Smad9 is a new type of transcriptional regulator in bone morphogenetic protein signaling

Tsukamoto, S.; Mizuta, T.; Fujimoto, M.; Ohte, S.; Osawa, K.; Miyamoto, A.; Yoneyama, K.; Murata, E.; Machiya, A.; Jimi, E.; Kokabu, S.; Katagiri, T.

doi:10.1038/srep07596

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Published: 23 December 2014

Smad9 is a new type of transcriptional regulator in bone morphogenetic protein signaling

S. Tsukamoto¹,
T. Mizuta¹,
M. Fujimoto¹,
S. Ohte¹,
K. Osawa¹,
A. Miyamoto¹,
K. Yoneyama¹,
E. Murata²,
A. Machiya¹,
E. Jimi³,
S. Kokabu^1,4 &
…
T. Katagiri¹

Scientific Reports volume 4, Article number: 7596 (2014) Cite this article

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Abstract

Smad1, Smad5 and Smad9 (also known as Smad8) are activated by phosphorylation by bone morphogenetic protein (BMP)-bound type I receptor kinases. We examined the role of Smad1, Smad5 and Smad9 by creating constitutively active forms (Smad^DVD). Transcriptional activity of Smad9^DVD was lower than that of Smad1^DVD or Smad5^DVD, even though all three Smad^DVDs associated with Smad4 and bound to the target DNA. The linker region of Smad9 was sufficient to reduce transcriptional activity. Smad9 expression was increased by the activation of BMP signaling, similar to that of inhibitory Smads (I-Smads) and Smad9 reduced BMP activity. In contrast to I-Smads, however, Smad9 did not inhibit the type I receptor kinase and suppressed the constitutively active Smad1^DVD. Smad9 formed complexes with Smad1 and bound to DNA but suppressed the transcription of the target gene. Taken together, our findings suggest that Smad9 is a new type of transcriptional regulator in BMP signaling.

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Introduction

Bone morphogenetic proteins (BMPs) are the largest subfamily of the transforming growth factor-β (TGF-β) family, which also includes growth and differentiation factors (GDFs) and activin. These proteins have multiple functions that are associated with growth, differentiation and cell death during embryonic development^1,2. The implantation of demineralized bone matrix or BMPs in skeletal muscle induces heterotopic bone formation in vertebrates³. This osteogenic activity is found in some BMPs and GDFs but not in TGF-βs, activin or other growth factors^4,5. The in vitro treatment of myoblasts with osteogenic BMPs inhibits myogenic maturation and induces the differentiation of myoblasts into cells expressing typical markers of mature osteoblasts, including alkaline phosphatase (ALP), parathyroid hormone receptor and osteocalcin⁶. In contrast, non-osteogenic members of the TGF-β family do not induce these phenotypic osteoblast markers in myoblast cultures in vitro, although non-osteogenic family members inhibit myogenesis⁶. Thus, osteogenic BMPs activate specific intracellular signaling pathways in target cells.

The intracellular signaling of TGF-β family members, including BMPs, is transduced by two types of transmembrane serine/threonine receptor kinases^7,8. Type II receptors phosphorylate the intracellular domain of type I receptors to activate receptor kinases, thereby activating downstream signaling pathways^7,8. Substitution mutations at the phosphorylation sites of type I receptors constitutively activate receptor kinases⁹. The overexpression of a constitutively activated form of type I BMP receptors (such as ALK2, BMPR-IA and BMPR-IB) induces the osteoblastic differentiation of myoblasts in cell culture in the absence of BMPs^10,11. Fibrodysplasia ossificans progressiva (FOP) is a rare genetic disorder that is characterized by heterotopic bone formation in skeletal muscle similar to the effects of BMP implantation^12,13. Our group and others have demonstrated that the ALK2 mutations in patients with FOP are natural gain-of-function mutants of type I BMP receptors^14,15,16,17. A kinase inhibitor of BMP type I receptors, LDN-193189, suppresses artificial bone formation in soft tissues in mice expressing constitutively activated ALK2¹⁸. These findings suggest that the substrates and downstream effectors of BMP type I receptors are critical for the bone-inducing activity of BMPs.

Smad proteins (Smads 1 through 9) are transcriptional regulators that are important for intracellular TGF-β family signaling^19,20,21. Smads are classified into 3 subgroups (R-Smad, Co-Smad and I-Smad) based on their structures and biochemical functions^20,22,23,24. Smads have conserved N-terminal MH1 and C-terminal MH2 domains that are connected by a linker region²⁵. Type I receptors phosphorylate R-Smads at the C-terminal serine-valine-serine (SVS) or serine-methionine-serine (SMS) motifs of the MH2 domains²⁶. Osteogenic BMPs induce the phosphorylation of Smad1, Smad5 and Smad9 (also known as Smad8), whereas, non-osteogenic members of the TGF-β family induce the phosphorylation of Smad2 and Smad3^20,27. I-Smads, Smad6 and Smad7, bind to type I receptors and inhibit receptor kinase activity^28,29,30,31. The expression of I-Smad mRNAs is increased in response to ligand stimulation, indicating that these mRNAs form negative feedback loops in the intracellular signaling among the TGF-β family³². Smad6 has been shown to interact with co-repressors, such as Hoxc-8 and CtBP, to inhibit BMP induced transcription activity^33,34. Phosphorylated R-Smads form complexes with Smad4, a Co-Smad and bind to the responsive element to regulate the transcription of target genes²⁰. The Id1, Id2 and Id3 genes are typical early-response genes in BMP signaling and have a conserved GGCGCC sequence in their 5′ upstream BMP-responsive elements (BREs) that is required for Smad binding^{35,36,37,38,39,40}. We identified a novel BMP-responsive non-coding RNA called BMP-inducible transcript-1 (Bit-1) by utilizing the conserved BMP-responsive elements among the Id genes, suggesting that the GGCGCC sequence is important for the BMP-induced transcription⁴⁰.

The specific role of each R-Smad on BMP activity remains unclear because the activation of a type I receptor simultaneously activates all the Smad and/or non-Smad signaling pathways^24,41. We reported a constitutively active mutant of Smad1, Smad1^DVD, in which the C-terminal SVS motif is mutated to an aspartic acid-valine-aspartic acid (DVD) sequence⁴². Smad1^DVD is recognized by an antibody that is specific for phosphorylated Smad1, Smad5 and Smad9 and activates BMP-induced biological events in vivo and in vitro in the absence of BMPs⁴². This constitutively active mutation allowed us to examine the specific role of Smad1 on BMP activity in the absence of the activation of the other signaling pathways. We used this strategy to examine the influence of Smad5 and Smad9 on BMP activity. We found that Smad9 was less active than Smad1 and Smad5 and that this lower activity depended on a deletion and a unique sequence in the linker region of Smad9. Moreover, Smad9 inhibited BMP signaling in cell culture similar to I-Smads. However, in contrast to I-Smads, Smad9 did not inhibit the phosphorylation of R-Smads by type I receptors. Smad9 formed complexes with BMP-regulated R-Smads and bound to the BRE but suppressed the transcriptional activity in a dominant negative manner. Thus, the inhibition of transcription is a previously uncharacterized role for Smad9 in BMP signaling.

Results

Construction and characterization of the constitutively active forms of R-Smads

We reported that the overexpression of Smad1^DVD under the control of the cytomegalovirus (CMV) promoter in the pFLAG-CMV2 vector induces the ligand-independent activation of BMP signaling, such as the induction of alkaline phosphatase (ALP) activity in C2C12 mouse myoblast cells⁴². The abundance of FLAG-Smad1^DVD and ALP activity were greater in the C2C12 cells that were transfected with Smad1^DVD in the pcDEF3 vector, which contains an elongation factor-1 promoter, compared to those transfected with pFLAG-CMV2 (Supplemental Fig. 1A and 1B). This increased signal-to-noise ratio enabled the identification of potentially weak activation. Therefore, we used the pcDEF3 vector.

We examined the effects of inhibitors on the BMP activity that is induced by a constitutively active BMPR-IA receptor or Smad1^DVD in C2C12 cells. In contrast to the constitutively active BMPR-IA, the activity of Smad1^DVD was almost unchanged, even in the presence of receptor inhibitors, including LDN-193189 (Supplemental Fig. 1C) and Smad6 (Supplemental Fig. 1D). However, the overexpression of a major component of NF-κB, p65, which inhibits the DNA binding of the Smad complex to a BRE⁴³, suppressed the activity of the constitutively active BMPR-IA and Smad1^DVD (Supplemental Fig. 1E). These data confirm our previous finding that Smad1^DVD acts as a constitutively active Smad1 in BMP signaling⁴².

To determine the role of each R-Smad in BMP signaling, we introduced serine-to-aspartate mutations in the residues that were homologous to the C-terminal SVS motif in Smad1 in other R-Smads, including Smad5, Smad9, Smad2 and Smad3. The overexpression of Smad1^DVD, Smad5^DVD and Smad9^DVD, but not Smad2^DMD or Smad3^DVD, in C2C12 cells increased the activity of two independent BMP-specific luciferase reporters, Id1WT4F-luc (Fig. 1A) and BIT1WT4F-luc (Fig. 1B), relative to that of the cells that were transfected with the empty vector or the corresponding wild-type Smad. The overexpression of the DVD forms of Smad1, Smad5 and Smad9 induced a typical marker of osteoblastic differentiation of C2C12 cells, ALP activity (Fig. 1C). These activities were lower with Smad9^DVD than with Smad1^DVD or Smad5^DVD, even under similar abundance conditions (Supplemental Fig. 2). As we previously reported for Smad1^DVD⁴², a specific antibody against phosphorylated Smad1, Smad5 and Smad9 recognized Smad1^DVD, Smad5^DVD and Smad9^DVD (Fig. 1D), suggesting that the C-terminal structures of these proteins mimic the conformational changes in the Smad proteins that were activated through phosphorylation.

Transcriptional activity of Smad9 is weaker than that of Smad1 and Smad5 despite having a greater DNA-binding activity

Because Smad9 showed lower activities than Smad1 or Smad5, we examined the biochemical characteristics of Smad9. The abundance of Smad9 was lower than that of Smad1 (Fig. 2A, bottom panel). However, more endogenous Smad4 was co-immunoprecipitated with FLAG-Smad9 than with FLAG-Smad1 from lysates of human embryonic kidney (HEK) 293 cells that were exposed to BMP-4 (Fig. 2A and 2B). FLAG-Smad9^DVD coprecipitated more endogenous Smad4 than did FLAG-Smad1^DVD and FLAG-Smad5^DVD (Fig. 2C). A specific antibody against Smad4 coprecipitated more endogenous Smad9 than endogenous Smad1 and Smad5 (Fig. 2E). The co-expression of Myc-Smad4 increased the transcriptional activity of Smad1^DVD, Smad5^DVD and Smad9^DVD, but that of Smad9^DVD was significantly lower (Fig. 2D). We further examined the DNA-binding capacity of Smad9 in a DNA pull-down assay using a BRE in the Id1 gene. Although the abundance of FLAG-Smad9 was less than that of FLAG-Smad1, more Smad9 than Smad1 bound to the BRE (Fig. 2F). Endogenous Smad9 also bound to the BRE in response to BMP-4 stimulation (Fig. 2G). The binding of both endogenous and FLAG-tagged Smad9 and Smad1 to the BRE was sequence-specific, as these proteins did not bind to the mutant sequence in which the Smad-binding GGCGCC core was substituted with TTTGCC (Fig. 2F and 2G).

The linker region of Smad9 suppresses the transcriptional activity of Smads

We constructed a series of chimeric Smad proteins in which the MH1 domain, linker region and MH2 domain of one Smad protein was replaced with those of another Smad protein. The overexpression of the chimeric Smad proteins in which the MH1 (Smad9-1-1^DVD) or MH2 (Smad1-1-9^DVD) domains in Smad1 were replaced with those from Smad9 resulted in an equivalent or increased activation of the Id1WT4F-luc reporter compared to that of Smad1^DVD (Fig. 3A and 3B), suggesting that the MH1 domain of Smad9 has a higher DNA-binding affinity than that of Smad1. In contrast, Smad1-9-1^DVD, which had the Smad9 linker region in Smad1^DVD, exhibited a significantly reduced ability to activate the reporter, similar to Smad9^DVD (Fig. 3A and 3B). We generated similar results with the overexpression of the chimeric Smads that were created using the domains of Smad5^DVD and Smad9^DVD (Fig. 3C and 3D), suggesting that the linker region of Smad9 suppressed its ability to activate BMP signaling.

Although Smad9, Smad1 and Smad5 share greater than 91% amino acid sequence similarity in the MH1 and MH2 domains, the linker regions share less than 40% similarity, including a deletion at the C-terminal portion of the linker in Smad9 compared to Smad1 and Smad5 (Supplemental Fig. 3). Thus, we inserted the sequence from this region of Smad1 [Smad9^DVD + S1L] or Smad5 [Smad9^DVD + S5L] into Smad9^DVD and found that the overexpression of either construct induced ALP activity (Fig. 3E and 3F) and activated the BMP reporter (Supplemental Fig. 4) to a similar degree as Smad1^DVD and Smad5^DVD. The PPAY motif in the linker region of Smad1 is recognized by E3 ubiquitin ligases and this motif is not present in Smad9⁴⁴. However, Smad9^DVD + S1L(AAAY), in which the PPAY motif has been substituted with the sequence AAAY, exhibited similar activity to Smad9^DVD + S1L (Fig. 3E and 3F), suggesting that the PPAY motif is not involved in Smad9 activity.

Smad9 mRNA is increased by BMP-4 stimulation similar to Smad6 mRNA

Transcription-dependent feedback loops are often involved in the regulation of signal transduction pathways, including BMP signaling³². We found that the expression of Smad9 increased in C2C12 myoblasts, C3H10T1/2 fibroblasts, MC3T3-E1 osteoblastic cells, 3T3-L1 adipocytes and primary calvarial osteoblasts that were exposed to BMP-4 (Fig. 4A). The expression of Smad9 mRNA, similar to Smad6 but not Smad1, Smad5, or Smad4, increased within one hour of BMP-4 stimulation in C2C12 cells (Fig. 4B). This increase was blocked by adding LDN-193189, a kinase inhibitor of BMP receptors, to the cell culture medium (Fig. 4C). In contrast, Smad7 but not Smad9 increased in the cells that were exposed to TGF-β1 (Fig. 4D), suggesting that Smad9 is induced by BMP-specific intracellular signaling. A BMP-induced increase in Smad9 protein levels was observed in C3H10T1/2 cells 24 h after stimulation (Fig. 4E). Furthermore, the increase in Smad9 mRNA levels was not blocked by pretreatment with cycloheximide, suggesting that Smad9 is an early response gene in the BMP-Smad axis (Supplemental Fig. 5). In addition, the abundance of Smad9 mRNA sharply decreased between E7.0 and E11.0 of developing mouse embryos, similar to Smad6 and Smad7 but not Smad1, Smad5 or Smad4 (Fig. 4F).

Smad9 inhibits BMP signaling

Our data suggest that Smad9 may act as an I-Smad similar to Smad6 and Smad7. Therefore, we tested whether the overexpression of wild-type Smad9 could inhibit the upstream activation of BMP signaling. As expected, the overexpression of Smad6 or Smad7 inhibited the BMP-specific Id1WT4F-luc activity that was induced by BMP-4 stimulation (Fig. 5A) or by the overexpression of a constitutively active form of BMPR-IA (Fig. 5B). Similarly, the overexpression of Smad9 suppressed the BMP activity that was induced by BMP-4 (Fig. 5A) or the constitutively active BMPR-IA (Fig. 5B). Overexpression of Smad9 suppressed the expression of endogenous Osterix mRNA induced by the constitutively active BMPR-IA and these results were similar to those observed with Smad6 (Supplemental Fig. 6).

We examined the effect of siRNA knockdown of endogenous Smad9 on BMP activity. The transfection of two independent siRNAs against Smad9 in C2C12 cells that were exposed to BMP-4 decreased Smad9 mRNA (Fig. 5C) and increased ALP activity (Fig. 5D).

Smad9 functioned as an inhibitor of the BMP-specific Id1WT4F-luc activity similar to Smad6 and Smad7. In contrast to Smad6 and Smad7, however, Smad9 was phosphorylated and did not inhibit the phosphorylation of Smad1 that was induced by the constitutively active BMPR-IA receptor (Fig. 5E). These results suggest that Smad9 suppressed BMP activity via a novel molecular mechanism.

Smad9 acts as a dominant negative Smad for Smad1

We examined the effects of Smad9 on the BMP activity that was induced by Smad1^DVD. The overexpression of either R-Smads (Smad1 and Smad5) or I-Smads (Smad6 and Smad7) inhibited the BMP-specific Id1WT4F-luc reporter activity that was induced by Smad1^DVD (Fig. 6A). However, Smad9 significantly suppressed the reporter activity in the presence of Smad1^DVD (Fig. 6A), suggesting that Smad9 functions downstream of the phosphorylation of R-Smads to inhibit BMP signaling.

To examine the molecular mechanisms of suppression by Smad9, we investigated whether Smad9 inhibited the ability of Smad1 to bind to a BRE in a DNA pull-down assay. More Smad9 bound to the BRE, even though the amount of Smad9 protein was lower than Smad1 (Fig. 6B). The amount of bound Smad9 was not decreased in Smad4-knockdown HEK293T cells, suggesting that the DNA-binding activity of Smad9 does not require Smad4 (Supplemental Fig. 7). The amounts of bound Smad9 and Smad1 were unchanged in the cells expressing both Smad9 and Smad1 simultaneously (Fig. 6B), suggesting that these proteins did not inhibit the DNA binding of each other.

We assessed whether Smad9 formed a complex with Smad1. We found that FLAG-Smad9 co-immunoprecipitated with Myc-Smad1 to a greater degree than FLAG-Smad1, even in Smad4-knockdown HEK293 cells (Fig. 6C). Endogenous Smad9 formed complexes with endogenous Smad1 or FLAG-Smad1 in BMP-4-stimulated cells (Fig. 6D). We examined the DNA-binding capacity of the Smad9-Smad1 complex. We co-immunoprecipitated FLAG-Smad9 and Myc-Smad1 from cell lysates using beads that were crosslinked with an antibody against the FLAG tag, eluted the complexes with an excess amount of FLAG peptide and then applied the eluents to a DNA pull-down assay using the BRE. Myc-Smad1 co-precipitated with the BRE in the presence of FLAG-Smad9 in HEK293 cells that were exposed to BMP-4 and decreased in the cells that were exposed to both BMP-4 and LDN-193189 (Fig. 6E), suggesting that the Smad9-Smad1 complex was formed and bound to the BRE in response to BMP signaling.

Discussion

In this study, we found that Smad9 is distinct from other Smads. The constitutively active form of Smad9 exhibited less activity than the BMP-regulated R-Smads, namely Smad1 and Smad5, even though more Smad9 associated with Smad4 and bound to the BRE. Moreover, we found several common characteristics among Smad9 and I-Smads, especially Smad6; 1) their transcription increased in response to BMP stimulation; 2) their mRNA levels sharply decreased during early embryonic development; 3) and they inhibited the BMP activity that was induced via type I receptors. These findings suggest that Smad9 shares biochemical characteristics with both BMP-regulated R-Smads and I-Smads. However, Smad9 and I-Smads exhibited distinct molecular mechanisms in the inhibition of BMP activity. I-Smads inhibit BMP activity by blocking the phosphorylation of R-Smads via direct interaction with type I receptors^28,29,30,31. Smad9 inhibited BMP activity without inhibiting the phosphorylation of R-Smads by BMP type I receptors. We found that Smad9 suppressed the transcription of a target gene via a novel mechanism. Because Smad9 itself exhibited less transcriptional activity, even though it bound to the BRE, the hetero-oligomeric complexes that formed with Smad1 or Smad5 would reduce the transcriptional activity in a dominant negative manner. Because Smad9 mRNA increased within 1 h of BMP stimulation, this novel type of inhibition may act as part of the negative feedback loop to regulate the fine-tuning of BMP signaling (Fig. 7). Taken together, our findings suggest that Smad9 is a new type of transcriptional repressor of BMP signaling that shares several biochemical characteristics with both BMP-regulated R-Smads and I-Smads but still has its own unique functions (Fig. 7). Further studies are required to elucidate the physiological role of Smad9 in BMP signaling.

Establishing the constitutively activated forms of Smads allowed us to examine the molecular mechanism of the low activity of Smad9. By constructing a series of chimeric Smad proteins, we found that the lower transcriptional activity of Smad9 was due to the shorter linker region compared to the regions of Smad1 and Smad5. The linker region of Smad9 inhibited the transcriptional activity of Smad1 and Smad5. The insertion of the fragment from the C-terminal linker region of Smad1 or Smad5 restored the transcriptional activity of Smad9. To examine the function of this fragment in the linker region, we deleted the short region from the Smad1 linker and analyzed its activity. Unexpectedly, the Smad1 containing the deletion still showed transcriptional activity in the BMP-specific reporter assay using Id1WT4F-luc (Supplemental Fig. 8), suggesting that the remaining portion of the linker region of Smad9 negatively affected BMP signaling. The linker region of Smad6 interacts with CtBP, a transcriptional co-repressor³⁴. Indeed, the linker region of Smad9 shares less than 40% homology with that of Smad1 and Smad5. Smad9, but not Smad1, interacts with ANA, which is a co-repressor in the TOB/BTG family that decreases the bone-inducing activity of BMP-2 in mice⁴⁵. Because more Smad9 interacted with Smad4, Smad1 and the BRE, the short linker likely allows Smad9 to interact with other molecules, including co-activators, co-repressors and target DNAs. The insertion of a fragment to the C-terminal region of the Smad9 linker may disrupt these interactions. Although mouse Smad9 is only a short form, the human genome has a short form and a long form of Smad9, Smad9b and Smad9a, respectively⁴⁶. Their linker regions share less than 51% homology with that of human Smad1 (Supplemental Fig. 9). Human Smad9a^DVD and Smad9b^DVD had significantly lower transcriptional activity of Id1WT4F-luc (Supplemental Fig. 10), supporting our hypothesis that the Smad9 linker region negatively affects BMP signaling. Additional studies are needed to elucidate the molecular mechanisms of the linker-dependent regulation of the transcriptional activities of Smad1, Smad5 and Smad9.

In conclusion, we found that although Smad9 shares common structures with positive effectors of BMP signaling, namely Smad1 and Smad5, it represses intracellular BMP signaling as a dominant negative Smad. Thus, Smad9 is a new type of transcriptional regulator in BMP signaling.

Experimental Procedures

Plasmids

A constitutively active form of Smad1, Smad1^DVD, in pFLAG-CMV2 has been previously reported⁴². The wild-type and the DVD or DMD (only Smad2) forms of the mouse Smad1, Smad2, Smad3, Smad5, Smad9, Smad4, Smad6, Smad7 (Smad4, Smad6 and Smad7 were wild-type only) and human Smad1, Smad9a (long form), Smad9b (short form) cDNAs except for the start codon (ATG), were amplified from a cDNA pool that was prepared from C2C12 cells and qPCR Human Reference Total RNA (Clontech, Palo Alto, CA), respectively, by standard RT-PCR using reverse primers containing wild-type, DMD or the DVD-mutated sequences (Supplemental Table 1) and inserted into pcDEF3 downstream of the FLAG-tag or Myc-tag sequence⁴⁷. Chimeric Smads with the structures that are described in the text and figures were constructed by ligating DNA fragments encoding each domain of the Smad in-frame into pcDEF3. Each Smad was divided into an MH1 domain, linker region and MH2 domain as follows (the numbers indicate the amino acid positions): Smad1, 2–132, 133–264 and 265–465; Smad5, 2–133, 134–264 and 265–465; and Smad9, 2–136, 137–227 and 228–428, respectively. The linker regions of Smad1(218–264) and Smad5(217–264) were inserted downstream of the linker of Smad9^DVD to obtain Smad9^DVD + S1L and Smad9^DVD + S5L, respectively. The linker region of human Smad1(218–264) was deleted in S1ΔLC^DVD. The final constructs were confirmed by DNA sequencing. The expression plasmids for p65 and constitutively active BMPR-IA were previously described^16,43. For the BMP-specific luciferase assay, Id1WT4F-luc³⁵ or BIT1WT4F-luc⁴⁰ was co-transfected with phRL-SV40 (Promega, Madison, WI).

Cell culture, transfection, luciferase assay and ALP assay

Mouse C2C12 myoblasts were cultured as previously described⁶. The cells were inoculated at 1.5 × 10⁴ cells/cm² in 96-well plates 1 day before transfection. After overnight culture, the cells were transfected with 200 ng of plasmid DNA using 0.5 μl of Lipofectamine 2000 reagent (Invitrogen, Carlsbad, CA) following the manufacturer's instructions and the culture medium was changed after 2.5 h³⁵. The luciferase assay was performed on day 1 using the Dual-Glo Luciferase Assay System (Promega) and the data are expressed as the luciferase activity of firefly relative to that of Renilla^15,35. The ALP activity was determined on day 3 using p-nitrophenylphosphate as the substrate as previously described⁴⁸. For the siRNA-mediated knockdown experiments, a scrambled control (si-Scr) (Invitrogen), two independent Smad9-specific (si-Smad9) siRNAs (#1: GGGCUUCUCUC AAAUGUAAACAGAA and #2: GGCACAUUGGAAAGGGUGUGGAUUU) (Invitrogen) or Smad4-specific (si-Smad4) siRNA⁴² were transfected using the Lipofectamine RNAi Max reagent (Invitrogen) according to the manufacturer's instructions.

Co-immunoprecipitation, DNA pull-down and Western blot analysis

Western blot analysis was performed as previously described^16,42. In brief, whole cell extracts were prepared using lysis buffer (10 mM Tris-HCl, pH 7.8, 1% Triton X-100 and protease inhibitor cocktail; Nacalai Tesque, Kyoto, Japan), separated using sodium dodecyl sulfate-containing polyacrylamide gel electrophoresis and transferred to polyvinylidene difluoride membranes.

For co-immunoprecipitation assays, the cell extracts were incubated with magnet beads that were conjugated to an antibody against the FLAG tag (Sigma-Aldrich, St. Louis, MO) for 12 h at 4°C. The FLAG-epitope-containing complex was precipitated using a magnet and eluted by adding FLAG peptide (Sigma-Aldrich).

For the DNA pull-down assays, whole cell extracts were incubated for 30 min on ice with biotinylated DNA of wild-type or mutant BMP-responsive element (BRE), which was prepared from Id1WT4F-luc or Id1MutB4F-luc³⁵, respectively, by standard PCR using a set of biotinylated primers⁴⁹. The DNA-protein complex was precipitated using streptavidin-conjugated magnet beads (Dynabeads M-280 streptavidin; Invitrogen) and applied to Western blot analysis as described above.

The following antibodies were used in these experiments: mouse monoclonal antibody against the FLAG tag (clone M2, Sigma-Aldrich), mouse monoclonal antibody against Smad4 (Cell Signaling Technology, Beverly, MA), mouse monoclonal antibody against the V5-tag (clone V5005; Nacalai Tesque), rabbit polyclonal antibody against the Myc-tag (Cell Signaling Technology), rabbit polyclonal antibody against phosphorylated Smad1, Smad5 and Smad9 (Cell Signaling Technology) and rabbit polyclonal antibody against α-tubulin (Cell Signaling Technology) and rabbit normal IgG (Cell Signaling Technology). The target proteins were detected using a horseradish peroxidase-conjugated antibody against mouse or rabbit IgG (Jackson ImmunoResearch Laboratories, West Grove, PA). Chemiluminescence was detected by a ChemiDoc XRS+ System (Bio-Rad, Hercules, CA).

Rabbit antiserum against mouse Smad9 was produced by regularly injecting a rabbit with a peptide corresponding to the mouse Smad9 linker region (a.a. 153–167).

Reverse transcription-polymerase chain reaction and quantitative RT-PCR

Total RNA was prepared using a NucleoSpin RNA kit (Macherey-Nagel, Düren, Germany) and reverse-transcribed with Superscript III (Invitrogen) according to the manufacturer's instructions. The PCR assays were performed with GoTaq DNA polymerase (Promega) with the listed primer sets (Supplemental Table 1). Quantitative RT-PCR (qRT-PCR) was performed using SYBR Premix Ex Taq (TaKaRa, Shiga, Japan) with a Thermal Cycler Dice TP800 (TaKaRa). The total RNA of mouse embryos was purchased from Clontech Laboratories (Mountain View, CA). The primers that were used are listed in Supplemental Table 1. The data are expressed as the relative expression levels compared to those of Atp5f1 as a control⁴⁹.

Statistical analysis

Comparisons were made using an unpaired ANOVA with Tukey-Kramer post-hoc test and Wilcoxon's signed rank test. The results are shown as the mean ± S.D. The statistical significance is indicated as follows: *, p < 0.05 and **, p < 0.01.

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Acknowledgements

We thank Seiya Suzuki, Mitsue Fujiwara and Ayaka Mizuguchi for their excellent technical assistance and the members of the Division of Pathophysiology for valuable discussion. We are grateful to Dr. J. A. Langer for kindly providing pcDEF3 and Dr. Michio Shiibashi for statistical analysis. This work was supported in part by the following sources: a Grant-in-Aid from the Ministry of Education, Culture, Sports, Science and Technology of Japan (to ST, 25893228; KY, 24592278; TK, 25293326 and 25670658; EJ: 23390424); a Grant-in-Aid from the Support Project for the Formation of a Strategic Center in a Private University from the Ministry of Education, Culture, Sports, Science and Technology of Japan (TK); and the Health and Labor Sciences Research Grants for Research on Measures for Intractable Diseases from the Ministry of Health, Labor and Welfare of Japan (TK).

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Division of Pathophysiology, Research Center for Genomic Medicine, Saitama Medical University, 1397-1 Yamane, Hidaka-shi, Saitama, 350-1241, Japan
S. Tsukamoto, T. Mizuta, M. Fujimoto, S. Ohte, K. Osawa, A. Miyamoto, K. Yoneyama, A. Machiya, S. Kokabu & T. Katagiri
School of Medical Technology and Health, Faculty of Health and Medical Care, Saitama Medical University, 1397-1 Yamane, Hidaka-shi, Saitama, 350-1241, Japan
E. Murata
Division of Molecular Signaling and Biochemistry, Department of Health Promotion, Kyushu Dental University, 2-6-1 Manazuru, Kokurakita-ku, Kitakyushu-shi, Fukuoka, 803-8580, Japan
E. Jimi
Department of Oral and Maxillofacial Surgery, Faculty of Medicine, Saitama Medical University, 38 Morohongo, Moroyama-machi, Iruma-gun, Saitama, 350-0495, Japan
S. Kokabu

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S. Tsukamoto
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T. Mizuta
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M. Fujimoto
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S. Ohte
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K. Osawa
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A. Miyamoto
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E. Murata
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A. Machiya
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E. Jimi
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Contributions

S.T. and T.K. designed the experiments. S.O., K.O., E.M., E.J., S.K. and T.K. conducted the experiments. S.T., T.M., M.F., S.O., K.O., A.M., K.Y. and T.K. performed the experiments and analyzed the data. S.T. and T.K. wrote the paper.

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dataset 1

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Tsukamoto, S., Mizuta, T., Fujimoto, M. et al. Smad9 is a new type of transcriptional regulator in bone morphogenetic protein signaling. Sci Rep 4, 7596 (2014). https://doi.org/10.1038/srep07596

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Received: 14 September 2014
Accepted: 03 December 2014
Published: 23 December 2014
DOI: https://doi.org/10.1038/srep07596

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