c-Src Controls Mouse Embryonic Osteogenic Differentiation Through Regulation of Stat1 Stability

by

Zahra Alvandi

A thesis submitted in conformity with the requirements for the degree of Doctor of Philosophy

Department of Laboratory Medicine and Pathobiology University of Toronto

© Copyright by Zahra Alvandi 2018

ii

c-Src Controls Mouse Embryonic Osteogenic Differentiation

Through Regulation of Stat1 Stability

Zahra Alvandi

Doctor of Philosophy

Department of Laboratory Medicine and Pathobiology University of Toronto

2018

Abstract

The protein tyrosine kinase Src is expressed ubiquitously and is involved in differentiation.

However, the role of Src in osteogenic differentiation is complex since it has been reported to

function both as repressor and activator. Using the small molecule inhibitor PP2, c-Src specific

siRNAs, and tet-inducible lentiviral vectors overexpressing constitutively active c-Src, I show c-

Src inhibitory role in mouse embryonic stem cells (mESCs) and mouse preosteoblast MC3T3-

E1s osteogenic differentiation. I further predicted that the increased level of osteogenic

differentiation is regulated through Runt-related transcription factor 2 (Runx2). Additional

analysis revealed that active c-Src (p-Y416-c-Src) lowers Runx2 nuclear localization and

transcriptional activity in mESCs without having any effect on Runx2 expression level. I provide

the first evidence linking inhibitory role of c-Src to Runx2 subcellular localization through signal

transducer and activator of transcription 1 (Stat1), the cytoplasmic anchoring partner of Runx2. I

discovered that higher level of active c-Src increases Stat1 protein half-life by inhibiting Stat1

proteasomal degradation. Therefore, by inhibition of c-Src activity, Stat1 protein level is reduced

resulting in less of interaction with Runx2. This in turn increases nuclear localization and

transcriptional activity of Runx2. Collectively, my study has defined a new mechanism, by

which c-Src activity inhibits transcriptional regulation of osteogenesis.

iii

Acknowledgments

First and foremost, I would like to express my sincere gratitude to my supervisor, Prof. Michal

Opas, for giving me the opportunity to pursue my PhD degree in his lab. Dr. Opas, you

welcomed me to your lab when I was extremely frustrated and hopeless and provided me with

the most peaceful environment so that I could focus on my research without any unwanted

distractions. And again, you supported me not only through years of research but you constantly

treated me with care I needed to overcome many challenging stages of my life in the past few

years. You made me a better researcher and you changed my vision in the most positive way

possible. Thank you!

My sincere thanks also go to my committee members, Prof. Jane Mitchel, and Prof. Christopher

McCulloch for their guidance and invaluable comments and suggestions which absolutely

improved the quality of my research and my thesis.

I am also grateful to former and current members of Opas lab, with especial thanks to Shirley

Yu, and Dr. Carlos Pilquil, for their help, guidance, and friendship.

I am also thankful to my wonderful friends, Bahareh, Shaghayegh, Saeed, Sheida, and Navid for

their amazing friendship, for letting me share my concerns, for being there when I needed them,

and for their kindness.

I want to dedicate this thesis to my wonderful parents, my lovely sister, Farnaz, and my one of a

kind brother, Koosha for being so understanding and supportive even though I could not be with

them during some of their toughest moments in their lives. This has been all possible because of

your constant support, care, and love all these years.

And finally, I would like to thank my husband, Farrokh, who supported me and helped me make

better decisions. Farrokh, you made this journey more exciting and efficient. Thank you for

toughening me up and pushing me constantly through the whole process.

iv

Table of Contents

Acknowledgments.......................................................................................................................... iii

Table of Contents ........................................................................................................................... iv

List of Abbreviations .................................................................................................................... vii

List of Tables ................................................................................................................................. ix

List of Figures ..................................................................................................................................x

List of Appendices ......................................................................................................................... xi

Chapter 1 ..........................................................................................................................................1

Introduction .................................................................................................................................1

1.1 Bone and Bone Formation ...................................................................................................1

1.1.1 Differentiation of Osteoblast Lineage ......................................................................3

1.2 Sources of Stem Cells for Osteogenic Differentiation .........................................................4

1.2.1 Embryonic Stem Cell Osteogenic Differentiation ...................................................7

1.3 Development of Osteoblast Phenotype ................................................................................9

1.3.1 BMP/ TGF β pathway ............................................................................................10

1.3.2 Wnt pathway ..........................................................................................................10

1.3.3 Ca2+ pathway ..........................................................................................................11

1.3.4 FGF pathway ..........................................................................................................11

1.3.5 Notch pathway .......................................................................................................12

1.4 Osteoblast Transcription Factors and ECM Proteins .........................................................12

1.5 Runx2 .................................................................................................................................14

1.5.1 Regulation of Runx2 Transcriptional Activity via Post-Translational Modifications .........................................................................................................15

1.5.2 Regulation of Runx2 Transcriptional Activity via Runx2-Interacting Partners ....16

1.6 Regulation of Osteogenic Differentiation by c-Src ...........................................................18

1.6.1 c-Src Structure and Function .................................................................................19

v

1.7 Hypothesis and Objectives .................................................................................................21

Chapter 2 ........................................................................................................................................22

Materials and Methods ..............................................................................................................22

2.1 22

2.1.1 Cell Culture ............................................................................................................22

2.1.3 Immunoprecipitation assay .......................................................................................23

2.1.4 Real-time PCR analysis ............................................................................................23

2.1.5 Chromatin Immunoprecipitation ...............................................................................23

2.1.6 Immunoblot analysis .................................................................................................24

2.1.7 Mineral deposition assays .........................................................................................25

2.1.8 Immunofluorescence (IF) staining ............................................................................25

2.1.9 Subcellular Fractionation ..........................................................................................26

2.1.10 Transfection ............................................................................................................26

2.1.11 Lentiviral virus production and transduction ..........................................................26

2.2 Statistical analysis ........................................................................................................27

Chapter 3 ........................................................................................................................................28

3.1 c-Src Activity Inhibits Osteogenic Differentiation in mESCs ............................................28

3.1.1 Osteogenic Differentiation in Mouse Embryonic Stem Cells ...................................28

3.1.2 c-Src Expression Profile in Mouse Embryonic Stem Cells During Osteogenic Differentiation ........................................................................................................31

3.1.3 Inhibition of c-Src at Different Periods During ES Cells Osteogenic Differentiation ........................................................................................................32

3.1.4 Overexpression of p-Y416-c-Src in MC3T3-E1s and its Effect on Osteogenic Differentiation ........................................................................................................41

3.2 c-Src Activity Inhibits Runx2 Nuclear Localization ...........................................................45

3.2.1 The Effect of c-Src Activity on Runx2 Target Genes’ Expression ..........................45

3.2.2 The Effect of c-Src Activity on Runx2 Expression ..................................................46

vi

3.2.3 c-Src Activity and its Effect on Runx2 Subcellular Localization .............................49

3.3.4 Inhibition of c-Src activity lowers Runx2 and Stat1 interaction ...............................51

3.3 c-Src Regulates Osteogenic Differentiation Through Stat1 ................................................53

3.3.1 c-Src effect on Runx2 localization and transcriptional activity during osteogenic differentiation is significantly reduced in the absence of Stat1 .............................53

3.3.2 c-Src Activity and its Effect on Stat1 Subcellular Localization ...............................55

3.3.3 The Effect of c-Src Activity on Stat1 Expression and its Half-life ..........................57

3.3.4 The Effect of c-Src Activity on Stat1 Degradation ...................................................61

Chapter 4 ........................................................................................................................................65

4 Discussion, Final Conclusions, and Future Directions ...............................................................65

4.1 Thesis summary ..................................................................................................................65

4.2 General Discussion and Conclusions ..................................................................................68

4.3 Future Directions .................................................................................................................73

4.3.1 Regulation of c-Src by Calreticulin ..........................................................................73

4.3.1 Regulation of Stat1 Degradation by ERK Downstream of c-Src .............................74

4.3.1 Stat1 Proteasomal Degradation by c-Src/SIAH2 ......................................................74

References ......................................................................................................................................75

Appendices .....................................................................................................................................88

98

vii

List of Abbreviations

AA Ascorbic Acid

ALP Alkaline Phosphatase

ARS Alizarin Red Staining

B-Gly β-glycerophosphate

BMP Bone Morphogenic Protein

BLC Bone Lining Cell

BSP Bone sialoprotein

CaM Calmodulin

CaMK

CRT

Calmodulin kinase

Calreticulin

CHX Cycloheximide

COL1 Type I Collagen

Dex Dexamethasone

DMEM Dulbecco’s modification eagle’s medium

EB Embryoid Bodies

EGTA Ethylene Glycol Tetraacetic Acid

ERK Extracellular Signal-Related Protein Kinase

ESC Embryonic stem Cell

FBS Fetal Bovine Serum

FGF Fibroblast Growth Factor

FITC Fluorescein Isothiocyanate

FZD Frizzled

GSK Glycogen Synthase Kinase

H3 Histone 3

HAT Histone Acetylase

HDAC Histone Deacetylase

HSC Hematopoietic Stem Cell

IF Immunofluorescence

IFN

iPS

Interferon

Induced Pluripotent Stem Cells

viii

LIF Leukemia Inhibitory Factor

MAPK Mitogen-Activated Protein Kinase

MEF Mouse Embryonic Fibroblast

mESC Mouse Embryonic Stem Cell

MSC Mesenchymal Stem Cell

NEAA Non-Essential Amino Acids

NFAT Nuclear Factor of Activated T Cell

NLS Nuclear-Localization Signal

NMTS Nuclear Matrix Targeting Signal

OC Osteocalcin

OPN Osteopontin

OSE Osteoblast-Specific Element

Osx Osterix

PB Polybrene

PST Proline/Serine/Threonine

PTH Parathyroid Hormone

PTHrP Parathyroid Hormone-Related Protein

Pi

QA

Inorganic Phosphate

Glutamine/Alanine

RA Retinoic Acid

Runx2 Runt Related Expression Factor 2

SFK c-Src Family Kinase

SH Src Homology

SHPII

Smad

Src Homology Phosphatase II

Similar to Mothers Against Decapentaplegic

STAT Signal Transducer and Activator of Transcription

TGF Transforming Growth Factor

Tyr Tyrosine

Ub Ubiquitin

YAP Yes-Associated Protein

ix

List of Tables

Table 1-1 Advantages and disadvantages of ESCs and MSCs for cell-based therapies………….6

x

List of Figures

Figure 1-1 Osteogenic differentiation protocol…………………………………………….9

Figure 1-2 The domain structure of inactive and active c-Src……………………….……20

Figure 3-1 Expression of osteogenic markers during osteogenic differentiation in mESC.29

Figure 3-2 Alizarin Red Staining (A) and Von Kossa staining (B) of day 21 osteo-nodules…………………………………………………………………………………30

Figure 3-4 Inhibition of c-Src activity between day 6-10 enhances mES osteogenesis......32

Figure 3-5 c-Src inhibitory effect on OC expression and mineralization is dose dependent………………………………………………………………………..……..37

Figure 3-6 Gene expression of osteogenic markers in MC3T3-E1……………………….39

Figure 3-7 c-Src downregulation in MC3T3-E1 increases osteogenic differentiation…....40

Figure 3-8 Overexpression of constitutively active c-Src reduced osteogenic differentiation………………………………………………………………...………...43

Figure 3-9 Inhibition of c-Src does not affect Runx2 expression in mESCs and MC3T3-E1s……………………………………………………………………………………...47

Figure 3-10 Runx2 nuclear localization is increased when c-Src activity is inhibite……..48

Figure 3-11 Inhibition of c-Src activity increases Runx2 transcriptional activity in mESCs………………………………………………………………………………….50

Figure 3-12 c-Src inhibition lowers Runx2-Stat1 interaction…………………….…….....52

Figure 3-13 c-Src inhibitory role on osteogenic differentiation is Stat1-dependent……....54

Figure 3-14 Stat1 phosphorylation at Y701 and its nuclear localization is increased in response to c-Src inhibition in mESCs…………………………………………………56

Figure 3-15 Stat1 protein stability is icreased when c-Src activity is inhibited……….…..59

Figure 3-16 Inhibition of c-Src activity increases ubiquitin-mediated Stat1 proteolysis………………………………………………………………………………63

Figure 4-1 c-Src activity inhibits osteogenic differentiation through regulation of Stat1 stability………………………………………………………………………………... 67

xi

List of Appendices

Figure A-1 Expression of pluripotency markers in early mES osteogenic differentiation…..88

Figure A-2 Inhibition of c-Src activity by pharmacological inhibitors……..……………......89

Figure A-3 Failed attempts for transfecting mESCs with c-Src siRNAs……..………...........90

Figure A-4 Stat1 phosphorylation status on Y701 in early days of osteogenic differentiation92

Figure A-5 Shp2 inhibition increases Stat1 Y701 phosphorylation and Runx2 localization...93

Figure A-6 Shp2 inhibition increases ERK activity……………...…………………………..94

Table A-1 Primer pairs used in qPCR analysis in this study.………………...…………..…..95

Table A-2 Primer pairs used in ChIP analysis in this study……………………...……..…....96

Table A-3 c-Src specific siRNA fragments used in this study………………..……...….…...97

1

Chapter 1

1 Introduction

1.1 Bone and Bone Formation

Current strategies in bone regenerative medicine are focused particularly on cell-based bone

substitutes. To develop an advanced bone substitute, it is necessary to understand the

osteogenesis processes in nature. Full understanding of signaling mechanisms responsible for

osteogenesis will optimize current strategies and create new approaches in bone regenerative

medicine.

Bone is a living, continuously self-renewing, specialized connective tissue with elastic and

strength properties which provides support and physical protection to various vital organs of the

body. It is estimated that in adult human body, the entire skeleton is renewed every 7 years. This

shows the dynamic balance of bone throughout the life. Bone is composed of two major cell

populations; osteoblast and osteoclast lineages in addition to the ECM composed of organic and

mineral phases (Aguila and Rowe, 2005). Bone formation by osteoblasts and resorption by

osteoclasts are tightly regulated processes responsible for continuous bone remodeling. The

dynamic balance of bone involves a complex coordination of multiple bone marrow cell types. In

adults, bone marrow mesenchymal stem cells (MSCs) differentiate through a series of stages in

which the progenitors give rise to the osteoblast lineage cells, later transferred into osteocytes

(Bianco, 2014) whereas osteoclasts originate from hematopoietic stem cell precursors (HSCs)

along the myeloid lineage differentiation (Feng and Teitelbaum, 2013). The imbalance between

bone formation and resorption results in various diseases, such as osteopetrosis, osteopenia, and

osteoporosis. Therefore, modulating osteogenic lineage commitment of stem cells could provide

effective therapeutic regime for related bone diseases.

During vertebrate embryogenesis, MSC gives rise to osteoblasts through two distinct processes

called intramembranous and endochondral ossification. During intramembranous ossification,

mesenchymal progenitors condense and directly differentiate into osteoblasts. The osteoblasts

from intramembranous ossification is limited to certain parts of the skull, as well as to a part of

2

the clavicle in mammals, whereas endochondral ossification produces osteoblasts in the rest of

the skeleton (Berendsen AD1, 2015). The endochondral bone tissue is generated through

cartilage intermediate consisting of many cells including mesenchymal- derived chondrocytes,

osteoblast and osteocytes (Akiyama and de Crombrugghe, 2009). The process of replacing

cartilage matrix produced by chondrocytes with trabecular bone which is made by osteoblasts

has been the subject of debate for years. The main source of osteoblasts in endochondral

ossification has been referred to progenitors in periosteum (Colnot, 2009). However, more

recently, chondrocytes have been also identified as a major source of osteoblasts contributing to

endochondral bone formation (Zhou et al. 2014).

Osteoblast lineage cells are a group of cells that includes mesenchymal progenitors, pre-

osteoblasts, osteoblasts (often called mature osteoblasts), bone-lining cells and osteocytes. A

brief description of these cells is provided below.

Mesenchymal progenitors or stem cells are adult stem cells that are found in bone marrow and

have the potential to differentiate to lineages of mesenchymal tissues including adipocytes,

chondrocytes, and osteoblast (Pittenger et al. 1999).

Preosteoblasts, are cells in the less mature stage of osteoblast differentiation and count as

precursor cells to osteoblasts. They are defined by expression of Runx2, the master regulator

of osteogenic differentiation. Endochondral and intramembranous ossifications are assumed to

be merged at the level of preosteoblasts and progress to differentiate.

Osteoblasts, the chief bone-making cells, have a capacity to produce a unique combination of

large amount of extracellular proteins including osteocalcin (OC), alkaline phosphatase (ALP),

and a large amount of type I collagen (COL1). These markers will be more extensively described

later in this chapter. When first deposited, the ECM which is rich in COL1 and not yet

mineralized is known as the osteoid. Osteoid is subsequently mineralized through the

accumulation of calcium phosphate forming a hard but lightweight composite material called

hydroxyapatite. Hydroxyapatite is the most stable form of calcium phosphate compound under

physiological condition that its application and properties has been reviewed elsewhere (Fihria

A., 2017).

3

Bone lining cells (BLCs) cover non-remodeling bone surface and have the potential to be

induced to proliferate and differentiate into osteogenic cells. Expression of CBFA1/Runx2 in

BLCs supports the notion that these cells belong to the osteoblastic lineage (WeinEmail M.N.,

2017). It has been suggested that they play roles in initiation of signaling involved in bone

resorption and bone remodeling (Miller et al. 1989).

Osteocytes, derived from osteoblasts, reside within bone and communicate extensively with each

other and other bone cell populations through gap-junctions to regulate bone metabolism. The

mature osteocyte expresses the protein sclerostin (SOST), a negative regulator of bone mass.

In normal physiologic states, SOST acts on osteoblasts at the surface of bone and is differentially

expressed in response to mechanical loading, and inflammatory molecules (Compton and Lee,

2014).

Differentiation of osteoblasts from mesenchymal osteo-progenitors is mainly evaluated based on

the expression of genes that are associated with the onset of osteoblast differentiation or control

the progression of differentiation and will be further described in the following (sub) sections.

1.1.1 Differentiation of Osteoblast Lineage

The differentiation process of osteoblasts often is mediated by mesenchymal progenitors,

preosteoblasts and osteoblasts. Different markers have been used to distinguish one group of

cells from others. For instance, osteoblasts are often characterized by the expression of OC, but

the molecular markers for the mesenchymal progenitors are a matter of debate. It has been

suggested that all osteo-chondroprogenitor cells derive from SOX9- expressing precursors during

mouse embryogenesis (Akiyama H1, 2005). Preosteoblasts encompass all cells transitioning

from progenitors to mature osteoblasts and therefore are, by definition, heterogeneous. However,

they normally express the transcription factor Runx2 or, at a more advanced stage of

differentiation, both Runx2 and osterix (Osx; also known as SP7). A subset of osteoblasts can

become osteocytes upon being incorporated within the bone matrix. The rest of the osteoblasts

are thought to either undergo apoptosis or transform into inactive BLCs (Bonewald, 2011).

There are three major stages of osteoblastogenesis: 1) proliferation, 2) extracellular matrix

(ECM) development and maturation, and 3) mineralization with characteristic changes in gene

expression. At early stage osteoblasts predominantly, express genes that guarantee competency

4

for proliferation and progression of cell cycle. ECM biosynthesis is also initiated at this stage. In

the next stage proliferated cells start their developmental transitions and exit from cell cycle.

Although the ECM biosynthesis will be continued, the expression of genes associated with

proliferation must be downregulated to further allow maturation of ECM. This is when maximal

expression of alkaline phosphatase (ALP) is coupled with osteopontin (OPN) expression, the

other phenotypic marker of osteoblast. After this stage cells are competent to become

mineralized and this is when the expression of OC elevates significantly (Owen TA., 1999;

Rutkovskiy A., 2016). Subsequently, enrichment of the organic scaffold with OC promotes

deposition of mineral substances which mainly includes calcium phosphate.

1.2 Sources of Stem Cells for Osteogenic Differentiation

Two major sources of stem cells for bone regeneration include MSCs and ESCs. Adult stem cells

and ESCs can be used for allogenic transplantation to alleviate the problem of shortage in donor

cells, tissues and organs in transplantation medicine for the treatment of a broad spectrum of

diseases.

MSCs are traditionally known to reside in bone marrow and are capable to differentiate to tissue

lineages including bone, cartilage, adipose, and muscle (Long F., 2001). Two different

differentiation models have been described for MSCs. In one model, MSCs are defined as

multipotent cells that could differentiate into all connective tissue cell types which under defined

environmental conditions can commit to specific differentiation pathways. In an alternative

model, MSCs have different differentiation potentials and the initial stage of differentiation

would limit the lineage potential (Baksh D., 2004). Trans-differentiation of other tissue types

derived from adult stem cells including adipogenic (Lee JA., 2003; Dragoo JL., 2003), myogenic

(Rauch C., 2002) and hematopoietic (Olmsted-Davis EA., 2003) towards osteogenic lineage has

been reported.

The other source of stem cells are ESCs which are derived from the inner cell mass of blastocyst

stage embryos (Thomson et al. 1998; Reubinoff et al. 2000). ESCs have the potential to

differentiate to any types of tissues and possibly to indefinitely proliferate in undifferentiated

status. Specific transcription factors have been associated with undifferentiated status of stem

cells. Sox2, Nanog, and Oct3/4 are crucial for the maintenance of pluripotency (Jaenisch and

Young, 2008) and their expression have been used to monitor pluripotency and initiation of

5

differentiation in this study. ES cells could be maintained in undifferentiation status by growing

on mouse embryonic fibroblasts (MEFs) or by treating them with leukemia inhibitory factor

(LIF). LIF is one of the feeder-cell-derived molecules which has been shown to play an

important role in the maintenance of ES pluripotency. Upon removal of mouse ES cells from

MEF or/and LIF these cells have the capacity to differentiate into hundreds of cell types and

could be induced to form any specific lineage/tissue. In general, three main approaches are used

to initiate ES cell differentiation. In one method, ES cells are cultured on stromal cells and go

through differentiation when they are in contact with these cells (Nakano et al. 1994). In another

protocol ES cells are cultured on a monolayer of ECM proteins to be induced to differentiate

(Keller G., 2005). The most common approach of ES differentiation methods though is

formation of 3D cell aggregates in suspension known as embryoid bodies (EBs) which offers the

advantage of an increase in cell-cell interactions (Keller G., 2005). It has been reported that loss

of pluripotency in EBs occurs reasonably rapidly within 6 days of EB development (Pineda et al.

2013). In our hands, this has been verified based on the expression of the pluripotent

transcription factors including Nanog, Oct3/4, and Sox2 which reaches to undetectable level

analyzed by qPCR after 6 days. However, advantages and disadvantages have been identified for

all described differentiation methods. For instance, ES cells grow better when are cocultured

with stromal cells. However, cell lineage screening would be extremely difficult due to technical

difficulty for cell isolation. Also, uncontrolled secretion factors from stromal cells may promote

ES cells toward differentiation to undesired cell types. Moreover, isolation of ES-derived cells

from culture at any stage of differentiation would be challenging. In terms of matrix proteins

monolayer as the base surface for ES cells could dramatically influence the differentiation of the

cells and may even bias development of certain cell types. In the case of EB formation for

differentiation although cell-cell interaction might be important for development, however, the

complex structure of EBs may lead to induction and production of some cytokines and growth

factors which would affect differentiation and certain developmental programs. Moreover, earlier

studies of ESCs differentiation included fetal bovine serum (FBS). Poorly defined combinations

of factors in FBS plus lot to lot variations generally make the optimization of differentiation

process to be challenging and reproducibly difficult (Murry and Keller, 2008). These limitations

add another level of complications when it comes to interpretation of results. New approaches

have been applied to increase the control over the formation of EBs to make them more suitable

for development studies. For instance, protocols have been developed and promoted to apply

6

serum-free differentiation conditions in which the effect of each inducing factor could be studied

(Kanke et al. 2014). However, these protocols present new challenges and limitations on their

own.

Mouse ES cells which served as our model for the current study are relatively easy to maintain in

culture in comparison to human ES (hES) which make them a feasible model for developmental

studies (Reviewed in (Langenbach and Handschel, 2013)). There are advantages and

disadvantages of ES applications in clinical setting (Kim and Park, 2017). These are listed in the

table 1-1.

Induced pluripotent stem cells (iPS) are also considered as a potential source for osteogenic

differentiation. Integration methods which were applied for reprograming of somatic cells into

iPS mainly included viral vectors (e.g. retroviral and lentiviral) to transfer selected genes into the

host genome. Despite reported high efficiency of the techniques, these methods have raised

safety concerns for clinical applications of iPS (Csobonyeiova et al. 2017).

ESCs MSCs

Advantages Disadvantages Advantages Disadvantages

Pluripotent Immunogenic Easier to direct them

to osteogenic lineage

Trans-differentiation to

other cells

Unlimited

proliferated capacity

Increased risk of

teratoma

formation

Availability Proliferative capacity

declines by age

less likely to carry

mutations

Difficult to

differentiate

uniformly

Easy to isolate and

expand

More genetic

abnormalities due to

exposure to

environmental factors

Low cost Ethical/Political

issues

Free from ethical

issues

Highly heterogeneous

Table 1-1 Advantages and disadvantages of ESCs and MSCs for cell-based therapies

7

1.2.1 Embryonic Stem Cell Osteogenic Differentiation

To use stem cells for bone repair in clinical settings it is essential to develop well-defined

protocols that allow differentiation of stem cells into osteogenic lineage. This will be necessary

to reduce the likelihood of spontaneous differentiation of stem cells into multiple divergent

lineages other than osteogenic lineage and formation of teratoma at the transplantation site

(Mackenzie and Flake, 2001; Trounson, 2002). Moreover, well differentiated osteogenic lineage

increases the efficiency of engraftment and better integration within recipient bone tissues.

Besides human clinical therapy, the development of such protocols would be a great source for

studying osteogenesis and bone development. An added advantage is that such protocols would

provide in vitro models for pharmokinetic and cytotoxicity testing which could reduce the need

for animal models and limits challenges over ethical issues.

The process of ES cell osteogenic differentiation requires the activity of specific transcription

factors which are expressed and function at distinct time points during the differentiation

process, thereby defining various developmental stages of osteoblast lineage (Long F., 2012). In

addition to expression level of osteogenic transcription markers, the deposition of ECM and

mineralization can be assessed by von Kossa or Alizarin Red S staining (ARS) to evaluate

osteogenic differentiation progress. In von Kossa staining, sliver nitrate reacts with phosphate in

the osteo nodules in the presence of acidic material and results in development of yellow to

brownish yellow color. This staining does not necessarily represent the level of Ca2+ deposit.

For this reason, combined with von Kossa staining, I applied ARS to evaluate and quantify the

level of calcification in osteo nodules. In ARS, Ca2+ forms an ARS-calcium complex in a

chelation process and visually turn to a bright red stain. This method is extremely popular for

mineralization assessment as absorbed dye can be extracted from the stained cells and quantified

by spectrophotometry. This way the mineralization levels of osteo-nodules could be compared in

less subjective manner.

In vitro osteogenic differentiation of ES cells requires the addition of several factors mimicking

those released by in vivo osteoblast’s microenvironment. Studies have considered and evaluated

many factors and their effects on ES cell osteogenic differentiation (Heng et al. 2004). Critical

components in majority of these protocols include ascorbic acid (AA) and β-glycerophosphate

(β-Gly) which was first demonstrated to induce mineralization by Buttery et al. (Buttery et al.

8

2001). AA, acts as a cofactor for enzymes that hydroxylate proline and lysine in pro-collagen

and therefore, is essential for the secretion of collagen into the ECM. Hydroxyapatite-containing

mineral is then deposited on this collagenous matrix if a source of inorganic phosphate such as β-

Gly is provided. Inorganic phosphate (Pi) acts as an intracellular signaling molecule to regulate

the expression of many osteogenic genes, including OPN (Fatherazi et al. 2009). A derivative

of AA, L-ascorbic acid-2- phosphate, has replaced AA in traditional protocols as it showed

higher stability in cell cultures (Gessin et al. 1993).

Dexamethasone (Dex), the other critical factor, is a glucocorticoid drug that has been shown to

have stimulatory effects on osteogenic differentiation. This may be due to its effect on MSCs

survival and proliferation (Wang et al. 2012). Some studies suggest that Dex induction of

osteogenic differentiation is Runx2 dependent. It was reported that Runx2 expression was

upregulated via canonical Wnt pathway in response to Dex treatment (Hamidouche et al. 2008).

Another study indicated a role for Dex in MKP-1 upregulation which in turn dephosphorylate

Runx2 at serine 125 and enhances Runx2 transcriptional activity (Phillips et al. 2006). More

studies are needed to clarify how these components induce osteogenic differentiation for us to

improve their application in osteogenic differentiation protocols.

Our lab has optimized a 21-day osteogenic differentiation protocol in which variety of factors

concentrations and duration of treatment have been applied to enrich osteo-lineage cells from R1

mouse ES cells (Yu et al. 2015). Four components that have been applied in our protocol

include retinoic acid (RA), AA, β-Gly, and Dex. In our protocol, RA is added to the culture

during the growth phase (day 3 to 5) in which EBs are in suspension. It has been shown that RA

promotes osteogenesis at least partially by increasing Runx2 expression in MSCs (Dingwall et

al. 2011). A detailed scheme of the optimized protocol that has been used in the current study is

shown in figure 1-1.

9

Figure 1-1 Osteogenic differentiation protocol used in this study.

EBs were formed from 250 mouse ES cells in hanging drops and were collected and transferred

to ultra-low attachment dishes at day 3 of differentiation. EBs were maintained in the floating

condition and underwent treatment with RA at 0.1 µM until day 5. At day 5, EBs were

transferred to cell culture dishes where they can adhere to pre-gelatinated surface. AA and β-Gly

were added at day 6 and kept throughout differentiation at 50 µg/ml and 10 mM, respectively.

0.1 μM Dex was then added at day 10 and kept throughout differentiation.

1.3 Development of Osteoblast Phenotype

Bone development involves a series of different stages which initiates by proliferation of osteo-

progenitor cells and is followed by their differentiation into osteoblasts. Both proliferation and

differentiation are tightly regulated by expression of regulatory genes and transcription factors.

At proliferation stage when cells are mitotically active, expression of genes involved in cell cycle

and growth including c-myc, c-fos, and c-Src are increased. Disrupted expression of c-Src has

been shown to reduce osteoblast proliferation (Marzia et al. 2000). Initiation and transition of

and from one stage to another is controlled by stage-specific transcription factors and production

of ECM proteins. Following down regulation of proliferation, cells start to express genes that are

associated more specifically to bone phenotype. Some of the genes of ECM including COL1 are

10

expressed in the proliferation stage. Accumulation of COL1 will contribute to the cessation of

proliferation and its expression will then decrease to the basal level during subsequent stage of

differentiation. ECM eventually undergoes some changes which makes it ready for

mineralization. During mineralization stage expression of some genes including BSPII and OC

starts to be induced and gradually peaks to the maximal level. Among genes which are induced

and expressed at high level during mineralization OC is the one which its expression starts to

increase in more mature osteoblasts. OC expressing cells are found only within mineralizing

nodules. Major signaling pathways involved in osteogenic differentiation and bone formation are

briefly described below.

1.3.1 BMP/ TGF β pathway

BMP/TGF-β pathway is a major signaling cascade involved in many cellular processes that are

responsible for bone formation during mammalian development. BMPs and TGF-β are cytokines

that belong to the TGF-β superfamily. Upon binding of TGF-β or BMP ligands to their receptor

complex, which consists of heterotetrameric combinations of type I and II serine/threonine

kinase receptors, signals will be transmitted to the cytoplasm through canonical or non-canonical

pathways. In canonical pathways, different types of Smad proteins are involved in which usually

Smads 1 and 5 are activated by BMP extracellular signals, and Smads 2 and 3 by TGF-β

extracellular signals. Activated Smads will then make a complex with Smad4 and translocate to

the nucleus where they interact with DNA promoter region and participate in transcription as a

transcriptional coactivator. Noncanonical TGF-β signaling also participates in osteogenesis

which involves many signaling molecules mainly from mitogen-activated protein kinase family

(MAPKs). In context of bone regeneration, the P38/MAPK activation has shown to play a role in

the expression of ALP and OC in osteoblastic cells. This has been reviewed more extensively

elsewhere (Rahman et al. 2015).

1.3.2 Wnt pathway

Wnt pathway which is activated by binding of a large family of Wnt ligands to the membrane-

spanning frizzled (FZD) receptors contributes to osteoblast differentiation and mineralization.

Wnt canonical pathway mediates signaling through the stabilization of β catenin, whereas the

noncanonical pathways are β catenin-independent. Canonical Wnt pathways play a critical role

in bone formation through regulation of osteoblast-specific gene expression and has been better

11

characterized regarding its role and therapeutic potential in bone disorders when compared to

other signaling pathways. In the absence of Wnt signal, cytoplasmic β-catenin is phosphorylated

by glycogen synthase kinase 3β (GSK3β), ubiquitinated and degraded. The major non-canonical

Wnt pathway is Ca2+-dependent in which upon binding of Wnt ligand to the FZD receptors a

signal cascade releasing intracellular Ca2+ are activated. The released Ca2+ in turn activates

downstream mediators including calcineurin and Ca2+/ calmodulin-dependent protein kinase II

(CaMKII). These mediators will then activate essential transcription factors for osteogenic

differentiation such as nuclear factor of activated T cells (NFATs) (reviewed in (Kim et al.

2013)).

1.3.3 Ca2+ pathway

Ca2+ signaling plays an important role in osteoblast proliferation and differentiation process. Ca2+

influx is induced by hormones and growth factors like vitamin D3 or through integrin-mediated

cell adhesion. Upon an increase in intercellular release of Ca2+ diverse signaling pathways are

activated. Calmodulin (CaM) is a ubiquitous Ca2+ binding protein that counts as a major

mediator of Ca2+. Ca2+/ CaM complex can interact with protein kinases and phosphatases to

transduce the message to downstream targets. Among them CaM kinase II (CaMKII) and

calcineurin are particularly important in osteoblast differentiation. It has been shown that the

Ca2+–CaM/CaMKII pathway leads to an increase in the phosphorylation of extracellular signal-

related protein kinase (ERK). Elevated level of phosphorylated ERK subsequently leads to

Runx2 activation which promotes osteoblast differentiation (Park et al. 2010). Ca2+/CaM

complex could also bind to calcineurin and activates it. Active calcineurin in turn

dephosphorylate NFAT transcription factors which leads to NFAT nucleic translocation and

subsequent promotion of osteoblast differentiation. Ca2+ signaling pathways and their effect on

osteoblast differentiation is extensively reviewed elsewhere (Zayzafoon, 2006).

1.3.4 FGF pathway

Fibroblast Growth Factor (FGF) involves several secreted signaling proteins that signal to

receptor tyrosine kinases and intracellular proteins. Several FGF molecules have been identified

to play regulatory roles in osteogenesis including FGF-2 and FGF-9 (Miraoui et al. 2008; Zou et

al. 2015). Binding of FGF ligands to FGFRs leads to sequential transphosphorylation of at least

six tyrosine residues. Activation of the FGFR tyrosine kinase domain allows binding and

12

phosphorylation of other adaptor proteins, including phospholipase Cγ/PKCα and signal

transducer and activator of transcription 1 (Stat1), Stat 3, and Stat 5 (Ornitz and Marie, 2015).

Activation of PKC subsequently induce Runx2 transcriptional activity and hence promotes

osteogenic differentiation. Stat1 branch of FGF signaling pathway is involved in chondrogenesis

in which upon activation, Stat1 translocate into the nucleus and activates cell cycle inhibitor P21

expression. Increased levels of P21 correlates with decreased capacity for chondrogenic

differentiation (Masson et al. 2015). FGF signaling pathways could also regulate osteogenesis

by modulating transcription factors that contribute to cell proliferation, differentiation, and

survival in cells of the osteoblast lineage through activation of ERK MAPKs, and PI3K/AKT

pathways (reviewed in (Ornitz and Marie, 2015)).

1.3.5 Notch pathway

Notch pathway requires cell-cell contact and facilitates short-range signaling between

neighboring cells, coordinating spatial and temporal regulation of cell fate during embryonic

development of tissues and organisms. It plays regulatory roles both in osteoblasts and

osteoclasts. In osteoblasts, Notch signaling is mediated via canonical (BMP/ Smads) as well as

noncanonical TGF-β/ MAPKs pathways. Inhibition of osteoblast differentiation by Notch

through sequence of mechanisms have been reported. These include; 1) Antagonizing Wnt

signaling, thus leading to the decreased abundance and activity of β-catenin, in part by allowing

its degradation by GSK3β, 2) Inhibiting NFATc1 transcriptional activity, 3) Suppressing Runx2

transcriptional activity via binding of protein complexes to Runx2 formed in response to Notch

activation. There are studies suggesting that Notch signaling promote proliferation of

intermediate osteoblast lineage cells, but impedes formation of mature osteoblasts (reviewed in

(Regan and Long, 2013) and (Ernesto Canalis, 2008)).

1.4 Osteoblast Transcription Factors and ECM Proteins

Induction, proliferation, differentiation, and maturation of osteoblasts are tightly regulated by

transcription factors (Komori, 2006; Komori, 2011). Expression and activation of large and

growing number of transcription factors involved in the regulation of the osteoblast phenotype

have been identified. However, the complete description of the transcription factor networks and

bone matrix proteins that regulate osteogenesis is beyond the scope of this introduction, but some

13

of the key regulators and ECM proteins that have been used as markers of osteogenic

differentiation in this study are briefly introduced here.

Runx2 (runt related transcription factor 2) is the master regulator/transcription factor

required to direct multipotent mesenchymal cells to the process of osteoblast

differentiation and inhibits them from differentiating into the adipocytic and chondrocytic

lineages (Ducy et al. 1997). Runx2 regulates the expression of its target genes in

osteogenesis by binding to osteoblast-specific element 2 (OSE2) in their promoters.

OSE1 has also been identified and its activity may be required for OSE2 promoter

activity. Runx2 DNA-binding domain interacts with major bone matrix genes’ promoters

including the COL1A1, BSPII, and OC and induces their expression (Komori, 2005).

Years after discovery of Runx2, its transcriptional target genes are still being investigated

to further identify the role of Runx2 in different biological contexts including bone

development (Stock et al. 2004; Stephens and Morrison, 2014). Runx2 is one of the key

players in this study and is reviewed more extensively later in this chapter.

Osx (osterix) is a zinc finger transcription factor that is considered as a major

transcriptional regulator of osteoblastogenesis. Significance of Osx was discovered when

generated bone from intramembranous ossification of Osx-null mice were entirely non-

mineralized (Nakashima et al. 2002). Osx expression is believed to be downstream of

Runx2, majorly because of normal expression of Runx2 in Osx-null mice, while Osx

expression is absent in Runx2-knockout mice (Huang et al. 2007; Sinha and Zhou,

2013). Osx function has been shown to be modulated by post-translational modifications.

In one study, it was demonstrated that phosphorylation of Osx by c-Src increases its

stability and enhances its transcriptional activity (Choi et al. 2015). More recently, it was

found that Osx in differentiating bone regulates BMP- induced bone formation and limits

the level of signaling to control bone growth rate in skull (Kague et al. 2016).

COL1 (collagen type 1) is the most abundant structural protein in animals and the most

prevalent fibrous protein of ECM encoded by two distinct genes COL1a1 and COL1a2.

Mutations in COL1 lead to several forms of bone abnormalities. Osteoblasts highly

express both genes and generate collagen fibrils which their cross-linking contributes to

bone strength (Viguet-Carrin et al. 2006). Runx2 has been shown to up-regulate COL1

14

expression that is considered as an early indicator of osteoblastic differentiation (Ducy et

al. 1997).

BSP (Bone sialoprotein) is a major non-collagenous glycoprotein abundantly expressed

in mineralized tissues. The protein is characterized by its ability to bind to hydroxyapatite

through polyglutamic acid sequences and play a role in the early mineralization of

osteoblasts. Knockout studies of BSP-/- mice showed a significant reduction in bone

formation (Malaval et al. 2008) due to the reduced number of osteoprogenitors (Bouet et

al. 2015).

OC (osteocalcin) also known as bone gamma-carboxyglutamic acid-containing protein

(BGLAP) is highly abundant bone protein secreted by osteoblasts. OC is strongly

expressed in more mature osteoblasts and is used as a specific marker in the process of

osteogenic differentiation. Gla (gamma-carboxyglutamate) domain of OC functions in

binding to Ca2+ and hydroxyapatite. Blocking of α2-integrin-ECM interactions has been

shown to block ascorbic acid dependent OSE2 activation, indicating that integrin

interactions may play a key role in the regulation of the osteocalcin gene. In post-

translational modification, OC is carboxylated and released from osteoblasts and deposits

in the bone matrix. The carboxylated OC recruits osteoclasts, promotes their maturation

and inhibits bone formation (reviewed in (Li et al. 2016)).

1.5 Runx2

Runx2 is highly conservative with homology of structure as high as 99% among animals. Runx2,

frequently described as the master regulator of osteoblastogenesis, is the first osteoblast-specific

transcription factor that play an important role throughout the induction, proliferation, and

maturation of osteoblasts. It is still the earliest transcription factors that regulates expression of

many osteoblast genes through interaction with their OSE2. To date, OSE2 like elements have

been identified in some osteogenesis related genes including OC, COL1, BSP and, OPN (Ducy

et al. 1997). The significance of Runx2 was revealed by in vivo studies when Runx2-/- generated

mice completely failed to make bone (Komori et al. 1997; Otto et al. 1997). In another attempt

Runx2-/- calvarial cells failed to differentiate to osteoblasts (Kobayashi et al. 2000). However,

double negative Runx2 did not affect the expression of its target genes including COL1A1 and

OC in mature osteoblasts (Maruyama et al. 2007). This indicates that Runx2 necessity is limited

15

to early stages of osteoblast differentiation. The protein Runx2 is composed of several domains

including glutamine/alanine (QA) rich region and a proline/serine/threonine (PST) rich region,

DNA-binding domain, a nuclear-localization signal (NLS), a nuclear matrix targeting signal

(NMTS), and a C-terminal VWRPY domain for TLE/Groucho interactions. Many of the

signaling pathways and transcription factors that influence osteoblast differentiation enforce their

effect through Runx2 expression and/or Runx2 function.

1.5.1 Regulation of Runx2 Transcriptional Activity via Post-Translational Modifications

Runx2 plays an important role in different aspects of osteogenesis by stimulating osteoblast

differentiation, promoting chondrocyte hypertrophy, and contributing to endothelial cell

migration and vascular invasion of developing bones. In our view, Runx2 is considered a focal

point for integration of a variety of signals affecting osteoblast activity. Therefore, it is not

surprising that Runx2 transcriptional activity is tightly regulated by numerous transcriptional co-

activators and co-repressors. Runx2 transcriptional activity is often not well correlated with its

expression level suggesting that this factor must be activated through post-translational

modification. Runx2 transcriptional activity is mainly regulated through phosphorylation,

acetylation, and ubiquitination.

Phosphorylation

Phosphorylation of Runx2 is one of the major process in regulation of Runx2 activity. Reports up

to date have identified several serine residues on Runx2 with both stimulatory and inhibitory

effects upon their phosphorylation. For instance, phosphorylation of Runx2 on S369, S373, and

S377 by GSK3β were found to inactivate Runx2 whereas its PKA-dependent phosphorylation of

S347 or on S301 and S319 have been shown to enhance Runx2 transcriptional activity and

stimulate the expression of osteoblast specific genes (reviewed in (Vimalraj et al. 2015)).

Acetylation

Acetylation of lysine residue within histone proteins reduces their affinity to bind to DNA which

subsequently results in an increase of DNA accessibility to transcription factors (Sterner and

Berger, 2000). Interaction of Runx2 with histone acetylases (HATs) and histone deacetylases

(HDACs) have been reported to affect Runx2 transcriptional activity. Runx2 protein stability and

16

transcriptional activity are induced by Smad1 and 5 downstream of BMP signaling which

facilitates Runx2 interaction with P300, a protein processing HAT activity (Jeon et al. 2006).

Furthermore, HDAC3 was shown to interact with the amino terminus of Runx2 and repress

Runx2 transactivation of the OC promoter (Schroeder et al. 2004).

Ubiquitination

Several mechanisms for ubiquitin-dependent regulation of Runx2 have been discovered. WW

domains in members of E3 ligases bind to the proline-rich domain in Runx2 and facilitate the

protein-protein interactions. These interactions will subsequently result in Runx2 ubiquitination

and proteasomal degradation. It is important to highlight that other post-translational

modifications that have been discussed earlier could have regulatory effects on ubiquitination

process as well. For instance, phosphorylation of a specific residue may provide a recognition

sequence for a ubiquitin E3 ligase that can target Runx2 for proteasomal degradation whereas

acetylation, may block the availability of a lysine residue to a ubiquitin moiety, resulting in an

increase of the protein stability. It has been shown that the reduced expression and function of

Runx2 upon its phosphorylation by GSK3β result from Runx2 ubiquitin-mediated degradation

through E3 ubiquitin ligase Fbw7α (Kumar et al. 2015). Recently it has been shown that mono

ubiquitination of Runx2 by an E3 ligase activated downstream of BMP pathway transactivates

Runx2 and enhances osteogenesis (Zhu et al. 2017).

1.5.2 Regulation of Runx2 Transcriptional Activity via Runx2-Interacting Partners

Runx2 interactions with other proteins affect Runx2 subcellular localization and transcriptional

activity. Two major identified Runx2 interacting partners include Stat1, and Yes-Associated

protein (YAP) which are discussed briefly below.

YAP

One pathway in which Runx2 activity is regulated through its cellular spatial distribution was

found when researchers were investigating the inhibitory role of c-Src kinase on osteogenic

differentiation (Zaidi et al. 2004). They found that YAP regulates Runx2 sub nuclear trafficking

upon phosphorylation and activation by c-Src. However, nucleo-cytoplasmic distribution of YAP

was not affected by inhibition of c-Src activity either by its specific inhibitor PP2 or in Src DN.

17

Furthermore, OC expression in YAP DN was still considerably lower than those treated with

PP2 (5 µM). Collectively, the inhibitory role of c-Src in bone formation via Runx2 is only

partially explained by YAP activity suggesting that other mechanisms are involved. It is also

important to note that some of these findings are generated within non-osseous cells (e.g. Hela

cells) which further complicate the interpretation of the outcome. Therefore, further

investigations of c-Src inhibitory/regulatory role in Runx2 activity and osteogenic differentiation

in osseous cells or/and osteoprogenitors are needed.

Stat1

The signal transducer and activator of transcription 1 (Stat1) has two isoforms, Stat1α (91 kDa)

and the shorter truncated version Stat1β (84 kDa). Stat1 was originally identified as a signaling

molecule downstream of interferon (IFN) signaling pathways. Stat1 usually localizes as an

inactive form in cell cytoplasm. However, upon activation of the JAK-Stat pathway by

interferons, Stat1 binds to the receptor complex via its src homology 2 domain (SH2) and

phosphorylates at Y701 residue. Phosphorylation of Stat1 leads to its dimerization and nuclear

translocation where it induces the expression of its target gene. Phosphorylation of Stat1 on a

serine residue (S727) has shown to increase the transcriptional activity of Stat1. Within the two

isoforms of Stat1, Stat1β lacks the serine 727 phosphorylation site in the C-terminus. Multiple

mechanisms are involved in downregulation of Stat1 and its transcriptional activity.

Dephosphorylation of Stat1 by protein phosphatase Src homology phosphatase II (SHP2) has

been reported to negatively regulate Stat1 activity (Wu et al. 2002). Proteasomal degradation of

Stat1 dimers following ubiquitination is another mechanism by which Stat1 is negatively

regulated. It has been demonstrated that ERK activity phosphorylates Stat1 at S727 which

contributes to its ubiquitination and degradation (Surinder M. Soond, 2008). In another study,

it was demonstrated that OPN targets Stat1 and mediates its degradation through ubiquitin-

proteasome system (Gao et al. 2007).

Recently it was found that Stat1 participates not only in immune regulations but also in

osteoclastogenesis and osteoblast differentiation (Tajima et al. 2010). It has been shown that

number of osteoclasts with enhanced bone resorption activity due to IFN-activated ISGF3

(Interferon-stimulated gene factor 3) were increased in Stat1-/- mice. However, Stat1-/- mice

showed an increase in bone mass indicating that Stat1 has an inhibitory effect on bone formation.

18

Further investigation revealed that unlike Stat1-/- mice, those with interferon regulatory factor 9

(IRF9) deletion showed no abnormalities in osteoblast parameters indicating an IFN-independent

role for Stat1 in osteogenesis. No significant difference was observed in apoptosis between

Stat1-/- and WT mice suggesting Stat1 role specifically in osteoblast differentiation. Moreover,

Runx2- dependent promoter activity of OC was affected only in Stat1-/-. Inhibition of Runx2

transcriptional activity by Stat1 was showed to be independent from Stat1 phosphorylation status

on Y701 or S727. However, activation of Stat1 due to IFN-γ stimulation reduced its interaction

with Runx2. This may suggest that the signaling downstream of Stat1 activation results in loss of

Runx2-Stat1 association. Finally, it was found that Stat1 interacts with Runx2 in the cytoplasm

of osteoblasts and inhibits Runx2 from being localized in the nucleus (Kim et al. 2003).

Although sufficient supporting results for Stat1 anchoring Runx2 were provided, the regulation

of this interaction during osteogenic differentiation is unclear.

1.6 Regulation of Osteogenic Differentiation by c-Src

c-Src family of non-receptor kinases (SFKs) have several members including Src, Yes, Fyn, Lyn,

Fgr, Hck, Lck, Yrk, and Blk. Although many members of this family have restricted patterns of

expression particularly in cells of hematopoietic system, the Src, Yes, Fyn, and Yrk are

ubiquitously expressed in mammals. Fyn and Yes are both highly expressed in brain, fibroblasts,

endothelial cells, and T cells whereas c-Src is highly expressed in brain, platelets, and

osteoclasts. Targeted disruption of c-Src only resulted in bone phenotype which made c-Src an

interesting target in bone development studies (Espada and Martin-Perez, 2017). A marked

decrease in the rate of bone resorption in mice with homozygous mutations in c-Src revealed

regulatory roles for c-Src in osteoclasts (Soriano et al. 1991). Almost a decade later, another

group of researchers showed that the increased level of bone in Src knockout mice was not only

due to a decrease in bone resorption but also an increased in osteoblast differentiation. Increased

differentiation of osteoblasts isolated from Src KO mice with elevated levels of OC mRNA and

other osteoblast markers, suggests that c-Src have inhibitory effect in osteoblast maturation

(Marzia et al. 2000). The inhibitory role of c-Src in osteogenesis has been the subject of other

studies (Zaidi et al. 2004; Peruzzi et al. 2012), however role of c-Src in bone development and

ES osteoblast differentiation is still poorly understood.

19

1.6.1 c-Src Structure and Function

Discovery of SFKs stems from work on the Rous sarcoma virus, a chicken tumor virus

discovered in 1911 by Peyton Rous (Rous, 1911). v-Src (a viral protein) is encoded by the avian

cancer-causing oncogene of Rous sarcoma virus. In contrast, c-Src (the normal cellular

homologue) is encoded by a physiological gene, the first proto-oncogene to be described and

characterized (Stehelin et al. 1976). From the N- to C-terminus, c-Src contains a membrane

anchoring SH4 N-terminal which contains 14- carbon myristoyl group, a unique domain, an SH3

with high affinity for proline-rich sequences, an SH2 domain which specifically interacts with

tyrosine phosphorylated residues in Src target proteins, an SH2-kinase linker, a protein-tyrosine

kinase domain (SH1), and a C-terminal regulatory segment. One of the most important functions

of SH2 and SH3 domains is that they constrain the activity of the enzyme through intramolecular

interactions. In fact, c-Src activity is regulated by phosphorylation of two distinct tyrosine

residues. The inactive state of Src is obtained by phosphorylation of tyrosine near the C-terminus

of c-Src (Tyr530 in mammals and Tyr527 in chicken) which enables its intermolecular

interaction with SH2. This cause SH3 domain interaction with a poly-proline motif of the linker

sequence which restricts access to the kinase domain and stabilizes the inactive conformation.

Under basal condition in normal growing fibroblasts a major portion of c-Src is inactive (Zheng

et al. 2000). The inactive conformation of c-Src can be released by several mechanisms

including (1) dephosphorylation of Y527 by tyrosine phosphatases like SHP2 (Roskoski, 2005),

and (2) mutation of Y527 or deletion of its C-terminus sequence which could turn c-Src to the

constitutively active state, as happens in the oncogenic form of SFKs. Dephosphorylation of c-

Src at Y527 is followed by autophosphorylation of c-Src tyrosine residue Y416 which in turn

makes c-Src enzymatically active. Schemes of c-Src structures in both active and inactive states

are shown figure 1-2.

20

Figure 1-2 The domain structures of inactive and active c-Src

The Src kinase architecture consists of four domains: the unique domain, followed by the

SH3, SH2, and tyrosine kinase domains. The activation loop of the kinase domain and the

activating (Tyr 416) and autoinhibitory (Tyr 527) phosphorylation sites are indicated. In the

autoinhibited form of c-Src kinase, upon phosphorylation of Y527 the SH2 domain binds the

phosphorylated C-terminal tail, and the SH3 domain binds the linker segment between the

SH2 and the kinase domain. These events keep c-Src in a conformation that inhibits its

phosphorylation at Y416 and Src activity.

21

1.7 Hypothesis and Objectives

It has been reported that c-Src is expressed in mESCs (Meyn et al. 2005) and hES cell lines

(Zhang et al. 2014a). The maintenance of c-Src expression throughout differentiation in hES

suggests roles for c-Src role in differentiation compared to many other family members that

might play critical roles in self-renewal property of stem cells. However, the role of c-Src in

osteogenic differentiation of ESCs has not been studied. Therefore, in this study we decided to

investigate how c-Src activity affects osteogenic differentiation in mESCs. To do so, I

hypothesized that c-Src kinase plays an important role in mouse ESC osteogenic differentiation

through regulation of Runx2. Considering the major role of c-Src family kinases in ESC

differentiation, to my knowledge, currently there is no published study investigating the effect of

c-Src on Runx2 regulation in mouse ESC osteogenic differentiation. Therefore, the objectives of

my research are as follows;

1. To demonstrate that c-Src kinase activity affects mouse ESC osteogenic differentiation

2. To verify whether or not Runx2 mediates the effect of c-Src

3. To explore signaling pathways downstream of c-Src which may affect osteogenesis of mouse ESCs

22

Chapter 2

2 Materials and Methods

2.1

2.1.1 Cell Culture

Mouse ES cells R1 derived from J1 129/Sv mice (Nagy et al. 1993) were maintained in their

undifferentiated status in Dulbecco’s modification eagle’s medium 1X (DMEM, Catalog No.

319-005-ES) supplemented with 10% FBS (FBS, Premium, Catalog No. 088150), 1% MEM

non-essential amino acids (MEM NEAA 100X, Gibco, Catalog No. 11140050), 10 nM 2-

mercaptoethanol (Bioshop, Catalog No. MER 002), and LIF. Drops containing 250 cells per

25 µl DMEM supplemented with 20% FBS were placed on the lids of tissue culture dishes for 3

days. After 3 days EBs formed in hanging drops were transferred into the floating cell culture

dishes containing medium supplemented with 0.1 µM retinoic acid (Sigma, Catalog No. R2625)

for 2 days. On day 5, the EBs were plated in tissue culture dishes coated with 0.1% gelatin. On

day 6, the differentiation medium was supplemented with 50 ng/ml L-ascorbic acid (Sigma,

catalog No. A5960) and 10 mM β-glycerophosphate disodium salt hydrate (Sigma, catalog No.

G9422) to promote osteogenic differentiation. 100 nM dexamethasone (Sigma, Catalog No.

D4902) was added on day 10 to further enrich cells of the osteogenic lineage. The medium was

changed every 2 days for the entire 21 days of differentiation.

MC3T3-E1 subclone 4 (ATCC® CRL2593™) were purchased from ATCC. Cells were

maintained in a-MEM with ribonucleotides, deoxyribonucleosides, 2 mM L-glutamine and 1

mM sodium pyruvate, but without ascorbic acid (Gibco, Custom Product, Catalog No.

A1049001) supplemented with 10%(v/v) FBS. For osteogenic differentiation, the cells were

cultured in a-MEM supplemented with 10% FBS, 50 mg/ml ascorbic acid, and 10 mM β-

glycerophosphate disodium salt hydrate for 21 days. Medium was renewed every 3 days.

23

2.1.2 Inhibition assay by pharmacological inhibitors

c-Src inhibitor PP2 (Calbiochem, Catalog No. 529573) and negative control PP3 (Calbiochem,

Catalog No. 529574) were applied where indicated. For protein half-life assay, Cycloheximide

(Sigma-Aldrich, Catalog No.C7698) was applied at 20 µM concentration for the indicated times.

Proteasome inhibitor MG132 was purchased from Millipore (Catalog No. 474790).

2.1.3 Immunoprecipitation assay

Cell extracts were harvested in a Pierce IP Lysis Buffer® (25 mM Tris-HCl pH 7.4, 150 mM

NaCl, 1% NP-40, 1 mM EDTA, 5% glycerol). 500 ug cell extracts were incubated with 1 ug

anti-RUNX2 (M-70, Santa Cruz, Catalog No.sc-10758), or Stat1 (Cell signaling, Catalog

No.9172S) at 4º C overnight. Immune complexes were recovered with True Blot® anti-rabbit Ig

beads (Rockland, Catalog No.00-8800-25) and subjected to SDS-PAGE.

2.1.4 Real-time PCR analysis

The Qiagen RNeasy Mini Kit (Qiagen, Catalog No. 74134) was used to extract total RNA

according to the manufacturer’s instructions. 500 ng RNA was reverse transcribed to cDNA

using iScript cDNA Synthesis Kit (Bio-Rad, Catalog No. 1708890) in a total reaction volume of

20 µl. To examine the mRNA expression of osteogenic markers, cDNA was amplified using

real-time qPCR. Real-time qPCR analysis was performed in Bio-Rad’s CFX384 Touch™

detection system. The cDNA levels were normalized against L32 gene. The primers sequences

used in this study are listed in table A-1.

2.1.5 Chromatin Immunoprecipitation

R1 differentiating cells were treated for 24 hours with 10 uM PP2 or PP3. The protein-DNA

complexes were cross-linked with 1% formaldehyde (10 min, RT), scraped, washed in PBS, and

harvested. Cells were then resuspended in ChIP lysis buffer (50 mM Tris-HCl pH 8, 150 mM

NaCl, 1mM EDTA, 1% Triton X-100, 0.1 % Na-deoxycholate containing protease inhibitor

cocktail (Sigma-Aldrich, Catalog No. P8340) and 1 mM DTT. Cells were sonicated 10 times for

15 seconds each time to shear DNA to less than 1 kb (verified by agarose gel analysis). The

soluble chromatin was collected by centrifugation and pre-cleared with protein-DNA- blocked

agarose beads for 1h. An aliquot of the pre-cleared chromatin was put aside as an input and the

supernatant were incubated overnight with 1 ug of Runx2 (M-70, Santa Cruz, Catalog No.sc-

24

10758), ChIP grade Rabbit IgG, polyclonal isotype control (Abcam, Catalog No. ab171870), and

RNA polymerase II (Abcam, Catalog No. ab5131) at 4º C. Protein-DNA- blocked agarose beads

were then added and incubate for 2 hours. After immunoprecipitation, beads were washed for 10

minutes once in 1mL of low-salt immune complex wash buffer (20 mM Tris-HCl [pH 8.0], 150

mM NaCl, 2 mM EDTA, 1% Triton X-100, 0.1% SDS), once in 1 ml of high-salt immune

complex wash buffer (20 mM Tris-HCl [pH 8.0], 500 mM NaCl, 2 mM EDTA, 1% Triton X-

100, 0.1% SDS), once in 1 ml of LiCl buffer (20 mM Tris-HCl [pH 8.0], 250 mM LiCl, 1 mM

EDTA, 1% NP-40, 1% Na-deoxycholate), and twice with 1 ml of TE (10 mM Tris-HCl [pH 8.0],

1 mM EDTA) in the order that was explained. Protein-DNA complexes were eluted in 100 ul of

fresh ChIP elution buffer (1% SDS, 0.1M NaCO3) for 1hr at 37°C in two settings of 50 µl of

elution buffer for 30 minutes. Cross-linked eluted complexes were then reversed by overnight

incubation using NaCl with the final concentration of 0.2 M at 65°C. The recovered DNA was

then treated with 1 ug of RNase A for 30 minutes at 37 ºC and 10 µg of Proteinase K for 2 hours

at 45 ºC. DNA was purified using a PCR purification kit (QIAGEN, Catalog No. 28104) and

eluted in 100 ul of DNase-RNase free water. ChIP DNA was amplified by qPCR using COL1A1,

OC, BSPII, and COL2A primer pairs. A list of ChIP primers is provided in table A-2.

2.1.6 Immunoblot analysis

Cells were collected in CST lysis buffer (Catalog No. 9803) containing 20 mM Tris-HCl [pH

7.5], 150 mM NaCl, 1 mM Na2EDTA, 1 mM EGTA, 1% Triton, 2.5 mM sodium

pyrophosphate, 1 mM β-glycerophosphate, 1 mM Na3 VO4, and 1 µg/ml leupeptin. The

Bradford method was used to quantify the concentrations of protein samples. 30 µg protein

sample were separated by SDS-PAGE and transferred to nitrocellulose membrane. For

immunoblotting, I used anti-Stat1 (Cell Signaling, catalog No.9172S), anti -p-Stat1-Y701 (Cell

Signaling, Catalog No.9171), anti-GAPDH (Cell Signaling, Catalog No.2118S), anti-c-Src (Cell

Signaling, Catalog No. 2108), anti-p-c-Src-Y416 (Cell Signaling, Catalog No.2101S), anti-p-c-

Src-Y527 (Cell Signaling, Catalog No.2105), anti- Runx2 (Cell Signaling, Catalog No.8486S),

anti-Ubiquitin (Biolegend, Catalog No.646301). The secondary antibody was goat polyclonal

secondary antibody to rabbit IgG - H&L (HRP) (Abcam, Catalog No. ab6721) or rabbit

polyclonal secondary antibody to mouse (HRP) (Abcam, Catalog No. ab6728). The secondary

antibody for immunoblot analysis of IP products was Rabbit True Blot® anti Rabbit IgG (HRP)

(Rockland, Catalog No.18-8816-31).

25

2.1.7 Mineral deposition assays

Differentiated day 21 nodules were examined for the presence of mineral deposits using Alizarin

Red S (ARS, sodium alizarin sulphonate) staining. 4 mM Alizarin Red S (Sigma, Catalog No.

A5533) was prepared in distilled water and the pH was adjusted to 4.0 using 10% ammonium

hydroxide. Cultures were fixed with 4% formalin for 15 minutes, triple washed with dH2O, and

stained with Alizarin Red S for 20 minutes. After removal of unincorporated excess dye with

distilled water, the mineralized nodules were stained as bright orange- red spots. Absorbed stains

by mineral nodules were then dissolved in 10% acetic acid and their absorbance of triplicates of

each sample were read at 405 nm with a plate reader and the concentration of ARS were

calculated using the equation of the trend line of ARS standard concentrations. For Von Kossa

staining, differentiated day 21 nodules were rinsed three times with PBS, fixed in 10% neutral

formalin buffer for 2 hours, and then washed 3 times with distilled water. Nodules were then

stained with 2.5% silver nitrate solution for 30 minutes. Finally, they were washed 3 times with

distilled water and were examined for mineralization.

2.1.8 Immunofluorescence (IF) staining

Nodules on coverslips were rinsed 3 times with PBS. They were then fixed in 3.7%

formaldehyde for 15 minutes, washed three times for 5 minutes each time in PBS, and

permeabilized with 0.1% Triton X-100 in buffered containing 100 mM 1,4

piperazinediethanesulfonic acid, 1 mM EGTA, and 4% (wt/vol) polyethylene glycol 8000 (pH

6.9) for 2 minutes. Nodules were then washed three times in PBS for 5 minutes and incubated

with 1% BSA, 22.52 mg/mL glycine in PBST (0.1% Tween 20) for 30 minutes to block

unspecific binding of the antibodies. After removal of blocking buffer nodules were incubated

with 1:1000 anti-Runx2 (Abcam, Catalog No. ab76956) at 4 ºC overnight. Next day nodules

were washed in PBS (3 times for 5 min) and incubated with the secondary antibody fluorescein

(FITC)-conjugated donkey anti-mouse (Jackson ImmunoResearch, Catalog No. 715-096-151) for

1hour at room temperature. Nodules were then washed 3 times in PBS for 10 minutes and

mounted in Pro® Gold Antifade Mountant (Thermo Fisher Scientific, Catalog No. P36931).

Imaging was performed using confocal LSM800 microscope.

26

2.1.9 Subcellular Fractionation

Nuclear and cytoplasmic fractions were prepared using the Nuclear Extraction Kit (Millipore,

Catalog No. 2900) as per manufacturer’s instruction. Briefly, cells were washed with PBS twice

and the pellet were resuspended in the cytoplasmic lysis buffer. After 15 minutes incubation on

ice, cells were centrifuged and the pellet were resuspended in fresh cytoplasmic lysis buffer

using a small gauge needle (27 gauge). Disrupted cell suspension was centrifuged at 8000 x g for

20 minutes. The supernatant was then collected as the cytoplasmic fraction. The remaining pellet

was then resuspended in the nuclear lysis buffer and centrifuged (16000 x g for 5 minutes). The

supernatant of this part was collected as the nuclear fraction.

2.1.10 Transfection

c-Src siRNAs (Catalog No. S238007, S238008, and S238009), Stat1 siRNA (Thermo Fisher

Scientific, Catalog No. AM16708), GAPDH siRNA as a positive control (Catalog No. 4390850),

and negative control siRNA (Ambion® Silencer® Negative Control #1, Catalog No. 4390843)

were purchased from Thermo Fisher Scientific. MC3T3 E1 cells were transfected with diluted

siRNA and lipofectamine RNAiMax reagent (Thermo Fisher Scientific, Catalog No. 13778030)

according to the manufacturer instructions. For ESCs transfection, Neon™ transfection system

was used following the recommended protocol for mESCs transfection. c-Src RNAi oligo

sequences are listed in table A-3.

2.1.11 Lentiviral virus production and transduction

Lenti-X 293T cell line (Clontech, Catalog No. 632180) was used for lentivirus production. The

cells were plated at 70% confluency the day before transfection on 100 mm culture dishes. 7 μg

lentiviral vectors containing constitutively active mouse c-Src (LV[Exp]-Tet3G:T2A:Puro

TRE3G>mSrc [NM_009271.3]*del:3xGGGGS:EGFP, VectorBuilder), or m-Cherry vector were

diluted in 600 µl of nuclease free sterile water. Dilute DNA was then added to one tube of Lenti-

X™ Packaging Single Shots Ecotropic, (TaKaRa, Catalog No. 631278) and was vortexed at high

speed for 20 seconds to dissolve the pellet completely. The mix was then incubated at room

temperature for 10 minutes and subjected to a quick spin for 2 seconds. Finally, the nanoparticle

complexes were added to Lenti-X 293T cells in 5 mls of serum-free, antibiotic-free DMEM

(Sigma, Catalog No. D5671) and incubated at 37°C supplied with 5% CO2. 24 hours post-

transfection 5 ml fresh serum-free, antibiotic-free media was added to the culture. 48 hours after

27

transfection supernatant containing viral particles was harvested and filtered through 0.45 µm

filters Millipore (Catalog No. SLHA033SS) and aliquots were stored at -80°C. Before being

used, the virus was rapidly thawed in a 37ºC water bath.

For transduction, media containing 10% Tet System Approved FBS (Takara, Catalog No.

631106) was prepared. 10 µl of Ecotropic Receptor Booster (TaKaRa, Catalog No. 631471)

along with the media containing 4 µg/ml polybrene (PB= hexadimethrine bromide; Sigma,

Catalog No. H-9268) was added to 60% confluent 6 well-plate cultures of MC3T3-E1s. Plates

were then centrifuged at 1200g for 20 minutes and incubated at 37°C for 2 hours. Media was

replaced with 2 ml complete PB containing media for all wells and viral containing supernatant

at MOI=10 was added to each well. After 16 hours, the virus-containing PB media was replaced

with 2 ml complete media. Next day, cells were treated with 2.5 µg/ml puromycin (Sigma,

Catalog No. P-8833) to select transduced cells for 5 to 7 days.

For doxycycline-induced protein over-expression, the transduced cells were incubated in 10%

FBS DMEM containing 2 µg/mL doxycycline (Sigma, Catalog No. D-9891). The medium was

refreshed every 24 hours for 3 days before collecting cell lysates for western blot analysis.

2.2 Statistical analysis

Statistical differences between two groups were analyzed by two-tailed Student’s t test. P values

of 0.05 or less were considered statistically significant. Multiple group statistical analysis was

performed using one-way ANOVA where indicated in which a summary of analysis is presented

as F and p values.

28

Chapter 3

3 Results

3.1 c-Src Activity Inhibits Osteogenic Differentiation in mESCs

3.1.1 Osteogenic Differentiation in Mouse Embryonic Stem Cells

To characterize the role of c-Src in ES cell osteogenesis we differentiated EBs under osteogenic

culture conditions described in the Methods. To monitor osteogenic differentiation, the

expression level of RNAs encoding Sox2, Oct3/4, and Nanog as markers of pluripotency along

with Runx2, Osx, COL1A1, BSPII, and OC as markers of osteogenesis were analyzed. To do

this, cells were harvested every day throughout the 21-day differentiation timeline. Extracted

total RNAs were subjected to cDNA synthesis and real time qPCR was performed. qPCR values

were normalized against corresponding L32 as housekeeping gene. Results showed that RNA

expression of pluripotency markers decreases to undetectable level around day 5 (Figure A-1)

which is around the time that Runx2 and Osx expression were started to show an increase. As for

COL1A1 and BSPII, higher level of RNA expression was detectable on day 9 and 12,

respectively. In terms of OC RNA expression, as the most specific osteogenic marker, a real

increase was recorded on day 18 which was maintained up to day 21 (Figure 3-1). Primer pairs

for qPCR analysis are listed in table A-1.

To examine mineralization, day 21 nodules were subjected to both Von Kossa and Alizarin Red

staining (ARS). For von Kossa staining, nodules were fixed with formalin solution and then

washed prior to the staining with silver nitrate. The interaction of the silver ions with phosphate

in the mineralized regions results in the formation of silver phosphate which was yellow to

brownish yellow and nonmineralized region remained colorless. As for ARS, nodules were fixed

and stained with Alizarin Red (PH 5.2). The reaction enables the formation of ARS- Ca2+ which

appeared optically red. Excess of dye was then washed away with water before imaging the

nodules. Two colonies from the same plate representing the nodules with low and high level of

mineralization are selected for both von Kossa and ARS methods and are shown in figure 3-2.

29

Figure 3-1 mRNA expression of osteogenic markers during 21 days of osteogenic

differentiation in mESCS

Gene expression of Runx2, Osx, BSPII, COL1A1, and OC of ESCs (day 0) and differentiating

cells (day1 to 21) which were induced for osteogenic differentiation were analyzed by real time

qPCR using the primer pairs listed in table A-1. Data shown represent the mean (±SD) of

triplicates. Among all Runx2 is the first gene showing an increase at day 5 of differentiation. OC

which is the marker of more mature osteoblast showed an induction more stably at the later

stages of differentiation starting at day 18. Results shown are representative experiments of three

independent assays.

Rel

ativ

e m

RN

A E

xpre

ssio

n

Days of osteogenic differentiation

30

Figure 3-2 Alizarin Red Staining (A) and Von Kossa staining (B) of day 21 osteo-nodules

Mineralization of osteo-nodules at the end of differentiation is assessed by ARS and von Kossa

staining. Two differentiated colonies per each staining are shown side by side to show lower

(image on the left) and higher (image on the right) level of mineralization in each type of

staining. (A) In ARS Ca2+ deposit reaction with Alizarin Red S were stained red. The colony on

the left showed less mineralization compared to the one on the right. (B) von Kossa staining

resulted in the formation of yellow to yellowish brown stain due to the accumulation of silver

phosphate in the mineralized area.

A

B

31

3.1.2 c-Src Expression Profile in Mouse Embryonic Stem Cells During Osteogenic Differentiation

Previous reports have suggested a role for c-Src in the regulation of osteoblast differentiation

(Marzia et al. 2000). However, the dynamic of c-Src activity during osteogenic differentiation of

ES cells was not studied. Therefore, I first examined the protein level for c-Src along with p-

Y416-c-Src as the representative of active state of c-Src every day during differentiation starting

at day 3 using immunoblotting analysis. 250 ES cells were seeded in the absence of LIF to

permit EB formation following hanging-drops method. Forming EBs were treated under

osteogenic culture for 21 days and were harvested for the indicated days starting at day 3 of

differentiation. Lysates were prepared and subjected to WB analysis. As shown in figure 3-3A,

results revealed that c-Src protein level does not undergo any major change in developing EBs.

In terms of c-Src activity, Y416 phospho-specific c-Src antibody detected variable level of active

c-Src indicating that c-Src activity varies during osteogenic differentiation of mESCs. More

specifically, p-c-Src-Y416 showed a decreased level in early stages of differentiation and was

slightly increased at the later stage starting at day 14 (Figure 3-3B). Note that results showed a

remarkable decrease in RNA expression of Oct3/4, Nanog, and SOX2 in differentiating mESCs

to almost undetectable level by day 6, consistent with the loss of pluripotent ES cells from the

culture. Collectively, these results provide further evidence for dual role for c-Src kinase in

signaling pathways that regulate ES renewal in earlier days of differentiation and potential role in

osteogenic differentiation in later days possibly from day 6. Clear downregulation of c-Src in

early stage of osteogenesis may suggest an inhibitory role for c-Src in early phase of the

differentiation.

32

A

3.1.3 Inhibition of c-Src at Different Periods During ES Cells Osteogenic Differentiation

The complex changes in c-Src activity observed during osteogenic differentiation suggested a

regulatory role for c-Src in osteogenic differentiation of ESCs. To test this idea, I used small-

molecule inhibitors for suppressing c-Src activity to determine whether suppression of c-Src

activity would affect ES cell osteogenic differentiation. I first compared multiple concentrations

of three distinct SFK inhibitors (SrcI-1, PP1, and PP2) on day 5 differentiating EBs for their

effects on inhibition of c-Src activity. Day 5 was chosen as it showed high abundance of p-Y416-

c-Src compared to many other days during osteogenic differentiation. One of the inhibitor

Figure 3-3 c-Src activity during 21 days of osteogenic

differentiation in mESCs

Cell lysates of differentiating cells were collected and subjected

to WB analysis for the indicated days to assess c-Src activity

based on its phosphorylation level at Y416. GAPDH served as

a loading control. Immunoblots showed an initial decrease of

activity at the earlier stage of differentiation starting at day 6,

and an increase at the later days starting at day 14 (A). p-c-Src-

Y416 band densities were quantified using ImageJ and were

normalized to corresponding total c-Src and then normalized to

corresponding GAPDH controls. Graphed values are shown

(B). The graph shows a representative experiment from three

independent assays.

B

33

examined was SrcI-1, a potent, competitive dual site (both the ATP- and peptide-binding) Src

kinase inhibitor. Src Inhibitor-1 is one of the "gold standards" for Src kinase inhibition with

IC50 = 44 nM for c-Src. The other one was PP1, which has been shown to have c-Src inhibitory

effect at IC50 = 170 nM. Finally, PP2 a potent, reversible, ATP-competitive, and selective

inhibitor of the Src family of protein tyrosine kinases capable of inhibiting c-Src at IC50 = 100

nM was included in the assay. To determine the effect of the tested inhibitors on overall c-Src

activity in vivo, day 5 differentiating cells were first treated with two different concentrations of

each inhibitor including 1 and 10 µM for 2 and 24 hours. Lysates were then collected and

subjected to immunoblot analysis. Judging by the levels of p-Y416-c-Src detected by p-Y416

phospho specific Src antibody, almost complete suppression of c-Src was achieved when cells

were treated with 10 µM of PP2 with no significant effect on cell growth. However, only partial

suppression of c-Src activity with the same or lower concentrations of SrcI-1 and PP1 was

possible without significant cytotoxicity effect (Appendix, A-2A, and A-2B). A wider range of

PP2 concentrations were then applied to further confirm the optimal amount of PP2 for

inhibitory assays. Therefore, day 5 differentiating cells were treated with this compound at 100

nM, 1 µM, 5 µM, 10 µM, or 20 µM followed by immunoblotting of whole-cell lysates with the

pY416 phospho-specific antibody. DMSO as the solvent control and PP3 as the inactive analog

of PP2 served as controls. Immunoblots are shown in figure A-2C. To facilitate visual

comparison of the discussed conditions band densities were quantified using ImageJ, measured

values of p-Y416-c-Src for each condition were normalized against corresponding DMSO and

graphed. Results clearly showed that PP2 treatment reduced p-Y416-c-Src immunoreactivity in a

dose-dependent fashion, with nearly complete inhibition at 10 µM whereas no remarkable

reduction of c-Src activity was detectable in the DMSO and PP3 control conditions.

To evaluate the impact of c-Src on mESC osteogenic differentiation, I performed c-Src inhibition

using PP2 (10 µM) for eight different time periods during osteogenic differentiation. Although

unlimited number of conditions were possible to do such assays, but I limited the numbers to

cover ups and downs of Runx2 expression as the earliest indicator of osteogenic differentiation.

A scheme shown in figure 3-4A summarized the inhibitory assays that have been conducted. PP3

(the inactive analog of PP2), and DMSO (solvent) served as controls for the indicated time

periods. At day 21, cells were collected and subjected to qPCR analysis to examine the effect of

c-Src activity for each time period on RNA expression level of OC which served as the specific

34

marker to evaluate osteoblast differentiation. OC qPCR values were normalized to corresponding

L32 for each condition. OC relative expression of PP2 treated cells were then normalized to their

corresponding DMSO condition and calculated values were graphed to show the fold change in

OC expression for every single period of treatment. One- way ANOVA analysis was then

performed to compare the OC expression among all tested conditions. Results showed inhibition

of c-Src activity during day 6 to 10 led to a statistically significant increase in OC mRNA

abundance (Figure 3-4B).

To assess the effect of c-Src inhibition on mineralization, day 21 osteo-nodules were stained with

both ARS and von Kossa methods. A marked increase in Ca2+ deposits evaluated by Alizarin

Red staining was observed in osteo-nodules in which c-Src activity was inhibited between day 6

to 10. Absorbed dye was dissolved in acetic acid and absorbance was read at 450 nm by

spectrophotometer. The concentration of Alizarin Red was then quantified, normalized to their

corresponding DMSO condition and their fold change was graphed as shown in figure 3-4C.

Statistical analysis by ANOVA resulted in a significant difference of Alizarin Red concentration

for day 6-10 PP2 treated cells when compared to every other PP2 condition. As for von Kossa

staining, more colonies of day 6-10 PP2 treated cells stained darker indicating a higher level of

mineralization when compared to other conditions. It is important to note that inhibition of c-Src

between days 3 to 5 of differentiation resulted in a significant reduction in final OC mRNA

expression and the overall mineralization which may suggest a regulatory role for c-Src in the

early stages of ESCs differentiation which is consistent with earlier studies investigating the role

of c-Src in early development of EBs.

35

Figure 3-4 Inhibition of c-Src activity between day 6-10 enhances mES osteogenesis.

c-Src activity was inhibited using PP2 (10 µM) for eight different time periods during osteogenic

differentiation of mESCs. PP3 (inactive analog of PP2) and DMSO served as controls. (B) Total

RNA of day 21 osteo-nodules was extracted and subjected to qPCR analysis using OC primer

pairs. Fold change in OC expression for each condition was calculated by normalizing PP2

treated values to their corresponding DMSO condition. Values representing the mean (±SD) of

triplicates were graphed and subjected to one-way ANOVA. (C) Day 21 osteo-nodules were

stained by Alizarin Red staining (ARS) method. Absorbed stains were extracted and the

concentration of Alizarin Red S was quantified using an equation of the standard trend line.

The graphs show pooled data from three independent experiments. Values were then normalized

to their corresponding DMSO control and graphed. Data shown represent the mean (±SD) of

triplicates. P values were calculated using one-way ANOVA (****p<0.0001)

A

B C

36

Next, I asked whether or not the effect of c-Src inhibition on level of OC RNA expression is

dose dependent. To answer this question, I used different concentrations of PP2 including 1, 5,

and 10 µM along with DMSO as the negative control to treat cells between day 6 to 10 of

osteogenic differentiation. Lysates were collected on day 21, the last day of our osteogenic

differentiation protocol, and subjected to qPCR analysis. Values for each condition of PP2

treatment was normalized against its corresponding DMSO condition to calculate fold change in

OC RNA level and graphed as shown in figure 3-5A. One-way ANOVA analysis was conducted

which confirmed a statistical significant difference in OC RNA level of cells treated with 10 µM

of PP2 when compared to 1 and 5µM concentrations. To assess whether or not the observed

phenotype is specific to osteogenic marker, COL2A expression level was also evaluated in

response to different concentrations of PP2. Values were quantified and graphed as described.

Statistical analysis using one-way ANOVA revealed no significant changes in COL2A RNA

level among different concentrations of PP2. Data are shown in figure 3-5B. Moreover,

mineralization of osteo-nodules was assessed by ARS staining which revealed significantly

higher ARS absorbance by colonies treated with higher concentrations of PP2 (Figure 3-5C).

37

A B

C

Figure 3-5 c-Src inhibitory effect on OC expression and

mineralization is dose dependent.

Three different concentrations of PP2 including 1, 5, and 10 µM

were applied between days 6 to 10 of osteogenic differentiation

in mESCs along with DMSO as negative control. (A) Lysates

were collected at the final day of differentiation (day 21) and OC

mRNA expression in response to different dosages of PP2 were

measured by real time qPCR and normalized to corresponding

DMSO controls. (B) Day 21 osteo-nodules were stained by ARS

method and absorbed stain was quantified, normalized to

corresponding DMSO condition and graphed as fold change.

The graphs show representative experiments from three

independent assays. Data shown represent the mean (±SD) of

triplicates. *p< 0.05, ***p<0.001, and ****p<0.0001.

38

A potential complication of studies using PP2, however, is nonspecific inhibition of other

protein-tyrosine kinases (Bain et al. 2003; Bain et al. 2007). To ensure that the phenotype

observed with PP2 was not a result of inhibition of other kinases, I attempted to transfect self-

renewing mESCs and differentiating cells with specific c-Src siRNAs using Neon electroporator

following the manufacturer instructions. Briefly, 107 cells were suspended in the transfecting

buffer subjected to electroporation using 10 nM, 100 nM, 200 nM of c-Src specific siRNA.

Three pulses at 1400 V with pulse width of 20ms were applied. GAPDH specific siRNA served

as a positive control at the same indicated concentrations along with the testing siRNA. The

commercial negative siRNA control served as the negative control to monitor any undesired

biological effect of the applied siRNAs. Further modifications of the protocol were applied

including using higher concentrations of siRNAs and higher voltages but no significant change

was observed. As shown in figure A-3A to A-3C no reduction in c-Src expression was observed

following the conducted transfections.

To overcome the aforementioned technical inefficiency of the experimented system, I was

advised to use MC3T3-E1 cells as an alternative model to study the specificity of c-Src effect on

osteogenic differentiation. MC3T3-E1 cells are osteoblast precursor cell line derived from mouse

calvarias. A series of subclones isolated from the cloned but phenotypically heterogeneous

MC3T3-E1 cell line exist. MC3T3-E1 subclone 4 has been reported to exhibit high levels of

osteoblast differentiation after growth in ascorbic acid and inorganic phosphate compared to

many other clones of MC3T3-E1s (Wang et al. 1999). To evaluate the osteoblastic

differentiation of MC3T3-E1s subclone 4 were cultured in AA containing media accompanied

with β-glycerophosphate as explained in the material and method for 21 days and cells were

collected for qPCR analysis every other day during the differentiation. mRNA measurements for

BSPII, Runx2, Osx, COL1A, and OC are shown in figure 3-6. Efficiency of transfection was

evaluated by WB analysis in which cells were transfected with 100 nM of c-Src specific siRNA

along with GAPDH as a positive control and a negative control siRNA. 48 hours post

transfection lysates were collected and subjected to WB analysis. As shown in figure 3-7A, c-Src

siRNA application resulted in a remarkable reduction in protein level of c-Src detected by

specific c-Src. To evaluate the specificity of c-Src inhibition and subsequently its effect on

osteogenic differentiation MC3T3-E1 cells were transfected with c-Src specific siRNA along

with Silencer™ Select negative control twice every second day of osteogenic differentiation.

39

Lysates were collected and subjected to qPCR analysis at day 14 and 21 of differentiation. Figure

3-7B shows fold change in mRNA expression of osteogenic markers including BSPII, COL1A,

and OC when normalized to their corresponding negative control which were increased in

response to c-Src depletion. Mineralization assessment of differentiating cells by ARS both at

day 14 and 21 showed a significant increase in comparison to their corresponding controls

(Figure 3-7C). This strongly suggests that c-Src activity at least in part is responsible for the

observed effects by PP2 treatment.

Days of osteogenic differentiation

Rel

ativ

e m

RN

A E

xpre

ssio

n

Figure 3-6 Gene expression of osteogenic

markers in MC3T3-E1

MC3T3-E1 cells underwent osteogenic

differentiation protocol for 21 days. Cells

were collected at the indicated days during

differentiation and lysates were prepared for

RNA extraction and qPCR analysis.

Expression of osteogenic markers including Runx2, Osx, BSPII, COL1A1, and OC were quantified

and normalized to their corresponding L32 housekeeping gene. Average of triplicate values (±SD)

were graphed. The graphs show a representative experiment from three independent assays.

40

B C

Figure 3-7 c-Src downregulation in MC3T3-E1 increases osteogenic differentiation

(A) MC3T3-E1 cells were transfected with c-Src specific siRNA along with GAPDH and

Negative select siRNAs as positive and negative control, respectively. Lysates were collected

and subjected to WB analysis using c-Src and GAPDH antibodies. (B) MC3T3-E1 cells were

transfected with specific c-Src or Negative select siRNAs twice every two days starting at day 0

of osteogenic differentiation. Lysates were collected at day 14 and day 21 of osteogenic

differentiation and extracted RNAs were subjected to qPCR analysis using BSPII, COL1A1, and

OC primer pairs. Values in triplicated for c-Src transfected cells were normalized against their

negative corresponding control and average values (±SD) were graphed as fold change. (C) Day

14 and day 21 differentiating MC3T3-E1 cells were subjected to ARS. Extracted stain was

quantified and normalized to the corresponding negative controls. Mean values of triplicates

A

41

(±SD) are shown as fold change in the graph. Graphs show a representative experiment from

three independent assays.

3.1.4 Overexpression of p-Y416-c-Src in MC3T3-E1s and its Effect on Osteogenic Differentiation

Mutational studies have clearly elucidated a closed, inactive conformation and an open, active

state as such when the C-terminal tyrosine at Tyr 527 is phosphorylated, c-Src is inactive; and

when dephosphorylated, it is active, with the potential for auto-phosphorylation at Tyr 416. So,

dephosphorylation of mouse c-Src at Tyr 527 can bring about activation, even when protein

levels are normal. To further assess the consequences of c-Src activity on osteogenic

differentiation I employed lentiviral vector over expressing mouse c-Src carrying deletion at Tyr

527. To create an optimal model system that will enable us to study the role of c-Src in

osteogenic differentiation, we pursued to obtain the tet-inducible lentiviral system. One major

benefit of employing an inducible system would be the ability to investigate the effect of the

protein expression and its activity at different time points during differentiation, as well as being

able to show reversibility and the specificity of an observation associated with the protein that is

being studied. In terms of the expression system, the Tet-On 3G Tetracycline-Inducible Gene

Expression Systems were applied which are the 3rd generation of the most powerful inducible

mammalian expression systems available which are widely in use in functional studies (Das et

al. 2016). It is important to note that the Tet-On 3G system offers a significant improvement

over Tet-On with significantly reduced basal expression and increased sensitivity to doxycycline,

a tetracycline analogue. The lentiviral constructs expressing a GFP control or GFP tagged mouse

c-Src del*Y527 were designed and obtained from Vectorbuilder. For the generation of the stable

cell lines to be used in our experiments, lentiviral packaging system Lenti-X Packaging Single

Shots (Ecotropic) were used along with the vector containing constitutively active c-Src or m-

Cherry vector to transfect Lenti-X 293T Cell Line. This lentiviral packaging systems that

produce high-titer lentivirus pseudotyped with the mCAT-1 envelope glycoprotein allow us to

limit transduction to mouse and rat cells. 7 µg of each vector was diluted in 600 µl of nuclease

free H2O and added to one vial of Packaging Single Shot containing the required packaging

proteins. The mix was then vortexed briefly and incubated at room temperature for 10 minutes.

After the incubation, the mix was added to a 10 cm 80% confluent Lenti-X 293T cells. The

image of cells 48 hours post-transfection along with control condition are shown in figure 3-8A.

42

Supernatant of the transfected cells containing viral particles were collected and subjected to

titration assays. A day before transduction MC3T3-E1 cells were seeded at the confluency of

80% in a 6 well plate. Next day, media was supplemented with 10% Tet System Approved Fetal

Bovine Serum (tetracycline free) and 4 µg/ml polybrene. To increase the efficiency of

transduction, 15 µl of Ecotropic Receptor Booster was added to the wells and the plate was

centrifuged at 1200g for 20 minutes. These receptors consist of exome-like particles coated with

mCAT-1 receptor. When applied to the target cells, fusion of the exome-like particles with host

cell plasma membrane temporarily increases surface mCAT-1 receptor, thus allowing for

efficient viral transduction. Cells were then incubated at 37°C for 2 hours prior to transduction.

MOI=10 of the prepared viral stock was used to transduce MC3T3-E1 cells. 48 hours post

transduction media was replaced and cells were incubated at 37°C for another 24 hours.

Puromycin resistant cells were selected by applying 2.5 µg/ml puromycin for 7 days. Cells were

then maintained and expanded in media containing 1µg/ml puromycin. To examine the effect of

active c-Src overexpression, these cells were exposed to AA containing media to be induced for

the osteoblastic differentiation and were exposed to 1 µg/mL doxycycline to induce the

expression of constitutively active c-Src for 14 days before being collected for any further

analysis. Initial WB analysis indicated that Dox induction of cells transduced with lentivirals

over expressing constitutively active c-Src had significant increase in p-c-Src-Y416 amount

when compared to those with no Dox treatment and more significantly to those with empty

vector (Figure 3-8B).

Next, qPCR analysis was performed to investigate the effect of c-Src activity in OC mRNA

expression at day 14 of osteogenic differentiation. Results showed a significant decrease in OC

mRNA expression (p<0.05) in the induced cells overexpressing p-c-Src- Y416 compared to the

controls (Figure 3-8C). To evaluate the effect of c-Src hyperactivity on mineralization, day 14

cells were stained following Alizarin Red staining protocol. Absorbed stain in each well were

extracted and concentrations were quantified. As shown in figure 3-8D, induced over expression

of p-c-Src- Y416 during osteogenic differentiation resulted in significant decrease of Ca2+

deposits. These results further confirmed the inhibitory effect of c-Src activity on osteogenic

differentiation in pre-osteoblasts MC3T3-E1s.

43

Figure 3-8 Overexpression of constitutively active c-Src reduces osteogenic differentiation.

(A) Lenti-X 293T cell line were co-transfected with EGFP linked lentiviral vector expressing

constitutively active c-Src along with ecotropic Lenti-X Pachaging Single Shot packaging

plasmid for 4 hours. Lenti-X 293T cells co-transfected with m-Cherry served as negative

controls. Cells were imaged 48 hours post-transfection. (B) Transfected cells for the indicated

conditions were lysed 48 hours after transfection and the lysates were subjected to WB analysis

using Y416 phospho-specific c-Src, and c-Src antibodies. GAPDH served as loading control.

MC3T3-E1 cells were transduced with MOI=10 of viral containing supernatants for

constitutively active c-Src over expression. M-Cherry viral particles were used to transduce

MC3T3-E1s as negative control. Expression of exogenous active c-Src was induced with 2

A B

p-Y416-c-Src

C D

0.0

0.5

1.0

1.5

2.0OC

Dox - + - +

m-Cherry p-Y416-c-Src

*

**

44

µg/ml of doxycycline for 14 days. (C) Lysates were collected and the extracted RNAs were

subjected to qPCR analysis for OC mRNA expression. Values from doxycycline- free condition

were used to normalize the induced condition. Data representing mean of triplicate values (±SD)

were graphed. (D) Day 14 differentiated MC3T3-E1 cells were subjected to ARS for

mineralization assessment. Extracted Alizarin Red S for the indicated conditions were collected

and their concentrations were calculated. Mean of triplicated values (±SD) are graphed. Graphs

are representative experiments from three independent assays.

Student t-test was performed for statistical analysis where * and **** indicates p<0.05 and

p<0.0001.

45

3.2 c-Src Activity Inhibits Runx2 Nuclear Localization

We showed earlier that inhibition of c-Src between day 6 to 10 significantly increases osteogenic

differentiation in ESCs evidenced by a significant increase in OC mRNA expression and level of

mineralization of day 21 osteo-nodules. We also confirmed the specificity of c-Src inhibition

using specific c-Src siRNAs in MC3T3-E1 cells which also resulted in an increase of the OC

mRNA level and overall mineralization level assess by ARS. Furthermore, using lenitiviral

system we showed that overexpression of constitutively active c-Src inhibits osteogenic

differentiation in MC3T3-E1 cells. Next, we addressed the questions of how c-Src activity may

inhibits osteogenic differentiation. More importantly, we were interested to know the

significance of c-Src in osteogenic differentiation between day 6 to 10. As earlier

discussed, Runx2 initiates osteogenesis in a manner that is precisely controlled temporally and

spatially, and loss of Runx2 expression at this early stage impairs osteogenic differentiation in

bone development. For this reason, Runx2 became an interesting target of regulatory role of c-

Src in osteogenic differentiation for further investigation.

3.2.1 The Effect of c-Src Activity on Runx2 Target Genes’ Expression

It has been shown by others that Runx2 is involved in the transcriptional activation of many

promoters including those of COL1a1, BSPII, and OC (Ducy et al. 1997; Kern et al. 2001).

Considering Runx2 significance in transcription regulation of osteogenic markers, I tested Runx2

target genes expression including BSPII, COL1A1, and OC in response to inhibition of c-Src

activity. To do so, differentiating mESCs were either treated with PP2, the specific inhibitor of c-

Src, or PP3 (the inactive analog of PP2), or DMSO (control solvent) as negative controls for 24

hours. Lysates were then collected and subjected to qPCR analysis using specific primers listed in

table A-1. Results showed that inhibition of c-Src activity by PP2 significantly increased mRNA

expression of BSPII and COL1A by almost 1.5-fold (p<0.01, and p<0.001, respectively) and of

OC by more than 2 folds (p<0.01) when compared to the DMSO condition (Figure 3-9A). Results

of PP2 treated cells were significantly different from PP3 condition, however there were no

significant difference between DMSO and PP3 conditions. Next, I investigated to see whether or

not this significant change of the mRNA expressions is due to an increased transcriptional activity

of Runx2 at the promoters of the tested Runx2 target genes. Therefore, treated cells with PP2, PP3,

or DMSO were subjected to ChIP analysis. Anti- Runx2 antibody was used to immunoprecipitate

46

Runx2 protein binding to the extracted chromatin, and anti- POL II and anti-IgG served as a

positive and a negative control in the ChIP assay, respectively. The efficiency of the antibodies

was examined in a separate IP assay following WB analysis (data not shown). PCR primers

targeting COL1A, OC, and BSPII promoters are listed in table A-2. COL2 primer served as a

negative control as its promoter activity is not directly regulated by Runx2. Results of the

conducted ChIP assay revealed that when c-Src activity is inhibited under PP2 treatment condition,

the promoter occupancy of the COL1A, BSPII, and OC by Runx2 are significantly increased by

3.9, 9.3, and 3.4, respectively when compared to the DMSO control condition (Figure 3-9B). No

significant changes were observed in the IgG level among the conditions for the tested genes.

Furthermore, Runx2 occupancy of COL2 promoter was below detection limit which confirmed the

specificity of the observed phenomenon for Runx2 target genes.

3.2.2 The Effect of c-Src Activity on Runx2 Expression

Next, I asked whether or not the increased level of Runx2 transcriptional activity is due to an

elevated level of Runx2 expression. Therefore, first to test whether c-Src would regulate Runx2

expression between day 6 to 10 of osteogenic differentiation in ES cells, I treated the

differentiating cells either with PP2 (c-Src specific inhibitor) or PP3 (the inactive analog of PP2)

and DMSO (control solvent) as controls between days 6 to 10. Next, I collected cells at day 5, 7,

9, and 11 for all the mentioned conditions and subjected the lysates to qPCR and WB analysis.

Inhibition of c-Src activity in mESCs did not affect Runx2 mRNA and protein level (Figures 3-

10A and 3-10B). To further confirm that the effects of c-Src inhibition on Runx2 were specific c-

Src, I used two different c-Src specific siRNAs to reduce c-Src production in MC3T3-E1 cells

following the protocol as described in the material and method section. I performed twice

transfections on MC3T3-E1 cells with c-Src specific siRNAs every second day during

osteogenic differentiation for 5 days. SilencerTM select negative control siRNA served as control.

Then, I collected the lysates and subjected them to western blotting. Results showed both

siRNAs successfully downregulated the expression of c-Src and subsequently p-Y416-c-Src.

However, lower c-Src activity due to the decreased level of c-Src phosphorylation at Y416 did

not result in any significant change in Runx2 protein level when compared to the control

condition (Figure 3-10C).

47

A B

Figure 3-9 Inhibition of c-Src activity increases Runx2 transcriptional activity in mESCs.

(A) Day 10 differentiating cells from mESCs were treated with PP2 (10 µM), or PP3 (10 µM),

or DMSO for 2 hours. Lysates were collected and their total RNA were extracted. mRNA

expression of Runx2 target genes osteogenic markers including BSPII, COL1A1, and OC were

quantified by qPCR analysis. Values for PP3 and PP2 treated cells were normalized to their

corresponding DMSO condition. Mean values of triplicates were graphed for the indicated

conditions. One-way ANOVA was conducted for each primer pairs and p values were calculated.

(B) Lysates from described assay were also subjected to ChIP analysis using Runx2 antibody.

Precipitated immunocomplexes were analyzed by qPCR using BSPII, COL1A1, and OC primer

pairs. Values for PP3 and PP2 treated cells were normalized to their corresponding DMSO

condition. Mean values of triplicates were graphed and subjected to one-way ANOVA for

statistical analysis. Graphs represent pooled data from three independent assays.

*, **, ***, and **** indicate p<0.05, p<0.01, p<0.001, and p<0.0001, respectively.

48

A B

C

Figure 3-10 Inhibition of c-Src does not affect Runx2

expression in mESCs and MC3T3-E1s.

c-Src activity was inhibited using PP2 (10 µM) in

differentiating mESCs during day to 10 of osteogenesis. PP3

(10 µM) and DMSO served as negative controls. Lysates were

collected at the indicated times. (A) RNA was extracted for

qPCR analysis using Runx2 primer pairs. All values were

normalized to the corresponding L32 housekeeping gene and

fold changes were calculated by normalizing the quantified

values against their DMSO corresponding condition and mean

values of triplicates (±SD) were graphed.

One-way ANOVA was performed for each day data set where calculated p values showed no

statistical significant difference among tested conditions. The graph shows pooled data from

three independent experiments. (B) Lysates from day 5, 7, and 10 were subjected to WB analysis

using Runx2 antibody. P-Y416-c-Src antibody was applied to assess c-Src activity among

indicated conditions. (C) MC3T3-E1 cells were transfected with two distinct specific c-Src

siRNA along with Silencer ™ select negative control. Lysates were collected 48 hours post-

transfection and subjected to WB analysis using the indicated antibodies.

49

3.2.3 c-Src Activity and its Effect on Runx2 Subcellular Localization

Since a change in Runx2 mRNA and protein level was not observed following c-Src inhibition

neither in ESCs nor in MC3T3-E1s. However, reported from other studies levels of Runx2 are

often not well correlated with its transcriptional activity (Franceschi et al. 2003). Therefore, I

decided to examine whether or not c-Src inhibition would have any effect on Runx2 subcellular

localization. To do so I employed immunofluorescence (IF) staining using anti-Runx2 antibody

in mESCs treated with PP2 (10 µM), or PP3 (10 µM), or DMSO for two hours. IF staining

showed that distribution of Runx2 is more restricted to the nucleus of differentiating mESCs

when c-Src activity was inhibited with PP2 in comparison to PP3 or DMSO control conditions

(Figure 3-11A). To confirm these results, I used a biochemical fractionation to separate the

cytoplasmic and nuclear protein fractions from the treated mESCs with PP2, PP3 and DMSO to

examine the sub-cellular localization of Runx2. Nuclear fractions were then subjected to WB

analysis using anti-Runx2 antibody. Histone 3 (H3) served as the loading control for nuclear

fractions. Anti-GAPDH antibody was applied to assess the level of contamination with

cytoplasmic fractions. Consistent with the IF results, Runx2 remained mostly cytoplasmic under

both control conditions (DMSO and PP3), whereas it was re-localized to the nucleus upon PP2

treatment (Figure 3-11B). To further investigate the specificity of c-Src contribution into nuclear

localization of Runx2, I used c-Src specific siRNAs to down regulate the expression and activity

of c-Src in MC3T3-E1 cells. SilencerTM select negative control siRNA was used to transfect the

cells to further test the specificity of the assay. After 48 hours of transfection, nuclear fractions

were extracted and subjected to WB analysis using anti-Runx2 antibodies. Results showed that

upon downregulation of c-Src, nuclear localization of Runx2 was increased in comparison to the

control condition (figure 3-11C).

50

Figure 3-11 Runx2 nuclear localization is increased

when c-Src activity is inhibited.

(A) Day 10 ES differentiating cells were treated with PP2

(10 µM), or PP3 (10 µM), or DMSO for 2 hours, fixed and

stained with Runx2 antibody. DAPI used as a marker for

nuclei staining. Runx2 nuclear localization was increased

upon inhibition of c-Src activity by PP2. (B) Day 10 ES

differentiating cells were treated with PP2 (10 µM), or PP3

(10 µM), or DMSO for 2 hours. Lysates were collected and

fractions were isolated. Extracted nuclear fractions were by WB using Runx2 antibody. H3 and GAPDH served as loading controls of nuclear and cytoplasmic

fractions, respectively. (C) MC3T3-E1 cells were either treated with PP2 (10 µM) or DMSO for two

hours or subjected to transfection with c-Src specific siRNA or silencer™ negative select. Lysates of all

for conditions were fractionated and analyzed by immunoblotting using Runx2 antibody.

Results shown are representative experiments from three independent assays.

10µm

51

3.3.4 Inhibition of c-Src activity lowers Runx2 and Stat1 interaction

One major mechanism that affects protein localization is through interaction with other protein(s)

that stably reside in a cellular compartment. Given the earlier observation regarding the

inhibitory effect of c-Src on Runx2 nuclear localization, I hypothesized that c-Src has regulatory

role on Runx2-Stat1 interaction. To examine my hypothesis, first I tested the interaction of

Runx2 and Stat1 when c-Src activity was inhibited with a co-IP assay. In this assay, day 10

differentiating EBs were treated with PP2 for 2 hours. PP3 and DMSO were served as controls.

Lysates were then prepared and subjected to an IP analysis using Runx2 antibody. Precipitated

complexes were then subjected to WB analysis with both Runx2 and Stat1 antibodies. As shown

in figure 3-12A, upon inhibition of c-Src activity with PP2, Runx2 interaction with Stat1 is

significantly reduced compared to the control conditions. To further examine the specificity of c-

Src inhibition and effect on Runx2-Stat1 interaction, I used specific c-Src siRNAs to inhibit c-

Src expression and activity this time in MC3T3-E1 cells. 48 hours post-transfection MC3T3-E1

cells were then collected and subjected to the co-IP assay in which immune complexes were

immunoprecipitated this time with Stat1 antibody to further confirm the specificity of the IP

assay. Precipitated immunocomplexes were subjected to WB analysis using both anti-Runx2 and

anti-Stat1 antibodies. Results are presented in the figure 3-12B which showed that although Stat1

antibody precipitated comparable level of Stat1 in both control and c-Src down-regulated

MC3T3-E1 cells, however Runx2 interacting with Stat1 was remarkably lower in cells with

downregulation of c-Src.

52

Figure 3-12 c-Src inhibition lowers Runx2-Stat1 interaction.

(A) c-Src activity was inhibited in day 10 of osteogenic differentiation in mESCs using PP2 (10

µM) for 2 hours. PP3 (10 µM) and DMSO served as controls. Lysates were collected and

subjected to IP assay using Runx2 antibody. Precipitated immunocomplexes were then analyzed

by WB using both Runx2 and Stat1 antibodies. (B) c-Src expression was down regulated in

MC3T3-E1 cells. Silencer™ select negative siRNA served as negative control. 48 hours post-

transfection cells were lysed and IP assay was performed using Stat1 antibody.

Immunocomplexes were subjected to WB analysis using Stat1 and Runx2 antibodies.

Results shown are representative experiments from three independent assays.

A B

53

3.3 c-Src Regulates Osteogenic Differentiation Through Stat1

3.3.1 c-Src effect on Runx2 localization and transcriptional activity during osteogenic differentiation is significantly reduced in the absence of Stat1

To test whether or not c-Src exerts its effect on osteogenic differentiation at least in part through

regulation of Stat1-Runx2 interaction, I decided to examine whether or not inhibition of c-Src

would still be effective on enhancement of osteogenic differentiation when Stat1 is depleted in

MC3T3-E1 cells. Therefore MC3T3-E1 cells were transfected by Stat1 specific siRNA or to

transiently downregulate Stat1. SilencerTM select negative control siRNA served as control. 48

hours post-transfection, transfected cells were either treated with DMSO or PP2 for 2 hours.

Cells were then collected and subjected to fractionation assay to examine the effect of c-Src

inhibition by PP2 on subcellular localization of Runx2 in absence and presence of Stat1. Nuclear

fractions were further analyzed by WB using Runx2 antibody. GAPDH and H3 served as

cytoplasmic and nuclear markers, respectively. Results showed that downregulation of Stat1

resulted in an increase in the level of nuclear Runx2 almost comparable to the control condition

upon inhibition in c-Src activity by PP2. Also, the results indicated that in the absence of Stat1,

the inhibition of c-Src activity was not as effective in terms of increase in Runx2 nuclear

localization (Figure 3-13A). To further examine the effect of c-Src activity in the absence and

presence of Stat1 on Runx2 transcriptional activity and osteogenic differentiation, I decided to

evaluate the expression of OC as the specific markers of osteoblasts and Runx2 target by the end

of a 14-day differentiation assay. Therefore, MC3T3-E1 cells were transfected with Stat1 every

other day for 3 times with either Stat1 specific siRNA or SilencerTM select negative control

siRNA. Transfected cells were treated with PP2 (10 µM) or DMSO for 6 days starting from day

3 (48 hours post-transfection). On day 14, the final day of osteogenic differentiation, total RNA

from differentiated MC3T3-E1 cells for the experimented conditions were extracted and

subjected to qPCR analysis to test the expression level of OC. Analyzed data were normalized to

the average value of control condition in which cells were transfected with negative control

siRNA and were treated with DMSO. Graphed data are shown in figure 3-13B. Data showed

that inhibition of c-Src activity with PP2 resulted in a statistically significant increase of OC

mRNA (p<0.01). Also, downregulation of Stat1 resulted in a significant higher expression of OC

mRNA with (p<0.01) or without (p<0.05) inhibition of c-Src activity by PP2. However, the

54

difference between OC mRNA level of -/+ PP2 treatment of transfected MC3T3-E1 cells with

specific Stat1 siRNA was not significant (p=0.2). The results of this experiment collectively

suggest that the regulatory effect of c-Src on Runx2 subcellular localization is through Stat1

protein as an interacting partner of Runx2.

10.0

0.5

1.0

1.5

2.0

2.5

OC

Fol

d ch

ange

Stat siRNA - - + + PP2 - + - +

**

***

Figure 3-13 c-Src inhibitory role on osteogenic differentiation is Stat1-dependent.

(A) MC3T3-E1 cells were transfected with Stat1 specific siRNA along with Silencer™ select

negative control siRNA. 48 hours post transfection cells were treated either with PP2 (10 µM) or

DMSO for 2 hours. Cells were lysed and subjected to fractionation assay. Nuclear fractions were

analyzed by WB analysis using Stat1 and Runx2 antibodies. H3 and GAPDH served as loading

controls for nuclear and cytoplasmic fractions. (B) MC3T3-E1 cells were subjected to

transfection three times every 48 hours. Transfected cells were either treated with PP2 (10 µM)

or DMSO for 6 days. Lysates were collected at day 14 of osteogenic differentiation and total

RNAs were extracted. RNAs were analyzed by qPCR using OC primer pairs. OC mRNA

expression was quantified and triplicate values for each condition were normalized to average of

triplicate values from DMSO treated negative siRNA transfected cells and graphed. One-way

ANOVA was conducted and p values were calculated. *, and ** indicate p<0.5 and p<0.1.

Results shown are representative experiment from three independent assays.

Stat1

55

3.3.2 c-Src Activity and its Effect on Stat1 Subcellular Localization

Our findings so far suggested that c-Src could have a regulatory role on Stat1 such that it

facilitates Stat1 interaction with Runx2 or prevents it. One possible way would be that c-Src

affects the phosphorylation of Stat1 which could in turn affect Stat1 interaction with other

molecules including Runx2. It is well established knowledge now that Stat1 undergoes

dimerization upon phosphorylation of Tyr 701 and translocate to the nucleus. Therefore, if c-Src

affects Stat1 phosphorylation at Y701, it could alter Stat1 sub cellular localization which

potentially affects Stat1 interaction with Runx2 in cytoplasm. Therefore, I asked whether or not

Stat1 phosphorylation status of Y701 is affected by c-Src activity. To address this question, I

conducted an experiment in which day 10 differentiating mESCs were treated with PP2 (10µM),

or PP3 (10 µM), or DMSO for 2 hours. Treated cells were then collected and subjected to WB

analysis using Y701 phospho-specific Stat1, Stat1, and GAPDH antibodies. Strikingly, inhibition

of c-Src by PP2 resulted in Stat1 phosphorylation at Y701 suggesting that c-Src activity

indirectly regulates Stat1 phosphorylation (Figure 3-14A).

To assess the subcellular localization of Stat1 in response to c-Src inhibition and Y701

phosphorylation, I evaluated Stat1 protein level in the nuclear fractions of the treated cells.

Extracted fractions were then subjected to WB analysis using Stat1 antibodies along with H3 and

GAPDH as loading controls. Shown in figure 3-14B, an increase in Stat1 was observed in

nuclear fractions of differentiating cells in which c-Src activity was inhibited by PP2 compared

to PP3 or DMSO treatments. I then extended my observation performing the same treatments to

evaluate Stat1 nuclear localization via IF analysis. Therefore, cells were treated with PP2

(10µM) for 2 hours. Cells were then fixed and subjected to immunostaining using anti Stat1

antibody. DAPI was used to stain nuclei. Results further confirmed that Stat1 nuclear

localization is increased when c-Src activity is inhibited by PP2 compared to PP3 (inactive

analogue of PP2) and DMSO (Figure 3-14C).

To better understand whether or not phosphorylation of Stat1 at Y701 might be critical in Stat1

interaction with Runx2 and/or osteogenic differentiation of mESCs, I examined Stat1 Y701

phosphorylation status in earlier days of during osteogenic differentiation. Lysates of

differentiating cells were collected every day starting from day 3 to day 12 and subjected to WB

analysis using Y701 phospho-specific Stat1 and Stat1 antibodies. GAPDH served as the loading

56

control. Interestingly, WB analysis revealed that when c-Src activity is higher, at day 3 and 5 for

instance, the level of Stat1 Y701 is barely detectable which further supports the notion that

inhibition of c-Src activity could increase Stat1 phosphorylation at Y701 and its nuclear

localization (Figure A-4).

Whole Cell Lysates

p-Y701-Stat1

Stat1

p-Y416-c-Src

c-Src

Figure 3-14 Stat1 phosphorylation at Y701

and its nuclear localization is increased in

response to c-Src inhibition in mESCs.

(A) c-Src activity was inhibited by PP2 (10 µM)

in day 10 differentiating mESCs for 2 hours.

PP3 (10 µM) and DMSO served as controls.

Lysates were prepared and subjected to WB

analysis using anti-Y701 phospho-specific Stat1

along with Stat1 and GAPDH antibodies. (B)

Nuclear fractions of cells with described

treatments were extracted and followed by WB

analysis using Stat1 antibody. (C) subcellular

localization of Stat1 in day 10 differentiating

mESCs were further assess by IF using anti

Stat1 antibody. DAPI was used to stain nuclei.

Results shown are representative experiments

from three independent assays.

10µm

A B

C

57

3.3.3 The Effect of c-Src Activity on Stat1 Expression and its Half-life

Kim et al. showed that mutant form of Stat1, Stat1 CYF, inhibited Runx2 transcriptional activity

almost as efficient as Stat1 WT, which suggest that interaction of Stat1 and Runx2 is

independent of Stat1 phosphorylation status at Y701(Kim et al. 2003). Therefore, other

regulatory mechanisms may exist which does not involve tyrosine phosphorylation modifications

of Stat1. This rationally led me to further explore Stat1 modifications in response to c-Src

activity. I first sought to assess whether c-Src activity would regulate Stat1 expression. I

examined the RNA expression of Stat1 in cells in response to inhibition of c-Src activity by

pharmacological inhibitors including PP2 and SrcI1 in MC3T3-E1 cells. After 24 hours of

treatment with the indicated inhibitors and DMSO as the solvent control cells from all three

conditions were collected and subjected qPCR analysis. Values of three independent assays were

normalized to their corresponding controls, pooled and graphed. As shown in figure 3-15A, there

were no statistical difference in Stat1 RNA expression level among cells with different level of

c-Src activity when compared and analyzed by one-way ANOVA. To examine the specificity of

c-Src inhibition, I applied two different c-Src specific siRNA to downregulate c-Src expression

and activity along with Silencer™ Select Negative Control as a negative control. 48 hours post-

transfection cells were collected, RNA was isolated and cDNA was synthesized. qPCR analysis

was performed using specific Stat1 primer pairs and values were normalized against their

corresponding controls. Graphed data is shown in figure 3-15B. Assessed by one-way ANOVA

there were no significant difference among the level of RNA among the explained conditions.

Next question I asked was whether or not c-Src activity would affect Stat1 protein level. To do

so, differentiating mESCs were treated with PP2 along with the PP3 control for 24 hours.

Lysates were collected after 4, 8, 12, and 24 hours of treatment and subjected to WB analysis

using Stat1 antibody. Results showed a remarkable decrease in Stat1 protein level after 24 hours

of treatment of cells with PP2. However, Stat1 level was almost unchanged throughout the assay

for the tested time points when they were treated with PP3 (Figure 3-15C). These results

suggested that inhibition of c-Src activity lowers Stat1 protein level.

The homeostasis of protein metabolism is maintained and regulated by the rates of its

biosynthesis and degradation in living system. The most common approach to determine

turnover or half-life of a protein in cultured cells is to measure the degradation of the protein

58

after blocking its biosynthesis. Cycloheximide (CHX), a protein synthesis inhibitor that acts

specifically on the 60S subunit of eukaryotic ribosome (Baliga BS, 2017), is widely used for this

purpose. Therefore, I tested the half-life of Stat1 in the presence and absence of c-Src activity

when Stat1 biosynthesis was inhibited by cycloheximide (10 µM). For this experiment, treated

differentiating EBs from mESCs either with 10 µM of PP2 or PP3 for 2 hours prior to

biosynthesis inhibition by CHX (20µM). Cells treated with CHX for 4, 8, 12, and 24 hours were

lysed. Lysates were subjected to WB analysis using Stat1 antibody. GAPDH served as loading

control. WB analysis revealed decreased level of Stat1 protein in response to biosynthesis

inhibition at the first collection time, 4 hours after treatment, when c-Src activity was inhibited

by PP2. However, in PP3 treated cells Stat1 is still detectable at the level comparable to CHX

free condition after 24 hours of CHX treatment (Figure 3-15C).

I also experimented the effect of c-Src inhibition on Stat1 protein level every 15 minutes for the

duration of 75 minutes in MC3T3-E1 cells with and without CHX. DMSO served as control in

this assay. Immunoblots are shown in figure 3-15D. Band densities of Stat1 for both conditions

were quantified and to the corresponding GAPDH using ImagJ. Normalized values were graphed

and half-life of Stat1 were estimated by applying linear regression method on log scale of

normalized band densities for each condition. Results showed that pharmacological inhibition of

c-Src activities significantly shortened the half-life of endogenous Stat1 in MC3T3-E1 by almost

5 times supporting that c-Src regulates Stat1 mainly through enhancing protein stability (Figure

3-15E). These results collectively suggested an important role for c-Src in regulation of Stat1

stability.

59

A B

C

D

E

60

Figure 3-15 Stat1 protein stability is icreased when c-Src activity is inhibited.

c-Src activity was inhibited in MC3T3-E1 cells using either (A) Src pharmacological inhibitors

including PP1, SrcI-1, and PP2 for 2 hours where DMSO served as negative control and (B)

three distinct c-Src specific siRNAs were down regulated with three different distinct siRNAs

where scilencer ™ select negative siRNA served as control for 48 hours. Lysates from both

experiments were collected and subjected to qPCR analysis using Stat1 primer pairs. Triplicate

values were normalized to the average values of control conditions and graphed. (C) mESCs

were treated with 10 µM of PP2 or PP3 as control for 24 hours. PP2 and PP3 treated cells were

then treated with CHX (10 µM) or DMSO and lysates were collected at the indicated times and

subjected to WB analysis using Stat1 antibody. GAPDH served as loading control. (D) MC3T3-

E1 cells were treated with PP2 or DMSO. CHX was applied at (10 µM) concentration for 75

minutes and lysates were collected every 15 minutes for the indicated time after CHX treatment.

Stat1 protein level was analyzed by WB. (E) Band densities of Stat1 for each treatment condition

was quantified by ImageJ and normalized to their corresponding GAPDH value. Normalized

values were graphed and trend line equation was predicted for Stat1 half-life estimation. Results

shown are representative experiment from three independent assays.

61

3.3.4 The Effect of c-Src Activity on Stat1 Degradation

c-Src activity effect on protein half-life of Stat1 raised the question whether or not c-Src

regulates protein stability of Stat1 through proteasomal degradation. Degradation of proteins

through proteasome involves enzymes that link chains of the polypeptide co-factor, ubiquitin, on

to proteins to mark them for degradation. Tagged proteins will then be recognized by the 26S

proteasome, a very large multicatalytic protease complex that degrades ubiquitinated proteins to

small peptides (Bhattacharyya et al. 2014). Therefore, to address this question, I examined Stat1

protein level in the absence and presence of MG132, a 26S proteasome inhibitor, when c-Src

activity is inhibited in MC3T3-E1 cells. In this assay first, c-Src activity was inhibited for 2

hours in MC3T3-E1s by two distinct inhibitors including SrcI-1 and PP2, both at the

concentrations of 10 µM. The cells were then either treated with 10 µM MG132 or DMSO

(solvent control) for 4 hours. Lysates were then collected and subjected to WB analysis to

evaluate Stat1 protein level in response to proteasome inhibition. Results showed that addition of

the 26S proteasome inhibitor, MG132, almost rescued the Stat1 loss caused by c-Src inhibition

with PP2 (Figure 3-16A), arguing that c-Src regulates Stat1 expression primarily via a

posttranslational mechanism. Next, to further confirm the specificity of c-Src inhibition I applied

three different c-Src specific siRNAs to down regulate c-Src expression in MC3T3-E1 cells.

Negative select siRNA served as control. 48 hours post- transfection, cells underwent treatment

with either MG132 (10 µM) or DMSO for 4 hours and lysates were collected. WB analysis of

this assay also showed that inhibition of proteasomal degradation by MG132 rescues the loss of

Stat1 comparable to the level observed in control condition, more notably for c-Src siRNA #1

and #2. Results are shown in figure 3-16B.

It has been shown by others that phosphorylation of Stat1 is required for its ubiquitination (Kim

and Maniatis, 1996). They showed that ubiquitination of Y701F mutant form of Stat1 in response

to activation of IFN signaling and MG132 treatment was not detectable while ubiquitination of

Stat1 WT was. Disappearance of Stat1 shortly after its phosphorylation was consistent with their

observations. We have showed that inhibition of c-Src activity results in an increased level in

Stat1 phosphorylation at Y701. We also showed that c-Src inhibition shortens half-life of Stat1

which could be rescued with MG132. Therefore, I postulated that inhibition of c-Src increases

Stat1 ubiquitination. To investigate this, MC3T3-E1s were either treated with proteasomal

inhibitor MG132 or DMSO as a solvent control for four hours. Two hours after the initiation of

62

the experiment treated cells were either exposed to Src inhibitor PP2(10 µM) or DMSO for

another two hours. Lysates were prepared and subjected to immunoprecipitation assay with Stat1

antibody. Precipitated immunocomplexes were then analyzed with WB using Stat1 antibody to

examine Stat1 ubiquitination status in each experimental condition. Accumulation of

ubiquitinated Stat1 was detected by WB analysis in cell lysates in which c-Src activity and

proteasomal degradation were inhibited These results strongly support our earlier hypothesis that

ubiquitination pathways contribute to degradation of Stat1 protein level in the absence of c-Src

activity (Figure 3-16C).

63

Figure 3-16 Inhibition of c-Src activity increases ubiquitin-mediated Stat1 proteolysis.

(A) c-Src activity was inhibited in MC3T3-E1 cells using 10 µM of PP2 and SrcI-1 for 2 hours

along with DMSO controls. After 2 hours, each treatment condition underwent a second

treatment either with MG132 (10 µM) or DMSO as a control for four hours. Lysates were

collected and subjected to WB analysis using Stat1and c-Src antibodies. GAPDH served as the

loading control. (B) c-Src activity was down-regulated in MC3T3-E1 cells using three different

c-Src specific siRNA along with silencer™ select negative siRNA control. After 48 hours, cells

in each condition were subjected to either MG132 (10 µM) or DMSO treatments for four hours.

Lysates were collected and analyzed is a WB assay using Stat1and c-Src antibodies. GAPDH

served as the loading control. Results shown are representative experiment from three

independent assays. (C) Proteasomal degradation was inhibited by MG132 in MC3T3-E1 cells

for two hours. DMSO served as the negative control. After two hours, MG132 and DMSO

A B

C

IP: S

tat1

Imm

unob

lot:

Sta

t1

64

treated cells underwent second treatments with PP2 (10 µM) or DMSO for another 2 hours.

Lysates were prepared and subjected to IP assay using Stat1 antibody. Precipitated

immunocomplexes were then analyzed by WB using both Stat1 antibody.

65

Chapter 4

4 Discussion, Final Conclusions, and Future Directions

4.1 Thesis summary

Src family of non-receptor tyrosine kinases has been shown to have regulatory effects on

osteogenic differentiation. However, its relevance to differentiation process during osteogenesis

in embryonic stem cells is not clearly understood. In this study, I first evaluated c-Src activity

during the 21 days of osteogenic differentiation in mECSs cultures. Results showed that the c-

Src activity assessed by the protein level of c-Src-Y416 drops significantly in early days of

osteogenic differentiation (day 6 to 10). Suppression of c-Src activity in mESCs by PP2 between

day 6 to 10 resulted in a significant increase of osteocalcin expression and mineralization in day

21 osteo-nodules. Furthermore, downregulation of c-Src with specific siRNA in MC3T3-E1 cells

in early days of differentiation resulted in significant increase of osteogenic differentiation

evaluated by osteocalcin expression and mineralization at day 14 and 21 of differentiation.

Finally, overexpression of constitutively active c-Src in MC3T3-E1 cells using lentiviral

particles dramatically decreased the osteogenic differentiation of preosteoblast MC3T3-E1s,

which further confirmed the inhibitory role of c-Src activity at least at some stages of osteogenic

differentiation. Altogether, I concluded that c-Src activity during early days of osteogenic

differentiation could hamper osteogenesis and its inhibition at specific time of differentiation

increases osteoblast differentiation and mineralization.

After establishing c-Src activity role in regulation of osteoblast differentiation, I investigated role

of Runx2 as the downstream molecule in mediating c-Src effect. I showed that c-Src activity has

no immediate effect on expression of Runx2 at mRNA or protein level. However, inhibition of c-

Src activity dramatically increased the nuclear localization of Runx2 assessed by WB and IF

analysis. I also showed that c-Src activity suppresses Runx2 transcriptional activity and therefore

downregulates the expression of Runx2 target genes including BSPII, COL1A, and OC which

subsequently hampers the osteogenic differentiation in mESCs. Finally, to understand how c-Src

activity regulates Runx2 subcellular localization I discovered that Stat1 is the mediating

molecule which anchor Runx2 in the cytoplasm when c-Src is active. IP analysis showed an

increase in Stat1-Runx2 interaction when c-Src is active in mESCs.

66

Moreover, I showed that c-Src activity has no influence on Runx2 subcellular localization when

Stat1 is downregulated suggesting that regulatory effect of c-Src on Runx2 localization is

mediated through Stat1. Further analysis revealed that upon inhibition of protein synthesis with

CHX in mESCs, reduction in Stat1 protein level is considerably faster when c-Src activity is

inhibited compared to control conditions. To be specific, Stat1 protein half-life was estimated to

be reduced from 480.8 minutes to 92.6 minutes when c-Src activity was inhibited in MC3T3-E1

cells which highly suggests a regulatory role for c-Src in Stat1 stability. I assessed c-Src effect

on Stat1 degradation through proteasome-ubiquitination pathway where I found that inhibition of

proteasome 26S by MG132 rescued Stat1 loss caused by c-Src inhibition by PP2 or c-Src

specific siRNAs. Finally, I showed that higher level of ubiquitin is associated with Stat1 in

mESCs treated by PP2 compared to PP3 or DMSO treated cells further supporting that c-Src

activity stabilizes Stat1. A summary of the findings is showed in figure 4-1.

67

Figure 4-1 c-Src activity inhibits osteogenic differentiation through regulation of Stat1

stability

c-Src activity stabilizes Stat1, which in turn increases its interaction with cytoplasmic Runx2 and

retains Runx2 in the cytoplasm. This lowers Runx2 level in the nucleus, which subsequently

decreases Runx2 transcriptional activity and reduction in Runx2 target genes involved is

progression of osteogenic differentiation. Inhibition of c-Src activity on the other hand, increases

Stat1 phosphorylation and its nuclear localization where it degrades. Therefore, less Stat1 would

be available to interact with Runx2 in the cytoplasm. This sets Runx2 free to translocate to the

nucleus and increases Runx2 target osteoblast maker genes’ expression and enhance osteogenic

differentiation.

68

4.2 General Discussion and Conclusions

The contradicting reports in regards of the role to c-Src in osteogenic differentiation (Marzia et

al. 2000; Choi et al. 2015) made us ask whether or not c-Src activity would play a role in

osteogenic differentiation of ESCs. Here, for the first time I showed the dynamic activity of c-

Src during osteogenic differentiation of R1 mESCs. Data showed that c-Src kinase activity

decreased almost monotonically for almost two weeks of differentiation following by a slight

increase during day 14 to day 21 of mESC osteogenic differentiation. The down-regulation of c-

Src kinase activity in the early phase of osteogenic differentiation may facilitate the osteogenesis,

since c-Src expression and activity have been shown to have inhibitory effect on HMSCs

differentiation toward osteoblasts (Marzia et al. 2000; Id et al. 2010). Increase of c-Src activity

later during osteogenic differentiation, starting from day 14, however, may suggest that c-Src

activity at later days might be beneficial in terms of osteogenic differentiation and its

maintenance. One study showed that c-Src interacts with and phosphorylates Osx and

subsequently increases its stability and transcriptional activity (Choi et al. 2015).

Differentiation of osteoblasts from osteoprogenitors could occur during multiple stages.

Different markers and signaling pathways may underlie these stages. Despite efforts in the field

it is still difficult to specify markers to distinguish each stage or to identify their significance.

Fluctuating level of p-Y416-c-Src during osteogenic differentiation may indicate c-Src activity

have inhibitory role in specific stages of osteogenic differentiation. This could explain at least

partially the identified contradicting roles of c-Src in osteogenic differentiation.

Sequence of steps for the cells with decreasing capacity of proliferation and increase in

differentiation have been postulated for osteoblast differentiation which includes immature and

mature osteoprogenitors, preosteoblast, mature osteoblast and osteocytes based on morphological

and histological studies (Aubin, 1998; Roeder et al. 2016). In my study, I selected eight different

time periods during osteogenic differentiation based on gene expression patterns of Runx2, Osx,

COL1A1, BSPII, and OC to test the influence of c-Src activity. The everyday mRNA profile of

the osteogenic markers showed an initial increase in Runx2 level between day 3-5 and between

day 6 to 10. Next, between day 10 to 14 there is a sharp drop in Runx2 level. Other markers

including BSPII and COL1A1 start to increase after day 10. Runx2 increases once more between

day 14 to 17. This could be due to the 1) increased number of premature and mature osteoblasts

69

and/ or 2) the expression of Runx2 from osteoprogenitors with delayed osteogenic differentiation

which could stem for the existence of heterogenous population of cells in the culture. I selected

overlapping time periods of c-Src inhibition which includes days 3-5, 5-10, 10-14, 10-17, 10-21,

14-17, 14-21, and 17-21. Our results showed that inhibition of c-Src activity in early days of

mESCS osteogenic differentiation between day 6 to 10 had the most dramatic effect favoring the

osteoblast differentiation. c-Src inhibition between day 6 to 10 of osteogenic differentiation

results in significantly higher level of OC, the specific osteoblast marker, and a remarkable

increase in mineralization of osteo-nodules assess by both ARS and von Kossa staining.

Continuous downregulation of p-Y416-c-Src at early days of mESCs osteogenic differentiation

starting at day 6 upon induction of osteogenesis by AA also supports the hypothesis that

inhibition of c-Src activity between day 6 to 10 increases osteogenic differentiation. In terms of

inhibition of c-Src activity in other tested time periods, day 3-5 inhibition resulted in a lower

level of osteogenic differentiation based on the lower expression of OC on day21 osteo-nodules

and their low level of mineralization assessed by ARS staining of day 21 osteo-nodules. It is

likely that decreased level in osteogenic differentiation resulted from lower proliferation due to

inhibition of c-Src at the stage that cells proliferate extensively. This is consistent with studies

that suggest c-Src activity promotes proliferation and initiation of differentiation (Zhang et al.

2014a). The second significant lower OC expression and mineralization was observed when c-

Src activity was inhibited between day 10 to 14. This may be due to the heterogeneous ES

osteogenic differentiation in culture. Therefore, the overall differentiation outcome of a

population of cells at proliferation phase for the indicated time were compromised. Also,

considering the potential role of c-Src activity in regulation of Osx transcriptional activity

combined by Osx expression pattern in our setting implies that c-Src inhibition of Osx could play

a role in down regulation of osteogenic differentiation (Choi et al. 2015).

Early inhibition of c-Src activity with specific siRNA in MC3T3-E1 preosteoblasts also resulted

in an increase of OC mRNA expression and mineralization. Previous studies have showed that

SFK inhibitor enhances osteoblast differentiation in MC3T3-E1 mainly through inhibition of c-

Src activity (Lee et al. 2010). However, the observed effect in terms of OC expression and

mineralization was not as profound when compared to the outcome of c-Src inhibition in ESCs.

One explanation for this could be that MC3T3-E1 cells are further advanced in terms of

osteogenic differentiation compared to ESCs and that inhibition of c-Src at such stage would not

70

be as effective as earlier stages. The other explanation is that since we used PP2 to inhibit c-Src

activity in ESCs, it is possible that unintentionally we inhibited other molecules whose activity is

also important in osteogenic differentiation. Understating mechanisms of how c-Src and other

members of SFK family affect osteogenesis would help reveal the reason behind such a

phenomenon.

Overexpression of constitutively active c-Src resulted in a significant decrease in OC mRNA

expression and remarkable reduction in mineralization of MC3T3-E1 after 14 days of osteogenic

differentiation supporting our earlier findings regarding inhibitory role of c-Src activity in

osteogenesis. In conclusion, our study provides evidence that inhibition of c-Src activity in early

ES osteogenesis leads to accelerated osteogenic differentiation, increased matrix production and

mineralization. This confirms that Src exerts a negative regulatory influence on bone formation

and is involved in the regulation of the early events of osteogenic differentiation.

The early commitment of stem cells to become osteoblasts requires expression of Runx2, a

master transcription factor that regulates several genes in osteoblasts, such as type I collagen,

BSP, OP, TGFβ, and OC (Long, 2012). Therefore, we investigated the potential role of c-Src in

regulation of Runx2 expression and transcriptional activity. Results revealed that inhibition of c-

Src activity in mESCs increases Runx2 nuclear localization and transcriptional activity. A link

between c-Src and Runx2 transcriptional activity has been reported previously in ROS 17/2.8

cells. They showed that the endogenous YAP interacts with the native Runx2 protein and

suppresses Runx2 transcriptional activity in a dose-dependent manner in Rat osteosarcoma ROS

17/2.8 (Zaidi et al. 2004). However, Zhang et al. recently demonstrated that the differentiation-

promoting activity of c-Src is antagonized by c-Yes in mES cells, despite their very close

phylogenetic relationship (Zhang et al. 2014b). Therefore, it might be argued that c-Src and Yes

may not be partnering in mES differentiation and that may be the regulation of Runx2

localization is independent of YAP pathway in mESCs. Here, we showed for the first time that c-

Src activity regulates Runx2 subcellular localization in osteoblasts. Furthermore, our results

indicated that Runx2 expression both at mRNA and protein level is not altered upon inhibition of

c-Src activity arguing that inhibition of c-Src activity most likely does not affect the number of

osteoprogenitors during differentiation or at least not for the duration of time that we studied.

71

There are a few mechanisms that could regulate subcellular distribution of a protein in response

to different cellular signaling. Two major factors that affect protein partitioning between the

cytosol and nucleus are interactions with the nuclear transport machinery and interactions with

anchor proteins that reside stably in the nucleus or cytosol. In the first case, signaling may

directly regulate the association of a protein with nuclear import and/or export factors.

Alternatively, a protein may contain a nuclear localization signal (NLS) but fail to enter the

nucleus at a significant rate because of a high affinity interaction with a cytosolic anchor protein

that does not enter the nucleus and may even occlude the NLS of its binding partner. Runx2

transportation into the nucleus is mediated by a NLS, which is located on the C-terminal side of

the ‘Runt domain’ (Thirunavukkarasu et al. 1998). Increase in nuclear localization of Runx2 due

to the inhibition of c-Src activity could have been result of one of the two mechanisms. In one

scenario c-Src, directly or indirectly, could affect NLS function and translocation of Runx2.

Direct or indirect phosphorylation of Runx2 by c-Src could also affect the conformation of the

protein and subsequently its NLS availability to facilitate or prevent Runx2 nuclear localization.

In the other possible scenario, a potential interacting partner of Runx2, Stat1could act as a

cytoplasmic attenuating partner of Runx2 (Sunhwa Kim, 2003). Enhanced transcriptional activity

of Runx2 in Stat1 -/- mice led Kim and his coworkers to discover that Runx2 localization is

regulated by Stat1 which act as Runx2 cytoplasmic anchoring protein. Since Stat1 and Runx2 are

both expressed throughout the osteogenic differentiation, tight regulation of their interaction

which subsequently affects Runx2 function would be necessary. However, regulatory mechanism

of Runx2 and Stat1 interaction during osteogenic differentiation is unclear. Our results showed

that inhibition of c-Src activity decreased the interaction of Runx2 with Stat1. More importantly,

our results confirmed the necessity of Stat1 for c-Src regulatory effect on Stat1-Runx2

interaction when downregulation of Stat1 by specific siRNAs resulted in loss of PP2 effect on

Runx2 nuclear localization. These collectively suggest that lower interaction of Runx2 with Stat1

in the cytoplasm in response to c-Src inhibition results in an increased level of nuclear Runx2

which enhances its transcriptional activity. Previous studies reported a stimulatory role for

Runx2 in early stages of osteogenic differentiation and an inhibitory role later in the process

when osteoblasts are matured (Komori, 2010). Early downregulation of c-Src activity with its

increase at later days during ES osteogenic differentiation is consistent with our findings. Here, I

propose a novel mechanism in which c-Src regulate mESCs osteogenic differentiation through

Runx2 subcellular localization.

72

I postulate mechanisms in which c-Src could directly affect Runx2-Stat1 interaction. One

possible mechanism could be through the phosphorylation by c-Src kinase activity.

Phosphorylation of one or possibly both proteins subsequently could affect protein-protein

interaction via two different scenarios. First, interacting proteins could directly bind to the

phosphorylation site via phosphopeptide binding sequences. SH2 domain of Stat1 for instance is

prone to dock to phosphorylated tyrosine residues on its interacting partners (Reich, 2013).

However, most of the previous reports have emphasized on the significance of Runx2 serine

residues phosphorylation in its cellular function and tyrosine phosphorylation of Runx2 is poorly

studied (Wee et al. 2002; Arumugam et al. 2018). Secondly, binding site of either of the

proteins could become hidden or exposed due to conformational changes resulting from

phosphorylation and subsequently affect protein-protein interactions. Our analysis revealed that

phosphorylation of Stat1 at Y701 is enhanced upon inhibition of c-Src activity and it increased

its nuclear localization. Earlier studies have shown that SHP2 phosphatase activity reduces Stat1

phosphorylation at Y701 (Wu et al. 2002). SHP2 on the other hand has shown to activate c-Src

through controlling Csk, the kinase of inhibitory Y527, access to c-Src (Zhang et al. 2004).

Using WB analysis in mESCs, I confirmed p-Y701-Stat1 is reduced in response to SHP2

inhibition by SHP2 specific inhibitor PHPS1 (Figure A5-A). Furthermore, our results showed

that SHP2 activity, assessed by p-Y580-SHP2 level, is decreased in response to PP2 treatment

(Figure A5-B). Runx2 nuclear level has also showed an increase in response to SHP2 inhibition

by PHPS1 (Figure A5-C). Collectively, our data suggest a notion of possible regulatory role on

Stat1 phosphorylation through the regulation of SHP2 activity. However, more investigation is

needed to clarify the mechanism.

In the context of osteogenesis, it could be argued that phosphorylation of Stat1 at Y701 affects

its conformation in a way that makes it less desirable interacting partner for Runx2. However,

mutagenesis analysis has been reported that phosphorylation status of Stat1 at tyrosine 701,

which controls its nuclear localization, has shown to be irrelevant to Stat1-Runx2 interaction

(Kim et al. 2003). Therefore, I postulate that other events downstream of Stat1 phosphorylation

at Y701 should be considered as alternative pathways of c-Src regulation of Stat1-Runx2

interaction. Mechanisms for proteins degradation in a phosphorylation-dependent manner have

been elucidated in recent years (Swaney et al. 2013). Our results indicating lower level of Stat1

protein in response to inhibition of c-Src activity combined by its increased level of

73

phosphorylation led us to speculate that c-Src regulates Stat1 protein stability. This hypothesis

was further confirmed when interruption of Stat1 biosynthesis by CHX in differentiating mESCs

showed remarkably faster decay of Stat1 when c-Src activity was inhibited by PP2 vs PP3

treated cells. Further analysis estimated that Stat1 half-life in PP2 treated MC3T3-E1 decreased

to 92.6 minutes from 480.8 minutes observed in control condition. These findings suggested that

Stat1 undergoes degradation in the absence of c-Src activity. Ubiquitin-proteasome pathway is

one the major known mechanism for regulation of activated Stat1 (Kim and Maniatis, 1996). Our

results showed that inhibition of proteasome activity by MG132 dramatically reduces loss of

Stat1 while c-Src was inactive either with application of c-Src inhibitor (PP2) or by c-Src

specific siRNA. These findings further confirmed that c-Src effect on Stat1 stability is regulated

through ubiquitin-proteasome pathway. Therefore, I examined Stat1 ubiquitination status in PP2

treated mESCs and where results showed higher level of Ub-associated Stat1 when c-Src activity

was inhibited. Collectively, our results highly suggest a regulatory role of c-Src in regulation of

Stat1 stability through proteasome-ubiquitination pathway.

In summary, it can be concluded that c-Src increases Stat1 stability by reducing Stat1

proteasomal degradation and increase the level of non-phosphorylated Stat1 in the cytoplasm.

Ample cytoplasmic Stat1 will in turn increase Runx2-Stat1 interaction in the cytoplasm and

inhibits Runx2 nuclear localization. Lower level of nuclear Runx2 hinders osteogenic

differentiation.

4.3 Future Directions

4.3.1 Regulation of c-Src by Calreticulin

We have shown that c-Src enhances Stat1 stability which in turn increases the interaction of

Stat1 and Runx2 in the cytoplasm. This affects Runx2 trafficking to the nucleus and is

accountable for the inhibitory effect of c-Src on osteogenic differentiation. Our previous work

suggests that calreticulin (CRT) regulates adipogenic and osteogenic differentiation of mouse ES

cells, whereby loss of CRT results in either abnormally high adipogenic or abnormally low

osteogenic differentiation. CRT is an ER protein with high Ca2+ binding capacity and is involved

in many cellular processes via its function as a chaperone and/or by regulating the intracellular

Ca2+ hemostasis (Michalak et al. 2002; Michalak et al. 2009). A former graduate student in our

lab established that CRT effect on Src kinase activity is via its Ca+2 buffering function such that

74

increasing cytosolic Ca+2 with ionomycin significantly decreases c-Src activity whereas Ca+2

chelation using BAPTA-AM enhances it via phosphorylation of p-Y416-c-Src. These findings

strongly suggest a regulatory role for CRT upstream of c-Src during osteogenic differentiation

and should be further investigated.

4.3.1 Regulation of Stat1 Degradation by ERK Downstream of c-Src

Phosphorylation of Stat1 on serine 727 is required for Stat1 transcriptional activity (O'Shea et al.

2002) and degradation (Soond et al. 2008). Active p42/p44 MAPK-ERK has shown to

phosphorylate STAT1 on serine 727 and targets it for proteasomal degradation (Soond et al.

2008). Using WB analysis, I showed that inhibition of c-Src activity resulted in an increase in

ERK activity (Figure A6). Therefore, enhanced activity of ERK in response to c-Src inhibition

potentially increases p-S727-Stat1 level and enhances Stat1 degradation. This observation may

suggest a role for ERK downstream of c-Src to regulate Stat1 stability and should be further

examined.

4.3.1 Stat1 Proteasomal Degradation by c-Src/SIAH2

E3 ubiquitin ligase seven-in-absentia-2 (SIAH2) has been shown to indirectly abrogate the

tyrosine phosphorylation of Stat1through tyrosine-kinase 2 (TYK2) degradation (Muller et al.

2014). Phosphorylation and activation of SIAH2 by c-Src has also been shown in an independent

study (Sarkar et al. 2012). c-Src regulatory effect on Stat1 proteasomal degradation through

SIAH2 should be further investigated.

74

References

Aguila, H.L., and Rowe, D.W. 2005. Skeletal development, bone remodeling, and hematopoiesis. Immunol Rev. 208: 7-18.

Akiyama, H. Kim, J.E., Nakashima, K., Balmes, G., Iwai, N., Deng J.M., Zhang, Z., Martin, J.F., Behringer, R.R., Nakamura, T., Crombrugghe, B. 2005. Osteo-chondroprogenitor cells are derived from Sox9 expressing precursors. Proc. Natl. Acad. Sci. U S A 102: 14665-70.

Akiyama, H. and de Crombrugghe, B. 2009. Transcriptional control of chondrocyte differentiation. 147-170.

Arumugam, B., Vairamani, M., Partridge, N.C., and Selvamurugan, N. 2018. Characterization of Runx2 phosphorylation sites required for TGF-beta1-mediated stimulation of matrix metalloproteinase-13 expression in osteoblastic cells. J Cell Physiol. 233: 1082-1094.

Aubin, J.E. 1998. Bone stem cells. J Cell Biochem Suppl. 30: 73-82.

Bain, J., McLauchlan, H., Elliott, M., and Cohen, P. 2003. The specificities of protein kinase inhibitors: an update. Biochem. J. 371: 199-204.

Bain, J., Plater, L., Elliott, M., Shpiro, N., Hastie, C.J., McLauchlan, H., Klevernic, I., Arthur, J.S., Alessi, D.R., and Cohen, P. 2007. The selectivity of protein kinase inhibitors: a further update. Biochem. J. 408: 297-315.

Baksh D., Song, L., Tuan, R.S. 2004. Adult mesenchymal stem cells: characterization, differentiation, and application in cell and gene therapy. J Cell Mol Med. 8: 301-16.

Berendsen A.D., Oslen, BR. 2015. Bone development. Bone 80: 14-18.

Bhattacharyya, S., Yu, H., Mim, C., and Matouschek, A. 2014. Regulated protein turnover: snapshots of the proteasome in action. Nat Rev Mol Cell Biol. 15: 122-133.

Bianco, P. 2014. "Mesenchymal" stem cells. Annu Rev Cell Dev Biol. 30: 677-704.

Bonewald, L.F. 2011. The amazing osteocyte. J Bone Miner Res. 26: 229-38.

Bouet, G., Bouleftour, W., Juignet, L., Linossier, M.T., Thomas, M., Vanden Bossche, A., Aubin, J.E., Vico, L., Marchat, D., and Malaval, L. 2015. The impairment of osteogenesis in bone sialoprotein (BSP) knockout calvaria cell cultures is cell density dependent. PLoS One 10: e0117402.

Buttery, L.D., Bourne, S, Xynos J.D., Wood, H, Hughes F.J., Hughes S.P., Episkopou, V, and Polak, J.M. 2001. Differentiation of osteoblasts and in vitro bone formation from murine embryonic stem cells. Tissue Eng. 7: 89-99.

Choi, Y.H., Han, Y., Lee, S.H., Cheong, H., Chun, K.H., Yeo, C.Y., and Lee, K.Y. 2015. Src enhances osteogenic differentiation through phosphorylation of Osterix. Mol Cell Endocrinol. 15: 85-97.

75

Colnot, C. 2009. Skeletal cell fate decisions within periosteum and bone marrow during bone regeneration. J Bone Miner Res. 24: 274-282.

Compton, J.T. and Lee, F.Y. 2014. A review of osteocyte function and the emerging importance of sclerostin. J Bone Joint Surg Am. 96: 1659-68.

Csobonyeiova, M., Polak, S., Zamborsky, R., and Danisovic, L. 2017. iPS cell technologies and their prospect for bone regeneration and disease modeling: A mini review. J Adv Res. 8: 321-327.

Das, A.T., Tenenbaum, L., and Berkhout, B. 2016. Tet-On Systems For Doxycycline-inducible Gene Expression. Curr Gene Ther. 16: 156-167.

Dingwall, M., Marchildon, F., Gunanayagam, A., Louis, C.S., and Wiper-Bergeron, N. 2011. Retinoic acid-induced Smad3 expression is required for the induction of osteoblastogenesis of mesenchymal stem cells. Differentiation 82: 57-65.

Dragoo J.L., Samimi, B., Zhu, M., Hame, SL. Thomas, B.J., Lieberman, J.R., Hedrick, M.K. Benhaim, P. 2003. Tissue-engineered cartilage and bone using stem cells from human infrapatellar fat pads. J Bone Joint Surg Br. 85: 740-7.

Ducy, P., Zhang, R., Geoffroy, V., Ridall, A.L., and Karsenty, G. 1997. Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell. 89: 747-54.

Canalis, E. 2008. Notch Signaling in Osteoblasts. Sci. Signal. 1: pe17.

Espada, J., and Martin-Perez, J. 2017. An Update on Src Family of Nonreceptor Tyrosine Kinases Biology. Int Rev Cell Mol Biol. 331: 83-122.

Fatherazi, S., Matsa-Dunn, D., Foster, B.L., Rutherford, R.B., Somerman, M.J., and Presland, R.B. 2009. Phosphate regulates osteopontin gene transcription. J Dent Res. 88: 39-44.

Feng, X., and Teitelbaum, S.L. 2013. Osteoclasts: New Insights. Bone Res. 1: 11-26.

Fihria A., Lenb, C., Varmac, RS., Solhy, A.L.2017. Hydroxyapatite: A review of syntheses, structure and applications in heterogeneous catalysis. Coordination Chemistry Reviews 347: 48-76.

Franceschi, R.T., Xiao, G., Jiang, D., Gopalakrishnan, R., Yang, S., and Reith, E. 2003. Multiple signaling pathways converge on the Cbfa1/Runx2 transcription factor to regulate osteoblast differentiation. Connect Tissue Res. 44: 109-16.

Gao, C., Guo, H., Mi, Z., Grusby, M.J., and Kuo, P.C. 2007. Osteopontin induces ubiquitin-dependent degradation of STAT1 in RAW264.7 murine macrophages. J Immunol. 178: 1870-81.

Gessin, J.C., Brown, L.J., Gordon, J.S., and Berg, R.A. 1993. Regulation of collagen synthesis in human dermal fibroblasts in contracted collagen gels by ascorbic acid, growth factors, and inhibitors of lipid peroxidation. Exp Cell Res. 206: 283-90.

76

Hamidouche, Z, Hay, E., Vaudin, P., Charbord, P., Schule, R., Marie, P.J., and Fromigue, O. 2008. FHL2 mediates dexamethasone-induced mesenchymal cell differentiation into osteoblasts by activating Wnt/beta-catenin signaling-dependent Runx2 expression. FASEB J. 22: 3813-22.

Heng, B.C., Cao, T., Stanton, L.W., Robson, P., and Olsen, B. 2004. Strategies for directing the differentiation of stem cells into the osteogenic lineage in vitro. J Bone Miner Res. 19: 1379-94.

Huang, W., Yang, S., Shao, J., and Li, Y.P. 2007. Signaling and transcriptional regulation in osteoblast commitment and differentiation. Front Biosci. 12: 3068-92.

Id, B.H., Lagneaux, L.F., Najar, M.F., Piccart, M.F., Ghanem, G.F., Body J.J., Journe, F. 2010. The Src inhibitor dasatinib accelerates the differentiation of human bone marrow-derived mesenchymal stromal cells into osteoblasts. BMC Cancer 17: 298.

Jaenisch, R., and Young, R. 2008. Stem cells, the molecular circuitry of pluripotency and nuclear reprogramming. Cell 132: 567-582.

Jeon, E.J., Lee, K.Y., Choi, N.S., Lee, M.H., Kim, H.N., Jin, Y.H., Ryoo, H.M., Choi, J.Y, Yoshida, M., Nishino, N., Oh, B.C., Lee, K.S., Lee, Y.H., and Bae, S.C. 2006. Bone morphogenetic protein-2 stimulates Runx2 acetylation. J Biol Chem. 281: 16502-11.

Kague, E., Roy, P., Asselin, G., Hu, G., Stanley, A., Albertson, C., Simonet, J., and Fisher, S. 2016. Osterix/Sp7 limits cranial bone initiation sites and is required for formation of sutures. Dev Biol. 413: 160-72.

Kanke, K., Masaki, H., Saito, T., Komiyama, Y., Hojo, H., Nakauchi, H., Lichtler, A.C., Takato, T., Chung, U.I., and Ohba, S. 2014. Stepwise differentiation of pluripotent stem cells into osteoblasts using four small molecules under serum-free and feeder-free conditions. Stem Cell Reports 2: 751-760.

Keller, G. 2005. Embryonic stem cell differentiation: emergence of a new era in biology and medicine. Genes Dev. 19: 1129-1155.

Kern, B., Shen, J., Starbuck, M., and Karsenty, G. 2001. Cbfa1 contributes to the osteoblast-specific expression of type I collagen genes. J Biol Chem. 276: 7101-7.

Kim, H.J. and Park, J.S. 2017. Usage of Human Mesenchymal Stem Cells in Cell-based Therapy: Advantages and Disadvantages. Dev Repord. 21: 1-10.

Kim, J.H., Liu, X., Wang, J., Chen, X., Zhang, H., Kim, S.H., Cui, J., Li, R., Zhang, W., Kong, Y., Zhang, J., Shui, W., Lamplot, J., Rogers, M.R., Zhao, C., Wang, N., Rajan, P., Tomal, J., Statz, J., Wu, N., Luu, H.H., Haydon, R.C., and He, TC. 2013. Wnt signaling in bone formation and its therapeutic potential for bone diseases. Ther. Adv. Musculoskelet. Dis. 5: 13-31.

77

Kim, S., Koga, T., Isobe, M., Kern, B.E., Yokochi, T., Chin, Y.E., Karsenty, G., Taniguchi, T., and Takayanagi, H. 2003. Stat1 functions as a cytoplasmic attenuator of Runx2 in the transcriptional program of osteoblast differentiation. Genes Dev. 17: 1979-1991.

Kim, T.K., and Maniatis, T. 1996. Regulation of interferon-gamma-activated STAT1 by the ubiquitin-proteasome pathway. Science 273: 1717-1719.

Kobayashi, H., Gao, Y.H., Ueta, C., Yamaguchi, A., and Komori, T. 2000. Multilineage differentiation of Cbfa1-deficient calvarial cells in vitro. Biochem Biophys Res Commun. 273: 630-6.

Komori, T. 2005. Regulation of skeletal development by the Runx family of transcription factors. J Cell Biochem. 95: 445-53.

Komori, T. 2006. Regulation of osteoblast differentiation by transcription factors. J Cell Biochem. 99: 1233-9.

Komori, T. 2010. Regulation of osteoblast differentiation by Runx2. Adv Exp Med Biol 658: 43-49.

Komori, T. 2011. Signaling networks in RUNX2-dependent bone development. J Cell Biochem. 112: 750-5.

Komori, T., Yagi, H., Nomura, S., Yamaguchi, A., Sasaki, K., Deguchi, K., Shimizu, Y., Bronson, R.T., Gao, Y.H., Inada, M., Sato, M., Okamoto, R., Kitamura, Y., Yoshiki, S., and Kishimoto, T. 1997. Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell. 89: 755-64.

Kumar, Y., Kapoor, I., Khan, K., Thacker, G., Khan, M.P., Shukla, N., Kanaujiya, J.K., Sanyal, S., Chattopadhyay, N., and Trivedi, A.K. 2015. E3 Ubiquitin Ligase Fbw7 Negatively Regulates Osteoblast Differentiation by Targeting Runx2 for Degradation. J Biol Chem. 290: 30975-87.

Langenbach, F., and Handschel, J. 2013. Effects of dexamethasone, ascorbic acid and beta-glycerophosphate on the osteogenic differentiation of stem cells in vitro. Stem Cell Res Ther. 4: 117.

Lee J.A., Parrett, B.M., Conejero, J.A., Laser, J., Chen, J., Kogon , A.J., Nanda, D., Grant, R.T., Breitbart, A.S. 2003. Biological alchemy: engineering bone and fat from fat-derived stem cells. Ann Plast Surg. 50: 610-7.

Lee, Y.C., Huang, C.F., Murshed, M., Chu, K., Araujo, J.C., Ye, X., deCrombrugghe, B., Yu-Lee, L.Y., Gallick, G.E., and Lin, S.H. 2010. Src family kinase/abl inhibitor dasatinib suppresses proliferation and enhances differentiation of osteoblasts. Oncogene 29: 3196-3207.

Li, J., Zhang, H., Yang, C., Li, Y., and Dai, Z. 2016. An overview of osteocalcin progress. J Bone Miner Metab. 34: 367-79.

78

Long, F. 2012. Building strong bones: molecular regulation of the osteoblast lineage. Nature Reviews Molecular Cell Biology 13: 27-38.

Long, M.W. 2001. Osteogenesis and bone-marrow-derived cells. Blood Cells Mol Dis. 27: 677-90.

Mackenzie, T.C. and Flake, A.W. 2001. Human mesenchymal stem cells persist, demonstrate site-specific multipotential differentiation, and are present in sites of wound healing and tissue regeneration after transplantation into fetal sheep. Blood Cells Mol Dis. 27: 601-4.

Malaval, L., Wade-Gueye, N.M., Boudiffa, M., Fei, J., Zirngibl, R., Chen, F., Laroche, N., Roux, J.P., Burt-Pichat, B., Duboeuf, F., Boivin, G., Jurdic, P., Lafage-Proust, M.H., Amedee, J, Vico, L., Rossant, J., and Aubin, J.E. 2008. Bone sialoprotein plays a functional role in bone formation and osteoclastogenesis. J Exp Med. 205: 1145-1153.

Maruyama, Z., Yoshida, C.A., Furuichi, T., Amizuka, N., Ito, M., Fukuyama, R., Miyazaki, T., Kitaura, H., Nakamura, K., Fujita, T., Kanatani, N., Moriishi, T., Yamana, K., Liu, W.F., Kawaguchi, H., Nakamura, K., and Komori, T. 2007. Runx2 determines bone maturity and turnover rate in postnatal bone development and is involved in bone loss in estrogen deficiency. Dev Dyn. 236: 1876-90.

Marzia, M., Sims, N.A., Voit, S., Migliaccio, S., Taranta, A., Bernardini, S., Faraggiana, T., Yoneda, T., Mundy, G.R., Boyce, B.F., Baron, R., and Teti, A. 2000. Decreased c-Src expression enhances osteoblast differentiation and bone formation. J Cell Biol. 151: 311-320.

Masson, A.O., Hess, R, O'Brien, K, Bertram, K.L., Tailor, P., Irvine, E., Ren, G., and Krawetz, R.J. 2015. Increased levels of p21((CIP1/WAF1)) correlate with decreased chondrogenic differentiation potential in synovial membrane progenitor cells. Mech Ageing Dev. 149: 31-40.

Meyn, M.A., Schreiner, S.J., Dumitrescu, T.P., Nau, G.J., and Smithgall, T.E. 2005. SRC family kinase activity is required for murine embryonic stem cell growth and differentiation. Mol Pharmacol. 68: 1320-30.

Michalak, M., Groenendyk, J., Szabo, E., Gold, L.I., and Opas, M. 2009. Calreticulin, a multi-process calcium-buffering chaperone of the endoplasmic reticulum. Biochem J. 417: 651-66.

Michalak, M., Robert Parker, J.M., and Opas, M. 2002. Ca2+ signaling and calcium binding chaperones of the endoplasmic reticulum. Cell Calcium. 32: 269-78.

Miller, SC, Saint-Georges, L, Bowman, BM, and Jee, WS. 1989. Bone lining cells: structure and function. Scanning Microsc. 3: 953-60.

Miraoui, H., Oudina, K., Petite, H., Tanimoto, Y., Moriyama, K., and Marie, P.J. 2008. Fibroblast growth factor receptor 2 promotes osteogenic differentiation in mesenchymal cells via ERK1/2 and protein kinase C signaling. J. Biol. Chem. 284: 4897-4904.

79

Muller, S., Chen, Y., Ginter, T., Schafer, C., Buchwald, M., Schmitz, L.M., Klitzsch, J., Schutz, A., Haitel, A., Schmid, K., Moriggl, R., Kenner, L., Friedrich, K., Haan, C., Petersen, I., Heinzel, T., and Kramer, O.H. 2014. SIAH2 antagonizes TYK2-STAT3 signaling in lung carcinoma cells. Oncotarget. 5: 3184-3196.

Murry, C.E. and Keller, G. 2008. Differentiation of embryonic stem cells to clinically relevant populations: lessons from embryonic development. Cell. 132: 661-80.

Nakano, T., Kodama, H., and Honjo, T. 1994. Generation of lymphohematopoietic cells from embryonic stem cells in culture. Science 265: 1098-101.

Nakashima, K., Zhou, X., Kunkel, G., Zhang, Z., Deng, J.M., Behringer, R.R., and de Crombrugghe, B. 2002. The novel zinc finger-containing transcription factor osterix is required for osteoblast differentiation and bone formation. Cell. 108: 17-29.

O'Shea, J.J., Gadina, M., and Schreiber, R.D. 2002. Cytokine signaling in 2002: new surprises in the Jak/Stat pathway. Cell 109: S121-31.

Olmsted-Davis, E.A., Gugala, Z., Camargo, F., Gannon, F.H., Jackson, K., Kienstra, K.A., Shine, H.D., Lindsey, R.W., Hirschi, K.K., Goodell, M.A., Brenner, M.K., Davis, A.R. 2003. Primitive adult hematopoietic stem cells can function as osteoblast precursors. Proc Natl Acad Sci U S A 100: 15877-82.

Ornitz, D.M. and Marie, P.J. 2015. Fibroblast growth factor signaling in skeletal development and disease. Genes Dev. 29: 1463-86.

Otto, F., Thornell, A.P., Crompton, T., Denzel, A., Gilmour, K.C., Rosewell, I.R., Stamp, G.W., Beddington, R., Mundlos, S., Olsen, B.R., Selby, P., and Owen, M.J. 1997. Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell. 89: 765-71.

Owen,T.A., Aronow, M., Shalhoub,V, Baone, L.M., Wilming, L., Tassinari, M.S., Kennedy, M.B., Pockwinse, S., Lian, J.B., Stein, G.S. 1999. Progressive development of the rat osteoblast phenotype in vitro: reciprocal relationships in expression of genes associated with osteoblast proliferation and differentiation during formation of the bone extracellular matrix. J Cell Physiol. 143: 420-30.

Park, O.J., Kim, H.J., Woo, K.M., Baek, J.H., and Ryoo, H.M. 2010. FGF2-activated ERK mitogen-activated protein kinase enhances Runx2 acetylation and stabilization. J Biol Chem. 285: 3568-74.

Peruzzi, B., Cappariello, A., Del Fattore, A., Rucci, N., De Benedetti, F., and Teti, A. 2012. c-Src and IL-6 inhibit osteoblast differentiation and integrate IGFBP5 signalling. Nat Commun. 3: 630.

Phillips, J.E., Gersbach, C.A., Wojtowicz, A.M., and Garcia, A.J. 2006. Glucocorticoid-induced osteogenesis is negatively regulated by Runx2/Cbfa1 serine phosphorylation. J Cell Sci. 119: 581-91.

80

Pineda, E.T., Nerem, R.M., and Ahsan, T. 2013. Differentiation patterns of embryonic stem cells in two- versus three-dimensional culture. Cells Tissues Organs 197: 399-410.

Pittenger, M.F., Mackay, A.M., Beck, S.C., Jaiswal, R.K., Douglas, R., Mosca, J.D., Moorman, M.A., Simonetti, D.W., Craig, S., and Marshak, D.R. 1999. Multilineage potential of adult human mesenchymal stem cells. Science 284: 143-147.

Rahman, M.S., Akhtar, N., Jamil, H.M., Banik, R.S., and Asaduzzaman, S.M. 2015. TGF-beta/BMP signaling and other molecular events: regulation of osteoblastogenesis and bone formation. Bone Res. 14: 15005.

Rauch C., Brunet, A.C., Deleule, J., Farge, E. 2002. C2C12 myoblast/osteoblast transdifferentiation steps enhanced by epigenetic inhibition of BMP2 endocytosis. Am J Physiol Cell Physiol. 283: C235-45.

Regan, J., and Long, F. 2013. Notch signaling and bone remodeling. Curr Osteoporos Rep. 11: 126-129.

Reich, N.C. 2013. STATs get their move on. JAKSTAT. 2: e27080.

Reubinoff, B.E., Pera, M.F., Fong, C.Y., Trounson, A., and Bongso, A. 2000. Embryonic stem cell lines from human blastocysts: somatic differentiation in vitro. Nat Biotechnol. 18: 399-404.

Roeder, E., Matthews, B.G., and Kalajzic, I. 2016. Visual reporters for study of the osteoblast lineage. Bone 92: 189-195.

Roskoski, R. 2005. Src kinase regulation by phosphorylation and dephosphorylation. Biochem Biophys Res Commun. 331: 1-14.

Rous, P. 1911. A Sarcoma of the fowl transmissible by an agent separable from the tumor cells. J Exp Med. 13: 397-411.

Rutkovskiy, A., Stenslokken, K., Vaage, I.J. 2016. Osteoblast Differentiation at a Glance. Med Sci Monit Basic Res. 22: 95-106.

Sarkar, T.R., Sharan, S., Wang, J., Pawar, S.A., Cantwell, C.A., Johnson, P.F., Morrison, D.K., Wang, J.M., and Sterneck, E. 2012. Identification of a Src tyrosine kinase/SIAH2 E3 ubiquitin ligase pathway that regulates C/EBP delta expression and contributes to transformation of breast tumor cells. Mol Cell Biol. 32: 320-332.

Schroeder, T.M., Kahler, R.A., Li, X., and Westendorf, J.J. 2004. Histone deacetylase 3 interacts with runx2 to repress the osteocalcin promoter and regulate osteoblast differentiation. J Biol Chem. 279: 41998-2007.

Sinha, K.M., and Zhou, X. 2013. Genetic and molecular control of osterix in skeletal formation. J Cell Biochem. 114: 975-84.

81

Soond, S.M., Townsend, P.A., Barry, S.P., Knight, R.A., Latchman, D.S., and Stephanou, A. 2008. ERK and the F-box protein betaTRCP target STAT1 for degradation. J Biol Chem. 283: 16077-83.

Soriano, P., Montgomery, C., Geske, R., and Bradley, A. 1991. Targeted disruption of the c-src proto-oncogene leads to osteopetrosis in mice. Cell. 64: 693-702.

Stehelin, D., Varmus, H.E., Bishop, J.M., and Vogt, P.K. 1976. DNA related to the transforming gene(s) of avian sarcoma viruses is present in normal avian DNA. Nature. 260: 170-3.

Stephens, A.S. and Morrison, N.A. 2014. Novel target genes of RUNX2 transcription factor and 1,25-dihydroxyvitamin D3. J Cell Biochem. 115: 1594-608.

Sterner, D.E. and Berger, S.L. 2000. Acetylation of histones and transcription-related factors. Microbiol Mol Biol Rev. 64: 435-59.

Stock, M., Schafer, H., Fliegauf, M., and Otto, F. 2004. Identification of novel genes of the bone-specific transcription factor Runx2. J Bone Miner Res. 19: 959-72.

Kim, S., Koga, T., Isobe, M., Kern, B.E., Yokochi, T, Chin, Y.E. Karsenty, G., Taniguchi, T. Takayanagi, H. 2003. Stat1 functions as a cytoplasmic attenuator of Runx2 in the transcriptional program of osteoblast differentiation. 17: 1979-1991.

Surinder M.S., Townsend, P.A., Barry, S.P., Knight, R.A., Latchman, D.S., Stephanou, A. 2008. ERK and the F-box Protein ßTRCP Target STAT1 for Degradation. J Biol Chem. 283: 16077-16083.

Swaney, D.L., Beltrao, P., Starita, L., Guo, A., Rush, J., Fields, S., Krogan, N.J., and Villen, J. 2013. Global analysis of phosphorylation and ubiquitylation cross-talk in protein degradation. Nat Methods. 10: 676-682.

Tajima, K., Takaishi, H., Takito, J., Tohmonda, T., Yoda, M., Ota, N., Kosaki, N., Matsumoto, M., Ikegami, H., Nakamura, T., Kimura, T., Okada, Y., Horiuchi, K., Chiba, K., and Toyama, Y. 2010. Inhibition of STAT1 accelerates bone fracture healing. J Orthop Res. 28: 937-41.

Thirunavukkarasu, K., Mahajan, M., McLarren, K.W., Stifani, S., and Karsenty, G. 1998. Two domains unique to osteoblast-specific transcription factor Osf2/Cbfa1 contribute to its transactivation function and its inability to heterodimerize with Cbfbeta. Mol Cell Biol. 18: 4197-208.

Thomson, J.A., Itskovitz-Eldor, J., Shapiro, S.S., Waknitz, M.A., Swiergiel, J.J., Marshall, V.S., and Jones, J.M. 1998. Embryonic stem cell lines derived from human blastocysts. Science 282: 1145-7.

Trounson, A. 2002. Human embryonic stem cells: mother of all cell and tissue types. Reprod Biomed Online 4: 58-63.

82

Viguet-Carrin, S., Garnero, P., and Delmas, P.D. 2006. The role of collagen in bone strength. Osteoporos Int. 17: 319-36.

Vimalraj, S., Arumugam, B., Miranda, P.J., and Selvamurugan, N. 2015. Runx2: Structure, function, and phosphorylation in osteoblast differentiation. Int J Biol Macromol. 78: 202-8.

Wang, D., Christensen, K., Chawla, K., Xiao, G., Krebsbach, P.H., and Franceschi, R.T. 1999. Isolation and characterization of MC3T3-E1 preosteoblast subclones with distinct in vitro and in vivo differentiation/mineralization potential. J Bone Miner Res. 14: 893-903.

Wang, H., Pang, B., Li, Y.F., Zhu, D., Pang, T., and Liu, Y. 2012. Dexamethasone has variable effects on mesenchymal stromal cells. Cytotherapy 14: 423-30.

Wee, H.J., Huang, G., Shigesada, K., and Ito, Y. 2002. Serine phosphorylation of RUNX2 with novel potential functions as negative regulatory mechanisms. EMBO Rep. 3: 967-974.

WeinEmail M.N. 2017. Bone Lining Cells: Normal Physiology and Role in Response to Anabolic Osteoporosis Treatments. Current Molecular Biology Reports 3: 79-84.

Wu, T.R., Hong, Y.K., Wang, X.D., Ling, M.Y., Dragoi, A.M., Chung, A.S., Campbell, A.G., Han, Z.Y., Feng, G.S., and Chin, Y.E. 2002. SHP-2 is a dual-specificity phosphatase involved in Stat1 dephosphorylation at both tyrosine and serine residues in nuclei. J Biol Chem. 277: 47572-47580.

Yu, Y., Al Mansoori, L., and Opas, M. 2015. Optimized osteogenic differentiation protocol from R1 mouse embryonic stem cells in vitro. Differentiation 89: 1-10.

Zaidi, S.K., Sullivan, A.J., Medina, R., Ito, Y., van Wijnen, A.J., Stein, J.L., Lian, J.B., and Stein, G.S. 2004. Tyrosine phosphorylation controls Runx2-mediated subnuclear targeting of YAP to repress transcription. EMBO J. 23: 790-799.

Zayzafoon, M. 2006. Calcium/calmodulin signaling controls osteoblast growth and differentiation. J Cell Biochem. 97: 56-70.

Zhang, S.Q., Yang, W., Kontaridis, M.I., Bivona, T.G., Wen, G., Araki, T., Luo, J., Thompson, J.A., Schraven, B.L., Philips, M.R., and Neel, B.G. 2004. Shp2 regulates SRC family kinase activity and Ras/Erk activation by controlling Csk recruitment. Mol Cell. 13: 341-355.

Zhang, X., Simerly, C., Hartnett, C., Schatten, G., and Smithgall, T.E. 2014b. Src-family tyrosine kinase activities are essential for differentiation of human embryonic stem cells. Stem Cell Res. 13: 379-389.

Zhang, X., Simerly, C., Hartnett, C., Schatten, G., and Smithgall, T.E. 2014a. Src-family tyrosine kinase activities are essential for differentiation of human embryonic stem cells. Stem Cell Res. 13: 379-389.

Zheng, X.M., Resnick, R.J., and Shalloway, D. 2000. A phosphotyrosine displacement mechanism for activation of Src by PTPalpha. EMBO J. 19: 964-78.

83

Zhou, X., von der Mark, K., Henry, S., Norton, W., Adams, H., and de Crombrugghe, B. 2014. Chondrocytes transdifferentiate into osteoblasts in endochondral bone during development, postnatal growth and fracture healing in mice. PLoS Genet. 10: e1004820.

Zhu, W., He, X., Hua, Y., Li, Q., Wang, J., and Gan, X. 2017. The E3 ubiquitin ligase WWP2 facilitates RUNX2 protein transactivation in a mono-ubiquitination manner during osteogenic differentiation. J Biol Chem. 292: 11178-11188.

Zou, L., Kidwai, F.K., Kopher, R.A., Motl, J., Kellum, C.A., Westendorf, J.J., and Kaufman, D.S. 2015. Use of RUNX2 expression to identify osteogenic progenitor cells derived from human embryonic stem cells. Stem Cell Reports 4: 190-198.

87

Appendices

88

Figure A-1

Figure A-1 Expression of pluripotency markers in early mES osteogenic differentiation

RNA expression of pluripotency markers including Oct3/4, Nanog, and Sox2 were measured by

qPCR in mESCs going through osteogenic differentiation for the indicated days. The primer

pairs are listed in table A-1. Data shown represent the mean (±SD) of triplicates. Data shown are

representative experiments of three independent assays.

Days of osteogenic differentiation

Rel

ativ

e m

RN

A E

xpre

ssio

n

89

Figure A-2

Figure A-2 Inhibition of c-Src activity by pharmacological inhibitors

(A and B) Three different inhibitors of c-Src were applied in the concentrations of 1µM and 10 µM in day

5 differentiating mES EBs for 2 and 24 hours. Lysates were collected and subjected to immunoblot

analysis using p-Y416-c-Src and c-Src antibodies. GAPDH served as loading control. (C) Day 5

differentiating EBs were treated with different concentrations of PP2, and PP3 (inactive analog PP2) of

including 100 nM, 1, 5, 10, and 20 µM for 2 hours. Adjusted volume of DMSO for each corresponding

concentration of PP2 served as the solvent control. Lysates were prepared and subjected to immunoblot

analysis using p-Y416-c-Src and c-Src antibodies. GAPDH served as loading control. (D) Band density

for each condition in the WB was quantified using ImageJ software and those for p-Y416-c-Src were

normalized to their corresponding GAPDH. Calculated values were graphed. Data shown are

representative experiments of three independent assays.

A B

C

D

D

90

Figure A-3

A

B

C

91

Figure A-3 Failed attempts for transfecting mESCs with c-Src siRNAs

Different c-Src siRNA from Ambion (A) and Thermo Fisher Scientific (B) along with GAPDH

specific siRNA as a positive control were applied using transfectamin to downregulate c-Src

activity in mESCs for the indicated concentrations. Lysates were collected and subjected to WB

analysis using c-Src antibody. GAPDH served as loading control. (C) mESCs were subjected to

transfection using electroporation with the indicated concentrations of c-Src and GAPDH

specific siRNAs. Lysates were analyzed by WB using c-Src and GAPDH antibodies. Data shown

are representative experiments of three independent assays.

92

Figure A-4

Figure A-4 Stat1 phosphorylation status on Y701 in early days of osteogenic differentiation

(A) Expression and phosphorylation status of proteins including p-Y701-Stat1, Stat1, p-Y527-c-

Src (inactive c-Src), p-Y416-c-Src (active c-Src), and c-Src were analyzed by immunoblotting.

GAPDH served as the loading control and is shown below its corresponding blots. (B) band

densities of p-Y701-Stat1 and p-Y416-c-Src were normalized to their corresponding total

protein, Stat1 and c-Src, and calculated values are shown in the graph. Data shown are

representative experiments of three independent assays.

Day (s) 3 5 6 7 8 9 10

p-Y416-c-

GAPDH

c-Src

Stat1

p-Y701-Stat1

GAPDH

p-Y527-c-

A B

93

Figure A-5

p-Y701-Stat1

Stat1

GAPDH

c-Src Y416

Whole cell lysates

Whole cell lysates

C

B A

Figure A-5 Shp2 inhibition increases Stat1 Y701

phosphorylation and Runx2 localization.

(A) Shp2 activity was inhibited using specific PHPS1

inhibitor (5 µM) for 2 hours. Expression and

phosphorylation status of proteins including p-Y701-

Stat1, Stat1, and p-Y416-c-Src were analyzed by

immunoblotting. GAPDH served as the loading

control. (B) Runx2 nuclear fraction in mESCs was

enhanced in response to Shp2 inhibition by PHPS1.

(C) Inhibition of c-Src activity by PP2 resulted in

reduced Shp2 activity (p-Y580-Shp2). GAPDH served

as the loading control and it is shown below each

corresponding blot.

94

Figure A-6

Figure A-6 Shp2 inhibition increases ERK activity.

Shp2 activity was inhibited using specific PHPS1 inhibitor (5 µM) for 2 hours. Expression and

phosphorylation status of proteins including p-Y416-c-Src, pERK, ERK were analyzed by

immunoblotting. GAPDH served as the loading control and it is shown below each

corresponding blot.

Whole Cell Lysates

95

Table A-1

Genes Forward primers Reverse Primers

L32 5’-CATGGCTGCCCTTCGGCCTC- 3’ 5’-CATTCTCTTCGCTGCGTAGCC-3’

Nanog 5’-AGGGTCTGCTACTGAGATGCTC-3’ 5’-CAACCACTGGTTTTTCTGCCAC-3’

Oct3/4 5’-TCTTCTGCTTCAGCAGCTTG-3’ 5’-GTTGGAGAAGGTGGAACCAA-3’

Sox2 5’-CACAACTCGGAGATCAGCAA-3’ 5’-CTCCGGGAAGCGTGTACTTA-3’

Runx2 5’-CCTCTGACTTCTGCCTCTGG-3’ 5’-TAAAGGTGGCTGGGTAGTGC-3’

Osx 5’-AAGTCCCACACAGCAGCTG-3’ 5’-AGCCGAGCTGCCAGAGTTTG-3’

BSPII 5’-AACAATCCGTGCCACTCA-3’ 5’-GGAGGGGGCTTCACTGAT-3’

COL1A1 5’-AGCAGGTCCTTGGAAACCTT-3’ 5’-AAGGAGTTTCATCTGGCCCT-3’

COL 2A1 5’-GCAAGATGAGGGCTTCCATA-3’ 5’-CTACGGTGTCAGGGCCAG-3’

OC 5’-GCCGGAGTCTGTTCACTACC-3’ 5’-GCGCTCTGTCTCTCTGACCT-3’

Stat1 5’-GGAGCACGCTGCCTATGATG-3’ 5’-CTCCAGAGAAAAGCGGCTGTA-3’

Table A-1 Primer pairs used in qPCR analysis in this study.

96

Table A-2

Genes Forward primers Reverse Primers

BSPII 5’-GCCTCAGTTGAATAAACATGAAA-3’ 5’-TCCTCACCCTTCAATTAAATCC-3’

COL1A1 5’-GCTTCCACGTTTACAGCTCTAAA-3’ 5’-GTCAGGAAAGGGTCATCTGTAGTC-3’

OC 5′-CTAATTGGGGGTCATGTGCT-3′ 5′-CCAGCTGAGGCTGAGAGAGA-3′

COL 2 5’-CACTTACATTGGCATGTTGCTAG-3’ 5’-GGGCTTTATTATTTTAGCACCAC-3’

Table A-2 Primer pairs used in ChIP analysis in this study.

97

Tab

le A-3 c-S

rc specific siR

NA

s used

in th

is stud

y.

Tab

le A-3