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Am J Physiol Heart Circ Physiol 283: H2458-H2465, 2002. First published August 29, 2002; doi:10.1152/ajpheart.00295.2002
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Vol. 283, Issue 6, H2458-H2465, December 2002

Endothelin-1 increases calcium and attenuates renin gene expression in As4.1 cells

Michael J. Ryan1, Thomas A. Black2, Susan L. Millard3, Kenneth W. Gross2, and George Hajduczok1

1 Department of Physiology and Biophysics and 3 Pharmacology and Toxicology, State University of New York at Buffalo, Buffalo 14214; and 2 Department of Molecular and Cellular Biology, Roswell Park Cancer Institute, Buffalo, New York 14263


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Endothelin-1 (ET-1) is a potent vasoconstrictor and blood pressure modulator. Renin secretion from juxtaglomerular (JG) cells is crucial for blood pressure and electrolyte homeostasis and has been shown to be modulated by ET-1; however, the cellular and molecular mechanism of this regulation is not clear. The purpose of this study was to gain a better understanding of the cellular and molecular pathways activated by ET-1 by using a renin-producing cell line As4.1. ET-1 caused an increase in As4.1 cell intracelluar Ca2+ concentration ([Ca2+]i) mediated by the ETA receptor as its antagonist, BQ-123, abolished the response. The nitric oxide donor nitroprusside, but not 8-bromo-cGMP, reduced the time necessary for successive ET-1 responses. Endothelin-3 had no effect on [Ca2+]i. ET-1 dose dependently increased total inositol phosphates with an EC50 of 2.1 nM. ET-1 reduced renin mRNA by 68% independently of changes in message decay. With the use of a renin-luciferase reporter system in As4.1 cells, ET-1 reduced luciferase activity by 51%, suggesting that renin gene transcription is directly modified by ET-1.

renin; endothelin; transcription; nitric oxide


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

THE PHYSIOLOGICAL CONTROL of renin production and release from juxtaglomerular (JG) cells is, in part, regulated by circulating vasoactive peptides such as angiotensin II. Endothelin (ET), a potent vasoconstrictor that binds to specific membrane receptors in smooth muscle to activate phospholipase C (PLC), inositol phosphate (IP), and intracellular calcium concentration ([Ca2+]i), has also been implicated in the control of renin production and release.

The effect of ET in the kidney appears to be localized to preglomerular arterioles (site of JG cells) as evidenced by its ability to cause a dose-dependent reduction in glomerular filtration (3). There are some reports that ET can regulate [Ca2+]i and inhibit cAMP-stimulated renin production and release (1, 19); however, less is known about the cellular pathways activated by ET and how they might lead to the regulation of renin at gene level. This lack of understanding has been confounded by the potential concurrent effect of the potent endothelium-derived vasodilator nitric oxide (NO), which by itself has been reported to modulate renin release (2, 12, 27). The interactions between NO and ET pathways (9, 18) may, therefore, complicate the effect of either agent alone on renin production and secretion. In addition, it has been shown that repeated application of ET causes receptor desensitization that can be reversed by NO donors (9, 18).

To date, the information pertaining to the cellular regulation of renin by ET has largely been obtained by using primary JG cell cultures from rats and mice. Therefore, the current study utilizes a different cellular model of renin-expressing cells, As4.1 (28), to further our understanding of ET and how it regulates renin from the receptors, to second messengers, to renin gene regulation.


    MATERIALS AND METHODS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Cell culture. As4.1 cells (ATCC No. CRL2193) are a renin-expressing clonal cell line derived from the kidney neoplasm of a transgenic mouse containing a renin promoter driven Simian virus-40 T-antigen transgene that demonstrated appropriate developmental, cell, and tissue-specific expression (28). Cells were maintained in humidified room air containing 5% CO2 at 37°C. As4.1 cells were cultured in DMEM supplemented with 10% fetal bovine serum. For measurement of [Ca2+]i, As4.1 cells were cultured onto 18-mm coverslips in serum-free DMEM 24 h before the experiment. To measure total intracellular IP metabolites, As4.1 cells were cultured 48 h before the experiments onto six-well cell culture dishes in DMEM with 10% fetal bovine serum. Similarly, cells were cultured onto six-well dishes 24 h before transfection experiments, whereas renin expression was measured from As4.1 cells grown on 100-mm petri dishes.

[Ca2+]i measurements. [Ca2+]i in individual As4.1 cells was measured using the Ca2+-sensitive dye fura 2 as previously described (25). As4.1 cells were washed with a Ca2+-free Ringer solution (in mM: 145 NaCl, 5 KCl, 1 MgCl2, 20 HEPES, 20 glucose, and 1 mM EGTA), followed by an incubation with 5 µM fura 2-AM for 20 min, in the dark, at room temperature. After the incubation, the cells were washed with normal Ringer solution (in nM: 145 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 20 HEPES, and 20 glucose) and placed in serum-free DMEM for 1 h at room temperature and 5% CO2.

The coverslips were mounted on the stage of an inverted microscope (Diaphot-TMB, Nikon) equipped for epifluorescence using a ×40 oil-immersion lens. The cells were superfused with normal Ringer or 0 Ca2+ Ringer, when specified, at a rate of 2-3 ml/min. The total bath volume was ~500 µl.

Fluorescence of the fura 2-loaded cells was measured at room temperature by using a digital image analysis program Metafluor (Universal Imaging). Images were taken at excitation wavelengths of 340 and 380 nm at 3-s intervals by using a silicon-intensified target camera (Hamamatsu). The average whole cell 340/380 ratio (R) values were converted to Ca2+ concentrations according to standard equations (10) by using rectangular glass capillary tubes containing known concentrations of Ca2+. [Ca2+]i was obtained using this ratio and the standard Ca2+ solutions in the following equation: [Ca]i = Kd (Sf/Sr)[(R - Rmin)/(Rmax - R)] where Kd is the dissociation constant for fura 2 of 224 nM at room temperature, Rmax is the 340/380 nm ration at saturating levels of [Ca2+]i, Rmin is the ratio at zero [Ca2+]i, Sf is the 380-nm excitation fluorescence intensity in zero [Ca2+]i, and Sr is the 380-nm excitation fluorescence intensity in saturating levels of [Ca2+]i.

As4.1 cell stimulation. Ca2+ experiments were performed while cells were superfused with either normal Ringer or Ca2+-free Ringer containing 50 nM bradykinin, 10 nM ET-1, 10 nM ET-3, or 10 nM ET-1 + 1 µM BQ-123 (ET receptor antagonist). The NO donor sodium nitroprusside (SNP, 500 µM) and the membrane-permeable analog of cGMP, 8-bromo-cGMP (1 mM), were also added to the above solutions and superfused at the indicated times.

Inositol phosphate measurement. Total cytosolic inositol phosphate was measured in As4.1 cells labeled with myo-[2-3H]inositol (15). Cells were incubated with myo-[2-3H]inositol (5 µCi/ml media) in serum-free, inositol-free DMEM 24 h before stimulation with ET. This allowed sufficient time for equilibration of the radiolabeled inositol with the membrane of the cell. To remove excess myo-[2-3H]inositol after the 24-h incubation, cells were washed twice with serum- and inositol-free DMEM. Cells were allowed to equilibrate in this medium for at least 1 h. Fifteen minutes before the start of an experimental protocol, medium was aspirated, followed by the addition of a Krebs solution (in mM: 118 NaCl, 4.6 KCl, 2.4 MgCl2, 1.2 K2HPO4, 24 NaHCO3, 11 glucose, 25 HEPES, and 2.5 CaCl2) containing 12 mM LiCl and 1 mM myoinositol. LiCl was necessary to prevent the reincorporation of IP from the cytosol to the membrane following activation of second messenger pathways.

After stimulation of the As4.1 cells with increasing doses (0.1-100 nM) of ET for 5 min each, 9% perchloric acid was added to each well to prevent the further hydrolysis of phosphotidylinositol 4,5-bisphosphate. The lysed As4.1 cell suspension was pelleted with the supernatant containing cytosolic fluid and metabolites. Pellets were saved and used in a Lowry assay to measure total protein. The supernatant was run on a Dowex column and washed with several solutions including myo-inositol (5 mM), sodium tetraborate (5 mM), and 1 M formate/0.1 M formic acid to collect the myo-[2-3H]inositol from the cytosolic fraction. Myo-[2-3H]inositol was measured with a scintillation counter and expressed as a percentage of control IP values.

RNA isolation and Northern blot analysis. Total RNA was isolated from As4.1 cells following application of the ET-1 by using the Ultraspec RNA isolation kit (Biotecx). The method was performed according to manufacturer's instructions and utilizes the guanidium thiocyanate, acid phenol, and chloroform approach to RNA purification. Ten-microgram samples of As4.1 cell RNA were used for Northern blot analysis as previously described in our laboratory (21). A mouse submandibular Ren 2d cDNA was used as a probe for renin (17), and GAPDH mRNA levels were analyzed and used as an internal control.

Renin message stability. As4.1 cell gene transcription was inhibited with actinomycin D (10 µg/ml). As4.1 cells were stimulated with ET-1 (10 nM) just before the start of the experiment. At the time of 0, 6, 13, and 20 h after the addition of ET-1, total cellular RNA was harvested, and renin expression was measured by using Northern blot analysis. The same experiment was run concurrently in the absence of ET-1 to serve as a time-matched control.

DNA transfections. The luciferase derivative of a DNA construct containing the renin proximal promoter and 4,000 bp of 5'-flanking sequence described previously (26) was introduced to As4.1 cells via liposome-mediated transfection using Fugene 6 transfection reagent (Boehringer Mannheim). This construct contains a 5'-flanking sequence of the Ren-1c gene fused with the Ren-2 proximal promoter. Briefly, Fugene 6 reagent, diluted in Optimem media, was added to the DNA. This mixture was added to As4.1 cells in culture on six-well cell culture dishes 2 h before ET-1 stimulation. Six microliters of Fugene 6 was used with a total DNA amount of 3.5 µg. To correct luciferase activity for variations in transfection efficiency, As4.1 cells were cotransfected with 200 ng of plasmid containing a Rous sarcoma virus (RSV) promoter driving beta -galactosidase (RSVbeta gal). As4.1 cells were transfected with a promoterless luciferase construct to determine the background luciferase activity.

Luciferase assay. Immediately after ET-1 stimulation, medium was aspirated from the six-well dishes, and cells were washed with phosphate buffer solution (in mM: 137 NaCl, 2.7 KCl, 4.3 Na2HPO4, and 1.4 KH2PO4; pH 7.3). Luciferase activity (chemiluminescence) was measured by using the Luciferase Reporter 1000 Assay System (Promega) in accordance with manufacturer's instructions. Briefly, 1× reporter lysis buffer was added to the cells to remove them from the cell culture dishes. The cells were collected in microcentrifuge tubes and subjected to a freeze-thaw cycle with dry ice and ethanol to lyse the cells completely. The lysed cells were then pelleted, and the supernatant luciferase activity was measured with a Monolight 2010 luminometer (Analytical Luminescence Laboratory). Results were expressed as percent luciferase activity of RSV luciferase-transfected cells.

beta -Galactosidase. beta -Galactosidase activity was measured using Galacto-Light Plus Chemiluminescence Reporter Assay (Tropix) in accordance with manufacturer's instructions.

Statistical analysis. All data are presented as means ± SE. Data are considered significantly different if P < 0.05, as evaluated by repeated-measures ANOVA or paired t-tests where specified. Statistical tests are noted with each figure along with the number of experiments.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

As4.1 cell calcium regulation by ET-1. Individual As4.1 cells loaded with the Ca2+-sensitive dye fura 2 were superfused with normal Ringer solution containing 10 nM ET-1 to measure changes in [Ca2+]i. ET-1 application causes a large transient increase in [Ca2+]i (Fig. 1A). Measurements from 26 cells stimulated with ET-1 revealed a mean increase in Ca2+ of 838 ± 81 nM above baseline values of 69 ± 3 nM (Fig. 1A, inset). To determine whether this response is mediated via intracellular or extracellular Ca2+ stores, the experiments were repeated with cells superfused in Ca2+-free containing Ringer solution (Fig. 1B). Despite the absence of extracellular Ca2+, ET-1 (10 nM) still elicited a rapid, transient increase in [Ca2+]i. Moreover, this response was not attenuated as evidenced by the 868 ± 76 nM increase over the basal [Ca2+]i (67 ± 3 nM, Fig. 1B, inset) and thus indicates that the response of As4.1 cells to ET-1 is mediated solely through mobilization of intracellular Ca2+ stores rather than through voltage-activated Ca2+ channels in the plasma membrane.


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  		Fig. 1.   A: endothelin-1 (ET-1, 10 nM) stimulates a rapid, transient increase of intracellular Ca2+ concentration ([Ca2+]i) in As4.1 cells. Shaded bar, application of ET-1. Mean increase (inset) in [Ca2+]i measured from 26 cells illustrates a peak calcium concentration of 838 ± 81 nM (P < 0.05, paired t-test). B: removal of extracellular calcium did not attenuate the ability of ET-1 to elicit the rise in [Ca2+]i. Calcium-free Ringer ([Ca2+]o, open bar) solution was used to superfuse the As4.1 cell while ET-1 (10 nM) in the same Ringer solution was applied (solid bar). Average [Ca2+]i in stimulated cells (inset) was 868 ± 76 nM and was significantly higher than basal [Ca2+]i (P < 0.05, paired t-test, n = 26). Data are represented as means ± SE.

To determine the ET receptor subtype mediating this response, we utilized various ET receptor agonists and antagonists. ET-1 is an agonist of both ETA and ETB receptors (20). Therefore, we stimulated As4.1 cells with ET-3 (10 nM), an agonist primarily of ETB receptors (23). No change in [Ca2+]i was observed, suggesting that the response is not mediated by ETB receptors. However, a subsequent dose of ET-1 did elicit a Ca2+ response (Fig. 2A). Mean changes in [Ca2+]i are depicted in Fig. 2A (inset) with an average increase of 587 ± 57 nM over basal levels of 68 ± 4 nM. To further elucidate which ET receptor was mediating the Ca2+ response in these cells, BQ-123 (1 µM), a specific ETA receptor antagonist (11), was utilized to determine whether the Ca2+ transients could be inhibited. When ET-1 was applied to As4.1 cells in the presence of BQ-123, no increase in [Ca2+]i was observed. Significantly, on removal of the BQ-123, application of ET-1 produced an elevated [Ca2+]i (Fig. 2B), thus supporting a role for ETA receptors in mediating the Ca2+ response in As4.1 cells.


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  		Fig. 2.   A: application of the ETB receptor agonist ET-3 (10 nM) did not change [Ca2+]i in As4.1 cells. Subsequent application of ET-1 (10 nM) resulted in an increased [Ca2+]i. Mean [Ca2+]i (inset) in ET-3-stimulated cells (69 ± 4 nM) was not different from basal levels (P < 0.05, ANOVA); however, subsequent application of ET-1 resulted in an increase of [Ca2+]i to 587 ± 57 nM (n = 17). B: ETA receptor antagonist BQ-123 (1 µM) inhibited ET-1 calcium stimulation of As4.1 cells. Cells were superfused with normal Ringer solution containing BQ-123 (1 µM) and ET-1 (10 nM). A significant increase (638 ± 47) in [Ca2+]i was observed when BQ-123 was removed from the bathing solution (inset) (P < 0.05, ANOVA). Data are represented as means ± SE (n = 17).

Figure 3A illustrates the occurrence of homologous desensitization of the [Ca2+]i response to repeated application of ET-1 (10 nM, 1 min duration). The time course of spontaneous recovery from desensitization to a second (test) application of ET-1 is shown in Fig. 3B. The test responses, as compared with the first application of ET-1 (control), are shown to recover by 35 min (Fig. 3B). However, when the control application of ET-1 is immediately followed by the superfusion of the NO donor SNP (500 µM), the [Ca2+]i response to the test application of ET-1 fully returns within 10 min (Fig. 4A). A similar recovery from desensitization was observed with the NO donor S-nitroso-N-acetyl penicillamine (1 µM) (data not shown). To investigate whether the recovery from desensitization in the presence SNP was due to a cGMP-dependent mechanism, 8-bromo-cGMP (1 mM) was substituted for SNP following the ET-1 control challenge. Figure 4B shows that the cGMP analog did not alter the recovery from ET-1 desensitization thus suggesting that NO may have a direct effect on the interaction of ET-1 with its receptor in renin-expressing cells. Importantly, activating two different receptor populations with successive doses of ET-1 and bradykinin (50 nM) (Fig. 5) resulted in an increased [Ca2+]i of similar magnitude either in the presence or absence of extracellular calcium. The mean changes in [Ca2+]i in the presence of calcium were 825 ± 34 and 804 ± 30 nM for ET-1 and bradykinin, respectively (n = 12). In the absence of extracellular Ca2+, the mean increase in [Ca2+]i was 718 ± 25 and 725 ± 21 nM for ET-1 and bradykinin, respectively (n = 18). These data suggest that the lack of a calcium response by successive doses of ET-1 are not due to depleted [Ca2+]i stores and supports the notion of ET receptor desensitization.


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  		Fig. 3.   A: homologous receptor desensitization in two As4.1 cells. Response in response to repeated application of ET-1 (solid bars). Only the first of two ET-1 stimuli results in an increased [Ca2+]i B: time course of recovery from desensitization of [Ca2+]i responses to test challenge of ET-1. Normalized responses compared with first application (control) of ET-1. All applications of ET-1 were of 1-min duration at 10 nM (n > 25 for each time period; data are means ± SE; *P < 0.01, ANOVA).



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  		Fig. 4.   A: modulation of ET-1 (10 nM)-induced [Ca2+]i transients (solid bars) by the nitric oxide (NO) donor sodium nitrorusside (SNP, 500 µM; open bar). SNP treatment prevented the occurrence of desensitization to a test ET-1 challenge. B: recovery from desensitization of [Ca2+]i responses to a test challenge of ET-1 is enhanced in the presence of SNP (500 µM) but not by 8-bromo-cGMP. Normalized responses compared with first application (control) of ET-1 (10 nM) only. Application of SNP or 8-bromo-cGMP (1 mM) began at the end of the initial ET-1 perfusion. Test ET-1 applications occurred at the indicated time points and are relative to end of first ET-1 challenge. (n > 22 for each time period; data are means ± SE, *P < 0.01, ANOVA).



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  		Fig. 5.   Successive doses of ET-1 (10 nM) and bradykinin (BK) are able to elicit increases in [Ca2+]i of similar magnitude in the presence of extracellular calcium (A) or the absence of extracellular calcium (B).

Phosphotidylinositol 4,5-bisphosphate hydrolysis by ET-1. We measured total IP in As4.1 cells incubated with various concentrations of ET-1. The results demonstrated a concentration-dependent increase in IP by ET-1 with an EC50 of 2.1 nM (Fig. 6). These data provide support for an ET-1-induced, IP-mediated release of [Ca2+]i stores in As4.1 cells. Subsequent experiments used a concentration of 10 nM ET-1 because it resulted in a near-maximal IP response.


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  		Fig. 6.   Dose-dependent increase in total As4.1 cytosolic inositol phosphate (IP) after incubation with ET-1. EC50 of the response to ET-1 occurs at a concentration of 2.1 nM.

Renin gene expression-transcription regulation by ET-1. Northern blot analysis was used to assess renin mRNA levels in control and ET-1-stimulated As4.1 cells. Cells were incubated for 24 h with 10 nM ET-1 or a super-maximal dose of 100 nM before harvesting the RNA. GAPDH mRNA was also quantified and used to correct for differences in gel loading. After stimulation of As4.1 cells with ET-1, renin expression was reduced to 32 ± 3 and 32 ± 6% of control at 10 and 100 nM, respectively (Fig. 7). These results demonstrate that ET-1 is able to modulate renin gene expression in the As4.1 cell as well as demonstrating that a maximal inhibitory response occurs at a concentration of 10 nM (maximal IP response occurred near this dose as well). After inhibiting transcription with actinomycin D (10 µg/ml), we used Northern blot analysis to investigate whether the kinetics of renin message decay, as a potential contributor to the observed decrease in expression, was altered by the presence of ET-1. The half-life of the renin message was unaffected by ET-1 (8.2 ± 0.8 h for control vs. 8.4 ± 0.7 h for 10 nM ET-1) (Fig. 8), thus demonstrating that message decay was not responsible for the decreased expression and supporting the direct influence of ET-1 on renin gene transcription.


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  		Fig. 7.   ET-1 inhibits renin expression in As4.1 cells. Twenty-four hours of incubation with either 10 or 100 nM ET-1 lead to a significant (*P < 0.05, ANOVA) decrease of renin expression to 32.6 ± 3.5% and 32.4 ± 6.6% of control expression levels, respectively. Each bar represents n = 3 of 10-µg samples of RNA quantified by Northern blot analysis. GAPDH expression was used to correct for variations in gel loading. Data are means ± SE.



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  		Fig. 8.   ET-1 does not alter the rate of renin message decay. As4.1 cells were incubated with 10 µg/ml actinomycin D and 10 nM ET-1. Northern blot analysis of 2-µg samples was performed at time of 0, 6, 13, and 20 h. In the absence of ET-1 (open circles, solid line), the renin mRNA half-life was 8.2 ± 0.8 and 8.4 ± 0.7 h in the presence of ET-1 (closed circles, dashed line) (n = 3, P > 0.55, paired t-test).

Recently, a 4,000-bp (-117 to -4100 relative to the transcription start site) sequence upstream of the renin proximal promoter (+6 to -117) was determined to contain an enhancer sequence as well as other important sequences required for renin gene regulation (16). Moreover, this construct may contain elements important for the effects of other physiological stimuli on renin gene transcription, including interleukin-1beta (17) and mechanical force (21). Therefore, we used the luciferase derivative of a DNA construct containing this 4,000-bp sequence fused to the renin proximal promoter and a luciferase reporter gene (construct c) to assess the role of ET-1 in the transcriptional regulation of the renin gene. Luciferase activity was measured as an estimate of transcriptional activity. Transfected As4.1 cells were incubated with 10 nM ET-1 for 48 h before luciferase measurements. The 48-h exposure to ET-1 was selected based on preliminary transfection experiments from our laboratory (K. W. Gross) utilizing the 4,000-bp renin gene sequence fused to a chloramphenicol acetyltransferase reporter gene (data not shown). Medium was changed after the first 24 h, and fresh ET-1 was added. beta -Galactosidase DNA was cotransfected with the luciferase construct and used to correct for variations in transfection efficiency. The results shown in Fig. 9 illustrate a reduction of transcriptional activity by 51% (P < 0.005) from control. Construct a (promoterless-luciferase construct) and construct b (luciferase plus the renin promoter) were used as negative controls.


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  		Fig. 9.   ET-1 inhibits renin gene transcription in As4.1 cells. Construct a (promoterless vector, PGL2 Basic) and construct b (promoter-luciferase) were used as negative controls because no difference was observed between ET-1-treated cells (hatched) and control cells (solid). After a 48-h incubation with 10 nM ET-1, construct c (4,000 bp, promoter-luciferase) caused a 51% decrease in renin gene transcription (P < 0.05, paired t-test).


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

The experiments described in the present study demonstrate that ET-1 stimulation of As4.1 cells results in the activation of a signal transduction pathway involving an increase in IP and [Ca2+]i. The increase in [Ca2+]i was dependent solely on intracellular stores and was mediated by the ETA receptor. Furthermore, receptor binding was altered by the presence of NO, because the time necessary for successive applications of ET-1 to elicit a rise in [Ca2+]i was markedly reduced in the presence of nitroprusside. These effects were likely not due to problems with refilling [Ca2+]i stores because successive doses of two different agonists ET-1 and bradykinin stimulated increases in [Ca2+]i both in the presence and absence of extracellular Ca2+. At the molecular level, incubation of As4.1 cells with ET-1 resulted in a reduction of renin gene transcription and message that occurred independently of changes in renin message decay.

As4.1 signal transduction. ET-1 stimulation of the As4.1 cells results in a sharp, transient rise in [Ca2+]i that is mediated solely by intracellular release. The absence of extracellular Ca2+ contribution to the Ca2+ increase in As4.1 cells is consistent with reports that JG cells do not possess any voltage-activated Ca2+ channels (14) that might lead to an influx of Ca2+ from the cell membrane. Our findings are also consistent with what has been reported in vivo where Schroeder et al. (24) recently demonstrated that ET causes a rapid transient rise in [Ca2+]i from vascular smooth muscle of preglomerular arteries (site JG cells). It is important to note that JG cells are postulated to be modified smooth muscle cells (4), and therefore one might expect renin-expressing cells and smooth muscle of preglomerular arteries to behave similarly when challenged with similar physiological stimuli. The release of [Ca2+]i by ET-1 in As4.1 cells is likely mediated by a phospholipase C-mediated second messenger pathway as evidenced by the concentration-dependent increase in total IP. Importantly, the EC50 for ET-1 in As4.1 cells is consistent with what has been reported in other cells types (6, 8).

ET receptor subtype. The study by Schroeder et al. (24) also indicated that the response of preglomerular artery vascular smooth muscle to ET-1 was mediated by the ETA receptor. This is consistent with our findings that ET-3, a selective ETB agonist, did not cause an increase in [Ca2+]i and the ETA receptor antagonist BQ-123 was able to inhibit the response of As4.1 cells to ET-1. In further support for the role of ETA in mediating the actions of ET-1 on renin, it has been reported that blockade of the ETA receptor in chronically instrumented dogs results in an increase in plasma renin activity. Although the current findings demonstrate a role for the ETA receptor in the regulation of renin, it should be noted that Kramer et al. (13) found ET to act through ETB receptors in primary JG cells. In addition, Endemann et al. (7) found that ET-3 could elicit an increase in [Ca2+]i in As4.1 cells. The difference may be explained by the higher concentration of ET-3 used (100 nM) by Endemann et al. (7) compared with the 10 nM used in the current study.

Given that repeated applications of ET-1 were unable to elicit a rise in [Ca2+]i for ~35 min after the initial stimulus, we speculated that homologous desensitization of the ET-1 receptor had occurred. This speculation was supported by our data showing that addition of SNP, a NO donor, was able to abrogate the desensitization to the secondary challenge of ET-1 by 10 min. The reversal was not mimicked by cGMP, suggesting that the presence of NO may be effecting the affinity that ET has for its receptor. Furthermore, this finding is consistent with the finding of Goligorsky et al. (9) that NO may play a role in the physiological termination of an ET-1 stimulus through displacing bound ET-1 from its receptor. Whereas these findings strongly support a role for receptor desensitization, they do not completely rule out the possibility that [Ca2+]i stores are depleted after the initial ET-1 stimulation. Furthermore, there is evidence that NO increases reuptake of calcium in the sarcoplasmic reticulum of vascular smooth muscle, making it possible that the effect of SNP to reduce the time between ET-1 responses is an artifact of refilling [Ca2+]i stores more rapidly (5). Therefore, we performed additional experiments to test whether successive doses of different agonists could elicit calcium responses in As4.1 cells. Our data demonstrating that successive doses of ET-1 followed by bradykinin doses indicate that [Ca2+]i stores are not depleted as each agonist leads to calcium responses of similar magnitude. This evidence further supports the occurrence of receptor desensitization by ET-1 that can be modulated by the presence of NO. Taken together the current findings are important for gaining a better understanding of how the effects of ET-1 on renin-expressing cells can be influenced by other physiologically important molecules.

Renin gene transcription. The data from the current study show that renin gene expression was markedly attenuated by the presence of ET-1. Moreover, this change in expression was not the result of posttranscriptional modification because it occurred independently of renin message decay kinetics. This by itself, suggests that the initial activation of signaling pathways by ET-1 ultimately leads to a decrease in renin gene transcription; however, to directly test this, we performed transient transfections in As4.1 cells with a 4,000-bp 5'-flanking region of the renin gene fused to a luciferase reporter gene. We selected this region based on studies done by our laboratory (K. W. Gross) where extensive promoter analysis revealed this to be a region of importance for the transcriptional regulation of the renin gene (16, 26). The current study demonstrates for the first time that ET-1 stimulation leads to a direct inhibition of renin gene transcription. Importantly, the level of transcriptional inhibition was commensurate with what was observed by Northern blot analysis.

As4.1 cell model. To date most of the data pertaining to cellular control of renin by ET has been obtained by using primary JG cultures. The current study is the first to perform a detailed investigation of the effects of ET-1 on As4.1 and the subsequent regulation of renin. The As4.1 cell has proven to be a particularly useful in vitro model for JG cells. It was derived by transgene-targeted oncogenesis of renin-expressing cells in the mouse kidney and therefore expresses and secretes high levels of the endogenous mouse renin gene over many months in culture. Morphologically, the As4.1 cell is similar to bona fide JG cells in vivo because it contains large renin-positive secretory granules. In addition to the transcriptional regulation of renin that has been elucidated by using the As4.1 cell line, physiological studies of renin regulation have been performed. For example, cAMP analogs cause an increase in renin secretion from As4.1 cells (28) and interleukin-1beta (17), and more recently, mechanical strain (21, 22) has been shown to regulate the renin gene in a manner consistent with what would be expected in vivo.

In conclusion, the current study outlines a cellular pathway for the regulation of renin by ET-1. Initiation of this signaling pathway likely leads to activation of other downstream targets in the nucleus that result in a decrease in renin gene expression over hours and days. These studies suggest that vascular endothelium in the preglomerular artery may play a crucial role in the regulation of renin in vivo because ET-1 and NO, both products of vascular endothelium, are involved in the regulation of renin through receptor interactions, second messengers, and transcriptional regulation. Interestingly, the cellular and molecular pathways activated by ET-1 are similar to those activated by mechanical strain in the As4.1 cell line thus demonstrating the complexity of renin gene regulation by various physiological stimuli. Further elucidating the downstream targets of the signaling pathways activated by ET-1 in As4.1 cells and how this leads to the regulation of the renin gene remains to be elucidated.


    ACKNOWLEDGEMENTS

We extend our sincere gratitude to Maureen Adolf for technical expertise with the inositol phosphate measurements. We thank Dr. Curt Sigmund (University of Iowa) for providing the DNA constructs used for transcription experiments.


    FOOTNOTES

This work was supported in part by National Heart, Lung, and Blood Institute Grants HL-49405 (to G. Hajduczok), HL-48459 (to K. W. Gross), and American Heart Association Grant 92-310G (to G. Hajduczok). This research also utilized core facilities supported in part by Roswell Park Cancer Institute's National Cancer Institute-funded Cancer Center Support Grant (CA16056). M. J. Ryan was partially supported by a Mark Diamond predoctoral grant.

Current address of S. L. Millard: Dept. of Pediatrics and Human Development, DeVos Children's Hospital, 330 Barclay Ave. NE, Suite 200, Grand Rapids, MI 49503.

Address for reprint requests and other correspondence: M. J. Ryan, Univ. of Iowa, Dept. of Internal Medicine, 3181 MERF, Iowa City, IA 52242 (E-mail: ryanm{at}physiology.uiowa.edu).

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.

August 29, 2002;10.1152/ajpheart.00295.2002

Received 3 April 2002; accepted in final form 26 August 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

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Am J Physiol Heart Circ Physiol 283(6):H2458-H2465
0363-6135/02 $5.00 Copyright © 2002 the American Physiological Society


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