NEUROGLIAL DIFFERENTIATION AND NEO-INNERVATION

IN EXTRACELLULAR MATRIX-BASED TISSUE

ENGINEERED INNERVATED SMOOTH MUSCLE SHEETS

BY

SHREYA RAGHAVAN

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

August 2014

Winston-Salem, NC

Approved By

Khalil N. Bitar, Ph.D., AGAF, Advisor and Chair

George J. Christ, Ph.D.

Aaron S. Goldstein, Ph.D.

Padmavathy Rajagopalan, Ph.D.

Mark E. Van Dyke, Ph.D.

ii

DEDICATION

This dissertation is dedicated to Amma and Appa – their unerring faith in me has

taught me that commitment and perseverance are all I need to achieve my dreams;

To Ram – you are my rock, and you complete me.

iii

ACKNOWLEDGEMENTS

This journey called graduate school has been a long and spectacular one that started in Ann

Arbor and ended in Winston-Salem. Graduate school has tested me immensely, has been

tenuous at times – but in retrospect (it’s also much easier to say this now), it has been quite

an amazing experience. It would also not have been possible without all the help,

encouragement, guidance, and support I received from my mentor, my committee, friends

and family.

I would like to express my gratitude for the support I’ve received from my mentor, Dr. Khalil

Bitar. He taught me the value of collaborative science and scientific rigor. He taught me to

respect the big picture, while working out the details of a scientific plan. He laid out this

incredible path for me, and I walked through it with his guidance. His mentorship has taught

me lessons in both my professional and personal life. Above all else, I have been made to

feel like a gifted student and an independent researcher.

My committee has kept me on my toes and on the scientific straight and narrow. My

discussions with them have always been productive and fruitful no matter what it was about

– rheometry, physiology, appropriate snow tires, or why google maps may not be the best

route indicator. Without their flexibility and unfathomable patience in the face of my email

barrage, calendar wars might have been an imminent danger to graduation for years.

Thank you for anchoring me in lab, Saranya - cheering me up with online shopping, and

teaching me how to do semi-dry transfers and western blots so they work every single time.

My thesis would be ~ 40% empty without this. There is no other person I’d rather share the

Stadie-Riggs slicer and dicer with. Elie saw me through failed experiments and celebrated

my mini-lab successes. Thank you for enabling my 2pm Kit Kat cravings, reminding me why

me and the elliptical are besties, keeping me focused on the big picture in life, and always

iv

listening to me whine. Sita and Bob taught an insufferable engineer about all things

biological. Steve, thank you for taking over and making this transition painless.

BiBi, you made Winston feel like home for me - who will make me fancy mason jar drinks

and fake pancakes when I go back to Ann Arbor? Hannah banana – meow ; such engineer,

so Matlab. Daniel, you are my grumpy cat enabler (yaysies), Without Sneha, Amritha, Kristy,

Sean, Jess, and the rest of the WFIRM gang, happy hour would not be the same. To the

competent core techs (Ray, Jay, Cathy and Brian) and the benevolent lab manager (Tom), I

would have had melt downs if equipment failure was thrown in this grad school mix. Beth

and Donna – thank you for fighting my calendar battles. I will truly miss WFIRM, even

though my pipettor went for a stroll every once in a while.

Momo: thank you for never once asking me when I was going to graduate. Thank you for

fueling me with midnight maggi and rendezvous crepes to keep me going for my 1am

timepoints and biochem crams. Neha, where would I be without the much-needed girl talk

through rough days? Savi, your spare bedroom was my data analysis haven. Deepi, Kapil,

Jans, Monkey, and Shobs – thank you for being there through the thick and thin of it all.

Bala, Vateez, and Samperv: seeing you endure graduate school made it easier for me to

know what to expect (while I was expecting? To graduate, that is).

Above all else, this experience would not have even been possible without my family by my

side. Malu and Prasad – I have looked up to you, and you motivate me to do more and be

more. Anna, you have been my inspiration, and I have always been in awe of how you

achieve things and make it look easy doing so. Lolly and Ajay – you are everything a little

sister can ask for. My parents-in-law, for listening to me patiently and always filling me with

positive thoughts. Chitthi and chittha- I did grow up to be a doctor (albeit of philosophy).

v

Amma and Appa – you are my biggest role models. I work every day to achieve greater

heights, because you dreamt this for me. Words cannot begin to describe how much your

support and encouragement has lit many a dark night for me through graduate school. Appa

– thank you for all that you have sacrificed for me, teaching me bountiful patience, and

inspiring me to be an engineer. Amma – you have singlehandedly set an example for the

endless possibilities for an independent woman, and have taught me to dream big. I hope I

can make you both proud.

Ram, needless to say, this journey would not have even been possible if not for you. Thank

you for always standing by my side, and for being there for me even when there was 500

miles between us. This hasn’t been easy on you either, but I truly appreciate the love,

support and steadfastness you have displayed through this. You are my strength, and it is

because of you that graduate school came to fruition. Thank you for willingly spending the

last three years living in two different cities, but never once making it feel like an imposition.

Thank you for sharing this journey with me (and for playing tennis with me even when I

couldn’t return a serve to save my life).

vi

TABLE OF CONTENTS

DEDICATION ........................................................................................................................................ ii

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

LIST OF FIGURES ........................................................................................................................... viii

LIST OF TABLES ............................................................................................................................... xi

LIST OF ABBREVIATIONS ............................................................................................................. xii

ABSTRACT........................................................................................................................................ xiii

CHAPTER 1: INTRODUCTION & SIGNIFICANCE ...................................................................... 1

CHAPTER 2: INTESTINAL REGENERATION: THE BIOENGINEERING APPROACH ....... 4

This manuscript was accepted for publication as Chapter 34, in Regenerative Medicine

Applications in Organ Transplantation, First Edition, Elsevier 2013 ....................................... 4

CHAPTER 3: SPECIFIC AIMS AND OBJECTIVES OF THE THESIS .................................... 25

CHAPTER 4: NEUROGLIAL DIFFERENTIATION OF ADULT ENTERIC NEURONAL

PROGENITOR CELLS AS A FUNCTION OF EXTRACELLULAR MATRIX COMPOSITION

............................................................................................................................................................. 30

vii

This manuscript was accepted for publication in Biomaterials 2013, 34(28): 6649-6658; . 30

CHAPTER 5: BIOENGINEERED THREE-DIMENSIONAL PHYSIOLOGICAL MODEL OF

COLONIC LONGITUDINAL SMOOTH MUSCLE IN VITRO ..................................................... 56

This manuscript was accepted for publication in Tissue Engineering Part C 2010; 16(5):

999-1009 ......................................................................................................................................... 56

CHAPTER 6: EXTRACELLULAR MATRIX MODULATES DIFFERENTIATION OF

NEURONAL SUBTYPES IN TISSUE ENGINEERED INNERVATED INTESTINAL

SMOOTH MUSCLE SHEETS ......................................................................................................... 81

This manuscript was accepted for publication in Biomaterials (in press, May 2014) .......... 81

CHAPTER 7: NEO-INNERVATION OF EXPERIMENTALLY DENERVATED COLON

USING TISSUE ENGINEERED INNERVATED LONGITUDINAL SMOOTH MUSCLE

SHEETS ............................................................................................................................................ 111

CHAPTER 8: FUTURE DIRECTIONS ......................................................................................... 129

CHAPTER 9: SUMMARY AND CONCLUSIONS ...................................................................... 134

SCHOLASTIC VITA ........................................................................................................................ 136

viii

LIST OF FIGURES

Figure 1: Schematic of the various sources of neural stem/progenitor cells ............. 14

Figure 2: Rabbit enteric neurospheres ..................................................................... 37

Figure 3: Schematic of enteric neurosphere differentiation as a function of ECM

composition .............................................................................................................. 38

Figure 4: Neuronal differentiation on individual coated coverslips ............................ 39

Figure 5: Neuronal differentiation on collagen-laminin substrates ............................ 41

Figure 6: Neuronal differentiation on composite collagen-laminin-heparan sulfate

(HS) substrates ........................................................................................................ 43

Figure 7: Glial differentiation on primary coated substrates ..................................... 45

Figure 8: Glial differentiation on collagen-laminin substrates ................................... 46

Figure 9: Glial differentiation on composite collagen-laminin-heparan sulfate (HS)

substrates ................................................................................................................. 47

Figure 10: Morphometric mesaurements .................................................................. 51

Figure 11: Process schematic for bioengineering rabbit Longitudinal Smooth Muscle

Cell Sheet ................................................................................................................. 65

Figure 12: Engineering highly aligned LSMC Sheets ............................................... 66

Figure 13: Immunofluorescence on isolated RLSMC ............................................... 67

ix

Figure 14: Passive tension and Signal to Noise Ratio (SNR) .................................. 68

Figure 15: Force generation in response to 1µM Acetylcholine in bioengineered

tissue and native longitudinal tissue ......................................................................... 69

Figure 16: Dose-Response to Acetylcholine in bioengineered tissue and native

longitudinal tissue .................................................................................................... 70

Figure 17: Response to Acetylcholine in Zero Calcium in bioengineered tissue and

rabbit longitudinal tissue .......................................................................................... 71

Figure 18: Inhibition of force generation by pre-incubation with VIP in bioengineered

tissue and rabbit longitudinal tissue ......................................................................... 71

Figure 19: Comparison of force generated in response to Acetylcholine ................. 72

Figure 20: Scanning electron micrographs of dehydrated ECM gels ....................... 89

Figure 21: Neuronal differentiation within tissue engineered sheets ........................ 91

Figure 22: Immunoblot analysis of tissue engineered longitudinal sheets ............... 92

Figure 23: Immunofluorescence for differentiated neurons within tissue engineered

sheets ...................................................................................................................... 93

Figure 24: Potassium chloride induced contraction of tissue engineered sheets ..... 95

Figure 25: Acetylcholine induced contraction ........................................................... 96

Figure 26: Electrical Field Stimulation induced relaxation in tissue engineered sheets

................................................................................................................................. 99

x

Figure 27: Inhibition of relaxation with L-NAME ..................................................... 100

Figure 28: Inhibition of relaxation with VIP-Ra ....................................................... 101

Figure 29: Enteric neurospheres from GFP mice ................................................... 114

Figure 30: Schematic of neo-innervation using tissue engineered innervated sheets

............................................................................................................................... 116

Figure 31: βIII Tubulin expression in sections of BAC treated, untreated and neo-

innervated colons ................................................................................................... 118

Figure 32: Immunoblot expression of βIII Tubulin in explant tissues ...................... 120

Figure 33: Immunofluorescence in explant tissues ................................................ 121

Figure 34: Electrical Field Stimulation (EFS) induced relaxation in explant colon

tissues .................................................................................................................... 123

Figure 35: Inhibition of nNOS in explant colon tissues ........................................... 124

Figure 36: Reduced Neuronal βIII Tubulin in BAC treated segment ....................... 130

Figure 37: Electrical Field Stimulation in BAC-treated segments in vivo ................ 131

Figure 38: Restoration of NO-mediated relaxation in BAC-treated neo-innervated

colon segments in vivo ........................................................................................... 132

xi

LIST OF TABLES

Table 1: Viscoelastic modulus of ECM gels ............................................................. 90

Table 2: Acetylcholine-induced contraction in tissue engineered sheets ................. 98

Table 3: EFS-induced relaxation in tissue engineered sheets ............................... 104

xii

LIST OF ABBREVIATIONS

Ach Acetylcholine

AU Arbitrary Units

AUC Area Under the Curve

BAC Benzalkonium chloride

ChAT Choline acetyltransferase

CNS Central nervous system

ECM Extracellular matrix

EFS Electrical Field Stimulation

ENS Enteric nervous system

GDNF Glial-cell derived neurotrophic factor

GFAP Glial fibrillary acidic protein

GFP Green fluorescent protein

GI Gastrointestinal

HBSS Hank’s balanced salt solution

HSCR Hirschsprung’s Disease

IAS Internal Anal Sphincter

KCl Potassium chloride

LES Lower esophageal sphincter

L-NAME Nω-Nitro-L-arginine methyl ester hydrochloride

nNOS neuronal nitric oxide synthase

NSC Neural Stem Cell

pLL poly-L-lysine

RLSMC Rabbit longitudinal smooth muscle cell

TTX Tetrodotoxin

VIP Vasoactive Intestinal Peptide

VIP-Ra [D-p-Cl-Phe6, Leu17]-Vasoactive Intestinal Peptide

xiii

ABSTRACT

Shreya Raghavan

NERUOGLIAL DIFFERENTIATION AND NEO-INNERVATION IN

EXTRACELLULAR MATRIX-BASED TISSUE ENGINEERED

INNERVATED SMOOTH MUSCLE SHEETS

Dissertation under the direction of

Khalil N. Bitar, PhD, AGAF

Excitatory and inhibitory motor neurons of the myenteric plexus in the enteric nervous

system regulate and coordinate smooth muscle motility. Enteric neurons and glia of the

myenteric plexus are organized into ganglia, that are surrounded by an extracellular matrix

composed primarily of collagen IV, laminin and heparan sulfate. Disruption of ganglionic

integrity, loss or damage of all or specific subsets of enteric neurons occur in several

pathological states resulting in intestinal dysmotility and reduced quality of life. Neural-crest

derived enteric neuronal progenitor cells persist in the adult mammalian gut, and contribute

to the plasticity observed in the adult enteric nervous system. Neural stem cell

transplantation is an emerging therapeutic paradigm aimed at repopulating enteric ganglia

and restoring intestinal motility. However, transplantation of neural stem cells within the gut

has demonstrated poor long-term phenotype maintenance and stability. We hypothesized

that extracellular matrix (ECM) microenvironments could be used to direct the differentiation

of enteric neuronal progenitor cells in vitro, prior to transplantation. The ECM plays an

enormous role developmentally, in the migration of neural crest cells into the gastrulating

mesoderm, and subsequent differentiation into neuroglial phenotypes appropriate for the

xiv

intestinal microenvironment. We evaluated the potential of ECM microenvironments in

modulating enteric neuroglial differentiation in 2D and 3D culture. We isolated enteric

neuronal progenitor cells in sufficient numbers from small intestinal biopsies of adult rabbits.

When presented with various ECM culture substrata, we observed that glial differentiation

was inhibited in surfaces promoting neuronal differentiation. A pre-ponderance of

differentiated glia was obtained in substrata that contained no ECM components. Neuronal

differentiation was enhanced in the presence of collagen IV, with the emergence of

branched neurons. Neurites on laminin coated substrata were the longest, averaging

~400µm. Heparan sulfate substrata promoted organization and preliminary neuronal

networking. A tissue engineering model of innervated intestinal smooth muscle sheets was

utilized to facilitate neuronal differentiation in viscoelastic hydrogels. Uniaxially aligned

smooth muscle monolayers were obtained using substrate microtopography. ECM hydrogels

were used to encapsulate enteric neuronal progenitor cells, generating innervated intestinal

smooth muscle sheets. By varying ECM hydrogel composition, proportions of differentiated

neurons were altered in the tissue engineered sheets. Sheets manufactured with collagen I

and laminin had enriched populations of excitatory cholinergic neurons with minimal

inhibitory neurons. Sheets manufactured with collagen IV had enriched populations of

inhibitory nitrergic neurons. Cholinergic and nitrergic neuronal mediated

contraction/relaxation matched their expression levels, when assessed using organ bath

studies and electrical field stimulation. A therapeutic application for tissue engineered

innervated sheets was evaluated as the potential to neo-innervate explant cultures of colon.

Benzalkonium chloride treatments were used to denervate explant colons. Tissue

engineered sheets were manufactured with enriched populations of nitrergic neurons, and

maintained in co-culture with the recipient denervated colons. Neo-innervation was

visualized following co-culture using immunohistochemistry and immunoblot expression for

xv

neuronal marker βIII Tubulin. Neuronal physiology was also improved following co-culture,

indicating functional neo-innervation.

To summarize, these studies demonstrate that enteric neuronal progenitor cells can be

differentiated into a pre-ponderance of either neurons or glia by modifying ECM culture

substrata. A functional diversity in motor neuronal phenotypes can be generated by

modifying ECM microenvironments in three-dimensional culture. Tissue engineered models

of innervated intestinal smooth muscle sheets were obtained, with enriched excitatory or

inhibitory neurons or a balanced expression of both excitatory and inhibitory subsets.

Constituent smooth muscle cells in the tissue engineered sheets retained a contractile

phenotype. Differentiated neurons within the tissue engineered sheets were physiologically

functional in mediating smooth muscle contraction and relaxation. Furthermore, these tissue

engineered sheets were capable of functional neo-innervation. Taken together, these

studies suggest that ECM microenvironments can be successfully used to modulate

neuroglial differentiation of enteric neuronal progenitor cells in vitro, prior to transplantation.

Transplantation of such pre-differentiated neurons will have a positive impact in remedying

dysganglionosis of the gastrointestinal tract.

1

CHAPTER 1: INTRODUCTION & SIGNIFICANCE

Gastrointestinal (GI) motor function is a coordinated interaction between the enteric

nervous system, interstitial cells of Cajal, smooth muscle cells, and the central nervous

system [1]. Disruption or absence of enteric ganglia can result in altered gut motility,

clinically presented as constipation, diarrhea or altered GI transit. In the spectrum of

neuromuscular disorders of the gut, the pathologies stem from the loss/damage of all

intrinsic innervation to the damage of specific neuronal subsets. Enteric neuropathies

including loss, degeneration, functional impairment or inadequate neuronal differentiation

lead to several gastrointestinal motor disorders [6].

The best characterized aganglionic disorder of the gut is Hirschsprung’s disease

(HSCR), occuring in 1 out of 5000 live births a year [3]. Enteric ganglia are absent in varying

lengths of the distal GI tract, resulting in functional obstruction [4]. Surgical resections of the

aganglionic bowel offer poor long-term outcomes, with persistent incontinence and lack of

adequate reinnervation and coordinated motility [5, 6].

Sphincteric achalasia, commonly occuring in the lower esophageal sphincter, results

in severe dysphagia with an incidence of 1 in 50,000. The underlying pathology in

sphincteric achalasia is the loss of inhibitory motor neurons in the myenteric plexus [7, 8].

Surgical treatment for achalasia includes injections of bulking agents and neurotoxins like

botulinum toxin, to reduce sphincteric tone and improve relaxation [9].

Enteric neuropathy can also be secondary to metabolic diseases like diabetes. In

diabetic gastric tissue, specific populations of neurons are reported missing including

nitrergic neurons in the fundus leading to impaired accomodation. Damaged cholinergic

neurons in the diabetic gastric antrum leads to improper antral contractions and loss of

agitated mixing required in the stomach [10-12]. Loss of nitrergic neurons in the diabetic or

aging pylorus sphincter result in delayed gastric emptying, thereby resulting in altered lower

2

GI motility and transit. Pharmacological therapies including pro-kinetic agents stimulate

cholinergic or nitrergic activity; however these therapies provide symptomatic relief without

addressing the underlying pathology.

Neural stem cell transplantation is an emerging therapeutic field, aimed at

repopulating enteric neuronal plexuses in order to restore gut motility. Cell-based therapies

utilize the fact that enteric neuronal progenitor cells exist post-natally in adult human gut,

including ganglionic bowel of patients with Hirschsprung’s disease [13-15]. These progenitor

cells undergo differentiation upon transplantation into embryonic aganglionic gut explants,

expressing markers for mature motor neurons and glia [16, 17]. Cell injection therapies in

vivo have primarily relied on the injection of progenitor cells itself, where these cells have

shown to migrate very limited distances and demonstrate preliminary electrophysiology

upon acquiring neurochemical coding appropriate for enteric neurons. However, with the

approach of direct injection of progenitor cells, there is no control over their eventual

phenotype or the maintenance of their phenotype. For diseases like Hirschsprung’s, where

all neuronal phenotypes are missing, such an approach could be beneficial [3, 6]. For

diseases like sphincteric achalasia, pyloric stenosis, or diabetic gastroparesis, specific

neurotransmitter deficiencies or damage/loss of nitrergic neurons occur [6, 18]. In such

cases, it is beneficial to identify methods of biasing neuronal differentiation into specific

neuronal subtypes prior to transplantation. Through the research outlined and performed in

this dissertation, an innovative method of terminally differentiating adult mammalian enteric

neuronal progenitor cells in vitro into specific neuronal and glial phenotypes is described. A

regenerative medicine based approach is evaluated to utilize these differentiated neurons to

neo-innervate aganglionic gut. In the long term, this approach could be optimized to deliver

specific neural subsets to the dysganglionic gut. Furthermore, protective microenvironments

that also provide adequate trophic support for neurons can be identified, to enhance

transplantation efficiency and survival.

3

References

[1] Hansen MB. The enteric nervous system I: organisation and classification. Pharmacol Toxicol. 2003;92:105-13. [2] De Giorgio R, Stanghellini V, Barbara G, Corinaldesi R, De Ponti F, Tonini M, et al. Primary enteric neuropathies underlying gastrointestinal motor dysfunction. Scand J Gastroenterol. 2000;35:114-22. [3] Arshad A, Powell C, Tighe MP. Hirschsprung's disease. BMJ. 2012;345:e5521. [4] Kessmann J. Hirschsprung's disease: diagnosis and management. Am Fam Physician. 2006;74:1319-22. [5] Wilkinson DJ, Edgar DH, Kenny SE. Future therapies for Hirschsprung's disease. Semin Pediatr Surg. 2012;21:364-70. [6] Baillie CT, Kenny SE, Rintala RJ, Booth JM, Lloyd DA. Long-term outcome and colonic motility after the Duhamel procedure for Hirschsprung's disease. J Pediatr Surg. 1999;34:325-9. [7] Park W, Vaezi MF. Etiology and pathogenesis of achalasia: the current understanding. Am J Gastroenterol. 2005;100:1404-14. [8] Sadowski DC, Ackah F, Jiang B, Svenson LW. Achalasia: incidence, prevalence and survival. A population-based study. Neurogastroenterol Motil. 2010;22:e256-61. [9] Pehlivanov N, Pasricha PJ. Achalasia: botox, dilatation or laparoscopic surgery in 2006. Neurogastroenterol Motil. 2006;18:799-804. [10] Takahashi T, Nakamura K, Itoh H, Sima AA, Owyang C. Impaired expression of nitric oxide synthase in the gastric myenteric plexus of spontaneously diabetic rats. Gastroenterology. 1997;113:1535-44. [11] Pasricha PJ, Pehlivanov ND, Gomez G, Vittal H, Lurken MS, Farrugia G. Changes in the gastric enteric nervous system and muscle: a case report on two patients with diabetic gastroparesis. BMC Gastroenterol. 2008;8:21. [12] Chandrasekharan B, Srinivasan S. Diabetes and the enteric nervous system. Neurogastroenterol Motil. 2007;19:951-60. [13] Schäfer Kh, Micci Ma, Pasricha PJ. Neural stem cell transplantation in the enteric nervous system: roadmaps and roadblocks. Neurogastroenterology & Motility. 2009;21:103-12. [14] Gershon MD. Transplanting the enteric nervous system: a step closer to treatment for aganglionosis. Gut. 2007;56:459-61. [15] Rauch U, Hansgen A, Hagl C, Holland-Cunz S, Schafer KH. Isolation and cultivation of neuronal precursor cells from the developing human enteric nervous system as a tool for cell therapy in dysganglionosis. Int J Colorectal Dis. 2006;21:554-9. [16] Natarajan D, Grigoriou M, Marcos-Gutierrez CV, Atkins C, Pachnis V. Multipotential progenitors of the mammalian enteric nervous system capable of colonising aganglionic bowel in organ culture. Development. 1999;126:157-68. [17] Lindley RM, Hawcutt DB, Connell MG, Almond SL, Vannucchi MG, Faussone-Pellegrini MS, et al. Human and mouse enteric nervous system neurosphere transplants regulate the function of aganglionic embryonic distal colon. Gastroenterology. 2008;135:205-16 e6. [18] Kraichely RE, Farrugia G. Achalasia: physiology and etiopathogenesis. Dis Esophagus. 2006;19:213-23.

4

CHAPTER 2: INTESTINAL REGENERATION: THE

BIOENGINEERING APPROACH

Khalil N. Bitar1,2 and Shreya Raghavan1,2

1Wake Forest Institute for Regenerative Medicine, Wake Forest School of Medicine, Winston-Salem NC 27101

2Virginia Tech-Wake Forest School of Biomedical Engineering and Sciences, Winston-Salem NC 27101

Running title: Neurodegenerative diseases of the GI tract

This manuscript was accepted for publication as Chapter 34, in Regenerative

Medicine Applications in Organ Transplantation, First Edition, Elsevier 2013

5

1. Introduction to the Enteric Nervous System (ENS)

Gastrointestinal function (motility, secretion, and maintenance of fluid and electrolyte

homeostasis) is controlled by three branches of the autonomic nervous system – the

sympathetic, parasympathetic and the enteric nerves. Intestinal secretomotor activity occurs

in the absence of extrinsic autonomic contributions, and is controlled predominantly by the

enteric nervous system (ENS). Hence, the ENS is called the intrinsic innervation of the gut.

The ENS assimilates information from local sensory input, muscle and mucosa and

determines the appropriate response of the gut. The ENS is the largest branch of the

autonomic nervous system. The ENS is contained within the walls of the gastrointestinal

tract and divided into two ganglionated plexi –Myenteric/Auerbach plexus between the

circular and longitudinal smooth muscle of the gut; Submucosal/Meissner plexus between

the mucosal layer and circular smooth muscle of the gut. The ENS is comprised of several

classes of functional neurons and enteric glia, similar to the central nervous system (CNS;

(1)).

The ENS demonstrates functional autonomy, capable of mediating gastrointestinal

secretomotor function independent of the CNS. ENS function can additionally be modulated

by sympathetic and parasympathetic ganglia of the autonomic nervous system arising from

the vagal, splanchnic or the sacral ganglia. Structurally, in the mammalian gut, the neurons

of the ENS (arising from either plexi) have axons >100nm away from effector smooth

muscle cells. Neurons lack a clearly defined ‘post-synaptic area’ and hence do not form

neuromuscular junctions, as observed in other parts of the peripheral nervous system (2).

Smooth muscle cells in the intestine exist in an electrical syncytium, and are innervated by

hundreds of excitatory and inhibitory motor neurons to effect gastrointestinal motility and the

peristaltic reflex.

6

Major functional groups of neurons in the mammalian myenteric plexus are: intrinsic

sensory neurons, motor neurons that result in smooth muscle contraction or relaxation, and

interneurons. Relaxant neurotransmitters include nitric oxide, vasoactive intestinal peptide

(VIP), pituitary adenylate cyclase activating peptide and purines. Contractile

neurotransmitters in the gut include acetylcholine and tachykinins (3). Depending on the

species and region of the gut, the composition, size, innervation patterns and connectivity of

internodal strands vary within the two plexi. The human ENS contains approximately 108

neurons, in par with the spinal cord (4).

The neurons and glia of the ENS are derived entirely from the neural crest. Migratory

cells enter the foregut mesenchyme from the vagal axis and colonize the developing gut in a

rostrocaudal fashion within 13 weeks of gestation in humans. Crest-derived cells from the

sacral level also contribute to the innervation of the rectum and hindgut (5, 6). GDNF is the

primary chemoattractant that mediates the migration of neural crest cells within and along

the gut. The development of the ENS and neural crest cell migration are complex and

dynamic processes. Multiple molecular signaling mechanisms involve neurotrophic factors

(GDNF, ET3, and Notch), long-range guidance molecules (Netrins, Semaphorins, Ephrins)

and extracellular matrix components (Laminins, Collagens, Heparan and Chondroitin

Sulphate proteoglycans). These cells differentiate into neurons and glia in the gut

microenvironment to express a full complement of ENS neurotransmitters by 24 weeks of

gestation. The ENS, however, continues to develop and differentiate up to two years post-

natally (7, 8).

2. Disorders of the enteric nervous system and current therapeutic paradigms

Disruption in the integrity of neural circuitry within the gut is described in several

neurodegenerative conditions that result in dysfunctional motility, secretion and

7

inflammation. Gastrointestinal motor function is intimately controlled by the intramural enteric

nervous system. It is a complex interplay between the smooth muscle of the muscularis

externa and the two enteric neuronal plexi (6). Depending on the region of the GI tract

where nerve dysfunction occurs, a wide variety of syndromes can present themselves that

include dysphagia, gastro-esophageal reflux disease, altered gastric emptying, stenosis and

sphincteric hypertrophy, abdominal pain, bloating, diarrhea, constipation, fecal incontinence,

megacolon, etc.

Achalasia was originally characterized as a failure of relaxation of the lower

esophageal sphincter (LES), clinically presenting itself as dysphagia and regurgitation. The

main pathogeny is the absence of intrinsic inhibitory neurons in the esophagus and the LES

but the preservation of cholinergic excitatory innervation, leading to tonic contraction and a

lack of transient relaxation (9, 10). In achalasia of the Internal Anal Sphincter (IAS), the

rectoanal inhibitory reflex in response to rectal distention is markedly absent. Absence of

nitrergic neurons by NADPH diaphorase activity has been reported in cases of Internal Anal

Sphincter (IAS) achalasia (11). Similar to achalasia, congenital hypertrophic pyloric stenosis

demonstrates an inhibitory denervation of the pylorus leading to gastric obstruction. Tonic

contraction maintained constantly evokes hypertrophy of the smooth muscle of the pylorus.

Sphincteric achalasia has been treated with intrasphincteric injections of botulinum toxin, to

temporarily inhibit the release of neuronal acetylcholine, thereby allowing transient reduction

in sphincteric tone. This has been reported in both LES and IAS achalasia (12, 13).

Pharmacological options include calcium channel blockers or long acting nitrates to reduce

sphincteric tone. Pneumatic dilation and surgical myotomy are other options for the

treatment of achalasia (14).

Neurogenic chronic intestinal pseudo-obstruction resulting in a derangement of

peristalsis and intestinal failure occurs due to a non-inflammatory degeneration of the ENS.

8

Several putative pathogenic mechanisms are implicated in the loss of intrinsic neurons

(altered calcium signaling, free radicals, etc.) (15). Enteric neuropathy is also secondary to

several other disorders (diabetes, Parkinson’s disease, inflammation) resulting in

gastrointestinal dysfunction (16, 17). Neurodegeneration has been associated with the aging

gut, with an age-related loss of enteric neurons resulting in altered intestinal motility and

gastric emptying (18). Pharmacological treatment is the mainstay of addressing symptoms

of dysmotility in several ENS disorders. Prokinetics are documented to improve intestinal

motility and control visceral sensitivity. Combination with antibiotics help reduce sepsis,

nausea and abdominal pain (19). Surgery is not a recommended option for chronic intestinal

pseudo-obstruction, because only rare cases benefit from surgical resections (15).

Hirschsprung’s disease or congenital megacolon is the best characterized disorder of

the ENS, where enteric neurons are absent in varying lengths of distal gut. Biopsies of

aganglionic segments demonstrate the absence of intrinsic inhibitory neurons leading to a

lack of relaxation of the diseased segment. The aganglionic gut thus remains tonically

contracted, obstructing the passage of intestinal contents leading to megacolon (20). A pull-

through surgery is essentially life-saving in Hirschsprung’s disease, but the surgery can

potentially disrupt anorectal innervation in the remaining ganglionic gut. Recurrent fecal

soiling and fecal incontinence has been reported as a long-term outcome (21). Enterocolitis

is also a serious complication post-surgical correction for Hirschsprung’s owing to the

alterations in the gut microbiome and/or the loss of the intestinal mucosal barrier. It has

been recently established that probiotic prophylaxis does not reduce Hirschsprung’s

associated enterocolitis (22).

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3. Tissue Engineering of the Intestinal Neuromusculature

In the following paragraphs, a current status and perspective of cell-based therapies for GI

disorders is provided. The initial focus is on neural stem cell transplantation, an emerging

and exciting therapeutic paradigm to reinstate neurogenic dysfunction of the intestine.

Several acellular biomaterials based approaches to intestinal regeneration and tissue

engineering have been successfully applied. A future perspective on the use of neural stem

cells in neuronal disorders of the gut is provided. Additionally, several approaches to smooth

muscle replacement to remedy intestinal dysfunction is also tackled in the following

paragraphs.

3.1 Introduction to Neural Stem Cell Transplantation

Neural stem cell therapy is an emerging therapeutic paradigm that ideally aims to

reinstate neuronal function and thus gastrointestinal motor function by repopulating the

enteric plexi. The hope is to restore neuronal function, rather than pharmacologically provide

palliative care. Several factors require to be taken into consideration while designing stem

cell based therapies to remedy aganglionic disorders or neurogenic disorders of the gut. Of

principal importance is the source of stem cells, and if they can be reliably and efficiently

isolated in adequate numbers from adults using minimally invasive procedures. A mode of

delivery needs to be determined to effectively colonize and repopulate dysfunctional enteric

ganglia. Furthermore, long term viability and parameters for successful engraftment and

phenotypic stability need also be contemplated before stem cell therapy can be translated

into the clinic successfully. Finally, in vitro research on phenotypes, trophic support and

directed differentiation could result in therapies designed to provide relief in particular GI

disorders: for example, an enriched population of nitrergic neurons could be used to relieve

10

gastric dysfunction and gastroparesis. Such a population of neurons could also theoretically

relieve sphincteric achalasia.

Neural stem cell transplantation is driven by two significant findings: i) neuroglial

progenitor cells can be isolated from adult mammalian gut, including ganglionated colon of

Hirschsprung’s patients (23, 24); and ii) progenitor cells can be induced to differentiate into

several neuronal subtypes and glia characteristic of the ENS. Several sources of neural

stem and progenitor cells have been explored – including CNS-derived stem cells, ENS-

derived stem cells, bone marrow derived stem cells that differentiate in to a neuronal

lineage, etc. The following paragraphs will expand on the various sources of neuronal stem

cells, their isolation and therapeutic delivery strategies used to transplant these cells in vivo.

4. Sources of smooth muscle and neuronal stem and progenitor cells

4.1 Isolation of smooth muscle stem and progenitor cells

In order to avoid immunosuppressive regimens, autologous stem cell sources are

ideal. Over the years, several tissue-resident organ-associated adult stem cells have been

identified in the human body (25). Bone marrow-derived stem cells have been demonstrated

to assume vascular and bladder smooth muscle phenotypes, and have thus been used in

regenerative medicine applications (26-28). This is an optimistic step towards producing

tissue engineered smooth muscle from autologous bone marrow-derived cells. However,

Hori et al. showed that bone marrow-derived mesenchymal stem cells seeded on collagen

scaffolds did not regenerate a smooth muscle layer upon transplantation in to the canine

intestine (29). Several populations of muscle derived-stem cells with clonal expansion and

self-renewal capabilities have been identified (30, 31). These adult muscle stem cells can

differentiate into myotubes as well as smooth muscle phenotypes (32). Within the post natal

gut, interstitial cells of Cajal have been shown to transdifferentiate into smooth muscle cells

upon blockade of Kit signaling, with increased expression of desmin and smooth muscle

11

myosin (33). Neural crest-derived nestin positive stem cells isolated from post natal murine

gut have also been demonstrated to differentiate into smooth muscle phenotypes (34).

4.2. Isolation of neuronal stem and progenitor cells

4.2.1 ENS as a source of neural crest stem and progenitor cells

The ENS is populated by migrating cells derived from the neural crest. Neural stem

or progenitor cells have been isolated from embryonic, fetal, post-natal and adult rodent

guts. Cells have been isolated from full-thickness gut tissue, mucosal gut biopsies or from

the muscularis externa layers containing the myenteric plexus alone (34-37). Enteric stem

and progenitor cells have also been isolated from human full thickness/muscularis/mucosal

gut biopsies (23, 38-40). Metzger et al. (23) also demonstrate the reliable isolation of enteric

neuronal progenitor cells from adult human gut up to 84 years of age. These cells have the

ability to differentiate into a number of mature enteric neuronal subtypes. Additionally,

neuroglial progenitor cells have also been isolated from ganglionated colons of

Hirschsprung’s patients (24). A variety of techniques have been used to isolate neuronal

stem cells from dissociated gut tissues. Embryonic mouse ENS and neonatal human ENS

derived progenitor cells have been isolated following trypsin-EDTA dissociation of caeca

(24, 38). Collagenase digestion in combination with trypsin has been used to obtain

neuronal progenitor cells from rodent muscularis externa (37). Enzymatic dissociations with

a combination of collagenase/dispase combinations, as well as collagenase XI/dispase have

been used to obtain human enteric neuronal progenitor cells from full thickness or mucosal

biopsies (23, 41). For information on further selection of isolated ENS neural stem cells, see

Table 1 in a review by Kulkarni et al.(42). In a majority of these cases, isolated cells are

cultured in non-adherent cultures, where these cells proliferate and aggregate to form

floating spherical colonies, called neurospheres. This process of the formation of floating

12

colonies of spherical aggregates has almost become a surrogate marker for “stemness” of

neural stem cells.

Neural crest derived enteric neuronal progenitor cells are positive for p75NTR and

have been immunoselected based on the expression of this low-affinity NGF receptor (23,

24, 38). Nestin, an intermediate filament protein expressed by neuroepithelial stem cells, is

also expressed in enteric neuronal progenitor cells isolated from rodent myenteric plexus

(36, 37, 43). Sox2 (SRY-related HMG transcription factor 2) is a marker of ENS progenitor

cells and glial derivates, known to be expressed in embryonic and adult rodent gut (44).

Sox10 is another Sox transcription factor that is vital to the maintenance of progenitor status

of these cells, so much so that normal Sox10 activity and endothelin signaling is essential

for retrieval of sufficient numbers of enteric neuronal progenitor cells and their ability to form

neurospheres. Mutations in the Sox10 gene has been implicated in the pathogenesis of

congenital megacolon associated with Hirschsprung’s disease (45, 46). Sox10 expression

is also an indicator of gliogenic potential of neural crest derived progenitor cells and is

important for glial fate acquisition(47).

4.2.2 Non-ENS sources of neural crest stem and progenitor cells

The first report of obtaining neural crest derivatives (peripheral sensory and

sympathetic neurons) from human embryonic stem cells was by Pomp et al. (48). Lee et al.

demonstrated the derivation of neural crest stem cells from human embryonic stem cells that

were capable of migration and differentiation akin to neural crest identity (49). Human

embryonic stem cell derived neuroepithelial stem cells have been transplanted into specific

regions of the rodent brain, where they demonstrated capabilities of axonal growth and

pathfinding (50). Induced pluripotent stem cells offer an alternative, keeping in mind the

ethical considerations of the use of human embryonic tissues. Through extensive genetic

reprogramming, skin fibroblasts are routinely converted into a pluripotent phenotype.

13

Directed differentiation of induced pluripotent stem cells in to multipotent neural crest stem

cells with high efficiency, under defined media and feeder-free cultures is now well

established (51). However, induced pluripotent stem cells still retain a risk for tumorigenicity

and their therapeutic potential is severely limited by the use of retroviral transductions to

derive them.

Post-migratory neural crest stem cells have been isolated from adult rodents from

several distinct regions including the gut, dorsal root ganglia, sciatic nerve, and even bone

marrow (52). Recently, however, neural crest stem cells isolated from the bone marrow have

been demonstrated to undergo in vivo tumorigenesis (53).

Neurogenesis has now been known to occur even in post-natal and adult

mammalian brain (54). Neural crest stem cells persist in restricted regions of adult brains,

including the subventricular zone and the subgranular layer of the dentate gyrus and the

external germinal layer of the cerebellum (55). Similar to ENS-derived neurospheres, adult

CNS-derived neural crest stem cells can be propagated in long term cultures in the

presence of epidermal and fibroblast growth factors. These cells retain their self-renewal

capacity and multi-potency (56).

CNS-derived neurospheres are often isolated from the subventricular zone of the

rodent fetal brain. It has been demonstrated that these NSCs can be nudged into enteric-like

neuronal subtypes when exposed to the gut microenvironment and gut derived soluble

factors, making them a candidate for transplantation (57). Therefore, CNS-derived neural

crest stem cells are a good alternative to ENS-derived stem cells. Additionally, it is also

hypothesized that adult neural crest stem cells from various sources retain enough plasticity

to assume ENS specific phenotypes and behaviour in response to the gut

microenvironment. A schematic of the various sources of neuronal progenitor cells derived

into enteric neurons/glia is shown in Figure 1.

14

Figure 1: Schematic of the various sources of neural stem/progenitor cells

Enteric neurons and glia can be derived from various sources of stem and progenitor cells arising from different parts of the body. Adult CNS-derived as well as ENS-derived neural stem cells have been transplanted in to mammalian gut, and have demonstrated differentiation into enteric neurons, enteric glia and smooth muscle cells.

5. A summary of delivery strategies for neural stem cells in the GI tract

Embryonic CNS-derived neural stem cells were injected bi-laterally into the mid-

pylorus of nNOS-/- mice. Grafted NSCs differentiated into neurons and glia and improved

gastric emptying of liquids and improved the relaxation of the pyloric sphincter (58). Fetal

and postnatal ENS-derived NSCs were injected into the external muscle layer of the distal

colon of wild-type mice. The grafted cells demonstrated several enteric neuron subtypes

including nNOS and choline acetyl transferase (59). Post-natal human enteric

15

neurospheres were transplanted into aganglionic mouse distal hindgut where they migrated

through pathways similar to neural crest cells developmentally. Furthermore, these cells

differentiated into NOS and VIP neurons and glia (24). Neuroepithelial cells were isolated

from the embryonic rodent neural tube and injected into chemically denervated distal colons

of rats. Grafted neuroepithelial stem cells were shown to differentiate into neurons and glia.

Intraluminal pressure readings indicated an inflation stimulated contraction as well as

electrical field stimulated responses (60). Post-natal human enteric neurospheres obtained

from intestinal mucosal biopsies were injected into explant cultures of human aganglionic

gut or chick embryos. Grafted cells colonized aganglionic chick or human hindguts and

generated ganglia-like structures with neurons and glia (39). A recent study by Hagl et al.

demonstrates that the microenvironment and smooth musculature of the aganglionic bowel

of HSCR patients supports neural stem cell differentiation (61), providing promise for the

development of neural stem cell transplantation as a therapeutic tool.

6. Smooth muscle intestinal tissue engineering

Several approaches have been used to re-engineer intestinal smooth musculature

(either sphincteric or non-sphincteric). Conventional tissue engineering approaches initially

used inert and/or prosthetic materials to replace a missing wall defect or conduit. Since then,

much progress has been made towards the use of biocompatible materials that allow

cellular infiltration, extensive remodeling and adsorption. In addition to a significant

repopulation of constituent cell types, ideal functional tissue engineering must also reinstate

peristalsis, motility and absorption/secretion. This requires functional smooth muscle cells

and epithelial cells in addition to a functional neuronal network.

Several synthetic polymeric combinations have been demonstrated to be

biocompatible, and capable of muscular ingrowth. Additionally, naturally occuring

extracellular matrix-derived polymers have also been used to reconstruct segments of the GI

16

tract. Autologous neo-esophagus constructs have been engineered using human

esophageal epithelial cells, aortic smooth muscle cells and dermal fibroblasts embedded

within human tendon collagen or polyglycolic acid meshes (62, 63). Saxena et al. used

basement membrane matrix coated scaffolds that demonstrated muscular ingrowth of the

smooth musculature as well as alignment and polarity of epithelial cells of the esophagus

(64).

Several mechanical devices were used to control reflux and GERD due to sphincteric

inadequacy, i.e. the lack of a sufficient high pressure from the lower esophageal sphincter.

To date, the only cell-based regenerative medicine approach in repairing defects of the LES

remains the injection of skeletal muscle-derived stem cells into the LES to augment

sphincteric pressure in canine models. While these cells integrated within the underlying

smooth musculature, there was no demonstration of a contractile smooth muscle phenotype

(65).

Collagen sponge scaffolds were used to repair gastric wall defects in which proton

pump positive cells were regenerated (66). Similar sponge scaffolds in combination with

synthetic biodegradable polymers have been seeded with autologous bone marrow

mesenchymal stem cells to repair gastric wall defects (67). Gastric epithelial organoid units

have been seeded on composite PLGA meshes to replace native stomach of rats (68). The

only regenerative medicine based approach to repair defects of the pyloric sphincter is the

injection of CNS-derived stem cells to reinstate nitric oxide neuronal function by Micci et al.

(58).

Several studies have been published to date, reporting tissue engineering of the

small and large intestines. Tissue engineering has offered an elegant solution to the bowel

lengthening surgeries commonly carried out in short bowel syndrome. Autologous smooth

17

muscle cells seeded within collagen sponge scaffolds have been used to successfully

regenerate villi structures of the small intestine (69). Tubular structure based approaches

have also had some limited success in replacing the smooth musculature of the small

intestine (70). (71, 72). Small intestinal submucosal ECM based scaffolds have been used

to replace tubular segments of bowel, where enteric neuronal plexuses were regenerated

and et basic physiological demands in a canine model (73).

Intestinal organoid units generated from sigmoid colon have been used to tissue

engineer tubular colonic structures (74). Such tissue engineered constructs have

demonstrated a significant absorptive capacity when implanted. Fibrin-based hydrogels

were used to generate three-dimensional concentrically aligned structures of circular smooth

muscle layers of the colon (75). Tissue engineered models of the longitudinal layer of

colonic smooth muscle has also been demonstrated (76). Individually, these structures

mimic native smooth muscle alignment and maintain aspects of colonic physiology like

peristalsis and response to contractile and relaxant neurotransmitters. Recently, the use of

chitosan to functionally support such tissue engineered colonic constructs for intestinal

tissue engineering was demonstrated (77).

Autologous skeletal-muscle derived stem cells were used to augment sphincteric

function of the internal anal sphincter as well. However, no evidence of the development of

functional smooth muscle was reported (78). Physiologically functional tissue engineered

constructs of rodent and human internal anal sphincters have been developed, that

demonstrate the spontaneous generation of myogenic basal tone (79-81). Pre-wired

bioengineered IAS constructs that included a functional innervation have also been

bioengineered. These constructs have been implanted into athymic rodents, where they

preserved neuronal networking as well as myogenic functionality (82).

18

7. Future perspectives

However, phenotypic stability, long term survival, and post-transplant fate all remain

to be optimized while moving forward with neural stem cell transplantation for clinical use

(42, 43). In order to provide trophic support and a permissive microenvironment, a more

fundamental understanding of factors that affect and maintain differentiation of enteric

neuronal progenitor cells is required.

As is true for all stem cells, their behavior and differentiation depends on their

location, microenvironment, and the establishment or reestablishment of interactions with

the extracellular matrix. Alternative to direct neural stem cell transplantation, grafting of

differentiated cells or progenitor cells committed to restricted phenotypes must be

considered. This is a potential work around to avoid differentiation in to undesirable

phenotypes. Additionally, in vitro differentiation may allow tailoring of cell-based

transplantation therapies to specific neurodegenerative diseases, for example, supplying

nitrergic neurons for sphincteric achalasia. Another area that needs intense focus is the

identification of trophic support required for the survival of transplanted stem and/or

progenitor cells. Parallel to this is the development of imaging modalities that non-invasively

track or image transplanted cells. Magnetic resonance imaging (MRI) offers a non-invasive

method to image, but labeling transplanted cells with contrast agents or small paramagnetic

iron oxide particles require cytotoxicity studies. Of paramount importance with cellular

labeling is that the biological activity of the stem/progenitor cell is not altered and their

differentiation capability is not affected. The complex interaction between the host

microenvironment and transplanted cells also needs to be considered, in order to improve

survival and for cell fate determination. Biomaterial based delivery can be used to provide

trophic support to transplanted neural stem cells or to manipulate host inflammatory

response. Extracellular matrix based hydrogel environments can also be utilized to provide a

19

protective microenvironment to transplanted cells. ECM molecules typical of the developing

gut, similar to the developing mesenchyme, may provide a microenvironment conducive to

sequestration and presentation of trophic factors like glial-cell derived neurotrophic factor, to

influence neuronal cell survival. Lastly, as a cautionary tale from clinical studies of

transplanting neural stem cells in several CNS-based neurological disorders, fundamental

research on molecular bases of therapeutic plasticity of neural stem cells is important.

Transplantation in to animal models of GI disorders will offer a rationale for the design of

future clinical studies.

8. Acknowledgements

This work was supported by NIH RO1 DK 071614 and RO1 DK 057020.

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58. Micci, M. A., Kahrig, K. M., Simmons, R. S., Sarna, S. K., Espejo-Navarro, M. R., and Pasricha, P. J. (2005) Neural stem cell transplantation in the stomach rescues gastric function in neuronal nitric oxide synthase-deficient mice, Gastroenterology 129, 1817-1824.

59. Hotta, R., Stamp, L. A., Foong, J. P. P., McConnell, S. N., Bergner, A. J., Anderson, R. B., Enomoto, H., Newgreen, D. F., Obermayr, F., Furness, J. B., and Young, H. M. (2013) Transplanted progenitors generate functional enteric neurons in the postnatal colon, The Journal of Clinical Investigation 0, 0-0.

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60. Liu, W., Wu, R. D., Dong, Y. L., and Gao, Y. M. (2007) Neuroepithelial stem cells differentiate into neuronal phenotypes and improve intestinal motility recovery after transplantation in the aganglionic colon of the rat, Neurogastroenterol Motil 19, 1001-1009.

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62. Hayashi, K., Ando, N., Ozawa, S., Kitagawa, Y., Miki, H., Sato, M., and Kitajima, M. (2004) A neo-esophagus reconstructed by cultured human esophageal epithelial cells, smooth muscle cells, fibroblasts, and collagen, ASAIO J 50, 261-266.

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73. Chen, M. K., and Badylak, S. F. (2001) Small bowel tissue engineering using small intestinal submucosa as a scaffold, J Surg Res 99, 352-358.

74. Grikscheit, T. C., Ochoa, E. R., Ramsanahie, A., Alsberg, E., Mooney, D., Whang, E. E., and Vacanti, J. P. (2003) Tissue-engineered large intestine resembles native colon with appropriate in vitro physiology and architecture, Ann Surg 238, 35-41.

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75. Hecker, L., Baar, K., Dennis, R. G., and Bitar, K. N. (2005) Development of a three-dimensional physiological model of the internal anal sphincter bioengineered in vitro from isolated smooth muscle cells, Am J Physiol Gastrointest Liver Physiol 289, G188-196.

76. Raghavan, S., Lam, M. T., Foster, L. L., Gilmont, R. R., Somara, S., Takayama, S., and Bitar, K. N. (2010) Bioengineered three-dimensional physiological model of colonic longitudinal smooth muscle in vitro, Tissue Eng Part C Methods 16, 999-1009.

77. Zakhem, E., Raghavan, S., Gilmont, R. R., and Bitar, K. N. (2012) Chitosan-based scaffolds for the support of smooth muscle constructs in intestinal tissue engineering, Biomaterials 33, 4810-4817.

78. Kang, S. B., Lee, H. N., Lee, J. Y., Park, J. S., and Lee, H. S. (2008) Sphincter contractility after muscle-derived stem cells autograft into the cryoinjured anal sphincters of rats, Dis Colon Rectum 51, 1367-1373.

79. Singh, J., and Rattan, S. (2012) Bioengineered human IAS reconstructs with functional and molecular properties similar to intact IAS, Am J Physiol Gastrointest Liver Physiol 303, G713-722.

80. Somara, S., Gilmont, R. R., Dennis, R. G., and Bitar, K. N. (2009) Bioengineered internal anal sphincter derived from isolated human internal anal sphincter smooth muscle cells, Gastroenterology 137, 53-61.

81. Raghavan, S., Miyasaka, E. A., Hashish, M., Somara, S., Gilmont, R. R., Teitelbaum, D. H., and Bitar, K. N. (2010) Successful implantation of physiologically functional bioengineered mouse internal anal sphincter, Am J Physiol Gastrointest Liver Physiol 299, G430-439.

82. Raghavan, S., Gilmont, R. R., Miyasaka, E. A., Somara, S., Srinivasan, S., Teitelbaum, D. H., and Bitar, K. N. (2011) Successful implantation of bioengineered, intrinsically innervated, human internal anal sphincter, Gastroenterology 141, 310-319.

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CHAPTER 3: SPECIFIC AIMS AND OBJECTIVES OF THE THESIS

The overarching hypothesis of this research was that extracellular matrix (ECM)

based microenvironments will influence neuroglial differentiation.

1. Role of the extracellular matrix in the developing ENS

The extracellular matrix plays an important role in the migration and development of

the enteric nervous system [1, 2]. Alterations in the extracellular matrix of the gut

mesenchyme have been documented in aganglionic regions of rodent gut, suggesting the

importance of a permissive extracellular environment to promote effective colonization and

differentiation of neural crest cells in the developing gut [2-5]. Laminin, and proteoglycans

are expressed within the embyonic gut to aid the colonization by vagal neural crest cells.

Collagen IV is distributed in the developing nervous system along the neural crest. It

encapsulates the adult myenteric and submucosal plexi in the mammalian gut [6]. Laminin

influences neuronal adhesion and neurite outgrowth through different peptides regions

within the laminin molecule [7]. In the gut, heparan sulfate proteoglycan is required for

GDNF signaling [8]. Glycosaminoglycan – growth factor interactions stabilize and increase

local bioavailability of growth factors, effectively inducing neurite outgrowth [9].

Several studies in the literature emphasize the role of utilizing extracellular matrix

based niches to control CNS-derived neural stem cell fate in vitro [10-12]. Preliminary

studies also demonstrate that the ECM microenvironment may play an important role in the

differentiation of post-natal ENS-derived neural stem- and progenitor cells [13, 14]. This

dissertation investigates the hypothesis that the developmental role played by ECM niches

in generating the diversity of the ENS translates to influencing differentiation of post-natal

ENS-derived progenitor cells in vitro.

26

2. Role of the extracellular matrix in the adult ENS microenvironment

The adult mammalian myenteric plexus is surrounded by an extracellular matrix

primarily composed of Collagen IV, Laminin and a heparan sulfate proteoglycan, with enteric

glia always in direct contact with the ECM. Enteric neurons also come in direct contact with

this ECM, though much less frequently than glia [6, 15]. Thus, the central hypothesis of this

dissertation is that the ECM can provide a background upon which cell-cell and cell-matrix

signaling can work to regulate phenotypes of differentiating enteric neuronal progenitor cells.

The research outlined in this proposal systematically evaluates the effect of

modulating culture substrata with neural ECM components like Collagen IV, Laminin and

heparan sulfate. Collagen I is additionally evaluated, owing to its favorable biological

properties and widespread use in neural tissue engineering [16-18]. Several nerve guidance

conduits using type I collagen based matrices to promote axonal regeneration are now

approved by the FDA [19].

3. Role of two-dimensional ECM culture substrata on neuroglial differentiation

Neuroglial differentiation was characterized by varying two dimensional culture

substrata with varying ECM components, individually or in combination. Through the studies

described within this specific aim, it was established that ECM composition could result in a

pre-ponderance of either glia or neurons. Neuronal and glial differentiation was often

complementary. The number of neurons, neurite lengths and preliminary neuronal network

formation were all enhanced in culture substrata that contained collagen IV, laminin and

heparan sulfate (Aim 1; Chapter 4).

27

4. Role of three-dimensional ECM microenvironments on neural subtype

specification

Tissue engineering offers an elegant tool to translate two-dimensional results into

three-dimensional microenvironments with relevant structural, architectural and rheological

cues. Here, a modified tissue engineered model (Chapter 5) of intestinal longitudinal

smooth muscle was utilized, to evaluate neuronal differentiation as a function of ECM

composition. Given that the cues for differentiation arising from a contractile phenotype of

smooth muscle cells would be constant, we hypothesized that ECM composition could

differentially regulate the generation of specific neuronal subsets. The objective was to

determine the presence of differentiated motor neurons obtained and investigate their ability

in mediating smooth muscle contraction/relaxation.

Through this experimental paradigm, differentiated neuronal subtypes were

identified. It was further established that ECM composition promoted the differentiation of

specific neuronal subtypes, resulting in enriched populations of excitatory or inhibitory motor

neurons. The ability of these differentiated neurons in mediating smooth muscle contraction

and relaxation was evaluated using real-time force generation measurements with

appropriate physiological stimuli and pharmacological inhibitors. The studies performed as

a part of this specific aim generated transplantable tissue engineered innervated intestinal

longitudinal sheets, with varying constituent neural subtypes ranging from highly cholinergic,

to highly nitrergic neurons (Aim 2; Chapter 6).

5. Neo-innervation potential of tissue engineered innervated intestinal sheets

Finally, the neo-innervation potential of enteric neuronal progenitor cells was

evaluated using a regenerative medicine approach. Organotypic cultures of experimentally

denervated gut were established as “recipient” tissues. Tissue engineered sheets

28

manufactured from composite collagen I/collagen IV to generate an enriched population of

nitrergic neurons. These tissue engineered innervated sheets were used as ‘donor’ tissues,

to supply neo-innervation to experimentally denervated colonic explants. Functional nitrergic

denervation and subsequent neo-innervation were investigated. These studies provided a

preliminary therapeutic potential for tissue engineered innervated sheets in their use for

replenishing specific neural subsets in intestinal dysganglionosis (Aim 3; Chapter 7).

The thesis concludes with a perspective on future directions of the research

(Chapter 8) and a summary of the dissertation (Chapter 9).

References

[1] Adams JC, Watt FM. Regulation of development and differentiation by the extracellular matrix. Development. 1993;117:1183-98. [2] Henderson DJ, Copp AJ. Role of the extracellular matrix in neural crest cell migration. J Anat. 1997;191 ( Pt 4):507-15. [3] Young HM, Anderson RB, Anderson CR. Guidance cues involved in the development of the peripheral autonomic nervous system. Auton Neurosci. 2004;112:1-14. [4] Parikh DH, Leibl M, Tam PK, Edgar D. Abnormal expression and distribution of nidogen in Hirschsprung's disease. J Pediatr Surg. 1995;30:1687-93. [5] Gershon MD. Transplanting the enteric nervous system: a step closer to treatment for aganglionosis. Gut. 2007;56:459-61. [6] Bannerman PG, Mirsky R, Jessen KR, Timpl R, Duance VC. Light microscopic immunolocalization of laminin, type IV collagen, nidogen, heparan sulphate proteoglycan and fibronectin in the enteric nervous system of rat and guinea pig. J Neurocytol. 1986;15:733-43. [7] Pomeranz HD, Sherman DL, Smalheiser NR, Tennyson VM, Gershon MD. Expression of a neurally related laminin binding protein by neural crest-derived cells that colonize the gut: relationship to the formation of enteric ganglia. J Comp Neurol. 1991;313:625-42. [8] Barnett MW, Fisher CE, Perona-Wright G, Davies JA. Signalling by glial cell line-derived neurotrophic factor (GDNF) requires heparan sulphate glycosaminoglycan. J Cell Sci. 2002;115:4495-503. [9] Hynes RO. The extracellular matrix: not just pretty fibrils. Science. 2009;326:1216-9. [10] Teixeira AI, Duckworth JK, Hermanson O. Getting the right stuff: controlling neural stem cell state and fate in vivo and in vitro with biomaterials. Cell Res. 2007;17:56-61. [11] Doetsch F. A niche for adult neural stem cells. Curr Opin Genet Dev. 2003;13:543-50. [12] Han S, Yang K, Shin Y, Lee JS, Kamm RD, Chung S, et al. Three-dimensional extracellular matrix-mediated neural stem cell differentiation in a microfluidic device. Lab Chip. 2012;12:2305-8. [13] Rauch U, Schafer KH. The extracellular matrix and its role in cell migration and development of the enteric nervous system. Eur J Pediatr Surg. 2003;13:158-62. [14] Schafer KH, Hagl CI, Rauch U. Differentiation of neurospheres from the enteric nervous system. Pediatr Surg Int. 2003;19:340-4.

29

[15] Gabella G. Innervation of the intestinal muscular coat. J Neurocytol. 1972;1:341-62. [16] Stang F, Fansa H, Wolf G, Reppin M, Keilhoff G. Structural parameters of collagen nerve grafts influence peripheral nerve regeneration. Biomaterials. 2005;26:3083-91. [17] Midha R, Shoichet MS, Dalton PD, Cao X, Munro CA, Noble J, et al. Tissue engineered alternatives to nerve transplantation for repair of peripheral nervous system injuries. Transplant Proc. 2001;33:612-5. [18] Dewitt DD, Kaszuba SN, Thompson DM, Stegemann JP. Collagen I-matrigel scaffolds for enhanced Schwann cell survival and control of three-dimensional cell morphology. Tissue Eng Part A. 2009;15:2785-93. [19] Kehoe S, Zhang XF, Boyd D. FDA approved guidance conduits and wraps for peripheral nerve injury: a review of materials and efficacy. Injury. 2012;43:553-72.

30

CHAPTER 4: NEUROGLIAL DIFFERENTIATION OF ADULT

ENTERIC NEURONAL PROGENITOR CELLS AS A FUNCTION OF

EXTRACELLULAR MATRIX COMPOSITION

Shreya Raghavan1,2, Robert R. Gilmont1, Khalil N. Bitar1,2

1Wake Forest Institute for Regenerative Medicine, Wake Forest School of Medicine, Winston-Salem NC 27101

2Virginia Tech-Wake Forest School of Biomedical Engineering and Sciences, Winston-Salem NC 27101

Keywords: Neural stem cell, neural tissue engineering, extracellular matrix, adult stem cell, enteric glia

This manuscript was accepted for publication in Biomaterials 2013, 34(28): 6649-6658;

31

ABSTRACT

Enteric neuronal progenitor cells are neural crest-derived stem cells that can be isolated

from fetal, post-natal and adult gut. Neural stem cell transplantation is an emerging

therapeutic paradigm to replace dysfunctional or lost enteric neurons in several aganglionic

disorders of the GI tract. The impetus to identify an appropriate microenvironment for enteric

neuronal progenitor cells derives from the need to improve survival and phenotypic stability

following implantation. Extracellular matrix composition can modulate stem cell fate and

direct differentiation. Adult mammalian myenteric ganglia in vivo are surrounded by a matrix

composed primarily of Collagen IV, Laminin and a Heparan sulfate proteoglycan. In these

studies, adult mammalian enteric neuronal progenitor cells isolated from full thickness rabbit

intestines were induced to differentiate when cultured on various combinations of neural

ECM substrates. Neuronal and glial differentiation was studied as a function of ECM

composition on coated glass coverslips. Poly- lysine coated coverslips (control) supported

extensive glial differentiation but very minimal neuronal differentiation. Individual culture

substrata (Laminin, Collagen I and Collagen IV) were conducive for both neuronal and glial

differentiation. The addition of laminin or heparan sulfate to collagen substrates improved

neuronal differentiation, significantly increased neurite lengths, branching and initiation of

neuronal network formation. Glial differentiation was extensive on control poly lysine coated

coverslips. Addition of laminin or heparan sulfate to composite collagen substrates

significantly reduced glial immunofluorescence. Various neural ECM components were

evaluated individually and in combination to study their effect of neuroglial differentiation of

adult enteric neuronal progenitor cells. Our results indicate that specific ECM substrates that

include type IV Collagen, laminin and heparan sulfate support and maintain neuronal and

glial differentiation to different extents. Here, we identify a matrix composition optimized to

tissue engineer transplantable innervated GI smooth muscle constructs to remedy

aganglionic disorders.

32

1. Introduction

Gastrointestinal motor function is intimately controlled by the intramural enteric

nervous system. It is a complex interplay between the smooth muscle of the muscularis

externa and the two enteric neuronal plexi [1]. Aganglionosis of varying lengths of distal gut

is the central pathology in Hirschsprung’s disease [2]. Enteric neuropathy is also secondary

to several other disorders (diabetes, Parkinson’s disease, inflammation) resulting in

gastrointestinal dysfunction [3, 4]. Neural stem cell therapy is an emerging therapeutic

paradigm that ideally aims to reinstate neuronal function and thus gastrointestinal motor

function by repopulating the enteric plexi. The research is driven by two significant findings:

i) neuroglial progenitor cells can be isolated from adult mammalian gut, including

ganglionated colon of Hirschsprung’s patients [5, 6]; and ii) progenitor cells can be induced

to differentiate into several neuronal subtypes and glia characteristic of the ENS upon

transplantation into explant cultures of aganglionic/aneural gut [5-9], or in vivo into distal

colo-rectums of adult rodents [10, 11]. However, phenotypic stability, long term survival, and

post-transplant fate all remain to be optimized while moving forward with neural stem cell

transplantation for clinical use [12, 13]. In order to provide trophic support and a permissive

microenvironment, a more fundamental understanding of factors that affect and maintain

differentiation of enteric neuronal progenitor cells is required. The studies described here

focus on in vitro differentiation of adult mammalian enteric neuronal progenitor cells,

particularly related to the effect of varying extracellular matrix composition of culture

substrata.

The extracellular matrix (ECM) plays an enormous role in dictating stem cell fate.

ECM composition, structure and mechanical properties can all modulate progenitor cell

differentiation [14, 15]. The adult mammalian myenteric plexus is surrounded by an

extracellular matrix primarily composed of Collagen IV, Laminin and a heparan sulfate

33

proteoglycan, with enteric glia always in direct contact with the ECM. Enteric neurons also

come in direct contact with this ECM, though much less frequently than glia [16, 17].

Laminin, fibronectin and proteoglycans are expressed within the embyonic gut to aid its

colonization by vagal neural crest cells. Collagen IV is distributed in the developing nervous

system along the neural crest. Additionally, laminin is implicated in both the central and

peripheral nervous system in promoting neural cell adhesion and axonal outgrowth [18].

Heparan sulfate is required for GDNF signaling in the gut, and has been known to stabilize

and influence neuronal differentiation in vitro [19-21].

In this paper, we describe the effect of components of neural ECM on the

differentiation of gut-derived neuronal progenitor cells of neural-crest lineage in vitro. Two

timepoints were defined to identify early and late differentiation events – day 5 (early) and

day 15 (late) based on previous experiments [11]. Immunohistochemistry for β III Tubulin

(neuron specific microtubule) and GFAP (Glial fibrillary acidic protein) was used to identify

differentiated neurons and glia on coated culture substrata. The principal objective of this

study was to identify the effect of extracellular matrix composition on the differentiation of

adult enteric neuronal progenitor cells in vitro.

2. Materials and Methods

2.1 Reagents

All tissue culture reagents were purchased from Invitrogen (Carlsbad, CA) unless specified

otherwise. Primary and fluorophore conjugated secondary antibodies were purchased from

Abcam (Cambridge, MA). Rat tail type I Collagen and natural mouse type IV Collagen were

purchased from BD Biosciences (Bedford, MA) and Laminin was from Invitrogen (Carlsbad,

CA). Heparan sulfate was purchased from Celsus Labs (Cincinnati, OH).

34

2.2 Isolation and culture of rabbit enteric neuronal progenitor cells and rabbit intestinal

smooth muscle cells.

New Zealand white rabbits were euthanized using ketamine/xylazine. Intestinal smooth

muscle cells were isolated and cultured using previously described protocols [22]. For the

isolation of enteric neuronal progenitor cells, 5 cm2 biopsies were dissected from the

jejunum, and retrieved in Hank’s Buffered Salt Solution (HBSS) with 2X

antibiotics/antimycotics and 1X gentamicin sulfate. Luminal content was cleaned and tissues

were washed extensively with HBSS. Enteric neuronal progenitor cells were isolated from

these tissues using a collagenase/dispase digestion method, reported by Almond et al. [6].

Cells were plated on to bacterial petri dishes in neuronal growth media (Neurobasal + 1X N2

supplement + 1X antibiotics) following filtration through a 40µm mesh.

2.3 Immunohistochemical characterization of rabbit enteric neurospheres

In order to characterize the initial phenotype of rabbit enteric neurospheres in culture,

neurospheres were harvested by centrifugation at 1000g for 10 minutes in microfuge tubes.

The growth media was gently aspirated, and neurospheres were fixed with 3.7% neutral

buffered formaldehyde and blocked with 10% horse serum. Primary antibodies for p75

(Millipore, Billerica MA), Sox2 and Nestin were incubated for 30 minutes at room

temperature. Unbound antibody was washed using phosphate buffered saline (PBS), and

appropriate fluorophore-conjugated secondary antibodies were incubated for an additional

30 minutes. Neurospheres were mounted using Prolong Gold antifade mounting medium

(Invitrogen, Carlsbad CA), and visualized using an inverted Nikon TiE fluorescent

microscope.

2.4 Differentiation of rabbit enteric neurosphere differentiation as a function of extracellular

matrix composition

35

22x11mm coverslips were washed in Neutrad (Decon Labs, King of Prussia PA) and rinsed

extensively in deionized water. Coverslips were sterilized by 70% ethanol, and subsequent

UV exposure for 45 minutes. Coverslips were coated with poly-L-lysine (pLL; 1 mg/ml), pLL+

10µg/cm2 type I Collagen, pLL + 10µg/cm2 type IV Collagen or pLL + 10µg/cm2 Laminin.

Composite coatings included:

i) 5µg/cm2 Collagen I + 5 µg/cm2 type IV Collagen;

ii) 5 µg/cm2 Collagen I + 5 or 10 µg/cm2 Laminin;

iii) 5 µg/cm2 Collagen IV + 5 or 10 µg/cm2 Laminin;

iv) 5 µg/cm2 Collagen I + 5 µg/cm2 Collagen IV + 0.1 µg/cm2 Heparan Sulfate (HS);

v) 5 µg/cm2 Collagen I + 5µg/cm2 Collagen IV + 5 µg/cm2 Laminin + 0.1 µg/cm2 HS.

Uncoated glass coverslips were seeded with rabbit colonic smooth muscle cells, and

allowed to reach confluence. Rabbit enteric neurospheres were harvested and treated with

Accutase to obtain a mixture of single cells as well as small neurospheres. 10,000 neuronal

progenitor cells were harvested and plated on to coated coverslips. To stimulate

differentiation induced via soluble smooth muscle factors, each plate was shared by one

confluent smooth muscle coverslip along with a coated coverslip containing adhered

neurospheres. Enteric neurospheres were allowed to differentiate for a period of fifteen

days, with a supplementation of neuronal differentiation medium every 2 days (Neurobasal-

A medium + 1X B27 supplement + 2% fetal calf serum + 1X antibiotics).

2.5 Immunohistochemical analysis of neuronal and glial differentiation of rabbit enteric

neurospheres on coated coverslips

Neuronal and glial differentiation was analyzed at two time points– Day 5 and Day 15 post

initiation of differentiation. Medium was aspirated and cells on coverslips were fixed with 3.7

neutral buffered formaldehyde. Cells were permeabilized with 0.15% Triton-X 100 and

36

blocked with 10% horse serum. βIII Tubulin was used to stain neuronal cells, and glial

fibrillary acidic protein (GFAP) was used to stain glial cells. Primary antibodies were

incubated for 1 hour at room temperature and unbound antibody was washed with PBS.

Fluorophore conjugated secondary antibodies (FITC-anti mouse and TRITC-anti rabbit)

were used to visualize fluorescence using an inverted Nikon TiE fluorescent microscope.

Staining with FITC-conjugated secondary antibody without the primary antibody was used

as a negative control. Confluent smooth muscle coverslips were stained with neuronal or

glial markers to avoid a false positive staining while identifying differentiated neurons or glia.

2.6 Data Analysis

Neurite lengths were measured from individual 10X micrographs obtained at the same

amplifier gain and exposure. Neurites were identified primarily by expression of

immunoreactivity for βIII Tubulin concurrently with neuronal morphology. Up to five

sequential fields of view were measured on each coverslip starting from one edge to the

other, covering the area of the coverslip. All cells were measured on each coverslip,

covering the entire area of the neuronal coverslip. Number of neurites measured for each

coverslip coating varied between 20-50 readings and is indicated next to a reported

measurement. The length of the longest neurite from each cell was measured using NIH

Image J using the freeform tool. Neurite lengths between coatings were compared using

one way ANOVA, with Bonferroni post-test to identify a significant difference (p<0.05) in

neurite lengths by varying culture substrata. GFAP immunofluorescence was quantified

using the Nikon Elements imaging software. Mean red (TRITC) fluorescence was calculated

from 10X micrographs, using a constant rectangular area tool that covered 100% of the field

of view. Multiple (at least 5) sequential fields of view at the same magnification were chosen

for each sample to obtain mean fluorescence. Mean red fluorescence indicated the

presence of GFAP expressing cells. One way ANOVA with Bonferroni post-test was used to

37

identify a significant difference in red fluorescent intensity between coated culture substrata.

GraphPad Prism 5.1 for Windows (San Diego, CA) was used to perform statistical analysis.

All statistics are from experiments between 3-5 individual sets, with multiple micrographs

within each set. Reported numbers are mean ± standard error of the mean.

3. Results

3.1 Initial phenotype of rabbit enteric neurospheres

Figure 2: Rabbit enteric neurospheres

(A) Phase contrast micrograph of rabbit enteric neurospheres in culture. Upon primary isolation and culture, progenitor cells proliferate and aggregate to form neurosphere-like bodies (enteric neurospheres). Immunohistochemistry for initial phenotype (B-D): Rabbit enteric neurospheres are p75NTR (A), Sox2 (B) and Nestin (C) positive – indicating that they are comprised of neural crest-derived neuronal and glial progenitor cells. Scale bar 100µm.

Upon digestion of rabbit jejunal biopsies with collagenase/dispase, near single cell

suspensions were obtained by filtration through 70µm and 40µm meshes. Single cells were

approximately 7µm in diameter. These cells were plated in non-adherent culture dishes.

Over the course of two weeks post plating, rabbit enteric neuronal progenitor cells

aggregated and proliferated in culture and formed floating spherical structures, called enteric

neurospheres (Figure 2A). Average neurospheres were 98.2 ± 8.3 µm (n=34) two weeks

post plating. The neurospheres continued to grow and aggregate, approaching 200-300µm,

whereupon they were broken down by trituration. Upon immunohistochemical examination,

the cells within enteric neurospheres were positive for the low affinity nerve growth factor

38

receptor p75NTR (Figure 2B). They were additionally also positive for Sox2 (Figure 2C, SRY

related homeobox factor 2) and Nestin (Figure 2D), a neuroepithelial stem cell marker. Our

results indicate that neurospheres derived from the rabbit intestine following this procedure

are comprised of neural crest-derived cells that are capable of differentiating in to enteric

neurons and/or enteric glia.

3.2 Neuronal differentiation on individual ECM substrates (Collagen I, Collagen IV or

Laminin)

Figure 3: Schematic of enteric neurosphere differentiation as a function of ECM composition

22x11mm coverslips were coated with pLL, Collagen I,

Collagen IV, Laminin or Heparan sulfate individually or

in combination. Enteric neurospheres were allowed to

adhered to the coated coverslips for 6 hours.

Separately, uncoated glass coverslips were seeded

with colonic smooth muscle cells, and allowed to grow

to confluence. In order to stimulate differentiation of

enteric neurospheres, a coverslip containing confluent

smooth muscle was placed within the same dish, so

the two coverslips shared soluble factors.

Poly-lysine (pLL) coating was a pre-requisite to enteric neurosphere adhesion to glass

coverslips. Glass coverslips that lacked any coating did not support enteric neurosphere

adhesion sufficiently to differentiate into neurons or glia. In order to maintain uniformity, all

coverslips were initially coated with pLL and additionally with Laminin, Collagen I or

Collagen IV. All coated coverslips required between 2-4 hours for enteric neurospheres to

attach.

Enteric neurospheres on coated coverslips were allowed to differentiate initially using

neuronal differentiation medium alone. Several sets of experiments demonstrated that these

cells did not undergo any differentiation under these conditions. Thereby, in order to render

39

the soluble environment conducive to differentiation, a confluent coverslip containing colonic

smooth muscle cells was placed in the same culture dish (Figure 3). The neuronal coverslip

(coated with ECM substrate and containing enteric neurospheres) and the smooth muscle

coverslips thereby shared soluble factors. The addition of the smooth muscle coverslip

marked the initiation of differentiation (day 0).

Morphological evidence of neuronal or glial differentiation was readily visible by day 5. A

later time point (day 15) was identified to study the development of mature neurons or glia in

vitro as a function of ECM composition. During the differentiation process, the culture dishes

remained undisturbed till the early time point (day 5) or the late time point (day 15), except

for medium supplementation. Neuronal differentiation was identified by immunofluorescent

staining of the neuronal coverslip at either day 5 or day 15 with an antibody directed against

βIII Tubulin.

Figure 4: Neuronal differentiation on individual coated coverslips

βIII Tubulin antibody (green) was used to visualize neurons on Day 5 (A-D) and Day 15 (E-

H) coverslips coated with pLL (A,E), Laminin (B,F), type I Collagen (C,G) and type IV

Collagen (D,H). Enteric neurospheres on pLL barely initiate neuronal differentiation even at

day 15. Neurospheres on Laminin, Collagen I and Collagen IV showed branching and

several neuronal processes both at the early and late time points in vitro.

40

Day 5 timepoint: Even in the presence of smooth muscle, enteric neurospheres on pLL

remained undifferentiated, with some progenitor cells within neurospheres expressing low

levels of βIII Tubulin (Figure 4A). However, with the addition of either laminin, Collagen I or

Collagen IV to pLL on the culture substrata, neuronal differentiation was evident by day 5

(Figure 4B-D). Neurite lengths varied non-significantly between 193.2 ± 9.9 µm and 288.2 ±

14. 5µm on ECM substrata at the early time point (Figure 10A). Neurons on Collagen IV

and Laminin coated coverslips demonstrated a higher level of branching (two or more

neurites per cell; Figure 4B,D).

Day 15 timepoint: At the day 15 timepoint, neurospheres on pLL coverslips barely initiated

neuronal differentiation, evidenced by a flatter morphology and the appearances of faint

tubulin-positive extensions (Figure 4E). With the addition of either laminin, Collagen I or

Collagen IV, neurite lengths were significantly longer compared to pLL (p<0.001).

Differentiated neurons on laminin coverslips demonstrated the longest neurite extensions

(326.9 ± 13.3µm, n=27; Figure 4F), significantly longer than Collagen I or Collagen IV

(p<0.05, Figure 10A). By day 15, neurons on Collagen I-coated coverslips still had no

significant branching compared to those on Collagen IV-coated coverslips (Figures 4G-H).

3.3 Neuronal differentiation on collagen-laminin substrates

In the next set of experiments, combinations of collagens and laminin were evaluated. Two

concentrations of laminin were evaluated to identify the minimum amount of laminin required

to influence neuronal differentiation. Coverslips were coated with either collagen I or

collagen IV with 5µg/cm2 or 10µg/cm2 of laminin. The addition of laminin enhanced neuronal

differentiation when compared to individual collagen substrates (compare Figure 5 with

Figure 4), but no significant difference was observed in neurite length between the two

concentrations of laminin.

41

Figure 5: Neuronal differentiation on collagen-laminin substrates

Neurons stained with βIII Tubulin (green) on Day 5 (A-D) and Day 15 (E-H) coverslips coated with type I Collagen and 5µg/cm2 Laminin (A,E) or 10µg/cm2 Laminin (B,F) or type IV Collagen with 5 (C,G) or 10 (D,H) µg/cm2 of Laminin. Addition of laminin to collagen substrates enhanced early and late neuronal differentiation, but no significant difference was observable between 5 and 10 µg/cm2 of laminin. Type IV Collagen substrates (C,D,G,H) demonstrated enhanced neuronal branching and differentiation compared to type I collagen (A,B,E,F) substrates.

Collagen I and Laminin: At the day 5 timepoint, addition of laminin to Collagen I increased

the number of progenitor cells undergoing neuronal differentiation, but did not alter neuronal

branching or neurite lengths significantly (Figure 5 A,B,E,F). At the day 15 timepoint,

significantly enhanced neuronal differentiation (p<0.05) was observed compared to Collagen

I only. Neurite lengths on collagen-laminin substrates at day 15 (279.7 ± 10.8 µm – 291.9 ±

8.1 µm; n=27-34) were longer than Collagen I substrates (235.5 ± 10.1 µm; Figure 10B).

42

Collagen IV and Laminin: The addition of laminin to Collagen IV enhanced neuronal

differentiation when compared to coverslips coated individually with Collagen IV only

(compare Figure 5C,D,G,H to Figure 4D,H). At the day 5 timepoint, the addition of laminin

increased the number of cells undergoing neuronal differentiation. No significant difference

was observed in neurite lengths at day 5 (247 ± 12.9 µm – 263.7 ± 9.7 µm; Figure 10B).

At the day 15 time point, coverslips coated with both collagen IV and laminin had

significantly (p<0.05) longer neurites compared to Collagen IV only (324.6 ± 16.5 µm

compared to 281.7 ± 8.6µm, Figure 5G-H). Initiation of inter-neuronal networking was also

observed. There was no observable or significant difference in neurite lengths between the

two concentrations of laminin used (5µg/cm2 or 10µg/cm2; Figure 10B).

3.4 Neuronal differentiation on composite ECM substrates with laminin and heparan sulfate

In this additional set, the effect of a combination of collagens on neuronal differentiation was

investigated. Composite coatings were evaluated with a 2:1 mix of Collagen I/Collagen IV as

the base. This composite collagen base was evaluated first. Additionally, neuronal

differentiation was evaluated on substrates that included laminin and/or heparan sulfate in

combination with composite collagen.

Composite Collagen I/Collagen IV: Several cells underwent neuronal differentiation (Figure

6A, E; similar to individual coatings of either Collagen I or Collagen IV), but neurite lengths

were significantly shorter on composite collagen substrates at day 5. Neurite lengths

measured at 156.1 ± 7.2 µm on day 5. At day 5, neuronal differentiation progressed and

individual neurons had multiple branches and long neurites (Figure 6E). Neurite lengths

averaged at 241.2 ± 9.4 µm on day 15 (Figure 10C).

43

Figure 6: Neuronal differentiation on composite collagen-laminin-heparan sulfate (HS) substrates

Neurons stained with βIII Tubulin (green) on Day 5 (A-D) and Day 15 (E-H) coverslips

coated with type I and IV Collagen with either 5µg/cm2 Laminin (B,F), 0.1µg/cm2 Heparan

Sulfate (HS; C,G), both (D,H) or none (A,E). Addition of laminin or HS to collagen substrates

enhanced early and late neuronal differentiation, with visible networking by day 15.

Substrates without laminin or HS demonstrated minimal neuronal differentiation (A,E).

Addition of Laminin: Addition of laminin to composite collagen substrates increased the

number of differentiated neurons visible by day 5 (Figure 6B). Neurite lengths were

significantly (p<0.05) longer on substrates containing laminin (215.1 ± 7.6 µm, n=43). At the

day 15 time point, substrates containing composite collagen and laminin demonstrated

significant clustering of neurons (Figure 6F), with increased neurite lengths averaging at

286.8 ± 9.5 µm, n=50.

Addition of heparan sulfate: Addition of heparan sulfate also dramatically increased the

number of progenitor cells undergoing neuronal differentiation by day 5 (Figure 5C).

Average neurite lengths on substrates containing heparan sulfate along with composite

collagen was 212.8 ± 9.5 µm, n=43 at day 5. At day 15, the initiation of neuronal networking

was visible with βIII Tubulin staining (Figure 6G).

44

Addition of laminin and heparan sulfate: The addition of laminin and heparan sulfate

together with the composite collagen increased the number of differentiated neurons as well

as the length of the individual neuronal processes and neurite branching (Figure 6D). At day

15, initiation of neuronal networking with significant clustering of neurons was observed

(Figure 6H). Neurite lengths were significantly longer (325.5 ± 19.4µm, n=36) compared to

composite collagen alone (Figure 10C).

3.5 Glial differentiation on individual ECM coatings (Collagen I, Collagen IV or Laminin)

In addition to neuronal differentiation studies described above, glial differentiation was also

studied as a function of ECM composition of culture substrata. Enteric neurospheres were

plated on to coated coverslips in duplicate, and one coverslip was used to evaluate neuronal

differentiation while a duplicate coverslip was used to evaluate glial differentiation. A primary

antibody directed against Glial fibrillary acidic protein (GFAP) was utilized to identify glial

differentiation. Fluorescent microscopy was used to visualize differentiated glia, using a

TRITC fluorophore conjugated secondary antibody. The Nikon documentation software was

used to calculate mean red fluorescence indicating the number of differentiated glia in a field

of view of constant area.

Day 5 timepoint: In the presence of smooth muscle, enteric neurospheres on pLL coated

coverslips demonstrated significant GFAP staining by day 5 (15.3 ± 1.3 AU; Figure 7A). In

contrast, enteric neurospheres on pLL coverslips did not demonstrate significant neuronal

differentiation at day 5, indicating the preferential differentiation in to glia at the early time

point on pLL coverslips. With the addition of laminin, enteric neurospheres demonstrated

highly significantly reduced GFAP staining (0.4 ± 0.2 AU). Undifferentiated neurospheres on

the laminin coverslips contained several progenitor cells that were positive for GFAP (Figure

7B). On the same laminin coated coverslips, neuronal differentiation was extensive at the

45

early timepoint, indicating an early preference for neuronal differentiation in the presence of

laminin (Figure 7B). Minimal glial differentiation was observed on either of the collagen

substrates (Figure 7C-D).

Figure 7: Glial differentiation on primary coated substrates

Glia stained with GFAP (red) on Day 5 (A-D) and Day 15 (E-H) coverslips coated with pLL

(A,E), Laminin (B,F), type I Collagen (C,G) and type IV Collagen (D,H). Enteric

neurospheres on pLL demonstrated maximal glial differentiation starting at day 5 up to day

15. While neurospheres on collagen substrates demonstrated good early and late glial

differentiation, laminin coated substrates showed no early glial differentiation by day 5 (B),

but differentiated subsequently by day 15 (F).

Day 15 timepoint: By the late day 15 timepoint, pLL coated coverslips had the highest

number of glia, indicated by a highly significant (p<0.0001) GFAP fluorescent intensity

averaging at 28. 6 ± 1.1 AU (Figure 7E, 10D). Glia were apparent on ECM coated coverslips

as well, but to a lower extent than on pLL. In contrast to day 5, laminin coated coverslips

demonstrated the presence of several glia at the day 15 time point, and a robust GFAP

fluorescent intensity was observed (Figure 6F, 16.5 ± 0.3 AU). Several glia were observed

46

by day 15 on either of the collagen substrates, with fluorescence ranging from 11.8 to 13.4

AU. (Figure 7G-H).

3.6 Glial differentiation on collagen-laminin substrates

Figure 8: Glial differentiation on collagen-laminin substrates

Glia stained with GFAP (red) on Day 5 (A-D) and Day 15 (E-H) coverslips coated with type I

Collagen and 5µg/cm2 Laminin (A,E) or 10µg/cm2 Laminin (B,F) or type IV Collagen with 5

(C,G) or 10 (D,H) µg/cm2 of Laminin. Addition of laminin to collagen substrates promoted

glial differentiation both at days 5 and 15. No significant difference was observable in

GFAP+ glial differentiation between 5 and 10 µg/cm2 of laminin.

Similar to neuronal differentiation, glial differentiation was evaluated on substrates that were

coated with either Collagen I or Collagen IV with laminin. The addition of laminin to collagen

coated coverslips did not inhibit glial differentiation. Several differentiated glia were

observed on day 5 (8.4 ± 0.8 – 14.1 ± 0.3 AU) on collagen-laminin substrates (Figure 8A-D).

There was no significant difference in the number of GFAP positive cells at the early time

point with the addition of laminin (5µg/cm2 or 10µg/cm2) to either Collagen I or Collagen IV

substrates Figure 10D). Robust GFAP expression (12.6 ± 1.3 – 14.2 ± 1.0 AU) was

47

observed at the day 15 time point on all collagen-laminin substrates, not significantly

different from one another (Figure 8E-H).

3.7 Glial differentiation on composite ECM substrates with laminin and heparan sulfate

Glial differentiation was evaluated by varying the culture substratum with a combination of

collagen I and IV. Additionally, the effect of the addition of laminin and/or heparan sulfate

was also studied on glial differentiation.

Figure 9: Glial differentiation on composite collagen-laminin-heparan sulfate (HS) substrates

Glia stained with GFAP (red) on Day 5 (A-D) and Day 15 (E-H) coverslips coated with type I

and IV Collagen with either 5µg/cm2 Laminin (B,F), 0.1µg/cm2 Heparan Sulfate (HS; C,G),

both (D,H) or none (A,E). Addition of laminin or HS to composite collagen substrates

demonstrated glial differentiation that peaked at day 5(B-D) that was sustained at the later

time point (E-H).

Composite Collagen I/Collagen IV: Glial differentiation peaked on day 5, on coverslips

coated with the collagen I/IV mixture (Figure 9A). Red fluorescence (15.8 ± 0.5 AU) was

comparable to that on pLL coated coverslips at day 5 (Figure 10D). In contrast, neuronal

differentiation on composite collagen coated coverslips was poor at the early time point,

48

indicating a preferential differentiation into glia early on. By day 15, initiation of clustering of

glial cells was observable (Figure 9E), with no significant increase in red fluorescence.

Addition of laminin and/or heparan sulfate: Early glial differentiation at day 5 was reduced

(10.2 ± 0.8 to 11.1 ± 0.5 AU) with the addition of laminin and/or heparan sulfate to composite

collagen substrates (Figure 9B-D). In contrast, these substrates supported neuronal

differentiation extensively (compare Figure 9A-D to Figure 6A-D), indicating a preferential

neuronal differentiation at the early time point. At the later day 15 time point, a non-

significant increase in the number of glia was observed (Figure 9E-H, Figure 10D).

4. Discussion

Enteric neuronal progenitor cells have been identified in the adult mammalian gut,

and have been isolated from humans >80 years of age [5, 23]. Previously, several groups

have shown that a self-renewing population of Sox 2 [8], Sox10 [23], Nestin [24] and p75

[25] positive neural-crest derived progenitor cells can be isolated either from full-thickness,

muscularis or mucosal biopsies of the adult mammalian gut [26-28]. These cells have been

demonstrated to have the potential to differentiate in to several neuronal subtypes including

inhibitory and excitatory motor neurons and glia. We have isolated neuronal progenitor cells

from full thickness biopsies of adult rabbit jejunums that aggregate in culture to form floating

spherical colonies, dubbed enteric neurospheres (Figure 2A). We confirmed that these

enteric neurospheres were comprised of cells positive for p75, Sox2 and Nestin (Figure 2 B-

D). The presence of p75NTR confirms the neural-crest lineage of the isolated cells. The

presence of Sox2 and Nestin confirm the progenitor status of the isolated cells, indicating

that these cells are similar to enteric neuronal progenitor cells previously isolated from the

gut that have the potential to differentiate into both neurons and glia.

49

The feasibility of transplantation of various types of neuronal progenitor cells (CNS-

derived, neural tube-derived, embryonic and adult ENS-derived) in explant cultures of

aneural gut is well established [5, 7, 9]. Conditions required for successful engraftment and

long-term phenotype maintenance, focusing on a permissive environment are yet to be

satisfactorily identified. Alterations in the extracellular matrix of the gut mesenchyme has

been documented in aganglionic regions of rodent gut, suggesting the importance of a

permissive extracellular environment to promote effective in utero colonization and

differentiation of neural crest cells in the developing gut [29-32]. Since transplantation and

subsequent functional neo-innervation is the clinical goal of neural stem cell transplantation,

in vitro studies must mimic developmental conditions in vivo, in terms of providing a

permissive and favorable ECM (preferably three-dimensional). A focus on the role of the

ECM in affecting neuroglial differentiation of adult enteric neuronal progenitor cells can

optimize the survivability and maintenance of a stable phenotype upon transplantation.

Previous studies by Schӓfer et al. demonstrate that fetal rodent enteric neurospheres

differentiate forming neurites on laminin coated culture surfaces, and secondary ganglion

like structures in composite ECM gels within 7 days in vitro [33].

Mammalian myenteric ganglia in vivo are surrounded by a matrix comprised

predominantly of type IV Collagen, laminin, heparan sulphate proteoglycan, and entactin

[17, 34]. The enteric plexus lacks large connective tissue spaces for blood vessels like the

peripheral nervous system. The studies in this paper sought to address the hypothesis that

a two-dimensional culture substratum will modulate neuronal and glial differentiation solely

based on ECM composition. In this study, glass coverslips were coated with combinations of

three different ECM components that enteric glia and neurons come in to contact with in vivo

in the adult myenteric plexus: Collagen IV, Laminin and Heparan sulfate [17]. Collagen I was

evaluated additionally, because of its extensive use in neural tissue engineering of the

50

peripheral nervous system [35, 36]. Poly-lysine (pLL) coated coverslips were used a control

because a coating was a prerequisite to cell adhesion and subsequent differentiation.

Neurospheres and neuronal progenitor cells attached to the pLL coated coverslips,

and stayed attached at day 5, but did not initiate neuronal differentiation. However, glial

differentiation was readily visible by day 5, and improved by day 15 (Figure 3A,E; Figure

6A,E). Enteric neurospheres on pLL substrates indicated a clear preference towards glial

differentiation versus neuronal differentiation. It is likely that progenitor cells or neurons were

only weakly attached to pLL in the absence of cellular adhesion recognition

sequences/molecules, as reported previously for hippocampal neurons and dorsal root

ganglion neurons [37, 38]. The severely reduced neurite outgrowth could have also been

caused by losing undifferentiated progenitor cells during medium supplementation or

immunohistochemical staining.

Addition of laminin to collagen substrates improved neurite outgrowth with longer

neurite lengths (compare 156.1 ± 7.2 µm to 215.1 ± 7.6 µm). Laminin has been long known

to stimulate neural cell attachment and neurite outgrowth [18]. Indeed, migratory neural

crest cells have been shown to acquire a neurally-related laminin receptor upon entering the

gut mesenchyme, that facilitates differentiation [39]. Our studies showed that while there

was an overall enhancement in neuronal differentiation as well as neurite outgrowth, there

was no significant difference between the additions of 5 or 10 µg/cm2 of laminin. This

empirical determination was important in determining a minimal amount of laminin that can

influence neuroglial differentiation without affecting neurite outgrowth adversely in a situation

that requires neo-innervation of denervated tissues.

51

Figure 10: Morphometric mesaurements

Neurite lengths were measured on coated culture substrata and compared using one way

ANOVA. Two bars for each substrate show mean neurite lengths at day 5 and day 15. (A)

Neurite lengths on pLL were significantly (***p<0.001) shorter than any primary coating

substrate. Laminin substrates had the longer neurites (*p<0.05). (B): No significant

difference was observed in neurite lengths with the addition of 5 or 10 µg/cm2 laminin. (C):

The addition of laminin or heparan sulphate significantly increased neurite lengths over

composite collagen substrata (*p<0.05). (D) Mean GFAP immunofluorescence was

quantified: pLL substrates (***p<0.001) and composite collagen substrates (*p<0.05)

supported extensive glial differentiation.

Addition of heparan sulfate to composite collagen mixtures improved neuronal

differentiation as well. Neuronal networking and neuronal clustering was visible at the later

time point. Heparan sulfate and its interaction with GDNF and other neurotrophic factors

stabilizes and makes these factors locally available, possibly modulating neurite outgrowth

and neuronal differentiation [19, 20]. Heparan sulfate interacts with both Collagen IV and

52

with Laminin, to positively modulate neuronal differentiation, evidenced by the enhanced

neurite lengths and initiation of neuronal networking (Figure 4 A-H). Mammadov et al.

recently demonstrated that self-assembled peptide nanofibers with heparan sulphate and

laminin mimetic sequences promoted neurite outgrowth of a PC12 cell line [20]. Composite

collagen substrates with laminin and/or heparan sulfate all maintained a low level of GFAP

positive glial cells, with initiation of astrocytic networking becoming more obvious at the later

time point. In general, substrates that supported neuronal differentiation demonstrated a

bare minimum of glial cells required to possibly support neuronal cell phenotype or survival.

Several studies in literature suggest that the presence of several axolemmal fragments can

arrest the proliferation of glia [40]. This is consistent with our observations of low levels of

GFAP immunofluorescence observed on substrates that supported extensive neuronal

differentiation. The only substrates that supported extensive differentiation of enteric

neuroglial progenitor cells into glia were pLL and individual coatings of Collagen I/IV.

Neuronal differentiation was present on these substrates, but not as extensively as any of

the other composite coatings that included laminin and heparan sulfate.

5. Conclusions

Individually, all evaluated culture substrates in these studies supported neuronal and

glial differentiation to varying degrees by day 15. Enteric neurospheres demonstrated a

clear tendency to differentiate into glia on pLL coated substrates as well as on composite

collagen substrates in the absence of laminin and heparan sulfate. In contrast, culture

substrates with laminin and heparan sulfate promoted extensive neuronal differentiation

while simultaneously supporting only a minimal glial cell population. Laminin and collagen IV

coated coverslips positively modulated neuronal differentiation by increased number of

neurites per neuron and longer neurite lengths compared to fibrillar Collagen I. These

results identify suitable 3D matrix compositions to deliver neuronal progenitor cells. Future

53

studies will focus on evaluating neuroglial differentiation within 3D matrices with variable

ECM composition, with an aim to tissue engineer intrinsically innervated sheets of smooth

muscle suitable for transplantation. Three dimensional hydrogel environments also provide

the mechanical cues for neural differentiation, more readily translatable to in vivo conditions

than infinitely stiff glass substrates.

6. Acknowledgements

This work was supported by NIH /NIDDK research grants RO1DK071614 and

RO1DK042876.

7. References

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[32] Gershon MD. Transplanting the enteric nervous system: a step closer to treatment for aganglionosis. Gut. 2007;56:459-61. [33] Schafer KH, Hagl CI, Rauch U. Differentiation of neurospheres from the enteric nervous system. Pediatr Surg Int. 2003;19:340-4. [34] Rauch U, Schafer KH. The extracellular matrix and its role in cell migration and development of the enteric nervous system. Eur J Pediatr Surg. 2003;13:158-62. [35] Stang F, Fansa H, Wolf G, Reppin M, Keilhoff G. Structural parameters of collagen nerve grafts influence peripheral nerve regeneration. Biomaterials. 2005;26:3083-91. [36] Dewitt DD, Kaszuba SN, Thompson DM, Stegemann JP. Collagen I-matrigel scaffolds for enhanced Schwann cell survival and control of three-dimensional cell morphology. Tissue Eng Part A. 2009;15:2785-93. [37] Liu B, Ma J, Gao E, He Y, Cui F, Xu Q. Development of an artificial neuronal network with post-mitotic rat fetal hippocampal cells by polyethylenimine. Biosens Bioelectron. 2008;23:1221-8. [38] Orr DJ, Smith RA. Neuronal maintenance and neurite extension of adult mouse neurones in non-neuronal cell-reduced cultures is dependent on substratum coating. J Cell Sci. 1988;91 ( Pt 4):555-61. [39] Pomeranz HD, Sherman DL, Smalheiser NR, Tennyson VM, Gershon MD. Expression of a neurally related laminin binding protein by neural crest-derived cells that colonize the gut: relationship to the formation of enteric ganglia. J Comp Neurol. 1991;313:625-42. [40] Eccleston PA, Bannerman PG, Pleasure DE, Winter J, Mirsky R, Jessen KR. Control of peripheral glial cell proliferation: enteric neurons exert an inhibitory influence on Schwann cell and enteric glial cell DNA synthesis in culture. Development. 1989;107:107-12.

56

CHAPTER 5: BIOENGINEERED THREE-DIMENSIONAL

PHYSIOLOGICAL MODEL OF COLONIC LONGITUDINAL SMOOTH

MUSCLE IN VITRO

S Raghavan1, MT Lam2, LL Foster1, RR Gilmont1, S Somara1, S Takayama2 and KN

Bitar1

GI Molecular Motors Lab, Department of Pediatrics, Gastroenterology1

Biomedical Engineering2

University of Michigan, Ann Arbor MI 48109

This manuscript was accepted for publication in Tissue Engineering Part C

2010; 16(5): 999-1009

57

ABSTRACT

Background: The objective of this study was to develop a physiological model of

longitudinal smooth muscle tissue from isolated longitudinal smooth muscle cells (LSMC)

arranged in the longitudinal axis. Methods: LSMC from rabbit sigmoid colon were isolated

and expanded in culture. Cells were seeded at high densities on to laminin coated Sylgard

surfaces with defined wavy micro-topographies. A highly aligned cell sheet was formed, to

which addition of fibrin resulted in delamination. Results: 1) Acetylcholine induced a dose-

dependent, rapid and sustained force generation. 2) Absence of extracellular calcium

attenuated the magnitude and sustainability of Ach- induced force by 50% and 60%

respectively. 3) Vasoactive Intestinal Peptide also attenuated the magnitude and

sustainability of Ach- induced force by 40% and 60% respectively. This data was similar to

force generated by longitudinal tissue. 4) Bioengineered constructs also maintained smooth

muscle phenotype and calcium dependence characteristics

Summary: This is a novel physiologically relevant in vitro 3-D model of colonic longitudinal

smooth muscle tissue. Bioengineered 3-D longitudinal smooth muscle presents the ability to

generate force, and respond to contractile agonists and relaxant peptides similar to native

longitudinal tissue. This model has potential applications to investigate the underlying

pathophysiology of dysfunctional colonic motility. It also presents as a readily implantable

“band-aid” colonic longitudinal muscle tissue.

1. INTRODUCTION

Colonic motility can be severely impacted in a variety of situations ranging from

idiopathic inflammatory bowel disorders to an individual’s lifestyle patterns and the amounts

of stress therein. Dysmotility presents itself as diarrhea and abdominal pain among other

symptoms, implicating the involvement of neural circuits and smooth muscle contractility (1-

4). The colon comprises of two principle smooth muscle layers– an inner circular layer

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wherein the smooth muscle cells are aligned concentrically around the lumen; and an outer

longitudinal layer wherein the smooth muscle cells run parallel to the length of the colon.

The two muscle layers, along with the enteric nervous system, modulate colonic motility

patterns. The muscle layers are the primary effectors of force generation in gut motility, with

their specific functionality arising from their cellular organization and alignment. Contraction

of the circular smooth muscle lowers the diameter of the luminal space, while contraction of

the longitudinal smooth muscle shortens the length of the colon. Duplication of the muscle

architecture has been a key aspect of developing a 3D in vitro model of muscle layers.

In vitro 3D models aid the study of muscle layers alone in the absence of mucosa

and neural inputs, eliminating the use of full thickness muscle preparations from animal

models. 3D growth milieu facilitates well defined aggregation geometries, mimics cellular

organization in vivo and thus displays functional superiority (5, 6). Structure can be directly

related to function in three dimensional models, which is of particular importance while

engineering smooth muscle tissue for force analyses.

Three dimensional primary muscle cultures were initially reported with muscle cells

bioengineered with the aid of scaffolds (7, 8). Over the years, the limitations of scaffold

based muscle tissue engineering were recognized and extensive efforts by many others

were made to bioengineer self-organizing skeletal muscle constructs, vascular constructs

and cardiac pressure constructs, etc. (9-14).

We modified the model described by Dennis & Kosnik (13) for mammalian skeletal

muscle constructs to culture three-dimensional self organizing colonic longitudinal smooth

muscle tissue. We provided a passive laminin-coated adhesion surface with micro-

topographies to promote cellular adhesion and orientation as described by Lam et al. (15,

16). A 10cm2 cell sheet was formed with longitudinal smooth muscle cells aligned along their

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longitudinal axis, similar to the alignment of longitudinal smooth muscle cells in vivo. For

ease of force measurement, the oriented cell sheet was overlaid with a fibrin matrix. This

coincided with the delamination and formation of bioengineered longitudinal smooth muscle

tissue in ~7-10 days. Bioengineered tissue retained their physiological properties of force

development and agonist-induced contractions and relaxations like native longitudinal

smooth muscle tissue.

This is the first report of in vitro three dimensional culture of isolated colonic

longitudinal smooth muscle without mucosal or neuronal cells. Since the bioengineered

tissue is from dispersed cells that are electrically uncoupled, the study of spontaneous

contractile activity of this muscle layer and its responses to peptide neurotransmitters,

contractile agonists without the sub-mucosal and mucosal inputs if any is possible.

Previously, longitudinal muscle activity was studied by immobilizing the adjacent

circular muscle layers in flat-sheet tissue preparations, or by implanting strain gauge

transducers with their measurement axis aligned to the axes of the muscle in a surgical

procedure in vivo. These aligned constructs could be conveniently hooked to a force

transducer, to effect in vitro force measurements sans a surgical procedure.

Apart from the convenience of force analysis and smooth muscle contractility studies

using the three-dimensional construct, the cells that make up the construct could also be

mutated with gene therapy and/or signal transduction pathway targets. Emerging evidence

suggests that modulation and control of the signal transduction pathways at the thin and

thick filament levels in smooth muscles could alter the contractility of the smooth muscle (17-

19). Thus, bioengineered tissue could be a convenient three-dimensional, physiologically

relevant in vitro tissue engineering model to study motility-related symptomatic defects at the

smooth muscle level.

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2. MATERIALS AND METHODS

Reagents

All tissue culture reagents including basal Dulbecco’s Modified Eagle’s Media

(DMEM), fetal bovine serum (FBS), media supplements like L-glutamine and antibiotics

were purchased from Invitrogen. (Carlsbad, CA). Acetylcholine and Vasoactive Intestinal

Peptide were purchased from Sigma (St.Louis, MO). Collagenase Type II was purchased

from Worthington. (Lakewood, NJ). Sylgard (Poly(dimethylsiloxane); PDMS) was from World

Precision Instruments (Sarasota, FL) and Epo-Tek Resin was from Epoxy Technology

(Boston, MA).

2.1 Isolation and culture of Rabbit Longitudinal Smooth Muscle Cells (RLSMC)

Rabbit sigmoid colon was removed by dissection, and relieved of fecal content. The

tissue was kept on ice and moist with Hank’s Balanced Salt Solution (HBSS) containing

antibiotics and sodium bicarbonate. The cleaned colon was slipped on to a plastic pipette.

Blood vessels and adherent fat were picked off with forceps. Kimwipe® (Kimberly-Clark, WI)

wetted with HBSS was used to wipe the outer layer of the colon.

Fine tip forceps were used to pick off the longitudinal muscle layer from the colon

and store them in ice-cold HBSS. The tissue was finely minced and digested twice with type

II collagenase (0.1%) at 32oC for 1 hour, filtered through a 500-μm Nytex® (Tetko Inc, NY)

mesh. The filtrate was washed three times and plated in DMEM with 10% Fetal Bovine

Serum, 1.5% Antibiotics and 0.5% L-glutamine on to regular tissue culture flasks.

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2.2 Preparation of replicating wavy molds

Master molds with sinusoidal wavy patterns were produced by thermal expansion,

oxidation and cooling of PDMS as described by Lam et al.(15). Briefly, 15:1 pre-polymer:

curing agent by weight was wrapped around a glass cylinder with a 2cm diameter. On

plasma oxidation with air at 60mTorr for 5-45 minutes, master molds with wavy patterns

6μm apart with depths of 1700nm were prepared (15).

Replicating molds were made by pouring Sylgard into 60mm plates, to a depth of 3mm.

After oven curing overnight at 60oC, a rectangular portion was cut out in the middle of the

plate and discarded. Epo-Tek was poured into the plate to fill the cut-out section and the

master wavy mold was placed wave-down and UV cured.

2.3 Preparation of wavy plates for cell culture

10:1 Sylgard was poured into the Epo-Tek replicating mold, and cured at room

temperature for 2 days. The rectangular section was then cut out, and a 35mm diameter

hole was punched out from it to fit regular 35mm culture plates. The round-cut wavy

subtrates were fixed to 35mm plates with the wavy surface facing upwards. This

arrangement was left to cure and detoxify at room temperature.

After 2 days, the wavy plates were sterilized under UV light and by ethanol for up to

90 minutes. 2µg/cm2 of Laminin was deposited evenly onto the tissue culture plate. Silk

suture anchors, 6mm wide, were dipped in dilute laminin solution, and were pinned onto the

wavy plates, 12mm apart. 2mL of Growth Medium (GM; DMEM + 10% FBS, 1% ABAM) was

added to the plate, and stored in the incubator under sterile conditions till the cells were

ready to be plated.

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2.4 Bioengineering aligned Cell Sheets

RLSMC were grown till they were 80-90% confluent. Cells were trypsinized as per a regular

passage. 500,000-600,000 cells are added to each wavy plate in 1.5mL of GM and allowed

to adhere and align along the wavy micro patterns on the laminin-coated PDMS substrate

for 3-4 days. Once the formation of the oriented cell sheet was microscopically confirmed,

4mg of fibrinogen and 5u of Thrombin and GM were added. When the cell sheet began to

delaminate, the growth media was switched to Differentiation Media (Medium 199, 5%

Horse Serum, 1% Antibiotics) to promote smooth muscle differentiation. The aligned cell

sheet delaminated and compacted, 2-4 days after gel addition, to form bioengineered

longitudinal smooth muscle tissue, anchored to the two silk sutures.

2.5 Immunofluorescence

Colonic longitudinal smooth muscle cells isolated from primary cultures were grown to

confluence on microscope chamber slides and treated with differentiation media for two

days. Cells were then fixed and assayed with antibodies for α-Smooth muscle Actin (F3777,

Sigma), smooth muscle specific heavy Caldesmon (c-4562, Sigma), and c-Kit (sc-168,

Santa Cruz Biotechnology). Negative control treatment was with secondary antibody only.

Immunofluorescence was visualized with an Olympus IX71 fluorescence microscope.

2.6 Physiological testing protocols

The physiological functionality of the aligned bioengineered RLSMC tissue was

evaluated in terms of force responses to various contractile agonists and antagonists. Force

measurements were carried out with a custom built magnetoresistive force transducer with

an attached vernier control. An insect pin was fixed in the center of the 4mL tissue bath, to

provide a hooking point. The complete set-up was housed in a vibration resistant table.

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Once the aligned cell sheets were formed, the holding pins were removed. The silk

suture anchors were gently teased away from the bioengineered tissue. One end of the

construct was hooked onto the movable sensing arm of the transducer, while the other end

hooked onto the stationary pin. It was ensured that the tissue remained immersed in fluid,

along with the mounting pins to avoid any air vibrations. For tissue experiments, a 1cm

piece of longitudinal tissue was isolated from the rabbit colon and directly tested in the

presence of tetrodotoxin.

The tissue was immersed in warm, oxygenated basal DMEM in regular calcium

experiments. In zero calcium experiments, the bath medium was replaced with Phosphate

Buffered Saline with 2mM EGTA. A 10% stretch was applied on the constructs by the

micromanipulator and this remained the length at which all further tests were carried out.

The baseline of force established upon equilibration at this length was arbitrarily set to zero.

The substances used were Acetylcholine (ACh) for cholinergic stimulation, and

Vasoactive Intestinal Peptide (VIP).The volume of these agonists were maintained uniform

at 100µL, avoiding any error from a non-uniform diffusion of the substance. It was also noted

that it took upto 60s for the uniform diffusion of the agonists in the tissue bath. Passive

tension was determined by measuring force with warmed basal DMEM alone without

hooking the tissue. To determine a signal to noise ratio (SNR), the bioengineered tissue was

attached to the measurement system without any stretch. For all other physiological force

measurements, the tissue was stretched an additional 10% of its original length. Baseline

was established prior to addition of any agonists. At the end of each experiment, or after 30

minutes (whichever was earlier), the longitudinal constructs were washed and supplied with

fresh warm oxygenated basal DMEM.

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Response to ACh was quantified by the addition of different doses ranging from 1nM

to 1µM. In addition, a single dose of 1µM was also independently tested on the constructs.

To study the inhibition of force generation in the presence of VIP – the constructs were first

treated with 1µM VIP and upon subsequent relaxation and recovery of baseline - 1µM of

ACh was added to the same tissue bath without a wash the subsequent response was

recorded.

2.7 Data Analysis

Raw data was acquired using LabScribe2 (iWorx, Hanover, NH). GraphPad Prism 5.01 for

Windows (GraphPad Software, San Diego CA, www.graphpad.com) was used for all further

data analysis. 2nd order Savitsky-Golay smoothing was applied on raw data. Values were

expressed as means and SEM of 4-7 experiments. Significant differences were tested using

2-way ANOVA with Bonferroni posttests to compare between means. A P value less than

0.05 was considered significant. Signal to noise ratio (SNR) was calculated by obtaining the

ratio of mean baseline force of a non-stretched tissue over the standard deviation of the

passive liquid tension. The traces of these measurements are shown in Figure 14.

3. RESULTS

3.1 Bioengineering Rabbit Longitudinal Smooth Muscle (RLSM) Tissue

Figure 11 outlines the progression of steps involved in bioengineering RLSM tissue.

Replicating wavy molds were made from Epo-Tek resin with defined wavy topographies.

These surfaces are characterized elsewhere (16). When 10:1 Sylgard was poured into these

replicating molds, they cured in 2-3 days to form a square shaped block with wavy patterns

embedded on the surface in contact with the replicating molds. These cured Sylgard blocks

were cut out and attached on to tissue culture plates with the wavy surface facing upwards.

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The plates were left to cure and detoxify on the bench top for 2-3 days, and sterilized further

in the biosafety cabinet with UV light and ethanol.

Figure 11: Process schematic for bioengineering rabbit Longitudinal Smooth Muscle Cell Sheet

Replicating molds with defined wavy surfaces were patterned with Epo-Tek using thermal expansion, oxidation and UV curing. PDMS based wavy culture plates were cured at room temperature for at least 2 days. 35mm circles were cut out from the cured PDMS and secured with more PDMS in a standard 35mm tissue dish. The plates were sterilized and prepped for culture. Laminin is deposited onto the plate, and silk sutures are pinned down. Longitudinal smooth muscle cells were cultured separately and seeded on to the plate, and allowed to align along the axis of the waves on the plate. Once alignment was microscopically confirmed (approximately four days post cell seeding), a fibrin hydrogel was added to promote delamination and subsequent aligned construct formation.

The micro patterns on the wavy plate were observed under a microscope, at 10x

magnification as shown in Figure 12A. Once sterilized, the plates were coated with 1-

2g/cm2 of laminin. Silk sutures dipped in dilute laminin were pinned down using insect pins

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as shown in Figures 12B. At this point, the plates were ready for cells to be seeded on to

them.

Figure 12C shows the cells on Day 0, 90 minutes after they have been seeded on to the

prepped wavy plates. The cells were randomly oriented and did not show any specific

alignment. Laminin promoted cell adhesion to the otherwise non-adhesive Sylgard surface.

Figure 12: Engineering highly aligned LSMC Sheets

A) 16x optical image of the PDMS mold fixed onto a 35 mm culture plate, with the wavy patterns seen as dark lines. B) The wavy plate was functionalized with laminin and silk sutures were pinned down 12mm apart. Growth media was added to the plate and left to sterilize under UV light before cells were plated on to them. C) 10x inverted microscope images show that cells were unaligned 1.5 hours after plating them, and show almost complete alignment on Day 3. Highly aligned cell sheet formation can be seen on Day 5; D) Addition of the fibrin gel on Day 5 promoted delamination (indicated by the black arrow) and the aligned cells contracted to form a highly longitudinally aligned construct 7 days after initial cell seeding, attached to the silk sutures.

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Cells showed partial alignment along the longitudinal axis of the mold 70 hours post initial

cell seeding as seen in Figure 12C. Between four and five days post initial cell seeding,

cellular alignment was complete, resulting in a 10cm2 cell sheet with completely aligned

RLSMC. The cell sheet formation was confirmed microscopically (Figure 12C Day 5) before

a fibrin gel was overlaid. The aligned cell sheet preferentially adhered to the hydrogel and

delaminated from the Sylgard surface and compacted into a bioengineered longitudinal

tissue (Figure 12D).

3.2 Immunofluorescence

Figure 13: Immunofluorescence on isolated RLSMC

A. α-Smooth Muscle Actin, a smooth muscle marker shows bright green stained stress fibers and smooth muscle actin.

B. Smooth muscle specific heavy Caldesmon shows bright green fluorescence. C. TRITC-conjugated ICC marker, c-Kit shows no positive areas, indicating the absence

of ICC cells in the longitudinal smooth muscle culture. D. Negative control of the TRITC-conjugated Rabbit secondary antibody, indicates

weak background staining like Tile c. Scale bar= 50µm

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Isolated RLSMCs were treated with Differentiation Media for 2 days, and assayed by

immunofluorescence to smooth muscle markers α-smooth muscle Actin (Figure 13A), and

smooth muscle specific heavy Caldesmon (Figure 13B). All the cells on the culture plate

stained brightly positive for both markers, indicating a population of smooth muscle cells.

Moreover, examination of the cells for interstitial cell marker (Figure 13C) showed a stark

negative, similar to the secondary antibody negative control (Figure 13D).

Figure 14: Passive tension and Signal to Noise Ratio (SNR)

(A) A trace of passive tension obtained from the bath medium without tissue. In order to determine the degree of system noise in the measurement, the bioengineered tissue was attached to the measurement system with no external stretch. (B) Active tension tracing. The SNR of the system, representing the degree of active tension is the ratio of mean signal strength to the standard deviation of the noise. SNR = ~250.

3.3 Acetylcholine Responses in RLSM bioengineered tissue

Addition of 1 M Acetylcholine (ACh) to the tissue bath with RLSM bioengineered tissue is

indicated by the arrows in Figure 15. Baseline was established prior to addition. Addition of

a compound into the 4mL tissue bath took up to 60s for its uniform diffusion.

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Figure 15: Force generation in response to 1µM Acetylcholine in bioengineered tissue and native longitudinal tissue

RLSMC sheets generated rapid-rising contractions of 94.4 ± 13.244 µN (n=7) in response to 1 µM Acetylcholine in regular calcium media. Contractions were sustained over a 12 minute period. Native longitudinal tissue also generated rapid rising contractions of 53.067 + 5.014 (n=6). Addition of ACh is indicated by the arrow at 180s. Representative tracings for bioengineered constructs and native tissue have been chosen.

The response of the bioengineered tissue (Figure 15A) is compared to the response

of native longitudinal tissue (Figure 15B). The average force generated in a bioengineered

tissue was 94.4 ± 13.244 N (n=7), while the average peak force in native tissue was 53.067

+ 5.014 (n=6). Bioengineered constructs (Figure 15A) and native tissue (Figure 15B) both

showed a rapid contraction, which was sustained for 12 minutes. Moreover, the tissue

displayed a dose-dependent response to Acetylcholine. This dose-dependence property

was maintained during the bioengineering process, as indicated by the similarity in response

of the bioengineered tissue and native tissue (Figure 16). Doses tested were between 1nM

and 1µM.

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Figure 16: Dose-Response to Acetylcholine in bioengineered tissue and native longitudinal tissue

Rabbit Longitudinal Tissue responded to Acetylcholine in a dose-dependent manner. Doses tested were between 1nM and 1µM. Bioengineered sheets from isolated longitudinal smooth muscle cells retained this dose-dependence property.

3.4 Response to Acetylcholine in the absence of extracellular Calcium

When regular Calcium media was replaced by Calcium-free PBS with 2mM chelating

agent EGTA, there was a ~60% inhibition in the magnitude of peak force generated in

response to 1µM Acetylcholine. Elevated magnitudes of forces were also sustained only

between 2.5-4 minutes, compared to the 12 minutes of sustained force in normal

extracellular calcium concentrations. The bioengineered tissue preserved the property of

native longitudinal smooth muscle’s classical dependency on extracellular calcium (20).

Representative tracings for the trend of force generation in response to 1µM Acetylcholine in

zero calcium is shown in Figure 17 for the bioengineered tissue, as well as native

longitudinal tissue.

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Figure 17: Response to Acetylcholine in Zero Calcium in bioengineered tissue and rabbit longitudinal tissue

Native longitudinal tissue generated only 16.57 + 2.095 µN (n=6) in response to 1µM ACh in the absence of extracellular calcium. This force was also sustained only for 2.5 minutes, versus the 12 minutes of force sustenance in regular calcium. Bioengineered tissue also showed force attenuated in magnitude by 50% and a reduction in time by 60%.

3.5 Response to Acetylcholine post Vasoactive Intestinal Peptide

Figure 18: Inhibition of force generation by pre-incubation with VIP in bioengineered tissue and rabbit longitudinal tissue

On treatment with VIP, both the native longitudinal tissue and the bioengineered tissue relaxed as indicated by the drop in baseline force. On subsequent stimulation with 1µM ACh, force generated was lowered by 40% in magnitude. Force was also sustained only for 5 minutes, versus the 12 minutes of force sustenance in control ACh responses.

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Treatment of bioengineered RLSMC cell sheets, as well as native longitudinal tissue,

with Vasoactive Intestinal Peptide (VIP) displayed a relaxation of baseline force. Pre-

incubation with VIP also attenuated the magnitude of force generated in response to 1µM

Acetylcholine by 40%. Bioengineered tissues generated an average peak force of 61.7 ± 8.7

µN (n=4) compared to 94.4 µN generated without the VIP pre-treatment. Moreover, elevated

forces were also sustained only for 5 minutes, versus the 12 minutes in control Acetylcholine

responses (50% inhibition). Representative traces are shown in Figure 18. A summary of the

values of force generated in response to Acetylcholine under various conditions is outlined

in the table accompanying Figure 19.

Figure 19: Comparison of force generated in response to Acetylcholine

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4. DISCUSSION

The myogenic component of altered colonic motility is smooth muscle contractility. In

altered motility resulting from inflammation, post-traumatic intestinal edema, etc., the

responses of circular smooth muscle layer have been characterized in detail previously. The

differences in response between the circular and longitudinal smooth muscle layers have

not been clearly outlined (21-23). This presents a strong requirement for an in vitro model of

longitudinal smooth muscle to enable the study of this muscle layer’s specific contribution to

abnormal colonic motility. We present a bioengineered physiologically responsive three-

dimensional reproduction of the longitudinal smooth muscle layer.

Common approaches in tissue engineering have utilized biodegradable, thermally

responsive, or even non-degradable scaffolds to engineer tissues (24). Regeneration of

tissues that have a primary functionality of force development requires scaffolds that

possess critical mechanical and structural properties resulting in complex and often

expensive fabrication processes. Drawbacks of conventional scaffold-based tissue

engineering apart from foreign body reactions also include precise control of porosity to

promote cell ingrowths and proliferation. Moreover, the use of biodegradable scaffolds

present the requirement of the scaffold to have controllable degradation rates in order to

regenerate tissue, before it degrades itself. Cells-in-gel culture techniques use extracellular

matrix (ECM) solutions as natural scaffolds to promote skeletal muscle tissue regeneration,

but these tissues lack physiological functionality(25, 26).

We report a physiologically functional model of longitudinal smooth muscle layer

from the colon. In our method of three-dimensional culture, we provide the longitudinal

smooth muscle cells with a micro-topographical surface to guide their alignment. This

alignment step was important to produce a cell sheet similar to the arrangement of

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longitudinal smooth muscle in vivo. The surfaces were patterned in a reproducible and

inexpensive manner using Sylgard (15, 16, 27) . Since cells present a lower adhesion on

hydrophobic Sylgard, the surface was coated with laminin to promote cellular attachment.

The surface topography aided in the alignment of the longitudinal cells to form a highly

oriented cell sheet. Figure 11 outlines the various steps involved in bioengineering the cell

sheet.

Original cell sheet engineering protocols used for skeletal muscle (16) were

optimized to fit the culture of longitudinal smooth muscle cells. Concentrations of laminin

coating the wavy plate surface and cell seeding densities were empirically determined and

optimized so that the smooth muscle cells adhered long enough to align themselves along

the axis of the micro patterns, mimicking alignment of longitudinal smooth muscle cells in

vivo. We used concentrations between 1-2 µg/cm2 to avoid chances of premature

delamination of the longitudinal cell layer, and seeded 500,000 cells per 35mm culture plate.

The cells saturated the surface of the culture plate and there was no room for proliferation.

Up to 100,000 cells were lost in media changes while the cells were allowed to align. Over 4

days, a highly aligned cell sheet was formed, in line with the wavy Sylgard topography

(Figure 12C).

The time in culture the fibrin hydrogel was overlaid was determined empirically for

different laminin concentrations. The optimal time for addition of the fibrin hydrogel was on

Days 4. At this point, the cells had adhered to the surface long enough to align along their

longitudinal axis and had begun to delaminate. Overlaying the aligned cells with the fibrin

gel coincided with the delamination, causing the cells to preferentially integrate and

proliferate in the fibrin gel layer. Subsequent contraction of the fibrin gel due to inherent

smooth muscle contractility and the resulting mechanical strain formed a compact

longitudinal smooth muscle cell sheet in 3-4 days post gel addition (Figure 12D). Fibrin gels

75

mimic the complexity of natural ECM and aid cell migration and ECM synthesis (28).

Functional skeletal muscle and cardiac pressure constructs were previously bioengineered

using fibrin scaffolds (10, 11, 29, 30). The overall rate of bioengineered tissue formation

depended on variables like coating laminin concentration, cell seeding density, overlaying

fibrin gel concentration and the time in culture at which the fibrin gel was overlain.

At the start of the delamination process, growth media was substituted with

Differentiation Media, primarily comprising of Medium 199 to promote the preservation of

smooth muscle phenotype. Phenotype maintenance was assayed on isolated RLSM cells

that were treated with differentiation media. The positive staining of α-smooth muscle actin

and h-Caldesmon revealed that the initial populations of cells used for bioengineering

maintained differentiated smooth muscle phenotype. The absence of staining with c-Kit

further indicated that our initial preparation was devoid of the presence of other any other

cell types, including Interstitial Cells of Cajal.

Bioengineered RLSMC tissues were compared with longitudinal smooth muscle

tissue directly isolated from the rabbit. 1cm strips were chosen, so as to match closely to the

dimensions of the bioengineered tissues.

Bioengineered three-dimensional RLSMC tissue responded to Acetylcholine (ACh) in

a functionally similar manner to longitudinal smooth muscle tissue directly isolated from the

rabbit. 1M ACh evoked a contractile response, generating 94.4 ± 13.244 N of force

(Figure 15). Bioengineered tissue also showed a dose-dependence property in its response

to ACh, like longitudinal tissue (Figure 16). Moreover, pre-incubation of the bioengineered

tissue with 1µM Atropine sulphate resulted in a complete inhibition of ACh-induced

contraction (data not shown). These results indicate not only that the bioengineered tissues

were capable of generating force, but also that they preserved receptor activated G-protein

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mediated responses to ACh. Various biochemical pathways resulting in smooth muscle

contraction in response to the contractile agonist, ACh, have been documented elsewhere

(31-34), with a commonality of downstream phosphorylation of Myosin Light Chain and

activation of Protein Kinase C (PKC). Bioengineered tissue also produced a rapid

contraction due to membrane depolarization by 30mM KCl and direct activation of PKC by

membrane permeable Phorbol ester, Phorbol dibutyrate (PdBU) (data not shown). This

demonstrates the ability of the bioengineered tissue to match the response of a native

tissue.

The driving force of smooth muscle contraction is the increase in intracellular calcium

concentration [Cai2+]. Key differences between circular and longitudinal smooth muscle arise

from their dependency on extra cellular calcium to mobilize intracellular calcium. In

longitudinal smooth muscle, agonist induced stimulation of Phospholipase A2 and

subsequent contraction requires a mandatory calcium influx step to produce calcium

induced calcium release from the sarcoplasmic reticulum mediated by Ryanodine receptors

(20). Incubation of bioengineered RLSMC tissue in zero-calcium buffer with 2mM EGTA

caused it to generate a significantly lower force in response to the same concentration of

ACh as before. The generation of an average force of 48 ± 9.47 µN (bioengineered tissue)

and 16.57 ± 2.09 µN (native tissue) even in the absence of extracellular calcium could

suggest the possible involvement of muscarinic receptor coupled G-protein activation of

Phospholipase D (PLD). PLD activation has been known to mediate the hydrolysis of

phosphatidylcholine, a ubiquitous phospholipid producing Diacylglycerol (DAG) without

accompanied production of IP3 (20).The marked absence in IP3 production would not

present any release of calcium from sarcoplasmic reticulum stores. DAG could lead to PKC

activation, thus leading to a downstream contraction in a manner that could be Calcium

77

independent (32, 35), in response to ACh. The bioengineered tissue behaved like native

tissue in its fundamental dependence on extracellular calcium.

Relaxation of longitudinal smooth muscle cells mediated by peptide neurotransmitter

VIP has been well documented (36-38). The bioengineered tissue relaxed to VIP like the

native longitudinal tissue (Figure 8). Pre-treatment of bioengineered RLSMC tissue with VIP

inhibited the magnitude of force generation elicited by 1M ACh by 40%. The initial

response to Acetylcholine was still maintained even in the presence of VIP, but forces were

sustained only for an overall 5 minutes, versus 12 minutes in ACh treatments that did not

include a VIP pre-treatment. This could indicate that VIP has a general inhibition pattern

without inhibiting a specific contraction pathway, thereby not affecting the initial contraction

response but producing a net overall inhibition of sustained contraction. This result is in line

with previous research, that shows VIP mediated Heat Shock Protein-20 phosphorylation

sustained over 30 minutes, causing relaxation and a subsequent “force suppression” in

colonic smooth muscle cells (17). The bioengineered tissue behaved in a functionally similar

manner to native tissue in response to VIP and post-VIP Acetylcholine induced force

generation. The physiology of the bioengineered tissue can be summarized as being

responsive to receptor-mediated cholinergic (Ach) and VIP-inergic stimulation. Additionally,

direct non-receptor mediated activation of Protein Kinase C-α by phorbol ester also

produced contractions. The response to potassium chloride was further testimony to the

preservation of physiologically relevant membrane ion-channels in the bioengineered tissue.

In conclusion, longitudinal smooth muscle cell sheets have been bioengineered,

allowing an extra cellular alignment step in culture, to mimic in vivo cellular alignment. This

is the first report of a three-dimensional colonic longitudinal smooth muscle model. The

aligned cell sheet promoted the formation of essential mature cellular connections between

smooth muscle cells. The cell sheet utilizes isolated longitudinal cells expanded in vitro

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using tissue culture methods. Force generation patterns revealed that bioengineered tissue

responded to physiologically relevant substances like Acetylcholine and VIP in a functionally

similar manner to longitudinal tissue isolated from the animal. This implies the preservation

of structural receptors, for agonist mediated G-protein coupled contraction/relaxation

pathways. Bioengineered tissues also maintain the characteristic of native longitudinal

smooth muscle tissue in their dependence on extra cellular calcium. The tissue engineered

cell-sheet could be used as a readily implantable “band-aid” tissue, along with local delivery

of FGF-2 to promote angiogenesis. This would ensure the maintenance of cell survival and

tissue viability. The three dimensional aligned LSMC cell sheet is also a reliable in vitro

model to study abnormal colonic motility.

Acknowledgements

This study was supported by National Institutes of Health grant RO1 DK-057020. We thank

Arti Dhiraaj for technical assistance.

References

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8. Zimmermann WH, Fink C, Kralisch D, Remmers U, Weil J, Eschenhagen T. Three-dimensional engineered heart tissue from neonatal rat cardiac myocytes. Biotechnology and Bioengineering.68:106-14. 1999. 9. Dennis RG, Kosnik PE, Gilbert ME, Faulkner JA. Excitability and contractility of skeletal muscle engineered from primary cultures and cell lines. American Journal of Physiology Cell Physiology.280. 2001. 10. Baar K, Birla R, Boluyt MO, Borschel GH, Arruda EM, Dennis RG. Self-organization of rat cardiac cells into contractile 3-D cardiac tissue. FASEB Journal.19:275-7. 2004. 11. Huang Y-C, Dennis RG, Larkin L, Baar K. Rapid formation of functional muscle in vitro using fibrin gels. Journal of Applied Physiology.98:706-13. 2005. 12. Hecker L, Baar K, Dennis RG, Bitar KN. Development of three-dimensional physiological model of the internal anal sphincter bioengineered in vitro from isolated smooth muscle cells. American Journal of Physiology Gastrointestinal and Liver Physiology.289:188-96. 2005. 13. Dennis RG, Kosnik PE. Excitability and isometric contractile properties of mammalian skeletal muscle constructs engineered in vitro In Vitro Cellular & Developmental Biology-Animal.36:327-35. 2000. 14. Kosnik PE, Faulkner JA, Dennis RG. Functional development of engineered skeletal muscle from adult and neonatal rats. Tissue Engineering.7. 2001. 15. Lam MT, Sim S, Zhu X, Takayama S. The effect of continuous wavy micropatterns on silicone substrates on the alignment of skeletal muscle myoblasts and myotubes. Biomaterials.27:4340-7. 2006. 16. Lam MT, Huang Y-C, Birla RK, Takayama S. Microfeature guided skeletal muscle tissue engineering for highly organized 3-dimensional free-standing constructs. Biomaterials.30:1150-5. 2009. 17. Gilmont RR, Somara S, Bitar KN. VIP induces PKA-mediated rapid and sustained phosphorylation of HSP20. Biochemical and Biophysical Research Communications.375:452-6. 2008. 18. Somara S, Bitar KN. Phosphorylated HSP27 modulates the association of phosphorylated caldesmon with tropomyosin in colonic smooth muscle. American Journal of Physiology Gastrointestinal and Liver Physiology.291:630-9. 2006. 19. Somara S, Pang H, Bitar KN. Agonist induced association of tropomyosin with protein kinase Cα in colonic smooth muscle. American Journal of Physiology Gastrointestinal and Liver Physiology.288:268-76. 2005. 20. Murthy KS. Signaling for contraction and relaxation in smooth muscle of the gut. Annual Reviews in Physiology.68:345-74. 2005. 21. Grossi L, McHugh KM, Collins SM. On the specificity of altered smooth muscle function in experimental colitis in rats. Gastroenterology.104. 1993. 22. Myers BS, Martin JS, Dempsey DT, Parkman HP, Thomas RM, Ryan JP. Acute experimental colitis decreases colonic circular smooth muscle contractility in rats American Journal of Physiology Gastrointestinal and Liver Physiology.273:928-36. 1997. 23. Moore-Olufemi SD, Padalecki J, Olufemi SE, Xue H, Oliver DH, Radhakrishnan RS, et al. Intestinal Edema: Effect of Enteral Feeding on Motility and Gene Expression. Journal of Surgical Research.2008:1-10. 2008. 24. Fedorovich NE, Alblas J, Wijn JRD, Hennink WE, Verbout AJ, Dhert WJA. Hydrogels as Extracellular Matrices for Skeletal Tissue Engineering: State-of-the-Art and Novel Application in Organ Printing. Tissue Engineering.13:1905-25. 2007. 25. Vandenburgh HH. Bioengineered muscle constructs and tissue-based therapy for cardiac disease. Progress in Pediatric Cardiology.21:167-71. 2005. 26. Okano T, Matsuda T. Tissue engineered skeletal muscle: preparation of highly dense, highly oriented hybrid muscular tissues. Cell Transplantation.7:71-82. 1998.

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27. Lam MT, Clem WC, Takayama S. Reversible on-demand cell alignment using reconfigurable microtopography. Biomaterials.29:1705-12. 2008. 28. Ye Q, Zuend G, Benedikt P, Jockenhoevel S, Hoerstrup SP, Sakyama S, et al. Fibrin gel as a three dimensional matrix in cardiovascular tissue engineering. European Journal of Cardio-thoracic Surgery.17:587-91. 2000. 29. Birla RK, Huang Y-C, Dennis RG. Development of a Novel Bioreactor for the Mechanical Loading of Tissue-Engineered Heart Muscle. Tissue Engineering.13:2239-48. 2007. 30. Shaikh FM, Callanan A, Kavanagh EG, Burke PE, Grace PA, McGloughlin TM. Fibrin: A Natural Biodegradable Scaffold in Vascular Tissue Engineering. Cells Tissues Organs.188:333-46. 2008. 31. An JY, Yun HS, Lee YP, Yang SJ, Shim JO, Jeong JH, et al. The intracellular pathway of the acetylcholine-induced contraction in cat detrusor muscle cells. British Journal of Pharmacology.137:1001-10. 2002. 32. Hillemeier AC, Deutsch DE, Bitar KN. Signal Transduction Pathways Associated with contraction during development of the feline gastric antrum. Gastroenterology.113:507-13. 1997. 33. Ding X, Murray PA. Acetylcholine activated Protein Kinase C-alpha in Pulmonary Venous Smooth Muscle. Journal of American Society of Anesthesiologists.106:507-14. 2007. 34. Patil SB, Bitar KN. RhoA- and PKCα-mediated phosphorylation of MYPT and its association with HSP27 in colonic smooth muscle cells. American Journal of Physiology Gastrointestinal and Liver Physiology.290. 2005. 35. Schmidt M, Huewe SM, Fasselt B, Homann D, Ruemenapp U, Sandmann J, et al. Mechanisms of phospholipase D stimulation by m3 muscarinic acetylcholine receptors: Evidence for involvement of tyrosine phosphorylation. European Journal of Biochemistry.225:667-75. 1994. 36. Kishi M, Takeuchi T, Suthamnatpong N, Ishii T, Nishio H, Hata F, et al. VIP- and PACAP-mediated nonadrenergic, noncholinergic inhibition in longitudinal muscle of rat distal colon: involement of activation of charybdotoxin- and apamin-sensitive K+ channels. British Journal of Pharmacology.119:623-30. 1996. 37. Kishi M, Takeuchi T, Katayama H, Yamazaki Y, Nishio H, Hata F, et al. Involvement of cyclic AMP–PKA pathway in VIP-induced, charybdotoxin-sensitive relaxation of longitudinal muscle of the distal colon of Wistar-ST rats. British Journal of Pharmacology.129. 2000. 38. Lefebvre RA, Smits GJ, Timmermans JP. Study of NO and VIP as non-adrenergic non-cholinergic neurotransmitters in the pig gastric fundus. British Journal of Pharmacology.116:2017-26. 1995.

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CHAPTER 6: EXTRACELLULAR MATRIX MODULATES

DIFFERENTIATION OF NEURONAL SUBTYPES IN TISSUE

ENGINEERED INNERVATED INTESTINAL SMOOTH MUSCLE

SHEETS

Shreya Raghavan1,2 and Khalil N. Bitar1,2

1Wake Forest Institute for Regenerative Medicine, Wake Forest School of Medicine, Winston-Salem NC 27101

2Virginia Tech-Wake Forest School of Biomedical Engineering and Sciences, Winston-Salem NC 27101

Running title: Neuronal differentiation in engineered intestinal sheets

Keywords: Neural stem cell, neural tissue engineering, extracellular matrix, adult stem cell, enteric nervous system

This manuscript was accepted for publication in Biomaterials (in press, June 2014)

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Introduction

An uninterrupted enteric nervous system with the preservation of myenteric ganglia

is required for intestinal motility and function [1, 2]. Motor neurons of the myenteric ganglia

pre-dominantly express acetylcholine/tachykinins (excitatory) or nitric oxide/inhibitory

peptides/purines (inhibitory) to mediate smooth muscle contraction and relaxation [3, 4].

Partial, selective or total loss of neurons is reported in several disorders including, but not

limited to, Hirschsprung’s disease, achalasia, and inflammation [5-8]. Neural-crest derived

enteric neuronal progenitor cells have been isolated from adult mammalian guts, including

ganglionic bowel of patients with Hirschsprung’s disease [9-12]. These cells have the ability

to differentiate into neuronal and glial phenotypes [13-15]. However, there is little information

and understanding of microenvironment driven differentiation and limited studies describing

subsequent functional behavior of these differentiated neurons in vitro [10, 11].

The ECM has been long known to provide permissive and non-permissive

environmental cues for migration and differentiation of neuronal progenitor cells [16].

Collagen IV, laminin and heparan sulfate proteoglycans play an important role in guiding the

migration of neural-crest derived cells and formation of the enteric nervous system [17-19].

Moreover, myenteric ganglia are surrounded by an ECM composed of type IV Collagen,

laminin and a heparan sulfate proteoglycan [20].

Type I fibrillar collagen has favorable biological properties and has been widely used

in neural tissue engineering [21, 22]. Collagen IV has been demonstrated to promote neurite

outgrowth in neuroepithelial progenitor cells and in sympathetic peripheral neurons in vitro

[23, 24]. Collagen IV aids the colonization of the embryonic gut and modulates selective

neurotrophic signaling [20, 25]. The IKVAV peptide in laminin interacts with a neurally

acquired receptor on post-migratory neural crest stem cells promoting their differentiation

[26]. In the gut, heparan sulfate proteoglycan is required for GDNF signaling [27].

Glycosaminoglycan – growth factor interactions additionally stabilize and increase local

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bioavailability of growth factors, thus inducing neurite outgrowth [28]. The ECM can provide

a background upon which cell-cell and cell-matrix signaling can work to regulate phenotypes

of differentiating enteric neuronal progenitor cells.

In previous studies using 2D culture substrata, we demonstrated that varying ECM

composition of 2D culture substrata influenced neuroglial differentiation of adult enteric

neuronal progenitor cells [29]. The number of neurons, neurite lengths and preliminary

neuronal network formation were all enhanced in culture substrata that contained collagen

IV, laminin and heparan sulfate. The previous studies, however, were carried out on 2D

coated glass coverslips. Moreover, we examined neuronal and glial differentiation, without

documenting the phenotype or functionality of the differentiated neurons. In the present

studies, we utilize tissue engineering as a tool to provide enteric neuronal progenitor cells

with 3D viscoelastic ECM microenvironments. Given that the cues for differentiation arising

from a contractile phenotype of smooth muscle cells would be constant, we hypothesized

that ECM composition could differentially regulate the generation of specific neuronal

subsets. The objective of the current undertaking was to determine the presence of

differentiated motor neurons obtained and investigate their ability in mediating smooth

muscle contraction/relaxation.

We utilize a previously described tissue engineered model of intestinal longitudinal

smooth muscle sheets, where uniaxial alignment of a smooth muscle monolayer was

facilitated by the use of substrate microtopography [30]. We innervated these tissue

engineered sheets using enteric neuronal progenitor cells embedded with hydrogels of

varying ECM composition. Differentiated neuronal composition (cholinergic, nitrergic,

peptidergic) within tissue engineered sheets was evaluated. Functional neuronal physiology

mediating smooth muscle contraction/relaxation was also evaluated in the tissue engineered

sheets using real-time force generation measurements.

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2. Materials and Methods

2.1 Materials

All tissue culture reagents (including media, supplements, and natural mouse laminin) were

purchased from Invitrogen (Carlsbad, CA). Rat tail type I collagen and mouse collagen type

IV were purchased from BD Biosciences (Bedford, MA). Heparan sulfate was purchased

from Celsus (Cincinnati, OH). Growth factors were purchased from Stemgent (Cambridge,

MA). All primary antibodies were purchased from Abcam (Cambridge, MA), unless specified

otherwise.

2.2 Isolation and primary culture from adult rabbit GI tissues

Enteric neuronal progenitor cells were isolated from jejunal biopsies of adult New Zealand

White rabbits using a collagenase/dispase digestion technique and cultured in neuronal

growth media, as described previously [29]. Cells aggregated to form enteric neurospheres.

Longitudinal smooth muscle cells were isolated from the adult rabbit sigmoid colon as

described previously [30]. Isolated longitudinal smooth muscle cells were expanded in

culture until confluency.

2.3 Composition and characterization of ECM hydrogels

ECM hydrogels were made with the following components:

i) Collagen I gels (800-1600µg/ml);

ii) Collagen I (800µg/ml) and Collagen IV (200µg/ml) gels composite gels;

iii) Collagen I and Collagen IV with laminin (5-10µg/ml);

iv) Collagen I and Collagen IV with laminin and heparan sulfate (10-20µg/ml).

Other components of the gel included: 1% fetal calf serum, 0.1X antibiotics in Dulbecco’s

modified Eagle’s medium. 0.1N Sodium hydroxide was used to adjust pH to ~7.4 for

gelation.

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2.3.1 Rheological characterization of ECM hydrogels: Oscillatory rheometry (ATS

RheoSystems) was used to measure viscoelastic moduli of ECM gels. 20mm parallel base

plates were used to perform a stress sweep of the sample at 1Hz. ECM gels were allowed to

gel in situ between the parallel plates at 37°C. The viscoelastic modulus was obtained from

a linear region of the stress-strain curve, at strains lower than 10%, within the sensitivity

ranges for torque and strain of the rheometer. 3-5 individually manufactured ECM gels were

measured to determine an average viscoelastic modulus. Compositions that resulted in a

matrix viscoelasticity within the range of 150-300Pa were utilized for further experimentation,

so as not to let stiffness be a variable in influencing neuroglial differentiation.

2.3.2 Characterization of ultrastructure of ECM hydrogels: Sample preparation of ECM

hydrogels for scanning electron microscopy was adapted from Stuart et al. [31]. Gels were

dehydrated through graded ethanol (10% to 100%). Hydrogels were dried at critical point

using carbon dioxide exchange. The resulting dehydrated ECM discs were mounted onto

metallic stubs with conducting carbon tape, sputter coated with gold, and visualized using an

AMRAY 1910 Field Emission Scanning Electron Microscope. Constant working distance and

magnification were maintained to image all samples. NIH Image J was used to measure and

compare fiber diameters. Porosity was determined using Image J from micrographs

obtained from at least three-independent samples of dehydrated ECM gels.

2.4 Tissue engineering innervated intestinal smooth muscle sheets

The tissue engineering process was adapted from Raghavan et al. [30]. Briefly, 500,000

longitudinal smooth muscle cells were aligned uniaxially for 4 days on 35mm diameter

circular Sylgard molds containing wavy microtopographies. Enteric neurospheres were

treated with Accutase to obtain single cell suspensions. 200,000 cells were resuspended in

the appropriate ECM solution and overlaid on the aligned smooth muscle monolayer. Upon

gelation, neuronal differentiation medium (neurobasal-A) was added, supplemented with

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B27 and 1% fetal bovine serum. Differentiation medium was exchanged every second day.

Enteric neuronal progenitor cells were allowed to differentiate within the hydrogel for a

period of 10 days. Smooth muscle cells compacted the ECM hydrogel over the next 10

days, forming ~1cm long innervated smooth muscle sheets, anchored between silk sutures.

Phase microscopy was used to image neuronal differentiation at the edge of the tissue

engineered sheets.

2.5 Biochemical characterization of neuroglial composition in tissue engineered sheets

At day 10, tissue engineered sheets were harvested in radioimmunoprecipitation buffer to

isolate protein. Protein concentration was estimated spectrophotometrically using the

Bradford assay. 20µg of protein from each sample was resolved electrophoretically and

transferred to polyvinylidene difluoride membranes. Membranes were blotted with antibodies

for neuronal βIII Tubulin, neuronal nitric oxide synthase (nNOS), choline acetyltransferase

(ChAT), and Smoothelin. β-Actin was used to confirm equal loading. HRP-conjugated

secondary antibodies were used to visualize proteins using enhanced chemiluminescence.

2.6 Immunohistochemical characterization of neuron composition in tissue engineered

sheets

Tissue engineered sheets were fixed in 4% formaldehyde and washed extensively in glycine

buffer. Immunohistochemical staining was performed following previously established

protocols utilized for staining differentiated neurons within bioengineered tissues [32].

Sheets were blocked with 10% horse serum and permeabilized in 0.15% Triton-X for 45

minutes. Permeabilized sheets were incubated with primary antibodies directed against

Vasoactive Intestinal Peptide (VIP), ChAT and nNOS for 60 minutes at room temperature.

Following antibody incubation, sheets were washed three times with phosphate buffered

saline, pH 7.4. Tissue engineered sheets were incubated with appropriate fluorophore

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conjugated secondary antibodies for 45 minutes, washed in phosphate buffered saline and

imaged using an inverted fluorescence microscopy (Nikon Ti-E, Japan). For a negative

control, incubation with the primary antibody was skipped, and only fluorophore conjugated

secondary antibodies were used to visualize background fluorescence.

2.7 Measurement of physiological function in innervated tissue engineered sheets

Myogenic and neuronal functionality were assessed using real-time force generation as

previously described [30, 33]. 4-5 individual tissue engineered sheets for each ECM

composition were tested. Tissue engineered sheets were anchored between a stationary pin

and measuring pin of a force transducer (Harvard Apparatus, Holliston MA) at 0% stretch.

The organ bath maintained temperature at 37°C. An additional 10% stretch was applied

using a vernier control. Tissues were immersed in 4ml of medium, which was exchanged at

the end of every experiment following a brief wash with fresh medium. Peak contraction or

maximal relaxation was quantified following pharmacological or electrical stimuli, and

compared between tissue engineered sheets with varying ECM compositions.

Before each treatment, tissues were washed in fresh warm medium and allowed to

equilibrate to a baseline. The following stimuli were used independently to assess

physiological functionality of the tissue engineered sheets: 1) 60mM Potassium chloride to

assess electromechanical coupling integrity of the smooth muscle; 2) 1µM Acetylcholine

(contractile agonist); 3) Electrical field stimulation (5Hz, 0.5ms, 40V) applied using parallel

plate platinum electrodes. Preincubation with neuronal blocker, tetrodotoxin (TTX) was used

to dissect myogenic and neuronal components of contraction/relaxation. Pre-incubation with

specific inhibitors were used to identify functional neuronal subtypes: 1) nNOS-blocker Nω-

Nitro-L-arginine methyl ester hydrochloride (L-NAME; 300µM); and 2) VIP-receptor

antagonist [D-p-Cl-Phe6, Leu17]-Vasoactive Intestinal Peptide (VIP-Ra; 2µM). Following

stimulation and subsequent contraction/relaxation and recovery, tissues were washed with

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fresh medium, and allowed to re-establish a baseline before the next treatment. Equilibrated

baseline was arbitrarily set to zero, to measure contraction/relaxation due to a stimulus.

2.8 Data Analysis and statistical methods

Densitometry on western blots was performed using BioRad Quantity One (Hercules, CA).

Raw data was acquired from the force transducer at 1000 samples/second. Second order

Savitsky-Golay smoothing was applied to data using GraphPad Prism 5.0 for Windows

(GraphPad Software, San Diego, CA). Area under the curve (AUC) was measured from the

time of addition of pharmacological agonist/electrical field to the end of the

contraction/relaxation response. Extent of inhibition by pharmacological inhibitors was

calculated by expressing the AUC of contraction/relaxation in the presence of the inhibitor

as a percentage of the AUC in the absence of the inhibitor. One way ANOVAs with Tukey

post-tests were used to compare means using GraphPad Prism. p ≤ 0.05 was considered

significant. Physiological evaluation and densitometry was carried out between 3-5 tissue

engineered sheets within each experimental set; all values are expressed as mean ± SEM.

3. Results

3.1 Ultrastructure and viscoelastic properties of ECM hydrogels

All compositions of ECM hydrogels gelled at 37°C within 30 minutes. Scanning electron

micrographs revealed a fibrous structure in type I Collagen gels (Fig 20A). The fibers were

randomly oriented, with diameters averaging at 478.3 ± 19.31µm. With the addition of type

IV Collagen, network-like structures were observed (Fig 20B).

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Figure 20: Scanning electron micrographs of dehydrated ECM gels

Images were obtained at constant magnification and constant working distance. (A)

Collagen I fibers were randomly oriented, and presented a dense fibrous structure; (B)

Composite Collagen I/IV sheets demonstrated evidence of the formation of network-like

structures; (C) There was no difference in ultrastructure with the addition of laminin; (D)

Evidence of cabling and cross-linking was observed with the addition of heparan sulfate.

Average porosity was determined and summarized in Table 1. Viscoelastic modulus was

calculated using oscillatory rheometry of ECM gels in their hydrated state, and tabulated in

Table 1. Scale bar 10µm.

Cables of fibers within the networked structures were thicker, with average diameters of

714.8 ± 36.67µm. Addition of laminin to the hydrogels did not alter the ultrastructure or the

networked suprastructure (Fig 20C). With the addition of heparan sulfate, the fibers within

the networked structures were pulled more tightly together and cabled (Fig 20D). The

dehydrated ECM gels displayed a porous appearance, with average porosity ranging from

40.77% - 43.95% (Table 1).

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Table 1: Viscoelastic modulus of ECM gels

Viscoelastic moduli were measured in hydrated ECM gels using oscillatory rheometry. Type

I Collagen gels had increasing viscoelastic moduli with increasing collagen concentration

ranging from 72.6 ± 4.86Pa (800µg/ml) to 182.3 ± 2.6 (1600µg/ml) to 424 ± 2 Pa

(3200µg/ml). The addition of 200µg/ml collagen IV to 800µg/ml collagen I increased the

modulus of the gels to 236 ± 13.53 Pa. The addition of laminin had no effect on viscoelastic

moduli (compare 236 ± 13.53 Pa to 220.7 ± 16.27 Pa). 10µg/ml of heparan sulfate caused

an increase in the modulus of ECM hydrogels (287 ± 20.11 Pa, p<0.05). Table 1

summarizes that the final ECM gels evaluated had viscoelastic moduli ranging from 182Pa

to 287Pa.

3.2 Neuronal differentiation in engineered innervated intestinal smooth muscle sheets

Uniaxially aligned smooth muscle cells compacted overlaying ECM hydrogels over 10 days

in culture as described before [30]. The resultant tissue engineered sheets were ~1cm long,

and a few cell layers thick. In the presence of smooth muscle, the enteric neuronal

progenitor cells differentiated within the ECM hydrogel. Neuronal differentiation was

identified morphologically by microscopic examination at day 10, demonstrating similar

differentiation profiles expressed by enteric neuronal progenitor cells, both in vitro [29] and

in tissue engineered constructs [32]. Several differentiated neurons were observed in tissue

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engineered sheets, regardless of the ECM composition (Fig 21). Arrows in the figures

indicate numerous instances of neuronal clustering and preliminary neuronal networking.

Figure 21: Neuronal differentiation within tissue engineered sheets

Phase contrast micrographs were obtained at the edge of the tissue engineered sheets. Evidence of neuronal differentiation and initiation of network formation was observed in all ECM compositions at day 10. The arrows indicate instances of preliminary neuronal networking. Scale bar 200µm.

3.3 Neuronal composition in engineered innervated intestinal smooth muscle sheets

Immunoblotting was used to assess neuronal composition within tissue engineered

innervated intestinal smooth muscle sheets. Blotting for β-actin demonstrated that equal

amounts of protein were assayed. Representative blots for each protein are shown,

indicating the approximate molecular weight at which they appear on the gels (Fig 22E).

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Contractile phenotype of constituent smooth muscle was demonstrated by the similar

expression of smoothelin, within the tissue engineered sheets (Fig 22B). The expression of

Smoothelin was constant, regardless of the ECM composition of the sheets, indicating that

the constituent smooth muscle cells maintained a contractile phenotype.

Figure 22: Immunoblot analysis of tissue engineered longitudinal sheets

Sheets were assessed for expression of neuronal differentiation, constituent smooth muscle phenotype, and excitatory and inhibitory neural markers. Densitometry was used to quantify band intensities, to quantify and compare expression. (A) Neuronal βIII Tubulin expression was similar amongst all four matrices suggesting that all ECM compositions supported neuronal differentiation; (B) Constituent smooth muscle within the tissue engineered sheets maintained contractile phenotype, demonstrated by similar Smoothelin expression; (C) Choline acetyltransferase (ChAT) expression was significantly (*p<0.05) elevated in Col I and Col I/IV/Laminin sheets compared to Col I/IV and Col I/IV/Lam/HS sheets; (D) Neuronal nitric oxide synthase (nNOS) expression was significantly lower (**p<0.001) in Col I sheets compared to elevated levels in all tissue engineered sheets containing Col4 with or without laminin and/or heparan sulfate. (E) Representative immunoblots are provided along with β Actin, demonstrating equal loading.

3.3.1 Neuronal differentiation: Pan neuronal marker βIII Tubulin expression was similar

amongst all tissue engineered sheets, despite the ECM composition (Fig 22A). This

suggested that irrespective of the ECM composition, neuronal differentiation of enteric

neurospheres proceeded similarly in the presence of smooth muscle cells. βIII Tubulin

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expression ranged from 21.65± 1.43AU – 28.98 ±0.85 AU. βIII Tubulin expression was

similar amongst various ECM gel compositions (ns; Fig 22A), indicating similar neuronal

differentiation.

3.3.2 Cholinergic neurons: Choline acetyltransferase (ChAT) expression was used to detect

the presence of cholinergic neurons (Fig 22C). Collagen I (33.73 ± 1.13 AU) and collagen

I/IV/laminin (28.82 ± 1.21 AU) sheets had a significantly elevated expression of ChAT

compared to sheets with composite collagen and/or heparan sulfate. Immunoblotting

demonstrated an enriched cholinergic neuron population in tissue engineered sheets

manufactured with collagen I only or composite collagen I/IV with laminin. The presence of

cholinergic neurons was additionally confirmed using immunohistochemistry (Fig 23E-H).

Figure 23: Immunofluorescence for differentiated neurons within tissue engineered sheets

Differentiated neurons within tissue engineered sheets were stained with markers for vasoactive intestinal peptide (VIP; red), choline acetyltransferase (ChAT; red) or neuronal nitric oxide synthase (nNOS; green). (A-D) Numerous differentiated VIP-ergic neurons were present in tissue engineered sheets. (E-H) Differentiated excitatory cholinergic neurons expressing ChAT were present within tissue engineered sheets; (I-L) Differentiated inhibitory nitrergic neurons expressing nNOS were present within tissue engineered sheets. Scale bar 100µm.

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3.3.4 Nitrergic inhibitory motor neurons: Neuronal nitric oxide synthase (nNOS) expression

was used to detect the presence of inhibitory nitrergic motor neurons (Fig 22D). Sheets with

collagen IV (with or without laminin/heparan sulfate) had a significantly higher nNOS

expression ranging from 26.37 ± 1.29 AU – 28.15 ± 2.69 AU. Conversely to ChAT, collagen I

sheets had minimal nNOS expression (11.33 ± 2.85 AU). Presence of nNOS was

additionally confirmed using immunohistochemistry (Fig 23I-L).

3.3.4 VIP-ergic inhibitory motor neurons: Vasoactive intestinal peptide (VIP) motor neurons

were identified using immunohistochemistry. VIP neurons were abundant, with increased

immunofluorescence in composite hydrogels with laminin and heparan sulfate (Fig 23A-D).

3.4 Agonist-induced contractility of tissue engineered innervated smooth muscle

sheets

3.4.1: Potassium Chloride-induced contraction

Electromechanical coupling integrity of constituent smooth muscle cells was first evaluated

using potassium chloride (KCl). KCl treatment elicited rapid contractions that were sustained

for ~5 minutes (Fig 24). Peak maximal contraction in response to KCl was similar between

the different tissue engineered sheets (Fig 24A-D), ranging from 279.5 ± 4.79µN to 296.5 ±

6.26µN. This correlated with the equivalent expression of contractile smooth muscle marker,

Smoothelin, indicating that the constituent smooth muscle cells within the tissue engineered

sheets maintained a contractile phenotype regardless of ECM composition. Furthermore,

KCl-induced contractions in tissue engineered sheets were similar to native rabbit intestinal

tissues (Fig 24E) in time course, but slightly reduced in magnitude. Peak KCl-induced

contractions in native tissue averaged 373.5 ± 10.63µN.

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Figure 24: Potassium chloride induced contraction of tissue engineered sheets

60mM Potassium chloride (KCl) was used to examine the electromechanical coupling integrity of the constituent smooth muscle cells within the tissue engineered sheets. The black traces indicate the contraction in response to the addition of KCl. The red traces indicate the addition of KCl in the presence of a neuronal blocker, TTX. Pre-treatment with TTX did not inhibit KCl-induced contraction. The ECM composition of the tissue engineered sheets did not affect smooth muscle contraction, evidenced by similar contractile patterns in response to KCl stimulation. A robust and immediate contraction was observed upon addition of KCl (indicated by the arrows) in all tissue engineered sheets (A-D), similar to native rabbit intestinal tissue (E). Peak contraction in response to KCl ranged between 279.5µN and 296.5µN in tissue engineered sheets, and averaged at 373.3 ± 10.63 in native tissue. (F) The area under the curve of KCl-induced contraction was quantified to demonstrate no significant (ns) difference in contraction in tissue engineered sheets, with a slightly elevated (*p<0.05) magnitude in native tissue.

KCl-induced contraction was unaffected by pre-treatment with neuronal blocker TTX (red

traces; Fig 24), indicating myogenic electromechanical coupling integrity. Figure 24F

demonstrates that the area under the curve of contraction was similar in all tissue

engineered sheets, and was significantly higher in native rabbit intestinal tissues. Although

reduced in magnitude compared to native tissue, KCl-induced contractions were similar

among the different tissue engineered sheets, indicating a robust contractile smooth muscle

phenotype unaffected by the ECM composition.

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3.4.2: Acetylcholine-induced contraction

Exogenous addition of 1µM Acetylcholine (Ach) was used to simulate agonist-induced

contraction. All tissue engineered sheets contracted in response to Ach, and sustained

contractions up to ~ 5 minutes post stimulation with Ach (Fig 25A-D). Tissue engineered

sheets with composite collagen I/IV with laminin had a significantly elevated peak maximal

Ach-induced contraction (Fig 25C; 232.9 ± 8.167µN), as well as an elevated area under the

curve of contraction (47606 ± 2054 AU).

Figure 25: Acetylcholine induced contraction

Addition of 1µM Acetylcholine (Ach; arrow) resulted in contraction of tissue engineered sheets, as well as native tissue. Red traces demonstrate Ach treatment in the presence of neuronal blocker, TTX. (A-E) Representative tracings of Ach-induced contraction in tissue engineered sheets and native tissue. Magnitude of Ach-induced contraction varied between tissue engineered sheets. Comparison of the area under the curve of contraction demonstrated that tissue engineered sheets approached 31.5% (Col4) - 67.6% (Laminin) of contraction observed in native tissue. In the presence of TTX, magnitude of Ach-induced contraction was attenuated. Quantification of inhibition (F) revealed that the degree of inhibition with TTX varied amongst the tissue engineered sheets with different ECM compositions. Highest inhibition was observed in Col i (72.77 ± 2.5%) and Col I/IV/Laminin (60.58 ± 1.7%) sheets, indicating an elevated presence of cholinergic neurons contributing to Ach-induced contraction. Significantly lower inhibition (*p<0.05; 48.36 ± 4.3 – 50.31 ± 4.2%) was seen in Col I/IV and Col I/IV/Lam/Heparan sulfate sheets. TTX pre-treatment attenuated Ach-induced contraction by 72.73 ± 3.7% in native tissue.

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Magnitude of Ach-induced contraction was still significantly lower compared to contraction in

native tissue (342.6 ± 3.15µN; 70448 ± 5876 AU). However, the time course of contraction

was very similar to native tissue in tissue engineered sheets containing laminin, reaching

maximal contraction within a minute of agonist stimulation. Collagen I sheets also had an

elevated Ach-induced contraction (Fig 25A; 238.9 ± 13.72 µN; 42668 ± 2172 AU)

corresponding to the elevated ChAT protein expression. However, the kinetics of contraction

did not match native tissue.

In order to estimate the smooth muscle (myogenic) component of Ach-induced contraction,

neurotoxin TTX was used as a pretreatment (red traces, Fig 25). Area under the curves of

contraction was compared with and without TTX pre-treatment in order to estimate

%inhibition (Fig 25F). %Inhibition of Ach-induced contraction in the presence of TTX was

highest in two ECM conditions: i) collagen I sheets (72.77 ± 2.45%); and ii) collagen

I/IV/laminin sheets (60.58 ± 1.66%). These values of %inhibition were similar to that

observed in native tissue (72.73 ± 3.66%) upon TTX-pretreatment. This increased neuronal

contribution to Ach-induced contraction also correlated with the elevated protein expression

of ChAT in collagen I and composite collagen I/IV/laminin sheets (Fig 22C). TTX-

pretreatment inhibited Ach-induced contraction to a significantly lower extent in collagen I/IV

± heparan sulfate sheets, ranging from 48.36 ± 4.36 % (Heparan sulfate) to 50.31 ± 4.22%

(collagen IV; Fig 25F).

3.5 Relaxation in engineered innervated sheets in response to electrical field

stimulation

Electrical field stimulation (EFS) at 5Hz, 0.5ms was used to stimulate neurons within the

tissue engineered sheets to produce relaxation of smooth muscle (Fig 26). The extent of

relaxation was quantified as area under the curve of relaxation.

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Table 2: Acetylcholine-induced contraction in tissue engineered sheets

Extent of relaxation significantly varied amongst the tissue engineered sheets with varying

ECM compositions. Sheets bioengineered with collagen IV, which displayed elevated nNOS

expression, had higher relaxation compared to sheets bioengineered with collagen I only

(compare 109693 ± 8465 AU in collagen I/IV sheets to 23142 ± 4921 in collagen I sheets).

Sheets containing laminin and/or heparan sulfate also had significantly elevated relaxation

compared to collagen I sheets (68395-69025 AU).

In response to EFS, native tissues relaxed generating 101550 ± 11279 AU. Tissue

engineered sheets with collagen IV and/or laminin and/or heparan sulfate additionally had a

time course of relaxation most similar to native tissue. Maximal relaxation was achieved

within 2 minutes of EFS, and a subsequent recovery of basal force was complete within 10

minutes. Upon pre-treatment with TTX, EFS-induced relaxation was inhibited entirely (red

traces, Fig 26).

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Figure 26: Electrical Field Stimulation induced relaxation in tissue engineered sheets

Electrical Field stimulation (EFS; shaded gray area) was used to stimulate relaxation in tissue engineered sheets (A-D) and native tissue (E). Red traces indicate TTX pre-treatment. EFS induced relaxation was significantly attenuated by TTX-pretreatment (90.9 ± 2.41% - 94.41 ± 0.93%), indicating that differentiated neurons within the tissue engineered sheets were capable of evoking smooth muscle relaxation. The magnitude of relaxation varied amongst the tissue engineered sheets (summarized in Table 3). (F) Quantification of the area under the curve of relaxation indicated that Col I sheets had a significantly low (***p<0.001; 23142 ± 4921AU) magnitude of relaxation. Relaxation in Col I/IV sheets (109693 ± 8465AU) were similar to those observed in native tissue (101550 ± 11279AU) in response to the electrical field, indicating the presence of elevated levels of inhibitory motor neurons capable of mediating relaxation. Relaxation was higher in Col I/IV/Laminin (69025 ± 7154AU) and Col I/IV/Lam/Heparan sulfate (68395 ± 8228AU) sheets, when compared to Col I, also indicating a similar increase in the presence of inhibitory motor neurons.

3.5.1 Inhibition of nitric oxide synthase

In order to identify the presence and functionality of nitrergic neurons, an inhibitor of nitric

oxide synthase (L-NAME) was used (green traces, Fig 27). %inhibition was determined by

comparing areas under the curves of maximal relaxation with and without the L-NAME pre-

treatment. %inhibition with L-NAME treatment was the lowest in collagen I sheets (33.37 ±

8.37%; green trace, Fig 27A). This corresponded to the low protein expression of nNOS in

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collagen I sheets compared to sheets containing collagen IV (Fig 22D). In contrast, the

inhibition of nNOS activity attenuated relaxation upto 61.71 ± 2.82% (green trace, Fig 27B)

in collagen I/IV sheets. In sheets containing laminin and heparan sulfate, %inhibition with L-

NAME varied between 62.28 ± 2.75% (laminin, Fig 26C) to 57.16 ± 1.91% (heparan sulfate).

This inhibition is significantly elevated compared to collagen I sheets, corresponding to the

increased expression of nNOS observed in the collagen I/collagen IV sheets (Fig 22C).

Native tissues had a higher %inhibition with L-NAME (78.02 ± 2.85%).

Figure 27: Inhibition of relaxation with L-NAME

The functionality of nitrergic neurons was studied by inhibiting EFS-induced relaxation with L-NAME, a non-metabolizable substrate for nNOS. The green traces indicate EFS in the presence of L-NAME. Pretreatment with L-NAME attenuated EFS-induced relaxation in all tissue engineered sheets (A-D) and native tissue (E). (F) Quantification of the area under the curve for relaxation indicated that the extent of L-NAME inhibition varied amongst the tissue engineered sheets. Col I sheets had a significantly lower %inhibition with L-NAME (*p<0.05; 33.4 ± 8.4%) corresponding to the lowest immunoblot expression of nNOS. The degree of L-NAME inhibition was higher in tissue engineered sheets containing Col I/IV and/or laminin and/or heparan sulfate (57.16% - 62.28%), corresponding to the higher immunoblot expression of nNOS. Attenuation of relaxation in the presence of L-NAME was 78 ± 2.9% in native tissue.

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3.5.2: Inhibition of the VIP-receptor:

The functionality of VIP-ergic neurons was assessed using a VIP receptor antagonist (VIP-

Ra). Pre-treatment with VIP-Ra inhibited maximal relaxation in all tissue engineered sheets

to varying extents ranging from 55.55 ± 3.92% - 65.92 ±5.38% (pink traces, Fig 28).

Inhibition of EFS-induced relaxation indicated the presence of differentiated VIP-ergic

neurons in tissue engineered sheets.

Figure 28: Inhibition of relaxation with VIP-Ra

The functionality of VIP-ergic neurons was studied by inhibiting EFS-induced relaxation with VIP-Ra, a VIP-receptor antagonist. Purple traces indicate EFS in the presence of VIP-Ra. In the presence of VIP-Ra, EFS-induced relaxation was attenuated in tissue engineered sheets (A-D) and in native tissue (E), indicating the presence of functional VIP-ergic neurons capable of mediating smooth muscle relaxation upon electrical field stimulation. Area under the curve of relaxation was quantified, to calculate the %inhibition of relaxation in the presence of VIP-Ra (F). Extent of VIP-Ra induced inhibition of relaxation varied from 56.55 ± 3.12% - 63.11 ± 3.2% in tissue engineered sheets, and averaged at 73.32 ± 3.23 % in native tissue.

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4. Discussion

Neural stem cell transplantation is a promising therapeutic approach to repopulate

neurons within enteric ganglia. A complete loss of neurons is reported in HSCR, and a

partial loss of selective neuronal subtypes is documented in achalasia and stenosis [7, 8,

34]. Several groups have injected enteric neuronal progenitor cells into experimental models

of aganglionosis, demonstrating the feasibility of transplantation [9, 12, 35]. However, there

is inadequate focus on differentiation of progenitor cells into mature neuronal subtypes, and

subsequent assessment of functionality. Here, we describe a novel method to bias

differentiation of enteric neuronal progenitor cells in vitro, prior to transplantation.

The ECM microenvironment, consisting of collagens, laminin and proteoglycans, not

only acts as a structural framework for cells, but also plays an active role in aiding

neurotrophic signaling [16, 36]. For these studies, we chose to evaluate four ECM

components (collagen I, collagen IV, laminin and heparan sulfate), three of which are known

to be present in adult myenteric ganglia [37]. Collagen IV has been documented to be

favorable for neurite outgrowth and neuronal differentiation [23, 24, 38]. Laminin has long

been known for its neurite promoting activity, in central, peripheral, and enteric neurons [26,

29, 39]. The role of the heparan sulfate proteoglycan in neuronal differentiation is also well

documented, both developmentally and in regenerative medicine applications [40-42].

Fibrillar Collagen I was used additionally in these studies for ease of gelation and

incorporation of other ECM components within a 3D hydrogel.

Apart from composition, substrate elasticity has been demonstrated to affect the

differentiation of adult neural stem cells, with neuronal differentiation reported between 100-

500Pa [43, 44]. ECM hydrogel compositions were adjusted in order to maintain their

viscoelastic modulus within the range suitable for neuronal differentiation (Table 1).

Structural architecture was verified using scanning electron microscopy, wherein the

addition of collagen IV demonstrated the presence of network structures, similar to self-

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assembled collagen IV in the mammalian basement membrane [45]. The addition of laminin

did not alter the ultrastructure, because it was expected to coat collagen fibers evenly.

Additionally, laminin was also not expected to alter the stiffness/viscoelasticity of the gels,

given the manner of its interaction with the collagens [46-48]. The glycosaminoglycan chains

of heparan sulfate are documented to cross link between laminin and collagen IV, thereby

pulling fibers into a more compact structure, and slightly increasing the viscoelasticity of

ECM gels [45, 49].

4.1 Smooth muscle cells within tissue engineered sheets drive the differentiation of enteric

neuronal progenitor cells

Tissue engineered sheets provided a good modality to assess variability of

differentiated neurons due to ECM composition as well as the functionality of differentiated

neurons. The proximity to smooth muscle promoted the differentiation of enteric neuronal

progenitor cells extensively. In vitro differentiation of neural stem cells in the presence of

gut-derived factors has been demonstrated previously by us and others [32, 33, 50].

Neurotrophic factors (NT-3, Neurturin, GDNF) and morphogens (BMP-2/4) capable of

driving enteric neuronal progenitor cell proliferation and differentiation have been

demonstrated to arise from the smooth muscle and mesenchyme of the developing and

adult gut [51-53]. Recently, the postnatal bowel was demonstrated to support the

differentiation of enteric neuronal progenitor cells, strengthening the fact that cues for

differentiation can be derived from the postnatal gut [54, 55]. Hence, it was expected that

smooth muscle cells within the tissue engineered sheets would drive the differentiation of

enteric neuronal progenitor cells. We evaluated all tissue engineered sheets to ensure that

the constituent smooth muscle cells demonstrated a contractile phenotype expressing

Smoothelin (Fig 22B). Smoothelin expression has been previously demonstrated to be

essential for contractility of smooth muscle [56]. In line with the equivalent expression of

smoothelin, myogenic electromechanical coupling integrity was also equivalent in the tissue

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engineered sheets (Fig 24A-D). Similar patterns of contractions were observed in tissue

engineered sheets in response to KCl, regardless of ECM composition.

4.2 ECM modulates differential neuronal subtypes while supporting overall smooth muscle-

driven neuronal differentiation

In the presence of the smooth muscle, enteric neuronal progenitor cells

differentiated, and expressed similar amounts of pan-neuronal marker βIII Tubulin (Figure

22A), suggesting that smooth muscle derived factors and substrate viscoelasticity were

suitable for overall neuronal differentiation. However, on closer examination of neural

subtypes, there was a differential expression of excitatory and inhibitory markers within

tissue engineered sheets with varying ECM compositions (Fig 22).

Sheets containing laminin had a balanced expression of both ChAT and nNOS.

Kinetics of Ach-induced contraction in laminin sheets was most similar to native tissue,

indicating the presence of an increased viable cholinergic neuronal component in composite

collagen/laminin sheets. Furthermore, attenuation of EFS-induced relaxation by L-NAME

(~62%) was also kinetically similar to native tissue (~78%), indicating the presence of a

nitrergic neuronal component.

Table 3: EFS-induced relaxation in tissue engineered sheets

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Collagen I, in the absence of any other matrix components, was the ECM of choice

when an enriched cholinergic neuronal population was required, with a significantly

diminished nitrergic neuronal population (Fig 22C-D). While these sheets demonstrated a

robust TTX-sensitive Ach-induced contraction commensurate with the heightened ChAT

protein expression, relaxation in response to an electrical field was diminished. Furthermore,

there was minimal attenuation of relaxation upon the inhibition of nNOS, correlating with the

low nNOS expression.

Composite Collagen I/IV sheets had an enhanced nNOS protein expression, with an

associated increase in EFS induced relaxation (Fig 26B). AUC of relaxation in composite

Collagen I/IV sheets was comparable to native intestinal tissue. However, both ChAT

expression and contraction was lower in composite collagen sheets.

4.3 ECM is a framework upon which smooth muscle derived factors regulate differentiation

of neural subtypes

Overall, the studies in this paper contribute towards understanding the impact of

environmental and molecular mechanisms associated with enteric neural subset activation.

Here we demonstrate a critical role of collagen I and collagen IV containing ECM

environments in promoting excitatory and inhibitory motor neurons, respectively. The ECM

microenvironment plays a role in modulating neurotrophic as well as morphogenetic

signaling. Morphogenetic signaling via the BMP family expressed in fetal gut is important for

the phenotypic diversity of enteric ganglia, including nitrergic and VIP-ergic neuron

differentiation [57] [58] [59, 60]. Collagen IV is documented to modulate BMP signaling, and

heparan sulfate modulates GDNF signaling in the gut [25, 27]. Immunoreactivity of

neurotrophic factor, NT-3, has been observed in ganglia and in the ECM molecules

surrounding them, suggesting a role for a Collagen IV-based ECM to modulate NT-3

signaling [61]. The constituent smooth muscle phenotype in tissue engineered sheets was

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contractile, expressing smoothelin, and generating contractions and relaxations approaching

~60% of those generated by native intestinal tissue. Differentiation cues arising from the

constituent smooth muscle cells drove enteric neuronal differentiation. Furthermore, it is

likely that the ECM could act as a framework for smooth muscle-derived factors, enhancing

or inhibiting their effects, resulting in the generation of differential neuronal phenotypes.

5. Conclusions

The studies in this paper used tissue engineering as a tool to evaluate the effect of

ECM composition on the differentiation of adult enteric neuronal progenitor cells into mature

neuronal subtypes. Furthermore, our studies indicated that different neuronal subtypes

emerged within microenvironments that varied in their ECM composition. A combination of

biochemical, immunohistochemical and physiological analyses revealed that several

functional differentiated neuronal subtypes were present in tissue engineered intestinal

sheets, capable of mediating smooth muscle contraction/relaxation. Neuronal populations

varied from being highly cholinergic (collagen I), highly nitrergic (collagen IV), or balanced

between the two (laminin and/or heparan sulfate). Additionally, peptidergic neurons with VIP

immunoreactivity were also detected. Our studies indicate that neuronal differentiation was

modulated by varying the composition of ECM microenvironments. Enriched populations of

differentiated neurons can be derived within transplantable tissue engineered sheets, using

ECM microenvironments. ECM microenvironments may also facilitate adequate trophic

support and phenotype maintenance of differentiated neurons.

6. Acknowledgements

This work was supported by NIH R01DK071614 and NIH R01DK042876. The authors would

like to thank Dr. Chris Pernell at the North Carolina State Food Science Rheology

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Laboratory for his expert technical assistance with rheometry. The authors declare no

conflicts of interest and no competing financial interests.

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[38] Tomaselli KJ, Damsky CH, Reichardt LF. Interactions of a neuronal cell line (PC12) with laminin, collagen IV, and fibronectin: identification of integrin-related glycoproteins involved in attachment and process outgrowth. J Cell Biol. 1987;105:2347-58. [39] Rogers SL, Letourneau PC, Palm SL, McCarthy J, Furcht LT. Neurite extension by peripheral and central nervous system neurons in response to substratum-bound fibronectin and laminin. Dev Biol. 1983;98:212-20. [40] Yamaguchi Y. Heparan sulfate proteoglycans in the nervous system: their diverse roles in neurogenesis, axon guidance, and synaptogenesis. Semin Cell Dev Biol. 2001;12:99-106. [41] Mammadov B, Mammadov R, Guler MO, Tekinay AB. Cooperative effect of heparan sulfate and laminin mimetic peptide nanofibers on the promotion of neurite outgrowth. Acta Biomaterialia. 2012;8:2077-86. [42] Sariola H, Saarma M. Novel functions and signalling pathways for GDNF. J Cell Sci. 2003;116:3855-62. [43] Banerjee A, Arha M, Choudhary S, Ashton RS, Bhatia SR, Schaffer DV, et al. The influence of hydrogel modulus on the proliferation and differentiation of encapsulated neural stem cells. Biomaterials. 2009;30:4695-9. [44] Saha K, Keung AJ, Irwin EF, Li Y, Little L, Schaffer DV, et al. Substrate modulus directs neural stem cell behavior. Biophys J. 2008;95:4426-38. [45] Kleinman HK, McGarvey ML, Hassell JR, Star VL, Cannon FB, Laurie GW, et al. Basement membrane complexes with biological activity. Biochemistry. 1986;25:312-8. [46] Deister C, Aljabari S, Schmidt CE. Effects of collagen 1, fibronectin, laminin and hyaluronic acid concentration in multi-component gels on neurite extension. J Biomater Sci Polym Ed. 2007;18:983-97. [47] Charonis AS, Tsilibary EC, Yurchenco PD, Furthmayr H. Binding of laminin to type IV collagen: a morphological study. J Cell Biol. 1985;100:1848-53. [48] Swindle-Reilly KE, Papke JB, Kutosky HP, Throm A, Hammer JA, Harkins AB, et al. The impact of laminin on 3D neurite extension in collagen gels. J Neural Eng. 2012;9:046007. [49] Yurchenco PD, Schittny JC. Molecular architecture of basement membranes. FASEB J. 1990;4:1577-90. [50] Kulkarni S, Zou B, Hanson J, Micci MA, Tiwari G, Becker L, et al. Gut-derived factors promote neurogenesis of CNS-neural stem cells and nudge their differentiation to an enteric-like neuronal phenotype. Am J Physiol Gastrointest Liver Physiol. 2011;301:G644-55. [51] Rossi J, Herzig KH, Voikar V, Hiltunen PH, Segerstrale M, Airaksinen MS. Alimentary tract innervation deficits and dysfunction in mice lacking GDNF family receptor alpha2. J Clin Invest. 2003;112:707-16. [52] Peters RJ, Osinski MA, Hongo JA, Bennett GL, Okragly AJ, Haak-Frendscho M, et al. GDNF is abundant in the adult rat gut. J Auton Nerv Syst. 1998;70:115-22. [53] Saffrey MJ, Burnstock G. Growth factors and the development and plasticity of the enteric nervous system. J Auton Nerv Syst. 1994;49:183-96. [54] Hotta R, Stamp LA, Foong JP, McConnell SN, Bergner AJ, Anderson RB, et al. Transplanted progenitors generate functional enteric neurons in the postnatal colon. J Clin Invest. 2013;123:1182-91. [55] Hagl CI, Rauch U, Klotz M, Heumuller S, Grundmann D, Ehnert S, et al. The microenvironment in the Hirschsprung's disease gut supports myenteric plexus growth. Int J Colorectal Dis. 2012;27:817-29. [56] Niessen P, Rensen S, van Deursen J, De Man J, De Laet A, Vanderwinden JM, et al. Smoothelin-a is essential for functional intestinal smooth muscle contractility in mice. Gastroenterology. 2005;129:1592-601.

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[57] Chalazonitis A, Pham TD, Li Z, Roman D, Guha U, Gomes W, et al. Bone morphogenetic protein regulation of enteric neuronal phenotypic diversity: relationship to timing of cell cycle exit. J Comp Neurol. 2008;509:474-92. [58] Anitha M, Shahnavaz N, Qayed E, Joseph I, Gossrau G, Mwangi S, et al. BMP2 promotes differentiation of nitrergic and catecholaminergic enteric neurons through a Smad1-dependent pathway. Am J Physiol Gastrointest Liver Physiol. 2010;298:G375-83. [59] Hendershot TJ, Liu H, Sarkar AA, Giovannucci DR, Clouthier DE, Abe M, et al. Expression of Hand2 is sufficient for neurogenesis and cell type-specific gene expression in the enteric nervous system. Dev Dyn. 2007;236:93-105. [60] Morikawa Y, Zehir A, Maska E, Deng C, Schneider MD, Mishina Y, et al. BMP signaling regulates sympathetic nervous system development through Smad4-dependent and -independent pathways. Development. 2009;136:3575-84. [61] Hoehner JC, Wester T, Pahlman S, Olsen L. Localization of neurotrophins and their high-affinity receptors during human enteric nervous system development. Gastroenterology. 1996;110:756-67.

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CHAPTER 7: NEO-INNERVATION OF EXPERIMENTALLY

DENERVATED COLON USING TISSUE ENGINEERED INNERVATED

LONGITUDINAL SMOOTH MUSCLE SHEETS

1. Introduction

The myenteric plexus of the enteric nervous system (ENS) is primarily responsible

for smooth muscle contraction/relaxation. Enteric neuropathies are central pathologies in

achalasia, and Hirschsprung’s disease; it is secondarily observed in diabetes, inflammation,

and Parkinson’s disease [1, 2]. In Hirschsprung’s disease, all enteric neural phenotypes are

lost. In diseases like sphincteric achalasia, pyloric stenosis, or diabetic gastroparesis,

specific damage/loss of nitrergic neurons occur [3, 4, 5]. In disorders where specific

neurotransmitter deficiencies occur, it is beneficial to identify methods of generating

enriched populations of specific neuronal subtypes for transplantation. In our studies, we

have utilized ECM-based hydrogel microenvironments to modulate the phenotype of enteric

neuronal progenitor cells in vitro.

Enteric neuronal progenitor cells have formed the basis of neural stem cell

transplantation, and have been transplanted into embryonic hindguts, where they migrate

and acquire several enteric neuronal phenotypes [6-11]. However, the embryonic

microenvironment is not similar to the adult intestinal microenvironment. Moreover, injection

of progenitor cells in vivo have poor engraftment and also migrate very limited distances [12,

13]. With the approach of injection of progenitor cells, there is no control over the eventual

phenotype of the differentiated cells. In cases like achalasia, where specific repopulation of

nitrergic neurons are required, injection of progenitor cells may not be beneficial in

supplementing nitrergic neuronal functionality. In such cases, it is beneficial to identify

methods of biasing neuronal differentiation into specific neuronal subtypes prior to

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transplantation. The objective of this study was to determine if functional nitrergic neo-

innervation could be facilitated in denervated colonic smooth muscle using tissue

engineered sheets with enriched nitrergic neurons.

We have demonstrated that we can reproducibly isolate neural crest derived enteric

neuronal progenitor cells from biopsies of rabbit jejunums. We generated enriched

populations of differentiated nitrergic neurons within tissue engineered sheets manufactured

with collagen IV hydrogels.

Serosal application of benzalkonium chloride (BAC) is an effective method to

chemically ablate the myenteric plexus in the colon. The earliest use of BAC for myenteric

denervation was described by Sato et al., where serosal application of BAC resulted in

specific myenteric denervation with no changes in the musculature [14-16]. It was postulated

that the cationic surfactant selectively damaged more depolarized neuronal cells faster than

affecting the less depolarized smooth muscle cells [17].

In these studies, we utilized tissue engineered sheets manufactured from composite

collagen I/collagen IV to generate an enriched population of nitrergic neurons. These tissue

engineered innervated sheets were used as ‘donor’ tissues, to supply neo-innervation to

experimentally denervated colonic explants. Following co-culture, neo-innervation was

investigated by studying the overall increase in neuronal marker expression via immunoblot,

and the specific presence of nitrergic neo-innervation using immunohistochemistry.

Additionally, the functionality of nitrergic neo-innervation was evaluated as its ability to

induce smooth muscle relaxation.

These studies will determine if functional nitrergic neo-innervation can be achieved

using pre-differentiated nitrergic neurons. Additionally, they will provide a preliminary

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therapeutic potential for tissue engineered innervated sheets in their use for remedying

dysganglionosis.

2. Materials and Methods

All tissue culture reagents were purchased from Life Technologies (Carlsbad, CA) unless

specified otherwise. Benzalkonium chloride was purchased from Sigma Aldrich (St. Louis,

MO). Collagen I and Collagen IV were purchased from BD Biosciences (San Jose, CA).

2.1 Isolation and denervation of rabbit colon

Sigmoid colons were dissected from rabbits and cleaned of fecal content. Adherent fat was

picked off using forceps. In order to denervate the colon, cleaned colon was wrapped in

tissue soaked in 0.1% benzalkonium chloride (BAC) solution in saline. The tissue was

rewetted with BAC solution every 15 minutes, for up to 1 hour. BAC treated tissues were

washed extensively in Hank’s Balanced Salt Solution (HBSS) to remove any traces of BAC.

Serosa, longitudinal smooth muscle and inner mucosal layer were removed using a sharp

scalpel. The remaining BAC treated colonic circular muscle was placed in immersion culture.

The immersion medium was 1X Dulbecco’s modified Eagle’s medium supplemented with

10% fetal bovine serum, 1.5X antibiotics/antimycotics.

Immersion medium was replaced every day. As a control, colon tissues not treated with BAC

were also placed in explant culture. These colons were processed in a similar manner,

except the soaking solution was replaced with saline instead of BAC. Denervation was

confirmed preliminarily using immunofluorescence on paraffin-processed cross sections of

tissues following BAC-treatment.

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2.2 Manufacturing nitrergic neuron-enriched tissue engineered intestinal longitudinal smooth

muscle sheets

Tissue engineered longitudinal intestinal smooth muscle sheets were manufactured

following protocols described previously in Chapters 5 and 6. In order to promote nitrergic

neuronal differentiation, a composite Collagen I/Collagen IV matrix was used in the

bioengineering process.

Rabbit intestinal longitudinal smooth muscle cells were isolated following protocols

described previously. Enteric neuronal progenitor cells were isolated from small intestines of

mice expressing GFP driven by the β Actin promoter. Isolated enteric neuronal progenitor

cells were immunofluorescent when probed with the GFP antibody (Figure 29).

Figure 29: Enteric neurospheres from GFP mice

Enteric neuronal progenitor cells were derived from mice expressing GFP driven by a β actin promoter. These cells aggregate in culture, similar to rabbit enteric neuronal progenitor cells, to form floating colonies of spherical structures (enteric neurospheres). Cells within the neurospheres are fluorescent, and express GFP. Scale bar 100µm.

Substrates with wavy microtopographies were used to achieve uniaxial alignment of rabbit

longitudinal smooth muscle cells. Following smooth muscle alignment, GFP-expressing

enteric neuronal progenitor cells were suspended in an ECM solution consisting of collagen

I/collagen IV. The ECM gel with GFP+ mouse enteric neuronal progenitor cells was overlaid

on top of the aligned smooth muscle monolayer. Neuronal differentiation medium was added

following gelation, and replaced every alternate day. This process resulted in the formation

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of ~1cm x 0.5cm innervated tissue engineered sheets that were anchored between two silk

sutures. Our previous data demonstrated that using this ECM composition, we were able to

derive an enriched population of differentiated nitrergic neurons within the tissue engineered

sheets.

2.3 Neo-innervation of explant colonic circular muscle

1cm x 0.5cm strips of explanted rabbit colonic circular muscle were pinned down to sterile

polydimethylsiloxane coated tissue culture dishes. To initiate neo-innervation, tissue

engineered intestinal longitudinal smooth muscle sheets were pinned onto the serosal side

of the explant circular muscle. Care was taken to match the dimensions of the tissue

engineered sheets and the explant colon tissues, in order to maximize the surface area of

contact and thereby maximize neo-innervation. The explant and tissue engineering sheet

co-culture was immersed in neuronal differentiation medium. Medium was exchanged every

alternate day, and co-cultures were maintained for 14 days. A schematic of this

experimental paradigm is shown in Figure 30.

2.4 Assessment of neo-innervation using immunoblotting

Explant tissues were removed from co-culture after 14 days and minced in RIPA buffer.

Proteins were extracted following sonication of lysate for 60s on ice. Extracted proteins were

separated electrophoretically in a 12% polyacrylamide gel and transferred to a

polyvinylidene difluoride membrane. Membranes were blocked with 5% milk and blotted with

primary antibodies directed against βIII Tubulin and β Actin as a loading control. HRP-

conjugated secondary antibodies were used to identify the expression of protein using

enhanced chemiluminescence. Control tissues that were untreated or BAC-treated tissues

that received no co-culture were also used for immunoblot analysis.

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Figure 30: Schematic of neo-innervation using tissue engineered innervated sheets

Tissue engineered longitudinal sheets were made using composite collagen I/collagen IV hydrogels. 1cm x 0.5cm strips of untreated or BAC treated explants were pinned serosal side up on to polydimethyl siloxane coated tissue culture dishes. In order to facilitate neo-innervation, tissue engineered sheets of the same dimensions were pinned abutting the serosal surface of BAC-treated explants. Explants were maintained in culture for 14 days, and harvested for immunoblots, immunofluorescence and physiological analyses.

2.5 Assessment of nitrergic neo-innervation using immunohistochemistry

At day 14, the explant tissues were removed from co-culture, fixed in 4% neutral buffered

formaldehyde, dehydrated, and embedded in paraffin. 8µm longitudinal sections were made

using a paraffin microtome. Longitudinal sections were treated with xylene to dissolve

paraffin, and rehydrated through graded alcohol. Hydrated sections were blocked and

permeabilized using 0.15% Triton-X and 10% horse serum. Sections were then incubated

for 60 minutes with primary antibodies directed against nNOS (Santa Cruz, rabbit polyclonal)

and GFP (Santa Cruz, mouse monoclonal). Appropriate fluorophore conjugated secondary

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antibodies were incubated for 60 minutes following a 15 minute wash with PBS to remove

unbound primary antibody. Nikon TiE inverted fluorescence microscope was used to

visualize fluorescence in the sections.

2.6 Assessment of physiological function of neo-innervation

At day 14, explant circular tissues were removed from co-culture and hooked on to a force

transducer, as described before in Chapter 6. Tissues were immersed in 4ml of basal

DMEM warmed to 37°C. Electrical field stimulation (EFS; 5Hz, 0.5ms) was applied using

parallel platinum plate electrodes set equidistant from the tissue, 8mm apart. EFS was

carried out following a 15 minute pre-incubation with L-NAME (300µM, nNOS inhibitor) to

identify the presence and functionality of nitrergic neo-innervation.

2.7 Data analysis and statistical methods

Densitometry on immunoblots were carried out using BioRad Quantity One, and normalized

to β Actin. One-way ANOVAs were used to identify a significant difference between means

of control untreated, BAC-treated and neo-innervated tissues. Physiology data was acquired

using Powerlab at a sampling rate of 1kHz, exported to GraphPad Prism 5.1 for Windows.

Second order Savitsky-Golay smoothing was applied to the data.

Area under the curve was estimated for the relaxation curves from the initiation of relaxation

to the retrieval of baseline. Inhibition of relaxation by L-NAME was quantified first by

calculating area under the relaxation curve, and expressing it as a % of the AUC without L-

NAME pre-treatment. %inhibition with L-NAME was compared by one-way ANOVAs

between means of control untreated, BAC-treated and neo-innervated tissues.

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3. Results

3.1 Neo-innervation of benzalkonium chloride (BAC) induced denervated explant circular

colon tissues

Serosal application of BAC for 60 minutes effectively denervated ~60% of myenteric ganglia

in explant colon tissues. Upon staining transverse sections of the serosal side of explant

tissues with βIII Tubulin, myenteric ganglia in BAC treated tissues displayed gaps that were

previously occupied by enteric neurons (Figure 31). Tubulin positive cells were still present,

because BAC-induced denervation was only ~60%. βIII Tubulin staining in untreated explant

tissues demonstrated full intact myenteric ganglia (Figure 31). The gaps in βIII Tubulin

staining indicated that BAC treatment denervated the explant tissues. Upon co-culture, the

myenteric ganglia appeared fuller. However, gaps were still present, indicating partial neo-

innervation, but not complete restoration of all the neurons solubilized with BAC treatment.

Figure 31: βIII Tubulin expression in sections of BAC treated, untreated and neo-innervated colons

6µm transverse sections were made from the serosal side of formalin fixed, paraffin embedded colonic circular smooth muscle tissues. Myenteric ganglia were visualized by staining with fluorophore conjugated βIII Tubulin. In BAC treated colons, myenteric ganglia had missing neuronal elements, with gaps observed in the ganglia (white arrows). In comparison, myenteric ganglia were full and undisturbed in control untreated colons. Neo-innervated colons had fuller myenteric ganglia compared to BAC-treated colons, but still contained gaps. This indicated a denervation of myenteric ganglia, and neo-innervation following co-culture.

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3.2 βIII Tubulin immunoblot expression in explant tissues

Western blots for βIII Tubulin were used to quantitate denervation and neo-innervation

(Figure 32). Control untreated explants, BAC treated denervated explants, and neo-

innervated explants were probed for the expression of neuronal marker βIII Tubulin. A

representative blot is provided, with a band for βIII Tubulin at ~50KDa, and a loading control

(β Actin, ~42KDa). Densitometry of the ~50KDa βIII Tubulin band indicated that with serosal

application of BAC, there was a reduction in neuronal marker expression by ~56%. The

reduction in βIII Tubulin expression indicated denervation of the explant tissues. The

duration of application and the concentration of BAC used did not denervate the explant

colons completely.

When explant tissues were co-cultured with ‘donor’ tissue engineering sheets for 14 days, a

20% increase in neuronal βIII Tubulin expression was observed. This significant increase in

neuronal marker expression indicated neo-innervation of BAC-treated tissues.

3.3 Presence of neo-innervated nitrergic neurons

Immunofluorescence for nNOS was used to identify the presence of nitrergic neurons. In

order to identify the source of the nitrergic neurons, longitudinal sections of explant tissues

were co-stained with both GFP as well as nNOS. The presence of GFP positive neurons

(Figure 33C) indicated that these neurons originated from tissue-engineered sheets.

GFP+ neurons were probed to confirm if they were nitrergic neurons. Some GFP+ neurons

expressed nNOS, which indicated that some neo-innervation originating from the tissue-

engineered sheets was nitrergic in nature (Figure 33A, yellow merged co-stains). Some

GFP+ neurons did not express nNOS (Figure 33C, green arrows), indicating that a small

percentage of neo-innervation was not nitrergic. These could be differentiated neurons of

another phenotype, possibly cholinergic or VIP-ergic.

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Figure 32: Immunoblot expression of βIII Tubulin in explant tissues

Proteins were extracted from untreated, BAC-treated or neo-innervated explants and resolved electrophoretically. Western blots demonstrated a clear band at ~50KDa for βIII Tubulin, and a band at ~42KDa for β actin. Densitometry of immunoblots demonstrated that with BAC treatment, βIII Tubulin expression reduced by 56% indicating the loss of myenteric neurons. In contrast, explants that were co-cultured with tissue engineered sheets had a significant ~20% increase (***p<0.001, n=4) in βIII Tubulin expression, indicating neo-innervation.

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Endogenous nNOS neurons in the explant tissue was also observed (Figure 33D-F) that

were not positive for GFP, indicating that BAC-induced denervation did not totally eliminate

endogenous neurons. Any endogenous reinnervation that occurred in vitro following BAC-

treatment may also be visualized.

Figure 33: Immunofluorescence in explant tissues

6µm longitudinal sections were made from neo-innervated explant tissues and co-stained with nNOS and GFP. (A-C) indicate GFP positive neo-innervation also expressing nNOS (yellow co-stain and arrows), indicating that the source of the nNOS neurons in this cluster was from the donor tissue engineered sheets. Also observed are GFP positive neurons (green arrows; C) that are not nNOS positive, indicating differentiation in to another neuronal phenotype. (D-F): Endogenous nNOS neurons that survived the BAC-denervation are not positive for GFP.

3.4 Neuronal functionality in explant tissues

1cm strips of explant tissues were hooked up to a force transducer to measure real-time

force generation. Electrical field stimulation (5Hz, 0.5ms) was used to evoke neuronally

mediated relaxation. Relaxation was abolished upon pre-treatment with neuronal blocker,

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Tetrodotoxin (TTX; red traces in Figure 34). Untreated control explant tissues demonstrated

a robust relaxation in response to EFS (Figure 34A). The magnitude of relaxation (-343.6 ±

17.3µN) as well as the area under the relaxation curve (109118 ± 6884AU) were maximal

compared to BAC-treated or BAC-treated and neo-innervated tissues. Upon BAC-induced

denervation, EFS-induced relaxation was significantly attenuated, averaging at -171.2 ±

8.96µN with an area under the curve of 32349 ± 8371AU (Figure 34B).

The reduced relaxation (both magnitude and duration accounted for in the area under the

relaxation curve) could be attributed to the loss of myenteric neurons upon BAC treatment.

When BAC-treated tissues were co-cultured with tissue engineered innervated sheets, an

increase in the magnitude of EFS-induced relaxation was observed in the recipient explant

tissues.

The improvement was significant, and relaxation averaged at -229.2 ± 20.46µN with an AUC

of 57650 ± 1679AU (Figure 34C). Comparison of relaxation is provided in Figure 34D. Our

results indicated that while BAC-induced denervation was not complete, sufficient

denervation and subsequent neo-innervation had occurred to result in a physiologically

significant difference in neuronal mediated smooth muscle relaxation.

Enhanced magnitudes of initial relaxation as well as the sustained relaxation, and a small

increase in the duration of relaxation were observed upon neo-innervation when compared

to BAC treated tissues. However, even with neo-innervation the duration of relaxation was

still ~100s shorter than control untreated tissues. This indicated that significant neuronal

subtypes required for EFS-induced relaxation were reintroduced with neo-innervation to

make a functional difference in physiology, but repopulation with all other neural types may

be incomplete.

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Figure 34: Electrical Field Stimulation (EFS) induced relaxation in explant colon tissues

1cm strips of untreated (A), BAC-treated (B) or neo-innervated (C) colon tissues were stimulated to relax using EFS (shaded grey area). Relaxation was completely inhibited in the presence of neuronal blocker, tetrodotoxin (red traces), indicating that relaxation was neuronally evoked. (A) Robust EFS-induced relaxation was observed in control untreated tissues, averaging at -343.6 + 17.3µN. (B) BAC-treated tissues had a significantly lower magnitude of EFS-induced relaxation compared to control untreated tissues (-171.2 + 8.96µN; ***p<0.001, n=4). (C) Neo-innervated explants had improved EFS-induced relaxation (-229.2 + 20.46µN; *p<0.05, n=4). (D) Area under the relaxation curve was quantified, and indicated that BAC treatment significantly lowered EFS-induced relaxation. Relaxation was increased upon neo-innervation.

Nitrergic neo-innervation: In order to identify a functional population of nitrergic neurons,

EFS-induced relaxation was repeated in the presence of L-NAME, an inhibitor of nNOS. In

the presence of L-NAME, EFS-induced relaxation was attenuated (green traces; Figure 35).

The extent of inhibition varied in the different tissues. Untreated explant tissues had the

maximum inhibition with L-NAME (77.74 ± 1.46%) indicating the presence of a robust and

functional nitrergic neuronal component that mediates EFS-induced relaxation. Upon BAC

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treatment, inhibition with L-NAME significantly decreased to 45.95 ± 3.084%, indicating the

partial loss of nitrergic neurons. When BAC-treated tissues were co-cultured with tissue

engineered sheets, a significantly elevated level of inhibition with L-NAME is observed

(54.01 ± 0.87%), indicating functional nitrergic neo-innervation.

Figure 35: Inhibition of nNOS in explant colon tissues

EFS-induced relaxation was inhibited by L-NAME pre-treatment (green traces) in 1cm strips of untreated (A), BAC-treated (B) or neo-innervated (C) colon tissues. (A) Inhibition of nNOS significantly diminished the magnitude of EFS-induced relaxation (77.74 + 1.46%, n=4) in untreated explants, indicating the presence of a functional nitrergic neuronal component contributing to EFS-induced relaxation. (B) Inhibition with L-NAME was significantly lower in BAC-treated tissues (45.95 + 3.08%, *p<0.05, n=4) indicating the loss of functional nitrergic neurons. (C) Neo-innervated explants had a significantly higher inhibition with L-NAME pre-treatment (54.01 + 0.86%, **p<0.01, n=4)

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4. Discussion

Enteric neuropathies are secondary to several disorders like diabetes, inflammation,

and obesity. Intestinal aganglionosis, i.e. the lack of enteric ganglia is the central pathology

of Hirschsprung’s disease. In a fully functioning enteric nervous system (ENS), neurons of

the submucosal plexus predominantly contribute to sensation and reflex based pathways,

while neurons of the myenteric plexus are primarily responsible for the motility patterns

observed in the gastrointestinal tract. Repopulation of myenteric neurons through a cell-

based therapeutic approach may positively modulate smooth muscle motility.

Several studies have demonstrated the feasibility of transplantation of ENS-derived

neuronal progenitor cells into the embryonic gut. More recently, it has also been established

that progenitor cells can be transplanted into the postnatal mouse gut where the cells

differentiate into neurons. These neurons demonstrate the neurochemical coding and

electrophysiological characteristics akin to enteric neurons [18]. However, transplantation of

progenitor cells into the gut is associated with an uncertainty in achieving the appropriate

neurochemical coding required of the pathology (for example: specific loss of inhibitory

motor neurons in achalasia).

Pre-differentiation of neurons in vitro, prior to transplantation, is of novel therapeutic

value because neurons are terminally differentiated. Our previous studies have

demonstrated that differentiated neurons derived using manipulation of extracellular

microenvironments are physiologically functional, and are capable of mediating smooth

muscle contraction and relaxation. The studies outlined in this aim evaluated the therapeutic

potential of pre-differentiated neurons using explant cultures of experimentally denervated

colon.

126

Our studies utilized serosal BAC application as a method to experimentally

denervate explant cultures of colon. BAC application ablates neurons selectively without

affecting the smooth muscle, resulting in a ~56% denervation of the explanted circular

smooth muscle of the colon. Specific myenteric denervation is reported upon serosal

application of BAC in several in vivo studies, with no alterations in the smooth musculature

of gut. The extent of denervation is reported to be closer to 80-90% in vivo due to immune

mechanisms that likely contribute to denervation [14-17]. In our in vitro studies, we observe

a 56% denervation that is due to the action of the cationic surfactant alone selectively on the

neural cells, with no immune contribution in the in vitro setting.

The experimental design included maintaining three different segments of tissue in

explant culture: i) untreated control tissues; ii) BAC treated denervated tissues; iii) BAC-

treated tissues that received donor tissue engineered innervated sheets. βIII Tubulin

expression was used as a marker to evaluate changes in innervation with BAC treatment or

co-culture. This paradigm accounts for any endogenous neuronal regeneration that may

have occurred in organotypic culture. Furthermore, we used enteric neuronal progenitor

cells derived from mice that constitutively express β Actin driven GFP. We were able to

discern neo-innervation from endogenous innervation using immunofluorescence.

Our results demonstrate that tissue engineered innervated longitudinal sheets can be

used to functionally innervate explant colon tissues. Neo-innervation was observed as an

increase in expression of neuronal marker βIII Tubulin by 20%. Neo-innervation was also

identified using GFP positive neurons within sections of explanted tissue. A majority of the

GFP positive neurons also expressed nNOS, indicating nitrergic neo-innervation of colon

explants. Neurons positive for GFP, but not nNOS represented the minority that

differentiated into phenotypes other than nitrergic.

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Although BAC-induced denervation was only 56%, there was significant effect on

neuronally mediated relaxation. Neo-innervation, though quantified to be an additional 20%,

was maximized by modulating extracellular matrix microenvironments to deliver nitrergic

neurons. The amount of nitrergic neo-innervation delivered resulted in a functional and

significant improvement in nitrergic neuron-mediated smooth muscle relaxation.

The implication of these studies is that pre-differentiated nitrergic neurons can graft

into explant colonic tissues, and contribute to smooth muscle relaxation. This additionally

suggests that the nitrergic neo-innervation established connectivity with existing neural

circuits. We have also demonstrated that experimentally denervated tissues support neo-

innervation. . Future studies will include confirming cholinergic contractile neo-innervation as

well as translating these findings to an in vivo model of aganglionosis (BAC-induced or

genetic).

5. References

[1] De Giorgio R, Camilleri M. Human enteric neuropathies: morphology and molecular pathology. Neurogastroenterol Motil. 2004;16:515-31. [2] Bitar K, Greenwood-Van Meerveld B, Saad R, Wiley JW. Aging and gastrointestinal neuromuscular function: insights from within and outside the gut. Neurogastroenterol Motil. 2011;23:490-501. [3] De Giorgio R, Stanghellini V, Barbara G, Corinaldesi R, De Ponti F, Tonini M, et al. Primary enteric neuropathies underlying gastrointestinal motor dysfunction. Scand J Gastroenterol. 2000;35:114-22. [4] Kessmann J. Hirschsprung's disease: diagnosis and management. Am Fam Physician. 2006;74:1319-22. [5] Kraichely RE, Farrugia G. Achalasia: physiology and etiopathogenesis. Dis Esophagus. 2006;19:213-23. [6] Bondurand N, Natarajan D, Thapar N, Atkins C, Pachnis V. Neuron and glia generating progenitors of the mammalian enteric nervous system isolated from foetal and postnatal gut cultures. Development. 2003;130:6387-400. [7] Kruger GM, Mosher JT, Bixby S, Joseph N, Iwashita T, Morrison SJ. Neural crest stem cells persist in the adult gut but undergo changes in self-renewal, neuronal subtype potential, and factor responsiveness. Neuron. 2002;35:657-69. [8] Raghavan S, Gilmont RR, Bitar KN. Neuroglial differentiation of adult enteric neuronal progenitor cells as a function of extracellular matrix composition. Biomaterials. 2013;34:6649-58.

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[9] Lindley RM, Hawcutt DB, Connell MG, Almond SL, Vannucchi MG, Faussone-Pellegrini MS, et al. Human and mouse enteric nervous system neurosphere transplants regulate the function of aganglionic embryonic distal colon. Gastroenterology. 2008;135:205-16 e6. [10] Metzger M, Caldwell C, Barlow AJ, Burns AJ, Thapar N. Enteric nervous system stem cells derived from human gut mucosa for the treatment of aganglionic gut disorders. Gastroenterology. 2009;136:2214-25 e1-3. [11] Rauch U, Hansgen A, Hagl C, Holland-Cunz S, Schafer KH. Isolation and cultivation of neuronal precursor cells from the developing human enteric nervous system as a tool for cell therapy in dysganglionosis. Int J Colorectal Dis. 2006;21:554-9. [12] Kulkarni S, Becker L, Pasricha PJ. Stem cell transplantation in neurodegenerative disorders of the gastrointestinal tract: future or fiction? Gut. 2012;61:613-21. [13] Schäfer Kh, Micci Ma, Pasricha PJ. Neural stem cell transplantation in the enteric nervous system: roadmaps and roadblocks. Neurogastroenterology & Motility. 2009;21:103-12. [14] Sato A, Yamamoto M, Imamura K, Kashiki Y, Kunieda T, Sakata K. Pathophysiology of aganglionic colon and anorectum: an experimental study on aganglionosis produced by a new method in the rat. J Pediatr Surg. 1978;13:399-435. [15] Hanani M, Ledder O, Yutkin V, Abu-Dalu R, Huang TY, Hartig W, et al. Regeneration of myenteric plexus in the mouse colon after experimental denervation with benzalkonium chloride. J Comp Neurol. 2003;462:315-27. [16] Yoneda A, Shima H, Nemeth L, Oue T, Puri P. Selective chemical ablation of the enteric plexus in mice. Pediatr Surg Int. 2002;18:234-7. [17] Sakata K, Kunieda T, Furuta T, Sato A. Selective destruction of intestinal nervous elements by local application of benzalkonium solution in the rat. Experientia. 1979;35:1611-3. [18] Hotta R, Stamp LA, Foong JP, McConnell SN, Bergner AJ, Anderson RB, et al. Transplanted progenitors generate functional enteric neurons in the postnatal colon. J Clin Invest. 2013;123:1182-91.

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CHAPTER 8: FUTURE DIRECTIONS

1. Identifying the mechanisms involved in neural subtype specification, including the

role of extracellular matrix based microenvironments

The mechanisms underlying neural subset specification remain unidentified. The data

generated in this thesis demonstrates that enteric neuronal progenitor cells differentiated

into enriched populations of motor neuronal subtypes when placed in microenvironments

with varying extracellular matrix compositions. Several factors may be involved in directing

the diversity of neuronal phenotypes, all thought to arise from a crosstalk between smooth

muscle cells and differentiating neuronal progenitor cells. Indeed, central nervous system

derived progenitor cells differentiate into appropriate enteric neuronal phenotypes in the

presence of gut-derived factors.

Several soluble factor candidates have been proposed to modulate the development of

neuronal subtypes, including Bone morphogenetic proteins (BMP 2/4, Glial-cell derived

neurotrophic factor (GDNF), Neuregulins, Endothelins, and neurotrophic factors NT-3, NT-

4/5. Extracellular matrix components like Collagen IV and heparan sulfate bind these

factors, modify their bioavailability and thereby modulate their signaling. Future studies will

involve identifying the factors involved in neural subset specification in the different ECM

microenvironments. These studies will involve ELISA-based quantitation of several different

soluble factors. Once these factors have been isolated and identified, a more

comprehensive switch to using defined neural differentiation medium will be carried out in

order to induce specific neural phenotypes.

Physically, cellular components signal through integrins with the extracellular matrix

microenvironment. Mutations in β1 integrins have demonstrated distal aganglionosis in

rodent models, indicating the central role of β1 integrins in the formation of the enteric

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nervous system. Our future studies will identify the roles specific αβ integrin combinations

will have on neuronal differentiation and neural subtype specification.

2. Translating neo-innervation in vivo

The data generated in Aim 3 of this thesis demonstrated that benzalkonium chloride

treatment denervated the myenteric plexus of rabbit colons. We re-supplied nitrergic neo-

innervation in denervated explants using innervated tissue engineered sheets. Furthermore,

the nitrergic neo-innervation was functional in response to electrical field stimulation,

mediating smooth muscle relaxation. In future studies, we propose to expand this to an

animal model in vivo. These studies will include rendering a portion of the distal colorectum

aganglionic during a survival surgery, and transplanting tissue engineered innervated

intestinal segments. Our preliminary data indicate a reduction in neuronal marker βIII

Tubulin following 14 days after a 30 minute exposure to benzalkonium chloride in vivo

(Figure 36).

Figure 36: Reduced Neuronal βIII Tubulin in BAC treated segment

Immunoblotting of whole tissue lysate from BAC-treated and untreated samples showed reduction in neuronal βIII Tubulin in BAC treated segment suggesting that there is loss of neurons upon treatment with BAC.

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Figure 37: Electrical Field Stimulation in BAC-treated segments in vivo

BAC treated colonic segment demonstrated very minimal relaxation (~165µN) compared to untreated colonic segment (~420 µN). This data demonstrates the loss of a functional neuronal component upon BAC treatment.

Physiological examination of harvested BAC-treated denervated segment and untreated

control rectal segments revealed that there was a loss of functional neuronal physiology,

indicated by an attenuated magnitude of electrical field stimulation induced relaxation

(Figure 37).

Innervated tissue engineered intestinal constructs were implanted around the denervated

rectum for 14 days. Following implantation and harvest, neuronal physiology was evaluated

using electrical field stimulation. The segment that received the implant demonstrated

significantly elevated magnitudes of relaxation, indicating a functional neo-innervation

(Figure 38). An L-NAME sensitive nitrergic neuronal component was also identified in the

neo-innervated segments in vivo.

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Figure 38: Restoration of NO-mediated relaxation in BAC-treated neo-innervated colon segments in vivo

Nitric oxide synthase inhibitor, L-NAME, was used to pretreat tissues before electrical field stimulation. nNOS inhibition in BAC treated tissues only resulted in a 20% attenuation of relaxation, indicating the absence of nitrergic neurons in the denervated segment. BAC treated segment receiving implant demonstrated a 70% inhibition of EFS-induced relaxation, indicating the reinstatement of functional nitrergic neurons mediating smooth muscle relaxation.

We propose to expand the findings from this thesis by two methods: i) examining the

mechanistic procedures through which the extracellular matrix microenvironment modulates

neural subtype specification; and ii) delivering enriched populations of differentiated neurons

into denervated segments of the intestine in order to study phenotype maintenance as well

as functional neo-innervation. This will include detailed studies involving the identification of

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the neurotrophic factors produced due to a reciprocal cross talk between smooth muscle

cells and enteric neuronal progenitor cells, and the involvement of the ECM in modulating

neurotrophic factor signaling. Additionally, studies will also identify the role of specific

integrin complexes that may modulate neuronal subtype specification. Finally, more

elaborate studies will be performed to establish a reliable in vivo model of denervation,

where nitrergic or cholinergic neo-innervation can be supplied to restore intestinal motility

and function.

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CHAPTER 9: SUMMARY AND CONCLUSIONS

The studies outlined and described within this thesis address the hypothesis that

extracellular matrix based microenvironments can be used to modulate the differentiation of

adult mammalian enteric neuronal progenitor cells. This hypothesis was first evaluated in

two dimensional culture substrata, using various combinations of collagen I, collagen IV,

laminin and heparan sulfate. Smooth muscle-derived soluble factors were used to drive the

differentiation of the enteric neuronal progenitor cells. In two dimensional cultures, enteric

neuronal progenitor cells differentiated into enriched populations of enteric glia, or enteric

neurons. Substrata that had no ECM coating generated a preponderance of differentiated

glia. Collagen I substrata also generated differentiated glia, in addition to differentiated

neurons that were minimally branched. Substrata that included collagen IV and laminin

differentiated into a preponderance of branched neurons, with minimal enteric glia. The

presence of heparan sulfate in culture substrata enhanced preliminary neuronal networking.

Substrata that were extremely suitable for neuronal differentiation included collagen IV,

laminin and heparan sulfate. These substrata supported glial differentiation minimally.

We translated these findings into three-dimensional ECM microenvironments using a

tissue engineered longitudinal sheet model. Here, we utilized substrate microtopography to

drive smooth muscle alignment. Enteric neuronal progenitor cells were encapsulated within

ECM hydrogels, whose composition and viscoelasticity mimicked neural ECM in vivo. The

overlay of the enteric neuronal progenitor cells over the aligned smooth muscle monolayer

generated innervated intestinal tissue engineered sheets.

By varying the composition of the ECM, we generated tissue engineered sheets with

a highly cholinergic motor neuronal population, enriched nitrergic motor neuronal population,

and a balanced expression of cholinergic and nitrergic neurons. The differentiated neurons

were capable of mediating smooth muscle contraction and relaxation upon appropriate

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physiological stimuli. Furthermore, additional pharmacological assays confirmed the

presence and physiological functionality of the differentiated neurons.

The final set of experiments evaluated the therapeutic potential of tissue engineered

innervated intestinal sheets in providing functional neo-innervation. Our experiments

demonstrated a preliminary proof of concept that innervated intestinal sheets tissue

engineered with enriched nitrergic neurons will functionally repopulate nitrergic neurons in

denervated smooth muscle. The neo-innervated nitrergic neurons additionally mediated

smooth muscle relaxation in response to electrical field stimulation. We confirmed neo-

innervation additionally in an in vivo rodent model using transplanted tissue engineered

innervated constructs.

Taken together, we have modulated the differentiation of adult mammalian enteric

neuronal progenitor cells to differentiate into a preponderance of neurons or glia using two

dimensional culture substrata. Additionally, we have tissue engineered innervated intestinal

tissue engineered sheets wherein we can have enriched cholinergic excitatory neurons,

enriched nitrergic inhibitory neurons, or a balanced population of excitatory and inhibitory

neurons. We have also further demonstrated that these tissue engineered innervated sheets

can be utilized to functionally neo-innervate denervate intestinal tissues, both in vitro and in

vivo.

Our results strongly demonstrate that extracellular matrix microenvironments are a

useful therapeutic tool to modulate differentiation of enteric neuronal progenitor cells into

defined phenotypes. These differentiated enteric neuronal phenotypes can be transplanted

in vivo in several dysganglionic disorders of the gastrointestinal tract, where they can

functionally neo-innervate denervated smooth muscle structures and reinstate

gastrointestinal motility and function.

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SCHOLASTIC VITA

EDUCATION

Virginia Tech-Wake Forest University School of Biomedical Engineering &

Sciences

PhD in Biomedical Engineering- 2010- 2014

Concentration: Cell and Tissue Engineering

University of Michigan, Ann Arbor

Masters in Biomedical Engineering – 2006-2008

Concentration: Biotechnology

Anna University, Chennai , India

Bachelors in Engineering- 2002-2006

Concentration: Instrumentation and Control Engineering

PEER-REVIEWED PUBLICATIONS

Raghavan S, Bitar KN – “The influence of extracellular matrix composition on the

differentiation of neuronal subtypes in tissue engineered innervated intestinal smooth

muscle sheets:, Biomaterials (2014), in press, June 2014.

Raghavan S, Miyasaka EA, Gilmont RR, Somara S, Teitelbaum DH, Bitar KN – “Peri-

anal implantation of bioengineered human internal anal sphincter constructs intrinsically

innervated with human neural progenitor cells” , Surgery (2014), 155(4): 668-674

Bitar KN, Raghavan S, Zakhem E – “Tissue Engineering in the Gut: Developments in

neuromusculature”, Gastroenterology (2014), 146(7): 1614-1624

Zakhem E, Raghavan S, Bitar KN – “Neo-innervation of a bioengineered intestinal

smooth muscle construct around chitosan scaffolds”, Biomaterials (2014), 35(6): 1882-

1889

Gilmont RR, Raghavan S, Somara S, Bitar KN – “Bioengineering of physiologically

functional intrinsically innervated human Internal Anal Sphincter constructs” – Tissue

Engineering Part A (2014), 20(11-12): 1603-1611

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Bitar KN and Raghavan S, - “Intestinal Regeneration: The Bioengineering Approach” –

Regenerative Medicine Applications in Organ Transplantation, First Edition, Elsevier

(2014)

Raghavan S, Gilmont RR, Bitar KN – “Neuroglial differentiation of adult enteric neuronal

progenitor cells as a function of extracellular matrix composition” – Biomaterials (2013),

34(28):6649-58

Bitar KN, Raghavan S – “Intestinal Tissue Engineering: Current concepts and Future

vision of regenerative medicine in the gut” – Neurogastroenterology and Motility (2012),

24(1):7-19

Bitar KN, Gilmont RR, Raghavan S, Somara S – “Cellular Physiology of Gastrointestinal

Smooth Muscle” –Physiology of the Gastrointestinal tract, Fifth Edition, Elsevier (2012)

Zakhem E, Raghavan S, Gilmont RR, Bitar KN-“Chitosan-based scaffolds for the

support of smooth muscle constructs in intestinal tissue engineering”-

Biomaterials(2012),33(19):4810-7

Fang Y, Frampton JP, Raghavan S, Sabahi-Kaviani R, Luker G, Deng CX, Takayama

S- “Rapid generation of multiplexed cell cocultures using acoustic droplet ejection

followed by aqueous two-phase exclusion patterning”-Tissue Engineering Part C(2012),

18(9):647-657

Raghavan S, Gilmont RR, Miyasaka EA, Somara S, Srinivasan S, Teitelbaum DH, Bitar

KN- “Successful implantation of Bioengineered, Intrinsically innervated, Human Internal

Anal Sphincter” – Gastroenterology (2011), 141(1):310-319

Miyasaka EA, Raghavan S, Gilmont RR, Mittal K, Somara S, Bitar KN, Teitelbaum DH –

“In vivo growth of a bioengineered internal anal sphincter: comparison of growth factors

for optimization of growth and survival” –Pediatric Surgery International (2011),

27(2):137-143

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Raghavan S, Lam MT, Foster LL, Gilmont RR, Somara S, Takayama S, Bitar KN -

“Bioengineered three-dimensional physiological model of colonic longitudinal smooth

muscle in vitro”, Tissue Engineering Part C (2010), 16(5):999-1009

Raghavan S, Miyasaka EA, Hashish MS, Somara S, Gilmont RR, Teitelbaum DH, Bitar

KN- “Successful implantation of physiologically functional bioengineered mouse internal

anal sphincter” – Am J Physiol Gastrointest Liver Physiol (2010), 299(2): G430-9

RESEARCH EXPERIENCE

Graduate Student Researcher, Wake Forest Institute for Regenerative Medicine (August

2011- May 2014)

Graduate Student Research Assistant & Research Technician II, University of Michigan

Medical School (May 2008 – July 2011)

Graduate Student Researcher, Department of Biomedical Engineering, University of

Michigan (December 2006- March 2008)

Research & Project Assistant, Biomedical Research Wing, Vijaya Health Center (August

2005-April 2006)

Summer Intern, Department of Biophotonics, Indian Institute of Sciences (April 2004-

August 2004)

PODIUM PRESENTATIONS

“Extracellular matrix composition modulates motor neuronal subtypes in tissue

engineered innervated intestinal smooth muscle sheets”, Distinguished abstract lectures,

Digestive Diseases Week 2014 (Chicago, IL)

“Extracellular matrix directs differentiation of motor neuronal subtypes in tissue

engineered innervated intestinal smooth muscle sheets”, Engineering Matrices for

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Tissue Regeneration, Tissue Engineering and Regenerative Medicine Americas 2013

(Atlanta, GA)

“Neuroglial differentiation of adult enteric mammalian progenitor cells as a function of

extracellular matrix composition”, Virginia Tech-Wake Forest School of Biomedical

Engineering & Sciences 2013 (Blacksburg, VA)

“In situ implantation preserves myogenic and neuronal responses in intrinsically

innervated bioengineered human internal anal sphincter tissue constructs”, AGA

Distinguished Abstracts Plenary Session, Digestive Diseases Week 2011 (Chicago, IL)

“Comparison of angiogenic growth factors for physiological functionality of implanted

bioengineered mouse internal anal sphincter”, Anorectal Motility Session, Digestive

Diseases Week 2011 (Chicago, IL)

“Successful implantation of physiologically functional intrinsically innervated

bioengineered human internal anal sphincter”, Anorectal Motility Disorders session,

Digestive Diseases Week 2010 (New Orleans, LA)

“Physiology of intrinsically innervated bioengineered construct from human internal anal

sphincter smooth muscle cells”, Anorectal Motiity Disorders session, Digestive Diseases

Week 2010 (New Orleans, LA)

“Successful implantation of physiologically functional bioengineered mouse Internal Anal

Sphincter” – Fecal Incontinence session, Digestive Diseases Week 2009 (Chicago,IL)

AWARDS AND ACHIEVEMENTS

Student Co-Chair, Imaging Technologies Session, TERMIS 2013 (Atlanta, GA).

Graduate Student Fellowship – School of Biomedical Engineering and Sciences, 2012

Gold medalist for Engineering - Anna University, 2006.

Sprachkurs Stipendiaten for Germanic languages– Goethe Institut, 2005.