PLCγ2 regulates osteoclastogenesis via its interaction with ITAM proteins and GAB2 (original) (raw)
Inhibition of PLCγ enzymatic activity blocks OC differentiation and disrupts actin ring formation. PLCγ isoforms are known activators of the NFAT family of transcription factors in lymphoid cells, downstream of immune receptor signaling via ITAMs (33). Since NFAT and ITAMs are both critical regulators of OC differentiation and cells lacking the ITAM-containing adapters Dap12/FcRγ display severe osteopetrosis (20), we asked whether PLCγ family members were also required for efficient OC differentiation and function. We therefore tested the capacity of the PLCγ inhibitor U73122 to block OC formation and bone resorption. Addition of U73122 to the culture media blocked OC development, as demonstrated by the absence of tartrate-resistant acid phosphatase–positive (TRAP-positive) cells, compared with the numerous multinucleated OCs obtained in wells cultured only with RANKL and M-CSF (Figure 1A). Replacing the media with fresh osteoclastogenic media lacking the inhibitor restored the ability of the cells to differentiate, indicating that the compound was not toxic to the cells (Figure 1A). Culture of OCs on dentin in the presence of U73122 blocked bone resorption (Figure 1B). Furthermore, addition of U73122 to mature resorbing cells rapidly disrupted the organization of the actin rings, indicating that this compound can block bone resorption independent of its capacity to affect differentiation (Figure 1C).
PLCγ inhibition blocks osteoclastogenesis and actin ring formation. (A) WT OCs were generated with RANKL (100 ng/ml) and M-CSF (10 ng/ml) in the presence of the PLC inhibitor U73122 (5 μM) for 4 days on plastic. In some wells, the media with the inhibitor was replaced with fresh osteoclastogenic media, and cells were allowed to differentiate for 4 more days (Withdrawal U73122). (B) WT OCs were grown on dentin with or without U73122 for 10 days. Cells were then removed and pits stained with hematoxylin red (Magnification, ×200). CTR, control. (C) WT OCs were generated on dentin in the absence of the inhibitor, then treated with vehicle or U73122 for 1 hour. Cells were fixed and actin stained using FITC-phalloidin (Magnification, ×200).
PLCγ2 regulates bone mass in vivo. To determine whether PLCγ1, PLCγ2, or both are required for OC formation, we turned to the analysis of mice lacking these proteins. Plcg1–/– mice die in utero by day 9, rendering the analysis of their bone phenotype difficult. Plcg2–/– mice, however, are viable. Thus, femurs from 4-month-old sex-matched WT and Plcg2–/– mice were subjected to histological examination (Figure 2, A–E) and μCT analysis (Figure 2, F–I). Plcg2–/– mice had a more than 3-fold increase in the percentage of trabecular bone volume versus total bone volume compared with their WT counterparts, as determined by histological examination (Figure 2B) and as shown in μCT reconstructed 3D images (Figure 2F). Histomorphometric analysis also indicated that the number of osteoblasts (OBs) per bone perimeter was similar in WT and Plcg2–/– mice (Figure 2C), while the number of OCs per bone perimeter and the OC surface were significantly decreased in the null animals (Figure 2, D and E). The number of the trabeculae (Figure 2G) and their thickness (Figure 2H) were significantly higher in the Plcg2–/– compared with WT mice, while the space between the trabeculae was smaller (Figure 2I). Thus, PLCγ2 regulates bone mass in vivo.
Osteopetrosis in mice lacking PLCγ2. (A) TRAP staining of decalcified histological sections of WT and Plcg2–/– proximal femurs. Magnification, ×40. (B–E) Quantitative analysis of bone parameters from histological sections of WT and Plcg2–/– bone femurs (n = 8) showing: percentage of bone volume versus total bone volume (BV/TV) (B); number of OBs (nOB) per bone perimeter (C); number of OCs (nOC) per bone perimeter (D); fraction of trabecular surface covered by OCs (E). (F) 3D reconstitution of μCT scans of WT and Plcg2–/– femurs. (G–I) 3D trabecular quantitative parameters of bone structure (n = 6). Graphs show mean ± SEM, with significant differences compared with WT indicated.
PLCγ2 is required for OC differentiation. To determine whether PLCγ2 modulates the capacity of OBs to form bone or to promote OC differentiation, primary OBs were isolated from newborn mice, and their capacity to mineralize extracellular matrix or induce osteoclastogenesis in a coculture system was examined. Plcg2–/– OBs were indistinguishable from their WT counterparts in their ability to form bone nodules in response to β-glycerolphosphate (WT, 14 ± 2.08 nodules/well versus Plcg2–/–, 14.6 ± 1.65 nodules/well) (Figure 3A). This finding was also supported by the in vivo analysis of bone mineral apposition determined by calcein double-labeling (Figure 3B), which revealed no significant difference in bone formation rates (BFRs) and mineral apposition rates (MARs) between WT (BFR, 0.8938 ± 0.1042 μm3/μm2/d; MAR, 0.5927 ± 0.0219 μm/d) and Plcg2–/– animals (BFR, 0.8450 ± 0.1629 μm3/μm2/d; MAR, 0.4015 ± 0.0876 μm/d). To determine whether Plcg2–/– OBs are capable of supporting OC differentiation, BM macrophages (BMMs) from WT mice were cocultured with either WT or Plcg2–/– OBs in the presence of 1,25 vitamin D3 (1,25 Vit D3). Cultures were stained for TRAP to identify OCs. WT BMMs became TRAP-positive, multinucleated cells when cultured with WT OBs (OCs, 9 ± 1.07/well) or null OBs (OCs, 7 ± 0.44/well), indicating that the expression of PLCγ2 in OBs is not required to support osteoclastogenesis. However, in a parallel experiment, BMMs from Plcg2–/– mice failed to form mature OCs in the coculture system, independent of the source of the OBs (Figure 3C). These results suggest that the increased in vivo bone mass observed in Plcg2–/– mice was not due to aberrant OB activity but was dependent on an intrinsic defect in the OC lineage. We therefore cultured primary BMMs with M-CSF and RANKL for 5 days, in the absence of OBs or stromal cells. The osteoclastogenic response of Plcg2–/– BMMs to RANKL and their capacity to resorb bone were arrested (Figure 3D; WT OCs, 33.2 ± 1.74/well versus Plcg2–/– OCs, not detected). Increasing the concentration of M-CSF from 10 to 100 ng/ml, a condition that partially rescued the osteoclastogenesis in cells lacking the ITAM-bearing adapter Dap12 (34), did not enhance Plcg2–/– OC differentiation (data not shown).
PLCγ2 is required for osteoclastogenesis. (A) Bone nodule formation in WT and Plcg2–/– OBs cultured with ascorbic acid and β-glycerolphosphate. (B) Double labeling of calvarial bones showing the degree of bone formation (BFR) of WT and Plcg2–/– mice injected on day 0 and day 7 with calcein (WT BFR, 0.8938 ± 0.1042 μm3/μm2/d; WT mineral apposition rate [MAR], 0.5927 ± 0.0219 μm/d; Plcg2–/– BFR, 0.8450 ± 0.1629 μm3/μm2/d; Plcg2–/– MAR, 0.4015 ± 0.0876 μm/d). (C) WT and Plcg2–/– BMMs cultured with WT and Plcg2–/– OBs in the presence of 10–8 M 1,25 Vit D3. After 14 days, cells were fixed and TRAP stained to detect the presence of multinucleated OCs. (D) TRAP-stained OCs generated with RANKL (100 ng/ml) and M-CSF (10 ng/ml) for 5 days (top panel) and bone resorptive pits generated from OCs plated on dentin for 7 days. Pits were stained with hematoxylin red. (E) WT and Plcg2–/– BMMs retrovirally transduced with vector alone (pMX) or with Flag-tagged PLCγ2 were allowed to differentiate in osteoclastogenic media for 5 days, then stained for TRAP. Magnification, ×200 (A, C–E); ×100 (B).
To determine which stage of OC differentiation is PLCγ2 dependent, the levels of early osteoclastogenic markers TRAP, cathepsin K, NFATc1, and calcitonin receptor, a late OC marker, were determined by real-time PCR analysis in BMMs cultured with RANKL (100 ng/ml) and M-CSF (10 ng/ml) up to 4 days. The expression of TRAP, cathepsin K, and NFATc1 increased within 24–48 hours after RANKL stimulation in WT but not in Plcg2–/– cells (not shown). On day 4, the levels of these genes were maximal in WT cells but dramatically lower in the null cells (Figure 4, A–D). Similarly, calcitonin receptor was detected after 4 days of treatment with RANKL only in WT OCs, suggesting that Plcg2–/– cells have an early block of OC development. Confirming mRNA expression, protein levels of NFATc1 were significantly higher in total cell lysates of day 4 WT OCs than in null cells, both under basal conditions and following 60 minutes of RANKL stimulation (Figure 4E). Together, these data indicate that PLCγ2 is required at early stages of OC development.
PLCγ2 regulates NFATc1 levels and the expression of early osteoclastogenic genes. (A–D) Real-time PCR analysis of osteoclastogenic markers in WT and Plcg2–/– cells in culture with M-CSF alone (d0) or with RANKL (100 ng/ml) plus M-CSF (10 ng/ml) for 4 days (d4). Data are normalized relative to β-actin. ND, not detected. Significant differences compared with WT are shown. (E) Expression levels of NFATc1 protein in total cell lysates from day 4 WT and Plcg2–/– OCs, starved for 6 hours and stimulated with RANKL for 0, 5, 20, and 60 minutes. PLCγ1 and PLCγ2 levels are shown. β-Actin served as loading control.
PLCγ2 is required for efficient RANKL-induced activation of AP1 and NF-κB in developing OCs. To further understand whether PLCγ2 is a downstream mediator of RANKL, M-CSF, or both during OC differentiation, we analyzed MAPK activation by BMMs in culture with M-CSF for 3 days. Activation of ERK and JNK in response to M-CSF occurred normally in BMMs lacking PLCy2 (Figure 5A). Similarly, ERK was normally phosphorylated in the null cells in response to RANKL (Figure 5B). In contrast, RANKL-mediated JNK phosphorylation was retarded and reduced in the absence of PLCγ2 (Figure 5B), and activation of c-Jun (by phosphorylation) was also decreased in both the cytoplasm and the nucleus (Figure 5, B and C). This result is particularly compelling, since JNK/c-Jun activates the AP1 complex, and inhibition of JNK activity blocks RANKL-induced OC development (19). Thus, to determine whether the decreased phosphorylation of JNK observed in Plcg2–/– BMMs stimulated with RANKL could result in deficient AP1 activation, nuclear extracts from WT and Plcg2–/– cells were subjected to EMSA analysis following RANKL stimulation. The activation of AP1 was abolished in RANKL-stimulated Plcg2–/– BMMs (Figure 5D, top panel).
PLCγ2 modulates RANKL-mediated signaling. (A) Activation of ERK and JNK in WT and Plcg2–/– BMMs in response to M-CSF (100 ng/ml). PLCγ1 and PLCγ2 levels are shown. (B) Western blot analysis of phospho-ERK, phospho-JNK, phospho–c-Jun, and phospho-IκBα in total cell lysates from WT and Plcg2–/– BMMs stimulated with RANKL (100 ng/ml) for the indicated times. β-Actin blots served as loading control for A and B. (C) Nuclear levels of phospho–c-Jun and p65 in nuclear extracts from WT and Plcg2–/– BMMs stimulated with RANKL. (D) The same samples used in C were subjected to AP1 and NF-κB nonradioactive EMSA analysis. To control for binding specificity, a concentration of unlabeled oligonucleotides (U.O.) 200-fold greater than that recommended by the manufacturer was added to nuclear extracts from WT cells stimulated with RANKL for 60 minutes. SP1 served as loading control for C and D.
Since NF-κB is the other major signaling pathway required for efficient osteoclastogenesis, and PLCγ isoforms have been shown to modulate T cell and B cell receptor–mediated NF-κB activation, we hypothesized that this pathway could also be regulated by PLCγ2 in OCs. Nuclear extracts from WT and Plcg2–/– BMMs stimulated with RANKL were subjected to EMSA analysis, and data show that in the null cells, activation of NF-κB was virtually absent (Figure 5D, bottom panel). In agreement with this result, IκBα phosphorylation and nuclear translocation of p65 were also decreased in Plcg2–/– BMMs (Figure 5, B and C). Collectively, these data suggest that PLCγ2 plays a central role in RANKL-mediated osteoclastogenesis by controlling activation of AP1 and NF-κB and upregulation of NFATc1.
PLCγ2 is phosphorylated by RANKL via Dap12-mediated costimulatory signals in a Src family kinase–dependent manner. Having shown that PLCγ2 is required for the activation of osteoclastogenic signaling pathways downstream of RANKL, we turned to the mechanism by which PLCγ2 itself is activated in OCs. In immune cells, PLCγ isoforms are tyrosine phosphorylated in a Src family kinase–dependent (SFK-dependent) manner (24). Incubation of WT cells with the SFK inhibitor PP2 (5 μM) completely blocked RANKL-mediated PLCγ2 phosphorylation and NFATc1 upregulation (Figure 6A), demonstrating a concordant dependence on SFK in OC lineage cells.
PLCγ2 is activated by RANKL via Dap12/FcRγ in an SFK-dependent manner. (A) WT OC precursors (preOCs; BMMs grown in RANKL-containing media for 2 days) cultured with the SFK inhibitor PP2 (5 μM) or vehicle (DMSO) were stimulated with RANKL and subjected to Western blot analysis to detect phosphorylated levels of PLCγ2, Src, and NFATc1. β-Actin served as control. (B) PLCγ1 and PLCγ2 phosphorylation in response to RANKL were measured by Western blot analysis in WT and Plcg2–/– preOCs. PLCγ1 and PLCγ2 levels are shown. (C) PLCγ1 and PLCγ2 phosphorylation in response to 5 minutes of treatment with either M-CSF or RANKL in WT and Plcg2–/– preOCs. β-Actin served as control. (D) Expression levels of endogenous Dap12 and Flag-tagged Dap12 retrovirally transduced in Dap12–/–FcR_γ_–/– BMMs are shown. ΔKO, Dap12–/–FcR_γ_–/–. (E) PLCγ2 phosphorylation was measured by Western blot analysis in WT, Dap12–/–FcR_γ_–/–, or Dap12–/–FcR_γ_–/– preOCs reconstituted with WT Dap12 stimulated with RANKL for the indicated times. Phospho-JNK is also shown. β-Actin served as loading control. (F) Nuclear localization of NFATc1 in WT and Dap12–/–FcR_γ_–/– OCs retrovirally transduced with pMX or Flag-tagged Dap12 is shown in red (left panels). Actin staining is shown in green, and nuclei, stained with DAPI, are shown in blue (right panels) (objective, ×20). Enlarged images (2.5-fold) show nuclear localization of NFATc1 (red) and nuclei stained with DAPI (blue) of representative cells located in the center of the photographed field.
To determine whether lack of PLCγ2 was abrogating the activation of PLCγ1 in response to RANKL and M-CSF, WT and Plcg2–/– OC precursors (BMMs grown in RANKL-containing media for 2 days) were stimulated with RANKL or M-CSF. Interestingly, RANKL-mediated PLCγ1 phosphorylation was barely measurable in both WT cells and cells lacking PLCγ2 (Figure 6B) despite detectable levels of PLCγ1 protein in both cell types. In contrast, a strong PLCγ1 phosphorylation signal was observed after M-CSF stimulation in WT and Plcg2–/– cells (Figure 6C).
Since costimulatory signals are required for RANKL-mediated calcium oscillation and NFATc1 expression, we determined whether PLCγ2 phosphorylation was dependent on the RANKL costimulatory receptors Dap12 and FcRγ. We found that PLCγ2 phosphorylation and NFATc1 nuclear localization were decreased in Dap12–/–FcR_γ_–/– cells treated with RANKL (Figure 6, E and F). Reexpression of Dap12, but not empty vector, in the double null OCs (Figure 6D) completely restored PLCγ2 phosphorylation (Figure 6E) and expression of NFATc1 in the nucleus (Figure 6F). Thus, PLCγ2 is phosphorylated downstream of Dap12 in an SFK-dependent manner, following RANKL stimulation.
PLCγ2 catalytic activity is required for NFATc1 upregulation but not for JNK and IκBα phosphorylation. Our data indicate that PLCγ2 regulates NFATc1 and activates JNK and IκBα. In contrast, Dap12/FcRγ modulates NFATc1 (Figure 6F) but not JNK (Figure 6E) or IκBα (20). Thus, we hypothesized that PLCγ2 regulates JNK and IκBα independent of Dap12/FcRγ. To determine whether PLCγ2 catalytic activity is required for JNK and IκBα phosphorylation, WT OC precursors, cultured in the presence of the PLC inhibitor U73122 (5 μM), were stimulated with RANKL for 0 to 60 minutes (Figure 7A). While dampening NFATc1 upregulation, the inhibitor had no effect on p-JNK and p-IκBα, suggesting that the catalytic activity of PLCγ2 is not required for phosphorylation of JNK and IκBα.
PLCγ2 forms a complex with GAB2 and modulates its activation. (A) WT preOCs cultured with or without the PLC inhibitor U73122 (5 μM) for 3 days were stimulated with RANKL and subjected to Western blot analysis for phospho-JNK, phospho-IκBα, and NFATc1. β-Actin served as control. (B) WT and Plcg2–/– BMMs retrovirally transduced with empty vector (pMX), WT PLCγ2, or catalytically inactive PLCγ2 (PLCγ2 H/F) were cultured with RANKL (100 ng/ml) and M-CSF (10 ng/ml) for 7 days, and multinucleated OCs were detected by TRAP staining. Objective, ×10. (C) The same cells as shown in B were subjected to RANKL stimulation and Western blot analysis to detect NFATc1 expression. (D) Plcg2–/– BMMs retrovirally transduced with pMX, WT PLCγ2, or PLCγ2 H/F were treated with RANKL, and phosphorylation of IκBα and JNK was determined by Western blot analysis. PLCγ2 expression levels are shown. β-Actin served as loading control in C and D. (E) PLCγ2 and GAB2 were reciprocally immunoprecipitated in WT and Plcg2–/– BMMs treated with RANKL and subjected to Western blot analysis using anti-PLCγ2 and anti-GAB2 Abs, respectively. TCL, total cell lysate. (F) GAB2 was immunoprecipitated in WT and Plcg2–/– BMMs and subjected to Western blot analysis using anti-phosphotyrosine Ab (clone 4G10), anti-RANK, and anti-PLCγ2.
Mutations of 2 histidine residues, conserved among all PLC family members, have been shown to inhibit the enzymatic activity of PLCγ1 by 90%–95% in vitro (35). Thus, we generated a PLCγ2 catalytic inactive construct carrying the double histidine 327/372 to phenylalanine mutation (PLCγ2 H/F). This mutant was proven to be lipase inactive compared with WT full-length PLCγ2 in 293 cells (data not shown). Plcg2–/– cells transduced with PLCγ2 H/F were incapable of generating mature, TRAP-positive OCs compared with cells expressing WT PLCγ2 (Figure 7B) or of promoting NFATc1 upregulation (Figure 7C). In contrast, the ability of PLCγ2 H/F to mediate JNK and IκBα phosphorylation was indistinguishable from that of WT PLCγ2 (Figure 7D). These data indicate that PLCγ2 can modulate RANK-mediated signaling independent of its catalytic activity by acting as an adapter molecule. One potential interacting protein is GAB2, which associates with RANK and mediates RANK-induced activation of NF-κB and JNK but not NFATc1 (16). Furthermore, GAB2 has been shown to bind PLCγ2 in mast cells (36). To determine whether PLCγ2 binds to GAB2 in OCs, we reciprocally immunoprecipitated GAB2 and PLCγ2 and found that the 2 molecules associated (Figure 7E). Immunoprecipitation with IgG control antibodies revealed the absence of nonspecific bands (data not shown). To further determine whether PLCγ2 modulates GAB2 activation or its recruitment to RANK, WT and Plcg2–/– BMMs, treated with RANKL, were immunoprecipitated with an anti-GAB2 Ab (Figure 7F) or with rabbit IgG control Ab (data not shown), followed by immunoblotting with phospho-tyrosine (4G10) and RANK. Results showed that PLCγ2 modulates GAB2 phosphorylation and its association with RANK, suggesting that this may be the mechanism by which PLCγ2 controls AP1 and NF-κB activation.
TNF-α cannot reverse the arrested capacity of PLCγ2-null cells to became OCs. The data presented above show that PLCγ2 is central in RANKL signaling. In OCs, TNF-α appears to be the dominant cytokine mediating inflammatory osteolysis and augments the osteoclastogenic potential of RANKL-exposed OC precursors (3). Due to the capacity of TNF-α to phosphorylate JNK and IκBα in BMMs and OCs (37), we tested its effect in modulating the above-mentioned signaling pathways in cells lacking PLCγ2. TNF-α–mediated phosphorylation of JNK and IκBα occurred similarly in WT and null cells (Figure 8A). Furthermore, TNF-α was not capable of inducing PLCγ2 phosphorylation (Figure 8A), suggesting that the mechanism by which TNF-α regulates JNK and IκBα is PLCγ2 independent. This result prompted us to determine whether TNF-α could rescue the osteoclastogenic defect of Plcg2–/– cells. Interestingly, while TNF-α augmented RANKL-mediated osteoclastogenesis of WT cells (WT OCs without TNF-α, 19.3 ± 1.5/well; WT OCs with TNF-α, 45.3 ± 1.99/well), the cytokine did not have any effect on the null cells, which remained blocked at the early stage of OC development (Plcg2–/– OCs, not detected) (Figure 8B). Consistent with this finding, TNF-α, compared with RANKL, was not capable of promoting NFATc1 upregulation in WT or in PLCγ2-null cells (Figure 8C).
TNF-α cannot correct the Plcg2–/– OC defect. (A) WT and Plcg2–/– BMMs were stimulated with TNF-α (10 ng/ml) with time. Phosphorylated JNK, IκBα, and PLCγ2 were examined. β-Actin served as control. (B) WT and Plcg2–/– BMMs were cultured for 3 days with RANKL (100 ng/ml) and M-CSF (10 ng/ml). On day 3 TNF-α was also added to the culture media. Cells were fixed and TRAP stained at day 7. Magnification, ×200. (C) WT and Plcg2–/– preOCs were stimulated with TNF-α or RANKL for the indicated times, and expression levels of NFATc1, PLCγ2, and β-actin were determined.