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INSERM U 478, Faculté de Médecine Xavier Bichat, 75870 Paris Cedex 18, France
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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5,6, resulting from an alternative splicing event
skipping exons 5 and 6 of the human MR gene. hMR5,6 mRNAs are
expressed in several human tissues at different levels compared with
wild-type human MR, as shown by real time PCR. Introduction of a
premature stop codon results in a 75-kDa protein lacking the entire
hinge region and ligand binding domain. Interestingly, hMR5,6 is
still capable of binding to DNA and acts as a ligand-independent
transactivator, with maximal transcriptional induction corresponding to
approximately 30–40% of aldosterone-activated wild-type human MR.
Coexpression of hMR5,6 with human MR or human GR increases their
transactivation potential at high doses of hormone. Finally, hMR5,6
is able to recruit the coactivators, steroid receptor coactivator 1,
receptor interacting protein 140, and transcription intermediary factor
1
, which enhance its transcriptional activity.
Ligand-independent transactivation and enhancement of both wild-type MR
and GR activities by hMR5,6 suggests that this new variant might
play a role in modulating corticosteroid effects in target
tissues. |
INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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We have previously shown that the human MR (hMR) gene is composed of 10
exons (6). Alternative transcription of two
5'-untranslated exons generates two mRNA isoforms, hMR and hMRß,
which are coexpressed in aldosterone target tissues (6, 7). hMR gene expression is controlled by two different
promoters, which differ by their basal activity as well as their
hormonal regulation (8). Recent experiments in transgenic
mice have shown a distinct tissue-specific utilization and activity of
the two hMR-regulatory regions in vivo (9).
Mineralocorticoids are mainly implicated in the maintenance of water
and salt homeostasis by regulating vectorial sodium transport in tight
epithelia, thus regulating blood pressure (10). Recently,
several other effects have been described, including a role for
aldosterone in the development of cardiac fibrosis
(11, 12, 13) and the differentiation of brown adipose tissue
(14). Given the pleiotropic effects mediated by MR and the
fact that the receptor possesses the same affinity for
glucocorticoids as for mineralocorticoids, the mechanisms mediating
cellular specificity become fundamental for the final transcriptional
response. In epithelial target tissues, specificity is acquired by the
presence of an enzyme, 11ß-hydroxysteroid dehydrogenase type II
(11HSD2), which converts glucocorticoids (circulating at 
1,000- fold
higher levels than mineralocorticoids) into inactive 11-dehydro
cogeners (15, 16). In nonepithelial tissues, such as the
brain, the heart, and brown adipose tissue, 11HSD2 activity appears
insufficient to allow for receptor selectivity, and it has been
postulated that MR might function as a high-affinity GR
(17). Nevertheless, specific aldosterone effects have also
been demonstrated in these tissues, indicating that other mechanisms
may exist regulating either hormonal access to the receptor or the
receptor response to a given hormone. In this context it has been shown
that MR can discriminate aldosterone from glucocorticoids independently
of 11HSD2 in terms of transcriptional response, due to differences in
the association/dissociation kinetics (18, 19). In
addition, heterodimerization between MR and GR, which could indeed
modulate the response to one or the other corticosteroid hormones in
target cells, has recently been reported (20, 21).
Finally, the existence of receptor splice variants seems to play a
major role in modulating receptor function. It has been shown that a
binding-incompetent 3'-splice variant of the human GR, hGRß, acts as
a dominant negative regulator not only of GR, but also of MR function
(22).
We have previously hypothesized the existence of other human MR (hMR)
transcripts, expressed in heart and epidermis (7). Given
the involvement of aldosterone in cardiovascular disease, we have
searched for new cardiac hMR isoforms, which could eventually modulate
hMR activity and hence aldosterone effects in the heart. In this paper
we describe a new hMR splice variant, hMR5,6, which lacks exons 5
and 6 of the hMR gene, resulting in a protein deleted of the entire
hinge region and ligand-binding domain. hMR5,6 is widely expressed
in human tissues and acts as a ligand-independent transcription factor,
capable of modulating hMR and hGR function. Finally, we show that
hMR5,6 interacts with different coactivators, which are able to
enhance its transcriptional potential.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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, numbering based on the nucleotide sequence published in Ref.
2). The clone contains 212 nucleotides of the
5-untranslated exon 1 and 477 nucleotides of 3'-untranslated
region. Comparison with the genomic sequence (6)
showed that the deleted fragment corresponds to exons 5 and 6,
indicating that the mRNA variant is generated by an alternative
splicing event. Deletion of the two exons introduces a frame-shift in
the hMR sequence, resulting in a premature termination codon 107 bp
downstream of the exon 4/intron D boundary (Fig. 2, A and B). The new mRNA codes for a
protein of 706 residues with a predicted size of 75 kDa, which
possesses the entire amino-terminal part of the receptor and the
DNA-binding domain, but has lost the hinge region and the
ligand-binding domain. Instead, it contains 35 additional residues
without homology to any other known sequence. In vitro
transcription/translation analyses confirmed that a protein with the
expected molecular mass is generated from this transcript (Fig. 3). No specific binding of tritiated
aldosterone or dexamethasone was detected, confirming the absence of a
competent ligand-binding pocket (data not shown). Some differences in
the sequence between hMR5,6 and the wild-type hMR have been
detected: at nucleotide position 221 (corresponding to the first
noncoding nucleotide of exon 2), G is changed to C in hMR5,6. An
A-to-G transition at positon 760 changes amino acid 180 from Ile to
Val, while a C-to-T transition at nucleotide position 944 changes codon
241 from Ala to Val. Finally, a silent C-to-T transition was found at
nucleotide position 1,719 (499 Asp/Asp). Substitution of Val 241 for an
Ala should not have a major influence on protein structure, since the
two residues have similar properties and are located outside the
putative hMR AF-1, which has been mapped between positions 328 and 382
(23). These nucleotide changes correspond to common
polymorphisms found in the normal population. In particular, C221C and
T1719 have frequencies of 39% and 15%, respectively
(24), and the Val241 mutation was reported to have
heterozygosity and homozygosity frequencies of 48% and 38%,
respectively (25), while it was not detected in another
study (24).
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5,6 Is Differentially Expressed among Tissues5,6, we
performed RT-PCR analysis on RNAs extracted from different human
tissues and two human immortalized cell lines using sense and antisense
primers located in exons 4 (S2136) and 7 (A2824), respectively (Fig. 4A). Two PCR products of the expected
size for the wild- type hMR (688 bp) and for hMR5,6 (192 bp) were
detected in all tissues analyzed. In this experiment, hMR5,6 was
highly expressed in the hippocampus and in the hepatic cell line SK
Hep1, whereas its expression seems very low in the colon. In all
other tissues, including lung, kidney, heart, and lymphocytes,
substantial amounts of hMR5,6 were detected. Therefore, both hMR
variants are expressed in human tissues. Interestingly, an additional
band of approximately 330 bp was also detected by RT-PCR in all
tissues, with the exception of the hepatic cell lines Hep3B and SK
Hep1. Given the size, we supposed that this band could result from the
amplification of another hMR splice variant lacking exon 5. Indeed, a
cDNA containing an internal 351-bp deletion comprising nucleotides
2,237–2,587 was originally isolated by Arriza et al.
(2) during the initial cloning of the hMR cDNA, and
alignment with the genomic sequence (6) has subsequently
shown that the deleted fragment corresponds to exon 5
(26). Sequencing of the approximately 330-bp amplification
product confirmed the existence of the variant lacking exon 5
(hMR5), which encodes for a predicted receptor lacking amino acids
672–788, including the whole hinge region and the N-terminal part of
the ligand-binding domain of the receptor. However, since the
full-length clone of this variant was not isolated during our library
screening, precise exon composition, i. e. presence of
untranslated exons 1 or 1ß, and its functional properties remain
to be established.
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5,6, we have
performed real-time PCR experiments using a Taqman probe. The hMR
and hMR5,6 common forward primer is located on exon 4, whereas
the reverse primers for the two hMR isoforms were chosen to overlap the
junction between alternatively spliced exons. The Taqman probe is also
located on exon 4. Thus, specific detection of each isoform during
real-time PCR is obtained exclusively by the specificity of the reverse
primer. Several tests of specificity were conducted. No fluorescent
signal was observed in PCR samples without a previous reverse
transcription. The PCR primers amplified only their cognate template
and produced no detectable amplification from the cDNA of the other
isoform (data not shown). Real-time PCR analysis allowed detection of
as little as 6 molecules of template, with a mean threshold cycle
(CT) of 37.51 for hMR5,6 and 36.15 for hMR.
Under these conditions, as illustrated in Fig. 4B
, the relative
hMR5,6 expression compared with that of hMR clearly differs among
tissues, with highest levels observed in the kidney. In three
independent experiments using kidney cDNA, the threshold cycle
CT for hMR5,6 was 33.33 ± 0.30 (n
= 8). Using the same primers, a fluorescent signal for hMR and
hMR5,6 was also detected in mouse kidney, indicating that expression
of hMR5,6 is conserved among species (data not shown).
The hMR5,6 Isoform Is Able to Bind to DNA
The DNA binding properties of hMR5,6 were next
investigated by EMSA. Whole-cell extracts were prepared from RCSV3
cells after transfection with the expression plasmids coding for
hMR5,6 and from Sf9 cells expressing hMR, and assayed for binding to
a consensus GRE sequence from the MMTV promoter. As previously shown
(3), hMR specifically bound to a consensus
32P-labeled GRE oligonucleotide (Fig. 5A, lane 8). hMR5,6 was also able to
bind to the consensus GRE in a specific manner, since binding was
efficiently competed for by excess amounts of unlabeled GRE (Fig. 5A, lanes 2–7). Comparison of competition experiments between the two
isoforms indicated that hMR5,6 possesses a similar affinity for the
GRE as hMR (data not shown). GRE-hMR5,6 complexes display a faster
mobility than GRE-hMR, consistent with the lower molecular mass of the
splice variant. The same results were obtained using in
vitro translated hMR5,6 (Fig. 5B). Coincubation of whole-cell
extracts (WCE) of hMR5,6 with different amounts of wild-type hMR
(data not shown) or cotranslation of hMR and hMR5,6 in the
reticulocyte lysate system (Fig. 5B, lanes 6 and 7) did not generate
additional retarded bands, indicating that in our experimental
conditions, hMR5,6 does not interact with hMR and does not form
heterodimers on the consensus GRE. Our results demonstrate that,
although hMR5,6 lacks the entire hinge region and ligand-binding
domain, it is competent for DNA binding.
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5,65,6, we tested its functional properties by transient
transfection assays in RCSV3 cells. For this purpose, different amounts
of a hMR5,6 expression vector were cotransfected with an
MMTV-luciferase reporter plasmid. In these experiments, hMR5,6 was
able to activate transcription from the MMTV promoter in a
dose-dependent manner, reaching a transactivation plateau with 0.5
µg of transfected plasmid (Fig. 6A).
Maximal activation corresponded to approximately 30–40% of luciferase
activity observed when transfecting wild-type hMR in the
presence of 10-9 M aldosterone.
Furthermore, as expected by the absence of the ligand-binding domain,
the transactivation activity of hMR5,6 was totally hormone
independent, since it was not modified by incubation with increasing
doses of aldosterone (Fig. 6B). These data were also confirmed by
transient transfections in CV1 cells (data not shown).
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5,6 could influence the activity of
the wild- type hMR. Three different amounts of hMR5,6 (0.05 µg,
0.5 µg, and 2.5 µg) were transfected in RCSV3 cells together with
0.5 µg of hMR and an MMTV-luciferase reporter plasmid (Fig. 7A). In the presence of increasing
concentrations of aldosterone, hMR activated transcription in a
dose-dependent manner, with an ED50 of
approximately 10-11 M, reaching a
plateau of activation at concentrations of aldosterone between
10-10 and 10-7
M. As expected, in the presence of hMR5,6,
transcriptional activation already occurred in the absence of hormone,
but only starting from a hMR5,6 to hMR ratio of 1. At lower ratios,
hMR seems to inhibit ligand-independent transcriptional activation by
hMR5,6. These data were also confirmed using 2.5 µg of hMR and
1.25 or 2.5 µg of hMR5,6 (data not shown). At a hMR5,6 to hMR
ratio of 0.1 and 5, the dose-response curve to aldosterone was not
markedly modified, although the maximum levels of transcriptional
activation corresponded to approximately 120% of hMR plus aldosterone
alone. At a 1:1 hMR5,6 to hMR ratio, a significant increase in
transactivation was observed (200% of hMR in the presence of
10-7 M aldosterone) with no evident
plateau. Therefore, these data indicate that in RCSV3 cells hMR5,6
modulates transcriptional activation of hMR depending on the relative
level of the splice variant.
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5,6 could
also influence hGR-mediated transactivation. We therefore performed
cotransfection assays using wild-type hGR alone or in the presence of
an equal amount of hMR5,6 (Fig. 7B). While the dexamethasone
dose-response curve of hGR decreased to 60% of maximal levels at high
doses of hormone (P < 0.001 for
10-7 M and
10-6 M dexamethasone
vs. 10-8
M), in the presence of hMR5,6 approximately
100% of activation was maintained even in the presence of high
concentrations of hormone. Note that very little relative
hormone-independent transactivation was observed with hMR5,6
(expressed as percentage of hGR plus dexamethasone); this is due to the
fact that hGR is a stronger transactivator than hMR. Altogether, these
results indicate that hMR5,6 is also able to potentiate hGR-mediated
transcriptional activation.
hMR5,6 Activates Transcription by Interacting with
Transcriptional Coactivators
The next question was how hMR5,6 was able to activate
transcription in the absence of a ligand-binding domain and a
functional AF-2. We speculated that hMR5,6 could interact with
transcriptional coactivators through its amino terminal domain, as
already reported for the PR (27), ER and ERß
(28, 29), and TRß2 (30). To test this
hypothesis, hMR5,6 was cotransfected in RCSV3 cells either alone or
with the expression plasmids for different transcriptional
coactivators, including steroid receptor coactivator (SRC)1a and SRC1e
[two isoforms of SRC1 diverging at their C termini
(31)], RIP140 (32), and TIF1
(33). A significant increase in transactivation was
observed in the presence of all four coactivators, although to a
different extent (Fig. 8). Whereas SRC1a
and RIP140 increased transcriptional activation of hMR5,6 only by
approximately 50%, SRC1e and hTIF1 enhanced the ability of
hMR5,6 to stimulate transcription by more than 2-fold. These results
are consistent with previous data showing that SRC1e enhanced the
ability of the ER to stimulate transcription to a greater extent than
SRC1a (31). Neither coactivator alone had any effect in
the absence or presence of aldosterone (data not shown).
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5,6 was tested by
glutathione S-transferase (GST) pull-down assays using
in vitro translated hMR5,6. As shown in Fig. 9, a strong interaction could be observed
when hMR5,6 was incubated with TIF1, whereas weaker, although
visible, signals are present with GST-SRC1 and GST-RIP140. As expected,
this interaction was independent of the addition of aldosterone. Taken
together, these results strongly suggest that hMR5,6 activates
transcription by recruiting transcriptional coactivators probably via
its amino-terminal domain.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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5,6, was
isolated, which arises from the skipping of exons 5 and 6 of the hMR
gene and is also observed in mice. hMR5,6 transcripts are present in
several human tissues, including lung, kidney, heart, liver,
lymphocytes, and brain, but their relative expression compared with the
wild-type hMR is variable. Relative quantitative real-time PCR analysis
has shown that the hMR5,6 to hMR ratio is approximately 5-fold
higher in the kidney than in the heart, with intermediary levels
observed in brain and liver. Although these results suggest that
hMR5,6 modulates corticosteroid effects in a tissue-specific manner,
extensive analysis of isoform expression in several individuals and in
normal and pathological states should give a more precise insight into
the mechanisms of regulation of hMR expression and splicing.
The 75-kDa hMR5,6 protein, which lacks the entire receptor hinge
region and the ligand-binding domain, but contains unique 35 residues
after the DNA-binding domain, not only binds to a consensus GRE but is
also able to activate transcription in a ligand-independent manner from
the MMTV promoter. In cotransfection assays, we have also shown that
hMR5,6 is able to potentiate the transcriptional activation induced
by wild-type hMR and hGR, particularly at high doses of hormone, where
these receptors alone either reach a transactivation plateau, or
display a progressive decrease of their transactivation potential.
Finally, transcriptional activation by hMR5,6 seems to involve
recruitment of transcriptional coactivators by the amino-terminal
domain of the receptor.
Different MR isoforms have already been described (26). In
addition to hMR and hMRß, three rat MR mRNA variants, rMR,
rMRß, and rMR
, have been identified, which are generated by
alternative transcription of different 5'-untranslated exons
(37). In addition, two MR splice variants changing the MR
coding region have recently been described. A 12-bp insertion resulting
from the use of a cryptic splice site at the exon 3/intron C splice
junction leads to an in-frame insertion of four amino acids to the
first zinc finger of the DNA-binding domain of the receptor
(38). Although this insertion variant might possess
slightly different DNA binding characteristics and transactivation
properties than MR, it is interesting to note that insertion of nine
amino acids between the two zinc fingers of the trout GR does not
significantly alter its functional characteristics (39, 40). Another MR variant with a 10-bp deletion results in a
truncated receptor lacking the C-terminal part of the steroid binding
domain. This isoform is largely expressed in rat and human tissues but
has no intrinsic activity nor does it modify transcriptional activity
of the wild-type MR (41).
Other than an exon 5-deleted ER splice variant (42),
ER5, whose role in carcinogenesis has been strongly suggested
(43), hMR5,6 is the only example of a naturally
occurring C-terminal deletion mutant acting as ligand-independent
transactivator and capable of positively modulating activity of the
wild-type receptor. Indeed, similar splice variants of TR
(44), VDR (45), or the recently identified
truncated form of PPAR (PPARtr) (46) exhibit a
dominant negative activity on the wild-type receptor. In particular, it
is interesting to note that PPARtr, which arises from skipping of
exon 6 of the human PPAR gene, possesses a structure very close to
that of hMR5,6 but is unable to bind to a consensus DNA element and
to activate transcription. This difference might be due to the fact
that for PPAR the C-terminal part of the receptor is required for
heterodimerization and therefore for DNA binding (47). In
MR, which binds as homodimer or MR/GR heterodimer to a GRE (3, 20, 21), one region involved in dimerization has been mapped to
the DNA-binding domain (21), which is conserved in
hMR5,6. However, no heterodimerization between hMR5,6 and hMR was
observed under our experimental conditions. This implies that either
hMR5,6 binds DNA and activates transcription as a monomer or that
elements conserved in the DNA-binding domain (DBD) are sufficient for
homodimerization but not for heterodimerization, which requires regions
located in the hinge region or in the ligand-binding domain (LBD).
Furthermore, our results indicate that the observed modulation of
wild-type hMR or hGR effects by hMR5,6 might result from synergistic
activation of response elements or from modification of intracellular
receptor localization, rather than from heterodimerization. Additional
experiments, investigating interactions of hMR5,6 with DNA and
intracellular trafficking of the receptors, are needed to fully
elucidate this aspect.
Concerning the particular functional characteristics of hMR5,6, it
is also worth noting that truncation of the ligand binding domain of
the GR yields a constitutively active protein with partial
transcriptional function (48). However, the presence of a
short amino acid stretch after the DNA-binding domain is required for
transactivation. In this context it might be hypothesized that the 35
additional C-terminal residues of hMR5,6, although unrelated, are
important for the correct folding of the two zinc fingers. The lack of
the C-terminal part of hMR not only abolishes the ligand- binding
domain, which in the absence of hormone normally locks the receptor in
an inactive conformation, but also the region interacting with heat
shock protein 90, which acts as a molecular chaperone for many steroid
receptors and could also play a role in the nucleo-cytoplasmic
shuttling (49). Indeed, whereas the other steroid
receptors are localized in the nucleus in the absence of hormone, MR
has been shown to reside predominantly in the cytoplasm in the absence
of ligand and to translocate into the nucleus upon addition of
aldosterone (50, 51). Given that a hormone-independent
nuclear localization signal is present in the second zinc finger of the
rat GR DBD (52), one could speculate that an analogous
signal allows hMR5,6, whose activity is no more repressed by the
LBD, to be present in the nucleus in the absence of ligand.
Our study has shown that hMR5,6 is capable of recruiting
coactivators such as SRC1, TIF1, and RIP140 that enhance its
transcriptional property. Although these factors were isolated for
binding to the LBD of nuclear receptors, including hMR
(53), in the presence of hormone, it has subsequently been
shown that SRC1 interacts also with the A/B domain of the PR in a
ligand-independent manner and that it is able to mediate
transcriptional enhancement by both the AF1 and AF2 of ER and GR to a
similar extent (27). Furthermore, SRC1 can be recruited in
a ligand-independent manner by ERß through phosphorylation of the AF1
(28). Two particular serine residues, one of which is
contained in a motif also present in other steroid receptors, are
critical for this interaction. By analogy, TR-ß2 binds CREB-binding
protein, SRC1, and pCIP in the absence of thyroid hormone through a
domain located between amino acids 1–50 of the receptor, whereas this
is not the case for TR-ß1 and TR-1 (30). Alignment of
the amino-terminal regions of ERß, TR-ß2, PR, GR, and MR shows some
amino acid conservation, in particular a GXP motif at position 232 and
a SP motif at position 283 of the hMR, the latter being localized in
the AF1 core of the GR (54). These motifs might correspond
to putative structural segments responsible for interaction with
coactivators. Indeed it was suggested that the proper assembly of the
individual activation functions AF1 and AF2 is necessary to render the
steroid receptor-DNA complex transcriptionally productive
(55), thus allowing functional synergism between AF1 and
AF2 (29), and that this assembly might depend on
coactivators (27).
In conclusion, it appears that corticosteroid function in target tissues is regulated by a complex cascade of events. In addition to regulation of intracellular hormone availability, differential transcriptional regulation of hMR expression by alternative promoter usage, and interaction of GR and MR variants, the generation of MR isoforms by alternative splicing appears to modulate the final transcriptional response. Although MR- and GR-mediated effects are not redundant, as shown by inactivation of single receptors in mice (56, 57), interaction between the mineralocorticoid- and glucocorticoid-signaling pathways certainly has major functional significance in many physiological processes. It will be of interest to analyze whether modifications of expression of hMR isoforms are involved in pathological processes.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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to screen a 96-well master plate containing
5,000 cDNA clones per well; the other primers were located in exons 6
and 8, respectively. hMR primers were as follows [23-mers, numbering
corresponds to the position of the 5'-nucleotide according to the
published hMR cDNA sequence (2), S identifies primers in
the sense orientation and A in the antisense orientation]: A180; S2601
and A2980; the vector-specific primer VP3 was
5'-GCAGAGCTCGTTTAGTGAACC-3'. Ninety six-well subplates containing
bacterial glycerol stocks of cDNAs contained in positive wells diluted
at 50 clones per well were subsequently screened by PCR using the same
primer pairs. PCR was performed using Platinum Taq DNA
polymerase (Life Technologies, Inc., Paisley, UK) in the
following conditions: one step at 94 C for 5 min followed by 35 cycles
of 94 C for 45 sec, 60 C or 56 C for 45 sec, and 72 C for 45 sec, and
one step at 72 C for 7 min. Bacterial glycerol stocks of positive wells
were seeded at a density of 5,000 colonies per Petri dish, grown at 30
C overnight and hybridized with hMR exon-specific probes or with a
1.7-kb EcoRI hMR restriction fragment using standard
techniques. DNA of positive clones was analyzed by restriction enzyme
digestion with NotI and EcoRI and sequencing.
RNA Extraction and RT-PCR Analysis
Total RNA was obtained from different human tissues using
standard techniques (Trizol, Life Technologies, Inc.), and
3 µg were first treated with DNAse I (Life Technologies, Inc.) to eliminate possible contaminations and then reverse
transcribed using Superscript Reverse Transcriptase and oligo
dT16 primer (Life Technologies, Inc.) in a final volume of 20 µl, as recommended by the
manufacturer. Primers used for PCR amplification were S2136 and A2824
(23-mers). cDNA was subsequently amplified using Platinum
Taq DNA polymerase (one step at 94 C for 5 min followed by
30 cycles of 94 C for 45 sec, 58 C for 45 sec, and 72 C for 1 min, and
one step at 72 C for 7 min). Products were separated on 3% agarose gel
and visualized by ethidium bromide staining.
Real-Time Quantitative PCR
Real-time quantitative PCR analysis of hMR variants was carried
out on an ABI7700 Sequence Detector (Applied Biosystems, Foster City,
CA). Taqman probe and primers (Eurogentec, Seraing, Belgium)
were as follows:
upper primer: 5'-TTATGTGCTGGAAGAAATGATTGC-3'
lower primers: hMR 5'-AACTTCTTTGACTTTCGTGCTCCT-3'
hMR5,6 5'-CAGACTGATGCATCTTCTCTCTCCTA-3'
TaqMan probe: 5'-FAM CATTGATAAGATTCGACGAAAGAATTGTCT TAMRA-3'.
cDNA was generated as described above using 250 ng of random hexamers
(Promega Corp., Madison, WI). For each experiment,
one-tenth of the reverse transcription reaction was used for PCR
in the presence of 5 mM MgCl2, 200
µM deoxynucleoside triphosphates (dNTPs), and 1.25
U Taq polymerase. Final primers and probe concentrations
were 400 nM for each primer and 100
nM probe. PCR reagents were from Eurogentec
(Seraing, Belgium). Reaction parameters were 95 C for 10 min followed
by 40 cycles at 95 C, 15 sec and 55 C, 1 min. These conditions were
chosen after an assay optimization study, in which different
concentrations of MgCl2, primers, and probe were
tested. For preparation of standards, restriction fragments covering
exon 4 to 7 of hMR and hMR5,6 were purified from agarose gel, and
DNA concentration was accurately quantified by spectrofluorometry using
Hoechst 33258 trihydrochlorid [Sigma, St. Louis, MO
(58)]. Standard curves were generated using serial
dilutions of purified fragments spanning 5 orders of magnitude,
yielding correlation coefficients of at least 0.98 in all experiments.
Each standard and sample values were determined in triplicate in one to
three independent experiments. hMR5,6 expression within a given
tissue was calculated relative to wild-type hMR, and results were
represented as relative hMR5,6/hMR expression compared with the
heart, which was chosen as the calibrator.
In Vitro Transcription and Translation
In vitro transcription and translation were
accomplished using the TNT Quick Coupled Transcription/Translation
system (Promega Corp., Madison, WI) according to the
manufacturer’s protocol. Recombinant pCMV6-hMR5,6 containing
hMR5,6 or recombinant pcDNA3-hMR containing a 3 kb hMR
XmaIII-AflII fragment inserted into pcDNA3
(Invitrogen, San Diego, CA) (59) was used as
a template for transcription with T7 polymerase followed by translation
with [35S]-methionine (1,000 Ci/mmol,
Amersham Pharmacia Biotech, Les Ulis, France). Radioactive
products were analyzed on a 10% SDS-polyacrylamide gel. Cold
methionine was used for translation of proteins used in EMSAs.
Cell Culture and Transfection Procedures
Rabbit RCSV3 cells derived from kidney cortical collecting duct
(60) were kindly provided by Dr. P. Ronco (Hôpital
Tenon, Paris, France). Cells were grown in a defined medium composed of
DMEM-Ham’s F12 supplemented with 5 µg/ml insulin, 5 µg/ml
transferrin, 2 mM glutamine, 100 IU/ml penicillin
and 100 µg/ml streptomycin, 20 mM HEPES, 50
nM sodium selenate, 50 nM dexamethasone, and
2% charcoal-stripped FCS. The cells were seeded in six-well plates at
a density of 5 x 105 cells per well at
least 6 h before transfection in fresh medium without any added
steroid. For all transfection experiments, RCSV3 cells were used
between passages 30 and 40.
Cells were cotransfected by the calcium phosphate method
(61) with plasmid pCMV6-hMR5,6 coding for hMR5,6 and
1.25 µg of an MMTV-luciferase reporter construct (pF31luc, gift of
Dr. H. Richard-Foy, Laboratoire de Biologie Moléculaire
Eucaryote, CNRS, Toulouse, France), with or without expression plasmids
coding for the human MR (pcDNA3-hMR) and GR (pRShGR) (gift of Dr. R.
Evans, Howard Hughes Medical Institute, La Jolla, CA).
Cotransfection of 0.5 µg pSVßgal (CLONTECH Laboratories, Inc.), a plasmid encoding for ß-galactosidase, was performed
to normalize for transfection efficiencies. The day after transfection,
cells were rinsed with PBS and steroids were added for 24 h. The
cells were rinsed twice with cold PBS and lysed in 25 mM
glycyl-glycine, pH 7.8, 1 mM EDTA, 1 mM
dithiothreitol, 8 mM MgSO4, 1%
Triton X100, 15% glycerol. Cellular extracts were assayed for
luciferase (62) and ß-galactosidase (63)
activities. Results were standardized for transfection efficiency and
expressed as the ratio of luciferase activity over ß-galactosidase
activity in arbitrary units.
For coactivator assays, 0.5 µg of hMR5,6 was cotransfected in the
same conditions as described above, together with 0.5 µg of
expression vectors for steroid receptor coactivators 1a (SRC1a) and 1e
(SRC1e), as well as RIP140 and hTIF1 (plasmids were kindly provided
by Drs. V. Cavaillès, INSERM 4148, Montpellier, France, and
M. G. Parker, Imperial Cancer Research Fund, London,
UK).
Aldosterone and dexamethasone were purchased from Sigma.
EMSA
For EMSA, WCE were prepared from RCSV3 cells previously
transfected with recombinant pCMV6-hMR5,6 by the lipofectamine
method (Life Technologies, Inc. Inc., Paisley, UK) and
from Sf9 cells infected by recombinant baculovirus AcNPV-hMR
containing the full-length hMR cDNA (64). Forty eight
hours after transfection, cells were washed with cold PBS and
homogenized in 20 mM HEPES, pH 7.9, 1,5 mM
MgCl2, 0,6 M NaCl, 0.2 mM
EDTA, 0.5 mM phenylmethylsulfonyl fluoride, 25% glycerol
and 0.1% protease, and phosphatase inhibitor cocktail
(Sigma) by 20 strokes in a glass-glass Potter apparatus at
4 C. The homogenates were centrifuged at 15,000 rpm for 30 min at 4 C,
and the supernatant was used as WCE. One microgram of recombinant
pcDNA3-hMR, 1 µg recombinant pCMV6-hMR5,6, or a combination of
them (0.5 µg each) were in vitro translated in the
presence of cold methionine, as described above. Gel mobility shift
assays were performed essentially as described in Ref. 3 .
Purified oligonucleotides were annealed and labeled with
[32P]dCTP (Amersham Pharmacia Biotech) using the Klenow fragment of DNA polymerase (Life Technologies, Inc., Paisley, UK) to a specific activity
of approximately 108 cpm/µg of DNA. Unlabeled
oligonucleotides were used as competitors. Oligonucleotides used in the
gel mobility shift experiments are as follows:
GREcon: 5'-AGCTGCTCAGCTAGAACACTCTGTTCTCTACT-3'
and 5'-AGCTAGTAGAGAACAGAGTGTTCTAGCTAGC-3'.
Protein-DNA complexes were separated from free DNA by electrophoresis on nondenaturing 4.5% polyacrylamide gel in 0.25 x Tris-borate-EDTA buffer at 200 V for 1 h. Gels were dried and exposed to x-ray film at -80 C.
GST Pull-Down Assays
Plasmids containing the GST (pGEX2TK), GST fused to RIP140
(GST-RIP140), and GST fused to hTIF1 amino acid sequence 630–854
(GST-hTIF1) were kindly provided by Dr. V. Cavaillès. GST-SRC1
encoding a fusion protein of GST with residues 570–780 of hSRC1
(common to both hSRC1a and hSRC1e isoforms) was provided by Dr. M.
G. Parker. GST and GST fusion proteins were expressed, and proteins
were prepared as previously described (31). An aliquot of
crude bacterial extract containing GST fusion proteins was incubated
for 30 min at 4 C with 100 µl glutathione-Sepharose beads, previously
washed three times in 1:1 (vol/vol) NETN (0.5% Nonidet P-40, 1
mM EDTA, 20 mM Tris-HCl, pH 8.0, 100
mM NaCl). Glutathione-Sepharose beads were then washed
three times with NETN. hMR5,6 was transcribed and translated
in vitro in the presence of
35S-methionine and incubated with the fusion
proteins on glutathione-Sepharose beads for 1 h at 4 C. The beads
were washed, suspended in 20 µl loading buffer, boiled for 3 min, and
analyzed by SDS-PAGE. Signals were amplified with Entensify, and gels
were autoradiographed at -80 C overnight.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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This work was supported in part by a grant of the European Section of Aldosterone Council (ESAC).
Abbreviations: DBD, DNA-binding domain; GST,
glutathione-S-transferase; GRE, glucocorticoid response
element; 11HSD2, 11ß-hydroxysteroid dehydrogenase type II; LBD,
ligand-binding domain; PPARtr, truncated form of PPAR; RIP140,
receptor interacting protein 140; SRC, steroid receptor coactivator;
TIF1, transcriptional intermediary factor 1; WCE, whole-cell
extract.
Received for publication September 6, 2000. Accepted for publication May 23, 2001.
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