Molecular Identification and Characterization of A and B Forms of the Glucocorticoid Receptor
Matthew R. Yudt and
John A. Cidlowski
Laboratory of Signal Transduction National Institute of
Environmental Health Sciences National Institutes of Health
Research Triangle Park, North Carolina 27709
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ABSTRACT
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The human glucocorticoid receptor (hGR
) is a
ligand-activated transcription factor that mediates the physiological
effects of corticosteroid hormones and is essential for life.
Originally cloned in 1986, the transcriptionally active hGR
was
reported to be a single protein species of 777 amino acids (molecular
mass = 94 kDa). Biochemical data, obtained using various
mammalian tissues and cell lines, however, have consistently revealed
an additional, slightly smaller, second hGR protein (molecular
mass = 91 kDa) that is not recognized by antibodies specific for
the transcriptionally inactive and dominant negative,
non-hormone-binding hGRß isoform. We report here that when a single
GR cDNA is transfected in COS-1 cells, or transcribed and translated
in vitro, two forms of the receptor are observed, similar
to those seen in cells that contain endogenous GR. These data suggest
that two forms of the hGR
are produced by alternative translation of
the same gene and are henceforth termed GR-A and GR-B. To test this
hypothesis, we have investigated the role of an internal ATG codon
corresponding to methionine 27 (M27) as a potential alternative
translation initiation site for the GR. Mutagenesis of this ATG codon
to ACG in human, rat, and mouse GR cDNA results in generation of a
single 94-kDa protein species, GR-A. Moreover, mutagenesis of the
initial ATG codon to ACG (Met 1 to Thr) also resulted in production of
single, shorter protein species (91 kDa), GR-B. Mutagenesis of the
Kozak translation initiation sequence strongly indicates that a leaky
ribosomal scanning mechanism is responsible for generating the GR-A and
-B isoforms. Western blot analysis using peptide-specific antibodies
show both the A and B receptor forms are present in human cell lines.
Both receptors exhibit similar subcellular localization and nuclear
translocation after ligand activation. Functional analyses of hGR-A
and hGR-B under various glucocorticoid-responsive promoters reveal the
shorter hGR-B to be nearly twice as effective as the longer hGR-A
species in gene transactivation, but not in transrepression.
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INTRODUCTION
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The glucocorticoid receptor (GR) mediates the physiological
effects of corticosteroid hormones in species from fish to mammals. GR
is a member of the nuclear hormone receptor superfamily of
ligand-activated transcription factors (1) and is essential for life
(2). The activity of the GR, as well as the progesterone (PR), androgen
(AR), and mineralocorticoid receptors (MR) is partially mediated
through a palindromic response element termed the glucocorticoid
response element (GRE) located in the promoter regions of target genes
(3). Unlike most members of the steroid receptor superfamily, the GR is
primarily cytosolic in the absence of ligand. Activation and nuclear
translocation of the GR after ligand binding proceeds through a complex
mechanism involving a loss of energy-dependent protein interactions
with hsp90, hsp70, and several other proteins (4). Although the
classical view of steroid action involves an increase in gene
transcription in response to receptor activation, in fact several
glucocorticoid target genes undergo a hormone-dependent repression (5).
Furthermore, in addition to GRE-dependent processes, a growing body of
literature indicates that many glucocorticoid responses involve protein
interactions with other transcription factors and likely proceed
through a mutual inhibitory antagonism involving direct protein-protein
interactions with other transcription factors including, for example,
nuclear factor-
B (NF-
B) and AP1 (6).
Our understanding of the complexity of nuclear receptor signaling
mechanisms has advanced significantly in recent years. The discovery
and characterization of receptor coactivators and corepressors bridge
the gap between the DNA-bound receptors and the general transcription
machinery (7, 8, 9). Similarly, our knowledge regarding the role of
chromatin structure in steroid receptor signaling has been enhanced in
recent years (10, 11). The three-dimensional structure of many nuclear
receptor ligand binding domains has not only revealed a common protein
fold and ligand binding symmetry among superfamily members, but exposed
the subtle ligand interactions and associated conformational changes
necessary for a mechanistic understanding of steroid action (reviewed
in Ref. 12). Furthermore, examples of ligand-independent activation
mechanisms in nuclear receptor signaling continue to multiply (13).
An additional level of complexity of steroid hormone receptor action is
the existence of multiple receptor subtypes and isoforms with unique
biological roles (14, 15, 16). For example, multiple genes encode different
forms of the estrogen, retinoid, and thyroid hormone receptors.
Alternative splicing of progesterone, glucocorticoid, and retinoid
receptor mRNA gives rise to multiple forms of these proteins. The
progesterone receptor (PR) exists as a mixture of A and B forms,
generated from the same gene by alternative translation initiation.
Although both PR isoforms can arise from a single mRNA (17), it appears
that specific promoters may also regulate mRNA production specific for
each PR isoform (18). Both forms of PRs are well known to display
distinct biochemical and physiological properties (19). This extensive
multiplicity within the nuclear receptor superfamily suggests that the
diversity of receptor expression may be an important component
mediating the various physiological actions of steroid hormones.
We report here that the GR
gene is subject to alternative
translation initiation from a downstream, in-frame ATG codon. Our data
suggest that a leaky ribosomal scanning mechanism (20) produces two GR
protein products, with the second initiating at an ATG codon
corresponding to methionine 27 in the hGR. We term the longer protein,
initiated from the first ATG codon (Met 1) as hGR-A, and the shorter
protein (751 amino acids) as hGR-B. We have constructed a GR-A-specific
antibody that, when used in conjunction with an antibody that
recognizes both protein species, permits the discrimination of
endogenous expression of the two hGR
isoforms. Interestingly, the
shorter hGR-B is twice as effective as the longer hGR-A isoform in
activating transcription from a GRE but has a similar efficacy in
repression of NF-
B/p65 transactivation. This discovery of an
alternative initiation site within the GR gene, and the functional
divergence observed, provides a new potential mechanism to explain the
diversity of glucocorticoid responses in different tissues.
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RESULTS
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Expression of Recombinant hGR
The cloning of the hGR
into mammalian and in vitro
expression vectors has allowed a direct study of this protein,
independent of the alternatively spliced GRß variant (21, 22). The
human hGR
and -ß variants differ by only 35 amino acids in length
at the extreme carboxy terminus. Although quantitative measurement of
their coexpression in human tissues or cell lines remains difficult
because of the relative abundance of hGR
to ß, the two isoforms
can be discriminated immunologically using specific antibodies (23, 24). Interestingly, the recombinant hGR
when expressed alone, either
in vitro with 35S methionine or in
COS-1 cells, known to be void of detectable endogenous GR, consistently
appears as a doublet of approximately equal intensities (Fig. 1A
).

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Figure 1. In Vitro and in Vivo
Expression of Recombinant hGR
A, The wild-type hGR was prepared either by in vitro
translation using 35S methionine and reticulocyte lysates
(left panel) or by transient transfection of COS-1 cells
(right panel). Approximately 25 µg of
protein were electrophoresed on an 8% polyacrylamide gel. The
35S-labeled receptor was detected by autoradiography of the
dried gel, while the COS-1-expressed proteins were transferred to
nitrocellulose and detected by Western blotting. The positions of the
molecular mass markers in kilodaltons are indicated. Electrophoresis
was carried out for an extended period to resolve the protein doublet
of approximately 94 and 91 kDa. B, Wild-type hGR (1777) and two
carboxy-terminal truncation mutants, hGR(1742) and hGR(1706), were
expressed in vitro using reticulocyte lysates.
Electrophoresis was carried out as in panel A to resolve the hGR
protein doublet. Data shown are representative of at least three
different experiments.
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There are several possible explanations for the origin of the observed
hGR
protein doublet. Although proteolysis could explain the
appearance of such a doublet, inclusion of several protease inhibitors
did not block production of the lower mol wt (Mr)
product, arguing against degradation as the source of the doublet.
Moreover, in vitro transcription and translation of two
carboxy-terminal truncation mutants, hGR(1742) and hGR(1706),
results in a similar doublet pattern (Fig. 1B
), suggesting that
carboxy-terminal degradation is not the source of the second band.
Another possible source of the doublet is phosphorylation; however,
phosphatase treatment of the reticulocyte or COS-1 lysate containing
hGR
does not affect the doublet pattern (data not shown).
Alternative Translation Initiation of the GR
To investigate the possibility of alternative translation
initiation of the GR as a source of the observed doublet, the hGR cDNA
was examined for downstream ATG start codons. Only one in-frame ATG
codon, corresponding to methionine 27, was found within the first 300
nucleotides of the initial hGR ATG translation start site. Translation
initiation from this internal ATG site would yield a protein almost 3
kDa shorter (apparent molecular mass = 91 kDa) than the
full-length hGR from residues 1777 (apparent molecular mass = 94
kDa). To test the hypothesis of alternative initiation as a source of
the protein doublet observed in GR expression systems, both the initial
ATG start codon (methionine 1) and the internal ATG (methionine 27)
were mutated to ACG (a threonine codon). Mutagenesis of the individual
ATG codons in the hGR
cDNA in both the in vitro
expression vector and the mammalian expression vector resulted in the
expression of a single hGR
species (Fig. 2
). We have termed the longer GR,
generated from the first ATG codon, GR-A. The shorter GR species,
translated from the internal ATG corresponding to methionine 27 (amino
acid 28 in rat and mouse GR), is designated as GR-B. A proteolytic
fragment common to both GR forms of approximately 83 kDa is
consistently observed at higher levels in cells expressing GR-B.
In every mammalian species in which GR has been cloned and sequenced,
except the guinea pig, an internal ATG was found 27 or 28 codons from
the initial ATG (Fig. 3A
). Interestingly,
the guinea pig has been shown to be relatively glucocorticoid resistant
in comparison to other mammals (25). In addition, neither the African
frog nor rainbow trout GR contain a second potential translational
start site near this position. Since doublets are detected in both
mouse (mGR) and rat GR (rGR) expression systems, a similar start site
mutagenesis analysis was carried out on these receptor species. As
observed for the hGR, mutagenesis of the potential start sites of the
mGR and rGR resulted in expression of single protein species in
vivo and in vitro (Fig. 3
, B and D). To compare the
native GR with the recombinant forms, mouse and rat liver samples were
analyzed for GR expression. Both mouse and rat liver GR are expressed
as doublets, directly corresponding to the A and B forms observed with
the recombinant proteins (Fig. 3
, C and E). This is a particularly
important observation considering neither mouse nor rat contain the ß
form of the GR (Bofetiado, D. M., and J. A. Cidlowski, unpublished
observations). These results suggest that the human, rat, and mouse GR
genes can produce two proteins via alternative initiation of the same
gene transcript.

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Figure 3. Species Comparison of GR Translation Initiation
Sites
A, Sequence alignments of residues 140 of GRs from various species.
The indicated sequences were compiled from NCBI databases (SWISS-PROT
accession numbers: human, P04150; squirrel monkey P79686; rat, P06536;
mouse, P06537; xenopus, P49844; rainbow trout, P49843). A
dot indicates a conserved residue, while a
dash indicates a single residue gap. B, The mouse GR
expression vector (pCMV-mGR) was mutated as in Fig. 2 for the hGR,
changing potential translation initiation codons (ATG) 1 and 28
individually to threonine codons (ACG) to yield M1T and M28T,
respectively. Protein expression in transfected COS-1 cells was
measured by Western blot with Ab57. A similar band pattern was observed
with in vitro translated wt and mutant mGR (not shown).
C, Fifty micrograms of mouse liver protein extract were subjected to
Western blot analysis with Ab57. The two major immunoreactive bands at
94 and 91 kDa correspond with the mGR-A and -B bands detected in panel
B. D, The rat GR expression vector (pSG5-rGR) was also mutated at the
potential rGR start sites (Met1 and Met 28) and subjected to in
vitro expression using reticulocyte lysates and
35S-methionine followed by autoradiography. An identical
pattern is observed when the same plasmids are expressed in COS-1 cells
(not shown). E, Fifty micrograms of rat liver protein were analyzed for
GR content by Western blotting with Ab57. The two major immunoreactive
bands at 94 and 91 kDa correspond directly with the rGR-A and -B
isoforms detected in panel D. All Western blots are representative of
at least three separate experiments.
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Detection of Endogenous hGR-A Using an hGR-A-Specific Antibody
To evaluate the endogenous expression pattern of the GR-A and -B
isoforms, an antibody was generated that is specific to the longer,
hGR-A isoform (Fig. 4A
). A dual Western
blot analysis was then carried out on COS-1-expressed hGR wild type
(wt) and the start site mutants, hGR-A (M27T) and hGR-B (M1T), using
both the hGR-A-specific antibody and Ab57, a polyclonal
epitope-purified antibody generated against a peptide sequence common
to both GR-A and GR-B (26). As seen in Fig. 4B
, the hGR-A-specific
antibody recognizes only the longer hGR-A species, while the Ab57
recognizes both isoforms. A similar immunoreactive evaluation was
carried out using various human cell culture lines (HeLa, CEM-C7, and
HEK-293) known to express endogenous hGR
. As observed with the
recombinant protein, the Ab57 detects the GR doublet while the
hGR-A-specific antibody detects only the top band (Fig. 4C
). The 83-kDa
band detected in HeLa cell extracts is a common GR degradation product
observed in this cell line. These data suggest both hGR-A and hGR-B are
endogenously expressed in several human cell lines.

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Figure 4. Western Analysis of hGR-A and hGR-B
A, hGR-A-specific antibody production. A polyclonal antibody was
prepared from rabbits, using a synthetic peptide antigen corresponding
to residues 322 of the hGR. The location of the Ab57 binding site
(residues 346357 of the mouse GR), which is present in both hGR-A and
hGR-B, is shown for comparison. B, The hGR-A-specific antibody
recognizes the hGR initiated from the methionine 1 codon but not the
GR-B form initiated from methionine 27 codon. COS-1 cells were
transfected with the wt hGR and the two start site mutants described
in Fig. 2 . As observed in Fig. 2 , Ab57 detects both forms of the hGR as
illustrated by Western blotting (left panel). In
contrast, the hGR-A specific antibody only detects the longer, hGR-A
form (right panel). C, Endogenous hGR production is a
mixture of the A and B GR isoforms. Endogenously produced hGR from
several cell lines (HeLa, HEK293, CEM-C7) were analyzed for hGR-A and
-B production by Western blotting with the GR-A-specific antibody.
Approximately 50 µg of total protein extract from the human cell
lines indicated, were probed with the Ab57 (left panel).
The same samples were probed with the GR-Aspecific antibody, in
which case only the longer, GR-A, form was detected. The 83-kDa band
detected in HeLa extracts is a commonly observed GR degradation
product. Blots shown are representative of three or more experiments.
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Unlike the majority of steroid hormone receptors, the GR is primarily
localized in the cytoplasm in the absence of ligand (27). However, in
response to hormone signal, the functional GR undergoes nuclear
translocation. To examine potential functional differences between the
GR-A and -B isoforms, immunocytochemistry was performed to evaluate the
nuclear and cytoplasmic localization of the two proteins in the absence
and presence of hormone. Using this method, no difference is observed
between the hGR-A (M27T) and hGR-B (M1T) forms in nuclear translocation
in response to dexamethasone (Dex) (Fig. 5
).

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Figure 5. Immunocytochemical Analysis of hGR-A and hGR-B
The nuclear translocation of the GR-A and -B isoforms in response to
Dex was compared by immunocytochemistry. COS-1 cells were transfected
with either hGR-A (top panels) or hGR-B (bottom
panels) and plated on chamber slides the following day.
Approximately 48 h after transfection, cells in chamber slides
were treated for 2 h with 100 mM Dex (+) or vehicle
(-) as indicated. Cells were fixed on slides and analyzed by
immunostaining as previously described (22 ). Both the Ab57 and
GR-A-specific antibodies were used in these experiments. Data shown are
representative of immunostained cells from one experiment, which was
reproduced three separate times.
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Gene Activation by hGR-A and -B
To compare the cellular functions of hGR-A and hGR-B,
transactivation studies were carried out on both receptor forms using
transient transfections of hGR-A and hGR-B in COS-1 cells. Utilizing
three GRE-driven reporter genes, a striking difference in
transactivation was observed between the two hGR
isoforms. As shown
in Fig. 6A
, the hGR-B form is more than
1.5-fold as effective in transactivation from a single GRE-driven
reporter gene (GRE1-CAT) than the longer hGR-A form (Fig. 6A
). When two
GREs are found in tandem (GRE2-luc), the maximal transactivation
activity of hGR-B is enhanced to nearly 2-fold that of hGR-A (Fig. 6B
).
Finally, this functional difference is also observed using a mouse
mammary tumor virus (MMTV) promoter reporter gene, where the GR-B
transactivation is at least 1.4-fold greater than in hGR-A (Fig. 6C
).
These data suggest that GR-B, lacking the first 27 residues of GR-A,
exhibits enhanced transcriptional activity in a variety of promoter
contexts and argue for a general mechanism, not strictly dependent on
promoter sequences. Although the EC50 values vary
depending on the promoter used, none of our transactivation assays are
sensitive enough to detect a difference between GR-A and GR-B within 5
nM.

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Figure 6. Functional Comparison of hGR-A and hGR-B
The transactivation function of hGR-A and -B in response to Dex
was measured using three glucocorticoid responsive reporter genes
containing single or multiple copies of GRE-binding sites. A schematic
of each reporter gene is shown at the top of the data
set in which it was used. A, COS-1 cells were transfected with a
constant amount of expression vector (50 ng) for either hGR-A or hGR-B
as well as a constant amount of the GRE1-CAT reporter (0.5 µg). After
transfection, cells were treated with increasing concentrations of Dex
ranging from 10 pM to 100 nM and harvested
approximately 1620 h later for CAT activity assay. Data shown are an
average of three separate experiments in triplicate with the indicated
SEM. B, Cells were transfected as in panel A, but with a
constant amount of the GRE2-luc reporter (0.5 µg). Cells were treated
with Dex as indicated and harvested approximately 1620 h later for
luciferase activity assay. Data shown are an average of five separate
experiments with the indicated SEM. C, COS-1 cells
transfected as in panels A and B, but with a constant amount of the
MMTV-CAT reporter (0.5 µg). Cells were treated and harvested for CAT
assay as in panel A. Data shown are an average of triplicate samples
from a representative of three separate experiments, with the indicated
SEM. D, To contrast the hormone titration experiments,
cells were transfected with increasing amounts of either hGR-A or hGR-B
together with the GRE2-luc reporter and treated with a constant amount
of Dex (100 µM). Cells were harvested and analyzed for
luciferase production as in panel B. Data shown are an average of two
individual experiments with the indicated SEM. E, To
measure relative expression levels of hGR-A and hGR-B, COS-1 cells were
transfected with 2.5 µg of wt hGR as a control (lane 1), and hGR-A
(lanes 24), or hGR-B (lanes 57) in 10-cm plates for 20 h.
Cells were harvested in ice-cold RIPA buffer containing 5
mM DTT and protease inhibitors and immediately subjected to
SDS-PAGE and Western blot analysis with Ab57. The approximate molecular
masses of detected bands are shown to the right. The
83-kDa band, commonly observed in both transient and endogenous
receptor expression systems, is likely a product of degradation.
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To further address the functional differences in transactivation,
experiments were carried out in which the expression vector
concentrations were varied and hormone levels were kept constant. When
the GR-A and B levels were varied under saturating concentration of Dex
(100 nM), hGR-B was approximately twice as effective as the
hGR-A in transactivating the GRE2-luc reporter (Fig. 6D
). Similar
results are observed when using a luciferase reporter construct
containing the MMTV promoter. To test whether the functional
differences can be attributed to differences in protein expression of
hGR-A and hGR-B, equivalent amounts of the expression vectors were
transfected in separate wells and subjected to Western blot analysis.
Despite the higher transactivation capacity observed for hGR-B, Fig. 6E
shows that transfection of equivalent amounts of expression vector
consistently results in a greater accumulation of expressed hGR-A
protein. These data suggest that the observed transactivation
differences may be underestimates of the transcriptional potential of
hGR-B when considering the actual levels of expressed protein.
Interestingly, the level of the 83-kDa degradation product is also
significantly higher when hGR-B vectors are used for receptor
expression, suggesting that hGR-B may be more susceptible to
proteolysis than is hGR-A (Fig. 6E
). However, since we do not know the
activity of the 83-kDa band, we cannot eliminate the possibility that
it does contribute to the transactivation levels observed in Fig. 6
, AD.
Gene Repression by hGR-A and -B
A growing body of data suggests that many GRmediated effects
occur independently of direct DNA (GRE) binding (28). These processes
occur via cross-talk with other signaling pathways and through protein
interactions independent of a GRE. One well studied cross-talk pathway
is the mutual repression observed between hormone-activated GR and the
transcription factor NF-
B (29). The ability of hGR-A and hGR-B to
repress transactivation of the NF-
B p65 subunit was evaluated.
Interestingly, both receptor isoforms appeared to antagonize p65
reporter activity to the same degree (Fig. 7
) in contrast to the observed difference
in GRE-dependent transactivation. These data support the hypothesis
that hGR activation and repression functions are contained within
separate regions of the protein and suggest a role for the first 27
residues in transactivation but not transrepression of NF-
B.
Mechanism of the Alternative Translation Initiation of GR
The cause of alternative initiation of the GR transcript may lie
within the sequence itself. Eukaryotic ribosomes appear to select the
start site for translation initiation by a scanning mechanism (reviewed
in Ref. 30). An AUG start codon is classified as strong or weak
depending on the adherence to a specific surrounding consensus sequence
(Kozak sequence). A purine at position -3 and a G at position +4
relative to the AUG (the A is considered as +1) are considered strong
initiator sequences. However, leaky scanning, in which the first AUG is
bypassed in favor of a nearby downstream AUG, is most predictably
caused by deviations from the strong sequence context within the first
AUG (31). The sequence contexts of both AUG start sites of human,
mouse, and rat GR are shown in Fig. 8A
.
For all three species (human, rat, and mouse), the first AUG does not
contain a purine at position -3, indicating it is a weak initiator
sequence. However, the second AUG in all three species of GR does
contain a purine at this position, indicating it is actually a stronger
translation initiation site, and could explain the high degree of leaky
ribosomal scanning and the production of two GR proteins from the same
mRNA.

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Figure 8. Translation Scanning of GR Transcripts
A, Sequence analysis of GR cDNA surrounding the first two ATG codons.
The consensus Kozak sequence, required for maximum efficiency of
protein translation initiation, is shown above the sequence surrounding
the first two ATG codons from human, mouse, and rat GR. The
underlined ATG sequence represents bases +1, +2, and +3
respectively. The bases at position -3 and +4 have been found to
signal either weak or strong initiator codons, as indicated. The second
ATG in human corresponds to Met 27, while in rat and mouse it is Met
28. B, Mutagenesis of the Kozak sequences surrounding the two hGR ATG
start sites. The weak Kozak sequence at the upstream (Met 1) ATG was
changed to a strong Kozak consensus site (M1 Kozak mutant). COS-1 cells
were transfected with this mutant along with WT hGR, and the two ATG
mutants described elsewhere (M1T and M27T). Cells were harvested and
analyzed by Western blotting for hGR with Ab 57.
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To test the hypothesis that leaky ribosomal scanning is the cause for
the production of GR A and B isoforms, we mutated the Kozak sequence of
the first AUG start site. When the "weak" consensus site of the
first AUG (Met 1) is changed to a "strong" one by a point mutation
at the -3 position (C to G), production of GR-B is completely lost
(Fig. 8B
). These data conclusively demonstrate that leaky ribosomal
scanning of a weak initiation sequence generates a second, B form, of
the GR.
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DISCUSSION
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The data presented in this paper demonstrate that the GR is a
product of alternative translation initiation, which results in the
production of both a GR-A and GR-B form. Both the A and B forms are
generated in approximately equivalent levels from a single cDNA when
expressed in vitro using reticulocyte lysates. However, the
GR-B form appears to be more susceptible to degradation, at least when
expressed in mammalian cells. Translation of an hGR
message from an
internal AUG codon, corresponding to methionine 27, results in a
protein with twice the amount of maximal transactivation as the longer
protein initiated from the first AUG codon. Interestingly, deletion of
the first 2530 residues of hPR-B also results in more effective
transcriptional activation (Horwitz, K., and L. Tung, personal
communication). These data suggest that the extreme amino termini of
steroid receptors may be involved in regulating receptor function.
One possible explanation for the increased activity of the shorter GR-B
is that the tertiary structure including the first 27 residues masks
the activation function(s) associated with other regions of the
receptor. A second possibility is that this region of the GR is
essential for an important protein interaction responsible for either
transcriptional silencing or repression. The first 27 residues of hGR,
however, do not appear to have significant homology with recognized
protein-protein interaction sites or other known functions. The fact
that the GR-B is a more effective transactivator on three separate
glucocorticoid-responsive promoters argues for a general mechanism
involving GR interactions with additional cellular factors. It remains
to be seen whether both isoforms homo- and heterodimerize equivalently,
bind DNA with the same affinities, and respond to different hormone
signals with the same relative potency. In addition, the tissue
distribution of the two isoforms remains to be elucidated.
It is now established that other steroid receptors produce N-terminal
truncation variants. For example, it is well known that two PRs are
derived from the same gene in humans and chickens (32). It was
originally suggested that the origin of the PR isoforms was alternative
translation initiation from the same message (17, 33). However,
additional studies with human breast cancer cell lines and hPR or cPR
cDNA suggest that the PR-A and -B isoforms may also be generated from
distinct promoters (18, 34, 35). The data presented in this paper
clearly show the production of two GR protein products from a single
cDNA source. In addition, Northern blot analysis of hGR
-transfected
COS-1 mRNA shows only a single hGR-specific message, further supporting
the one-message/twoprotein hypothesis (36). The androgen receptor
(AR) has also been shown to exist as multiple forms, differing at the
amino terminus, which are expressed in a tissue-specific manner (37).
Alternative initiation has been implicated as the mechanism responsible
for the generation of these two AR forms (38).
The mechanism of alternative initiation of the GR is shown to be under
the translational direction exerted in the sequence surrounding the ATG
codons (31). As would be expected, creation of a strong Kozak consensus
site (-3 C to G) by a point mutation 3' of GR Met1 AUG completely
abolished ribosomal read-through and production of GR-B. This new GR
expression construct contains no coding region mutations and functions
similarly to the M27T hGR used in these studies. That a point mutant in
the noncoding, promoterless region of the GR cDNA had such a drastic
effect on protein expression and function is remarkable and
unprecedented in the nuclear receptor field. Recent interest in mRNA
regulation mechanisms, such as splicing, stability, and the role of
structure, are likely to lead to increased analysis of translational
control mechanisms as identified here.
Alternative translation initiation produces two functionally distinct
forms of the GR. The presence of both forms in several human cell lines
and rodent tissues suggests that the generation of these two protein
species may be a general phenomenon. However, the ultimate
physiological significance of these receptor isoforms remains to be
established. Although our data, utilizing both in vitro
translation and transient expression systems, suggest both the GR-A and
-B are being expressed via leaky ribosomal scanning from a single cDNA
and corresponding mRNA, we cannot rule out the existence of alternative
promoters regulating expression in vivo, as has been shown
for the PR (35). We have presented evidence suggesting that both GR
products are generated in vivo and in a variety of mammalian
cell lines and that a significant functional difference exists between
the two. It is intriguing to speculate whether differences exist
between GR-A and GR-B in tissue distribution and/or expression during
development, aging, or cell death. Such studies, however, will require
currently unavailable antibodies that selectively recognize the shorter
GR-B form in the presence of GR-A. For example, it is now known that
the PR-A and -B isoforms function in a tissue-specific fashion (39) and
that both isoforms regulate a distinct subset of genes (40). Regulated
expression of either GR isoform in favor of the other would suggest a
physiological role of alternative translation initiation of the GR.
Indeed, there are several reports that present evidence for
physiological regulation of alternative translation initiation of
critical transcription factors and cell cycle regulators (41, 42). The
potential for differential regulation of functionally distinct GR
isoforms, at the level of translation, is an area that clearly needs
further inquiry.
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MATERIALS AND METHODS
|
---|
Materials, Antibody Production, and Plasmids
trans-35S-label (1,108
Ci/mmol) was purchased from ICN Biochemicals, Inc.
(Irvine CA). [14C]Chloramphenicol (4060
mCi/mmol) was obtained from NEN Life Science Products
(Boston, MA). Dex was supplied by Steraloids (Wilton, NH).
Acetyl-coenzyme A and protease inhibitors were purchased from
Roche Molecular Biochemicals (Indianapolis, IN).
Oligonucleotide primers for mutagenesis and PCR were synthesized by
Oligos Etc. (Bethel, ME). The hGR-A-specific antibody was produced by
Covance Laboratories, Inc.(Denver, PA) using a synthetic
peptide antigen corresponding to residues 322 of the hGR synthesized
at the University of North Carolina at Chapel Hill. The GR-A- specific
antibody was purified on a peptide-linked sepharose affinity column as
previously described for antibody purification (24). Characterization
of this antibody is described below. The peroxidase-labeled secondary
antibodies and enhanced chemiluminescence (ECL) reagents were purchased
from Amersham Pharmacia Biotech (Piscataway, NJ).
Use of the hGR mammalian (pCMV-hGR
) and in vitro
(pT7-hGR
) expression vectors was described in a previous publication
(23). The mouse GR mammalian and in vitro expression vectors
were also used as previously described (43). The rat GR expression
vector, pRSV-rGR, was a gift of Dr. Trevor Archer and was used for both
in vitro and COS-1 expression systems. The reporter plasmids
GRE1- and GRE2-CAT (44), and pGCMS-CAT (45) have been described
elsewhere. All site-specific mutagenesis was done with the Quick Change
Mutagenesis kit (Promega Corp., Madison WI), following
their protocol for primer design. The carboxyterminal hGR
truncation mutants, hGR(1742) and hGR(1706), were created by
mutating the codons at positions 743 and 707 to TGA stop codons. All
mutants were verified by DNA sequencing.
Cell Culture, Transfections, Luciferase, and Chloramphenicol
Acetyltransferase (CAT) Assays
COS-1 and HEK293 cells were maintained in DMEM with high glucose
containing 2 mM glutamine and 10% (vol/vol) mixture of
heat-inactivated FCS/calf serum (1:1). For transactivation assays,
cells were incubated for 12 days in media containing dextran-coated
charcoal-stripped sera to remove endogenous steroids. HeLa cells were
maintained in Eagles MEM supplemented with glutamine and 10%
FCS/calf serum. CEM-C7 cells were grown in suspension in RPMI 1640
medium supplemented with 2 mM glutamine, 10% (vol/vol)
heat-inactivated FCS, and 0.1 M HCl. All cell culture media
contained 100 IU/ml penicillin and 100 mg/ml streptomycin. Cell
cultures were maintained in a 5% CO2 humidified
incubator at 37 C and passaged every 34 days. All transfections were
carried out with Mirus TransIT LT-1 reagent according to the
manufactures protocol (Pan Vera, Madison WI). An appropriate amount
of TransIT reagent (3 µl per µg of transfected plasmid) was added
to OPTIMEM (Life Sciences, Inc., St. Petersburg, FL) for 5 min.
Purified plasmid DNA was then added and allowed to complex for 30 min
at room temperature, before being added to cells with media containing
stripped serum. Six to eight hours after transfection, the media were
replaced with fresh serum-stripped media containing vehicle or Dex.
Transfected cells were incubated in the presence or absence of the
indicated amount of Dex for 1824 h before harvesting. Cells for
luciferase assays were harvested in 1x Reporter Lysis Buffer
(Promega Corp.). Total protein was measured using the
Bio-Rad Protein Assay reagent (Bio-Rad Laboratories, Inc.,
Hercules, CA) according to the manufactures protocol, and equivalent
amounts of total protein were used for luciferase activity assays. The
luciferase activity was measured using the 96-well plate format with an
MLX automated microtiter plate luminometer from Dynex.
CAT assays were carried out essentially as described previously (45).
Approximately 0.10.2 mg of protein extracts were incubated overnight
at 37 C with 1 mM acetyl-coenzyme A and 0.1 µCi of
[14C]chloramphenicol in Tris-EDTA (TE).
Samples were extracted in mixed xylenes and then back extracted one
time with TE, pH 8.0, before liquid scintillation counting. A standard
curve was generated using commercially available, purified CAT as
described by the manufacturer (Promega Corp.). All
experiments were conducted under conditions in which substrate was in
excess and the relationship of counts per min to CAT activity was
linear. Data are expressed as counts per min per microgram of total
protein.
Animals
Male Sprague Dawley rats (23 months old) and C57BL mice (6
months old) were used in all experiments. All animals were maintained
under controlled conditions of temperature (25 C) and lighting and
allowed free access to food and saline. All experimental protocols were
approved by the animal review committee at the institute and were
performed in accordance with the guidelines set forth in the NIH Guide
for the Care and Use of Laboratory Animals published by the USPHS. Rats
were killed by decapitation and mice were asphyxiated under
CO2, before removal of liver tissue. Liver tissue
fragments were homogenized on ice for 30 sec at maximum speed with a
Tekmar Tissuemizer in a radioimmunoprecipitation assay (RIPA) buffer
(50 mM Tris-HCl, pH 8.0, 0.1% SDS, 1% Triton x-100,
0.5% sodium deoxycholate, 2 mM EDTA, and 150
mM NaCl) containing 5 mM dithiothreitol (DTT)
and protease inhibitors. After a brief low-speed centrifugation to
remove tissue debris, extracts were incubated for an additional 20 min
on ice before centrifugation at 20,000 x g for 20 min.
The resulting supernatants were measured for total protein
concentration (typically 1020 mg/ml) and subjected to Western
blotting as described below.
Immunocytochemistry and Western Blotting
Immunocytochemistry was carried out essentially as previously
described (22). The day after transfection, cells were split on
two-chamber glass slides. Approximately 48 h after transfection,
the cells were treated with 100 nM Dex or vehicle for
2 h. The cells were fixed in 2% paraformaldehyde, washed in PBS,
and permeabilized with 0.2% Triton X-100. Cells were again washed in
PBS, treated with 2% normal goat serum, washed in PBS, and incubated
with epitope-purified Ab57 (1:7500) for 20 h at 4 C. The cells
were washed in PBS and incubated with biotinylated goat antirabbit IgF
(1:400) for 1 h at room temperature. Immunoreactivity was
visualized by staining with avidin-biotin-peroxidase.
For Western blotting, cells were lysed for 20 min on ice in RIPA buffer
containing 150 mM NaCl, 5 mM DTT, and protease
inhibitors (0.1 mM Pefa Block, 1 µM
leupeptin, and 1 µM pepstatin). After a high-speed
spin to remove cellular debris, total protein was measured using the
Bio-Rad Laboratories, Inc. detection kit. Unless indicated
otherwise, 50 µg of protein extract were then separated on precast
8% Tris-glycine gels (Novex, San Diego CA) and
transferred to nitrocellulose. The membranes were washed in TBST
(Tris-buffered saline with 0.1% Tween-20) and blocked in TBST
containing 5% nonfat milk for a minimum of 2 h at room
temperature. Blots were next incubated in the same solution
supplemented with affinity-purified primary antibodies, Ab57 (1:2,500)
and GR-A specific (1:5,000), overnight at 4 C. After extensive rinsing
and washing in TBST (three times, 1015 min), the blots were probed
with peroxidase-conjugated goat antirabbit secondary antibody
(1:10,000) for 2 h at room temperature. Bands were visualized
using ECL reagents (Amersham Pharmacia Biotech).
Repression of NF-
B/p65
GR-mediated repression of NF-
B/p65 transactivation was
studied as reported previously (44). A constant amount (12.5 ng) of the
NF-
B-p65 subunit expression vector (pCMVp65) was used, in
conjunction with the NF-
B-luciferase reporter, 3XMHC-luc (0.5 µg),
to measure p65-mediated transactivation. The repression of p65 activity
by GRs was measured by cotransfecting increasing amounts (10, 50, and
250 ng) of the hGR-A or hGR-B expression vectors. After transfections,
cells were treated without or with 100 nM Dex.
Approximately 1620 h after hormone treatment, cells were harvested
and luciferase activity was determined as described above. Equivalent
amounts of total protein were assayed for luciferase activity as
described above, and data from individual experiments were averaged and
presented along with the SEM.
 |
ACKNOWLEDGMENTS
|
---|
We wish to thank Dr. Trevor Archer for the rGR plasmid, Dr. Turk
E. Rhen for supplying the rat liver tissue, and Dr. Ester Carballo-Jane
for the mice. The authors would also like to thank Chris Jewell for
technical assistance and discussions, and Drs. J. Hall and H. Kinyamu
for reviewing the manuscript.
 |
FOOTNOTES
|
---|
Address requests for reprints to: Dr. John Cidlowski, Laboratory of Integrative Biology, National Institute of Environmental Health Sciences, National Institutes of Health, 111 TW Alexander Drive NC F307, Research Triangle Park, North Carolina 27709-2233. E-mail:
cidlowski{at}niehs.nih.gov
Received for publication November 22, 2000.
Revision received March 15, 2001.
Accepted for publication April 2, 2001.
 |
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