Advertisement

Periostin pp 49-61 | Cite as

Periostin in Bone Regeneration

  • Oriane Duchamp de Lageneste
  • Céline ColnotEmail author
Chapter
Part of the Advances in Experimental Medicine and Biology book series (AEMB, volume 1132)

Abstract

Bone regeneration is an efficient regenerative process depending on the recruitment and activation of skeletal stem cells that allow cartilage and bone formation leading to fracture consolidation. Periosteum, the tissue located at the outer surface of bone is now recognized as an essential player in the bone repair process and contains skeletal stem cells with high regenerative potential. The matrix composition of the periosteum defines its roles in bone growth, in cortical bone modeling and remodeling in response to mechanical strain, and in bone repair. Periostin is a key extracellular matrix component of the periosteum involved in periosteum functions. In this chapter, we summarize the current knowledge on the bone regeneration process, the role of the periosteum and skeletal stem cells, and Periostin functions in this context. The matricellular protein Periostin has several roles through all stages of bone repair: in the early days of repair during the initial activation of stem cells within periosteum, in the active phase of cartilage and bone deposition in the facture callus, and in the final phase of bone bridging and reconstitution of the stem cell pool within periosteum.

Keywords

Periosteum Periostin Bone repair Stem cell 

6.1 Introduction

Following a fracture, bone regeneration is initiated. Bone tissue has a great capacity to regenerate, creating new bone which is indistinguishable from the uninjured bone, without leaving a scar. Although the steps of bone regeneration largely recapitulate those observed during bone development, bone regeneration is also regulated by other factors such as the inflammatory response and the mechanical environment that can influence the repair process [1, 2, 3]. The origins of skeletal stem cells and the factors regulating their functions in bone repair have been investigated using mouse models. While systemic recruitment of cells is minimal, bone marrow stromal cells/skeletal stem cells (BMSCs) have a local osteogenic potential and mostly have trophic and immunomodulatory roles within the bone marrow compartment. Periosteum is another local source of skeletal stem cells for bone repair [4, 5, 6].

The osteogenic potential of the periosteum has been highlighted since 1742 when Duhamel placed a periosteum on a silver ring around a bone and observed new bone formation coming from the periosteum [7]. One century later, Dupuytren suggested that periosteum and bone marrow played a role in cartilage formation in the fracture callus [8]. It is now well established that the periosteum plays many roles during bone development, growth, repair and aging, and its functions may be affected in some pathologies. The ECM composition of the periosteum is likely to influence its functions. The matricellular protein Periostin is expressed within periosteum and periodontal ligament and was first identified as Osteoblast-specific factor 2 (OSF-2) in a mouse osteoblastic calvarial cell line (MC3T3-E1) and in primary osteoblasts [9, 10, 11]. In bone, Periostin acts as a structural component of the matrix regulating collagen cross-linking and as a signaling molecule via interaction with integrin receptors [12]. Functional analyses of periosteal cells and Periostin using Periostin KO mice have now revealed several roles of Periostin in bone regeneration.

6.2 Bone Regeneration after Fracture

Bone regeneration generally occurs through a combination of endochondral and intramembranous ossification. The regeneration process can be summarized into four critical stages: the inflammation phase (hematoma formation and skeletal stem cell recruitment), the soft callus phase (neo-angiogenesis and cartilage/bone formation), the hard callus phase (cartilage resorption and/or active bone deposition) and the remodeling phase [13, 14]. Following a fracture, cells coming from the local bone marrow and the vasculature form the hematoma and the inflammatory process is initiated [15]. Pro-inflammatory mediators are secreted and initiate the repair cascade [16]. Neutrophils are first mobilized at the fracture site and secrete interleukins and chemokines like IL-6 (interleukin-6) and CCL2 (chemokine ligand 2) [15, 17]. Interleukins-1β (IL-1β) [18], IL-6 [19, 20], IL-11 [15], IL-17 [21], IL-18 [15], IL-23 [18] as well as Tumor Necrosis Factor-α (TNF-α) [22] promote angiogenesis and attract inflammatory cells like macrophages that remove necrotic tissues [16]. The inflammatory phase of healing is generally completed after 7 days following a fracture in mice [23]. In cases of bacterial infection, the inflammation phase persists and the healing process fails [24]. The inflammatory cells, macrophages and platelets secrete growth factors like Platelet-Derived Growth Factor (PDGF), Vascular Endothelial Growth Factor (VEGF) and Bone Morphogenetic Proteins (BMPs) to initiate the recruitment of skeletal stem/progenitor cells at the fracture site that are activated around day 3 [13, 16, 25]. During the soft callus phase, skeletal stem cells differentiate into chondrocytes or directly into osteoblasts depending on the mechanical tensions. In the center of the callus, where the mechanical tensions are high, bone regeneration occurs via endochondral ossification while at the periphery, bone regeneration mainly occurs through intramembranous ossification. During endochondral ossification, skeletal stem cells first differentiate into chondrocytes, proliferate and secrete a cartilaginous matrix rich in Type II Collagen and Aggrecan. These cells are then replaced by osteoblasts that produce a bone matrix rich in Type I Collagen. During the hard callus phase, the cartilage matrix is mineralized. Chondrocytes undergo hypertrophy and secrete VEGF to allow vascular invasion of hypertrophic cartilage. Due to osteoclast activity, cartilage is resorbed and replaced by woven bone. While it was generally recognized that hypertrophic chondrocytes die by apoptosis allowing vascular invasion and endochondral ossification, recent studies have shown that some hypertrophic chondrocytes can transdifferentiate into osteoblasts as observed at the level of the growth plate [26, 27, 28, 29, 30]. The cascade of events during this transition between mineralized cartilage and new bone is strictly regulated and any disruption of this transition leads to delayed or impaired healing [31, 32]. During the remodeling phase of repair, primary bone in the callus is degraded by osteoclasts and replaced by lamellar bone to restore the anatomical structure of bone that supports mechanical loads [23]. In mice, the replacement of cartilage by bone is complete by day 28 and the bone remodeling process carries on until after 2 months.

6.3 Role of Periosteum in Bone Repair

The periosteum from Greek peri: surrounding and osteon: bone, is a thin membrane of connective tissue on the external surface of most bones. The periosteum is attached to the bone cortex through Sharpey’s fibers and serves as an attachment site for the muscles, tendons and ligaments [33]. The periosteum is composed of two layers, an outer fibrous layer rich in fibroblasts, Type I Collagen and Elastin fibers and an inner layer named the « cambium » layer in direct contact with the bone cortex [5, 34, 35](Fig. 6.1). The cambium layer is rich in osteoblasts, fibroblasts, osteoprogenitor cells and putative skeletal stem cells in a sparse collagenous matrix. The periosteum is highly vascularized and innervated [36]. The periosteal arteries supply blood to the adjacent bone and muscles [37]. The presence of sensory nerves endings that penetrate from the periosteum into bone canals makes the bone sensitive to pain [38].
Fig. 6.1

Composition of the periosteum. The periosteum is composed of two layers, an outer fibrous layer containing fibroblasts and microvessels, and an inner (cambium) layer highly vascularized and innervated, containing skeletal stem cells and in contact with the bone surface covered by pre-osteoblasts. (Adapted from Refs. [5, 35])

Due to its external localization on bone, the periosteum is extremely sensitive to mechanical stimuli. When mechanical tensions or increased loading are applied, new bone formation deriving from the periosteum is observed, as well as periosteal hypertrophy, DNA synthesis, and cell proliferation [39, 40, 41, 42]. During childhood and through adulthood, periosteal appositions allow bone to grow in width and cortical bone modeling/remodeling in concert with osteoclasts maintain bone structure in response to mechanical strain. In the context of fracture, it has been long known that removal/stripping of the periosteum impairs bone regeneration [43]. In models of diaphyseal fracture in rats, disruption of the periosteum impairs early chondrogenesis [43, 44]. Transplantation of Rosa26-LacZ segmental grafts into critical bone defects or periosteum grafts into fractures showed that periosteum was a major contributor to cartilage and bone in the callus compared to bone marrow and endosteum [4, 45, 46, 47].

The periosteal response to the fracture can be observed histologically as early as 24 h following injury with a cellular proliferation leading to thickening of the periosteum near the fracture site. The size and the cellular content of the callus depend on the periosteal response to the mechanical environment. If the fracture is left unstabilized, the periosteal response to mechanical strain is very efficient, leading to bone regeneration via endochondral ossification. In the presence of mechanical stability or if the fracture is reduced, the periosteal response is less efficient leading to direct ossification [2, 32, 48]. These studies highlight the high regenerative potential of the periosteum. However, the absence of specific marker(s) to follow periosteum-derived cells (PCs) during bone regeneration makes it difficult to distinguish its endogenous potential compared to bone marrow. Therefore, many studies have aimed to isolate cells from the periosteum to study their in vitro and in vivo properties .

6.4 Periosteal Cells Express Periostin in Response to Bone Injury

Bone marrow stromal cells/skeletal stem cells (BMSCs) are the most described skeletal stem cells in the literature. In vitro, BMSCs can differentiate into osteoblasts, adipocytes and chondrocytes, and form colonies at low density. In vivo, the regenerative potential of BMSCs has been tested using cell transplantation in subcutaneous sites, where they form bone ossicles and support hematopoiesis [49]. However, these ectopic sites do not recapitulate the environment of a fracture (i.e. inflammation and mechanical stimuli). Self-renewal was shown using clonal expansion in vitro followed by ectopic subcutaneous transplantation. Many studies report transplantation of BMSCs in bone defects or at injury sites, but few include lineage tracing to follow their fate. BMSCs rapidly disappear after transplantation and have poor osteogenic potential. BMSCs rather stimulate repair via their ability to secrete growth factors and inflammatory factors [50, 51]. Several lineage studies using Cre reporter mice have aimed to define markers for BMSCs and follow these cells during bone repair [52, 53, 54]. However, there is no clear evidence regarding the specificity of these markers that may also label cells in periosteum and other tissues. Indeed, periosteal cells (PCs) express markers described initially for BMSCs such as PDGFRα, Gremlin 1, Cxcl12 and Nestin. PCs and BMSCs exhibit similar differentiation characteristics in vitro, but PCs have increased clonogenicity and cell growth. After transplantation in fractures, PCs have a higher integration potential in cartilage and bone in the callus compared to BMSCs and persist within bone indicating the higher regenerative potential of PCs compared to BMSCs [55].

Microarray analyses of PCs and BMSCs have revealed an over expression of Periostin (Postn) and Periostin-linked genes in PCs in response to injury (Fig. 6.2). Some of these genes encode matricellular proteins (Postn and Thrombospondin 2) and other extracellular matrix (ECM) proteins (Col3a1, Lumican, Asporin, Decorin, Fibrillin 1 and Dermatopontin) (Fig. 6.2). Studies have also reported Postn up regulation in the fractured tibias along with other genes encoding ECM proteins such as SPARC, Biglycan and Decorin [55, 56, 57, 58]. The expression of matricellular proteins is tightly regulated throughout life. Their constitutive expression in adult tissues is limited, but is elevated during development and in response to injury or stress [59]. Postn expression can be detected in the developing embryo by E9.5 in the first bronchial arch epithelium that is later the site for odontogenesis [60, 61]. Postn and Periostin-like-factor (PLF) are expressed at E12.5 in and around mesenchymal cells that condensate to form cartilage templates in the vertebrae, by E13.5 in cartilaginous vertebrae and ribs and by E16.5 in proliferating and hypertrophic chondrocytes in the limbs [62]. In the adult, Periostin is specifically expressed is collagen-rich tissues submitted to mechanical stress like the periodontal ligament and periosteum [11].
Fig. 6.2

Periostin is a marker defining the periosteum response to bone injury. (a) Schematic representation of primary cultures of periosteal cells (PCs) and bone marrow stromal cells (BMSCs) from un-injured (day 0, d0) and injured (day 3, d3) mouse tibia. (b) Comparative molecular response of PCs and BMSCs to bone injury (d0 versus d3) via microarray analysis: (left) Venn diagram showing 203 genes defining the periosteum response to injury (PRI); (right) Five significant functions are identified via GSEA analyses in the PRI “response to external stimulus”, “extracellular space”, regulation of external stimulus”, “matrisome” and “stem cell”. (c) (left) Periostin (Postn) and 8 other genes are found in common among the five significant functions. (right) Six genes including Postn, Egln1 (Endoglin1), Col3a1 (Collagen3a1), Lum (Lumican), Aspn (Asporin) and Dcn (Decorin) belonging to the list of Postn linked genes are also found in the PRI. (d) Periostin expression (left) in the intact periosteum and (right) in the activated periosteum at day 3 post-fracture. Scale bar: 0.5 mm. SO: SafraninO staining, c: cortex, m: muscle, bm: bone marrow, po: periosteum, cl: cambium layer, fl: fibrous layer, orange arrow indicates the fracture site (Adapted from Ref. [55])

6.5 Periostin Is Required for Bone Repair

Matricellular proteins have diverse functions such as promoting cell adhesion that may induce cell migration, they interact with growth factors that facilitate cell-matrix interactions, and bridge inorganic matter and proteins of the ECM [63]. Matricellular proteins are crucial regulators of cell phenotype, and consequently tissue function. While mice knockout models of structural ECM proteins generally induce severe phenotypes that can be lethal during development, gene deletion of matricellular proteins induces mild phenotypes that are exacerbated upon injury [64]. Like other matricellular proteins, Postn gene deletion in mice does not lead to embryonic lethality [65]. Postn KO mice show no apparent phenotype at birth, but exhibit growth retardation and weight loss within 1 month. The Postn KO bone phenotype is marked by the reduction of the size of the skull, ribs and cartilaginous growth plates and less trabecular bone in the limbs suggesting a role of Postn in bone development and homeostasis [65]. The absence of Postn also leads to loss of alveolar bone in the tooth probably due to increased osteoclasts activity in the periodontium. Thus the phenotype of Postn KO mice might at least, in part, be due to feeding disabilities caused by lesions in the in periodontal ligament, and under soft diet, growth retardation was attenuated [65].

Mice lacking Postn have impaired bone healing that is characterized by decreased callus and bone volumes throughout the stages of repair, as well as decreased cartilage formation and delayed cartilage removal (Fig. 6.3) [55]. During the remodelling phase of repair, Postn KO mice show absence of consolidation with reduced bone bridging in the callus (Fig. 6.3). This is correlated with defective periosteum shown by decreased contribution to bone repair of GFP labelled Postn KO periosteum grafts transplanted at the fracture site of wild type hosts compared to wild type grafts (Fig. 6.4) [55]. The absence of Postn that is normally highly expressed in the cambial layer of the activated periosteum compared to un-injured periosteum thus impairs periosteal activation and callus formation (Figs. 6.2 and 6.4). Downstream pathways regulated by Postn and mediating skeletal stem cell (SSC) activation still remain to be analysed. Cells maintain their normal functions in tissues by adhering to ECM proteins through integrin receptors. Postn may regulate SSC function in periosteum via its ability to bind to integrin receptors at the cell surface that has been previously described to enhance cell proliferation, survival and migration [66, 67, 68, 69].
Fig. 6.3

Periostin is required for bone repair. (a) Histomorphometric analyses showing callus, cartilage and bone volumes at days 7 through 28 post-fracture in wild type and Periostin KO mice. (b) Representative sections of fracture calluses stained with Picrosirius (PS) red showing callus bridging in wild type callus at day 28 but decreased bone bridging in Periostin KO mice and fibrosis deposition. Scale bar: 1 mm. Statistical differences between the groups were determined using Mann-Whitney test (*p ≤ 0.05, **p < 0.001, ***p < 0.0005) (n = 3 to 5). All data represent mean ± SD. (Adapted from Ref. [55])

Fig. 6.4

Periostin regulates the skeletal stem cell niche within periosteum. (Top) Experimental design for the transplantation of periosteum grafts from wild type GFP donor or Postn KO-GFP donor mice at the fracture site of wild type hosts. (Left panels) Safranin-O (SO) and DAPI/GFP staining on longitudinal callus sections at day 14 showing contribution of wild type GFP grafts to cartilage and bone. At day 28 post-fracture, rare periosteum-derived GFP+ cells integrate in the new bone (nb) to form osteocytes (white arrow) and in the new periosteum (po, white arrowheads). After a second fracture, abundant periosteum-derived GFP+ cells are found in the callus and form cartilage (white arrowheads) and bone (white arrows) at day 7 and few GFP+ cells reintegrate the new periosteum at day 28 (white arrow heads). After a third fracture, periosteum-derived GFP+ cells can again form cartilage efficiently in the callus by day 7 indicating SSC self-renewal in periosteum. (Right panels) Few GFP+ Postn KO grafts-derived cells are detected in the cartilage at day 14 but no GFP+ Postn KO cells can be detected in the new periosteum at day 28. After a second injury, no GFP+ Postn KO chondrocytes contribute to the callus by day 7 indicating absence of SSC self-renewal. These Postn KO-GFP grafts induce fibrosis at the fracture site of wild type hosts. SO Scale bars = 1 mm. po: periosteum, nb: new bone, bm: bone marrow (Adapted from Ref. [55])

During the soft and hard callus phase of bone repair, Postn is also expressed in hypertrophic cartilage at the junction between cartilage and bone within the callus and in osteoblasts within new bone trabeculae [55]. Postn inactivation may thus impact bone formation directly during bone repair as Postn has been shown to regulate osteoblast proliferation, adhesion and differentiation [11, 60]. Although we did not detect expression of Postn in TRAP-positive osteoclasts in the fracture callus (data not shown), Postn deposition in ECM may also indirectly influence osteoclast function [66]. A conditional knockout approach may therefore be useful to determine the role of Postn in various cell populations during bone repair. At later stages of repair, fibrosis is observed in the callus of Postn KO mice (Fig. 6.3) [55]. Postn has been shown to promote fibrosis in allergic and respiratory diseases [70, 71]. The cellular origin of this fibrosis and the specific role of Postn in fibrosis deposition and remodelling in the fracture callus remain to be established.

6.6 Periostin Regulates the Skeletal Stem Cell Niche Within Periosteum

In addition to regulating periosteal cell activation in the inflammatory phase of repair, and periosteal cell differentiation during the soft callus phase of repair, Postn also regulates the niche of periosteal cells during the remodelling phase of repair. This was first suggested by the expression of Postn in the inner layer of the newly formed periosteum [55]. A niche for stem cells regulates their quiescence, self-renewal and differentiation potential in normal tissues and in response to injury. To demonstrate the presence of stem cells with self-renewing potential within periosteum, GFP labelled periosteal grafts were transplanted at the fracture site and graft contribution to repair was evaluated over 3 consecutive cycles of injuries. The results show that PCs within periosteum could be activated to participate in callus formation, reintegrate their niche in the newly formed periosteum at the end of the repair process and be reactivated after several injuries to contribute again to bone repair (Fig. 6.4, left panels) [55]. To test whether the absence of Postn had an impact on the capacity of PCs to reintegrate their niche and reconstitute a pool of SSCs, GFP labelled periosteum grafts from Postn KO mice were transplanted in wild type hosts. Although the fracture callus was mostly wild type host-derived, bone regeneration was impaired around the Postn KO graft with no cartilage formation and local induction of fibrosis. Moreover, Postn KO PCs could not reintegrate their niche in the new periosteum at later days and did not contribute to cartilage and callus formation after a second injury (Fig. 6.4, right panels). PCs isolated from Postn KO mice show less clonogenicity as well as decreased osteogenic and adipogenic potential in vitro. Together these data indicate that Postn is required in the periosteal niche to reconstitute the pool of PCs at the end of the repair process. Interestingly, transplantation of Postn KO graft in Postn KO hosts caused an aggravation of the repair phenotype after a second injury, further supporting the role of Postn in re-establishing the stem cell pool in periosteum [55].

The nature and role of ECM proteins and cells that constitute the SSC niche within periosteum are unknown. The periosteum is composed of many cell types (fibroblasts, pre-osteoblasts, pericytes, skeletal stem cells and osteoprogenitor cells) entrapped in the ECM. Changes in other ECM molecules are found in Postn KO PCs, suggesting complementary roles of these ECM factors in regulating SSC activation and periosteum function during bone repair [55]. Further investigation on Postn and identification of other ECM molecules that regulate PCs activation in response to injury and long-term maintenance in the periosteum will be crucial to better understand periosteum and PC functions in bone homeostasis and repair (Fig. 6.5).
Fig. 6.5

Schematic representation of Periostin functions during bone repair [1]. Periostin is up-regulated in the activated periosteum in response to injury, [2] is required for callus, cartilage and bone formation, and [3] allows self-renewal of SSCs within the periosteum. (Adapted from Ref. [55])

6.7 Clinical Applications Using Periosteal Cells and Periostin in Bone Repair

The increased regenerative capacities of PCs compared to BMSCs make the periosteum a good target to stimulate repair or an attractive source of cells for therapy. For clinical applications in orthopaedics, several concerns are raised regarding the use of SSCs such as their site and method of isolation, ex vivo expansion, mechanism of activation as well as their osteogenic potential and long term integration after transplantation. For cell-based approaches aiming to treat complex fractures or critical size defects, a two-step surgery to isolate PCs would be necessary. The resection methods as well as culture conditions could influence stem cell number and regenerative potential. Moreover, types of bones from which periosteum could be isolated and inter-individual differences (age, sex) could also impact PCs potential [72, 73]. Cell-based approaches using PCs need to be explored or these cells could be reactivated in their native environment (periosteal niche) using pharmacological drugs that would act on signalling pathways to activate these cells or to modulate the ECM niche.

The enhanced regenerative potential of PCs marked by an augmented molecular response to injury and Periostin up regulation is essential for adequate bone repair, PCs self-renewal and the reconstitution of the periosteal niche [55]. Periostin could be used therapeutically to stimulate cell migration and contribution to repair in cell-based approaches. In a model of critical size defect in rabbit, Zhang and collaborators (2017) transplanted osteogenic-induced modified BMSCs and showed increased bone formation, bone mineral density and bone volume concomitant with increased Periostin/β-catenin protein expression. In vitro, Periostin increased ALP and alizarin red S production by BMSCs with increased levels of associated proteins (p-LRP-6, p-GSK3, β-catenin and Runx2) [74]. This suggests that Periostin may be delivered to directly activate Wnt/β-catenin signaling to stimulate bone formation in the callus [75, 76]. In another study, recombinant Periostin stimulated human adipose tissue-derived stromal cells (hASC) proliferation and migration, and augmented hASC-mediated repair in a critical size calvarial defect model [68]. The multiple roles of Periostin in regulating matrix composition and cellular functions during bone repair may be mediated by various isoforms, therefore the roles of these different isoforms need to be defined to target Periostin or select appropriate Periostin fragments for tissue engineering approaches in bone repair.

Notes

Acknowledgements

This work is supported by Osteosynthesis and Trauma Care Foundation, ANR-13-BSV1-001, ANR-18-CE14-0033 and NIH R01AR072707.

References

  1. 1.
    Claes L, Recknagel S, Ignatius A (2012) Fracture healing under healthy and inflammatory conditions. Nat Rev Rheumatol 8:133–143CrossRefGoogle Scholar
  2. 2.
    Thompson Z, Miclau T, Hu D, Helms JA (2002) A model for intramembranous ossification during fracture healing. J Orthop Res 20:1091–1098CrossRefGoogle Scholar
  3. 3.
    Miclau T et al (2007) Effects of delayed stabilization on fracture healing. J Orthop Res 25:1552–1558CrossRefGoogle Scholar
  4. 4.
    Colnot C, Zhang X, Knothe Tate ML (2012) Current insights on the regenerative potential of the periosteum: molecular, cellular, and endogenous engineering approaches. J Orthop Res 30:1869–1878CrossRefGoogle Scholar
  5. 5.
    Ferretti C, Mattioli-Belmonte M (2014) Periosteum derived stem cells for regenerative medicine proposals: boosting current knowledge. World J Stem Cells 6:266–277CrossRefGoogle Scholar
  6. 6.
    Roberts SJ, van Gastel N, Carmeliet G, Luyten FP (2015) Uncovering the periosteum for skeletal regeneration: the stem cell that lies beneath. Bone 70:10–18CrossRefGoogle Scholar
  7. 7.
    Duhamel HL (1742) Sur le développement et la crue des os des animaux. Acad Roy des Sci Paris Mém 55:354–357Google Scholar
  8. 8.
    Dupuytren G (1847) On the injuries and diseases of bone. The Sydenham Society, LondonGoogle Scholar
  9. 9.
    Takeshita S, Kikuno R, Tezuka K, Amann E (1993) Osteoblast-specific factor 2: cloning of a putative bone adhesion protein with homology with the insect protein fasciclin I. Biochem J 294(Pt 1):271–278CrossRefGoogle Scholar
  10. 10.
    Ducy P, Zhang R, Geoffroy V, Ridall AL, Karsenty G (1997) Osf2/Cbfa1: a transcriptional activator of osteoblast differentiation. Cell 89:747–754CrossRefGoogle Scholar
  11. 11.
    Horiuchi K et al (1999) Identification and characterization of a novel protein, periostin, with restricted expression to periosteum and periodontal ligament and increased expression by transforming growth factor beta. J Bone Miner Res 14:1239–1249CrossRefGoogle Scholar
  12. 12.
    Bonnet N, Garnero P, Ferrari S (2016) Periostin action in bone. Mol Cell Endocrinol 432:75–82CrossRefGoogle Scholar
  13. 13.
    Dimitriou R, Tsiridis E, Giannoudis PV (2005) Current concepts of molecular aspects of bone healing. Injury 36:1392–1404CrossRefGoogle Scholar
  14. 14.
    Einhorn TA, Gerstenfeld LC (2015) Fracture healing: mechanisms and interventions. Nat Rev Rheumatol 11:45–54CrossRefGoogle Scholar
  15. 15.
    Mountziaris PM, Mikos AG (2008) Modulation of the inflammatory response for enhanced bone tissue regeneration. Tissue Eng Part B Rev 14:179–186CrossRefGoogle Scholar
  16. 16.
    Kon T et al (2001) Expression of osteoprotegerin, receptor activator of NF-kappaB ligand (osteoprotegerin ligand) and related proinflammatory cytokines during fracture healing. J Bone Miner Res 16:1004–1014CrossRefGoogle Scholar
  17. 17.
    Xing Z et al (2010) Multiple roles for CCR2 during fracture healing. Dis Model Mech 3:451–458CrossRefGoogle Scholar
  18. 18.
    Al-Sebaei MO et al (2014) Role of Fas and Treg cells in fracture healing as characterized in the fas-deficient (lpr) mouse model of lupus. J Bone Miner Res 29:1478–1491CrossRefGoogle Scholar
  19. 19.
    Wallace A, Cooney TE, Englund R, Lubahn JD (2011) Effects of interleukin-6 ablation on fracture healing in mice. J Orthop Res 29:1437–1442CrossRefGoogle Scholar
  20. 20.
    Yang X et al (2007) Callus mineralization and maturation are delayed during fracture healing in interleukin-6 knockout mice. Bone 41:928–936CrossRefGoogle Scholar
  21. 21.
    Nam D et al (2012) T-lymphocytes enable osteoblast maturation via IL-17F during the early phase of fracture repair. PLoS One 7:e40044CrossRefGoogle Scholar
  22. 22.
    Gerstenfeld LC et al (2003) Impaired fracture healing in the absence of TNF-alpha signaling: the role of TNF-alpha in endochondral cartilage resorption. J Bone Miner Res 18:1584–1592CrossRefGoogle Scholar
  23. 23.
    Gerstenfeld LC, Cullinane DM, Barnes GL, Graves DT, Einhorn TA (2003) Fracture healing as a post-natal developmental process: molecular, spatial, and temporal aspects of its regulation. J Cell Biochem 88:873–884CrossRefGoogle Scholar
  24. 24.
    Motsitsi NS (2008) Management of infected nonunion of long bones: the last decade (1996-2006). Injury 39:155–160CrossRefGoogle Scholar
  25. 25.
    Barnes GL, Kostenuik PJ, Gerstenfeld LC, Einhorn TA (1999) Growth factor regulation of fracture repair. J Bone Miner Res 14:1805–1815CrossRefGoogle Scholar
  26. 26.
    Bahney CS et al (2014) Stem cell-derived endochondral cartilage stimulates bone healing by tissue transformation. J Bone Miner Res 29:1269–1282CrossRefGoogle Scholar
  27. 27.
    Hu DP et al (2017) Cartilage to bone transformation during fracture healing is coordinated by the invading vasculature and induction of the core pluripotency genes. Development 144:221–234CrossRefGoogle Scholar
  28. 28.
    Park J et al (2015) Dual pathways to endochondral osteoblasts: a novel chondrocyte-derived osteoprogenitor cell identified in hypertrophic cartilage. Biol Open 4:608–621CrossRefGoogle Scholar
  29. 29.
    Yang L, Tsang KY, Tang HC, Chan D, Cheah KS (2014) Hypertrophic chondrocytes can become osteoblasts and osteocytes in endochondral bone formation. Proc Natl Acad Sci U S A 111:12097–12102CrossRefGoogle Scholar
  30. 30.
    Zhou X et al (2014) Chondrocytes transdifferentiate into osteoblasts in endochondral bone during development, postnatal growth and fracture healing in mice. PLoS Genet 10:e1004820CrossRefGoogle Scholar
  31. 31.
    Behonick DJ et al (2007) Role of matrix metalloproteinase 13 in both endochondral and intramembranous ossification during skeletal regeneration. PLoS One 2:e1150CrossRefGoogle Scholar
  32. 32.
    Colnot C, Thompson Z, Miclau T, Werb Z, Helms JA (2003) Altered fracture repair in the absence of MMP9. Development 130:4123–4133CrossRefGoogle Scholar
  33. 33.
    Benjamin M et al (2006) Where tendons and ligaments meet bone: attachment sites (‘entheses’) in relation to exercise and/or mechanical load. J Anat 208:471–490CrossRefGoogle Scholar
  34. 34.
    Allen MR, Hock JM, Burr DB (2004) Periosteum: biology, regulation, and response to osteoporosis therapies. Bone 35:1003–1012CrossRefGoogle Scholar
  35. 35.
    Chang H, Knothe Tate ML (2012) Concise review: the periosteum: tapping into a reservoir of clinically useful progenitor cells. Stem Cells Transl Med 1:480–491CrossRefGoogle Scholar
  36. 36.
    Hohmann EL, Elde RP, Rysavy JA, Einzig S, Gebhard RL (1986) Innervation of periosteum and bone by sympathetic vasoactive intestinal peptide-containing nerve fibers. Science 232:868–871CrossRefGoogle Scholar
  37. 37.
    Chanavaz M (1995) Anatomy and histophysiology of the periosteum: quantification of the periosteal blood supply to the adjacent bone with 85Sr and gamma spectrometry. J Oral Implantol 21:214–219PubMedGoogle Scholar
  38. 38.
    Garcia-Castellano JM, Diaz-Herrera P, Morcuende JA (2000) Is bone a target-tissue for the nervous system? New advances on the understanding of their interactions. Iowa Orthop J 20:49–58PubMedPubMedCentralGoogle Scholar
  39. 39.
    Kearney CJ, Lee JY, Padera RF, Hsu HP, Spector M (2011) Extracorporeal shock wave-induced proliferation of periosteal cells. J Orthop Res 29:1536–1543CrossRefGoogle Scholar
  40. 40.
    Pead MJ, Skerry TM, Lanyon LE (1988) Direct transformation from quiescence to bone formation in the adult periosteum following a single brief period of bone loading. J Bone Miner Res 3:647–656CrossRefGoogle Scholar
  41. 41.
    Raab-Cullen DM, Thiede MA, Petersen DN, Kimmel DB, Recker RR (1994) Mechanical loading stimulates rapid changes in periosteal gene expression. Calcif Tissue Int 55:473–478CrossRefGoogle Scholar
  42. 42.
    Simon TM, Van Sickle DC, Kunishima DH, Jackson DW (2003) Cambium cell stimulation from surgical release of the periosteum. J Orthop Res 21:470–480CrossRefGoogle Scholar
  43. 43.
    Utvag SE, Grundnes O, Reikeraos O (1996) Effects of periosteal stripping on healing of segmental fractures in rats. J Orthop Trauma 10:279–284CrossRefGoogle Scholar
  44. 44.
    Ozaki A, Tsunoda M, Kinoshita S, Saura R (2000) Role of fracture hematoma and periosteum during fracture healing in rats: interaction of fracture hematoma and the periosteum in the initial step of the healing process. J Orthop Sci 5:64–70CrossRefGoogle Scholar
  45. 45.
    Zhang X et al (2005) Periosteal stem cells are essential for bone revitalization and repair. J Musculoskelet Neuronal Interact 5:360–362PubMedGoogle Scholar
  46. 46.
    Zhang X et al (2005) Periosteal progenitor cell fate in segmental cortical bone graft transplantations: implications for functional tissue engineering. J Bone Miner Res 20:2124–2137CrossRefGoogle Scholar
  47. 47.
    Colnot C (2009) Skeletal cell fate decisions within periosteum and bone marrow during bone regeneration. J Bone Miner Res 24:274–282CrossRefGoogle Scholar
  48. 48.
    Lu C et al (2005) Cellular basis for age-related changes in fracture repair. J Orthop Res 23:1300–1307CrossRefGoogle Scholar
  49. 49.
    Sacchetti B et al (2007) Self-renewing osteoprogenitors in bone marrow sinusoids can organize a hematopoietic microenvironment. Cell 131:324–336CrossRefGoogle Scholar
  50. 50.
    Granero-Molto F et al (2009) Regenerative effects of transplanted mesenchymal stem cells in fracture healing. Stem Cells 27:1887–1898CrossRefGoogle Scholar
  51. 51.
    Fernandes MB et al (2014) The effect of bone allografts combined with bone marrow stromal cells on the healing of segmental bone defects in a sheep model. BMC Vet Res 10:36CrossRefGoogle Scholar
  52. 52.
    Mizoguchi T et al (2014) Osterix marks distinct waves of primitive and definitive stromal progenitors during bone marrow development. Dev Cell 29:340–349CrossRefGoogle Scholar
  53. 53.
    Worthley DL et al (2015) Gremlin 1 identifies a skeletal stem cell with bone, cartilage, and reticular stromal potential. Cell 160:269–284CrossRefGoogle Scholar
  54. 54.
    Zhou BO, Yue R, Murphy MM, Peyer JG, Morrison SJ (2014) Leptin-receptor-expressing mesenchymal stromal cells represent the main source of bone formed by adult bone marrow. Cell Stem Cell 15:154–168CrossRefGoogle Scholar
  55. 55.
    Duchamp de Lageneste O et al (2018) Periosteum contains skeletal stem cells with high bone regenerative potential controlled by Periostin. Nat Commun 9:773CrossRefGoogle Scholar
  56. 56.
    Nakazawa T et al (2004) Gene expression of periostin in the early stage of fracture healing detected by cDNA microarray analysis. J Orthop Res 22:520–525CrossRefGoogle Scholar
  57. 57.
    Hirakawa K et al (1994) Localization of the mRNA for bone matrix proteins during fracture healing as determined by in situ hybridization. J Bone Miner Res 9:1551–1557CrossRefGoogle Scholar
  58. 58.
    Kawaguchi H et al (1994) Stimulation of fracture repair by recombinant human basic fibroblast growth factor in Normal and Streptozotocin-diabetic rats. Endocrinology 135:774–781CrossRefGoogle Scholar
  59. 59.
    Bornstein P (2009) Matricellular proteins: an overview. J Cell Commun Signal 3:163–165CrossRefGoogle Scholar
  60. 60.
    Litvin J et al (2004) Expression and function of periostin-isoforms in bone. J Cell Biochem 92:1044–1061CrossRefGoogle Scholar
  61. 61.
    Kruzynska-Frejtag A et al (2004) Periostin is expressed within the developing teeth at the sites of epithelial-mesenchymal interaction. Dev Dyn 229:857–868CrossRefGoogle Scholar
  62. 62.
    Zhu S et al (2008) Immunolocalization of Periostin-like factor and Periostin during embryogenesis. J Histochem Cytochem 56:329–345CrossRefGoogle Scholar
  63. 63.
    Murphy-Ullrich JE, Sage EH (2014) Revisiting the matricellular concept. Matrix Biol 37:1–14CrossRefGoogle Scholar
  64. 64.
    Alford AI, Hankenson KD (2006) Matricellular proteins: extracellular modulators of bone development, remodeling, and regeneration. Bone 38:749–757CrossRefGoogle Scholar
  65. 65.
    Rios H et al (2005) Periostin null mice exhibit dwarfism, incisor enamel defects, and an early-onset periodontal disease-like phenotype. Mol Cell Biol 25:11131–11144CrossRefGoogle Scholar
  66. 66.
    Cobo T et al (2016) Role of Periostin in adhesion and migration of bone remodeling cells. PLoS One 11:e0147837CrossRefGoogle Scholar
  67. 67.
    Sonnenberg-Riethmacher E, Miehe M, Riethmacher D (2015) Promotion of periostin expression contributes to the migration of Schwann cells. J Cell Sci 128:3345–3355CrossRefGoogle Scholar
  68. 68.
    Heo SC et al (2011) Periostin mediates human adipose tissue-derived mesenchymal stem cell-stimulated tumor growth in a xenograft lung adenocarcinoma model. Biochim Biophys Acta 1813:2061–2070CrossRefGoogle Scholar
  69. 69.
    Matsuzawa M et al (2015) Periostin of human periodontal ligament fibroblasts promotes migration of human mesenchymal stem cell through the alphavbeta3 integrin/FAK/PI3K/Akt pathway. J Periodontal Res 50:855–863CrossRefGoogle Scholar
  70. 70.
    Naik PK et al (2012) Periostin promotes fibrosis and predicts progression in patients with idiopathic pulmonary fibrosis. Am J Physiol Lung Cell Mol Physiol 303:L1046–L1056CrossRefGoogle Scholar
  71. 71.
    Takayama G et al (2006) Periostin: a novel component of subepithelial fibrosis of bronchial asthma downstream of IL-4 and IL-13 signals. J Allergy Clin Immunol 118:98–104CrossRefGoogle Scholar
  72. 72.
    O’Driscoll SW, Saris DB, Ito Y, Fitzimmons JS (2001) The chondrogenic potential of periosteum decreases with age. J Orthop Res 19:95–103CrossRefGoogle Scholar
  73. 73.
    Ferretti C et al (2015) Human periosteal derived stem cell potential: the impact of age. Stem Cell Rev 11(3):487–500CrossRefGoogle Scholar
  74. 74.
    Zhang F et al (2017) Periostin upregulates Wnt/beta-catenin signaling to promote the osteogenesis of CTLA4-modified human bone marrow-mesenchymal stem cells. Sci Rep 7:41634CrossRefGoogle Scholar
  75. 75.
    Bonnet N, Conway SJ, Ferrari SL (2012) Regulation of beta catenin signaling and parathyroid hormone anabolic effects in bone by the matricellular protein periostin. Proc Natl Acad Sci U S A 109:15048–15053CrossRefGoogle Scholar
  76. 76.
    Robinson JA et al (2006) Wnt/beta-catenin signaling is a normal physiological response to mechanical loading in bone. J Biol Chem 281:31720–31728CrossRefGoogle Scholar

Copyright information

© Springer Nature Singapore Pte Ltd. 2019

Authors and Affiliations

  1. 1.INSERM UMR1163, Imagine InstituteParis Descartes UniversityParisFrance

Personalised recommendations