Am J Physiol Heart Circ Physiol 283: H2458–H2465, 2002.
First published August 29, 2002; 10.1152/ajpheart.00295.2002.
Endothelin-1 increases calcium and attenuates renin
gene expression in As4.1 cells
MICHAEL J. RYAN,1 THOMAS A. BLACK,2 SUSAN L. MILLARD,3
KENNETH W. GROSS,2 AND GEORGE HAJDUCZOK1
1
Department of Physiology and Biophysics and 3Pharmacology and Toxicology,
State University of New York at Buffalo, Buffalo 14214; and 2Department of Molecular
and Cellular Biology, Roswell Park Cancer Institute, Buffalo, New York 14263
Received 3 April 2002; accepted in final form 26 August 2002
Ryan, Michael J., Thomas A. Black, Susan L. Millard,
Kenneth W. Gross, and George Hajduczok. Endothelin-1
increases calcium and attenuates renin gene expression in
As4.1 cells. Am J Physiol Heart Circ Physiol 283:
H2458–H2465, 2002. First published August 29, 2002;
10.1152/ajpheart.00295.2002.—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 reninproducing 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-bromocGMP, 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
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,
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: [email protected]).
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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
Cell culture. As4.1 cells (ATCC No. CRL2193) are a reninexpressing 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
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HEPES, 20 glucose, and 1 mM EGTA), followed by an incubation with 5 ␮M 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 ␮l.
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 ␮M BQ-123 (ET receptor
antagonist). The NO donor sodium nitroprusside (SNP, 500
␮M) and the membrane-permeable analog of cGMP, 8-bromocGMP (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
␮Ci/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.
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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 ␮g/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 ␮g. 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 ␤-galactosidase (RSV␤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.
␤-Galactosidase. ␤-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
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
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(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
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.
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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 voltageactivated Ca2⫹ channels in the plasma membrane.
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 ␮M), 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.
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 ␮M), 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 ␮M) (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
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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.
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
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-3stimulated 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 ␮M) inhibited ET-1 calcium stimulation of
As4.1 cells. Cells were superfused with normal Ringer solution
containing BQ-123 (1 ␮M) 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).
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 ␮M; 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 ␮M) 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).
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
␮g/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.
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
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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).
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-1␤ (17)
and mechanical force (21). Therefore, we used the lu-
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.
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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. ␤-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.
DISCUSSION
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-␮g samples of RNA quantified by Northern blot analysis. GAPDH expression was used to correct for variations in gel loading. Data are means ⫾ SE.
ciferase 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
Fig. 8. ET-1 does not alter the rate of renin message decay. As4.1
cells were incubated with 10 ␮g/ml actinomycin D and 10 nM ET-1.
Northern blot analysis of 2-␮g 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).
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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
Fig. 9. ET-1 inhibits renin gene transcription in As4.1 cells. Construct a (promoterless vector, PGL2 Basic) and construct b (promoterluciferase) 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).
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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 experAJP-Heart Circ Physiol • VOL
iments 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 transgenetargeted 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-1␤ (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
283 • DECEMBER 2002 •
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RENIN INHIBITION BY ENDOTHELIN
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.
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.
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 Institutefunded 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.
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