The Human Glucocorticoid Receptor (hGR) ß Isoform Suppresses the Transcriptional Activity of hGR{alpha} by Interfering with Formation of Active Coactivator Complexes

Evangelia Charmandari, George P. Chrousos, Takamasa Ichijo, Nisan Bhattacharyya, Alessandra Vottero, Emmanuil Souvatzoglou and Tomoshige Kino

Pediatric and Reproductive Endocrinology Branch (E.C., G.P.C., T.I., A.V., T.K.), National Institute of Child Health and Human Development, and Diabetes Branch (N.B.), National Institute of Diabetes & Digestive & Kidney Diseases, National Institutes of Health, Bethesda, Maryland 20892

Address all correspondence and requests for reprints to: Evangelia Charmandari, MD, Pediatric and Reproductive Endocrinology Branch, National Institute of Child Health and Human Development, National Institutes of Health, 10 Center Drive, Building 10, Room 9D42, Bethesda, Maryland 20892-1583. E-mail: charmane{at}mail.nih.gov.


    ABSTRACT
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
The human glucocorticoid receptor (hGR) ß, a splicing variant of the classic receptor hGR{alpha}, functions as a dominant-negative inhibitor of hGR{alpha}. We explored the mechanism(s) underlying this effect of hGRß by evaluating the interactions of this isoform with known steroid receptor coactivators. We found that hGRß suppressed the transcriptional activity of both activation function (AF)-1 and AF-2 of hGR{alpha}, indicating that hGRß may exert its dominant-negative effect by affecting the function of coactivators that are attracted to these transactivation domains. hGRß bound to one of the p160 coactivators, the glucocorticoid receptor-interacting protein 1 (GRIP1) via its preserved AF-1 but not via its defective AF-2 in vitro. In a chromatin immunoprecipitation assay, hGRß prevented coprecipitation of GRIP1 with hGR{alpha} tethered to glucocorticoid response elements of the endogenous tyrosine aminotransferase promoter, whereas deletion of the AF-1 of hGRß abolished this effect. In further experiments, overexpression of GRIP1 attenuated the suppressive effect of hGRß on hGR{alpha}-mediated transactivation of the mouse mammary tumor virus promoter. Competition for binding to glucocorticoid response elements or heterodimerization with hGR{alpha} via the D loop dimerization interface occurred, but they were not necessary for the suppressive effect of hGRß on the transcriptional activity of hGR{alpha}. Our findings suggest that hGRß suppresses the transcriptional activity of hGR{alpha} by competing with hGR{alpha} for binding to GRIP1, and possibly other p160 coactivators, through its preserved AF-1. These findings suggest that participation of hGRß in the formation of a coactivator complex renders this complex ineffective.


    INTRODUCTION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
GLUCOCORTICOIDS REGULATE a variety of biologic processes and exert profound influences on many physiologic functions by virtue of their diverse roles in growth, development, and maintenance of basal and stress-related homeostasis (1, 2). At the cellular level, their actions are mediated by a 94-kDa intracellular receptor protein, the glucocorticoid receptor (GR), which belongs to the superfamily of steroid/thyroid/retinoic acid receptor proteins that function as ligand-dependent transcription factors (3, 4, 5, 6). Alternative splicing of the human (h) GR gene in exon 9 generates two highly homologous receptor isoforms, termed {alpha} and ß. These are identical through amino acid 727, but then they diverge, with hGR{alpha} having an additional 50 amino acids and hGRß having an additional, nonhomologous 15 amino acids (7) (Fig. 1AGo).



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Fig. 1. The Human GR

A, Genomic DNA and cDNA, and protein structures of the hGR. The hGR gene consists of 10 exons. Exon 1 is an untranslated region, exon 2 codes for the NTD, exons 3 and 4 for the DBD, and exons 5–9 for the hinge region and the LBD. The glucocorticoid receptor gene contains two terminal exons 9 (exon 9{alpha} and 9ß) alternatively spliced to produce the hGR{alpha} and hGRß isoforms. B, Schematic representation of the plasmids used for the experiments described in this study. {cjs2110}, GAL4 DBD; {blacksquare}, hGR DBD; *, the A458T mutation introduced in the DBD of hGRß; Deletions are indicated by single horizontal lines.

 
hGR{alpha} is ubiquitously expressed in almost all human tissues and cells and represents the classic GR that functions as a ligand-dependent transcription factor. In the absence of ligand, hGR{alpha} resides mostly in the cytoplasm of cells as part of a large multiprotein complex (8). Upon hormone binding, hGR{alpha} translocates into the nucleus, where it binds as a homodimer to glucocorticoid response elements (GREs) and activates or represses the transcription of glucocorticoid-responsive genes in a hormone-dependent manner (9). The receptor may also modulate transcription independently of binding to GREs, by physically interacting with other transcription factors, such as activator protein-1 (AP-1) and nuclear factor-{kappa}B (NF-{kappa}B) (10, 11). Two regions of hGR{alpha} possess intrinsic transcriptional activation function (AF): AF-1, which is located at the amino-terminal domain (NTD) and is glucocorticoid independent, and AF-2, which is located at the carboxyl-terminal domain [ligand binding domain (LBD)] and is glucocorticoid dependent (5). To initiate transcription, hGR{alpha} uses its transcriptional activation domains as surfaces to recruit chromatin remodeling factors or adapter proteins, termed coactivators, that serve to link response element-bound transcription factors, such as the hGR{alpha}, with general transcription factors (GTFs) of the RNA polymerase complex (12, 13, 14). Coactivators include the histone acetyltransferase family of proteins and the switching/sucrose nonfermenting family. A well-characterized group of histone acetyltransferase coactivators is the p160 family of proteins, which includes the steroid receptor coactivator 1 and the GR-interacting protein 1 (GRIP1). The p160 coactivators interact directly with both the AF-1 and the AF-2 of hGR{alpha}, respectively, through their carboxyl-terminal domain and multiple amphipathic LXXLL helical motifs located in their nuclear receptor-binding domain (NRB), forming coactivator complexes (15, 16, 17). In addition to coactivators, there are corepressors, such as the nuclear receptor corepressor and silencing mediator of retinoid and thyroid hormone receptor, which function in opposition to coactivators, retaining the hGR{alpha} in a transcriptionally inactive state. These corepressors recruit and activate complexes with histone deacetylation function, forming hGR{alpha} transrepression complexes (17, 18). The deacetylation of core histones enhances nucleosomal condensation of chromatin and supercoiling of DNA, thus preventing transcription factors from accessing DNA and interfering with the initiation of transcription.

hGRß is a 742-amino acid polypeptide also ubiquitously expressed in tissues, usually at lower concentrations than hGR{alpha}, with the exception of epithelial cells and neutrophils (19, 20, 21, 22). In contrast with hGR{alpha}, hGRß interacts poorly with heat-shock proteins, resides primarily in the nucleus of cells independently of the presence of ligand, does not bind glucocorticoids or antiglucocorticoids, and is transcriptionally inactive (23, 24, 25). hGRß exerts a dominant-negative effect on hGR{alpha}, inhibiting hGR{alpha}-mediated transactivation of target genes in a dose-dependent manner (19, 20, 21, 22, 26). The mechanism(s) underlying this inhibition have not been fully elucidated but may involve competition between hGR{alpha} and hGRß for binding to GREs, formation of hGR{alpha}-hGRß heterodimers that are transcriptionally inactive, and/or titration or squelching of coactivators necessary for hGR{alpha} transcriptional activation (19, 20, 21, 22, 26). Recent studies have localized the dominant-negative activity of hGRß to two amino acids within its unique carboxyl-terminal region and demonstrated that nuclear localization is a critical feature of its dominant-negative activity (27). The ability of hGRß to antagonize the function of hGR{alpha} suggests that hGRß may play a critical role in regulating target tissue sensitivity to glucocorticoids (28, 29, 30, 31, 32). Increased expression of hGRß has been documented in generalized and tissue-specific glucocorticoid resistance and leads to a reduction in the ability of hGR{alpha} to bind to GREs (28, 29, 30, 31). Therefore, an imbalance in hGR{alpha} and hGRß expression may underlie the pathogenesis of several clinical conditions associated with glucocorticoid resistance, such as rheumatoid arthritis, systemic lupus erythematosus, or ulcerative colitis (33).

In the present study, we examined further the molecular mechanisms underlying the dominant-negative effect of hGRß on the transcriptional activity of hGR{alpha}. Our findings suggest that these mechanisms primarily involve competition at the level of coactivators, whereas heterodimerization via the carboxyl-terminal domain dimerization interface and the formation of transcriptionally inactive or weakly active hGR{alpha}-hGRß heterodimers might be additional factors contributing to this process.


    RESULTS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
hGRß Suppresses the Transcriptional Activity of hGR{alpha} Independently of Binding to GREs
To examine whether hGRß exerts its dominant-negative activity on hGR{alpha} by competing with hGR{alpha} for binding to GREs, we employed the pRSVG-gal-G plasmid, in which the DNA binding domain (DBD) of hGR{alpha} has been replaced by the GAL4-DBD (Fig. 1BGo). Therefore, this chimeric construct expresses the amino-terminal and carboxyl-terminal domains of hGR{alpha} and has the ability to stimulate the transcription of the GAL4-responsive promoter 17mer-tk in response to dexamethasone. CV-1 cells were transfected with pRSVG-gal-G or GAL4-SP1 (0.05 µg/well), p17mer-tk-luc (0.5 µg/well), and pSV40-ß-gal (0.1 µg/well). G-gal-G stimulated the transcriptional activity of 17mer-tk by 10-fold in response to dexamethasone. Coexpression of hGRß (0.5 µg/well) suppressed the dexamethasone-induced transactivation of G-gal-G by 2-fold, whereas it had no effect on the transcriptional activity of GAL4-SP1 (Fig. 2Go). These findings indicate that hGRß may suppress the transcriptional activity of hGR{alpha} independently of binding to GREs.



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Fig. 2. hGRß Suppresses the Transcriptional Activity of hGR{alpha} Independently of Binding to GREs

CV-1 cells were transfected with pRSVG-gal-G or GAL4-SP1, p17mer-tk-luc, and pSV40-ß-gal. Coexpression of hGRß decreased the transcriptional activity of G-gal-G in response to dexamethasone by 2-fold but had no effect on the transcriptional activity of GAL4-SP1. Bars represent mean ± SEM of at least three independent experiments.

 
hGRß Suppresses both AF-1 and AF-2 of hGR{alpha}
Having demonstrated that hGRß suppresses the transcriptional activity of G-gal-G, which expresses both the NTD and LBD of hGR{alpha}, we examined the effect of hGRß on either of those domains separately. In CV-1 cells transfected with GAL4-GR-NTD or GAL4-GR{alpha}-LBD (0.05 µg/well) (Fig. 1BGo), p17mer-tk-luc (0.5 µg/well) and pSV40-ß-gal (0.1 µg/well), coexpression of hGRß (0.5 µg/well) suppressed the transcriptional activity of both GAL4-GR-NTD (Fig. 3AGo) and GAL4-GR{alpha}-LBD (Fig. 3BGo). These results suggest that hGRß suppresses the transcriptional activity of both the NTD and LBD of hGR{alpha} independently, and may, therefore, exert a suppressive effect on both AF-1 and AF-2 of hGR{alpha}.



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Fig. 3. The Effect of hGRß on the Transcriptional Activity of (A) the NTD and (B) Carboxyl-Terminal Domain (LBD) of hGR{alpha}

CV-1 cells were transfected with GAL4-GR-NTD or GAL4-GR{alpha}-LBD. Coexpression of hGRß decreased the transcriptional activity of both the NTD and LBD of hGR{alpha}. C, The effect of hGRß on AF-1 and AF-2 of hGR{alpha}. CV-1 cells were transfected with plasmids expressing the wild-type (WT) hGR{alpha} or any of the mutants hGR{alpha}({Delta}9–385), hGR{alpha}({Delta}204–290) and hGR{alpha}(1–515). Coexpression of hGRß suppressed the transcriptional activity of both AF-1 and AF-2 of hGR{alpha}. Bars represent mean ± SEM of at least three independent experiments.

 
To test the latter hypothesis further, we employed a panel of hGR{alpha} mutants, in which either AF-1 or AF-2 is deleted, and tested the effect of hGRß on their transactivation of the mouse mammary tumor virus (MMTV) promoter. CV-1 cells were transfected with pMMTV-luc (0.5 µg/well), pSV40-ß-gal (0.1 µg/well) and pRShGR{alpha} (0.05 µg/well) or any of the following hGR{alpha}-related constructs: pRShGR{alpha}({Delta}9–385), pRShGR{alpha}({Delta}204–290), and pRShGR{alpha}(1–515) (0.05 µg/well). Of those, pRShGR{alpha}({Delta}9–385) and pRShGR{alpha}({Delta}204–290) have a deletion over the AF-1 domain, whereas pRShGR{alpha}(1–515) lacks the entire carboxyl-terminal domain and, hence AF-2. Coexpression of hGRß (0.5 µg/well) decreased the transcriptional activity of both the AF-1- and AF-2-defective hGR{alpha} expressing plasmids (Fig. 3CGo), indicating that hGRß suppresses the activity of both AF-1 and AF-2 of hGR{alpha} independently.

hGRß Displays a Distinct Interaction with the GRIP1 Coactivator in Vitro and Prevents Tethering of GRIP1 to hGR{alpha} in Vivo
In view of the suppressive effect of hGRß on both AF-1 and AF-2 of hGR{alpha}, and the importance of AF-1 and AF-2 for interaction with coactivators in initiating transcription, we explored the interaction among hGR{alpha}, hGRß, and one of the p160 coactivators, the GRIP1 coactivator. GRIP1 contains two sites that bind to hGR{alpha}: one site, the NRB site, is located between amino acids 542 and 745 and contains three LXXLL signature motifs through which GRIP1 interacts with the AF-2 of hGR{alpha} in a ligand-dependent fashion; the other site is located at the carboxyl terminus, between amino acids 1121 and 1250, and binds to the AF-1 of hGR{alpha} in a ligand-independent fashion (34, 35, 36). An additional binding site for hGR{alpha}, termed auxiliary nuclear receptor-interacting domain (NIDaux), is located between the NRB site and the carboxyl-terminal AF-1 binding site (37) (Fig. 4AGo).



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Fig. 4. hGRß Competes with hGR{alpha} for Binding to GRIP1 via Its Preserved AF-1

A, Linearized GRIP1 molecule and distribution of its functional domains. AD1, Activation domain 1; AD2, activation domain 2; HLH, helix-loop-helix; NIDaux, auxiliary nuclear receptor-interacting domain; PAS, period arylhydrogen receptor and single-minded (adapted with permission from Vottero, A., et al. J Clin Endocrinol Metab 87:2658–2667, 2002 (55 ). © The Endocrine Society.] B and C, Interaction of hGR{alpha}, hGRß and hGRß({Delta}77–261) with GRIP1 in a GST pull-down assay. Both hGR{alpha} and hGRß bound to full-length and carboxyl-terminal fragments of GRIP1 independently of the presence of ligand. Although hGR{alpha} bound

 
We first performed a glutathione-S-transferase (GST)-pull-down assay to investigate the interaction of hGR{alpha} and hGRß with the full-length GRIP1 [GRIP1(1–1462)], its amino terminal fragment GRIP1(597–774), which contains the AF-2 interacting site, and its carboxyl-terminal fragment GRIP1(740–1217), which contains the AF-1-interacting site. Both hGR{alpha} and hGRß bound to full-length GRIP1(1–1462), with the former interaction being partially ligand-dependent and the latter interaction being ligand independent. Also, both hGR{alpha} and hGRß bound to GRIP1(740–1217) independently of ligand. However, although hGR{alpha} interacted with GRIP1(597–774) in a ligand-dependent fashion, there was no interaction between hGRß and GRIP1 (597–774) (Fig. 4BGo). Deletion of the AF-1 of hGRß abolished its interaction with GRIP1(740–1217) (Fig. 4CGo). These findings suggest that hGRß interacts with the GRIP1 coactivator in vitro only through its AF-1.

We subsequently investigated the effect of hGRß on the interaction between hGR{alpha} and GRIP1 on the GREs of the endogenous glucocorticoid-responsive tyrosine aminotransferase (TAT) promoter in a chromatin immunoprecipitation assay (ChIP). GRIP1 was successfully coprecipitated with TAT GREs via hGR{alpha} in a ligand-dependent fashion in the absence of hGRß (Fig. 4DGo, upper panel, lines 9–12). Coexpression of hGRß abolished this effect, and GRIP1 was no longer associated with GREs (Fig. 4DGo, upper panel, lines 15–18), whereas coexpression of hGRß({Delta}77–261) did not interfere with the coprecipitation of GRIP1 with the TAT GREs (Fig. 4DGo, upper panel, lanes 21–24). In the same experiment, hGR{alpha} coprecipitated with TAT GREs both in the absence and in the presence of hGRß or hGRß({Delta}77–261), although this association was weaker in the latter case (Fig. 4DGo, lower panel, lines 27–30, 39–42, 45–48). These results indicate that hGRß inhibits the accumulation of GRIP1 to the GRE-bound hGR{alpha} in vivo via its preserved AF-1, and that the AF-1 of hGRß plays an important role in its dominant-negative activity on hGR{alpha}-mediated transactivation of target genes. The dominant-negative effect of hGRß on the transcriptional activity of hGR{alpha} in H4IIE cells is illustrated in Fig. 4EGo.

The AF-1 Region of hGRß Is Necessary for its Dominant-Negative Activity upon the Transcriptional Activity of hGR{alpha}
To investigate further the role of the AF-1 of hGRß, we examined the effect of wild-type hGRß and its AF-1-defective mutant, hGRß({Delta}77–261), on the transcriptional activity of the NTD of hGR{alpha}. CV-1 cells were transfected with GAL4-GR-NTD (0.05 µg/well), p17mer-tk-luc (0.5 µg/well) and pSV40-ß-gal (0.1 µg/well). Coexpression of hGRß (0.5 µg/well) suppressed the transcriptional activity of GAL4-GR-NTD, whereas deletion of the AF-1 of hGRß abolished this effect (Fig. 5AGo, columns 2–4). These results indicate that the AF-1 of hGRß is necessary for its dominant-negative effect on hGR{alpha}-mediated transactivation of the GAL4-responsive promoter.



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Fig. 5. The Effect of the AF-1 of hGRß on Its Dominant-Negative Activity upon the Transcriptional Activity of hGR{alpha}

A, The effect of AF-1 of hGRß on GRIP1-enhanced transcriptional activity of hGR{alpha}. CV-1 cells were transfected with GAL4-GR-NTD, and a GRIP1-expressing plasmid. Coexpression of hGRß decreased both basal and GRIP1-enhanced hGR-NTD-mediated transactivation of the promoter, whereas deletion of the AF-1 of hGRß abolished this effect. Bars represent mean ± SEM of at least three independent experiments. B, hGRß, hGRßA458T and hGRß({Delta}77–261) were expressed in CV-1 cells. Expected molecular masses: hGRß WT and hGRßA458T: 94 kDa, hGRß({Delta}77–261): 66 kDa to the NRB fragment of GRIP1 in a ligand-dependent fashion, there was no such interaction between hGRß and GRIP1 (597–774). Furthermore, there was no interaction between the AF-1 defective mutant of hGRß and the carboxyl-terminal fragment of GRIP1. D, Effect of hGRß expression on the accumulation of GRIP1 to the hGR{alpha}-bound GREs in ChIP assay: hGR{alpha} was successfully coprecipitated with GRIP1 in a ligand-dependent fashion in the absence of hGRß. Coexpression of hGRß, but not its AF-1 defective mutant, interfered with binding of GRIP1 to hGR{alpha}. E, The effect of hGRß on the transcriptional activity of hGR{alpha} in H4IIE cells.

 
In the same experiment, overexpression of GRIP1 (0.05 µg/well) enhanced the transcriptional activity of GAL4-GR-NTD (Fig. 5AGo, columns 2 and 5), a finding consistent with previous reports on the association between GRIP1 and AF-1 domain of hGR{alpha} (37). hGRß suppressed the GRIP1-enhanced transcriptional activity of GAL4-GR-NTD, whereas deletion of the AF-1 of hGRß abolished this effect (Fig. 5AGo, columns 6 and 7). These findings suggest that hGRß inhibits the functional interaction between hGR{alpha} and the GRIP1 coactivator by binding to the carboxyl-terminal fragment of GRIP1 via its AF-1 domain, further underscoring the importance of AF-1 for the dominant-negative effect of hGRß on hGR{alpha}-mediated transactivation of glucocorticoid-responsive genes. Expression of hGRß and hGRß({Delta}77–261) in CV-1 cells is shown in Fig. 5BGo.

Overexpression of GRIP1 Attenuates the Dominant-Negative Effect of hGRß on the Transcriptional Activity of hGR{alpha}
We then examined whether GRIP1 attenuates the dominant-negative effect of hGRß on hGR{alpha}-mediated transactivation of the MMTV promoter. CV-1 cells were transfected with pRShGR{alpha} (0.05 µg/well), pMMTV-luc (0.5 µg/well) and pSV40-ß-gal (0.1 µg/well). Although hGRß (0.2 µg/well) suppressed the transcriptional activity of hGR{alpha} (Fig. 6Go, columns 1, 2, 5, and 6), GRIP1 (0.25 or 0.5 µg/well) enhanced it (Fig. 6Go, columns 1–4). Coexpression of hGRß and GRIP1 resulted in attenuation of the suppressive effect of hGRß on hGR{alpha}-mediated transactivation of the MMTV promoter (Fig. 6Go, columns 7 and 8), possibly by squelching limited amounts of hGRß by the overexpressed GRIP1. These results provide additional evidence that the dominant-negative effect of hGRß on the transcriptional activity of hGR{alpha} is dependent upon GRIP1 for interaction with hGR{alpha} and enhancement of its transcriptional activity.



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Fig. 6. The Effect of Overexpression of GRIP1 on the Dominant-Negative Activity of hGRß

CV-1 cells were transfected with hGR{alpha} and hGRß-expressing plasmids. Overexpression of GRIP1 [0.25 µg/well (+) or 0.5 µg/well (++)] resulted in attenuation of the hGRß suppressive effect on hGR{alpha}-mediated transactivation of the MMTV promoter. Bars represent mean ± SEM of at least three independent experiments.

 
Deletion of the LBD of hGRß Does Not Abolish Completely Its Dominant-Negative Effect on the Transcriptional Activity of hGR{alpha}
To examine whether the LBD of hGRß accounts for its dominant-negative activity on hGR{alpha}, CV-1 cells were transfected with pRShGR{alpha} (0.05 µg/well), pRShGRß or pRShGRß(1–559) (0.5 µg/well), pMMTV-luc (0.5 µg/well), and pSV40-ß-gal (0.1 µg/well) (Fig. 1BGo). hGRß(1–559), which lacks LBD, suppressed the hGR{alpha}-mediated transactivation of the MMTV promoter, but to a lesser extent than the wild-type hGRß, whereas this molecule, unlike GRß, had some agonist activity (Fig. 7Go). These results suggest that hGRß preserves its dominant-negative activity even when it is devoid of its LBD. However, the underlying mechanisms of the dominant-negative activity of a LBD-containing hGRß, and an LBD-devoid glucocorticoid receptor mutant may be distinct.



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Fig. 7. The Role of the LBD in the Dominant-Negative Activity of hGRß

CV-1 cells were transfected with pRShGR{alpha}. Coexpression of hGRß(1–559) or wild-type hGRß suppressed the hGR{alpha}-mediated transactivation of the MMTV promoter. hGRß(1–559) had some inherent transactivation activity. Bars represent mean ± SEM of at least three independent experiments.

 
Heterodimerization via the DBD-Dimerization Domain Is Not Necessary for the Suppressive Effect of hGRß on the Transcriptional Activity of hGR{alpha}
To investigate whether hGRß exerts its dominant-negative effect on hGR{alpha} via heterodimerization through the DBD, we introduced the A458T mutation, in which alanine is replaced by threonine at amino acid 458, into the pRShGRß plasmid to prevent the formation of hGR{alpha}-hGRß heterodimers. Wild-type hGRß (0.5 µg/well) suppressed the hGR{alpha}-mediated (0.05 µg/well) transactivation of the MMTV promoter, whereas the suppressive effect of hGRßA458T (0.5 µg/well) was less prominent (Fig. 8Go). These results indicate that heterodimerization of hGRß with hGR{alpha} via their DBD-dimerization interface is not necessary for the dominant-negative activity of hGRß. The expression of hGRßA458T in CV-1 cells is shown in Fig. 5BGo.



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Fig. 8. The Effect of Heterodimerization via the DBD-Dimerization Domain in the Dominant-Negative Activity of hGRß

CV-1 cells were transfected with pRShGR{alpha}. Coexpression of both hGRß and hGRßA458T suppressed the hGR{alpha}-mediated transactivation of the MMTV promoter, however, the suppressive effect of hGRßA458T was less prominent. Bars represent mean ± SEM of at least three independent experiments.

 

    DISCUSSION
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
We explored the molecular mechanisms underlying the dominant-negative activity of hGRß upon the transcriptional activity of hGR{alpha}. We have demonstrated that hGRß suppresses both the AF-1 and AF-2 of hGR{alpha}, interacts with the GRIP1 coactivator in vitro only through its AF-1 and prevents tethering of hGR{alpha} to GRIP1 in vivo, whereas overexpression of GRIP1 attenuates the dominant-negative effect of hGRß on the transcriptional activity of hGR{alpha}. Competition for binding to GREs or heterodimerization with hGR{alpha} via the DBD dimerization domain are not necessary for the suppressive effect of hGRß on hGR{alpha}-mediated transactivation of target genes. These findings indicate that the dominant-negative activity of hGRß is most likely due to competition with hGR{alpha} for binding to GRIP1 and possibly other p160 coactivators through its conserved AF-1, leading to the formation of an inactive glucocorticoid receptor-coactivator complex.

One of the previously proposed mechanisms for the dominant-negative activity of hGRß is competition with hGR{alpha} for binding to GREs, a phenomenon that occurs in vitro (19). Here we demonstrated that binding to DNA is not necessary for the dominant-negative activity of hGRß, given that hGRß suppressed the transcriptional activity of G-gal-G, in which the DBD of hGR{alpha} is replaced by the DBD of GAL4. This finding is also supported by the fact that overexpression of hGRß does not significantly affect the transactivation of the progesterone (PR) or androgen (AR) receptors by similar hormone-response elements (26). Within the steroid/thyroid/retinoic receptor superfamily, the GRs, mineralocorticoid receptors (MRs), PRs, and ARs comprise a family based on sequence conservation (38). The DBDs of these receptors are highly conserved, and as a result, each receptor recognizes the same or similar hormone response elements (HREs) (39). That hGRß only weakly represses the PR and AR transcriptional activity, despite its ability to bind with their HREs, suggests that its inhibitory effect on hGR{alpha} transcriptional activation does not result simply from competition for binding to the same response elements, especially so because these receptors have lower affinity for these HREs than hGR{alpha} (40). In addition, preliminary analysis of mutations in the DBD shows no effect on the ability of hGRß to function as a dominant-negative inhibitor of hGR{alpha} (27).

Like other transcriptional modulators, hGR{alpha} uses its transcriptional activation domains AF-1 and AF-2 as surfaces to recruit coactivators and chromatin remodeling factors, and to interact with GTFs (41). AF-2, which maps to the carboxyl terminus, is glucocorticoid dependent, whereas AF-1 is located at the amino terminus of the receptor and is glucocorticoid independent. hGRß suppressed both AF-1 and AF-2 of hGR{alpha} in independent experiments of our study, possibly suggesting competition between hGRß and hGR{alpha} for binding to the same coactivators. Therefore, we investigated the in vitro interaction between hGRß and the GRIP1 coactivator. GRIP1 contains two sites that interact with the transactivation domains of hGR{alpha}: an AF-1-interacting site, which is located at the carboxyl-terminal domain of GRIP1, and an AF-2-interacting site located at its NRB domain (34, 35, 36). In a GST pull-down assay, hGRß bound to the full-length and carboxyl-terminal fragment of GRIP1, but not to the NRB fragment of GRIP1. On the other hand, however, the AF-1-defective mutant hGRß({Delta}77–261) displayed no interaction with the carboxyl-terminal fragment of GRIP1, indicating that hGRß interacts with GRIP1 through its AF-1 domain. In view of these findings, we investigated whether hGRß competes with hGR{alpha} for binding to GRIP1. In ChIP, GRIP1 was successfully coprecipitated with TAT GREs via hGR{alpha} in a ligand-dependent fashion in the absence of hGRß, whereas coexpression of hGRß, but not of hGRß({Delta}77–261), abolished this effect. Our findings indicate that competition of hGRß with hGR{alpha} for binding to GRIP1, or possibly other coactivators, might interfere with the initiation and propagation of transcription and may account for the dominant-negative activity of hGRß. In further experiments, overexpression of GRIP1 attenuated the inhibitory effect of hGRß on hGR{alpha}-mediated transactivation of the MMTV promoter, further underscoring the importance of these proteins in facilitating transcriptional enhancement of hGR{alpha}. These data are compatible with formation of an ineffective coactivator complex rendered inactive by the presence of GRß.

Granted that different steroid hormone receptors use different sets of coactivators for exerting their transcriptional activities (42, 43, 44), a specific, potent, dominant-negative effect of hGRß on hGR{alpha}, but not on PR or AR, might be so because of selective use of coactivators among these receptors. On the other hand, the great structural homology between hGR{alpha} and the human MR (45) may explain why hGRß suppresses the transcriptional activity of the latter quite potently compared with that of PR or AR, possibly by formation of an ineffective coactivator complex containing hGRß, MR, and a p160 coactivator molecule, in a fashion similar to the one we propose for its effect on hGR{alpha} (46, 47).

Finally, we demonstrated that heterodimerization of hGR{alpha} and hGRß (20) is not necessary for the suppressive effect of hGRß on the transcriptional activity of hGR{alpha}. The ability of the receptor to dimerize depends on the D loop, a stretch of five amino acids located in the carboxyl-terminal region of DBD. Several contacts by D loop residues at the dimerization interface stabilize receptor dimers, thereby allowing DNA binding (48, 49). A mutant hGR{alpha} with an amino acid substitution located in the D loop (A458T) fails to bind DNA and cannot transactivate GRE-dependent promoters (50, 51). In addition to the D loop in the DBD, another region in the LBD has also been shown to be involved in homodimerization of the receptor. This domain is located near the carboxyl terminus and consists of a heptad repeat of hydrophobic residues and additional ones located in intermediate positions (52, 53). Interestingly, hGR{alpha} and hGRß diverge in the middle of this latter dimerization domain. Therefore, it is likely that the preservation of an intact proximal half of the carboxyl-terminal dimerization domain in hGRß, despite the presence of the A458T mutation, still allows the formation of transcriptionally impaired hGR{alpha}-hGRß heterodimers, which might explain the suppressive effect of hGRßA458T on hGR{alpha}-mediated transactivation of the MMTV promoter in our study. This concept is further supported by the recently published crystal structure of the LBD of hGR{alpha}, which suggests that this dimerization interface involves H4 and ß-strands 3 and 4, located on the opposite face of the folded domain from H10 to H12. Therefore, the structural changes in H11 and H12 of hGRß would not be expected to perturb the dimer interface found in hGR{alpha} or to interfere with the hGR{alpha}-hGRß heterodimerization (54).

Recent studies have demonstrated that the 15 unique amino acid sequence and the absence of H12 in hGRß account for its dominant-negative activity (26, 27). The absence of H12 renders the receptor unable to bind ligand or transactivate reporter genes, whereas the 15 unique amino acids help keep the receptor constitutively localized in the nucleus. Both nuclear localization of hGRß and physical interaction with hGR{alpha} are important prerequisites for maximum dominant-negative activity (27). Interestingly, the dominant-negative function of hGRß was localized in just two amino acids within the unique hGRß sequence (27). A possible explanation for these findings would be that these two amino acids are important for interaction with corepressor molecule(s) or other factors involved in transrepression. Indeed, upstream deletions of the LBD of hGRß, including these two amino acids, lead to loss of dominant-negative activity (27). Furthermore, we have documented that the pathologic natural hGR{alpha} mutants hGR{alpha}I559N, hGR{alpha}I747M and hGR{alpha}L773P also exert a dominant-negative effect upon the wild-type receptor (25, 55, 56). Therefore, it is possible that the amino acid residues reported by Yudt et al. (26, 27) may not be the only ones to account for the dominant-negative activity of hGRß, and that a change in the structure of hGRß LBD may influence the transcriptional activity of its NTD and/or knock out the interaction between hGRß and GRIP1.

We conclude that the mechanisms underlying the dominant-negative activity of hGRß upon the transcriptional activity of hGR{alpha} primarily involve competition at the level of coactivators, whereas heterodimerization via the carboxyl-terminal dimerization domain and the formation of transcriptionally inactive or weakly active hGR{alpha}-hGRß heterodimers might be additional factors contributing to this process. Competition for binding to GREs or heterodimerization with hGR{alpha} via the DBD dimerization domain is not necessary for the suppressive effect of hGRß on the transcriptional activity of hGR{alpha}. Further studies are required to explore the interaction between hGRß and other coactivators that enhance the transcriptional activity of hGR{alpha}, as well as the role of hGRß in accentuating the inhibitory effect of corepressors on hGR{alpha}-mediated transactivation of target genes.


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 
Plasmids (Fig. 1BGo)
The plasmids pRShGR{alpha} and pRShGRß express the hGR{alpha} and hGRß isoform, respectively, under the control of the Rous sarcoma virus (RSV) promoter. The plasmids pRShGR{alpha}({Delta}9–385), pRShGR{alpha}({Delta}204–290), and pRShGR{alpha}(1–515) were constructed from the pRShGR{alpha} plasmid after deletion ({Delta}) of the regions 9–385, 204–290, and 516–777, respectively, and they express the corresponding fragments of hGR{alpha}. Of those, pRShGR{alpha}({Delta}9–385) and pRShGR{alpha}({Delta}204–290) lack the AF-1 domain, whereas pRShGR{alpha}(1–515) lacks almost the entire LBD and, therefore, the AF-2 domain. The pRSVG-gal-G plasmid was constructed by replacing the DBD of hGR{alpha} by the DBD of the yeast transcription factor GAL4 and expresses the NTD and LBD of hGR{alpha} (Fig. 1BGo). All plasmids were gifts from Dr. R. M. Evans (The Salk Institute, La Jolla, CA).

The pRShGRßA458T plasmid was constructed by introducing the A458T mutation into the pRShGRß plasmid using PCR-assisted site-directed mutagenesis (Stratagene, La Jolla, CA). The A458T mutation, in which alanine is replaced by threonine at amino acid position 458, impairs dimerization and, therefore, GRE-dependent transactivation of hGR{alpha}, but it does not interfere with functions that involve cross talk with other transcription factors (57). The pRShGRß({Delta}77–261) and pBK/CMV-hGRß({Delta}77–261) plasmids were constructed by digestion of pRShGRß and pBK/CMV-hGRß, respectively, using the enzyme BglII, followed by auto-ligation, which results in deletion of cDNA sequence of amino acids 77–261 of these plasmids. The plasmid pRSV-erbA–1, which contains a thyroid receptor cDNA in inverse orientation, was used as a negative control for all hGR{alpha}- and hGRß-related plasmids (Dr. R. M. Evans) (Fig. 1BGo).

The plasmids GAL4-GR{alpha}-LBD and GAL4-GR-NTD express, respectively, the hGR{alpha}-LBD (amino acids 490–777) and hGR-NTD (amino acids 1–420); they were constructed by subcloning the corresponding nucleotides of hGR into the pM vector (CLONTECH, Palo Alto, CA) to produce GAL4-GR fusion proteins (Fig. 1BGo). GAL4-SP1, which expresses the GAL4-DBD-fused SP1 transcription factor, was used as a negative control for all GAL4-GR-related plasmids and was a gift from Dr. Y. Shi (Harvard Medical School, Boston, MA).

The plasmids pGEX4T3-GRIP1(1–1462), pGEX4T3-GRIP1(597–774), and pGEX4T3-GRIP1(740–1217), which express the GST-fusion proteins GRIP1(1–1462) (full-length of GRIP1), GRIP1(597–774) (NRB fragment of GRIP1) and GRIP1(740–1217) (carboxyl-terminal fragment of GRIP1), respectively, were constructed by subcloning the corresponding cDNA fragments of GRIP1 into the pGEX4T3 plasmid (Amersham Pharmacia Biotech, Piscataway, NJ) as previously described (34, 36, 37). The pSG5-GRIP1(1–1462) plasmid was a gift from Dr. M. G. Stallcup (University of Southern California, Los Angeles, CA). The vector pSG5 (Stratagene) was used as negative control for the GRIP1-related constructs.

The pMMTV-luc plasmid, which expresses luciferase under the control of the glucocorticoid-responsive MMTV promoter, was a gift from Dr. G. L. Hager (National Cancer Institute, National Institutes of Health, Bethesda, MD). The p17mer-tk-luc contains the luciferase gene under the control of the four GAL4-response elements cloned upstream the proximal portion of the herpes simplex virus thymidine kinase promoter, and was a gift from Dr. M. J. Tsai (Baylor College of Medicine, Houston, TX). The pSV40-ß-gal encodes the ß-galactosidase gene under the control of simian virus (SV) 40 promoter (Promega, Madison, WI).

Cell Cultures
CV-1 and COS-7 embryonic African green monkey kidney cells, and H4IIE rat hepatoma cells were grown in DMEM supplemented with 10% fetal bovine serum and antibiotics. Cells were incubated in a humidified atmosphere of 5% CO2 at 37 C and passaged every 3–4 d. Twenty-four hours before transfection, subconfluent cells were removed from their flasks by trypsinization, resuspended in supplemented medium and plated in six-well plates (CV-1, H4IIE), 75 cm2 flasks (COS-7) or 150-mm-diameter dishes (H4IIE) at a concentration of 1.5 x 105 cells/well, 1 x 106 cells/flask and 2.5 x 106 cells/dish, respectively.

Transient Transfection Assays
CV-1 and COS-7 cells were transfected using the lipofectin method (Invitrogen Life Technologies, Gaithersburg, MD) as previously described (58). H4IIE cells were transfected using FuGENE 6 reagent according to the instructions of the manufacturer (Roche Diagnostics Corp., Indianapolis, IN).

Transactivation Assays
CV-1 or H4IIE cells were seeded in six-well plates at a concentration of 1.5 x 105 cells/well. Twenty-four hours later, cells were cotransfected with pRShGR{alpha} or hGR-related plasmids (0.05 µg/well), pMMTV-luc or 17mer-tk-luc (0.5 µg/well) and pSV40-ß-gal (0.1 µg/well). In experiments designed to examine the effect of hGRß on the transcriptional activity of hGR{alpha}, pRShGRß, or hGRß-related plasmid(s) were added to the transfection medium at a concentration of 0.5 µg/well. In experiments designed to examine the effect of GRIP1 on the transcriptional activity of hGR{alpha}, pSG5-GRIP1(1–1462) was added to the transfection medium at a concentration of 0.25–0.5 µg/well. The plasmids pRSV-erbA–1, pM and pSG5 were used as negative controls in the appropriate experiments to maintain a constant amount of DNA per well. Twenty-four hours (CV-1) or 6 h (H4IIE) after transfection, the transfection medium was replaced with supplemented DMEM. After a further 24-h period, dexamethasone (Sigma Chemical Co., St. Louis, MO) or vehicle (100% ethanol) was added to the medium at a concentration of 10–6 M.

Luciferase and ß-Galactosidase Assays
Seventy-two hours after transfection, cells were washed with PBS twice, and lysed using a reporter lysis buffer (Promega). Luciferase activity in the cell lysates was determined in a Monolight 3010 Luminometer (BD PharMingen, San Diego, CA) as previously described (59). ß-Galactosidase activity was determined in the same samples using a ß-galactosidase enzyme assay system (Galacto-Light Plus, Tropix, Bedford, MA) according to the instructions of the manufacturer. Luciferase activity was divided by ß-galactosidase activity to account for transfection efficiency. All experiments were repeated at least three times.

Western Blot Analyses
CV-1 and COS-7 cells were seeded in 75 cm2 flasks at a concentration of 1 x 106 cells/flask and grown in supplemented DMEM. Subconfluent cells were transfected with pRShGRß, pRShGRßA458T, pRShGRß({Delta}77–261), GAL4-GR{alpha}-LBD, or GAL4-GR-NTD (20 µg/flask) using the lipofectin method. Twenty-four hours (CV-1) or 6 h (COS-7) later, the transfection medium was replaced with supplemented DMEM. Thirty hours after transfection, cells were rinsed with ice-cold PBS (three times), gently scraped from flasks, centrifuged briefly, and lysed using a lysis buffer that consisted of 100 mM Tris-HCl (pH: 8.5), 250 mM NaCl, 1% Nonidet P-40 (pH 7.2) and protease inhibitors (1 Tab/50 ml of Complete; Roche Applied Science, Indianapolis, IN). The homogenates were centrifuged (11,000 rpm at 4 C) for 30 min to obtain whole cell extracts. Whole cell extracts were mixed with Tris-glycine sodium dodecyl sulfate (SDS) sample buffer (2x) (Invitrogen Life Technologies, Carlsbad, CA), heated to 95 C for 3–5 min, and electrophoresed alongside molecular weight prestained markers (SeeBlue, Novex) through 8% Tris-glycine gel (Invitrogen Life Technologies). After electroblotting (25 V/0.8 mA/cm2) onto Hybond C membranes (Amersham Pharmacia Biotech, Buckinghamshire, UK), proteins were incubated with blocking solution (5% milk powder/PBS/0.05% Tween 20) for 4 h. Immunoblotting was performed at 4 C overnight. hGRß, hGRßA458T, and hGRß({Delta}77–261) were detected using a purified specific rabbit polyclonal anti-GRß antibody (Affinity BioReagents, Golden, CO) at 10 µg/ml. GAL4-GR{alpha}-LBD, and GAL4-GR-NTD were detected by an anti-GAL4-DBD antibody (Santa Cruz Biotechnology Inc., Santa Cruz, CA) (10 µg/ml). After washing with PBS three times, membranes were incubated with horseradish peroxidase-conjugated goat antirabbit IgG at 1:1000 dilution at room temperature for 1 h. hGRß, hGRßA458T, hGRß({Delta}77–261), GAL4-GR{alpha}-LBD, and GAL4-GR-NTD were visualized using the ECL Plus Western Blotting Detection System (Amersham Pharmacia Biotech) and exposed to high performance chemiluminescence film (Hyperfilm ECL, Amersham Pharmacia Biotech).

GST Pull-Down Assay
GST-fusion protein expression vectors [GST-fused GRIP1(1–1462), GRIP1(559–774) and GRIP1(740–1217)] were grown in Escherichia coli BL21 cells for 3–4 h and induced with IPTG (0.5–1 mM) for an additional 2 h. Cells were resuspended in 1/50 volume in PBS and extracts were generated by pulse sonication on ice (10 sec). Cell debris was then pelleted at 10,000 rpm for 10 min at 4 C. Glutathione-Sepharose beads (500 µl) were washed with PBS and the supernatant (5 ml) obtained from the cell extracts was allowed to bind by gentle agitation in PBS for 5 min. The beads with the bound GST fusion protein were then pelleted and washed three times with 5 ml of the same buffer and analyzed by SDS-PAGE for amount of protein bound to the GST beads.

Coupled in vitro transcription/translation reactions (TNT Quick Coupled Transcription/Translation System, Promega) were used to produce 35S-labeled hGR{alpha}, hGRß, and hGRß({Delta}77–261) in rabbit reticulocyte lysate by using pBK/CMV-hGR{alpha}, pBK/CMV-hGRß and pBK/CMV-hGRß({Delta}77–261), respectively. 35S-labeled hGR{alpha} hGRß and hGRß({Delta}77–261) (20 µl of crude translated protein) were incubated with GST fusion proteins bound to Glutathione-Sepharose beads (Amersham Pharmacia Biotech) for 2–3 h at 4 C in binding buffer [50 mM Tris (pH 7.2), 1 mM EDTA, 1 mM dithiothreitol, 150 mM NaCl, and 0.1% Triton X-100], washed, eluted and fractionated by SDS-PAGE. Samples were loaded and electrophoresed on an 8% SDS-PAGE gel. The percent of starting material loaded in input lanes was 10 or 20%. The gel was fixed, treated with Enlightning buffer (NEN Life Science Products, Inc., Boston, MA) and dried. Radioactivity was detected by exposing a film on the gel (60).

ChIP Assay
H4IIE rat hepatoma cells were seeded in 150-mm diameter dishes at a concentration of 2.5 x 106 cells/dish and grown in supplemented DMEM. Subconfluent cells were transfected using FuGENE 6 reagent according to the instructions of the manufacturer (Roche Diagnostics Corp.). Cells were cotransfected with pRShGR{alpha} (15 µg/dish) and pSG5-GRIP1(1–1462) (15 µg/dish) in the presence or absence of pRShGRß or pRShGRß({Delta}77–261) (15 µg/dish). Four hours later, the transfection medium was replaced with supplemented DMEM. Sixteen hours after transfection, cells were treated with dexamethasone (10–6 M) or vehicle (100% ethanol) for 24 h. Cells were fixed and cross-linked with 1% formaldehyde for 10 min, harvested and suspended in ice-cold lysis buffer [10 mM Tris-HCl (pH 7.4), 3 mM CaCl2, 2 mM MgCl2]. Swollen cells were resuspended with equal volumes of lysis buffer and Nonidet P-40 (NP-40) lysis buffer [10 mM Tris-HCl (pH 7.4), 10 mM NaCl, 3 mM MgCl2, 0.5% NP-40], homogenized in a Dounce homogenizer (Bellco Glass, Inc., Vineland, NJ), and centrifuged at 1500 rpm for 10 min at 4 C. Nuclear pellets were stored in a buffer containing 50 mM Tris-HCl (pH 8.0), 25% glycerol, 5 mM (CH3COO)2Mg, 0.1 mM EDTA, and 12 mM ß-mercaptoethanol at –80 C (61).

Equal amounts of nuclei (300 µg) were used for each immunoprecipitation experiment. Nuclei were diluted in 500 µl of ChIP dilution buffer [16.7 mM Tris-HCl (pH 8.1), 167 mM NaCl, 0.01% SDS, 1.1% Triton X-100, 1.2 mM EDTA, protease inhibitors], sonicated (Misonix sonicator, Farmingdale, NY) and centrifuged at 13,000 rpm for 5 min. Supernatants were precleared and the amount of chromatosomes present in each sample was measured at 260 nm. Equal amounts of chromatosomes (200 µg) were immunoprecipitated with the indicated antibodies for 14 h at 4 C. Immunocomplexes were captured on salmon sperm DNA/protein A-agarose (Upstate, Charlottesville, VA), washed sequentially with low salt wash buffer [20 mM Tris-HCl (pH 8.1), 150 mM NaCl, 0.1% SDS, 1% Triton X-100, 2 mM EDTA], high salt wash buffer [20 mM Tris-HCl (pH 8.1), 500 mM NaCl, 0.1% SDS, 1% Triton 10 X-100, 2 mM EDTA], LiCl wash buffer [10 mM Tris-HCl (pH 8.1), 0.25 M LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA] and twice with TE buffer [10 mM Tris-HCl (pH 8.0), 1 mM EDTA]. Samples were eluted with freshly prepared elution buffer [50 mM Tris-HCl (pH 8.1), 1 mM EDTA, 1% SDS] at 65 C for 10 min and, after adjusting the concentration of NaCl to 200 mM, cross-linking was reverted at 65 C for 4 h. After treatment with ribonuclease A (10 mg/ml) and proteinase K (20 mg/ml) for 1 h at 45 C, genomic DNA fragments were extracted with phenol/chloroform and precipitated with ethanol.

The antibodies used were anti-hGR{alpha} (Affinity Bioreagents, Golden, CO) and anti-GRIP1 (Santa Cruz Biotechnology). Primer sets were designed to amplify the rat tyrosine aminotransferase (TAT) promoter region, which contains tandem GREs and is located approximately 2500 bps upstream the transcription initiation site (forward primer: 5'-TCTTCTCAGTGTTCTCTATCAC-3'; reverse primer: 5'-CAGAAACCGACAGGC GACTACG-3') (62).

Equal volumes of DNA were used for PCR amplification of the rat TAT promoter region. Initiation was performed for 7 min at 94 C, followed by 30 cycles of denaturation at 94 C for 1 min, annealing at 50 C for 1 min and extension at 72 C for 1 min, and a final period of extension at 72 C for 7 min. PCR amplified products were electrophoresed on 2% agarose gel. All experiments were repeated at least three times.


    ACKNOWLEDGMENTS
 
The authors thank Drs. R. M. Evans (The Salk Institute, La Jolla, CA), L. P. Freedman, G. L. Hager (National Cancer Institute, National Institutes of Health, Bethesda, MD), Y. Shi (Harvard Medical School, Boston, MA), M. G. Stallcup (University of Southern California, Los Angeles, CA), and M. J. Tsai (Baylor College of Medicine, Houston, TX) for providing plasmids for some of the experiments described in this study.


    FOOTNOTES
 
First Published Online September 30, 2004

Abbreviations: AF, Activation function; AP-1, activator protein-1; AR, androgen receptor; ChIP, chromatin immunoprecipitation; DBD, DNA binding domain; GR, glucocorticoid receptor; GRE, glucocorticoid response element; GRIP1, glucocorticoid receptor-interacting protein 1; GST, glutathione-S-transferase; GTF, general transcription factor; hGR, human glucocorticoid receptor; HRE, hormone response elements; LBD, ligand binding domain; MMTV, mouse mammary tumor virus; MR, mineralocorticoid receptor; NF-{kappa}B, nuclear factor-{kappa}B; NP-40, Nonidet P-40; NRB, nuclear receptor-binding domain; NTD, amino-terminal domain; PR, progesterone receptor; RSV, Rous sarcoma virus; SDS, sodium dodecyl sulfate; TAT, tyrosine aminotransferase.

Received for publication March 16, 2004. Accepted for publication September 20, 2004.


    REFERENCES
 TOP
 ABSTRACT
 INTRODUCTION
 RESULTS
 DISCUSSION
 MATERIALS AND METHODS
 REFERENCES
 

  1. Tsai MJ, O’Malley BW 1994 Molecular mechanisms of action of steroid/thyroid receptor superfamily members. Annu Rev Biochem 63:451–486[CrossRef][Medline]
  2. Lazar, M.A. 2003. Mechanism of action of hormones that act on nuclear receptors. In: Larsen PR, Kronenberg HM, Melmed S, Polonsky KS, eds. Williams textbook of endocrinology. Philadelphia: W. B. Saunders; 35–44
  3. Carson-Jurica MA, Schrader WT, O’Malley BW 1990 Steroid receptor family: structure and functions. Endocr Rev 11:201–220[Abstract]
  4. Picard D, Yamamoto KR 1987 Two signals mediate hormone-dependent nuclear localization of the glucocorticoid receptor. EMBO J 6:3333–3340[Abstract]
  5. Hollenberg SM, Evans RM 1988 Multiple and cooperative transactivation domains of the human glucocorticoid receptor. Cell 55:899–906[Medline]
  6. Dalman FC, Scherrer LC, Taylor LP, Akil H, Pratt WB 1991 Localization of the 90-kDa heat shock protein-binding site within the hormone-binding domain of the glucocorticoid receptor by peptide competition. J Biol Chem 266:3482–3490[Abstract/Free Full Text]
  7. Lu NZ, Cidlowski JA 2004 The origin and functions of multiple human glucocorticoid receptor isoforms. Ann NY Acad Sci 1024:102–123[Abstract/Free Full Text]
  8. Pratt WB 1993 The role of heat shock proteins in regulating the function, folding, and trafficking of the glucocorticoid receptor. J Biol Chem 268:21455–21458[Free Full Text]
  9. Bamberger CM, Schulte HM, Chrousos GP 1996 Molecular determinants of glucocorticoid receptor function and tissue sensitivity to glucocorticoids. Endocr Rev 17:245–261[Abstract]
  10. Jonat C, Rahmsdorf HJ, Park KK, Cato AC, Gebel S, Ponta H, Herrlich P 1990 Antitumor promotion and antiinflammation: down-modulation of AP-1 (Fos/Jun) activity by glucocorticoid hormone. Cell 62:1189–1204[Medline]
  11. Scheinman RI, Gualberto A, Jewell CM, Cidlowski JA, Baldwin Jr AS 1995 Characterization of mechanisms involved in transrepression of NF-{kappa}B by activated glucocorticoid receptors. Mol Cell Biol 15:943–953[Abstract]
  12. McKenna NJ, Xu J, Nawaz Z, Tsai SY, Tsai MJ, O’Malley BW 1999 Nuclear receptor coactivators: multiple enzymes, multiple complexes, multiple functions. J Steroid Biochem Mol Biol 69:3–12[CrossRef][Medline]
  13. McKenna NJ, O’Malley BW 2002 Combinatorial control of gene expression by nuclear receptors and coregulators. Cell 108:465–474[Medline]
  14. Auboeuf D, Honig A, Berget SM, O’Malley BW 2002 Coordinate regulation of transcription and splicing by steroid receptor coregulators. Science 298:416–419[Abstract/Free Full Text]
  15. Heery DM, Kalkhoven E, Hoare S, Parker MG 1997 A signature motif in transcriptional co-activators mediates binding to nuclear receptors. Nature 387:733–736[CrossRef][Medline]
  16. Collingwood TN, Urnov FD, Wolffe AP 1999 Nuclear receptors: coactivators, corepressors and chromatin remodeling in the control of transcription. J Mol Endocrinol 23:255–275[Abstract/Free Full Text]
  17. Glass CK, Rosenfeld MG 2000 The coregulator exchange in transcriptional functions of nuclear receptors. Genes Dev 14:121–141[Free Full Text]
  18. Ito K, Barnes PJ, Adcock IM 2000 Glucocorticoid receptor recruitment of histone deacetylase 2 inhibits interleukin-1ß-induced histone H4 acetylation on lysines 8 and 12. Mol Cell Biol 20:6891–6903[Abstract/Free Full Text]
  19. Bamberger CM, Bamberger AM, de Castro M, Chrousos GP 1995 Glucocorticoid receptor ß, a potential endogenous inhibitor of glucocorticoid action in humans. J Clin Invest 95:2435–2441[Medline]
  20. de Castro M, Elliot S, Kino T, Bamberger C, Karl M, Webster E, Chrousos GP 1996 The non-ligand binding ß-isoform of the human glucocorticoid receptor (hGR ß): tissue levels, mechanism of action, and potential physiologic role. Mol Med 2:597–607[Medline]
  21. Oakley RH, Sar M, Cidlowski JA 1996 The human glucocorticoid receptor ß isoform. Expression, biochemical properties, and putative function. J Biol Chem 271:9550–9559[Abstract/Free Full Text]
  22. Oakley RH, Webster JC, Sar M, Parker Jr CR, Cidlowski JA 1997 Expression and subcellular distribution of the ß-isoform of the human glucocorticoid receptor. Endocrinology 138:5028–5038[Abstract/Free Full Text]
  23. Hollenberg SM, Weinberger C, Ong ES, Cerelli G, Oro A, Lebo R, Thompson EB, Rosenfeld MG, Evans RM 1985 Primary structure and expression of a functional human glucocorticoid receptor cDNA. Nature 318:635–641[Medline]
  24. Giguere V, Hollenberg SM, Rosenfeld MG, Evans RM 1986 Functional domains of the human glucocorticoid receptor. Cell 46:645–652[Medline]
  25. Kino T, Stauber RH, Resau JH, Pavlakis GN, Chrousos GP 2001 Pathologic human GR mutant has a transdominant negative effect on the wild-type GR by inhibiting its translocation into the nucleus: importance of the ligand-binding domain for intracellular GR trafficking. J Clin Endocrinol Metab 86:5600–5608[Abstract/Free Full Text]
  26. Oakley RH, Jewell CM, Yudt MR, Bofetiado DM, Cidlowski JA 1999 The dominant negative activity of the human glucocorticoid receptor ß isoform. Specificity and mechanisms of action. J Biol Chem 274:27857–27866[Abstract/Free Full Text]
  27. Yudt MR, Jewell CM, Bienstock RJ, Cidlowski JA 2003 Molecular origins for the dominant negative function of human glucocorticoid receptor ß. Mol Cell Biol 23:4319–4330[Abstract/Free Full Text]
  28. Leung DY, Hamid Q, Vottero A, Szefler SJ, Surs W, Minshall E, Chrousos GP, Klemm DJ 1997 Association of glucocorticoid insensitivity with increased expression of glucocorticoid receptor ß. J Exp Med 186:1567–1574[Abstract/Free Full Text]
  29. Shahidi H, Vottero A, Stratakis CA, Taymans SE, Karl M, Longui CA, Chrousos GP, Daughaday WH, Gregory SA, Plate JM 1999 Imbalanced expression of the glucocorticoid receptor isoforms in cultured lymphocytes from a patient with systemic glucocorticoid resistance and chronic lymphocytic leukemia. Biochem Biophys Res Commun 254:559–565[CrossRef][Medline]
  30. Longui CA, Vottero A, Adamson PC, Cole DE, Kino T, Monte O, Chrousos GP 2000 Low glucocorticoid receptor {alpha}/ß ratio in T-cell lymphoblastic leukemia. Horm Metab Res 32:401–406[Medline]
  31. Hauk PJ, Goleva E, Strickland I, Vottero A, Chrousos GP, Kisich KO, Leung DY 2002 Increased glucocorticoid receptor ß expression converts mouse hybridoma cells to a corticosteroid-insensitive phenotype. Am J Respir Cell Mol Biol 27:361–367[Abstract/Free Full Text]
  32. Hauk PJ, Hamid QA, Chrousos GP, Leung DY 2000 Induction of corticosteroid insensitivity in human PBMCs by microbial superantigens. J Allergy Clin Immunol 105:782–787[CrossRef][Medline]
  33. Chrousos GP 1995 The hypothalamic-pituitary-adrenal axis and immune-mediated inflammation. N Engl J Med 332:1351–1362[Free Full Text]
  34. Ma H, Hong H, Huang SM, Irvine RA, Webb P, Kushner PJ, Coetzee GA, Stallcup MR 1999 Multiple signal input and output domains of the 160-kilodalton nuclear receptor coactivator proteins. Mol Cell Biol 19:6164–6173[Abstract/Free Full Text]
  35. Ding XF, Anderson CM, Ma H, Hong H, Uht RM, Kushner PJ, Stallcup MR 1998 Nuclear receptor-binding sites of coactivators glucocorticoid receptor interacting protein 1 (GRIP1) and steroid receptor coactivator 1 (SRC-1): multiple motifs with different binding specificities. Mol Endocrinol 12:302–313[Abstract/Free Full Text]
  36. Webb P, Nguyen P, Shinsako J, Anderson C, Feng W, Nguyen MP, Chen D, Huang SM, Subramanian S, McKinerney E, Katzenellenbogen BS, Stallcup MR, Kushner PJ 1998 Estrogen receptor activation function 1 works by binding p160 coactivator proteins. Mol Endocrinol 12:1605–1618[Abstract/Free Full Text]
  37. Hong H, Darimont BD, Ma H, Yang L, Yamamoto KR, Stallcup MR 1999 An additional region of coactivator GRIP1 required for interaction with the hormone-binding domains of a subset of nuclear receptors. J Biol Chem 274:3496–3502[Abstract/Free Full Text]
  38. Evans RM 1988 The steroid and thyroid hormone receptor superfamily. Science 240:889–895[Medline]
  39. Ham J, Thomson A, Needham M, Webb P, Parker M 1988 Characterization of response elements for androgens, glucocorticoids and progestins in mouse mammary tumour virus. Nucleic Acids Res 16:5263–5276[Abstract]
  40. Rundlett SE, Miesfeld RL 1995 Quantitative differences in androgen and glucocorticoid receptor DNA binding properties contribute to receptor-selective transcriptional regulation. Mol Cell Endocrinol 109:1–10[CrossRef][Medline]
  41. Chen JL, Attardi LD, Verrijzer CP, Yokomori K, Tjian, R 1994 Assembly of recombinant TFIID reveals differential coactivator requirements for distinct transcriptional activators. Cell 79:93–105[Medline]
  42. Bramlett KS, Wu Y, Burris TP 2001 Ligands specify coactivator nuclear receptor (NR) box affinity for estrogen receptor subtypes. Mol Endocrinol 15:909–922[Abstract/Free Full Text]
  43. McInerney EM, Rose DW, Flynn SE, Westin S, Mullen TM, Krones A, Inostroza J, Torchia J, Nolte RT, Assa-Munt N, Milburn MV, Glass CK, Rosenfeld MG 1998 Determinants of coactivator LXXLL motif specificity in nuclear receptor transcriptional activation. Genes Dev 12:3357–3368[Abstract/Free Full Text]
  44. Darimont BD, Wagner RL, Apriletti JW, Stallcup MR, Kushner PJ, Baxter JD, Fletterick RJ, Yamamoto KR 1998 Structure and specificity of nuclear receptor-coactivator interactions. Genes Dev 12:3343–3356[Abstract/Free Full Text]
  45. Thornton JW 2001 Evolution of vertebrate steroid receptors from an ancestral estrogen receptor by ligand exploitation and serial genome expansions. Proc Natl Acad Sci USA 98:5671–5676[Abstract/Free Full Text]
  46. Liu W, Wang J, Sauter NK, Pearce D 1995 Steroid receptor heterodimerization demonstrated in vitro and in vivo. Proc Natl Acad Sci USA 92:12480–12484[Abstract]
  47. Bamberger CM, Bamberger AM, Wald M, Chrousos GP, Schulte HM 1997 Inhibition of mineralocorticoid activity by the ß-isoform of the human glucocorticoid receptor. J Steroid Biochem Mol Biol 60:43–50[CrossRef][Medline]
  48. Luisi BF, Xu WX, Otwinowski Z, Freedman LP, Yamamoto KR, Sigler PB 1991 Crystallographic analysis of the interaction of the glucocorticoid receptor with DNA. Nature 352:497–505[CrossRef][Medline]
  49. Dahlman-Wright K, Wright A, Gustafsson JA, Carlstedt-Duke J 1991 Interaction of the glucocorticoid receptor DNA-binding domain with DNA as a dimer is mediated by a short segment of five amino acids. J Biol Chem 266:3107–3112[Abstract/Free Full Text]
  50. Heck S, Kullmann M, Gast A, Ponta H, Rahmsdorf HJ, Herrlich P, Cato AC 1994 A distinct modulating domain in glucocorticoid receptor monomers in the repression of activity of the transcription factor AP-1. EMBO J 13:4087–4095[Abstract]
  51. Adams M, Meijer OC, Wang J, Bhargava A, Pearce D 2003 Homodimerization of the glucocorticoid receptor is not essential for response element binding: activation of the phenylethanolamine N-methyltransferase gene by dimerization-defective mutants. Mol Endocrinol 17:2583–2592[Abstract/Free Full Text]
  52. Kumar V, Chambon P 1988 The estrogen receptor binds tightly to its responsive element as a ligand-induced homodimer. Cell 55:145–156[Medline]
  53. Fawell SE, Lees JA, White R, Parker MG 1990 Characterization and colocalization of steroid binding and dimerization activities in the mouse estrogen receptor. Cell 60:953–962[Medline]
  54. Bledsoe RK, Montana VG, Stanley TB, Delves CJ, Apolito CJ, McKee DD, Consler TG, Parks DJ, Stewart EL, Willson TM, Lambert MH, Moore JT, Pearce KH, Xu HE 2002 Crystal structure of the glucocorticoid receptor ligand binding domain reveals a novel mode of receptor dimerization and coactivator recognition. Cell 110:93–105[Medline]
  55. Vottero A, Kino T, Combe H, Lecomte P, Chrousos GP 2002 A novel, C-terminal dominant negative mutation of the GR causes familial glucocorticoid resistance through abnormal interactions with p160 steroid receptor coactivators. J Clin Endocrinol Metab 87:2658–2667[Abstract/Free Full Text]
  56. Charmandari E, Kino T, Tiulpakov A, Zachman K, Raji A, Chrousos GP, A novel, carboxyl-terminal point mutation of the human glucocorticoid receptor gene causing glucocorticoid resistance: decreased affinity for ligand, reduced transactivation, and dominant negative activity upon the wild-type receptor. Oral presentation at the 86th Annual Meeting of The Endocrine Society, New Orleans, LA, 2004
  57. Reichardt HM, Kaestner KH, Tuckermann J, Kretz O, Wessely O, Bock R, Gass P, Schmid W, Herrlich P, Angel P, Schutz G 1998 DNA binding of the glucocorticoid receptor is not essential for survival. Cell 93:531–541[Medline]
  58. Felgner PL, Gadek TR, Holm M, Roman R, Chan HW, Wenz M, Northrop JP, Ringold GM, Danielsen M 1987 Lipofection: a highly efficient, lipid-mediated DNA-transfection procedure. Proc Natl Acad Sci USA 84:7413–7417[Abstract]
  59. Brasier AR, Tate JE, Habener JF 1989 Optimized use of the firefly luciferase assay as a reporter gene in mammalian cell lines. Biotechniques 7:1116–1122[Medline]
  60. Kino T, Gragerov A, Kopp JB, Stauber RH, Pavlakis GN, Chrousos GP 1999 The HIV-1 virion-associated protein Vpr is a coactivator of the human glucocorticoid receptor. J Exp Med 189:51–62[Abstract/Free Full Text]
  61. Bhattacharyya N, Dey A, Minucci S, Zimmer A, John S, Hager G, Ozato K 1997 Retinoid-induced chromatin structure alterations in the retinoic acid receptor ß 2 promoter. Mol Cell Biol 17:6481–6490[Abstract]
  62. Jantzen HM, Strahle U, Gloss B, Stewart F, Schmid W, Boshart M, Miksicek R, Schutz G 1987 Cooperativity of glucocorticoid response elements located far upstream of the tyrosine aminotransferase gene. Cell 49:29–38[Medline]