Department of Medicine, Ottawa Hospital, and the Kidney Research Centre, Ottawa Health Research Institute, University of Ottawa, Ottawa, Ontario, Canada K1H 8L6
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ABSTRACT |
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Angiotensin II (ANG II) subtype 2 (AT2) receptors are expressed in the adult kidney, but the
effects of AT2 receptor activation are unclear. The
proximal tubule cell line LLC-PK1 was transfected with a
plasmid containing cDNA for the rat AT2 receptor. In
transfected cells, specific binding of 125I-labeled ANG II
was detected (dissociation constant = 0.81 nM), with inhibition by
the AT2 antagonist PD-123319, and no effect of the
AT1 antagonist losartan. ANG II (107 M)
significantly inhibited mitogen-activated protein kinase (MAPK) activity in transfected cells, associated with decreased
phosphorylation of the extracellular signal-related kinases ERK1 and
ERK2. ANG II stimulated phosphotyrosine phosphatase activity within 5 min, an effect blocked by PD-123319 and the phosphatase inhibitor
vanadate. In transfected cells, ANG II inhibited epidermal growth
factor-stimulated [3H]thymidine incorporation, an effect
reversed by vanadate. In contrast, vanadate did not block ANG
II-stimulated apoptosis of transfected cells. In summary,
AT2 receptors in proximal tubule cells inhibit MAPK
activity and stimulate phosphotyrosine phosphatase. AT2
receptor-induced inhibition of mitogenesis is mediated by phosphatase
activation, whereas effects on apoptosis are insensitive to
phosphatase inhibition. The data suggest that AT2 receptors inhibit cell growth via distinct signaling pathways in the proximal tubule.
mitogen-activated protein kinase; tyrosine phosphatase; apoptosis; LLC-PK1; angiotensin II subtype 2 receptor
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INTRODUCTION |
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ANGIOTENSIN II (ANG II) exerts a wide variety of effects in the kidney, including arteriolar vasoconstriction, mesangial cell contraction, elaboration of extracellular matrix proteins, stimulation of proximal tubule sodium and bicarbonate transport, and tubular cell hypertrophy or hyperplasia, among others. These well-described effects occur via interaction with G protein-coupled angiotensin subtype 1 (AT1) receptors, present in smooth muscle cells and in cells of virtually all nephron segments (9). The cDNA for the angiotensin subtype 2 (AT2) receptor was cloned in 1993 and encodes a seven transmembrane-domain protein of 363 amino acids, with 34% homology to the AT1 receptor (11, 20). Expression of the intrarenal AT2 receptor occurs at its highest levels in the fetal kidney, with a rapid decline in expression after birth (26). Recent studies, however, have immunohistochemically localized AT2 receptors to the adult kidney, within glomeruli, tubules, and interstitial cells (23).
The precise physiological functions of intrarenal AT2 receptors have been difficult to demonstrate, at least partly because AT2 receptors appear to be expressed at low levels compared with AT1 receptors. In the postischemic rat kidney, we have demonstrated an upregulation of AT2 receptor mRNA in proximal tubule (12), and increased AT2 receptor expression has been reported after injury in other tissues (22, 36), suggesting a potential role for these receptors in modulating repair responses. In nonrenal cells, activation of AT2 receptors inhibits cell growth (16, 21) or induces apoptosis (34, 35). Indeed, in a model of rat obstructive uropathy, AT2 receptors appear to inhibit development of interstitial fibrosis, suggesting that in renal interstitial cells, these receptors may decrease formation of extracellular matrix proteins such as collagen IV (18). Furthermore, intrarenal AT2 receptors may stimulate natriuresis. In animal studies, AT2 receptors protect against the hypertensive and sodium-retaining properties of AT1 receptor activation, an effect caused by AT2-mediated production of intrarenal bradykinin and nitric oxide (NO), with resultant increased generation of cGMP (27-29).
In the present study, we focused on the potential growth regulatory properties of AT2 receptors in cells of the renal proximal tubule, a nephron segment that is particularly susceptible to ischemic and toxic injury and which exhibits activation of growth pathways during the repair phase. To do this, we created a proximal tubule cell line that selectively and stably expresses ANG II AT2 receptors. In these cells, we determined that coupling of ANG II to AT2 receptors inhibits mitogen-activated protein kinase (MAPK) activity and stimulates phosphotyrosine phosphatase, associated with impaired mitogenic responses to epidermal growth factor (EGF). ANG II also increased apoptosis via a phosphatase-insensitive pathway. The data indicate that AT2 receptors negatively modulate renal epithelial cell growth.
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METHODS |
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Cell culture and transfection. To create a permanent cell line expressing ANG II AT2 receptors, cDNA encoding the rat AT2 receptor was cloned, and cells of the porcine proximal tubule-like cell line LLC-PK1 (American Type Culture Collection, Rockville, MD) were transfected with an expression plasmid containing the AT2 receptor cDNA. Briefly, cDNA encoding a 1,163-bp fragment of the rat AT2 receptor containing the entire open reading frame (11, 20) was isolated from 0.5 µg of rat kidney cortex total RNA by RT-PCR, using 100 pmol of a forward primer [5'-AGGATTGGGAGTCTCTGACAGTTC-3', corresponding to 113-136 bp of the rat AT2 receptor cDNA (11)] and 100 pmoles of a reverse primer (5'-CTCCAAACCATTGCTAGGCTGATTAC-3', corresponding to 1251-1276 bp). After reverse transcription, and after 5 min of denaturation at 94°C, PCR was carried out for 40 cycles of 94°C for 30 s, 63°C for 30 s, and 72°C for 45 s, followed by extension at 72°C for 10 min. The 1163-bp cDNA product was then reamplified by PCR (30 cycles), using a phosphorylated form of the same 5' forward primer (generated with T4 polynucleotide kinase; Invitrogen, Carlsbad, CA) and the unphosphorylated reverse primer. After quantification on an ethidium bromide-stained 2% agarose gel, the 5'-phosphorylated 1,163-bp cDNA product was purified (Geneclean Kit, Bio/Can Scientific, Mississauga, ON, Canada) and ligated at 14°C for 16 h into the linearized plasmid vector pCR 3.1-Uni, which contains the neomycin resistance gene (Invitrogen), using T4 DNA ligase. Use of the 5' phosphorylation method of PCR ensured that the cDNA product was inserted unidirectionally into the vector pCR 3.1-Uni, in the correct orientation for expression of the AT2 receptor.
After ligation, Escherichia coli bacterial cells (TOP-10F, Invitrogen) were transformed with the plasmid ligation product for large-scale preparation of plasmid DNA (Qiagen maxi kit, Qiagen, Chatsworth, CA). PCR on DNA isolates documented the presence of the desired 1163-bp product, and correct orientation was confirmed by restriction enzyme digestion with Apa1, which generated the expected fragments of 971 and 5,236 bp. LLC-PK1 cells were grown to 90% confluence in Dulbecco's modified Eagle's medium nutrient mixture-Ham's F-12 (DMEM/F-12), supplemented with 10% fetal bovine serum, penicillin (100 U/ml), and streptomycin (100 µg/ml), at 37°C in a humidified atmosphere of 95% room air-5% CO2. Transfection was performed by calcium phosphate precipitation, essentially as we have described (4), with 20 µg of rat AT2 receptor expression plasmid added to each plate of cells. Control cells underwent mock transfection with sterile water. Forty-eight hours after transfection, the antibiotic G-418 (GIBCO, Burlington, ON, Canada) was added to the cells (500 µg/ml) to select for stable transfectants. Within 14 days, no mock-transfected cells survived in the presence of G-418. In contrast, cells transfected with the AT2 expression plasmid reached ~50% confluence at 14 days. These cells were trypsinized, plated into 75-cm2 cell culture flasks, and used for further studies. Clonal selection of cells was not performed, as initial studies demonstrated significant binding of 125I-labeled ANG II to AT2 receptors in the transfected cells. For all experiments, cells were grown in DMEM/F-12 with 10% fetal bovine serum, penicillin/streptomycin, and G-418 and used at passages 1-10. Preliminary experiments revealed that wild-type LLC-PK1 cells demonstrated no significant specific binding of 125I-ANG II, indicating the absence of AT1 or AT2 receptors. In transfected LLC-PK1 cells, mRNA for AT2 receptors was readily detected by RT-PCR (not shown). In DNA isolated from transfected LLC-PK1 cells, DNA sequencing (University of Ottawa DNA Sequencing Facility) confirmed the presence of the 1163-bp AT2 receptor cDNA, in the correct orientation.125I-ANG II binding.
Binding of 125I-ANG II was determined in transfected cells
grown in 24-well plastic culture dishes. Cells were incubated in PBS with 0.5% albumin (BSA, Sigma, St. Louis, MO), 1 mM EDTA, 0.25 mM
phenanthroline, supplemented with 125I-ANG II (0.1-8.0
nM, 2,000 Ci/mmol, Amersham, Oakville, ON) with or without unlabeled
ANG II (106-10
12 M), losartan
(10
6-10
8 M) or the AT2
receptor antagonist PD-123319 (10
6-10
12 M)
for 1 h at 37°C. Incubation buffer was then rapidly removed, and
cells were washed four times with ice-cold PBS with 0.5% BSA and
solubilized in 0.25 N NaOH-0.1% SDS, and cell-associated radioactivity was measured in a gamma counter. Nonspecific binding was determined in
the presence of 10
6 M unlabeled ANG II. Scatchard
analysis of AT2 receptors was performed using the Prism
graphics software program.
Measurement of cAMP and cGMP.
Cells were grown to confluence on 24-well dishes and then incubated for
16 h in serum-free DMEM/F-12 medium. For assays of cAMP or cGMP,
cells were incubated at 37°C for 15 or 60 min in DMEM/F-12,
supplemented with IBMX (5 × 104 M), 0.5% BSA, in
the presence or absence of ANG II or other agonists. Medium was then
aspirated and replaced with ice-cold 10% TCA (vol/vol). After a
further 30 min, samples were extracted four times with four volumes of
water-saturated ether and brought to pH 7 with Tris. Aliquots were
assayed for cAMP or cGMP, using radioligand competitive binding
assay kits containing [3H]cAMP (Intermedico, Markham, ON)
or [3H]cGMP (Amersham), as we have performed (4,
25).
Measurement of cytosolic calcium concentration. Cytosolic calcium concentration ([Ca2+]i) was measured in transfected LLC-PK1 cells, essentially as we have described (3). Cells were grown to confluence on glass coverslips and loaded with 5 µM of fura 2-acetoxymethyl ester (fura 2-AM) in the presence of 0.005% Pluronic-F127 for 45 min at room temperature. Cells were then continuously perfused at 37°C with a solution consisting of (in mM) 105 NaCl, 24 NaHCO3, 2 Na2HPO4, 5 KCl, 1.0 MgSO4, 1.5 CaCl2, 4 lactic acid, 5 glucose, 1 alanine, 10 HEPES (pH 7.3), and 0.2% BSA, and agonists were added at various times. Fluorescence was measured from dual monochromators set at 340 and 380 nm, using a computer-linked analytical system (Photon Technology International, South Brunswick, NJ). Fluorescence emission was measured by photon counting, and the corrected fluorescence emission intensity ratio, from 340- and 380-nm excitation, was monitored continuously in selected cells and used as an indicator of [Ca2+]i.
Measurement of p42/p44 MAPK activity.
MAPK activity was determined in transfected cells, using an assay that
measures phosphorylation of a synthetic peptide substrate containing
the Thr669 phosphorylation site of the EGF receptor
(Amersham). Cells were grown to confluence on 24-well plastic dishes,
then rendered quiescent in serum-free media for 16 h. Cells were
incubated in the presence or absence of ANG II for 5 min at 37°C, the
medium was aspirated, and cells were lysed in a buffer consisting of 10 mM Tris, 150 mM NaCl, 2 mM EGTA, 2 mM dithiothreitol (DTT), 1 mM
orthovanadate, 1 mM phenylmethylsulfonyl fluoride, 10 µg/ml
leupeptin, and 10 µg/ml aprotinin, pH 7.4, at 4°C (buffer
A). Cellular debris was removed by centrifugation at 15,000 g for 20 min. Cell supernatants were then incubated with
peptide substrate, reaction buffer, and a magnesium-ATP buffer
containing 200 µCi/ml [-32P]ATP (Amersham). The
reaction was terminated after 30 min, and aliquots of the reaction
mixture were spotted onto binding paper discs. After being washed two
times with 75 mM phosphoric acid and three times with distilled water,
bound phosphorylated peptide was measured by scintillation
spectrometry. In each experiment, blanks were performed in the absence
of peptide substrate. Cell proteins were quantified by Bradford assay
(Bio-Rad, Montreal, QC, Canada). Results are expressed as
femtomoles of phosphate transferred per microgram protein per minute.
Western blotting of extracellular signal-related kinases 1 and 2. Phosphorylation of extracellular signal-related kinases ERK1 (p44) and ERK2 (p42) was measured by Western blotting. After incubation with or without ANG II, cells were lysed in buffer A. After quantification of proteins, equal amounts of protein lysates (3 µg) were run on 10% SDS-polyacrylamide gels and transferred to nitrocellulose membranes (Bio-Rad). The membranes were blocked with 10% skim milk in Tris-buffered saline (pH 7.6) containing 0.1% Tween 20 (TBS-T) for 1 h at room temperature. The membranes were then incubated for 16 h at 4°C with a 1:1,000 dilution of polyclonal antibody to phosphorylated rat ERK1 and ERK2 (Santa Cruz Biotechnology, Santa Cruz, CA), followed by incubation with a 1:2,000 dilution of anti-rat secondary antibody conjugated to horseradish peroxidase (Amersham). After washing of membranes, phosphorylated proteins were detected by enhanced chemiluminescence (ECL, Amersham) on Hyperfilm (Amersham). Prestained standards were used as molecular weight markers (Bio-Rad), and, to control for protein loading, all membranes were stripped and reprobed with polyclonal antibody to unphosphorylated rat ERK1 and ERK2 (Santa Cruz Biotechnology). Signals on Western blots were quantified by densitometry and corrected for unphosphorylated ERK1 and ERK2 levels, using an image-analysis software program (Kodak Densitometer 1S440CF).
Measurement of cytosolic phosphotyrosine phosphatase activity.
Cytosolic phosphotyrosine phosphatase activity was measured in cells
treated for 5 or 30 min with or without ANG II, by measuring the
dephosphorylation of myelin basic protein (Sigma), previously phosphorylated by protein tyrosine kinase,
p60c-src (Oncogene, Cambridge, MA), essentially
as described (7). Equal amounts of cell lysate proteins
were incubated with myelin basic protein that had been phosphorylated
by p60c-src in the presence of
[-32P]ATP (10 mCi/ml, Amersham) in a solution
containing 5 mM EDTA, 25 mM HEPES (pH 7.3), and 10 mM DTT, for 15 min
at 37°C. The reaction was terminated by addition of ice-cold
charcoal, followed by centrifugation at 15,000 g for 5 min.
The supernatants, containing liberated 32P, were removed
and counted by scintillation spectrometry. Phosphate release was
expressed as femtomoles of phosphate released per microgram protein.
Measurement of DNA synthesis.
DNA synthesis was assayed by measurement of [3H]thymidine
incorporation, as we have previously done (4). After 48-h
quiescence, transfected cells were exposed to agonists [EGF,
108 M], with or without ANG II
[10
7-10
10 M], and/or the phosphatase
inhibitor sodium orthovanadate (vanadate; 10
6 M, Sigma)
for 16 h in serum-free medium, followed by labeling for 2 h
with 2 µCi/ml of [3H]thymidine (Amersham). The cells
were then washed five times with ice-cold PBS plus 0.5% BSA and lysed
in 0.25 N NaOH-0.1% SDS, and DNA-incorporated thymidine was counted by
scintillation spectrometry.
Assay of apoptosis.
Apoptosis was measured in cells by in situ terminal
deoxynucleotidyl transferase nick-end labeling (TUNEL) assay, using a commercial kit (Trevigen, Gaithersburg, MD). Cells grown on 12-mm glass
coverslips and rendered quiescent for 48 h were incubated in
serum-free DMEM/F-12 for an additional 48 h, with or without ANG
II (107 M), or with ANG II plus vanadate
(10
6 M). After fixation of cells in 3.7% formaldehyde
and treatment with 20 µg/ml proteinase K and 2%
H2O2, TUNEL staining was performed using
terminal deoxynucleotidyl transferase, with detection by incubation
with streptavidin-horseradish peroxidase (Amersham) and
3,3'-diaminobenzidine chromogen solution (ESBE Scientific, Montreal,
QC, Canada). Cells were counterstained with methyl green, and
TUNEL-positive cells were counted on coverslips mounted on microscope
slides, using a Zeiss Axioplan microscope. For quantitation of
apoptosis, the total number of cells in 10 separate microscopic fields at ×200 magnification was counted, with the viewer blinded to
the origin of the slides.
Data analysis. Results are presented as means ± SE of experiments performed in duplicate. Significance was determined by Student's t-test or by ANOVA for cases with multiple comparisons. Significance is considered as P < 0.05.
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RESULTS |
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125I-ANG II binding studies.
In LLC-PK1 cells transfected with the AT2
receptor pCR3.1-Uni expression plasmid, specific binding of
125I-ANG II was demonstrated, with concentration-dependent
inhibition by unlabeled ANG II or the AT2 receptor
antagonist PD-123319. The AT1 receptor antagonist losartan
(106-10
8 M) had no significant effect on
125I-ANG II binding in these cells (Fig.
1A). Scatchard analysis revealed the presence of high-affinity ANG II receptors, with a
receptor dissociation constant of 0.81 nM and a maximum number of
binding sites (Bmax) of 4.80 fmol/mg (Fig.
1B). These studies confirmed the expression of rat
AT2 receptor protein on the plasma membranes of transfected
cells, with binding characteristics similar to those previously
reported (11, 19, 20).
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Effect of ANG II on cAMP, cGMP, and
[Ca2+]i.
In transfected cells, administration of ANG II
(107-10
11 M) for 15 or 60 min had no
significant effect on cellular levels of cAMP or cGMP
(n = 3-5; Fig. 2).
Similarly, although arginine vasopressin caused significant increases
in calcium concentration in fura 2-AM-loaded transfected cells, ANG II
(10
7 M) had no effect on cell calcium concentration
(n = 5; Fig. 3).
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MAPK activity, phosphorylation of ERK1 and ERK2, and
phosphotyrosine phosphatase activity.
Activation of MAPK is associated with stimulation of downstream cell
growth responses, including mitogenesis, and AT2 receptor activation has been reported to inhibit MAPK activity in other cell
types (2). In transfected LLC-PK1 cells,
incubation with ANG II (107 M) for 5 min caused a
significant inhibition of MAPK activity, an effect reversed by
PD-123319 {Fig. 4, control 3,475 ± 435 vs. ANG II 2,513 ± 196 fmol of phosphate
transferred · µg
protein
1 · min
1
(P < 0.001, n = 7) vs. ANG II + PD-123319 3,342 ± 256 fmol of phosphate
transferred · µg
protein
1 · min
1 [P = not significant (NS) vs. control, n = 7]}. Lower
concentrations of ANG II did not induce significant inhibition of MAPK
activity [ANG II 10
9 M: 8.2 ± 3.9% inhibition,
P = NS vs. control (n = 4)].
Incubation of cells with PD-123319 alone had no effect on MAPK
activity. Inhibition of MAPK activity was associated with a significant reduction in phosphorylation of the p44 and p42 kinases, ERK1 and ERK2,
respectively, determined by Western blot analysis, with no effect
on expression of the unphosphorylated enzymes [Fig. 5 (n = 8)].
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Effect of ANG II on EGF-stimulated mitogenesis.
Activation of phosphotyrosine phosphatase and inhibition of MAPK
activity by ANG II in transfected cells suggested that AT2 receptors might induce altered growth responses. In serum-starved transfected cells, ANG II caused a modest but significant inhibition of
EGF-stimulated DNA synthesis (Fig.
7A). As an index of the biological relevance of this inhibitory response, a similar degree of
inhibition of EGF-stimulated mitogenesis was also observed by
preincubation of cells with the tyrosine kinase inhibitor genistein (5 × 106 µM) [19.4 ± 0.9% inhibition
(n = 3)]. Preincubation of cells with vanadate
(10
6 M) blocked the inhibitory effect of ANG II on
EGF-stimulated mitogenesis [Fig. 7B, EGF 418 ± 21.8 dpm/µg vs. EGF + ANG II (10
7 M) 324 ± 22.1 dpm/µg, P < 0.01 vs. EGF (n = 5),
vs. EGF + ANG II (10
7 M) + vanadate 379 ± 19.5 dpm/µg, P = NS vs. EGF (n = 5)].
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Effect of ANG II on apoptosis.
To determine if AT2 receptors are linked to
apoptosis in these cells, cells were serum deprived for 48 h and treated with ANG II (107 M) for an additional
48 h, followed by TUNEL assay. In the absence of ANG II, there was
no significant difference in the numbers of apoptotic cells in
wild-type vs. transfected cells (not shown). In transfected cells, ANG
II significantly increased the numbers of apoptotic cells [Fig.
8, control 25.5 ± 5.6 vs. ANG II
40.2 ± 4.2 apoptotic cells/10 fields; P < 0.025 (n = 8)]. However, preincubation of cells with
vanadate (10
6 M) had no effect on ANG II-stimulated
apoptosis [Fig. 8, ANG II + vanadate 37.6 ± 3.9 apoptotic cells/10 fields; P = NS vs. ANG II
(n = 8)].
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DISCUSSION |
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ANG II AT1 receptors are expressed in abundance in the
adult kidney, in virtually all nephron segments (9), and
mediate the well-known intrarenal actions of ANG II, including
vasoconstriction, enhanced tubular sodium reabsorption, and stimulation
of cell growth (1). Recent studies reveal, however, that
although ANG II AT2 receptor mRNA is of low abundance in
the adult kidney (26), AT2 receptor protein is
expressed within glomeruli, tubules, and interstitial cells
(23). To examine signaling and growth responses mediated
by AT2 receptors in the proximal tubule, in the present studies we transfected LLC-PK1 cells with an expression
plasmid encoding the rat AT2 receptor. The cells were
selected by their ability to grow in the presence of the antibiotic
G-418, and our results indicate reproducible expression of
AT2 receptor protein up to at least 10 cell passages, with
no presence of AT1 receptors. In these renal epithelial
cells, activation of AT2 receptors by ANG II
(107 M) stimulates a phosphotyrosine phosphatase,
associated with inhibition of MAPK activity and dephosphorylation of
ERK1 and ERK2. Because we did not subclone cells in this study, it is
possible that some cells in the transfected population express higher
levels of surface AT2 receptors, and therefore may respond
similarly to even lower concentrations of ANG II. Furthermore, ANG II
inhibits EGF-stimulated DNA synthesis in these cells, in a
vanadate-sensitive fashion, whereas ANG II-stimulated apoptosis
is not blocked by vanadate. This suggests that AT2
receptor-induced phosphatase activation is involved in inhibition of
mitogenic responses, but not in mediation of apoptosis.
Mice with targeted deletion of the AT2 receptor gene demonstrate elevated blood pressures compared with wild-type mice and have an enhanced hypertensive response to ANG II infusions associated with decreased urinary sodium excretion (6, 30). This suggests that AT2 receptors are linked to natriuresis. On the basis of a number of elegant studies by Siragy and Carey (27-29), it has been postulated that intrarenal AT2 receptors mediate vasodilatation and natriuresis through stimulation of intrarenal bradykinin production, which increases local NO release and cGMP levels. In the present studies, we observed no significant effect of AT2 receptor activation on cellular levels of cAMP, cGMP, or calcium. In the proximal tubule, increases in cAMP are associated with inhibition of apical Na+/H+ exchange activity (5, 33), and cytosolic calcium may regulate the antiporter via effects on calcium/calmodulin-dependent kinase activity (32). Our results suggest that AT2 receptors in proximal tubule may not be linked to direct stimulation of guanylate cyclase and cGMP formation, which has been reported to cause inhibition of tubular sodium transport (25).
In contrast to these negative data, activation of AT2 receptors in transfected cells caused significant inhibition of MAPK activity and blocked ERK1 (p44) and ERK2 (p42) phosphorylation. This was associated with stimulation of a vanadate-sensitive phosphotyrosine phosphatase activity. These results contrast with those of Haithcock et al. (8), who reported that AT2 receptor-stimulated arachidonic acid release in renal epithelial cells mediated activation of ERK1 and ERK2. It is difficult to explain these differences in data, although it must be noted that our transfected LLC-PK1 cells express AT2 receptors but not AT1 receptors, and so any potential involvement of AT1-mediated pathways is excluded in these cells. Furthermore, our data are in agreement with studies on AT2 receptor signaling in pheochromocytoma (PC12W) and neuroblastoma cells, which demonstrated AT2-mediated inhibition of MAPK and activation of SHP-1 tyrosine phosphatase, which leads to dephosphorylation of the anti-apoptotic protein Bcl-2 (2, 10).
In transfected cells, ANG II inhibited EGF-stimulated [3H]thymidine incorporation and caused a stimulation of apoptosis. An antiproliferative effect of AT2 receptor activation has also been reported in cultured coronary endothelial cells (31), and AT2 receptors induce apoptosis in cultured PC12W cells and mouse fibroblasts (35). Few studies have focused on the effects of AT2 receptor activation on renal cell growth responses. In this regard, AT2 receptors inhibit growth of renal medullary interstitial cells in culture (15), and, in rats with unilateral ureteral obstruction, treatment with the AT2 antagonist PD-123319 inhibits apoptosis of tubular cells and increases interstitial collagen IV formation in the obstructed kidney (18). In mice with targeted deletion of the AT2 receptor gene, ureteral obstruction is associated with increased interstitial fibrosis, an abundance of interstitial fibroblasts, and decreased tubulointerstitial cell apoptosis compared with wild-type mice (14). Taken together, these data are consistent with the results of our study, and support a growth inhibitory effect of AT2 receptors in renal tubular cells.
A study by Miura and Karnik (17) revealed that expression of AT2 receptors in fibroblasts, Chinese hamster ovary cells, and vascular smooth muscle cells (VSMC) stimulates apoptosis, even in the absence of ANG II, suggesting that ligand-independent receptor activation of apoptosis-signaling pathways occurs. In our study, we observed no significant difference in apoptosis between wild-type and AT2 receptor-transfected cells in the absence of ANG II, indicating that binding of ANG II to the AT2 receptor is required to induce apoptosis in these cells. Of interest, however, is that the stimulation of apoptosis by ANG II was not reversed by vanadate, suggesting that activation of phosphotyrosine phosphatase does not mediate apoptosis in these cells. This contrasts with studies in PC12W cells, where AT2 receptor-stimulated phosphatase activity is proposed to induce apoptosis via dephosphorylation of Bcl-2 and production of ceramides (13). The AT2 receptor-mediated signaling pathways for apoptosis in LLC-PK1 cells will require further study, and in this regard p38 MAPK activation has been implicated in AT2 receptor promotion of apoptosis in embryonic VSMC, in the absence of ligand (17).
The coexistence of AT1 and AT2 receptors in proximal tubules in vivo (23, 24) suggests that growth responses to ANG II will be affected by the relative abundance of each receptor class. Upregulation of intrarenal AT2 receptors has been reported with sodium depletion (23), and we have previously demonstrated increased proximal tubular AT2 receptor mRNA expression after renal ischemia/reperfusion injury (12). Enhanced AT2 receptor expression has also been demonstrated in the heart after myocardial ischemia (22) and in ischemic brain (36). Our results suggest the possibility that enhanced expression of tubular AT2 receptors after injury may inhibit tubular cell mitogenesis in response to endogenous growth factors such as EGF.
In summary, we have created a renal proximal tubule cell line that stably expresses AT2 receptors. In these cells, ANG II inhibits MAPK activity and phosphorylation, and activates a vanadate-sensitive phosphotyrosine phosphatase, associated with inhibition of EGF-stimulated mitogenesis. This effect, along with the AT2 receptor-dependent stimulation of apoptosis, is in contrast to the well-known growth stimulatory responses mediated by AT1 receptors. The establishment of this cell line will provide a useful model to study further the effects of AT2 receptors on tubular epithelial cell function.
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ACKNOWLEDGEMENTS |
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This work was supported by a grant from the Kidney Foundation of Canada to K. D. Burns. Parts of this work were presented in abstract form at the annual meeting of the American Society of Nephrology, Miami, FL, in November 1999, and have appeared in abstract form (J Am Soc Nephrol 10: A2456, 1999).
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FOOTNOTES |
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Address for reprint requests and other correspondence: K. D. Burns, FRCPC, Associate Professor and Head, Div. of Nephrology, The Ottawa Hospital and Univ. of Ottawa, 501 Smyth Rd., Rm. LM-18, Ottawa, Ontario, Canada K1H 8L6 (E-mail: kburns{at}ottawahospital.on.ca).
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.
Received 18 September 2000; accepted in final form 6 April 2001.
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