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Interactions between Undifferentiated and
Osteogenically Differentiated Mesenchymal
Stromal Cells during Osteogenesis
Yinghong Zhou BDS, MDS
Institute of Health and Biomedical Innovation
School of Chemistry, Physics and Mechanical Engineering (CPME)
Faculty of Science & Engineering
Queensland University of Technology
Thesis submitted for the award of Doctor of Philosophy
February 2013
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KEYWORDS
Bone regeneration, bone tissue engineering, cell-based therapy, cell interaction, cell
recruitment, mesenchymal stromal cells (MSCs), osteogenesis, osteogenic
differentiation, vascular endothelial growth factor (VEGF), CXCL12/CXCR4,
phosphoinositide 3-kinase (PI3K) /Akt
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ABSTRACT
Despite tremendous efforts that have been made to develop new therapeutic strategies,
including the fabrication of novel bioactive scaffolds, the application of various
growth factors and gene-modified cells in pre-clinical or clinical settings, bone defect
healing still poses a great challenge to orthopaedic surgeons.
Increasing evidence supports that cell-based therapy has emerged as one of the most
promising therapeutic approaches for tissue repair and regeneration due to the
inherent characteristics of mesenchymal stromal cells (MSCs) in respect to their self-
renewal capacity and multipotent differentiation potential. More interestingly, recent
experimental evidence suggests that implanted MSCs have a close communication
with host cells, and the contribution of donor cells is beyond their direct conversion
into bone forming cells. Collectively, these findings indicate that transplanted donor
cells play an important role in bone defect healing. However, how transplanted donor
cells initiate osteogenesis, their role in the recruitment of host MSCs to form
functional structures and the mechanisms behind these processes remain largely
undefined. Motivated by the issues mentioned above, the purpose of this thesis was to
unveil the interactions between donor cells and host cells during osteogenesis and in
particular the involvement of VEGF, the CXCL12/CXCR4 axis and the PI3K/Akt
signaling pathway in cell recruitment and donor-host cell interactions. This project
was designed as three separate but closely-linked segments which, in the end, led to
three independent papers.
The first segment of this project investigated how osteogenically differentiated human
mesenchymal stromal cells (O-hMSCs) were involved in the osteogenic processes and
how they interacted with the host cells after implantation. Histology,
immunohistochemistry and in situ hybridisation were some of the techniques applied
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in this part of the study to identify the interaction between donor cells and host cells in
vivo. Results revealed that in both ectopic (subcutaneous pocket) and orthotopic (skull
defect) sites of severe combined immunodeficiency (SCID) mice, donor O-hMSCs
could recruit host cells to initiate osteogenesis. We demonstrated that the ratio of
donor cells and host cells inside the implants was significantly dependent on the local
micro-environment and directly correlated with osteogenesis. The findings from this
study provided clear evidence that donor O-hMSCs play an integral role in host cell
recruitment during osteogenesis.
Osteogenesis and angiogenesis are two closely correlated processes during functional
bone formation and regeneration. As the best characterised angiogenic mediator,
VEGF is believed to play a crucial role in skeletal development by enhancement of
the coupling of angiogenesis and osteogenesis. The second segment of this project
was designed to investigate the role of VEGF in the interaction between
undifferentiated and osteogenically differentiated bone marrow stromal cells (MSCs
and OMSCs) in vitro and in vivo. VEGF secretion increased dramatically in MSCs
after osteogenic differentiation. Furthermore, the secretion of VEGF from OMSCs
altered the expression pattern of CXCL12/CXCR4 in MSCs in vitro, which may be
responsible for the increased cell migration in vivo. In situ hybridisation demonstrated
that donor OMSCs recruited host cells during the osteogenic processes at the
orthotopic sites, and blocking VEGF with neutralizing antibody resulted in a
significant decrease of host cell recruitment and new bone formation. These findings
suggest that the interaction between donor OMSCs and host MSCs might be mediated
by the paracrine effect of VEGF and the activation of CXCL12/CXCR4 axis,
resulting in enhanced host MSCs migration to the implantation site.
Based on the findings of the second segment, subsequent experiments were designed
to determine the possible mechanisms that drive the host cells to migrate to the defect
site. Our co-culture studies demonstrated that OMSCs induced phenotypic changes in
MSCs via dimerization of platelet-derived growth factor receptor (PDGFR), which
then activated the signal transduction. The PI3K/Akt signaling pathway was probed
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by using targeted PI3K inhibitor LY294002, and the activity was monitored by
Western blotting analysis using total- and phospho-Akt specific antibodies.
Downstream targets of the PI3K/Akt signaling pathway, mTOR, p70S6k
and PAK1,
which regulate actin organization and cell motility, were also activated in the presence
of OMSCs-derived conditioned medium. The findings from this segment
demonstrated that VEGF secreted by OMSCs plays a critical role in recruiting host
cells to accelerate osteogenesis via the activation of PDGFRs, which then activate the
PI3K/Akt signaling pathway.
In summary, the body of work presented in this dissertation has demonstrated that the
interactions between donor cells and host cells were critical for the initiation of
ectopic and orthotopic osteogenesis and that this process was mediated by the VEGF,
CXCL12/CXCR4 and the PI3K/Akt signaling pathway. The findings from this
dissertation may provide a scientific rationale for the development of novel
therapeutic strategies in the treatment and management of bone defects.
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TABLE OF CONTENTS
KEYWORDS ............................................................................................................. i
ABSTRACT .............................................................................................................. ii
TABLE OF CONTENTS .......................................................................................... v
LIST OF FIGURES ................................................................................................. vii
LIST OF ABBREVIATIONS ................................................................................... ix
STATEMENT OF ORIGINAL AUTHORSHIP ...................................................... xii
LIST OF PUBLICATIONS .................................................................................... xiii
LIST OF CONFERENCE PRESENTATIONS ........................................................ xv
ACKNOWLEDGEMENTS ................................................................................... xvii
CHAPTER 1 INTRODUCTION................................................................................. 1
1.1 Preface ........................................................................................................ 1
1.2 Main purpose of this study .......................................................................... 2
1.3 Questions to be addressed............................................................................ 2
1.4 Possible outcomes and significance ............................................................. 3
CHAPTER 2 OVERVIEW OF PROJECT DESIGN .................................................... 4
2.1 The effect of pre-programmed donor cell on host cell recruitment during
osteogenesis ........................................................................................................... 4
2.2 Possible regulatory factors that lead to the recruitment of host cells ............. 5
2.3 Potential cell signaling pathways involved in the host cell recruitment ........ 6
CHAPTER 3 LITERATURE REVIEW ....................................................................... 7
3.1 Overview .................................................................................................... 7
3.2 Cell-based therapy for bone defects ............................................................. 8
3.3 Cell sources for cell-based bone tissue engineering ................................... 10
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3.4 The contribution of donor cells in cell-based bone tissue engineering ........ 15
3.5 The involvement of host cells in cell-based bone tissue engineering .......... 16
3.6 Chemokines and growth factors related with cell migration ....................... 19
3.7 Signaling pathways related with cell migration .......................................... 27
3.8 Summary ................................................................................................... 34
CHAPTER 4 RESEARCH REPORTS ...................................................................... 35
REPORT ONE..................................................................................................... 37
Implantation of osteogenically differentiated donor mesenchymal stromal cells
causes recruitment of host cells ............................................................................ 37
REPORT TWO .................................................................................................... 55
VEGF induced CXCL12/CXCR4 axis in the recruitment of mesenchymal stromal
cells (MSCs) during osteogenesis......................................................................... 55
REPORT THREE ................................................................................................ 81
Involvement of VEGF/PDGFR and the PI3K/Akt signaling pathway in OMSCs-
induced MSCs migration ..................................................................................... 81
CHAPTER 5 CONCLUSIONS AND PERSPECTIVES ............................................103
5.1 General discussion ..................................................................................104
5.2 Project summary .....................................................................................113
5.3 Future direction .......................................................................................115
REFERENCES.......................................................................................................117
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LIST OF FIGURES
Figure 1. Schematic of cell-based therapy strategy .................................................... 9
Figure 2. Schematic of osteoblast differentiation..................................................... 13
Figure 3. Schematic of endogenous cells homing to the damaged tissues for in situ
tissue regeneration ................................................................................................... 17
Figure 4. CXCL12/CXCR4 signaling pathway ....................................................... 22
Figure 5. Vascular endothelial growth factor (VEGF) family members and their
receptors .................................................................................................................. 24
Figure 6. Simplified schematic of the platelet-derived growth factor (PDGF) system
................................................................................................................................ 26
Figure 7. Signaling through phosphatidylinositol (3,4,5)-triphosphate .................... 29
Figure 8. Schematic of the three mammalian mitogen-activated protein kinase
(MAPK) signaling pathways ................................................................................... 33
Figure 9. hMSCs characterisation and in vitro osteogenic differentiation ................ 45
Figure 10. In vitro migration assay. ........................................................................ 46
Figure 11. Osteogenesis in subcutaneous and skull defect implants ......................... 48
Figure 12. Distribution of donor human cells and host mouse cells in the
subcutaneous implants revealed by ISH ................................................................... 49
Figure 13. Distribution of O-hMSCs and host mouse cells in the skull defect implants
and bone marrows, revealed by ISH ........................................................................ 51
Figure 14. Histological evaluation of a skull defect model after a 4-week implantation
of OMSCs ............................................................................................................... 68
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Figure 15. VEGF secretion from OMSCs ............................................................... 69
Figure 16. VEGF neutralizing antibody interfered with the migration of MSCs
towards OMSCs. ..................................................................................................... 71
Figure 17. Morphological and cytoskeletal changes in MSCs cultured in CMOMSC
with or without the addition of VEGF neutralizing antibody .................................... 72
Figure 18. CMOMSC
increased the CXCL12/CXCR4 expression of MSCs ............... 73
Figure 19. CXCL12/CXCR4 expression of MSCs stimulated by VEGF.................. 74
Figure 20. Blocking VEGF with neutralizing antibody resulted in a significant
decrease of MSCs recruitment and new bone formation ........................................... 76
Figure 21. Schematic of the proposed mechanism by which donor OMSCs initiate a
paracrine signaling cascade that results in altered phenotype and enhanced motility of
host MSCs in a skull defect model ........................................................................... 79
Figure 22. The expression of VEGFR and PDGFR mRNA transcripts was examined
by RT-qPCR analysis, using HUVECs as positive control ....................................... 90
Figure 23. CMOMSC
-induced tyrosine phosphorylation of PDGFR-α and PDGFR-β in
MSC. ....................................................................................................................... 92
Figure 24. The levels of protein expression of PDGFR-α and PDGFR-β in MSCs in
response to different stimulations ............................................................................ 93
Figure 25. The involvement of PDGFR in MSCs migration induced by CMOMSC
.... 94
Figure 26. Akt and Erk1/2 phosphorylation status in MSCs in response to various
stimulations ............................................................................................................. 96
Figure 27. Activation of the PI3K/Akt signaling is required for CMOMSC
induced
MSCs migration ...................................................................................................... 97
Figure 28 The phosphorylation status of downstream effectors of the PI3K/Akt
signaling pathway. ................................................................................................... 98
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LIST OF ABBREVIATIONS
ALP alkaline phosphatase
BM bone marrow
BMP bone morphogenetic protein
BMSC bone marrow stromal cell
BSA bovine serum albumin
BSP bone sialoprotein
COLI Type-I collagen
CXCL12 chemokine (C-X-C motif) ligand 12
CXCR4 chemokine (C-X-C motif) receptor 4
DAB 3, 3'-diaminobenzidine
DMEM Dulbecco's Modified Eagle Medium
DNA deoxyribonucleic acid
EC endothelial cell
ECM extracellular matrix
EDTA ethylenediaminetetraacetic acid
EGM endothelial cell growth media
EPC endothelial progenitor cell
ESC embryonic stem cell
FACS fluorescence activated cell sorting
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FBS fetal bovine serum
HRP horseradish peroxidase
HUVEC human umbilical vein endothelial cell
IHC immunohistochemistry
ISH in situ hybridisation
kDa kilodalton
MSC mesenchymal stromal cell
OB osteoblast
OCN osteocalcin
O(-h)MSC osteogenically differentiated (human) mesenchymal
stromal cell
OPN osteopontin
PBS phosphate buffered saline
PDGF platelet-derived growth factor
PDGFR platelet-derived growth factor receptor
PI3K phosphoinositide 3-kinase
P/S penicillin and streptomycin
RNA ribonucleic acid
RNase ribonuclease
RT-qPCR real time quantitative reverse transcription polymerase
chain reaction
RUNX2 Runt-related transcription factor 2
SCID severe combined immunodeficiency
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SDS-PAGE sodium dodecyl sulfate polyacrylamide gel
electrophoresis
TBS Tris-buffered saline
VEGF vascular endothelial growth factor
VEGFR vascular endothelial growth factor receptor
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LIST OF PUBLICATIONS
The following is a list of published, accepted or submitted manuscripts that have
arisen from the work in this thesis or are relevant to the work performed in this
PhD project:
1. Zhou Y, Fan W, Prasadam I, Crawford R, Xiao Y. Implantation of osteogenic
differentiated donor mesenchymal stem cells causes recruitment of host cells. Journal
of Tissue Engineering and Regenerative Medicine, 2012. doi: 10.1002/term.1619.
(IF: 3.278)
2. Wu C, Zhou Y, Fan W, Han P, Chang J, Yuen J, Zhang M, Xiao Y. Hypoxia-
mimicking mesoporous bioactive glass scaffolds with controllable cobalt ion release
for bone tissue engineering. Biomaterials, 2012. 33(7):2076-85. (IF: 7.404)
3. Zhou Y, Chakravorty N, Xiao Y, Gu W. Mesenchymal stem cells and nano-
structure surfaces. Book chapter in Methods in Molecular Biology, 2013. Springer.
Accepted.
4. Zhou Y, Crawford R, Xiao Y. VEGF induced CXCL12/CXCR4 axis in the
recruitment of mesenchymal stromal cells (MSCs) by osteogenically differentiated
MSCs (OMSCs) during osteogenesis. Manuscript submitted.
5. Zhou Y, Crawford R, Xiao Y. Involvement of VEGF/PDGFR and PI3K/Akt
signaling pathway in OMSCs induced MSCs migration. Manuscript submitted.
The following is a list of publications that are not relevant to the work performed
in this PhD project but published during the PhD candidature:
1. Wu C, Zhou Y (co-first author), Xu M, Han P, Chen L, Chang J, Xiao Y.
Copper-containing mesoporous bioactive glass scaffolds with multifunctional
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properties of angiogenesis capacity, osteostimulation and antibacterial activity.
Biomaterials, 2012. 34(2):422-33. (IF: 7.404)
2. Wu C, Zhou Y (co-first author), Lin C, Chang J, Xiao Y. Strontium-containing
mesoporous bioactive glass scaffolds with improved osteogenic/cementogenic
differentiation of periodontal ligament cells for periodontal tissue engineering. Acta
Biomaterialia, 2012. 8(10):3805-15. (IF: 4.865)
3. Zhou Y, Wu C, Xiao Y. The stimulation of proliferation and differentiation of
periodontal ligament cells by the ionic products from Ca7Si2P2O16 bioceramics. Acta
Biomaterialia, 2012. 8(6):2307-2316. (IF: 4.865)
4. Wu C, Fan W, Zhou Y, Luo Y, Gelinsky M, Chang J, Xiao Y. 3D-Printing of
highly uniform β-CaSiO3 ceramic scaffolds: preparation, characterization and in vivo
osteogenesis. Journal of Materials Chemistry, 2012. 22:12288-12295. (IF: 5.099)
5. Fan W, Wu C, Han P, Zhou Y, Xiao Y. Porous Ca-Si-based nanospheres: a
potential intra-canal disinfectant-carrier for infected canal treatment. Materials
Letters, 2012. 81:16-19. (IF: 2.307)
6. Wu C, Zhang Y, Zhou Y, Fan W, Xiao Y. A comparative study of mesoporous
glass/silk and non-mesoporous glass/silk scaffolds: Physiochemistry and in vivo
osteogenesis. Acta Biomaterialia, 2011. 7(5):2229-2236. (IF: 4.865)
7. Zhang Y, Fan W, Nothdurft L, Wu C, Zhou Y, Crawford R, Xiao Y. In vitro
and in vivo evaluation of adenovirus combined silk fibroin scaffolds for bone
morphogenetic protein-7 gene delivery. Tissue Engineering Part C: Methods, 2011.
17(8):789-797. (IF: 4.636)
8. Wu C, Zhang Y, Fan W, Ke X, Hu X, Zhou Y, Xiao Y. CaSiO3 microstructure
modulating the in vitro and in vivo bioactivity of poly (lactide-co-glycolide)
microspheres. Journal of Biomedical Materials Research: Part A, 2011. 98(1):122-
131. (IF: 3.044)
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LIST OF CONFERENCE PRESENTATIONS
The following is a list of conference presentations during the PhD candidature:
1. Zhou Y, Fan W, Prasadam I, Han P, Zhang X, Crawford R, Xiao Y.
Recruitment of host mesenchymal stromal cells by osteogenic differentiated donor
osteoblasts via VEGF regulated pathway. IHBI Inspires Postgraduate Student
Conference. 22nd
-23rd
November 2012, Gold Coast, Australia. Oral presentation.
2. Zhou Y, Fan W, Prasadam I, Crawford R, Xiao Y. VEGF induced
CXCL12/CXCR4 axis in the recruitment of mesenchymal stem cells (MSCs) by
osteogenic differentiated bone marrow stromal cells (O-BMSCs). 3rd
Tissue
Engineering & Regenerative Medicine International Society (TERMIS) World
Congress 2012. 5th-8
th September 2012, Vienna, Austria. Oral presentation.
3. Zhou Y, Crawford R, Xiao Y. The migration of host cells towards osteogenic
differentiated donor mesenchymal stromal cells in ectopic and orthotopic
osteogenesis. 1st Asia-Pacific Bone and Mineral Research Meeting with the
Australian & New Zealand Bone & Mineral Society (ANZBMS) 22nd
Annual
Scientific Meeting. 2nd
-5th
September, 2012, Perth, Australia. Poster presentation.
4. Zhou Y, Crawford R, Xiao Y. Implantation of osteogenic differentiated donor
mesenchymal stromal cells causes recruitment of host cells. Australian & New
Zealand Orthopaedic Research Society (ANZORS) 18th
Annual Scientific
Meeting. 30th
August-1st September 2012, Perth, Australia. Oral presentation.
5. Zhou Y, Fan W, Crawford R, Xiao Y. Hypoxia maintains stemness of BMSCs,
PDLCs and DPCs. IHBI Inspires Postgraduate Student Conference. 24th-25
th
November 2011, Brisbane, Australia. Oral presentation.
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6. Zhou Y, Fan W, Crawford R, Xiao Y. Stem cell reprogramming genes are
differentially expressed in BMSCs, PDLCs and DPCs in response to hypoxic
environment. Australian & New Zealand Orthopaedic Research Society
(ANZORS) 17th
Annual Scientific Meeting. 1st-2
nd September 2011, Brisbane,
Australia. Oral presentation.
7. Zhou Y, Fan W, Zhang Y, Crawford R, Xiao Y. Localisation of human bone
marrow-derived osteoblasts in both ectopic and orthotopic osteogenic process. IHBI
Inspires Postgraduate Student Conference. 25th-26
th November 2010, Gold Coast,
Australia. Poster presentation.
8. Zhang Y, Fan W, Nothdurft L, Wu C, Zhou Y, Crawford R, Xiao Y. In vitro
and in vivo evaluation of adenovirus combined silk fibroin scaffolds for bone
morphogenetic protein-7 gene delivery. Tissue Engineering & Regenerative
Medicine International Society (TERMIS) 2010 Asia Pacific Meeting. 15th-17
th
September 2010, Sydney, Australia. Poster presentation.
9. Zhou Y, Fan W, Zhang Y, Crawford R, Xiao Y. Localisation of human bone
marrow-derived osteoblasts in both ectopic and orthotopic osteogenic process. Tissue
Engineering & Regenerative Medicine International Society (TERMIS) 2010
Asia Pacific Meeting. 15th-17
th September 2010, Sydney, Australia. Poster
presentation.
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ACKNOWLEDGEMENTS
At the end of my PhD candidature, I would like to extend my appreciation and
gratitude to the following individuals, for without them, this thesis would simply be
impossible.
First and foremost, I would like to express my sincere gratitude to my principal
supervisor, Professor Yin Xiao, for his invaluable guidance, innovative ideas and
continuous support throughout my PhD journey. I really cherish and appreciate his
motivation, enthusiasm and his willingness to extend support not only in my study but
also in my life. I am grateful to my associate supervisor, Professor Ross Crawford, for
his expertise and stimulating clinical skills. His optimism and encouragement has
inspired me to tackle the challenges during the candidature. I consider myself
extremely fortunate to have been granted such a great opportunity to work with both
of them.
My appreciation also goes to all the former and current colleagues in the Bone Group,
especially Dr Chengtie Wu, Dr Wei Fan, Dr Indira Prasadam, Ms Wei Shi, Ms
Pingping Han and Ms Xufang Zhang. Learning and working with them has always
been so enjoyable. The friendship established during my PhD journey has been very
important to me.
I would also like to show my gratitude to Dr Michael Doran, Dr Travis Klein, Dr
Ying Dong, Dr Roland Steck, Dr Siamak Saifzadeh, and other staff at the Institute of
Health and Biomedical Innovation and the Medical Engineering Research Facility.
Thanks for the great help and precious suggestions to my study.
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A special note of appreciation goes to IHBI supporting staff, especially Mr David
Smith, Mr Dod Roshanbin, Mr Robert Smeaton, and Mr Scott Tucker, who keep our
lab running smoothly and well-organized.
I would also like to take this opportunity to thank my wonderful friends, especially Dr
Xuan Hu and Ms Ellen Fong. Their valued friendship and support has been a
motivation in completion of this project.
A heartfelt gratitude goes to Matthew Tuch for his generous patience and
encouragement in these important years of my life. Thanks for believing in me and
being there for me with a heart warming smile.
At last, I wish to dedicate my thesis to my parents for their forever and unconditional
care and love. They have worked hard and spared no effort to provide the best for me
to thrive.
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CHAPTER 1 INTRODUCTION
1.1 Preface
Orthopaedic surgeons have long held the aspiration to heal bone defects resulting
from traumatic insult, oncological resection, congenital deformities, and progressive
degenerative diseases. To realize this ambition, innumerable studies have been
conducted to advance autogeneic, allogeneic and even xenogeneic bone graft
therapies [1-3]. Although some favourable results have been achieved in animal
studies or clinical trials [4-6], the outcomes are far from satisfactory due to limited
availability, donor site morbidity, insufficient integration and immunological
repulsive reactions in the recipient against allo- or xenogeneic bone grafts [7].
Advances in materials science, cell and molecular biology, especially in the
knowledge of the underlying mechanisms of bone healing have thrown a new light on
the repair and regeneration of bone defects [8].
Cell-based therapy for bone regeneration offers a paradigm shift that may provide
alternative solutions from the traditional invasive surgeries. Significant efforts have
been devoted to characterizing the potential cell sources that are relatively abundant
and easily accessible. However, there has not been a universal understanding as to
which expansion conditions are optimal for the manufacture of MSCs intended for
bone repair and regeneration. For example, different cell seeding density [9-11],
vascular endothelial growth factor (VEGF) application [12-15] and low oxygen
tension [16-18] have all been reported to improve the longevity and the osteogenic
potency of MSCs. The induction of MSCs towards osteoblast-like cells prior to the
implantation is also found to favour more bone regeneration [19-21]. In previous
studies the concept of cell-based bone tissue engineering has been tested using
collagen type I matrices seeded with cells of osteogenic potential and implanted into
sites with osseous damage [22, 23]. New bone formation was found in defect sites
implanted with bone marrow derived-osteoblasts (OBs) at 4 weeks, while no obvious
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new bone formation was identified at the same time point without OBs transplantation
[23]. Therefore, focus has been directed on elucidating the intrinsic controls that keep
the transplanted donor cells alive and direct them along particular differentiation
pathways, which may interact with the microenvironment where the host
mesenchymal stromal cells (MSCs) reside [24]. MSCs exist in highly specialized
microenvironments in which their maintenance, proliferation, migration and
differentiation involve a complex interplay of many local and systematic signals
between MSCs and adjacent cells [25]. Even in simplified in vitro model systems, the
molecular and microenvironmental cues which are necessary to induce bone cell
differentiation have not yet been fully identified. Furthermore, concurrent studies
have suggested that an activation and chemotactic migration of host MSCs adjacent to
the defect sites is favourable for bone regeneration [26]. However, the interaction
between donor cells and host cells, and the recruitment mechanisms of host MSCs to
osseous defects are not clear and need to be further demonstrated. In addition, the
differentiation state of the donor cells, which may have significant implications for
the effectiveness of cell transplantation, has not been sufficiently elucidated.
1.2 Main purpose of this study
The main purpose of this study is to investigate the interactions between donor cells
and host cells, and to identify the possible molecular mechanisms in the initiation of
bone formation upon donor cell implantation, which is essential to developing optimal
strategies for clinical application of cell-based therapy in bone tissue engineering.
1.3 Questions to be addressed
1. Do transplanted donor cells play a role in recruiting host MSCs to the
bone defect sites to initiate osteogenesis? How do donor cells interact
with host MSCs?
2. What are the possible regulatory factors from osteogenically
differentiated MSCs (OMSCs) that lead to the recruitment of host
MSCs?
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3. What are the potential signaling pathways involved in the
communication between donor cells and host cells during osteogenesis?
1.4 Possible outcomes and significance
Based on earlier attempts to deliver osteoprogenitor cells that are capable of
responding to the local microenvironment and facilitate new bone formation in a
robust and rapid fashion, the concept and application of cell-based therapy for bone
tissue engineering has been emphasized in this project. More knowledge about the
role of pre-programmed donor cells in bone defects healing, the interaction between
donor cells and host cells during osteogenesis has been revealed. Furthermore, this
project is believed to be the first to unveil the possible molecular pathways which may
induce the host cells’ response to the signals from the implanted exogenous cells, and
activate the bone repair and regeneration process upon cell implantation. The new
findings from this project will be of significant use in facilitating the application of
pre-programmed donor cells and hopefully extend the field of clinical applications of
cell-based therapy in bone tissue engineering.
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CHAPTER 2 OVERVIEW OF PROJECT DESIGN
2.1 The effect of pre-programmed donor cells on host cell
recruitment during osteogenesis
2.1.1 In vitro cell migration
In order to investigate the effect of manipulation of donor cells on the motility of
MSCs, an in vitro migration assay was performed using transwell system. Transwell
inserts seeded with MSCs were assembled onto the companion plates, which were
seeded with osteogenically differentiated MSCs (OMSCs) or undifferentiated MSCs.
After 8 h of incubation, cell motility was quantified by counting the number of cells
that migrated through the pores in response to different stimulations from the bottom
companion plates, and the differences were compared between groups.
2.1.2 In vivo host cell recruitment
Collagen scaffolds carrying OMSCs or MSCs were implanted subcutaneously or into
skull defects in severe combined immunodeficiency (SCID) mice. After samples were
harvested and fixed, micro-computed tomography (μCT) scanning was performed to
reconstruct three-dimensional (3D) images of the implants, and to measure the
calcification volume for statistical analysis. After samples were processed and
sectioned, the ectopic and orthotopic osteogenesis was assessed by subsequent
histological and immunohistochemical (IHC) staining. In situ hybridisation (ISH)
against human Alu sequences was performed to distinguish donor human cells from
host mouse cells, and to investigate the recruitment tendency of host cells in different
microenvironments. The localisation of transplanted human cells identified by the Alu
probe in both subcutaneous and skull defect scaffolds was observed, recorded and
analysed.
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2.2 Possible regulatory factors that lead to the recruitment
of host cells
2.2.1 Comparison between OMSCs and MSCs
In order to unveil the secreted factors from donor OMSCs that may lead to the
migration of host MSCs, further study in the new bone forming area was performed.
Triple staining technique revealed that donor OMSCs strongly expressed VEGF,
which might exert a paracrine effect resulting in host cell recruitment and their
involvement in new bone formation. This finding formed the foundation for the
subsequent project development. Alizarin Red S staining was performed to compare
the mineralization, while real time quantitative-PCR (RT-qPCR) analysis, enzyme-
linked immunosorbent assay (ELISA) and Western blotting analysis were performed
to compare the VEGF secretion between OMSCs and MSCs.
2.2.2 The in vitro regulatory effect of VEGF
Indirect co-culture models were applied to study the effect of VEGF derived from
OMSCs on the motility, the cell morphology and the CXCL12/CXCR4 expression of
MSCs. In vitro migration assay and real-time xCELLigence system were performed to
investigate the motility of MSCs in response to different stimulations, and the
responding cell morphology changes were further examined by fluorescent staining.
Furthermore, the expression pattern of CXCL12/CXCR4, which is the best known
axis that contributes to cell migration, was studied extensively and the effect of VEGF
on cell responses was probed using a specific neutralizing antibody.
2.2.3 The in vivo regulatory effect of VEGF
The regulatory effect of VEGF on host cell recruitment was examined by the
application of neutralizing VEGF antibody in vivo, after OMSCs implantation into the
skull defects in SCID mice. Histological staining and µCT scanning were performed
to observe the new bone formation. The recruitment of host cells to the bone defect
sites was monitored by ISH staining.
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2.3 Potential cell signaling pathways involved in the host
cell recruitment
2.3.1 Functional receptors profile
MSCs were stimulated with different concentrations of VEGF, or conditioned
medium derived from MSCs (CMMSC
) or OMSCs (CMOMSC
) to establish the receptors’
activation profile. The functional role of platelet-derived growth factor receptors
(PDGFRs) was further investigated using phospho-receptor tyrosine kinase (RTK)
array, Western blotting analysis and in vitro migration assay.
2.3.2 The involvement of cell signaling pathways
In order to unveil the potential regulatory cell signaling pathway involved in the host
cell recruitment, MSCs were stimulated in different conditions with or without the
interference of the dimerization of PDGFRs. The involvement of PI3K/Akt and
Erk1/2 in OMSCs-induced MSCs migration was monitored by relevant
phosphorylated antibodies and inhibitors. The time course phosphorylation status of
the PI3K/Akt signaling pathway was further investigated by Western blotting analysis.
2.3.3 Downstream effectors of the PI3K/Akt signaling pathway
The stimulatory effect of VEGF secreted from OMSCs on the activation of the
PI3K/Akt signaling pathway was further investigated by detecting the
phosphorylation status of the downstream effectors, mTOR, p70S6K
, and PAK1, by
Western blotting evaluation of specific antibodies.
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CHAPTER 3 LITERATURE REVIEW
3.1 Overview
Tissue engineering is an emerging interdisciplinary field involving principles of the
life sciences and engineering with the expectation of constructing a biological
substitute for damaged tissues, both in vitro and in vivo, in order to restore, maintain,
or improve normal tissue structure and function [27]. Rapid expansion of knowledge
regarding the potential for tissue engineering, and more specifically progenitor cell-
based tissue engineering, offers new insights to the treatment of an array of diseases,
such as graft-versus-host disease (GVHD) [28], cardiovascular failure [29, 30], liver
diseases [31], injury or degeneration of the spinal cord [32, 33], tendon [34], cartilage
[35], bone [36] and numerous others. Among the aforementioned conditions and
diseases, bone defects resulting from trauma or pathological and physiological
resorption represent a significant global health issue that deserves optimal treatment.
Although many of the conventional modalities for repairing bone defects are able to
restore certain levels of function, various drawbacks ranging from limited donor tissue
to procedure-related complications remain a major clinical challenge [7, 37]. Unlike
traditional surgeries that need to sacrifice tissues from the donor site, the tissue
engineering approach can achieve the goal of bone repair and regeneration without the
necessity of donor site morbidity [38]. Recent studies have demonstrated the
feasibility of generating tissue-engineered bone for repairing bone defects [39-41],
and the outcomes of ongoing clinical trials are very promising [36, 42, 43].
The concept of cell-based tissue engineering has fuelled investigations of various
progenitor cell sources that can be manipulated, either in vitro or in vivo, to repair or
replace damaged tissues. Given the complex three-dimensional structure of bone
tissues, cells, extracellular matrix, cell-cell communications, cell-matrix interactions,
growth factors, cytokines and other cell components have to be combined in a well-
coordinated space- and time-dependent manner to achieve successful results in bone
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tissue engineering. In the past few years, there have been significant advances in the
understanding of the molecular mechanisms of cell fate determination, an extremely
complex process regulated by multi-mutual interplay of signaling pathways [44-47].
Therefore, cell-based therapies with potential to stimulate endogenous progenitor cells
during osteogenesis have become a prime translational goal.
This review focuses on the current status of knowledge as well as the emerging
challenges in the area of cell-based therapy and potential cell sources related to bone
tissue engineering. Additionally, the critical role of the transplanted donor cells, the
regulatory factors, and the cell signaling pathways leading to the recruitment of host
cells, have been thoroughly reviewed.
3.2 Cell-based therapy for bone defects
The treatments for a multitude of skeletal deficits arising from tumors, infections,
trauma, biochemical disorders, and abnormal skeletal development have been
conducted clinically through the development of surgical techniques for autologous
bone graft, allogeneic tissue transplantation and the implementation of biocompatible
materials [48]. However, each of these reconstructive options has its disadvantages.
The limited availability of autologous bone graft that can be harvested, and
considerable morbidity at the donor site remains inherent limitations of autologous
transplantation [49]. Conversely, although the use of allograft or xenograft avoids
impairments to the donor site, it is sometimes associated with the risks of infection,
and unfavorable immune response from the host [50, 51]. In addition, certain bone-
substitute materials can lead to poor integration, adverse reactions and eventual bone
resorption [52, 53]. Recently, encouraging results from in vitro [54] and in vivo [55,
56] studies, as well as early human clinical applications [57, 58], have aroused intense
interest in cell-based therapy as an alternative to the traditional techniques for bone
defect repair and regeneration.
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Figure 1. Schematic of cell-based therapy strategy. The cell transplantation/delivery strategy
involves tissue biopsies, ex vivo expansion of cells from the biopsies, and subsequent transplantation of
the cells into the recipient. Adapted from [59].
The traditional concept of cell-based therapy involves the harvest of suitable cell
sources from autologous/allogenic donors, ex vivo cultivation and expansion to a large
population (Figure 1). These cells are either transplanted back into to the patient by
injection, or seeded onto a three-dimensional scaffold and then implanted into the
recipient [60, 61]. Alternatively, the cell-seeded scaffold can be incubated in a
bioreactor, where the cells are exposed to certain biological, chemical or physical
stimuli that promote the formation of the appropriate tissue prior to the implantation
[62, 63]. The rationale for cell-based therapy to induce bone regeneration is based on
the high osteogenic potency of osteoinductive cells, which may actively induce
adjacent progenitor cells to form new bone tissue [64]. Recent investigations probing
progenitor cell populations have produced a large amount of evidence to support the
clinical viability of cell-based bone tissue engineering [25]. Most of the studies have
focused on the selection of cell sources that are safe, less costly and more easily
available. However, a key unresolved question is whether these cells themselves
participate in the regenerative process mainly by differentiating into osteoblasts or
facilitating tissue repair through the production of various trophic factors (cytokines)
that may modulate the host tissue microenvironment. This may facilitate the
differentiation of resident stem cells and lead to the cell recruitment through blood
vessels, bringing even more of the host cells to the bone defect sites. Recent research
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has also revealed that donor cell number [65], nutrition and matrix acidity in the
recipient site [66, 67] can also have different effects on the regenerative process.
3.3 Cell sources for cell-based bone tissue engineering
3.3.1 Mesenchymal stromal cell (MSC)
Regardless of their original sources, stem cells share two characteristic properties.
Firstly, they have the capacity for prolonged or unlimited self-renewal under certain
conditions, and secondly they retain the potential to differentiate into a variety of
more specialized cell types [68]. Undoubtedly, human embryonic stem cells (hESCs)
are able to differentiate down an osteogenic lineage in an efficient manner and
maintain an adequate proliferative capacity post-harvest [69]. However, the political
and ethical issues associated with hESCs have cast a shadow of doubt over their
realistic potential for implementation in tissue engineering strategies [70]. Thus,
MSCs have emerged as a promising cell source in the ongoing research of bone tissue
engineering [71]. Numerous studies have demonstrated that MSCs derived from
various origins, including skin, hair follicle, periosteum, bone marrow, adipose tissue,
and umbilical cord blood, are able to differentiate into osteoprogenitors and
osteoblasts in vitro and in vivo [72-77]. The osteogenic capacity of MSCs can be
further enhanced by the systemic or local administration of bone morphogenic
proteins (BMPs) or other osteoinductive biomolecules, either in slow-release systems
or viral/non-viral based gene delivery systems [78-82]. The addition of supporting
biomolecules has gained increasing popularity especially when bone repair in older,
osteoporotic or diabetic individuals is considered, because the differentiation capacity
of autologous MSCs in these conditions is known to be compromised [83-85].
Although the initial enthusiasm surrounding MSC-based bone tissue engineering is
still growing enormously, research towards the goal of defining a cell surface antigen
profile unique to MSCs has only achieved marginal success. According to various
research, MSCs express positive for a long list of cell surface markers, such as CD29,
CD44, CD73, CD90 (Thy-1), CD105, CD117, CD133, CD166 (activated leukocyte-
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cell adhesion molecule, ALCAM), and CD186 [86, 87]. Other markers including Sca-
1 (stem cell antigen-1), SCF R/c-kit (stem cell factor), SH2, SH3, SH4, STRO-1, and
HLA Class I have also been identified [88-90]. Furthermore, it is generally agreed
that the immunodepletion of the hematopoietic cells using anti-CD11b, CD34 and
CD45, forms the fundamental concept for negative selection method of MSCs
purification [91, 92].
In addition, the systemic analysis of cell surface molecules of MSCs has revealed that
MSCs express receptors for numerous extracellular matrix (ECM) proteins including
collagen (α1β1-, α2β1-integrin), laminin (α6β1-, α6β4-integrin), fibronectin (α3β1-,
α5β1-integrin) and vitronectin (αvβ1-, αvβ3-integrin) [93-96]. These specific
expression patterns of adhesion molecules suggest the potential interactions between
MSCs and other cell types in vivo [97-99]. Owing to the complexity of the stem cell
niche, the study of dynamic interactions between stem cells and differentiated
neighbouring cells in their physiological microenvironment has only recently been
explored [100-103]. Depending on the ECM and signal molecules in the
microenvironment, MSCs can develop into adipocytes, osteoblasts, chondrocytes,
myoblasts or other phenotypes [104-107]. However, no single marker has yet been
identified that definitively delineates MSCs in vivo and hence there is a lack of
thorough understanding of the mechanisms underlying stem cell renewal and its
functional differentiation characteristics.
3.3.2 Bone marrow stromal cell (BMSC)
The first MSC population to gain significant attention from researchers and provide
the largest contribution to the current knowledge regarding the osteogenic potential of
MSCs is derived from the bone marrow cavity, which is in tight contact with the
hematopoietic compartment [108]. Bone marrow stromal cells (BMSCs) have been
identified as a population of organized hierarchical postnatal stem cells with the
potential to undergo osteogenic, as well as chondrogenic and adipogenic
differentiation when exposed to appropriate microenvironmental cues [76].
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Because of their multipotent nature, BMSCs have become one of the most important
cell sources for cell-based therapy and tissue regeneration [109]. Their osteogenic
differentiation potential has been well characterised in many in vitro studies [110-
112]. In addition, animal model-based studies also reveal that BMSCs have
significant capability for healing bone defects of critical size in vivo when implanted
with proper biomaterials [113, 114], indicating their great potential for bone tissue
engineering applications. In terms of human studies, the use of in vitro expanded
autologous BMSCs in conjunction with porous hydroxyapatite ceramic scaffolds has
been reported in the treatment of large bone defects [115]. Progressive integration of
the implants with the native bone was observed as well as new bone formation inside
the bioceramic pores and vascular ingrowth. A favorable integration of the implants
with the pre-existing bone was maintained during the long-term follow-up period and
no major adverse reactions were found, illustrating the durability achieved by the cell-
based bone tissue engineering approach [116].
Osteogenic induction may enhance the probability of in vivo bone formation and
contribute to the reliability of bone tissue engineering. Several recent investigations
demonstrated that in vitro osteogenic induction of BMSCs before implantation could
significantly promote subsequent ectopic bone formation compared to
undifferentiated BMSCs [19]. Therefore, it may be feasible to obtain a relatively
small amount of autologous BMSCs from patients, expand and pre-differentiate them
into an appropriate number of osteoblastic cells and implant them back to the donors
to facilitate the repair and regeneration of bone defects. Recent research has also
addressed the importance of a proper microenvironment for BMSC transplantation
[117]. It is hypothesized that injured tissues produce the appropriate signals necessary
for cell transplantation and that the local microenvironment may eventually induce
particular phenotypes and functions of the transplanted cells [118-120]. However, the
specific microenvironment (i.e., the unique combination of matrix, growth factors,
and cell adhesion cues), which is critically important for the control of progenitor cell
maintenance, proliferation, and differentiation, and is required to determine the final
fate of the transplanted BMSCs in vivo, remains unknown.
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3.3.3 Osteoblast (OB)
Understanding bone growth regulation, the continued remodeling of bone throughout
life, and the regeneration of injured tissues are the basic biological concerns of
clinicians for the treatment of skeletal diseases. The major event that triggers
osteogenesis is the transition of MSCs into bone forming osteoblasts (OBs). OBs are
mononuclear, not terminally differentiated cells that are responsible for the synthesis,
deposition and mineralization of ECM. Therefore, they play a pivotal role in creating
and maintaining skeletal architecture. MSCs need to undergo several transitional steps
before becoming mature OBs. Each transition requires complex activation or
suppression of critical molecular elements for the commitment and differentiation of
OBs to occur (Figure 2).
Figure 2. Schematic of osteoblast differentiation. Adapted from [121].
Over the last decade, molecular and genetic studies have broadened our understanding
of OB differentiation. Ducy and Karsenty conducted the initial study on the regulation
of osteocalcin, which is the most specific gene of OBs [122]. The earliest
osteoprogenitor cells express Runt-related transcription factor 2 (RUNX2), a bone
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specific transcription factor, which is required for the early stages of OB
differentiation [123]. Osteoblastogenesis consists of three major stages, proliferation,
matrix maturation, and mineralization, which are characterised by sequentially
expressed distinctive markers from OBs. Alkaline phosphatase (ALP), type I collagen
(COLI), osteocalcin (OCN), osteopontin (OPN) and PTH-related protein (PTHrP)
receptor are the most frequently used markers during osteoblastogenesis [124, 125].
Generally, ALP and COLI are early markers of OBs’ differentiation, while OCN and
PTHrP receptors appear late. OPN reaches its peak during proliferation and in the
later stages of differentiation.
Following the initial lineage commitment, there is a phase of expansion. The initial
cell division is asymmetric, giving rise to another stem cell and a committed
osteoprogenitor. The pre-osteoblasts are at an intermediate stage. They still proliferate
and then develop into OBs that synthesize bone matrix. The mature OBs lie adjacent
to the newly formed osteoid and express ALP, OPN, BSP, and OCN. The cumulative
effect of the recruitment of stem cells and their expansion, and the functional capacity
of mature OBs, are measured by rates of bone formation in vivo. There are three fates
upon completion of the synthetic phase in the terminal stage of the remodeling cycle.
OBs may become osteocytes upon entrapment within the mineralized matrix, evolve
into inactive lining cells that protect the bone matrix from osteoclasts, or undergo
apoptosis [126].
Immature OBs derived from MSCs may provide more consistency in forming bone
than precursor populations with multipotentiality in terms of cell-based therapy for
bone defect healing [127, 128]. However, the isolation procedure of OBs is
considerably labour-intensive and time consuming. In addition, only a relatively small
number of cells are available after the dissociation of the tissue and their expansion
rates are comparatively low. Consequently, the generation of clinically adequate
numbers of OBs for cell-based therapy is a significant challenge.
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3.4 The contribution of donor cells in cell-based bone tissue
engineering
Although the outcome of cell-based therapy using the aforementioned cell sources has
been encouraging, the precise mechanisms by which the donor cells enhance tissue
repair and regeneration are not clearly understood. In most cases, the intent of cell-
based therapy is to repair damaged tissues with donor cells capable of carrying out
structural and functional recovery [128]. However, the extent of donor cell
contribution to the cells/tissues of interest has not been fully elucidated. It becomes
apparent that in many situations, the donor cells improve tissue regeneration at the site
of damage without significant engraftment or differentiation [129]. Studies in which
MSCs were transplanted into children with severe forms of osteogenesis imperfecta
(OI) revealed the level of donor cell engraftment was extremely low and could not
justify the growth acceleration and the bone fracture rate reduction [130-132]. In other
related studies, transplantation of BMSCs into animal models of myocardial infarction
(MI) led to significantly improved cardiac function due to the secreted factors from
donor cells, rather than direct engraftment or differentiation [133, 134]. These
findings lead to the speculation that besides direct cell-to-cell contact, the donor cells
may secrete proinflammatory cytokines (such as IL-1 and IL-6), mitogens (such as
IGF, FGF and PDGF), morphogens (BMPs), and angiogenic factors (VEGF and
angiopoietins) [135, 136], which exert paracrine activities and modulate the
microenvironment for neighboring stem cells to carry out the reparative process [137-
140]. In addition, the broad spectrum of bioactive molecules that are produced by the
donor cells may lead to the tropism of endogenous cells from their original niches.
In the field of cell-based bone tissue engineering, a recent study from Cancedda’s
team has shed some light on the commitment of donor cells by using a murine model
of ectopic bone formation. This particular study demonstrated that the origin of newly
formed bone was dependent on the maturation status of the implanted cells [141]. An
intramembranous ossification directly performed by the donor cells was observed
when mature OBs were seeded onto a porous ceramic. In contrast, the implantation of
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undifferentiated MSCs led to the formation of new bone of host origin via
endochondral ossification. Researchers have also shown that MSCs are able to
accelerate the wound healing process by stimulating cell differentiation [142], and
releasing pro-angiogenic factors that promote new blood vessel formation [143]. The
newly formed vasculature is not only critical for the transportation of oxygen and
nutrients to the centre of the implants, but also fundamental to the recruitment of host
cells through the connection with the host circulating system. This creates a
communicative network between the recipient organism and the engineered graft. In
addition, the implanted cell/scaffold constructs may create a transient inflammatory
microenvironment, in which an innate response is mediated by a mixed cell
population, mainly composed of the recruited monocytes, macrophages, and
lymphocytes. The released factors from the inflammatory cells in turn act as signals
and impact on the way donor cells contribute to the tissue repair and regeneration
process in vivo [144-146]. In this respect, although the molecular mechanisms are yet
poorly understood, it is reasonable to hypothesize that donor cells can act indirectly as
“signaling centres”, orchestrating the host response to the skeletal defects, in addition
to their direct role as osteoprogenitors and osteoblasts.
3.5 The involvement of host cells in cell-based bone tissue
engineering
There is no doubt that cell-based therapy is becoming one of the most promising
strategies to repair irreversibly-affected tissues. However, the therapeutic efficacy
may not only depend on the involvement of donor cells, but also the activation and
homing of endogenous cells from their original niches through the circulation to the
injured tissues. Within a complex niche, endogenous cells are exposed to a variety of
spatially and temporally controlled biochemical mixtures of insoluble transmembrane
receptor ligands, proteases, adhesion molecules (such as selectins and integrins),
ECM molecules, as well as soluble chemokines, cytokines and growth factors [147,
148]. In the field of stem cell biology, the term “homing” is often defined as the
recruitment of endogenous cells to the damaged tissues or their navigation to other
targeted anatomic destinations following mobilization [149], often over great
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distances via the bloodstream. In addition, a highly proliferative population of
progenitor cells residing in the neighboring healthy tissues may also be recruited to
the defect sites [59] (Figure 3).
Figure 3. Schematic of endogenous cells homing to the damaged tissues for in situ
tissue regeneration. The homing of endogenous cells involves actively recruiting host stem cells
from either the blood or a tissue specific niche to the site of injury. Adapted from [59].
Host cell mobilization is not only essential in a wide range of physiological and
pathological phenomena, but also important to biotechnological applications such as
regenerative medicine and tissue engineering. Previous work has demonstrated that
endogenous biological resources and environmental conditions can be used for in situ
tissue regeneration, suggesting the possibility to reduce the intensive labour for donor
Exit tissue
niche to
enter blood
18 | P a g e
cell procurement and manipulation [150]. Applications of in situ tissue regeneration
are primarily based on the local release of bioactive molecules, such as cytokines and
growth factors, to enhance the mobilization of host cells and control the fate of the
recruited endogenous cells for efficient tissue regeneration. Mao and colleagues have
successfully fabricated a TGFβ3-infused scaffold which could induce homing of
endogenous cells to regenerate the entire articular surface of the rabbit synovial joint
[151]. A more recent study instead emphasized the efficiency of combined systemic
and local delivery of stem cell recruiting factors for tissue engineering, showing that
the combined system could significantly enhance the recruitment of host cells which
were capable of differentiating into multiple lineages [152].
Likewise, in cell-based bone tissue engineering, a continuous source of cell
recruitment to the bone defect site is ensured by the endogenous cells. The research
by Tasso and coworkers showed that two consecutive waves of host cell migration
were induced by the donor cells seeded on the porous ceramic scaffolds [153]. The
first wave, observed 7 days after the implantation, was enriched in CD31+ endothelial
cells, whereas the second wave, observed at day 11, was enriched in CD146+
perivascular cells, potentially responsible for the newly formed bone. Another study
has also demonstrated that osteoblastic precursors can be systemically recruited from
remote bone marrow cavities via the bloodstream and become involved in fracture
healing [154].
The full comprehension of the host response to implanted donor cells is of great
importance to the development of novel clinical strategies where host cells could
contribute to regenerate the appropriate tissue. What remains unclear is the
mechanism of how the endogenous cells are recruited to the damaged
tissue/transplantation site, and what potential role the recruited host cells play in the
process of tissue repair and regeneration. Therefore, it is necessary to further
understand the phenomenon of host/donor cells interaction, which may benefit the
optimization of treatment alternatives for nonunion fractures and other pathologic
skeletal deficits.
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3.6 Chemokines and growth factors related with cell
migration
Cell migration is a fundamental component of embryogenesis, vasculogenesis, wound
healing, immune surveillance, as well as tumor cell metastasis. It is a mechanically
integrated molecular process regulated by a series of coordinated mechanisms [155].
Cells become asymmetrical and polarized during migration, with front and back ends
formed to direct movement [156]. Broad lamellipodia and spike-like filopodia take
place primarily around the cell front, whereas, stress fibers, which are involved in cell
adhesion, are formed or deformed at the back [157]. These multi-step processes
involve morphological changes that result from the rearrangement of actin
cytoskeleton, the emergence of protrusive membrane structures followed by the
contraction of the cell body, the detachment of the uropod and the secretion of matrix
degrading enzymes [157, 158]. A multitude of chemokines, cytokines and growth
factors play critical roles in regulating the mobilization and homing of MSCs for
effective regeneration of functional tissue.
3.6.1 CXCL12
Chemokines (Chemotactic cytokines) are a family of small (8-10 kDa) proteins able to
chemically direct responsive cells, such as lymphocytes, neutrophils, hematopoietic
cells and other cell types to the targeted tissue compartment [159]. In addition,
chemokines are thought to play an important role in cell activation, differentiation and
survival [160-162]. Members of the chemokines are divided into four subfamilies,
namely CXC, CC, C and CX3C chemokines, depending on the spacing of their first
two cysteine residues [159, 163]. Of these, CXC and CC chemokines are the two
major groups.
Chemokine (C-X-C motif) ligand 12 (CXCL12) is the most predominant stem cell
homing factor to date [164]. It was first identified as stromal cell-derived factor-1
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(SDF-1) in the supernatant of BMSCs [165]. CXCL12 is strongly expressed in bone
marrow and produced by osteoblasts, endothelial cells and reticular cells that are
scattered through the bone marrow stroma [166]. CXCL12 is highly conservative and
has 99% homology between mouse and human, allowing CXCL12 to act across
species barriers. Upon stimulation with CXCL12, MSCs are able to rearrange the
actin cytoskeleton and increase phosphorylation of the focal adhesion adapter protein
paxillin, which is required for focal adhesion turnover and ultimately migration [167].
The critical role for CXCL12 in the homing of endogenous cells to damaged sites has
been unraveled by studies in animal models of liver, limb, and heart damage [168-
171]. One of the most important functions of CXCL12 is related to tissue repair and
regeneration. Recent investigations have shown the repair of ischemic injuries
involves the selective recruitment of circulating or resident progenitor cells. Hypoxia-
inducible factor-1 (HIF-1), a central regulator of tissue hypoxia, induces CXCL12
expression in ischemic areas in direct proportion to reduced oxygen tension in vivo
[172, 173]. The elevated CXCL12 then recruits the circulating progenitor cells to
repair the damaged tissues.
Chemokine (C-X-C motif) receptor 4 (CXCR4) is the best characterised receptor for
CXCL12 and an essential mediator for the chemotaxis responses induced by CXCL12
in many responsive cells [174]. Previous studies have shown the CXCR4-deficient
mice died in utero or perinatally due to multiple defects in developing hematopoietic
and nervous system [175-177], leading to the conclusion that CXCR4 is indispensible
to several physiological processes during late embryonic and early postnatal stages. A
recent study has also provided evidence that CXCR4 functions in postnatal bone
development by regulating OB development in cooperation with BMPs signaling,
indicating CXCR4 acts as an endogenous signaling component necessary for bone
formation [178]. In bone marrow, CXCL12 availability has been shown to be
regulated by the uptake of CXCL12 via CXCR4 positive cells, which then translocate
the chemokine into the bone marrow. This transporter function is characteristic of
both endothelial and stromal cells [179]. The binding of CXCL12 to CXCR4 leads to
receptor structural changes and G-protein activation, which then stimulates signaling
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cascades (e.g., PI3K and MAPKs), and induces intracellular Ca2+
release, resulting in
cell migration, proliferation and survival (Figure 4).
The interaction of CXCL12 with CXCR4 is thought to regulate trafficking of the
hematopoietic stem cells, endothelial progenitor cells (EPCs) and in prostate and
breast cancer metastasis to bone and lung [180, 181]. The pivotal role of the
CXCL12/CXCR4 axis in the mobilization and homing of stem cells to injured or
stressed tissue is further investigated by numerous researchers [182-184]. Previous
work has demonstrated that applying a neutralizing CXCR4 antibody after induction
of cerebral ischemia in a rat model compromised BMSC homing, resulting in poor
neurological recovery after stroke [185]. On the other hand, transfection or
transduction of MSCs with CXCR4 is reported to enhance cell migration toward
CXCL12 in vitro [186, 187] and toward infracted myocardium in rats [188], whereas
almost no migration is observed to normal myocardium [188]. Recent studies have
emphasized the clinical effect of CXCR4 antagonist. The bicyclam AMD3100, also
known as Plerixafor (Mozobil™), is a competitive selective CXCR4 antagonist. It
blocks the binding of CXCR4 to CXCL12 and has been demonstrated to interfere
with a number of pathophysiological processes [189], such as rheumatoid, allergic
and malignant diseases. Physiologically, bone marrow has a higher CXCL12
concentration than any other tissues, therefore, circulating MSCs home to bone
marrow proficiently. AMD3100 has been shown to mobilize CD34+ stem cells from
the bone marrow into the blood circulation and augment the migration of bone
marrow-derived EPCs into sites of neovascularization after myocardial infarction
[190]. Currently, AMD3100 is actively pursued as a stem cell mobilizer for
autologous transplantation in patients with multiple myeloma and non-Hodgkin's
lymphoma [189], and has great potential for the treatment of other hematological
malignancies and non-hematological malignancies.
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Figure 4. CXCL12/CXCR4 signaling pathway. Binding of CXCL12 to CXCR4 leads to
receptor structural changes and G-protein activation. Activated G-proteins induce multiple signaling
pathways, such as PI3K, MAPKs and NFκB, and the release of intracellular Ca2+. These signaling
pathways are associated with cell proliferation, migration and survival.
3.6.2 VEGF
As most members from the VEGF/platelet-derived growth factor (PDGF) family,
VEGF is the potent mitogen for endothelial cells (ECs), and one of the key regulators
of angiogenesis, vasculogenesis and developmental hematopoiesis. In addition to its
well-documented effects on blood vessel development and formation [191, 192],
VEGF also plays a critical role in promoting osteogenesis [193]. VEGF is of great
importance for the patterning of the bone growth plate [194, 195] by stimulating the
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differentiation of OBs [196] and the crosstalk between OBs and ECs [197]. In a
mouse femur fracture study, treatment with exogenous VEGF enhanced not only
blood vessel formation but also ossification and new bone formation, whereas VEGF
inhibition impaired callus mineralization [198]. Various cell types secrete VEGF,
including tumor cells, macrophages, OBs and osteoblast-like cells, and several growth
factors, hormones, and cytokines regulate its expression. It has been demonstrated that
VEGF secretion from osteoblastic cells is at low levels in the early stage of osteogenic
differentiation [199] and markedly increases as osteoblastogenesis proceeds [200].
This is significantly regulated by OB-specific transcription factor Osterix, as reported
in a recent study [201]. Interestingly, VEGF has been proved to be chemoattractive
for primary human osteoblasts, providing first evidence that it may also be involved in
local recruitment of osteoprogenitor cells in the course of endochondral bone
formation or remolding [202, 203].
There are five structurally related mammalian VEGF ligands, namely, VEGF-A,
VEGF-B, VEGF-C, VEGF-D and placental growth factor (PlGF) [204]. Each VEGF
ligand has several different variants due to either alternative splicing or processing.
Being the most abundant and active member of the VEGF family, VEGF-A
undergoes alternative splicing, which results in at least six isoforms (VEGF-A121,
VEGF-A145, VEGF-A165, VEGF-A183, VEGF-A189 and VEGF-A206), among which
VEGF-A165 is the most predominant and biologically potent. The VEGFs are
produced by many different cell types and typically act locally in a paracrine manner.
However, VEGF-A produced by ECs may act in an autocrine manner to stimulate
vessel survival [205].
The VEGFs bind to three structurally related VEGFR tyrosine kinases, VEGFR-1
(Flt-1), VEGFR-2 (KDR/Flk-1) and VEGFR-3 (Flt-4), each of which forms a
homodimer on ligand binding [206] (Figure 5). VEGF-A binds to VEGFR-1 and
VEGFR-2, but not VEGFR-3. The majority of VEGF signaling is mediated through
VEGFR-2 in endothelial cells [206, 207]. VEGF-A165 can also interact with
neuropilin-1 (NRP-1) and neuropilin-2 (NRP-2), which are cell surface
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transmembrane glycoprotein receptors [208]. NRPs have a short cytoplasmic domain,
thus it is speculated that they do not mediate the signal independently, but function
only as co-receptors [208]. It has been reported that NRP-1 can associate and form a
complex with VEGFR-2, leading to an enhanced VEGFR-2 signaling induced by
VEGF-A165 binding to NRP-1 [209]. One interesting finding is that VEGF-A can
stimulate MSCs proliferation and mobilization to damaged tissues by binding to and
activating PDGF receptors, even though MSCs express low basal level of VEGFRs
[210].
Figure 5. Vascular endothelial growth factor (VEGF) family members and their
receptors. The mammalian VEGF family consists of five structurally related glycoproteins, VEGF-A,
VEGF-B, VEGF-C, VEGF-D and placental growth factor (PlGF). The VEGFs bind to three
structurally related VEGFR tyrosine kinases. VEGF-A binds to both VEGFR-1 and VEGFR-2. VEGF-
B and PlGF bind exclusively to VEGFR-1. VEGFR-3 is a specific receptor for VEGF-C and VEGF-D.
VEGF-C and VEGF-D can be proteolytically processed to allow binding to VEGFR-2 as well. In
addition, neuropilin (NRP)-1 and NRP-2 act as co-receptors for specific isoforms of VEGF family
members, and increase the binding affinity of these ligands to their respective receptors. Adapted from
[211].
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3.6.3 PDGF
PDGFs are the primary mitogens for the cells of mesenchymal and neuroectodermal
origin and are members of the evolutionarily conserved, structurally and functionally
related VEGF/PDGF family [212]. First identified in 1970s as a serum growth factor
that stimulates proliferation of fibroblasts, smooth muscle cells (SMCs) and glial cells
[213-215], the PDGF family is now one of the best characterised growth factor
systems.
The PDGF family consists of four different isoforms, the traditional PDGF-A and
PDGF-B, and more recently discovered PDGF-C and PDGF-D [216]. PDGF-A and
PDGF-B are processed intracellularly and are secreted in their active forms, while
PDGF-C and PDGF-D are secreted as latent factors and require extracellular cleavage
of the CUB domain. This is carried out by the tissue-type plasminogen activator (tPA)
for PDGF-C, and the urokinase-type plasminogen activator (uPA) or matriptase for
PDGF-D [217-220]. The PDGF ligands exist as disulphide-bonded polypeptide chains,
including the four homodimers PDGF-AA, PDGF-BB, PDGF-CC and PDGF-DD,
and one heterodimer, PDGF-AB [221]. The ligands exert their biological activities via
two distinct but structurally related, membrane bound receptor tyrosine kinases
(RTKs), PDGF receptor-α (PDGFR-α) and PDGF receptor-β (PDGFR-β). Particularly
strong expression of PDGFR-α has been noticed in subtypes of mesenchymal
progenitors in lung, skin and intestine, as well as in oligodendrocyte progenitors,
while PDGFR-β is expressed in the mesenchyme, especially in vascular SMCs
(vSMCs) and pericytes [222]. Ball et al. also reported that MSCs in vitro have a high
cell surface ratio of PDGFR-α/PDGFR-β [223], and PDGFR-α appears to be a
characteristic of undifferentiated MSCs. The PDGFRs have common domain
structures, including five extracellular immunoglobulin (Ig) loops and a split
intracellular tyrosine kinase domain [222]. The binding of ligand to the receptor
triggers receptor homodimerization or heterodimerization, resulting in
autophosphorylation of specific tyrosine residues within the intracellular domains and
kinase activation. Once activated, intracellular mediators dock to phosphotyrosine
residues in the receptor, leading to the activation of downstream signaling pathways,
e.g. PI3K, MAPK and PLC-γ. The ability of each of the five dimeric PDGF ligands to
26 | P a g e
bind and activate the PDGF receptors is quite specific as summarized in Figure 6.
PDGF-AA can only interact with PDGFR-α, whereas PDGF-BB has a greater binding
affinity for PDGFR-β. However, PDGFR-β is also capable of binding PDGFR-α, and
thus can induce all three receptor homodimer and heterodimer. PDGF-AB and PDGF-
CC can both bind to PDGFR-α, and initiate the formation of homodimer and
heterodimer. PDGF-DD binds predominantly to PDGFR-β and forms a homodimer
PDGFR-ββ, but also may induce lower levels of the heterodimeric PDGFR-αβ [222,
224].
Figure 6. Simplified schematic of the platelet-derived growth factor (PDGF) system.
PDGF-A and-B are secreted as active homo- or heterodimers. PDGF-C and –D are secreted as
homodimers and latent factors which need extracellular cleavage of the CUB domain by tPA (PDGF-
C), uPA (PDGF-D) or matriptase (PDGF-D) for receptor binding and activation. PDGF-AA is the most
selective member of the PDGF family and activates PDGFR-αα exclusively. PDGF-BB is the universal
ligand. PDGF-AB and PDGF-CC assemble and activate PDGFR-αα and PDGFR-αβ. PDGF-DD
activates PDGFR-ββ, and under certain circumstances, PDGFR-αβ. PDGF binding results in
autophosphorylation and activation of different signaling pathways, such as JAK/STAT, PI3K, PLC-γ
or MAPK. Downstream signaling promotes gene expression and mediates the biological functions of
the PDGF isoforms, e.g. proliferation, migration, and survival. Adapted from [225].
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Since being originally isolated due to its ability to promote cell proliferation, PDGF
has been appreciated to initiate additional cellular responses such as cell survival and
migration. PDGF signaling is undoubtedly required for the spreading of various
populations of cells during embryonic development, such as oligodendrocyte
precursors in the spinal cord, neural crest mesenchymal cells toward the branchial
pouches and cardiac outflow tract, and pericytes along newly formed angiogenic
sprouts [222]. The mechanisms by which PDGF regulates the process of cell
spreading remain largely undefined in most of the situations. However, recent studies
in Drosophila, Xenopus, and mouse have demonstrated a role for PDGF family
members in directed cell migration in certain developmental processes in vivo. PDGF-
AA induced chemotaxis is well documented [226], and PDGFR-α promotes a
chemotactic response in various cell types [227]. Nagel et al. demonstrated PDGF-
A/PDGFR-α signaling is required for directional migration of mesoderm on its
endogenous matrix substratum, as well as for cell orientation in the embryo [228].
Knockout of the genes encoding PDGFR-β or PDGF-B in mice causes deficient mural
cell recruitment, leading to widespread microvascular bleeding [229-231]. PDGF-BB,
the main ligand of PDGFR-β, is a potent stimulant of SMCs recruitment during
neointimal hyperplasia following vascular injury [222]. PDGFR signaling has also
emerged as a predominant pathway in the recruitment of perivascular MSCs, which
play fundamental roles in angiogenesis, wound repair and tissue regeneration [210,
232-234]. A recent study has further identified the synergistic relationship between
α5β1-integrin and PDGFR-β. Cell adhesion to fibronectin induced PDGFR-β
phosphorylation in an α5β1-integrin dependent manner, leading to the activation of
the PI3K/Akt pathway, actin reorganization and recruitment of MSCs [235].
3.7 Signaling pathways related with cell migration
In order to participate in tissue repair and regeneration, MSCs have to be mobilized
and migrate to the target sites and integrate with the local tissues. Understanding the
potential signaling pathways in relation to the endogenous cell homing can offer new
insights into the management of bone defect healing.
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3.7.1 PI3K/Akt signaling pathway
Phosphoinositide 3-kinases (PI3Ks) were initially discovered as lipid kinases
associated with viral oncoproteins about 20 years ago [236]. Since then, the
association between PI3Ks and cancer has been further elucidated [237]. Over the last
decade, growing evidence has indicated that preferential activation of PI3Ks is
important for defining the leading edge in a wide range of motile cell types [238-240].
PI3K has been the focus of attention with respect to its activation by chemokine
receptors and the role it plays in regulating cell migration [241, 242]. The PI3K
family can be divided into three main classes according to their in vitro lipid substrate
specificity, enzyme structure and products formed [243]. The heterodimeric class I
PI3Ks consist of two structurally closely related subgroups. Class IA enzymes share
one of the common regulatory subunits (p85α and -β, p50α, and p55α and -γ) and
different catalytic subunits (p110α, -β, and -γ) [244]. Class IB enzymes comprises a
p101 or p84 regulatory subunit as well as the p110γ catalytic isoform [245, 246].
Classes IA and IB are the most extensively studied PI3Ks and, together with their
lipid product phosphatidylinositol (3,4,5)-triphosphate (Ptdlns(3,4,5)P3), have been
widely implicated in controlling cell migration and polarity [247] (Figure 7). Class II
PI3Ks lack a regulatory subunit and contain three isoforms (C2α, -β, and -γ), which
are capable to phosphorylate Ptdlns and Ptdlns(4)P to Ptdlns(3)P and Ptdlns(3,4)P2,
respectively. Recent studies have suggested that class II PI3Ks may play a positive
role in the regulation of cell adhesion and actin reorganization during cell migration
and wound healing in non-immune cell systems [248, 249]. The sole member of class
III PI3K is vacuolar protein sorting 34 (Vps34), which consists of a regulatory subunit,
p150, for its activity. Vps34 produces only Ptdlns(3)P from Ptdlns and is involved in
multiple steps of membrane trafficking. However, no defined role of Vps 34 in cell
migration has been reported so far [250].
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Figure 7. Signaling through phosphatidylinositol (3,4,5)-triphosphate. On stimulation of
class I PI3Ks together with their lipid product phosphatidylinositol-3,4,5-trisphosphate
(Ptdlns(3,4,5)P3), several phosphatases are involved in subsequent dephosphorylation steps (dashed
arrows). PTEN hydrolyses the 3-phosphate of Ptdlns(3,4,5)P3 to regenerate phosphatidylinositol-4,5-
bisphosphate (Ptdlns(4,5)P2), thus terminating PI3K signaling. 5-phosphatases such as SHIP, however,
generate Ptdlns(3,4)P2 that, like Ptdlns(3,4,5)P3, can facilitate the activation of Akt. The key reaction
steps outlined are integrated into complex signaling networks in which Akt and other targets of 3-
phosphorylated phosphoinositides, together with further downstream components, regulate cell growth,
proliferation, survival and motility. Adapted from [251].
The serine/threonine kinase Akt, also known as protein kinase B (PKB), is the best
characterised downstream effector of class І PI3Ks. The Akt family consists of three
isoforms, namely Akt1 (PKBα), Akt2 (PKBβ) and Akt3 (PKBγ). They are expressed
ubiquitously, share similar activation mechanisms and typically initiate signal relay to
secondary signaling cascades by phosphorylating an overlapping subset of particular
proteins [252]. Akt has emerged as one of the key regulators for cell survival,
proliferation, angiogenesis and glucose metabolism [253-256]. A growing body of
evidence indicates that Akt may also affect cytoskeletal changes and promote cell
migration in different organisms [257-260]. However, the role of isoform-specific Akt
family members in the regulation of cell migration has been difficult to ascertain.
Activation of Akt1 has been found to negatively influence mammary epithelial cell
migration and epithelial-mesenchymal transition (EMT) [261], whereas siRNA
knockdown of Akt1, but not Akt2, results in a considerable increase in cell migration,
30 | P a g e
which was related to the activation of extracellular-signal-regulated kinase-1 and -2
(ERK1/2) [261]. The inhibitory effect of Akt1 on cell motility and invasion has also
been investigated in various breast cancer cell lines, in which degradation of the
nuclear factor of activated T cells (NFAT) transcription factors was involved [262].
On the contrary, other reports have claimed a distinctive role of Akt1 in promoting
cell migration and invasion [263, 264]. Similarly, the positive effect of Akt2 on cell
motility has been well established. Recent investigations have demonstrated that the
reduction of Akt2 inhibits chemotaxis signal transduction in glioma cells [265] and
breast cancer cells [266]. On the other hand, Akt2 has been found to be a negative
regulator of Pak1 [267]. Akt2 knock-out mouse embryonic fibroblasts have increased
migration, especially invasion through ECM, suggesting a negative role of Akt2 in
regulation of Rac/Pak signaling, the actin cytoskeleton and cell migration [267].
Interestingly, an inhibitory role for both Akt1 and Akt2 in prostate cancer migration
and invasion has been reported in a recent study [268]. However, the inconclusive
effects of Akt1 and Akt2 on cancer cell migration may not translate to other cell types.
These studies demonstrate the importance of crosstalk between the PI3K/Akt pathway
and other pathways and highlight the cell type-specific actions of Akt kinases in the
regulation of cell motility.
Several PI3K/Akt stimulators have been documented to enhance cell migration [269-
272]. The role of CXCL12, one such stimulator, in cancer cell invasion has been well
studied. Binding of CXCL12 to CXCR4 leads to the stimulation of a signaling
cascade that involves activation of multiple targets, including ERK1/2, p38 and Akt
[273-276]. It has been reported previously that CXCL12 could stimulate a rapid and
transient accumulation of PtdIns(3,4,5)P3 in leukaemic T cell lines and peripheral
blood-derived lymphocytes within 15 sec, and the elevated accumulation returned to
basal level 5 min after chemokine treatment [277]. Other work has further
demonstrated the cell migration induced by CXCL12 can be abrogated by the PI3K
inhibitors wortmannin and LY294002 [241, 242, 273]. Hypoxia pre-conditioning has
also been shown to enhance MSCs migration and improve their tissue regenerative
potential via the activation of the PI3K/Akt signaling pathway [278].
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A number of phosphorylation targets for Akt are now emerging. Enomoto and
colleagues have recently identified a novel substrate of Akt, designated Girdin
(girders of actin filaments), which is an actin binding protein [279]. The accumulation
of phosphorylated Girdin at the leading edge of migrating cells suggests that Girdin is
essential for the remodeling of the actin cytoskeleton that propels the cells forward
and provides a direct link between Akt and cell motility [279, 280]. The p70 S6
kinase (p70S6K
) is also a downstream effector of the PI3K/Akt pathway, which
regulates cytoskeleton dynamics via the actin filament cross-linking proteins. The
overexpression of p70S6K
in ovarian cancer cells has been reported to induce a marked
reorganization of the actin cytoskeleton and promote directional cell migration [281].
Researchers have demonstrated that p70S6K
can stimulate the rapid activation of Rho
family small GTPases Cdc42 and Rac1, as well as their major downstream effector,
p21-activated kinase 1 (PAK1) [281, 282]. PAK1 is the first discovered member of
the mammalian PAK family, which comprises six proteins divided in group І (PAK1-
3) and group II (PAK4-6) according to their structural and functional features [283,
284]. PAK1 plays a fundamental role in controlling cell motility by linking a variety
of extracellular signals to changes in actin cytoskeleton organization, cell shape and
adhesion dynamics [285-287].
3.7.2 MAPK signaling pathway
Mitogen-activated protein kinases (MAPKs) belongs to a large family of highly
conserved serine/threonine protein kinases, which are found in all eukaryotic cells,
and expressed in virtually all tissue types. To date, six distinct classes of MAPKs have
been characterised, namely, the extracellular signal-regulated kinases (ERK) 1 and 2
(ERK1/2), ERK3/4, ERK5, ERK7/8, p38 (α/β/γ/δ) MAPK, and c-Jun N-terminal
kinases (JNK1/2/3) [288-290]. JNK1/2/3 and p38 (α/β/γ/δ) MAPK are collectively
called stress-activated protein kinases (SAPK). ERK5 is also known as big MAPK
(BMK1) [291, 292]. Very little is known about the atypical MAPK enzymes, ERK3/4
and ERK7/8. However, all the members of MAPKs consist of a homodimer of
molecular mass of about 80 kDa and contain the signature sequence T–X-Y, where T
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and Y are threonine and tyrosine, and X denotes glutamate in ERK, proline in JNK,
and glycine in p38, respectively [293, 294].
MAPKs are pivotal components of a complex intracellular signal transduction
pathway that functions to integrate the cells’ response to extracellular stimuli (Figure
8). The activation of MAPKs proceeds through a series of upstream molecules in an
orderly fashion. Various extracellular stimulations, e.g. hormones, growth factors,
cytokines and stress signals, are relayed from the cell surface through an elegant
cascade of phosphorylation events leading to the cell nucleus, where a diverse array of
intracellular responses occur [295]. In general, ERK1/2 are preferentially activated in
response to growth factors that act through receptor tyrosine kinases, e.g. epidermal
growth factor (EGF), fibroblast growth factor (FGF) and PDGF, whereas the
JNK1/2/3 and p38 (α/β/γ/δ) MAPKs are more responsive to pro-inflammatory
cytokines such as tumor necrosis factor (TNF)-α and interleukin (IL)-1β or stress
stimuli ranging from osmotic shock to ionizing radiation [289, 290, 295].
Consistent with their critical roles in key cellular activities, including cell proliferation,
differentiation, survival and apoptosis [296], the MAPK signaling pathway has also
been implicated in the migration of various cell types [297]. Accumulating data
suggest that MAPKs are essential in regulating the dynamics of focal adhesion and
the reorganization of microtubules and filamentous actin that is required for cell
morphogenesis and migration [297]. ERK1/2 is the most extensively studied
subfamily of MAPKs and has been implicated in the migration of a wide range of cell
types. Numerous studies have reported the ERK pathway inhibitors PD98059 and
U0126 suppress cell migration in response to ECM, such as fibronectin, vitronectin
and collagen [298, 299], growth factors, such as EGF, FGF, PDGF and VEGF [300-
303], and other stimuli, such as fetal bovine serum and urokinase-type plasminogen
activator (uPA) [304, 305].
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Figure 8. Schematic of the three mammalian mitogen-activated protein kinase
(MAPK) signaling pathways. Extracellular stimuli activate the MAPK pathways through
mechanisms mediated by GTPases, including RAS, RAC, CDC42 (cell-division cycle 42) and RHO
(RAS homologue). Once MAPKKKs (MAPK kinase kinases), such as RAF, MEKK (MAPK/ERK
kinase kinase) and TAK (TGFβ-activated kinase), are activated, they phosphorylate MAPKKs (MAPK
kinases) on two serine residues. MAPKKs in turn phosphorylate the MAPKs ERK (extracellular-
signal-regulated kinase), JNK (JUN N-terminal kinase) and p38 on both threonine and tyrosine
residues, which results in the catalytic activation of these MAPKs. Activated MAPKs can translocate to
the nucleus to phosphorylate a number of transcription factors, such as ternary complex factor (TCF)
family members and components in the activator protein 1 (AP1) complexes, including JUN and
activating transcription factor 2 (ATF2), thereby altering gene transcription. TCF forms a complex with
serum response factor (SRF) to regulate Fos induction. AP1 is involved in the transcription of a wide
variety of genes. Adapted from [306].
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3.8 Summary
Although small bone defects may heal through the body’s normal reparative process,
large skeletal deficits overwhelm this system and may fail to reach a satisfied
structural and functional recovery, despite the use of conventional reconstructive
approaches. With the recent advanced research in regenerative medicine and tissue
engineering, cell-based therapy has emerged as one of the most promising approaches
for tissue repair and regeneration. Cell-cell interaction between host and donor cells is
believed to play a critical role in enhancing the healing process after in vivo cell
transplantation. However, little is known about how donor cells lead to tissue repair
and regeneration after transplantation or how they interact with the host cells in vivo.
In addition, a comprehensive understanding of the host response to the transplanted
donor cells is crucial for developing novel clinical strategies based on the active host
cell contribution to the regeneration of damaged tissues. The recruitment process of
endogenous cells may be facilitated by external stimuli, such as the manipulation of
donor cells and the application of homing factors, which may offer new therapeutic
options to harness the host’s innate capacity for the repair and regeneration of bone
defects. Migration of MSCs is controlled by complicated signal networks.
Understanding the molecular mechanisms of MSC migration will help to develop
strategies to facilitate the tissue forming capacity of cell delivery therapy and to
optimize cell-based bone tissue engineering.
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CHAPTER 4 RESEARCH REPORTS
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REPORT ONE
Implantation of osteogenically differentiated donor mesenchymal
stromal cells causes recruitment of host cells
Yinghong Zhou, Wei Fan, Indira Prasadam, Ross Crawford, Yin Xiao
Suggested Statement of Contribution of Co-Authors for Thesis by
Published Papers:
Contributors Statement of contribution*
Yinghong Zhou Involved in the conception and design of the project, performed
laboratory experiments, data analysis and interpretation. Wrote
the manuscript.
Wei Fan Involved in sample collection, and assisted with manuscript
preparation.
Indira Prasadam Assisted with manuscript preparation.
Ross Crawford Involved in the conception and design of the project, assisted in
collection of clinical samples, and reviewed the manuscript.
Yin Xiao Involved in the conception and design of the project, and
reviewed the manuscript.
Principal Supervisor Confirmation
I have sighted email or other correspondence from all co-authors confirming
their certifying authorship.
Name Signature Date
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ABSTRACT
Background: The interaction between host and donor cells is believed to play an
important role in bone tissue engineering. However, it is still unclear how donor cells
behave and interact with host cells in vivo. The purpose of this study was to track the
interactions between transplanted osteogenic cells and donor cells during osteogenesis.
Methods: An in vitro migration assay was carried out to investigate the ability of
osteogenically differentiated human mesenchymal stromal cells (O-hMSCs) to recruit
undifferentiated mesenchymal stromal cell (MSCs). At the in vivo level, O-hMSCs
were implanted subcutaneously or into skull defects in severe combined
immunodeficient (SCID) mice. New bone formation was observed by µCT and
histological procedures. In situ hybridisation (ISH) against human Alu sequences was
performed to distinguish donor cells from host cells.
Results: The in vitro migration assay revealed an increased migration potential of
MSCs by co-culturing with O-hMSCs. In agreement with the results of in vitro studies,
ISH against human Alu sequences showed that host mouse MSCs migrated in large
numbers into the transplantation site in response to O-hMSCs, suggesting that donor
cell characteristics may help to recruit the host MSCs for transplantation success.
Interestingly, host cells recruited by O-hMSCs were the major cell populations in
newly formed bone tissues, indicating that O-hMSCs can trigger and initiate
osteogenesis when transplanted in orthotopic sites.
Conclusions: The observations from this study demonstrated that in vitro induced O-
hMSCs were able to attract host MSCs in vivo and were involved in the osteogenesis
together with host cells, which may be of importance for bone tissue engineering
applications.
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INTRODUCTION
Rapid expansion of knowledge regarding progenitor cell-based tissue engineering
offers new insights into the treatment of various kinds of diseases [24], especially
those related to bone defects as a result of tumour, infection, trauma, biochemical
disorders, and abnormal skeletal development [307]. Cell transplantation is one of the
most promising strategies used to enhance the tissue-healing process, as it introduces
functioning cells directly into affected areas [308, 309]. The concept of cell-based
therapy involves cell isolation from autologous, allogenic, or xenogenic sources, cell
proliferation and differentiation in vitro, and application in vivo [23, 310, 311].
Recently, this concept has been tested in large bone defect healing in several studies
[39, 312].
Cell-cell interactions between host and donor cells are believed to play an important
role in enhancing the healing process after in vivo cell transplantation [313]. In cell
therapy for bone defect healing, the delivery of osteogenic cells into the defect area is
often considered to be of direct help to the new bone formation [314]. Due to the
reduced multipotency compared with progenitor cells, osteoblasts may be more
committed to form new bone tissue [64]. A study on the rabbit mandible showed that
transplantation of osteoblast-like cells has the potential to promote maturity of the
distracted callus [315]. Another study reported that transplantation of bone marrow
stromal cells contributed to new bone formation in vivo [316]. However, until now,
cell therapy for bone tissue engineering has focused on determining a source of
exogenous cells with osteogenic capacity, without paying attention to the possibility
that the recipient itself might provide the cell elements necessary for tissue repair.
Little evidence shows how and to what extent the transplanted donor osteogenic cells
behave and interact with host mesenchymal stromal cells (MSCs) during the new
bone formation process [317]. The purpose of this study was to investigate how
transplanted osteogenically differentiated human MSCs (O-hMSCs) were involved in
osteogenic processes and how they interact with host MSCs after transplantation.
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MATERIALS AND METHODS
Cell culture
Human bone marrow was sourced from patients (n=5, three females and two males)
in the Department of Orthopaedics at Prince Charles Hospital, with informed consent
and ethics approval from the Ethics Committee of Queensland University of
Technology. The average age of the patients involved in this study was 69.6 ± 3.7
(range 65-75) years. The hMSCs for this study were isolated by density gradient
centrifugation over Lymphoprep™ (Axis-Shield PoC AS, Oslo, Norway) according to
the manufacturer’s protocol [318, 319]. The cells were then seeded in culture flasks in
Dulbecco’s modified Eagle’s medium (DMEM; Gibco®, Life Technologies Pty Ltd.,
Australia) containing 10% (v/v) fetal bovine serum (FBS; In Vitro Technologies,
Australia) and 1% (v/v) penicillin/streptomycin (P/S; Gibco®, Life Technologies Pty
Ltd., Australia). Following 3 days incubation in a humidified atmosphere of 5% CO2
at 37ºC, the non-adherent cells were washed away leaving behind the adherent cell
population that was growing in clusters. Upon reaching 70-80% confluence, the
attached hMSCs were subcultured after treatment with 0.25% trypsin (Gibco®, Life
Technologies Pty Ltd., Australia) and 1mM EDTA (Gibco®, Life Technologies Pty
Ltd., Australia). In vitro osteogenic differentiation of hMSCs was performed in
osteogenic medium (DMEM supplemented with 10% FBS, 1% P/S, 10 mM β-
glycerophosphate, 50 µM ascorbic acid and 100 nM dexamethasone; Sigma Aldrich,
Australia) for 2 weeks.
Characterisation of hMSCs
To confirm the phenotype of hMSCs obtained through in vitro culture, flow
cytometric analysis against the cell surface markers CD34, CD45, CD73, CD90 and
CD105 (BD Biosciences, North Ryde, NSW, Australia; Clontech, Palo Alto, CA,
USA) was performed [320]. Briefly, after incubation with primary antibodies against
the aforementioned cell markers and repeated washing with 3% PBS/bovine serum
albumin (BSA), hMSCs were resuspended and incubated with secondary antibody
(phycoerythrin-conjugated goat anti-mouse immunoglobulin G; Jackson
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ImmunoResearch Laboratories, West Grove, PA, USA), then resuspended in 3%
PBS/BSA and read through in a flow cytometry machine (Becton-Dickinson, Cowley,
Oxford, UK). In the negative control, the primary antibody was omitted and replaced
by a mouse isotype IgG.
In vitro migration assay
hMSCs (1×104) were seeded on polyethylene terephthalate (PET) track-etched
membrane transwell inserts (10 mm effective diameter of membrand with 8 µm pore
size; Becton-Dickinson Labware, Franklin Lakes, NJ, USA). After 24 h of cell
attachment, the transwell inserts were assembled onto the 12-well companion plates
cultured with 1×104 hMSCs, which were subjected to osteogenic differentiation (O-
hMSCs) or retained in undifferentiated condition (hMSCs) for 2 weeks prior to the
migration assay. All co-cultures were performed in serum-free medium to rule out the
effect of serum in complete medium on the migration of hMSCs. Cell migration was
allowed to proceed for 8 h. Cells that had migrated to the other side of the insert
membrane were fixed, and then stained with crystal violet. Images were taken with an
inverted microscope (Eclipse TS100, Nikon Australia Pty Ltd.). The migration was
quantified by counting the number of cells that migrated through the pores in six
randomly selected fields on each transwell insert membrane, and values from three
independent experiments, with triplicate samples in each experiment, were averaged
and showed as mean ± SD.
In vitro osteogenic differentiation of hMSCs on collagen scaffolds
Collagen scaffolds were fabricated from a 1mm thick bovine type I collagen sheet
(Medtronic Australia Pty Ltd, North Ryde, NSW, Australia), using a hole puncher 5
mm across and sterilized with 75% ethanol, followed by three rinses of PBS prior to
cell seeding. For the in vitro osteogenic differentiation, 6×104
hMSCs were first
seeded onto the collagen membrane and cultured in osteogenic medium for 2 weeks.
Collagen with no cells and collagen with undifferentiated hMSCs were used as
controls.
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In vivo implantation of osteogenically differentiated hMSCs
Nine 5 week-old male severe combined immunodeficient (SCID) mice (Animal
Resources Centre, Canning Vale, WA, Australia) were used for the skull defect model
[78, 321], and another three SCID mice were used for subcutaneous cell implantation.
For the skull defect implantation, an incision was made longitudinally along the
centre line of the shaved skull skin. A 2.5mm diameter defect was then made through
the skull, using a dental burr. The defect was treated as follows with three mice in
each group: i.e. defects filled with collagen scaffold, collagen carrying
undifferentiated hMSCs, or collagen carrying O-hMSCs. For the subcutaneous cell
implantation, three longitudinal incisions were made in each animal along the central
line of the shaved dorsum, approximately 1 cm apart. Each individual pocket received
one scaffold from each group, using a complete randomized design for cell-free
collagen scaffolds, collagen scaffolds carrying hMSCs, or collagen scaffolds carrying
O-hMSCs. The SCID mice were sacrificed 2 weeks after subcutaneous implantation
and 4 weeks after the skull defect implantation.
Micro-computed tomography (μCT) assessment
All the samples harvested were fixed in 4% paraformaldehyde overnight at room
temperature and then washed in PBS. Cross-sectional scans parallel to the cell-
seeding surface of the implants were performed in a µCT scanner (µCT40, Scanco
Medical AG, Brüttisellen, Switzerland) at a resolution of 12 µm and three-
dimensional (3D) images of the implants were reconstructed. The calcified area inside
the implant on each scanned slice was circled out and the volume of calcification
within the implants was measured using the inbuilt scanner software package and
recorded for statistical analysis.
Histology and immunohistochemistry (IHC)
All harvested subcutaneous samples were embedded in paraffin blocks for sectioning,
while the skull defect samples were first decalcified in 10% EDTA solution for 4
43 | P a g e
weeks and then embedded in paraffin. 5 µm serial sections were cut from the
embedded samples, using a microtome (Leica Microsystems GmbH, Wetzlar,
Germany). Only sections close to the centre of the samples were used for the staining.
For haematoxylin and eosin (H&E) staining, tissue slices were dewaxed in xylene and
rehydrated in descending concentrations of ethanol (100%, 90% and 70%), then
stained with Mayer’s haematoxylin (HD Scientific Supplies Pty Ltd., NSW, Australia)
for 2 min before rinsing with tap water for 5 min. The slides were dehydrated in
ascending concentration of ethanol (70%, 90% and 100%) then stained with eosin
(HD Scientific Supplies Pty Ltd., NSW, Australia) for 15 s, cleared with xylene and
mounted with DPX mounting medium (Fronine, Laboratory Supplies, NSW,
Australia).
IHC staining was performed for the assessment of new bone formation in the skull
defects. In brief, sections were blocked with 10% swine serum for 1 h, then incubated
with primary alkaline phosphatase antibody (ALP; 1:400, anti-human; Sigma-Aldrich,
Australia) overnight at 4°C. The sections were washed free of primary antibody and
incubated with biotinylated secondary antibody (Dako, CA, USA) for 15 min,
followed by incubation with horseradish peroxidise (HRP)-conjugated streptavidin-
biotin complex (Dako, CA, USA) for 15 min. Antibody complexes were visualized by
the addition of diaminobenzidine (DAB) for 3 min. The sections were then lightly
counterstained with haematoxylin and mounted with mounting medium.
von Kossa staining
In order to observe the calcification in subcutaneous implants, samples were subjected
to 5% silver nitrate (Sigma-Aldrich, Australia) and exposed to light for 30 min. The
sections were then washed in distilled water and dehydrated in ethanol before being
mounted with cover slips and mounting medium.
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In situ hybridisation (ISH)
To trace the donor cells (i.e O-hMSCs) in skull defect sites and subcutaneous implants,
human Alu probe was obtained from PanPath (Budel, the Netherlands). Briefly,
hybridisation of the Alu probe was carried out for 16 h at 37°C in a humidified
chamber after denaturation at 95°C for 5 min, followed by post-hybridisation washing
procedure. 3-Amino-9-ethylcarbazole (AEC) was then applied to visualize the
hybridisation. The sections were then counterstained with haematoxylin, and mounted
with Crystal MountTM
aqueous mounting medium (Sigma-Aldrich, Australia).
The localisation of transplanted human cells identified by the Alu probe in both
subcutaneous and skull defect scaffolds was observed, recorded and analysed. The
lining cells around the initial bone were counted as osteoblasts and those trapped in
the bone matrix and isolated in lacunae were counted as osteocytes. Three sections
from each group were included and the cell percentage of both donor human cells and
host mouse cells in three randomly selected areas on each section within the implant,
newly-formed bone tissues and bone marrows adjacent to the implants or from the
femur was calculated and recorded for further analysis.
Statistical analysis
Non-parametric Kruskal-Wallis test was performed to distinguish the differences
among all the groups in the in vitro migration assay and the µCT quantified data
analysis. The differences of cell percentage between donor and host cells inside
samples were subjected to Wilcoxon test. Significant difference was considered at
p<0.05.
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RESULTS
hMSCs characterisation and in vitro osteogenic differentiation
To verify the cells derived from the human bone marrow are MSCs, a panel of
antibodies against stem cell markers was chosen to evaluate the phenotypes of the
hMSCs used in the present study. Flow cytometry against CD34, CD45, CD73, CD90,
and CD105 revealed that the in vitro expanded hMSCs were negative for
haematopoietic cell surface markers of CD34 and CD45, while positive for selected
mesenchymal cell surface markers of CD73, CD90 and CD105 (Figure 9A). The
hMSCs derived from the bone marrow showed typical fibroblastic morphology
(Figure 9B). von Kossa staining showed calcium precipitation by hMSCs after in
vitro osteogenic differentiation, confirming the mineralization potential of hMSCs
derived from bone marrow (Figure 9C).
Figure 9. hMSCs characterisation and in vitro osteogenic differentiation. (A) Flow
cytometric analysis of cell surface antigen expression in hMSCs showed negative expression of
haematopoietic cell surface markers (CD34 and CD45) and positive expression of mesenchymal cell
surface markers (CD73, CD90 and CD105). (B) Microscopic observation of undifferentiated hMSCs
showed typical fibroblastic morphology. (C) von Kossa staining showed the mineralization potential of
hMSCs after osteogenic differentiation.
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In vitro migration assay
The effect of O-hMSC on the migration of hMSCs was assessed using a transwell
device containing a cell culture insert with 8 µm pores. After 8 h of incubation, the
number of hMSCs which migrated through the transwell membrane increased
significantly in the co-culture with O-hMSCs (Figure 10A, D, G) compared with the
serum-free medium (Figure 2A, B, E) and the co-culture with undifferentiated hMSCs
(Figure 10A, C, F) (p<0.05).
Figure 10. In vitro migration assay. (A) Comparison of the number of hMSCs that migrated to
the other side of the insert membrane among different groups. SF, serum-free medium; hMSC,
undifferentiated hMSCs were seeded in the companion plate; O-hMSC, osteogenically differentiated
hMSCs were seeded in the companion plate (*p<0.05). (B)-(D) Schematics representation of co-culture
groups. (E)-(G) Images of hMSCs that migrated to the other side of the insert membrane: (E) Serum-
free medium; (F) hMSCs were seeded in the companion plate; and (G) O-hMSCs were seeded in the
companion plate.
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Osteogenesis in subcutaneous and skull defect implants
H&E staining on ectopic implants showed no histological signs of new bone
formation (Figure 11I-K). However, µCT and von Kossa staining showed
mineralization along collagen fibres in the scaffolds carrying O-hMSCs (Figure 11C,
D, L). There was no mineralization in the scaffolds carrying undifferentiated hMSCs
or in the cell-free scaffolds (Figure 11A, B, D).
H&E staining revealed significant new bone formation, showing a trabecular bone-
like structure within the implants carrying O-hMSCs in the skull defect areas (Figure
11O). µCT scanning and 3D reconstruction revealed that a substantial amount of
mineralization was found in the implants carrying O-hMSCs, showing the greatest
volume of the calcified areas (Figure 11G, H). However, there was almost no new
bone formation within the cell-free or undifferentiated hMSCs-carrying implants in
the skull defect areas (Figure 11E, F, H, M, N). The newly formed bone in the O-
hMSCs implants was further confirmed by IHC staining against ALP (Figure 11P).
Cell migration in ectopic sites
In the subcutaneous implants, ISH against human Alu sequences revealed that host
mouse cells (Alu-negative) migrated into the centre of the implants, gathered together
and were then surrounded by donor O-hMSCs. This showed a close relationship
between the transplanted human cells and the host mouse cells in the osteogenic
process (Figure 12D). On the other hand, in the cell-free or undifferentiated hMSCs-
loaded scaffolds (Figure 12B, C), the host mouse cells distributed sparsely in the
transplanted scaffolds. A higher host:donor cell ratio was found inside the scaffolds
when O-hMSCs were implanted subcutaneously (Figure 12A; p<0.05).
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Figure 11. Osteogenesis in subcutaneous and skull defect implants. (A)-(C) 3D images of
subcutaneous implants with scaffold only, scaffold+hMSCs or scaffold+O-hMSCs. (D) Quantification
of the bone volume of the calcified areas within the subcutaneous implants (*p<0.05). (E)-(G) 3D
images of skull defect treated with scaffold only, scaffold+hMSCs or scaffold+O-hMSCs. (H)
Quantification of bone volume of the calcified areas within the skull defect implants (*p<0.05). (I)-(K)
H&E staining of the subcutaneous implants showed no new bone formation in all three groups. (L) von
Kossa staining revealed calcification within the subcutaneous implants carrying O-hMSCs. H&E
staining of skull defect filled with scaffold alone (M) and scaffold+hMSCs (N), showing no new bone
formation. (O) H&E staining of skull defect filled with scaffold+O-hMSCs showing new bone
formation. (P) ALP staining revealed active osteoblasts in the new bone tissues in skull defect treated
with scaffold +O-hMSCs.
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Figure 12. Distribution of donor human cells and host mouse cells in the
subcutaneous implants revealed by ISH. (A) Quantification and comparison of host:donor cell
ratio between the hMSCs and O-hMSCs groups hMSC, undifferentiated hMSCs; O-hMSC,
osteogenically differentiated hMSCs (*p<0.05). (B) Implants with collagen scaffold only. (C) Implants
with hMSCs showing both the donor (red, arrow) and host cells (blue, arrowhead). (D) Implants with
O-hMSCs showing both the donor (red, arrow) and host cells (blue, arrow head).
Cell migration in orthotopic sites
In the skull defect areas, ISH against human Alu sequences revealed that more than
60% of total cells in the observed areas were host mouse cells within the O-hMSCs
implants (Figure 13A; p<0.01). In newly formed bone tissues, O-hMSCs were found
to turn into both osteoblasts and osteocytes (Figure 13D-F). However, the number of
human-originated osteoblasts and osteocytes were significantly less than the cells
from the host, indicating more host cells were involved in the newly formed bone
tissues (Figure 13B-F; p<0.05).
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Alu-positive cells in the newly formed bone marrow
In the bone marrow of the newly formed bone within the implants carrying O-hMSCs,
Alu-positive cells were identified coexisting with host marrow cells and occupied
about 30% of total bone marrow cells (Figure 13H). Some Alu-positive cells were
round in shape and mixed with Alu-negative host cells in the newly formed bone
marrow, while others were found attached to the endosteal bone surfaces (Figure
13G). In the bone marrow close to the defect, no Alu-positive cells were found
(Figure 13I).
To investigate whether transplanted donor cells migrated into the circulation system,
femur samples from the same animal after cell transplantation were collected and
processed. No Alu-positive cells were found in bone marrow collected from the femur
samples, indicating that transplanted O-hMSCs were only involved in local
osteogenesis (Figure 13J).
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Figure 13. Distribution of O-hMSCs and host mouse cells in the skull defect implants
and bone marrows, revealed by ISH. (A)-(C) Quantification and comparison of cell percentage
between donor and host cells (O-hMSC: osteogenically differentiated hMSCs): (A) General cell
percentage; (B) Cell percentage as osteoblasts; (C) Cell percentage as osteocytes (*p<0.05, **p<0.01).
(D)-(F) Skull defect implants showed both the donor (red) and host cells (blue); arrows, donor cells-
derived osteoblasts on new bone surface; arrowhead, donor cells-derived osteocytes. (G) Bone marrow
in skull defect showing both donor human cells (red) and host mouse cells (blue); arrow, marrow cell-
like transplanted donor cells; arrowhead, endosteal cell-like transplanted donor cells. (H)
Quantification and comparison of cell percentage between donor O-hMSCs and host mouse cells in
bone marrow (*p<0.05). (I) Bone marrow adjacent to the skull defect. (J) Bone marrow in the femur of
the mouse with cell-transplanted skull defect.
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DISCUSSION
Cell therapies have tremendous potential to treat a wide array of diseases, and the
major strategy of cell transplantation is via scaffolds, which are intended to serve as
templates for tissue formation [322]. However, integration of templated tissues with
host tissues remains a significant challenge. Since MSCs were first isolated from bone
marrow [323], they have been considered to be among the most important stem cell
sources for cell therapy, due to their multilineage differentiation capacity [116, 324,
325]. Although the osteogenic differentiation potential of MSCs has been well
characterised in many in vitro and in vivo studies [110, 326-328], no study has been
conducted so far to clarify how the donor cells interact with the host cells in the
osteogenic process.
In the present study, the in vivo behaviour of O-hMSCs was investigated using the
ISH technique against human Alu sequences in both subcutaneous (ectopic) and skull
defect (orthotopic) osteogenesis. The results showed that O-hMSCs played a
significant role in host cell recruitment in both ectopic and orthotopic sites. The cell
distribution and the ratio of donor:host cells were significantly different in ectopic and
orthotopic sites.
Subcutaneous transplantation of O-hMSCs revealed that the host cells migrated into
the centre of O-hMSCs and exhibited almost a 50:50 donor:host cell ratio. This
phenomenon may indicate that ectopic osteogenesis of transplanted O-hMSCs needs
the involvement of a certain amount of host cells to generate an optimal local
microenvironment for initial calcification. The host cells formed cell clusters or
nodules among the transplanted cells, indicating close cell-cell communication
between host and donor cells.
In the skull defect areas, a substantial of newly formed bone tissue was found in
implants carrying O-hMSCs. It is very interesting to find that these transplanted O-
53 | P a g e
hMSCs were involved in the new bone formation and differentiated to both
osteoblasts and osteocytes at a similar donor:host cell ratio. Our finding of a large
number of recipient cells at the defect sites confirmed cell migration into the defect.
More host cells than donor cells differentiated into osteoblasts and osteocytes in skull
defect areas. This is different from the subcutaneous implant, which showed a nearly
equivalent number between the host and donor cells. This difference might be
attributed to the different local environment (ectopic and orthotopic) and different
osteogenic process (calcification and new bone formation). The smaller number of O-
hMSCs compared with host cells as osteoblasts and osteocytes in skull defect areas
may indicate that transplanted donor cells played a more important role in initiating
osteogenesis, rather than in maintaining the bone formation process. Different
proliferation rates between the human and mouse cells, postoperative complications
present at the defect sites, and other factors, such as immunological reactions, may
also tip the balance in favour of the recipient cells.
Another interesting aspect of tracking the transplanted donor cells is to investigate the
role of donor cells in the reconstitution of bone marrow in the newly formed bone,
and whether they could enter the blood flow and circulate into the bone marrow cell
compartment. In this study, only in the bone marrow cavities in the skull defects were
the donor cells found. This means that at the 4 week time point, the donor cells could
recruit host haematopoietic cells to form bone marrow in newly formed bone.
However, donor cells did not travel to the far end of body extremities. In the new
bone marrow, transplanted cells took up a lower cell percentage than the host marrow
cells. Considering the similarly low percentage of donor O-hMSCs as osteoblasts and
osteocytes in newly formed bone tissues, in vivo osteogenesis seems to require more
involvement of host cells than donor cells. In the bone marrow, some transplanted
cells presented the round shape of host marrow cells, but some were attached to the
endosteal bone cavity surface. This phenomenon suggests some transplanted cells
could become endosteal cells.
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The reason why O-hMSCs recruited more MSCs compared to undifferentiated
hMSCs is not clear. It is possible that cytokines released by O-hMSCs may have
stimulated MSCs to travel toward O-hMSCs. A recent study demonstrated that SDF-
1, a member in the chemokine family, is important for the migration of MSCs to bone
marrow [164]. Many studies have shown that during tissue repair, local SDF-1
expression is upregulated and acts as a signal to recruit the CXCR4 (SDF-1 ligand)
expressing cells from circulation and/or bone marrow. Apart from chemokines, other
molecular factors, such as fibroblast growth factor (bFGF) [329], matrix
metalloproteinases (MMPs) [330], and TLR ligation [331] were known to increase the
migratory homing activity of MSCs to the damaged tissue. It is possible that one or
more of the factors described above, or other factors unknown, are induced by O-
hMSCs, giving rise to a positive feedback of the host MSCs, which in turn trigger the
bone formation.
This study demonstrated that in both ectopic and orthotopic sites, transplanted O-
hMSCs could recruit host cells to initiate osteogenesis involving the formation of
osteoblasts, osteocytes, and bone marrow components. The ratio of donor:host cells
inside the transplants were significantly dependent on the local environment and
directly correlated with osteogenesis. This new finding provides the groundwork for
future studies about the interaction between transplanted donor cells and host cells in
bone defect healing.
ACKNOWLEDGEMENTS
The authors declare that there is no conflict of interest and gratefully acknowledge the
contribution of Ms Wei Shi to this study. The work was supported by Queensland
University of Technology.
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REPORT TWO
VEGF induced CXCL12/CXCR4 axis in the recruitment of mesenchymal
stromal cells (MSCs) during osteogenesis
Yinghong Zhou, Wei Fan, Indira Prasadam, Ross Crawford, Yin Xiao
Suggested Statement of Contribution of Co-Authors for Thesis by
Published Papers:
Contributors Statement of contribution*
Yinghong Zhou Involved in the conception and design of the project, performed
laboratory experiments, data analysis and interpretation. Wrote
the manuscript.
Wei Fan Involved in sample collection, and assisted with manuscript
preparation.
Indira Prasadam Assisted with manuscript preparation.
Ross Crawford Involved in the conception and design of the project, and assisted
in collection of clinical samples.
Yin Xiao Involved in the conception and design of the project, and
reviewed the manuscript.
Principal Supervisor Confirmation
I have sighted email or other correspondence from all co-authors confirming
their certifying authorship.
Name Signature Date
56 | P a g e
ABSTRACT
Background: Cell-cell interaction is believed to play important roles in the cell-based
therapy for bone defect healing. However, it is still unclear what factors are involved
in the interaction between donor cells and host cells during the bone healing process.
The purpose of this study was to investigate the potential effect of vascular
endothelial growth factor (VEGF) on the expression pattern of CXCL12/CXCR4 with
respect to the interactions between undifferentiated and osteogenically differentiated
mesenchymal stromal cells (MSCs and OMSCs) in vitro and in vivo.
Methods: Indirect co-culture models were applied to study the effect of VEGF
derived from OMSCs on the motility, the cell morphology and the CXCL12/CXCR4
expression of MSCs in vitro. The role of VEGF in the regulation of the host cell
recruitment was monitored by the application of neutralizing VEGF antibody in vivo,
after OMSC implantation into the skull defects in SCID mice.
Results: Our data showed that VEGF secretion increased dramatically in MSCs after
osteogenic differentiation in vitro and the secretion of VEGF promoted the motility of
MSCs with elongated actin filament and enhanced the expression of
CXCL12/CXCR4. In vivo findings revealed remarkable new bone formation within
the defects implanted with OMSCs. Furthermore, blocking VEGF with neutralizing
antibody resulted in significant decrease of MSCs recruitment and new bone
formation.
Conclusions: The observations from this study demonstrate that VEGF secreted by in
vitro induced OMSCs plays a pivotal role in MSC recruitment possibly via the
activation of the CXCL12/CXCR4 axis during osteogenesis.
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INTRODUCTION
Bone defect remains one of the most prevalent clinical problems even though various
strategies to stimulate bone healing have been explored [78, 332-335]. Due to
morbidity issues in donor sites and the limited availability of autografts, allograft
transplantation has become an attractive alternative treatment option. However, the
clinical results are limited and sometimes associated with poor clinical predictability
and severe adverse reactions [336]. In order to overcome these limitations, it is of
great interest to develop novel biological strategies, which require elucidation of the
molecular signals responsible for successful bone regeneration.
Cell-based therapy, such as mesenchymal stromal cell (MSC) transplantation, is a
promising strategy for the treatment of bone defects. MSCs are attractive candidates
for cell-based bone regeneration [337], because of their capacity to differentiate into
osteogenic lineage following in vitro expansion and osteogenic induction. The
development, remodelling and repair of bone are controlled by the proliferation,
differentiation, migration and apoptosis of various cell types, including osteoblasts,
osteoclasts and haematopoietic cells [338]. Although a lot of studies have focused on
the ability of implanted cells to differentiate within the defect sites, more recent
research suggests other important factors may be involved in the bone tissue
regeneration, such as the regulation of direct or indirect cell-cell interactions [339,
340], as well as the release of soluble factors [341]. However, the paracrine factors
and molecular mechanisms that contribute to the interaction between host cells and
donor cells during osteogenesis are poorly understood.
Recent studies have shown that cellular movement and relocalisation are essential for
many fundamental physiologic properties, including angiogenesis, chemotaxis, and
osteogenesis [342, 343]. Osteogenesis and angiogenesis are two closely correlated
processes during bone formation and regeneration [344]. Vascular endothelial growth
factor (VEGF) is highly expressed in vascularised tissues and can promote endothelial
cell proliferation, differentiation and migration from pre-existing vasculatures. As the
best characterised angiogenic mediator, VEGF is believed to play a crucial role in
skeletal development by enhancement of angiogenesis [345, 346]. On the other hand,
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a key feature regulating cell movement is chemotaxis, which induces migration via
signaling molecules termed chemokines [347, 348]. Chemokines are a large family of
small, basic polypeptides that have mainly been characterised as chemoattractants.
They navigate trafficking and localisation for various cell type in tissue compartments
[349]. The interaction between chemokine (C-X-C motif) ligand 12 (CXCL12) and its
receptor CXCR4 is believed to be the most important contributor to MSC homing to
areas of tissue damage and is involved in tissue regeneration [350-354]. However,
there is little information to be found regarding the potential effect of VEGF on the
expression pattern of CXCL12/CXCR4 with respect to the recruitment of MSCs
during new bone formation. We therefore investigated whether VEGF secretion by
osteogenically differentiated MSCs (OMSCs) was associated with CXCL12/CXCR4
expression alterations, which may lead to the migration and recruitment of MSCs in
the osteogenic process.
MATERIALS AND METHODS
Cell culture
Human bone marrow samples were obtained from patients (n=3) undergoing elective
knee replacement surgery in the Department of Orthopaedics at Prince Charles
Hospital with informed consent from each patient and ethics approval from the Ethics
Committee of Queensland University of Technology. Bone marrow was sourced from
the femoral canal. After draining of synovial fluid, a femoral drill hole was made
through the distal femur and an intramedullary rod was passed into the femoral canal
allowing marrow aspirates to be collected with a syringe. Bone marrow-derived
MSCs for this study were isolated by density gradient centrifugation over
Lymphoprep (Axis-shield PoC AS, Oslo, Norway) according to the manufacturer’s
protocol. The cells were then seeded in culture flasks in low glucose Dulbecco’s
Modified Eagle Medium (DMEM; Gibco®, Life Technologies Pty Ltd., Australia)
containing 10% (v/v) fetal bovine serum (FBS; In Vitro Technologies, Australia) and
1% (v/v) penicillin/streptomycin (P/S; Gibco®, Life Technologies Pty Ltd., Australia)
at 37°C, 5% CO2. The medium was changed twice weekly to wash out all non-
adherent cells. Adherent bone marrow-derived MSCs were then passaged once grown
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to 70%-80% confluence. Cell characterisation for MSCs has been carried out in our
previous study [318, 355].
Osteogenic differentiation of bone marrow-derived MSCs
Isolated MSCs were subjected to a 2-week osteogenic differentiation in osteogenic
medium (DMEM supplemented with 10% FBS, 1% penicillin/streptomycin, 10 mM
β-glycerophosphate, 50 µM ascorbic acid and 100 nM dexamethasone; Sigma Aldrich,
Australia) for further experiments. Medium was changed twice weekly.
Alizarin Red S staining
In order to identify mineralization nodules, Alizarin Red S staining was performed on
day 14 after MSCs and OMSCs had grown to a confluent monolayer in a 6-well plate.
The medium was removed and the cells were washed with PBS and fixed with 100%
methanol for 10 min at room temperature. After rinsing with distilled water, the cells
were stained in a solution of 1% Alizarin Red S (Sigma-Aldrich, Australia) at pH 4.1
for 20 min and the excessive solution was then washed away. The samples were air
dried and photos were taken under a microscope (Eclipse TS100, Nikon Australia Pty
Ltd.).
Preparation of conditioned medium (CM)
CM was obtained from 1×107
MSCs (CMMSC
) or OMSCs (CMOMSC
). The cells were
washed twice with PBS, and then incubated with 10 mL of serum-free DMEM at
37oC for 48 h. The CM was then collected, subjected to centrifugation at speed of
1000 g for 5 min at 4oC and filtration through a 0.2 µm low protein binding filter
(Millipore Corporation, Billerica, MA, USA) to remove cell debris. The supernatants
were stored at -80oC for subsequent experiments.
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Enzyme-linked immunosorbent assay
The concentration of VEGF in CMMSC
and CMOMSC
was tested using ELISA assay kit
for VEGF (R&D Systems Inc., Australia) according to the manufacture’s instruction.
Briefly, all the samples and standards were incubated in the assigned wells of the 96-
well plate for 2 h at room temperature. After three washings, 200 µL VEGF conjugate
was added into each well and incubated for 2 h at room temperature. After washing,
the amount of conjugate bound to each well was determined by the addition of 200 µL
substrate solution. The reaction was quenched by adding 50 µL stop solution per well
and measured immediately at 450 nm with a microplate reader (SpectraMax, Plus 384,
Molecular Devices, Inc., USA). The test was performed in triplicate, and the VEGF
levels were determined using the generated standard curve. All the results were
expressed as the amount (pg) of VEGF in per µL CM.
Indirect co-culture
The CM from MSCs or OMSCs was mixed with fresh culture medium in a ratio of
1:1 before applying to MSCs with or without the addition of 20 ng/mL VEGF
neutralizing antibody (R&D Systems Inc., Australia). The medium was changed every
three days. MSCs were harvested for RNA and protein on day 6 for subsequent
experiments.
In vitro migration assay
MSCs (5×104) were seeded on transwell inserts with 8 µm pore size (Becton-
Dickinson Labware, Franklin Lakes, NJ, USA). After 24 h of cell attachment, the
transwell inserts were assembled onto the 6-well companion plates cultured with
5×104 MSCs, which were subjected to osteogenic differentiation (OMSCs) or retained
in undifferentiated condition (MSCs) for 2 weeks prior to the migration assay. All co-
cultures were performed in serum-free medium with or without the addition of 20
ng/mL VEGF neutralizing antibody. Cell migration was allowed to proceed for 8 h.
Cells that had migrated to the other side of the insert membrane were fixed, and then
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stained with crystal violet. Images were taken with an inverted microscope (Eclipse
TS100, Nikon Australia Pty Ltd.). The migration was quantified by counting the
number of cells that migrated through the pores in six randomly selected fields on
each transwell insert membrane, and values from three independent experiments were
averaged and shown as mean ± SD.
xCELLigence system monitored cell migration
The real-time migration tendency of MSCs in response to CMMSC
and CMOMSC
was
further monitored using the CIM-Plate 16 and xCELLigence system RTCA DP
Instrument (Roche Products Pty Ltd., Australia). The CIM-Plate 16 is a 16-well
modified Boyden chamber composed of an upper chamber (UC) and a lower chamber
(LC). The UC is sealed at the bottom by a microporous polyethylene terepthalate
(PET) membrane. These micropores allow the physical translocation of cells from the
upper part of the UC to the bottom side of the membrane, which is covered with
interdigitated gold microelectrode sensors. As migrated cell attach and spread on the
electrode sensors, it leads to an increase in electrical impedance [356]. The LC
contains 16 wells, each of which serves as a reservoir for a chemoattractant solution.
In the present study, 100 µL of cell suspension (4×104 cells) were added to each well
of the UC after serum starvation for 4 h. The UC was then placed on the LC of the
CIM-Plate 16 containing 160 µL CMMSC
and CMOMSC
respectively, with or without
VEGF neutralizing antibody as an attractant. Cell migration was monitored over a
period of up to 12 h. The generated impedance signal was displayed as a
dimensionless parameter termed cell index, which was directly proportional to the
total area of the cell-culture well.
VEGF stimulation of MSCs
In order to further investigate the effect of VEGF on the expression pattern of
CXCL12/CXCR4, cell culture medium supplemented with 20, 40, 80 ng VEGF
(R&D Systems Inc., Australia) per mL was applied to the MSCs. The medium was
replenished every three days through a 6-day culture period, after which the cells were
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harvested and subjected to real time quantitative PCR (RT-qPCR) and Western
blotting analysis.
Fluorescent staining for morphological changes
2×103 MSCs were seeded on each coverslip (ProSciTech, QLD, Australia) placed in a
24-well plate. After overnight culture in serum-free medium, CMMSC
, and CMOMSC
with or without VEGF neutralizing antibody, the MSCs were fixed with 4%
paraformaldehyde (PFA) for 30 min. Fixed cells were permeabilized with 0.1% Triton
X-100 in PBS for 5 min and washed twice with 0.1% BSA/PBS. The nuclei were
stained with 0.1 µg/mL DAPI (Life Technologies Pty Ltd., Australia) for 30 min, and
the actin filaments were labelled with rhodamine phalloidin (Life Technologies Pty
Ltd., Australia). After washing, the coverslips were mounted with ProLong® Gold
(Life Technologies Pty Ltd., Australia) and secured to glass slides with three spots of
nail varnish. Morphological characteristics were identified under a fluorescent
microscope. The shapes of the fluorescent signals induced by the conditioned medium
were analysed using the integrated morphometry shape factor analysis of MetaMorph
image analysis software version 7.1.2 (Universal Imaging, Downingtown, PA, USA).
In the present study, we used a standard shape factor defined by (area/perimeter2) ×
4π for quantitative analysis of cell shape [357]. This algorithm assigns a value from 0
to 1, describing the shape of a perfect circle attains a value of 1 and a line is assigned
as 0.
RNA extraction, cDNA synthesis, and real time quantitative-PCR (RT-qPCR)
The mRNA expression of alkaline phosphatase (ALP), osteopontin (OPN) and
osteocalcin (OCN) and VEGF was measured in MSCs and OMSCs to detect the
difference before and after osteogenic differentiation. The expression of CXCL12 and
CXCR4 in MSCs was also measured after co-culture with different CM and VEGF
treatment. Total RNA was extracted from the cells with TRIzol Reagent (Ambion®,
Life Technologies Pty Ltd., Australia). Complementary DNA was synthesized from 1
μg of total RNA using DyNAmo™ cDNA Synthesis Kit (Finnzymes, Genesearch Pty
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Ltd., Australia) following the manufacturer’s protocol. The RT-qPCR primers (Table
1) were designed based on cDNA sequences from the NCBI Sequence database and
the primer specificity was confirmed by BLASTN searches. RT-qPCR was performed
on an ABI Prism 7300 Sequence Detection System (Applied Biosystems, Australia)
using SYBR® Green detection reagent (Applied Biosystems, Australia). In brief,
cDNA samples (2.5 μL of 1:10 diluted template for total volume of 25 μL per reaction)
were analysed for the genes of interest and for the GAPDH house keeping gene. The
cycling conditions were set as follows: 2 min at 95°C for activation followed by 45
cycles of denaturation at 95°C for 5 sec, annealing at 60°C for 10 sec, and extension
at 72°C for 15 sec. The specificity of amplification was verified by a single peak in
the melting curve. All reactions were run in triplicate for three independent
experiments. Dissociation curve analysis was performed to validate the absence of
primer dimmers and nonspecific PCR products. The mean cycle threshold (Ct) value
of each target gene was normalized against Ct value of GAPDH and the relative
expression calculated using the following formula: 2-(normalized average Cts)
×104 [358].
Table 1. Primer sequences for the gene observed in this study
Gene
name
Forward sequences Reverse sequences
ALP 5’TCAGAAGCTCAACACCAACG 5’TTGTACGTCTTGGAGAGGG
C
OCN 5’CACCGAGACACCATGAG 5’TGGAGAGGAGCAGAACTG
OPN 5’CCAAGTAAGTCCAACGAAAG 5’GGTGATGTCCTCGTCTGTA
VEGF 5’GCTGTCTTGGGTGCATTGG 5’GCAGCCTGGGACCACTTG
CXCL12 5’GCTACAGATGCCCATGCCGA
TTC
5’CCTGAATCCACTTTAGCTTC
GGGTC
CXCR4 5’CCAGAAGAAACTGAGAAGC
ATGACGG
5’CCAGACGCCAACATAGACC
ACC
GAPDH 5’ TCAGCAATGCCTCCTGCAC 5’ TCTGGGTGGCAGTGATGGC
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Protein extraction and Western blotting
Total protein was harvested by lysing the cells in a lysis buffer containing 20 mM
HEPES (pH 7.4), 10% glycerol, 1% Triton X-100, 2mM EGTA and a protease
inhibitor cocktail (Roche Products Pty. Ltd., Dee Why, NSW, Australia). The protein
concentration was determined by the BCA Protein Assay Kit (Thermo Fisher
Scientific, VIC, Australia). 20 µg of protein from each sample was separated by 8%
SDS-PAGE gels. The protein was then transferred onto a nitrocellulose membrane
(Pall Corporation, East Hills, NY, USA). After blocking for 1 h with blocking buffer
(1×Tris-buffered saline with 5% skim milk and 0.01% Tween-20), the membranes
were probed with primary antibodies against VEGF (1:1000 rabbit anti-human),
CXCL12 (1:1000, rabbit anti-human), CXCR4 (1:1000, rabbit anti-human) and α-
Tubulin (1:5000, rabbit anti-human) overnight at 4°C. The primary antibodies were
all obtained from Abcam (Sapphire Bioscience Pty. Ltd., Waterloo, NSW, Australia).
The membranes were washed three times for 20 min each in TBS containing 0.01%
Tween-20, and then incubated with horseradish peroxidase (HRP)-conjugated goat
anti-rabbit secondary antibody (Abcam; Sapphire Bioscience Pty. Ltd., Waterloo,
NSW, Australia) at 1:2000 dilution for 1 h at room temperature. Targeted proteins
were visualized using the SuperSignal West Femto Maximum Sensitivity Substrate
(Thermo Fisher Scientific, Australia) and exposed on x-ray film (Fujifilm, Australia).
Bands intensities were quantified by scanning densitometry and analysed using
ImageJ software [359].
Animal study and drug administration
Collagen scaffolds were fabricated from a 1mm-thick bovine Type-I collagen sheet
(Medtronic Australia Pty Ltd, North Ryde, NSW, Australia) using a hole puncher and
sterilized by soaking in 75% ethanol, followed by three rinses of PBS prior to cell
seeding. For the in vitro osteogenic differentiation, 6×104 MSCs were firstly seeded
onto collagen membrane and cultured in osteogenic medium for 2 weeks.
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Twelve 5-week old male SCID mice (Animal Resources Centre, Canning Vale, WA,
Australia) were used for the skull defect model [78, 321] and subcutaneous cell
implantation, respectively. For the skull defect implantation, an incision was made
longitudinally along the centre line of the shaved skull skin after anaesthesia. A 3 mm
defect was then made through the skull using a dental bur. The collagen membrane
carrying OMSCs were inserted into the defects of each mice. For the subcutaneous
cell implantation, a longitudinal incision was made in each animal along the central
line of the shaved dorsal. Each individual pocket received one collagen scaffold
carrying OMSCs. Six mice were assigned to experimental group using a complete
randomized design, which received local injection of 20 ng VEGF neutralizing
antibody in 100 uL PBS for two weeks. The other six mice in the control group
received 100 uL of PBS. All of the SCID mice were sacrificed after four weeks and
the implants were retrieved.
Micro-computed tomography (μCT) assessment
All the samples harvested were fixed in 4% PFA overnight at room temperature and
then washed in PBS. Cross-sectional scans parallel to the cell seeding surface of the
implants were performed in a micro-CT scanner (µCT40, SCANCO Medical AG,
Brüttisellen, Switzerland) at a resolution of 12 µm and three-dimensional (3D) images
of the implants were reconstructed. The calcified area inside the implant on each
scanned slice was circled out and the volume of calcification within the implants was
measured using the inbuilt scanner software package and recorded for statistical
analysis.
Histology and immunohistochemistry (IHC)
All harvested subcutaneous samples were embedded in paraffin block for slicing,
whereas the skull defect samples were firstly decalcified in 10% EDTA solution for
four weeks and then embedded in paraffin. 5 µm serial slices were cut from the
embedded samples using a microtome (Leica Microsystems GmbH, Wetzlar,
Germany). Only slices close to the centre of the samples were used for the staining.
66 | P a g e
In order to observe the calcification in subcutaneous implants, sample was subjected
to von Kossa with H&E staining. The slices were dewaxed in xylene and then
hydrated in 100%, 90%, and 70% ethanol for 3 min each. After the application of 5%
silver nitrate solution (Sigma-Aldrich, Australia), the slices were exposed to light for
1 h. After rinsing in distilled water, 5% sodium thiosulphate was added to the slices
for 5 min. The slices were then washed in tap water, placed in Harris’s haematoxylin
(Sigma-Aldrich, Australia) for 2 min then rinsed in tap water before dipping 3 times
in acid ethanol. After dehydration, eosin (HD Scientific Supplies Pty Ltd., NSW,
Australia) was applied to the slices for 15 sec. The slices were then rinsed in 100%
ethanol twice and cleared in xylene before being mounted with cover slips.
IHC staining was performed for assessment of new bone formation in the skull
defects. Briefly, endogenous peroxidase activity was quenched by incubating with 3%
H2O2. After washing with PBS, all the sections were blocked with 10% swine serum
for 1 h. Sections were then incubated with optimal dilution of primary antibodies
against ALP (1:400, anti-human, Sigma-Aldrich, Australia) overnight at 4°C. Sections
were washed free of primary antibodies and then incubated with a biotinylated
secondary antibody (Dako, CA, USA) for 15 min, followed by 15 min incubation with
HRP-conjugated streptavidin-biotin complex (Dako, CA, USA). Antibody complexes
were visualised after the addition of a buffered diaminobenzidine (DAB) for 3 min.
In situ hybridisation (ISH)
To verify the origin of cells in skull defect sites, ISH was performed using a human
specific digoxigenin (DIG)-labelled oligonucleotide probe (PanPath, Budel, the
Netherlands), which detects Alu repeat sequences found in human chromosomes.
Briefly, tissue sections were dewaxed and rehydrated through a descending ethanol
series with RNase-free reagents, and then treated with proteinase K for 10 min. One
drop of the ready-to-use Alu probe solution was added onto each section and the
slices were covered with a coverslip. After heating at 95oC for 5 min, the slices were
incubated at 37°C in a humidified chamber overnight followed by post-hybridisation
67 | P a g e
washing procedure. 3-Amino-9-ethylcarbazole (AEC) was then applied to visualise
the bound probe. After rinsing in distilled water, the sections were lightly
counterstained with haematoxylin, then mounted with Crystal MountTM
aqueous
mounting medium (Sigma-Aldrich, Australia).
For ISH and IHC double staining, after the donor human cells were detected using the
Alu DNA probe, which was visualised with NiCl2+DAB (staining colour is black),
the first primary antibody (VEGF) was applied and visualised with DAB (staining
colour is brown). The host mouse cells were then stained with Nuclear Fast Red
(Sigma-Aldrich, Australia). After rinsing with tap water, the sections were dehydrated
with ascending concentrations of ethanol, cleared with xylene and mounted with a
cover slip.
Statistical analysis
Statistical evaluation was performed using SPSS software (SPSS, Sigma Stat, Erkrath,
Germany). Differences between two groups were assessed by Student t-test.
Differences in the treatment outcomes from different conditions were assessed by
analysis of variance (ANOVA) with Bonferroni post-hoc test. Values of p<0.05 were
considered statistically significant.
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RESULTS
OMSCs express high level of VEGF
Histology images showed a close interaction between donor human OMSCs and host
mouse cells in the newly formed bone tissues from the skull defect model (Figure 14).
Furthermore, the ISH together with IHC staining revealed that the donor OMSCs
strongly expressed VEGF, giving a clue that VEGF secreted by donor OMSCs may
play a role in the host cell recruitment (Figure 14A and B).
Figure 14. Histological evaluation of a skull defect model after a 4-week implantation
of OMSCs. (A) Representative image showing host mouse cells were recruited to the bone defect
area and involved in the new bone formation with donor human cells (20×). (B) 40× magnification of
the selected area, showing the VEGF secretion from OMSCs, as well as the close interaction between
donor cells and host cells. Host mouse cells were stained with Nuclear Fast Red (pink colour, dashed
arrow pointed); Donor human cells were stained with NiCl2+DAB (black colour, arrow pointed);
VEGF was stained with DAB (brown colour, arrowhead pointed). NB: newly formed bone.
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Alizarin Red S staining was performed and the result showed that the mineralization
in MSCs was significantly enhanced after osteogenic differentiation (Figure 15A).
The mRNA level of ALP, OPN and OCN also showed significant increase in OMSCs
(Figure 15B, p<0.05). VEGF gene expression of OMSCs increased remarkably
(Figure 15C, p<0.05). ELISA result showed that VEGF secretion was significantly
elevated in MSCs after exposure to osteogenic medium (Figure 15D, p<0.01).
Western blotting analysis further confirmed the elevated VEGF protein level (Figure
15E, p<0.05).
Figure 15. VEGF secretion from OMSCs. (A) Alizarin Red S staining showed the
mineralization in MSCs was significantly enhanced after cultured in osteogenic medium for 2 weeks.
(B) mRNA transcripts of ALP, OCN and OPN, showed a significant increase in OMSCs (*p<0.05). (C)
VEGF gene expression of MSCs increased remarkably after osteogenic differentiation (*p<0.05). (D)
ELISA result showed that VEGF secretion significantly elevated in OMSCs (**p<0.01). (E) and (F)
Western blotting analysis showed greater VEGF expression in OMSCs in the protein level, as indicated
by the significantly higher band density (*p<0.05).
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VEGF secreted by OMSCs induced the migration of MSCs
After 8 h incubation, the migration of MSCs through the transwell membrane was
significantly enhanced in the co-culture with OMSCs compared with the serum-free
medium or the co-culture with MSCs (Figure 16C, D and I, p<0.01). However, the
number of migrated MSCs dropped dramatically when the OMSCs were treated with
VEGF neutralizing antibody (Figure 16D and J, p<0.01). The increased motility of
MSCs toward OMSCs was further confirmed using the xCELLigence system. The
cell index of MSCs in CMOMSC
was elevated throughout the experimental period, but
dropped down to the level as that in the CMMSC
when VEGF neutralizing antibody
was applied (Figure 16K, p<0.05). The data demonstrated that VEGF secreted by
OMSCs serves as a chemoattractant to MSCs.
Morphological changes of MSCs cultured in CMOMSC
As described above, enhanced migration of MSCs was observed in the indirect co-
culture with OMSCs, indicating a more mobile MSC phenotype was induced. We
then further investigated the effect of CMOMSC
on the morphological and cytoskeletal
changes in MSCs. Figure 17 shows MSCs cultured in CMOMSC
underwent remarkable
morphological alterations compared with those cultured in CMMSC
over the same
period of time. Actin filaments organized along the length of the cell, making the
general appearance of cell as an elongated rod shape (Figure 17B). However, the
application of VEGF neutralizing antibody in CMOMSC
led to a reversed effect (Figure
17C). Cell shape factor changed from around 0.5 to 0.1, indicating a transformation
from a more regular and round shape to an irregular and elongated cell shape when
MSCs were grown in the CMOMSC
instead of CM
MSC. Additionally, when VEGF
neutralizing antibody was added into the CMOMSC
, the cells tended to regain a higher
shape factor, indicative of the importance of VEGF in maintaining the migratory
phenotype (Figure 17D, p<0.05).
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Figure 16. VEGF neutralizing antibody interfered with the migration of MSCs
towards OMSCs. (A)-(D) and (I) The migration capacity of MSCs towards OMSCs was
significantly superior to the control groups (SF, serum-free medium; MSC, undifferentiated
mesenchymal stromal cells; OMSC, osteogenically differentiated mesenchymal stromal cells;
**p<0.01). (A)-(D) and (J) The number of migrated MSCs decreased significantly with the addition of
20 ng/mL VEGF neutralizing antibody (*p<0.05). (K) The increased motility of MSCs toward OMSCs
was further confirmed by the xCELLigence system (*p<0.05).
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Figure 17. Morphological and cytoskeletal changes in MSCs cultured in CMOMSC
with or without the addition of VEGF neutralizing antibody. (A)-(C) MSCs cultured in
CMOMSC showed elongated actin filaments. The application of VEGF neutralizing antibody in CMOMSC
led to a reversed effect. (D) The shape factor of MSCs cultured in CMOMSC was significantly lower than
that in CMMSC and CMOMSC + Abvegf (*p<0.05).
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CMOMSC
increased the CXCL12/CXCR4 expression of MSCs
We then further tested the effect of VEGF secreted by OMSCs on the expression
pattern of CXCL12/CXCR4, which is believed to be the most important axis that
contributes to MSCs homing to areas of tissue damage. When MSCs were cultured in
CMOMSC
, the CXCL12/CXCR4 mRNA expression showed a significant elevation
(Figure 18A and B, p<0.05) compared to those cultured in CMMSC
. Furthermore, the
expression of CXCL12/CXCR4 decreased when VEGF neutralizing antibody was
added into CMOMSC
(Figure 18A and B, p<0.05). The change of CXCL12/CXCR4
expression was further confirmed by the Western blotting results (Figure 18C-E,
p<0.05).
Figure 18. CMOMSC
increased the CXCL12/CXCR4 expression of MSCs. (A) and (B)
mRNA levels of CXCL12 and CXCR4 were significantly elevated in MSCs cultured in CMOMSC
(*p<0.05). However, the expression of CXCL12/CXCR4 decreased when VEGF neutralizing antibody
was applied (*p<0.05). (C)-(E) The elevated expression of CXCL12/CXCR4 was further confirmed by
the Western blotting analysis on the protein level, as indicated by the significantly higher band density
(*p<0.05). CMMSC
, conditioned medium derived from undifferentiated mesenchymal stromal cells;
CMOMSC, conditioned medium derived from osteogenically differentiated mesenchymal stromal cells;
Abvegf, VEGF neutralizing antibody.
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CXCL12/CXCR4 expression in MSCs was stimulated by VEGF
The effect of VEGF on the expression of CXCL12/CXCR4 in MSCs was further
investigated by culturing the MSCs in medium supplemented with increasing
concentration of VEGF. The results showed that the CXCL12/CXCR4 gene
expression in MSCs enhanced significantly when stimulated by VEGF in a dose
dependant manner (Figure 19A and B, p<0.05). Western blotting analysis showed a
similar trend on the protein level (Figure 19C-E, p<0.05).
Figure 19. CXCL12/CXCR4 expression of MSCs stimulated by VEGF. (A) and (B)
mRNA levels of CXCL12 and CXCR4 in MSCs treated with different dosage of VEGF, showing a
steady increase in a dose dependent manner (*p<0.05). (C)-(E) The elevated expression of
CXCL12/CXCR4 in MSCs stimulated with increasing dosage of VEGF was further confirmed by the
Western blotting analysis on the protein level, as indicated by the significantly higher band density
(*p<0.05).
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Inhibition of VEGF abrogated the effect of OMSCs on the recruitment of host cells
To determine whether VEGF secreted by OMSCs plays a role in host cell recruitment
in vivo, donor cells were implanted into skull defects of SCID mice with or without
the local administration of VEGF neutralizing antibody. The von Kossa with H&E
staining revealed the mineralization found in the subcutaneous implants carrying
OMSCs was suppressed when the VEGF neutralizing antibody was applied (Figure
20A and B). For the skull defect samples, µCT scanning and 3D reconstruction
confirmed that VEGF neutralizing antibody significantly inhibited the bone healing
induced by the implantation of the OMSCs. (Figure 20E and F). IHC staining against
ALP revealed less new bone tissues in the skull defects after the local application of
VEGF neutralizing antibody (Figure 20C and D). The results suggested that VEGF
was involved in the accelerated bone healing induced by OMSC implantation. ISH
against human Alu sequences revealed that host mouse cells (Alu negative) were
recruited to the skull defect area and actively involved in the bone regeneration
process together with the donor human OMSCs (Figure 20G). On the contrary, fewer
host cells were found in the implants when VEGF was neutralized (Figure 20H).
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Figure 20. Blocking VEGF with neutralizing antibody resulted in a significant
decrease of MSC recruitment and new bone formation. (A) von Kossa with H&E staining
showed mineralization in the subcutaneous implants carrying OMSCs. (B) von Kossa with H&E
staining detected no mineralization when the VEGF neutralizing antibody was applied. (C)
Immunohistochemical staining against ALP revealed newly formed bone in skull defect implanted with
collagen scaffold carrying OMSCs. (D) Less new bone formation was detected in the skull defects after
the local application of VEGF neutralizing antibody. (E) and (F) 3D images of skull defects implanted
with collagen scaffold carrying OMSC, with or without the application of VEGF neutralizing antibody.
(G) ISH staining revealed that host mouse cells (stained in purple) were recruited to the skull defect
area and involved in the bone regeneration together with the donor human OMSCs (stained in red). (H)
Fewer host cells were recruited to the implants when VEGF was neutralized.
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DISCUSSION
Cell implantation is currently regarded as an important and promising regime for
repairing and restoring the biological functions of bone defects [307, 308]. Bone
marrow-derived MSCs are promising candidates for such cell-based therapy, due to
their immune tolerance, self-renewal capacity and ease of use [360-362]. Chemotactic
migration of host mesenchymal progenitor cells (MPCs) from bone marrow or other
tissues have been identified as a crucial event during various physiological and
pathological processes [351, 353, 354]. Increasing evidence indicates that MSCs
communicate with other cells in vivo and appear to “home” to areas of defect and
injury in response to cellular signals. However, even though the therapeutic potential
of MSCs is an area of great excitement and promise, the factors that trigger MSCs’
response and the underlying mechanisms that guide the host MSCs migrate to the
bone defect sites are still poorly understood. In this study we have demonstrated that
VEGF secreted by OMSCs led to the altered expression pattern of CXCL12/CXCR4
in MSCs, and we have also provided some insights into the communication between
host cells and donor cells during osteogenesis.
MSCs secrete numerous cytokines including VEGF, bFGF, IGF-1 and HGF [363].
Among all these, VEGF is the key regulator in a cascade of molecular and cellular
events that ultimately lead to the development of the vascular system [364]. VEGF is
not only a critical mediator in physiological angiogenesis, but also a vital factor in
skeletal growth [365]. Previous studies suggested that VEGF plays an important role
in MSCs migration [366-368], as well as in the recruitment and proliferation of
endothelial cells (ECs) [369]. The data from our study showed that the VEGF
secretion increased dramatically in bone marrow-derived MSCs after osteogenic
differentiation in vitro. Similar results have been reported previously that VEGF is
expressed in osteoblast-like cells in a differentiation-dependent manner, indicating
that VEGF plays a positive role in the regulation of osteoblast activity [199]. A recent
study has also demonstrated that Osterix (Osx), an essential osteoblast-specific
transcription factor required for osteoblast differentiation, directly targets VEGF
expression [201], suggesting a close link between osteogenesis and angiogenesis. The
enhanced secretion of VEGF from OMSCs was further confirmed by the ISH and
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IHC staining, which showed a strong expression of VEGF in OMSCs correlated with
greater recruitment of host MSCs.
We then demonstrated the positive effect of VEGF on the motility of MSCs in vitro
using transwell migration assay and real-time xCELLigence system. The results
suggested the secreted factors derived from OMSCs were a potent chemoattractant for
MSCs. The enhanced motility of MSCs was correlated with the changes in actin
organization and distribution from azimuthal symmetry around the cell rim to
concentration at a particular region [370]. In addition to morphological changes,
MSCs underwent specific alterations in gene expression patterns in response to the
exterior stimuli. Exposure to CMOMSC
enhanced migration of MSCs and this was
accompanied by an increased expression pattern of CXCL12/CXCR4. In agreement
with the gene expression profile, CXCL12/CXCR4 protein levels also increased in
MSCs upon exposure to CMOMSC
. However, the addition of VEGF neutralizing
antibody abrogated the effect of OMSCs on the actin filament arrangement as well as
the CXCL12/CXCR4 expression of MSCs. We further confirmed a dose-dependent
chemoattractive effect of VEGF on MSCs. VEGF stimulation remarkably increased
the expression of CXCL12/CXCR4, which is the best known axis that contributes to
cell migration [352]. Additionally, the expression of CXCL12/CXCR4 in MSCs was
down regulated when VEGF neutralizing antibody was applied, suggesting the
secretion of VEGF from OMSCs played a crucial role in inducing the MSCs
recruitment.
The interaction between donor cells and host cells is an emerging area of research,
and it is important to understand the mechanisms that govern this interaction. Our data
has shown that there was differential regulation of CXCL12/CXCR4 in MSCs
exposed to different microenvironments and that these changes influenced chemotaxis,
an important function of MSCs. Based on the previous finding, our in vivo study
further confirmed that OMSCs played a significant role in the host cells recruitment.
It is very interesting to find that the transplanted OMSCs were involved in the new
bone formation. Furthermore, the function of OMSCs in host cell recruitment was
79 | P a g e
supported by the elevated secretion of VEGF, which then led to the markedly
increased expression of CXCL12/CXCR4 and changes of morphology in host MSCs
(Figure 21). CXCL12 may signal through its receptor, CXCR4, and function in an
autocrine manner to support chemotaxis of host cells towards the transplanted donor
cells in the bone defect sites. The newly formed blood vessels, donor cell- and host
cell-derived osteoblasts may work together to restore the damaged bone tissue.
Further studies are needed to elucidate the full repertoire of CXCL12/CXCR4
signaling in MSCs and to identify other pathways that are critical to the
microenvironment-specific MSC biology. This will be crucial to the understanding of
clinically important processes, such as cell transplantation, bone marrow engraftment
and wound healing.
Figure 21. Schematic of the proposed mechanism by which donor OMSCs initiate a
paracrine signaling cascade that results in altered phenotype and enhanced motility of
host MSCs in a skull defect model. The interaction between OMSCs and MSCs is centrally
regulated by VEGF secreted by OMSCs, and involves the participation of the CXCL12/CXCR4 axis,
which may accelerate the host cell recruitment process.
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CONCLUSION
We demonstrated the VEGF secreted by OMSCs plays a significant role in the
expression pattern of CXCL12/CXCR4 of undifferentiated bone marrow-derived
MSCs. The involvement of CXCL12/CXCR4 in the VEGF-mediated host cell
recruitment towards skull defect was confirmed for the first time. The present study
offers insights into the donor/host cell interaction and is of clinical significance for the
application of cell-based therapy for bone repair and regeneration.
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REPORT THREE
Involvement of VEGF/PDGFR and the PI3K/Akt signaling pathway in
OMSCs-induced MSCs migration
Yinghong Zhou, Ross Crawford, Yin Xiao
Suggested Statement of Contribution of Co-Authors for Thesis by
Published Papers:
Contributors Statement of contribution*
Yinghong Zhou Involved in the conception and design of the project, performed
laboratory experiments, data analysis and interpretation. Wrote
the manuscript.
Ross Crawford Assisted in collection of clinical samples and reviewed the
manuscript.
Yin Xiao Involved in the conception and design of the project, and
reviewed the manuscript.
Principal Supervisor Confirmation
I have sighted email or other correspondence from all co-authors confirming
their certifying authorship.
Name Signature Date
82 | P a g e
ABSTRACT
Background: Although it has been shown that mesenchymal stromal cells (MSCs)
are attracted to vascular endothelial growth factor (VEGF) secreted by osteogenically
differentiated MSCs (OMSCs) in our previous study, it is still not known that how
VEGF signals through MSCs which are expected to have low levels of VEGF
receptors (VEGFRs) and what downstream pathways are involved in the recruitment
of MSCs.
Methods: MSCs were stimulated with different concentrations of VEGF, conditioned
medium derived from MSCs (CMMSC
) or OMSCs (CMOMSC
) to establish the receptors’
activation profile. The role of platelet-derived growth factor receptors (PDGFRs) was
further investigated using phospho-receptor tyrosine kinase (RTK) array, Western
blotting analysis and in vitro migration assay. The involvement of the PI3K/Akt
signaling pathway in OMSCs-induced MSC migration was monitored by relevant
phosphorylated antibodies and inhibitors.
Results: We found that VEGF secreted by OMSCs was capable of inducing migration
of MSCs which are VEGFR deficient, and this induction was mediated through a
molecular crosstalk of PDGFRs and the PI3K/Akt signaling kinase. The VEGF-
induced MSC migration paralleled with the enhanced expression of PDGFRs and the
activation of the PI3K/Akt kinase. Blocking of PDGFRs activity impaired VEGF-
induced MSC migration as well as PI3K/Akt kinase activation. Moreover, a PI3K/Akt
specific inhibitor, LY294002, resulted in the suppression of VEGF-induced MSC
migration and dephosphorylation of downstream effectors of the PI3K/Akt signaling
pathway, including Akt, mTOR, p70S6K
and PAK1, further supporting the role of the
PI3K/Akt pathway in the MSC recruitment.
Conclusions: The present study established a mechanistic link among VEGF,
PDGFRs and PI3K/Akt in the regulation of MSC recruitment that eventually may be
beneficial to the bone repair and regeneration.
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INTRODUCTION
The molecular interactions between donor and host cells are indispensable in the
process of host mesenchymal stromal cells (MSCs) recruitment for tissue repair and
regeneration. Guidance of cell migration depends critically on subcellularly localised
perception and transduction of various signals. As has been widely reported, vascular
endothelial growth factor (VEGF) proved to be chemoattractive for primary human
osteoblasts [202, 203]. This provides the first evidence that VEGF may also be
involved in local recruitment of osteoprogenitor cells in the course of endochondral
bone formation or remodeling. VEGF exerts its effects on vascular cells by binding to
three structurally related VEGF receptor (VEGFR) tyrosine kinases, VEGFR-1,
VEGFR-2 and VEGFR-3, each forms homodimers on ligand binding [206]. VEGF
and platelet-derived growth factor (PDGF) are closely related members, as evidenced
by sequence analysis which predicts that VEGF and PDGF evolved from a common
ancestor [207]. VEGFRs and PDGF receptors (PDGFRs) are also structurally related,
characterised by extracellular immunoglobulin-like domains with an intracellular
tyrosine kinase domain interrupted by a non-catalytic region [371]. Bone marrow-
derived MSCs, which can differentiate into vascular cells, may be recruited by VEGF
during the process of angiogenesis and osteogenesis. However, several studies have
reported that there is limited expression of VEGFR in MSCs [200, 372], leading to the
speculation that VEGF may signal through PDGFRs and regulate the migration of
MSCs [210].
Our previous study (REPORT TWO) has demonstrated that VEGF secreted from
osteogenically differentiated mesenchymal stromal cells (OMSCs) played a
significant role in the recruitment of host cells by regulating the expression pattern of
CXCL12/CXCR4, an important axis for cell migration. The present study aimed to
identify the downstream signaling pathways associated with the interaction between
OMSCs and MSCs. Recent studies have reported that binding of CXCL12 to CXCR4
may lead to stimulation of signaling cascade that involves activation of multiple
targets, including Erk1/2, p38 and Akt [273-276]. Akt, also known as protein kinase B
(PKB), is one of the best characterised targets of phosphoinositide 3-kinases (PI3K)
signaling pathway, which regulates cell survival, cytoskeletal rearrangements and cell
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invasion [252, 253]. Accumulating data suggest that major downstream effectors of
the PI3K/Akt signaling pathway, mTOR, p70 S6 kinase (p70S6K
) and p21-activated
kinase 1 (PAK1), are associated with changes in actin cytoskeleton organization, cell
shape and adhesion dynamics, and play critical roles in cell motility regulation [281,
282, 285-287].
Here in the present study, we found that OMSCs stimulated the migration of MSCs by
activating PDGFRs. Furthermore, our findings demonstrated VEGF/PDGFR mediated
the activation of the PI3K/Akt signaling cascade, which led to the phosphorylation of
mTOR, p70S6K
and PAK1, and was significantly associated with the cell migration
induced by OMSCs.
MATERIALS AND METHODS
Cells and Cell culture
Human bone marrow samples were collected from patients (n=3, 1 female and 2
males) undergoing elective knee replacement surgery in the Department of
Orthopaedics at Prince Charles Hospital, Brisbane, Queensland, Australia after
obtaining informed consent from each donor and ethics approval from the Ethics
Committee of Queensland University of Technology. Bone marrow-derived MSCs for
this study were isolated by density gradient centrifugation over Lymphoprep (Axis-
shield PoC AS, Oslo, Norway) according to the manufacturer’s protocol. The cells
were then seeded in culture flasks supplemented with low glucose Dulbecco’s
Modified Eagle Medium (DMEM; Gibco®, Life Technologies Pty Ltd., Australia)
containing 10% (v/v) fetal bovine serum (FBS; In Vitro Technologies, Australia) and
1% (v/v) penicillin/streptomycin (P/S; Gibco®, Life Technologies Pty Ltd., Australia)
at 37°C, 5% CO2. The medium was changed twice weekly to wash out all non-
adherent cells. Adherent bone marrow-derived MSCs were then passaged once grown
to 70%-80% confluence. Cell characterisation for MSCs has been carried out in our
previous study [318, 355]. Human umbilical vein endothelial cells (HUVECs;
Clonetics™, Lonza Australia Pty Ltd., Mt Waverley, VIC, Australia) were used as
85 | P a g e
positive control when required. HUVECs were routinely cultured in EGM™-2
BulletKits™ (Lonza Australia Pty Ltd., Mt Waverley, VIC, Australia) composed of
endothelial cell basal medium-2 (EBM-2) supplemented with 2% FBS,
gentamicin/amphotericin-B, VEGF, human epidermal growth factor (hFGF)-B,
insulin-like growth factor-1 (IGF-1), hEGF, ascorbic acid, hydrocortisone, and
heparin, as described by the manufacturer (Lonza Australia Pty Ltd., Mt Waverley,
VIC, Australia).
Osteogenic differentiation of bone marrow-derived MSCs
Isolated MSCs were subjected to osteogenic differentiation for 2 weeks in osteogenic
medium (DMEM supplemented with 10% FBS, 1% penicillin/streptomycin, 10 mM
β-glycerophosphate, 50 µM ascorbic acid and 100 nM dexamethasone; Sigma Aldrich,
Australia) for further experiments.
Preparation of conditioned medium (CM)
CM was obtained from 1×107
MSCs (CMMSC
) or OMSCs (CMOMSC
). The cells were
washed twice with PBS, and incubated with 10 mL of serum-free DMEM at 37 oC for
48 h. The CM was then collected, subjected to centrifugation at speed of 1000 g for 5
min at 4oC and filtration through a 0.2 µm low protein binding filter (Millipore
Corporation, Billerica, MA, USA) to remove cell debris. The supernatants were stored
at -80oC for subsequent experiments.
Stimulation of MSCs
In order to investigate the effect of VEGF on the expression pattern of VEGFRs and
PDGFRs, cell culture medium supplemented with 20, 40, 80 ng VEGF (R&D
Systems Inc., Australia) per mL was applied to HUVECs and MSCs. The cells were
harvested and subjected to real time quantitative PCR (RT-qPCR) analysis after 24 h.
MSCs were also stimulated with serum-free medium, CMMSC
, CMOMSC
with or
86 | P a g e
without the supplementation of 20 ng/mL VEGF neutralizing antibody (R&D Systems
Inc., Australia) to further investigate the expression pattern of various receptors.
For the functional tests, MSCs were pre-treated with 5 µM PDGFR tyrosine kinase
inhibitor (Merck Pty Ltd., Australia), 10 µM LY294002 (Cell Signaling Technology,
Inc. Danvers, MA, USA) or DMSO as control 30 min prior to the stimulation of
CMMSC
and CMOMSC
. MSCs were harvested for RNA and protein at different time
points (e.g. 15 min, 30 min and 60 min) according to subsequent experiments.
In vitro migration assay using transwell chambers
MSCs migration was monitored using 6-well transwell cell culture Boyden chambers
with 23.1 mm effective diameter polyethylene terephthalate (PET) membrane
containing 8 µm pores (Becton-Dickinson Labware, Franklin Lakes, NJ, USA). 5×104
MSCs were firstly seeded on to each transwell chamber for the initial attachment.
After serum-starved overnight, the transwell chambers were assembled onto the 6-
well companion plates supplemented with CM from MSCs or OMSCs. To attenuate
the PDGFR in MSCs, 5 µM PDGFR tyrosine kinase inhibitor was applied. Non-
specific effects of the inhibitor were ruled out by repeating the migration assay in the
presence of an unrelated antibody against an intracellular antigen IgG. Cell migration
was allowed to proceed for 8 h. Cells that had migrated to the other side of the
transwell membrane were fixed, and stained with crystal violet. Images were taken
with an inverted microscope (Eclipse TS100, Nikon Australia Pty Ltd.). The
migration was quantified by counting the number of cells that migrated through the
pores in six randomly selected fields on each transwell membrane, and values from
three independent experiments, with triplicate samples in each experiment, were
averaged and shown as mean ± SD.
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Phospho-receptor tyrosine kinase (RTK) array
A human phospho-receptor tyrosine kinase (RTK) array kit (R&D Systems Inc.,
Australia) was used to simultaneously detect the relative tyrosine phosphorylation
levels of 49 different RTKs in the untreated or CM-treated MSC lysates. Each array
contains duplicate validated controls and captures antibodies for specific RTKs.
MSCs were serum-starved for 12 h and then stimulated with CMMSC
or CMOMSC
for
15 min at 37°C in a humidified atmosphere with 5% CO2, then immediately placed on
ice, washed twice with chilled PBS, and harvested using the lysis buffer that comes
with the RTK array kit. Total protein concentration was quantitated using the BCA
Protein Assay Kit (Thermo Fisher Scientific, VIC, Australia). RTK array analysis was
performed according to the manufacturer’s protocol. In brief, the array membranes
were blocked, and then incubated with 250 µg MSC lysate overnight at 4°C. After
washing, the membranes were incubated with anti-phospho-tyrosine-HRP for 2 h at
room temperature, then washed again, and developed with detection reagent. RTK
spots were visualized on x-ray film (Fujifilm, Australia).
RNA extraction, cDNA synthesis, and real time quantitative-PCR (RT-qPCR)
Total RNA was extracted from the cells with TRIzol® Reagent (Ambion
®, Life
Technologies Pty Ltd., Australia). Complementary DNA was synthesized from 1 μg
of total RNA using DyNAmo™ cDNA Synthesis Kit (Finnzymes, Genesearch Pty
Ltd., Australia) following the manufacturer’s protocol. The RT-qPCR primers (Table
2) were designed based on cDNA sequences from the NCBI Sequence database and
the primer specificity was confirmed by BLASTN searches. RT-qPCR was performed
on an ABI Prism 7300 Sequence Detection System (Applied Biosystems, Australia)
using SYBR® Green detection reagent Applied Biosystems, Australia). In brief,
cDNA samples (2.5 μL of 1:10 diluted template for total volume of 25 μL per reaction)
were analysed for the genes of interest and for the GAPDH house keeping gene. All
reactions were run in triplicate for three independent experiments. Dissociation curve
analysis was performed to validate the absence of primer dimmers and nonspecific
PCR products. The mean cycle threshold (Ct) value of each target gene was
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normalized against Ct value of GAPDH and the relative expression calculated using
the following formula: 2-(normalized average Cts)
×104 [358].
Table 2. Primer sequences for the gene observed in this study
Gene
name
Forward sequences Reverse sequences
VEGFR-1 5’CCCTTATGATGCCAGCAAGT
G
5’CCAAAAGCCCCTCTTCCAA
VEGFR-2 5’CAAACGCTGACATGTACGG
TCTA
5’CCAACTGCCAATACCAGTG
GAT
VEGFR-3 5’GTCAGGGGAGATCATCGGG
AC
5’GGAGCTGGTGGTGAATGTG
CCC
PDGFR-α 5’GATAGCTTCCTGAGCCACCA
C
5’CATAGAGTGATCTCTGGAT
GTCGG
PDGFR-β 5’GGAGACCCGGTATGTGTCA
GAGC
5’CCAGCACTCGGACAGGGAC
ATTG
GAPDH 5’ TCAGCAATGCCTCCTGCAC 5’TCTGGGTGGCAGTGATGGC
Protein extraction and Western blotting
Total protein was harvested by lysing the cells in a lysis buffer containing 20 mM
HEPES (pH 7.4), 10% glycerol, 1% Triton X-100, 2mM EGTA and a
phosphatase/protease inhibitor cocktail (Roche Products Pty. Ltd., Dee Why, NSW,
Australia). The protein concentration was determined by the BCA Protein Assay Kit
(Thermo Fisher Scientific, VIC, Australia). 15 µg of protein from each sample was
separated by 8% SDS-PAGE gels, and then the separated protein was transferred to
nitrocellulose membrane (Pall Corporation, East Hills, NY, USA). After blocking for
1 h with Odyssey® blocking buffer (Millennium Science, Australia), the membranes
were incubated with the primary antibodies against PDGFR-α (1:1000, rabbit anti-
human), PDGFR-β (1:1000, rabbit anti-human), Akt (1:2000, mouse anti-human),
phospho-Akt (Ser473) (1:2000, rabbit anti-human), Erk1/2 (1:2000, mouse anti-
human), phospho-Erk1/2 (1:2000, rabbit anti-human), mTOR (1:1000, mouse anti-
89 | P a g e
human), phospho-mTOR (1:1000, rabbit anti-human), p70S6K
(1:1000, rabbit anti-
human), phospho-p70S6K
(Thr389) (1:1000, rabbit anti-human), PAK1 (1:1000, rabbit
anti-human), phospho-PAK1 (Ser144)/PAK2 (Ser141) (1:1000, rabbit anti-human),
and α-Tubulin (1:4000, rabbit anti-human) overnight at 4oC. The primary antibodies
were all obtained from Cell Signaling (Cell Signaling Technology, Inc. Danvers, MA,
USA). The membranes were washed three times for 20 min each in TBS containing
0.01% Tween-20, and then incubated with corresponding fluorescent secondary
antibodies (Cell Signaling Technology, Inc. Danvers, MA, USA) at 1:2000 dilution
for 1 h at room temperature. Targeted proteins were visualized using the Odyssey®
infrared imaging system. Band intensities were quantified by scanning densitometry
and analysed using ImageJ software [359].
Statistical analysis
Statistical evaluation was performed using SPSS software (SPSS Inc., Chicago, Il,
USA). Values are expressed as mean ± standard deviation (SD). One-way analysis of
variance followed by Student’s t-test was used to determine statistically significant
differences between experimental and control samples. Significant difference was
considered at p<0.05.
RESULTS
Expression profile of VEGFR and PDGFR in MSCs
RT-qPCR analysis was performed using total RNA isolated from MSCs, with human
umbilical vein endothelial cells (HUVECs) as positive control. Only a very low level
of VEGFR-1, VEGFR-2, or VEGFR-3 mRNA transcripts were determined in MSCs
compared to HUVECs, with or without the stimulation of different concentration of
VEGF, reflecting the lack of VEGFRs in MSCs (Figure 22). On the contrary, both
VEGFR-1 and VEGFR-2 were readily detected in HUVECs, showing a steady
increase after exposure to VEGF (Figure 22). Interestingly, the treatment of MSCs
with different concentration of VEGF under serum-free condition significantly
augmented the expression of PDGFR-α and PDGFR-β in MSCs (Figure 22, p<0.05).
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Figure 22. The expression of VEGFR and PDGFR mRNA transcripts was examined
by RT-qPCR analysis, using HUVECs as positive control. H: HUVECs; B: bone marrow-
derived mesenchymal stromal cells; SF: serum-free medium; BCM: conditioned medium from bone
marrow-derived MSCs; OCM: conditioned medium from osteogenically differentiated bone marrow-
derived MSCs; 20v: 20 ng/mL VEGF; 40v: 40 ng/mL VEGF; 80v: 80 ng/mL VEGF.
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Furthermore, the co-culture with CM derived from OMSCs enhanced the expression
of PDGFRs, but failed to reveal an increase in the levels of VEGFRs. Importantly, the
application of 20 ng/mL VEGF neutralizing antibody impaired the PDGFRs
expression in MSCs induced by CMOMSC
. These data indicated that PDGFR-α and
PDGFR-β may be the key players in VEGF dependent or CMOMSC
-induced migration
of MSCs, whereas VEGFRs, although required for VEGF-mediated endothelial cells
(ECs) migration [373], are unlikely to be involved.
CMOMSC
-induced PDGFR-α and PDGFR-β tyrosine phosphorylation in MSCs
Having demonstrated the enhanced expression profile of PDGFRs in MSCs after
stimulated with CMOMSC
, we next examined whether CMOMSC
could lead to PDGFR
tyrosine autophosphorylation by using a human phospho-RTK array containing 49
different specific RTK antibodies. RTK screening of CMOMSC
treated MSCs revealed
distinct PDGFR-α and PDGFR-β tyrosine phosphorylation (Figure 23C). In
comparison, the cell lysate from serum-free medium or CMMSC
treated MSCs showed
a barely detectable level of tyrosine phosphorylation (Figure 23A and B). The
neutralization of VEGF was accompanied by a significant decrease in CMOMSC
-
induced PDGFRs activation (Figure 23D). Interestingly, no VEGFR tyrosine
phosphorylation was detected, further validating that the MSCs biological activity
stimulated by CMOMSC
was not mediated through VEGFRs.
Western blotting analysis also demonstrated that, compared to CMMSC
, CM
OMSC
enhanced PDGFR-α and PDGFR-β expression of MSCs in protein level (Figure 24A).
The supplementation of VEGF neutralizing antibody led to a substantial decrease of
PDGFR expression in MSCs (Figure 24A-C, p<0.05), indicating that CMOMSC
-
induced MSCs activity may be dependent on a PDGFR-mediated mechanism.
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Figure 23. CMOMSC
-induced tyrosine phosphorylation of PDGFR-α and PDGFR-β in
MSCs. Human phospho-RTK array was used to examine RTK phosphorylation levels in MSCs lysate
samples. RTK array analysis of (A) MSCs cultured in serum-free medium without supplementation of
exogenous growth factor; (B) MSCs cultured in CMMSC
; (C) MSCs stimulated with CMOMSC
; (D)
MSCs cultured in CMOMSC supplemented with VEGF neutralizing antibody.
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Figure 24. The levels of protein expression of PDGFR-α and PDGFR-β in MSCs in
response to different stimulations. (A) Western blotting evaluation of PDGFR-α and PDGFR-β
expressions of MSCs in protein level after stimulation with CMMSC and CMOMSC with or without the
addition of VEGF neutralizing antibody. (B) and (C) Quantitative measurement of the protein band
density of PDGFR-α and PDGFR-β. The expressions of PDGFR-α and PDGFR-β elevated when MSCs
were stimulated with CMOMSC. The addition of VEGF neutralizing antibody in CMOMSC led to reduced
levels of protein expression of both PDGFR-α and PDGFR-β (*p<0.05). CMMSC, conditioned medium
derived from undifferentiated mesenchymal stromal cells; CMOMSC, conditioned medium derived from
osteogenically differentiated mesenchymal stromal cells; Abvegf, VEGF neutralizing antibody.
Role of PDGFRs in CMOMSC
-induced MSC migration
To examine whether PDGFRs participate in CMOMSC
-induced MSC migration which
has been proven in the previous study (REPORT TWO), the activity of PDGFRs was
blocked by treating the MSCs with PDGFR tyrosine kinase inhibitor for 8 h. In vitro
transwell migration assay revealed that pre-treatment of PDGFR tyrosine kinase
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inhibitor mediated inhibition of PDGFRs action, which significantly impaired
CMOMSC
-induced MSC migration, resulting in substantial reduction of migrated cells
(Figure 25E, F and G, p<0.05). On the other hand, pre-treatment with either VEGFR-
1 or VEGFR-2 inhibitor did not alter CMOMSC
-induced MSC migration (p>0.05; data
not shown).
Figure 25. The involvement of PDGFR in MSC migration induced by CMOMSC
. (A)-(C)
Schematics representation of in vitro migration assay settings. The bottom chambers were
supplemented with (A) CMMSC, (B) CMOMSC and (C) CMOMSC with PDGFR tyrosine kinase inhibitor.
(D)-(F) Representative images of MSC migration induced by (D) CMMSC, (E) CMOMSC and (F) CMOMSC
with PDGFR tyrosine kinase inhibitor. (G) The migration capacity of MSCs was significantly
stimulated by CMOMSC compared to CMMSC. PDGFR tyrosine kinase inhibitor was applied to attenuate
the PDGFR bioactivity and the number of migrated MSCs decreased dramatically (*p<0.05). CMMSC,
conditioned medium derived from undifferentiated mesenchymal stromal cells; CMOMSC, conditioned
medium derived from osteogenically differentiated mesenchymal stromal cells; PDGFRi, PDGFR
tyrosine kinase inhibitor.
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Activation of the PI3K/Akt signaling is required for CMOMSC
-induced MSC
migration
To determine the signaling pathways involved in the CMOMSC
-induced MSC
migration, we then examined the effect of PDGFR tyrosine kinase inhibitor on the
phosphorylation of Akt and Erk1/2 in the co-culture system. MSCs were stimulated
with CMMSC
and CMOMSC
with or without PDGFR tyrosine kinase inhibitor after
serum-starved overnight. Following treatment, the phospho- and total levels of Akt
and Erk1/2 were determined by Western blotting analysis. As shown in Figure 26,
MSCs displayed a very low basal Akt phosphorylation after exposure to CMMSC
.
After stimulation with CMOMSC
, a significant increase of Akt phosphorylation was
produced, but the addition of PDGFR tyrosine kinase inhibitor resulted in a decreased
phosphorylation. However, there was no major difference in the expression profile of
phospho-Erk1/2.
In order to further confirm the involvement of the PI3K/Akt signaling pathway in
CMOMSC
-induced MSC migration, MSCs were pre-treated with 10 µΜ LY294002 for
30 min prior to their exposure to CMOMSC
for different time points (e.g. 15 min, 30
min, and 60 min). As shown in Figure 27, CMOMSC
induced a dramatic and rapid
elevation of phospho-Akt, reaching a maximum within 30 min of exposure followed
by a gradual decrease observed 60 min after CMOMSC
stimulation. No significant
change in total Akt expression was observed over the course of the experiment. The
interference with the PI3K/Akt signaling by LY294002 abolished the signal
transduction (Figure 27A and B, p<0.05). Furthermore, the inhibition of the
PI3K/Akt signaling pathway was associated with significantly reduced migration of
MSCs induced by CMOMSC
as shown in the in vitro migration assay (Figure 27C,
p<0.05).
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Figure 26. Akt and Erk1/2 phosphorylation status in MSCs in response to various
stimulations. (A) Western blotting evaluation of phospho- and total levels of Akt and Erk1/2 in
MSCs after stimulation with CMMSC and CMOMSC with or without the addition of PDGFR tyrosine
kinase inhibitor. (B) and (C) Quantitative measurement of the protein band density of phospho-Akt and
phospho-Erk1/2. Compared to CMMSC, CMOMSC induced a substantial phosphorylation of Akt. The pre-
treatment of PDGFR tyrosine kinase inhibitor attenuated the Akt phosphorylation induced by CMOMSC.
No significant difference was noticed in the phosphorylation status of Erk1/2 in MSCs after different
treatments (*p<0.05). CMMSC, conditioned medium derived from undifferentiated mesenchymal
stromal cells; CMOMSC, conditioned medium derived from osteogenically differentiated mesenchymal
stromal cells; PDGFRi, PDGFR tyrosine kinase inhibitor.
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Figure 27. Activation of the PI3K/Akt signaling is required for CMOMSC
induced
MSCs migration. (A) Western blotting evaluation of phospho- and total levels of Akt in CMOMSC-
stimulated MSCs at different time points with or without the pre-treatment of 10 µM LY294002. (B)
Quantitative measurement of the protein band density of phospho-Akt. CMOMSC induced the activity of
Akt within 30 min at highest level with gradual decrease till 60 min after stimulation. LY294002, a
specific PI3K/Akt inhibitor, suppressed the CMOMSC
-induced phosphorylation of Akt; (C) In vitro
migration assay showed that CMOMSC-induced MSC migration was significantly interfered by the
addition of LY294002 (*p<0.05). CMOMSC, conditioned medium derived from osteogenically
differentiated mesenchymal stromal cells.
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Activation of the PI3K/Akt signaling pathway downstream effectors
The phosphorylation status of downstream effectors of the PI3K/Akt signaling
pathway was evaluated by Western blotting analysis. As shown in Figure 28 A and B,
CMOMSC
significantly increased the phosphorylation of mTOR, p70S6K
, and PAK1 in
MSCs, compared to serum-free medium and CMMSC
. However, the phosphorylation
status of mTOR, p70S6K
, and PAK1 was dramatically suppressed in response to the
pre-treatment of LY294002.
Figure 28 The phosphorylation status of downstream effectors of the PI3K/Akt
signaling pathway. (A) Western blotting evaluation of phospho- and total levels of mTOR, p70S6K
and PAK1 in MSCs in response to various stimulations. (B) Quantitative measurement of the protein
band density of phospho-mTOR, phospho-p70S6K, and phospho-PAK1. CMOMSC induced significantly
increased phosphorylation of mTOR, p70S6K, and PAK1 in MSCs, compared to SF and CMMSC.
LY294002 suppressed the CMOMSC-induced phosphorylation of mTOR, p70S6K, and PAK1 (*p<0.05).
SF, serum-free medium; CMMSC, conditioned medium derived from undifferentiated mesenchymal
stromal cells; CMOMSC, conditioned medium derived from osteogenically differentiated mesenchymal
stromal cells.
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DISCUSSION
Targeted migration of endogenous cells to the required site is an essential aspect for
potential therapeutic success. However, the means by which donor cells trigger the
necessary signals required for the recruitment of host MSCs to the defect sites have
remained a puzzling gap in knowledge. For the last decade, considerable effort has
been directed toward identifying the signaling pathways that contribute to host cell
migration and the regeneration of damaged tissues. Our previous studies (REPORT
ONE and REPORT TWO) have shown that donor OMSCs attract host MSCs by
releasing VEGF among a variety of growth factors. Only little is known, however,
about how VEGF secreted by OMSCs signal through the downstream pathways that
instruct the recruitment of MSCs.
As a member from the VEGF/PDGF family, VEGF is the potent mitogen for
endothelial cells (ECs). VEGF not only regulates angiogenesis/vasculogenesis and
vascular permeability, but also induces and enhances ECs proliferation and migration
[374]. Numerous studies have suggested that the chemoattractant activity of VEGF
could be mediated through VEGFR-1, VEGFR-2 and VEGFR-3 [375-378]. However,
a number of studies have reported that limited VEGFR expression was found in
MSCs, indicating VEGF may signal through other receptors in MSCs [200, 372, 379].
In fact, previous reports have also demonstrated that VEGF and PDGF members are
closely related, and VEGFRs and PDGFRs exhibit structural similarities [223, 371].
The PDGF ligands exert their biological activities via two distinct but structurally
related receptors, PDGF receptor-α (PDGFR-α) and PDGFR-β. Particularly strong
expression of PDGFR-α has been noticed in subtypes of mesenchymal progenitors in
lung, skin and intestine, while PDGFR-β is reported to express in the mesenchyme,
especially in vascular SMCs (vSMCs) and pericytes [222]. The present study revealed
that although MSCs only expressed low levels of VEGFR-1, VEGFR-2 and VEGFR-
3, they did express high levels of PDGFR-α and PDGFR-β, which were activated by
the stimulation of VEGF and CM derived from OMSCs. Furthermore, blocking the
biological activity of PDGFR-α and PDGFR-β in MSCs by PDGFR tyrosine kinase
inhibitor prevented MSCs migration induced by CMOMSC
. However, the function of
VEGFRs had no impact on MSC directional migration. Our results suggested that
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functional cell surface PDGFR-α and PDGFR-β are both crucial determinants in
mediating MSC migration induced by OMSCs.
Having demonstrated that VEGF promotes mobilization of MSCs towards OMSCs
via PDGFRs, a series of experiments were performed to identify the downstream
signaling pathways participating in this process. The PI3K/Akt signal transduction has
been shown to be a crucial element in multiple cellular function, including cell
survival, proliferation, migration and invasion induced by VEGF among other
cytokines or growth factors [380]. This knowledge prompted us to investigate the
PI3K/Akt signaling in the context of what influence OMSCs have on the motility of
MSCs. The protein kinase Akt, a multifunctional regulator of cell survival, is the key
downstream effector of PI3K. One of the major functions of Akt is to phosphorylate
and activate mTOR, which is a protein kinase of the PI3K/Akt signaling pathway with
a central role in the regulation of cell proliferation, survival and mobility [381].
Subsequently, activated mTOR regulates p70S6K
and PAK1 phosphorylation. The Akt-
mTOR-p70S6K
-PAK1 cascade has been considered not only a central regulatory
pathway of the protein translation involved in regulating cell proliferation, growth,
differentiation and survival, but also a crucial step leading to actin cytoskeleton
reorganization. In the present study, detailed pathway analyses revealed that the
interaction of OMSCs with MSCs led to a rapidly increased PI3K/Akt
phosphorylation via the activation of PDGFRs. We also observed that the downstream
effectors of the PI3K/Akt signaling pathway, mTOR, p70S6K
and PAK1 were
significantly activated when MSCs were stimulated with CMOMSC
. However, the
pharmacological suppression of the PI3K/Akt activity resulted in a complete
attenuation of CMOMSC
-induced cell morphology change. These results suggested that
the PI3K/Akt kinase was activated and served as an essential downstream signaling
molecule in OMSCs-induced MSCs migration.
CONCLUSION
In summary, we demonstrated that MSCs possess potent migratory capacity after
stimulated by OMSCs, which can be applied in clinical setting to maximize the
regeneration effects of the cell-based therapeutic approaches. Our results also
indicated that the PI3K/Akt signaling pathway plays an important role in the
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recruitment of MSCs. More intensive studies are required to identify the responsive
host cells and to further explore the cellular and molecular mechanisms that govern
the directional migration of the host cells, thereby allowing for optimisation of the
therapeutic potential of pre-osteogenic differentiation of donor cells to be employed
for bone regeneration.
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CHAPTER 5 CONCLUSIONS AND PERSPECTIVES
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5.1 General discussion
Bone is a dynamic, mineralized tissue that continuously undergoes modeling and
remodeling, and possesses the intrinsic capacity for regeneration in response to injury
[382]. However, large bone defects resulting from trauma, tumor, and infection
cannot heal properly without intervention, and thus the treatment for these skeletal
deficits represents a major challenge in orthopaedic medicine. Although autologous
bone grafting is widely used in the clinic and has been the gold standard due to its
osteogenic, osteoinductive, and osteoconductive capacity [383], it is often associated
with limited tissue availability, chronic pain, and considerable morbidity at the donor
site [384, 385]. Allogenic or xenogenic bone grafting can be alternative options, but
these strategies have certain drawbacks that include inferior osteoactivity compared to
autologous bone grafting, donor incompatibility, and an increased risk of pathogen
transmission [383, 386].
In view of the limitations of conventional bone graft strategies, tissue engineering
recently has emerged as a promising approach for bone repair and regeneration. The
fundamental concept of bone tissue engineering is to combine progenitor cells, such
as MSCs or osteogenically differentiated/mature cells (for osteogenesis) seeded onto
biocompatible scaffolds and ideally in three-dimensional structures (for
osteoconduction and vascular ingrowth), with appropriate growth factors (for
osteoinduction) to trigger and utilize the endogenous biological capacity to generate
functional bone structures.
5.1.1 Scaffold materials for bone tissue engineering
Tissue engineering scaffolds with a well-defined architecture are designed to
influence the physical, chemical and biological environment to facilitate cell
attachment, proliferation and differentiation. Growth factors and other biomolecules
can be incorporated into the scaffold to guide the regulation of cellular function
during tissue regeneration [387]. The application of the tissue engineering scaffolds
provide the temporary support structure for tissue forming cells to synthesize desired
tissue and to allow nutrient diffusion. Scaffolds for bone tissue engineering are
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commonly constructed from biodegradable polymeric materials, either synthetic or
natural [388]. However, for the regeneration of load-bearing bones, the use of
biodegradable polymer scaffolds is challenging because of their low mechanical
strength. Attempts have been made to reinforce the biodegradable polymers with
biocompatible inorganic materials, such as calcium phosphate-based bioceramics and
bioactive glass.
5.1.1.1 Ceramics and bioactive glasses
Ceramics have long been recognized as potent osteoconductive materials for bone
tissue engineering. Calcium phosphate-based ceramics, such as hydroxyapatite (HA)
and β-tricalcium phosphate (β-TCP) are the inorganic materials which attract most
attention for bone repair application. When compared to β-TCP, HA shows minimal
biodegradation, which blocks the formation of new bones and remodeling, and results
in poor local stability or permanent stress concentration [389]. β-TCP is relatively
balanced between bone formation and scaffold absorption. It is also a good
biodegradable ceramic material which can supply a large quantity of calcium ion and
sulfate ion for bone regeneration [389]. However, the mechanical strength of β-TCP is
relatively weaker than that of HA [390, 391]. Furthermore, single β-TCP lacks of
osteoinductivity and osteogenicity, which restricts its application in bone tissue
engineering [389, 391]. In order to optimize the application of both HA and β-TCP,
some researchers have manipulated different ratio of a more stable phase (HA) and a
more soluble one (β-TCP), allowing the formation of biphasic calcium phosphate
(BCP) with a preferable degradation rate and mechanical properties [392].
Ever since the report of the bone-bonding properties of 45S5 Bioglass® more than 40
years ago [393], bioactive glasses (BG) based on silicate [394], phosphate [395] or
borate [396] represent another essential group of inorganic, bioactive scaffold
materials used for bone tissue engineering. The mechanism of the bonding of BG to
bone has been attributed to the formation of an HA-like layer on the glass surface in
contact with the body fluid [397]. It is believed that the initial formation of HA-like
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layer involves the adsorption of growth factors, followed by attachment, proliferation
and differentiation of osteoprogenitor cells [390]. 45S5 Bioglass® has been the gold
standard for the application of BG in bone tissue engineering, however, as a scaffold
material, it has several limitations. The difficulty of processing 45S5 Bioglass®
into
porous 3D scaffolds results in low mechanical strength [398]. Furthermore, the low
degradation rate of 45S5 Bioglass®
makes it difficult to match the rate of new bone
tissue formation [397]. The conversion of the scaffold to an HA-like layer is often
incomplete, thus a portion of unconverted glass remains in the scaffold, raising
uncertainty about its effect on the local microenvironment in vivo.
5.1.1.2 Synthetic degradable polymers
Synthetic degradable polymers possess various advantages such as predictable batch
to batch uniformity, being free from concerns of immunogenicity, and having a
reliable source of raw material [388]. Polyglycolide (PGA), polylactide (PLA) and
poly(lactide-co-glycolide) (PLGA) belong to the saturated aliphatic polymeric
materials, which are one of the ancient and most frequently used groups of materials
in the field of bone tissue engineering.
PGA is a rigid, thermoplastic material with a highly crystallinity (45-55%), and
therefore it exhibits high tensile modules with extremely low solubility in organic
solvents [399]. Due to its excellent fiber forming ability, PGA was initially
investigated as one of the best candidates for absorbable sutures. The first
biodegradable synthetic suture, Dexon™, which was approved by the US Food and
Drug Administration in the 1960s, was based on PGA. Sutures made from PGA lose
about 50% of their strength after two weeks, 100% after four weeks and are
completely absorbed within 4-6 months [400]. Non-woven PGA fabrics have been
extensively used as scaffold materials for tissue regeneration due to their excellent
degradability and good initial mechanical properties. However, the accumulation of
the degradation product, glycolic acid, may worsen the pH environment surrounding
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the cells and cause damage to the tissues, which may limit the biomedical application
of PGA.
PLA is synthesized by the dimerization of lactic acid. Lactic acid is a chiral molecule
with two optical isomers, the naturally occurring L (Levorotatory) isomer, and the D
(Dextrorotatory) isomer. Therefore, the polymers are referred to as PLLA and PDLA,
respectively. There is also a racemic mixture of D,L-PLA named PDLLA [401]. PLLA
is more commonly used, because the biodegradation product of PLLA is naturally
available in the human body. PLA has linear structure, and one pendent methyl group
which makes it more amorphous and hydrophobic than PGA [388]. However, being
more hydrophobic, the degradation rate of PLA is relatively low. It has been reported
that high molecular weight PLLA can take more than five years to be completely
absorbed, whereas about one year is needed for PDLLA [402]. Being a low strength
polymer with faster degradation rate compared to PLLA, PDLLA is the preferred
candidate for developing drug delivery vehicles and as low strength scaffold material
for tissue regeneration.
PLGA is synthesized through co-polymerizing lactide and glycolide, the mechanical
and biodegradation properties of which are based on the ratio of individual monomers
in the composite. Co-polymers of 25-75% L-LA with glycolide are amorphous due to
the disruption of the regularity of the polymer chain. PLGA polymers containing
50:50 ratio of lactide and glycolide show much greater degradation rate than those
containing higher proportion of either of the two monomers [403]. PLGA
demonstrates good cell adhesion and proliferation, making it a favorable candidate for
tissue engineering applications, especially in blood vessel, bone and cartilage
regeneration [404-406]. A major concern about the application of the synthetic
polymers is the chemicals (additives, traces of catalysts and inhibitors) and monomers
(lactic acid and glycolic acid) released from the degradation of polymer, resulting in
local and systemic host reactions that may cause clinical complications [407]. One of
the by-products, lactic acid, is reported to potentially create an adverse cellular
response at the implant site by reducing the local pH, leading to the release of
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prostaglandin E2 (PGE2), a bone resorbing and inflammatory mediator, by human
synovial fibroblasts and murine macrophages [408]. The toxic residues resulting from
the chemical cross-linking also make these synthetic polymers less desirable for
implantable devices [409, 410].
5.1.1.3 Natural degradable polymers
Natural polymers can be considered as the first biodegradable scaffold biomaterials
used clinically. Most of the naturally occurring polymers undergo enzymatic
degradation, and their degradation rate varies significantly with the site of
implantation, depending on the availability and concentration of the enzymes. The
most commonly investigated natural scaffold materials include alginate, collagen,
hyaluronic acid, fibrin, silk and chitosan. Much of the interest in these natural
polymers stems from their superior biocompatibility, relative abundance, the ability to
present receptor-binding ligands to cells, the susceptibility to cell-triggered proteolytic
degradation, and the possibility to mimic the innate microenvironment for bone tissue
regeneration.
Collagen is the most abundant protein in mammals, constituting up to one-third of
total body protein [411]. The individual polypeptide chains of collagen contain 20
different amino acids and the precise composition varies among different tissues [412].
Variations in the amino acid sequence give rise to the different types of collagen,
among which type I collagen is the most predominant type found in the extracellular
matrix (ECM), especially in tissues such as tendon, periosteum and bone [413].
Collagen has been among the most widely used biomaterials for bone tissue
engineering due to its abundance, biocompatibility, high porosity, low antigenicity
and absorbability in the body [414, 415].
The main disadvantage of collagen is its poor mechanical strength, especially
compared with native bone. To enhance its mechanical property, collagen is cross-
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linked by a variety of chemical agents or physical treatments. However, the
commonly used chemical agents such as glutaraldehyde and polyepoxy are cytotoxic
at certain concentrations, thus their use has been limited due to the residual cross-
linking compounds remaining in the collagen scaffold [416]. On the other hand,
overexposure to physical treatments, such as dehydrothermal treatment, ultraviolet
irradiation, gamma irradiation and microwave irradiation may lead to collagen
degradation [415]. Another major concern about the application of allogenic or
xenogenic collagen is its potential antigenicity and immunogenicity [417]. However,
reports have shown that the adverse immunological responses only occur in rare cases
and generally settle within a few months, especially in the application of type I
collagen [417, 418]. Premature absorption and other adverse effects of collagen
implantation have been rare and the combined use with immunosuppressant has been
shown to be effective [419].
5.1.2 Growth factors delivery
With improved understanding of fracture healing and bone regeneration at the
molecular level [135], recent research has demonstrated that the incorporation or
release of growth factors, such as basic fibroblast growth factor (bFGF), bone
morphogenetic proteins (BMPs), insulin-like growth factor (IGF), platelet-derived
growth factor (PDGF), transforming growth factor β (TGF-β), and vascular
endothelial growth factor (VEGF) can improve the biological performance of
scaffolds as cell carriers. Of these molecules, BMPs are the most extensively studied,
as they are potent osteoinductive factors. More than 30 BMPs have been identified so
far. Of the known BMPs, BMP-2, -4, and -7 are each individually able to induce new
bone formation at ectopic and orthotopic sites [78, 420-424]. However, the clinical
use of BMPs has been limited because of the expensive and difficult purification
process. Furthermore, the proteins isolated from allogenic or xenogenic sources may
carry potential health risks. The application of recombinant human BMP (rhBMP) has
been reported in a number of animal models as well as a few human clinical trials
[425-427]. The therapeutic potential of rhBMPs-2, -4, and -7 has been proven in
selected fracture repair and bone regeneration process. However, the safety issues
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risen from the supraphysiological concentrations of growth factors needed to obtain
the desired osteoinductive effects, the high cost of treatment, and more importantly,
the possibility of local inflammation and edema [428], ectopic bone formation [429]
and carcinogenicity [430] may have reduced the clinical research enthusiasm.
Unlike bone graft transplants where there is a pre-existing vascular supply, synthetic
bone constructs are devoid of vasculature. Researchers have been trying to address
this issue of whether it is a pre-vascularised scaffold developed in vitro or the release
of angiogenic factors from scaffold that promote angiogenesis in situ that will
enhance optimal bone regeneration. VEGF has been one of the most used growth
factors in the application of bone tissue engineering, because it is the key regulator in
a cascade of molecular and cellular events that ultimately lead to the development of
the vascular system [365]. Even though supplying VEGF alone was not sufficient to
initiate the cascade of bone regeneration in the critical-sized calvarial defects, it has
been reported that the combination of angiogenic and osteogenic factors can stimulate
bone healing [431]. Dual release of VEGF and BMP-2 from biodegradable scaffolds
seeded with BMSCs has been demonstrated to be effective for bone repair and
regeneration in a murine femur bone defect model [432]. PLGA scaffolds containing
plasmids encoding DNA for BMP-4 and VEGF have also shown greater bone forming
ability when implanted subcutaneously [433]. Hence, the combined delivery system
of growth factors at different rate kinetics locally from biodegradable scaffolds may
be a promising approach to enhance the reparative mechanism of bone defects.
Although these findings are interesting, the possible application of VEGF in bone
tissue engineering is limited by the short biological half-life, the need for repeated
examinations for optimum dosage, and the expensive cost for the provision of a
sustained concentration at the site of bone regeneration.
5.1.3 Cell sources
Recent advances in stem cell biology and regenerative medicine enable the
development of cell-based therapy. Cell-based bone tissue engineering approaches
entail the seeding of a scaffold with progenitor cells or mature cells that will be
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delivered to and retained within the lesion site. It has been reported that stem cell
delivery is likely to produce more reliable and effective results in the management of
bone defect compared to the traditional treatments [434, 435]. MSCs can be easily
isolated from readily accessible bone marrow among other tissues. Additionally, in
vitro expansion potential enables the generation of a sufficient number of bone
marrow-derived MSCs for transplantation [436]. Immunosuppressive and anti-
inflammatory functions [437, 438], homing ability to injured sites [439, 440] and
capability to modify the microenvironment by paracrine factors [441] enable MSCs to
become a promising candidate for cell-based bone tissue engineering.
It has been noted that stem cells alone are not sufficient for a long lasting regenerative
effect. The acceleration of endogenous regenerative mechanism that involves host cell
recruitment, a biological process also known as cell homing, has been considered as a
highly useful and practical approach for bone tissue regeneration [151, 310].
Although there are reports claiming that the MSCs from systemic sources which
express CXCR4 at their surface actively participate in bone regeneration [442, 443], it
is generally accepted that the skeletal progenitors within the local environment play a
more predominant role [444]. These skeletal progenitors may come from the bone
marrow within the injured bone, from the surrounding periosteum, and from soft
tissues in close proximity with the bone. However, all these tissues are closely linked,
which makes it difficult to distinguish their participation during bone repair and
regeneration.
There is a significant body of evidence that demonstrates the transplanted cells can
either differentiate into bone forming cells or attract other cell types to build up bone
tissue and simultaneously resorb cell carrier or bone substitute material [445-447].
Thus, the development of donor cell delivery approaches which can establish key
interactions with host cells in ways that unlock the endogenous power for robust
regeneration becomes increasingly important. The systemic delivery of donor cells
does not require invasive surgical intervention, and the donor cells could potentially
be injected at multiple time points. Furthermore, systemic delivery may be more
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beneficial to certain bone disorders affecting the whole body [448, 449]. However,
when delivered systemically, the donor cells contribute modestly to bone forming
cells [444]. Therefore, systemic delivery may not be ideal to direct exogenous cells
toward osteogenesis in a bone defect site. To augment the therapeutic effects of donor
cells, a number of strategies have been developed to deliver donor cells locally and
support bone regeneration. It has been reported that the volume of mineralized callus
is related with the number and concentration of fibroblast colony-forming units in the
bone marrow graft [450], indicating that the number of donor cells may play a part in
the osteogenic stimulation. Osteogenic differentiation of donor cells prior to
implantation may favor superior bone regeneration [19, 451, 452], but it decreases the
expansion capacity. Many efforts are also focused on the design of scaffolds to create
a biocompatible environment and provide proper vascularization. Scaffolds loaded
with angiogenic growth factors have been employed initially [453, 454]. Currently,
the implantation of composites of MSCs mixed with endothelial progenitor cells
(EPCs) is considered as a promising alternative [455].
5.1.4 Animal models for bone repair investigations
It is generally accepted that an ideal animal model for bone repair and regeneration
should mimic clinical conditions of bone injury, be able to utilize fixation, allow
mechanical load, create a permissive microenvironment for diffusion of nutrients and
growth factors, and permit angiogenesis [456]. The model of ectopic bone formation
is commonly employed because it is the least invasive and more suitable for
preliminary screening of donor cells, scaffolds and growth factors [457]. Nearly any
mammal can be chosen, from mouse and rat to rabbit, dog, and pig among numerous
others. To date, the rodent model is the most popular and widely used due to the low
cost, lax skin and availability of immunodeficient animals. A wide variety of locations
have been used for ectopic bone formation model, including subcutaneous,
intramuscular, and kidney capsule transplantation [458]. Subcutaneous implantation is
the simplest of all experimental models of ectopic bone formation. However, the
subcutaneous model is largely devoid of appropriate mechanical stimulation, which
may lead to eventual degradation of the newly formed bone. The physical
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identification of the implant can be challenging, especially when the newly formed
bone shares similar color with the surrounding dermal tissues. The bone forming
capacity is actually inferior in subcutaneous models compared to other ectopic models
due to reduced vascularization and blood flow. The lax skin may also allow
potentially significant migration of the implants, making it difficult for sample
collection.
For non-weight bearing testing, the bone defect can be created within the calvaria, rib
or mandibles, which have comparatively low mechanical forces. In this model, most
scaffold materials can be designed to fit the size of the defect and do not require
further fixation. For weight bearing testing, the defect can be made in long bones,
such as femur or tibia, using an osteotomy approach or a traumatic approach [456].
Osteotomy can surgically remove the required length of bone from a predetermined
site, producing a consistent defect in all subjects. However, this does not reflect the
actual clinical conditions following traumatic injury which produce a jagged cut edge
and traumatize both the bone and surrounding soft tissue. The disadvantage of the
traumatic approach is the potential for larger variation in defect size among subjects.
In a weight bearing site, the bone defect is reconstructed with the test material alone
or in combination with cells and growth factors and fixed using external or internal
fixation. Since the animal will start to move after recovery, the biomechanical
properties of the selected scaffold are crucial for optimum bone regeneration in a
weight bearing bone defect. Scaffolds need to be strong enough to allow weight-
bearing during the initial stage of the regenerative process but also biodegradable at
the right time to allow bone remodeling [457]. The contribution of host and donor
cells to bone formation in these complex animal models still remains to be elucidated.
5.2 Project summary
At the time when this project was first conceived, very few attempts had been made to
understand the relationship between host cells and donor cells, although an apparent
correlation between these two counterparts is present at a clinical scale [130-132].
The project sought to advance the knowledge and understanding of the interactions
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between donor cells and host cells during osteogenesis, the involvement of host cells
in the bone healing process, and the role of pre-programmed donor cells in the
recruitment of host cells. The ultimate goal is to unveil the potential molecular
mechanisms involved during this process at a cellular level. This project was designed
with three separate but closely–linked segments, which in the end led to three
independent reports summarized as follows.
Firstly, in REPORT ONE, we highlighted the concept of cell-based therapy and the
discrepancies that exist in the literature with respect to the interaction and
involvement of host cells and donor cells during osteogenesis. Having demonstrated
the enhanced migration tendency of MSCs towards OMSCs in the in vitro migration
assay, we then further investigated the role of donor cells during osteogenesis in a
small animal model. Using type-I collagen as a cell-carrier, donor cells were
implanted subcutaneously or into skull defects. Pre-manipulation of the donor cells
was found to significantly enhance mineralization and bone formation. The most
interesting finding was that in both ectopic and orthotopic sites, the implantation of
OMSCs could attract host cells and initiate osteogenesis. These findings provide some
insight into the paracrine effect of the secreted factors from OMSCs on the host cell
recruitment in vivo.
Using a combined in situ hybridisation and immunohistochemical staining technique,
our histological images revealed that the donor OMSCs strongly expressed VEGF,
giving a clue that VEGF secreted by donor OMSCs may play a role in the host cell
recruitment. Therefore, in REPORT TWO we particularly focused on the potential
effect of VEGF. Our data showed that VEGF secretion increased dramatically in
MSCs after osteogenic differentiation in vitro. The secretion of VEGF promoted the
motility of MSCs with elongated actin filament and enhanced the expression of
CXCL12/CXCR4, which is believed to be the most important axis that contributes to
MSCs homing to areas of damaged tissue. In vivo findings revealed that blocking
VEGF with neutralizing antibody led to a remarkable decrease of MSC recruitment
and new bone formation. Thus, the results from this study for the first time showed
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that VEGF secreted by transplanted OMSCs altered the cell morphology and
CXCL12/CXCR4 expression pattern of MSCs, triggering the downstream signaling
pathway, which may explain the host cell recruitment phenomenon observed in our
previous study.
In REPORT THREE, we first demonstrated that MSCs express limited functional
VEGFRs, which is in agreement with other reports [200, 372, 379]. As VEGF and
PDGF are closely related members from the VEGF/PDGF super family, VEGFRs and
PDGFRs are also structurally related, so we speculated that VEGF may exert its
paracrine effect through the activation of PDGFRs. Therefore, we furthered our study
and the results showed that the function of VEGFRs had no impact on MSC
directional migration, instead, the functional cell surface PDGFRs were crucial
determinants in mediating MSC migration induced by OMSCs. In order to unveil the
root causes of the enhanced host cell recruitment, our study probed cell signaling
pathways that are highly related to cell polarization and migration, such as PI3K/Akt
and Erk1/2. Our data showed that OMSCs induced a swift activation of PI3K/Akt in
MSCs, leading to the phosphorylation of downstream effectors, such as mTOR,
p70S6K
, and PAK1. This segment of study showed that the VEGF secreted from
OMSCs promoted MSCs directional migration via the activation of PDGFRs, which
then led to the activation of the PI3K/Akt signaling pathway.
5.3 Future direction
In this thesis, we mainly focused on the paracrine effect of VEGF secreted from
OMSCs, however, there are other cytokines derived from OMSCs which may also
play a significant role in host cell recruitment. Therefore, further studies are justified
to identify such soluble factors, probably by using proteomic mapping of the
conditioned medium derived from OMSCs. Also, during the last decade, growing
evidence has indicated that preferential activation of PI3K/Akt at the side of the cell
facing the chemoattractant gradient is critical for establishing a new leading edge, cell
polarization, and directional movement in a wide range of motile cell types [240, 459,
460]. Thus the clear recognition that the PI3K/Akt signaling pathway underlines the
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recruitment of host cells towards bone defect sites makes the clarification of
downstream targets of central importance to our understanding of bone defect healing.
More advanced animal models, such as knock-ins encoding catalytically inactive Akt
mutants, will likely provide a more accurate picture of the role of the PI3K/Akt
signaling pathway in bone tissue regeneration driven by the recruited endogenous
cells. It will also be important to further define the role of crosstalk between distinct
downstream branches of the PI3K/Akt signaling pathway, as well as the networking
with other signaling pathways, in the ultimate outcome of optimal bone remodelling
and regeneration.
Taken together, the observations from this work suggest that host cells and donor cells
interact closely in the bone healing process after cell transplantation. We have
reported for the first time that the implanted OMSCs play an integral role in recruiting
host cells towards the implantation sites, and more importantly, we have demonstrated
the factors secreted from donor cells (e.g. VEGF) can induce cell morphology
changes and enhance expression of CXCL12/CXCR4. We also reported that VEGF
secreted by OMSCs can signal through PDGFRs in MSCs. The signal is then
amplified by the activation of the PI3K/Akt kinase, ultimately leading to endogenous
cell recruitment and robust bone regeneration. This highlights the novelty of the
obtained results and therefore provides the scientific rationale to define new
therapeutic approaches for bone defect healing.
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