In Vitro Assessment of Osteoblast Behavior in

Craniosynostosis

by

Tatiana Karine Simon Cypel

A thesis submitted in conformity with the requirements for the degree of Master

of Science

The Institute of Medical Science

University of Toronto

© Copyright by Tatiana Karine Simon Cypel – 2011

ii

IN VITRO ASSESSMENT OF OSTEOBLAST BEHAVIOR IN

CRANIOSYNOSTOSIS

Tatiana Karine Simon Cypel

Masters of Science

The Institute of Medical Science

University of Toronto

2011

ASTRACT

Introduction: The objective of this study is to investigate the role of osteoblasts in

the pathophysiology of premature suture fusion in infants.

Hypothesis: Regional variations in osteoblast function and cell signalling exist in

calvaria of infants with craniosynostosis.

Methods: Bone and periosteal tissue from fused and patent cranial sutures and

adjacent bone were harvested from infants undergoing surgery for craniosynostosis and used

to develop primary osteoblast cell cultures. Dural tissue was obtained from neurosurgical

procedures in order to generate an osteoblast-dural co-culture. Osteoblast proliferation,

differentiation, mineralization, protein expression (Noggin, BMP3 and Runx2) and response

to exogenous FGF2 stimulation were assessed.

Results: Cell cultures demonstrated significant (p<0.05) regional variations in

osteoblast proliferation, alkaline phosphatase and in vitro bone nodule formation. The

iii

expression of anti-osteogenic molecules (Noggin and BMP3) was decreased in osteoblasts

from fused suture regions. Expression of Runx2 was increased in fused suture osteoblasts in

dural co-culture.

Conclusion: The creation of a pro-osteogenic environment through the decreased

expression of anti-osteogenic signalling molecules and increased expression of osteogenic

factors may be responsible for premature suture fusion in infants.

iv

AKNOWLEDGEMENTS

I would like to thank Dr. Christopher Forrest for giving me the opportunity to work in the

exciting and very important field of craniofacial care and research. Dr. Forrest has always

taken time to ensure that I had all the support required to complete my work and he was

always keen to provide me opportunities in order to increase my level of knowledge.

Furthermore, his qualified guidance and enthusiastic supervision in my research project,

along with his commitment to patient care, have provided me with a role model of a

contemporary surgeon.

I would like to also thank Dr. Cho Pang for these two years. Dr. Pang taught me how to think

and work as a scientist.

Drs Iona Leong, Cho Pang and Peter Dirks, as members of my Program Advisory

Committee, have been invaluable in providing their expert guidance and ideas to further

enrich my research and make it productive.

The research outlined here would not have been possible without the technical and

intellectual support of my colleagues in the Craniofacial Surgery Department ( Dr. John

Phillips and the craniofacial clinical fellows) and Neurosurgery Department (Dr. Rutka and

v

Dr. Drake). Homa Ashrafpour and Ning Huang, as our lab manager and technician

respectively, have done excellent work in keeping the lab efficient and supporting my

experiments. Thanks to Balram Sukhu, his expertise in bone cell culture and osteoblast

behavior made this work possible.

I also would like to thank Dr. Rinaldo De Angeli Pinto, chair of the Division of Plastic

Surgery (Federal University of Rio Grande do Sul) where I performed my plastic surgery

training in Brazil. Dr. De Angeli was for me, an example of a superb, ethical, and highly

competent surgeon. He created the surgical “personality” I currently have and hopefully I

will carry that for my entire career.

I would like to acknowledge the contribution of Nicole Gojska in assisting with technical

work in the conditioned culture medium project.

No research would be possible without sufficient resources. I would like to thank the

Craniofacial Care and Research Funding, The SickKids Start up Funding, The Komedyplast

and Amercian Society of Craniofacial Surgery, The Physician’s Service Incorporate

Foundation and the American Society of Maxillofacial Surgeons for supporting and making

this work possible.

vi

DEDICATION

This thesis is dedicated to my family: my parents and two sisters who have always supported

me and stood behind me and my husband Marcelo, who has shared my challenges and

successes with the greatest of understanding, patience and support.

vii

TABLE OF CONTENTS

Page

I. List of Tables x

II. List of Figures x

III. List of Appendices xiii

IV. List of Abbreviations xiv

V. Introduction 1

(a) Overview of Craniosynostosis 2

- Classification 3

- Functional Problems Associated with Craniosynostosis 5

- Surgical Treatment 7

(b) Pathogenesis of Craniosynostosis 9

- Embryology of Cranial Suture 9

- Normal Skull and Suture Growth 10

- Normal Suture Fusion 10

- Historical Theories of Craniosynostosis 11

- Current Theories 12

viii

- The Role of Anti-osteogenic Signalling 13

- The Role of Dura Mater 17

- The Role of Runx2 18

- The Role of FGFs 20

- Experimental Models for Craniosynostosis Research 22

- Effect of Culture Medium Composition on Osteoblast Function 23

VI. Hypothesis 27

VII. Material and Methods 30

- In Vitro Human Osteoblast Cell Culture Model 31

- Statistical Analyses 38

VII. Results

- Demographics 40

- Histology 41

- Collagen I Expression 42

- Validation of Cell Culture Model 43

- Medium Composition 45

- Osteoblast Proliferation 47

- Runx2 Expression 52

- Alkaline Phosphatase Activity 54

- Mineralization 58

- Transmission Electron Microscopy 63

- Expression of Anti-osteogenic Signalling Molecules 64

- Dura Mater Expression of FGF2 and TGF-β1 70

ix

- Recombinant Human FGF2 Stimulation 71

VIII. Discussion 77

IX. Conclusion 93

X. Future Directions 96

XI. References 99

XII. Appendices 110

- Detailed Protocols 111

x

LIST OF TABLES

Page

Table 1: Classification of Craniosynostosis 3

Table 2: Demographics of Patients with Craniosynostosis Enrolled in the Study 40

LIST OF FIGURES

Page

Figure 1: Single-suture Synostosis Phenotypic Expression 4

Figure 2: Mechanism that BMP3 and Noggin Cause Inhibition of Bone Formation 16

Figure 3: The Role of Runx2 in Osteogenic Differentiation 19

Figure 4: Potential Mechanisms of Craniosynostosis 26

Figure 5: Hematoxylin-and eosin Staining for Bone Tissue Sampling 41

Figure 6: Immunohistochemistry Staining for Collagen I 42

Figure 7: Evidences of Osteoblasts in our Cell Culture Model 43

Figure 8: MTT Assays of Human Cranial Suture-derived Osteoblasts 46

Figure 9: Cellular Growth Prior and Post-subculture 47

Figure 10: Osteoblast Proliferation Rates (MTT) 48

Figure 11: Proliferation Rates for Syndromic Patients 49

xi

Figure 12: Osteoblast Proliferation Rates in Co-culture with Dura Mater 50

Figure 13: Expression of Runx2 Demonstrated by Immunohistochemistry 52

Figure 14: Expression of Runx2 by Western Blot 53

Figure 15: Alkaline Phosphatase Assay Assessing Osteoblast Differentiation Rates 54

Figure 16: Alkaline Phosphatase Activity 55

Figure 17: Analysis by qRT-PCR of Alkaline Phosphatase and Osteocalcin 56

Figure 18: AP for Osteoblast Co-cultured with Dura Mater Cells 57

Figure 19: Bone Nodule Formation – Alizarin Red Assay 59

Figure 20: Bone Nodule Formation at Days 14 and 18 60

Figure 21: Mineralization at Days 21 and 28 61

Figure 22: Mineralization for Osteoblasts Co-cultured with Dura Mater 62

Figure 23: Transmission Electron Microscopy of Bone Nodules 63

Figure 24: Immunohistochemistry Analysis of Noggin in Tissue Samples 64

Figure 25: Graphic Analysis for Noggin in the Tissue Samples 65

Figure 26: Immunohistochemistry Analysis of BMP3 in Tissue Samples 66

Figure 27: Graphic Analysis for BMP3 in the Tissue Samples 67

Figure 28: Western Blot Analysis of BMP3 and Noggin 68

xii

Figure 29: Expression of Pro-osteogenic Molecules in the Conditioned Medium from

Dura Mater Cells 70

Figure 30: Proliferation Rates of Osteoblasts Stimulated with FGF2 72

Figure 31: Proliferation Rates after Stimulation with FGF2 (MTT) 73

Figure 32: AP Staining after Stimulation with FGF2 75

Figure 33: APA after Stimulation with FGF2 76

xiii

LIST OF APPENDICES

Page

I. Calvarial Osteogenic Cell Culture 111

II. Medium Composition Study 114

III. MTT Assay 116

IV. Alkaline Phosphatase Activity 117

V. Alkaline Phosphatase Staining 118

VI. Alizarin Red Staining 119

VII. Transmission Electron Microscopy 119

VIII. Histology 120

IX. Immunohistochemistry and Immunofluoresence 120

X. Western Blot 121

XI. Real Time PCR 124

XII. Materials 126

xiv

LIST OF ABREVIATIONS

AMEM (αAMEM) Alpha Modified Eagle Medium

ANOVA Analysis of Variance

AP Alkaline Phosphatase

APA Alkaline Phosphatase Activity

BMP2 Bone Morphogenic Protein 2

BMP3 Bone Morphogenic Protein 3

BMP4 Bone Morphogenic Protein 4

BMP7 Bone Morphogenic Protein 7

CT Computed Tomography

DMEM Dulbeco Modified Eagle Medium

Cbfa1 Core binding factor alpha 1

DNA Deoxyribonucleic acid

FBS Fetal Bovine Serum

FGFs Fibroblast Growth Factors

FGFR Fibroblast Growth Factor Receptor

FGF2 Fibroblast Growth Factor 2

FS Fused Suture

IHC Immunohistochemistry

I-SMADS Inhibitory Smads

MTT Methylthiazoletetrazolium

PS Patent Suture

xv

qRT-PCR Real Time Quantitative Reverse Transcripition PCR

REB Research Ethical Board

RNA Ribonucleic Acid

SMAD Sma and Mad Related Proteins

TEM Transmission Electron Microscopy

TGF-β Transforming Growth Factor-β

WB Western Blot

1

Introduction

2

OVERVIEW OF CRANIOSYNOSTOSIS

Craniosynostosis, the premature fusion of one or more cranial sutures, is a relatively

common congenital disorder, affecting as many as 1 in 2000 to 2500 live births worldwide1.

Hippocrates provided the first description of this anomaly in 100 B.C. He noted the variable

appearance of the calvarial deformity and correlated it with the pattern of cranial suture

involvement. In 1851, Virchow was the first one to recognize that cranial sutures were

responsible for growth of the calvarium at right angles to the suture and that premature fusion

resulted in growth arrest at right angles to the suture with compensatory growth at patent

sutures2.

Premature ossification of cranial sutures may lead to a number of serious

morphologic and functional consequences, such as increased intracranial pressure,

developmental delay, visual and hearing impairment and can require major staged

reconstructive procedures to correct the condition. Despite its prevalence, the cause of

craniosynostosis still remains largely unknown.

Craniosynostosis may be classified by the number of involved sutures, etiology or

whether the condition is associated with a syndrome or not (Table 1).

3

Table 1: Classification of Craniosynostosis

Anatomical

Single Suture Multiple Sutures

Sagittal Any combination possible – usually bi-coronal

Metopic

Unicoronal

Lambdoid

Minor suture (fronto-sphenoid, zygomatico-temporal)

Etiology

Primary

Non-Syndromic Syndromic

Scaphycephaly (sagittal) Apert’s syndrome

Trigonocephaly (metopic) Crouzon’s syndrome

Anterior plagiocephaly (unicoronal) Pfeiffer syndrome

Posterior plagiocephaly (lambdoid) Jackson-Weiss syndrome

Turribrachycephaly (bi-coronal) Carpenter syndrome

Oxycephaly (delayed bicoronal) Muenke syndrome

Saethre-Chotzen syndrome

Secondary

Biomechanical (shunt)

Bone metabolic disorders

Nutritional

The major cranial sutures involved (in order of decreasing frequency) are the sagittal,

metopic, coronal and lambdoid sutures and each give a characteristic shape to the cranial

vault (Figure 1).

4

Figure 1: Single-suture Synostosis Phenotypic Expression: Single suture non-

syndromic craniosynostosis phenotypic and radiologic (CT) features.

Single suture craniosynostosis is usually an isolated phenomenon and has a low

incidence of familial occurrence, generally considered to be less than 5%. The highest

incidence of familial occurrence is with sagittal synostosis, which has been reported to be as

high as 8%3.

While the majority of craniosynostosis occur due to unknown causes,

craniosynostosis can occur as a result of metabolic disorders (e.g., hyperthyroidism),

malformations (e.g., holoprosencephaly, microcephaly, shunted hydrocephalus,

5

encephalocele), drug exposure (e.g., valproic acid, phenytoin) or mucopolysaccharidosis

(e.g., Hurler’s syndrome, Morquio’s syndrome).

FUNCTIONAL PROBLEMS ASSOCIATED WITH CRANIOSYNSTOSIS

Premature fusion of the cranial sutures may be associated with a variety of clinical

problems ranging from cranial facial dysmorphism to increased intracranial pressure due to

cranio-cerebral disproportion. The severity and extent of involvement is dependent upon the

number of affected sutures and the presence or absence of an associated syndrome. This

section details the potential functional problems associated with craniosynostosis.

Increased Intracranial Pressure: Elevated intracranial pressure may be associated

with craniosynostosis4. Renier et al. published the first major study to measure the

preoperative and postoperative differences in intracranial pressure in infants with

craniosynostosis. Nonsyndromic patients overall had a 14% incidence of elevated intracranial

pressure before surgery. The number of involved sutures affects the probability that

intracranial pressure will be elevated. Preoperatively, 8.9% of the patients with one suture

affected had elevated intracranial pressure but those with multiple sutures involved had a

45% incidence of intracranial hypertension. The effect of craniosynostosis, elevated

intracranial pressure, and syndromic disease on intelligence is less clear. In general, for most

nonsyndromic patients, intelligence is normal with single-suture involvement (>90% IQ ≥

90) but decreases with multiple suture involvement (78% IQ ≥ 90)5. For syndromic patients,

the degree of developmental delay is highly variable, depending on the syndrome and

severity of disease.

6

Orbits: Exorbitism is a significant component for syndromic patients with

craniosynostosis. Hypoplastic orbits and a retruded midface can cause globe and corneal

exposure, which can result in corneal injury and exposure keratitis. The majority of orbital

problems occur relatively early, in the first 5 years of life which represents the main period of

orbital growth.

Airway: Syndromic craniosynostosis may be associated with airway compromise that

requires early recognition and treatment to prevent cardiopulmonary and neurologic

sequelae6. This may be due to midface retrusion and choanal atresia resulting in airway

obstruction. The reported incidence of airway obstruction in syndromic craniosynostosis

ranges from 40% to 100%6. Many of these children will additionally have lower airway

anomalies that include tracheomalacia, complete cartilaginous tracheas, and granulations.

Airway obstruction and hypoxia during sleep may present as snoring, noisy respirations,

apneic episodes, paradoxical chest movements, persistent restlessness, feeding difficulties,

failure to thrive, hypertension, daytime fatigue, and cardiopulmonary or neurologic

impairment7. Complications of persistent airway obstruction include respiratory infections,

cor pulmonale, neurologic dysfunction and brain damage.7

Neurodevelopment Particularly problematic is the issue of intelligence and

neurocognitive development. Virtanen et al8 contended that in a series of patients undergoing

early operative correction for sagittal craniosynostosis, certain indices of neurocognitive

performance were below those of age-matched control subjects and remained delayed

throughout the period of examination after surgery. Other authors contended that although

young children with craniosynostosis are often normal from a mental standpoint, there is an

increase in frequency of psychomotor problems as they develop5.

7

Aesthetic/Psychosocial Aesthetic considerations are more difficult to quantify than

the objective values of protecting vision, reducing intracranial pressure, or improving

occlusion. Infants with craniosynostosis have visibly altered craniofacial appearance, which

varies in relation to the location and extent of suture involvement. The possibility of

permanent abnormality in facial or cranial appearance seems to greatly

affect parents’

decisions to have their infant undergo craniofacial surgery. Many surgeons believe that the

primary indication for cranioplasty in isolated synostosis is cosmetic

rather than functional

9.

There is clear evidence in other groups of children that even mild deviations from typical

facial appearance can have significant impact on psychological adjustment. Congenital

defects involving an infant’s face and skull seem to evoke particularly strong emotional

responses from parents, who must contend with a host of potentially stressful

events and

circumstances, including the infant’s unusual

appearance, potentially life-threatening

surgeries and other medical procedures, and the possibility of future neuropsychological

and

educational problems. All of these factors can potentially affect parents’ responsiveness

and

adaptation to the infant with craniofacial abnormality10

.

SURGICAL TREATMENT

The field of craniofacial surgery has existed for only a short time in the overall

history of medicine. Of course, the patients have always been there, with a birth prevalence

of craniosynostosis of approximately 1 in 2000 live births1; but it is only since Paul Tessier

began his pioneering work in the late 1960s that the field of craniomaxillofacial

reconstruction developed. Since the 1970s the specialty of craniofacial surgery has grown to

include multidisciplinary teams treating a wide variety of patients.

8

Indications for Treatment

In nonsyndromic patients, individual cranial sutures may be fused, resulting in an

abnormality of shape requiring cranio-orbital reshaping, but the midface is generally

unaffected. In syndromic craniosynostosis, in addition to cranial suture abnormalities, other

facial skeletal anomalies exist including a shortened cranial base, orbital hypoplasia, midface

and zygomatic retrusion to name a few. The reasons for surgical intervention in the

nonsyndromic group may be primarily aesthetic; in the syndromic group, they are

multifactorial. Aesthetic considerations are more difficult to quantify. Nevertheless, the

benefits of improving appearance on psychosocial adjustment have been evaluated across

many different populations11

.

Timing of Surgery

Advances in pediatric anesthesia and intensive care have allowed extensive cranial

reconstructions in infancy that previously were available only to adolescents and adults.

Concerns for the negative effect of early intervention on skeletal growth were reduced by

studies demonstrating that in syndromic patients, the midface was deficient whether patients

had early surgery or not12, 13

.

McCarthy and Cutting14

proposed that the first procedure, cranial vault remodeling

and fronto-orbital advancement, be performed between 6 and 9 months of age. At an earlier

age, the bone is more fragile; at a later age (>18 months), residual calvarial defects will fail

to reossify. Early surgery is also important because of rapid growth of the brain, which more

than doubles in volume during the first year of life15

.

9

Pathogenesis of Craniosynostosis

During cranial development, adjustment to the expanding brain takes place by bone

deposition at sutural margins and on the ectocranial surface of the calvarium, and by bone

resorption on the endocranial surface16

. The cranial sutures, as primary areas of growth

during the expansion of the cranium16, 17

play a pivotal role. Although a descriptive

knowledge of suture morphogenesis and function is well reported16, 18

, the spatial and

temporal regulation of bone deposition, resorption, and remodeling are not well understood19

.

Embryology of Cranial Suture

Sutural morphogenesis occurs in midgestation when enlarging bone plates in the

primordial cranium come into apposition16, 17

. Mesenchymal cell populations along the

expanding osteogenic fronts differentiate into osteoblasts and contribute to the formation of

osteoid and the mineralization of new bone17, 18

, bringing opposing bones into close

approximation. An intervening zone of immature mesenchymal tissue is thereby created

between them, forming the blastema of the suture18, 20

Suture mesenchymal cells continue to

proliferate and organize within the fibrous suture matrix, which acquires the characteristic

appearance of the mature, multilamellar suture17, 18

. Simultaneously, populations of

mesenchymal cells bordering the suture along the osteogenic fronts continue to differentiate

into osteoblasts, contributing to the formation of new bone17, 18

. From the time of their

formation, sutures are extremely active centers of cellular proliferation, cellular

differentiation, tissue synthesis, and remodeling. The intricate regulation of these processes

allows for the completion of cranial morphogenesis while preventing the formation of bone

10

across the sutural space, an event which occurs only in the pathologic condition of

craniosynostosis or part of normal suture fusion later in adult life19, 21, 22

.

Normal Skull and Suture Growth

Intramembranous ossification of the skull begins at the end of the second month of

gestation. A center of osteogenesis develops directly in vascularized mesenchyme. Expansion

of the ossification center proceeds rapidly via appositional growth. Initially cancellous bone

forms, but as trabeculae thicken and the bone becomes less porous, it become compact bone.

Eventually each intramembranous cranial bone has enlarged to the point at which it

articulates with an adjacent bone via a syndesmosis or sutures. Growth then proceeds at the

sutures19

.

Growth at the suture area is a secondary, compensatory, and mechanically obligatory

event following the primary growth of the enclosed brain and ocular globes. The bones of the

calvaria are displaced outward by the enlarging brain. Each bone of the domed skull roof

responds to the expansion of the brain by depositing new bone at the contact edges of the

sutures.

Normal Suture Fusion

Functioning sutures are the sites of continuous bone deposition and resorption.

Initially, sutures are straight edges of bone separated by connective tissue. Gradually,

interdigitations develop and become more prominent with time19

. For interdigitations to

form, develop, and interlock, the distribution of osteoblasts along sutural bone must be

uneven with clumps of osteoblasts at the tip of each interdigitation23

. Sutural interdigitations

11

may permit adjustive movements and/or stress reduction. Their architecture may depend on

the types and distribution of forces.

Suture closure has been attributed to vascular, hormonal, genetic, mechanical, and

local factors. Biomechanical factors have been a perennial favorite mechanism24

. The cause

of suture closure is still unclear. There may be one or possibly more than one mechanism.

The relationship between suture closure, cessation of growth, and functional demands across

sutures poses questions about various biological relationships. Does cessation of growth lead

to suture fusion? The growth of the human brain ceases prior to the onset of osseous fusion

of the cranial sutures. With this is a delay from completion of brain growth to sutural fusion

in the 20s and 30s.

Historical Theories of Craniosynostosis

Although certain cranial deformities arise from mechanical or functional causes (e.g.,

plagiocephaly and hydrocephaly), the molecular basis of the majority of craniofacial

abnormalities is becoming increasingly evident through advancements in molecular biology.

Early explanations of cranial suture fusion included anectodal associations with intrauterine

constraint, uterine malformations, decreased amniotic fluid, or breech presentation. Ozaki et

al25

performed ultrastructural analysis of sagittal sutures in the process of fusion. Their

analysis revealed several new facts:

1. Premature fusion of sutures was found to begin centrally in the suture.

2. It began endocranially as opposed to both endocranially and ectocranially in

normal sutures.

12

3. It exhibited a disorganized ultrastructure of lower density than in normal sutures.

Molecular biology has now taken us beyond the speculative explanations of

mechanical causes to the roots of abnormal suture development. This progression is

particularly evident in autosomal dominant syndromic craniosynostosis subtypes.

Current Theories

Recent work has demonstrated that fusion of the calvarial sutures is mediated by

locally elaborated soluble growth factors, leading some to speculate that external

biomechanical forces play little role in suture development. Historically, the theory that fetal

head constraint may play a critical role in the pathogenesis of many cases of nonsyndromic

craniosynostosis has been supplanted by humoral theories although it is possible that cranial

biomechanical stresses experienced in fetal and early life might be the trigger that leads to

dural cytokine signalling involved with suture fusion and/or patency26

.

On the other hand, research focused on the molecular mechanisms underlying normal

cranial suture fusion has demonstrated the importance of dura mater mediated cell signalling

in the complex process of fusion of normal cranial sutures. It is hypothesized that the dura

mater acts as a regionally specific endogenous tissue engineer, releasing growth factors in a

specific orchestrated fashion that cause the overlying cranial suture to close in a predictable

fashion. Possible candidates for these growth factors include fibroblast growth factor (FGF),

and transforming growth factor-β (TGF-β) isoforms.

In addition to the effects of these locally released growth factors, cranial suture

development has been shown to be influenced by anti-osteogenic signalling molecules such

13

as Noggin and Bone Morphogenic Protein 3 (BMP3), which are upregulated in patent sutures

during the normal process of suture fusion thereby maintaining the cranial suture patency27,

28. Furthermore, the role of Runx2, a transcription factor that is a marker of osteoblast

differentiation, has been implicated in the process of normal cranial suture fusion. Runx2 is

found in osteogenic fronts and sutural mesenchyme and has been demonstrated to be

upregulated in fusing sutures during the process of normal suture fusion. Runx2 up regulation

enhances differentiation and bone production leading to earlier suture fusion. This factor has

been shown to regulate the expression of a number of proteins, including osteocalcin,

produced by the mature osteoblast and responsible for its bone formation29

.

The Role of Anti-osteogenic Signalling

Noggin

Noggin, a secreted BMP2/4 antagonist produced by osteoblasts and released in the

extracellular matrix, is important in the process of normal suture fusion27

. Noggin is a

polypeptide that inhibits Transforming Growth Factor – β (TGF-β) signal transduction by

binding to TGF-β family ligands and preventing them from binding to their correspondent

receptors (Figure 2). Down-regulation of pro-osteogenic BMP signalling (part of the TGF-β

superfamily) is then observed which maintains suture patency27

. On the other hand, down-

regulation of Noggin expression results in disinhibition of pro-osteogenic BMP signalling

(BMP2,4,7) increasing bone formation which leads to suture fusion and may be one of the

molecular mechanisms involved in the pathophysiology of craniosynostosis.

Research findings in a murine model of normal suture fusion demonstrate that down-

regulation of Noggin expression in the normally fusing posterior frontal suture increased

14

bone formation with resulting suture fusion. In a normal non-fusing suture (sagittal),

increased Noggin expression results in suture patency27

. It has also been shown that the

suture-specific dura mater is an independent source of Noggin. Cultured dura mater cells

from patent sutures expressed high levels of Noggin protein, whereas the dura mater from the

fusing posterior frontal suture expressed almost undetectable levels of Noggin30

.

Experiments in a rabbit model of congenital coronal craniosynostosis also

demonstrated the interaction between Noggin and premature suture fusion. Fusing sutures

showed low Noggin expression29

. Underexpression of Noggin was also found in the dura and

coronal mesenchyme prior to suture fusion. In contrast, in the same model, the patent coronal

and sagittal sutures expressed normal levels of Noggin leading to suture patency29

.

Despite the current findings strongly suggesting an important role for Noggin in

maintaining suture patency in animal models of normal suture fusion, there is a lack of

understanding about Noggin expression and its interactions in infants with craniosynostosis.

Bone Morphogenic Proteins

The BMPs are growth factors secreted by osteoblasts and released in the extracellular

matrix. They are part of the TGF-β superfamily, which are well known for their ability to

induce the formation of bone and cartilage. The actions of these growth factors are highly

concentration dependent and influence a number of cellular processes. For instance, BMP2, 4

and 7 have been shown to promote cellular chemotaxis and proliferation at low extracellular

concentrations and to induce cellular differentiation and bone formation at high extracellular

concentrations31

.

15

BMP3 is an antagonist of BMP2 and BMP4 32

. Rather than impeding BMP signalling

of bone formation by binding to a ligand and preventing specific ligand-receptor interactions

(as does Noggin), BMP3 activates a TGF-β/activin–specific response pathway33

(Figure 2).

Activin is a member of the TGF-β superfamily, and antagonizes the BMP pathway by

competing for SMAD proteins; SMAD proteins are transcription factors that regulate the

expression of genes involved in the modulation of the activity of TGF-β ligands involved in

osteoblast differentiation and bone formation34

.

BMP3 has been implicated in the process of normal suture fusion in mice28

. Altough

the source of BMP3 during normal suture fusion is not clear, its expression pattern is

consistent with that of an antagonist playing a role in suture fusion and patency. BMP3 levels

decreased in the posterior frontal suture during suture fusion and were maintained or

increased in the patent sagittal suture35

. It is speculated that BMP3 may be negatively

regulated by osteogenic factors such as FGF2 and TGF-β1 which are differentially expressed

in the fusing posterior frontal and sagittal suture complexes36, 37

. These factors are noted to

increase in the dura mater underlying the fusing posterior frontal suture during fusion when

compared with the patent sagittal suture30

.

16

Activin ResponsePathway

BMP2

Noggin

SMAD Activation andNuclear TranslocationResulting in Bone Formation

Role of Noggin and BMP3 Signalling

Promotes bone formation

Binds to BMP2 blocking the

stimulus for bone formation

Antagonize bone formation by

competing for SMADS protein

BMP-3 activates a TGF-β/activin-specific response antagonizing osteogenic signaling

BMP3

No signalling

Figure 2: Mechanism by which BMP3 and Noggin cause inhibition of bone formation.

Schematic of Noggin and BMP3-mediated antagonism of bone morphogenetic protein (BMP)

signalling. Antagonists such as Noggin bind to BMP ligand and prevent ligand-receptor interactions.

BMP3 binds to TGF-β/activin receptors and blocks BMP signalling downstream of activated BMP

receptor complexes.

Despite the insights in Noggin and BMP3 expression during the normal suture fusion

process, it is not clear if down-regulation of these anti-osteogenic molecules during

premature suture fusion is a cause or the effect of pro-osteogenic activation leading to

premature ossification in craniosynostosis. Additional investigation of the expression of

osteogenic antagonists and their regulation will further advance our knowledge of the

17

complex cascades regulating suture fusion and patency in infants with syndromic and non-

syndromic craniosynostosis.

The Role of Dura Mater

Central to many studies of cranial sutures has been the role of the dura mater.

Historically, dura mater was thought to be a conduit for tensile forces transmitted from the

expanding neurocranium38, 39

. The formation of sutures was seen as a byproduct of this

mechanical phenomenon, forming along dural reflections in both normal and disease states39

.

While several animal models have established the importance of dura mater in the

regeneration of normal cranial bone40

and sutures in developing animals, evidence suggests

that cell signalling and humoral mechanisms are more important than biomechanical forces

with respect to bone regeneration41

.

Recent findings suggest that dura mater modulates calvarial ossification in many

ways, including providing a source of osteoblastic precursors and/or supplying osteogenic

cytokines42

. Evidences suggest that the underlying dura mater also influences the behavior of

the overlying suture complex by means of paracrine signalling43

. The dura mater underlying

the cranial suture complex is one of several sources of FGF2 and TGF-β1cytokines in vivo;

however it remains unclear whether the levels of these growth factors produced by dura

mater are capable of down-regulating anti-osteogenic molecules expression in osteoblasts to

favor a pro-osteogenic enviroment and promote premature suture fusion44

.

Li et al44

demonstrated direct evidence for a paracrine effect of juvenile dura mater

cells on osteoblasts by showing that dura mater derived FGF2 mediates mitogenic activity in

calvarial osteoblasts which is inhibited by neutralizing FGF245

. Osteoblasts demonstrated

significantly increased proliferation when combined with juvenile dura mater cells in co-

18

culture or when dura mater cell-conditioned medium was applied to them. Moreover high

levels of FGF2 protein were detected in juvenile dura mater cells and their conditioned

medium. In contrast low levels of FGF2 protein were detected in adult dura mater cells and

not detectable levels in their conditioned medium. This study reinforced the idea that FGF2

might be an important paracrine signalling factor in vivo supplied by the underlying dura

mater to stimulate the overlying calvarial osteoblasts44

.

The idea that dura mater derived from immature animals is osteoinductive and/or

osteogenic in nature was further supported by studies in which heterotopic transplantation of

the dura mater into epithelial mesenchymal pockets in adult rats caused ectopic bone

formation42

. Furthermore, when dura mater from adult guinea pigs (18 months of age) was

grafted into the base of calvarial defects created in syngeneic infant guinea pigs (3 to 4 weeks

old), incomplete reossification was observed40

. In contrast, dura mater taken from an infant

rat and placed into the calvarial defect of an adult rat markedly enhanced reossification46

.

Therefore immature dura mater seems to have a strong influence on the development

of bone formation in vivo and as such we may expect the dura mater to have similar

importance in suture regulation. Despite the advancements in the understanding of the pivotal

role of dura mater in premature suture fusion there still is a lack of information regarding the

interactions of dura mater pro-osteogenic signalling and anti-osteogenic molecules leading to

premature calvarial ossification in infants with craniosynostosis.

The Role of Runx2

Polyomavirus enhancer binding protein 2/core binding factor Alpha 1 (Cbfa1) or

currently denominated Runx2 is a master transcription factor that has been shown to regulate

19

osteoblast differentiation stimulating osteogenic gene transcription through a cascade,

starting with BMP-2 binding to its receptor (BMPR-II). This binding activates a SMAD

(signal transducers for the members of the transforming growth factor-beta superfamily)

signalling cascade, ultimately activating Runx2 and stimulating osteogenesis (Figure 3). It

also regulates the expression of a wide variety of genes responsible for the osteoblast

phenotype and function including osteocalcin and TGF-β47

. The latter provides a direct link

between transcriptional regulation and growth factor activity.

Figure 3: Role of Runx2 in osteogenic differentiation. BMP2 binds to its receptor

(BMPR-II). This binding activates a SMAD signalling cascade, ultimately activating Runx2

and stimulating osteogenic gene transcription.

Reflecting its major role in bone formation, Runx2 levels have been shown to be

elevated in areas of normal suture formation in mice48

. Mutations in which Runx2 is absent

demonstrate defects in osteogenesis49

. These studies provide a sound basis for an effort to

20

examine the activity of specific transcription factors such as Cbfa1/Runx2 in osteoblasts

derived from fused sutures and to determine whether they play a causative role in sutural

closure associated with craniosynostosis.

The Role of Fibroblast Growth Factors

Fibroblast Growth Factor 2 is a member of the fibroblast growth factor family

involved in angiogenesis, wound healing, and embryonic development50

. The FGFs are

heparin-binding proteins and interactions with cell-surface associated heparin sulfate

proteoglycans have been shown to be essential for FGF signal transduction. FGFs are key

players in the processes of proliferation and differentiation of wide variety of cells and

tissues. Several observations indicate that FGFs may play an important role in the control of

osteogenesis during skeletal development. FGF2 is a potent mesodermal inducer during

embryogenesis and FGF receptors (FGFRs) are strongly expressed in developing bones50

.

Studies in bovine calvaria cells showed that FGF2 is produced by osteoblasts and

accumulates in the bone matrix51

. In bovine and rodent calvaria-derived cells, the effects of

FGF on bone cell proliferation and differentiation appear to be opposite. FGF1 and FGF2

stimulate cell proliferation but inhibit alkaline phosphatase (AP) activity and reduce collagen

type I (ColI) and osteocalcin (OC) expression, indicating that FGF2 has independent effects

on calvarial cell proliferation. The effects of FGFs on osteoblastic cell differentiation and

bone matrix formation in long-term culture are however conflicting, since positive52, 53

and

negative effects54

have been reported, depending on the cell culture system.

Considering that dura mater cells may be a source of FGF2 and anti-osteogenic

molecules such as Noggin may be regulated by FGF2, it is important to know if this

21

molecule is the key element in the pathophysiology of craniosynostosis, which may

corroborate with the hypotheses that dura mater influences osteoblast behavior at the fused

suture site through release of FGF2 leading to decreased Noggin and BMP3 expression. It is

also unclear if osteoblasts from different sites are defective and not able to respond equally to

FGF2 stimulation, or if there is an up-regulation of this molecule at the fused suture site.

While blocking FGF signalling30

or FGF2 activity55

prevents cranial suture fusion or

osteogenesis, respectively, the findings suggest that exogenous FGF signalling is capable of

suppressing Noggin expression during cranial suture fusion.

Many researchers have concentrated efforts on investigating the genetic basis of

syndromic craniosynostosis and the functional consequences of mutations involving the

fibroblast growth factor receptor (FGFR) gene. Recent findings shown that point mutations

in FGFR-1 and FGFR-2 induce premature cranial ossification suggesting that FGF is an

important regulator of bone-forming cells during human calvaria (HC) osteogenesis56

.

Mutations of three FGFRs account for most causes of syndromic craniosynostosis, including

Crouzon’s, Crouzon’s with acanthosis nigricans, Pfeiffer’s, Apert’s, Muenke’s, Beare-

Stevenson, and Jackson-Weiss syndromes57

. On activation, the FGFR immunoglobulin-like

bonding regions form dimers, activating the intracellular tyrosine kinase. Subsequent

downstream effects on the nucleus influence cellular proliferation, differentiation, and

migration. Characterization of specific mutations in genes that cause craniosynostosis is a

step forward understanding the mechanism of normal and abnormal development of calvarial

bone. However, various approaches of research in this area are still needed to help unravel

the complex interaction of gene products that participate in signalling pathways58

.

22

Taken all findings together, FGF2 may guide suture fate (patency versus fusion). It is

also important to clarify the role of FGF2 in proliferation and differentiation, and to

investigate if different concentrations at the suture site are responsible for the regional

differences in osteoblast behavior. Furthermore, it is important to elucidate whether dura

mater is the source of FGF2 and evaluate the ability of osteoblast from different sites to

respond to FGF2 stimulation, clarifying one step of cascade that leads to craniosynostosis in

infants.

Experimental Models for Craniosynostosis Research

Over the past several years, investigation of the biology underlying programmed

posterior frontal suture fusion in rats and mice has been taken as a means of understanding

the pathology seen clinically in human craniosynostosis44

. The murine model has been

thought to be an excellent system with which to study suture development and molecular

specification between mice and humans45

. In the mouse, the posterofrontal suture lies

between two frontal bones, and the sagittal suture lies between the two parietal bones. The

posterofrontal suture undergoes fusion in a predictable manner on postnatal days 8 to 10,

whereas other sutures remain patent28

. This is analogous to humans, in which the metopic

suture fuses in infancy and the other sutures remain patent well into adulthood. Thus, the

murine posterofrontal and the sagittal sutures taken in juxtaposition, as exemplars of normal

suture fusion and patency, respectively, allow for insight into both the normal coordination of

suture fusion and possible mechanisms of craniosynostosis19

.

Currently, the most representative animal model of craniosynsostosis is the rabbit

craniosynostosis strain from the University of Pittsburgh58

. In this model, pathologic suture

23

fusion begins in utero, causing cranial vault deformities such as plagiocephaly in unilateral

coronal suture synostosis and brachycephaly in bilateral synostosis59

. This model has made it

possible to investigate the biomolecular mechanisms involved in craniosynostosis, including

the role of anti-osteogenic molecules such as Noggin and pro-osteogenic factors such as

Runx2.

Although there are few studies utilizing a limited number of discarded samples from

patients with craniosynostosis60

, to date there are no reliable models of cell cultures derived

from human calvarial bone using a large number of patients and well representing the wide

spectrum of craniosynostosis in infants.

Effect of Culture Medium Composition on Osteoblast Function

The use of normal and fused suture osteoblasts derived from cranial bones of patients

with syndromic and non-syndromic craniosynostosis provides a valuable model for

investigating molecular and cellular defects associated with this significant disorder in

infants60

. However, there is little uniformity in the conditions used in human osteoblast cell

cultures, particularly the concentration of reagents present in the media. Initial studies by

Coelho et al61

analyzed the effects of two widely used culture media, Dulbecco’s modified

Eagle’s medium (DMEM) and minimum essential medium Eagle Alpha modification (α-

MEM) on human osteoblastic characteristics, including cell viability and alkaline

phosphatase (AP) activity61

. These studies demonstrated that DMEM, a less nutrient-rich

medium when compared to α-MEM, appears to demonstrate higher values of cell

proliferation and growth61, 62

. Subsequently, many studies have found that the optimal

concentration of fetal bovine serum (FBS) to supplement the culture medium is FBS 10%,

24

for it produces the highest proliferation rates62, 63

. The success of cranial suture biology in

explaining the pathophysiological mechanism of craniosynostosis is predicated in replicable

and efficient cell culture systems that are representative of in vivo cellular dynamics.

Therefore, the standardization of culture conditions, especially the medium and the presence

of essential compounds, is critical for developing future applications of cranial suture

research in reconstructive medicine.

Summary of Research

Current research has proposed the molecular basis of craniosynostosis based on

normal suture fusion animal models. It has been shown that the patency of the suture line

depends on the balance between pro-osteogenic and anti-osteogenic signalling molecules,

and the imbalance between those may be responsible for premature suture fusion. However,

is not clear which changes in the cranial suture enviroment are responsible for this

phenomenon. To elucidate the mechanisms locally involved in premature suture fusion we

propose to create a human model of osteoblast cell culture by obtaining calvarial bone

samples from patent sutures, fused sutures and adjacent bone from infants affected by this

condition. First, we propose to evaluate the samples histologically to confirm regional

variations between suture sites and adjacent bone. We then aim to search in the surrounding

enviroment for factors that may contribute to the imbalance between pro- and anti-osteogenic

signalling. It has been shown that dura mater may be the source of growth factors, such as

FGF2 and TGF-β that orchestrate this complex process of suture fusion and patency.

However, the interaction between osteoblast and the underlying dura mater has not been well

characterized, especially in humans. By combining dura mater cells and osteoblasts from

humans (using a co-culture model) we hope to be able to better evaluate osteoblast behavior.

25

We plan to evaluate important mediators, some pro-osteogenic (FGF2) and other anti-

osteogenic (Noggin and BMP3) that may be regulated by dura mater paracrine signalling or

may be imbalanced at the suture site affecting bone formation. Noggin and BMP3 have been

shown to downregulate ossification at the suture site in order to maintain suture patency

during the normal suture fusion process. Alternatively, it has been hypothesized that the

excessive bone formation at the fused suture site may be due a defect in differentiation,

function or both in fused suture osteoblasts, independent of humoral signalling from the

surrounding microenvironment. A potential candidate to explain these defects is the

transcriptional factor Runx2. Runx2 controls osteoblast differentiation and expression of

proteins such as osteocalcin responsible for bone forming function. As such we plan to

evaluate protein levels of Runx2 in fused and patent sutures.

The strength of this research is based on the establishment of a human model of

osteoblast cell culture harvested from patients affected by craniosynostosis. We also plan to

develop a model that combines human osteoblasts with human dura mater cells, establishing

a more physiological environment to evaluate osteoblast behavior. Using this model, we have

the possibility to surpass the normal suture fusion animal model and search in more detail the

mechanisms of craniosynostosis counting on a large number of human samples representing

the wide spectrum of this condition.

26

Figure 4. Potential mechanisms of craniosynostosis

Figure 4: The patency of the suture line depends on the balance between pro-

osteogenic and anti-osteogenic signalling molecules. It is possible that an imbalance between

the pro-osteogenic and anti-osteogenic signalling molecules is responsible for the

development of craniosynostosis as seen in the panel on the right.

27

Hypothesis

Regional variations in osteoblast function and cell signalling exist in calvaria of

infants with craniosynostosis.

SPECIFIC AIMS:

I) To develop a reliable osteoblast cell culture from calvaria of infants

undergoing surgery for craniosynostosis repair.

Rationale: The use of osteoblast cells derived from cranial bones of patients with

craniosynostosis may provide a valuable model for investigating the molecular and cellular

abnormalities associated with this disorder. In order to develop a valid technique for

osteoblast cell culture, the effects of differing media will be used to determine the optimal

growth conditions. To validate our bone cell culture model and demonstrate the presence of

osteoblasts, we will assess cellular proliferation, Runx2 expression, alkaline phosphatase,

osteocalcin, collagen I expression and mineralization. To further confirm our findings of in

vitro bone formation, ultrastructural analysis of the samples will be performed by

Transmission Electron Microscopy.

II) To assess regional variations in osteoblast behavior with and without dura

mater cells in co-culture.

Rationale: Although variations in osteoblast activity have been shown in murine models of

normal suture fusion, no studies have been performed using human osteoblasts and dura

mater cells in vitro from infants with craniosynostosis. Osteoblast behavior will be studied in

28

osteogenic monocultures and co-cultures with dura mater cells by MTT (proliferation rates),

immunocytochemistry and Western Blot for Runx2 expression, alkaline phosphatase assay

(differentiation), mineralization assay, and Transmission Electron Microscopy (TEM).

Quantitative Real Time – Polymerase Chain Reaction (qRT-PCR) will be performed to

evaluate the DNA expression of alkaline phosphatase and osteocalcin to confirm the cellular

level of differentiation. It is anticipated that these experiments will demonstrate regional

variations in osteoblast proliferation and differentiation which will provide the basis for aim

III.

III) To investigate the role of anti-osteogenic signalling on human osteoblasts in

vitro with and without dura mater cells in co-culture.

Rationale: It has been demonstrated that Noggin and BMP3 are important signalling

molecules in models of normal suture fusion but their role in craniosynostosis is unknown.

Noggin and BMP3 both down-regulate bone formation around the suture promoting suture

patency. These experiments (Western blot and immunohistochemistry) will investigate the

expression of these molecules in patent and fused sutures in infants with craniosynostosis in

order to determine if regional variations in expression exist. This findings may have

significance with respect to understanding the pathophysiology of craniosynostosis in

humans.

IV) To investigate the role of dura mater paracrine signalling in the

pathophysiology of craniosynostosis in humans.

Rationale: Dura mater underlying the cranial suture complex is one of several sources of

FGF2 and TGF-β1cytokines in vivo. However, it remains unclear whether growth factors

29

produced by dura mater are capable of down-regulating BMP3 and Noggin expression in

osteoblasts and up-regulate Runx2 expression thereby influencing osteoblast behavior. The

aim of these experiments is to determine if dura mater cells in co-culture influence

osteoblasts through paracrine signalling through secretion of FGF2 and TGF-β cytokines.

V) To assess the effects of exogenous administration of FGF2 on osteoblast

function in vitro.

Rationale: It has been demonstrated that FGF2 signalling is of central importance for

premature cranial suture fusion and might be an important paracrine signalling factor from

underlying dura mater to overlying calvarial osteoblasts. These experiments will investigate

the effects of exogenous administration of FGF2 to our cell culture and compare with the

effects produced by dura mater cells in a co-culture model.

30

Material and Methods

31

Tissue Sampling:

Patient Population: Patients sequentially selected with syndromic and non-syndromic

craniosynostosis (3 months - 3 years old) scheduled to undergo elective cranial vault

reshaping for craniosynostosis at The Hospital for Sick Children between July 2008 and

September 2010 were enrolled in this study. Informed consent was obtained. This research

received REB approval. Patients with previous cranial vault surgery were excluded from the

study.

Bone samples: During surgery, bone samples (5x5mm) and periosteum (1x1cm) were

obtained from fused and patent cranial sutures and non-suture bone, during the normal course

of the procedure. The sex, age, side, suture site and type of craniosynostosis was to be

registered. Tissue samples from fused suture, patent suture and adjacent non sutural bone

were processed for routine histology to confirm the origin of each sample, confirming the

patency or fusion of the suture line, and collagen I staining (See Appendix pg.120)60

.

Dura Mater: Samples of dura mater (patients between 3 to 17 years old) were obtained

from patients undergoing surgery for epilepsy and used to develop cell co-culture models.

Aim I) To develop a reliable osteoblast cell culture from infants undergoing

surgery for craniosynostosis repair.

Bone and periosteal tissue samples were taken from the operating room,

transported in ice in 50ml tubes containing αMEM and 5x penicillin-streptomycin and

processed immediately for cell culture. Samples were sequentially digested in a collagenase

(Sigma/Aldrich C0130) mixture at 37º C for 20 min and centrifuged at 700 rpm for 8

minutes. The pellet was resuspended in αMEM (Wisent Bio-Cat# 310010) containing 10-7

M

32

dexamethasone (Sigma/Aldrich Cat# D8893) and 15% FBS and plated in 75cm2 tissue

culture flasks (SARSTEDT – Cat#83.1813.002) (See Appendix pg.111). At subconfluence,

cultures were trypsinized (0.05% Trypsin-EDTA-Wisent Bio Cat# 325042) and seeded into

the tissue culture plates – 24 wells/5000 cells per well and 96 wells/1000 cells per well

(SARSTEDT – cat# 83.1835) for analysis (See Appendix pg.112). Medium changes were

done every 2-3 days. At 1 week after subculture, the medium was additionally supplemented

with 1mM ß-glycophosphate (Sigma/Aldrich cat# G6251) and 50μg/ml Ascorbic acid

(Sigma/Aldrich Cat# A2218)64

. Identification of cultured cells was performed by phase

contrast microscopy.

In order to validate our bone cell culture model and demonstrate the presence of

osteoblasts, we assessed cellular proliferation, Runx2 expression, alkaline phosphatase

staining, osteocalcin and collagen I expression and mineralization. To further confirm our

findings of in vitro bone formation, ultrastructural analysis of the samples was performed by

TEM.

i) Proliferation Rates: To assess proliferation in cells derived from infants with

craniosynotosis, the standard MTT assay (Sigma – Ref. 5655) - (See Appendix

pg.116) was employed at days 1,3,5 and 7 following subculture. This assay

assesses mitochondrial dehydrogenase activity and can serve as an indirect

measure of cellular proliferation65

.

ii) Differentiation Rates: At the same time points, alkaline phosphatase activity

was analyzed using the standard ρ-nitrophenil phosphate assay (pNP, Sigma –

Ref. 104-0) - (See Appendix pg.117). The final alkaline phosphatase activity

was adjusted per protein content (µg) and time of assay incubation (h)64

. After

33

confluence in cell culture, imunnocytochemistry for Runx2 (abcan 54868) was

performed (See Appendix pg.120)58

.

iii) qRT-PCR: Osteoblast differentiation was also evaluated by gene expression

of Osteocalcin, which represents the latest marker of osteoblast differentiation

(Primers - Forward: GGCAGCGAGGTAGTGAAGAG and Reverse:

CTGGAGAGGAGCAGAACTGG) and Alkaline Phosphatase (Primers –

Forward: CGTGGCTAAGAATGTCATCATTGTT and Reverse:

TGGTGGAGCTGACCCTTGA) examined by real time PCR. HPRT was

used as the housekeeping gene (See Appendix pg.124)66

.

iv) IHC: Collagen I (Mouse monoclonal to Colagen I – abcan 6308) expression

was evaluated by Immunocytochemistry (See Appendix pg.120).

v) Mineralization assay: Subsequently, cells were grown for 21 and 28 days

following subculture and were analyzed for mineralized bone nodule

formation via Alizarin Red S assay (Sigma Ref. A5533) - (See Appendix

pg.119)64

.

vi) Transmission Electron Microscopy: Mineralized nodules grown on

coverslips were fixed in 2% glutaraldehyde and processed for Transmission

Electron Microscopy to confirm bone formation and structure (See Appendix

pg 119)64

.

The effects of differing media composition on cell culture was assessed in order to

optimize the culture settings for osteoblast growth and differentiation. Human osteogenic

cells from patients (n=7) with craniosynostosis were cultured in αMEM (Wisent Bio-Cat#

310010) containing 10-7

M dexamethasone (Sigma/Aldrich Cat# D8893), supplemented with

34

i) 1% FBS, ii) 10% FBS, iii) 15% FBS or iv) ascorbic acid and β-glycophosphate.

Experimental culture conditions were compared on the basis of active cell growth (MTT

reduction assay) and differentiation (AP assay).

Aim II) To assess regional variations in osteoblast behavior with and without

dura mater cells in co-culture.

Osteoblasts obtained from regions of fused suture, patent suture and adjacent non

sutural bone and periosteum were cultured in αMEM containing 10-7

M dexamethasone,

supplemented with 15% FBS for 7 days and then supplemented with ascorbic acid 50µg/1ml

of medium and 1% β-glycophosphate. Proliferation rates, differentiation, including alkaline

phosphatase, collagen I and osteocalcin and mineralization were assessed in all 3 regions. In

order to determine if dura mater cells exerted any influence on osteoblast behavior, co-

cultures were established with dura mater samples from neurosurgical patients and

proliferation rates, differentiation and mineralization were assessed. Results of proliferation

were also stratified in syndromic and non-syndromic patients.

Dura mater sample was taken from the operating room, transported in ice in 14ml

tube containing αMEM and 5x penicillin-streptomycin and processed immediately for cell

culture. Samples were sequentially digested in a collagenase (Sigma/Aldrich C0130) mixture

at 37º C for 20 min and centrifuged at 700 rpm for 8 minutes. The pellet was resuspended in

αMEM (Wisent Bio-Cat# 310010) containing 10-7

M dexamethasone (Sigma/Aldrich Cat#

D8893) and 1% FBS and plated in 25cm2 tissue culture flasks (SARSTEDT –

Cat#83.1813.002) (See Appendix pg.111)44

. At subconfluence, cultures were trypsinized

35

(0.05% Trypsin-EDTA-Wisent Bio Cat# 325042) and seeded into the tissue culture plates – 6

wells/50000 for analysis (See Appendix pg.112). Medium changes were done every 2-3 days.

i) Co-Culture Model: We plated 5.0x104

first-passage osteoblasts per well in

six-well tissue culture plates (SARSTEDT Cat# 83.1839) and 2.0x104

dura

mater cells onto correspondenting co-culture filter inserts (VWR Cat# 62406-

163). The inserts have a pore size of 0.4µm. Cells are cultured separately in

standard medium until both cell populations were confluent. Dura mater cell-

seeded filter inserts are then combined with the six-well plates of osteoblasts

and cultured in αMEM containing 10-7

M dexamethasone, supplemented with

1% FBS for 10 days. The medium is changed every other day up to 10 days

when cells and medium were collected for Western blot analysis. Osteoblasts

cultured with empty co-culture inserts serve as a control67

.

Aim III) To investigate the role of anti-osteogenic signalling on human

osteoblasts in vitro with and without dura mater cells in co-culture.

Tissue samples from fused suture, patent suture and adjacent non sutural bone were

sent for immunohistochemical analysis of Noggin and BMP3 expression. These molecules

were investigated in tissue samples to obtain true representation of tissue expression of

Noggin/BMP3 in vivo without the influence of cell culture. Osteoblasts obtained from

regions of fused suture, patent suture and adjacent non sutural bone, and periosteum were

cultured as described in Aim II. In order to determine the influence of dura mater cells on

Noggin and BMP3 expression, a co-culture was established and medium from osteoblasts

36

alone or in co-culture was collected at day 10. Protein expression was measured by Western

Blot.

i) Investigation of Noggin and BMP3 expression by immunohistochemistry

Samples were demineralized with EDTA, fixed in 10% formalin, embedded in

paraffin, microtomized (5µm) and stained with rabbit polyclonal antibody to

Noggin (abcam) used in dilution 1:20 or rabbit polyclonal antibody to BMP3

(R&D System) used in dilution 1:5 to analyze their spatial expression patterns

in fused and patent sutures and non-suture bones. Detection was performed

with Goat Polymer (Biocaremedical) for BMP3 and ABC Ellite System

(Vector) for Noggin and Collagen I. 3,3’Diaminobenzidine (DAB) was used

as chromogen. Semi-quantitative analysis of the staining was carried out. The

staining intensity in the extracellular matrix (Noggin and BMP3) was

evaluated using semi-quantitative scoring system: no staining (0), low staining

(1), intermediate staining (2), and strong staining (3). The results were

evaluated by 2 independent investigators and averaged.

ii) Detection of Noggin and BMP3 by Western Blot : Western blot analysis for

Noggin (abcam 16054) and BMP3 (abcam71500) was carried out in

osteoblasts alone and in co-culture with dura mater cells (See Appendix

pg.121). β-actin was used as a positive control. Results were analyzed by

densitometry and expressed as protein density29

.

37

Aim IV) To investigate the role of dura mater paracrine signalling in the

pathophysiology of craniosynostosis in humans.

In order to search for potential growth factors responsible for the dura mater paracrine

signalling, dura mater cells were grown for 10 days and conditioned medium was collected at

different time points to determine the expression of FGF2 and TGF-β, both described in the

literature as the primary growth factors secreted by dura mater that may influence osteoblast

behavior.

i) Detection of FGF2 and TGF-β by Western blot : Conditioned Medium from

dura mater cells culture was collected at days 3,5 7 and 10 after subculture.

Western blot analysis was carried out for FGF2 (ab57059) and TGF-β (ab27969)

expression (See Appendix pg.121). β-actin was used as a positive control. Results

were analyzed by densitometry and expressed as protein density29

.

Aim V) To assess the effects of exogenous administration of FGF2 on osteoblast

function in vitro.

In Aim IV we examined the expression of endogenous FGF2 in dura-mater cells. The

aim in these experiments was to evaluate osteoblast behavior under stimulation by exogenous

human recombinant FGF2. Osteoblasts obtained from regions of fused suture, patent suture

and adjacent non-sutural bone were cultured alone or with increasing doses of human

recombinant FGF2. In order to evaluate osteoblast capacity to respond to stimulation,

proliferation and differentation rates were assessed.

i) Human Recombinant FGF2 Stimulation: Cells were plated onto 6 and 24-well

plates at a density of 50000 and 5000 cells/well respectively. After overnight

attachment, cells were treated with osteogenic differentiation media (αMEM

38

containing 10-7

M dexamethasone, supplemented with 15% FBS, ascorbic acid

50µg/1ml of medium and 1% β-glycophosphate) supplemented with human

recombinant FGF2 (5, 10, 50 and 100ng/ml)58

or only osteogenic media as a

control. Medium was changed every 2-3 days. FGF2 was added at each medium

change. MTT and Alkaline Phosphatase were performed at days 1, 3, 5 and 7.

Alkaline Phosphatase staining was performed at 1 week to assess early osteogenic

differentiation.

Statistical Analyses

Statistical analyses were performed using Graph Pad Prism and data are expressed as

mean ± standard deviation (SD). One-way analysis of variance (ANOVA) was applied for

comparison of sample groups in the MTT, Alkaline Phosphatase and Mineralization Assays,

Western Blot density, Immunohistochemistry density and qRT-PCR. Differences in values

between groups were evaluated using Tukey’s test. Two-way analysis of variance (ANOVA)

with repeated measurements was applied when more variables were evaluated for all three

groups. Significance was established at p < 0.05.

Research Ethics Board: This research has been approved by the Research Ethical Board at

The Hospital for Sick Children (1000013036). Consent was obtained from parents in all

cases.

39

Results

40

Demographics

Forty-five patients were enrolled in the study from July of 2008 to September of

2010. Twenty-eight were male and seventeen female. Mean age of the non-syndromic

patients was 7.5 ± 2.5 months (range from 3 months old to 13 months old). Forty patients had

single suture non-syndromic craniosynostosis. Samples were obtained from five syndromic

patients (3 Apert’s Syndrome, 1 Crouzon Syndrome and 1 patient with chromosome 7p

deletion). The sagittal suture was the most frequently involved followed by the metopic

suture (Table 2). All surgeries were performed uneventfully and there were no mortalities or

significant morbidities.

Cases n=45 Age

(months)

Male Female Sagittal Coronal Metopic Lambdoid

Syndromic

n=5

12.6±5.2 2 3 1 3 1 -

Non-

syndromic

n=40

7.5±2.5 26 14 19 9 11 1

Table 2: Demographics of patients with craniosynostosis enrolled in the study.

Histology

Histology was performed in tissue samples using hematoxylin-and-eosin staining to

confirm the tissue architecture of the cranial site from where the samples were harvested.

Histological evaluation of the patent, fused and adjacent bone confirmed the clinical

observations of presence or absence of a suture. Hematoxylin- and eosin-stained sections of

the control bone showed mature lamellar bone and hematopoietic marrow. Sections of patent

sutures showed fibrous connective tissue flanked on both sides by calvarial bony plates.

41

Sections of fused sutures showed an absence of the fibrous connective tissue zone, which

was replaced by lamellar bone. Some sites showed remnants of fibrous tissue in areas of

partial osseous obliteration and areas of bone remodeling (Figure 5).

Control Patent Suture Fused Suture

DAPI

H&E x100 magnification

H&E x50 magnification

LB

HM FT

FT

B

B

LB

HM

Figure 5: Histological sections of control bone, patent and fused suture bone with H&E

stain. The control bone showed lamellar bone (LB) and hematopoietic marrow (HM). The

patent suture showed fibrous connective tissue (FT) flanked on both sides by calvarial bony

plates (not seen) representing the normal patent suture. The fused suture showed the absence

of the fibrous connective tissue zone, replaced by bone (B).

42

Collagen I Expression

Collagen I is secreted by mature bone in the extracellular matrix. Collagen I

expression in tissue samples from non-sutural adjacent bone, patent and fused sutures was

assessed by IHC. Staining for Collagen I was positive for all three sites. The control bone

showed areas of mature bone. The patent suture showed an intense staining at the fibrous

connective tissue representing the normal patent suture. Stained cross-section of fused suture

showed intense collagen I staining, with areas of mature bone and remodeling (Figure 6).

Negative Control

100x

Fused Suture

Control Bone

Patent Suture

Figure 6: Immunohistochemistry staining for Collagen I. Stained sections show

positive expression of collagen I in fused bone, patent bone and adjacent non-sutural bone,

demonstrating formation of extracellular bone matrix (Magnification 100x).

43

Aim I) To develop a reliable osteoblast cell culture from calvarial of infants

undergoing surgery for craniosynostosis repair.

After culture of bone samples and periosteum, osteoblast growth was evaluated daily

by phase contrast microscopy up to confluence (usually 5 to 7 days after culture). Bone

samples and periosteum demonstrated the same potential for osteoblast retrival (4.5 x 106

versus 4.2 x 106) after 5 days. As such, osteoblasts from bone samples were preferred for all

experiments due to the in vivo relation between dura mater and calvarial bones.

Developing a valid bone cell culture model from calvaria of infants with craniosynostosis

In order to demonstrate the presence of osteoblasts in our cell culture model, we

assessed cellular proliferation, Runx2, alkaline phosphatase, collagen I and osteocalcin

expression and mineralization. To further confirm our findings of in vitro bone formation,

ultrastructural analysis of the samples was performed, demonstrating features of in vitro

osteogenesis, including mineral depositions and collagen fibrils in the extracellular matrix. A

ring pattern for electron diffraction studies was characteristic of normal hydroxyapatite

(Figure 7).

44

45

Figure 7: Evidences of osteoblasts in our cell culture model and in vitro bone

formation: Patent suture cells were able to proliferate in vitro (A) and differentiate in

osteoblasts as shown by the expression of differentiation markers such as alkaline

phosphatase, Runx2, collagen I and osteocalcin (B). Mineralization and Bone Formation was

achieved after 28 days in osteogenic conditions. Ultrastructural analysis of the samples was

performed, demonstrating features of in vitro osteogenesis, including mineral depositions

and collagen fibrils in the extracellular matrix. A ring pattern for electron diffraction studies

was characteristic of normal hydroxyapatite (C).

Effect of medium composition on cellular proliferation

Fused suture osteoblasts, independent of the experimental conditions or time-point,

demonstrated higher growth rates than the patent and control sutures. The addition of

ascorbic acid and β-glycerophosphate to the osteogenic medium resulted in significantly

higher growth rates for the fused suture osteoblasts when compared αMEM (1% FBS) and

αMEM (15% FBS) on day 3 (p<0.05) (Figure 8A). There was no significant differences in

proliferation rates at days 5,7 and 10 independent of medium composition.

46

Figure 8: MTT assays of human cranial suture-derived osteoblasts. All cultures in

1%, 10%, 15% FBS or 15% FBS + ascorbic acid (50µg)/1ml of medium. Values are

mean±SD; n=7, *p<0.05.

Differences in FBS concentration did not significantly affect the growth of cranial

suture-derived osteoblasts from fused suture, patent suture and adjacent bone. Moreover,

there was no significant difference in alkaline phosphatase activity independent of the

medium composition and therefore we chose for the model system the medium consisting of

αMEM containing 10-7

M dexamethasone, supplemented with 15% FBS for 7 days and then

supplemented with ascorbic acid (50µg)/1ml of medium and 1% β-glycerophosphate for the

experiments.

47

Aim II) To assess regional variations in osteoblast behavior with and without

dura mater cells in co-culture.

Osteoblast Proliferation

Cells were grown until confluence and then were subcultured (Figure 9). Proliferation

rates of control bone, patent suture and fused suture osteoblasts were evaluated in triplicate

by MTT assay.

Figure 9: Cellular growth prior and post-subculture. A) Cranial suture-derived

culture prior to subculture three days after harvesting. Osteoblasts from fused sutures

achieved confluence earlier than those from patent sutures or non-sutural adjacent bone. B)

Cranial suture-derived subculture at day 3.

48

The osteoblasts from the fused sutures exhibited a significant (p < 0.01) increased rate

of growth, compared with those derived from the control and patent suture at days 5 and 7

(Figure 10).

All Cases

Day 1 Day 3 Day 5 Day 70

5

10

15Control Bone

Patent Suture

Fused Suture

* *

* p<0.02n=33 per group

*

OD

/1000 c

ells

Figure 10: Osteoblast proliferation rates (MTT): Fused suture cells showed a

significantly higher rate of proliferation at time-point 3 (p<0.05) when compared to control

bone and time-points 5 (p<0.001) and 7 (p<0.001) when compared with control and patent

suture osteoblasts at the same time-points.

Proliferation rates were significantly lower for syndromic cases in all three groups

when compared with non-syndromic patients (Figure 11). For this reason we decided to

exclude samples from syndromic patients for the subsequent experiments in order to not

confound the findings.

49

Non-Syndromic Cases

Day 1 Day 3 Day 5 Day 70

5

10

15Control Bone

Patent Suture

Fused Suture

* *

* p<0.02n=28 per group

A)

*

OD

/1000 c

ells

Syndromic Cases

Day 1 Day 3 Day 5 Day 70

5

10

15Control Bone

Patent Suture

Fused Suture

p=0.8n=5 per group

B)

OD

/1000 c

ells

Figure 11: Proliferation Rates for Syndromic Patients: Proliferation rates were

significantly higher for non-syndromic patients (A) when compared with syndromic patients

(B). Also the difference in proliferation rates between sites was not significant (p=0.8) in the

syndromic patients.

50

Co-Culture of Osteoblasts and Dura Mater Cells: Proliferation Rates

Osteoblasts from control bone, fused suture and patent suture co-cultured with dura

mater were compared with their counterparts without dura mater cells (Figure 12). Adding

dural cells to the osteoblast culture does not change significantly the proliferation rates for

fused suture and control bone. However, patent suture osteoblasts in co-culture with dura

demonstrated a significantly greater (p=0.001) proliferation rate at day 7 when compared

with patent suture osteoblasts without dura.

Patent Suture

Day 1 Day 3 Day 5 Day 70

5

10

15

Subculture with Dura

Subculture without Dura

p=0.001 (two-way ANOVA)n= 28 each group without duran= 6 each group with dura

B)

OD

/1000 c

ells

*

Control Bone

Day 1 Day 3 Day 5 Day 70

5

10

15Subculture without Dura

Subculture with Dura

p=0.27 (two-way ANOVA)n= 28 each group without duran= 6 each group with dura

A)

OD

/1000 c

ells

51

Fused Suture

Day 1 Day 3 Day 5 Day 70

5

10

15Subculture without Dura

Subculture with Dura

C)

p=0.36 (two-way ANOVA)n= 28 each group without duran= 6 each group with dura

OD

/1000 c

ells

Figure 12: Osteoblast proliferation rates in co-culture with dura mater: Co-culture

with dura mater cells does not change significantly (p>0.05) the proliferation rates for fused

suture and control bone. Patent suture osteoblasts in co-culture with dura mater

demonstrated a significantly greater (p=0.001) proliferation rate at day 7 when compared

with patent suture osteoblasts without dura mater.

Runx2 Expression

Runx2 is a transcriptional factor that controls osteoblast differentiation through the

regulation of osteogenic proteins, including osteocalcin and osteopontin.

Immunocytochemical analysis of Runx2 showed expression of this molecule in all three

groups, demonstrating that cells from adjacent bone, patent suture and fused suture are able

to differentiate in osteoblasts (Figure 13).

52

Negative Control Control Bone Patent Suture Fused Suture

400x

400x

Dapi

Runx-2

Figure 13: Expression of Runx2 demonstrated by Immunocytochemistry (nuclear

staining). Control bone, patent suture and fused suture cells were similarly positive for

Runx2, indicating that cells from all three groups were able to differentiate.

However, Western blot analysis of Runx2 showed an up regulation of this molecule

in the control group and fused suture group with and without dura mater when compared

with patent suture. This difference was significantly (p<0.05) greater when osteoblasts

53

derived from fused suture were compared without and with co-culture with dura mater cells

(Figure 14).

57kDaRunx-2

SaOs2 C C+D P P+D F F+D

ß-actin 43kDa

Runx2 Expression

Contr

ol Bone

Contr

ol Bone

+ Dura

Mat

er

Pat

ent S

uture

Pat

ent S

uture

+ D

ura M

ater

Fused S

uture

Fused S

uture

+ D

ura M

ater

0

50

100

150

% R

ela

tive C

on

tro

l

p=0.024

n=6 per group

*

SaOs2: Human osteosarcoma cell line

C: Control Bone

C+D: Control Bone + Dural cells

P: Patent Suture

P+D: Patent Suture + Dural cells

F: Fused Suture

F+D: Fused Suture + Dural cells

Figure 14: Expression of Runx2 by Western Blot. Runx2 showed up regulation in

the control and fused suture group when compared with patent suture. Expression of Runx2

was significantly (p=0.024) increased when osteoblasts from the fused suture were combined

in co-culture with dura mater cells compared with monoculture.

54

Alkaline Phosphatase Activity

Alkaline Phosphatase activity increased steadily for the first 3 days. Rapid

progression was noticed at day 5 and 7 for all three groups. This finding, taken together with

other markers, demonstrated that cells grown in culture differentiate into osteoblasts. The

increased levels of AP are consistent with the increase in cell numbers. Osteoblasts from the

fused sutures showed higher levels of AP expression compared to control and patent suture

(Figure 15).

Alkaline Phosphatase

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Control Bone

Patent Suture

Fused Suture

p<0.05n=28 per group

*

*****

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

Figure 15: Alkaline Phosphatase Assay: Results of AP measurement, taken together

with other markers of differentiation, suggest that osteoblasts are present in culture. The

55

increased levels of AP are consistent with the increased number of cells at days 5 and 7.

(*p<0.05 control versus patent suture;

** p<0.05 fused suture versus patent suture and

***p<0.05 fused suture versus control and patent bone). Final AP concentration is expressed

as mmol of pNP per hour per µg of protein.

Figure 16: Alkaline Phosphatase Activity (Fast Blue BB salt - dark blue staining)

in cell cultures: Osteoblasts derived from the fused suture demonstrated multiple layers and

an increased number of cells at day 7.

56

qRT-PCR

AP Osteocalcin0

2

4

6Control Bone

Patent Suture

Fused Suture

Exp

ressio

n o

f m

RN

A

p=0.66 (AP)

p=0.11 (Osteocalcin)

n=6 per group

Figure 17: Analysis by qRT-PCR of alkaline phosphatase and osteocalcin gene

expression: Expression of alkaline phosphatase and osteocalcin mRNA are not significantly

different between osteoblasts from various regions of the infant calvarium. mRNA levels

were measured by qRT-PCR and normalized to HPRT mRNA.

57

Co-Culture of Osteoblasts and Dura Mater Cells: Alkaline Phosphatase

AP was expressed by control bone, patent suture and fused suture osteoblasts when

combined with dura mater cells and expression increased with time (Figure 18). Adding dural

cells to the control bone and patent suture osteoblasts enhanced their differentiation when

compared with their culture without dura mater.

Control Bone

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Culture with Dura

Culture without Dura

p=0.0005 (two-way ANOVA)n= 28 each group without duran= 6 each group with dura

A)

*

*

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

*

Patent Suture

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Culture with Dura

Culture without Dura

p=0.002 (two-way ANOVA)n= 28 each group without duran= 6 each group with dura

B)

*

*

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

58

Fused Suture

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Culture with Dura

Culture without Dura

p<0.04 (two-way ANOVA)n= 28 each group without duran= 6 each group with dura

C)

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

Figure 18: AP in osteoblasts co-culture with dura mater cells: Osteoblasts co-

cultured with dura mater cells demonstrate increased AP expression over time. Co-culture

with dura mater cells significantly enhanced control bone (A) differentiation at days 3,5 and

7 and patent suture osteoblasts (B) differentiation at days 3 and 7 compared with osteoblasts

without dura mater and reduced differentiation of fused suture osteoblasts (C). Final AP

concentration is expressed as mmol of pNP per hour per µg of protein.

Mineralization

In cultures containing β-glycerophosphate (10mM) and ascorbic acid (50µg/ml), the

secretion of extracellular matrix leads to progressive mineralization and eventual bone

nodule formation by day 14 (Figure 20). Osteoblasts from all three sites were capable of

forming bone nodules in vitro, as evidenced by extensive Alizarin Red Staining. However, at

28 days osteoblasts from fused sutures demonstrated significantly (p<0.05) greater

mineralization compared to osteoblasts from patent sutures (Figure 19). Bone nodules

were detected earlier in osteoblasts from fused sutures (11±3 days) compared with control

(13±4 days) and patent sutures (16±4 days) osteoblasts (Figure 20). Formation of bone

59

nodules was significantly (p<0.0001) more robust for osteoblasts from fused suture (65±9

mm2 – mean area ± SD) when compared with control (34±4 mm

2) and patent suture

osteoblast (22±2 mm2).

Mineralization

Day 21 Day 280

1

2

3

4

5Control Bone

Patent Suture

Fused Suture

OD

/1000 c

ells

*

*p<0.05

n=21

**

Figure 19: Bone Nodule Formation (Alizarin Red Assay). Osteoblasts from control

bone demonstrated significantly (*p<0.01) greater mineralization when compared with

patent suture at day 28. Osteoblasts from the fused suture demonstrated significantly

(**p<0.05) greater mineralization when compared with control bone and patent suture

osteoblasts at day 28 post subculture.

60

Figure 20: Bone Nodule Formation at days 14 and 18 (Alizarin Red S Staining):

Formation of bone nodules in cultures derived from fused sutures was detected earlier (11±3

days) compared with that of control (13±4 days) and patent sutures (16±4 days). Arrow

shows bone nodule formation.

61

Control – 28 days Patent Suture – 28 days Fused Suture– 28 days

Control – 21 days Patent Suture – 21days Fused Suture– 21days

100x 100x 100x

100x 100x 100x

Figure 21: Mineralization at days 21 and 28 (Alizarin Red Staining). Formation of

bone nodules in cultures derived from fused sutures was significantly (p<0.001) more robust,

compared with that of normal sutures and control areas at day 28. Arrow shows bone nodule

formation.

62

Co-Culture of Osteoblasts and Dura Mater Cells: Mineralization

Mineralization rates in osteoblasts co-cultured with dura mater cells were not

different between groups. The addition of dural cells to culture resulted in qualitative

differences in all three groups with reduced nodule formation and mineralization (Figure 22).

Figure 22: Bone nodule formation in osteogenic cultures with and without dural co-

culture at day 28 (Alizarin Red S Staining). Mineralization in osteoblasts co-cultured with

dura mater cells was not different between groups.

63

Transmission Electron Microscopy

Electron microscopy of cell cultures of control bone, patent suture and fused sutures

revealed ultrastructural features of in vitro osteogenesis, including mineral depositions

and collagen fibrils in the extracellular matrix (Figure 23).

Figure 23: Transmission Electron Microscopy of Bone Nodule Formation: Sections

showing ultrastructural features of in vitro osteogenesis, including collagen fibrils,

mineral and osteoblasts. Cell cultures from control bone, patent suture and fused suture

were able to form bone.

Aim III) To investigate the role of anti-osteogenic signalling on human

osteoblasts in vitro with and without dura mater cells in co-culture.

64

Expression of anti-osteogenic signalling molecules

The anti-osteogenic molecules Noggin and BMP3 were analyzed in tissue samples

from control bone, patent suture and fused suture by IHC. The expression of Noggin and

BMP3 in the osteoblasts from patent suture was greater than expression levels in control

bone and fusing suture (Figure 24).

Figure 24: Immunohistochemistry analysis of Noggin in tissue samples. Noggin

expression (expressed as a brown pigment in the extracellular matrix) was found to be

greater in the osteoblasts from patent suture when compared to control bone or fused suture.

65

Noggin Expression

Contr

ol Bone

Pat

ent S

uture

Fused S

uture

0

1

2

3

4

Sem

i-q

uan

tita

tive s

co

re p<0.0001n=6

* *

Figure 25: Quantification of Immunohistochemical analysis for Noggin in the

tissue samples. Expression of Noggin was significantly (p<0.0001) greater in osteoblasts

from patent sutures when compared with the control bone and fused suture. The staining

intensity was evaluated using semi-quantitative scoring system: no staining (0), low staining

(1), intermediate staining (2), and strong staining (3). The results were evaluated by 2

independent investigators and averaged.

.

[Type a quote from the document or the summary of an interesting point. You can position the

text box anywhere in the document. Use the Text Box Tools tab to change the formatting of the

pull quote text box.]

66

Negative Control Patent Suture

Control Bone Fused Suture

200x 200x

200x 200x

Figure 26: Immunohistochemistry analysis of BMP3 in tissue samples. BMP3

expression was greater in the extracellular matrix of patent sutures when compared to fused

sutures.

67

BMP3 Expression

Contr

ol Bone

Pat

ent S

uture

Fused S

uture

0

1

2

3

4S

em

i-q

uan

tita

tive s

co

re

p<0.0001

n=6 per group

**

Figure 27: Quantification of Immunohistochemical analysis for BMP3 in the

tissue samples. BMP3 expression was significantly (p<0.0001) higher in patent sutures when

compared with control and fused suture. The staining intensity was evaluated using semi-

quantitative scoring system.

In order to further confirm these findings and to evaluate the influence of dura mater

cells in co-culture on Noggin and BMP3 expression, Western Blot analysis was performed in

control bone, patent suture and fused suture samples after cell culture. BMP3 expression

(protein expression) was significantly up regulated in the patent suture when co-cultured with

dura mater cells but not in control and fused suture osteoblasts (Figure 28A). Noggin

expression was enhanced in all three groups when in co-culture with dura mater cells (Figure

68

28B). The significant differences previously observed in the immunohistochemical analysis

of calvarial tissue (figures 25 and 27) were not observed in the WB analysis of the three sites

without dura mater.

BMP-3 Expression

Contr

ol Bone

Contr

ol Bone

+ Dura

Mate

r

Paten

t Sutu

re

Paten

t Sutu

re +

Dura

Mat

er

Fused S

uture

Fused S

uture

+ D

ura M

ater

0

50

100

150

200

% R

ela

tive

Co

ntr

ol

p<0.0001

n=6

Mean SD

112.01 16.8

*

A) BMP-3 53kDa

C C+D P P+D F F+D

ß-actin 43kDa

SaOs2

SaOs2: Human osteosarcoma cell line

C: Control Bone

C+D: Control Bone + Dural cells

P: Patent Suture

P+D: Patent Suture + Dural cells

F: Fused Suture

F+D: Fused Suture + Dural cells

69

B)C C+D P P+D F F+D

ß-actin 43kDa

SaOs2

SaOs2: Human osteosarcoma cell line

C: Control Bone

C+D: Control Bone + Dural cells

P: Patent Suture

P+D: Patent Suture + Dural cells

F: Fused Suture

F+D: Fused Suture + Dural cells

Noggin 26kDa

Noggin Expression

Contr

ol Bone

Contr

ol Bone

+ Dura

Mat

er

Paten

t Sutu

re

Paten

t Sutu

re +

Dura

Mat

er

Fused S

uture

Fused S

uture

+ D

ura M

ater

0

50

100

150

200

% R

ela

tive C

on

tro

l

**

*

p<0.0001

n=6

Mean SD

116.6 33.9

Figure 28: Western Blot analysis of BMP3 and Noggin after 10 days of co-culture

in osteogenic medium. WB analysis of the osteogenic medium showed an increased in

Noggin and BMP3 protein expression when control bone, patent suture and fused suture

were compared. A) BMP3 expression was significantly (p<0.0001) up regulated only in

patent suture osteoblasts co-cultured with dura mater cells. B) Noggin expression showed

significantly (p<0.0001) increased expression in all three groups when osteoblasts were co-

cultured with dura mater cells compared with monocultures.

Aim IV) To investigate the role of dura mater paracrine signalling in the

pathophysiology of craniosynostosis in humans.

70

Dura Mater Expression of FGF2 and TGF-β1

In order to understand the influence of dura mater cells on osteoblast behavior, we

then searched for candidate molecules that may be responsible for dura mater paracrine

signalling modifying osteoblast behavior. We chose FGF2 and TGF-β1 to examine. Western

Blot of the conditioned medium obtained at days 3,5,7 and 10 from dural cultures

demonstrate the presence of FGF2 and TGF-β1. TGF-β1 levels remained consistent

throughout the study period whereas FGF2 levels declined after day 3 (Figure 29).

FGF2

TGF-ß1 44kDa

47.5kDa

D3 D5 D7 D10 D3 D5 D7 D10

Dura from 3 year old patient Dura from 11 years old patient

FGF2

Day 3 Day 5 Day 7 Day 100

10

20

30

40

FG

F-2

Den

sit

y

TGF- B1

Day 3 Day 5 Day 7 Day 100

10

20

30

40

TG

F-

B1 D

en

sit

y

Mean SD

Day 3: 14.8 2.82

Day 5: 12.8 0.28

Day 7:7.35 5.3

Day 10: 10.1 0.424

Mean SD

Day 3: 22.05 0.05

Day 5: 11.6 3.8

Day 7:10.85 4.150

Day 10: 5.5 0.5

Actin 43KDa

Figure 29: Expression of pro-osteogenic molecules in the conditioned medium

from dura mater cells. TGF-ß1 did not show significant variation between days. FGF2 had a

two-fold higher expression at day 3 when compared with the subsequent days. Western blot

analyses by densitometry.

71

Aim V) To assess the effects of exogenous administration of FGF2 on osteoblast

function in vitro.

Recombinant Human FGF2 Stimulation

FGF2 was expressed in higher concentration in the medium from dura mater cells

when compared with TGF-β1. We then decided to add exogenous FGF2 to control bone,

patent suture and fused suture osteoblast cell cultures to evaluate the ability of osteoblasts to

respond to this stimulation. The addition of FGF2 to culture medium enhanced cellular

proliferation at all 4 concentrations for the three groups in a dose dependent fashion.

Osteoblasts from fused suture were more sensitive to lower doses of stimulation (5ng/ml –

p<0.05) at day 7 than the other two groups.

72

Proliferation

Control Bone

5ng/ml 10ng/ml 50ng/ml 100ng/mlNo stimulation

200x

Patent Bone

Fused Bone

200x

200x

Figure 30: Proliferation of osteoblasts stimulated with FGF2. At 3 days of

incubation, an increase in proliferation was observed for all three groups independent of the

FGF2 concentration compared with non-stimulated osteoblasts shown in the photos of cell

culture plates.

73

MTT-Control Bone

Day 1 Day 3 Day 5 Day 70

5

10

15Control Bone

5 ng/ml

10ng/ml

50ng/ml

100ng/ml

A)

** *

p<0.0001(two-way ANOVA)n=4

Co-culture with Dura Mater

* *

OD

/1000 c

ells

MTT-Patent Suture

Day 1 Day 3 Day 5 Day 70

5

10

15Patent Suture

5ng/ml

10ng/ml

50ng/ml

100ng/ml

B)

*

*

p<0.0001(two-way ANOVA)n=4

Co-culture with Dura Mater

OD

/1000 c

ells

MTT-Fused Suture

Day 1 Day 3 Day 5 Day 70

5

10

15Fused Suture

5ng/ml

10ng/ml

50ng/ml

100ng/ml

C)

**

**

***

*

p<0.0001(two-way ANOVA)n=4

Co-culture with Dura Mater

OD

/1000 c

ells

Figure 31: Proliferation Rates after Stimulation with FGF2 (MTT): Stimulation

with recombinant human FGF2 significantly (p<0.0001) increases the rate of proliferation

74

for all three groups (in a dose dependent manner) compared with osteoblasts without

stimulation. A) In the control bone group proliferation is significantly greater with 50ng/ml

(p<0.05) at days 5 and 7; 100ng/ml (p<0.01) at days 5 and in co-culture with dura mater

cells at days 5 (p<0.001) and 7 (p<0.01). B) In the patent suture group proliferation is

significantly greater in co-culture with dura mater cells at day 5 (p<0.05) and 7 (p<0.001)

C) In the fused suture group proliferation is significantly greater with 5ng/ml at day

7(p<0.05); 50 ng/ml at days 3(p<0.05),5 (p<0.01)and 7(p<0.05) and with 100ng/ml at days

5 and 7(p<0.0) and in co-culture with dura mater cells at days 5 and 7 (p<0.001).

75

Differentiation

Osteoblasts cultured with exogenous FGF2 demonstrated an increase in alkaline

phosphatase expression in all three groups at days 3,5 and 7. However the differences

between groups were not significant.

Control Bone

Patent Bone

No stimulation 5ng/ml 10ng/ml 50ng/ml 100ng/ml

Fused Bone

200x

200x

200x

Figure 32: AP expression after stimulation with FGF2. At 3 days of incubation,

increased alkaline phosphatase staining is seen in all three groups treated with FGF2

independent of the FGF2 concentration.

76

Alkaline PhosphataseControl Bone

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Control Bone

5 ng/ml

10ng/ml

50ng/ml

100ng/ml

A)

Co-culture with Dura Mater

p<0.01(two-way ANOVA)n=4

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

Alkaline PhosphatasePatent Suture

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Patent Suture

5ng/ml

10ng/ml

50ng/ml

100ng/ml

Co-culture with Dura Mater

B)

p=0.1(two-way ANOVA)n=4

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

Alkaline PhosphataseFused Suture

Day 1 Day 3 Day 5 Day 70.0

0.2

0.4

0.6

0.8Fused Suture

5ng/ml

10ng/ml

50ng/ml

100ng/ml

Co-culture with Dura Mater

C)

p=0.27(two-way ANOVA)n=4

mm

ol o

f p

NP

/h/µ

g o

f p

rote

in

Figure 33: APA after stimulation with FGF2: Osteoblasts cultured with exogenous

FGF2 demonstrated an increase in alkaline phosphatase in all three groups at days 3,5 and

7 compared with day 1. Final AP concentration is expressed as mmol of pNP per hour per

µg of protein.

77

Discussion

78

The work developed herein validates our hypotheses that regional variations in

osteoblast behavior in infants with craniosynostosis do exist.

We first developed a human model of osteoblast cell culture by obtaining calvarial

bone samples from patent sutures, fused sutures and adjacent bone from infants with

craniosynostosis. Histologically, we confirmed regional variations between suture sites and

adjacent bone. The ability to obtain osteoblasts capable of forming bone was demonstrated

by cell proliferation, presence of Runx2 and alkaline phosphatase, collagen I expression,

osteocalcin production, mineralization and transmission electron microscopy. Cells derived

from fused suture sites clearly demonstrated greater osteogenic potential with higher rates of

proliferation and differentiation when compared with cells from control bone and patent

suture. We also searched for factors in the surrounding enviroment that may contribute to the

imbalance between pro- and anti-osteogenic signalling. Noggin and BMP3, both anti-

osteogenic molecules were significantly greater expressed by patent sutures than by fused

sutures and control bone. Runx2 expression was upregulated in the fused suture osteoblasts

combined with dura mater demonstrating that osteoblasts from fused sutures may be more

differentiated that their counterparts. Dura mater cells were shown to mainly influence

osteoblast differentiation and mineralization, downregulating those parameters in fused

suture cells and enhancing proliferation and differentiation in patent suture cells. On the other

hand, exogenous FGF2 appeared to regulate proliferation in a dose dependent fashion, with

osteogenic cells from fused sutures being more sensitive to lower doses of FGF2. Taken

together, our findings demonstrated interactions between calvarial osteogenic cells and dura

mater and also validate our hypothesis that regional variations exist in the calvarial bones of

infants with craniosynostosis.

79

Development of a valid model of osteogenic culture derived from cranial suture sites

Over the past several years, many studies have used animal models to investigate the

biology underlying premature suture fusion28, 29,48

. Specifically, the sagittal suture in a

murine model has been utilized as a model of normal patent human suture biology and the

fusing posterior frontal suture as a model of premature fusion. A rabbit cranial suture model

from a colony of congenitally fused rabbits29

has also been studied in an attempt to

understand the mechanisms underlying syndromic craniosynostosis.

Despite extensive research based on animal models, there is little literature on the use

of human cranial tissue samples to investigate the biomolecular mechanisms involved in

craniosynostosis. Recent studies have been limited by the number of samples obtained from

non-syndromic patients60

. We were able to develop a valid human model of osteogenic cell

culture derived from cranial suture sites, and obtain a large number of samples representing

the wide spectrum of this condition. Our study is the largest one compared to those in the

current literature. We obtained samples from 45 patients with syndromic and non-syndromic

craniosynostosis representative of the epidemiology of craniosynostosis with regards to with

age, suture affected and sex. We also obtained samples from different regions of the cranial

vault in all patients, representing the fused suture, the patent suture and adjacent non-sutural

bone.

In order to validate our bone cell culture model and demonstrate the presence of

osteoblasts in cell culture, we assessed cellular proliferation, Runx2 expression, alkaline

phosphatase activity, collagen I and osteocalcin expression and mineralization. Cells isolated

from patent suture, fused suture and adjacent bone demonstrated characteristics of the

osteoblast phenotype with rapid proliferation, presence of alkaline phosphatase activity,

80

expression of Runx2, and in vitro bone formation. The earliest stages of osteoblast

differentiation can be detected through the measurement of alkaline phosphatase and Runx2.

Although alkaline phosphatase can be expressed by other cell types such as fibroblasts,

Runx2 is a specific marker of osteoblast differentiation and was expressed by all cells of our

culture model, confirming that cells were able to differentiate in osteoblasts. In cultures

containing ascorbic acid and β-glycerophosphate, the secretion of extracellular matrix by

differentiated osteoblasts was followed by progressive mineralization and bone nodule

formation. To further confirm our findings of in vitro bone formation, ultrastructural analysis

of the samples was performed, demonstrating features of in vitro osteogenesis, including

mineral depositions and collagen fibrils in the extracellular matrix. A ring pattern for electron

diffraction studies was characteristic of normal hydroxyapatite.

Taken all together, the above findings allow us to conclude that the human bone cell

culture model described herein is valid and reliable.

We determined the most favorable conditions for osteogenic culture by varying

medium composition based on animal models previously described. The development of

osteoblast function and morphology in vivo strongly depends on the composition of the

culture medium, particularly essential factors that influence various stages of proliferation

and differentiation during the process of osteogenesis68

. Early studies of osteoblastic cell

lines have demonstrated that osteoblasts undergo a temporal sequence of differentiation,

featuring active cell proliferation, expression of osteoblastic markers, synthesis and

deposition of a collagenous extracellular matrix, as well as mineralization, in order to

develop normal osteoblast phenotype and function68

.

81

Osteoblast proliferation rates were higher in the fused sutures, independent of

experimental culture conditions. This observation is consistent with a study that showed bone

formation activity at the fused suture site is increased in non-syndromic craniosynostosis,

leading to premature ossification69

. The same study in humans showed that at all ages (3 – 18

months), the proliferation of osteoblasts obtained from fused suture locations was similar to

that from patent sutures69

and both AP activity and osteocalcin production by osteoblast cells

originating from fused sutures were significantly higher than that shown in cells derived from

normal sutures, upon stimulation with vitamin D69

. The difference in findings between the

previous work69

and our study may be attributed to different growth conditions used in the

two investigations. Studies by Coelho et al. have shown that the presence of dexamethasone

in combination with ascorbic acid and β-glycerophosphate resulted in significantly increased

cell growth and AP activity68

.

The effects of ascorbic acid have been well defined by previous studies, which have

emphasized its critical role in collagen biosynthesis, particularly in the production of the

collagenous bone extracellular matrix68

due to its role as a cofactor for proline hydroxylase

and lysine hydroxylase, which are involved in the hydroxylation of collagen. A Na+-

dependent transporter specific for ascorbic acid is present in the plasma membrane of

osteoblasts and is essential for maintenance of intracellular ascorbate concentrations70

. One

pathway that mediates ascorbate’s effect on osteoblast growth and differentiation is the

collagen receptors or integrins70

. In addition, osteoblast differentiation as mediated by 1,25

(OH)2 Vitamin D3, retinoic acid, and bone morphogenic proteins is also affected by the

concentration of ascorbic acid. Thus, ascorbic acid is essential for normal bone formation70

.

Our study showed that during a 10 day time interval, regional variations in osteoblast

82

proliferation rates are evident in infants with craniosynostosis. Fused suture cell cultures,

supplemented with 15% FBS + ascorbic acid + α-glycerophosphate demonstrated

significantly higher growth rates, in comparison to both αMEM (1% FBS) and αMEM

(15%FBS) on day 3. Studies by Prader et al., which used similar ascorbic acid

supplementation, demonstrated that an optimal amount of ascorbic acid (50µg/mL)

stimulates osteoblastic differentiation, as well as dose-dependent collagen synthesis, leading

to higher AP values62

. This relationship between cell differentiation rates and ascorbic acid

was investigated by Franceschi in 1992, who proposed that the collagen matrix synthesized

by cells treated with ascorbic acid, provides a more favourable environment for tissue-

specific gene expression in mesenchymal-derived tissues like bone, resulting in higher

differentiation rates.

Cell culture models are a critical tool in cranial suture biology research. This study

has described the effects of culture medium composition on proliferation and differentiation

of human calvarial osteogenic cells. Furthermore, for human calvarial osteogenic cells,

αMEM with 10-7

M dexamethasone, supplemented with 15% FBS and ascorbic acid proved to

be the best culture medium, resulting in high growth and differentiation rates. These results

reveal the importance in defining the experimental conditions in prospective osteoblast cell

studies68

.

Assessment of regional variations in osteoblast behavior with and without dura mater

cells in co-culture.

In the current study, MTT and alkaline phosphatase activities, as indices of bone cell

proliferation and differentiation, respectively, were greater in osteogenic cultures from fused

sutures compared with patent sutures and adjacent non-sutural bone. Fused suture cells

83

proliferated faster and differentiated earlier then control and patent suture cells. Proliferation

rates were also higher in younger patients when compared with patients over 12 months of

age (seen in the syndromic group) which is in accordance with the literature71

. Hassler et al71

demonstrated that after surgical treatment of sagittal synostosis in children aged 6 months or

younger, reossification usually started 2 weeks postoperatively and was complete within 6

months. In children aged 7 to 12 months, reossification was prolonged and lasted for 12

months or longer and in children older than 12 months reossification was incomplete with

persistent pseudosutures, compromising surgical results. Patients with syndromic

craniosynostosis in our study were older (mean 12.6 months old) than patients with non-

syndromic craniosynostosis (7.5 months old). The delayed surgical procedure is usually

associated with other comorbidities found in syndromic patients. Cells retrieved from older

patients may not retain the same potential for proliferation and differentiation once they

reach maturity. Animal studies corroborate the influence of age on osteoblast proliferation44

.

It also may suggest that osteoblasts from younger patients are more susceptible to the dura

mater paracrine effect and the paracrine effects may last longer in vitro. The influence of age

in osteoblast behavior will be investigated in depth in further studies using our cell culture

model.

Osteoblasts from non-sutural bone, patent and fused sutures also showed a

significant difference in their alkaline phosphatase activity during the early period of culture;

alkaline phosphatase activity was significantly greater (p<0.05) during this period in the

fused suture cultures.

Formation of bone nodules in cultures derived from fused sutures was not only

detected earlier (11±3 days) compared with control (13±4 days) and patent sutures (16±4

84

days) osteoblasts, but also individual nodules in these cultures were two to three times larger

in diameter than those derived from control and patent sutures (p<0.0001). These results

suggest that osteoblasts derived from fused sutures exhibit an aggressive pattern of bone

formation, compared with their normal counterparts. TEM revealed ultrastructural features of

in vitro osteogenesis, including mineral depositions and dissected collagen fibrils in the

extracellular matrix.

In view of the ability of fused suture osteoblasts to aggressively form bone in vitro,

we examined the levels of Runx2 (also known as Cbfa-1). Runx2, a transcription factor that

is a marker of osteoblast differentiation, has been localized to the critical area of cranial

suture fusion. Runx2 is found in the osteogenic fronts and sutural mesenchyme and has

variable expressivity in patent versus fusing sutures in normal development29

. By

immunofluorescence we were able to show that Runx2 was expressed by adjacent bone,

patent suture and fusing suture, demonstrating that cells from all three sites have the ability to

differentiate. WB analyses demonstrated up-regulation of Runx2 expression at the fused

suture site when osteoblasts were combined with dura mater cells. Based on our findings,

Runx2 did not show significant differences when osteoblast from different sites were

evaluated, suggesting that it may not regulate suture patency and fusion itself, but it may

interact with other molecules or respond to dura mater paracrine signalling determining the

osteoblast differentiation in premature suture fusion.

The findings of accelerated in vitro bone formation parallel the clinical and

histological observations of partial or complete suture fusion suggesting important regional

variations in osteoblast behavior in infants with craniosynostosis.

85

The Role of Dura Mater

Over the last decade, numerous animal studies have suggested that the immature dura

mater is the critical factor governing successful calvarial ossification72, 73

. More recently,

studies have begun to identify the biomolecular and cellular differences between immature

and mature dura mater74

. Collectively, these studies suggest that the immature dura mater

supplies growth factors and/or osteoblast-like cells that participate in successful calvarial

reossification. Longaker67

et al., using a novel co-culture system, demonstrated, for the first

time, that osteoblast proliferation, gene expression, and bone nodule formation can be

directly modulated by paracrine signals from immature, non–suture-associated dural cells.

Furthermore, that study demonstrated that immature, non–suture-associated dura mater,

isolated or in co-culture, produced high levels of osteoinductive cytokines like TGF-β1 and

FGF2.

Although clinical observations and animal studies have demonstrated that young dura

mater (3-days-old mice) is both osteogenic and osteoconductive67

, to date no study has

clarified the interaction between human cranial sutures and human dura mater in

craniosynostosis. To better understand the contributions of dura mater and osteoblasts to

calvarial ossification, we co-cultured mature human dura mater with osteoblasts to examine

the effects of dural cells on the biologic activity of co-cultured osteoblasts. Our co-culture

model allowed us to investigate the in vitro interactions between human dura mater and the

calvarial bones. The dura mater cells were obtained from neurosurgical procedures. A

potential limitation of our study includes the age of neurosurgical patients undergoing

surgery for epilepsy. Although we attempted to age-match our patients, craniofacial patients

were younger at the time of the surgery (3-13 months old) compared with neurosurgical

86

patients (3 to 17 years old). We were also not able to select specific sites where dural samples

were harvested. Specifically we could not match the dural sites to the cranial suture sites and

samples from fused sutures could not be obtained. However, despite these shortcomings,

dural co-culture demonstrated that osteoblasts were influenced by the presence of dura in the

following ways: enhancing proliferation and differentiation for osteoblasts from patent

sutures; enhancing differentiation for osteoblasts from control bone and promoting the same

pattern of mineralization for all three groups.

Herein we demonstrated that osteogenic cultures from the fused suture and the control

bone co-cultured with dura mater showed no difference in proliferation rates compared with

osteogenic monocultures from the same areas cultured without dura. We speculate it may be

due to the age of patients from whom dura mater was harvested. It may influence

proliferation rates, with less mature dura mater from age-matched patients perhaps enhancing

proliferation rates for all three groups. However, co-cultured control bone and patent suture

osteogenic cultures expressed significantly (p<0.05) greater levels of alkaline phosphatase

when compared with osteogenic cultures without dura. There were no differences in pattern

of mineralization seen at day 28 in osteoblasts co-cultured with dura mater cells when

comparing control, patent suture and fused suture, suggesting that interaction with dura mater

cells resulted in down-regulation of mineralization in vitro. These findings suggest that

paracrine signalling from dura mater controls mineralization at the suture site by up or

downregulating osteoblast differentiation.

87

The role of anti-osteogenic signalling on human osteoblasts in vitro with and without

dura mater cells in co-culture

Our data suggest an important role for Noggin in pathologic suture fusion. Low

expression of Noggin was found in the fused suture when osteoblasts were cultured alone. In

the same system, patent suture osteoblasts expressed higher levels of Noggin. Interestingly,

Noggin was expressed in the conditioned medium from all three sites when combined with

dura mater cells, suggesting that dura mater cells may be the source of Noggin, maintaining

the patency of the suture line. These findings are in concordance with the literature. Warren

et al27, 75

have examined postnatal suture mesenchyme in an attempt to determine Noggin’s

role in normal suture fusion in a murine model. They found that Noggin was expressed by

the patent sagittal suture but not by the fused posterior frontal suture. They also found that

expression of Noggin was decreased by FGF2 and that overexpression of Noggin, induced by

gene transfection, resulted in suture patency of the normally fused suture.

BMP3 has also been implicated in the process of normal suture fusion in mice. Our

results confirm the findings from the normal suture fusion model. BMP3 was strongly

expressed by the patent suture as demonstrated by IHC, but BMP3 was not expressed by the

fused suture. Expression of BMP3 was increased in co-cultures of dura mater cells with

patent suture cells, which did not occur in fused suture or control sites osteogenic co-cultures.

This may suggest that Noggin and BMP3 do not have the same responses to dura mater

paracrine signalling. Based on our findings, co-culture with dura mater cells was able to

change only the pattern of Noggin expression leading us to speculate that dura mater could

be a source of Noggin regulating suture patency and fusion. This will be further investigated

in our laboratory.

88

Further studies evaluating the influence of dura mater and/or Noggin in Runx2

expression, are necessary to elucidate osteoblast behavior and regional variations in

craniosynostosis.

The role of dura mater paracrine signalling in the pathophysiology of craniosynostosis

To better understand the influences of dura mater and the biomolecular mechanisms

mediating the observed changes in proliferation, gene expression, and bone nodule formation

of co-cultured osteoblasts, we examined the expression of TGF-β1 and FGF2 in the

conditioned medium from dural cell cultures at days 3, 5, 7 and 10.

We focused on these candidate molecules because evidence suggests dura mater is a

source of FGF2 and TGF-β1 and they promote osteogenesis and osseous repair in vivo72, 76

.

The actions of these growth factors are highly concentration dependent and influence a

number of cellular processes77

. Our results demonstrated dural cells expressed both TGF-β1

and FGF2. It is possible that FGF2 released by dura mater cells may be a key growth factor

responsible for the paracrine effects of dura mater in controlling suture patency and fusion.

Considering we were able to identify the expression of these molecules in the

medium from dura mater cells, we questioned whether the levels of dural cell TGF-β1 and

FGF2 were responsible for the observed changes in proliferation and differentiation of co-

cultured osteoblasts. The literature concerning the effect of TGF-β1 on osteoblasts in vitro

contains many conflicting reports. TGF-β1 may cause either an increase or decrease in

osteoblast proliferation depending on the type of cell and cellular density78, 79

. Irrespective of

its effects on osteoblast proliferation, TGF-β1 is a potent chemoattractant for osteoblast-like

89

cells. Thus, the osteoinductive effects of TGF-β1 in vivo may instead be related to its ability

to recruit osteoprogenitor cells80

.

Interestingly, several studies have demonstrated that TGF-β1 has conflicting effects

on osteoblast type I collagen and alkaline phosphatase expression81-83

. Most notably, in

contrast to its in vivo activity, in vitro TGF-β1 seems to retard osteoblast differentiation and

decrease osteoblast expression of osteocalcin and the formation of mineralized bone

nodules81

. Thus, it is unlikely that TGF-β1 alone could account for the dramatic phenotypic

differences observed in co-cultured osteoblasts. Based on these reports, we did not

investigate the influence of TGF-β1 in osteoblast behavior. We decided to focus on FGF2

effects on osteoblast proliferation and function.

Fibroblast growth factors are a highly conserved family of at least 19 closely related

monomeric peptides. FGF2 is the most abundant ligand, and it has been shown to stimulate

osteoblast proliferation and enhance bone formation in vivo and in vitro84

. FGF2 expression

is elevated during fracture healing, and exogenously applied FGF2 accelerates osteogenesis

in critically sized bone defects and at fracture sites85

. Despite its in vivo osteoinductive

properties, FGF2 had been thought, until recently, to decrease osteoblast proliferation and

expression of markers of differentiation, such as type I collagen, alkaline phosphatase, and

osteocalcin86

. However, a recent study by Debiais et al58

demonstrated that an osteoblast’s

response to FGF2 is stage specific. Immature osteoblasts (3 days old mice) show significant

increase in proliferation under FGF2 stimulation in comparison to mature osteoblasts (30

days old mice). Continuous (7-day) FGF2 treatment of differentiating human calvarial–

derived osteoblasts (immature cells) resulted in increased osteocalcin production and matrix

mineralization. The above-mentioned studies on the isolated effects of TGF-β1 and FGF2 are

90

interesting, but osteoblasts in a calvarial defect are exposed simultaneously to a variety of

cytokines. Could TGF-β1 and FGF2 interact to affect osteoblast activity? In one of the only

studies to investigate the effect of simultaneous TGF-β1 and FGF2 stimulation on osteoblast

behavior, Globus et al87

demonstrated that these cytokines acted in a dose-dependent,

synergistic manner to increase the rate of bovine osteoblast proliferation.

Effect of exogenous human recombinant FGF2 administration on osteoblast function in

vitro

The increased rate of proliferation of our co-cultured patent suture osteoblasts may

have resulted from the synergistic effects of TGF-β1 and FGF2 elaborated by the dural cells.

To date no studies have adequately examined the effect of simultaneous (long-term) TGF-β1

and FGF2 treatment on osteoblast gene expression and differentiation.

To elucidate the effects of FGF2 on osteoblast behavior in short-term cultures we

supplemented our standard osteogenic media with human recombinant FGF2 (5, 10, 50 and

100ng/ml)45

. Cells derived from control bone, patent suture and fused suture demonstrated a

significant (p<0.05) increase of proliferation rates over time and compared with unstimulated

osteoblasts, showing that FGF2 is a potent mitogenic factor in human calvarial cells. In the

FGF2 stimulation experiments, unstimulated osteogenic cells from all three groups

demonstrated similar rates of proliferation at days 5 and 7 when compared with our previous

experiments on proliferation where fused suture osteoblasts demonstrated significantly

higher rates of proliferation. Cell cultures used for FGF2 experiments were derived from

older patients (mean age 11.7 months old) and mainly from coronal sutures, whereas cell

cultures used for the proliferation experiments were mainly from sagittal sutures and younger

91

patients (5.1 months old). These findings support the influence of age on osteoblast behavior

and may also suggest that osteoblasts from different sutures behave differently.

In accordance with previous observations, quantification of alkaline phosphatase was

not significantly increased when comparing the osteoblasts unstimulated with osteoblasts

stimulated by different concentrations of FGF2. When comparing results obtained with FGF2

stimulation and co-culture with dura mater cells, the latest was responsible for a more

consistent enhancing in osteoblast proliferation. These findings suggest that the effects of

FGF2 are dose dependent and may be also depend of the timing of exposure to growth

factors. In vivo, during osteogenesis, osteoblasts are exposed to dura mater influence since

the first trimester, which may explain the behavior of cells at different suture sites.

Unfortunately, we have not been able to develop such an ideal condition to confirm those

suppositions.

These observations highlight the multifactorial nature of calvarial bone development,

suggesting that premature cranial suture fusion most likely results from activation of

osteoprogenitors and osteoblasts in combination with the proliferation and differentiation of a

subpopulation of osteoblast-like cells residing within the dura mater itself.

In summary, the study reported here demonstrated that dura mater can affect

proliferation, rate and degree of differentiation, and bone nodule formation. Furthermore, we

demonstrated that non–suture-associated dura mater in isolation produced high levels of

osteoinductive cytokines (i.e., TGF-β1 and FGF2). Although these candidate cytokines cause

conflicting effects on osteoblasts in vitro and in vivo, they are interesting candidates for the

changes in behavior observed in co-cultured osteoblasts. As a future direction of this work

we aim to decipher and understand the exact nature of paracrine signalling responsible for the

92

observed changes in phenotype and function of co-cultured osteoblasts and include Noggin

as potential candidate for the paracrine signalling by dura mater cells.

93

Conclusion

94

The work herein demonstrates that regional variations in osteoblast behavior exist in

the calvaria of infants affected by craniosynostosis. A successful cell culture technique for

generating a reliable osteoblast culture was developed. Co-culture with dura mater

demonstrated that there was no significant effect on osteoblast proliferation rates from

regions of fused suture and control bone. However, dural influence in vitro was noted in AP

expression and mineralization rates suggestive of a paracrine effect. This was demonstrated

by assays detecting important cytokines (FGF2 and TGF-β1) in the media of the dura mater

cultures.

The increased expression of anti-osteogenic molecules, Noggin and BMP3 in patent

sutures osteoblasts suggests that they play a role in the maintenance of suture patency and

down-regulation at levels similar with control bone was noted in the fused suture osteoblasts

acting as further evidence of their role.

The creation of a pro-osteogenic environment that would support premature fusion of

the cranial sutures was further corroborated with the Runx2 studies. These studies

demonstrated that Runx2, a transcription factor that is important in osteoblast differentiation

was expressed by fused suture cells. This would suggest that these cells are more responsive

and amenable to an environment that would promote osteogenesis and hence, fusion of the

sutures.

Exogenous FGF2 was demonstrated to selectively enhance growth of fused suture

cell cultures. This pro-osteogenic molecule did not have the same impact on cells from patent

sutures, thereby confirming the presence of regional variations in osteoblast behavior.

95

In summary, we would support the theory that activation of the fibroblast growth

factor signalling pathway secreted by dura mater potentiates suture osteogenesis by inhibiting

the expression of osteogenic antagonists such as Noggin and BMP3, allowing agonists such

as bone morphogenic proteins (BMP2 and 4) to increase osteoblastic differentiation from

cells in the suture mesenchyme leading to premature suture fusion.

A better understanding of the mechanism underlying of this condition could lead to

the development of new therapeutic modalities. For example, if nonsyndromic

craniosynostosis derives from aberrant growth factor production or action, correction of this

problem might be possible through selective drug therapy. Gene therapy to correct the

molecular defect(s), however, will clearly require a thorough understanding of the complex

cascades regulating suture fusion and patency.

96

Future Directions

97

As a future direction of this work we aim to decipher and understand the exact nature

of paracrine signalling responsible for the observed changes in phenotype and function of co-

cultured osteoblasts.

Although we demonstrated important differences in osteoblast behavior depending on

the suture site and the interaction with dura mater cells, we were limited by the age of

patients who underwent neurosurgical procedures. In the future, studies obtaining dura mater

samples from age-matched patients and from the fused suture site would shed light on the

mechanisms involved in the local paracrine signalling by dura mater cells. The current co-

culture model described in our work also does not account for the effects of cell-cell

interaction or the effects of conditioned medium collected from dura mater. Those

interactions would be further investigated based on our human bone culture model of

craniosynostosis. By enhancing our understanding of the cellular events which comprise

suture formation and growth, it may be possible to recapitulate, in an experimental setting,

the normal development of a suture at a specific calvarial site.

Based on the findings presented in this work, a further step would be to investigate in

greater depth the influence of age in osteoblast behavior. Osteoblasts from non-syndromic

patients demonstrated regional variations not seen in the osteoblasts from syndromic patients.

Furthermore, proliferation rates were equal for all three groups in the FGF2 experiments

when osteoblasts were not stimulated. Reviewing patients’ age, osteoblasts for those

experiments were retrieved from older patients (mean age 11.7 months) which could explain

the lack of differences. Taken all together, investigation of the influence of age in osteoblast

behavior could shed light some of the contradictory findings of this work.

98

Another important step forward would be to understand in greater depth the role of

Noggin in the pathophysiology and therapeutics of craniosynostosis. Further experiments

will investigate the expression of Noggin by dura mater cells. Noggin also will be added to

cultured cells, as well as, anti-Noggin antibody to neutralize its effects and parameters of

proliferation and function will be reassessed. Patients will be also stratified by age to

determine the ideal time point where cells would be able to respond to a therapy.

Though at present we are just beginning to understand the phenomenon of premature

suture fusion, ultimately, this may lead to a new therapeutic modality as an alternative to

invasive procedures which would restore both the structure and function of the developing

cranium.

99

References

100

1. Shen K, Krakora SM, Cunningham M, et al. Medical treatment of craniosynostosis:

recombinant Noggin inhibits coronal suture closure in the rat craniosynostosis model. Orthod

Craniofac Res 2009;12:254-62.

2. Delashaw JB, Persing JA, Broaddus WC, Jane JA. Cranial vault growth in

craniosynostosis. J Neurosurg 1989;70:159-65.

3. Cohen MM, Jr., Kreiborg S. Birth prevalence studies of the Crouzon syndrome:

comparison of direct and indirect methods. Clin Genet 1992;41:12-5.

4. Renier D, Sainte-Rose C, Marchac D, Hirsch JF. Intracranial pressure in

craniostenosis. J Neurosurg 1982;57:370-7.

5. Kapp-Simon KA, Figueroa A, Jocher CA, Schafer M. Longitudinal assessment of

mental development in infants with nonsyndromic craniosynostosis with and without cranial

release and reconstruction. Plast Reconstr Surg 1993;92:831-9; discussion 40-1.

6. Moore MH. Upper airway obstruction in the syndromal craniosynostoses. Br J Plast

Surg 1993;46:355-62.

7. Lo LJ, Chen YR. Airway obstruction in severe syndromic craniosynostosis. Ann Plast

Surg 1999;43:258-64.

8. Virtanen R, Korhonen T, Fagerholm J, Viljanto J. Neurocognitive sequelae of

scaphocephaly. Pediatrics 1999;103:791-5.

9. Hayward R, Jones B, Evans R. Functional outcome after surgery for trigonocephaly.

Plast Reconstr Surg 1999;104:582-3.

10. Speltz ML, Greenberg MT, Endriga MC, Galbreath H. Developmental approach to

the psychology of craniofacial anomalies. Cleft Palate Craniofac J 1994;31:61-7.

101

11. Pertschuk MJ, Whitaker LA. Psychosocial considerations in craniofacial deformity.

Clin Plast Surg 1987;14:163-8.

12. Coccaro PJ, McCarthy JG, Epstein FJ, Wood-Smith D, Converse JM. Early and late

surgery in craniofacial dysostosis: a longitudinal cephalometric study. Am J Orthod

1980;77:421-36.

13. Kreiborg S, Aduss H. Pre- and postsurgical facial growth in patients with Crouzon's

and Apert's syndromes. Cleft Palate J 1986;23 Suppl 1:78-90.

14. McCarthy JG, Cutting CB. The timing of surgical intervention in craniofacial

anomalies. Clin Plast Surg 1990;17:161-82.

15. DiRocco C, Caldarelli M, Ceddia A, Iannelli A, Velardi F. [Craniostenosis. Analysis

of 161 cases surgically treated during the first year of life]. Minerva Pediatr 1989;41:393-

404.

16. Moss ML. Growth of the calvaria in the rat; the determination of osseous

morphology. Am J Anat 1954;94:333-61.

17. Pritchard JJ, Scott JH, Girgis FG. The structure and development of cranial and facial

sutures. J Anat 1956;90:73-86.

18. Markens IS. Embryonic development of the coronal suture in man and rat. Acta Anat

(Basel) 1975;93:257-73.

19. Cohen MM, Jr. Sutural biology and the correlates of craniosynostosis. Am J Med

Genet 1993;47:581-616.

20. Johansen VA, Hall SH. Morphogenesis of the mouse coronal suture. Acta Anat

(Basel) 1982;114:58-67.

102

21. Kokich VG. Age changes in the human frontozygomatic suture from 20 to 95 years.

Am J Orthod 1976;69:411-30.

22. Winograd JM, Im MJ, Vander Kolk CA. Osteoblastic and osteoclastic activation in

coronal sutures undergoing fusion ex vivo. Plast Reconstr Surg 1997;100:1103-12.

23. Koskinen L, Isotupa K, Koski K. A note on craniofacial sutural growth. Am J Phys

Anthropol 1976;45:511-6.

24. Moss ML. Fusion of the frontal suture in the rat. Am J Anat 1958;102:141-65.

25. Ozaki W, Buchman SR, Muraszko KM, Coleman D. Investigation of the influences

of biomechanical force on the ultrastructure of human sagittal craniosynostosis. Plast

Reconstr Surg 1998;102:1385-94.

26. Kirschner RE, Gannon FH, Xu J, et al. Craniosynostosis and altered patterns of fetal

TGF-beta expression induced by intrauterine constraint. Plast Reconstr Surg 2002;109:2338-

46; discussion 47-54.

27. Warren SM, Brunet LJ, Harland RM, Economides AN, Longaker MT. The BMP

antagonist noggin regulates cranial suture fusion. Nature 2003;422:625-9.

28. Nacamuli RP, Fong KD, Lenton KA, et al. Expression and possible mechanisms of

regulation of BMP3 in rat cranial sutures. Plast Reconstr Surg 2005;116:1353-62.

29. Gabbay JS, Heller J, Spoon DB, et al. Noggin underexpression and runx-2

overexpression in a craniosynostosis rabbit model. Ann Plast Surg 2006;56:306-11.

30. Greenwald JA, Mehrara BJ, Spector JA, et al. In vivo modulation of FGF biological

activity alters cranial suture fate. Am J Pathol 2001;158:441-52.

31. Urist MR. Bone morphogenetic protein: the molecularization of skeletal system

development. J Bone Miner Res 1997;12:343-6.

103

32. Hino J, Matsuo H, Kangawa K. Bone morphogenetic protein-3b (BMP-3b) gene

expression is correlated with differentiation in rat calvarial osteoblasts. Biochem Biophys

Res Commun 1999;256:419-24.

33. Daluiski A, Engstrand T, Bahamonde ME, et al. Bone morphogenetic protein-3 is a

negative regulator of bone density. Nat Genet 2001;27:84-8.

34. Candia AF, Watabe T, Hawley SH, et al. Cellular interpretation of multiple TGF-beta

signals: intracellular antagonism between activin/BVg1 and BMP-2/4 signaling mediated by

Smads. Development 1997;124:4467-80.

35. Fong KD, Song HM, Nacamuli RP, et al. Apoptosis in a rodent model of cranial

suture fusion: in situ imaging and gene expression analysis. Plast Reconstr Surg

2004;113:2037-47.

36. Mehrara BJ, Mackool RJ, McCarthy JG, Gittes GK, Longaker MT.

Immunolocalization of basic fibroblast growth factor and fibroblast growth factor receptor-1

and receptor-2 in rat cranial sutures. Plast Reconstr Surg 1998;102:1805-17; discussion 18-

20.

37. Greenwald JA, Mehrara BJ, Spector JA, et al. Regional differentiation of cranial

suture-associated dura mater in vivo and in vitro: implications for suture fusion and patency.

J Bone Miner Res 2000;15:2413-30.

38. Smith DW, Tondury G. Origin of the calvaria and its sutures. Am J Dis Child

1978;132:662-6.

39. Moss ML. Inhibition and stimulation of sutural fusion in the rat calvaria. Anat Rec

1960;136:457-67.

104

40. Hobar PC, Schreiber JS, McCarthy JG, Thomas PA. The role of the dura in cranial

bone regeneration in the immature animal. Plast Reconstr Surg 1993;92:405-10.

41. Mossaz CF, Kokich VG. Redevelopment of the calvaria after partial craniectomy in

growing rabbits: the effect of altering dural continuity. Acta Anat (Basel) 1981;109:321-31.

42. Yu JC, McClintock JS, Gannon F, Gao XX, Mobasser JP, Sharawy M. Regional

differences of dura osteoinduction: squamous dura induces osteogenesis, sutural dura induces

chondrogenesis and osteogenesis. Plast Reconstr Surg 1997;100:23-31.

43. Slater BJ, Kwan MD, Gupta DM, Amasha RR, Wan DC, Longaker MT. Dissecting

the influence of regional dura mater on cranial suture biology. Plast Reconstr Surg

2008;122:77-84.

44. Li S, Quarto N, Longaker MT. Dura mater-derived FGF-2 mediates mitogenic

signaling in calvarial osteoblasts. Am J Physiol Cell Physiol 2007;293:C1834-42.

45. James AW, Xu Y, Wang R, Longaker MT. Proliferation, osteogenic differentiation,

and fgf-2 modulation of posterofrontal/sagittal suture-derived mesenchymal cells in vitro.

Plast Reconstr Surg 2008;122:53-63.

46. Guzel MZ, Yildirim AM, Yucel A, Seradjmir M, Dervisoglu S. Osteogenic potential

of infant dural grafts in different recipient beds. J Craniofac Surg 1995;6:489-93.

47. Geoffroy V, Corral DA, Zhou L, Lee B, Karsenty G. Genomic organization,

expression of the human CBFA1 gene, and evidence for an alternative splicing event

affecting protein function. Mamm Genome 1998;9:54-7.

48. Nacamuli RP, Fong KD, Warren SM, et al. Markers of osteoblast differentiation in

fusing and nonfusing cranial sutures. Plast Reconstr Surg 2003;112:1328-35.

105

49. Otto F, Thornell AP, Crompton T, et al. Cbfa1, a candidate gene for cleidocranial

dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell

1997;89:765-71.

50. Kimelman D, Kirschner M. Synergistic induction of mesoderm by FGF and TGF-beta

and the identification of an mRNA coding for FGF in the early Xenopus embryo. Cell

1987;51:869-77.

51. Globus RK, Plouet J, Gospodarowicz D. Cultured bovine bone cells synthesize basic

fibroblast growth factor and store it in their extracellular matrix. Endocrinology

1989;124:1539-47.

52. Noff D, Pitaru S, Savion N. Basic fibroblast growth factor enhances the capacity of

bone marrow cells to form bone-like nodules in vitro. FEBS Lett 1989;250:619-21.

53. Pitaru S, Kotev-Emeth S, Noff D, Kaffuler S, Savion N. Effect of basic fibroblast

growth factor on the growth and differentiation of adult stromal bone marrow cells: enhanced

development of mineralized bone-like tissue in culture. J Bone Miner Res 1993;8:919-29.

54. Tang KT, Capparelli C, Stein JL, et al. Acidic fibroblast growth factor inhibits

osteoblast differentiation in vitro: altered expression of collagenase, cell growth-related, and

mineralization-associated genes. J Cell Biochem 1996;61:152-66.

55. Moore R, Ferretti P, Copp A, Thorogood P. Blocking endogenous FGF-2 activity

prevents cranial osteogenesis. Dev Biol 2002;243:99-114.

56. Wilkie AO, Morriss-Kay GM, Jones EY, Heath JK. Functions of fibroblast growth

factors and their receptors. Curr Biol 1995;5:500-7.

57. Robin NH, Falk MJ, Haldeman-Englert CR. FGFR-Related Craniosynostosis

Syndromes. 1993.

106

58. Debiais F, Hott M, Graulet AM, Marie PJ. The effects of fibroblast growth factor-2

on human neonatal calvaria osteoblastic cells are differentiation stage specific. J Bone Miner

Res 1998;13:645-54.

59. Mooney MP, Losken HW, Siegel MI, et al. Development of a strain of rabbits with

congenital simple nonsyndromic coronal suture synostosis. Part II: Somatic and craniofacial

growth patterns. Cleft Palate Craniofac J 1994;31:8-16.

60. Shevde NK, Bendixen AC, Maruyama M, Li BL, Billmire DA. Enhanced activity of

osteoblast differentiation factor (PEBP2alphaA2/CBFa1) in affected sutural osteoblasts from

patients with nonsyndromic craniosynostosis. Cleft Palate Craniofac J 2001;38:606-14.

61. Coelho MJ, Cabral AT, Fernande MH. Human bone cell cultures in biocompatibility

testing. Part I: osteoblastic differentiation of serially passaged human bone marrow cells

cultured in alpha-MEM and in DMEM. Biomaterials 2000;21:1087-94.

62. Pradel W, Mai R, Gedrange T, Lauer G. Cell passage and composition of culture

medium effects proliferation and differentiation of human osteoblast-like cells from facial

bone. J Physiol Pharmacol 2008;59 Suppl 5:47-58.

63. Reuther T, Kettmann C, Scheer M, Kochel M, Iida S, Kubler AC. Cryopreservation

of osteoblast-like cells: viability and differentiation with replacement of fetal bovine serum in

vitro. Cells Tissues Organs 2006;183:32-40.

64. Gevorgyan A, La Scala GC, Sukhu B, et al. An in vitro model of radiation-induced

craniofacial bone growth inhibition. J Craniofac Surg 2007;18:1044-50.

65. Carmichael J, DeGraff WG, Gazdar AF, Minna JD, Mitchell JB. Evaluation of a

tetrazolium-based semiautomated colorimetric assay: assessment of chemosensitivity testing.

Cancer Res 1987;47:936-42.

107

66. VanGuilder HD, Vrana KE, Freeman WM. Twenty-five years of quantitative PCR for

gene expression analysis. Biotechniques 2008;44:619-26.

67. Spector JA, Greenwald JA, Warren SM, et al. Co-culture of osteoblasts with

immature dural cells causes an increased rate and degree of osteoblast differentiation. Plast

Reconstr Surg 2002;109:631-42; discussion 43-4.

68. Coelho MJ, Fernandes MH. Human bone cell cultures in biocompatibility testing.

Part II: effect of ascorbic acid, beta-glycerophosphate and dexamethasone on osteoblastic

differentiation. Biomaterials 2000;21:1095-102.

69. De Pollack C, Renier D, Hott M, Marie PJ. Increased bone formation and osteoblastic

cell phenotype in premature cranial suture ossification (craniosynostosis). J Bone Miner Res

1996;11:401-7.

70. Ganta DR, McCarthy MB, Gronowicz GA. Ascorbic acid alters collagen integrins in

bone culture. Endocrinology 1997;138:3606-12.

71. Hassler W, Zentner J. Radical osteoclastic craniectomy in sagittal synostosis.

Neurosurgery 1990;27:539-43.

72. Bradley JP, Han VK, Roth DA, Levine JP, McCarthy JG, Longaker MT. Increased

IGF-I and IGF-II mRNA and IGF-I peptide in fusing rat cranial sutures suggest evidence for

a paracrine role of insulin-like growth factors in suture fusion. Plast Reconstr Surg

1999;104:129-38.

73. Bradley JP, Levine JP, McCarthy JG, Longaker MT. Studies in cranial suture

biology: regional dura mater determines in vitro cranial suture fusion. Plast Reconstr Surg

1997;100:1091-9; discussion; 100-2.

108

74. Greenwald JA, Mehrara BJ, Spector JA, et al. Biomolecular mechanisms of calvarial

bone induction: immature versus mature dura mater. Plast Reconstr Surg 2000;105:1382-92.

75. Zimmerman LB, De Jesus-Escobar JM, Harland RM. The Spemann organizer signal

noggin binds and inactivates bone morphogenetic protein 4. Cell 1996;86:599-606.

76. Song HM, Fong KD, Nacamuli RP, et al. Mechanisms of murine cranial suture

patency mediated by a dominant negative transforming growth factor-beta receptor

adenovirus. Plast Reconstr Surg 2004;113:1685-97.

77. Bostrom MP, Saleh KJ, Einhorn TA. Osteoinductive growth factors in preclinical

fracture and long bone defects models. Orthop Clin North Am 1999;30:647-58.

78. Uneno S, Yamamoto I, Yamamuro T, et al. Transforming growth factor beta

modulates proliferation of osteoblastic cells: relation to its effect on receptor levels for

epidermal growth factor. J Bone Miner Res 1989;4:165-71.

79. Robey PG, Young MF, Flanders KC, et al. Osteoblasts synthesize and respond to

transforming growth factor-type beta (TGF-beta) in vitro. J Cell Biol 1987;105:457-63.

80. Lucas PA. Chemotactic response of osteoblast-like cells to transforming growth

factor beta. Bone 1989;10:459-63.

81. Harris SE, Bonewald LF, Harris MA, et al. Effects of transforming growth factor beta

on bone nodule formation and expression of bone morphogenetic protein 2, osteocalcin,

osteopontin, alkaline phosphatase, and type I collagen mRNA in long-term cultures of fetal

rat calvarial osteoblasts. J Bone Miner Res 1994;9:855-63.

82. Noda M, Rodan GA. Type-beta transforming growth factor inhibits proliferation and

expression of alkaline phosphatase in murine osteoblast-like cells. Biochem Biophys Res

Commun 1986;140:56-65.

109

83. Centrella M, McCarthy TL, Canalis E. Transforming growth factor beta is a

bifunctional regulator of replication and collagen synthesis in osteoblast-enriched cell

cultures from fetal rat bone. J Biol Chem 1987;262:2869-74.

84. Nakamura K, Kurokawa T, Kawaguchi H, et al. Stimulation of endosteal bone

formation by local intraosseous application of basic fibroblast growth factor in rats. Rev

Rhum Engl Ed 1997;64:101-5.

85. Kawaguchi H, Kurokawa T, Hanada K, et al. Stimulation of fracture repair by

recombinant human basic fibroblast growth factor in normal and streptozotocin-diabetic rats.

Endocrinology 1994;135:774-81.

86. Canalis E, Centrella M, McCarthy T. Effects of basic fibroblast growth factor on bone

formation in vitro. J Clin Invest 1988;81:1572-7.

87. Globus RK, Patterson-Buckendahl P, Gospodarowicz D. Regulation of bovine bone

cell proliferation by fibroblast growth factor and transforming growth factor beta.

Endocrinology 1988;123:98-105.

110

Appendices

111

Calvarial osteogenic cell culture

Calvarial osteogenic cell culture preparation

The bone sample is kept on ice in a 50 ml Falcon tube with αMEM and 5xpenicillin-

streptomycin until digestion. Samples are processed within 30 min after taken from the

operating room. It is then transferred to a 65-mm Petri dish (Falcon™, Becton, Dickinson

and Co., Franklin Lakes, NJ –Ref. 35002) with 6 ml collagenase mixture (See appendix pg.

126). The bone is diced with scissors into as small as possible pieces (< 0.5 mm3). The

mixture with small bone fragments is transferred to a 14 ml tube (Falcon, Ref. 35-2059) and

incubated at 37° C for 20 minutes for digestion in collagenase mixture (See appendix pg.

126). During digestion, the tube is removed from the incubator and mixed on Vortex for 30

seconds every 5 minutes. After 20 minutes of incubation, the tube is removed from the

incubator and left for the larger pieces to settle to the bottom of the tube. The upper layers of

the mixture are transferred to a new 50 ml Falcon tube, already containing 15 ml of FSGM

(See appendix pg.127). Then, 6 ml of collagenase mixture is added to tube 1, containing

small bone pieces for further digestion at 37° C for 20 minutes.

The above procedure is repeated to a total of 3 times to digest out as many cells as

possible. Cells collected from each digestion are added to the same 50 ml Falcon tube. After

the final digestion, the 50 ml Falcon tube is centrifuged at 4° C, 700 rpm (80.5× g) in IEC

Centra MP4R (International Equipment Company, Needham Heights, MA – Ref. 2438; 28-

tubes, 4 × 250 ml rotor head, Ref. 224) for 15 minutes. The supernatant is removed and

discarded. The cell pellet is diluted in 15 ml FSGM, resuspended and plated into a T75 tissue

culture flask (SARSTEDT Ref. 83.1813.002). The medium is changed every 2-3 days until

confluence.

112

Calvarial osteogenic cell subculture

At confluence, the culture medium is aspirated with a sterile Pasteur’s pipette and

discarded. The flasks are rinsed with 5 ml PBS without Ca and Mg (Wisent Bioproducts Ref.

80450) to remove any residual medium and the liquid is aspirated. 5 ml Trypsin-EDTA 1x

(Wisent Bioproducts Ref. 325042) reconstituted in PBS without Ca and Mg and the flask is

incubated at 37° for 5 minutes. Then, the flask is removed from the incubator and observed

under the phase contrast microscope to insure that all cells are detached. If any cells are

remaining attached, tap the side of the flask to detach the cells. If necessary, a cell scraper

(SARSTEDT Ref. 83.1830) is used to remove any residual cell colony attached to the plate.

Neutralize Trypsin with 5 ml of FSGM, pipetting up and down and washing the

bottom of the flask. The content of the flask is moved to a 14 ml tube (Falcon – Ref. 35-

2098) and the later is centrifuged at 800 rpm (105.2x g), 4°C for 7 minutes [relative

centrifugal force (RCF) value (g) = 1118 × 10E-8 × R × N2, where R is the radius in cm from

the centre of the rotor to the point at which the RCF value is required (for 4, 28 and 48 tubes

=14.7 cm), and N = speed of centrifuge in rpm]). The supernatant is carefully aspirated with

a sterile Pasteur’s pipette and discarded, leaving very little liquid on top of the cell pellet.

Then, precisely 5 ml of FSGM is added to the cell pellet and pipetted up and down 10-20

times to obtain single-cell suspension. If multiple primary cell cultures exist, the suspensions

are combined in a larger tube. If only a small number of cells are expected to be retrieved

from the pellet, little amount (1-2 ml) of FSGM is used.

After resuspending the pellet, 0.1-0.2 ml of the suspension is transferred to an

Eppendorf tube. Cells are counted in a haemocytometer and plated at densities of 1000 cells

per well in flat bottomed 96-well plates (SARSTEDT – Ref. 83.1835) for alkaline

113

phosphatase (AP) (including protein) and mineralization assays. Three columns, comprising

of 4 wells is used for each AP and protein assays on the same plate and 4 wells for

mineralization assay on a separate plate. Concurrently, cells are seeded into 24-well plates, 4

wells per plate for each of AP/Protein and mineralization assays. For collagen type I assay,

50,000 cells per well are seeded in a flat bottomed 6-well tissue culture plate (SARSTEDT

Cat# 83.1839).

Medium change in 96-well plates

Medium (α-MEM with 15% FCS, 10-7 Dexamethasone) was changed on Monday,

Wednesday and Friday. The old medium was removed with a 20G (at later days, especially

with folding, 25-26G) sterile needle connected to a wall suction system, suctioning the fluid

form the edge of the left lower quadrant of the culture well. A new needle was used for each

plate, in order to reduce the risk of contamination. Using a multichannel (8) pipette, 100 μl

(150 μl for the weekend) of medium was delivered to each of the wells in the 96-well plates,

changing the tips between plates, depending on the medium change interval and cell

concentration. For the 6-well plates, 2 ml (3 ml for the weekend) of medium was added. At

around day 7 (1 week in subculture), prior to medium change, vitamin C (Ascorbic acid –

Sigma/Aldrich Cat# A2218), 0.33 μl stock solution (50 μg) / 1 ml of medium, necessary for

collagen synthesis, and β-glycerophosphate 1% (Sigma – Ref. G6251) were added to

promote mineralization.

114

Medium Composition Study

Cells were subcultured by trypsinization (5 mL Trypsin-EDTA) and neutralized in 5 mL

of FSGM to obtain single-cell suspension that was transferred to an Eppendorf tube17

.

Subsequently, cells were counted in a haemocytometer and plated at densities of 1000 cells

per 96-well culture plates. On day 3, 5 and 7 of culture, old medium (αMEM with 15% FBS)

was removed and 200 μl of either, (αMEM containing 10-7

M dexamethasone, supplemented

with 15%, 10%, 1% of FBS, or an osteogenic medium supplemented with ascorbic acid and

β-glycophosphate was added to each well to generate four culture medium conditions. Cells

were incubated at 37°C and the medium was changed every 2-3 days.

Preparation of the periosteum-derived osteoblast cell culture

The periosteum was kept in α-MEM + RNA + DNA + 1× antibiotic on ice until

digestion and then transfered to 3 ml collagenase mixture (for composition, see Materials pg.

120) in a 60-mm Petri dish (Falcon – Ref. 4-1007-0). The periosteum was diced with scissors

into small pieces (< 1 mm3) and transfered to a 14 ml tube (Falcon – Ref. 35- 2059) and

incubated at 37° C for 15'. 5 ml of α-MEM + RNA + DNA + 1× antibiotic (penicillin-

streptomycin) + 15 % FBS + 10-7 M Dexamethasone (FS-GM) were added and mixed to

allow the larger pieces to settle to the bottom of the tube (Tube 1) The upper layers of the

mixture were transferred to a new tube, centrifugated at 4° C and 700 rpm (80.5× g) in IEC

Centra MP4R (International Equipment Company, Needham Heights, MA – Ref. 2438; 28-

tubes, 4 × 250 ml rotor head, Ref. 224) for 8' and then removed and discarded the supernatant

containing collagenase. The cell pellet was diluted with 5 ml FS-GM and seeded into T25

flask with 0.2 μm vented plug cap and incubated at 37° C. 3 ml of collagenase was added to

115

tube 1, containing small periosteal pieces for further digestion at 37° C for 15'. After the

second “redo” 5 ml FS-GM and was added to the tube and centrifuged at 4° C and 700 rpm

(80.5× g) for 8'. After removing and discarding the supernatant containing collagenase, the

pellet was diluted in 5 ml FS-GM, and plated. Medium was changed every 2-3 days until

confluence.

Subculture of the periosteum derived cell culture

The culture medium was aspirated with sterile Pasteur’s pipette and discarded from

flasks. Cells were washed with 5 ml PBS without Ca, Mg (Wisent Bioproducts – Ref

311011) to remove any residual medium.PBS is then removed with sterile Pasteur’s pipette.

3-5 ml Trypsin-EDTA 1x was added (Wisent Bioproducts – Ref. 425042), reconstituted in

PBS without Calcium and Magnesium), and incubate at 37° for 5 minutes. After removal

from the incubator flasks were tapped at the side to detach the cells and a cell scraper

(SARSTEDT Ref. 83.1830) was used to remove any residual cell colony attached to the

plate. Trypsin was neutralized with 5 ml of fully supplemented growth medium (FSGM),

pipetting up and down, and washing the bottom of the flask. All the content was aspirated

and placed into a 50-ml tube (Falcon – Ref. 35-2098) and then centrifugated at 800 rpm

(105.2x g), 4°C for 7 minutes [relative centrifugal force (RCF) value (g) = 1118 × 10E-8 × R

× N2, where R is the radius in cm from the centre of the rotor to the point at which the RCF

value is required (for 4, 28 and 48 tubes =14.7 cm), and N = speed of centrifuge in rpm]).

With a sterile Pasteur’s pipette the supernatant was carefully aspirated and discarded, leaving

some liquid on top of the cell pellet.

116

Exactly 5 ml of FSGM were added to the pellet and pipette up and down 10-20 times to

obtain single cell suspension. If multiple plates exist from the same specimen, the suspension

was transferred to the next pellet-containing tube. If only a small number of cells was

expected to be retrieved from the pellet, less than 5 ml of FSGM (1 or 2 ml) was used. After

the last pellet has been suspended, cells were counting using 0.1-0.2 ml of the liquid.

• Cells were counted as previously described and plated at densities of 500 (mineralization),

and 1,000 per well in a flat bottomed 96-well tissue culture plate (SARSDETD – Ref.

83.1835), 8 wells per dilution) and 50,000 per well in a flat bottomed 6-well tissue culture

plate (Falcon – Ref. 35-3046) for collagen assay.

MTT Assay

3-[4,5-Dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide; Thiazolyl blue – kept in the

freege (4C) The MTT test is a simple, accurate, reproducible means for measuring the

activity of living cells via mitochondrial dehydrogenase activity (Mosmann, T. Rapid

colorimetric assay for cellular growth and survival: application to proliferation and

cytotoxicity assays). The reactant will form formazan crystals in living cells mitochondria

when incubated at 37° for 3-5 hours, and this will be released in solution by the DMSO.

Formazan solution absorbs light at 550-570 nm but not at 620-650 (690) nm, which

absorbance results from cell debris & well imperfections. Final optical density (OD) obtained

from formazan formation can be calculated OD=L1-L2 or automatically by the μQUANT

spectrophotometer.

Reagent

• 50mg of MTT (Sigma – Ref. M2128) dissolved in 10 ml PBS and filtered.

Technique

117

30 μl off the mixture were added to each well (analysis in triplicate), without removing the

medium and then incubated at 37°C for 5 hours. Wells contents were aspirated and 100 μl of

DMSO (Sigma – Ref. D-8779) were added to wells + one “blank” well, left for 5 minutes

and shaked (cells-containing wells produce a purple color).

Reading

In a spectrophotometer, results were read at @ 570nm (primary), with reference

wavelength of 690nm.

Alkaline phosphatase activity, current technique: p-Nitrophenyl

Phosphate (expressed per amount of protein per time of incubation of AP assay)

Reactant

Paranitrophenyl Phosphate (pNP, Sigma – Ref. 104-0) (kept in the -200C freezer) 9

mg/ml was dissolved in 10 ml of Tris buffer at pH=9.0, mix well.

Technique

Medium was removed from wells (analysis in triplicate). Wells were flushed twice

with 100 μl PBS and 200 μl of the reactant solution were added to each well + 1 for “blank”

measurement. Cells were incubated at 37°C, checking for color changes (when many cells,

significant color change in 1 hour, otherwise a few hours may be necessary). Results were

read on Titertek Multiskan MCC/340 MK II (Titertek, Huntsville, AB) at Filter 2 (@

405nm).

118

Protein absorbance

Medium was removed from wells (analysis in duplicate). Wells were flushed twice

with 100 μl NaHCO3 (Sigma – Ref. S-6014) and 100 μl of Bio-Rad protein assay reagent

(Bio-Rad Laboratories, Hercules, CA – Ref. 500-0006) diluted 1:5 in NaHCO3 at pH 7.4

were added to sample wells + to 1 “blank” well. After 10 minutes absorption was measured

on Titertek: Same procedure as above, use Filter 7 (@ 620nm)

Calculation of real values

APA: slope: 324.4 intercept: - 1.383 real value = Abs × 324.4 – 1.383

Proteins: slope: 246 intercept: 0.671 real value = Abs × 246 + 0.671

Final AP concentration is expressed as mmol of pNP per hour per μg of protein.

Alkaline phosphatase staining: Fast Blue BB Salt

Reactant

Naphtol was dissolved as phosphate (Sigma – Ref. N-5625), 3 mg in 50 μl

dimethylformammide (Sigma – Ref. D-4254) and then again in 10 ml of Tris buffer 0.2 M at

pH = 9.0 (Sigma-Fluka – Ref. 98306). 20 mg of Fast Blue BB salt (Sigma – Ref. F-3378)

was added. Wells were shake and left at room temperature for 10 minutes. As soon as the

colour changes to yellow-brownish, it was filtered using non-sterile syringe filter to get rid of

particulate matter.

Technique

100 μl of the reactant solution was added to each well and placed in the incubator at

37°C. Assessment for possible color change was done q 5 minutes (cells stained in dark blue

are considered AP positive cells [osteoblast-like cells]). For 6-well plates, 1 ml was added

119

(just to cover the cell surface) of the reactant solution to each well, placed in the incubator at

37°C, and analyzed similarly.

Alizarin Red Staining for mineralization in 96 well plates

Medium was removed and 100 µl of formalin (Sigma – Ref. HT-50-1-320) added and

left for 5 minutes. Formalin was then removed and washed twice with 150 μl of

distilled water. 50 μl of Alizarin Red S (Cat. No. A5533, Sigma Chemical Co., St.

Louis, MO) 2% staining solution were added and left for 5 minutes (added also to

“blank” well).The staining solution was removed from samples and blank and washed

twice with 150 μl of distilled water. Results were read @ 525nm on Titertek

Multiskan MCC/340 MK II.

Transmission Electron Microscopy

Electron microscopy was used to analyze bone-specific ultrastructure in vitro. Cells were

grown in 24-well plates up to day 28 in subculture, then fixed using 2% glutaraldehyde in 0.1

mol/L sodium cacodylate buffer, and postfixed in 1% osmium tetroxide in 0.1 mol/L sodium

cacodylate buffer. The samples were dehydrated through a graded ethanol series, followed by

propylene oxide, and embedded in Quetol-Spurr resin. Sections 100nm thick were cut on an

RMC MT6000 ultramicrotome, stained with uranyl acetate and lead citrate and viewed in an

FEI Tecnai 20 TEM.

120

HISTOLOGY

The bone samples were fixed in 10% phosphate buffered formalin for 48 hours at 4°C and

then transferred to Bouin’s fixative for 48 hours at 4°C, washed in distilled water, decalcified

in acetic acid/4% formaldehyde/0.85% saline, dehydrated in graded ethanol, and embedded

in paraffin. Serial sections were cut for routine hematoxylin and eosin staining to define areas

of complete ossification, fibrous tissue and remodeling bone.

IMMUNOHISTOCHEMISTRY AND IMMUNOFLUORESENCE

The samples were fixed in 3-4% paraformaldehyde in PBS pH 7.4 for 15 min at room

temperature and then washed twice with ice cold PBS. Samples were incubated for 10 min

with PBS containing 0.25% Triton X-100. Triton X-100 is the most popular detergent for

improving the penetration of the antibody. However, it is not appropriate for the use of

membrane-associated antigens since it destroys membranes. Cells were washed in PBS three

times for 5 min. Cells were then incubated with 1% BSA in PBST for 30 min to block

unspecific binding of the antibodies Following, the same cells were incubated in the diluted

antibody in 1% BSA in PBST in a humidified chamber for 1 hr at room temperature or

overnight at 4°C. The solution was decanted and cells washed three times in PBS, 5 min each

wash. Cells were then incubated with the secondary antibody in 1% BSA for 1 hr at room

temperature in dark. The secondary antibody solution was also decanted and cells washed

three times with PBS for 5 min each in dark. Cells were then incubated on 0.1-1 μg/ml

Hoechst or DAPI (DNA stain) for 1 min and rinsed with PBS. The coverslip was mounted

with a drop of mounting medium and sealed with nail polish to prevent drying and movement

under microscope. Slides were stored in dark at -20 or 4°C.

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Formalin-fixed, paraffin-embedded tissue sections (5µm) were mounted on positively

charged microscope slides. Tissue samples were then deparaffinized in xylene, 3% H2O2

was used to clear endogenous pigment. Antigen retrieval was then performed with pepsin at

37°C and tissue sections were blocked with 0.5% BSA with casein. Rabbit polyclonal to

Noggin (Abcam) was used in dilution 1:20, mouse monoclonal to Collagen I (Abcam) was

used in dilution 1:400 or rabbit polyclonal to BMP3 (R&D System) was used in dilution 1:5.

Detection was performed with Goat Polymer (Biocaremedical) for BMP3 and ABC Ellite

System (Vector) for Noggin and Collagen I. DAB was used as chromogen.

WESTERN BLOT

Cells were subcultured into 6 well plates to confluence, 500 µl of medium was taken

from each well and centrifuged at 2000 RPM for 10 min. 300 µl was taken and placed

in a new 1.5 ml tube and added 100 µl of 4x sample buffer, boiled at 100°C for 5 min.

The 200 µl were left for protein assay. Residual medium was discarded. 500 µl of

lyses buffer was added to each well, cells were collected with a 3ml Syringe and 25G

needle to small tubes, centrifuged at 14,000 RPM for 6 minutes and transferred to

new tubes. 120 µl of 4x sample buffer was added and samples were boiled at 100°C

for 5 min.

SDS

Resolving (8%) and stacking gels (4%) were prepared just before pouring the gel

between the glasses. Resolving gel (8 ml for mini gel) was poured and the upper surface was

quickly covered with water, and let polymerize for about 30min. Water on top was

eliminated, and then stacking gel (1.5 ml) was poured on top of the resolving gel. Combs

were inserted avoiding bubbles and let polymerize for about 15min. The comb was taken

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away and inserted the glasses in the Western blotting apparatus. Running buffer was added.

25 or 50 l samples supernatants and cell pellets (cell lysates: loading buffer +1:1)

previously boiled for 2 min were loaded. Proteins were separated by electrophoresis 2 h at

100V. At the end of the electrophoresis gel was directly transferred onto PVDF membrane

which was treated with MeOH for 1 min. The cassette for transferring was prepared in the

following order: black side, sponge, filter paper, gel, PVDF membrane, filter paper, sponge

and clear side. Blotting tank was placed in the cold room overnight at 36V (100mA). After

transfer, the membrane was washing once with 1X TBST and kept in 4oC.

Western blotting

The membrane was treated with MeOH for 1 min then washed with 1X TBST. The

membrane was then blocked with 4% BSA or 5% milk in 1X TBST for 1 hour at room

temperature and then washed 2X with 1X TBST. The membrane was incubated with the first

antibody in 1X TBST for 1h and washed 3 X 5min with 1X TBST. Next, the membrane was

incubated with the second antibody in 1X TBST for 1 hour at room temperature and washed

6X 5min with 1X TBST. Detection was done with Western Lightening Chemiluminescence

Reagent.

The membrane was stripped with stripping buffer and blocked with 4% BSA in

1XTBST overnight at 4oC and then washed the membrane with 1XTBST buffer. The

membrane was incubated with anti-β-actin* (1:3000) dilution in 4% BSA-TBST for 1 hour at

room temperature (*anti-β-actin, Sigma, cat# A5441, arise in Mouse, MW 47KDa) and

washed 3 X 5 min with 1X TBST. Then it was incubated with horseradish peroxidase

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conjugate anti-rabbit IgG (1:10,000) in 1X TBST for 1h and washed 5 X 5min with 1X

TBST. Detection was done with Western Lightening Chemiluminescence Reagent.

SDS PAGE Gels

Resolving Gel (35ml)

5% 6% 7% 8% 10% 12% 15% 20%

40%

Acrylamide

4.38 5.25 4.38 5.25 4.38 5.25 4.38 5.25

1.5M Tris,

pH 8.8

8.75 8.75 8.75 8.75 8.75 8.75 8.75 8.75

1% APS 2.5 2.5 2.5 2.5 2.5 2.5 2.5 2.5

dH2O 19.4 18.5 19.4 18.5 19.4 18.5 19.4 18.5

*Add 25l of TEMED

Stacking Gel (8ml)

5% 4% 3%

40% Acrylamide 1 0.8 0.6

0.5M Tris, pH6.8 2 2 2

1% APS 1 1 1

dH2O 4 4.2 4.4

*Add 8l of TEMED

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10X TBST (1L)

100ml 1M Tris pH 7.4 (121.14g Tris= 1L dH2O)

87.7g NaCl

5ml Tween 20

Adjust volume to 1L with dH2O

1X Transfer buffer

Tris 6.06g

Glycine 28.82g

Methanol 200ml

Adjust volume to 2L with dH2O

REAL TIME PCR

Cells were growth for 10 days. After remove the medium from wells incubate cells

with TRIZOL at 15-30° C for 5 minutes. 200 µl of chloroform were added per 1 ml of

TRIZOL used. Tubes were vortex for 15 seconds and then incubate at 15-30° C for

10-15 minutes with vortex half way through. Samples were centrifuge at 12,000 PRM

for 15 minutes at 2-8°C (cold room). For RNA precipitation the aqueous phase was

transferred to an eppendorf tube carefully, using a P-200 pipettor. Lower, DNA phase

was frozen -80°C. 500µl of Isopropyl Alcohol was added, mixed gently, and

incubated at room temperature for 10 minutes. Tubes were centrifuged at 12,000 X g

for 10 minutes at 4°C. The RNA precipitate formed a gel-like pellet at the bottom of

the tube. The supernatant was carefully pulled off with a P-200. The cell pellet was

125

washed once with 1ml of 75% Ethanol (ICE Cold) and centrifuge at 12,000 x g for 15

minutes at 4°C. Again, the supernatant was carefully pulled off with a P-200. The

pellet was air dried completely so there was no residual ethanol. RNA was dissolved

in Rnase- free water (DEPC WATER), volume dependent on number of cells, or size

of pellet (50µl). RNA concentration was checked by spec.

Reverse transcription was carried out with the SuperScript First-Strand

Synthesis System for RT-PC. The following RNA/primer mixture was prepared in

each tube: Total RNA 5µg, Random hexamers (50ng/µ) 3µl, 10 mM dNTO mix 1µl

and DEPC water to 10µl. Samples were incubated at 65°C for 5 min and then on iced

for at least 1 min. The reaction master mixture (10x TR buffer 2µl, 25 mM MgCl2

4µl, 0.1 M DTT 2µl and RNAase OUT 1µl) was added to the RNA/primer mixture,

mixed briefly, and then placed at room temperature for 2 min. Then 1µl (50 units) of

SuperScript II RT was added to each tube, mixed and incubated at 25°C for 10 min.

The tubes were then incubated at 42°C for 50 min, heat inactivated at 70°C for 15

min, and then chilled on ice. 1 µl RNase H was added and incubatet at 37°C for 20

min. The 1st strand cDNA was stored at -20°C until use for reat-time PCR.

The primer concentrations were normalized and mixed-specific forward and

reverse primers pair. Each primer concentration in the mixture was 5ρmol/µl. The

following mixture was prepared for each optical tube: 25 or 12.5 µl SYBR Green Mix

(2x), 0.5 or 0.2 cDNA, 2 or 1 µl primer mix pair (5ρmol/µl each primer), 22.5 or 11.3

µl water. The experiment and the following PCR program were set on ABI Prism

SDS 7000 according to the primers used: 94°C for 90 sec, 60°C for 30 sec and 72°C

for 30 sec – 40 cycles. After PCR was finished, the PCR specificity was examined by

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3% agarose gel using 25 µl from each reation. The real-time PCR results were

analyzed with a SDS 7000 software.

MATERIALS

Collagenase Incubation Mixture

Substance Supplier 200 ml 300 ml 400 ml Storage

Collagenase Sigma C 0130 or Sigma C9891 0.6 g 0.9 g 1.2 g -20°C

D-sorbitol Fisher S 459 or Sigma S-3889 (500 g) 3.644 g 5.466 g 7.288 g Shelf

(room t°)

Chondroitin

Sulphate

Fluka 27043 or Sigma C-4384 1.2 g 1.8 g 2.4 g +4°C

DNAse Sigma D 4513 0.8 ml

1.2 ml

1.6 ml

-20°C

The above chemicals were combined and added the corresponding amount of Krebs’ II

buffer with Zn2+. Stored at – 20° C.

Krebs’ II A buffer with Zn2+

Substance Concentration 1 litre 2 litres

NaCl 111.2 mM 6.4965 g 12.993 g

Tris Buffer (Base) 21.3 mM 2.5802 g 5.1604 g

Glucose 13.0 mM 2.3421 g 4.6842 g

KCl 5.4 mM 0.4026 g 0.8052 g

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MgCl2 1.3 mM 0.2643 g 0.5286 g

ZnCl2 0.5 mM 0.0682 g 0.1364 g

The first five chemicals were combined in 900 or 1800 ml of distilled water, pH adjusted to

7.4. ZnCl2 was added in and made up the volume in a 1 or 2 litre volumetric flask. The

solution was filtered sterile and stored at 4° C.

Mediums

Complete medium (15% FCS, Dexamethasone 10E-7

) from α-MEM:

α-MEM volume was divided by 0.85 to obtain the volume of the final solution, the initial α-

MEM volume was subtracted to obtain FCS volume.

(i.e.: α-MEM 500 ml: 500/0.85 = 588 ml; subtract 500 = 88 ml (vol. of FCS to add)).

Dexamethasone is provided in a 10E-3

solution: to obtain 10E-7

: (10E-7

* Volume in ml) /

10E-3

(i.e. (10E-7

* 588) / 10E-3

= 0.0588 ml [58.8 μl] of Dexamethasone 10E-3

solution)

FSGM (1% β-Glycerophosphate 15% FCS, Dexamethasone 10E-7

) from α-MEM:

Alpha-MEM volume was divided by 0.84 to obtain the volume of the final solution, divided

by 100 to obtain the volume of β-GP, subtracted this + the initial α-MEM volume to obtain

FCS volume. (i.e.: α-MEM 500 ml: 500/0.84 = 595 ml; subtract 500 + 6 (vol β-GP) = 89 ml

(vol. of FCS to add).Vitamin C (Ascorbic Acid) stock solution, 0.33 μl (50 μg) / 1 ml

medium only at the time of use.

128