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Céline Brand,2 Philippe Lefebvre,2* and Keiko Ozato1
Laboratory of Molecular Growth Regulation, National Institute of Child Health and Human Development, National Institutes of Health, Bethesda, Maryland 20892,1 INSERM U459 and Ligue Nationale contre le Cancer, Faculté de Médecine Henri Warembourg, 59045 Lille Cedex, France2
Received 29 August 2001/ Returned for modification 12 October 2001/ Accepted 29 November 2001
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ABSTRACT |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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INTRODUCTION |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Ligand-mediated activation of RAR is dependent on a carboxy-terminal 
-helical region (activating function-2 AD) located in the ligand-binding domain (LBD). Crystal structures of several unliganded and agonist-bound LBDs have shown that upon ligand binding, the activating function-2 AD is folded against the LBD, creating a new interface suitable for coactivator binding (49). Biochemical and expression cloning approaches have led to the identification of several families of coactivators. These include proteins from the basal transcriptional machinery and several transcriptional intermediary factors (29). In addition, ligand binding leads to the release of nuclear receptor corepressors SMRT/TRAC and NCoR/RIP13 (33, 52, 54). Many of these proteins appear to function as components of large, multiprotein complexes with several enzymatic activities (acetylases and kinases).
Despite these advances, how transcription from retinoid-responsive promoters is regulated in the chromatinized state in vivo has remained incompletely understood. We have been interested in the transcription of the RARß gene because it plays a critical role in mediating ligand-dependent growth inhibition and apoptosis in many cell lines. Moreover, its transcriptional state is thought to be relevant to certain cancers in which RARß likely exerts antiproliferative effects (10, 26, 31, 35, 43, 44, 53, 64).
In P19 embryonal carcinoma (EC) cells, the RARß gene is one of the immediate-early genes induced by all-trans-retinoic acid (atRA) (22). This gene is under the control of a promoter that contains a canonical (proximal) RARE composed of a DR-5 element. It also contains a cyclic AMP (cAMP) response element and an auxiliary (distal) RARE (58, 62) (Fig. 1A).
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Comparison of transcription patterns between transiently and stably transfected reporters is a useful means to unravel molecular mechanisms of nuclear receptor (NR)-controlled transcription (59). Previous structural studies have indicated that, while stably integrated replicating promoters are fully organized into a nucleosomal array, transiently transfected promoters are not fully chromatinized (5, 50). Functional comparisons between transient and stable reporters demonstrated that transcriptional activation patterns differ greatly in the two systems. For example, a transiently transfected human immunodeficiency virus promoter is readily activated by Tat, but the stably integrated promoter has additional requirements for activation (11).
For the more extensively studied mouse mammary tumor virus (MMTV) promoter, transiently transfected promoters are constitutively transcribed, while in the stable, replicating templates, transcription is seen only after ligand addition (50, 60). In addition, while the progesterone receptor efficiently stimulates transient promoters, it poorly stimulates stably integrated reporters (60). Also notable is the marked difference in the kinetics of MMTV transcription by glucocorticoid: in a replicating template, ligand-activated transcription is followed by a sharp attenuation of transcription in the presence of ligand, while in a transient promoter, transcription continues throughout (4, 39). Some of these differences may be accounted for by the activity of chromatin-remodeling machines (27) and posttranslational modifications of histone H1 (40). Moreover, sensitivity to histone deacetylase inhibitors has been shown to differ markedly in some transient and stable promoters, including the MMTV promoter (6, 9, 18, 45).
Here, we studied RA-activated transcription from two types of RA-responsive promoters, the naturally occurring RARß and a synthetic promoter, that were introduced either transiently or stably into P19 cells. Transiently introduced reporters supported continuous, cooperative activation of transcription by RAR (retinoid) and RXR (rexinoid) ligands. In contrast, stable reporters displayed a complex biphasic pattern of transcription in which the initial cooperative activation by retinoids and rexinoids was followed by a rexinoid-insensitive state, which could be translated into RXR-mediated repression under certain circumstances. Furthermore, loss of cooperativity was completely relieved upon treatment with histone deacetylase inhibitors with a time course similar to that of the onset of loss of cooperativity.
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MATERIALS AND METHODS |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Plasmids. The reporter plasmid carrying the mRARß promoter-luciferase/GL2 and the deletion mutants characterized in Fig. 7 have been described elsewhere, as well as (DR5)2 Ld40 and DR5 Ld40 (8, 23, 48) pBKCMV SRC-1 was obtained from B. W. O'Malley.
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minimal essential medium supplemented with 10% fetal calf serum plus 1,000 U of penicillin and 10 µg of streptomycin per ml. Transient transfections were performed using the Lipofectamine Plus reagent (Gibco-BRL). Luciferase assays were performed with the dual luciferase assay system (Promega, Madison, Wis.) according to the manufacturer's guidelines. To obtain stably transfected clones, P19 cells were transfected as follows. The mRARß2 +124/+14 Luc plasmid (23) and mutant constructs were cotransfected with the pSV2neo plasmid in a 1:10 ratio and selected with G418 (200 µg/ml) for 10 to 15 days. Overexpression of SRC-1 in P19 cells was achieved by selecting cellular clones obtained as follows. P19 cells were transfected as described above using a 1:1 ratio of pBKCMV SRC-1 and mRARß promoter-luciferase DNAs. From 200 to 500 colonies of G418-resistant cells were pooled and used for analysis. Luciferase activity was normalized against protein concentration.
RNA preparation and RT-PCR. P19 EC cells were grown as above and treated with the indicated retinoic acid receptor ligands for 20 h. Total RNA was prepared using RNAble reagent (Eurobio, Les Ulis, France) according to the manufacturer's protocol. Total RNA (50 µg) was then treated with 10 U of RNase-free DNase I (Genhunter, Nashville, Tenn.) for 1 h at 37°C to digest genomic DNA. The purified RNA was adjusted to 1 µg/µl and checked for integrity by standard agarose gel electrophoresis.
Reverse transcription (RT) was performed using oligo(dT) primers as recommended by the manufacturer (Promega, Charbonnières, France). After 1 h at 37°C, PCRs were carried out as follows: 94°C for 1 min, 56°C for 1 min, and 72°C for 1 min. Numbers of cycles were adjusted for each gene to ensure that amplification reactions were in a linear range. Primers were designed to amplify cDNAs fragments ranging in size from 300 to 600 bp and were as follows: actin primers, 5'-ATCATGTTTGAGACCTTCAA-3' and 5"-CATCTCTTGCTCGAAGTCCA-3"; RARß primers, 5"-AAGTGGTAGGAAGTGAGCTG-3" and 5"-CTACATTGAGCAGTATGCCG-3".
Real-time PCR. After purification of RNAs and reverse transcription as described above, the synthesized cDNAs were analyzed by PCR amplification using the TaqMan PCR master mix (Applied Biosystems, Foster City, Calif.) and the appropriate mix of primers. Typically, a mix of 18S and luciferase primers or mRARß promoter primers was used. The 18S primers were purchased from Applied Biosystems. The fluorescein (FAM)/6-carboxymethylrhodamine (TAMRA) probe and forward and reverse primers for the luciferase transcript were ACACCCCAACATCTTCGACGCAGG, GAATTGGAATCCATCTTGCTCC (Luc 1357F), and GGAAGTTCACCGGCGTCA (Luc 1442B), respectively. The FAM/TAMRA probe and forward and reverse primers for the mRARß transcript were CAGCACCGGCATACTGCTCAA, TCAGTGGATTCACCCAGGC (RAR468F), and TCGGGACGAGCTCCTCAG (RAR557B), respectively. Reactions (40 cycles) and data analysis were carried out with an ABI Prism 7700 (Perkin-Elmer).
Nuclear extracts and Western blotting. Nuclear extracts from P19 EC and P19 cells stably transfected with SRC-1 were prepared according to Dignam and al. (24). Proteins were resolved by sodium dodecyl sulfate-10% polyacrylamide gel electrophoresis (SDS-10% PAGE) and transferred onto a nitrocellulose membrane. Immunodetection was performed using an anti-SRC-1 antiserum (Santa Cruz Biotechnology, Inc.) as described previously (41).
Micrococcal nuclease digestion. P19 cells were collected in ice-cold 1x phosphate-buffered saline (PBS). The cell pellet was resuspended in 5 volumes of buffer A (20 mM Tris-HCl [pH 7.5], 1 mM EDTA, 50 mM KCl, 1% Triton X-100, 0.15 mM spermine, 0.5 mM spermidine, 5 mM sodium butyrate, and protease inhibitor cocktail [Sigma; dilution, 1:100]). Cells were lysed by several strokes in a Dounce homogenizer with pestle A, and the homogenate was layered onto a 0.8 M sucrose cushion in buffer A. Nuclei (ca. 3 to 5 optical density units at 260 nm) were digested with increasing amounts of micrococcal nuclease (Worthington Biochemicals Corp., Lakewood, N.J.) in a final volume of 300 µl. The reaction was stopped by adding an equal volume of stop buffer (20 mM Tris-HCl [pH 7.5], 1% SDS, 25 mM EDTA, and 5 mg of proteinase K [Promega] per ml) and incubated for 8 h at 37°C. DNA was then purified, and 50 to 80 mg was digested with 25 U of PstI (RAR) or NotI and EcoNI (luciferase).
Cleaved DNAs were analyzed with native 1% agarose gels and stained with ethidium bromide. Gels were transferred to nylon membranes (Hybond N+; Amersham), and DNA was UV-cross-linked to the membrane. Hybridization was carried out with a 614-bp RAR 32P-labeled probe (Rediprime IIp; Amersham Pharmacia Biotech), which was amplified from mouse genomic DNA (SmaI-PstI). or with a 559-bp NcoI/EcoNI fragment from pGL3 (luciferase gene). After hybridization, blots were washed and radioactive material was detected using a Storm 860 fluorophosphorimager.
ChIP assays. For the chromatin immunoprecipitation (ChIP) assays, 2 x 106 cells were grown in a 20-cm dish and treated with retinoids and/or TSA as indicated. Cells were fixed with formaldehyde (1% final concentration) at 37°C for 15 min, and the reaction was stopped by adding glycine at 200 mM final concentration. Cells then were washed with ice-cold PBS and lysed by adding 400 µl of lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris-HCl [pH 8], 1x protease inhibitors [Roche Molecular Biochemicals, Indianapolis, Ind.]) to the cellular pellet. DNA was sheared by sonication, and cell debris was eliminated by centrifugation for 15 min. Supernatants were collected and diluted 10-fold in immunoprecipitation (IP) buffer (20 mM Tris-HCl [pH 8.0], 1% Triton X-100, 2 mM EDTA, 150 mM NaCl, protease inhibitors) followed by immunoclearing with 100 µl of a 50% protein A-Sepharose slurry (equilibrated in IP buffer with 200 µg of salmon sperm and 1 mg of bovine serum albumin per ml) for 2 h at 4°C.
Immunoprecipitations were performed overnight with specific antibodies (anti-acetylated histones were from Upstate Biotech, Inc., Lake Placid, N.Y., and the anti-RNAP antibody C-21 was from Santa Cruz). Complexes were collected by incubation with 60 µl of 50% protein A-Sepharose for 2 h at 4°C. Beads were washed sequentially for 10 min each in buffer A (20 mM Tris-HCl [pH 8], 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 150 mM NaCl), buffer B (20 mM Tris-HCl [pH 8.0], 0.1% SDS, 1% Triton X-100, 2 mM EDTA, 500 mM NaCl), buffer C (0.25 M LiCl, 1% NP-40, 1% sodium deoxycholate, 1 mM EDTA, 20 mM Tris-HCl [pH 8]), and twice in TE buffer (10 mM Tris-HCl [pH 8], 1 mM EDTA). Immunocomplexes were eluted off the beads by incubation with 200 µl of 1% SDS-0.1 M NaHCO3 and heated at 65°C for at least 6 h to reverse formaldehyde cross-linking.
DNA was purified with a QIAquick Spin kit (Qiagen). The -124/+31 region was amplified with the following primers: forward 5' RARß promoter, 5"-GGGAGTTTTTAAGCGCTGTGAG-3', and reverse RARß promoter, 5"-GGAGCAGCTCACTTCCTACC-3". The +165/+524 region was amplified using 5"-TAGGACCCGCGCGCTCCCGGAG-3" and 5"-ATTGAGCAGTATGCCGGTGCT-3" as forward and reverse primers, respectively. The number of PCR cycles was adjusted to maintain the amplification rate in the linear range (typically between 20 and 30 cycles).
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RESULTS |
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As expected, 9-cis-RA, an RAR-RXR panagonist, stimulated the activity of the transiently introduced reporter more potently than atRA, an RAR-specific ligand (21-fold versus 10-fold) (Fig. 1B). A similar cooperativity was observed when P19 cells were treated for 20 h by RAR subsaturating (0.1 nM) or saturating (100 nM) concentrations of TTNPB, a synthetic retinoid, in combination with 1 µM SR11237, a rexinoid. The overall level of activity was, however, lower when 0.1 nM TTNPB was used, since this ligand concentration promoted an inframaximal luciferase activity.
As shown in Fig. 1C, the endogenous mRARß promoter responded strongly, much like the transiently expressed reporter gene, to 100 nM TTNPB but not detectably to 0.1 nM TTNPB or 1 µM SR11237. In contrast, the two ligands failed to give cooperative activation of the mRARß promoter (Fig. 1C), and this lack of cooperativity was also observed for different concentrations of TTNPB and SR11237 (B. Lefebvre and K. Ozato, unpublished observations). Rather, coaddition of the two ligands led to a slight but reproducible decrease (15 to 20%) in endogenous mRARß promoter activity relative to that by TTNPB alone, indicating different mechanisms of transcription controlling transient and stable promoters.
We then asked whether the mRARß promoter is organized into nucleosomes when transiently transfected into P19 cells. In the experiment shown in Fig. 1D, nuclei from P19 cells transiently transfected with the RARß luciferase reporter gene were treated with micrococcal nuclease. Purified DNA was then Southern blotted using a probe corresponding to the RARß gene or to the luciferase gene (Fig. 1D). The endogenous RARß promoter gene showed a clear nucleosomal repeat (Fig. 1D), similar to that reported previously with the stably integrated reporter (7). In contrast, the luciferase DNA was digested evenly upon micrococcal nuclease treatment without producing a repeated pattern. These contrasting patterns gave us the impetus to study the correlation between chromatin organization of RA-sensitive genes and cooperativity between RXR and RAR.
Loss of cooperativity between RXR and RAR is also observed in P2 cells. P2 cells are a subclone of P19 cells in which 25 to 30 copies of the chimeric mRARß promoter-luciferase construct (Fig. 2A) have been introduced. atRA-induced chromatin structure alterations in this cell line recapitulate those observed in P19 cells and paralleled the ability of retinoids and rexinoids to induce transcription (7, 32).
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2-fold). On the other hand, the RAR-specific ligand TTNPB (100 nM) strongly stimulated the integrated reporter activity. Coaddition of the two ligands at these concentrations failed to give cooperative activation in the stably integrated reporter but again led to a decrease (10 to 15%) in reporter activity relative to that triggered by TTNPB alone. These results indicated that retinoids and rexinoids do not cooperatively activate stably integrated RARß promoters, in contrast to transiently transfected promoters. Furthermore, we noted that, similar to the results with P19 cells, RAR and RXR cooperatively stimulated RARß promoter activity in NIH 3T3 cells for transiently introduced reporters, but not for the same stably introduced reporter (Lefebvre and Ozato, unpublished), indicating that the different patterns of mRARß promoter activation are not a property restricted to P19 cells. Finally, this cooperativity was not specific for these two ligands, since other synthetic ligands elicited the same response profile (Lefebvre and Ozato, unpublished).
Cooperativity between RAR and RXR is abrogated in stably integrated promoters in a time- and dose-dependent manner.
Since P2 cells faithfully recapitulated events leading to transcriptional activation of the mRARß promoter, we carried out a combined dose-response and kinetic analysis of the activity of this promoter in P2 cells in response to various concentrations of TTNPB, which displays an affinity for RAR of about 10 to 20 nM. The luciferase activity was assayed in cell extracts after treatment with the indicated combinations of ligands (0.1 nM, 1 nM, 10 nM, and 100 nM TTNPB and 1 µM SR11237) for various periods of time (3 h to 20 h) (Fig. 3A).
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To examine whether the status of RAR activation influences RXR/RAR cooperativity, we compared the reporter activity at 100 nM TTNPB to that observed at 0.1 nM TTNPB, a 1,000-fold-lower concentration than the optimal concentration of this ligand, upon coaddition of 1 µM SR11237. Results were expressed as the ratio of the luciferase activity measured with RAR and RXR ligands to that in the presence of RAR ligand alone, thereby reflecting the cooperativity between the two receptors (Fig. 3B). As shown above, a 20-h treatment of P2 cells with RXR and RAR ligands yielded a response inhibited by 15 to 20% compared to that with RAR ligand alone, reflected by a 0.8 cooperativity ratio. In contrast, 0.1 nM TTNPB allowed a cooperative response (factor, 1.8 to 2.0).
As expected, the transiently transfected promoter showed cooperativity when stimulated with 0.1 nM TTNPB and 1 µM SR11237, with a cooperativity ratio of about 2.2 to 2.5. A similar result was observed with 1 nM TTNPB (Lefebvre and Ozato, unpublished) and 100 nM TTNPB (Fig. 3B). These results indicate that the stably integrated promoter can mount cooperative activation by the two ligands only when the level of activation by RAR ligand is low and that cooperativity is lost when the promoter is strongly activated by RAR.
Cooperativity between RAR and RXR is sensitive to histone deacetylase inhibitors. Since the loss of cooperativity occurred only for the stably integrated or endogenous RARß promoters and thereby suggested an active role of chromatin in this process, we next tested the effect of TSA and sodium butyrate, two potent histone deacetylase inhibitors, on RAR/RXR cooperativity. Note that control experiments showed that histone deacetylase inhibitors did not modify luciferase mRNA and protein stabilities (P. Lefebvre and C. Brand, unpublished data). At low TTNPB concentrations (0.1 and 1 nM), TSA and sodium butyrate maintained the cooperativity between TTNPB and SR11237, and similar results were found in NIH 3T3 cells (Lefebvre and Ozato, unpublished). Interestingly, loss of cooperativity was counteracted by sodium butyrate and TSA, restoring a degree of cooperativity similar to that observed at early stages, suggesting an active role of an acetylation-deacetylation phenomenon in this process.
Next, we examined the kinetics of TSA action on this cooperative process (Fig. 3C). In the early stages, TTNPB alone yielded a low reporter activity (Fig. 3A) and TSA treatment did not modify the level of cooperativity (Fig. 3C). Subsequently, TSA treatment maintained the two-ligand cooperativity. Indeed, while treatment of P2 cells with 100 nM TTNPB and 1 µM SR11237 did not lead to cooperative activation after 20 h, TSA again allowed cooperative activation of the promoter in these conditions, as well as sodium butyrate (Lefebvre and Ozato, unpublished). These results indicated that histone deacetylase inhibitors prevented the loss of cooperativity that begins at a later stage of transcription, although it has no effect on ligand cooperativity that takes place at an early stage. Thus, loss of cooperativity is caused by a late-acting mechanism sensitive to histone deacetylase inhibitors.
Histone deacetylase-sensitive loss of cooperativity is independent of H3 and H4 acetylation status at the RARß promoter. The above results raised the possibility that the observed loss of cooperativity involved an histone acetylation-deacetylation mechanism. To address this possibility further, we next examined the histone acetylation status of the RARß promoter by using chromatin immunoprecipitation assays (Fig. 4A). Antibodies specific to acetylated histones H3 and H4 were used to immunoprecipitate formaldehyde-cross-linked, sonicated chromatin from P19 cells following the different ligand treatments. Quantitative PCR analysis of DNA input or bound to immunocomplexes were carried out to detect a fragment of 179 bp of the endogenous mRARß promoter (Fig. 4B). ChIP analysis with antibodies to acetylated H3 and acetylated H4 demonstrated a high constitutive H3 and H4 histone acetylation at the mRARß promoter in naive cells. Adding TTNPB (100 nM or 1 µM) alone or in the presence of SR11237 (1 µM) caused no significant changes in the histone acetylation status of the promoter after 30 min, and similar results were observed after longer treatment (20 h). Ro41-5353, an RAR antagonist, did not modify the H3 and H4 acetylation level (Lefebvre and Ozato, unpublished).
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Histone deacetylase inhibition prevents RNAP recruitment to transcribed regions. To better understand the molecular mechanism underlying the different steps in activation of the RARß promoter, we examined the engagement of RNA polymerase II (RNAP) using the ChIP assay with an antibody against the N terminus of the RNAP rpb1-encoded subunit, which recognizes this polypeptide irrespective of posttranslational modifications. As shown in Fig. 5, the ChIP assay revealed that a considerable amount of RNAP is constitutively associated with the RARß promoter in the absence of ligand. Adding 1 µM TTNPB alone for 15 min or longer did not significantly increase RNAP recruitment, showing that RNAP is already engaged on the promoter in basal conditions. Challenging cells with both TTNPB and SR11237 led to an increase in RNAP loading to the promoter only at a very short incubation time (15 min).
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We next asked how TSA affected RNAP loading at prolonged incubation times, conditions in which this histone deacetylase inhibitor restores the RXR-RAR cooperativity (see Fig. 3). TSA treatment considerably reduced RNAP association in the presence of TTNPB (10-fold decrease), which was further sensitive to SR11237 and increased by 2-fold. In contrast, RNAP loading to the promoter region was not affected in these conditions. These results indicated that the activation of the RARß promoter takes place following a two-step process. The first step is characterized by the low ability of TTNPB to promote RNAP engagement on the mRARß gene, a step during which SR11237 can generate a cooperative activation. The second and later stage, inhibited by histone deacetylase inhibitors, is characterized by the lack of cooperativity between RXR and RAR and by maximal engagement of RNAP on the mRARß gene.
Histone deacetylase inhibitor-mediated inhibition of the mRARß promoter is an RAR-controlled event. The inhibition of RNAP loading at the second stage of promoter activation by histone deacetylase inhibitors suggested that RAR is the molecular relay of a histone deacetylase inhibitor-controlled mechanism. We thus investigated whether RAR transcriptional activity could be altered by TSA. The mRARß promoter activity was monitored by real-time PCR detection of RARß or luciferase transcripts in P19 or P2 cells. P19 cells were first treated with increasing concentrations of TSA or of TSA and Am580, an RAR-specific ligand exhibiting properties strictly similar to atRA and TTNPB in this system. While TSA only weakly induced the basal transcriptional activity of the mRARß promoter at high concentrations (250 and 500 nM) and did not affect the response to rexinoids, Am580 promoted a very strong induction of the promoter activity. This RAR-mediated induction was inhibited in a dose-dependent manner by TSA, with a 50% inhibitory concentration (IC50) of about 10 to 50 nM.
The effect of histone deacetylase inhibition on transiently introduced templates and on stably integrated promoter activities (Fig. 6B through E) was monitored by assaying mRNA transcripts originating from each type of promoter. The transiently introduced reporter gene was strongly inducible by atRA (and TTNPB; data not shown), and TSA did not alter its responsiveness to an RAR-specific ligand (atRA) (Fig. 6B). In the same cellular background, we observed that the endogenous mRARß promoter activity was also highly inducible by atRA (Fig. 6C) as well as in P2 cells (Fig. 6D) but that, in contrast to its transiently transfected counterpart, it was strongly repressed by TSA. Similarly, the activity of the stably integrated chimeric reporter gene was highly sensitive to TSA-mediated inhibition (Fig. 6E).
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To assess this possibility, limited micrococcal nuclease digestion of P19 cell nuclei was carried out after or without TSA treatment and compared to the digestion pattern of the transiently introduced mRARß promoter-luciferase reporter gene (Fig. 6F). Interestingly, no nucleosomal ladder was detected in TSA-treated chromatin, suggesting a general unpacking of the nucleosomal array on the mRARß promoter.
A second RARE is required for inhibition of RXR-RAR cooperativity. To gain insight into the molecular basis of loss of cooperativity, we tested a series of mutant RARß reporters stably integrated into P19 cells (Fig. 7). A mutant reporter with an altered cAMP response element behaved in a manner similar to the wild type, as no cooperativity was observed after 20 h, which was fully restored upon histone deacetylase inhibition (Fig. 6A). A reporter with a mutation in the proximal RARE (DR5) failed to respond to either ligand alone or in cooperativity, underlining the predominant role of this response element in the regulation of mRARß promoter activity. More informative was a mutant reporter from which both the distal RARE and cAMP response element were deleted. This mutant displayed clear cooperativity in response to concomitant RXR and RAR stimulation. Since this mutant contains only one RARE, it was of importance to ascertain whether another promoter containing only one RARE also sustained cooperative activation in response to both types of ligands.
To this end, a synthetic reporter, (DR5)2 Ld40, consisting of a duplicated RARE connected to a basal promoter (a TATA box and initiator element) (Fig. 7A) with altered spacing compared to the mRARß promoter (22 versus 14 bp), was stably introduced into P19 cells. Promoter activity was tested following stimulation with RAR and/or RXR ligands. Responses obtained with this promoter paralleled those obtained with the RARß promoter: coaddition of 100 nM TTNPB and 1 µM SR11237 yielded no cooperative response (Fig. 7A), although cooperativity was restored upon histone deacetylase inhibition and observed at lower concentrations of TTNPB (Lefebvre and Ozato, unpublished).
The DR5 Ld40 promoter, containing only one consensus DR5 RARE, was then integrated into the P19 cell genome, and its activity was assayed as above. As observed for the RAREp mutant, this construct also displayed a cooperative response in response to RAR and RXR ligands (Fig. 7A). These results show that loss of cooperativity is not a special feature of the RARß promoter and requires at least two RAREs.
Finally, to ascertain whether loss of cooperativity is conditioned by chromatin assembly on the promoter, the activity of the synthetic reporters (DR5)2 Ld40 and DR5 Ld40 was tested in transient transfections. Again, results with these synthetic promoters paralleled those of the natural RARß promoter (Fig. 2B), as cotreatment with the two ligands led to a cooperative activation of transcription (Fig. 7B). These data thus indicate that loss of cooperativity occurs only in chromosomal promoters possessing more than one RARE, but not in those harboring a single RARE.
Increased coactivator recruitment and loss of cooperativity. In contrast to transiently introduced templates, chromosomal templates exhibited decreased activity in conditions thought to favor coactivator recruitment to the promoter. We thus examined the effect of stable overexpression of SRC-1, a member of the p160 coactivator family involved in the retinoid response (37). Western blot analysis confirmed that levels of SRC-1 expression were higher in these cells than in those transfected with the empty vector (Fig. 8A). In transient-cotransfection assays, SRC-1 displayed the expected coactivator activity, since it significantly enhanced retinoid-induced RARß reporter activity in response to TTNPB alone, as well as in response to TTNPB and SR11237 (Fig. 8B). A similar result was obtained with the RAREp construct, which harbors the proximal RARE as a unique cis-acting element.
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As a control for specific effect for multiple responsive elements, we next tested promoter activity with P19 cells stably transfected with the RAREp construct and the SRC-1 expression vector. No change in the cooperativity between RAR and RXR ligands was observed. Thus, increased expression of SRC-1 led to a negative regulation of the RARß promoter in the presence of an RXR ligand and required a tandem repeat of RARE.
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DISCUSSION |
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The idea that mRARß promoter engagement critically affects the extent of loss of cooperativity is supported by two sets of evidence. First, loss of cooperativity is not observed at suboptimal retinoid concentrations, and rexinoids cooperatively enhance transcript accumulation even beyond a 20-h stimulation for chromosomal reporters in these conditions. These results are consistent with previous reports showing a cooperative effect of retinoids and rexinoids on cellular differentiation and apoptosis only at subsaturating retinoid concentrations, emphasizing the biological importance of loss of cooperativity (13, 46, 48). Second, SRC-1 overexpression leads to an RXR-mediated negative regulation of the promoter. We also observed that activation of the cAMP-dependent protein kinase A pathway potentiated retinoid-induced transcription on transiently transfected templates. In contrast, cAMP elicited a stronger repression of mRARß promoter activity in the presence of both RXR and RAR ligands (cooperativity factor of 0.6 versus 0.8; Lefebvre and Ozato, unpublished). These data support the view that increased NCoA recruitment to the promoter (through either multiple RAREs, overexpression of nuclear coactivator A, or costimulation of several signaling pathways) is likely to be a triggering factor for loss of cooperativity.
Effects of histone deacetylase inhibitors on RARß transcriptional activation. Histone deacetylase inhibitor (TSA and sodium butyrate) effects are observed at the second stage of ligand activation, indicating that their effects are delayed and follow a kinetics similar to that of loss of cooperativity. Although these results suggested a link between histone acetylation and loss of cooperativity, H3 and H4 acetylation levels before and after ligand addition remained unchanged in the promoter vicinity, which is in contrast to results reported for other retinoid-regulated genes (14). However, the lack of histone hyperacetylation during remodeling and promoter activation has recently been shown for several yeast promoters, including PHO8 and HO (20).
The constitutive loading of RNAP on the mRARß promoter, which is also detected on transiently transfected templates (Lefebvre and Brand, unpublished), was unexpected considering previous studies documenting the absence of factor binding prior to ligand addition (15, 23). However, a lack of in vivo footprinting could reflect different binding stabilities that can be detected more readily by formaldehyde cross-linking. More importantly, a direct correlation between H3 and H4 hyperacetylation, transcriptional activity of the promoter, and RNAP loading was found in a region mapping to exon 3 of the RARß gene. Thus, RARß transcriptional activity is correlated to RNAP progression along the gene but not to its favored tethering to the promoter. This view is further substantiated by the correlation between increased progression of RNAP to exon 3 upon simultaneous and cooperative activation of RAR and RXR. Interestingly, RNA elongation on nucleosomal DNA is strongly influenced by histone acetylation, most notably by a decreased pause site residence time (55).
Cooperative activation was restored upon histone deacetylase inhibitor-mediated inhibition of RAR activity, with a measured IC50 in agreement with the reported IC50 of histone deacetylase 1, 4, or 6 by TSA in vitro (28, 38). While this observation does not easily fit with our current understanding of NR-mediated transcription, there are some precedents for our observations. The MMTV promoter again provides a very interesting point of comparison, since its glucocorticoid-induced activation is selectively inhibited by histone deacetylase inhibitors (6, 9). Of note is the occurrence of two functional glucocorticoid response elements in this viral promoter, which however responded biphasically to histone deacetylase inhibitors (6). The insensitivity of the endogenous or integrated mRARß promoters to histone deacetylase inhibitors in the early phase is in some aspects very similar to that of the CREB-regulated somatostatin gene (45). In this model, chromatin-specific regulation of CREB phosphorylation was found to occur at late stages of promoter activation, consistent with the view that the nucleosomal architecture limits access of activating protein kinase A to nucleosomal DNA-bound CREB.
It is still unclear how increasing the acetylation rate of a cellular component would lead to a partially decreased RAR activity, which depends on both template integration into the genome and the number of response elements. RAR could be a direct target for acetylases, but in vitro acetylation assays using purified RAR and RXR failed to reveal significant acetylation of these two NRs, in conditions where H2A, H2B, H3, and H4 are readily modified (Lefebvre and Brand, unpublished). Histone deacetylase inhibitors may also activate the MEK pathway (25), but no significant activation of the MEK/ERK pathway by histone deacetylase inhibitors was detected in P19 cells, and several protein kinase inhibitors, including PD98059, a specific inhibitor of MEK1 and MEK2 (1), were unable to reverse the histone deacetylase inhibitor effect on mRARß promoter activity (Lefebvre and Brand, unpublished).
Interestingly, acetylation of ACTR, an NR coactivator, negatively regulates its association with the estrogen receptor, providing a link between transcriptional attenuation and acetylation of components of the transcription machinery. However, this inactivation process was evidenced with a transiently transfected reporter gene (14). p300/CBP but not pCAF may acetylate ACTR and TIF2 in vitro, raising the possibility that acetylation or other posttranslational modifications occur as a function of the molecular composition of the cofactor complex. This raises the issue as to the possible role of chromatin assembly on the RARß promoter in dictating this molecular composition and the ability of RXR-RAR dimer to favor RNA elongation through hyperacetylated chromatin. Indeed, we observed that TSA is able to significantly decrease the quality of the nucleosomal array on the mRARß promoter, indicating an altered nucleosome accessibility and spacing. Along those lines, transcription from chromosomal MMTV templates is uniquely dependent on the interaction between the glucocorticoid receptor and the chromatin remodeling complex BRG1/BAF (27).
In summary, our work points to three important conclusions. First, we showed that maximal activation of the mRARß promoter, probably through increased tethering of coenzyme A to the promoter region, converts rexinoids into inactive states or even into being repressors of a chromatinized promoter activity. These results raise the possibility that this histone deacetylase-regulated negative control provides a means of ensuring highly gradual hormonal response and subsequent attenuated transcription. Second, our previous in vitro results showing that histone acetylation is required for RAR/RXR binding to a nucleosomal RARE (42) further suggest that receptors are constitutively bound to DNA in the P19 background. In support of these observations, ChIP experiments performed with anti-RXR or anti-RNAP antibodies showed constitutive binding of these proteins to the promoter in all conditions (P. Lefebvre, data not shown). Thus, histone deacetylase inhibitors are likely to affect a step downstream of transcription factor loading per se, and preliminary data showing that retinoids induce Ser 5 hyperphosphorylation of the RNAP C-terminal domain would implicate C-terminal domain kinases in this process. Third, it also emphasizes the general importance of studying each promoter as an integrated component in which transcriptional activation results from mutual interaction between the different transcription factors and is regulated by the chromatin environment.
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ACKNOWLEDGMENTS |
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Part of this work was financed by grants from INSERM, Association pour la Recherche sur le Cancer, and Ligue Nationale contre le Cancer (Comité du Nord-Pas de Calais).
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FOOTNOTES |
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Present address: Laboratoire de Dynamique Nucléaire et Plasticité du Génôme (UMR 218 du CNRS), Institut Curie/Section de Recherche, 75231 Paris Cedex 05, France.
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