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Molecular and Cellular Biology, December 2005, p. 10591-10603, Vol. 25, No. 23
0270-7306/05/$08.00+0     doi:10.1128/MCB.25.23.10591-10603.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.

Impaired Retinoic Acid (RA) Signal Leads to RARß2 Epigenetic Silencing and RA Resistance

MingQiang Ren, Silvia Pozzi, Gaia Bistulfi, Giulia Somenzi, Stefano Rossetti, and Nicoletta Sacchi*

Department of Cancer Genetics, Roswell Park Cancer Institute, Buffalo, New York 14263

Received 25 May 2005/ Returned for modification 13 July 2005/ Accepted 16 September 2005


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ABSTRACT
 
Resistance to the growth-inhibitory action of retinoic acid (RA), the bioactive derivative of vitamin A, is common in human tumors. One form of RA resistance has been associated with silencing and hypermethylation of the retinoic acid receptor ß2 gene (RARß2), an RA-regulated tumor suppressor gene. The presence of an epigenetically silent RARß2 correlates with lack of the RA receptor {alpha} (RAR{alpha}). Normally, RAR{alpha} regulates RARß2 transcription by mediating dynamic changes of RARß2 chromatin in the presence and absence of RA. Here we show that interfering with RA signal through RAR{alpha} (which was achieved by use of a dominant-negative RAR{alpha}, by downregulation of RAR{alpha} by RNA interference, and by use of RAR{alpha} antagonists) induces an exacerbation of the repressed chromatin status of RARß2 and leads to RARß2 transcriptional silencing. Further, we demonstrate that RARß2 silencing causes resistance to the growth-inhibitory effect of RA. Apparently, RARß2 silencing can also occur in the absence of DNA methylation. Conversely, we demonstrate that restoration of RA signal at a silent RARß2 through RAR{alpha} leads to RARß2 reactivation. This report provides proof of principle that RARß2 silencing and RA resistance are consequent to an impaired integration of RA signal at RARß2 chromatin.


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INTRODUCTION
 
Cells of different histotypes seem prone to lose the ability to respond to the growth-inhibitory action of retinoic acid (RA), the potent bioactive derivative of vitamin A. RA regulates fundamental cellular processes, such as growth, differentiation, and apoptosis (7). Previously, we and others showed a correlation between a common form of RA resistance and repressive epigenetic changes (at both the histone and DNA levels) in the RA receptor ß2 gene (RARß2) (5, 33, 34, 40).

RARß2 is an RA-regulated tumor suppressor gene (19, 26, 32). Detection of aberrant RARß2 methylation in tumors of different histotypes raised the question of whether this epigenetic change is critical for silencing this tumor suppressor gene. Previously, we proposed that aberrant RARß2 inactivity might induce repressive epigenetic changes at RARß2, leading to RARß2 silencing and RA resistance (33, 34). RARß2 transcription is normally regulated by dynamic histone changes in the presence and absence of RA (9, 14, 29, 41). Therefore, we hypothesized that the impaired integration of RA signal at RARß2 can create a state of exacerbated-protracted RARß2 transcriptional inactivity and attract chromatin-repressive changes, including DNA methylation. The conversion of RARß2 from a state permissive for transcription into a stable state nonpermissive for transcription would cause biological RA resistance. Our hypothesis hinges on the original supposition of Ng and Bird (28) that chromatin inactivity, the prerequisite for epigenetic silencing of genes on chromosome X (18), could also lead to silencing of genes on other chromosomes. Thus, an aberrant inactive RARß2 chromatin status would be the prerequisite for RARß2 epigenetic silencing.

RARß2 DNA methylation and silencing were shown to be induced by active recruitment of repressor proteins by an oncogenic fusion protein in leukemic cells (13). However, to our knowledge, this oncoprotein has not been demonstrated in epithelial cancer cells and tumors of the breast, prostate, colon, lung, and head and neck, where RARß2 has also been found silenced (33, 34). In contrast, cancer epithelial cells and tumors appear to have either a low intracellular concentration of RA or a lack or derangement of proteins involved in either RA metabolism and homeostasis or RARß2 transcriptional regulation. Thus, RA resistance might be the consequence of an exacerbated-protracted RARß2 transcriptional repression caused by a defective integration of RA signal at RARß2, which might be induced by genetic, epigenetic, metabolic, and environmental factors capable of shutting off the "communication" between RA and RARß2 chromatin.

We identified and tested as a possible cause of aberrant RARß2 inactivity the lack of functional RAR{alpha}, the upper regulator of RARß2 transcription. RAR{alpha} has the role of keeping the chromatin of its direct target genes, such as RARß2, poised for transcription yet inactive. Upon binding of RA to RAR{alpha}, the chromatin status of the target genes is converted from inactive into active because of the exchange of corepressor complexes with coactivator complexes, which would rapidly induce histone changes, chromatin remodeling, and transcription activation (14, 29).

In the course of our studies of RA-resistant breast and prostate cancer cell lines, we observed the following: (i) the presence of low or negligible binding of RAR{alpha} at RARß2 in RA-resistant breast and prostate cancer epithelial cells carrying RARß2 nonpermissive alleles (we define as nonpermissive the alleles that cannot be transcriptionally activated by RA and as permissive the alleles that are poised for transcription yet inactive in the absence of RA but capable of transcription in the presence of RA); (ii) the presence of RARß2 unmethylated (U), permissive alleles in RAR{alpha}-positive cells, which contain many other methylated (M) genes (20), pointing at RAR{alpha} as a critical factor that can spare RARß2 chromatin from falling into a nonpermissive status; and (iii) the presence of a minimal stretch of methylated CpGs in the first RARß2 exon—corresponding to exon 5 of the RARß locus (38)—in methylated alleles, suggesting that CpG methylation originates in a specific epicenter from unmethylated yet nonpermissive alleles.

In this study we simulated possible genetic, epigenetic, and metabolic scenarios that could impair the flow of RA signal at RARß2 chromatin via RAR{alpha}. Using three different strategies—a dominant-negative RAR{alpha} lacking the RA-binding domain, downregulation of RAR{alpha} by RNA interference, and RA antagonists acting specifically at RAR{alpha}—we induced the conversion of RARß2 permissive alleles into nonpermissive alleles in RA-sensitive human cells. The RARß2 nonpermissive alleles developed a significant load of repressive histone tail modifications and failed to recruit RNA polymerase II at the region containing the transcription start site. Only a percentage of nonpermissive alleles developed CpG hypermethylation, thus showing that aberrant hypermethylation is not an absolute requirement for RARß2 silencing. In this report we also demonstrate that restoring RA signal through RAR{alpha} at an epigenetically silent RARß2 is critical to reestablishing a RARß2 status compatible with transcription. RARß2 epigenetic silencing has been described as being associated with an RA-resistant phenotype (26, 32). Here we prove that RA resistance is the consequence of RARß2 epigenetic silencing.


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MATERIALS AND METHODS
 
Cell lines. Breast and prostate cancer cell lines (ATCC, Manassas, VA) and COS-1 cells (ATCC) were grown using standard protocols.

Drugs. All-trans-RA, 5-aza-2'-deoxycytidine (5-Aza), a demethylating agent (10), and trichostatin A (TSA), a histone deacetylase inhibitor (42), were all from Sigma (St. Louis, MO). These drugs were dissolved and stored as described previously (34). The RAR{alpha} antagonist ER50891 was a kind gift of Kouichi Kikuchi, Discovery Research Laboratories, Ibaraki, Japan, and the RAR{alpha} antagonist RO414253 was a kind gift from Salvatore Toma, National Cancer Institute, Genoa, Italy. Treatments with these drugs were all performed in medium supplemented with 5% charcoal-dextran-stripped serum.

Colony formation assay. Exponentially growing cells were seeded at 5 x 102 cells/well in six-well plates in triplicate and allowed to attach to the substrate. Cells were left untreated or treated with drugs for 24 h, and then the medium was replaced with drug-free medium and the cells were grown for 14 to 21 days. Colonies were fixed with methanol, stained with Giemsa (Sigma), and counted to establish the colony formation index as described previously (34). The statistical significance was calculated by Student's t test for three independent experiments.

Cell transfections. The RAR{alpha} dominant-negative mutant RAR{alpha}403 was subcloned by PCR into the FLAG-containing pCMV-tag vector (Stratagene, La Jolla, CA) with primers introducing EcoRI and XhoI restriction sites (sense, 5'-TATGAATTCATGGCCAGCAACAGCAGCTC-3'; antisense, 5'-ATACTCGAGGGGATCTCCATCTTCAGCGT-3'), and the empty pCMV-tag vector was transfected in T47D by using Lipofectamine Plus (Invitrogen, Carlsbad, CA). The LNasRARß2VI vector, which harbors six copies of RARß2 antisense (asRARß2) and the control empty vector LNSX (kindly provided by S. Y. Sun, University of Texas M.D. Anderson Cancer Center, Houston, TX) (37) were also transfected in T47D by Lipofectamine Plus. Stable clones were selected and maintained with G418 (Invitrogen) at 0.8 mg/ml. Stable MDA-MB-231 clones overexpressing RAR{alpha} 1 were obtained by cotransfecting cells with pSG5-hRAR{alpha} 1 vector (kindly provided by Fausto Andreola, National Cancer Institute, Bethesda, MD) and G418-resistant pcDNA3.1(+) vector (Invitrogen) by use of Lipofectamine Plus. Control cells were cotransfected with the empty vector pSG5 (Promega, Madison, WI) and pcDNA3.1(+). Stably transfected cells, selected with increasing concentrations of G418 (0.5 to 2.5 mg/ml), were tested for expression of exogenous RAR{alpha} 1 by Western blotting with the C-20 anti-RAR{alpha} antibody (Santa Cruz Biotechnology).

Retroviral infections. Supernatants containing either the RAR{alpha} dominant-negative LXRAR{alpha}403SN or the empty LXSN retroviral particles (kindly provided by Fausto Andreola, National Cancer Institute, Bethesda, MD) were used to infect T47D cells in the presence of 4 mg/ml Polybrene (Sigma) as described previously (39). Infected cells were selected with 0.8 mg/ml G418. Single clones were isolated after 14 to 21 days and screened for the presence of either the LXSN or the LXRAR{alpha}403SN construct by reverse transcription-PCR (RT-PCR). Positive clones were maintained in 0.8 mg/ml G418.

RAR{alpha} RNA interference. The 19-nucleotide sequence (AGCGCACCAGGAAACCTTC) corresponding to nucleotides 680 to 699 of RAR{alpha} exon 5 (GenBank accession no. NM_000964) was inserted into pSUPER-retro (OligoEngine, Seattle, WA) according to the manufacturer's instructions. The silencing efficiency of the resulting construct, pSUPER-RAR{alpha}, was first tested on exogenous RAR{alpha} by transiently cotransfecting COS-1 cells with pSG5-hRAR{alpha}1, encoding the human RAR{alpha}1 and pSUPER-RAR{alpha} at different ratios. Exogenous RAR{alpha} levels, normalized to GAPD (glyceraldehyde-3-phosphate dehydrogenase) expression levels, were estimated by Western blotting with the C-20 anti-RAR{alpha} and anti-GAPD (Santa Cruz Biotechnology, Santa Cruz, CA) antibodies and appropriate horseradish peroxidase-conjugated secondary antibodies (Santa Cruz Biotechnology; Amersham, Piscataway, NJ). Stable transfections with pSUPER-RAR{alpha} and pSUPER-retro (control) were performed with Lipofectamine Plus. Transfected T47D clones were selected with 2 µg/ml puromycin (Sigma). Clones displaying significant silencing of RAR{alpha} by real-time RT-PCR and Western blotting were chosen for further analysis.

RAR{alpha} antagonist experiments. Cells (5 x 102) were seeded in six-well plates in triplicate in medium containing 5% charcoal-dextran-stripped serum, allowed to attach to the plastic substrate, and treated with ER50891 (10 µM) and RA (0.1 µM), either alone or in combination, for 24 h. After this time, the medium was replaced by drug-free medium. Cells were grown until colonies became visible (14 to 21 days). Colonies were stained with Giemsa to establish the colony formation index (34). Pools of colonies derived from cells that survived each treatment (125 colonies from one well with no drugs, 118 clones from one well treated with ER50891 alone, and 115 clones from one well treated with ER50891 plus RA) were used to isolate genomic DNA that was used for methylation-specific PCR (MSP) analysis with M4 primers and U4 primers. In a parallel replica experiment, we instead used cloning cylinders to isolate six independent clones that survived the treatment with either ER50891 alone (which we designated ER clones) or ER50891 in combination with RA (which we designated ER/RA clones). All clones from each group that were expanded in drug-free medium did not show RA-induced RARß2 transcription by real-time RT-PCR and were shown to be U or M by MSP. One clone from each of the groups ER-C6 and ER/RA-C5 was further expanded in drug-free medium.

Real-time RT-PCR. Total RNA was obtained using Trizol (Invitrogen), treated with DNase I (Ambion, Austin, TX), retrotranscribed with a SuperScript first-strand synthesis system (Invitrogen), and amplified by real-time RT-PCR on an iCycler apparatus (Bio-Rad, Hercules, CA) by using iQ SYBR green Supermix (Bio-Rad) and specific primers for RARß2 (sense, 5'-GACTGTATGGATGTTCTGTCAG-3'; antisense, 5'-ATTTGTCCTGGCAGACGAAGCA-3'), RAR{alpha} (both isoforms 1 and 2) (sense, 5'-TGTGGACTTCGCCAAGCA-3'; antisense, 5'-CGTGTACCGCGTGCAGA-3'), RAR{alpha} 1 (sense, 5'-GCCAGGCGCTCTGACCACTC-3'; antisense, 5'-CAGGCGCTGACCCCATAGTGGT-3'); and GAPD (sense, 5'-GAAGGTGAAGGTCGGAGTC-3'; antisense, 5'-GAAGATGGTGATGGGATTTC-3'), with the appropriate annealing temperature according to standard protocols. The RAR{alpha} and RARß2 transcription levels were normalized to the GAPD transcription level. The statistical significance was calculated using Student's t test for three independent determinations.

Luciferase assay. The luciferase reporter assay was performed essentially as described previously using the luciferase reporter vector RARß2-pGL2 (12) (kindly donated by K. Ozato, National Institutes of Health, Bethesda, MD) and the control vector pRL-TK (Promega). Vector DNAs were cotransfected with Lipofectamine Plus in cells grown in a 12-well plate; 24 h after transfection, the medium was replaced with medium with or without RA (1.0 µM). Luciferase activity was measured by a dual luciferase reporter assay system (Promega) according to the manufacturer's instructions. The values represent the averages (normalized to the control) of three independent experiments, each performed in triplicate.

ChIP. Quantitative chromatin immunoprecipitation (ChIP) was performed using reagents purchased from Upstate (Lake Placid, NY) following the manufacturer's protocol. Chromatin was immunoprecipitated with antibodies against acetyl-histone H4 (Ac-H4) (Upstate) (1:400), acetyl-lysine (K) 9 histone H3 (H3-Ac-K9) (Upstate) (1:400), dimethyl-K4 histone H3 (H3-Me-K4) (Upstate) (1:400), trimethyl-K9 histone H3 (H3-Tri-Me-K9) (Upstate) (1:200), RAR{alpha} C terminus (Santa Cruz Biotechnology) (1:60), RAR{alpha} N terminus (Biolegend, San Diego, CA) (1:100), RARß (Santa Cruz Biotechnology [1:70]; Active Motif, Carlsbad, CA [1:600]), RNA polymerase II (Upstate) (1:200), and FLAG epitope (Sigma) (1:350). Re-ChIP was performed with the anti-RAR{alpha} C-terminus antibody on chromatin immunoprecipitated with an anti-Ac-H4 antibody after elution with re-ChIP elution buffer (10 mM EDTA, 50 mM Tris-HCl [pH 8.0], 0.7 M NaCl, 20 mM dithiothreitol). Control ChIPs were without the respective antibodies. The immunoprecipitated DNA was amplified by real-time PCR with either RARß2 primers (sense, 5'-GGTTCACCGAAAGTTCACTCGCAT-3'; antisense, 5'-CAGGCTTGCTCGGCCAATCCA-3') or the GAPD primers (sense, 5'-GGTGCGTGCCCAGTTGAACCA-3'; antisense, 5'-AAAGAAGATGCGGCTGACTGTCGAA-3'). The RARß2 DNA relative enrichment was calculated by normalizing the RARß2 PCR signal to the PCR signals obtained both from the input DNA (total chromatin fraction) and the GAPD DNA. Statistical significance was determined using the Student's t test on three independent determinations.

DNA methylation analysis. Genomic DNA extracted with DNAzol (Invitrogen) was modified with sodium bisulfite as described previously (33). Modified DNA was used for both MSP and sequencing analyses. For sequencing analysis, a 635-bp region, encompassing 27 RARß2 CpG sites in the RARß2-regulatory region, was amplified by nested PCR (40) using first the primer set RARß2 sense 1 (5'-GTATAGAGGAATTTAAAGTGTGGGTTGGG-3') and RARß2 antisense 1 (5'-CCTATAATTAATCCAAATAATCATTTACC-3') and subsequently the primer set RARß2 sense 2 (5'-GTAGG(C/T)GGAATATTGTTTTTTAAGTTAAG-3') and RARß2 antisense 2 (5'-AATCATTTACCATTTTCCAAACTTACTC-3'). The PCR products were either directly sequenced or sequenced after subcloning in the pCR4-TOPO plasmid vector (Invitrogen). In the latter case, we sequenced a minimum of 20 to a maximum of 50 clones corresponding to the sample specified in Results. MSP was performed by amplifying bisulfite-modified DNA with the first RARß2 primer set (see above). Then, 2 µl of a 1:1,000 dilution of the first amplification product was reamplified with the U4 sense- and U4 antisense-specific primers or with the M4 sense- and M4 antisense-specific primers (33) or with the different combinations of U4 and M4 primers. RARß2 alleles detected with both U4 primers were classified as U, and the alleles amplified with either both M4 primers (these products are the ones discussed in Results) or the sense M4 primer and the U4 antisense primer were classified as M alleles. No product was amplified with the U4 sense primer and the M4 antisense primer.


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RESULTS
 
Lack of RAR{alpha} correlates with RARß2 epigenetic silencing in RA-resistant cancer cells. Analysis of a panel of breast and prostate cell lines showed different RA-induced RARß2 transcription levels in correlation with differential growth inhibition by RA after 24 h of treatment (Fig. 1A, top and bottom, respectively). DNA methylation analysis using two complementary techniques, bisulfite sequencing and MSP, identified cell lines (i) homozygous for RARß2 M alleles (MCF7, MDA-MB-231, and LNCaP carry 100% of M alleles), (ii) homozygous for RARß2 U alleles (T47D carries 100% of U alleles), and (iii) heterozygous for U and M alleles (DU145) (Fig. 1B). As determined on the basis of sequencing of 20 independent DU145 RARß2 alleles, 30% were U alleles and 70% were M alleles. The distribution of methylated CpGs in the M alleles of all cell lines involved at least a common stretch of CpGs in the first RARß2 exon (Fig. 1B, bottom). Lack of RA-induced RARß2 transcription was also detected in cells (DU145) with U alleles, which must therefore be interpreted as nonpermissive. Moreover, RARß2 was spared by methylation in T47D, a cell line with several other hypermethylated genes (20).



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  		FIG. 1. Lack of RAR{alpha} correlates with RARß2 epigenetic silencing in RA-resistant cells. (A) Real-time RT-PCR (top) and colony formation assay (bottom) on a panel of breast and prostate cell lines showed a positive correlation between RA-induced RARß2 transcription and RA-induced growth inhibition. (B) Methylation-specific PCR (MSP, top) and bisulfite sequencing (bottom) show the presence of cells homozygous for RARß2 M alleles (MCF-7, LNCaP, and MDA-MB-231), heterozygous for U and M alleles (DU145), and homozygous for U alleles (T47D) and the localization of methylated CpGs in M alleles. (C) ChIP analysis shows a significant increase in the levels of both histone Ac-H4 (P < 0.05) and H3-Ac-K9 (P < 0.01) in T47D RARß2 chromatin but not in DU145 in response to RA. The levels of H3-Tri-Me-K9 and H3-Me-K4 were significantly higher (P < 0.01) and lower (P < 0.01), respectively, in DU145 relative to T47D. The dotted lines represent what we defined as "the histone modification threshold" required for RARß2 transcription. (D) Schematic diagram showing that RAR{alpha} binds the RARß2 RARE region in the absence of RA and that transcription is induced in the presence of RA (top) (14). RAR{alpha} transcription (bottom left) and binding of RAR{alpha} protein at RARß2 (bottom right) are significantly higher (P < 0.05) in cells with RARß2-permissive alleles (T47D) than in cells with RARß2 nonpermissive alleles (DU145 and MDA-MB-231).

Analysis of the chromatin associated with RARß2 alleles enabled us to quantitate the level of a few histone modifications—considered a hallmark of either repressive or active chromatin (4, 22)—in the RARß2 region encompassing the RA-responsive element (RARE), the TATA box, the transcription start site, and the common stretch of CpGs in the first exon. We detected significant differences in the levels of acetylation of both histone H4 (Ac-H4) and histone H3 at lysine (K) 9 (H3-Ac-K9) in response to RA treatment for 24 h in T47D RARß2 chromatin but not in DU145 chromatin (Fig. 1C, top left and bottom left). Moreover, both in the absence (baseline) and in the presence of RA, the level of methylation of histone H3 on K4 (H3-Me-K4)—a parameter of chromatin activity—was higher in T47D than in DU145 chromatin (Fig. 1C, bottom right), while the opposite was true for H3-K9 methylation (H3-Tri-Me-K9)—a parameter of chromatin inactivity (Fig. 1C, top right). Thus, the overall quantity of histone modifications, which we define as the "histone modification threshold" (Fig. 1C, dotted lines), as well as the quality of histone modifications, seems to be critical in order to "switch on" RARß2 transcription in response to RA. RA induced both qualitative and quantitative modifications in nonpermissive alleles as well but, apparently, unless a critical accumulation of histone changes hit the threshold, the transcription remained "off."

Normally, RAR{alpha} bound to RARß2 keeps the chromatin poised for transcription (yet inactive) (14) (Fig. 1D, top left) and ready for activation by RA (Fig. 1D, top right). RA binding to RAR{alpha} would induce histone modifications capable of activating RARß2 very rapidly (29). Once RARß2 is induced, it could regulate its own transcription (8). We observed that the presence of RARß2 nonpermissive alleles (regardless of their DNA methylation status) always correlated with a lower level of RAR{alpha} transcription (Fig. 1D, bottom left). Consistent with this observation, quantitative ChIP with anti-RAR{alpha} antibodies, followed by RARß2 DNA amplification of the region encompassing the RARE (schematic diagram in Fig. 1B), detected significantly more RAR{alpha} associated with RARß2 in T47D cells than in MDA-MB-231 and DU145 cells (Fig. 1D, bottom right). Of note, by ChIP with anti-RARß antibodies we found RARß2 binding only at the T47D RARß2 promoter (data not shown).

Altogether, these observations made us hypothesize that a defective integration of RA signal at RARß2 due to lack of functional RAR{alpha} can convert RARß2 into inactivity, marked by repressive epigenetic changes at histone and DNA level, and RA resistance.

Induction of RARß2 epigenetic silencing by a dominant-negative RAR{alpha} in RA-sensitive cells. First, we simulated a genetic scenario whereby an RAR{alpha} mutation with dominant-negative features occurs in RA-sensitive, RAR{alpha}-positive cells homozygous for RARß2-permissive alleles. We used a well-characterized dominant-negative RAR{alpha} mutant, RAR{alpha}403, which lacks the C-terminal RA-binding domain but retains the capacity to heterodimerize with the retinoid X receptor and bind to the RARE regions (Fig. 2A) (11, 15, 39). RAR{alpha} 403 should compete with wild-type RAR{alpha} in the heterodimerization with retinoid X receptor (23). It might also cause an impaired turnover of corepressor complexes at RAR{alpha} target genes, because it lacks the C terminus (27, 29).



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  		FIG. 2. Functional RAR{alpha} inhibition by a dominant-negative RAR{alpha} in RA-sensitive cells leads to RARß2 silencing marked by repressive epigenetic changes. (A) Schematic diagram representing the structure of the RAR{alpha}-dominant-negative RAR{alpha}403 (bottom) missing part of the RA-binding domain. (B) The FLAG-tagged RAR{alpha}403, transiently transfected into T47D cells, binds the RARß2 promoter concomitant with a significant (P < 0.05) decrease of RA-induced RARß2 transcription (middle). ChIP experiments with antibodies (Ab) directed against the RAR{alpha} C and N termini, showing that the C-Ab, but not the N-Ab, detects remarkably less wild-type RAR{alpha} at RARß2 in DN C8 cells, carrying both the dominant-negative and wild-type RAR{alpha} isoforms, than LX C5 cells carrying only wild-type RAR{alpha} (right). (C) In contrast to LX C5, DN C8 does not significantly express both luciferase from an exogenous RARß2 promoter (P < 0.05) (left) and endogenous RARß2 (P < 0.05) (middle) in response to the presence of RA, consistent with lack of recruitment of RNA polymerase II at the promoter (right). (D) RARß2 ChIP analysis showing that RA significantly (P < 0.01) increases the level of activating histone modifications (Ac-H4, H3-K9-Ac, and H3-Me-K4) to the "threshold" (dotted line) associated with transcription activation (on) only in LX C5 chromatin and not in DN C8 chromatin. Consistently, the level of H3-Tri-Me-K9 is higher in DN C8 chromatin than in LX C5 chromatin. (E) MSP (left) and bisulfite sequencing (right) show that DN C8 is heterozygous for U and M RARß2 alleles, with the methylated CpGs mostly localized in exon 1.

At 24 h after transient transfection with a FLAG-RAR{alpha}403 construct, T47D breast cancer cells (which express mainly endogenous RAR{alpha}1; data not shown) showed the presence of the dominant-negative mutant at the RARß2 promoter (ChIP; Fig. 2B, bottom left), concomitant with significant RARß2 transcriptional downregulation in response to the presence of RA (Fig. 2B, bottom middle). Thus, RARß2 silencing seems to be initiated when RAR{alpha}403 resides at the RARß2 promoter region. The dominant-negative RAR{alpha} was also found at the RARß2 of a T47D clone expressing the dominant-negative protein (DN C8). This was deduced on the basis of a ChIP experiment with an antibody directed against the RAR{alpha} C terminus; this antibody detected remarkably less wild-type RAR{alpha} at RARß2 than an antibody directed against the RAR{alpha} N terminus in DN C8 relative to LX C5 (Fig. 2B, bottom right). RA treatment (24 h) failed to induce luciferase transcription from a transiently transfected RARß2 promoter-luciferase construct (Fig. 2C, left), thus proving the dominant-negative effect of RAR{alpha}403 on endogenous RAR{alpha}. Finally, in response to the presence of RA, DN C8 did not show endogenous RARß2 transcription (real-time RT-PCR; Fig. 2C, middle). This finding was mirrored by a lack of RNA polymerase II at RARß2 (Fig. 2C, right). The repressed transcriptional status in the DN C8 clone was paralleled by repressive quantitative and qualitative histone modifications, which, in response to RA treatment, did not reach the threshold necessary for "switching on" RARß2 transcription (Fig. 2D; dotted line). Interestingly, MSP analysis of DN C8 RARß2 DNA with M4 and U4 primers (see Materials and Methods) showed U and M alleles. After bisulfite sequencing of 50 alleles, we found that DN C8 contained 60% U and 40% M alleles with a few methylated CpGs (Fig. 2E). In contrast to what was reported for cells carrying another dominant-negative RAR{alpha} with an intact RA-binding domain (13), RA treatment (1 µM, 72 h) did not reverse RARß2 DNA methylation. Only treatment with 5-Aza (0.8 µM for 72 h) led to demethylation and RA-induced RARß2 transcription in DN C8 (data not shown).

RARß2 epigenetic silencing confers an RA-resistant phenotype. Concomitant with RARß2 epigenetic silencing, DN C8 cells developed stable resistance to the growth-inhibitory effect of RA, as assessed by colony formation (Fig. 3A). Because RAR{alpha} controls several RA-responsive target genes, we tested whether the observed RA resistance phenotype was indeed caused by RARß2 silencing. To this end we targeted T47D RARß2 transcription with an antisense RARß2 (37). Stable T47D clones transfected with LNasRARß2VI that carry multiple copies of the RARß2 antisense, like the As-C3 and As-C4 clones (where "As" represents "antisense") (Fig. 3B, left), were significantly less sensitive to RA-induced growth inhibition than the cognate control clone EV-C3 carrying the empty vector LNSX (Fig. 3B, right). Thus, it is conceivable that the RA-resistant phenotype developed by DN C8 cells is the consequence of RARß2 epigenetic silencing.



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  		FIG. 3. RARß2 epigenetic silencing confers an RA-resistant phenotype. (A) Colony formation assay showing that DN C8 cells, but not LX C5 cells, display RA resistance. (B) The RARß2 antisense construct LNasRARß2VI (top left) is detected in the T47D As-C3 and As-C4 clones, as shown by PCR (bottom left). Colony formation assay showing that As-C3 and As-C4, but not the control EV-C3 clone carrying the empty vector, display RA resistance (right).

RARß2 DNA methylation is not necessary to confer a nonpermissive transcriptional status. Because RA did not induce transcription from both RARß2-U and RARß2-M nonpermissive alleles in DN C8 cells, we hypothesized the existence of RA-resistant DN C8 cells homozygous for nonpermissive unmethylated alleles. We isolated 20 independent DN C8 subclones which were resistant to RA and unable to reexpress the RARß2 transcript in response to RA. Screening for the presence of methylation in the first exon by MSP identified 18 clones heterozygous for both U and M alleles and 2 clones homozygous for U alleles (Fig. 4A). The CpG methylation-free status of the U alleles was confirmed by bisulfite sequencing (20 alleles of one of the two subclones are shown in Fig. 4B). The level of Ac-H4 did not increase significantly in response to RA in the RARß2 chromatin of this subclone as it did in the RARß2 chromatin of LX C5 (Fig. 4C, top) that is instead homozygous for RARß2 U alleles that are permissive (Fig. 2E, top). The level of H3-Tri-Me-K9 in the DN C8 subclone chromatin was in the range observed in the parental DN C8 clone (Fig. 4C, bottom). Apparently, RARß2 silencing also can be imposed in the absence of DNA methylation.



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  		FIG. 4. RARß2 DNA methylation is not necessary to confer a nonpermissive RARß2 transcriptional status. (A) RARß2 methylation and transcription in 20 DN C8 subclones shows that 2 clones do not present CpG methylation and RA-induced RARß2 transcription. (B) MSP and bisulfite sequencing analysis of one of the two DN C8 subclones homozygous for U alleles. (C) ChIP analysis shows that the chromatin of the DN C8 subclone homozygous for U alleles and the parental DN C8 is marked by comparable levels of Ac-H4 (top) and H3-Tri-Me-K9 (bottom) in the presence and absence of RA.

Knocking down RAR{alpha} by RNA interference also triggers RARß2 epigenetic silencing and RA resistance. It can be argued that RARß2 silencing in DN C8 cells is due to the recruitment of histone-modifying enzymes or DNA methyltransferases (DNMTs) at RARß2 by the RAR{alpha}403 protein, as shown for the PML-RAR{alpha} protein (13). While nonrandom dominant-negative RAR{alpha} mutations were never to our knowledge reported in cancer epithelial cells, RAR{alpha} expression was reported to be low or absent in these cells (30). This could be due to loss of heterozygosity or RAR{alpha} epigenetic silencing or both. Thus, we tested the effect of RAR{alpha} knockdown on RARß2 transcription by using RAR{alpha}-specific RNA interference in T47D cells.

Stable expression in T47D cells of a short hairpin RNA (targeting a sequence common to both RAR{alpha}1 and RAR{alpha}2; Fig. 5A, top left), selected for efficient knockdown of exogenous RAR{alpha}1 overexpressed into COS cells (Fig. 5A, bottom left), silenced the endogenous RAR{alpha} in prototypic clones such as SI{alpha} C7 (Fig. 5A, top right). As a result, RA-induced RARß2 transcription (Fig. 5A, bottom right) was abrogated in association with development of a RARß2 chromatin status unable to integrate the RA signal (Fig. 5B). Indeed, RA failed to induce the level of both Ac-H4 and H3-Ac-K9 up to the threshold (Fig. 5B, left and middle; dotted line) associated with RARß2 transcription. Moreover, SI{alpha} C7 chromatin showed a significantly higher level of H3-Tri-Me-K9 than the control pSC2 chromatin (Fig. 5B, right). MSP analysis showed the presence of M alleles. Bisulfite sequencing of 25 alleles evidenced, as it did in DN C8, either U alleles or M alleles methylated in just a few CpGs (Fig. 5C). Surprisingly, profound RAR{alpha} downregulation seems sufficient to create a stable nonpermissive RARß2 status, apparently marked by an accumulation of histone-repressive changes but not always by CpG methylation. We do not know yet how lack of RAR{alpha} makes the RARß2-chromatin a "prey" of repressor proteins. Also, in this case, when RARß2 falls into silencing, cells apparently acquire resistance to RA-induced growth inhibition (Fig. 5D).



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  		FIG. 5. Stable RNA interference of RAR{alpha} triggers RARß2 epigenetic silencing and RA resistance. (A) The RAR{alpha} targeting sequence (top left), cloned in the pSUPER-retro vector and selected for efficient knockdown of exogenous RAR{alpha} (encoded by pSG-hRAR{alpha} 1) in COS cells (bottom left), efficiently knocks down the endogenous RAR{alpha} protein in the T47D clone SI{alpha} C7 (top right) consistent with significant transcriptional downregulation (P < 0.01) (bottom right). (B) Quantitative ChIP analysis showing that RA induces in SI{alpha} C7 RARß2 chromatin significant lower levels of both Ac-H4 (P < 0.05) (left) and H3-Ac-K9 (P < 0.05) (middle) than in pS C2 chromatin, well below the threshold (dotted line) required for transcription. The level of H3-Tri-Me-K9 (right) is also significantly higher (P < 0.05) in SI{alpha} C7 RARß2 chromatin than in pS C2 chromatin. (C) MSP (right) and bisulfite sequencing (left) show CpG hypermethylation in a few SI{alpha} C7 RARß2 alleles and the localization of the CpGs. (D) Colony formation assay showing that SI{alpha} C7 cells are RA resistant.

Induction of RARß2 epigenetic silencing by RAR{alpha} antagonists. It was reported that the intracellular RA level is lower in cancer epithelial cells than in their normal counterparts (17). Thus, we set out to test whether interfering with RA availability at endogenous RAR{alpha} with RAR{alpha} antagonists can force RARß2 into a deep state of inactivity.

First we tested that the RAR{alpha}-specific antagonist ER50891 (25) (Fig. 6A, top) was able to significantly inhibit the induction of RARß2 transcription in response to RA in a time course experiment in which T47D cells were grown in the presence of ER50891 (10 µM) alone, RA (0.1 µM) alone, or ER50891 and RA in combination for 4, 8, and 24 h (Fig. 6A, bottom). By using ChIP analysis with a RAR{alpha}-specific antibody we observed that the levels of endogenous RAR{alpha} occupancy of the RARß2 region containing the RARE did not differ significantly during the 24 h in cells grown in the presence of the antagonist (Fig. 6B, top). In contrast, we observed that in the same cell samples the levels of H3-Tri-Me-K9 (hallmark of repressive chromatin) and H3-Ac-K9 (hallmark of active chromatin) associated with RARß2 DNA increased and decreased, respectively (Fig. 6B, bottom two panels). We also detected by both MSP analysis and bisulfite sequencing the appearance of M alleles (Fig. 6C, top), with methylation emerging once again in a few CpGs in the first exon (Fig. 6C). This experiment suggests that upon the RAR{alpha} antagonist binding to endogenous RAR{alpha}, a few histone and DNA-modifying enzymes might be rapidly recruited at RARß2 to impose repressive changes. Our supposition is corroborated also by the observation that T47D cells treated up to 96 h with another RAR{alpha} antagonist, RO415253 (10 µM) (1), developed DNA methylation within the first 4 h (Fig. 6D). This is consistent with previous reports showing that a dominant-negative RAR{alpha} mutant transfected in cells carrying an unmethylated RARß2 triggered the appearance of RARß2 DNA methylation within a few hours of transfection (13). Thus, aberrant RARß2-repressive changes seem to occur as rapidly as normal chromatin changes (29). However, we do not know yet which chromatin repressor proteins are recruited and in which order at RARß2 in response to the RAR{alpha} antagonist.



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  		FIG. 6. Induction of RARß2 epigenetic silencing by RAR{alpha} antagonists. (A and B) (A) Time course analysis showing that the RAR{alpha} antagonist ER50891 (top) abrogates RA-induced RARß2 transcription in T47D cells (bottom) concomitant with (B) induction of a significant (P < 0.05) increase of repressive H3-Tri-Me-K9 (middle) and a significant (P < 0.05) decrease of activating H3-Ac-K9 (bottom), while RAR{alpha} resides at RARß2 (top). (C) MSP (top) and bisulfite sequencing (bottom) show CpG methylation already at 4 h after treatment with ER50891. (D) Time course analysis of RARß2 methylation with the RAR{alpha} antagonist RO415253, showing rapid occurrence of CpG methylation (MSP, middle; bisulfite sequencing, bottom). (E) Colony formation assay showing that ER50891 can rescue T47D cells from RA-induced growth inhibition (top). MSP analysis of pools of clones derived from cells that survived treatment with ER50891 alone or in combination with RA detected both U and M alleles (bottom). (F) Stability of RARß2 epigenetic silencing in ER-C6 and ER/RA-C5, showing heterozygosity for U and M alleles (top left), RA resistance (bottom left), lack of RA-induced RARß2 transcription (top right), and histone H4 hypoacetylation (bottom right).

It can be argued that RARß2 epigenetic silencing was observed because of the constant presence in the cell of inductive factors (such as the dominant-negative RAR{alpha}, the RAR{alpha}-RNA interference construct, and the RAR{alpha} antagonists).

Here we show that RARß2-repressive chromatin changes are retained by T47D cells after removal of the RAR{alpha} antagonist. T47D cells treated for up to 24 h with ER50891 (10 µM), alone or in combination with RA (0.1 µM), were grown in drug-free medium until we observed the appearance of discrete colonies. A clonogenicity assay showed that ER50891 could rescue cells from RA-induced growth inhibition (Fig. 6E, top). DNA was extracted by the entire pool of clones that survived treatment with ER50891 alone or in combination with RA (for details, see Materials and Methods). MSP showed the presence in both pools of clones of RARß2 alleles with and without CpG methylation (Fig. 6E, bottom). MSP analysis of two independent clones that survived ER50891 treatment (clone ER-C6) or combined ER50891 and RA treatment (clone ER/RA-C5) showed the presence of both U and M alleles (Fig. 6F, top left). The chromatin associated with the silent RARß2 alleles in these clones was marked by histone H4 hypoacetylation (Fig. 6F, bottom right), consistent with lack of RA inducibility of RARß2 transcription (Fig. 6F, top right) and RA resistance (Fig. 6F, bottom left). Apparently, RAR{alpha} was no longer bound at RARß2 (S. Pozzi, unpublished observations). We conclude that after removal of the inductive factor (RAR{alpha} antagonist), the RARß2-repressive epigenetic changes remain stable in at least some of the cells that were originally exposed to the inductive factor.

RARß2 reactivation requires restoration of RA signal at a silent RARß2 through RAR{alpha}. Our work, as well as the work of others, has shown that treatment of cells carrying a silent RARß2 with either histone deacetylase inhibitors or demethylating agents, alone or in combination, can resensitize an epigenetically silent RARß2 to RA (6, 24, 34). A few preliminary observations pointed at RAR{alpha} as a critical factor for RA-induced RARß2 reactivation from an epigenetically silent promoter. We observed specifically that (i) the occurrence of reactivation of RA-induced RARß2 transcription in a RAR{alpha}-negative SG5C1 clone (derived from MDA-MB-231) treated with TSA (330 nM) and/or 5-Aza (0.8 µM), alone or in combination (Fig. 7A, bottom left), was concomitant with reactivation of endogenous RAR{alpha} (Fig. 7A, top left), (ii) the drug-induced endogenous RAR{alpha} was found at the RARß2 promoter in response to the presence of RA (Fig. 7A, middle), and (iii) RA-induced RARß2 transcription after treatment with the different drugs was abrogated by the RAR{alpha} antagonist ER50891 (Fig. 7A, right).



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  		FIG. 7. Restoring RA signal at a silent RARß2 through RAR{alpha} leads to RARß2 reactivation. (A) Concomitant reactivation of RAR{alpha} (top left) and RARß2 (bottom left) with TSA and 5-Aza alone or in combination in MDA-MB-231 SG5C1 leads to a significant increase of RAR{alpha} binding at RARß2 (middle). The RAR{alpha} antagonist ER50891 significantly abrogates RA-induced RARß2 transcription after TSA and 5-Aza treatment alone (P < 0.05) or in combination (P < 0.001) (right). Ab, antibody. (B) Exogenous expression of RAR{alpha} in MDA-MB-231 RAR{alpha}C21 (top left) leads to significantly (P < 0.001) higher RA-induced RARß2 transcription (top middle) concomitant with significant recruitment (P < 0.05) of RNA polymerase II at RARß2 (top right) and a significant increase of Ac-H4 (P < 0.05) in the RARß2 chromatin (bottom left). Apparently RAR{alpha} associates significantly (P < 0.05) more with the reacetylated chromatin in the presence of RA (middle bottom). RARß2 DNA associated with RAR{alpha} is methylated (bottom right). (C) Real-time RT-PCR shows that the level of RA-induced RARß2 transcription after treatment with TSA is significantly (P < 0.01) higher in RAR{alpha}C21 than in SG5C1 (top left), and paralleled by a significant (P < 0.05) increase in Ac-H4 in the RARß2 chromatin (middle left), as well as binding of both RAR{alpha} and RNA polymerase II at RARß2 (top right and bottom right, respectively), in the absence of demethylation of RARß2 DNA (bottom left). Demethylation was observed only in the samples treated with 5-Aza (bottom left). (D). Schematic diagram based on the findings presented in panels A, B, and C showing that RAR{alpha}—either drug-induced endogenous (en-RAR{alpha}) or exogenous (ex-RAR{alpha})—plays an active role in reestablishing a RARß2 status, which is permissive for transcription.

Next, we mechanistically tested the involvement of RAR{alpha} in the reactivation of transcription from a silent RARß2 in an MDA-MB-231 clone expressing exogenous RAR{alpha} (RAR{alpha} C21) (Fig. 7B, top left). We observed that RARß2 transcriptional reactivation (Fig. 7B, top middle) and recruitment of RNA polymerase II at RARß2 (Fig. 7B, top right) in response to the presence of RA (1 µM, 24 h) occurred with significant reacetylation of histone H4 (Fig. 7B, bottom left). By immunoprecipitating with anti-RAR{alpha} antibodies the reacetylated chromatin, we found a significantly higher level of RAR{alpha} bound at RARß2 in response to RA (see Re-ChIP panel, Fig. 7B, bottom middle). MSP analysis of the RARß2 DNA immunoprecipitated by RAR{alpha} showed persistence of RARß2 CpG methylation (Fig. 7B, bottom right). Apparently, reestablishing the RA signal at RARß2 via exogenous RAR{alpha} by inducing RARß2 chromatin reacetylation enabled transcription from the methylated RARß2.

Interestingly, the level of RA-induced RARß2 transcription obtained with exogenous RAR{alpha} was comparable to the level of RA-induced transcription obtained with the two drugs in SG5C1 (Fig. 7C, top left, arrows) and correlated with the increase of histone H4 reacetylation at RARß2 up to the threshold enabling transcription (Fig. 7C, middle left, arrows) rather than with demethylation of RARß2 DNA (Fig. 7C, bottom left). The increment in the level of drug-induced endogenous RAR{alpha} in SG5C1 and drug-induced endogenous plus exogenous RAR{alpha} in RAR{alpha}C21 at RARß2 (Fig. 7C, top right) was mirrored by an increment of RNA polymerase II recruitment at RARß2 (Fig. 7C, bottom right). This latter increment, paralleled by the increment in RARß2 transcription, might reflect an involvement of RARß2 itself, which, once induced by RA, would contribute to its own transcription by a positive-feedback autoregulatory loop (8, 21, 36). However, we were not able to prove it by ChIP analysis with anti-RARß2 antibodies as we did instead in control T47D cells (data not shown).

Based on the overall findings that we summarized in the schematic diagram in Fig. 7D, RAR{alpha} appears to be critical in the restoration of a permissive transcriptional status at a silent RARß2.


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DISCUSSION
 
We proposed previously that hormone-regulated genes, whose transcription is normally regulated by dynamic changes of the chromatin status in the presence and absence of the specific hormone (9, 14, 29, 41), might be ideal models for testing the etiology of aberrant epigenetic silencing in cancer and aging (33). In particular, we hypothesized that an aberrant chromatin-repressive status, consequent to lack of either RA or a crucial component of the machinery necessary to mediate the integration of RA signal at RARß2, could explain why RARß2 is so frequently epigenetically silenced in RA-resistant cancer cells (33) and RA-resistant tumors (34).

In the first part of this report we prove that impairing the integration of RA signal through RAR{alpha} at RARß2 leads to RARß2 epigenetic silencing. Conversely, we show that reintegration of RA-RAR{alpha} signaling at a silent RARß2 leads to transcriptional reactivation. RAR{alpha}, the upper regulator of RARß2 transcription, is expressed in RA-sensitive cells where RARß2 can be induced by RA and is homozygous for unmethylated RARß2 alleles (Fig. 1). When we interfered with RA signaling at RARß2 by three alternative strategies, namely, a dominant-negative RAR{alpha} lacking the RA-binding domain (Fig. 2), RAR{alpha}-RNA interference (Fig. 5), and RAR{alpha} antagonists (Fig. 6) in RA-sensitive cells, we always induced the conversion of RARß2 alleles permissive for transcription into a nonpermissive (unresponsive to RA) status. The chromatin of nonpermissive alleles was marked by repressive histone modifications (H3-Tri-Me-K9, hypoacetylation of histone H4, and H3-K9), and also by CpG methylation, but in only a fraction of alleles, which remained unresponsive to RA. RA could not influence the level of critical histone modifications to reach what we define as the "threshold" of histone modifications required for transcription (as it happens instead in cognate control clones) even when used at a pharmacological concentration (1 µM). Remarkably, a nonpermissive RARß2 status can be conferred without CpG methylation. Consistently, we identified RA-resistant cells, which were homozygous for RARß2 unmethylated alleles yet nonpermissive for transcription in response to RA (Fig. 4).

We further demonstrated that the RARß2 nonpermissive (nonresponsive to RA) status was stable also in the absence of the inductive factor. Specifically, we demonstrated that this is the case by attenuating the RA signal at RARß2 with a RAR{alpha} antagonist and showing that the RARß2 nonpermissive status—marked by repressive histone H4 hypoacetylation and, in some alleles, also CpG methylation—was maintained for a long time after the RAR{alpha} antagonist was removed (Fig. 6E and 6F).

It was beyond the scope of this study to show which repressor proteins (including histone-modifying enzymes and DNA methyltransferases) initiated the exacerbation of the RARß2-repressed status in response to different inductive factors. However, from the initial repressive events induced by either the dominant-negative RAR{alpha} or the RAR{alpha} antagonist ER50891 we speculate that both histone and DNA-modifying enzymes might have been recruited at RARß2 while there was persisting occupancy of the promoter by either the dominant-negative (Fig. 2B) or endogenous wild-type RAR{alpha} (Fig. 6B). The order in which repressive changes at histone and DNA level accumulate at gene promoters has been addressed in a few studies (2, 35). In the case of RARß2, the accumulation of repressive histone modifications appears to precede CpG methylation. We inferred that this might be the case because we found that only a fraction of nonpermissive alleles developed CpG methylation and in only a few CpGs in the first exon (Fig. 2, 5, and 6). It is conceivable that this region is the epicenter of CpG methylation. These findings suggest indirectly that DNMTs are recruited after other repressive critical proteins. This is because the nonpermissive status can also be achieved in the absence of CpG methylation.

We also demonstrated that induction of a silent, nonpermissive RARß2 status is indeed the cause of biological RA resistance. Directly targeting RARß2 transcription with a RARß2 antisense in RA-sensitive T47D cells led to the same RA-resistant phenotype (Fig. 3) observed after induction of RARß2 epigenetic silencing with any of the three strategies used to functionally inactivate RAR{alpha}. In summary, impairment of RA signal at RARß2 through RAR{alpha} in RA-sensitive cells appears to lead to an exacerbation of the RARß2 chromatin status, stable silencing, and, ultimately, RA resistance (Fig. 8).



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  		FIG. 8. The critical role of RA-RAR{alpha} signaling in RARß2 epigenetic silencing and RA resistance. (A) Normally, in the absence of RA, RAR{alpha} keeps RARß2 chromatin poised for transcription; upon RA binding at RAR{alpha} the chromatin is converted to an active state (14). RARß2 in turn enhances its own transcription (8, 21, 36). Induction of the RARß2 tumor suppressor results in cell growth inhibition. (B) Impaired RA signal through RAR{alpha} at RARß2 apparently converts RARß2 chromatin from a poised to a severely repressed, silenced state no longer compatible with transcription. This state is characterized by epigenetic repressive modifications at the histone and DNA level. The biological consequence of the conversion of RARß2 into a silent state is the resistance to the growth-inhibitory action of RA. (C) Conversely, reconnecting RA signal at a silent RARß2 chromatin through RAR{alpha} restores a chromatin state again compatible with transcription. RARß2 reactivation results in the reestablishment of responsiveness to the growth-inhibitory action of RA.

Conversely, we prove that reconnecting RA signal through an exogenous RAR{alpha} at a silent, heavily hypermethylated RARß2 in RA-resistant cells results in transcriptional reactivation concomitant with RARß2 chromatin histone H4 reacetylation but not demethylation (Fig. 7C, left). Interestingly, the level of RA-induced RARß2 reactivation in the presence of exogenous RAR{alpha} was comparable to the level induced with TSA in the presence of endogenous RAR{alpha}. Also, in this case RARß2 reactivation was achieved without demethylation. Apparently, RA binding to RAR{alpha}, which is known to actively recruit coactivator complexes with histone acetyltransferase activity (29), is sufficient to convert RARß2 from a silent to a permissive state, as we show in the schematic diagram in Fig. 7D. A consequence of restoring RA-RAR{alpha} signaling at RARß2 likely is the reactivation of the RARß2 receptor itself, which is expected to sustain its own transcription (8, 21, 36). Because we did not detect RA-induced RARß2 reactivation after treatment with TSA and 5-Aza alone or in combination when we abrogated RAR{alpha} function with the RAR{alpha} antagonist (Fig. 7A, right), we conclude that RARß2 can play a role in its own transcription but only after its transcription is triggered by RAR{alpha}, as we show in the schematic diagram in Fig. 7D. In summary, reintegration of RA signal at an epigenetically silent RARß2 through RAR{alpha} in RA-resistant cells would restore a chromatin status enabling transcription of the tumor suppressor gene in response to the presence of RA and, consequently, sensitivity to the growth-inhibitory action of RA (Fig. 8).

Lack of functional integration of RA signal at RARß2 through RAR{alpha} might occur in vivo. RAR{alpha} is not expressed in a high percentage of tumors (31). This leads to the hypothesis that in vivo RARß2 silencing can be a consequence of RAR{alpha} loss or silencing. In support of this hypothesis, we observed that reactivation of RARß2 by TSA and 5-Aza also restored RAR{alpha} (Fig. 7A, left). Interestingly, estrogen receptor alpha (ER{alpha}) is often epigenetically silenced in RAR{alpha}-negative tumors (16). RAR{alpha} is regulated, at least in part, by ER{alpha} (30). This makes us speculate that epigenetic silencing of a few hormonally regulated genes could occur in a "domino" fashion. We continue to assert (33, 34) that lack of intracellular RA availability at RAR{alpha} could also trigger a repressive epigenetic domino effect. Recently we found that RARß2 silencing leads to the silencing of downstream genes involved in RA metabolism, thus suggesting the existence of epigenetic networks (unpublished observations).

A few translational considerations come to mind at the end of this report. Detection of RARß2 hypermethylation, a test for early breast cancer (3) and other cancers, apparently underestimates epigenetic RA resistance, because it may miss cells homozygous for silent, yet unmethylated, RARß2 alleles (Fig. 4). Therapeutic strategies aimed at resensitizing RA-resistant cells to RA by restoring RARß2 would require drugs powerful enough to reinduce simultaneously both RARß2 and critical RARß2 regulators. But more than anything else, this report highlights the importance of identifying the intrinsic and extrinsic factors that in vivo might force genes like RARß2 into aberrant or protracted inactivity. Identification of these factors can lead to novel cancer prevention strategies.


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ACKNOWLEDGMENTS
 
We thank James Herman, David Kowalski, André Hoogeveen, and the anonymous reviewers for helpful suggestions and constructive criticism and Ellen Sanders for manuscript editing.

This work was supported by an AIRC grant (Italy) (N.S.), U.S. Army grant DAMD17-02-01-0432 (N.S.), a Roswell Park Cancer Institute Alliance grant (N.S.), the Graduate Program of Molecular Medicine, University of Milan (S.P., G.B., and G.S.), and the CISI Center of Excellence, University of Milan (S.R.).


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FOOTNOTES
 
* Corresponding author. Mailing address: Roswell Park Cancer Institute, Elm & Carlton Streets, C&V Bldg., RM 226, Buffalo, NY 14263. Phone: (716) 845-1053. Fax: (716) 845-1741. E-mail: nicoletta.sacchi{at}roswellpark.org. Back


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Molecular and Cellular Biology, December 2005, p. 10591-10603, Vol. 25, No. 23
0270-7306/05/$08.00+0     doi:10.1128/MCB.25.23.10591-10603.2005
Copyright © 2005, American Society for Microbiology. All Rights Reserved.



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