CUES BY JOHN BRADFORD SCOTT A Dissertation Submitted … · CUES BY JOHN BRADFORD SCOTT A Dissertation Submitted to the Graduate Faculty of ... Perfusion loop schematic ... in tissue

DEVELOPMENT OF A TISSUE ENGINEERED SKELETAL MUSCLE REPAIR

CONSTRUCT FEATURING BIOMIMETIC PHYSICAL, CHEMICAL, AND MECHANICAL

CUES

BY

JOHN BRADFORD SCOTT

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

May 2015

Winston-Salem, North Carolina

Approved By:

George J. Christ, Ph.D., Advisor & Chair

Martin K. Childers, Ph.D.

Carolanne E. Milligan, Ph.D.

Aaron M. Mohs, Ph.D.

Justin M. Saul, Ph.D.

Abby R. Whittington, Ph.D. / Aaron S. Goldstein, Ph.D.

ii

TABLE OF CONTENTS

ACKNOWLEDGEMENTS ............................................................................................... iii

LIST OF FIGURES AND TABLES .................................................................................. iv

ABBREVIATIONS .......................................................................................................... vi

ABSTRACT .................................................................................................................. viii

CHAPTER I – INTRODUCTION ...................................................................................... 1

CHAPTER II – THE PROMOTION OF AXON EXTENSION IN VITRO USING

POLYMER-TEMPLATED FIBRIN SCAFFOLDS ...................................................... 37

CHAPTER III – IN VITRO AGRIN-INDUCED ACETYLCHOLINE RECEPTOR

CLUSTERING ON SKELETAL MUSCLE CELLS SEEDED ON A TUNABLE FIBRIN-

BASED SCAFFOLD ................................................................................................ 70

CHAPTER IV – ADVANCES TOWARD ENABLING THREE-DIMENSIONAL FIBRIN

SCAFFOLDS FOR TISSUE ENGINEERED MUSCLE REPAIR ............................. 109

CHAPTER V – PRELIMINARY EVALUATION OF INNERVATION AND REMODELING

OF A TISSUE ENGINEERED SKELETAL MUSCLE REPAIR CONSTRUCT IN VIVO

.............................................................................................................................. 152

CHAPTER VI – DISCUSSION, CONCLUSIONS, AND FUTURE DIRECTIONS .......... 173

CURRICULUM VITAE ................................................................................................. 183

iii

ACKNOWLEDGEMENTS

First, I would like to thank my brothers and friends – near and far, past and

present. Despite my best efforts to the contrary, you have kept me (mostly) sane through

difficult times, provided a constant reminder that life exists beyond my professional

commitments, and at times challenged me to think differently and grow beyond my

previous limitations.

I also acknowledge my parents – despite (and sometimes because of) our

diferrent points of view, the way I view and interact with the world has continually

evolved. Thank you for your encouragement and tolerance.

To those who have worked alongside me in the completion of this dissertation –

members of the Christ and Saul labs, WFIRM core technicians, you know who you are –

thank you for your constant assistance, willingness to discuss ideas, and academic

support. This work would have been impossible without you.

To members and staff of WFIRM, SBES, and the WFU Graduate School – thank

you for providing the framework and logistical support for my continuing education. I

would like especially to express my gratitude to members of my advisory committee,

who have shown remarkable patience during my candidacy.

Finally, I would like to thank the faculty and staff of Wake Forest who have

educated me about alternative careers in science. Your work gives hope to often

clueless graduate students on the cusp of entering a competitive world.

iv

LIST OF FIGURES AND TABLES

CHAPTER I

Fig. 1. Schematic of final tissue engineered muscle construct fabrication and

evaluation .......................................................................................................... 20

CHAPTER II

Fig. 1. Scaffold fabrication process .......................................................................... 41

Table 1. Template fiber fabrication parameters and resulting diameter .................... 43

Fig. 2. Scaffold morphology ..................................................................................... 51

Fig. 3. Conduit alignment within scaffolds ................................................................ 52

Fig. 4. Quantification of fiber template diameter relative to measured resulting

conduit diameter ................................................................................................ 53

Table 2. Scaffold porosity ........................................................................................ 54

Fig. 5. Scaffold mechanical properties ..................................................................... 55

Fig. 6. Axon infiltration of scaffolds .......................................................................... 57

Fig. 7. Quantification of axon infiltration ................................................................... 58

CHAPTER III

Fig. 1. Schematic of 3D tissue engineered muscle construct fabrication, seeding, and

preconditioning .................................................................................................. 84

Fig. 2. Efficacy of agrin treatment in 2D ................................................................... 88

Fig. 3. AChR clustering adjacent to agrin-presenting microparticles in 2D ............... 90

Fig. 4. Dramatic impact of bioreactor preconditioning on AChR expression and

organization in 3D tissue engineered constructs seeded with rat MDCs ............ 92

Fig. 5. Representative examples of AChR clustering in a bioreactor-preconditioned

agrin-presenting tissue engineered construct ..................................................... 93

v

CHAPTER IV

Fig. 1. Perfusion loop schematic ............................................................................ 117

Fig. 2. Cell-infiltrated fibrin scaffold ........................................................................ 126

Fig. 3. Cell seeding of patterned fibrin immediately after perfusion ........................ 128

Fig. 4. Cell density in rMDC-seeded fibrin scaffolds over time................................ 129

Fig. 5. Fibrin scaffolds with 12 μm diameter conduits after cell seeding with a 4.8

mL/min perfusion of rMDCs ............................................................................. 130

Fig. 6. Effect of directionality of perfusion and conduit lumen diameter on cell seeding

........................................................................................................................ 131

Fig. 7. Example melt extruded PVOH fiber ............................................................ 132

Table 1. Water solubility and composition of melt-extruded fibers .......................... 133

Fig. 8. Initial perfusion-seeded, microparticle-presenting 30 mm constructs........... 134

Fig. 9. Optimized microparticle loading of fibrin scaffold......................................... 135

Fig. 10. Initial perfusion cell seeding of 10-15 mm long fibrin scaffolds .................. 136

Fig. 11. rMDC perfusion of 15 mm long CA-templated fibrin scaffolds mounted in

silicone tubing .................................................................................................. 136

CHAPTER V

Fig. 1. Schematic overview of rat neurotization study ............................................ 157

Fig. 2. Overview of host response to construct implantation .................................. 164

Fig. 3. Glycogen content of explants in the construct area ..................................... 166

Fig. 4. Potential Interaction of immobilized femoral nerve stump with suture.......... 167

vi

ABBREVIATIONS

2D: two-dimensional

3D: three-dimensional

α-BTX: alpha bungarotoxin

ACh: acetylcholine

AChR: acetylcholine receptor

AJ1: 41 μm diameter pMMA fiber formed using the Alex James & Associates extruder

ANOVA: analysis of variance

BAM: bladder acellular matrix

BSA: bovine serum albumin

CA: cellulose acetate

DAPI: 4',6-diamidino-2-phenylindole

DI: deionized

DMEM: Dulbecco's modified Eagle's medium

DRG: dorsal root ganglia / ganglion

E: Young’s modulus

ECM: extracellular matrix

EDC: N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride

FBR: foreign body response

FBS: fetal bovine serum

FDU: 5-fluoro-2′-deoxyuridine

H&E: hematoxylin and eosin

HS: horse serum

ID: inner diameter

LD: latissimus dorsi

vii

Lrp4: low-density lipoprotein receptor-related protein 4

(r)MDC(s): (rat) muscle-derived cell(s)

MEP: motor end plate

MMP(s): matrix metalloproteinase(s)

MuSK: muscle-specific kinase

MW: molecular weight

OD: outer diameter

NBF: neutral-buffered formalin

NGF: nerve growth factor

NMJ: neuromuscular junction

PAS: periodic acid - Schiff

PBS: phosphate-buffered saline

PEO: poly(ethylene oxide)

pMMA: poly(methyl-methacrylate)

PVOH: polyvinyl alcohol

s-NHS: N-hydroxysulfosuccinimide

SD: standard deviation

SEM: scanning electron microscopy

TEMR: tissue engineered muscle repair

viii

ABSTRACT

John Bradford Scott

DEVELOPMENT OF A TISSUE ENGINEERED SKELETAL MUSCLE REPAIR

CONSTRUCT FEATURING BIOMIMETIC PHYSICAL, CHEMICAL, AND MECHANICAL

CUES

Dissertation under the direction of

George J. Christ, Ph.D.

Volumetric muscle loss (VML), an injury to skeletal muscle which permanently

impairs appearance and function [1], commonly results from trauma, congenital defect,

or surgical side effect, and significantly affects quality of life. Unfortunately, clinical

treatments for VML are lacking. The "gold standard" treatment remains autografting,

which is often limited by donor site availability and morbidity as well as incomplete repair

of function or restoration of cosmesis. In the current work, we describe an in vitro

approach to tissue engineered skeletal muscle repair combining biomaterial-seeded

cells with pharmacological and mechanical cues. The overall goal of this work was to

develop and characterize a tissue engineered construct which could eventually form the

basis for a clinical alternative to autografting in VML.

This approach began with a fibrin scaffold patterned with an aligned array of

cylindrical pores. This scaffold was evaluated for its mechanical properties and its ability

to support neurite ingrowth and muscle cell seeding in vitro. The approach further

incorporated exogenous agrin, which was designed to mimic in vitro the neuronal

pharmacological cue initiating native assembly of the motor end plate (MEP) in vivo.

Agrin efficacy was evaluated both alone and in conjunction with mechanical stretch by

histological labeling of acetylcholine receptor (AChR) clusters, the presumptive in vitro

ix

analog of MEPs. Next, the ability of the construct to maintain these AChR clusters in the

complex biological environment in vivo was assessed. Constructs were implanted

subcutaneously in a rodent model and connected to the proximal stump of the femoral

nerve. This experiment was designed to assess the biocompatibility and initial

remodeling of the implanted construct as well as the potential formation and function of

neuromuscular junctions between host nerves and construct muscle cells.

These studies demonstrated that patterned scaffolds supported neurite ingrowth

and muscle cell seeding in vitro. Moreover, the combination of agrin and stretch in vitro

resulted in dramatic enhancement of AChR clustering, which is a key phenotypic aspect

of MEPs on innervated skeletal muscle in vivo. Taken together, these results represent

an important first step toward development of a novel tissue engineered muscle repair

treatment for potentially expanded clinical applications to VML injuries.

[1] Grogan BF, Hsu JR, Skeletal Trauma Research Consortium. Volumetric Muscle

Loss. J Am Acad Orthop Surg 2011;19:S35–7.

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CHAPTER I

INTRODUCTION

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1. Clinical significance of volumetric muscle loss and peripheral nerve injuries

In humans, muscles constitute nearly half of the body's mass [1] and are critical

to daily life. Skeletal muscle, the most common type in the body, is responsible for up to

85% of heat production and, through its anchorage to other structural elements - notably

the skeleton - movement of body parts leading to ambulation, breathing, and other

essential tasks [1]. To enable these functions, muscles are spread throughout the body

and are a particularly prevalent component in the extremities, which serve as the drivers

of locomotion.

Unfortunately, extremity injury is incredibly common in the current clinical setting

[2]. Advances in motor vehicle restraint systems in the civilian setting, and similar

advances in body armor in the military setting, have saved lives of patients who would

have otherwise died of their wounds - but these survivors now often present to the clinic

with extremity trauma [2,3]. This places a large financial burden on the health care

system. For example, when ranking resource utilization by injury in casualties from US

armed conflicts in Iraq and Afghanistan, extremity trauma comprised 65% [2]. This

proportion was estimated to correspond to costs totaling roughly $1.2 billion [2].

The vast majority of military injuries are caused by explosions [3], which are

associated with significant damage to extremity soft tissues [4], including muscle.

Injuries of this type can be so severe that they result in persistent functional impairment,

which is termed volumetric muscle loss (VML) [5]. Though the single greatest cause of

damage to skeletal muscle is trauma [3,5–8], it can also result from congenital or

acquired disease or tumor resection [9]. This places VML as a common, and costly,

problem in both military and civilian settings.

There are also over 50,000 surgical procedures for peripheral nerve injuries

reported annually [10] in both civilian and military settings due to motor vehicle

accidents, knife injuries, gunshot wounds, and other sources of trauma [11,12]. Injuries

3

to peripheral nerves are also quite debilitating, resulting in impairment or loss of function

in tissue they innervate — for example, weakness or paralysis of a body part by inability

to signal skeletal muscles to contract.

Co-morbid injuries of a critical size involving both skeletal muscle and the

innervating peripheral nerve represent an especially aggravated class of VML termed

total compartment loss [5]. Unsurprisingly, the prognosis for an injury of this extent is

extremely poor, with bracing by an external load-bearing device the only currently-

envisioned treatment strategy [5].

2. Development of the neuromuscular junction

The neuromuscular synapse is a highly organized structure designed to

efficiently transfer an electrical depolarization propagated down a nerve axon into a

resultant electrical event in the target muscle fiber by way of the chemical transmitter

acetylcholine (ACh) [1]. To establish and maintain this elegant function, the synapse

must be assembled by a carefully coordinated series of events. This begins in the

developing embryo, where differentiated muscle fibers appear prior to innervation of the

region by motor nerve axons [13]. These myofibers, and indeed less mature muscle cells

called myotubes (see Section 4) which are long, multinucleated cells that lack the size

and striations of myofibers, express acetylcholine receptors (AChRs) within their cell

membranes [14]. AChRs are initially "prepatterned" into a concentration of clusters near

the middle of the cell [13,15,16]. Formation of these prepatterns require the molecules

low-density lipoprotein receptor-related protein 4 (Lrp4) and muscle-specific kinase

(MuSK) [17]. Upon migration of the motor axon to the myofiber, these prepatterns have

been suggested to dictate where the axon grows and where the synapse between the

two cells will form [15]. Though a muscle fiber may initially be innervated by more than

4

one axon early in development, shortly after birth these excess connections will have

disappeared, leaving one axon to innervate each fiber [16].

Agrin is a heparan sulfate proteoglycan which, in synaptogenesis, is one of the

primary molecules responsible for accumulation of muscle AChRs directly opposite the

axon terminal. Agrin is also expressed in many other tissues and features many

isoforms. Splice differences near agrin's C-terminal dramatically alter the effects of the

molecule, the most relevant of which for the purpose of this review is an amino acid

insert at a location termed the z-site in mammals [14,18]. This z-site insertion is critical to

the behavior of neuronal agrins [14,18], and agrins carrying it are referred to as z+ agrin

where appropriate to facilitate discussion of this differential status. The work described

here is concerned solely with z+ agrin; therefore, any further references to "agrin"

indicate this splice variant. Many suppliers (e.g. R&D Systems, Minneapolis, MN)

produce recombinant C-terminal agrin fragments (sometimes referred to as mini-agrins)

based on this z+ isoform which are suitable for experimentation.

Agrin is synthesized within the cell body of the motor neuron but is subsequently

transported down the axon and anchored within the synaptic basal lamina [14,17,18].

This agrin then binds to Lrp4 on the muscle fiber surface, which in turn associates with

MuSK [17,19]. This association results in the concentration of AChRs underneath the

axon terminal [14], in the area of the muscle membrane that becomes the post-synaptic

structure or motor end plate (MEP). This process likely occurs by at least four different

mechanisms, all of which are linked to the agrin signal to some extent: first, AChRs

which normally randomly diffuse within the membrane become trapped when they

contact a growing aggregate of other AChRs [14]; second, the average time of individual

AChR molecule residence in the muscle membrane increases [14]; third, muscle nuclei

near the synapse specialize to express genes encoding AChR subunit proteins, thereby

increasing total AChR count in the membrane nearby [14,18]; fourth, AChR subunit

5

expression is suppressed in nuclei distant from the membrane [14], potentially due to

cell-wide inhibition resulting from ACh stimulation [16].

The initial assembly of the synapse leads to further differentiation both of the

presynaptic neuron and the postsynaptic muscle fiber. Lrp4 acts as a muscle-derived

retrograde differentiation signal to the motor axon [20]. Similarly, downstream effects of

MuSK activation by agrin-Lrp4 cause the cell membrane at the MEP to remodel into an

arrangement of invaginated junctional folds, with AChRs concentrated at high density at

the crests of the folds [14]. The structures - both nerve and muscle - comprising the

mature synaptic apparatus are collectively called the neuromuscular junction (NMJ).

Within the NMJ, the two apposing cell membranes remain separated by a space roughly

60-100 nm wide known as the synaptic cleft [1]. This cleft is isolated from fluid of the

surrounding tissue by a Schwann cell, which wraps the entire NMJ [1].

In summary, agrin signaling is a powerful regulator of neuromuscular synapse

assembly and, therefore, of vital importance to proper function of motor units that are

required for ambulation, breathing, and other required bodily functions.

3. Relevant aspects of structure and function in skeletal muscle and peripheral

nerve

Each skeletal muscle is composed primarily of numerous contractile muscle

cells, called myofibers, which can be oriented within the overall structure in a wide

variety of ways depending on the function of the muscle [1]. For example, myofibers

within fusiform muscles such as the biceps brachii have a largely parallel arrangement

designed to produce force in a single direction, while those within a circular muscle such

as the orbicularis oculi are arranged in a loop in order to produce an opening or closing

motion [1].

6

Within the muscle, these microscopic myofibers are sheathed in loose connective

tissue termed the endomysium [1]. Groups of myofibers together form fascicles - parallel

strands of muscle large enough to be visible to the unaided eye - which are in turn

sheathed in thicker connective tissue known as perimysium [1]. Surrounding the entire

muscle compartment is yet another layer of connective tissue termed the fascia, which

connects to the perimysium via an interstitial layer called epimysium [1]. These layers of

connective tissue surround and penetrate throughout the muscle to offer organization

and mechanical support, but are also continuous with muscle attachment structures,

such as tendons, which transmit force from the muscle (which generates force by

contraction) to the skeleton (which typically moves as a result) [1].

Within the structure of the skeletal muscle are other tissue types - notably nerve

and blood vessels [1]. Though veins, arteries, and large nerves typically lie wholly

outside the muscles, smaller branches thereof are carried inside muscle compartments,

with larger structures within the perimysium and capillaries and terminal nerve fibers

within the epimysium [1]. Blood vessels are required to be spread throughout muscle in

order to supply nutrients to and eliminate waste from individual muscle cells [1] due to

the limits of diffusion, which in the case of metabolically active cells may be inadequate

at distances greater than 100 μm [21]. Likewise, nerves must heavily infiltrate muscle in

order to signal via the NMJ, a structure which requires proximity between the muscle

and nerve cells on the order of tens of nanometers as discussed above.

Much like muscle, peripheral nerves are composed of numerous nerve fibers, or

axons, aligned into a larger whole [22]. Nerves also feature connective tissue -

endoneurium surrounding each fiber, perineurium around fascicles composed of several

fibers, and epineurium surrounding the entire nerve [22] - in structure and arrangement

incredibly similar to those of skeletal muscle, if not necessarily similar in size.

7

One motor neuron and all the skeletal muscle fibers it innervates are collectively

known as a motor unit [1]. All muscle fibers of a motor unit contract simultaneously when

stimulated by their innervating neuron [1] in a process known as excitation-contraction

coupling. When a nerve impulse reaches the NMJ, the neurotransmitter ACh is released

from synaptic vesicles into the synaptic cleft [1]. ACh diffuses across the cleft and binds

to AChRs accumulated at the MEP, beginning a cascade which causes the postsynaptic

muscle cell to contract [1]. ACh in the synapse is rapidly broken down by

acetylcholinesterase present in the synaptic muscle membrane and basal lamina,

ensuring that muscles relax as soon as the nerve ceases releasing ACh [1].

4. The native wound healing process in injuries to skeletal muscle

Skeletal muscle possesses an impressive capacity for self-repair from traumatic

injury [23–25]. Though often described between sources using slightly different

nomenclature, the progression of endogenous muscle repair can be broken down into

phases, which can overlap but always proceed in a specific order. These are, in order,

the destruction / inflammatory phase, the activation / repair phase, and the maturation /

remodeling phase [24,25]. The initial insult in the destruction phase ruptures myofiber

cell membranes. This rupture would result in the necrosis of the entire length of the

myofiber (sometimes up to centimeters in length) if not for a specialized structure known

as the contraction band, which restricts the damage only to the site of insult [26]. The

rupture of the cell membrane releases intracellular components to the extracellular

space, which in turn attracts inflammatory macrophages to the injury site [27]. At the

injury site, macrophages first remove cell debris by phagocytosis but then undergo a

switch to an anti-inflammatory phenotype, releasing factors that activate the proliferation

and differentiation of satellite cells [24,25,27].

8

Satellite cells are stem cells which normally reside in a quiescent state under the

basal lamina (i.e. the layer of extracellular matrix (ECM) below the periphery of

myofibers) [28]; for review, see [29]). Unlike myofibers themselves, satellite cells are

capable of asymmetric division, giving rise both to more satellite cells [30] and to

progeny called myoblasts that incorporate into myofibers [31–33]. Numerous studies

have demonstrated the impressive ability of satellite cells to self-renew [30,34–36], in

some cases giving rise to ~11 times the original number of satellite cells and a number

of myofiber nuclei besides [30]. Though other stem cell populations [37] or even de-

differentiated myofiber nuclei [38] have been proposed to contribute to skeletal muscle

repair, subsequent studies have demonstrated that satellite cells are required for this

process [29].

Myoblasts can either fuse with each other to form myotubes [31] or with existing

myofibers to replace nuclei lost during injury [33], thereby enabling the myofiber stumps

to extend inward from the injury edges [23,24]. While cellular muscle repair is

proceeding from the injury borders, the gap left by the rupture of myofibers first fills with

a hematoma before being invaded by macrophages and, subsequently, fibroblasts

[25,26,39]. These form a non-contractile connective tissue scar [24,25] composed

initially of fibronectin and type III collagen but later of an increasing proportion of type I

collagen [40]. The repair phase concludes when the extending myofiber stumps come

into contact with the expanding fibrotic scar [24,25]. Thereafter, maturation of the repair

occurs through the creation of new myotendinous junction-like structures linking the

myofibers to the scar to create a load-transferring structure [41,42], while over time the

repair will remodel by the reduction in thickness of the scar [24,41].

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5. Limitations of the native wound healing process in injuries to skeletal muscle

and peripheral nerve

Unfortunately, there are several steps during native skeletal muscle wound

healing that can be radically altered by the infliction of a large (i.e. VML) injury. The first

is that the proper progression of the steps above assumes that the basal lamina

surrounding the injury remains intact to serve as a physical guide for cellular activity [25].

However, larger injuries - especially those resulting from tissue transection, or VML

injuries such as ablation or excision, rather than blunt impact - are associated with

destruction of portions of the basal lamina along with more extensive necrosis of

myofibers. Without the basal lamina, the normal repair process can still lead to aberrant

anatomy due to the absence of appropriate environmental cues. For example, separate

myotubes formed by satellite cells within the same region can fuse incompletely, forming

a larger fiber which "forks" into two smaller fibers [25].

Further, stem cells within the injury site can differentiate toward a myofibroblast

lineage instead of the preferred myogenic lineage - thus, instead of fusing with the

stumps of injured myofibers, these cells act to contract the wound bed and overproduce

extracellular matrix that increases the size of the connective tissue scar [43]. Though

cells with this phenotype appear as early as 1 week after injury, their proportion within

the population increases over time until, after ~5 weeks, they have almost completely

supplanted myogenic cells in the injured site [43]. Thus, large injuries that take longer to

heal are more likely to develop an extensive, non-contractile scar where there was once

functional muscle. Importantly, this is the definition of VML - the "traumatic or surgical

loss of skeletal muscle with resultant functional impairment" [5].

Though peripheral nerves also have a significant innate capacity for self-repair

due to the ability of axons to migrate through endoneurial tubes left empty by Wallerian

degeneration distal to the injury [44], more severe injuries often leave a physical gap or

10

barrier of scar tissue that axons cannot easily cross. These cases require surgical

intervention for function to be restored [10,44]. Small segmental injuries can be repaired

via surgical coaptation of nerve ends; however, surgical repair placing tension on the

nerve has been linked to poor clinical outcomes [10,44]. This requirement for tensionless

repair defines the critical size for nerve injuries, beyond which native wound healing is

insufficient to repair tissue loss.

Peripheral nerve injury incidence and healing are of interest in VML not just

because muscle cannot be stimulated to contract without innervation, but also because

the regeneration of muscle injury on a cellular level can only partially progress in the

absence of innervation [24,45]. Moreover, the connective tissue scar formed after

muscle injury must be navigated by axon sprouts if innervation of distal myofibers is to

be accomplished (see Fig. 7 of [25]). By contrast, nerves typically regenerate by

migration of the axon growth cone through a favorable environment consisting of

neurotrophic and other environmental cues in the basal lamina left behind by Wallerian

degeneration [44]. Taken together, these indicate that fibrosis observed in VML injuries

could serve as a barrier to reinnervation and, therefore, optimal functional repair of the

muscle. This would be aggravated in co-morbid injuries of both muscle and an adjacent

innervating nerve.

6. Current clinical treatment of VML

Surgical treatment of VML is possible in some cases by free tissue transfer

[5,46–48], a procedure in which muscle tissue is harvested from one site within the

patient and then implanted at the injured site [48]. Transfers of this type are

advantageous because they can be completed within one procedure period and can

support innervation of the transferred tissue [49]. One requirement for successful free

tissue transfer is the presence of a vascular pedicle consisting of an artery and one or

11

more veins of appropriate length and caliber [48]. The implant is revascularized and

potentially reinnervated at the recipient site by connecting to local blood vessels of the

appropriate type and a source of donor axons, respectively, thereby providing blood

supply and an opportunity for neo-innervation to the graft [48].

Unfortunately, free muscle transfer is limited by the ability to locate donor tissue

of sufficient vascularity [48] and innervation [48,49] in an already-injured patient, and by

the associated, likely permanent morbidity at the harvest site [5,48]. Wound geometry is

a concern, as muscles available for transfer with large coverage area (e.g. the latissimus

dorsi at sizes up to 25 cm wide x 40 cm long, [48]) may be thin, and therefore incapable

of filling a thick defect. Even with modern advanced microsurgical techniques [48],

complication rates can approach 50% [49] and cosmetic scarring may limit patient

satisfaction [48].

Another, more limited type of tissue transfer is the rotational muscle flap. This

procedure is similar to free flap transfer, and has many of the same limitations [50], but

in this case a relatively dispensable donor muscle can be located in close proximity to

the injured site [51]. This allows the donor muscle to be dissected from most of the

surrounding tissue, but not completely removed [51]. Critically, the primary blood vessel

and, if possible, a nerve supplying the muscle remain attached and serve as the point

around which the flap is rotated to cover the injured site [51]. Another substantial

limitation of rotational flaps, beyond the requirement for donor and injury sites in

proximity, is that they typically cannot cover large defects [52]. Examples of this

technique include rotation of the inferior segment of the trapezius onto a craniofacial or

neck defect [51], rotation of the latissmus dorsi onto a deltoid defect [53], and rotation of

the sartorius to cover a femoral defect [54].

Beyond these already significant limitations, flap transfer is a technically complex

procedure, requiring an experienced surgical team and careful patient selection [5,48]. In

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complex pathophysiologies that do not meet these criteria, amputation is the default

alternative [48]. These limitations in the standard of care for VML injuries combine with

their frequent presentation, as described in Section 1, to create a significant unmet need

in modern medicine for muscle reconstruction, repair, and replacement.

7. Single-component approaches in development for the repair of VML injuries

Several strategies are under investigation in an attempt to improve the standard

of care for VML injuries. The use of various biomaterial scaffolds alone has been

reported in multiple models of skeletal muscle repair [55–59], with varying results. One

potential advantage of scaffolds derived from ECM is that they can contain molecules

that provide mechanical support and mediate cell adhesion [60]. Moreover, these

scaffolds can be degraded and remodeled by endogenous processes at the implant site

[60]. In an example ECM-derived approach, a sponge composed of type I (~70%) and

type III (~30%) collagen had hemostatic and anti-inflammatory properties when placed

into a wounded muscle, but did not induce significant regeneration of muscle fibers [56].

One notable class of scaffold-only studies implanted decellularized muscle matrix

in an injury model. This approach was reported in some cases to be ineffective at

promoting functional recovery without the addition of cells [57], instead being remodeled

into fibrous tissue by native processes [55]. However, though the new tissue generated

by remodeling was non-contractile, it was somewhat counterintuitively shown on at least

one occasion to recover one-third the functional deficit created by the injury [59]. This

mechanism was termed "functional fibrosis," and presumably operates by shielding

remaining muscle mass from overload injury [59].

Of note, one scaffold-only repair has been performed in a human [61]. In this

study, an ECM scaffold derived from porcine small intestine submucosa was implanted

into a VML injury in the thigh of a marine. At 16 weeks postoperatively, torque generated

13

by the affected limb increased by a modest amount, while that generated by the

contralateral limb decreased slightly - presumably as it was forced to compensate less

[61]. At 36 weeks postoperatively, some new tissue formation was observed at the

implant site [61]. However, even this degree of success was higher than most studies in

this category - as described above - and was doubtless affected by careful patient

selection and a highly specialized surgical staff.

Similarly, pharmacological and / or genetic approaches (i.e., regenerative

pharmacology) have been investigated as a potential therapeutic for skeletal muscle

repair [62–66]. One study investigated mitigating the inflammatory response following

muscle injury by using anti-inflammatory or antioxidant small molecules, but concluded

that these drugs had no significant effect on function [62]. Another report demonstrated

that intramuscular injection of transgenic DNA-delivering microspheres could stably

induce transgene expression in muscle cells in vivo, though the DNA in this case

encoded for luciferase so no commentary could be made regarding changes to muscle

function [63]. In a further study using models of atrophic muscle in vitro and in vivo,

delivery of exogenous macrophage colony-stimulating factor increased functional protein

expression, potentially through enhancement of the anti-inflammatory macrophage

phenotype [64]. Losartan, a medication normally prescribed for hypertension, was shown

in a murine injury model to histologically and functionally improve muscle repair following

administration post-injury [65]. Though not a traditional pharmacologic per se, hyperbaric

oxygen treatment has been used in athletes with muscular injuries and was shown in a

rat injury model to increase force production and the prevalence of markers for satellite

cell proliferation and myofiber maturation [66]. Finally, a number of growth factors have

been investigated as candidates for exogenous delivery to supplement the native muscle

repair process (for review, see [67]). These have typically been hypothesized either to

14

promote myogenesis or to inhibit TGF-β1-induced fibrosis, and largely had the desired

effect, though a risk of side effects was documented.

8. Multi-component tissue engineering approaches in development for the repair

of VML injuries

Due to the potential limitations of materials, cells, or pharmacologics alone,

tissue engineered grafts or their analogs in vitro often feature two or more of these

components ([57,68–85]; see reviews [86,87]). Such approaches attempt to mimic the

structure and efficacy of autologous tissue transfer without the limitations associated

with harvest from a donor site. However, the degree to which these approaches

incorporate multiple components varies greatly.

In one example, tissue engineered skeletal muscle was created in vitro by

culturing a monolayer of isolated muscle cells, treating the monolayer with low-serum

differentiation medium supplemented with TGF-β1, and then initiating self-assembly of a

muscle strip by presenting the monolayer with opposing tendon fragments [83]. This

strategy represented the combination of a single cell type treated with pharmacologics,

but in the absence of a biomaterial scaffold. Similarly, an acellular combination

biomaterial-pharmacologic strategy designed to recruit and direct myogenic host cells

from the surrounding environment was evaluated in vivo by loading gelatin with

exogenous growth factors before implanting the construct in a rat model [88].

The vast majority of tissue engineering approaches, however, consist of a cell-

seeded biomaterial scaffold. These scaffolding materials are often designed to degrade

in vivo, being broken down and removed from the implant site over a time course that

corresponds to new tissue formation during wound healing / regeneration / repair [87].

Depending on the biomaterial, this remodeling of the scaffold may release myogenic

degradation products composed of the scaffold subunits themselves or, in the case of

15

scaffolds derived from tissue, of growth factors and other molecules previously trapped

within the material [89].

One important design consideration when selecting a biomaterial for tissue

engineering is its mechanical stiffness, which is quantified by the elastic or Young's

modulus (abbreviated E). This matrix stiffness has been shown to directly affect

adhesion and differentiation of muscle cells [79]. Another consideration is the chemical

composition of the scaffold surface, which has been shown to affect expression of

muscle proteins [80] and muscle cell force generation over time [75], presumably

mediated by the differing strength of cell-matrix interactions among substrate

chemistries.

This cell-seeded biomaterial construct can be further functionalized with one or

more additional cues designed to alter cell behavior. Pharmacologics, as described

above, are often incorporated, and may interact synergistically with other cues presented

by the biomaterial [87]. Critically, however, few studies to date have incorporated

exogenous agrin into muscle tissue engineering strategies [78,82]. One notable study

evaluated the presentation of soluble agrin to tissue engineered muscle constructs in

vitro [78]. In this case, agrin presentation had no effect on cell number, myosin

expression, or calcium dynamics, but increased the number of AChR clusters in scaffold-

resident cells, and most importantly increased contractile force generation of the

construct [78]. In a separate study, soluble agrin was used to condition C2C12 mouse

myoblast cells grown within a fibrin gel in vitro, which were subsequently implanted in

vivo in contact with a transected nerve [82]. Histological evaluation in this study indicated

that pre-treatment with agrin may have enhanced angiogenesis and NMJ formation (i.e.,

colocalized areas of neurofilament and AChR staining) in the area of the implant, but

critically did not result in well-oriented myofibers [82].

16

As such, initial efforts indicate that agrin may be a promising pharmacological

supplement for muscle tissue engineering. However, it has yet to be used in a non-

soluble embodiment analogous to its stably tethered location in normal NMJs [18], and

further has not been evaluated as an implanted component of the tissue engineered

construct in vivo - both of which may be required to provide optimal benefit to muscle

function.

Additionally, physical cues provided by the biomaterial such as microfibers [81],

microgrooves [70,79,80], and high aspect ratio pores [71], are another class of

functionalization. Importantly, aligned physical cues have been demonstrated to not only

orient skeletal muscle cells in a desired direction - such as a preferred direction of

contraction - but also to encourage myoblast fusion and expression of contractile

proteins [79–81]. In one noteworthy example, a cell culture substrate interrupted by a

series of posts was used to grow extensive hydrogel-cell networks [72]. In this case, the

size, spacing, and shape of the posts - which patterned pores within the resulting

construct - was found to affect the viability, alignment, and protein expression of

scaffold-resident muscle cells [72]. Thus, scaffold patterning is an incredibly powerful

means to modify muscle cell behavior for tissue engineering applications.

9. The importance of mechanical stretch in modulating muscle cell phenotype in

vitro

Like patterning, mechanical stretch applied in vitro is a cue which has been found

to enhance myogenesis within engineered muscle [68,90–95]. Early experimentation in

mechanical alteration of muscle cell phenotype aimed to mimic tensile forces

experienced in vivo [91,92,96]. In these studies, cyclic mechanical force was applied to

elastic biomaterials seeded with myoblasts, leading to stretch of the culture substrate

and the attached cells. Results indicated that mechanical stretch enhanced multiple

17

markers of muscle cell proliferation and maturation, including DNA content, protein

production, myoblast fusion, average myotube cross-sectional area (hypertrophy), total

myotube cross-sectional area (above the increase in per-cell area, indicating increase in

myotube count), and striations [68,91].

Recent efforts of our lab have expanded upon these findings using a

decellularized porcine bladder submucosa scaffold referred to as bladder acellular matrix

(BAM) [93–95]. This scaffold lends itself well to cell seeding and subsequent

preconditioning (i.e., cyclic mechanical stretch) in a bioreactor. We refer to this

multidisciplinary strategy as tissue engineered muscle repair (TEMR). Initial

experimentation using the TEMR construct indicated that sufficient preconditioning not

only increased the contractile force generated by TEMR constructs upon electrical

stimulation in vitro, but also that preconditioned constructs implanted in vivo for 2-4

weeks also generated more force than unconditioned controls [93].

Further studies evaluated the ability of TEMR to morphologically and functionally

repair a VML defect in the mouse latissimus dorsi (LD) [94,95]. The first of these found

that TEMR significantly restored 60-70% contractile function to injured muscles when

compared to unrepaired and scaffold-only controls [94]. Histologically, explanted TEMR

constructs also evidenced significant increases in markers of muscle repair and blood

vessel infiltration of the injured site, and may have supported axon infiltration at the

interface between TEMR and the remaining LD [94]. The second study confirmed the

essential nature of mechanical preconditioning to functional repair of an injury by

comparison to a cell-seeded but statically-cultured control [95]. Further, this study found

that an additional cell seeding step performed mid-way through preconditioning

enhanced TEMR cellularity, multinucleated cell count, and relevant protein expression

prior to implantation, as well as force production by explants at longer time points post-

18

implantation [95]. These observations were presumably due to the creation of distinct

"proliferating" and "differentiating" cell sub-populations within the construct [95].

Therefore, the TEMR strategy has documented utility in the repair of model VML

injuries. Creation of a platform of skeletal muscle tissue engineering technologies,

though, could enable treatment of a wider variety of injuries, such as thicker defects that

do not correspond in shape and size to seeded BAM. Further, additional

functionalizations as described above could be incorporated into the TEMR strategy to

repair the remaining 30-40% functional deficit between TEMR and the uninjured muscle.

As a final note, there are currently few tissue engineering strategies for muscle

repair that incorporate neural innervation / re-innervation as part of the core technology.

Given the importance of neural input to muscle formation, repair, physiology, and

function, this seems to be a key technological barrier to more widespread clinical

applications of tissue engineering for muscle repair. Moreover, an approach designed to

repair both components of total compartment loss injuries would be a huge step forward

for this unmet medical need.

10. Overview of Dissertation

The objective of this dissertation was to create a tissue engineered alternative to

autografting, lacking donor site morbidity and having the potential to restore function by

physical guidance of nerve axons and/or muscle progenitors and pharmacological

maintenance of a muscle phenotype similar to that seen in innervated myofibers. To this

end, we proposed a multidisciplinary tissue engineering approach incorporating

biomimetic materials, cell supplementation, and pharmacological signaling that

would represent an important step toward expanded clinical indications for

treatment of VML injuries. The basis of this technology was utilization of a biomaterial

with suitable physical properties for tunable patterning to a desired architecture, in order

19

to better recapitulate the anatomy of muscle tissue. Within this context, cell phenotype

could be further modulated by mechanical loading, delivery of exogenous cues, or a

combination of the two.

The central hypothesis of this dissertation was that biomimetic cues could be

leveraged to create a tissue engineered construct in vitro that would accelerate skeletal

muscle tissue formation and function following implantation in vivo, and furthermore,

maintain the skeletal muscle phenotype in vivo during reinnervation of VML injuries. The

specific cues under investigation included physical patterning to guide recapitulation of

native microanatomy, as well as chemical signaling and mechanical strain to promote

and maintain a more mature cellular phenotype in vitro and in vivo.

Therefore, this dissertation featured three broad objectives:

Objective 1: Construct a three-dimensional biomaterial scaffold with architecture

that would support aligned tissues such as nerve and muscle to encourage tissue

formation resembling native anatomy. The material would be characterized and its

performance evaluated by using an in vitro model for axon extension.

Objective 2: Functionalize the patterned biomaterial for skeletal muscle repair by

pharmacologic / chemical induction and maintenance of acetylcholine receptor clusters

with agrin, a hallmark of a more mature, innervated muscle phenotype in vitro.

Objective 3: Apply the fibrin construct featuring agrin-stimulated muscle cells to

a rodent neurotization model to assess its ability to maintain AChR clusters and support

innervation in the complex environment seen in vivo.

A summary of this approach is presented schematically in Fig. 1.

10.1. Rationale and research design of Objective 1

The templated fibrin biomaterial that served as the basis for the tissue

engineered muscle repair strategy described here was first developed in the context of a

20

Fig. 1. Schematic of final tissue engineered muscle construct fabrication and evaluation. (A) Pharmacologic delivery vehicles were formed by immobilization of agrin on a microcarrier surface (see Chapter III). These agrin carriers were then be suspended in a solution of fibrinogen, which was (B) polymerized to fibrin around a sacrificial template of polymer fibers (see Chapters II & III). Selective dissolution of the template resulted in a patterned hydrogel material that presented agrin via bound microcarriers. The patterning process created a porous microarchitecture within the scaffold, which was (C) seeded via perfusion of a suspension of muscle-derived cells (see Chapter IV). Resident cells then received a combination of physical cues from scaffold conduit pores, pharmacologic cues from scaffold agrin, and (D) mechanical cues from cyclic stretch in a bioreactor (see Chapter III). The final construct was then (E) evaluated in vivo using a rodent neurotization model (see Chapter V).

21

potential nerve graft. However, due to similarities in connective tissue architecture

between nerve and muscle as discussed in Section 3, it was also an ideal candidate for

muscle repair (see Objective 2). Further, the ability to support axon infiltration would be

useful in models of innervation in vivo (see Objective 3). Thus, Objective 1 centered on

characterization of patterned fibrin scaffolds and evaluation of their support of axon

extension in vitro.

Objective 1 featured two subordinate goals. The first was to adapt a fibrin

templating approach [97,98], which had been previously used by members of our team,

for creation of a biomaterial scaffold featuring biomimetic aligned pores. As sacrifical

templating requires only that the elected biomaterial and template feature differing

solubility in a chosen solvent, numerous natural and synthetic polymers are compatible

with this approach [99–102]. However, fibrin was elected for numerous reasons, as

follows:

• It has been highly studied as a tissue engineering substrate, not only for repair of

peripheral nerve [103–107] but also for skeletal muscle

[69,72,73,75,76,84,108,109].

• Fibrin serves as an endogenous provisional scaffold after injury and is therefore

a component of the native wound healing process; in peripheral nerve

regeneration, it is laid down in a cable-like fashion ahead of the growth cone as a

template for neural growth [110].

• It is naturally degradable into non-toxic byproducts and is capable of being

autologously sourced [97] to minimize potential foreign body response.

The sacrificial templating approach, in turn, has the advantage of generating a

customizable porous network corresponding to any possible arrangement of the

templating material. Wallerian degeneration leaves a series of axially-aligned lumens

formed by the connective tissue of the endoneurium, which serves to guide native

22

reinnervation after injury [44]. Aligned cylindrical conduits patterned within fibrin could

mimic the shape of these endoneurial tubes and potentially provide a substrate

conducive to axon migration.

The second goal of Objective 1 was to measure neurite length within these

scaffolds over time in vitro in order to determine the potential effect of conduit diameter

on the rate of neurite growth. This was relevant because the supportive expression of

growth associated genes in the neuron is transient after axon insult [111], making speed

of axon migration through a tissue engineered graft toward any resident muscle cells to

be innervated a critical concern. To evaluate this rate, we employed an embryonic

chicken dorsal root ganglia (DRG) model [112].

As a final design consideration, we aimed to incorporate scaffolds with elastic

modulus similar to that of native soft tissue - initially nerve, and later skeletal muscle -

because matrix stiffness has been shown to modulate differentiation of adherent cells

[113,114]. The findings of this objective were designed to inform material suitability for

later objectives, in terms of scaffold physical and mechanical properties for Objective 2

and estimated axonal ingrowth in vivo for Objective 3.


Though we have discussed scaffold fabrication primarily in terms of its

application to nerve grafting, it was also expected to result in parameters suitable for

muscle repair. Sacrificial patterning, for example, has the potential to produce scaffolds

with porosity in excess of 75%-90% [97], which would offer high surface area within

pores to enable robust cell seeding. Moreover, many skeletal muscle defects are

irregularly shaped, not only because of the inherent variability in anatomy amongst

human muscles (see Section 3) but also because they are often traumatic in origin [3,5–

8]. The enzymatic polymerization of fibrin could be leveraged to form the scaffold and its

23

associated pore network into any shape that can be molded and patterned, respectively,

a distinct advantage over more geometrically restrictive fabrication methods. Finally, use

of a fibrin substrate for muscle cell seeding has been shown to result in tissue

engineered muscle bundles with promising force generation in vitro [75].

Objective 2 featured four subordinate goals. First, we attempted to elicit AChR

clustering in vitro, which occurs in muscle cells presented with neural agrin in normal

physiology (see Section 2). Denervated muscle cells undergo significant atrophy in

cases of chronic denervation [24]. Moreover, clinical expectations for axon growth during

reinnervation of injured tissue are roughly 1 mm per day [111], resulting in potential

denervation for weeks or even months in sites substantially distant from the source of re-

innervation (e.g., the bulk of large injuries). If maintenance of AChR clustering created in

vitro (the presumptive correlates of MEPs in vivo) could be maintained in vivo, then

perhaps this approach could be used to counteract the atrophic effects of muscle

denervation during processing in vitro and colonization of the construct by host axons in

vivo after implantation. In this scenario, the capacity of the engineered construct to

restore function following implantation in vivo could ultimately be enhanced.

As a first step towards this goal, we investigated incorporating exogenous agrin

into the fibrin material in a manner that maintained its bioactivity on scaffold-seeded

muscle cells in vitro. As a vehicle to present agrin to cells, we selected microparticles

with surface-bound agrin molecules. This embodiment was chosen due to its similarity to

normal physiology, wherein neural agrin molecules are stably inserted within the

synaptic basal lamina after being synthesized by the nerve axon [18], an arrangement

mimicked more closely by surface-bound signal than by, for example, a soluble signal.

This strategy was initially evaluated in a 2D analog of the fibrin scaffold using a C2C12

mouse myoblast model system.

24

Second, we seeded 3D fibrin scaffolds as fabricated in pursuit of Objective 1 with

cells isolated from rat muscle explants. A tissue engineering strategy relevant for human

use would preferentially incorporate autologously sourced cells in order to minimize the

risk of immune rejection. Compared to the C2C12 cell line, then, expanded primary cells

were thought to be a more representative pre-clinical model. Additionally, higher cell

seeding densities have previously resulted in increased myoblast fusion and eventual

development of striated myofibers [84]. Accomplishing this robust seeding in 3D with

muscle cells, however, was expected to be challenging, as traditional static addition

(e.g., manual pipetting) of a cell suspension has been shown to be inefficient at seeding

cells within the interior of a porous scaffold [115]. Our strategy to overcome this obstacle

was through the use of flow perfusion of cell suspensions through the scaffold pore

network. In the literature, perfusion has most often been reported as a mechanism to

enhance nutrient exchange or modulate cellular phenotype in pre-seeded cells [116–

122], but has also been used to seed cells more deeply and uniformly within scaffolds

than static seeding [21,123–126]. Importantly, however, these prior studies evaluated

other combinations of cell type, scaffold material, and / or scaffold architecture than that

dicussed here, making the current effort a novel investigation.

The third goal of Objective 2 was to wed the advances of the first two goals,

combining agrin presentation with a highly cellularized muscle construct. To accomplish

this, we immobilized agrin-bound microparticles within the walls of the fibrin scaffold,

predicting that some proportion of these particles would extend partially into the pore

lumens and there make contact with seeded cells. This strategy would permit control of

agrin "dosing" as well as the stoichiometry of agrin particles relative to seeded myoblasts

and / or myotubes - both could be accomplished by adjustment of the microparticle

density incorporated into the fibrin scaffold.

25

Finally, as our fourth goal, we evaluated the effect of bioreactor preconditioning

via mechanical stretch on the cell-seeded agrin-presenting tissue engineered construct.

The rationale for this approach is related to observations (see Section 9) that mechanical

stimulation in vitro can encourage a more mature muscle phenotype as measured by: 1)

skeletal muscle protein-related second messenger activity [92], 2) increased average

muscle cell diameter [68,91], and 3) sum cross-sectional area of myofibers [68]. Thus,

we hypothesized that it might also affect AChR clustering either alone or synergistically

in the presence of agrin.


In Objective 3, we evaluated the characteristics of the optimized tissue

engineered muscle construct developed in pursuit of Objective 2 following implantation

in vivo using a rat neurotization model. The primary goal of this study was to determine if

agrin presentation would maintain AChR clusters in the complex in vivo environment.

Accordingly, we hypothesized that, due to prior optimization in vitro (Objectives 1 and 2),

the construct would better support axon ingrowth and augment muscle tissue formation

following implantation in vivo. To this end, we coapted the construct to a transected

nerve to see if it would yield self-assembled neuromuscular junctions (NMJs) linking

implanted muscle cells to the host nervous system. In this rat neurotization model, the

proximal stump of the femoral nerve was diverted to our ectopically implanted optimized

muscle construct (Fig. 1). This preliminary study was designed to provide important

information regarding the potential utility of the current technology to restore function in a

more complex VML injury model in vivo.

26

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CHAPTER II

THE PROMOTION OF AXON EXTENSION IN VITRO USING POLYMER-TEMPLATED

FIBRIN SCAFFOLDS

John B. Scott, Mehdi Afshari, Richard Kotek, Justin M. Saul

The following chapter has been published in Biomaterials (2011 Jul;32(21):4830-9).

Citations, figure captions, and other stylistic elements of this chapter conform to the

demands of the publishing journal, and as such may differ from those found elsewhere in

this dissertation. The published version can be seen online at

http://www.sciencedirect.com/science/article/pii/S0142961211003061.

Elsevier, the publisher of Biomaterials and copyright holder of this chapter, has granted

John Scott permission to reproduce it as part of this dissertation (see “Author Use” at

http://www.elsevier.com/journal-authors/author-rights-and-responsibilities#author-use).

As primary author, John Scott performed experiments and prepared the manuscript for

all content in Chapter II except template fiber fabrication and the authorship of methods

relevant thereto (see first paragraph of Section 2.1), which were performed by Mehdi

Afshari and Richard Kotek.

38

Abstract

Biomaterial nerve cuffs are a clinical alternative to autografts and allografts as a

means to repair segmental peripheral nerve defects. However, existing clinical

biomaterial constructs lack true incorporation of physical guidance cues into their design.

In both two- and three-dimensional systems, it is known that substrate geometry directly

affects rates of axon migration. However, the ability to incorporate these cues into

biomaterial scaffolds of sufficient porosity to promote robust nerve regeneration in three-

dimensional systems is a challenge. We have developed fibrin constructs fabricated by a

sacrificial templating approach, yielding scaffolds with multiple 10–250 μm diameter

conduits depending on the diameter of the template fibers. The resulting scaffolds

contained numerous, highly aligned conduits, had porosity of ~ 80%, and showed

mechanical properties comparable to native nerve (150–300 kPa Young’s modulus). We

studied the effects of the conduit diameters on the rate of axon migration through the

scaffold to investigate if manipulation of this geometry could be used to ultimately

promote more rapid bridging of the scaffold. All diameters studied led to axon migration,

but in contrast to effects of fiber diameters in other systems, the rate of axon migration

was independent of conduit diameter in these templated scaffolds. However, aligned

conduits did support more rapid axon migration than non-aligned, tortuous controls.

Keywords:

Fibrin, Micropatterning, Nerve guide, Nerve tissue engineering, Scaffold

39

1. Introduction

There are over 50,000 surgical procedures for peripheral nerve injuries reported

annually [1] in both civilian and military settings due to motor vehicle accidents, knife

injuries, gunshot wounds, and other sources of trauma [2,3]. Though small segmental

injuries can be repaired via surgical coaptation of nerve ends, the implantation of a graft

is required if direct suture would result in tension on the nerve [1].

The current clinical gold standard of autografting has well-known detriments

including multiple surgical sites and donor site morbidity [4,5]. Allografts have been

employed [1,6], but concerns persist regarding the potential for disease transmission [7]

and ethical concerns [8]. Biomaterial nerve cuffs based on collagen have also been

employed at the clinical level. While these nerve cuff materials avoid drawbacks of

human tissue grafting, it is not clear that their performance matches autografts and

allografts [4]. In addition to the absence of neurotrophic factors present in autografts and

allografts, nerve cuffs possess only one lumen, and therefore have far less surface area

per volume than, for example, decellularized constructs.

At the pre-clinical level, a number of filler materials, often hydrogels [9–15], have

been incorporated into nerve cuff materials. These filler materials are believed to provide

a physical matrix onto which infiltrating cells can attach. However, achieving robust

regeneration with these filler materials is generally believed to require the presentation of

soluble or immobilized chemical cues such as growth factors or cell stimulating peptide

sequences [16,17].

Besides chemical cues, there are multiple lines of evidence suggesting that

physical cues play an important role in promoting axon migration. The efficacy of

regeneration through autografts and allografts may be due to the presence of aligned

conduits in conjunction with neurotrophins [18]. Further, decellularization of nerve tissue

results in hollow, aligned conduits that support nerve regeneration [19,20]. It is also

40

known that axons will track along fibers formed by extrusion or electrospinning in a

fashion dependent on the diameter of the fibers [21,22]. Importantly, neural scaffolds

consisting only of electrospun fibers are known to affect regenerative processes in vitro

[22] and have led to axonal bridging of large neural defects [23].

From a materials perspective, one drawback to bundled fiber approaches is that

the overall porosity of nerve guidance channels packed with these fibers is expected to

be quite low. In fact, approaches to provide physical cues with fibers while providing

sufficient porosity are being actively investigated [24]. Here, we describe a different

approach to the development of biomaterial scaffolds that can be fabricated de novo,

provide physical guidance cues, and provide sufficient porosity to allow robust axonal

growth.

The goals of the present study were twofold. The first goal was to fabricate highly

porous scaffolds containing large numbers of axially-aligned luminal conduits within a

biomaterial scaffold having mechanical properties suitable for neural repair. To

accomplish this, we utilized a sacrificial templating approach within the context of a

degradable fibrin hydrogel (see Fig. 1). Numerous natural and synthetic polymer

scaffolds are compatible with this type of approach [25–29]. However, we focused on the

use of fibrin in these studies in part because fibrin is a component of native peripheral

nerve repair, being laid down in a cable ahead of the growing axon bundle as a template

for growth [30]. Moreover, the sacrificial templating approach has the advantage of

generating aligned cylindrical conduits similar in morphology to denervated endoneurial

tubes left after Wallerian degeneration is complete [31], which we reasoned would

provide a conducive substrate to axon migration. As a final design consideration, we

aimed to incorporate scaffolds with mechanical integrity similar to that of native nerve so

they would handle well and could ultimately integrate properly with target tissue.

41

Fig. 1. Scaffold fabrication process. (A) Sacrificial poly(methyl methacrylate) (pMMA) template fibers, shown in gray, were bundled within a Teflon® mold. (B) A solution of fibrinogen in PBS was back-filled around the aligned fiber bundle via centrifugation and polymerized to fibrin, shown in brown, using thrombin. (C) pMMA was dissolved using acetone, leaving an array of aligned empty conduits, shown in white, within the fibrin matrix. (For interpretation of the references to color in this figure legend, the reader is referred to the web version of this article.)

The second goal was to assess the rate of axonal migration through the conduits

of these scaffolds in order to determine the effect of the conduit diameter on the rate of

axonal extension. We employed an embryonic chicken dorsal root ganglia (DRG) model

to assess rates of axon migration [32]. Speed of axon migration is a critical metric for

nerve regeneration as more positive functional outcomes are expected with more rapid

neural regeneration.

Employing sacrificial template fibers of well-known diameter allowed for the

fabrication of scaffolds with conduits that appeared to span the full length of the scaffold

in a wide range of diameters. This allowed us to assess the effects of the conduit

diameters on the rate of axon migration within the context of a three-dimensional

construct of mechanical properties comparable to native nerve. This report describes the

42

fabrication and characterization of the scaffolds as well as the axonal response as a

function of conduit diameter in these scaffolds.

2. Methods

2.1. Template fiber fabrication and characterization

Monofilament strands of poly(methyl-methacrylate) (pMMA) were formed by melt

extrusion at the North Carolina State University College of Textiles. 100 kDa pMMA

(Polysciences, Warrington, PA) was melt extruded on two different extruder systems to

achieve a range of fiber diameters by variation of system parameters including type of

extruder, temperature, rate of extrusion, and collection godet speed (see Table 1). One

set of extrusions was performed on an Alex James and Associates, Incorporated melt

extruder with a spinneret containing 64 holes. Each orifice had a diameter of 0.6 mm and

a length:diameter ratio of 2.3:1. The process temperatures were controlled at five

different zones in the extruder. The temperatures from the feed to the die zone were

210, 220, 240, 245, and 250 °C. Undrawn filaments were taken up at 200 or 800 m/min

(Table 1). Melt spinning was also performed on the Bradford piston system with a one

hole spinneret 0.5 mm in diameter and length:diameter ratio of 2. The piston

temperature was maintained between 240 and 250 °C (Table 1). The undrawn

monofilament was taken up at 27.5, 80 or 96.5 m/min (Table 1).

Cellulose acetate (CA) fibers (generous donation of Eastman Chemical Co., Kingsport,

TN) were also used. Brightfield imaging of both CA and pMMA fibers was performed on

a Zeiss Axiovert II microscope, and micrographs were captured with AxioVision software

(both Carl Zeiss, Inc. North America, Thornwood, NY). Images were imported into

ImageJ software (National Institutes of Health, Bethesda, MD) and fiber diameter was

43

Table 1 Template fiber fabrication parameters and resulting diameter. Extrusion Method

Speed of Alex James Motor (RPM) or Piston (mm/min)

Extrusion Temperature (°C)

Godet Speed (m/min)

Fiber Diameter (μm) Average ± SD

Cellulose Acetate

a a a 11.7 ± 2.5

Alex James 15 250 800 18.5 ± 3.7

Alex James 15 250 200 40.5 ± 3.3

Piston 2.5 250 96.5 105.4 ± 6.8

Piston 3 250 80 161.3 ± 24.0

Piston 3.6 240 27.5 247.7 ± 3.3 a Donated cellulose acetate fibers were manufactured using a proprietary technique and are therefore only characterized by diameter.

measured for at least N = 5 samples. Average and standard deviation of fiber diameters

were recorded for use in studying the relationship between template fiber size and fibrin

scaffold microarchitecture.

2.2. Scaffold fabrication

Fibrin scaffolds with highly aligned conduits were fabricated by using a polymer-

templating approach, summarized graphically in Fig. 1, in which polymeric fibers were

used as sacrificial templates. To begin, aligned pMMA fibers or cellulose acetate (CA)

fibers were bundled tightly and inserted into a Teflon® tube (McMaster-Carr, Los

Angeles, CA). Scaffolds were fabricated in Teflon® tubing with either 3 mm or 8 mm

inside diameter, depending on the nature of the experiment (see below). Care was taken

to fill as much free space within the mold as possible in order to maximize porosity of the

final scaffold. CA fibers averaged 11.7 μm in diameter, while five different diameters of

pMMA fiber were used, averaging 18.5 μm, 40.5 μm, 105.4 μm, 161.3 μm, and 247.7

μm. Each scaffold was templated with fibers of a uniform diameter, resulting in a range

of six scaffold types for use in further experiments. A 200 mg/mL solution of bovine

fibrinogen (Sigma–Aldrich, St. Louis, MO) in PBS was then back-filled around the fiber

44

bundle by centrifugation in a Sorvall Legend RT benchtop centrifuge (Kendro Laboratory

Products, Newtown, CT) at 50 rcf. The fibrinogen surrounding the pMMA templates was

then polymerized for at least 16 h in a thrombin working solution consisting of 26 parts

PBS, two parts 5.5 mg/mL CaCl2 (Thermo Fisher Scientific, Waltham, MA), and two

parts 250 U/mL thrombin (Sigma). After polymerization, excess fibrin was removed from

each end of the fiber bundle with a razor blade. The resulting scaffold material was then

placed in acetone (Fisher) to selectively dissolve the sacrificial CA or pMMA fibers.

Three acetone washes were used lasting two hours, four hours, and 16 hours,

respectively. This acetone step also proved sufficient to sterilize scaffolds for use in

tissue culture experiments.

Random porous and non-porogen control scaffolds were fabricated similarly. To

create scaffolds incorporated with a random porous network, a template was formed by

filling the Teflon® mold with 180–212 μm diameter pMMA beads (Polysciences,

Warrington, PA). This size range of beads was obtained by sieving. The bead-filled

molds were vigorously tapped to settle beads into a close-packed arrangement and

heated at 140 °C for 22 h to sinter the beads together at points of contact [25] and [33].

Fibrinogen back-filling, polymerization into fibrin, and template removal with acetone

were performed as above for the sacrificial fibers. Non-porogen scaffolds were created

by omitting the inclusion of a polymer template. Instead, fibrinogen was pipetted into the

Teflon® mold, a wedge was removed from the wall of the mold with a razor blade, and

the entire assembly was immersed in thrombin working solution as described above to

polymerize the fibrin block. The remaining Teflon® was then removed, and acetone

washes were performed as above to sterilize the scaffolds. These resulting non-porogen

scaffolds were essentially a fibrin hydrogel plug.

45

2.3. Characterization of scaffold morphology

Scaffold morphology was evaluated by using scanning electron microscopy

(SEM) of scaffolds consisting of each different sacrificial template diameter. Scaffolds

fabricated as described above were cut in half longitudinally with a scalpel, hydrated in

sterile PBS, dried in a model EMS850X critical point dryer (Electron Microscopy

Sciences, Hatfield, PA), mounted on SEM chucks for longitudinal or cross-sectional

imaging, and sputter coated with an Au-Pd mixture using a Hummer 6.2 sputter coater

(Anatech Ltd., Battle Creek, MI). Scaffolds were imaged with an S-2600 N environmental

scanning electron microscope (Hitachi High Technologies America, Inc., Schaumburg,

IL) in either longitudinal- or cross-section. Images of the longitudinally-sectioned

scaffolds were imported into ImageJ software to quantify the degree of conduit alignment

within the scaffold. Five regions within the micrograph were randomly selected and,

within each region, the angular deviation from the scaffold longitudinal axis was recorded

for all observed conduits. Similarly, the diameters of the conduits were determined by

ImageJ analysis of the cross-sectional SEM images by measuring five separate regions

of a single scaffold.

2.4. Scaffold porosity determined by mercury porosimetry

Porosity was measured for scaffolds made with each fiber template by mercury

porosimetry. 8 mm diameter scaffolds of different luminal conduit dimensions were

prepared as described above. Samples were then trimmed to 1 cm length, weighed, and

evaluated using an AutoPore IV mercury porosimeter (Micromeritics, Norcross, GA).

Mercury intrusion was performed over a range of 0.10–33000 psia, and total intrusion

volume was measured over this range. Total pore volume was calculated automatically

by the bundled software package (AutoPore IV 9500 v1.09, Micromeritics) by subtracting

intrusion volume at maximum pressure from intrusion volume at zero pressure [34].

46

Sample bulk volume was assumed to equal total penetrometer volume minus intrusion

volume at zero pressure. Sample percent porosity was then calculated from bulk volume

and pore volume by software using the equation:

P% = (Vpore/Vbulk) ∗ 100% [34].

N = 3 were evaluated for each scaffold type, and analysis of variance (ANOVA)

was performed on resulting averages using Prism 5 (GraphPad Software, La Jolla, CA)

to determine if any significant differences existed between scaffold types using a value

of P < 0.05 to determine statistical significance.

2.5. Mechanical properties determined by tensile testing

To determine stiffness and strength, scaffolds were fabricated as described

above, re-hydrated in PBS for at least six hours, and trimmed to 3 cm length. Scaffolds

of each type (n = 3) were mounted into the grips of a Model 5544 tension tester (Instron,

Canton, MA) equipped with a 100 N load cell, and scaffold diameter and length were

measured. Samples were subjected to 10 cycles of 0.3 mm maximum extension at

0.1667 mm/s as a preconditioning step prior to tensile testing to failure. Raw data were

exported to MS Excel 2007 (Microsoft, Redmond, WA) and plotted as tensile load vs.

extension and tensile stress vs. strain. Maximum load was directly located in tabular raw

data and reported. The linear region of the stress vs. strain plot was selected manually,

and linear regression was performed on this data set. The slope of the resulting line was

reported as the scaffold Young’s Modulus (E). ANOVA and post-hoc Tukey’s test (where

indicated by positive ANOVA) of calculated E and maximum load were performed using

Prism 5. N = 3 samples were evaluated for each scaffold and P < 0.05 was taken as

statistically significant.

47

2.6. DRG seeding in vitro

To determine the rate of axon infiltration through the scaffolds in response to

scaffold architecture, we employed an embryonic chicken dorsal root ganglia (DRG)

model [32]. DRG culture medium was composed of HyClone high-glucose Dulbecco’s

modified Eagle’s medium (Fisher), 10% v/v fetal bovine serum (GIBCO Invitrogen,

Carlsbad, CA), 1% v/v 5-fluoro-2′-deoxyuridine (FDU, Sigma), 1% v/v Uridine (Sigma),

1% v/v penicillin-streptomycin (Invitrogen), 20 μg/mL bovine aprotinin (Sigma), and 50

ng/mL nerve growth factor (NGF, BD Biosciences). Aprotinin was included to reduce

fibrinolytic scaffold degradation, and NGF was used to stimulate axonal extension [32].

In particular, we note that FDU and uridine were added to minimize proliferation of non-

neuronal cells, especially Schwann cells. The purpose of inhibiting Schwann cell

proliferation was to allow us to focus on axonal extension in response to the conduit

diameters alone rather than in response to Schwann cells and other cell-based trophic

cues.

3 mm diameter aligned conduit scaffolds and control scaffolds, fabricated as

described above, were trimmed to 5 mm in length and washed three times in sterile PBS

for at least two hours, four hours, and 16 hours respectively to remove acetone. Small

amounts of 100 mg/mL fibrinogen and thrombin working solution were used as a glue to

attach scaffolds to the wall of individual wells in 24-well Falcon polystyrene culture

dishes (BD Biosciences, Franklin Lakes, NJ) under aseptic conditions. Scaffolds were

kept wetted with PBS to prevent contraction and retain normal morphology of the fibrin

matrix until DRG seeding.

DRG were dissected from E7-E8 chicken embryos (Tyson Chicken, Hays, NC)

and stored on ice in sterile 1% penicillin/streptomycin in PBS for a time not exceeding

one hour. Ten DRG were seeded on the top surface of each scaffold to increase the

likelihood of obtaining a longitudinal section from the scaffold that had a DRG and

48

associated axons (see below). DRG culture medium was added to a height just below

the seeded scaffold top to allow DRG attachment to the scaffolds. This cell cluster was

allowed to attach to the scaffold for 24 h, after which additional medium was added to

cover the top of the scaffold and DRG. Samples were removed after 1, 2, or 3 days, with

n = 3 for each combination of scaffold type and time point. Removed scaffolds were

briefly washed once in PBS, oriented in a plastic tray for longitudinal sectioning, and

then snap frozen in OCT compound (Sakura Finetek USA, Torrance, CA) with liquid

nitrogen.

2.7. Immunohistochemistry

Axon infiltration into fibrin scaffolds was determined by immunohistochemically

staining longitudinal sections for neurofilament. 50 μm-thick sections were cut from OCT-

embedded scaffolds along longitudinal planes using a CM 1950 cryostat (Leica

Microsystems, Bannockburn, IL) and immobilized on silane-coated glass slides

(LabScientific, Livingston, NJ). Sections containing DRG were fixed for 3 min in 10%

neutral-buffered formalin (Leica) and then washed to remove excess fixative and OCT

compound. Sections were blocked using Dual Endogenous Enzyme Block (Dako North

America, Carpinteria, CA) and Serum-Free Protein Block (Dako). Sections were then

treated with mouse monoclonal antibody to 160 kDa and 200 kDa neurofilament

(AbCam, Cambridge, MA) or to mouse IgG (isotype) control at 1:100 dilution in Antibody

Diluent Solution (Dako) for 30 min. Serum-Free Protein Block was applied a second time

before then treating sections with biotinylated goat anti-mouse IgG or horse anti-mouse

IgG (Vector Laboratories, Burlingame, CA) at 1:200 dilution in Antibody Diluent Solution.

Sections were then treated with ABC-AP reagent (Vector) before staining with Vector

Red Alkaline Phosphatase Substrate (Vector). Coverslips were mounted to slides using

Fluoromount-G (Southern Biotech, Birmingham, Alabama) and dried overnight at 4 °C.

49

2.8. Image analysis

Staining was imaged via brightfield microscopy using a DM4000 B microscope

(Leica) with attached Retiga 2000RV camera (QImaging, Surrey, BC, Canada), coupled

with ImagePro 6.2 software (Media Cybernetics Inc., Bethesda, MD). Maximum axon

length was measured using ImageJ software. After the longest axon was identified, a

line parallel to the scaffold longitudinal axis was drawn between the axon tip and scaffold

surface and the length of the line was measured. One image from each seeded scaffold

was analyzed and average maximum axon length calculated for each combination of

time point and scaffold type, resulting in N = 3 for each reported data point. This method

of quantification of maximum length of axon extension is consistent with those reported

by others [21]. Statistical analysis of these averages was performed in Prism 5 using

ANOVA and post-hoc pairwise Bonferroni comparisons where appropriate. P < 0.05 was

used as the significance threshold.

3. Results

3.1. Template fiber characterization

To fabricate scaffolds and assess the effects of conduit diameter on the rate of

axon extension, it was necessary to acquire polymeric fiber templates of a range of

diameters. Acetone-soluble polymer fibers were either custom extruded or donated to

serve as these templates. 12 μm templates were obtained by donation of cellulose

acetate fibers, but to obtain template fibers of other diameters, processing parameters

for extrusion were modulated to obtain fibers diameters from approximately 19–250 μm

as determined by light microscopy and summarized in Table 1. We anticipated that this

50

wide micro-scale range would provide a good basis for evaluating the effects of conduit

diameter on axon growth in three dimensions.

3.2. Scaffold morphology

One goal of these studies was to fabricate three-dimensional scaffolds with well-

defined conduit dimensions to assess the effect of conduit diameters on axon infiltration

rate. We employed scanning electron microscopy (SEM) to qualitatively and

quantitatively assess the physical architecture of the scaffolds. Fig. 2 shows SEM

micrographs of scaffolds in both the longitudinal and transverse planes at magnifications

of 25–50X. All scaffolds had the same general architecture with densely-packed hollow

conduits that were axially-aligned. It can be clearly observed that with small diameter

templates, the conduit diameters are much smaller (e.g., 12 μm template fibers)

compared to larger diameter template fibers (e.g., 250 μm template fibers). Higher

magnification images are provided in Fig. 2 for smaller diameter templates to

demonstrate the presence of conduits difficult to see at lower magnification. Based on

the longitudinal sections of all scaffolds, the conduits appear to run the full length of the

scaffolds, indicating the ability of axons to migrate through the length of the material (see

below). We noted artifacts in the SEM images of 19 and 41 μm templates that we

believe may be a slight contamination of the pMMA templates with higher molecular

weight polymers. This contamination did not lead to any observable effects in other

studies and appears to be a small fraction of the total polymer template used.

To quantitate the axial alignment of the scaffolds, we measured the deviation of

these conduits from the main axis of the scaffold. Fig. 3 shows the alignment of the

conduits within the scaffolds measured by this technique. Quantification of conduit

orientation revealed that all patterned channels fell within 16° of the scaffold axis and

assumed a roughly Gaussian distribution centered near 0°. The results of this

51

Fig. 2. Scaffold morphology. SEM micrographs of scaffolds in cross-section (top images) and longitudinal section (bottom images) are shown for each template type. For smaller diameter templates (top set of panels), the middle row shows higher magnification images to demonstrate the presence of the conduits present. Micrographs depict conduits of uniform diameter traveling the length of the scaffold. Scale is shown in the bottom right of individual images.

52

Fig. 3. Conduit alignment within scaffolds. Histograms represent number of conduits as a function of angle from the scaffold midline. All histograms are centered near 0°, indicating a high degree of alignment among all scaffold conduits.

characterization indicate that the scaffolds have highly aligned and densely-packed

conduits.

As described above, we had measured the diameters of the templating fibers,

which should provide an indication of the resulting conduits. However, we also estimated

the diameters of the conduits by measurement from the SEM images. Fig. 4 shows the

results of these SEM measurements compared to the template diameters. We note that

in the larger diameter scaffolds the SEM measurements are less than the templating

diameters. However, we observed significant contraction of the scaffolds during critical

point drying for these larger scaffolds (e.g., see Fig. 2F), which is the reason for the

difference. Error bars denote standard deviations of about 5–10% of the average,

indicating that the pores were quite uniform in their diameter for a given template. We

53

also noted that the scaffolds did have “memory” in that after contraction due to

dehydration, scaffolds immersed in aqueous media did return to normal size through

swelling of the hydrogel.

3.3. Scaffold porosity

To further characterize the observation of the packing density of the conduits

from the SEM studies, we performed mercury porosimetry. Table 2 summarizes the

scaffold porosity. These results indicate that the average porosity for all templates fell

near 80% with little variability. This is important for studies below with axon migration.

Because all scaffolds had nearly the same porosity, differences in axon migration could

Fig. 4. Quantification of fiber template diameter (white bars) relative to measured resulting conduit diameter (black bars). Results show the ability to generate scaffolds over a range of conduit diameters with good homogeneity of template and conduit diameters. Note that 161 μm and 248 μm template diameters showed contraction during critical point drying and conduit diameters are therefore underestimated.

54

Table 2 Scaffold porosity.

Template Diameter (μm) Porosity (%)

11.7 79.13 ± 2.00

18.5 78.62 ± 12.16

40.5 82.36 ± 1.41

105.4 82.39 ± 1.90

161.3 80.74 ± 4.28

247.7 83.09 ± 3.81

be attributed to the conduit diameters or pattern (aligned vs. random porous) but not to

differences in scaffold porosity.

3.4. Mechanical testing

A second design parameter for these studies was to develop scaffolds that would

be suitable in mimicking the mechanical characteristics of native tissue and to ensure

that they would have sufficient mechanical integrity. To determine the mechanical

properties of the fabricated scaffolds, we performed tensile tests on scaffolds to

determine the elastic modulus and maximal load as a function of the conduit dimensions.

Fig. 5 summarizes the tensile test data from these studies. We observed that the

maximal load fell between 0.5 N and 1.2 N for all scaffolds, with significant difference

observed between only one pair. The elastic moduli of scaffolds featuring smaller

conduits were shown to be significantly higher than that of scaffolds fabricated with

larger conduits. We note that elastic moduli in the range of 100–330 kPa are comparable

in magnitude to the elastic modulus of native nerve [35].

3.5. Axon infiltration in vitro

Others have previously shown in fiber-based systems that the rate of axon

migration is dependent upon the dimensions of the fiber on which axons are tracking [21]

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Fig. 5. Scaffold mechanical properties. (A) Young’s modulus and (B) maximum load data are shown for scaffolds fabricated using five template diameters, as average ± SD with n = 3 for each data point. Pound signs (#) represent significant differences from 12 μm diameter conduit scaffolds, while asterisks (∗) indicate significant differences from 41 μm diameter conduit scaffolds; in each case, one symbol corresponds to a significant result (p < 0.05), while two symbols correspond to a highly significant result (p < 0.01). Scaffolds fabricated with smaller conduits featured significantly increased stiffness compared to those with larger conduits.

and [22]. The fabrication of scaffolds with varying conduit diameters but similar porosities

allowed us to explore the effect of conduit dimensions on axon infiltration within the

context of a three-dimensional, porous scaffold. We employed an embryonic chicken

DRG model, as this provided a system in which a full neural explant could be cultured

with the scaffolds.

We utilized antimitotics (uridine and FDU) in order to isolate effects of the

scaffold on axons alone rather than adding a confounding factor related to the issue of

Schwann cell migration. Moreover, preliminary studies revealed that scaffold

degradation by DRG occurred on a time scale similar to that of axon infiltration,

potentially complicating axon length measurements. For this reason, we included

aprotinin to the culture media to halt fibrinolytic degradation of the scaffold and allow for

reliable measurements of axon infiltration.

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To assess axonal response to the scaffolds, we used immunohistochemical

staining with anti-neurofilament antibodies. A colorimetric stain (Vector Red Alkaline

Phosphatase) was used instead of fluorometric staining methods due to the levels of

autofluorescence from the high fibrinogen (and ultimately fibrin) concentration used to

fabricate the scaffolds. A representative image from each scaffold type after one day in

culture is shown in Fig. 6. Similar results were obtained with anti-betaIII-tubulin

antibodies (images not shown), though staining was more intense with neurofilament. As

can be seen in Fig. 6, DRG attached to scaffolds after 24 h in culture and axons began

to migrate both across the scaffold edge and into the conduits of the scaffold.

We employed two control scaffolds to contrast the behavior with the axially-aligned

conduits. Fibrin plugs (non-porogen controls) were used to demonstrate the need for

porous patterning within dense fibrin hydrogels created using 200 mg/mL fibrinogen. As

shown in Fig. 6, no axon infiltration was observed into the non-porogen fibrin hydrogel

control. A fibrin scaffold with randomly oriented spherical pores (rather than axially-

aligned conduits) was also used to determine the effect of the axial alignment. As can be

seen in Fig. 6, we did not observe the same degree of axon infiltration into these

spherically-templated scaffolds. Dotted lines were added to Fig. 6 using Photoshop

Elements 6.0 (Adobe Systems Incorporated, San Jose, CA) to highlight the length of

axonal extension into the scaffolds.

To quantify these observations, we measured axonal extension into each

scaffold type. We collected images of immunohistochemically-stained scaffold sections

after 1, 2, or 3 days in culture to assess the rate of axonal migration into the scaffold.

Fig. 7B–D shows averaged maximum axon length for each template size at all three time

points evaluated, and the method used to determine these lengths is shown on a

57

Fig. 6. Axon infiltration of scaffolds. A representative image from each experimental and control scaffold type is shown after 1 day cultured in vitro. Dotted lines have been added to highlight the progression of axon infiltration. Aligned-conduit scaffolds supported faster axon ingrowth to the scaffold matrix than random porous controls. Axons could not infiltrate non-porogen controls, though some growth along the scaffold exterior was observed. Brightness and contrast of photos were adjusted and dotted lines were added using Photoshop Elements 6.0 (Adobe Systems Incorporated, San Jose, CA). An image of a scaffold (41 μm template) stained with an isotype IgG control antibody is shown to demonstrate the specificity of neurofilament staining observed in other images. Similar isotype controls were used for all scaffolds but only a representative image is shown. Scale bar shown in the bottom right of each image represents 500 μm.

58

Fig. 7. Quantification of axon infiltration. (A) Representative image of a scaffold after 1 day of culture and stained for neurofilament as in Fig. 7. The method of measuring the longest axon present in each histological image is shown. Length was measured parallel to the longitudinal axis of the scaffold, as shown by the black bracket in the example subfigure. The scale bar at the bottom right measures 500 μm. Maximum axon lengths are shown as average ± SD for culture in vitro after (B) 1 day, (C) 2 day, and (D) 3 day time points over the range of scaffold types studied, with n = 3 for each data point. Filled circles (●) represent experimental aligned conduit scaffolds, open squares (□) on each plot represent the random porous control, and open triangles (▵) represent the non-

porogen control scaffold. Note that non-porogen control scaffolds (▵) show no growth and are located near the origin. Asterisks report significant differences between experimental scaffolds and random porous controls, with one symbol corresponding to a significant result (p < 0.05) and two symbols corresponding to a highly significant result (p < 0.01). Results from all experimental and random porous scaffolds were significantly different (at least p < 0.05) from those of nonporous controls. Aligned-conduit scaffolds supported more rapid axon growth than both controls at 1 and 2 days over the entire range of conduit diameters. Similarly, nearly all conduit diameters showed significantly longer axon extension after 3 days than non-porogen and random porous control scaffolds.

59

representative scaffold from a 24-hour time point (Fig. 7A). This quantification reinforced

the observation of little growth into non-porogen control scaffolds (data point located

near origin), with axonal migration into the non-porogen scaffolds being highly

significantly different than axially-aligned scaffolds at all time points (P < 0.01). Though

axons were observed growing into random porous controls, the length of these axons

was significantly (P < 0.05) shorter than those infiltrating all aligned conduits at one- and

two-day time points.

Of particular interest was the comparison between axially-aligned scaffolds. In

contrast to previous studies indicating a role for the dimensions of aligned fiber

templates [21] and [22], in our sacrificial template system, we found no significant

differences between any of the template diameters investigated (ranging from 12 to 250

μm diameter) at any of the time points observed (1, 2, and 3 days).

4. Discussion

The success of autografts for neural regeneration likely involves the presentation

of physical guidance cues through the endoneurial channels and chemical factors [18]

(e.g., neurotrophins) in the graft secreted by Schwann cells. Several strategies for the

presentation of chemical cues to regenerating cells in biomaterial constructs have been

developed including delivery through soluble and immobilized factors [36]. The

incorporation of physical cues has been demonstrated [21], [23], [26], [27] and [37], but

relatively few studies have been directed at incorporating these cues into biomaterial

scaffolds or to the understanding of the geometrical requirements of these cues (e.g.,

effects of diameter on axonal regeneration). Several methods of incorporating physical

cues into scaffolds exist, including (1) alignment of fiber bundles [21] and [23] and

formation of aligned multi-luminal constructs via (2) sacrificial templating [26] and [29],

60

(3) phase separation [37], (4) lithography [38] and [39], and (5) solid free-form fabrication

[40].

For aligned fiber constructs it has been demonstrated that the diameter of the

fiber guides directly affect the rate of axon migration, providing an approach to increase

rates of axonal regeneration. However, these aligned fiber constructs have often been

used in two-dimensional systems (e.g. [22]) and translation of this technology to a three-

dimensional system will likely be problematic due to the low porosity of resulting three-

dimensional constructs. Sacrificial template methods provide an architecture similar to

native nerve and offer the potential for a more porous system. Existing needs for

materials formed by this approach include achieving sufficient porosity to support robust

axonal infiltration, provision of mechanical properties comparable to native nerve, and an

understanding of the effect of the conduit diameters on rates of axonal migration.

In these studies, we developed a fibrin-based construct in an effort to begin to

address these questions. Fibrin was selected because it is one of the earliest

components of the native nerve repair process [41] and [42], and several groups have

used this material as a filler for nerve cuff technology to enhance regeneration [43], [44],

[45] and [46] and/or deliver neurotrophic factors [47] and [48] or cells [49]. In addition,

the use of a sphere-templating approach to fabricate scaffolds from this material has

previously been reported [25].

We fabricated different scaffolds with a range of conduit diameters in an effort to

identify the relationship between conduit diameter and rate of axon regeneration in this

three-dimensional system. This was done because although the diameter (caliber) of the

endoneurium is known for mature native nerve, it is not clear that these diameters are

optimal for a regenerative process following injury.

The scaffolds fabricated by the sacrificial fiber approach in the current study had

a physical resemblance similar to native nerve tissue following decellularization or post-

61

injury Wallerian degeneration (compare Fig. 2 with [31]). This approach therefore offers

the ability to achieve important structure–function relationships of native tissue while

optimizing the system for regeneration. Although contraction during processing for SEM

was observed for larger diameter fiber templates (see Fig. 2), it is clear that conduit

diameter was directly related to the fiber template diameter. Further, the conduits were

highly aligned and SEM images suggest that the conduits spanned the length of the

scaffolds. We also note that there was no observed impediment to axon infiltration and

we were able to visually observe, for the large diameters, that conduits ran the full length

of the scaffold.

As a control for the role of alignment, we also incorporated porosity into the

scaffolds by the use of spherical templates [25] that created a more tortuous path than

the aligned fiber templates. The more tortuous spherical templates resulted in axon

infiltration into the scaffolds, but with a rate of extension significantly less than the less

tortuous aligned scaffold conduits (300 μm per day compared to 600–900 μm per day).

These results demonstrate the importance of non-tortuous paths for axonal extension in

three dimensional scaffolding systems and are consistent with previous results indicating

that oriented patterns are superior in directing rapid axon extension in two-dimensional

systems [50].

Importantly, the scaffolds were also highly porous, with porosity values of

approximately 80% regardless of the template fiber diameter. In addition, the mechanical

properties of these scaffolds (Fig. 5) were similar to native nerve, with an elastic

modulus of approximately 100–330 kPa compared to native nerve with an elastic

modulus of 580 kPa [35]. This combination of a highly porous scaffold material with

mechanical properties similar to native tissue is of considerable importance for several

reasons. Although the fibrin hydrogel alone has good mechanical properties, the porosity

provided by the conduits is necessary to allow the migration of axons through the

62

material. In addition, the material has sufficient mechanical strength to prevent collapse

of the conduits during handling, indicating the ability to handle these scaffolds within a

surgical setting. Lastly, the similarity of the mechanical properties of these scaffolds

relative to native nerve would likely minimize scarring that could occur due to mechanical

mismatch at site of implantation. Thus, the fibrin scaffolds fabricated by this approach

have material properties suitable and favorable for nerve grafting applications.

We evaluated the rate of axon migration into the scaffolds as a function of

conduit diameter by immunohistochemistry with histomorphometry. For these in vitro

studies we employed an embryonic chicken DRG model. This model was well-suited to

these studies as the size of the DRG allowed them to be placed on the top of the

scaffold (similar to the proximal end of a nerve injury) for all conduit diameters tested.

Dissociation of DRG to isolate neuronal cells was not feasible due to the small size of

the neurons and the large diameter of some conduits. We therefore elected to use whole

DRG explants for these studies. To focus on the axon extension through the conduits in

response to conduit diameters, we included uridine and FDU in the culture medium to

inhibit mitotic growth of Schwann cells. We observed during pilot studies that the smaller

diameter conduits (e.g., 12 and 19 μm) showed lower numbers of Schwann cells

because these cells were excluded by the diameter of the conduits. To achieve similar

trophic conditions for all scaffold types, we therefore inhibited Schwann cell mitosis with

uridine and FDU. While Schwann cells play a key role in the functional nerve recovery in

vivo, the hydrogel nature of these scaffolds offers the potential to provide exogenous

trophic support through controlled release until Schwann cells can ultimately associate

with the regenerated axons. We reason that if more rapid axon extension can be

achieved in vivo this will provide more favorable long-term functional recovery. Aprotinin

was also added to the culture to prevent degradation of the scaffold by matrix

metalloproteinases (MMPs) [49]. We also added exogenous NGF to the media to

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stimulate axonal extension from the DRG. We recognize that future strategies utilizing

this approach may require means to prevent proteolytic degradation of fibrin and to

provide controlled release of growth factors to promote axonal extension. We have found

that the hydrogel nature of these scaffolds is compatible with the slow release of small

molecule proteins such as aprotinin and growth factors (data not shown).

We evaluated the maximum distance of axon migration through the scaffolds as

reported by others [21]. Other approaches to quantify number of axons were not used

because of variability observed in DRG spreading on the scaffold surfaces (e.g., see Fig.

2).

A key finding from these studies was that no significant differences in axon length

with time were observed over the wide range of conduit diameters studied. We had

hypothesized that smaller diameter conduits would lead to increased rates of axonal

extension based on previous studies that indicated changes in length of axon extension

at a given time point (indicating differences in the rate of extension [21] and [22]). Our

results may indicate that physical shape of the curvature observed by axonal growth

cones (i.e., convex for a fiber compared to concave for a conduit) affects axonal

migration. The results could also indicate that the porosity of the material, such as the

conduit scaffolds used in these studies, is a more important parameter than the

geometrical dimensions of the physical guidance cues. We speculate that one

advantage to the smaller diameter conduits is that they may prevent infiltration of

inflammatory cells (e.g., neutrophils and monocytes/macrophages), allowing unimpeded

infiltration of axons.

We note that other studies looking at effects of geometrical parameters on axon

migration have utilized different time points (5 or 7 days) rather than the 3 time points in

our studies [21] and [22]. It is conceivable that differences could occur at longer time

points in our system. However, because we observed multiple time points (1, 2, and 3

64

days), we can see the linear rate of axon migration, indicating that similar results would

be expected at 5 or 7 days. Unfortunately a direct comparison to these other studies was

not possible. Although the fibrin scaffolds can be fabricated in any length, the

mechanical properties were not sufficient to allow their use in the culture system used in

our study; that is, the scaffolds did not have sufficient rigidity to remain upright when

glued to the culture plate. Wen and Tresco were able to evaluate at longer time points,

presumably because the more rigid nature of their scaffolds allowed them to employ

longer scaffolds in a similar culture system [21]. It is also noteworthy that in the three-

dimensional fibrin system in our study the axon extension length was approximately

1500 μm at 3 days (∼500 μm/day), compared to Wang et al, who saw extension of 1400

μm over 5 days (∼280 μm/day) in the same chick DRG system. However, differences in

culture conditions and DRG model make direct comparisons to other studies challenging

[21] and [22].

5. Conclusions

In these studies, we have described an approach to the fabrication of fibrin

scaffolds suitable for peripheral nerve repair applications. Although the sacrificial

templating approach should be compatible with other materials, the mechanical

properties achieved with fibrin scaffolds are comparable to native nerve and are

expected to support axonal migration due to the role of fibrin in the natural repair

process following nerve injury. An important finding from these studies was that the use

of highly aligned conduits supports more rapid axon migration through the scaffolds than

tortuous, non-aligned pores. However, in contrast to two-dimensional fiber-based

systems, the diameters in these three-dimensional scaffolds were not found to

significantly affect the rate of axon migration through the materials. The approach used

for the fabrication of these scaffolds represent a useful means by which to incorporate

65

physical guidance cues and mechanical integrity into biomaterials useful for peripheral

nerve repair.

Acknowledgments

This work was funded by the Wake Forest University Health Sciences Venture

Fund, the Department of Biomedical Engineering, and the Wake Forest Institute for

Regenerative Medicine. The authors gratefully acknowledge the laboratory of Dr.

Carolanne Milligan, Wake Forest University Health Sciences Department of

Neurobiology and Anatomy, for assistance with isolation and culture of embryonic

chicken DRG.

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CHAPTER III

IN VITRO AGRIN-INDUCED ACETYLCHOLINE RECEPTOR CLUSTERING ON

SKELETAL MUSCLE CELLS SEEDED ON A TUNABLE FIBRIN-BASED SCAFFOLD

John B. Scott, Benjamin T. Corona, Catherine L. Ward, Micael R. Deschenes, Benjamin

S. Harrison, Justin M. Saul, and George J. Christ

The following chapter was written in a style acceptable for submission to a peer-

reviewed journal. As such, citations, figure captions, and other stylistic elements of this

chapter may differ from those found elsewhere in this dissertation.

As primary author, John Scott performed all experiments and prepared the manuscript

for Chapter III.

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Abstract

Volumetric muscle loss (VML) injuries are particularly debilitating, and commonly

result from trauma, infection, congenital defect, or surgical side effect. Unfortunately,

current medical treatments for these injuries are lacking. The "gold standard" for

treatment of critical-size defects remains autografting, which is restricted by donor site

availability and morbidity, and further is often associated with incomplete repair of

function or restoration of cosmesis. In the current work, we describe an in vitro tissue

engineering approach incorporating biomimetic materials, cell supplementation,

pharmacologic signaling, and mechanical strain that recapitulates some phenotypic

aspects of motor end plates (MEPs) observed on innervated skeletal muscle in vivo.

C2C12 cells cultured on a 2D fibrin biomaterial and treated with neural (Z+) agrin

exhibited clusters of acetylcholine receptors (AChRs). When agrin was presented bound

to the surface of a carrier microparticle, this response was spatially restricted to areas of

cell – microparticle contact. Moreover, AChR clusters were observed from 16 – 72 hours

after treatment when Z+ agrin was adsorbed to the surface of microparticles, but this

time frame was extended to 120 hours and beyond when agrin was covalently linked to

the microparticle surface. Similar AChR clustering was observed both when covalent

agrin microparticles were allowed to settle from a culture medium suspension above a

cell-seeded fibrin surface and when C2C12 cells were seeded on an agrin microparticle-

containing fibrin surface. The latter microparticle delivery method enabled construction of

3D agrin-presenting fibrin scaffolds, which were seeded with expanded primary rodent

muscle derived cells (MDCs). Incorporation of agrin-presenting microparticles in 3D fibrin

constructs with surface-seeded rat MDCs resulted in little or no expression of AChR

clusters. However, cyclic stretch of an agrin-presenting 3D fibrin scaffold seeded with

MDCs resulted in dramatic enhancement of AChR clustering relative to samples lacking

72

either agrin or stretch, providing evidence that mechanical strain and agrin

supplementation may act synergistically in stimulating MEP-like structures in vitro. Taken

together, these results describe a multidisciplinary approach leading to an in vitro

phenotype that mimics some key structural features observed in innervated muscle in

vivo. This represents an important first step toward preservation of native muscle

phenotype following tissue engineered construct implantation in vivo.

Keywords:

Agrin, Bioreactor, Cyclic stretch, Drug delivery, Fibrin, Microparticles, Skeletal muscle

tissue engineering, Scaffold, Volumetric muscle loss

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1. Introduction

The ability to voluntarily control skeletal muscle is critically important in daily life,

as it is responsible not only for mobility and manipulation but also for life-sustaining

functions such as breathing and swallowing. Skeletal muscle possesses an impressive

capacity for self-repair with a well-defined regenerative response, beginning with

inflammation of the wounded area and progressing, in turn, to satellite cell mobilization

and, ultimately, maturation of myofibers and remodeling of muscle architecture [1–4]. In

particularly extensive injuries to skeletal muscle, however, the native repair process is

insufficient to functionally regenerate the wound, which is instead covered over by scar

tissue [2]. As the connective tissue composing a scar is not contractile, and can often

serve as a barrier to reinnervation [4], a critical size of injury exists from which muscle

function can never be fully restored by native wound healing.

An injury of this type, in which muscle injury results in permanent loss of function,

is termed volumetric muscle loss (VML) [5]. Though VML can be caused by congenital

abnormalities, acquired disease, or tumor resection, the single greatest cause of

damage to skeletal muscle is trauma [5], largely because of its proportion in the human

body and location in extremities. Traumatic injuries are incredibly common among

military personnel [6,7], and while advances in personal protective equipment are saving

the lives of soldiers that would otherwise suffer fatal wounds, these individuals are often

left with significant injuries [8] including VML [5]. Moreover, similar phenomena are seen

in civilian settings, such as severe injuries among those in automobile collisions whose

lives were saved by safety mechanisms or improved emergent trauma care [8]. Thus,

VML is common in the current clinical setting.

Though surgical treatment of VML is possible in some cases by autologous

tissue transfer [9–11], the limitations of donor site size and morbidity remain and

treatment is largely incapable of restoring normal function or appearance. Alternative

74

strategies to directly repair VML or enhance the native healing process exist in various

stages of investigation or use, including cell transplantation [12,13], platelet-rich plasma

[14], growth factors [15], hyperbaric oxygen inhalation [16], gene therapy [17], and

traditional pharmacologics [18,19], but all feature characteristic limitations and none are

yet capable of restoring full function to critically injured skeletal muscle tissue. This,

combined with the frequent presentation of these injuries, represents a significant unmet

clinical need.

Several muscle tissue engineering strategies are under investigation in an

attempt to remedy the shortfall in standard of care for VML injuries. The use of various

biomaterial scaffolds has been reported in multiple models of skeletal muscle repair [20–

28], with varying results. For this and similar reasons, materials are often coupled with

relevant cells in an attempt to engineer skeletal muscle tissue in vitro or in vivo [22,29–

40]. Though these results overall have shown promise, there is still room for

therapaeutic improvement. Thus, many recent efforts have incorporated additional cues

into combination cell-material scaffolds, including mechanical stretch prior to

implantation [41–46] and chemical motifs [47–54], in an attempt to further enhance

tissue engineering strategies by better recapitulating the complex native skeletal muscle

microenvironment.

One important aspect of this microenvironment is signaling provided by axons

which innervate muscle, forming neuromuscular junctions [55–58]. Any tissue

engineered skeletal muscle implant must integrate with the host nerve supply to fully

restore voluntary motion. Muscle cells undergo significant atrophy if chronically

denervated [2]; moreover, clinical expectations for axon growth during reinnervation of

injured tissue are roughly 1 mm per day [59]. In this scenario, incorporation of

technologies that would maintain skeletal muscle phenotype and function during the host

75

re-innervation of treated VML injuries would be a valuable adjunct to prevent atrophy

during the interim period associated with lack of neuronal signals.

One promising candidate for pharmacological supplementation of tissue

engineered muscle repair strategies is neural agrin, a heparan sulfate proteoglycan

noteworthy for its ability to induce clustering of plasma membrane-bound actylcholine

receptors (AChRs) in skeletal muscle cells [55–58,60–62]. This clustering behavior is

unique to agrin isoforms featuring a splice insert at the Z-site that is specific to nerve-

derived (Z+) agrin [61], and is a critical first step in the assembly of a mature

postsynaptic apparatus, or motor end plate (MEP), on the muscle cell in normal

physiology [55–58]. Studies to date exploring neural agrin functionalization for skeletal

muscle tissue engineering applications [50,53] have shown promising results. However,

these studies have exclusively employed soluble agrin conditioning in vitro, which would

not be capable of maintaining the agrin signal within the tissue engineered construct if

implanted in vivo. By contrast, neural agrin is immobilized in the synaptic basal lamina in

normal physiology and is confined very tightly to areas of nerve-muscle contact [58].

In the current work, we describe efforts to develop a technology that generates a

cellular phenotype in vitro including the presence of AChR clusters via incorporation of

agrin into a fibrin biomaterial. Evidence of these AChR clusters in vitro would be an

important step in the development of tissue engineered skeletal muscle, as it would be

more reminiscent of the MEPs seen in innervated skeletal muscle in vivo. Such a

phenotype would be of significant utility in a tissue engineering approach, as it could

counteract the effects of muscle denervation during construct creation as well as after

construct implantation. In contrast to previous strategies, this approach utilizes

biomaterial microparticles as delivery vehicles exclusively for non-soluble, locally

immobilized neural agrin, which may produce more desirable results by bio-mimicry.

Further, we discuss potential synergy between mechanical stretch and agrin

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presentation which may in the future be combined with a previously successful tissue

engineered skeletal muscle repair technology [44–46] to create a next-generation

strategy with expanded clinical applications for the treatment of devastating VML

injuries.

2. Methods

2.1. Culture medium formulation and preparation of agrin-presenting and control

materials

Four different culture medium formulations were used in subsequent

experiments, with all components expressed as % v/v and sourced from Thermo Fisher

Scientific (Waltham, MA) unless otherwise specified. C2C12 growth medium consisted

of 89% high-glucose Dulbecco’s modified Eagle’s medium (DMEM), 10% heat-

inactivated fetal bovine serum (FBS), and 1% penicillin/streptomycin. Differentiation

medium consisted of 97% 1:1 DMEM:F12 mixture, 2% horse serum (HS), and 1%

antibiotic-antimycotic. Myogenic medium consisted of 68% high-glucose DMEM, 20%

FBS, 10% horse serum, 1% chicken embryo extract (Sera Laboratories International Ltd,

West Sussex, UK), and 1% antibiotic-antimycotic. Rat muscle derived cell (rMDC)

growth medium consisted of 84% low-glucose DMEM, 15% FBS, and 1%

antibiotic/antimycotic.

Experimental cell cultures in vitro described below were supplemented with agrin

either dissolved within culture medium or bound to the surface of a microcarrier.

Solution-phase agrin culture medium was prepared by diluting a carrier-free recombinant

rat agrin (R&D Systems, Minneapolis, MN) stock at 100 μg/mL (~1.11 μM) in sterile PBS

by using an appropriate culture medium (as specified per-experiment below) to a final

agrin concentration of 50 ng/mL (~0.56 nM).

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Agrin-adsorbed microcarriers were prepared from either 10 μm diameter

Polybead® microspheres or 10 μm diameter Fluoresbrite® yellow green microspheres

(both Polysciences, Inc., Warrington, PA). Microspheres were first centrifuged from stock

into a pellet and then sterilized by resuspension to a concentration of ~34 million

particles/mL in 95% v/v ethanol in water and agitation on a lab roller for at least 15

minutes at ~22°C, with subsequent steps carried out in a biological safety cabinet to

maintain sterility. Microspheres were centrifuged and washed in sterile water twice to

remove residual ethanol before being resuspended from a pellet to a concentration of

~84 million particles/mL in 100 μg/mL agrin stock and agitated on a lab roller for at least

1 hour at ~22°C to allow for agrin adsorption. Finally, microcarriers were centrifuged and

washed in sterile water three times to remove non-adsorbed agrin and then

resuspended to a functional microcarrier stock concentration of ~48 million particles/mL

in sterile water, in which form they were stored statically at 4°C for up to 2 months before

use.

Covalently-bound agrin microcarriers were prepared by using the zero-length

crosslinker N-(3-dimethylaminopropyl)-N′-ethylcarbodiimide hydrochloride (EDC, Sigma-

Aldrich, St. Louis, MO) to link amine groups on agrin molecules to either 10 μm diameter

Fluoresbrite® yellow green carboxylate microspheres (Polysciences) for 2D fibrin

systems, or 10 μm diameter SiO2 microspheres (Corpuscular, Cold Spring, NY) for 3D

fibrin systems. Microspheres were first centrifuged from stock into a pellet and then

washed twice in MES buffer, pH 5.7, consisting of 0.1 M MES (Acros Organics, Geel,

Belgium) and 0.5 M NaCl (Sigma). Microspheres were then centrifuged and

resuspended to ~42 million particles/mL in an intermediary crosslinking solution

consisting of MES buffer with 10 mg/mL (52 mM) EDC and 10 mg/mL (46 mM) N-

hydroxysulfosuccinimide sodium salt (sulfo-NHS, Sigma). This intermediary suspension

was agitated on a lab roller for 15 minutes at ~22°C before being quickly centrifuged and

78

washed twice in phosphate buffered saline (PBS, Thermo Fisher Scientific), pH 7.4.

These functionalized microspheres were centrifuged to a pellet and crosslinking to agrin

was accomplished by resuspending to a concentration of ~42 million particles/mL in a 50

μg/mL (~560 nM) solution of agrin in PBS and agitating on a lab roller for at least 6 hours

at ~22°C. Agrin-bound microcarriers were subsequently centrifuged, resuspended at a

concentration of ~34 million particles/mL in a solution of 95% v/v ethanol in water, and

agitated on a lab roller for at least 30 minutes at ~22°C to both remove any agrin simply

adsorbed to the microsphere surface as well as to sterilize the microcarriers. Finally,

microcarriers were centrifuged and washed twice with sterile water in a biological safety

cabinet to remove residual ethanol and then resuspended to a functional microcarrier

stock concentration of ~48 million particles/mL in sterile water, in which form they were

stored statically at 4°C for up to 3 months before use.

Control materials were prepared similarly. As a negative control to solution-phase

agrin in culture medium, culture medium of the appropriate type was used without added

agrin. As a negative control to agrin-adsorbed microcarriers, 10 μm diameter Polybead®

microspheres were sterilized in ethanol and washed in sterile water as above, but were

not subjected to further manipulation. As a negative control to covalently-bound agrin

microcarriers in 2D model systems, Polybead® carboxylate microspheres or

Fluoresbrite® yellow green carboxylate microspheres, as appropriate, were sterilized in

ethanol and washed in sterile water as above, but were not subjected to further

manipulation. As a negative control to covalently-bound agrin microcarriers in 3D model

systems, covalently-bound bovine serum albumin (BSA, Sigma) microcarriers were

fabricated. Polybead® carboxylate microspheres were treated as above, with the sole

change that the agrin solution was replaced with BSA in a 36.7 μg/mL (~560 nM)

solution in 50% v/v PBS/deionized water.

79

2.2. 2D Fibrin-seeded C2C12 cell response in vitro to agrin-adsorbed microcarriers

Two-dimensional fibrin biomaterial gels were created by immobilizing a thin sheet

of fibrin on a glass coverslip. 12 mm diameter round glass coverslips (Leica

Microsystems, Buffalo Grove, IL) were mechanically roughened with sandpaper to

provide an adherent surface before being rinsed with DI water and dried. A fibrin gel was

then applied to one side of each coverslip by mixing 7 μL fibrinogen (Sigma) at 10

mg/mL (~29 μM) in PBS with 10μL of a thrombin working solution containing 16.7 U/mL

thrombin and 367 μg/mL (~3.3 mM) CaCl2 (both Sigma) in PBS and spreading the

mixture evenly across the glass surface. After gel polymerization, coverslips were placed

into individual wells of a 24-well dish and then sterilized by exposure to ≥1 MRad γ

radiation.

AChR clustering in C2C12 cells in response to solution-phase agrin and agrin-

adsorbed microcarriers was evaluated by adding materials to a layer of fibrin-adherent

C2C12 cells. To initially seed fibrin gels, C2C12 mouse myoblasts (ATCC, Manassas,

Virginia) were suspended in C2C12 growth medium to a concentration of 50,000

cells/mL and then 1 mL of this suspension was added to each well containing a fibrin-

coated coverslip. Cells were allowed to settle by gravity, attach to fibrin, and multiply

over 16-24 hours. Growth medium was then removed and replaced with differentiation

medium supplemented with aprotinin (Sigma) at 20 μg/ml (~3.1 μM) to inhibit fibrinolysis.

This supplemented differentiation medium was exchanged every 2-3 days. After 6 days

of culture in differentiation medium, either (1) aprotinin-supplemented differentiation

medium alone, (2) a solution of agrin in aprotinin-supplemented differentiation medium,

(3) a suspension of control microparticles at ~1.19e6 particles / mL in aprotinin-

supplemented differentiation medium, or (4) a suspension of agrin-adsorbed

microcarriers at ~1.2 million particles/mL in aprotinin-supplemented differentiation

medium was added to fibrin-adherent C2C12 cells to evaluate cellular response. Cells

80

were then left for a final 16-24 hours to allow any AChR clustering to occur before being

evaluated by histology and epifluorescence microscopy as described in Sections 2.6 and

2.7 below.

2.3. 2D Fibrin-seeded C2C12 cell response in vitro to covalently bound agrin

microcarriers

Preliminary evaluation of AChR clustering in C2C12 cells in response to

covalently-bound agrin microcarriers was evaluated similarly to the previous experiment.

Aprotinin supplementation, medium changes, and histology were performed as in

Section 2.2 above, but experimental particles presenting covalently immobilized agrin or

control microparticles were added to C2C12 cell-seeded fibrin gels after only 2 days of

culture in differentiation medium, allowing for 5 days of cell-microcarrier contact.

Staining, evaluation of AChR localization using confocal microscopy, and image

processing were performed as described in Sections 2.6 and 2.7 below.

AChR clustering in C2C12 cells cultured on a fibrin-presenting surface was then

evaluated. Fibrin gels were polymerized on glass coverslips as above, with the following

two exceptions. First, fibrinogen was filter-sterilized by passing through a 0.2 μm-pore

filter before being used in gel polymerization and all other components were combined in

sterile form, eliminating the need for γ-irradiation of gels. Second, 10 μl of fibrinogen

solution containing a suspension of ~124 million mL-1 covalently-bound agrin

Fluoresbrite® particles or of untreated control Fluoresbrite® particles was polymerized to

fibrin using 2 μL of a solution consisting of 125 U/mL thrombin and 2.75 μg/mL (~25 mM)

CaCl2 in PBS. These agrin-presenting fibrin materials and controls were seeded with

C2C12 cells, cultured, stained, and imaged as above. Because microcarriers were

immobilized within the gel prior to cell seeding, no microparticles of any kind were added

in suspension.

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2.4. Isolation of rat muscle derived cells

rMDCs used in 3D cell culture were obtained from 4-6 week old male Lewis rats

(Charles River, Raleigh, NC) and were isolated as previously described [45,46]. Briefly,

tissue culture plates were prepared by incubating 15 cm diameter tissue culture

polystyrene (Fisher) with Matrigel (Corning, Corning, NY) diluted 1:50 in PBS at 37°C for

≥4 hours.

Meanwhile, tibialis anterior and soleus muscles were removed from euthanized

donor rats, rinsed in 10% povidone iodine solution (McKesson Corporation, San

Francisco, CA), and transferred to a sterile environment where they were subsequently

washed in PBS, manually minced to a suspension of fine particles in low-glucose DMEM

supplemented with collagenase (Worthington Biochem, Lakewood, NJ) at 770 U/mL,

and incubated under static conditions at 37°C for 2 hours. Digested tissue suspensions

were centrifuged, the supernatant was removed, and tissue was resuspended in

myogenic medium at a proportion of all tissue from one animal in 100 mL medium. Dilute

Matrigel was removed from treated plates and resuspended tissue was then added at 20

mL/plate.

After 2 days, myogenic medium was removed and replaced with rMDC growth

medium. After a further 2 days, cells were dissociated from plates by incubating in a

solution of EDTA and 0.05% trypsin-EDTA (both Invitrogen Life Technologies, Grand

Island, NY) for 5-10 min at 37°C and mechanically agitating plates. Cells were then

replated on untreated 15 cm diameter tissue culture polystyrene in rMDC growth

medium at 1.5-2.5 million cells/plate. At ~70% confluence, usually after 3-4 days, cells

were again dissociated from plates and used for experimentation as described in Section

2.5 below.

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This isolation protocol was approved by the Wake Forest University Institutional

Animal Care and Use Committee and complies with animal use guidelines established

by the American Physiological Society.

2.5. 3D in vitro model system for evaluation of rat muscle derived cell response to

combined agrin presentation, fibrin materials, and mechanical strain

AChR clustering in rMDCs in response to agrin presentation and mechanical

strain was evaluated in the context of a 3D fibrin scaffold. Scaffolds were fabricated

using methods similar to those previously described [63]. Cellulose acetate (CA) fibers

(generous donation of Eastman Chemical Company, Kingsport, TN) ~12 μm in diameter

were arranged into an aligned mat roughly 35 mm long x 1 mm thick, in a width

appropriate to the number of scaffolds desired. Experimental covalently-bound SiO2

agrin microcarriers or control covalently-bound BSA microcarriers were suspended in

200 mg/mL fibrinogen at a density of ~24 million particles/mL, and this suspension was

added to the fibers until the mat was completely impregnated.

Fibrinogen was polymerized to fibrin by adding a solution of 125 U/mL thrombin

and 2.75 μg/mL (~25 mM) CaCl2 in PBS and leaving undisturbed for at least 4 hours at

~22°C. This large scaffold was then trimmed into individual smaller scaffolds measuring

roughly 30 mm long x 3 mm wide x 1 mm thick using a razor blade. These were then

sequentially washed, first in acetone and then in sterile PBS, as described previously

[63] to yield sterilized 3D scaffolds of known mechanical properties [63].

One 30mm long x 3 mm wide plane of each scaffold was then seeded with

rMDCs by manually pipetting a ~30 μL suspension per scaffold of ~30 million cells/mL in

rMDC growth medium over a closely packed side-by-side arrangement of scaffolds. After

allowing an initial 30 minute period for cell attachment, ~700 μL rMDC growth medium

per scaffold was added to the arrangement to totally immerse cell-seeded scaffolds.

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One group of cell-seeded scaffolds (consisting of two subgroups featuring

experimental agrin and control BSA microcarrier types, respectively) were left

mechanically undisturbed for 6 days, while a second group (also consisting of agrin and

BSA subgroups) was removed from static culture after allowing 16-24 hours for full cell

attachment and immediately clamped into the mounts of a custom-fabricated bioreactor.

This bioreactor was similar to those previously described [44–46], consisting of a

medium reservoir, a computer-controlled electromagnetic actuator, and a series of

opposing clamps to suspend scaffolds within the medium reservoir while anchoring one

scaffold end to a fixed point and the other to the linear actuator. This design allowed

repeated application of uniaxial cyclic stretch with high spatiotemporal control. The

bioreactor was used to “precondition” cell-seeded scaffolds by subjecting them to 10%

mechanical strain parallel to the long (i.e. 30 mm) axis of scaffolds 3 times per minute for

the first 5 minutes of every hour over a total time of 5 days. This experiment is depicted

schematically in Fig. 1.

In all groups, rMDC growth medium was exchanged every 2-3 days, and was

supplemented with aprotinin starting on day 3. After 6 days, AChR localization in

scaffold-resident muscle cells was evaluated by using histology and imaging methods

described in Section 2.6 and 2.7 below.

2.6. Histological preparation of samples

2D cultures on fibrin gels were evaluated histologically while still attached to

coverslips. Gel-adherent cells were fixed in 10% neutral-buffered formalin (Leica) for 3-5

min, washed three times for 5 minutes each in tris-buffered saline with Tween (TBST,

Dako, Glostrup, Denmark), blocked against non-specific binding using a blocking buffer

consisting of 5% v/v HS in TBST for at least 30 minutes at ~22°C, stained for AChR

localization using α-bungarotoxin conjugated with Alexa Fluor 594 (α-BTX, Invitrogen) at

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Fig. 1. Schematic of 3D tissue engineered muscle construct fabrication, seeding, and preconditioning. (A) Pharmacological delivery vehicles were created by covalent immobilization of agrin on a microparticle surface. These agrin microcarriers were then suspended in a solution of fibrinogen, which was (B) polymerized to fibrin around a sacrificial template of polymer fibers, as previously described [63]. Selective dissolution of the template resulted in a patterned hydrogel material of known mechanical properties [63] that presented agrin via embedded microcarriers. (C) Scaffolds were then seeded by manually pipetting a suspension of muscle-derived cells over the scaffold surface. Resident cells then received a combination of pharmacological cues from scaffold agrin and/or (D) mechanical cues from cyclic stretch in a bioreactor.

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100 nM dilution in blocking buffer for at least 1 hour at ~22°C in the dark, washed a

further three times in TBST, mounted to glass slides using Vectashield with DAPI

(Vector Laboratories, Burlingame, CA) and dried overnight at 4°C.

3D cell-seeded scaffolds were removed whole from bioreactors, fixed by immersion in

4% paraformaldehyde in PBS for 16-24 hours under static conditions at 4°C, and

paraffin processed in an ASP300 tissue processor (Leica) by using the following steps:

1) 60% isopropyl alcohol in water (IPA), 45 minutes;2) 70% IPA, 45 minutes; 3) 80%

IPA, 45 minutes; 4) 95% IPA, 45 minutes; 5) 95% IPA, 1 hour; 6) 100% IPA, 45 minutes;

7) 100% IPA, 45 minutes; 8) 100% xylenes, 45 minutes; 9) 100% xylenes, 45 minutes;

10) paraffin, 45 minutes; 11) paraffin, 1 hour; 12) paraffin, 1 hour.

Processed samples were then mounted into paraffin blocks and longitudinally

sectioned on a microtome to 15 μm thickness, with the plane of the section

perpendicular to the 30 mm x 3 mm plane of the scaffold. Tissue sections were

immobilized on glass slides, dried for 16-48 hours at 60°C, and then left indefinitely at

~22°C until stained. Prior to staining, slides were deparaffinized by sequential washes in

xylenes, ethanol, and tap water. Staining was then performed as for 2D samples,

beginning with the initial three washes of TBST.

2.7. Imaging of samples and subsequent image processing

Epifluorescence and confocal microscopy were used to evaluate AChR staining

in 2D samples. Epifluorescence images were obtained by using a DM4000 B

microscope (Leica) with attached Retiga 2000RV camera (QImaging, Surrey, BC,

Canada), coupled with ImagePro 6.2 software (Media Cybernetics Inc., Bethesda, MD).

Confocal micrographs were captured on an FV10i confocal microscope (Olympus

America, Center Valley, PA). Three-dimensional projections of sequential confocal z-

stacks were obtained using FIJI image processing software [64]. Surface renderings

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(see Fig. 3B, 3D, and 3F) were obtained by importing 3D projections of individual color

channels obtained in FIJI to ImageJ 3D Viewer [65] and manually adjusting each color

threshold until structures appeared visually similar to the native 3D projection.

AChR localization in 3D samples was imaged only with confocal microscopy.

Methods were identical to those above, except that areas for imaging were selected by

viewing DAPI and autofluorescence signals only before images of the α-BTX stain were

captured. Prior to 3D projection, a representative region of uniform intensity in the area

of fibrin material autofluorescence was located from one confocal z-slice from each

sample. Intensity within each region was then quantified in each color spectrum (405 nm

wavelength for DAPI, 473 nm for the fibrin scaffold, and 635 nm for α-BTX) by selecting

the region in FIJI and recording the mean value from the selection’s histogram. These

intensities were then averaged within each treatment group for each color, the treatment

group with all color averages nearest the center of the possible intensity range was

identified, and differences in selection intensity between each sample and this center

average were calculated for each color.

To allow direct comparison of all samples, brightness was normalized across

samples by adjusting maximum intensity in the FIJI B&C (Brightness & Contrast) window

for each color of each image in each confocal z-stack by the calculated difference. This

assured that α-BTX labeling of AChRs would not be differentially represented between

groups, and that DAPI and fibrin autofluorescence would not overpower any AChR

signal present. 3D projection was then performed in FIJI by using normalized z-stacks,

as described above. Scale bars and other image labels were applied in Powerpoint

(Microsoft, Redmond, WA).

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3. Results


C2C12 cells readily attached to fibrin gels and took on an elongated morphology

over 7 days of culture in differentiation medium. As observed by fluorescent labeling with

α-BTX, these elongated cells often expressed AChRs diffusely within their membranes

(Fig. 2A); however, in the absence of exogenous agrin supplementation, there was no

clustering or other observable spatial organization of these receptors. In samples treated

with agrin dissolved within the culture medium, AChR clusters were observed throughout

numerous cells within 16 hours after treatment, though without spatial organization or

predictability (Fig. 2B).

Polystyrene microparticles were observed to spontaneously settle by gravity from

suspension over cell cultures, often coming to rest near elongated C2C12 cells. Though

these particles were visible alongside cells under fluorescence microscopy, presumably

due to autofluorescence of the polystyrene material, no change in cellular AChR quantity

or location was observed resulting from treatment with microparticles that did not carry

adsorbed agrin (Fig. 2C). Conversely, when agrin-adsorbed microcarriers were added to

the culture, elongated cells near these agrin-presenting particles displayed focal areas of

strong AChR labeling at areas of potential cell-microparticle contact after as little as 16

hours (Fig. 2D), indicative of AChR clustering. Though it was not clear from

epifluorescence micrographs if these clusters only occurred at areas where

microparticles and cells fully came into contact or if proximity was sufficient, AChR

clustering was exceedingly rare in portions of cells substantially distant from an agrin

microcarrier. This indicated that microparticles did not act as depots for the more general

release and diffusion of soluble agrin in the cell culture media.

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Fig. 2. Efficacy of agrin treatment in 2D. Epifluorescent micrographs of differentiated C2C12 cells stained with fluorescent α-bungarotoxin summarize the response of cultured muscle cells to agrin presented for 16-24 hours prior to staining. (A) Unconditioned cells are diffusely fluorescent when stained, indicating the presence of AChRs within the membrane. (B) When treated with agrin-conditioned culture medium, AChRs cluster together in a spatially uncoordinated fashion. (C) Though untreated polystyrene microparticles are visible after addition in suspension due to autofluorescence, there is no AChR clustering response. (D) AChR clusters are visible in the membranes of cells contacting microparticles carrying surface-adsorbed agrin in a spatially coordinated fashion. Arrows highlight examples of AChR clustering. Scale bar represents 15 μm.

To observe C2C12 cell response to microparticle-mediated agrin delivery in the

context of these planar 2D fibrin surfaces, samples were imaged with confocal

microscopy and reconstructed in ImageJ in all three dimensions. This technique allowed

resolution of the planar material, adherent cells, and nearby microparticles across their

vertical thickness, especially after combining horizontal planar “slice” images into a 3D

rendered image. 3D confocal images showed that, while cells sometimes exhibited

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AChR clusters (red) a small distance away from agrin-adsorbed microcarriers after 16-

24 hours of treatment, this never occurred in cells that did not contact a microcarrier

(Fig. 3A). Applying a thresholding algorithm using ImageJ 3D viewer allowed structures

visualized in native confocal microscopy, which are by necessity somewhat translucent,

to be viewed as opaque surfaces. This technique demonstrated even more clearly the

contact between cellular AChR cluster and agrin microcarrier (Fig. 3B, which represents

the exact location shown in Fig. 3A).


microcarriers

Agrin delivery over longer time periods was investigated using covalently-bound

agrin microcarriers. After delivery to fibrin-adherent C2C12 cells over the last 5 days of

culture in differentiation medium, AChR clustering was observed in nearby cells (Fig. 3C-

3D). Though this effect was similar to that seen after 1 day of treatment with agrin-

adsorbed microcarriers, AChR clusters near covalent agrin particles were both more

expansive and more diffusely spread from the area of cell-microparticle contact.

Subsequently, microcarriers with covalently bound agrin were embedded into the

underlying fibrin gel to explore the potential for agrin signaling of cells from the substrate

biomaterial itself without the necessity to further manipulate constructs after cell seeding.

This embedding was selected as it would be a more relevant embodiment of agrin

presentation for implantation of these materials in vivo in future studies. Imaging of these

samples (Fig. 3E-3F) revealed that C2C12 cells attached to the modified fibrin gel,

elongated, and exhibited AChR clusters similar to their behavior in previous

experiments. The embedded fluorescent polystyrene microparticles were also visible,

confirming the suitability of the fabrication method for immobilization of microparticles

over the full 8 days of culture.

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Fig. 3. AChR clustering adjacent to agrin-presenting microparticles in 2D. (A-B) After 16-24 hours of presenting microparticles by suspension within culture medium, AChR clusters (red) are present in differentiated C2C12 cells cultured on fibrin and adjacent to microparticles (green) presenting surface-adsorbed agrin, even in complex cell topographies. (C-D) Similar behavior is observed after 5 days of suspension treatment in cultured cells adjacent to microparticle surfaces which have been covalently bound to agrin. (E-F) This behavior is maintained when covalently-bound agrin microparticles are embedded within the underlying fibrin gel for the entire 8-day period of culture. Successful agrin presentation via biomaterial-embedded particles enables the construction of true 3D scaffolds for simultaneous presentation of defined mechanical and chemical cues. A, B, and C represent selected rotational views of a 3D rendered confocal image stack, while B, D, and F represent the same 3D renders after application of a thresholding algorithm to visually reconstruct microscopic surfaces. Scale bar represents 10 μm.

Of note, all experiments featuring agrin microcarriers also included a control

group of microparticles that did not present agrin. Though cells and microparticles could

always be seen upon examination of these control samples, no cells exhibited any AChR

clustering, instead appearing similar to Fig. 2C. This strongly indicated that the surface-

bound agrin was responsible for the AChR clustering that was observed.

3.3. 3D in vitro model system for evaluation of combined agrin presentation, fibrin

materials, and mechanical strain

To further evaluate the utility of these materials for skeletal muscle tissue

engineering, rMDCs were used as a cell model in place of the C2C12 cells used in prior

experimentation. Moreover, insoluble agrin and uniaxial mechanical stretch were

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incorporated to explore their respective effects and possible synergy on AChR clustering

in rMDCs.

Microscopy of cell-seeded scaffold sections showed that fibrin, like polystyrene,

is autofluorescent. This allowed visualization of the scaffold (green) in resulting images

without requiring a specific label for fibrin, and controlled for potential non-specific

fluorescence in spectra used for labels when channels were overlaid. rMDCs seeded on

3D fibrin materials with embedded covalently-bound BSA control microcarriers, which

thereby lacked exogenous agrin, exhibited little AChR labeling of any kind and no AChR

clustering when not subjected to cyclic strain (Fig. 4A). This was consistent with a lack of

AChR expression. rMDCs seeded on 3D fibrin with embedded covalently-bound agrin

microcarriers exhibited a similar lack of AChRs and AChR clusters when cultured

statically (Fig. 4B). This suggested that agrin alone is insufficient to induce AChR

expression or clustering in rMDCs in the 3D fibrin environment.

In contrast, rMDCs seeded on BSA-presenting 3D fibrin scaffolds exhibited diffuse AChR

labeling (Fig. 4C) after bioreactor preconditioning, similar to elongated C2C12 cells in

the 2D model (Fig. 2A). However, these AChRs were spatially unorganized, indicating

that mechanical strain alone cannot induce AChR clustering. By contrast, rMDCs seeded

on agrin-presenting 3D fibrin scaffolds displayed extensive tracts of clustered AChRs

after bioreactor conditioning at all observed cell-seeded areas (Fig. 2D-2F). Interestingly,

these clusters were neither spatially limited to discrete areas of cell- particle contact nor

elliptical in shape as was often observed in 2D culture (compare to Fig. 2D and Fig 3A-

3F). Instead, they featured large, interconnected, complex geometry (Fig. 5A-5D

represent magnified views of Fig. 5E-5F, which are in turn selected areas from Fig. 4D

and Fig. 4F).

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Fig. 4: Dramatic impact of bioreactor preconditioning on AChR expression and organization in 3D tissue engineered constructs seeded with rat MDCs. (A) Very little AChR labeling (red) was observed in unstretched fibrin constructs (green autofluorescence) with embedded microparticles presenting covalently immobilized BSA, indicating an immature phenotype in cultured cells (blue nuclei). (B) AChR labeling is likewise absent from unstretched constructs with embedded agrin-presenting microparticles, indicating that agrin alone does not induce significant differentiation of rat cells under these conditions, and further, apparently cannot induce clustering. (C) Diffuse labeling of AChRs is seen in constructs with BSA-presenting microparticles after conditioning with cyclic mechanical strain in a bioreactor. There is no observed AChR clustering. (D-F) Bioreactor preconditioning induces robust appearance of AChRs in cells cultured on constructs with embedded agrin-presenting microparticles. We consistently observed AChR expression as well as organization of these receptors into large, tract-like clusters, indicating that mechanical stretch and agrin presentation may act synergistically. Arrows indicate example SiO2 microparticles embedded within the fibrin scaffold. Scale bar represents 20 μm.

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Fig. 5: Representative examples of AChR clustering in a bioreactor-preconditioned agrin-presenting tissue engineered construct. Shown are rMDCs (blue nuclei) seeded on the surface of fibrin scaffolds (green autofluorescence) carrying embedded SiO2 microparticles with covalently surface-bound agrin. Cells exhibited AChR clustering (red) after constructs were subjected to 5 days of cyclic mechanical strain. In contrast to the discrete geometry of AChR clusters observed under static culture with C2C12 cells (see Figs. 2-3), AChRs in these rMDCs cultured on bioreactor-conditioned agrin-presenting scaffolds clustered into extensive structures of complex geometry (A-D) more reminiscent of the appearance of native MEPs in vivo. By contrast, AChR clusters were never observed under static 3D culture conditions (see Fig. 4), indicating mechanical stretch and agrin presentation may act synergistically in the formation of mature MEP-like structures in vitro. A-D represent selected magnified views from a 3D-projected confocal z-stack, and are shown in the context of the larger section (E-F, compare to Fig. 4D & 4F). Scale bars represent 20 μm.

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4. Discussion

4.1. Rationale and hypothesis

We previously described [45,46] efforts to functionally reconstruct rat skeletal

muscle in vivo using a tissue engineering approach combining biomaterials, cells, and

preconditioning via mechanical stimulation. We referred to that technology as a Tissue

Engineered Muscle Repair (TEMR) construct. That approach succeeded in restoring 60-

70% of native skeletal muscle force generation within 2 months of implantation of TEMR

in a surgically created rodent VML injury model. These improvements in contractile force

were significantly greater, with respect to the rate and / or magnitude of functional

recovery, to those observed following no repair of the VML injury [45,46], implantation of

a decellularized scaffold without cells [45], or a cell-seeded scaffold without a

preconditioning step [46]. However, as noted, no intervention fully restored injured

muscles to an uninjured native level of force generation.

These studies illustrate the need for further improvements in tissue engineered

skeletal muscle repair.The current study represents another step toward developing

additional technology platforms that may provide further functional improvements for

repair of VML injuries. Notably, though AChR clustering has been described in response

to adsorbed agrin microcarriers in the literature [60], the current approach critically

evaluated not only adsorption but also covalent linkage of agrin to microcarriers, both in

a biomaterial context. Fibrin was the selected biomaterial in the current study as it could

be assembled around agrin microcarriers, allowing their immobilization within the cell

seeding surface. Further, fibrin degrades by native bodily processes in vivo on the order

of weeks [66], and can be fabricated to possess mechanical stiffness similar to native

skeletal muscle (Young’s modulus: 110 – 320 kPa fibrin [63] vs. 7 – 127 kPa muscle

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[67]). This combination of parameters suggested that fibrin would be an ideal material for

both this proof-of-concept study in vitro and potential further application in vivo.

The central hypothesis was that biomaterial-mediated agrin supplementation to

muscle cells in vitro, either alone or in combination with mechanical stimulation, would

result in creation of a nominally more native-like muscle phenotype as characterized by

the presence of agrin-mediated AChR clustering.


To begin, we formed 2D (planar) fibrin gels on cover glass and then seeded

these gels with C2C12 mouse myoblasts. After 7 days of culture, microparticles were

suspended in culture medium, which settled via gravity onto the cell layer. These

particles often aggregated near elongated cells (see Fig. 2C-2D), likely due to

topography – essentially “valleys” between cell edges and the biomaterial or between

parallel cells which discouraged lateral movement. Control microparticles lacking agrin

(Fig. 2C) and agrin microcarriers distant from cells (elements of Fig. 2D) shared a

uniformly fluorescent appearance, indicating autofluorescence of the polystyrene

material.

Following fluorescent α-BTX staining, spatially uncoordinated AChR clusters

were observed in cells treated with agrin in solution (Fig. 2B). By contrast, agrin

microcarriers adjacent to cells (Fig. 2D, see arrows) were often observed to have a

bright spot somewhere within the microparticle periphery. This indicated the presence of

a cluster of AChRs adjacent to or underneath the microparticle, as nonspecific artifacts

related to particle material or shape (e.g., refraction through a polstyrene sphere) would

have been observed independently of cells or agrin. Also of note was that all C2C12

cultures, regardless of exposure to agrin and / or polystyrene, contained cells which

expressed AChRs after 7 days of culture in differentiation medium. This result indicated

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that neither material had a qualitatively-detectable effect on overall AChR expression,

only on clustering of AChRs.

We next employed confocal microscopy, which enabled 3D reconstruction of the

particle-cell interface. These experiments used fluorescent microparticles, as

polystyrene autofluorescence was less intense in confocal images than it was under

epifluorecence. Agrin microcarriers were observed to directly contact cells after settling

from suspension. Further, AChR clusters were closely associated with the cell-particle

interface (Fig. 3A-3B).

These results suggested the suitability of polymeric agrin microcarriers in the

context of a fibrin scaffold. While it is well-known that hydrogels can serve as depots for

protein delivery [68–73], this design would presumably lead to diffusion-based release of

the protein [73] and its eventual depletion from the system. By contrast, coupling agrin to

the surface of a microcarrier would allow spatial control of both location and dose, and

could focus a relatively small amount of protein to therapeutically high concentrations by

preventing diffusion. Moreover, in normal physiology, agrin is immobilized within the

synaptic basal lamina [58], a spatial arrangement more closely emulated by an agrin-

presenting surface.

Similar results were observed if agrin microcarriers were added during the last 1-

3 days of culture. However, no AChR clusters were observed if microcarriers were

added earlier (data not shown), which corresponded to an exchange of culture medium

after agrin addition. This may indicate that agrin desorbed from the microparticle surface,

either spontaneously or by a Vroman-like exchange [74], and was subsequently lost

upon media removal. Instead of a number of days, we reasoned that AChR clusters

should ideally be observed over weeks of culture. This is because presentation of

endogenous neural cues to implanted tissue engineered constructs could require weeks

to months at the clinically-expected reinnervation rate of 1 mm/day [59].

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microcarriers

This mismatch presented a limitation which we reasoned could be overcome by

covalently bonding agrin to the microcarrier surface. We used the commonly-used

crosslinker, EDC, to bind agrin to commercially-available carboxylate microparticles. We

had previously noted that no AChR clustering was observed if agrin-adsorbed

microcarriers were treated with ethanol prior to their addition in culture (data not shown),

supporting the hypothesis that adsorbed agrin was displaced by a Vroman-like

exchange. Ethanol sterilization of covalently-bound agrin microcarriers, then, additionally

inactivated or washed away any agrin that had simply adsorbed to the particle surface.

This allowed conclusive evaluation of covalent agrin immobilization.

Covalently-bound agrin microcarriers were added to C2C12 cells on 2D fibrin for

the last 5 days of culture, which included one or more exchanges of culture medium.

Though the AChR clusters observed were less spatially restricted to areas of cell-particle

contact than those observed near adsorbed agrin microcarriers (compare Fig. 3C-3D to

Fig. 3A-3B), they were often larger, potentially as a result of longer-term agrin

presentation. This demonstrated suitability of covalently coupling agrin to microcarriers

for longer-term signaling of muscle cells, despite the inability to specifically crosslink

agrin in non-active sites using EDC.

Our next goal was to immobilize agrin microcarriers within fibrin to create an

agrin-presenting substrate for cell attachment. This was accomplished by suspending

microcarriers within the fibrinogen solution before scaffold polymerization. In contrast to

prior experiments, no movement or loss of microparticles was observed during medium

exchanges or histological processing of samples fabricated using this method. This

indicated microparticles were successfully embedded within fibrin.

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Imaging of stained scaffolds revealed AChR clusters near immobilized

microparticles (Fig. 3E-3F). Of note were fluorescent “halos” around many microparticles

(Fig. 3E) which manifested as presumptive artifacts in surface renders (Fig. 3F). These

likely resulted from light refracting at the interface between polystyrene and fibrin, as

they did not fuly encircle the microparticles. Despite this signal, AChR clusters were

apparent, indicating that immobilized agrin microcarriers could be easily incorporated

into 3D scaffolds.

4.4. 3D in vitro model system for evaluation of combined agrin presentation, fibrin

materials, and mechanical strain

In the 2D model system, we used the C2C12 cell line due to its ready availability

and straightforward differentiation in vitro. In the 3D model system, we wished to

evaluate the effects of agrin instead on rMDCs. First, this was due to our extensive prior

experience in 3D skeletal muscle tissue engineering approaches using this cell type [44–

46]. Second, future evaluation of the tissue-engineered construct in vivo could be

performed in immune-competent animals (in this case, Lewis rats) if it incorporated an

isogeneic cell source.

The 3D fibrin scaffold was formed around a template bundle of sacrificial fibers

as previously described [63], as this method provided a material with well-understood

mechanical properties suitable for bioreactor preconditioning. These scaffolds

incorporated SiO2 covalently-bound agrin microparticles that, unlike their polystyrene

analogs, could also survive scaffold patterning in acetone. However, as these particles

are made from a non-degradable material, alternatives to SiO2 may be required for any

future use of this technology in vivo. In contrast to the 2D model, microcarriers covalently

bound to BSA served as a control in the unlikely event that previously-observed AChR

clustering was not specific to agrin. Though this patterned scaffold could potentially

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support cell attachment, growth, and ideally alignment within its pores, we elected to

seed rMDCs on the scaffold exterior. Efficient seeding of the scaffold interior with cells

represents a possible future development of the tissue engineering approach described

here, but may require a strategy to enhance mass transport through the construct.

Prior results [46] indicated that bioreactor preconditioning has a significant effect

on muscle-specific protein expression in rMDCs. Based on this observation, we

hypothesized preconditioning would also affect AChR clustering in rMDCs or their

sensitivity to exogenous agrin. To this end, the effect of mechanical preconditioning on

AChR clustering in seeded constructs was evaluated, both alone (i.e., in BSA-presenting

scaffolds) and in combination with agrin.

Critically, to minimize the risk of observing a carefully selected clustering event

instead of representative phenomena, areas within α-BTX-stained scaffold sections were

selected for imaging using the DAPI and fibrin autofluorescence spectra alone. Sections

were then imaged post hoc for labeling of AChRs. This ensured that imaged areas would

contain cells - and that their relation to the scaffold could be determined - but that

presence or absence of AChR clusters would not bias data collection.

As we hypothesized, little AChR expression was observed in statically cultured

(i.e., non-preconditioned) controls, regardless of the absence (Fig. 4A) or presence of

agrin (Fig. 4B). Some AChR labeling was observed in preconditioned constructs

incorporating BSA microcarriers. However, this labeling was diffuse, similar to that

observed in differentiated C2C12 cells that were not treated with agrin (compare Fig. 4C

to Figs. 2A & 2C). By contrast, AChR clustering was exclusively observed in

preconditioned, agrin-presenting constructs (Figs. 4D-4F) and was widespread across

the cell-seeded surface. Combined with the randomized method of selecting areas for

imaging, these results suggested that AChR clustering may result from a synergistic

effect of mechanical stretch and exogenous agrin supplementation.

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In normal skeletal muscle physiology, the accumulation of AChRs in the MEP of

mature synapses takes on a tortuous morphology [75]. Interestingly, the AChR clusters

observed in preconditioned agrin constructs were not confined only to the cell-

microparticle interface as was often observed in 2D (see Fig. 2D and Fig. 3). Instead,

AChRs clustered into extensive structures of complex geometry (Fig. 5) more

reminiscent of the appearance of native MEPs in vivo. This reinforced the indication that

mechanical stretch and agrin presentation may act synergistically in the formation of

more mature MEP-like structures in vitro. However, the exact mechanisms giving rise to

the complex morphology of these AChR clusters are unclear at this time.

These results critically expand upon existing studies from the literature

documenting agrin supplementation of skeletal muscle tissue engineering approaches

[50,53]. Primarily, all studies of this nature to date have evaluated the effects of soluble

agrin stimulation of muscle constructs, either in vitro alone [50] or in vitro prior to

implantation in vivo [53]. These studies demonstrated increased force production in vitro

by engineered constructs [50] and, potentially, accelerated innervation of constructs

implanted in vivo [53] following agrin stimulation. The current study documents a method

by which the agrin signal to construct-resident cells can be maintained over longer

periods of time, including after implantation in vivo, a potentially useful extension to

previous approaches. Further, results described here document potential synergy

between agrin presentation and cyclic mechanical strain on AChR clustering. Future

studies may confirm that effects on contractile function and construct innervation, shown

elsewhere after soluble agrin stimulation [50,53], also result from immobile agrin

presentation alone or in conjunction with cyclic stretch. Indeed, as exogenous agrin

maintained in immobile contact with muscle cells more closely mimics the spatial

arrangement found in native synapses, it may enhance relevant markers of skeletal

muscle construct function relative to an otherwise equivalent soluble signal.

101

5. Conclusions

In this work, we describe AChR clustering in muscle cells cultured in vitro on

agrin-presenting biomaterials suitable for skeletal muscle tissue engineering

applications. AChR clusters were observed in a C2C12 cell line in a 2D model system

and in a more relevant 3D model system using expanded rat MDCs. These models

demonstrated the utility of microparticles as an immobile, non-diffusion based platform

for presentation of agrin to cells over several days in culture. Moreover, extensive AChR

clustering may result from synergy between mechanical strain and exogenous agrin. The

3D model embodied a tissue-engineered muscle repair construct in vitro, consisting of

fibrin and microparticle biomaterials, pharmacological stimulation via agrin, and cell

signaling via mechanical stretch. Though promising, these results merit further

investigation in vivo to determine if agrin-presenting biomaterials described here will

enhance functional outcomes in tissue engineered repair of clinically intractable VML

injuries.

Acknowledgements

This work was funded by an award from the Telemedicine and Advanced

Technology Research Center, by the Wake Forest University Department of Biomedical

Engineering, and by the Wake Forest Institute for Regenerative Medicine. The authors

gratefully acknowledge the assistance of Hannah Baker, Christopher Bergman, Hayden

Holbrook, Venu Kesireddy, Juliana Passipieri, Pooja Patil, Mevan Siriwardane, Claire

Staley, and Taylor Zak in performing experiments described herein, as well as Cathy

Mathis and Cynthia Zimmerman for their assistance with histology and Christopher

Booth for his assistance in confocal microscopy.

102

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CHAPTER IV

ADVANCES TOWARD ENABLING THREE-DIMENSIONAL FIBRIN SCAFFOLDS FOR

TISSUE ENGINEERED MUSCLE REPAIR

John B. Scott

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1. Introduction

Prior Chapters explored fibrin biomaterials as a mechanical support and physical

guide for neurite extension from peripheral nerve (Chapter II) and as a substrate for the

presentation of insoluble agrin to adherent muscle cells in a tissue engineered muscle

repair construct (Chapter III). These results demonstrated that fibrin hydrogels can be

fabricated to withstand maximum loads over 1 N in magnitude and feature elastic moduli

between 110-330 kPa. Additionally, they can incorporate particulate drug delivery

vehicles and a porous network of aligned conduits.

However, Chapter II only demonstrated the use of fibrin in the guidance of nerve,

though many of the above parameters also suggested its use as a substrate for skeletal

muscle. For example, native skeletal muscle elastic modulus (79-126 kPa [1]) is similar

to that of the patterned fibrin. Further, native muscle is composed of myofibers aligned

along a preferred axis of contraction, surrounded in turn by layers of connective tissue

similar to the micro-architectural patterning of these scaffolds. This arrangement echoes

anatomy of a peripheral nerve (compare connective tissue components of Fig. 1.1 from

[2] and Fig. 1 from [3]) and cross-sections of patterned fibrin (Chapter II, Fig. 2). Also,

physical cues such as nano- and micro-fibers and microgrooves [4–9] have been

previously used to guide muscle cell alignment and / or mophology. This indicates that

patterned fibrin could also organize seeded muscle cells into the alignment necessary

for macroscopic force production.

Therefore, this Chapter describes efforts to create 3D tissue engineered muscle-

like constructs using patterned fibrin. One major goal was the robust seeding of muscle

cells within patterned fibrin scaffolds, which could potentially be achieved by perfusion of

muscle cell suspensions through the conduit network. Flow perfusion of engineered

tissue constructs has been significantly described in the literature, though typically as a

method to enhance mass transport and / or to modulate cellular behavior [10–16]. Some

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studies, though, have explored perfusion of cell suspensions as a method to enact rapid

and uniform cell seeding within porous scaffolds [17–21]. The majority of investigation on

scaffold perfusion has focused on bone tissue engineering [10–12,14–18,20,22], though

it has also been applied to other tissues such as kidney [21], cartilage [23,24], tendon

[13], and cardiac muscle [25,26]. Of note, few perfusion approaches have attempted to

seed scaffolds with parallel longitudinal pores [19], and none of which we are aware

have done so using skeletal muscle cells.

A second goal of this study was to incorporate agrin microcarriers, studied

extensively in 2D using polystyrene microparticles, into the fibrin scaffold. This was

challenging, as polystyrene is dissolved by the same acetone immersion used to remove

sacrificial fibers during scaffold fabrication. To overcome this challenge, two parallel

strategies were employed. First, water soluble polymers - notably polyvinyl alcohol

(PVOH) and poly(ethylene oxide) (PEO) - were evaluated for the formation of fibers

which could pattern fibrin scaffolds without damaging polystyrene agrin microcarriers.

Second, a microparticle material which did not dissolve in acetone - silica - was

employed to fabricate agrin microcarriers compatible with existing scaffold fabrication

methods.

It is important to note that, though presented here after Chapter III due to

preparation of that Chapter for manuscript submission, experiments described below

were performed between the 2D and 3D experiments in Chapter III. The current Chapter

represents efforts to incorporate agrin delivery as described in 2D experiments of

Chapter III into 3D fibrin scaffolds, and further, to enable cell seeding throughout the

interior of such a scaffold. Due to continued difficulties in meeting the second of these

goals, as described below, the surface seeding methods described in 3D experiments in

Chapter III were elected to evaluate proof of concept of fibrin-mediated agrin delivery.

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Nevertheless, the results discussed below represent important findings on which to base

future development of these technologies.

2. Methods

Unless otherwise noted, all aqueous formulations are described as % v/v.

Further, all methods were performed at ambient pressure on a lab bench at a room

temperature of ~22°C unless otherwise noted.

2.1. Surface seeding of 3D planar fibrin scaffolds

To evaluate the suitability of patterned fibrin for supporting the attachment and

growth of muscle cells, a flat (planar) block of fibrin was fabricated using methods similar

to those described in Chapter II. To begin, aligned cellulose acetate (CA) fibers

(generous donation of Eastman Chemical Company, Kingsport, TN) ~12 μm in diameter

were arranged into a mat ~1 mm thick. A solution of bovine fibrinogen (Sigma-Aldrich,

St. Louis, MO) at 200 mg/mL in PBS was added to the fibers and left undisturbed until it

completely impregnated the mat. Fibrinogen was polymerized to fibrin by adding a

solution of 125 U/mL thrombin and 2.75 μg/mL (~24.8 mM) CaCl2 (both Sigma) in PBS,

wrapping the wetted mat in Parafilm (Bemish Flexible Packaging, Oshkosh, WI) to

prevent evaporation, and leaving undisturbed for at least 4 hours.

This fibrin sheet was then sectioned into smaller scaffolds ~3 mm wide x 3 mm

long using a razor blade before being washed in acetone four times (at least 2 hr, 2 hr, 4

hr, and 8 hr in duration, respectively) to remove the CA fiber component and sterilize the

scaffolds. To remove acetone, scaffolds were then washed four times in sterile PBS

(same durations as acetone) in a biological safety cabinet and stored in the final sterile

wash at 4°C until use.

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Scaffolds were then seeded with C2C12 mouse myoblasts to evaluate their

response to fibrin. One scaffold each was placed into the wells of a 24-well polystyrene

tissue culture plate (Thermo Fisher Scientific, Waltham, MA). C2C12 growth medium

consisted of 89% high-glucose Dulbecco’s modified Eagle’s medium (DMEM), 10%

heat-inactivated fetal bovine serum (FBS), and 1% penicillin/streptomycin (all Thermo

Fisher Scientific) and was supplemented with aprotinin (Sigma) at 20 μg/ml (~3.07 μM)

to inhibit fibrinolysis. A 1 mL suspension of C2C12 cells (ATCC, Manassas, VA) in this

growth medium at a concentration of 50,000 cells / mL was then added to each well, and

cultures were left undisturbed at 37°C and 5% CO2 in an incubator to allow cells to settle

by gravity onto scaffolds.

After 1 day, growth medium was aspirated and replaced with differentiation

medium consisting of 97% 1:1 DMEM:F12 mixture, 2% horse serum (HS), and 1%

antibiotic-antimycotic (all Thermo Fisher Scientific), which was also supplemented with

aprotinin as above. This medium was exchanged every 2-3 days. After 6 days in

differentiation medium, cultures were supplemented with carrier-free recombinant rat

agrin (R&D Systems, Minneapolis, MN) to a final concentration of 50 ng/mL (~5.55e-10

M) and left for one final day in culture to evaluate acetylcholine receptor (AChR)

clustering in scaffold-resident cells.

After the total 8-day culture period, scaffolds were removed and fixed overnight in

10% neutral buffered formalin (NBF). Fixed scaffolds were paraffin processed in an

ASP300 tissue processor (Leica Microsystems, Buffalo Grove, IL) using the following

steps: 1) 60% isopropyl alcohol in water (IPA), 45 minutes; 2) 70% IPA, 45 minutes; 3)

80% IPA, 45 minutes; 4) 95% IPA, 45 minutes; 5) 95% IPA, 1 hour; 6) 100% IPA, 45

minutes; 7) 100% IPA, 45 minutes; 8) 100% xylenes, 45 minutes; 9) 100% xylenes, 45

minutes; 10) paraffin, 45 minutes; 11) paraffin, 1 hour; 12) paraffin, 1 hour.

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Processed samples were then mounted into paraffin blocks and sectioned to 15

μm thickness on a microtome. Tissue sections were immobilized on glass slides, dried

for 16-48 hours at 60°C, and then stored indefinitely until stained. Prior to staining, slides

were deparaffinized by sequential washes in xylenes, ethanol, and tap water.

The staining protocol was as follows. Slides were: 1) Washed three times for 5

minutes each in tris-buffered saline with Tween (TBST, Dako, Glostrup, Denmark); 2)

Blocked against non-specific binding using a blocking buffer consisting of 5% HS in

TBST for at least 30 minutes; 3) Stained for AChR localization using α-bungarotoxin

conjugated with Alexa Fluor 594 (α-BTX, Invitrogen Life Technologies, Grand Island,

NY) at 1e-7 M dilution in blocking buffer for at least 1 hour in the dark; 4) Washed a

further three times in TBST, 5 min each; 5) Mounted with cover glass using Vectashield

with DAPI (Vector Laboratories, Burlingame, CA) and dried overnight at 4°C.

Epifluorescence images were obtained with a DM4000 B microscope (Leica) with

attached Retiga 2000RV camera (QImaging, Surrey, BC, Canada), coupled with

ImagePro 6.2 software (Media Cybernetics Inc., Bethesda, MD).

2.2. 3D seeding of patterned fibrin via uni-directional perfusion of muscle cell

suspensions

Scaffolds were again fabricated by using the sacrificial templating method

described in Chapter II. Briefly, CA fibers 12 μm in diameter were arranged into a dense,

aligned bundle and placed into a cylindrical mold 4.5 mm in diameter. Fibrin was

polymerized around the fibers, the fibers were selectively removed with acetone,

patterned scaffolds were trimmed to 4 mm in length, and scaffolds were then rehydrated

in successive washes of sterile PBS.

Rat muscle derived cells (rMDCs) used throughout this Chapter were isolated,

expanded, and removed for use as described in Chapter III, Section 2.4. Likewise,

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culture medium formulations for rMDC growth medium and differentiation medium were

identical to those used in Chapter III, Section 2.1.

To seed scaffolds with rMDCs, a closed perfusion loop (Fig. 1) was created by

linking a medium reservoir, a peristaltic pump, and a scaffold chamber in series by using

3.2 mm inner diameter (ID) x 4.8 mm outer diameter (OD) Tygon® silicone tubing

(United States Plastic Corp., Lima, OH) and barbed male-to-male plastic fittings of

appropriate diameter (McMaster-Carr, Atlanta, GA). The medium reservoir was an open-

topped 50 mL conical tube (Corning Life Sciences, Tewksbury, MA), the peristaltic pump

was an Ismatec model CP78016-10 (Cole-Parmer, London, UK), and the scaffold

chamber was fabricated by placing a 4 mm length of 4.5 mm ID x 6.4 mm OD Teflon®

tubing of the same type used in scaffold fabrication within a 6 mm length of 6.4 mm ID x

9.5 mm OD Silastic® tubing (Cole-Parmer).

The reusable scaffold chamber consisted of an outer silicone tube to contain flow

and two adapters to connect this outer tube to the rest of the flow loop. This chamber

allowed scaffolds to be placed at an accessible location at a known, reproducible

distance from the pump outlet. Basic versions of this design were found to poorly

immobilize the scaffold within the chamber, instead allowing it to be drawn into the

downstream flow loop were it could become stuck and block flow or wash out into the

medium reservoir. To correct this problem, the outer tube was made larger than the rest

of the flow tubing (as listed above) and Spectra/Mesh polypropylene screen (Cole-

Parmer) was placed at both ends of the scaffold to block it from entering the smaller

perfusion tubing. In this arrangement, the 4.5 mm ID mounting tube placed within the

outer tube served both to laterally immobilize the scaffold as well as to trap the

Spectra/Mesh against the plastic adapter fittings perpendicular to the flow direction.

This perfusion loop was sterilized by piecewise exposure to either γ radiation or ethylene

oxide and then assembled aseptically in a biological safety cabinet, with the final step

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being to place a hydrated scaffold within the scaffold chamber. A 10 mL suspension of

rMDCs in rMDC growth medium at 500,000 cells/mL was then placed into the medium

reservoir and the pump was turned on for 30 minutes. The perfusate exiting the scaffold

was returned to the medium reservoir for recirculation.

Cell suspensions were perfused in two experimental groups at 1.2 mL/min and

4.8 mL/min to evaluate the effect of flow rate on cell seeding. To provide a baseline for

comparison, a static seeding control group was included by placing fibrin scaffolds

vertically in standard tissue culture polystyrene dishes and manually pipetting a

suspension of 5 million rMPCs in 100 μL growth medium over one face of the open

microarchitecture.

Regardless of treatment, scaffolds were briefly (~15 min) left undisturbed to allow

for initial cell attachment and then either immediately submerged in rMDC growth

medium to allow for cell expansion and migration or fixed (see below) for a first time

point. A second time point was taken after 24 hours in growth medium, after which

scaffolds were fixed as above. For the third time point, growth medium was exchanged

for differentiation medium after 24 hours total culture and seeded scaffolds were cultured

for a further 48 hours. At the end of all time points, experimental and control scaffolds

were fixed, embedded in paraffin, sectioned, dried, deparaffinized, and rehydrated as in

Section 2.1. Sections were then mounted using Vectashield with DAPI to allow

visualization of nuclei and counting of cell density.

Epifluorescence Images were captured as in Section 2.1. Images were analyzed

by separating each imaged scaffold section into three segments of equal length (top,

middle, and bottom) in ImageJ software, with the scaffold edge nearest the pump output

considered “top.” The area of each segment was measured and the total number of cells

in each segment was manually counted. These data were then used to calculate cell

density in each segment. As section area varies with depth of the section within the

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Fig. 1. Perfusion loop schematic. A peristaltic pump was used to perfuse a recirculating suspension of muscle cells through a closed loop, ideally resulting in uniform cell seeding within the microarchitecture of a patterned fibrin scaffold.

cylindrical scaffold, it was important to report cell density to accurately evaluate cell

seeding.

For each combination of experimental / control group and time point, three

scaffolds each were seeded and three sections from each scaffold were stained and

imaged, for a total of n=9 replicates. Statistical analysis was performed as described

below (see Section 2.8). For the immediate time point, cell densities were compared

across experimental groups within segments as well as over entire scaffold sections. For

later time points, densities were compared only within the middle segment, which the

immediate time point showed to be the most difficult to seed.

2.3. 3D seeding of patterned fibrin via bi-directional perfusion of muscle cell suspensions

Cells were again perfused through scaffolds; however, in this experiment flow

direction was reversed halfway through the seeding step. Scaffold fabrication, rMDC

culture prior to seeding, perfusion loop assembly, scaffold fixation, sectioning, staining,

and imaging, and statistical analysis were performed as in Section 2.2, with the

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exception that two groups of scaffolds were templated – one using 12 μm diameter CA

fibers and the second using 41 μm diameter poly(methyl-methacrylate) (pMMA) fibers.

Also as in Section 2.2, 5 million rMDCs were suspended in 10 mL rMDC growth

medium and perfused through a sacrificially-templated fibrin scaffold for a total of 30

min. In the first flow condition, the direction of flow in the continuous loop was simply

reversed at 15 min, leaving no delay. As this reversal had the potential to displace some

cells that had just been seeded, the flow was paused in the second group for 15 minutes

between direction changes to allow for cell attachment. Immediately after perfusion, all

seeded scaffolds were taken for an immediate time point. Cell density was counted only

for middle segments, as in later time points in Section 2.2.

Each of the four resulting experimental groups (12 μm template with no delay

between flow directions, 41 μm template with no delay, 12 μm template with a 15 minute

delay before reversing flow direction, and 41 μm template with a 15 minute delay) was

evaluated over three scaffolds. Three non-sequential sections were taken from each

scaffold for imaging and analysis, again totaling n=9 replicates.

2.4. Fabrication of water-soluble template fibers for 3D patterning

Polymers were then melt extruded, with the overall goal of producing water-

soluble fibers over a similar range of diameters to those used previously (i.e. 10–250

μm). All experimentation in Section 2.4 was performed using a Dynisco Laboratory

Mixing Extruder (LME) with a round 1/8” diameter output orifice. To extrude fibers,

polymer was melted and then forced through the heated orifice to form a preliminary

fiber. The cooling fiber was then drawn through ambient air into a smaller diameter by

tensile force using a rotating spool called the take-up system. The primary variables

when using this system, therefore, were polymer type (which dictated ideal temperature

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settings within the LME), polymer flow rate through the orifice, and collection speed of

the take-up system.

An initial experimentation period was performed using polyvinyl alcohol (PVOH,

Polysciences, Inc., Warrington, PA), of MW ~25,000 Da and 88 mole % hydrolyzed. The

goal was to identify the bounds of settings capable of extruding uniform fibers.

With the range of feasible extruder temperatures, rotor (i.e. molten polymer)

output, and take-up speeds established, the next goal was to quantify fiber properties at

discrete settings within these ranges. Extruder temperatures, which were dictated by the

polymer, were only varied in minute amounts as required to maintain good polymer flow

through the orifice. Two rotor output settings – 10% and 20% of total capacity – were

evaluated. Likewise, four take-up speed settings – corresponding to roughly equally-

spaced increments of the middle third of total capacity – were used. Fibers were

produced using all 8 combinations of these two parameters, and then three samples

from each fiber were selected from widely-spaced areas on the strand to control for

variability over time. These samples were mounted to scanning electron microscope

(SEM) chucks, sputter coated with a mixture of gold/palladium, and imaged. Fiber

diameter was measured from SEM images using ImageJ. A further portion of each

created fiber type was immersed in water to test its solubility.

A second experiment was then performed to increase ease of extrusion and

uniformity of the resulting fiber diameter. In this experiment, pMMA (Polysciences) and

PVOH pellets in varying proportions by weight were mixed well before being extruded as

in the first experiment. Again, ideal extrusion temperatures were identified. Rotor output,

take-up speeds, SEM preparation, imaging, and diameter measurement were as

described above. To evaluate water solubility of resulting fibers, a portion of each fiber

type was immersed first in acetone and then in water.

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Third, poly(ethylene oxide) (PEO, Polysciences) was evaluated by using a similar

range of extrusion parameters. After another initial evaluation of extruder temperatures

for this new polymer, rotor output, take-up speed, SEM preparation, imaging, diameter

measurement, and water solubility testing were performed as in the pure PVOH

experiment above.

Fourth, PVOH, again at ~25,000 Da MW but instead with 98 mole % hydrolyzed

(Polysciences), was evaluated over a similar range of extrusion parameters. This

material was evaluated both alone and mixed with pMMA as in the first and second

experiments above, with new extruder temperatures determined first but subsequently

using identical rotor output and take-up speeds. The resulting fibers were subjected to

SEM preparation, imaging, and diameter measurement as well as water- and acetone /

water-solubility testing as appropriate, as above.

Fifth, 88% hydrolyzed PVOH granules were extensively dried under vacuum prior

to adding them to the extruder. Moreover, extruder heat was increased, which was

predicted to be enabled by removal of surface water and would allow the take-up system

to draw the resultant fiber to a smaller diameter. The resulting fibers were subjected to

SEM preparation and analysis, as well as water solubility testing, as above.

A sixth and final experiment was then undertaken in an attempt to extrude fibers

by using varying amounts of dried 98% hydrolyzed PVOH with smaller amounts of dried

88% hydrolyzed PVOH. After again evaluating feasible extruder temperatures, fibers

were extruded using identical rotor output to that above, but were collected using a

custom-fabricated replacement take-up system which was capable of much faster

speeds and, ideally, production of smaller-diameter fibers. The resulting fibers were

again subjected to SEM preparation and analysis, as well as water solubility testing, as

above.

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2.5. Perfusion seeding and subsequent cyclic stretch of 30mm long patterned fibrin

scaffolds containing agrin-presenting SiO2 microspheres

The goal of the following experiment was to fabricate fibrin scaffolds with

embedded covalently-bound agrin microcarriers to a length suitable for preconditioning

in a cyclic stretch bioreactor, and then to seed these scaffolds with cells. To this end, the

following combination of parameters was used: 1) Covalently-bound silica (SiO2) agrin

microcarriers, immobilized within 2) fibrin scaffolds sacrificially patterned using CA

template fibers, 3) seeded via perfusion of a suspension of rMDCs, and subsequently 4)

preconditioned using a bioreactor designed to apply cyclic mechanical strain.

Agrin was covalently linked to the surface of carboxylate SiO2 microparticles

(Corpuscular, Cold Spring, NY) using the N-(3-dimethylaminopropyl)-N′-

ethylcarbodiimide hydrochloride (EDC) / N-hydroxysulfosuccinimide (sulfo-NHS) zero-

length crosslinking reaction described in Chapter III, Section 2.1 (reagents: MES from

Acros Organics, Geel, Belgium; NaCl, EDC, and sulfo-NHS from Sigma). Fibrin scaffolds

were fabricated to 30 mm in length and 3 mm in diameter as described in Chapter II, but

using a fibrinogen solution containing a suspension of covalently-bound silica agrin

microcarriers (see methods, Chapter III, Section 2.5). For these 3 mm diameter

scaffolds, the chamber designed to immobilize the scaffold required a thicker Teflon®

mounting tube with ID = 3.2 mm to successfully immobilize the smaller smaffold. The

overall length of the scaffold chamber was adjusted to snugly accommodate the new

scaffold size.

Calculations of nuclear density based on previous perfusion results indicated that

a density of 17,300 microparticles / mm3 of scaffold would provide one agrin microcarrier

per nucleus, which was selected for microparticle loading. As in Chapters II and III,

scaffolds were patterned with 12 μm diameter CA fibers, which were removed by

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sequential washes of acetone. Micropatterned, agrin-presenting fibrin scaffolds were

then rehydrated with sequential washes of sterile PBS.

rMDCs were then seeded as in Section 2.2 of this Chapter, with a uni-directional

30-minute perfusion of 5 million cells suspended in 10 mL growth medium. Cell-seeded

constructs were then subjected to bioreactor preconditioning as in Chapter III. Briefly,

constructs were aseptically removed from the perfusion loop and mounted into the

clamps of a cyclic stretch bioreactor, leaving a 20 mm gap between clamps. Constructs

were then subjected to 10% mechanical strain once every 20 seconds for the first five

minutes of every hour. rMDC growth medium was supplemented with 20 μg/mL aprotinin

to prevent fibrinolysis of the scaffold and was exchanged every 2-3 days. After 7 days,

preconditioned constructs were fixed, embedded in paraffin, sectioned, stained for

cellular location with DAPI and for AChR presence and organization with fluorescently-

labeled α-BTX, and imaged under epifluorescence as described in Section 2.1 of this

Chapter.

2.6. Optimization of microsphere immobilization density within fibrin scaffolds

Alternative methods to embed agrin microcarriers were then explored in order to

increase their density within the scaffold. Template CA fibers were packed in Teflon®

molds similarly to Section 2.5 of this Chapter, but with roughly 70% the quantity of fiber.

Moreover, untreated silica microparticles were included at twice the previous

concentration in fibrinogen solution. Scaffold fabrication and perfusion cell seeding were

performed as in Section 2.5, but the bioreactor preconditioning step was omitted.

Cell-seeded constructs were also processed as in Section 2.5, and imaged using

a FITC fluorescence filter (for scaffold and particle autofluorescence) as well as a DAPI

filter (for cell nuclei). This combination of parameters was selected to evaluate the

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hypothesis as simply and rapidly as possible, leaving the inclusion of agrin and

mechanical stretch for later evaluation.

2.7. 3D seeding of 15 mm long patterned fibrin via perfusion of muscle cell suspensions

The final perfusion seeding study consisted of two experiments. The first

experiment was performed to compare ten different experimental conditions, each

featuring one replicate. The goal in this case was to identify one or more parameters that

were common across highly-cellular groups, which could be used in the second

experiment. Again, the scaffold chamber length was adjusted to accommodate the new

scaffold size.

One parameter that had never been evaluated was stiffness of the mounting

material placed between the fibrin scaffold and the outer tubing. Any non-uniformities in

pump output could result in pulsatile changes in pressure. A scaffold mounted within a

rigid tube, such as the Teflon® employed by previous designs, could be compressed or

deformed by these pulses. By comparison, a less rigid mounting material such as

silicone tubing, could absorb some of the pressure pulses and potentially make flow

more uniform at the site of the scaffold. However, as the scaffold mounting material had

never been varied before, it was unknown if this new parameter would interact

unpredictably with others, giving rise to many possible experimental conditions.

The ten conditions evaluated in the first experiment were as follows:

1) More rigid (Teflon®) mount, 12 μm diameter (CA) fiber, 10 mm scaffold length, 30

min perfusion, 4.8 mL/min flow rate, bi-directional perfusion (with flow direction

changed immediately halfway through the total duration)

2) Teflon® mount, CA fiber, 15 mm scaffold length, 60 min perfusion, 4.8 mL/min flow

rate, bi-directional perfusion

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3) Teflon® mount, 41 μm diameter Alex James pMMA (AJ1) fiber, 10 mm scaffold

length, 30 min perfusion, 4.8 mL/min flow rate, bi-directional perfusion

4) Teflon® mount, AJ1 fiber, 15 mm scaffold length, 60 min perfusion, 4.8 mL/min flow


5) Teflon® mount, AJ1 fiber, 15 mm scaffold length, 60 min perfusion, 9.6 mL/min flow

rate, uni-directional perfusion

6) More compliant (silicone) mount, CA fiber, 10 mm scaffold length, 30 min perfusion,

4.8 mL/min flow rate, bi-directional perfusion

7) Silicone mount, CA fiber, 15 mm scaffold length, 60 min perfusion, 9.6 mL/min flow


8) Silicone mount, AJ1 fiber, 10 mm scaffold length, 30 min perfusion, 4.8 mL/min flow

rate, uni-directional perfusion





The silicone mounting tube, where indicated, replaced the 3.2 mm ID x 6.4 mm

OD Teflon® mounting tube within the scaffold chamber of the perfusion loop, and had

identical dimensions.

The second experiment consisted of the following experimental groups (with

number of replicates shown in parentheses), using only scaffolds 15 mm in length

templated with CA fibers and mounted in silicone:

1) 30 min perfusion, 4.8 mL/min flow rate, bi-directional perfusion (n=2)




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5) 60 min perfusion, 9.6 mL/min flow rate, uni-directional perfusion (n=3)

Microspheres were not included in scaffolds used in these experiments in order

to aid cell quantification by nuclear stain, as silica microspheres are autofluorescent and

similar in size to cell nuclei. Otherwise, scaffolds were fabricated as in Section 2.6

above. Perfusion of 5 million passage 2 rMDCs in 10 mL growth medium per scaffold

was carried out in a closed flow loop as in previous experiments. After perfusion,

scaffolds were processed and stained with DAPI as in Section 2.6 above. Again, cell

density was counted in the middle third of the scaffold length and used to evaluate

perfusion performance.

2.8. Statistical analysis and presentation of results

Statistical analysis was performed in GraphPad PRISM 5. ANOVA was first used

in each study to determine presence or absence of statistically significant differences

across an entire data set. When indicated by ANOVA, Tukey’s Test was used as post-

hoc analysis to identify individual pairwise differences. For all comparisons, p<0.05 was

considered statistically significant. All numerical results are presented as mean ±

standard deviation (SD) unless otherwise noted.

3. Results


The planar fibrin material supported attachment and growth of C2C12 cells, as

shown by presence of numerous nuclei stained with DAPI (blue) in Fig. 2. Cells were

observed not just on top of the scaffold, but also throughout the scaffold thickness (Fig.

2A). Therefore, cells appear to have migrated by native processes into the patterned

conduits, in some cases a few millimeters in distance, over a total period of 8 days.

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Fig. 2. Cell-infiltrated fibrin scaffold. (A) After 1 day of attachment and 7 days of differentiation, C2C12 cells uniformly colonized the full thickness of a patterned fibrin scaffold as visualized with DAPI nuclear stain (blue), demonstrating compatibility between the material (green autofluorescence) and muscle cells. Cells with elongated morphology were found near the (B) top, (C) middle, and (D) bottom of the scaffold thickness. Moreover, these cells expressed acetylcholine receptors, as shown by staining with fluorescent α-bungarotoxin (red). Positive z-direction (opposite the force of gravity) shown by arrow. Scale bars represent 15μm.

Moreover, α-BTX labeling showed that cells near the top (Fig. 2B), middle (Fig.

2C), and bottom (Fig. 2D) of the scaffold cross-section expressed AChRs (red) and

featured elongated morphology, phenomena that are characteristic of differentiating

C2C12 cells. However, unlike C2C12 cells cultured on 2D fibrin analogs (see Chapter III,

Section 3.1), C2C12 cells grown on this pseudo-3D fibrin sheet did not exhibit AChR

clusters in response to exogenous soluble agrin.

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suspensions

Uni-directional perfusion of cell suspensions through patterned fibrin succeeded

in seeding cells within scaffolds, as confirmed by DAPI nuclear stain of scaffold sections.

Fig. 3 shows cell density by scaffold segment (top, middle, or bottom third), as well as

overall cell density in the section, for perfused and statically-seeded scaffolds at the

immediate time point. No difference in cell distribution was observed between static

seeding and the 1.2 mL/min flow rate. However, an increased perfusion flow rate of 4.8

mL/min increased cell colonization of the scaffold in all segments in statistically

significant fashion (see asterisks in Fig. 3).

Analysis of this metric at the 1 day growth and 1 day growth plus 2 day

differentiation time points is shown in Fig. 4. Surprisingly, the number of cells declined

significantly in the 4.8 mL/min group between the immediate time point and the 1 day + 2

day time point.


Both uni-directional and bi-directional perfusion of cell suspensions resulted in

seeding of rMDCs within fibrin scaffolds, as shown in Fig. 5. However, ANOVA indicated

no significant effect of perfusion direction (uni-directional vs. bi-directional) or conduit

diameter (12 μm vs. 41 μm) on seeded nuclear density in the middle segment of

analyzed perfused scaffold sections (Fig. 6).


The initial PVOH experimentation period determined that ideal extruder

temperatures were 162°C for the rotor and 177–181°C for the header. Low tensile force

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Fig. 3. Cell seeding of patterned fibrin immediately after perfusion. (A) Bar graphs depict cell density for individual thirds of overall scaffold length, as well as average total density over the entire scaffold, for experimental perfusion-seeded scaffolds as well as controls seeded by addition of static cell suspensions. Values are shown as average ± SD for n=9 replicates (3 separate sections from each of 3 distinct fibrin scaffolds). Counts were obtained by observation of scaffold sections labeled with DAPI nuclear stain. Oneway ANOVA revealed statistically significant differences among the 3 treatment groups at every region of the scaffold examined (p<0.0003 in all cases). Posthoc multiple pairwise comparisons indicated in all cases that perfusion of cell suspensions through scaffolds at a rate of 4.8 mL/min resulted in significantly greater engrafted cell density than either 1.2 mL/min flow rate perfusion or static seeding, which were indistinguishable. Asterisk denotes statistically significant difference from both the static seeded scaffolds as well as those seeded at the 1.2 mL/min flow rate; p<0.05, Tukey. (B) A representative DAPI-stained section of a 4.8 mL/min perfusionseeded fibrin scaffold illustrating cellular infiltration.

from the take-up system was necessary to avoid severing the fiber as it was extruded,

while high take-up speed was found to decrease fiber diameter. In addition, rotor output

was found to be positively correlated with fiber diameter. As such, extrusion was

accomplished between 10% and 20% of total rotor output, and over a range of take-up

speeds corresponding to roughly the middle third of overall capacity. Settings outside

these ranges rendered fibers produced, if any, unusable – including failure to force

polymer through the orifice, polymer decomposition by excess heat, the extrusion of

fibers of irregular diameter, or fiber breakage by excessive take-up tension.

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Fig. 4. Cell density in rMDC-seeded fibrin scaffolds over time. Shown are average cell densities within the middle third of the scaffold length, which is the most difficult to seed being farthest from the open ends of the conduit microarchitecture. 4.8 mL/min flow rate perfusion was statistically superior to static seeding and 1.2 mL/min flow rate perfusion immediately after seeding. However, cell density declined in the 4.8 mL/min group over time, and was not significantly different than alternative seeding methods at later time points. In fact, cell density significantly declined in this group after 1 day in growth medium and a further 2 days in differentiation medium. Each bar represents average ± SD for n=9 replicates.

This initial experiment did produce fibers that dissolved in water, as desired. However,

melt extrusion of PVOH was found to be very sensitive to environmental and physical

factors such as extruder temperature and ambient moisture. The resulting fibers under

these circumstances were much larger than fibers used previously - ~250 μm in

diameter. Attempts to draw them to a finer size using low rotor speeds or high take-up

speeds caused the strand to break.

In the second experiment, fibers roughly 100 μm in diameter were produced. These

fibers dissolved in water after an acetone pretreatment removed the pMMA component.

However, the resulting fibers were porous, and dissolved very rapidly in water at room

temperature. This was problematic as fibers dissolved when fibrinogen in aqueous

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Fig. 5. Fibrin scaffolds with 12 μm diameter conduits after cell seeding with a 4.8 mL/min perfusion of rMDCs. (A) Uni-directionally perfused scaffold (as shown in Fig. 3 above, for comparison). (B) Bi-directionally perfused scaffold, with no delay between change of flow direction. Cells are shown via DAPI nuclear label.

solution was added.

In the third experiment, PEO did not extrude into usable fibers using any

combination of parameters attempted.

The fourth experiment using 98% hydrolyzed PVOH yielded continuous fibers

only with the addition of pMMA. However, as in the second study, these fibers dissolved

in water too quickly after acetone pre-treatment.

In the fifth experiment, pure PVOH fibers ~80 μm in diameter (Fig. 7) were

extruded by pre-drying the polymer granules and increasing extruder temperature.

However, this diameter was 2-8 times larger than fibers used in perfusion seeding

experiments. Moreover, the resulting fibers also dissolved very rapidly in contact with

aqueous fibrinogen solution.

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Fig. 6. Effect of directionality of perfusion and conduit lumen diameter on cell seeding. As above, cell density results are shown for the middle third of the scaffold length. Contrary to expectation, no significant difference was observed between scaffolds patterned with 12 μm diameter and 41 μm diameter conduits, nor between scaffolds perfused with a suspension of cells uni-directionally or bi-directionally. Each bar represents average ± SD for n=9 replicates.

The sixth experiment employed PVOH with a mixture of 88% and 98%

hydrolyzed polymer, and resulted in small-diameter PVOH fibers in conjunction with an

improvised, faster-rotating take-up system. At 65 μm ± 5 μm in diameter, these were

almost as small as the 41 μm pMMA template fibers used in prior perfusion studies.

Moreover, these PVOH fibers were soluble in water, though only at 60°C. However,

fibers swelled larger than their original size in water even at room temperature. If these

fibers were tightly packed into a mold and used as a potential template for fibrin scaffold

patterning, the aqueous fibrinogen solution would similarly cause the fibers to swell,

taking up all empty space between fibers in the bundle and preventing full impregnation

with fibrinogen.

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Fig. 7. Example melt extruded PVOH fiber. SEM shows a water-soluble fiber roughly 80 μm in diameter, with uniform cylindrical structure.

Polymer compositions that resulted in successful fiber extrusion, and their water

solubility, are summarized in Table 1.



Fibrin scaffolds were successfully fabricated from fibrinogen containing a

suspension of microparticles and mounted in the stretch bioreactor. Though some

unavoidable macroscopic damage to the ends of the scaffold resulted from mechanical

compression in the mounting clamps, no slippage of the scaffold was noted, and no

gross defects or other macroscopic structural alterations were observed in the center of

the construct following cyclic mechanical strain (Fig. 8A).

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Table 1. Water solubility and composition of melt-extruded fibers Polymer 1 Polymer 2 Mix %

(v/v) Dissolution in H2O (at 37°C / 60°C)

PVOH, 88% hydrol. pMMA 90 / 10 No / No PVOH, 88% hydrol. --- 100 / 0 Rapid / Rapid PVOH, 88% hydrol. PVOH, 98% hydrol. 75 / 25 Slow / Rapid PVOH, 88% hydrol. PVOH, 98% hydrol. 50 / 50 No / Slow PVOH, 88% hydrol. PVOH, 98% hydrol. 25 / 75 No / Slow --- PVOH, 98% hydrol. 0 / 100 No / Slow

Moreover, an intact patterned microarchitecture was observed via

autofluorescence of the fibrin material under microscopy (Fig. 8B-C). Unfortunately, cell

seeding within the scaffold interior was greatly decreased in both density and uniformity

compared to previous results from Sections 3.2 and 3.3 of this Chapter, presumably due

to the increased length of the scaffolds. In addition, no microparticles were visible in the

scaffold interior. These results suggested that future efforts would require alternative

methods to fabricate scaffolds with immobilized microparticles for presentation of agrin

to cells. Finally, no AChR clusters were noted.


Fibrin scaffolds were successfully fabricated using a less-dense bundle of

sacrificial template fibers. The top ~10 mm of a representative scaffold, which

corresponds to the end of the fiber bundle where the particle-loaded fibrinogen solution

was added to the mold before scaffold polymerization, is shown in Fig. 9. Particles were

most dense in the top ~2 mm of the scaffold (see Fig. 9B). However, loading was

appreciable and appeared uniform in the deeper ~8mm portion of the scaffold, as shown

in Fig. 9C. Micrographs from the lower 20 mm of the scaffold's length showed a decline

in particle density (data not shown). Similar to Section 3.5, little cell seeding was

observed.

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Fig. 8. Initial perfusion-seeded, microparticle-presenting 30mm constructs. (A) The gross construct was largely intact after cell seeding, mounting in the bioreactor, and undergoing 7 days of cyclic stretch. Scaffold ends were deformed by bioreactor mounting clamps, an unavoidable consequence of exerting adequate force to keep scaffold ends in place during cyclic stretch. The damaged areas could be easily trimmed away prior to implantation. (B) A DAPI-stained section of one end of the scaffold, displaying this compression on a microscopic scale. (C) A similar section near the middle of the scaffold length. The scaffold micro- and macro-architecture were preserved over the course of the experiment, indicating compatibility of the scaffold with the bioreactor protocol. In this initial experiment, microparticle loading was low, and the longer scaffold caused a sharp decline in cell seeding of the scaffold interior.


In the first experiment, cell seeding density in the middle third of the scaffold was

highly variable, as shown in Fig. 10. However, the more compliant silicone mounting

material was a

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Fig. 9. Optimized microparticle loading of fibrin scaffold. By decreasing template fiber packing and doubling microparticle quantity suspended in fibrinogen solution before scaffold formation, microparticle density in the top 10 mm of the resulting scaffold (A) was substantially elevated. Insets are shown of the scaffold top (B), and of the uniformly seeded area (C). Though particle loading was elevated at the top, loading abruptly decreased roughly 2 mm deep into the scaffold. This indicated that trimming away and discarding the top 2 mm could produce a scaffold uniformly loaded with microparticles. Moreover, this result conclusively showed compatibility of the silica particles with the required acetone treatment. Scale bar represents 200 μm.

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Fig. 10: Initial perfusion cell seeding of 10-15 mm long fibrin scaffolds. A wide array of parameters was attempted in multiple combinations with single replicates in an attempt to rapidly identify promising conduit diameter, mounting material, flow rate, or flow direction. Identifying one or more with particular promise would make a later study, with multiple replicates, feasible. As shown, several scaffolds mounted in more compliant silicone tubing were found to be promising. Cell density was reported in the middle third of scaffold length, which is the region characteristically most difficult to robustly seed with cells.

Fig. 11: rMDC perfusion of 15 mm long CA-templated fibrin scaffolds mounted in silicone tubing. Parameters including perfusion duration, directionality of perfusion, and perfusion flow rate, were found to have no significant effect on seeded cell density in the middle third of the scaffold (n=2 for bi-directionally perfused scaffold types, n=3 for uni-directionally perfused scaffold types; p < 0.05 used to determine significance of one-way ANOVA). Data presented as average ± standard error of the mean.

137

common parameter among several of the most densely seeded scaffolds. No other

parameters were consistently observed in scaffolds densely seeded with cells. Overall,

Teflon®-mounted groups were characterized by low cellularity (~2 to ~30 cells / mm2)

compared to silicone-mounted groups (~4 to ~150 cells / mm2). Moreover, scaffolds 10

mm in length were not appreciably more densely seeded than those 15 mm in length.

Results from the second experiment are shown in Fig. 11. All experimental

groups supported cell seeding of the middle third of the scaffold to a density between 80

and 100 cells / mm2, with no significant differences reported by ANOVA.

4. Discussion


One objective of the current study was to develop a scaffold seeding

methodology applicable to the creation of a tissue engineered muscle repair construct

featuring complex 3D geometries. In a first effort toward this goal, we hypothesized that

patterned fibrin scaffolds (see Chapter II) would readily support surface attachment and

growth of C2C12 mouse myoblast cells. To evaluate this hypothesis, we fabricated thin

(~1 mm thick) fibrin sheets and seeded them by addition of a suspension of C2C12 cells.

Presumably, this would have resulted in C2C12 cells seeded exclusively on the scaffold

exterior.

However, after 8 days of culture, cells were observed not just at the top surface

of the scaffold as expected, but also within the microscopic conduits (see Fig. 2). This

result was unexpected, as the porous microarchitecture was patterned parallel to the top

surface of the scaffold, and was nominally open only on two of the four 3 mm wide x 1

mm thick (i.e. side) faces of the scaffold. Moreover, aprotinin was added to culture

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medium to prevent cells from accessing the conduit lumens by degrading the conduit

walls. Therefore, we concluded that cell infiltration resulted from cellular motility.

In addition, C2C12 cells were elongated and strongly oriented parallel to the

direction of the conduit lumens which housed them (Fig. 2B-2D), indicating that the

sacrificial patterning successfully guided muscle cell alignment. As force production by

muscle cells is critically tied to their directionality, inducing alignment of scaffold seeded

cells could accelerate tissue engineered construct processing.

These observations collectively supported our original hypothesis.


suspensions

Thus, cells could be uniformly seeded throughout a scaffold by static seeding

followed by cell migration. However, observed cell density was low. Moreover, it is likely

that cells were initially distributed at the scaffold periphery and slowly migrated inwards.

As discussed in Chapter III, denervated muscle undergoes progressive atrophy [27].

Therefore, accelerating muscle cell seeding of scaffolds is necessary.

To this end, we hypothesized that perfusing a cell suspension through the

scaffold architecture via fluid flow would increase cell seeding density and uniformity of

cell distribution within the scaffold over a short period of time. In our initial perfusion

experiment, a suspension of rMDCs in culture medium was circulated through a fibrin

scaffold using a peristaltic pump and a closed flow loop, as depicted schematically in

Fig. 1.

Unsurprisingly, the scaffold edge first contacted by the cell suspension (the

scaffold "top") was most heavily seeded with cells, regardless of the experimental

condition (Fig. 3A). Indeed, in statically-seeded control samples and scaffolds perfused

at a lower flow rate of 1.2 mL/min, the top was the only area appreciably seeded with

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cells 30 minutes after cells were first added. This was unsurprising, as the diameter of

conduits in this case (12 μm) was likely similar to that of the cells, resulting in many cells

becoming caught at the nearest edge of the scaffold. The middle third of the scaffold

was the least densely seeded area, indicating that this region is the most difficult to

seed. Because of our goal to uniformly seed scaffolds, and because quantification of cell

seeding density was a labor-intensive process, seeding in the middle third of the scaffold

was chosen as a metric for further perfusion experimentation.

4.8 mL/min perfusion significantly increased cell seeding in the scaffold interior

relative to 1.2 mL/min perfusion or static seeding. Using this treatment, cell seeding

density over the entire 15 μm thick scaffold section averaged ~500 cells / mm2,

indicating that slightly over 2 million of the total 5 million cells were captured within these

scaffolds. Qualitatively, DAPI labeling showed a relatively uniform distribution of cells

seeded within the scaffolds (Fig. 3B).

However, investigation of seeded scaffolds after further culture revealed that high

seeded cell density was not maintained over time by standard static culture in an

incubator (Fig. 4). In fact, cell density significantly declined over 1 day in growth medium

and 2 days in differentiation medium in the 4.8 mL/min perfusion group. At the final time

point, interior cell density had fallen below 80 cells / mm2 in all groups, and no significant

differences were observed. Collectively, these observations may indicate that the

scaffold represents a diffusional barrier which, combined with the significant nutrient

requirement of densely-seeded cells in 3D, leads to necrosis of cells seeded within the

scaffold interior.

These results strongly supported our hypothesis that perfusion of a cell

suspension could more densely and uniformly seed cells within a 4 mm long x 4.5 mm

diameter 3D scaffold.

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Though quantitative results from the uni-directional perfusion seeding study were

encouraging, qualitative analysis of images indicated that cell seeding across the width

of the scaffold may have been less uniform than overall cell density would indicate. In

Fig. 3B for example, longitudinal tracts of cell nuclei can be seen within certain conduits.

We reasoned that the scaffold fabrication method used may have created some conduits

that were better accessed from one end than the other, due to the inherent variability of

fabricating scaffolds by hand. We hypothesized that perfusing the cell suspension into

both open ends of the scaffold would further increase uniformity and density of cell

seeding.

Simultaneously, as viscous drag on cells was previously identified as a significant

factor, we hypothesized that perfusion seeding could be further optimized by varying

conduit diameter. To this end, experimental groups were included which featured two

different fiber template diameters of 12 μm (CA) and 41 μm (pMMA). These sizes were

the smallest and largest which resulted in a scaffold of identical bulk stiffness (see

Chapter III, Fig. 5A). We wished to stay within this range to minimize potential

confounding factors when comparing to uni-directional perfusion results.

Fig. 5 shows representative sections of scaffolds seeded via both uni-directional

(Fig. 5A) and bi-directional (Fig. 5B) perfusion. Qualitatively, the only observable

difference in cell engraftment within the scaffold was an increase of seeding at the

bottom edge, which became the first area contacted by the cell suspension after reversal

of flow direction. Like in uni-directionally perfused scaffolds, cells in bi-directionally

perfused scaffolds were still arranged into longitudinal tracts, with intervening areas of

relatively low seeding density. This allowed us to reject the first part of our hypothesis

predicting improved seeding uniformity using bi-directional perfusion.

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Further, as shown in Fig. 6, no significant difference in cell seeding density was

seen resulting from altered conduit size, direction of perfusion, or delay between reversal

of flow direction using our prior metric within the middle third of the scaffold. This allowed

us to reject the second part of our hypothesis, which predicted an increase in seeded

cell density.

This lack of observed differences was unexpected, but potentially of interest for

future work. For further experimentation, CA fibers were selected for scaffold

fabrications, as they are readily available in larger quantity than pMMA fibers. We

elected to continue varying directionality of flow in some further experiments, as it was

technically straightforward and had potential to interact unpredictably with other

parameters yet to be evaluated.


To this point, all fibrin scaffolds were templated with acetone-soluble CA or

pMMA fibers. Simultaneously, significant experimentation in agrin delivery to this point

(see Chapter III, Sections 2.2-2.3 and 3.1-3.2) utilized agrin delivery vehicles fabricated

using polystyrene (PS) microspheres, which are likewise soluble in acetone.

Incorporating agrin-presenting particles to patterned fibrin would therefore require a

change in one material.

One method to form a polymer into a usable template fiber is melt extrusion, by

which molten polymer is forced through a die of desired cross-sectional shape and then

drawn under tension into a long strand of smaller diameter. Melt extruded acetone-

soluble fibers used previously were generous gifts or created in collaboration with the

North Carolina State University College of Textiles.

By contrast, in the experiments described here, we chose to attempt melt

extrusion of the water-soluble polymers poly(vinyl alcohol) (PVOH) and poly (ethylene

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oxide) (PEO). Though a series of experiments were performed, each with its own

specific hypothesis, the overall hypothesis of the series was that melt extrusion could be

employed to produce water-soluble fibers which would in turn allow incorporation of PS

microparticles into templated fibrin scaffolds.

During prior experimentation, we found that scaffolds templated using sacrificial

fibers 105 μm in diameter or larger were difficult to suture (data not shown) and had

lower stiffness than scaffolds used for muscle cell culture previously. Thus, water-soluble

polymers would ideally be in the 12 - 41 μm diameter range, which could be used to

fabricate scaffolds of understood perfusion kinetics and preferred mechanical properties.

Through sequential experimentation, relatively uniform water soluble fibers as

small as 65 μm were produced (an example fiber 80 μm in diameter is shown in Fig. 7).

Though larger than acetone-soluble fibers used previously to template fibrin, these fibers

demonstrated that melt extrusion of PVOH was feasible using the equipment described.

However, while nominally water soluble, these fibers often swelled during degradation,

or only dissolved in water when substantially heated. When attempting to polymerize

fibrin around these templates, fiber swelling would fill free space in the mold required for

fibrinogen and defeat, to an extent, the fabrication of smaller-diameter fibers. Further, we

reasoned that elevated temperatures required to dissolve some created fibers would

quickly denature any agrin present in a combined system.

In conclusion, though our experiments resulted in water-soluble fibers, they were

unsuitable for creation scaffold templating using methodology described. We were

therefore unable to support or reject the hypothesis that water-soluble fibers could be

used in the templating of a polystyrene-containing fibrin scaffold. As such, we elected an

alternative strategy simultaneously utilizing acetone-soluble CA or pMMA fibers and

acetone-stable agrin microcarriers made from silica. Though we recognized that silica is

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not biodegradable, and therefore not an ideal biomterial for this application, it enabled

proof of concept at this early stage.



Lacking suitable water-soluble fibers, we hypothesized that acetone-insoluble

agrin microcarriers could be incorporated within fibrin scaffolds fabricated using prior

methods. If this were correct, these scaffolds could be seeded with cells by perfusion

and resulting constructs could undergo cyclic mechanical strain in a bioreactor based on

earlier sections of this Chapter and prior work [28,29], respectively. To this end, we

obtained microparticles made of SiO2, or silica, which is acetone-insoluble. These

particles were commercially available in identical sizes to PS microparticles used

previously, with the option of surface carboxyl groups which could take advantage of our

existing covalent coupling technique (see Chapter III, Section 2.1).

Covalently-bound agrin microparticles were fabricated using the zero-length

crosslinker EDC, which created a peptide bond between agrin molecules and SiO2

microparticle surfaces. Similarly to Chapter III, Section 2.5, these microparticles were

mixed into fibrinogen solutions used to fabricate fibrin scaffolds. Further, scaffold length

was increased to 30 mm and scaffold diameter was decreased to 3 mm, as this

represented a scaffold size which could be readily mounted in our available stretch

bioreactors.

30 mm long x 3 mm diameter fibrin scaffolds successfully underwent cell

seeding, cyclic stretch in the bioreactor preconditioning protocol, and histological

preparation. Imaged scaffold sections revealed intact conduit microarchitecture, which

showed for the first time that these fibrin scaffolds were mechanically compatible with

the stretch bioreactor. Though expected gross defects occurred at the scaffold ends

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resulting from compression in bioreactor mounting clamps (Fig. 8A), the fibrin material

was easily trimmed, allowing damaged ends to be removed prior to any further use.

However, few cells or microparticles were observed by microscopy within the

conduits or conduit walls, respectively (Fig. 8B-8C). As cells were observed at the

scaffold periphery, the sharp decline in interior seeding likely resulted from fluid dynamic

changes in the perfusion process brought about by use of much longer scaffolds.

Moreover, the observed lack of AChR clusters was expected due to a corresponding

lack of cells and agrin microparticles in close proximity to one another.

Post hoc calculations predicted that average conduit wall thickness of 80%

porous scaffolds fabricated using these parameters was likely less than 2 μm. Therefore,

tight packing of sacrificial fibers within the scaffold mold likely excluded the 10 μm

diameter microparticles from the scaffold interior.

Without cells and microcarriers in proximity, the bioactivity of acetone-treated

agrin microcarriers could not be verified. Therefore, the hypothesis could not be

supported nor rejected in this experiment. Further experiments therefore refined the

incorporation of agrin presentation and 3D cell seeding portions of the hypothesis.


As before, the goal of this experiment was to fabricate a patterned fibrin scaffold

containing microparticles made from SiO2. However, we hypothesized for the current

experiment that microparticle loading within the scaffold interior could be increased by

packing template fibers less densely within the Teflon® mold before the addition of

fibrinogen, as discussed in Section 4.5. Again, a cell perfusion step was included for the

dual purposes of enabling cell seeding and to evaluate whether any successfully

immobilized microparticles would be washed out of the scaffold under fluid flow. By

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contrast, we simplified the experiment by removing the linkage of agrin to microparticles

and the bioreactor preconditioning of scaffolds.

Though the resulting scaffolds were still poorly seeded with rMDCs using the

perfusion methods specified, they featured three distinct areas of microparticle density:

the top ~2mm, which was heavily laden with particles (Fig. 9B); the ~8 mm below that,

which featured a relatively uniform and substantial density of embedded microparticles

(Fig. 9C); and the bottom ~20 mm, characterized by a decline in immobilized

microparticle density (not shown). This result supported the hypothesis that less

densely-packed fibers would allow elevated particle loading within the scaffold. Further,

it showed not only effective particle loading in the fabrication step and stability of silica

particles in acetone, but also that fluid flow through the scaffold did not displace

immobilized particles.

However, the preference for uniform microparticle loading recommended that the

top and bottom portions of the scaffold not be used in the final construct. As mounting

into the cyclic stretch bioreactor necessitated loss of scaffold ends, this was an

acceptable compromise. In addition, decreased fiber packing directly resulted in a less

porous scaffold, in turn reducing the potential 3D seeding area for cells. This

represented a necessary compromise to enable fabrication of an agrin-presenting

scaffold.

The decline in microparticle density in the ~20 mm of the scaffold below the area

shown in Fig. 9, along with difficulty in cell seeding of longer constructs, recommended

use of shorter constructs. However, scaffolds longer than the 4 mm previously perfused

were required for further experimentation due to loss of construct ends when clamped

into cyclic stretch bioreactors.

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As a compromise between these size requirements and the constraints of

perfusion - chiefly poor cell density in longer perfusion-seeded scaffolds as stated, but

also the number of rMDCs required to seed longer scaffolds - 3 mm diameter cylindrical

scaffolds either 10 mm or 15 mm in length were selected for further experimentation.

This represented a total increase in scaffold volume of only 11% or 67%, respectively,

relative to 4 mm long x 4.5 mm diameter scaffolds previously seeded successfully by

perfusion, while increasing the length of the scaffold roughly two- to four-fold and

thereby reaching a size more relevant for future studies.

The additional inclusion of alternative scaffold mounting materials presented a

large array of potential combinations of experimental variables in concert with the new,

shorter scaffold length. To evaluate some of the most promising combinations as rapidly

as possible, we elected a two-phase experimental design. First, we evaluated common

parameters amongst densely cell-seeded scaffolds using groups with single replicates.

We then performed a second experiment with more replicates in fewer groups featuring

the common parameters. We hypothesized for this two-part experiment that 3 mm wide

x 10-15 mm long scaffolds could be seeded to similarly high interior cell densities as 4.5

mm wide x 4 mm long scaffolds.

Use of a more compliant silicone scaffold mounting material was suggested to be

of benefit by the small initial experiment. However, the follow-on experiment identified no

statistically significant differences between any groups evaluated. Moreover, seeded cell

densities in these scaffolds were ~75-95 cells / mm2, substantially lower than the ~250

cells / mm2 seen previously in 4.5 mm diameter x 4 mm long scaffolds (Fig. 3A).

Assuming similar distribution of cells throughout the entire scaffold, this indicated that 15

mm long x 3 mm diameter scaffolds captured 1 million to 1.25 million of the 5 million

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cells in suspension, while 4 mm long x 4.5 mm diameter scaffolds captured over 2

million.

Therefore, it was concluded that the decreased cell density observed was not

simply a function of larger available surface area for cell seeding within the scaffold

microarchitecture, but instead a proportionally reduced efficiency in seeding longer,

narrower scaffolds. As such, the hypothesis in this experiment was rejected.

As the only parameter which could not be varied, we now hypothesize that the

scaffolds fabricated by hand using our current methodology possess interior architecture

that, while apparently uniform under light and electron microscopy as shown in the

original proposal, features subtle inconsistencies which undermine repeatability in

perfusion outcomes and which compound as the scaffold length increases.

Given these findings, we reevaluated advances to date and compared to the

remaining goal of evaluating cell-seeded agrin-presenting fibrin materials in vivo. In

Chapter III, we observed AChR clustering behavior in C2C12 cells resident on a 2D

fibrin layer featuring embedded agrin-presenting particles. Moreover, we demonstrated

that 3D fibrin scaffolds could incorporate immobilized agrin microcarriers. rMPCs

surface-seeded onto 3D scaffolds also exhibited extensive AChR clustering following

cyclic stretch.

Considering persistent difficulties reliably seeding cylindrical scaffold interiors via

perfusion of cell suspensions, we elected to evaluate planar, surface-seeded scaffolds

(as described in Chapter III) in vivo using a rat neurotization model. This model and the

results thereof are discussed in Chapter V.

5. Conclusions

In this Chapter, we describe efforts to translate promising fibrin biomaterials and

agrin pharmacologics from 2D forms, as successfully demonstrated in the first portion of

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Chapter III, to 3D embodiments. These efforts were primarily twofold: first, to create a 3D

fibrin construct with patterned pores for muscle cell attachment and guidance which

presented an immobilized agrin signal; second, to densely seed the interior of such a

scaffold with muscle cells. The overall objective of this study was to create a highly

cellularized tissue engineered muscle repair construct which would be suitable for

implantation in vivo.

Toward this end, we found that fibrin biomaterials supported the attachment and

growth of both a C2C12 muscle cell line and expanded rMDCs. Moreover, agrin-

presenting microparticles were successfully embedded into fibrin in a manner compatible

with sacrificial templating methods.

Unfortunately, attempts to robustly seed the interior of such scaffolds only met

with success in scaffolds which were too short to feasibly use in conjunction with

mechanical preconditioning in a bioreactor. However, future efforts may yield improved

results, either through refinements of techniques described here, or through alternatives

such as assembly of pre-seeded scaffold components, enhancement of native cellular

motility into the scaffold, or other approaches not yet envisioned.

Acknowledgements




gratefully acknowledge the assistance of Hannah Baker, Christopher Bergman, Hayden

Holbrook, Venu Kesireddy, Juliana Passipieri, and Mevan Siriwardane in performing

experiments described herein, as well as Cathy Mathis and Cynthia Zimmerman for their

assistance with histology.

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CHAPTER V

PRELIMINARY EVALUATION OF INNERVATION AND REMODELING OF A TISSUE

ENGINEERED SKELETAL MUSCLE REPAIR CONSTRUCT IN VIVO

John B. Scott

153

1. Introduction

Chapters II through IV describe sequential advances in the development and

evaluation in vitro of a biomimetic construct aimed at tissue engineered repair of

volumetric muscle loss (VML) injuries. Results showed that the construct could direct

neuronal process ingrowth, support muscle cell attachment, and incorporate

pharmacologic (immobilized neural agrin) and mechanical (bioreactor preconditioning)

cues designed to elicit acetylcholine receptor (AChR) clusters, the presumptive in vitro

analog of motor end plates (MEPs). This Chapter describes the implantation of these

constructs in vivo using a model of innervation, a necessary process in the healing of

large muscle injuries.

This study utilized a rat neurotization model wherein the cell-seeded scaffolds

were implanted in the subcutaneous space of a rat hindlimb and then sutured to the

proximal stump of the transected femoral nerve. Based on results from a study in the

literature [1] and our previous evaluation of neurite in-growth into the scaffold in vitro

(see Chapter II), axons were expected to grow out from the proximal femoral stump onto

the constructs.

Prior to implantation, experimental constructs were formed by mechanically

preconditioning surface rMDC-seeded, agrin-presenting planar fibrin scaffolds in vitro.

This embodiment was previously shown to exhibit substantial AChR clustering (see

Chapter III). We hypothesized first that neo-innervation of constructs would result in de

novo NMJ formation between the host nervous system and the tissue engineered

muscle construct, and second that AChR clusters formed prior to implantation could

enhance either these NMJs or ectopic muscle resulting from host remodeling of the

implant.

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2. Methods

Unless otherwise noted, all culture medium formulations are described as % v/v

and all histological staining reagents are expressed as % w/v in deionized (DI) water.

Further, all methods were performed at ambient pressure on a lab bench at a room

temperature of ~22°C unless otherwise noted.

2.1. Culture medium formulations and isolation of rat muscle derived cells

Culture medium formulations for myogenic medium and rat muscle derived cell

(rMDC) growth medium were identical to those used in Chapter III, Section 2.1. Briefly,

these were myogenic medium: 68% high-glucose DMEM, 20% FBS, 10% horse serum,

1% chicken embryo extract, and 1% antibiotic-antimycotic; rMDC growth medium: 84%

low-glucose DMEM, 15% FBS, and 1% antibiotic/antimycotic.

rMDCs used in this Chapter were isolated and expanded using identical methods

to those described in Chapter III, Section 2.4. Briefly, Matrigel®-coated tissue culture

plates were prepared prior to isolation. Meanwhile, tibialis anterior and soleus muscles

were removed from euthanized donor rats, rinsed in 10% povidone iodine solution, and

transferred to a sterile environment where they were subsequently washed in PBS,

manually minced to a suspension of fine particles in collagenase-supplemented culture

medium, and incubated at 37°C. After 2 hours, digested tissue suspensions were

centrifuged, the supernatant was removed, and tissue was resuspended in myogenic

medium and added to Matrigel®-coated plates.

After 2 days, myogenic medium was removed and replaced with rMDC growth

medium. After a further 2 days, cells were dissociated from plates and then replated on

untreated tissue culture polystyrene in rMDC growth medium. At ~70% confluence,

usually after 3-4 days, cells were again dissociated from plates and used for

experimentation as described below.

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This isolation protocol was approved by the Wake Forest University Institutional

Animal Care and Use Committee and complies with animal use guidelines established

by the American Physiological Society.

2.2. Preparation, cell seeding, and bioreactor preconditioning of agrin-presenting fibrin

scaffolds

3D patterned fibrin scaffolds embedded with covalently-bound agrin microcarriers

were fabricated, surface seeded with rMDCs, and preconditioned by cyclic mechanical

strain in a bioreactor as described in Chapter III, Section 2.5.

Briefly, the zero-length crosslinker N-(3-Dimethylaminopropyl)-N′-

ethylcarbodiimide hydrochloride (EDC, Sigma-Aldrich, St. Louis, MO) was used to

covalently bond either recombinant rat agrin (R&D Systems, Minneapolis, MN) or bovine

serum albumin (BSA, Sigma-Aldrich) molecules to 10 μm diameter SiO2 microspheres

(Corpuscular, Cold Spring, NY). Subsequently, cellulose acetate (CA) fibers (generous

donation of Eastman Chemical Company, Kingsport, TN) ~12 μm in diameter were

arranged into an aligned mat. Protein-bound microcarriers were suspended in fibrinogen

(Sigma-Aldrich) and this suspension was added to the fiber mat. Fibrinogen was

polymerized to fibrin in the presence of thrombin and CaCl2 in PBS, creating a large

fibrin scaffold which was then trimmed into individual smaller scaffolds measuring

roughly 30 mm long x 3 mm wide x 1 mm thick. These were then sequentially washed

four times each in acetone and then sterile PBS to yield scaffolds suitable for cell

seeding.

One 30mm long x 3 mm wide flat plane of each scaffold was then seeded with

cells by manually pipetting a suspension of rMDCs in growth medium over scaffold

surfaces. After allowing a brief initial period for cell attachment, growth medium was

added to totally immerse cell-seeded scaffold constructs.

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Constructs were removed from static culture after allowing 16-24 hours for full

cell attachment and clamped into the mounts of a custom-fabricated bioreactor designed

to repeatedly apply uniaxial cyclic stretch to growth medium-immersed tissue constructs.

The bioreactor was used to precondition cell-seeded scaffolds by subjecting them to

10% mechanical strain parallel to the long axis of scaffolds, 3 times per minute for the

first 5 minutes of every hour over a total time of 5 days. In all groups, rMDC growth

medium was exchanged every 2-3 days, and was supplemented with aprotinin starting

on day 1. After completion of the 5 day preconditioning step, scaffolds were removed

from the bioreactor. Scaffold ends deformed by mounting in the bioreactor were

removed with sterile surgical scissors, and scaffolds were immediately implanted in vivo

(see Section 2.3).

2.3. Implantation of tissue engineered muscle repair constructs in vivo

After predoncitioning, constructs were implanted in vivo to histologically evaluate

structural and functional innervation by the host. The model is represented schematically

in Fig. 1.

The following experimental groups were included in this experiment. All groups

featured bioreactor-preconditioned fibrin constructs evaluated at a single 2 week time

point using histology (see Section 2.4 below).

• Covalently-bound agrin microcarriers, surface-seeded rMDCs (n=3)

• Covalently-bound BSA microcarriers, surface-seeded rMDCs (n=3)

• Covalently-bound agrin microcarriers, no rMDCs (n=3)

To begin, 12-16 week old male Lewis rats (Charles River, Raleigh, NC) were

placed under general anesthesia by isoflurane inhalation. The surgical site at the medial

left hindlimb was shaved and sterilized using sequential scrubs of povidone-iodine

(McKesson Corporation, San Francisco, CA) and ethanol, and the site was covered in a

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Fig. 1: Schematic overview of rat neurotization study. (A) A longitudinal incision was made in the rat hindlimb to expose the femoral nerve (thin white structure between black arrowheads). (B) The femoral nerve was transected, and a bioreactor-preconditioned fibrin construct (pink structure between white brackets) was placed in contact with the proximal nerve stump. Suture loops (black arrowheads) were anchored in the underlying fascia and used to immobilize the construct in place, as well as to maintain contact between nerve and construct. (C) At a two week time point, the surgical site was reopened. Some construct integration with and remodeling by adjacent native tissue was observed, but constructs were still grossly visible (pale structure between white brackets) at this early time point. (D) Half of all animals were subjected to electrical stimulation of the femoral nerve (black arrowhead) proximal to the construct, which would deplete glycogen reserves of any newly innervated muscle in the construct (white bracket). Constructs and the adjacent femoral nerve were then explanted for histological analysis. (E) Sections for histology were parallel to the long axis of the construct with the aim to capture a plane including both the remodeled construct and the underlying nerve. (F) Visual observation of H&E staining was used to identify sections including both construct and nerve. Ideally, the nerve would have robustly innervated the construct area. Unfortunately, in most samples the nerve (gray) was disconnected from the remaining scaffold (orange), though in some samples it interfaced with the remodeled tissue (purple) surrounding the remaining scaffold. To best represent results and provide landmarks for reference, subsequent imaging (dashed box) was performed at the remaining scaffold nearest the nerve.

158

sterile drape. A longitudinal incision was made in the skin to expose the femoral nerve

(Fig. 1A).

One fibrin construct per animal was removed from the bioreactor, trimmed (as

described in Section 2.2), and immediately immobilized subcutaneously by anchoring

sutures into the nearby underlying muscle fascia and then looping these sutures around

the construct. The femoral nerve was then dissected away from the femoral vein and

artery and transected. The proximal stump of the femoral nerve was immobilized against

the scaffold surface by placing the stump within the proximal suture loop holding the

construct. The anchoring suture loops were then tightened and knotted securely in place

(Fig. 1B). The surgical site was sutured closed, and the animals were allowed to recover.

At the 2 week post-implantation time point, animals were again placed under

general anesthesia and the implant site was re-exposed (Fig. 1C) using similar methods.

To assess functional formation of neuromuscular junctions (NMJs) between host nerve

axons and engineered muscle constructs, half of all implanted constructs were

electrically stimulated by placing needle electrodes across the femoral nerve proximal to

its interface with the construct (Fig. 1D), subjecting the nerve to 100ms trains with 0.1ms

pulses at 100Hz every 0.5 seconds over a period of 15 minutes, and then explanting the

construct and attached nerve stump. This approach was previously shown to deplete

glycogen stores of innervated muscle [2], but would have no effect on non-innervated

muscle. Comparison of histological labeling of glycogen between stimulated and

unstimulated samples, then, could be used to assess functional innervation (see Section

2.9). The remaining half of implanted constructs were explanted immediately after

exposing the implant site, receiving no electrical stimulation. Animals were then

humanely euthanized.

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2.4. Histological preparation of samples

All explanted constructs were prepared for histology identically to 3D cell-seeded

scaffolds previously evaluated in vitro (see Chapter III, Section 2.6). Briefly, immediately

after being explanted, samples were fixed overnight in 4% paraformaldehyde in PBS at

4°C before being paraffin processed, longitudinally sectioned to 15μm thickness (Fig.

1E), mounted to glass slides, dried at least 48 hours at 60°C, and stored indefinitely

before use.

Slides were deparaffinized and hydrated to water immediately before staining.

Histological staining included hematoxylin and eosin (H&E) to allow evaluation of

morphology of tissues and constructs, Holmes silver nitrate to label axons, Masson's

trichrome to label muscle and collagen, and periodic acid-Schiff (PAS) stain to label

glycogen. Unless otherwise noted, all histology reagents were sourced from Sigma-

Aldrich.

2.5. Hematoxylin and eosin staining and location of nerve in explants

H&E staining was performed first on every 13th sequential tissue section. After

being deparaffinized and rehydrated, sections were stained using the following steps: 1)

Immersed in Gill's hematoxylin (PolyScientific R&D Corp., Bay Shore, NY), 2 minutes; 2)

Washed twice in DI water, 2.5 minutes and 2 minutes, respectively; 3) Washed in Scott's

tap water (Leica Microsystems, Buffalo Grove, IL), 1 minute; 4) Immersed in 95%

ethanol, 1 minute; 5) Immersed in eosin; 20 seconds; 6) Dehydrated in two changes of

95% ethanol and three changes of 100% ethanol; 7) Cleared in two changes of xylenes;

8) Mounted with MM 24 mounting medium (Leica); 9) Allowed to dry.

H&E stained sections were then imaged to locate areas from each sample, if

any, where the femoral nerve stump remained attached to the construct at the end point

(see Fig. 1F). These identified areas (which numbered n=3 for the agrin+rMDCs group,

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n=2 for the BSA+rMDCs group, and n=2 for the agrin+acellular group) were selected,

and where necessary, an area containing some of the remaining scaffold material was

selected from the remaining replicate.

Sequential sections near the areas selected using H&E were stained using the

other three labels to histologically evaluate n=3 from each group. Constructs could

become detached from the nerve at many points during the experiment, including by

surgeon error at implantation or explantation, by animal activity during the implantation

period, or by growth or infiltration of tissue between the nerve and the construct.

Therefore, it was necessary to evaluate this attachment and the experimental endpoint

after all sample processing was complete.

2.6. Holmes silver nitrate label for axons

Hydrated slides with 3 sections each were stained using the Holmes silver nitrate

label using the following steps: 1) Placed in 20% silver nitrate in the dark, 1 hour; 2)

Washed in 3 changes of DI water, 10 minutes each; 3) Placed in an impregnating

solution (0.11% boric acid, 0.075% sodium tetraborate decahydrate, 0.0017% silver

nitrate, and 0.083% pyridine) and incubated in the dark at 37ºC, overnight; 4) Removed

from solution and carefully drained; 5) Placed in a reducing solution (1% hydroquinone

and 10% sodium sulfite), 90 seconds; 6) Washed in running water, 3 minutes; 7) Rinsed

briefly in DI water; 8) Toned in 0.2% gold chloride, 3 minutes; 9) Quickly rinsed in DI

water; 10) Placed in 2% oxalic acid, 3 to 10 minutes; 11) Observed until axons began

turning blue-black; 12) Immediately placed into DI water to halt the reaction; 13) Further

rinsed in several changes of DI water; 14) Placed in 5% sodium thiosulfate, 5 minutes;

15) Washed in gently running tap water, 5 minutes.

After labeling all 3 sections with Holmes, PAS and Masson's staining were each

performed on one section of each slide.

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2.7. Periodic acid - Schiff stain for glycogen

PAS staining was performed on one section of each slide already stained using

the Holmes method. Sections were first oxidized in 0.5% periodic acid solution for 5

minutes and then rinsed briefly in DI water. Sections were then placed in fresh Schiff

reagent (Leica) for 15 minutes before being washed in lukewarm tap water for 5 minutes.

Sections were counterstained with Gill’s hematoxylin as in H&E staining for 1 minute and

washed in running tap water for 5 minutes.

2.8. Masson's trichrome stain for collagen and muscle

Masson's trichrome stain was likewise applied after Holmes, on a different

section than that stained via PAS. Sections were stained by using the following steps: 1)

Immersed in Gill's hematoxylin, 2 minutes; 2) Rinsed in running tap water, 10 minutes; 3)

Quickly rinsed in 2-3 changes of DI water; 4) Stained briefly in Biebrich scarlet-acid

fuchsin solution (90 parts 1% Biebrich scarlet solution, 10 parts 1% acid fuchsin solution,

and 1 part glacial acetic acid); 5) Rinsed in 3-4 changes of DI water; 6) Differentiated in

a solution (2.5% phosphomolybdic acid and 2.5% phosphotungstic acid) until red dye

was removed from collagen (usually 5-15 minutes); 7) Drained; 8) Transferred briefly

into aniline blue solution (2.45% aniline blue in a mixture of 2 parts glacial acetic acid

and 100 parts DI water); 9) Rinsed briefly in 4-6 changes of DI water to remove excess

stain; 10) Differentiated in 1% acetic acid solution, 1-2 minutes.

After all three stains were completed, slides were dehydrated in two changes

each of 95% ethanol and 100% ethanol, cleared in xylenes, mounted with MM 24, and

allowed to dry. This left each slide with one sectioned labeled using Holmes alone, a

second section co-labeled with Holmes and PAS, and the third section co-labeled with

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Holmes and Masson's trichrome. The sequential nature of scaffold sections allowed

comparison of the same observed structures using different labels.

2.9. Imaging and annotations

Brightfield images were obtained by using a DM4000 B microscope (Leica) with

attached Retiga 2000RV camera (QImaging, Surrey, BC, Canada), coupled with

ImagePro 6.2 software (Media Cybernetics Inc., Bethesda, MD). Images were cropped

and supplemented with scale bars and other annotations in Powerpoint (Microsoft,

Redmond, WA). Labels were added by investigators to aid readers in identifying the

fibrin scaffold and surrounding structures in presented images.

Images were analyzed for the presence of co-localized areas of Holmes and

Masson's staining (Fig. 2), which would indicate potential neuromuscular connections

between host axons and implanted constructs. Further, PAS-stained samples of

stimulated constructs were compared to those of unstimulated constructs within the

same experimental group (Fig. 3) to discern presence of functional innervation by the

femoral nerve.

3. Results

As indicated by gross examination of the exposed surgical site (representative

image in Fig. 1C), all samples histologically featured regions where the fibrin scaffold

was still largely intact (regions marked “Construct” in Fig. 2). All samples likewise

featured regions surrounding the scaffold where the host tissue had integrated with and

begun to remodel the implanted construct (regions marked “Native” in Fig. 2). Holmes

staining indicated that host axons may have innervated the construct surface (red

arrowheads in Fig. 2B, 2E, and 2H). In cell-seeded constructs, the integrated region

featured tissues which stained red via Masson’s trichrome, indicating potential immature

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muscle (red arrowheads in Fig. 2C and 2F). These structures were observed much less

frequently in explants which were not seeded with MDCs prior to bioreactor conditioning

and implantation (see example in Fig. 2I).

No evidence of striations or cellular / structural alignment was seen in or amongst

the red-stained muscle areas in either the agrin-presenting or control BSA-presenting

samples. This suggested a lack of mature muscle phenotype regardless of treatment,

which was not unexpected at this early time point. However, all explants were

characterized by substantial infiltration of presumptive inflammatory cells (see multiple

round nuclei throughout areas of Fig. 2, especially in the integrated region labeled

“Native”), indicating that an aspect of the construct - likely the non-degradable silica

particles, which were present in all scaffolds - elicited an immune response. Significant

blue labeling was further observed in the partially-remodeled areas of trichrome-stained

sections (Fig. 2C, 2F, and 2I), indicative of collagen deposition / fibrosis.

PAS staining indicated no differences in glycogen content between stimulated

and unstimulated samples in any experimental group (compare Fig. 3A to Fig. 3B-3C,

Fig. 3D to Fig. 3E-3F, and Fig. 3G-3H to Fig. 3I). This suggested that functionally

innervated, contractile muscle did not exist in the explant.

One interesting result was seen in PAS-stained samples. Constructs which had

been cell-seeded in vitro were found to have substantial (~800 μm or greater) native

tissue ingrowth between the nerve stump and the remaining fibrin material. By contrast,

two out of three non-cell-seeded constructs featured remaining construct material within

~200 μm of the nerve. Using the nerve as a landmark, this result suggests that surface-

seeding constructs with MDCs may accelerate degradation of the scaffold.

Finally, one unexpected phenomenon was observed near sutures in some samples. In

these, the proximal femoral nerve stump was associated very closely with the suture

used to anchor it in place (Fig. 4). As shown by Holmes staining in one sample, axons

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Fig. 2: Overview of host response to construct implantation. Shown are representative images from bioreactor-preconditioned constructs after explant in each experimental group: cell-seeded agrin-presenting constructs (A-C), cell-seeded BSA-presenting constructs (D-F), and agrin-presenting constructs not pre-seeded with cells (G-I). Remaining construct and areas remodeled by native tissue ingrowth within the samples have been labeled appropriately to aid visualization. In each sub-image, the nearby nerve is beyond the bottom of the field of view (see Fig. 1F). Each sample is presented with three different labels on near-sequential sections to mirror structures as closely as possible. Hematoxylin and eosin (A, D, G) were used to label overall morphology of cells (pink) and nuclei (purple), Holmes silver nitrate (B, E, H) was used to label axons (red-black), and Masson’s trichrome (C, F, I) was used to label collagen (blue) and muscle (red). Note that trichrome sections were also labeled with Holmes, that fibrin in the remaining construct area non-specifically stains with eosin, Holmes, and trichrome, and that some nuclei – notably those of presumptive inflammatory cells – also non-specifically label with Holmes (round red structures in middle column). Constructs elicited a substantial inflammatory response, as seen by significant nuclear infiltration (A-B, D-E, G-H), especially in the native-remodeled area. Though some axonal ingrowth (long dark structures at the native/construct interface, red arrowheads in B, E, H) may have occurred, and some muscle tissue may have formed in the native-remodeled area of cell-seeded constructs (red-stained tissue, red arrowheads in C and F, but not observed in I), these areas also featured significant collagen deposition (blue-stained tissue in C, F, I) and a lack of tissue organization and alignment or striated muscle fibers. Taken in combination, these factors strongly suggest a lack of mature muscle tissue in all experimental groups. Scale bar represents 50 μm.

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extended from the outside of the explant toward the suture, and then abruptly stopped

before reaching the construct area (Fig. 4B). Though this was only directly observed in

one sample, the close association between nerve and suture was observed more

frequently.

4. Discussion

4.1. Overview

To evaluate the hypothesis in this study, we employed numerous histological

markers which would allow comparison of tissue morphology, co-localization of nerve

axons and muscle cells, as well as formation of functional neuromuscular connections

amongst treatment groups. It was not necessary for this purpose that host nerves

specifically innervate AChR clusters which had developed in vitro prior to implantation of

the construct. Successful proof-of-concept at this early time point would justify further

refinement of 3D seeding techniques and development of biodegradable agrin particles

that, in concert, would enable future evaluation of this technology over longer time

periods and perhaps in models of VML.

4.2. Histological characterization of explants

At the 2-week end point, implants had partially - but not fully - integrated with the

surrounding tissue (compare freshly implanted construct in Fig. 1B to re-opened implant

site in Fig. 1C), a result which was expected at this early time point. This was confirmed

via histology, as remaining fibrin material with relatively intact patterned architecture was

observed in many tissue sections (see right side of sub-images in Figs. 2 & 3).

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Fig. 3: Glycogen content of explants in the construct area. Shown are three samples from each experimental group, stained with periodic acid-Schiff to label glycogen (pink / light purple) and counterstained with hematoxylin to label nuclei (dark purple / black). Electrical stimulation applied to the femoral nerve, as performed in half the samples here, would have depleted innervated muscle of glycogen content. As in Fig. 2, remaining construct and areas remodeled by native tissue ingrowth within the samples have been labeled (all samples). In most samples (A-F, I), the nearby nerve is beyond the bottom of the field of view. However, in two samples that lacked cellular seeding in vitro before implantation, the remaining construct area was not substantially displaced from the femoral nerve stump (see nearby areas labeled “Construct” and “Nerve” in G-H). In each experimental group, at least one electrically unstimulated (A, D, G-H) and one electrically stimulated (B-C, E-F, I) sample are presented. Note that samples were also labeled with Holmes (see Fig. 2) and that remaining fibrin non-specifically labels with PAS (aligned purple structures to the right of each image). Despite the potential presence of nerve and muscle at the interface between construct and native tissue (see Fig. 2), no differences in glycogen content were observed in this region between stimulated and unstimulated samples in any group. This suggested a lack of functional connections between the innervating femoral nerve and any contractile muscle tissue in the implant, though this was likely due to a lack of mature muscle in any explant group. The presence of remaining fibrin material in proximity to the femoral nerve stump in some non-cell-seeded constructs may indicate that cell seeding enhances host remodeling of the scaffold. Scale bar represents 50 μm.

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Fig. 4: Potential interaction of immobilized femoral nerve stump with suture. Shown is a sample wherein the suture loop anchoring the nerve to the construct may have had a deleterious effect on innervation of the implant. As in Fig. 2, three different labels – hematoxylin and eosin (A), Holmes silver nitrate (B), and Masson’s trichrome (C) – were performed on sequential sections to mirror structures as closely as possible. In the H&E label (A), the femoral nerve stump can be seen in close proximity to both the anchoring suture and the area surrounding the construct. In the Holmes label (B), axons, visible as elongated dark structures highlighted with black arrowheads, can be seen extending from the outside of the explant (top left) around the suture and abruptly ceasing (red arrowhead) on the side of the suture loop facing the construct. They failed to successfully innervate the construct, instead leaving an intervening space (asterisks). This may have resulted from undue compression on the nerve by tying the anchoring suture too tightly, which could have prevented axon extension. Alternatively, this may indicate that the convex shape of the suture fibers served as a topographical guidance cue for axon extension, and that the axons preferentially grew into the anchoring suture instead of along the implanted construct exterior. Though this phenomenon was not often observed, it may inform further studies involving surgical coaptation of nerves to biomaterials or the use of patterned biomaterials for peripheral nerve repair. Scale bar represents 50 μm.

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Holmes labeling showed numerous axons within the remodeled area, including

some near the surface of remaining fibrin material (see middle column of sub-images,

Fig. 2), in accordance with expectations. Trichrome staining revealed potential muscle

tissue within samples which had been pre-seeded with rMDCs before implantation, often

in areas near Holmes-labeled axons (compare middle and right columns of sub-images,

Fig. 2). Though no striations or other morphological characteristics of mature muscle

fibers were observed within these areas, this was not surprising at such an early time

point. Silica microparticles, though difficult to observe reliably due to their optical

transparency and similarity in size to cells and/or nuclei, were present throughout the

remaining scaffold material.

4.3. Presumptive inflammatory and fibrotic responses

Unfortunately, significant nuclear infiltration was observed by the counterstain of

many labels (Fig. 2). This suggested that one or more aspects of the implant - such as

the rigid, non-biodegradable silica microparticles - induced an inflammatory response

from the host. The magnitude of this response may have created a challenging

environment for de novo muscle formation at this early time point.

Tissue surrounding scaffolds in the implanted area also stained substantially for

collagen (blue label in right column of sub-images, Fig. 2). Any muscle tissue within this

area, at this or any subsequent time point, would likely produce little useful motion due to

the relative stiffness of the surrounding collagen network, even if successfully innervated

by the observed axons to form a functional motor unit.

4.4. Evaluation of functional motor units by glycogen depletion

To evaluate the potential presence of functional motor units despite the

unfavorable implant environment, we imaged PAS staining in electrically stimulated and

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unstimulated samples to compare relative levels of glycogen in and around the construct

(Fig. 3). Though Holmes and Masson's labeling suggested the possibility for connections

between the innervating host nerve and implanted muscle cells, glycogen content of

explants was qualitatively similar among all samples. The stimulation protocol used

would have thoroughly exhausted the glycogen supply of any innervated contractile

fibers.Therefore, we concluded that either the innervating axons we observed had not

assembled functional synapses with construct muscle cells, or more likely, that any

muscle tissue that was present was not sufficiently mature to undergo contraction.

Unfortunately, the morphology of the explant sections prevented unequivocal

determination of the precise mechanisms responsible. In any event, the utility of agrin-

presenting constructs was not supported by this model system at this time point.

4.5. Potential nerve interaction with suture

During image analysis, we noticed an unexpected phenomenon - axons growing

out of the femoral nerve stump in some samples remained in the area of the nearby

suture instead of colonizing the construct (Fig. 4). The exact mechanism impeding axon

outgrowth was not clear, but could be due to nerve compression by tying the suture loop

too tightly during surgical implantation of the construct. Alternatively, nerve axons have

been previously shown to preferentially track along small-diameter fibers [3], and the

presence of multifilament suture so close to a re-growing nerve may have presented a

more competitive guidance cue to axons than did the construct. Regardless, this result

suggests that future studies may require an alternative method of construct

immobilization, perhaps by microsurgically anchoring monofilament suture between the

end of the nerve stump and the construct itself.

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4.6. Fate of construct AChR clusters formed prior to implantation, evaluation of

hypothesis, and prospectives

In light of the observed histological characteristics and lack of functional

innervation - i.e., the lack of detectable differences in PAS staining between samples

receiving nerve stimulation in vivo and those that did not - the presence of significant

AChR clusters within the explants seemed improbable and was not examined herein. As

such, the maintenance in vivo post-implantation of the intricately clustered AChR

structures observed in vitro (see Chapter III, Fig. 3 & 4) remains to be determined.

Ultimately, pre-seeding of scaffolds with cells prior to bioreactor preconditioning was

shown to be important by the presence of muscle tissue in cellularized constructs (Fig.

2C & 2F), but not in acellular controls (Fig. 2I). Conversely, no differences were

observed between scaffolds fabricated with agrin-presenting microparticles (Fig. 2A-2C)

and those with control BSA-presenting particles (Fig. 2D-2F). Results, then, only partially

supported our original hypothesis – namely, that exogenous agrin presentation could

enhance either NMJs or ectopic muscle resulting from host remodeling of the implanted

tissue engineered muscle construct.

Despite the challenges ahead, these valuable preliminary data do indicate that

future pursuit of this promising technology may require strategies to reduce inflammation

likely resulting from non-degradable silica microparticles. Analysis of later time points

would also be informative regarding the duration and magnitude of the inflammatory

response to the implants fabricated using these methods. Such future studies might

utilize treatments of the fibrin biomaterial and / or development of less rigid,

biodegradable agrin-delivering microparticles designed to reduce immunogenicity of the

combined biomaterial, and thus, promote more functional remodeling of the implanted

construct.

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5. Conclusions

We previously described a biomimetic tissue engineering approach which

combined a pharmacologic cue (agrin delivery) with mechanical cues (patterned fibrin

and uniaxial strain). In vitro, substantial AChR clustering was observed in muscle cells

cultured using this approach over prolonged time points. The objective of the current

study was to further demonstrate the potential of this combinatorial technology using a

model of muscle innervation in vivo, a critical process in the healing of skeletal muscle

injuries.

Unfortunately, the results of this study were not as encouraging as previous work

in vitro, as no indication of functional connections between new muscle, either delivered

by the implant or induced from the surrounding tissue, and host nerve could be found.

Yet histological evidence of innervation of the partially remodeled construct and of

potential muscle tissue in the surrounding area were observed in constructs seeded with

cells prior to implantation, suggesting that this technology merits further investigation.

The data from the proof-of-concept in vivo study presented here will provide valuable

groundwork for studies to come, and may in the future contribute to repair of devastating

volumetric muscle loss injuries by a tissue engineering approach.

Acknowledgements




gratefully acknowledge the assistance of Hannah Baker and Juliana Passipieri in

performing surgical procedures described herein, as well as Cathy Mathis, Cynthia

Zimmerman, and Christopher Bergman for their extensive assistance with histology.

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References [1] Dhawan V, Lytle IF, Dow DE, Huang Y-C, Brown DL. Neurotization Improves


[2] Górski J, Krawczuk I, Górska M, Rutkiewicz J. Inhibition of glycogenesis in rat

muscles partially depleted of glycogen. Am J Physiol 1991;261:C305–9. [3] Wen X, Tresco PA. Effect of filament diameter and extracellular matrix molecule

precoating on neurite outgrowth and Schwann cell behavior on multifilament entubulation bridging device in vitro. J Biomed Mater Res A 2006;76A:626–37. doi:10.1002/jbm.a.30520.

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CHAPTER VI

DISCUSSION, CONCLUSIONS, AND FUTURE DIRECTIONS

John B. Scott

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1. Summary of doctoral dissertation

In summary, in this work we have described a skeletal muscle tissue engineering

approach combining a structural fibrin biomaterial and seeded isogeneic cells with

physical (porous patterning), chemical (neural agrin), and mechanical (cyclic stretch)

cues for the eventual application to repair of volumetric muscle loss injuries.

In Chapter II, we demonstrated that the scaffold featured stiffness of 100-320

kPa, similar to the 79-126 kPa reported for native skeletal muscle [1]. Further, the

scaffold supported ingrowth of axons at a rate of 600-900 μm / day, which was

comparable to clinical expectations of reinnervation at 1 mm / day [2] despite the lack of

specific chemical or cellular trophic support in the scaffold.

In Chapters III & IV, we found using a 2D model in vitro that linkage of agrin to

microparticle surfaces enabled pharmacological signaling to be incorporated into fibrin

scaffolds. Acetylcholine receptor (AChR) clustering, the outcome measure for these

studies, was observed after 1 week or more when agrin was covalently coupled to

microcarriers. Further experimentation using a 3D model in vitro indicated that AChR

clustering may be enhanced by a synergistic effect of agrin and mechanical stretch. In

addition, we described a perfusion method which was capable of uniformly seeding

small (4.5 mm diameter x 4 mm long) 3D scaffolds with cells in densities up to 30 million

cells / cm3.

In Chapter V, we found that tissue engineered muscle repair constructs

implanted in vivo for 2 weeks were partially remodeled by the host environment. These

constructs may have supported innervation by host axons, as predicted by results from

Chapter II, and formation of new muscle tissue, as predicted by those from Chapter III.

As stated in Chapter I, the central hypothesis of this dissertation was that

biomimetic cues could be leveraged to create a tissue engineered construct in vitro that

would accelerate skeletal muscle tissue formation and function following implantation in

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vivo, and furthermore, maintain the skeletal muscle phenotype in vivo during

reinnervation of VML injuries. The observations summarized above provided substantial

support for some aspects of this hypothesis. Chief among these aspects was that AChR

clusters, the observable marker which defined the specified skeletal muscle phenotype,

were observed in vitro within the context of an implantable, 3D, biomimetic tissue

engineered construct. However, other aspects of the hypothesis could not be tested by

results described here due to technical challenges related to creating larger, more

robustly cell-seeded 3D scaffolds, and must be revised or investigated further. These

aspects, and potential avenues for future investigation, are discussed below.

2. Key limitations of the described work and resulting future directions

2.1. Biodegradable agrin microcarriers

Though the results described herein represent a significant proof-of-concept of

the envisioned skeletal muscle tissue engineering strategy, this approach is not without

limitations. The first is that the desired muscle phenotype was evaluated in vitro solely

using histochemical markers for AChR localization. The advantage of the α-bungarotoxin

label, and therefore the reason it was selected as the primary outcome measure in these

studies, is that it provides data on both the presence and, critically, location of AChRs.

Establishing the presence of AChR clustering in the vicinity of agrin-presenting

microparticles was a key aspect of the development of this technology. However, the

physiological relevance of these AChR clusters - specifically for skeletal muscle function

as stated in the central hypothesis - was not established by this dissertation. This was

primarily due to the inflammatory response elicited by constructs implanted in vivo as

discussed in Chapter V.

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As fibrin has been employed by prior skeletal muscle tissue engineering

strategies in vivo [3–5], it is likely that the non-degradable SiO2 microparticles were

responsible for a large portion of the inflammation seen in Chapter V. Ideally, these

microparticles could be replaced by analogs which would degrade at a similar rate to

that of new tissue formation, thereby minimizing the elicited foreign body response

(FBR) [6,7]. As native innervation would obviate the need for exogenous agrin

presentation, the rate of microparticle degradation should also be correlated, if possible,

to innervation of the construct.

2.2. Acetylcholine release from microparticles

Moreover, degradable pharmacologic microparticles would eventually need to be

multi-functional. In normal physiology, acetylcholine (ACh) is believed to cause cell-wide

destabilization of AChR clusters, an effect which is offset at the synapse by the

presentation of agrin [8]. This results in further phenotypic maturity, as it prevents the

muscle from being stimulated to contract by an ectopic AChR cluster.

In Chapter III, AChR clusters apposing covalently-bound agrin microcarriers were

often larger than, and somewhat diffusely located around, areas of cell-particle contact.

Release of soluble ACh from an interior depot within agrin-presenting microparticles

could encourage remodeling of the AChR clusters within the construct in biomimetic

fashion. Nominally, this would maintain the appropriate physiological status of the

implanted and regenerating muscle until native re-innervation could occur.

Combining this ACh release with agrin presentation by a degradable microcarrier

would require a microparticle with very specific properties. As the presented agrin would

preferentially be immobilized on the microparticle surface rather than released as a

soluble signal, the exterior of the microparticle would ideally be the last component to

degrade. Simultaneously, release of soluble ACh would be most straightforward to

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accomplish by diffusion out of an interior depot, which could be extended over a longer

time period if the outer layer of the microparticle served as a diffusion barrier.

Therefore, one potential embodiment of the microparticle would consist of a

faster-degrading polymer core containing ACh molecules surrounded by a slower-

degrading outer polymer shell with covalently-bound surface agrin. Alternatively, if the

degradation products of the core were found not to diffuse out through the shell

alongside the ACh, a composite microparticle could be designed. This would feature a

number of small-diameter, slowly-degrading microparticles with covalently-bound agrin

embedded within a faster-degrading bulk polymer containing diffusible ACh to form a

larger microparticle. Either of these strategies could immobilize an insoluble agrin signal

in one location (analogous to an immobile nerve terminal in vivo) while slowly releasing

ACh, all in the context of a degradable microparticle.

2.3. Modulation of fibrin degradation

Though evaluated at an early time point, the in vivo experiment herein (Chapter

V) was noteworthy in that it was our first significant use of a fibrin scaffold in an animal

model. Though fibrin is a common substrate for muscle tissue engineering, it is

commonly polymerized from fibrinogen solutions of much lower concentration than that

used in the studies discussed here [3,5,9–13]. The host response to implanted

constructs is therefore difficult to predict, even if incorporating degradable microparticles.

Further study on fibrin degradation dynamics in vivo may be required to completely

minimize the FBR. Fortunately, fibrin degradation can be accelerated by decreasing the

component fibrinogen concentration or delayed by chemically crosslinking the network

after gel formation, such as by treatment with genipin [10].

In short, mitigation of the FBR is the logical next step in the development of this

tissue engineering platform. Further characterization or optimization of constructs in vitro

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could be wasted if it is later found that agrin-presenting fibrin elicits unavoidable

inflammation in vivo.

2.4. Functional characterization of tissue engineered muscle repair constructs

As stated above, skeletal muscle function following implantation in vivo

could not be evaluated. However, assuming a mild host response to the implanted

construct, one method of evaluating function is by measurement of force generated by

the construct in response to a contractile stimulus. We have extensively measured force

in response to electrical stimuli [14–16], an outcome measure which is applicable not

only to tissue explants but also to constructs generated in vitro prior to implantation.

Single-cell electrophysiology and / or calcium imaging also represent methods

which could be used to evaluate function. In this case, function refers to development of

excitation-contraction coupling, which can be evaluated even if the contractile apparatus

has not yet matured. However, these are more technically challenging than force

measurement in the context of a fibrin biomaterial because cells to be measured or

imaged must be located within the relative opacity of the fibrin scaffold.

2.5. Physical patterning for modulation of cell behavior

As noted in Chapter I, aligned physical cues have been demonstrated -

primarily in 2D culture in vitro - to encourage maturation of skeletal muscle cells [17–19].

The fibrin scaffold in the current study was designed to take advantage of this

phenomenon, incorporating an array of aligned parallel conduits that additionally

provided porosity for cell seeding. In Chapter IV, we did observe that 3D scaffolds

supported the static seeding and infiltration of C2C12 cells over 8 days of culture, and

that these cells morphologically elongated in the direction of the pores. However, to truly

comment on the potential effects of aligned pores would require:

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1) A control pattern of similar porosity with cues of a different size or shape, such as

the random interconnected pore network in control scaffolds described in Chapter

I; and

2) A method of uniformly and repeatably seeding relevant cells within the pore

networks.

Flow perfusion of rat muscle derived cells (rMDCs), as described in Chapter IV,

filled the second requirement in 4.5 mm diameter x 4 mm long scaffolds. Though these

seeded constructs were too short for bioreactor preconditioning, they would be

sufficiently long for evaluation of physcial cues. Unfortunately, rMDCs which were initially

densely seeded in 3D by perfusion declined in number over 3 days of subsequent

culture. This was presumably due to mass transport limitations and resulting cell

necrosis within statically-cultured constructs.

However, future studies could overcome these limitations. For example, several

approaches to overcome mass transport limitations within other tissue engineering

strategies have been described in the literature. These include incorporation of oxygen-

generating materials [20,21], in vitro perfusion of culture medium after cell seeding of

constructs [22,23], and prevascularization of constructs before implantation in vivo [3].

Alternatively, or in parallel, enabling the robust perfusion seeding of scaffolds 15

mm or more in length would allow truly 3D-seeded scaffolds to be mechanically

preconditioned in the cyclic stretch bioreactor. We hypothesize that preconditioning itself

could enhance cell survival, either by the convection of culture medium through the

construct during actuation or by phenotypic modulation of the cells in question. Robust

3D seeding would also allow more conclusive determination of phenomena observed

herein using the surface-seeded proof of concept model.

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3. Conclusion

In conclusion, this dissertation has described a significant proof-of-concept in this

biomimetic skeletal muscle tissue engineering approach. This concept holds great

potential, and numerous avenues exist for future development either in series or parallel.

It is our hope that the advances described herein one day enhance clinical treatment of

currently devastating and intractable volumetric muscle loss injuries.

References [1] Levinson SF, Shinagawa M, Sato T. Sonoelastic determination of human skeletal


[2] Gordon T, Brushart TM, Chan KM. Augmenting nerve regeneration with electrical

stimulation. Neurol Res 2008;30:1012–22. doi:10.1179/174313208X362488. [3] Bach AD, Arkudas A, Tjiawi J, Polykandriotis E, Kneser U, Horch RE, et al. A new


[4] Dhawan V, Lytle IF, Dow DE, Huang Y-C, Brown DL. Neurotization Improves




[6] Cezar CA, Mooney DJ. Biomaterial-based delivery for skeletal muscle repair. Adv

Drug Deliv Rev 2014. doi:10.1016/j.addr.2014.09.008. [7] Wolf MT, Dearth CL, Sonnenberg SB, Loboa EG, Badylak SF. Naturally derived

and synthetic scaffolds for skeletal muscle reconstruction. Adv Drug Deliv Rev 2014. doi:10.1016/j.addr.2014.08.011.

[8] Witzemann V. Development of the neuromuscular junction. Cell Tissue Res

2006;326:263–71. doi:10.1007/s00441-006-0237-x. [9] Bian W, Bursac N. Engineered skeletal muscle tissue networks with controllable


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[13] Hinds S, Bian W, Dennis RG, Bursac N. The role of extracellular matrix

composition in structure and function of bioengineered skeletal muscle. Biomaterials 2011;32:3575–83. doi:10.1016/j.biomaterials.2011.01.062.













[20] Oh SH, Ward CL, Atala A, Yoo JJ, Harrison BS. Oxygen generating scaffolds for

enhancing engineered tissue survival. Biomaterials 2009;30:757–62. doi:10.1016/j.biomaterials.2008.09.065.

[21] Ward CL, Corona BT, Yoo JJ, Harrison BS, Christ GJ. Oxygen generating

biomaterials preserve skeletal muscle homeostasis under hypoxic and ischemic conditions. PloS One 2013;8:e72485. doi:10.1371/journal.pone.0072485.

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prevascularized cell-based constructs for myocardial tissue engineering. Stem Cells Int 2014;2014:434169. doi:10.1155/2014/434169.

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CURRICULUM VITAE

JOHN BRADFORD SCOTT, B.S. Ph.D. Candidate

Virginia Tech – Wake Forest University School of Biomedical Engineering and Sciences Wake Forest Institute for Regenerative Medicine

EDUCATION Ph.D. Wake Forest University, Winston-Salem, NC

Biomedical Engineering 2008-Current

B.S. The University of Tennessee, Knoxville, TN

Biomedical Engineering Summa Cum Laude, with Honors 2004-2008

RESEARCH EXPERIENCE 2008-2015 Wake Forest Institute for Regenerative Medicine, Winston-Salem, NC

Graduate Research Associate, George J. Christ Laboratory (2011-2015) and Justin M. Saul Laboratory (2008-2011) • Directed $350,000 US Army-funded project aimed at healing

devastating, intractable muscle injuries. • Designed experiments by synthesizing literature findings and personal

expertise. • Developed material constructs incorporating biomimetic patterning,

drug delivery, and mechanical cues. 2007 The University of Tennessee REU Program, Knoxville, TN Undergraduate Researcher, Anthony E. English Laboratory

• Quantified effects of potential anti-cancer drugs. • Validated drug success using wavelet theory.

PEER-REVIEWED PUBLICATIONS 1. Scott JB, Ward CW, Corona BT, Deschenes MR, Harrison BS, Saul JM, Christ GJ. In

vitro development of a novel tissue engineered muscle repair construct. 2015. In preparation.

2. Scott JB, Afshari M, Kotek R, Saul JM. The promotion of axon extension in vitro using polymer-templated fibrin scaffolds. Biomaterials. 2011 Jul;32(21):4830-9.

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PRESENTATIONS AND REPORTS TO RESEARCH SPONSORS

1. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Final Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); December 2014

2. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Quarter 7 Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); July 2014

3. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Quarter 6 Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); April 2014

4. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, WFIRM Annual Retreat, February 10, 2014

5. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Quarter 5 Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); January 2014

6. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, NCTERMS Annual Meeting, October 28, 2013

7. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Annual Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); October 2013

8. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Quarter 3 Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); July 2013

9. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, SBES Student Research Symposium, May 16, 2013

10. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Quarter 2 Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); April 2013

11. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function" (podium), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, Monthly Meeting Winston-Salem IEEE Chapter, April 10, 2013

12. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, WFU Graduate Student Research Day, March 21, 2013

13. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, WFIRM Annual Retreat, March 19, 2013

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14. "Novel Methods for Enhanced Formation and Stabilization of Motor Endplates in Tissue Engineered Skeletal Muscle," Quarter 1 Status Report; prepared by John B. Scott and George J. Christ, PhD (PI); January 2013

15. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function" (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, BMES Annual Meeting, October 2012

16. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function" (poster), John B. Scott, Catherine L. Ward, Benjamin T. Corona, Benjamin S. Harrison, Justin M. Saul, and George J. Christ, BMES Annual Meeting, September 2012

17. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Justin M. Saul, and George J. Christ, NCTERMS Annual Meeting, September 10, 2012

18. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Justin M. Saul, and George J. Christ, SBES Student Research Symposium, May 10, 2012

19. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function” (poster), John B. Scott, Justin M. Saul, and George J. Christ, WFIRM Annual Retreat, March 5, 2012

20. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function In Vitro and In Vivo” (podium), John B. Scott, Justin M. Saul, and George J. Christ, TERMIS – North America Annual Meeting, December 13, 2011

21. “Development of a Novel Tissue Engineered Muscle Repair Construct with Potential for Enhanced Motor End Plate Formation and Function In Vitro and In Vivo” (poster), John B. Scott, Justin M. Saul, and George J. Christ, NCTERMS Annual Meeting, November 4, 2011

22. “Intraluminal Nerve Scaffolds Promote Axon Extension In Vitro” (poster), John B. Scott and Justin M. Saul, SBES Student Research Symposium, May 12, 2011

23. “Intraluminal Nerve Scaffolds Promote Axon Extension In Vitro” (poster), John B. Scott and Justin M. Saul, WFU Graduate Student Research Day, March 22, 2011

24. “Polymer-templated fibrin scaffolds promote axon extension in vitro” (podium), John B. Scott and Justin M. Saul, WFIRM Annual Retreat, March 7, 2011

25. “Intraluminal Nerve Scaffolds Promote Axon Extension In Vitro” (poster), John B. Scott and Justin M. Saul, WFIRM Annual Retreat, March 7, 2011

26. “Intraluminal Nerve Scaffolds Promote Axon Extension In Vitro” (poster), John B. Scott and Justin M. Saul, TERMIS – North America Annual Meeting, December 5, 2010

27. “Templated Biomaterial Scaffolds for Directed Neural Regeneration” (podium), John B. Scott and Justin M. Saul, Monthly Meeting Winston-Salem IEEE Chapter, August 11, 2010

28. “Intraluminal Nerve Scaffolds Promote Axon Extension and Cell Infiltration In Vitro” (poster), John B. Scott and Justin M. Saul, SBES Student Research Symposium, May 13, 2010

29. “Intraluminal Nerve Scaffolds Promote Axon Extension and Cell Infiltration In Vitro” (poster), John B. Scott and Justin M. Saul, WFU Graduate Student Research Day, March 23, 2010

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30. “Intraluminal Nerve Scaffolds Promote Axon Extension and Cell Infiltration In Vitro” (poster), John B. Scott and Justin M. Saul, WFIRM Annual Retreat, March 8, 2010

31. “Templated Biomaterial Scaffolds for Directed Neural Regeneration” (podium), John B. Scott and Justin M. Saul, Monthly Meeting Winston-Salem IEEE Chapter, January 13, 2010

32. “Development of Polymer-Templated Intraluminal Nerve Scaffolds” (poster), John B. Scott and Justin M. Saul, NCTERMS Annual Meeting, November 2009

33. “Development of Polymer-Templated Intraluminal Nerve Scaffolds” (poster), John B. Scott and Justin M. Saul, BMES Annual Meeting, October 7, 2009

34. “Aligned Conduit Scaffolds Direct Neurite Extension In Vitro” (poster), John B. Scott and Justin M. Saul, WFU Graduate Student Research Day, April 6, 2009

35. “Aligned Conduit Scaffolds Direct Neurite Extension In Vitro” (poster), John B. Scott and Justin M. Saul, WFIRM Annual Retreat, March 23, 2009

36. “Development of Axially-Aligned Scaffolds for Optimization of Neural Regeneration” (poster), John B. Scott and Justin M. Saul, NCTERMS Annual Meeting, November 2008

AWARDS

• Outstanding Poster, North Carolina Tissue Engineering and Regenerative Medicine Society Annual Meeting. November 2011

• Best Presentation, Predoctoral Category, Wake Forest Institute for Regenerative Medicine Annual Retreat. March 2011

• Achieved rank of Eagle Scout after participation in Boy Scouts of America since age 5. May 2004

PROFESSIONAL EXPERIENCE 2014-Current Technology Commercialization Intern

Wake Forest Innovations, Winston-Salem, NC

• Interviewed Wake Forest inventors to assess IP protection and marketing potential of disclosed inventions.

• Analyzed geographic markets for solar technology to inform patent nationalization decisions.

• Drafted marketing materials to engage potential licensees for nanomaterials, oncology, and RFID inventions.

• Met with industry partners to discuss technologies and licensing options, both non-confidentially and under CDA.

2006 Reliability Technology Intern Eastman Chemical Company, Kingsport, TN

• Located historically unreliable equipment. • Isolated root causes of recurring malfunctions.

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PATENT 2012 Application PCT/US2012/057094 “Bioscaffolds for Formation of Motor Endplates”

Co-Inventor ACTIVITIES 2013-2014 First-Ever Breast Cancer Startup Challenge

Member: Wake Forest Team 3

2009-2014 Biomedical Engineering Society Member 2009-2013 Wake Forest Institute for Regenerative Medicine Summer Scholars

Program, Minority Access to Research Careers Program, and Wake Forest University Work-Study Program

Mentor of seven visiting student researchers 2005-2008 Theta Tau Professional Engineering Fraternity, Chi Gamma Chapter

Founding Brother, Scribe, and Author of Colony Bylaws SKILLS Research Design

• Experimental design • Literature review

Technology Commercialization

• Intellectual property protection • Patent technology review • Market analysis • Drafting of non-confidential marketing materials • Legal agreements

Biomaterials Processing and Characterization

• Biomaterial scaffold design • Bioreactor design and operation • Covalent crosslinking • Tensile Testing • Critical point drying • Metallic sputtering • Scanning electron microscopy • Polymer melt extrusion

Cell Culture

• Aseptic technique • Primary cell isolation and expansion

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Histology

• Immunohistochemistry • Special stains • Confocal microscopy • Fluorescence and brightfield microscopy

In Vivo Experimentation

• Rodent surgeries and explants • Electrophysiological assessment

Other

• Authorship of GLP / GMP compliant standard operating procedures and master batch records

• Microsoft Windows • Microsoft Office • FIJI / ImageJ • GraphPad PRISM

REFERENCES George Christ, Ph.D. Academic Advisor, 2011-Current

Professor, Wake Forest University School of Medicine [email protected] 434-924-5794

Charlie Shaw, Ph.D. Technology Commercialization Internship Mentor, 2014-Current Commercialization Associate, Wake Forest Innovations [email protected] 336-716-3729

Stephen Susalka, Ph.D., CLP Professor, Principles of Intellectual Property Development, 2014 Executive Director, Association of University Technology Managers [email protected] 336-546-7977

Peter Sheldrake, Ph.D. Professor, Commercializing Innovation, 2014 Adjunct Instructor, Wake Forest University School of Business [email protected] 336-705-0735

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Justin Saul, Ph.D. Academic Advisor, 2008-2011 Associate Professor, Miami University College of Engineering & Computing [email protected] 513-529-0769

Documents

CUES BY JOHN BRADFORD SCOTT A Dissertation Submitted … · CUES BY JOHN BRADFORD SCOTT A Dissertation Submitted to the Graduate Faculty of ... Perfusion loop schematic ... in tissue