Deficiency of the intermediate filament synemin reduces bone mass in vivo

Abstract

While the type IV intermediate filament protein, synemin, has been shown to play a role in striated muscle and neuronal tissue, its presence and function have not been described in skeletal tissue. Here, we report that genetic ablation of synemin in 14-wk-old male mice results in osteopenia that includes a more than 2-fold reduction in the trabecular bone fraction in the distal femur and a reduction in the cross-sectional area at the femoral middiaphysis due to an attendant reduction in both the periosteal and endosteal perimeter. Analysis of serum markers of bone formation and static histomorphometry revealed a statistically significant defect in osteoblast activity and osteoblast number in vivo. Interestingly, primary osteoblasts isolated from synemin-null mice demonstrate markedly enhanced osteogenic capacity with a concomitant reduction in cyclin D1 mRNA expression, which may explain the loss of osteoblast number observed in vivo. In total, these data suggest an important, previously unknown role for synemin in bone physiology.

intermediate filaments (IFs) are structural proteins of the nuclear membrane and the cytosol that determine cellular architecture and cytoplasmic integrity (6, 14, 21, 26). IFs also serve as a scaffold for enzymes involved in intracellular signaling (16, 41) and in directing the assembly of microtubules (9), thereby modulating a range of cellular activities, including motility (16, 20, 41). More than 65 different genes code for IF proteins (22) of 6 different classes (10). These include the keratins (types I and II); desmin, vimentin and glial fibrillar acidic protein (type III); neurofilament proteins, internexin and synemin (type IV); the nuclear lamins (type V); and other, tissue-specific filament proteins, such as phakinin, filensin, and nexin (type VI). When purified, most of these proteins form homopolymers (e.g., vimentin, desmin) or stoichiometric heteropolymers (e.g., types I and II) (39).

The type IV intermediate filament protein, synemin, does not homopolymerize (4) or form 1:1 heteropolymers with other filament subunits. Rather, it coassembles into desmin or vimentin filaments (17, 53) and can associate with keratin filaments (23). Synemin is encoded by a single gene (Synm) and is primarily expressed as either α or β isoforms, with molecular masses of ∼220 and 180 kDa, respectively. Unlike other intermediate filament proteins, synemin is an A-kinase anchoring protein, or AKAP (44, 45), which allows it to modulate signaling cascades, including the phosphorylation of nearby proteins such as desmin (53).

We and others have been studying the roles of intermediate filaments in striated muscle. Like striated muscle, bone is of mesenchymal origin and expresses keratins and vimentin (27, 28, 49, 54), the type III intermediate filament protein that in muscle is replaced by desmin as development proceeds (12, 50). Although the literature contains a number of reports on the effects on skeletal muscle of the elimination of one or more intermediate filaments, including synemin, by homologous recombination (2, 3, 13, 15, 29, 31, 34, 38, 40, 43, 46–48, 51, 52), it is mute regarding the effects of these “knock-outs” on bone. Indeed, to the best of our knowledge, the expression of synemin in bone cells has never been reported. Here we report¹ that mice lacking synemin (Synm−/−, Synm-null) are osteopenic, with a more than 50% reduction in trabecular bone density, associated with a decrease in trabecular thickness and an increase in trabecular spacing in the distal femur. In addition, they show a subtle cortical phenotype, with a reduction in the periosteal and endosteal perimeter of the femoral diaphysis. Our studies of cells in vivo and in vitro and of the serum markers of bone turnover indicate that these defects in Synm−/− bone tissue are likely due to changes in osteoblasts. Small increases in osteoclast activity may also occur.

METHODS

Animal care.

Synemin-null mice were generated on the C57Bl/6 background by homologous recombination [Synm^{tm1.1(KOMP)Vlcg}; MGI: 2661187; The International Mouse Phenotyping Consortium] and genotyped as described (13). We used male, age-matched C57Bl/6 control (WT) and Synm-null mice from 13–14 wk of age for the long bone extracts and microcomputed tomography (microCT) studies, and 4 wk of age for the primary osteoblast culture experiments. Our animal protocols were approved by the Institutional Animal Care and Use Committee of the University of Maryland School of Medicine.

X-ray imaging.

Lateral and AP radiographs (35 kV, 10 s) were taken of the mice, post euthanasia, with a Faxitron digital X-ray system, as described (18).

MicroCT.

Femurs were dissected from 14-wk-old male WT and Synm^−/− mice, fixed in 4% paraformaldehyde for up to 4 days and then transferred to 70% ethanol. Three-dimensional microCT was performed on the femurs (n = 7/genotype) for gross morphological assessment using a Bruker SkyScan 1172 microCT scanner. Bone morphology and microarchitecture were assessed at the distal femoral metaphysis for trabecular parameters [trabecular bone volume/tissue volume (BV/TV), trabecular number (Tb.N), trabecular thickness (Tb.Th.), and trabecular separation (TbSp)] and the middiaphysis for cortical parameters [periosteal perimeter (Ps.Pm), endocortical perimeter (Ec.Pm), cortical cross sectional thickness (Cs.Th) and mean polar moment of inertia (MMI)]. The skeletal parameters assessed by microCT followed the nomenclature guidelines as outlined by Bouxsein et al. (5). Femurs were scanned with 2K resolution, 10-μm voxel size, 0.5 Al filter at 60 kV and 167 μA. Trabecular bone was delineated manually, in a region of interest 0.2 mm to 2.0 mm proximal to the distal femoral growth plate. For cortical bone parameters, analysis was performed at the femoral diaphysis beginning at 56% of the femoral length (measured from the head of the femur) extending 0.5 mm distally.

ELISA.

Serum was collected from animals at the time of euthanasia. Serum COOH-terminal telopeptide of type 1 collagen (CTX) and procollagen type 1 NH₂-terminal propeptide (P1NP) levels were quantitated using IDS assay for P1NP production and RatLaps for CTX production according to the manufacturer's specifications [Immunodiagnostic Systems (IDS) Gaithersburg, MD].

Histology and bone histomorphometry.

Fixed tissue was decalcified in 14% EDTA (pH 8.0). Subsequently, the tissue was embedded in paraffin, and 5-μm serial, longitudinal sections were prepared. For osteoclasts, sections were stained for tartrate-resistant acid phosphate (TRAP) using a leukocyte acid phosphatases kit (Sigma), as described (35). For osteoblasts, sections were stained with Goldner's trichrome. Osteoblasts and osteoclasts adjacent to the bone surface in a fixed region of interest within the secondary spongiosa were quantified using the Bioquant osteomeasure histology software, as described (7, 8).

Primary osteoblast isolation and cell culture.

Primary osteoblasts were isolated as previously described (1, 7). Briefly, femurs and tibiae were dissected from mice of each genotype, and bone marrow was flushed with sterile saline after removing the epiphyses. The remaining diaphyseal bones were cut into chips and incubated with collagenase solution for 2 h at 37°C, washed with αMEM containing 10% fetal bovine serum, penicillin (50 IU/ml), streptomycin (50 μg/ml), and gentamycin (50 μg/ml), and plated to allow cells to migrate from the bone chips. After 3 days, cells were treated with mineralization medium, which contains 50 ng/ml ascorbic acid-2-phosphate and 10 mM glycerol-2-phosphate. Cells were maintained by changing the medium three times per week. After 14 days of culture under mineralizing conditions, cells were lysed for RNA isolation and quantitative RT-PCR or stained for mineralization with Alizarin Red S. MC3T3 cells were cultured as described (30).

Immunofluorescence and Western blotting.

Primary osteoblasts were fixed with cold 4% paraformaldehyde for 15 min, permeabilized with 0.25% Triton X-100 for 10 min, and incubated 1 h in Superblock (Thermo Scientific, Waltham, MA). Samples were then incubated for 1.5 h in affinity-purified anti-synemin (diluted 1:25 in blocking solution). The generation and validation of this antibody has been described (13). Subsequently, the samples were incubated for 1 h in goat anti-rabbit IgG-Cy3 and phalloidin 488 (1:100; Chemicon International, Temecula, CA), rinsed, and mounted. Samples were observed with a LSM 510 confocal laser scanning microscope (Carl Zeiss, Oberkochen, Germany). For tissue extracts, tibiae were dissected from both genotypes, cleaned of soft tissue, the marrow cavity flushed with sterile saline, and lysed in a modified RIPA buffer [50 mM Tris (pH 8.0), 150 mM NaCl, 1.0% NP-40, 0.5% sodium deoxycholate, 0.1% SDS, 10 mM Na₄P₂O₇, 10 mM β-glycero-PO₄, 10 mM NaF, 10 mM EDTA, 1 mM EGTA, 1X HALT phosphatase/protease inhibitor]. Samples were homogenized using 1-mm-diameter stainless steel beads and a Qiagen tissue lyser LT at 50 MHz for up to 10 min. Samples were pelleted, and the supernatant was kept for protein expression analysis by Western blotting. MC3T3 whole cell extracts were prepared as described (36).

Quantitative real time RT-PCR.

RNA was isolated using TRIzol, reverse transcribed, and used for quantitative PCR as described (19). Genes were simultaneously normalized to Gapdh, Rpl13, and Hprt (19, 37). The synemin primers were 5′-GGC TGA GGA TCA GGC ACT CT-3′ and 5′-GTG GAC CAG TTC AGT TCA TCT AGC T-3′. These primers do not discriminate isoforms.

Alizarin Red S staining.

Confluent cultures of primary osteoblasts, maintained for 14 days in mineralizing media, were rinsed in HBSS and fixed in 10% neutral buffered formalin prior to staining mineralized nodules with 1 mg/ml Alizarin Red S (pH 4.2) solution, as described (33).

Statistics.

Data were compared between genotypes by a two-tailed t-test for unpaired samples. A P value < 0.05 indicated statistical significance.

RESULTS

At 14 wk of age, wild-type (Synm+/+) and knockout (Synm−/−) mice were virtually indistinguishable except for a slight decrease in body weight (Fig. 1). However, despite gross morphologic similarity, 14-wk-old male Synm−/− mice are severely osteopenic, with a more than twofold decrease in trabecular bone mass (BV/TV) relative to wild-type controls, as determined by microCT of the distal femur (Fig. 2A). This loss of trabecular bone fraction was due to changes in bone microarchitecture, which included decreased trabecular number, decreased trabecular thickness, and a concomitant increase in trabecular separation relative to wild-type control mice. Likewise, a subtle cortical bone phenotype was observed at the femoral middiaphysis, including a reduced cross-sectional area of the diaphysis due to an attendant reduction in both the periosteal and endosteal perimeter (Fig. 2B). This diaphyseal narrowing results in a 30% decrease in calculated mean polar moment of inertia in the Synm−/− mice relative to controls.

Fig. 1.

Synemin-null mice appear grossly normal relative to control mice. In comparison to age- and sex-matched wild-type (+/+) control mice, 14 wk old, male synemin-null (−/−) mice appear grossly normal with no overt skeletal defects observed by digital X-ray. While there is a slight decrease in body weight in Synm−/− mice relative to controls, there is no significant difference in body length (nose to tip of the tail) or tibial length. n = 7/genotype. Bar graphs display means ± SD. *P < 0.05; n.s., no significant difference.

Fig. 2.

Synemin-null mice have a trabecular and cortical bone phenotype. A: longitudinal digital X-ray of the femur. The approximate region of interest used for microCT quantification of trabecular (Tb.ROI) and cortical (Ct.ROI) bone is shown. Representative 3-dimensional models of the reconstructed microCT data for 14-wk-old, male wild-type (+/+) control mice and age- and sex-matched synemin-null (−/−) mice are shown for trabecular bone at the metaphysis of the distal femur (B) or cortical bone at the femoral middiaphysis (C). Scale bars, 0.4 mm. Parameters of bone microarchitecture are shown as measured by microCT of the distal femur of 14-wk-old, male wild-type (+/+) control mice and age- and sex-matched synemin-null (−/−) mice. Trabecular (B) and cortical (C) parameters are shown (n = 7/genotype). Bar graphs display means ± SD. *P < 0.05; n.s., no significant difference.

To understand the underlying mechanism driving the observed osteopenia, we examined serum markers of bone turnover using commercial ELISAs for bone formation (type I collagen N-propeptide, P1NP) and bone resorption (type I collagen C-telopeptide, CTX) (Fig. 3A). Genetic ablation of synemin resulted in a nearly twofold reduction in P1NP levels, suggestive of reduced bone formation. In contrast, serum CTX levels, a marker of osteoclastic bone resorption, were statistically unchanged in Synm−/− mice relative to wild-type controls. Static bone histomorphometric analysis of the skeletal cellularity in the trabecular compartment revealed a nearly twofold reduction in osteoblast number in Synm−/− mice (Fig. 3, B and C). The osteoclast surface/bone surface trended toward an increase in Synm−/− mice but did not reach statistical significance (Fig. 3, B and C).

Fig. 3.

Synemin-null mice exhibit reduced serum bone formation markers and reduced osteoblast number in vivo. A: serum levels of markers of bone formation (P1NP) and bone resorption (CTX) were determined in 14-wk-old, male wild-type (+/+) control mice and age- and sex-matched synemin-null (−/−) mice (n = 7/genotype). B: ratios of osteoblast surface/bone surface (OB.S/BS) and osteoclast surface/bone surface (OC.S/BS) were determined in serial sections of Goldner's trichrome- or TRAP-stained trabecular bone at the distal femur, respectively (n = 3/genotype). C: representative images of stained sections from wild-type and Synm−/− mice are shown. Scale bars, 0.4 mm. Insets show TRAP staining for osteoclasts (red arrowheads) or Goldner's trichrome staining revealing osteoblasts (yellow arrowheads) present on trabecular bone from serial sections. Bar graphs display means ± SD. *P < 0.05; n.s., no significant difference.

Given the apparent alteration in osteoblast number and activity in vivo, we determined if there was a cell autonomous defect in osteoblast differentiation using primary, long bone-derived osteoblasts from wild-type and Synm−/− mice. Immunofluorescence microscopy of cultured primary osteoblasts revealed the expression of synemin protein in wild-type, but not Synm−/− cells (Fig. 4A). In wild-type cells, synemin staining appeared concentrated in small, roughly circular spots that were not regularly organized with respect to actin stress fibers. These structures, perhaps podosomes, were unlabeled in Synm−/− cells. Quantitative real-time RT-PCR confirmed expression of Synm mRNA in wild-type cells and its absence in Synm−/− osteoblasts (Fig. 4B). Western blotting detected a ∼180-kDa band corresponding to synemin protein in tissue extracts from the tibia (flushed of marrow) of wild-type mice and in a MC3T3 mouse osteoblast cell line, confirming its expression in the osteoblast lineage (Fig. 4C).

Fig. 4.

Synemin-null mice show a cell autonomous defect in osteoblast differentiation and proliferation. A: immunofluorescence of primary osteoblasts from 4-wk-old wild type (+/+) or synemin-null (−/−) mice stained with phalloidin (actin) and antibodies to synemin. Scale bar, 10 μm. B and D: quantitative real-time PCR cDNA isolated from 4-wk-old primary wild-type (+/+) or synemin-null (−/−) mice. Bar graphs display means ± SD. n = 3/genotype. *P < 0.05. C: Western blot from tibial extracts or whole cell extracts from cultured MC3T3 cells probed with anti-synemin antibodies. E: Alizarin Red S staining of mineralization by primary osteoblasts from mice of the indicated genotype. Triplicate wells of a 96-well plate are shown for each genotype.

Examination of osteoblast differentiation markers in osteoblasts cultured from wild-type and Synm−/− mice revealed a paradoxical increase in osteoblast genes, including Runx2, bone sialoprotein, and osteocalcin (Fig. 4D). Interestingly, mRNA levels of the proliferative marker, cyclin D1, was significantly reduced in Synm−/− osteoblasts (Fig. 4D). This finding is consistent with the decreased osteoblast number observed in vivo. Mineralization was elevated in Synm−/− cultures relative to wild-type controls (Fig. 4E). In total, these ex vivo data suggest that osteoblast proliferation may be impaired in Synm−/− osteoblasts, while osteogenic differentiation is elevated.

DISCUSSION

Previous studies of the effects of synemin deletion have shown modest, but significant, changes in skeletal muscle (13, 29). Here, we examine the consequence of the absence of synemin on bone, where the effects are much more pronounced. Long bones lacking synemin are osteopenic with a profound loss of trabecular microarchitecture and changes in cortical bone geometry. These major changes suggest that synemin plays a direct role in bone homeostasis, as such a striking change in bone would not be predicted by the mild changes in skeletal muscle in Synm−/− mice.

The skeletal phenotype detected in these mice points to a cell autonomous defect in osteoblasts. We observed reduced osteoblast number and reduced levels of a serum marker of bone formation (P1NP) in the Synm−/− mice. Paradoxically, markers of differentiation and the mineralizing capacity of osteoblasts isolated from Synm−/− mice were markedly enhanced. As the bone mass is reduced, it seems likely that progenitor cell proliferation is defective, leading to more rapid differentiation and less expansion. In support of this notion, cyclin D1 mRNA levels are reduced in Synm−/− osteoblasts in vitro, and osteoblast number is reduced in vivo. A nonmutually exclusive possibility is that increased osteoclast-mediated bone resorption may also contribute to the osteopenic phenotype. While we did not find a statistically significant change in circulating CTX or osteoclast number in vivo, both values were slightly elevated in Synm−/− mice. Indeed, it is possible that an elevation of bone resorption preceded the time point that we examined.

The molecular mechanism by which synemin affects bone cell function is unclear. Our immunofluorescence staining in primary osteoblasts shows punctate cytoplasmic staining. This staining pattern seems unusual for an IF, but a similar pattern has been reported for synemin in transfected SW13-cl2 adrenal cortex carcinoma cells (24). Further, the validity that the antibody recognizes synemin is reinforced by the absence of staining in the Synm−/− mice. It remains unclear if synemin is coassembled into intermediate filaments at sites of cell-substrate adhesion in osteoblasts, or if it concentrates at those sites due to its interactions with other proteins. In addition to its role as an intermediate filament, synemin can also function as an A-kinase anchoring protein (AKAP) (44). AKAPs are signaling scaffolds that are involved in the spatiotemporal control of signaling pathways, including cAMP-dependent signaling (11, 56). Protein kinase A and cAMP-dependent signaling pathways are fundamental to the osteoblast and osteoclast lineages and the coordination of bone remodeling (25, 32, 42, 55). Accordingly, it is possible that synemin's function in bone is facilitated by its role as an AKAP rather than as an intermediate filament. In any case, the skeletal phenotype of Synm−/− mice raises intriguing questions about the role of synemin in the biology of osteoblasts that merit further study.

Ours is the first report of a role for synemin in bone, where it plays a cell autonomous role in osteoblast differentiation. Future studies are needed to systemically examine its role throughout skeletal growth and maturation, as well as more thoroughly examine the effects on both osteoblast and osteoclast lineages.

GRANTS

This work was supported by Grants R01-AR-063631 (J. P. Stains) and R01-AR-055928 (R. J. Bloch) from the National Institutes of Health; F31-AR-064673 (A. M. Buo) from the National Institute of Arthritis and Musculoskeletal and Skin Diseases; and from a Physiological Genomics Fellowship from the American Physiological Society (K. P. Garcia-Pelagio) and from CONACYT (K. P. Garcia-Pelagio).

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the author(s).

AUTHOR CONTRIBUTIONS

M.C.M., A.M.B., and K.P.G.-P. performed experiments; M.C.M., A.M.B., K.P.G.-P., J.P.S., and R.J.B. analyzed data; M.C.M., A.M.B., K.P.G.-P., J.P.S., and R.J.B. interpreted results of experiments; M.C.M., A.M.B., K.P.G.-P., and J.P.S. prepared figures; M.C.M., J.P.S., and R.J.B. drafted manuscript; M.C.M., A.M.B., K.P.G.-P., J.P.S., and R.J.B. edited and revised manuscript; M.C.M., A.M.B., K.P.G.-P., J.P.S., and R.J.B. approved final version of manuscript; J.P.S. and R.J.B. conception and design of research.