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Molecular and Cellular Biology, March 2007, p. 2266-2282, Vol. 27, No. 6
0270-7306/07/$08.00+0 doi:10.1128/MCB.01439-06
Human Glucocorticoid Receptor ß Binds RU-486 and Is Transcriptionally Active
Laura J. Lewis-Tuffin,1,
Christine M. Jewell,1
Rachelle J. Bienstock,2
Jennifer B. Collins,3 and
John A. Cidlowski1*
Laboratory of Signal Transduction,1 Scientific Computing Laboratory,2 NIEHS Microarray Facility, National Institute of Environmental Health Sciences, National Institutes of Health, Department of Health and Human Services, 111 TW Alexander Drive, Research Triangle Park, North Carolina 277093
Received 4 August 2006/ Returned for modification 1 September 2006/ Accepted 5 January 2007
 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Human glucocorticoid receptor (hGR) is expressed as two alternately spliced C-terminal isoforms,
and ß. In contrast to the
canonical hGR, hGRß is a nucleus-localized orphan
receptor thought not to bind ligand and not to affect gene
transcription other than by acting as a dominant negative to
hGR. Here we used confocal microscopy to examine the cellular
localization of transiently expressed fluorescent protein-tagged
hGRß in COS-1 and U-2 OS cells. Surprisingly, yellow
fluorescent protein (YFP)-hGRß was predominantly located in the
cytoplasm and translocated to the nucleus following application of the
glucocorticoid antagonist RU-486. This effect of RU-486 was confirmed
with transiently expressed wild-type hGRß. Confocal microscopy
of coexpressed YFP-hGRß and cyan fluorescent
protein-hGR in COS-1 cells indicated that the receptors move
into the nucleus independently. Using a ligand binding assay, we
confirmed that hGRß bound RU-486 but not the hGR
ligand dexamethasone. Examination of the cellular localization of
YFP-hGRß in response to a series of 57 related compounds
indicated that RU-486 is thus far the only identified ligand that
interacts with hGRß. The selective interaction of RU-486 with
hGRß was also supported by molecular modeling and computational
docking studies. Interestingly, microarray analysis indicates that
hGRß, expressed in the absence of hGR, can regulate
gene expression and furthermore that occupation of hGRß with
the antagonist RU-486 diminishes that capacity despite the lack of
helix 12 in the ligand binding
domain. |
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Glucocorticoids are an important class of natural and synthetic steroid hormone signaling molecules whose physiological actions can affect nearly every part of the human body. The effects of glucocorticoids are mediated by the glucocorticoid receptor (GR), a hormone binding transcription factor of the steroid family of nuclear receptors. The gene for human GR (hGR) was originally cloned in 1985 and consists of nine exons (12). This structural organization produces two isoforms of GR, hGR
and
hGRß, by the alternative splicing of exon 9
(22). Exon 9 encodes the
carboxy-terminal end of the ligand binding domain (LBD) for GR, as well
as the 3' untranslated region. Thus, the hGR and
-ß isoforms are identical up through amino acid 727, at which
point they diverge. hGR has an additional 50 amino acids that
encode helices 11 and 12 of the ligand binding domain. In contrast,
hGRß has only an additional 15 distinct amino acids.
Consequently, hGRß is missing helix 12 of the ligand binding
domain and possesses a unique sequence in helix 11 compared to
hGR. hGR is the classical GR and is found in the
cytoplasm in the absence of ligand. Upon ligand binding, hGR
translocates to the nucleus, where it affects gene transcription. In
contrast, immunohistochemistry studies have shown that hGRß is
constitutively nuclear and does not bind agonists
(22,
23). In addition,
hGRß is thought to affect gene transcription only by acting as
a dominant negative to hGR and altering the ability of
hGR to signal (1,
21,
22,
38).
The
distribution and relative expression of hGR versus
hGRß have been examined in a number of human cell lines
(23), as well as in both
healthy and diseased human cells and tissues. While there is general
agreement that the expression of hGR is greater than the
expression of hGRß in all cells and tissues, the actual extent
of hGRß expression is less clear. Although mRNA for
hGRß has been found in a variety of human tissues
(1,
22,
25), hGRß protein
has been shown to have a more restricted cellular distribution. Most
frequently, hGRß protein has been found in healthy T
lymphocytes, macrophages, neutrophils, eosinophils, and endogenous
peripheral blood mononuclear cells
(9,
10,
33). In addition,
hGRß protein has been reported in brain, lung, and heart tissue
(23), although there is a
contradictory report
(25). Interestingly, the
expression of hGRß has also been shown to be increased in
glucocorticoid-resistant forms of asthma
(3,
9,
16,
30), ulcerative colitis
(13), nasal polyposis
(10), and chronic
lymphocytic leukemia (19,
29). In addition, we have
previously shown that the expression of hGRß can be activated
in cells by proinflammatory cytokines
(36). Under these
conditions, hGRß is the predominant receptor in the cells and a
state of glucocorticoid resistance ensues. These reports suggest a
potential physiological consequence to changes in hGRß
expression. Thus, hGRß may be a key modulator of the
progression of certain immune-related glucocorticoid-resistant
diseases. In this report, we describe the identification of the
antiglucocorticoid/antiprogestin compound RU-486 as a ligand for
hGRß which causes nuclear translocation of both transiently and
stably transfected hGRß in COS-1 and U-2 OS cell lines. In
addition, we show that hGRß introduced into U-2 OS cells in the
absence of hGR can widely regulate gene expression and that
this action of the receptor is modulated by the glucocorticoid receptor
antagonist RU-486 despite the absence of a helix 12 in
hGRß.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Plasmids. The plasmids pEYFP-hGR
and
pECFP-hGR were previously described
(27). Plasmid
pCMV-hGRß was previously described
(22). The plasmid
pEYFP-hGRß was made by replacing the ClaI/BamHI fragment of
pEYFP-hGR (containing the hGR-specific 3'
coding sequences) with the ClaI/BamHI fragment from pCMV-hGRß
(containing the hGRß-specific coding sequences as well as 1,430
bp of hGRß 3' untranslated region). The plasmids
pTET-OFF and pTRE2hyg were obtained from BD Biosciences Clontech
(Mountain View,
CA).
Compounds.
The following compounds
were purchased from Steraloids, Inc. (Newport, RI): cortexolone
(4-pregnen-17,21-diol-3,20-dione), corticosterone
(4-pregnen-11ß,21-diol-3,20-dione), cortisol
(4-pregnen-11ß,17,21-triol-3,20-dione), cortisone
(4-pregnen-17,21-diol-3,11,20-trione), deltafludrocortisone
(1,4-pregnadien-9-fluoro-11ß,17,21-triol-3,20-dione),
desoximetasone
(1,4-pregnadien-9-fluoro-16-methyl-11ß,21-diol-3,20-dione),
dexamethasone
(1,4-pregnadien-9-fluoro-16-methyl-11ß,17,21-triol-3,20-dione),
dexamethasone 21-mesylate
(1,4-pregnadien-9-fluoro-16-methyl-11ß,17,21-triol-3,20-dione-21-methanesulfonate),
17ß-estradiol [1,3,5(10)-estratrien-3,17ß-diol],
prednisolone(1,4-pregnadien-11ß,17,21-triol-3,20-dione), progesterone
(4-pregnen-3,20-dione), RU-486 (4,9-estradien-17-propynyl,
11ß-[4-dimethylamino]phenyl-17ß-ol-3-one), testosterone
(4-androsten-17ß-ol-3-one), triamcinolone
(1,4-pregnadien-9-fluoro-11ß,16,17,21-tetrol-3,20-dione),
and triamcinolone acetonide
(1,4-pregnadien-9-fluoro-11ß,16,17,21-tetrol-3,20-dione-16,17-acetonide).RU-486 was also purchased from Sigma-Aldrich (St. Louis, MO). Compound
3
{2'-(3-pyridyl)-11ß,17,21-trihydroxy-16-methyl-20-oxopregn-4-eno[3,2-c]pyrazole},
compound 6
{2'-(4-iodophenyl)-11ß,17,21-trihydroxy-16-methyl-20-oxopregn-4-eno[3,2-c]pyrazole},
compound 11
{2'-(4-bromophenyl)-11ß,17,21-trihydroxy-16-methyl-20-oxopregn-4-eno[3,2-c]pyrazole},
compound 12
{2'-(4-fluorophenyl)-11ß,17,21-trihydroxy-16-methyl-20-oxopregn-4-eno[3,2-c]pyrazole},
and compound 16b
{2'-(2-chloro-3-pyridyl)-11ß,17,21-trihydroxy-16-methyl-20-oxopregn-4-eno[3,2-c]pyrazole}
were kind gifts from R. Hochberg (Yale University, New Haven, CT).
Deacylcortivazol
{2'-(phenyl)-11ß,17,21-trihydroxy-6,16-dimethyl-20-oxopregn-4,6-dieno[3,2-c]pyrazole}
was a kind gift from S. Simons (NIDDK, NIH, Bethesda, MD).
RU-28362(1,4,6-androstatrien-6-methyl-17-propynyl,
11ß,17-diol-3-one) was a kind gift from P. Housley(University of South Carolina School of Medicine, Columbia, SC).
ZK98299
(4,9-gonadien-11ß-[4-dimethylamino]phenyl-17-ol-17ß-[3-hydroxypropyl]-13-methyl-3-one)was a kind gift from T. Archer (NIEHS, NIH, Research Triangle Park,
NC). C. E. Cook (Research Triangle Institute, Research
Triangle Park, NC) and D. McDonnell (Duke University, Durham, NC)
kindly provided RTI compounds as follows: RTI 3021-002, -003, -020,
-021, and -023 and RTI 6413-001, -002, -006, -009a, -015, -016, -018,
-028, -029E, -029Z, -030, -031, -039, -042, -043, -043ox, -044, -045,
-045ox, -046a, -046b, -049b, -050a, -050b, -051a, -051b, -052, -054,
-055, -056, -057, and -058. [3H]RU-486 was obtained from
American Radiolabeled Chemicals, Inc. (St. Louis, MO).
[3H]dexamethasone was obtained from Perkin-Elmer Life
Sciences (Woodbridge, Ontario,
Canada).
Generation of U-2 OS cell lines stably expressing hGRß. U-2 OS cells were transfected with the pTET-OFF regulatory plasmid to establish the U-OFF parental cell line (20). In these cell lines, protein expression can be regulated by the addition of tetracycline to the medium. MluI and EcoRV ends were added onto the coding region (amino acids 1 to 742) of hGRß by PCR amplification of the pCMVhGRß plasmid. The pTRE2hyg vector was digested with MluI and EcoRV, and the two DNAs were ligated to form the pTRE2hGRß plasmid. The pTRE2hGRß plasmid was then transfected into the U-OFF cells, and clones that stably express hGRß were selected using 200 µg/ml of Geneticin and 500 µg/ml of hygromycin. Two hundred micrograms of hygromycin per milliliter was used for maintenance. Several clones were obtained, and the hGRß receptor levels were compared using Western blot analysis with an hGRß-specific antibody. Clone identity was further confirmed by isolating total RNA and performing reverse transcription-PCR (RT-PCR) with hGRß-specific primers. The resulting PCR products were sequenced.
Cell culture and transfection.
COS-1 cells
were maintained in Dulbecco modified Eagle medium (DMEM) with high
glucose (Invitrogen Life Technologies) supplemented with 10% fetal calf
serum-calf serum, 50 units/ml penicillin, and 0.05 mg/ml streptomycin
(Sigma-Aldrich). U-2 OS cells (wild type) were maintained in
DMEM-F-12 medium (Invitrogen Life Technologies) supplemented
with 5% heat-inactivated fetal calf serum, 50 units/ml penicillin, 0.05
mg/ml streptomycin, and 2 mM L-glutamine. U-OFF stable cells
were maintained in DMEM-F-12 medium supplemented with 10% fetal
calf serum-calf serum, 50 units/ml penicillin, 0.05 mg/ml streptomycin,
2 mM L-glutamine, and 0.2 mg/ml Geneticin (Invitrogen Life
Technologies). U-2 OS and U-2 OSß stable cells were
maintained in U-OFF medium with 0.2 mg/ml hygromycin B (Invitrogen Life
Technologies). All cells were grown at 37°C and 5%
CO2 in a humidified incubator and passaged every 3 to 7
days, as they approached confluence.
One day prior to transfection, COS-1 or U-2 OS cells were transferred to 78.5-cm2 dishes (7.5 x 105 cells/dish). Cells were transfected with TransIt-LT1 reagent (Mirus, Madison, WI) as described by the manufacturer using 20 µl TransIt-LT1 and 1.5 µg DNA per dish. The next day, cells were transferred to 9.6-cm2 glass-bottomed dishes (MatTek Corp., Ashland, MA), 1.5 x 105 cells/dish in medium containing charcoal-stripped serum. Microscope imaging was done the following day.
Confocal microscopy.
On the day of imaging, transfected
COS-1 or U-2 OS cells were treated with 1 µM of steroid for 3
to 6 h. Subsequently, cells expressing yellow fluorescent
protein (YFP)-hGR or YFP-hGRß were observed using a
Zeiss LSM 510 confocal laser scanning microscope as previously reported
(27). Cells expressing
cyan fluorescent protein (CFP)-hGR were observed on the same
microscope, exciting fluorescence with an argon laser at 458 nm and
collecting emission with a 470- to 500-nm band-pass filter.
Quantitative receptor localization analysis was carried out using the
Zeiss LSM 510 software to determine the fluorescence intensity of the
receptor in an equivalently sized region in both the nucleus and the
cytoplasm of at least 10 cells per treatment condition per experiment.
Experiments were repeated at least twice. The average ratio of nuclear
to cytoplasmic intensity was then used as a measure of receptor
distribution throughout the
cell.
Immunocytochemistry and Western blot analysis.
Immunocytochemistry was carried out
on COS-1 or U-2 OS cells transfected with CMV-hGR or
CMV-hGRß and on U-2 OSß cells, as previously described
using affinity-purified anti-GR#57 antibody prepared in our laboratory
(37,
38). Western blot assays
for hGR and hGRß were carried out on protein extracts
from U-OFF, U-2 OS, and U-2 OSß cells. Cell pellets
were sonicated in low-detergent buffer (20 mM Tris-Cl, pH 7.5, 2 mM
EDTA, 150 mM NaCl, 0.5% Triton X-100, with one protease inhibitor
tablet [Roche; no. 1 836 153] per 10 ml buffer added immediately prior
to use) for 30 seconds and then centrifuged for 15 min at 12,800
x g at 4°C. The protein concentration of the
supernatants was determined using Bio-Rad protein assay reagent
(Bio-Rad Laboratories, Hercules, CA) against bovine serum albumin
standards. Protein samples were heated in 1x Laemmli loading
buffer plus 2-mercaptoethanol for 5 min at 100°C prior to being
electrophoretically resolved on an 8% precast Tris-glycine gel
(Invitrogen Life Technologies, Carlsbad, CA). Proteins were transferred
to an 0.2-µm nitrocellulose membrane, blocked in 10% nonfat dry
milk in TBS-T (50 mM Tris, 150 mM NaCl, 0.5% Tween 20, pH 7.5)
overnight at 4°C, washed in TBS-T, and then
incubated with anti-ß-actin (1:10,000; catalog no.
MAB1501; Chemicon, Temecula, CA) plus either anti-GR#57
(1:1,000) or BShGR (1:1,000; catalog no. PA3-514; Affinity
BioReagents) in TBS-T for 1 h at room temperature. After
being washed in TBS-T, the blot was incubated with goat anti-rabbit
peroxidase-conjugated antibody (ECL Western blotting analysis system;
Amersham Biosciences, Buckinghamshire, England) diluted 1:10,000 in
TBS-T for 1 h at room temperature. After further washing,
bands were visualized with enhanced chemiluminescence detection
reagents (ECL; Amersham Biosciences) as specified by the
manufacturer.
Ligand binding assays. (i) Column binding assay. One day prior to the assay, cells were plated at 1.5 x 107 cells/145-cm2 dish in medium containing charcoal-stripped serum. On the day of assay, cells were harvested by incubation for 10 min in Versene (2.68 µM KCl, 1.47 µM KH2PO4, 137 mM NaCl, 0.54 µM EDTA, 8.09 mM Na2HPO4 · 7H2O) at 37°C and 5% CO2 followed by scraping. Cells were resuspended in 1 ml ice-cold phosphate-buffered saline, radiolabeled steroid was added with or without 500- to 1,000-fold-excess unlabeled steroid, and cells were incubated on ice for 2 h with gentle agitation. Cells were collected by centrifugation at 4°C, resuspended in an equal volume of ice-cold buffer A (20 mM sodium phosphate, pH 7.0, 50 mM NaCl, 10% glycerol, 2 mM ß-mercaptoethanol, 1 mM EDTA, pH 8.0), and homogenized at 4°C using a prechilled homogenizer with three 10-second bursts interspersed with 10-second rests on ice. Extracts were centrifuged at 165,000 x g for 1 h at 2°C. The supernatant was applied at 4°C to a Sephadex G-25 HiTrap desalting column (Amersham Biosciences Corp., Piscataway, NJ) previously equilibrated with buffer A according to the manufacturer's instructions, followed by 7.5 ml buffer A to elute. One-hundred-microliter fractions were collected and counted in a scintillation counter.
(ii) Ethanol extraction assay. U-2 OSß cells were plated in medium containing charcoal-stripped serum plus 1 µM dexamethasone 1 day prior to assay at 1 x 106 cells/well in six-well plates. Duplicate wells were incubated with 1, 5, 10, 25, 50, or 100 nM 3H-labeled RU-486 in the absence (total binding) or presence (nonspecific binding) of 10 µM cold RU-486 for 2 h at 37°C. Cells were then washed five times with 1 ml/well ice-cold phosphate-buffered saline and incubated at room temperature for 30 min in 1-ml/well 100% ethanol to extract the RU-486. Ethanol was then removed and counted in 5 ml scintillation fluid in a scintillation counter. For each assay, count data were converted to pmol and normalized to the total protein in one well of cells cultured in parallel. Total and nonspecific pmol/mg data were analyzed using GraphPad Prism 4 (GraphPad Software, Inc., San Diego, CA). Global curve fitting was performed simultaneously on the total and nonspecific binding data. The nonspecific curve was then subtracted from the total curve to obtain a curve of specific ligand binding which was fitted using the nonlinear regression curve-fitting function for a hyperbola (one-site binding) in this program. This generates a best-fit curve from which Bmax and Kd are determined. This assay was repeated four times; Kd values from the four assays were averaged to obtain the reported Kd.
Computational studies: molecular modeling and ligand docking.
The development of a homology
molecular model for hGRß was reported previously
(38). This model was
employed for computational docking studies. The Schrodinger
Glide v.3.5 docking software was used for all
ligand docking studies, and GlideScore was used as
the docking scoring function for ligand docking evaluation and
comparison. The Glide docking method and scoring function for ligands
have been found to compare favorably with most docking methods and have
been demonstrated capable of predicting ligand-receptor binding
interactions (7,
8). Four potential
hGRß ligands which were evaluated experimentally were
computationally docked with the hGRß model receptor:
dexamethasone, ZK98299, RTI 6413-001, and RU-486. The starting
three-dimensional (3D) structures for the dexamethasone and RU-486
ligands for docking were extracted, respectively, from the solved
crystal structures for hGR, Protein Data Bank files 1M2Z and
1NHZ. The structure for ZK98299 was obtained as a two-dimensional sdf
file from PubChem, and the structure for RTI 6413-001 was built and
optimized beginning with the 3D structure for RU-486, which was
modified using the BioMedCAche software. All starting ligand structures
were prepared for docking using the Schrodinger Ligprep software to
generate energy-minimized correct 3D ligand structures for docking
including tautomeric, stereochemical, and ionization variations. The
hGRß active site was defined for the ligand docking (volume of
the receptor searched when attempting to dock a ligand) by
superimposing the hGRß receptor model structure on that of the
solved crystal structure of the hGR receptor with bound ligand
(Protein Data Bank files 1M2Z, 1NHZ, and 1P93). The receptor grid
binding box for hGRß was defined as the area superimposed on
the ligand binding site within the hGR receptor crystal
structure. Ligand docking of the same four ligands performed with the
wild-type hGRß receptor was repeated with a model of the mutant
Q642V hGRß receptor, and the same computational methodology was
used for docking with the mutant receptor
model.
Microarray analysis. U-OFF and U-2 OSß cells were cultured in charcoal-stripped serum medium for 24 h prior to treatment. Total RNA was extracted from 5 x 10 6 U-OFF or U-2 OSß cells treated with either ethanol vehicle or 1 µM RU-486 for 6 h using the RNAqueous total RNA isolation kit (Ambion Inc. Austin, TX) according to the manufacturer's instructions. RNA was treated with DNase using the DNA-free DNase treatment and removal reagents (Ambion Inc.) according to to manufacturer's instructions prior to use with the microarray. Four pairs of RNA (vehicle versus RU-486 treated) were harvested for each cell type to yield four biological replicates for gene expression analysis.
Linear amplification label protocol and feature extraction. Gene expression analysis was conducted using Agilent Human1Av2 arrays (Agilent Technologies, Palo Alto, CA). Total RNA was amplified using the Agilent Low RNA Input Fluorescent Linear Amplification kit protocol. Starting with 500 ng of total RNA, Cy3- or Cy5-labeled cRNA was produced according to the manufacturer's protocol. For each two-color comparison, 750 ng of each Cy3- and Cy5-labeled cRNA was mixed and fragmented using the Agilent In Situ Hybridization kit protocol. Hybridizations were performed for 17 h in a rotating hybridization oven using the Agilent 60-mer oligonucleotide microarray processing protocol. Slides were washed as indicated in this protocol and then scanned with an Agilent scanner. Data were obtained using the Agilent Feature Extraction software (v7.5), using defaults for all parameters.
Rosetta Resolver (v5.0). Images and GEML files, including error and P values, were exported from the Agilent Feature Extraction software and deposited into Rosetta Resolver (version 5.0) (Rosetta Biosoftware, Kirkland, WA). The resultant ratio profiles were combined into ratio experiments as described in the work of Stoughton and Dai (32). In total, three ratio experiments were built containing eight arrays each (four biological replicates each with dye swaps) for U-OFF vehicle versus U-2 OSß vehicle, U-2 OSß vehicle versus RU-486, and U-OFF vehicle versus RU-486. Based on these experiments, lists of genes that were determined to be statistically differentially expressed using Resolver's error model at P < 0.001 were saved. The lists were then combined into a single list and the genes clustered hierarchically using Rosetta Resolver.
In a separate series of
experiments, total RNA was isolated from U-OFF and U-2 OS
cells cultured in charcoal-stripped serum medium and processed and
analyzed as described above. Three sets of RNA were isolated for three
biological replicates.
Quantitative RT-PCR analysis. U-OFF and stably hGRß-expressing cell lines were cultured in charcoal-stripped serum medium for 24 hours prior to treatment and then treated with vehicle or 1 µM RU-486 for 6 hours. Total RNA was isolated using the QIAGEN RNeasy minikit. Real-time PCR was performed using the 7900HT Sequence Detection System and predesigned primer/probe sets available from Applied Biosystems (Foster City, CA) following the manufacturer's instructions. The signal obtained from each gene primer/probe set was normalized to that of the unregulated housekeeping gene cyclophilin B primer/probe set (also available from Applied Biosystems). Each primer/probe set was analyzed in triplicate and with at least three different sets of RNA isolated from U-OFF cells, U-2 OSß vehicle-treated cells, and U-2 OSß RU-486-treated cells and normalized to cyclophilin B.
Statistical analysis. Statistical analysis for Fig. 1, 3, and 6 was performed using JMP5.0.1 software (SAS, Cary, NC). One-way analysis of variance (ANOVA) was performed for Fig. 1 and 3, and two-way ANOVA for Fig. 6, to identify the existence of statistically significant treatment differences for each cell and receptor type. Where significance was indicated, post hoc testing using Dunnett's test (comparison versus vehicle control) was carried out. Statistical analysis for Fig. 2 and 4 was performed by Shyamal Peddada of the NIEHS Biostatistics Branch. For each cell and receptor type, we tested the null hypothesis that all three treatment groups have the same probability of observing any given category against the two-sided alternative that each of the treated groups was different from the control (vehicle-treated) group. We tested the above hypothesis using a Dunnett-type test statistic along the lines of the work of Peddada et al. (24). The P values were determined using the bootstrap methodology (6). In all cases statistical significance was accepted at P < 0.05.
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FIG. 1. YFP-hGRß
translocates into the nucleus of transfected cells in response to
RU-486. (A) COS-1 and U-2 OS cells were transiently
transfected with plasmids expressing YFP-hGR or
YFP-hGRß and treated with ethanol vehicle or 1 µM
RU-486 or dexamethasone (Dex) for 3 h before being examined
with confocal microscopy. Both treatments caused nuclear translocation
of YFP-hGR in both cell lines; only RU-486 caused nuclear
translocation of YFP-hGRß. (B) COS-1 cells were
transiently transfected with a plasmid expressing YFP and treated with
1 µM RU-486 for 3 h before imaging. RU-486 had no
effect on the cellular localization of YFP. (C) The
localization of YFP-hGR and YFP-hGRß in response to
steroid treatment was quantified by determining the ratio of the
fluorescence intensity in an area of the nucleus divided by the
fluorescence intensity in a similarly sized area of the cytoplasm.
Black bars indicate no treatment; striped bars indicate 1 µM
RU-486; white bars indicate 1 µM dexamethasone. This analysis
confirmed that RU-486 but not dexamethasone caused nuclear
translocation of YFP-hGRß in both cell lines. Data are means
± SEMs (n  30 cells per treatment condition);
*, significantly different at P < 0.05 versus
vehicle for that receptor-cell type combination. (D) COS-1
cells were transfected with a plasmid expressing YFP-hGRß and
treated for 0, 0.25, 0.5, 0.75, 1, 2, 3, 4, 5, 6, 7, 8, 9, or
10 h with 1 µM RU-486. Cells were then imaged and
quantified as in panel C. Nuclear localization was clearly evident by
2 h of treatment. Data are means ± SEMs.
(E) COS-1 cells were transfected with a YFP-hGRß
expression plasmid and treated for 3 h with 1, 10, 100, 250,
500, 750, or 1,000 nM of RU-486. Cells were then imaged and quantified
as in panel C. Nuclear translocation of YFP-hGRß first became
evident with 250 nM RU-486. Data are means ±
SEMs.
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FIG. 3. RU-486-dependent
nuclear translocation of hGRß is not due to cytoplasmic
heterodimerization with hGR. (A) COS-1 cells were
transiently transfected with equal amounts of plasmids expressing
CFP-hGR and YFP-hGRß and treated with ethanol vehicle
or 1 µM RU-486 or dexamethasone (Dex) for 3 h before
being examined with confocal microscopy. Blue indicates localization of
CFP-hGR; yellow indicates YFP-hGRß localization; in
the merged image, green indicates areas where both receptors are
located. Both RU-486 and dexamethasone caused nuclear translocation of
CFP-hGR; only RU-486 caused translocation of YFP-hGRß.
(B) Localization of CFP-hGR was quantified by
determining the ratio of the fluorescence intensity in an area of the
nucleus divided by the fluorescence intensity in a similarly sized area
of the cytoplasm in the absence or presence of cotransfected
YFP-hGRß, with or without steroid treatment. Black bars
indicate vehicle treatment; striped bars indicate 1 µM RU-486;
white bars are 1 µM dexamethasone. Both dexamethasone and
RU-486 caused nuclear translocation of CFP-hGR, regardless of
the presence of YFP-hGRß. Data are means ± SEMs
(n 30 cells per treatment condition); *,
significant difference at P < 0.05 versus vehicle
treatment for that receptor-treatment combination. ANOVA indicated no
statistically significant effect of YFP-hGRß on the
localization of CFP-hGR. (C) Localization of
YFP-hGRß was quantified as in panel B
in the
absence or presence of cotransfected CFP-hGR, with or without
steroid treatment. Black bars indicate vehicle treatment; striped bars
indicate 1 µM RU-486; white bars are 1 µM
dexamethasone. RU-486 caused nuclear translocation of YFP-hGRß
to the same extent, and dexamethasone had no effect on translocation,
regardless of the presence of CFP-hGR. Data are means
± SEMs (n 30 cells per treatment condition);
*, significant difference at P < 0.05 versus
vehicle treatment for that receptor-treatment combination. ANOVA
indicated no statistically significant effect of CFP-hGR on
the localization of
YFP-hGRß.
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FIG. 6. Computer
modeling of the hGRß ligand binding domain docked with RU-486,
RTI 6413-001, ZK98299, or dexamethasone. (A) Illustrations of
the highest GlideScore poses for the ligands RU-486, RTI 6413-001,
ZK98299, and dexamethasone computationally docked into the model of the
hGRß ligand binding domain. Amino acids labeled on the diagrams
are predicated to be within 3 angstroms of the respective ligands;
boxed amino acids are predicted to form hydrogen bonds with the
respective ligands. The C-terminal tail of the hGRß ligand
binding domain can be seen to curl snugly around the dimethylaniline
substituent of RU-486, RTI 6413-001, and ZK98299, while dexamethasone
seems to make little contact with the hGRß C-terminal tail.
(B) Conformation of RU-486, RTI 6413-001, ZK98299, and
dexamethasone (Dex) when computationally docked in the hGRß
ligand binding domain (middle and right-hand structures); the
conformation of RU-486 and dexamethasone obtained from the solved
crystal structures of the hGR ligand binding domain (Protein
Data Bank files 1NHZ and 1M2Z, respectively) is shown for comparison
(left-hand structures). The positions of the D ring and carbons 11 and
17 are indicated. (C) (Left) Illustration of the ligand
RU-486 computationally docked into the model of a Q642V hGRß
mutant ligand binding domain. Amino acids labeled on the diagrams are
predicated to be within 3 angstroms of the ligand. No amino acids are
predicted to form hydrogen bonds with RU-486 in the ligand binding
domain of this mutant hGRß. (Right) Conformation of RU-486 when
computationally docked in the Q642V versus wild-type hGRß
ligand binding domains. Note that the orientation of the hydroxyl group
on carbon 17 (indicated) differs between the two
structures.
|
 View larger version (47K): [in a new window] |
FIG. 2. Wild-type
hGRß translocates into the nucleus in response to RU-486.
(A) COS-1 (left) and U-2 OS (right) cells were transiently
transfected with CMV-hGR or CMV-hGRß plasmids, which
express wild-type hGR or hGRß, respectively, and
treated with ethanol vehicle or 1 µM RU-486 or dexamethasone
(Dex) for 3 h before being processed for immunocytochemistry
with the GR#57 antibody, which recognizes both hGR and
hGRß. (B) Schematic illustration of the scoring
system used to quantitate the localization of the receptors with
different treatments. A number value was assigned to each cell based on
the relative amount of cytoplasmic and nuclear receptor as indicated
(N, nuclear receptor; C, cytoplasmic receptor): N << C, 1; N
< C, 2; N = C, 3; N > C, 4; N >> C, 5.
(C) Frequency histograms of the resulting localization scores
are plotted (n 130). Black bars indicate receptor
localization with vehicle treatment; striped bars indicate 1 µM
RU-486; white bars indicate 1 µM dexamethasone. Both ligands
caused statistically significant changes in the frequency histogram of
hGR, reflecting its nuclear translocation: the percentage of
cells scored as 5 is higher for the RU-486 and dexamethasone treatments
than for the vehicle treatment, while there are more vehicle-treated
cells scoring at 4 or below. In contrast, only RU-486 caused nuclear
translocation of hGRß: there was little difference in the
number of vehicle-treated cells versus dexamethasone-treated cells at
any score, while the number of RU-486-treated cells with a score of 5
was greater than the number of vehicle-treated cells with that score.
Bars, 25
µm.
|
 View larger version (32K): [in a new window] |
FIG. 4. Expression
and RU-486-dependent nuclear translocation of wild-type hGRß in
the U-2 OSß stable cell line. (A) The U-2 OS cell
line U-2 OSß, stably expressing hGRß under the control
of a Tet-OFF promoter system, was created. U-2 OS, U-2
OSß, and the U-OFF parental cell line were treated for at least
3 h with 1 µM RU-486 and then harvested for Western
blot assays. The GR#57 antibody recognizes both hGR (94 kDa)
and hGRß (90 kDa); the BShGR antibody recognizes hGRß
only; the actin antibody was used to demonstrate equal loading of the
lanes. The U-OFF cells did not express detectable levels of GR, while
U-2 OS and U-2 OSß exclusively expressed hGR
or hGRß, respectively. Numbers in the middle are
molecular masses in kilodaltons. (B) U-2 OSß cells
were treated with ethanol vehicle or 1 µM RU-486 ordexamethasone (Dex) for 3 h before being processed for
immunocytochemistry with the GR#57 antibody. Bar, 25 µm. RU-486
but not dexamethasone caused nuclear translocation of stably expressed
hGRß. (C) Quantification of the immunocytochemical
localization of hGRß receptor was carried out by assigning
localization scores and plotting a frequency histogram as in Fig.
2B. This analysis
confirmed that RU-486 but not dexamethasone caused the wild-type
receptor to undergo nuclear translocation. Black bars indicate vehicle
treatment; striped bars indicate 1 µM RU-486; white bars
indicate 1 µM dexamethasone; n
290.
|
|
TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The hGRß glucocorticoid receptor isoform undergoes nuclear translocation in response to RU-486. Our first indication that RU-486 might interact with hGRß came from confocal imaging studies. Plasmids expressing hGR
or hGRß that were tagged with YFP at
the amino terminus of GR (YFP-hGR and YFP-hGRß) were
transiently transfected into COS-1 cells. This cell line was chosen
because it does not express detectable levels of endogenous GR.
Transfected cells were treated for 3 h with 1 µM
dexamethasone or RU-486 and then imaged live using confocal laser
scanning microscopy (Fig.
1A). Untreated COS-1 cells expressing YFP-hGR showed the expected
cytoplasmic distribution of unliganded receptor. Also, as predicted,
both dexamethasone and RU-486 caused complete nuclear translocation of
YFP-hGR. However, untreated COS-1 cells expressing
YFP-hGRß also showed a largely cytoplasmic receptor
distribution. This pattern differs from the primarily
nuclear distribution of non-YFP-tagged hGRß in COS-1 cells that
we have previously reported
(23). Surprisingly,
YFP-hGRß translocated into the nucleus of cells in response to
treatment with RU-486 but not in response to dexamethasone, suggesting
for the first time that hGRß may be able to bind
ligand.
Previous reports have suggested a nuclear distribution of
the wild-type hGRß receptor, regardless of the presence of
agonist (23). Since it
was possible that the RU-486-stimulated nuclear translocation of
YFP-hGRß was a cell-type-specific response, we examined this
issue in the human osteosarcoma cell line U-2 OS, which also does not
express detectable levels of endogenous GR (Fig.
1A). Again,
YFP-hGR was located in the cytoplasm of U-2 OS cells in the
absence of treatment and translocated to the nucleus in response to
treatment with RU-486 or dexamethasone. Although YFP-hGRß was
located primarily in the nucleus of these cells in the absence of
treatment, the receptor was clearly present in the cytoplasm as well.
Nuclear translocation of YFP-hGRß occurred in response to
treatment with RU-486 but not dexamethasone. This RU-486-dependent
nuclear translocation of YFP-hGRß was not due to an effect of
RU-486 on the YFP tag since COS-1 cells that transiently expressed YFP
alone showed no change in YFP distribution with 3 h of 1
µM RU-486 treatment (Fig.
1B).
To facilitate
direct comparison of glucocorticoid receptor distribution in multiple
cells across experiments, the subcellular distribution in the two cell
lines was quantified. At least 30 cells for each cell, receptor, and
treatment type were examined. For each cell, the fluorescence intensity
of an area in the nucleus and the fluorescence intensity of an equally
sized area in the cytoplasm were used to create a ratio, which was then
combined to give an average ± standard error of the mean (SEM)
for each cell type and treatment condition (Fig.
1C). With this ratio,
numbers less than 1 indicate primarily cytoplasmic receptor
localization, numbers equal to 1 indicate equal receptor
distribution across the cell, and numbers greater than 1 indicate
primarily nuclear localization. This quantification confirmed that both
dexamethasone and RU-486 caused complete nuclear translocation of
YFP-hGR in both cell types, while only RU-486 promoted nuclear
translocation of YFP-hGRß in the two cell types examined. Thus,
in two different cell lines, treatment with RU-486 promotes nuclear
translocation of transiently expressed YFP-hGRß.
The time
course of RU-486-dependent YFP-hGRß nuclear translocation was
next determined by treating COS-1 cells transiently expressing
YFP-hGRß with 1 µM RU-486 for various times and then
imaging the cells and quantifying them as in Fig.
1A and C (Fig.
1D). These results
indicated that changes in receptor distribution occurred with as little
as 30 min of treatment. YFP-hGRß localization was primarily
nuclear by 2 h of treatment and was maximal by 6 h.
The RU-486 dose dependence of the YFP-hGRß nuclear
translocation was also determined. YFP-hGRß-expressing COS-1
cells were treated for 3 h with various concentrations of
RU-486 before being imaged and quantified (Fig.
1E). Significant receptor
nuclear translocation occurred at a concentration of 100 nM of RU-486
and reached maximum at 750 nM RU-486. Thus, the kinetics of
hGRß translocation are slower than those observed for
hGR with either agonists or antagonists.
We next
evaluated if the nuclear translocation that we observed for
YFP-hGRß in response to RU-486 also occurred with wild-type
hGRß. For these experiments, COS-1 and U-2 OS cells were
transiently transfected with plasmids expressing wild-type hGR
or hGRß (cytomegalovirus [CMV]-hGR or
CMV-hGRß, respectively), treated for 3 h with 1
µM RU-486 or dexamethasone, and then fixed and analyzed by
immunocytochemistry for the localization of the receptors
using the GR#57 antibody that recognizes both receptor
isoforms (Fig.
2A). Results were similar to those obtained with the
YFP-tagged receptors. In both cell types, wild-type hGR was
located in the cytoplasm in the absence of treatment and translocated
to the nucleus with RU-486 or dexamethasone treatment. Similarly, in
both cell types, wild-type hGRß was located in both the nucleus
and the cytoplasm in the absence of treatment. RU-486 facilitated
nuclear translocation of hGRß whereas dexamethasone did not. To
facilitate direct comparison of these results in multiple cells across
experiments, receptor localization was quantified by assigning a number
value to each cell based on the relative amount of cytoplasmic and
nuclear receptor: specifically, where N = nuclear and C
= cytoplasmic receptor, N << C, 1; N < C, 2; N
= C, 3; N > C, 4; N >> C, 5 (Fig.
2B). At least 130 cells
were analyzed per cell type, receptor, and treatment; localization
scores were then plotted as frequency histograms and analyzed for
statistical differences (Fig.
2C). This analysis
confirmed that, in both cell types, wild-type hGR responded
with nuclear translocation to both RU-486 and dexamethasone: the
percentage of cells scored as 5 is higher for the RU-486 (striped bars)
or dexamethasone (white bars) treatment than for the vehicle treatment
(black bars). In contrast, the localization of wild-type hGRß
changed only in response to RU-486: there was little difference in the
number of vehicle- versus dexamethasone-treated cells (black versus
white bars, respectively) at any score, while the number of RU-486
(striped bars)-treated cells increased as the scores indicated
progressively nuclear
localization.
hGRß moves into the nucleus independently of hGR.
Since the hGRß isoform of hGR
has not been previously demonstrated to bind ligand, the mechanism
underlying the observed RU-486-dependent nuclear translocation of
hGRß was unclear. One possibility was an RU-486-dependent
heterodimerization of hGRß with a small amount of hGR
in the cytoplasm, followed by nuclear translocation of the receptor
dimer. Alternatively, hGRß might bind RU-486, followed by
conformational changes in hGRß that stimulate nuclear
translocation, similar to the canonical response of hGR to
ligand binding.
If cytoplasmic heterodimerization of hGRß
with hGR is the mechanism by which nuclear
translocation of hGRß occurs, in COS-1 and U-2 OS cells it does
so in the presence of limiting quantities of hGR, since the
hGR protein is not detected in these cells by Western blotting
(see Fig. 4A and data not
shown). This could explain the incomplete nuclear translocation of
hGRß observed in Fig.
1A and
2A: more hGR
protein might be needed to obtain complete hGRß translocation.
To determine if increased hGR expression could result in
complete nuclear translocation of hGRß, COS-1 cells were
transiently cotransfected with equal amounts of plasmids expressing
CFP-hGR and YFP-hGRß, treated with 1 µM RU-486
or dexamethasone for 3 h, and then imaged live using confocal
microscopy (Fig.
3). Tagging the two receptors with different spectral
variants of GFP made it possible to simultaneously determine the
localization of the two receptors. As with YFP-hGR,
CFP-hGR was present in the cytoplasm in the absence of
treatment and translocated to the nucleus upon treatment with RU-486 or
dexamethasone (Fig. 3A,
top). Similarly, YFP-hGRß was found in the cytoplasm in the
absence of treatment and underwent nuclear translocation in response to
RU-486 but not dexamethasone (Fig.
3A, middle). The merged
images show where in the cell the two receptor isoforms colocalized, as
indicated by the green color (Fig.
3A, bottom).
To
facilitate direct comparison of these receptor distributions in
multiple cells across experiments, at least 30 cells for each receptor
and treatment type were quantified by determining the ratio of nuclear
to cytoplasmic fluorescence intensity, as before (Fig.
1C). To determine the
effect of YFP-hGRß on the cellular localization of
CFP-hGR, the localization ratios for CFP-hGR were
compared in the presence and absence of YFP-hGRß (Fig.
3B). Both RU-486 and
dexamethasone caused complete nuclear translocation of
CFP-hGR, in the presence and absence of YFP-hGRß.
ANOVA indicated no statistically significant effect of YFP-hGRß
on the localization of CFP-hGR. Similarly, the effect of
CFP-hGR on the cellular localization of YFP-hGRß was
determined (Fig. 3C).
RU-486, but not dexamethasone, caused nuclear translocation of
YFP-hGRß in both the presence and absence of CFP-hGR.
Again, nuclear translocation of YFP-hGRß was not complete, even
in the presence of similar amounts of CFP-hGR. ANOVA indicated
no statistically significant difference in the extent of this receptor
localization due to the presence of CFP-hGR. Thus, nuclear
translocation of hGRß in response to RU-486 is not likely due
to heterodimerization of the receptor with hGR in the
cytoplasm, followed by hGR-facilitated nuclear translocation.
Rather, hGRß is able to undergo nuclear translocation on its
own in response to RU-486.
hGRß is selective for RU-486. The observation that hGRß is able to undergo nuclear translocation in response to the ligand RU-486 suggests that other ligands may cause nuclear translocation as well. Since nuclear translocation of the receptor is an easily observed biological process, we used this method to screen for other potential ligands of hGRß. Accordingly, COS-1 cells were transfected with the YFP-hGRß expression plasmid, treated with various ligands, and then observed live using fluorescence microscopy for nuclear localization of YFP-hGRß (Table 1). Several classes of ligands were examined, including 10 glucocorticoids; four antiglucocorticoids; six analogs of cortivasol (14, 15); and ligands for the estrogen, progesterone, and androgen receptors. In addition, 37 antiprogestins with structural similarities to RU-486 were also tested (26, 35). Of the 57 compounds tested, only RU-486 promoted nuclear translocation of YFP-hGRß, suggesting that hGRß is highly selective for RU-486.
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View this table: [in a new window] |
TABLE 1. Observation
of ligand-dependent nuclear translocation of receptor
|
in a U-2 OS stable cell line and
endogenous GR expression in the U-OFF parental cell line used to make
the U-2 OSß cells
(20) (Fig.
4A). As expected, endogenous hGR was not detected in the U-OFF parental cell
line by this method. Only hGR was detected in the U-2
OS cell line, while only hGRß was detected in the U-2
OSß cell line. Treatment of U-2 OSß cells for
3 h with 1 µM RU-486 or dexamethasone, followed by
immunocytochemical localization of hGRß using the GR#57
antibody indicated that hGRß undergoes RU-486-dependent but not
dexamethasone-dependent nuclear translocation in this cell line (Fig.
4B). Quantification of
these results was carried out as in Fig.
2 on at least 290 cells
per treatment and confirmed that only RU-486 caused statistically
significant nuclear translocation of hGRß in the U-2
OSß cells (Fig.
4C). These results were
also confirmed in a second, independent clone of this stable cell line
(data not shown).
The U-OFF, U-2 OS, and U-2 OSß
stable cell lines were then used to determine the ability of
hGR and hGRß to bind RU-486 versus dexamethasone.
Whole-cell ligand binding assays were performed by incubating each cell
type with 3H-labeled RU-486 or [3H]dexamethasone
for 2 h at 0°C in the presence or absence of excess
unlabeled steroid. Cell lysates were then prepared and applied to a
Sephadex G-50 column, and fractions were collected to separate the
early-eluting bound steroid from the late-eluting free steroid (Fig.
5). The U-OFF cells exhibited a small amount of binding to both
[3H]dexamethasone and [3H]RU-486, which was
competed by the presence of excess cold ligand, indicating that the
binding was saturable (Fig.
5A). This small amount of
binding may be due to trace amounts of endogenous hGR or perhaps
progesterone receptor expression in these cells that is not
detectable by other methods such as Western blotting (for
example, Fig. 4A). The U-2
OS cells also exhibited binding to both
[3H]dexamethasone and [3H]RU-486 (Fig.
5B). However, the extent
of binding in U-2 OS cells was 5 to 7.5 times that seen in the
U-OFF cell line, indicating that the binding was due to the greatly
increased expression of hGR in these cells compared to the
U-OFF cells. Finally, the U-2 OSß cells exhibited a small
amount of [3H]dexamethasone binding comparable to that seen
with the U-OFF parental cell line (Fig.
5C). In contrast, in two
clones of the U-2 OSß cell line (Fig.
5C and data not shown),
the U-2 OSß cells exhibited six times the amount of
[3H]RU-486 binding that was observed with the U-OFF cells,
indicating that hGRß is able to bind the ligand RU-486. This
binding was confirmed with a second type of whole-cell ligand binding
assay in which the bound ligand was extracted from the cells with
ethanol (see Materials and Methods; data not shown). Using the ethanol
extraction assay, we determined the Kd of RU-486
binding to hGRß to be approximately 138 nM (L. Lewis-Tuffin and
J. Cidlowski, unpublished results), whereas the Kd
of RU-486 binding to hGR has been reported to be between 5 and
10 nM (27). Thus, the
affinity of RU-486 for hGRß is clearly lower than that for
hGR.
 View larger version (30K): [in a new window] |
FIG. 5. Wild-type
hGRß binds RU-486. (A) Whole-cell ligand binding
assays were performed with U-OFF cells treated with 100 nM
[3H]dexamethasone (closed triangles) or 100 nM
[3H]RU-486 (closed squares) for 2 h in the
presence or absence of excess cold steroid (50 µM dexamethasone
[open triangles] or 100 µM RU-486 [open squares]). Cells were
disrupted and applied to a Sephadex G-50 column, and fractions were
collected to separate early-eluting bound steroid from late-eluting
free steroid. The small elution between fractions 10 and 20 indicates
ligand binding to the small amount of endogenous hGR or progesterone
receptor (PR) present in these cells; this binding was
effectively competed away with excess cold ligand, indicating that it
was not nonspecific binding. (B) The experiment in panel A
was repeated using U-2 OS cells treated with 20 nM
[3H]dexamethasone (closed triangles) or 100 nM
[3H]RU-486 (closed squares) with and without 20 µM
and 10 µM unlabeled steroid (open symbols), respectively. Both
[3H]dexamethasone and [3H]RU-486 bound to
hGR in these cells; this binding was effectively competed away
with excess cold ligand. (C) The experiment in panel A was
repeated using U-2 OSß cells treated with 100 nM
[3H]dexamethasone (closed triangles) or 100 nM
[3H]RU-486 (closed squares), with or without 50 µM
or 100 µM unlabeled steroid (open symbols), respectively. As
with the U-OFF cells, the small elution between fractions 10 and 20
with [3H]dexamethasone treatment indicates ligand binding to
the small amount of endogenous hGR or PR present in these cells. In
contrast, [3H]RU-486 showed sixfold-higher amounts of
binding, which were effectively competed away with excess cold ligand
in these cells, indicating binding of [3H]RU-486 to
hGRß.
|
In addition to the dimethylaniline on position 11, the substituents present on the 17 position of the steroid ring D system and their stereochemical orientation also appear to be of significant importance for the hGRß ligand. Computational docking indicates that the RU-486 substituent groups in the 17 alpha and beta positions on the steroid ring D participate in favorable H-bonding interactions with hGRß residue Q642 (Fig. 6A). In contrast, ZK98299 and RTI 6413-001 have different chemical substituents present on position 17 compared with RU-486 (Fig. 6B). It appears that the larger 17ß-C-O-CH3 substituent present in RTI 6413-001 compared with the smaller 17ß-OH in RU-486 causes these substituents to dock into the receptor (in the highest ranked docked poses, Fig. 6A) with opposite stereochemical orientations (Fig. 6B), precluding formation of an H bond between hGRß residue Q642 and RTI 6413-001. Thus, the computational docking studies suggest that a combination of the dimethylaniline group on position 11 of the C ring and the 17ß-OH in a stereochemical orientation that facilitates its interaction with Q642 may underlie the productive interaction of RU-486 with the hGRß ligand binding domain.
It has previously
been shown that the glutamine 642 valine (Q642V) point mutation in
hGR dramatically decreases the interaction of the receptor
with ligands containing a 17-OH group (such as dexamethasone)
(18) but does not affect
interaction with ligands that do not have this group. To assess the
effect of this mutation on the interaction of hGRß with the
17ß-OH group of RU-486, computational docking studies were
repeated with a molecular model of a mutant Q642V hGRß receptor
(Fig. 6C). The size and
chemical composition differences between the glutamine and valine side
chains indicate that the terminal oxygen atom of the glutamine side
chain, which is 2.0 angstroms from the hydrogen of the 17ß-OH
group of RU-486 and forms a hydrogen bond with it, has been replaced
with one of the terminal hydrogens on the valine side chain.
Consequently, the orientation of the OH substituent on the RU-486 D
ring changes so that now it is the O of this group that is closest to
the terminal hydrogen of the V642 side chain. The distances between the
three terminal V642 hydrogens and the RU-486 OH group are now 2.6, 3.0,
and 3.6 angstroms, respectively. These distances are longer and less
favorable for hydrogen bond formation between RU-486 and the Q642V
mutant residue than is the distance between the wild-type hGRß
glutamine oxygen and the RU-486 ligand hydrogen. Additionally, this
mutation alters the conformation of the RU-486 substituent
17ß-OH group within the hGRß ligand binding pocket. Our
data suggest that the orientation of this group is critical for ligand
docking with hGRß. Thus, the modeling results suggest that a
Q642V hGRß receptor would not bind RU-486 while the Q642V
hGR receptor should.
Regulation of gene expression by hGRß.
The ability of hGRß to regulate
gene expression is unknown, and reporter assays have consistently
suggested that hGRß can regulate expression only by
antagonizing the action of hGR, regardless of the presence of
RU-486 (1,
22). However, the
regulation of transiently transfected reporter constructs may not
reflect the regulation of endogenous genes in the context of chromatin.
Based on our results indicating that RU-486 can bind to hGRß,
we hypothesized that RU-486 might modulate the functional activity of
hGRß and perhaps even be an agonist for this receptor isoform.
To determine if hGRß was capable of regulating gene expression
on its own in a chromatin context, we performed a total genome
microarray analysis. U-OFF and U-2 OSß cells were cultured in
charcoal-stripped serum medium for 24 h prior to treatment
and subsequent RNA isolation. Total RNA was prepared from U-OFF and U-2
OSß cells treated with vehicle or 1 µM RU-486 for
6 h, and microarray analysis was performed on four biological
replicates for each cell and treatment type. For each biological
replicate, three comparisons were made: U-OFF vehicle versus U-2
OSß vehicle, U-2 OSß vehicle versus U-2 OSß
RU-486, and U-OFF vehicle versus U-OFF RU-486. The results from each of
the biological replicates were combined, and the data were then
examined for genes that were differentially regulated at P
< 0.001 in any one of the three comparisons (5,622 genes total
combined). Figure
7A shows a cluster analysis of these three comparison groups. Genes shown
in green are repressed down to –0.3 (twofold repressed), and
genes shown in red are induced up to 0.3 (twofold induced). Those genes
represented in gray had a P value of >0.001 and
therefore did not pass our stringency requirement for significance. The
top cluster analysis shown in Fig.
7A, the comparison of
U-OFF vehicle with U-2 OSß vehicle, reveals the ability of
hGRß to regulate gene expression on its own, that is, in the
absence of hGR and in the absence of ligand. In this
comparison, 5,152 genes met our criteria for significant regulation. Of
these genes, 2,685 were induced and 2,467 were repressed by
hGRß expression (Fig.
7B). These results
indicate that hGRß is able to regulate gene expression in the
absence of hGR, a property of hGRß that was not
previously known. The middle cluster analysis, comparing U-2
OSß vehicle with U-2 OSß RU-486, shows that in
hGRß-expressing cells treated with RU-486 only 997 genes were
significantly regulated (Fig.
7A). This is far less than
the number of genes regulated by hGRß alone. Of the 997 genes,
only 260 were induced while a larger number of genes (737) were
repressed (Fig. 7B). The
third comparison, U-OFF vehicle versus U-OFF RU-486, shows that in the
absence of exogenous glucocorticoid receptor RU-486 treatment
significantly regulated 114 genes: 44 genes were induced and 70 genes
were repressed. Thus, RU-486 modulates the ability of hGRß to
regulate gene expression in a manner consistent with its binding to
hGRß. Interestingly, RU-486 appears to behave as an antagonist
to hGRß-mediated gene regulation.
 View larger version (39K): [in a new window] |
FIG. 7. hGRß
can regulate gene expression. (A) Microarray analysis was
performed on RNA from four biological replicates of U-OFF and U-2
OSß cells treated for 6 h with either ethanol vehicle
or 1 µM RU-486. Genes that were significantly different at
P < 0.001 were identified for each comparison: U-OFF
vehicle versus U-2 OSß vehicle, U-2 OSß vehicle versus
RU-486, and U-OFF vehicle versus RU-486. These genes were combined into
one list that was subjected to cluster analysis. Red indicates genes
that were induced for each comparison; green indicates genes that were
repressed. Fold changes are presented on a log-scale continuum with 0
(black) indicating no change for a given comparison. Any genes with
P > 0.001 did not meet our stringency requirements for
significance and are shown in gray. (B) Genes that were
identified as being significantly regulated in panel A for each
comparison were separated into induced versus repressed. Of the 5,152
genes that were statistically significantly regulated by hGRß
expression, 2,685 were induced and 2,467 were repressed (left graph).
Of the 997 genes that met our criteria for being statistically
significantly regulated by hGRß plus RU-486 treatment, 260 were
induced and 737 were repressed (middle graph). U-OFF cells treated
with RU-486 yielded 114 significantly regulated genes with
44 genes induced and 77 repressed (right graph). (C) Genes
from the combined gene lists are depicted using Venn diagrams. Genes
that are common between each of the three comparison groups are
represented in the overlapping
circles.
|
We next compared
genes that were regulated by hGRß to those regulated by
hGR. For this comparison, we isolated RNA from both the U-OFF
parental cells and those stably expressing hGR (U-OFF versus
U-2 OS), which were cultured for 24 h in
charcoal-stripped serum medium. However, prior to experimental
analysis, these cells were continuously cultured in medium containing
non-charcoal-stripped fetal calf serum; therefore, we cannot rule out a
residual effect of glucocorticoids from the fetal calf serum in the
case of hGR. Microarray analysis was performed on three
biological replicates, and the results from the combined gene list
showed that a total of 6,040 genes were regulated by hGR at
P < 0.001. These 6,040 genes were then compared to the
5,152 genes that were regulated by the hGRß-expressing cells
(U-OFF versus U-2 OSß) using human chromosome mapping (Fig.
8A). These maps show the physical position of the genes with known loci. The
structure of each chromosome is depicted in green, induced genes are
red, and repressed genes are blue. The color bar on the right shows the
expression level of these genes ranging from 5.0 (highly induced) to
0.01 (highly repressed). This analysis illustrates that there are
significant differences in the genes that are affected by the
expression of these two glucocorticoid receptors across the human
genome. Analysis of the lists by Venn diagram (Fig.
8B) illustrates that a
significant number of genes were both commonly and uniquely regulated
by these two glucocorticoid receptor isoforms. Additionally, we
analyzed gene regulation by the two active forms of the receptor,
hGR treated with 100 nM dexamethasone and hGRß vehicle
treated. Although these experiments were performed at different times
with slightly different parameters, we found 
1,000 genes
commonly regulated (data not shown).
 View larger version (37K): [in a new window] |
FIG. 8. Comparison
of gene regulation between hGR and hGRß. (A)
Microarray analysis was performed on RNA from three biological
replicates of U-OFF and U-2 OS cells. Genes that were
significantly different at P < 0.001 were identified
and combined into one list. This gene list was compared to the combined
gene list of the hGRß-expressing cells (U-OFF versus U-2
OSß) using human chromosome mapping. These maps show the
physical positions of the genes with known loci. The structure of each
chromosome is depicted in green with induced genes in red and repressed
genes in blue. The color bar on the right shows the expression level of
these genes ranging from 5.0 (highly induced) to 0.01 (highly
repressed). (B) The Venn diagram illustrates the genes that
are commonly regulated by both hGR and hGRß.
(C) Quantitative RT-PCR was performed using the 7900HT
sequence detection system using predesigned primer/probe sets for SAA1,
SPINK6, and INHBA and a custom primer/probe for cyclophilin B available
from Applied Biosystems. Each primer/probe set was analyzed in
triplicate and with at least three different sets of RNA isolated from
U-OFF cells, U-2 OSß vehicle-treated cells, and U-2 OSß
RU-486-treated cells and normalized to cyclophilin
B.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The canonical view of hGRß has been that it is barely expressed, does not bind ligand, and controls transcription only via a dominant-negative effect on hGR
-induced transcription.
Therefore, early studies suggested that it was generally of little
physiological importance. However, increasing evidence, both previously
published and presented here, suggests that this view is limited and
incorrect. Our work presented here indicates that hGRß is able
to control transcription without hGR involvement. We also
demonstrate that hGRß can interact with at least one ligand
(RU-486) and shed some light on factors that may govern ligand
interaction with the hGRß LBD. Furthermore, the increased
expression of hGRß in the development of
glucocorticoid-resistant forms of immune-related diseases is
increasingly well documented, particularly in asthma and ulcerative
colitis (17).
Importantly, these studies together suggest that it is the relative
ratio of hGR to hGRß that is the critical factor.
Thus, although hGRß expression may be low in noninflammatory
cells, increases in its expression during inflammation may produce
significant effects on glucocorticoid sensitivity. The development of
glucocorticoid resistance is a serious complication for diseases such
as asthma in which the most effective treatments exploit the
anti-inflammatory and immunomodulatory actions of glucocorticoids. A
ligand for hGRß could potentially reverse its contribution to
glucocorticoid insensitivity and restore the effectiveness of
glucocorticoid treatments.
The LBD of hGRß is identical
to that of hGR up through amino acid 727, which corresponds to
the end of helix 10 in the hGR LBD. From there, the two
receptors differ significantly. Although the crystal structure of the
hGRß LBD has not yet been determined, a comparative model of
the hGRß LBD was developed based on the X-ray crystal structure
of the hGR LBD and the structures of previously solved
homologous nuclear receptor LBDs
(38). This model
indicates that in addition to being truncated prior to the hGR
LBD helix 12, the last 15 amino acids of the hGRß LBD form a
somewhat flexible structure that does not resemble the highly ordered
helix 11 of hGR. Together, these features are thought to
underlie the inability of hGRß to bind ligand. Although ligands
may be able to enter the LBD of both hGR and hGRß, the
resulting conformational changes that serve to retain ligands in the
hGR LBD cannot occur with the hGRß LBD. Thus, ligands
that enter the hGRß LBD may occupy it for such a brief time
that they would be considered to have low affinity and not affect
hGRß activity. We report here for the first time that
hGRß can bind a ligand, RU-486, in such a way as to change the
cellular localization of the receptor and alter its ability to regulate
gene expression.
Given the predicted structural differences
between the hGRß and hGR LBDs, the observation that
hGRß binds RU-486 and changes its cellular localization was
completely unexpected. Indeed, previous work in our laboratory failed
to observe binding of RU-486 to hGRß
(22). However, this early
work was done using cells transiently transfected with hGRß and
with a low concentration of RU-486 (50 nM), the combination of which
may explain the lack of binding observed in that study. Our
dose-response results for the nuclear translocation of YFP-hGRß
indicate that at least 100 nM RU-486 is necessary for translocation,
which is consistent with the estimated Kd of 138 nM
for RU-486 binding to hGRß. It is therefore likely that in our
original experiment we did not observe binding of RU-486 to
hGRß because the concentration of 3H-ligand was too
low (18). Because the
RU-486 affinity for hGRß is low compared to the interaction of
compounds such as dexamethasone or RU-486 with hGR, we were
concerned that the effects of RU-486 on hGRß might instead be
due to an unidentified contaminant in the RU-486 stock. To address this
issue, we confirmed that freshly prepared RU-486 from two different
sources (Steraloids and Sigma) had identical effects on hGRß
cellular localization (data not shown). In addition, mass spectrometry
analysis confirmed that the only appreciable impurity in either stock
solution was 0.5% of an 11ß-[4-monomethylamino] phenyl version
of RU-486 (data not shown). Therefore, it is unlikely that a
contaminating compound caused the effects on hGRß localization
seen with RU-486 treatment.
The interaction between RU-486 and
hGRß was initially suggested by studies of the cellular
localization of transiently transfected YFP-hGRß in COS-1
cells. YFP-hGRß is found primarily in the cytoplasm in COS-1
cells, in contrast to the primarily nuclear localization of
YFP-hGRß in another cell line, U-2 OS (Fig.
1), as well as of
wild-type hGRß in COS-1 cells (Fig.
2) and other cell types
(23). This unusual
localization of YFP-tagged hGRß in COS-1 facilitated the
observation of nuclear translocation in response to RU-486 but not
dexamethasone. This translocation may not necessarily indicate a
ligand-hGRß interaction. As was noted, one possibility is that
RU-486 facilitates heterodimer formation between hGR and
hGRß as a result of binding to hGR, which results in
cotranslocation of the receptors. While this may be possible, two
observations make this mechanism unlikely to account for the
RU-486-dependent hGRß nuclear translocation. First, transiently
transfected hGRß is expressed far in excess of any endogenous
hGR in COS-1 and U-2 OS cells. It is hard to explain how the
very small amounts of endogenous hGR in these cells could
heterodimerize with and cause translocation of such an excess of
hGRß over a time course of minutes to hours, especially in
COS-1 cells, where the majority of YFP-hGRß is initially found
in the cytoplasm. If such a mechanism did exist, it would be expected
that the extent of nuclear translocation of YFP-hGRß would
increase in the presence of increased hGR expression. This was
not the case, however; the same, incomplete nuclear translocation of
YFP-hGRß occurred in response to RU-486 in the presence of
equivalent levels of CFP-hGR (Fig.
3). Furthermore,
CFP-hGR did undergo complete nuclear translocation in response
to RU-486 in these cells: the differential distribution of the two
receptors in the same cell is highlighted in the merged images in Fig.
3. Together, these results
strongly indicate that heterodimerization of hGRß with
hGR does not occur in the cytoplasm and is not responsible for
the RU-486-dependent nuclear translocation of
hGRß.
Although ligands may exist for hGRß that do
not cause cytoplasmic-to-nuclear translocation of YFP-tagged
hGRß in COS-1 cells, this is nevertheless a convenient approach
to screen for other candidate hGRß ligands. Accordingly, we
examined a total of 57 compounds for their ability to cause nuclear
translocation of hGRß that might indicate receptor-ligand
binding. RU-486 has several features that make it a unique
glucocorticoid receptor ligand (as seen in Fig.
6B). In particular, both
the nuclear localization studies and the computational docking studies
support the idea that the 11ß-dimethylaniline group,
the 17-propynyl group, and the placement of the hydroxyl group
in the 17ß (rather than 17) position all may
contribute to the unique properties of RU-486 as a ligand for
hGRß. Several of the ligands that we examined contained these
features, though only RU-486 contained them all. For example, ZK98299
has the 11ß-dimethylaniline group, RU-28362 has the
17-propynyl and 17ß-hydroxl groups, RTI 6413-001 has
the 11ß-dimethylaniline and 17-propynyl groups, and
RTI 3021-002 has the 11ß-dimethylaniline and
17ß-hydroxl groups. When all of these groups are present in the
ligand, they may form unique interactions with amino acids lining the
pocket of the hGRß LBD that could facilitate prolonged
occupation of the LBD even in the absence of helix 12.
Indeed,
our computational docking studies support the importance of these
substituents in potential hGRß ligands, as evidenced by the
good van der Waals interactions exhibited between the RU-486 ligand and
hGRß receptor, which were absent in the case of the docked
dexamethasone ligand. To begin with, the hGRß C terminus seems
to wrap around and interact with the 11ß-dimethylaniline group
when this substituent is present in a potential ligand. Furthermore, it
appears that the highest-ranked docked pose for RU-486 in the
hGRß structure has the stereochemistry on position 17 reversed
from that of docked RU-486 in the solved hGR crystal structure
(Protein Data Bank 1NHZ) (Fig.
6B). This finding can be
attributed to differences in the ligand interactions with the vastly
different hGR and hGRß C-terminal structures. The
17-propynyl substituent may facilitate this specific
stereochemical orientation of the RU-486 17-OH group within the
hGRß LBD, permitting hydrogen bond interactions with Q642 in
the computational docking studies with the hGRß model
structure. The importance of hydrogen bond interactions between ligand
and Q642 has already been established for hGR, both with
modeling data (18) and
experimentally (2,
18,
28). Computational
docking studies of a Q642V mutant of hGRß support the
importance of this residue in hydrogen bond formation and the
stereochemistry of substituent 17 on the RU-486 ligand. Experimentally,
only RU-486 was able to bind to hGRß and cause nuclear
translocation of the receptor. Such ligand specificity on the part of a
receptor is a highly sought feature in drug development. It will be
important to learn more about the basis for this specificity, as well
as how it can be exploited to produce higher-affinity hGRß
ligands.
Previous work with transiently overexpressed
hGRß in COS-1 cells and endogenous hGRß in HeLa S3
cells indicates that hGRß is primarily found in the nucleus
(23). It is not known if
this localization is the result of an endogenous ligand or is a
property inherent to hGRß. It is interesting, however, that
substitution of amino acids Lys733 and Pro734 found in helix 11 of
hGRß into hGR also leads to a nuclear localization of
hGR in the absence of ligand
(38). Of greater
importance is the ability of the receptor to mediate changes in gene
expression, which might be modulated by binding RU-486. Accordingly, we
performed microarray analysis on cells that stably express hGRß
in the absence of hGR to determine the ability of hGRß
to affect gene expression directly. Our results indicate that
hGRß can regulate gene expression by itself, a novel finding
that may be especially important in disease states in which
hGRß expression is up-regulated. Furthermore, this regulatory
activity of hGRß can be altered by RU-486. Additionally, the
resemblance that is emerging between hGRß and two other members
of the nuclear receptor superfamily of proteins—CAR and
ERR—is interesting
(34). All three of these
receptors appear to be ligand-independent activators
of gene expression. As of yet, they do not appear
to have endogenous ligands but can bind synthetic drugs such
as TCPOBOP for CAR (34),
tamoxifen for ERR
(4), and now RU-486 for
hGRß.
When used as an abortifacient, RU-486 is administered as a single 600-mg dose (31). Other uses for this potent antiprogestin, antiglucocorticoid agent are also being explored, which would likely entail different doses and administration plans (31). Due to its low rate of metabolic clearance in humans, even a single 100-mg dose of RU-486 persists at micromolar concentrations in the blood up to 72 h after administration (11). Thus, although the affinity of hGRß for RU-486 is relatively low, it is possible that standard use of this drug results in modification of the activity of endogenous hGRß. Furthermore, our demonstration of the ability of hGRß to bind a ligand suggests that it may be possible to design other ligands with higher affinity for this receptor. Taken together, this work suggests that hGRß may have a more important physiological role than has previously been thought, both as an endogenous manipulator of gene expression and as a pharmaceutical target.
We thank Paul Housley, Nick Lu, and Brian Necela for invaluable discussions of this work. We thank Richard Hochberg, Stoney Simons, Paul Housley, Donald McDonnell, and Ed Cook for kindly providing many of the ligands screened in Table 1. We thank Danica Ducharme of the NIEHS Microarray Facility for performing the microarray hybridizations. We thank Fred Lih of the NIEHS Mass Spectrometry Work Group for analysis of the purity of our RU-486 stocks.
* Corresponding author. Mailing address: National Institute of Environmental Health Sciences, P.O. Box 12233, MD F3-07, Research Triangle Park, NC 27709. Phone: (919) 541-1564. Fax: (919) 541-1367. E-mail: cidlows1{at}niehs.nih.gov.
Published ahead of print on 22 January 2007.
Present
address: Department of Cancer Biology, Mayo Clinic, 4500 San Pablo
Road, Jacksonville, FL 32224.
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