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Endocrinology Vol. 139, No. 4 1952-1964
Copyright © 1998 by The Endocrine Society

ARTICLES

Conditionally Immortalized Murine Bone Marrow Stromal Cells Mediate Parathyroid Hormone-Dependent Osteoclastogenesis in Vitro1

B.-Y. Liu, J. Guo, B. Lanske, P. Divieti, H. M. Kronenberg and F. R. Bringhurst

Endocrine Unit, Massachusetts General Hospital, and Harvard Medical School, Boston, Massachusetts 02114

Address all correspondence and requests for reprints to: F. R. Bringhurst, M.D., Endocrine Unit/Wellman 5, Massachusetts General Hospital, Boston, Massachusetts 02114.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
PTH recruits and activates osteoclasts to cause bone resorption. These actions of PTH are thought to be mediated indirectly via type 1 PTH/PTH-related peptide receptors (PTH1Rs) expressed by adjacent marrow stromal or osteoblastic cells, although some evidence suggests that PTH may act directly on early hematopoietic osteoclast progenitors. We have established clonal, conditionally immortalized, PTH-responsive, bone marrow stromal cell lines from mice that harbor both a transgene encoding a temperature-sensitive mutant of the simian virus 40 large T antigen and deletion of a single allele of the PTH1R gene. Of 60 stromal cell lines isolated, 45 expressed functional PTH1Rs. During coculture with normal murine spleen cells, 5 of 42 such cell lines could support formation of tartrate-resistant acid phosphatase-positive, multinucleated cells (TRAP+ MNCs) in response to 1,25-dihydroxyvitamin D3, but only 2 of these did so in response to PTH. One of these, MS1 cells, expressed numerous cytokines and proteins characteristic of the osteogenic lineage and showed increased production of interleukin-6 in response to PTH. MS1 cells supported dose-dependent induction by rat (r) PTH-(1–34) (0.1–100 nM) of TRAP+ MNCs that expressed calcitonin receptors and formed resorption lacunae on dentine slices. This effect of PTH, which required cell to cell contact between MS1 and spleen cells, was mimicked by coadministration of cAMP analog and phorbol ester but only partially by either agent alone. The carboxyl-terminal fragment rPTH-(53–84) also induced osteoclast-like cell formation, but the maximal effect was only 30% as great as that of rPTH-(1–34). Importantly, rPTH-(1–34) induced TRAP+ MNC formation even when PTH1R-/- osteoclast progenitors (from fetal liver of mice homozygous for ablation of the PTH1R gene) were cocultured with MS1 cells. We conclude that activation of PTH1Rs on cells of the osteoclast lineage is not required for PTH-(1–34)-induced osteoclast formation in the presence of appropriate PTH-responsive marrow stromal cells. MS1 cells provide a useful model for further study of PTH regulation of osteoclastogenesis.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
PTH IS A major regulator of mineral ion metabolism and acts on bone and kidney to alter net fluxes of calcium and phosphate into the extracellular fluid. The effects of PTH on bone are complex, in that continuous exposure to high PTH concentrations causes net bone resorption, whereas intermittent exposure to lower concentrations in vivo increases net bone formation (1). The manner in which PTH concurrently and differentially regulates the formation and activity of osteoclasts and osteoblasts in these settings is incompletely understood.

Mature multinucleated osteoclasts arise by fusion of committed precursors that are derived, in turn, from hematopoietic progenitors of the monocyte-macrophage lineage (granulocyte-macrophage colony-forming units) (2, 3). PTH increases osteoclast number, activity, and survival (4, 5), yet mature mammalian osteoclasts seem not to express functional type 1 PTH/PTH-related peptide (PTHrP) receptors (PTH1Rs) (6, 7). Much evidence suggests that PTH-stimulated osteoclastic bone resorption requires interactions between osteoclasts, or their progenitors or precursors, with accessory mesenchymal marrow stromal cells or osteoblasts (2, 6, 8, 9, 10, 11). Thus, the full resorptive effects of PTH on bone generally are believed to require the production by osteoblasts or marrow stromal cells of secreted or membrane-anchored factors, which, in turn, function as downstream effectors to recruit and activate osteoclasts (2, 3, 6, 8, 9, 12). Of these soluble factors, macrophage colony-stimulating factor (M-CSF), interleukin-6 (IL-6), and IL-11 are believed to play critical roles in PTH-mediated osteoclast formation (13, 14, 15, 16). At the same time, evidence from several systems indicates that PTH may exert direct effects on isolated osteoclast progenitors (17, 18, 19, 20). Such findings have raised questions regarding the role(s) of accessory mesenchymal cells in supporting earlier steps in osteoclast differentiation.

The amino-terminus of PTH binds to PTH1Rs expressed on osteoblasts and activates multiple intracellular second messenger signals signals (21, 22). The roles of the PTH1R and the various intracellular messenger signals generated by it in mediating osteoclast formation and activation by PTH are not clearly understood, and available data are somewhat contradictory (11, 20, 23). Certain evidence also points to a role in osteoclastogenesis for peptides derived from the carboxyl-terminal portion of the PTH-(1–84) molecule (18), which presumably exert their effects via receptors distinct from the PTH1R (24). Expression of the PTH1R is not an absolute general requirement for osteoclastogenesis, as osteoclasts still are formed in fetal mice in which PTH1R expression has been ablated by gene targeting (25). The role of this receptor in the various target cells involved in PTH-dependent osteoclastogenesis remains undefined.

Few in vitro model systems are suitable for direct studies of the cellular mechanisms and interactions involved in PTH-induced osteoclastogenesis. Several transformed human and rodent osteoblastic cell lines have been employed to investigate the actions of PTH on mature osteoblasts, including the induction of osteoclast differentiation or activation during cocultures with mixed spleen or bone marrow cells used as sources of osteoclast progenitors (11, 12, 15, 26), but no suitable marrow-derived stromal cell lines are available for studies of PTH-dependent osteoclastogenesis. Moreover, the osteoblastic cell lines used in reported studies are constitutively transformed and thus may not adequately reflect the responses of normal osteoblastic cells in vivo. One approach to this problem, previously employed successfully by others to establish clonal cell lines from bone and bone marrow, has been to derive such cell lines from normal tissues of mice that express a transgene encoding a temperature-sensitive mutant of the simian virus 40 (SV40) viral large T antigen (tsTAg) (27, 28, 29). Such cells may be conveniently isolated under permissive culture conditions and then caused to assume a more differentiated, nonproliferative phenotype at nonpermissive temperatures (27, 28, 29).

To address the role of the PTH1R in osteoclastogenesis, we sought to isolate, from the stromal layer of murine bone marrow cultures, conditionally immortalized clonal cell lines that could support PTH-dependent osteoclastogenesis. To enable future in vitro selection for PTH1R-null subclones, we also wanted to obtain such cells from animals (PTH1R+/-) in which one allele of the PTH1R gene had already been disrupted and replaced with a selectable marker (i.e. neo resistance) (25). We report here the successful isolation of such conditionally immortalized, clonal marrow stromal cells that can support the generation, from cocultured normal spleen cells and in a PTH-dependent manner, of multinucleated cells (MNCs) that express tartrate-resistant acid phosphatase (TRAP) and other features of mature osteoclasts. These cells exhibit an osteogenic phenotype and appear to induce osteoclastogenesis in response to activation of both protein kinase A (PKA)- and protein kinase C (PKC)-dependent signaling pathways. Further, these cells support PTH-(1–34)-stimulated osteoclastogenesis even when the required osteoclast progenitors are derived from mice homozygous for deletion of the PTH1R gene.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Materials
Synthetic rat (r) PTH-(1–34), human (h) PTH-(53–84), and salmon calcitonin (sCT) were obtained from Bachem (Torrance, CA), 1,25-dihydroxyvitamin D3 [1,25-(OH)2D3] was obtained from Biomol (Plymouth Meeting, PA), and recombinant mouse interferon-{gamma} (IFN-{gamma}) was obtained from Genzyme (Boston, MA). Eagle’s MEM (EMEM) and {alpha}MEM were supplied by the Media Kitchen at the Massachusetts General Hospital. FBS, penicillin, and streptomycin were obtained from Life Technologies (Grand Island, NY). Dexamethasone, 8-bromoadenosine cAMP (8-BrcAMP), 12-O-tetradecanoylphorbol 13-acetate (TPA), naphthol AS-BI phosphoric acid, and red violet LB salt were products of Sigma Chemical Co. (St. Louis, MO). All other chemicals used were of analytical grade. Transwell culture inserts (6.5 mm in diameter) with membrane filters (0.4 µm) and 24-well culture plates were purchased from Becton Dickson Labware (Franklin Lakes, NJ), and multichambered slides were obtained from Lab-Tek (Naperville, IL).

Animals
Transgenic mice were derived from matings of H-2Kb-tsA58 transgenic mice (Immortomouse, Charles River, Wilmington, MA) with mice heterozygous for ablation of the PTH1R gene. Genotypes of mice were confirmed by Southern blot or PCR of total DNA, using probes (or primers) specific for the PTH1R, the PTH1R knock-out allele, and the SV40 TAg (25). C57BL/6 mice were obtained from Jackson Laboratories (Bar Harbor, ME). Animals were maintained in facilities operated by the Office of Laboratory Animal Research of the Massachusetts General Hospital in accordance with the NIH Guide for the Care and Use of Laboratory Animals and were employed using protocols approved by the institution’s subcommittee on animal care.

Establishment of bone marrow stromal cell lines
Bone marrow stromal cells were obtained from the femurs and tibias of 4- to 6-week-old H-2KbtsA58 transgenic/PTH1R+/- mice. The mice were killed by carbon dioxide asphyxiation and cervical dislocation, the femurs and tibias were dissected free of adherent soft tissue, the metaphyses were removed, and the marrow was flushed from the marrow cavities with EMEM via a 23-gauge needle. The marrow cells from one lower limb were washed twice and cultured in 25-cm2 flasks with EMEM supplemented with 10% heat-inactivated FBS, 100 U/ml penicillin, and 100 mg/ml streptomycin at 33 C in a humidified atmosphere of 5% CO2 in air. Because the tsTAg gene expressed by these mouse cells was under the control of a IFN-{gamma}-inducible H-2Kb promoter, 5 U/ml mouse IFN-{gamma} were added to the culture medium to maximize proliferation. After incubation for 3 days, the cells were trypsinized and replated in new flasks at a density of 1 x 104 cells/cm2. The cultures were fed fresh medium every 3–4 days and subcultured by trypsinization every 10–14 days. After the third passage, the cells were plated at a cloning density of 100 cells/10-cm dish. After 4 weeks of further cultivation, individual colonies became apparent and were isolated by direct aspiration with a pipette.

To study growth, stromal cells were inoculated in 35-mm culture dishes under permissive conditions (33 C in the presence of IFN-{gamma}), semipermissive conditions (37 C in the absence of IFN-{gamma}), and nonpermissive conditions (39.5 C in the absence of IFN-{gamma}) in EMEM with 10% heat-inactivated FBS. At the appropriate times, cells were washed twice with PBS, trypsinized, and counted with a hemocytometer.

cAMP measurements
Confluent clonal stromal cells in 24-well plates were washed with assay buffer (135 mM NaCl, 6 mM KCl, 1 mM MgCl2, 2.8 mM glucose, 1.2 mM CaCl2, and 20 mM HEPES, pH 7.4) and incubated with the same buffer containing 0.1% heat-inactivated BSA, 1 mM isobutylmethylxanthine, and agonist or vehicle at 37 C for 15 min The buffer then was rapidly aspirated, the plates were immediately frozen in liquid nitrogen for 1 min, and the frozen cells were subsequently thawed directly into 0.5 ml 50 mM HCl. Total cellular cAMP in the acidic extracts was measured using a commercial RIA kit (New England Nuclear Corp., Boston, MA). Results were expressed as picomoles of cAMP produced per well over 15 min.

PTH/PTHrP receptor binding
Confluent clonal stromal cells in 24-well plates were washed twice with 0.5 ml binding buffer (100 mM NaCl, 5 mM KCl, 2 mM CaCl2, and 50 mM Tris-HCl, pH 7.8) before incubation with 125I-labeled [Tyr36]hPTHrP-(1–36)NH2 (100,000 cpm/well), prepared by the chloramine-T method and purified by HPLC as previously described (21), in 0.5 ml complete buffer (binding buffer plus 5% heated-inactivated horse serum) with or without competing rPTH-(1–34) ligand at 15 C for 4 h. After incubation, cells were washed four times with cold binding buffer, dissolved in 0.5 ml 0.1% SDS, and aliquoted for measurement of cell-associated radioactivity.

Inositol phosphate accumulation
Stromal cells in six-well dishes were incubated for 24 h with 3 mCi/ml [3H]myo-inositol (Amersham Corp., Arlington Heights, IL) in inositol-free DMEM-Ham’s F-12 medium supplemented with 10% heat-inactivated FBS before incubation for 40 min with rPTH-(1–34) or vehicle alone, added in fresh inositol-free medium containing 30 mM LiCl. Water-soluble inositol polyphosphates were isolated and separated by ion exchange chromatography, as described previously (30).

TRAP+ MNC formation
Stromal cells (4 x 104 cells/well) were plated in 24-well plates and cultured for 24 h before being overlaid with spleen cells (106 cells/well) from 8- to 11-week-old C57BL/6 male mice. Cells were cultured in 0.5 ml {alpha}MEM supplemented with 10% heat-inactivated FBS and 10-7 M dexamethasone with or without 1,25-(OH)2D3 (10-8 M), various fragments of PTH (10-7-10-11 M), 8-BrcAMP (10-7-10-3 M), or TPA (10-6-10-10 M) at 37 C for 3 weeks. All cultures were refed by half-changes of fresh medium every 2 days. The cells then were fixed in ethanol-acetone (50:50, vol/vol) and stained for TRAP. TRAP+ cells containing 3 or more nuclei were scored as osteoclast-like MNCs (TRAP+ MNCs). Cells were counted at x20 magnification in 30 contiguous fields along 2 orthogonal pathways in each well, a method previously employed to account for the nonuniform distribution of cells within wells (31). The number of TRAP+ MNCs contained in these 30 fields (representing approximately 0.25 cm2, or one eighth, of the surface area of the well) was expressed as the number per well. To examine the importance in osteoclast development of cell to cell contact between MS1 cells and osteoclast progenitors, cell culture inserts with a membrane filter (0.4 µm) were placed in each well of a 24-well plate, and the spleen cells and stromal cells were separated by the membrane filter. The cultures were refed every 2 days for 3 weeks, as described above, until TRAP staining was performed.

Cocultures of cloned marrow stromal cells and fetal mouse liver
Mice heterozygous for the PTH/PTHrP receptor gene deletion were interbred to obtain homozygous mice, i.e. PTH1R-/-. Embryos of 15.5 days gestation were obtained by cesarean section, and genotyping of the embryos was performed subsequently by PCR. Fetal livers were excised from 8 PTH1R-/- and 10 heterozygous (PTH1R+/-) embryos and dissected into fragments of approximately 1 mm3. One fetal liver fragment per well was placed on an MS1 cell layer that had been plated 1 day previously at a density of 4 x 104 cells/well in a 24-well plate. These cocultures were maintained and refed as described above.

Enzyme histochemistry
After being cultured for the indicated times (usually 21 days), cells were stained for TRAP after washing with PBS, air-drying, and fixing in ethanol-acetone (50:50, vol/vol) for 1 min. The cells then were stained for TRAP by incubating for 1 h at 37 C in 0.1 M sodium acetate buffer (pH 5.2) containing naphthol AS-BI phosphate as a substrate and red violet LB salt as a stain for the reaction product in the presence of 10 mM sodium tartrate. TRAP-positive MNCs containing three or more nuclei were scored as osteoclast-like MNCs.

Demonstration of CT receptors
Stromal cells and spleen cells were cocultured on eight-well multichambered slides for 3 weeks before being washed with binding buffer and incubated with [125I]sCT (0.2 nM; 106 cpm/ml) at room temperature for 1 h in complete binding buffer (binding buffer plus 5% heated-inactivated horse serum). CT was iodinated by the chloramine-T method. Nonspecific binding was assessed on parallel slides in the presence of 300 nM sCT. After this incubation, the slides were fixed and stained for TRAP as described above, dipped in Kodak NTB-2 nuclear emulsion (Eastman Kodak, Rochester, NY), and stored at 4 C for 3 weeks before developing for autoradiography.

Bone resorption pit formation assay
Human dentine slices were supplied by Dr. J. T. Wang, National Taiwan University (Taiwan, Republic of China). Dentine slices (4 x 4 mm, 130–180 µm thick) were prepared with an ethanol-cooled, low speed diamond saw (Isomet, Buehler Co., Lake Bluff, IL). The slices were cleaned twice by ultrasonication in 70% ethanol for 5 min, rinsed twice in distilled water after each sonication, dried under UV light for 1 h on each side, degassed under vacuum for 24 h, and incubated in culture medium overnight before use. Stromal cells were seeded on the dentine slices in 48-well plates at a density of 2 x 104 cells/well and cultured for 24 h. Then, 5 x 105 primary mouse spleen cells prepared from 8- to 11-week-old mice were overlaid on the dentine slices in 0.4 ml culture medium and refed as described above. Cultures were continued in the presence of 1,25-(OH)2D3, rPTH-(1–34), or vehicle alone for 2–3 weeks. At the end of the culture period, the slices were washed twice with distilled water, and the cells were removed from the slices by immersion in bleach solution (6% NaOCl-5.2% NaCl) for 10 min. The slices were etched in 5% (wt/vol) aluminum sulfate for 10 min, rinsed with distilled water, stained with Coomassie brilliant blue (0.5% in 45% methanol-9% acetic acid) for 2 min, rinsed in water, and photographed using an Olympus BH2 microscope (Olympus Corp., New Hyde Park, NY). For some experiments, cocultures were established on multiwell slides coated with a film of calcium phosphate (Osteologic MultiTest, Millenium Biologix, Kingston, Canada), which subsequently were processed according to the manufacturer’s recommendations and viewed after von Kossa staining.

Alkaline phosphatase (ALP) staining
Cytochemical staining of ALP was performed on cells grown to confluence at 33 C and then incubated at 33, 37, or 39.5 C for an additional 3–7 days. The cells were washed with PBS, fixed in 10% formalin in acetone at room temperature, rinsed with distilled water, reacted with naphthol AS-MX for 15 min at room temperature (Sigma, 86-R), rinsed again with water, and then stained with hematoxylin. Positive ALP staining was revealed as a red-violet color.

Bone nodule formation
Cells were seeded at 5 x 105 cells/well in six-well plates and grown to confluence at 33 C. Thereafter, medium was changed to mineralization medium ({alpha}MEM containing 10% heat-inactivated FBS, 1% penicillin-streptomycin, and 10 mM ß-glycerophosphate). For the assessment of effects of glucocorticoid and hormone on mineralization, dexamethasone (10-11-10-7 M), ascorbic acid (50 mg/ml), or rPTH-(1–34) (10-11-10-7 M) were added separately or in combination. Cultures then were returned to 33 C or transferred to 37 C and maintained with half-changes of medium twice a week. At the end of the treatment, the cultures were stained with a modified von Kossa method to assess the formation of mineralized nodules.

Immunocytochemistry for the SV40 tsTAg
Cells were seeded at 2.5 x 104/cm2 into multichambered slides and allowed to proliferate for 3 days under permissive conditions. The medium was changed, and the cells were incubated under either permissive or nonpermissive conditions for an additional 3–14 days. Immunocytochemistry for the SV40 TAg was performed using a mouse monoclonal antibody directed against the wild-type TAg (Pab 101, Santa Cruz Biotechnology, Santa Cruz, CA) according to the manufacturer’s suggestion, with modification. Briefly, the cells were washed with cold calcium/magnesium-free PBS, fixed with methanol at -20 C, air-dried, and rinsed with PBS. The fixed cells then were blocked with 1.5% (vol/vol) rabbit blocking serum in PBS at room temperature, rinsed with PBS containing 1% (wt/vol) BSA (PBS-BSA), and incubated for 30 min with 1 mg/ml specific antibody in PBS-BSA or with the same concentration of a nonspecific mouse IgG. UMR106–01 cells cultivated at 37 C were employed as a negative control. After washing three times with PBS, the cells were incubated with a biotin-conjugated rabbit antimouse IgG for 30 min in 1.5% blocking serum-PBS, rinsed with PBS, and subsequently incubated for 30 min with avidin-biotin enzyme reagent (all from Santa Cruz). The cells were further washed with PBS followed by 0.5% Triton X-100 in PBS before incubation with 0.01% (wt/vol) 3,3-diaminobenzoidine and a few drops of 30% H2O2 in PBS for 2–5 min. Finally, the cells then were washed extensively with distilled water, air-dried, and photographed using an Olympus BH2 microscope.

IL-6 production
Cells were cultured on 24-well plates and grown to confluence under permissive conditions before transfer to EMEM with 0.5% FBS. They were refed with the same medium 24 h later, and experiments were initiated by the addition of rPTH-(1–34) after an additional 24 h of culture. IL-6 concentrations were measured in cell-free supernatants collected 1, 2, 4, 6, 12, or 24 h after the addition of rPTH-(1–34) (10-7 M), using a commercially available enzyme-linked immunosorbant assay (ELISA kit, Genzyme, Cambridge, MA) according to the manufacturer’s recommendation. The sensitivity of this assay is less than 5 pg/ml.

RT-PCR analysis of messenger RNA (mRNA) expression
RT-PCR was performed on total RNA extracted from MS1 cells by the acid-guanidinium thiocyanate-phenol-chloroform method, using a GeneAmp RNA/PCR kit (Perkin-Elmer, Norwalk, CT) with a Peltier Thermal Cycler (MJ Research, Watertown, MA). The oligonucleotides specific for glyceraldehyde-3-phosphate dehydrogenase, various cytokines, osteopontin, osteocalcin, and type I collagen {alpha}1 were synthesized by the Biopolymer Facility at Massachusetts General Hospital. The sequences of these oligonucleotides are described in Table 1Go. RNA primed with specific oligonucleotides was reverse transcribed using cloned Moloney leukemia virus reverse transcriptase (200 U), 1 mM of each deoxy-NTP, and 5 U ribonuclease inhibitor in a final volume of 50 µl. Controls were included in which the reverse transcriptase or the RNA template was omitted. The reaction was run at room temperature for 10 min before the temperature was raised to 42 C for 30 min to complete the extension reaction. The reaction mixture then was heated to 95 C for 5 min to denature the resulting RNA-complementary DNA hybrids and quickly chilled on ice. Subsequent PCR was performed using pairs of specific primers for each gene product. The PCR conditions were as follows: 1 min at 94 C for denaturation, 1 min at 54 C for primer annealing, and 2 min at 72 C for primer extension for 35 cycles, then a final 10-min extension at 72 C. The PCR products were electrophoresed in 1.5% agarose gels and visualized using ethidium bromide staining.


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  		Table 1. Oligonucleotide primers used for PCR analysis

 
Statistical analysis
Results were expressed as the mean ± SD for groups of separate culture wells. Significance of differences was analyzed by ANOVA, followed by Bonferroni correction of t tests for multiple comparisons against the same control group. Differences between treatments were analyzed using the Student-Newman-Keuls test.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
Conditionally immortalized marrow stromal cell lines were isolated from 4-week-old mice that were the progeny of matings between H-2KbtsA58 transgenic mice and mice that retained only one functional allele of the PTH1R gene. Adherent marrow cells were seeded at low density and maintained in growth medium for 4–6 weeks, after which individual colonies were isolated and subsequently subcloned. To identify PTH-responsive subclones, cell lines first were screened for evidence of increased cAMP accumulation in response to rPTH-(1–34) (100 nM). Forty-five of 60 clones tested exhibited such cAMP responsiveness to PTH (range, 5- to 100-fold over the basal value), whereupon the expression of functional PTH1Rs was confirmed by demonstration of specific binding of 125I-labeled [Tyr36]hPTHrP-(1–36)NH2 (not shown).

TRAP+ MNC formation in cocultures of marrow stromal cell lines and normal mouse spleen cells
Of 42 clonal stromal cell lines that expressed functional PTH1Rs, only 5 could support TRAP+ MNC formation in response to 1,25-(OH)2D3 or PTH when cocultured for 3 weeks with normal mouse spleen cells. All 5 of these cell lines supported 1,25-(OH)2D3-dependent TRAP+ MNC formation in this assay, whereas only two, MS1 and MS2, supported TRAP+ MNC formation in response to rPTH-(1–34) (100 nM; Fig. 1Go). The maximal responses to PTH in cocultures involving MS1 cells were in the range of 80–120 TRAP+ MNCs/30 high power fields (~0.25 cm2; see Materials and Methods), whereas few (i.e. <10) TRAP+ MNCs were observed in control cocultures in the absence of 1,25-(OH)2D3 or PTH. No TRAP+ MNCs were observed in cultures of spleen cells or stromal cells alone or when cocultures were conducted in the absence of dexamethasone. When cocultures were maintained at the permissive temperature (i.e. 33 C), few TRAP+ MNCs were observed, even in the presence of 1,25-(OH)2D3 or PTH. In cocultures conducted at the fully nonpermissive temperature (39.5 C), few spleen cells survived for the duration of the experiments (3 weeks), and no TRAP+ MNCs were formed. Accordingly, subsequent coculture experiments were performed at 37 C.



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  		Figure 1. Formation of TRAP+ MNCs in cocultures of clonal marrow stromal cells and normal spleen cells. Clonal marrow stromal cells and normal murine spleen cells were cocultured in the presence of rat PTH-(1–34) (100 nM; PTH), 1,25-(OH)2D3 (10 nM; Vit D), or vehicle alone (Control) for 21 days before staining for TRAP and enumeration of TRAP+ MNCs, as described in Materials and Methods. A, Appearance of TRAP+ MNCs formed in response to PTH (x400 magnification). B, Induction of TRAP+ MNCs by PTH and 1,25-(OH)2D3 in MS1 and MS2 cells. Each bar represents the mean ± SD of the number of TRAP+ MNC enumerated in each of 12 replicate cocultures (note that only about 12% of the surface of each well was examined; see Materials and Methods). In each cell line, both PTH- and vitamin D-treated groups differed significantly from controls (P < 0.01).

 
In further studies with MS1 cells, the TRAP+ MNCs formed during MS1/spleen cell cocultures were evaluated for expression of CT receptors and for their ability to resorb mineralized substrate, two specific features of osteoclasts (2, 3). As shown in Fig. 2Go, specific binding of [125I]sCT, fully displaced by 300 nM nonradioactive sCT, was localized over TRAP+ MNCs (and mononuclear cells) in both 1,25-(OH)2D3- and PTH-treated cocultures. These TRAP+ MNCs also resorbed bone in response to PTH or 1,25-(OH)2D3, as demonstrated by the appearance of numerous resorption lacunae during cocultures conducted on dentine slices (Fig. 3Go) or on slides coated with calcium phosphate (Osteologic; not shown). Resorption lacunae rarely were observed on dentine slices cultured with either MS1 cells or spleen cells alone or with MS1/spleen cell cocultures in the absence of the hormones. A requirement for cell-cell contact between MS1 and mouse spleen cells was demonstrated when no TRAP+ MNC cells were formed in response to 1,25-(OH)2D3 or PTH in cocultures in which spleen cells were seeded inside transwell inserts and MS1 cells were plated in the outer wells (or vice versa; not shown).



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  		Figure 2. Autoradiography of [125I]sCT binding to cocultures of MS1 and spleen cells. MS1 and spleen cells were cocultured on multichambered slides for 21 days in the presence of rPTH-(1–34) (100 nM). Cells then were incubated with [125I]sCT at room temperature for 1 h in either the absence (A) or presence (B) of excess nonradioactive sCT (300 nM), stained for TRAP, and processed for autoradiography as described in Materials and Methods (x400 magnification).

 


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  		Figure 3. Resorption of dentine by TRAP+ MNCs formed in MS1/spleen cell cocultures. Cocultures were conducted as described in Fig. 1Go, except that cells were deposited onto slices of human dentine (see Materials and Methods). A, Resorption lacunae are visible in Coomassie-stained dentine, after removal of adherent cells, from a culture treated with rPTH-(1-34) (100 nM) (x400 magnification). B, Dentine slice from untreated control coculture (x100 magnification).

 
In PTH-treated MS1/spleen cell cocultures, TRAP+ MNCs first appeared at 5 days, reached a peak at 21 days, and then rapidly declined in number (Fig. 4AGo). Rat PTH-(1–34) induced formation of TRAP+ MNCs in a dose-dependent manner, with a minimally effective concentration of 0.1 nM, after 21 days of coculture (Fig. 4BGo). Although 1,25-(OH)2D3 and PTH are believed to stimulate osteoclast formation by different mechanisms and to exert interactive effects in some systems (32), addition of 1,25-(OH)2D3 (0.01 nM to 10 nM) together with PTH at 10-fold higher concentrations (i.e. 0.1–100 nM) did not lead to any additive effects in MS1/spleen cell cocultures (data not shown).



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  		Figure 4. Time course and concentration dependence of PTH-induced TRAP+ MNC formation. MS1 and spleen cells were incubated for the indicated number of days with 100 nM rPTH-(1–34) (A) or for 21 days with the indicated concentrations of rPTH-(1–34) (B) before staining for TRAP and enumeration of TRAP+ MNCs. Data are expressed as the mean ± SD of six cultures. *, Significantly different from vehicle-treated controls (P < 0.05)

 
Phenotypic characteristics of MS1 stromal cells
MS1 cells grew rapidly under permissive conditions (33 C), with a doubling time of approximately 40 h. Growth ceased within 3–5 days of transfer to 37 C, however, and none occurred after 1–2 days at 39.5 C. Expression of SV40 TAg, readily detected by immunocytochemistry in nuclei of cells maintained at 33 C, also was greatly reduced within several days of transfer to 39.5 C, and cell viability, as assessed by trypan blue exclusion, declined from more than 90% at 33 C to approximately 60% within 1 week of culture at 39.5 C. MS1 cells have maintained the capacity to support TRAP+ MNC formation in cocultures with normal spleen cells despite continuous passage and culture at permissive conditions for more than 2 yr.

Expression of ALP (by cytochemical analysis) was uniform among MS1 cells incubated at 33 or 37 C and was not influenced by exposure to dexamethasone (10-7 M) or ß-glycerophosphate (10 mM) for up to 7 days. In the presence of ß-glycerophosphate (10 mM), MS1 cells formed mineralized nodules (von Kossa staining) within 3 weeks at 37 C (but not at 33 C). Nodule formation was accelerated (to within 10 days) in a dose-dependent manner by dexamethasone (0.01–100 nM; Fig. 5Go) and was not observed in the absence of ß-glycerophosphate. MS1 cultures did not stain with Alcian blue and showed no evidence of adipocytic differentiation, as assessed by microscopy and Sudan black staining, even after prolonged (>4 week) exposure to dexamethasone (10-7 M) at 37 C.



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  		Figure 5. Mineralization of MS1 cultures in the presence of dexamethasone. MS1 cells were maintained at 37 C for 5 days (A) or 10 days (B) in either unsupplemented {alpha}MEM (a) or with added ß-glycerophosphate (10 mM) plus dexamethasone at concentrations of 10-11 M (b), 10-10 M (c), 10-9 M (d), 10-8 M (e), and 10-7 M(f) before von Kossa staining.

 
MS1 cells expressed mRNAs for several bone matrix proteins, including type I collagen and the noncollagenous proteins osteopontin and osteocalcin, as detected by RT-PCR (Fig. 6Go). mRNAs for M-CSF, IL-1{alpha}, IL-6, tumor necrosis factor-{alpha}, IL-6 receptor, and IL-11 receptor also were readily detected in cells grown at permissive or semipermissive conditions, with or without prior hormone treatment. By this semiquantitative method, no evident hormonal regulation of these transcripts was noted. Increased secretion of IL-6 protein by MS1 cells was detected within 1 h and peaked by 4 h after the addition of rPTH-(1–34) (0.1–100 nM; not shown).



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  		Figure 6. Expression of mRNA for bone matrix proteins, cytokines, and cytokine receptors by MS1 marrow stromal cells. Total RNA isolated from MS1 cells maintained for 7 days at 37 C was analyzed by RT-PCR for the presence of specific mRNAs, using the primers described in Table 1Go (see Materials and Methods). Control PCR reactions (not shown), performed in the absence of RNA or by using the same samples but omitting the reverse transcriptase, showed no specific bands. The sizes of the mol wt markers (base pairs) applied to the lateral lanes are shown.

 
PTH1R expression and function in MS1 cells
PTH1R expression by MS1 cells was further characterized by radioligand binding analysis, using iodinated [Tyr36]hPTHrP-(1–36)NH2 as radioligand. As shown in Fig. 7AGo, binding of hPTHrP-(1–36) to confluent monolayers of MS1 cells was specifically displaced by increasing concentrations of rPTH-(1–34). Scatchard analysis demonstrated a single class of binding sites (~50,000/cell) with an apparent dissociation constant (Kd) of approximately 2 nM. Rat PTH-(1–34) (0.01–100 nM) elicited dose-dependent stimulation of intracellular cAMP (to a maximum of 60-fold over basal values) with an EC50 of approximately 0.1 nM (Fig. 7BGo). No changes in inositol polyphosphate production could be detected in response to as much as 1000 nM rPTH-(1–34), however, in either MS1 cells or nine other PTH1R-expressing clonal stromal cell lines tested. Treatment with dexamethasone (10 nM) or ß-glycerophosphate (10 mM) for 3 days did not alter PTH-stimulated cAMP production (not shown).



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  		Figure 7. PTH1R expression and cAMP responsiveness in MS1 cells. A, MS1 cells were incubated with 125I-labeled [Tyr34]hPTHrP-(1–36)amide (140,000 cpm/well) in the presence of rPTH-(1–34) at the indicated concentrations for 6 h at 15 C before measurement of cell-associated radioactivity, as described in Materials and Methods. Each point is the mean ± SD of specific binding for duplicate wells, expressed as a percentage of maximal binding in the absence of added PTH (2.6% of added counts per min). Nonspecific binding in the presence of 1000 nM rPTH was 0.5% of the added radioactivity. (the inset shows Scatchard analysis, where the ordinate is the bound/free ratio and the abscissa is specific binding). B, MS1 cells were incubated with rPTH-(1–34) at the concentrations shown for 15 min at 37 C in the presence of isobutylmethylxanthine (IBMX; 1 mM) before extraction of the cell layer for measurement of cAMP by RIA, as described in Materials and Methods. Values shown are the mean ± SD, expressed as picomoles per well.

 
To examine the possible roles of the two major PTH1R-activated intracellular signaling pathways in stimulating osteoclast formation, MS1/spleen cell cocultures were treated with 8-BrcAMP and TPA, pharmacological activators of PKA and PKC, respectively. As shown in Table 2Go, TRAP+ MNC formation was increased dose dependently in response to either 8-BrcAMP or TPA alone, but addition of both drugs (at optimal concentrations of 0.01 mM and 10 nM, respectively) was required to fully mimic the response to rPTH-(1–34). At higher concentrations, combinations of these agents exerted microscopically evident cytotoxic effects.


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  		Table 2. Effects of 8-BrcAMP and TPA on TRAP+ MNC formation in MS1/spleen cell cocultures

 
To determine whether carboxyl-terminal PTH fragments previously reported to support formation of osteoclast-like cells from murine marrow progenitors PTHR (18) might be active in the system described here, MS-1/spleen cell cocultures were treated for 3 weeks with hPTH-(53–84). As shown in Table 3Go, hPTH-(53–84) modestly stimulated TRAP+ MNC formation in a dose-dependent manner, but at the highest concentration tested (1000 nM), the response was less than 30% of that seen with the N-terminal fragment rPTH-(1–34) (cf. Figs. 1Go and 4Go and Table 2Go). The hPTH-(53–84) fragment (1000 nM) did not increase cAMP accumulation in MS1 cells (not shown).


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  		Table 3. Effects of carboxyl-terminal fragment hPTH-(53-84) on TRAP+ MNC formation in MS1/spleen cell cocultures

 
Role of PTH1R expression by osteoclast progenitors in PTH-mediated osteoclastogenesis
Evidence of PTH binding to osteoclast progenitors (17) and of direct actions of N-terminal PTH peptides to cause formation of osteoclast-like cells from putative osteoclast progenitors in certain in vitro systems (17, 19, 20) has indicated that PTH-(1–34) may act directly on such cells, possibly by activating PTH1Rs, to induce osteoclast formation and bone resorption independent of its effects on stromal cells or osteoblasts. We approached this issue directly by coculturing the PTH1R-expressing MS1 cells with osteoclast precursors obtained from mice in which both alleles of the PTH1R gene had been deleted (i.e. PTH1R-/- mice). Because homozygous ablation of the PTH1R gene is neonatally lethal (25), spleen or marrow could not readily be employed as a source of osteoclast progenitors from such animals. Instead, we employed fetal liver as an alternate source of osteoclast precursors (33, 34). For these experiments, MS1 stromal cells were cocultured with fragments of liver removed from 15.5-day-old fetuses that subsequently were determined to be either heterozygous or homozygous for ablation of the PTH1R gene. In each case, formation of TRAP+ MNCs was analyzed after the addition of 1,25-(OH)2D3 (10 nM), rPTH-(1–34) (100 nM), or vehicle alone. Liver tissue from a total of 18 different 15.5-day-old fetuses (10 heterozygotes and 8 homozygotes) was used for these studies.

As shown in Table 4Go, TRAP+ MNCs were formed in response to rPTH-(1–34) regardless of whether fetal liver fragments from heterozygous PTH1R+/- or homozygous PTH1R-/- mice were cocultured with MS1 stromal cells. The numbers of TRAP+ MNCs formed in these fetal liver cocultures were much lower than those obtained in cocultures with dispersed spleen cells, but the responses to PTH were comparable to those induced by 1,25-(OH)2D3 (10 nM). In fact, the numbers of TRAP+ MNCs formed in MS-1 cell/fetal liver cocultures were the same in response to rPTH-(1–34) [or 1,25-(OH)2D3] regardless of the genotype of the liver tissue used as the source of osteoclast progenitors. All treatment groups differed significantly from the corresponding controls incubated in the absence of PTH or 1,25-(OH)2D3, in which TRAP+ MNCs rarely appeared (Table 4Go). Subsequent autoradiographic studies demonstrated that the TRAP+ MNCs generated from the fetal liver cocultures did express CT receptors (data not shown).


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  		Table 4. TRAP+ MNC formation in cocultures of fetal livers and MS1 cells

 

   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Osteoblasts and bone marrow stromal cells are regarded as essential accessory cells in osteoclastogenesis (2, 6, 8, 9, 10, 11). Several marrow stromal cell lines support 1,25-(OH)2D3-induced osteoclast development in vitro (9, 28, 35), but none are reported to do so in response to PTH. To facilitate direct study of PTH-induced osteoclast formation in vitro, we sought to establish conditionally transformed clonal marrow stromal cell lines that could mediate PTH-dependent osteoclast differentiation from hematopoietic precursors in spleen or marrow. The MS1 cells we isolated, when cocultured with murine spleen cells, supported PTH-dependent formation of multinucleated cells that exhibited major functional criteria for osteoclasts: i.e. they were TRAP+, expressed calcitonin receptors, and formed resorption lacunae on dentine slices. Moreover, cell-cell contact with MS1 cells was absolutely required for osteoclast formation, as has been reported for other systems (2, 8, 9, 10, 36).

We observed basal PTH1R expression in most (75%) of the unselected cell lines we initially isolated from murine marrow. In contrast, among 5 (of 120) marrow stromal cell lines found to support 1,25-(OH)2D3-dependent osteoclast formation, Feuerbach et al. detected no PTH-dependent cAMP production unless the cells were pretreated with osteogenic protein-1 (28). The explanation for this disparity in the frequency of basal PTH1R expression is unclear, but could relate to differences in transgene constructs, animal strains, or culture conditions or to the fact that the animals we used were heterozygous also for ablation of the PTH1R gene, which may somehow have altered the stromal cell composition of their marrow.

Of the stromal cell lines we isolated, only a few (12%) were competent to support osteoclastogenesis in response to 1,25-(OH)2D3 during cocultures with normal spleen cells, and only a subset of these (i.e. 5% of all of the PTH1R-expressing cell lines tested) could do so in response to PTH as well. The marrow cell lines we isolated were not subjected to any intentional selection in vitro, and their proliferation was dependent upon expression of the tsA58 transgene that was driven, in turn, by the ubiquitously active MHC II promoter. Thus, the 60 clones we isolated were expected to constitute a representative sampling of the marrow stromal cell population. Our findings are not dissimilar to those of Chambers et al. (27) and Feuerbach et al. (28), who found that 30% and 4%, respectively, of the cell lines they isolated could support 1,25-(OH)2D3-dependent osteoclastogenesis. Collectively, these results indicate that only a minor fraction of the murine marrow stromal population normally is competent to mediate osteoclast differentiation in response to osteotropic hormones.

Our results also indicate that neither PTH1R expression per se nor the particular repertoire of stromal cell responses required for 1,25-(OH)2D3-dependent osteoclastogenesis is sufficient to enable a marrow stromal cell to support PTH-induced osteoclast formation. Thus, three of our clonal stromal cell lines could mediate osteoclast formation in response to 1,25-(OH)2D3, but not PTH (despite expression of PTH1Rs), and in cocultures involving MS1 cells, the effects of PTH and 1,25-(OH)2D3 were not additive. These findings suggest that PTH may engage some or all of the cellular responses that mediate 1,25-(OH)2D3-dependent osteoclastogenesis, but that additional events must be required for the PTH response. The number of PTH1Rs expressed per cell, which was not measured in all of our cell lines, may modulate the overall signaling response to the hormone (30) and thus could contribute to these differences in accessory cell function among PTH1R-expressing cells. More likely, differences in other stromal cell characteristics, including expression or secretion of particular cytokines, critical integrins, cytokine receptors, other cell surface molecules, or unknown soluble mediators (6, 15), may underlie the unique functional phenotypes represented by MS1 and MS2 cells. MS1 cells did express mRNAs encoding several cytokines previously implicated in osteoclastogenesis, including M-CSF, IL-1{alpha}, IL-6, IL-11, and tumor necrosis factor-{alpha} (2, 3, 13, 14, 16). Among these, secretion of IL-6 protein by MS1 cells was up-regulated by PTH, as has been reported in other systems (2, 15). In this regard, it is of interest that, like preadipocytic ST2 and MC3T3-G2/PA6 cells (9), the osteoclastogenic activity of MS1 cells was glucocorticoid dependent, as Udagawa and colleagues recently have shown that glucocorticoid induction of IL-6 receptor {alpha}-subunit expression by murine osteoblastic cells is required for an osteoclastic response in marrow or spleen cell cocultures (37).

We and others (28) observed no stromal cell-dependent osteoclastogenesis unless the tsTAg was inactivated, which suggests that these conditionally transformed stromal cells must assume a more differentiated, nontransformed phenotype to effectively support osteoclastogenesis. MS1 cells seem to be committed to the osteoblastic lineage, as they express ALP, type I collagen, and the noncollagenous bone matrix proteins, osteopontin and osteocalcin. Thus, they may be related to the osteogenic subtype(s) of marrow stromal cells previously described in primary isolates of marrow cells or in colony-forming assays of so-called colony-forming unit-fibroblastic (38, 39, 40, 41). Their osteogenic character also was evidenced by the formation of mineralized nodules during prolonged culture at semipermissive conditions. The up-regulation of MS1 cell mineralization by glucocorticoid is of interest. Unlike rat, rabbit, or human marrow or osteoblastic cells, murine osteogenic cells, in the few studies that have addressed this issue, are reported not to require or even to be inhibited by glucocorticoids (42, 43, 44). The effects of dexamethasone appear to be complex, however, and may depend upon the stage of osteoblastic differentiation as well as the basal production of inhibitory cytokines, the secretion of which is inhibited by glucocorticoids (45, 46). In any event, this effect of dexamethasone suggests that MS1 cells may represent a subpopulation of osteogenic cells distinct from those that ordinarily give rise to mineralized nodules in mixed primary murine marrow cultures.

Previous reports have implicated the cAMP/PKA pathway as the primary signaling cascade responsible for the osteoclastogenic effects of PTH (11, 15, 20, 26). In MS1/spleen cell cocultures, TRAP+ MNC formation was induced by direct pharmacological activation of either PKA or PKC, but the maximal response fell short of that observed with rPTH-(1–34) unless both 8-BrcAMP and TPA were added together. These results could indicate that both signaling pathways must be activated by the PTH1R to elicit the full osteoclastogenic response. On the other hand, other cells within the heterogeneous spleen cell population may have served as the targets of one or both of these drugs in these experiments. Thus, determination of the roles of specific PTH1R signals generated within MS1 cells in mediating the osteoclastic response will require further study. This analysis should be facilitated by efforts, ongoing in our laboratory, to isolate PTH1R-/- subclones of MS1 cells in which responses mediated by transfected signal-selective PTH1Rs (47) could be directly analyzed.

It is widely believed that the osteoclastogenic effects of PTH are mediated indirectly via its actions on accessory stromal or osteoblastic cells, a view consistent with the absence of PTH binding to mature mammalian osteoclasts, the requirement for cell-cell contact, and the detection of soluble factors released by osteoblasts in response to PTH (2, 3, 6, 8, 9, 10, 11, 36). Recent studies have implicated direct actions of PTH on isolated osteoclast progenitors, however (17, 18, 20, 26), which could be mediated by the PTH1R or by alternate receptor species, such as the type 2 PTH receptor or PTH receptors with specificity for C-terminal regions of the ligand. Indeed, we did observe osteoclast formation in response to hPTH-(53–84), which is known not to bind to or activate the PTH1R (24). The target cell(s) for hPTH-(53–84) in our cocultures is not yet known, although we could not detect binding to MS1 cells of the C-terminal radioligand [125I]hPTH-(19–84) (unpublished observations), and previous reports have implicated direct actions of such fragments on osteoclast progenitors (18).

We directly addressed the possible role of the PTH1R in cells of the osteoclast lineage by coculturing MS1 cells with fetal hepatic osteoclast progenitors that lacked PTH1Rs (i.e. PTH1R-/- progenitors). We found that formation of osteoclasts was induced by rPTH-(1–34) as readily from PTH1R-/- as from PTHR+/- fetal hepatic progenitors. Moreover, the numbers of TRAP+ MNCs formed in response to 1,25-(OH)2D3 were similar regardless of the genotype of the donor liver, which suggests that vitamin D-dependent osteoclastogenesis does not require expression of PTH1Rs on osteoclast progenitors. Because no TRAP+ MNCs formed in cultures of either MS1 cells or fetal liver alone, these results indicate unequivocally for the first time that expression of PTH1Rs on osteoclast progenitors is not required for PTH-mediated osteoclast-like cell formation in the presence of PTH-responsive stromal cells. This result is not inconsistent with previous reports of PTH-(1–34) actions on osteoclast progenitors (17, 18, 20, 26), but it does suggest that such actions, if mediated by type 1 PTH1Rs, are not essential for PTH-induced osteoclastogenesis.


   Footnotes
 
1 This work was supported by NIH Grants DK-11794 and DK-47038. Back

Received September 8, 1997.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 References
 

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Parathyroid Hormone Controls Receptor Activator of NF-{kappa}B Ligand Gene Expression via a Distant Transcriptional Enhancer.
Mol. Cell. Biol., September 1, 2006; 26(17): 6453 - 6468.
[Abstract] [Full Text] [PDF]

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S. Kim, M. Yamazaki, L. A. Zella, N. K. Shevde, and J. W. Pike
Activation of Receptor Activator of NF-{kappa}B Ligand Gene Expression by 1,25-Dihydroxyvitamin D3 Is Mediated through Multiple Long-Range Enhancers.
Mol. Cell. Biol., September 1, 2006; 26(17): 6469 - 6486.
[Abstract] [Full Text] [PDF]