2 Division of Nephrology/Hypertension, University of Berne, 3010 Berne, Switzerland; and 1 Medizinische Klinik IV, University of Erlangen-Nuremberg, Erlangen 8520, Germany
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ABSTRACT |
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Angiotensin II (ANG
II) and nitric oxide (NO) have contrasting vascular effects, yet both
sustain inflammatory responses. We investigated the impact of ANG II on
lipopolysaccharide (LPS)/interferon- (IFN)-induced NO production in
cultured rat mesangial cells (MCs). LPS/IFN-induced nitrite production,
the inducible form of nitric oxide synthase (NOS-2) mRNA, and protein
expression were dose dependently inhibited by ANG II on coincubation,
which was abolished on ANG II type 2 (AT2) receptor
blockade by PD-123319. Homology-based RT-PCR verified the presence of
AT1A, AT1B, and AT2 receptors. To
shift the AT receptor expression toward the type 1 receptor, two sets
of experiments were performed: LPS/IFN preincubation for 24 h was
followed by 8-h coincubation with ANG II; or during 24-h coincubation
of LPS/IFN and ANG II, dexamethasone was added for the last 6-h period.
Both led to an amplified overall expression of NOS-2 protein and NO
production that was inhibitable by actinomycin D in the first setup.
Induced NO production was enhanced via the AT1 receptor;
however, it was diminished via the AT2 receptor. In
conclusion, induced NO production is negatively controlled by the
AT2, whereas AT1 receptor stimulation enhanced
NO synthesis in MCs. The overall NO availability depended on the onset
of the inflammatory stimuli with respect to ANG II exposure and the
available AT receptors.
inducible nitric oxide synthase; glomerular inflammation; angiotensin II type 1 and type 2 receptor; dexamethasone
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INTRODUCTION |
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IMMUNE COMPLEX FORMATION within glomeruli initiates chemical and cellular responses mediating inflammatory signals, such as NO.
NO is a radical gas and signaling molecule. Its production is catalyzed by NO synthase (NOS) isoforms converting L-arginine and molecular oxygen to L-citrulline and NO. If adequately stimulated by endotoxin or cytokine, mesangial cells (MCs) express the inducible NOS isoform (NOS-2) (35). Nanomolar concentrations of NO generated by NOS-2 in MCs are sufficient to sustain inflammatory responses, influence glomerular perfusion (3, 45), or even evoke substantial cell damage leading to apoptosis or cytotoxic cell death (42, 43).
Increased NO production is part of the inflammatory glomerular response of various glomerulonephritis models (4, 26, 37) and has been localized to MCs (5). This is further supported by an amelioration of experimental glomerulonephritis observed with inhibition of NO synthesis (49).
ANG II leads to renal vasoconstriction, regulating glomerular perfusion
as well as filtration (25), and directly influences the
mesangial constrictive response (44). ANG II is a potent regulator of MC phenotypical changes, such as migration
(28), proliferation (19), or protein kinase C
(PKC) expression (1). This supports its role as a growth
factor, profibrogenic and proinflammatory mediator (50),
and regulator of additional proinflammatory mediators, such as
transforming growth factor- (TGF-
) (54) or
monocyte-chemoattractant protein-1 (51). The effects of
ANG II are transmitted via at least three different receptor subtypes
in rats, the AT1A, the AT1B, and the
AT2 receptors, with very diverse downstream signaling mechanisms and temporospatial distributions (reviewed in Ref. 23).
Animal models of experimental glomerulonephritis exploiting ANG I-converting enzyme inhibitors and ANG II receptor blockers support the concept of ANG II being an important proinflammatory mediator via the AT1 receptor (36, 41, 54).
MCs present an important regulatory element for vascular conductance as well as the ultrafiltration surface area within the glomerulus. They are critically controlled by vasodilators as well as vasoconstrictors. During periods of glomerular inflammation, ANG II- and NO-related phenotypical changes of MCs could determine glomerular function as well as inflammatory responses. ANG II and NO have been postulated to interact closely, at least in the regulation of renal hemodynamics (14, 32).
Conflicting results have been published regarding the influence of ANG II on induced NO production in conditions of sepsis or in glomerulonephritis. Some authors report a downregulation or inhibition of the induced NO production in cultured rat astroglial cells by certain proinflammatory stimuli (29) and similarly in renal cells such as mouse proximal tubular cells (52) and rat MCs (27, 40). Others were able to demonstrate an augmented lipopolysaccharide (LPS)- or cytokine-induced NO production in rat cardiac myocytes (21, 53).
The aim of the present study was to examine the influence of ANG II on
LPS/interferon- (IFN)-induced NO production in cultured rat MCs. We
were able to demonstrate that the resulting NO response to ANG II
depended on the activation of discrete ANG II receptor subtypes. The
AT1 receptor subtypes transferred stimulation of NO
production, whereas the AT2 receptor subtype inhibited
induced NO production. The overall NO availability depended on
the onset of the inflammatory stimuli with respect to ANG II exposure
and the available ANG II receptors. Thus vascular effects of ANG II could be counteracted by a modulated NO production; however,
proinflammatory properties of ANG II mediated via AT1
receptor activation could be potentiated via an enhanced NO production.
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MATERIALS AND METHODS |
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Materials. DMEM, L-glutamate, insulin, penicillin, streptomycin, LPS (Escherichia coli serotype 026:B6), IFN, NG-monomethyl-L-arginine (L-NMMA), actinomycin D, and ANG II were all purchased from Sigma (Deisenhofen, Germany). DUP-753 and PD-123319 were from Biotrend (Köln, Germany). FCS was from Boehringer (Mannheim, Germany), and tissue culture plastic was from Falcon (Becton-Dickinson, Heidelberg, Germany).
Cell culture and isolation of MCs.
For preparation and culture of glomerular MCs from male Sprague-Dawley
rats, standard techniques were used as described previously (34,
35). Briefly, the kidneys were excised, and the cortex was
separated from the medulla and homogenized using razor blades. Glomeruli were isolated from the homogenate by sequential sieving and
collected on a 75-µm sieve. After treatment with 500 U/ml collagenase
(type IV, Sigma) in PBS for 30 min, the glomerular remnants consisting
predominantly of endothelial cells and MCs were seeded in tissue flasks
(30,000 glomeruli/flask) containing DMEM supplemented with 20% FCS, 5 µg/ml bovine insulin, 100 U/ml penicillin, and 100 µg/ml
streptomycin. After three to four passages using 0.05% trypsin-0.02%
EDTA, stable and homogenous cultures of MCs were obtained. The
outgrowing cells were characterized as MCs by positive
immunocytochemical staining for Thy-1.1 (Serotec; Blackthorn, Bicester,
UK), smooth muscle cell actin, and myosin showing typical MC
morphology. MCs were further cultured in DMEM supplemented with
10% heat-inactivated FCS (56°C for 1 h), 2 mM glutamate, 5 ng/ml insulin, 100 U/ml penicillin, and 1 mg/ml streptomycin. Cells
were used for experiments at subconfluence or passaged at confluence
with trypsin/EDTA (0.05%-0.02% wt/vol). To obtain quiescent cells,
MCs were maintained in medium containing 0.5% FCS for 3 days before
cytokine treatment. MCs were used between passages 15 and 25. To
stimulate NOS-2 induction, we used LPS (E. coli serotype
026:B6, 10 µg/ml medium) and IFN (100 U/ml medium). Subconfluent quiescent MCs were coincubated according to the experimental conditions with ANG II (106-10
8 M), DUP-753
(10
5-10
7 M), PD-123319
(10
5-10
7 M), dexamethasone
(10
6 M), and/or TGF-
(5 ng/ml; Calbiochem, Germany).
Measurement of NO production by detection of nitrite in supernatants of cultured MCs. MCs were grown to subconfluence either in 24-well plates or, to accommodate protein or RNA extraction, on 60- and 100-mm plastic culture plates. The cells were conditioned in standard culture medium between 24 h and 32 h, according to the experimental protocol. The nitrite content was measured using the Griess colorimetric method (35). Briefly, 200 µl of the supernatant as well as control medium supplemented with known NaNO2 concentrations serving as standard were mixed with 75 µl Griess reagent including HCl [50 µl of Griess reagent (25 mM sulfanilamide, and 25 mM naphthylethylenediamine), and 25 µl 6 M HCl], and incubated for 30 min in the dark. Optical density was measured at 550 nm with an enzyme-linked immunosorbent assay (ELISA) reader, and the nitrite content of the conditioned medium was calculated according to the NaNO2 standard curve obtained.
Preparation of radioactive cDNA probes (rat NOS-2, rat GAPDH). The glyceraldehyde-3-phosphate dehydrogenase (GAPDH) probe was gained using a homology-based RT-PCR (30 cycles, annealing at 50°C) of rat MC cDNA using the following primers (synthesized by Life Technologies): sense, 5'-AATGCATCCTGCACCACCAA-3' (position 469-488 of M17701); and antisense, 5'-GTCATTGAGAGCAATGCCAGC-3' (position 919-939 of M17701).
We then used a 1,639-bp rat NOS-2 fragment which we obtained by homology-based RT-PCR (40 cycles, annealing at 60°C) of LPS/IFN-induced (24 h) cultured rat MCs, using the following primers (synthesized by Life Technologies): sense, 5'-CTGAATTCTGCATGGACCAGTATAAGGCAAGC-3' (position 1877-1900 of M84373); and antisense, 5'-CTGGATCCACCTGCTCCTCGCTCAA-3' (position 3479-3495 of M84373) (added restrictions sites are marked with underscore). 32P labeling was performed for all probes using the random prime labeling kit (Boehringer).Northern blot hybridization analysis.
Total RNA from rat MCs grown to subconfluence in 100-mm culture dishes
was obtained as described earlier (35). Twenty micrograms of total RNA were electrophoretically size-fractionated under denaturing conditions, using 1% agarose including 1.8% formaldehyde. The separated RNA was capillarily transferred by 20× SSC to nylon membranes (Amersham), and fixed through ultraviolet cross-linking. Next, the RNA was prehybridized (5× Denhardt's solution, 5× SSC, 50 mM Na3PO4, 0.1% SDS, 250 mg herring sperm DNA,
and 50% formamide) for at least 2 h at 42°C and then hybridized
with the 32P-labeled specific NOS-2 probe for 24 h,
using the same conditions. The membrane was washed at 42°C twice for
15 min with 2× SSC and 0.1% SDS, for 30 min with 0.1× SSC and 0.1%
SDS, and exposed to X-ray film (Kodak, XAR-5 supplied by Sigma,
Deisenhofen, Germany) at 80°C for at least 24 h. To rectify
for RNA transfer and content, filters were stained with bromophenol
blue (0.04% in 500 mM sodium acetate, pH 5.5), and rehybridized with a
32P-labeled rat GAPDH probe.
ANG II receptor subtype analysis by RT-PCR. One microgram total RNA of quiescent cultured rat MCs either grown under control conditions or exposed to LPS/IFN was subjected to an RT reaction using oligo-(dT)16 according to the protocol of the Perkin-Elmer Cetus GeneAmp RNA PCR kit. Specific primers for the homology-based PCR (36 cycles of 1 min of denaturation at 95°C, 30 s of annealing at 59°C, and 1 min of extension at 72°C) produced PCR products that were size separated on ethidium bromide-stained agarose gels, revealing bands at the expected respective sizes. Negative controls for the RT and the PCR reagents remained negative. Primer (synthesized by GIBCO) sequences were as follows for the AT1 receptors, sense, 5'-TCG AAT TCC ACC TAT GTA AGA TCG CTT C-3' (for the AT1A receptor position 554-573 of M74054, and for the AT1B receptor position 448-467 of X64052); and antisense, 5'-TCG GAT CCG CAC AAT CGC CAT AAT TAT CC-3' (for the AT1A receptor position 998-978 of M74054, and for the AT1B receptor position 892-872 of X64052); and for the AT2 receptor, sense, 5'-TCG AAT TCT TGC TGC CAC CAG CAG AAA C-3' (position 3070-3089 of RNU22663); and antisense, 5'-TCG GAT CCG TGT GGG CCT CCA AAC CAT TGC TA-3' (position 4172-4195 of RNU22663) as reported before (added restriction sites are underlined) (30, 38). Contamination of sample RNA by genomic DNA was excluded by directly subjecting the sample RNA to PCR amplification without RT step. A suitable EcoR I restriction site (30) within the AT1A receptor PCR product was used to distinguish between AT1A and AT1B PCR products.
Western blot analysis of NOS-2 protein. Preparation of MC lysates and immunoblot analysis were performed using time-matched 60-mm plates of MCs. They were incubated with complete medium containing 0.5% FCS, medium supplemented with LPS (10 µg/ml medium) and IFN (100 U/ml medium), or medium supplemented additionally with reagents according to the experimental protocol. At the end of the incubation period, the medium was removed and assayed for the nitrite content, as described above. The plates were washed twice with ice-cold PBS. Thereafter, 0.5 ml of boiling lysis buffer solution (1% SDS and 10 mM Tris, pH 7.4) was added to the cells. The cell lysates were collected in centrifuge tubes after boiling and spinning at 12,000 g for 5 min at 4°C. Protein contents were determined using the bicinchoninic acid protein assay reagent (BCA; Pierce, Munich, Germany). The samples (10 µg) were diluted in electrophoresis sample buffer (250 mM Tris, pH 6.8, 4% SDS, 10% glycerol, 0.006% bromphenol blue, and 2% mercaptoethanol), boiled for 5 min, quenched on ice, and resolved by electrophoresis through 0.1% SDS-10% polyacrylamide gels. Proteins were electrophoretically transferred to nitrocellulose membranes (Amersham, Braunschweig, Germany). The transfer efficiency was determined by staining the membranes with Ponceau S. After destaining in distilled water, the membranes were quenched in blocking solution [5% powdered dried low-fat milk in washing solution (10 mM Tris, pH 7.5; 100 mM NaCl, 0.1% Tween 20)] for 1 h at room temperature. The blocking solution was decanted, and membranes were incubated for 1 h at room temperature with the primary antibody (1:1,000, anti-NOS-2, rabbit, polyclonal; Affiniti, Nottingham, UK). Membranes were washed for 30 min at room temperature with several changes of the washing solution. The secondary detecting antibody was horseradish peroxidase-conjugated anti-rabbit IgG (1:2,000; Serva, Heidelberg, Germany) which was used with the enhanced chemiluminescence protocol (Amersham).
Statistical analysis. When suitable, means ± SE were determined. To test for statistically significant differences, we used Student's t-test or analysis of variance, if multiple comparisons were made against a single control, whenever applicable. Significance was assigned at P < 0.05.
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RESULTS |
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LPS/IFN-induced nitrite production is inhibited by ANG II.
The stable metabolite of NO, nitrite, was utilized to assess NO
production resulting from NOS-2 induction in cultured rat MCs.
Incubation with LPS/IFN for 24 h increased nitrite accumulation in
the cell culture supernatant (27.2 ± 1.7 µmol/ml medium)
compared with untreated controls (0.6 ± 0.3 µmol/ml medium).
Coincubation with ANG II (107 M) inhibited nitrite
accumulation (16.2 ± 1.0 µmol/ml medium) (Fig.
1A). The dose response for ANG
II (10
8-10
6 M) revealed the inhibition
to be at maximum with concentrations of ANG II of 10
7 M. This concentration was also applied in successive experiments (Fig.
1B). The NOS inhibitor L-NMMA (10
4
M) was unable to reverse the nitrite production in all experimental conditions (data not shown).
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Twenty-four-hour coincubation of ANG II with LPS/IFN reduced
cumulative NOS-2 mRNA and protein expression.
To assess regulatory mechanisms involved in the effects of the
coincubational downregulation of LPS/IFN-induced NO synthesis by ANG
II, total RNA was subjected to size fractionation. No cumulative NOS-2
mRNA expression was detected by Northern blot analysis in cultured MCs
in basal conditions; however, 24-h incubation with LPS/IFN induced
NOS-2 mRNA, which was markedly diminished in the presence of ANG II
(107 M) (Fig.
2A). The inhibitory effect of
TGF-
on LPS/IFN-induced NOS-2 mRNA expression was chosen as control.
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NOS-2 inhibition is ANG II receptor subtype specific.
Using nitrite accumulation to determine NOS-2-derived NO production, we
evaluated ANG II receptor subtype specificity. Incubation with the
AT1- and AT2-specific inhibitors DUP-753 and
PD-123319, respectively, (both at 106 M) did not
influence LPS/IFN-induced nitrite production. The inhibition by ANG II
(10
7 M) was not abolished by DUP-753; however, PD-123319
completely reversed the ANG II-dependent inhibition of NO production,
suggesting an AT2-mediated inhibitory effect under these
experimental conditions (Fig. 3).
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Presence of ANG II receptor subtypes in cultured rat MCs. Homology-based RT-PCR reactions for AT1 and AT2 receptors generated PCR products of the expected size on separation on ethidium bromide-stained agarose gels. Their identity was verified by restriction analysis. A suitable EcoR I restriction site within the AT1A PCR product (30) was used to identify AT1A and AT1B receptor subtypes (data not shown).
Twenty-four-hour preincubation with LPS/IFN followed by 8-h
coincubation with ANG II enhanced NOS-2 protein expression and NO
production in cultured rat MCs.
The NOS-2 protein expression increased with 32 h of LPS/IFN
stimulation compared with experiments lasting only 24 h. The
addition of ANG II (107 M) for the final 8-h period
following 24-h preincubation with LPS/IFN alone augmented the NOS-2
protein expression even further compared with 32-h incubation with
LPS/IFN alone (Fig. 4).
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Dexamethasone converts coincubational inhibition by ANG II into
stimulation of nitrite production via the AT1 receptor.
Dexamethasone is known to enhance AT1A receptor expression
and ANG II binding to AT1 receptors in several tissues
including MCs (8, 33, 46). To verify our finding of an
AT1-mediated potentiation of NO production, we stimulated
cultured MCs for 24 h with LPS/IFN; however, dexamethasone was
added only after 18 h for the remaining 6-h period to allow an
induced nitrite production, since dexamethasone is known to inhibit
NOS-2 expression on coincubation. The coincubation with ANG II
(107 M) for the whole experimental period of 24 h
led to a significant increase in induced nitrite production (161.5 ± 25% of LPS/IFN and dexamethasone). The inhibition of the
AT1 receptor abolished the ANG II effect completely to
92.3 ± 15.4% of LPS/IFN and dexamethasone. In support of an
AT1-mediated NO synthesis as well as a downregulatory action of AT2 activation, inhibition of the AT2
receptor resulted in a pronounced increase in nitrite production
(461.3 ± 46.3% of LPS/IFN and dexamethasone) (Fig.
6).
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DISCUSSION |
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In glomerulonephritis, MCs are exposed to several proinflammatory stimuli such as cytokines which are either secreted by invading inflammatory cells, by resident cells themselves, or by other mediators such as systemically or locally synthesized ANG II.
Glomerular MCs play a critical role in the perpetuation of glomerular diseases (47). ANG II takes part in the inflammatory response of MCs (51, 54). Therefore, we analyzed whether ANG II regulates induced NO production in the inflammatory range resulting from expression of active NOS-2 protein.
We chose a model of inflammation simulated by exposure of MCs to LPS
and IFN. These mediators lead to PKC/mitogen-activated protein
kinase-dependent and -independent activation of nuclear factor-B
(NF-
B) and IFN regulatory factors, resulting in induced NO
levels within the inflammatory range (9, 10, 11). Our experiments suggest ANG II-dependent inhibition of LPS/IFN-induced production of NO in MCs during simultaneous exposure of the cells to
LPS/IFN and ANG II. This effect of ANG II was dose dependent within a
physiological concentration range. Analysis of cumulative NOS-2 mRNA as
well as of NOS-2 protein expression indicated a transcriptional regulation.
In an evaluation of the ANG II receptor subtype involved, neither
specific receptor blocker (DUP-753 for the AT1 subtype, and
PD-123319 for the AT2 subtype) exerted nonspecific effects on nitrite production at concentrations of 106 M if
incubated without ANG II. This finding also excluded a significant contamination by exogenous ANG II. The inhibitory effect of ANG II was
unaffected by coincubation with the AT1 but was completely abolished with the AT2 receptor blocker. This suggests an
inhibitory effect mediated via AT2 receptors. We were
unable to demonstrate a significant direct stimulatory effect through
AT1 receptors in this coincubation setup, although a slight
tendency toward an increased nitrite production was visible. ANG II
binding studies and the demonstration of AT1A and
AT1B receptor subtype mRNA (6, 7, 8) proposed
the predominance of AT1 receptors in humans and rats,
preferentially of the AT1B subtype in rat MCs. Conversely, Ernsberger et al. (13) and Goto et al. (15)
demonstrated by pharmacological and molecular biological means,
respectively, the existence of AT2 receptors in rat MCs.
Ardaillou et al. (2) found a marked expression of
AT2 receptors in mouse MCs. To demonstrate the
receptor expression in our MC cultures, we performed RT-PCR for the
AT1 and the AT2 receptor subtypes, as well as
restriction analysis for the AT1A and AT1B
subtypes, in support of our functional data regarding receptor subtype
activity and specificity.
In all studies performed so far addressing the potential interaction of
ANG II and induced NO production, coincubation experiments were
performed omitting the impact of a sequential occurrence of stimuli
modulating not just the inflammatory response but also ANG II receptor
subtype expression. ANG II may act differentially with respect to the
AT receptors present. The AT receptor expression is subject to changes
in the presence of inflammatory stimuli or glucocorticosteroids. We
hypothesized that ANG II would act proinflammatory via the
AT1 receptor by enhancing induced NO production in
inflammatory conditions. Inflammation, as induced by LPS and cytokines,
led to downregulation of the AT2 receptor, as was
previously described in rat fibroblasts (48). An enhanced
AT1 receptor-mediated signal transduction (24)
and an LPS-mediated induction of the cyclooxygenase (COX-2) via the
AT1 receptor has been demonstrated in rat vascular smooth
muscle cells (39). COX-2 activity then stimulates a
cAMP-dependent PKA activation of the NF-B pathway, a factor which
results in a profound production of induced NO (9, 10,
11).
ANG II added to cultured MCs preexposed to LPS/IFN further stimulated NOS-2 protein and NO production. This increase was abolished by AT1 blockade in a dose-dependent fashion, even below control inductions, indicative of a downregulatory AT2 effect. In contrast to the coincubation experiments, a stimulatory effect of AT1 receptor activation was now observed on blockade of the AT2 receptor. This supported our assumption of a potential AT1 receptor-related stimulation of induced NO production, however, controlled via AT2 receptors.
To further clarify the role of the AT1 receptor,
dexamethasone was applied to stimulate AT1A receptor
expression (8) and binding of ANG II and to reduce
AT2 receptor expression (48). We
exploited the fact that the AT1A receptor promotor, in
contrast to the AT1B and the AT2 receptor,
contains three putative glucocorticoid-responsive elements,
one of which has already been found to be dexamethasone responsive at
concentrations of 106 M (17). An
AT1-related stimulatory effect of ANG II on LPS/IFN-induced NO production was present despite the inhibitory effect of
dexamethasone on NOS-2 transcription via upregulated
inhibitor-
B
activity and decreased nuclear p65
translocation (12). This is consistent with previous
observations in a model of interleukin-1
(IL-1
)-dependent induced
NO production with either
1-adrenergic- or
adenosine-related stimulation in rat vascular smooth muscle cells and
cardiac myocytes (20, 22).
The reduction of NO synthesis on coincubation with LPS/IFN and ANG II
is in agreement with studies performed by Kihara et al.
(27) and Wolf et al. (52) in cultured MCs and
tubular cells, respectively. Neither study demonstrated ANG II receptor expression. Kihara et al. (27) demonstrated an inhibition
of IL-1-induced NO production on coincubation with ANG II. In
contrast to our findings, the AT1 receptor antagonist
CV-11974, but not the AT2 blocker PD-123319, was able to
reverse the ANG II-dependent inhibition at concentrations of
10
5 M, which may already exert unspecific effects on
AT2 receptors (18). No NOS-2 transcriptional
downregulation was observed in either study (27, 52). Wolf
et al. (52) suggested posttranscriptional mechanisms
beyond mRNA stability. However, in our experiments the LPS/IFN-induced
steady-state NOS-2 mRNA and protein expression were reduced by ANG II
at concentrations of 10
7 M on coincubation, indicating
mechanisms involving transcription or mRNA stability. We were able to
demonstrate that actinomycin D was able to almost completely abolish
the ANG II-dependent stimulation of nitrite production of MCs when
preincubated with LPS/IFN for 24 h. This suggested a
transcriptional regulation, although additional mechanisms may still be active.
In line with our findings, an IL-1- and LPS-mediated NOS-2 induction
has been demonstrated in rat and rabbit cardiac myocytes in which
AT1 receptor-mediated augmentation of NO and cGMP
production led to contractile depression (21, 53). In
further support of our own finding of an AT2
receptor-dependent downregulation of induced NO production is a
preliminary report by Peters et al. (40), who found
inhibition of LPS/IFN-mediated NOS-2 induction by ANG II and ANG II
fragments in cultured rat MCs (fragments 1-7 and 3-8).
In conclusion, we describe a regulatory mechanism by which ANG II acts on glomerular mesangial NO production in a time- and ANG II receptor-dependent manner, and we provide evidence for a significant contribution of renal AT2 receptors. This supplements reports of AT2-related renal responses, such as pressure natriuresis (16, 31). ANG II interacts with induced NO production upon proinflammatory stimuli. An inflammatory response may be enhanced following AT1 receptor stimulation by an augmented NO production. In contrast, AT2 receptor activation restricts the amount of induced NO synthesis. Once the AT1/AT2 receptor balance is shifted toward the AT1 receptor, the NO response is augmented most likely transcriptionally. Further work will need to be directed toward elucidating the related downstream signals. The reported ANG II-dependent regulation of induced NO production opens new insight into understanding renal damage and functional deterioration in the presence of glomerular inflammation, yet the extent of its contribution still needs to be defined.
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ACKNOWLEDGEMENTS |
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We thank Dr. Michael Wiederkehr for critical review of this manuscript and C. Staiger for excellent technical assistance.
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FOOTNOTES |
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This work was supported in part by the Deutsche Forschungsgemeinschaft Klinische Forschergruppe "Molekulare Regulationsmechanismen in glomerulären Zellen der Niere" and by a grant from the Swiss National Foundation, both to M. G. Mohaupt.
Address for reprint requests and other correspondence: M. G. Mohaupt, Univ. Hospital, Berne, Division of Nephrology/Hypertension, 3010 Berne, Switzerland (E-mail: markus.mohaupt{at}insel.ch).
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 27 March 2000; accepted in final form 11 August 2000.
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