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Am J Physiol Endocrinol Metab 292: E1790-E1800, 2007. First published February 20, 2007; doi:10.1152/ajpendo.00708.2006
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Aldosterone-induced EGFR expression: interaction between the human mineralocorticoid receptor and the human EGFR promoter

Claudia Grossmann,1 Alexander W. Krug,2 Ruth Freudinger,1 Sigrid Mildenberger,1 Katharina Voelker,1 and Michael Gekle1

1Physiologisches Institut der Universitaet Wuerzburg, Wuerzburg; and 2Universitaetsklinikum Carl Gustav Carus Universitaet Dresden, Medical Clinic III, Dresden, Germany

Submitted 22 December 2006 ; accepted in final form 15 February 2007


   ABSTRACT
TOP
 ABSTRACT
EXPERIMENTAL PROCEDURES
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Aldosterone plays a key role in cardiovascular and renal injury. The underlying mechanisms are not completely understood. Because the epidermal growth factor receptor (EGFR) is involved in the development of fibrosis and vascular dysfunction, upregulation of EGFR expression by aldosterone-bound mineralocorticoid receptor (MR) is an attractive hypothesis. We investigated the effect of aldosterone on EGFR expression in the aorta of adrenalectomized rats and in human aorta smooth muscle cells (HAoSMC) in primary culture. Aldosterone, but not dexamethasone, stimulated EGFR expression in vivo in the aorta as well as in HAoSMC. EGFR degradation was not affected. Aldosterone-induced EGFR expression in HAoSMC was dose dependent and prevented by spironolactone. Furthermore, incubation of HAoSMC with aldosterone led to enhanced EGF-induced ERK1/2 phosphorylation and an EGFR-dependent increase in media fibronectin. EGFR promoter reporter gene assay as well as chromatin immunoprecipitation data indicate that MR interacts with the EGFR promoter. With deletion constructs we gained evidence that this interaction takes place between the hMR and the EGFR promoter regions 316–163 (stronger activation site, EC50 ~1.0 nM) and 163–1 (weaker activation site, EC50 ~0.7 nM), which do not comprise canonical glucocorticoid response elements and are not activated by the human glucocorticoid receptor. The interactions require in part the NH2-terminal domains of MR. ELISA-based transcription factor DNA binding assay with in vitro synthesized hMR suggest direct binding to region 163–1. Our results indicate that aldosterone leads to enhanced EGFR expression via an interaction with the EGFR promoter, which is MR specific and could contribute to the aldosterone-induced increase in fibronectin abundance.

epidermal growth factor receptor; vascular smooth muscle cell

RECENTLY, IT HAS BECOME EVIDENT that the importance of aldosterone and the mineralocorticoid receptor (MR) extends beyond regulation of salt and water homeostasis. In animal models, aldosterone promotes cardiovascular fibrosis and hypertrophy due to tissue remodeling as well as endothelial dysfunction independently of its effects on blood pressure or electrolyte control (3, 45, 49). Several mechanisms have been proposed for the pathological actions of aldosterone, including enhanced collagen and fibronectin synthesis, formation of reactive oxygen species, decreased NO availability, and interaction with peptide hormone signaling (23). The pathophysiological significance of aldosterone for humans has received impressive support from clinical studies, especially RALES and EPHESUS, in which low doses of MR antagonists led to a dramatic improvement of mortality in patients with impaired left ventricular function (34, 35). Unfortunately, the underlying mechanisms of this important action of aldosterone and the MR are not completely understood.

Classically, aldosterone binds to the cytosolic MR, which then translocates to the cell nucleus and acts as a transcription factor (1). Additionally, steroids, including aldosterone, have the ability to interact with peptide hormone signaling (19). One important peptide signaling "system" concerning aldosterone is the epidermal growth factor (EGF) and its receptor (EGFR) (10, 19). Pharmacological in vivo studies have shown that mineralocorticoids enhance EGF-induced contraction of arteries (9) and that the MR antagonist spironolactone reduces the expression of EGFR mRNA after cerebral ischemia (6). Interestingly, EGFR expression supports fibrosis in cardiovascular and renal tissue (8, 16, 44).

The importance of EGFR in mediating pathophysiological effects of heterologous signaling systems is supported by its role in endothelin-induced fibrogenic effects (8). Endothelin-induced phosphorylation of the mitogen-activated protein kinase and endothelin-induced increase in collagen I gene activity were completely prevented by an inhibitor of the EGFR kinase. Angiotensin II (ANG II) also "uses" the EGFR to elicit certain effects in cardiovascular cells. For example, Bokemeyer et al. (4) reported that ANG II-induced growth of vascular smooth muscle cells requires activation of EGFR, and Mazak et al. (24) showed that ANG II- or aldosterone-induced stimulation of ERK1/2 in vascular smooth muscle cells depends on EGFR. In cardiomyocytes, type 1A angiotensin receptor promotes hypertrophy via transactivation of the EGFR (2, 17, 46). Thus, EGFR not only transduces the signal of its classical ligand EGF but also serves as a heterologous transducer for other pathological signaling pathways like endothelin-1, ANG II, and catecholamines, all of which have been linked to cardiovascular remodeling (13).

Previously, aldosterone has been shown to stimulate the expression of the EGFR in different cultured cell types, the kidneys, the heart, and, on the mRNA level, also the aorta (6, 9, 18, 27, 31); however, it is not clear whether aldosterone also enhances EGFR protein expression in cells from other tissues. Furthermore, the underlying mechanism(s) is not yet known. However, understanding the events leading to enhanced EGFR expression could help to explain aldosterone-induced cardiovascular inflammation and fibrosis.

In the present study, we first determined EGFR protein expression in rats receiving aldosterone via osmotic minipumps to test whether there is an in vivo effect. To determine whether aldosterone exerts a direct effect on vascular cells and whether human cells are also sensitive, we assayed aldosterone-induced EGFR expression in human aorta vascular smooth muscle cells (HAoSMC) in primary culture. As a preliminary marker for the potential pathophysiological relevance of EGFR expression, we determined fibronectin secretion from HAoSMC. For the further investigation of underlying mechanisms, EGFR promoter activity was tested (pER reporter gene assay) in human embryonic kidney (HEK) cells with and without cotransfected hMR. Subsequently, we established a chromatin immunoprecipitation (ChIP) assay for hMR, which allows the determination of hMR binding to segments of intact DNA despite unidentified response elements to show the interaction of hMR with the EGFR promoter. Generating several deletion constructs of the EGFR promoter, we identified two regions of interaction with hMR that are not sensitive to stimulation of the glucocorticoid receptor (GR). Finally, we developed an ELISA-based transcription factor DNA binding assay to determine direct binding of hMR to DNA. Our data demonstrate that EGFR is an aldosterone-inducible protein in vasculature and investigate the molecular mechanism of enhanced EGFR expression caused by aldosterone, exploring a possible new target for prevention of renal and cardiovascular remodeling.


   EXPERIMENTAL PROCEDURES
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animal study. Experiments were performed according to the legal rules of the Federal Republic of Germany essentially as described before (18). They were approved by the Regierung von Unterfranken, Wuerzburg, Germany. Male Munich-Wistar rats weighing between 200 and 250 g (Charles River, Sulzfeld, Germany) underwent adrenalectomy (ADX) under NarcorenR anesthesia using two lumbodorsal incisions. Subsequently, an osmotic minipump was implanted (model 2ML1; Alzet, Palo Alto, CA). Eighteen animals were subdivided into three groups: 1) vehicle (ADX), 2) aldosterone (ADX + Aldo; 36 µg/100 g body wt and day), and 3) dexamethasone (ADX + Dex; 36 µg/100 g body wt and day). The animals were maintained on a standard rat pellet diet and were offered 0.9% NaCl solution. Six days after ADX, the animals were anesthetized with Inactin (120 mg/kg body wt; Byk-Gulden, Konstanz, Germany), and the organs were removed and frozen in liquid nitrogen until further processing. Blood samples were used to determine Na+, K+, Cl–, and aldosterone (kindly performed by the Central Laboratory of the Medical School, University of Wuerzburg and the Department of Endocrinology, University of Duesseldorf, Germany).

Cell culture. HAoSMC were obtained from PromoCell (Heidelberg, Germany) and cultivated using the media and instructions of the manufacturer. HEK-293 cells were acquired from American Type Culture Collection (Rockville, MD) and cultivated as described before (12). HEK-293 cells, which do not express endogenous hMR (12), were transfected with Polyfect Reagent (Qiagen, Hilden, Germany), according to the manufacturer's instructions, in medium without supplements. Stable cell clones were attained by selection with G418 (600 mg/l) and dilution cloning. Twenty-four hours before all cell culture experiments, cells were made quiescent by incubation in medium without supplements.

EGFR and fibronectin ELISA. EGFR expression was determined by an EGFR sandwich ELISA (R&D Systems, Minneapolis, MN) according to the manufacturer's protocol. Tissues were homogenized with 20 strokes of a Janke&Kunkel (Staufen, Germany) homogenizer at 4°C in extraction buffer [10 mM Tris, pH 7.4, 100 mM NaCl, 1 mM EDTA, 1 mM EGTA, 1 mM NaF, 20 mM Na4P2O7, 2 mM Na3VO4, 1% Triton X-100, 10% glycerol, 0.1% SDS, 0.5% deoxycholate, and protease inhibitor cocktail (1:1,000); Calbiochem, Bad Soden, Germany]. HAoSMC were lysed in cell extraction buffer. Protein content was determined using the BCA reagent from Pierce (Rockford, IL). Fibronectin was measured according to Vesey et al. (48).

Quantification of ERK1/2 phosphorylation by ELISA. For the quantification of ERK1/2 phosphorylation, we performed an ELISA according to Versteeg et al. (47) with minor modifications described previously (19). After stimulation as indicated, the cells were fixed with 4% formaldehyde in PBS and permeabilized with 0.1% Triton X-100. Cells were blocked with 10% fetal calf serum in PBS-Triton for 1 h and incubated overnight with the primary antibody (rabbit anti-phospho-ERK1/2, 1:1,000, New England Biolabs). Subsequently, cells were incubated with secondary antibody (peroxidase-conjugated mouse anti-rabbit antibody, dilution 1:10,000) in PBS-Triton with 5% BSA for 1 h at room temperature. Finally, the cells were incubated with 50 µl of a solution containing 0.4 mg/ml o-phenylenediamine, 11.8 mg/ml Na2HPO4, 7.3 mg/ml citric acid, and 0.015% H2O2 for 15 min at room temperature in the dark. The resulting signal was detected at 490 nm with a multiwell multilabel counter (Victor2; Wallac, Turku, Finland). Protein content in the wells was determined with trypan blue (19).

EGFR Western blot. HAoSMC were lysed in RIPA buffer as described elsewhere (18). Tissues were homogenized with 20 strokes of a Janke&Kunkel homogenizer at 4°C in RIPA buffer. Western blots were performed as described before the use of EGFR antibody sc-03 (1:1,000, Santa Cruz Biotechnology, Santa Cruz, CA) (18).

EGFR promoter reporter gene assay. We studied the activation of the EGFR promoter with pERLuc, possessing a firefly luciferase under control of the EGFR promoter (generously provided by A. Johnson) (15). Luciferase activity was assessed with the Luciferase Assay System (Promega, Mannheim, Germany). beta-Galactosidase encoded by pcDNA3.1 Hislacz (Invitrogen, Karlsruhe, Germany) was cotransfected as an internal transfection control, and its activity was assessed with a beta-Gal Assay Kit (Invitrogen).

Glucocorticoid response element reporter gene assay. To measure the transactivation activity of the hMR and its deletion constructs, we used pGRE-SEAP from Clontech (Mountain View, CA), consisting of three glucocorticoid response elements (GRE) coupled to a secretory alkaline phosphatase. To determine the activity of the EGFR promoter fragment 316–163, we cloned this fragment into pSEAP-basic. For quantification of transactivation we utilized the AttoPhos System (Promega). As an internal transfection control we cotransfected pEGFP-C1 (Clontech) and quantified the fluorescence.

pER and hMR deletion plasmids. Different deletion mutants of the EGFR promoter were constructed by restriction and PCR cloning (Table 1). The constructs were inserted into either the pLuc reporter or the pSEAP reporter, containing the secretory alkaline phosphates. Truncated versions of the pEGFP-hMR, which lack either the NH2-terminal AB-domain (CDEF-hMR) or the AB-domain and the DNA-binding domain (DEF-hMR), were constructed by cutting pEGFP-hMR [a kind gift from N. Farman consisting of the hMR-tagged NH2-terminally with an enhanced green fluorescent protein (EGFP)] with BglII and HindIII and then inserting appropriate PCR fragments (Table 1) (32). To exclude the possibility that the EGFP tag of the hMR changes the characteristics of the receptor, we compared its GRE activation and nuclear translocation properties with that of the untagged hMR but could not find significant differences (12, 32).


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  		Table 1. Primers and restriction enzymes for deletion constructs

 
Immunoprecipitation and EGFP-hMR Western blot. Cells were incubated for 24 h and then washed, harvested, and lysed in RIPA buffer (21). Lysates were then centrifuged at 11,000 rpm at 4°C for 10 min, and the supernatant was incubated overnight with EGFP antibody (sc-8334, Santa Cruz Biotechnology) with end-over-end rotation and then with A/G plus agarose for another 24 h. After 10 min of centrifugation at 10,000 rpm at 4°C, the pellet was mixed with 40 µl of Laemmli buffer and separated with an 8% SDS-page-gel. For Western blot, cells were lysed in Laemmli buffer, homogenized with a 21-gauge needle, and separated by 8% SDS-PAGE. The MR was detected with an antibody directed against its EGFP tag (Jl-8, BD Clontech, 1:2,500). As a secondary antibody, peroxidase-coupled anti-mouse-IgG was used and detected with an ECL kit (Amersham, Arlington Heights, IL).

ChIP. HEK cells were transfected and stimulated for 24 h. Proteins and DNA were cross-linked by incubation in 1% formaldehyde for 10 min at 37°C. Cells were then harvested, lysed in SDS lysis buffer (1% SDS, 10 mM EDTA, 50 mM Tris·HCl, pH 8.1), and sonicated in an ultrasound bath with in-between cooling on ice. Cell lysates were then centrifuged for 10 min at 13,000 rpm at 4°C, and the supernatant was diluted 1:10 in dilution buffer (0.01% SDS, 1% Triton X-100, 1.2 mM EDTA, 16.7 mM Tris·HCl, pH 8.1, 167 mM NaCl). Each sample (50 µl) was saved as input control, whereas the rest was precleared with A/G plus agarose. After preclearing, lysates were incubated with EGFP antibody (sc-8334, Santa Cruz Biotechnology) overnight at 4°C. A/G plus agarose was used for immunoprecipitation followed by rigorous washing with low-salt buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris·HCl, pH 8.1, 150 mM NaCl), high-salt buffer (0.1% SDS, 1% Triton X-100, 2 mM EDTA, 20 mM Tris·HCl, pH 8.1, 500 mM NaCl), LiCl buffer (0.25 M LiCl, 1% NP-40, 1% deoxycholate, 1 mM EDTA, 10 mM Tris·HCl, pH 8.1), and TE buffer (10 mM Tris·HCl + 1 mM EDTA, pH 8.0). For elution, 2x 250 µl of buffer containing 1% SDS and 0.1 M NaHCO3 were utilized. Cross-linking of proteins to DNA was reversed with 20 µl of 5 M NaCl and incubated for 4 h at 65°C. Proteins were then digested with proteinase K, and DNA was recovered by phenol-chloroform extraction and ethanol precipitation. PCR was performed, with aggtgccagaacatttctcta (sense) and aacacgcagatgcagtcg (antisense) to detect pGRE-SEAP and ggaccctcggactttagagc (sense) and aggagcagaggaggaggaga (antisense) to detect the EGFR promoter.

ELISA-based transcription factor DNA binding assay. For this sandwich assay, biotinylated DNA probes were captured in streptavidin-coated wells and the transcriptions factors bound were detected by specific antibodies. This assay was performed with hMR (or LacZ as negative control) synthesized in vitro using the TNT T7 Quick Coupled Transcription/Translation system (Promega). For this purpose, hMR was cloned into the pcDNA3-HisC vector from Clontech, introducing an XPRESS-flag at the NH2-terminal end (the lacZ construct contains the same XPRESS-flag). Biotinylated fragments of the EGFR promoter (pER-163, pER163–316) were used as probes. A biotinylated canonical GRE-GRE-GRE probe (ggtacattttgttctagaacaaaatgtaccggtacattttgttct) was used as positive control. First, hMR and LacZ were incubated with aldosterone for 1 h at 4°C. Biotinylated probes were immobilized in streptavidin-coated wells. Wells without biotinylated probes were used as blanks. Subsequently, hMR (or LacZ or no protein) was transferred to streptavidin-coated wells with the biotinylated probes and incubated for 2 h at room temperature with an equal volume of blocking buffer (5% BSA in PBS-Tween 0.05%). After three washes and a 2-h incubation with anti-XPRESS (1:1,000, Invitrogen) and three further washes and a 1-h incubation with horseradish peroxidase-coupled secondary antibody, the color reaction was started by the addition of 0.5 mg/ml o-phenylenediamine, 11.8 mg/ml Na2HPO4·2H2O plus 7.3 mg/ml citric acid and 0.015% H2O2.

Sequencing. All PCR products were sequenced (MWG Biotech) to confirm sequence identity.

Materials. Aldosterone and dexamethasone were purchased from Sigma (Munich, Germany).

Statistics. The data are presented as mean values ± SE. Significance of difference was tested by ANOVA. Differences were considered significant if P < 0.05; n = number of animals or cell culture dishes. Cells from at least two different passages were used.


   RESULTS
TOP
 ABSTRACT
 EXPERIMENTAL PROCEDURES
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
Aldosterone stimulates EGFR expression in the aorta of ADX rats. First, we determined EGFR protein expression in ADX rats receiving aldosterone via osmotic minipumps to test whether there was an in vivo effect of aldosterone on EGFR protein expression. In ADX control rats, aldosterone plasma concentration was below the detection limit, confirming successful surgical procedure. Application of aldosterone, resulting in plasma aldosterone concentrations of ~4.9 nmol/l, led to enhanced Na+ and reduced K+ concentrations in serum and therefore to an altered Na+/K+ ratio (Table 2). Figure 1 shows that aldosterone, but not dexamethasone, enhanced EGFR expression significantly in the aorta, whereas adipose tissue and liver showed no change in EGFR expression. In ventricular homogenates, we also observed a significant increase in EGFR expression with aldosterone (vehicle: 26.6 ± 1.7 pg/mg; aldosterone: 45.1 ± 1.5 pg/mg; dexamethasone: 30.9 ± 2.3 pg/mg; n = 5).


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  		Table 2. Serum parameters of the adrenalectomized animals

 

Figure 1
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  		Fig. 1. Effect of in vivo aldosterone administration on epidermal growth factor receptor (EGFR) expression in different rat tissues measured by ELISA (A, C, D) or Western blot (B). Aldosterone enhanced EGFR expression in tissue from aorta (A and B) but not from liver or adipose tissue (C and D). ADX, adrenalectomized; DMSO, DMSO (vehicle) application via osmotic minipump; Aldo, aldosterone application via osmotic minipump; Dexa, dexamethasone application via osmotic minipump. *P < 0.05, n = 6 for each group.

 
Aldosterone stimulates EGFR expression and increases media fibronectin abundance EGFR dependently in HAoSMC. To determine whether aldosterone exerts a direct effect on vascular cells and whether human cells are also sensitive, we assayed aldosterone-induced EGFR expression in HAoSMC in primary culture. As shown in Fig. 2, A and B, 72 h of exposure to aldosterone stimulated EGFR expression in HAoSMC, which was prevented by spironolactone. Results after 48-h exposure were similar. EGFR expression was 119 ± 4% of control (n = 9, P < 0.05 vs. control) after 24 h. Figure 2C shows the concentration dependence of the stimulation of EGFR expression. Half-maximum stimulation was observed at ~0.3 nmol/l. To test whether enhanced EGFR expression is functional (i.e., EGFR expressed in the cell membrane, coupled to signaling cascades), EGF-induced ERK1/2 phosphorylation (10 µg/l EGF for 10 min) was determined by ELISA. HAoSMC incubated for 48 h with vehicle showed an EGF-induced increase of ERK1/2 phosphorylation to 197 ± 5% of control (n = 6). HAoSMC incubated for 48 h with 10 nmol/l aldosterone showed an EGF-induced increase of ERK1/2 phosphorylation to 251 ± 5% of control (n = 6), which was significantly larger compared with the vehicle-treated cells. Thus, the 60% increase in EGFR expression is paralleled by a ~60% increase in EGF-induced ERK1/2 phosphorylation. The glucocorticoid dexamethasone (100 nmol/l, 72 h) led to a slight reduction of EGFR expression (67 ± 3% of control; n = 4). To rule out the possibility of reduced EGFR degradation in the presence of aldosterone, we measured EGFR expression in the presence of 10 µmol/l cycloheximide [inhibitor of protein synthesis (Fig. 2D)]. Under these conditions, EGFR abundance decreased significantly under control conditions as well as in the presence of aldosterone, whereas in the absence of cycloheximide aldosterone induced a significant increase. Thus, the effect of aldosterone cannot be explained by reduced EGFR degradation.


Figure 2
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  		Fig. 2. A and B: 72-h incubation with 10 nmol/l aldosterone stimulated EGFR expression in human aorta smooth muscle cells (HAoSMC), as determined by Western blot (A) and ELISA (B) (n = 8, *P < 0.05). The effect was inhibited by 1 µM spironolactone (spiro). C: effect of different concentrations of aldosterone as measured by ELISA (n = 8, *P < 0.05). D: in the presence of protein synthesis inhibitor cycloheximide (chx, 10 µmol/l), EGFR abundance decreases to a similar extent under control conditions and with 10 nmol/l aldosterone (n = 4, *P < 0.05 vs. vehicle). E: aldosterone- and EGFR-induced fibronectin abundance in HAoSMC media measured by ELISA (n = 12–20, *P < 0.05). This induction was inhibited by the EGFR kinase inhibitor AG-1478 (100 nmol/l).

 
As a preliminary marker for the potential pathophysiological relevance of EGFR expression, we determined fibronectin abundance in media, because fibronectin appears to be responsible for increased arterial stiffness (28). Aldosterone increased media fibronectin abundance per se slightly and enhanced EGF-induced fibronectin secretion in HAoSMC, an effect that was inhibited by the EGFR kinase inhibitor AG-1478 (Fig. 2E).

EGFR promoter activity is increased by ligand-bound MR. Previous studies reported an aldosterone-induced increase of EGFR mRNA (9, 27, 31). One way to account for the enhanced expression of EGFR in the presence of aldosterone-bound MR is stimulation of the EGFR promoter with enhanced transcriptional activity. For further investigation of underlying mechanisms, EGFR promoter activity was tested in HEK cells with and without cotransfected hMR (Fig. 3). First, we tested whether aldosterone enhances EGFR expression in HEK cells transiently transfected with hMR. As shown in Fig. 3A, in the presence, but not in the absence, of hMR, aldosterone enhanced EGFR expression. Because HEK cells show substantial basal EGFR expression and cells were transfected only transiently, the relative increase of EGFR expression is smaller compared with HAoSMC. When HEK cells were transfected with hMR, aldosterone elicited a concentration-dependent increase in EGFR reporter activity (Fig. 3B), indicating that aldosterone is able to enhance EGFR promoter activity. When cells were transfected with the empty vector instead of hMR, aldosterone had no effect on promoter activity. The presence of hMR without aldosterone led to a small but not significant rise in EGFR promoter activity.


Figure 3
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  		Fig. 3. A: 72-h incubation with aldosterone stimulated EGFR expression in HEK cells transfected with human mineralocorticoid receptor (hMR) as determined by ELISA (n = 4, *P < 0.05). Interaction between hMR and EGFR promoter assayed by pER reporter gene assay and chromatin immunoprecipitation (ChIP). B: HEK cells transiently transfected with hMR and pER show increased EGFR promoter activity with rising concentrations of aldosterone (n ≥ 5), whereas cells transfected with empty vector (pEGFP) and pER show no response to aldosterone (n = 3, *P < 0.05, **P < 0.01). Right: fold induction achieved by rising aldosterone concentrations. C: Western blot (WB) and immunoprecipitation (IP) of EGFP-hMR with the EGFP antibody used for ChIP. Control ChIP of HEK cells transfected with pGRE-SEAP and either EGFP-hMR or EGFP (IP with anti-EGFP antibody, PCR directed against a region flanking of GRE; PCR controls: water, negative; pGRE-SEAP, positive). D: ChIP of HEK cells transfected with either EGFP-hMR (hMR) or enhanced green fluorescent protein (EGFP; IP with anti-EGFP antibody, PCR directed against endogenous EGFR promoter).

 
ChIP is able to identify DNA fragments bound to hMR. To further investigate whether aldosterone-bound hMR interacts with the EGFR promoter, we established a ChIP assay for the hMR. This technique allows the determination of hMR binding to segments of intact DNA despite unidentified response elements. Because of the lack of suitable antibodies against the hMR, we utilized EGFP-tagged hMR and employed an anti-EGFP antibody for the immunoprecipitation. As a control experiment for our assay we transfected HEK cells with pGRE-SEAP containing three GREs known to bind hMR. Additionally, the cells were transfected with either pEGFP-hMR or empty vector pEGFP and stimulated with aldosterone. Next, we performed ChIP with an EGFP antibody. The DNA bound to the precipitate was assayed using primers annealing adjacent to the GREs. Aliquots of the cell lysates with sheared DNA prior to ChIP were included in the PCR reaction (= input) to ensure that the samples contained equal amounts of DNA fragments. In our experiment, the PCR signals from the input were of equal intensity. After ChIP, only the aldosterone-stimulated samples with hMR showed a substantial PCR band detecting the transfected GREs (Fig. 3C). We concluded that our assay is able to detect direct interactions between the hMR and GREs and is therefore suitable for detecting direct interactions between the hMR and other DNA fragments as well.

Interaction between hMR and the EGFR promoter can be shown by ChIP. Having established ChIP for hMR, we investigated whether the hMR binds to the endogenous EGFR promoter of HEK cells. We repeated the ChIP experiment, this time transfecting the HEK cells only with either pEGFP-hMR or pEGFP and stimulating them with aldosterone. The PCR was designed to amplify a 523-bp region within the EGFR promoter (Fig. 3D). After ChIP, EGFR promoter DNA was detected only in cells transfected with pEGFP-hMR but not in cells transfected with pEGFP, whereas the signals of the input samples were of comparable intensity. This suggests that the hMR binds directly to the EGFR promoter of HEK cells.

EGFR promoter regions involved in interaction with hMR. To narrow down the EGFR promoter part interacting with the hMR, we created deletion constructs of the promoter and measured their responsiveness to aldosterone/hMR. The basal activity decreased for constructs shorter than pER-460 (Fig. 4A), indicating that the promoter region –163 to –460 is important for ~75% of the basal activity.


Figure 4
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  		Fig. 4. A: basal activity and activity in the presence of stimulated hMR of the different EGFR promoter constructs (n = 5–8, *P < 0.05 vs. control). The complete EGFR promoter or its fragments were cotransfected with pEGFP-MR or pEGFP into HEK cells. Promoter activity was determined after 24-h stimulation with 100 nM aldosterone. B: fold induction by aldosterone/hMR of the different promoter constructs.

 
The stimulatory effect of aldosterone/hMR on the constructs containing 1,118 (= pER), 864 (= pER-864), or 510 bp (= pER-510) upstream of the translation start site (Fig. 4, A and B) was of comparable degree. For the constructs, pER-460, pER-352, and pER-316 induction by aldosterone increased significantly (Fig. 4B) and was highest for pER-316. Additional removal of bp –316 to –163 (pER-163) resulted in a significant decline of promoter activation by aldosterone. Thus, the promoter region –316 to –163 seems to be required for a strong stimulatory action of aldosterone. Nevertheless, for pER-163 we still observed a small but significant activation by aldosterone, albeit lower than that of pER-316. Because the construct pER{Delta}510–163 showed a similar degree of activation as pER-163 (Fig. 4B), the promoter region between –1,118 and –510 apparently plays no major role for the interaction with MR.

Our data obtained so far can only be explained by the existence of two sites of interaction between hMR and the EGFR promoter. We tried to simulate the behavior of the basal activity as well as of the activation of the contructs pER, pER-316, and pER-163 (see supplementary Figs. 1–3). The experimental data could be nicely simulated, assuming positive interaction of hMR at two regions of the promoter; one site of strong activation is located between bp –316 and –163, and the second site of weaker activation is located between bp –163 and the translation start site. pER-163 is stimulated by aldosterone dose dependently only in the presence of hMR (Fig. 5, A and B). By contrast, pER-163 was not stimulated by the human GR (hGR, transfected under the same experimental conditions as hMR) stimulated with 100 nmol/l dexamethasone (Fig. 5A). Although our data obtained so far show that pER-163 per se is sufficient to mediate part of the stimulation, this has not been shown for pER-316–163. To confirm that this region per se is able to mediate aldosterone-induced stimulation of promoter activity, we generated an additional construct containing only the region 316–163. Figure 5C shows that pER-316–163 is not stimulated by unbound hMR (i.e., in the absence of aldosterone) or by aldosterone in the absence of hMR. However, when aldosterone was added in the presence of hMR, we observed a dose-dependent stimulation of pER-316–163 activity with an EC50 concentration of ~1 nmol/l (Fig. 5D). hGR did not stimulate pER-316–163 (Fig. 5C). The activity of pER-316–163 was induced approximately fourfold by aldosterone compared with an ~2.5-fold induction of pER-163, supporting our prediction of a stronger and a weaker site. From these data we concluded that the stronger interaction site of the hMR is located in the 316–163 region of the EGFR promoter. Sequence analysis of this region as well as of pER-163 did not reveal any classical GRE element (search performed with AliBaba 2.1; http://darwin.nmsu.edu/~molb470/fall2003/Projects/solorz/), in accord with the lack of effect of hGR. In future studies, we hope to determine the precise MR interaction sequences and assess whether MR binds directly to this site or requires additional factors.


Figure 5
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  		Fig. 5. A: promoter fragment 163–1 (pER-163) is stimulated by aldosterone in the presence, but not in the absence, of hMR. Human glucocorticoid receptor [hGR activated by 100 nmol/l dexamethasone (dexa)] does not stimulate pER-163. B: aldosterone induced dose-dependent stimulation of pER-163 activity with half-maximum effect at ~1 nmol/l in the presence of hMR (n = 9, *P < 0.05 vs. control). C: promoter fragment 316–163 (pER-316–163) is stimulated by aldosterone in the presence, but not in the absence, of hMR. hGR (activated by 100 nmol/l dexa) does not stimulate pER-316–163. D: aldosterone induced dose-dependent stimulation of pER-316–163 activity with half-maximum effect at ~1 nmol/l in the presence of hMR (n = 6, *P < 0.05 vs. control). E: ELISA-based transcription factor DNA binding assay using in vitro synthesized hMR shows direct binding to the positive control [glucocorticoid response element (GRE)] and to pER-163, but not to pER-316–163 (n = 5–6, *P < 0.05 vs. LacZ).

 
To test whether hMR interacts directly with pER-163 or pER-316–163, we performed ELISA-based transcription factor DNA binding assays using in vitro synthesized hMR (or LacZ as negative control) and the EGFR promoter fragments 163–316 and 1–163, respectively, as probes (see EXPERIMENTAL PROCEDURES). The results obtained indicate that there is a direct interaction between hMR and the EGFR promoter region 1–163 but not region 316–163 (Fig. 5E).

NH2-terminal domain of hMR is involved in interaction with EGFR promoter. Subsequently, we created deletion constructs of the hMR lacking the NH2-terminal activator function-1 (AF1) or AF1 plus DNA-binding domain C. We then compared the transactivation capacity of the complete MR, CDEF-hMR (deletion construct containing domains C, D, E, and F of the hMR), and DEF-hMR (deletion construct containing domains D, E, and F of the hMR) in a GRE reporter gene assay and observed that the transactivation activity of the truncated CDEF-hMR, as expected, exceeded that of the complete hMR, whereas DEF-hMR showed no transactivation activity above control levels (Fig. 6B) (39). There was no significant difference in the dose-response curves of CDEF-hMR and hMR in the GRE reporter gene assay (Fig. 6B, inset). Figure 6C shows that CDEF-hMR still possesses the ability to translocate into the nucleus. DEF-hMR can also translocate into the nucleus (data not shown). Next, we tested pER activation induced by hMR, CDEF-hMR, and DEF-hMR in the presence of aldosterone. As shown in Fig. 6D, CDEF-hMR was significantly less effective than the full-length hMR in activating the EGFR promoter. DEF-hMR had no effect at all. Thus, the NH2-terminal domain is required for full stimulation of the EGFR promoter. Finally, we tested whether the NH2-terminal domain is also required for pER-163 or pER-316–163 activation. As shown in Fig. 6D, pER-316–163 was not stimulated by CDEF-hMR, and stimulation of pER-163 by CDEF-hMR was significantly smaller compared with hMR.


Figure 6
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  		Fig. 6. A: characterization of hMR deletion constructs CDEF- and DEF-hMR. CDEF-hMR contains MR domains C, D, E, and F; DEF-hMR possesses regions D, E, and F but lacks DNA-binding domain C (= DBD); LBD, ligand-binding domain. B: transactivation activity of hMR and CDEF- and DEF-hMR tested in HEK cells with GRE-SEAP reporter gene assay in the presence of 100 nmol/l aldosterone. Inset: GRE-SEAP transactivation activities of hMR and CDEF-hMR with varying aldosterone concentrations (n = 8). C: unstimulated CDEF-hMR is located mainly in the cytosol of transfected HEK cells and translocates into the nucleus when 1 nM aldosterone is added. D: CDEF-hMR stimulates the EGFR promoter significantly less than hMR. DEF-hMR is completely inactive (incubation with 100 nmol/l aldosterone for 24 h; n = 6–9, *P < 0.05 vs. vector, **P < 0.05 vs. hMR). CDEF-hMR exerted no effect at promoter fragment 316–163 (pER-316–163) but induced a small but statistically significant stimulation at promoter fragment 163–1 (pER-163).

 

   DISCUSSION
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Our results suggest that aldosterone, but not dexamethasone, can lead to an upregulation of EGFR expression in rat aorta and HAoSMC, which is not explained by reduced EGFR degradation. Furthermore, aldosterone enhances EGF-induced ERK1/2 phosphorylation in HAoSMC, showing that functional EGFR is expressed, similar to our earlier results obtained in CHO cells. According to our data in HEK cells, aldosterone modulates EGFR expression through an interaction between ligand-bound hMR and the regions 316–163 and 163–1 bp of the EGFR promoter. Because dexamethasone/hGR did not elicit a stimulatory effect in our experiments and did not stimulate pER in a previous study (18), we assume that the effect is MR specific and does not involve the GR. This conclusion is supported by the fact that spironolactone inhibited EGFR induction in primary culture. Overall, our data are in agreement with previous pharmacological studies (6, 9, 18, 27, 31).

The aldosterone concentrations achieved in the in vivo experiments are higher than control aldosterone concentrations in healthy animals. Yet, under certain conditions, like for example salt depletion (30) or renal dysfunction (38), plasma aldosterone can rise to levels similar to those in our study. Furthermore, the dose-response curve in HAoSMC shows that lower concentrations of aldosterone can be also effective and that the half-maximum concentrations are close to the EC50 value of hMR. Previous in vivo studies in rats have already demonstrated that no significant elevations in blood pressure occur after only 5 days of aldosterone infusion, ruling out that our results are due to changes in blood pressure (29, 33). Concerning the vascular cell types involved, our experiments with cells in primary culture suggest that EGFR expression is enhanced, at least in VSMCs, which therefore behave like certain other cell types investigated before (6, 9, 18, 27, 31).

Promotion of EGFR expression can have severe consequences for the affected tissue because transactivation of the EGFR is used by a variety of mediators with potential pathophysiological relevance like GPCR-signaling peptides, cytokines, and growth factors (13, 19, 20, 42). For VSMC and cardiomyocytes, GPCR-signaling peptides like ANG II, phenylephrine, and endothelin have been shown to induce mitogenic and proliferative effects via enhanced phosphorylation of the EGFR, ultimately leading to cardiovascular remodeling independently of blood pressure (2, 5, 8, 14, 22, 26, 36, 41, 46). Various groups have shown that, in VSMC, aldosterone- or ANG II-mediated ERK phosphorylation is dependent on EGFR transactivation and that EGFR kinase inhibitors lead to attenuation of growth, migration, or ANG I-converting enzyme expression (4, 17, 24, 25, 25, 40, 43). Interestingly, EGFR expression also supports fibrosis in cardiovascular and renal tissue (8, 16, 44). For example, endothelin-induced phosphorylation of the mitogen-activated protein kinase and endothelin-induced increase in collagen I gene activity were completely prevented by an inhibitor of the EGFR kinase. Therefore, EGFR transactivation seems to be a key factor promoting GPCR-mediated growth and remodeling, and its upregulation by aldosterone in vasculature could enhance the deleterious effects of various vasoactive peptides. In this respect it, is worth mentioning that endothelin receptor antagonism prevents aldosterone-induced vascular remodeling (37) and that the aldosterone synthase inhibitor FAD286 ameliorates ANG II-induced end organ damage (7). In our study, aldosterone led to an increase in fibronectin abundance and enhanced the effect of EGF on fibronectin abundance in media from HAoSMC. Both effects could be prevented by the EGFR kinase inhibitor AG-1478. This supports the hypothesis that aldosterone furthers vascular remodeling via EGFR expression. In addition, we (11) showed recently that aldosterone potentiates H2O2-induced collagen abundance in HAoSMC and that this effect depends on EGFR. This hypothesis relies at the moment on the correlation of EGFR expression and alterations in matrix homeostasis on the one side and the preventive action of AG-1478 on the other side. It could be strengthened by future demonstration that silencing of EGFR prevents the mentioned effects of aldosterone.

Various cellular signaling pathways are involved in the regulation of EGFR expression, and therefore we tried to determine the mechanism(s) employed by aldosterone/MR. For this purpose we used HEK cells, which can be readily transfected and where we are able to control hMR expression. The observed increase in EGFR promoter activity by ligand-bound MR supports the hypothesis of an interaction between aldosterone/MR and the EGFR promoter; however, it does not allow distinguishing between a direct and an indirect interaction. To address this question, we established a ChIP assay for the hMR using a canonical GRE, which has been shown to bind hMR (39). We then applied this ChIP assay to investigate MR-EGFR promoter interactions and could show that there is indeed a direct interaction.

To narrow down the site of hMR-EGFR promoter interaction we used truncated constructs of the EGFR promoter and modeled the experimental results obtained (see supplementary figures). Our model predicted two independent regions of interaction, a stronger one between position –316 and –163 of the promoter and a weaker one between position –163 and –1. These predictions were confirmed by our experimental data for pER-316–163 and pER-163. Although the two hMR-responsive regions seem to interact independently with hMR, we cannot exclude at the moment that additional transcription modulators also play a role during hMR-induced pER stimulation. This issue will be investigated in future studies. Both hMR-responsive regions were completely unresponsive to the hGR. Because CDEF-hMR, lacking the AF1 domain, was significantly less effective than hMR, the NH2-terminal domain seems to be crucial for the interaction.

Finally, ELISA-based transcription factor DNA binding assays, using in vitro synthesized hMR, support the hypothesis that the promoter region 1–163 directly interacts with hMR, although no canonical GRE element could be identified, in agreement with the lack of stimulation by the GR. This seems not to be the case for region 163–316, which possibly needs additional transcription factors mediating the interaction.

The exact mineralocorticoid response element in pER is not yet known, but we are currently addressing this question in detail to determine the precise hMR interaction sequence by fine mapping of the promoter segment. Once this element has been further defined, it will be interesting to see to what extent it discriminates between hMR and hGR and which other promoters contain this element. This information could help us to understand mineralocorticoid selectivity and to find further aldosterone-regulated genes. Furthermore, it will be important to determine whether the transcriptional action of hMR at the EGFR promoter requires the presence of additional transcription factors and, if so, which.


   GRANTS
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 ABSTRACT
 EXPERIMENTAL PROCEDURES
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 GRANTS
REFERENCES
 
This study was supported by the Deutsche Forschungsgemeinschaft (DFG Ge905/8-1).


   ACKNOWLEDGMENTS
 
We thank Drs. Farman and Johnson for providing the pEGFP-hMR and pERLuc. We also thank Dr. A. Fiebeler for helpful discussions.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. Gekle, Physiologisches Institut, Universitaet Wuerzburg, Roentgenring 9, 97070 Wuerzburg, Germany (e-mail: michael.gekle{at}mail.uni-wuerzburg.de)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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