Thyroid-stimulating hormone induces a Wnt-dependent, feed-forward loop for osteoblastogenesis in embryonic stem cell cultures

Abstract

We have shown that the anterior pituitary hormone, thyroid-stimulating hormone (TSH), can bypass the thyroid to exert a direct protective effect on the skeleton. Thus, we have suggested that a low TSH level may contribute to the bone loss of hyperthyroidism that has been attributed traditionally to high thyroid hormone levels. Earlier mouse genetic, cell-based, and clinical studies together have established that TSH inhibits osteoclastic bone resorption. However, the direct influence of TSH on the osteoblast has remained unclear. Here, we have used a model system developed from murine ES cells, induced to form mature mineralizing osteoblasts, and show that TSH stimulates osteoblast differentiation primarily through the activation of protein kinase Cδ and the up-regulation of the noncanonical Wnt components frizzled and Wnt5a. We predict that a TSH-induced, fast-forward short loop in bone marrow permits Wnt5a production, which, in addition to enhancing osteoblast differentiation, also stimulates osteoprotegerin secretion to attenuate bone resorption by neighboring osteoclasts. We surmise that this loop should uncouple bone formation from bone resorption with a net increase in bone mass, which is what has been observed upon injecting TSH.

Until recently, thyroid-stimulating hormone (TSH) was thought solely to regulate thyroid follicle development and thyroid hormone secretion (1). Although radio-ligand binding and mRNA studies had suggested that TSH receptors (TSHRs) were expressed more ubiquitously (2, 3), it was only in 2003 that we definitively established that TSHRs were localized to bone cells and that absent TSHR signaling caused high-turnover bone loss (4). Subsequent rodent and clinical studies have since pointed to a role for TSHRs in normal physiology, as well as in the pathophysiology of bone loss in the hyperthyroid state (5–9). A recent study, in which a woman with isolated TSH deficiency developed not only myxedema coma, but also severe osteoporosis, is more evidence for a fundamental role for TSH in skeletal homeostasis (10). Less compelling, but nonetheless important, is that individuals with activating TSHR polymorphisms but with normal thyroid function have a higher bone mass than matched controls (11).

TSH inhibits bone resorption directly by acting on osteoclastic TSHRs (4, 12), as well as indirectly by suppressing the production of the osteoclastogenic cytokine tumor necrosis factor-α (TNF-α) from macrophages (13). However, when administered in vivo, TSH not only prevents bone loss via an antiresorptive action (14), but also stimulates bone formation in certain rodent models, such as aged rats, to restore the lost bone even 28 wk post ovariectomy (14, 15). These studies have led to the speculation that, in addition to its osteoclast-inhibitory actions seen in mice, rats, and humans, TSH may also stimulate bone formation through a direct action on the osteoblast, which we have shown possesses TSHRs in abundance (4).

We know that children with congenital hypothyroidism have runted skeletons. Although this skeletal defect has been attributed solely to low thyroid hormone levels, the emerging function of TSH in skeletal physiology begs the question whether TSH also plays a role in skeletal morphogenesis and bone growth. In other words, can a high TSH cause premature ossification in growing children, or at least contribute to it? Hence, our use of an ES cell model, which we have found expresses TSHRs (16). That a glycoprotein hormone receptor, such as the TSHR, should appear so early in development is a testament to its general importance in embryology.

Hence, the studies described here serve two distinct, but related purposes. First, we describe an experimental system in which TSHR-positive ES cells can be induced to form mature, mineralizing osteoblasts. Second, and of equal importance, is that we show clearly that TSH can functionally stimulate the osteoblast differentiation in these ES cell cultures. These data suggest that TSH could indeed be a hormonal (or local) regulator of bone accretion during skeletal morphogenesis and growth.

Results

We first studied the effect of incubating murine ES cells with osteogenic differentiation factors on the formation of osteoblast colonies during a 30-d culture. We found a significant decline in the expression of the stem cell renewal genes Rex-1, Oct-4, and Sox-2 (Fig. 1A); this suggested that ES cells had begun to lose their “stemness.” In parallel, the cultures began to display alkaline phosphatase (ALP)-positive colonies, colony-forming units-fibroblastoids (CFU-fs) (Fig. 1B). PCR revealed the expression of both early and late osteoblast differentiation genes, namely ALP, type 1 collagen, and osteocalcin (Fig. 1C). In separate experiments, mature mineralized colonies, colony forming units-osteoblasts (CFU-obs), were seen with von Kossa (Fig. 3A) and Alizarin red staining (Fig. 3B). Mineralization was confirmed by increased calcium and phosphorous content of the colonies, respectively (Fig. 3 C and D).

Fig. 1.

Murine ES cell cultures can be induced to form osteoblasts. ES cell culture in differentiation medium (DF) (Materials and Methods) resulted in (A) a marked reduction in the expression of the stem cell marker genes Rex-1, Sox-2, and Oct-4 at day 30 compared with day 2 (by qPCR). *P < 0.005, **P < 0.0001, ***P < 0.0004. (B) The appearance of alkaline phosphatase (ALP)-positive colonies and CFU-fs. (C) The expression of osteoblast genes, namely ALP, type 1 collagen, osteocalcin (OC), and osteoprotegerin (OPG). Data are representative of two separate experiments.

Fig. 3.

TSH stimulates osteoblastogenesis and mineralization. (A and B) Von Kossa-positive and Alizarin-positive colony formation in ES cell cultures incubated in differentiating medium (DF) for 30 d (Materials and Methods). (a) Untreated and (b) enhanced mineralization with differentiation and (c) influence of TSH (1 mU/mL). TSH failed to induce differentiation of ES cells in the absence of DF (not illustrated). (C and D) Quantity of calcium and phosphate in the colonies was enhanced with DF and TSH (1 mU/mL). (E) DF and TSH (0, 1, 5, and/or 10 mU/mL) enhanced the expression of (a) collagen type-1, (b) alkaline phosphatase (AP), and (c) osteocalcin (OC) at day 30 as assessed by qPCR.

We next studied whether the TSHR was expressed in these cultures, and if so, whether its activation affected osteoblast differentiation. Fig. 2A shows TSHR mRNA in ES cells throughout the 30-d cultures. The addition of TSH appeared to dampen TSHR expression. TSHR protein expression was confirmed by flow cytometry using the anti-TSHR monoclonal antibody M1 directed to residues 381–385 (Fig. 2B). Seventy-three percent of cells displayed TSHR expression.

Fig. 2.

TSH receptors (TSHRs) are expressed in differentiating murine ES cell cultures. RT-PCR (A) and flow cytometry (B) were performed on ES cell cultures after 30 d of incubation in differentiation medium (DF) (Materials and Methods). (A) DF stimulated, with or without TSH (10 mU/mL) and TSHR mRNA expression in long-term cultures. FRTL5 cells treated with 5 hormone was used as the positive control and GAPDH as the loading control. (B) Approximately 73% of cells were TSHR-positive when stained with anti-TSHR antibody (M1) (blue) compared with an isotype IgG control (red) (Materials and Methods).

Activation of the TSHR in ES cells enhanced osteoblast differentiation markedly. Von Kossa- and Alizarin red-positive CFU-obs increased significantly in the presence of TSH compared with control cultures containing differentiation medium only (Fig. 3 A and B). Enhanced mineralization was confirmed by calcium and phosphate measurements (Fig. 3 C and D). Importantly, however, TSH triggered a dramatic, up to 200-fold, concentration-dependent increase in type 1 collagen. It also enhanced the expression of ALP and osteocalcin (Fig. 3E). The data together demonstrate that TSH stimulates osteoblastogenesis by acting on the TSHR in differentiating ES cell cultures. This pro-osteoblastic action of TSH is consistent with studies from our group and others, wherein TSH injections are anabolic in vivo. Notably, injected TSH restores the lost bone post ovariectomy and enhances bone formation in calcein-labeling studies (14, 15).

To probe the mechanism of TSH-induced osteoblastogenesis, we studied the expression of molecules involved in Wnt signaling, a dominant pathway regulating bone formation and bone mass. Fig. 4A shows a small enhancement in the expression of the canonical Wnt receptor LRP5 upon differentiation, but this was attenuated by TSH, confirming previous results (4). LRP6 remained relatively unaffected by differentiation or by TSH (Fig. 4B). The target canonical Wnt transducer β-catenin likewise remained relatively unaltered upon differentiation and by TSH (Fig. 4C). In contrast, the noncanonical coreceptor frizzled (Frz) was up-regulated dramatically by TSH (Fig. 4D), in a concentration-dependent manner (data not shown). Likewise, quantitative PCR and Western blotting showed a strong induction by TSH of the noncanonical agonist Wnt5A (Fig. 4 E and F). The data demonstrated that the noncanonical arm of the Wnt signaling pathway, consisting of Frz and Wnt5a, was the primary transducer of TSH effects on osteoblastogenesis. Consistent with this pathway, the production of osteoprotegerin (OPG) was also enhanced as a function of osteoblast differentiation and with TSH by ∼50-fold (Fig. 4F). This increase was confirmed on Western blotting (Fig. 4G).

Fig. 4.

TSH stimulates noncannonical Wnt signaling. The effects of differentiation media (DF) and TSH (0, 5, or 10 mU/mL) on components of the Wnt-signaling pathway (assessed by real time PCR), namely Lrp5 (A), Lrp6 (B), β-catenin (C), Frizzled (Frz) (D), Wnt5a (E), and osteoprotegerin (OPG) (F) in 30-d ES cell cultures. *P < 0.05 by ANOVA one-way analysis. (G) Densitometric values from Western blots showing the up-regulation of Wnt5a but not of β-catenin in the cell lysates of cultures treated with DF+/TSH+. β-Actin is shown as the loading control.

Finally, we sought to explore proximal pathways: notably, whether the induction of osteoblastogenesis by TSH was mediated by PKA or PKCδ, both of which are known to be downstream of TSHR activation, but only one of which, namely PKCδ, has been implicated in mediating osteoblastogenesis (17). The Western blot in Fig. 5A shows that PKCδ phosphorylation was markedly enhanced upon ES cell differentiation and with TSH, whereas PKA phosphorylation was not. To assess a possible function for PKCδ in TSH action, we studied whether the known PKCδ inhibitor rotterlin could inhibit TSH-induced osteoblastogenesis. Thus, whereas TSH stimulated expression of the differentiation genes osteopontin and type 1 collagen, rotterlin inhibited this response (Fig. 5 B and C). This established that PKCδ was a dominant downstream mediator of TSH-induced osteoblastogenesis.

Fig. 5.

TSH stimulates protein kinase Cδ phosphorylation. (A) Differentiation medium (DF) (Materials and Methods) with and without TSH (0 and 1 mU/mL) triggered the phosphorylation of protein kinase Cδ (PKC-δ), but not of protein kinase A (PKA) as shown in this Western blot. β-Actin was the loading control. n = 2. (B and C) PKC-δ inhibitor, rotterlin (10 μM), inhibited the increase in type1 collagen and osteopontin (OPN) expression triggered by TSH (1 mU/mL) as assessed by qPCR.

Discussion

These results underscore important concepts relating to osteoblast differentiation from pluripotent ES cells. First, we find that mature, mineralizing osteoblasts can indeed form from ES cells without the intermediacy of embyroid bodies, which is consistent with previous reports (16). This pattern is also consistent with the reduced expression of stem cell renewal genes and with the enhanced expression of osteoblast marker genes. Second, the data establish that TSH stimulates osteoblastic differentiation from ES cells. This TSH-induced osteoblastogenesis is concordant with, and might in fact explain, the anabolic action of TSH in vivo. Third, we show that TSH potently stimulates the production of OPG from ES cell-derived osteoblasts. This observation is supported by data from primary cultures, wherein mature osteoblasts synthesize OPG and immature cells produce mainly RANKL (18).

At the mechanistic level, we show that the action of TSH is exerted primarily through the noncanonical Wnt pathway downstream of PKCδ. Wnt signaling is known to increase bone mass using two separate mechanisms: directly, by enhancing osteoblastogenesis and bone formation and, indirectly, by stimulating the production of OPG from osteoblasts, which, in turn, inhibits RANK-l-induced osteoclastogenesis and bone resorption (19). Of note is that ES cell-derived osteoblasts produce abundant OPG that is further enhanced by TSH. This suggests that OPG may, at least in part, be responsible for the indirect action of TSH in inhibiting bone resorption and increasing bone mass. That said, TSH also inhibits osteoclasts directly and, in addition, suppresses the production of TNFα, a proresorptive cytokine (4, 13).

The 19 secreted Wnts bind to 1 of 10 Frz receptors and one of two LRPs (20). Of the four known intracellular signaling pathways, the best characterized is the canonical Wnt/β-catenin pathway that signals through LRP5/6, GSK-3β, and β-catenin to ultimately activate the nuclear transcription complex lymphoid enhancement factor/T cell factor (20). Gain-of-function mutations of this pathway increase osteoblastogenesis, bone formation, and bone mass (21). Loss-of-function mutations cause diseases characterized by bone loss, such as the osteoporosis-pseudoglioma syndrome (22). Our results show modest increases in LRP5, and no increases in LRP6 or β-catenin expression. LRP5 levels dropped in response to TSH, which might suggest a modest inhibitory effect of TSH on canonical signaling. This is not unexpected, however, as (i) TSH decreased osteoblastogenesis in primary osteoblast cultures from adult mice, and (ii) LRP5 expression was elevated in osteoblasts from TSHR^−/− mice (4).

In contrast to the canonical pathway, mutations of the molecular components of the noncanonical pathway, abundantly expressed in bone cells, impair bone formation during skeletal development (23). In differentiating ES cells, the already high Frz levels were further up-regulated with TSH to ∼50-fold of basal. Furthermore, the prototypical agonist for noncanonical Wnt-signaling Wnt5a was dramatically enhanced by both differentiation and TSH. Wnt5a is known to increase bone mass and to drive OPG production, mainly from mature osteoblasts (24).

Finally, TSH activated PKCδ, rather than PKA, in ES cell cultures. Importantly, rotterlin, a specific PKCδ inhibitor, strongly attenuated the osteoblastogenic response to TSH. This suggested that PKCδ mediated, at least in part, the actions of TSH on the osteoblast. Of note is that PKCδ is downstream of parathyroid hormone effects on osteoblast differentiation (25), and also, together with Runx2, mediates anabolic responses to connexin-43 and FGF-2 (26). Thus, the stimulation by TSH of PKCδ phosphorylation further confirms the induction of an anabolic pathway.

In summary, we show that ES cells are fully capable of differentiating into mature mineralizing osteoblasts; that TSH promotes osteoblast differentiation; that TSH stimulates OPG production, which may indirectly affect osteoclastogenesis; that a noncanonical pathway, mainly involving Frz and Wnt5a, mediates the osteoblastogenic effect of TSH; and that PKCδ is a downstream mediator of TSH action. We thus propose a short feed-forward loop in which TSH stimulates Wnt5a production that not only enhances osteoblastogenesis, but also produces OPG, which, in turn, dampens osteoclastic resorption to further increase bone mass (Fig. 6). Because osteoblasts and osteoclasts are in close anatomical proximity, and because their functions are tightly coupled in time and space (18), it is possible that TSH uncouples bone formation from bone resorption, using Wnt5a. The question remains, however, whether systemic, pituitary-derived, TSH is responsible for this modulation or whether local, bone marrow TSH, such as a TSH-β subunit variant (27), is the key to this permissive function.

Fig. 6.

TSH induces a local fast-forward loop for the regulation of bone remodeling. TSH stimulates Wnt5a secretion that enhances osteoblastogenesis and mineralization, but also, through its action on frizzled (Frz), stimulates the production of osteoprotegerin (OPG). OPG, in turn, inhibits bone resorption through its attenuation of RANKL signaling. These effects underscore the bone mass-enhancing effects of TSH that we have noted in vivo (13).

Materials and Methods

Cell Culture.

Mouse ES cell line (WT 9.5) was maintained in a permanent pluripotent state with the addition of leukemia inhibitory factor (LIF) (10 ng/mL; Stem Cell Technologies) on gelatin-coated dishes in Dulbecco's modified Eagle medium, (Invitrogen Life Technologies, Inc.) supplemented with 10% FBS, 1% penicillin–streptomycin, and 1.5 × 10⁴ M monothioglycerol (Sigma). Cultures were maintained in a humidified chamber in a 5% CO₂/air mixture at 37 ° C. ES cell cultures were monitored daily, and the cells were passaged at 1:3 ratios every 2 d. For all experiments, cells were plated at the appropriate densities without LIF for differentiation into its mesenchymal lineage, followed by osteogenic differentiation factors (DFs). Lineage differentiation was accomplished over 5 consecutive days, followed by osteogenic differentiation for another 30 consecutive days with treatments containing differentiation (+) or no differentiation (−) factor with or without TSH. Media were changed and treatments refreshed every 3 d. All osteoblast cultures were terminated and analyzed on day 30 for the entire experiment except for CFU-fs, which was terminated and stained on day 12 of culture. All cultures were treated with osteogenic differentiation factors containing 10 mM β-glycerophosphate, 1 mM dexamethazone, 50 μg/mL ascorbic acid, and 5 × 10⁻⁸ M vitamin D3. Vitamin D3, calciotrophic hormone 1,25-dihydroxyvitamin D(3) [1,25-(OH)(2)D(3)] was initally added on day 21 of culture.

CFU-f Staining for Alkaline Phosphatase.

Cells were plated (1.0 × 10⁶) in six-well plates (Falcon, BD Biosciences) and fixed with 10% formalin. At day 3, multicellular fibroblastoid colonies (CFU-fs), appeared that became increasingly alkaline phosphatase positive over the following week. Upon further incubation, the colonies mineralized to form CFU-obs (1). CFU-fs colonies were stained for alkaline phosphatase using a kit as per the manufacturer's recommendations (Sigma Aldrich) on day 12.

Staining for Mineralization of Osteoblasts.

Mineralization of osteoblasts was ascertained by (i) von Kossa staining (28) and (ii) Alizarin red staining. For von Kossa staining, cells were plated and fixed as described above for alkaline phosphatase staining. Cells were stained with 2% silver nitrate followed by 2.5% sodium thiosulfate (Sigma Aldrich). For Alizarin red staining, cells were similarly plated and fixed and stained with 2% Alizarin red stain (28) (pH 4.2) for 10 min, washed with dH2O several times, and finally allowed to air dry at room temperature. Measurement of Alizarin red stain eluted from these cells was used as an index of calcium release from osteoblast cells in culture. Cells were grown in a 24-well dish with 1 × 10⁵ cells/well, the stain was eluted by shaking in 100% iso-propanol for 10 min, and the OD was measured at 540 nm.

Measurement of Calcium and Phosphorous in Osteoblasts.

Cells were cultured in a 24-well dish at a density of 1 × 10⁵ cell/well and allowed to differentiate. On day 30, cells were washed twice with 1× PBS, and 0.1 N NaOH was added to each well for 24 h. Cells were then washed twice again with 1× PBS and centrifuged, and the supernatant was collected and tested for calcium and phosphorous, as per the manufacturer's instructions (Sigma Aldrich kit).

Detection of TSH Receptors in Osteoblasts by Flow Cytometry.

Briefly, cells were plated at a density of 6 × 10⁶ cell/10-cm dish and then treated with osteogenic stimulation for 30 d. Cells were trypsinized, washed once with FACS buffer (PBS + 0.02% sodium azide), and then stained with TSH receptor-specific antibody M1 to residues 381–385 (kindly supplied by B. Rees Smith, RSR Ltd., Cardiff, UK) for 1 h at room temperature. Following primary antibody staining, the cells were washed with buffer and stained further with an anti-mouse Fab PE-conjugated antibody (Jackson ImmunoResearch) at 1:200 dilution. Stained cells were then analyzed in the red channel (FL2) for PE staining. Negative controls stained with primary and secondary isotypes were also analyzed the same way.

RNA Isolation.

Total RNA was isolated from cells using TRIZOL reagent (Invitrogen), and chromosomal DNA was removed in accordance with the manufacturer's instructions. The RNA concentration was determined on the basis of absorbance at 260 nm, and its purity was evaluated by the ratio of absorbance at 260:280 nm (>1.9). RNAs were kept frozen at −70 °C until analyzed. After digestion of genomic DNA by treatment with Ambion's Turbo DNA-free DNase I (Ambion, Inc.), total RNA (1 μg) was reverse-transcribed into cDNA with random hexamers using Advantage RT-for-PCR kit (Clontech Laboratories, Inc.).

Semiquantitative PCR.

RT-PCRs for stem cell and osteoblast markers was performed using Titanium Taq polymerase (Clontech Laboratories). Details of primer sequences are shown in Table S1. Cycling conditions were as follows: 94 °C for 1 min, followed by 30 cycles of amplification (94 °C denaturation for 0.5 min; annealing for 1 min, annealing temperature dependent on primers; 72 °C elongation for 2 min) with a final incubation at 72 °C for 7 min. The amplified PCR products were separated on a 2% agarose gels and visualized by ethidium bromide staining.

Quantitative RT-PCRs.

Quantitative RT-PCRs (qRT-PCRs) were performed using the Applied Biosystems StepOne Plus real-time PCR system (Applied Biosystems). The reactions were established with a Power SYBR Green master mix (Applied Biosystems), 0.4 μL (2 μM) sense/antisense gene-specific primers, 2 μL cDNA, and diethylpyrocarbonate-treated water to a final volume of 20 μL. The PCR mix was denatured at 95 °C for 60 s before the first PCR cycle. The thermal cycle profile was the following: denaturizing for 30 s at 95 °C, annealing for 30 s at 57–60 °C (dependent on primers), and extension for 60 s at 72 °C. A total of 40 PCR cycles were used. PCR efficiency, uniformity, and linear dynamic range of each qRT-PCR assay were assessed by the construction of standard curves using DNA standards. An average threshold cycle from triple assays was used for further calculation. For each target gene, the relative gene expression was normalized to that of the glyceraldehyde-3-phosphate dehydrogenase (GAPDH) housekeeping gene using Applied Biosystems Step One Plus Real-time PCR systems software. Data presented (mean) are from three independent experiments in which all sample sets were analyzed in triplicate. The primers used are detailed in Table S1.

Western Blot Analyses.

The cellular protein was extracted in the lysis buffer (pH 7.5) containing Tris⋅HCl (50 mM), Triton X-100 (0.5%), EDTA (2 mM), NaCl (150 mM), and 1 mM PMSF. The protein was measured by the method of Lowry (29): 50 μg of protein was electrophoresed in 12% SDS-polyacrylamide gel under reducing conditions and electrotransfered to nitrocellulose membranes (Millipore). After blocking in 5% BSA, the membranes were incubated with anti–β-catenin (1:2,000), anti-actin at a dilution of 1:2,000, anti-OPG, anti-Wnt5a at a dilution of 1: 500 (Santa Cruz Biotechnology), and anti–PKC-δ and anti-PKA antibodies at a dilution of 1:2,000 (Cell Signaling) and then probed with anti-rabbit, anti-goat, or anti-mouse secondary antibody conjugated with peroxidase (Santa Cruz Biotechnology). The signals were detected with an ECL Western blotting detection system (Amersham Biosciences).

Osteoblast Signaling.

Osteoblast cultures were grown in six-well plates at a density of 1 × 10⁶ cells/well and cultured as described above (Cell Culture) with DFs and TSH (1 mU/mL). On day 5 or day 30, media were refreshed with serum-free media with or without rotterlin for 1 h before differentiation treatment with or without TSH for another hour. Cultures were then terminated after the 2-h treatment.

Statistical Analyses.

Differences resulting from treatments were assessed by Student's t-test or ANOVA. All differences were considered statistically significant if P < 0.05. All data are expressed as mean ± SEM. The statistical analyses were performed using Graph Pad v4.0.