Coordinated fibroblast growth factor and heparan sulfate regulation of osteogenesis

Gene 379 (2006) 79 – 91 www.elsevier.com/locate/gene

Coordinated fibroblast growth factor and heparan sulfate regulation of osteogenesis Rebecca A. Jackson a,1 , Victor Nurcombe a,b,2 , Simon M. Cool a,b,⁎ a

Laboratory of Stem Cells and Tissue Repair, Institute of Molecular and Cell Biology, Proteos, 61 Biopolis Drive, 138673, Singapore b Department of Orthopaedic Surgery, National University of Singapore, Singapore Received 22 February 2006; received in revised form 17 April 2006; accepted 20 April 2006 Available online 19 May 2006 Received by A.J. van Wijnen

Abstract Growth and lineage-specific differentiation constitute crucial phases in the development of stem cells. Control over these processes is exerted by particular elements of the extracellular matrix, which ultimately trigger a cascade of signals that regulate uncommitted cells, by modulating their survival and cell cycle progression, to shape developmental processes. Uncontrolled, constitutive activation of fibroblast growth factor receptors (FGFR) results in bone abnormalities, underlining the stringent control over fibroblast growth factor (FGF) activity that must be maintained for normal osteogenesis to proceed. Mounting evidence suggests that FGF signalling, together with a large number of other growth and adhesive factors, is controlled by the extracellular glycosaminoglycan sugar, heparan sulfate (HS). In this review, we focus on FGF activity during osteogenesis, their receptors, and the use of HS as a therapeutic adjuvant for bone repair. © 2006 Elsevier B.V. All rights reserved. Keywords: FGF; HS; Bone; Fracture repair; Heparin-binding growth factor; Osteoblast

1. Introduction Abbreviations: FGF, fibroblast growth factor; FGFR, fibroblast growth factor receptors; HS, heparan sulfate; HSPGs, heparan sulfate proteoglycans; GAG, glycosaminoglycan; HBGF, heparin-binding growth factor; ALP, alkaline phosphatase; OC, osteocalcin; OPN, osteopontin; ON, osteonectin; Runx2, runtrelated transcription factor 2; MSC, mesenchymal stem cells; GlcA, glucuronic acid; GlcN, glucosamine; IdoA, iduronic acid; BMP, bone morphogenetic protein; TGF, transforming growth factor; IGF, insulin-like growth factor; VEGF, vascular endothelial growth factor; GH, growth hormone; HGF, hepacyte growth factor; MAPK, the mitogen-activated protein kinase; ERK1/ 2, extracellular signal-related kinases 1/2; SAPK/JNK, stress-activated protein kinase/c-Jun N-terminal kinase; PKC, protein kinase C; PI3K, phosphatidylinositol 3 kinase; CMDBS, carboxymethyl benzylamide sulfonate; RGTA, regenerating agents. ⁎ Corresponding author. Laboratory of Stem Cells and Tissue Repair, Institute of Molecular and Cell Biology, Proteos, 61 Biopolis Drive, Singapore 138673. Tel.: +65 6586 9714; fax: +65 6779 1117. E-mail addresses: [email protected] (R.A. Jackson), [email protected] (V. Nurcombe), [email protected] (S.M. Cool). 1 Tel.: +65 6586 9709; fax: +65 6779 1117. 2 Tel.: +65 6586 9708; fax: +65 6779 1117. 0378-1119/$ - see front matter © 2006 Elsevier B.V. All rights reserved. doi:10.1016/j.gene.2006.04.028

More than five years ago, the World Health Organisation officially launched “The Bone and Joint Decade, 2000–2010” in an attempt to raise awareness of the growing burden of musculoskeletal disorders on society, and to help alleviate the suffering and costs associated with joint disorders through improvement of diagnosis and treatment. A key step in this endeavour is to improve the fundamental understanding of bone formation and, in particular, the intricate play among the molecular elements that contribute to this process. Bone formation is highly coordinated, beginning with the commitment of mesenchymal stem cells (MSCs) to an osteogenic fate and their subsequent differentiation and maturation into the major bone-forming cells, the osteoblasts (Fig. 1). This sequential progression is regulated, among other influences, by a diverse repertoire of growth and adhesive factors acting in autocrine/paracrine manners at specific developmental stages. Of particular interest are the fibroblast growth factor (FGF) family and their receptors (FGFR), which interact with cell-

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Fig. 1. Osteoblast lineage progression. The progression of bone progenitor cells through to the osteoblast phenotype is tightly controlled by a diverse repertoire of fibroblast growth factors (FGF) and their receptors (FGFR). This diagram illustrates the sequential stages of osteogenic commitment and differentiation into preosteoblasts that are responsible for cell growth, followed by their subsequent maturation into the major bone forming cells, osteoblasts. Osteoblasts will later become surrounded and separated from other osteoblasts by the matrix they produce, and terminally differentiate into osteocytes. At each stage, different FGF ligands are important in bone formation. In particular, FGFs-2, -9 and -18 have been shown to act at each of the stages of proliferation, differentiation and maturation, and FGF2 protects cells against apoptosis.

surface heparan sulfate proteoglycans (HSPGs) to coordinate cell-fate decisions. 2. FGFs and FGFRs in osteogenesis 2.1. FGFs in controlling osteoblast phenotype FGFs comprise a family of at least 23 structurally-related members that are expressed by almost all tissues during the key stages of development. FGFs are typically between 20 and 35 kDa and have long been recognised as important regulators of osteogenesis (Ornitz and Marie, 2002). In vitro, FGFs-2, -4, -9 and -18 have all been shown to regulate the growth of a spectrum of cell types within growing bone, in a host of species (Canalis et al., 1988; Iseki et al., 1999; Rice et al., 2000; Shimoaka et al., 2002; Walsh et al., 2003). More recently, FGFs have been shown to have mitogenic effects on not only precursor cells, but also on maturing osteoblasts (Fakhry et al., 2005). In addition to their role in cell proliferation, FGFs are also known to affect the various stages of osteoblast differentiation, directly influencing the expression of various matrix proteins, including alkaline phosphatase (ALP) and osteocalcin (OC), as well as the transcription factor, Runx2. ALP is an early marker of osteoblast differentiation and its expression is enhanced by FGF-2 stimulation in rat bone marrow precursor cells (Noff et al., 1989; Pitaru et al., 1993; Zhang et al., 2002). Runx2, also known as cbfa-1, is a member of the Runt-related family of transcription factors and has been shown to be essential for both skeletal patterning during embryogenesis and the progression of osteoblast differentiation (Ducy et al., 1997, 1999; Komori et al., 1997). Runx2−/− mice

die immediately after birth, with a complete lack of skeletal ossification (Komori et al., 1997; Otto et al., 1997), and previous studies have shown that Runx2 expression precedes that of OC and collagen (Ducy et al., 1997). Runx2 enhances OC transcription by binding to its cognate binding site, OSE2, on the OC promoter (Ducy et al., 1997; Xiao et al., 1997), and similar binding sites for Runx2 have been subsequently identified on the collagen α1 gene (Kern et al., 2001). Runx2 expression is stimulated by FGFs-2, -4 and -8 (Xiao et al., 2000; Zhang et al., 2002; Kim et al., 2003b), and FGFs-2 and -4 can enhance the transcriptional activity and DNA binding properties of Runx2 (Xiao et al., 2000; Kim et al., 2003b). Interestingly, the interaction between Runx2 and OSE2 is obligate for FGF-2 induction of OC expression (Xiao et al., 2002). FGF-9 can also increase OC gene expression and maintain Runx2 levels in calvarial cells (Fakhry et al., 2005). FGF-2 is also important in matrix mineralisation (Noff et al., 1989), and Fgf-2−/− mice show a marked reduction in trabecular architecture, as well as decreased trabecular bone volume, mineral apposition, and bone formation rates (Montero et al., 2000). Contrastingly, some earlier in vitro studies in calvarial cells and osteoblast cell lines demonstrated negative effects of FGF treatment on the expression of osteoblast markers. FGF-2 caused a decrease in the levels of ALP, OC and collagen mRNA in rat osteosarcoma cells in a dose-dependent manner (Rodan et al., 1989). In another study, FGF-2 transiently enhanced proline incorporation into collagen type I, but only 24–48 h after the stimulus had been withdrawn, whereas continuous FGF-2 stimulation inhibited proline incorporation (Canalis et al., 1988). The effects of FGFs on cell proliferation have also been inconsistent. Whilst FGF-2 stimulates cell proliferation in

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bovine (Globus et al., 1989), murine (Mansukhani et al., 2000) and human osteoblasts (Debiais et al., 1998) as well as in bone marrow precursor cells (Noff et al., 1989), FGF-2 treatment in vivo can lead to reduced proliferation and increased suture fusion in mice (Iseki et al., 1999). The reasons for these conflicting results remain unclear; however there is evidence to suggest that these discrepancies arise due to the stage-specific effects of FGF ligands. Treatment of immature osteoblasts with FGF-2 causes an increase in proliferation, whereas treatment of more mature osteoblasts with FGF-2 causes an increase in OC and matrix mineralisation in human cells (Debiais et al., 1998). Similarly, FGF-9 can also influence osteoblasts at various stages by increasing proliferation rates in mature osteoblast cells, increasing differentiation marker expression and modulating matrix mineralisation (Fakhry et al., 2005). FGF-18, which also regulates cell colony formation, has been shown to delay suture closure in FGF-18−/− mice (Ohbayashi et al., 2002). During calvarial bone formation, FGF-18 is expressed by both MSCs and differentiating osteoblasts; gene targeting experiments show that, in addition to delayed suture closure, mice deficient of FGF-18 show decreased proliferation and delayed terminal osteoblast differentiation (Ohbayashi et al., 2002). Fig. 1 illustrates the various stages of bone development and indicates the FGF ligands important during osteogenesis, demonstrating that several FGF ligands can influence cells at many stages. FGFs can also regulate the expression of other growth factors both in vitro and in vivo. FGF-2 and FGF-9 increase the expression of transforming growth factor (TGF)-β and bone morphogenetic protein (BMP)-2 in osteoblasts in vitro (Noda and Vogel, 1989; Fakhry et al., 2005), and FGF-2 has been shown to increase insulin-like growth factor (IGF)-I expression both in vitro (Zhang et al., 2002) and in vivo (Power et al., 2002). In addition, TGF-βs can promote the mitogenic effects of FGF-1 and FGF-2 in fetal calf calvarial cells (Globus et al., 1988) and in turn FGF-1 can increase TGF-β expression in proliferative and maturing osteoblast cells (Tang et al., 1996). Cell survival can also be regulated by FGF stimulation. Treatment of cells for 24–48 h with FGF-2 prior to the removal of serum can protect osteoblasts from apoptosis through an inhibition of caspase-2 and caspase-3 activity (Debiais et al., 2004). Furthermore, under hypoxic culture conditions, transfection of MSCs with FGF-2 before their transplantation into infarcted myocardia was shown to increase cell viability by three-fold, and reduce apoptosis by increasing the expression of the anti-apoptotic gene, Bcl2 (Song et al., 2005). 2.2. FGFs signal through MAP kinase activation Successful bone formation is dependent upon the interplay between the local expression of FGFs (Naski and Ornitz, 1998) and the spatiotemporal expression of four tyrosine kinase FGF receptors (FGFRs1–4) and their splice variants. FGFRs typically contain an extracellular ligand-binding domain and a highly conserved intracellular signalling domain connected by a transmembrane region. The extracellular domain contains 3 cysteine-flanked immunoglobulin-like domains (Ig loops), an acid box, a heparin-binding region and a cell adhesion molecule

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(CAM) recognition domain. The FGF binding site is thought to be situated between the second and third Ig loops, with the third Ig loop responsible for controlling FGF affinity (Szebenyi and Fallon, 1999). Splice variants of the receptors form through alternate splicing in the third Ig loop to create IIIb and IIIc isoforms. It is thought that the IIIb isoforms are restricted to ectodermal tissues whereas the IIIc isoforms are found in mesenchymal tissues (Kettunen et al., 1998; Szebenyi and Fallon, 1999; Beer et al., 2000). The intracellular region is the most highly conserved domain within the receptor family, containing kinase inserts and a number of tyrosine residues that are responsible for receptor signalling. The binding of FGF to the extracellular portion of its FGFR causes receptor dimerisation and subsequent autophosphorylation of intrinsic tyrosine residues (Fig. 2). This, in turn, causes downstream activation of specific signal transduction pathways (Szebenyi and Fallon, 1999). A large proportion of FGFR signalling is transduced through the activation of the mitogen-activated protein kinase (MAPK) pathways, including the extracellular signal-related kinases 1/2 (ERK1/2; also known as p42/44 MAPK), p38 MAPK, stress-activated protein kinase/c-Jun N-terminal kinase (SAPK/JNK), protein kinase C (PKC) and phosphatidylinositol 3 kinase (PI3K) pathways (Gardner and Johnson, 1996; Kanai et al., 1997; Maher, 1999; Delehedde et al., 2000; Mansukhani et al., 2000; Suzuki et al., 2000; Cailliau et al., 2001). Signalling is initiated following tyrosine phosphorylation of the FGF receptor substrate 2 (FRS2) protein, that interacts with the growth factor receptor-bound protein 2/Son of sevenless protein (Grb2/Sos) complex (Schlessinger, 1994; Kouhara et al., 1997) to activate signalling via the ERK1/2 pathway. Activated ERK1/2 is then translocated to the nucleus to induce changes in gene expression and activation of transcription factors. FRS2 functions as a lipid-anchored docking protein, mediating between the receptor and Grb2/Sos, which it recruits to the plasma membrane in response to FGF stimulation to induce activation of MAPK and other signalling pathways (Kouhara et al., 1997). FGF-2 activation of ERK1/2 has been shown to increase Runx2 phosphorylation and the expression of OC and osteopontin (OPN) (Boudreaux and Towler, 1996; Xiao et al., 2002; Kim et al., 2003b). FGF-2 also induces a phosphorylation of SAPK/JNK that is important for the release of vascular endothelial growth factor (VEGF) (Tokuda et al., 2003), and increases PI3K activity without a contiguous phosphorylation of Akt or the downstream effector p70 S6 kinase (Debiais et al., 2004). Other studies have shown that FGF-2 increases ERK1/2 activation independent of PI3K, as its specific inhibitor, wortmannin, could not inhibit FGF-2-mediated increases in ERK1/2 (Chaudhary and Hruska, 2001). PKC has recently been shown to increase the FGF-2-directed stimulation of Runx2 expression, as well as the post-translational modification of Runx2 protein, thereby increasing its transcriptional activity (Kim et al., 2003a). Furthermore, inhibition of PKC can completely block FGF-2-induced Runx2 expression, an effect that cannot be replicated with MAPK inhibitors (Kim et al., 2003a). In addition to FGF-2, FGF-18 can also up-regulate ERK1/2 phosphorylation in osteoblasts as well as both ERK1/2

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Fig. 2. FGF/FGFR signalling in osteogenesis. FGF signalling is mediated through its binding to high affinity receptor tyrosine kinases, FGFR. Ligand binding induces receptor dimerisation and the phosphorylation of intrinsic kinase residues. This, in turn, activates FRS2 which recruits the Grb2/SOS complex to the plasma membrane and causes the subsequent induction of a number of MAPK-mediated pathways. FGFs have been shown to activate ERK1/2, p38 MAPK, SAPK/JNK, PKC and PI3K pathways to induce a plethora of changes in a number of different tissue types.

and p38 MAPK phosphorylation in chondrocytes, all of which can be blocked by the use of specific kinase inhibitors (Shimoaka et al., 2002). 2.3. Mutations in FGFRs can lead to disfiguring bone disorders The importance of FGFRs for bone development was recently highlighted with the discoveries that mutations within the amino acid coding sequences of FGFRs1–3 were responsible for the formation of skeletal dysplasias characterised by craniosynostoses, limb abnormalities, and/or dwarfism (Muenke and Schell, 1995; Wilkie et al., 1995a; Burke et al., 1998; Naski and Ornitz, 1998). These gain-of-function mutations cause constitutive, uncontrolled activation of FGFR signalling, that can be independent of ligand binding (Wilkie et al., 1995b; Yamaguchi and Rossant, 1995; Naski and Ornitz, 1998). Point mutations in FGFRs1–3, which give rise to craniosynostosis syndromes such as the Apert, Pfeiffer, Crouzon and Jackson–Weiss syndromes, are primarily characterised by premature fusion of cranial sutures due to unregulated intramembranous bone formation (Webster and Donoghue, 1997). In contrast, dwarfism syndromes, such as achondroplasia, are mostly caused by mutations in FGFR3, which disrupt normal and regulated chondrocyte proliferation and differentiation during skeletogenesis (Peters et al., 1993). These disorders demonstrate the importance of maintaining stringent control over FGF activity for normal bone growth and development.

2.4. The roles of FGFRs during osteogenesis It has previously been shown that FGFR1 is expressed predominantly during osteoblast differentiation (Iseki et al., 1999; Zhou et al., 2000), whereas FGFR2 is involved in regulating osteoblast proliferation (Iseki et al., 1999; Yu et al., 2003). More recently, a role for FGFR2 in osteoblast differentiation has been suggested, as FGFR2IIIc−/− mice have a delayed onset of ossification, together with a decrease in Runx2 transcription (Eswarakumar et al., 2002). In contrast, FGFR3 has been shown to act as a negative regulator of osteogenesis; its primary function appears to be associated with the control of chondrocyte proliferation during endochondral ossification (Deng et al., 1996; Molteni et al., 1999; Murakami et al., 2004). However, it has been shown that inhibition of FGFR3 expression by twist inhibits the differentiation of osteoprogenitor cells into osteoblasts (Funato et al., 2001), suggesting that FGFR3 may also be involved in actively regulating osteoblast differentiation. Studies also reveal that young adult mice lacking FGFR3 are osteopenic, developing severe osteomalacia as a consequence of the altered expression of FGFRs and other osteoblast differentiation markers (Valverde-Franco et al., 2004). Furthermore, constitutive, ligandindependent activation of FGFR3, resulting from a Gly369Cys homozygous mutation, leads to enhanced osteoclast activity at the hypertrophic cartilage/trabecular bone interface, increased bony collar formation in the growth plates of the knee joints, and increased expression levels of OPN, osteonectin (ON) and

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OC in the long bone trabeculae (Chen et al., 1999). This FGFR3 mutation corresponds to the Gly375Cys mutation in humans that is responsible for achondroplasia, the most common form of dwarfism, characterised by shortened limbs, primarily due to reduced chondrocyte activity in long bones. Unlike the other FGFRs, FGFR4 has yet to be linked to a skeletal abnormality and thus, until recently, almost nothing was known about its role in bone. Preliminary studies have shown that FGFR4 is expressed at high levels in rudimentary bone and at sites of intramembranous ossification (Cool et al., 2002). 3. The heparan sulfate proteoglycans — co-receptors for FGF/FGFR interactions 3.1. Heparan sulfate proteoglycans Creating yet further complexity for FGF/FGFR signalling is the obligate requirement for heparan sulfate proteoglycans (HSPGs) (Rapraeger et al., 1991; Yayon et al., 1991; Esko and Selleck, 2002) to form a tripartite complex with FGF and its receptor (Kan et al., 1993). HSPGs are highly abundant, strongly anionic molecules, ubiquitously distributed in the extracellular matrix and across the surfaces of nearly all cells. They are composed of a protein core to which heparan sulfate (HS) glycosaminoglycan (GAG) sugar side chains of varying length and number are covalently bound (Kjellen and Lindahl, 1991; Turnbull et al., 2001). Within this complex, the sugar chains of HSPGs not only protect the ligand from normal proteolytic degradation (Gospodarowicz and Cheng, 1986), but also enhance and stabilise ligand/receptor interactions on the cell surface (Esko and Selleck, 2002). The recent discovery that mutations in HS biosynthesis result in significant developmental defects in mice as well as in Drosophila has emphasised the importance of HSPGs during stages of normal tissue patterning and growth (Paine-Saunders et al., 2000). In contrast, mutations within the HSPG core proteins are relatively benign developmentally, suggesting that the great bulk of any HSPG's biological activity can be ascribed to its sugar complement.

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3.2. Sub-classes of HSPGs In addition to the many specialised forms of HSPGs, such as agrin, there are three broad categories of tissue HSPG, based primarily on their core protein composition: the perlecans, glypicans and syndecans (Fig. 3A). The large perlecans, approximately 460 kDa, are secreted by cells into the extracellular environment and are the major PG of basement membranes. The protein core consists of 5 protein domains, the first of which contains Ser-Gly attachment sites for 3 HS chains. Perlecan side chains have been shown to be involved in FGF metabolism (Aviezer et al., 1994). Perlecan null mice (Hspg2−/−) die either at about embryonic day 10.5 or shortly after birth, displaying defective cephalic development and skeletal abnormalities in the long bones, thorax and craniofacial regions (Hassell et al., 2002); emphasising the importance of perlecans for successful osteogenesis. Glypicans have also been revealed as important for normal bone formation. Glypicans comprise a family of 6 structurallyrelated members, ranging from 57 to 84 kDa, that are attached to the cell surface through a glycosyl-phosphatidylinositol (GPI) anchor. The extracellular region contains multiple GAG attachment sites, and can carry from 2 to 6 HS chains, some of which can interact with FGFs (Galli et al., 2003). Glypican 1 is highly expressed in the skeletal system (Litwack et al., 1998) and mutations in the glypican 3 gene result in Simpson– Golabi–Behmel Syndrome, a disorder characterised by pre- and post-natal overgrowth, cleft palate, congenital heart and renal defects, vertebral and rib defects and postaxial polydactyly (Hughes-Benzie et al., 1992; Paine-Saunders et al., 2000). The expression patterns of syndecans have been studied in bone in relation to their localisation with FGFRs. There are four transmembrane syndecan proteins varying between 22 and 43 kDa, with several GAG attachment sites in their ectodomain. Although HS is the major GAG chain attached to syndecans, syndecans 1 and − 4 can be also modified by chondroitin sulfate (CS) GAG chains (Rapraeger et al., 1985; Shworak et al., 1994). Syndecans have a short, highly conserved cytoplasmic domain

Fig. 3. HSPG classifications and HS structure. A) Heparan sulfate proteoglycans can be broadly classified as glypicans, perlecans and syndecans, depending on their distribution within and around the cell. Glypicans are attached to the cell membrane by GPI anchors, whereas syndecans are transmembrane proteins. Perlecans are actively secreted into the pericellular matrix and are predominantly found in the basement membrane. B) Heparan sulfate is composed of repeating disaccharide units of glucuronic acid and glucosamine of varying length that can be modified at irregular intervals along the chain. These modifications cluster the sulfation patterns on the HS chain into discrete protein-binding domains and ultimately lead to different growth factor-binding specificities of different HS species.

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and may be responsible for osteoblast cell–cell adhesion via their interaction with CAMs and cadherins interacting with focal contacts and the cytoskeleton, and in mediating bone cell– matrix interactions. In particular, syndecan 1 may be an integral part of a “beneficial” regulatory loop that inhibits bone loss, as syndecan 1 can not only decrease osteoclast formation, but can also increase osteoblast development, so decreasing the incidence of lytic bone disease in SCID mice (Dhodapkar et al., 1998). Syndecan 4 interacts with dynamin II to mediate focal adhesion and stress-fiber formation, whereas syndecan 2 is not thought to be involved in focal adhesion complexes (Woods and Couchman, 1994; Yoo et al., 2005). Despite the differences in activity of syndecans 1 and − 4, the HS chains isolated from these core proteins in murine mammary gland cells are almost indistinguishable from each other (Zako et al., 2003). During chondrogenic development, chondrocytes in the femoral condyles have been shown to express syndecan 4 coincidently with FGFR3 expression, with low levels of syndecans 1 and − 2 (Molteni et al., 1999). In contrast, during osteoblast differentiation, the expression patterns of syndecans 2 and − 4 coincide with those of FGFRs1–3. These spatiotemporal expression patterns further suggest an important, modulating role for syndecans on FGFRs during bone development. 3.3. The interaction between HS and the FGF–FGFR complex The interaction of HS with FGF and its receptor is complicated, and the precise conformation and stoichiometry of molecules interacting within the complex are still being elucidated. Three potential models that have been proposed to describe the interaction of FGF/FGFR/HS. It has been suggested that a single HS molecule binds 2 FGFs which, in turn, binds 2 FGFRs to induce signal transduction (DiGabriele et al., 1998). It has also been posited that a trimeric complex of 1 HS, 1 FGF and 1 FGFR must form first, before dimerising with a second trimeric complex to induce signalling (Schlessinger et al., 2000). A third hypothesis suggests that the binding of 1 HS with 1 FGF and 1 FGFR causes the dimerisation of a second FGFR for signal transduction (Springer et al., 1994). HS crystallisation studies have provided a variety of possible orientations of the elements within the signalling complex, and each model has supporting evidence. One major complication has been the use of Heparin, rather than tissue-specific HS, in the complexes that have been studied; the relative promiscuity of the mast cell-derived Heparin provides less constraint than less charge-dense forms of the HS sugar. 3.4. HS chains mediate different cell phenotypes Despite a lack of understanding of the molecular detail of exactly how HS interacts with ligands and receptors, its requirement for various cellular processes, including the mediation of FGF signalling, has been well established. Cells that lack HS are hampered in their interactions with many growth factors (Schlessinger, 2004). Independent of their core proteins, HS sugars have been implicated in cell–cell adhesion, cell–matrix adhesion, migration, proliferation, blood coagula-

tion, inflammation, tissue regeneration, tumour progression, lipoprotein metabolism and viral entry into cells (Kolset and Salmivirta, 1999; Tumova et al., 2000; Shukla and Spear, 2001). HS chains are linear, highly sulfated GAG co-polymers comprised of repeating disaccharide units of glucuronic acid (GlcA) and glucosamine (GlcN), with variable N- and O-sulfated groups attached along their length (Fig. 3B). The component disaccharides can be sulfated by a number of sulfotransferases, most commonly by O-linking at the C2 of the GlcA, by O-linking at the C3 or C6 of the GlcN and/or N-sulfation at the C2 of the GlcN residue; this latter substitution is essential for charged domain formation (Gallagher, 1997), and is often accompanied by epimerisation of the carboxyl group of GlcA to yield iduronic acid (IdoA). HS chain sulfation tends to be irregular, existing in clustered, short, functional domains separated by longer, unsulfated (acetylated) regions (Gallagher, 2001). Each tissue therefore has a unique, characteristic HS profile that may also vary at different stages of tissue development (Brickman et al., 1998; Lindahl et al., 1998; Allen et al., 2001). These variations in sulfation result in altered HS/FGF specificities within different cell types (Nurcombe et al., 1993; Rosenberg et al., 1997), which subtly alter the receptor–ligand complex and the downstream signals that lead to changes in cell phenotype. The current preponderance of evidence suggests that HS is expressed in tissue- and developmental stage-specific forms (Perrimon and Bernfield, 2000; Gallagher, 2001) that are required for embryonic organ formation (Badylak, 2002). The specific interaction of HS with a number of proteins demonstrates its importance in many biological processes. It has been shown that cells from both the osteoblast and osteoclast lineages synthesise HSPGs, and that HSPGs are localised to both the membrane and matrix of bone tissue. (Nakamura and Ozawa, 1994). Different HS species have been shown to bind to a number of different growth factors, collectively termed heparin-binding growth factors (HBGFs), including the FGFs, the VEGFs, hepatocyte growth factor (HGF) and members of the TGF-β superfamily, including the sub-family of BMPs. The terms HBGF I and II are historically synonymous with FGF-1 and FGF-2 and were often used interchangeably. The vital importance of HS for mediating individual cell responses to susceptible growth factors has been demonstrated in experiments where disruption of HS with the specific enzyme heparinase inhibits the FGF-2-mediated proliferation of smooth muscle cells in vitro (Yayon et al., 1991) as well as their proliferation in injured carotid arteries (Kinsella et al., 2004). Previous studies from our laboratory have shown that HS isolated from neuronal tissue at different embryonic stages can induce altered phenotypes in pre-confluent breast cancer cells when added in combination with different FGF ligands. HS species isolated from mice at embryonic day 11 (HS1) and day 9 (HS2) (Brickman et al., 1998) were examined in combination with FGF-1 and FGF-2 for their ability to influence breast cancer cell phenotype (Nurcombe et al., 2000). It was shown that FGF-1 in combination with its potentiating HS, HS1, could induce migration of MCF-7 and MDA-MB-231 breast cancer cells, which did not occur when HS1 was delivered in combination with FGF-2. Similarly, FGF-2 in combination with HS2 could

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induce proliferation in the same breast cancer cell lines in a dosedependent manner, an effect that could not be replicated with FGF-1. It therefore appears that FGF signalling can be carefully and deliberately regulated by the application of appropriate HS chains. More recently, we have shown that HS derived from adult neural tissue can be used to induce a neuronal phenotype in primary osteoblast cells (Chipperfield et al., 2002), demonstrating that the regulatory impact that different HS species impart on different tissue types depends less on tissue specificity than ligand-receptor specificity. We have shown that HS isolated from the conditioned media, cell surface and matrix compartments of differentiating osteoblasts show different sulfation patterns along their chains. HS from the cell surface compartment showed a higher degree of N-sulfation than those HS species isolated from the media and matrix compartments (our unpublished data). Furthermore, these HS species could increase the expression of Runx2 and ALP in pre-confluent MC3T3-E1 cells, and when combined with FGF-2, HS isolated from the matrix compartment could increase OPN mRNA expression levels above those seen with FGF-2 alone (our unpublished data). Runx2 has been shown to be important in the progression of osteoblast differentiation (Ducy et al., 1997, 1999; Komori et al., 1997), and has been found to play a role in the switch between proliferative and differentiative cell phenotypes that is essential for successful bone formation. Our findings suggest that HS may be able to induce a shift between proliferation and differentiation by increasing Runx2, which supports previous findings of the importance of stage-specific HS in controlling cell phenotype. The ability of HS to augment growth factor activity, such as FGF-2, through both specific signalling pathways and in stimulating osteoblast-specific transcription factors, has great ramifications for bone tissue manipulation, as it creates a secondary level of control of extracellular matrix protein and, therefore, osteogenesis. 4. Fracture therapies using growth factors and their co-factors molecules The successful treatment of fractures and bone defects that result from trauma or surgical resection is a major concern in orthopaedics. As immunological rejection limits the use of allogenic or xenogenic bone grafting, there has been a significant shift within the field of orthopaedics towards bone tissue engineering and its potentially more sophisticated techniques for promoting osteogenesis. One particular avenue being explored is the harnessing of the unique properties of growth factors and their co-factor HS molecules. Fracture healing proceeds through a cascade of cellular events in which MSCs respond to regulatory changes in the environment by proliferating, differentiating, and synthesising extracellular matrix. Whilst it is still unclear from exactly where these cells originate, evidence suggests that the periosteum, endosteum and cells from within the bone marrow are the major contributors, as well as cells from the surrounding muscle and soft tissues (Caplan, 1991). Fracture healing is dependent upon the initial stages of hematoma formation, inflammation and the

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formation of granulation tissue, and studies often achieve successful increases in bone formation from a single application of growth factors acting at these early stages (Nakamura et al., 1998a,b; Einhorn et al., 2003). Furthermore, successful bone tissue engineering has also been achieved through the delivery of bioactive growth factors in bioresorbable scaffolding matrices (Lisignoli et al., 2002; Srouji et al., 2004). These methods are particularly attractive as they boost local osteoprogenitor cell recruitment, proliferation and differentiation. 4.1. Fracture repair using growth factors Many of the growth factors currently being investigated for fracture healing are HBGFs that are produced by the osteoblasts and stored within the matrix of bone. Of these, FGF-1 and FGF2 have been extensively investigated as candidates for fracture healing. FGF-1 has been shown to aid in the bridging of a parietal bone critical-sized defect (Cuevas et al., 1997) and to increase the bone-implant interface when used in combination with titanium-based scaffolds (McCracken et al., 2001). FGF-2 has been more extensively studied for its use in fracture healing, and has been shown to increase callus size, bone formation, and mechanical stability in rats, rabbits and baboons (Nakamura et al., 1995; Radomsky et al., 1999; Pun et al., 2000; Nakajima et al., 2001; Iwaniec et al., 2002). Continuous supplementation of FGF-2 for up to 2 weeks can enhance bone formation in rats with as little as 3 ng of FGF-2 or as much as 4 mg per dose, depending on the mode and frequency of delivery (Nakamura et al., 1995; Pun et al., 2000; Iwaniec et al., 2002). In contrast, a single application of FGF-2 can stimulate callus formation at 4 weeks and increase bone mineral content by 8 weeks (Nakamura et al., 1998b), indicating that sustained delivery of FGF-2 is not obligatory. Furthermore, FGF-2 increases bone formation in a dose-dependent manner (Inui et al., 1998; Kimoto et al., 1998). Kimoto and others surgically created a subperiosteal pouch and filled a dome-shaped filter with demineralised bone matrix mixed with FGF-2 at concentrations of 0, 1, 10, or 100 ng in a collagen mini-pellet. They showed that after 4 weeks, there was a considerable increase in bone formation with 1 ng of FGF-2, whereas bone formation was inhibited in the 100 ng group. As well as the FGFs, members of the TGF-β superfamily have been shown to be successful in enhancing osteogenesis. Both TGF-β1 and -β2 are known to increase differentiation of committed osteoprogenitor cells, and are potent stimulators of bone repair in calvarial and long bone defects when administered alone (Joyce et al., 1990) or in combination with either IGF-I or IGF-I/growth hormone (GH) (Schmidmaier et al., 2002). In addition, several growth factors can directly regulate the availability of other growth factors; for example, FGF-2 can regulate the production of IGF-I in vitro and in vivo as well as the expression of TGF-β. FGFs have also been shown to regulate the expression of VEGF and HGF, factors that are also mitogenic for osteoprogenitor cells (Marie, 2003). Interestingly, one study showed that the systemic delivery of GH combined with the local release of IGF-I and TGF-β from coated intramedullary rods could not synergistically enhance

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bone formation compared to the separate delivery of each growth factor (Schmidmaier et al., 2002). However, a more recent investigation demonstrated that the combined local delivery of IGF-I in combination with TGF-β1 in a hydrogel scaffold could significantly enhance bone formation in a rat tibial segmental defect over IGF-I alone (Srouji et al., 2004), suggesting that perhaps the mode of delivery also affects the potential for growth factor-mediated bone healing. These findings also demonstrate that a universally applicable combination of growth factors has yet to be established for fracture therapy. Of the growth factors currently being tested, BMPs are successful in augmenting bone formation in critical-sized defects (Geesink et al., 1999; Friedlaender et al., 2001; Govender et al., 2002). BMPs are structurally-related to the TGF-β superfamily and have the capacity to regulate the commitment and differentiation of osteoprogenitor cells (Wozney and Rosen, 1998). Originally identified for their ability to form ectopic bone, BMP-2 and -7 (also known as osteogenic protein-1, OP-1) have been widely used to increase osteogenesis in bone defect sites, and have been successful in helping unite critical-sized defects in the long bones of rats, rabbits, dogs, sheep, primates and even in humans (Boden et al., 2002; Govender et al., 2002; Murakami et al., 2002; Bragdon et al., 2003; Schmoekel et al., 2004; Sumner et al., 2004). Several randomized, controlled studies have been conducted in patients with tibial and fibular defects or non-unions. The results showed recombinant human (rh) BMP-2 and BMP-7 not only increase fracture healing to levels achievable with bone grafting, but also reduce the risk of failure, reduce the requirement for surgical invasion following the surgery and eradicate the problems previously associated with donor site morbidity (Geesink et al., 1999; Friedlaender et al., 2001; Govender et al., 2002). Furthermore, rhBMP-2 on absorbable collagen sponge has been approved by the Food and Drug Administration (FDA) for use in open long bone fractures and as a replacement for autogenous bone grafting when used inside titanium-tapered anterior spinal fusion cages [for review see (Boden, 2005)]. Although the clinical use of growth factors is increasing, their efficacy in orthopaedics is complicated by virtue of their inherent instability. Growth factors are rapidly cleared in vivo as well as being highly susceptible to proteolytic degradation (Babensee et al., 2000). In addition, large quantities of growth factors are usually required in order to achieve a successful outcome, as their exposure to sterilisation procedures in the unbound state, as well as exposure to organic solvents, alkaline pH or storage at room temperature all appear to reduce their bioactivity (Westall et al., 1983; Caccia et al., 1992; Berscht et al., 1994). Furthermore, all of the aforementioned growth factors are proto-oncogenic, which is increasingly becoming an important translational problem (Hunter and Avalos, 2000). Thus, the long-term considerations about the safety of growth factors as a therapy are still being debated. Taken together, these findings suggest that an alternative, bioactive agent that can protect, localise and/or enhance the effects of exogenously applied growth factors in vivo would be a great advantage in orthopaedics as it would reduce the dosage of exogenous

growth factors required. Furthermore, an agent that can be used independently to augment in vivo bone formation by harnessing the potential of endogenously-produced growth factors would be even more therapeutically desirable. 4.2. Accelerated fracture repair with HS and HS-like molecules The binding and protective capabilities of HS sugars for a wide range of growth factors involved in mediating osteogenesis, particularly the FGFs, TGF-βs and BMPs, suggest that HS may provide a significant adjuvant within fracture repair strategies. Sugar-based therapies are not a new approach. HSlike molecules have been tested over the last decade for their ability to augment bone formation, as well as in the healing of other tissue types. Dextran derivatives substituted with carboxymethyl benzylamide sulfonate (CMDBS) have been shown to mimic heparin/HS by providing a somewhat similar protection and stabilisation capability for growth factors as imparted by HS, albeit with less affinity. Otherwise known as ‘regenerating agents’ (RGTAs), these HS-like molecules have been shown to stimulate tissue repair in skin, bone, muscle and the cornea (Blanquaert et al., 1995; Meddahi et al., 1996; Logeart-Avramoglou and Jozefonvicz, 1999; Papy-Garcia et al., 2002), which is presumed to occur through a process of enhanced growth factor activity within the wound site. CMDBSs have been shown to induce complete bony bridging of a rat critical-sized skull defect, with the formation of only fibrous tissue in the controls (Blanquaert et al., 1995). Similarly, RGTA1507, another form of CMDBS, was shown to enhance collagen accumulation and epithelial cell proliferation when administered to hamsters that had been induced with periodontitis (Escartin et al., 2003). RGTA1507 has a particular affinity for FGF-1 and TGF-β, and Escartin and co-workers have suggested that these increases in tissue formation may be brought about by the specific interaction of RGTA1507 with these particular growth factors (Escartin et al., 2003). In addition, RGTA11 delivered in collagen plasters enhanced the proportion of ALP-positive osteoprogenitors as well as the formation of bony nodules adjacent to craniotomy defects not seen in the control (Lafont et al., 2004). Blanquaert et al. have suggested that the effects of these HS-like molecules may be, in part, due to the protecting and potentiating effects of HS for endogenous growth factors (Blanquaert et al., 1995). Enhancing and prolonging growth factor expression within a callus site might, in turn, promote the expression of a number of genes important in the control of osteogenesis. Previous tissue culture studies have shown that E9, another form of derivatized dextran, can increase osteoblast proliferation in a dose-responsive manner, inhibiting proliferation at high doses, and can also increase the expression of ALP, collagen I, OC and ON (Berrada et al., 1994). Therefore, it is assumed that HS-like molecules exert their effect by increasing both cell proliferation and differentiation within the wound site, presumably by enhancing growth factor-mediated signals. Although HS-like molecules have the potential to accelerate fracture repair, HS molecules themselves are only now being investigated as a potential treatment therapy in bone. We

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Fig. 4. Schematic representation of the potential interactions of HS in fracture repair. HS may increase fracture repair by sequestering a number of different growth factor (GF) ligands to the cell surface and increase their receptor (Rc) binding. HS has been shown to bind to various growth factors (represented as GF1, GF2 etc) which can either soak up the local growth factors or enhance the local concentration of growth factor on the cell surface and facilitate enhanced ligand/receptor interactions. HS harvested from osteoblast cells increases the expression of a number of genes involved in osteoblast differentiation as well as the expression of a number of growth factors present within the callus. This increase in growth factor expression levels further enhances osteoblast signalling and bone formation through increased growth factor signalling.

recently demonstrated that HS harvested from differentiating MC3T3-E1 cells can increase trabecular bone volume by 20% from a once-off application into a mid-diaphyseal femoral fracture (Jackson et al., 2006). Furthermore, HS supplementation increased the expression of ALP and Runx2, as well as the expression of a number of HBGFs present within the callus. We hypothesise that the increases in bone formation and markers of osteoblast differentiation were triggered by the enhanced presentation of growth factors within the callus caused by the sequestering capabilities of HS. Furthermore, the ability of HS to increase the expression of other growth factors suggests that HS can not only increase presentation of already present growth factors, but activate signalling pathways to increase production of other growth factors important in mediating bone formation. This potential mechanism of HS activity is outlined in Fig. 4. Together, these findings show that HS and HS-like molecules appear to have the remarkable potential to create an environment that enhances growth factor signalling, thereby increasing the expression of markers important in the progression of osteogenesis. 5. Heparin versus heparan sulfate A common misconception surrounding the use of HS and HS-like molecules in bone therapies is drawn from the discovery that long-term heparin therapy induces osteoporosis (Barbour et al., 1994). The anti-coagulant heparin is a hypersulfated form of HS, isolated from mast cell-rich tissue such as lung and gut, and is used to inhibit blood clotting as well as in the treatment of a broad spectrum of thrombotic blood vessel,

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heart, and lung conditions. Consequently, heparin is properly regarded as one of the most successful biomedical therapies ever developed. However, in addition to its benefits, one-third of patients receiving continuous heparin treatment also shows a decrease in bone density which does not appear to be rapidly reversible (Shaughnessy et al., 1999). Studies with rabbits indicate that injections of 1000 IU/kg of heparin per day can lead to improper lamellar bone structure and a large, uncalcified bone matrix (Turan et al., 2003). HS-based therapies will not necessarily have similar effects to heparin, predominantly due to structural differences between these molecules which can alter growth factor and growth factor receptor interactions. Clinical grade, low molecular weight heparin is approximately 10–12 kD, with an extraordinarily high charge density, and thus cross-binding activity, whereas HS from other tissues can be as large as 70 kD, with much more variation in chain sulfation. Although other HS forms may also have anti-coagulant activity, particularly those derived from endothelial cells, these do not have the same efficacy as heparin. Heparin has more than 85% of the GlcN residues in the N-deacetylated and N-sulfated forms, whilst the GlcN residues of HS are only 40–60% Nconverted. In addition, approximately 70% of the GlcA in heparin is in the IdoA form, whereas HS has only 30–50% IdoA content. These structural differences appear to underlie the reason why heparin has such long-term disadvantages for bone formation, whereas other forms of HS may, in fact, aid in promoting osteogenesis. 6. Conclusions The FGFs are an important regulator of bone formation, having potent effects at all stages of the osteogenic lineage. HS, as a specific co-factor and catalyst of the FGFs, may represent an important therapeutic adjuvant for fracture repair. A greater understanding of the interaction between FGF, its receptor and HS at various developmental stages, and the preferences the FGFs display for different HS structures, may provide important clues as to the most advantageous combination for the purpose of specifically enhancing osteogenesis. Acknowledgements This paper was funded by the Agency for Science Technology and Research (A-STAR) at the Institute of Molecular and Cell Biology (IMCB), Singapore. References Allen, B.L., Filla, M.S., Rapraeger, A.C., 2001. Role of heparan sulfate as a tissue-specific regulator of FGF-4 and FGF receptor recognition. J. Cell Biol. 155, 845–858. Aviezer, D., Hecht, D., Safran, M., Eisinger, M., David, G., Yayon, A., 1994. Perlecan, basal lamina proteoglycan, promotes basic fibroblast growth factor-receptor binding, mitogenesis, and angiogenesis. Cell 79, 1005–1013. Babensee, J.E., McIntire, L.V., Mikos, A.G., 2000. Growth factor delivery for tissue engineering. Pharm. Res. 17, 497–504. Badylak, S.F., 2002. The extracellular matrix as a scaffold for tissue reconstruction. Semin. Cell. Dev. Biol. 13, 377–383.

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