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NATURAL POLYMERIC CARRIERS FOR LOCAL DELIVERY

OF THERAPEUTIC AGENTS TO PROMOTE ENHANCED

BONE REGENERATION

BY

CHRISTINE JANE KOWALCZEWSKI

A Dissertation Submitted to the Graduate Faculty of

WAKE FOREST UNIVERSITY GRADUATE SCHOOL OF ARTS AND SCIENCES

in Partial Fulfillment of the Requirements

for the Degree of

DOCTOR OF PHILOSOPHY

Biomedical Engineering

December 2014

Winston-Salem, North Carolina

Approved By:

Justin M. Saul, Ph.D., Advisor

Examining Committee:

Karl-Erik Andersson, M.D., Ph.D.

Aaron S. Goldstein, Ph.D.

Thomas L. Smith, Ph.D.

Mark E. Van Dyke, Ph.D.

Stephen J. Walker, Ph.D.

ii

TABLE OF CONTENTS

ACKNOWLEDGEMENTS……………………………………..………….………….....…iii-vi

LIST OF ILLUSTRATIONS AND TABLES……………………………………...……vii - viii

ABBREVIATIONS…………………………………………………………………………. ix-x

ABSTRACT…………………………………………………………………………..…….xi-xii

CHAPTER I

Introduction into the Current Treatment and Microenvironmental Challenges to

Regenerative Medicine for Large Bone Defects………………………………….……….1

CHAPTER II

Current Strategies to Balance Therapeutic Delivery of Bone Morphogenetic Protein-2

from Natural Polymers………………………………………………...………..……..…20

CHAPTER III

Modulation of Kerateine Material Properties for the Controlled Release of

rhBMP-2………………………………………………………………………………....41

CHAPTER IV

Reduction of Ectopic Bone Growth in Critically-Sized Rat Mandible Defects by Delivery

of rhBMP-2 from Kerateine Biomaterials ……………………………...…………....….70

CHAPTER V

Surface-Mediated Delivery of siRNA Using Polycationic Complexes for the Knockdown

of Inhibitory Molecules……………...……………………….…………………..…..…107

CHAPTER VI

Knockdown of Bone Morphogenetic Protein-2 Binding Antagonist Noggin by Delivery

of siRNA from Fibrin Hydrogels………………………...………………………...…146

CHAPTER VII

Conclusion and Prospectus……………………………………..………………………167

CURRICULUM VITAE…………………………………………………………………..……176

iii

ACKNOWLEDGEMENTS

“Any road followed precisely to its end leads precisely nowhere.”

-Frank Herbert, DUNE

The road of my PhD has been a meandering journey from starting at WFU to moving to

Miami University and starting new projects. Even though I had my reservations in the beginning

of the move to Miami, I look back and appreciate the wonderful experience I would not have had

if I decided to stay at WFU. I know that it would not have been as enjoyable if it were not for the

people who helped me along my path.

To my advisor and mentor Dr. Saul (aka “boss”) who always needed more cowbell, thank

you for knowing when to test me and when to give me free reign. For checking me if I started to

stray too far away when I lost track of the big picture. Also, for kicking me out of lab when I

needed to get away and telling me to STOP when I needed to stop. But most importantly, for giving

me the opportunity to work with you. I couldn’t imagine anyone else who would have guided me

through this journey better; my thesis work would not have been as enjoyable without your

guidance. I want to also thank Sara Saul, who I know was behind the scenes supporting Dr. Saul

as well as me. Thank you for reminding him to give positive reinforcement every once in a while

and for being his support so he could spend time working on the science he loves.

To my committee: KE – Thank you for always suggesting the next book I should read that

was not a science article! For always making sure that all of your students balanced their love of

science with a love for life and a good bottle of wine. AG – For keeping your sense of humor,

even in the middle of my Preliminary Exam to make me laugh in the middle of one of the most

iv

difficult milestones in this journey. Your constant sense of enthusiasm for science was always

shining through and infectious whether it was in class or just catching up at the Symposium. TS

– Thank you for helping me develop my animal model and reminding me to keep the clinicians in

mind when thinking about R&D. For always being so positive and supportive. MVD – Thank you

for always taking the time to be such an amazing teacher and to make sure I understood what was

going on, even if you had to resort to metaphors to get it through my thick skull! My appreciation

for keratin’s diversity wouldn’t be what it is now if it hadn’t been for you being there when I

started my first keratin experiments. Also, for welcoming into the keratin group and offering to

help me those 6 months Dr. Saul was already at Miami. SW – I will always be thankful of your

open door (me usually frantically running through it) and helping to guide me through the

beginnings of my RNA work. I doubt I would have enjoyed my siRNA research if I hadn’t had

your guidance at the very beginning.

To the Yin to Dr. Saul’s Yang, Seth Tomblyn thank you for being the counter- weight who

I could unhesitatingly talk to and feel as though I could sound silly with you never thinking that I

was. Which brings me to Mary Ellenburg, I don’t know how I would have made those six months

at WFU before I joined Dr Saul at Miami without you. Thank you for always double checking my

math and giving me the support to push through all of those assays.

My master’s PI Claudia Fischbach-Tesclh, thank you for starting me on my path. I know

I wouldn’t be here if it weren’t for your guidance and faith in me. To the rest of the Fischbach

Lab during my time; Emily, Dan, CC, Scott, Sid, Rahmi, Chris, Bo – I loved the time I spent with

you. You all taught me so much about research and provided the hands on fundamentals which got

me through those early days of my graduate work.

v

To my WFU family - Joanne, Tom, Jan, Cathy, Prof. Wilson, Tamer, Hadi, The D’s (Dan,

Dipen, Dave, and Doug), Oscar (best long term sofa-crasher/roommate), Katie, Emma, and

Hannah. Thank you for always listening when I became frustrated and for talking me through it.

The lab was a place I always loved and it was because you were all there. The hardest part of

leaving WFU was because I was leaving you. Pedro and Amy – to the best roomies ever! I miss

your positive and supportive attitudes. You guys were always there for me. Johnny-boy – for being

my partner in crime. Roche – for always helping everyone and for showing me the tricks to the

keratin trade.

To my Miami family - Nancy, Mina, Steve, Jeff, Matt, Ash, Patricia, Ishmial. Miami would

not have been a home if it weren’t for you guys. I am so thankful that I pushed and bullied my

way into your lives. Our family dinners were always the best part of my week and you kept me

laughing. I have to give a special thanks to Jeff (and Mom and Dad Triplett) for bringing me in

and providing unconditional support and love. Jeff, for being amazing: driving me to my prelim,

listening to all my practice talks, proof-reading….on and on.

To the Miami Coworkers – Judy, Candace, and Laurie – I already miss our lunch dates.

Judy – Thank you so much for all of your work and keeping up the fun energy at work. The lab

wouldn’t flow as calmly if you weren’t there. Dave, Ryan, Sangheon, Sarah, Salma, Michiko,

Martha, Rachael, Ken, Trevor, Jay-Paul – You guys were amazing. I was so luck that I had such

a great group of lab mates.

To my family, for understanding that I never knew when I would be home. Mommy –

Thank you for being the strong woman who never gave up on making sure that I reach my potential

and for being an example of someone who loves their work. I don’t think I would have went back

to lab in the middle of the night for time points if I didn’t see that it was ok to have fun at work

vi

and be a little crazy. Daddy fatty – I think you gave me the best tool I had during my PhD.

Growing up you never answered questions right out and would tell me to “Go look it up.” I don’t

think I would have been as independent and curious if it wasn’t for those four little words. Andy

– I’m so lucky that my best friend on this planet turned out to be my brother. Danger Siblings till

the end. Garett – Thanks for always making me smile and sticking by my side during these past 7

years of graduate school. I am so lucky to have a partner who helped me anyway he could,

including the dishes.

vii

LIST OF ILLUSTRATIONS AND TABLES

CHAPTER I

No Illustrations or Tables

CHAPTER II


CHAPTER III

Figure 1. Schematic of extraction method and fabrication of kerateine biomaterials…………..51

Figure 2. Rheological characterization of kerateine hydrogel…………………………..…..…..52

Figure 3. In vitro rhBMP-2 carrier characterization……………………………………..……..54

Figure 4. µCT images of bone regeneration in critically-sized rat mandibular defect………....55

Figure 5. Quantification of percent bone regeneration and bone mineral density…..………..…56

Figure 6. Quantification of ectopic bone formation……………………………...…………..….57

Figure 7. Histological images of bone regeneration………………………………………….....59

CHAPTER IV

Figure 1. Electrophoretic mobility shift assay of siRNA transfection agents……………..….....85

Table 1. Dynamic light scattering measurements of siRNA complex diameters………………..85

Figure 2. siRNA complex adsorption and size distribution on fibrin hydrogel surface……..….87

Figure 3. Release kinetics of siRNA complexes from fibrin hydrogel surface……...........….…88

Figure 4. Schwann cell internalization of siRNA complexes…………………………………..90

Table 2. Transfection efficacy of surface-mediated siRNA into Schwann cells………...….......92

Figure 5. Flow cytometry gate analysis of Schwann cells………………………………...........92

Figure 6. Schwann cell viability after surface-mediated siRNA transfection…….……….….....93

Figure 7. qRT-PCR of MAG mRNA after surface-mediated siRNA transfection……….....…..94

viii

CHAPTER V

Figure 1. Schematic of surface-mediated transfection experimental design……..........………114

Figure 2. FE-SEM images and size characterization of surface adsorbed Lipofectamine

siRNA complexes to a fibrin hydrogel surface…………...……………………....…124

Figure 3. Release kinetics of Lipofectamine siRNA complexes from fibrin surface………....126

Figure 4. Transfection efficacy of Lipofectamine siRNA complexes into MC3T3-E1………..127

Figure 5. MC3T3-E1 internalization of Lipofectamine siRNA complexes…………….……...128

Figure 6. Noggin mRNA expression changes with rhBMP-2 dose………………………...….129

Figure 7. MC3T3-E1 cytotoxic effects of Lipofectamine siRNA complexes………….……...130

Figure 8. MC3T3-E1 noggin mRNA levels with rhBMP-2 challenge after

surface-mediated transfection of Lipofectamine siRNA

complexes………..…………..…………………………………………………..…..132

CHAPTER VI


APPENDIX A

Figure S1. Keratin extraction and biomaterial formation……………………………….....…..168

Table S1. Kerateine hydrogel formulations………………………………………………..…..169

Figure S2. Effect on weight percent and alpha:gamma on kerateine hydrogel

material properties...................................................................................................172


degradation…...........................................................................................................173


release of rhBMP-2……..........................................................................................175

ix

LIST OF ABBREVIATIONS

α Alpha Keratin Fraction

α-MEM Alpha Modified Eagle’s Medium

ACS Absorbable Collagen Sponge

ANOVA Analysis of Variance

BMP Bone Morphogenetic Protein

CaCl2 Calcium Chloride

DBM Demineralized Bone Matrix

DI Water Deionized Water

DLS Dynamic Light Scattering

DMEM Dulbecco's Modification of Eagle Medium

DNA Deoxyribonucleic Acid

η’ Viscosity

EMSA Electrophoretic Mobility Shift Assay

ECM Extracellular Matrix

FBS Fetal Bovine Serum

FGF Fibroblast Growth Factor

γ Gamma Keratin Fraction

G* Complex Modulus

G’ Elastic Modulus

G” Viscous Modulus

IGF Insulin-like Growth Factor

KTN Kerateine

KOS Keratose

MAG Myelin-Associated Glycoprotein

µg Microgram

mg Milligram

x

µL Microliter

mL Milliliter

mM Millimolar

NOG Noggin

N:P Nitrogen : Phosphate

PBS Phosphate Buffered Saline

PDGF Platelet-Derived Growth Factor

PEI Polyethylenimine

pDNA Plasmid DNA

qRT-PCR Quantitative Reverse-Transcription Polymerase Chain Reaction

rhBMP-2 Recombinant Human Bone Morphogenetic Protein-2

RNA Ribonucleic Acid

RNAi RNA Interference

siRNA Small Interfering RNA

SEM Scanning Electron Microscopy

SEM Standard Error of the Mean

siRNA-AF488 Alexa 488 Tagged siRNA

siRNA-NT Non-Targeted siRNA

siRNA-GFP Green Fluorescent Protein targeting siRNA

siRNA-MAG Myelin-Associated Glycoprotein targeting siRNA

siRNA-NOG Noggin Targeted siRNA

STDEV Standard Deviation

Tan δ Ratio of Viscous Modulus to Elastic Modulus

TERM Tissue Engineering and Regenerative Medicine

TGFβ Transforming Growth Factor Beta

USD United States Dollar

VEGF Vascular Endothelial Growth Factor

w/v Weight/Volume

xi

ABSTRACT

Bone has the innate ability to heal, but on occasion defects surpass a critical size capable

of spontaneously healing. These critically-sized bone defects pose significant challenges for

reforming bone continuity and often require surgical intervention to facilitate bone regeneration.

Recently, products combining biomaterials with recombinant human bone morphogenetic protein

2 (rhBMP-2) have proven to be a successful clinical alternative to autografts in promoting bone

regeneration. While collagen rhBMP-2 carriers have shown robust regenerative capabilities,

adverse side effects such as ectopic bone growth are likely caused by a burst release of

supraphysiological levels of rhBMP-2 from the carrier. We hypothesize that improved bone

regeneration can be achieved through tailored rhBMP-2 release from keratin carriers or through

fibrin surface-mediated delivery of siRNA targeting rhBMP-2 antagonists. For this reason, we

developed an alternative natural polymer rhBMP-2 carrier from keratin. Keratin can be formed

into a number of biomaterials which have advantages over collagen including controlled rates of

degradation, flexible material properties, and tunable rates of rhBMP-2 delivery. In a critically-

sized rat mandibular model the keratin biomaterials reduced ectopic bone growth, and achieved

comparable levels of bone healing to existing collagen-based clinical alternative for craniofacial

injuries. However, our keratin carriers still delivered a supraphysiological concentration of

rhBMP-2. We have separately observed the ability of small interfering RNA (siRNA) delivered

from fibrin carriers to knockdown myelin-associated glycoprotein (a molecular inhibitor to nerve

regeneration). We therefore attempted to apply the same approach to bone regeneration. In order

to lower the therapeutic rhBMP-2 dose, targeting rhBMP-2 antagonists such as noggin through

substrate-mediated delivery of siRNA complexes can potentially increase local rhBMP-2 efficacy.

xii

Fibrin, a natural polymer involved in bone healing, is able to electrostatically bind siRNA

complexed with polycationic transfection agents to its surface in order to locally control molecular

inhibitors of bone regeneration. Sustained transfection of sequence specific siRNA targeting

noggin was able to achieve dose-dependent reduction of noggin mRNA levels after administering

a supraphysiological dose of rhBMP-2. In conclusion, regeneration of critically-sized bone defects

can be enhanced through the delivery of therapeutic agents to control local molecular cues from

natural polymers.

1

CHAPTER I

INTRODUCTION TO THE CURRENT TREATMENTS AND

MICROENVIRONMENTAL CHALLENGES IN REGENERATIVE MEDICINE FOR

LARGE BONE DEFECTS

Christine J. Kowalczewski

2

1. Clinical Significance

Bone tissue is a dynamic system which is continuously being remodeled on a day

to day basis [1]. When an injury occurs the normal healing process restores the functional

(biological and mechanical) properties of the system. Unfortunately, the native healing

potential of bone is occasionally unable to mend itself due to a number of factors including:

smoking, malnutrition, congenital disease, and cancer. Further, in some cases healthy non-

pathologically weakened individuals experience the inability for bone closure and a return

to normal function due to large defects caused by trauma. For example, it is estimated that

10% of bone fractures in the United States result in impaired or incomplete healing [2]. A

lack of osteogenic cells, signaling molecules, stability, osteoconductive scaffold, and

inadequate vascularity all contribute to the inability of bone to heal [3].

Bone defects which surpass a size that can spontaneously heal during the lifetime

of the individual are known as critical-size defects. These critically-sized defects frequently

lead to secondary complications and development of a non-union [4]. In order for a large

defect to heal and be restored to function, a surgical intervention and placement of a bone

graft at the injury site is required to attempt to bridge the defect area. Bone grafts are

currently the second most commonly transplanted tissue in the United States with an

estimated 500,000 to 1.5 million bone-grafting procedures performed yearly accumulating

in a $1.6-$2.5 billion market [5,6].

3

2. Current Clinical Strategies

According to the American Academy of Orthopedic Surgeons the “ideal bone graft”

should be “biocompatible, bioresorbable, osteoconductive, osteoinductive, structurally

similar to bone, easy to use and cost-effective.” [7] Osteoconductive materials support bone

healing through vascularization, architecture, chemical composition (e.g. calcium sulphate,

calcium phosphate, calcium hydroxyapatite, etc), and surface charge [8]. Osteoinduction

refers to the process that supports the migration and proliferation of mesenchymal stem

cells as well as promoting differentiation of preosteocytes [9].

2.1 Autografts, Allografts, and Xenografts

Autologous bone grafts remain the clinical “gold standard” for bone defects

because it has both osteoconductive and osteoinductive properties. Autografts are most

often harvested form the iliac crest or other bone [7]. Even though the practice of autograft

transplantation has no immunogenic response, it has a limited supply, donor site morbidity,

and up to 30% failure rate [10]. One solution to overcoming secondary trauma and limited

supply of autogenic bone grafts is to use allogenic or xenogenic bone sources.

Allografts harvested from cadaveric sources have the advantage of being

osteoconductive and osteoinductive while being readily available through regional tissue

banks in a variety of sizes and shapes. However, there is a limited supply, and allograft

treatments are associated with risk of rejection and transmission of infectious disease [11].

Tissue processing and sterilization through freezing or 25 kGy gamma irradiation virtually

eliminates the possibility of disease transmission. Unfortunately, irradiation causes a

number of adverse effects on tissue properties such as weakening its strength [12].

4

Xenografts are primarily harvested from bovine origins and are

osteoinductive/conductive. Research stemming from clinical trials has determined

xenografts, primarily from bovine origins, to be unsuitable due to increased risk of

infection, immunogenicity, and host rejection [13,14].

2.2 Demineralized Bone Matrix

Demineralized bone matrix (DBM) products derived from allograft sources once

comprised ~50% of the bone graft market [5] but since the emergence of a number of FDA

approved tissue engineered bone substitutes DBM now accounts for ~20% of the grafting

market [15]. Commercial procurement of the allografts from tissue banks (to create DBM)

is variable due to donor age, health, and sterilization techniques which are not standardized,

resulting in a highly variable source material. FDA regulations mandate complete records

be obtained including family consent, donor medical records and social behavior which

would identify exclusionary risk factors. DBM human-derived tissue product or

alloimplant is processed to have no viable cells. Powder, putty, pastes, and flexible sheets

often used as an accessory or filler for other bone substitutes are not often used alone in

regenerative applications. The DBM must be reconstituted with a viscous carrier such as

a water-soluble polymer or glycerol. Osteogenic activity and clinical outcomes using DBM

are highly variable because of the inherent differences in donor material [16]. Non-

standardized production and lack of precise control conditions needed for storage to

maintain bioactivity has resulted in inconsistent results with different products as well as

product lot numbers [15,16]. For this reason, the use of bone substitutes have become

increasingly popular in the orthopedic applications due to their predicable outcomes.

5

2.3 Current Bone Substitutes

Bone substitutes have been developed and evolved in response to the shortcomings

of autografts, allografts, and DBM products. Bone substitutes currently are the fastest

growing segment in the bone graft market and account for over 50% of sales according to

the Hospital Purchasing Database [5]. They are classified into ceramic-based, polymer-

based, or growth factor-based bone grafts.

Ceramic-based products are relatively inexpensive, uniform in quality, unlimited in

supply, and lack the risk of disease transmission; however they are strictly osteoconductive.

On their own they do not play a biological role in regulating osteoinduction. Calcium

sulfate is absorbed within one to three months resulting in a mismatch to the rate of bone

depositing. This mismatch is particularly detrimental for the regenerative potential of large

bone defects which often take more than 3 months to successfully heal [17]. After calcium

sulfate matrix is broken down, severe cases of inflammatory reaction which interfere with

new bone formation are believed to be caused by the calcium rich environment [18].

Hydroxyapatite (calcium phosphate) is another ceramic-based graft which is commonly

derived from sea coral and used clinically as a bone void filler. Unfortunately,

hydroxyapatite resorbs too slowly (time scale of years) and delays new bone ingrowth.

Ceramic-based grafts pore size was within the optimal diameter for cell infiltration, but the

overall porosity is very low [17]. There are a number of studies which show no statistical

differences between bone defects treated with the ceramic-based grafts versus autografts

[19,20] with a greater incidence of post-operative fracture [17].

The best known and successful polymer-based product, polymethyl methacrylate

(pMMA), has been used since the 1940s. Typically it is used as a bone cement and filler

6

in a number of orthopaedic applications. It functions to increase load-bearing of

reconstructed areas. As with ceramic-based products, pMMA is very dense with low

porosity thus preventing bone ingrowth [21]. In the past, exothermal polymerization of

MMA in vivo has been associated with thermal necrosis of bone and toxicity due to non-

polymerized monomers [22]. Fortunately, modern PMMA products have seen vast

improvement on in vivo exothermicity and less monomer leakage [23]. On its own PMMA,

is not bioactive and is often supplemented with hydroxyapatite, bioglass, or autologous

bone shards.

To address the disadvantages of ceramic and polymer-based bone substitutes, tissue

engineering techniques have been applied to develop a number of growth factor-based

products. Bone regeneration is controlled by a number of molecules which will be

elaborated upon further in this thesis chapter. Currently, only two commercially produced

growth factors have been FDA approved for clinical use: bone morphogenetic protein-2

(BMP-2) and bone morphogenetic protein-7 (BMP-7) (also known as osteogenic protein-

1(OP-1)). Primarily, BMPs act as cytokines mediating the differentiation of mesenchymal

cells into cartilage and bone forming cells [24]. BMP-2 and -7 (OP-1) in particular have

been identified as playing critical roles in bone formation and healing by their ability to

induce osteoblast differentiation [25]. The genetic sequence of BMP was first identified in

1988 which subsequently allowed for the commercial production of various BMPs through

the use of recombinant gene technology [26]. This marked a notable increase in the use of

BMPs in bone tissue engineering. BMP-2 and -7 are highly osteoinductive growth factors

which are incorporated into an osteoconductive biomaterial, such as collagen, in order to

7

overcome difficulties in rhBMP-2 efficacy such as a short half-life [27] and receptor-

binding antagonists [24].

Given that collagen is the main organic component of bone, it already provides the

appropriate characteristics of an osteoconductive matrix needed for the ingrowth of cells

and mineralization. Absorbable collagen sponges (ACS) and collagen particles were the

first generation of tissue engineered products on the market with FDA approval. Initial

clinical trials conducted with rhBMP carriers such as collagen for spinal fusion applications

were not only successful but outperformed autologous bone grafts [28].

Like many first generation products there have been a number of drawbacks to

rhBMP-2 collagen carriers. Positioning of the ACS is often difficult and secondary

displacement of the collagen sponge is commonly observed. Collagen is quickly degraded

in vivo, causing voids to develop within the new bone matrix due to the inability of collagen

to provide a structure which lasts long enough for cell migration [29]. The main issue of

ACS is the initial burst release of a supra-physiological dose of rhBMP-2 into the local

environment which leads to heterotrophic ossification [30]. One other side-effect to this

burst release is the activation of osteoclasts in the surrounding environment resulting in

bone resorption of adjoining healthy tissue. Surrounding mesenchymal lineage progenitor

cells in adjacent musculature receive the BMP-2 signal to differentiate into osteoblasts

which leads to mineral deposition within the muscle [31]. This burst release would not be

an issue with physiologically relevant levels of rhBMP-2, but in order to reach the

therapeutic threshold, current clinical applications use 10-1000 fold higher concentrations

than that of native BMP-2 levels found in the body [32]. For this reason, a number of

8

natural polymers are now under investigation as alternatives to ACS for rhBMP-2 delivery

and will be discussed in Chapter II.

2.4 Financial Considerations

With the growing concern over medical cost expenditures of orthopaedic treatments

and bone grafts there is an increasing concern with the growing population. Many patients

are now considering the cost-to-benefit ratio of many of these products. Allogenic bone

grafts in the form of bone chips cost around $400 USD while long bone grafts can range

from $500-$1700 USD. DBM products on average cost between $750-$1,000 USD per

treatment depending on the brand selected. For ceramic-based products, the average

calcium sulfate treatment costs $700 USD while hydroxyapatite will cost $1,500 USD.

Currently, the most expensive bone substitutes are those growth-factor based

products which deliver BMP-2 ($3,500-$5000 USD) or BMP-7 ($5000 USD) [5,33,34].

On average, the cost of treatment with BMP carriers is higher (~7%) than autografts; most

of this cost (~41%) attributed to the actual price of BMPs [35]. Although the growth-factor

based products are the most expensive initially, the effectiveness of the product and the risk

of complications leading to revision surgery must be taken into account. Future bone graft

developers must be sensitive to the final product cost when designing their treatment.

Although the cost of the bone substitute is important to keep in mind, its price is still

overshadowed by the cost of the overall procedure.

9

3. Molecular Influences of Bone Regeneration

3.1 Bone Healing Process

Bone fracture healing is a complex cascade of local and systemic events which

involve a number of regulatory factors including cytokines, hormones, extracellular matrix

proteins, and a number of cell types. Since the phases of bone healing were first described

[36], they have essentially have remained classified into the inflammatory phase, repair

phase, and remodeling phase. These phases are not clearly distinct biologically but they

merge into one another throughout the healing bone. The inflammatory phase begins

immediately after injury when the bone defect is initially filled with a fibrin-rich

hematoma. Entrapped platelets release a number of cytokines which then recruit a number

of inflammatory cells to the site of injury. Infiltration of inflammatory cells (macrophages,

monoctyes, and lymphocytes) and fibroblasts into the hematoma over time replace the

hematoma with granulation tissue and promote ingrowth of vasculature through

angiogenesis. The neo-vasculature allows for migration of mesenchymal cells into the

wound site where they begin to proliferate and synthesize a matrix of fibrous tissue

comprising of random collagen fibers. Newly differentiated osteoblasts mineralize the

unorganized collagen network forming woven bone. During the remodeling phase, the

unorganized woven bone will be replaced by a mature bone matrix. Resorption and

deposition over time restores the appropriate microarchitecture organization in response to

the mechanical stress placed on the bone [3,37]. Throughout the phases of bone healing

and formation a number of molecular cues are expressed in order to control the process.

10

3.2 Molecular Promoters of Bone Regeneration

A number of molecules have been identified in the bone healing process. Particular

interest has been placed on molecular promoters of bone regeneration which can be

incorporated into an osteoconductive biomaterial in order to enhance the osteoinductive

potential of bone substitutes. These molecular cues are expressed at different times

throughout the healing process with various effects on target cells and are divided into three

categories: 1. pro-inflammatory cytokines, 2. angiogenic factors, and 3. the transforming

growth factor beta (TGF- β) superfamily and other growth factors.

Pro-inflammatory cytokines interleukin-1 (IL-1), interleukin-6 (IL-6), and tumor

necrosis factor alpha (TNF-α) are secreted within the first 24 hours of bone damage by

macrophages which causes the initiation of the repair phase of bone regeneration. These

cytokines initiate the downstream responses which initiate the induction of both angiogenic

factors and growth factors. TNF-α activates osteoclast activity for the removal of bone

debris and promotes recruitment of mesenchymal stem cells. The expression of these pro-

inflammatory cytokines is highest during the first 24 hours of bone healing and the

cytokines are subsequently expressed in smaller quantities during repair and remodeling

phase [37].

The initial hypoxic conditions during fracture healing stimulates the demand for

angiogenesis. Initiation and successful ingrowth of vasculature is crucial for successful

bone healing [38]. Platelet-derived growth factor (PDGF) is released in the early stages

of bone healing from platelets entrapped within the hematoma and upregulates vascular

endothelial growth factor (VEGF) [39]. The expression of VEGF [40] and PDGF [41] by

pre-osteogenic cells has been shown to be a crucial component in regulating the rate of

11

neo-angiogenesis to correlate with the rate of bone formation [42]. In addition PDGF acts

as a potent mitogen of inflammatory cells and mesenchymal stem cells (MSCs) [43].

Vascular ingrowth is also regulated by fibroblast growth factor (FGF) secreted by

macrophages, mesenchymal stem cells, chondrocytes, and osteoblasts. FGF-1 and FGF-2,

being the most commonly expressed growth factors in bone regeneration, partake by

increasing callus formation and osteoblast activity [44]. Although angiogenic factors are

crucial for effective bone healing, alone they are not osteoinductive.

A number of endogenous growth factors play a role in varying stages of bone

healing, and possibly none are as important as the transforming growth factor-beta (TGF-

β) superfamily which acts upon a broad range of cells, influencing cellular activity, growth,

differentiation, and extracellular matrix production. After initial blood clot formation,

platelets release TGF-β to stimulate the proliferation of periosteal cells. Even though TGF-

β plays a role in the production of extracellular proteins for callus formation, its

osteoinductive potential is limited and does not have an effect on mineralization [45].

Insulin-like growth factors (IGF) play critical roles in skeletal development as well as

fracture healing by promoting bone matrix formation such as collagen type 1 [46]. IGF-1

is the most potent in the IGF family and is localized in healing fractures [47]. IGF-1

stimulates chemotaxis and activity of osteoblasts, and has the greatest effect on bone

formation when it is used in combination with TGF-β [48]. TGF-β’s most important roles

may lay in initiating the production of BMPs in osteoprogenitor cells while inhibiting

osteoclast activation [43].

The most important members of the TGF-β superfamily are the BMPs. BMPs are

pleiotropic regulators of growth [49], differentiation [50], and apoptosis [51] of a variety

12

of cell types. One property of BMPs is their ability to be osteoinductive by themselves

(especially BMP-2,-6,-7, and -9 [52]), which makes them attractive therapeutically for

tissue engineering products. They are strong promoters of differentiation of

osteoprogenitor cells into osteoblasts. BMPs are expressed at various points along the

phases of bone healing. BMP-2, -6, and -9 have been found to be the most potent inductors

of pluripotent MSCs to differentiate into osteoblasts. Most other BMPs act more to support

the terminal differentiation and maturation of osteocytes [53]. As previously discussed

(Chapter 1, 3.3), to date only BMP-2 and -7 have been approved for clinical use in the

United States for bone substitutes.

3.3 Molecular Inhibitors of Bone Regeneration

Molecular control over fracture healing follows developmental osteogenesis very

closely [54]. Patterning and maintenance of tissue is overseen through not only molecular

promoters, such as those discussed in the previous section, but is also greatly regulated by

molecular inhibitors. Recent interest has focused on bone regeneration inhibitors in

particular due to their mechanistic role in which negative feedback and crosstalk decrease

the cellular exposure of the molecular promoters of bone regeneration. The effectiveness

of these signaling regulators and how they can have significant negative impact on BMP

efficacy can be appreciated when considering the therapeutic dose of BMPs in bone

substitutes (10-1000x physiological concentrations [32]). BMP carriers are loaded with

supraphysiological concentrations in order to overcome the regulating factors of BMP

inhibitors to elicit a therapeutic response. These inhibitors are along these levels of the

BMP signaling cascade: 1. intracellular, 2. pseudo-receptors, and 3. extracellular.

13

Intracellular inhibitors of BMP signaling include inhibitory SMADs which are

dormant in the nucleus until BMP stimulation at which time they are released into the

cytoplasm. After BMP binds to its receptor, SMADs inhibit the signal transduction by

interacting with the BMP receptor. SMAD-6 specifically acts to inhibit BMP signaling

while SMAD-7 targets the general TGF-β superfamily [55]. Another intracellular regulator

is SMAD ubiquitin regulatory factor (SMURF), which controls intracellular BMP signal

transduction by binding and degrading various positive signaling molecules or by

degrading the BMP receptor [56].

BMP and activin membrane bound inhibitor (BAMBI) is a pseudo-receptor which

presents an extracellular domain similar to a BMP receptor domain; the difference is that

BAMBI lacks the intracellular domain. Therefore, when BMPs bind to a pseudo-receptor

it cannot form an active receptor complex in order to propagate the signal [46].

Extracellular inhibitors, mainly produced by osteoblasts, are secreted proteins that

act as BMP binding antagonists which prevent BMP from binding with its receptors. An

increase in extracellular inhibitor expression is directly correlated with an increase in local

BMP levels. The majority of the binding antagonists focus on BMPs with the strongest

osteoinductive potential (BMP-2,-6, and -9 [52]). Even though BMPs are mainly

associated with being molecular promoters of bone regeneration, BMP-3 is an antagonist

of osteogenic BMPs. Ironically, it is the most abundantly expressed BMP in adult bone

[46]. The differential screening-selected gene aberrative in neuroblastoma (DAN) family

proteins include gremlin and sclerostin. Gremlin binds and blocks BMP-2, -4, and -7.

Gremlin is responsible for inhibiting osteoblast differentiation and reducing bone

remodeling [57]. Sclerostin is produced by osteoclasts and directly competes with BMP-

14

2,-4,-6, and -7 in binding to their receptors to inhibit osteoblast differentiation and bone

remodeling. Sclerostin not only decreases MSC differentiation and osteoblast activity but

also induces apoptosis in bone cells. Follistatin neutralizes BMP-2,-4,-15, and joins with

high affinity to BMP-7 by forming a trimeric complex between itself, BMP, and the

receptor [58]. In embryogenesis, follistatin is known to inhibit all aspects of BMP activity

[59], but its role in adult bone healing is yet to be completely understood. Chordin binds

BMP-2,-4, and -7 and blocks their ability to bind to BMP receptors and acts similarly to

gremlin [60]. Noggin has the ability to bind to the greatest number of molecular promoters

of bone regeneration: BMP-2,-4,-5,-6, and -7 and prevents them from binding to BMP-2

receptors. Noggin works in a complementary fashion with gremlin to develop a local zone

which is devoid of BMP [61].

4. Conclusions

In order to address the challenges and drawbacks of current augmentation strategies

for critically-sized bone defects, tissue engineered bone substitutes were designed in mind

to be both osteoconductive (collagen carrier) and osteoinductive (rhBMP-2). Like many

first generation products, there have been a number of drawbacks to rhBMP-2 collagen

carriers such as edema and ectopic bone growth. In order to develop the next generation

of bone substitutes it is important to understand the biological action and temporal

expression during the healing cascade. Currently the focus has been placed on only

incorporating molecules which promote bone regeneration (BMPs) without acknowledging

the innate molecular controls achieved with inhibitory molecules (e.g., Noggin, Gremlin).

Therefore, in order to successfully decrease the therapeutic concentration of BMPs, the

negative feedback signaling caused by BMP antagonists must be addressed.

15

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20

CHAPTER II

CURRENT BONE TISSUE ENGINEERING STRATEGIES TO BALANCE LOCAL

THERAPEUTIC DELIVERY FROM BIOMATERIALS


21

1. Biomaterials for Bone Regeneration

Tissue engineering and regenerative medicine (TERM) is a multidisciplinary field

which encompasses material science, biology, chemistry, pharmacology, medicine and

engineering. Traditionally, the tissue engineering approach combines a triumvirate of cells

(e.g. mesenchymal stem cells), biomaterial scaffolds (e.g. polymers), and signals (e.g.

growth factors) in order to create a tissue substitute that can be implanted into a patient. In

the end, translational medicine’s goal is to bring the experience and knowledge from

“bench to bedside”[1].

TERM therapies have been centered on ex vivo cell-based therapies for decades and

the addition of cells was once thought to be the only way a product would be successful.

However, limited availability of cell sources, difficulties in ex vivo cell expansion, cost,

and regulatory hurdles have driven the focus on endogenous cell recruitment [2]. A host’s

cell population includes stem cells or progenitor cells that are actively attracted to sites of

injury [3]. In situ tissue regeneration aims to capitalize the body’s innate ability to heal by

using biomaterials and signaling molecules to recruit host stem or tissue-specific progenitor

cells to the site of injury, thereby allowing for regeneration without the need for cell

transplantation.

Great effort has been placed on developing biomimetic scaffolds which provide a

three dimensional matrix for cell migration and proliferation. Although modern

biomaterials lack the temporal and spatial complexity to fully mimic the native

extracellular matrix (ECM) [4], they capture many of the essential elements. Through their

mechanical, material, and chemical cues, biomaterials are able to influence a number of

cellular functions (e.g. cell-matrix interactions) including proliferation, differentiation,

22

apoptosis, and signaling [4]. Ideally, these biomaterials should not only be non-toxic but

biodegradable as well so that the artificial matrix will degrade over time and give way to

new tissue being formed within at the site where the material was implanted. For this

reason, this thesis is primarily concerned with degradable polymers. Hydrogel biomaterials

have been of particular interest since they mimic the ECM’s high surface to volume ratio,

can be fabricated from a number of materials, and can be loaded with a variety of

molecules.

Small molecules and soluble factors are often incorporated into a biomaterial in

order to recruit or influence the behavior of cells in and around the matrix. Growth factors,

nucleic acids, and other molecular cues can be incorporated into biomaterial scaffolds in

order to influence cellular outcomes. Ultimately, the aim of TERM is to restore tissue and

organ function.

1.1 Synthetic Materials for Bone Regeneration

Synthetic polymers can be either non-degradable or degradable in nature and have

the advantage of being able to be manufactured in large quantities and customizable to

show a wide range of material properties. A number of synthetic polymers are already

FDA-approved as scaffolds. Poly(a-hydroxy acids); poly(lactic acid) (PLA), poly(glycolic

acid) (PGA), and poly(lactic-co-glycolide) (PLGA) [5] have been used for decades in

orthopaedic applications [6,7]. These polymers undergo bulk degradation by hydrolysis of

which the rate of degradation can be tailored to the demands of the tissue. Their broken

down components, lactic acid and glycolic acid, can be processed and excreted from the

body; regrettably, these components can decrease the pH of the local environment [8].

There is a multitude of synthetic polymers (e.g. polyanhydrides, polyurethanes, etc.) but

23

they can all cause immunogenicity and toxicity due to their chemical crosslinkers and

polymerizers [9,10]. These polymers are also hydrophobic which hinders cell attachment

[11] and use as hydrogels; for that reason, other polymers such as poly(ethylene glycol)

(PEG) are added. Unfortunately, PEG reduces serum protein adsorption thereby reducing

cell adhesion to the polymer matrix [11]. Synthetic materials are biologically inert due to

a lack of cell binding domains such as Arginine:Glycine:Aspartic Acid (RGD) [12]. For

this reason a number of strategies have been developed to improve the bioactivity of

synthetic polymers such as incorporating cell binding motifs (e.g. RGD sequence) [13]. In

order to make these synthetic polymers more osteoconductive they are combined with

calcium phosphates (e.g. hydroxyapatite) [14] and natural polymers like collagen [15,16].

For synthetic polymers to achieve both osteoconductive and osteoinductive properties they

must be a combination product composed of the synthetic polymer, calcium phosphate, a

natural polymer or cell binding motif, and osteogenic growth factor [17,18].

1.2 Natural Materials for Bone Regeneration

Interest in natural polymers has been increasing due to a number of beneficial

properties such as low cytotoxicity, low immunogenicity, and similarity to the ECM. Living

organisms synthesize a number of macromolecular components which a biological

environment can recognize and degrade hydrolytically or metabolically [19]. Poly

saccharides (e.g. alginate, chitosan) and proteins (e.g. collagen, silk, fibrin, and keratin) are

extracted from renewable plant [20], algae [21], animal [20,22] or human [23,24]

resources. The majority of natural polymers are hydrophilic which make them prime

candidates for hydrogel applications. This thesis utilizes two fibrous natural polymers:

keratin and fibrin.

24

1.2.1 Keratin

Keratins are intermediate filament and intermediate filament associated proteins

which are widely found in nature and are best known as being the structural proteins

referred to as hard or soft keratins. Hard keratins can be found in wool, hair, and hooves

while soft keratins are found in skin. There are over 20 keratin proteins which form

filamentous polymers able to form intramolecular and intermolecular disulfide bonds

through cysteine residues. Wool and human hair are commonly used as sources for

biomaterials due to their ready availability, renewable source, non-toxic, ease of

sterilization, and ability to be hydrolytically degraded. According to the literature, three

groups of keratin proteins can be extracted: (1.) alpha-, (2.) beta, and (3). gamma. Alpha

(α) and gamma (γ) keratins are of particular interest for biomedical applications because of

their ability to be processed into a number of biomaterials such as electrospun fibers [25],

scaffolds [26,27], and films [28]. α-keratins have a molecular mass of approximately 60-

80kDa and serve a structural purpose due to their alpha-helical tertiary structure. γ-keratins

are high sulfur containing globular protein with molecular mass of approximately 15kDa.

One advantage of keratin compared to other natural polymers is that its material properties

can be modified. Depending on the extraction process used, the material properties of

keratin will change due to the chemical nature of cysteine residues (Figure 1)[29]. Keratin,

extracted through an oxidative extraction in which cysteine is converted into cysteic acid,

is termed keratose. The cysteic acid groups of keratose are unable to form disulfide bonds

with neighboring polymer chains and hydrogels only form through physical entanglements.

Due to this and its hydrophilic nature, biomaterials composed of keratose degrade in a

matter of weeks in vivo and in vitro [30]. Keratin extraction through a reductive process

25

in which the cysteine is capable of reforming disulfide bridges between and within

polymers without exogenous toxic cross-linkers are called kerateines. The lack of cysteic

groups cause kerateine to be less polar and mechanically more stable than keratose, thereby

making it less susceptible to hydrolytic degradation an enabling kerateine to remain in vivo

for months [31].

Even though keratin biomaterials have been in use for decades, keratin is often

overlooked as a natural polymer, even after successful applications in drug delivery [32]

and tissue engineering [33]. Keratin is a unique material for bone fracture applications

because it can be used as a traditional sponge [27,34] or injectable gel [32,35] which allows

for minimally invasive administration able to contour to any defect shape. Reconstituted

keratin spontaneously forms into a network consisting of interconnected pores with pore

diameter between 30-70µm [31,36] which is favorable for non-load-bearing bone

regeneration [37]. Due to keratin’s isoelectric point range (pH 5-6) [38] it can immobilize

polycationic molecules such as rhBMP-2 and polycationic siRNA complexes. A number of

studies have already shown keratin’s ability to achieve robust healing in injured bone as a

carrier for hydroxyapatite. [39–41] One study which used a keratin sponge as a carrier for

rhBMP-2 did not investigate how keratin material properties could be adjusted to tune

rhBMP-2 release kinetics [42]. Due to kerateine’s ability to form disulfide bonds and

remain for months in vivo [31] (time scale required for bone regeneration) we believe it

could be successfully used as a rhBMP-2 carrier for bone regeneration.

1.2.2 Fibrinogen/Fibrin

Fibrin is a fibrous protein involved in hemostasis when soluble fibrinogen (340kDa

molecular weight) is polymerized in the presence of the enzyme thrombin (prothrombin

26

activated by Ca2+), which is activated in response to injury. Orthopaedic surgeons have

long appreciated the significantly positive affect blood clots (which include fibrin) has on

bone regeneration. Even though fibrin is not a regular component of the uninjured ECM,

it serves as a temporary matrix for all wounded tissue until remodeling can being to replace

it with tissue-specific ECM. Not only does fibrin provide a structure, but it also provides a

repository for a of signaling cues to direct cell behavior [43]. Alone, fibrin induces

osteogenic differentiation [44] and angiogenesis [45]. Thrombin-mediated release of

fibrinopeptides A and B [46] attracts inflammatory cell migration to the site of injury [47]

and induces proliferation [48]. Fibrin has numerous binding sites for not only cells, but for

growth factors and other ECM components as well. Fibrin sequesters a multitude of growth

factors which are essential for bone regeneration (Chapter I) including FGF, VEGF, PDGF,

IGF, and the TGF-β superfamily including BMP-2 [49].

In the 1940s, technology was developed that allowed fibrin could be purified in

large quantities [50], thus opening the door for its use as a biomaterial. Fibrin is degraded

(fibrinolysis) into nontoxic components by the serine protease plasmin, which is initiated

by the coagulation cascade. The rate of degradation depends on the fibrin fiber thickness

and density; a fibrin network comprised of dense thin fibers degrades much more slowly

than a network of thick loose fibers [51]. Fiber thickness has also been shown to play a

large role in cell-specific signaling molecule expression [52]. Fibrin material properties

can be altered through polymerization by adjusting the concentrations of fibrinogen,

thrombin, and Ca2+/salt concentrations [53]. In general, increasing fibrinogen

concentration leads to increasingly dense plug formation. However, but care must be taken

not to make the hydrogel too dense or else it will inhibit cell migration [54]. Decreasing

27

thrombin concentration leads to increased material modulus, ultimate tensile strength, and

increasing fibrin fiber diameters [55]. NaCl concentration has an effect on hydrogel

compressive modulus and fiber diameter, which in turn affects MSC differentiation and

alkaline phosphatase expression [56]. With all of these factors in mind, fibrin biomaterials

have been used for decades for cell and biomolecule delivery [57].

Fibrin biomaterials have been successfully used in a number of TERM healing

strategies for nerve [58], muscle [59], skin [60], and bone [61]. Fibrin is FDA approved as

a wound sealant, and clinically, fibrin sealants have been mixed with hydroxyapatite for

craniofacial applications [62]. Isoelectric points of fibrinogen (pH 5.5) and fibrin (pH 5.6)

[63] would cause fibrin to have negative surface charge at the neutral pH, which would

allow for rhBMP-2 “entrapment” and adsorption of nucleic acid complexes [64].

2. Controlled Delivery Strategies for BMP-2

One of the main benefits of hydrogels is that bioactive molecules and therapeutics

can be easily incorporated or physically entrapped into the network. In these

circumstances, the therapeutic agent is transported through the hydrogel and is released by

diffusion and/or hydrogel degradation. Release rates of biomolecules are driven by a

concentration gradient that is influenced by interactions between the molecule and

hydrogel matrix. In cases where there is no or little interaction between the therapeutic

agent and the hydrogel network, an early rapid burst release is followed by tapering into

smaller amounts at later time points (as the concentration lowers) [65]. This profile is

typical of the relationship between absorbable collagen sponge and rhBMP-2. In order to

limit the initial burst release and control the rate of rhBMP-2 release from the hydrogel

28

network a number of strategies have been investigated including encapsulation,

immobilization, and controlled carrier degradation.

rhBMP-2 can be encapsulated via micro, macro, and nanoparticles alone [66] or

within a scaffold [67]. In this way, rhBMP-2 release is regulated by the rate of particle

degradation. When rhBMP-2 microspheres are entrapped in a hydrogel network they retain

rhBMP-2 for significantly longer periods of time (compared to adsorption) resulting in

prolonged bioactivity [68]. Synthetic polymers are commonly used for this purpose due to

their predictable rates of degradation and release [69,70]. Natural polymers such as

chitosan [17,71,72] , alginate [73], and gelatin [74] are being increasingly used for rhBMP-

2 microsphere encapsulation.

Another method for controlling rhBMP-2 presentation to cells is through direct

immobilization of rhBMP-2 to the carrier system. Temporal control of growth factor

release can be achieved with biomaterials through protease degradable tethers [75];

mimicking the ECM’s native role in BMP-2 delivery. Other approaches such as chemically-

conjugating BMP-2 directly to a polymer matrix, have been shown to maintain BMP-2

activity for extended periods of time [76,77]. One other approach is through

electrospinning, where rhBMP-2 is directly embedded into polymer fibers for long-term

delivery [78].

One important design parameter to regulate carrier degradation as well as rhBMP-

2 release is to modify the mesh size of the polymer network through the crosslink density.

When the overall crosslink density of a hydrogel network is increased, it decreases the

molecular distance between crosslinks, resulting in a smaller hydrogel mesh size. This in

turn lowers the diffusivity of the therapeutic agent by increasing the interactions between

29

the growth factor and the hydrogel network. In this manner tunable rates of release can be

achieved by varying the crosslink density of a hydrogel [65]. This is one approach many

hydrogel systems utilize since it is the simplest method for successful encapsulation of

rhBMP-2. Examination of key parameters has allowed elucidation of these parameters

what allow for near zero-order release to be achieved [79]. Another method involves

increasing BMP-2 affinity for the carrier system through succinylation [80], alkylation

[81], or use of glycosaminoglycan conjugates like heparin [82–84]. A number of strategies

like our own are simply investigating protein alternatives to ACS.

3. Controlled Delivery Strategies for siRNA

A major challenge in regenerative medicine is the means to promote and direct cells

functions such as proliferation, differentiation, and migration in engineered constructs.

Gene therapy offers a means to introduce exogenous genetic material into cells in order to

modify expression of desired proteins. Small interfering RNA (siRNA) is a relatively new

gene-silencing mechanism by which post-transcriptional gene silencing can occur [85].

Delivery of siRNA alone is not successful due to its susceptibility to degradation and

overall negative charge which prevents siRNA from passing through the cell membrane.

To overcome these challenges and allow siRNA to be effective in the cytoplasm of targeted

cells, siRNA can be packaged into viral or non-viral vectors. Although viral vectors have

greater transfection efficiency than non-viral vectors, there are regulatory challenges to

viral vectors which impede their translation into clinical applications. Non-viral vectors

such as liposomes are simple packages which are composed of cationic compounds that

sequester siRNA [86]. Both cationic polymers such as polyethylenimine (PEI)[87] and

cationic liposome/lipid-based systems such as DOTMA [88] are able to electrostatically

30

package siRNA. We refer to polymer-based complexes of siRNA as polyplexes and lipid-

based complexes of siRNA as lipoplexes. There has been great success with solution-phase

transfection for targeting sequence-specific knockdown has led to increased understanding

of development [89] and signaling pathways [90,91].

Krebs, MD et al. showed for the first time the ability of a three dimensional

hydrogel to locally retain and deliver siRNA in a sustainable fashion [92]. Since this work

was published, substrate-mediated deliver of siRNA has been applied to a number of

TERM applications including skin [93] and nerve [94]. Local delivery strategies now

involve layer-by-layer [95], nanoparticle embedding [96], and direct incorporation into a

cationic hydrogel [97].

Recently, there has been interest in targeting inhibitory molecules along their

signaling pathway to aid in bone regeneration. Primary, focus has been on co-delivery of

inductive molecules such as BMP-2 and siRNA targeting suppressive molecules. Since

substrate-mediated siRNA delivery is still a relatively (<10 years) new approach many

researchers “pre-treat” their cells by the classic means of solution-phase transfection before

seeding the transfected cells onto the synthetic bone matrix. However, the overall take-

home message is the same: reducing the expression of osteogenic inhibitors enhances the

action of pro-regenerative molecules. Targeting intracellular regulators of BMP-2, such as

Smurf1 [98], resulted in improved osteogenic activity of MSCs. “Pre-treatment” with

noggin siRNA has also led to significantly increased expression of osteogenic markers in

in vitro, and of bone regeneration in vivo [99]. Substrate-mediated siRNA delivery for bone

application has just begun to make an impact on the scientific community. The use of

chitosan hydrogels as a reservoir for siRNA delivery has shown successful down regulation

31

of osteoclast activity in vitro [97]. Substrate-mediated delivery of noggin siRNA from a

synthetic polymer has successfully enhanced osteogenic activity in vitro [100]. Delivery of

noggin siRNA from a natural polymer hydrogel has yet to be studied. Currently, there is a

need to locally regulate BMP-2 antagonist (such as noggin) activity within the bone healing

microenvironment after BMP-2 carrier treatment.

4. Thesis Overview

This dissertation work’s emphasized the ability of natural polymers to be modified

in order to deliver molecular cues to balance local effects of pro-growth and anti-inhibitory

molecules for bone regeneration. Kerateine was investigated to evaluate its potential as an

alternative rhBMP-2 carrier (Chapter III). This work was important for understanding the

unique relationship between keratin material properties and rhBMP-2 release by

modulating the hydrogel weight to volume percentage or sub-fraction (α and γ) ratio to

develop an injectable rhBMP-2 carrier. Two kerateine formulations were evaluated as

rhBMP-2 carriers and compared to ACS as an injectable gel or implantable scaffold. These

rhBMP-2 carriers (kerateine gel, kerateine scaffold, or ACS) were then used to treat a

critically-sized rat mandibular defect to test the ability of rhBMP-2 kerateine carriers to

regenerate bone (Chapter IV). These findings demonstrate the ability to tailor natural

polymers through their innate characteristics and may be broadly applicable to a number

of different wound healing applications besides bone regeneration.

Emphasis has traditionally been placed on growth-supporting molecular cues in

TERM products; including this dissertation’s work (Chapter III and IV). Even though

regeneration is successful, it is hindered by the underlining inhibitory molecule. To support

regeneration we explored the opportunity of removing locally expressed inhibitory

32

molecules, through substrate-delivery of siRNA. Peripheral nerve injuries are one system

in which a continuously expressed protein, myelin-associated protein (MAG), inhibits

regeneration after injury. A proof-of-principle study was first conducted targeting an

“inhibitory” molecule which is continuously expressed in order to evaluate the feasibility

of delivering siRNA complexes from a fibrin surface (Chapter V). We applied these

findings to the field of bone regeneration in which the expression of inhibitory molecules

(noggin) was dependent on the expression levels of molecular promotors (rhBMP-2)

(Chapter VI). Taken together these findings show novel approaches to lower the rhBMP-

2 dose in carriers by either controlling the rate of rhBMP-2 release or by increasing rhBMP-

2 efficacy by down-regulating rhBMP-2 antagonists.

The work discussed in this dissertation showcases the ability of natural polymers to

locally deliver pro-regenerative therapeutics to augment bone regeneration. These findings

support the idea that molecular promoters and inhibitors must be balanced in order to

achieve the greatest amount of tissue regeneration with the lowest amount of adverse side

effects. Understanding the natural molecular pattern of development and wound healing in

order to incorporate the information from basic science to the clinical theater will improve

the next generation of TERM products.

33

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41

CHAPTER III

REDUCTION OF ECTOPIC BONE GROWTH IN CRITICALLY-SIZED RAT

MANDIBLE DEFECTS BY DELIVERY OF RHBMP-2 FROM KERATEINE

BIOMATERIALS.

Christine J. Kowalczewski, Seth Tombyln, David C. Wasnick, Michael R. Hughes, Mary

D. Ellenburg, Michael F. Callahan, Thomas L. Smith, Mark E. Van Dyke, Luke R.

Burnett, Justin M. Saul

Manuscript was written and prepared by Christine J. Kowalczewski. Justin M. Saul served

in an advisory capacity and all authors served in an editing capacity. Text and figure

formatting variations were due to journal guidelines. This work was published in

Biomaterials. 2014;35:3220–8.

42

______________________________________________________________________

Abstract

Absorbable collagen sponges (ACS) are used clinically as carriers of recombinant

human bone morphogenetic protein 2 (rhBMP-2) to promote bone regeneration. ACS

exhibit ectopic bone growth due to delivery of supraphysiological levels of rhBMP-2,

which is particularly problematic in craniofacial bone injuries for both functional and

aesthetic reasons. We hypothesized that hydrogels from the reduced form of keratin

proteins (kerateine) would serve as a suitable alternative to ACS carriers of rhBMP-2. The

rationale for this hypothesis is that keratin biomaterials degrade slowly in vivo, have

modifiable material properties, and have demonstrated capacity to deliver therapeutic

agents. We investigated kerateine hydrogels and freeze-dried scaffolds as rhBMP-2

carriers in a critically-sized rat mandibular defect model. ACS, kerateine hydrogels, and

kerateine scaffolds loaded with rhBMP-2 achieved bridging in animals by 8 weeks as

indicated by micro-computed tomography. Kerateine scaffolds achieved statistically

increased bone mineral density compared to ACS and kerateine hydrogels, with levels

reaching those of native bone. Importantly, both kerateine hydrogels and kerateine

scaffolds had significantly less ectopic bone growth than ACS sponges at both 8 and 16

weeks post-operatively. These studies demonstrate the suitability of keratins as rhBMP-2

carriers due to equal regenerative capacity with reduced ectopic growth compared to ACS.

KEYWORDS: bone regeneration, collagen, hydrogel, scaffold, in vivo test, keratin

43

1. Introduction

High-energy craniofacial trauma are seen in 75% motor vehicle accidents [1] and

29% of battlefield injuries from improvised explosive devices [2]. The resulting injuries

are irregularly shaped and affect a number of tissues including muscle, nerve, and bone [3].

Irregularly shaped craniofacial bone injuries are often too large to heal spontaneously

without external intervention (i.e., they are critically-sized). Over the past few years the

number of maxillofacial reconstructive procedures has dramatically increased; with over

200,000 procedures performed in 2012 alone [4]. Complications arising from the use of

auto- and allogeneic bone grafts as well as the difficulty of their use in critically-sized bone

defects in the head, face, and jaw have led to increased clinical interest in the use of bone

substitutes [5].

Bone substitutes often incorporate potent exogenous agents such as recombinant

human bone morphogenetic protein 2 (rhBMP-2) to recruit marrow and periosteal pre-

osteoblasts to differentiate and bridge the defect site [6]. Exogenous delivery of rhBMP-2

requires a carrier system to overcome a short systemic half-life and prevent rapid diffusion

of the rhBMP-2 away from the implant (injury) site. FDA-approved collagen-based

rhBMP-2 carriers such as Medtronic’s Infuse ® have been successful in restricted

applications, including maxillofacial bone grafts [7]. Unfortunately, the large dose of

rhBMP-2 required to elicit a therapeutic response causes an over activation of osteogenic

differentiation near the treatment area due to the effects of rhBMP-2 on preosteogenic cells

of mesenchymal lineage found in the neighboring muscle and periosteal [8,9]. Further, the

supraphysiological release of rhBMP-2 caused by the inability to retain the rhBMP-2 and

the enzymatic degradation of the collagen sponge results in side effects in the surrounding

44

area including ectopic bone growth, bone resorption, and edema [10,11]. Due to these

clinically observed adverse events and the irregular geometry of craniofacial bone defects,

it is clear that alternative carrier systems for rhBMP-2 are needed. Natural polymers are

of particular interest as rhBMP-2 carriers because they are often bioresorbable, elicit

limited immune response, have low cytotoxicity, and unlike synthetic polymers they can

be cleared and metabolized by the host with minimal inflammatory response [12].

Keratins are intermediate filament proteins widely found in nature and are best

known as being the structural proteins found in wool, hooves, and hair. Keratins are

commonly extracted from wool and human hair because these are a renewable source for

biomaterials. Although there are a number of keratinases produced by bacteria [13] and

fungi [14], keratinase enzymes have not been identified in humans, making keratin

biomaterials attractive for long-term drug delivery due to slow in vivo resorption. Keratins

are also attractive as biomaterials and as carriers of rhBMP-2 because they are easily

sterilized, are hydrolytically degraded, and have material properties that can be modified

[15–17]. Modification of keratin’s material properties is dependent on the method of

extraction, the two general methods of extraction: oxidative or reductive. Each respective

method leads to different material properties of the resulting processed biomaterials due to

differences in the chemical nature of the cysteine residues that result from these extraction

chemistries [18,19]. The oxidative form, keratose, lacks disulfide cross-links due to

sulfonic acid residues on cysteine [16]. This results in a material that rapidly degrades;

therefore, keratose was not used in the studies described in this manuscript. Kerateine, the

form of keratin resulting from reductive extraction (see Figure 1, step 1), acquires material

properties from both physical entanglements and the formation of covalent disulfide bonds

45

between cysteine residues (i.e., cystine), which results in a stable, cross-linked hydrogel

[17]. Extracted keratin proteins (both keratose and kerateine) can be further processed to

obtain two sub-fractions: alpha (α) keratin and gamma (γ) keratin. α-keratin is the key

structural component due to its alpha-helical tertiary structure, and γ-keratin is a high sulfur

containing globular protein (see Figure 1, step 2) [20].

In the work reported here, we explored the use of kerateine (reductive extraction)

as a carrier for rhBMP-2 as an alternative to absorbable collagen sponges for application

to craniofacial defects. Our rationale was that the presence of disulfide bonds in kerateine

(reductive form of keratin) would lead to in vivo stability on the order of months, which is

a timescale appropriate for bone regeneration. Furthermore, we have previously observed,

and confirmed in this report, a correlation between the rate of hydrogel degradation and the

rate of therapeutic agent delivery [21,22]. For these reasons, we hypothesized that kerateine

biomaterials could be formulated to support controlled delivery of rhBMP-2 that would

improve measures of bone regeneration (bridging and bone mineral density) while reducing

negative side effects (ectopic bone growth) associated with current clinically-used

absorbable collagen sponges. We specifically investigated two types of kerateine carriers:

(1) an injectable hydrogel and (2) an implantable (freeze-dried hydrogel) scaffold that was

fabricated to the geometry of the defect. After initial characterization of mechanical,

scaffold/gel degradation, and rhBMP-2 release, we then applied the kerateine carriers to a

critically-sized rat mandibular defect model to assess measures of healing including bone

regeneration and ectopic bone growth. We compared the kerateine biomaterials directly to

the current clinical system of rhBMP-2 delivery from absorbable collagen sponges.

46

2. Materials and Methods

2.1 rhBMP-2 Carrier Preparation

Lyophilized and sterile (via 2 MRad gamma-irradiation) kerateine powder (see

Figure 1, Step 3) was obtained from KeraNetics, Winston-Salem, NC. Under sterile

conditions, a 6% (weight/volume) kerateine powder at an 80:20 ratio of α: γ kerateine

(which we refer to as 6% 80:20 kerateine) was reconstituted in water with or without

100µg/mL rhBMP-2 (Pfizer, New York, NY) (see Figure 1, step 4). The mixture was then

vortexed, centrifuged, and incubated overnight at 37˚C to form gels. Kerateine scaffolds

were formed by freeze-drying 5mm circular biopsy punches of hydrogels of a thickness of

2.5mm (see Figure 1, step 5). Based on mass balance, 5µg of rhBMP-2 was present in each

kerateine gel or scaffold used for in vitro and in vivo experiments. A 5mm diameter

absorbable collagen sponge (ACS) was obtained from a XX-small Infuse kit (Medtronic,

Minneapolis, MN) by biopsy punch. 5 g total rhBMP-2 was loaded onto each ACS. This

is consistent with manufacturer’s recommended method of preparation with the exception

of a reduced rhBMP-2 dosage and scaffold size.

2.2 Scanning Electron Microscopy

Lyophilized kerateine scaffolds were gold sputter coated (Denton Desk II Cold

Sputter Coater, Moorestown, NJ) and imaged at 5,000x magnification with a Zeiss Supra

35VP FEG SEM (Oberkochen, Germany).

2.3 Rheology

A 6% 80:20 kerateine powder in water without rhBMP-2 was vortexed, centrifuged,

loaded into a 22 x 1.2 mm cylindrical teflon template, and then incubated overnight at 37˚C

to make a homogenous gel (n=3). Samples were then loaded onto a Bohlin CS-10

Rotational Rheometer (Malvern, Worcestershire, UK) system with parallel plate geometry

47

(gap width = 1mm) for assessment of gel rheological characteristics at room temperature

(25˚C). A dynamic oscillation stress sweep at a constant frequency of 1 Hz was used to

determine the linear viscoelastic region (LVER) of the kerateine biomaterial. A frequency

sweep from 0.1-10 Hz was then performed at a constant stress (25 kPa) within the LVER.

2.4 In Vitro rhBMP-2 Carrier Characterization

Release of rhBMP-2 from each carrier was determined in vitro over the course of 4

weeks. rhBMP-2 carriers (kerateine gel, kerateine scaffold, or collagen sponge) loaded

with 5g rhBMP-2 were prepared as described above and placed in a 1.5mL conical tube.

Data show the results of a single experiment (n=3) that was conducted twice. Carriers

without rhBMP-2 served as negative controls. 100μL of sterile PBS (Gibco, Grand Island,

NY) was added to each conical tube. Tubes were then incubated at 37°C. At pre-assigned

time points the PBS was removed and replaced with fresh 100µL PBS. Collected samples

were frozen at -80˚C until analysis was performed by DC Protein Assay (Bio-Rad,

Hercules, CA) for carrier degradation and rhBMP-2 ELISA (Peprotech, Rocky Hill, NJ)

for rhBMP-2 release.

2.5 Critically-Sized Rat Mandible Defect Model

All animal studies were conducted with the approval and guidance of the Miami

University Institutional Animal Care and Use Committee (IACUC). 350g male Sprague

Dawley rats (Harlan, Indianapolis, IN) that were approximately 10 weeks old were

anesthetized with isoflurane. Under aseptic conditions, an incision was made through the

masseter muscle, which was retracted to expose the bone of the masseteric fossa. A full

thickness 5mm diameter circular critically-sized defect was created with a sterile 2.4 mm

circular engraving cutter (Dremel, Racine, WI) with intermittent rinses of PBS (Gibco,

48

Grand Island, NY). The defect was then treated with carriers either containing or lacking

5g rhBMP-2. The ACS and kerateine scaffolds were placed directly into the surgical

defect site while kerateine hydrogels were injected from a sterile 1mL syringe. After

application of treatment groups, the masseter muscle held the treatment in place and was

closed with resorbable POLYSORB suture (Syneture, Norwalk, CT). The skin was closed

with nonresorbable SURGIPRO suture (Syneture, Norwalk, CT) and was removed one

week later. Control animals receiving “no treatment” were closed without implanting or

injecting materials to the mandibular defect. Five animals with bilateral defects (10 total

defects) were used per treatment group. Five animals that did not undergo the surgical

procedure served as controls for normal, age-matched bone.

2.6 µCT Image Analysis of Bone Regeneration and Ectopic Growth

Non-invasive assessment of in vivo bone regeneration was determined by 52 micron

resolution (voxel size 0.00014mm3) micro-Computed Tomography (µCT) (Imtek

MicroCAT II, Siemens, USA) in the Imaging Research Center at the Cincinnati Children’s

Hospital Medical Center at 8 and 16 weeks post operatively. µCT images were

reconstructed in Osirix DICOM-viewer (Pixmeo, Geneva, Switzerland) with 3D surface

rendering (Osirix threshold for bone = 625 pixel). When un-bridged defect or ectopic

growth were observed, Regions of Interest (ROI) were drawn around respective boundaries

on every tenth image and interpolated to obtain volume renderings of the defect or ectopic

growth. Percent bone regeneration was calculated by normalizing the volume of the un-

bridged defect ROI by the original defect volume. Bone mineral density of treatment area

was calculated from known grey scale standards run with each -CT scan and determined

by greyscale analysis in ImageJ (NIH, USA). Bone mineral density of regenerated tissue

49

within the defect area was normalized to the bone mineral density of controls of normal,

age-matched bone of the masseteric fossa.

2.7 Tissue Processing and Staining

After the 16 week µCT scan, rats were humanely euthanized. The mandibles were

immediately harvested and hemi-mandibles separated. Tissue was fixed for three days in

10% neutral buffer formalin (Shandon Formal Fix, Thermo Scientific, Waltham, MA),

decalcified for three days in formic acid (Immunocal, Decal Chemical Corporation,

Tallman, NY), and decalfication stopped with Cal-Arrest (Decal Chemical Corporation,

Tallman, NY). Tissue was then soaked in PBS for 3 days, placed in 30% sucrose overnight,

and embedded in paraffin. Tissue was sectioned in the coronal plane by microtome

(Microm HM 355s, Thermo Scientific, Florence, KY) into 10µm sections on positively

charged slides (Leica, DE). After drying overnight at 60°C, slides were stained with

Masson’s Trichrome (American Master Tech, Lodi, CA). Slides were imaged 4x objective

on an AX70 Olympus (Center Valley, PA) microscope and images captured with a Nikon

D300 (Tokyo, Japan). Due to the size of the defect, multiple images of each respective

defect were combined to form panoramic images by using Adobe Elements (Adobe

Systems, San Jose, CA).

2.8 Statistical Analysis

In vitro data was analyzed by using one-way analysis of variance (ANOVA) with

Tukey’s post-hoc comparisons. In vivo analysis of bone regeneration, bone mineral

density, and ectopic bone growth was performed by using mixed model ANOVA with

planned post-hoc comparison using Bonferroni correction. An arcsine transformation was

applied to the percent bone regeneration prior to model fitting to stabilize variance.

50

Statistically significant p-values are presented in both the text and figure legends. Analyses

were performed by using Minitab version 16 (Minitab, State College, PA) or SAS® version

9.2 for Windows (SAS Institute, Cary, NC).

3. Results


Several kerateine formulations were tested for rheology and degradation profiles to

determine a material that would have suitable mechanical and degradation properties (data

not shown). We report on the final formulation selected, which was 6 % kerateine (w/v) for

a hydrogel with an 80:20 ratio of α:γ. Figure 1B shows an SEM image of the resulting

freeze-dried scaffold material for this formulation, demonstrating the porous nature of the

materials.

51

Figure 1. Schematic of extraction method and fabrication of kerateine biomaterials.

(A) Reductive extraction of human hair results in the reduced form of keratin known as

kerateine. Further purification separates α and γ fractions of the kerateine extracts which

can be freeze-dried and reconstituted to form a number of biomaterials including injectable

hydrogels, which can be further processed into freeze-dried scaffolds (SEM images of hair

reproduced with permission from de Guzman et al.[16]). (B) FE-SEM internal structure

of a kerateine scaffold at 250x magnification.

52

3.2 Rheology

Kerateine hydrogel physical characteristics were determined by oscillatory

rheology within the LVER. Kerateine material properties can be modified by either weight

percent or the ratio of alpha to gamma kerateine in the material formulation. At a constant

shear stress (τ = 25 Pa) a frequency sweep from 0.1 – 10 Hz shows little effect on the

complex modulus of the hydrogel (Figure 2). The elastic modulus (G’) is greater than the

viscous modulus (G”), the phase angle (˚) is close to 0˚, and tan δ is less than 1 along the

frequency sweep. These findings are indicative of formation of a stable cross-linked

hydrogel network showing characteristics of an elastic solid gel.

Figure 2. Rheological characterization of kerateine hydrogel. 6% 80:20 kerateine

hydrogel in the LVER acting with the typical behavior of a stable hydrogel network where

G’ and G” are independent of frequency and tan δ is close to 0 (Experimental n=3, Mean ±

SD).

53

3.3 In Vitro rhBMP-2 Carrier Characterization

Analysis of carrier degradation showed that both the kerateine gel and scaffold at

the initial time point (1.5 hours of incubation) had 10.0% of the total protein erode from

each of the carriers (Figure 3A). At the end of the four week period the kerateine gel had

the greatest cumulative erosion (32.4 ± 3.2%) followed by the kerateine scaffold (23.7 ±

1.9%) while the ACS had the least cumulative erosion (17.1 ± 1.2%). rhBMP-2 release was

measured over a four week period to compare kerateine gels, kerateine scaffolds, and ACS

carriers. To observe if rhBMP-2 kerateine carriers have an initial burst release like ACS, a

rhBMP-2 release kinetic study was performed (Figure 3B). Linear regression over initial

(linear) release period of two days showed that ACS had a cumulative release of 6.8% of

total rhBMP-2 per day whereas kerateine scaffolds released 3.0% of rhBMP-2 per day and

kerateiene gels released 2.3% of rhBMP-2 per day. The initial rate of rhBMP-2 release

from kerateine gels and scaffolds demonstrated 66.0% and 55.0% lower rhBMP-2 release

rates, respectively, compared to ACS when measuring the linear regression over the first

two days. The kerateine gel had a 25.0% lower rate of rhBMP-2 release compared to the

kerateine scaffold. The greatest total rhBMP-2 release over the total 4 week period was

from the ACS (32.1 ± 2.6%) followed by the kerateine scaffolds (15.9 ± 1.2%). If the

initial 6.0% burst release at the first time point (1.5 hours) after rehydration of the kerateine

scaffolds is subtracted, then the kerateine scaffolds show a release profile similar to

kerateine gels (8.0 ± 0.7%).

54

Figure 3. In vitro rhBMP-2 carrier characterization. (A) Percentage of cumulative

carrier erosion and (B) rhBMP-2 release from kerateine gel, kerateine scaffold, or ACS

loaded with 5g of rhBMP-2 over a period of 4 weeks. Pairwise statistical comparisons

were made between each group. (Experimental n=3 Mean ± SD; * p<0.05 kerateine gel vs

kerateine scaffold, ** p<0.05 for both kerateine carriers vs ACS.)

3.4 Bone Regeneration and Ectopic Bone Growth Analysis

µCT images of bone regeneration sixteen weeks after surgical creation of bilateral

5mm critically-sized circular mandibular defects (Figure 4A) were compared to µCT

reconstruction of normal mandibular bone (Figure 4B). 3D renderings of μCT images

indicate that untreated rat mandibles (Figure 4C) as well as those treated with kerateine

gels (Figure 4E) or scaffolds (Figure 4G) without rhBMP-2 did not spontaneously heal by

16 weeks, highlighting the critical-sized nature of the defect. In contrast, 3D reconstruction

of week 16 week µCT images demonstrates the ability of ACS (Figure 4D), injectable

kerateine gels (Figure 4F) and scaffolds (Figure 4H) loaded with rhBMP-2 to elicit

complete bridging of the defect.

55

Figure 4. µCT images of bone regeneration in critically-sized rat mandibular defect. (A) Schematic of initial 5mm defect size and location imposed on a sketch of a normal rat

mandible. (B-H) Representative µCT images of bone regeneration sixteen weeks after

surgical creation of a bilateral 5mm critically-sized circular mandibular defect in (B)

normal, uninjured, age-matched mandible; (C) rats receiving no treatment, (E) kerateine

gel without rhBMP-2 or (G) kerateine scaffold without rhBMP-2 were unable to bridge the

defect (arrows). Administration of (D) ACS with 5 g rhBMP-2, (F) kerateine gel with 5

g rhBMP-2, or (H) kerateine scaffold with 5µg rhBMP-2 were able to close the defect

with varying degrees of undesired ectopic bone growth (arrowheads).

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Measurements of defect volumes show that, at 8 and 16 weeks, groups not treated

with rhBMP-2 resulted in statistically less bone regeneration compared to rhBMP-2

treated groups (Figure 5A). Interestingly, kerateine gels alone (without rhBMP-2) were

able to achieve statistically more bone regeneration than no treatment and scaffold only

after 16 weeks. rhBMP-2 carriers (ACS, kerateine gel, and kerateine scaffold) were

statistically equivalent to one another and were able to bridge the defect gap after 8

weeks (Figure 5A, 8 week data).

Figure 5. Quantification of percent bone regeneration and bone mineral density. (A)

Bone regeneration was normalized to initial defect volume with complete defect bridging

represented by 100%. (B) Regenerated bone mineral density of the defect area was

normalized to normal uninjured bone at the same location. (n=5 animals with bilateral

treatment averaging, Mean ± SD; *p<0.0001 vs No Treatment, #p<0.05 vs ACS,

@p<0.0001 vs Normal Bone)

Differentiation between cartilaginous and bone tissue was determined by

thresholding the CT images as well as measuring the bone mineral density (BMD) within

the defect area (Figure 5B). At 8 and 16 weeks, treatment groups without rhBMP-2 had

statistically lower BMD than normal bone and rhBMP-2 carrier groups. Kerateine rhBMP-

2 loaded gels performed statistically as well as the ACS carriers at 8 weeks and had

57

statistically greater BMD by 16 weeks. Kerateine scaffolds loaded with rhBMP-2 achieved

statistically greater BMD than ACS at both time points. More importantly, the kerateine

scaffold rhBMP-2 carrier was the only treatment group to achieve BMD values that were

statistically equivalent to normal, uninjured, age-matched bone.

Reconstruction of DICOM images revealed formation of ectopic bone growth in

all experimental groups containing rhBMP-2, as shown by arrowheads in Figures 4D, 4F,

and 4H. Groups not treated with rhBMP-2 (no treatment Figure 4C; kerateine gel Figure

4E; and kerateine scaffold Figure 4G) showed no evidence of ectopic growth.

Quantification of ectopic growth in rhBMP-2 treated animals was measured by using

Osirix 3D volume rendering. As can be observed in Figures 4D, 4F, and 4H and

quantified in Figure 6, kerateine gels and scaffolds used as carriers for rhBMP-2 had

significantly less ectopic growth than ACS at the same rhBMP-2 dose at both 8 and 16

week measurements.

Figure 6. Quantification of ectopic bone formation. Ectopic bone growth volume for

rhBMP-2 carrier system as determined by volumetric analysis of CT images. (n=5

animals with bilateral treatment averaging; Mean ± SD; #p<0.0001 vs. ACS)

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3.5 Histology

Histological evaluation of experimental treatment groups was performed in order

to evaluate the regenerated tissue within the defect site (Figure 7). Masson’s trichrome

stains mature bone matrix and endogenous keratin (intracellular not from implanted

materials) red while fibrous tissue, osteoid, and collagenous tissue stain blue. Differences

in observed defect area (dashed and solid lines) are a result of taking images at different

areas along the defect site. No treatment (Figure 7B), kerateine scaffold (no rhBMP-2;

Figure 7C) and kerateine gel (no rhBMP-2; Figure 7D) showed clear defect boundaries

with infiltration of soft collagen matrix (dashed line). Kerateine, which was not fully

degraded and remains brown in color, can be seen in the middle of the defect area in both

kerateine gel and scaffolds not loaded with rhBMP-2 (Figure C and D inserts). All rhBMP-

2 carrier groups (ACS, Figure 7F; kerateine scaffold, Figure 7G; and kerateine gel, Figure

7H) showed bridging of the defect site with newly formed trabecular (T), woven (W), and

mature (M) bone.

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Figure 7. Histological images of bone regeneration. Histological images of bone

regeneration in section of rat hemi-mandibles 16 weeks post-surgery. Mandibles were

sectioned along the sagittal plane of the decalcified defect area and stained with Masson’s

Trichrome. (A) Normal bone and (E) a schematic of the anatomical orientation. Animals

administered (B) no treatment, (C) kerateine scaffold, or (D) kerateine gel without rhBMP-

2 were unable to bridge the defect (indicated by dashed line). (F) Absorbable Collagen

Sponge, (G) kerateine scaffold, or (H) kerateine gel loaded with 5µg rhBMP-2 were able

to close the defect (solid line) with mature (M), woven (W) and trabecular (T) bone. Each

main image is stitched from images collected with a 4x objective; scale bar denotes 2mm.

For B, C, D, F, G, and H inserts are images taken with 20x objective (scale bar indicates

0.2mm). Inserts for no treatment (B) and kerateine carriers without rhBMP-2 (C &D)

highlight the collagen fiber orientation as well as remaining kerateine carrier after 16 weeks

(arrows). Inserts for rhBMP-2 treatment groups highlight varying stages of bone

formation.

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4. Discussion

There is a lack of suitable rhBMP-2 biomaterial carriers for the treatment of bone

defects in the head, face, and jaw. The goal of these studies was to investigate the suitability

of kerateine hydrogels or scaffolds as alternative carriers for rhBMP-2 in craniofacial or

mandibular bone defects. The primary functions of BMP-2 carriers are (1) to increase the

efficacy of rhBMP-2 by preventing rapid diffusion away from the injury site while

promoting a chemoattractive gradient and (2) to provide a matrix for migrating cells.

From the in vitro rhBMP-2 release studies, even though the ACS does not

hydrolytically erode as quickly as the kerateine carriers, it does have a greater release of

rhBMP-2. There is a strong positive correlation between high protein isoelectric points

(rhBMP-2 pI ~ 8.5) and retention times within a carrier system [10,23] with mild acidic

conditions [24]. Because keratin has an isoelectric range less than neutral (pH 5-6, where

range indicates that multiple keratin protein are present) [19], it can immobilize molecules

with opposite charge (at neutral pH) such as rhBMP-2. This may explain why the rhBMP-

2 release plateaus after five days compared to collagen (pI ~8.26) [25], which continues a

steady release of rhBMP-2 in vitro. Unlike synthetic polymers which are homogenous,

naturally derived polymers such as collagen and kerateine are a heterogeneous mixture of

protein fractions with a range of molecular weights and isoelectric points [17,26].

Kerateine’s interaction with other proteins is also dependent on the amounts of kerateine

sub-fractions (α- and γ-kerateine) and keratin associated proteins. After the initial release

of soluble rhBMP-2 was observed for both the kerateine gel and scaffold, the release profile

plateaued after the third day. This result suggests that the remainder of the rhBMP-2

remains within the kerateine carrier matrix. While rhBMP-2 release from the kerateine

61

carriers plateaus, rhBMP-2 released from the ACS continues to release without plateauing

during the course of our studies. These differences could be the result of the heterogeneous

nature of collagen and kerateine [17,26]. Smaller molecular weight polymer fractions of

both collagen and kerateine degrade more quickly than the larger molecular weight (more

hydrolytically stable) polymer chains; thereby, a relatively more homogenous protein

mixture remains. This could be a potential explanation for rhBMP-2 release mirroring

kerateine degradation which both plateau after three days. After three days, the remaining

kerateine could have a lower isoelectric point (than the initial heterogeneous mixture)

resulting in an overall more negatively charged carrier which interacts more favorably with

rhBMP-2, therefore increasing kerateine’s potential to sequester rhBMP-2. On the other

hand, as collagen degrades it also becomes more homogeneous possibly resulting in a more

over all positive charge which would repel rhBMP-2. This would explain why rhBMP-2

release from the ACS did not plateau during the course of our in vitro study. Differences

in degradation and BMP-2 release seen between kerateine gel and scaffold could be

associated with the lyophilization process to fabricate the kerateine scaffolds which has

been observed previously causing changes of swelling ratio and degradation [27]. It is

important to note that this in vitro behavior will likely be different than in vivo behavior in

regards to carrier degradation and thus rhBMP-2 release. Mammals are known to express

collagenase enzymes that will degrade ACS sponges in several weeks. This would lead to

rapid resorption of the ACS, causing subsequent increased release of rhBMP-2. This rapid

resorption of collagen would lead to loss of the provisional matrix provided by the material,

likely causing the commonly observed voids within the new bone matrix which is

confirmed with our histological findings (Figure 7F)[28,29]. Recent finding suggest these

62

voids are comprised of adipogenic tissue as a result of a high dose of BMP-2 [30]. In

contrast, we are not aware of the presence of keratinase enzymes that specifically degrade

keratins in mammals, indicating that the mechanism of keratin degradation in vivo would

be hydrolytic and/or non-specific proteolytic degradation. This would lead to a more

sustained rhBMP-2 release in vivo and longer time frame for the kerateine carrier to serve

as a matrix to support infiltration of bone matrix forming cells. This is supported by

histological images, which show the presence of implanted (not endogenous) kerateine

materials at the defect site after 16 weeks in vivo (Figure 7 C and D inserts).

While both ACS and kerateine gels or scaffolds loaded with rhBMP-2 led to

bridging of the critically-sized gap, only kerateine scaffolds with rhBMP-2 led to bone

mineral densities statistically equivalent to native bone by the 16 week time point. The

presence of trabecular, woven, and mature bone observed in groups treated with rhBMP-2

carriers (kerateine and ACS) clearly indicates that the rhBMP-2 remained bioactive

(osteoinductive) after biomaterial preparations.

Importantly, there was a significant difference in the levels of ectopic bone growth

between the two carrier systems (ACS vs. kerateine). This was grossly evident upon

explantation of the jaws for tissue recovery and histology as large amounts of ectopic

growth with the collagen carrier was readily observed. Ectopic growth is also

demonstrated by both qualitative (Figure 4D vs. Figures 4F and 4H; Figure 7F vs. Figure

7G and 7H) and quantitative (Figure 6) analysis of CT scans and histological images. It

might be argued that a lower dose of rhBMP-2 would have reduced the ectopic growth seen

from the ACS sponges (as well as kerateine gels and scaffolds). However, we used a

consistent dosage (5 g of rhBMP-2 per defect site) that is in the range reported by other

63

groups working in rat models [31,32]. This is an important result in that kerateine carriers

(gels and scaffolds) of rhBMP-2 led to equivalent or better bone regeneration

characteristics (bridging, bone volume, and bone mineral density) while at the same time

achieving a significant reduction in ectopic bone growth compared to the ACS. We also

note that in a previous study using oxidative form of keratin (keratose) similar levels of

bone regeneration were achieved compared to ACS, but there were no observable

differences in ectopic growth [22]. However, in the current study using the reductive form

of keratin (kerateine), similar levels of bone regeneration were achieved compared to ACS,

but there was a significant reduction in ectopic growth. Although the animal models are

different, this indicates a potential role for the form of the keratin carrier in controlling

rhBMP-2 release and ectopic bone formation.

The histological evidence underscores the role that rhBMP-2 plays as an

osteoinductive protein in bone regeneration but also highlights the less-appreciated role of

the carrier. Location of osteoids in regions within the woven and mature bone matrix

suggests active bone remodeling within all rhBMP-2 carrier groups (Figure 7F, G, and H

inserts). These findings are indicative of the repair phase of bone healing seen in the first

few months after injury. In the absence of rhBMP-2, defect sites treated with kerateine

carriers alone did not calcify the collagenous tissue. The main issue with ACS is the initial

increased release rate of a supra-physiological dose of rhBMP-2 into the local environment,

which leads to ectopic bone growth [33]. This initial release would not be an issue with

physiologically relevant levels of rhBMP-2, but in order to reach the therapeutic threshold

current clinical applications use rhBMP-2 concentrations that are 10 – 1000 fold higher

than native (endogenous) levels of BMP-2 [34]. The rate of biomaterial degradation is

64

important not only for providing a structure for an appropriate length of time (in this case

months) but also for controlling drug release. Equally important is the prevention of rapid

rhBMP-2 diffusion away from the injury site, which promotes osteogenic cell proliferation

and differentiation within the defect area.

It has been recently suggested that an initial burst release followed by a more

sustained release of rhBMP-2 promotes more favorable bone healing [33]. Taken together,

our results suggest such a mechanism occurs in which kerateine carriers are able to promote

a more favorable mandibular bone regeneration with reduced ectopic growth. The relative

lack of burst release by kerateine gels and scaffolds, compared to ACS, is sufficient to

initiate bone healing but low enough to ultimately reduce levels of ectopic growth. From

the in vitro data, it is clear that a large fraction of rhBMP-2 remains within the kerateine

carrier. Therefore, there would be less of a diffusional gradient away from the carrier,

leading to less differentiation of mesenchymal or muscle progenitor cells near but outside

the carrier as a result of reduced levels of soluble rhBMP-2, thereby reducing ectopic bone

growth [35]. This conclusion is supported by our observation during explant of the

mandibles where the ectopic growth around ACS carriers had a gradient appearance

ranging from bone closer to the implant site to more cartilaginous tissue further away from

the implant, as would be expected from a decreasing rhBMP-2 gradient [36]. We also note

that ectopic bone formation for ACS groups was aligned with the direction and location of

the surgical incision, indicating that the ACS does not sequester the rhBMP-2 to the

intended site of action. In other words, it is possible that the rhBMP-2 retained by the

carrier is equally important to regeneration in the defect site as the soluble rhBMP-2

released from the carrier.

65

The result of similar or improved bone regeneration in combination with the

reduced ectopic bone growth observed when using kerateine carriers is particularly

important in craniofacial applications where it essential to preserve the aesthetics of the

facial structure by limiting ectopic bone formation. The tunable nature of kerateine

formulations (e.g., by weight percent or α: γ ratios) lends itself to tailoring formulations to

specific needs for a particular injury (e.g., rapid or sustained matrix degradation/rhBMP-2

delivery).

6. Conclusion

Kerateine biomaterials offer a naturally-based polymeric system that can provide

tailored rhBMP-2 release profiles and degradation. In a critically-sized rat mandibular

model the kerateine biomaterials reduced ectopic bone growth, and achieved comparable

levels of bone healing to existing collagen-based clinical alternatives for mandible injuries.

In addition, fabrication of rhBMP-2 carriers into easily deliverable injectable hydrogels or

implantable scaffolds such as the kerateine carriers described in this report would reduce

the invasive nature of reconstructive procedures, thereby reducing potential subsequent

complications such as infection. The benefit of an injectable gel is the potential to conform

directly to the irregularly shaped defect or a mold that fits the geometry of the craniofacial

injuries. Given the challenges related to ectopic bone formation observed with ACS, the

analogous healing profiles and reduction in ectopic bone formation observed with keratin-

based carriers highlight the potential clinical utility of these materials in bone regeneration.

66

Conflicts of interest

Some authors have potential conflicts interest to disclose as co-founder (MVD),

shareholders (MVD, LB, ME, ST), employees (LB, ME, ST), or received

research/institutional support from KeraNetics LLC (JMS, MVD, TS, CJK). Wake Forest

University Health Sciences has a potential financial interest in KeraNetics, LLC through

licensing agreements. The terms of this arrangement have been reviewed and approved by

Wake Forest University Health Sciences in accordance with its conflict of interest policy.

Acknowledgements

rhBMP-2 used for this work was generously donated by Pfizer. We acknowledge

Cincinnati Children’s Hospital Medical Center (Ronald Pratt, John Pearce, and Diana

Lindquist) for µCT imaging and the Center for Applied Microscopy and Imaging (Richard

Edelmann and Matthew Duley) for histological and SEM imaging. This work was partially

supported by the National Institutes of Health (JMS; R01AR061391) and the content is

solely the responsibility of the authors and does not necessarily represent the official views

of the National Institutes of Health. This work was also partially supported by the US Army

Medical Research and Materiel Command under Contract No.W81XWH-10-C-0165

(LB). The views, opinions and/or findings contained in this report are those of the

author(s) and should not be construed as an official Department of the Army position,

policy or decision unless so designated by other documentation.

67

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CHAPTER IV

SURFACE-MEDIATED DELIVERY OF SIRNA USING POLYCATIONIC

COMPLEXES FROM FIBRIN HYDROGELS FOR THE KNOCKDOWN OF

INHIBITORY MOLECULES.

Christine J. Kowalczewski and Justin M. Saul


in an advisory and editing capacity. Text and figure formatting variations were due to

journal guidelines. This work will be submitted to PlOS ONE.

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______________________________________________________________________________

Abstract

Tissue engineering/regenerative medicine (TERM) has offered the possibility of

generating new tissues following injury or disease. However, the regenerative potential of

tissues in humans is often hindered by the expression of inhibitory molecules. In the case

of nerve injuries, one such molecule is myelin-associated glycoprotein (MAG), which is

up-regulated following injury to the nerve. Traditional TERM approaches focus on

incorporating cells and/or molecules (e.g., growth factors) into a biomaterial (“scaffold”)

in an effort to promote tissue regeneration. Such growth-promoting have led to significant

steps forward in achieving tissue regeneration in nerve and in other tissues. However, less

emphasis has been placed on promoting growth by removal of inhibitory molecule barriers.

One approach to regulate the local expression of inhibitory molecules is through delivery

of small interfering RNA (siRNA). In this study, we investigated the ability to deliver

siRNA directly to cells via delivery from a fibrin hydrogel surface (surface-mediated

delivery) after complexation of the siRNA with polycationic materials. We investigated

complexation with two lipid-based complexing agents (Dharmafect and Lipofectamine)

and one polymer-based complexing agent (25kDa linear polyethylenimine; PEI). SEM

images and release kinetics of the siRNA complexes confirmed stable adsorption of all

three complexes to the fibrin surface. After successful adsorption of siRNA complexes,

S16 cells (a MAG-expressing Schwann cell line) were seeded onto the fibrin to measure

uptake, cytotoxicity, and bioactivity of the siRNA at a 48 hour time point. For all

complexing agents, nearly 100% of S16 seeded on fibrin gels coated with siRNA

complexes showed cell uptake within the first four hours, and high levels of siRNA uptake

were observed in cells. 48 hours after surface-mediated delivery of MAG siRNA

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complexed with DharmaFECT, Lipofectamine, and PEI N:P=20 were able to knockdown

92.47±0.44%, 69.54±1.33%, and 85.9±0.59% of normal MAG mRNA expression. Some

off-target effects were observed depending on the control siRNA used. Local delivery of

siRNA complexes from fibrin hydrogels remains a potential way to temporally regulate

molecular inhibitors of tissue regeneration.

Keywords: biomaterial, gene therapy, temporal, myelin-associated glycoprotein (MAG)

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1. Introduction

Over the past decade, RNA-Interference (RNAi) technology has provided the

biomedical sciences a novel tool to selectively target and silence protein expression by

cleaving the respective sequence-specific complimentary mRNA. This process normally

occurs in the body when Dicer cleaves double stranded RNAs which then binds with RNA

induced silencing complex (RISC). The RISC complex binds and cleaves complimentary

mRNA. There are no endogenous small interfering RNA (siRNA); exogenous siRNA

bypasses DICER and is able to directly associate with RISC [1]. siRNA has been of

particular interest due to the ability to tailor synthetic siRNA to predetermined target

sequences of the mRNA of interest which would provide the greatest efficacy of gene

silencing [2]. Inopportunely, siRNA’s phosphate end groups causes an overall negatively

charged molecule which, along with its relatively large size (~13kDa), limits its ability to

passively diffuse across the cell membrane [3]. To overcome these delivery barriers, a

number of transfection vehicles (viral and non-viral) have been developed to package the

siRNA into nano-sized complexes for cellular delivery. Systemic delivery of these

complexes has been successfully used to treat a wide range of diseases and cancer in animal

models leading to a number of clinical trials [4]. Targeting local areas of interests after

systemic delivery siRNA for regenerative medicine and tissue engineering platforms has

proven to be more difficult due to rapid dispersion, low transfection efficiency, and short-

term silencing in cells. To this end, researchers have begun to incorporate siRNA

complexes into a number of synthetic [5,6] and natural [7,8] biomaterials. Biodegradable

biomaterials have long been utilized to provide a matrix for cells and a reservoir for

therapeutic agents in order to support wound healing until normal physiological function

is restored and are therefore a logical starting point as siRNA carriers for regenerative

74

medicine applications. However, alternative delivery strategies to those previously used

are required in order to increase cellular uptake and enhance the effects of the siRNA.

Fibrin is a natural biodegradable polymer that has been used for therapeutic

delivery for decades due to its ability interact and bind other proteins and promote cell

adhesion [9,10]. Fibrin has long been utilized for therapeutic biomedical applications

including successful delivery of non-viral nucleic acids [11], including our group’s

previous work [12]. Fibrin-siRNA technology is of particular interest in applications where

the healing potential of an injury site is often diminished by locally expressed inhibitory

molecules [13], like seen in cases of nerve injury [14]. Nerve injuries are one example in

which changes in gene and protein expression following injury interferes with the tissue’s

ability to regenerate after injury [15].

Inhibitory molecules that hinder neurite growth are present in both the central and

peripheral nervous systems [16,17]. Myelin-associated glycoprotein (MAG) is an integral

membrane protein which functions as a cell adhesion molecule expressed by myelinating

glial cells. MAG also plays a crucial role in the long-term maintenance and health of axons

and the surrounding myelin [18] in non-injury conditions. After nerve injury, proteins such

as MAG are a major concern since they are part of a milieu of both physical and chemical

barriers. MAG and other proteins thus inhibit the ability of the growth cone to pass the site

of injury [19,20] and reestablish connection to the distal nerve end. Fibrin conduits for

central [21] and peripheral nerve regeneration [22] have achieved promising results,

showing ability to promote nerve growth through the scaffolds either alone or loaded with

growth factors, but these strategies do not address the issue of inhibitory molecules which

dampen their regenerative potential.

75

Our goal was to investigate the ability to directly deliver siRNA from the surface

of fibrin hydrogels to (Schwann) cells seeded onto the surface of fibrin. We complexed

siRNA with three commercially available, cationic, non-viral transfection reagents

(DharmaFECT 3, Lipofectamine 2000, and linear 25kDa polyethylenimine) and adsorbed

these to fibrin hydrogels. We hypothesized that the polycation-siRNA complexes would

remain associated with fibrin, thus allowing for direct cellular uptake of the complexes

from the fibrin surfaces. Moreover, this approach would provide a means to achieve local

molecular control of inhibitory molecules through a fibrin siRNA complex carrier system

in situations of nerve injury. To this end, we chose the MAG-expressing immortalized

Schwann cell line S16 [23] to measure sustained siRNA complex uptake, transfection

efficiency, and inhibitory molecule (MAG) mRNA knockdown 48 hours after cells were

associated with the fibrin hydrogels coated in complexed siRNA.


2.1 Complexation of siRNA with Polycationic Transfection Vehicles

Accell eGFP siRNA (Thermo Fisher Scientific, Waltham, MA), Accell Control

siRNA #1 Non Targeting (Thermo Fisher Scientific, Waltham, MA), All Stars Negative

Control - Alexa 488 siRNA (Qiagen, Venlo, NL), and Accell SmartPool MAG siRNA

(Thermo Fisher Scientific, Waltham, MA) suspending in sterile RNase-free water for a

final stock concentration of 20mM were used throughout this study. siRNA was complexed

with two commercially available lipid transfection agents, DharmaFECT 2 (Thermo Fisher

Scientific, Pittsburgh, PA ) and Lipofectamine 2000 (Life Technologies, Grand Island,

NY), and one polymer transfection agent, 25 kDa linear polyethylenimine (Polysciences,

Warrington, PA) suspended in sterile RNase-free water at a stock concentration of 1mg/mL.

20mM siRNA was complexed to the polycationic lipids using manufacturer’s directions

76

for both DharmaFECT and Lipofectamine (known as lipoplexes). PEI was complexed with

siRNA with increasing N:P (nitrogen : phosphate) (known as polyplexes) to establish the

minimum N:P ratio necessary for siRNA complexation. After initial testing an N:P = 1:20

was used for subsequent experiments with PEI. siRNA and transfection agents were

diluted into working volumes with sterile RNase-free water (Thermo Fisher Scientific,

Waltham, MA) under aseptic conditions. For delivery of 3µg siRNA (n=3; 1 µg per 24

well): 4.41µL Lipofectamine 2000, 35µL DharmaFECT, or 7.71 µL PEI stock solution was

added to RNase-free water for a total volume of 50µL in a sterile 1.5mL conical tube. In a

different sterile 1.5mL conical tube 11.28µL of 20µM siRNA stock solution was added of

a total volume of 50µL of RNase-free water. The respective Lipofectamine and siRNA

solutions were then mixed together (100uL total) and allowed to complex at room

temperature for 20 minute at room temperature. After 20 minutes an additional 350 µL

RNase-free water was added to each respective transfection vehicle to a total volume of

450 µL containing 3 µg siRNA complexes (n=3; 1 µg per 24 well). 150 µL of each then

immediately used for subsequent experiments. For lower and higher siRNA doses,

transfection agents and siRNA volumes were adjusted accordingly to scale.

2.2 Electrophoretic Mobility Shift Assay

siRNA complexation by the three transfection agents was verified by

electrophoretic mobility shift assay (EMSA). Transfection agents were complexed with

0.5 µg eGFP siRNA as described above. A 1% (weight/volume) agarose gel was prepared

by mixing biomolecular grade agarose (Thermo Fisher Scientific, Waltham, MA) and

ethidium bromide (Thermo Fisher Scientific, Waltham, MA) in tris(hydroxymethyl)

aminomethane (Tris)-acetic acid-Ethylenediaminetetraacetic acid (EDTA) (TAE) buffer

77

(Thermo Fisher Scientific, Waltham, MA), which was cast and allowed to cool to room

temperature. Uncomplexed eGFP siRNA served as an experimental control to evaluate if

any siRNA was not successfully complexed by the transfection vehicle. A 1kb DNA ladder

marker (Promega, Madison, WI) was used to track the progression of free eGFP siRNA

which was not bound by the transfection agent as it migrates through the gel. 10µL of each

experimental group was mixed with 2 µL loading dye, loaded into a well, and EMSA was

run in TAE buffer for 30 min at 100mV and visualization was performed under UV light.

2.3 Dynamic Light Scattering (DLS)

siRNA (for GFP) complexed with each polycation was evaluated for diameter

measured by dynamic light scattering (DLS). Each siRNA complexing agent nanoparticle

formulation was brought to a final concentration of 1µg/mL solution in sterile DI water (n

= 3 for each formulation). This solution was gently suspended and loaded into a UV-

transparent cuvette (Sarstedt, Sarstedt, Germany) for DLS particle size analysis on a

Malvern Nanosizer (Malvern, Worcestershire, United Kingdom).

2.4 Preparation of Fibrin Hydrogel

Fibrinogen powder from bovine plasma (Sigma-Aldrich, St. Louis, MO) was

dissolved with sterile RNase-free water under aseptic conditions at 200 mg/mL and placed

in a 37˚C water bath to dissolve. The fibrinogen solution was used to coat the bottom of

22mm square glass coverslips (50 µL fibrinogen solution), 48 (25 µL fibrinogen solution),

24 (50µL fibrinogen solution), or 6 well plate (240µL fibrinogen solution), depending on

the experiment (see below). One part fibrinogen solution was mixed with three parts

thrombin working solution which consists of 13 parts PBS to one part 5.5mg/mL CaCl2

(Thermo Fisher Scientific, Waltham, MA) and one part 250U/mL bovine plasma thrombin

78

(Sigma-Aldrich, St. Louis, MO) to generate the fibrin hydrogels. The resulting 5% (w/v)

fibrin hydrogel was allowed to polymerize overnight at 37°C in a cell incubator (humidified

atmosphere of 5% CO2).

2.5 Scanning Electron Microscopy (SEM)

Fibrin hydrogel formulations prepared in a 24 well dish as described above were

cut with a 5mm biopsy punch and placed into a 1.5mL conical tube. Sterile RNase-free

water containing each siRNA-complex formulation (with PEI prepared at an N:P=20 ratio)

were adsorbed to fibrin hydrogels for 4 hours at a total amount of siRNA of 5 g per well.

RNase-free water and eGFP siRNA alone (not complexed with a transfection agent) served

as controls. Hydrogels were then taken through alcohol dehydration steps, critical point

dried, and gold sputter coated (2 minutes ~90 nm thickness). Adsorbed siRNA complexes

or appropriate controls were imaged with Zeiss Supra 35 VP FE-SEM (Jena, Germany) at

15k x magnification. Adsorbed siRNA complex particle size was measured using ImageJ

software to measure the complex size distribution between 5 images of the fibrin hydrogel

surface.

2.6 siRNA Complex Desorption Kinetics

Fibrin hydrogels were prepared as described above in a 48 well dish. Sterile RNase-

free water containing each siRNA-complex type were allowed to adsorb the fibrin gel

surface for hours (n = 3 for each experimental group). For these experiments, 0.5 g of

siRNA was used in each well. Alexa 488 siRNA was used because the siRNA can be

detected by fluorescence reading. Sterile RNase-free water served as a blank for

subtraction of any autofluorescence associated with the degradation of the fibrin hydrogel.

After 4 hours, the solution was collected to determine the amount of Alexa 488 siRNA that

79

did not adsorb to the surface (time = 0). The hydrogel was washed once with 200µL of

PBS and then added to the previously collected non-adsorbed siRNA to determine the

overall amount of siRNA that was no longer present. 200µL of fresh PBS was added to

each well and at pre-assigned time points 100µL the PBS was collected and replaced with

100µL of fresh PBS. All time point samples were frozen and stored at -20°C until thawed

for analysis. A final concentration of 1% TritonX-100 (Sigma Aldrich, St. Louis, MO) and

2% SDS (Sigma Aldrich, St. Louis, MO) was added to each sample in order to release the

Alexa 488 siRNA from the transfection vehicle. Fluorescent readings were then taken of

each sample and were correlated to a standard curve of known Alexa488 siRNA

concentrations on a BioTek plate reader (Winooski, VT) at 485nm/525 nm Ex/Em.

2.7 Cell Culture

S16 Schwann cells were obtained from ATCC (ATCC CRL-2941, Manassas, VA)

and cultured as recommend by using Dulbecco's Modified Eagle's Medium (DMEM)

supplemented with 10% Fetal Bovine Serum (FBS), and 1% Antibiotic-Antimycotic (A-

A, Gibco, Life Technologies Grand Island, NY) at 37°C in a cell incubator (humidified

atmosphere of 5% CO2). Cells were cultured on 150mm cell culture dishes (Thermo Fisher

Scientific,Waltham, MA) treated with 15µg/mL poly-L-lysine (Sigma-Alrich, St. Louis,

MO) until they were 60-70% confluent. At this time media was removed and cells were

washed with 10mL PBS and detached with 0.25% Trypsin-EDTA (Gibco, Life

Technologies Grand Island, NY) for 5 minutes in a cell incubator. 5mL of the complete

growth medium described above (DMEM + 10% FBS + 1% A-A) was added and cells

were centrifuged for 5 minutes. Cells were resuspended in fresh complete growth media

80

and plated 1:8 with complete growth media. In the following experiments, “transfection

media” refers to DMEM supplemented with 10% FBS alone without the addition of A-A.

2.8 Confocal Microscopy

22mm round glass coverslips were thinly coated with fibrin hydrogels as describe

above and were allowed to polymerize overnight in a 6 well plate. For this study, 1ug

Alexa 488 siRNA was complexed with Lipofectamine 2000, DharmaFECT, or PEI

(N:P=20) in sterile RNase-free water for 20 minutes. Fibrin hydrogels were prewashed

with sterile RNase-free water. Normal untreated S16 cells cultured on fibrin hydrogels

treated only with water (no siRNA treatments) or with uncomplexed Alexa 488 siRNA

suspended in sterile RNase-free water (no complexing agents) served as negative controls.

150µL of each siRNA-complexing agent was added drop wise to the top of the fibrin coated

coverslip and allowed to adsorb for four hours. After four hours, the coverslips were

washed with 200µL sterile PBS. S16 were passaged in transfection media (DMEM; 10%

FBS; 0% A-A) and 10,000 cells (in 150µL transfection media) were seeded onto each fibrin

hydrogel (on coverslip). The cells were allowed to attach for four hours after which the

transfection media was removed and replaced with normal culture media. After 24 hours,

coverslips were washed with PBS, fixed in 10% formaldehyde (methanol-free) for 10

minutes. Cells were counter stained by using manufacture’s guidelines for Alexa 555

phalloidin stain for F-Actin (Invitrogen, Carlsbad, CA) and DAPI for nuclei (Invitrogen,

Carlsbad, CA). Coverslips were then mounted to glass slides with MM83 mounting media

(Leica,Wetzlar, Germany) and allowed to dry overnight in the dark at 4°C. Confocal

imaging (Ziess 710 Laser Scanning Confocal System, Jena, Germany) of experimental

81

groups was performed at 20x magnification. Zeiss LSM Image Browser (Zeiss, Jena,

Germany) was used for image processing.

2.9 Flow Cytometry

Fibrin hydrogels were coated on the bottom of a 6 well plates as described above

for these experiments. A prewash of 1mL of sterile RNase-free water was added to the

hydrogel surface. Alexa 488 siRNA was complexed with Lipofectamine, DharmaFECT, or

PEI (N:P=20) in order to deliver 4µg of Alexa 488 siRNA per well in a total volume of 300

L. Uncomplexed Alexa 488 or water alone served as negative controls. All groups were

tested in triplicate (n = 3). siRNA-complexes were allowed to adsorb for 4 hours to the

fibrin hydrogel surface. After four hours the Alexa 488 siRNA solutions were removed

and the hydrogel surface was washed once with PBS. Immediately afterwards 250,000

S16 cells were seeded onto the hydrogel surface with 300µL transfection media (DMEM;

10%FBS;0%A-A) for four hours. The transfection media was removed and the hydrogel

surface was washed once with PBS. Then 1mL of culture media (DMEM; 10%FBS; 1%A-

A) was added to each well and maintained until reaching pre-established time points of 4

and 48 hours. At each preassigned time point, media was removed and cells were washed

once with PBS. Cells were detached from the hydrogel surface with two 5 minute

incubations of 0.25% trypsin (Gibco, Life Technologies Grand Island, NY). Wells were

washed once with culture media to ensure collection of cells. Cells were collected into

15mL conical tubes and centrifuged for 5 minutes. Supernatant was removed and the cell

pellet was resuspended in 1mL PBS to serve as a wash step. Cells were then centrifuged

again for 5 minutes, the PBS supernatant was removed, and the cell pellet was suspended

in 500µL of sterile Flow Fluid (0.1% BSA in PBS) or BD FACsFlow sheath fluid (BD,

82

Franklin Lakes, NJ). Each 500µL cell suspension was then pipetted through a 0.4µm cell

strainer (BD, Franklin Lakes, NJ) into a polystyrene flow cytometer tube (BD, Franklin

Lakes, NJ) and placed on ice until analysis was performed. Alexa 488 siRNA uptake was

measured by using a BD FACScan™ or BD FACSCalibur™ (BD, Franklin Lakes, NJ) at

488nm excitation laser. Cells only (without Alexa 488 siRNA treatment) served as negative

controls to determine acquisition parameters. 10,000 events were captured for each

treatment group (n = 3). BD Cell Quest Pro software (BD, Franklin Lakes, NJ) was used

for data acquisition and analysis.

2.10 Cell Viability (MTS)

For cell viability assays, fibrin hydrogels were prepared in 48 well dishes as

described above. Sterile RNase-free water containing 0.5µg MAG siRNA complexed with

Dharmafact, Lipofectamine, or PEI (N:P = 20) was allowed to adsorb to the fibrin gel

surface for 4 hours. Uncomplexed 0.5µg MAG and sterile RNase-free water served as

negative controls. All groups were tested in triplicate (n = 3). After four hours, wells were

washed with sterile PBS and 10,000 S16 were seeded onto each well with transfection

media. After another four hours transfection media was removed and replaced with 200µL

culture media. Cells were incubated for 48 hours at which point the culture media was

removed and replaced with 200µL of CellTiter 96® Aqueous Non-Radioactive Cell

Proliferation Assay (Promega, Madison, WI) for one hour in a humidified 37°C, 5% carbon

dioxide (CO2) incubator according to the manufacturer’s directions. The reaction was

stopped by cell lysis with a 10% SDS solution. 200µL of the cell lysate was transferred to

a 96 well plate and 490nm absorbance readings were immediately taken on a BioTek plate

reader (Winooski, VT) and then normalized to normal S16 (mean ± SD).

83

2.11 Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)

For qRT-PCR experiments, fibrin hydrogels were coated onto 24-well plates and

coated with either MAG or NT siRNA to deliver 1 g of siRNA/well as described above.

30,000 S16 cells were seeded with 200µL transfection media per well and allowed to attach

for 4 hours. Afterwards, the transfection media was removed and replaced with 200µL of

normal culture media. Cells were incubated at 37°C in a cell incubator (humidified

atmosphere of 5% CO2) for 48 hours at which time the cells were harvested.

At each pre-assigned time point, media was removed and cells were washed once

with PBS. Cells were detached from the hydrogel surface with two 5 minute incubations

of 0.25% trypsin (Gibco, Life Technologies Grand Island, NY). Wells were then washed

once with culture media to ensure collection of cells, collected into 15mL conical tubes,

and centrifuged for 5 minutes. Supernatant was removed and the cell pellet was

resuspended in 1mL PBS to serve as a wash step. Cells were then centrifuged for 5 minutes,

the PBS supernatant was removed, and the cell pellet was stored at -20°C until RNA

extraction was performed.

RNA extraction with an RNeasy Mini Kit (Qiagen, Venlo, NL) was followed by

RNA quantification with a NanoDrop 2000 (Thermo Scientific, Wilmington, DE).

Samples were normalized to 50ng RNA and immediately reverse transcribed into cDNA

by Omniscript RT kit according to the manufacturer’s instructions (Qiagen, Venlo,

Netherlands) and frozen at -20° C until analyzed by qPCR. cDNA samples were mixed

with QuantiTech SYBR Green PCR kit (Qiagen, Venlo, Netherlands) and primers

(QuantiTech Primer Assay, Qiagen, Venlo, Netherlands) targeting rat MAG (QT00195391)

or beta-actin control (QT00193473). Quantitative PCR was performed on a Bio-Rad

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iCycler (Bio Rad, Hercules, CA) for 40 cycles. Data was analyzed using Biogazelle

QBase+ software (Zwijnaarde, Belgium) and MAG expression was compared to reference

gene expression β-actin) then MAG expression was normalized to normal S16 seeded on

fibrin hydrogels.


Statistical analysis was performed by using one way analysis of variances

(ANOVA) followed by the Tukey’s post hoc test to compare differences between groups.

p<0.05 was considered statistically significant.

3. Results

3.1 Electrophoretic Mobility Shift Assay

We used an electrophoretic mobility shift assay (EMSA) in order to ensure that the

siRNA was complexed by each complexing agent at the amount of material used.

Dharmafect and Lipofectamine were used at levels recommended by the manufacturer for

siRNA, but we tested several N:P ratios for PEI due to some variability in the literature

regarding the appropriate N:P ratio to use. The uncomplexed siRNA control migrated

uninhibited through the agarose gel while DharmaFECT and Lipofectamine 2000 both

were able to complex the eGFP siRNA. The lowest N:P ratio able to complex siRNA with

PEI was N:P=5, with increasing N:P ratios allowing less siRNA to move through the gel

(Figure 1). We selected N:P = 20 for further experiments because this would ensure

complex formation, pilot qRT-PCR and flow cytometry studies, and because we have

previously had success in delivering plasmid DNA at this N:P ratio [12].

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Figure 1. Electrophoretic mobility shift assay of siRNA transfection agents. GFP

siRNA successful complexation with DharmaFECT, Lipofectamine, and PEI with a

N:P=5 or greater was verified by Electron Shift Mobility Assay. GFP siRNA alone

served as negative control.

3.2 Dynamic Light Scattering

Complex diameter size distribution was measured by dynamic light scattering

(DLS) in order to determine the diameters of the siRNA-complexes before their adsorption

to the fibrin hydrogel surfaces. All experimental groups demonstrate a Gaussian

distribution between 100-200nm (Table 1). This is in the range expected for such siRNA

complexes and falls within the diameter suitable for cellular uptake via endocytosis.

Indications of aggregation (bimodal distribution with one peak centered around a few

micron and not nanometer diameter) that were not observed by DLS.

Table 1. Dynamic light scattering measurements of siRNA complex diameters.

siRNA complex Diameter Measurements by Dynamic Light Scattering.

Transfection

Vehicle

Complex Diameter

(nm ± SD)

DharmaFECT 119.66 ± 17.66

Lipofectamine 128.17 ± 30.12

PEI N:P =20 197.06 ± 39.9

86


A key aspect to this approach of surface-mediated transfection is the stable

adsorption of the siRNA-complexes to the fibrin hydrogel surface. To ensure that this

adsorption process did not disrupt the complexes or lead to aggregation, we used SEM to

visualize the surfaces and also quantify the size of the siRNA complexes. Images were

taken of the fibrin hydrogel surface after allowing either RNase-free water or complexes

to adsorb for four hours. As expected, fibrin surfaces treated with only RNAase-free water

showed the typical fibrous structure but no nanoparticles inherent to the gels that could be

misinterpreted as siRNA complexes (Figure 2A). eGFP complexed with Lipofectamine,

DharmaFECT, or PEI N:P=20 (Figure 2B, 2C, 2D) show successful complex adsorption to

the fibrin surface (Figure 2B, 2C, 2D arrows). ImageJ measurements for each of 5 separate

areas were collected and are represented individually (Figure 2E, 2F, 2G, 2H) by

histograms. The diameter of the adsorbed DharmaFECT, Lipofectamine, and PEI N:P=20

compexes were measured to be 143±27nm, 141±28nm, and 138±30nm respectively

(Figure 2E, 2F, 2G, 2H inserts). These measurements are consistent with DLS

measurements of complex diameter in solution (Table 1), indicating that the adsorption did

not lead to major changes in the complexes. In addition, minimal levels of aggregation of

the complexes were observed.

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Figure 2. siRNA complex adsorption and size distribution on fibrin hydrogel surface. FE-SEM images of (A) fibrin alone or adsorbed siRNA complexed with (B) DharmaFECT,

(C) Lipofectamine, or (D) PEI N:P=20. 5 random images of siRNA complexes adsorbed

to the fibrin surface (white arrows) analyzed by ImageJ to determine particle diameter size

(µm) of adsorbed complexes of (E) fibrin no siRNA, (F) DharmaFECT, (G) Lipofectamine,

(H) PEI N:P=20. Insert is the average diameter ± STDEV.


To measure the ability of the complexes to remain on the hydrogel surface to allow

for surface-mediated transfection over a period of time, the rate of siRNA complex

desorption was be measured (Figure 3). After three days the majority of siRNA complexes

remained adsorbed to the hydrogel surface. The transfection agent with the greatest rate

of release was the PEI (N:P=20), which showed a statistically different rate of release

compared to the cationic lipid transfection reagents (Dharmafect and Lipofectamine) after

3 hours. Of the cationic lipids, Lipofectamine showed the slowest rate of release from the

fibrin surface. After 24 hours PEI Lipofectamine, and Dhamafect each had statistically

different release profiles from each other. At the end of three days 93.8±1.27% PEI N:P=20,

96.37±0.45% DharmaFECT, and 98.7±0.73% Lipofectamine siRNA-AF488 complexes

remained adsorbed to the fibrin hydrogel surface.

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Figure 3. Release kinetics of siRNA complexes from fibrin hydrogel surface. Release

kinetics of siRNA-AF488 complexes after being allowed to adsorbed to a fibrin surface

for four hours (n=3; mean ± SD). *p<0.05 PEI N:P = 20 versus DharmaFECT and

Lipofectamine, *p<0.05 Between all treatment groups


Confocal microscopy was performed 24 hours post surface-mediated transfection

in order to visualize the internalization and distribution of siRNA delivered from the

various complexing agents to S16 cells. Alexa 488 siRNA was used for these studies in

order to allow visualization of the siRNA for the various treatment groups. Normal S16 (no

siRNA present) seeded on a fibrin surface showed the normal morphology with actin

filaments and nuclei (Figure 4A), and we did not observe deviation from this morphology

in S16 cells in the presence of any of the complexing agents. Uncomplexed Alexa 488

siRNA was not observed to be internalized into S16 (Figure 4B). Surface-mediated

transfection of siRNA complexes showed positive uptake and internalization of Alexa 488

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siRNA complexed with DharmaFECT (Figure 4C) and Lipofectamine (Figure 4D).

Positive uptake of Alexa 488 siRNA PEI N:P=20 (Figure 4E) was not observed and is

believed to be due to PEI complexation causing signal quenching. Because the confocal

microscope is able to visualize planes of view within the cell, these results also demonstrate

that the cells have internalized the siRNA and that the siRNA is not simply on the surface

of the fibrin underneath the cells (which would be difficult to determine with an

epifluorescence microscope image).

90

Figure 4. Schwann cell internalization of siRNA complexes. Confocal images of S16

cells 24 hours after being seeded on the surface of (A) fibrin only or treatments adsorbed

to the surface for four hours: (B) uncomplexed Alexa 488 siRNA, (C) DharmaFECT, (D)

Lipofectamine, or (E) PEI N:P = 20. (blue = DAPI nucleus; red = F-actin; green =

siRNA) Scale bar denotes 100 µm. Arrow highlights internalized siRNA complexes.

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3.6 Flow Cytometry

We reasoned that in order to achieve biological effect, a high percentage of cells

would require uptake of siRNA. In order to quantify the percentage of the cell population

that was taking up the siRNA for each of the various formulations or controls, we used

flow cytometry to track the number (or percentage) of cells that had associated siRNA at 4

and 48 hours (Table 2). Normal S16 cells (no siRNA present) was used to establish a gate

for detection of Alexa 488 siRNA. Uncomplexed Alexa 488 siRNA achieved the lowest

transfection efficiency with only 3.81±0.85% of cells showing uptake after 4 hours. The

percentage of cells taking up siRNA complexes was slightly higher at 4 hours and then

decreased at 48, likely due to “dilution” of the siRNA as the cells proliferated (Figure 5).

siRNA delivered from each of the complexes also decreased over the course of the 3 days

(data not shown), again likely due to this dilution effect as the cells proliferated.

All three complexing agents achieved nearly 100% transfection efficiency at the 4

hour time point. PEI initially showed the greatest number of transfected cells

(99.47±0.36%) which decreased on the second day (87.11±4.45%). Both cationic lipid

agents (DharmaFECT and Lipofectamine) were statistically equivalent in their ability to

successful transfect S16 cells after 48 hours. On the last day, DharmaFECT (96.22±0.45%)

and Lipofectamine (86.15±21.09%) retained statistically greater transfection compared to

both the uncomplexed siRNA and PEI.

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Table 2. Transfection efficacy of surface-mediated siRNA into Schwann cells. Flow

Cytometry data showing the percentage of S16 Schwannoma cells associated with

fluorescently tagged siRNA complexes following 4 and 48 hours of incubation (n = 3; mean

± SD). *p<0.05 Uncomplexed siRNA versus siRNA complexes.

Treatment 4 Hours 48 Hours

Uncomplexed siRNA 3.81 ± 0.85 * 0.13 ± 0.01 *

DharmaFECT 98.68 ± 0.75 96.22 ± 0.45

Lipofectamine 99.8 ± 0.06 86.15 ± 21.09

PEI N:P = 20 99.47 ± 0.36 87.11 ± 4.45

Figure 5. Flow cytometry gate analysis of Schwann cells. Representative flow

cytometer gates after 48 hours of S16 seeded on (A) fibrin only (no siRNA-AF488), (B)

uncomplexed siRNA-AF488, or siRNA-AF488 complexed with (C) DharmaFECT, (D)

Lipofectamine, and (E) PEI N:P=20. Events falling in gate R2 were counted as positive

for siRNA-488 and were tabulated to create Table 2.

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3.7 Cell Viability (MTS)

Given the relatively high amounts of uptake achieved by surface-mediated

transfection, we wanted to investigate the potential toxicity of the internalized complexes

on the cells, given the known toxicities associated with solution-phase delivery

(transfection) for these and other transfection reagents [24]. Toxicity was measured by

comparing the metabolic activity of normal S16 cells seeded on the fibrin hydrogel surface

and compared with each respective agent after two days (Figure 6). All data were

normalized to these untreated S16 cells at each specific time point. After 48 hours, the

uncomplexed MAG siRNA as well as DharamFECT and Lipofectamine had no effect on

cell viability, but PEI showed a 15.78±4.63% decrease in cell activity compared to

untreated cells.

Figure 6. Schwann cell viability after surface-mediated siRNA transfection. S16 cell

viability was determined by MTS Assay after 48 hours after being seeded on the surface of

fibrin hydrogels alone, treated with siRNA alone, or complexed with either DharmaFECT,

Lipofectamine, or PEI. Cell viability was normalized to normal cells after 48 hours of

being seeded on the fibrin hydrogel surface (n = 3; mean ± STDEV). *p < 0.05 versus

normal cells.

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3.8 Quantitative Real-Time Polymerase Chain Reaction (qRT-PCR)

In order to determine the ability of the surface-mediated transfected siRNA to

knockdown MAG mRNA levels, qRT-PCR was performed (Figure 7). MAG mRNA

expression was normalized to MAG mRNA levels of normal untreated S16 seeded on the

fibrin hydrogel. At 48 hours, Lipofectamine continued to reduce MAG mRNA levels

(69.54±2.49%) but was still unable to achieve the same level of MAG knockdown as both

DharmaFECT (92.47±0.44% and PEI N:P=20 (85.9±1.33%) when complexed with MAG

siRNA. At this time, off-target (down-regulation of unwanted targets) MAG knockdown

was not measured with PEI N:P=20 GFP but was with DharmaFECT GFP.

Figure 7. qRT-PCR of MAG mRNA after surface-mediated siRNA transfection. qRT-

PCR of MAG mRNA expression in S16 cells 48 hours after surface-mediated siRNA

transfection from a fibrin hydrogel surface. Relative MAG mRNA expression is

normalized to normal MAG mRNA levels. (n=3; mean ± SEM) *p<0.05 versus Normal;

**p<0.05 versus uncomplexed siRNA MAG; %p<0.5 versus PEI N:P=20 GFP; #p<0.05

DharmaFECT GFP; @p<0.05; @p<0.05 versus Lipofectamine MAG.

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4. Discussion

Nature performs a cascade of regulatory tasks during tissue development, which we

are beginning to learn from in order to design regenerative medicine platforms to better

mimic developmental cues for repair of injured tissues. An important lesson from

developmental biology is that molecules cannot be classified into simply “pro” and “con”

regenerative molecules. Thus, some molecules may play one role in one situation (i.e.,

healthy tissue) and another role in a different situation (i.e., injured tissue). Perhaps in no

other tissue is this delicate balance so clearly seen as in the nervous system. Although a

number of inhibitory molecules are up-regulated following nerve injuries (and these are

different in injuries to the central versus peripheral nervous system), myelin-associated

glycoprotein (MAG) is in interesting model and case study.

MAG plays in an important role in healthy tissue, providing support to neurons

from glial cells via direct interactions [18,25]. However, it is also well established that

MAG plays an inhibitory role in nerve regeneration following injury [20]. Despite MAG

expression on supporting glia causing an inhibitory effect on the migrating growth cone of

the axon after nerve injury, once the growth cone has passed the barrier, glia support and

production of MAG should return in order to maintain homeostasis in the regenerated

nerve. MAG is required for successful nerve regeneration due to its role in supporting

neuron survival, enhancing long-term axon-myelin stability, structuring nodes of Ranvier,

and regulating the axon cytoskeleton [25]. For these reasons, it is important to remember

that MAG function must be restored to resume its supportive role if and when the nerve

gap has been “bridged.”

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In this regard, siRNA technology is promising in that it would provide the

opportunity to knock down MAG’s inhibitory effects but do so in a transient fashion to

allow MAG to perform its role in healthy tissue. Given the many studies that have

investigated neural conduits with growth factors to promote nerve guidance across gaps

[26], it is not difficult to envision a system in which siRNA is used alongside these growth

factors to simultaneously knock down inhibitory molecules. If siRNA were adsorbed to

biomaterial scaffolds being used to promote bridging of the nerve injury (i.e., a “neuro-

conductive” material), it could temporarily down-regulate inhibitory molecules during the

bridging process but allow their levels to return to those dictated by the regenerated tissue

following bridging. Thus, siRNA would, in theory, provide a window of opportunity for

the migrating axon to pass through the area without the MAG barrier typically found in the

injured microenvironment. Over time, the migrating cells would take up the siRNA

complexes until it is depleted at which time the silencing effect of the siRNA will decrease,

allowing for the production of MAG in order to resume its supportive role and improve the

potential for nerve regeneration.

The concept of the gene-activated matrix for plasmid DNA delivery to achieve

transgene expression of growth factors is not new [27]. Here we sought to conduct

fundamental preliminary studies on the feasibility of a similar approach for the delivery of

siRNA from biomaterial scaffolds. The goal of this study was to investigate: 1) the ability

of three commercially available siRNA transfection agents (2 cationic lipids and one

polycation) to be adsorbed to a fibrin hydrogel; 3) uptake and internalization of the siRNA

complexes into cells; and 3) mRNA knockdown of an inhibitory molecule target.

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To assess endosomal uptake, siRNA was complexed with three common non-viral

transfection reagents for nucleic acids. EMSA verified stable formation of siRNA

lipoplexes (DharmaFECT and Lipofectamine) and stable polyplexes (linear PEI) when the

N:P ratio was ≥ 5. PEI at an N:P ratio of 20 was chosen for subsequent experiments based

on our own pilot studies, which agreed with previously published work on N:P ratio effect

on knockdown with limited cytotoxic outcome [28]. DLS measurements of respective

distribution of siRNA complex diameters in suspension were <200 nm diameter typically

associated for cellular uptake (Table 1). When adsorbed to the fibrin surface, complexes

retained their size distribution (Figure 2E, 2F, 2G, 2H).

Due to the isoelectric points of fibrinogen (pH 5.5) and fibrin (pH 5.6), the fibrin

hydrogels would be expected to have a negative surface charge at the neutral pH under

which our experiments were conducted. Thus, it is possible to anchor molecules/particles

with an overall positive charge through electrostatic interactions [29]. This has been

utilized with success in a number of nucleic acid delivery systems, leading to research to

enhance the electrostatic interaction [30]. SEM images confirm both lipoplexes and

polyplexes anchor to the fibrin surface (Figure 2A, 2B, 2C, 2D arrows) without matrix

modification to tether the complexes. When release kinetics were measured over the

course of three days, no burst release was observed from the hydrogel. These

measurements indicate that > 95% of each siRNA complexed with lipids (Dharmafect and

Lipofectamine) and > 90% of the siRNA complexed with polymer (PEI) remained

anchored to the fibrin surface and thus available for cellular uptake.

MAG-producing Schwann cells (cell line S16) were seeded onto fibrin hydrogels

coated with uncomplexed siRNA or the three transfection vehicles. Confocal images of

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the S16 cells confirmed uptake and internalization of Alexa 488 siRNA complexes from

the hydrogel surface (Figure 5C, 5D, 5E). PEI packaging of the Alexa 488 siRNA caused

fluorescence signal quenching [31] and was therefore difficult to observe in confocal

images. These findings support the need for siRNA packaging into a transfection vehicle

in order to pass the cell membrane, which uncomplexed Alexa 488 siRNA was unable to

achieve (Figure 5B). Flow cytometry data supported these findings, indicating cellular

uptake of the complexed siRNA. Remarkably, almost 100% of cells were transfected by

DharmaFECT, Lipofectamine, and PEI, while <5% of cells were transfected by

uncomplexed Alexa 488 siRNA within the first 4 hours of contact with the treated hydrogel.

A drop after 24 hours in positively expressing cells could be a result of signal quenching;

in earlier time points there were enough internalized polyplexes to have a strong enough

signal for detection as can be seen in the shift in the gated cell population (Figure 4). As

cells proliferated, the internalized siRNA complexes were likely split between the two

daughter cells. With a confluent plate, all of the available siRNA complexes would have

been taken by and then split amongst the cells. In previous studies, cell migration along the

siRNA complex adsorbed surface has been shown to maintain elevated levels of

transfection [32]. Our current findings agree with surface-mediated pDNA complex

transfection studies which report 2D systems having greater transfection over 3D

encapsulation/incorporation into the biomaterial. Research suggests that this could be due

to greater complex presentation to the cell seen in both lipid-based complexes [11] and

polymer-based complexes [12] in 2D environments. Cell adhesion to the substrate has a

substantial impact on success of substrate-mediated delivery and gene silencing [33] and

99

potentially is one of the main reasons for fibrin’s high transfection rate of nucleic acid

delivery.

With such high transfection rates, cytotoxicity often results [3]. After the first 48

hours of transfection, there was slight, but non-significant effect on cell activity expect for

PEI N:P=20. Even though decreases in cell activity were observed, none of the transfection

vehicles went below 80% of normal untreated cells (Figure 5). Thrombin and fibrinogen

concentrations have been shown to positively affect cell proliferation and migration [34]

which can explain why greater cytotoxic affects were not measured in this system.

The release kinetics and flow cytometry data suggests two possible routes of

transfection: (1) solution-based uptake by which desorbed complexes from the fibrin

hydrogel released into solution are taken up by cells or (2) direct uptake from the surface

(surface-mediated uptake) as the cell migrate over the adsorbed siRNA complexes. In our

model, the low levels of siRNA complex desorption we believe would not achieve the level

of MAG mRNA knockdown which was measured in our system. For this reason, we

believe that knockdown was achieved by surface-mediated transfection. Uptake through

clathrin-mediated endocytosis as well as micropinocytosis are believed to be the two main

methods of siRNA complex internalization. The mechanism of endosome escape is still not

fully understood but are a number of hypotheses: escape during the early stage endosome

of macropinosomes which are “leaky” [35], lipid merging between the lipoplex and

endosome [36], “proton-sponge” effect [37], or formation of transient endosome

membrane pores [38]. No matter which route of endosomal escape occurs, when siRNA

releases into the cytosol it must maintain bioactivity to bind with RNA-induced silencing

complex (RISC) and cleave sequence specific mRNA.

100

In our model, a significant reduction of MAG mRNA was achieved with both

surface adsorbed lipid-based complexes and polymer-based complexes after 48 hours

(Figure 7). Though Uncomplexed siRNA was unable to reduce MAG mRNA levels, likely

due to low levels of cell uptake previously discussed. Unspecific knockdown of our gene

of interest mRNA was observed using non-targeting siRNA. Off-target effects have long

plagued siRNA therapies and can be due to a number of reasons. Non-viral transfection

agents such as lipids and PEI have both resulted in non-specific knockdown [39,40]. Off-

target knockdown can also be due to non-specific siRNA such as those without a

physiological target (ex GFP or non-targeting siRNA), which have been shown to have

sequence-specific off-target effects [41]. Also, siRNA dose concentrations have been

shown to play a role in off-target effects [42]. Solution-phase siRNA transfection doses are

lower than substrate-mediated delivery, but multiple “low dose” solution-phase

applications are needed to the sustain transfection over time. Our current study siRNA

dose correlates with other published substrate-mediated siRNA delivery systems [33,43].

It is important to keep in mind that even though substrate-mediated systems are initially

loaded with higher concentrations then typical solution-phase systems, not all of the

substrate-mediated siRNA is take up immediately by the cells. Off-target knockdown has

also been observed to vary between cell types; therefore, S16 cells may be more sensitive

to RISC-dependent off-target knockdown. Nonetheless, given the levels of knockdown

achieved and the enhanced knockdown compared to non-target sequences, this approach

appears to be a plausible method for delivery to Schwann cells as a means to decrease

levels of inhibitory molecules.

101

Our current work reports the effects of surface-mediated transfection from fibrin

hydrogels. Further studies are required to evaluate this system for applications for longer

term (weeks) gene silencing and return of function of the target molecule. In previous

studies, surface-mediated transfection of non-targeting siRNA complexes from template

nerve conduits had no significant effect on glial cell motility or proliferation, and did not

induce an inflammatory response [44]. Therefore, transient silencing of gene targets can

be applied in a number of regenerative medicine applications as well as loss/gain of

function experiments.

5. Conclusion

These studies demonstrate the ability to achieve surface-mediated delivery of

siRNA from fibrin hydrogels when siRNA is complexed with polycationic polymers and

lipids. A high rate of transfection was maintained during the course of the study with only

slight cytotoxic effects. Importantly, the siRNA complexes achieved significant

knockdown of the target molecules (MAG) after 48 hours. Off-target effects emphasize the

need for contained research in this mainly misunderstood phenomenon. Therefore, these

studies show the viability of a surface-mediated approach to the delivery of siRNA from

biomaterial scaffolds to promote more favorable regeneration by targeted knockdown of

inhibitory molecules.

102

Acknowledgments

The authors acknowledge assistance from the Center for Nanotechnology and

Molecular Materials at Wake Forest University (SEM imaging), Dr. Richard Edelmann and

Matthew Duley at the Miami University Center for Advanced Microscopy and Imaging

(confocal microscopy). This work was partially supported by the National Institutes of

Health (JMS; R01AR061391) and the content is solely the responsibility of the authors and

does not necessarily represent the official views of the National Institutes of Health. This

work was also partially supported by The Wake Forest University Health Sciences

Translational Sciences Institute Pilot Grant and The Wake Forest Graduate School.

103

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CHAPTER V

KNOCKDOWN OF THE BONE MORPHOGENETIC PROTEIN-2 BINDING

ANTAGONIST NOGGIN VIA SURFACE-MEDIATED DELIVERY OF SIRNA FROM

FIBRIN HYDROGELS.

Christine J. Kowalczewski and Justin M. Saul


in an advisory and editing capacity. Text and figure formatting variations were due to

journal guidelines. This work will be submitted to Biomaterials.

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________________________________________________________________________

Abstract

The bone morphogenetic protein 2 (BMP-2) bone substitute market is estimated to

account for $858 million annually. Unfortunately, clinically reported adverse side effects

have been observed, likely owing to the requirement for supraphysiological doses of BMP-

2. As such, approved clinical applications have been limited to lumbar spinal fusion, tibial

fracture, and some craniofacial applications. One reason for the required supraphysiologic

BMP-2 dose may be to overcome the opposing effects of noggin, a BMP-2 binding

antagonist that is known to be up-regulated with increasing BMP-2 concentrations. We

hypothesize that in the long-term, it may be possible to reduce the required doses of BMP-

2 if antagonists such as noggin could be at least temporarily removed or knocked down.

One approach to lower noggin levels would be to deliver siRNA against noggin mRNA.

We have investigated this possibility by conducting pilot studies on delivery of siRNA via

non-viral cationic lipid (Lipofectamine) delivered directly from a fibrin surface (surface-

mediated delivery) to MC3T3-E1 pre-osteoblasts that have been stimulated by

supraphysiologic doses of BMP-2. Surface-mediated delivery of siRNA complexes was

confirmed by confocal microscopy and flow cytometry, which led to uptake in 98.5±0.1%

of MC3T3-E1 cells after 48 hours. The internalized siRNA complexes did not affect

MC3T3-E1 viability. A dose-dependent reduction in noggin mRNA levels was achieved

in MC3T3-E1 cells after the cells were cultured with a 10 g/mL dose of BMP-2. Two

treatments, 0.5µg and 1µg noggin siRNA, were able to achieve noggin mRNA levels which

are statistically equivalent to normal MC3T3-E1 noggin mRNA levels (i.e., in cells that

have not been exposed to BMP-2). Surface-mediated siRNA transfection offers an

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efficacious and controlled approach to reduce BMP-2 antagonists which may lower the

required therapeutic dose of BMP-2; thereby reducing negative side effects of BMP-2 such

as ectopic bone growth.

Keywords: Fibrin, natural polymer, gene therapy, mRNA

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1. Introduction

Two million bone graft procedures are performed annually to repair injury to bone

sustained from trauma, tumor resections, and orthopedic surgery [1]. The therapeutic gold

standard uses autologous bone grafts, and allogeneic bone grafts such as demineralized

bone matrix are also widely used. However, due to limited donor tissue and donor site

morbidity (autografts) and concerns about immunological rejection (allografts) a number

of alternative bone grafts have been developed and approved by the FDA [1]. These devices

aim to provide a material component (typically, collagen) that acts as an osteoconductive

matrix for cell attachment and migration. They also provide an osteoinductive mitogenic

agent or growth factor to promote bone regeneration [2].

One growth factor of particular interest for bone regeneration, due to its

osteoinductive capability, is bone morphogenetic protein-2 (BMP-2) [3]. In native bone

regeneration, BMP-2 is uniquely required for initiation of bone healing [4] as well as for

long-term duration of the healing process by its continuous expression [5]. BMP-2 acts as

a chemoattractant of mesenchymal progenitor cells and stimulates their differentiation into

osteoblasts [6]. Administration of exogenous BMP-2 alone can promote the bone healing

process [7], but its short half-life [8], the presence of receptor-binding antagonists [9], and

its potent morphogenetic potential require that it be incorporated into or onto a material to

confine the effects of BMP-2 to the local implant site.

For FDA-approved products, recombinant human BMP-2 (rhBMP-2) is adsorbed

to collagen-based osteoconductive matrices [10]. Unfortunately, the supra-physiological

dose of rhBMP-2 [11] delivered into the local environment from these products has led to

contraindications including heterotrophic ossification, seroma, and edema [12–14]. The

high doses of rhBMP-2 need to overcome the therapeutic threshold stems from endogenous

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self-limiting regulation of BMP-2 expression through extracellular inhibitors and

antagonists [9,15]. Signaling regulators such as noggin are produced by local osteoblasts

and have a high affinity to bind BMP-2 thus preventing it from binding to its receptor [16].

Consequently, BMP-2 efficacy is greatly decreased due to a temporal and spatial increase

of noggin expression as a result of increases in BMP-2 levels [17]. This negative feedback

mechanism provides a way to self-regulate BMP-2 levels under healing conditions and in

clinical examples when rhBMP-2 is delivered exogenously as part of a synthetic bone graft.

Considerable effort, including from our group [18], has been focused on the exploration of

controlling the amount of rhBMP-2 delivered at the implant site either through alternative

carrier systems [19,20] or through the use of other delivery strategies [21–23].

Unfortunately, even with significant reduction of rhBMP-2 concentration, the

amount of rhBMP-2 currently delivered is still higher than physiological levels [11]. This

may be due, in part, to the fact that reducing the amount of rhBMP-2 alone does not address

the underlying role that antagonists, such as noggin, play in mitigating effects of BMP-2.

For these reasons, we are interested in exploring the delivery of small interfering RNA

(siRNA) as an alternative strategy to mitigate the effects of noggin.

Since its discovery, siRNA has created great excitement in the medical community

for its potential in therapeutic applications ranging from anti-cancer therapeutics [24] to

treatment of chronic neurological conditions [25] by targeting specific mRNAs that code

for proteins of interest [26,27]. A key limitation to successful application of siRNA as a

therapeutic strategy has been delivery to the site of action. Systemic delivery is challenging

because siRNA is unstable in blood. This necessitates use of complexing agents to protect

the siRNA and achieve long circulation times. Other challenges include achieving

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localization to the target tissue, localization to the target cells, and achieving cellular uptake

of the siRNA [28,29]. However, for tissue engineering/regenerative medicine applications

these barriers are less problematic as the siRNA can be directly implanted at the site of

action (i.e., tissue in need of repair). Recent progress has led to success in incorporating

siRNA into synthetic [30–33], natural [34,35], and composite [36,37] biomaterials for local

delivery of sequence-specific mRNA targets.

In terms of bone regeneration, siRNA targeting guanine nucleotide-binding protein

alpha-stimulating activity polypeptide 1 (GNAS1) and prolyl hydroxylase domain-

containing protein 2 (PHD2) have been administered directly to mesenchymal stem cells

to promote their differentiation towards an osteogenic phenotype [38]. siRNA has also been

used in directing transcription factor and osteogenic pathway regulation to accelerate bone

fracture healing [38,39]. Noggin has been investigated as a siRNA target due to its well-

known antagonistic role against BMP-2. Several groups have used transfection methods

including non-viral carriers [40], electroporation [41], or viral gene delivery strategies [42].

Others have developed novel carrier systems for siRNA to be delivered in conjunction with

rhBMP-2 [43]. Not only has knockdown of noggin inhibition been shown to increase

osteogenesis but it also lowers the effective rhBMP-2 dose required to elicit a favorable

response [41,43].

In the study reported here, we sought to use an approach in which siRNA could be

delivered locally and directly to cells from the surface of an osteoconductive natural

polymeric biomaterial carrier. A number of natural polymers have been used for bone

regeneration applications due to their inherent ability to promote cell attachment,

biodegradation, and minimal toxicity. These include collagen [44], gelatin [45], keratin

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[18,46,47], and fibrin [48,49]. In these studies, we have used fibrin as the material due to

its role in native bone healing [50], previous use as a BMP-2 carrier for bone regeneration

[48,49], and its suitability for local delivery of gene therapy agents including nucleic acids

[51,52]. Our approach was to complex the siRNA with a commercially available

complexing agent (Lipofectamine, a cationic liposome) adsorbed to the surface of a

biomaterial. We sought to conduct a pilot study to determine: (1) the ability to adsorb

siRNA complexes with a polycationic transfection vehicle to a fibrin hydrogel surface, (2)

the uptake of these complexes into preosteoblast cells seeded onto the surface of the fibrin

hydrogel, and (3) the effects on cell viability and knockdown of the target noggin mRNA

in the preosteoblasts in conjunction with a supraphysiological dose of rhBMP-2 (study

outline summarized in Figure 1).

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Figure 1. Schematic of surface-mediated transfection experimental design.

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2.1 Complexation of siRNA with Polycationic Transfection Vehicles

Several types of siRNA were used for the experiments described here. Accell eGFP

siRNA (Thermo Fisher Scientific, Waltham, MA) was used for complexing assays and

particle size analysis. Accell Control siRNA #1 Non Targeting (Thermo Fisher Scientific,

Waltham, MA) and All Stars Negative Control - Alexa 488 siRNA (Qiagen, Venlo,

Netherlands) were used for non-target controls. Accell SmartPool Noggin siRNA (Thermo

Fisher Scientific, Waltham, MA) was used as the sequence to target noggin mRNA. The

same techniques were used to form complexes with Lipofectamine 2000 (Life

Technologies, Grand Island, NY) for each type of siRNA used. Briefly, 20mM siRNA and

Lipofectamine 2000 transfection agent were diluted into working concentrations

(0.1µg/mL, 0.25µg/mL, 0.5µg/mL, 1µg/mL) with sterile RNase-free water (Thermo Fisher

Scientific, Waltham, MA) under aseptic conditions. siRNA complexes were formed by

using manufacturer’s directions for Lipofectamine 2000. For delivery of 1µg siRNA (n=3),

1.47µL Lipofectamine 2000 was added to RNase-free water for a total volume of 25µL in

a sterile 1.5mL conical tube. In a different sterile 1.5mL conical tube 3.76µL of 20µM

siRNA stock solution was added of a total volume of 25µL of RNase-free water. The

respective Lipofectamine and siRNA solutions were then mixed together and allowed to

complex at room temperature for 20 minute then immediately used for subsequent

experiments. For lower siRNA doses, Lipofectamine and siRNA volumes were adjusted

accordingly. The Lipofectamine-siRNA complexes were measured for particle diameter

by dynamic light scattering (DLS). For these measurements, the particles were measured

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by dynamic light scattering at an siRNA concentration of 1 g/mL and measured on a

Zetasizer, Nano (Malvern, Worcestershire, United Kingdom).

2.2 Fibrin Hydrogel Preparation

Fibrinogen powder from bovine plasma (Sigma-Aldrich, St. Louis, MO) was

dissolved with sterile RNase-free water under asceptic conditions in a 37˚C water bath to

achieve a 200mg/mL fibrinogen solution. Fibrinogen solution was used to coat the bottom

of glass coverslips (50 µL), 48 (25 µL) well plates, 24 (50 µL) well plates, or 6 (240 µL)

well plates depending on the experiment. One part fibrinogen solution was mixed with

three parts thrombin working solution which consists of 13 parts PBS to one part 5.5mg/mL

CaCl2 (Thermo Fisher Scientific,Waltham, MA) and one part 250U/mL bovine plasma

thrombin (Sigma-Aldrich, St. Louis, MO). The resulting 5% (w/v) fibrin hydrogel was

allowed to polymerize overnight at 37°C in a cell incubator (humidified atmosphere of 5%

CO2 /95% air).


Scanning electron microscopy was used to assess siRNA complex adsorption and

particle diameter after adsorption to fibrin hydrogels. Fibrin hydrogels were fabricated as

described above within a 24 well plate. A 5mm biopsy punch was taken and placed into a

1.5mL conical tube. Sterile RNase-free water containing Lipofectamine-siRNA complexes

prepared as described above were added to the top of the fibrin hydrogels and allow to

adsorb for 4 hours. RNase-free water and siRNA alone (not complexed with Lipofectamine

2000) served as controls. Hydrogels were then taken through alcohol dehydration steps,

critical point dried, and gold sputter coated. Adsorbed siRNA complexes were imaged with

Zeiss Supra 35 VP FE-SEM (Jena, Germany) at 5,000 x and 15,000 x magnification. Five

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images were taken of separate areas of the fibrin surface at 15,000 x magnification were

imaged and Lipofectamine-siRNA complexes were measured for particle diameter in

Image J software.


To determine the stability of siRNA complex adsorption to the fibrin hydrogels, we

conducted a desorption (release) study in a 37°C in a cell incubator (humidified atmosphere

of 5% CO2 /95% air). Fibrin hydrogels were fabricated as described above in a 48 well

plate. Lipofectamine 2000 complexed with 0.5µg Alexa 488 siRNA in sterile RNAase-free

water was allowed to adsorb to the fibrin gel surface for 4 hours. The hydrogel was washed

once with 200uL of PBS and then added to a 1.5mL conical tube to quantify the amount of

siRNA that did not absorb to the surface. 200uL of fresh PBS was added to each well and

at preassigned time points (1.5, 3, 6, 12, 24, 48, 72 hours) 100uL was removed from each

well for analysis and was replaced with 100uL of fresh PBS. All collected samples were

frozen at -20°C until thawed for analysis. A final concentration of 1% TritonX-100 (Sigma

Aldrich, St. Louis, MO) and 2% SDS (Sigma Aldrich, St. Louis, MO) was added to each

sample in order to release the Alexa 488 siRNA from the Lipofectamine (decomplexation).

Samples were added to a 96-well black plate and fluorescence readings were made on a

BioTek platereader at 485nm/525 nm (Ex/Em).

2.5 MC3T3-E1 Cell Culture

Mouse preosteoblast MC3T3-E1 Subclone 4 cells (ATCC® CRL2593™;

Manassas, VA) were cultured as recommend in complete growth media; Alpha Minimum

Essential Medium (α-MEM) supplemented with 10% Fetal Bovine Serum, and 1%

Antibiotic-Antimycotic (Gibco, Life Technologies Grand Island, NY) at 37°C in a cell

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incubator (humidified atmosphere of 5% CO2). Cells were cultured on 150mm cell culture

plates (Thermo Fisher Scientific,Waltham, MA) until 60-70% confluent. At this point

media was removed, cells were washed with 10mL PBS, and detached with 0.25% Trypsin-

EDTA (Gibco, Life Technologies Grand Island, NY) for 5 minutes in the cell culture

incubator. Complete growth medium (5mL) was added and cells were centrifuged at 1500

rpm for 5 minutes. Cells were re-suspended and plated 1:8 with complete growth media.

In the following experiments, “transfection media” refers to α-MEM supplemented with

10% FBS alone without the addition of Antibiotic-Antimycotic.

2.6 Flow Cytometry

Flow cytometry was used to assess the uptake of siRNA complexes (or

uncomplexed siRNA control) in cells. For these studies siRNA labeled with Alexa 488

was used to allow detection by the fluorescence detectors. Fibrin hydrogels were coated

on the bottom of a 6 well plate as described earlier and allowed to polymerize overnight.

After one wash in PBS, Alexa 488 siRNA complexes were adsorbed to the hydrogel surface

as before. No siRNA (water only) or uncomplexed siRNA (no Lipofectamine 2000) served

as controls. 300uL containing 0.5ug Alexa 488 siRNA (in Lipofectamine 2000 or

uncomplexed) was allowed to adsorb to the fibrin hydrogel surface for 4 hours. At this

time, siRNA solutions were removed and the hydrogel surface was washed once with PBS.

Immediately afterwards 40,000 MC3T3-E1 cells were seeded to the hydrogel surface with

300 µL transfection media (αMEM with 10%FBS) for 4 hours. The transfection media was

removed and the hydrogel surface was washed once with PBS. Then 1mL of culture media

(αMEM; 10%FBS;1%A-A) was added to each well and cells were placed into culture for

for 48 hours. After 48 hours, media was removed and cells were washed once with PBS

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and detached from the hydrogel surface by two 5 minute washes of 0.25% trypsin (Gibco,

Life Technologies Grand Island, NY). Wells were washed once with culture media to

ensure collection of cells. Cells were collected into 15mL conical tubes and centrifuged for

5 minutes at 1500 rpm, the supernatant was removed, cells were washed in 1mL of PBS

and centrifuged again, the PBS supernatant was removed, and the cell pellet was suspended

in 500 L of sterile BD FACsFlow sheath fluid (BD, Franklin Lakes, NJ). Each 500 µL

cell suspension was then pipetted through a 0.4 µm cell strainer (BD, Franklin Lakes, NJ)

into a polystyrene flow cytometer tube (BD, Franklin Lakes, NJ) and placed on ice. Cell-

associated Alexa 488 siRNA uptake was then measured with a BD FACScan (BD, Franklin

Lakes, NJ). Cells only (without Alexa 488 siRNA) treatment served as negative controls

to determine acquisition parameters and establish gating regions. BD Cell Quest Pro

software (BD, Franklin Lakes, NJ) was used for data acquisition and analysis.


Confocal microscopy was used to assess cellular uptake and distribution of siRNA

in MC3T3-E1 cells. 22mm round glass coverslips were thinly coated with fibrin hydrogels

as described above and allowed to polymerize overnight in a 6 well plate. 0.5ug Alexa 488

siRNA was complexed with Lipofectamine 2000 as described above. Fibrin hydrogels

washed with sterile RNase-free Sterile RNase-free water (no siRNA) or uncomplexed

Alexa 488 siRNA (no Lipofectamine 2000) suspended in sterile RNase-free served as

controls. 150µL of solution (water, siRNA alone, or siRNA complexed with

Lipofectamine) was added drop-wise to the top of the fibrin-coated coverslip and allowed

to adsorb for 4 hours. After 4 hours, the coverslips were washed with 200µL sterile PBS.

MC3T3 were passaged in transfection media (αMEM; 10%FBS;0%A-A) and 20,000 cells

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(150uL transfection media) were seeded onto each coverslip. The cells were allowed to

attach for 4 hours after which the transfection media was removed and replaced with

normal culture media. After 48 hours coverslips were washed with PBS, fixed in 10%

formaldehyde (methanol free) for 10 minutes. Cells were counter-stained for F-actin with

an Alexa 555 phaoloidin dye (Invitrogen, Carlsbad, CA) and for nuclei with 300nM DAPI

(Invitrogen). Coverslips were then mounted to glass slides with MM83 mounting media

(Leica,Wetzlar, Germany) and allowed to dry overnight in the dark at 4°C. Confocal

imaging (Ziess 710 Laser Scanning Confocal System, Jena, Germany) of experimental

groups was performed at 20x magnification. The Zeiss LSM Image Browser (Zeiss, Jena,

Germany) was used for image processing.

2.8 rhBMP-2 Dose Effect on MC3T3 Noggin Expression

It is known that noggin expression is up-regulated in response to increased BMP-2

concentration. We assessed expression of noggin mRNA in response to increasing doses

of BMP-2 to ensure appropriate cell response to BMP-2. Fibrin hydrogels were coated on

24 well plates as previously described and allowed to polymerize overnight at 37°C

followed by a RNase-free water wash. MC3T3-E1 passaged with transfection media were

seeded at 20,000 cells/well and allowed to adhere for 4 hours. The transfection media was

then removed and replaced with 200uL of culture media supplemented with varying doses

of rhBMP-2: 0, 0.001, 0.01, 0.1, 1, and 10 µg/mL rhBMP-2 (n=3 for each). After 24 hours

the cells were harvested with 0.25% trypsin by two 5 minute incubations at 37°C and then

washed with culture media. Trypsin and media washes were collected into 15mL conical

tubes and centrifuged for 5 minutes at 1500 rpm. Supernatant was removed, cells were

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resuspended in 500µL PBS, and centrifuged for 5 minutes at 1500rpm. Supernatant was

removed and cell pellet was frozen at -20°C until RT-qPCR was performed (section 2.11).

2.9 Cell Activity (MTS Assay)

MTS assay was used to determine effects of the siRNA and siRNA complexes on

cell viability. These viability assays were conducted in the presence of rhBMP-2 (used in

this section 2.9 and section 2.10) to achieve up-regulation of noggin mRNA expression.

Fibrin hydrogels were allowed to form overnight at 37˚C within a 48 well dish as described

above. Sterile RNase-free water containing Lipofectamine 2000 (complexed with either

0.25 µg Noggin (NOG) siRNA or Non-Targeting (NT) siRNA as previously described) and

was allowed to adsorb to the fibrin gel surface for 4 hours (n=3). Uncomplexed (0.25 µg

NOG or NT siRNA) and sterile RNase-free water served as negative controls (n=3). After

4 hours, wells were washed with sterile PBS and 10,000 MC3T3-E1 were seeded onto each

well with transfection media. After another 4 hours, transfection media was removed and

replaced with 100µL culture media alone or containing 10µg/mL rhBMP-2. Cells were

incubated for 48 hours at which point the culture media was removed and replaced with

200µL of CellTiter 96® Aqueous Non-Radioactive Cell Proliferation Assay (Promega,

Madison, WI) for 1 hour in a humidified 37°C, 5% CO2 incubator using manufacturer’s

directions. Reaction was stopped by lysis of cells with a 10% SDS solution. 200µL of the

cell lysate was transferred to a 96 well plate and 490nm absorbance readings were

immediately taken and correlated with a standard curve of known MC3T3-E1 cell number

absorbance values. Cell numbers were normalized to MC3T3-E1treated with no siRNA

and 10µg/mL rhBMP-2 (± SEM).

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2.10 siRNA Dose Effect on MC3T3 Noggin Expression

Fibrin hydrogels were coated on 24 well plates as previously described and allowed

to polymerize overnight at 37°C followed by a RNase-free water wash. Lipofectamine was

complexed as previously described with either 0.5µg NT (non-target control), 0.5µg Alexa

488 (non-target control), or NOG siRNA at various doses per well: 0µg, 0.1µg, 0.25µg,

0.5µg, 1µg. Water, 2.2µLLipofectamine alone (vehicle-only concentration which would

complex 0.5µg siRNA), or uncomplexed NOG siRNA served as negative controls. Each

treatment (or control) was allowed to adsorb to the fibrin surface for 4 hours in RNAase-

free water. After 4 hours, wells were washed with sterile PBS and 20,000 MC3T3-E1 were

seeded onto each well with transfection media. After another 4 hours transfection media

was removed and replaced with 200µL culture media alone or containing 10µg/mL rhBMP-

2. Cells were then incubated for 48 hours at which time they were harvested with 0.25%

trypsin by two 5 minute incubations at 37°C and washed with culture media. Trypsin and

media washes were collected into 15mL conical tubes and centrifuged for 5 minutes at

1500 rpm. Supernatant was removed, cells were re-suspended in 500µL PBS, and

centrifuged for 5 minutes at 1500 rpm. Supernatant was removed and cell pellet was frozen

at -20°C until RT-qPCR was performed.

2.11 RT-qPCR

RNA extraction was performed by using Maxwell 16 LEV simplyRNA Tissue Kit

(Promega, Madison, WI) followed by RNA quantification by using a NanoDrop 2000

(Thermo Scientific, Wilmington, DE). 50ng of sample RNA and immediately reverse

transcribed into cDNA by Omniscript RT kit according to the manufacturer’s instructions

(Qiagen, Venlo, Netherlands) and frozen at -20° C until analyzed by qPCR. cDNA samples

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were mixed with QuantiTech SYBR Green PCR kit (Qiagen, Venlo, Netherlands) and

appropriate primers (QuantiTech Primer Assay, Qiagen, Venlo, Netherlands) targeting

mouse Noggin (QT00256585), Beta-Actin (QT00095242), and GAPDH (QT01658692).

Quantative PCR was performed on a Bio-Rad iCycler (Bio Rad, Hercules, CA) for 40

cycles. Data were analyzed using Biogazelle QBase+ software (Zwijnaarde, Belgium) and

noggin expression was normalized to normal MC3T3-E1 without rhBMP-2 treatment.


All statistical analysis was performed by using Minitab 16 (Minitab, State College,

PA). Normalized values (± SEM) were compared using One-Way ANOVA using Tukey’s

post hoc test.

3. Results

3.1 siRNA Complex Adsorption and Size Analysis

Cellular uptake of Lipofectamine complexes (and other non-viral gene delivery

vehicles) is based partly on the size of the complex. To ensure that the adsorption process

did lead to aggregation of the siRNA complexes, we assessed particle diameter by DLS

before adsorption and by SEM image analysis after adsorption. DLS of the Lipofectamine

complexes showed particles of 128nm±30nm (mean diameter ± STDEV). SEM images of

the surface of the fibrin hydrogel at (5k x magnification) in the presence of RNase-free

water (no siRNA; Figure 2A), uncomplexed siRNA (Figure 2B), and Lipofectamine-

siRNA complexes (Figure 2C) are shown after 4 hours of adsorption. As expected, no

complexes were observed when water only or when uncomplexed siRNA were used.

Lipofectamine-siRNA complexes were adsorbed to the fibrin surface (Figure 2C, arrows).

ImageJ measurements of the adsorbed complexes were 140±28nm (mean diameter ±

STDEV) (Figure 2C insert) with minimal aggregation of the complexes observed. It

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therefore appears that adsorbtion of Lipofectamine-siRNA complexes to the fibrin

hydrogel did not affect the diameter size as determined by comparison to the DLS

measurements of complexes in solution.

Figure 2. FE-SEM images and size characterization of surface adsorbed

Lipofectamine siRNA complexes to a fibrin hydrogel surface. FE-SEM images of the

fibrin hydrogel surface after being treated for 4 hours with: (A) RNase-free water, (B)

uncomplexed siRNA, or (C) Lipofectamine siRNA-GFP complexes (arrows). ImageJ

measurement of the diameter size distribution of the adsorbed siRNA complexes showed

complex diameters centered on 200nm (5k x magnification; scale bar denotes 1µm). (D)

ImageJ particle size analysis of adsorbed Lipofectamine siRNA-GFP complexes on the

fibrin surface correlates with DLS analysis of solution suspension of Lipofectamine siRNA

nanoparticles (insert).

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3.2 Release Kinetics

The goal of our approach is to achieve direct delivery of siRNA to cells via surface-

mediated uptake when the cells contact the biomaterial. To assess the stability of the

adsorbed siRNA, we performed an experiment with fluorescently-labeled siRNA to

measure the amount of siRNA eluting from the surface after adsorption. The amount of

siRNA not eluted was therefore the amount remaining stably associated with the

biomaterial surface. The rate of siRNA complex release was measured over the course of

48 hours, which exceeds the length of time used for the cell studies reported below. Figure

3A shows the release profiles of siRNA from the fibrin surface when complexed with

siRNA. After three days only 2% of the siRNA was released, and most of the release

occurred after an initial short burst in the first several hours, likely due to some complexes

that were not strongly adsorbed. Thus 98% of Lipofectamine siRNA complexes remained

adsorbed to the fibrin hydrogel surface after 72 hours, indicating stable adsorption.

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Figure 3. Release kinetics of Lipofectamine siRNA complexes from fibrin surface.

Release kinetics of Lipofectamine siRNA-AF488 complexes from fibrin hydrogel (n=3,

error bars denote ±STDEV).

3.3 Flow Cytometry

In order to successfully knock down noggin, we reasoned that a large percentage of

the cells would require uptake of the Lipofectamine-siRNA complexes. To quantify the

number of cells with associated siRNA, we performed flow cytometry on the cells after

culture on the fibrin surfaces in the presence of fluorescently-labeled siRNA. We used

cells cultured on fibrin without siRNA to gate (Figure 4A) and compared cell-association

with uncomplexed siRNA (Figure 4B) and Lipofectamine-siRNA complexes (Figure 4C).

The number of cells falling into the gated region was measured after 48 hours of culture

with the fibrin surfaces containing siRNA and the number of cells (“events”) was

determined by using the appropriate fluorescence channel for AF488. Results show that

48 hours after surface-mediated delivery of Alexa 488 siRNA, only 7.18±0.62% of cells

had taken up the siRNA and the fluorescence intensity in these cells was low. In contrast,

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98.45±0.06% of cells were positive for Alexa 488 siRNA that had been complexed with

Lipofectamine. These results demonstrate the importance of complexation for surface-

mediated uptake of the siRNA by MC3T3-E1 cells.

Figure 4. Transfection efficacy of Lipofectamine siRNA complexes into MC3T3-E1.

Transfection efficiency was quantified using flow cytometery 48 hours after seeding on

fibrin hydrogels. (A) Normal MC3T3-E1 were seeded on fibrin only to serve as a negative

control. (B) Uncomplexed siRNA-AF488 or (C) Lipofectamine siRNA-AF488 complexes

were then measured to record the percentage of cells which expressed AF488 (n = 3; mean

± SEM).


To further assess the distribution of siRNA in MC3TE-E1 cells following uptake,

we used confocal microscopy. Fluorescently-labeled siRNA (Alexa 488 siRNA) and

MC3T3-E1 cells were used in the same manner as in the flow cytometry study. Phalloidin

was used to label actin filaments to demark intracellular siRNA and cells were also

counterstained with DAPI for nuclear staining. Because confocal, rather than

epifluorescence, microscopy was used, we were able to ensure that observed siRNA was

inside the cell rather than on the fibrin surface underneath the cell. Figure 5A shows

MC3T3-E1 cells in the absence of siRNA. Figures 5B (uncomplexed siRNA) and 5C

(siRNA complexed with Lipofectamine) show no noticeable change in cell morphology or

actin filament arrangement. Uncomplexed Alexa 488 siRNA was not observed to be

internalized by the MC3T3 (Figure 5B) while surface-mediated delivery with

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Lipofectamine-siRNA complexes showed clear uptake and internalization (Figure 5C) of

the Alexa 488-Lipofactamine complexes (green arrow) after 24 hours. Qualitatively, the

results are consistent with the flow cytometry data where all cells had some level of siRNA

uptake when delivered with Lipofectamine, and these cells clearly had greater fluorescence

signal than when siRNA was uncomplexed.

Figure 5. MC3T3-E1 internalization of Lipofectamine siRNA complexes. Confocal

images of MC3T3 were taken 48 hours after seeding on fibrin hydrogels (20x

magnification; scale bar denotes 100 µm) to visualize (A) cells alone, (B) the inability to

internalize uncomplexed Alexa 488 siRNA, or (C) internalization of surface adsorbed

Lipofectamine – Alexa 488 siRNA complexes (green arrows).

3.5 rhBMP-2 Dose Effect on MC3T3 noggin Expression

Noggin expression is up-regulated in response to increases in BMP-2. We

performed RT-qPCR on noggin expression in response to increasing BMP-2 doses to

confirm this up-regulation in our hands and also to determine the concentration of rhBMP-

2 to use for the knockdown experiments (section 3.7). For these experiments, increasing

concentrations of rhBMP-2 given in solution were used. RT-qPCR values were normalized

to MC3T3-E1 that were not treated with rhBMP-2 (0µg/mL). After 24 hours,

129

concentrations above 0.01µg/mL rhBMP-2 resulted in significantly increased levels of

Noggin mRNA expression in MC3T3-E1 cells (Figure 6). Each 10x increase above

0.001µg/mL rhBMP-2 resulted in a statistically significant difference in Noggin mRNA

between respective dose levels. The greatest noggin mRNA expression was 20.82 ± 1.47

fold, which resulted from the bigest rhBMP-2 concentration (10µg/mL). Therefore, we

chose to use 10µg/mL rhBMP-2 for subsequent experiments on noggin knockdown with

surface-mediated delivery of siRNA.

Figure 6. Noggin mRNA expression changes with rhBMP-2 dose. Noggin Expression

Levels with BMP-2 Dose 48 hours after MC3T3-E1 were seeded on fibrin hydrogels.

Values were normalized to basal rhBMP-2 untreated MC3T3-E1 (0µg/mL BMP-2) Noggin

mRNA expression levels. Error bar denotes Relative Expression ± SEM; *p < 0.05 vs

0µg/mL BMP-2, **p < 0.05 between treatment groups.

3.6 MTS Assay

MTS assay was performed 48 hours after MC3T3 were seeded on fibrin hydrogels

(Figure 7). These experiments were conducted after delivery of rhBMP-2 at 10 g/mL.

130

Cell viability values were normalized to MC3T3 cells treated with 10µg/mL rhBMP-2 only

(no siRNA). MC3T3 cells without treatment (no siRNA, no rhbMP-2) did not show a

difference in metabolic activity compared to the MC3T3 cells treated with 10µg/mL

rhBMP-2. Uncomplexed NOG siRNA and NT siRNA as well as NOG siRNA and NT

siRNA complexed with Lipofectamine did not have a significant effect on the viability of

MC3T3-E1 cells.

Figure 7. MC3T3-E1 cytotoxic effects of Lipofectamine siRNA complexes. MTS assay

was preformed 48 hours after MC3T3-E1 were seeded on fibrin hydrogels with or without

adsorbed 0.25µg siRNA alone or complexed with Lipofectamine 2000. MTS reaction was

incubated for one hour at 37°C before the reaction was stopped by lysing cells. Absorbance

readings were correlated to average cell number and then normalized to MC3T3 treated

with no siRNA; 10µg/mL rhBMP-2. Error bars denote ± SEM; no statistical differences

were measured.

3.7 siRNA Dose Effect on MC3T3 Noggin Expression

We sought to determine the ability to achieve knockdown of noggin expression up-

regulated by the presence of 10 g/mL of rhBMP-2. Noggin expression in MC3T3-E1

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cells was assessed by RT-qPCR 48 hours after delivery of 10 g/mL of BMP-2 when the

cells were cultured in the presence of siRNA. siRNA for NOG was either uncomplexed or

complexed with Lipofectamine. Several control siRNA groups were used. A non-target

(NT) sequence was used as was an Alexa 488 siRNA sequence. Both were used in

uncomplexed form or complexed with Lipofectamine. Lipofectamine only (no siRNA;

vehicle control) was also used. Basal Noggin mRNA expression of MC3T3-E1 not

supplemented with rhBMP-2 (0 µg/mL rhBMP-2) showed 91.5 ± 3.81% lower Noggin

mRNA expression than MC3T3-E1 cells treated with 10µg/mL rhBMP-2, consistent with

the results shown in Figure 7 but in this case at a 48 hour (instead of 24 hour) time point.

As shown in Figure 8A, 0.5µg of uncomplexed siRNA-NOG, Lipofectamine vehicle only

(no siRNA), and 0.5 µg siRNA-AF488 complexed with Lipofectamine expressed

statistically equivalent noggin mRNA to cells treated with 10µg/mL rhBMP-2 only. The

NT siRNA complexed with Lipofectamine led to a 20% decrease in noggin expression,

which we interpret to be off-target effects, though this was not statistically different than

the MC3T3-E1 cells receiving 10 g/mL rhBMP-2 with no siRNA treatment. In contrast

the MC3T3-E1 cells culture in the presence of 0.5 g NOG siRNA complexed with

Lipofectamine led to a significant decrease in noggin expression compared to cells treated

with only 10 g/mL rhBMP-2. Thus knockdown was approximately 80.4%.

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Figure 8. MC3T3-E1 noggin mRNA levels with rhBMP-2 challenge after surface-

mediated transfection of Lipofectamine siRNA complexes. MC3T3-E1 Noggin mRNA

expression levels with rhBMP-2 challenge after surface-mediated transfection was

quantified by qRT-PCR after 48 hours (n=3; Mean relative quantity ± SEM). Noggin

mRNA levels were normalized to 10µg/mL BMP-2; no siRNA. (A) Control

groups.*p<0.05 versus Normal, **p<versus 0.5µg siRNA-NT (B) Lipofectamine siRNA-

NOG complex dose response. ap<versus Normal, bp<versus 10µg/mL BMP-2; no siRNA, cp<0.05 versus 0.25µg siRNA-NOG.

We also conducted a dose-response experiment for a range of NOG siRNA

concentrations (Figure 8B). There is a clear trend: with increasing the amount of surface-

adsorbed Lipofectamine-siRNA-NOG complexes there is an increase knockdown of

Noggin mRNA after rhBMP-2 challenge in MC3T3-E1 cells. Surface-mediated delivery

of 0.1µg, 0.25µg, 0.5µg, and 1µg siRNA-NOG respectively resulted in 26.95±2.68%,

42.63±3.72%, 80.4±19.59%, and 90.61±0.58% knockdown of Noggin mRNA when treated

with 10µg/mL rhBMP-2 for 48 hours. MC3T3-E1 cells treated with 0.5µg or 1µg of

siRNA-NOG and challenged with 10 µg/mL rhBMP-2 had mRNA levels of noggin reduced

to those statistically equivalent to noggin mRNA expression in cells not challenged with

rhBMP-2 (0µg/mL rhBMP-2). This demonstrates at the mRNA level that noggin

expression can be reduced to near basal levels via siRNA delivery in a surface-mediate

fashion.

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4. Discussion

The use of FDA-approved rhBMP-2 devices to enhance bone regeneration has

increased over the past decade, bringing to light a number of drawbacks. First, the initial

cost of single treatment ($5,000-$8,000 USD) is due, in part, to the cost associated with

production of rhBMP-2 to be delivered (about 1000 times more than present naturally in

the body) [11]. This supraphysiological rhBMP-2 dose directly leads to the larger

secondary problems of adverse side effects such as edema and ectopic bone growth [53].

In order to improve current rhBMP-2 devices, an understanding of the intrinsic controls of

bone regeneration must be taken into account. The regenerative potential of bone at a site

of injury is often diminished by locally expressed inhibitory molecules such as the BMP-2

antagonist noggin. The current clinical strategy to overcome the therapeutic threshold set

by BMP-2 inhibitors is to incorporate supra-physiological concentrations of rhBMP-2 into

the device. Previous studies, including our own [18], have focused on lowering loading

concentrations or using alternative biomaterials to control the rate BMP-2 release from the

carrier system [19,46,54].

While this emphasis on the delivery of molecules such as growth factors are pro-

regeneration (or promote regeneration) is common and rational for tissue engineering, the

approach described here is fundamentally different. That is, this approach seeks to promote

tissue regeneration by delivery of molecules such as siRNA that can knockdown molecules

typically inhibitory to regeneration. In this case, we investigated the ability to knock down

the BMP-2 antagonist noggin through the use of siRNA. Because molecules that promote

(bone) tissue formation such as rhBMP-2 can be adsorbed to biomaterials for delivery to

cells, we were interested in investigating this approach for the delivery of siRNA.

134

In order for this approach to be effective several criteria had to be met. First, we

required a material to which we could adsorb siRNA complexes. In terms of the polymer

carrier, we investigated fibrin for these studies. Fibrin is a native biopolymer in wound

healing and functions to provide a provincial matrix for cell infiltration prior to production

of type 1 collagen, the main extracellular matrix component of bone. The capacity of fibrin

to adsorb 6.7 times more serum proteins (including fibronectin and heparin) than collagen

aids in fibrin’s role in adhesion, proliferation, differentiation, and maturation of osteoblasts

[55,56]. Fibrin is a negatively charged fibrillar protein at neutral pH [57]. We can take

advantage of the overall positive charge of the siRNA complexed with the polycationic

transfection vehicles to adsorb the nanoparticles to the fibrin hydrogel. In principle this

approach could be used with any polymeric material bearing a negative surface charge at

neutral pH, though we have had previous success with fibrin for the delivery of plasmid

DNA complexed with PEI with an overall positive zeta potential [51].

A second criteria that we investigated was the ability of the siRNA complexes to

remain adsorbed to the fibrin surface. This is necessary to allow cells to take up the

complexes in a surface-mediated fashion. Adsorption of the complexes to fibrin did not

greatly alter their diameters compared to their diameter in solution (determined by DLS),

remaining at ~ 100-200nm in diameter, which is of a sufficiently small diameter to allow

for endosomal uptake by cells [58]. Investigation into siRNA complex desorption from the

fibrin hydrogel surface showed the majority of Lipofectamine-siRNA (fluorescently

labeled with Alexa 488) complexes remained on the hydrogel surface after 3 days. Due to

the limited release of complexes these findings suggest a method of surface-mediated

transfection (also known as reverse transfection or solid phase delivery) [59] over the

135

traditional solution-phase (“forward”) transfection. Surface-mediated transfection of

nucleic acids has shown the benefits of reduced mass transport and minimal transfection

agent aggregation, thereby having greater transfection efficacy [60] while using less

material. This approach is akin to growth factor delivery from material surfaces and

therefore of potential use in tissue engineering/regenerative medicine applications for local

implantation of biomaterial-based constructs.

We also reasoned that it would be necessary for nearly all cells to take up the siRNA

complexes in order to achieve knockdown. If a small fraction of the cells were to take up

the complexes, the overall effect might be great on those cells, but minimal for the overall

local cell population. Results of the flow cytometry study show that virtually all cells has

siRNA associated with them (98%) when the siRNA was complexed with Lipofectamine.

This is in contrast to uncomplexed siRNA, which showed only 7% of cells in the gated

region with uptake and all at a relatively low signal for Alexa 488. This suggests that the

coating of the surface with the complexes was effective for mediating delivery to cells. We

investigated whether the cells were actually internalizing the siRNA through confocal

microscopy. These images clearly demonstrated intracellular siRNA when delivered via

Lipofectamine complexes as evidenced by their presence within the actin-stained region.

The siRNA was not located in the nucleus, but we cannot rule out the possibility that they

were located in the endosomes given the punctate staining patterns observed. Nonetheless,

these confocal images were taken after 4 hours of cell incubation, allowing sufficient time

for endosomal escape to occur to before the 48 hour time point of the RT-qPCR studies.

Effective nucleic acid delivery through transfection reagents is often sacrificed in

order to minimize cellular toxicity [61]. We therefore investigated whether the cellular

136

uptake led to observable toxicity on cells. Even though 98.45±0.06% of MC3T3-E1 were

positive for siRNA uptake when complexed with Lipofectamine (flow cytometry results),

this did not lead to observable or statistically significant reductions in cell viability as

determined by the MTS assay. This may be because the amount of siRNA delivered to the

cells via the surface-mediated uptake while efficient, may be lower than observed for

solution-phase non-viral transfection. The time frame of uptake by the surface-mediated

approach would also differ from solution-phase transfer. Internalization of siRNA would

not only occur at the time of initial cell attachment to the hydrogel surface, but would

essentially be “re-transfected” as the cells divide and/or migrate on the fibrin surface to

areas previously unoccupied by cells but still containing siRNA complexes. Not delivering

the entire siRNA dose at once, as with solution-phase transfection, but throughout time not

only allows the cells to recover but negates the need for retreating the cells every few days

to maintain adequate knockdown levels. Most commercially available transfection vehicles

alone using traditional solution-phase transfection is highly dependent on cell type and on

average achieve ~70% transfection. In difficult to transfect cell lines and primary cells the

transfection efficacy decreases to ~30%. The surface-mediated approach has been

recognized for its benefits in cell microarray technology [62] but has seen limited success

being translated to tissue engineering/regenerative medicine applications. Our technique

offers reverse transfection method that can be easily applied to a number of tissue

engineering applications, including our primary target is this manuscript: bone

regeneration.

In our present study, the MC3T3-E1 pre-osteoblasts attached to a fibrin hydrogel

up-regulate noggin mRNA expression levels with increasing amounts of BMP-2 as

137

expected [17]. This may be akin to the mechanism by which the body responds to

supraphysiological doses of rhBMP-2 in current technology that leads to ectopic bone

formation. After 48 hours, siRNA targeting against noggin expressed a dose-dependent

reaction in noggin mRNA levels with the highest surface concentration of Noggin siRNA

causing the greatest amout of Noggin mRNA knockdown (Figure 8). Interestingly, the two

highest Noggin siRNA surface concentrations (0.5µg and 1µg /well) reached statistically

equivalent Noggin mRNA levels to MC3T3-E1 cells which were never exposed to BMP-

2. Uncomplexed (naked) siRNA-NOG, Lipofectamine transfection reagent alone, and

0.5µg of siRNA-AF488 did not have an effect on Noggin mRNA expression. Even though

off-target effects of siRNA-NT were observed [63–65], the 0.5µg siRNA-NT dose did not

achieve knockdown levels as the equivalent siRNA-NOG dose. Rather, the siRNA-NT

was statistically similar to the 0.1µg siRNA-NOG concentration.

One potential concern related to this study is the use of Lipofectamine, which is not

recognized as suitable for systemic delivery of nucleic acids such as siRNA [66]. This is

because Lipofectamine is rapidly cleared when delivered systemically. However, for this

local delivery system, this cationic liposome system should be suitable for in vivo use. We

do note that two other commercially available cationic materials (Dharmafect and linear

polyethylenimine) were investigated but did not achieve the levels of knockdown of noggin

mRNA expression observed with Lipofectamine as the complexing agent.

Noggin regulation through siRNA delivery has been successful previously [40–43],

but to our knowledge our work is the only studies to show a controlled dose-dependent

regulation of noggin mRNA which we attribute to the high surface-mediated transfection

efficiency seen in our system. This dose behavior is of particular interest due to the

138

dualistic effects of both noggin and BMP-2 on osteogenic behavior of a number of cell

types. Even though BMP-2 induces differentiation in MSCs, pre-osteoblasts, and immature

osteoblasts [67] there are recent findings that show BMP-2 induces apoptosis in mature

osteoblasts (but not MSCs) in a dose dependent manner [68]. Along these lines, up-

regulation of noggin inhibits apoptosis caused by BMP-2 [69]. On the other hand,

complete knockdown of noggin decreases BMP-2 induced osteogenic differentiation of

human MSCs [70]. Therefore, it is suggested that some noggin expression is necessary

for suitable and controlled bone regeneration. Taken together, there must be a balance

struck between noggin and BMP-2 regulation. This siRNA-based approach offers that

possibility given that the siRNA knockdown is temporary. siRNA provides a short-term

silencing of target genes for about one week, which correlates to the time when a

supraphysiological burst release of rhBMP-2 is observed with current FDA approved

collagen devices. Surface-mediated siRNA transfection would provide a finite controlled

amount of noggin silencing which would taper-off with time thus allowing the native

Noggin-BMP-2 control mechanism to be slowly reset to normal physiological function.

Once the siRNA-NOG is depleted, noggin expression can return to thus control BMP-2

levels and prevent adverse effects like heterotopic ossification. Although we have not

investigated the translational aspects of mRNA knockdown on protein expression, this

study provides convincing evidence for the mechanism of siRNA delivery and the large

amount of mRNA knockdown that can be achieved.

139

Conclusion

Our method to locally deliver regulatory siRNA complexes which are adsorbed to

a fibrin hydrogel surface offers a simple, non-toxic method of surface-mediated delivery

resulting in nearly 100% cellular uptake in MC3T3-E1 cells. Due to this high level of

cellular uptake we were able to achieve controlled noggin mRNA knockdown with varying

concentrations of siRNA-NOG, including reaching basal levels of noggin mRNA in BMP-

2 treated MC3T3-E1 cells with the two highest surface concentrations. Ultimately, this

approach may offer the potential to use reduced levels of rhBMP-2 for in vivo bone

regeneration via knockdown of one BMP-2 antagonist.

Acknowledgments

The authors acknowledge assistance from the Center for Nanotechnology and

Molecular Materials at Wake Forest University (SEM imaging), Dr. Richard Edelmann and

Matthew Duley at the Miami University Center for Advanced Microscopy and Imaging

(confocal microscopy), and Dr. Andor Kiss at the Miami University Center for

Bioinformatics and Functional Genomics (RT-qPCR). This work was partially supported

by the National Institutes of Health (JMS; R01AR061391) and the content is solely the

responsibility of the authors and does not necessarily represent the official views of the

National Institutes of Health. This work was also partially supported by The Wake Forest

University Health Sciences Translational Sciences Institute Pilot Grant and The Wake

Forest Graduate School.

140

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CHAPTER VI

CONCLUSIONS AND PROSPECTUS


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1. Summary of Findings

As the field of regenerative medicine progresses, research must take into

consideration the endogenous factors that influence wound healing. Continued

understanding of developmental biology, wound healing, and normal physiology can be

brought together in order to develop novel tissue engineered products. Overall, this thesis

work examined the ability of natural polymer carriers to locally deliver pro-regenerative

therapeutics. Biomaterial carriers fabricated from polymers naturally produced by the

human body are of particular interest because they are readily recognized by infiltrating

cells through cell-binding motifs [1,2]. Therefore, we studied the ability of two fibrous

polymers, keratin and fibrin, to be used as hydrogels for therapeutic agent delivery. Efforts

focused on bone healing due to the expression of both positive and negative molecular cues

that influence healing outcome. Investigation centered on developing two different

strategies to enhance bone regeneration. The first strategy involved a “classic” TERM

approach in which a potent growth factor was loaded into a biomaterial in order to

stimulate/augment healing. The second strategy investigated a less common strategy to

remove/dampen inhibitory molecules which hinder the potential of the “classic” TERM

approach. For this dissertation, these studies were studied within the context of bone tissue

engineering or neural tissue engineering.

1.1 Controlled Delivery of rhBMP-2 from Kerateine Biomaterials

Current clinical tissue engineered devices for healing bone defects is a successful

example of how a natural polymer carrier (collagen) can locally deliver a pro-regenerative

therapeutic (rhBMP-2). Unfortunately, like many first generation products there are a

number of drawbacks to this device which have resulted in adverse side effects [3]. These

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drawbacks primarily stem from the choice of collagen as the rhBMP-2 carrier. The

inability of the collagen sponge to sequester rhBMP-2 as well as the rapid enzymatic

degradation of the collagen matrix results in a burst release of rhBMP-2 into the local

microenvironment leading to heterotrophic ossification [4]. Also, the rapid enzymatic

degradation of the collagen matrix (in a matter of weeks) does not correlate with the rate

of bone regeneration (months) resulting in voids and non-closure of the bone defects [5].

Therefore kerateine, an alternative natural protein whose hydrolytic degradation rate

coincides with the rate of bone formation, was chosen to investigate its ability as a rhBMP-

2 carrier.

Previously, keratin biomaterials have been successfully used in a number of

applications including bone regeneration [6–8]. These previous studies used crude keratin

extracts which did not separate the alpha from the gamma keratin sub-fractions. Unlike

other natural polymers, keratin material properties can be altered through weight:volume

percent (w/v %) or proportion of sub-fractions (alpha(α):gamma(γ)) (Appendix A) [9].

This unique control over material properties allows for the control of the rate of degradation

and drug release from kerateine biomaterials. In order to take advantage of this

distinguishing feature and to investigate the potential of kerateine (the reduced form of

keratin) to function as a rhBMP-2 carrier to heal critically-sized defects. The goal of this

project was to develop a product which would be able to fill the irregular geometry of a

bone defect and be easily used within an operating room. Therefore, emphasis was placed

on developing a hydrogel that could be directly injected into the defect site or into a mold

to be further processed into an implantable scaffold (Appendix A). Kerateine biomaterials

provide a temporary matrix with porosity within the optimal parameters for

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osteoconductive materials. Rheological characterization of kerateine hydrogels confirm

w/v% and sub-fraction ratios effect on material properties. Hydrogels with greater w/v%

and containing higher ratios of α-fraction showed less “flowable” characteristics. Only

hydrogels which had stable crosslink network with “flowable” characteristics were chosen

for investigation of carrier degradation and rhBMP-2 release kinetics. Results show that

increasing the w:v% and α-kerateine ratio results in a lowering of the rate of kerateine

biomaterial degradation and rhBMP-2 release from the carrier. From our in vitro findings

a 6 % (w/v) 80:20 (α:γ) was chosen to investigate the bioactivity and regenerative potential

of rhBMP-2 kerateine carriers in a critically-sized rat mandibular defect.

To investigate the ability of both an injectable kerateine hydrogel and implantable

scaffold as rhBMP-2 carriers to heal a critically-sized bone defect they were compared to

the current clinically applied tissue engineered device (absorbable collagen sponge - ACS)

(Chapter III). Carriers were loaded with rhBMP-2 and in vitro comparisons between

carriers revealed differences in both carrier erosion as well as rhBMP-2 release. Although

the ACS showed slower rate of carrier degradation (likely due to lack of collagenases), the

hydrolytic degraded kerateine biomaterials had a more representative degradation profile

for the in vivo characteristic release profile due to the lack of known human keratinases.

Kerateine scaffolds and injectable gels showed significantly smaller rhBMP-2 burst release

than ACS which plateaued after 48 hours. This might be attributed to the interaction

between BMP-2 and the heterogeneous mixture of keratin proteins and keratin-associated

proteins making up the keratin biomaterial [10–12]. Previous work done by our group

measured a statistically increased binding affinity between rhBMP-2 and kerateine over

collagen [12]. In order to test if rhBMP-2 delivered from kerateine materials was bioactive,

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treatments with (ACS, kerateine gel, and kerateine scaffold) or without (kerateine gel and

kerateine scaffold) 5µg rhBMP-2 were administered to bilateral critically-sized mandibular

defects. Treatment groups which did not deliver rhBMP-2 were unable to bridge the gap

while every rhBMP-2 carrier completely closed the defect with neo-bone tissue,

highlighting the importance of rhBMP-2 for application in bone regeneration. Bone

mineral density of rhBMP-2 kerateine gel was equivalent to ACS while the rhBMP-2

scaffold outperformed the current tissue engineered bone substitute. Most importantly,

rhBMP-2 kerateine scaffolds reached bone mineral density at the same levels as normal,

uninjured bone by the end of the study. µCT analysis of bone mineral density is supported

by histological evidence which shows the formation of mature bone with large voids within

the ACS regenerated bone which correlates with adverse clinical findings. The formation

of the observed “void” spaces has been previously observed to be dependent on the

concentration of rhBMP-2 within the carrier with lower rhBMP-2 resulting in formation of

less void space [13]. On the other hand, both kerateine carriers showed woven and mature

bone formation with limited areas of void space. Since both ACS and kerateine carriers

were loaded with equal concentrations of rhBMP-2 suggesting that rhBMP-2 concentration

is not the only factor in void formation. One factor which may attribute to the differences

in void formation is the effect of enzymes on the rate of biomaterial degradation. The

histological findings highlight the advantage of having a lack of keratinases within the

body, which would cause rapid carrier degradation resulting in void space within the newly

formed bone. Also, the lack of keratinases prevents the rapid carrier degradation that leads

to rhBMP-2 burst release into the surrounding tissue leading to ectopic bone formation.

Therefore, kerateine offers an attractive alternative natural polymer alternative to collagen

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for the delivery of rhBMP-2 for bone regeneration. Unfortunately, these keratin carriers

still require delivery of a supraphysiological dose of rhBMP-2, which results in commonly

seen adverse ectopic bone growth in the dose equivalent in humans. These findings

highlight the necessity of developing techniques to reduce rhBMP-2 loading concentrations

by addressing the reduction of rhBMP-2 efficacy caused by local inhibitory molecules, as

was addressed in Chapters IV and V of this dissertation work.

1.2 Knockdown of Inhibitory Molecules by Delivery of siRNA Complexes from Fibrin

Hydrogels

In order to explore the ability to reduce the expression of inhibitory molecules, a

proof-of-concept study was developed to deliver siRNA from the surface of fibrin

hydrogels. Successful siRNA complex adsorption and limited complex release from the

fibrin surface suggested substrate-mediated transfection was possible. In order to study the

feasibility of our substrate-mediated transfection approach, a cellular target was chosen

based on its constant expression. For this reason, we initially investigated the ability to

regulate myelin-associated glycoprotein (MAG) expressed by the S16 Schwann cell line

(Chapter IV). In normal physiology MAG provides an important supportive role for axons

in the peripheral nervous system [14], but after injury MAG impedes axon regeneration

[15]. S16 cells were seeded onto fibrin surface with adsorbed siRNA complexes resulting

in almost 100% transfection efficiency while causing little cytotoxicity. siRNA alone was

unable to transfect the cells; these findings highlight the need to complex siRNA with a

cationic transfection agent in order for the complexes to be adsorbed to the negative fibrin

surface (as indicated by the in vitro release study) and for the siRNA to cross the cell

membrane (as indicated by confocal microscopy). Internalized siRNA complexes

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remained biologically active and were able to knockdown MAG mRNA expression when

delivered with polycationic transfection agents. Even though higher than expected off-

target knockdown was measured, we applied this approach for targeting a rhBMP-2

antagonist because the targeted siRNA achieved greater knockdown than off-target effects

for at least one of the transfection reagents (PEI).

1.3 Knockdown of Noggin by Delivery of siRNA Complexes from Fibrin Hydrogels

The efficacy of rhBMP-2 treatments is often diminished by its short half-life [16]

as well as the negative feedback mechanism of endogenous rhBMP-2 antagonists [17].

This has led to loading supraphysiological concentrations of rhBMP-2 into bone substitutes

in order to surpass the therapeutic threshold set by rhBMP-2 short half-life and inhibitors.

By using the technique previously applied to knockdown MAG for peripheral nerve

applications, we investigated the ability of substrate-mediated transfection to control the

mRNA expression of a rhBMP-2 binding antagonist (Chapter V). When pre-osteoblast

cells were seeded on fibrin surface coated with siRNA complexes, nearly 100% of cells

were transfected. Pre-osteoblasts seeded on fibrin matrix exhibited a rhBMP-2 dose

dependent effect on noggin mRNA expression, with the supraphysiological dose of

10µg/mL rhBMP-2 resulting in twenty times more noggin mRNA expression than in cells

not treated with rhBMP-2. Therefore, all subsequent experiments were dosed with

10µg/mL rhBMP-2. Surprisingly, unlike any solution-phase transfection experiments,

siRNA complexes had no cytotoxic effect on pre-osteoblasts when delivered from the fibrin

surface. This could be due to the lack of acute bolus siRNA complex delivery which is seen

in solution-phase transfection. When noggin siRNA transfected pre-osteoblasts were

challenged with 10µg/mL rhBMP-2, surface-mediated transfection showed a dose

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dependent effect to inhibit up-regulation of noggin mRNA. This technique has the potential

to lower the therapeutic threshold caused by rhBMP-2 antagonists thereby reducing the

amount of rhBMP-2 that needs to be loaded into the carrier. This lowered rhBMP-2 dose

would theoretically result in a decrease in clinically-observed adverse side effects. The

local knockdown of noggin expression would allow for bone formation within the site of

injury. If exogenous rhBMP-2 should diffuse away from the carrier and into the

surrounding healthy tissue, the endogenous BMP-2 antagonists, such as noggin, of the

surrounding tissue could potentially neutralize the rhBMP-2 signal and prevent/limit

heterotrophic ossification.

2. Limitations and Prospective Work

The work presented in this dissertation has presented two feasible strategies to

enhance bone regeneration. One by controlling the rate of BMP-2 release from kerateine

carriers, and the other by locally targeting a BMP-2 antagonist (noggin) with siRNA

complexes from the surface of a fibrin carrier. Although our findings show great potential

there are a number of limitations which may be approved upon including: (1.) kerateine

properties, (2.) geometry, (3.) biomimetic cues.

Even though our findings depict that kerateine hydrogel material properties can be

altered by α:γ ratios and weight percent; it also illustrates the limited range of properties

which can successfully be used for an injectable rhBMP-2 carrier product. Kerateine is

spontaneously capable of forming disulfide bonds when free thiols become oxidized. While

providing hydrogel network stability and allowing for kerateine hydrogels to remain for a

month in vivo these disulfide bonds restrict the practical working range of kerateine

formulations for the delivery of rhBMP-2. As previously seen only 6% and 7% kerateine

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gels at limited α:γ ratios were practical for rhBMP-2 delivery for craniofacial defects

(Appendix A). Therefore to increase the range of keratiene material properties it may be

possible to modify the levels of free thiols available for disulfide bonding.

One approach to achieve this is through the mixing of keratose and kerateine to

modulate disulfide bonds. Due to the inability of keratose to form disulfide bonds [18], it

can be mixed with kerateine as a method for controlling crosslinking in a keratin gel

system. Increasing the ratio of keratose in the hydrogel system would in turn leave fewer

cysteine residues capable of forming disulfide bonds. Another method to control the

crosslink density of kerateine is by preventing the disulfide bonds from reforming through

the use of an alkylating agent such as iodoacetamide or acrylamide. These alkylating agents

would “cap” the free thiol groups along the kerateine polymer chain. This method has

successfully be used to increase the range of kerateine degradation and release kinetics

[12]. These are two techniques which can be utilized to control and increase the range of

material properties thereby creating a greater range of rates of carrier degradation and

therapeutic agent release.

In vivo evidence suggests that the rhBMP-2 loading concentration within the

kerateine carriers is still too high (supraphysiological). Even though ectopic formation was

greatly reduced when compared to the ACS, the ectopic bone growth volume was still on

the scale of cm3 on a bone whose thickness is 0.05cm. One potential reason for the ectopic

bone formation was due to our delivery constraints. For example, the thickness of the rat

mandible at the site of injury is 0.5mm thick but unfortunately we were unable to fabricate

scaffolds of such small thickness. Therefore a 2.5mm thick scaffold was fabricated in order

to provide enough support for handling requirements. The injectable rBMP-2 carrier

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showed less ectopic bone formation but was not statistically different than what was

measured with the kerateine scaffold. The ectopic bone growth observed with the kerateine

gel could potentially stem from injecting 50µL per defect when a smaller volume should

have been used. Using a higher initial rhBMP-2 concentration would have allowed for the

delivery of the same total amount of rhBMP-2 but delivered in a smaller volume. A smaller

delivered volume would potentially reduce the ectopic bone formation by limiting the

amount of kerateine gel spreading away from site of action.

Even with success with reducing ectopic bone formation, our current kerateine

carrier delivery of a single growth factor (BMP-2) does not recapitulate the dynamic

physiological cell signaling seen during bone healing (Chapter I). In order to better mimic

the native healing environment future work may focus on delivering a cocktail of molecular

promoters of bone regeneration. In our work, the rat mandible is a relatively thin bone and

concerns with vascularization was not directly addressed. When scaling up into larger

bones, regeneration is if often obstructed by the lack of neo-vasculature formation. Studies

have previously shown the importance of angiogenesis and its ability to promote bone

regeneration [19,20]. Research on dual delivery of angiogenic and osteogenic growth

factors from the same biomaterial in on-going [21]. One angiogenic factor of particular

interest is vascular endothelial growth factor (VEGF) not only because it increases

endothelial cell differentiation, proliferation, and vascular tube formation [22] but also

contributes to the cross-talk between endothelial cells and osteoblasts [23]. Dual delivery

of VEGF and BMP-2 has been shown to significantly enhance bone regeneration compared

to delivering VEGF and BMP-2 separately [20,24–26]. Though careful consideration must

be placed on the both the concentration [27,21] as well as the respective rate of release [21]

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in order to elicit a therapeutic response. Keratin hydrogels have been show to

simultaneously deliver a cocktail of VEGF, bFGF, and IGF-1 in vitro [28], which are all

growth-promoting molecules involved in the bone healing cascade [29]. For future BMP-

2 kerateine carriers to be successful in regenerating larger bone defects, investigation into

the addition of angiogenic molecules such as VEGF should be considered.

Surface-mediated siRNA has the potential to be a potent tool for applications

wherein cells migrate across the biomaterial; such as in conduits for peripheral nerve

regeneration [30]. Proof-of-principle studies demonstrate the ability of pre-osteoblasts to

take up siRNA complexes adsorbed to a fibrin surface (Chapter IV). In order for this

technique to be successfully translated into a bone substitute it must be incorporated

directly into the substrate matrix. Fibrin has previously been successfully embedded with

pDNA complexed with PEI and this technique can be used to encapsulate siRNA

complexes [31].

One potential downfall of noggin siRNA delivery is the initial infiltration of

inflammatory cells which would take up the siRNA complexes in vivo; therefore, noggin

might not be the best target. The normal bone healing cascade pro-inflammatory mediators

are required to initiate bone healing [32]. Unfortunately, when any foreign biomaterial is

implanted into the body they induce varying degrees of a host inflammatory response [33–

35]. Up-regulation of inflammatory cytokines has shown to inhibit bone healing [36] and

could serve as potential targets during the first stages of bone regeneration. Clinical

concerns have also been raised due to inflammatory swelling by the up-regulation of IL-6

and TNFα caused by rhBMP-2 from collagen carriers [3]. rhBMP-2 induced inflammation

has been shown to be triggered in a dose-dependent manner (i.e., the greater rhBMP-2 dose

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the greater the pro-inflammatory cytokine expression) [37]. Also, a number of pro-

inflammatory cytokines have been identified to impair bone regeneration including

Interleukin (IL)-1,-6, and TNFα [38,39]. Pro-inflammatory cytokines also have shown to

reduce the osteoinductive efficacy of BMP-2 [40,41]. The greatest rhBMP-2 burst release

from carriers (increasing cytokine production) occurs within the first few days post-

implantation, which correlates with the greatest expression levels of pro-inflammatory

cytokines thereby compounding the inflammatory response which impedes bone

regeneration [42]. Also, increased edema and cytokine production in vivo has shown to be

dose-dependent with increasing concentrations of BMP-2 [37]. These facts highlights the

need to locally reduce inflammatory molecules to enhance bone regeneration. The addition

of drugs which modulate the pro-inflammatory response (e.g., corticosteroids and

nonsteroidal anti-inflammatory drugs) have been demonstrated to improve bone fracture

healing [43,42]. Thus, recent success in modulating pro-inflammatory cytokine expression

by local deliver of siRNA [44,45] has significant implications for the potential application

in bone defect healing.

A second potential application of substrate-mediated siRNA delivery is to promote

angiogenesis in order to enhance bone regeneration. As previously discussed in this

chapter, establishment of neo-vasculature is critical for bone regeneration. Once again, the

classic tissue engineering and regenerative medicine (TERM) approach for delivering pro-

regenerative factors, such as VEGF, is one method to promote angiogenesis. Recent work

has investigated the ability to target anti-angiogenic factors such as intracellular adaptor

protein (Lnk) [46] and prolyl hydroxylase domain-containing protein 2 (PHD2) [47] which

has resulted in increased angiogenic factors such as VEGF and Angiopoietin-1 [46,47].

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When applied in vitro in a mouse fracture model, local delivery of siRNA targeting Lnk

mRNA was able to significantly accelerate bone healing when compared to control siRNA

groups [46]. Therefore, there are a number of molecular inhibitors within the bone healing

mechanism which can be locally down regulated through substrate-mediated siRNA

delivery in order to enhance bone regeneration.

In the present studies, only mRNA expression levels were measured. No functional

protein measurements were preformed such as a MAG or noggin-specific ELISA or

Western blot. Even though there was a significant reduction of MAG and noggin mRNA,

this does not ensure that sufficient reduction of mRNA was achieved to reduce protein

expression. For this reason, future studies should include mRNA and protein expression

over a longer period of time to assess the effectiveness of fibrin-mediated delivery of

siRNA. This is particularly true in systems like bone in which regeneration takes a number

of weeks [32]. Our previous work highlights our ability to regulate noggin mRNA levels

in a dose dependent manner. Next, functional osteogenic outcome measures should also

be investigated to investigate the impact noggin mRNA/protein knockdown has on BMP-

2 efficacy. In line with our current results, we hypothesize that BMP-2 efficacy should

increase in a complimentary dose-dependent manner to noggin knockdown (more noggin

knockdown, more BMP-2 efficacy). To test this the same groups of BMP-2 can be used:

0µg/mL BMP-2 (negative control), 10µg/mL BMP-2 with no noggin siRNA (positive

control), and then 10µg/mL BMP-2 with varying doses of noggin and control siRNA.

BMP-2 efficacy can be tested by BMP-2’s ability to bind to its receptor and initiate a

signaling cascade by measuring downstream activation of osteogenic transcription factors

such as runt-related transcription factor-2 (RUNX2) (pre-osteoblasts) [48,49], also known

159

as core-binding factor subunit alpha-1 (CBFα1), and/or Osterix (Osx), also known as

Sp7(mature osteoblasts) [50]. Functional outcome studies such as Alkaline Phosphatase

(measure osteoblast activity) and Alizarin Red (measure calcific deposition) will then

provide information on the further downstream efficacy of BMP-2. To investigate if BMP-

2 or noggin down-regulation caused chondrogenic activity, Alcian Blue staining (stains

sulfated proteoglycans deposits) can be performed. Once proof-of-principle has been

established using surface-mediated delivery of noggin siRNA, the siRNA complexes must

be incorporated into the fibrin (or other) hydrogels for delivery before in vivo experiments

can be conducted. Unfortunately, this means additional in vitro testing must be performed

since the application is moving from a 2-D into a 3-D system and BMP-2 and siRNA doses

will need to be adjusted.

Another potential downfall of using fibrin for bone regeneration is that unmodified

fibrin might not be the best carrier system due to rapid enzymatic degradation [51,52]. In

terms of noggin siRNA controlling the initial burst release of rhBMP-2 from a carrier this

may not be a problem, but unfortunately the fibrin (without modifications) would degrade

too quickly to provide a suitable matrix for infiltrating cells [51], leading to delayed union,

non-union, or development of voids within the regenerated bone. Therefore, an alternative

natural polymer maybe more suitable. Interestingly, based on our results described in

Chapter III and above, keratin would be a suitable alternative natural polymer for substrate-

mediated siRNA delivery for bone applications due to its overall charge and degradation

rate which matches the rate of bone tissue formation (months). Combining both BMP-2

and noggin siRNA complexes in a dual delivery system using a kerateine carrier system

could be the next step in furthering the knowledge presented in this dissertation.

160

Experimental groups would have to cover: kerateine gel alone –BMP2 -siRNA, kerateine

gel +BMP-2 –siRNA, kerateine gel –BMP-2 +noggin siRNA, kerateine gel-BMP-2

+control siRNA, kerateine gel +BMP-2 +noggin siRNA, and kerateine gel +BMP-2

+control siRNA. Also, various ratios of BMP-2 to noggin siRNA would need to be

examined in order to elicit the greatest osteogenic potential with the lowest BMP-2 dose.

Natural-based biomaterials present a number of challenges including predictability

of degradation, batch-to-batch variability and denaturation during fabrication and

processing. When proceeding with natural polymers for future TERM products, strict

control over resource material and good manufacturing practice must in be in place to

mitigate variability as much as possible [53].

Outside the scope of bone regeneration, the two approaches investigated (BMP-2

kerateine carriers and siRNA fibrin carriers) can be adapted to a number of regenerative

application for critically-sized wounds (e.g., volumetric muscle loss [54]) since all tissues

are affected by both positive and negative regenerative cues. Our system can also

potentially be applied in prevention/reduction of scarring in skin [55,56] and abdominal

adhesions [57,58]. Overall, this thesis has provided insight into controlling short-

term/temporary molecular cues to enhance wound healing until normal physiological

function is restored.

3. Conclusion

This dissertation work highlights the potential of natural polymers (keratin and

fibrin) as carriers for pro-regenerative molecules (rhBMP-2 and noggin siRNA) in bone

regeneration. This work provides an insight into enhancing bone regeneration through

controlled delivery of growth factors whose efficiency could be increased by the short term

161

knockdown of inhibitory molecules. Further work to optimize both of these strategies to

enhancing TERM bone regeneration products is needed. Most importantly, these

approaches have the potential to be tailored and applied to enhance any wound healing

system that can be treated by tissue engineering/regenerative medicine approaches.

162

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APPENDIX A

MODULATION OF KERATEINE MATERIAL PROPERTIES FOR THE CONTROLLED

RELEASE OF RECOMBINANT HUMAN BONE MORPHENOGENETIC PROTEIN-2.

1. Overview

First generation recombinant human bone morphogenetic protein-2 (rhBMP-2)

carriers for the regeneration of large bone defects have a number of adverse side effects

stemming from the inability to adequately control the rate of rhBMP-2 release from the

carrier. A number of alternative natural polymers have been under investigation in a variety

of tissue engineering applications for a number of years including keratin, a fibular protein

extracted from hair. Variation in keratin biomaterial material properties result from not only

the extraction method used (oxidative or reactive) but also the separation of protein sub-

fractions (alpha (α) and gamma (γ)) respectively possessing their own unique properties

(Figure S1). We hypothesized that the reduced form of keratin (kerateine) native sub-

fractions (α and γ kerateine) can be mixed in order to change the keratin material properties

to control rhBMP-2 release. This appendix shows the in vitro investigation leading to

choosing the kerateine formulation (6% 80:20) for our in vivo work (Chapter III). Our

findings showed kerateine (KTN) hydrogel material properties are affected by both weight-

volume (w/v) percentage as well as the ratio of α-KTN to γ-KTN (α:γ) (Figure S2). The

change in KTN material properties had an impact on the both the rate of KTN degradation

(Figure S3) as well as the rate of rhBMP-2 released (Figure S4) from the respective

hydrogels. Taken together KTN hydrogels provide an interesting alternative rhBMP-2

carrier system for bone regeneration applications.

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Figure S1. Keratin extraction leads to either keratose or kerateine production through

oxidative or reductive processes, respectively. Alpha and gamma fractions are separated

by isoelectric precipitation, which can then be lyophilized and store until reconstituted into

desired formulations.

2. Methods

2.1 rhBMP-2 Preparation

rhBMP-2 was obtained by the generous donation of lyophilized powder from Pfizer

(New York, NY). rhBMP-2 acquired from Pfizer was reconstituted with sterile DI water,

aliquoted, and stored at -20°C until it was used.

169

2.2 Biomaterial Preparation

Lyophilized and sterile (via 2 MRad gamma-irradiation) kerateine powder (Figure

1) was obtained from KeraNetics (Winston-Salem, NC). In order to explore the influence

of weight percent and α:γ ratio kerateine fractions; a number of kerateine gel formulation

were made (STable 1). Pre-determined amounts of sterile α- and γ-kerateine powders were

weighed and vigorously vortexed in order to uniformly mix the components. The premixed

kerateine powders were then reconstituted with either sterile DI water, 100µg/mL rhBMP-

2 (10ug total dose per 100µL gel), vortexed until well mixed, and allowed to gel overnight

at 37°C (humidified atmosphere of 5% CO2 /95% air).

STable 1. Kerateine Hydrogel Formulations

Total

Weight/Volume

Alpha

Fraction

(α)

Gamma

Fraction

(γ)

5% 100

90

80

70

0

10

20

30

6% 100

90

80

70

0

10

20

30

7% 100

90

80

70

0

10

20

30

8% 100

90

80

70

0

10

20

30

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2.3 Rheological Characterization of Kerateine Hydrogel Material Properties

Kerateine formulations were formed as previously described. Briefly, powder in

sterile DI water was vortexed, centrifuged, loaded into a 22 x 1.2 mm cylindrical teflon

template, and then incubated overnight at 37˚C to make a homogenous gel (n=3). Samples

were then loaded onto a Bohlin CS-10 Rotational Rheometer (Malvern, Worcestershire,

UK) system with parallel plate geometry (gap width = 1mm) for assessment of gel

rheological characteristics at room temperature (25˚C). A dynamic oscillation stress sweep

at a constant frequency (f) of 1 Hz was used to determine the linear viscoelastic region

(LVER) of the kerateine biomaterial. A frequency sweep from 0.1-10 Hz was then

performed at a constant stress (25 kPa) within the LVER.

2.4 Kerateine Formulation Degradation and rhBMP-2 Release

Release of rhBMP-2 from each carrier was determined in vitro over the course of 4

weeks. rhBMP-2 carriers without rhBMP-2 served as negative controls. 100μL of sterile

PBS (Gibco, Grand Island, NY) was added to each conical tube. Tubes were then incubated

at 37°C. At preassigned time points the PBS was collected and replaced with fresh 100µL

PBS. Collected samples were frozen at -80˚C until analysis was performed by DC Protein

Assay (Bio-Rad, Hercules, CA) and rhBMP-2 ELISA (Peprotech, Rocky Hill, NJ) for

carrier erosion and rhBMP-2 release, respectively. rhBMP-2 carriers without rhBMP-2

served as negative controls for rhBMP-2 ELISAs. Carrier degradation was normalized to

total initial carrier protein weight while rhBMP-2 release was normalized to initial rhBMP-

2 loading dose.

171

3. Results and Discussion

3.1 Rheological Characterization of Kerateine Hydrogel Material Properties

To decrease the potential number of experimental groups we first examined the

material properties of the various kerateine formulations (STable 1). Hydrogel selection

criteria requirements were that the hydrogel formulations are able to form stable cross-link

networks (Elastic Modulus (G’) > Viscous Modulus (G’’)). Kerateine gels (STable 1) were

formed and allowed to gel overnight in a Teflon template designed to allow the hydrogel

to fit the dimensions of the rheometer testing area. Due to viscoelastic properties of

hydrogels, samples were tested at room temperature on a 1mm gap parallel plate shear

rheometer. A dynamic oscillation stress sweep at a constant frequency of 1 Hz was used

to determine the linear viscoelastic region (LVER) of the kerateine biomaterial. Samples

were then tested at constant stress (25 kPa) which lies within the LVER over a frequency

sweep from 0.1-10 Hz.

Rheological evidence illustrates that increasing ratio of γ-fraction kerateine

decreases the complex modulus of the hydrogel (Figure S2A). Increasing weight percent

of kerateine increases the complex modulus of the hydrogel (Figure S2B). 5% gels were

characterized by physical chain entanglements and were unable form covalent bonds

between polymer chains (Figure S2C). We verified that kerateine hydrogels were able to

form stable disulfide cross-links over the entire frequency ranged for 6% and 7% gels

(Figure S2D). The 8% w/v gels would crumble when shear force was applied and the

material properties were not affected with increasing γ-fraction. For these reasons we are

able to exclude the 5% and 8% weight/volume (w/v) formulations of kerateine hydrogels.

Therefore, both 6% and 7% kerateine gels chosen for future experiments since they were

172

able to form stable hydrogel networks whose material properties could be modified through

varying α:γ ratios.

Figure S2. Determination of kerateine complex modulus as a function of α:γ ratio (A) and

weight percent (B). Determination of stable kerateine networks by elastic and viscous

moduli through frequency. Kerateine pseudogels (5% 80:20) (C) were excluded while

stable hydrogel networks (7% 80:20) (D) were included for future experiments. From the

rheological evidence, kerateine hydrogel material properties can be altered by changing α:γ

ratio and weight percent.

3.2 Degradation Profile of Kerateine Biomaterials

In an ideal material for tissue engineering, the rate of biomaterial degradation

should correlate with the rate of which the tissue can migrate and proliferate into the defect

area. The repair stage duration of fractures spans several weeks to form woven bone with

the biomaterial needing to provide structural support for that length of time. This is a

potential advantage of using kerateine biomaterials which last months both in vitro and in

vivo. To further investigate this, rates of degradation were once again be tailored by

173

kerateine weight percent and varying α- and γ-kerateine fraction ratios. Kerateine gels

were formed as previously described in the bottom of 1.5mL round bottom tubes and

allowed to gel overnight. From the previous rheological data we have excluded 5% and 8%

w/v kerateine gels. The following day an equal volume of phosphate buffer saline (PBS)

was added to the top of the formed gels and incubated at 37˚C. Preassigned time points

were collected over a course of several weeks. Samples were collected and fresh PBS was

replenished on top of the kerateine hydrogel formulations. The total amount of protein in

each sample was determined by Lowry protein assay.

The results of the in vitro data suggest these kerateine hydrogels would persist

through the repair stage of fracture healing. Importantly, ~50% of the total mass percent

of kerateine hydrogels remained after 35 days. Increasing amounts of γ-fraction kerateine

resulted in increased degradation rates of the hydrogel (Figure S3A) while increasing

weight percentage lowered the degradation rates of the kerateine hydrogel (Figure S3B).

Figure S3. Determination of kerateine degradation rate as a function of α:γ ratio (A) and

weight percent (B). From the lowry protein assay evidence, kerateine hydrogel degradation

rates can be altered by changing α:γ ratio and weight percent.

174

3.3 rhBMP-2 Release Kinetics from Kerateine Biomaterials

For regenerative medicine applications it is important that growth factors are able

to be released and diffuse away from their biomaterial carrier to create a chemoattractive

gradient for migrating cells to follow. Once cells have migrated to the biomaterial carrier,

it is important that there is a growth factor reservoir remaining within the carrier system to

promote regeneration. For this purpose we investigated if rhBMP-2 release profiles can be

altered by changing kerateine material properties through α:γ fraction ratios and weight

percent. Kerateine gel formulations (STable 1) were reconstituted with a sterile DI water

solution (control) or containing 100µg/mL rhBMP-2. 100µL of gel was loaded into the

bottom of 1.5mL round bottom tubes where they polymerized overnight. From the

previous rheological data we have excluded 5% and 8% w/v kerateine gels. The following

day, 100µL of phosphate saline buffer was added to the top of the formed gels and

incubated at 37˚C over a course of several weeks. Samples were collected at preassigned

time points and fresh PBS was replenished on top of the kerateine hydrogel formulations.

The total amount of rhBMP-2 in each sample was determined by rhBMP-2 specific

Enzyme-linked Immunosorbent Assay (ELISA).

Findings show that increasing the gamma composition of the kerateine hydrogel

causes an increase of rhBMP-2 release from the hydrogel (Figure S4A). Interestingly,

rhBMP-2 is released from 7% and 6% kerateine gels did not show a difference (Figure

S4B).

175

Figure S4. Determination of rhBMP-2 release from kerateine hydrogels as a function of

α:γ ratio (A) and weight percent (B). From the ELISA evidence, kerateine hydrogel

degradation rates can be altered by changing α:γ ratio while weight percent did not.

4. Conclusions

Our findings support our ability to modify kerateine material properties to control

rhBMP-2 delivery. Changing kerateine material properties by α:γ kerateine fraction ratios

and weight percent resulted in varying mechanical, degradation, and rhBMP-2 release

properties. The degradation rate of the kerateine hydrogels indicates that the rhBMP-2

carrier could remain for enough time to allow and support cellular migration into the site

of injury. From our findings and the ease of which the formulation could be injected, we

selected the 6% 80:20 kerateine formulation to be used in the critically-sized rat mandibular

defect model. Even though our data depicts kerateine hydrogel material properties can be

altered by α:γ ratios and weight percent, it also illustrates the limited range of properties

which can be achieved.

A B

176

CHRISTINE JANE KOWALCZEWSKI

33 Lynn Batts Lane, Apt 5108• San Antonio, TX

(607) 738-1976 • [email protected]

EDUCATION

Ph.D. Biomedical Engineering (2009 – 2014 expected)

Virginia Tech/Wake Forest University, Winston-Salem, NC

Advisor: Dr. Justin M. Saul

Thesis: Natural polymeric carriers for local delivery of therapeutic agents to promote

enhanced bone regeneration.

M. Eng. Biomedical Engineering (2008 - 2009)

Cornell University, Ithaca, NY

Advisor: Dr. Claudia Fischbach-Teschl

B.S. Biomedical Engineering (2003 - 2007)

Rensselaer Polytechnic Institute, Troy, NY

EXPERIENCE

Institute for Surgical Research

Postdoctoral Fellow (July 2014 – present)

Worked as part of an interdisciplinary team for Dr Robert Christy

Develop porcine model to study the effects of antimicrobial loaded hydrogel

wound dressing

Miami University Research Associate (January 2012 – June 2014)

Oversaw planning and implementation of a new research laboratory

Supervised 2 undergraduate summer scholars, 3 masters students, trained 2 lab

technicians, and 6 undergraduates

Preformed surgical procedure of critically-sized mandibular defect in rats to

evaluate bone regeneration and ectopic bone growth of rhBMP-2 keratin carriers

compared to a commercially available product

Developed a substrate mediated polycationic based siRNA delivery system from

fibrin hydrogels for improved rhBMP-2 efficacy in vitro by the down regulation

of a BMP-2 antagonist in pre-osteoblasts

Modulated keratin hydrogel crosslinks to control keratin hydrogel degradation

and therapeutic agent release

o Developed a protocol to mimic enzymatic degradation in bone

microenvironments

Wake Forest University

Wake Forest Institute for Regenerative Medicine (WFIRM)

Research Assistant (July 2009 – January 2012)

Supervised 1 undergraduate summer scholar and 1 first year rotational PhD

student

Developed a surface mediated polycationic based oligonucleotide delivery system

from fibrin hydrogels for peripheral nerve applications

mailto:[email protected]

177

o Evaluated transfection efficiency and targeted mRNA sequence knockdown of

inhibitory molecules in schwannoma cells

Manufactured PEGylated liposomes loaded with fluorescently tagged BSA

Worked as part of an interdisciplinary team for an industry (KeraNetics, LLC)

funded research project to develop an injectable keratin gel and implantable

keratin scaffold as a carrier for rhBMP-2 for bone regeneration

o Interacted with industry PI on regular basis and gave monthly project

milestone updates

o In vitro characterization and validation of keratin hydrogel formulations

o Developed a clinically-relevant mandibular critically-sized bone defect model

in rats

Cornell University

Master’s Design Project (June 2008 – May 2009)

Supervised one undergraduate student

Application of a novel PLG 3D cell culture system designed to mimic the native

bone structure to study the effects of environmental factors which influence

paracrine signaling between tumor and native healthy cells on osteolysis.

Assessed that the mineral constituent hydroxylapatite (HA) of the bone

microenvironment plays a role in promoting osteoclast activation by stimulating

secretion of cytokines by mammary tumor cells

Publications

Kowalczewski, C., Tombyln, S., Wasnick, D., Hughes, M., Ellenburg, M.,

Callahan, M., Smith, T., Van Dyke, M., Burnett, L., Saul, J. “Reduction of Ectopic

Bone Growth in Critically-Sized Rat Mandible Defects by Delivery of rhBMP-2

from Kerateine Biomaterials.”

Tomblyn, S., Kneller, E., Ellenburg, M., Kowalczewski, C., Walker, S., Burnett,

L., Van Dyke, M., Saul, J. “Simultaneous Delivery of Exogenous Growth Factors

and Muscle Progenitor Cells with keratin hydrogels.” Submitted 2014

Kowalczewski, C., Saul, J. “Knockdown of the Bone Morphogenetic Protein-2

Binding Antagonist Noggin Through Surface-Mediated Delivery of Noggin

siRNA from Fibrin Hydrogels.” In preparation 2014.

Kowalczewski, C., Saul, J. “Surface-Mediated Delivery of siRNA Using

Polycationic Complexes for the Knockdown of Inhibitory Molecules in Peripheral

Nerve Regeneration.” In preparation 2014.

Kowalczewski, C., Saul, J. “Naturally Derived Polymer Delivery Systems for

Regenerative Pharmacology.” In preparation by invitation of Frontiers in

Pharmacology. 2014

Wilson, P., Kowalczewski, C., Lu, B., Mack, D., Henninger, J. Stem Cell

Engineering Handbook. Chapter 3 Engineering Pluripotency: Induced Pluripotent

Stem Cells (ISBN: 1420090054). In press. June 2014.

Pathi SP, Kowalczewski C, Tadipatri R, Fischbach C (2010) A Novel 3-D

Mineralized Tumor Model to Study Breast Cancer Bone Metastasis. PLoS ONE

5(1):e8849. doi:10.1371/journal.pone.0008849

178

Podium Presentations

Biomedical Engineering Society 2014 Annual Meeting (October 22-25, 2014)

Kowalczewski, C. and Saul, JM. “Delivery of siRNA from Fibrin Hydrogels for

mRNA Knockdown of the BMP-2 Antagonist Noggin”

Henry B. Gonzalez Convention Center, San Antonio, TX

Orthopaedic Research Society 2013 Annual Meeting (January 26-29, 2013)

Kowalczewski, C. et al. “Controlled Release of rhBMP-2 from Keratin

Biomaterials Promotes Bridging of Critically-Sized Rat Mandibular Defect While

Minimizing Ectopic Bone Growth.”

Henry B. Gonzalez Convention Center, San Antonio, TX

Tissue Engineering and Regenerative Medicine International Society (TERMIS)

(December 5-8, 2010)

Kowalczewski, C. Saul, J. “Surface-Mediated Delivery of siRNA Using

Polycationic Complexes for the Knockdown of Inhibitory Molecules in Nerve

Regeneration.”

Hilton Walt Disney World Resort, Orlando, FL

Institute of Biological Engineering Regional Conference (October 17-18, 2008)

Kowalczewski, C. Pathi, S. Fischbach, C. “A Novel 3-D Mineralized Tumor

Model to Study the Release of Interleukin-8 in Breast Cancer Bone Metastasis.”

Cornell University, Ithaca, NY

AWARDS

Wake Forest Medical School Alumni Student Travel Award

(December 2010, December 2012)

1st Place Poster Institute of Biological Engineering Regional Conference

(April 17, 2009)

Rensselaer Alumni Scholarship

(2003-2007)

PROFESSIONAL SOCIETIES

Tissue Engineering & Regenerative Medicine International Society (TERMIS)

(2009-present)

o TERMIS-SYIS Student Co-chair

(July 2011- February 2012)

Biomedical Engineering Society (BMES)

(2009-present)

Society for Biomaterials

(2009-present)

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