insights into fracture repair using a murine model …...identified. fracture repair is delayed...
TRANSCRIPT
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INSIGHTS INTO FRACTURE REPAIR USING A MURINE
MODEL OF OSTEOFIBROUS DYSPLASIA
by
Wei Xiang Xie
A thesis submitted in conformity with the requirements
for the degree of Master of Science
University of Toronto
© Copyright by Wei Xiang (William) Xie (2019)
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Abstract
Insights into Fracture Repair Using a Murine Model of
Osteofibrous Dysplasia
Wei Xiang Xie
Master of Science
Institute of Medical Science
University of Toronto
2019
Osteofibrous dysplasia (OFD) is a rare disease characterized by the development of
radiolucent lesions at the tibial periosteal surface, where it causes non-healing fractures.
We previously identified gain-of-function mutations in the MET gene as a cause for
OFD. Fracture tissue samples from OFD patients exhibit aberrant MET activity and
delays in osteoblast differentiation. We hypothesized gain-of-function MET
mutations result in delayed bone repair ability due to reduced osteoblast
differentiation. MetΔ15-HET mice exhibit aberrant and prolonged upregulation of MET
signaling and total β-catenin levels similar to OFD patients. MetΔ15-HET osteoblasts
demonstrate a differentiation defect in vitro though no gross skeletal defects were
identified. Fracture repair is delayed in MetΔ15-HET mice, with decreased bone
formation 2-weeks post fracture-inducing surgery. Our data is consistent with a novel
role for MET-mediated signaling regulating osteogenesis and open up the possibility of
modulating the MET pathway to augment fracture healing.
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Acknowledgments
I would like to thank my everyone who has helped me throughout the completion of this
degree. Firstly, I would like to thank my supervisor Dr. Peter Kannu, for allowing me the
opportunity to pursue this degree and project and assisting in the writing of this thesis.
Secondly, I would like to thank my Program Advisory Committee Members, Drs. Morris
Manolson and Jane Mitchell for their insights, guidance and constructive criticism along
the way.
I would like to thank Dr. Kashif Ahmed for assisting and guiding me through the design
and execution of the experiments and always making himself available to assist me
when needed. Our summer student Lisa Vi in assistance with the skeletal preps and
sectioning of histological slides. I would also like to thank The Centre for
Phenogenomics for assistance in assisting with colony management.
Personally, I would like to thank my friends and family for their continuous support and
encouragement in pursuing and accomplishing my goals. Without them I would not be
the person I am today – for that I am eternally grateful.
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Contributions
William Xie (author) solely prepared this thesis. All aspects of this thesis, including
planning, execution, analysis and writing of the original research contained within was
performed by the author. The contributions by other individuals are formally and
inclusively acknowledged:
Dr. Peter Kannu (Supervisor and Committee Member) – Mentorship, laboratory space,
guidance on the planning, execution, analysis and presentation of all experiments in
addition to thesis preparation.
Dr. Jane Mitchell (Committee Member) – Mentorship, guidance in interpretation of
results and thesis preparation.
Dr. Morris Manolson (Committee Member) – Mentorship, guidance in interpretation of
results and thesis preparation.
Drs. Irina Voronov and Ralph Zirngibl (Collaborators) – Assistance in execution of
experiments and analysis of results in Figure 4.4.
Dr. Simon Kelley (Collaborator) – Assistance in performing the tibial fracture surgeries in
Figure 4.10.
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Table of Contents
Acknowledgments ................................................................................................................ iii
Contributions ........................................................................................................................ iv
List of Figures ........................................................................................................................ ix
List of Abbreviations ............................................................................................................. xi
1 Introduction ........................................................................................................................ 1 Figure 1A. Protein expression analysis of healthy bone versus osteofibrous dysplasia bone ............................. 3 Figure 1B. Regulation of MET Signaling ............................................................................................................... 4
1.1 Osteofibrous Dysplasia ................................................................................................................. 5 Figure 1C. Osteofibrous Dysplasia lesions at the tibia ......................................................................................... 8 Figure 1D. Osteofibrous Dysplasia fractures at the tibia ..................................................................................... 9 Figure 1E. Summary of Characteristics of Osteofibrous Dysplasia .................................................................... 10 Figure 1F. Genetic Summary of Osteofibrous Dysplasia .................................................................................... 11 Figure 1G. Zonal Architecture of Osteofibrous Dysplasia Lesions ..................................................................... 12
1.2 Neurofibromatosis Type 1 (NF-1) ................................................................................................ 13
1.3 Dysregulation of MET to METΔ14 ratios causes disease: Osteofibrous Dysplasia ....................... 14
1.4 Characterization of Mesenchymal-Epithelial Transition Gene’s Structure .................................. 14
1.5 MET Signaling and Function ........................................................................................................ 16
1.6 MET’s Role in Development ........................................................................................................ 17
1.7 CBL-Dependent Regulation of MET Signaling .............................................................................. 17
1.8 Other Means of Regulating MET Signaling .................................................................................. 18
1.9 Mouse Model of Osteofibrous Dysplasia .................................................................................... 19 Figure 1H. MET Exon 14 Skipping Mutation....................................................................................................... 20
1.10 Chondrocyte Differentiation ..................................................................................................... 21
1.11 Osteoblast Differentiation ........................................................................................................ 22
1.11 Osteocyte Differentiation ......................................................................................................... 23
1.12 Osteoclast Differentiation ........................................................................................................ 24 Figure 1I. Stages of Osteoblast Differentiation .................................................................................................. 25
1.13 Selected Molecular Factors Affecting Osteoblast Differentiation ............................................. 26
1.14 Key Signaling Pathways Involved in Bone ................................................................................. 26 Figure 1J. Summary of Genes and Factors Affecting Osteoblast Differentiation ............................................... 30
1.15 Bone Development ................................................................................................................... 31 Figure 1K. Overview of Intramembranous and Endochondral Ossification ....................................................... 32
1.16 Fracture Healing ....................................................................................................................... 33 Figure 1L. Overview of Stages of Fracture Repair .............................................................................................. 35 Figure 1M. Genes Involved in Fracture Repair ................................................................................................... 36
1.17 What is known about the MET-HGF pathway in Bone .............................................................. 37
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2 Research Aims, Hypothesis, and Summary Plan ................................................................. 38
2.1 Rationale .................................................................................................................................... 39
2.2 Hypothesis .................................................................................................................................. 39
2.3 Objectives ................................................................................................................................... 40
2.4 Clinical Significance..................................................................................................................... 42
3 Methods ........................................................................................................................... 43
3.1 Generation of a Met exon 15 splice donor mutant allele by CRISPR/Cas9-mediated genome editing .............................................................................................................................................. 43
3.2 Embryo Microinjection ............................................................................................................... 43
3.3 Confirmation of Exon 15 Skipping ............................................................................................... 43
3.4 Maintenance and Genotyping of Mice........................................................................................ 44
......................................................................................................................................................... 45
......................................................................................................................................................... 45
3.5 Murine Embryonic Fibroblast Generation................................................................................... 45
3.6 MET Pathway Analysis ................................................................................................................ 46
3.7 Activation of MET pathway ........................................................................................................ 46
3.8 Western Blot Analysis ................................................................................................................. 46
3.9 Colony Forming Unit – Fibroblast ............................................................................................... 47
3.10 Colony Forming Unit – ALP and Osteoblast............................................................................... 47
3.11 RT-PCR: Gene Expression Analysis ............................................................................................ 48
3.12 Skeletal Prep Staining ............................................................................................................... 48
3.13 Epiphyseal Growth Plate Staining ............................................................................................. 49
3.14 Visualization and Measurement of Long Bones and Epiphyseal Growth Plates ........................ 49
3.15 Tibia Fracture Generation ......................................................................................................... 50
3.16 Histological Staining of Fracture Callus ..................................................................................... 51
3.17 Statistical Analyses ................................................................................................................... 51
4 Results .............................................................................................................................. 53
4.1 Met Δ15-HET mouse embryonic fibroblast exhibit higher levels of MET protein and upregulated MET signaling ................................................................................................................................... 53
Figure 4.1. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts......................... 55
4.2 Met Δ15-HET mouse embryonic fibroblast exhibit upregulated and prolonged MET signaling after HGF stimulation................................................................................................................................ 56
Figure 4.2. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes............................................................................................ 58 Figure 4.2 Contd. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes.................................................................................... 59
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Figure 4.2 Contd. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes.................................................................................... 61
4.3 Met Δ15-HET MEFs exhibit upregulated levels of β-catenin.......................................................... 62 Figure 4.3 Met Δ15-HET MEFs exhibit gene expression dysregulation resulting in upregulation of β-catenin ... 64 Figure 4.3 Contd. ................................................................................................................................................ 65 Figure 4.4. Met Δ15-HET bone marrow stromal cells exhibit reduced mineralization ability ........................... 68 Figure 4.4 Contd. Met Δ15-HET bone marrow stromal cells exhibit reduced mineralization ability ................... 69
4.5 Mature Met Δ15-HET osteoblasts display dysregulation of osteoblast specific gene markers ...... 70 Figure 4.5 Met Δ15-HET mature osteoblasts display dysregulation of osteoblast specific gene markers ........... 73
4.6 Body weight and preliminary comparisons of WT and Met Δ15-HET mice ................................... 74 Figure 4.6 Body weight and preliminary comparisons of WT and Met Δ15-HET mice ........................................ 75 Figure 4.6 Contd. ................................................................................................................................................ 76 Figure 4.6 Contd. ................................................................................................................................................ 77 Figure 4.6 Contd. ................................................................................................................................................ 78
4.7 Skeletal Staining of P5 WT and Met Δ15-HET mice ....................................................................... 79 Figure 4.7. 5-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates.......................................................................................................................................................... 82 Figure 4.7. 5-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates.......................................................................................................................................................... 83
4.8 Skeletal Staining of P21 WT and Met Δ15-HET mice ..................................................................... 84 Figure 4.8. 21-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates.......................................................................................................................................................... 87 Figure 4.8. 21-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates.......................................................................................................................................................... 88
4.9 Histological analysis of P21 WT and Met Δ15-HET mice epiphyseal growth plate......................... 89 Figure 4.9 Met Δ15-HET mice exhibit no differences in epiphyseal growth plate lengths in comparison to WT littermates.......................................................................................................................................................... 90 Figure 4.9 Contd. Met Δ15-HET mice exhibit no differences in epiphyseal growth plate lengths in comparison to WT littermates ............................................................................................................................................... 91
4.10 Overactivation of MET signaling impairs fracture healing ........................................................ 92 Figure 4.10 Aberrant MET signaling impairs fracture healing in 12-week old mice .......................................... 93 Figure 4.10 Contd. .............................................................................................................................................. 94 Figure 4.10 Contd. .............................................................................................................................................. 95 Figure 4.10 Contd. .............................................................................................................................................. 96 Figure 4.10 Contd. .............................................................................................................................................. 97
5 Discussion ......................................................................................................................... 98
5.1 MetΔ15-HET mice exhibit higher MET protein levels and upregulated MET signaling .................. 98
5.2 Overactivation of MET signaling causes osteoblast differentiation defects .............................. 100
5.3 MetΔ15-HET MEFs exhibit dysregulation of β-catenin activity ................................................... 100
5.4 Overactivation of MET signaling does not affect postnatal skeletal development ................... 101
5.5 MetΔ15-HET mice experience delayed fracture healing ability ................................................... 102
5.6 MetΔ15-HET osteoblasts display dysregulation of β-catenin ...................................................... 104
5.7 Limitations to the Study ........................................................................................................... 107
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5.8 Proposed Mechanism for the Dysregulation of β-catenin Signaling in Osteoblast Differentiation ....................................................................................................................................................... 110
Figure 5.1. Proposed Mechanism for the Dysregulation of β-catenin Signaling in Osteoblast Differentiation ......................................................................................................................................................................... 111
6 Conclusions ..................................................................................................................... 112
7 Future Directions ............................................................................................................. 115
9 References ...................................................................................................................... 118
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List of Figures
Figure 1A Protein expression analysis of healthy bone versus osteofibrous dysplasia bone …………………………………………………………………………………………….13
Figure 1B Regulation of MET Signaling……………………………………………………..19 Figure 1C Osteofibrous Dysplasia lesions at the tibia …………………………………….23
Figure 1D Osteofibrous Dysplasia lesions at the tibia……………………………………..24
Figure 1E Summary of Characteristics of Osteofibrous Dysplasia ………………………25 Figure 1F Genetic Summary of Osteofibrous Dysplasia…………………………………..26
Figure 1G Zonal Architecture of Osteofibrous Dysplasia………………………………….27
Figure 1H MET Exon Skipping Mutation…………………………………………………….31 Figure 1I Overview of Stages of Osteoblast Differentiation……………………………….36
Figure 1J Summary of Genes and Factors Affecting Osteoblast Differentiation………,.41
Figure 1K Overview of Intramembranous and Endochondral Ossification………………43 Figure 1L Overview of Stages of Fracture Repair………………………………………….47
Figure 1M Genes Involved in Fracture Repair……………………………………………...48 Figure 4.1 Protein expression analysis of WT and Met Δ15-HET………………………….66
Figure 4.2 Protein expression analysis of WT and Met Δ15-HET mouse embryonic
fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes ……………..69 Figure 4.13 Met Δ15-HET BMSCs exhibit differential gene expression at day 21 of
osteoblast differentiation………………………………………………………………………75
Figure 4.3 Met Δ15-HET MEFs exhibit gene expression dysregulation resulting in
upregulation of β-catenin……………………………………………………………………...82
Figure 4.4 Met Δ15-HET bone marrow stromal cell exhibit reduced mineralization
ability…………………………………………………………………………………………….87
Figure 4.5 Mature Met Δ15-HET osteoblasts display dysregulation of osteoblast specific
gene markers…………………………………………………………………………………..91
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Figure 4.6 Body weight and preliminary comparisons of WT and Met Δ15-HET mice…..93
Figure 4.7 5-day old Met Δ15-HET mice exhibit no differences in long bone lengths in
comparison WT littermates………………………………………………………………….101
Figure 4.8 21-day old Met Δ15-HET mice exhibit no differences in long bone lengths in
comparison WT littermates………………………………………………………………….106
Figure 4.9 Met Δ15-HET mice exhibit no differences in epiphyseal growth plate lengths in
comparison to WT littermates…………………………………………………………….108
Figure 4.10 Aberrant MET signaling impairs fracture healing in 12-week old mice………………………………………………………………………………………...….111
Figure 5.1 Proposed Mechanism for the Dysregulation of β-catenin Signaling in Osteoblast Differentiation……………………………………………………………………111
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List of Abbreviations
ALP Alkaline Phosphatase
BMP Bone Morphogenetic Protein
BMSC Bone Marrow Stromal Cell
BSP Bone Sialoprotein
CFU-F Colony Forming Unit–Fibroblast
CFU-O Colony Forming Unit-Osteoblast
ColI Collagen Type I
ERK1/2 Extracellular Signal-Regulated Protein Kinases 1 and 2
HGF Hepatocyte Growth Factor
IL Interleukin
LRP6 Low Density Lipoprotein Receptor-Related Protein 6 MAPK Mitogen-
Activated Protein Kinases
MAPK Mitogen-Activated Protein Kinase
M-CSF Macrophage Colony Stimulating Factor
MET Mesenchymal-Epithelial Transition Gene
MMP Matrix Metalloproteinase
MSC Mesenchymal Stem Cell
OCN Osteocalcin
OPG Osteoprotegerin
OPN Osteopontin
OSX Osterix
PI3K Phosphatidylinositol-3-Kinase
RANK Receptor Activator of Nuclear Factor-κβ
RANKL Receptor Activator of Nuclear Factor-κβ Ligand
RUNX2 Runt-related transcription factor 2
SSC Skeletal Stem Cell
SOX9 SRY (Sex Determining Region Y)-Box-9
STAT1 Signal Transducers and Activators of Transcription 1 TD Thanatophoric
Dysplasia
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TGF-β Transforming Growth Factor Beta
TNF-α Tumor Necrosis Factor-α
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1 Introduction
Osteofibrous dysplasia (OFD) is a rare (<0.2% of bone tumors) non-neoplastic condition
which can cause tibial bowing prior to weight bearing in affected individuals. OFD can
also progress to spontaneous non-healing fractures resulting in impaired mobility and
constant pain, significantly affecting quality of life. Through exome sequencing, our lab
and others have previously identified gain-of-function mutations in the tyrosine kinase
MET as the cause of the OFD in four unrelated familial cases. The gain of function MET
mutations results in upregulated MET levels, changes in downstream MET effectors and
changes in β-catenin levels (Fig 1A). We have also shown impaired osteoblast
mineralization and decreased osteoclast function in affected patient bone tissue (Gray
et al., 2015). However, the mechanistic explanation connecting these observations
remains to be clarified.
Surprisingly little is known about MET in fracture repair despite our identification of a
human single gene disorder manifesting abnormal bone repair secondary to gain-of-
function MET mutations. Interestingly, we have also found increased MET expression
in a sub-set of human fracture non-union tissue. Fracture non-union is defined as an
inability to form a bridging callus at the fracture site. Delays in fracture repair causing
non-unions are a significant health problem with approximately 10% of all long bone
fractures healing with delay and 7.5% resulting in non-unions, despite improved surgical
techniques (Mills, Aitken, & Simpson, 2017).
There is currently no cure for OFD, and treatment is limited to pain management of
affected individuals and management of symptoms. Understanding the underlying
mechanisms resulting in reduced bone healing ability may potentially uncover novel
therapeutic targets to treat OFD and improve bone repair in general. The knowledge
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gained from such an endeavor would be transferable to managing fracture non-unions
from a variety of other causes.
Here we look to determine the role of MET signalling in the osteoblast differentiation and
fracture healing. In order to study these processes, we have created a murine mouse
model which reproduces one of the gain-of-function MET mutations identified in OFD.
Our genetic mouse model appears phenotypically normal but has yet to be
characterized and studied in detail. The purpose of this project is to determine the
underlying signaling changes secondary to the Met mutation in our murine model,
characterize the skeletal phenotype and utilize the model to study fracture repair in vivo.
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Actin
β-catenin
Actin
AKT
p-AKT
Actin
MET
Control
Bone OFD
Bone
Figure 1A. Protein expression analysis of healthy bone versus osteofibrous dysplasia bone
Representative western blots of healthy and osteofibrous dysplasia patient bone protein expression analysis. Healthy bone samples were obtained from
the iliac crest of healthy patients during surgical intervention, while osteofibrous dysplasia bone samples were obtained during excision of affected lesional tissue. Osteofibrous dysplasia patients exhibit upregulated
MET protein along with upregulation of its downstream effectors AKT and β-catenin in comparison to healthy bone samples. In addition, osteofibrous dysplasia patients exhibit upregulated phosphorylation at AKT indicative of
upregulated MET signaling.
A
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Regulation of MET Signaling
Regulation of MET Signaling
Figure 1B. Regulation of MET Signaling MET receptor without HGF binding (A). HGF binding to wild-type MET receptor leads to receptor dimerization and auto-phosphorylation. CBL is a E3 ubiquitin ligase involved in downregulation of the receptor (B). Depicts the structure of the exon 14 exclusion in MET-HET
mice resulting in loss of the regulatory juxtamembrane domain and CBL binding site (Y1003) (C).
A B
C
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1.1 Osteofibrous Dysplasia Osteofibrous dysplasia (OFD) (OMIM #607278), also known as ossifying fibroma of the
long bone, is a developmental skeletal disorder characterized by benign fibro-osseous
lesions of the bone (Hitachi et al., 2018). First described as congenital osteitis fibrosa in
1921 by Frangenheim, Kempson later reported affected individuals displayed similar
physical characteristics to fibrous dysplasia. In 1976 Campanacci characterized the
condition as “osteofibrous dysplasia of the tibia and fibula” to describe its anatomical
position, origin of development, and histologic similarities to fibrous dysplasia (Park,
Unni, McLeod, & Pritchard, 1993).
OFD makes up approximately 0.2% of all primary bone tumors, commonly affecting
children under the age of 10, with no sex preference (Most, Sim, & Inwards, 2010) (Fig
1E). OFD is frequently sporadic (not inherited) affecting one limb but has more rarely
been described in familial cases where the disease is bilateral (Fig 1F). OFD is
characterized by the development of radiolucent lesions at the periosteal surface of the
diaphyseal cortex (Fig 1C) (Gray et al., 2015) with a strong predilection for the mid-
diaphysis of the tibia. While almost exclusively occurring in this bone, the fibula, radius
and ulna may also be affected as well (Gray et al., 2015; Hitachi et al., 2018; Kahn,
2003; Park et al., 1993; Taylor et al., 2012). OFD lesions exhibit an elongated shape,
involving 10 – 40% of the bone length (Park et al., 1993).
OFD typically presents as asymptomatic lower leg swelling (Hitachi et al., 2018; Most et
al., 2010). Affected individuals suffer anterior tibial bowing prior to weight bearing and
face an increased risk of fracture. Approximately 31% of affected individuals present
with pain, 19% with pathologic fracture and 13% with tibial bowing. In addition, bone
lesions may be discovered incidentally during examination and imaging for unrelated
issues (Fig 1E) (Gleason et al., 2008; Park et al., 1993). Lesions may resolve
spontaneously during skeletal maturation (Gray et al., 2015; Most et al., 2010), but if a
fracture develops, fracture non-union often complicates the healing process. Fracture
non-union is defined as the inability to form a bridging callus at the fracture site. The
fracture non-union phenotype in OFD appear very similar to the radiological appearance
of a tibial pseudoarthrosis which complicates another single gene disorder,
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Neurofibromatosis Type 1 (NF-1) (OMIM #162200) (Ghadakzadeh, Kannu, Whetstone,
Howard, & Alman, 2016).
Histologically, OFD is characterized by a well-defined zonal architecture (Fig 1G) (Gray
et al., 2015) with a central osteolytic region, abundant in fibrous tissue and spindle-
shaped fibroblast (Gray et al., 2015; Park et al., 1993). More peripherally, cells of
osteoblastic lineage predominate (Gray et al., 2015). Additionally, OFD exhibits
immature woven bone trabeculae at the center of the lesion, which is more abundant
moving outwards (Most et al., 2010). Immature woven bone trabeculae eventually
undergoes anastomosis with one another and merges with the bone of the outer and
inner cortices (Park et al., 1993). Moving towards the outer edge of the lesions, the
trabeculae are larger and greater in number, eventually transitioning to organized
parallel layers of bone, or lamellae.
OFD presents with two fundamental histological patterns: fibrous tissue surrounded by
boney trabeculae rimmed by active osteoblasts and a zonal architecture (Park et al.,
1993). Cells found at the center of the lesions exhibit undifferentiated mesenchymal cell
markers with a subset of these cells positive for both osteoblast and epithelial cell
markers (Gray et al., 2015; Park et al., 1993). Conversely, bridging zones of the osteoid
are rich in surface osteoblasts and embedded osteocytes interspersed between lesions,
pointing to an osteoblast differentiation defect at the lesions (Gray et al., 2015).
The histological appearance of OFD is somewhat similar to that of fibrous dysplasia,
with both entities displaying variably shaped spicules of woven bone separated by
fibrous tissue (Kahn, 2003). OFD’s distinctive osteoblast rimming of the boney
trabeculae and zonal architecture distinguishes it from fibrous dysplasia, thus allowing
histological delineation between the two conditions (Fig 1G) (Gray et al., 2015; Most et
al., 2010).
Radiologically, adamantinomas (ADM) are typically considered in the differential
diagnosis of OFD (Karol, Brown, Wise, & Waldron, 2005; Park et al., 1993).
Adamantinomas are low-grade malignant tumours occurring predominantly in mature
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skeletons with a predilection for the tibial midshaft. Similar to OFD, affected individuals
report pain, anterior bowing of the tibia, fractures and fracture non-unions. Histologically,
OFD and ADM lesions exhibit similar cytokeratin immunoprofiles and cytogenetic
changes (Kahn, 2003). The commonalities in clinical presentation along with similar
histological features previously led many to hypothesize that OFD progresses into ADM,
though this has been proven to be untrue (Park et al., 1993). Unlike ADM, OFD lesions
do not undergo neoplastic transformation and are self-limiting in nature. Therefore, the
delineation between these two conditions is imperative since they vary greatly in
treatment (Most et al., 2010). Untreated or undertreated ADM results in tumour
metastasis and fatality (Kahn, 2003).
8
OFD Lesions at the Tibia
Tibia Tibia
Femur Femur
Figure 1C. Osteofibrous Dysplasia lesions at the tibia
Anteroposterior (A) and lateral (B) radiographs of diaphyseal tibial osteofibrous dysplasia. Osteofibrous dysplasia is characterized by the development of fibro-osseous lesions demonstrated by the differences in radiopacity (arrows). These lesions
commonly develop at the mid-diaphysis of the tibia and in some cases affecting the fibula as well.
A B
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OFD Fractures at the Tibia
Figure 1D. Osteofibrous Dysplasia fractures at the tibia
Anteroposterior radiographs of the left and right tibias of a patient suffering from familial bilateral case of osteofibrous dysplasia. The patient presents with spontaneous non-healing tibial fractures (arrows) prior to weight bearing. Diaphyseal tibial osteofibrous
dysplasia is indicated by the radiolucent gaps.
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Adapted with Permission from Bethapudi et al., 2014
Figure 1E. Summary of Characteristics of Osteofibrous Dysplasia
Brief summary of the common characteristics and presentations of osteofibrous
dysplasia. Table describes the nature, age, sexual preference, commonly affected sites, clinical symptoms, histopathological symptoms, metastatic status and inheritance patterns of the disease.
11
DNA Mutation Protein Mutation Bones Affected Bilateral or Unilateral
Family 1, Familial
c.3010_3028+8del
p.Leu964_Asp1010del
Tibia, Fibula, Ulna
Bilateral
Family 2, Familial
c.3028+1G>T
p.Leu964_Asp1010del
Tibia, Fibula Bilateral
Family 3, Familial
c.3028+1G>T
p.Leu964_Asp1010del
Tibia, Fibula Bilateral
Family 4, Simplex
c.3028+1G>T
p.Leu964_Asp1010del
Tibia, Fibula Bilateral
Sporadic c.3008A>C
p.Tyr1003Ser
Tibia, Fibula Unilateral
Figure 1F. Genetic Summary of Osteofibrous Dysplasia
Selected cases of genetic mutations resulting in osteofibrous dysplasia. All cases described here result in the loss of negat ive feedback regulatory control through either the loss of the juxtamembrane regulatory domain or the key Tyrosine residue 1003, within the juxtamembrane domain required for its function. Loss of this key tyrosine residue results in loss of CBL-mediated
degradation of the receptor post-activation and therefore elevated downstream MET signaling.
Adapted with Permission from Gray et al.,2015
12
Osteofibrous Dysplasia Architecture
Figure 1G. Zonal Architecture of Osteofibrous Dysplasia Lesions Haemotoxylin & Eosin stain of osteofibrous dysplasia lesional tissue excised at surgery from a 9-
year old patient exhibiting familial osteofibrous dysplasia. The stain demonstrates spindle-shaped cells at the center of lesions (arrow) surrounded by osteoid (o). Immunohistochemical staining of lesional tissue for alkaline phosphatase (ALP) show it to be maximally expressed at the periphery
of lesions while absent at the center (arrows). Osteofibrous dysplasia lesions are characterized by rimming of osteoblasts staining positive for the osteoblastic marker, osterix (OSX), and osteoclastic markers tartrate resistant acid phosphatase (TRAP).
Reproduced with Permission from Gray et al., 2015
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1.2 Neurofibromatosis Type 1 (NF-1) NF-1 (OMIM #162200) is associated with a number of different skeletal complications
including the development of tibial bowing and non-healing tibial fractures (tibial
pseudoarthrosis). The clinical presentation of OFD thus shares many similarities with
NF-1. This autosomal dominant disorder is caused by mutations in the neurofibromin-1
(NF1) protein leading to overactivation of RAS signaling and dysregulation of β-catenin.
Similar to OFD, NF-1 is also characterized by defects in osteoblast differentiation; cells
at the NF-1 tibial fracture site do not undergo osteoblast differentiation (Ghadakzadeh et
al., 2016). The development of NF-1 related bone disease has been shown to result
from RAS pathway overactivation and upregulation of downstream ERK1/2 signaling
(Sharma et al., 2013). More recently, Ghadakzadeh et al., (2016) determined that β-
catenin levels were elevated in NF-1 pseudoarthrosis. Furthermore, localized
inactivation of the Wnt pathway with the antagonist Dickkopft-1 (Dkk1) at the fracture
site of Nf1-deficient mice resulted in improved fracture healing and osteoblast
differentiation. This study’s data is consistent with previous findings indicating β-catenin
protein levels must be tightly regulated for normal osteoblast differentiation and any
deviations result in reduced differentiation (Y. Chen et al., 2007). The similarities
between OFD and NF-1 pseudoarthrosis are uncanny and it is therefore reasonable to
hypothesize that these two single gene diseases may share similar disease
mechanisms.
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1.3 Dysregulation of MET to METΔ14 ratios causes disease: Osteofibrous Dysplasia Through exome sequencing our lab in collaboration with other groups has identified
somatic and germline mutations in the MET gene cause familial and simplex
osteofibrous dysplasia (Gray et al., 2015). Curiously, the identical germline MET exon
14 skipping mutation which causes OFD has also been reported as a somatic mutation
in lung and gastric cancer. In lung and gastric cell lines, the mutation results in an
upregulation of a MET downstream effector, mitogen-activated protein kinase (MAPK)
signaling (Heist et al., 2016). MET germline mutations different from those which cause
OFD have previously been shown to cause papillary renal cell carcinomas (OMIM
#605074). In hereditary papillary renal cell carcinoma, the MET receptor tyrosine kinase
domain undergoes autoactivating amino acid substitutions, which promotes cellular
transformation. Autosomal recessive MET mutations are also thought to be the cause of
a rare form of deafness (OMIM #616705).
All identified OFD somatic and germline mutations result in exon 14 skipping (Gray et
al., 2015). As previously mentioned MET’s exon 14 encodes for the regulatory
juxtamembrane domain of the MET receptor where the ubiquitin ligase CBL binds (Fig
1H). One of the identified OFD mutations results in alternative splicing of MET and
exclusion of the juxtamembrane domain pY1003 ubiquitination target. Loss of the CBL
pY1003 docking site leads to reduced internalization and MET receptor degradation.
The receptor’s signaling function remains competent in downstream signal transduction
resulting in aberrant MET signaling (Gray et al., 2015). These results suggest the
mutations underlying OFD stabilize the MET receptor and confer a gain-of-function to
the protein.
1.4 Characterization of Mesenchymal-Epithelial Transition Gene’s Structure The mesenchymal-epithelial transition gene (MET) is a proto-oncogene found on
chromosome 7 band 7q21–q31. Its expression is regulated by Ets (E-twenty six), Pax3
(paired box 3), AP2 (activator protein-2) and Tcf-4 (transcription factor 4) (Boon et al.,
2002). MET is synthesized as a 170 kDa precursor which undergoes post-translational
modification into a mature 190 kDa transmembrane receptor (Parikh et al., 2018). The
protein product of this gene, MET (also called c-MET) tyrosine kinase, is actively
15
expressed during embryogenesis and adulthood on epithelial cells of many organs such
as the liver, blood vessels, muscle, and bone marrow. MET receptor expression can
also be found on cells involved in bone homeostasis such as osteoblasts, osteoclasts,
fibroblasts, and chondrocytes (Gray et al., 2015; Guévremont et al., 2003; Organ &
Tsao, 2011).
The MET receptor is composed of an and -subunit with the subunit composed
solely of extracellular segments while the -subunit encompasses intracellular and
extracellular domains. The extracellular domain of the MET receptor comprises an N
terminal domain, plexin-semaphorin-integrin (PSI) domain, and transmembrane domain.
The N-terminal domain is composed of a 500-residue semaphorin domain, sharing
homologies with domains found in the semaphorin and plexin families. This region
encompasses the extracellular subunit and a portion of the transmembrane -
subunits, linked together by disulfide bonds. The PSI domain follows the semaphorin
domain, spanning approximately 50 residues with four disulfide bonds (Organ et al.,
2011). The PSI domain connects the extracellular region of the receptor to the
transmembrane helix via four immunoglobulin-plexin-transcription (IPT) domains (Organ
et al., 2011).
Intracellularly, the MET receptor is composed of a tyrosine kinase catalytic domain
flanked by a distinctive regulatory juxtamembrane domain and carboxy-terminal
sequences essential for substrate docking and downstream signalling (Liu et al., 2008;
Organ et al., 2011). There are numerous key tyrosine residues along the intracellular
domain of the receptor, critical for normal cellular function and regulation of receptor
signaling. Of note, catalytic tyrosine residues, found within the catalytic domain, Tyr1234
and Tyr1235 positively enhance downstream enzyme activity (Organ & Tsao, 2011).
Negative regulation of the MET-receptor occurs at Tyr1003 of the juxtamembrane
domain, in a negative feedback fashion with the recruitment of ubiquitin ligase casitas B-
lineage lymphoma (CBL) (Gray et al., 2015; Organ et al., 2011) (Fig 1B).
MET is a prototypic member of the receptor tyrosine kinase (RTK) family but
differentiates itself from the rest of the family through its distinct structure. It is also the
16
only high-affinity receptor for hepatocyte growth factor (HGF) also known as scatter
factor (SF) (Christensen et al., 2005). In a similar fashion to the MET receptor, HGF is
synthesized as an inactive precursor and is converted to a two-chain, disulfide-linked
heterodimer through proteolytic cleavage (X. Liu et al., 2008). HGF is sequestered in its
active form in the extracellular matrices of most tissues by heparin-like proteoglycans
(T. Kobayashi et al., 1994). Cells of mesenchymal origin, such as osteoblasts,
osteoclasts, and fibroblasts are the main source of HGF production, with osteoblasts
being the preeminent source of HGF in bones (Grano et al., 2002; Vallet et al., 2016).
RTKs have been shown to be intricately involved in key processes of mammalian
development, cell function and tissue homeostasis. Dysregulation of RTKs affect crucial
processes such as cell growth and survival, organ morphogenesis, neovascularization
and tissue repair and regeneration (Christensen et al., 2005).
1.5 MET Signaling and Function Stimulation of the MET receptor by HGF, results in receptor dimerization or
multimerization and phosphorylation of tyrosine residues along the juxtamembrane,
catalytic and cytoplasmic tail domains allowing for regulation of internalization, catalytic
activity, and docking of regulatory substrates respectively (X. Liu et al., 2008; Naldini et
al., 1991). Some of these phosphorylation events result in activation of cellular growth
programs, promoting mitogenesis, motility, invasion and morphogenesis (Frampton et
al., 2015). Upon activation, phosphorylation of tyrosine residues found in the catalytic
loop of the kinase domain, Tyr1234 and Tyr1235, ensues (Birchmeier & Gherardi,
1998). Phosphorylation at these sites triggers a conformational change in the receptor
resulting in phosphorylation events at Tyr1349 and Tyr1356 of the C-terminal domain
allowing them to act as docking sites for adaptor and effector proteins with SH2-
containing domains, phosphotyrosine binding (PTB) domains, and MET binding
domains (MBDs) (Sattler et al., 2009). These docking site are crucial for activating
downstream pathways such as RAS/mitogen-activated protein kinase (RAS/MAPK),
phosphatidylinositol 3-kinase/protein kinase B (PI3K/AKT), signal transducer and
activator of transcription (STATs), phospholipase C (PLC), and proto-oncogene
tyrosine kinase Src (c-Src) (Furge et al., 2000; Ponzetto et al., 1994).
17
The tyrosine kinase domain of the MET receptor is also responsible for the initiation of
various downstream signalling pathways, chiefly through the following major pathway
regulators: MAPK, PI3K/AKT, and STAT (Parikh et al., 2018). Under normal
physiological conditions these pathways regulate cell proliferation, motility, and survival
among other numerous cellular processes (Parikh et al., 2018). MET activation also has
a crucial role in epithelial-mesenchymal interactions during injury repair (Lesko, 2007).
MET mediates epithelial cell dissociation, migration towards the site of injury,
proliferation and reconstruction of the epidermal layer (Lesko, 2007). Additional post-
translational and receptor domain modifications further contribute to the regulation of
biological functions downstream of the activated MET receptor (Sattler et al., 2009).
1.6 MET’s Role in Development Interactions between HGF and its receptor have been previously shown to play an
essential role in mammalian embryogenesis, muscle development, nervous system
formation, hematopoietic cell differentiation, bone remodeling, angiogenesis and
organization of three-dimensional tubular structures (e.g. renal tubular cells) (Birchmeier
et al., 1998; Comoglio et al., 2001). In embryogenesis, HGF mediated MET signaling
produces instructions critical for growth and survival of hepatocytes and trophoblast
cells. Hgf and Met knockout embryos display underdeveloped liver and placental
labyrinths caused by defects in epithelial-mesenchymal transition (Schmidt et al., 1995).
This results in early embryonic lethality due to compromised placental mediated
exchange between maternal and fetal blood (Schmidt et al., 1995; Trusolino, Bertotti, &
Comoglio, 2010). These mice also exhibit reduced hypaxial muscles, muscles ventral to
the horizontal septum of the vertebrae, due to loss of MET mediated proliferation and
motility (Lesko, 2007). Lastly, loss of MET signalling’s anti-apoptotic effects leads to
increased apoptosis in neuronal cells leading to impaired axon bundling and neuronal
defects (Trusolino et al., 2010).
1.7 CBL-Dependent Regulation of MET Signaling In addition to downstream pathway activation and signalling post-ligand binding, the
HGF/MET receptor complex is quickly internalized post-phosphorylation of Tyr1003 in
the juxtamembrane domain of the receptor into clathrin-coated vesicles (Abella et al.,
18
2005). This phosphorylation modification recruits the Casitas B-lineage Lymphoma
(CBL) protein, an E3 ubiquitin-protein ligase, which facilitates polyubiquitination of the
receptor complex creating ubiquitin recognition motifs for trafficking and degradation of
the internalized receptor (Baldanzi et al., 2014; Sattler et al., 2009). CBL also acts as an
adaptor for endophilin, a protein crucial for receptor internalization, trafficking and
degradation (Kjaerulff et al., 2011). Internalized receptors continue on to two distinct
fates: they are recycled back to the plasma membrane or degraded via the lysosomal
pathway (Teis et al., 2003). This negative feedback mechanism allows for modulation of
MET signaling and associated downstream signalling. Disruption of this mechanism
could result in cellular transformation through increased recycling of the receptor back to
the plasma membrane causing hyper-signalling or increased degradation of the
internalized receptor leading to hypo-signalling (Abella et al., 2005).
1.8 Other Means of Regulating MET Signaling Ubiquitin mediated lysosomal degradation of the MET receptor is the major determinant
of receptor sensitivity and regulator of signal activity, though it is not the only regulatory
mechanism (Trusolino et al., 2010). Numerous cytokines such as interleukin-1, -6 and
transforming growth factor β (TGFβ) have been shown to enhance transcriptional
activity of HGF in fibroblasts and macrophages and MET in epithelial cells. These
factors also upregulate the production of plasmin and matriptase proteases, to enhance
cleavage of pro-HGF to its active form (Bhowmick, Neilson, & Moses, 2004;
Michalopoulos & DeFrances, 2005). Proteasomal degradation can also occur through
CBL-interacting protein 85 (CiN85 or SH3KbP1) endophilins, which acts as a scaffold
molecule for CBL to promote internalization of the MET receptor through contact
dependent means (Oved & Yarden, 2002). Lastly, the MET receptor can be regulated
through proteolytic means as well. A disintegrin and metalloprotease (AdAM) mediated
shedding of the extracellular domain can also regulate receptor stability, releasing a
soluble N-terminal fragment with a cytoplasmic tail remaining anchored to the
membrane. The remaining cytoplasmic tail is proteolytically cleaved by γ-secretase and
degraded by proteasomal means (Foveau et al., 2009). Proteolytic degradation does not
require activation of the receptor like CBL-mediated degradation does. In addition, it
leaves behind the extracellular domain of the receptor, allowing for sequestration of
19
HGF and impairs dimerization with full-length MET receptors causing inhibition of MET
signaling (Trusolino et al., 2010; Zhang, Graveel, Shinomiya, & Vande Woude, 2004).
1.9 Mouse Model of Osteofibrous Dysplasia The regulatory juxtamembrane domain found in the human MET receptor is coded by
exon 15 in mice (exon 14 in humans). In order to study OFD, we created a MetΔ15-HET
mouse model (through the CRISPR-Cas9 system), with an in-frame deletion of the
juxtamembrane segment containing Y1003 to replicate the mutation seen in human
disease (Fig 1B). Mice homozygous for the MetΔ15 mutation are embryonically lethal at
around embryonic day 10.5 (E10.5) and are not the scope of this project. Heterozygous
MetΔ15 mice are used in this study. MetΔ15-HET mice are viable and fertile and do not
develop liver abnormalities or tumors. A semi-stabilized tibial fracture model is used to
induce representative mid-diaphyseal fractures seen in humans since Met Δ15-HET mice
do not develop spontaneous fractures.
20
Figure 1H. MET Exon 14 Skipping Mutation Schematic diagram of MET showing a point mutation of G>T at the exon-intron boundary of Exon
14 resulting in the skipping of Exon 14. Exon 14 codes for a regulatory juxtamembrane domain crucial in negative feedback control of MET signaling post-activation. This mutation was found in family 2, 3 and 4 as described in Figure 1F.
21
1.10 Chondrocyte Differentiation
Chondrogenesis is a critical process in the development and maintenance of healthy
skeletal physiology in vertebrates during embryonic and postnatal development
(Kozhemyakina, Lassar, & Zelzer, 2015). Though chondrocytes also play a critical role
in bone physiology and endochondral ossification, this study is focused on the effects of
aberrant MET signaling on osteoblasts and therefore the review of chondrogenesis is
more rudimentary in nature.
Chondrogenesis involves the creation of chondrocytes through progressive steps in the
commitment and differentiation of mesenchymal stem cells (MSCs) to produce new
cartilage tissue. MSCs are unique in that they are pluripotent, meaning they have the
ability to differentiate into a limited variety of cell types, such as adipocytes, osteoblast,
chondrocytes and endothelial cells (Pez Ponte et al., 2007). Chondrogenesis starts with
the migration and condensation of mesenchymal stem cells (mediated by cell-cell
interactions) at prospective skeleton sites (Oberlender & Tuan, 1994; Sun & Beier,
2014). Members of the SOX transcription factor family commit MSCs to immature
chondrocytes after cellular condensation has occurred. This is followed by proliferation
and hypertrophy of the cells. At the end of chondrogenesis, terminal chondrocytes can
either undergo apoptosis or transdifferentiate to osteoblasts. This transdifferentiation
process provides an additional source of osteoblasts distinct from the periosteum
derived pool during endochondral ossification (Yang, Tsang, Tang, Chan, & Cheah,
2016; Zhou et al., 2014).
22
1.11 Osteoblast Differentiation MSCs have been shown to be heavily involved in the fracture healing process and
regeneration of bone post-injury (Garg et al., 2017). Under the influence of injury-
mediated chemokines, MSCs found in bone marrow, periosteum, walls of blood vessels
and circulation, and muscle are recruited to the fracture site. The divergent
differentiation MSCs undergo is strongly influenced and regulated by several key growth
factors though the full understanding of the mechanism behind this recruitment remains
unclear (Ito, 2011). (Kratchmarova et al., 2005).
Osteoblast are cells of mesenchymal origin which play a critical role in maintaining bone
homeostasis. Their involvement can be broken down into two main categories involving
the formation and resorption of bone. In their formative role, osteoblasts produce
extracellular matrix proteins, regulate matrix mineralization and control bone remodeling
(Corrado et al., 2017). Osteoblasts fulfil their role in bone resorption through expression
of RANKL and OPG. RANKL is an essential factor for the recruitment, differentiation,
activation and survival of osteoclastic cells through its interactions with the RANK
receptor found on immature and mature osteoclastic cells (Langdahl, Ferrari, &
Dempster, 2016). OPG is a soluble receptor for RANKL, which inhibits the promotion of
osteoclastic differentiation and activity, by sequestering and preventing RANK-RANKL
interactions (Langdahl et al., 2016; Wei Liu & Zhang, 2015).
Osteoblast commitment, differentiation and growth is controlled by several local and
systemic pathways which act through paracrine and/or autocrine manners in order to
regulate the activity of key transcriptional factors (Aubin et al., 1995). Some of these
factors include bone morphogenetic proteins (BMPs), hedgehog proteins, cell growth
factors, hormones, cytokine modulators and canonical Wnt/β-catenin signaling (Vimalraj
et al., 2015). The effect of these factors in promoting or inhibiting osteoblast
differentiation relies on the stage of maturation and development (Canalis et al., 1992).
This complex differentiation process can largely defined by three distinct stages:
proliferation, maturation and extra-cellular matrix synthesis, and matrix mineralization
(Neve, Corrado, & Cantatore, 2011) (Fig 1I).
23
Proliferation:
Mesenchymal stem cells become committed to the osteoblastic differentiation pathway
under Runx2 expression. RUNX2 directs mesenchymal progenitor cells to begin
differentiation to pre-osteoblasts, while inhibiting their differentiation to adipocytes and
chondrocytes (Toshihisa Komori, 2006). Proliferating osteoblasts begin expressing
fibronectin, TGFβ Receptor 1 and osteopontin (Rutkovskiy, Stensløkken, & Vaage,
2016), markers of early stage osteoblast differentiation.
Maturation and Extracellular Matrix Synthesis:
As pre-osteoblasts mature, they exit the cell cycle and differentiate to immature
osteoblasts which produce extracellular matrix proteins such as alkaline phosphatase
and type 1 collagen (Col1a1). The immature osteoblast stage is characterized by
maximal expression of alkaline phosphatase (ALP) a membrane-bound enzyme
required for bone matrix mineralization (Huang, 2007). The expression of ALP occurs
post type 1 collagen production and reaches maximal levels just prior to mineralization.
ALP can also be found in other tissues such as the liver, intestines, spleen and kidneys.
In bone, it is a marker of late-stage osteoblast differentiation (Heino & Hentunen, 2008).
Matrix Mineralization:
Mature osteoblast are cuboidal in shape, with enlarged Golgi complexes and a well-
defined endoplasmic reticulum (Heino & Hentunen, 2008). These cells begin expression
and secretion of the following matrix markers: osteopontin, bone sialoprotein and
osteocalcin (Huang, 2007).
1.11 Osteocyte Differentiation Osteoblasts which have embedded themselves within the bone matrix differentiate to
osteocytes. Osteocytes are spider-shaped cells derived from MSCs which have
undergone the osteoblast differentiation process. During the transition from osteoblast
to osteocyte, the once prominent Golgi complexes and endoplasmic reticulum decrease
in size (Capulli, Paone, & Rucci, 2014). Osteoblastic markers such as BSP, OC, ALP
and type 1 collagen which were once abundantly expressed are downregulated and
replaced with osteocytic markers (Bellido, 2014). Osteocytes are the most abundant
24
cells found in bone, making up 90 – 95% of the total number of cells. Osteocytes
possess branched processes which extend throughout the mineralized matrix, thus
allowing them to form an interconnected network. This allows them to sense and
respond to stimuli to regulate bone remodelling and adaptation. These effects are
mediated through both cell-cell interactions and signaling with osteoclasts and
osteoblasts (Goldring, 2015).
1.12 Osteoclast Differentiation Osteoclasts are multinucleated bone reabsorbing cells differentiated from hematopoietic
stem cells found in the bone marrow. Osteoclasts rely on the presence of osteoblast
and osteocyte secreted macrophage-colony stimulating factor (M-CSF) and RANKL for
their survival, proliferation, differentiation and activation (Park-Min, 2018). In the
presence of M-CSF, expression of the RANK receptor is increased in osteoclast
precursor cells allowing for increased sensitivity to RANKL activation (Katagiri &
Takahashi, 2002). RANKL-RANK signaling promotes the differentiation and fusion of
preosteoclasts to multinucleated mature osteoclasts. Osteoclasts exert their resorptive
function through attachment to the bone matrix and forming an integrin mediated seal
around the resorption zone, to separate the resorptive microenvironment from the
extracellular space (Soysa & Alles, 2016). This is followed by the targeted secretion of
HCl to dissolve the hydroxyapatite crystals allowing for secreted proteolytic enzymes to
degrade the collagenous bone matrix. Degraded products are removed via a
transcytosis pathway from the ruffled border of the sealing zone to the secretory
domain, where it is released into the extracellular matrix (Väänänen, Zhao, Mulari, &
Halleen, 2000).
25
Figure 1I. Stages of Osteoblast Differentiation Brief overview of the stages of osteoblast differentiation and select markers secreted at the indicated stages. Progenitor cells are
committed to the osteo-progenitor fate by RUNX2 expression. Osteo-progenitor cells transition to immature osteoblast under the regulation of RUNX2 and OSX. Immature osteoblast begin secretion of extracellular proteins such as alkaline phosphatase, bone sialoprotein and type 1 collagen. These factors are critical for matrix mineralization. Mature osteoblast express and secret Osteonectin, osteopontin, and
osteocalcin during the mineralization stage. As mature osteoblast become embedded within the bone matrix, they differentiate to
osteocytes.
26
1.13 Selected Molecular Factors Affecting Osteoblast Differentiation
RUNX2 is a key transcription factor in bone formation
The RUNX family of transcription factors are defined by a runt DNA binding domain.
RUNX2 functions as a platform protein to regulate bone specific genes, coregulatory
proteins, and chromatin remodelling factors. RUNX2 has the ability to enhance DNA
binding of itself, through upregulation of PI3K and AKT expression (Thomas et al.,
2004). RUNX2 transcriptional activity can also be regulated through phosphorylation
events, of note, ERK1/2 mediated phosphorylation of RUNX2 enhances its association
with coactivators to upregulate osteoblast specific genes (Greenblatt et al., 2010).
Mice deficient in Runx2 exhibit a complete lack of bone (T. Komori et al., 1997; Otto et
al., 1997). In contrast, Runx2 over-expression induces non-osteogenic cells to express
osteoblastic markers (Yamaguchi et al., 2000). In addition to its role in bone formation,
RUNX2 regulates RANKL expression, an inducer of osteoclastogenesis (Vimalraj et al.,
2015). RUNX2 can also form complexes with STAT1, TWIST and HEY1 to disrupt
binding of key downstream osteogenic gene promoter regions, resulting in inhibition of
osteoblast differentiation (Bialek et al., 2004; S. Kim et al., 2003; Thomas et al., 2004).
Osterix is important in the differentiation of pre-osteoblasts to immature
osteoblasts
The commitment of pre-osteoblasts to immature osteoblasts is partly regulated by
osterix (OSX). Osterix null mice are completely void of osteoblasts, indicating the
essential role of this transcription factor in osteoblast differentiation (Huang, 2007; Koga
et al., 2005). Osterix’s role in osteoblast differentiation is downstream of RUNX2; Runx2
expression is present in the mesenchymal stem cells of osterix null mice, but osterix
expression is absent in Runx2 null mice (Huang, 2007; Toshihisa Komori, 2006).
1.14 Key Signaling Pathways Involved in Bone WNT Pathway
The canonical WNT pathway is important in the formation of bone and a tight regulation
of its activity is required to ensure the terminal differentiation of osteoblasts. Canonical
27
WNT signaling involves the stabilization of β-catenin by inhibiting glycogen synthase
kinase 3 beta (GSK-3β), a critical component of the β-catenin degradation complex
(Nelson & Nusse, 2004; Stamos & Weis, 2013). GSK-3β is also inhibited by
adrenomedullin (ADM); ADM is produced in bone tissues and acts as an important
regulator of osteoblast cells through modulation of Wnt signaling (Lausson & Cressent,
2011). Inhibition of GSK-3β mediated β-catenin phosphorylation results in the
accumulation of unphosphorylated β-catenin in the cytoplasm, translocation of β-catenin
to the cytosol and activation of downstream gene targets such as Axin2. Axin2 is a
scaffolding protein which promotes the phosphorylation of β-catenin by GSK-3β and its
subsequent degradation (Jho et al., 2002).
β-catenin is essential for osteoblast differentiation in both endochondral and
intramembranous ossification. While the inactivation of β-catenin blocks osteoblast
differentiation and results in a chondrocyte fate(Day et al., 2005), β-catenin appears not
to be crucial in the initial commitment stages of osteoblast differentiat ion. β-catenin
regulates the pre-osteoblast to immature osteoblast transition stages (Hill et al., 2005;
Hu, 2004) by enhancing Runx2 expression, thus promoting commitment to the
osteoblastic lineage and inhibiting the adipocyte and chondrocyte fates (Gaur et al.,
2005).
RAS-MEK1/2-ERK1/2 Signaling
RAS proteins act as molecular switches for many signaling cascades involved in cell
biology. RAS activation occurs downstream of MET receptor signaling and has been
previously shown to upregulate osteoprogenitor cell proliferation resulting in increased
osteoblastic and stromal cell descendants (Papaioannou, Mirzamohammadi, &
Kobayashi, 2016). RAS activation and associated activation of PI3K, regulates AKT and
ERK1/2 signaling activity to stimulate osteoblast differentiation (Ghosh-Choudhury,
Mandal, & Choudhury, 2007; Yamashita et al., 2008). RAS protein activator like 3
(RASAL3) is a part of the Ras protein family and a negative regulator of RAS signaling.
RASAL3 inactivates RAS proteins by converting active RAS-GTP to inactive RAS-GDP.
Knockdown of RASAL3 increases RAS-GTP levels and phosphorylated ERK, a marker
of RAS activity (Muro et al., 2015).
28
Downstream effectors of RAS signaling such as mitogen-activated protein kinase kinase
(MEK1/2 or MAP2K) and mitogen-activated protein kinase 1 (ERK1/2 or MAPK1),
regulate Runx2 expression through modulation of canonical Wnt signaling (Li, Ge, &
Franceschi, 2017; R.T., C., G., H., & D., 2009). RAS signaling can also increase
stabilization and modulation of β-catenin directly (Jeong, Ro, & Choi, 2018). Through
these studies, it is clear that the regulation of RAS and its downstream effectors is
crucial for the osteoblast differentiation process.
PI3K-AKT-mTOR Signaling
Activation of phosphoinositide 3-kinase (PI3K) occurs directly downstream of the MET
receptor, its activation creates a docking site for protein kinase B (AKT) thus allowing
access for PI3K to activate AKT through phosphorylation. AKT directly phosphorylates
and activates mammalian target of rapamycin (mTOR), in the mammalian target of
rapamycin complex 1 (mTORC1) (Hemmings & Restuccia, 2012). PI3K-AKT-mTOR
kinase pathway activation directly promotes osteoblast differentiation and proliferation
through. There is also a connection with β-catenin since activated AKT signalling
phosphorylates β-catenin at Serine 552 to enhance its stabilization and nuclear
translocation, where it upregulates the expression of Runx2 and Osterix (Raucci,
Bellosta, Grassi, Basilico, & Mansukhani, 2008). AKT activation of mTOR also
upregulates Runx2 expression(Dai et al., 2017). Thus, it is clear that regulation of PI3K
signaling and its downstream effectors is crucial for osteoblast differentiation.
29
30
Figure 1J. Summary of Genes and Factors Affecting Osteoblast Differentiation
Effects of selected published genes and factors affecting osteoblast differentiation and the model utilized to determine their effect.
31
1.15 Bone Development
The adult skeleton consists of 206 bones subcategorized into the axial skeleton (skull,
ribs and spine) and appendicular skeleton. Though morphologically and functionally
diverse, all bones form through either intramembranous and endochondral ossification.
Intramembranous ossification (IO) forms the flat bones which make up the skull, the
mandible and the clavicles. IO is characterized by the direct differentiation of
mesenchymal stem cells into osteoblasts, eventually producing bone (Fig 1K) (E. M.
Thompson, Matsiko, Farrell, Kelly, & O’Brien, 2015). Appendicular bones such as the
tibia, the preeminent site affected by OFD, are formed through endochondral
ossification (EO) and therefore will be the main focus of this section. EO relies on the
formation of an intermediate cartilaginous template which is replaced by bone through a
series of processes (Shapiro, 2008) (Fig 1K). EO begins with the condensation and
differentiation of mesenchymal stem cells to chondrocytes, which secrete proteoglycans
and extracellular matrix forming the hyaline cartilaginous template (E. M. Thompson et
al., 2015). Once the cartilaginous structure is completed, chondrocytes stop secretion
and while embedded in the cartilage template, begin to proliferate (Farrell et al., 2011).
Chondrocytes then align into lacunae columns, mature and undergo hypertrophy
followed by apoptosis or transdifferentiation to osteoblast (Long & Ornitz, 2013; Yang et
al., 2016). This is followed by vascularization and calcification of the transitory cartilage
template (Long & Ornitz, 2013). While the cartilage template continues to grow
interstitially, mesenchymal stem cells of the perichondrium begin differentiation to
osteoblasts, secreting osteoid to make a cortical bone collar, transforming the
perichondrium into the periosteum (E. M. Thompson et al., 2015). This cortical bone
acts as a dense protective outer layer of bone surrounding the inner cavity (E. M.
Thompson et al., 2015). Osteoprogenitor cells occupying the cavity at the center of the
cartilage template formed by chondroclasts will now differentiate to osteoblasts and form
a primary ossification center. These osteoblasts produce and secrete osteoid forming
spongy trabecular bone.
32
Figure 1K. Overview of Intramembranous and Endochondral Ossification Intramembranous ossification occurs with the direct differentiation of mesenchymal stem cells to osteoblasts and subsequent f ormation of an ossification center and bone formation (A). Endochondral ossification begins with the condensation and differentiation of mesenchymal stem cells to chondrocytes which forms a cartilaginous template resembling the final bone’s structure. In the final stages of endochondral
ossification osteoblasts mineralize this cartilaginous template resulting in the formation of new bone (B).
33
1.16 Fracture Healing
A fracture is defined by a lack of continuity in the bone. Fracture healing under the right
circumstances is a robust process. However, approximately 4 to 10% of all fractures
develop in a non-union fashion (Hak et al., 2014). A fracture non-union is the incomplete
consolidation of a fracture, without the formation of a boney and cartilaginous bridge
across the disconnected ends of bone also known as a callus, after approximately 4 – 8
months post-injury (Tall, 2018).
Fracture non-unions present in two distinct forms; hypertrophic and atrophic non-unions
(Kostenuik & Mirza, 2017). Hypertrophic non-unions are caused by an unstable fracture
site due to excessive fracture site motion resulting in large amounts of non-bridging
callus at the fracture site (Hak, 2011; Tall, 2018). Atrophic non-unions develop due to a
deficiency in vascularization post-fracture caused by injury to the soft-tissue envelope
depriving fractures of normal blood supply and reduced vascular in-growth resulting in
fibrous tissue at the fracture site (Hak, 2011; Hankenson, Dishowitz, Gray, & Schenker,
2011; Tall, 2018).
Fracture non-unions result in poor mobility compromising ones activities of daily living,
resulting in a loss of productivity associated mental health burdens (Hak et al., 2014).
While the management of a fracture non-union is largely surgical there have been
promising results utilizing recombinant human bone morphogenetic proteins (BMPs) to
enhance bone formation and improve bridging of fracture callus (Kostenuik & Mirza,
2017). There remains significant challenges associated with sequestering BMPs at the
fracture site and an opportunity to develop new treatment modalities (Marsell & Einhorn,
2011).
Primary, or direct healing is extremely rare and only typically occurs with rigid support of
the fracture and formation of cortical bone without a transitory cartilaginous intermediate
(Z. Thompson, Miclau, Hu, & Helms, 2002). Secondary, or indirect healing is much more
common as it does not rely on stabilization and rigidity of the fracture site (Marsell &
Einhorn, 2011). Fracture healing of bones in the appendicular skeleton, such as the
34
tibia, heal through secondary healing; a combination of intramembranous and
endochondral ossification (Runyan & Gabrick, 2017).
Secondary fracture healing can be described in the 3 following phases (Fig 1L):
1) Reactive Phase: Inflammation occurs directly after a fracture breaks the blood vessels
and disrupts the blood supply to the bone marrow, periosteum, bone and surrounding
soft tissue (Jha, Blau, & Bhattacharyya, 2016). First, a hematoma forms secondary to
the activation of the plasma coagulation cascade and exposure of platelets to the
extravascular environment (Loi et al., 2016). These events in addition to the local
microenvironment, trigger the condensation of mesenchymal cells from multiple sources
(periosteum, endosteum, and bone marrow) and their differentiation to chondrocytes
and osteoblasts (Loi et al., 2016). Since muscle and periosteum overlying the bone are
sources of mesenchymal stem cells, fractures with poor soft tissue coverage may result
in delayed or inhibited healing (Chan, Harry, Williams, & Nanchahal, 2012).
2) Reparative Phase: The hematoma, acts as a scaffold for callus formation, in which the
recently recruited mesenchymal stem cells differentiate to chondrocytes and osteoblasts
(Jha et al., 2016). Chondrocytes begin laying down cartilage, forming a soft callus which
is crucial in bridging the gap and restoring some of the bone’s original strength and
temporary stability. In the last stages of soft callus formation, chondrocytes mineralize
the soft matrix. IO occurs at regions of healthy vasculature, directly adjacent to the distal
and proximal ends of the fracture, resulting in formation of a hard callus (Marsell &
Einhorn, 2011). In parallel, EO occurs between the ends of the fracture and external to
periosteal sites (Marsell & Einhorn, 2011).
3) Remodeling Phase: This phase is characterized by osteoblast and osteoclast
cooperation and communication mediated through RANK-RANKL interactions and
osteoprotegerin (OPG) expression. The hard callus is gradually remodeled by
osteoclasts and osteoblasts until normal bone geometry and integrity is re-established
(Kostenuik & Mirza, 2017).
35
Figure 1L. Overview of Stages of Fracture Repair
Brief overview of the stages of fracture repair. There are four stages in the repair of a broken bone: (A) Formation of a hematoma post injury. (B) Formation of a fibrocartilaginous soft callus. (C) The formation of a bony hard callus and (D) Remodelling and addition of
compact bone.
36
Figure 1M. Genes Involved in Fracture Repair Schematic diagram of the cells important in endochondral ossification mediated fracture repair; chondrocytes, osteoblasts and osteoclast.
Osteoblasts and osteoclasts have the ability to express MET and its one and only ligand, HGF. Furthermore, previous studies have implicated downstream effectors of MET, RAS, MAPK, and AKT to be involved in the regulation of key transcription factors involved in the differentiation process of these cells.
37
1.17 What is known about the MET-HGF pathway in Bone MET and its only ligand HGF are expressed in osteoblasts and osteoclasts. HGF may
be a coupling factor between osteoblasts and osteoclasts (Fig 1M) (Grano et al., 2002;
Lee et al., 2018). The activation of the MET receptor in osteoblasts results in cell cycle
progression. In vitro studies show loss of MET in osteoblast resulted in suppressed
osteoclastogenesis (Lee et al., 2018). Furthermore, exogenous HGF stimulation of
human mesenchymal stem cells inhibited bone-morphogenic protein-2 (BMP-2) and
osteopontin, and other markers of osteoblast differentiation, via the MET receptor and
its downstream effectors (Standal et al., 2007). Others have reported HGF stimulation of
human mesenchymal stem cells promotes osteogenic marker expression and the
inhibition of it, results in reduced matrix mineralization (Aenlle, Curtis, Roos, & Howard,
2014; H. Te Chen, Tsou, Chang, & Tang, 2012; Tsai, Huang, Yang, & Tang, 2012).
While these data imply a regulatory role for MET signaling and HGF in the osteoblast
differentiation process, the explanation for the contradictory results remains unclear.
38
2 Research Aims, Hypothesis, and Summary Plan
MET signaling regulates a diverse number of functions including but not limited to
proliferation and cell survival. MET protein levels have been shown to be elevated in the
fracture non-union tissue of OFD patients (Gray et al., 2015). Previously published data
also indicate abnormalities of osteoblast differentiation in OFD patient cells. MET has
previously been shown to be expressed at the periosteum of long bones, a crucial
source of mesenchymal stem cells during fracture repair (Gray et al., 2015; Mara et al.,
2011). Its’ expression can be found in osteoblasts, osteoclasts, chondrocytes and
fibroblasts, which all play their own crucial role in fracture repair (Fig 1M) (Grano et al.,
2002; Vallet et al., 2016). Our group and others have identified three different germline
MET mutations having a causative role in OFD; these novel autosomal dominant gain-
of-function mutations all result in the loss of MET’s juxtamembrane domain, which is
crucial for downregulation of MET signaling. The cause of the reduced fracture healing
seen in OFD patients is still largely unknown therefore investigating the MET receptor
and osteoblast behaviour can provide insight into new mechanisms by which long bones
grow and heal. Taken together, these findings suggest MET signaling may have a
critical role in normal bone biology and fracture repair. We hypothesize that gain-of-
function MET mutations result in delayed bone repair ability due to reduced
osteoblast differentiation. Since the cause of the reduced fracture healing seen in
OFD patients is still largely unknown this study will focus on the role of MET in
osteoblasts and during fracture repair.
The following experiments will be completed to test the hypothesis. Pathway analysis at
the protein level will be conducted to visualize any dysregulations in key signaling
pathways downstream of MET such as RAS, AKT, and β-catenin which have been
previously shown to regulate osteoblast differentiation (Y. Chen et al., 2007;
Papaioannou et al., 2016). This is followed by examination of osteoblast differentiation
in vitro through colony forming unit (CFU) assays to determine osteogenic potential of
WT and Met Δ15-HET bone marrow stromal cells (BMSCs). The length of the long bones
and epiphyseal growth plates of WT and MetΔ15-HET mice will then be examined to
expose any gross skeletal phenotypes present in mutants. We will shift our focus to
39
examining fracture healing in vivo. We will utilize a semi-stabilized tibial fracture model
to induce fractures since our genetic mouse model does not develop spontaneous
fractures. Fractures generated through this method have been shown to heal through
intramembranous and endochondral ossification (Y. Chen et al., 2007).
2.1 Rationale We have identified a unique human condition in which gain-of-function MET mutations
result in non-healing fractures with increased MET expression at the bone fracture site.
In mice MET is expressed at the periosteum, which is a thin membranous structure
involved in the maintenance of cortical bone and fracture healing. Surprisingly, little else
is known about MET in fracture repair or its role in bone cell differentiation. Given that
many of the downstream pathways of MET have been previously implicated in delayed
fracture healing and osteoblast differentiation, dysregulation of MET signaling may be
responsible for the delays in fracture healing seen in OFD patients (Y. Chen et al., 2007;
Gray et al., 2015). In addition, the inhibition of the MET receptor has previously been
shown to stimulate osteoblast differentiation and bone regeneration in vitro by multiple
groups (Fioramonti et al., 2017; J. W. Kim et al., 2017; Kokabu, 2013; Shibasaki et al.,
2015). However, the effects of aberrant and upregulated MET signaling on osteoblast
differentiation and bone repair is still unclear.
2.2 Hypothesis Previous research in the Kannu lab has implicated MET exon 14 skipping mutations to
cause osteofibrous dysplasia in humans. Forced induction of this exon-exclusion event
results in retarded osteoblast differentiation and reduced bone-matrix mineralization in
vitro but the mechanism underlying this phenotype is still largely unknown (Gray et al.,
2015). We hypothesize that gain-of-function MET mutations result in delayed bone
repair ability due to reduced osteoblast differentiation.
40
2.3 Objectives
Objective 1: Investigating MET pathway irregularities in Met Δ15-HET mice
The first aim of this study was to investigate whether there are similar MET pathway
irregularities in Met Δ15-HET mice as what is seen in human OFD. The utilization of a
mouse model was critical as patient samples and cells are difficult to obtain. Pathway
analysis was conducted utilizing WT and Met Δ15-HET mouse embryonic fibroblast cell
lines as they share many similarities with osteoblasts: both are a part of the connective-
tissue cells family, secrete the MET receptor and HGF, collagenous extracellular matrix
rich in type-1 collagen and are responsible for the structural framework of the body
(Grano et al., 2002; Vallet et al., 2016). In addition, patient samples are few and far in-
between, and are extremely difficult to come by. For these reasons we felt mouse
embryonic fibroblasts were a suitable alternative in place of osteoblasts to perform the
MET signalling analysis reported here. Protein was extracted from WT and Met Δ15-HET
murine embryonic fibroblast for protein expression analysis of downstream effectors of
MET signaling, along of key players which have been previously implicated in osteoblast
differentiation defect phenotypes. WT and Met Δ15-HET murine embryonic fibroblasts
were also treated with exogenous HGF in timed experiments to examine whether the
mutation was ligand (HGF) dependent.
Gene expression analysis was also conducted on WT and Met Δ15-HET osteoblasts
during the terminal stages of differentiation. This was done as Gray et al., (2015)
reported aberrant MET signaling to dysregulate late stage markers of osteoblast
differentiation while leaving early stage markers unaffected. Since this was completed in
a cell line, we wanted to confirm these results in primary osteoblasts.
Objective 2: Skeletal Characterization of WT and in Met Δ15-HET mice
Human OFD patients develop tibial bowing which may progress to non-healing
fractures. The second aim of this study was to determine if there were any
endochondral or intramembranous bone differences between WT and Met Δ15-HET
littermate mice. We first examined the osteogenic potential of bone marrow stromal cells
(BMSCs), which are mesenchymal stem cells originating from the bone marrow stroma.
41
We investigated the osteoblast differentiation process since it is involved in both forms
of ossification.
While Met Δ15-HET mice appear to be phenotypically normal, they have not been
extensively studied and characterized to examine bone differences. Full-body
dissections and skeletal preparations of male mice at the pup age (P5) and the more
skeletally mature age (P21) were performed to visualize any gross defects or
differences. These timepoints were chosen as P5 mice acted as a pre-pubescent young
immature skeleton, representing the age group (<10 years old) which is most commonly
affected by OFD. P21 mice acted as the post-pubescent adult mature skeletons,
(representing adult humans) and used to visualize longer term effects of the MET
mutation. Stained skeletons were photographed and further dissected to isolate long
bones (femur, tibia, humerus, radius) for manual and digital measurements by two-
blinded individuals. Measurements were analyzed to observe any statistical
significance. This was followed by examination of the long bone epiphyseal growth plate
zones of WT and Met Δ15-HET P21 littermate mice to examine if there were any
differences in zone heights.
Objective 3: Examination of Fracture Healing in WT and Met Δ15-HET mice in vivo
The third aim of this study was to examine the effects of the Met exon 15 skipping
mutation on fracture healing in vivo. This was done to test our hypothesis that the
phenotype seen in humans may be a stress induced one, as healthy tissue surrounding
OFD lesions do not experience defects in osteoblast differentiation. Transverse tibial
fractures were induced in 12-week old male mice utilizing a semi-stabilized tibial fracture
model. Progression of fracture healing was followed at 14-days post-fracture induction.
This timepoint was chosen as osteoblasts are involved in three key pathways occurring
simultaneously at this point in fracture repair: intramembranous ossification,
endochondral ossification, and callus remodeling. In addition, the 14-days post fracture
timepoint marks the peak of hard callus formation (Marsell & Einhorn, 2011). Quality of
healing was examined by analyzing callus size and mineralization at the aforementioned
time-point.
42
2.4 Clinical Significance
There is currently no cure for osteofibrous dysplasia, a quality of life impacting disease,
with treatment limited to pain management, stabilization of weakened tibias, or
osteotomies to correct severe deformities. Since OFD patients present with reduced
fracture healing abilities, understanding the underlying mechanisms responsible for their
phenotype may enable the identification of therapeutic targets and development of
novel therapeutic treatments. Furthermore, the knowledge gained from this study could
also be applied to other diseases displaying similar pseudarthrosis phenotypes such as
neurofibromatosis-1, or in the elderly where fracture healing abilities are reduced. By
improving the clinical outcomes of fracture healing, we hope to improve quality of life by
enabling patients to walk independently and perform their everyday activities.
43
3 Methods
3.1 Generation of a Met exon 15 splice donor mutant allele by CRISPR/Cas9-mediated genome editing Cas9 guide RNAs targeting the splice donor sequence of mouse Met exon 15 were
identified using the crispr.mit.edu site. Two guide RNAs targeting the exon 15 splice
donor sequence were selected for activity testing. Guide RNAs were cloned into a T7
promoter vector followed by in vitro transcription and spin column purification. Guide
RNA functional testing was performed by an in vitro cleavage assay incubating Cas9
protein and guide RNA with PCR-amplified target site. The guide RNA selected for
genome editing in embryos was Met-g68B (protospacer sequence 5’-
GTAAACTGAATTATACCTTC-3’). The donor oligonucleotide used to insert the splice
site mutation was Met-KI-B (5’-
AGCTCACAGAGGTCTATGTATAGATATTTCTCAGGATAGTAAACTGAATTATAACTT
CTGGAAAAGTAGCTCTGTAGTCTACAGACTCATTTGAAACCATCTCTGTAG-3’).
Cas9 mRNA was produced by T7 in vitro transcription.
3.2 Embryo Microinjection C57BL/6J zygotes were microinjected with Mix1 (10 ng/ul Cas9 mRNA, 5 ng/ul guide
RNA and 100 ng/ul donor oligonucleotide) or Mix2 (10 ng/ul Cas9 mRNA, 5 ng/ul guide
RNA and 10 ng/ul donor oligonucleotide) and implanted in recipient pseudopregnant
females. Resulting pups were screened by PCR and sequencing for the presence of the
mutant allele. One male founder from Mix1 and 1 male and 1 female founder from Mix2
were found to harbor the desired mutation.
3.3 Confirmation of Exon 15 Skipping The exon 15 skipping event was confirmed in our genetic mouse model by examining
the Met transcript for the presence or absence of exon 15. This was done by designing
PCR primers targeted to Exon 14 and 16 of the Met transcript. Met transcripts
containing exon 15 would result in formation of a 469 bp PCR product while the
exclusion of exon 15 would result in a 319 bp PCR product. Gel electrophoresis
44
mediated resolution was utilized to distinguish the differently sized PCR products.
Primers are as follows: Forward Primer: GGAAGCAAGCAGTCTCTTCAAC, Reverse
Primer: TGCTGAACTGCTTGGACCAG.
3.4 Maintenance and Genotyping of Mice Mice used in this study were housed in standardized cages at The Centre for
Phenogenomics in Toronto, ON according to their standards and guidelines. Mouse
genotyping was completed DNA extraction from mice tail clips done using DNA
extraction reagent from QuantaBio’s Extracta DNA Prep for PCR-Tissue (84158). 50 ul
of DNA extraction reagent is used per tail clipping. Incubated at 95oC for 35 minutes. 50
ul of DNA stabilizing buffer from QuantaBio’s Extracta DNA Prep for PCR-Tissue
(84159) was added post incubation. Concentration of the DNA extracted was
determined by nanodrop. Concentrations are used to calculate the dilution required of
extracted DNA to 5ng/mL. qPCR Master Mix Composition Per Sample: 10 uL of
TaqMan 2X Universal PCR Master Mix (4324018), 0.5 uL of Custom Taqman™ SNP
Genotyping Assay (4332077), 0.5 uL H2O. Primers are as follows: Forward Primer:
CCAACTACAGAGATGGTTTCAAATGAGT, Reverse Primer:
CACAGCTCACAGAGGTCTATGTATAGATAT. Probes are as follows: Probe 1:
CTTTTCCAGAAGGTATAATT, Probe 2: CTTTTCCAGAAGTTATAATT
45
3.5 Murine Embryonic Fibroblast Generation WT and Met Δ15-HET murine embryonic fibroblasts were derived following WiCell’s
“Derivation of Mouse Embryonic Fibroblasts (MEFs)” protocol. In short, timed-pregnant
mice were sacrificed on postnotum day E11.5, where 0.5 is the day of detection of a
copulation plug, by cervical dislocation. Pregnant mice were prepped by dissection by
sterilization of mice abdomen. An incision in the peritoneal wall was made until
exposure of the uterine horns. Uterine horns were extracted from the mother’s carcass,
washed with PBS. Embryos were released from embryonic sacs and washed with PBS,
followed by separation of visceral tissue from the embryos. Embryos were minced using
curved dissecting scissors into grain sized pieces (minced for approximately 5 – 1
minutes), followed by 2 ml of trypsin and additional mincing to ensure pieces were
further reduced in size. Lastly, add 5 ml more trypsin, pipetting cells vigorously up and
down to freeing of the cells from minced tissue. Incubate cells in a T75 flask, allowing
for growth and adherence, when cells reached 90% the culture were harvested to be
Heterozygous Mutant: GT
Homozygous Mutant: TT
Negative Control
(Water)
Homozygous
WT: GG
Typical allelic discrimination plot. Real-time PCR instrument software displaying the results of the
allelic discrimination data as a plot wild-type (GG) versus Met homozygous (TT).
46
stored in canonical tubes with Cryopreservation media. Cells were aliquoted into 1.5 ml
cryovials and flash-frozen with dry ice to be stored in -80oC freezer overnight and
moved to liquid nitrogen storage the following day for long term storage. Vials were
defrosted, re-suspended and cultured as required with Dulbecco’s Modified Eagle
Medium F/12 (Gibco) with 5% anti-anti and 10% fetal bovine serum.
3.6 MET Pathway Analysis WT and Met Δ15-HET murine embryonic fibroblasts were allowed to be cultured until
90% confluency with Dulbecco’s Modified Eagle Medium F/12 (Gibco) with 5% anti-anti
and 10% fetal bovine serum in a T75 flask. Culture media was removed and washed
with PBS to ensure removal of all residual media. Cells were detached and freed with
trypsin to form cell pellets in conical tubes for RNA and protein extraction.
3.7 Activation of MET pathway WT and Met Δ15-HET murine embryonic fibroblasts were allowed to be cultured until
50% confluency with Dulbecco’s Modified Eagle Medium F/12 (Gibco) with 5% anti-anti
and 10% fetal bovine serum in a T25 flask. 10 ng/ml HGF treatment was directly applied
to the culture media for 0, 5, and 30 minutes. Following these timepoints, HGF culture
media was removed and cells were washed with PBS to ensure removal of all residual
media. Cells were detached and freed with trypsin to form cell pellets in conical tubes
for RNA and protein extraction.
3.8 Western Blot Analysis
To examine protein levels of MET and its downstream effectors, protein was extracted
from WT and Met Δ15-HET MEFs. Cell cultures were pelletized, and protein extracted
using radioimmunoprecipitation assay buffer (RIPA buffer). Equal amounts of total
protein were resolved utilizing SDS-polyacrylamide gel-based electrophoresis, then
transferred to nitrocellulose membranes and immunoblotted at 4oC overnight with the
indicated primary antibodies: MET (Santa Cruz Biotechnology), phospho-MET (Cell
Signaling Technology), AKT (Cell Signaling Technology), phospho-AKT (Cell Signaling
Technology), mTOR (Cell Signaling Technology), phospho-mTOR (Cell Signaling
Technology), MEK (Cell Signaling Technology), phospho-MEK, ERK (Cell Signaling
47
Technology), phospho-ERK (Cell Signaling Technology), RAS (Cell Signaling
Technology), and β-catenin (BD Transduction Laboratories). Horseradish Peroxidase
(HRP) tagged secondary antibodies and ChemiLuminescence were utilized to detect
hybridization. Expression of the targeted proteins were quantified relative to the
expression of Actin or Vinculin as the housekeeping proteins. Quantification of Met Δ15-
HET protein expression was performed utilizing Image Lab quantification tools relative
to WT levels. Western blot analysis was carried out in triplicates to make sure the
experiments are reproducible.
3.9 Colony Forming Unit – Fibroblast Femora and tibiae from 12-week old male WT and Met Δ15-HET mice were isolated and
debrided of soft tissue. Bone marrow was flushed into BMSC culture media containing
α-modification of eagle’s medium (α-MEM) (Wisent Inc, Cat# 310-012-CL, St. Bruno,
CA), 10% FBS (Gibco Life Technology, Cat# 16000-044, Burlington, ON) and 1x
antibiotic and antimycotic (Wisent Inc, Cat# 450-115-EL). The cell suspensions were
passed through an 18G needle and 70μm cell strainer to dissociate clumps of cells.
Single cell suspensions were plated at a density of 1x106cells/cm2 and 5x106cells/cm2
surface area on 6-well plates, in BMSC culture medium for 7 days, with 50% of the
media changed at day 4. At day 7, plates were stained with crystal violet in 25%
methanol for 15 minutes. CFUs were counted manually using a microscope. CFU-F
colonies were defined as a discrete colony that contained 30 or more cells with the
majority of cells in each colony staining positively for crystal violet. All quantification
performed with 3 wells per mouse, and minimum of 3 mice per genotype.
3.10 Colony Forming Unit – ALP and Osteoblast Femora and tibiae from 12-week old male WT and Met Δ15-HET mice were isolated and
debrided of soft tissue. Bone marrow was flushed into BMSC culture media containing
α-modification of eagle’s medium (α-MEM) (Wisent Inc, Cat# 310-012-CL, St. Bruno,
CA), 10% FBS (Gibco Life Technology, Cat# 16000-044, Burlington, ON) and 1x
antibiotic and antimycotic (Wisent Inc, Cat# 450-115-EL). The cell suspensions were
passed through an 18G needle and 70μm cell strainer to dissociate clumps of cells.
Single cell suspensions were plated at a density of 1x106cells/cm2 and 5x106cells/cm2
48
surface area on 6-well plates, in BMSC culture medium for 7 days, with 50% of the
media changed at day 4. At day 7, the media was changed for osteoblastic
differentiation media, which consisted of BMSC culture media, as described above, with
the addition of 50μg/ml Ascorbic Acid (Sigma, Cat# A4544), 10-8M Dexamethasone
(Sigma, Cat# D8893), 8mM β-Glycerol phosphate (Sigma, Cat# G9891). The
osteoblastic differentiation media was subsequently changed every 2-3 days for the
duration of the experiment. On days 14 and 21 with OB differentiation media cells were
rinsed with PBS and fixed with 10% formalin for 30 mins and rinsed twice with water.
Alkaline phosphatase staining (Fast Red TR/Naphthol AS-MX Tablets: Sigma#F4523,
manufactory protocol) and Von Kossa staining (2.5% (w/v) Silver Nitrate: Sigma
#S8157, 30 minutes at room temperature on light box) were performed to assess for
osteoblastic differentiation and mineralization. CFUs were counted manually using a
microscope. CFU-ALP and CFU-OB colonies were defined as a discrete colony that
contained 30 or more cells with the majority of cells in each colony staining positively for
alkaline phosphatase and Von Kossa respectively. All quantification performed with 3
wells per mouse, and minimum of 3 mice per genotype.
3.11 RT-PCR: Gene Expression Analysis RNA was extracted from cell pellets using Monarch Total RNA Miniprep Kit (T2010S)
and used to create cDNA using LunaScript RT SuperMix Kit (E3010L), following
manufacturer’s protocol and instructions. cDNA was targeted with primers targeting
genes of interest from RNA-Seq and western blot analysis data. qPCR analysis was
carried out in triplicates to make sure the experiments are reproducible.
3.12 Skeletal Prep Staining For staining of bone and cartilage, whole skeletons were dissected from mice and fixed
in 95% ethanol for 48 hours in room temperature. Murine skeletons were submerged in
Alcian blue staining solution (2.5ml 0.3% Alcian Blue SGS (Sigma), 10ml glacial acetic
acid, and 40ml ethanol) for 48 hours at 37°C. Alcian blue staining solution was replaced
with 95% ethanol daily over 3 days. Skeletons were then submerged in Alizarin red
staining solution (0.5ml 0.2% Alizarin Red S (Sigma), 5ml 10% KOH, and 45ml distilled
water) for 24 hours in room temperature. Alizarin red staining solution was then
replaced with 20% glycerol, 1% KOH for 3 days in room temperature and then 50%
49
glycerol, 1% KOH until muscle and fat tissue was dissolved. This dissolving step varied
in time depending on the mouse size and age. Skeletons were then stored in 80%
glycerol for 24 hours and then 100% glycerol for long-term storage.
3.13 Epiphyseal Growth Plate Staining Mouse limbs were harvested in PBS and stored in 70% ethanol prior to paraffin
embedding and sectioning. Limbs were then paraffin embedded and sectioned. Slides
were rehydrated in the series of following ethanol concentrations: 100%, 90%, 70%,
50%, then washed twice in distilled water for 3 minutes. For H&E staining, slides were
then immersed in 0.3% ammonium hydroxide for 20 dips and rinsed twice in water for 1
minute, followed by immersion in Eosin Y certified biological stain (Fisher Scientific) for
10 dips. For Toluidine blue staining, slides were then immersed in 1% Toluidine Blue O
(Sigma) for 10 minutes. For Safranin-O staining, slides were then immersed in Weigert’s
iron hematoxylin solution (Sigma) for 10 minutes and rinsed in distilled water followed by
immersion in 1% Safranin O (Sigma) for 5 minutes. After all staining procedures, slides
were washed in water thrice for 1 minute, dehydrated in a 95%, 100% ethanol and
xylene series, then mounted and covered.
3.14 Visualization and Measurement of Long Bones and Epiphyseal Growth Plates Skeletal staining preparations of mice were photographed with a Canon DSLR over a
backlight while utilizing a custom cardboard aperture to maintain the same
magnification. A ruler was placed in each shot for measurement references, and images
were rotated in ImageJ and PowerPoint. Measurements of limbs were performed using
a Mastercraft digital caliper. The measurements were performed blinded by two
individuals and with each bone measured twice and its averaged value reported for
statistical analyses.
For P5 mice, humerus bones were measured from the most proximal staining point (at
the articulation with the distal glenoid cavity) to the most distal staining point (at the
articulation with the proximal ulnar aspect). Radius bones were measured from the most
proximal staining point (at the head of the radius) to the most distal staining point (at the
condyle). Femur bones were measured from the most proximal staining point (at the
50
articulation with the distal acetabulum) to the most distal staining point (preceding the
knee femoral cartilage). Tibia bones were measured from the most proximal staining
point (proceeding the knee tibial cartilage) to the most distal staining point (at the
articulation with the talus bone).
For P21 mice, humerus bones were measured from the most proximal staining point (at
the head of the humerus) to the most distal staining point (at the condyle). Following
suit, radius bones were measured from the most proximal staining point (at the head of
the radius) to the most distal staining point (at the condyle). Femur bones were
measured from the most proximal staining point (the femoral head and greater
trochanter) to the most distal staining point (at the femoral condyles). Tibia bones were
measured from the most proximal staining point (the head and tuberosity) to the most
distal staining point (at the malleoli).
Epiphyseal plates were visualized and photographed microscopically, then transferred
to ImageJ and PowerPoint for cropping. Measurements of epiphyseal plate zones and
cellularity were performed using ImageJ, then recorded using the Region of Interest
(ROI) Manager. The measurements were also performed blinded by two individuals and
each replicate was measured twice with its averaged value reported for statistical
analyses.
3.15 Tibia Fracture Generation Semi-stabilized tibial fractures were generated on 12-week-old male WT and Met Δ15-
HET mice following a protocol previously described (Y. Chen et al., 2007). Briefly, the
mice were anaesthetized using inhalational isoflurane general anesthesia. The left hind
limb of each mouse was surgically prepared by shaving and cleaning with povidone-
iodine as disinfectant. A small anterior midline incision was made over the knee joint
and proximal tibia. A 0.7 mm pilot hole was made at the proximal tibial epiphysis, medial
to the insertion of the patella tendon using a hollow 27G needle. Intramedullary fixation
was completed via a 0.7mm Anticorro insect pin (Fine Science Tools,
http://www.finescience.ca) was then inserted into the medullary cavity of the intact tibia
and advanced to the distal tibia. A transverse fracture was then induced at the mid-shaft
of the tibia using blunt scissors. The insect pin was cut 5-7mm proud of the proximal
tibial epiphysis and the skin incision closed with a series of absorbable vicryl rapide
51
sutures and metallic wound clips. Analgesic given subcutaneously (buprenorphine,
0.1mg/kg/twice per day) was administered for 3 days post-surgery. Previous data shows
that a fracture generated in this manner heals through both intramembranous and
endochondral ossification (Hiltunen, Vuorio, & Aro, 1993; Le, Miclau, Hu, & Helms,
2001). The animals were allowed to maintain free and full weight bearing in their cages
following surgery. At specific time points (14 days) after the fracture, samples of
fractured and unfractured tibiae were harvested following euthanasia with inhaled
carbon dioxide. A minimum of 3 mice per genotype, per time point, were used for further
analysis.
3.16 Histological Staining of Fracture Callus Fractured tibiae were harvested on post-fracture day 14, fixed in 10% formalin following
by decalcification with 20% (w/v) EDTA pH 8.0 or formic acid bone decalcifier (Decal
Chemical Corp, Tallman, NY) and embedded in paraffin. Serial 5 μM sections of paraffin
embedded tissues were deparaffinized and rehydrated through an alcohol gradient to
water. Sections were stained with Safranin-O and counter stained with fast
green/Mayer’s haematoxylin. Here red staining confirms the presence of proteoglycans,
which indicates cartilaginous tissue, and green staining indicates bone (Camplejohn &
Allard, 1988), or stained with Tartrate Resistant Acid Phosphatase (TRAP)
(manufacturers protocol, TRAP Staining- 387A-1KT, Sigma, St. Louis, MO) to evaluate
osteoclast activity. Here red staining confirms the presence of proteoglycans, which
indicates cartilaginous tissue, and green staining indicates bone (Camplejohn & Allard,
1988), or stained with Tartrate Resistant Acid Phosphatase (TRAP) (manufacturers
protocol, TRAP Staining- 387A-1KT, Sigma, St. Louis, MO) to evaluate osteoclast
activity.
3.17 Statistical Analyses Data (mouse weights, bone measurements, epiphyseal plate measurements,
chondrocyte experiments, etc.) were analyzed using the Student’s two-tailed
heteroscedastic t test by comparing all test groups (heterozygous mutant) to
corresponding control groups (wildtype). The ANOVA test was not utilized as multiple
group comparisons were not performed in any analyses. Significance was defined as
52
the p-value (P), with *P <0.05. Means and error bars were graphed using Microsoft
Excel chart and error bar formatting tools.
53
4 Results
4.1 Met Δ15-HET mouse embryonic fibroblast exhibit higher levels of MET protein and upregulated MET signaling WT and Met Δ15-HET mouse embryonic fibroblasts (MEFs) were subjected to protein
expression analysis to observe modulations and activity of downstream effectors of the
MET signaling pathway. This experiment was designed to test whether our mouse
model demonstrated similar disruptions in MET signaling seen in OFD patients. Murine
embryonic fibroblasts were cultured until 90% confluency in a T75 flask then prepared
for protein extraction. Protein was extracted and separated by SDS-PAGE and probed
for MET and its downstream effectors: AKT, mTOR MEK, ERK and their activated
phosphorylated versions.
MET was chosen to test if similar changes to the protein level as seen in human OFD
(Figure 1A) was replicated in the Met Δ15-HET mouse model, which mimics the identical
loss of regulatory control as humans. Protein kinase B (AKT) is a downstream MET
effector and was chosen as it has been implicated in the regulation of Runx2, a key
marker of early osteoblast differentiation. AKT regulates RUNX2 through Wnt/β-catenin
dependent and independent signaling pathways. Mammalian target of rapamycin
(mTOR), a downstream effector of AKT, was chosen as it is also involved in the
regulation of Runx2 and Wnt/β-catenin signaling through the mTORC1 complex.
Mitogen-activated protein kinase kinase (MEK1/2 or MAP2K) and mitogen-activated
protein kinase 1 (ERK1/2 or MAPK1), downstream effectors of RAS signaling (which is
connected to MET as shown previous in Figure 1A), have also been previously been
shown to directly regulate Runx2 expression and osteoblast differentiation. This
experiment tested whether signaling changes in Met Δ15-HET mice are similar to those
observed in human OFD. The utilization of a mouse model was required to overcome
the difficulties in obtaining OFD patient bone samples and cells.
Met Δ15-HET MEFs exhibited higher phosphorylation at tyrosine residues (Y1234 and
Y1235) found in the catalytic domain of the receptor, indicative of greater MET catalytic
54
signaling activity in comparison to WT MEFs (Fig 4.1 A). Accordingly, downstream
effectors of MET: AKT, mTOR, MEK1/2, ERK1/2 followed suit and also exhibited
upregulated signaling activity with increased levels of phosphorylation in Met Δ15-HET
MEFs (Fig 4.1 B & C). Surprisingly, Met Δ15-HET MEFs also displayed higher levels of
unphosphorylated MET, mTOR, MEK, ERK1/2 in comparison to their WT counterparts
(Fig 4.1 A – C).
55
WT HET WT HET WT HET
Y1234/Y1235
Figure 4.1. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts
Representative images of western blot protein analysis of MET (A) and its downstream effectors: AKT, mTOR, MEK, ERK1/2 (B & C) and their phosphorylated versions (B & C) in WT and MetΔ15-HET mouse embryonic fibroblasts (MEFs). MetΔ15-HET MEFs display
upregulated MET protein along with phosphorylation indicative of increased signaling activity. Accordingly, downstream effectors of MET display also display upregulated
signaling in MetΔ15-HET MEFs. Actin and Vinculin are the loading controls. N=3
A B C
56
4.2 Met Δ15-HET mouse embryonic fibroblast exhibit upregulated and prolonged MET signaling after HGF stimulation
WT and Met Δ15-HET mouse embryonic fibroblasts (MEFs) were treated with 10ng/ml
HGF for 0, 5, and 30 minutes. HGF is the only known ligand for MET. The experimental
timepoints were chosen as the half-life of HGF is approximately 3 – 5 minutes, therefore
peak signaling activity should be seen at 5 minutes and returned to basal levels at the
30-minute timepoint (Chang et al., 2016). Treated cells were subjected to protein
expression analysis to observe modulation, activity, and duration of activity in
downstream effectors of the MET signaling pathway. The results of this experiment
would allow us to examine if the dysregulation in signaling is ligand dependent.
Murine embryonic fibroblasts were cultured until 50% confluency in a T25 flask then
treated with 10ng/ml HGF for the aforementioned durations of time and prepared for
protein extraction. Protein was extracted and separated by SDS-PAGE and probed for
MET and its downstream effectors: AKT, mTOR, RAS, ERK1/2, β-catenin and their
activated phosphorylated versions.
MET, AKT, mTOR, RAS, and ERK1/2 were chosen for their involvement in regulation of
Runx2 expression and the osteoblast differentiation process as described in the
previous experiment. We chose to investigate β-catenin since the tight spatiotemporal
regulation of β-catenin signaling is crucial during osteoblast differentiation and
deviations compromise the differentiation process (Y. Chen et al., 2007). β-catenin’s
Serine 552 phosphorylation activity was examined as it is a phosphorylation target of
AKT. Phosphorylation at this site enhances β-catenin stability and transcriptional
activity.
Consistent with our previous data Met Δ15-HET MEFs exhibited greater phosphorylation
at tyrosine residues (Y1234 and Y1235) found in the catalytic domain of the receptor
prior to HGF stimulation (Fig 4.2 A & D). The higher levels of unphosphorylated MET
protein in comparison to WT MEFs may be due to reduced ability for the degradation
and terminal of signalling. At 5-minutes post HGF treatment, Met Δ15-HET MEFs
57
exhibited greater upregulation in phosphorylation at the Y1234 and Y1235 catalytic
tyrosine residues indicating increased MET signaling activity in comparison to WT MEFs
(Fig 4.2 A). Accordingly, downstream effectors of MET: AKT, mTOR, ERK and β-catenin
followed suit and also exhibited greater upregulated activity with increased levels of
phosphorylation 5-minutes post HGF treatment (Fig 4.2 A - M). This is further proof of
upregulated MET signaling pathway activity. Surprisingly, Met Δ15-HET MEFs displayed
higher unphosphorylated protein levels of MET, AKT, mTOR, RAS, ERK, β-catenin post
HGF treatment in comparison to their WT counterparts as well (Fig 4.2 A - M). This is
surprisingly as we previously believed the Met Δ15-HET mutation would only alter
phosphorylation activity at downstream effectors of MET. This indicates aberrant MET
signaling must modulation expression of these proteins as well, in addition to changes in
signaling activity.
Met Δ15-HET MEFs exhibit prolonged MET signaling following HGF stimulation; we
found prolonged phosphorylation activity at the MET receptor and its aforementioned
downstream effectors 30-minutes post HGF stimulation (Fig 4.2 A & B). This contrasts
the behaviour of WT MEFs, which returned to basal levels of phosphorylation activity at
the 30-minute timepoint, exhibiting similar activity to the 0-minute timepoint where no
HGF stimulation was applied (Fig 4.2 A & B).
58
MET
p-MET
AKT
p-AKT
Actin
mTOR
p-mTOR
p-AKT
AKT
MET
p-MET
Actin
Y1234/Y1235
Figure 4.2. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes
Representative images of western blots of MET and its downstream effectors: AKT, mTOR and their activated phosphorylated versions (A) in WT and MetΔ15-HET MEFs. When HGF stimulation is applied MetΔ15-HET MEFs display upregulated MET protein along with
phosphorylation indicative of increased signaling activity. In addition, MetΔ15-HET MEFs
display prolonged signaling in comparison to their WT counterparts. Accordingly, downstream effectors of MET display also display upregulated and prolonged signaling in MetΔ15-HET MEFs. Actin is the loading control. N=3
A
59
β-Catenin
p-β-Catenin (Ser 552)
Actin
ERK
RAS
Figure 4.2 Contd. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes
Representative images of western blots of downstream effectors of MET: RAS, ERK, β-catenin and their phosphorylated versions (B) in WT and MetΔ15-HET MEFs. When HGF stimulation is applied MetΔ15-HET MEFs display upregulated MET protein along with
phosphorylation indicative of increased signaling activity. In addition, MetΔ15-HET MEFs
display prolonged signaling in comparison to their WT counterparts. Accordingly, downstream effectors of MET display also display upregulated and prolonged signaling in
MetΔ15-HET MEFs. Actin is the loading control. N=3
B
60
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Figure 4.2 Contd. Protein expression analysis of WT and Met Δ15-HET mouse embryonic fibroblasts treated with 10ng/ml HGF stimulation for 0, 5, and 30 minutes Quantification of western blots of downstream effectors of MET: AKT, mTOR, RAS, ERK, β-
catenin and their phosphorylated versions normalized to WT levels (C – M). Student’s two-tailed t test was performed for statistical analysis comparing WT to MET Δ15-HET
measurements, with significance level set at *P<0.05. N=3
*
* *
*
*
*
* *
*
N.S.
N.S.
N.S. N.S.
N.S. N.S.
62
4.3 Met Δ15-HET MEFs exhibit upregulated levels of β-catenin
RNA-sequencing and protein expression analysis uncovered candidate genes of
interest for reverse transcription polymerase chain reaction (RT-PCR) validation
analysis. The following genes were chosen to be examined as previous experiments
revealed the potential for them to be a part of the explanation for delayed fracture
healing in OFD. WT and Met Δ15-HET MEFs were cultured until 80% confluency and
processed for RNA extraction followed by cDNA synthesis and RT-PCR analysis for the
following genes: Met, Axin2, β-catenin, Erk, Akt.
Since we saw higher levels of MET at the protein level in Met Δ15-HET MEFs, we wanted
to confirm this was caused by a failure to degrade the receptor rather than upregulated
Met expression caused by aberrant MET signaling. Therefore, Met expression was
examined between WT and Met Δ15-HET MEFs to observe any potential differences.
Here we saw no significant changes in Met expression between WT and Met Δ15-HET
mutants, indicating the differences in MET at the protein level may be due to the failure
to degrade the receptor rather than changes in gene expression (Fig 4.3 B). β-catenin,
Erk, and Akt were chosen as we saw upregulation of their protein levels in Met Δ15-HETs
during western blot analysis (Fig 4.2 A & B) and therefore wanted to check whether
there were associated gene expression changes. These three proteins are also
important in the osteoblast differentiation process requiring tight regulation of their
expression and activity (Ghadakzadeh et al., 2016; Greenblatt et al., 2010). Here we
saw a 6.43-fold, 6.88-fold, and 7.4-fold upregulation in expression of β-catenin, Erk, and
Akt respectively confirming our previous results indicating the MET mutation also affects
gene expression (Fig 4.3 D – F).
While examining the dysregulation of β-catenin levels, one must also examine if there is
any aberrant degradation of it as well, therefore Axin2 was chosen as it is a critical
scaffold protein of the β-catenin degradation complex consisting of GSKβ and APC
(Shang, Hua, & Hu, 2017). Here we saw a 12-fold decrease in Axin2 expression (Fig 4.3
C), indicating reduced degradation of β-catenin in Met Δ15-HET MEFs along with the
increased expression of β-catenin.
63
Overall these results indicate aberrant MET signaling may be caused by the failure to
degrade the MET receptor in a negative-feedback fashion and not caused by elevated
Met gene expression. As we observed higher levels of MET protein and activation of
downstream effectors in Met Δ15-HETs despite no changes in Met expression. They also
show the dysregulation of β-catenin levels in Met Δ15-HET MEFs occurs through multiple
diverse pathways.
64
A
Figure 4.3 Met Δ15-HET MEFs exhibit gene expression dysregulation resulting in upregulation of β-catenin RT-PCR gene expression analysis of Met, Axin2, β-catenin, Erk, Akt. Expression
changes shown as log2 fold change.
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*
Figure 4.3 Contd. RT-PCR gene expression analysis of Met, Axin2, β-catenin, Erk, Akt. Expression
changes shown as fold-change normalized to WT expression (B – F). Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET
measurements, with significance level set at *P<0.05. N=3
66
4.4 Met Δ15-HET bone marrow stromal cells exhibit reduced osteoblast differentiation and bone mineralization
In order to visualize differences in osteogenic capacity in vitro, we utilized bone marrow
stromal cells (BMSCs) isolated from 10-week-old male WT and Met Δ15-HET mice’s
femurs. Only male mice were used in this experiment to account for sex-specific
differences in osteogenic potential. 10-week-old aged mice were used as 8 – 12-week-
old mice provide the most reliable source of bone marrow stromal cells (Beane,
Fonseca, Cooper, Koren, & Darling, 2014; Nadri et al., 2007). BMSCs were plated at
1x106 cells/well in a 6-well dish and induced to undergo osteoblast differentiation after 7
days of rest using osteoblast (OB) differentiation media consisting of ascorbic acid,
dexamethasone, and β-Glycerol phosphate.
Colony forming units – fibroblasts (CFU-F) were quantified at day 7 of BMSC rest by
staining with crystal violet and examining colonies staining positively for crystal violet.
This assay provides a means to assess the proliferation and cologenic capacity of the
cells when expanded in culture (Nadri et al., 2007). Osteoblast cultures were stained for
alkaline phosphatase (ALP) 14 days after addition of OB differentiation media. ALP acts
as a marker of bone matrix secretion and early stage marker of osteoblast
differentiation. At day 21, cultures were stained with Von Kossa, a late stage marker of
osteoblast differentiation, to visualize matrix mineralization. Colony forming units –
alkaline phosphatase (CFU-ALP) were quantified as colonies staining positively for
alkaline phosphatase. Colony forming units – osteoblast (CFU-OB) were quantified as
colonies staining positively for Von Kossa on day 21 of osteoblast differentiation. The
number of bone nodules formed, represents the number of osteoprogenitors in a bone
marrow sample as each colony forms from a single cell precursor (Aubin, 1998). CFU-F,
CFU-ALP, and CFU-OB colonies were defined as discrete colonies which contained 30
or more cells.
Surprisingly, Met Δ15-HET BMSC colonies exhibited a reduced ability to form CFU-
fibroblast. Indicating Met Δ15-HET BMSCs may have a reduced capacity for proliferation,
as each colony is derived from a single-cell precursor (Bianco & Robey, 2000). Met Δ15-
HET BMSCs also formed 23% less CFU-F colonies in comparison to WT BMSCs. They
67
also exhibited 35% less CFU-ALP colonies and 80% less CFU-OB colonies in
comparison to WT. This indicates Met Δ15-HET BMSCs display reduced osteoblast
differentiation and bone mineralization capacity caused by aberrant MET signaling.
These results also show our Met Δ15-HET mouse model mimics the same osteoblast
differentiation defect seen in human OFD patients and may thus exhibit the same
reduced fracture healing ability as well.
68
Met Δ15-HET
WT
Figure 4.4. Met Δ15-HET bone marrow stromal cells exhibit reduced mineralization ability Bone marrow stromal cells were obtained from 12-week old male mice. Representative wells of WT and Met Δ15-HET bone marrow stromal cells plated at 1x106 cells/well treated
with osteoblast differentiation media and stained for alkaline phosphatase and Von Kossa 21 days after initiation of the differentiation process (A). N=3
A
69
Nu
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CFU-F CFU-ALP CFU-OB
** * ***
Figure 4.4 Contd. Met Δ15-HET bone marrow stromal cells exhibit reduced mineralization ability
Quantification of colony forming units (CFU) plated at 1x106 cells/well normalized to WT to demonstrate reduced ability of Met Δ15-HET BMSCs to form CFU-Fibroblast (CFU-F;
p=0.011), CFU-Alkaline Phosphatase (CFU-ALP; p=0.005) measured on day 14, and CFU-Osteoblast (CFU-OB; p=2.95x10-5) measured on day 21 (B). Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET measurements, with
significance level set at *P<0.05. N=3
B
70
4.5 Mature Met Δ15-HET osteoblasts display dysregulation of osteoblast specific gene markers
In order to gain a better understanding of the underlying mechanistic changes in the
osteoblast differentiation defect observed in the previous experiment, we examined the
expression of osteoblast specific gene markers at day 21 of differentiation. This
timepoint was chosen as we saw a more severe and striking reduction in osteoblast
differentiation at day 21 of our osteoblast assay experiments, suggesting the Met Δ15-
HET mutation may be interfering with the late stages of the differentiation process.
Here we utilized BMSC induced osteoblast differentiation cell cultures as described
previously. RNA was extracted and cDNA synthesized on day 14 of osteoblast
differentiation, followed by RT-PCR analysis for the following genes: β-catenin, Runx2,
Alp. The same was done for day 21 of osteoblast differentiation, followed by RT-PCR
analysis for the following genes: β-catenin, Axin2, Runx2, Osteocalcin, Adrenomedullin,
Rasal3.
β-catenin, Axin2 were chosen for their involvement in the osteoblast differentiation
process as previously described. At day 14 of osteoblast differentiation we see no
differences in β-catenin expression while we found a 22-fold increase in β-catenin
expression at day 21 of osteoblast differentiation (Fig 4.5 A & B). This indicates there is
a dysregulation of β-catenin expression in day 21 Met Δ15-HET osteoblasts. In addition,
the 6.4-fold reduction in Axin2 expression indicates reduced degradation of β-catenin in
day 21 Met Δ15-HET osteoblasts as well (Fig 4.5 C). Together, these two results
correspond with the upregulation of β-catenin at the protein level in Met Δ15-HET
mutants seen during our western blot analyses (Fig 4.2 B).
Runx2 was chosen as the persistence of its expression during the later stages of
differentiation inhibits terminal differentiation of osteoblasts. In addition, β-catenin
positively regulates Runx2 expression. Therefore, if there is dysregulation in β-catenin
levels, Runx2 should follow suit as well. Alkaline phosphatase and osteocalcin
expression were examined they act as early and late markers of osteoblast maturation
71
respectively. Thus, we would expect to see a reduction in alkaline phosphatase and
osteocalcin expression in Met Δ15-HET osteoblasts if there was a defect in terminal
differentiation. Met Δ15-HET osteoblasts exhibited a non-significant 1.03-fold increase in
Runx2 expression at day 14 of differentiation, while a 27-fold increase in Runx2
expression was seen at day 21 of differentiation (Fig 4.5 D & E). This upregulation
occurs at a stage at which Runx2 expression should be at its minima. As expected, we
saw a 1.2-fold reduction in alkaline phosphatase expression in day 14 Met Δ15-HET
osteoblasts (Fig 4.5 F). This reduction was present at day 21 of differentiation as well,
with Met Δ15-HET osteoblast exhibiting a 1.3-fold reduction in alkaline phosphatase (Fig
4.5 G). In addition, a 4-fold reduction in osteocalcin expression was seen in day 21 Met
Δ15-HET osteoblasts, indicative of a failure in the terminal osteoblast differentiation
process (Fig 4.5 H).
Adm and Rasal3 were chosen for their involvement in the regulation of β-catenin
stability. (Vagin & Beenhouwer, 2016). We saw a 24.6-fold increase in Adm expression
in day 21 Met Δ15-HET osteoblasts, adrenomedullin acts as a regulator of the inhibition
of GSK3β activity (Fig 4.5 I) (Lausson & Cressent, 2011). In addition, a 1.29-fold
decrease in Rasal3 expression is seen in day 21 Met Δ15-HET osteoblasts, which
functions in the inactivation of RAS activity and β-catenin (Fig 4.5 J) (Muro et al., 2015).
Overall these results indicate aberrant MET signaling secondary to the gain-of-function
MET mutation results in dysregulation of β-catenin mediated Runx2 expression, leading
to inhibition of terminal differentiation in Met Δ15-HET osteoblasts.
72
C
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*
Figure 4.5 Met Δ15-HET mature osteoblasts display dysregulation of osteoblast specific gene markers Bone marrow stromal cells were obtained from 12-week old male mice. RT-PCR gene expression analysis of β-catenin, Runx2, Alp on day 14 of osteoblast differentiation and β-catenin, Axin2, Runx2, Alp, Osteocalcin, Adm, Rasal3 on day 21 of osteoblast differentiation.
Expression changes shown as fold-change normalized to WT expression (A – E). Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET
measurements, with significance level set at *P<0.05. N=3
* *
J
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74
4.6 Body weight and preliminary comparisons of WT and Met Δ15-HET mice
We examined the body weights of WT and Met Δ15-HET mice to rule out a generalized
growth defect secondary to the gain-of-function MET mutation. Mice were selected at
the pup/neonatal age (P5), juvenile age (P21) and adult age (3 month) for imaging and
examination of body weight. These timepoints were chosen to represent pre-pubescent,
adult and adult mice respectively.
At the P5 timepoint, mice were not distinguished based on sex, as sex determination
has been described to be unreliable at this time point (Schlomer et al., 2013). At the P21
and 3-month timepoints, only male mice were used to avoid confounding factors relating
to differences in body weight due to sex.
At P5, decapitated WT and Met Δ15-HET mice do not appear to differ significantly in
body size or average weight (Fig 4.6 A & D). At P21, WT and Met Δ15-HET mice also
appear similar superficially (Fig 4.6 B & D). The same is true at the 3 months old time
point when examining WT and Met Δ15-HET mice (Fig 4.6 C & D). Weights charted over
time also revealed a consistent linear body weight velocity between WT and Met Δ15-
HET mice (Fig 4.6 E). These results preliminarily suggest the Met Δ15-HET mutation
does not cause a developmental phenotype affecting overall size.
75
WT Met Δ15-HET
Figure 4.6 Body weight and preliminary comparisons of WT and Met Δ15-HET mice
Side-by-side comparison of physical appearance of WT and Met Δ15-HET at P5 to observe
gross superficial differences at pup stage (A). N=6
A 5cm
76
WT Met Δ15-HET
Figure 4.6 Contd. Side-by-side comparison of physical appearance of WT and Met Δ15-HET at P21 to observe
gross superficial differences at the juvenile stage (B). N=6
B 10cm
77
WT Met Δ15-HET
Figure 4.6 Contd. Side-by-side comparison of physical appearance of WT and Met Δ15-HET at 3 months old to
observe gross superficial differences at the juvenile stage (C). N=4
C 10cm
78
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Figure 4.6 Contd. Representative images of WT vs Met Δ15-HET mice at P5 (A), P21 (B), 3 months old (C). Body weight
measurements of WT and Met Δ15-HET mice at P5, P21, 3 months of age. N=6, 6, and 4 respectively (D). Average body weights of WT and Met Δ15-HET plotted over time (E). Student’s two-tailed t test was
performed for statistical analysis, with significance level set at *P<0.05.
D
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79
4.7 Skeletal Staining of P5 WT and Met Δ15-HET mice
We previously showed Met Δ15-HET BMSCs exhibit reduced ability to form osteoblasts
and mineralization nodules. Osteoblasts are heavily involved in the endochondral and
intramembranous ossification processes. Therefore, we determined if there were any
discrepancies in bones formed through these processes. In addition, we examined for
gross skeletal phenotypic differences present in 5-day old mice. P5 mice were sacrificed
by cervical dislocation and dissected for whole-mount skeletal staining. The staining
protocol was used to digest excess muscle and fat tissue, stain bone with Alizarin red
and stain cartilage with Alcian blue to delineate skeletal regions for greater accuracy
when measuring.
No gross body skeletal defects were seen between WT and Met Δ15-HET mice at 5 days
old (Fig 4.7 A). Isolation of the upper limbs showed no significant differences in humerus
and radius lengths between the two (Fig 4.7 C & E). Isolation of the lower limbs
displayed no significant differences in femur and tibia lengths in WT and Met Δ15-HET
mice (Fig 4.7 D & E). These results indicate this mutation does not affect endochondral
ossification and long bone development at 5-days old. No gross differences in skull
length or size were seen at P5 (Fig 4.7 B & F). This suggests the rate of
intramembranous ossification, the process in which skulls are formed, is not affected by
the Met Δ15-HET mutation at this timepoint.
80
50 m
m
50 m
m
WT Met Δ15-HET A
81
WT Met Δ15-HET
WT Met Δ15-HET
B
C
10 mm
10 mm
82
WT Met Δ15-HET
D
10 mm
Figure 4.7. 5-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates
Representative images of 5-day old WT and Met Δ15-HET whole-body, condylobasal lengths,
humerus and radius bones, and femur and tibia bones skeletal preps stained with Alcian Blue
and Alizarin Red for cartilage and bone identification respectively (A-D). Red lines indicate observable ossified regions in each body part.
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Figure 4.7. 5-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates
Average measurements of P5 WT and MET Δ15-HET femur, tibia, humerus, radius (E). Average measurements of P5 WT and Met Δ15-HET condylobasal lengths (F). Student’s two-tailed t test
was performed for statistical analysis comparing WT to Met Δ15-HET measurements, with
significance level set at *P<0.05. N = 6.
0
1
2
3
4
5
6
7
8
Avg Femur Avg Tibia Avg Humerus Avg Radius
Ave
rag
e L
en
gth
(m
m)
WT
HET
N.S.
N.S.
N.S.
N.S. E
0
5
10
15
20
25
WT HET
Av
era
ge
Co
nd
ylo
ba
sa
l L
en
gth
(m
m)
F
N.S.
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4.8 Skeletal Staining of P21 WT and Met Δ15-HET mice
No gross body skeletal defects were seen between WT and Met Δ15-HET mice at 21-
days-old (Fig 4.8 A). Isolation of the upper limbs showed no significant differences in
humerus and radius lengths between the two (Fig 4.8 C & E). Isolation of the lower
limbs displayed no significant differences in femur and tibia lengths in WT and Met Δ15-
HET mice (Fig 4.8 D & E). These results indicate this mutation does not affect
endochondral ossification and long bone development at 21-days old either. No gross
differences in skull length or size were seen at P21 (Fig 4.8 B & F). This suggests the
rate of intramembranous ossification, the process in which skulls are formed, is not
affected by the MetΔ15-HET mutation at this timepoint as well.
85
WT Met Δ15-HET 10
0 m
m
10
0 m
m
A
86
WT Met Δ15-HET
WT Met Δ15-HET
B
C
20 mm
10 mm
87
WT Met Δ15-HET
D
10 mm
Figure 4.8. 21-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates
Representative images of 21-day old WT and Met Δ15-HET whole-body, condylobasal lengths, humerus
and radius bones, and femur and tibia bones skeletal preps stained with Alcian Blue and Alizarin Red
for cartilage and bone identification respectively (A-D). Red lines indicate observable ossified regions in each body part.
88
0
2
4
6
8
10
12
14
Avg Femur Avg Tibia Avg Humerus Avg Radius
Ave
rag
e L
en
gth
(m
m)
WT
HET
Figure 4.8. 21-day old Met Δ15-HET mice exhibit no differences in long bone lengths in comparison to WT littermates Average measurements of P21 WT and MET Δ15-HET femur, tibia, humerus, radius (E). Average
measurements of P21 WT and Met Δ15-HET condylobasal lengths (F). Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET measurements, with significance level
set at *P<0.05. N = 6.
N.S.
N.S.
N.S.
N.S.
E
0
5
10
15
20
25
WT HET
Av
era
ge
Co
nd
ylo
ba
sa
l L
en
gth
(m
m)
F N.S.
89
4.9 Histological analysis of P21 WT and Met Δ15-HET mice epiphyseal growth plate
We previously discovered no differences in endochondral and intramembranous bone
lengths, despite osteoblast being critical in both processes. The epiphyseal growth plate
is where chondrocytes progressively enlarge, hypertrophy and either differentiate to
osteoblasts or undergo apoptosis and mineralization. This is very similar to the
processes which occur in endochondral ossification. Thus, we analyzed the growth plate
to determine if there were irregularities in growth plate morphology between WT and
Met Δ15-HET mice.
Proximal tibia epiphyseal growth plates of 21-day old male WT and MET Δ15-HET
littermates were sectioned and stained with Safranin-O and fast green to visualize
cartilage and bone respectively.
Here we find Met Δ15-HET mice exhibit no significant differences in height or cellular
organization at the resting, proliferating, prehypertrophic and hypertrophic zones (Fig
4.9 A & B). No significant differences in overall epiphyseal growth plate height was seen
either between WT and Met Δ15-HET mice at 21-days old. Overall these results confirm
and further suggest this mutation does not affect long bone development at 21-days old.
90
A
Figure 4.9 Met Δ15-HET mice exhibit no differences in epiphyseal growth plate lengths in comparison to WT littermates
Safranin-O staining of P21 WT vs Met Δ15-HET tibial epiphyseal growth plates (composed of the
resting, proliferative, prehypertrophic and hypertrophic zones). Representative images shown
here (A).
Resting Zone
Proliferating Zone
Prehypertrophic Zone
Hypertrophic Zone
91
0
50
100
150
200
250
RZ PZ PHZ HZ Full
Len
gth
(μ
m)
Epiphyseal Growth Plate Zone
B
Figure 4.9 Contd. Met Δ15-HET mice exhibit no differences in epiphyseal growth plate lengths in comparison to WT littermates
Average measurements of resting zone (RZ), proliferating zone (PZ), prehypertrophic zone (PHZ), hypertrophic zone (HZ) and full zone (Full) (B). Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET measurements, with significance level set
at *P<0.05. N = 3
N.S.
Zones: RZ = Resting
PZ = Proliferating PHZ = Prehypertrophic HZ = Hypertrophic
N.S.
N.S.
N.S.
N.S.
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4.10 Overactivation of MET signaling impairs fracture healing
The lack of a skeletal phenotype despite a reduction in osteoblast differentiation ability
in Met Δ15-HET mice led us to believe the phenotype seen in OFD patients may be
stress induced. We then investigated the fracture healing ability of WT and Met Δ15-HET
12-week-old mice. Since our Met Δ15-HET mouse model do not develop spontaneous
fractures, here we utilized a semi-stabilized tibial fracture induction model which has
been shown to heal through a combination of endochondral and intramembranous
ossification (Y. Chen et al., 2007). Mice were sacrificed 14-days post-fracture for
histological analysis to evaluate fracture healing. This time-point was chosen as it is the
mid-way point of fracture repair where multiple processes (intramembranous
ossification, endochondral ossification, and callus remodelling) involving osteoblasts can
be examined.
At 14-days post-fracture we found significantly more unmineralized cartilage at the
callus site, stained by Safranin-O in red, in Met Δ15-HET mice (Fig 4.10 A). Accordingly,
histomorphometric analysis showed Met Δ15-HET mice fracture sites had significantly
less bone volume/total callous tissue volume (BV/TV) and more osteoid volume/bone
volume (OV/BV) in comparison to their timepoint matched WT counterparts (Fig 4.10 B
& C). This indicates Met Δ15-HET possessed less mineralized bone and more
unmineralized tissue (osteoid) at the fracture site, alluding to a delay in fracture healing.
We also see significantly less tartrate resistant acid phosphatase (TRAP) positive
staining, the histochemical marker of osteoclasts, in Met Δ15-HET mice 14-days post
fracture (Fig 4.10 D) (Minkin, 1982). This was confirmed through colony counting
quantification of the TRAP positive osteoclasts, indicating Met Δ15-HET mice had
significantly reduced number of osteoclasts per defined region of interest (Fig 4.10 E).
Overall, these results indicate Met Δ15-HET mice are at an earlier stage in the fracture
repair timeline in comparison to WT mice 14-days post-fracture.
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Safranin-O staining for Cartilage (red)
A
Figure 4.10 Aberrant MET signaling impairs fracture healing in 12-week old mice Representative images of tibial fractures of WT and Met Δ15-HET mice stained with
Safranin-O and fast green to visualize cartilage and bone respectively. Images display the region of interest at the fracture callus. N = 3 mice per group
WT Met Δ15-HET
94
0
10
20
30
40
WT HET
Perc
en
tag
e
Bone Volume/Tissue Volume [%]
* B
Figure 4.10 Contd.
Quantification of the relative proportion of mineralized bone cartilage and total callous tissue volume at the fracture site using histomorphometric analysis. Student’s two-tailed t test was performed for statistical analysis comparing WT to
Met Δ15-HET measurements, with significance level set at *P<0.05. N=3 mice per
group.
95
*
0
3
6
9
12
15
WT HET
Perc
en
tag
e
Osteoid Volume/Bone Volume [%]C
Figure 4.10 Contd. Quantification of the relative proportion of unmineralized tissue (osteoid) and total
mineralized bone tissue at the fracture site using histomorphometric analysis. Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET measurements, with significance level set at *P<0.05. N=3 mice per
group.
*
96
WT Met Δ15-HET D
Figure 4.10 Contd. Representative images of WT and Met Δ15-HET tibial fracture callous sites stained
for tartrate-resistant acid phosphatase (TRAP) positive cells (osteoclasts). Arrows indicate TRAP positive cells. N = 3 mice per group.
97
0
5
10
15
20
25
30
35
40
WT HET
Nu
mb
er
of
Oste
ocla
sts
/Reg
ion
of
Inte
rest
* E
Figure 4.10 Contd.
Quantification of the number of osteoclasts per region of interest. Student’s two-tailed t test was performed for statistical analysis comparing WT to Met Δ15-HET
measurements, with significance level set at *P<0.05. N=3 mice per group.
98
5 Discussion
Fracture healing is a highly regulated and complex process. Dysregulation at any one of
the many steps may result in delayed healing and predispose to the non-union seen in
osteofibrous dysplasia. Fracture healing is dependent on osteoblasts to mineralize the
cartilaginous matrix and restore bone function and structural integrity (Marsell &
Einhorn, 2011). Therefore, we believe that a defect in osteoblast differentiation, as seen
in MetΔ15-HET mice, results in delayed fracture healing. Determining and defining the
mechanisms underlying this process in osteofibrous dysplasia may lead to the discovery
of novel therapeutic targets for non-invasive therapies to augment fracture repair.
5.1 MetΔ15-HET mice exhibit higher MET protein levels and upregulated MET signaling
In this project, we used a mouse model specifically created to mimic one of the human
OFD mutations which results in a MET gain-of-function mutation. We first determined if
the model exhibited a similar dysregulation in signalling pathways previously observed
in osteofibrous dysplasia. To accomplish this, we tested MET and its immediate
downstream effectors’ activity at the protein level using MEFs. MetΔ15-HET MEFs
exhibited elevated basal phosphorylation activity at MET catalytic phosphorylation sites
and its downstream effectors: AKT, mTOR, MEK1/2 and ERK1/2 in comparison to WT
MEFs (Fig 4.1A – C). Higher levels of basal signaling activity at these proteins was
consistent with what was previously identified in OFD patients and previously reported
by Gray et al. (2015) ; they demonstrated the competency of the mutant OFD MET
receptor to initiate phosphorylation and downstream signal transduction despite the loss
of the juxtamembrane domain. We also observed higher MET protein levels in mutant
MEFs, but no changes in Met gene expression between WT and MetΔ15-HET MEFs (Fig
4.3 B). Together this data indicates the higher levels of MET protein may be secondary
to diminished receptor internalization and degradation since Met gene expression was
unchanged. Since our results demonstrate similar changes in MET and its downstream
effectors as what was previously reported in human OFD patients, we were satisfied
with using this model for our other proposed experiments.
99
Next, we determined if the MetΔ15-HET mutation was ligand dependent by following
receptor activation after stimulation with its one and only ligand, HGF. Since the half-life
of HGF is approximately 3 – 5 minutes, peak signalling activity should be observed at
the 5-minute time-point (Chang et al., 2016). When the MET receptor was exogenously
activated, we observed a greater upregulation in catalytic phosphorylation activity at
MET and its downstream effectors (RAS, AKT, mTOR, ERK and β-catenin) 5-minutes
post HGF addition in mutant MEFs versus WT (Fig 4.2A – B). This activity level was
also maintained for a longer duration in MetΔ15-HET MEFs. There was a persistence of
phosphorylation activity 30-minutes post-addition of HGF by which time WT MEFs had
returned to basal activity levels. The reduction in WT MEF signaling activity 30-minutes
post-HGF is consistent with the expected degradation of the MET receptor and signal
termination post receptor activation. Together, these two results suggest the
upregulation of the signaling is due to a failure to internalize and degrade the receptor
post-activation, and consistent with a deletion of the ubiquitin ligase CBL binding site.
The MetΔ15-HET mutation appears to be ligand dependent as increasing HGF
concentrations above basal levels resulted in upregulation of signaling activity. This
observation is consistent with what has been reported by other groups who describe
mutant MET-mediated transformations to be ligand-dependent and inhibited by HGF
antagonists (Michieli et al., 1999). Overall, these results indicate the MetΔ15-HET
mutation is a gain-of-function mutation which is ligand dependent. Our data is also
consistent with the MetΔ15-HET receptor exhibiting reduced degradation.
Mutant MEFs also exhibited higher protein levels of downstream MET effectors: AKT,
mTOR, MEK, ERK1/2 and RAS, replicating what was seen in human OFD lesional
tissue (Fig 1A). In addition, RT-PCR analysis confirmed upregulation of AKT and
ERK1/2 expression in METΔ15-HET MEFs suggesting modulation at the gene
expression level. We have shown that a key osteoblast transcriptional factor, Runx2, is
upregulated in MetΔ15-HET MEFs. Others have shown that an upregulation of Runx2
expression can increase PI3K, AKT and mTOR expression (Cohen-Solal, Boregowda, &
Lasfar, 2015; Fujita et al., 2004). We speculate that Runx2 expression may be changing
AKT and ERK1/2 expression in our animal model but agree that the mechanism behind
100
these gene expression changes is not yet well understood and will be a critical point of
interest for future experiments aimed at deciphering the relationship between MET
signaling and downstream effectors.
5.2 Overactivation of MET signaling causes osteoblast differentiation defects
After confirming our mouse model recapitulates the dysregulations in signaling seen in
human OFD patients, we tested if the MetΔ15-HET mutation produces an osteoblast
differentiation defect. Here we compared the osteogenic potential of our WT and MetΔ15-
HET mice utilizing colony forming unit assays. We showed MetΔ15-HET BMSCs have a
reduced ability to form fibroblast colonies in comparison to WT bone marrow (Fig 4.4 B).
CFU-F are used as an approximation of skeletal stem cell concentration in bone marrow
(Kuznetsov, Mankani, Bianco, & Robey, 2009). Thus, this data indicates MetΔ15-HET
mice have reduced skeletal stem cells (a subset of BSMCs) in comparison to their WT
counterparts. We then showed MetΔ15-HET mice formed a reduced number of CFU-ALP
and CFU-OB consistent with a reduced ability for osteoblast differentiation and
mineralization (Fig 4.4 B). This finding was similar to what was described by Gray et al.,
(2015) in OFD patients. The number of bone nodule per number of cultured cells
provides an estimation and measurement of the bone forming capacity of the bone
marrow sample (Aubin, 1998). We began to see the reductions in osteoblast
differentiation capacity at day 14 of differentiation through the CFU-ALP assay, but the
reductions in colony number were much more severe at day 21. This suggests the
MetΔ15-HET mutation must have a greater impact on differentiation between day 14 and
21 than between day 1 and 14 of the differentiation timeline. Osteoblasts undergo
maturation between day 14 and 21, suggesting the MetΔ15-HET mutation may be
targeting its effects at the maturation stage of the differentiation process.
5.3 MetΔ15-HET MEFs exhibit dysregulation of β-catenin activity
Through pathway analysis at the protein and gene expression level, we found
dysregulation in multiple pathways leading to the upregulation of β-catenin and Runx2
expression levels in MetΔ15-HET MEFs.
101
MetΔ15-HET MEFs exhibited higher levels of phosphorylated ERK1/2, indicative of
increased ERK1/2 signaling activity (Fig 4.1C & 4.2 B). ERK1/2 signaling can
phosphorylate LDL-related protein 6 (LRP6), a coreceptor of WNT proteins and key
positive regulator of Wnt/β-catenin signaling. This phosphorylation event enhances the
cellular response to WNT proteins, upregulating Wnt/β-catenin signaling (Cervenka et
al., 2010). We also saw upregulated AKT signaling and corresponding phosphorylation
of the Ser552 residue in β-catenin (Fig 4.2 A & B). Phosphorylation at this residue is
AKT dependent and promotes β-catenin stability and translocation to the nucleus where
it regulates downstream target gene expression (Raucci et al., 2008). Increased AKT
signaling also upregulates downstream pathways such as mTOR, which acts through
the mTORC1 complex to directly upregulate the key osteoblast transcription factor
Runx2 (Dai et al., 2017) (Fig 4.2 A). We also identified a down-regulation in Axin2
expression. Axin2 is required for GSK-3β to phosphorylate β-catenin and target it for
degradation (Jho et al., 2002). Thus, we expect a reduction in Axin2 to increase the
stability of β-catenin as was shown in our data.
In conclusion, we believe that the combination of events affecting ERK1/2 and AKT
signaling along with dysregulations in gene expression of regulators of β-catenin
stability result in the constitutively higher β-catenin levels observed in MetΔ15-HET mice.
Osteoblast differentiation requires tight regulation of β-catenin to achieve terminal
differentiation (Ghadakzadeh et al., 2016). Thus, changes to the spatial and temporal
control of β-catenin levels may result in the osteoblast differentiation seen in MetΔ15-
HET osteoblasts.
5.4 Overactivation of MET signaling does not affect postnatal skeletal development
MetΔ15-HET mice appear to be phenotypically normal in our breeding facility but have
not been thoroughly examined previously. Therefore, we determined their skeletal
phenotype using standard skeletal preparations. Despite seeing reductions in osteoblast
differentiation potential in vitro, MetΔ15-HET mice exhibited no gross skeletal
102
abnormalities in comparison to WT at 5-days old and 21-days old. We did not identify
any changes in long bones lengths or abnormalities of the cranial skeleton (formed
through endochondral and intramembranous ossification respectively) (Fig 4.7 E & F,
4.8 E & F). In addition, there were no differences in tibial epiphyseal growth plate zone
lengths or overall growth plate height between WT and MetΔ15-HET mice. Overall, this
indicates murine skeletal development and growth does not appear to be affected by the
MetΔ15-HET.
In OFD, there is abnormal osteoblast differentiation at the lesional tissue (shown by the
presence of cells positive for both osteoblast and fibroblast cell markers). The healthy
tissue surrounding these lesions and the rest of the skeleton are normal (Gray et al.,
2015). It is plausible that the osteoblast differentiation defect in MetΔ15-HET BMSCs
does not affect normal development but does affect bone repair. Another plausible
explanation for the lack of a skeletal phenotype in mice may be due to their more robust
regenerative ability (Haffner-Luntzer, Kovtun, Rapp, & Ignatius, 2016). Lastly, the
absence of a skeletal phenotype could be because MetΔ15-HET mice still exhibit partial
regulatory control of Met signaling and not a complete block in osteoblast differentiation.
This notion is supported by the embryonic lethality seen in MetΔ15-HOMO mice, which
may be caused by the inability to form an organized cartilaginous skeletal precursor
during embryogenesis (Ben-Zvi, Shilo, & Barkai, 2011).
5.5 MetΔ15-HET mice experience delayed fracture healing ability
To test whether the in vitro osteoblast differentiation defect in MetΔ15-HET mice impairs
bone healing, we examined fracture healing in vivo. Our MetΔ15-HET mice do not
develop spontaneous fractures and we thus induced fractures using a stabilized mid-
diaphyseal tibia fracture model. In mice fractures are typically healed 28 days post-
fracture, with peak soft callus formation occurring approximately 7 – 9 days post-
fracture, accompanied by a peak in proteoglycan content while peak hard callus
formation is reached 14 days post-fracture (Marsell & Einhorn, 2011).
103
We decided to first assess the healing process 14-days post fracture, as this timepoint
allows for visualization of intramembranous and endochondral ossification and
remodelling which both require osteoblasts. In comparison to WT mice, MetΔ15-HET
mice still exhibit large amounts of cartilage callus 14-days post fracture, indicated by the
greater amount of Safranin-o staining (Fig 4.10 A). This suggests MetΔ15-HET fracture
sites have yet to transition from a soft to hard boney callus, a process in which
osteoblast are heavily involved. Accordingly, MetΔ15-HET fractures also displayed less
bone volume and more unmineralized matrix (osteoid) volume at the fracture site when
compared to WT mice at the same timepoint (Fig 4.10 B & C). Endochondral ossification
and osteoblasts are critical for the transition from soft callus to hard callus therefore a
defect in osteoblast differentiation would hinder this transition. In summary our data
indicates delayed fracture healing in MetΔ15-HET mice in comparison to their WT
counterparts.
Osteoclasts are intimately involved in the remodeling of the hard callus, a late stage
process in fracture healing, which starts approximately 13 – 14 days post-fracture.
Osteoclasts are typically in abundance at the fracture site 14-days post-fracture.
However, the MetΔ15-HET callus contained 50% less osteoclasts in comparison to WT
counterparts 14-days post fracture (Fig 4.10 D & E). We believe that this is further proof
of delayed fracture repair in the MetΔ15-HET mouse.
There is an intimate relationship between osteoclasts and osteoblasts, in which co-
regulation occurs through RANK-RANKL interactions. As previously mentioned,
osteoblasts secrete RANKL which binds the RANK receptor on osteoclast to facilitate
their maturation. Therefore, a defect in terminal osteoblast differentiation reducing
osteoblast formation will also reduce RANKL expression which is needed to induce
osteoclastogenesis (Katagiri & Takahashi, 2002).
The reduction in osteoclast differentiation could also be attributed to the upregulation of
adrenomedullin expression we previously identified. Along with ADM’s role in the
inhibition of GSK3β, it is also involved in the inhibition of RANK-RANKL interactions (Y.
Liu et al., 2017). Therefore, diminished osteoclast numbers seen in MetΔ15-HET fracture
104
calluses could be the result of reduced terminally differentiated osteoblasts (available to
produce RANKL) and increased ADM-mediated suppression of RANKL-RANK
interactions (Y. Liu et al., 2017).
5.6 MetΔ15-HET osteoblasts display dysregulation of β-catenin
When examining the gene expression profile of osteoblasts at day 21 of differentiation,
we found a dysregulation of osteoblast specific gene markers in MetΔ15-HETs. We saw
a 22-fold increase in β-catenin along with a 6.4-fold reduction in Axin2 expression, a
critical scaffold protein in the β-catenin degradation complex (Fig 4.5 B & C) (Jho et al.,
2002; Yan et al., 2009). The combination of increased expression of β-catenin and
reduced ability to degrade the β-catenin protein would explain the higher levels of β-
catenin and its phosphorylated version seen in MetΔ15-HETs. The elevated levels of β-
catenin occur at a timepoint at which β-catenin requires tight regulation for normal
osteoblast differentiation. Any deviations from this would result in terminal osteoblast
differentiation defect (Ghadakzadeh et al., 2016).
One consequence of elevated β-catenin levels is an upregulation of downstream target
genes such as Runx2. Here we report MetΔ15-HET osteoblasts exhibit a 27-fold increase
in Runx2 expression at the end of in vitro osteoblast differentiation (day 21) (Fig 4.5 E).
In addition, we saw a 1.3-fold reduction in alkaline phosphatase and 4-fold reduction in
osteocalcin, late stage osteoblastic markers, in MetΔ15-HET osteoblasts at day 21 (Fig
4.5 G & H). This suggests that the MET gain of function mutation signaling causes
inhibition of terminal osteoblast differentiation through dysregulation of β-catenin and
Runx2. This is consistent with what has been reported by multiple groups; the
overexpression of Runx2 during the maturation stages of osteoblast differentiation
prevents terminal differentiation (T. M. Liu & Lee, 2012; Wenguang Liu et al., 2001). In
addition, this also fits with what was described by Gray et al., (2015) who found no
changes in early markers of osteoblast differentiation but observed reductions in late
stage osteoblastic markers when the MET exon 15 skipping mutation was induced in an
osteoblastic cell line. In summary, these results provide a potential mechanism
105
explaining the observed in vitro osteoblast differentiation defect and the diminished
callus mineralization in MetΔ15-HET mice fractures (Fig 4.4 A & B, Fig 4.10 A – E).
Day 21 MetΔ15-HET osteoblasts were also found to exhibit a 24.6-fold increase in
adrenomedullin (ADM) expression (Fig 4.5 I), which is involved in the inactivation of
GSK3β. Since GSK3β is involved in the degradation of β-catenin, we would expect
higher levels of β-catenin and RAS, which we did indeed observe in MetΔ15-HET MEFs
(Fig 4.2 B) (Jeong et al., 2018). The upregulation of RAS was also accentuated by
downregulation of Rasal3 expression (Fig 4.5 J), an inactivator of RAS signaling (Muro
et al., 2015). The elevated levels of RAS protein and activity promotes increased
stabilization of β-catenin through ERK1/2-dependent and independent pathways (Jeong
et al., 2018).
Through comprehensive protein and gene expression analysis, we were able to
implicate a dysregulation of β-catenin as a potential reason behind the osteoblast
differentiation defect seen in our in vitro and in vivo experiments. Our findings are
consistent with the idea that β-catenin requires tight spatiotemporal regulation during
osteoblast differentiation and fracture healing. In the early stages of fracture repair,
precise regulation of β-catenin is required for mesenchymal stem cell differentiation to
osteoblasts. In the early stages of osteoblast differentiation, β-catenin positively
regulates osteoblasts through upregulation of Runx2 expression (Y. Chen et al., 2007).
In the late maturation stages, Runx2 expression is inhibited and terminal differentiation
to mature osteoblasts ensures. We propose, β-catenin levels are constitutively elevated
in MetΔ15-HET mice, resulting in upregulation of Runx2 during the maturation stages
hindering terminal differentiation.
Interestingly, others have also described a dysregulation of RAS and β-catenin levels in
pseudarthrosis secondary to neurofibromatosis-1 (NF-1) and fibrous dysplasia
(Ghadakzadeh et al., 2016; Regard et al., 2011). These bone diseases present similar
phenotypes to osteofibrous dysplasia and we propose they also share similar causative
mechanisms; through the dysregulation of β-catenin mediated Runx2 expression. Liu et
al., (2001) showed osteoblast specific overexpression of Runx2 in vivo resulted in a
106
maturational blockage in osteoblasts in vivo; mice exhibited reduced number of
terminally differentiated osteoblasts and osteocalcin expression. We report similar
findings in our mature MetΔ15-HET osteoblasts, which displayed upregulations in Runx2
expression, downregulation of osteocalcin, and a reduced ability to terminally
differentiate. Additionally, when Liu et al., 2001 overexpressed Runx2 in osteoblasts in
vivo, it resulted in osteopenia and the development of fractures. The loss of the
spatiotemporal control of the regulatory relationship between Wnt/β-catenin signaling
and Runx2 expression could be the commonality between diseases presenting with
pseudarthrosis phenotypes.
Accordingly, we believe manipulation of the Wnt/β-catenin signaling pathway could be a
potential therapeutic target in treating OFD patients exhibiting non-healing tibial
fractures. This approach has been previously tested in vivo in treating NF-1 related tibial
fracture. Dickkopf-1 mediated inhibition of Wnt/β-catenin signaling led to the union of
fractures and improved osteoblast differentiation (Ghadakzadeh et al., 2016).
In this study we were able to characterize and validate the use of a MetΔ15-HET genetic
mouse model to replicate the aberrant signaling seen in human osteofibrous dysplasia.
Through pathway analysis at the protein and gene expression level, we were able to
confirm aberrant MET signaling is caused by the failure to degrade the MET receptor
post-activation. We were also able to replicate the osteoblast differentiation defect seen
in human disease in vitro utilizing our MetΔ15-HET mouse model. In addition, we
propose a potential mechanism explaining the osteoblast differentiation defect seen in
OFD. MetΔ15-HET mouse skeletal characterization revealed an absence of gross
defects. When examining the fracture healing ability of WT and MetΔ15-HET mice, we
found MetΔ15-HETs displayed delayed fracture repair in comparison to WT mice.
Overall, these results support the hypothesis that gain-of-function MET mutations result
in diminished bone repair ability due to reduced osteoblast differentiation. We believe
this defect is caused by upregulated signaling at downstream effectors of aberrant MET
signaling resulting in constitutively elevated levels of β-catenin leading to loss of
spatiotemporal regulation of Runx2 expression and inhibition of osteoblast terminal
differentiation.
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Patients currently affected by OFD experience severe pain and disabilities due to non-
healing fractures, resulting in gait deformation and punishing reduction in quality of life.
Since the role of MET in bone formation is largely unstudied, we hope our data will pave
the way for further interventions to improve fracture healing in these patients.
5.7 Limitations to the Study
Through in vitro and in vivo experiments, we have been able to implicate MET’s role in
bone development and fracture repair, however, there were limitations in this study
which are discussed here.
Firstly, we used a model where the MET mutation was expressed in all cell types. This
makes it difficult to delineate and define the clear effects of the mutation on a specific
cell type such as osteoblasts. This limitation is especially important when examining
complex processes such as fracture repair which involve numerous cell types and
signaling pathways acting in concert. In the face of this limitation, the MetΔ15-HET
mouse model we have created and utilized does provide compelling evidence for the
role of MET in bone development and fracture repair. The use of an osteoblast targeted
conditional MetΔ15 mouse model would allow for more precise conclusions of the effect
of the osteoblast differentiation defect on fracture healing, it may not allow for us to
examine any broad changes in the fracture repair process.
This leads to the next limitation of this study; we did not extensively explore the role of
osteoclasts but rather focused on osteoblasts function. The harmonic relationship
between osteoblasts and osteoclasts has been well defined; they are known to co-
regulate and communicate through RANK-RANKL interactions. In addition, MET and
HGF also are involved in reciprocal signaling between osteoblasts and osteoclasts.
Osteoclast function may be impaired by the MetΔ15-HET mutation independent of
RANK-RANKL interactions. Investigation into the effects of the MetΔ15-HET mutation on
osteoclast differentiation and function may provide alternative explanations and
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mechanisms for why we see reduced osteoclast number at the fracture site 14-days
post-fracture.
Various animals models with enhanced angiogenesis have shown improved bone
regeneration abilities post-fractures, making modulation of vascularization post-fracture
a viable therapeutic approach in improving fracture outcomes (Hankenson et al., 2011).
We did not examine the effects of the MET mutation on vascularization, a key
component of the fracture repair process (Hankenson et al., 2011). Vascular invasion of
the fracture site supplies oxygen, nutrients and inflammatory cells to the fracture site
and contributes to the formation of a hematoma which acts as the template for the
formation of a vascular callus. Failure to vascularize the fracture site has been shown to
result in diminished repair and greater incidence of non-unions (Dickson, Katzman,
Delgado, & Conteras, 1994; Hankenson et al., 2011). This crucial process is regulated
by numerous growth factors, but most notably by VEGF signaling which has been
shown to be regulated by MET’s downstream pathways: PI3K/AKT, MAPK and STAT3
(Matsumura et al., 2013). Therefore, it is highly plausible the MET mutation may affect
vascularization during the fracture repair process.
Many of the pathway analysis experiments utilized MEFs instead of osteoblasts;
although the results may be representative of what is seen in osteoblasts, there still may
be subtle differences which cannot be ignored. This is despite the fact they share many
similarities such as both being a part of the connective-tissue cell family, secrete type-1
collagen and are responsible for the structural framework of the body (Mackie, Ahmed,
Tatarczuch, Chen, & Mirams, 2008). Osteoblast differentiation is a dynamic process
with multiple key stages such as the commitment of MSCs to pre-osteoblasts and
maturation of immature osteoblasts. These stages involve distinctive regulation of key
genes and signaling pathways therefore utilizing MEFs results in loss of temporal
information and cannot capture the dynamic properties of the process. Nonetheless,
MEFs allow for a broad overview of the potential differences in signalling and
expression between WT and MetΔ15-HET mutants, and therefore still provide valuable
knowledge on the effects of aberrant MET signaling. In addition, the technical benefits
and advantages of using fibroblasts due to their resilient characteristics and ability to
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thrive, making them one of the easiest cells to grow in cell culture, were why they were
chosen in favour of osteoblasts (Alberts et al., 2002).
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5.8 Proposed Mechanism for the Dysregulation of β-catenin Signaling in Osteoblast Differentiation
During the maturation stages of osteoblast differentiation:
1) Loss of juxtamembrane domain results in diminished degradation of the MET
receptor causing upregulation of signaling
2) Upregulation of MET signaling upregulates downstream pathways: AKT-mTOR
and RAS-MEK-ERK1/2 through upregulated phosphorylation events
3) Aberrant MET signaling results in upregulated ADM mediated inhibition of
GSK3β activity and downregulation of RASAL3 mediated inactivation of RAS
activity
Overall Effect: inappropriate elevated expression of Runx2 at a timepoint at which
Runx2 inhibits terminal osteoblast maturation resulting in a differentiation defect.
I. Upregulated AKT signaling results in
upregulation of phosphorylation of β-catenin
at Ser552, enhancing its stability and
promoting translocation to the nucleus to
upregulate Runx2 expression through
interactions with LEF/TCF transcription factors
II. Upregulated AKT activity increases mTOR
activity
III. Upregulated mTOR signaling directly
upregulates Runx2 expression through the
mTORC1 complex
I. At the same time, upregulation of RAS
signaling enhances β-catenin stability and
translocation to the nucleus resulting
upregulation of Runx2
II. Upregulated RAS activity increases ERK1/2
activity
III. Upregulated ERK1/2 signaling activity, can
result in upregulation of LRP6 phosphorylation
leading to inhibition of β-catenin degradation
complex activity. Therefore β-catenin stability
and translocation to the nucleus is enhanced
resulting in upregulation of Runx2
Pathway 1 Pathway 2
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Figure 5.1. Proposed Mechanism for the Dysregulation of β-catenin Signaling in Osteoblast Differentiation
112
6 Conclusions
We initially hypothesized that gain-of-function MET mutations result in diminished bone
repair due to reduced osteoblast differentiation. To test this hypothesis, we 1)
determined MET pathway irregularities between WT and MetΔ15-HET mice, 2)
characterized the skeletal phenotype of WT and MetΔ15-HET, and 3) compared fracture
healing between WT and MetΔ15-HET mice in vivo.
We first validated the pathway signaling profile of our MetΔ15-HET mouse model to
ensure it captured the same dysregulations in signaling seen in OFD patients. MetΔ15-
HET MEFs exhibit upregulated protein levels of MET, its downstream effectors and their
phosphorylated versions in comparison to WT MEFs (Fig 4.1A – C). The absence of an
upregulation in MET gene expression suggests the elevated protein levels is due to a
failure in degradation of the receptor, while elevated levels of downstream effectors
indicated elevated basal signaling activity.
When HGF stimulation was applied, MetΔ15-HET MEFs exhibited higher upregulation of
signaling activity and longer sustained signaling activity in comparison to WT MEFs.
This is indicative of a diminished ability for MetΔ15-HETs to degrade the MET receptor
and terminate signaling activity. Overall these results confirm the MetΔ15-HET mouse
model replicates the dysregulations in signalling seen in humans as described by Gray
et al., (2015).
Next, we examined osteoblast differentiation through CFU assays of primary BMSCs
induced to differentiate osteoblasts. Here we found MetΔ15-HET BMSCs displayed a
reduced ability for differentiation and mineralization in comparison to WT counterparts,
forming 35% less CFU-ALP colonies and 80% less CFU-OBs at day 21 (Fig 4.4B). This
demonstrates an osteoblast differentiation defect in MetΔ15-HET BMSCs.
We then examined if there were any differences in gene expression profiles between
WT and MetΔ15-HET mutants. We found multiple pathways and factors with
dysregulated signaling and expression resulting in the upregulation of β-catenin levels
and Runx2 expression during the late stages of osteoblast differentiation (osteoblast
maturation). These two factors require strict spatiotemporal regulation of expression and
113
activity in order to achieve terminal osteoblast differentiation. β-catenin mediated
upregulation of Runx2 expression during a time (day 21 of osteoblast differentiation) at
which Runx2 inhibits the progression of osteoblast differentiation leads us to believe
dysregulation of β-catenin signaling activity is the cause of the differentiation defect
seen. This conclusion is supported by the literature, as diseases with similar bone
phenotypes have also been shown to be caused by dysregulations in spatiotemporal
regulation of β-catenin (Ghadakzadeh et al., 2016).
Since we found dysregulation of multiple signaling pathways and an osteoblast
differentiation defect in MetΔ15-HET mutants, we examined if these mutant mice
exhibited any gross skeletal phenotypes. Through whole skeletal prep and epiphyseal
growth plate analysis, we found there were no differences in intramembranous and
endochondral bone size (Fig 4.7 & 4.8). These results suggest perhaps the defect seen
in humans is a stress induced defect caused by fractures. This fits in line with the
characterization of OFD in which osteoblast differentiation defects are only seen in
lesional tissue but not the surrounding healthy tissue resulting in a distinctive zonal
architecture.
We then moved on to examination of fracture healing in vivo and found 12-week old
male MetΔ15-HET mice exhibited delayed fracture healing in comparison to WT
counterparts at 14-days post-fracture. We saw increased cartilage at the callus site
during a timepoint at which there should be minimal, as the soft callus should have been
converted to hard callus. Furthermore, we saw minimal osteoclast number at the callus
in MetΔ15-HET mice confirming the remodeling process has yet to begin, while WT
calluses were abundant in osteoclasts already. This points to MetΔ15-HET mice being at
a less advanced stage of fracture repair due to a delay in the mineralization of the hard
callus, a process in which osteoblasts are intimately involved in. This leads us to
conclude overactivation of MET signaling delays fracture healing due the inability for
osteoblasts to terminally differentiate and mineralize the soft callus.
Overall, this study demonstrates overactive MET signaling results in stabilization of β-
catenin and upregulated Runx2 expression causing loss of the tight spatiotemporal
regulation required for terminal osteoblast differentiation. The inability for osteoblasts to
114
terminally differentiate results in the delays in fracture healing seen in our mouse model
and OFD patients. Further interventional studies against the MET and β-catenin
signaling pathway may result in the development of novel therapeutic strategies against
OFD and similar diseases before resorting to invasive surgical means.
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7 Future Directions
Osteofibrous Dysplasia (OFD) is a rare benign non-neoplastic condition, caused by
mutations in the MET gene, in which patients experience reduced fracture healing and
osteoblast differentiation ability. The MET receptor is a regulator of numerous
proliferation and differentiation pathways critical for osteoblast differentiation. In this
study, we demonstrate the loss of spatiotemporal control of Runx2 expression due to
dysregulation of β-catenin levels results in an osteoblast differentiation defect and
delayed fracture healing. This further supports and reiterates the need for tight control of
β-catenin levels during osteoblast differentiation and coordinated expression of Runx2
depending on the stage of osteoblast differentiation (Ghadakzadeh et al., 2016).
To better understand the dysregulated pathways in osteoblast differentiation some
future directions would include pathway analysis of MET and its downstream effectors
throughout the differentiation process. These results may reveal the exact stage of
osteoblast differentiation the dysregulation of β-catenin occurs at. This would be done
by examining protein and gene expression levels at the following timepoints: day 2 of
osteoblast differentiation (commitment to osteoblast differentiation), day 7 (pre-
osteoblast to immature osteoblast transition), day 14 (maturation of immature osteoblast
begins) and day 21 (terminally differentiated). The results of these experiments would
provide a comprehensive timeline of the modulations and dysregulations which occur
due to the MetΔ15-HET mutation.
To further confirm the role of MET signaling in osteoblast differentiation, rescue
experiments would need to be performed using selective MET inhibitors such as ARQ
197 (Jeay et al., 2007). MET inhibitors would be applied during the osteoblast
differentiation process to determine if the defect seen in Met Δ15-HET mice can be
improved. This would be measured by colony forming unit assays, protein expression,
and gene expression analysis of osteoblastic markers such as Runx2, ALP and
osteocalcin. A successful rescue would show improvements in CFU-ALP and CFU-OB
number and restoration of the aforementioned osteoblast differentiation markers to WT
levels. The ligand-dependency of aberrant MET signaling could be further examined by
treatment of Met Δ15-HET MEFs with NK4. NK4 is a HGF specific antagonist which
116
competitively inhibits the binding of HGF to the MET receptor and prevents
transphosphorylation post-activation (Date, Matsumoto, Shimura, Tanaka, & Nakamura,
1997; Du et al., 2007). Reduction of the elevated levels of phosphorylated MET and its
downstream effectors seen in Met Δ15-HET MEFs, would reconfirm the ligand-dependent
nature of this mutation.
Fracture repair is a complex process involving a delicate balance between osteoblast
and osteoclast activity. Though the main focus of this study was focused on the role of
osteoblasts, the role of osteoclasts cannot be ignored. In this study we demonstrate the
MET exon 15 skipping mutation may result in an osteoclast differentiation defect as well
with a reduction in osteoclast number at the callus site 14-days post fracture in Met Δ15-
HET mice. This is not so far-fetched as the primary method of osteoclastogenesis
induction is through the communication of osteoblast and osteoclasts through RANK-
RANKL interactions (Park-Min, 2018). In addition, MET and HGF participate in
reciprocal signaling between osteoblasts and osteoclasts as well, therefore it is not
unlikely osteoclast function and differentiation may also be impacted by this mutation.
This would be done by performing the pathway analysis experiments highlighted in this
study utilizing BMSC induced osteoclast cultures (Bradley & Oursler, 2008). In addition,
a bone resorption functional assay would be performed to examine any potential
differences in osteoclast function. The results of these experiments could also provide
alternative explanations and mechanisms explaining the reduction in osteoclast number
seen at the callus site of Met Δ15-HET mice 14-days post fracture.
Lastly, a crucial and necessary step in this line of research would be to transition from
mouse and animal model experiments to human clinical trials. The transability and
practicality of targeting some of these elucidated pathway effectors would need to be
examined as well. Only then would we be closer to developing novel therapeutic
treatment plans against reduced fracture healing in osteofibrous dysplasia patients.
117
8 Supplementary Results 8.1 Confirmation of Exon 15 Skipping Event
The occurrence of the Exon 15 skipping event was confirmed in our Met Δ15-HET mouse
model utilizing PCR primers targeting Exon 14 and Exon 16 followed by PCR
amplification. Therefore, in complete Met transcripts (with Exon 15), Exon 14 – 16 would
be amplified by PCR, resulting in the formation of a 469 bp product. The exclusion of
Exon 15 would result in the formation of a 319 bp PCR product. Through gel
electrophoresis mediated resolution, we were able to confirm the exon skipping event by
differentiating PCR product sizes. Here we found WT osteoblasts to exhibit both
complete Met transcripts along with a small proportion of Met transcripts excluding Exon
15 as well. This coincides with what was reported by Gray et al., (2015) who found the
Met exon 15 skipping event to occur during development in mice. In addition, as
expected we see MetΔ15-HET osteoblasts exhibiting both complete Met and MetΔ15
transcripts confirming the exon skipping event in our genetic mouse model.
WT MetΔ15-HET
100
200
300
400
500
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