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Molecular Endocrinology, doi:10.1210/me.2006-0342
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Molecular Endocrinology 21 (2): 376-387
Copyright © 2007 by The Endocrine Society

The Orphan Receptor Tyrosine Kinase Ror2 Promotes Osteoblast Differentiation and Enhances ex Vivo Bone Formation

Yan Liu, Ramesh A. Bhat, Laura M. Seestaller-Wehr, Shoichi Fukayama, Annamarie Mangine, Robert A. Moran, Barry S. Komm, Peter V. N. Bodine and Julia Billiard

Women’s Health and Musculoskeletal Biology, Wyeth Research, Collegeville, Pennsylvania 19426

Address all correspondence and requests for reprints to: Julia Billiard, Ph.D., Women’s Health and Musculoskeletal Biology, Wyeth Research, 500 Arcola Road, Collegeville, Pennsylvania 19426. E-mail: billiaj{at}wyeth.com.


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Ror2 is a receptor tyrosine kinase, the expression of which increases during differentiation of pluripotent stem cells to osteoblasts and then declines as cells progress to osteocytes. To test whether Ror2 plays a role in osteoblastogenesis, we investigated the effects of Ror2 overexpression and down-regulation on osteoblastic lineage commitment and differentiation. Expression of Ror2 in pluripotent human mesenchymal stem cells (hMSCs) by adenoviral infection caused formation of mineralized extracellular matrix, which is the ultimate phenotype of an osteogenic tissue. Concomitantly, Ror2 over-expression inhibited adipogenic differentiation of hMSCs as monitored by lipid formation. Ror2 shifted hMSC fate toward osteoblastogenesis by inducing osteogenic transcription factor osterix and suppressing adipogenic transcription factors CCAAT/enhancer-binding protein {alpha} and peroxisome proliferator activated receptor {gamma}. Infection with Ror2 virus also strongly promoted matrix mineralization in committed osteoblast-like MC3T3-E1 cells. Expression of Ror2 in a human preosteocytic cell line by stable transfection also promoted further differentiation, as judged by inhibited alkaline phosphatase activity, potentiated osteocalcin secretion, and increased cellular apoptosis. In contrast, down-regulation of Ror2 expression by short hairpin RNA essentially abrogated dexamethasone-induced mineralization of hMSCs. Furthermore, down-regulation of Ror2 expression in fully differentiated SaOS-2 osteosarcoma cells inhibited alkaline phosphatase activity. We conclude that Ror2 initiates commitment of MSCs to osteoblastic lineage and promotes differentiation at early and late stages of osteoblastogenesis. Finally, using a mouse calvariae ex vivo organ culture model, we demonstrate that these effects of Ror2 result in increased bone formation, suggesting that it may also activate mature osteoblasts.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
ROR2 BELONGS TO a family of receptor tyrosine kinases that, in mammals, consists of two members, Ror1 and Ror2. Ror receptors are most closely related to Trk neurotrophin receptors and muscle-specific kinase (1). To date, Ror2 has been known mainly for its role in developmental morphogenesis, particularly of the cartilage-derived skeleton. Disruption of mouse Ror2 leads to profound skeletal abnormalities with all endochondrally derived bones foreshortened or misshapen (2, 3, 4), and mutations within the Ror2 gene in humans are responsible for short stature, limb bone shortening, and segmental defects of the spine (5, 6, 7, 8). Furthermore, Ror2 has been shown to functionally interact with BRI-b, a bone morphogenetic protein receptor that modulates chondrogenesis (9). In addition to their skeletal phenotype, mice lacking Ror2 exhibit ventricular septal defects and respiratory dysfunction resulting in neonatal lethality (2, 3). More recently, Ror2 has been identified as a prosurvival kinase in HeLa cervical carcinoma cells (10) and shown to modulate neurite extension in central neurons (11).

Osteoblasts arise from mesenchymal stem cells (MSCs) and undergo further differentiation to either lining cells or osteocytes (12, 13, 14, 15, 16). The transition from stem cells to mature osteoblasts is characterized by the formation of mineralized extracellular matrix (17) and is controlled by key osteogenic transcription factors, Runx2 (18, 19, 20) and osterix (21), as well as dozens of hormones, growth factors, and cytokines.

From multiple models that represent distinct stages of osteoblastogenesis, we chose several for our studies. As a model of lineage commitment and early differentiation, we used human (h) MSCs, which are multipotent and can give rise in vitro to osteoblasts, adipocytes, or chondrocytes depending on the culture conditions (22, 23). Well-characterized murine MC3T3-C1 cells (24, 25) and human SaOS-2 osteosarcoma cells (26, 27) were used as more mature osteoblast-like models. Finally, a human HOB-01–09 preosteocytic cell line developed in our laboratory was used as a model representing a late stage of differentiation. HOB-01–09 cells were developed by infecting primary human bone cells with an adenovirus that encoded a temperature-sensitive simian virus 40 (SV40) large T antigen and passaging them beyond the crisis point (i.e. passages 15–20) to obtain an immortal line suitable for stable transfections (17, 28, 29). These cells exhibit a preosteocytic phenotype, as verified by cell morphology and expression of distinct protein markers. In addition, they proliferate at the permissive temperature (34 C) but stop dividing at the nonpermissive temperature (39 C) when the T antigen is inactivated.

Using some of the models described above, we have previously shown that Ror2 expression is strongly regulated during osteoblastic differentiation, being virtually undetectable in pluripotent stem cells, peaking in preosteoblasts, and then declining as cells progress to osteocytes (30). This strongly suggests that Ror2 plays a role in osteoblastogenesis.

Here we investigate a possible involvement of Ror2 in osteoblast lineage commitment and differentiation. We find that overexpression of Ror2 in hMSCs by adenoviral infection induces expression of osteogenic transcription factors, osterix and Runx2, and causes formation of mineralized extracellular matrix. Concomitantly, Ror2 overexpression inhibits adipogenic differentiation of hMSCs. Ror2 also promotes matrix mineralization in the committed osteoblast-like MC3T3-E1 cells and induces further differentiation of HOB-01–09 preosteocytes upon stable overexpression. In contrast, inhibition of endogenous Ror2 expression suppresses hMSC mineralization caused by a well-known osteogenic agent dexamethasone (dex) (31, 32) and inhibits alkaline phosphatase (AP) activity in SaOS-2 osteosarcoma cells. Thus, Ror2 initiates commitment of hMSCs to the osteoblastic lineage and promotes differentiation at both early and late stages of osteoblastogenesis. Interestingly, the majority of these effects are independent of the tyrosine kinase activity of the receptor, because they are also mediated by the kinase domain mutant Ror2KD. Finally, we demonstrate that these effects of Ror2 result in increased bone formation in mouse calvariae during ex vivo organ culture.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
We have previously shown that expression of Ror2 mRNA is strongly induced during osteogenic differentiation of hMSCs (30). To test whether Ror2 plays a role in osteoblastogenesis, we assessed functional consequences of Ror2 overexpression and down-regulation on hMSC differentiation. For Ror2 overexpression, we generated two adenoviruses encoding Ror2 and the kinase domain mutant (Ror2KD), each containing a COOH-terminal flag epitope tag. In the Ror2KD, three lysines at positions 504 (in the putative ATP-binding domain), 507, and 509 were replaced with isoleucines (30, 33), resulting in a significantly reduced tyrosine kinase activity (Fig. 1AGo). Because mutant Ror2 forms may be misfolded and retained in the endoplasmic reticulum (ER) (34), we verified Ror2 and Ror2KD expression in the plasma membrane, but not in the SIL-1-positive ER-containing fraction of the cell extract (Fig. 1BGo). hMSCs were infected with the Ror2 or Ror2KD viruses, and the level of Ror2 protein expression was monitored for 12 d. As shown in Fig. 1CGo, the expression was detectable within 24 h after infection, reached its peak approximately 6 d later, and declined slowly thereafter. For control, hMSCs were infected with the ß-galactosidase (ß-gal) expression cassette in the same adenoviral background. X-gal staining and immunocytochemistry with anti-flag antibody revealed greater than 90% infectivity 3–6 d after infection (see Fig. 1DGo for d 6). When uninfected or ß-gal-infected hMSCs were cultured in the presence of ascorbic acid and ß-glycerophosphate (ß-GP) for up to 15 d, we observed no mineralized matrix formation (Fig. 1EGo and data not shown). Under the same conditions, the hMSCs infected with Ror2 adenovirus formed mineralized matrix after 15 d in culture as evidenced by both alizarin red-S and von Kossa staining (Fig. 1EGo). Overexpression of the Ror2KD mutant produced a somewhat lower degree of mineralization than the wild-type protein (Fig. 1EGo). Quantitation of alizarin red-S incorporation revealed 18-fold induction by Ror2 and 14-fold induction by the Ror2KD mutant. Neither Ror2 nor Ror2KD affected total cell number or the apoptosis rate for hMSCs when compared with ß-gal infection (total cell numbers assessed on d 3 post infection; data not shown). Ror2 and Ror2KD viruses also strongly promoted matrix mineralization in the mature osteoblast-like murine MC3T3-E1 cells (Fig. 2Go). Thus, Ror2 promotes commitment of MSCs to osteoblastic lineage as well as enhances differentiation of a committed osteoblast cell line. The tyrosine kinase activity of the receptor seems to be largely dispensable for this effect.


Figure 1
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  		Fig. 1. Ror2 Overexpression Causes Mineralized Matrix Formation in hMSCs

A, Kinase activity of Ror2 and Ror2KD proteins. U2OS cells were infected with Ror2 or Ror2KD adenoviruses, and the whole-cell extracts were immunoprecipitated on flag affinity agarose and subjected to in vitro autophosphorylation assay. B, Ror2 and Ror2KD are expressed in the plasma membrane and not in the ER. U2OS cells were infected with Ror2 or Ror2KD adenoviruses before isolation of membrane and ER-containing protein fractions. Subsequently, 25 µg/lane membrane (M) and 16 µg/lane ER-containing (ER) fractions were used in Western immunoblotting for the flag epitope tag and for the ER-resident protein SIL-1. C, Time course of Ror2 and Ror2KD overexpression in hMSCs. hMSCs were infected with Ror2 or Ror2KD adenoviruses on d 0 and maintained in growth medium for the indicated periods of time before isolation of whole-cell protein extracts and Western immunoblotting for the flag epitope tag (5 µg protein/lane). D, Infection efficiency observed in hMSCs. hMSC cultures were infected with ß-gal, Ror2, or Ror2KD adenoviruses, incubated for 6 d in MSCGM supplemented with 0.05 mM ascorbic acid and 10 mM ß-GP, and subjected to X-gal staining (left panel) or immunocytochemistry for the flag epitope (center and right). E, Mineralized matrix formation in hMSCs upon infection. hMSCs were infected as in D, incubated for 15 d in MSCGM supplemented with 0.05 mM ascorbic acid and 10 mM ß-GP, and subjected to Alizarin red-S or von Kossa staining for mineralized nodule formation. WB, Western blot.

 

Figure 2
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  		Fig. 2. Ror2 Overexpression Promotes Mineralized Matrix Formation in MC3T3-E1 Osteoblast-Like Cells

A, MC3T3-E1 cultures were infected with ß-gal, Ror2, or Ror2KD adenoviruses and 6 d later subjected to X-gal staining (left panel) or immunocytochemistry for flag epitope tag (center and right). B, MC3T3-E1 cells were infected as in A, grown to confluence, and incubated for 15 d in growth medium supplemented with 10 mM ß-GP and ascorbic acid (12.5 µg/ml for the first feeding and 25 µg/ml for subsequent feedings). Alizarin red-S staining for mineralized nodule formation was performed as described in Materials and Methods.

 
To address the mechanism by which Ror2 promotes mineralization, we assessed its effects on the expression of the key osteogenic transcription factors, osterix and Runx2. Compared with ß-gal-infected control cells, Ror2 overexpression led to a 4-fold increase in osterix levels by d 3 and a 160-fold increase in osterix by d 9, just before matrix mineralization. Ror2KD overexpression led to a somewhat weaker, but still very substantial, induction of osterix (Fig. 3AGo). Overexpression of Ror2 also caused a moderate (~2 fold), but reproducible increase in Runx2 expression at d 3. This Runx2 induction was not observed in the Ror2KD-infected cells (Fig. 3BGo), indicating that the increase in Runx2 is not required for matrix mineralization. In contrast, there seems to be a good correlation between the extent of osterix induction and matrix mineralization in response to Ror2 and Ror2KD, suggesting that osterix may underlie the ability of this tyrosine kinase receptor to promote commitment of hMSCs to osteogenesis (Fig. 3AGo). Ror2-induced differentiation resulted in increased expression of several osteoblast-specific genes, including type I collagen, osteopontin, and mineralized tissue-specific bone sialoprotein (BSP) after 15 d (Fig. 3CGo), but not the osteocytic markers dentin matrix protein 1 (35) or sclerostin (36) (data not shown). In contrast, the HOB-01–09 preosteocytic cells expressed high levels of sclerostin, but no detectable BSP (data not shown). Thus Ror2 induced differentiation of hMSCs to mature osteoblasts that produce the mineralized matrix, but did not promote differentiation to osteocytic stage.


Figure 3
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  		Fig. 3. Ror2 Promotes Expression of Osteogenic Transcription Factors in hMSCs

hMSCs were infected with ß-gal, Ror2, or Ror2KD adenoviruses and incubated for the indicated times in MSCGM supplemented with 0.05 mM ascorbic acid (AA) and 10 mM ß-GP. At indicated times, total cellular RNA was isolated and subjected to real-time RT-PCR analysis for osterix (A) or Runx2 (B). The levels of mRNA were normalized to the expression of cyclophilin B in each sample, and the relative mRNA expression in Ror2-infected cells (solid bars) and Ror2KD-infected cells (open bars) was compared with the expression in ß-gal-infected cells, which was set at 1 at each time point. The figure shows representative traces of at least three independent experiments. C, hMSCs were infected with ß-gal or Ror2 adenoviruses and incubated for 15 d in MSCGM supplemented with 0.05 mM ascorbic acid (AA) and 10 mM ß-GP. Total cellular RNA was isolated and subjected to real-time RT-PCR analysis for BSP, type I collagen (col 1), and osteopontin (OPN). The levels of mRNA were normalized to the expression of cyclophilin B in each sample, and the relative mRNA expression in ß-gal-infected cells was set at 1.

 
We next assessed effects of Ror2 on an alternate fate of hMSCs, i.e. adipogenesis, induced by incubation in indomethacin- and 3-isobutyl-1-methylxanthine-containing adipogenic cocktail. Both Ror2 and Ror2KD mutant inhibited expression of the major adipogenic transcription factors, CCAAT/enhancer-binding protein (C/EBP){alpha} and peroxisome proliferator activated receptor (PPAR){gamma} (Fig. 4AGo), and caused a marked decrease in the ability of hMSCs to form oil red O-positive lipid-producing adipocytes (Fig. 4BGo). Thus, Ror2 alters the cell fate of MSCs by shifting the balance of transcription factors to favor osteoblastogenesis.


Figure 4
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  		Fig. 4. Ror2 Overexpression Inhibits Adipogenic Differentiation of hMSCs

A, hMSCs were infected with ß-gal, Ror2, or Ror2KD adenoviruses and incubated in MSCGM supplemented with adipogenic cocktail for 8 d. Total cellular RNA was isolated and subjected to real-time RT-PCR analysis for adipogenic transcription factors C/EBP{alpha} and PPAR{gamma}. The levels of mRNA were normalized to the expression of cyclophilin B in each sample, and the relative mRNA expression in ß-gal-infected cells was set at 100%. B, hMSCs were infected with ß-gal, Ror2, or Ror2KD viruses, incubated in MSCGM supplemented with adipogenic cocktail for 21 d, and then subjected to oil red O staining. C, Quantification of oil red O staining from three independent experiments (*, P < 0.05). WB, Western blot.

 
To assess whether the rise in Ror2 expression during hMSC differentiation is critical for osteoblastogenesis, we performed dex-induced differentiation when endogenous Ror2 expression was inhibited. To this end, hMSCs were infected with the adenovirus containing Ror2-specific short hairpin RNA (shRNA), which strongly inhibited the dex-induced rise in Ror2 protein expression when compared with enhanced green fluorescent protein (EGFP)-specific control shRNA (Fig. 5AGo). Infection with Ror2 shRNA essentially abrogated the ability of dex to induce matrix mineralization (Fig. 5Go, B and C), suggesting that the rise in Ror2, at least in part, mediates dex-induced osteoblastic differentiation of hMSCs.


Figure 5
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  		Fig. 5. Down-Regulating Ror2 Inhibits dex-Induced Osteogenic Differentiation of hMSCs

hMSCs were infected with adenoviral expression vectors containing shRNA specific for Ror2 or EGFP (control) and incubated in MSCGM supplemented with 0.05 mM ascorbic acid, 10 mM ß-GP, and 100 nM dex. After 9 d of incubation, 50 µg of the whole-cell protein extracts was subjected to Western immunoblotting for the endogenous Ror2 protein (A). After 11 d of incubation, alizarin red-S staining was performed (B) and quantified (C) to assess the extent of mineralized matrix formation. In panel C, the amount of alizarin red-S incorporated in presence of EGFP shRNA was set at 100%. The results in B and C are representative of three independent experiments. WB, Western blot.

 
To obtain further evidence for osteogenic potential of the endogenous Ror2 protein, we used the SaOS-2 osteosarcoma cell line that exhibits high levels of Ror2 protein expression, secretes high levels of alkaline phosphatase (AP), and allows high transfection efficiency. We were able to achieve approximately 90% reduction of Ror2 protein in SaOS-2 cells by transfection with Ror2-specific short interfering RNA (siRNA) (Fig. 6AGo). This down-regulation of endogenous Ror2 caused approximately 45% suppression of AP activity produced by SaOS-2 cells (Fig. 6BGo). Thus, down-regulation of Ror2 causes partial reversal of the osteoblast phenotype in SaOS-2 cells.


Figure 6
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  		Fig. 6. Down-Regulating Ror2 Inhibits AP Activity in SaOS-2 Osteosarcoma Cells

SaOS-2 cells were transfected with Ror2 siRNA or nonspecific siRNA, and 48 h later whole-cell protein extracts were subjected to Western immunoblotting for the endogenous Ror2 protein using 50 µg of extract per lane (A). AP activity was assessed 72 h after transfection and normalized to the amount of total cellular protein (B). The results in B are means ± SE of at least three independent experiments (*, P < 0.05). WB, Western blot.

 
We have previously shown that where Ror2 expression increases from stem cells to osteoblasts, it declines during further differentiation and essentially disappears in osteocytes (30). To investigate the effect of Ror2 on later stages of osteoblast differentiation, we stably overexpressed this receptor in HOB-01–09 preosteocytic cell line characterized by low endogenous levels of Ror2. The preosteocytic phenotype of HOB-01–09 was confirmed by a 18-fold increase in NO secretion in response to mechanical stress (Fig. 7Go) and by expression of high levels of an osteocyte-specific gene sclerostin (Fig. 3Go). Figure 8Go, A and B, illustrates Ror2 mRNA and protein overexpression obtained after stable transfection with Ror2-flag or Ror2KD-flag when compared with endogenous levels observed in pcDNA3.1-expressing cells. We generated several other cell lines overexpressing the untagged and flag-tagged wild-type Ror2 receptor by 3- to 5-fold but were unable to achieve higher overexpression, suggesting that higher levels of Ror2 are not well tolerated by preosteocytic cells. In contrast, the kinase domain mutant was overexpressed by 30-fold. All experiments were performed in three independent Ror2 and Ror2-flag lines with similar results, but only the results for one Ror2-flag line are reported. Ror2 overexpression caused marked changes in cellular appearance, with cells aggregating in nodules typical of later stage osteoblasts (Fig. 8CGo). We also found that Ror2, but not Ror2KD overexpression, resulted in a 3-fold increase in osteocalcin accumulation in the medium (Fig. 8DGo) and that both Ror2 and Ror2KD almost completely abrogated AP activity in HOB-01–09 cells (Fig. 8EGo). In addition, both Ror2 and Ror2KD caused a 60% increase in the rate of HOB-01–09 apoptosis, as assessed by DNA fragmentation (Fig. 8FGo). All of the changes affected by Ror2 in HOB-01–09 lines are indicative of further differentiation of preosteocytic cells toward osteocytes.


Figure 7
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  		Fig. 7. Mechanical Stress Causes NO Production in HOB-01–09 Cells

The cells were grown and subjected to mechanical stimulus as described in Materials and Methods. The results are means ± SE of four independent experiments.

 

Figure 8
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  		Fig. 8. Ror2 Overexpression Promotes Further Differentiation of HOB-01–09 Preosteocytes

A, Real-time RT-PCR analysis of the Ror2 mRNA expression in HOB-01–09 cells overexpressing Ror2-flag (R2), Ror2KD-flag (KD), or empty vector (pcDNA 3.1, DNA). The levels of mRNA were normalized to the expression of 18S rRNA in each sample, and the relative mRNA expression in HOB-01–09-pcDNA cells was set at 1. B, Flag-tagged Ror2 and Ror2KD proteins were immunoprecipitated out of 1 mg of the whole-cell protein extracts on anti-flag affinity agarose before Western immunoblotting with the anti-flag antibody. C, Alizarin red-S staining was performed on the HOB-01–09 cells overexpressing the indicated constructs. D, E, and F, HOB-01–09 cells overexpressing the indicated construct were incubated for 48 h in BSA-medium at 39 C, and osteocalcin secretion (D), AP activity (E), and DNA fragmentation (F) were assessed as described in Materials and Methods. Values in D and E were normalized to total cellular protein. In F, the level of DNA fragmentation observed in HOB-01–09-pcDNA cells was set at 1. The results are means ± SE of at least three independent experiments (*, P < 0.05). IP, Immunoprecipitation; WB, Western blot.

 
Finally, we asked whether the ability of Ror2 to promote osteoblast differentiation in vitro translates into increased bone formation in organ culture. Calvariae from 4-d-old mouse pups were excised and either left uninfected or infected with ß-gal, Ror2, or Ror2KD adenoviruses. The total bone area was assessed 7 d later after hematoxylin and eosin staining (Fig. 9AGo). As shown in Fig. 9BGo, infection with the Ror2 virus resulted in a 72% increase in the total bone area compared with uninfected or ß-gal-infected calvariae (Fig. 9AGo) without a change in total osteoblast number, suggesting that Ror2 increased bone formation by existing osteoblasts. Infection with the Ror2KD virus did not increase the total bone area, implying that whereas kinase activity was dispensable for the Ror2 function in vitro, it is required for the Ror2-induced osteogenesis in organ culture (Fig. 9BGo). Thus, Ror2 not only promotes osteoblast differentiation in vitro but also is a potent bone-forming agent ex vivo.


Figure 9
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  		Fig. 9. Ror2 Overexpression Increases Total Bone Area of Neonatal Mouse Calvariae

Calvarial bones of 4-d-old mouse littermates were either left uninfected (control) or infected with ß-gal, Ror2, or Ror2KD adenoviruses. After 7 d of incubation in the presence of adenovirus, calvariae were stained with hematoxylin and eosin and photographed under the light microscope (A). Subsequently, total bone area (open bars) and osteoblast number (solid bars) were determined for each calvaria and the values observed in uninfected cultures were set at 100% (B). The results are means ± SE of four to five calvariae per condition (*, P < 0.0001 compared with ß-gal infection). The results are representative of three independent experiments.

 

   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
MATERIALS AND METHODS
 REFERENCES
 
Our data provide strong evidence that Ror2 initiates commitment of MSCs to osteoblastic lineage as well as promotes differentiation at both early and late stages of osteoblastogenesis. Furthermore, we demonstrate that these effects of Ror2 lead to increased bone formation in ex vivo organ cultures.

A reciprocal relationship exists between bone and fat formation in vivo with an increase in marrow adipocytes observed in all conditions associated with a loss of bone mass, such as ovariectomy, immobilization, or treatment with glucocorticoids (37). This relationship is also observed between osteogenic and adipogenic differentiation of marrow stromal cells in vitro (37, 38), where several signaling cascades have been shown to potentiate one fate and inhibit the other. These include the MAPK cascade (39) and canonical Wnt signaling (40), which promote osteogenesis at the expense of adipogenesis, and PPAR{gamma} signaling that increases adipogenesis and decreases osteogenesis (41, 42, 43). We can now add Ror2 to the factors that shift hMSC fate toward osteoblastogenesis. The shift occurs through induction of osteogenic transcription factor osterix and suppression of adipogenic transcription factors C/EBP{alpha} and PPAR{gamma}, but the mechanism by which Ror2 regulates mRNA expression of these transcription factors remains to be fully elucidated.

In addition to the most obvious marker of differentiation, formation of a mineralized matrix, overexpression of Ror2 in hMSCs induced expression of a mineralized tissue-specific marker BSP (44) but not of the osteocytic marker sclerostin (36). In contrast, HOB-01–09 preosteocytes expressed high levels of sclerostin, suggesting that hMSCs never reach the osteocytic stage of differentiation in our system, but go only as far as mature osteoblasts that synthesize the bone matrix.

We observed a very modest induction of the Runx2 expression that was abolished in the Ror2KD mutant. Because the Ror2KD mutant induces strong matrix mineralization, we conclude that an increase in Runx2 expression is not a prerequisite for this effect. However, osterix, a gene that is downstream of Runx2 (21), was dramatically enhanced by both Ror2 and Ror2KD overexpression, and the extent of this enhancement correlated with the degree of matrix mineralization. It is possible that Ror2 enhances Runx2 activity at the posttranscriptional level, because translational and posttranslational modifications have been shown to be the main mechanism of Runx2 activation in multiple systems (45, 46, 47). Alternatively, Ror2 may regulate osterix expression through Runx2-independent pathways such as activation of the Dlx5 transcription factor that mediates BMP2-induced osterix production (48).

Some of the growth factors known to modulate hMSC differentiation belong to the Wnt family of secreted glycoproteins. Wnts bind to a membrane receptor complex composed of a G protein-coupled receptor, Frizzled, and a low-density lipoprotein receptor-related protein, and this binding activates either the canonical ß-catenin pathway or one of several noncanonical pathways (49, 50). Ror2 has been shown to modulate both canonical and noncanonical Wnt signaling. For the canonical Wnts, Ror2 potentiates Wnt1-mediated transcription and inhibits Wnt3 signaling (30). Interestingly, despite a well-documented role of canonical Wnt signaling in inducing osteogenesis in humans and mice (51, 52, 53, 54), and in mouse and rat stromal cells in vitro (55, 56, 57), it appears that continuous activation of canonical Wnt signaling actually inhibits osteogenic differentiation of hMSCs in vitro (Refs. 58 and 59 , and our unpublished data). These observations raise a possibility that Ror2 promotes hMSC mineralization by down-regulating Wnt3 signaling. However, the Ror2KD mutant retains only about 40% of Wnt3-inhibitory potential (30), but approximately 80% of its mineralization potential (Fig. 1Go), making it unlikely that Ror2 functions in hMSCs solely through Wnt3 inhibition. Ror2 has also been shown to bind noncanonical Wnt5a and to enhance its signaling to c-Jun N-terminal kinase (60). In hMSCs, Wnt5a does not promote mineralization (59), which is the phenotype opposite to that elicited by Ror2. Thus, it appears unlikely that the ability of Ror2 to promote hMSC differentiation is connected solely to its function in the Wnt signaling pathway.

In addition to its effects on hMSC commitment, Ror2 promoted differentiation in several models of later stages of osteoblastogenesis, including committed osteoblast-like MC3T3-E1 and SaOS-2 cells. Furthermore, introducing Ror2 into preosteocytic cells by stable transfection also elicited a more mature phenotype. Indeed, further differentiation at these late stages of human osteogenesis is characterized by a decrease in AP activity, an increase in osteocalcin secretion and apoptosis, and a change in cellular morphology (17). Ror2 overexpression affected all of these changes.

Our results indicate that the tyrosine kinase activity of Ror2 is largely dispensable with regard to its role in in vitro osteoblast differentiation, but seems to be required for its effects on ex vivo bone formation. We have confirmed the loss of the tyrosine kinase activity in Ror2KD by both in vitro kinase assays and the loss of its ability to modulate canonical Wnt signaling (30). However, we cannot exclude the possibility that the minimal residual kinase activity accounts for the observed effects of Ror2KD in vitro after 15 d of incubation, whereas it is not sufficient over the 7-d course of our ex vivo studies. On the other hand, Ror2 is known to possess some kinase-independent functionality. Kinase-dead mutants of the Caenorhabditis elegans Ror protein CAM1 retain some, although not all, of the CAM1 functions and can mediate cell migration (61). The membrane-anchored extracellular domain of XRor2 inhibits convergent extension and retains some ability to prevent neural plate closure in Xenopus embryos (33). However, the secreted form of the extracellular domain of XRor2 loses these signaling capabilities (33), suggesting that an additional function of Ror2, perhaps as a cell adhesion molecule, is required for signaling. However, tyrosine kinase activity of Ror2 is required during embryonic development in humans and mice. Indeed, in humans, point mutations in the Ror2 tyrosine kinase domain cause Robinow syndrome as severe as that caused by deletion of almost the entire protein (5), and mice lacking the entire Ror2 receptor (3), as well as those lacking only the tyrosine kinase domain (4), appear to exhibit indistinguishable phenotypes. Thus, it appears that in different cellular contexts, Ror2 has both tyrosine kinase-dependent and -independent functions.

In conclusion, we have identified the Ror2 tyrosine kinase receptor as a novel stimulator of osteoblast commitment and differentiation. We further demonstrated that these effects of Ror2 translate into increased bone formation during organ culture, thus making Ror2 an attractive drug target to manipulate for the treatment of osteoporosis and other bone diseases.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
REFERENCES
 
Materials and Tissue Culture
Except where noted, tissue culture reagents were purchased from Invitrogen Corp. (Carlsbad, CA); other reagents and chemicals were purchased from either Sigma Chemical Co. (St. Louis, MO) or Invitrogen. Anti-flag M2 mouse monoclonal antibody and anti-flag M2 affinity agarose were obtained from Sigma; antihuman Ror2 goat polyclonal antibody was purchased from R&D Systems (Minneapolis, MN); and polyclonal antibody against resident ER SIL-1 protein was from Abcam, Inc. (Cambridge, MA). Horseradish peroxidase-conjugated and Alexa Fluor-conjugated secondary antibodies were from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA) and Molecular Probes, Inc. (Eugene, OR), respectively.

The hMSCs were purchased from Cambrex, Inc. (Baltimore, MD) and maintained at 37 C in a 5% CO2-95% humidified air incubator using hMSC growth medium (MSCGM, Cambrex). The MC3T3-E1 murine osteoblast-like cells and C3H10T1/2 murine mesenchymal cells were kept at 37 C in MEM containing 10% heat-inactivated fetal bovine serum (FBS), 1% penicillin-streptomycin, and 2 mM glutaMAX-I. The SaOS-2 and U2OS human osteosarcoma cells were maintained in McCoy’s 5A modified medium, containing 10% heat-inactivated FBS, 1% penicillin-streptomycin, and 2 mM glutaMAX-I. The human embryonic kidney (HEK) 293A cell line was maintained in DMEM, containing 10% heat-inactivated FBS, 1% penicillin-streptomycin, and 2 mM glutaMAX-I.

Adenoviruses
Generation of the flag-tagged Ror2 and flag-tagged Ror2KD mutant expression plasmids was previously described (30). Entire transcription units were cut out of these plasmids as MluI–SalI fragments containing the cytomegalovirus (CMV) promoter driving the open reading frames and SV40 late polyadenylation signal sequence. These were cloned into the corresponding sites in a modified pENTR1A plasmid (Invitrogen). The resulting pENTR1A plasmids were mixed with pAD/PL-DEST Gateway vector (Invitrogen), and recombination was performed using the LR clonase enzyme (Invitrogen). The plasmid pAD/PL-DEST contains the human Ad5 sequence with the deletion of E1a and E3 region and is suitable for the construction of recombinant viruses for protein expression. The recombinant plasmids were linearized by digestion with PacI, purified by phenol/chloroform extraction, and precipitated. The DNA was dissolved in OptiMEM and transfected into HEK 293A cells using Lipofectamine 2000 (Invitrogen) as recommended by the vendor. After incubation for 24 h, the transfected cells were overlaid with growth medium containing 5% FBS and 1.25% (wt/vol) SeaPlaque agarose (Cambrex). The cells were fed with fresh agar overlay every third day. The recombinant plaques were isolated, amplified in HEK 293A cells, and confirmed by restriction analysis of the viral DNA. The recombinant viruses were plaque purified once before the amplification of the viral stock.

For construction of ß-gal adenovirus, the transcription unit containing CMV promoter, the ß-gal coding sequence, and SV40 polyadenylation signal was cloned as a SnaB1–SalI fragment into the Ad5{Delta}E1a plasmid containing adenovirus sequence within map units 0–17 with the deletion of E1a region between map units 1.4 and 9.1. The Ad5 recombinant plasmid was linearized and transfected along with the ClaI fragment of Ad5 with the deletion in the E3 region (80–88 map units) into HEK 293A cells using a calcium phosphate precipitation method. The virus plaques generated by homologous recombination were isolated, amplified, and verified by restriction analysis of the viral DNA. The virus was further plaque purified, and the virus stock was prepared in HEK 293A cells.

For the human coxsackie adenovirus receptor (hCAR) adenovirus, the hCAR open reading frame was amplified from HEK 293 cells RNA by RT-PCR using the following primers: 5'-AAAGGTACCACCATGGCGCTCCTGCTGTGCTTCGTG-3' (forward primer) and 5'-AATCTAGACTATACTATAGACCCATCCTTGCTCTG-3' (reverse primer). The Asp718–XbaI fragment was cloned into a modified pENTR Gateway vector. This expression plasmid contained between the recombination sites the complete transcription unit including the CMV promoter, hCAR open reading frame, and SV40 late poly A signal sequence. The hCAR adenovirus was isolated using the ViraPower Adenoviral expression system (Invitrogen). The hCAR entry plasmid was built into pAd/PL-DEST plasmid using the LR clonase reaction. The resulting pAD/CAR plasmid was digested with PacI to remove the vector sequence, and purified DNA was transfected into HEK 293A cells using Lipofectamine 2000. The virus plaques were isolated and plaque purified, and virus stocks were prepared and titered in HEK 293 cells.

Adenoviruses containing shRNA sequences specific for human Ror2 or for EGFP (control) were obtained from Galapagos, Inc. (Mechelen, Belgium). The shRNA target sequences were as follows: 5'-TGTGAGCAACGCCCGCTAC-3' (Ror2) and 5'-GCTGACCCTGAAGTTCATC-3' (EGFP).

Generation of Stable Cell Lines
HOB-01–09 postsenescent preosteocytic cell line was engineered as previously described (28, 29) using the temperature-sensitive SV40 large T antigen mutant. Cells immortalized with this large T antigen mutant proliferate and express a transformed phenotype at 34 C when the mutant protein is active but revert to a nontransformed phenotype at 39 C when the mutant protein is inactive. Ror2, Ror2-flag, and Ror2KD-flag in pcDNA3.1 or empty pcDNA3.1 were transfected into HOB-01–09 cells by electroporation. For each transfection, 10 µg of plasmid DNA was added to 8 x 106 cells in PBS and incubated on ice for 5 min. The mixture was electroporated in 2-mm gap cuvettes (BTX, San Diego, CA) using ECM 600 electroporator (BTX) with the following parameters: 100 µF, 48 Ohms, 150 V (pulse duration ~10 msec). Cells were cooled 5 min on ice, transferred to a 150-mm culture flask and maintained at 34 C in a 5% CO2-95% humidified air incubator using DMEM/F-12 medium containing 10% FBS, 1% penicillin-streptomycin, and 2 mM glutaMAX-I (HOB growth medium) supplemented with 500 µg/ml G418 (Invitrogen) until isolated colonies of G418-resistant cells were formed. Colonies were trypsinized and transferred one per well onto 96-well plates. Colonies were grown at 34 C in HOB growth medium supplemented with 125 µg/ml G418, and levels of Ror expression were assessed by real-time RT-PCR and Western immunoblotting. For experimental use, 24 h after plating cells were switched to the nonpermissive temperature of 39 C.

Calvarial Organ Culture and Infection
Mice were used in accordance with the Animal Welfare Act and the NIH Guide for the Care and Use of Laboratory Animals. Calvariae were excised from 4-d-old mouse littermates, cut along the sagittal suture, and incubated for 24 h in serum-free BGJ medium containing 0.1% BSA. Each half of the calvaria was then placed with the concave surface downward on a stainless steel grid (Small Parts, Inc., Miami, FL) in a well of a 12-well plate. Each well contained 1 ml BGJ medium with 1% FBS, without or with ß-gal, Ror2, or Ror2KD adenoviruses (3.75 million viral particles per well). Calvariae were incubated in a 5% CO2-95% humidified air incubator, and the medium and adenoviruses were changed after 4 d.

After 7 d of incubation in the presence of adenovirus, calvariae were fixed in 10% neutral phosphate-buffered formaldehyde at room temperature (RT) for 72 h and then decalcified for 6 h in 10% EDTA in PBS. Calvariae in each group were embedded in parallel in the same paraffin block, and 4-µm sections were stained with hematoxylin-eosin. Consistent bone areas (200 µm away from frontal sutures) were selected for histomorphometric analysis. In brief, a 200-µm square grid was placed on each calvaria, and the total bone area was determined with the Osteomeasure System (Osteometrics, Inc., Atlanta, GA).

Viral Infection
hMSCs were seeded at 6000/cm2 in 96-, 12-, or six-well plates and allowed to adhere and proliferate overnight. Cells were infected for 24 h in 0.4 ml/cm2 MSCGM using Ror2, Ror2KD, or ß-gal adenoviruses at multiplicity of infection (MOI) = 750 in the presence of hCAR (MOI = 750) to improve infection efficiency. After 24 h, cells were washed once in PBS and MSCGM, MSCGM supplemented with 0.05 mM ascorbic acid and 10 mM ß-GP, or MSCGM containing adipogenic supplements (PT-3004, Cambrex) were added. Where indicated, 100 nM dex was added to the medium. For shRNA infection, cells were seeded at 6000/cm2 in 12- or six-well plates and allowed to adhere and proliferate for 3 d. Cells were infected for 72 h in 0.4 ml/cm2 MSCGM using adenoviruses coding for Ror2-specific shRNA or EGFP-specific shRNA at 4000 viral particles per cell (based on original seeding density) in the presence of hCAR (MOI = 750). After 72 h, cells were washed once in PBS and MSCGM or MSCGM supplemented with 0.05 mM ascorbic acid, 10 mM ß-GP, and 100 nM dex was added. Media were changed every 5 d.

MC3T3-E1 cells were seeded at 6000/cm2 in 12-well plates and allowed to adhere and proliferate overnight. Cells were infected for 24 h in 0.4 ml/cm2 MC3T3-E1 growth medium containing reduced amount of FBS (1% vol/vol). Ror2, Ror2KD, or ß-gal adenoviruses were used at MOI = 3000 in the presence of hCAR (MOI = 750). After infection, cells were washed once in PBS and transferred into growth medium. At confluence (d 0), differentiation was induced by addition of growth medium supplemented with 10 mM ß-GP and ascorbic acid (12.5 µg/ml for the first feeding and 25 µg/ml for subsequent feedings). The medium was changed every 48 h until fixation and alizarin red-S staining.

U2OS cells were seeded at 75,000/cm2 in six-well plates and infected 24 h later with Ror2 or Ror2KD adenovirus at MOI = 100. Infection was allowed to proceed for 24 h, and cell extracts were collected another 24 h later.

siRNA Transfection
SaOS-2 cells were plated on six-well plates at 52,000 cells/cm2 and transfected 24 h later with 25 nM Ror2 siRNA or nonspecific siRNA (both from Dharmacon, Inc., Lafayette, CO) using 10 µl Lipofectamine 2000 reagent according to the manufacturer’s instructions.

Cell Fractionation, Immunoprecipitation, in Vitro Kinase Assay, and Western Blotting Analysis
To obtain total cellular protein, cells were solubilized in whole-cell lysis buffer [150 mM NaCl, 50 mM Tris-HCl (pH 7.5), 1% Triton X-100, protease and phosphatase inhibitor cocktails (both from Sigma)] and the extracts were clarified by centrifugation. For the membrane preparation, the cells were homogenized by electronic polytron, and the membranes were collected by centrifugation at 25,000 x g for 30 min at 4 C. The membrane proteins were extracted from the pellets by incubation with whole-cell lysis buffer for 20 min at 4 C and clarified by centrifugation. To separate the ER-containing fraction, cells were incubated in the hypotonic lysis buffer [10 mM Tris (pH 7.4); 200 µM MgCl2; protease and phosphatase inhibitor cocktails], homogenized in Dounce homogenizer and clarified at 100,000 x g for 90 min at 4 C.

For the in vitro autophosphorylation assay, U2OS cells were infected with Ror2 or Ror2KD adenoviruses, and the total cellular proteins were isolated and precipitated on M2 flag affinity agarose. The kinase assay was performed on precipitated proteins as previously described (30).

For detection of Ror proteins in HOB-01–09-stably transfected cell lines, flag-tagged Ror2 or Ror2KD was immunoprecipitated out of 1 mg of the whole-cell protein extract on anti-flag affinity agarose before Western immunoblotting.

For immunoblotting, protein extracts were resolved by SDS-PAGE under denaturing and reducing conditions, transferred to nitrocellulose membranes, and probed with specific antibodies.

Immunocytochemistry and X-gal Staining
For immunocytochemistry, the cells were fixed in 4% (wt/vol) paraformaldehyde, permeabilized with 0.25% Triton X-100, and blocked in 10% BSA. Monoclonal anti-flag M2 antibody (1:1000 in PBS plus 3% BSA) was applied for 2 h at RT, followed by Alexa 568-conjugated goat antimouse IgG (1:250) and 100 ng/ml Hoechst dye for 1 h in the dark. Cells were viewed by fluorescent microscopy (Nikon Eclipse TE 300; Nikon, Melville, NY), and images were captured with a Nikon digital camera DXM1200 and Nikon ATC-1 software and processed using Photoshop 7.0 (Adobe Systems, San Jose, CA). For X-gal staining, cells were fixed as above and incubated at 37 C overnight in 0.1% X-gal (Invitrogen) in PBS containing 5 mM K3Fe(CN)6, 5 mM K4Fe(CN)6, and 2 mM MgCl2. Cells were examined and imaged as for immunocytochemistry except using a visible light source.

Histochemical Staining
Formation of mineralized nodules by hMSC and MC3T3-E1 cells was determined on 12-well plates by alizarin red-S histochemical staining (28) or by von Kossa (3% wt/vol AgNO3) histochemical staining (62). To quantify the level of alizarin red-S staining, the dye was eluted with 1 ml/well of 10% (wt/vol) cetylpyridinium chloride. Alizarin red-S in the eluted samples was quantified (vs. a standard curve of 0–800 µM dye) at 562 nm with a microplate reader.

Adipogenesis was monitored in hMSCs on 12-well plates by oil red O histochemical staining. The cells were fixed at RT for 2 h with 10% neutral buffered formalin, washed with PBS, and stained for 10 min at RT with 18 mg/ml oil red O in 60% isopropanol, pH 7. The stained cells were washed with PBS and photographed. Oil red O staining was quantified at 510 nm, after elution with isopropyl alcohol.

RNA Isolation and Real-Time PCR Analysis
Total cellular RNA was isolated using the RNeasy Kit (QIAGEN, Valencia, CA) following the manufacturer’s instructions and subjected to real-time RT-PCR analysis using the ABI PRISM 7700 Sequence Detection System as described in Ref. 30 . All mRNA levels were normalized to the levels of housekeeping genes, cyclophilin B or 18S rRNA. Primers and probes for human C/EBP{alpha}, PPAR{gamma}, bone sialoprotein, sclerostin, osteopontin, dentin matrix protein 1, and 18S rRNA were purchased from Applied Biosystems (Foster City, CA); sequences of all other primers and probes are listed in Table 1Go.


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  		Table 1. Primers and Probes Used in the Real-Time PCR Analysis of Human Genes

 
AP and Osteocalcin Assays
HOB-01–09 stably transfected cells were plated at 50,000/cm2 in 12-well plates in HOB growth medium and allowed to proliferate at 34 C overnight. The cells were transferred into DMEM/F-12 containing 0.25% (wt/vol) BSA, 1% penicillin-streptomycin, 2 mM glutaMAX-I, 50 µg/ml ascorbic acid, and 10 nM menadione sodium bisulfite (BSA-medium) and incubated at 39 C for 48 h. At the end of this incubation, conditioned media were assayed for secreted osteocalcin using intact human osteocalcin ELISA kit (Biomedical Technologies, Inc., Stoughton, MA) according to manufacturer’s instructions except the protein standards were dissolved in BSA-medium. After the conditioned media were removed and the cells were washed with PBS, 60 µl of 0.1 M Tris buffer, pH 9.8, containing 0.2% (vol/vol) Triton X-100 (AP buffer) were added to each well, and the cells were stored at –80 C. After thawing, 40 µl of the cell lysate was assayed for AP activity as previously described (63). The amount of osteocalcin and AP activity in each well was normalized to the amount of total cellular protein in the same well determined by the bicinchoninic acid assay. SaOS-2 cells were seeded at 52,000 cells/cm2 in six-well plates, transfected with the indicated siRNA, and incubated at 37 C for 48 h. AP activity was assessed as described above except that 200 µl of AP buffer was used per well, and 1 µl of that was assayed for activity.

DNA Fragmentation Analysis
HOB-01–09 stably transfected cells were plated as for the AP activity assay and incubated in BSA-medium at a nonpermissive temperature of 39 C for 48 h. At the end of this incubation, the extent of cellular apoptosis was assessed by measuring DNA fragmentation using a cell death detection ELISAplus kit (Roche Clinical Laboratories, Indianapolis, IN) as described by the manufacturer.

Mechanosensory Stimulation
HOB-01–09 cells were seeded onto BioFlex type I collagen-coated six-well plates and incubated at 34 C for 24 h followed by 39 C for 24 h. The cells were subjected to a strain of 3400 microstrain, 2 Hz, 7200 cycles at 37 C using an FX-3000 Flexercell Strain Unit (Flexercell International, Hillsborough, NC). The conditioned medium was removed 4.5 h later and analyzed for the presence of nitric oxide (NO) using a kit manufactured by R&D Systems (Minneapolis, MN).

Statistical Analysis
Data are presented as means ± SE. Statistical significance was determined using one-way ANOVA or Student’s t test. Results were considered statistically different when P < 0.05.


   FOOTNOTES
 
Disclosure Summary: Yan Liu, Ramesh A. Bhat, Laura M. Seestaller-Wehr, Shoichi Fukayama, Annamarie Mangine, Robert A. Moran, Barry S. Komm, Peter V. N. Bodine, and Julia Billiard are full-time employees of Wyeth Research.

First Published Online November 9, 2006

Abbreviations: AP, Alkaline phosphatase; BSP, bone sialoprotein; CAR, coxsackie adenovirus receptor; C/EBP, CCAAT/enhancer-binding protein; CMV, cytomegalovirus; EGFP, enhanced green fluorescent protein; FBS, fetal bovine serum; ß-gal, ß-galactosidase; ß-GP, ß-glycerophosphate; HEK, human embryonic kidney; MOI, multiplicity of infection; MSC, mesenchymal stem cell; MSCGM, MSC growth medium; PPAR, peroxisome proliferator activated receptor; RT, room temperature; shRNA, short hairpin RNA; siRNA, short interfering RNA; SV40, simian virus 40.

Received for publication August 18, 2006. Accepted for publication November 2, 2006.


   REFERENCES
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 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
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