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Original Studies |
Mothers and Babies Research Center, Endocrine Unit, John Hunter Hospital, Newcastle, New South Wales 2310, Australia
Address all correspondence and requests for reprints to: Dr. Roger Smith, Mothers and Babies Research Center, Endocrine Unit, John Hunter Hospital, Newcastle, New South Wales 2310, Australia. E-mail: mdrsm{at}mail.newcastle.edu.au
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Abstract |
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Introduction |
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Glucocorticoids, a major subclass of steroid hormones, exert profound effects on cell growth, development, differentiation, and homeostasis through their receptors by modulating the expression of many genes (17, 18, 19). Glucocorticoid receptors (GRs), with bound ligand, act as hormone-dependent transcription factors that recognize a specific glucocorticoid response element (GRE) located either upstream or downstream from the transcription initiation site of target genes, resulting in positive or negative effects on transcription (17, 18, 20, 21). However, glucocorticoid modulation of target gene expression is far more complex than was apparent at the time the genes for the GRs were isolated (22), especially if the targeted gene does not contain a consensus GRE, as is the case for the CRH gene. GRs can also interact with components of the transcription initiation complex (22, 23, 24) and cross-talk with other signaling pathways (22, 25, 26, 27, 28). Indeed, it has recently been suggested that all the important physiological functions of GR may be reliant on protein-protein interactions (29).
Recently, a number of studies on the transcriptional regulation of CRH gene expression by glucocorticoids have been carried out in the transfected mouse corticotroph tumor cell line, AtT-20 (30, 31, 32, 33). Adler and colleagues demonstrated that the synthetic glucocorticoid dexamethasone decreased basal CRH messenger ribonucleic acid (mRNA) level by 40–50% and repressed forskolin-stimulated CRH messenger RNA by 70% in stably transfected AtT-20 cells (31). Both Van’s (30) and Guardiola-Diaz’s (32) groups demonstrated that dexamethasone reduced cAMP stimulation of hCRH promoter activity by more than 50% in transiently transfected AtT-20 cells. More recently, Malkoski and colleagues (33) localized a DNA sequence capable of binding the GR in vitro, which is responsible for dexamethasone-dependent repression of cAMP-stimulated CRH promoter activity in AtT-20 cells. Taken together, these studies show that glucocorticoids inhibit CRH gene or promoter activities in transfected AtT-20 cells despite the absence of a consensus GRE.
In the placenta, in contrast to AtT-20 cells, Robinson and colleagues reported that glucocorticoids can stimulate CRH gene expression at protein and messenger RNA levels in human primary trophoblastic cells (13), but, to date, how the CRH gene is regulated by glucocorticoids in placental cells remains unclear.
In this report, primary cultures of human cytotrophoblasts have been used for the first time as a model to characterize the molecular mechanisms involved in glucocorticoid stimulation of human CRH (hCRH) gene expression. We show that in the presence of dexamethasone, transcription of the hCRH gene is increased via a process that is at least in part a primary transcriptional response, occurs at physiological concentrations, and is directed through a cAMP response element (CRE) located in the gene’s promoter region.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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All constructs used luciferase as the reporter for assaying transcriptional activation of the promoters under study. The orientation and sequence of all constructs were confirmed by DNA sequencing. The human CRH genomic clone, CRH1001+, was the gift of Joseph Majzoub, Harvard University Medical School (Boston, MA) (31). The CRH 5'-flanking DNA was subcloned into the promoterless Photinus (firefly) luciferase reporter vector pGL3-Basic (Promega Corp., Madison, WI). To make pCRH(5500)-GL3, the 5.5-kb upstream region was isolated by XbaI and HaeII double digestion and filled to blunt ends with Klenow enzyme (Promega Corp.), then ligated into the SmaI site of pGL3-Basic vector with T4 DNA ligase (Promega Corp.). To create pCRH(4300)-GL3, the 4300-bp fragment was isolated from the pCRH(5500)-GL3 construct with SacI and NheI double digestion, then ligated into the SacI and NheI sites of pGL3-Basic vector in a forward orientation. To construct pCRH(663)-GL3, a 790-bp promoter region (including 663 bp of hCRH promoter region and 127 bp of first exon region) was isolated by a PstI digest, then subcloned into pGL3-Basic vector in the same manner.
Additional deletions of the CRH promoter were made by stepwise removal of 5'-flanking DNA with exonuclease III and S1 nuclease. Briefly, hCRH(663 bp)pGL3-Basic vector was double digested with KpnI and NheI, which generates a 5'-overhang at the NheI site in the vector 5' to the hCRH promoter sequences that is suitable for exonuclease III digestion and a 3'-overhang at the KpnI site, located 15 bp 5' of the NheI site, that is resistant to exonuclease III digestion. After treatment with exonuclease III, S1 nuclease was added to remove the single strand DNA overhangs, and the different length promoter-vector DNAs were recircularized with ligase.
To create a plasmid with the ß-globin promoter driving the luciferase reporter the rabbit ß-globin promoter sequence (-109 to +10 bp) was removed from pGLOB-CAT (34) by BamHI and BglII double digestion, and ligated into the BglII site of the pGL3-Basic vector to make the GLOB-pGL3 vector. To construct the CRE-globin promoter plasmid, the BglII site of a luciferase construct containing the hCRH promoter region from -340 to -215 bp (which had been constructed by linking a hCRH PCR fragment into the BglII and [I]MluI sites of the pGL3-promoter vector) was converted to a XhoI site using a linker oligonucleotide. MluI and XhoI double digestion removed the hCRH promoter sequences from this plasmid, and the fragment was ligated into MluI- and XhoI-digested pGLOB-GL3 to create pCRE-GLOB-GL3.
Mutagenesis
Oligonucleotide-directed mutagenesis of the CRE, hybrid steroid response element (HRE), and ecdysone response element (EcRE) were carried out in the pCRH(663)-GL3 construct using the Quik Change Site-Directed Mutagenesis Kit (Stratagene, La Jolla, CA). Mutated base pairs, confirmed by DNA sequencing, are at positions -229, -226, and -225 bp in mtCRE; at -272, -269, and -267 bp in mtEcRE; and at -215 and -213 bp in mtHRE. The oligonucleotides used are as follows with the mutated nucleotides underlined (the complementary strand sequence is not shown): mtCRE, 5'-ccttccattttagggctcgctgcagtcaccaagaggcg-3'; mtEcCR, 5'-ctcattcaagaatttttctcgagggacaagtcataagaagcccttc-3'; and mtHRE, 5'-ggcctttcatagtaagaggcctatatgttttcacacttggg-3'.
Placental cell isolation and cultures
Human term placentas were obtained from normal pregnant women after spontaneous vaginal delivery or elective cesarian. Collection of placentas was performed with the approval of the Hunter Area Health Service, New South Wales, Australia, and the University of Newcastle human research ethics committees. Cytotrophoblasts were obtained according to Kliman’s method (35). Briefly, chorionic villi tissue obtained from the maternal side of the placenta was dispersed with trypsin and deoxyribonuclease I, a highly purified fraction of cytotrophoblasts was obtained by repeated Percoll gradient centrifugations, and cells were maintained in DMEM (Life Technologies, Inc., Gaithersburg, MD). The purity of the cytotrophoblasts was determined by immunohistochemical staining with markers specific to syncytiotrophoblast (CRH), epithelium (cytokeratin), endothelium, and fibroblasts (vimentin). All experiments were carried out in preparations of cytotrophoblasts with purity greater than 95%.
Transfection
Standard transfection methods were as follows: freshly isolated cytotrophoblasts were plated in six-well plates (Falcon, Becton Dickinson, Bedford, MA) at 2.5 x 106 cells/well. Cells were incubated with plasmid-liposome complexes comprised of 20 µg DNA and 0.5 µg control DNA (pRL-TK vector, Promega Corp.) with 20 µg freshly prepared liposomes (1 mg/mL each of dioleyphosphatidylethanolamine and dimethyldioctadecylammonium bromide) (36, 37) in a humidified atmosphere of 5% CO2 at 37 C. Four hours later, cells were fed with 10% charcoal-stripped FBS with or without dexamethasone. Transfections without hormone received the same volume of ethanol vehicle. A luciferase assay was carried out 48 h thereafter with the dual luciferase assay kit (Promega Corp.).
A 1% transfection efficiency of primary placental cells was routinely obtained as determined by in situ detection of ß-galactosidase activity after transfection with the pSV-ß-galactosidase control reporter vector (Promega Corp.).
AtT-20 cells (D16–16) were obtained from Karen Sheppard (Baker Medical Research Institute, Melbourne, Australia) and maintained in DMEM with 10% charcoal-stripped FBS in a humidified atmosphere of 5% CO2 at 37 C. For transient transfection, cells were plated on six-well plates at a density of 5 x 105/well. Transfections were carried out at 40–60% confluence 24 h after plating. Cells were transfected as described above for 3 h.
Nuclear run-on in vitro transcription assay
Nuclei were isolated, and the nuclear run-on in vitro
transcription was performed essentially as described previously (34, 38, 39). Primary placental cells (3 x 107)
were collected by scraping into phosphate-buffered saline, followed by
gentle resuspension in cell lysis buffer [10 mmol/L Tris-HC (pH 7.5),
10 mmol/L NaCl, and 2 mmol/L MgCl2] and lysis in
0.5% Nonidet P-40. The crude nuclear pellet was formed by
centrifugation at 250 x g for 5 min, then washed with
lysis buffer. The final nuclear pellet was gently resuspended and
stored in 50 mmol/L Tris-Cl (pH 7.5), 5 mmol/L
MgCl2, 0.1 mmol/L ethylenediamine tetraacetate
(EDTA), and 40% glycerol. The fresh placental cell nuclei were
suspended in 25 mmol/L Tris-HCl (pH 7.5), 2.5 mmol/L
MgCl2, 0.05 mmol/L EDTA, and 20% glycerol. Forty
microliters of nuclei were incubated in a 100-µL reaction mixture
containing 125 mmol/L Tris-Cl (pH 7.5); 50 mmol/L NaCl; 350 mmol/L
(NH4)2SO4;
5 mmol/L MgCl2; 0.2 mmol/L EDTA; 1 mg/mL heparin;
0.5 mmol/L concentrations of ATP, GTP, and UTP; and 150 µCi
[
-32P]CTP for 45 min at 32 C, then 100 µg
transfer RNA were added to the mixture. The reaction was treated with
deoxyribonuclease (125 µg/mL) and proteinase K (125 µg/mL) for 30
min at 37 C. After mixing with 50 µL 200 mmol/L EDTA and 50 µL 10%
SDS, the mixture was extracted with phenol/chloroform. The aqueous
phase was precipitated in 10% trichloroacetic acid, then washed three
times with 5% trichloroacetic acid. The pellet was resuspended in 25
mmol/L Tris-C1 (pH 7.5), 1 mmol/L EDTA, then ethanol-precipitated. The
pellet was resuspended in distilled water, and aliquots were used
to assay specific RNA level by hybridization to filter-bound DNA.
Linear plasmid carrying the human CRH genomic clone, CRH1001+ (31), or control DNA (PUC18 plasmid) dissolved in 0.1 mol/L Tris, pH 7.5, was boiled for 10 min and cooled on ice, 1 vol 20 x SSC (standard saline citrate) was added and loaded onto the slot blot apparatus. The nitrocellulose was dried at 80 C in a gel dryer (model 583, Bio-Rad Laboratories, Inc., Richmond, CA) for 2 h. After prehybridization overnight, nitrocellulose filters were hybridized for 48 h in 50% formamide, 3.3 x SSC, 20 mmol/L sodium phosphate, 0.1% SDS, 1 x Denhardt’s solution, 20 µg/mL transfer RNA, 40 µg/mL heparin, and 107 cpm in vitro synthesized RNA. The same quantity of radioactivity was used for all hybridizations, carried out in parallel. After hybridization, the nitrocellulose filters were washed four times in 4 x SSC and 0.1% SDS at 65 C, treated with ribonuclease A (10 µg/mL), and washed several times again. After air-drying, the nitrocellulose filters were exposed to Kodak Biomax MS film (Eastman Kodak Co., Rochester, NY) with an intensifying screen for 24–72 h.
Electrophoretic mobility shift assay (EMSA)
EMSAs were performed using as a probe 5-32P-labeled, blunt ended, double stranded oligonucleotides (wtCRE) identical to the CRE motif of the hCRH promoter region from -232 to -217 bp. The probe (10,000 cpm) was incubated for 1 h at 4 C in 15 µL 10% glycerol-10 mmol/L Tris-HCl (pH 7.8), 10 mmol/L KCl, and 1 mmol/L dithiothreitol containing 10 µg nuclear protein extract after preincubation for 30 min with 2 µg poly(dI-dC) and 0.5 µg BSA. Competition was performed with 50 ng unlabeled wtCRE oligonucleotides or oligonucleotides identical to the CRE region of the CRH promoter from -248 to -211 bp (comCRE) or containing mutations at positions -229, -226, and -225 bp (mtCRE), which were added simultaneously with the labeled probe. The samples were loaded on a 4% polyacrylamide gel in 0.25% Tris-glycine and 1 mmol/L EDTA (pH 8.3). The gel was run for 2 h at 4 C.
The DNA sequences of the oligonucleotides used in EMSA are as follows (the complementary strand sequence is not shown): wtCRE, 5'-TCGTTGACGTCACCAA-3'; comCRE, 5'-CCTTCCATTTTAGGGC-TCGTTGACGTCACCAAGAGGCG-3'; and mtCRE 5'-CCTTCCATT- TTAGGGCTCGCTGCAGTCACCAAGAGGCG-3'.
CRH RIA
CRH immunoreactivity in the culture medium was extracted using activated Vycor (Corning, NY) glass. Frozen culture medium samples (1 mL) were thawed at room temperature and adsorbed onto Vycor silica glass powder (200 mg glass powder/1 mL medium sample). The sample was then washed first with 3 mL deionized water and then with 2 mL 1 mol/L HCl (BDH Chemicals, Poole, UK) before the adsorbed material was eluted with 2 mL 60% acetone (BDH Chemicals). The eluate was transferred to polycarbonate tubes, dried, and stored at -20 C for RIA. CRH RIA was performed as previously described (14). Human CRH-(1–41) (Sigma, St. Louis, MO) was used as the standard, and radioligand was prepared with the chloramine-T method and purified by HPLC. The anti-CRH antibody Y2B0 was a gift from Phil Lowry (University of Reading, Reading, UK). The concentration of CRH IR was expressed as picograms per 2.5 x 106 cells/24 h.
Statistical analyses
Induction was defined as the fold increase over a baseline level of 1.0 or 100%. The values are expressed as the mean ± SEM. P < 0.05 was considered statistically significant. Statistical analysis for dexamethasone-dependent induction was determined by paired t test, and the difference in hormonal response for various plasmid/promoter constructs was assessed by unpaired t tests. Multiple comparisons were performed by one-way ANOVA or repeated measures ANOVA together with post-hoc pairwise comparisons using Excel 97 (Microsoft Corp., Redmond, WA).
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Results |
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To investigate whether the increase in CRH is a transcriptional
response to dexamethasone and to determine the requirement for ongoing
protein synthesis in this glucocorticoid-dependent induction of CRH
gene expression, nuclear run-on transcription was performed with nuclei
from human primary placental cells in the presence or absence of
cycloheximide, which blocks peptide bond formation. As shown in Fig. 1
, A and B, dexamethasone stimulated
endogenous CRH gene expression 2-fold, and prior exposure to
cycloheximide did not affect the level of dexamethasone stimulation of
endogenous CRH gene expression. These results indicate that the
dexamethasone-mediated up-regulation of CRH expression occurs at the
transcriptional level in a manner that does not require synthesis of
new protein.
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To determine whether DNA sequences within the 5'-flanking region
of the CRH gene are involved in glucocorticoid-mediated transcriptional
regulation, primary cultures of human placental cells were transfected
with plasmids containing a luciferase reporter gene fused to 5500 bp
[pCRH(5500)-GL3] or 663 bp
[pCRH(663)-GL3] of hCRH 5'-flanking DNA
sequences. Dexamethasone (10-7 mol/L) increases
hCRH promoter activity 2-fold (Fig. 1C
), and this increase parallels
the elevation in endogenous CRH gene expression (Fig. 1D) as determined
by RIA for CRH in culture medium of transfected cells. This result is
consistent with the nuclear run-on result and shows that dexamethasone
induces transcription of the hCRH gene through DNA sequences in the
promoter region of the gene.
Time-course studies show that hCRH promoter activity is increased by
12-h exposure to dexamethasone, with maximal stimulation occurring by
36 h (see Fig. 2A). Dose-response
studies indicate that a maximal stimulation of 2-fold over basal
levels occurs at 10-8 mol/L dexamethasone (see
Fig. 2B).
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To localize the regulatory elements required for dexamethasone
stimulation of hCRH gene expression, we transfected luciferase reporter
plasmids containing progressively shorter sections of the hCRH gene
into human primary placental cells (Fig. 3). This analysis of these 5'-deletions
of the hCRH promoter sequences indicates that dexamethasone
responsiveness is lost when the DNA sequences between -342 to -213 bp
are removed. This indicates that this region of the promoter is
required for dexamethasone-mediated up- regulation of hCRH gene
expression in human placental cells.
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To determine whether dexamethasone modulates cAMP-stimulated CRH
promoter activity, we examined the coeffects of dexamethasone and
8-bromo-cAMP on expression of different hCRH gene promoter constructs
in human primary placental cells. Treatment with dexamethasone,
8-bromo-cAMP, and both agents increases hCRH promoter activity 2-, 5-,
and 8-fold, respectively, using the longer hCRH gene promoter
constructs in these cells (see Fig. 4).
However, deletion of the -342 to -213 bp promoter region not only
destroys cAMP inducibility of the hCRH promoter, but also abolishes
dexamethasone responsiveness.
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A search of the hCRH DNA sequence against the TRANSFAC database (40) revealed that the region between -342 and -213 bp contains a consensus CRE, a hybrid steroid HRE (41), and a sequence with identity to the insect steroid hormone (42), EcRE.
To further characterize this promoter region as a critical regulatory
element for dexamethasone up-regulation of hCRH gene expression, we
made hCRH promoter-luciferase constructs in which the CRE, HRE, and
EcRE consensus sequences were specifically mutated. Dexamethasone
stimulates expression from the wild-type promoter (see Fig. 5A), whereas mutation of the CRE not only
decreases basal hCRH promoter activity by 30% (P <
0.05), but abolishes dexamethasone responsiveness as well. This
indicates that dexamethasone stimulation of hCRH gene expression
requires a functional cAMP regulatory element in the placental cells.
Mutation of the HRE has no effect on basal expression or dexamethasone
induction, but mutation of the EcRE results in a 2-fold increase in
basal expression and maintains dexamethasone induction.
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To test whether the hCRH CRE region can transfer dexamethasone
up-regulation to a heterologous promoter, we created a plasmid with the
hCRH CRE region inserted 5' of the rabbit ß-globin promoter, linked
to the luciferase reporter. The effects of dexamethasone on this
chimeric construct were studied in transiently transfected human
primary placental cells. Dexamethasone has no effect on the native
rabbit ß-globin promoter driving the luciferase reporter (see Fig. 5B), but significantly induces the expression of the chimeric construct
containing the hCRH CRE linked to the ß-globin promoter. Furthermore,
the CRE also increases basal rabbit ß-globin promoter activity. This
study shows that the hCRH CRE can transfer dexamethasone up-regulation
to a heterologous promoter in placental cells, indicating that the hCRH
CRE region alone is sufficient to confer dexamethasone-mediated
transcriptional induction in the placental cell environment.
A placental nuclear protein binds specifically with CRE in vitro
To show that the regulation of the hCRH gene in primary placental
cells involves specific interaction of transcription factor protein
binding to the CRE we used EMSA to detect nuclear protein-DNA
complexes. Figure 6 shows that a specific
DNA-protein complex (Fig. 6, at position of arrow) is
detected using 32P-labeled wild-type CRE (wtCRE)
oligonucleotide probe (lane 2). This complex can be specifically
competed away with cold wtCRE oligonucleotides or another
oligonucleotide pair containing the hCRH CRE plus more of the hCRH
flanking sequences (comCRE; lanes 6 and 7), but not with
oligonucleotides containing a mutated CRE (mutCRE; lane 8). No
difference in complex formation was observed under these conditions
with nuclei isolated from cells exposed to dexamethasone, cAMP, both
agents, or no treatment (compare lanes 2–5 and 9).
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To test whether CRE-mediated dexamethasone up-regulation of CRH
expression is unique to the placenta and distinct from mechanisms
controlling CRH gene expression in AtT-20 cells, we compared expression
of the hCRH (663 bp)-luciferase gene construct in placental cells and
AtT-20 cells. Dexamethasone stimulates CRH gene expression in
transfected primary cultures of human placental cells (Fig. 7A), whereas it blocks cAMP stimulation
of hCRH gene expression in AtT-20 cells (Fig. 7B), suggesting that the
differential regulation of CRH gene in placenta and hypothalamus is
most likely due to tissue-specific transcriptional factor differences
rather than to the CRH gene itself.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The hypothalamic hormone CRH is also expressed at many other sites in the central nervous system (8, 9) and in peripheral tissues, including the placenta (10, 11, 12). Glucocorticoids can exert either inhibitory or stimulatory effects on CRH mRNA levels depending on the cell model examined. In AtT-20 cells, glucocorticoids inhibit CRH mRNA accumulation, whether measured using stable or transient transfection with CRH gene or promoter constructs (31, 43, 44), whereas glucocorticoids stimulate CRH mRNA accumulation in human primary trophoblast cultures (12). Guardiola-Diaz and colleagues indicate that glucocorticoid repression of cAMP-activated CRH promoter activity is modulated via glucocorticoid receptor interference with CRE-binding protein, and glucocorticoid receptor binding to putative GR-binding sites is not required (32). In contrast, Malkoski and colleagues suggest that glucocorticoids repress cAMP-stimulated, but not basal, CRH promoter activity through direct glucocorticoid receptor interaction with DNA, and a functional CRE is not required (33). Exactly how glucocorticoids modulate CRH gene activity is still unclear, and the different results reported may reflect differences in the cell lines examined. It should be noted that AtT-20 cells are a transformed mouse corticotroph cell line that does not express endogenous CRH, although it can accurately express and secrete CRH after transfection with CRH gene constructs (31). Thus, interpretations of results from transcriptional analyses with cells exhibiting a partially differentiated phenotype must be made with caution, and extrapolation to in vivo events may be difficult.
It is well established that human villous trophoblasts in vitro differentiate from cytotrophoblastic cells into syncytiotrophoblasts (35), which express both endogenous CRH (12, 13) and glucocorticoid receptor (45). Thus, human trophoblast primary cultures provide a good model to characterize endocrine factors involved in transcriptional regulation of hCRH gene expression within the placenta. However, it is notoriously difficult to transfect such primary placental cells. Recently, Golos and colleagues reported that freshly isolated primate placental cytotrophoblasts can take and express exogenous genes during fusion to syncytiotrophoblasts (36). Our results are consistent with those of Golos et al. (36) in demonstrating transient transfection of human primary placental cells with CRH promoter constructs. Furthermore, we show that dexamethasone increases both basal and cAMP-mediated hCRH promoter activity in transfected human primary placental cells, and this correlates well with the measured response of CRH peptide production. Dexamethasone stimulates transfected hCRH promoter activity in a dose-dependent manner, with the effective dose for dexamethasone ranging from 10-9–10-8 mol/L. We conclude that primary cultures of human placental cells provide a good model to characterize transcriptional regulation of the hCRH gene by glucocorticoids in the human placenta.
Genes regulated by glucocorticoids have been classified into two groups, primary and secondary response genes (46). According to this classification, the primary genes are defined as those that respond relatively rapidly to glucocorticoid and do not require ongoing protein synthesis, whereas secondary response genes are those whose induction is delayed from hours to days and are dependent upon new protein synthesis. We have measured transcription by in vitro extension of in vivo initiated RNA (nuclear run-on) to demonstrate that dexamethasone stimulation of hCRH gene expression in primary human placental cells does not require ongoing protein synthesis. In time-course studies, hCRH promoter activity increased in transfected primary placental cells after 12-h exposure to dexamethasone, whereas maximal stimulation occurred by 36 h. These results are consistent with those of Guardiola-Diaz et al. in AtT-20 cells (32) and Rosen et al. (44) in NPLC cells. Taken together, these data show that the hCRH gene belongs to the primary glucocorticoid response class, implying that glucocorticoid-mediated effects on hCRH gene transcription are modulated by direct or indirect interaction of GR with the hCRH promoter region in a manner not requiring the synthesis of new protein.
Sequence analysis of the proximal hCRH promoter region did not reveal
the presence of consensus palindromic GREs (47), even though
glucocorticoids stimulate placental CRH secretion (13, 15) and mRNA
expression (15). Work with AtT-20 cells indicates that glucocorticoids
repress cAMP-stimulated, but not basal, hCRH promoter activity (30, 32, 33). Our data suggest that dexamethasone stimulates placental hCRH gene
expression through its interaction with the cAMP signaling pathway.
Indeed, cross-talk between the steroid receptor signaling pathway and
membrane receptor signals such as activator protein-1, signal
transducer and activator of transcription-5, and nuclear factor-
B
has been established (22, 27, 28, 48, 49, 50). This type of regulation does
not depend on the presence of a GR-binding site in the promoter and
potentially explains glucocorticoid-mediated transcription of certain
genes (22, 27, 50), such as hCRH, whose promoters lack GREs or negative
GREs (47, 51). Recently, Stauber and colleagues demonstrated that a
mutual cross-interference between glucocorticoid receptor and
CRE-binding protein was important for transcription regulation of the
glycoprotein hormone -subunit gene in human placental cells (26). We
now show that deletion and site-directed mutagenesis of the CRE in the
hCRH promoter abolishes dexamethasone responsiveness, whereas a hCRH
promoter fragment containing a functional CRE confers glucocorticoid
responsiveness to a heterologous promoter. These results clearly
indicate that a functional CRE is necessary and adequate for
glucocorticoid-mediated stimulation of hCRH gene expression in human
primary placental cells. This is consistent with the findings of
Guardiola-Diaz and colleagues for glucocorticoid-mediated repression of
CRH expression in the AtT-20 cell line (32). Gel shift analysis shows
that a nuclear protein from primary placental cells specifically binds
to the CRE. Taken together, these data imply that up-regulation of hCRH
gene expression in response to glucocorticoids in human placental cells
occurs through an interaction with the cAMP signaling pathway.
Finally, we compared the transfection results from human primary placental cells with those from AtT-20 cells using chimeric hCRH(663 bp)luciferase constructs. Our observations from AtT-20 cells that dexamethasone decreased cAMP-mediated, but not basal, hCRH promoter activity by nearly 50% are consistent with the findings of other groups (30, 31, 32, 33). Dexamethasone exerts an opposite effect on CRH gene expression in placental compared with AtT-20 cells, suggesting that there are unique mechanisms within placental cells by which glucocorticoids modulate hCRH gene expression. CRH is a single copy gene that is highly conserved; thus, the differential regulation of the hCRH gene in placental and AtT-20 cells is probably due to tissue-specific differences in trans-acting factors.
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Acknowledgments |
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Footnotes |
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2 Present address: Division of Endocrinology, Children’s Hospital
Medical Center, 3333 Burnet Avenue, Cincinnati, Ohio 45229-3039.
Received October 11, 1999.
Revised January 4, 2000.
Accepted January 4, 2000.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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B: what language. TIBS. 23:233–235.
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