The authors state that they have no conflicts of interest.
Article first published online: 24 MAR 2008
Copyright © 2008 ASBMR
Journal of Bone and Mineral Research
Volume 23, Issue 8, pages 1238–1248, August 2008
How to Cite
Zhang, X. and Zanello, L. P. (2008), Vitamin D Receptor–Dependent 1α,25(OH)2 Vitamin D3–Induced Anti-Apoptotic PI3K/AKT Signaling in Osteoblasts. J Bone Miner Res, 23: 1238–1248. doi: 10.1359/jbmr.080326
Published online on March 24, 2008;
- Issue published online: 4 DEC 2009
- Article first published online: 24 MAR 2008
- Manuscript Accepted: 21 MAR 2008
- Manuscript Revised: 8 FEB 2008
- Manuscript Received: 17 JUL 2007
- vitamin D
Osteoblast apoptosis plays a crucial role in bone remodeling. Physiological doses of 1α,25(OH)2-vitamin D3 (1,25D) protect osteoblasts against apoptosis by means of mechanisms only partially understood. We studied activation of an Akt survival cascade downstream of 1,25D nongenomic stimulation of phosphatidylinositide-3′-kinase (PI3K) in osteoblastic cells. We measured a dose- and time-dependent 1,25D induction of Akt phosphorylation (p-Akt) in cultured osteoblastic cells. Maximal response was achieved with 10 nM 1,25D after 5 min. We found that staurosporine (STSP)-induced apoptosis was significantly reduced in 1,25D-pretreated osteoblasts. 1,25D prosurvival effects were abolished when cells were preincubated with inhibitors of PI3K activation. By means of siRNA silencing, we proved that 1,25D induction of p-Akt requires a classic vitamin D receptor (VDR) in osteoblasts. Furthermore, non-osteoblastic CV-1 cells transfected with an enhanced green fluorescent protein (EGFP)-VDR construct responded to 1,25D treatment with a rapid p-Akt response associated with increased cell survival not detected in native, nontransfected cells. We measured increased levels of p-Akt substrates p-Bad and p-FKHR and significantly reduced activity of caspases 8 and 3/7 after 1,25D treatment. In addition, 1,25D-induced protection against apoptosis was abolished when osteoblasts were preincubated with pertussis toxin. We conclude that anti-apoptotic effects of 1,25D in osteoblasts occur through nongenomic activation of a VDR/PI3K/Akt survival pathway that includes phosphorylation of multiple p-Akt substrates and reduction of caspase activities.
1α,25(OH)2-vitamin D3 (1,25D), the hormonally active metabolite of vitamin D3, is essential for the production and maintenance of the adult skeleton.[1, 2] 1,25D regulates systemic levels of calcium and phosphate, affecting the state of mineralization of bone. Skeletal defects found in mice lacking a functional vitamin D receptor (VDR), for example, can be corrected by normalizing mineral ion homeostasis through the diet. In addition, 1,25D has direct effects on osteoblasts in vitro.[4, 5] This steroid hormone promotes osteoblast differentiation, cell death and survival, and osteogenic activities in vitro. 1,25D control over the osteoblast to osteoclast ratio might be seen as one possible mechanism to explain bone anabolic properties of the hormone.
The direct effects of 1,25D on osteoblasts involve two different molecular mechanisms. On one hand, 1,25D binds to a nuclear VDR highly expressed in osteoblasts. Ligand-bound VDR functions as a transcription factor for the expression of genes involved in osteoblast differentiation, proliferation, and bone matrix production.[5, 6] On the other hand, 1,25D exerts VDR-dependent extranuclear actions that lead to stimulation of plasma membrane activities[7-9] and signaling cascades that ultimately modulate nuclear control of the cell cycle, programmed cell death, and survival.[7-11] Cytoplasmic VDR coupling to protein kinases has been proposed to occur through G proteins in osteoblasts; however, the precise molecular mechanisms remain mostly unknown.
Hormone regulation of bone cell apoptosis seems to be a crucial mechanism in the control of the osteoblast to osteoclast cell ratio, and therefore, the state of mineralization of the bone. In particular, 1,25D has been shown to have anti-apoptotic effects on primary osteoblasts and osteoblastic cell lines in vitro.[13-15] These effects involve nongenomic stimulation of a phosphatidylinositide-3′-kinase (PI3K)/c-Jun N-terminal kinase (JNK) pathway that leads to suppression of induced DNA fragmentation in human osteoblastic cells.[12-15] Although 1,25D-induced anti-apoptotic effects in osteoblasts have been documented by several groups, the underlying signaling pathways remain only partially understood. Identification of the signaling molecules involved in 1,25D-induced protection against apoptosis has potential therapeutic value for the treatment of bone diseases characterized by decreased bone mass and mineralized matrix.[16-18]
The PI3K/Akt pathway is one of the most critical signaling pathways involved in the regulation of cell survival.[19, 20] Here we describe 1,25D induction of Akt activation (phosphorylation) leading to protection against induced apoptosis in osteoblasts and downstream phosphorylation of Bad protein and members of the Forkhead (FKHR) family of transcription factors.[21, 22] In addition, we study the inhibitory effects of 1,25D pretreatment on large and short prodomain caspase activities after induction of apoptosis with staurosporine (STSP), which increases mitochondrial permeability.[23-25]
The initiation of the signal transduction that couples a ligand-bound extranuclear VDR to specific cytoplasmic cascades remains mostly unknown.[8, 12, 26, 27] Rapid modulation of kinase cascades by 1,25D in osteoblasts has been shown to be sensitive to pertussis (PTX) toxin. Here we describe for the first time 1,25D activation of a VDR-dependent, PTX-sensitive nongenomic PI3K/Akt pathway that leads to suppression of STSP-induced apoptosis in osteoblastic cells in culture. Although it would be premature to conclude that in vitro 1,25D induction of osteoblast survival explains in vivo bone anabolic effects of the hormone, our findings represent a contribution to the understanding of the molecular mechanisms by which the hormone helps to maintain the adequate osteoblast population needed for the production of mineralized matrix in the remodeling adult bone.
MATERIALS AND METHODS
1,25D (Biomol Research Laboratories, Plymouth, PA, USA) was stored as a stock solution in pure ethanol at –20°C, in the dark. Wortmanin, PTX, LY294002, and STSP were purchased from Sigma (St Louis, MO, USA).
Rat osteosarcoma ROS 17/2.8 cells (kindly provided by AW Norman, University of California-Riverside) and human osteosarcoma SaOS-2 cells (American Type Culture Collection; ATCC, Manassas, VA, USA) were cultured in Ham F-12 nutrient mixture (Sigma), with the addition of 1 mM Glutamax (Invitrogen, Carlsbad, CA, USA), 5% FBS (Fisher, Pittsburgh, PA, USA), 5% Serum Plus (JRH Biosciences, Woodland, CA, USA), penicillin (100 U/ml), streptomycin (100 μg/ml), and 1.1 mM CaCl2 at 37°C in a humidified 5% CO2 atmosphere, essentially as described previously. CV-1, green monkey kidney cells (ATCC), and undifferentiated NG108-15 neuroblastoma-glioma cells (kindly provided by ME Adams, University of California-Riverside) were cultured in modified Eagle's MEM (Sigma), containing 10% FBS and antibiotics. Cells were plated at low density (105 cells/ml) in 100 × 20-mm tissue culture dishes and used 3–5 days after passage, at ∼80% confluence. Culture medium was replaced by serum-free medium 24 h before treatment with 1,25D and/or the specified reagent.
For apoptosis assays, PI3K inhibitors wortmannin and LY294002 and the G protein inhibitor PTX were added to serum-free cultures at a final concentration of 50 nM, 10 μM, and 100 mg/ml, respectively, for 30 min (for PI3K inhibitors) and 5 h (for PTX) before the addition of hormone. Cells were incubated with 10 nM 1,25D for an additional hour, followed by 10-h exposure to 100 nM STSP to induce apoptosis.
Establishment of a stable VDR knockdown ROS 17/2.8 cell line
Native ROS 17/2.8 cells were grown in 6-well plates for 24 h before transfection. Cells were transfected at 90–95% confluence with 4 μg GenEclipse VDR siRNA construct or control vector DNA in 250 μl Ham F-12 serum-free medium containing 10 μl Lipofectamine 2000 (Invitrogen, Carlsbad, CA, USA), according to the manufacturer's protocol and basically as described before for SaOS-2 cells. Transfection medium was replaced by fresh serum-containing medium after 6 h. Cultures were maintained for additional 30 h. Cells were subcultured at a 1:10 ratio in medium supplemented with 1.6 mg/ml geneticin (Gibco, Auckland, New Zealand) to select for the stably transfected colonies.
Enhanced green fluorescent protein-VDR transfection of CV-1 cells
CV-1 cells were transfected with an enhanced green fluorescent protein (EGFP)-VDR construct or an EGFP control vector DNA (kindly provided by AW Norman, University of California-Riverside) in 250 μl serum-free medium containing 10 μl Lipofectamine 2000 (Invitrogen). Transfection medium was replaced by fresh serum-containing medium after 6 h. Cells were maintained for an additional 30 h and subcultured at a 1:10 ratio in medium supplemented with 1.2 mg/ml geneticin (Gibco) to select for stably transfected colonies. Positively transfected cells were screened with an epifluorescence Olympus IX50 microscope.
After STSP treatment, cells were washed with ice-cold PBS buffer once and trypsinized (0.25% trypsin; Invitrogen). A cell pellet was obtained by centrifugation at 1000 rpm for 6 min. Cells were resuspended in 0.5 ml PBS buffer supplemented with 1% FBS and fixed overnight with methanol at 4°C. Fixed cells were stained with 0.2 mg/ml propidium iodide (PI; Sigma) at 37°C for 2 h. Percentage of apoptotic cells was measured with a FACScan flow cytometer (Becton Dickinson, Franklin Lakes, NJ, USA). Apoptotic cells were identified on the basis of fragmented DNA stained with PI (DNA content < 2N).
Western blot analysis
Cell lysates were obtained in lysis buffer composed of 50 mM Tris-HCl (pH 7.4), 150 mM NaCl, 2 mM EDTA, 25 mM NaF, 0.25% (wt/vol) sodium deoxycholate, 1% (vol/vol) NP-40, 0.2 mM Na3VO4, 1 mM 4-(2-aminoethyl)benzenesulfonyl fluoride hydrochloride (Pefabloc), 2 μg/ml leupeptin, and 2 μg/ml aprotinin (Sigma). Equal volumes of cell lysate (equivalent to 20 μg protein) were loaded and separated on 7.5% or 10% SDS-PAGE gels and transferred to PVDF membranes. After blockade with 5% nonfat milk in Tris-buffered saline plus 0.1% Tween 20 (TBST), membranes were incubated with primary antibodies against p-Akt (anti-Ser-473), p-FKHR (anti-Ser-256; Cell Signaling Technology, Beverly, MA, USA), p-Bad (anti- Ser-136), Bad, Akt, FKHR, or VDR (Santa Cruz Biotechnology, Santa Cruz, CA, USA), raised in rabbit, at 1:1000 dilutions. Each blot was probed with antibodies against α-vinculin or β-actin as a reference for protein loading between samples. Primary antibodies were detected with horseradish peroxidase–conjugated secondary anti-rabbit antibodies and enhanced chemiluminescence. Blot density was digitalized and analyzed with the UnScan-It gel software (Silk Scientific, Orem, UT, USA).
Immunofluorescence and cell imaging
ROS 17/2.8 cells grown on glass coverslips were fixed with 3.7% formaldehyde (Sigma) for 20 min, permeabilized with ice-cold 95% ethanol for 5 min, and incubated in 5% goat serum in PBS (pH 7.4) for 1 h to block any nonspecific binding of the antibodies. Cells were incubated overnight with a primary anti-p-FKHR antibody at a 1:100 dilution, rinsed with PBS, and incubated with an anti-rabbit Cy3 fluorescent secondary antibody conjugate (Sigma, 1:500 dilution) for 1 h. Cell nuclei were labeled with 1 μg/ml DAPI (Sigma) in mounting solution. Subcellular localization of p-FKHR was studied with an Olympus IX50 inverted microscope. Images were captured with a Spot digital camera (Diagnostic Instruments, Sterling Heights, MI, USA) and analyzed with Simple PCI C-Imaging Systems software (Compix, Cranberry Township, PA, USA).
Caspase activity assay
Carboxyfluorescein fluorochrome inhibitor of caspases (FLICA) kits for detection of poly-caspase and caspase 8, 9, and 3/7 (Immunochemistry Technologies, Bloomington, MN, USA) activities were used according to the manufacturer's protocols. The assay uses a FLICA that binds covalently to active caspases. Briefly, 3-day-old ROS 17/2.8 cells were serum-starved for 24 h, and apoptosis was induced with STSP as described before. Caspase activities were measured in 96-well plates with a Synergy HT multiplate reader (BioTeck Instruments, Winoosky, VT, USA). Briefly, 300 μl of cell suspension (∼0.5 × 106 cells/ml) were incubated with the caspase detection reagent for 1–2 h at 37°C under 5% CO2. Fluorescence intensity, which is proportional to caspase activity, was measured in black-walled, clear bottom wells. For fluorescence microscopy staining, 105 cells were seeded onto sterile glass coverslips in 6-well plates, and apoptosis was induced after 24 h. Cells were incubated with FLICA solution for 1 h at 37°C under 5% CO2. FLICA-containing medium was replaced with Hoechst staining medium (Immunochemistry Technologies) for detection of apoptotic nuclei and incubated for 5 min at 37°C under 5% CO2, followed by two washes. Cells were observed under an Olympus IX50 epifluorescence microscope. The green fluorescence signal of caspase-positive cells was detected with excitation at 490 nm and emission at 520 nm.
Cell proliferation assay
Cells were seeded into 96-well plates at 1000 cells/well for 24 h. Cells at 80% confluency were treated with 100 nM STSP for 10 h to induce apoptosis, with or without a 1-h pretreatment with either 10 nM 1,25D or vehicle (≤0.05% ethanol) for 3 days. STSP was washed off with fresh culture medium, and cells were allowed to proliferate for an additional 48 h. We used a CyQUANT NF cell proliferation assay kit (Invitrogen) that quantifies the fluorescence intensity of dye binding to cellular DNA, with excitation at 485 nm and emission at 530 nm. The extent of proliferation was determined by comparing relative fluorescence units in 1,25D-pretreated versus control untreated samples.
Secreted alkaline phosphatase activity assay
Secreted alkaline phosphatase activity (SEAP) activity was measured in native controls and siRNA VDR-transfected ROS 17/2.8 cells with a commercial chemiluminescent reporter assay kit for secreted alkaline phosphatase enzyme (Applied Biosystems, Bedford, MA, USA), according to the manufacturer's protocol. Briefly, cells were harvested and transferred to 96-well plates after a 24-h treatment with 1,25D, where they were treated with the reagents provided in the kit. The relative intensity of luminescent signals was recorded was a Synergy microplate reader (Bioteck Instruments).
Data were expressed as means ± SE. Statistically significant differences between two treatments were determined by two-tailed unpaired Student's t-test and one-way ANOVA to compare means of more than three treatments.
1,25D induces Akt activation in a dose- and time-dependent manner
To study the hypothesis that 1,25D potentiates Akt phosphorylation downstream of PI3K activation in osteoblasts subjected to an apoptotic stimulus, we measured relative levels of p-Akt with and without 1,25D stimulation in rat osteoblastic ROS 17/2.8 cells treated with 100 nM STSP for 10 h to induce apoptosis.
Quantitative Western blot analyses was performed in 4- to 5-day-old ROS 17/2.8 cell cultures with a polyclonal antibody that specifically recognizes a phosphorylated Ser-473 residue in the Akt molecule (Fig. 1). Cells were serum-starved for 24 h before 1,25D incubation. Cell lysates were obtained from cultures treated with 1,25D in a concentration range of 0.1–1000 nM for 10 min. Relative amounts of phosphorylated (p-Akt) versus total Akt were estimated by measuring the density of the immunoblots. Phosphorylation levels were normalized respect to the total amount of protein loaded in the gel. Relative levels of p-Akt in 1,25D-treated cells were expressed as fold increase with respect to vehicle-treated cells (0 nM 1,25D), as shown in Fig. 1A. Maximum p-Akt increase with respect to vehicle (2.4 ± 0.3-fold) was obtained with a concentration of 10 nM 1,25D. Next, we studied the time course of the p-Akt response obtained with 10 nM 1,25D. 1,25D increased relative levels of p-Akt within 5 min of treatment (Fig. 1B). Maximum increase was achieved at 10 min and remained significantly elevated for the entire duration of the experiment (1 h). We found similar 1,25D induction of p-Akt in human SaOS-2 osteoblastic cells and non-osteoblastic, undifferentiated NG108-15 neuroblastoma-glioma cells (Table 1). Our results thus show that transient 1,25D treatment stimulates Akt activity in a time- and concentration-dependent manner in osteoblastic and (undifferentiated) non-osteoblastic cell systems.
1,25D treatment reduces STSP-induced apoptosis through a PI3K/Akt signaling in ROS 17/2.8 osteoblasts
To study the hypothesis that rapid upregulation of p-Akt by nanomolar concentrations of 1,25D leads to attenuation of apoptosis in osteoblasts, we measured the percentage of apoptotic ROS 17/2.8 cells with or without pretreatment with 10 nM 1,25D for 1 h. Apoptosis was induced with 100 nM STSP applied for 10 h after pretreating cells with either 10 nM 1,25D or vehicle (0.01% ethanol) for 1 h. As shown in Figs. 2A and 2B, 100 nM STSP induced a significant 2-fold increase in the percentage of apoptotic cells in osteoblast cultures at 80% confluence with respect to mock (untreated) cells. We found that pretreatment of ROS 17/2.8 cells with 10 nM 1,25D for 1 h before apoptosis induction decreased the number of apoptotic cells by ∼35% with respect to values obtained in the absence of hormone treatment (Table 1). To verify our hypothesis that a PI3K/Akt pathway is involved in 1,25D anti-apoptotic functions, we pretreated the cells with PI3K inhibitors wortmannin (50 nM) and LY294002 (10 μM) for 30 min before incubation with 10 nM 1,25D. We found that pretreatment of ROS 17/2.8 cells with these inhibitors abolished 1,25D anti-apoptotic effects (Fig. 2B), thus confirming that 1,25D protection against apoptosis occurs through PI3K activation.
Next, we measured cell densities of STSP-treated ROS 17/2.8 cultures for 3 days with or without pretreatment with 10 nM 1,25D. As shown in Fig. 2C, cell counts obtained from cultures pretreated with 1,25D were ∼30% higher than cultures that developed in the absence of hormone (Table 1). These data agree with the apoptosis results shown in the same figure and further support our observations that 1,25D promotes survival of this osteoblastic cell line.
1,25D treatment induces phosphorylation (inactivation) of prodeath proteins Bad and FKHR downstream of PI3K/Akt activation
With the purpose to identify signal molecules involved in 1,25D-induced suppression of osteoblast apoptosis, we measured relative phosphorylation levels of PI3K/Akt downstream effectors known to be involved in the cell survival pathway in the presence and absence of 1,25D treatment. Akt kinase targets various downstream effectors involved in the control of cell apoptosis. Among them are proapoptotic Bad protein and members of the Forkhead (FKHR) family of transcription factors, which are negatively regulated by phosphorylation catalyzed by Akt, and therefore inhibit apoptosis while present in the cytoplasm in their inactive (phosphorylated) form. Here, we performed Western blot analysis with antibodies against p-Bad (at Ser-136) and p-FKHR (at Ser-256), relative to their unphosphorylated forms, in the presence and absence of hormone treatment. ROS 17/2.8 cell lysates were obtained after treating 4-day-old cultures with 10 nM 1,25D for the indicated time periods. As shown in Figs. 3A and 3B, total amounts of unphosphorylated Bad and FKHR remained stable for the entire length of the experiment (up to 3 and 6 h, respectively). However, the relative amount of their inactive (phosphorylated) forms showed a significant increase during the time course of 1,25D treatment. p-Bad levels, on one hand, increased significantly by 1.6-fold rapidly after 5 min of 1,25D treatment and returned to initial levels at 30 min (Fig. 3A). On the other hand, 1,25D stimulation of FKHR inactivation (phosphorylation) was detected after 30 min and remained elevated for at least 6 h (Fig. 3B). Similar responses were found for 1–20 nM 1,25D (data not shown). This time course of FKHR phosphorylation on 1,25D treatment agrees with previous observations that Akt may translocate to the cell nucleus where it interacts with and phosphorylates FKHR at Ser-256. This suppresses FKHR transcriptional activity as a regulator of the expression of genes implicated in the control of cell death.
We further studied whether 1,25D treatment induces p-FKHR cytoplasmic retention. By means of immunofluorescence, we detected that, in the absence of 1,25D treatment, osteoblasts exhibited basal low levels of p-FKHR (Fig. 3B, top left image). However, a 1-h treatment with 10 nM 1,25D stimulated a robust increase in the fluorescence signal corresponding to an increase in p-FKHR levels in the cytoplasm (Fig. 3B, top right image). Pretreatment of ROS 17/2.8 cells with PI3K inhibitors LY294002 and wortmannin reduced 1,25D-enhanced cytoplasmic p-FKHR signal (Fig. 3B, bottom image). These results are consistent with the increase in p-FKHR levels detected by Western blot with a 1-h-long 1,25D treatment (Fig. 3B) and support the hypothesis that FKHR phosphorylation occurs as a consequence of 1,25D activation of the PI3K/Akt survival pathway. Taken together, results shown in Fig. 3 indicate that 1,25D suppresses STSP-induced apoptosis through phosphorylation of effectors Bad and FKHR downstream of PI3K/Akt activation in osteoblasts.
1,25D treatment inhibits caspase activities in osteoblasts
Increased caspase activities are required for successful completion of cell apoptosis. Proteolytic caspase activities are implicated in the development of the apoptotic phenotype, such as cellular dismantling.[31, 32] We investigated the hypothesis that 1,25D protection against STSP-induced apoptosis involves attenuation of caspase activities. 1,25D decreases caspase-8 activity in a Fas-induced apoptotic pathway in osteoblasts. Here we measured total and caspase 8, 9, and 3/7 activities in STSP-treated ROS 17/2.8 cells, with and without a 1-h preincubation with 10 nM 1,25D. Initiator caspases 8 and 9, the latter a constituent of the apoptosome, are involved in the extrinsic and intrinsic apoptotic pathways, respectively. Caspases 3/7 are effector caspases at the end of both the intrinsic or mitochondrial and extrinsic apoptotic pathways. As shown in Fig. 4A, 1,25D treatment blocked ∼53% of STSP-induced total active caspases compared with vehicle-treated ROS 17/2.8 osteoblasts. In agreement with this result, fluorescence imaging performed for the active form of total caspases in STSP-induced apoptotic cells showed that 1,25D pretreatment markedly decreased the activation of total caspases with respect to a vehicle-treated sample (Fig. 4D).
The STSP mechanism of induction of apoptosis is not fully known. STSP seems to activate components of the extrinsic and intrinsic pathways in different cell systems. We found that STSP treatment of ROS 17/2.8 cells increases the activities of caspases 8 and 9 (Fig. 4C). Preincubation of cells with 10 nM 1,25D significantly decreased caspase 8 activation compared with vehicle-treated cells but not caspase 9 activation (Fig. 4C), suggesting that 1,25D suppression of STSP-induced apoptosis involves components of the extrinsic pathway. On the other hand, a 43% reduction of caspases 3/7 activities was measured in ROS 17/2.8 cells treated with 1,25D compared with cells treated with vehicle (Fig. 4B). Although there is no published evidence up to date of Akt direct phosphorylation of caspases 3 or 7, protein database analysis showed that caspases 3/7 contain Akt consensus phosphorylation sites. Taken together, our results suggest that 1,25D attenuates effector caspases 3/7 activities, likely through an extrinsic apoptotic pathway induced by STSP treatment that seems to involve the initiator caspase 8, but not caspase 9.
Classic VDR is required for the rapid induction of Akt phosphorylation by 1,25D
The understanding of the molecular mechanisms underlying 1,25D induction of transcription-independent VDR functions is still controversial. Although some groups have identified non-VDR proteins that seem to function as a distinct 1,25D membrane-related receptor,[35, 36] there is increasing evidence that a classic VDR plays a nongenomic role in 1,25D induction of cytoplasmic signal transduction in addition to its well-known genomic actions.[8, 11, 27, 37, 38] We studied the involvement of a classic VDR in 1,25D induction of PI3K/Akt activation in three cell lines that express different levels of the native VDR protein (Fig. 5A). To study the hypothesis that a classic VDR is required for nongenomic 1,25D induction of Akt activation, we performed quantitative Western blot analysis of p-Akt/Akt levels before and after 1,25D treatment in osteoblastic rat ROS 17/2.8 and human SaOS-2 cells in which VDR expression levels were reduced by siRNA VDR silencing and in non-osteoblastic CV-1 cells transfected with a EGFP-VDR (Fig. 5; Table 1).
We showed before (Fig. 1) that 10 nM 1,25D significantly increases p-Akt/Akt levels in ROS 17/2.8 osteosarcoma cells within 5 min. Here we found that native CV-1 cells expressing low endogenous levels of VDR protein did not show significant p-Akt upregulation after a 10-min treatment with 0.1–100 nM 1,25D (Fig. 5C, top Western blot panel). To study the requirement of a VDR in 1,25D induction of Akt phosphorylation, we stably silenced VDR expression in rat ROS 17/2.8 and human SaOS-2 cells by means of an siRNA strategy. To verify that the newly created siRNA VDR-transfected osteoblastic subclones carried a functional VDR deficiency, we measured 1,25D induction of SEAP in siRNA VDR, and siRNA control vector-transfected cells. We found that our VDR knockdown subclones had significantly reduced VDR genomic functions, as shown in Fig. 5B for ROS 17/2.8 cells (for the SaOS-2 siRNA VDR subclone). Next, we measured 1,25D nongenomic induction of Akt phosphorylation in siRNA VDR and siRNA control vector-transfected cells. As shown in Fig. 5B, we found that 1,25D induction of p-Akt levels was significantly reduced in VDR knockdown osteoblasts compared with p-Akt levels measured in siRNA control vector-transfected cells for 10 and 100 nM of the hormone. Table 1 summarizes the fold increase values for p-Akt/Akt levels obtained with 10 and 100 nM 1,25D in native, control vector-transfected, and siRNA VDR ROS 17/2.8 and SaOS-2 cells, and the associated reduction of apoptosis and increased survival rates obtained with 10 nM of the hormone.
To further study whether 1,25D failure to increase p-Akt levels in CV-1 cells was caused by low native VDR levels of this cell line, we studied the hypothesis that expression of an exogenous VDR in CV-1 cells can restore 1,25D ability to activate native Akt under 1,25D treatment and the cells' survival response to hormone treatment. We transiently transfected CV-1 cells with an EGFP-VDR construct and comparatively measured 1,25D induction of Akt phosphorylation and cell proliferation in native, control EGFP-transfected, and EGFP-VDR–transfected cells (Fig. 5C; Table 1). We measured significantly higher p-Akt levels in EGFP-VDR transfected CV-1 cells with respect to EGFP-control transfected cells treated with 10 nM 1,25D (Table 1). In addition, we measured a significant activation of p-Akt by 10 and 100 nM 1,25D and increase in cell survival in the neuroblastoma-glioma NG108-15 cell line, which expresses the VDR (Table 1). Last, we measured an increase in CV-1 cell survival in cells transfected with EGFP-VDR and treated with 10 nM 1,25D for 3–4 days but not in nontransfected and EGFP-control–transfected cells. It is interesting to note that, as reflected by the Western blots shown in Fig. 5, basal levels of p-Akt in siRNA VDR osteoblastic cells and native CV-1 cells expressing low VDR levels were relatively lower than basal p-Akt levels in each respective control cell type expressing higher VDR levels (control siRNA-transfected ROS 17/2.8 cells and EGFP-VDR–transfected CV-1 cells). From our transfection experiments, it seems that the reduction in VDR levels on VDR silencing in ROS 17/2.8 cells results in a marked increase in basal Akt phosphorylation; however, these levels are insensitive to 1,25D treatment. Similarly, native CV-1 cells expressed higher basal p-Akt levels that did not respond to 1,25D. These higher basal p-Akt levels measured in the absence of a VDR may reflect the outcome of other cellular pathways indirectly inhibited by VDR activation. Taken together, our data showed that a classic VDR, likely localized in the cell cytoplasm in close proximity to the plasma membrane, is required for the rapid activation of a prosurvival Akt signaling in osteoblasts and non-osteoblastic cells.
1,25D induction of PI3K/Akt activation and subsequent protection against apoptosis is inhibited by PTX
There is growing evidence that some plasma membrane steroid receptors regulate nongenomic effects by coupling to a G protein.[39, 40] We investigated the hypothesis that 1,25D activation of Akt phosphorylation may involve a G protein–coupled pathway. PI3K activation can occur by coupling of its 110-kDa catalytic subunit to a G heterotrimeric protein. We found that 1,25D stimulation of p-Akt levels is abolished in ROS 17/2.8 cells that have been preincubated with 100 ng/ml PTX for 5 h (Fig. 6A). This suggests that 1,25D-induced Akt activation may occur through a PTX-sensitive subunit, likely a Gαi protein expressed in osteoblasts, which couples to a cytoplasmic VDR. In addition, we studied whether 1,25D activation of a G protein/PI3K/Akt pathway leads to suppression of STSP-induced apoptosis in ROS 17/2.8 cells. As shown in Fig. 6B, 1,25D treatment significantly reduced ROS 17/2.8 cells apoptosis only in the absence of PTX pretreatment. Our results indicate that a heterotrimeric G protein seems to be involved in 1,25D stimulation of a survival signaling upstream of a PI3K/Akt pathway.
Nongenomic 1,25D activation of prosurvival PI3K/Akt pathways has been shown previously in vitro in a variety of cell types.[43, 44] We showed here that 1,25D upregulation of p-Akt leads to suppression of osteoblast apoptosis and increased cell survival. Although anti-apoptotic effects of 1,25D in osteoblasts had been shown previously to involve a VDR/PI3K signaling, a detailed characterization of this PI3K cascade remained to be elucidated.
Protein kinase Akt is one of the best-characterized targets of PI3K activation. Several downstream targets of PI3K/Akt activation have been implicated in cell survival, including the prodeath protein Bad, effector caspases 3/7, and transcription factors of the Forkhead family.[20-22] Rapid, nongenomic 1,25D activation of the PI3K/p-Akt signaling inhibits an STSP-induced apoptotic cascade at multiple points (Fig. 7) and inhibits the development of cellular mechanisms leading to induced programmed cell death. Our results on 1,25D reduction of initiator caspase 8 levels, but not caspase 9 and effector caspases 3/7, suggest that hormone-induced anti-apoptotic signaling interferes with an extrinsic cell death pathway induced by STSP.
In addition to its anti-apoptotic effects, 1,25D is well known for its anti-proliferative actions in vitro.[11, 43, 45] We showed recently that prolonged treatment of cultured osteoblasts with 1,25D (3 or more days) induces cell death. In contrast, anti-apoptotic effects can be measured after short-term exposure of osteoblast cultures to 1,25D (<2 days). At the cellular level, availability of 1,25D molecules—as a function of concentration, solubility, receptor/ligand binding affinities, diffusion time, and distances outside and inside of the cytosol—will determine the interaction with specific molecular targets and the type of cellular response. In this paper, we showed that, under our experimental conditions, protection against STSP-induced apoptosis is achieved within a 1-h pretreatment with 10 nM 1,25D. This survival signaling seems to include a PTX-sensitive G protein upstream of PI3K activation, which may couple to a membrane-associated VDR. Rapid, nongenomic actions of steroids have, in some cases, been associated with the activation of novel membrane receptors that couple to a G protein.[47, 48] We showed here, however, that 1,25D nongenomic activation of PI3K/Akt occurs only in the presence of a functional extranuclear VDR and is sensitive to PTX, suggesting that a classic receptor may couple to a Gαi to initiate 1,25D rapid functions.
We therefore propose that activation of a membrane-associated or cytoplasmic VDR, likely to couple to a heterotrimeric G protein, results in the recruitment of a PI3K isoform to the inner surface of the plasma membrane (Fig. 7). After diffusing across the osteoblast plasma membrane, 1,25D binds to a classic VDR that resides at the cytoplasm in close proximity to the phospholipid bilayer. Ligand-bound VDR rapidly transduces the hormone signal to a PTX-sensitive Gα protein complex, which interacts with the PI3K catalytic subunit, resulting in stimulation of PI3K. Activated PI3K catalyzes PIP3 production, which facilitates Akt recruitment at the plasma membrane and subsequent phosphorylation (activation) of Akt. As it moves through the cytoplasm, activated p-Akt phosphorylates its downstream apoptosis regulators, including pro-apoptosis Bad (within 5–10 min of 1,25D treatment), and FKHR in the proximity to the cell nucleus (at 30 min). In addition, the activities of the initiator caspase 8 and effector caspases 3/7 present in the cytoplasm are reduced. These synchronized molecular responses downstream of a ligand-bound cytoplasmic VDR lead ultimately to attenuation of STSP-induced apoptosis. We propose that 1,25D functions as a survival factor in osteoblasts by promoting, on one hand, genomic suppression of cell death through inactivation (phosphorylation) of FKHR transcription factors and by reducing proteolytic caspase and prodeath Bad protein activities on the other. Overall, 1,25D induction of osteoblast survival may help maintain, at the tissue level, an osteoblast to osteoclast ratio favorable for adult bone production. Identification of the molecular transducers of 1,25D prosurvival signaling has potential pharmacological value for the design of new therapeutic agents for the treatment of bone diseases.
The authors thank Barbara Walter for technical support with flow cytometry and Elmer Hillo for assistance with the graphics. This work was funded by NIH Grant DK071115 to LPZ.
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