocompatibility ...... 120

Future Directions ...... 122

Conclusions ...... 124

REFERENCES ...... 125

LIST OF TABLES TABLE

1. Theoretical applications of user defined hydrogels for the treatment of neurological disorders ...... 7

2. Matrix of 25 hydrogels studied ...... 55

3. Brain tissue mechanical properties ...... 68

4. Summary of selected results from in vitro studies (Chapter III) ...... 92

LIST OF FIGURES FIGURE

1. Hydrogels can be used to encapsulate cells, microparticles, or therapeutics for multiple delivery purposes ...... 16

2. Sample parameters for selected hydrogel properties as a function of time (t) ...... 20

3. Survival and fate of peripheral cell types as a function of biomaterial stiffness ...... 22

4. Survival and fate of neural stem cells as a function of biomaterial stiffness ...... 24

5. Hydrogel pore size changes as a function of precursor molecular weight ...... 27

6. PEG-PLA-based hydrogels implanted into the rat brain induced a long term neuroimmune response similar to that observed in sham brains penetrated with a needle...... 41

7. Immunolabeling of undifferentiated NPC prior to hydrogel encapsulation ...... 60

8. Hydrogel physical properties ...... 67

9. Hydrogel mechanical properties ...... 70

10. Presence of fNPC at 24 hours and three weeks ...... 71

11. Fetal-NPC differentiation ...... 74

12. Presence of aNPC at 24 hours and three weeks ...... 75

13. Adult-NPC differentiation ...... 77

14. Morphology of differentiated NPC three weeks post-hydrogel encapsulation ...... 83

15. BrdU+ cells were found in the brains of all animals regardless of implantation method ...... 100

16. The greatest migration along the rostral-caudal axis away from the implant track was seen in Hydrogel “N” implanted brains ...... 101

17. Implanted NPC at the striatal and corpus callosal interface express both BrdU and Nestin...... 103

18. Both astrocyte and microglia presence indicates that the hydrogels were biocompatible ...... 104

19. GFAP and Iba1 immunoreactivity were used as indicators of biocompatibility ...... 105

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LIST OF ABBREVIATIONS

ANOVA Analysis of variance

aNPC Adult-derived neural progenitor cells

AP Anterior-posterior

bFGF Basis fibroblast growth factor

βTubIII Beta-Tubulin III

BrdU 5-bromo-2’-deoxyuridine

CC Corpus callosum

CTX Cortex

DAB 3,3’-diaminobenzidine

DAPI 4',6-diamidino-2-phenylindole

DIV Days in vitro

DV Dorsal-ventral

ECM Extracellular matrix

EGF Epidermal growth factor

fNPC Fetal-derived neural progenitor cells

GFAP Glial fibrillary acidic protein

HA Hyaluronic acid

HBSS Hanks’ balanced salt solution

Iba1 Ionized calcium binding adaptor molecule 1

ICC Immunocytochemistry

IHC Immunohistochemistry

kPa Kilopascals

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MAP2 Microtubule-associated protein 2

ML Medial-lateral

NPC Neural progenitor cell

PB Phosphate buffer (0.1M)

PBS Phosphate buffer saline

PEG Poly(ethylene glycol)

SEM Standard error of the mean

ST Striatum

SVZ Subventricular zone

TCP Tissue culture polystyrene

wt% Percent weight

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CHAPTER I

INTRODUCTION

Recently, neural tissue engineering has incurred greater interest by the neuroscience and bioengineering fields as an approach to treating neurological disorders. The goal of neural tissue engineering is to recreate damaged brain tissue by combining biomaterials, designed to act as the extracellular matrix, with neural cells and/or other therapeutic components. Theoretically, this could result in a biomaterial-cell composite prepared in the lab that resembles neural tissue, which could then be placed into the site of damage. The ideal composite would allow for functional integration of the contained cells and the surrounding tissue, potentially mitigating the effects of the disorder.

Of these biomaterials, polymer-based hydrogels can be developed with certain definable properties, which can be tailored by adjusting the polymer chemistry, to make them applicable for neural tissue engineering. However, very few studies have addressed how these properties affect the biocompatibility with tissues of the central nervous system. A material is defined as biocompatible if it is non-toxic to the cells with which it is in contact and it does not provoke an immune response beyond a normal reaction. Conflicting results from scientists using hydrogels that are not thoroughly characterized has demonstrated the importance of investigating hydrogel properties and establishing biocompatibility in vitro and in vivo before considering a material suitable. For example, Matrigel has been used extensively for cell culturing and even for in vivo tumor studies;

1 however, when used in the brain, it provokes a substantial neuro-immune response.

Thus, to make biomaterials a reasonable tool for neural tissue engineering, the goal of this research was to identify hydrogels with properties that were physiologically and functionally relevant to brain tissue, and further, to identify hydrogels which could be translated from in vitro to in vivo applications while remaining biocompatible.

To this end, the studies described in this thesis first aimed to identify ratios of hyaluronic acid and poly(ethylene glycol) polymers which form hydrogels that had physical properties applicable to neural tissue engineering and mechanical properties that matched those of brain tissue. The physical properties of polymerization and degradation were explored. A functional polymerization process can allow for the encapsulation of neural cells, while degradation is necessary for the hydrogel to be biocompatible with brain tissue. The mechanical property of compressive modulus was used to match hydrogel stiffness to the stiffness of relative brain tissues, as it has been established that the matching of mechanical properties can improve biocompatibility.

To study the effects of hydrogel properties on neural cells in vitro, neural progenitor cells derived from fetal and adult rats were encapsulated within the hydrogels. The survival and differentiation of these cells as a function of hydrogel properties was explored. For example, previous studies by other researchers have demonstrated the contributions of hydrogel compressive modulus to the survival and maturation of neuronal cells vs. glial cells. Thus, the studies in this

thesis were designed to elucidate which formulation of hydrogel(s) had a

compressive modulus which supported neuronal differentiation over glial

differentiation and vice-versa. Also of particular interest was how differences in the age of the animals from which the cells were derived affected the response of

the encapsulated cells to the hydrogel.

Finally, the in vivo biocompatibility of the hydrogels was explored by

implanting two biologically-relevant hydrogels containing neural cells into the

brain. The hydrogels were chosen based on the physical and mechanical

properties and on the results achieved in vitro. The outcomes of hydrogel-

mediated delivery of neural progenitor cells into the brain were used as a

measure of in situ cytocompatibility. It was hypothesized that the hydrogels might

protect the encapsulated cells from the implant procedure and natural

inflammatory reaction, resulting in increased survival of implanted cells. The

neuroimmune response to the implanted material was used as a measure of

histocompatibility with the brain. Establishing this hydrogel biocompatibility allows

for the ability to directly translate hydrogel use from in vitro to in vivo and

contributes to the development of clinically applicable biomaterials for neural

tissue engineering.

Much of the information found in this thesis has been previously published

as review and original research journal articles. Chapter II is an integration of two

review pieces published as Aurand, et al. (2012) Neuroscience Research, 72(3):

199-213; and Aurand, et al. (2012) Journal of Functional Biomaterials, 3(4): 839-

863. The information found in Chapter III has been published as Aurand, et al.

(2014) Stem Cell Research, 12(1): 11-23. Finally, the information found in

Chapter IV is in preparation to be published as Aurand and Bjugstad, (2014).

CHAPTER II

LITERATURE REVIEW1

Introduction

Millions of individuals in the United States are affected by neurological disorders [1]. From stroke and traumatic brain injury, to Alzheimer’s and

Parkinson’s diseases, the causes and manifestations of these disorders vary greatly, as do the impacts and outcomes. Likewise, treatments for these disorders are numerous and diverse; however, very few treatments result in complete recovery and many diseases have limited treatment options and no cure.

One of the reasons for the lack of effective therapies for neurological disorders is the limited regenerative abilities of the central nervous system [2, 3].

For example, in individuals who have suffered a stroke, there is a sustained loss of functional tissue and thus treatments are often limited to rehabilitation of lost functions, such as speech or movement. Similarly, the damage to cells and brain structures associated with neurodegenerative disorders such as Parkinson’s and

Alzheimer’s diseases, for the most part, are permanent. Current attempts to replace lost neural cells by implantation in disorders like Parkinson’s disease and stroke have had limited success and enormous variability between implanted patients [4-9]. That there have been some successful outcomes provides proof of concept that replacing lost neural cells through neural grafting or implantation

1 All or part of this literature review has been previously published as Aurand, et al. (2012) Neuroscience Research, 72(3): 199-213; and Aurand, et al. (2012) Journal of Functional Biomaterials, 3(4): 839-863. 5

can be a viable treatment option. However, the large scale loss of grafted cells in

the days following implantation is most likely the biggest contributor to their

variable success [10-14]. To improve the outcomes of cell implantation into the

central nervous system (CNS), there needs to be some method of protecting the

neural grafts from the host response to the implant procedure and for guiding

graft growth and integration. The incorporation of cells into protective

biomaterials could enhance and universalize the success of neural implants.

Biomaterials, such as polymer-based hydrogels, can be used as delivery tools not only for cell grafting, but also to deliver drugs, viral constructs, DNA, growth factors, and other therapeutics, with precise delivery into a defined brain region and with specified temporal release [15-25]. Considering the diversity of options polymers provide, their application to neurological disorders may be limited only by the creativity of the researcher (Table 1). For example, a multi- delivery hydrogel could be formulated to help treat pediatric neuronal ceroid lipofuscinoses (Batten’s disease). Batten’s disease results from a lysosomal enzyme deficiency [26]. Previous attempts to treat Batten’s disease included enzyme replacement therapies, gene therapies to transduce functional gene copies, and neural stem cell implants which produce operative enzymes [26].

Because it is possible to have control over multiple delivery options at once, it is conceivable that a single hydrogel could incorporate all three treatment options providing immediate enzyme delivery to at-risk host cells, to insert grafted cells with functional enzyme, and to induce host cells to produce the enzyme themselves. Through the functional modification and customization of hydrogels

with cells and therapeutics, neural tissue engineering strategies could achieve

greater levels of success.

Table 1. Theoretical applications of user-defined hydrogels for the treatment of neurological disorders.

Neurological Theoretical Hydrogel Application Disorder

Spinal Cord Injury, • Hydrogel tube lined with factors to enhance axonal Parkinson’s bridging across spinal lesion or between the Disease substantia nigra and striatum.

Stroke, Traumatic • Hydrogel provides architectural support in the lesion Brain Injury, cavity, maintaining structural integrity and protecting Gliomas overlying structures.

Traumatic Brain Injury, Stroke, • Neural progenitor cells encapsulated by a hydrogel Alzheimer’s are protected from the wound healing process invoked Disease, by the surgical procedure. Huntington’s Disease, • Encapsulation of progenitor cells and differentiation Parkinson’s factors for in situ differentiation. Disease

• Temporally and spatially control the application of Down Syndrome growth factors during critical windows of neural development.

• Load hydrogel insert with functional enzyme and gene Glutaric Acidemia containing vectors as a preventative step against metabolic crises that result in striatal lesioning.

In attempting to re-engineer neural tissues of the CNS using hydrogels,

two specific characteristics must be taken into account that make this system

unique compared to tissues found in the rest of the body. First, the CNS is

considered an “immune-privileged” tissue. It has an independent immune

response governed by resident cells of the brain rather than the immune cells of 7

the body. Second, the CNS has a limited capacity to repair damage and grow

new cells. These two characteristics combine to make neural tissue engineering

difficult. To be most effective, re-engineering damaged brain tissue involves

implanting replacement neural cells. Unfortunately, the act of implanting these cells engages the neuroimmune system to respond to the damage created by penetrating the brain. The goal of the neuroimmune response, similar to the immune response in the body, is to mitigate the damaging elements inducing neuroinflammation, to contain the damage by building a glial scar, and finally, to activate the healing process to repair damaged, but still surviving, neural cells.

Most replacement cells, along with neighboring host cells, are lost during the

acute neuroinflammatory response [10-14, 27, 28]. Cell survival can range from less than 1%, to only as high as 10% [10-12, 14, 29-32], depending on the source of the replacement cells and where they are implanted. The ability to improve survival and promote replacement neural cell integration into the CNS would greatly advance neural tissue engineering.

Because of this compromised survival of implanted cells, the use of hydrogels to deliver cells to a site of damage within the brain or spinal cord is of significant interest. Replacement neural cells encapsulated within a hydrogel may be protected from the acute neural inflammatory response, and thus, much more likely to survive [27, 33-38]. Because hydrogels can be formulated from many different types of polymers and with a broad variety of material properties, they are an ideal material for replicating the three dimensional extracellular matrix (ECM) of neural tissue [39-50]. However, before hydrogels can be used to

overcome the obstacles of replacement neural cell implantation, they must be appropriately designed to be biocompatible with the encapsulated neural cells and with the CNS environment. If the hydrogel is not biocompatible, the neuroimmune response to the implant may create additional damage to that the

brain is not equipped to repair.

Polymers and Hydrogels

Polymers are large chains of repeating structural units called monomers.

By this broad definition, DNA and RNA are considered natural polymers with their

repeating monomers of nucleotides; and, while it is more common to think of

polymers as rubber or plastics, their industrial uses are much broader, including

skin creams, toothpastes, contact lenses, and joint replacements. Because

polymers are incredibly versatile in their composition and formation, research is

currently focused on better understanding polymer properties and potential

applications. This includes expanding the library of existing materials to explore

novel functions and increased control over material properties. Of particular

interest are applications of polymers as biomaterials for treating brain disorders,

through neural tissue engineering [46, 51-56], site specific drug delivery [16, 19,

21, 57-59], or as carriers for neural cell implantation [54, 60-64].

Hydrogels are one type of polymer with chemical and physical properties

that make them highly suited for use in biological systems. Hydrogel polymers

have a high water content (i.e. >90% water) due to their hydrophilic nature and,

because of this, have attracted attention for cell culture and tissue regeneration.

Hydrogels can be pliant and flexible like soft tissues, rigid like cartilage or bone, or elastic to mimic skin or blood vessels [50, 65-76]. The ECM of the brain, which provides the macroscopic architecture and supports neural cell survival, migration, and differentiation, is formed from a hyaluronic acid-based hydrogel backbone [77-80]. Compared to hydrogels composed of only natural materials, however, synthetic hydrogels can be created to more accurately imitate the physical and mechanical characteristics of the ECM. The advantage of synthetic hydrogels is the ability to tightly control polymerization, degradation, and biocompatibility. Additionally, cells, drugs, or other therapeutic agents can be incorporated into the synthetic hydrogel to fill a functional need [25, 42, 48, 54,

81-84].

Hydrogels most often used in neuroscience applications not only have a high water content, but also fall under the definition of chemically cross-linked synthetic hydrogels with viscoelastic properties. Synthetic hydrogels, unlike those based on natural materials such as gelatin, agarose, or Matrigel, are better chemically defined, and are biologically inert. This reduces data variability between in vitro studies and decreases potential immunorejection when inserted into the brain. Having viscoelastic properties provides the hydrogel with the ability to retain its polymerized shape and retain the strain resistance of a viscous liquid.

Retaining three-dimensional (3D) shape is an important characteristic for tissue engineering and will be discussed in later sections.

This literature review will cover properties of hydrogels that are believed to be important considerations before using them in living tissues, specifically the

10 brain, with a focus on 3D neural tissue engineering. Polymerization, or gelation, is the transitioning process from a liquid to a solid. How a hydrogel polymerizes and how quickly this process happens can determine how the hydrogel can be used. Degradation is the process of breaking the polymer network bonds, resulting in the erosion of the hydrogel and the release of monomer and oligomer precursors. Similar to polymerization, how a hydrogel degrades, and how quickly the process occurs, can determine how the hydrogel is used therapeutically. For example, a fast degrading hydrogel could be used to quickly deliver an immunosuppressive drug to brain tissue for immediate action against the inflammatory response, whereas a slowly degrading hydrogel could protect encapsulated cells from this same initial inflammatory insult and provide a longer- lasting physical scaffold for rebuilding tissue. A major benefit of using hydrogels is the ability to incorporate different types of molecules and to encapsulate in 3D a variety of different cell types. The performance of a hydrogel in brain tissue and the release of cells or therapeutics from the hydrogel depend on the mechanical and chemical properties of the hydrogel and how it is defined during polymerization and degradation. Finally, biocompatibility, specifically within the brain, is an especially important consideration, as the brain is a partially immune- privileged site reacting independently of the peripheral immune system.

Biocompatibility also includes the viability and function of cells encapsulated within the hydrogel carrier. It is important to note that hydrogels can behave differently in vitro and in vivo. For example, in vitro degradation is usually dependent on hydrolysis through unlimited access to water and/or a single

11 enzyme-based degradation (e.g. hyaluronidase), whereas in vivo access to water is limited and multiple enzymes could be present to degrade the hydrogel. Thus, researchers should consider translational aspects, from in vitro to in vivo, when designing a hydrogel.

Polymerization

Polymerization is the chemical process by which monomer precursors are reacted to form a polymeric structure. This process can be used to create well- hydrated, covalently cross-linked hydrogels from monomer precursors, such as poly(ethylene glycol) (PEG) or hyaluronic acid (HA) [41, 81, 85-87]. It should be noted that in many cases, including the two examples above, the hydrogel precursors begin as independent polymers, rather than simply monomers.

Polymerization generates covalent bonds between monomers, called cross-links

[86, 87]. Covalent cross-links allow precise control of the final cross-link density – the number of bonds between monomers in the final hydrogel product. The cross-link density can alter the degradation rate, mechanical properties, and functionality of the hydrogel [82]. The density of functional groups, such as acrylates, thiols, or polyesters, on the monomer backbone also alters the number and space between cross-links made during the process of polymerization. An increase in the density of functional groups will result in a greater number of cross-links in the hydrogel and tighter average mesh structure. Alternatively, decreasing the length of the cross-linker can also decrease the average mesh size. In addition to chemical cross-linking, another class of hydrogels can be

formed by the physical association of components, often by hydrogen-bonding at

the molecular level. Examples of physically cross-linked hydrogels include alginate, agarose, Matrigel, collagen, and a recently described two-component protein hydrogel [33, 88-91].

Hydrogel formation can be temperature-dependant (thermopolymerization)

[92-97], pH-dependent [92, 94, 98], or visible or ultraviolet (UV) light-dependent

(photopolymerization) [86, 99-101]. Both temperature- and pH-dependant polymerization are forms of physical cross-linking and can be reversible. The benefits of a reversible hydrogel are best demonstrated in the field of diabetes research: a pH-dependent hydrogel was created to release insulin in response to excess glucose and gluconic acid, which alter the pH of the tissue surrounding the hydrogel [102]. Thus, this hydrogel formulation allows insulin to be released only when it is needed, just as release would occur in normal tissue. Many temperature- and pH-dependant hydrogels are formulated to gel under physiological conditions, which can be ideal for in situ polymerization [103]. In

this process, the un-polymerized hydrogel solution is injected into the tissue site

and subsequently polymerizes in response to the difference in body temperature

or pH. This can allow the hydrogel to fill an irregularly shaped lesion site or coat a

surface. However, if the conditions during hydrogel injection exceed standard

physiological conditions, the hydrogel may not polymerize. If the precursors are

not adequately controlled before or during delivery, the materials can have

unintended behaviors, such as clogging the syringe and/or causing some

discomfort to the patient at the site of injection [92]. Hydrogels that polymerize in

13 situ can be limited in their ability to encapsulate cells and/or proteins or may fail to provide a sometimes necessary macroscopic architecture. In contrast, chemically cross-linked synthetic polymers provide researchers with highly tunable materials whose gelation behavior, and therefore physical and chemical properties, can be carefully adjusted with intricate control.

Photopolymerization is a common laboratory process because of the rapid bond formation that occurs when exposed to white or UV light. The addition of a photoinitiator allows the hydrogel to polymerize by generating reactive species that catalyze the polymerization process. UV exposure and the generation of reactive species can be a concern, especially when cells are to be incorporated into the hydrogel, because this can, in some cases, result in decreased viability of the cells [104-106]. The addition of lactic acid, a commonly used functional unit in biomaterials, can mitigate some of the effects of the reactive species on neural stem cells [104, 107]. Despite this, the advantages of photopolymerization may override any concerns regarding how the hydrogel is polymerized.

Ex vivo polymerization of such materials allows for the precise control over size and shape of the hydrogel generated. For example, the amorphous shape produced by the in vivo polymerization of a hydrogel would not benefit peripheral nerve regeneration, which needs a strand or tubular shape implanted parallel to the nerve to increase axonal outgrowth [60, 108]. The tubular shape of the hydrogel implant mimics the Schwann cell column that allows axons to grow protected on the inside of the tube or column [60, 109-111]. Additionally, culturing

Schwann cells onto a grooved inner surface of a polymer tube further enhances

peripheral nerve regeneration by guiding the direction of outgrowth and

concentrating the adhesion molecules and growth factors inside the tube

adjacent to the desired area of neuronal growth [112].

One of the more exciting properties of an ex vivo polymerizing hydrogel is the ability to fabricate a complex or layered hydrogel form (Figure 1, Table 1) [20,

58, 113-119]. This process can be beneficial if finer control over the spatial or temporal aspects of the hydrogel is desired. For example, in an attempt to control the temporal release of growth factors, Lampe et al [58], formulated several hydrogel strands in which two distinct microparticle formulations were incorporated into one hydrogel strand. Each type of microparticle carried a different protein and was formulated for a different release rate (Figure 1; see the

Drug Delivery section in this review for more details). Some complex hydrogel

formations are based on the “layer-by-layer” (LbL) assembly of thin, multiple

layered films [20, 113-116] (Figure 1). These films can carry DNA, RNA,

oligonucleotide constructs, or almost any therapeutic agent for precise spatial

exposure of agents to cell surfaces.

Degradation

Many hydrogels formulated for tissue engineering and/or the release of

therapeutics are designed to be biodegradable (or bioresorbable) to reduce the

complications of tissue scarring and glia tumor formation from permanent

Figure 1. Hydrogels can be used to encapsulate cells, microparticles, or therapeutics for multiple delivery purposes. The hydrogel material should be formulated to provide a specific benefit to the encapsulate or host, such as protection from the immune response for encapsulated cells, spatial seclusion of trophic factors, or temporal release of therapeutic compound. A. Illustrates the use of a hydrogel strand to deliver two different types of microparticles in a spatially- and temporally-defined manner. Lampe, et al. [58] employed PLGA- based microparticles loaded with BDNF (circles) or GNDF (stars) with differing release kinetics encapsulated within PEG-based hydrogel strands to demonstrate defined release of protein from the hydrogel into the tissue (see text for more details). B. Wilson, et al. [114] used the layer-by-layer formulation technique to encapsulate pancreatic islets (pentagons) within layers of different formulations of hydrogel (encompassing lines). The authors fluorescently labeled the different hydrogel formulations to distinguish layers around the islets. The ability to form multiple unique layers of hydrogel around an encapsulate suggests the potential for each layer to contribute a specific, distinct function for both the encapsulate and the host tissue. C. Using centrifugal casting methods, Mironov, et al. [119] used HA-based hydrogel seeded with cadiovascular progenitor cells (small circles) to produce hydrogel tubes lined with cells. The overall shape of hydrogel structures, such as tubes or conduits, allow for the production of specialized tissue structures, such as blood vessels or conduits for axon bundles.

implants, such as those seen around implanted electrodes [120-123]. The process of hydrogel degradation is the cleavage of labile bonds within the polymer and the monomer components, resulting in smaller chemical groups that

can be cleared from the implant site. Often, the degradation of a hydrogel occurs

through the hydrolytic or enzymatic cleavage of bonds made during

polymerization [57, 62, 86, 107, 124-126]. The addition of degradable monomeric units, such as poly(α-hydroxy acids) (i.e. poly(dl-lactic acid) and poly(glycolic acid)), and poly(ε-caprolactone), allow the hydrogel monomers to separate when the bonds are cleaved by water [15, 57, 62, 81, 86, 87, 104, 107, 127]. In hydrogels with poly(α-hydroxy acids), the ester bonds between poly-lactic or

poly-glycolic acid units are hydrolyzed, leading to the production of the simple

products of lactic or glycolic acid [86, 104, 128]. Hydrogels can also be

engineered with enzyme specific cleavage sites by adding sequences of specific

peptides between macromers. This allows hydrogel degradation in the presence

of an appropriate, tissue specific enzyme [18, 44, 62, 81, 124, 129-132]. For example, Rice, et al. [131], formulated hydrogels with poly(caprolactone) units which, in addition to hydrolysis, were enzymatically degraded upon addition of exogenous lipase. This allowed the authors to design the hydrogel chemistry with a degradation rate that matches the rate of production of ECM components by encapsulated chondrocytes [131, 133]. This type of degradation is often the focus for cell-mediated degradation, especially by encapsulated cells, and cell- specific controlled release of bioactive molecules [44, 81, 129, 132-135]. Cell- mediated enzymatic degradation may be helpful to encapsulated neural cells that could preferentially degrade the hydrogel material at sites of neurite extension, allowing for increased neuritic growth and pathway formation. A hyaluronic-acid based hydrogel developed by Park, et al. [136], was designed with matrix

17 metalloproteinase (MMP) sensitivity which allows MMPs produced by encapsulated cells to degrade the hydrogel scaffold. Along with an IKVAV (Ile-

Lys-Val-Ala-Val)peptide sequence and brain-derived neurotrophic factor (BDNF), these hydrogels allowed for scaffold remodeling by encapsulated mesenchymal stem cells and stimulated neuronal differentiation and neurite outgrowth [136].

Finally, the rate at which a hydrogel degrades over time can be determined in part by the number of degradable units incorporated by the researcher; typically, the greater number of degradable moieties, the faster the rate of degradation [57,

86, 87, 101, 107, 124, 131, 135, 137].

Hydrogel degradation can occur via surface or bulk erosion [138, 139].

The type of degradation which occurs is dependent on the amount of incorporated degradable units, cross-link density, and access of water and/or enzymes to the interior of the hydrogel [138]. Surface degradation occurs when water and enzymes cannot penetrate the interior of the hydrogel, due to high cross-link density or limited access to cleavage points, forcing the surface or exterior bonds to cleave first. Conversely, bulk erosion is degradation that occurs essentially homogeneously throughout the hydrogel, with interior and exterior bonds being cleaved simultaneously. Bulk erosion is common in hydrogels because of their high internal water content and fast diffusion and is even more likely to occur in hydrogels with a lower cross-link density and those with a greater amount of degradable polymer [138, 139]. In cases of nerve repair where the structural support is necessary even in a degradable hydrogel, surface erosion can be preferred, as it will allow the hydrogel to continue acting as a

conduit scaffold [140]. Similar to polymerization, the rate of degradation and

process by which it occurs influences mechanical properties of the hydrogel and

will be further discussed in the next section.

As suggested earlier, degradation can be especially important in

decreasing the immunological response to an implanted hydrogel. The temporary

presence of a biodegradable hydrogel is most often preferable to a permanent

implant, which can encourage a lasting immune response and glial scarring.

When designing a hydrogel for a specific application, the degradation rate can be

imperative to both the function of the hydrogel and the response of a host. As a

scaffold, a slower degradation rate gives the cells the time to develop their own

extracellular matrix and to extend processes and reintegrate into the host neural

circuitry. However, faster clearance of the hydrogel can be beneficial in vivo, as

quick degradation contributes to a reduced immune response. It is necessary to

balance the benefit of a hydrogel as a scaffold for developing cells and tissues

with the response of a host to a foreign body/material.

Mechanical Properties and Physical Architecture

Mechanical and physical features which are helpful to consider when constructing a hydrogel include the strength and stiffness, the mesh size and porosity, the overall architecture, and physical dimensions of the hydrogel. Many of these characteristics can significantly contribute to the effectiveness of the hydrogel, from cytocompatibility with encapsulated cells, to biocompatibility within tissues, and how therapeutic agents are released from the hydrogel.

The mechanical environment with which cells may interact can affect the viability and behavior of the cells; thus, the use of a hydrogel with specific mechanical properties can affect the function of encapsulated cells or the subsistence of a hydrogel in tissue [66-69, 141-144]. As mentioned above, the chemical properties of a hydrogel can determine the cross-link density, which in turn contributes to defining the mechanical and physical properties of a hydrogel

– specifically the stiffness of the polymerized product (Figure 2) [50, 84, 145].

Figure 2. Sample parameters for selected hydrogel properties as a function of time (t). Compressive modulus (solid line) is a measure of the hydrogel strength and increases as the hydrogel polymerizes. During polymerization, cross-links form between monomers, increasing the cross-link density (dotted line). The number of cross-links declines as the bonds are hydrolyzed or cleaved enzymatically during degradation. This also results in a decrease of compressive modulus. The time to degradation (t) is dependent on the chemical composition of the hydrogel and physical properties, such as the incorporation of pores. In most hydrogels, the compressive modulus remains constant during the time between polymerization and degradation. The figure illustrates an example of some sample rates – actual rates depend on a myriad of factors, including chemical and physical properties, incorporations, and external environment.

The compressive modulus is a measure of mechanical strength or

stiffness of a hydrogel and can be most easily varied by changing the percent

composition of monomers to directly affect cross-link density [39, 57].

Alternatively, cross-link density, and thus compressive modulus, can be modified by changing the molecular weight of the monomer or varying the total amount of cross-linker [39, 86, 95, 130, 145]. In general, a higher cross-linking density produces a more rigid hydrogel, such as one that might be suitable for re-

engineering bone, while fewer cross-links make for softer hydrogels that are ideal for brain and other soft tissues. When using hydrogels for the engineering of specific tissues, variations in compressive modulus can be critical to how well the

hydrogel functions. For instance, using primary neuronal cell cultures, Lampe et

al. [141], found that the cells survived better in softer hydrogels, with a

compressive modulus less than or equal to 3.8kPa, compared to cells in

hydrogels with a stiffer compressive modulus (>19kPa). The compressive

modulus for brain tissue observed by Lampe et al. and others [47, 141], ranges

from 2.6-5.7kPa. In contrast, a study by Chatterjee, et al. [68] using osteoblasts,

demonstrated that osteogenesis was improved when the cells were grown in

hydrogels with a higher compressive modulus, similar to that of mineralizing

bone. Tissues such as bone can have a compressive modulus from 100-300kPa,

and as high as 1.5x107kPa [146, 147].

These studies suggest there is no single mechanical strength which works

best for all cell types and that the stiffness of the culture environment needs to be

tuned to the cell type for optimal performance (Figure 3). More specifically, with

Figure 3. Survival and fate of peripheral cell types as a function of biomaterial stiffness. This is a general summary of results obtained across a variety of biomaterials, therefore it is important to remember that biomaterial composition can also have an effect on cell survival and proliferation. A. Chondrocytes appear to grow on a broad range of stiffnesses; however, the differential expression of collagens and ECM components (ie. aggrecan, glycosaminoglycans) could be a function of the stiffness of the biomaterial [69, 148-151]. B. Based on available information, fibroblasts may need a less stiff biomaterial for survival and proliferation, compared to studies that have used chondrocytes [41, 152, 153]. Based on two in vivo studies, encapsulated fibroblasts inserted subcutaneously survive and produce ECM components in biomaterials with both softer and stiffer characteristics [152, 153]. C. Mesenchymal stem cells (MSC) also appear to survive and proliferate better on softer materials; however, changes in stiffness alter the MSC differentiation. In softer materials, MSC begin to express neuronal markers. As the stiffness increases, MSC may begin to express adipogenic or myogenic markers in addition to increasing their expression of factors important to angiogenesis (i.e. VEGF). At the higher range of stiffness, MSC express osteogenic markers [143, 154-159].

regards to neural cell types, neurons prefer to grow on substrates with lower compressive moduli (0.1-1.0kPa), compared to relatively stiffer surfaces (0.5-

10kPa) preferred by astrocytes and oligodendrocytes; meanwhile the compressive properties ideal for neurite branching and extension are even more limited (Figure 4) [47, 143, 144, 160-164]. The impact of these studies indicates that finely controlled specificity of the mechanical properties of the hydrogel environment is especially important in directing neural cell lineage, proliferation, and growth. To add a final consideration, the compressive modulus of a hydrogel decreases as a function of degradation and the loss of cross-links, resulting in a loss of mechanical integrity (Figure 2) [57, 125, 165]. This can be beneficial if the hydrogel is designed to degrade on the same time scale during which the encapsulated cells produce their own ECM to replace the tissue, theoretically allowing the cells to create their own mechanically robust ECM as the hydrogel degrades.

Additional mechanical measures include elasticity and stretch, measured by the tensile, or Young’s, modulus, and the ability to withstand straining or shearing forces, measured by the shear modulus. The shear and Young’s moduli, along with the compressive modulus, are measures of the elastic properties of the material as defined by Hooke’s law. Mimicking these mechanical properties in the creation of a hydrogel allows tissue engineers to imitate peripheral tissues, such as skin or blood vessels, which are frequently exposed to these types of mechanical stresses [76, 166, 167]. In a study which

Figure 4. Survival and fate of neural stem cells as a function of biomaterial stiffness. The fate of neural stem cell (NSC) populations grown on (2- dimensional) or in (3-dimensional) a polymer change as result of varying mechanical properties. Neural cells do not survive well in a biomaterial that is very soft (less stiff) or on a very stiff material. However, those which do survive at the lower stiffness tend towards a neuronal cell fate, whereas astrocytes develop more predominantly at higher stiffness. Neuritic extension is also best observed when the stiffness is lower, but NSC migration is more optimal at slightly higher stiffness [47, 107, 128, 143, 144, 161, 163, 168]. A. Post-natal day 6 rat neural stem cells (NSC) developed into >50% neurons, ~25% astrocytes, ~15% oligodendrocytes, and ~10% remained undifferentiated (nestin +) at this approximate level of stiffness [162]. B-C. Sensory spinal neurons had long neurite extensions but branching was favored on the less stiff material (B) compared to the more rigid material (C). Astrocyte survival was poor on both types of materials [144]. D-F. Embryonic day 13.5 midbrain-derived NSC, at 24 hours post-encapsulation, had ~54% survival with spheres of mixed populations of neuronal and astrocytic cells (D), ~47% survival (E), and ~31% survival (F), with segregated spheres of neuronal and astrocytic cells. At 21 days, no NSC were surviving in the stiffer hydrogel composition (F) [47].

exemplifies the unique tunable characteristics of hydrogels, HA and PEG-based hydrogels designed by Young and Engler [66] were designed to become stiffer

(an increase in elastic modulus) over time in a manner similar to the temporal change in stiffness observed in developing heart muscle. The change in elastic modulus from ~0.5kPa, similar to mesoderm tissues, to~10kPa, similar to mature heart muscle, induced immature cardiac cells to differentiate into mature cardiomyocytes [66]. In adult brain tissue, which is not routinely exposed to 24

stretching and shearing stresses except under extreme conditions, the

compressive modulus may be the more critical mechanical characteristic to

mimic in the hydrogel. Compressive changes in the brain as a whole can occur

with even mild brain trauma, neuroinflammation, and even hypertension [169,

170]. Highly localized compressive changes can also occur during neural cell

migration, which requires remodeling of the ECM [171].

Mesh size describes a nanoscopic physical characteristic which arises from the chemical features of the hydrogel described previously and contributes to mechanical properties, such as hydrogel stiffness. It is defined as the distance between the cross-link points in a hydrogel and is typically measured in angstroms (Å), with sizes sometimes ranging from 10 to 150Å [69, 130, 141,

149]. Mesh size is especially important to consider when encapsulating cells into a polymer matrix because the cells must be able to exchange nutrients and wastes with the external tissue across the polymer border. Additionally, diffusion rates depend on the size of the molecule and the mesh size of the hydrogel – the

smaller the mesh size, the slower diffusion occurs, and vice-versa. The ability for

fluids and small molecules to move in and out of a hydrogel depends initially on

diffusion, while degradation plays a role in the later movement of larger

molecules and cells [172, 173]. Water, a molecule of about 2Å, easily diffuses in and out of hydrogels thus increasing the rate of hydrolysis and contributing to the bulk erosion of the hydrogel [86, 138]. However, the diffusion of larger molecules, such as proteins, specifically neurotrophic factors, may be limited if a hydrogel is

not designed with a suitable mesh size for their free diffusion to cells

encapsulated within a hydrogel.

The macroscopic architecture of a hydrogel, such as the incorporation of

interconnected pores and the overall shape and size, can determine its function

within a tissue.The incorporation of pores in a hydrogel can be an important

factor to consider, as some cells and tissues rely on larger spaces within the

hydrogel to function properly. While the terms mesh size and pore size (or

porosity) are sometimes used interchangeably, pores are defined by their larger

size, typically in the micron (µm) range in diameter (Figure 5) [75, 149, 174-176].

Pores can occur naturally depending on hydrogel chemistry (such as molecular weight of monomer precursors) [149], or can be created intentionally.

The creation of pores can be achieved by constructing a complex hydrogel with regions of polymer with faster degradation kinetics than the rest of the hydrogel, resulting in interconnected pockets within the hydrogel [174, 175], while other processes include much more complex methods of fabrication [75, 135, 176-

178].

The purposeful incorporation of pores into the hydrogel architecture can serve to drive cell-specific growth and differentiation [176, 179]. For example, neurite extension can be guided by constructing hydrogels with interconnected pores that allow for growth before the hydrogel degrades [27, 175]. Namba, et al.

[175], demonstrated an increased number of neurites and neuritic branching from primary neurons encapsulated in PEG scaffolds with approximately 1.6 µm diameter pores compared with non-porous PEG scaffolds. This type of polymer

Figure 5. Hydrogel pore size changes as a function of precursor molecular weight. Lin, et al. [149] has demonstrated variation in hydrogel pore size based on the molecular weight of the polymer constituents. Using polyethylene glycol diacrylate (PEGDA) monomer precursors with molecular weights of 3.4kDa (A, box is magnified in a), 6kDa (B, box is magnified in b), 10kDa (C), and 20kDa (D), the authors produced hydrogels with pores (arrows) ranging in size from 9- 13µm for hydrogels made from 3.4kDa PEGDA (A), to 38-42µm for 20kDa PEGDA hydrogels (D). The authors observed a similar increase in the nanoscopic property of mesh size depending on molecular weight (45.1Å for 3.4kDa hydrogels and up to 130.9Å for 20kDa hydrogels). These scanning electron microscopy images are of freeze-dried PEGDA hydrogels. It would be unwise to assume that the physical characteristics of the dried hydrogel resemble that of a fully hydrated hydrogel. A fully hydrated hydrogel would be swollen with water and have an amorphous and fibrillary internal meshwork. It is possible that the pore sizes, once hydrated, would be considerable smaller. Scale bar represents 50µm. The above image has been modified from the original by inserting magnification boxes (a, b) to better appreciate the smaller pore sizes. Reproduced with kind permission from Springer Science+Business Media: Pharmaceutical Research, Influence of Physical Properties of Biomaterials on Cellular Behavior, vol. 28, 2011, p.1426, Lin, S., et al.

architecture may contribute to developing hydrogels for the purpose of neural circuit reconstruction.

Similarly, many tissue engineering hydrogel applications, such as producing skeletal or cardiovascular tissues, require the in-growth of host blood vessels to achieve incorporation of the cell-loaded hydrogel into the tissue and to provide oxygen and nutrients to encapsulated cells [64, 177, 180]. This may also become necessary in the brain if a large volume of hydrogel is introduced – the existence of pores may allow for vascularization of the implanted hydrogel. To use vascularization of tumors as an example, an implant volume of greater than

1 to 2 mm3 would require new vascularization for encapsulated cell survival [181].

Superporous hydrogels, like those developed by Keskar, et al. [176, 177], demonstrate this permeability by illustrating the in-growth of vascular structures and blood cells within the interconnected pores of the hydrogel. While the presence of pores will likely contribute to the function and success of a hydrogel, there is little evidence that pores contribute directly to biocompatibility of encapsulated cells in small hydrogel scaffolds [179].

The overall shape and size of a hydrogel should also be considered. The majority of hydrogels can be formed in any shape desired. For example, a hydrogel could be used to fill the amorphous shape of a lesion site, such as a stroke cavity in the brain [27, 97, 182], by implanting before polymerization or by using a hydrogel with very low compressive modulus (and therefore less “solid”).

In contrast, a hydrogel formed as a strand could be used to link two remote

regions to reconstruct a neural circuit, such as the nigrostriatal pathway in

Parkinson’s disease [58]. Similarly, the hydrogel strand could be used as a

bridge across a glial scar to help repair a spinal cord injury [43]. Size is also

important in the brain because of the limited open space within the tissue and the

confines presented by the skull. For example, a large volume hydrogel could

negatively impact the brain by increasing pressure in the tissue surrounding the

implanted hydrogel. Swelling of a hydrogel is an additional consideration in vitro

because unconstrained hydrogel degradation results in an increase in the overall

size of a hydrogel. Hydrogel studies often describe swell ratios to give an idea of

the water and fluid uptake of a hydrogel during degradation, as swell ratio can be

a contributing factor to defining degradation rate. Swelling in the brain, however,

is less of a concern because the counter-forces of the surrounding tissue and

relatively slow degradation confines the increase in overall size due to swelling

[58]. Size and shape can also impact the biocompatibility of a hydrogel, because

increased surface area can contribute to a greater tissue-material interface which

increases accessibility to the hydrogel for inflammatory and immune cells [183,

184].

Surface texture, or topography, of a hydrogel material can also affect

neural cell morphology and migration [112, 185-187]. Particularly, neural stem cells undergo migration, development, and polarization patterns based on contact guidance cues, which can be mimicked though complex structuring of the polymer surface [112, 186-190]. A grooved pattern can be used to guide neurite extension; however, optimal neuritic growth is dependent on the depth and width

of the groove [187]. Altering the texture of the surface also alters the total

exposed area, which is likely to lead to an increased immune response.

In designing a hydrogel to enhance or replace neural tissue, hydrogel

engineers must keep in mind the desired site of implant and the function and

specific anatomy of the surrounding tissue. These features will likely constrain

the design choices for the hydrogel. For example, the spinal cord

neuroarchitecture is highly organized with parallel tracts of neurites and axons

and thus may benefit from a hydrogel with organized architecture, such as

nanofibers or nanotubes [46, 108-110, 112, 191]. Comparatively, a cerebral stroke cavity may require a hydrogel with well-defined properties of polymerization so that a hydrogel can be implanted prior to polymerizing to fill the cavity [27, 36, 38, 40, 94, 192]. Implantation of hydrogels into white matter, such as in the spinal cord or corpus callosum, will limit access to free water and thus slow the hydrolytic degradation [57, 139, 193, 194].

Drug Delivery

One way to increase the effectiveness of a hydrogel is to incorporate

condition-specific drugs or molecules which can function to sustain encapsulated cells or provide support for the tissue surrounding an implanted hydrogel. These

can include growth factors and differentiating factors to specify the fate of

encapsulated stem cells, or anti-inflammatory agents to suppress the immune

system of the host. One well known example of drug delivery via polymers to the

CNS is the release of the chemotherapeutic agent, carmustine (BCNU), from

implantable polymer wafers to combat brain tumors (Gliadel™ system) [195,

196]. Major advantages of encapsulating carmustine include stabilizing the drug,

which normally has a short half-life, and providing precise control over the localization of drug release, limiting collateral damage to healthy brain tissue and reducing side-effects [16, 195, 196]. A number of studies have demonstrated the growth and differentiation of neural cells by incorporating neurotrophic factors into polymers and hydrogels. For example, ciliary neurotrophic factor (CNTF) has been incorporated to promote neural cell proliferation, differentiation, and neurite outgrowth [19, 197]. Similar results have been seen with neurotrophin-3 (NT3), platelet-derived growth factor (PDGF), glial-derived neurotrophic factor (GDNF), and nerve growth factor (NGF) [198, 199]. Lee, et al. [200] used a 3D lithographic printing technique to incorporate layers of hydrogel containing neural stem cells on top of hydrogel layers containing vascular endothelial growth factor (VEGF) to demonstrate neural stem cell migration towards the growth factor. Additionally, monomers and peptides, such as a reactive oxygen species-binding polymerizable superoxide dismutase (SOD) mimetic metalloporphyrin macromer

(MnTPPyP-Acryl; [201]) or a peptide antagonist to tumor necrosis factor-α

(TNFα), can be added to the surface of the hydrogel to impede the host inflammatory system from targeting encapsulated cells [201, 202].

Though there are still many variables to consider, drug delivery mediated by hydrogel is perhaps the most common hydrogel application. Additionally, some of the first clinical uses of hydrogels were as drug delivery-tools. As suggested, the incorporation of growth factors and trophic molecules into a

hydrogel system allows researchers targeted site application along with temporal

control over release. Gelatin-based hydrogels were used to deliver dopamine to

the striatal region of Parkinsonian rats [203], while the immense variation afforded with synthetic PEG hydrogels have allowed the tailored release of neurotrophic factors over a matter of weeks to months [19]. Many proteins and molecules that are difficult to deliver due to stability or kinetics, or compounds that are toxic systemically, have been the focus of research into hydrogel-based drug delivery.

Release of therapeutics from a hydrogel is dependent not only on hydrogel-defined factors, such as degradation and mesh size, but it is also dependent on the therapeutic molecule itself. How the therapeutic molecule is incorporated, for example tethering or encapsulation in the hydrogel or within microparticles, and the actual size of the molecule can define the release profile of the molecule from a hydrogel.

The size and chemical identity of incorporated molecules can affect the release kinetics from the hydrogel. Specifically, a small mesh size hydrogel may prevent larger molecules from readily diffusing from the hydrogel and thus, may require the hydrogel to degrade before they are released. For instance, small molecules, such as the drug diltiazem (used to block calcium channels) at ~5Å in diameter, could readily diffuse out of most hydrogels [204]. In contrast, glial- derived neurotrophic factor (GDNF), a rectangular molecule of about 30 x 36 x

80Å (~23kDa), must be incorporated into a hydrogel with a mesh size of 80Å or larger in order for it to easily escape the hydrogel by diffusion [205]. Larger

molecules and cells often require degradation of the hydrogel before they can

progress into the environment (the average neuron soma ranges from 4-100µm,

or 40,000-1,000,000Å, in diameter [206]). Because diffusion can occur more quickly than degradation, molecules incorporated into the hydrogel that are smaller can reach their target more quickly than larger molecules that must wait for the hydrogel to dissolve, resulting in two or more different release rates.

These differing release kinetics can be ideal for a hydrogel designed with both encapsulated trophic factors or immunosuppressors and cells, where the diffusible factors can go to work immediately to provide a host environment that is well-suited for, or more closely matched to, the encapsulated cells, which are released later as the hydrogel degrades. Lastly, the polarity of molecules can also affect how they disperse from a hydrogel. Jeong, et al. [192], demonstrated that hydrophilic drugs, such as ketoprofen, are released more readily from a hydrogel, where the release rate is determined by diffusion; compared to hydrophobic drugs, such as spironolactone, where release requires hydrogel degradation.

Tethering of molecules to the polymer itself can also greatly impact the release and distribution of molecules from a hydrogel [126]. In the case of tethered molecules, release kinetics are dependent on the degradation

(hydrolysis or enzymatic cleavage) of the bonds between the therapeutic molecule and hydrogel backbone. Tethering can be accomplished much in the same way that the hydrogel is formed – by introducing the molecule during the polymerization process, the same bonds that connect hydrogel monomers can

connect drugs and other proteins (i.e. ester bonds between thiol and ether

groups) [126]. Certain molecules can also be used to tether drugs to the hydrogel, such as heparin, which binds PEG backbones and has been shown to have a reversible affinity with a number of growth factors [130]. Tethering can also be achieved through enzyme-sensitive oligopeptide tethers, such as matrix metalloproteinase-sensitive tethers bound to vascular endothelial growth factor

(VEGF), which when released induces angiogenesis [207]. However, it is important to consider that tethered molecules must withstand the polymerization procedures and the tethers themselves should be biocompatible, as the activity of the molecule could be decreased if the bonding or tethering molecules block active sites or remain attached to the drug post-release [130].

Incorporating drugs first into a smaller polymer structures, or microparticles, which are then incorporated into a larger hydrogel structure, is another way to control drug release [19, 58, 174, 184, 208-210]. Microparticles can be used to carry trophic factors and a variety of small molecule drugs, proteins, and peptides, such as siRNAs. Interesting examples of the use of microparticles include encapsulating antigens for the development of systemic immunity [211-213]. In the central nervous system, microparticles have been used to deliver dopamine and norepinephrine into the striatum of rats to suppress the symptoms of Parkinson’s disease [214]. Microparticles (or microspheres) are often made of polymer materials and are subject to variation in polymerization and degradation chemistry and kinetics, particle size, and loading density. Like hydrogels and other degradable polymers, polymer microparticles can be

designed with varying rates of degradation. This can be advantageous for

applications in which molecule release from a microparticle is designed to occur

at a different time point than hydrogel degradation. Such design might be

warranted if a hydrogel were being used to implant cells but there was need for

extended release of supporting factors – the hydrogel would be degraded,

allowing the cells to incorporate into the surrounding tissue while still receiving

trophic support from factors incorporated into microparticles.

Microparticles can also be used when more than one therapeutic agent is

needed and each needs to be released in its own time and location. In a recent

study, hydrogel strands carrying two formulations of poly(lactic-co-glycolic acid)

(PLGA)-based microparticles were implanted into the rat brain [58]. One group of

PLGA-based microparticles were loaded with brain-derived neurotrophic factor

(BDNF) and designed to degrade slowly. The other formulation of PLGA- microparticles was loaded with glial cell-derived neurotrophic factor (GDNF) and designed to degrade more quickly. The fast releasing microparticles released all the GDNF within a 28 day window, whereas the slow releasing microparticles released BDNF consistently for at least 2 months. The study demonstrated that the rate of protein release can be controlled by altering the rate of degradation of the microparticles, without changing the properties of the overall hydrogel strand

[58]. This could be beneficial in treating Parkinson’s disease where BDNF release from slower degrading microparticles into the striatum could encourage neurite outgrowth [215, 216], while GDNF release from faster degrading

microparticles into the substantia nigra from the same hydrogel strand could

provide immediate cell support for grafted neurons [217, 218].

Biocompatibility

The term biocompatibility can be ambiguous and is often used in a variety

of contexts. Throughout this thesis biocompatibility is presented in two contexts: the histocompatibility of an implanted hydrogel with regards to the local and/or systemic response of the host and the cytocompatibility of a hydrogel with encapsulated cells, both in vitro and in vivo. Hydrogel biocompatibility in the

brain, an immuno-privileged site mostly independent of the peripheral immune

system, is especially focused on the neuro-inflammatory and neuro-immune

response of the brain as identified by the reactions of the resident glia.

CNS Environment and Immune Response of the Brain

The CNS is composed of two main neural cell types: neurons and glia.

The limited capacity for the brain and spinal cord to self-repair lies in that mature neurons are post-mitotic and thus, cannot give rise to new cells. Glial cells, however, do proliferate and can be an obstacle in neural tissue engineering.

There are two main types of glial cells: astrocytes and oligodendrocytes. Under normal conditions, astrocytes perform a wide variety of supportive functions in the CNS. Through their interaction with the vascular system, astrocytes form a restrictive barrier through which they exchange nutrients and wastes with the rest of the body [219, 220]. This astrocytic barrier, along with endothelial cells and

36 pericytes, forms the blood-brain barrier (BBB). Oligodendrocytes in the CNS produce the myelin sheath which wraps neuronal axons to moderate signal transduction [219, 221, 222]. Oligodendrocyte impairment is the basis of neurodegenerative disorders such as multiple sclerosis and the leukodystrophies

[223-226].

The undamaged BBB prevents blood and immune cells in the body from entering the CNS. Thus, the brain and spinal cord then, must attend to immunogenic stimuli using resident cells, rather than relying on the immune system of the body. Microglia cells are the resident neuroimmune cells in the

CNS. During fetal development, microglia differentiate from the same cell lineage as macrophages, the immune cells of the body. Both cell types have similar immune functions, but reside in two distinct areas [227-229].

In addition to the neurons and glia, a very small population of neural stem cells can also be found in the subventricular zone of the lateral ventricles and the subgranular zone of the dentate gyrus in adult brains [3, 171, 230-232]. These neural stem cells produce neurons and glia that replace cells of the olfactory bulb and the hippocampus where neurogenesis is necessary for continued function [3,

171, 230-232]. However, this production of mature neural cells is very limited and cannot compensate for the amount of damage typically seen in most CNS injuries and disorders, thus the need to improve neural cell replacement strategies [3, 171, 231].

Because the brain is mostly isolated from the periphery by the blood-brain- barrier, it has a similar but slightly different response to tissue damage and

foreign materials implanted than peripheral tissues. Brain tissue damage

produced during the implantation process can trigger a limited infiltration of

macrophages and foreign-body giant cells of the peripheral immune system from

damaged blood vessels, however, the bulk of the neuroimmune response is

carried by the resident microglia and astrocytes [172, 221, 222, 233-235]. The

acute neuroinflammatory response is initiated by microglia; this includes antigen presentation and the initiation of cytotoxic pathways occuring over the first few days following injury [172, 183, 221, 222]. It is during this time that some healthy cells will become collateral damage, overwhelmed by the cytotoxic forces of the microglia [10, 236, 237]. In the days to weeks that follow the acute response, the astrocyte population increases in an attempt to repair damage and to isolate any offending materials in contact with the tissue by building a glial scar [183, 220,

235]. This glial scar is similar to the fibroblast scar tissue that develops in peripheral tissues, however, in the brain and spinal cord, the glial scar becomes a barrier to neuritic and axonal extensions [183, 223, 235, 238]. In the days and weeks following tissue damage, cytokine activation in both microglia and astrocytes turns from cytotoxicity to neuroprotection, through the secretion of anti-inflammatory agents (i.e. TGF-β, TNF-α, and thrombospondin) and neurotrophic factors (BDNF, GDNF, nerve growth factor (NGF)) [183, 220, 222].

Extracellular matrix proteins secreted by cells at the wound site contribute to the development of the glial scar. In the spinal cord, the scar prevents axons from reconnecting across the site of damage, limiting the recovery of motor function. In the brain, the effects of glial scarring are less well understood.

Haberler et al., [239] analyzed brains from eight Parkinson’s disease patients

implanted with electrodes for deep brain stimulation. In each case, the brain

tissue surrounding the electrode did not appear to have any long term glial

activation but a glial scar encased the electrode leaving a well defined cavity in

the brain. Chondroitin-sulfated proteoglycans (CSPG), laminin, collagen, and

fibronectin can all be found in the glial scar. CSPGs are inhibitory to the

outgrowth of neuronal axons [240-242]. Of note is the presence of laminin,

collagen, and fibronectin in the glial scar. These three components are abundant in the ECM of bodily tissues however they are generally not found in the undamaged CNS tissue because of their overly fibrous nature. However, in vitro studies have shown that they promote axonal outgrowth from neurons plated on laminin, collagen, and/or fibronectin coated plates [77, 79, 240, 243-246]. The role of these fibrous components on axonal outgrowth, nerve repair, and glial scarring has yet to be firmly established. Studies by Davies, et. al. [247, 248] suggest that the type of astrocyte involved in forming the glial scar is the primary determinant of growth permissiveness. Some astrocytes produce fewer CSPGs when activated and tend to promote fibrous alignment within the scar tissue, which seems to be more conducive to axonal growth across the scar [248].

In addition to an overall glial scarring, the number and degree of reactivity of the surrounding astrocytes and microglia involved in the immune response can be considered an indicator of the level of immunorejection [183] and can therefore be used as a tool to determine in vivo biocompatibility [58, 172, 249].

For example, rat brains implanted with a more quickly degrading poly(ethylene

39 glycol) (PEG)-based hydrogel had fewer microglia but more astrocytes in the surrounding brain than brains implanted with a more slowly degrading hydrogel of the same composition [172]. However, most of the astrocytes and microglia had a reactive morphology surrounding the more quickly degrading hydrogel suggesting the neuroimmune response was still activated [172]. Yang, et. al.[89] also demonstrated the in vivo reactivity of astrocytes and microglia in response to a biocompatible hydrogel. Levels of astrocyte reactivity around the implanted hydrogel were shown to be comparable to those around a saline injection. As well, microglia were shown to be nearly absent around the implanted hydrogel after eight weeks, similar to the saline injection [89]. In traumatic brain injury, the microglial presence can remain elevated long after the initial injury [250-253].

Thus, to determine the biocompatibility of a biomaterial with brain or spinal cord tissue, the acute, as well as the long-term behavior of the glial cells should be determined for as long as the desired biomaterial will be in situ.

Because of the differences in the neuroimmune response in the brain compaired to other tissues, certain considerations should be attended to when designing or using a hydrogel that will be implanted into the brain. Interestingly, many studies of hydrogel biocompatibility with the CNS suggest that the immune reaction is in direct response to the mechanical trauma of implantation and that the material itself does not contribute to the immediate immune response (Figure

6) [172, 183]. As such, this characteristic is advantageous when using hydrogels to implant encapsulated cells into the brain because the hydrogels can be used to increase graft survival during the immediate, implantation-induced

Figure 6. PEG-PLA-based hydrogels implanted into the rat brain induced a long term neuroimmune response similar to that observed in sham brains penetrated with a needle. Two months after needle penetration (A-C) or hydrogel implant (D-F), a glial response remains. A, D. GFAP+ astrocytes appear to be more abundant in the tissue surrounding the needle penetration (circled in A) whereas there are fewer astrocytes in the tissue surrounding the hydrogel implant (D). There does appear to be an increase in astrocytes at the hydrogel-brain interface with some GFAP+ processes piercing the hydrogel (D, *). B, E. Microglial cells, identified by positive CD68 reactivity, remain in both the sham brain (B) and in the hydrogel implanted brain (E). While there is not an observable difference in the number of microglia in the surrounding tissues, there is a profusion of microglia that have infiltrated the hydrogel (*). C, F. MAP2, labeling identifying neurons, shows a paucity of MAP2 presence in the area surrounding the needle penetration (circled in C) whereas in the hydrogel brain, MAP2+ cells and neurites can be found closely positioned to the hydrogel (*). Circles in A-C indicate the center of the needle penetration. * in D-F, indicate the center of the hydrogel implant. Arrows (A-C and D-F) indicate corresponding blood vessels found in the adjacent tissue sections. Images were taken between the cerebral peduncle and the subthalamic nucleus. Scale bar indicates 200µm. Quantified data from this study can be found in detail in Bjugstad et al. [172].

neuroimmune response [27, 202, 254, 255]. The long term compatibility of the material with the host tissue contributes to the effectivness of the implantation while a subsequent immune reaction may depend on the degradation by- products, therapeutic agents, and/or cells the hydrogel delivers. Particularily,

hydrogel chemistry and mechanical properties significantly contribute to the

biocompatibilty and must be tuned appropriately for the tissue type being

engineered. Fortunately, most synthetic hydrogels have been developed to be

biologically inert and thus chemically biocompatible.

Contributions of Hydrogel Properties to Biocompatibility

Hydrogel chemisty is important in determining the biocompatibility of a

hydrogel. The high water content of hydrogels contributes to their biocompatibility

and is one major innate advantage of using these types of polymers as

biomaterials. The hydrogel by-products, resulting from the degradation process, are probably the most important chemical considerations. Degradation by-

products arise as the bonds within the hydrogel are cleaved: hydrolysis and

enzymatic cleavage results in the hydrogel breaking down into the various

individual monomer or oligomer units from which the hydrogel is comprised.

Some studies have demonstrated the adverse effects of toxic degradation by-

products, such as poly(methylidene malonate 2.1.2) and poly(propylene

fumarate), in various tissues [183, 256-258]. Recently, however, many groups

using hydrogels for tissue engineering have considered the biocompatibility of

their by-products when formulating the polymer constituents and, as such, most

hydrogels in use today as biomaterials are considered chemically non-toxic. The

use of poly(α-hydroxy acids), such as lactic and glycolic acids, as degradable

units are a good choice because they have been shown to be biocompatible and

can be metabolically recycled by the body [86, 104, 107]. Likewise, the use of

hyaluronic acid and poly(ethylene glycol) as polymer backbones can be

beneficial because hyaluronic acid is a natural component of the ECM [42, 50]

and poly(ethylene glycol), though synthetic, is biologically inert [86]. Most

synthetic hydrogels have minimal cell adhesion and low protein adsorption, which

limits their recognition by immune cells, increasing their biocompatibility [52].

The ability for a hydrogel to degrade is necessary in the brain, because glial scarring can occur around permanent implants [120-123]. Scarring inhibits the repair and reconnection of neural circuitry, which is especially apparent in spinal cord injury [259-261]. The ability to degrade is dependent on precursor chemistry. The molecular weight of the individual hydrogel components impacts the degradation rate: typically, higher molecular weight polymers produce denser hydrogels, thus slowing degradation. In a similar fashion, hydrogels with greater total percent polymer content degrade more slowly [41, 57, 125, 138, 139, 145,

154, 262]. Simply, the rate at which a hydrogel degrades can alter the neuroimmune response. In rat brain tissue, it was found that faster degrading hydrogels invoked a neuroimmune response similar in degree and rate of resolution as observed in brains given a needle penetration only [172]. By comparison, slowly degrading hydrogels invoked less of an acute response, but during the chronic phase, maintained a larger non-reactive glial population [172].

This effect is also demonstrated by Tysseling-Mattiace, et. al. [263] using self- assembling peptide amphiphile nanofibers with the laminin epitope IKVAV. The authors demonstrate the regeneration of spinal cord axonal fibers across an injury site following the degradation of the nanofibers. While nanofiber

degradation was necessary for axonal outgrowth, the authors suggest that the

nanofibers also may have primed the lesion site for this regeneration [263].

Some chemical additions can contribute to the cell-containing functionality of the hydrogel, such as enhancing cell adhesion or encapsulation. Cell adhesion can be controlled by supplementing the hydrogel chemistry with oligopeptide sequences and tethering monomers. The most commonly referenced additions are heparin or gelatin, which mimic many ECM proteoglycans, and the amino acid sequence Arg-Gly-Asp (RGD) [27, 152, 153, 264-269]. The addition of

heparin, gelatin, or RGD sequences can facilitate cell attachment to the hydrogel,

increasing the ability for neural cells to extend processes, migrate, and

differentiate [63, 160, 267, 270-272]. For example, the commonly used

fibronectin-derived RGD peptide sequence, when incorporated into the hydrogel, has been shown to improve cell viability and migration [134, 264, 265, 267, 273,

274]. Additionally, as RGD sequence density in the hydrogel increases, neural stem cell adherance and neurite outgrowth increases [266, 268]. Even the addition of electrically charged monomers, such as sodium methacrylate, have been shown to impact cell behavior (including differentiation) through the interaction of the cells with a charged hydrogel surface [270, 275]. While hydrogels can be formulated with additional proteins, such as collagen, to further mimic the ECM [276, 277], encapsulated cells themselves have been shown to secrete their own environmental molecules, such as fibronectin and collagen

[128, 149, 278]. Chondrocytes especially, have been shown to function well

within hydrogels, with evidence that extended in vitro culture in PEG-based

hydrogels can lead to formation of complete cartilage-like tissues [149, 279].

Growth or trophic factors can be added to a hydrogel to further encourage

proliferation, migration, neurite extension, and differentiation [19, 35, 136, 197-

200, 280-282]. Other chemical groups may be added to counter-act a property which negatively impacts biocompatibility. For example, the addition of lactic acid to a photopolymerizing PEG hydrogel may help to neutralize free-radicals and improve neural cell survival [104, 107]. The additions can be permanently incorporated by attachment to the hydrogel polymer backbone, or freely incorporated into the hydrogel and released by diffusion and/or degradation [40,

134, 267, 273].

It is important that all hydrogel components and additions are applicable to

CNS tissue. A common biomaterial based on peripheral ECM components,

collagen, laminin, and fibronectin, is Matrigel, derived from a murine sarcoma cell

line. It is successfully used for in vitro culture of neural cells and many other cell

types. Further, Matrigel has been used successfully when injected

subcutaneously for studies of angiogenesis and revascularization [283]. The

effects of Matrigel used as a biomaterial for brain tissue repair however, are still

unclear. In animal models of stroke/injury, both Lu, et. al. [284] and Jin, et. al.

[36] found that Matrigel reduced the volume of the stroke/injury lesion only when

it encapsulated cells. Matrigel without cells was no better at reducing the lesion

volume than cells alone or buffer-injected control brains. Unfortunately, neither of

these studies evaluated the glial response to the Matrigel. A study by Uemura, et.

45 al. [33], however, demonstrated increased apoptosis of neural precursor cells encapsulated in Matrigel compared with unencapsulated controls. In these studies, neural precursor cells encapsulated into growth factor-reduced Matrigel were implanted into the normal mouse striatum. The number of TUNEL-positive cells was significantly higher in encapsulated cells than in unencapsulated cells for at least 72 hours post implant [33]. The preservation of the implanted cells is a primary function for biomaterials in brain tissue engineering. In addition, photomicrographs revealed a dense core of peripheral leukocytes in the Matrigel with encapsulated cells [33]. As with other studies, the authors did not evaluate biocompatibility with regards to the glial reaction. These results continue to underscore the need to ensure that the hydrogel and its components are designed to promote success both to the individual cells and to the surrounding tissues.

In addition to the significant effects hydrogel chemistry can have on encapsulated cells and neural tissue, most cell types also perform better in hydrogel environments in which the mechanical properties (i.e. stiffness, elasticity, etc.) more closely resemble their host tissue. Indeed, mechanical environment can play a key role in cell survival and tissue formation [143].

Mechanical properties of biomaterials have been shown to play a greater role in the growth and development of CNS cells than previously realized. Neural cell processes, from attachment and migration to differentiation and maturation, have been shown to be dependent on culture substrate stiffness and elasticity (Figure

4) [40, 142-144, 168, 185]. Neural cells have been shown to grow best in softer

46 hydrogels where the mechanical properties more closely match those found in native neural tissue [40, 47, 143, 144, 160-164, 168, 185]. Furthermore, increasing the compressive modulus to 20kPa and higher and exceeding the modulus observed in brain tissue with a stiffness closer to that of tissue culture polystyrene, induces a preferential survival of astrocytes and an increase in gene expression for ECM proteins associated with glial scar formation [141].

Designing a hydrogel for the replacement of dopamine neurons for

Parkinson’s disease would require the less stiff mechanical properties necessary for neuronal survival and neurite extension. In comparison, a hydrogel designed for rebuilding a damaged spinal cord tract should be relatively stiffer to support astrocyte and oligodendrocyte survival [40, 47, 128, 143, 144, 161, 163, 168].

Such subtle variations in mechanical properties impact the functionality of a hydrogel to guide encapsulated cell fate and survival, while larger variations can alter the biocompatibility of the material to brain or spinal cord as a whole.

Matching of the hydrogel mechanical property to host tissue is important to minimize contact stresses and the phenomenon of stress shielding. These phenomena have rarely been studied in soft tissues, such as the brain and spinal cord [285]. Biran, et al. [121] demonstrated the effects of the incompatibility of chronically implanted silicon microelectrodes. A significant microglial response and loss of neuronal cells bodies in the region around the microelectrode illustrates a foreign body immune response in the brain. While the authors did not elucidate as to the exact cause of the neuroimmune response, it could easily be suggested that the difference in mechanical nature between the implant and

47 surrounding tissue is likely the cause [121]. To achieve biocompatibility and successful integration of a hydrogel into neural tissue, it must be designed with the mechanical properties of the brain in mind and the forces to which tissues of the CNS might be exposed. For example, the spinal cord must be flexible enough to move, bend, or twist within the vertebrae under normal movement.

However, under extreme conditions the spinal cord is susceptible to stretching, shearing, and compacting, as would be experienced during a car accident.

Decades of neural cell replacement strategies used for treating the neurodegenerative disorder Parkinson’s disease revealed that neuons survive better in the host when they are implanted as a piece of donor neural tissue, rather than as dissociated cell suspensions [286-289]. This demonstrates the impact of retaining a three-dimensional (3D) environment in which cell attachments are maintained. As well, donor neural tissue pieces preserve components of the brain ECM, which can provide chemical and physical supprt for the implanted cells. Thus, as in other tissue engineering strategies, the ability to mimic the 3D ECM environment becomes a key element to successful tissue engineering. Unfortunately, political and ethical complications of using whole neural tissue have limited its use, and as such, current tissue engineering strategies focus on using neural stem/progenitor cell lines as a source for replacement neural cells. The use of cell lines, grown as single cell suspensions, makes the need for an artificial ECM more important.

Throughout this thesis, the words “encapsulated” or “incorporated” have been used to describe neural cell populations that reside within a 3D hydrogel

composition, as compared to cells which are grown on the surface of a hydrogel

composition (two-dimensional; 2D). A study by Lampe et al., [141] showed that

when the mechanical and chemical properties of a hydrogel are all equal, neural

cells survive and thrive better when grown in a 3D environment as compared to

2D. Even when the compressive modulus exceeds that which is preferred by

neural stem cells, in the range of 15-20kPa, neural cells grown in 3D environments survive significantly better than those on a 2D environment with a compressive modulus closer that that of the brain (e.g. 1.4-3.8 kPa) [141].

Summary

While further development of these systems for neural cells and tissue is

still needed, hydrogels provide a promising avenue for tissue engineering and

cell transplantation, especially in the brain where the regenerative abilities can be

limited.

The methods of polymerization and degradation of these biomaterials

contribute to the general functionality of the hydrogel. These two processes are

directly affected by the chemical characteristics of the hydrogel, which also

govern the mechanical and physical characteristics, as well as the overall shape

of a hydrogel. The properties of a synthetic hydrogel can be easily modified to

mimic the ECM of most tissues of the body, including the brain. Lastly, all of

these characteristics will directly influence the capacity of the material to perform

the task it was designed for – whether it be drug delivery, cell encapsulation for

implantation, circuit reconstruction, or a combination of these therapies.

From cross-link density and mesh size, to degradation rate and erosion mechanism, the chemical and mechanical properties of the hydrogel determine the successful integration of neural cells into the host brain tissue and the release profiles of molecules or drugs. This can be beneficial for timed release of therapeutics – for example, the immediate release of fast acting compounds contributing to the attenuation of the inflammatory reaction or the more gradual release of trophic factors acting to support long-term cell growth and survival. By incorporating cells, drugs, and/or trophic factors, researchers can be one step closer to developing tissues ex vivo or in situ for brain repair. This allows for multiple levels of complexity to be achieved with a hydrogel, including the potential for directed cell differentiation and vascularization within the hydrogel and formation of composite neural tissues. Three-dimensional tissue engineering allows for the advanced study of biological and physiological processes in a way heretofore unexplored. As well, advanced hydrogel technologies are allowing for new methods of cell-based drug screening and localized drug delivery to human tissues. The first long-term results of hydrogel implementation are being realized and show great promise for the future of this technology – in the brain and beyond.

Acknowledgement

I would like to acknowledge the authors of Lin et al. [149] and Springer

Science and Business Media, for their kind permission to use the images found in Figure 5 of this Literature Review Chapter.

CHAPTER III

HYDROGEL FORMULATION DETERMINES CELL FATE OF FETAL AND

ADULT NEURAL PROGENITOR CELLS2

Introduction

The high water content and highly customizable nature of hydrogels

makes these materials well suited for tissue engineering, especially in brain and

spinal cord. Unfortunately, too many studies use hydrogels without reporting the

tunable properties. As a consequence, they may not recognize how these

properties can alter their final results. Hydrogel chemistry, including the

molecular weight (mw) of the components, the type of polymerizing

modifications, and the total polymer content, all contribute to the customization of

hydrogel behaviors (e.g. the physical properties, such as polymerization and

degradation, and the mechanical properties). These behaviors will determine the

success of a hydrogel used for tissue engineering. For example, two studies

treating spinal cord injury used chemically similar hydrogels but reported

substantially different results: Park, et al. [136] found significant repair after spinal

cord injury using a hyaluronic acid (HA; mw 170kDa) hydrogel with an identified

shear storage modulus (G’) of 0.3kPa, but an undefined final weight percent

(wt%) of HA and an unspecific HA modification to allow for polymerization. In

contrast, Horn, et al. [290] found no repair of the spinal cord using a thiol-

modified HA-based hydrogel at a 0.5 or 1.0wt%, but failed to report the molecular weight of the HA or the mechanical properties of the hydrogel. While the

2 All or part of this thesis is taken with permission from Aurand, et al. (2014) Stem Cell Research, 12(1): 11-23. 51 hydrogels used in these two studies are very similar in chemistry, neither study provides enough detail about the tunable properties to be independently replicated. Hydrogel chemistry and subsequent physical and mechanical properties all have unique contributions to successful tissue engineering, specifically with regard to the reaction of the host tissue to the hydrogel and how replacement cells respond to hydrogel encapsulation [40, 291]. A comprehensive exploration of hydrogels well suited for neural tissue engineering, composed of commercially available materials with defined tunable properties may help standardize the use of hydrogels for neural tissue repair.

Hydrogels comprised of HA and poly(ethylene glycol) (PEG) provide both the natural element (HA) for neural cell interaction and the synthetic element

(PEG) for customization and functionalization. HA is the main polymer backbone of the extracellular matrix (ECM) of the brain and is degraded by hyaluronidases produced by both neurons and glia in vivo [292, 293]. Biologically inert PEG provides additional control over hydrogel physical properties and helps to enhance functionality through prefabrication of more complex polymers, allowing for the attachment of cells or incorporation of growth factors or drugs [19, 40, 58,

66, 86, 130, 149, 202, 291]. My study employs only these two components, without further modifications, to assess baseline biocompatibility and explore how changes in hydrogel polymer ratio and subsequent physical and mechanical properties affect the fate of encapsulated neural progenitor cells (NPC).

Many studies have explored the use of neural cells and tissue for their use in treating neurological disorders [294-296]. Clinical trials have been undertaken

implanting neural stem cells and NPC to treat a number of neurological diseases,

including Parkinson’s, Batten disease, and cerebral palsy [297-301]. Because of the ethical controversies surrounding the destruction of a fetus, research has also begun to explore the potential use of NPC derived from adult brain [302,

303]. Currently published studies often treat NPC derived from fetal or adult brains as the same type of cells. Indeed, Pollard et al. [304] found NPC from adult and fetal sources express the same neural progenitor markers (e.g., nestin, sox2, blbp, olig2, etc.) and respond similarly to basic fibroblast growth factor

(bFGF) and epidermal growth factor (EGF). While the major molecular NPC qualities are common to both cell types, few studies have compared the fates of these two cell types in vitro and even fewer studies have addressed the fate of hydrogel-encapsulated NPC using cells from either source.

Hydrogel biocompatibility is improved when the mechanical properties of the hydrogel are matched to the host tissue [40, 47, 143, 160, 161, 164, 291,

305]. Since tissue-matching is an important quality of a successful hydrogel, I investigated the fate of both fetal- and adult-derived NPC in twenty-five different

HA and PEG hydrogels. Because fetal-derived NPC are extracted from a brain with an inherently weaker mechanical integrity than adult-derived NPC, where the brain is stiffer [306], I hypothesized that the survival and differentiation of

NPC would be optimal when encapsulated into hydrogels that more closely match the originating tissue. My study is unique because of this comparison between fetal tissue-derived NPC and adult tissue-derived NPC encapsulated in

a range of biologically relevant hydrogels with mechanical properties of both

juvenile and adult brain.

Methods

Hydrogel Formulations and Polymerization

Polymer hydrogels were formulated from thiol-modified carboxy- methylated hyaluronic acid (CMHAS; “HA”) and thiol-reactive poly(ethylene glycol) diacrylate (PEG). Both products were purchased from Glycosan

BioSystems, Inc (Cat.# GS222 and GS700). The HA had a molecular weight of

250±30kDa and a company-reported degree of methylation of approximately 75-

85%; the PEG had a molecular weight of 3.4 kDa and the company has reported a 95% or greater degree of acrylation. Lyophilized HA and PEG solids were reconstituted with degassed, deionized water (DG water; cat# GS241, Glycosan

BioSystems, Inc) to 2%(w/v HA) and 10%(w/v PEG) stock solutions. To achieve desired formulations in Table 2, additional DG water was added to dilute stock materials as needed. For degradation experiments and mechanical testing, trypan blue was added to the DG water to visualize the hydrogels (final trypan blue concentration <0.00002%). For in vitro experiments, DG water was replaced with Hanks’ Balanced Salt Solution (HBSS; Gibco) to improve cytocompatibility with encapsulated cells.

Hydrogels were made in 25 different formulations as based on the final polymer ratios of HA%(w/v) to PEG%(w/v) (Table 2). HA ranged from 0.2% to

1.0% in 0.2% increments and PEG ranged from 0.6% to 3.0% in 0.6%

increments. The range of polymer concentrations were chosen based on

company instructions for preparations, on achieving a range which encompassed

mechanical properties of brain tissue, and on practicality of use. Hydrogels with

greater than 1.0% HA and 3.0% PEG polymerized too quickly (< 30 seconds) for

practical use. Hydrogels with less than 0.2% HA retained a liquid status, making

these materials ineffective at encapsulating cells three-dimensionally or to

undergo mechanical testing with the described methods.

Table 2. Matrix of 25 hydrogels studied. HA (wt%)

0.2 0.4 0.6 0.8 1.0 3.0 U P K F A 2.4 V Q L G B PEG 1.8 W R M H C (wt%) 1.2 X S N I D 0.6 Y T O J E Final hydrogel formulations based on the HA-to-PEG ratio were formulations based on the HA-to-PEG ratio were designated by the letters A-Y. For example, the hydrogel identified as formulation “M” had a final polymer concentration of 0.6%(w/v) HA and 1.8%(w/v) PEG. HA: hyaluronic acid, PEG: poly(ethylene glycol), wt%: percent weight by volume of polymer in final hydrogel formulation.designated by the letters A-Y. For example, the hydrogel identified as formulation “M” had a final polymer concentration of 0.6%(w/v) HA and 1.8%(w/v) PEG. HA: hyaluronic acid, PEG: poly(ethylene glycol), wt%: percent weight by volume of polymer in final hydrogel formulation.

Hydrogels were formed by adding DG water and stock PEG in discrete

volumes to a vial of stock HA. Materials were then transferred via pipette to the

respective polymerization vessels. Formulations were made in HA dilution series

(i.e. 1.0% to 0.2%HA) with the PEG concentration kept constant by adding

additional stock PEG and DG water to the vial of HA. For example, formulation

“A” was made starting with stock HA and adding PEG and DG water to create a

1.0%HA and 3.0%PEG hydrogel solution. Material was removed to the polymerization vessel and additional DG water and PEG were added to the remaining HA to create formulation “F” (0.8%HA:3.0%PEG), and so on until the five formulations using 3.0% PEG were obtained. Volumes varied based on experiment.

Polymerization of the hydrogel materials occurs through covalent bond formation between the thiol modifications on the HA and the thiol-reactive portions of the diacrylated PEG. This reaction is independent of low pH or temperature steps and does not require ultraviolet exposure. The polymerization rate (or gelation time) was determined per manufacturer’s instructions and based on published literature [95, 307-310]. This rate was recorded as the amount of time (in minutes) from the combination of materials to form a solid product as determined by inversion and indentation.

Hydrogel Degradation

Hydrogel degradation was measured using the samples from mechanical testing. Each hydrogel sample was quartered and each quarter placed in one well of a 12-well plate containing 1mL of ~100u/mL hyaluronidase ([41]; cat.#

H3506, Sigma-Aldrich) in HBSS. Hydrogels in enzyme solution were incubated in a 37°C humidified chamber. Half the degradation solution was replaced every other day, with care taken not to disturb the hydrogel. Hydrogels were observed twice daily for 72-hours and then once daily until complete degradation.

Complete degradation was determined as the point at which the hydrogel was no

longer visibly present to the naked eye and no solid product could be found in the solution. The time to complete degradation was measured in days.

Mechanical Testing

Compressive modulus was used as a measure of hydrogel stiffness. For each formulation, 400µL of hydrogel solution was polymerized at room temperature in a 10mm x 10mm x 4mm biopsy mold (Electron Microscopy

Sciences). Polymerized hydrogels were incubated at 37°C in a humidified chamber for 24 hours prior to testing. Hydrogels were brought to room temperature and extracted from the mold using a thin metal spatula. Hydrogel dimensions were measured to ensure uniform size and for modulus calculations.

Testing was done using an MTS Insight 5-SL uni-axial mechanical tester (MTS

Systems) fitted with a 5N load cell. Samples were compressed at a rate of

200µm/min to an end-strain of 40% of initial height. Compressive modulus (kPa) was calculated as the slope at 20% strain of an exponential fit to a true stress- true strain curve. The curve was constructed from data that were filtered with a sliding average filter, window size five, to reduce noise and truncated to contain values between 0-30% strain. Data are presented as the mean ± SEM compressive modulus in kPa of four to six measures (samples) per formulation.

For comparison, brain tissue samples were measured for compressive modulus in a similar manner. Whole brains were extracted immediately following

CO2 euthanasia from rat pups (postnatal day 7, n=3), adult rats (at least two

months of age, n=9), and adult mice (2-3 months, n=5) into ice cold HBSS. Adult

57 tissue was trimmed to match hydrogel dimensions, resulting in the removal of olfactory bulbs, cerebellum, and dorsal cortex, when necessary; sub-cortical structures were kept intact. Juvenile brains were measured whole. Tissues were brought to room temperature and tested using the same parameters as hydrogel samples.

Fetal NPC Derivation

Fetal NPC (fNPC) were derived from embryonic day 15 (e15) rats. Timed pregnant Sprague Dawley females (Harlan Laboratories) were terminally anesthetized and both uterine horns removed. Embryos from each horn were excised into ice cold Hanks’ Balanced Salt Solution (HBSS; Gibco) and decapitated. Mean crown-rump length was 13.74 ± 0.07mm. Whole brains, minus the olfactory bulbs and cerebella, were washed three times in ice cold HBSS.

Tissues were manually dissociated to single cell suspension by using first a

10mL pipette tip, followed by a 1mL pipette tip. The cell solution was divided into uncoated Petri dishes with approximately three brains per dish. Cultures were grown in standard serum-free medium (3:1 DMEM/F12 (Mediatech, Inc/HyClone

Laboratories, Inc), 1X B27 supplement (Life Technologies), 100U/mL penicillin

(Fisher Scientific), 1µg/mL streptomycin (Fisher Scientific), 2mM L-Glutamine

(Fisher Scientific), and 20ng/mL basic fibroblast growth factor (bFGF) and epidermal growth factor (EGF)) at 37°C and 5% CO2. One half of the culture medium was replaced every other day. Six days after plating (DIV6), cells were dissociated and frozen in standard medium with 10% DMSO (Fisher Scientific)

58 and stored in liquid nitrogen. For encapsulation into hydrogels, fNPC were thawed and plated in standard medium in Petri dishes. The fNPC were maintained in dishes for four days prior to encapsulation. To ensure the cells were NPC, a small sample (at least 2000 cells) was plated onto a gelatin coated slide, and immunolabeled for nestin (neural progenitor cell marker), beta-tubulin

III (βTubIII; neuronal marker), and glial fibrillary acidic protein (GFAP, glial marker). Prior to encapsulation, the cells were consistently >99% nestin-positive and only 1-5 cells were ever found to be positive for βTubIII or GFAP (Figure 7A-

7C).

Adult NPC Derivation

Adult NPC (aNPC) were generated from the hippocampi and subventricular zones (SVZ) of adult female Sprague Dawley rats. Animals were euthanized by CO2 asphyxiation and decapitated. Whole brains were removed and the hippocampus was dissected bilaterally into ice cold sterile HBSS. To obtain the SVZ, a coronal section, rostral to the hippocampus and containing the lateral ventricle, striatum, and septum was excised. Using a surgical blade, the striatal and septal tissues were trimmed to approximately 3mm from the lateral ventricle on all sides to issolate the ependymal and subependymal layers of the ventricle. All tissues were combined, cut into smaller pieces, and washed three times in sterile HBSS. Tissue pieces were manually dissociated to a cell solution

Figure 7. Immunolabeling of undifferentiated NPC prior to hydrogel encapsulation. NPC were immunolabeled for markers associated with neural progenitor cells (nestin) and neuronal and glial differentiation (βTubIII and GFAP). A-C. Fetal-derived NPC (fNPC) were almost entirely nestin+ (A), however, very sparsely, cells were βTubIII+ (B; arrows) or GFAP+ (C; arrows). D- E. Adult-derived NPC (aNPC) were also nestin+ with almost none of the cells labeling βTubIII+ (D) or GFAP+ (E). Nestin (red), βTubIII and GFAP (green), DAPI (blue). Scale bars are 100μm (A-C) and 50μm (D-E).

with a 10mL pipette tip, followed by a 1mL pipette tip. This cell solution was split

into uncoated Petri dishes (2-3 brains per dish) and cells were cultured in

standard medium as described for fNPC. Every other day, one half of the

medium was replaced from the top of the dish. Cells were cultured at 37°C and

5% CO2 for 25 days (DIV25) and aNPC were allowed to attach to the dish. At

DIV25 cells were collected, manually dissociated, and frozen in standard medium

with 10% DMSO and stored in liquid nitrogen. For encapsulation into hydrogels,

aNPC were thawed and plated into standard medium in Petri dishes. The aNPC

were maintained in dishes for four days prior to encapsulation. As with the fNPC,

a sample of aNPC was immunolabeled for nestin, βTubIII, and GFAP. Prior to

encapsulation, >99 % of cells were found to be nestin-positive with only 2-3 cells

positive for either βTubIII or GFAP (Figure 7D-7E).

NPC Encapsulation

NPC were encapsulated at DIV10 for fNPC and DIV29 for aNPC. For

aNPC, attached cells were dissociated from the petri dishes using a cell scraper;

fNPC were collected as floating neurospheres. All NPC were collected into a

15mL tube and washed in HBSS. NPC were then manually dissociated to single

cell solution by gently pipetting using a 200µL pipette tip. Cells were re-

suspended in HBSS at a concentration of 10,000 viable cells (vc)/µL. For NPC

encapsulation, hydrogels were fabricated in a manner similar to those used for

mechanical testing, except cell solution and additional HBSS were used in place

of DG H2O. Cell solution was combined with reconstituted HA prior to addition of

PEG and gently vortexed, to ensure even three-dimensional (3D) encapsulation.

Hydrogel series were made by adding additional cell solution, HBSS, and PEG to achieve the desired hydrogel formulation (Table 2) with a final concentration of

1000vc/µL for each hydrogel sample.

NPC 24-hour Survival

NPC survival was assessed 24 hours post-encapsulation. Fetal or adult

NPC were encapsulated into the hydrogels as described. A 96-well plate was

used to polymerize 50µL of hydrogel-cell material per well, with four to eight wells

per formulation for fNPC and three to eight wells per formulation for aNPC.

Hydrogels were allowed to polymerize fully for approximately 40 minutes before

100μL of standard medium was added to each well. As a control, wells with 50µL

of dissociated NPC alone (1000vc/µL) in standard medium were also cultured

(TCP controls; n=9). The approximate cell density, based on the surface area of

TCP wells, was 1,562 vc/mm2. Plates were incubated for 24 hours at 37°C and

5% CO2. The MultiTox-Fluor Multiplex Cytotoxicity Assay (Promega, cat.#G9200,

Madison, WI) was used to measure cell viability per manufacturer’s instructions.

Medium was removed from atop the hydrogel and hydrogels were incubated with

50µL assay reagent for 3 hours at 37°C. Controls wells and a cell curve (50µL

per well, fNPC or aNPC cultured the day of measurements) were incubated with

50µL of assay reagent for 30 minutes. Hydrogels were incubated longer to allow

reagent to fully permeate the hydrogel, but within the time frame suggested by

the manufacturer. Live cell fluorescence was measured using a Bioteck Synergy

HT multi-modal microplate reader at 400nm excitation and 508nm emission. Data from all reads were combined by formulation and total cell numbers were determined using an averaged cell curve. Any outliers, as determined by the statistical software, were excluded from the analysis; outliers were rare and excluded 3 of 148 data points for fNPC and 7 of 104 data points for aNPC. Total cell numbers, representing survival, were calculated in a volume of 50µL.

Cell Numbers and Differentiation at Three Weeks

To determine the long-term effects of hydrogel encapsulation on cell number and differentiation, a low density of NPC were cultured for three-weeks in serum-based media to drive differentiation. This lower density was chosen to limit the influence of cell-to-cell factors driving differentiation, enhancing the influence of the hydrogel. To do this, CultureWell™ multiwell chambered coverslips (Grace

BioLabs, item# 103380; Bend, OR) were coated with 15µg/mL poly-L-ornithine and incubated at 37°C overnight, at which time remaining poly-L-orthine was aspirated off. Coverslips were washed three-times with HBSS and air dried.

Twenty microliters of hydrogel solution containing NPC (1000vc/μL) was added to each to each well and allowed to polymerize at 37°C for approximately 40 minutes. As a control, 20μL of unencapsulated NPC solution was also plated directly onto the coverslip (TCP control). Chambered coverslips were placed in

100mm petri dishes (2 per dish) and dishes were filled with differentiation medium (1:1 DMEM/F12, 100U/mL penicillin, 1µg/mL streptomycin, 2mM L-

Glutamine, and 10% fetal bovine serum (FBS; Fisher Scientific, Fair Lawn, NJ)).

Plates were incubated at 37°C and 5% CO2. No additional growth factors were

added to direct NPC differentiation towards a specific cell fate. Half of the

differentiation medium was replaced every other day. After three weeks, cells

were fixed with 4% paraformaldehyde and processed for immunocytochemistry.

Each hydrogel-NPC formulation and unencapsulated TCP controls had 3-4 wells.

Based on the surface area of the TCP controls, the cell density was 707vc/mm2,

approximately half the cell density of the 24 hour cultures.

Immunocytochemistry and Microscopy

To identify NPC that either differentiated into astrocytes, neurons, or

remained undifferentiated, all chambered coverslips were processed for

immunocytochemistry (ICC). Standard ICC methods were followed except that

washes and incubation periods were extended to assure materials penetrated

the hydrogels. All washes were done in PBS three times for 10-20 minutes each.

Each well was incubated in protein block (5% goat serum, 1% bovine serum

albumin, and 0.3% Triton X-100 in PBS) for two hours. To label astrocytes, anti-

glial fibrilary acidic protein (GFAP; rabbit anti-GFAP; Zymed Laboratories,

Carlsbad, CA, Cat# 18-0063; 1:500) was used and wells were incubated

overnight at 4°C. The next day a Cy-3 goat anti-rabbit secondary antibody was

used (Jackson ImmunoResearch Laboratories, West Grove, PA, Cat# 11-165-

144; 1:400) and wells were incubated for four hours at room temperature in the dark. To label neurons, anti-beta tubulin III (rabbit anti-β-tubulin III), conjugated to

Alexa Fluor 488 (βTubIII; Millipore, Temecula, CA, Cat# AB15708A4; 1:400) was used and incubated overnight at 4°C. All cell nuclei were labeled with 4',6- diamidino-2-phenylindole (DAPI; Fisher Scientific,Fir Lawn, NJ) at 100ng/mL in

PBS for one hour at room temperature, followed by one wash overnight in PBS at

4°C. Following ICC, coverslips were removed from the petri dishes and placed on glass slides with anti-fade mounting medium (0.1M propyl gallate in 9:1 glycerol/PBS).

An Olympus BX61 microscope complete image analysis system with

CellSens suite image analysis software was used to create images of hydrogels

and TCP control wells. Z-stacks were made of hydrogels with a height greater

than seven microns. Four images (most of which were stacked images) were

taken per well.

To assess the number of cells still present at three weeks post-

encapsulation, the total number of DAPI+ cells was counted per image. The use

of DAPI at this time point allowed us to identify the total numbers of whole and

intact cells still incorporated into the hydrogel. The number of DAPI positive cells

was used to determine the average density of cells per mm3 for hydrogels or

mm2 for TCP controls. The average density was multiplied by the total volume

(with remaining hydrogel height given by the Z-stack) or surface area of each well to achieve a total number of cells for the well. Data are reported as the mean total number of cells by formulation.

To assess differentiation, cells in each image were counted as either

GFAP+ (glia/astrocytes), βTubIII+ (neurons), or as GFAP and βTubIII negative

(undifferentiated NPC), resulting in three outcome groups (“cell types”). Cell populations were determined as a density of cells per mm3 volume of hydrogel,

with hydrogel height given by the Z stack. Differentiation data are reported as a

percent of the total cells counted.

Data Analysis and Statistics

All data were analyzed using the Statistica software (Statistica 7.0,

Statsoft, Inc.). Two-way ANOVAs were used to compare levels of HA, levels of

PEG, and the interaction between HA and PEG, on the rates of polymerization,

degradation, mechanical properties, and NPC survival. NPC differentiation was

analyzed using a 3-way ANOVA with cell types as a repeated measure.

Individual cell counts for survival and differentiation were correlated with mean

compressive modulus for each formulation. Fetal NPC and adult NPC were

analyzed separately. When significant differences were found, a Fisher Least

Significant Difference post-hoc test was used to identify differences between

specific means. Significance was determined at p<0.05. Data were graphed as

color-coded contour graphs for interactions between HA and PEG. The levels of

HA were plotted on the x-axis, the levels of PEG on the y-axis, and the measured

data are represented in the contours of the z-axis. Contours were derived from the raw data using a distance-weighted least squares calculation.

Results

Polymerization

Polymerization rate was measured as the number of minutes required for

a hydrogel formulation to solidify, as indicated by the inversion method.

Polymerization rates ranged from 2 to 40 minutes. Hydrogels containing less

than 0.4% HA were excluded from the statistical analysis because they failed to

form a solid product. There was a significant effect of HA content (F(3,

50)=40.90, p<0.0001), as well as PEG content (F(4, 50)=15.43, p<0.0001), on

the polymerization rate. When either HA or PEG content in the hydrogels was

increased, the polymerization rate increased (occurred more quickly) (Figure 8A).

There was no significant interaction between the HA and PEG on polymerization

66 rates (F(12, 50)=1.58, p>0.05), suggesting they both contributed to polymerization, but their effects were not additive.

Figure 8. Hydrogel physical properties. Polymerization and degradation rates for hydrogels were determined in vitro. A: Polymerization rate was measured in minutes. Time to polymerization decreased (purple contours) as HA and PEG concentrations both increased. B: Degradation rate of hydrogels submerged in hyaluronidase solution was measured in days. Hydrogels with greater amounts of total polymer took longer to degrade (red contours); at higher PEG contents (1.8% to 3.0%), degradation was dependent on the HA content of the hydrogel. HA: hyaluronic acid, PEG: poly(ethylene glycol), wt%: percent weight by volume of polymer in final hydrogel formulation.

Degradation

Hydrogels were subjected to enzymatic degradation in a hyaluronidase solution. While not necessarily reflective of in vivo degradation or that initiated by cells in vitro, this was done to determine a relative timeline of degradation.

Degradation rates ranged from less than 12 hours (0.5 days) to 12 days.

Conversely to the polymerization rates, increasing the HA content significantly decreased degradation rate (F(4, 87)=370.39, p<0.0001), as did increasing the

PEG content (F(4, 87)=101.94, p<0.0001). There was also a significant interaction between HA and PEG content (F(16, 87)=40.50, p<0.0001) (Figure

8B). At low levels of HA (0.2% and 0.4%), none of the PEG contents tested 67

changed the degradation rate. However, as HA content increased, higher levels

of PEG (1.8%, 2.4%, and 3.0%) substantially slowed degradation. Therefore, the

degradation rates of hydrogels with HA content of 0.8% and 1.0% could be

manipulated by altering the PEG content.

Compressive Modulus

Compressive modulus was measured in kPa. Hydrogels had a mean

range from 0.12±0.06kPa to 31.30±1.92kPa. These measures overlapped the measures of fresh brain tissue (Table 3). In brain tissue, the compressive modulus varied based on species and age. In general, rats had a higher compressive modulus than mice of the same age. Furthermore, brains from older rats (≥2 months) also had a higher compressive modulus than brains from young rats (post-natal day 7).

Table 3. Brain tissue mechanical properties. Compressive Modulus Species* Age N (Mean ±SEM) Mouse 2-3 mo 3.48 ± 0.92 kPa 5 P7 2.02 ± 0.15 kPa 3 Rat 2-3 mo 8.50±1.22 kPa 3 4 mo 9.71 ± 2.50 kPa 4 *All tissues were harvested from female rodents and tested at room temperature.

For the hydrogels, compressive modulus was significantly influenced by

HA, with a modulating effect by PEG content (Figure 9). Overall, as HA content

increased, there was a corresponding increase in the compressive modulus (F(4,

86)=115.43, p<0.0001). Likewise, as PEG content was increased, there was an

increase in the compressive modulus (F(4, 86)=19.16, p<0.0001). Similar to

degradation, the combined effect of HA and PEG significantly altered

compressive modulus (F(16, 86)=5.60, p<0.0001). At low levels of HA, increasing

levels of PEG had minimal effect on compressive modulus (p>0.05). However, at

the higher levels of HA (0.6%, 0.8%, and 1.0%), increased PEG content

significantly enhanced the increase in compressive modulus (p<0.05). Thus, like

degradation, hydrogels with HA content of 0.8% and 1.0% were the most

amenable to manipulating the compressive modulus, simply by changing the

PEG content.

Fetal NPC Survival and Differentiation

Fetal NPC (fNPC) survival was measured 24 hours post-encapsulation.

Cells encapsulated in hydrogels, regardless of formulation, survived better than unencapsulated fNPC in TCP controls (mean survival for all 25 hydrogels =

61137.66±1346.63; mean TCP survival = 17311.75±1736.67). Comparisons

between the 25 hydrogels indicates survival of hydrogel-encapsulated fNPC was increased as HA content increased (F(4, 120)=78.97, p<0.0001), but decreased when PEG content was increased (F(4, 120)=281.71, p<0.0001). The combined effects of both HA and PEG created a significant interaction (F(16, 120)=47.54, p<0.0001). At the highest concentration of HA (1.0%), survival rates were fairly

3.0

2.4 30 25 1.8 20 kPa 15 10 PEG (wt%) PEG 5 1.2 0

0.6 0.2 0.4 0.6 0.8 1.0 HA (wt%) Figure 9. Hydrogel mechanical properties. Compressive modulus was measured for all 25 HA and PEG hydrogel formulations. Hydrogels with the lowest compressive modulus were those containing 0.2% HA and higher concentrations of PEG, as indicated by the purple contours. The highest compressive modulus was measured in the 1.0% HA and 3.0% PEG hydrogel, as indicated by the red colored contour.

consistent regardless of the amount of PEG. As HA content decreased, the

amount of PEG in the hydrogel had a more variable effect, with the lowest survival rates measured in those hydrogels with 2.4% and 3.0% PEG (Figure

10A).

A correlation between compressive modulus and survival was used to

address how the mechanical properties of a hydrogel affect biocompatibility. At

24 hours, the hydrogel compressive modulus appeared not to affect the survival

Figure 10. Presence of fNPC at 24 hours and three weeks. The total number of fNPC was determined at 24 hours (A) and at three weeks (B) post- encapsulation. A. Measures indicated hydrogels with lower PEG content have increased survival at 24 hours. Survival decreased as PEG content increased, mostly at lower concentrations of HA (purple and blue contours). B. Long-term cell numbers indicated that hydrogels with lower amounts of HA, but higher amounts of PEG had the greatest number of cells (red and yellow contours). Generally, hydrogels with lower concentrations of PEG had the lowest cell numbers at three weeks post-encapsulation, regardless of HA. Note that the plating density of the three week studies was half that of the 24 hour studies, thus direct comparisons cannot be made between these two time-points.

of fNPC (r=0.22, p>0.05, n=25), suggesting cell survival at this time-point is dependent on the hydrogel composition and not the relative stiffness.

The persistence of cells encapsulated in the hydrogel was also measured at three weeks post-encapsulation in conditions supporting NPC differentiation.

The long-term cell number and degree of differentiation for fNPC and aNPC in hydrogels containing less than 0.4% HA were excluded from the statistical analysis because they did not form a solid hydrogel. These hydrogels degraded too quickly, preventing accurate measurement of outcomes in the described culture conditions.

Results from remaining hydrogels suggest that during this three-week

period, the number of fNPC retained within the hydrogels was decreased

compared with TCP controls (mean hydrogel cell number = 3627.33±198.83;

mean TCP cell number = 24851.59±2718.65). The mean number of persisting

cells in the hydrogels was approximately 14% of that in TCP. Across the

different hydrogel compositions, the long-term cell retention was impacted by both HA and PEG (F(3,54)=3.66 and F(4,54)=22.04, respectively, p<0.05) and by the interaction between the two hydrogel components (F(12,54)=5.30, p<0.0001).

The number of fNPC still present at three weeks was highest when lower concentrations of HA were paired with higher concentrations of PEG, achieving cell numbers that were 20-25% of that measured in TCP (Figure 10B). This interaction between HA and PEG is in contrast to cell numbers measured at 24 hours, where low HA and high PEG had lower cell numbers. The long-term persistence of fNPC within the hydrogels did not correlate with the compressive modulus (r=-0.09, p>0.05, n=20).

The differentiation of fNPC into young neurons and/or astrocytes was assessed at three weeks. The percentage of neuronal cells (βTubIII+), glial cells

(GFAP+), or undifferentiated cells (DAPI+, βTubIII- and GFAP-) in each hydrogel was determined. After three weeks in differentiation medium, many cells differentiated within the hydrogel. Neuronal βTubIII+ cells were the smallest population at 2.80±0.54%, GFAP+ cells comprised an average of 21.14±1.31%, and undifferentiated cells comprised the remaining 76.06±1.32%. The TCP controls, while similar, had slightly fewer βTubIII+ and GFAP+ cells, at

1.34±0.10% and 19.38±1.08% respectively, while most cells (79.28±1.04%) were

not positive for either differentiation marker.

Hydrogel formulation had significant effects on fNPC differentiation

(F(24,108)=2.51, p=0.0007). Most cells which differentiated expressed GFAP

(21.14±1.31%) (Figure 11A). The percent of glial cells ranged from less-than

10% to almost 40% of the total cell populations. Increases in GFAP+ cell density was mostly measured in hydrogels with the mid-range levels of HA and lower levels of PEG, although a small peak of GFAP+ cells were seen in the hydrogels with 1.0% HA and 3.0% PEG (“A” in Table 2). Neuronal cells made up less than

6% of cells in most hydrogel formulations, with the exception of the 0.4% HA and

1.2% PEG hydrogel (“S” in Table 2; Figure 11B). In this hydrogel, βTubIII+ cells made up 14.46±1.98% of the population. This increase in neuronal differentiation in hydrogel “S” compared to overall neuronal means (14.46% vs. 2.80% respectively) appears to have occurred at the cost of cells differentiating into glial cells (7.93% vs. 21.14%) rather than a decrease in the overall number of cells which remained undifferentiated (77.61% vs. 76.06%; Figure 11C). For fNPC, population changes in glial cell density were related to changes in the hydrogel stiffness. GFAP+ cells increased with increasing compressive modulus (r=0.52, p<0.05, n=20). The differentiation into βTubIII+ cells was not influenced by this mechanical property (r=-0.25, p>0.05, n=20).

Figure 11. Fetal-NPC differentiation. Differentiation of fNPC was assessed at three weeks post-encapsulation. A. When differentiation occurred, most cells were positive for GFAP, particularly in hydrogels with 0.6% and 0.8% HA and 1.2% PEG (green contours). B. Neuronal differentiation, as indicated by βTubIII immunoreactivity, was low and found primarily at 0.4% HA and 1.2% PEG (blue contours). C. The majority of fNPC were not positive for either βTubIII or GFAP, indicating they probably remained undifferentiated. Undiff: undifferentiated cells (GFAP-/βTubIII-).

Adult NPC Survival and Differentiation

Overall, hydrogel encapsulation did not improve 24-hour survival of aNPC

compared with TCP controls (mean hydrogel survival = 93884.68±3031.04; mean

TCP survival = 130485.22±13027.96). Most hydrogels had fewer surviving aNPC

at 24 hours than TCP. Within the hydrogel groups alone, there were significant

effects of both HA (F(4,72)=9.44, p<0.0001) and PEG (F(4, 72)=41.79,

p<0.0001), as well as a combined effect of HA and PEG (F(16,72)=5.11,

p<0.0001). Generally, the greatest effects on survival were measured with

increased PEG content which significantly decreased survival (Figure 12A).

Increases in HA content also decreased survival, especially at higher PEG

concentrations (1.8-3.0% PEG). The most optimal acute survival conditions were measured in hydrogels with both low HA and PEG concentrations. However,

Figure 12. Presence of aNPC at 24 hours and three weeks. Total remaining aNPC numbers were determined at 24 hours (A) and at three weeks (B) post- encapsulation. A. At 24-hours, survival of aNPC was best at lower concentrations of HA and PEG, although there was an increase in survival at 0.2% HA and 2.4% PEG (red contours). B. Analysis of aNPC cell numbers after three weeks of hydrogel encapsulation indicated aNPC were present in greater numbers in hydrogels with higher polymer content. Adult-NPC cell numbers were optimal at 0.8% and 1.0% HA with 2.4% PEG (red contour). Note that the plating density of the three week studies was half that of the 24 hour studies, thus direct comparisons cannot be made between these two time-points.

there was an independent peak in survival observed in hydrogels with 0.2% HA and 2.4% PEG (“V” in Table 2). Like fNPC, the survival of aNPC at 24 hours was not correlated to hydrogel compressive modulus (r=-0.36, p>0.05, n=25). At three weeks, the number of aNPC remaining in some hydrogel formulations was better than that of the TCP controls while some were below control measures (mean hydrogel cell number = 5177.48±383.27; mean TCP cell number =

6104.01±809.90). Comparing the hydrogel formulations, significant interactive effects of both HA and PEG were found on the number of persisting cells at three weeks (F(12,59)=4.12, p=0.0001). At higher concentrations of PEG, long-term cell numbers increased, especially with accompanying increases in HA content.

The highest cell numbers were measured in hydrogels with 0.8% HA and 1.8%

and 2.4% PEG, and were nearly twice that measured in TCP (Figure 12B). In general, hydrogels with the lowest amounts of PEG (0.6% and 1.2%) had long- term aNPC cell numbers below TCP controls. This effect was seen regardless of

HA content. Unlike the long-term cell numbers of fNPC, the number of aNPC at three weeks was positively correlated with hydrogel mechanical properties.

Within the compressive modulus range I measured, as hydrogel compressive modulus increased, there was an increase in the number of aNPC (r=0.56, p<0.05, n=20).

After three weeks exposed to conditions which induce differentiation, the majority of encapsulated aNPC remained undifferentiated (80.02±2.12%),

17.87±1.87% expressed βTubIII, and only 2.11±0.40% expressed GFAP. By comparison, the TCP controls had a differentiation rate of 30.45±2.35% for undifferentiated cells, while 66.30±2.04% of cells expressed βTubIII, and

3.25±1.14% expressed GFAP. The level of overall differentiation in aNPC is consistent with that observed with encapsulated fNPC; however, the cell fates

(βTubIII vs. GFAP) are in sharp contrast to fNPC, where most differentiated cells

were GFAP+.

When considering the effects of the hydrogel composition, HA and PEG

had a significant combined effect on aNPC differentiation (F(24,118)=11.54,

p<0.0001). Overall, as HA concentration decreased, more cells were found to

express GFAP and βTubIII (Figure 13A and 13B); however, as both HA and PEG

increased, cells remained undifferentiated (negative for βTubIII and GFAP;

Figure 13C). The highest levels of overall differentiation were measured in

Figure 13. Adult-NPC differentiation. Differentiation of aNPC was assessed at three weeks post-encapsulation. A. Only a small population of aNPC differentiated to a glial lineage, as indicated by GFAP immunoreactivity (blue contours). B. The majority of differentiated cells adopted a neuronal fate, suggested by βTubIII immunoreactivity. Note the strong presence of green and blue contours, suggesting up to 40% of cells were βTubIII-positive. Increased differentiation was seen in hydrogels with 0.4% HA and 1.2% PEG, for both cell types assessed. C. Similar to fNPC, the majority of aNPC were neither βTubIII nor GFAP positive, suggesting they likely remained undifferentiated at three weeks.

hydrogels with 0.4% HA and 1.2% PEG (“S” in Table 2), with 50.37±3.16% of cells expressing βTubIII, 13.91±1.02% expressing GFAP, and only 35.72±3.50% appearing to remain in their progenitor state, a rate similar to that observed in

TCP controls. An elevated number of βTubIII cells (57.02±3.16%) were also found in hydrogels with 0.4% HA and 3.0% PEG (“P” in Table 2). In this particular formulation though, there was no increase in GFAP+ cells, suggesting a formulation which favors neuronal differentiation rather than general cell maturation.

In aNPC, the compressive modulus was correlated with βTubIII+ cell density (r=-0.68, p<0.05, n=20) but not GFAP+ cell density (r=-0.37, p>0.05,

77 n=20). The negative correlation between βTubIII expression and compressive modulus suggests aNPC tend to differentiate into neurons on hydrogels which are less stiff.

Discussion

My results indicate there are multiple variables that can affect the interactions between hydrogel and NPC, and the choice of which hydrogel to use is dependent on the function to which the hydrogel will be applied. Both HA and

PEG made contributions to polymerization and degradation rates, cell survival, and cell differentiation. Interestingly, HA was the main contributor to compressive modulus, whereas PEG provided slight modifying changes. Furthermore, the impacts of HA and PEG on cell fate was often determined by the age of the neural progenitor donor tissue.

Acute hydrogel biocompatibility, as measured by 24-hour survival, indicated fetal-derived NPC survived hydrogel encapsulation well. This may be a result of greater plasticity found in fetal-derived cells [171, 231, 311]. Greater plasticity could allow fNPC to equilibrate more quickly to an environment which more closely resembles developing neural tissue. Alternatively, fNPC may adapt more quickly to the hydrogels because this environment is similar to the neurosphere formations these cells adopted when cultured prior to encapsulation. Meanwhile, the aNPC prior to encapsulation attached as individual cells to the TCP surface and thus, initially, fewer cells may adapt to the

3D hydrogel environment. Adult-derived NPC survival was dependent on the

78 hydrogel formulation, and was generally lower at 24 hours than tissue culture controls. In general, however, both cell types appeared to show better survival at

24 hours in hydrogels with lower polymer content (Figures 10A and 12A).

At three weeks, cell numbers were reversed, with fNPC encapsulated in hydrogels markedly lower than in controls and retention of encapsulated aNPC in some hydrogels better than control cell numbers. The effects of hydrogel composition at three weeks were also reversed. Unlike the 24-hour survival, long-term persistence of encapsulated fNPC and aNPC suggests both cell types were retained well within hydrogels with higher polymer concentrations, especially the aNPC (Figures 10B and 12B).

The differences between 24-hour survival and three-week cell numbers could result from the initial polymerization process negatively affecting the aNPC and/or long-term habituation to the microenvironment affecting the fNPC. For example, studies using mouse embryonic fibroblasts encapsulated in HA and

PEG hydrogels found there was significant cell loss immediately after encapsulation [41, 312]. These researchers attribute the cell loss to the polymerization of polymer hydrogel components. This polymerization process could account for the low cell numbers in the aNPC-hydrogel complexes at 24 hours. The increased plasticity of the fNPC may have protected these cells from the polymerization process [171, 231, 311].

After the initial polymerization process ends, the qualities of the microenvironment likely determine the long-term presence of the cells. I found that increased stiffness of the hydrogel was associated with increased cell

79 numbers at three weeks. This may suggest that, once polymerized, hydrogels with greater polymer content provide a more favorable microenvironment, either by supporting long-term cell survival or supporting the retention of cells in the hydrogel. This microenvironment may be more favorable simply because it provides good structural support while still allowing for the diffusion of nutrients and wastes. The diffusion of solutes in a hydrogel, or in the brain extracellular matrix, is determined by the distance between the crosslinked polymers [313].

Smaller molecules pass more easily through the crosslinks, whereas larger molecules or cells can be entrapped. Over time, encapsulated cells can modify their microenvironment in vitro through the expression of native ECM proteins and hyaluronidases [154, 155, 314, 315]. This cell guided restructuring of the hydrogel environment could also permit pooling and binding of pro-survival growth factors in the medium and secreted from encapsulated cells [79]. This environmental restructuring may also provide the NPC with cues for proliferation and/or differentiation [79, 316-319].

One particular cue, contact mediated inhibition of cell growth, may account for the differences in cell numbers between the TCP and the hydrogels at three weeks when using fNPC. My data suggests that there was greater long-term persistence of the fNPC in the TCP compared to those encapsulated in the hydrogels. Since proliferation is generally guided by cell contact, the low cell density at which the TCP cells were grown could have induced a level of proliferation not found in the hydrogel grown cells. The cells encapsulated into

the hydrogel may have undergone contact mediated inhibition due to the 3D

nature of their environment.

Additionally, while fNPC in the TCP controls showed an increase in cell

number over the three-week culture period (mean cell number is approximately

124% of the starting cell number), fNPC in the hydrogels reached approximately

18% of the starting population. This could be due to cell death, indicating a lack

of support provided by the hydrogels, or a release of cells over time from the

degrading hydrogel. In the latter scenario, surviving cells may have been freed

from encapsulation as the hydrogel degraded and removed from culture during

media changes, resulting in the decrease in cell number. This process may also

be the case with the aNPC after three weeks, as suggest by the observance that

there were greater cell numbers in hydrogels which degrade more slowly. The

addition of modified gelatin, denatured collagen, or other attachment factors

might have prevented this cell loss. These additions are often used in studies in

which cells are grown on the surface of the HA and PEG hydrogels [152, 153,

320]. However, because collagen is not typically found in the brain ECM [77, 79,

243, 245], and because I wanted to address baseline biocompatibility of the most basic hydrogel, I did not include additional attachment factors. Future studies will inevitably decide to add factors to customize the hydrogel to improve cellular attachment and/or cell survival. The current study, has allowed a foundation from which that customization can occur with confidence.

Following the encapsulation of undifferentiated nestin-positive cells, I initiated the differentiation of fetal- and adult-derived NPC using serum-

supplemented medium. These conditions were free of additional trophic factors

that can guide cell fate, such as retinoic acid or cililary neurotrophic factor

(CNTF), which are used to induce neurons and astrocytes respectively [161,

321]. Thus, the differentiation of my cells was determined by fundamental properties of the basic hydrogel and cell-intrinsic mechanisms [322].

Fundamental attributes of the hydrogel, such as the mechanical properties, are known to drive cell fate [40, 47, 143, 160, 161, 168, 323]. For example, mechanical properties that best mimic the nature of brain tissue have been found to induce neuronal differentiation in mesenchymal stem cells and embryonic stem cells [143, 324]. In committed neurons and astrocytes, hydrogel stiffness can alter cell morphology, inducing neuritic branching from neurons grown on softer hydrogels and supporting the multi-processed morphology of astrocytes when grown on harder substrates [160, 325]. Most of my cells from both fNPC and aNPC remained undifferentiated (Figures 11C and 13C). The incomplete state of differentiation is consistent with previous results in both hydrogels and standard tissue culture conditions [47, 161, 164, 326-328].

Nonetheless, depending on hydrogel formulation, I observed a mixture of neuronal and glial differentiation in both fNPC and aNPC (Figures 11 and 13). I also observed a mixture of differentiated cell morphologies (Figure 14). Most of the βTubIII+ fetal-derived cells had a more mature neuronal phenotype with neuritic branching (Figure 14 A, B), in contrast with the βTubIII+ adult-derived cells in which the labeling was mostly compact and cytoplasmic (Figure 14 E, F).

Figure 14. Morphology of differentiated NPC three weeks post-hydrogel encapsulation. Fetal-derived NPC (fNPC; A-D): A-B. βTubIII+ fNPC typically had more mature morphologies, as indicated by long neurite extension (arrows). In many cases, these neurites extended into the surrounding hydrogel (A). C-D. GFAP+ glial cells had both mature (arrows indicating extended processes) and immature morphologies (arrowhead in D). Cells were sometimes observed in clusters (A, B, D), with extensions surrounding the cluster (B and D). This clustering was not necessarily hydrogel specific, as both A and C are the same hydrogel compositions (“S”), but there are clearly clustered cells (A) and individual cells (C). Adult-derived NPC (aNPC; E-G): E-F. aNPC typically had more immature morphologies with βTubIII+ cytoplasm (arrowheads) and little or no neuritic extensions (E). When neuritic extensions were present, they tended to be thin and short (arrow in F). G. Conversely, GFAP+ cells had a mature, branched morphology (red labeled cell; arrow indicates process extension). Note the interaction between the GFAP+ cells and the smaller βTubIII+ cells (green, arrow head) in this photomicrograph (G). A: Formulation “S”, scale bar: 25μm; B: Formulation “N”, scale bar: 10μm; C: Formulation “S”, scale bar: 25μm; D: Formulation “T”, scale bar: 25μm; E: Formulation “S”, scale bar: 50μm; F: Formulation “P”, scale bar: 25μm; G: Formulation “S”, scale bar: 50μm. DAPI (blue), βTubIII (green), GFAP (red).

GFAP+ cells from both sources showed more mature phenotypes, also indicated

by branching, especially the adult-derived cells (Figure 14 C, D, and F).

The three-week differentiation of fNPC revealed GFAP+ cells were the

dominant cell type in all hydrogels, but in hydrogels with a compressive modulus

that approached adult brain tissue measures, this glial differentiation substantially 83 increased (Figure 11A). There was also increased neuronal differentiation in a small subset of hydrogels with a low compressive modulus, closer to that observed for juvenile brain tissue (Table 3). My results confirm what Seidlits, et al. [47] also reported. While their data were not quantified, they reported an increase in neuronal differentiation in fNPC encapsulated in hydrogels that match the mechanical properties of fetal brain, and an increase in GFAP+ cells in those hydrogels representative of adult brain [47].

In contrast, my aNPC had a greater tendency to differentiate into neurons than the fNPC, specifically in hydrogels that had compressive moduli similar to both adult and juvenile brain tissue (Figure 13). The hydrogel results are consistent with previous findings using adult-derived NPC [161, 164]. The findings reported by Saha, et al. [161], are interesting because they compared not only mechanical properties of the biomaterials, but also the effects of differentiation media. Using media designed to induce neuronal differentiation, they found only 40% of their cells expressed beta-tubulin III on hydrogels with an elastic modulus similar to adult brain tissue. This was similar to the levels measured in their tissue culture plates. By comparison, using a standard serum- based medium, similar to that used here, Saha, et al. were able to increase the number of beta-tubulin III-expressing cells to 60% or greater, and reduce the expression of GFAP-positive cells to below 30% on hydrogels with mechanical properties similar to adult and fetal brain tissue. Their study suggests adult- derived NPC may be susceptible to both mechanical and chemical cues for neuronal differentiation. My study reveals the contrastive differentiation profiles of

the fetal-derived and adult-derived NPC, suggesting these two cell types are not

interchangeable for neural tissue repair applications. This further confirms the

impact of hydrogel mechanical properties on encapsulated cell fate and

emphasizes the need to select hydrogels for more than just their chemical

biocompatibility.

Interestingly, both NPC populations had the highest levels of neuronal

differentiation when grown in hydrogels with 0.4% HA and 1.2% PEG

(formulation “S”; Figures 11B and 13B). This differentiation may likely occur because this ratio of HA and PEG produces an ideal chemical, physical, and mechanical environment to support neural differentiation in both fNPC and aNPC. The “S” formulation may be an excellent “introductory” hydrogel for those interested in entering the field of neural tissue engineering. Alternatively, for studies desiring specific cell fates, such as the development of astrocytes for spinal cord repair [329], hydrogels “N” or “I” encapsulating fetal-derived NPC may provide the researcher with the desired astrocytic cell type. However, driving differentiation of complete cell populations will require more than simply changing the mechanical properties and picking the right cell source.

In conclusion, this study has established some of the basic properties and biocompatibility for twenty-five hydrogel compositions using various combined levels of HA and PEG in order to identify basic hydrogels suitable for neural tissue engineering. I further explored the contribution of these hydrogels to the fates of encapsulated NPC derived from fetal and adult brain. Now that the primary qualities have been defined for the basic HA and PEG hydrogel, the next

85 step will be to functionalize these hydrogels for a specific purpose. This can be done by either passively or actively incorporating factors into the hydrogel, through the addition of attachment factors or immune-modulators, or by producing specific geometries to guide cell morphology [40].

CHAPTER IV

IN VIVO BIOCOMPATIBILITY OF HYALURONIC ACID AND POLY(ETHYLENE

GLYCOL) HYDROGELS

Introduction

As we learn more about the causes and pathologies of neurological

disorders, we have begun to better understand the potential that reconstructing

neural tissue might have in the treatment and repair of these disorders.

Hydrogels, a combination of polymers with high water content, have become

useful bioengineering tools for cell delivery, structural tissue support, and/or the

local delivery of drugs or trophic factors [40]. Because of their soft, customizable

nature, hydrogels present an ideal way to combine tissue engineering techniques

with neural cells to recreate brain tissue. For example, a hydrogel combining

neural cells and growth factors can be injected into a stroke cavity to fill the

space left by tissue necrosis and provide a scaffold for tissue repair [27, 36, 38,

59, 330]. The addition of hydrogels could facilitate the repair outcome by

increasing implanted cell survival and integration. Many studies have shown that

the majority of cells implanted into the brain become victims of the initial

inflammatory reaction triggered by the implantation procedure and die within the

first weeks after implantation [10-14, 27, 28]. Hydrogel encapsulation of cells has been shown to protect cells from this immune response, resulting in higher rates of implanted cell survival [27, 33, 34, 36, 331].

Even with the potential of these materials, hydrogel biocompatibility must

be addressed. In the context of the brain, biocompatibility means that

87 encapsulated cells survive the implant process and that individual host cells adjacent to the hydrogel survive well. Furthermore, the host brain tissue should not initiate an immunorejection response. Therefore, hydrogel material must be cytocompatible and histocompatible [183, 291]. Numerous factors contribute to hydrogel biocompatibility, including the chemical, physical, and mechanical nature of the hydrogel material, and factors specific to the cells and tissue of interest [40, 291]. An example of these factors is represented by several studies using poly(methylidene malonate 2.1.2) (PMM 2.1.2) [256, 332]. PMM 2.1.2 was found to be biocompatible when it was introduced intravenously and was non- toxic to encapsulated fibroblast cells [332]. However, when it was used to make microspheres which were implanted into the brain, biocompatibility was not achieved [256]. Initially, the PMM 2.1.2 microspheres were well tolerated; however, once the microspheres began to degrade, the degradation products of

PMM 2.1.2 were found to be highly toxic to neural tissue [256]. These contradictory responses between peripheral exposure and brain tissue demonstrate the changing nature of hydrogel biocompatibility over time and illustrate the necessity for full characterization of the biocompatibility of the hydrogel in the tissue of interest (e.g. brain tissue).

One method of increasing the potential for brain biocompatibility is to develop a hydrogel material that mimics, as closely as possible, the extracellular matrix (ECM) of the brain. The ECM of the brain is different than most other tissues in the body. It is especially soft and composed on a backbone of hyaluronic acid (HA) rather than collagen or fibronectin, as seen in peripheral

tissues [77-80]. HA, and the ECM in general, play major roles in fetal brain

development, wound healing, and cell attachment [77, 333-338]. Recent studies have also suggested that HA and the ECM are involved in some disease neuropathologies, such as Alzheimer’s and traumatic brain injury [260, 339, 340].

Thus, HA-based hydrogels would better mimic the chemistry of the brain ECM than a collagen or fibronectin-based hydrogel.

In the brain ECM, HA is paired with various proteoglycans to form a hydrogel. For tissue engineering purposes, HA is often paired with poly(ethylene glycol) (PEG) to form a hydrogel with tunable properties, including the ability to manipulate the compressive modulus (a measure of stiffness). PEG is a synthetic, biologically inert, polymer that has been used in a variety of tissue engineering settings, including the brain and spinal cord [86, 172, 249]. In addition to matching the brain ECM chemically, hydrogels designed for neural tissue should be degradable and have a compressive modulus which approximates that of the brain [40, 47, 160, 161, 163, 164, 168, 185, 291]. By varying the polymer composition (i.e. the ratio of HA to PEG) hydrogels can be tailored to possess specific, definable, properties, such as compressive modulus

[39, 40, 44, 50, 57, 82, 84, 262].

The matching of hydrogel properties to the tissue, as a whole, is an

important factor in achieving brain biocompatibility; however, these properties

can also affect the fate of individual neural cells [40, 142-144, 162, 164, 185,

291]. For example, in vitro, relatively stiffer hydrogels have been shown to

support glial cell types, while neuronal cells survive better on softer hydrogels

[40, 47, 143, 144, 160-164]. Using neural progenitor cells (NPC) derived from

fetal or adult brains, I recently demonstrated a relationship between hydrogel

stiffness and the age of the donor cells on the fate of NPC. When comparing the

differentiation of hydrogel-encapsulated fetal-derived rat NPC to that of adult-

derived NPC, I found that when the hydrogel stiffness was increased, adult-

derived NPC had fewer neuronal phenotypes. By comparison, fetal-derived NPC, under the same conditions had an increase in glial cell types [341]. Thus, the

same hydrogels provoked different differentiation pathways in fetal-derived NPC compared to adult-derived NPC. This further illustrates the importance of carefully evaluating the hydrogel material and not assuming that a single material will produce the same results under slightly different conditions.

The present study was designed to assess the biocompatibility of

hyaluronic acid (HA) and poly(ethylene glycol) (PEG) hydrogels with

encapsulated fetal-derived NPC when implanted into the adult rat brain. By

encapsulating the NPC three-dimensionally into the hydrogel, my goal was to

improve NPC survival post-implantation and demonstrate the effects of hydrogel

encapsulation on the fate of implanted cells. Further, the neuroimmune response

of the brain to the hydrogel was examined. HA and PEG were chosen because of

their previously demonstrated biocompatibility as independent precursors [49, 86,

136, 172, 244, 249, 260, 340, 342]. I chose to use HA and PEG together without

additional attachment or trophic factors to determine the baseline biocompatibility

of this hydrogel with a discrete chemistry. I implanted hydrogel-cell complexes

into the intact striatum, a region of the brain often targeted for neural

transplantation in models of Parkinson’s disease and stroke [12, 214, 287, 343-

349]. My previous work established the biocompatibility of these materials with

NPC in vitro [341]; here I seek to establish the hydrogel biocompatibility in vivo.

Methods

Hydrogel and NPC Preparation

The hydrogels employed in this study were made using thiol-modified

carboxy-methylated hyaluronic acid (HA) and thiol-reactive poly(ethylene glycol)

diacrylate (PEG) (Glycosan BioSystems, Inc), reconstituted with degassed,

deionized water to form a 2% (w/v) stock HA solution and 10% (w/v) stock PEG

solution. The two hydrogel formulations used were based on formulations

amended from the Glycosan BioSystems (BioTime, Inc) standard protocol for the

production of Hystem hydrogels and earlier studies performed by Aurand, et al.

[341].

Two hydrogel preparations, denoted as “N” and “S” here and in my

previous study [341], were used based on my previous work. These particular

hydrogels were chosen because of their similarity during in vitro cell survival

studies (114% of initial plating for Hydrogel “N” and 111% of initial plating for

Hydrogel “S”; Table 4)[341]. Based on these in vitro studies, Hydrogel “N” without

cells had a compressive modulus (stiffness) of 9.86kPa, while the compressive

modulus of Hydrogel “S” without cells was 4.28kPa, both well within the range

measured for adult rat brain (8.5-9.7kPa) and a neonatal rat brain (2.0kPa)

(Table 4; [341]). When encapsulated NPC were differentiated in vitro, hydrogel

Table 4. Summary of selected results from in vitro studies (Chapter III). In Vitro In Vitro Compressive Survival HA PEG Differentiation Modulus (% of (wt%) (wt%) Neuronal Glial (kPa) initial (βTubIII+) (GFAP+) plating) Hydrogel 0.6 1.2 9.86* 114% 2.8% 37.5% “N” Hydrogel 0.4 1.2 4.28** 110% 14.5% 7.9% “S” Similar to brain compressive modulus of *4 month adult rat (9.71 ± 2.5 kPa) and **post-natal day 7 rat pup (2.02 ± 0.2 kPa).

“N” had a relatively high number of glial cells (37.5%) with low neuronal

differentiation (2.8%), whereas hydrogel “S” had highest rate of neuronal

differentiation of hydrogels tested (14.5%), but low glial differentiation (7.9%)

(Table 4) [341]. Thus, I chose two hydrogels with similar survival rates but different long term encapsulation effects.

Hydrogel “N” was formulated to have a final concentration of 0.6% HA and

1.2% PEG. The final concentrations for Hydrogel “S” were 0.4% HA and 1.2%

PEG (Table 4). NPC were encapsulated into both hydrogels at 4000 viable cells

(vc)/μl. Briefly, a stock HA solution was combined with a concentrated cell

solution in Hanks’ Balanced Salt Solution (HBSS; Gibco), followed by the

addition of stock PEG solution and mixed gently, but thoroughly, by vortexing at a

low speed. Hydrogel-cell complexes were allowed to fully polymerize, and were

kept on ice until implanted.

Fetal neural progenitor cells (NPC) were derived from the whole brain,

minus olfactory bulbs and cerebellum, of embryonic day 15 rats. Briefly, timed

pregnant female rats were terminally anesthetized and embryos from each 92

uterine horn were excised into cold HBSS. Brain tissues were manually

dissociated to a single cell solution that was divided into uncoated petri dishes.

NPC were grown as neurospheres in serum-free medium (3:1 DMEM/F12

(Mediatech, Inc/HyClone Laboratories, Inc), 1X B27 supplement (Life

Technologies), 100U/mL penicillin (Fisher Scientific), 1µg/mL streptomycin

(Fisher Scientific), 2mM L-Glutamine (Fisher Scientific), and 20ng/mL basic

fibroblast growth factor (bFGF) and epidermal growth factor (EGF)) at 37°C and

5% CO2 for six days; 50% of culture medium was replaced every other day. After

six days, cells were dissociated and frozen in medium containing 10% DMSO

(Fisher Scientific). NPC were stored in liquid nitrogen. Four days prior to

implantation, NPC were thawed and re-plated. Twenty-four hours after re-plating,

1μL of 10mΜ stock BrdU was added to the Petri dishes per every 1mL of standard medium. NPC were treated with BrdU for 48 hours. The BrdU medium was removed and NPC were given fresh standard medium for 24 hours prior to encapsulation.

For encapsulation/implantation, NPC were rinsed with HBSS, dissociated, and re-suspended as a concentrated cell solution in HBSS. This concentrated cell solution was further diluted in HBSS to 4000 vc/µL for Cell Alone Implants, or was added to hydrogel precursor components for encapsulation. All NPC treated groups received an implant total of 20,000 viable cells (5μL material at

4000vc/μL).

Implantation Surgery

Twenty-six intact, adult male rats (Sprague-Dawley; mean weight 271.2 ±

2.9g) received one of four implantation procedures: the Sham control group

received needle penetration and injection of 5μL of HBSS (n=6 animals); the

Cells Alone control group received 4000vc/μL in 5μL of HBSS (n=6); the

Hydrogel “N” group received 4000vc/μL in 5μL of 0.6% HA and 1.2% PEG

hydrogel (n=7); the Hydrogel “S” group received 4000vc/μL in 5μL of 0.4% HA

and 1.2% PEG hydrogel (n=7).

Rats were anesthetized with 80mg/kg ketamine and 10mg/kg xylazine and placed into a stereotaxic device. Materials were loaded into a 10μL Hamilton syringe fitted with an 18 gauge needle and were implanted unilaterally at a ventral projection into the following coordinates of the striatum: +0.96mm AP and

+3.0mm ML from Bregma, and -4.5mm DV from the brain surface. All animals were monitored post-operation until normal movement and behavior was regained. Prior to and post-operatively, all animals were housed in pairs and

given access to food and water ad libitum. All manipulations and housing were

approved by the Institutional Animal Care and Use Committee at the University of

Colorado – Anschutz Medical Campus in accordance with the National Institutes

of Health Guidelines for the Care and Use of Laboratory Animals. Every effort

was made to minimize pain and distress of the animals.

Tissue Preparation and Immunohistochemistry

At two months post-implantation, all animals were euthanized with

160mg/kg ketamine and 20mg/kg xylazine and transcardially perfused with

250mL of ice cold isotonic 0.9% saline followed by 250mL ice cold 4%

paraformaldehyde in phosphate buffer. Brains were removed and stored in fresh

4% paraformaldehyde at 4°C for one week and then transferred into 30%

sucrose until brains sunk and tissue could be sectioned. Tissues were cut

coronally at 30μm into free-floating sections and stored at -20°C.

Immunohistochemistry (IHC) was used to determine cell survival

(cytocompatibility) and differentiation and the host tissue immune response

(histocompatibility). Anti-BrdU antibody (1:50; BD Biosciences, cat. #347580) was used to identify implanted cells in sections taken every 180μm (every 6th

section). For double labeling, anti-MAP2 (1:500; Millipore, cat. #AB5622) and

anti-GFAP (1:500; Zymed, cat. #18-0063) antibodies were used in conjunction

with the anti-BrdU to determine differentiation of implanted cells in sections at

every 360μm (every 12th section). Anti-Nestin antibody (1:200; Millipore, cat.

#MAB353) was also used to determine differentiation of implanted cells in

sections at every 360μm. Anti-GFAP and anti-Iba1 (1: 500; Wako, cat. #019-

19741) antibodies were used to determine the endogenous immune response in

sections at every 360μm. For each label, a negative antibody control was

performed by omitting the primary antibody. As well, for BrdU labeling, tissue

from a Sham implanted animal (no cells) was used as a control for implanted

cells.

Briefly, tissues were washed in phosphate buffer (PB; 0.1Μ, pH 7.4)

followed by 1.5% endogenous peroxidase block, a 5% protein block, and biotin

and avidin blocking steps. For BrdU labeling, an antigen retrieval step was

necessary, entailing a 7min incubation in 2N HCl prior to the peroxidase blocking

step. Tissues were incubated in primary antibody overnight at 4°C. The following

morning, tissues were washed with PB and incubated in secondary antibody for

1.5-2hrs at room temperature. For BrdU-alone, Nestin, GFAP-alone, and Iba1

labeling, biotinylated secondary antibodies against the primary antibody host

animal were used (1:200; Vector, cat. #s BA-9200 (BrdU and Nestin labeling),

BA-1000). For BrdU/MAP2 and BrdU/GFAP double labeling, fluorescently tagged secondary antibodies were used. Brdu was labeled with DyLight 488 (green

fluorescence; 1:400; Vector, cat. #DI-2488) and MAP2 and GFAP were labeled

with DyLight 594 (red fluorescence; 1:400; Vector, cat. #DI-1594). Following

secondary antibody incubation, tissues were again washed in PB. Double labeled

tissues were then mounted on gelatin-coated slides and coverslipped with

Vectashield® hard set mounting media (Vector). Biotinylated secondary-tagged

sections were incubated in an ABC-peroxidase (Vector) solution for 1hr at room

temperature, washed in PB, water, sodium acetate buffer, and briefly incubated

in a DAB solution containing nickel sulfate for visualization. Sections were then

mounted on gelatin coated slides and coverslipped using Permount. BrdU-alone,

labeled sections were counterstained with methyl green prior to coverslipping so

that brain regions could be more readily identified.

Microscopy and Statistics

An Olympus BX61 microscope complete image analysis system with

CellSens suite image analysis software was used to locate cells of interest and to

obtain digital images under white light and fluorescence light. All statistics and

graphing were done using Statistica 7.1 software (StatSoft, Inc). Statisical

significance was established at p< 0.05.

For implanted cell survival, starting at the olfactory bulbs and moving caudally to the cerebellum, all BrdU+ cells were counted and the anatomical location was recorded. The number of BrdU+ cells counted was multiplied by six to achieve the total number of surviving cells per animal, as every 6th section was

used. For further analysis, the number of BrdU+ cells was categorized by

anatomical location, i.e. cortex or striatum. Means ± SEM by treatment group are

reported and a one-way ANOVA was used to determine significant differences in

NPC survival.

BrdU+ cell migration along the rostral-caudal axis was determined by locating the approximate center of the implantation site. This was identified as the tissue section in which the bore hole of the needle was the widest. For many animals, this location also corresponded to the section in which the greatest number of BrdU+ cells was counted. This section was identified as the zero point

(the bolded line at 0mm on the x-axis of Figure 16). The distance of each section

containing BrdU+ cells rostral or caudal to this zero point was identified. To

determine significant differences in migration distance, the distance of each

section was weighted by the total number of cells found in that section, thus a

section 750μm away from the zero point with 35 BrdU+ cells carried more weight

in the statistical analysis than a section at the same distance but which only had

2 BrdU+ cells. Data was then analyzed by one-way ANOVA and a post-hoc

Fisher least-significant differences test was used to identify the significant

differences between the means.

Differences in NPC differentiation were based on the percentage of implanted cells expressing both BrdU+ and MAP2 or GFAP. NPC were considered to be differentiated if they had a BrdU+ nucleus (green fluorescence)

and a MAP2+ or GFAP+ cytoplasm with mature cell morphology (i.e. neurite

extension and branching; red fluorescence). At least 100 BrdU+ cells per animal

were identified and digitally photographed using the microscope’s green/FITC

fluorescence filter. Once photographed, the fluorescence filter was switched to

the red/TRITC and again digitally photographed. Green and red images were

merged and the number of differentiated NPC were counted.

Biocompatibility of the implanted material with the host tissue was

determined by the glial reactivity surrounding the implant site. Digital images

were taken of GFAP+ or Iba1+ immunoreactivity in the striatum medial to the

implantation track. Using the microscope image analysis software, a 200μm by

200μm box was overlaid onto the images and positioned adjacent to the track.

The total number of immunopositive cells within the box was counted. At least

three boxes per animal were used. The mean number (±SEM) of cells per animal

by group was reported and a one-way ANOVA was used to determine

significance.

Four animals were excluded from analysis – two animals from the Sham

group, and one animal each from the Cells Alone and Hydrogel “S” groups – due

to excessive tissue damage in which the cortex appeared have a large cavitation

98 from the 18 guage needle. This relatively large needle was needed in order to load the polymerized hydrogel-cell complexes. All other animals appeared to have orderly transplant tracks with intact brain tissue. As such, the resulting group sizes were: Sham: N=4, Cells Alone: N=5, Hydrogel “N”: N=7, and

Hydrogel “S”: N=6.

Results

Cytocompatibility

BrdU+ cell counts indicate that cell survival after implantation was high.

On average, 34.35% (± 10.16%) of implanted cells survived up to two months in vivo. There was no significant difference in NPC survival between the groups

(F(2,15)=0.701, p>0.05; Figure 15A); however, the group implanted with

Hydrogel “N” had an average survival of 49.53% (± 23.89%). The Hydrogel “S” implanted group, with 26.91% (± 10.66%) survival, was more like the Cells Alone group which had an average survival of 22.04% (± 8.75%).

Because BrdU+ cells were found throughout the implanted hemisphere, the percent of BrdU+ cells in the cortex, corpus callosum, striatum, and subventricular zone was determined (Figure 15B). There were significant differences in the percent of implanted cells that were found in the different areas

(F(3,45)=16.405, p<0.0001) regardless of group. At least half of the BrdU+ cells were found in the overlying cortex (52.29% ± 4.94%), with the remaining cells found mostly in the corpus callosum and striatum (20.20% ± 4.33%and 24.30%

±4.87%, respectively). Only a small fraction were found in the neurogenic niche,

Figure 15. BrdU+ cells were found in the brains of all animals regardless of implantation method. A. The total number of BrdU+ cells suggests that all implanted animals had good NPC survival post-implantation. While no significant differences were found between the groups (p> 0.05), in brains implanted with hydrogel-encapsulated cells, the total number of BrdU+ cells found post-mortem tended to be higher than those implanted with cells alone. The left axis of the graph corresponds to the mean number of BrdU+ cells (± SEM). The right axis depicts this mean as a percentage of the total number of NPC implanted (20,000 cells). B. When BrdU+ cells were identified by location, most cells were found in the cortex, followed by the striatum and corpus callosum. No significant differences were found between groups as a function of brain region (p> 0.05). In all groups, the subventricular zone had the lowest percentage of BrdU+ cells. CA: Cells Alone, “N”: Hydrogel “N”, “S”: Hydrogel “S”, CTX: cortex, CC: corpus callosum, ST: striatum, SVZ: subventricular zone.

the SVZ (3.21% ± 2.04%). Between the treatment groups, there were no

differences in cell survival based on location (F(6,45)=0.384, p>0.05). All groups

had a similar location pattern described above, with most cells being located in

the cortex. Even though few were found in the SVZ, cells from Hydrogel “N”

were three times more likely to be found in the SVZ than cells which were

encapsulated in Hydrogel “S” or the Cells Alone groups (6.19% ± 5.15%

compared to 0.91% ± 0.62% and 1.82% ± 1.43%, respectively). No BrdU+ cells

were found in any other brain regions, including the hippocampus, olfactory

bulbs, or cerebellum.

100

The extent to which BrdU+ implanted cells were found along the rostral-

caudal axis was used as a measure of migration (Figure 16). Significantly greater

migration was found in the Hydrogel “N” implanted animals (F(2,275)=12.551,

p<0.0001). In this group, BrdU+ cells were found as far as 1.8mm rostral and 3.4

mm caudal. In comparison, the Hydrogel “S” and Cells Alone groups had a

narrow range of migration, especially in the caudal axis with a maximum

migration of 1.8mm.

Figure 16. The greatest migration along the rostral-caudal axis away from the implant track was seen in Hydrogel “N” implanted brains. Most BrdU+ cells were found at or near the implant track. In particular, the Cells Alone and the Hydrogel “S” treatment groups had a short rostral-caudal migration range, while cells encapsulated in Hydrogel “N” migrated further caudally. X-axis: millimeters from implant track (bolded line: 0 mm). Y-axis: Treatment groups. Z- axis: the number of BrdU+ cells in a section. Each symbol represents one brain section.

101

Co-labeling of tissue with markers for BrdU in conjunction with MAP2 or

GFAP demonstrated little differentiation of implanted cells. Only three of the 18 animals implanted with NPC had BrdU/MAP2 co-labeled cells; one animal from the Cells Alone group and two animals from the Hydrogel “S” group. In these three animals, fewer than 3% of cells were positive for BrdU and MAP2, suggesting that neuronal differentiation was extremely low. Eleven of 18 animals had cells positive for BrdU and GFAP suggesting glial differentiation. While this double labeling was seen in animals from all three groups, no group had greater than 2% BrdU/GFAP co-labeled cells (Cells Alone: 0.87% ± 0.59%; Hydrogel “N”:

0.97% ± 0.65%; Hydrogel “S”: 1.20% ± 0.58%). When present, co-labeled cells

(MAP2 and GFAP) were found randomly in the implanted hemisphere. Because of the limited neuronal and glial differentiation observed, adjacent tissue sections were labeled for Nestin as a marker of neural progenitors. In all animals, Nestin labeling followed similar distribution patterns as BrdU labeling (Figure 17).

Histocompatibility

To assess the biocompatibility of the hydrogels tested in the brain, the tissue was immunolabeled for GFAP+ astrocytes and Iba1+ microglia alone. The number of GFAP+ cells indicates that there were no significant differences in the density of astrocytes adjacent to the implant track between the treatment groups

F(3,18)=0.784, p>0.05; Figure 18A). However, there was a slight increase observed in the number of GFAP+ cells in the Sham control animals (35.29 ±

4.57/200μm2), compared with the other groups (Cells Alone: 24.44 ±

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Figure 17. Implanted NPC at the striatal and corpus callosal interface express both BrdU and Nestin. A. BrdU+ cells (black) were found in the corpus callosum (CC) on the lateral border of the striatum (St). For BrdU labeling, sections were counterstained with methyl green (light gray). B. Nestin+ cells (black) were almost always found in the same location as BrdU+ cells in adjacent sections (arrows). Almost no cells were found to co-label for BrdU and GFAP or BrdU and MAP2 (not shown). Scale bar is 100μm for both images.

5.89/200 μm2; Hydrogel “N”: 28.06 ± 4.86/200μm2; Hydrogel “S”: 31.13 ±

3.16/200μm2). Furthermore, the cell morphology in the Sham and Hydrogel “S”

groups suggests there were more reactive GFAP+ cells, as designated by large

cell bodies and short-thick processes (Figure 19A-D)[220, 291, 350]. By

comparison, GFAP+ cell morphology in the Cells Alone and Hydrogel “N” groups

demonstrated more quiescent-looking cells, with smaller soma and long-thin processes (Figure 19A-D)[220, 291, 350].

Iba1 immunolabeling for microglia revealed similar results as those found for astrocytes. There were no significant differences in the density of Iba1+ cells adjacent to the implant track between groups (F(3,18)=0.200, p>0.05; Figure

18B). Sham, Cells Alone, and Hydrogel “S” treatments all had an average 25

103

Figure 18. Both astrocyte and microglia presence indicates that the hydrogels were biocompatible. A. GFAP immunoreactivity indicated that the number of astrocytes surrounding the implant track were equivalent in all groups. The Sham group, as an indicator of standard wound healing, actually had a slightly higher number of GFAP+ cells. B. Iba1 immunoreactivity, a marker for microglia, also found no significant differences between treatment groups. CA: Cells Alone, “N”: Hydrogel “N”, “S”: Hydrogel “S”.

Iba1+ cells per 200µm2 (Sham: 25.01 ± 3.55; Cells Alone: 25.18 ± 3.84; Hydrogel

“S”: 25.38 ± 2.20). Morphologically, the Iba1+ cells looked to be quiescent, as

indicated by round cell bodies and thin processes (Figure 19E-H) [350, 351]. The lack of significant immune response to the hydrogels, as measured by astrocyte and microglial responses, compared with the Sham controls, suggests the hydrogels do not provoke greater immune response than a simple needle penetration and are thus likely biocompatible with brain tissue.

104

Figure 19. GFAP and Iba1 immunoreactivity were used as indicators of biocompatibility. GFAP immunoreactive astrocytes (A-D) were seen in all treatment groups, with a greater number of GFAP+ cells associated with the needle tract (right boarder of each photomicrograph). For Sham (A) and Hydrogel “S” (D) groups specifically, many GFAP+ cells were observed to have a strong immunoreactive morphology (closed arrows in A-D), while in the Cells Alone (B) and Hydrogel “N” (C) groups, a greater number of cells demonstrated a more non-reactive/quiescent morphology (open arrows in A-D). Iba1 immunoreactive microglia (E-H) were fewer compared to GFAP+ astrocytes. Additionally, all groups appear to have similar microglial morphologies with small round cell bodies and fewer and thinner processes, suggesting a resting/quiescent cell morphology (open arrows in E-H). In all photomicrographs, the needle tract forms the right border of the image. Each image is the best, most representative image for that group. The images were chosen based on the number of immunopositive cells shown in the photo which corresponds to the mean for that group; i.e. the image depicted in D was counted to have 32 GFAP+ cells per 200μm2, closely matching the mean for Hydrogel “S” GFAP labeling, which was approximately 31 cells/200μm2. A, E: Sham; B, F: Cells Alone; C, G: Hydrogel “N”; D, H: Hydrogel “S”. Scale bar is 50μm for all images.

Discussion

The results of the present study indicate that HA and PEG hydrogels

supported encapsulated NPC survival in vivo and allowed them to migrate out of

the hydrogel into the host environment without generating a neuroimmune

response in the host brain. The data reveal that there were no statistically 105

significant differences in in vivo cytocompatibility or histocompatibility, between

hydrogel implanted animals and controls (HBSS-implanted sham or cells alone).

Furthermore, NPC encapsulated in the higher HA-PEG content hydrogel (“N”)

ultimately had a greater migration distance than NPC implanted without hydrogel

or NPC which had been encapsulated in the “S” hydrogel. This study

suggests that these two hydrogels can be used successfully to deliver NPC into

the brain environment and therefore are viable tools for neural tissue

engineering.

Surviving implanted cells labeled with BrdU were seen in all animals. The

survival of implanted NPC, at 22-50%, was greater than what is typically seen in

implantation studies [10, 12, 31, 344, 348, 352-359]. Often the total number of

surviving cells is so small it is not reported, but of the eight studies referenced

here which report rates of survival, the average is 6.5% [10, 12, 31, 352, 355,

357-359]. In addition to the high rate of survival, I observed no differences in

survival between cells implanted without hydrogel compared with those

encapsulated within the two hydrogel formulations. My previous study has shown

that the survival of NPC in vitro is improved when the cells are encapsulated in

the Hydrogel “N” and “S” formulations compared with standard culture conditions

(114% and 111%, respectively, compared to 34.6% survival on tissue culture

polystyrene at 24 hours; Table 4) [341]. Thus, these hydrogels, based on HA and

PEG, appear to be viable materials for both in vivo and in vitro uses in brain

applications. In contrast to my results, which exhibited good cytocompatibility

with the hydrogels, a study by Uemura, et al. demonstrated poor

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cytocompatibility in vivo [33]. They showed that Matrigel, a commonly used hydrogel-like culture substrate derived from the heterogenous secretion of mouse sarcoma cells, supported NPC in vitro, but Matrigel-encapsulated cells had a high rate of death when transplanted into the brain [33]. This illustrates the importance of establishing the cytocompatibility of the biomaterial both in vitro and in vivo.

Because both hydrogel formulations used in this study had similar survival rates in vitro [341], I may relate any differences in NPC survival in vivo to the environment of living brain tissue. For example, I observed a greater degree of cell survival with Hydrogel “N” (49.53%), compared to Hydrogel “S” (26.91%).

The first few weeks following implantation results in the majority of cell death due to the host immune response to needle penetration (a.k.a. the wound healing response) [10-14, 27, 28]. Hydrogel “N”, which degrades more slowly [341], may provide greater initial protection of the encapsulated cells from this inflammatory reaction, thus resulting in more surviving cells long term. Hydrogel “S”, shown to degrade more quickly in vitro [341], may begin to degrade too quickly in vivo, exposing the encapsulated cells to the inflammatory response and thus resulting in a survival rate more similar to the Cells Alone treatment.

Despite the target of implanting the NPC into the striatum, the majority of surviving BrdU+ cells were found in the cortex, regardless of treatment. This may suggest that the implanted material (whether saline, cells alone, or hydrogel-cell

complexes) was placed in the cortex rather than in the striatum. These

conclusions are contradicted by five hydrogel implanted animals, in which a small

107 amount of un-degraded hydrogel material was only found in the striatum and not in the cortex (data not shown), indicating that the implant was correctly placed into the striatum. It is possible that the tissue damage caused by the needle might be responsible for the high numbers of implanted cells in the cortex.

Studies addressing neurological damage, both pathological and trauma-induced, have shown that NPC migrate to the site of damage and may take part in the tissue repair process [11, 13, 28, 59, 171, 237, 355, 360-365]. The cells not found in the cortex were predominately found in the striatum and corpus callosum, the majority being found at the lateral striatal-callosal interface, suggesting the lateral white matter tracks may be possible routes of passive migration. It is important to note that my implantation target was more lateral than central in the striatum. This location likely reduced medial migration toward the lateral ventricles and subventricular zones as there was a more readily accessible route via the corpus callosum. Access to the corpus callosum would allow for the migration of the NPC to all areas in which BrdU+ cells were found, including the very few cells found in the subventricular zone. The location of many BrdU+ cells in the corpus callosum, and a few in the subventricular zone, may indicate that the implanted cells were beginning to follow endogenous migration patterns often seen with host NPC [361, 363, 366, 367]; however, none of the implanted cells were found in the target areas of these endogenous migration routes, such as the contralateral hemisphere or the olfactory bulbs

[171, 363, 368-370]. Indeed, I also found it interesting that very few of the implanted cells were found in the subventricular zone which, as a neurogenic

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niche [171, 318, 371, 372], might have been the most hospitable and nearest

region for the implanted NPC.

The varied locations of the BrdU+ cells suggest that the hydrogels were not retaining the cells at the implant site. This lack of retainment may be explained by the lack of adhesion molecules in my hydrogels, such as RGD

peptides or gelatin, which are used in vitro to anchor the cells onto the material

[40, 291, 373]. Upon examination of the entire rostral-caudal extent of the brain, I

found differences in the distances BrdU+ cells migrated from the center of the

implant site. Cells encapsulated within Hydrogel “N” were spread over a

significantly longer distance along the rostral-caudal axis than BrdU+ cells from

Hydrogel “S” or the Cells Alone treatments. At the furthest extent, twenty to thirty

cells from Hydrogel “N” were found in the striatum at the level at which the

fimbria fornix emerges. In comparison, the furthest that cells from Hydrogel “S”

migrated was within the striatum at a level where the septum is still prominent

and only two cells were found at this location. This spread of implanted cells may

suggest that the Hydrogel “N” composition has properties that permit or

encourage implanted cell migration. As suggested above, the slower degradation

rate of Hydrogel “N” may segregate these cells from the initial inflammatory

reaction of the implantation procedure. This may also result in isolating the cells

from chemical cues in the injured tissue which typically promote the retention of

cells at the injury site [171, 220, 221, 223, 360, 374]. Thus, the later release of

cells from Hydrogel “N” into the tissue without such “damage” cues (or a

reduction in cue strength) may allow for further unguided migration from the

109 implantation site. In contrast, cells implanted without the encapsulating hydrogel or the more quickly degrading Hydrogel “S” may be subject to cues driving their involvement in the immune response and early repair process, keeping them nearer to the implant site.

An alternative explanation to the differences in migration distance may be that the mechanical properties of the hydrogels play a role in NPC migration by offering dissimilar hydrogel-to-tissue transitions. For example, the compressive modulus of Hydrogel “S” more closely matches the stiffness of neonatal brain tissue (Table 4). While the encapsulated cells may migrate well within the hydrogel, the mechanical differences between the hydrogel and the adult brain tissue acts as a barrier, impeding the cells from readily traveling into the brain, resulting in a smaller range of migration. In contrast, the stiffer Hydrogel “N” more closely resembles that of the host adult brain tissue, and thus there is less of a mechanical transition, allowing greater migration out of the hydrogel.

Despite the degree of migration away from the site of implantation, I found minimal differentiation of my implanted cells to mature neuronal or glial phenotypes, regardless of treatment. Only a very small fraction of the BrdU+ cells had adopted the mature protein expression and morphology associated with differentiation of either cell type. Other studies have shown similar nominal differentiation of implanted cells [31, 317, 343, 345, 348, 349, 359, 363, 375,

376]. For example, Fricker, et al. observed that the majority of NPC which were implanted into the striatum remained undifferentiated, mainly expressing markers of immature progenitors [370]. This lack of differentiation could be due to the

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decline in the ability of the adult brain to produce the cues which mature NPC

into committed adult cells – compared to the fetal brain from which my cells were

derived – as has been established [317, 371, 377-381]. I observed very slight

increases in the number of differentiated cells in animals implanted with Hydrogel

“S” compared with Cells Alone and Hydrogel “N” implanted groups. I previously

showed in vitro that Hydrogel “S” encapsulated NPC were more likely to adopt a

neuronal fate compared with other hydrogel formulations tested, including

Hydrogel “N” (Table 4) [341]. Interestingly, this current study indicates that both neuronal and glial differentiation were slightly increased in vivo in cells which were encapsulated within Hydrogel “S”, though neither to the levels which were attained in vitro. Hydrogel “S” may have a different rate of differentiation compared to Hydrogel “N” because it is less stiff and more closely resembles the mechanical properties of the fetal brain [341] where there is greater propensity for differentiation than in the adult brain [379, 381]. However, because the number of differentiated cells was so small, similar mechanical properties alone cannot compensate for cues that may promote or inhibit cellular differentiation in vivo as it did in vitro (Table 4).

My study of the hydrogel histocompatibility with host brain tissue

demonstrates the glial responses to the hydrogels are no different than the

response measured in the Cells Alone and Sham implants. In this study, I was

most interested in the long-term immune response, when the tissue would be

exposed to the degradation products of the hydrogel, rather than an acute or

wound healing response. The Sham implanted group was included as an

111 indicator of the standard wound healing process [89, 382] which typically resolves into a small but long-term increase in the local resident glial population

[222, 250, 252, 383]. An exacerbated long-term immune response would be indicative of the brain’s immunorejection of the NPC, the hydrogels, or their degradation products. For my study I chose a two-month time point to allow the initial inflammatory reaction to recede, and for the assembly, if any, of a clinically relevant immunorejection response to the implanted hydrogel, the degradation products, or the implanted cells. This would be indicated by a persistent recruitment and activation of a large number of astrocytes and microglia at the implant track [172, 183, 249, 291]. GFAP+ astrocytes within the tissues from all groups demonstrated both quiescent and reactive morphologies, indicating a mild response to the implant [183, 220, 223, 374, 384]. The continued lack of differences in microglia between groups was unsurprising, as the activity of microglia is more classically associated with the initial inflammatory reaction rather than the long-term repair response [183, 221]. The results of astrocyte and microglia recruitment and activation indicate that the hydrogels were not inciting an immune response beyond that of the implantation procedure itself and provides convincing evidence that the hydrogels tested here are biocompatible

[291].

In conclusion, the results suggest that the hydrogel formulations tested in this study are biocompatible in vivo with both the encapsulated NPC and the brain tissue into which they were implanted. Measures of implanted cell survival indicate that the hydrogels were useful in delivering NPC into the brain and did

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not prevent migration of implanted cells away from the hydrogel and into the

surrounding tissue. However, modifications to the hydrogels, such as the addition

of various cell attachment factors, may further improve survival and/or confine

cells to a specific site within the tissue, if those characteristics are desirable. The

prevention of migration may be an important property in treating some “localized”

disorders, such as Parkinson’s disease, in which a specific region of the brain is

affected. Differentiation results indicate that the pre-differentiation of cells prior to

encapsulation or the incorporation of differentiation cues into the hydrogel may

be necessary to produce more mature phenotypes, as the unassisted maturation

appears to be occurring in a very limited fashion in vivo. The insignificant glial reaction which paralleled the response to the control implant treatments suggests the selected hydrogels are well-suited for use in the brain. However, as previous studies have shown, just because these hydrogels were biocompatible with adult rat brain should not be proof of biocompatibility for all species of animal, at all ages – the importance of property matching to host tissue remains [40, 291]. The

results of this study further support the use of an HA and PEG hydrogel for brain

applications and advocates, with additional specialization, the use of these

materials as a viable tool for clinical implementation in the future.

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CHAPTER V

SUMMARY AND CONCLUDING REMARKS

The characterization of hydrogels biocompatible for neural tissue engineering involves three basic steps: 1. defining the tunable properties of the hydrogel, 2. establishing the material cytocompatibility with neural cells, and 3. establishing the histocompatibility with host brain tissue. This thesis presented a series of studies which established these three characteristics using combinations of hyaluronic acid (HA) and poly(ethylene glycol) (PEG) to produce customizable hydrogels with definable chemical, physical, and mechanical characteristics. Initially, twenty-five hydrogels were generated and their tunable properties were defined. The interaction between hydrogel polymerization, degradation, and compressive modulus (the tunable properties) on the survival and fate of two populations of neural progenitor cells (NPC) was studied. Finally, an in vivo investigation of the hydrogel interaction with NPC and surrounding tissue in the brain provided indications as to the biocompatibility of the material in its intended use. Through thorough characterization of these HA and PEG hydrogels, this thesis demonstrates the production of biocompatible materials that can be used in numerous settings to study and potentially treat neurological diseases and disorders.

Hydrogel Properties

The tunable properties measured in this thesis suggest that the HA and

PEG hydrogels used here can be formulated with properties physiologically

114 relevant to neural tissue engineering. The relatively simple, two-component chemistry of HA-PEG hydrogels was chosen with the chemistry of the brain extracellular matrix (ECM) in mind. HA is a prudent material for use in hydrogels geared toward the brain since it is both a major component of the brain ECM and can be degraded in vivo by endogenous hyaluronidase [292, 293, 334]. Though a synthetic material, PEG is biologically inert and its use provided for additional control over the properties of the hydrogel, including hydrogel stiffness. The use of HA and PEG is in contrast to the majority of hydrogels designed for peripheral tissue engineering, which frequently contain polymer components not regularly found in the brain, such as collagen or fibronectin [77, 79, 243, 245].

The hydrogels investigated here, which were comprised of relatively low concentrations of HA and PEG, demonstrated a polymerization process feasible for neural cell encapsulation. The hydrogels polymerized between two and forty minutes at room temperature. This rate of polymerization occurred quickly enough that the cells were not put at risk by being outside the tissue culture conditions for an extended period, but was also slow enough to allow for the distribution of the cells three-dimensionally within the hydrogel prior to complete polymerization. Importantly, these hydrogels polymerized without the addition of a chemical or thermal catalyst. More complex polymerization processes induced by catalysts are potentially harmful to encapsulated cells. For example, the UV photopolymerization process can produce reactive chemical species damaging to cells [104-106]. Whether for three-dimensional culture or for implantation, cells must survive the polymerization process. The modified HA and PEG polymers

115 used here form hydrogels by covalent bonding under bench top conditions, and thus had a greater likelihood of being biocompatible.

Hydrogel degradation is also a necessary function to attain biocompatibility in the brain. With non-degradable materials, such as chronically implanted electrodes, a significant glial scar emerges and eventual immunorejection occurs, causing disruptive tissue damage [120-123]. When biomaterials are degradable, the glial response has the opportunity to resolve itself over time.

The HA-PEG hydrogels used in this thesis were demonstrated to have a range of in vitro degradation rates from less than 12 hours, up to 12 days in hyaluronidase solution. Slower degradation rates were observed in hydrogels with higher concentrations of HA and PEG. Hyaluronidase is an enzyme produced by both neurons and glia in the brain to specifically degrade HA. Thus,

HA and PEG hydrogels are susceptible to degradation in vivo via the same hyaluronidase activity that could be produced by both the surrounding tissue and encapsulated cells. While it is assumed that the relative degradation rates in vitro reflect the degradation rates in vivo, two months post-implant six of the fourteen animals implanted with hydrogel had some un-degraded material remaining - two from the Hydrogel “S” group and four from the Hydrogel “N” group. Hydrogel “N” had a higher polymer content than Hydrogel “S” suggesting that, though the time course is altered, the trend of higher polymer content hydrogels degrading more slowly continues to hold true in vivo.

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The mechanical properties of the hydrogels tested encompassed those of

rodent brain tissue from animals of different ages and species. Here, it was

demonstrated that hydrogel stiffness was mostly dependent on HA concentration

and that PEG contributed to small modifying effects. As HA concentration

increased, and at high HA concentrations, as PEG increased, the compressive

modulus of the hydrogel was increased. A number of the hydrogels

demonstrated compressive moduli similar to those measured of rat brain tissue

samples in animals ranging from post-natal day 7 to more than 4 months of age, and the compressive moduli of adult mouse brain tissue. Brain tissue samples demonstrated increasing compressive modulus with age, indicating that as the animal matures, the brain becomes stiffer. Considering the findings here, and since HA is a major component of the brain ECM, one might be tempted to think that an increase in HA in the brain may contribute to this increased brain compressive modulus as in the hydrogels. However, this is contradicted by studies which have shown that HA concentration decreases during maturation

[385, 386]. Further study of the chemistry and components of the natural brain

ECM is needed to understand the contributions of these molecules on the increasing brain compressive modulus and how hydrogel chemistry might be better tailored to the ECM of animals of different ages.

Though they were chosen because of their physiological relevancy, the hydrogels used for the implantation studies in this thesis had different compressive moduli. Hydrogel “N” had a compressive modulus that more closely matched the adult brain, while the compressive modulus of Hydrogel “S” was

117

closer to that of a younger animal. That the two hydrogels implanted into the

brain did not demonstrate significant differences in biocompatibility suggests that

the brain may be tolerant of slight differences in material compressive modulus.

The matching of hydrogel stiffness to tissue stiffness has been continually

demonstrated to improve biocompatibility [40, 47, 143, 160, 161, 164, 291, 305].

Thus, it was expected that a hydrogel with a compressive modulus closely

matching younger tissue may not be as compatible as a hydrogel that matches

mature tissue when implanted into the adult brain. Future studies of the in vivo

biocompatibility of HA-PEG hydrogels with a wider range of compressive moduli may expose the extent to which compressive modulus can differ from the brain tissue before becoming incompatible.

In Vitro Biocompatibility

The hydrogels demonstrated good in vitro biocompatibility with

encapsulated NPC, as measured by survival and differentiation. Survival of

encapsulated NPC was dependent on the hydrogel formulation, with the highest

levels of survival at the lowest concentrations of HA and PEG. Interestingly, fetal-

derived NPC (fNPC) and adult-derived NPC (aNPC) responded differently to the

same hydrogel formulations. For example, in hydrogels with higher compressive

moduli, fNPC survival was still relatively high, while aNPC did not survive as well

in these stiffer hydrogels. Survival at this stage (after only 24-hours of

encapsulation) is likely a function of hydrogel chemistry, polymerization,

encapsulation, and to a lesser extent, though still relevant, compressive modulus.

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At this time point, hydrogel degradation presumably has little contribution to

survival. Overall, based on 24-hour survival, fNPC responded better to

encapsulation when compared to fNPC tissue culture polystyrene (TCP) controls,

in contrast to aNPC which did not survive encapsulation as well when compared

to aNPC TCP controls. This may suggest that either the polymerization process or the act of being encapsulated is more difficult for adult-derived cells, thus decreasing the biocompatibility between the hydrogels and aNPC. For this reason, fNPC were chosen for the implantation studies.

Encapsulated NPC differentiated into neurons and glia within the hydrogels after three-weeks in vitro. Given that this differentiation was unaided by cues from trophic factors added to the culture conditions suggests the hydrogel environment was sufficient to drive differentiation. Differences in NPC differentiation based on hydrogel formulation were also observed. There were two distinctive trends of differentiation related to hydrogel compressive: glial differentiation of fNPC was increased in stiffer hydrogels, while neuronal differentiation of aNPC was increased in softer hydrogels. These results were congruous with previous studies, in which stiffer hydrogels provided greater support for glial cells, while neuronal populations did better on softer hydrogels

[47, 143, 144, 160-164]. None of these reports, however, directly compared the culture of fetal-derived and adult-derived cell populations within hydrogels of the same formulations, as done in this thesis.

An interesting finding, which may only be indirectly related to hydrogel encapsulation, was the difference in fate potential of the cells based on their

119

source. More specifically, the majority of fetal-derived cells which differentiated became glial cells regardless of the hydrogel formulation. In contrast, the majority of differentiated adult-derived cells were neuronal. This suggests that, though both cell types were predominantly nestin-positive populations prior to encapsulation, the differentiation capactity of these cells might be different independent of environment. Regardless of these differences, that the hydrogels support differentiation indicates that they can support the long-term maintenance of both types of NPC, making them biocompatible in vitro.

In Vivo Biocompatibility

Establishing the in vitro biocompatibility of HA-PEG hydrogels with neural

cells and delineating the fates of encapsulated cells helps to transition these

materials directly into an in vivo model. The data from these studies

demonstrated that the HA-PEG hydrogels maintain their biocompatibility when

transitioned from in vitro to in vivo studies, making them applicable for potential

clinical use. Not all materials used for in vitro purposes can be transitioned to use

in vivo. Matrigel, which is commonly used as a three-dimensional culture

substrate, cannot be effectively used for neural implantation studies because of

its unknown chemistry and propensity to cause an inflammatory response in the

brain [33]. Thus, materials that are more thoroughly defined, such as those used

in this thesis, stand a greater chance of use in the brain, especially after

demonstrating good compatibility with neural cells in vitro.

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HA-PEG hydrogels demonstrated good biocompatibility in both cyto- and histo-compatibility measures when implanted into the brain. The hydrogels were used to successfully deliver encapsulated fetal-derived NPC, with 20-50% survival at two months post-implantation. This is greater than what is typically seen of implanted cell survival without hydrogel encapsulation [10, 12, 31, 344,

348, 352-359]. However, in these studies encapsulated cell survival was not significantly greater than the cells alone control, which had ~20% survival. The high rate of survival in all groups, including the cells alone control group, might have been a function of the needle size used for implantation. A large needle was necessary for delivery of the hydrogels, but the increased internal diameter may also have decreased the shearing stresses on the cells during the procedure, contributing to good survival. Unfortunately, the large needle size also resulted in significant tissue damage in the cortex. Nevertheless, survival of implanted cells encapsulated within the hydrogels was high enough to indicate good in situ cytocompatibility.

Independent of survival, implanted cells were observed to have migrated from the site of implantation into the surrounding brain tissue. Cells encapsulated in Hydrogel “N” migrated significantly further into the brain than cells encapsulated in Hydrogel “S” or from the Cells Alone group. Given this singular, two-month time point, it is hard to say how the implanted cells might be transitioning from the hydrogel to reside in the surrounding tissue. This could be occurring passively through degradation of the hydrogel, subsequently causing release of the cells, or through the active migration of cells out of the hydrogel

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prior to degradation. Future studies at earlier time points following implantation

may address the mode of implanted cell migration. The migration of cells

indicates that the hydrogel material does not present a barrier to the integration

of encapsulated cells into the brain.

No differences were observed in the immune response of the host brain tissue to the implanted material, further confirming the biocompatibility of HA-

PEG hydrogels. The number of astrocytes or microglia adjacent to the site of

implantation in hydrogel-implanted tissue and in the tissue up to 200 microns

away, was similar compared to sham-penetrated controls. That the glial reaction at the implantation site was no greater than the standard wound healing reaction observed in the control tissues suggests that the hydrogels and the degradaded components were non-toxic. Since the hydrogels were chosen for the physiological relevancy of their chemical and mechanical properties, the data confirmed expectations.

Future Directions

For the application of these hydrogels in a clinical setting, further

characterization would be required. Properties such as conversion efficiency

during polymerization (the efficiency of bond formation and the presence of

unreacted polymer), additional mechanical properties, such as elasticity and

response to shearing, and rates of in vivo degradation could be explored. The

more basic chemistry of the HA-PEG hydrogels studied here make this process

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of complete characterization more simple, as every addition to a hydrogel adds

complexity and additional steps to the characterization process.

Cell survival, both in vitro and in vivo may be improved by the addition of attachment motifs, such as the RGD peptide sequence. Nervous system cells in particular, survive better when they have something to attach to. While the 3D environment of the hydrogel can provide the structural physical surroundings to support survival, the chemical bonds formed when a cell attaches to a substrate are likely to improve survival. However, the addition of attachment factors may also affect other cellular processes such as migration and differentiation.

Growth factor-modified hydrogels could be used for specific, therapeutic purposes. The addition of growth factors into the hydrogel may further improve the survival of implanted cells, increase the rates of progenitor differentiation, or encourage reinnervation of damaged areas. For example, the repair of tissue in a stroke-infarcted brain will require replacement of both neuronal and glial cells and an environment which encourages infiltration of blood vessels. This is in contrast to a parkinsonian brain, in which a specific population of neuronal cells is damaged. In the former condition, a hydrogel that supports a variety of cell types and contains growth factors stimulating formation of blood vessels would be preferred, while in the latter, a hydrogel containing additional cues that results in differentiation of encapsulated cells to dopaminergic neurons and the reconstruction of the nigrostriatal tract through growth factor-guided neurite extension would be more beneficial. Customization of HA-PEG hydrogels by the addition of growth factors and the encapsulation of specific cell populations could

123 potentially produce both of these environments using the same hydrogel foundation; however, increasing the complexity of the hydrogel chemistry is more likely to contribute to a greater immune response from the host tissue. Thus, any modification of hydrogel materials requires thorough investigation as to the biocompatibility prior to application.

Conclusions

The studies described in this thesis were designed to investigate the formulation of HA and PEG hydrogels with properties matching that of the brain tissue ECM environment and which would be biocompatible in vitro and in vivo.

Through completion of this research, I have defined twenty-five HA and PEG hydrogels with properties which are physiologically relevant to the brain. The hydrogels were demonstrated to be cytocompatible with neural progenitor cells derived from fetal and adult rats, where the fate of the encapsulated cells was a function of the hydrogel formulation and the source of the cells. Finally, the HA-

PEG hydrogels chosen for implantation into the brain were determined to be biocompatible with brain tissue and did not prevent cells from migrating into the surrounding brain tissue. Further exploration of these hydrogels, including additions and modifications to make the materials more functionally relevant, could have great impact on improving neural tissue engineering techniques in the future.

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