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Notch signaling is a central mechanism for controlling embryogenesis. However, its in vivo function during mesenchymal cell differentiation, and specifically, in bone homeostasis remains largely unknown. Here, we show that osteoblast-specific gain of Notch function causes severe osteosclerosis due to increased proliferation of immature osteoblasts. Under these pathological conditions, Notch stimulates early osteoblastic proliferation by up-regulating Cyclin D, Cyclin E, and Osterix. Notch also regulates terminal osteoblastic differentiation by directly binding Runx2 and repressing its transactivation function. In contrast, loss of all physiologic Notch signaling in osteoblasts, generated by deletion of Presenilin 1 and 2 in bone, is associated with late onset, age-related osteoporosis resulting from increased osteoblast-dependent osteoclastic activity due to decreased production of Osteoprotegerin. Together, these findings highlight the potential dimorphic effects of Notch signaling in bone homeostasis and may provide direction for novel therapeutic applications.
Evolutionarily conserved Notch signaling plays a critical role in cell fate determination, and various developmental processes by translating cell-cell interactions into specific transcriptional programs1, 2. Temporal and spatial modulation of this pathway can significantly affect proliferation, differentiation and apoptotic events3. Moreover, the timing of Notch signaling can lead to diverse effects within the same cell lineage 4, 5. In mammals, activation of up to four Notch receptors by membrane-bound ligands initiates a process leading to presenilin-mediated cleavage and release of the Notch intracellular domain (NICD) from the membrane that then traffics to the nucleus. NICD subsequently regulates the expression of genes in cooperation with the transcription factor RBP-Jκ and Mastermind-like proteins.
The observation that mutations in the Notch ligand Delta homologue-3 (Dll-3) and γ-secretase Presenilin1 both cause axial skeletal phenotypes originally linked Notch signaling with skeletal development6, 7. Recently, several in vitro studies with conflicting results implicated the Notch pathway in the regulation of osteoblast differentiation, but the in vivo role of Notch signaling in bone homeostasis still remains unknown8–12.
In this study, we investigate the tissue, cellular, and molecular consequences of both gain and loss of function of Notch signaling in committed osteoblasts.
To determine the pathological consequences of in vivo gain of Notch function during bone formation and homeostasis, we generated transgenic mice expressing the Notch1 intracellular domain (N1ICD) under the control of the type I collagen (Col1a1) promoter (Suppl. Fig. 1a,b). Here, gain of Notch function would occur in committed osteoblastic cells since this marker is both an early and late marker of the osteoblastic lineage. Interestingly, founder mice expressing high levels of the transgene, were small at birth and showed progressive growth retardation. Analysis of three established lines showed increased bone mass on radiographs at 4 weeks and a thickened, osteosclerotic appearance following skeletal preparations (Fig. 1a and Suppl. Fig. 1c). Histologically, marrow spaces in 4 week-old transgenic mice were largely filled with trabecular bone composed predominantly of immature woven rather than lamellar bone and surrounded by fibrotic marrow containing cells with morphologic features of early osteoblastic precursors, suggesting increased proliferation of these cells (Fig. 1b). The cortices of the bones were also composed of woven bone, and this phenotype was also present in 11 week-old mice. Toluidine blue staining of 11 week-old transgenic mice indicated increased number of osteoblasts (Fig. 1c). Quantitative histomorphometry of an established mouse line confirmed the significant increase in trabecular bone volume and osteoblast surface, consistent with the high bone mass being due to increased osteoblastic activity (Fig. 1d). This increased osteoblastic activity led to increased production of osteoid (Fig. 1d) and bone formation (Suppl. Fig. 1d)
Since bone formation and resorption are coupled in vivo, we analyzed the status of osteoclasts by staining for tartrate-resistant acid phosphatase (TRAP) activity in osteoclasts from bone sections of 4 week-old mice. Although total TRAP staining was qualitatively increased in the limb sections of transgenic mice (Suppl. Fig. 1e), consistent with increased bone mass and remodeling, the osteoclastic parameters normalized to bone surface, i.e., osteoclast number per millimeter of bone surface and osteoclast surface, were significantly decreased in trabecular bone of transgenic mice (Suppl. Fig. 1e). Together, these data support that gain of Notch function in committed osteoblastic lineage cells stimulates the proliferation of early osteoblastic precursors that differentiate into immature osteoblasts producing increased amounts of immature woven bone. While osteoclastic activity was secondarily stimulated by this massive osteoblastic proliferation, bone formation much greatly outweighed bone resorption leading to an osteosclerotic phenotype.
To determine the underlying cellular and molecular mechanism for the increase in early osteoblastic precursors in transgenic bone, we cultured P6 calvarial osteoblasts and found significantly increased numbers of BrdU-positive cells, consistent with increased cellular proliferation (Fig. 1e). Quantitative real time RT-PCR (Q-RT-PCR) of 1 month-old calvarial RNA showed an increased abundance of early osteoblastic differentiation markers, including Osterix (Osx), Alkaline Phosphatase (Alp) and Bone Sialoprotein (Bsp). In contrast, later markers of osteoblast differentiation, including osteocalcin were down-regulated (Fig. 1f). To exclude that the increased bone mass was due to decreased osteoclastic activity, we assessed the expression of markers that regulate macrophage differentiation along the osteoclastic lineage in the forelimbs of 4 weeks-old transgenic mice. RANK Ligand (Rankl), Osteoprotegerin (Opg) and Macrophage Colony Stimulation Factor (M-Csf) were all highly expressed suggesting that the hyper-proliferation of the early osteoblastic pool was associated with increased production of both pro- (Rankl and M-CSF) and anti-osteoclastic (Opg) differentiation factors by osteoblasts (Fig. 1g). Hence, on both histological and molecular levels, gain of Notch signaling in committed osteoblastic precursors resulted in their proliferation as well as secondary stimulation of differentiation of the monocytic lineage of osteoclastic precursors, but far fewer than the magnitude osteoblast proliferation. Thus, the net effect is an increase of immature woven bone formation resulting in severe clinical osteosclerosis.
How Notch signaling regulates these processes on a biochemical level is unknown. Osteoblast differentiation from mesenchymal stem cells and subsequent maturation steps require the function of the runt domain transcription factor Runx2 and the zinc finger transcription factor Osterix. Runx2 is required for commitment of mesenchymal osteochondroprogenitors to the osteoblastic lineage, differentiation into mature osteoblasts, and terminal differentiation into osteocytes. In contrast, Osx is important in expansion of the early osteoblastic pool19. While Bsp and Alp are markers of early osteoblasts, Osteocalcin is a marker of later, mature osteoblasts. To determine the mechanistic basis of Notch action in this context, we tested the effects of Notch expression on these key transcriptional regulators of osteoblast differentiation and maturation. Notch1 ICD alone was able to directly bind Runx2 and repress its transactivation of a reporter Osteocalcin enhancer in vitro (in Cos7 and in rat osteosarcoma Ros17/2.8 cells) (Fig. 2a–c, Suppl. Fig. 1f). Electrophoretic mobility shift assays (EMSA) showed that NICD could inhibit RUNX2 binding to a target cis element in the Type X collagen promoter (Suppl. Fig. 1g). Interestingly, there was significant down-regulation of Runx2 protein in P2 calvaria of transgenic mice (Fig. 2d). Hence, the down- regulation of Osteocalcin and the delay in late osteoblast differentiation in vivo is likely due in part, to direct repression of Runx2 by Notch at the protein level. At the same time, we observed up-regulation of Osx expression in the P2 calvaria of transgenic mice. Moreover, Notch1 ICD activated the Osx promoter in transient transfection studies in C2C12 cells that were induced to differentiate into osteoblasts with BMP2 treatment (Fig. 2e). These data suggest that Notch can induce proliferation of committed osteoblast precursors by directly up-regulating transcription of Osx, while it inhibits their maturation by repressing the function of Runx2.
To further understand the biochemical basis of Notch on osteoblastic proliferation, we analyzed the expression of cell cycle markers and detected increased RNA expression of Cyclin D and Cyclin A by Q-RT-PCR in osteoblasts over-expressing Notch1 ICD (Fig. 2f). This correlated with increased Cyclin D and Cyclin E expression at the protein levels (Fig. 2g). We did not, however, observe significant differences in the levels of two other important cell cycle regulators implicated in bone homeostasis, p53, and Rb. Interestingly, it has been shown with in vitro and ex vivo studies that Runx2 can suppress osteoblast proliferation, and promote osteoblast maturation by supporting exit from the cell cycle20, 21. Moreover, CyclinD1-Cdk4 can induce Runx2 ubiquitination and degradation, thus Runx2 activity can be regulated by the cell cycle machinery22. Hence, gain of Notch can inhibit osteoblast maturation by direct repression of Runx2 function as well as by repressing Runx2’s anti-proliferative effects via Cyclin D1 up-regulation.
To determine if the pathological effects of gain of Notch signaling reflect a physiological function during bone homeostasis, we generated a tissue-specific model of loss of Notch signaling in osteoblasts. Because all Notch receptors are expressed in osteoblasts (data not shown), we abolished Notch signaling by generating null mice for both Presenilin1 (Ps1) and Presenilin 2 (Ps2). Since Ps2 null mice are viable and fertile, we generated double homozygotes for the Ps2 null allele and the Ps1 floxed allele, but heterozygous for the type I collagen Cre recombinase transgene (Ps1 f/f; Ps2 −/−/; Col1a1Cre/+ or DKO). DKO mice were compared to their Ps1 f/f; Ps2 −/− littermates as controls and efficient deletion of the Ps1 f/f allele (approximately 92%) was confirmed by RT-PCR for Presenilin RNA expression and genomic PCR for DNA recombination in calvarial osteoblasts and tail DNA, respectively (Suppl. Fig. 2a,b). Moreover, we confirmed that this led to decreased Notch1 ICD processing on western analysis (Suppl. Fig. 2c). Histomorphometric analyses of 6 month-old, but not 3 month-old, DKO mice showed they were osteoporotic, a decreased tissue bone mass phenotype opposite that of the osteosclerotic tissue phenotype in gain of Notch function transgenic mice (Fig. 3a–c). Bone formation rates (BFR), osteoblast surfaces (Ob.S/BS), and mineralized surfaces (MS/BS) in vertebrae and long bones in the DKO mice were similar to those in control mice (Suppl. Fig. 3). However, osteoclast numbers, osteoclast surfaces, and eroded surfaces were increased in DKO vertebrae and long bones at only 6 months but not at 3 months (Fig. 3d,e; Suppl. Fig. 3). These findings suggest that loss of Ps1 and Ps2, and hence, all Notch signaling in osteoblasts, led to osteoporosis through activation of osteoclastogenesis, resulting in increased bone resorption over bone formation with age-related penetrance.
Activated osteoblasts support osteoclast formation and differentiation from osteoclast precursors (OCPs) by expressing M-CSF and RANKL, but they also inhibit this process through Osteoprotegerin (Opg), which binds to and inactivates RANKL. To further examine the effects of loss of Ps1/Ps2 in osteoblasts on osteoclastogenesis, we performed osteoblast/osteoclastic precursor (OCP) co-culture studies. In this ex vivo assay, P7 DKO calvarial osteoblasts stimulated formation of more osteoclasts from wild type spleen-derived OCPs than did wild type osteoblasts, suggesting that Ps1/Ps2 deletions can affect osteoclastogenesis in a non-cell-autonomous fashion (Fig. 4a). To determine whether this effect was specific for Notch signaling, we tested whether heterologous expression of Notch1 ICD after lentiviral transduction of DKO osteoblasts could suppress osteoclastogenesis in co-cultures studies (Fig. 4b, Suppl. Fig. 4a). Compared to control vector expressing EGFP, Notch1 ICD lentiviral transduction of Ps1/Ps2 mutant osteoblasts was able to suppress osteoclastogenesis suggesting that the DKO phenotype was due primarily to Presenilin activation of Notch signaling.
The in vivo relevance of this was confirmed by flow-cytometric marker analysis of bone marrow cells from 3 month-old DKO mice. This showed increased staining of early OCPs in total cells (cFMS+) and in more differentiated OCPs (CD11b+/Gr-1−/lo) compared to controls, indicating an expansion of the OCP pool in DKO mice (Fig. 4c). To determine if this increase of osteoclast differentiation was due to an imbalance of osteoblastic inductive (Rankl and M-csf) vs. suppressive signals (Opg), we analyzed their RNA expression in DKO vs. control bone at P4. We found comparable expression of Rankl in DKO mice, but expression of Opg was significantly decreased (Fig. 4 d). Similarly, we found decreased Opg production cultured DKO vs. WT calvarial osteoblasts (Suppl. Fig 4b). Hence, under physiological conditions, Notch signaling enabled by Ps1/Ps2 function in osteoblasts represses osteoclast differentiation by regulating Opg expression.
Together, these in vivo gain and loss of function studies support for the first time a central role of Notch and Presenilin signaling in regulating both osteoclastogenesis and immature osteoblastic proliferation during bone homeostasis (Fig. 5).
Until now, few primary signaling mechanisms regulating osteoblast differentiation and function during bone homeostasis have been identified in vivo by genetic and biochemical studies. Wnt signaling via LRP5/6 co-receptors and canonical β-catenin activity are required for osteoblast lineage commitment and function23–25. Activation of this pathway leads to high bone mass26, 27. Activating mutations in TGFβ in humans is associated with increased bone formation and inhibition of bone resorption28. However, not unexpectedly, apparently discrepant results in vivo have been observed depending on the timing of gain vs. loss of TGFβ function. Similarly, Notch signaling likely exhibits temporal and spatial dependence.
In bone, our data suggest that Notch and Presenilin signaling may be important in the physiological regulation of osteoclastogenesis by osteoblasts. Moreover, it raises the question of whether loss of Notch signaling contributes to age-related osteoporosis, since this type of osteoporosis is associated with increased resorption over bone formation as is seen in our DKO model29. We discovered that the one function of Notch in committed osteoblasts is to regulate osteoclastogenesis via regulation of Opg production. The magnitude of Opg dysregulation and the age-related penetrance of the osteoporosis in the loss of function mouse phenotype correlate well with epidemiological data in humans where age-related osteoporosis has been associated with changes in OPG production30–33. Furthermore, the report that heterozygote Opg mutant mice exhibit an age-related osteoporotic phenotype suggests that this mechanism is sufficient for disease pathogenesis34. What is unclear is whether Opg dysregulation is due to direct regulation by Notch1 ICD or by its target transcription factors, given the still poorly characterized Opg regulatory region. Further studies showing chromatin immunoprecipitation (ChIP) analysis on a well defined functional OPG promoter with Notch1 ICD or with it target genes would help to address this issue. Similarly, our studies do not address the potential role of Notch signaling prior to osteoblastic commitment in the mesenchymal stem cell (Fig. 6). Here, Runx2 plays the central role in osteoblastic commitment. Our data on the Notch-Runx2 interaction suggest that early loss of function of Notch would actually lead to increase commitment to the osteoblastic lineage and perhaps depletion of the mesenchymal stem cell compartment.
In a pathological disease context, our findings show that activation of Notch signaling in the committed osteoblastic lineage leads to expansion of an immature osteoblast pool. The primary mode of action is transcriptional up regulation of the early osteoblast transcription factor Osterix, and increase of Cyclin D and E proteins. These data raise the question of the potential contribution of activation of Notch signaling in human diseases related to osteoblastic proliferation such as in bone pathologies like human osteosarcomas. The significant up-regulation of Cyclin D1 in the transgenic mice correlates with the observation in humans where 10% of osteosarcomas show amplification of the chromosomal region encoding Cyclin D135. While our data suggest that Notch can directly interact with Runx2 to inhibit its binding to target cis elements and its pro-differentiation function, this is not likely the main determinant of the gain of function phenotype in mice.
Finally, our data have important therapeutic implications. There are few anabolic bone agents for the treatment of osteoporosis, with most therapies targeted at inhibition of bone resorption. Up-regulation of Notch signaling may represent a potential approach for increasing bone formation over bone resorption as well as for inhibiting osteoclastogenesis. However, it is clear that temporal effects of Notch on other cellular compartments such as the mesenchymal stem cell pool would have to be considered, i.e., Notch inhibition of Runx2 function could inhibit mesenchymal stem cell commitment to the osteoblastic lineage. Second, in opposing fashion, inhibition of Notch signaling may be a therapeutic option to investigate for the treatment of proliferative disorders of the osteoblast such as in osteosclerotic diseases or bone cancers.
From a mechanistic perspective, the function of Notch signaling in bone constitutes a rare example of a signaling pathway capable of regulating both osteoblastic and osteoclastic lineages but differently when considering gain of function vs. loss of function scenarios. The other in vivo example for this is Ephrin B2 signaling where reverse signaling through ephrin B2 ligand expressed by osteoclasts suppress osteoclast precursors, whereas forward signaling through EphB4 receptor expressed by osteoblasts enhances osteoblast formation38,39. Together, our data point to a dimorphic role for Notch signaling in osteoblast biology, i.e., the stimulation of osteoblastic precursors in a pathological context, and the inhibition of osteoclastogenesis in the physiological regulation of bone mass and homeostasis.
Myc-His tagged Notch1 ICD including amino acid position 1760–2556 (gift of Tom Kadesch) was cloned under the control of 2.3 kb osteoblast specific Col1a1 promoter in a coat color vector containing tyrosinase minigene and the WPRE posttranscriptional sequences40. Transgenic founders were generated by pronuclear injections according to standard techniques. All transgenic lines were maintained on a FVB/N background. The transgenic mice were identified at birth by eye pigmentation and confirmed by PCR using primers specific for the WPRE. Previously published Ps1f/f and Ps2 −/− mice were crossed with Col1a1-Cre mice (Gerard Karsenty), to generate osteoblast specific Ps1/Ps2 double knock out (DKO) mice.
We cleared and stained skeletons from 1 month-old mice with alcian blue for cartilage and alizarin red for bone as described41. Mice were sacrificed, and the whole skeleton was fixed in 10% neutral-buffered formalin for 18 hours. For radiographic analyses, the skeletons were analyzed by contact radiography with a Faxitron X ray cabinet (Faxitron Xray Corp., Wheeling, IL). Paraffin embedded tissues were sectioned at 4–7μM thickness, and stained with hematoxylin and eosin. Toluidine blue, Von Kossa and Goldner’s stains were performed on 5–7 micron undecalcified lumbar vertebral plastic sections by using standard protocols. All static and dynamic histomorphometry analyses were performed according to standard protocols using the OsteoMeasure histomorphometry system (Osteometrics Inc. GA). Histomorphometric analyses were performed on 4 week-old transgenic mice and 6 month-old knockout mice with n=3 and n=5–7, respectively in each group. Micro-CT scanning of the trabecular bone of the distal femur was analyzed by the micro-CT system (μCT-40; Scanco Medical, Bassersdorf, Switzerland).
Osterix-luciferase was a gift of Mark S. Nanes. For lentivirus vector production, plasmids pHIV-N1-IRES-eYFP was constructed by inserting a FLAG-tagged version of intracellular activated form of Notch1 (N1), just upstream of the 1.4 kb IRES-eYFP cassette of pHIV-IRES-eYFP42. For the primary osteoblasts, lentiviral vectors used were self-inactivating and had the 0.5 kb mouse phosphoglycerate kinase (PGK) promoter inserted upstream of either N1-IRES-eYFP or IRES-eYFP43. VSV G-pseudotyped vector supernatants were produced as previously described44. After 72 hours, cell culture supernatants were harvested and clarified. Typical titers after concentration by ultracentrifugation were in excess of 108 IU/ml, for the two SIN vectors as assessed on HOS cells by epifluorescence microscopy. Titers of unconcentrated non-SIN vectors were in excess of 107 IU/ml.
Calvarial osteoblasts from one week-old mice (n=3) were co-cultured with spleen cells at a number of 5×103 and 5×104 per well respectively in 96-well plates for 7 days in the presence 10−8 M of VitD3. The cells were then stained for TRAP activity, and counted as described previously45. For the lentiviral rescue experiment 5×103 osteoblasts isolated from calvaria of 10 day-old Ps1/Ps2 DKO mice were cultured in a 96-well plate for 2 days. The cells were then either infected with 5 μl Notch1 ICD lentivirus or the YFP- lentiviral vector supernatant for 24 hours in 100 μl α-MEM containing 10 % FBS and 8 μg polybrene/ml. The infected cells were then co-cultured with 5×104 spleen cells from 10 day-old WT mouse for 7 days in the presence of 10−8 M vitamin D3.
After lysis of erythrocytes with ammonium chloride solution, 2×106 cells of bone marrow or spleen were incubated 5 minutes with anti-murine CD16/32 to block Fc receptor-mediated antibody binding followed by triple staining of anti-mouse CD11b-APC, and Gr-1-FITC and c-Fms-PE antibodies for 30 minutes. The cells were then subjected to FACS to analyze the CD11b+/Gr-1−/lo cells that contain osteoclast precursors and c-Fms+ cells in both total gated and CD11b+/Gr-1−/lo population.
Osteoblasts from calvaria of P6 transgenic mice and wild type littermates (n=5 each group) were isolated as previously described41. 48 hours after the initial culture, cells were re-plated and expanded an additional day. Cells were treated with BrdU labeling reagent according to manufacturer’s instructions for 6 hours, washed with PBS, and fixed with 70% ethanol for 25 minutes at 4°C (Zymed). Three to five areas for each genotype (n=3 slides) were counted by two independent observers blinded to genotype. BrdU positive cells over total cells were scored visually and with Automeasure software (Zeiss Axiovision).
Proteins were extracted from P2 mice by homogenizing the calvaria (n=3 each group) in a buffer containing 5% SDS, and 0.0625 M Tris.HCl. Western blot analyses were performed by using, anti-P53 antibody (gift of Larry Donehower), anti-Runx2 PEBP2αA (M70) polyclonal antibody (Santa Cruz Biotechnology), anti-CyclinD1 antibody H-2953 (Santa Cruz Biotechnology), anti-cyclin E antibody ab-7959 (Abcam). Protein content was normalized with anti-γ-tubulin mouse monoclonal antibody (Sigma).
Glutathione-S-transferase (GST), GST-NICDTAD, and GST-NICDRA (gift of Tom Kadesch) were expressed in the BL21 strain of Escherichia coli (Stratagene). GST proteins were induced with 0.2 mM isopropyl-b-d-thiogalactopyranoside (IPTG) (Promega), and the bacteria were allowed to grow an additional 4 to 5 hours. Following induction, cells were lysed by sonication. GST proteins were bound to glutathione resin (Amersham Bioscences, NJ). 625μM Methionine-labeled flag tagged Runx2 proteins were generated by a T7 in vitro transcription/translation kit (Novagen) and incubated with GST or GST-NICDTAD, GST-NICDRA immobilized on glutathione-Sepharose beads at 4°C for 2 hours. The beads were then washed five times with TNN buffer containing 1% NP40, boiled in 2×SDS sample loading buffer, and separated by SDS-PAGE. Western blot was performed to detect the Flag- tagged Runx2 protein, by using anti-flag M2 monoclonal antibody (Sigma).
Total RNA was extracted using TRIzol reagent (Invitrogen) from calvaria and forelimbs of P4 and 4 week-old mice (n=5 and n=3 each group, respectively). cDNAs were synthesized from extracted RNA by using Superscript III First Strand RT-PCR kit (Invitrogen). Real-time quantitative PCR amplifications were performed on LightCycler (Roche) and with TaqMan assay (Applied Biosystems probe HS00172878-M1). β-actin and β2-microglobulin genes were used as internal controls for the quantity and quality of the cDNAs in real time PCR assays.
Cos7 and Ros17/2.8 cells were transfected with the 6XOSE2-luc reporter gene by using Lipofectamine Plus according to manufacturer’s recommendations (Invitrogen). Luciferase and β-galactosidase activities were assayed 48 hrs after transfection. C2C12 cells were transfected with –1269/91 Osx-p-luc (gift of Mark S. Nanes) by using Fugene6 according to manufacturer’s instruction (Roche). 24 hrs after the transfection, the cells were induced with 300ng/ml recombinant human BMP-2 (R&D Systems), and cells were harvested and assayed the following day. All transfections were performed in triplicates with pSV2βgal as an internal control for transfection efficiency.
Data are expressed as mean values ± standard deviation (SD). Statistical significance was computed using Student’s paired t test. A P value < 0.05 was considered statistically significant.
Total RNA was extracted using the TRIzol reagent (Invitrogen) from P7 osteoblast cultures. cDNAs were synthesized by using Superscript III First Strand RT-PCR kit (Invitrogen). Primers specific to Ps1 exon 4 (5′CTTGACAACCCTGAGCCAAT3′) and exon7 (5′GAAATCACAGCCAAGATGAGC3′) were used to to amplify the Ps1 floxed allele (422bp). The β-actin gene was used as an internal control of the quantity and quality of the cDNAs.
Proteins were extracted from P2 transgenic mice or P4 Ps1 DKO control mice by homogenizing the calvaria (n=3 each group) in a buffer containing 5% SDS, and 0.0625 M Tris-HCl. Western blot analyses were performed by using anti-Notch1 ICD antibody Val 1744 (Cell Signaling). Protein content was normalized with anti-γ-tubulin mouse monoclonal antibody (Sigma). For EMSA, labeling of oligonucleotide probes, incubation of in vitro translated proteins, and EMSA were performed as previously described40.
HeLa cells (6×106 cells/dish) were plated in 10 cm dishes in DMEM+10%FBS for transient transfection. Forty-eight hours later, cells were harvested and lysed in lysis buffer (20 mM Tris pH 8.0, 200 mM NaCl, 0.5% Triton X-100) supplemented with protease inhibitors. Lysates were subjected to immunoprecipitation with anti-FLAG (Sigma), anti-Myc (Invitrogen), anti-Mouse IgG (Santa Cruz) and protein G agarose at 4°C overnight. Immunoprecipitates were then washed three times in lysis buffer and subjected to SDS-PAGE followed by western blot analysis for anti-FLAG (Sigma) or anti-Myc (Invitrogen) antibodies.
Skeletons from 2-day old mice were prepared as described and stained with alcian blue 8GX for cartilage and alizarin red S for bone41. Toluidine blue and TRAP staining were performed on 5–7 micron undecalcified lumbar vertebral plastic sections by using standard protocols. All static and dynamic histomorphometric analyses were performed according to standard protocols using the OsteoMeasure histomorphometry system (Osteometrics Inc. GA). Double labeling was performed by intraperitoneal calcein (Sigma) injection twice with an interval of 7 days. Mice were sacrificed 2 days after the last injection. Calcein labeling was assessed in the vertebrae using formalin fixed undecalcified 5–7 micron-thick plastic sections. For ELISA, the co-culture medium was collected at day 2 and day 4 and analyzed according manufacturer’s protocol (R&D System).
Supplemental Figure 1. Histomorphometry in osteoblast-specific Notch gain of function transgenic mice. a, Transgenic construct expressing Notch1 ICD. b, Expression of the transgene by Q-RT-PCR, and western blot of calvarial extracts showing increased Notch1 ICD RNA and protein in transgenic mouse line Tg1. c, Skeletal preparations from 3 different transgenic lines (Tg1, Tg2, Tg3) expression Notch1 ICD. Arrows indicate the thickening of the ribs, clavicles and humerus. Scale bar, 500 mm. d, Calcein labeling of 4 week-old spinal trabeculae of WT and Tg mice showing dramatically increased calcein labeling due to increased but disorganized woven bone formation. Scale bar, 100 μm. e, TRAP stained hind limbs of 4 week-old transgenic and wild type mice show absolute increase in relative staining consistent with absolute increase in number of osteoclasts. Scale bar, 500 μm. However, histomorphometry of 4 week-old transgenic hind limbs (n=3) stained with TRAP shows decreased osteoclast number (Oc.N) and surface (Oc.S) per bone surface (BS) in secondary spongiosa of tibia consistent with relative decrease in osteoclast per area of bone in transgenic vs. wild type mice. * p<0.05 between WT and Tg. f, Notch1 ICD co-immunoprecipitates with Runx2. HeLa cells were transfected with plasmids expressing either Myc-His epitope tagged Notch1 or Flag-tagged Runx2. Upper panel: Immunoprecipitation (IP) performed using anti-Myc antibody followed by Western blot (WB) with anti-Flag antibody. Lower panel: IP with anti-Flag antibody followed by WB with anti-Myc antibody. g, Notch1 ICD represses DNA binding of Runx2 in EMSA. COL10A1 promoter element was bound to: : Lane1, no protein; lane 2, RUNX2 alone; lanes 3–5, RUNX2 with increasing amounts of in vitro transcribed-translated Notch1 ICD (lane 3: 1X, lane 4: 2X; lane 5: 3X); lane 6, in vitro transcribed-translated luciferase; lane 7, mutated probe which is unable to bind Runx2. Runx2-DNA protein complex is shown by arrow. Free probe is below. Increasing concentration of Notch1 ICD decreases formation of Runx2-DNA complex. Negative control luciferase protein had no effect on this complex.
Supplemental Figure 2. Normal osteoblast formation in Ps1/Ps2 deleted mice. a, Genomic PCR showing Ps1, Ps2, and Cre alleles from tail DNA of various genotypes. b, Semi- quantitative RT-PCR for Ps1 expression from Ps1f/f deletion using RNA from cultured P7 osteoblastic cells. cDNA product from floxed (undeleted) Ps1 allele is noted by the band (Ps1) at increasing PCR cycles. cDNA content was normalized with β-actin. C; Control, D; DKO. c, Western blot shows the expression of Notch1 ICD in Ps1/Ps2 DKO or single knockout Ps1f/f; Ps2−/− (SKO) mice.
Supplemental Figure 3. a, 6 month-old (n=7) lumbar vertebral sections stained with toluidine blue and doubly-labeled with calcein showed no significant changes in osteoblast surface (Ob.S), mineral apposition rate (MAR), and bone formation rate (BFR), respectively. b, 3 month-old (n=6) lumbar vertebral sections stained with toluidine blue and doubly-labeled with calcein showed no significant changes in mineral apposition rate (MAR), bone formation rate (BFR) and mineralized surface per bone surface (MS/BS), respectively. c, Histomorphometry of 6 month-old (n=6) mice lumbar vertebrae revealed decreased bone volume (BV/TV), osteoclast number per bone surface (N.Oc/BS), and osteoclast surface per bone surface (Oc.S/BS) in DKO mice. * p<0.05 between WT and DKO.
Supplemental Figure 4. a, Western analysis using anti-FLAG antibody on lysates of 293 cells infected with lentivirus expressing NICD (right lane) tagged with FLAG or with control GFP expressing virus (left lane). b, ELISA measurement of OPG levels in pooled medium of Ps1/Ps2 null (DKO) vs. wild type (WT) calvarial osteoblasts from co-culture experiments with wild type splenocytes after 2 and 4 days of culture.
We thank Melih Acar and Olga Sirin for technical assistance. This work was supported by NIH grants ES11253 (B. Lee), HD22657 (B. Lee), DE016990 (B. Lee), AR43510 (B. Boyce).