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Article |
Cdk2 and Pin1 negatively regulate the transcriptional corepressor SMRT
Correspondence to H.-Y. Kao: hxk43{at}case.edu
Silencing mediator for retinoic acid and thyroid hormone receptor (SMRT) is a transcriptional corepressor that participates in diverse signaling pathways and human diseases. However, regulation of SMRT stability remains largely unexplored. We show that the peptidyl-prolyl isomerase Pin1 interacts with SMRT both in vitro and in mammalian cells. This interaction requires the WW domain of Pin1 and SMRT phosphorylation. Pin1 regulates SMRT protein stability, thereby affecting SMRT-dependent transcriptional repression. SMRT phosphorylation at multiple sites is required for Pin1 interaction, and these sites can be phosphorylated by Cdk2, which interacts with SMRT. Cdk2-mediated phosphorylation of SMRT is required for Pin1 binding and decreases SMRT stability, whereas mutation of these phosphorylation sites abrogates Pin1 binding and stabilizes SMRT. Finally, decreases in SMRT stability occur in response to the activation of Her2/Neu/ErbB2, and this receptor functions upstream of both Pin1 and Cdk2 in the signaling cascade that regulates SMRT stability and cellular response to tamoxifen.
© 2008 Stanya et al. This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.jcb.org/misc/terms.shtml). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).
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Introduction |
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SMRT contains at least three different types of functional domains. Near the N terminus are two Swi/Ada/N-CoR/TFIID motifs in addition to two receptor interaction domains near the C terminus (Privalsky, 2004). SMRT also contains at least four independent repression domains (RDs; I–IV). Because diverse proteins are recruited to these RDs, we sought to identify novel regulators of SMRT by using RDs III and IV as bait in a yeast two-hybrid screen. We identified the peptidyl-prolyl cis-trans isomerase, Pin1, as a SMRT-interacting protein.
Pin1 is comprised of an N-terminal protein-binding WW domain and a C-terminal peptidyl-prolyl isomerase (PPIase) domain (Lu et al., 1996; Yeh and Means, 2007). The WW domain of Pin1 binds preferentially to phospho-Ser-Pro– (pS-P) or phospho-Thr-Pro (pT-P)–containing peptide sequences (Ranganathan et al., 1997; Yaffe et al., 1997), and the enzyme domain also preferentially isomerizes the prolyl bond after pT-P or pS-P. Pin1 is frequently localized to nuclei and serves as a regulatory protein for a variety of proteins associated with transcription, including cyclin E1, c-Myc, p53, SRC-3, and the retinoic acid receptor (Zacchi et al., 2002; Zheng et al., 2002; Yeh et al., 2004, 2006; Brondani et al., 2005; Yi et al., 2005; van Drogen et al., 2006). In the case of all of these transcription factors, the binding of Pin1 to a phosphorylated motif regulates the stability of its target protein. In this study, we characterize the interaction between SMRT and Pin1. We find that Pin1 binds to phosphorylated SMRT, identify the relevant protein kinase as Cdk2, and show that Cdk2 and Pin1 facilitate the degradation of SMRT. We also demonstrate that Cdk2 and Pin1 are required for ErbB2-mediated degradation of SMRT protein. Together, our data reveal a novel mechanism by which SMRT is regulated in cells.
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Results |
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-HA antibodies, we were able to coimmunoprecipitate FLAG-Pin1 with HA-SMRT (1,178–1,823; Fig. 1 A, lane 2), indicating that these proteins interact. This interaction was confirmed by showing that endogenous SMRT was able to bind GST–Pin1 (Fig. 1 B, lane 3) but not GST alone (Fig. 1 B, lane 2). Because both Pin1 and 14-3-3 proteins interact with substrates in a phosphorylation-dependent manner (Yaffe and Elia, 2001), we tested whether 14-3-3 proteins were also able to interact with SMRT (Fig. 1 C). GST–Pin1 (Fig. 1 C, lane 2) but not GST–14-3-3
(Fig. 1 C, lane 3) was able to pull down HA-SMRT (1,178–1,823). Finally, Pin1 was also able to coimmunoprecipitate endogenous SMRT (Fig. 1 D, lane 3), whereas protein A beads alone were unable to do so (Fig. 1 D, lane 2). These data confirm Pin1 to be a SMRT-interacting protein both in vitro and in mammalian cells and verify the interaction originally observed in the yeast two-hybrid screen.
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The importance of the WW domain of Pin1 for SMRT interaction suggested that phosphorylation of SMRT might be involved in this interaction. As an initial test of this hypothesis, extracts of HeLa cells expressing HA-SMRT (1,178–1,823) were incubated with increasing concentrations of calf intestinal phosphatase before incubation with GST–Pin1. Fig. 1 H shows that phosphatase treatment of extracts dramatically reduced the association of SMRT with Pin1 in vitro (lanes 5 and 6). These data indicate that phosphorylation of SMRT is very likely to play a critical role in Pin1 binding.
Pin1 affects SMRT stability
To examine the functional significance of the SMRT and Pin1 association, we tested whether ectopic expression of Pin1 affected the transcriptional repressor activity of SMRT. Transient transfections showed that Gal4-SMRT (1,050–1,823) potently repressed basal transcription of a Gal4 reporter (Fig. 2 A, lane 1), but coexpression of Pin1 reduced SMRT repression activity in a dose-dependent manner (Fig. 2 A, lanes 2–4).
Because Pin1 compromised SMRT repressor activity, we hypothesized that Pin1 might modulate SMRT-mediated repression by affecting its steady-state levels, as has been shown for other nuclear targets of Pin1. To test this possibility, SMRT (1,012–2,507) and Pin1 were coexpressed in CV-1 cells. As shown in Fig. 2 B, expression of Pin1 had a negative, dose-dependent effect on SMRT protein levels. We also evaluated the effect of Pin1 on the expression of endogenous SMRT by immunofluorescence microscopy. SMRT levels were significantly decreased in cells transfected with FLAG-Pin1 (Fig. 2 C, compare c with f), whereas the levels of the closely related corepressor N-CoR did not change (Fig. 2 C, i and l). We confirmed these observations using siRNA targeting Pin1. Fig. 2 D shows that when endogenous Pin1 levels were decreased by siRNA, endogenous SMRT protein levels increased (lanes 2 and 3), whereas HDAC3 levels showed negligible differences. Together, these data suggest that Pin1 regulates SMRT abundance.
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To further examine the effect of ectopic Pin1 on SMRT, we used Pin1–/– mouse embryonic fibroblast (MEF) cells (Yeh et al., 2006). Consistent with previous data, ectopic expression of FLAG-Pin1 in Pin1–/– MEFs dramatically decreased HA-SMRT (1,178–1,823) protein levels in a dose-dependent manner (Fig. 2 G), indicating that this mechanism is active in MEFs. We then determined whether the presence of endogenous Pin1 altered SMRT stability by performing metabolic pulse-chase labeling assays to measure the half-life of SMRT in Pin1+/+ and Pin1–/– MEFs. MEF cells were pulse labeled with [35S]methionine and [35S]cysteine followed by a chase with unlabeled amino acids. Cell extracts were subjected to immunoprecipitation with SMRT antibodies and autoradiography. Fig. 2 H shows that SMRT was more stable in Pin1–/– MEFs than in Pin+/+ MEFs. To confirm that the effect on SMRT half-life was because of Pin1, we transfected HeLa cells stably expressing short hairpin RNA against Pin1 (Reineke et al., 2008) with either WT or mutant Pin1 (C113A/A118T). Fig. 2 I shows that WT Pin1 decreased SMRT stability, whereas the mutant had little effect. From these data, we conclude that Pin1 is a SMRT-interacting protein that regulates SMRT protein levels by modulating SMRT half-life.
Mapping Pin1 interaction sites
To elucidate the mechanism by which Pin1 modulates SMRT stability, we first mapped the pS/T-P motifs in SMRT responsible for Pin1 binding. SMRT deletion constructs fused to the yeast Gal4 DBD (Fig. 3 A, left) were used in yeast two-hybrid assays.
As shown in Fig. 3 A, we found that Pin1 does not interact with the N- or C-terminal regions of SMRT; rather, Pin1 specifically interacts with amino acids 1,178–1,823 of SMRT (containing RDs III and IV). These results were confirmed by GST pull downs (Fig. 3 B) showing that Pin1 interacts specifically with the SMRT fragment corresponding to RDs III and IV (Fig. 3 B, lane 6) but neither the N- nor C-terminal fragments (Fig. 3 B, lanes 3 and 9).
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). Because loss of two potential Cdk sites abrogated GST–Pin1 interaction (S1241 and T1445; Fig. 3 D), we next investigated whether Pin1 could interact with SMRT phosphorylated at those three Cdk sites. We performed pull downs with GST–Pin1 and immunoblotted using phosphospecific antibodies for the three Cdk sites (Fig. 3 E). We found that GST–Pin1 interacted with HA-SMRT (1,178–1,823) that was phosphorylated at these three Cdk sites (Fig. 3 E, lanes 6, 9, and 12). Collectively with the phosphatase data (Fig. 1 H), these results reveal that phosphorylation of SMRT is critical for its interaction with Pin1.
Because deletion of Cdk sites in SMRT leads to loss of Pin1 binding, we next investigated the importance of these sites by generating a Ser
Ala and ThrAla triple mutant SMRT (SMRT 3x; Fig. 3 C, ). The phosphospecific antibodies were unable to detect this mutant (Fig. 4 A, lane 2), confirming the specificity of the antibodies.
SMRT 3x (1,012–2,507) was much less able to interact with GST–Pin1 (Fig. 4 B, lane 6) than the WT, indicating that intact Cdk consensus sites are critical for interaction with Pin1. In addition, we also generated double and single Cdk mutants in SMRT and found that single mutations had no significant effect on Pin1 binding, whereas the double mutants showed only slight decreases in Pin1 binding (unpublished data). SMRT 3x (1,178–1,823) was also more stable than the WT protein, as shown by cycloheximide (CHX) treatment in HeLa cells (Fig. 4 C). These data indicate that consensus Cdk sites in SMRT are phosphorylated and critical for the SMRT–Pin1 interaction.
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ErbB2 destabilizes SMRT protein level
Because ErbB2/Her2/Neu has been shown to increase Cdk activity and up-regulate Pin1 expression in certain breast cancers (Ryo et al., 2002; Timms et al., 2002), we next investigated whether ErbB2 activity could modulate SMRT half-life. Using heregulin, a small protein activator of ErbB2, and AG825, a chemical inhibitor of ErbB2, we analyzed SMRT half-life. MCF-7 cells show a decrease in SMRT half-life when treated with heregulin as compared with the vehicle control (Fig. 7 A).
Furthermore, in BT-474 cells that overexpress ErbB2, both transfected HA-SMRT (1,012–2,507; Fig. 7 B, right) and endogenous SMRT (Fig. 7 B, left) half-life decreased upon heregulin treatment. Conversely, upon treatment with the ErbB2 inhibitor AG825, both SMRT half-life (Fig. 7 B) and SMRT protein levels increased (Fig. 7 C). Finally, to test whether ErbB2 acts upstream of Cdk2 and Pin1, we used siRNA targeting either Pin1 or Cdk2 to evaluate SMRT protein levels. BT-474 breast cancer cells were transfected with siRNAs followed by treatment with heregulin. Fig. 7 D shows that heregulin decreased SMRT protein levels by half (lanes 1 and 2), whereas siRNA targeting either Pin1 or Cdk2 blocked this effect (lanes 3–6). Collectively, these results indicate that ErbB2 signaling upstream of Cdk2 and Pin1 is a potential regulatory cascade involved in regulating the stability of SMRT.
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gene targets such as c-Myc and progesterone receptor (PR). As shown in Fig. 7 E, tamoxifen treatment repressed both c-Myc and PR expression but not the control gene 36B4 (lanes 1 and 2), whereas knockdown of SMRT compromised tamoxifen-mediated repression of both c-Myc and PR expression (lanes 3 and 4), and knockdown of Pin1 or Cdk2 increased repression of both genes (lanes 5–8). Interestingly, knockdown of Pin1 led to increases in c-Myc and PR transcription, whereas Cdk2 knockdown increased PR transcript levels. These data indicate that SMRT is critical for ER-dependent gene repression in response to tamoxifen. Because tamoxifen suppresses cell proliferation of BT-474 cells, we also examined whether knockdown of SMRT, Pin1, or Cdk2 affected proliferation rates. Fig. 7 F shows that knockdown of SMRT increased cell proliferation when compared with siControl, whereas Cdk2 showed a slight decrease in cell proliferation compared with the control. Together, these data indicate that control of SMRT stability likely has multiple cellular effects, including gene expression and cell proliferation. |
Discussion |
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In tamoxifen-responsive breast cancer cells, tamoxifen-bound ER recruits corepressor complexes to inhibit ER-target gene expression (Smith et al., 1997; Fleming et al., 2004). Decreased levels of corepressors correlate with acquired tamoxifen resistance (Lavinsky et al., 1998), whereas high expression of coactivators favors an agonist effect for tamoxifen (Shang and Brown, 2002; Shou et al., 2004). Overexpression of SMRT or N-CoR promotes the antagonist activity of tamoxifen in transient transfection assays (Jackson et al., 1997). The antiproliferative activity of tamoxifen therefore depends on the relative abundance of corepressors and coactivators (Graham et al., 2000a,b; Fujita et al., 2003). Indeed, both SMRT and N-CoR are required for the antiproliferative effects of tamoxifen in MCF-7 cells (Keeton and Brown, 2005). Additionally, certain coactivators such as AIB1/SRC-3 have been shown to be up-regulated in ErbB2-positive breast tumors, which would further disrupt the corepressor/coactivator ratio (Osborne et al., 2003), and it is interesting that SRC-3 turnover is also regulated by Pin1 (Yi et al., 2005) and reversible phosphorylation (Wu et al., 2007). Here we show that SMRT protein levels can be modulated by the oncogene Her2/Neu/ErbB2 to alter the ratio of corepressors to coactivators, thereby elucidating part of the potential mechanism underlying tamoxifen resistance and aberrant transcriptional regulation in breast cancers.
Our studies clearly demonstrate that SMRT has distinct functions from N-CoR, as knockdown of SMRT is sufficient to desensitize cells to tamoxifen-mediated inhibition of PR and c-Myc expression and cell growth. Furthermore, overexpression of Pin1 in MCF-7 cells decreases SMRT protein levels but not N-CoR. Indeed, recent studies show that these two corepressors can be targets of different cellular pathways (Jepsen et al., 2000, 2007; Jonas and Privalsky, 2004; Yu et al., 2006), including growth factor signaling, DNA damage response, and normal mammalian development, and likely regulate some nonoverlapping genes. As mentioned previously, knockdown of both SMRT and N-CoR is required to overcome the antiproliferative effects of tamoxifen in MCF-7 cells (Keeton and Brown, 2005). Furthermore, N-CoR protein levels are decreased in tamoxifen-resistant tumors from a mouse model (Lavinsky et al., 1998). This indicates that a global decrease in corepressor expression is required for breast cancer progression and that there is an analogous pathway leading to the accelerated degradation of N-CoR in ErbB2-positive/tamoxifen-resistant breast cancers. It is probable that ErbB2-positive/tamoxifen-resistant breast cancers exploit a normal cellular pathway leading to the degradation of SMRT.
Pin1 has been shown to collaborate with several kinases to modulate protein stability, including MAPK, GSK3β, Cdk1, and Cdk2 (Ryo et al., 2001; Yeh et al., 2004, 2006; Pastorino et al., 2006). Our observation that Cdk2 phosphorylates SMRT to regulate its abundance supports the idea that Pin1 exerts diverse functions in different signaling pathways. We hypothesize that compartmental and temporal regulation of the association between Pin1 and its targets fine tunes Pin1 activity. For example, several Pin1-mediated degradation pathways are likely to be cell cycle dependent, as multiple cell cycle regulatory proteins are involved (Yeh and Means, 2007). It will be highly informative to explore whether SMRT protein levels are regulated in a cell cycle–dependent manner, as has been suggested previously (Park et al., 1999). Like SMRT, N-CoR also contains seven consensus Cdk phosphorylation sites. However, of the three sites identified here, only serine 1469 (S1469) is not conserved between SMRT and N-CoR. Intriguingly, S1469 is the phosphorylation site that does not change in response to Cdk2 overexpression. Therefore, it is likely that there are other kinases that may play a role in this complex regulatory pathway.
Pin1 is thought to be an important regulator of tumorigenesis and is overexpressed or underexpressed in different tumors (Bao et al., 2004; Mantovani et al., 2007; Yeh and Means, 2007); thus, it can potentially act as either an oncogene or a tumor-suppressor gene, depending on cellular context. ErbB2 activation stimulates Pin1 transcription, and in turn Pin1 stimulates the transformative properties of ErbB2-positive cells (Ryo et al., 2002). Interestingly, although activation of ErbB2 decreases SMRT protein levels in BT-474 cells, Pin1 knockdown does not affect SMRT levels (Fig. 7 D). However, knockdown of Pin1 sensitizes cells to tamoxifen-mediated repression of c-Myc and PR expression without affecting BT-474 cell proliferation (Fig. 7, E and F). These data suggest that Pin1 may affect tamoxifen sensitivity through other pathways independent of controlling SMRT protein levels.
Pin1 is capable of promoting both stabilization and degradation of several target proteins (Yeh and Means, 2007); however, in many cases, the mechanisms underlying these opposing activities remain largely unexplored. In the case of c-Myc, Pin1 promotes a conformational change that allows rapid dephosphorylation of a specific threonine residue that is required for the binding of a ubiquitin E3 ligase (Yeh et al., 2004). In other cases, the conformational switch to certain prolyl isomers (cis or trans) also makes the target protein better substrates for particular E3 ubiquitin ligases (and therefore better substrates for degradation) or protein phosphatases (Yi et al., 2005; van Drogen et al., 2006; Yeh et al., 2006; Wu et al., 2007; Yeh and Means, 2007). Proteins such as c-Myc, cyclin E, and SRC-3 use a common Pin1-interacting motif to facilitate turnover by the E3 ligase Fbw7. As this motif does not exist in SMRT, it is likely that a different E3 ligase may be involved. Thus, identification of components of the degradation machinery responsible for Pin1- and Cdk2-dependent SMRT degradation warrants further investigation and may reveal potential new therapeutic targets for treatment of certain breast cancers.
Phosphorylation of SMRT by Cdk2 may participate in some integral aspect of cell cycle regulation. Certain genes required for cell cycle progression may require removal or degradation of SMRT from their promoters in order for cells to proliferate. Another interesting hypothesis is that SMRT may play roles other than that of a corepressor as recent studies have indicated, including involvement in DNA damage repair pathways and cell cycle regulation (Li et al., 2006; Yu et al., 2006). Additionally, it will be important to explore other potential regulators of SMRT stability and activity that may play a role in human development and disease generation or progression. Our observation that cyclin E coimmunoprecipitated SMRT but not Cdk2 is intriguing. This could be caused by the inaccessibility of the epitope by cyclin E antibodies. Alternatively, but not exclusively, these data suggest that the association of cyclin E with SMRT is independent of Cdk2. It will be interesting to further investigate the functional significance of the association between SMRT and cyclin E.
The ability of Pin1 to control SMRT protein levels appears to be cell type dependent, as Pin1 knockdown significantly increases SMRT protein steady-state levels in HeLa but not in BT-474 cells (Fig. 2 D and Fig. 7 D). In contrast, knockdown of Cdk2 increases SMRT protein levels in both HeLa and BT-474 cells (Fig. 6 E and Fig. 7 D). These observations suggest that Cdk2 may use Pin1-dependent or -independent pathways to promote SMRT degradation. ErbB2 activation stimulates Cdk2 activity, and knockdown of Cdk2 significantly increases steady-state levels of SMRT proteins and blocks ErbB2-mediated SMRT degradation, indicating that Cdk2 is an integral component in the ErbB2-dependent SMRT degradation pathway. Notably, knockdown of Cdk2 abolishes ErbB2-activated gene expression (Fig. 7 E) and cell proliferation (Fig. 7 F). Our data favor an oncogenic role for Cdk2 in SMRT destabilization and suggest that decreases in SMRT protein levels correlate with relief in gene repression and increases in proliferation, both indicators of tamoxifen resistance, whereas increases in SMRT levels correlate with tamoxifen sensitivity.
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Materials and methods |
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Plasmid construction
pGBT9-SMRT yeast two-hybrid plasmids have been previously described (Kao et al., 2000). Pin1, Cdk2, and cyclin A expression vectors were generated by PCR from a HeLa library and subcloned into the CMX-1F or -1H vector (Gao et al., 2006). HA full-length SMRT and HA-SMRT (1–1,178) were derived from previously described plasmids (Park et al., 1999); HA-SMRT (1,012–2,507), HA-SMRT (1,823–2,507), and HA- and FLAG-SMRT (1,178–1,823) were derived from previously described plasmids (Chen and Evans, 1995). GST–Pin1, GST-SMRT (1,178–1,578), and GST-SMRT (1,560–1,823) were generated from HA-Pin1 and HA-SMRT (1,178–1,823), respectively, and cloned into the pGEX vector (GE Healthcare). GST–14-3-3, reporter constructs, and β-galactosidase expression vectors have been previously described (Kao et al., 2001). Site-directed mutagenesis was performed using the QuikChange kit (Stratagene).
Cell culture and transfection
HeLa, MCF-7, and CV-1 cells were grown in standard DME (Sigma-Aldrich) supplemented with 10% FBS, 50 U/ml penicillin G, and 50 µg/ml streptomycin sulfate at 37°C in 5% CO2. MEF cells were grown in DME (Invitrogen) supplemented with 10% heat-inactivated FBS. BT-474 cells were grown in DME (American Type Culture Collection) supplemented with 10% FBS. Transfections were performed using either Lipofectamine (CV-1) or Lipofectamine 2000 (HeLa, MCF-7, MEFs, and BT-474; Invitrogen) and harvested 48 h after transfection. Transfections with siRNA (Thermo Fisher Scientific) were performed using Lipofectamine 2000, and cells were harvested 72 h after transfection. Cells were treated with 20 nM calyculin A (BIOMOL International, L.P.) to inhibit phosphatase activity for 1 h before harvest where noted. Cells were treated with 100 µg/ml CHX (Sigma-Aldrich) for the indicated times before harvest where noted.
In vitro protein–protein interaction assays
GST fusion proteins, GST, GST–Pin1 (WT or mutant), and GST–14-3-3 were expressed in an Escherichia coli DH5 strain and affinity purified on glutathione–Sepharose 4B beads. In vitro pull-down assays were performed by incubating GST–Pin1 with nuclear extracts (Dignam et al., 1983) or whole cell lysates from HeLa cells according to our published protocol (Kao et al., 2000) for 1 h at 4°C. After extensive washes, SDS-PAGE sample buffer was added to the beads, boiled, and separated by SDS-PAGE. For phosphatase treatments, extracts prepared from HA-SMRT– expressing cells were treated with increasing concentrations of calf intestinal phosphatase (Roche) for 30 min at 30°C before GST pull-down assays. His6-SMRT (1,178–1,823) was purified from E. coli BL-21 cells using nickel–nitrilotriacetic acid agarose beads (QIAGEN).
Coimmunoprecipitations
Transiently transfected HeLa whole cell extracts (WCEs) were prepared according to our published protocol (Kao et al., 2000). Immunoprecipitations for transfected samples were performed using anti-Flag or anti-HA M2 beads (Sigma-Aldrich). Endogenous coimmunoprecipitations were performed using the indicated antibodies (anti-Cdk2, cyclin A, cyclin E, and 14-3-3 [Santa Cruz Biotechnology, Inc.]; anti-Pin1 [Millipore]) with HeLa nuclear extracts. Extracts were incubated with antibodies and protein A beads (RepliGen Corp.) for 3 h at 4°C. Immunopellets were washed extensively and subjected to SDS-PAGE followed by immunoblot analyses with the antibodies anti-Flag (Sigma-Aldrich), anti-HA (Roche), and anti-Pin1 (Millipore) and anti-Cdk2, anti–cyclin A, anti–14-3-3, and anti-HDAC3 (Santa Cruz Biotechnology, Inc.). SMRT and N-CoR antibodies were purified in our laboratory; phosphospecific SMRT antibodies were generated and purified by Affinity BioReagents.
Transient transfection reporter assays
CV-1 cells were cotransfected with 16.6–66.6 ng pCMX-Gal4 and pCMX-Gal4-SMRT (1,060–1,823) constructs (Nagy et al., 1997), 100 ng pMH100–thymidine kinase–Luc, and 100 ng pCMX-LacZ in DME growth medium using Lipofectamine. The amount of DNA in each transfection was kept constant by the addition of parachlorometaxylenol. Cells were harvested and assayed for luciferase activity 36–48 h after transfection. The luciferase activity was normalized to the β-galactosidase activity. Each transfection was performed in triplicate and repeated at least two times.
In vitro kinase assays
GST-SMRT (1,178–1,578 and 1,560–1,823) constructs were purified from E. coli and incubated with purified Cdk2 (New England Biolabs, Inc.) according to the manufacturer's protocol. In brief, equal amounts of purified GST-tagged proteins were incubated with 100 U of purified Cdk2–cyclin A complexes in the associated buffer (NEB) for 30 min at 30°C. For autoradiography, [32P]ATP (PerkinElmer) was used. Reactions were stopped with SDS-PAGE sample buffer, separated by SDS-PAGE, transferred to polyvinylidene difluoride membrane, and subjected to autoradiography, immunoblotting (anti-GST; Santa Cruz Biotechnology, Inc.), or Coomassie staining.
Mass spectrometry
HeLa WCEs transfected with FLAG-SMRT (1,178–1,823) and treated with 20 nM calyculin A were subjected to coimmunoprecipitations using anti-FLAG affinity beads (Sigma-Aldrich) and separated by SDS-PAGE. The gel was Coomassie stained, and protein bands were cut out, washed and destained (50% ethanol and 5% acetic acid), reduced and alkylated (DTT and iodoacetamide), dehydrated (acetonitrile), and dried in a speed vac. Samples were digested using trypsin (20 ng/µL in 50 mM ammonium bicarbonate), and peptides were analyzed by liquid chromatography-mass spectrometry (linear ion trap; Finnigan LTQ).
Pulse-chase labeling
Cells were starved 10 min before labeling with DME lacking cysteine and methionine. Cells were then pulse labeled for 30 min with DME containing [35S]Met/Cys (PerkinElmer), washed twice, and chased in DME with unlabeled Met/Cys for the indicated times. Before labeling, cells were treated with either 100 ng/ml heregulin (R&D Systems) or 0.1 µM AG825 (EMD) for 30 min. Cells were lysed, and WCEs were subjected to immunoprecipitation overnight with either anti-SMRT (our own) or anti-HA antibodies (Sigma-Aldrich), separated by SDS-PAGE, and analyzed by autoradiography. Bands were quantified using an imaging system (VersaDoc; Bio-Rad Laboratories).
Immunofluorescent microscopy
Immunofluorescent staining was performed at room temperature as described previously (Reineke et al., 2008) using the antibodies anti-SMRT (ABR) and anti-FLAG (Santa Cruz Biotechnology, Inc.). N-CoR antibodies were purified in our laboratory. Secondary antibodies (conjugated to Alexa Fluor 488 or 594) were purchased from Invitrogen. Coverslips were mounted using mounting medium with DAPI (Vectashield; Vector Laboratories). Cells were analyzed using a fluorescent microscope (DMLB; Leica) using a 40x lens (Leica) with a numerical aperture of 506744. Images were captured with a camera (7.2 Color Mosaic; Diagnostic Instruments, Inc.) and acquired using SPOT Advanced (Diagnostic Instruments, Inc.).
RT-PCR
BT-474 cells were transfected with siRNA as described in Cell culture and transfection. 48 h after transfection, cells were treated with 100 nM 4-hydroxytamoxifen (Sigma-Aldrich) or ethanol vector control. 72 h after treatment, total RNA was harvested using PrepEase RNA Spin kits (USB). cDNA was generated using Superscript II reverse transcription and oligo-dT primers (Invitrogen). PCR reactions were performed using the following primers: 36B4 forward 5'-TGTTTCATTGTGGGAGCAGAC-3', 36B4 reverse 5'-AAGCACTTCAGGGTTCTAGAT-3', c-Myc forward 5'-ATGAAAAGGCCCCCAAGGTAGTTAT-3', c-Myc reverse 5'-GCATTTGATCATGCATTTGAAACAA-3', PR forward 5'-CCATGTGGCAGATCCCACAGGAGTT-3', and PR reverse 5'-TGGAAATTCAACACTCAGTGCC-3'. Quantification was performed in Photoshop (Adobe).
Cell proliferation
BT-474 cells were transfected with siRNA as described for RT-PCR. 48 h after transfection, cells were counted, and 1,500 cells were plated into 96-well plates. Cells were treated with either 100 nM 4-hydroxytamoxifen or ethanol vehicle control. The day 0 time point was taken 
12 h after plating. Cell proliferation was measured every 24 h using the CyQUANT NF Cell Proliferation Assay kit (Invitrogen).
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Acknowledgments |
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This work was supported by the American Cancer Society (grant RSG-04-052-01-GMC) and the Ohio Cancer Research Associates (grant to H.-Y. Kao) and by the National Institutes of Health (grant CA082845 to A.R. Means and grant T32GM08056 to K.J. Stanya). H.-Y. Kao is a recipient of the James T. Pardee–Carl A. Gerstacker Assistant Professor of Cancer Research Faculty Chair in Cancer Research at Case Western Reserve University Cancer Center.
Submitted: 30 June 2008
Accepted: 4 September 2008
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