Requirement for the SnoN Oncoprotein in Transforming Growth Factor β-Induced Oncogenic Transformation of Fibroblast Cells

Cell culture, antisera, and constructs.

Mouse NIH 3T3 fibroblasts, NRK fibroblasts, and the rat intestinal epithelial cell line RIE-1 (ATCC) were maintained in Dulbecco's modified Eagle's medium supplemented with 10% fetal bovine serum (FBS). Hep3B human hepatoma cells (ATCC) were maintained in minimal essential medium supplemented with 10% FBS. The mouse embryonic cell line AKR-2B (ATCC) was grown in McCoy's 5A medium supplemented with 10% FBS.

Antibodies for phospho-Smad2 and phospho-Smad3 were generous gifts from Aristidis Moustakas. Antisera against Smad3 (FL425 and I-20), Smad4 (C-20), Smad2 (S-20), SnoN (H317), and p300 (N15) were purchased from Santa Cruz Biotechnology. Antisera against Smad2 were purchased from BD, Transduction Laboratories. The antibody against hexamethylene bisacetamide-induced protein 1 (HEXIM1) was a gift from Qiang Zhou.

All reporter constructs containing deletions, truncations, or mutations in the snoN promoter were generated by PCR and cloned into the pGL₂ basic vector (Promega). All mutations were verified by sequencing.

Transfection and transcriptional reporter assays.

Hep3B cells were transiently transfected with the reporter constructs using LipofectAMINE Plus (Invitrogen) (16). For luciferase assays, a total of 2.5 μg of DNA (0.5 μg of the luciferase reporter construct and 2 μg of Smad proteins) was used for each transfection. Luciferase activity was measured 48 h later.

Northern blotting.

Hep3B, RIE-1, NIH 3T3, NRK, and AKR-2B cells were serum starved for 16 h and then treated with 200 pM TGF-β1 in the presence or absence of 100 μg/ml cycloheximide or 1 μg/ml actinomycin D for the indicated times. Total RNA was prepared from these cells using the RNeasy Total RNA kit (QIAGEN). Twenty micrograms of total RNA was resolved on a 1% formaldehyde gel, transferred to a nylon membrane, and analyzed by Northern blotting with a human or mouse snoN cDNA probe. All of the snoN bands in the Northern blots were quantified by Image J.

DNase I footprinting.

A DNase I footprinting reaction was performed as described in the protocol of the “Core Footprinting System” (Promega). One microgram of a fusion protein consisting of glutathione S-transferase (GST) and Smad3 or Smad4 was preincubated on ice for 10 min in binding buffer (50 mM Tris, pH 8.0, 100 mM KCl, 12.5 mM MgCl₂, 1 mM EDTA, 20% glycerol, and 1 mM dithiothreitol), following by a 20-min incubation on ice with a ³²P-labeled DNA probe (1 × 10⁶ cpm/μg). The samples were then treated with 5 μl DNase I (1 U/μl; Promega) for exactly 2 min at room temperature and analyzed on a 6% denaturing polyacrylamide gel. The relative positions of the protected bands on the gel were determined by a G reaction using the protocol of Maxam and Gilbert (28).

EMSA.

DNA fragments containing regions of the snoN promoter between kb −2.8 and −2.4 (−2.8/−2.4) or between kb −2.4 and −2.0 (−2.4/−2.0) were amplified by PCR, end labeled with T4 polynucleotide kinase, and gel purified. The ability of recombinant Smad3, Smad4, or nuclear extracts to bind to these DNA fragments was analyzed by electrophoretic mobility shift assays (EMSA) as described previously (15, 46).

"V体育官网入口" ChIP.

Hep3B cells were stimulated with 200 pM TGF-β1 and cross-linked with formaldehyde for 10 min, and chromatin immunoprecipitation (ChIP) was carried out as described previously (4) with the following modifications. The chromatin fractions from 2 × 10⁷ cells were precleared with protein A-Sepharose for 30 min at 4°C and then subjected to immunoprecipitation with 5 μg specific antibodies (anti-Smad3 [I-20], -Smad4 [C-20], -Smad2 [S-20], and -p300 [N15]) at 4°C. For PCR, 3 μl from a 30-μl immunoprecipitate or 3 μl of a 1:30 dilution from a 60-μl input sample was used as a template. The primer pairs used to amplify TGF-β-responsive regions of the snoN and PAI-1 promoters are as follows: for the snoN promoter, 5′-CTCGAGGAAGGAAGGAGGG-3′ (forward) and 5′-TACACACAGCCTCTGACGTC-3′ (reverse); for the PAI-1 promoter, 5′-CCAGCACACCTCCAACCTC-3′ (forward) and 5′-CCCTGCAGCCAAACACAGC-3′ (reverse).

RNA interference.

Small interfering RNAs (siRNAs) were designed to specifically target either murine snoN, murine smad3, or human smad3 and smad2 in accordance with the guidelines provided (OligoEngine). Oligonucleotides carrying the siRNAs are as follows (sense strands shown): murine snoN, 5′-CTCCATTCTGCAGAGGAAG-3′; human smad3-a, 5′-CGTCAACACCAAGTGCATC3′; human smad3-b, 5′-GGCCATCACCCAGAAC-3′; human smad2, 5′-ACAGGACGATTAGATGAGC-3′; negative-control sequence, 5′-CACCTGTACAGTTGCTAGT-3′. The oligonucleotide pairs were cloned into the pSUPER vector (OligoEngine).

Generation of AKR-2B cell lines stably expressing a siRNA.

Oligonucleotide pairs encoding the siRNA for murine snoN or an unrelated sequence in the pSUPER.retro.puro vector (OligoEngine) were transfected into Phoenix-Eco cells with LipofectAMINE Plus (Invitrogen). Forty-eight hours posttransfection, a total of 3 × 10⁵ AKR-2B cells were infected with the viral supernatant. Forty-eight hours later, the infected cells were switched into a medium containing 1.5 μg/ml puromycin to start selection. To confirm the expression of the siRNA, individual clones of infected cells were treated with 100 pM TGF-β1 for 1 h. Total RNA was extracted from these cells, and the levels of snoN mRNA were analyzed by Northern blotting.

Soft-agar assay.

A 2-ml volume of growth medium containing 0.66% Bacto agar (Difco) was poured into a 6-well dish to form the bottom layer. A total of 5,000 AKR-2B cells or 50,000 NRK cells were suspended in 2 ml of medium containing 0.4% agar and 2 nM (for AKR-2B cells) or 10 ng/ml (for NRK cells) epidermal growth factor (EGF), with or without 100 pM TGF-β1, and this suspension was overlaid on the hardened bottom layer. A 1-ml volume of fresh medium containing 0.4% agar and 2 nM or 10 ng/ml EGF, with or without 100 pM TGF-β1, was added to the dish every week. After 3 weeks of incubation, colonies were visualized by staining with 0.5 mg/ml 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyl-tetrazolium bromide (MTT) (Sigma) and scanned on a Hewlett-Packard ScanJet 5500C to visualize the colonies. For treatment with the TβRI inhibitor SB-431542, 1 ml of medium with or without 10 μM SB-431542 (Tocris) was added on top of the soft agar at the indicated time point post-TGF-β1 addition. The medium was replaced every 2 days after weekly soft-agar feeding.

V体育平台登录 - Immunoprecipitation and Western blotting.

Endogenous Smads were analyzed by Western blotting as described previously (26). The densities of bands in Western blots were quantified by Image J. For immunoprecipitation of endogenous SnoN, a polyclonal antibody to SnoN was raised against a peptide located at the N terminus of human SnoN (DDGSPPAKKMITDIH). The anti-peptide antibody was covalently coupled to protein A-Sepharose and incubated with nuclear extracts from AKR-2B cells stimulated with or without TGF-β1. SnoN was analyzed by Western blotting with anti-SnoN (Santa Cruz).

TGF-β induces prolonged snoN expression in two fibroblast cell lines that are transformed by TGF-β1.

We have shown previously that in Hep3B cells, TGF-β induces a marked increase in snoN gene expression within 2 hours of stimulation (47). To determine whether this is a general phenomenon that also occurs in other cell types, especially in AKR-2B and NRK fibroblasts that are transformed by TGF-β either alone or together with EGF (29, 37), a time course of TGF-β1 treatment was performed for several cell lines including two epithelial cell lines, RIE-1 (Fig. 1A) and HaCaT (data not shown), and three fibroblast cell lines, NIH 3T3, NRK, and AKR-2B (Fig. 1A). Like Hep3B cells, RIE-1 and HaCaT cells undergo cell cycle arrest in response to TGF-β. In contrast, the growth of the three fibroblast cell lines is stimulated by TGF-β. Total RNA was prepared from these cells, and the levels of snoN mRNA were measured by Northern blotting. As shown in Fig. 1A, induction of snoN mRNA by TGF-β was observed in all cell lines. The increase in snoN mRNA levels occurred within 1 h of TGF-β1 stimulation and peaked by 2 h. However, the duration and magnitude of the induction differ from cell line to cell line. The snoN mRNA level remained high for at least 8 to 24 h in the two TGF-β-transformed fibroblast lines, while it rapidly decreased within 2 to 4 h in NIH 3T3 cells and within 4 to 6 h in the epithelial cell lines (Fig. 1A and data not shown). Interestingly, the strongest and longest induction of snoN expression was observed in AKR-2B cells, which display a strong transforming phenotype in response to TGF-β (as evidenced by the formation of large numbers of fast-growing colonies on soft agar). In NRK cells, where TGF-β induces a moderate level of transformation, the duration of induction of snoN expression was correspondingly shorter.

To confirm that the upregulation of snoN mRNA resulted in an increase in SnoN protein, Western blot analysis was carried out using an antibody that recognizes mouse SnoN. In AKR-2B cells, TGF-β induced an increase in the SnoN protein level at 2 h after stimulation, and the SnoN level remained high for at least 8 h after stimulation (Fig. 1B). Taken together, our data indicate that induction of snoN expression by TGF-β occurs in all cell lines examined, but only those cell lines that are transformed by TGF-β appeared to exhibit a stronger and prolonged induction.

TGF-β activates snoN gene expression at the level of transcription.

To examine whether the induction of snoN expression, especially at later time points, in AKR-2B and NRK cells requires protein synthesis, cells were pretreated with the protein synthesis inhibitor cycloheximide. The induction of snoN gene expression by TGF-β in either AKR-2B (Fig. 1C) or Hep3B and NRK (data not shown) cells in the presence or absence of cycloheximide was then examined by Northern blotting. As shown in Fig. 1C, pretreatment with cycloheximide did not affect the levels of snoN mRNA at any time after TGF-β stimulation in AKR-2B cells or other cell lines (data not shown), indicating that this upregulation is due to direct activation of TGF-β signaling pathways.

We next examined snoN mRNA expression following TGF-β stimulation in the absence or presence of the transcription inhibitor actinomycin D. As shown in Fig. 1D, induction of snoN expression was effectively blocked by actinomycin D, suggesting that TGF-β activates snoN expression mainly at the level of transcription. To determine whether alteration in the stability of SnoN contributes to the prolonged induction of SnoN, AKR-2B cells were treated with TGF-β1 for 2 h to induce SnoN expression. Cycloheximide was then added, and the treatment (with both TGF-β1 and cycloheximide) was allowed to continue for another 6 h. As shown in Fig. 1E, cycloheximide completely blocked SnoN induction by TGF-β after 8 h, suggesting that induction of SnoN expression is entirely due to the increase in SnoN synthesis. Consistent with this, in a pulse-chase assay, the half-life of TGF-β-induced, newly synthesized SnoN was not altered after 6 additional hours of TGF-β treatment in AKR-2B cells (data not shown).

To map the TGF-β-responsive elements in the snoN promoter, a 4.8-kb murine snoN genomic DNA fragment containing sequences immediate upstream of the translation start site was cloned upstream of the luciferase reporter gene. When transfected into Hep3B or AKR-2B cells, this 4.8-kb fragment was sufficient to mediate TGF-β-induced transcriptional activation (Fig. 1F). Because Hep3B cells are easier to transfect than AKR-2B cells, we carried out the major part of the promoter-mapping experiments with Hep3B cells. The key experiments were later repeated with AKR-2B cells.

A series of 5′ deletion mutants of the snoN promoter was generated from the 4.8-kb snoN genomic fragment and tested for ability to mediate TGF-β responses (Fig. 1G). Deletions of sequences from kb −4.8 to −2.8 had no effect on TGF-β-induced transcriptional activation of the snoN promoter (Fig. 1G). However, further deletions to kb −2.4 completely abrogated TGF-β responsiveness, suggesting that the TGF-β-responsive element is located between kb −2.8 and −2.4 of the snoN promoter. In agreement with this, when cloned upstream of a heterologous promoter, this 400-bp fragment (sno2.8/2.4) was able to mediate TGF-β-induced activation of the reporter construct to an extent comparable to that exhibited by the natural 2.8-kb snoN promoter fragment (Fig. 1G).

TGF-β induces prolonged phosphorylation of Smad2 in AKR-2B and NRK cells.

Since Smad proteins are critical mediators of TGF-β signaling, we next examined whether they are involved in TGF-β-induced snoN expression. For this purpose, the inhibitory Smad, Smad7, was cotransfected with sno2.8 into Hep3B cells. Overexpression of Smad7 efficiently blocked TGF-β-inducible transcription from the snoN promoter (Fig. 2A), suggesting that this upregulation is mediated by a Smad-dependent pathway.

To examine whether the difference in the duration of snoN induction reflects a difference in the activities of the Smad proteins, we next measured the relative amounts of active Smad2 and Smad3 in various cell lines. AKR-2B, NRK, NIH 3T3, and RIE-1 cells were treated with TGF-β1 for various periods. The phosphorylation of Smad2 and Smad3 was detected by Western blotting with antibodies specific for phospho-Smad2 or phospho-Smad3. In all cell lines, TGF-β1 induced a rapid increase in phosphorylation of Smad2 and Smad3 (Fig. 2B). In NIH 3T3 and RIE-1 cells, levels of phospho-Smad2 reached a maximum at 1 h after stimulation and declined back to the basal level after 4 h (Fig. 2B), while levels of phospho-Smad2 remained high in AKR-2B and NRK cells for 8 to 24 h (Fig. 2B and C). In contrast, the duration of phospho-Smad3 was not significantly different in the four cell lines (Fig. 2B). This suggests that phosphorylation of Smad2 and phosphorylation of Smad3 are regulated separately in these cells and that a longer induction of snoN expression by TGF-β correlates with a prolonged phosphorylation and activation of Smad2.

To further strengthen the notion that the sustained activation of Smad2 is responsible for the prolonged induction of snoN expression by TGF-β in AKR-2B and NRK cells, a specific inhibitor of the TβRI kinase, SB-431542, was employed (7, 19, 21). AKR-2B (Fig. 2C) or NRK (data not shown) cells were stimulated with TGF-β for various periods. SB-431542 was added to the culture at 3 h or 6 h after TGF-β addition. The levels of phospho-Smad2 were then measured by Western blotting with anti-phospho-Smad2, and snoN expression was examined by Northern blotting. As shown in Fig. 2C, SB-431542 effectively shortened the duration of receptor-mediated phosphorylation of Smad2. Interestingly, the induction of snoN expression by TGF-β also showed a marked decrease upon addition of SB-431542, and the timing of the inhibition of snoN induction correlated with that of the inhibition of Smad2 phosphorylation (Fig. 2C and data not shown). This finding confirms that in AKR-2B and NRK-2B cells, the sustained Smad2 phosphorylation in response to TGF-β, instead of the stability of mRNA, is responsible for the prolonged induction of snoN expression. It also suggests that the sustained phosphorylation of Smad2 is likely due to the activity of TβRI kinase, not to a reduced phosphatase activity in these cells.

Binding of Smad complexes to the TGF-β-responsive elements is essential for TGF-β-induced activation of snoN transcription. (VSports最新版本)

Since induction of snoN expression correlated with activation of Smad2 (Fig. 2C), we next examined whether the Smad2/Smad4 complex binds directly to the 400-bp TGF-β-responsive region in the snoN promoter to activate snoN expression. To locate the binding sites of Smads within this TGF-β-responsive promoter region, DNase I footprinting was carried out using two 200-bp end-labeled DNA probes, containing sequences from kb −2.8 to −2.6 and from kb −2.6 to −2.4 of the snoN promoter, respectively. Recombinant Smad4 generated four protected regions within the 400-bp TGF-β-responsive region (Fig. 3A). All of them contain a CAGA or TCTG core sequence that was previously identified as the SBE (42). In an EMSA using the four Smad4-protected sequences as probes, Smad4 bound to all four SBEs (Fig. 3B). These binding activities were disrupted by point mutations that changed the core SBE sequences. Interestingly, although all four probes contain the same core SBE sequence, the affinities of Smads for the four regions differed, indicating that the flanking sequences of SBE also affect binding of the Smad proteins.

To determine the importance of these Smad binding elements in TGF-β-induced snoN expression, mutations altering the four SBEs either individually or together were introduced into the snoN promoter. In a luciferase assay (Fig. 3C), mutations of individual SBEs decreased TGF-β inducible transcription only slightly. In contrast, alteration of all SBEs markedly abrogated TGF-β-induced snoN transcription but did not affect the level of basal transcription. This indicates that all four Smad binding sites together mediate TGF-β-induced transcription of the snoN gene.

Finally, to confirm that this 400-bp TGF-β-responsive region in the snoN promoter mediates binding of the endogenous Smad proteins in vivo, ChIP assays were carried out. A pair of primers spanning the 400-bp TGF-β-responsive region was employed in the PCR to measure the endogenous proteins bound to this region of the snoN promoter. As an example of a typical TGF-β-responsive promoter, the plasminogen activator inhibitor type I (PAI-1) promoter was analyzed in parallel. Primers flanking the 800-bp fragment from the PAI-1 promoter that contains the TGF-β-responsive elements were included in the PCRs. In the absence of TGF-β, Smad4, as well as very low levels of R-Smads, was found to bind to the TGF-β-responsive regions of both snoN and PAI-1 promoters (Fig. 3D). Treatment of cells with TGF-β1 for 1 h resulted in an increase in the binding of Smad2 and the coactivator p300/CBP to the two promoter sequences. Surprisingly, association of Smad3 with the snoN promoter was not detected in the presence of TGF-β. In contrast, the binding of Smad3 to the PAI-1 promoter was dramatically increased by TGF-β treatment. Thus, Smad2 and Smad3 may play different roles in regulation of snoN transcription, and the effect of Smad3 on snoN gene expression may differ from that on the expression of typical TGF-β-responsive genes such as PAI-1.

Smad2 and Smad3 play opposite roles in regulation of snoN expression.

To investigate the effects of Smad2 and Smad3 on the activation of snoN expression, Smad2 or Smad3 was cotransfected with Smad4 and the sno2.8 reporter into Hep3B or AKR-2B cells. As a control, pLUC800, which contains the 800-bp TGF-β-responsive element from the PAI-1 promoter, was tested in a parallel experiment. As shown in Fig. 4A, expression of Smad2 and Smad4 caused a marked increase in transcription from both the snoN and PAI-1 promoters, even in the absence of TGF-β1. Surprisingly, overexpression of Smad3 and Smad4 inhibited activation of the snoN promoter induced by TGF-β in both cell lines (Fig. 4A). This inhibition appears to be specific to the snoN promoter, because expression of Smad3 greatly activated transcription from the PAI-1 promoter both in the presence and in the absence of TGF-β. Thus, Smad2 enhances snoN gene expression, while Smad3 blocks it.

To further confirm this, we employed the siRNA duplex specific for either human Smad2 or human/murine Smad3 and introduced it into Hep3B or AKR-2B cells using the pSUPER vector (6). In transfected Hep3B or AKR-2B cells, the siRNA duplexes specifically reduced the expression of Smad proteins (Fig. 4B, bottom panels). When introduced together with sno2.8, the siRNA for Smad2 decreased snoN transcription in response to TGF-β, while expression of Smad3 siRNAs resulted in a marked increase in snoN transcription (Fig. 4B). As a control, expression of an unrelated siRNA did not affect the expression of Smad proteins or TGF-β-induced snoN transcription (Fig. 4B). Taken together, these results indicate that Smad2 and Smad3 exert different effects on snoN gene expression.

The DNA binding activity and oligomerization of Smad3 are required for inhibition of TGF-β-induced snoN transcription by Smad3.

In order to understand how Smad3 negatively regulates snoN gene expression, we next examined whether the DNA binding and oligomerization activities of Smad3 are required for this inhibition. A mutant Smad3 (R74A) deficient in binding to DNA (42) was cotransfected with Smad4 and sno2.8 into Hep3B cells. While wild-type (WT) Smad3 effectively blocked activation of snoN transcription by TGF-β, Smad3R74A failed to do so (Fig. 4C). Not only was this mutant Smad3 incapable of inhibiting snoN expression; it actually activated transcription both in the absence and in the presence of TGF-β, as did Smad2 (Fig. 4C). This result suggests that the DNA binding ability of Smad3 is critical to its inhibitory activity.

Recently, an alternatively spliced form of Smad2 that lacks exon 3, Smad2(Δexon3), has been shown to mediate most of TGF-β signaling events during embryonic development (11, 55). Smad2(Δexon3) contains the DNA binding activity and has been speculated to function in manner similar to Smad3 (55). We therefore tested the role of Smad2(Δexon3) in the regulation of snoN induction by TGF-β. When Smad2(Δexon3) was cotransfected with Smad4 and sno2.8 into either AkR-2B or Hep3B cells, it transactivated snoN promoter as efficiently as full-length Smad2 (Fig. 4C). This indicates that, at least in this case, Smad2(Δexon3) functions like full-length Smad2 [Smad2(FL)], not Smad3, and that the DNA binding ability of Smad is necessary but not sufficient for the inhibitory function. In addition, the expression levels of Smad2(Δexon3) in all cell lines described in this study are much lower than those of Smad2(FL) (data not shown). This suggests that Smad2(FL) plays a major role in regulation of snoN expression in these cell lines.

Oligomerization of Smad3 with Smad4 has been shown to be essential for the transcription activity (40). To examine whether this ability is required for the inhibition of snoN expression by Smad3, mutant Smad3 (S3V277D) or Smad4 (S4V370D) defective in oligomerization was examined in the luciferase reporter assay (Fig. 4C). When introduced together with WT Smad4 or Smad3, both mutants failed to inhibit snoN expression (Fig. 4C). Rather, they activated transcription from the snoN promoter as efficiently as Smad2. This suggests that hetero-oligomerization of Smad3 with Smad4 is essential for its inhibitory activity. Interestingly, some of the mutations of Smad4 found in human cancer cells disrupt the ability of Smad4 to heterodimerize with Smad3 (40). Our results suggest that in these cancer cells (39, 50), Smad3 activates, instead of inhibiting, snoN expression. This may account for the elevated level of snoN expression in these cancer cells.

A Smad-inhibitory element (SIE) mediates inhibition by Smad3.

Internal deletions within the 2.8-kb snoN promoter fragment were generated and used to determine the region in the snoN promoter that mediates the inhibitory function of Smad3. As shown in Fig. 5A, deletion of sequences up to kb −2.0 had no effect on the inhibitory response to Smad3. In contrast, further deletion to kb −2.4 or an internal deletion of sequences between kb −2.0 and −2.4 abolished the suppressive effects of Smad3/Smad4. These two deletion mutants also exhibited reduced basal transcription but still retained the ability to respond to TGF-β. This suggests that the region between kb −2.4 and −2.0 is critical for both basal transcriptional activity and Smad3-dependent transcriptional inhibition.

Since the DNA binding activity of Smad3 was important for its inhibition of snoN transcription, we speculated that Smad3 may bind directly to a specific inhibitory element in this region of the snoN promoter. A DNase I footprinting assay was therefore carried out to determine the Smad3 binding site using the promoter fragment from kb −2.4 to −2.0 promoter fragment as a probe. A protected region with the sequence 5′-CGGCGACGGCGACGGCGACGGCG-3′ was identified at bp −2179 to −2201 (Fig. 5B). This sequence is a triple-repeat of the CGACGG box that displays no resemblance to the typical SBE motif. In an EMSA, purified recombinant Smad3 and Smad4 both bound to the WT but not the mutant triple CGACGG sequence (Fig. 5C), confirming that this sequence can indeed mediate Smad binding.

To determine whether this novel Smad binding element can mediate inhibition of snoN expression by Smad3/Smad4, the triple CGACGG box was mutated in sno2.8. When this mutant was cotransfected with Smad3 and Smad4 into Hep3B cells, transcription from the mutant snoN promoter (SIEm) was not repressed by the Smad3/Smad4 complex (Fig. 5D). These results demonstrate that this CGACGG box is a novel Smad3-dependent inhibitory element (SIE) and that Smad3/Smad4 inhibits TGF-β-induced transcription of the snoN gene by binding to the SIE.

Taken together, our molecular analysis of snoN transcription indicates that the Smad2/Smad4 complex binds directly to the SBE sequences in the snoN promoter to activate its expression, while Smad3/Smad4 inhibits snoN expression through binding to an inhibitory element. Thus, Smad2 and Smad3 play opposite roles in the regulation of snoN expression, and this also provides an explanation for our earlier observation that the duration of snoN induction by TGF-β correlates positively with activation of Smad2 but not with that of Smad3.

Induction of snoN expression by TGF-β is required for TGF-β-induced transformation of AKR-2B and NRK fibroblasts.

TGF-β has been shown to promote the proliferation of many fibroblast cell lines (2). However, only AKR-2B and NRK cells undergo anchorage-independent growth in response to TGF-β1 (29, 37). We showed earlier that induction of snoN expression in these two cell lines has a longer duration and a higher intensity than in other cell lines (Fig. 1A). To determine whether this induction is required for TGF-β-induced transformation of AKR-2B cells, we first introduced a siRNA specific for the murine snoN gene stably into AKR-2B cells to reduce snoN expression. As a negative control, an unrelated siRNA was also stably introduced into AKR-2B cells (control cells). Several stable clones that showed a significant decrease in both basal and TGF-β-induced snoN expression were obtained (Fig. 6A and data not shown). Two of these clones, sisnoN#6 and -#13, were analyzed further.

In the absence of TGF-β, the sisno cells grew more slowly than AKR-2B cells and control cells (data not shown), suggesting that SnoN acts to promote cell proliferation. Interestingly, TGF-β stimulated the proliferation of both the parental AKR-2B and control cells but inhibited that of sisno cells (Fig. 6B). To examine the effect of reducing snoN expression on the ability of TGF-β to induce anchorage-independent growth of AKR-2B cells, sisno cells were subjected to a soft-agar assay. Cells were suspended in 0.4% agar medium in the absence or presence of 100 pM TGF-β1 and overlaid on the bottom agar. After 3 weeks, while both the parental AKR-2B and control cells readily formed soft-agar colonies in the presence of TGF-β, the two stable cell lines expressing the snoN siRNA failed to do so (Fig. 6C). In a parallel control experiment, none of the cell lines grew in the soft agar in the absence of TGF-β (Fig. 6C). These results indicate that SnoN expression is required for TGF-β-induced oncogenic transformation of AKR-2B fibroblasts. Thus, SnoN is an important mediator of the transforming activity of TGF-β.

Since AKR-2B and NRK cells displayed a prolonged activation of snoN expression by TGF-β and can be transformed by TGF-β, we next asked whether this prolonged induction of snoN expression is responsible for the transformation of these two cell lines. For this purpose, the TβRI inhibitor SB-431542 was employed in the soft-agar assay. AKR-2B or NRK cells were suspended in 0.4% agar medium in the presence of 100 pM TGF-β1 and overlaid on the bottom agar. After 2.5 h or 6 h of stimulation with TGF-β1, a medium containing 10 μM SB-431542 was added on top of the soft agar. As shown previously (29, 37), AKR-2B and NRK cells formed colonies in soft agar in the presence of TGF-β1. Addition of SB-431542 after 2.5 h or 6 h of TGF-β1 treatment markedly reduced the formation of soft-agar colonies (Fig. 6D). Since the addition of SB-431542 at these times allowed only transient activation of Smad2 and upregulation of snoN expression (Fig. 2C), similar to that observed in other fibroblast and epithelial cell lines, these results again support the notion that prolonged induction of snoN expression is required for TGF-β-induced oncogenic transformation of AKR-2B and NRK cells. However, this result should be interpreted with caution, since treatment of cells with SB-431542 may also shorten the activation of other Smad-independent pathways of TGF-β (53). This could also contribute to the impaired transforming phenotype.