Skip to main content
  • ASM
    • Antimicrobial Agents and Chemotherapy
    • Applied and Environmental Microbiology
    • Clinical Microbiology Reviews
    • Clinical and Vaccine Immunology
    • EcoSal Plus
    • Eukaryotic Cell
    • Infection and Immunity
    • Journal of Bacteriology
    • Journal of Clinical Microbiology
    • Journal of Microbiology & Biology Education
    • Journal of Virology
    • mBio
    • Microbiology and Molecular Biology Reviews
    • Microbiology Resource Announcements
    • Microbiology Spectrum
    • Molecular and Cellular Biology
    • mSphere
    • mSystems
  • Log in
  • My alerts
  • My Cart

Main menu

  • Home
  • Articles
    • Current Issue
    • Accepted Manuscripts
    • Archive
    • Minireviews
  • For Authors
    • Submit a Manuscript
    • Scope
    • Editorial Policy
    • Submission, Review, & Publication Processes
    • Organization and Format
    • Errata, Author Corrections, Retractions
    • Illustrations and Tables
    • Nomenclature
    • Abbreviations and Conventions
    • Publication Fees
    • Ethics Resources and Policies
  • About the Journal
    • About MCB
    • Editor in Chief
    • Editorial Board
    • For Reviewers
    • For the Media
    • For Librarians
    • For Advertisers
    • Alerts
    • RSS
    • FAQ
  • Subscribe
    • Members
    • Institutions
  • ASM
    • Antimicrobial Agents and Chemotherapy
    • Applied and Environmental Microbiology
    • Clinical Microbiology Reviews
    • Clinical and Vaccine Immunology
    • EcoSal Plus
    • Eukaryotic Cell
    • Infection and Immunity
    • Journal of Bacteriology
    • Journal of Clinical Microbiology
    • Journal of Microbiology & Biology Education
    • Journal of Virology
    • mBio
    • Microbiology and Molecular Biology Reviews
    • Microbiology Resource Announcements
    • Microbiology Spectrum
    • Molecular and Cellular Biology
    • mSphere
    • mSystems

User menu

  • Log in
  • My alerts
  • My Cart

Search

  • Advanced search
Molecular and Cellular Biology
publisher-logosite-logo

Advanced Search

  • Home
  • Articles
    • Current Issue
    • Accepted Manuscripts
    • Archive
    • Minireviews
  • For Authors
    • Submit a Manuscript
    • Scope
    • Editorial Policy
    • Submission, Review, & Publication Processes
    • Organization and Format
    • Errata, Author Corrections, Retractions
    • Illustrations and Tables
    • Nomenclature
    • Abbreviations and Conventions
    • Publication Fees
    • Ethics Resources and Policies
  • About the Journal
    • About MCB
    • Editor in Chief
    • Editorial Board
    • For Reviewers
    • For the Media
    • For Librarians
    • For Advertisers
    • Alerts
    • RSS
    • FAQ
  • Subscribe
    • Members
    • Institutions
Research Article

Role of SMPD3 during Bone Fracture Healing and Regulation of Its Expression

Garthiga Manickam, Pierre Moffatt, Monzur Murshed
Garthiga Manickam
aFaculty of Dentistry, McGill University, Montreal, Quebec, Canada
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Pierre Moffatt
bShriners Hospital for Children, McGill University, Montreal, Quebec, Canada
cDepartment of Human Genetics, McGill University, Montreal, Quebec, Canada
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
Monzur Murshed
aFaculty of Dentistry, McGill University, Montreal, Quebec, Canada
bShriners Hospital for Children, McGill University, Montreal, Quebec, Canada
dDepartment of Medicine, McGill University, Montreal, Quebec, Canada
  • Find this author on Google Scholar
  • Find this author on PubMed
  • Search for this author on this site
DOI: 10.1128/MCB.00370-18
  • Article
  • Figures & Data
  • Info & Metrics
  • PDF
Loading

ABSTRACT

Sphingomyelin phosphodiesterase 3 (SMPD3), a lipid-metabolizing enzyme present in bone and cartilage, has important roles in the developing skeleton. We previously showed that SMPD3 deficiency results in delayed extracellular matrix (ECM) mineralization and severe skeletal deformities in an inducible knockout mouse model, Smpd3flox/flox; Osx-Cre mice, in which Smpd3 was ablated in Osx-expressing chondrocytes and osteoblasts during early skeletogenesis. However, as shown in the current study, ablation of Smpd3 postnatally in 3-month-old Smpd3flox/flox; Osx-Cre mice resulted in only a mild bone mineralization defect. Interestingly, though, there was a marked increase of unmineralized osteoid in the fractured tibiae of 3-month-old Smpd3flox/flox; Osx-Cre mice. As was the case in the embryonic bones, we also observed impaired chondrocyte apoptosis at the fracture sites of Smpd3flox/flox; Osx-Cre mice. We further examined how Smpd3 expression is regulated in ATDC5 chondrogenic cells by two major regulators of chondrogenesis, bone morphogenetic protein 2 (BMP-2) and PTHrP. Our data show that BMP-2 positively regulates Smpd3 expression via p38 mitogen-activated protein kinase. Taken together, our findings show that SMPD3 plays a significant role in ECM mineralization and chondrocyte apoptosis during fracture healing. Furthermore, our gene expression analyses suggest that BMP-2 and PTHrP exert opposing effects on the regulation of Smpd3 expression in chondrocytes.

INTRODUCTION

The majority of bones in our body are formed through endochondral ossification, which is a complex process regulated by the concerted activities of various signaling molecules and downstream transcription factors. Endochondral ossification is initiated by the condensation of mesenchymal stem cells (MSCs), which initially express Sox9, a key regulator of chondrogenesis (1, 2). SOX9 and RUNX2, another critical transcription factor regulating skeletal development, induce the differentiation and maturation of chondrocytes (3–5), which ultimately give rise to four distinct cellular zones, collectively known as the growth plate. Each of these zones carries chondrocytes at a particular stage of differentiation: (i) the resting zone of chondrocyte precursors, (ii) the proliferating zone with chondrocytes that produce type II collagen among various other extracellular matrix (ECM) proteins, (iii) the prehypertrophic zone, and (iv) the hypertrophic zone where the terminally differentiated cells synthesize type X collagen before going through programmed cell death. Our group recently reported important roles of a lipid metabolizing enzyme, sphingomyelin phosphodiesterase 3 (SMPD3), in growth plate development and bone mineralization. We generated an inducible knockout mouse model, Smpd3flox/flox; Osx-Cre, to selectively ablate Smpd3 in both Osx-expressing chondrocytes and osteoblasts in the embryonic skeleton. Characterization of the skeletal phenotype of these embryos showed that SMPD3 activity in both of these cell types is essential for the initiation of ECM mineralization. Furthermore, SMPD3 is also required for apoptosis of hypertrophic chondrocytes in the developing growth plate (6, 7).

Despite SMPD3’s critical functions during growth plate development, the regulation of its expression in chondrocytes is still not well understood. Previous studies reported that bone morphogenetic protein (BMP) signaling induces Smpd3 expression in C2C12 myoblasts and chondrocyte cultures via the upregulation of Runx2 (8, 9). We recently showed that treatment of ATDC5 chondrogenic cells with forskolin, a PTHrP surrogate, markedly downregulates Smpd3 expression despite a significant upregulation of Runx2 expression (7). This observation implies that Smpd3 expression can be regulated independent of RUNX2 in chondrocytes.

Based on our knowledge of the critical role of SMPD3 in endochondral bone development and considering that nonrigid bone fracture healing recapitulates the process of endochondral ossification, we hypothesize that the loss of SMPD3 function in chondrocytes and osteoblasts would adversely affect the process of fracture healing in our experimental model. In the present study, we tested this hypothesis. When the tibiae of Smpd3flox/flox; Osx-Cre mice, in which Smpd3 was ablated postnatally, were surgically fractured, there was a marked increase of unmineralized bone (osteoid) at the fracture sites in comparison to the control mice. We also observed an increase in cartilaginous ECM and impaired chondrocyte apoptosis at the fracture sites of these mice.

In order to understand the regulation of Smpd3 expression in chondrocytes, we investigated the downstream events of both BMP and PTHrP signaling pathways in ATDC5 chondrogenic cells. We show that BMP-2 can upregulate Smpd3 expression in ATDC5 cells via p38 mitogen-activated protein kinase (MAPK) and its downstream transcription factors. In contrast, PTHrP treatment suppresses BMP-2-mediated induction of Smpd3. Here, we propose a model in which BMP signaling and PTHrP signaling, two major signaling events regulating chondrogenesis, converge on p38 MAPK to exert opposing effects on Smpd3 expression.

Our present study suggests that during nonrigid fracture healing involving endochondral ossification, SMPD3 plays a critical role as seen in the developing growth plate. Furthermore, p38 MAPK acts as a major signaling node to regulate Smpd3 expression in chondrocytes.

RESULTS

Generation of viable Smpd3flox/flox; Osx-Cre mice using the inducible Cre system.Smpd3flox/flox; Osx-Cre mice, in which Smpd3 was conditionally ablated in the chondrocytes and osteoblasts during the early stage of skeletogenesis, exhibit severe congenital skeletal deformities resulting in perinatal lethality (7). Although osterix (OSX) was initially identified as an osteoblast-specific transcription factor, later studies showed its expression in the growth plate of the developing skeleton (10–13). To validate this further, we performed immunofluorescence with an anti-OSX antibody and detected OSX protein in prehypertrophic chondrocytes in the developing growth plate (Fig. 1A to C), which also express SMPD3.

FIG 1
  • Open in new tab
  • Download powerpoint
FIG 1

Generation of viable Smpd3flox/flox; Osx-Cre mice. (A) Superimposed light and fluorescence microscopy images of the humerus of an E15.5 embryo showing the different zones in the developing growth plate: resting chondrocytes (RC), proliferating chondrocytes (PC), prehypertrophic chondrocytes (PHC), hypertrophic chondrocytes (HC), and the midshaft (M) region. The blue stain represents the H33258-stained nuclei. (B and C) Fluorescence microscopy shows the OSX-positive nuclei in the differentiating chondrocytes (arrow) (B) shows and osteoblasts in the midshaft (arrow) and the periosteum (*) (C). (D) Breeding scheme to generate Smpd3flox/flox; Osx-Cre mice and table depicting the survival of Smpd3flox/flox; Osx-Cre pups with or without doxycycline treatment of the pregnant dames. *, number of carcasses retrieved for genotyping. (E) In the absence of doxycycline, E18.5 Smpd3flox/flox; Osx-Cre embryos show a severe skeletal phenotype, where the limbs are shorter and bent, and the calvaria is undermineralized. This was prevented by doxycycline treatment, since treated Smpd3flox/flox; Osx-Cre embryos are similar to control Smpd3flox/flox embryos.

In order to generate viable Smpd3flox/flox; Osx-Cre mice, the inducible Osx-Cre transgene expression was suppressed by treating the pregnant dames with doxycycline in water and diet from embryonic day 10.5 (E10.5) to postnatal day 7 (P7) (13). This treatment was sufficient to generate viable Smpd3flox/flox; Osx-Cre mice (Fig. 1D). We then compared the skeletal phenotypes of Smpd3flox/flox; Osx-Cre E18.5 embryos obtained from the pregnant dames fed with (DOX+) or without (DOX–) doxycycline by performing whole-mount alizarin red and alcian blue staining. We observed that Smpd3flox/flox; Osx-Cre embryos from the DOX+ group were indistinguishable from that of Smpd3flox/flox embryos since no skeletal deformities were detected. On the contrary, Smpd3flox/flox; Osx-Cre embryos from the untreated group exhibited the previously reported skeletal deformities (i.e., bent limbs and undermineralized calvaria) (Fig. 1E).

Postnatal ablation of Smpd3 in Smpd3flox/flox; Osx-Cre mice results in a mild bone mineralization defect.We next examined how SMPD3 deficiency in Osx-Cre-expressing cells affects the adult skeleton. For this purpose, we analyzed the bones of 3-month-old Smpd3flox/flox; Osx-Cre mice (viable due to postbirth ablation of Smpd3) by micro-computed tomography (micro-CT) and histology. Micro-CT analyses of the proximal tibiae did not reveal any significant differences in the trabecular and cortical parameters (Fig. 2A and B). However, von Kossa and van Gieson (VK/VG)-stained histological sections of the vertebral bones showed a mild but significant increase of osteoid volume over bone volume (OV/BV) (Fig. 2C and D). This suggests that SMPD3 function is critical for normal skeletal development in the embryos but has only a minor role in the adult skeleton.

FIG 2
  • Open in new tab
  • Download powerpoint
FIG 2

Characterization of skeletal parameters of adult Smpd3flox/flox; Osx-Cre mice. (A) Bone surface over bone volume (BS/BV), trabecular thickness (Tb/Th), bone volume over tissue volume (BV/TV), and trabecular number (Tb.N) were not significantly different between WT, Smpd3flox/flox, and Smpd3flox/flox; Osx-Cre mice. (B) Total tissue area (Tt.Ar), cortical area (Ct.Ar), cortical area over total tissue area (Ct.Ar/Tt.Ar), and cortical thickness (Ct.Th) were not significantly different between WT, Smpd3flox/flox and Smpd3flox/flox; Osx-Cre mice. (C) Histological sections of vertebra from 3-month-old WT, Osx-Cre, Smpd3flox/flox, and Smpd3flox/flox; Osx-Cre mice stained with VK/VG at high magnification showing the presence of osteoid (arrow). (D) Quantification showing a mild but significant increase in OV/BV in Smpd3flox/flox; Osx-Cre vertebra compared to the WT, Osx-Cre, and Smpd3flox/flox vertebra. Error bars represent standard deviations. **, P < 0.01; ***, P < 0.001.

Increased unmineralized bone in the calli of Smpd3flox/flox; Osx-Cre tibia.In order to investigate further whether SMPD3 is required in the adult bone after injury, we performed rodded-immobilized fracture surgery on 2-month-old wild-type (WT), Osx-Cre, Smpd3flox/flox, and Smpd3flox/flox; Osx-Cre tibiae. First, we used a PCR-based approach using primers external to the loxP sites (Fig. 3A) to verify whether the “floxed” Smpd3 sequence is excised by the Osx promoter-driven Cre-recombinase enzyme. The presence of an ∼200-bp amplicon in the amplified DNA from the calli of Smpd3flox/flox; Osx-Cre mice (Fig. 3B, upper panel) confirmed that the “floxed” sequence was deleted. As expected, control Smpd3flox/flox callus DNA did not show this band. PCR performed using Elastin (non-“floxed”) gene-specific primers (amplicon size, ∼500 bp) from the same DNA samples were presented as a loading control (Fig. 3B, lower panel). We next analyzed the callus size and mineralized tissue volume using micro-CT at 14 and 28 days postsurgery and did not find a significant difference among WT, Osx-Cre, Smpd3flox/flox, and Smpd3flox/flox; Osx-Cre tibiae at any of these time points. Similarly, when the mineralized tissue volume was normalized by the callus size, no significant differences were observed among the four genotypes (Fig. 3C and D).

FIG 3
  • Open in new tab
  • Download powerpoint
FIG 3

Excision of Smpd3 gene in fracture calli and analyses by micro-CT. (A) Schematic representation of the targeted Smpd3 locus. A neomycin resistance cassette (Neor) flanked by two loxP sites (triangles) and a separate third loxP site were inserted into intron 8 and exon 9 downstream of the stop codon, respectively. The primers (F1 and R1) used for detecting the excision of the “floxed” Smpd3 sequence are indicated. (B) The ∼200-bp amplicons representing the altered Smpd3 alleles after excision are obtained only from the Smpd3flox/flox; Osx-Cre but not from control Smpd3flox/flox callus DNA (upper panel). A ∼500-bp fragment of a non-“floxed” Eln gene was amplified as a loading control (lower panel). Three callus samples were analyzed for each genotype. (C and D) Quantification of callus size, mineralized tissue volume, and ratio of mineralized tissue volume to callus size in WT, Smpd3flox/flox, Osx-Cre, and Smpd3flox/flox; Osx-Cre fractured tibiae showing no difference in any of the parameters at 2 weeks (C) and 4 weeks (D) postsurgery. Error bars represent standard deviations.

We next processed the scanned samples for histological analyses. VK/VG staining showed delayed healing hallmarked by the increased presence of osteoid at the fracture sites in Smpd3flox/flox; Osx-Cre tibiae compared to that of WT, Osx-Cre, and Smpd3flox/flox tibiae analyzed 14 days postsurgery (Fig. 4A). At 28 days postsurgery, the increased osteoid volume in the Smpd3flox/flox; Osx-Cre callus appeared to be less severe (Fig. 4B). We further verified these observations by quantifying the amount of OV/BV and showed that there was a significant increase of OV/BV in Smpd3flox/flox; Osx-Cre tibiae at 14 days postsurgery (Fig. 4C). Although the OV/BV at the Smpd3flox/flox; Osx-Cre fracture site decreased markedly by 28 days postsurgery, it remained significantly higher than that of WT, Osx-Cre, and Smpd3flox/flox tibiae (Fig. 4D).

FIG 4
  • Open in new tab
  • Download powerpoint
FIG 4

Increased osteoid volume in Smpd3flox/flox; Osx-Cre fractured tibia postsurgery. (A) VK/VG stained sections of a 2-week-postsurgery tibia showing an increased amount of osteoid (pink stain indicated by arrow) in Smpd3flox/flox; Osx-Cre mice compared to that of WT, Smpd3flox/flox, and Osx-Cre mice. The bottom panels show magnified views of the boxed area. (B) VK/VG-stained sections of 4-week-postsurgery tibiae showing that the amount of osteoid (arrow) in Smpd3flox/flox; Osx-Cre callus is modestly higher in comparison to that of WT, Smpd3flox/flox, and Osx-Cre fractured tibiae. (C and D) Quantification showing a significant increase in OV/BV in Smpd3flox/flox; Osx-Cre fracture sites at 2 weeks postsurgery compared to that of the WT, Osx-Cre, and Smpd3flox/flox mice. By 4 weeks postsurgery, a modest but significant increase of osteoid volume was observed in Smpd3flox/flox; Osx-Cre fracture sites. Error bars represent standard deviations. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Developmental anomalies of the skeleton are recapitulated during fracture healing in Smpd3flox/flox; Osx-Cre mice.Alcian blue- and van Gieson (AB/VG)-stained histological sections of the fractured tibiae at 2 weeks postsurgery showed an increased amount of cartilage matrix (blue stain) in Smpd3flox/flox; Osx-Cre mice compared to the control fractured tibiae (Fig. 5A). We next stained the histology sections with a nuclear stain, H33258 (Hoechst stain), and quantified the number of cells with a ring structure, which are indicative of nuclear chromatin condensation leading up to cellular apoptosis. We noticed that the number of apoptotic chondrocyte-like cells within the Smpd3flox/flox; Osx-Cre fractured tibia was significantly lower compared to the controls (Fig. 5B and C). Although we consistently observed a tendency for an increase of the cartilage matrix volume over callus volume at the Smpd3flox/flox; Osx-Cre fracture sites, histomorphometric analyses revealed that this was not statistically significant (Fig. 5D).

FIG 5
  • Open in new tab
  • Download powerpoint
FIG 5

Increased cartilage matrix and delayed chondrocyte apoptosis in the fracture calli of Smpd3flox/flox; Osx-Cre mice. (A) AB/VG-stained sections of 2-week-postsurgery tibiae showing an increase in the amount of cartilage matrix (blue) in Smpd3flox/flox; Osx-Cre fracture sites compared to that of WT, Osx-Cre, and Smpd3flox/flox mice. The bottom panels show magnified views of the boxed areas in the corresponding upper panels. (B) Hoechst staining showed the presence of condensed chromatin ring structures in the nuclei at WT, Osx-Cre, and Smpd3flox/flox fracture callus tissues. Nuclei with such structures are fewer in number in the Smpd3flox/flox; Osx-Cre fracture callus. (C) Quantification showing significant decrease in the number of apoptotic cells with condensed chromatin in the nucleus in an Smpd3flox/flox; Osx-Cre callus compared to WT, Osx-Cre, and Smpd3flox/flox callus tissues. (D) Quantification showing a tendency for an increase in cartilage matrix volume over callus volume at the Smpd3flox/flox; Osx-Cre fracture sites compared to that of WT, Osx-Cre, and Smpd3flox/flox mice. Error bars represent standard deviations. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Upregulation of Bmp-2 and Smpd3 expression during fracture healing.Gene expression analyses of the tissues at the fracture sites from 3-month-old WT mice indicated a significant upregulation of Bmp-2, a critical signaling molecule for endochondral bone development, at 13 days postsurgery (Fig. 6A). Similarly, we also observed a significant upregulation in Smpd3 expression in the same samples (Fig. 6B). We validated the latter finding by immunohistochemistry and found that at 14 and 28 days postsurgery, the fracture callus expressed more SMPD3 compared to the same region of the bone in the nonfractured contralateral leg (Fig. 6C).

FIG 6
  • Open in new tab
  • Download powerpoint
FIG 6

Regulation of Smpd3 expression in chondrocytes by BMP-2. (A and B) Gene expression analyses of 3-month-old fractured WT bones showing a significant upregulation of Bmp-2 expression (A) and Smpd3 expression (B) in the callus. Expression analyses by qRT-PCR were performed 13 days postsurgery. (C) Immunohistochemistry with anti-SMPD3 antibody shows an increased presence of SMPD3 in the callus of WT fractured tibia at 14 and 28 days postsurgery. A bone section of the unfractured contralateral tibia was used as a control. (D) ATDC5 chondrogenic cells treated with 300 ng/ml of BMP-2 for 48 h show a significant upregulation of chondrogenic marker Col2a1, Col10a1, Sox6, Sox9, Osx, Mef2c, and Smpd3 expression. (E) Differentiated ATDC5 cells (cultured in the presence of ascorbic acid and β-glycerophosphate for 6 days) when treated with 2 μM dorsomorphin (Dorso) show significant downregulation of Col2a1, Col10a1, Sox5, Sox6, Osx, Mef2c, and Smpd3 expression. Cells were treated with dorsomorphin for 30 min every 2 days, before the medium was changed. Error bars represent standard deviations. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

The observation that both Bmp-2 expression and its downstream target SMPD3 was significantly upregulated in the fracture callus prompted us to investigate the regulatory cues downstream of the BMP-2 signaling pathway inducing Smpd3 expression. We first treated ATDC5 chondrogenic cells with BMP-2 for 48 h and examined the expression of various chondrocyte markers by quantitative real-time PCR (qRT-PCR). We observed a significant upregulation of Col2a1, Col10a1, Sox6, Sox9, Osx, and Mef2c (Fig. 6D). There was also a marked upregulation of Smpd3 expression. Interestingly, there was no effect of BMP-2 on the expression of Sox5 and Runx2 at this time point.

We next cultured ATDC5 cells in a differentiation medium supplemented with ascorbic acid and β-glycerophosphate for 6 days. Cells were treated with or without dorsomorphin, a selective inhibitor of BMP signaling. At the end of the culture period, gene expression analyses were performed by qRT-PCR. Dorsomorphin treatment downregulated Col2a1, Col10a1, Sox5, Sox6, Osx, and Mef2c expression significantly, while a mild downregulation of Sox9 and Runx2 expression was observed at this time point. As expected, Smpd3 expression was markedly downregulated in the treated cells (Fig. 6E).

BMP-2 regulates Smpd3 expression through a noncanonical pathway.We next examined whether BMP-2 had an immediate direct effect on the proximal promoter of Smpd3. We transfected ATDC5 cells with a reporter construct, pSmpd3-luc2, which contains a 1.9-kb Smpd3 proximal promoter fragment upstream of the firefly luciferase gene. Transfection of this construct into ATDC5 cells resulted in an almost 30-fold increase (compared to empty vector-transfected cells) in luciferase activity at 8 h and an almost 50-fold increase at 48 h. However, when the transfected cells were treated with BMP-2 for 8 or 48 h, we did not observe any further upregulation of the Smpd3 promoter activity at any of these time points, whereas the control luciferase expression vector, pIDWT4F-luc, carrying four tandem repeats of a BMP-responsive element present in the Id1 gene was activated in the presence of BMP-2 (Fig. 7A). In agreement with these findings, in silico analyses using MatInspector did not reveal any consensus BMP response elements within the 5-kb Smpd3 proximal promoter (data not shown).

FIG 7
  • Open in new tab
  • Download powerpoint
FIG 7

BMP-2 regulates Smpd3 through the noncanonical pathway. (A) ATDC5 cells were transiently transfected with a luciferase expression vector driven by a 1.9-kb Smpd3 proximal promoter, pSmpd3-luc. There was no significant induction of Smpd3 promoter activity as detected by luciferase activity at 8 and 48 h after BMP-2 treatment, whereas cells transfected with a control luciferase expression vector, pIDWT4F-luc, showed significantly upregulated luciferase activity in the presence of BMP-2. (B) Cells were treated with 20 μM PD98059, an ERK1/2 inhibitor, with or without BMP-2. There was a significant upregulation of Smpd3 expression upon this treatment. (C) Cells were treated with 5 μM BI-87G3, a JNK inhibitor, with or without BMP-2. This treatment led to a significant upregulation of Smpd3 expression. (D) Cells were treated with 10 μM SB202190, a p38 inhibitor, with or without BMP-2. There was a significant downregulation of Smpd3 expression. Error bars represent standard deviations. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Our data suggested that the canonical arm of BMP signaling involving SMADs may not have a direct effect on the transcriptional regulation of Smpd3. Considering this, we next investigated the effect of the noncanonical arm of BMP signaling involving the mitogen-activated protein kinases (MAPKs). We treated ATCD5 cells with the following pharmacological inhibitors: (i) 20 μM PD98059 (extracellular signal-regulated kinase 1/2 [ERK1/2] inhibitor), (ii) 5 μM BI-87G3 (c-Jun N-terminal kinase [JNK] inhibitor), or (iii) 10 μM SB202190 (p38 MAPK inhibitor). The cells were pretreated with the inhibitor of interest for 1 h and then treated with or without BMP-2 for 48 h. Cells treated with ERK1/2 inhibitor PD98059 or JNK inhibitor BI-87G3, in the presence of BMP-2, showed a significant upregulation in Smpd3 expression (Fig. 7B and C). In contrast to that, we observed a significant reduction in BMP-2-induced Smpd3 expression when cells were treated with p38 inhibitor SB202190 (Fig. 7D).

Transcription factors downstream of p38 MAPK regulate Smpd3 expression.We treated the BMP-2-induced ATDC5 cultures with SB202190 and examined how p38 MAPK inhibition affects the expression of Mef2c, Sox6, Sox9, and Osx. The expression levels of all of these transcription factors were significantly downregulated (Fig. 8A to D). In a follow-up experiment, ATDC5 cells were transfected with Mef2c, Sox6, Sox9, and Osx expression vectors. Overexpression of all these transcription factors resulted in a significant upregulation of endogenous Smpd3 expression (Fig. 8E to H). The transfection efficiency (∼50%) was analyzed by quantifying the green fluorescent protein (GFP)-positive cells transfected by a GFP expression vector under identical conditions (Fig. 8I).

FIG 8
  • Open in new tab
  • Download powerpoint
FIG 8

p38 MAPK upregulates key transcription factors that regulate Smpd3 expression. (A to D) ATDC5 cells treated with BMP-2 and SB202190, a p38 inhibitor, showed a reduction of Mef2c (A), Sox6 (B), Sox9 (C), and Osx (D) expression. (E to H) ATDC5 cells separately transfected with the expression vectors Mef2c (E), Sox6 (F), Sox9 (G), or Osx (H) showed an upregulated expression of the corresponding transcription factors (left), as well as significant upregulation of Smpd3 expression (right). (I) A representative image shows cells transfected with a GFP expression vector under the transfection conditions described above. Quantification of the GFP-positive cells shows that ∼48.6% cells were transfected. Error bars represent standard deviations. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

PTHrP suppresses BMP-2-induced Smpd3 expression.We previously showed that PTHrP suppresses Smpd3 expression by downregulating Sox9 expression (7). In the present study, we investigated how PTHrP and BMP signaling pathways may cross talk to regulate the expression of major transcription factors upstream of Smpd3. To address this, we examined the status of Smpd3 expression in ATDC5 cultures treated with both PTHrP (0.1 μM) and BMP-2 (300 ng/ml). We observed a downregulation of Mef2c, Sox6, Sox9, and Osx expression (Fig. 9A to D). In agreement with these findings, we also found that the BMP-2-induced Smpd3 expression was markedly reduced by PTHrP (Fig. 9E).

FIG 9
  • Open in new tab
  • Download powerpoint
FIG 9

PTHrP suppresses BMP-2 induced Smpd3 expression. (A to E) Gene expression analysis of cells treated with PTHrP and BMP-2 showing a suppression of Mef2c (A), Sox6 (B), Sox9 (C), Osx (D), and Smpd3 (E) expression. (F) Scheme of the 2-kb Smpd3 proximal promoter showing the presence of SRY binding sites (AACAAA). (G) The major cluster of SRY binding sites spanning nucleotides –595 to –646 in the Smpd3 proximal promoter was deleted to generate a luciferase reporter construct (pSmpd3ΔSRY-luc2). Cotransfection of HEK293 cells with this and a Sox9 expression vector shows a significant reduction of Smpd3 proximal promoter activity, as detected by the luciferase activity in compared to cells transfected with the Sox9 expression vector alone. Error bars represent standard deviations. *, P < 0.05; **, P < 0.01; ***, P < 0.001.

We earlier reported the absence of consensus SOX9 binding sites in the proximal Smpd3 promoter and predicted that the cluster of SRY binding sites (AACAAA) spanning nucleotides –595 to –646 of the Smpd3 gene (Fig. 9F) might recruit SOX9. To investigate this further, we deleted the cluster using an in vitro PCR-based method and the mutated promoter-luciferase construct was used with or without the Sox9 expression vector to transfect HEK293 cells, which normally do not express SOX9. A luciferase assay demonstrated that the deletion of the SRY cluster significantly abolishes the SOX9-mediated induction of the Smpd3 proximal promoter (Fig. 9G).

DISCUSSION

SMPD3 has emerged as a key regulator of skeletal development. SMPD3 deficiency affects the mineralization of both intramembranous and endochondral bones. In addition, the loss of SMPD3 activity affects normal apoptosis of hypertrophic chondrocytes in the developing skeleton. The prosurvival roles of SMPD3 in the skeletal cells is demonstrated by the perinatal lethality of pups with conditional ablation of Smpd3 in chondrocytes and osteoblasts in Smpd3flox/flox; Osx-Cre mice.

We show here that viable Smpd3flox/flox; Osx-Cre mice can be generated by using the regulatable Cre recombinase expression feature in Osx-Cre transgenic mice (13). By feeding pregnant mice with DOX, we turned off the expression of Cre recombinase in Smpd3flox/flox; Osx-Cre embryos from E10.5 to P7, thereby preventing the deletion of the “floxed” Smpd3 sequence encoding the active site of the enzyme during this period. This delayed ablation of Smpd3 was sufficient to keep all the Smpd3flox/flox; Osx-Cre mice alive with no visual skeletal deformities and apparently normal life span.

Despite a critical role of SMPD3 in the embryonic skeleton, this enzyme appears to play a less critical role in the adult skeleton since no effects on static bone parameters were observed when Smpd3flox/flox; Osx-Cre mice were analyzed at 3 months of age. Only a mild increase of osteoid was observed in these mice. These findings are in agreement with the reported analyses of the bones of fro/fro mice, in which Smpd3 is ablated globally (6, 7, 14). Collectively, our findings from past and current studies suggest that the mechanism of ECM mineralization during the early developmental stages in which SMPD3 plays a critical role may differ from that of the postnatal and adult skeleton.

The observation that Smpd3 expression and protein levels are upregulated in the fracture callus prompted us to further investigate the role of this enzyme during fracture healing. Since the steps of endochondral ossification are recapitulated during secondary fracture healing (15), ablation of Smpd3 in both chondrocytes and osteoblasts was necessary for our experiments. This requirement justifies the use of Smpd3flox/flox; Osx-Cre mice in this study since Smpd3 is ablated in both of these cell types in these mice. It should be noted here that in order to avoid the reported skeletal anomalies seen in young Osx-Cre mice (16), we performed all our experiments after the animals reached 2 months of age. In addition, we included control Osx-Cre mice in our experiments when appropriate.

Although there was a mild increase of osteoid in adult Smpd3flox/flox; Osx-Cre mice, the amount of osteoid was markedly increased by 2 weeks postsurgery at the fracture site of the healing bones. In general, an increase of osteoid at the fracture site is seen during healing (17). However, the accumulation of osteoid is further accentuated in Smpd3flox/flox; Osx-Cre fractures in comparison to the control mice. This can be explained by the combined effects of increased synthesis of new ECM at the fracture site and its impaired mineralization caused by the loss of SMPD3 activity. By 4 weeks postsurgery, although the osteoid volume is still significantly higher, the severity of this trait decreases markedly. This is most likely caused by the normalization of the bone formation rate at the fracture site toward the end of the healing period. It is also possible that alternative pathways compensate for SMPD3 deficiency during the later stages of fracture healing.

In agreement with the notion that nonrigid fracture healing mimics endochondral ossification and our previous work showing a reduced apoptosis of SMPD3-deficient hypertrophic chondrocytes, we expected a decrease of the number of apoptotic chondrocytes at the fracture sites in Smpd3flox/flox; Osx-Cre mice. Our analyses confirmed that this is indeed the case. Taking our results together, for the first time, we demonstrate here that although in the adult skeleton SMPD3 does not play a major role (as shown by a mild bone mineralization defects), during fracture healing, locally produced SMPD3 plays an important role in the cartilage and bone matrix mineralization. This is an important novel finding that shows the reprogramming of the expression and function of SMPD3, an enzyme primarily required for embryonic skeletogenesis, during the healing of adult bone fracture.

Recently, several intracellular enzymes have been shown to play critical roles during the early phases of skeletal tissue mineralization. These intracellular enzymes include (i) SMPD3, which generates ceramide and phosphocholine from sphingomyelin, (ii) choline kinases, which generate phosphocholine by phosphorylating choline in the cytosol, and (iii) phosphatase, orphan 1 (PHOSPHO1), which cleaves phosphocholine to liberate inorganic phosphate. Interestingly, all these enzymes are involved in phosphocholine metabolism (18, 19). Indeed, we recently reported a similar impairment of fracture healing in Phospho1–/– mice as seen in Smpd3flox/flox; Osx-Cre mice, further suggesting a metabolic link between these two enzymes (17).

Until now only a few studies examined how Smpd3 expression is regulated in skeletal cells. A thorough understanding of how various signaling molecules and downstream transcription factors may regulate this process is still missing. Earlier reports demonstrated that BMP-2 induces Smpd3 expression in chondrocyte cultures (8, 9). Likewise, it is possible that upregulation of BMP-2 induces Smpd3 expression at the fracture site. In addition, Smpd3 expression in the callus can be induced by cytokines that are released during the inflammatory stages. These cytokines can then activate MAPKs, particularly p38 MAPKs. Considering that p38 MAPKs are activated by both inflammatory cytokines and BMP signaling, it is likely that p38 MAPKs act as a major signaling node to integrate multiple regulatory pathways to induce Smpd3 expression in the fractured bone (20–22).

Our group reported earlier that 1 h of PTHrP (0.1 μM) treatment of ATDC5 cells differentiated for 6 days can markedly suppress Smpd3 expression. Interestingly, the expression of RUNX2, a key transcription factor for skeletal development and an inducer of Smpd3 expression, was not downregulated in these cells. This observation prompted us to examine whether other transcription factors regulating chondrogenesis may affect Smpd3 expression. Indeed, we identified SOX9 as a major regulator of Smpd3 expression in ATDC5 cells. In the present study, we performed further in-depth analyses to understand the mode of interactions between BMP and PTHrP signaling, two major pathways regulating chondrogenesis, on the regulation of Smpd3 expression.

Our in silico analyses showing the absence of the BMP response element in the 5-kb Smpd3 promoter and in vitro experiments using promoter constructs at two different time points suggest that the canonical arm of BMP signaling pathway may not be a direct regulator of Smpd3 expression. We therefore focused on the MAPKs involved in the noncanonical BMP signaling pathway. We found that cultures induced by BMP-2, when treated with pharmacologic inhibitors of ERK1/2 or JNK, upregulate Smpd3 expression. Since these two MAPKs are actually induced by BMP signaling, we concluded that they are not responsible for the BMP-mediated upregulation of Smpd3 expression. On the contrary, pharmacologic inhibition of p38 resulted in a marked downregulation of Smpd3 expression in BMP-2-treated cells, suggesting that p38 acts as a major downstream inducer of this gene.

p38 MAPK signaling has been shown to play a major role in endochondral bone development, particularly in the maturation and apoptosis of chondrocytes in the developing growth plates (20). Furthermore, inhibition of p38 activity in chondrocytes isolated from the articular cartilage of patients with osteoarthritis (OA) reduced the number of apoptotic chondrocytes in vitro (23). Our data identifying p38 as a major regulator of Smpd3 expression imply that SMPD3 may act as a downstream effector of p38 to regulate chondrocyte apoptosis. This possibility is supported by the observation that the loss of SMPD3 activity reduces the number of apoptotic hypertrophic chondrocytes (6–8).

The major transcription factors acting downstream of p38 MAPK have been identified. Among them, based on the relevance to the present study, we examined MEF2C, SOX6, SOX9, and OSX for their role in the regulation of Smpd3 expression. Tew et al. reported that stimulation of p38 MAPK activity in chondrocytes results in an increase of Sox9 mRNA (24, 25). In addition, SOX6 has been shown to be regulated by BMP signaling (26). In agreement with these findings, we show here that, as is the case with BMP-2 signaling and p38 activation, transfection of ATDC5 cells by Mef2c, Sox6, Sox9, or Osx expression vectors upregulates Smpd3 expression.

Stanton et al. showed that p38 MAPK via MEF2C regulates gene expression in hypertrophic chondrocytes (20, 21). The in vivo ablation of Mef2c in chondrocytes impairs chondrocyte hypertrophy, which results in an accumulation of hypertrophic chondrocytes in the growth plate (27). This observation is similar to that seen in Smpd3-deficient chondrocytes. In addition, we found that OSX can upregulate Smpd3 expression in ATDC5 cells. This observation has physiologic significance since, in addition to osteoblasts, OSX is also expressed in prehypertrophic chondrocytes, the primary cell types expressing Smpd3 in the developing growth plates.

PTHrP signaling is known to inhibit p38 kinase activity in chondrocytes (28). Our data presented here show that BMP-2-mediated induction of Smpd3 expression can be suppressed by PTHrP. Indeed, p38 can act as a common target for both BMP and PTHrP signaling in chondrocytes to modulate the antagonistic effects of these pathways. This notion is further supported by the observation that the expression of transcription factors acting downstream of BMP-2 were regulated inversely in ATDC5 cells treated with PTHrP.

In a previous article, we reported that SOX9 regulates Smpd3 expression via intronic enhancer sequences (7). Interestingly, in vitro analyses showed that the 1.9-kb Smpd3 proximal promoter without the enhancer element can also be induced when cotransfected with a Sox9 expression vector. Although there was no consensus SOX9 binding element present in the proximal promoter, there was a cluster of SRY binding sites (AACAAA), which are known to bind SOX9, albeit at a lower affinity (29). Our in vitro mutagenesis experiment showed that indeed this cluster plays an important role in Smpd3 gene expression in chondrocytes. However, it is possible that SOX9 is not the only transcription factor to bind to this cluster to regulate Smpd3 expression.

Based on our findings, we propose a scheme depicting the involvement of several factors that regulate Smpd3 expression in ATDC5 cells (Fig. 10). Previous studies have shown that BMP-2 can act through both canonical and noncanonical pathways to upregulate Runx2, which then upregulates Smpd3 expression (8, 9). We show that PTHrP can inhibit Smpd3 expression via downregulation of Sox6 and Sox9. Among different MAPKs involved in the BMP-2 signaling pathway, both ERK1/2 and JNK inhibit Smpd3 expression. On the other hand, p38 MAPK upregulates OSX, MEF2C, SOX6, and SOX9, which in turn upregulate Smpd3 expression.

FIG 10
  • Open in new tab
  • Download powerpoint
FIG 10

Model depicting the regulation of Smpd3 expression. Previous studies show that BMP-2 acts through SMAD1/5/8 and MAPK signaling pathways to upregulate RUNX2, which in turn induces Smpd3 promoter activity. Here, we show that BMP-2 can regulate Smpd3 expression through ERK1/2, JNK, and p38 kinase directly. ERK1/2 and JNK negatively regulates Smpd3 expression, while p38 induces Smpd3 expression by upregulating transcription factors MEF2C, SOX6, SOX9, and OSX. PTHrP inhibits p38 kinase activity and negatively regulates the expression of its downstream transcription factors to downregulate Smpd3 expression. Among the downstream transcription factors, SOX9 regulates Smpd3 expression via regulatory sequences present in intron 1 of the Smpd3 gene and a cluster of SRY binding sites present in the Smpd3 proximal promoter.

In conclusion, understanding the regulation of Smpd3 expression and the linked metabolic pathways may identify the mediators which can be targeted through pharmacological means to enhance skeletal tissue mineralization in cases like fracture healing. On the other hand, Smpd3 expression can be suppressed in cases such as OA, where apoptosis of hypertrophic chondrocytes is problematic. Further work is needed to understand how various transcription factors cross talk to regulate Smpd3 expression and whether there are additional signaling pathways and transcription factors involved in this process.

MATERIALS AND METHODS

Materials.Recombinant human BMP-2 was purchased from GenScript (Piscataway, NJ), and dorsomorphin (P5499), PD98059 (P215), BI-87G3 (SML0489), SB202190 (S7067), parathyroid hormone-related peptide (PTHrP; SRP4651), and forskolin (F6886) were obtained from Sigma-Aldrich (Saint Louis, MO).

Mice.The generation and genotyping of Smpd3flox/flox mice were described previously (7). The Osx-Cre mice were obtained from the Jackson Laboratory (stock number 006361). Genotypes were determined by PCR on genomic DNA isolated from tail biopsy specimens. The presence of the “floxed” allele in the gene-targeted mice was confirmed by using the primers 5′-TCAGAGAACGGAGGCTGATC-3′ and 5′-AAGAGACCGTTGGGAATACC-3′ (amplicon sizes, 106 bp [wild type] and 140 bp [“floxed” allele]). The Cre transgene was detected using the primer pair 5′-GCCTGCATTACCGGTCGATGCAACGA-3′ and 5′-GTGGCAGATGGCGCGGCAACACCCATT-3′ (amplicon size, ∼700 bp). In order to detect Cre-mediated deletion, DNA were isolated by using the phenol-chloroform method from tissue collected from the fracture callus of Smpd3flox/flox and Smpd3flox/flox; Osx-Cre mice 4 weeks postsurgery. A multiplexed PCR was performed on an equal amount of template DNA using F1 (5′-CATCATTCCCACAGCCTGAA-3′) and R1 (5′-AAGAGACCGTTGGGAATACC-3′) primers (amplicon size, ∼200 bp). The mouse Elastin gene was amplified (amplicon size, ∼500 bp) to be used as a loading control. All animal procedures were reviewed and approved by the McGill Institutional Animal Care and Use Committee following the guidelines of the Canadian Council on Animal Care.

Doxycycline treatment.Drinking water containing 1% dextrose was supplemented with 0.4 mg/ml doxycycline hyclate (Sigma, D9891) to prepare DOX water. A commercially available diet (S38888-DOX Diet; Bio-Serv) containing 200 mg/kg doxycycline was used as the DOX diet. Pregnant female mice were switched to DOX water and diet on day postcoitum (d.p.c.) 10.5, and this regimen was continued until the pups were 7 days old.

Intramedullary nailing surgical procedure.The intramedullary nailing surgery was performed on the right tibia at 2 months of age. The analyses of the fractured bones were performed 2 and 4 weeks postsurgery. The surgical procedures were done as previously described (17). Briefly, the mice were first injected subcutaneously with 0.04 ml of buprenorphine (0.3 mg diluted in 12 ml of normal saline) before the surgery. Mice were then anesthetized using isoflurane gas and oxygen and kept under anesthesia throughout the full length of the procedure. Once mice were anesthetized, they were injected with carprofen (5 μl/g) and normal saline (20 μl/g) subcutaneously. The right leg was then shaved to remove all fur surrounding the tibia. The surgical site was prepped with 2% chlorhexidine and 70% ethanol solution. A 3-mm vertical skin incision was made above the knee, and the patellar ligament was exposed where a vertical medial parapatellar incision (approximately 1 to 1.5 mm in length) was made on the tibial plateau. A 25-gauge (25G) needle was inserted through the incision into the tibial canal and through the needle, the internal wire core guide (Quincke BD 25G 3″ spinal needle, catalog no. 450170; Becton Dickinson, Franklin Lakes, NJ) was inserted. The 25G needle was then removed while keeping the internal wire core guide in the tibial canal. The protruded part of the wire core guide was clipped and bent at 90° at the proximal end and tucked in under the patellar ligament to avoid puncture of the ligament. Using the same skin incision, a mid-tibial shaft osteotomy was made using an extra fine Bonn scissors (catalog no. 14084-08; Fine Science Tools, Vancouver, Canada). The incision was sutured with a single 6-0 vicryl suture (Johnson & Johnson, Skillman, NJ), and 1 to 2 drops of analgesia, lidocaine/bupivacaine, was added to the fracture site. The skin was sutured using three to four horizontal mattress sutures. After the surgery, the mice were placed in their cages on a heated pad and allowed to move around freely. The mice were then given subcutaneous injections of 0.05 ml of buprenorphine and carprofen (5 μl/g) at 24 and 48 h postsurgery. The animals were closely monitored, and any mice showing signs of severe pain, bleeding, or swelling were euthanized. All procedures were performed under aseptic conditions in the procedure room of the animal facility at the Shriners Hospital for Children, Montreal, Quebec, Canada.

Sample collection.The mice were sacrificed using isoflurane and CO2 at 2 or 4 weeks postsurgery. The tibiae were dissected at the level of the knee and ankle. The intramedullary nail was carefully pulled out using a needle holder. Tibiae were then fixed in buffered formalin overnight and transferred to 70% ethanol for micro-CT and histological analyses.

Micro-CT.The SkyScan 1272 micro-CT machine (SkyScan, Bruker, Belgium) was used to image samples. Tibiae were harvested and imaged in plastic tubes containing 70% ethanol. The scans were performed using the SkyScan 1272 software at a source voltage of 60 kV and a source current of 166 μA. The acquisition parameters were as follows: pixel size, 5.00 μm; rotation step, 0.4; aluminum, 0.5-mm filter; and image size, 2,452 by 1,640 pixels. The generated images were used to reconstruct the three-dimensional image of the scanned bone using the NRecon Reconstruction Software (SkyScan, Bruker, Belgium) using a beam-hardening correction of 15%, a ring artifact correction at 6, smoothing at 2, and a dynamic range of 0.00 to 0.12. The reconstructed images were then rotated using DataViewer and analyzed using CTAn (both from SkyScan).

Skeletal preparation.Skeletal tissues from newborn and adult mice were fixed overnight in 95% ethanol, stained by 0.015% alcian blue dye (Sigma-Aldrich) in a 1:4 solution of glacial acetic acid and absolute ethanol for 24 h, and treated with 2% potassium hydroxide until the soft tissues were dissolved. The mineralized tissues were stained by 0.005% alizarin red (Sigma-Aldrich) solution in 1% potassium hydroxide and clarified in 1% potassium hydroxide–20% glycerol for >2 days.

Histology and histomorphometry.Fractured and nonfractured tibia were fixed overnight in 4% paraformaldehyde–phosphate-buffered saline (PBS), embedded in methyl methacrylate, sectioned (7-μm thickness), and processed for von Kossa and van Gieson (VKVG) staining. Stained bone sections were analyzed for osteoid volume/bone volume (OV/BV) using the Osteomeasure software (Osteometrics, Inc.). The region of interest (ROI) was selected within the fracture healing site. Plastic sections were also decalcified in a Cal-Ex decalcifier (Fisher, Pittsburgh, PA) overnight and processed for alcian blue and van Gieson (AB/VG) staining. AB/VG-stained bone sections were then analyzed for cartilage matrix volume/callus volume using the Osteomeasure software. Images were taken at room temperature using a light microscope (DM200; Leica) with a 20 (numerical aperture of 0.40) or 40 (numerical aperture of 0.65) objective. All histological images were captured using a camera (DP72; Olympus), acquired with DP2-BSW software (XV3.0; Olympus), and processed using Photoshop (Adobe).

Gene expression analysis.Gene expression analysis was performed using a qRT-PCR system (model 7500; Applied Biosystems). Total RNA was extracted from fractured and nonfractured tibia at 13 days postsurgery from 3-month-old WT male mice with TRIzol reagent (Ambion, catalog no. 15596018) and subjected to DNase I (Applied Biosystems, M0303S) treatment. The first-strand cDNA synthesis and qRT-PCR were performed using a high-capacity cDNA reverse transcription kit (Applied Biosystems) and SYBR green quantitative PCR master mix (Maxima; Fermentas), respectively. Relative gene expression analysis was performed as described previously (30). The primer sequences for Runx2, Sox9, Osx (Sp7), Col10a1, Smpd3, Alpl, and Col2a1 were the same as those published previously (6, 30). The remaining primer sequences are presented in Table 1.

View this table:
  • View inline
  • View popup
TABLE 1

Primer sequences

Immunofluorescence and immunohistochemistry.For immunofluorescence, E15.5 humeri were embedded in paraffin and sectioned (7 μm). Sections were blocked with 5% bovine serum albumin (BSA; Fisher, Pittsburgh, PA) in Tris-buffered saline (TBS)–Triton X-100, followed by incubation with antiosterix (anti-OSX) antibody from Abcam (Cambridge, MA). Localization of the primary antibody was detected with Alexa Fluor 488-conjugated affinity-purified donkey anti-rabbit antibody from Jackson ImmunoResearch Laboratories, Inc. (West Grove, PA). After three washes with PBS, the nuclei were visualized with Hoechst stain (H33258; Sigma-Aldrich). Images were captured by the EVOS FL cell imaging system (Life Technologies). For immunohistochemistry, plastic sections were blocked with 5% BSA (Fisher) in TBS–Triton X-100, followed by incubation with anti-SMPD3 antibody (Santa Cruz Biotechnology). The samples were further incubated with horseradish peroxidase (HRP)-conjugated goat secondary antibody (1:1,000; Abcam), and then a colorimetric reaction was carried out with 3,3-diaminobenzidine and 0.02% H2O2, followed by counterstaining with hematoxylin. Light microscopy images were captured as described above.

DNA constructs.For Smpd3 promoter studies, we used the pSmpd3-luc reporter construct that we generated previously (7). To study the role of SRY binding sites, we deleted a cluster of SRY-binding sites spanning nucleotides –595 to –646 within the 1.9-kb mouse Smpd3 proximal promoter. The modified promoter was then cloned into the SmaI and Sal I sites in the multiple cloning site of the pGL4.10 (luc2) vector (Promega, Madison, WI).

Cell culture.ATDC5 cells were cultured in alpha minimal essential medium (Invitrogen, Carlsbad, CA) supplemented with 10% fetal bovine serum (PAA Laboratories) and 100 U/ml of penicillin-streptomycin at 37°C under 5% CO2 in a humidified incubator. Cells were treated with 300 ng/ml of BMP-2 for 48 h and then subjected to gene expression analysis. To induce chondrogenic differentiation, cells were cultured in differentiation medium containing ascorbic acid (100 μg/ml) and β-glycerophosphate (5 mM) for 6 days. To inhibit BMP signaling, the cells were treated with 2 μM dorsomorphin for 30 min and washed with PBS, and differentiation medium was added to the culture. The medium was changed in this manner every 2 days and, on the sixth day, the cells were subjected to gene expression analysis.

In order to study the role of MAPK signaling, cells were treated with the indicated inhibitor for 1 h and washed with PBS, and then either inhibitor alone or inhibitor and BMP-2 were added to the medium for a total of 48 h; the cells were then subjected to gene expression analysis. HEK293 cells were cultured in Dulbecco modified Eagle medium (Invitrogen).

To investigate the effect of both BMP-2 and PTHrP on Smpd3 expression, we treated ATDC5 cells with 0.1 μM PTHrP for 24 h and then cotreated the cells with 0.1 μM PTHrP and 300 ng/ml of BMP-2 for another 48 h. The cells were then subjected to gene expression analysis.

To study transcriptional regulation of Smpd3 expression, we transfected ATDC5 cells with 2.5 μg of Mef2c, Sox6, Sox9, or Osx expression vectors using a Lipofectamine 3000 transfection kit (Invitrogen) according to the supplier’s instructions. The cells were then subjected to gene expression analysis after 48 h. The transfection efficiency was determined by transfecting ATDC5 cells with a GFP expression vector (pMax-GFP). The nuclei were visualized with Hoechst stain (Sigma-Aldrich). Images were captured by the EVOS FL cell imaging system at ×20 (Life Technologies). GFP-positive cells were quantified.

Luciferase assay.ATDC5 cells were cotransfected with 500 ng of Smpd3 reporter plasmids or the pIDWT4F-luc reporter plasmid (31) and 50 ng of Renilla luciferase (R-Luc) plasmids using the Lipofectamine 3000 transfection kit (Invitrogen). The pIDWT4F-luc reporter plasmid was used as a control for BMP-2 since the promoter contains four tandem repeats of a BMP-responsive element found in Id1, a direct target of BMP signaling. After 24 h, cells were treated with 300 ng/ml of BMP-2 for 8 or 48 h. HEK293 cells were cotransfected with 500 ng of the plasmid combinations as indicated, as well as 10 ng of R-Luc plasmid. The luciferase activity was measured after 48 h using a Dual-Luciferase reporter assay system (Promega), as described previously (30).

Statistical analysis.All results are shown as means of the standard deviation. Statistical analyses were performed by the Student t test or analysis of variance (Tukey’s multiple-comparison test).

ACKNOWLEDGMENTS

We declare no conflicts of interest.

We thank Mia Esser and Louise Marineau for animal husbandry, Jingjing Li for technical support, Corine Martineau for training to perform the intramedullary nailing surgical procedure, and the help of Martin Pellicelli and support from the FRQS Network for Oral and Bone Health Research (RSBO) for the micro-CT analyses and equipment.

This study was funded by a research grant from the Natural Sciences and Engineering Research Council of Canada to M.M. (G245390).

FOOTNOTES

    • Received 21 July 2018.
    • Returned for modification 7 August 2018.
    • Accepted 6 November 2018.
    • Accepted manuscript posted online 10 December 2018.
  • Copyright © 2019 American Society for Microbiology.

All Rights Reserved.

REFERENCES

  1. 1.↵
    1. Lefebvre V,
    2. Huang W,
    3. Harley VR,
    4. Goodfellow PN,
    5. de Crombrugghe B
    . 1997. SOX9 is a potent activator of the chondrocyte-specific enhancer of the pro α1(II) collagen gene. Mol Cell Biol 17:2336–2346.
    OpenUrlAbstract/FREE Full Text
  2. 2.↵
    1. Mori-Akiyama Y,
    2. Akiyama H,
    3. Rowitch DH,
    4. de Crombrugghe B
    . 2003. Sox9 is required for determination of the chondrogenic cell lineage in the cranial neural crest. Proc Natl Acad Sci U S A 100:9360–9365. doi:10.1073/pnas.1631288100.
    OpenUrlAbstract/FREE Full Text
  3. 3.↵
    1. Komori T
    . 2003. Requisite roles of Runx2 and Cbfb in skeletal development. J Bone Miner Metab 21:193–197. doi:10.1007/s00774-002-0408-0.
    OpenUrlCrossRefPubMedWeb of Science
  4. 4.↵
    1. Komori T,
    2. Yagi H,
    3. Nomura S,
    4. Yamaguchi A,
    5. Sasaki K,
    6. Deguchi K,
    7. Shimizu Y,
    8. Bronson RT,
    9. Gao YH,
    10. Inada M,
    11. Sato M,
    12. Okamoto R,
    13. Kitamura Y,
    14. Yoshiki S,
    15. Kishimoto T
    . 1997. Targeted disruption of Cbfa1 results in a complete lack of bone formation owing to maturational arrest of osteoblasts. Cell 89:755–764. doi:10.1016/S0092-8674(00)80258-5.
    OpenUrlCrossRefPubMedWeb of Science
  5. 5.↵
    1. Otto F,
    2. Thornell AP,
    3. Crompton T,
    4. Denzel A,
    5. Gilmour KC,
    6. Rosewell IR,
    7. Stamp GW,
    8. Beddington RS,
    9. Mundlos S,
    10. Olsen BR,
    11. Selby PB,
    12. Owen MJ
    . 1997. Cbfa1, a candidate gene for cleidocranial dysplasia syndrome, is essential for osteoblast differentiation and bone development. Cell 89:765–771.
    OpenUrlCrossRefPubMedWeb of Science
  6. 6.↵
    1. Khavandgar Z,
    2. Poirier C,
    3. Clarke CJ,
    4. Li J,
    5. Wang N,
    6. McKee MD,
    7. Hannun YA,
    8. Murshed M
    . 2011. A cell-autonomous requirement for neutral sphingomyelinase 2 in bone mineralization. J Cell Biol 194:277–289. doi:10.1083/jcb.201102051.
    OpenUrlAbstract/FREE Full Text
  7. 7.↵
    1. Li J,
    2. Manickam G,
    3. Ray S,
    4. Oh CD,
    5. Yasuda H,
    6. Moffatt P,
    7. Murshed M
    . 2016. Smpd3 expression in both chondrocytes and osteoblasts is required for normal endochondral bone development. Mol Cell Biol 36:2282–2299. doi:10.1128/MCB.01077-15.
    OpenUrlAbstract/FREE Full Text
  8. 8.↵
    1. Kakoi H,
    2. Maeda S,
    3. Shinohara N,
    4. Matsuyama K,
    5. Imamura K,
    6. Kawamura I,
    7. Nagano S,
    8. Setoguchi T,
    9. Yokouchi M,
    10. Ishidou Y,
    11. Komiya S
    . 2014. Bone morphogenic protein (BMP) signaling up-regulates neutral sphingomyelinase 2 to suppress chondrocyte maturation via the Akt protein signaling pathway as a negative feedback mechanism. J Biol Chem 289:8135–8150. doi:10.1074/jbc.M113.509331.
    OpenUrlAbstract/FREE Full Text
  9. 9.↵
    1. Chae YM,
    2. Heo SH,
    3. Kim JY,
    4. Lee JM,
    5. Ryoo HM,
    6. Cho JY
    . 2009. Upregulation of Smpd3 via BMP2 stimulation and Runx2. BMB Rep 42:86–90.
    OpenUrlCrossRefPubMed
  10. 10.↵
    1. Nishimura R,
    2. Wakabayashi M,
    3. Hata K,
    4. Matsubara T,
    5. Honma S,
    6. Wakisaka S,
    7. Kiyonari H,
    8. Shioi G,
    9. Yamaguchi A,
    10. Tsumaki N,
    11. Akiyama H,
    12. Yoneda T
    . 2012. Osterix regulates calcification and degradation of chondrogenic matrices through matrix metalloproteinase 13 (MMP13) expression in association with transcription factor Runx2 during endochondral ossification. J Biol Chem 287:33179–33190. doi:10.1074/jbc.M111.337063.
    OpenUrlAbstract/FREE Full Text
  11. 11.↵
    1. Zhou X,
    2. Zhang Z,
    3. Feng JQ,
    4. Dusevich VM,
    5. Sinha K,
    6. Zhang H,
    7. Darnay BG,
    8. de Crombrugghe B
    . 2010. Multiple functions of Osterix are required for bone growth and homeostasis in postnatal mice. Proc Natl Acad Sci U S A 107:12919–12924. doi:10.1073/pnas.0912855107.
    OpenUrlAbstract/FREE Full Text
  12. 12.↵
    1. Jing J,
    2. Hinton RJ,
    3. Jing Y,
    4. Liu Y,
    5. Zhou X,
    6. Feng JQ
    . 2014. Osterix couples chondrogenesis and osteogenesis in post-natal condylar growth. J Dent Res 93:1014–1021. doi:10.1177/0022034514549379.
    OpenUrlCrossRefPubMed
  13. 13.↵
    1. Rodda SJ,
    2. McMahon AP
    . 2006. Distinct roles for Hedgehog and canonical Wnt signaling in specification, differentiation and maintenance of osteoblast progenitors. Development 133:3231–3244. doi:10.1242/dev.02480.
    OpenUrlAbstract/FREE Full Text
  14. 14.↵
    1. Coleman RM,
    2. Aguilera L,
    3. Quinones L,
    4. Lukashova L,
    5. Poirier C,
    6. Boskey A
    . 2012. Comparison of bone tissue properties in mouse models with collagenous and non-collagenous genetic mutations using FTIRI. Bone 51:920–928. doi:10.1016/j.bone.2012.08.110.
    OpenUrlCrossRef
  15. 15.↵
    1. Marsell R,
    2. Einhorn TA
    . 2011. The biology of fracture healing. Injury 42:551–555. doi:10.1016/j.injury.2011.03.031.
    OpenUrlCrossRefPubMed
  16. 16.↵
    1. Davey RA,
    2. Clarke MV,
    3. Sastra S,
    4. Skinner JP,
    5. Chiang C,
    6. Anderson PH,
    7. Zajac JD
    . 2012. Decreased body weight in young Osterix-Cre transgenic mice results in delayed cortical bone expansion and accrual. Transgenic Res 21:885–893. doi:10.1007/s11248-011-9581-z.
    OpenUrlCrossRefPubMed
  17. 17.↵
    1. Morcos MW,
    2. Al-Jallad H,
    3. Li J,
    4. Farquharson C,
    5. Millán JL,
    6. Hamdy RC,
    7. Murshed M
    . 2018. PHOSPHO1 is essential for normal bone fracture healing. Bone Joint Res 7:397–405. doi:10.1302/2046-3758.76.BJR-2017-0140.R2.
    OpenUrlCrossRef
  18. 18.↵
    1. Li Z,
    2. Wu G,
    3. Sher RB,
    4. Khavandgar Z,
    5. Hermansson M,
    6. Cox GA,
    7. Doschak MR,
    8. Murshed M,
    9. Beier F,
    10. Vance DE
    . 2014. Choline kinase beta is required for normal endochondral bone formation. Biochim Biophys Acta 1840:2112–2122. doi:10.1016/j.bbagen.2014.03.008.
    OpenUrlCrossRef
  19. 19.↵
    1. Houston B,
    2. Stewart AJ,
    3. Farquharson C
    . 2004. PHOSPHO1—a novel phosphatase specifically expressed at sites of mineralization in bone and cartilage. Bone 34:629–637. doi:10.1016/j.bone.2003.12.023.
    OpenUrlCrossRefPubMed
  20. 20.↵
    1. Stanton LA,
    2. Sabari S,
    3. Sampaio AV,
    4. Underhill TM,
    5. Beier F
    . 2004. p38 MAP kinase signalling is required for hypertrophic chondrocyte differentiation. Biochem J 378:53–62. doi:10.1042/BJ20030874.
    OpenUrlAbstract/FREE Full Text
  21. 21.↵
    1. Stanton LA,
    2. Underhill TM,
    3. Beier F
    . 2003. MAP kinases in chondrocyte differentiation. Dev Biol 263:165–175.
    OpenUrlCrossRefPubMedWeb of Science
  22. 22.↵
    1. Thouverey C,
    2. Caverzasio J
    . 2015. Focus on the p38 MAPK signaling pathway in bone development and maintenance. Bonekey Rep 4:711. doi:10.1038/bonekey.2015.80.
    OpenUrlCrossRef
  23. 23.↵
    1. Takebe K,
    2. Nishiyama T,
    3. Hayashi S,
    4. Hashimoto S,
    5. Fujishiro T,
    6. Kanzaki N,
    7. Kawakita K,
    8. Iwasa K,
    9. Kuroda R,
    10. Kurosaka M
    . 2011. Regulation of p38 MAPK phosphorylation inhibits chondrocyte apoptosis in response to heat stress or mechanical stress. Int J Mol Med 27:329–335. doi:10.3892/ijmm.2010.588.
    OpenUrlCrossRefPubMed
  24. 24.↵
    1. Tew SR,
    2. Hardingham TE
    . 2006. Regulation of SOX9 mRNA in human articular chondrocytes involving p38 MAPK activation and mRNA stabilization. J Biol Chem 281:39471–39479. doi:10.1074/jbc.M604322200.
    OpenUrlAbstract/FREE Full Text
  25. 25.↵
    1. Tew SR,
    2. Clegg PD
    . 2011. Analysis of posttranscriptional regulation of SOX9 mRNA during in vitro chondrogenesis. Tissue Eng Part A 17:1801–1807. doi:10.1089/ten.TEA.2010.0579.
    OpenUrlCrossRefPubMed
  26. 26.↵
    1. Fernandez-Lloris R,
    2. Vinals F,
    3. Lopez-Rovira T,
    4. Harley V,
    5. Bartrons R,
    6. Rosa JL,
    7. Ventura F
    . 2003. Induction of the Sry-related factor SOX6 contributes to bone morphogenetic protein-2-induced chondroblastic differentiation of C3H10T1/2 cells. Mol Endocrinol 17:1332–1343. doi:10.1210/me.2002-0254.
    OpenUrlCrossRefPubMedWeb of Science
  27. 27.↵
    1. Arnold MA,
    2. Kim Y,
    3. Czubryt MP,
    4. Phan D,
    5. McAnally J,
    6. Qi X,
    7. Shelton JM,
    8. Richardson JA,
    9. Bassel-Duby R,
    10. Olson EN
    . 2007. MEF2C transcription factor controls chondrocyte hypertrophy and bone development. Dev Cell 12:377–389. doi:10.1016/j.devcel.2007.02.004.
    OpenUrlCrossRefPubMedWeb of Science
  28. 28.↵
    1. Zhen X,
    2. Wei L,
    3. Wu Q,
    4. Zhang Y,
    5. Chen Q
    . 2001. Mitogen-activated protein kinase p38 mediates regulation of chondrocyte differentiation by parathyroid hormone. J Biol Chem 276:4879–4885. doi:10.1074/jbc.M004990200.
    OpenUrlAbstract/FREE Full Text
  29. 29.↵
    1. Mertin S,
    2. McDowall SG,
    3. Harley VR
    . 1999. The DNA-binding specificity of SOX9 and other SOX proteins. Nucleic Acids Res 27:1359–1364.
    OpenUrlCrossRefPubMedWeb of Science
  30. 30.↵
    1. Li J,
    2. Khavandgar Z,
    3. Lin SH,
    4. Murshed M
    . 2011. Lithium chloride attenuates BMP-2 signaling and inhibits osteogenic differentiation through a novel WNT/GSK3-independent mechanism. Bone 48:321–331. doi:10.1016/j.bone.2010.09.033.
    OpenUrlCrossRefPubMed
  31. 31.↵
    1. Katagiri T,
    2. Imada M,
    3. Yanai T,
    4. Suda T,
    5. Takahashi N,
    6. Kamijo R
    . 2002. Identification of a BMP-responsive element in Id1, the gene for inhibition of myogenesis. Genes Cells 7:949–960.
    OpenUrlCrossRefPubMedWeb of Science
PreviousNext
Back to top
Download PDF
Citation Tools
Role of SMPD3 during Bone Fracture Healing and Regulation of Its Expression
Garthiga Manickam, Pierre Moffatt, Monzur Murshed
Molecular and Cellular Biology Feb 2019, 39 (4) e00370-18; DOI: 10.1128/MCB.00370-18

Citation Manager Formats

  • BibTeX
  • Bookends
  • EasyBib
  • EndNote (tagged)
  • EndNote 8 (xml)
  • Medlars
  • Mendeley
  • Papers
  • RefWorks Tagged
  • Ref Manager
  • RIS
  • Zotero
Print

Alerts
Sign In to Email Alerts with your Email Address
Email

Thank you for sharing this Molecular and Cellular Biology article.

NOTE: We request your email address only to inform the recipient that it was you who recommended this article, and that it is not junk mail. We do not retain these email addresses.

Enter multiple addresses on separate lines or separate them with commas.
Role of SMPD3 during Bone Fracture Healing and Regulation of Its Expression
(Your Name) has forwarded a page to you from Molecular and Cellular Biology
(Your Name) thought you would be interested in this article in Molecular and Cellular Biology.
CAPTCHA
This question is for testing whether or not you are a human visitor and to prevent automated spam submissions.
Share
Role of SMPD3 during Bone Fracture Healing and Regulation of Its Expression
Garthiga Manickam, Pierre Moffatt, Monzur Murshed
Molecular and Cellular Biology Feb 2019, 39 (4) e00370-18; DOI: 10.1128/MCB.00370-18
del.icio.us logo Digg logo Reddit logo Twitter logo CiteULike logo Facebook logo Google logo Mendeley logo
  • Top
  • Article
    • ABSTRACT
    • INTRODUCTION
    • RESULTS
    • DISCUSSION
    • MATERIALS AND METHODS
    • ACKNOWLEDGMENTS
    • FOOTNOTES
    • REFERENCES
  • Figures & Data
  • Info & Metrics
  • PDF

KEYWORDS

BMP-2
chondrocytes
fracture
mineralization
PTHrP
SMPD3
Sox9
p38 MAPK

Related Articles

Cited By...

About

  • About MCB
  • Editor in Chief
  • Editorial Board
  • Policies
  • For Reviewers
  • For the Media
  • For Librarians
  • For Advertisers
  • Alerts
  • RSS
  • FAQ
  • Permissions
  • Journal Announcements

Authors

  • ASM Author Center
  • Submit a Manuscript
  • Article Types
  • Ethics
  • Contact Us

Follow #MCBJournal

@ASMicrobiology

       

ASM Journals

ASM journals are the most prominent publications in the field, delivering up-to-date and authoritative coverage of both basic and clinical microbiology.

About ASM | Contact Us | Press Room

 

ASM is a member of

Scientific Society Publisher Alliance

 

American Society for Microbiology
1752 N St. NW
Washington, DC 20036
Phone: (202) 737-3600

Copyright © 2021 American Society for Microbiology | Privacy Policy | Website feedback

Print ISSN: 0270-7306; Online ISSN: 1098-5549