Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2009 Sep 2.
Published in final edited form as: Oncogene. 2007 Mar 12;26(38):5596–5605. doi: 10.1038/sj.onc.1210339

Pituitary tumor transforming gene interacts with Sp1 to modulate G1/S cell phase transition

Y Tong 1, Y Tan 1, C Zhou 1, S Melmed 1
PMCID: PMC2736684  NIHMSID: NIHMS121897  PMID: 17353909

Abstract

Pituitary tumor transforming gene (PTTG1) was isolated from rat pituitary tumor cells, and subsequently identified as a securin protein as well as a transcription factor. We show here a global transcriptional effect of PTTG1 in human choriocarcinoma JEG-3 cells by simultaneously assessing 20 000 gene promoters using chromatin immunoprecipitation (ChIP)-on-Chip experiments. Seven hundred and forty-six gene promoters (P<0.001) were found enriched, with functions relating to cell cycle, metabolic control and signal transduction. Significant interaction between PTTG1 and Sp1 (P<0.000001) was found by transcriptional pattern analysis of ChIP data and further confirmed by immunoprecipitation and pull-down assays. PTTG1 acts coordinately with Sp1 to induce cyclin D3 expression ∼threefold, and promotes G1/S-phase transition independently of p21. PTTG1 deletion was also protective for anchorage-independent cell colony formation. The results show that PTTG1 exhibits properties of a global transcription factor, and specifically modulates the G1/S-phase transition by interacting with Sp1. This novel signaling pathway may be required for PTTG1 transforming activity.

Keywords: PTTG1, pituitary, cell cycle

Introduction

Pituitary tumor transforming gene (PTTG1) isolated from rat pituitary tumor cells (Pei and Melmed, 1997), was subsequently identified as a securin protein (Zou et al., 1999) and a transcription factor (Dominguez et al., 1998). It also plays a role in organ development and metabolism. PTTG1 is required for rat liver regeneration, human fetal brain development (Boelaert et al., 2003) and telencephalic neurogenesis (Tarabykin et al., 2000). PTTG1-null mice exhibit testicular, splenic, pancreatic beta cell and pituitary hypoplasia (Wang et al., 2001, 2003; Chesnokova et al., 2005). Disrupted PTTG1 results in insulinopenic diabetes in adult mice, with male-selective hyperglycemia (Wang et al., 2003).

Enhanced PTTG1 abundance correlates with tumor development and size (McCabe et al., 2003). PTTG1 is induced in the early stages of estrogen-induced rat prolactinoma pathogenesis (Heaney et al., 1999) and has been suggested as a prognostic marker for differentiated thyroid cancer, lymph node invasion and breast cancer recurrence (Solbach et al., 2004), and colon cancer invasiveness and vascularity (Heaney et al., 2000).

Mechanisms of PTTG1 action include binding to p53, inhibiting its transcriptional activity and interacting with Ku, the regulatory subunit of DNA-dependent protein kinase (Wang and Melmed, 2000; Pei, 2000), activating c-Myc (Pei, 2001) and basic fibroblast growth factor (bFGF) (Chien and Pei, 2000), and acting as a securin to inhibit separase, which cleaves sister chromatids with fidelity during mitosis (Zou et al., 1999). However, it is unclear how PTTG1 overexpression mediates tumorigenesis and little is known of the functional mechanisms underlying cellular PTTG1 action.

Because of widespread PTTG1 functions, we surveyed PTTG1 effects on 20 000 gene promoters. We show here that PTTG1 acts as a global transcription factor, and specifically modulates the G1/S-phase transition by interacting with Sp1, which may underlie its transforming activity.

Results

ChIP-on-Chip experiments and microarray image analysis

The chromatin immunoprecipitation (ChIP)-on-Chip assay showed that 746 gene promoters (3.73% of all gene promoters studied) were significantly enriched (P<0.001) among 20 000 genes by immunoprecipitation with PTTG1 antibody (Supplemental Data 2) and the remaining 13 000 genes were used as background promoter set. The results of ChIP-on-Chip assay were confirmed by ChIP polymerase chain reaction (PCR) as described (Li et al., 2003). Briefly, 19 gene promoters, including ATPAF1, HUMCYT2A, C6orf102, KNTC1, COCH, PP2447, CSNK2B, RAD51L3, EIF4B, RASGRF1, FLJ31153, RPL32, GABARAPL2, SREBF2, GALNT5, SUCLG2, GCDH, TNFRSF13C and GOLGA1, were selected and conventional ChIP PCR assay used to detect enrichment of the promoters in the immunoprecipitated chromatins. The negative primers used were from the ChIP-IT kit (Active Motif). The results showed that at least 95% of the observed enriched genes are true positives (Figure 1a).

Figure 1.

Figure 1

Open in a new tab

ChIP-on-Chip and transcription pattern analysis. (a) Nineteen genes were randomly selected from the ChIP-on-Chip results. JEG-3chromatin was immunoprecipitated by PTTG1 antibody and regular PCR assay carried out using ChIP-IT kits (Active Motif) to detect and confirm promoter enrichment. All 19 gene promoters were confirmed as positive. (b) Transcription pattern analysis: frequency distribution of occurrence of the Sp1 motif in the 1000 randomly selected background promoter data sets. Solid red line represents frequency of occurrence of the Sp1 motif in the enriched promoter data set. Frequency of Sp1 in the PTTG1-enriched set was significantly (P<0.000001) higher than expected by chance. (c) Frequency distribution of occurrence of the Sp1 motif along the promoter sequence (10 intervals, each 200 bp long, ranging from −1800 bp to 200 bp) for both the enriched and background promoter set (containing the remaining non-enriched promoters). The largest difference appeared to be −200 bp from the TSS.

Transcriptional pattern analysis

The transcriptional pattern analysis as described in Materials and methods was then performed to unravel transcriptional patterns yielded by the ChIP-on-Chip results. This approach allowed us to test functional relationships of PTTG1 with other transcription factors and to identify transcriptional regulation modules controlling gene expression. The same transcriptional pattern analysis was also applied to published c-Myc ChIP-on-Chip data (Li et al., 2003) and the results showed that the method appropriately identified reported Myc/Max interaction partners TFIIB, Sp1, E2F and Ap-2 (Supplemental Data 3).

Among the 746 enriched gene promoters, 618 were well characterized from the promoter sequence database (http://biowulf.bu.edu/zlab/PromoSer/promoser.html) and the corresponding sequences extracted for transcriptional motif analysis as stated above. Analysis of PTTG1 ChIP-on-Chip data and the background data sets revealed a significant interaction between PTTG1 and Sp1 (P<0.000001) (Figure 1b). To further identify the binding location of SP1, the promoter sequence of each gene in both the enriched and background promoter set was evenly divided into 10 intervals, each comprising 200 bp (from −1800 bp upstream to 200 bp downstream of transcription starting site (TSS)) and the frequency of motif occurrence in each interval was calculated and plotted in Figure 1c. The results show that the greatest difference of Sp1 binding location between enriched and background promoter sets occurs at −200 to 0 bp, suggesting that the Sp1 binding site may approximate the TSS position (Figure 1c).

PTTG1 interacts with Sp1

We used agarose-conjugated Sp1 antibody to immunoprecipitate the Sp1 complex, and as shown in Figure 2a, PTTG1 associated with Sp1 in both JEG-3and HCT116 cells, suggesting that they interact in vivo. We confirmed PTTG1 and Sp1 interaction by His-tag pulldown assay and further mapped their interaction domains using poly-His-tagged PTTG1, Sp1 and their respective fragments (Figure 2b and c). The PTTG1 N-terminus (aa 1–120) and Sp1 C-terminus (aa 401–785) bind to the full-length protein, respectively, suggesting the location of protein domains required for their physical interactions (Figure 2b, c and d).

Figure 2.

Figure 2

Open in a new tab

PTTG1 interacts with Sp1. (a) Protein complex was immunoprecipitated using anti-Sp1 antibody from JEG-3and HCT116 cell lysate and then detected by PTTG1 antibody. Results show that PTTG1 is present in the Sp1 complex in both cell lines. (b) PTTG1 was preyed by Sp1 and its C-terminus fragments (aa 401–785), showing that PTTG1 binds to Sp1 C-terminus; (c) Sp1 was preyed by PTTG1 and its N-terminus (aa 1–120), showing that Sp1 binds to PTTG1 N-terminus; (d) mapping of Sp1 and PTTG1 interaction domains.

PTTG1 regulates Sp1 binding to its motif and colocalizes with Sp1 on cyclin D3 promoter

We further examined Sp1 binding to its motif in the presence of PTTG1 and PTTG1 fragments. In Figure 3a1, we show that PTTG1 and the consensus Sp1 binding sequence in the absence of Sp1 protein did not cause a shift, suggesting that PTTG1 does not bind the Sp1 binding sequence. Incubation of Sp1 with the Sp1 binding sequence is shown to shift the band, suggesting that Sp1 binds to this sequence as expected. Incubation of Sp1, PTTG1 and the consensus Sp1 binding sequence resulted in both a shifted and a super-shifted band. The shifted band likely comprises Sp1 and consensus Sp1 binding sequence complex. As only two proteins are present in this system, the super-shifted bands appear to reflect a formation of PTTG1-Sp1-consensus Sp1 binding sequence complexes.

Figure 3.

Figure 3

Open in a new tab

PTTG colocalized with Sp1 on cyclin D3promoter and modulates Sp1 binding to its motif. Long arrow, shifted band upon adding of Sp1; short arrow, super-shifted band upon adding PTTG1 or PTTG1.F1 to Sp1–DNA. (a1) EMSA showed that full-length PTTG1 dose dependently enhanced Sp1 binding to its binding motif according to the supershifting Sp1–DNA band. (a2) PTTG1. F1 N-terminus (101–202) did not enhance Sp1 binding to its binding motif. (a3) Density of super-shifted bands (short arrow head) were quantified and compared with the band super-shifted by 50 ng PTTG1 or PTTG1 fragment (mean±s.d., n=3, P<0.01 vs control); (b) Chromatin was immunoprecipitated by Sp1 antibody, then by PTTG1 antibody and finally detected by primers targeting the cyclin D3 promoter. Results show colocalization of Sp1 and PTTG1 on the cyclin D3 promoter in both cell lines assessed.

Interestingly, although both PTTG1 and its fragment F1 bind to Sp1 and caused a super-shift, only intact PTTG1 dose dependently shifted the Sp1–DNA complex in 6% non-denaturing gels (Figure 3a1) compared with the truncated PTTG1 fragment (Figure 3a2). The supershifted bands were semiquantified and compared with bands shifted by 50 ng PTTG1 protein (Figure 3a3). PTTG1 is a bipolar protein, with a basic N-terminus and an acidic C-terminus. Intact but not truncated PTTG1 fragments enhanced Sp1 binding to its binding sites, suggesting that the PTTG1 basic N-terminus (F1) binds to Sp1, whereas its acidic C-terminus (F2) may be important for modulating Sp1 activity.

Cyclin D3was enriched in the ChIP-on-Chip assay, and has been reported to regulate the G1/S-phase transition and may also be regulated by Sp1 (Wang et al., 1999). As described above, we showed that PTTG1 interacts with Sp1, and therefore examined the effects of PTTG1 and Sp1 on cyclin D3. We first immunoprecipitated JEG-3and HCT116 cell chromatin with Sp1 antibody and subsequently with PTTG1 antibody and used primers targeting the cyclin D3 promoter (−1800 to 200 bp) to detect localization. As shown in Figure 3b, sequential ChIP by Sp1 and PTTG1 antibody revealed the presence of the cyclin D3 promoter, suggesting colocalization of the two proteins on the cyclin D3 promoter in both cell lines tested (characterization of the cyclin D3 promoter and the primers targeting cyclin D3 promoter are shown in Supplemental Data 4).

PTTG1 and Sp1 act co-ordinately in the G1/S-phase transition by regulating cyclin D3 expression, independently of p21

PTTG1 has been shown to interact with Sp1 in previous experiments and as Sp1 acts as a G1 cell cycle phase-specific transcription factor (Grinstein et al., 2002), also regulating cancer cell proliferation (Safe and Abdelrahim, 2005), we examined the role of PTTG1 on the G1/S-phase transition. A treatment matrix is shown in Figure 4a. We tested the effects of transfections on PTTG1 and Sp1 levels, and showed that the plasmid readily increases, whereas small interfering RNA (siRNA) reduces, the corresponding protein level (Figure 4b). PTTG1 overexpression or transfection with Sp1 plasmids both enhanced accumulation of cells in S phase (∼15% increase in treated vs 42.6% in control) (Figure 4c). siRNA-targeting PTTG1 or Sp1 resulted in attenuated G1/S-phase transition with increased G1 and decreased S phase, respectively (Figure 4c). Increased G1 and decreased number of cells in S phase, and subsequent inhibition of cell proliferation upon PTTG1 siRNA treatment suggests a delayed G1/S-phase transition. These results indicate that PTTG1 overexpression may facilitate the G1/S-phase transition, which is further confirmed by the observed cell cycle changes with PTTG1 overexpression. Co-transfection of PTTG1 plasmid + Sp1 siRNA or PTTG1 siRNA + Sp1 plasmid reverses the effect of each other on S phase cells (Figure 4c). These results suggest that PTTG1 and Sp1 act co-ordinately to modulate the G1/S-phase transition.

Figure 4.

Figure 4

Figure 4

Open in a new tab

Effects of PTTG1 and Sp1 on G1/S-phase transition. (a) The treatment matrix applies to panels b-f. (b) PTTG1 and Sp1 overexpression via plasmid transfection; knockdown via siRNA transfection was confirmed by Western blot. (c) JEG-3and p21−/− HCT116 cells were treated as indicated and then subjected to cell cycle analysis. PTTG1 and Sp1 overexpression increased S phase cells in both cell lines (mean ± s.d., n = 3, P<0.01 vs control). This effect was neutralized by PTTG1 si-RNA or by Sp1 siRNA. (d) Sp1 or PTTG1 overexpression enhanced cyclin D3 mRNA and this effect was neutralized by respective si-RNA for PTTG1 or Sp1 (mean ± s.d., n = 3, P<0.01 vs control). (e1) Western blot confirmed that Rb phosphorylation levels correlated with cyclin D3 expression levels in both JEG-3 and P21−/− HCT116 cell lines. (e2) The density of the bands showed in e1 were semiquantified and normalized to actin or Rb levels, respectively (mean ± s.d., n = 3, P<0.01 vs control). (f1) Dual treatment of Sp1 and PTTG1 had a synergistic effect on cyclin D3 expression in both cell lines. (f2) Bands shown in Figure 3 f1 were semiquantified and normalized by actin levels (mean ± s.d., n = 3, P<0.01 vs single treatments). (g) Proposed role of PTTG1 on regulating the G1/S-phase transition.

The G1/S-phase transition has been characterized as a balance of cyclins and p21 proteins; cyclins promote S phase, whereas p21 maintain cells in G1 arrest. Cyclin D3 was enriched in the ChIP-on-Chip assay, and has been reported to regulate the G1/S-phase transition and is potentially regulated by Sp1 (Wang et al., 1999). We therefore examined cyclin D3 levels as well as its downstream signaling pathway by modulating PTTG1 and Sp1 levels. As shown in Figure 4d and e, cyclin D3 mRNA and protein levels were enhanced two- to threefold after Sp1 or PTTG1 overexpression, and increased cyclin D3 levels were accompanied by enhanced Rb phosphorylation, thus promoting S phase (Figure 4e). Transfection of Sp1 or PTTG1 siRNAs, respectively, showed opposing effects with reduced cyclin D3 levels and reduced Rb phosporylation (Figure 4e). Co-transfection with PTTG1 plamids + Sp1 siRNA or Sp1 plasmids + PTTG1 siRNA neutralized cyclin D3 levels (Figure 4e) and co-transfection with PTTG1 + Sp1 plasmids enhanced the effect compared with transfection alone (Figure 4f) further implicating co-ordinated PTTG1 and Sp1-controlled cyclin D3 expression and downstream pathway.

We also employed p21−/− HCT116 cells to investigate the requirement for p21 in this pathway. As shown in Figure 4c–f, similar results were achieved in p21−/− HCT116 cells, suggesting that PTTG1 overexpression regulates G1/S-phase transition occurs independently of p21.

PTTG1 role in G1/S-abberant cell growth and transformation

We assessed the role of PTTG1 on G1/S genetically aberrant cells using p21−/− HCT116 cells which exhibit enhanced cell growth as well as enhanced S phase (data not shown). Knockdown of PTTG1 levels in p21−/− HCT116 cells with PTTG1 siRNA reduced cell growth by ∼60% after 4 days (Figure 5a1). Similar to cell growth inhibition, colony formation of p21−/− HCT116 cells treated with PTTG1 siRNA was reduced by ∼65% (Figure 5a2 and a3). To test effects of PTTG1 depletion on cell transformation, we employed Rb +/− and Rb +/− PTTG1−/− mouse embryo fibroblasts (MEFs) infected with retrovirus carrying different sets of oncogenes, including Ras + TAG, Ras + Myc and Ras + E1A. Rb +/− MEFs were readily transformed and showed increased cell growth and colony formation. Rb +/− PTTG1−/− MEFs exhibited reduced cell growth and colony formation compared with Rb +/− cells (Figure 5b1–7), suggesting that PTTG1 deletion is protective for cell transformation.

Figure 5.

Figure 5

Open in a new tab

Role of PTTG1 in G1/S phase genetically aberrant cells. Proliferation and colony formation assays were performed as described. Cell and colony numbers were counted manually. (a1) p21−/− HCT116 cell proliferation was reduced when treated with PTTG1 siRNA (mean ± s.d., n = 3, P<0.01 vs control). (a2) Colony formation of p21−/− HCT116 cells was reduced by PTTG1 siRNA. (a3) Colony formation of p21−/− HCT116 cells was reduced when treated with PTTG1 siRNA. Colonies counted manually (mean ± s.d., n = 3, P<0.01 vs control). (b) Ras + T-Ag, Ras + E1A, Ras + Myc, oncogenes were used to transform MEFs. (b1) Ras + T-Ag oncogenes transformed PTTG1 +/+, Rb +/− had higher proliferation rates and (b2) more colony formation compared with PTTG1−/−, Rb +/− MEFs, (b3) Colonies were counted manually (mean ± s.d., n = 3, P<0.01 vs control). (b4) Ras + E1A transformed PTTG1 +/+, Rb +/− had higher proliferation rates and (b5) more colony formation compared with PTTG1−/−, Rb +/− MEFs (mean ± s.d., n = 3, P<0.01 vs control). (b6) Ras + Myc transformed PTTG1 +/+, Rb +/− had higher proliferation rates and (b7) more colony formation compared with PTTG1−/−, Rb +/− MEFs (mean ± s.d., n = 3, P<0.01 vs control).

Discussion

PTTG1 is induced in the early stages of estrogen-induced rat prolactinoma pathogenesis (Heaney et al., 1999) and has been suggested as a prognostic marker for differentiated thyroid cancer, lymph node invasion and breast cancer recurrence (Solbach et al., 2004), and colon cancer invasiveness and vascularity (Heaney et al., 2000). Enhanced PTTG1 tumor abundance correlates with tumor development, size and malignancy (Saez et al., 1999; Honda et al., 2003). Our previous work showed that PTTG1−/− mice were viable (Wang et al., 2001), and are protected against Rb heterozygous tumor formation (Chesnokova et al., 2005), suggesting that PTTG1 knockdown could be a potential option for cancer treatment. However, improved understanding of molecular mechanisms for PTTG1 action on cell replication is still required.

ChIP-on-Chip enabled us to survey thousands of gene promoters simultaneously (Li et al., 2003), and create a snap-shot of the transcription processes taking place under specific conditions. Results of the microarray image analysis showed that PTTG1 binds to 746 gene promoters (P<0.001), indicating a global transcriptional effect. Although the transcription process is temporal and varies from cell to cell, the underlying pattern is consistent and the method provides useful information for assessing transcription factor function. Transcriptional pattern analysis showed that PTTG1 might interact with Sp1 (P<0.000001). The interaction was further confirmed by co-immunoprecipitation and His-pulldown assay and mapped interaction domains between PTTG1 and Sp1. Electrophoretic mobility shift assay (EMSA) further suggested that full-length PTTG1 rather than PTTG1 N-terminus positively regulates Sp1 activity by enhancing binding to its motif, suggesting that the PTTG1 N-terminus binds to Sp1, whereas its C-terminus is important for modulating Sp1 activity. Although the results show that PTTG1 regulates Sp1 binding to its motif, whether or not PTTG1 directly regulates Sp1 binding on the cyclin D3 promoter requires further investigation, and we cannot yet exclude the possibility that the two proteins act independently.

Sp1 is a universal transcription factor in several cancers (Safe and Abdelrahim, 2005), a G1 cell cycle phase-specific transcription factor (Grinstein et al., 2002), and regulator of cell growth and cycle progression, purine/pyrimidine synthesis and metabolism, angiogenesis and antiapoptosis related genes (Khan et al., 2003). The importance of Sp1 in mediating hormone-induced MCF-7 cell proliferation was confirmed by RNA interference of Sp1 (Abdelrahim et al., 2002), and the pivotal role of Sp1 in the G1 phase was confirmed in both HBL-100 and HeLa cells (Ryuto et al., 1996). Moreover, expression of dominant-negative Sp1 in HeLa cells increased the percentage of cells in G1 phase. As the interaction between PTTG1 and Sp1 suggests a role for PTTG1 in regulating the G1/S cell cycle phase transition, we examined the role of PTTG1 on the cell cycle. The results showed that PTTG1 and Sp1 act co-ordinately in G1/S-phase transition. In this report, we describe the role of PTTG1 in regulating Sp1 transcriptional activity. Sp1 has also been reported to regulate PTTG1 gene transcription and the PTTG1 promoter has four potential Sp1 binding sites (Clem et al., 2003). These results therefore suggest the presence of an auto-feedback back mechanism between Sp1 and PTTG1. However, in our experiments, overexpression of Sp1 had little effect on PTTG1 expression and knockdown of Sp1 modestly decreased PTTG1 expression (Figure 4b, lane F). This result may due to an already high PTTG1 expression level in cancer cells.

The G1/S-phase transition is regulated mainly by a balance of cyclins and p21 activity. Cyclins promote S-phase entry, whereas p21 keeps cells arrested in G1, and Sp1 activates cyclin Ds expression. In our study of PTTG1 targeted genes, we showed that the cyclin D3 promoter was enriched by both PTTG1 and Sp1 antibody suggesting that PTTG1 and Sp1 might coordinately control cyclin D3 expression and may be responsible for PTTG1 induced G1/S-phase transition. Our results in Figure 4 support this conclusion. As a further support of this postulate, we surveyed JEG-3 gene expression profiles by knocking down PTTG1 levels using siRNA and the Genechip microarray U133 plus 2.0. Decreased cyclin D3 expression levels were shown to be one of the most prominent changes among more than 50000 genes (data not shown), suggesting a dominant role for this pathway. G1/S transition regulation could be mediated by other pathways. For example, PTTG1 may bind to p53 and inhibit p21 transactivation (Bernal et al., 2002) which facilitates Rb phosphorylation, enabling the G1/S transition. c-Myc, induced by PTTG1, may in turn also suppress p21 expression (Pei, 2001). However, by using p21−/− HCT116 cells, we observed similar results, suggesting that the pathway functions independently of p21. Other pathways proposed for PTTG1 function include c-Myc activation in PTTG1 transformed NIH-3T3 cells, induction of bFGF expression, and binding to p53 and modulating its function. Importantly, PTTG1 exhibits securin function by binding to separase, and its increased expression may dysregulate chromatid separation and result in chromosome instability (Zou et al., 1999). The results shown here are likely effective of an integration of different pathways.

To further investigate the role of PTTG1 as a G1/S-phase regulator in cell transformation, G1/S phase genetically aberrant cells including p21−/− HCT116 colon cancer cells and Rb +/− MEFs were used. The results confirmed the PTTG1 requirement for cell transformation and further suggest a G1/S-phase role for PTTG1 in enabling cell transformation. These results support the potential use of PTTG1 as an antineoplastic therapeutic target.

Although it has been reported that PTTG1 functions as a securin and plays an important role in the G2/M-phase transition, this study demonstrates that PTTG1 also plays a role in the G1/S-phase transition. However, besides inducing cell proliferation by PTTG1 effects on G1/S-phase transition, cell proliferation may in fact also be inhibited by other PTTG1 effects on the cell cycle (Yu et al., 2000). This finding adds to our knowledge of PTTG1 action and may further elucidate the role of PTTG1 in cancer development.

In summary, using ChIP-on-ChiP technology, our results suggest a global transcriptional role for PTTG1, which may underlie its widespread functions. Based on the transcriptional pattern analysis of ChIP data, interaction between PTTG1 and Sp1 was found and further confirmed by immunoprecipitation and pulldown assays. We also describe a novel signaling pathway showing that PTTG1 interacts with Sp1 and modulates the G1/S-phase transition independently of p21. PTTG1 regulation of the G1/S-phase transition may be required for PTTG1 transforming activity.

Materials and methods

ChIP-on-Chip experiments

ChIP-on-Chip experiments were performed using H20K-chips (Aviva Systems Biology, San Diego, CA, USA) according to the manufacturer's protocol (details of the ChIP-on-Chip experiments can be found in the Supplemental Data 5).

Microarray image analysis and transcription pattern analysis

Analysis of microarray scanning images was performed according to the published protocol (Ren et al., 2000). Enrichment was considered to be significant if P<0.001 (enriched positive promoter set), and the remaining genes were used as the background promoter set. Each motif was queried against the enriched positive promoter set as well as 1000 random background promoter sets (each set containing the same number of genes randomly selected from the complete background promoter set). The corresponding frequency of occurrence of the query motif in each promoter set (enriched promoter set plus 1000 random promoter sets) was calculated and compared. Motifs with P<0.000001 were counted as significant. To further identify motif binding site locations, the promoter sequence of each gene was divided evenly into 10 intervals (each of 200 bp long, ranging from −1800 bp to 200 bp) for both the enriched and background promoter sets. The frequency of occurrence of a motif in these 10 different intervals along the promoter sequences was calculated and plotted to visually show the binding location (details of the transcriptional pattern analysis can be found in the Supplemental Data 1 and a source code is available upon request).

Other experiments

Details of the pull-down assay, co-immunoprecipitation, cell cycle analysis, real-time quantitative PCR, EMSAs and Growth curve and transformation assays can be found in the Supplemental Data 5.

Acknowledgments

p21−/− HCT116 cells were kindly provided by Dr Bert Vogelstein, Johns Hopkins University. Mouse embryos were kindly provided by Vera Chesnokova. Retroviral plasmids pBabe-ras, pWZL-E1A, pWZL-Myc and pWZL-T-Ag were from Dr Pandolfi, Memorial Sloan-Kettering Cancer Center. This work was supported by NIH Grant CA 75979 (SM) and The Doris Factor Molecular Endocrinology Laboratory.

Abbreviations

ChIP

chromatin immunoprecipitation

MEF

mouse embryo fibroblast

PTTG

pituitary tumor transforming gene

Footnotes

Supplementary Information accompanies the paper on the Oncogene website (http://www.nature.com/onc).

References

  1. Abdelrahim M, Samudio I, Smith R, III, Burghardt R, Safe S. Small inhibitory RNA duplexes for Sp1 mRNA block basal and estrogen-induced gene expression and cell cycle progression in MCF-7 breast cancer cells. J Biol Chem. 2002;277:28815–28822. doi: 10.1074/jbc.M203828200. [DOI] [PubMed] [Google Scholar]
  2. Bernal JA, Luna R, Espina A, Lazaro I, Ramos-Morales F, Romero F, et al. Human securin interacts with p53 and modulates p53-mediated transcriptional activity and apoptosis. Nat Genet. 2002;32:306–311. doi: 10.1038/ng997. [DOI] [PubMed] [Google Scholar]
  3. Boelaert K, Tannahill LA, Bulmer JN, Kachilele S, Chan SY, Kim D, et al. A potential role for PTTG/securin in the developing human fetal brain. FASEB J. 2003;17:1631–1639. doi: 10.1096/fj.02-0948com. [DOI] [PubMed] [Google Scholar]
  4. Chesnokova V, Kovacs K, Castro AV, Zonis S, Melmed S. Pituitary hypoplasia in Pttg−/− mice is protective for Rb+/− pituitary tumorigenesis. Mol Endocrinol. 2005;19:2371–2379. doi: 10.1210/me.2005-0137. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Chien W, Pei L. A novel binding factor facilitates nuclear translocation and transcriptional activation function of the pituitary tumor-transforming gene product. J Biol Chem. 2000;275:19422–19427. doi: 10.1074/jbc.M910105199. [DOI] [PubMed] [Google Scholar]
  6. Clem AL, Hamid T, Kakar SS. Characterization of the role of Sp1 and NF-Y in differential regulation of PTTG/securin expression in tumor cells. Gene. 2003;322:113–121. doi: 10.1016/j.gene.2003.08.012. [DOI] [PubMed] [Google Scholar]
  7. Dominguez A, Ramos-Morales F, Romero F, Rios RM, Dreyfus F, Tortolero M, et al. hpttg, a human homologue of rat pttg, is overexpressed in hematopoietic neoplasms. Evidence for a transcriptional activation function of hPTTG. Oncogene. 1998;17:2187–2193. doi: 10.1038/sj.onc.1202140. [DOI] [PubMed] [Google Scholar]
  8. Grinstein E, Jundt F, Weinert I, Wernet P, Royer HD. Sp1 as G1 cell cycle phase specific transcription factor in epithelial cells. Oncogene. 2002;21:1485–1492. doi: 10.1038/sj.onc.1205211. [DOI] [PubMed] [Google Scholar]
  9. Heaney AP, Horwitz GA, Wang Z, Singson R, Melmed S. Early involvement of estrogen-induced pituitary tumor transforming gene and fibroblast growth factor expression in prolactinoma pathogenesis. Nat Med. 1999;5:1317–1321. doi: 10.1038/15275. [DOI] [PubMed] [Google Scholar]
  10. Heaney AP, Singson R, McCabe CJ, Nelson V, Nakashima M, Melmed S. Expression of pituitary-tumour transforming gene in colorectal tumours. Lancet. 2000;355:716–719. doi: 10.1016/S0140-6736(99)10238-1. [DOI] [PubMed] [Google Scholar]
  11. Honda S, Hayashi M, Kobayashi Y, Ishikawa Y, Nakagawa K, Tsuchiya E. A role for the pituitary tumor-transforming gene in the genesis and progression of non-small cell lung carcinomas. Anticancer Res. 2003;23:3775–3782. [PubMed] [Google Scholar]
  12. Khan S, Abdelrahim M, Samudio I, Safe S. Estrogen receptor/Sp1 complexes are required for induction of cad gene expression by 17beta-estradiol in breast cancer cells. Endocrinology. 2003;144:2325–2335. doi: 10.1210/en.2002-0149. [DOI] [PubMed] [Google Scholar]
  13. Li Z, Van CS, Qu C, Cavenee WK, Zhang MQ, Ren B. A global transcriptional regulatory role for c-Myc in Burkitt's lymphoma cells. Proc Natl Acad Sci USA. 2003;100:8164–8169. doi: 10.1073/pnas.1332764100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  14. McCabe CJ, Khaira JS, Boelaert K, Heaney AP, Tannahill LA, Hussain S, et al. Expression of pituitary tumour transforming gene (PTTG) and fibroblast growth factor-2 (FGF-2) in human pituitary adenomas: relationships to clinical tumour behaviour. Clin Endocrinol (Oxford) 2003;58:141–150. doi: 10.1046/j.1365-2265.2003.01598.x. [DOI] [PubMed] [Google Scholar]
  15. Pei L. Activation of mitogen-activated protein kinase cascade regulates pituitary tumor-transforming gene transactivation function. J Biol Chem. 2000;275:31191–31198. doi: 10.1074/jbc.M002451200. [DOI] [PubMed] [Google Scholar]
  16. Pei L. Identification of c-myc as a down-stream target for pituitary tumor-transforming gene. J Biol Chem. 2001;276:8484–8491. doi: 10.1074/jbc.M009654200. [DOI] [PubMed] [Google Scholar]
  17. Pei L, Melmed S. Isolation and characterization of a pituitary tumor-transforming gene (PTTG) Mol Endocrinol. 1997;11:433–441. doi: 10.1210/mend.11.4.9911. [DOI] [PubMed] [Google Scholar]
  18. Ren B, Robert F, Wyrick JJ, Aparicio O, Jennings EG, Simon I, et al. Genome-wide location and function of DNA binding proteins. Science. 2000;290:2306–2309. doi: 10.1126/science.290.5500.2306. [DOI] [PubMed] [Google Scholar]
  19. Ryuto M, Ono M, Izumi H, Yoshida S, Weich HA, Kohno K, et al. Induction of vascular endothelial growth factor by tumor necrosis factor alpha in human glioma cells. Possible roles of SP-1. J Biol Chem. 1996;271:28220–28228. doi: 10.1074/jbc.271.45.28220. [DOI] [PubMed] [Google Scholar]
  20. Saez C, Japon MA, Ramos-Morales F, Romero F, Segura DI, Tortolero M, et al. hpttg is over-expressed in pituitary adenomas and other primary epithelial neoplasias. Oncogene. 1999;18:5473–5476. doi: 10.1038/sj.onc.1202914. [DOI] [PubMed] [Google Scholar]
  21. Safe S, Abdelrahim M. Sp transcription factor family and its role in cancer. Eur J Cancer. 2005;41:2438–2448. doi: 10.1016/j.ejca.2005.08.006. [DOI] [PubMed] [Google Scholar]
  22. Solbach C, Roller M, Fellbaum C, Nicoletti M, Kaufmann M. PTTG mRNA expression in primary breast cancer: a prognostic marker for lymph node invasion and tumor recurrence. Breast. 2004;13:80–81. doi: 10.1016/j.breast.2003.09.008. [DOI] [PubMed] [Google Scholar]
  23. Tarabykin V, Britanova O, Fradkov A, Voss A, Katz LS, Lukyanov S, et al. Expression of PTTG and prc1 genes during telencephalic neurogenesis. Mech Dev. 2000;92:301–304. doi: 10.1016/s0925-4773(00)00243-4. [DOI] [PubMed] [Google Scholar]
  24. Wang Z, Melmed S. Pituitary tumor transforming gene (PTTG) transforming and transactivation activity. J Biol Chem. 2000;275:7459–7461. doi: 10.1074/jbc.275.11.7459. [DOI] [PubMed] [Google Scholar]
  25. Wang Z, Moro E, Kovacs K, Yu R, Melmed S. Pituitary tumor transforming gene-null male mice exhibit impaired pancreatic beta cell proliferation and diabetes. Proc Natl Acad Sci USA. 2003;100:3428–3432. doi: 10.1073/pnas.0638052100. [DOI] [PMC free article] [PubMed] [Google Scholar]
  26. Wang Z, Yu R, Melmed S. Mice lacking pituitary tumor transforming gene show testicular and splenic hypoplasia, thymic hyperplasia, thrombocytopenia, aberrant cell cycle progression, and premature centromere division. Mol Endocrinol. 2001;15:1870–1879. doi: 10.1210/mend.15.11.0729. [DOI] [PubMed] [Google Scholar]
  27. Wang Z, Zhang Y, Lu J, Sun S, Ravid K. Mpl ligand enhances the transcription of the cyclin D3 gene: a potential role for Sp1 transcription factor. Blood. 1999;93:4208–4221. [PubMed] [Google Scholar]
  28. Yu R, Ren SG, Horwitz GA, Wang Z, Melmed S. Pituitary tumor transforming gene (PTTG) regulates placental JEG-3cell division and survival: evidence from live cell imaging. Mol Endocrinol. 2000;14:1137–1146. doi: 10.1210/mend.14.8.0501. [DOI] [PubMed] [Google Scholar]
  29. Zou H, McGarry TJ, Bernal T, Kirschner MW. Identification of a vertebrate sister-chromatid separation inhibitor involved in transformation and tumorigenesis. Science. 1999;285:418–422. doi: 10.1126/science.285.5426.418. [DOI] [PubMed] [Google Scholar]

RESOURCES