MOLECULAR AND CELLULAR BIOLOGY, June 2011, p. 2287–2298 0270-7306/11/$12.00 doi:10.1128/MCB.01381-10 Copyright © 2011, American Society for Microbiology. All Rights Reserved.
Vol. 31, No. 11
A Novel Transcription Complex That Selectively Modulates Apoptosis of Breast Cancer Cells through Regulation of FASTKD2䌤 Kay T. Yeung,1 Sharmistha Das,1 Jin Zhang,1 Alejandro Lomniczi,2 Sergio R. Ojeda,2 Chong-Feng Xu,1 Thomas A. Neubert,1 and Herbert H. Samuels1* Department of Pharmacology, New York University School of Medicine, New York, New York 10016,1 and Division of Neuroscience, Oregon National Primate Research Center/Oregon Health and Science University, Beaverton, Oregon 970062 Received 1 December 2010/Returned for modification 28 January 2011/Accepted 15 March 2011
We previously reported that expression of NRIF3 (nuclear receptor interacting factor-3) rapidly and selectively leads to apoptosis of breast cancer cells. DIF-1 (also known as interferon regulatory factor-2 binding protein 2 [IRF-2BP2]), the cellular target of NRIF3, was identified as a transcriptional repressor, and DIF-1 knockdown leads to apoptosis of breast cancer cells but not other cell types. Here, we identify IRF-2BP1 and EAP1 (enhanced at puberty 1) as important components of the DIF-1 complex mediating both complex stability and transcriptional repression. This interaction of DIF-1, IRF-2BP1, and EAP1 occurs through the conserved C4 zinc fingers of these proteins. Microarray studies were carried out in breast cancer cell lines engineered to conditionally and rapidly increase the levels of the death domain (DD1) region of NRIF3. The DIF-1 complex was found to repress FASTKD2, a putative proapoptotic gene, in breast cancer cells and to bind to the FASTKD2 gene by chromatin immunoprecipitation. FASTKD2 knockdown prevents apoptosis of breast cancer cells from NRIF3 expression or DIF-1 knockdown, while expression of FASTKD2 leads to apoptosis of both breast and nonbreast cancer cells. Thus, regulation of FASTKD2 by NRIF3 and the DIF-1 complex acts as a novel death switch that selectively modulates apoptosis in breast cancer. release of cytochrome c, which activates the caspase-9 pathway (8, 13, 25, 38). Several years ago we identified a nuclear hormone receptor coactivator which we refer to as nuclear receptor interacting factor 3 (NRIF3) (22). Expression of NRIF3 specifically and rapidly leads to caspase-2-dependent apoptosis in breast cancer cells but not other cell types (5, 21, 34). Time lapse photography documented changes characteristic of apoptosis (5). This effect of NRIF3 on mediating the apoptosis of breast cancer cells was mapped to a short ⬃30-amino-acid region (amino acids 20 to 50) of NRIF3. We refer to this region as death domain-1 (DD1), since it is necessary and sufficient to mediate apoptosis of breast cancer cells (5, 21). The change of Ser28 to Ala28 abrogated the ability of DD1 to mediate apoptosis (5, 21), suggesting that phosphorylation of Ser28 is important for this biological effect of NRIF3/DD1. In a subsequent study we cloned an intracellular target of NRIF3/DD1 and refer to this factor as DD1 interacting factor-1 (DIF-1) (34). DIF-1 was found to be identical to interferon regulating factor-2 binding protein 2A (IRF-2BP2A), which had been reported to act as a repressor which mediated the inhibitory effect of IRF-2 on gene expression (3). We found that DIF-1 appears to act as a repressor through Sirt1, a class III histone and protein deacetylase (34). NRIF3/DD1 acts by binding to DIF-1 and reversing the repression mediated by DIF-1 (34). Our studies suggested that DIF-1 acts to selectively repress one or more proapoptotic genes in breast cancer cells which are activated when DIF-1-mediated repression is reversed. The notion that DIF-1 represses proapoptotic genes in breast cancer cells is further supported by the finding that knockdown of DIF-1 by small interfering RNA (siRNA) leads to apoptosis of breast cancer cells but not other cell types; including MCF-10A cells, an immortalized normal breast epi-
Programmed cell death or apoptosis, a fundamental process in growth and development, can be targeted in the treatment of various tumors. Apoptosis is mediated through activation of “initiator” caspases (e.g., caspase-2, -8, -9, and -10). The active initiator caspase then cleaves and activates “effector” caspases (e.g., caspase-3, -6, and -7) (7, 15, 31), which leads to cleavage of a wide variety of protein components in the cell. Although the “initiator-effector” caspase cascade is best exemplified by the “extrinsic” pathway involving surface membrane death receptors (e.g., Fas, tumor necrosis factor alpha [TNF-␣], and TRAIL), apoptosis also occurs through an intrinsic pathway initiated by intracellular stress signals, such as DNA damage and radiation (7, 19, 31). The intrinsic pathway involves changes in the permeability of the mitochondrial outer membrane that release apoptogenic factors, such as cytochrome c, which complexes with the WD-40 repeats of Apaf-1 to form oligomers which recruit procaspase-9 (37). These structures (apoptosomes) activate procaspase-9, which then activates effector caspases, such as caspase-3. Until recently the role of caspase-2 in apoptosis has been considered minor since caspase-2 knockout mice exhibit only minor phenotypic changes (4). In addition, unlike caspase-9, activated caspase-2 does not appear to cleave other caspases, leading to a proteolytic cascade (35). However, studies indicate an important role for caspase-2 in stress and DNA damageinduced mitochondrial permeability (20, 26, 38). The caspase2-mediated changes in mitochondrial permeability leads to the
* Corresponding author. Mailing address: Department of Pharmacology, MSB 424, New York University School of Medicine, 550 First Ave., New York, NY 10016. Phone: (212) 263-6279. Fax: (212) 2637133. E-mail:
[email protected]. 䌤 Published ahead of print on 28 March 2011. 2287
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thelial cell line (34). Thus, DIF-1 acts as a “death switch” whose activity can be attenuated by the binding of NRIF3/ DD1, leading to proapoptotic gene expression in breast cancer cells. To understand how DIF-1 might function to modulate apoptosis, we generated stable lines of HeLa cells and T-47D breast cancer cells expressing DIF-1 containing N-terminal FLAG and influenza hemagglutination (HA) epitope tags. We previously used this approach to identify the protein composition of a complex containing a transcription factor (NIF-1) by mass spectrometry (11). Purification of DIF-1 protein complexes by sequential FLAG and HA immunoprecipitation followed by mass spectrometry identified interferon regulatory factor-2 binding protein 1 (IRF-2BP1; we refer to this as BP1 in this study) and enhanced at puberty 1 (EAP1; also known as c14orf4) as components of a DIF-1 complex. BP1 has been reported to promote the activity of transforming growth factor  (TGF) in cells (9). Thus far, the only physiological context in which EAP1 has been shown to participate is in the neuroendocrine control of female puberty and female reproductive cyclicity (14). Knockdown of EAP1 targeted to the preoptic region of the hypothalamus results in delayed puberty and disrupted estrous cyclicity. DIF-1, BP1, and EAP1 contain highly conserved (⬎90% identical) N-terminal C4 zinc fingers and C-terminal RING fingers. These regions exhibit close to 85% identity to orthologues found in Drosophila and Caenorhabditis elegans, suggesting that these proteins may mediate essential functions, such as cell survival. BP1 and EAP-1 were initially recognized as related factors, based on sequence homology, when IRF-2BP2A (DIF-1) was first described (3). However, it was not known that these proteins could act as a complex. In this study we show that BP1 and EAP1 are components of a DIF-1 complex that is stabilized through the interactions of the conserved N-terminal C4 zinc fingers of these proteins. Knockdown of BP1 and/or EAP1 disrupts the integrity of the complex, indicating that these factors are essential for complex formation. Furthermore, the repressive effect of DIF-1 is highly dependent on EAP1 and BP1 since knockdown of one or both factors reversed DIF-1mediated repression. Consistent with these factors acting as a complex, DIF-1, BP1, and EAP1 colocalize in the nucleus in cell lines and coexpress in the specific regions of the hypothalamus where EAP1 was reported to exert its facilitatory effect on female reproductive function (14). By utilizing stable cell lines conditionally expressing DD1 in microarray and expression studies, we identified FASTKD2 (Fas-activated serine-threonine kinase domain 2) as a target gene of the DIF-1 complex in breast cancer cells. This is further supported by chromatin immunoprecipitation (ChIP) studies in T-47D cells, indicating that the DIF-1 complex binds to the 5⬘ untranslated first exon of the FASTKD2 gene. Although little is known about FASTKD2, it has been reported to localize to the inner mitochondrial membrane and to possibly play a role in apoptosis (12). We demonstrate that rapid derepression of the FASTKD2 gene in breast cancer cells is a crucial step in mediating apoptosis due to DIF-1 knockdown or NRIF3/DD1 expression. Taken together, our findings indicate that the interplay of DIF-1, BP1, EAP1, and FASTKD2 plays a central role in determining whether breast cancer cells survive or undergo apoptosis.
MOL. CELL. BIOL. MATERIALS AND METHODS Plasmids. cDNA clones of DIF-1 and BP1 in pEFplink2 vectors were a generous gift from Kay Childs and Stephen Goodbourn (3). Full-length DIF-1 in various vectors was described previously (34). High-fidelity DNA polymerase (Pfusion; Qiagen) was used to generate full-length BP1 and EAP1 cDNAs by PCR, and the fragments were cloned into pLPC. pLPC is a retrovirally based vector with a puromycin resistance gene and contains one FLAG and two HA epitope sequences upstream of the multiple cloning site (11). BP1 and EAP1 were subcloned into a pEGFP-C3 or pEGFP-C1 vector (Clontech) to generate green fluorescent protein (GFP)-BP1 or GFP-EAP1. Gal4-BP1 and Gal4-EAP1 were generated by excising the cDNAs from the corresponding pEGFP constructs and subcloning then into pSG424M, a modified pSG424 vector expressing the yeast Gal4 DNA binding domain (27). We also constructed mutants of the C4 zinc finger (Zm) or the RING finger (Rm) by changing the two cysteines, crucial for secondary structure in each region, to alanine by site-directed mutagenesis. The expressed mutants are DIF-1-Zm (the DIF-1 C16A/C19A mutant), DIF1-Rm (the DIF-1 C506A/C509A mutant), BP1-Zm (the BP1 C12A/C15A mutant), BP1-Rm (the BP1 C502A/C505A mutant), EAP1-Zm (the EAP1 C14A/ C17A mutantt), and EAP1-Rm (the EAP1 C715A/C718A mutant) in either a pEGFP or pSG424M (GAL4 DNA-binding domain [DBD] fusion) vector. The pLPC-DD1-ERT2 or pLPC-DD1(S28A)-ERT2 mutant was generated by cloning the PCR-amplified fragments of the mutated estrogen receptor-␣ (ERT2) (10), which is activated by 4OHT, and DD1 or DD1(S28A) into a pLPC vector. Full-length FASTKD2 cDNA or ⌬M-FASTKD2 lacking the N-terminal mitochondrial import sequence was generated by PCR and cloned into p3xFLAGCMV-14 (Sigma) to yield FASTKD2-FLAG or ⌬M-FASTKD2 with a C-terminal 3⫻ FLAG tag. All constructs were confirmed by sequencing. Vectors expressing pEGFP-DD1 and pEGFP-DD1(S28A) were described previously (21). Stable cell lines. 293T cells, seeded in 15-cm dishes at 5 million cells per dish, were transfected with A retroviral packaging vector and either the pLPC retrovirally based vector or pLPC-DIF-1, pLPC-DD1-ERT2, or pLPC-DD1(S28A)ERT2 by calcium phosphate precipitation. The retroviral supernatant was collected at 36 h and 60 h posttransfection. The supernatant was then filtered through a 0.45-um sterile filter and added to HeLa cells or T-47D cells for infection. Forty-eight hours postinfection, cells were selected through resistance to 2 g/ml puromycin for 2 weeks. Single colonies of each of the stable cell lines were isolated by serial dilution and screened for the expression of FLAG–HA– DIF-1 by immunocytochemistry using anti-HA antibody at a 1:200 dilution. Expression of FLAG–HA–DIF-1, FLAG–HA–DD1–ERT2, or FLAG–HA– DD1(S28A)–ERT2 in the isolated clones was confirmed by Western blotting. Whole-cell lysis and extract preparation for dual-tag affinity purification. HeLa or T-47D cell lines stably expressing FLAG–HA–DIF-1 or only the FLAG-HA tag were used. Approximately 109 adherent cells growing in monolayer in 15-cm dishes were collected for each dual-tag purification. The cells were washed in ice-cold phosphate-buffered saline and pelleted. The cell pellet was swollen in an equal volume of hypotonic buffer (20 mM Tris-HCl [pH 7.4], 20 mM KCl, 1.5 mM MgCl2, 0.2 mM EDTA, 25% glycerol, 0.2 mM phenylmethylsulfonyl fluoride [PMSF]) and homogenized 15 times with a glass pestle (Kontes). After homogenization, an equal volume of high-salt buffer (20 mM Tris-HCl [pH 7.4], 1.2 M KCl, 1.5 mM MgCl2, 0.2 mM EDTA, 25% glycerol, 0.2 mM PMSF) was added slowly, and the mixture was rotated at 4°C for 1 h. The lysed cell mixture was then centrifuged at 13,000 rpm (SS-34 rotor) for 30 min. The floating lipid layer was removed, and the supernatant collected. Three volumes of hypotonic buffer containing 0.1% Triton X-100 and one minicomplete protease inhibitor cocktail tablet (Roche) were added to the supernatant and mixed overnight at 4°C. The whole-cell lysate was then centrifuged at 4,000 rpm for 30 min the next day to remove any protein precipitates. The protein concentration was determined, and the lysates were stored at ⫺80°C for future affinity purification. Dual-tag affinity purification of the DIF-1 complex. The complex was purified by first incubating the whole-cell lysate (usually about 100 mg protein) with 300 l of anti-FLAG-M2 affinity gel (Sigma) for 4 h at 4°C. The agarose beads were then washed five times before three rounds of elution with an equal volume of 0.1 mg/ml 1⫻ FLAG peptide (Sigma) for 1 h per elution. The eluates were incubated with 100 l of anti-HA affinity gel (Sigma) overnight at 4°C. The agarose beads were then washed three times with Tris-based saline (50 mM Tris-HCl [pH 7.4] and 0.15 M NaCl) containing 0.01% Triton X-100. This amount of Triton X-100 (16) was sufficient to maintain complex formation and was below the critical micelle concentration, making it compatible with analyzing the sample by mass spectrometry (11). Proteins that remained bound were eluted twice at room temperature with an equal volume of wash buffer containing protease inhibitors
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and 1.2 mg/ml HA peptide (Covance) for 1 h per elution. Ten percent of the final total eluates was saved for SDS-PAGE followed by silver staining to visualize the isolated proteins (see Fig. 1 posted at http://hdl.handle.net/2451/29921). The remaining eluates were identified by nanoflow liquid chromatography-mass spectrometry (LC-MS/MS) (ThermoFisher Scientific LTQ-Orbitrap) after tryptic digestion in solution as previously described (11). Hybridization immunohistochemistry. This procedure was performed as previously described (1, 29) using riboprobes transcribed from EAP1, NRIF3, BP1, and DIF-1 cDNA templates. The cDNAs were generated by reverse transcription-PCR (RT-PCR) of total RNA extracted from rat or mouse brain and were cloned into the plasmid pGEM-T (Promega, Madison, WI) for subsequent in vitro transcription. The plasmids used were as follows: EAP1 cDNA, a 381-bp fragment corresponding to nucleotides (nt) 754 to 1134 in rat EAP1 mRNA (XM_234429); NRIF3 cDNA, a 401-bp fragment corresponding to nt 183 to 584 in rat NRIF3 mRNA (NM_001013213); BP1 cDNA, a 380-bp fragment corresponding to nt 1440 to 1820 in rat BP1 mRNA (NM_0011007483); and DIF-1 cDNA, a 458-bp fragment corresponding to nt 708 to 1165 in mouse DIF-1 mRNA (XM_284454). To perform the in situ hybridization procedure, the brains of immature (28- to 30-day-old) female rats were fixed by transcardiac perfusion of 4% paraformaldehyde-borate buffer, pH 9.5 (29). Thereafter, the brains were postfixed for 16 to 20 h at 4°C in the same fixative containing 10% sucrose, blocked coronally, frozen on dry ice, and stored at ⫺85°C until use. The blocks were sectioned at 30-m intervals using a freezing sliding microtome, and the sections were processed for hybridization as reported (1, 29). A double label in situ hybridization procedure (1) combining a digoxigenin-labeled EAP1 cRNA with 35S-UTP-labeled NRIF3, BP1, and DIF-1 cRNA probes was used to determine if EAP1 mRNA-containing neurons also express NRIF3, BP1, and/or DIF-1 mRNAs. The hybridization procedure employed was that recommended by Simmons et al. (29), with the modifications previously reported (1). The digoxigenin-labeled EAP1 probe was prepared exactly as described previously (1), using 500 ng of linearized plasmid template and SP6 RNA polymerase in a 10-l reaction volume. The 35S-UTP-labeled probes were prepared by transcribing linearized NRIF3, BP1, and DIF-1 cDNA templates with either T7 (DIF-1) or SP6 RNA polymerase (NRIF3 and BP1) (both enzymes from Promega) in the presence of 35S-UTP (Amersham Biosciences, Buckinghamshire, England). Following labeling, the probes were purified using a Nick column (Amersham Biosciences). Sense probes were prepared using the same templates but transcribing them from the opposite direction. Following an overnight hybridization at 55 to 56°C, the slides were washed and processed for digoxigenin detection of EAP1 mRNA (1). The sections were then dehydrated, and the slides were dipped in Ilford K5 emulsion (without defatting), instead of the NTB-2 emulsion used for isotopic hybridization, and were exposed to the emulsion for 3 weeks at 4°C in the dark. At this time the slides were developed, quickly dehydrated, dried, and coverslipped (without prior counterstaining) for microscopic examination. Immunofluorescence. Cells were plated, treated, and fixed as described below for the experiments for the terminal deoxynucleotidyltransferase-mediated dUTP-biotin nick end labeling (TUNEL) assays. Anti-FLAG-M2 antibody (Sigma) and anti-mouse fluorescein isothiocyanate (FITC) antibody (Zymed) were used to stain for FLAG-DD1-ERT2 or FASTKD2-FLAG expression in fixed cells. After treatments and/or transfections, cells were fixed and permeabilized with 1⫻ PBS with 0.2% Triton X-100 for 10 min at 25°C. After washing the cells three times with 1⫻ PBS, the cells were blocked with 3% bovine serum albumin (BSA) in 1⫻ PBS for 45 min at 25°C and then incubated with 3 g/ml of FLAG-M2 antibody (Sigma) in 3% BSA in 1⫻ PBS. After the primary antibody incubation, the cells were washed three times in 1⫻ PBS. The cells were then incubated with 7.5 g/ml of the secondary anti-mouse FITC antibody (Zymed) for 1 h at 25°C. The cells were finally washed three times in 1⫻ PBS, stained with 4⬘,6-diamidino-2-phenylindole (DAPI) to visualize nuclei, mounted on slides, examined by fluorescent microscopy, and digitally imaged. ChIP and qRT-PCR for ChIP. ChIP was performed with a few modifications of a protocol described previously (30). One 10-cm dish of approximately 9 million cells was fixed by 1% formaldehyde in 1⫻ PBS at 25°C for precisely 10 min. A final concentration of 125 mM glycine was added directly to the cells for 5 min at 25°C to quench formaldehyde and stop cross-linking. Cells were then harvested and pelleted for 5 min at 1,000 rpm at 4°C. The supernatant was aspirated, and the pellet was resuspended in 3 ml (to achieve a cell concentration of no more than 3 million cells/ml buffer) of ice-cold LB1 (50 mM HEPES [pH 7.5], 140 mM NaCl, 1 mM EDTA [pH 8], 10% glycerol, 0.5% NP-40, 0.25% Triton X-100) for 10 min in 4°C. The mixture was then centrifuged at 1,000 rpm for 5 min at 4°C to collect the nuclei. Subsequently, the pelleted nuclei were resuspended in 3 ml LB2 (200 mM NaCl, 1 mM EDTA [pH 8], 0.5 mM EGTA [pH 8], 10 mM Tris-Cl pH 8) for 10 min at 4°C and then centrifuged again at 1,000 rpm for 5 min to collect the chromatin. Finally, the chromatin pellet was
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resuspended in 3 ml LB3 (10 mM Tris-Cl, 1 mM EDTA [pH 8], 0.5 mM EGTA [pH 8]). Sodium-sarcosyl was added (final concentration of 0.5%) to the resuspended chromatin before sonication in a Bioruptor sonicator. Chromatin was sonicated in 1-ml aliquots in 15-ml Falcon tubes three times for 15 min each, with cycles of 30 s on and 30 s off. The sonicated chromatin was treated with proteinase K followed by phenol-chloroform extracted to isolate DNA. The ChIP assays utilized chromatin when the DNA was found to be ⬃150 to 250 bp as determined by agarose gel electrophoresis. Three milliliters of the sonicated samples was precleared with 40 l of packed mouse IgG beads overnight at 4°C. Subsequently, 1 ml (approximately 1 mg of chromatin) of precleared samples was incubated with a 20-l packed volume of anti-FLAG M2 affinity gel (Sigma) overnight at 4°C. One hundred microliters of the precleared sonicated samples was saved as inputs. The next day, the beads were washed 5 times for 15 min at 4°C using LB3. Then, 50 l of 0.2 mg/ml of 3⫻ FLAG peptide in 1⫻ Trisbuffered saline (TBS) was incubated with beads twice for 1 h at 4°C to elute bound proteins. The eluted samples and saved inputs were then incubated overnight at 65°C in a volume of 200 l buffer with a final concentration of 50 mM Tris, 10 mM EDTA, and 1% SDS. The next day, the samples were digested with RNase A at 37°C and proteinase K at 65°C and then extracted with phenolchloroform. The resultant DNA from the ChIP experiments was dissolved in 50 l double-distilled water, and 2 l of the dissolved DNA was used for quantitative RT-PCR (qRT-PCR) using Maxima SYBR green/fluorescein qPCR master mix (Fermentas) and the indicated primer sets. Sequences of the primers are listed in Table 3 posted at http://hdl.handle.net/2451/29921. Parallel ChIP studies using only mouse IgG-agarose beads showed no differences among the various cell lines.
RESULTS BP1 and EAP1 interact with DIF-1 and are components of the DIF-1 complex(es). A double affinity purification, similar to that previously described for the NIF-1 complex (11) and followed by in-solution digestion, was used to isolate and identify components of the DIF-1 complex(es) by mass spectrometry. Cell extracts from approximately 1 billion HeLa or T-47D cells stably expressing FLAG-HA-tagged DIF-1 or just the FLAG-HA tag were used to immunoprecipitate and purify a DIF-1 complex(es) (Fig. 1A). A fraction of the purified proteins was resolved in a 10% SDS-polyacrylamide gel and silver stained (see Fig. 1 posted at http://hdl.handle.net/2451/29921). Mass spectrometry revealed a number of proteins that associate with DIF-1 and some that may be specific for each cell line (see Table 1 at the URL mentioned above). A Mascot protein score of 50 was applied as a cutoff for 95% confidence identification. BP1 and EAP1 had the highest Mascot scores for both cell types as specific DIF-1 interactors. A similar double affinity purification was performed using FLAG-HA-DD1 in HeLa cells. DIF-1, BP1, and EAP1 were among the highest-scoring proteins identified (A. A. Tinnikov and H. H. Samuels, unpublished data). These results indicate that DIF-1, BP1, EAP1, and NRIF3/DD1 may interact to form a complex in cells and prompted us to study the role of BP1 and EAP1 on DIF-1 function. To confirm the mass spectrometry results, we carried out expression studies to examine for interaction between DIF-1, BP1, and EAP1 in mammalian cells. HeLa cells were transiently transfected with expression vectors encoding GFP– DIF-1, GFP-BP1, or GFP-EAP1 along with vectors expressing FLAG–HA–DIF-1, FLAG-HA-BP1, or FLAG-HA-EAP1. By immunoprecipitation using FLAG-M2 agarose beads, DIF-1, BP1, and EAP1 were validated to form homo-oligomers and hetero-oligomers (Fig. 1B). In addition, GFP–DIF-1, GFPBP1, and GFP-EAP1 each show a similar nuclear localization pattern with nucleoli exclusion when expressed in HeLa cells
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FIG. 1. Isolation and characterization of DIF-1-containing protein complexes from HeLa and T-47D cells. (A) Schematic for the isolation of DIF-1 complexes. HeLa and T-47D cell lines stably expressing FLAG–HA–DIF-1 or only a FLAG-HA tag were established by retrovirally mediated gene transfer. One hundred milligrams protein from whole-cell extracts derived from one billion cells was incubated with FLAG-M2 antibody agarose beads for 4 h. The agarose matrix was then washed 7 times and then incubated twice with FLAG peptide. The FLAG peptide eluted proteins were incubated with HA antibody affinity matrix overnight and then eluted twice with HA peptide. Ten percent of the final HA eluates were analyzed by SDS-PAGE followed by silver staining (see Fig. 1 posted at http://hdl.handle.net/2451/29921). The remaining HA eluates were used for protein identification by mass spectrometry after in-solution tryptic digestion (11). (B) DIF-1, BP1, and EAP1 interact in HeLa cells. HeLa cells were transiently transfected to express FLAG-HA-tagged and GFP-tagged proteins as indicated. Thirty-six hours later, cell lysates were incubated with FLAG-M2 agarose for 16 h. The beads were then washed 5 times, boiled in 2⫻ SDS loading buffer, electrophoresed in 12% SDS-PAGE gels, and immunoblotted using FLAG or GFP antibodies.
(see Fig. 2A posted at http://hdl.handle.net/2451/29921). A similar nuclear localization pattern for all three proteins was also found in T-47D and HEK293 cells (data not shown). EAP1 was previously shown to be widely expressed in the brain (14). Since EAP1 forms a complex with DIF-1 and BP1, and NRIF3 interacts with DIF-1, we sought to determine whether the mRNAs encoding these proteins are expressed in the same cells. Within the hypothalamus, EAP1 localizes to areas involved in reproductive control: the medial preoptic area, arcuate nucleus, and the ventromedial nucleus (14). Double hybridization histochemistry indicates that EAP1 mRNAcontaining neurons also express NRIF3, BP1, and DIF-1 mRNA (see Fig. 2B posted at http://hdl.handle.net/2451 /29921). This coexpression is also observed with other brain regions, as illustrated by the presence of NRIF3, BP1, and DIF-1 mRNA transcripts in EAP1 mRNA-containing cells of the piriform cortex (see Fig. 2B at the URL mentioned above). Thus, all the components of the DIF-1 repressive complex appear to express in brain cells in situ. BP1 and EAP1 are crucial for maintaining DIF-1 complex stability and DIF-1-mediated transcriptional repression. After verification of interaction and colocalization, we further characterized the DIF-1 complex(es) by size exclusion gel filtration chromatography. FLAG–HA–DIF-1 complexes were affinity purified from HeLa or T-47D cells stably expressing the protein using FLAG-M2-antibody beads. The size distribution of the DIF-1 complex was then analyzed by Superose 6 column chromatography by immunoblotting the fractions using antiFLAG, anti-BP1, and anti-EAP1 antibodies. DIF-1, BP1, and EAP1 form a distinct complex which eluted between the 660kDa and 440-kDa protein standards, with some trailing lowermolecular-weight species (Fig. 2A). Since the approximate sizes of DIF-1, BP1, and EAP1 range from 65 to 95 kDa, this indicates that either the complex contains other proteins not
yet identified and/or the complex contains multiple homo- or hetero-oligomers of DIF-1, BP1, and EAP1. To determine the role of BP1 and EAP1 within the DIF-1 complex, we knocked down BP1 and/or EAP1 by siRNA and observed a major shift in the DIF-1 complex (Fig. 2B). This shift was not a result of possible siRNA-mediated apoptosis since it occurred in the presence of zVAD-fmk (pan-caspase inhibitor) and zVDVAD 䡠 FMK (caspase-2 inhibitor). The siRNAs used against BP1 (BP1 no. 3) and EAP1 (EAP1 no. 2) (see Table 2 at http://hdl.handle.net/2451/29921) yielded more than 70% knockdown efficiency without affecting the expression of DIF-1. Forty-eight hours after knockdown of BP1 and/or EAP1 in HeLa cells stably expressing FLAG–HA– DIF-1, the DIF-1 complex was found to decrease from ⬃600 kDa to ⬃300 kDa (Fig. 2B). A similar-size reduction of the complex after BP1 and/or EAP1 knockdown was also observed with T-47D cells (Fig. 2B). Interestingly, the decreased complex sizes were similar whether BP1 or EAP1 or both were knocked down, suggesting that each protein is essential for the complex stability. To further study this, the affinity-purified input and the fractions containing FLAG–HA–DIF-1 from the column shown in Fig. 2A were analyzed for protein composition by silver stain. Results are shown for HeLa cells (see Fig. 3 posted at http: //hdl.handle.net/2451/29921), although the general pattern was similar for T-47D cells. The affinity-purified input shows that knockdown of either BP1 or EAP1 resulted in the loss of a number of proteins in the FLAG–HA–DIF-1 complex(es) (e.g., of 30 kDa and 45 kDa) (see Fig. 3A at the URL mentioned above). Knockdown of both BP1 and EAP1 showed a major change in the proteins found in the FLAG–HA–DIF-1 input. Consistent with differences in the protein composition of the input, the FLAG–HA–DIF-1-containing fractions from cells which received control siRNA showed a large number of
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FIG. 2. BP1 and EAP1 are components of and stabilize the DIF-1 complex. (A) Western blotting confirmation of components and size of the isolated DIF-1 complex in HeLa and T-47D cells. DIF-1-associated proteins, purified using FLAG-M2-agarose beads followed by FLAG peptide elution, were fractionated in a Superose 6 gel filtration column. The collected fractions were analyzed by Western blotting (after electrophoresis in 8% SDS gels) with the antibodies indicated on the left. Positions of protein standards are indicated on the top (thyroglobulin, 660 kDa; ferritin A, 440 kDa; aldolase, 160 kDa). (B) Knockdown of BP1 or EAP1 reduces the size of a DIF-1 complex(es) in HeLa and T-47D cells. BP1 siRNA and/or EAP1 siRNA were transfected in HeLa and T-47D cells stably expressing FLAG–HA–DIF-1. Cells also received zVAD-fmk and zVDVAD-fmk in case apoptosis might occur as a result of knockdown of BP1 and/or EAP1. The DIF-1 protein complex(es), isolated by FLAG-M2 beads followed by elution with FLAG peptide, was then fractionated in a Superdex 200 column. The collected fractions were analyzed by Western blotting with FLAG antibody. Positions of the protein standards are indicated on the top.
proteins (see Fig. 3B at the URL mentioned above). Knockdown of either BP1 or EAP1 (see Fig. 3C and D, respectively, at the URL mentioned above) resulted in a reduction in the number of proteins in the DIF-1 complex(es) and a shift of proteins in the 40- to 70-kDa range from the 660- to 440-kDa fractions to fractions corresponding to 160 kDa and below. Knockdown of both BP1 and EAP1 (see Fig. 3E at the URL mentioned above) eliminated almost all the proteins in the DIF-1 complex, which raises the possibility that disruption of the complex leads to the degradation or dissociation of protein components of the DIF-1 complex (although DIF-1 could still be identified by silver stain). DIF-1 in the double knockdown eluted in a fraction much greater than the predicted size for DIF-1 alone, suggesting that DIF-1 can self-interact to form oligomers, which is consistent with the results shown in Fig. 1B. Previous studies by Childs and Goodbourn (3) and our laboratory (34) indicated that DIF-1 or BP1 can mediate repression as GAL4-DBD fusion proteins. Here, we compared the extent of transcriptional repression mediated by DIF-1, BP1,
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and EAP1 as a GAL4-DBD fusion on a pG5-SV-BCAT reporter (36) in HeLa cells (Fig. 3A). T-47D cells also showed a similar transcriptional repression pattern (data not shown). The amount of plasmid used in all Gal4 transactivation experiments reflects the concentration at which maximal repression occurs, which was determined in separate dose-response studies. Interestingly, DIF-1-mediated repression was found to be almost fully reversed when BP1 or EAP1 expression was knocked down by RNA interference (RNAi) in HeLa cells (Fig. 3B) or T-47D cells (Fig. 3C). Thus, BP1 and EAP1 are important for both DIF-1 complex stability and DIF-1-mediated repression. Zinc finger structural motifs of DIF-1, BP1, and EAP1 play an important role in protein-protein interaction and transcriptional repression. After establishing that DIF-1, BP1, and EAP1 exist as a complex which mediates transcriptional repression, we further investigated the structural basis of the interaction. All three proteins have high sequence homology at the N terminus and C terminus, where, respectively, a predicted C4 zinc finger and a C3HC4 RING finger-like domain are localized (Fig. 4A). While C4 zinc finger domains are thought to be DNA binding or protein-protein interacting motifs (24), RING fingers in proteins are thought to mediate E3 ubiquitin or SUMO ligase activity or to act as protein-protein interaction domains (2). Thus, we examined whether the zinc or RING fingers of DIF-1, BP1, and EAP1 play an important role in protein-protein interactions and/or transcriptional repression. Two cysteines crucial for the C4 zinc finger (Zm) or RING finger (Rm) secondary structure in each region were changed to alanine by site-directed mutagenesis (see Materials and Methods for details). As shown in Fig. 4B, the zinc finger mutants of DIF-1, BP1, and EAP1 do not interact with wildtype DIF-1, BP1, or EAP1 while the RING finger mutants interacted with their wild-type counterparts. In addition, the zinc finger mutants exhibit a decrease in transcriptional repression, while the RING finger mutants were similar to their wild-type counterparts (Fig. 4C), indicating that the zinc finger is essential for protein-protein interactions and important for transcriptional repression mediated by DIF-1, BP1, or EAP1. FASTKD2 is a target gene of the DIF-1 complex and NRIF3/ DD1 in breast cancer cells. To understand how transcriptional regulation by the DIF-1 complex leads to apoptosis, we sought to identify proapoptotic genes that are repressed by DIF-1 and activated by NRIF3/DD1 in breast cancer cell lines but not in other cell lines. We developed a conditional system in which DD1, which inhibits DIF-1-mediated transcriptional repression, is rapidly activated, leading to expression of DIF-1-repressed target genes which are identified in microarray studies. We generated T-47D, MCF7, and HeLa stable cell lines expressing FLAG-DD1-ERT2 or FLAG-DD1(S28A)-ERT2 expressing the DD1 region of NRIF3 fused to a mutant ligand binding domain of the human estrogen receptor-␣ (ERT2) (10), which is selectively and rapidly activated by 4-hydroxytamoxifen (4OHT). Within 1 h of 4OHT (1 uM) incubation, DD1-ERT2 rapidly accumulates in the nucleus (Fig. 5A). Breast cancer stable cell lines, but not the HeLa stable cell line, expressing DD1-ERT2 exhibit apoptosis (TUNEL) after 3 h to 4 h of 4OHT treatment (Fig. 5A; see also Fig. 4 posted at http://hdl.handle.net/2451/29921). In addition, T-47D cells ex-
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FIG. 3. BP1 and EAP1 are important for transcriptional repression mediated by DIF-1. (A) DIF-1, BP1, and EAP1 mediate transcriptional repression. HeLa cells were transfected with 100 ng of the pG5-SV-BCAT reporter (36) along with 200 ng of vector expressing Gal4-DIF-1, Gal4-BP1, or Gal4-EAP1. Chloramphenicol acetyltransferase (CAT) activity was determined 24 h later. The extent of repression was studied in a separate dose-response study, and the plasmid concentration used in this study reflects the concentration at which maximal repression was obtained. Results are presented as the mean ⫾ standard error of the mean (SEM) from three representative experiments. (B) Repression by DIF-1 is dependent on BP1 and EAP1 in HeLa cells. HeLa cells were transfected with either BP1 siRNA, EAP1 siRNA, or control siRNA. One hour before introduction of the siRNAs, cells also received zVAD-fmk and zVDVAD-fmk in case apoptosis might occur as a result of knockdown of BP1 and EAP1. Forty hours later, 100 ng of pG5-SV-BCAT and 200 ng of vector expressing Gal4-DIF-1 or 150 ng (equal molar amount of plasmid) of vector expressing only the Gal4-DBD were transfected. Twenty-four hours later, the cells were harvested for CAT activity. Results are presented as the mean ⫾ SEM from three representative experiments. (C) Repression by DIF-1 is dependent on BP1 and EAP1 in T-47D cells. T-47D cells were transfected and processed for CAT assay as described in the legend for panel B. Data shown are presented as the mean ⫾ SEM from three representative experiments.
pressing DD1(S28A)-ERT2 did not exhibit apoptosis after 24 h of 4OHT treatment despite 4OHT-induced DD1(S28A)ERT2 nuclear translocation (see Fig. 4 at the URL mentioned above). Microarray studies were performed (6) with mRNA from the T-47D DD1-ERT2 or DD1(S28A)-ERT2 cell lines 4 h after incubation with 1 uM of 4OHT or an ethanol vehicle (EtOH) control. These results were compared with those for 4OHT-treated HeLa cells expressing DD1-ERT2. FASTKD2 was identified as one of the genes that was rapidly induced by 4OHT in the DD1-ERT2-expressing T-47D cells but not the DD1(S28A)-ERT2 T-47D or DD1-ERT2 HeLa cell lines. qRT-PCR validated this finding along with MCF7 DD1-ERT2expressing cells and further demonstrated that FASTKD2 expression was unchanged after 4OHT treatment of all of the stable cell lines expressing the inactive DD1(S28A)-ERT2 (Fig. 5B). FASTKD2 is essential for DD1 expression-mediated or DIF-1 knockdown-mediated apoptosis of breast cancer cells. Since FASTKD2 appears to be a target gene of DIF-1, and DD1 induces derepression of the gene, we assessed its role in the DD1/DIF-1-mediated apoptotic pathway by knockdown using siRNAs (Fig. 5C). siRNAs targeting transiently ex-
pressed FASTKD2 were tested for efficiency, in the presence of caspase inhibitors to block apoptosis. FASTKD2 siRNA no. 1 was used for studies in this paper (see Table 2 posted at http://hdl.handle.net/2451/29921), although siRNA no. 2 gave similar results. Knockdown of endogenous FASTKD2 completely protected T-47D and MCF7 stable breast cancer lines from DD1-mediated apoptosis (Fig. 5C). Figure 5 at the URL mentioned above shows similar results with SKBR3 breast cancer cells in which GFP-DD1 was transiently expressed. FASTKD2 knockdown also protected breast cancer cells from DIF-1 knockdown-mediated apoptosis (Fig. 6A; see also Fig. 5 at the URL mentioned above) as well as that mediated by GFP-NRIF3 (not shown). This protective effect appears limited to DD1/NRIF3 expression or DIF-1 knockdown-mediated apoptosis, as knockdown of FASTKD2 did not protect cells from etoposide-mediated apoptosis (Fig. 6A). FASTKD2 is a proapoptotic protein localized in the mitochondria. FASTKD2 was previously identified as an inner mitochondrial compartment protein that exhibited a nonsense mutation in both alleles in a family with infantile mitochondrial encephalomyopathy (12). The predicted domain structure of FASTKD2 is shown at the top of Fig. 5B. FASTKD2 contains
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FIG. 4. The C4 zinc finger plays an important role in protein-protein interaction and transcriptional repression by DIF-1, BP1, and EAP1. (A) The N-terminal and C-terminal regions of DIF-1, BP1, and EAP1 are highly conserved. Alignment of DIF-1, BP1, and EAP1 using a multiple sequence alignment program, ClustalW, demonstrates a single conserved N-terminal C4 zinc finger and a C-terminal C3HC4 RING finger. (B) The zinc finger motif is essential for interactions between DIF-1, BP1, and EAP1. HeLa cells were transfected with vectors expressing a FLAG-HA tag or FLAG-HA tag fusions of DIF-1, BP1, or EAP1. The cells were also transfected to express GFP-tagged fusions of DIF-1 C16A/C19A (DIF-Zm; DIF-1 zinc finger mutant), DIF-1 C506A/C509A (DIF-1 Rm; DIF-1 RING finger mutant), BP1 C12A/C15A (BP1-Zm; BP1 zinc finger mutant), or BP1 C502A/C505A (BP1-Rm; BP1 RING finger mutant) or EAP1 C14A/C17A (EAP1-Zm; EAP1 zinc finger mutant) or EAP1 C715A/C718A (EAP1-Rm; EAP1 RING finger mutant), as indicated. Thirty-six hours later cells were harvested and processed for immunoprecipitation and immunoblotting as described in the legend to Fig. 1B. (C) The zinc-finger motif is important for DIF-1-, BP1-, and EAP1-mediated transcriptional repression. HeLa cells were transfected with 100 ng of the pG5-SV-BCAT reporter along with 150 ng of vector expressing the Gal4-DBD and 200 ng of vector expressing Gal4-DBD fusion proteins of the full-length wild type (WT), Zm (zinc finger mutant), or Rm (RING finger mutant) of DIF-1, BP1, or EAP1. CAT activity was assayed 24 h later. Data shown are presented as the mean ⫾ SEM from three representative experiments.
an N-terminal ⬃50-amino-acid mitochondrial import sequence, two transmembrane domains (TM), and predicted Cterminal FAST kinase-like domains. C-terminal to the FAST kinase-like domains is a Rap domain thought to be involved in RNA binding (not shown). Mitochondria from patients with this form of infantile mitochondrial encephalomyopathy exhibit low cytochrome oxidase activity, and fibroblasts from these patients demonstrated a less efficient apoptotic response to staurosporine than wild-type fibroblasts (12). FASTKD2 mRNA is ubiquitously expressed in tissues, particularly those with abundant mitochondria (12, 28). Interestingly, we found that FASTKD2 mRNA was rapidly induced by DD1 in breast cancer cells but not HeLa cells (Fig. 5B).
FASTKD2-mediated apoptosis might be specific for breast cancer cells, or its expression could lead to apoptosis of other cell lines when expressed. To examine this, we transiently expressed C-terminal FLAG-tagged FASTKD2 (FASTKD2FLAG) in HeLa and T-47D cells, and 24 h later the cells were examined for cell localization by immunofluorescence with anti-FLAG-M2 antibody and apoptosis by a TUNEL assay (Fig. 6B). Both cell lines showed extensive apoptosis along with a perinuclear punctuate distribution pattern of FASTKD2FLAG that was characteristic of mitochondrial localization similar to that previously described (12, 28). FASTKD2mediated apoptosis was blocked by the caspase inhibitors zVAD-fmk and zVDVAD-fmk (not shown). Thus, FASTKD2
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FIG. 5. FASTKD2 expression is regulated by DD1 and is essential for DD1-mediated apoptosis. (A) T-47D cells stably expressing DD1-ERT2 undergo apoptosis after 3 h of 4-hydroxytamoxifen (4OHT) incubation. T-47D FLAG-DD1-ERT2 stable cells were treated with either ethanol control (EtOH) for 4 h or 1 uM of 4OHT at 1, 2, 3, and 4 h. Cells were then fixed and permeabilized for immunofluorescence for FLAG (green) or TUNEL (red) assay. Nuclei are stained with DAPI (blue). (B) FASTKD2 mRNA is rapidly induced by 4OHT in breast cancer cells expressing DD1-ERT2 but not DD1-ERT2-expressing HeLa cells. Expression of the DIF-1 target proapoptotic gene FASTKD2 was examined by qRT-PCR after 5 h of 1 uM of 4OHT or EtOH treatment in T-47D, MCF7, or HeLa stable cell lines expressing FLAG-DD1-ERT2 or FLAG-DD1(S28A)ERT2. Fold induction represents the FASTKD2 expression values of 4OHT-treated cells relative to those of EtOH-treated cells. Data represent the mean ⫾ SEM from three representative experiments. Structural domains of FASTKD2 are also shown at the top. (C) Knockdown of FASTKD2 protects breast cancer cell lines from DD1-mediated apoptosis. FASTKD2 siRNA was transfected 24 h prior to 4OHT or EtOH control treatment in T-47D and MCF7 cell lines expressing FLAG-HA-DD1-ERT2. Twenty-four hours later, cells were fixed and processed for FLAG expression (green) and for TUNEL (red) assay. Nuclei were stained with DAPI (blue).
appears to be a general caspase-dependent proapoptotic factor, which in breast cancer cells is rapidly induced in the NRIF3/DIF-1-mediated apoptotic pathway. Mitochondrial localization is not required for FASTKD2mediated apoptosis. An important question is whether apoptosis mediated by FASTKD2 is dependent on its mitochondrial localization. Apoptosis initiated by a rapid increase in mitochondrial FASTKD2 might result from an altered mitochondrial function, leading to changes in mitochondrial permeability. Alternatively, a rapid increase in cytoplasmic FASTKD2, which occurs prior to mitochondrial localization, could mediate the apoptotic response. To explore these alternative possibilities, we generated a vector expressing FASTKD2-FLAG
lacking the N-terminal mitochondrial import sequence (⌬MFASTKD2-FLAG). Expression of ⌬M-FASTKD2-FLAG in T-47D and HeLa cells leads to a diffuse distribution of the protein, without evidence of mitochondrial localization (Fig. 6C). Nevertheless, expression of ⌬M-FASTKD2-FLAG leads to apoptosis in both cell lines similar to that found with mitochondrially localized FASTKD2 (Fig. 6B and C). The DIF-1 complex associates with the FASTKD2 gene. To address whether the DIF-1 complex might directly regulate FASTKD2 expression, T-47D and HeLa cell lines stably expressing FLAG–HA–DIF-1 (T-47D DIF-1 and HeLa DIF-1) were used in chromatin immunoprecipitation (ChIP) studies (Fig. 7). T-47D and HeLa cell lines stably expressing only the
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FIG. 6. FASTKD2 expression leads to apoptosis but does not require mitochondrial localization. (A) Knockdown of FASTKD2 protects breast cancer cell lines from DIF-1 knockdown-mediated apoptosis. FASTKD2 siRNA was transfected 24 h prior to DIF-1 siRNA transfection (24 h) or 100 uM etoposide treatment (5 h). Cells were then fixed and processed for TUNEL (red) assay. Nuclei were stained with DAPI (blue). (B) Ectopic expression of FASTKD2 induces apoptosis in both T-47D and HeLa cell lines. Cells were transfected to express FASTKD2-FLAG. Control cells were transfected with a control vector. Twenty-four hours later the cells were fixed and processed for FLAG (green) and for TUNEL (red) assay. Nuclei were stained with DAPI (blue). (C) Ectopic expression of ⌬M-FASTKD2 induces apoptosis in HeLa and T-47D cells. HeLa cells, in duplicate wells, were transfected to express FASTKD2-FLAG or ⌬M-FASTKD2-FLAG lacking the N-terminal mitochondrial import sequence. Twenty-four hours later the cells were fixed and processed for FLAG (green) or for TUNEL (red) assay. Nuclei were stained with DAPI (blue). Identical results were found for T-47D cells (not illustrated).
FLAG-HA tag served as controls (indicated as T-47D and HeLa in Fig. 7). Sonicated chromatin was incubated with FLAG-M2 beads, and DNA from the immunoprecipitated chromatin was amplified using specific primers against regions around the predicted transcription start site of the FASTKD2 gene (Fig. 7A). DIF-1 was found to associate with a region ⬃150 to 300 bp downstream of the transcription start site of FASTKD2 (primer set B; ⫹170 to ⫹294) (Fig. 7B). In contrast, DIF-1 did not associate with the same region of the gene in the HeLa FLAG–HA–DIF-1 stable cell line (Fig. 7B). Figure 6 posted at http://hdl.handle.net/2451/29921 shows no signal with primer set A (⫹262 to ⫹389) or primer set D (⫺129 to ⫺13), while primer set C (⫹81 to ⫹243), which overlaps with primer set B, shows a weak signal. Interestingly, knockdown of BP1 or EAP1 completely abrogated the association of DIF-1 with the FASTKD2 gene (Fig. 7C), consistent with the finding that BP1 and EAP1 are essential for both DIF-1 complex formation and transcriptional repression. Taken together, these findings sug-
gest that the DIF-1 complex directly regulates FASTKD2 nearby the transcription site. DISCUSSION DIF-1 is a highly conserved protein (⬃90% identical in various mammalian species; Xenopus DIF-1 is ⬃70% identical to mammalian DIF-1). In addition, the zinc and RING fingers of DIF-1, BP1, and EAP1 exhibit close to 85% identity in orthologues found in Drosophila and C. elegans, although the role of these factors in these organisms has not been established. The high degree of conservation of the zinc and RING fingers suggests that these proteins mediate essential functions in the organism, and DIF-1 has been shown to play an important role in cell survival (18, 34). Previously, Tinnikov et al. (34) established that DIF-1 functions as an anti-apoptotic factor in breast cancer cells and not other cell lines. Based on these and other studies, we concluded that DIF-1 acts by repressing the expres-
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FIG. 7. The DIF-1 complex associates with the FASTKD2 gene in T-47D cells. (A) Position of primer sets used in qRT-PCR. After chromatin immunoprecipitation (ChIP), DNA from the bound chromatin was amplified using each of these primer sets. Sequences of the primers can be found in Table 3 posted at http://hdl.handle.net/2451/29921. TSS, predicted transcription start site of FASTKD2. (B) DIF-1 associates with a region approximately 150 to 300 nucleotides downstream of the transcription start site of FASTKD2. Sonicated short fragments of chromatin (150 to 300 bp) from T-47D or HeLa stable cell lines expressing either the FLAG-HA tag control (T-47D and HeLa) or FLAG-HA-DIF-1 (T-47D DIF-1 and HeLa DIF-1) were used for ChIP. This was performed using FLAG-M2 agarose beads followed by qRT-PCR using primer set B to amplify the ⫹170 to ⫹294 region of the FASTKD2 gene. ChIP-qRT-PCR results shown are presented as the mean ⫾ SEM from three independent experiments. See Materials and Methods for details. (C) DIF-1 association with the FASTKD2 gene is dependent on expression of BP1 and EAP1. Prior to ChIP-qRT-PCR analysis, FLAG-HA-DIF1 stable cell lines were transfected with either control siRNA, DIF-1 siRNA, BP1 siRNA, or EAP1 siRNA. Cells were analyzed by ChIP using FLAG-M2 beads and quantitated by qRT-PCR using primer set B. The ChIP results shown represent the mean ⫾ SEM from three independent experiments.
sion of proapoptotic genes and this effect can be reversed by knockdown of DIF-1 or by the binding of NRIF3/DD1 to DIF-1 (34). This notion is further supported by the finding that pretreatment of T-47D cells with ␣-amanitin completely abrogated apoptosis mediated by NRIF3/DD1 in breast cancer cells under conditions in which NRIF3/DD1 was expressed (34). In another study, U2OS osteosarcoma cells exhibited a greater degree of apoptosis in response to doxorubicin or actinomycin D after knockdown of IRF-2BP2A (DIF-1) (18). In addition, DIF-1 was reported to be a p53 target gene, and DIF-1 impedes transactivation of the p53 target genes, p21 and Bax (18). To further elucidate how DIF-1 acts in cells, we purified and analyzed components of DIF-1 complexes by mass spectrometry. We identified and characterized two DIF-1 interactors which contribute to DIF-1 activity, BP1 and EAP1. Both proteins were shown to strongly interact with DIF-1 and to form a distinct complex in both HeLa and T-47D breast cancer cells that elutes in gel filtration chromatography at about 600 kDa. BP1, also referred to as PCTA, has been reported to play a role as an antagonist and competitor
for tumor growth interacting factor (TGIF) binding of cytoplasmic promyelocytic leukemia (cPML), which leads to the release of cPML to the cytoplasm. cPML then promotes TGF-mediated signaling and transcriptional activation (9). BP1 has also recently been suggested to be a JDP2 ubiquitin ligase which mediates repression of ATF-2-dependent transcription (17). EAP1 has been implicated as a transcriptional regulator of female reproductive function through actions exerted in the preoptic region of the basal forebrain (14). Knockdown of EAP1 targeted to this region of the brain compromises both the timing of puberty and the cyclicity of female reproductive function. However, the cellular mechanism(s) underlying these deficits is unknown. Since EAP1 is necessary for repression by DIF-1, knockdown of EAP1 might act to derepress one or more genes which interfere with female puberty. As such, our findings reveal a possible mechanism by which EAP1 (and the DIF-1 complex) could control a cellular process of broad physiological significance. DIF-1 and BP1 were originally identified through a yeast twohybrid screen to act as interactors and corepressors for
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IRF-2 (3). However, we did not detect IRF-2 in our DIF-1 complex in either HeLa or T-47D cells (our unpublished data). The reason for this is not clear, although the binding of IRF-2 to the DIF-1/BP1/EAP1 complex may not be sufficiently stable to remain bound during the conditions used for isolation for mass spectrometry. Due to the high sequence homology at the N-terminal C4 zinc finger and C-terminal C3HC4 RING finger regions, we considered the possibility that DIF-1 and BP1 interact through these regions. We found that the N-terminal C4 zinc fingers of DIF-1, BP1, and EAP1 are essential for this interaction. Zinc finger mutants of DIF-1, BP1, and EAP1 fail to homo-oligomerize or hetero-oligomerize in coimmunoprecipitation studies (Fig. 4B). All three proteins mediate transcriptional repression as Gal4 fusions through the zinc finger domain. This is in agreement with previous studies showing that the RING finger regions of DIF-1 and BP1 are not sufficient to mediate repression (17, 18). Furthermore, our RNAi studies showed that transcriptional repression mediated by DIF-1 requires BP1 and EAP1 for activity (Fig. 3). In contrast, BP1- and EAP1mediated transcriptional repression is not dependent on DIF-1 (see Fig. 7 posted at http://hdl.handle.net/2451/29921). This suggests that BP1 and EAP1 may have biologic roles independent of DIF-1, such as those described previously in other studies (9, 14, 17). The finding that BP1 and/or EAP1 are necessary for DIF1-mediated repression also raises the possibility that DIF-1 might have a bifunctional role in which the composition of DIF-1 complexes in different cells may dictate whether DIF-1 acts as a repressor or activator. We have not found DIF-1 to act as an activator in the various breast and nonbreast cell lines under the conditions we studied. However, it is of interest that Teng et al. (32) reported that cardiac muscle contains very high levels of IRF-2BP2 protein compared with other tissues. The level of IRF-2BP2 protein in muscle is further increased in ischemia and was reported to stimulate expression of vascular endothelial growth factor A. This apparently occurs through the interaction of IRF-2BP2 in cardiac cells with the TEA domain transcription factor TEAD4 and VGLL4, a member of the vestigial-like family or regulators (32). Thus, like other transcription factors (e.g., nuclear hormone receptors), IRF2BP2 may have the potential to act as a repressor or activator, depending on the cell and possibly promoter context. Through expression, ChIP, and functional studies we identified FASTKD2 as a target gene of DIF-1 and NRIF3/DD1 in breast cancer cells. FASTKD2, recently identified as an inner mitochondrial compartment protein (12), contains a conserved kinase-like region of the eukaryotic Fas-activated serine-threonine kinase (FAST), a constitutively phosphorylated kinase that is known to be rapidly dephosphorylated and activated during Fas receptor ligation (33). Based on sequence homology (NCBI search), the FAST kinase-like domain is found in five human proteins (FASTKD 1 to 5) which contain two FAST kinase-like domains and a putative RNA-binding domain (RAP) near the C terminus. Although, the biological functions of all these factors are not fully known, a very recent study indicated that FASTKD 1 to 5 localize to mitochondria and FASTKD3 influences basal and stress-induced mitochondrial oxygen consumption (28). The original FAST kinase is an anti-apoptotic factor that is
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FIG. 8. Schematic model of the regulation of the FASTKD2 gene by the DIF-1 complex and NRIF3 in breast cancer cells. The DIF-1/ EAP1/BP1 complex binds to and selectively represses the FASTKD2 gene in breast cancer cells. Repression of the FASTKD2 gene by the DIF-1 complex is reversed through increased expression and/or phosphorylation of Ser28 of NRIF3. Furthermore, the model explains why apoptosis also results from knockdown of DIF-1 by siRNA (Fig. 6A).
essential for cell survival (23). Consistent with the function of the FAST kinase acting as a survival protein is the finding that RNAi knockdown of FAST leads to apoptosis and that overexpression of FAST protects cells from apoptosis (23). In contrast with that of FAST, knockdown of FASTKD2 protects cells from apoptosis, while increased expression of FASTKD2 leads to apoptosis (Fig. 6). While our results support the notion that FASTKD2 is proapoptotic, we further demonstrate that the DIF-1 complex is a transcriptional regulator of this factor in breast cancer cells and that knockdown of FASTKD2 is protective against apoptosis specifically initiated by NRIF3/ DD1 expression or DIF-1 knockdown. The discovery of FASTKD2 as a DIF-1 complex target gene and as a mediator of apoptosis by DIF-1/NRIF3 provides insight into a novel pathway that selectively modulates cell survival. Currently, the physiological roles of FASTKD2 and its related family members are not well understood. FASTKD2 is expressed in a wide variety of tissues (12, 28). Thus, its level of expression must be tightly controlled to prevent apoptosis resulting from high levels of FASTKD2 expression. Figure 8 depicts such regulation of the FASTKD2 gene by the DIF-1 complex in breast cancer cells where the DIF-1 complex represses the FASTKD2 gene and NRIF3/DD1 activation reverses this repression, leading to FASTKD2 expression and apoptosis. Since FASTKD2 expression leads to apoptosis, we explored whether FASTKD2 might mediate apoptosis after mitochondria import or, alternatively, act in the cytoplasm if rapidly synthesized, such that its level exceeds an “apoptotic threshold” prior to mitochondrial import. We generated a vector expressing FASTKD2 lacking the N-terminal mitochondrial import signal and found that FASTKD2 can mediate apoptosis independent of mitochondrial importation (Fig. 6C). Thus, our findings are consistent with a model in which a rapid and transient increase in FASTKD2 expression can lead to apoptosis, presumably through phosphorylation, leading to ac-
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tivation of proapoptotic factors or inhibition of antiapoptotic factors that reside on the surface or outside mitochondria. In summary, we have identified and characterized components and a target gene of a novel protein complex, involved in transcriptional repression, that controls whether cells survive or undergo apoptosis via the caspase-2 pathway. Although we have not identified all the protein components of the DIF-1 complex(es), our study clearly documents an important and fundamental role of this complex in cell survival. It is noteworthy and of interest that all the breast cancer cell lines we have studied (whether estrogen receptor positive or negative) undergo apoptosis when DIF-1 is knocked down or inactivated by the binding of NRIF3/DD1. An important question is which cellular events control the activity of DIF-1, which modulates FASTKD2 expression and determines cell survival. Ser28 of NRIF3 is essential for NRIF3/DD1-mediated apoptosis, suggesting that phosphorylation of Ser28 may be necessary for NRIF3-dependent apoptosis through inhibition of DIF-1 activity. Thus, a kinase(s) that phosphorylates Ser28 of NRIF3 may act as a “cell sensor” and initiate apoptosis through phosphorylation of NRIF3. Identifying the factors that regulate the NRIF3/DIF-1/BP1/EAP1 pathway and understanding how the DIF-1 complex selectively associates with and regulates the FASTKD2 gene in breast cancer cells, should provide insight into possible new approaches for the treatment of breast cancer as well as regulation of physiological events such as the development of female puberty. ACKNOWLEDGMENTS We thank Kay S. Childs and Stephen Goodbourn for providing us with cDNAs for IRF-2BP2A and IRF-2BP1. This work was supported by a grant from the NIH to H.H.S. (DK16636), an Entertainment Industry Foundation grant (to H.H.S.), a NY State Peter T. Rowley Breast Cancer Research Award (to H.H.S.), and NIH Shared Instrumentation Grant 1S10 RR017990 and NCI Cancer Institute Core Grant P30CA016087-239025 to T.A.N. K.T.Y. was supported by a Pharmacological Sciences Training Grant from the NIGMS (T32GM066704). S.R.O. was supported by NIH grants HD25123 and U54 HD18185 through a cooperative agreement as part of the Specialized Cooperative Centers Program in Reproduction and Infertility Research (NICHD) and RR000163 for operation of the Oregon National Primate Research Center. REFERENCES 1. Berg-von der Emde, K., et al. 1995. Neurotrophins and the neuroendocrine brain: different neurotrophins sustain anatomically and functionally segregated subsets of hypothalamic dopaminergic neurons. J. Neurosci. 15:4223– 4237. 2. Borden, K. L. 2000. RING domains: master builders of molecular scaffolds? J. Mol. Biol. 295:1103–1112. 3. Childs, K. S., and S. Goodbourn. 2003. Identification of novel co-repressor molecules for interferon regulatory factor-2. Nucleic Acids Res. 31:3016– 3026. 4. Colussi, P. A., and S. Kumar. 1999. Targeted disruption of caspase genes in mice: what they tell us about the functions of individual caspases in apoptosis. Immunol. Cell Biol. 77:58–63. 5. Das, S., et al. 2007. The nuclear receptor interacting factor-3 transcriptional coregulator mediates rapid apoptosis in breast cancer cells through direct and bystander-mediated events. Cancer Res. 67:1775–1782. 6. Das, S., et al. 2007. Farnesyl pyrophosphate is a novel transcriptional activator for a subset of nuclear hormone receptors. Mol. Endocrinol. 21:2672– 2686. 7. Elmore, S. 2007. Apoptosis: a review of programmed cell death. Toxicol. Pathol. 35:495–516. 8. Enoksson, M., et al. 2004. Caspase-2 permeabilizes the outer mitochondrial membrane and disrupts the binding of cytochrome c to anionic phospholipids. J. Biol. Chem. 279:49575–49578.
MOL. CELL. BIOL. 9. Faresse, N., et al. 2008. Identification of PCTA, a TGIF antagonist that promotes PML function in TGF-beta signalling. EMBO J. 27:1804–1815. 10. Feil, R., J. Wagner, D. Metzger, and P. Chambon. 1997. Regulation of Cre recombinase activity by mutated estrogen receptor ligand-binding domains. Biochem. Biophys. Res. Commun. 237:752–757. 11. Garapaty, S., et al. 2009. Identification and characterization of a novel nuclear protein complex involved in nuclear hormone receptor-mediated gene regulation. J. Biol. Chem. 284:7542–7552. 12. Ghezzi, D., et al. 2008. FASTKD2 nonsense mutation in an infantile mitochondrial encephalomyopathy associated with cytochrome c oxidase deficiency. Am. J. Hum. Genet. 83:415–423. 13. Guo, Y., S. M. Srinivasula, A. Druilhe, T. Fernandes-Alnemri, and E. S. Alnemri. 2002. Caspase-2 induces apoptosis by releasing proapoptotic proteins from mitochondria. J. Biol. Chem. 277:13430–13437. 14. Heger, S., et al. 2007. Enhanced at puberty 1 (EAP1) is a new transcriptional regulator of the female neuroendocrine reproductive axis. J. Clin. Invest. 117:2145–2154. 15. Hengartner, M. O. 2000. The biochemistry of apoptosis. Nature 407:770– 776. 16. Jin, J., et al. 2005. A mammalian chromatin remodeling complex with similarities to the yeast INO80 complex. J. Biol. Chem. 280:41207–41212. 17. Kimura, M. 2008. IRF2-binding protein-1 is a JDP2 ubiquitin ligase and an inhibitor of ATF2-dependent transcription. FEBS Lett. 582:2833–2837. 18. Koeppel, M., et al. 2009. The novel p53 target gene IRF2BP2 participates in cell survival during the p53 stress response. Nucleic Acids Res. 37:322–335. 19. Lahm, A., A. Paradisi, D. R. Green, and G. Melino. 2003. Death fold domain interaction in apoptosis. Cell Death Differ. 10:10–12. 20. Lassus, P., X. Opitz-Araya, and Y. Lazebnik. 2002. Requirement for caspase-2 in stress-induced apoptosis before mitochondrial permeabilization. Science 297:1352–1354. 21. Li, D., S. Das, T. Yamada, and H. H. Samuels. 2004. The NRIF3 family of transcriptional coregulators induces rapid and profound apoptosis in breast cancer cells. Mol. Cell. Biol. 24:3838–3848. 22. Li, D., F. Wang, and H. H. Samuels. 2001. Domain structure of the NRIF3 family of coregulators suggests potential dual roles in transcriptional regulation. Mol. Cell. Biol. 21:8371–8384. 23. Li, W., M. Simarro, N. Kedersha, and P. Anderson. 2004. FAST is a survival protein that senses mitochondrial stress and modulates TIA-1-regulated changes in protein expression. Mol. Cell. Biol. 24:10718–10732. 24. Matthews, J. M., and M. Sunde. 2002. Zinc fingers—folds for many occasions. IUBMB Life 54:351–355. 25. Paroni, G., C. Henderson, C. Schneider, and C. Brancolini. 2002. Caspase-2 can trigger cytochrome c release and apoptosis from the nucleus. J. Biol. Chem. 277:15147–15161. 26. Robertson, J. D., et al. 2004. Processed caspase-2 can induce mitochondriamediated apoptosis independently of its enzymatic activity. EMBO Rep. 5:643–648. 27. Sadowski, I., and M. Ptashne. 1989. A vector for expressing GAL4(1–147) fusions in mammalian cells. Nucleic Acids Res. 17:7539. 28. Simarro, M., et al. 2010. Fast kinase domain-containing protein 3 is a mitochondrial protein essential for cellular respiration. Biochem. Biophys. Res. Commun. 401:440–446. 29. Simmons, D. M., J. L. Arriza, and L. W. Swanson. 1989. A complete protocol for in situ hybridization of messenger RNAs in brain and other tissues with radiolabeled single-stranded RNA probes. J. Histotechnol. 12:169–181. 30. Takahashi, Y., J. B. Rayman, and B. D. Dynlacht. 2000. Analysis of promoter binding by the E2F and pRB families in vivo: distinct E2F proteins mediate activation and repression. Genes Dev. 14:804–816. 31. Taylor, R. C., S. P. Cullen, and S. J. Martin. 2008. Apoptosis: controlled demolition at the cellular level. Nat. Rev. Mol. Cell Biol. 9:231–241. 32. Teng, A. C., et al. 11 August 2010. IRF2BP2 is a skeletal and cardiac muscle-enriched ischemia-inducible activator of VEGFA expression. FASEB J. [Epub ahead of print.] 10.1096/fj.10-167049. 33. Tian, Q., J. Taupin, S. Elledge, M. Robertson, and P. Anderson. 1995. Fas-activated serine/threonine kinase (FAST) phosphorylates TIA-1 during Fas-mediated apoptosis. J. Exp. Med. 182:865–874. 34. Tinnikov, A. A., K. T. Yeung, S. Das, and H. H. Samuels. 2009. Identification of a novel pathway that selectively modulates apoptosis of breast cancer cells. Cancer Res. 69:1375–1382. 35. Troy, C. M., and M. L. Shelanski. 2003. Caspase-2 redux. Cell Death Differ. 10:101–107. 36. Witzgall, R., E. O’Leary, R. Gessner, A. J. Ouellette, and J. V. Bonventre. 1993. Kid-1, a putative renal transcription factor: regulation during ontogeny and in response to ischemia and toxic injury. Mol. Cell. Biol. 13:1933–1942. 37. Yoshida, H. 2003. The role of apaf-1 in programmed cell death: from worm to tumor. Cell Struct. Funct. 28:3–9. 38. Zhivotovsky, B., and S. Orrenius. 2005. Caspase-2 function in response to DNA damage. Biochem. Biophys. Res. Commun. 331:859–867.
