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UNIVERS ITY OF OULU P.O.B . 7500 F I -90014 UNIVERS ITY OF OULU F INLAND
A C T A U N I V E R S I T A T I S O U L U E N S I S
S E R I E S E D I T O R S
SCIENTIAE RERUM NATURALIUM
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ISBN 978-951-42-9317-7 (Paperback)ISBN 978-951-42-9318-4 (PDF)ISSN 0355-3221 (Print)ISSN 1796-2234 (Online)
U N I V E R S I TAT I S O U L U E N S I S
MEDICA
ACTAD
D 1034
ACTA
Annina Sipola
OULU 2009
D 1034
Annina Sipola
EFFECTS OF VASCULAR ENDOTHELIAL GROWTH FACTOR (VEGF-A) AND ENDOSTATIN ON BONE
FACULTY OF MEDICINE,INSTITUTE OF BIOMEDICINE,DEPARTMENT OF ANATOMY AND CELL BIOLOGY,UNIVERSITY OF OULU
A C T A U N I V E R S I T A T I S O U L U E N S I SD M e d i c a 1 0 3 4
ANNINA SIPOLA
EFFECTS OF VASCULAR ENDOTHELIAL GROWTHFACTOR (VEGF-A) AND ENDOSTATIN ON BONE
Academic dissertation to be presented with the assent ofthe Faculty of Medicine of the University of Oulu forpublic defence in Auditorium A101 of the Department ofAnatomy and Cell Biology (Aapistie 7 A), on 4 December2009, at 2 p.m.
OULUN YLIOPISTO, OULU 2009
Copyright © 2009Acta Univ. Oul. D 1034, 2009
Supervised byProfessor Juha Tuukkanen
Reviewed byProfessor Heikki KrögerProfessor George Sándor
ISBN 978-951-42-9317-7 (Paperback)ISBN 978-951-42-9318-4 (PDF)http://herkules.oulu.fi/isbn9789514293184/ISSN 0355-3221 (Printed)ISSN 1796-2234 (Online)http://herkules.oulu.fi/issn03553221/
Cover designRaimo Ahonen
OULU UNIVERSITY PRESSOULU 2009
Sipola, Annina, Effects of vascular endothelial growth factor (VEGF-A) andendostatin on bone. Faculty of Medicine, Institute of Biomedicine, Department of Anatomy and Cell Biology,University of Oulu, P.O.Box 5000, FI-90014 University of Oulu, Finland Acta Univ. Oul. D 1034, 2009Oulu, Finland
AbstractAngiogenesis is essential for the replacement of cartilage by bone during skeletal growth andregeneration. Vascular endothelial growth factor-A (VEGF-A) is a key regulator of angiogenesiswhereas endostatin, a potent inhibitor of endothelial cell proliferation and migration, is a naturalantagonist of VEGF-A. The regulatory roles of these peptides in angiogenesis, bone formation andbone cells were investigated in this study.
In the present work we studied the effects of VEGF-A, delivered with an adenoviral vector, onthe recovery of bone drilling defects in rat femur. Our data confirm the important role of VEGF inbone healing and that adenoviral VEGF gene transfer may modify bone defect healing in a rodentmodel.
We studied the effects of VEGF-A and endostatin on bone resorption activity. It was found thatVEGF-A is a potent stimulator of bone resorption and osteoclast differentiation in vitro andendostatin can antagonize this stimulatory effect when acting directly on bone cells. This suggeststhat endostatin is indeed a regulator of bone resorption, but not a critical one.
In the present study we induced ectopic bone formation in the hamstring muscles of adult mice.The effects of VEGF-A and endostatin in the ectopic bone formation assay were evaluated by thesimultaneous delivery of both peptides with recombinant adenoviral vectors. It was found thatendostatin retards the cartilage phase in endochondral ossification that subsequently reduces boneformation. We conclude that bone growth and healing, which share features with ectopic boneformation, may be regulated by endostatin.
To confirm in vivo effects on bone formation we further investigated the effects of endostatinand VEGF-A on mouse pre-osteoblastic cells in vitro. Finally the effects of endostatin on bonewere studied in transgenic mouse lines overexpressing endostatin, and mice lacking collagenXVIII.
Keywords: bone regeneration, endostatin, osteogenesis, VEGF-A
To my family
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Acknowledgements
This research was carried out in the Department of Anatomy and Cell Biology, at
the University of Oulu.
I would like to express my sincere gratitude to my supervisor, professor Juha
Tuukkanen, DDS, PhD. His guidance, positive attitude and patience have been
the most valuable throughout this project.
I also wish to express my appreciation to Docent Timo Hautala, MD, PhD for
constructive guidance.
I am grateful to Joanna Ilvesaro PhD, for teaching me the basics of lab work
and helping me in so many other ways at the beginning of this study.
I want to thank the former Chair of the Department of Anatomy and Cell
Biology, Professor Hannu Rajaniemi and also the present Chair Petri Lehenkari
for providing me this opportunity to work in the department.
Professor Heikki Kröger, MD, PhD and Professor George Sándor, MD, DDS,
PhD are acknowledged for their critical and valuable comments on this thesis,
and Dr. Deborah Kaska for the careful and expert English revision of this thesis.
I wish to thank all the co-authors: Elli Birr, Pekka Jalovaara, Timo Jämsä,
Ralf Pettersson, Taina Pihlajaniemi, Tatu Tarkka, Lotta Seppinen, Ylermi Soini,
Frej Stenbäck and Seppo Ylä-Herttuala for their contribution to the work.
I would like to warmly thank the whole staff of the Department of Anatomy,
especially the assistants who I worked with during those afternoon hours teaching
group works. I wish to thank Marja Juustila, Taina Poikela and Anna-Maija
Ruonala for their skillfull technical assistance and also Risto Helminen and Matti
Paadar for their help with the a number of computer problems.
I wish to thank all the members of Juha’s group for the opportunity to work
with them, especially my roommate Katri Nelo for all those nice conversations
and help with numerous practical matters, Mari Murtomaa, for pushing me
forward to finishing my thesis with your hilarious style, and Hanna Kokkonen for
sharing the practical arrangements of the dissertation and organization of the
dissertation party. I also warmly thank my dear friend Virpi Muhonen, for being a
priceless help in every work related problem I have had and sharing moments
outside work.
I warmly thank my great friends Antti and Marika, Jussi and Anu, Jone and
Pia for all the enjoyable moments that I have spent in your relaxing company. I
wish to thank my dear friends Heini and Jarno, Hannaleena and Mikko, Marja
and Jani for many unforgettable moments. Your positive energy makes me feel
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that anything in this world is possible. I wish also to thank my almost like sisters
Suvi, Jonna and Johanna for all the fun times since we were little girls.
I want to express my gratitude to my lovely parents Annikki and Timo for all
the support and love they have given in my life. I owe my loving thanks to my
sister and soul mate Marika and her husband Antti, who is like a brother to me,
and their lovely son Luka for being always there for me. I wish everybody would
have friends like you.
My husband Saku deserves my deepest thanks for his love, support and
amazing never-ending optimism. There is nobody quite like him. Our son Oliver
has taught me what is the meaning of this life. I love you both.
This work was financially supported by the National Graduate School for
Musculo-Skeletal Diseases (TBGS), the Finnish Dental Society Apollonia and the
Emil Aaltonen Foundation.
Oulu, November 2009
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Abbreviations
α-MEM α-modified minimum essential medium
αVβ3 Vitronectin receptor
ALP Alkaline phosphatase
BMP Bone morphogenetic protein
BMU Bone multicellular unit
BRU Bone remodeling unit
BSA Bovine serum albumin
BSP Bone sialoprotein
CFU Colony-forming unit
DMEM Dulbecco’s modified Eagle’s medium
ECM Extracellular matrix
ERK Extracellular signal-regulated kinase
FBS Fetal bovine serum
FGF Fibroblast growth factor
E Embryonic day
EC Endothelial cell
ENDO Endostatin
ES Endostatin
F-actin Filamentous actin
FBS Fetal bovine serum
FCS Fetal calf serum
FITC Fluorescein isothiocyanate
FSD Functional secretory domain
HSPG Heparan sulfate
IGF Insulin-like growth factor
IL Interleukin
M-CSF Macrophage colony-stimulating factor
MMP Matrix metalloproteinase
µM Micromolar, 10–6 g/mol
NCP Non-collagen protein
MSC Mesenchymal stem cell
NO Nitric oxide
OCN Osteocalcin
OPG Osteoprotegerin
OPN Osteopontin
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Osx Osterix
PBS Phosphate-buffered saline
PCR Polymerase chain reaction
PDGF Platelet-derived growth factor
PFA Paraformaldehyde
pQCT Peripheral quantitative computed tomography
PTH Parathyroid hormone
RANK Receptor activator of NF-κB
RANKL Receptor activator of NF-κB ligand
RGD Arginine-glycine-aspartic acid
Runx2 Runt-related transcription factor 2
TGF-β Transforming growth factor β
TNF Tumor necrosis factor
TRACP Tartrate-resistant acid phosphatase
SZ Sealing zone
VEGF Vascular endothelial growth factor
WGA Wheat germ agglutinin
1,25(OH)2D3 1,25-dihydroxyvitamin D3
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List of original publications
This thesis is based on the following articles, which are referred to in the text by
their Roman numerals:
I Tarkka T, Sipola A, Jämsä T, Soini Y, Ylä-Herttuala S, Tuukkanen J & Hautala T (2003) Adenoviral VEGF-A gene transfer induces angiogenesis and promotes bone formation in healing osseous tissue. J Gene Medicine 5: 560–566.
II Sipola A, Nelo K, Hautala T, Ilvesaro J & Tuukkanen J (2006) Endostatin inhibits VEGF-induced osteoclastic bone resorption in vitro. BMC Musculoskelet Disord 13(7): 56.
III Sipola A, Ilvesaro J, Birr E, Jalovaara P, Petterson RF, Stenbäck F, Ylä-Herttuala S, Hautala T & Tuukkanen J (2007) Endostatin inhibits endochondral ossification. J Gene Medicine: 1057–1064
IV Sipola A, Seppinen L, Pihlajaniemi T & Tuukkanen J (2009) Endostatin affects osteoblast behaviour in vitro, but collagenXVIII/endostatin is not essential for skeletal development in vivo. Calcified Tissue International 85(5): 412–420.
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Contents
Abstract Acknowledgements 7 Abbreviations 9 List of original publications 11 Contents 13 1 Introduction 17 2 Review of the literature 19
2.1 Bone ........................................................................................................ 19 2.1.1 Bone formation ............................................................................. 20 2.1.2 Endochondral bone formation ...................................................... 20 2.1.3 Bone matrix .................................................................................. 21
2.2 Bone cells ................................................................................................ 22 2.2.1 Osteoblasts.................................................................................... 22 2.2.2 Differentiation of osteoblasts ........................................................ 23 2.2.3 Wnt-signaling and bone ................................................................ 24 2.2.4 Osteocytes .................................................................................... 26 2.2.5 Osteoclasts .................................................................................... 27 2.2.6 Osteoclast differentiation .............................................................. 27 2.2.7 Osteoclastic bone resorption ......................................................... 28
2.3 Bone remodeling ..................................................................................... 29 2.3.1 Bone remodeling cycle ................................................................. 31
2.4 Vascularization in bone ........................................................................... 33 2.4.1 Formation and structure of blood vessels ..................................... 33 2.4.2 Bone endothelium ......................................................................... 34
2.5 VEGF ...................................................................................................... 35 2.5.1 VEGF receptors ............................................................................ 37 2.5.2 VEGF in endochondral bone formation ....................................... 38 2.5.3 VEGF and chondrocytes ............................................................... 39 2.5.4 VEGF and osteoclasts/chondroclasts ............................................ 40 2.5.5 VEGF and osteoblasts .................................................................. 40
2.6 XVIII collagen and endostatin ................................................................ 41 2.6.1 XVIII collagen .............................................................................. 41 2.6.2 Endostatin ..................................................................................... 42 2.6.3 Collagen XVIII /Endostatin inhibits angiogenesis ....................... 43 2.6.4 Endostatin, bone and cartilage ...................................................... 44
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2.6.5 Mechanism of endostatin action ................................................... 44 2.7 Gene therapy as a method of Growth Factor Delivery ............................ 45
3 Aims of the study 47 4 Materials and methods 49
4.1 Reagents (I-IV)........................................................................................ 49 4.1.1 Recombinant proteins (II, IV) ...................................................... 49 4.1.2 Recombinant adenoviruses (I, III) ................................................ 49
4.2 Experimental animals in vivo (I, III, IV) ................................................. 49 4.2.1 Bone defect healing model (I) ...................................................... 49 4.2.2 Endochondral ossification animal model (III) .............................. 50 4.2.3 Transgenic mice and knockout mice (IV) ..................................... 50
4.3 Cell Cultures ........................................................................................... 51 4.3.1 Osteoclast isolation and culture (II) .............................................. 51 4.3.2 Osteoclast differentiation assay (II) .............................................. 52 4.3.3 Osteoblast MC3T3 culture (IV) .................................................... 52
4.4 Histology, histomorphometry and immunohistochemistry (I, III,
IV) ........................................................................................................... 53 4.4.1 Haematoxylin-eosin (HE) staining (I, III, IV) .............................. 53 4.4.2 Immunostaining of capillaries by FVIII (I, III) ............................ 54 4.4.3 Collagen XVIII antibodies (IV) .................................................... 54 4.4.4 TRACP staining for identifying osteoclasts (II, III) ..................... 55 4.4.5 Area Measurement of Excavated Pits (II) .................................... 55
4.5 Biological assays ..................................................................................... 55 4.5.1 Determination of cell proliferation (WST-1) (IV) ........................ 55 4.5.2 Alizarin red S staining and quantification of calcium
deposition (IV) ............................................................................. 56 4.6 Radiographical analysis .......................................................................... 56
4.6.1 Radiographic evaluation of bone formation (III) .......................... 56 4.6.2 Peripheral quantitative computed tomography (pQCT) (I,
IV) ................................................................................................ 57 4.6.3 Faxitron X-ray imaging (IV) ........................................................ 57
4.7 Statistical analysis (I-IV) ......................................................................... 57 5 Results 59
5.1 Bone defect healing model (I) ................................................................. 59 5.1.1 Bone mineral content evaluated with peripheral
quantitative computed tomography (pQCT) (I) ............................ 59 5.1.2 Induction of vascularization (I) .................................................... 59
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5.1.3 Histological evaluation (I) ............................................................ 59 5.2 Effects of VEGF-A and endostatin on osteoclastic bone
resorption and differentiation in vitro (II) ............................................... 62 5.2.1 Bone resorption assay (II) ............................................................ 62 5.2.2 Osteoclast differentiation (II) ....................................................... 62
5.3 Endochondral ossification model (III) .................................................... 62 5.3.1 Radiographic evaluation of bone formation (III) .......................... 62 5.3.2 Induction of vascularization (III) .................................................. 63 5.3.3 Bone histology (III) ...................................................................... 64 5.3.4 The number of cartilage-resorbing chondroclasts (III) ................. 66
5.4 The effects of endostatin on the MC3T3-E1 osteoblastic cell line.......... 66 5.4.1 Osteoblast proliferation in vitro .................................................... 66 5.4.2 Matrix mineralization in vitro ....................................................... 66
5.5 Transgenic mice and knockout mice ....................................................... 67 5.5.1 pQCT and faxitron results ............................................................ 67 5.5.2 The formation of secondary ossification centers .......................... 67 5.5.3 Localization of collagen XVIII..................................................... 68
6 Discussion 71 6.1 Adenoviral VEGF-A gene transfer induces angiogenesis and
promotes bone formation in healing osseous tissues (I) .......................... 71 6.2 Endostatin inhibits VEGF-A induced osteoclastic bone
resorption in vitro (II) .............................................................................. 73 6.3 Endostatin inhibits endochondral ossification (III) ................................. 74 6.4 Endostatin inhibits proliferation of MC3T3-E1 osteoblasts and
together with VEGF-A suppress mineralization (IV) .............................. 77 6.5 The role of collagen XVIII/endostatin on bone development in
vivo in transgenic mice overexpressing endostatin and mice
lacking collagen XVIII ............................................................................ 78 7 Conclusions 81 References 83 Original publications 103
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1 Introduction
Throughout life, bones continuously alter their internal structure by remodeling
unlike other durable structures, such as teeth, tendons, and cartilage. Bone uses
this process to adapt to mechanical stress, repair damaged bone and maintain ion
homeostasis.
During bone growth, development, and remodeling, angiogenesis as well as
osteogenesis are closely associated processes. Physiological bone angiogenesis
occurs primarily during development and fracture healing. The majority of bones
in the vertebrate skeleton are formed by endochondral ossification, and blood
vessel invasion into cartilage is a crucial first step in the process whereby
cartilage is replaced by bone. This vasculature is also required for the transport of
nutrients and precursor cells, such as precursors of chondroclasts and osteoclasts
to the renewing bone tissue. Vascular endothelial growth factor, VEGF, is a key
mediator of physiological and pathophysiological angiogenesis. Bone remodeling
is intimately associated with vascular sinusoids and VEGF has an important role
in mediating interactions between the bone and the vasculature. Bone
endothelium has been found to signal in unique ways to other bone cells,
including bone- resorbing osteoclasts, bone forming osteoblasts, and their
progenitors.
Endostatin, a C-terminal fragment of collagen XVIII that is known to
antagonize VEGF-A, is found in basal membranes and is expressed by human
cartilage and fibrocartilage. Endostatin is capable of neutralizing the endothelial
cell migration, neovascularization, and vascular permeability induced by VEGF.
Resorbing osteoclasts express αv-integrins, which are receptors for endostatin,
thus providing a potential machanism for endostatin to control of osteoclast
function. Angiogenic and antiangiogenic factors play important roles in
ossification. It is hypothesized that endostatin expression by mature cartilage may
suppress osteoclasts while endostatin-deficient ossifying cartilage allows
osteoclastic activity and endochondral ossification.
The aim of this study was to explore the roles of VEGF-A and endostatin in
bone and also in processes that are not directly related to angiogenesis. We
approached these issues by using in vitro cell cultures and in vivo experiments.
We studied the role of VEGF, delivered with a first generation adenoviral vector,
in bone healing of drilling defects in rat femur. We also studied the effects of
VEGF-A and endostatin on bone resorption activity with a rat pit formation assay
and mouse osteoclast differentiation assay. Furthermore, we employed an
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endochondral ossification model in the mouse hind muscle pouch stimulated by
BMPs, and overexpressed endostatin with or without VEGF-A using recombinant
adenoviral vectors. Finally, we investigated the influences of endostatin and
VEGF-A on mouse pre-osteoblastic cells (MC3T3-E1). The role of endostatin in
bone formation in vivo was also studied on transgenic mouse lines overexpressing
endostatin (ES-tg), and mice lacking collagen XVIII (Col18a1-/-).
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2 Review of the literature
2.1 Bone
Bone has properties that make it ideal for its functions. It is mechanically strong
having very high tensile and compressive strength, but at the same time being
relatively light and elastic. Bone tissue has three main functions: First, it has a
mechanical function giving shape and rigidity to the body and providing
attachment sites for muscles as simple mechanical lever systems to produce body
movement. Second, bones provide a rigid framework that supports and protects
the soft organs of the body. The skull, for example, protects the brain. Third, it
serves as a metabolic tissue maintaining calcium and phosphate homeostasis.
Bone also provides an environment for hematopoiesis.
Anatomically, bones in the skeleton can be divided into flat bones, such as
skull bones, mandible and scapula and long bones, like humerus, femur and tibia.
In long bones, the shaft, composed mainly of compact bone, is called the
diaphysis and the region between the growth plate and the expanded end of the
bone, the epiphysis. The metaphysis is the junctional region between the growth
plate and the diaphysis. There are two morphologically distinct types of bone.
Cortical (compact) bone forms the outer thick and protective layer whereas
trabecular (cancellous) bone, found inside the bone, has a sponge-like structure
that serves a metabolic function due to its large surface area for remodeling.
(Marks & Odgren 2002)
Based on bone matrix arrangement, bone tissue can be classified as lamellar
or woven. Mature and cortical bone are typically lamellar in that with collagen
fibers are arranged in lamellae. In cortical bone, they are concentrically organized
around a neurovascular canal, termed a Haversian canal, commonly referred to as
osteons. Adjacent osteons are linked together by lateral canals. In trabecular bone
lamellae are arranged differently (parallel to each other) compared to those in the
denser compact bone. Woven bone, observed in the developing or immature bone,
is the type initially laid down during osteogenesis. Bones have an outer fibrous
sheath called the periosteum, and an inner surface, referred to as the endosteum,
which is in direct contact with the marrow. The periosteum covers all bone
surfaces except at the joints and is anchored to the bone by strong, collagenous
fibers called Sharpeys’ fibers that penetrate into the bone tissue. The endosteum is
a membranous sheath that lines the marrow cavity and also envelopes the surface
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of cancellous bone and lines the blood vessel canals (Volkman’s canals) that run
through the bone. They both contain an abundant supply of blood vessels, nerves,
nerve endings and osteoprogenitor cells. Bone has a rich vascular supply,
receiving 10–20% of the cardiac output. Blood vessels not only supply oxygen
and nutrition to forming and growing bones, but they play an active role in bone
formation. (Marks & Odgren 2002)
2.1.1 Bone formation
Bone formation can occur through two mechanisms, by intramembranous or
endochondral ossification. In either case, bone develops by the replacement of a
preexisting connective tissue. The skeleton is formed by mesenchymal cells,
derived from cranial neural crest, somites and lateral plate mesoderm. These cells
condense and form the future skeletal elements. (Hall & Miyaki 1992, Zelzer &
Olsen 2005) Intramembranous ossification is the process by which a mesenchymal
template is replaced by bone. It takes place mainly during the development of
some flat bones of the skull and facial bones, when mesenchymal precursor cells
differentiate directly into bone-forming osteoblasts and begin to secrete bone
matrix. Endochondral ossification is responsible for the formation of bones of the
extremities and vertebral column. Bone tissue replaces a preexisting hyaline
cartilage, the anlagen of the future bone.
2.1.2 Endochondral bone formation
Endochondral ossification is initiated when mesenchymal progenitor cells
condense and differentiate into chondrocytes to form the calcified cartilaginous
scaffold for new bone formation. Cells in the connective tissue surrounding the
cartilage (perichondrium) differentiate into osteoblasts that deposit a mineralized
bone matrix and give rise to the bone collar, the cortical bone. Capillaries invade
the perichondrium that surrounds the future diaphysis and transform it into the
periosteum.
Hypertrophic chondrocytes in the central region of the cartilage secrete
VEGF to induce the invasion of blood vessels. Vascularization of the calcified
cartilage brings osteoclasts that resorb the cartilage, osteoblasts that deposit bone
matrix and also other mesenchymal cells into the cartilage from the
perichondrium (Carlevaro et al. 2000, Zelzer et al. 2001, Colnot et al. 2004). This
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invasion results in the formation of a primary ossification center in the midshaft,
and cartilage is eroded and replaced by bone marrow and trabecular bone in the
ossification center. The process of endochondral ossification is seen in figure 1.
Later, secondary ossification centers of ossification appear within the
cartilaginous epiphysis by a mechanism that is similar to the formation of the
primary center (Karaplis 2002). Discrete layers of residual chondrocytes form
growth plates between the epiphyseal and metaphyseal bone centers to support
further postnatal longitudinal bone growth (Maes & Carmeliet 2008).
Endochondral ossification occurs during development as well as in pathological
osteogenesis such as fracture repair and ectopic ossification. The association of
angiogenesis, VEGF and endochondral bone formation will be discussed more in
chapter 2.5.2.
Fig. 1. A schematic presentation of an endochondral process leading to the formation
of the mouse tibia, where the differentiation of chondrocytes from mesenchymal cells
results in the formation of an avascular cartilage model of the future bone.
ED = embryonic day. Modified from (Zelzer & Olsen 2005).
2.1.3 Bone matrix
Bone tissue consists of mineralized extracellular matrix (ECM) and bone cells,
including bone-forming osteoblasts, bone-lining cells, osteocytes and bone-
resorbing osteoclasts. The ECM can be divided to organic and inorganic
components, which constitutes 35% and inorganic which constitutes 65% of bone
tissue, respectively. Inorganic bone matrix consists mainly of hydroxyapatite. The
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association of these substances gives bone its hardness and resistance. The
organic matrix is composed primarily of the most abundant protein in the human
body, type I collagen, and several non-collagenous proteins. Type I collagen is the
major protein in bone matrix, representing about 90% of the organic matrix. Type
I collagen is a triple-helical structure, which contains two α1 chains and one α2
chain. These assemble into fibrils, which are then arrenged in layers. The network
of type I collagen fibers provides the structure on which bone mineral is
deposited. The remaining 10–15% of the bone total protein is composed of non-
collagenous proteins (NCPs), which can be divided into cell-attachment proteins,
proteoglycans and gamma-carboxylated proteins.
The interaction of cells with the extracellular matrix is essential for their
anchorage, proliferation, migration and differentiation. Virtually all of the bone
matrix proteins are modified post-translationally to contain either N- or O-linked
oligosaccharides, being glycoproteins. In bone matrix, there are multiple
glycoproteins, including fibronectin, thrombospondin, vitronectin, osteoadherin,
osteopontin (OPN) and bone sialoprotein (BSP) that contain the integrin-binding
amino acid sequence Arg-Gly-Asp (RGD) and may therefore mediate cell
attachment to the bone surface (Robey 2002).
2.2 Bone cells
2.2.1 Osteoblasts
Osteoblasts are bone-forming cells located on the surface of bone or osteoid.
Morphologically, osteoblasts are cuboidal and columnar in shape and have an
extremely well developed rough endoplasmic reticulum, a large nucleus, and an
enlarged Golgi complex due to the high level of protein synthesis (Marks &
Odgren 2002). There are four maturational stages of osteoblast development:
preosteoblast, osteoblast, osteocyte and bone-lining cell (Aubin & Liu 1996).
Osteoblasts are polarized cells: they have an architecture in which different
structures are located at each end, based on the cell’s activity. Osteoblasts are
responsible for synthesizing the organic components of bone matrix, type I
collagen and non-collagenous proteins and regulate the formation of
hydroxyapatite crystals in the newly formed bone. The production of type I
collagen by active osteoblasts occurs after osteoblast precursor proliferation,
followed by alkaline phosphatase (ALP) expression that is needed for
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mineralization of bone matrix. Several non-collagenous proteins are deposited in
the matrix, like bone sialoprotein (Bsp), osteopontin (OPN), osteocalcin (OCN)
and also proteoglycans and glycoproteins.
2.2.2 Differentiation of osteoblasts
Osteoblasts arise from the same mesenchymal stem cells (MSC) as chondroblasts,
adipocytes, myoblasts and fibroblasts (Bennet et al. 1991, Grigoriadis et al. 1988,
Yamaguchi et al. 1991). The proliferation and differentiation of cells of the
osteoblast lineage occur under the influence of a number of a transcription
factors, growth factors and hormones (Aubin & Triffit 2002).
The transcription factors, Runx2 (runt related transcription factor, also known
as core binding factor 1, CBFA1) and Osterix (Osx), which functions downstream
from Runx2 regulate osteoblast differentiation (Karsenty & Wagner 2002,
Karsenty 2003, Nakashima et al. 2002). In the absence of either Runx2 or osterix,
no osteoblasts are formed. Runx2 null mice have an almost perfectly patterned
skeleton composed of cartilage, but no bones (Komori et al. 1997, Otto et al. 1997). Also chondrocyte differentiation to hypertrophy is diminished, except in
the distal appendicular skeleton (tibia-fibula, radius-ulna), where chondrocytes do
differentiate to hypertrophy. In addition, there is no blood vessel invasion into
hypertrophic cartilage in Runx2 null mice (Inada et al. 1999, Kim et al. 1999,
Zelzer & Olsen 2005). Runx2 can be phosphorylated and activated by the
mitogen-activated protein kinase (MAPK) pathway. This pathway can be
stimulated by a variety of signals, including bone morphogenic proteins (BMPs).
Bone morphogenetic proteins (BMPs), members of a family of secreted growth
factors, provide important tissue-specific signals to preosteoblasts that are
essential for full osteogenic differentiation (Chen et al. 2004, Mundy 2002). The
MAPK pathway can be also stimulated by signals initiated by the extracellular
matrix (ECM), fibroblast growth factor (FGF-2), mechanical loading and
hormones such as parathyroid hormone (PTH) (Franceschi & Xiao 2003). Many studies have highlighted the important role of Wnt signaling in bone
formation. The formation and proliferation of preosteoblast cells requires
signaling through the Wnt-frizzled-Lrp5 (low density lipoprotein receptor-related
protein) β-catenin signaling pathway (He et al. 2004).
A number of factors support the growth and differentiation of
osteoprogenitors and the commitment of mesenchymal stem cells to an osteoblast
lineage. This regulation is a complex phenomenon taking place in multiple points
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in the maturational sequence. The hormones, growth factors and cytokines
participating in this process may have biphasic or opposite activities at different
times or on different bone compartments (Aubin et al. 2006). FGFs, TGFβ, and
BMPs are potent osteoinductive factors at the early stages of osteoprogenitor
differentiation and proliferation. In addition, PTH, PTHRrP, IGFs,
glucocorticoids, vitamin D, and estrogen are important regulators of osteoblast
differentiation and maturation (Aubin & Triffitt 2002, Aubin et al. 2006, Hill
1998).
2.2.3 Wnt-signaling and bone
Wnts are a family of 19 secreted proteins that participate in the development and
maintenance of many organs and tissues, including bone (Wodarz & Nusse 1998,
Logan & Nusse 2004, Moon et al. 2004). Wnt binds to membrane receptor
complexes including low-density lipoprotein receptor-related protein (LRP)-5/6
as well as frizzled proteins (Logan & Nusse 2004, Moon et al. 2004). The
formation of this ligand-receptor complex initiates a number of signaling
cascades that include the canonical/beta-catenin pathway as well as several
noncanonical pathways. Canonical Wnt signaling has been reported to play a
significant role in the control of bone formation.
Briefly, the canonical pathway is transduced through stabilization of the β-
catenin protein, which is the key mediator of the Wnt pathway. Upon Wnt
activation it cannot be phosphorylated by GSK-3 leading to its cytoplasmic
accumulation and translocation into the nucleus where it triggers gene expression.
The absence of Wnt expression or inhibition of binding to its membrane receptors
leads to degradation of ß-catenin and inactivation of the signaling cascade (Liu et al. 2002).
Wnt signaling regulates bone formation through a number of mechanisms
and through different mechanisms at different stages of life (Krishnan et al. 2006). It controls mesenchymal stem cells and it regulates most aspects of
osteoblast physiology (Reya & Clevers 2005). It dictates osteoblast specification
from osteo-/chondroprogenitors and stimulates osteoblast proliferation. In
addition it enhances osteoblast and osteocyte survival (Khosla et al. 2008).
ß-catenin is required for the early and terminal differentiation proliferation and
function of osteoblasts (Hill et al. 2005, Rodda & McMahon 2006). The Wnt
pathway regulates the expression of certain transcription factors, such as Runx2,
Osterix and dlx5, and strongly stimulates osteoblastogenesis through the
25
interaction of various components (Bennett et al. 2005, Yavropoulou & Yovos
2007).
β-catenin has been shown to be essential for determining whether
mesenchymal progenitors become chondrocytes or osteoblasts. Precursors lacking
ß-catenin develop into chondrocytes (Hill et al. 2005, Day et al. 2005) but the
maturation of chondrocytes requires the presence of Wnt signaling (Church et al. 2002, Chimal-Monroy et al. 2002). Wnt signaling has a role in both embryonic
cartilage development and postnatal endochondral bone formation (Yang 2008).
Wnt signaling regulates osteoclast formation and bone resorption. β-catenin
regulates osteoclastogenesis through effects on the expression of osteoprotegerin
and RANKL (Glass et al. 2005, Holmen et al. 2005).
In clinical cases, mutations have been found in the Wnt genes, receptors and
inhibitors of the Wnt signaling pathway that are associated with changes in bone
formation and turnover. Wnt signaling also plays a role during bone regeneration
following fracture (Yavropoulou & Yovos 2007).
Fig. 2. A schematic presentation of the role of canonical Wnt signaling in skeletal
development. Plus signs indicate positive effects of Wnt; minus signs indicate
inhibitory effects of physiological canonical Wnt signaling. Wnt signaling facilitates
osteoblast specification from mesenchymal progenitors at the expense of
adipogenesis. It cooperates with Runx2 and osterix to maintain and promote
osteoblast maturation. Wnt signaling influences osteoclast maturation by regulating
RANKL levels in osteoblasts (Khosla et al. 2008). Modified from (Liu et al. 2008).
26
2.2.4 Osteocytes
Osteocytes are terminally differentiated bone-forming cells that comprises the
majority, 90–95% of bone cells (Bonewald 2008). When osteoblasts have
completed their matrix-forming function, approximately 50–70% die by
apoptosis, and the remainder are either incorporated into the newly formed
osteoid matrix and become osteocytes or remain on the surface as bone lining
cells (Dempster 2006). Bone lining cells can revert back to osteoblasts under
certain circumstances, for example after stimulation by PTH or mechanical force
(Dobnig & Turner 1995). Osteocytes have a very particular location in bone, they
are spaced throughout the mineralized matrix.
Dendritic-shaped osteocytes are highly branched cells that occupy small spaces
between lamellae, called lacunae. From these cell bodies, long cytoplasmic
processes radiate in all directions. Osteocytes form a unique three-dimensional
organization, that is a metabolically and electrically active network via
cytoplasmic processes that pass through the bone matrix via small channels, the
canaliculi, which course through the lamellae and interconnect neighboring
lacunae and the lining cells on the bone (Nijweide et al. 2002, Knothe Tate et al. 2004).
Several functions of osteocytes and their possible roles in the process of bone
remodeling have been proposed (Noble 2008). Based on the morphological
characteristics of osteocytes, the sensation of mechanical stress loaded onto bone
is suspected to be one of their functions. The flow of extracellular fluid in
response to mechanical forces throughout the canaliculi induces shear stress and
deformation of the cell membranes. It has been suggested that primary cilia
mediate mechanosensing in bone cells (Malone et al. 2007, Xiao et al. 2006). It
has also been hypothesized, that osteocytes have a role in regulating local bone
remodeling. A recent finding, that osteocytes are the source of sclerostin, a
molecule that regulates osteoblast function, supports this hypothesis (Noble
2008). In addition, osteocytes participate in calcium homeostasis (Nijweide et al. 2002).
The life span of osteocytes is probably largely determined by bone turnover:
it is thought that osteocytes are phagocytozed by osteoclasts during bone
resorption (Nijweide et al. 2002).
27
2.2.5 Osteoclasts
Osteoclasts are cells of hematopoietic origin (Suda et al. 1992). They function in
bone resorption during bone growth for the resorption of calcified cartilage and
modeling of growing bone. Resorption is also needed during fracture healing, as
well as tooth eruption.
Osteoclasts are giant (20–100 µm) cells, enableing them to cover a relatively
large matrix area and thus operate more efficiently (Bar-Shavit 2007). They are
multinuclear cells having an average of 4–20 nuclei, but around 100 nuclei are
found in osteoclasts in Paget's disease patients (Roodman & Windle 2005).
Osteoclasts are also characterized by numerous mitochondria that are needed to
provide the energy required for the resorption process (Mundy 1990). Osteoclasts
cycle between resorbing and nonresorbing phases. To resorb bone, they attach to
the bone matrix, their cytoskeleton reorganizes and they assume a highly
polarized morphology containing several different plasma membrane domains
(Baron 1989). The characteristics of osteoclasts also include the expression of
calcitonin receptors, vitronectin receptors and tartrate resistant acid phosphatase
(TRACP) (Takahashi et al. 2002)
2.2.6 Osteoclast differentiation
Osteoclasts are multinucleated bone resorbing cells derived from hematopoietic
cells of the monocyte-macrophage progenitor cell lineage in the bone marrow
(Suda et al. 1992, Takahashi et al. 2002). After proliferating in the bone marrow,
mononuclear preosteoclasts fuse into multinucleated cells. They reach the bone
surface through the blood circulation. Mononuclear precursors of osteoclasts also
circulate in the peripheral blood (Väänänen 2005). Under physiological
conditions, osteoclast formation is supported by cell-to-cell contact by
osteoblast/stromal cells, which express the receptor activator of nuclear factor
kappa B (NF-κB) ligand, RANKL, as a membrane bound factor (Suda et al. 1999). The differentiation of osteoclasts is dependent on the presence of RANKL
and macrophage colony-stimulating factor (M-CSF) that are expressed by
osteoblasts and stromal cells (Yasuda et al. 1998). RANKL together with M-CSF,
induces osteoclast differentiation without the need for supportive cells. RANKL
also increases the bone-resorbing activity of mature osteoclasts and enhances
their survival (Yasuda et al. 1998). M-CSF is crucial for the proliferation and
survival of precursor cells of osteoclasts as well as macrophages. Signals are
28
transmitted through the cFMS, which belongs to the receptor tyrosine kinase
superfamily. The op/op mouse lacks functional M-CSF and has osteoclast-
deficient osteopetrosis (Yoshida et al. 1990).
RANKL and RANK are members of the tumor necrosis factor (TNF) and
TNF receptor superfamilies. RANKL is a type II transmembrane protein found on
the surface of expressing cells and as a proteolytically released soluble form
(Lacey et al. 1998, Wong et al. 1997, Anderson et al. 1997). RANK is a type I
homotrimeric transmembrane protein.
Osteoprotegerin (OPG), also released by osteoblasts, but not exclusively, is a
soluble decoy receptor for RANKL. It blocks RANK-RANKL interaction and
thus protects the skeleton from excessive bone resorption (Simonet et al. 1997,
Lacey et al. 1998). OPG expression is regulated by most of the factors that induce
RANKL expression by osteoblasts. In general, upregulation of RANKL is
associated with downregulation of OPG. A recent finding suggests, that OPG
expression is regulated by Wnt/β-catenin signaling in osteoblasts, the same
pathway that regulates osteoblastic bone formation (Boyce et al. 2005, Glass et al. 2005, Holmen et al. 2005).
Systemic and local factors control the recruitment, differentiation and activity
of osteoclasts, mainly by affecting the RANK-RANKL-OPG system through
stromal cells and osteoblasts. Disorders in this system seem to affect the rate of
bone resorption in a several pathological states (Hofbauer & Heufelder 2001).
Parathyroid hormone (PTH), 1,25 dihydroxyvitamin D3, the biologically active
form of vitamin D3 tumor necrosis factor-α (TNF- α), and interleukins (ILs) have
been implicated as factors that enhance osteoclast formation and resorption
(Alsina et al. 1996). Estrogen, TGF-β, calcitonin, IGF and ILs have been shown
to have inhibitory effects (Roodman 2001). Recently, it was found that estrogen
prevents bone loss via the estrogen receptor α and induction of the Fas ligand
(Nakamura et al. 2007). Bone resorption is also regulated by the immune system,
where T-cell expression of RANKL may contribute to pathological conditions
such as periodontitis and autoimmune arthritis (Katagiri & Takahashi 2002,
Schett et al. 2005).
2.2.7 Osteoclastic bone resorption
After osteoclasts migrate to the resorption site, they initiate a multistep process
called the resorption cycle (Väänänen & Zhao 2002). First, mononuclear
precursors fuse to form a polykaryon and move to the site of resorption. Then,
29
osteoclasts attach tightly to the mineralized bone matrix and generate an isolated
resorption site. The part of osteoclast cell membrane, the sealing zone, seals the
cell onto the bone matrix (Väänänen & Horton 1995). The primary adhesion
mediating structures of osteoclasts are actin-rich structures known as podosomes
(Lakkakorpi et al. 1991, Jurdic et al. 2006). avß3 integrin activation has an
important role in the osteoclast attachment leading to podosome formation
(Väänänen 2005). The sealing zone surrounds the ruffled border, which is a
resorbing organelle. Bone demineralization involves acidification of the isolated
extracellular enviroment. pH ~4.5 is created by fusion of acidic vesicles with the
ruffled border and by an electrogenic proton pump (H+-ATPase) coupled to a Cl-
channel (Blair et al. 1989, Väänänen et al. 1990). In addition to these domains,
osteoclasts also have a basolateral membrane facing the bone marrow and a
fourth functional secretory domain (FSD) which is located in the upper part of the
cell opposite to the ruffled border (Bar-Shavit 2007).
Osteoclasts degrade both the mineral and organic component of bone matrix
in resorption lacunae under ruffled border membranes. The acidic milieu first
dissolve bone mineral (hydroxyapatite), exposing the demineralized organic
component of bone for lysosomal proteases. It seems, that cathepsin K is the main
bone matrix-degrading enzyme (Gowen et al. 1999). Metalloproteinases,
especially MMP9s, are also released into resorption lacuna to degrade collagen
and noncollagen proteins (Everts et al. 1998, Xia et al. 1999).
Bone degradation products are endocytosed, endocytic vesicles are
transported to the functional secretory domain and degradation products are
released at the cell’s antiresorptive surface (Nesbitt & Horton 1997, Salo et al. 1997). TRACP has been shown to be localized in transcytotic vesicles and able to
destroy collagen and other proteins (Halleen et al. 1999). Finally, the osteoclast
detaches from the bone and loses its polarized structure. Then, the osteoclast
either relocates to a new resorption site or undergoes apoptosis (Bar-Shavit 2007).
2.3 Bone remodeling
Bones continually alter their internal structure by remodeling. The remodeling
rate varies in different types of bones: trabecular bone is remodeled at a higher
rate than cortical bone in a healthy adult (Parfitt 2002). At any one time,
approximately 20% of the cancellous bone surface is undergoing remodeling, and
at any one surface location, remodeling will occur on average every 2-years
(Parfitt et al. 1997). The bone remodeling processes involve the resorption of the
30
old bone and the formation of new bone, which are coupled to one another and
are a strictly controlled. Most adult skeletal diseases are due to an excess of
osteoclastic activity, leading to an imbalance in bone remodeling which favors
resorption. Such diseases include osteoporosis, periodontal diseases, rheumatoid
arthritis, multiple myeloma, and metastatic cancers (Boyle et al. 2003).
Osteoporotic bone loss is the result of high bone turnover in which bone
resorption outpaces bone deposition (Rodan & Martin 2000, Teitelbaum 2007)
while osteopetrosis, which is a metabolic bone disease characterized by decreased
bone resorption, that leads to the accumulation of excessive amounts of bone.
Although having higher bone mass, osteopetrotic bones are easier to fracture
(Tuukkanen et al. 2000).
Bone remodeling occurs in discrete locations and involves a group of
different kinds of cells. The basic multicellular unit (BMU), which comprises
osteoblasts and osteoclasts within the bone-remodeling cavity, is located in close
proximity to a blood vessel, as seen in figure 3. In cortical bone, the capillary
grows along the excavated tunnel, and on trabecular surfaces, small capillaries are
frequently seen adjacent to osteoblasts (Ott 2000). Another model proposed for
the relationship of osteoblasts, osteoclasts, and capillaries, involves bone
remodeling compartments (BRCs). A BRC comprises the BMU, as well as the
layer of flat lining or stromal cells positive for osteoblast markers associated with
the capillary (Hauge et al. 2001, Parfitt 2000), as seen in figure 4. They are
connected to the osteocyte network through gap junctions. It has been thought
that BRCs may be structures through which mechanosensory signals from the
osteocyte network are transmitted to osteoclasts and osteoblasts so that they can
change their activity on trabecular surfaces (Eriksen et al. 2007).
A BMU is not a permanent structure. It forms in response to a signal or
stimulus, performs its function and disbands. The skeleton contains millions of
BMUs, all at different stages and they all are geographically and chronologically
separated from other BMUs (Hill 1998). The bone remodeling cycle involves a
complex series of steps that are highly regulated and always follows certain
phases. It starts with activation of osteoclast precursors, and osteoclastic
resorption. Then on the reversal phase it continues with apoptosis of osteoclasts
and smoothing of the erosion cavities by mononuclear cells. Finally,
mesenchymal cells differentiate into functional osteoblasts that lay down matrix
mineralization (Frost 1973, Frost 1986).
31
Fig. 3. Illustration of the basic multicellular (BMU) model of bone resorption in cortical
bone. Modified from (Cackowski & Roodman 2007).
2.3.1 Bone remodeling cycle
Activation is a continuous process in BMUs. When a BMU spreads, new surfaces
undergo activation. Activation involves the recruitment of osteoclast precursors to
the bone and their differentiation and fusion into functional osteoclasts. It is not
clear what attracts the resorptive cells to the remodeling site. It has been
suggested that osteoclast precursors (monocytes) are mobilized by chemotaxis,
and the chemoattractants responsible for this activity are derived from the bone
matrix, or in the case of collagen and osteocalcin, directly from the osteoblasts
that produce them (Malone et al. 1982, Bar-Shavit 2007). MMPs produced by
osteoclast lineages have been found to be important for the migration of precursor
cells (Delaisse et al. 2003).
Heino and colleagues have suggested that healthy osteocytes inhibit
resorption, and when these signals cease, the osteoclast precursors migrate toward
the site (Heino et al. 2002). Alternatively, it has been proposed that apoptotic
osteocytes may secrete regulatory factors that induce osteoclast differentiation
and initiate bone repair (Hedgecock et al. 2007).
During the resorption phase, osteoclasts remove both mineral and organic
components of the bone matrix (Blair et al. 1986). Multinucleated osteoclasts are
active for approximately 12 days (Parfitt et al. 1996), and after the osteoclasts
32
have finished resorption, the phase ends with osteoclast apoptosis followed by a
reversal phase, which lasts approximately 9 days (Reddy 2004).
During the reversal phase osteoblast-like cells remove the remaining organic
matrix from the resorption lacuna and it is populated by osteoblast precursors that
proliferate and differentiate into osteoblasts (Everts et al. 2002, Mulari et al. 2004). During reversal, all-important coupling signals are also sent out to
osteoblasts to replace the bone that has been removed (Olsen 2006).
After that, the osteoblasts begin to synthesize the osteoid and finally 15 days
after the organic matrix is secreted, it begins to mineralize. Osteoblasts secrete
small membrane bound vesicles, called matrix vesicles that establish suitable
conditions for initial mineral deposition by the concentration of calcium and
phosphate ions (Anderson 2003).
Fig. 4. Remodeling cycle. A) Activation of osteoclastic precursors. B) Resorption by
activated multinuclear osteoclasts. C) Reversal, when osteoblast like cells remove the
remaining organic matrix from the bottom of the resorption lacuna before deposting
new collagen fibrils. Osteoblast precursors proliferate and differentiate into
osteoblasts and then migrate into the resorption lacuna. D) Formation and
mineralization. Osteoblasts continue to deposit new bone matrix and mineralize
osteoid until the cavity is filled. E) Resting. Once osteoblasts are embedded in
osteoid, they mature into terminally differentiated osteocytes. Modified from (Matsuo
& Irie 2008).
33
2.4 Vascularization in bone
Bone is a highly vascularized tissue. The significance of capillary growth in bone
modeling and remodeling is well established in bone development and fracture
healing (Stevens & Williams 1999). The vasculature is required for the transport
of oxygen, nutrients, growth enhancing molecules and precursor cells, such as
precursors of chondroclasts and osteoclasts, for the renewing bone tissue.
The majority of bones in the vertebrate skeleton are formed by endochondral
ossification, and blood vessel invasion into the cartilage is an important
phenomenon in endochondral ossification. In embryonic skeletal tissue,
osteogenesis and angiogenesis are temporally related and a lack of VEGF activity
arrests the bone development (Gerber et al. 1999, Ensig et al. 2000). There are
numerous skeletal pathologies such as osteoporosis, osteopetrosis and
inflammatory bone loss that are related to modifications in blood supply (Brandi
& Collin-Osdoby 2006).
Angiogenesis is thought to depend on a finely tuned balance between
endogenous stimulators and inhibitors (Polverini 1995). Molecules that serve as
positive regulators of angiogenesis include fibroblast growth factor (FGF),
transforming growth factor (TGF)-α, TGF-β, hepatocyte growth factor (HGF),
tumor necrosis factor (TNF)-α, angiogenin, interleukin (IL)-8, and the
angiopoietins and particularly members of the vascular endothelial growth factor
(VEGF) family (Folkman & Shing 1992, Yancopoulos et al. 2000, Ferrara et al. 2003, Dai & Rabie 2007).
2.4.1 Formation and structure of blood vessels
The formation of a vascular network is implemented by both vasculogenesis and
angiogenesis. Vasculogenesis is a process whereby vessels are formed de novo
from endothelial cell (EC) precursors, known as angioblasts (Gonzalez-Crussi
1971). Angiogenesis is defined as the growth of new capillaries from a pre-
existing vascular network (Risau 1997). The blood vessel system is hierarchically
organized to deliver oxygen, soluble factors, and various types of cells to all
tissues in a carefully regulated manner. The inside of a blood vessel is lined with
endothelial cells, adhering to each other and to specific matrix molecules. The
endothelium functions as a barrier that limits the movement of cells and
molecules between the circulation and tissues (Risau 1995). The endothelial layer
is attached to the basement membrane (BM), which is composed of collagen IV,
34
XV, XVIII, laminins, heparan sulfate proteoglycans, nidogen-entactin and other
matrix components (Timpl 1996). Angiogenesis is an invasive process that
requires controlled activity of extracellular proteases and their inhibitors (Liotta et al. 1991). As a result the BM dissolves causing the EC to come in contact with
proteolytic fragments of the constituents of the basement-membrane.
Specialized smooth muscular cells called pericytes envelop the surface of the
vascular tube. The interaction between pericytes and the EC is important for the
maturation, remodeling and maintenance of the vascular system. Gap junctions
provide direct connections between the cytoplasm of pericytes and endothelial
cells, and they enable the exchange of ions and small molecules (Bergers & Song
2005). Pericytes and supporting mural cells sense the external environment
through integrin receptors that promote cell survival, growth, migration and tube
formation (Hall et al. 2006). Integrins have an important role in the angiogenic
activity of endothelial cells because their proliferation is anchorage-dependent.
Inhibitors of various integrins have all demonstrated anti-angiogenic effects. The
endothelium signals between surrounding cells through a host of humoral and
growth factors, cytokines and chemokines, reactive metabolites, and polarized
surface-associated molecules (Cleaver & Melton 2003). Pericytes have VEGF
receptors and are capable of differentiating to chondrocytes and osteoblasts
(Doherty & Canfied 1999). Thus they provide a pool of stem cells in
endochondral ossification and bone healing.
2.4.2 Bone endothelium
Bone endothelium has been found to signal to other bone cells, including
osteoclasts, osteoblasts, and their progenitors. Bone remodeling sites associate
intimately with vascular sinusoids, and histological findings suggest that
osteoblasts and osteoprogenitor cells develop concomitantly with endothelial cells
in blood vessels adjacent to sites where new bone is formed (Dai & Rabie 2007).
Osteoclasts, which are derived from hematopoietic precursors are present in both
the bone marrow and the peripheral circulation (Roodman 1999). They must
migrate through the endothelium to reach future sites of bone resorption. It has
been shown, that endothelial cells can stimulate the formation of osteoclasts by
several mechanisms, including RANKL expression, and also promote bone
resorption in vitro (Formigli et al. 1995, Collin-Osdoby et al. 2001) There is also
some evidence, that osteoclasts stimulate angiogenesis (Tanaka et al. 2007). It has
35
been shown that osteoblast lineage cells are also present in the circulation
(Eghbali-Fatourechi et al. 2005).
Vascular endothelial cells synthesize and secrete many factors known to have
an effect on bone cells, including M-CSF, RANKL, various growth factors,
cytokines, chemokines, prostanoids, free radicals, small peptides, adhesion
molecules, and matrix constituents. Additionally, bone endothelial cells are also
able to respond to bone modulators, including PTH, sex steroids, and
proinflammatory cytokines (Brandi et al. 1993, Brandi & Collin-Osdoby 2006).
Two key molecules that have well defined roles are VEGF and angiopoietins.
The receptors for both these ligands are predominantly expressed on ECs and
they regulate the proliferation and survival of these cells. For example,
angiopoietin-1 (Ang-1), expressed predominantly upon ECs, regulates the
proliferation and survival of endothelial cells. Ang-1 is expressed also in
osteoblasts. It binds to the tyrosine kinase receptor, Tie-2, which is expressed in
endothelial cells and also in hematopoietic stem cells (HSC) (Huang & Bao 2004,
Huang et al. 2007). Tie-2 positive HSC are localized to the bone surface where
they are in contact with Ang-1 expressing osteoblasts. It was recently discovered
that a subset of osteoblasts function as a key component of the HSC niche,
controlling HSC numbers. In addition to osteoblasts, HSC interact with other
stromal cells too, including ECs (Yin & Li 2006).
2.5 VEGF
Vascular endothelial growth factor, VEGF, generally referred to as VEGF-A is an
angiogenic factor that is important for vascular development and maintenance in
all mammalian organs and therefore a key mediator of physiological
angiogenesis, for example during embryogenesis, reproductive functions and
skeletal growth. In addition, VEGF-A has a crucial role in pathophysiological
angiogenesis that is associated with numerous malignant, ischemic, inflammatory,
infectious and immune disorders (Carmeliet 2005). Numerous different strategies
that target VEGF receptor signal transduction pathways have been designed to
inhibit pathological angiogenesis in several malignancies (Ferrara et al. 2003).
Additionally, there is also clinical interest in promoting angiogenesis in
conditions such as stroke and ischemia (Olsson et al. 2006).
VEGF is also called the vascular permeability factor (VPF), because in 1983
it was first discovered as a permeability factor based on its ability to induce
vascular leakage (Senger et al. 1983). In the late 1980s VEGF was discovered as
36
an angiogenic factor that was found to stimulate endothelial cells to proliferate,
migrate, and survive in a serum poor environment (Leung et al. 1989).
The importance of VEGF-A during angiogenesis has been demonstrated in
transgenic mouse gene deletion studies. Mice lacking a single Vegf allele resulted
in embryonic lethality, due to deficient and abnormal vascular development
(Carmeliet et al. 1996). Similar results were obtained with mice carrying
deletions of the VEGF receptor allele VEGFR-1 (Fong et al. 1995) or VEGFR-2
(Shalaby et al. 1995).
The VEGF family members are secreted dimeric glycoproteins, which belong
to a family of proteins that include VEGF-A, VEGF-B, VEGF-C, VEGF-D, and
placenta-like growth factor (PLGF). The predominant form, VEGF-A, is
alternatively spliced into at least five different isoforms in humans. VEGF121,
VEGF 145, VEGF 165, VEGF 189, and VEGF 206, that have different biological
activities such as different abilities to interact with the VEGF co-receptors
heparan sulfate proteoglycans (HSPGs) and neuropilin (Houck et al. 1991,
Poltorak et al. 2000). VEGF164 and VEGF188 show increased binding to heparin
sulfate containing proteoglycans present on the cell surface and extracellular
matrix. The VEGF120 splice form is completely soluble and unable to bind
heparin (Ferrara & Davis-Smyth 1997, Hall et al. 2006). Murine VEGF-As are
shorter than the corresponding human isoforms of 121, 145, 165, 189, and 206
amino acids (termed VEGF120, VEGF164, etc.)(Tamayose et al. 1996).
Proteolytic processing of VEGF-A splice variants affects their ability to interact
with the VEGF co-receptors (Lee et al. 2005).
VEGF165, used in this study, is physiologically the most abundant isoform of
VEGF-A (Tischer et al. 1991). VEGF120 is diffusible, VEGF188 binds to the cell
surface and extracellular matrix and VEGF164 combines these properties.
VEGF165 exists in both a diffusible form and as an extracellular matrix bound
form, which can be released or activated by plasmin, lactate (Kumar et al. 2007),
or matrix metalloproteinases (MMPs) (Lee et al. 2005, Breen 2007). Many major
growth factors including epidermal growth factor, TGF-a, TGF-ß, keratinocyte
growth factor, insulin-like growth factor-1, FGF, and platelet-derived growth
factor upregulate VEGF mRNA expression, suggesting that paracrine or autocrine
release of such factors cooperates with local hypoxia in regulating VEGF release
in the microenviroment (Ferrara et al. 2003).
37
2.5.1 VEGF receptors
The biological functions of VEGF are mediated through the binding to specific
receptors and co-receptors that are primarily expressed in the vascular and lymph-
vessel endothelial cells, but are also located on non-endothelial cell types
including monocytes and macrophages, hematopoietic stem cells, epithelial cells,
fibroblasts, smooth muscle cells, and myogenic precursor cells (satellite cells)
(Olsson et al. 2006, Breen 2007).
VEGF ligands bind to three receptor tyrosine kinases (RTKs) denoted VEGFR-1 (Flt-1), VEGFR-2 (KDR/Flk-1), and VEGFR-3 (Flt-4) as well as to co-
receptors. Co-receptors, including heparan sulfate proteoglycans (HSPGs) and
neuropilins (NRP), lack tyrosine kinase activity but may modify the binding to
tyrosine kinase containing VEGFRs or bind VEGF directly to signal a cellular
response (Breen 2007). All isoforms bind VEGFR1 and VEGFR2, whereas only
VEGF164 has been shown to bind NRP-1 (Soker et al. 1998, Neufeld et al. 2002).
The deletion of a single VEGF allele or receptor gene allele, VEGFR-1 or
VEGFR-2, has been shown to result in fatal vascular defects in the embryo (Fong
et al. 1995, Shalaby et al. 1995, Carmeliet et al. 1996, Ferrara et al. 1996).
VEGFR1 is expressed on many types of endothelial cells, monocytes (Barleon et al. 1996) and dendritic progenitors (Wu et al. 1996). VEGFR1 ligands are
chemoattractive for monocytes and osteoclasts (Ensig et al. 2000, Clauss et al. 1990). VEGFR1 acts as a decoy through its ability to bind VEGF-A and regulate
the level of VEGF available to bind to VEGFR2 and initiate vessel formation
during embryogenesis. A soluble form of VEGFR1 also acts as a natural VEGF
trap since it is expressed through alternative transcription and binds to VEGF-A
to prevent angiogenesis (Maynard et al. 2003, Breen 2007). The expression of
VEGFR2 is almost exclusively limited to vascular endothelial cells and their
progenitors (Asahara et al. 1997). VEGFR2 is associated with the integrin-
dependent migration of endothelial cells, since it can form a complex with
integrin αVß3 and induce of endothelial cell proliferation, migration, and in vivo
angiogenesis (Hutchings et al. 2003, Dai & Rabie 2007). VEGFR3 has been
shown to be important for lymphatic- endothelial cell development and function
(Kaipainen et al. 1995).
38
2.5.2 VEGF in endochondral bone formation
Over 40 years ago Trueta and others concluded that blood vessels have an active
role in bone formation (Trueta & Amato 1960, Trueta & Buhr 1963, Trueta &
Trias 1961). Skeleton formation starts with mesenchymal condensation and the
transcription factor SOX9 has been found to be an important regulator in this
process (Bi et al. 1999). Inactivation of SOX9 results in complete absence of
mesenchymal condensation leading to failure of cartilage formation. Recent
findings provide evidence for the involvement of SOX9 in the regulation of
VEGF expression in condensing mesenchyme, whereas VEGF expression has
been found to regulate limb vasculature via a direct long-range mechanism
(Eshkar-Oren et al. 2009)
Harada and others were first to document the expression of VEGF in bone
tissue and osteoblasts (Harada et al. 1994). Vessel invasion has a key role in
endochondral bone formation and VEGF plays an important role in this
phenomenon. VEGF may mediate interactions between endothelial cells and bone
cells. Endothelial cells, chondrocytes (Carlevaro et al. 2000), and osteoblasts
(Wang et al. 1997) secrete endogenous VEGF that may stimulate chondrocytes
(Carlevaro et al.2000), osteoblasts (Mayr-Wohlwart et al. 2002, Deckers et al. 2000) and osteoclasts (Nakagawa et al. 2000). In endochondral ossification, an
avascular cartilage model is first formed. Immature and proliferating
chondrocytes secrete and express angiogenic inhibitors (Moses et al. 1999).
Chondrocytes in the cartilage template and later in the growth plate first
proliferate and then differentiate into mature hypertrophic cells. They express
high levels of VEGF (Gerber et al. 1999).
VEGF expression in the hypertrophic cartilage causes the upregulation of
VEGFR1 and VEGFR2 expression in the perichondral endothelium (Colnot et al. 2001, Zelzer et al. 2001, Zelzer et al. 2002) and thereupon induces vessels to
invade the hypertrophic cartilage from the perichondrium (Colnot et al. 2004).
Zelzer and Olsen have speculated that high levels of VEGF near hypertrophic
chondrocytes maintain endothelial cells in a sprouting mode. This has been
explained by the fact, that sprouting growth plate capillaries contain no basement
membranes or pericytes but, in the region where bone matrix is being
synthesized, endothelial cells are surrounded by BMs and pericytes (Zelzer &
Olsen 2005).
VEGF upregulation is maintained in the sprouting vessels under the growth
plates during all stages of bone development (Gerber et al. 1999). Concurrent
39
with vascular invasion, the hypertrophic cartilage matrix is degraded by invading
osteoclasts/chondroclasts. Osteoblasts and marrow cells start to populate and
ossify two eventual centers of ossification: the primary ossification center on
midshaft and a secondary center on the epiphysis (Maes & Carmeliet 2008). It has
been shown, that MMP-9 specifically regulates proteolysis of nonmineralized
cartilage and the release of ECM-bound VEGF, which has a direct chemotactic
affect on osteoclasts (Engsig et al. 2000).
A study by Gerber and colleagues showed that blocking the VEGF receptors
inhibited vascular invasion of the cartilage and bone formation and resorption in
juvenile mice. In their study, blood vessel invasion was almost completely
suppressed, concomitant with impaired trabecular bone formation and expansion
of the hypertrophic chondrocyte zone (Gerber et al. 1999). In addition, transgenic
mice having chondrocyte specific inactivation of VEGF-A, show embryonic
lethality, disturbance in the blood vessel development and aberrant endochondral
bone formation (Haigh et al. 2000). The specific contributions of the VEGF
isoforms have been studied by using models in mice expressing only one of the
major isoforms. These studies demonstrate a significant role of VEGF at both an
early and late stage of cartilage vascularization. The matrix-binding isoforms
mediate metaphyseal angiogenesis and thereby regulate both trabecular bone
formation and growth plate morphogenesis during endochondral bone formation.
The soluble isoforms diffuse to the perichondrium and are required for epiphyseal
vascularization and secondary ossification in growing bone (Zelzer et al. 2002,
Maes et al. 2002, Maes et al. 2004).
2.5.3 VEGF and chondrocytes
VEGF is expressed at high levels in hypertrophic chondrocytes and low levels in
maturing chondrocytes in rats (Carlevaro et al. 2000). VEGF-A secretion by
hypertrophic chondrocytes is not only a paracrine process that attracts and
stimulates proliferation of endothelial cells, but is also an autocrine process that is
needed for chondrocyte survival (Dai & Rabie 2007). Pufe and others have shown
that VEGF affects the proliferation of chondrocytes in a dose dependent manner
with the C28/12 immortalized chondrocyte cell line (Pufe et al. 2004a).
It has been suggested, that Runx2 is involved in the regulation of VEGF in
hypertrophic chondrocytes. There is no blood vessel invasion into hypertrophic
cartilage in Runx2 null mice, since the upregulated expression of VEGF is absent
in the hypertrophic cartilage of Runx2-deficient mice (Zelzer 2001). The culture
40
of chondrocytes under hypoxic conditions up-regulates VEGF-A expression via
HIF-1a (Pfander et al. 2003). VEGF expression via HIF-1a is also induced by
mechanical overload of cartilage discs (Pufe et al. 2004b).
2.5.4 VEGF and osteoclasts/chondroclasts
Osteoclasts share a common hematopoietic precursor with monocytes and
macrophages, and, like them, osteoclasts express FLT1 as their main VEGF
receptor (Barleon et al. 1996, Niida et al. 1999). However, the expression of KDR
by cultured osteoclasts has also been reported (Nakagawa et al. 2000, Tombran-
Tink & Barnstable 2004). VEGF induces osteoclast differentiation in combination
with RANK from spleen-or bone marrow-derived precursors in culture and
further, a single injection of recombinant human VEGF-A (rhVEGF-A) into
osteopetrotic op/op mice (lack of MCSF-1) induced osteoclast recruitment and
survival, while stimulating osteoclastic bone resorption (Niida et al. 1999). VEGF
has been shown to stimulate the resorption activity of osteoclasts in vitro
(Nakagawa et al. 2000) and to be a chemoattractant for osteoclasts invading into
developing long bones (Ensig et al. 2000). VEGF and RANK stimulate the
migration of the osteoclasts into hypertrophic cartilage through the extracellular
signal-regulated kinases 1 and 2 (ERK1/2) pathway (Henriksen et al. 2003, Min
et al. 2003). VEGF also enhances dose- and time-dependent osteoclastic bone
resorption in a culture of highly purified rabbit mature osteoclasts partially by
enhancing the survival of the cells (Nakagawa et al. 2000).
2.5.5 VEGF and osteoblasts
VEGF has been implicated in various aspects of osteoblast function. Osteoblasts
have been found to express VEGF and its receptors in vitro with the highest
expression levels being found at the late differentiation stages (Harada et al. 1994, Deckers et al. 2000, Harper et al. 2001).
VEGF is a central molecule in osteoblast-stimulated angiogenesis, having
both autocrine and paracrine mechanisms of action (Grellier et al. 2009). Several
factors with important roles in regulating bone formation also induce the
expression of VEGF by osteoblasts. Prostaglandins E1 and E2, BMP-4, BMP-6,
BMP-7, FGF-2, TGF-b, endothelin-1, IGF-1, and vitamin D3 can all induce
VEGF expression in osteoblasts by activating a variety of signaling pathways, but
41
VEGF expression is inhibited by bone catabolic factors such as glucocorticoids
(Zelzer and Olsen 2005).
VEGF acts as a potent chemoattractant for rat (Nakagawa et al. 2000) and
human (Mayr-Wohlwart et al. 2002) osteoblasts, as well as bone-marrow-derived
mesenchymal progenitor cells (Fiedler et al. 2005). A number of studies indicate,
that VEGF induces the differentiation of osteoblastic cells (Midy & Plouet 1994,
Deckers et al. 2000, Street et al. 2002). Zelzer and coworkers showed that VEGF
has a stimulatory effect on bone formation. During calvaria organ culture,
treatment with VEGF led to an increase in the thickness of parietal bone (Zelzer
et al. 2002).
Furthermore, it has been reported that adenovirus-mediated VEGF gene
transfer induced bone formation by increasing osteoblast number and osteoid
forming activity in vivo (Hiltunen et al. 2003). VEGFR-1 signaling has been
shown to be critical for osteoblast activity during bone formation through
participation in osteoclastogenesis, as well as in the maintenance of a bone
marrow hematopoiesis supportive microenvironment. Mice that were deficient in
flt-1 signaling were shown to have very depleted numbers of osteoclasts and
osteoblasts, and also a deficiency in bone marrow cavity formation (Niida et al. 2005). Recently, also VEGFR3 has been found to be important for osteoblast
differentiation (Orlandini et al. 2006).
2.6 XVIII collagen and endostatin
2.6.1 XVIII collagen
Type XVIII collagen has been localized in basement membrane (BM)
prominently in vascular and epithelial BM throughout the body (Muragi et al. 1995). Collagen XVIII is a heparan sulfate proteoglycan (HSPG) (Halfter et al. 1998). HSPGs are composed of a core protein and heparin sulfate (HS) side
chains. Collagen XVIII belongs to the multiplexin (multiple triple-helix domains
and interruptions) subclass of the non-fibrillar collagen family, having ten
interrupted triple-helical (COL) domains that vary in length, separated by 11 non-
collagenous (NC) regions, flanked by N- and C-terminal non-collagenous
domains (Erickson & Couchman 2000). Collagen XVIII is expressed as three
N-terminal variants. These are transcribed from two different promoters,
promoter 1 being responsible for the production of the short variant, and promoter
42
2 for the two longer variants. The middle variant is generated from the long
variant by alternative splicing (Muragi et al. 1995, Rehn & Pihlajaniemi 1995,
Saarela et al. 1998). The C terminus includes a trimerization region, a hinge
region, which provides protease-sensitive sites, and an anti-angiogenic fragment,
endostatin.
2.6.2 Endostatin
Endostatin is a 20kDa carboxy-terminal fragment of the non-collagenous domain
of the collagen XVIII α1 chain. It was originally identified from murine
hemangioendothelioma cells (EOMA) (O´Reilly et al. 1997).
Structurally, endostatin is characterized by compact globular folding with
surface exposed arginine-rich patches (Hohenester et al. 1998). They are involved
in binding to heparin sulfate, which in turn may be important for endostatin’s
anti-angiogenic function (Kreuger et al. 2002, Sasaki et al. 1999, Dixelius et al. 2003).
Endostatin is liberated from the non-collagenous domain by the action of
various proteases, including matrix metalloproteinases (MMP) Cathepsin L, and
the serine protease elastase (Wen et al. 1999, Felbor et al. 2000). The released
forms can be found in the vessel wall, in plateles and freely circulating.
According to a number of studies, the normal mean circulating endostatin plasma
level is in the range of 10–50ng/ml (Dixelius et al. 2003).
43
Fig. 5. A schematic presentation of the structure of collagen XVIII. Ten collagenous
domains are interrupted and flanked by 11 non-collagenous domains. These are
numbered from the carboxy-to the amino terminus, NC1- through NC-11. Collagen
XVIII is expressed as three N-terminal variants. These are transcribed from two
different promoters. Modified from Dixelius et al. 2003, Bix et al. 2005).
2.6.3 Collagen XVIII /Endostatin inhibits angiogenesis
The effects of endostatin are broad, since in endothelial cells it inhibits
endothelial cell proliferation (O´Reilly et al. 1997), migration (Yamaguchi et al. 1999), proteinase activity (Wickström et al. 2001) and vessel stabilization
(Dixelius et al. 2000). Furthermore, endostatin induces endothelial cell apoptosis
(Dhanabal et al. 1999, Dixelius et al. 2000). Endostatin also affects tumor
angiogenesis indirectly by suppressing the endogenous expression of VEGF
(Hajitou et al. 2002).
Collagen XVIII deficient mice survive and do not display obvious vascular
abnormalities or altered tumor growth (Fukai et al. 2002). They suffer from eye
abnormalities, having defects in almost all ocular structures that express collagen
XVIII. These abnormalities are similar to those observed in patients with
Knoblochs syndrome, which is caused by an inactivating mutation in the collagen
α1(XVIII) gene (Fukai et al. 2002).
44
2.6.4 Endostatin, bone and cartilage
Only little is known regarding endostatins role in bone and cartilage. Pufe and
others have shown that endostatin/collagen XVIII is expressed in human cartilage
and fibrocartilage and they suggest it might reflect the importance of endostatin
for the development and maintenance of avascular zones. According to their
hypothesis, endostatin expression by mature cartilage may suppress osteoclasts
while endostatin-deficient ossifying cartilage allows osteoclastic activity and
endochondral ossification (Pufe et al. 2004c). Lau and others found that
endostatin may have a role in the anabolic effect of mechanical stimuli on bone
since endostatin can prevent the effect of fluid shear stress on osteoblasts (Lau et al. 2006). There is evidence that endostatin inhibits Wnt-signaling, acting
possibly at the level of ß-catenin in the signaling pathway (Hanai et al. 2002).
2.6.5 Mechanism of endostatin action
The exact mechanism of endostatin mediated inhibition has not yet been
elucidated. Endostatin binds to a number of cell surface molecules, but the signal
transduction events following binding to these receptors that result in the
inhibition of angiogenesis have so far been poorly characterized (Wickström et al. 2005, Delaney et al. 2006). Endostatin modulates many intracellular pathways.
Endostatin has been shown to be a heparin binding protein (Hohenester et al. 1998, Sasaki et al. 1999). This suggests that endostatin may competitively inhibit
the binding of angiogenic growth factors to heparan sulfate proteoglycans, which
are known to act as co-receptors for a number of cytokines (Hohenester et al. 1998). Dixelius and colleagues showed that heparin binding is needed for the
anti-angiogenic effects of endostatin. Endostatin mutants lacking heparin-binding
ability failed to block growth factor induced angiogenesis, in contrast to wild –
type endostatin in the chicken chorioallantoic membrane assay (Dixelius 2000).
Endostatin binding with low affinity to the heparan sulfate proteoglycans
glypican-1 and glypican-4 or with a high affinity to an as yet unidentified receptor
has been reported (Karumanchi et al. 2001).
It has been shown, that endostatin’s anti-angiogenic effects are mediated at
least in part, by interactions with integrins. Integrins play an essential role in the
angiogenic activity of endothelial cells because their proliferation is anchorage-
dependent. Inhibitors of various integrins have all demonstrated anti-angiogenic
effects (Bix & Iozzo 2005). Inhibitory action by endostatin has been proposed to
45
involve binding to the receptor α5β1 (Sudhakar et al. 2003). According to studies
of Wickström and colleagues, endostatin interacts with endothelial cells α5β1
integrin and perhaps concomitantly, with cell surface, heparan sulfate
proteoglycans and caveolin (Wickström et al. 2002, Wickström et al. 2003),
which leads to complex signaling cascades (Wickström et al. 2002, Bix & Iozzo
2005).
Interestingly, endostatin blocks both the VEGFR1 and VEGFR2 receptors of
VEGF and by that mechanism inhibits VEGF signaling (Kim et al. 2002). Recent
evidence shows that endostatin can also bind to VEGFR3 (Kojima et al. 2008). In
addition, endostatin’s inhibitory action is involved with several signaling
pathways, including Wnt/β-catenin (Hanai et al. 2002) and ERK1/2 signaling
(Schmidt et al. 2006). The anti-angiogenic activity of endostatin also requires the
presence of endothelial cell E-selectin, an endothelial cell-specific membrane
glycoprotein that is important for leukocyte attachment (Yu et al. 2004).
Furthermore, endostatin binds tropomyosin 3, a member of a large family of
actin-interacting proteins (MacDonald et al. 2001).
2.7 Gene therapy as a method of Growth Factor Delivery
VEGF has been used to promote angiogenesis of ischemic muscle tissues and
accelerate fracture healing, whereas endostatin has been found to inhibit the
growth of primary tumors and their metastases in a number of animal models and
human subjects in clinical trials (Spector et al. 2000, Folkman 2006, Keramaris et al. 2008, Uchida & Haas 2009).
The application of exogenous proteins may be limited by their short
biological half-lives and because of difficulties associated with obtaining
sufficient quantities for optimal therapeutic benefit. A single dose of recombinant
protein may not be sufficient to increase or decrease angiogenesis or osteogenesis
(Spector et al. 2000). A better strategy for protein delivery may be gene therapy,
which involves the transfer of genetic information to cells, and can be completed
in vivo or ex vivo. For in- vivo gene therapy, a vector carrying the therapeutic
DNA is directly implanted or injected into the target tissue so that host cells are
transfected and express the protein. For ex vivo therapy, gene transfer can be
delivered through transfection of cultured cells, which are then implanted or
injected into the target tissue. The duration of protein synthesis after gene therapy
depends on the techniques used to deliver the gene to the cell. (Kofron &
Laurencin 2006)
46
Vectors are agents that enhance the entry and expresion of DNA in a target
cell. Ideal vector have a high transfection efficiency, low toxicity, and consistent
gene expression. They may be of viral and nonviral origin. Currently, the most
common types of vectors are viruses that have been genetically altered to carry
non-viral DNA. Viruses have evolved specialized molecular mechanisms to
efficiently transport their genomes inside the cells they infect. Some of the
different types of viruses used as gene therapy vectors are for example,
adenoviruses, retroviruses, adeno-associated viruses and lentiviruses.
Adenoviruses can be produced in high titer and can transfect dividing and non-
dividing cells. Adenoviral DNA does not integrate into the genome and is not
replicated during cell division, with the result that transfected cells are gradually
diluted out of the population. The primary disadvantage of adenoviral vectors is a
potential host immune response, which targets the virus and cells transfected by
the virus. (Lieberman et al. 2002, Franceschi 2005, Kofron & Laurencin 2006)
47
3 Aims of the study
Angiogenesis is a fundamental part of osteogenesis as well as bone remodeling.
Since there is an intimate relationship between the vasculature and bone cells
within the bone remodeling site, the role of the basement membranes of the
vascular endothelium in the recruitment of bone cells was studied. One
component of basement membrane is collagen XVIII, a proteoglycan containing
heparan sulfate, and endostatin is a fragment of collagen XVIII, therefore, the
hypothesis was that endostatin could have a role in the activation of bone cells.
The affects of the major angiogenic factor, VEGF, on bone cells was studied.
The aims of the present study were:
1. to investigate the feasibility of a first-generation adenoviral vector to deliver
VEGF overexpression locally at the site of bone injury in order to alter the
course of bone regeneration in the rat femur.
2. to study the possible role of endostatin and VEGF-A on osteoclasts in vitro
by using a classical resorption pit assay, and to investigate osteoclast
mediated bone resorption and osteoclastogenesis in a differentiation assay
using bone marrow hematopoietic stem cells.
3. to follow the bone formation cascade in vivo in an endochondral ossification
model where ectopic bone formation is stimulated by BMPs in the mouse
hind muscle pouch, in the presence of overexpressed endostatin with or
without VEGF-A using a recombinant adenoviral vector.
4. to further investigate the influences of endostatin on osteoblastic behavior in vitro. The role of endostatin in bone was studied in vivo using a transgenic
mouse line overexpressing endostatin (ES-tg) and knockout mice lacking
collagen XVIII (Col18a1-/-).
48
49
4 Materials and methods
4.1 Reagents (I-IV)
4.1.1 Recombinant proteins (II, IV)
VEGF-A
recombinant human VEGF165 produced in Spodoptera frugiperda
(Calbiochem) was diluted in sterile PBS containing ≥ 0.1% BSA.
Endostatin
recombinant human endostatin polypeptide produced in Pichia pastoris
(Calbiochem) was diluted in sterile PBS.
4.1.2 Recombinant adenoviruses (I, III)
Recombinant E1-E3-deleted adenoviral vectors encoding VEGF164 (AdVEGF-
A) (Breier et al.1992) and nuclear targeted β-galactosidase (AdlacZ) under the
CMV promoter were constructed and produced in 293 human embryonic
kidneycells as previously described (Laitinen et al. 1998). Adenoviruses were
analyzed to be free from helper viruses, lipopolysaccharide and bacteriological
contaminants (Laitinen et al. 1998, Puumalainen et al. 1998).
To create a recombinant adenovirus expressing rat endostatin, the expression
cassette was cloned into the pQBI-AdCMV5-IRES-GFP vector (Quantum
Biotechnologies, Montreal, Quebec, Canada). The vector and the plasmid pJM17
(Microbix Biosystems, Toronto, Ontario, Canada) were then cotransfected into
293 cells, and the recombinant adenovirus was isolated, amplified, and purified
according to standard procedures (Pulkkanen et al. 2002).
4.2 Experimental animals in vivo (I, III, IV)
4.2.1 Bone defect healing model (I)
Male Sprague-Dawley rats (350–410 g) were anesthetized by intraperitoneal
injection of fentanyl citrate 0.315 mg/ml, midazolam 5 mg/ml, and fluanisone 10
mg/ml (1 : 1 : 1). A cylindrical 3-mm osteoperiosteal defect was drilled in distal
50
femurs approximately 5 mm from the epiphyseal plate. The surrounding muscle
was closed around the lesion and 2 × 108 plaque-forming units (pfu) of
adenoviral vector encoding vascular endothelial growth factor (AdVEGF-A) were
injected into the muscle in a 10-µl volume. The contralateral femurs served as
controls and received equal amounts of β-galactosidase adenoviral vector
(AdlacZ). The animals were sacrificed 1, 2, or 4 weeks (at least seven animals at
each time point) following the procedure and the femurs were collected and fixed
in 4% paraformaldehyde in 0.15 M sodium phosphate (pH 7.2) for further
analysis.
4.2.2 Endochondral ossification animal model (III)
Balb/c mice, age 10–12 weeks, were anesthetized by intraperitoneal injection of
ketamine midazolam (0.08 ml/10 g body weight). Ectopic bone formation was
induced by implanting native reindeer (Rangifer tarandus) BMP extract combined
with gelatine gel (5 mg of each) in 0.9% saline and recombinant adenoviral
vectors designed to overexpress endostatin (AdEndostatin), VEGF-A (AdVEGF-
A) ,or nuclear targeted β-galactosidase (AdlacZ). The mixture (100 µl) was
injected (1 ml syringe, 20 G needle) into both thigh muscle pouches in the
bilateral hind legs. The mice used in this study (total of 76) were divided into four
groups. The control group received BMP and gelatin gel only (n = 16) while the
second group received BMP combined with the gelatin gel and 1.2 × 109 pfu of
AdVEGF-A (n = 19). The third and the fourth groups were injected with BMP
combined with gelatin gel and 1.5 × 109 pfu Ad-endostatin only (n = 20) or in
conjugation with 1.2 × 109 pfu AdVEGF-A (n = 20), respectively. In addition,
four animals served as controls and received equal amounts, 1.6 × 109 pfu of
AdlacZ in BMP combined with gelatine gel. The animals were sacrificed in a
chamber with carbon dioxide 1, 2 or 3 weeks after the operation and the hind legs
were collected for further analysis. The animal tests were performed after
approval by the National Laboratory Animal Center. All aspects of animal care
complied with the Animal Welfare Act and the recommendations of the NIH-PHS
Guide for the Care and Use of Laboratory animals.
4.2.3 Transgenic mice and knockout mice (IV)
J4 mice overexpressing monomeric endostatin in the keratinocytes under the
keratin-14 promoter (ES-tg) with age- and sex-matched FVB/N wild type
51
controls, and knock-out mice deficient in collagen XVIII (Col18a1-/-) with age-
and sex-matched C57BL/6J wild type controls were used to study the effects of
collagen XVIII/endostatin on bone development in vivo. The generation of
collagen XVIII knock-out and J4 mice has been described in previous reports
(Fukai et al. 2002, Elamaa et al. 2005). Groups of 6–10 mice containing equal
numbers of males and females were used for the experiments, and the mice were
sacrificed at the age of 14 or 30 days. All the experimentats were approved by the
Animal Care and Use Committee of the University of Oulu, and in the case of the
Col18a1-/- mice also by the State Provincial Office in Oulu.
4.3 Cell Cultures
4.3.1 Osteoclast isolation and culture (II)
The procedure for the isolation and culture of osteoclasts described earlier by
Boyde et al. and by Chambers et al. was modified slightly and has been described
in detail previously (Boyde et al. 1984, Chambers et al. 1984). Briefly,
mechanically harvested osteoclasts from the long bones of 1- or 2-day-old
Sprague-Dawley rat pups were allowed to attach to ultrasonicated bovine cortical
bone slices (0.125 cm2 or 0.5 cm2). After 30 minutes, the nonattached cells were
rinsed away, and the remaining cells on the bone slices were cultured in
Dulbecco’s modified Eagle’s medium (α-MEM) buffered with 20 mM HEPES
and containing 0.84 g of sodium bicarbonate/liter, 2 mM L-glutamine, 100 IU of
penicillin/ml, 100 μg of streptomycin/ml and 7–10% heat-inactivated fetal calf
serum (FCS), at +37°C (5% CO2 and 95% air). The cells were divided into ten
groups: the control group had α-MEM-FCS as their culture medium with no
added substances (later referred to as control), the VEGF-A-treated cells had 100
ng/ml, 50 ng/ml or 10 ng/ml VEGF-A in α-MEM-FCS (later referred to as
VEGF). The endostatin groups had 0.02, 0.2 or 2 µg/ml endostatin in α-MEM-
FCS (later referred to as ENDO) and the last groups had both VEGF-A and
endostatin added into the complete culture media (later referred to as
VEGF+ENDO). The last groups had 100 ng/ml VEGF-A and 2 µg/ml endostatin,
50 ng/ml VEGF-A and 0.2 µg/ml endostatin or 10 ng/ml VEGF-A and 0.02µg/ml
endostatin. The cells were allowed to grow for 48 h, after which the bone slices
were fixed with freshly made refrigerated 3% paraformaldehyde (PFA) and 2%
sucrose in phosphate-buffered saline (PBS) for 10 minutes at room temperature.
52
The data shown were gathered from three independent experiments, each of
which gave identical results.
4.3.2 Osteoclast differentiation assay (II)
The procedure for osteoclast differentiation described earlier by Takahashi et al. was slightly modified (Takahashi et al. 1988). Mouse (C57BL/6, 8–12 weeks)
bone marrow cells were isolated from mouse long bones using a syringe and
transferred to a petri dish. After incubation at +37 °C for 2 hours, non-attached
cells were collected from petri dish and seeded in 24-well plates containing
sonicated cortical bovine bone slices (0.125–0.5 cm2) at a concentration of 1 ×
106 cells/well. Cells on the bone slices were cultured in four groups: control,
VEGF (100ng/ml), ENDO (2µg/ml) and VEGF + ENDO respectively. The
control group was cultured in α-MEM medium (Sigma, UK) with 10% heat-
inactivated fetal calf serum (FCS), 2 mM L-glutamine, 100 IU/ml penicillin, 10
μg/ml streptomycin and 20 mM Hepes, a medium later referred to as α-MEM-
FCS. The cells were cultured in 500 μl α- MEM containing 30 ng/ml RANKL
(Peprotech Ec., UK) and 10 ng/ml M-CSF (R&D Systems). Half of the medium
was replaced every 3rd day. The cells were cultured at +37 °C (5% CO2, 95% air)
for 7 days, after which the cultures were stopped by fixing the cells with 3%
paraformaldehyde (PFA) / 2% sucrose in PBS.
4.3.3 Osteoblast MC3T3 culture (IV)
MC3T3-E1 cells are pre-osteoblasts derived from murine calvarias (American
Type Culture Collection, ATCC, USA). They were maintained in α-MEM
supplemented with 10% FCS, 2nM L-glutamine, 100 IU 7ml penicillin and 10
µg/ml streptomycin (α-MEM-FCS), and passaged every 2–3 days using standard
techniques. After pre-culturing the cells were seeded at a density of 10.000 cells /
cm2 in the presence of α-MEM with 50 µg/ml ascorbic acid and 10 mM β-
glycerophosphate (mineralization medium). The cells were cultured in six groups
and the medium was supplemented as follows: 1) control group (α-MEM-FCS as
a culture medium with no added substances), 2) VEGF-A 100 ng/ml (recombinant
human VEGF165 produced in Spodoptera frugiperda, Calbiochem), 3) endostatin
2 μg/ml (recombinant human endostatin polypeptide produced in Pichia pastoris,
Calbiochem), 4) endostatin 20 μg/ml, 5) VEGF-A + endostatin 2 μg/ml and
6) VEGF-A + endostatin 20 μg/ml. The cells were cultured at +37 °C (5% CO2,
53
95% air) and the medium freshly supplemented with VEGF-A, endostatin or
VEGF-A and endostatin together was replaced after 1, 3, 7 and 10 days of culture.
Each experiment was repeated at least twice.
4.4 Histology, histomorphometry and immunohistochemistry (I, III, IV)
For histological analysis to evaluate bone defect healing, the femurs were fixed in
phosphate-buffered 4% paraformaldehyde and decalcifiedin 10%
ethylenediaminetetraacetic acid (EDTA) (I). For evaluation of evaluating of
ectopic new bone, the collected specimens were fixed in 10% neutral formalin
solution and decalcified in 5% formic acid. After decalcification, the bones were
embedded in paraffin (III). When studying transgenic mouse lines (IV), the bone
specimens were fixed in 70% ethanol, decalcified and embedded in paraffin.
4.4.1 Haematoxylin-eosin (HE) staining (I, III, IV)
The decalcified bones were cut into 5-µm tissue sections and stained with
hematoxylin and eosin. Morphological analysis was performed with a digital
image analysis system MCIDM4 (Imaging Research Inc., St Catharines, Canada)
using a Nikon Optiphot II microscope and a Sony DXC-930P color camera.
Study (I): The histology of the tissue was analyzed in order to demonstrate
differences in healing between the VEGF-treated and control femurs. The
proportion of fibrous reparative tissue was measured histomorphometrically from
the cross sections of the defect area 1, 2 or 4 weeks after the procedure. The
amount of cartilage at the periosteal rim of the defect was also measured 1 and 2
weeks after the procedure.
Study (III): Evaluation of ectopic new bone was based on the proportional area of
remaining cartilage in the ossicle 2 and 3 weeks after the injection determined
with morphometrical analysis. The ossicle was defined as the tissue surrounded
by fibrous capsule and containing a cortical outer bone shell and developing bone
marrow around the remaining endochondral cartilage.
Study (IV): Femur samples were studied for the analysis of secondary ossification
centers in 2 and 4 week old Col18a1-/-, C57BL/6J, ES-tg and FVB/N mice.
54
4.4.2 Immunostaining of capillaries by FVIII (I, III)
The VEGF (I, III) and endostatin (III) effect in bone was demonstrated by
immunostaining of capillaries using a polyclonal rabbit anti-human antibody to
FVIII-related antigen (Dako, Dakopatts, Denmark). The primary antibody was
applied on the slides for 1 h (dilution of 1 : 50) followed by a secondary anti-
rabbit antibody (Dako), after which the ABC-complex was applied. The color was
developed with diaminobenzidine. In study (I) the vascular organization was
evaluated and density was estimated as the number of positively stained blood
vessels per high power field (HPF). In each section, a minimum of five HPFs was
analyzed. In study (III) morphometrical analyses of the factor VIII positive blood
vessels were done from three microscopic fields (digital image analysis system
MCID M4 using a Nikon Optiphot II microscope and a Sony DXC- 930P color
camera) of the highest vessel density persection in a continuous fashion. The
numbers of the positively stained blood vessels were counted as proportional area
per microscopic field.
4.4.3 Collagen XVIII antibodies (IV)
Paraffin-embedded 5 μm sections were dewaxed, washed in PBS and stained with
collagen XVIII antibodies using a tyramide signal amplification (TSA) kit
(PerkinElmer). The following antibodies were used in immunohistochemistry: a
polyclonal rabbit-anti mouse antibody recognizing the N-terminal part of type
XVIII collagen (anti-all moXVIII) and a polyclonal rabbit-anti mouse antibody
recognizing the N-terminal part of the two longer variants of type XVIII collagen
(anti-long moXVIII) (Saarela et al., unpublished data) Heat-induced epitope
retrieval was carried out in citrate buffer (10mM sodium citrate, 0.05% Tween,
pH 6.0) for 10 min in a microwave oven. The slides were then incubated
overnight at + 4 ºC with either anti-all moXVIII (4 μg/ml) or anti-long moXVIII
(10 μg/ml) antibody, diluted in TNB buffer (0.1 M Tris-HCl (pH 7.5), 0.15 M
NaCl, 0.5% Blocking Buffer). This was followed by a 30-minute incubation with
a biotinylated secondary anti-rabbit antibody (Vector Laboratories) at room
temperature. Counterstaining was performed with haematoxylin.
55
4.4.4 TRACP staining for identifying osteoclasts (II, III)
The cells were stained for tartrate-resistant acid phosphatase (TRACP), a
commonly accepted marker of osteoclasts (Minkin, calcify tissue int 1982) using
a Sigma TRACP kit (no. 386-A, Sigma Chemicals, St.Louis, MO) according to
the manufacturer’s instructions. To visualize the nuclei, the cells were incubated
with the DNA-binding fluorochrome Hoechst 33258 (1 mg/ml stock diluted 1:800
in PBS, Sigma Chemical Co, St. Louis, MO) for 10 minutes at room temperature.
Study (II): The numbers of multinucleated TRACP-positive cells on each
bone slice were counted, using a Nikon Eclipse 600 microscope and Nikon Plan
Fluor 20x/0.50 objective with an appropriate filterset.
Study (III): The numbers of the multinucleated TRACP-positive cells in
contact with cartilage or newly formed bone trabeculae were counted from five
microscope fields using a Nikon Eclipse 600 microscope with a 40x/0.50
objective with an appropriate filterset.
4.4.5 Area Measurement of Excavated Pits (II)
Prior to the staining of the pits, total detachment of the cells from the bone slices
was ensured by wiping the cellular surface of the slices with a soft brush. The pits
were stained with peroxidase-conjugated wheat germ agglutinin-lectin (WGA; 20
μg/ml, Sigma Chemical Co., St.Louis, MO) for 20 minutes, washed with PBS,
and incubated for 5 minutes in diaminobenzidine (0.5 mg/ml) +0.03% H2O2.
Resorption pits were visualized as brown-colored areas after staining with WGA.
Morphometric analysis of the resorption pits was performed with an MCID image
analyzer, utilizing M2 software (Imaging Research Inc., Brock University,
Ontario, Canada) to the quantify areas of the resorption lacunae.
4.5 Biological assays
4.5.1 Determination of cell proliferation (WST-1) (IV)
MC3T3-E1 cell proliferation was measured by a WST-1 assay (Roche, Germany),
which quantifies the reduction of the tetrazolium salt WST-1 by mitochondrial
dehydrogenases in viable cells. Analyses were performed 3, 7, 10 and 14 days
after the cells were plated. WST-1 reagent (20µl) and α-MEM with ascorbic and
β-glycerophosphate (200 μl) were added to the cells and the culture incubated at
56
+37°C (5% CO2, 95% air) for 1 hour. The absorbance of the reaction product was
measured with a 1420 Victor2 Multilabel counter (Wallac, Turku, Finland) at a
wavelength of 450 nm.
4.5.2 Alizarin red S staining and quantification of calcium deposition (IV)
The mineralization of matrix and nodules was determined using Alizarin red S
staining. Briefly, MC3T3-E1 cell cultures were fixed at culture day 14 with 3%
PFA/ 2% sucrose in PBS for 10 minutes at room temperature, washed with PBS
and stained for 30 min with a 1% solution of Alizarin red S (Amresco, USA) pH
4.2 at room temperature with rotation. The cultures were washed three times with
deionized water and once with 70% ethanol before the addition of 100 nM
cetylpyridinium chloride (Sigma) for 1 hour to solubilize and release calcium-
bound Alizarin red into the solution. The solution was collected and the Alizarin
red S concentration was determined by absorbance measurements at 570 nm
using an Alizarin red S standard curve in the same solution.
4.6 Radiographical analysis
4.6.1 Radiographic evaluation of bone formation (III)
Standard lateral position radiographs (100 mA, 20 kV, 0.08 s/exp; Mamex de
Maq, Soredex, Orion, Finland) were taken from all the hind legs of the mice.
Calibration of the equipment for the measurement of optical density was
performed using an aluminum wedge. New bone formation was evaluated as the
area (in mm2) of calcified tissue visible in the radiographs with a digital image
analysis system MCID M4 (Imaging Research Inc., St Catharines, Canada). The
radiographs were digitized on a light table with a ccd camera (Dage MTI 72E,
Dage- MTI, Inc., Michigan City, IN, USA) using a Micro Nikkor 55mm objective
(Nikon, Tokyo, Japan).
57
4.6.2 Peripheral quantitative computed tomography (pQCT) (I, IV)
Bones were scanned with a Stratec XCT 960 A peripheral quantitative computed
tomography (pQCT) system with software version 5.21 (Norland Stratec
Medizintechnic GmbH, Birkenfeld, Germany).
Study (I): The bone mineral content of the femurs treated with AdVEGFAor
AdlacZ was analyzed at 1, 2, and 4 weeks after the operation. Distal femurs were
scanned for three to five consecutive cross-sectional images with slice distances
of 0.5 mm. Bones were scanned with a voxel size of 0.148 × 0.148 × 1.25 mm3. A
slice representing the central area of the defect was chosen for analysis, and the
total bone mineral content (BMC, mg/mm) was recorded.
Study (IV): To evaluate to effects of endostatin in vivo, a voxel size of 0.148
× 0.148 × 1.25 mm3 (transgenic mice overexpressing endostatin and FVB/N
control mice) and 0.092 × 0.092 × 1.25 mm3 (mice deficient in collagen XVIII
(Col18a1-/-) and C57BL/6J control mice) were used for the measurements. The
scout view of the pQCT system determined the middle point of each bone, from
which one cross-sectional slice was scanned and the total mineral content and
density measured.
4.6.3 Faxitron X-ray imaging (IV)
Radiographs of the dissected bones were taken in anteroposterior (AP) and lateral
projection on a Faxitron Specimen Radiography System (Model MX-20,
Wheeling, IL, USA) with a high-resolution algorithm. The imaging parameters
were identical for all the specimens and the following settings were used: 6 s, 30
kV for the 30 day old mice and 20 kV for the 14 day old mice, pixel matrix
1024 × 1054; resolution 200 pixels mm-1. The images were transferred to a
personal computer for analysis. The lengths and areas of tibias and femurs were
measured from the radiographs. As the weight of the mice varied from litter to
litter in both mutant and control mice, we normalized the bone density data by the
length and area of the mouse bones measured from the Faxitron images.
4.7 Statistical analysis (I-IV)
The values are the mean ± SEM. Significance was calculated using one-way
ANOVA followed by Student’s t-test with the statistical package Origin 6.0 or
7.5. (Microcal Software Inc., Northampton, MA)
58
59
5 Results
5.1 Bone defect healing model (I)
5.1.1 Bone mineral content evaluated with peripheral quantitative computed tomography (pQCT) (I)
The bone mineral content (BMC) at the injury site was analyzed by peripheral
quantitative computed tomography (pQCT) revealing a significant increase in
BMC in the VEGF-treated femurs (13.9 mg/mm vs.12.1 mg/mm, difference
14.9%, p < 0.001) 2 weeks after the operation. At the 1-week and 4 week time
points, there was no statistically significant difference between the two sides.
5.1.2 Induction of vascularization (I)
The number of capillaries was significantly increased at the 1-week time point
only (5.5 in control group vs. 7.5 in VEGF group, 36.4% difference, p < 0.01),
after which the difference was no longer significant.
5.1.3 Histological evaluation (I)
The histology of the tissue sections was analyzed in order to demonstrate
differences in healing between the VEGF-treated and the control femurs. One
week after the procedure (Figures 6A and 6B), both sides still had large
hematomas with surrounding seams of mesenchymal stem cells. The
morphological difference in the amount of non-ossified callus is most obviously
seen at the 2-week time point (Figures 6C and 6D), indicating amplified
ossification in the VEGF-treated femurs. The proportion of fibrous tissue
representing residual defect was significantly smaller in the VEGF group 2 weeks
after the operation (19.5% in VEGF group vs. 31.0% in control group, 59.0%
difference, p < 0.05). At the 4-week time point (Figures 6E and 6F), the defect
was difficult to localize due to almost complete healing and little difference was
seen between the VEGF overexpression and the control sides.
60
Fig. 6. Example of histology of bone regeneration in the drilling defects in the VEGF-
treated (right panel) and the control femurs (left panel) 1 (A, B), 2 (C, D), and 4 weeks
(E, F) after the operation. HE-stained decalcified sections. Scale bar: 1mm.
Endochondral ossification took place at the periosteal rim of the drilling defect
whereas direct ossification occurred at the central part of the healing metaphyseal
defect. The amount of hyaline cartilage was high at the early phases of healing,
61
but the VEGF treatment stimulated rapid replacement of the cartilage with mature
bone. At 2 weeks, the amount of cartilage was significantly lower in the VEGF
group compared with the control (0.015 mm2 vs. 0.065 mm2, 3.3-fold difference,
p < 0.01). At 4 weeks, no cartilage was detected in the defect area. The data
support the hypothesis that VEGF overexpression significantly enhanced the
endochondral ossification (Figure 7).
Fig. 7. Examples of cartilage at the periosteal defect side. The control defect 1 week
after the operation (A) had a lower amount of cartilage compared with the VEGF-
treated femur (B). At 2 weeks, the amount of cartilage was significantly higher in the
control (C) compared with the VEGF group (D). Hematoxylin-eosin staining of paraffin-
embedded tissue. Scale bar 50 µm.
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5.2 Effects of VEGF-A and endostatin on osteoclastic bone resorption and differentiation in vitro (II)
5.2.1 Bone resorption assay (II)
The number of osteoclasts in each sample group was counted by analyzing
multinuclear TRACP positive cells in the resorption assay. No significant
differences in the number osteoclasts were found between the different groups.
The total resorbed area was significantly increased in response to VEGF-A
treatment and the effect was dose dependent. Endostatin alone had no inhibitory
effect on the resorbed area when compared to the basal level of osteoclastic bone
resorption. When both endostatin and VEGF-A were supplemented, the
stimulatory effect by VEGF-A on bone resorption was limited to the basal level.
The resorption activity was even below basal level with the highest
concentrations (p < 0.05 in ctrl vs. VEGF100 and endostatin). Thus, endostatin
could block the VEGF-A-induced stimulatory effect on osteoclastic bone
resorption.
5.2.2 Osteoclast differentiation (II)
VEGF-A was added to the medium containing M-CSF and RANKL in order to
achieve the maximal differentiation stimulus. Endostatin inhibited the maximal
osteoclastogenesis alone or in combination with VEGF.
5.3 Endochondral ossification model (III)
5.3.1 Radiographic evaluation of bone formation (III)
The effect of the recombinant adenoviral endostatin gene transfer with or without
VEGF-A in bone formation was first evaluated from radiographs. X-ray images
of the dissected legs displayed significant bone formation in all treatment groups
at the 2- and 3-week time points. Two weeks after the injection, there were no
significant differences between the study groups. At the 3-week time point,
however, we could see retardation of bone formation in radiographs of the
endostatin-treated animals compared to the VEGFA- treated and the control
63
groups. In addition, the retardation was clearly visible in the animals that received
a combination of VEGF-A and endostatin.
5.3.2 Induction of vascularization (III)
The samples were processed for immunohistochemistry in order to demonstrate
vascular structures positive for the FVIII-related antigen. At the 1-week time
point, previously existing vessels (Figures 8A and 8G) were easily identified due
to the well-defined endothelium. At all time points, endothelial staining at the
bone formation site was increased in the VEGF-A group compared to the control.
After 2 and 3 weeks, endostatin blunted the VEGF-A effect. Endostatin alone,
however, did not decrease the basal neovascularization (Figure 8). At the 2- and
3-week time points, staining for the factor VIII-related antigen indicated intense
and partially premature vascularization in the VEGF-A-stimulated ossification.
The new vessels were morphologically more mature in the endostatin group
compared to the VEGF-A stimulated angiogenesis.
64
Fig. 8. Bones were decalcified, cut into tissue sections, and immunostained for FVIII-
related antigen. The figure shows examples of BMP control (A–C), VEGF-A (D–F),
endostatin (G–I) and VEGF-A+endostatin (J–L) treated ossicles 1 week (A, D, G, J), 2
weeks (B, E, H, K) and 3 weeks (C, F, I, L) after the injection. Pre-existing vessels were
nicely stained all over the induction site. Examples of these old vessels are seen in (A)
and (C). Scale bar: 100 μm.
5.3.3 Bone histology (III)
The histology of the tissue sections was analyzed in order to demonstrate
differences in endochondral bone development. At the 2-week time point, there
was no difference in the proportional ossification of the cartilage phase.
Significant differences were seen in the area of cartilage remnants of the bone
nodules in different test groups 3 weeks after the injection. We could demonstrate
that endostatin inhibited cartilage resorption compared to control and VEGF-A-
65
treated animals. The proportional area of bone was lower in the endostatin group
compared to the control (Figure 9).
Fig. 9. (A) Example of new bone formation induced by BMP injection. Ectopic new
bone was evaluated as a proportional area of bone tissue/osseous nodule 2 and 3
weeks after the injection. (Arrow indicates the cortical shell of the ossicle and ∗
indicates cartilage). (B) At the 3-week time point endostatin inhibited the proportional
area of bone tissue. At 1 or 2 weeks, no difference in proportional area of bone
tissue/ossicle was seen. ∗∗p < 0.01. Scale bar: 200 μm.
66
5.3.4 The number of cartilage-resorbing chondroclasts (III)
Cartilage resorption was studied by identifying the number of TRACP-positive
chondroclasts/osteoclasts. VEGF-A alone stimulated osteoclasts as demonstrated
by the increased number of TRACP-positive cells 3 weeks after the operation.
Endostatin gene transfer decreased the amount of TRACP-positive cells
compared to the VEGF-A group at the 2- and 3-week time points. Endostatin also
decreased the VEGF-A stimulated osteoclastogenesis at the 2-week time point.
5.4 The effects of endostatin on the MC3T3-E1 osteoblastic cell line
5.4.1 Osteoblast proliferation in vitro
MC3T3-E1 cell proliferation was measured by a WST-1 assay. On days 3–10
there were no statistical differences between the groups. On day 14 of culture
endostatin 20 μg/ml decreased osteoblast proliferation compared to the basal
control level (p < 0.05) and VEGF-A group (p < 0.05). Endostatin 2 μg/ml also
decreased osteoblast proliferation, but not significantly. Endostatin 2 μg/ml
together with VEGF-A decreased osteoblast proliferation compared to the basal
level (p < 0.001), VEGF-A (p < 0.001) and endostatin 2 μg/ml (p < 0.05).
Interestingly, VEGF-A together with the higher dose of endostatin, 20 μg/ml did
not significantly affect proliferation.
5.4.2 Matrix mineralization in vitro
The immature osteoblast cell line, MC3T3-E1, up-regulates osteoblast-specific
genes and produces mineralized nodules undergoing the developmental stages
associated with differentiating osteoblasts. The process of mineralization is
initiated around day 10. A low dose of endostatin 2 μg/ml together with VEGF-A
decreased the mineralisation compared either to the basal control level, VEGF-A
or endostatin alone (p < 0.05). Endostatin 20 μg/ml together with VEGF-A does
not significantly affect mineralization when compared to the basal control level,
but if compared to treatment with VEGF-A alone, a decrease in matrix
mineralization can be observed (p < 0.05).
67
5.5 Transgenic mice and knockout mice
5.5.1 pQCT and faxitron results
The bone mineral content or density, as analyzed with pQCT, was not altered in
ES-tg or Col18a1-/- mice as compared to their FVB/N or C57BL/6J controls.
Moreover, there were no significant differences in bone sizes between the
Col18a1-/- mice and their controls. The lengths of tibias and femurs were
measured from radiographs. In the ES-tg mice the lengths of femurs and tibias
were smaller at the age of 14 days, compared to FVB/N controls (p < 0.01). This
delay of growth was transient, as no difference was seen at 30 days of age.
5.5.2 The formation of secondary ossification centers
In the bones of 14-day-old Col18a1-/- mice we detected a delay in the formation
of secondary ossification centers. In all C57BL/6J mice the secondary ossification
centers were already clearly visible in the epiphyseal areas of femurs. However,
in Col18a1-/- mice the secondary ossification centers were either missing, or very
small (Figure 10). At the age of one month, no clear differences could be
observed between the knock-out and control mice: in both, the ossification of the
epiphyses had occurred. The growth plate was still visible in both mouse lines at
this age. In ES-tg mice we could see no difference in the ossification of the
epiphyseal area as compared to their FVB/N controls.
68
Fig. 10. In 2-week-old C57BL/6J control mouse femurs (A) the formation of secondary
ossification centers is clearly visible in the epiphyseal areas, whereas in the Col18a1-
/- mutants (B) the secondary ossification centers are either very small, or, as in the
case of some samples, totally absent at this timepoint. In 2-week-old FVB/N (C) and
ES-tg (D) mice the secondary ossification centers are well developed, and no obvious
differences can be detected between these two mouse lines.
5.5.3 Localization of collagen XVIII
Immunohistochemistry was used to study the localization of collagen XVIII in
bone tissue. Collagen XVIII was localised in the blood vessels of bone marrow:
staining was seen with the antibody against all three variants (anti-all moXVIII).
The short variant of collagen XVIII was responsible for this immunosignal, as the
antibody that stains the two longer variants of collagen XVIII (anti-long
moXVIII) did not stain the vessels (Figure 11). Staining with collagen XVIII
antibodies also gave a signal in the epiphyseal cartilage, surrounding the
chondrocytes, but this signal proved to be unspecific as similar staining could be
seen in cartilage of mice lacking collagen XVIII.
69
Fig. 11. Collagen XVIII was localized in the central vessels and some smaller vessels
of the bone marrow (A and B) and vessels of the periosteum (C) and at the skeletal
muscle attachment site (D). As expected, the collagen XVIII KO used as negative
controls showed no staining (E and F). The samples are from 14-day-old mice.
70
71
6 Discussion
6.1 Adenoviral VEGF-A gene transfer induces angiogenesis and
promotes bone formation in healing osseous tissues (I)
Angiogenesis has a crucial role in bone healing. VEGF is expressed within the
first weeks of bone healing (Uchia et al. 2003) and it may enhance osteoblast
differentiation (Deckers et al. 2000). Pufe and colleagues have shown that VEGF
is expressed in fracture callus at the beginning of fracture healing after which
expression decreases by day 5 (Pufe et al. 2002). We studied the effect of VEGF-
A on bone healing and the feasibility of first –generation adenoviral vector to
deliver VEGF overexpression at the bone injury site.
The exogenous VEGF protein is an unstable, short-lived protein in vivo and
moreover, costly to produce (Keramaris et al. 2008). The use of gene therapy for
bone regeneration enables delivery of VEGF at physiological levels using natural
cellular mechanisms. In order to achieve fracture healing, the required therapeutic
level of VEGF must be maintained for a period of time. A first-generation
adenoviral vector typically produces expression that lasts for 1–2 weeks in vivo,
after which the virus may be eliminated by the immune response (Cao et al. 2002,
Ylä-Herttuala et al. 2000, Okubo et al. 2001). The rodent model is useful for
studying bone healing properties, since rodent bone healing is a very rapid
process. Bone regeneration occurs within 4 weeks even in the absence of
interventions (Uusitalo et al. 2001).
In this study, the defects were healed finally in both groups, but the healing
process was accelerated in the VEGF treated group. In order to monitor bone
recovery, we first measured the bone mineral content (BMC) using peripheral
quantitative computed tomography (pQCT). A number of studies have used
pQCT in mice and rats to observe cortical bone geometry along with cortical and
trabecular bone mineral density (BMD). This imaging method has been used to
monitor such changes after pharmacologic or mechanical interventions and to
monitor fracture healing (Breen et al. 1998, Armamento-Villareal et al. 2005,
Gasser et al. 2008, McCann et al. 2008). Locally VEGF-A overexpression
increased the bone mineral content measured with (pQCT) 2 weeks after the
operation. At the 1-week time point, there was as yet no statistically significant
difference between the two sides and at the 4-week time point, the mineral
72
content of both VEGF-treated and control bones was decreased due to the
reorganization of the callus at the injury location. Endochondral ossification took place at the periosteal rim of the drilling
defect whereas intramembranous ossification occurred at the central part of the
healing metaphyseal defect. The importance of angiogenesis and VEGF on
endochondral ossification has been pointed out in the study of Gerber and
colleagues (Gerber et al. 1999). Increased vasculature stimulates
intramembranous ossification as well (Collin-Osdoby 1994, Zelzer et al. 2002,
Horner et al. 1999).
Factor VIII expression is normally restricted to vascular endothelial cells and
therefore Factor VIII immunostaining is considered to be a reliable method to
detect blood vessels in dense connective tissue. In the present study, the number
of FVIII-related capillaries was significantly increased at the 1-week time point
providing increased blood supply to the healing bone and accelerated healing of
the defect. At 2 or 4 weeks, the difference could not be seen anymore. This could
be due to VEGF production at the early phases of bone healing and viral
expression may shut down soon after the virus delivery. Furthermore, skeletal
muscle is not an ideal target tissue for the recombinant adenoviral vector due to
the low number of viral receptors, the quiescence of myoblasts in mature muscle,
and physical barriers (Cao et al. 2002, Nalbantoglu et al. 2001). Our results thus
support previous results that increased vasculature also stimulates
intramembranous ossification (Collin-Osdoby 1994, Zelzer et al. 2002). The
capillaries were more disorganized in the VEGF-A stimulated condition
compared with the control. The induction of disorganized vascularization
indicates that the viral gene transfer resuted in increased VEGF-A activity. It
could be possible to adjust VEGF dosing with other growth factors to reduce this
phenomenon.
In this study, a first-generation adenoviral vector produced VEGF
concentrations with biological effects on bone metabolism and was able to
modify the histological features of bone healing. The morphological difference in
the amount of non-ossified callus was most obviously seen at the 2-week time
point, where the proportion of fibrous reparative tissue was more efficiently
cleared and replaced with bone in the VEGF-A group, indicating amplified
ossification in the VEGF-A treated femurs. We also measured the amount of
hyaline cartilage at the periosteal rim, and at the early phases of healing the
amount of cartilage was high. However, VEGF treatment stimulated rapid
replacement of the cartilage with mature bone. At the 4-weeks time point the
73
defect was difficult to localize due to almost complete healing, and only little
differences were seen between groups.
6.2 Endostatin inhibits VEGF-A induced osteoclastic bone resorption in vitro (II)
The intimate contact of the bone remodeling site with vascular sinusoids indicates
interplay between the vascular endothelium and the bone cells. A continuous
supply of bone-degrading osteoclasts is needed during bone development.
Osteoclasts are derived from hematopoietic precursors present in both the bone
marrow and the peripheral circulation (Roodman 1999). Osteoclast precursors
need to adhere to and migrate through endothelium to reach future sites of bone
resorption. It has been reported that VEGF is potentially a monocyte
chemoattractant (Clauss et al. 1996). Monocytes have been shown to express
VEGFR1, but not VEGFR2 (Barleon et al. 1996). Soluble Flt-1 blocks invasion
of osteoclasts into the marrow cavity of developing long bones in vivo (Niida et al. 1999). Mature osteoclasts express VEGFR1 and VEGFR2 on the cell surface
(Gerber et al. 1999, Niida et al. 1999, Nakagawa et al. 2000). Endostatin is a
known antagonist of VEGF-mediated endothelial cell migration and proliferation
(Yamaguchi et al. 1999, Taddei et al. 1999). Interestingly, the recruitment and
invasion of osteoclast precursors can be disturbed by endostatin in the metatarsal
model, in which fusion and migration of osteoclast precursors is required for
osteoclastic bone resorption (Henriksen et al. 2003).
Only little is known about the effects of endostatin on other cells and thus we
investigated the possibility that VEGF and endostatin may control
osteoclastogenesis and osteoclastic bone resorption in vitro. Endostatin has been
shown to bind αv- and α5-integrins (Rehn et al. 2001), and resorbing osteoclasts
express at least αv-integrins (Chuntharapai et al. 1993, Duong et al. 2000).
Therefore it is possibile that endostatin could control osteoclast function.
We used a classical resorption pit assay and we analysed the number of
osteoclasts by counting the multinuclear TRACP-positive cells. However
significant differences between the different groups were not detected. Next we
measured osteoclastic bone resorption. In response to VEGF-A treatment the total
resorbed area was significantly increased and the response was dose dependent.
Endostatin alone had no inhibitory effect on the resorbed area when compared to
the basal level of osteoclastic bone resorption, but when both endostatin and
VEGF-A were present, the stimulatory effect by VEGF-A was reduced to the
74
basal level. Our finding concerning the increased resorption achieved by VEGF is
consistent with the result of Nakagawa and colleagues (Nakagawa et al. 2000).
However, they demonstrated that VEGF-A enhanced the survival of pure rabbit
osteoclasts, although in our resorption pit assay containing osteoblasts and
osteoclasts, the number of TRACP-positive cells remained unchanged. The data
show that the increased total resorbed area in our model is not due to enhanced
survival rate. The observed effect is more likely a result of a direct VEGF-A
stimulation of osteoclastic bone resorption.
Since the basal resorption was not affected by endostatin in the pit assay, we
investigated wheter endostatin could inhibit the differentiation of osteoclasts.
Bone marrow hematopoietic cells differentiate into osteoclasts in the presence of
RANKL and M-CSF. When VEGF-A was added to the medium containing
M-CSF and RANKL, endostatin inhibited osteoclastogenesis compared to the
VEGF induced maximal stimulus. Interestingly, we could see the inhibitory effect
also in the absence of VEGF-A.
It seems, however, that the regulatory effect of endostatin is not critical since
endostatin alone cannot modify the basal bone resorption. In the study of
Henriksen and colleagues, endostatin completely inhibited VEGF mediated
osteoclast invasion but had no effect on RANKL or M-CSF induced migration. In
cultures of purified osteoclasts both RANKL and VEGF induced phosphorylation
of ERK1/2 MAP kinase (Henriksen et al. 2003). It is possible that endostatin may
not only limit the migration of osteoclast precursors, it may also inhibit VEGF-A-
induced osteoclast activation. Pufe and colleagues have detected a similar
influence of endostatin on hyaline articular cartilage. Endostatin alone had no
effect on the basal levels of the phosphorylation of ERK1/2, whereas co-
incubation with endostatin blocked VEGF-induced ERK1/2 phosphorylation in
immortalized human chondrocytes (C28/I2) (Pufe et al. 2004c, Pufe et al. 2004d).
6.3 Endostatin inhibits endochondral ossification (III)
Angiogenesis is essential for the replacement of cartilage by bone during skeletal
growth and regeneration as well as on ectopic ossification. The function of
endostatins during the angiogenesis process of bone formation is quite unknown.
In our previous studies we observed that VEGF-A stimulates osteoclasts and
osteoclastogenesis thus regulating the duration of the cartilage phase in this
process. In addition, endostatin inhibited osteoclastogenesis and osteoclast
75
function in vitro indicating that endostatin may coordinate angiogenesis and
ossification in vivo.
It is hypothesized that endostatin expression by mature cartilage may
suppress osteoclasts while endostatin-deficient ossifying cartilage allows
osteoclastic activity and endochondral ossification (Pufe et al. 2004c). Moreover,
it has been suggested that endostatin has a homeostatic function in cartilage
metabolism (Feng et al. 2005). Recently, endostatin has been found to repress the
development and volume of tissue grafts in mice with rheumatoid arthritis, which
is characterized by an inflammatory erosive synovitis, where invasion of
inflammatory cells into the joint synovium is accompanied by excessive
angiogenesis causing destruction of the cartilage and fibrous tissue (Matsuno et al. 2002, Yin et al. 2002).
Since the role of endostatin in the presence of existing stem cells in vivo
requires further exploration, we employed an endochondral ossification model in
the mouse hind muscle pouch stimulated by BMPs. Endostatin was overexpressed
with or without VEGF-A using a recombinant adenoviral vector. BMPs have been
used to heal bone defects in experimental animal tests, but also in clinical
settings, mainly in orthopedic and oral surgery (Pekkarinen et al. 2005).
The muscle pouch model of mouse has been a standard method for studying
the activity of BMP since its discovery and has been shown to be an appropriate
animal model (Sampath & Reddi 1981). In this study, we used injectable BMP
implants, which require only a minimally invasive technique. Previously, they
have been investigated in many animal experiments with promising results
(Forslund et al. 1998, Bax et al. 1999, Muschik et al. 2000, Blokhuis et al. 2001,
Li et al. 2002). The native reindeer BMP extract we used (Pekkarinen et al. 2003)
contains a cocktail of purified proteins consisting of multiple BMPs and other
non-BMP proteins combined with gelatin gel (Aldinger et al. 1991, Iwata et al. 2002, Jortikka et al. 1993a, Jortikka et al. 1993b). Although individual rhBMPs
are able to induce ectopic new bone formation, purified BMP extract from human
matrix has been demonstrated to produce new bone more efficiently than rhBMP-
2 (Bessho et al. 1999). This suggests that the effect of native BMP extract
represents synergistic activity between different BMP proteins. We observed strong ectopic bone formation in all our study groups that was
induced with BMP extract combined with gelatin gel in conjugation with the
viruses. First we evaluated the new bone formation from the X-ray images taken
from the dissected legs at 2- and 3-week time points. At the 1-week time point,
the mineralization of cartilage had not yet started. At 2 weeks, the difference
76
between the study groups was not significant, but we observed less bone
formation in endostatin-treated animals at the 3 weeks time point. Contrary to our
previous findingwith the bone drilling defect model, VEGF overexpression did
not enhanced bone formation. This may indicate that the surrounding tissue may
have regulatory properties that control ossification.
Peng and others have described a synergistic enhancement of VEGF and
BMP-4 or BMP-2 on bone formation (Peng et al. 2002, Peng et al. 2005).
However, VEGF alone did not promote bone regeneration when using a model
where the healing of cortical defects in the parietal bones of mice was studied.
They also reported that the beneficial effect of VEGF on bone healing elicited by
BMP-4 depends critically on the ratio of VEGF to BMP4, with an improper ratio
leading to unfavorable effects on bone healing, while disruption of the BMP-
2/VEGF ratio has less negative effects on bone formation (Peng et al. 2002). Furthermore, Kakudo and others reported an increased amount of VEGF
induced ectopic bone formation seen in X-rays. They implanted human
recombinant BMP-2 in rat calf muscle using a disc implant (Kakudo et al. 2006).
Thus it is possible that species differences and the mode of application of the
BMP and VEGF/BMP ratio influence bone formation. Moreover, Murphy and
colleagues suggested that the increased vascularization achieved with VEGF
enhances mineralized tissue generation, but not necessarily osteoid formation
when using VEGF with the bio-mineralized scaffold compared to mineralized
scaffold alone (Murphy et al. 2004).
In this study, we could not see increased endochondral bone formation caused
by VEGF-A overexpression in the mesenchyme of the muscular space indicating
that the amount of BMP determines the quantity of bone to be formed at the
ectopic site. VEGF-A however, stimulated the number of cartilage resorbing
osteoclasts and the number of FVIII-related antigen-positive blood vessels thus
accelerating the endochondral phase. Endostatin overexpression alone or in
combination with VEGF-A decreased the number of TRACP-positive cells
compared to animals with VEGF-A overexpression at 2- and 3-week time points
suggesting that cartilage resorption by chondroclasts is decreased in response to
endostatin even in the presence of VEGF-A. These results are in line with our
previous results showing that endostatin inhibits VEGF-A-stimulated osteoclastic
bone resorption in vitro independent of VEGF-A. In addition, as expected,
endostatin prevented the formation of vessels and thus inhibition of vascular
development. Endostatin could reduce the amount of BMP-induced bone
77
formation, since cartilage resorption and osteoblast function are dependent on
blood vessel formation.
6.4 Endostatin inhibits proliferation of MC3T3-E1 osteoblasts and together with VEGF-A suppress mineralization (IV)
We investigated how endostatin affects the behaviour of osteoblasts. A number of
studies have pointed out the significant role of canonical Wnt signaling in the
control of osteoblastogenesis and bone formation (Yavropoulou & Yovos 2007)
and recently it was reported that endostatin is a potent inhibitor of Wnt signaling
(Hanai et al. 2002). Thus, endostatin may have a role in the anabolic effect of
bone. Mechanical loading generates interstitial fluid flow that exerts a shear stress
at surfaces of osteoblasts and osteocytes. The canonical Wnt pathway is one of
the pathways that is up-regulated in response to the fluid shear stress, thus
translating fluid shear signals into biological effects in bone cells. Endostatin can
prevent the effect of fluid shear stress on osteoblasts completely (Lau et al. 2006). In the present study we used MC3T3-E1 cells, which behave as immature,
committed osteoblastic cells and go on to differentiate in response to intracellular
and extracellular cues. We measured MC3T3-E1 cell proliferation using a WST-1
assay and the mineralization of matrix and bone nodules was determined using
Alizarin red S staining. VEGF alone had no effect on MC3T3-E1 cell
proliferation, nor a significant effect on mineralization, which is consistent with
previous findings (Kitagawa et al. 2005). Proliferative effects of VEGF, at least in
endothelial cells, are mediated by Flk-1/KDR (Guo et al. 1995, Quinn et al. 1993), that is not expressed by MC3T3-E1 cells, which could explain why VEGF
did not have a proliferative effect on these cells. Kitagawa and others suggested
that since VEGF has earlier been shown to induce markers of osteoblast
differentiation, e.g. alkaline phosphatase and osteocalcin expression, but not to
increase mineralization, other factors are needed in addition to VEGF for the
induction of mineralization (Kitagawa et al. 2005). However, treatment of cells
with endostatin alone decreased the proliferation of osteoblasts. Endostatin 20
μg/ml decreased osteoblast proliferation on day 14 of culture, and endostatin 2
μg/ml also slightly decreased osteoblast proliferation but this difference was not
statistically significant. The addition of endostatin alone did not affect
mineralization either. Interestingly, again we could see the highest inhibitory effect of endostatin
when it was used together with VEGF. A low dose of endostatin (2 μg/ml)
78
together with VEGF decreased osteoblast proliferation compared to the basal
level, VEGF-A, and endostatin 2 μg/ml groups. Likewise, matrix mineralization
was inhibited when a low dose endostatin (2 μg/ml) added together with VEGF-A
was used compared to control, VEGF-A, or endostatin groups. With a higher
concentration of endostatin (20 μg/ml) combined together with VEGF-A, this
inhibitory effect could not seen. It is not known why the inhibitory effect of
endostatin is highest when it is combined with VEGF. The concentration of
endostatin may be crucial for obtaining a certain effect. A recent report indicates
that the doses of endostatin required for meaningful effects in vitro and in animal
models vary over a wide scale (Sun et al. 2009). It has also been shown, that
endostatin’s effects on angiogenesis and tumor growth are biphasic, operating
over a U-shaped curve (Celik et al. 2005). Endostatin has been shown to block the interaction of VEGF with both Flt-1
and Flk-1/KDR receptors (Kim et al. 2002). Recently endostatin has been found
to bind to VEGFR3 (Kojima et al. 2008), which has a role in osteoblast
differentiation (Orlandini et al. 2006). Furthermore, endostatin has been shown to
affect the VEGF-mediated signaling pathway (Kim et al. 2002) and also ERK1/2
signaling (Schmidt et al. 2006). Since MC3T3-E1 cells do not express Flk-
1/KDR receptors, endostatin can only interact with the Flt-1 or VEGFR3, and
VEGF-A only with the Flt-1 receptor and therefore more endostatin peptides may
be available for binding to VEGFR3.
6.5 The role of collagen XVIII/endostatin on bone development in vivo in transgenic mice overexpressing endostatin and mice
lacking collagen XVIII
Our previous studies and those of others, have suggested that endostatin seems to
have a number of effects on bone and therefore, we wanted to further explore its
role in bone development in vivo using transgenic mice overexpressing endostatin
(ES-tg) and mice lacking collagen XVIII (Col18a1-/-). However, it has been
reported that collagen XVIII/endostatin does not have a role as a critical negative
regulator of angiogenesis during development and postnatal growth, since a lack
of collagen XVIII and endostatin in mice and humans does not increase
angiogenesis in major organs (Marneros & Olsen 2005).
In this study, we detected a delay in the formation of secondary ossification
centers in the bones of 14-day-old Col18a1-/- mice. At the age of one month the
ossification of the epiphyses had already occurred and clear differences were not
79
observed. Delayed ossification was also not detected at embryonic stages E15.5
and E18.5. In ES-tg mice no difference in the ossification of the epiphyseal area
could not be seen. Moreover, the bone mineral content or density, as analyzed
with pQCT was not altered and there were no significant differences in bone sizes
between the Col18a1-/- mice and their controls.
The possible involvement of endostatin in Wnt signaling may explain the
delay in the formation of the ossification centers in Col18a1-/- mice. The longest
variant of collagen XVIII contains a Frizzled-module, which has been shown to
be able to inhibit Wnt/beta-catenin signaling by binding to Wnt-molecules and
hence inhibiting their binding to cell-membrane receptors. The lack of the longest
variant could result in altered Wnt signalling (Quelard et al. 2008). In mice that
constitutively overexpress active β-catenin in cartilage, growth plates were totally
disorganized, and the maturation of chondrocytes and endochondral ossification
were inhibited. The activation of Wnt/β-catenin in mature chondrocytes
stimulates hypertrophy, matrix mineralization, and the expression of VEGF,
MMP-13, and several other MMPs among others (Tamamura et al. 2005). It has
been found, that the phenotype of collagen XVIII deficient mice is similar to mice
expressing the VEGF120 and VEGF188 isoforms only. They have been shown to
have differences in solubility and matrix binding properties and also in their
ability to bind to heparin and heparan sulfate proteoglycans. Since collagen XVIII
is a heparan sulfate proteoglycans it has proposed that collagen XVIII may be
involved in VEGF function and is required for the binding of to its receptor
(Hurskainen et al. 2005).
In ES-tg mice we observed a delay in the growth of bone length: femurs and
tibias were smaller at the age of 14 days. This difference could not be seen in 30-
day-old mice suggesting that this delay was temporary. Endostatin overexpression
takes place in the keratinocytes of ES-tg mice, but they show a seven-fold
increase in the concentration of endostatin in their circulation. It is possible that
the endostatin concentration in the bones is not high enough to cause any longer
lasting or permanent defects, but it only leads to a slight delay in the growth rate.
We used immunohistochemistry to determine the localization of collagen XVIII
in bone tissue. Collagen XVIII was detected surrounding the vessel structures in
bone marrow and periosteum, and at the skeletal muscle attachment site by the
antibody against all three variants. The short variant of collagen XVIII was shown
to be responsible for this immunosignal. Pufe and colleagues reported that
collagen XVIII/endostatin was expressed in human cartilage and fibrocartilage
(Pufe et al. 2004c). In present study, there was a signal in the cartilage, but this
80
proved to be unspecific as staining was also seen in the mice lacking collagen
XVIII. However, this can result from unspecific attachment of our primary
antibodies to the matrix surrounding the chondrocytes and thus it is still possible
that collagen XVIII is expressed in mouse cartilage.
Although mild delays in both mutant mouse lines were detected in the bone
development process, collagen XVIII/endostatin does not seem to have a crucial
role in skeletal development. Endostatin is one of the several regulators of
angiogenesis and it is likely that other factors may compensate for the loss of
collagen/endostatin in most tissues.
81
7 Conclusions
Our results confirmed the important role of VEGF in bone healing. VEGF
overexpression was delivered locally to the bone injury site using adenoviral
VEGF-A gene transfer, which was able to modify the histological features of
bone healing in a rodent model. The endochondral phase was completed earlier,
and also bone mineral content was enhanced in animals overexpressing VEGF.
The data from pit assay studies showed that VEGF-A induces bone resorption
and endostatin can suppress this VEGF-A induced resorption to the basal level. In
addition, osteoclastogenesis can be inhibited by endostatin. However, our findings
suggest that the regulatory effect of endostatin is not critical, since endostatin
alone does not modify the basal bone resorption.
The results from the endochondral ossification model, where the mouse hind
muscle pouch was stimulated by BMPs and endostatin was overexpressed with or
without VEGF-A using a recombinant adenoviral vector, showed that endostatin
retards the cartilage phase and subsequently reduces bone formation.
Endostatin has minor effects on the behavior of osteoblasts in cell culture
conditions. These effects are enhanced when cells are treated together with
VEGF.
Finally, we investigated the role of endostatin on bone development in vivo
using transgenic mice overexpressing endostatin and mice lacking collagen
XVIII. Mild delays in both mutant mouse lines were detected in the bone
development process. The formation of secondary ossification centers in the
bones of mice lacking collagen XVIII was delayed, as well as the growth of bone
lenght in ES-tg mice. On the basis of these results, we conclude that collagen
XVIII/endostatin does not seem to have crucial role in skeletal development.
As an overall conclusion, VEGF-A overexpression accelerates bone defect
healing. Endostatin retards bone formation and seems to inhibit VEGF-A
stimulated effects on bone cells. However, collagen XVIII/endostatin does not
seem to have a crucial role in skeletal development.
82
83
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Original publications
I Tarkka T, Sipola A, Jämsä T, Soini Y, Ylä-Herttuala S, Tuukkanen J & Hautala T (2003) Adenoviral VEGF-A gene transfer induces angiogenesis and promotes bone formation in healing osseous tissue. J Gene Medicine 5: 560–566.
II Sipola A, Nelo K, Hautala T, Ilvesaro J & Tuukkanen J (2006) Endostatin inhibits VEGF-induced osteoclastic bone resorption in vitro. BMC Musculoskelet Disord 13(7): 56.
III Sipola A, Ilvesaro J, Birr E, Jalovaara P, Petterson RF, Stenbäck F, Ylä-Herttuala S, Hautala T & Tuukkanen J (2007) Endostatin inhibits endochondral ossification. J Gene Medicine: 1057–1064
IV Sipola A, Seppinen L, Pihlajaniemi T & Tuukkanen J (2009) Endostatin affects osteoblast behaviour in vitro, but collagenXVIII/endostatin is not essential for skeletal development in vivo. Calcified Tissue International 85(5): 412–420.
Reprinted with permissions from BioMed Central (II), John Wiley & Sons (I, III)
and Springer Science + Business Media (IV).
Original publications are not included in the electronic version of the dissertation.
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