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J. Biol. Chem., Vol. 280, Issue 15, 14545-14555, April 15, 2005
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From the Thyroid Section, Division of Endocrinology, Diabetes and Hypertension, Department of Medicine, Brigham and Women's Hospital, Harvard Institute of Medicine, Boston, Massachusetts 02115
Received for publication, October 8, 2004 , and in revised form, January 26, 2005.
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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1 mutant. More interestingly, NCoR acted as a co-activator to enhance TR-mediated basal transactivation in the absence of T3. This effect was eliminated by removal of TR or NCoR binding. Most strikingly, T3 induced a remarkable increase in TR·DNA binding at 40–60 min after T3 exposure that rapidly returned to basal levels, suggesting a T3-induced remodeling of chromatin structure at the early stage of T3 stimulation resulting in repression. Therefore, we propose a mechanism by which NCoR, GAF, and TR interact with the CD44 negative T3-responsive element to enhance basal transactivation, whereas T3 induces the remodeling of chromatin structure for repression. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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and TR
, each of which consists of a DNA binding domain (DBD), a ligand-binding domain, co-activator/co-repressor binding domains, and a transcription activation domain. The P-box of the zinc fingers within the DBD of TR recognizes the AGGTCA sequence of the thyroid hormoneresponsive element (TRE), which is diversely arranged in direct repeat (DR+4), palindrome, and inverted palindrome forms (1, 2). When unliganded TR binds to TRE, it recruits co-repressors (NCoR/SMRT) and histone deacetylases for repression. On the other hand, T3 binding to TR induces conformational changes in TR to release co-repressors and recruit co-activators such as CBP/p300, pCAF, and SRC-1 for transactivation (3–8). Thus, although the mechanism of transcriptional regulation for the positive TRE (pTRE) is relatively well studied, T3-dependent negative gene regulation is still poorly understood and is limited to a few genes. It has not been possible to identify a consensus sequence of negative TRE (nTRE).
Recently, Wondisford and colleagues (9) provided evidence that direct binding of hTR2 to nTRE is required for T3-dependent negative regulation. This group generated a non-TRE binding hTR2 mutant (TR2-GS125) in which two amino acids (EG at 125 and 126 in the zinc finger region of DBD of TR2) were mutated to GS, preventing recognition of the TRE sequence. When TR2-GS125 was transfected into 293T cells, T3-dependent negative regulation of the hTSH gene was abolished (9). Subsequently, Wondisford et al. showed that TR2 bound to the TRE sequence in the hTSH fragment elicited a negative response to T3. The physiological significance of TR·DNA binding was further confirmed by the finding that TR2-GS125 knock-in homozygous mutant (TRGS/GS) mice displayed abnormal T3 regulation of the hypothalamic-pituitary thyroid axis and retina, identical to abnormalities previously observed in TR KO (TR–/–) mice, although TRGS/GS mutant mice showed much less severe defects in hearing and outer hair cell loss than do TR–/–mice (9). These results indicate that TR·DNA binding is necessary for negative transcriptional regulation of the TSH gene.
Other studies suggest that TR binds directly to TREs or TRE-like sequences for negative regulation by T3 (9, 10) and that various types of nTRE sequences resembling TREs are located in the promoters (trkB and amyloid precursor protein) (11–13), the first exons (c-myc) (14), and the 3'-untranslated regions (rat growth hormone and Clone 144) (15). For example, the TSH promoter contains palindromic TREs necessary for negative regulation by T3 (16), whereas mouse TSH has a DR+2 TRE that includes a TRE near the TSS (9, 17, 18). RSV-LTR, Cyp7A1, and SCD1 promoters have different structures (10, 19, 20). However, no nTRE consensus sequence distinct from pTRE was clearly identified, raising the possibility of a novel nTRE different in sequence from these positively regulated genes.
On the other hand, there are also data that T3-dependent negative regulation can occur without direct TR·DNA interaction. Kushner and colleagues (21, 22) showed that TR interacted with jun/fos through the AP1 site by a protein-protein interaction that was subsequently regulated by T3. Another recent study from our group indicated that JEG-3 cell-specific transcription factor-DNA binding induced T3-dependent negative regulation without direct TR·DNA binding (23). Jameson and colleagues indicated that TSH gene expression might be negatively regulated by T3 through a squelching mechanism (24, 25). This model proposed that, in the absence of T3, unliganded TR bound to co-repressors, forming a complex that was subsequently sequestered from the promoter region, resulting in increased histone acetylation and transactivation through the recruitment of more co-activators to the promoter. However, in the presence of T3, T3-TR formed a complex with co-activators such as CBP/p300 that was segregated from the TSH promoter, giving co-repressors NCoR/SMRT and histone deacetylases easy access to the promoter region for histone deacetylation and repression (25). When NCoR or SMRT was overexpressed in the presence of TRH and TSH and - promoter-Luc, TR-mediated basal transactivation of these promoters was enhanced by TR·NCoR interaction (9). These results suggest that negative regulation by T3 could be induced without direct TR·DNA interaction (25). However, this model did not explain how the specificity of negative regulation is conferred. Thus, the question of whether TR·DNA interaction with the negatively regulated genes is essential for T3-dependent negative regulation is still not resolved.
We approached this problem by screening the rat genome for other genes whose expression is suppressed by T3. We determined that CD44 is negatively responsive to T3 at a directly transcriptional level in various tissues and cell types. CD44 is a cell adhesion molecule involved in diverse biological processes, including angiogenesis, lymphogenesis, wound healing, inflammation, and cancer metastasis (26). Recent reports indicate that CD44 plays an important role in generation of papillary thyroid carcinomas (PTCs), the most prevalent malignancy of the thyroid gland. Some PTCs are thought to be initiated by the RET/PTC chimeric oncogene, which is generated by rearrangements of the RET receptor tyrosine kinase (26). RET/PTC signaling induces up-regulation of osteopontin and CD44, resulting in proliferation and invasion of transformed PC Cl 3 thyrocytes (26). Another interesting and relevant finding is that Alzheimer's disease-associated 
-secretase cleaves CD44, generating intracellular domains for nuclear signaling and CD44 -peptides, whose functions are unknown. This cleavage pattern is similar to that of -amyloid precursor protein by the -secretase, which produces an amyloid precursor protein intracellular domain for nuclear signaling and an amyloid -peptide for -amyloid plaque formation (27). Notch, low density lipoprotein receptor-related protein, E-cadherin, and ErbB-4 are also the same family proteins (28–36). All of these results suggest a correlation between regulation of CD44 expression and progress of these diseases.
Here we provide evidence that, during negative regulation by T3, TR binds weakly but directly to a novel CD44 nTRE different from the pTRE. In addition, we identified two proteins (GAF and NCoR) required for T3-dependent negative regulation of CD44 gene expression. Of special note, cooperative interaction among TR, NCoR, and GAF was essential for enhanced unliganded TR-mediated basal expression, whereas repression was induced by T3 through a transient, but strong in vivo TR·DNA interaction, probably also bringing about remodeling of the chromatin structure. These results suggest a novel mechanism for T3-dependent negative regulation.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Plasmid Constructions and Mutagenesis—Standard techniques were used for all plasmid constructions, as described previously. A 5-kb 5'-FR genomic fragment, PCR-amplified 0.1- to 1.8-kb DNA fragments of 5'-FR, or synthesized double-stranded oligonucleotides were subcloned into KpnI and XhoI restriction sites of the pGL3-basic luciferase-expressing reporter vector (Promega, Madison, WI). A 177-bp CD44 promoter in pGL3BS-Luc (177-bp CD44-Luc) was mutated using the QuikChange site-directed mutagenesis kit (Stratagene, La Jolla, CA) as indicated in figure legends. To generate a non-TRE binding GS71 sequence as already described in JEG-3 cells (23), mTR1 cDNA was inserted in a CDM expression vector in which two bases from TSS of the mTRa1 cDNA were mutated: 614 (A to G) and at 616 (G to A). All constructs were confirmed by automated sequencing.
Constructs for siRNA expression were generated by cloning hairpin siRNA sequences into pSilencer2 vectors (Ambion, Austin, TX) according to the manufacturer's protocol. Target sequences for the expressed siRNA were 814–834 (5'-AAAGGAGTGATTCTCGATCAC-3') for NCoR, 377–398 (5'-AACCAGAACGGCGCCGAAGGC-3') for GAF, 343–361 (5'-GGTTATGTACAGGAACGCA-3') for GFP, and 5'-AAGATCGATCGATCGATCGAT-3' for random sequence siRNA. All sequences of cloned constructs were also confirmed by automated sequencing.
Transfection and Luciferase (Luc) Assay—Transfections were performed as previously described (23) with FuGENE 6 transfection reagent (Roche Applied Science) in COS-7 cells. Each plate was transfected with 2 µg of Luciferase reporter plasmids and 0.5 µg of TKGH, which constitutively expresses human growth hormone (hGH), as a control for transfection efficiency. 0.1 µg of CDM8 vector expressing mouse TR1 (wild-type or mutant) or pCDNA3 vector expressing NCoR were transfected with and without 0.1–0.3 µg of pSilencer 2.1-U6 neo (Ambion) expressing siRNAs of NCoR and GAF, as described in the figure legends. CDM8 and pCDNA3 empty vectors or GFP and random oligonucleotide siRNA-expressing vectors were used as controls. After transfection, cells were incubated in 60-mm culture dishes in the absence of thyroid hormone for 2 days and then exposed to 50 nM T3 for 0–24 h at 37 °C. A luciferase assay was performed according to the manufacturer's protocol (Promega) and was normalized to hGH expression in the absence and presence of T3 to calculate T3 responsiveness. Transfection data are mean ± S.D. of a minimum of triplicate samples.
Electrophoretic Mobility Shift Assay—EMSA was performed as described previously (23). Double-stranded oligonucleotides of wild-type and mutant CD44 nTRE were radiolabeled with [32P]dCTP (PerkinElmer Life Sciences) by fill-in reaction using Klenow DNA polymerase, which was subsequently gel-purified (23). Chicken TR1 and human retinoid X receptor were overexpressed in Escherichia coli and purified as described previously (23). Radiolabeled probes were reacted with bacterially expressed cTR and/or human retinoid X receptor and resolved by non-denaturing PAGE.
Methylation Interference Assay—Dimethyl sulfate methylation interference assay was performed using a standard protocol as described previously (23).
Chromatin Immunoprecipitation Assay—ChIP assay was performed according to manufacturer's protocol (Upstate, Lake Placid, NY) (38, 39). Briefly, 177-bp wild-type (177-bp CD-Luc) and mutant (177-bp CD-Dbl-Luc) constructs were co-transfected with TR and/or NCoR expression vectors into COS-7 cells and incubated in the presence or absence of 50 nM T3 for 0–24 h as represented in Fig. 8. Cells were cross-linked by 1% formaldehyde for 10 min and harvested in the presence of protease inhibitor (EDTA-free Complete, Roche Applied Science). These cells were then lysed and sonicated to generate 200- to 500-bp DNA fragments, which were subsequently confirmed by agarose gel electrophoresis. TR and NCoR binding fragments were immunoprecipitated using rabbit polyclonal anti-TR and goat polyclonal anti-NCoR antibodies (Santa Cruz Biotechnology, Santa Cruz, CA). As a control, immunoglobulin G (IgG) was used to determine the background level. Nonspecific interaction was minimized by thorough washing. Specifically bound DNA fragments were purified and quantified by real-time PCR using specific primers. The sense primer was pGL3P-4952S1 (pGL3 Promoter-Luc vector, –4952 to –4971: 5'-CTAGCAAAATAGGCTGTCCC-3'), and the antisense primer was 31CD described in Fig. 8 (CD44, –31 to –51: 5'-CCAGGCTTTGAAAGAGTGACC-3').
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The eight genes identified as strongly negatively regulated by T3 were tested to determine whether they are transcriptionally or post-transcriptionally regulated by T3. To identify which of the cDNAs were transcriptionally regulated by T3,2 µM actinomycin D was used to block transcription, and the half-life of the mRNA was measured in the presence and absence of T3. Genes that showed no changes in mRNA half-life by T3 and actinomycin D exposure were categorized as transcriptionally regulated (data not shown). To identify those directly responsive to T3, 30 µM cycloheximide was added to hypothyroid GC cells 1 h prior to T3 treatment. Cells were taken for RNA isolation after 0, 3, and 6 h of exposure to T3, and the changes in the specific RNAs were compared with control cells not exposed to cycloheximide. Genes that showed no changes in mRNA level by T3 and cycloheximide exposure were categorized as directly responsive to T3 (data not shown). Four genes met both criteria.
To identify which of these four genes was negatively regulated by T3 in a general, as opposed to a tissue-specific fashion we examined mRNA of nine tissues from hypothyroid, euthyroid, and hyperthyroid (4-h and 24-h T3-treated) rats. These included blood, lung, anterior pituitary (AP), large intestine (L. Int.), stomach (STM), kidney, liver, heart, and hypothalamus (HPT) (Fig. 1). Serum T4 and T3 measurements were also performed to confirm the thyroid status of these animals. Examination of T3-dependent mRNA expression of the four genes in these tissues by quantitative real-time PCR showed that two were generally negatively regulated, whereas the other two were negatively regulated in a tissue-specific manner (data not shown). Because the CD44 gene was negatively regulated in most tissues (Fig. 1), it was selected as a model gene with which to study the T3-dependent negative regulation mechanism.
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1 (mTR1) expressing plasmid into COS-7 cells, luciferase expression was decreased 2-fold with T3 treatment (Fig. 2A). Similar results were obtained using other cell lines such as anterior pituitary (GC) and hypothalamic (HT22) cells (data not shown), confirming that this 5-kb 5'-FR fragment was negatively responsive to T3. Serial deletion mutation analysis showed that the negative response to T3 was maintained until the gene fragment was reduced to 177-bp (–129 to +48 bp from the transcriptional start site (TSS)) (Fig. 2A). The effect of TR1 was similar to that of TR1 (data not shown). When the 5'-FR was further reduced to 113-bp (–65 to +48 bp), the repression ratio (+T3 Luc/–T3 Luc) of the 113-bp fragment was significantly reduced to 0.71 (Fig. 2A), and the transcriptional potency of the construct in the absence of T3 was also substantially lowered to near-background level (Fig. 2B). Furthermore, when GAGA factor (GAF) binding site (TCTC at –1to +3 bp) within the 113-bp fragment was mutated to TTTT, the weak residual TR-response was completely abolished (Fig. 2B, (–65
+48)-GAF). These results indicate that the GAF binding site and the sequences located between –129 and –66 bp are critically important for negative regulation of CD44 gene expression. This suggests that the sequences between –129 and –66 bp include TR binding sites for a negative response to T3.
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1. EMSA showed that TR1 strongly bound the 89-bp fragment (–129 to –41 bp) but not with the 88-bp fragment, as expected (data not shown). Subsequent dimethyl sulfate footprinting assay with the 89-bp fragment showed that G-residues at –80, –78, –52, and –51 of the top strand and at –76, –75, –49, and –47 of the bottom strand were specifically protected by cTR1 (Fig. 3A). Interestingly, the protected regions consisted of two positive TRE half-sites with "everted" palindromic orientation separated by 20 bp (Fig. 3B). Each protected region contained an additional protected Gly residue at the end of the TRE half-site that was different from the typical pTRE, which consists of a direct repeat of a 4-bp sequence separation (DR+4).
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and RSV, and the 170-bp fragment of rD2 conferring a negative response to T3 showed much lower affinity to TR1 than did the 45-bp CD44 fragment (Fig. 3C, lanes 8–10). This raises the question of whether the presence of high affinity TREs in CD44 is required for negative regulation by T3. The High Affinity TR Binding of TREs within the 64-bp Fragment (–129 to –66 bp) Was Not Required for T3-dependent Negative Regulation—We sought to determine whether these high affinity TREs were directly involved in negative regulation by T3. First, we generated several mutations at or around the TREs, as depicted in Fig. 4A. EMSA showed that mutations of both TRE sites resulted in a complete loss (M1) or a significant decrease (M2) in TR binding (Fig. 4B). However, shortening of the spacer length without changing the TR binding sites did not decrease the binding affinity (M3–M5) but actually increased binding in M6 (Fig. 4B). This again confirmed that these two TREs are high affinity TR binding sites (Fig. 4B).
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1-expressing construct into COS-7 cells, none of them showed disrupted T3-dependent negative regulation (Fig. 4D). The total luciferase expression by mutated constructs (in the absence and presence of T3) was decreased 2- to 3-fold compared with the wild type, but the changes in the repression ratios were small compared with those noted with the wild type (Fig. 4D, WT). When empty vector lacking mTR1 was used as a control, it did not respond to T3 (Fig. 4D, CNT). These results indicated that high affinity TR binding TREs are not necessary for T3-dependent negative regulation of CD44 gene expression. This raises the possibility that weak or transient TR binding sites are present within the 64-bp CD44 fragment conferring negative response to T3, as illustrated in hTSH (9) and RSV nTRE (10, 40) fragments (Fig. 3C). Direct TR·DNA binding is required for T3-dependent negative regulation of the mutant 177-bp CD44-Dbl fragment that lost its capacity for high affinity binding to TR1.
To determine whether there is direct TR·DNA interaction in vivo for negative regulation by T3, we generated a non-DNA binding mutant GS71. The amino acids at 71 and 72 in the P-box of DNA binding zinc-finger domain of mTR1 were converted from EG (Glu and Gly) to GS (Gly and Ser) of mTR1 (Fig. 5, A and B), which eliminated DNA binding capability, as was the case in the hTR2-GS125 mutant (9).
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1 expression construct into COS-7 cells, Luc expression was decreased by T3 as expected (Fig. 5C, TR1). However, transient expression of GS71 instead of TR1 eliminated any T3-dependent negative regulation (Fig. 5C, GS71). Similar results were obtained using the wild-type 177-bp CD44-Luc (data not shown) as well as the 170-bp rD2-Luc and 1.2-kb hTSH-Luc conferring a negative response to T3 (Fig. 5, D and E). To demonstrate the effects of these two receptors on these constructs, we included a potent positively responding reporter, a TK-Luc plasmid containing three copies of the consensus DR+4 TRE (41) from the hdio1 gene were used. These plasmids, along with an empty vector control, were transfected into COS-7 cells. The results of luciferase expression relative to an internal transfection efficiency control (TKGH), with and without T3, are shown in Fig. 5F. These data indicated that the potent positive regulation by the TR was markedly decreased in GS71. In sum, these data indicated that the direct weak or transient TR binding site for T3-responsive repression was located within the 177-bp rCD44-Dbl DNA fragment, even in the absence of high affinity TR binding sites.
NCoR Enhances TR-mediated Basal Transactivation of the rCD44 Gene—The indication that direct but weak TR·DNA binding is necessary for negative regulation by T3 raises the possibility that other co-repressors interact with TR on the nTRE. Therefore, we addressed whether NCoR is involved in this negative regulation mechanism. When the 177-bp CD44-Dbl-Luc plasmid was co-transfected with mTR1 or mTR1 plus human NCoR-expressing constructs, NCoR further enhanced mTR1-mediated basal transactivation in the absence of T3, which then returned to basal or lower levels by T3 (Fig. 6A, lane 2 versus 5). Consistently, expression of the C-terminal region of NCoR was enough to generate the same effect on the enhanced mTR1-mediated basal transactivation as did by the full-length NCoR (data not shown), suggesting that transactivation domain is localized within the C-terminal region. This result was further supported by the knockdown of NCoR expression. When NCoR siRNA was co-expressed, NCoR-mediated basal transactivation was completely abolished (Fig. 6A, lanes 4 and 7), indicating the abrogation of TR-mediated basal transactivation. Decrease of NCoR mRNA expression after siRNA treatment was confirmed by PCR (data not shown). Co-expression of green fluorescence protein siRNA as a control did not significantly affect T3-dependent negative regulation (Fig. 6A, lane 8). Notably, when NCoR siRNA was co-expressed with mTR1 without NCoR, TR-mediated basal transactivation was completely abolished (Fig. 6A, lane 4), suggesting not only that basal transactivation by TR1 on this promoter is regulated by endogenously expressed NCoR, but also that NCoR is required for TR-mediated basal transactivation. Negative regulation by T3 was not observed when a non-DNA binding mutant GS71 was used (Fig. 6A, lane 3 and 6), suggesting that enhanced basal transactivation by NCoR was mediated through TR bound to DNA. As a control, 3XhD1-Luc containing three copies of human D1 DR+4 TRE was co-transfected with the NCoR expression plasmid; TR-mediated transactivation was repressed by NCoR (Fig. 6B). This was consistent with published data on the positive TRE (25, 42, 43). Thus, these data support the concept that NCoR acts as a co-activator to enhance TR-mediated basal transactivation of this negatively T3-responsive promoter. Unlike the mechanism responsible for the squelching of NCoR and histone deacetylases by TR, T3-dependent negative regulation of the CD44 promoter requires direct TR·DNA binding for the NCoR effect.
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–101 bp), 29 bp (–100–72 bp), and 30 bp (–65–36 bp) were incubated with bacterially expressed cTR1, a specific retarded band appeared in the band of one fragment (–100–72 bp) (Fig. 7B, lane 3; Fig. 7C, lane 3), even though its binding affinity was about 100-fold less than that of DR+4-positive TRE (Fig. 7B, lane 1; note the difference in loading amount). The retarded band was significantly reduced either by addition of the 50x unlabeled 29-bp (–100–72bp) fragment (Fig. 7C, lane 2) or by a mutation of the 6 bases at –82–87 (Fig. 7A, Dbl-M1; Fig. 7C, lane 4). These data suggest that the interaction between TR and the TTTGGG sequence at –82–87 bp might be necessary for T3-dependent negative regulation.
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–87 bp was sufficient to bring about negative regulation by T3. When either the weak TRE at –82–87 alone (Fig. 8A, Dbl-M1) or a GAGA factor (GAF) binding site at –1+3 bp alone (Fig. 8A, Dbl-M2) was mutated within the 177-bp CD44-Dbl-Luc plasmid, T3-dependent negative regulation was not significantly affected (Fig. 8B, Dbl-M1 and Dbl-M2). However, when both the GAF and the weak TR binding sites were mutated, transcriptional activity was almost abolished, showing a >2000-fold reduction compared with 177-bp CD44-Dbl (Fig. 8, A and B, Dbl-M3). In addition, when GAF expression was knocked down by transfecting 0.1 or 0.3 µg of GAF siRNA-expressing plasmid in COS-7 cells, NCoR-enhanced TR-mediated basal transactivation was abolished (Fig. 8D, siGAF/0.1 and siGAF/0.3). As a control, a plasmid that produces random oligonucleotide siRNA was transfected at the same time but elicited no significant response (Fig. 8D, siRDM/0.3). All of these data indicate that binding of both GAF and weak TR to sites at –1+3bp and –82–87 bp (respectively) is required for NCoR-dependent TR-mediated basal transactivation and T3-dependent repression of CD44 gene expression. Transient Interaction of TR and NCoR with CD44 nTRE in Vivo—It has recently been shown that the ChIP assay can be applied to the study of endogenous, integrated, or transiently transfected TR-regulated genes (9, 38, 39, 44). Therefore, we decided to perform ChIP assays to determine whether we could detect TR·DNA interaction in vivo using a transient transfection system. First, we optimized a protocol to quantitate protein·DNA binding by a combination of ChIP assay and quantitative real-time PCR (quantitative real-time PCR) as described in detail under "Materials and Methods."
First, as a control experiment, we determined TR·DNA interaction on the high affinity TRE of the wild-type 177-bp CD44 promoter. Wild-type 177-bp CD44-Luc was co-transfected with TR1 (or GS71) and NCoR expression constructs into COS-7 cells with and without 50 nM T3 for 24 h, followed by cross-linking with 1% formaldehyde to fix the cells, and ChIP assay was performed using a specific anti-TR1 antibody. The TR-bound DNA was analyzed by quantitative real-time PCR using promoter-specific primers as depicted in Fig. 9A. Transfection efficiency was normalized to hGH by co-transfection of TKGH plasmid. As an immunoprecipitation control, immunoglobulin G (IgG) was used to establish a nonspecific background against which normalization of specific signals was measured. Ratios of signals to background by IgG are illustrated in Fig. 9B.
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1 (Fig. 9B, lanes 2 versus 4). This result was consistent with the expected high affinity TR·DNA interaction on the wild-type CD44 promoter, validating this protocol as a suitable method by which to analyze in vivo protein-DNA interaction.
We then examined whether TR·DNA interaction could be detected using a 177-bp CD-Dbl-Luc that contained no high affinity TREs using the same protocol. In the absence of T3, TR1-DNA interaction was compared with GS71 as a control. As expected, we detected a weak but direct TR1·DNA interaction that was significantly higher than that with GS71, even though it was just above the background nonspecific interaction level by IgG (Fig. 9C, lane 2, TR1 versus GS71). These results indicate that the difference in DNA binding affinity between TR1 and GS71 represents actual TR1-DNA binding, because nonspecific binding to DNA by IgG was relatively higher than the back ground level by GS71. This suggests that TR interacts weakly but specifically with the CD44 promoter. Thus, the in vivo result was consistent with the in vitro results using EMSA (Fig. 7, B and C). Similar results were also obtained with NCoR, suggesting that NCoR interacts with the TR·DNA complex (Fig. 9C, lane 3).
We extended this experiment to examine the effects of T3 on TR·DNA interaction. When ChIP assay was performed with COS-7 cells that were co-transfected with 177-bp CD44-Dbl-Luc plasmid, and TR1- and NCoR-expressing constructs, and were subsequently exposed to T3 for 0 min, 10 min, 20 min, 30 min, 40 min, 50 min, 60 min, and 24 h, T3 surprisingly induced transient interaction of TR·DNA over 60 min with increases seen at 10, 30, 50, and 60 min, and decreases at 20 and 40 min (Fig. 9D). Interestingly, the peak of the strong TR·DNA interaction appeared between 50–60 min (Fig. 9D). When we repeated the same experiment with 20-min intervals of T3 stimulation, TR·DNA binding was decreased at 20 min of T3 exposure but remarkably increased at 40 min. TR·DNA interaction then decreased and maintained a steady lower level over 24 h (data not shown). In further experiments, the same pattern of transient TR·DNA interaction was reproduced, even though the peak of the strong TR·DNA interaction was slightly shifted (possibly due to small differences in experimental conditions). This clearly indicates that TR interacted with DNA strongly during the early stages of T3 stimulation. Transfection efficiency was monitored by hGH expression, maintaining a steady level during all transfections, with 8% standard deviation. Amounts of DNA used for immunoprecipitation did not vary significantly between samples, as measured by ethidium bromide staining of the agarose gel. This transient interaction suggested that T3-TR recruited chromatin remodeling enzymes to generate repression at the early stages of T3 exposure. Thus, all of these results suggest that TR interacts with a potential nTRE sequence at –82–87 bp of the CD44 promoter, weakly in the absence of T3 but transiently strongly at the early stage of T3 stimulation. This again confirms that TR binds to CD44 nTRE in vivo in T3-dependent negative regulation.
Interestingly, NCoR showed a similar pattern of dynamic interaction, although much weaker than TR binding (Fig. 8D). This suggested that NCoR also interacted weakly with the TR·DNA complex.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Localization of the Negative T3-responsive Fragment—Interestingly, the effect of TR1 on T3-dependent negative regulation of luciferase expression from the core CD44 fragment was higher or equivalent to that of TR (data not shown). Similar results were also obtained using 1.2-kb hTSH-Luc and 0.65-kb rD2-Luc reporter constructs that conferred a negative response to T3 (data not shown). This indicates that TR1 acts as a potent negative regulator of nTRE. This contradicts a previous report that TR1 was a more potent activator on pTRE than TR that has a preference for negative regulation by T3 (45). It is likely that negative regulation by TR2 is rather tissue-specific, especially to anterior pituitary and hypothalamus (46). Therefore, we chose TR1 to study the general mechanism of T3-dependent negative regulation.
After demonstrating that a negative T3 response and thus an nTRE is located within the 5-kb 5'-FR, we performed deletion analysis and identified a minimal fragment for negative regulation by T3. We initially obtained a 177-bp DNA fragment (–129+48 from the TSS) that conferred a negative response to T3 (Fig. 2A). Further 5' deletion to the 113-bp fragment (–65+48 bp) reduced but did not completely eliminate the negative response to T3 (Fig. 2A). The transcriptional activity of the 113-bp fragment was largely abolished to just above background levels (Fig. 2B). This suggests that TR still interacted (although weakly) with the 113-bp fragment, which was almost non-functional. We found that this residual response to T3 was caused by the GAF binding to a site –1+3 bp from the TSS (Fig. 8B). When the GAF binding site of the 113-bp fragment was mutated to be non-functional, the residual T3 response was completely abolished (Fig. 8C). This suggests that the GAF bound to the 113-bp fragment interacts weakly with TR and is responsible for the residual weak negative response to T3. Thus, these results support the notion that a major TR binding site for negative regulation by T3 is located within a 64-bp (–129–65 bp) fragment.
Weak TR and GAF Binding Sites Are Required for Negative Regulation by T3—Initially, we evaluated whether an nTRE could consist of the same sequences as a pTRE, because two high affinity TREs were identified within a –129–65 bp fragment conferring negative response to T3. Two half-sites of TRE consensus sequences were located at –79–74 bp (AGGACA) and –53–48 bp (AGGTCA) with a 20-bp separation. However, mutational analysis indicated that these TREs were not necessary for T3-dependent negative regulation (Fig. 4D). This result contradicts published results suggesting the existence of positive TRE-like sequences within negative T3-responsive promoters (10, 13, 16–18, 23, 40, 47–50). For example, hTSH contains a TRE sequence and RSV-nTRE two everted pTRE-like sequences with an 8-bp separation (10). This suggests that TR might not interact with pTRE for T3-dependent negative regulation, but rather that these high affinity TREs are involved in the increase of total transcriptional activity of the 177-bp CD44 promoter (Fig. 4D). In fact, EMSA showed very weak TR binding to the other nTRE fragments chosen for study (Fig. 3C), even though the hTSH and RSV promoters contain pTRE sequences in the core negative fragment (9, 10).
This result led us to re-examine whether there was any weak or transient TR binding site on the 177-bp CD44-Dbl that was mutated at the high affinity TR binding site. Functional analysis of the non-DNA binding mutant GS71 showed that TR binding was required for T3-dependent negative regulation of the 177-bp CD44-Dbl-Luc. Similar results were obtained using 1.2-kb hTSH and 170-bp rD2, which conferred a negative response to T3 (Fig. 5, D and E). Consistently, EMSA also showed that TR bound weakly to the specific sequence TTTGGG at –82–87 bp. In the same context, sequences of hTSH and RSV and 170-bp rD2 conferring a negative response to T3 also showed a weak interaction with TR1 (Fig. 3C). This interaction was further supported by ChIP assay of the 177-bp CD44-Dbl fragment (Fig. 9). Thus, it is likely that weak TR·DNA interaction is a general characteristic of the nTREs.
However, the weak TR·DNA interaction alone was not sufficient to elicit negative regulation by T3 as shown in Fig. 8B. This suggested that interaction of unknown TR-associated proteins to TR bound to the nTRE was required for the cooperative repression. We identified one of them, GAF, a DNA binding transcription factor. Mutation of both the TR binding sequence TTTGGG and the GAF binding site at –1+3 bp almost completely abolished transcriptional activity, whereas mutation of either one alone had no significant effect. This suggests that there is a putative interaction among nTRE, TR, and GAF to confer negative regulation by T3. Interestingly, the same sequence is also present in the core nTRE of hTSH, TTTGGGTCA (–4+5 bp from the TSS). When GGG was mutated to AAA in the nTRE of hTSH, EMSA and ChIP assays resulted in a loss of not only TR binding but also functional repression by T3 (9). This again supports the contention that TTTGGG might be a novel nTRE consensus sequence.
Both the GAGA-binding factor (GAF) and the GAF-responsive element have been extensively investigated, including demonstrations that GAF functions in transcriptional regulation and chromatin remodeling (51). Our functional analysis of the CD44 promoter showed that GAF binding was required for negative regulation by T3. It is also possible that GAF is generally involved in the T3-dependent negative regulation mechanism, not only with the CD44 gene. Sequence data indicated that other potential GAF binding sites were located at –207–204 bp and at +52+55 bp of the hTSH gene. It will be interesting to determine whether TR bound to TRE at –1+5 bp of hTSH interacts with GAF at one of these binding sites.
NCoR-mediated Basal Transactivation—Different from the "squelching" model, which proposed that NCoR enhanced unliganded TR-mediated basal transactivation without TR·DNA interaction, our results indicate that T3-dependent negative regulation requires TR·nTRE interaction, even though its interaction was weak in the absence of T3 (demonstrated by EMSA, ChIP, and transfection assays with GS71). In addition, both GAF and NCoR were required for T3-dependent negative regulation, possibly through NCoR, GAF-DNA binding, and TR·nTRE interaction. These results support a novel mechanism underlying T3-dependent repression.
It would be especially valuable to characterize the dual functions of NCoR, which appear to be dependent on the promoter context. Thus, on the positive TRE, NCoR acts as co-repressor, while on the negative TRE, as a co-activator, possibly by interacting with GAF and TR. When the C-terminal region of NCoR, which contains CoRNR boxes that interact with nuclear receptors for repression, was transiently expressed in COS-7 cells, the effect was the same as that generated by the full-length NCoR (data not shown). This raises the interesting question of whether the C-terminal domain of NCoR also contains TR and co-activator interaction domains for TR-mediated basal transactivation of T3-dependently negatively regulated genes.
Alternatively, the varying function of NCoR depending on the promoter context may be explained by its acting as a bridging molecule, rather than as a co-activator or co-repressor. It is possible that the TR·nTRE complex has a high affinity to certain domains in the C-terminal NCoR to recruit co-activators such as GAF for enhanced transactivation, whereas TR·pTRE complex binds to CoRNR boxes of NCoR to induce repression.
Transient Interaction of TR with DNA during T3 Exposure— TR·DNA interaction on the CD44 promoter was also confirmed in vivo by ChIP assay. In the absence of T3, TR weakly but significantly interacted with DNA. Surprisingly, in the presence of T3, TR bound to DNA in a pattern of transient interaction at early stages of T3 stimulation. TR·DNA interaction peaked 50–60 min after the addition of T3 in vivo (Fig. 9D). TR-associated nuclear proteins are probably recruited in vivo onto the nTRE, enhancing TR·DNA interaction, with a peak 50–60 min after T3 exposure. If true, this suggests that the most important result of TR·DNA interaction was to initiate a repressed state in the chromatin structure for repressed function. This again implies that there was transient but strong direct TR·DNA interactions in vivo on the CD44 promoter, despite its lack of high affinity TRE binding sites. The generation of peak interaction at such an early stage of T3 stimulation is a novel phenomenon.
Other interesting questions remain to be answered in broadening our understanding of the T3-dependent repression mechanism. What is the basal mechanism underlying T3-dependent repression? How are histone molecules modified by acetylation, methylation, and phosphorylation? Which other chromatin remodeling proteins, such as FACTs (facilitates chromatin transcription) and NURFs (the ATP-dependent nucleosome remodeling factors), are recruited?
The working model we propose is depicted in Fig. 10. In the absence of T3, TR interacts weakly with nTRE (Fig. 10, part a). Without NCoR or GAF, TR·nTRE interaction allows basal expression. When NCoR and GAF are present, GAF binds to its recognition site at –1+3 bp and subsequently forms the TR·NCoR·GAF complex, containing the DNA loop structure that enhances transactivation (part b). At this stage, basal transactivation is further enhanced through the TR·nTRE interaction (part b). Addition of T3 causes repression through a transient TR·DNA interaction (parts c and d). After 50–60 min of T3 stimulation, TR·nTRE interaction is maximized, possibly recruiting chromatin remodeling proteins such as NCoR/SMRT, histone deacetylases, and NURFs (part c). As time goes on, chromatin structure is remodeled to its repressed state, and TR dissociates from DNA (part d). This state is maintained until the T3 level is reduced (part e). When the T3 level falls to hypothyroid concentrations, TR·nTRE interaction resumes, supporting basal transactivation (part a).
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To whom correspondence should be addressed: Thyroid Section, Division of Endocrinology, Diabetes and Hypertension, Dept. of Medicine, Brigham and Women's Hospital, Harvard Institute of Medicine, 77 Ave. Louis Pasteur, Boston, MA 02115. Tel.: 617-525-5710; Fax: 617-731-4718; E-mail: swkim{at}rics.bwh.harvard.edu or swkim58a{at}yahoo.com.
1 The abbreviations used are: T3, 3,5,3'-triiodothyronine; T4, thyroxine; 5'-FR, 5'-flanking region; TR, thyroid hormone receptor; TRE, thyroid hormone-responsive element; pTRE, positive TRE; nTRE, negative TRE; GAF, GAGA binding factor; TSS, transcription start site; NCoR, nuclear receptor corepressor; DBD, DNA binding domain; TSH, thyrotropin; PTC, papillary thyroid carcinoma; siRNA, small interference RNA; TKGH, TK promoter-growth hormone plasmid; hGH, human growth hormone; EMSA, electrophoretic mobility shift assay; cTR, chicken TR; ChIP, chromatin immunoprecipitation assay; RVS, Rous sarcoma virus; GFP, green fluorescent protein.
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ACKNOWLEDGMENTS |
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-Luc and hTR2-GS125 constructs. |
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