Functional expression of the renin-angiotensin system in human podocytes
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《美国生理学杂志》
1Department of Medicine, Division of Nephrology and General Medicine, University Hospital Freiburg, Freiburg
2Department of Medicine D, Division of General Internal Medicine and Nephrology, University Hospital Münster, Münster
3Department of Pathology, University of Freiburg, Freiburg
4Department of Anatomy and Cell Biology, University of Heidelberg, Heidelberg, Germany
5Children's Renal Unit and Academic Renal Unit, Southmead Hospital, University of Bristol, Bristol, United Kingdom
ABSTRACT
Experimental and clinical studies impressively demonstrate that angiotensin-converting enzyme inhibitors (ACEI) and angiotensin receptor blockers (ARB) significantly reduce proteinuria and retard progression of glomerular disease. The underlying intraglomerular mechanisms are not yet fully elucidated. As podocyte injury constitutes a critical step in the pathogenesis of glomerular proteinuria, beneficial effects of ACEI and ARB may partially result from interference with a local renin-angiotensin system (RAS) in podocytes. The knowledge of expression and function of a local RAS in podocytes is limited. In this study, we demonstrate functional expression of key components of the RAS in differentiated human podocytes: podocytes express mRNA for angiotensinogen, renin, ACE type 1, and the AT1 and AT2 angiotensin receptor subtypes. In Western blot experiments and immunostainings, expression of the AT1 and AT2 receptor was demonstrated both in differentiated human podocytes and in human kidney cortex. ANG II induced a concentration-dependent increase in cytosolic Ca2+ concentration via AT1 receptors in differentiated human podocytes, whereas it did not increase cAMP. Furthermore, ANG II secretion was detected, which was blocked by neither the ACEI captopril nor the renin inhibitor remikiren nor the chymase inhibitor chymostatin. ANG II secretion of podocytes was not increased by mechanical stress. Finally, ANG II was found to increase staurosporine-induced apoptosis in podocytes. We speculate that ACEI and ARB exert their beneficial effects, in part, by interfering with a local RAS in podocytes. Further experiments are required to identify the underlying molecular mechanism(s) of podocyte protection.
cytosolic Ca2+; human; captopril; AT1 receptor; AT2 receptor; apoptosis; mechanical stress
THE RENIN-ANGIOTENSIN SYSTEM (RAS) plays a crucial role in controlling blood pressure, fluid balance, and electrolyte homeostasis (48, 51). Enzymatic cleavage by renin and angiotensin-converting enzyme (ACE) leads to the conversion of angiotensinogen via ANG I to its main effector molecule ANG II. Apart from its hemodynamic action, ANG II is capable of inducing a multitude of nonhemodynamic effects such as induction of oxygen radicals, cytokines, stimulation of collagen synthesis, apoptosis, proliferation, and hypertrophy (20, 51). Within the glomerulus, ANG II reduces the ultrafiltration coefficient and modulates glomerular capillary permselectivity leading to proteinuria, initiating and increasing subsequent tubulointerstitial injury (3, 26, 42). Acting as growth hormone, ANG II contributes to the pathogenesis of glomerulosclerosis (20).
Clinical studies have impressively demonstrated that ACE inhibitors (ACEI) markedly reduce proteinuria and progression of glomerular disease (28, 31, 43). Furthermore, angiotensin type 1 receptor blockers (ARB) have clearly been shown to retard the progression of diabetic nephropathy (6, 29). Most recently, the addition of ARB to ACEI treatment further reduced proteinuria (2, 27).
The podocyte is the most differentiated cell type within the glomerulus. Synthesizing various components of the glomerular basement membrane and the slit diaphragm, it is crucially involved in the maintenance of the glomerular filtration barrier (40). Podocyte injury leads to alterations of the foot process/slit diaphragm complex with loss of glomerular permselectivity, resulting in proteinuria (40).
Interestingly, little is known about the expression and function of the RAS in podocytes. In this regard, one may hypothesize that the antiproteinuric effects of ACEI or ARB may partially result from interference with a local RAS in podocytes. This prompted us to specifically investigate expression and function of components of the RAS in differentiated human podocytes.
METHODS
Cell culture. The conditionally immortalized human podocyte cell line has been developed as recently described by transfection with the temperature-sensitive SV40 T-gene (44). Proliferating at a temperature of 33°C, these cells transform into a quiescent, differentiated phenotype after transfer to 37°C. Cells then stain positive for the in vivo podocyte markers synaptopodin, nephrin, podocin, CD2AP, ZO-1, -, -, -catenins, and P-cadherin (44). For experiments, cells between passage 10 and 18 were seeded at 37°C into 6-well plates, 24-well plates, or flasks and cultured in standard RPMI media containing 10% fetal calf serum (Biochrom, Berlin, Germany), 2.5 mM L-glutamin, 0.1 mM sodium pyruvate, 1 g/l nonessential amino acids (all Seromed, Berlin, Germany), and insulin-transferrin-sodium selenite supplement (Boehringer, Mannheim, Germany), 100 U/ml penicillin, and 100 mg/ml streptomycin (GIBCO, Eggenstein, Germany) for at least 7 days until cells were differentiated showing an arborized morphology.
RT-PCR for components of the RAS. The RNA preparation, the reverse transcription, and the PCR were performed according to the method recently described (14). In brief, the total RNA from cultured human podocytes was isolated with guanidinium/acid phenol/chloroform extraction and the amount of RNA was measured with spectrophotometry. For first-strand synthesis, total RNA from podocytes was mixed in 5x reverse transcription buffer and completed with 0.5 mM dNTP, 10 μM random hexanucleotide primer, 10 mM dithiothreitol, Moloney murine leukemia virus 0.02 U RNAse inhibitor/ng RNA, and 100 U MMLV reverse transcriptase/μg RNA. PCR amplification of RT reactions without reverse transcriptase revealed no PCR product, thereby excluding amplification of genomic DNA. RT was performed at 42°C for 1 h, followed by a denaturation at 95°C for 5 min. PCR was performed in duplicate with a total volume of 20 μl, each containing 40 ng RNA per 4 μl of RT template, 16 μl of PCR master mixture, and 10 pmol each of sense and antisense primer. The mixture was overlaid with mineral oil and heated for 2 min at 94°C. The cycle profile consisted of 1 min of denaturation at 94°C, annealing for 1 min at 60°C (renin: 58°C; ACE1: 59°C), and extension for 1 min at 72°C. For amplification, different numbers of cycles were used (see Table 1). The amplification products of 10 μl of each PCR reaction were separated on a 1.5% agarose gel, stained with ethidium bromide, and visualized by UV irradiation. The primer oligonucleotides were selected from published cDNA sequences and are depicted in Table 1. All primers are noted in 5'-3' direction.
Immunohistochemistry of kidney cortex. Normal kidney cortex was obtained from nephrectomy specimens. Fixation and preparation of tissue for immunohistochemical analyses were performed using standard techniques. In brief, kidneys were perfused after removal with cold (4°C) PBS followed by 4% formaldehyde. Kidneys were then incubated for 24 h at 4°C in 4% formaldehyde solution, embedded in paraffin, and cut into 4-μm-thick slices. Slices were deparaffinized in xylol for 1 h, gradually hydrated through graded alcohols (100 to 70%), and washed in deionized water. After incubation in 1% H2O2 for 30 min, slices were rehydrated with PBS, and antigen unmasking was performed by boiling slices in 10 mM citrate buffer (pH 6) for AT1 and WT1 or EDTA (pH 9) for AT2 for 2 x 10 min. Blocking was performed using a 1% BSA solution for 10 min.
For staining of WT1 and AT1, sections were then incubated overnight in a humidified chamber at 4°C with antibodies (Abs) to the angiotensin AT1 receptor (rabbit anti-human, 200 μg/ml, 1:200 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) and WT1 (cat anti-human, 100 μg/ml, 1:100 dilution; Santa Cruz Biotechnology). The slices were washed extensively with PBS and incubated for 90 min with a secondary antibody using a commercially available ABC-Kit (Vectastain Mouse Peroxidase, Vectorlabs, Burlingame, CA). Slices were washed with PBS, incubated with avidin-biotin for 45 min, and stained with 3-amino-9 ethylcarbazole (red) or fast blue BB salt/naphtol AS-MX phosphate (blue; Sigma, Steinheim, Germany). Sections were examined with a conventional light microscope (Zeiss LSM 510; Zeiss, Oberkochen, Germany) and photographed using a digital camera. Negative controls were performed by elimination of the primary Abs. For "double-labeling" experiments, serial ultrathin sections of human kidneys were stained with Abs to the angiotensin AT1 receptor and WT1 and pictures of the same glomerulus were projected on top of each other using commercially available computer programs.
For staining of WT1 and AT2, sections were incubated for 30 min (WT1) or 60 min (AT2) in a humidified chamber at 4°C with Abs to the angiotensin AT2 receptor (goat anti-human, 200 μg/ml, 1:50, Santa Cruz Biotechnology) and WT1 (mouse anti-human, 200 μg/ml, 1:100 dilution; Santa Cruz Biotechnology). AT2 was then detected using a labeled streptavidin-biotin detection kit (ChemMate Detection Kit, Alkaline Phosphatase/Red, Code K 5005, Dako; red), and WT1 was detected using an ABC Kit (StreptABComplex/HRP Duet, Code K 0492, Dako, Hamburg, Germany; brown). All sections were then also stained using Mayer's hemalaun. Sections were examined with a conventional light microscope (Zeiss Axioskop 50) and photographed using a digital camera (Sony MC-3210/II/PM). Negative controls were performed by elimination of the primary Abs.
Immunocytochemistry of cultured human podocytes. Immunolabeling was done as previously described (34). Briefly, coverslips were fixed with 2% paraformaldehyde and 4% sucrose in PBS for 10 min and were then permeabilized with 0.3% Triton X-100 (Sigma-Aldrich, Munich, Germany) in PBS for 10 min. Nonspecific binding sites were blocked with 2% FCS, 2% bovine serum albumin, and fish gelatine (Sigma-Aldrich) in PBS for 30 min. Primary (rabbit anti-AT1 and rabbit anti-AT2, both Santa Cruz Biotechnology) and secondary antibodies were applied in a 1:100 dilution according to standard techniques, and the coverslips were mounted on glass slides with 15% Mowiol (Calbiochem, La Jolla, CA) and 50% glycerol in PBS. After being washed with PBS, bound primary Abs were detected using FITC-conjugated cat anti-rabbit IgG (Santa Cruz Biotechnology). As controls, primary Abs were either omitted or boiled at 99°C for 10 min and used thereafter. Micrographs were taken on a microscope equipped for epiluminescence (Zeiss Axiophot).
Western blotting. Western blotting was performed using standard techniques. In brief, cells were washed once with PBS, scraped with lysis buffer containing 2 mM EDTA, 2 mM EGTA, 100 mM NaCl, 20 mM Tris, 0.1% SDS, 1% NP-40, 2 mM PMSF, and proteinase inhibitor cocktail (Roche Diagnostics, Mannheim, Germany), and sonicated. The samples were resuspended in Laemmli sample buffer, boiled (5 min), and subjected to SDS-PAGE and transfer electrophoresis. The transblots were stained with Ponceau solution to prove equal amounts of protein were loaded on the membrane and probed with primary Abs (rabbit anti-AT1 and goat anti-AT2, both either with or without blocking peptide 5:1; both Santa Cruz Biotechnology) followed by peroxidase-labeled secondary antibodies (donkey anti-rabbit, 1:4,000; Amersham Pharmacia, Piscataway, NJ; rabbit anti-goat, Dako) and detected by chemiluminescence detection reagents (ECL, Amersham Pharmacia).
Measurements of the intracellular calcium activity. Measurements of cytosolic calcium (Cai2+) with the Ca2+-sensitive dye fura 2 (Sigma-Aldrich) were performed in podocytes on an inverted fluorescence microscope setup (38). The system allows fluorescence measurements at the single-cell level at three excitation wavelengths. The field of measurement can be set between a diameter of 2 and 300 μm with an adjustable pinhole. A time resolution of up to 200 Hz was achieved by using a high-speed filter wheel and a single-photon counting tube (Hamamatsu H63460 [GenBank] –04; Hamamatsu, Herrsching, Germany). The autofluorescence signal of cells that had not been loaded with fura 2 was measured and subtracted from the results obtained in fura 2-loaded cells. This had no effect on the bandwidth of the measurements. A calibration of the fura 2 fluorescence signal was attempted at the end of each experiment by using Ca2+ ionophore ionomycin (1 μM) and low- and high-Ca2+ buffers. Cai2+ concentration ([Ca2+]i) was calculated from the fluorescence ratio 340:380 nm according to the equation described by Grynkiewicz et al. (15).
Measurements of intracellular cAMP. Podocytes were cultured in six-well plates. They were kept at 37°C and rinsed with a physiological Ringer solution. After preincubation with 0.5 M 3-isobutyl-I-methylxanthine (IBMX; RBI, Cologne, Germany) for 5 min, cells were exposed to the added agents. To terminate the assay, the supernatants were rapidly removed and cells were rinsed with ice-cold ethanol. After an ethanol extraction, cAMP concentrations were measured with an ELISA (Amersham Buchler, Braunschweig, Germany).
Measurements of ANG II secretion. Podocytes were cultured in six-well plates. They were kept at 37°C, and the medium was changed from 10 to 0% FCS. The cells were then incubated either with vehicle or with remikiren (a kind gift of Dr. D. Müller, Berlin, Germany), captopril, chymostatin (both Sigma), or a combination of the three. After 24 h, the supernatant was taken and centrifuged at 10,000 rpm for 5 min. Concentrations were then measured with an ELISA (Spibio, Massy Cedex, France).
Mechanical stress experiments. Differentiated human podocytes were seeded in six-well plates with a flexible bottom (Bioflex, Flexcell International, Hillsborough, NC). The flexible silicone membranes were coated with collagen IV to facilitate cell attachment. After 3 days, the six-well plate was mounted on a Plexiglas manifold connected to a custom-built apparatus, which induced cyclic variations in air pressure below and above atmospheric pressure. Cyclic pressure variations caused upward and downward motion of the silicone membranes. Pressure amplitude was chosen to give a maximum up- and downward deflection of the membrane center of 6 mm, being equal to an increase in membrane area by 11%, or 5% mean linear cell strain. Cycle frequency was adjusted to 0.5 Hz.
Measurements of lactate dehydrogenase release from podocytes. To assess cytotoxicity, lactate dehydrogenase (LDH) release was measured with a routine autoanalyzer (Modular I, Hitachi, Tokyo, Japan). Total LDH content of the cells was measured after incubation of cells in 1% Triton X-100.
Quantification of apoptosis and necrosis by flow cytometry. Apoptosis was determined by detecting annexin V binding using flow cytometry as described previously (24). Evaluation of cell necrosis was performed by simultaneous detection of propidium iodide (PI) uptake by the nonpermeabilized cells. Immortalized podocytes, after 24-h incubation with the indicated stimuli, were labeled with annexin V-FITC (Bender Medsystems, Vienna, Austria) and PI (5 μg/ml) in staining buffer (containing 1% BSA in 50 mM HEPES buffer, pH 7.4) for 15 min on ice. After being stained, cells were washed in PBS and fixed in 4% paraformaldehyde to apply flow cytometry. Cells were analyzed on a FACScan flow cytometer (Becton Dickinson, Mountain View, CA).
TUNEL procedure. DNA strand breaks were identified by TUNEL (TdT-mediated dUTP nick-end labeling) reaction using an in situ cell death detection kit (Roche) according to the manufacturer's instructions. First, podocytes were cultured on coverslips incubated for indicated times, then washed with PBS fixed in 4% paraformaldehyde at room temperature for 30 min. Subsequently, the cytoplasmic membrane was permeabilized with 0.1% Triton X-100 for 2 min on ice. After being washed two times, cells were incubated with TUNEL reaction mixture for 60 min at 37°C. Samples were directly analyzed under a fluorescence microscope by detection at 550 nm. Cells labeled in the absence of TdT were always used as negative controls, whereas cells pretreated with DNase after permeabilization were used as positive controls.
Chemicals. Unless otherwise indicated, chemicals were purchased from Sigma.
Statistical analyses. Data are given as means ± SE, where n refers to the number of experiments. Student's t-test was used to compare mean values within one experimental series. A P value 0.05 was considered statistically significant.
RESULTS
mRNA expression of key components of the RAS in differentiated human podocytes. Differentiated human podocytes express mRNA for angiotensinogen, renin, ACE type 1, and both the AT1 and AT2 angiotensin receptor subtypes (Fig. 1). PCR primers and conditions are depicted in Table 1.
AT receptor expression in differentiated human podocytes. Western blot analyses consistently revealed protein expression of both the AT1 and AT2 receptor in human podocytes (AT1 receptor: Fig. 2A, lanes 1 and 2; AT2 receptor: Fig. 2B, lanes 1 and 2), the specificity of which was confirmed using respective blocking peptides (AT1 receptor: Fig. 2A, lanes 3 and 4; AT2 receptor: Fig. 2B, lanes 3 and 4). In additional immunofluorescence experiments, expression of AT1 and AT2 receptors was detected (Fig. 3). Here, expression of the AT1 receptor was more prominent than AT2 receptor expression. Immunostainings most often detected angiotensin receptors as intracellular clusters. Similar expression patterns of angiotensin receptors have also been recently shown in various other cell types (37, 50).
Angiotensin receptor expression in human kidney cortex. Merged pictures of serial sections of human kidney cortex stained with antibodies against AT1 receptors (blue) and the podocyte-specific protein WT1 (red) demonstrate colocalization of AT1 receptors and WT1 in human podocytes ex vivo (Fig. 4, A-C).
In separate experiments, AT2 receptor expression was detected. Double stainings of human kidney cortex revealed colocalization of AT2 receptors (red) and WT1 (brown) in human podocytes ex vivo (Fig. 4, D-E).
ANG II increases [Cai2+] in differentiated human podocytes via AT1 receptors. In microfluorescence experiments, ANG II reversibly increased [Ca2+]i in podocytes (original experiment, Fig. 5A). This [Ca2+]i increase was concentration dependent (Fig. 5B; EC: 60 nM). The ANG II-induced [Ca2+]i increase was reversibly blocked by the AT1 receptor antagonist losartan (Fig. 5C).
ANG II does not increase the cAMP concentration in differentiated human podocytes. Recently, it has been reported that ANG II elevates the cAMP concentration in rat glomerular epithelial cells in culture (46, 47). To address this issue in differentiated human podocytes, cAMP concentration was measured in separate experiments. Within a time course of 2 h, ANG II (1 μM) did not increase cAMP production in human podocytes (Fig. 6A). To exclude rapid cAMP degradation by phosphodiesterases during the experiments, the specific phosphodiesterase inhibitor IBMX (500 μM) was added. Again, no elevation of cAMP levels was observed (Fig. 6B). In the presence of IBMX, the membrane-permeable adenylate cyclase activator forskolin concentration dependently increased cAMP production. In this setting, ANG II had no additive effect (Fig. 6C).
Differentiated human podocytes secrete ANG II. After having detected mRNA expression of key components of a local RAS, we were interested in secretion of ANG II as the final RAS effector by differentiated human podocytes. As detected by ELISA, human podocytes secrete ANG II (Fig. 7A). However, neither the specific renin inhibitor remikiren (10 μM), the ACE inhibitor captopril (10 μM), the chymase inhibitor chymostatin (100 μM), nor a combination of the three inhibitors significantly reduced ANG II secretion of podocytes (Fig. 7A). Furthermore, ANG II secretion of podocytes was not altered by mechanical stress (Fig. 7B).
ANG II does not increase LDH release in differentiated human podocytes. To test for podocyte damage induced by ANG II, LDH release experiments were performed. Even on prolonged stimulation at high ANG II concentration (24 and 48 h, 1 μM), LDH release was not significantly increased (n = 6–12; Fig. 8).
ANG II increases staurosporine-induced apoptosis in differentiated human podocytes. As measured by flow cytometry, staurosporine (1 μM, 24 h) induced apoptosis of differentiated human podocytes. Stimulation of podocytes with ANG II (1 μM) in addition to staurosporine increased the rate of apoptosis induced by staurosporine only (1 μM). However, stimulation with ANG II only did not alter the rate of apoptosis (Fig. 9). These findings were confirmed by TUNEL assay (data not shown).
DISCUSSION
Based on both experimental and clinical studies, there is clear evidence that the RAS plays a pivotal role in the pathophysiology of cardiovascular and renal disease (48, 51). Furthermore, to its prominent hemodynamic actions, the key effector ANG II causes numerous nonhemodynamic renal effects: ANG II is a growth factor of glomerular mesangial and endothelial cells, tubular epithelial cells, and fibroblasts (18, 51). ANG II causes cellular hypertrophy (51) and induces apoptosis (4). ANG II increases the expression of TGF- and stimulates synthesis of extracellular matrix proteins such as collagen type IV (18, 51). ANG II acts as proinflammatory cytokine and promotes oxidative stress (48, 51). Given this multitude of ANG II effects, local angiotensin-generating systems are of special interest (1).
Within the glomerulus, podocytes are critically involved in maintaining the glomerular filtration barrier and in counteracting glomerular capillary wall distension (23, 40). Podocyte damage leads to proteinuria and initiates glomerulosclerosis, finally resulting in progressive loss of kidney function (22, 40).
Indirect evidence suggests that ANG II is capable of disturbing podocyte biology. Spontaneous proteinuria in Munich- Wistar-Froemter rats has been attributed to an altered distribution of the slit diaphragm protein zonula occludens-1, both of which were prevented by the ACEI lisinopril (30). In experimental diabetic nephropathy, podocyte foot process broadening was attenuated by both the ACEI ramipril and the ARB valsartan (32). In animal models of diabetic nephropathy, reduction of nephrin expression was attenuated by the ACEI perindopril (19) and by the ARB valsartan and irbesartan (5, 8). Selective overexpression of the AT1 receptor in podocytes in rats has most recently been reported to lead to protein leakage and structural podocyte damage, progressing to glomerulosclerosis (17). In a recent renal biopsy study, placebo-treated patients with diabetic nephropathy showed a marked reduction of glomerular nephrin expression compared with nondiabetic control subjects, whereas nephrin expression in patients treated with the ACEI perindopril was similar to the control group (25). Another renal biopsy study revealed reduction of glomerular nephrin expression in both type 1 and type 2 diabetic patients compared with controls (10). These findings suggest direct rather than indirect ANG II effects on podocytes, which furthermore are independent of its hemodynamic actions.
We show for the first time that differentiated human podocytes express all RAS components required to generate ANG II. Furthermore, secretion of ANG II by resting human podocytes was observed. ANG II production has most recently been reported to occur in primary mouse podocytes exposed to mechanical strain (11). We exposed podocytes to mechanical strain under conditions known to affect podocytes (12) but did not find a significant rise in ANG II secretion. It is of note that ANG II production in both studies was not inhibited by the ACEI captopril, suggesting predominant involvement of non-ACE pathways (9) to generate ANG II in podocytes. Similar findings have been reported in both renal and extrarenal tissues (1). As neither a renin inhibitor nor a chymase inhibitor significantly changed ANG II secretion, in podocytes ANG II seems to be produced via an as yet unidentified alternative pathway.
Within the proximal tubular fluid, ANG II concentrations of 6–10 pmol/ml have been measured (7, 36). These high concentrations cannot be explained by filtration of circulating ANG II (36, 45). Given the data provided here, ANG II secretion by podocytes may contribute to the high intratubular ANG II concentration. The extent of this contribution remains to be estimated using a suitable experimental approach. However, ANG II in the tubular fluid appears to be predominantly formed from precursors secreted into the tubular lumen by proximal tubular cells (36).
ANG II secreted by podocytes may well exert autocrine effects. 1) ANG II secreted by podocytes may directly activate respective membrane angiotensin receptors. 2) AT1 receptor-mediated endocytosis of intact ANG II may occur, leading to intracellular ANG II accumulation (36). 3) Intact intracellular ANG II may further regulate gene transcription (41). 4) ANG II might stimulate angiotensinogen mRNA expression under certain conditions (36).
Ex vivo and in vitro studies in rodents revealed functional expression of the AT1 angiotensin receptor in podocytes (11, 13, 16, 39, 46, 49). In some studies, AT2 receptor expression similarly was detected in podocytes (11, 46, 49). We demonstrate by RT-PCR, Western blotting, and immunostaining experiments that differentiated human podocytes express both the AT1 and AT2 angiotensin receptor subtype. As the Cai2+ increase elicited by ANG II in human podocytes was completely blocked by the AT1 receptor antagonist losartan, we conclude from these findings that AT2 receptor activation is not significantly involved in the functional responses to ANG II investigated here. We speculate that in vivo effects of ANG II in podocytes under physiological and pathophysiological conditions are predominantly mediated by AT1 receptor activation.
Recently, ANG II has been reported to induce a concentration-dependent increase of cAMP in rat podocytes, which could only be blocked by simultaneous use of both AT1 and AT2 receptor blockers (46). In patch-clamp experiments of podocytes in intact rat glomeruli, we recently were not able to demonstrate cellular responses to forskolin or 8-(4-chlorophenylthio)-cAMP (13). In the current study, differentiated human podocytes were stimulated either with ANG II alone or in the presence of the phosphodiesterase inhibitor IBMX to exclude rapid cAMP degradation. cAMP production was repetitively measured over a time course of 120 min. In these experiments, we did not observe induction of significant cAMP production by ANG II in differentiated human podocytes. Using the adenylate cyclase activator forskolin in combination with IBMX, marked cAMP production was observed. However, this again was not augmented by addition of ANG II. We conclude from these experiments that ANG II does not induce cAMP production in human podocytes.
Within the plethora of its biological effects, ANG II has been claimed to induce cell growth, proliferation, and apoptosis (9, 48, 51). Here, it has been shown most recently that ANG II enhanced DNA synthesis in differentiated mouse podocytes, an effect that was markedly inhibited by the AT1 receptor antagonist losartan, whereas the AT2 receptor antagonist PD-123319 was without effect (49). In contrast, using a similar model of differentiated mouse podocytes another group reported exogenous ANG II to induce apoptosis (11). Podocyte apoptosis induced by mechanical strain was significantly ameliorated by the AT1 receptor antagonist losartan (11). In the current study, LDH release and cell-sorting experiments did not indicate podocyte damage even on prolonged ANG II stimulation. However, ANG II did increase staurosporine-induced apoptosis. One may therefore speculate that once a pathological, e.g., inflammatory, environment has induced podocyte damage, this may be markedly enhanced by local ANG II. In this context, it is of specific note that intrarenal often exert systemic ANG II concentrations under pathophysiological conditions (45). In these circumstances, a functional RAS in human podocytes may transmit damage to this unique cell type.
Podocytes play a key role in the maintenance of the glomerular filtration barrier. Here, we show for the first time that human podocytes possess a functionally active local RAS. They not only respond to ANG II by activation of AT1 receptors, but are further capable of directly producing ANG II. This RAS appears to be involved not only in physiological mechanisms but also in pathological responses (11). Inhibition of ANG II effects in podocytes might lower podocyte tonus, decrease the rate of apoptosis, and increase glomerular permselectivity (21). Based on these grounds, podocytes constitute an important target for ACE inhibitors and AT1 receptor blockers (5, 8, 19, 25, 30, 32). The combination of these two pharmacological drug classes has recently been shown to result in additive renoprotective effects in both nondiabetic and diabetic disease (33, 35).
Compelling evidence has accumulated that ANG II exerts a number of pathophysiological events in podocytes. Here, nephroprotection by ACE inhibitors and AT1 receptor blockers, in part, can be spelled "podocyto-protection."
GRANTS
This work was supported by Deutsche Forschungsgemeinschaft Grant Pa 483/5–2.
ACKNOWLEDGMENTS
We thank P. Daemisch, C. Meyer, T. Kilic, C. Kocksch, P. Kulick, and B. Weinhold for excellent technical assistance.
FOOTNOTES
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.
H. Pavenstdt and K.-G. Fischer contributed equally to this work.
REFERENCES
Bader M, Peters J, Baltatu O, Muller DN, Luft FC, and Ganten D. Tissue renin-angiotensin systems: new insights from experimental animal models in hypertension research. J Mol Med 79: 76–102, 2001.
Berger ED, Bader BD, Ebert C, Risler T, and Erley CM. Reduction of proteinuria; combined effects of receptor blockade and low dose angiotensin-converting enzyme inhibition. J Hypertens 20: 739–743, 2002.
Blantz RC, Konnen KS, and Tucker BJ. Angiotensin II effects upon the glomerular microcirculation and ultrafiltration coefficient of the rat. J Clin Invest 57: 419–434, 1976.
Bonnet F, Cao Z, and Cooper ME. Apoptosis and angiotensin II: yet another renal regulatory system Exp Nephrol 9: 295–300, 2001.
Bonnet F, Cooper ME, Kawachi H, Allen TJ, Boner G, and Cao Z. Irbesartan normalises the deficiency in glomerular nephrin expression in a model of diabetes and hypertension. Diabetologia 44: 874–877, 2001.
Brenner BM, Cooper ME, de Zeeuw D, Keane WF, Mitch WE, Parving HH, Remuzzi G, Snapinn SM, Zhang Z, and Shahinfar S. Effects of losartan on renal and cardiovascular outcomes in patients with type 2 diabetes and nephropathy. N Engl J Med 345: 861–869, 2001.
Cervenka L, Wang CT, Mitchell KD, and Navar LG. Proximal tubular angiotensin II levels and renal functional responses to AT1 receptor blockade in nonclipped kidneys of Goldblatt hypertensive rats. Hypertension 33: 102–107, 1999.
Davis BJ, Cao Z, de Gasparo M, Kawachi H, Cooper ME, and Allen TJ. Disparate effects of angiotensin II antagonists and calcium channel blockers on albuminuria in experimental diabetes and hypertension: potential role of nephrin. J Hypertens 21: 209–216, 2003.
Dinh DT, Frauman AG, Johnston CI, and Fabiani ME. Angiotensin receptors: distribution, signalling and function. Clin Sci 100: 481–492, 2001.
Doublier S, Salvidio G, Lupia E, Ruotsalainen V, Verzola D, Deferrari G, and Camussi G. Nephrin expression is reduced in human diabetic nephropathy: evidence for a distinct role for glycated albumin and angiotensin II. Diabetes 52: 1023–1030, 2003.
Durvasula RV, Petermann AT, Hiromura K, Blonski M, Pippin J, Mundel P, Pichler R, Griffin S, Couser WG, and Shankland SJ. Activation of a local tissue angiotensin system in podocytes by mechanical strain. Kidney Int 65: 30–39, 2004.
Endlich N, Kress KR, Reiser J, Uttenweiler D, Kriz W, Mundel P, and Endlich K. Podocytes respond to mechanical stress in vitro. J Am Soc Nephrol 12: 413–422, 2001.
Gloy J, Henger A, Fischer KG, Nitschke R, Mundel P, Bleich M, Schollmeyer P, Greger R, and Pavenstdt H. Angiotensin II depolarizes podocytes in the intact glomerulus of the rat. J Clin Invest 99: 2772–2781, 1997.
Greiber S, Münzel T, Kstner S, Müller B, Schollmeyer P, and Pavenstdt H. NAD(P)H oxidase activity in cultured human podocytes: effects of adenosine triphosphate. Kidney Int 53: 654–663, 1998.
Grynkiewicz G, Poenie M, and Tsien RY. A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260: 3340–3350, 1985.
Henger A, Huber T, Fischer KG, Nitschke R, Mundel P, Schollmeyer P, Greger R, and Pavenstdt H. Angiotensin II increases the cytosolic calcium activity in rat podocytes in culture. Kidney Int 52: 687–693, 1997.
Hoffmann S, Podlich D, Hahnel B, Kriz W, and Gretz N. Angiotensin II type 1 receptor overexpression in podocytes induces glomerulosclerosis in transgenic rats. J Am Soc Nephrol 15: 1475–1487, 2004.
Inagami T. Molecular biology and signaling of angiotensin receptors: an overview. J Am Soc Nephrol 10, Suppl 11: S2–S7, 1999.
Kelly DJ, Aaltonen P, Cox AJ, Rumble JR, Langham R, Panagiotopoulos S, Jerums G, Holthofer H, and Gilbert RE. Expression of the slit-diaphragm protein, nephrin, in experimental diabetic nephropathy: differing effects of anti-proteinuric therapies. Nephrol Dial Transplant 17: 1327–1332, 2002.
Kim S and Iwao H. Molecular and cellular mechanisms of angiotensin II-mediated cardiovascular and renal diseases. Pharmacol Rev 52: 11–34, 2000.
Kriz W. Podocytes as a target for treatment with ACE inhibitors and/or angiotensin-receptor blockers. Kidney Int 65: 333–334, 2004.
Kriz W, Elger M, Nagata M, Kretzler M, Uiker S, Koeppen-Hagemann I, Tenschert S, and Lemley KV. The role of podocytes in the development of glomerular sclerosis. Kidney Int 45: S-64–S-72, 1994.
Kriz W, Hackenthal E, Nobiling R, Sakai T, and Elger M. A role for podocytes to counteract capillary wall distension. Kidney Int 45: 369–376, 1994.
Lang D, Dohle F, Terstesse M, Bangen P, August C, Pauels HG, and Heidenreich S. Downregulation of monocyte apoptosis by phagocytosis of platelets: involvement of a caspase-9, caspase-3, and heat shock protein 70-dependent pathway. J Immunol 168: 6152–6158, 2002.
Langham RG, Kelly DJ, Cox AJ, Thomson NM, Holthofer H, Zaoui P, Pinel N, Cordonnier DJ, and Gilbert RE. Proteinuria and the expression of the podocyte slit diaphragm protein, nephrin, in diabetic nephropathy: effects of angiotensin converting enzyme inhibition. Diabetologia 45: 1572–1576, 2002.
Lapinski R, Perico N, Remuzzi A, Sangalli F, Benigni A, and Remuzzi G. Angiotensin II modulates glomerular capillary permselectivity in rat isolated perfused kidney. J Am Soc Nephrol 7: 653–660, 1996.
Laverman GD, Navis G, Henning RH, De Jong PE, and de Zeeuw D. Dual renin-angiotensin system blockade at optimal doses for proteinuria. Kidney Int 62: 1020–1025, 2002.
Lewis EJ, Hunsicker LG, Bain RP, and Rohde RD. The effect of angiotensin-converting-enzyme inhibition on diabetic nephropathy. The Collaborative Study Group. N Engl J Med 329: 1456–1462, 1993.
Lewis EJ, Hunsicker LG, Clarke WR, Berl T, Pohl MA, Lewis JB, Ritz E, Atkins RC, Rohde R, and Raz I. Renoprotective effect of the angiotensin-receptor antagonist irbesartan in patients with nephropathy due to type 2 diabetes. N Engl J Med 345: 851–860, 2001.
Macconi D, Ghilardi M, Bonassi ME, Mohamed EI, Abbate M, Colombi F, Remuzzi G, and Remuzzi A. Effect of angiotensin-converting enzyme inhibition on glomerular basement membrane permeability and distribution of zonula occludens-1 in MWF rats. J Am Soc Nephrol 11: 477–489, 2000.
Maschio G, Alberti D, Janin G, Locatelli F, Mann JF, Motolese M, Ponticelli C, Ritz E, and Zucchelli P. Effect of the angiotensin-converting enzyme inhibitor benazepril on the progression of chronic renal insufficiency. The angiotensin-converting enzyme inhibition in progressive renal insufficiency study group. N Engl J Med 334: 939–945, 1996.
Mifsud SA, Allen TJ, Bertram JF, Hulthen UL, Kelly DJ, Cooper ME, Wilkinson-Berka JL, and Gilbert RE. Podocyte foot process broadening in experimental diabetic nephropathy: amelioration with renin-angiotensin blockade. Diabetologia 44: 878–882, 2001.
Mogensen CE, Neldam S, Tikkanen I, Oren S, Viskoper R, Watts RW, and Cooper ME. Randomised controlled trial of dual blockade of renin-angiotensin system in patients with hypertension, microalbuminuria, and non-insulin dependent diabetes: the candesartan and lisinopril microalbuminuria (CALM) study. BMJ 321: 1440–1444, 2000.
Mundel P, Reiser J, and Kriz W. Induction of differentiation in cultured rat and human podocytes. J Am Soc Nephrol 8: 697–705, 1997.
Nakao N, Yoshimura A, Morita H, Takada M, Kayano T, and Ideura T. Combination treatment of angiotensin-II receptor blocker and angiotensin-converting-enzyme inhibitor in non-diabetic renal disease (COOPERATE): a randomised controlled trial. Lancet 361: 117–124, 2003.
Navar LG, Harrison-Bernard LM, Nishiyama A, and Kobori H. Regulation of intrarenal angiotensin II in hypertension. Hypertension 39: 316–322, 2002.
Nithipatikom K, Holmes BB, McCoy MJ, Hillard CJ, and Campbell WB. Chronic administration of nitric oxide reduces angiotensin II receptor type 1 expression and aldosterone synthesis in zona glomerulosa cells. Am J Physiol Endocrinol Metab 287: E820–E827, 2004.
Nitschke R, Frbe U, and Greger R. Antidiuretic hormone acts via V1 receptors on intracellular calcium in the isolated perfused rabbit cortical thick ascending limb. Pflügers Arch 417: 622–632, 1991.
Nitschke R, Henger A, Ricken S, Gloy J, Müller V, Greger R, and Pavenstdt H. Angiotensin II increases the intracellular calcium activity in podocytes of the intact glomerulus. Kidney Int 57: 41–49, 2000.
Pavenstdt H, Kriz W, and Kretzler M. Cell biology of the glomerular podocyte. Physiol Rev 83: 253–307, 2003.
Re RN. On the biological actions of intracellular angiotensin. Hypertension 35: 1189–1190, 2000.
Remuzzi G and Bertani T. Pathophysiology of progressive nephropathies. N Engl J Med 339: 1448–1456, 1998.
Ruggenenti P, Perna A, Gherardi G, Garini G, Zoccali C, Salvadori M, Scolari F, Schena FP, and Remuzzi G. Renoprotective properties of ACE inhibition in nondiabetic nephropathies with nonnephrotic proteinuria. Lancet 354: 359–364, 1999.
Saleem MA, O'Hare MJ, Reiser J, Coward RJ, Inward CD, Farren T, Xing CY, Ni L, Mathieson PW, and Mundel P. A conditionally immortalized human podocyte cell line demonstrating nephrin and podocin expression. J Am Soc Nephrol 13: 630–638, 2002.
Seikaly MG, Arant BS Jr, and Seney FD Jr. Endogenous angiotensin concentrations in specific intrarenal fluid compartments of the rat. J Clin Invest 86: 1352–1357, 1990.
Sharma M, Sharma R, Greene AS, McCarthy ET, and Savin VJ. Documentation of angiotensin II receptors in glomerular epithelial cells. Am J Physiol Renal Physiol 274: F623–F627, 1998.
Sharma R, Lovell HB, Wiegmann TB, and Savin VJ. Vasoactive substances induce cytoskeletal changes in cultured rat glomerular epithelial cells. J Am Soc Nephrol 3: 1131–1138, 1992.
Volpe M, Savoia C, De Paolis P, Ostrowska B, Tarasi D, and Rubattu S. The renin-angiotensin system as a risk factor and therapeutic target for cardiovascular and renal disease. J Am Soc Nephrol 13, Suppl 3: S173–S178, 2002.
Wang L, Flannery PJ, and Spurney RF. Characterization of angiotensin II receptor subtypes in podocytes. J Lab Clin Med 142: 313–321, 2003.
Wennemuth G and Aumüller G. Angiotensin II-mediated calcium signals and mitogenesis in human prostate stromal cell line hPCPs. Br J Pharmacol 144: 3–10, 2005.
Wolf G, Butzmann U, and Wenzel UO. The renin-angiotensin system and progression of renal disease: from hemodynamics to cell biology. Nephron Physiol 93: 3–13, 2003.(Max C. Liebau, D. Lang, J. Bhm, N. Endli)
2Department of Medicine D, Division of General Internal Medicine and Nephrology, University Hospital Münster, Münster
3Department of Pathology, University of Freiburg, Freiburg
4Department of Anatomy and Cell Biology, University of Heidelberg, Heidelberg, Germany
5Children's Renal Unit and Academic Renal Unit, Southmead Hospital, University of Bristol, Bristol, United Kingdom
ABSTRACT
Experimental and clinical studies impressively demonstrate that angiotensin-converting enzyme inhibitors (ACEI) and angiotensin receptor blockers (ARB) significantly reduce proteinuria and retard progression of glomerular disease. The underlying intraglomerular mechanisms are not yet fully elucidated. As podocyte injury constitutes a critical step in the pathogenesis of glomerular proteinuria, beneficial effects of ACEI and ARB may partially result from interference with a local renin-angiotensin system (RAS) in podocytes. The knowledge of expression and function of a local RAS in podocytes is limited. In this study, we demonstrate functional expression of key components of the RAS in differentiated human podocytes: podocytes express mRNA for angiotensinogen, renin, ACE type 1, and the AT1 and AT2 angiotensin receptor subtypes. In Western blot experiments and immunostainings, expression of the AT1 and AT2 receptor was demonstrated both in differentiated human podocytes and in human kidney cortex. ANG II induced a concentration-dependent increase in cytosolic Ca2+ concentration via AT1 receptors in differentiated human podocytes, whereas it did not increase cAMP. Furthermore, ANG II secretion was detected, which was blocked by neither the ACEI captopril nor the renin inhibitor remikiren nor the chymase inhibitor chymostatin. ANG II secretion of podocytes was not increased by mechanical stress. Finally, ANG II was found to increase staurosporine-induced apoptosis in podocytes. We speculate that ACEI and ARB exert their beneficial effects, in part, by interfering with a local RAS in podocytes. Further experiments are required to identify the underlying molecular mechanism(s) of podocyte protection.
cytosolic Ca2+; human; captopril; AT1 receptor; AT2 receptor; apoptosis; mechanical stress
THE RENIN-ANGIOTENSIN SYSTEM (RAS) plays a crucial role in controlling blood pressure, fluid balance, and electrolyte homeostasis (48, 51). Enzymatic cleavage by renin and angiotensin-converting enzyme (ACE) leads to the conversion of angiotensinogen via ANG I to its main effector molecule ANG II. Apart from its hemodynamic action, ANG II is capable of inducing a multitude of nonhemodynamic effects such as induction of oxygen radicals, cytokines, stimulation of collagen synthesis, apoptosis, proliferation, and hypertrophy (20, 51). Within the glomerulus, ANG II reduces the ultrafiltration coefficient and modulates glomerular capillary permselectivity leading to proteinuria, initiating and increasing subsequent tubulointerstitial injury (3, 26, 42). Acting as growth hormone, ANG II contributes to the pathogenesis of glomerulosclerosis (20).
Clinical studies have impressively demonstrated that ACE inhibitors (ACEI) markedly reduce proteinuria and progression of glomerular disease (28, 31, 43). Furthermore, angiotensin type 1 receptor blockers (ARB) have clearly been shown to retard the progression of diabetic nephropathy (6, 29). Most recently, the addition of ARB to ACEI treatment further reduced proteinuria (2, 27).
The podocyte is the most differentiated cell type within the glomerulus. Synthesizing various components of the glomerular basement membrane and the slit diaphragm, it is crucially involved in the maintenance of the glomerular filtration barrier (40). Podocyte injury leads to alterations of the foot process/slit diaphragm complex with loss of glomerular permselectivity, resulting in proteinuria (40).
Interestingly, little is known about the expression and function of the RAS in podocytes. In this regard, one may hypothesize that the antiproteinuric effects of ACEI or ARB may partially result from interference with a local RAS in podocytes. This prompted us to specifically investigate expression and function of components of the RAS in differentiated human podocytes.
METHODS
Cell culture. The conditionally immortalized human podocyte cell line has been developed as recently described by transfection with the temperature-sensitive SV40 T-gene (44). Proliferating at a temperature of 33°C, these cells transform into a quiescent, differentiated phenotype after transfer to 37°C. Cells then stain positive for the in vivo podocyte markers synaptopodin, nephrin, podocin, CD2AP, ZO-1, -, -, -catenins, and P-cadherin (44). For experiments, cells between passage 10 and 18 were seeded at 37°C into 6-well plates, 24-well plates, or flasks and cultured in standard RPMI media containing 10% fetal calf serum (Biochrom, Berlin, Germany), 2.5 mM L-glutamin, 0.1 mM sodium pyruvate, 1 g/l nonessential amino acids (all Seromed, Berlin, Germany), and insulin-transferrin-sodium selenite supplement (Boehringer, Mannheim, Germany), 100 U/ml penicillin, and 100 mg/ml streptomycin (GIBCO, Eggenstein, Germany) for at least 7 days until cells were differentiated showing an arborized morphology.
RT-PCR for components of the RAS. The RNA preparation, the reverse transcription, and the PCR were performed according to the method recently described (14). In brief, the total RNA from cultured human podocytes was isolated with guanidinium/acid phenol/chloroform extraction and the amount of RNA was measured with spectrophotometry. For first-strand synthesis, total RNA from podocytes was mixed in 5x reverse transcription buffer and completed with 0.5 mM dNTP, 10 μM random hexanucleotide primer, 10 mM dithiothreitol, Moloney murine leukemia virus 0.02 U RNAse inhibitor/ng RNA, and 100 U MMLV reverse transcriptase/μg RNA. PCR amplification of RT reactions without reverse transcriptase revealed no PCR product, thereby excluding amplification of genomic DNA. RT was performed at 42°C for 1 h, followed by a denaturation at 95°C for 5 min. PCR was performed in duplicate with a total volume of 20 μl, each containing 40 ng RNA per 4 μl of RT template, 16 μl of PCR master mixture, and 10 pmol each of sense and antisense primer. The mixture was overlaid with mineral oil and heated for 2 min at 94°C. The cycle profile consisted of 1 min of denaturation at 94°C, annealing for 1 min at 60°C (renin: 58°C; ACE1: 59°C), and extension for 1 min at 72°C. For amplification, different numbers of cycles were used (see Table 1). The amplification products of 10 μl of each PCR reaction were separated on a 1.5% agarose gel, stained with ethidium bromide, and visualized by UV irradiation. The primer oligonucleotides were selected from published cDNA sequences and are depicted in Table 1. All primers are noted in 5'-3' direction.
Immunohistochemistry of kidney cortex. Normal kidney cortex was obtained from nephrectomy specimens. Fixation and preparation of tissue for immunohistochemical analyses were performed using standard techniques. In brief, kidneys were perfused after removal with cold (4°C) PBS followed by 4% formaldehyde. Kidneys were then incubated for 24 h at 4°C in 4% formaldehyde solution, embedded in paraffin, and cut into 4-μm-thick slices. Slices were deparaffinized in xylol for 1 h, gradually hydrated through graded alcohols (100 to 70%), and washed in deionized water. After incubation in 1% H2O2 for 30 min, slices were rehydrated with PBS, and antigen unmasking was performed by boiling slices in 10 mM citrate buffer (pH 6) for AT1 and WT1 or EDTA (pH 9) for AT2 for 2 x 10 min. Blocking was performed using a 1% BSA solution for 10 min.
For staining of WT1 and AT1, sections were then incubated overnight in a humidified chamber at 4°C with antibodies (Abs) to the angiotensin AT1 receptor (rabbit anti-human, 200 μg/ml, 1:200 dilution; Santa Cruz Biotechnology, Santa Cruz, CA) and WT1 (cat anti-human, 100 μg/ml, 1:100 dilution; Santa Cruz Biotechnology). The slices were washed extensively with PBS and incubated for 90 min with a secondary antibody using a commercially available ABC-Kit (Vectastain Mouse Peroxidase, Vectorlabs, Burlingame, CA). Slices were washed with PBS, incubated with avidin-biotin for 45 min, and stained with 3-amino-9 ethylcarbazole (red) or fast blue BB salt/naphtol AS-MX phosphate (blue; Sigma, Steinheim, Germany). Sections were examined with a conventional light microscope (Zeiss LSM 510; Zeiss, Oberkochen, Germany) and photographed using a digital camera. Negative controls were performed by elimination of the primary Abs. For "double-labeling" experiments, serial ultrathin sections of human kidneys were stained with Abs to the angiotensin AT1 receptor and WT1 and pictures of the same glomerulus were projected on top of each other using commercially available computer programs.
For staining of WT1 and AT2, sections were incubated for 30 min (WT1) or 60 min (AT2) in a humidified chamber at 4°C with Abs to the angiotensin AT2 receptor (goat anti-human, 200 μg/ml, 1:50, Santa Cruz Biotechnology) and WT1 (mouse anti-human, 200 μg/ml, 1:100 dilution; Santa Cruz Biotechnology). AT2 was then detected using a labeled streptavidin-biotin detection kit (ChemMate Detection Kit, Alkaline Phosphatase/Red, Code K 5005, Dako; red), and WT1 was detected using an ABC Kit (StreptABComplex/HRP Duet, Code K 0492, Dako, Hamburg, Germany; brown). All sections were then also stained using Mayer's hemalaun. Sections were examined with a conventional light microscope (Zeiss Axioskop 50) and photographed using a digital camera (Sony MC-3210/II/PM). Negative controls were performed by elimination of the primary Abs.
Immunocytochemistry of cultured human podocytes. Immunolabeling was done as previously described (34). Briefly, coverslips were fixed with 2% paraformaldehyde and 4% sucrose in PBS for 10 min and were then permeabilized with 0.3% Triton X-100 (Sigma-Aldrich, Munich, Germany) in PBS for 10 min. Nonspecific binding sites were blocked with 2% FCS, 2% bovine serum albumin, and fish gelatine (Sigma-Aldrich) in PBS for 30 min. Primary (rabbit anti-AT1 and rabbit anti-AT2, both Santa Cruz Biotechnology) and secondary antibodies were applied in a 1:100 dilution according to standard techniques, and the coverslips were mounted on glass slides with 15% Mowiol (Calbiochem, La Jolla, CA) and 50% glycerol in PBS. After being washed with PBS, bound primary Abs were detected using FITC-conjugated cat anti-rabbit IgG (Santa Cruz Biotechnology). As controls, primary Abs were either omitted or boiled at 99°C for 10 min and used thereafter. Micrographs were taken on a microscope equipped for epiluminescence (Zeiss Axiophot).
Western blotting. Western blotting was performed using standard techniques. In brief, cells were washed once with PBS, scraped with lysis buffer containing 2 mM EDTA, 2 mM EGTA, 100 mM NaCl, 20 mM Tris, 0.1% SDS, 1% NP-40, 2 mM PMSF, and proteinase inhibitor cocktail (Roche Diagnostics, Mannheim, Germany), and sonicated. The samples were resuspended in Laemmli sample buffer, boiled (5 min), and subjected to SDS-PAGE and transfer electrophoresis. The transblots were stained with Ponceau solution to prove equal amounts of protein were loaded on the membrane and probed with primary Abs (rabbit anti-AT1 and goat anti-AT2, both either with or without blocking peptide 5:1; both Santa Cruz Biotechnology) followed by peroxidase-labeled secondary antibodies (donkey anti-rabbit, 1:4,000; Amersham Pharmacia, Piscataway, NJ; rabbit anti-goat, Dako) and detected by chemiluminescence detection reagents (ECL, Amersham Pharmacia).
Measurements of the intracellular calcium activity. Measurements of cytosolic calcium (Cai2+) with the Ca2+-sensitive dye fura 2 (Sigma-Aldrich) were performed in podocytes on an inverted fluorescence microscope setup (38). The system allows fluorescence measurements at the single-cell level at three excitation wavelengths. The field of measurement can be set between a diameter of 2 and 300 μm with an adjustable pinhole. A time resolution of up to 200 Hz was achieved by using a high-speed filter wheel and a single-photon counting tube (Hamamatsu H63460 [GenBank] –04; Hamamatsu, Herrsching, Germany). The autofluorescence signal of cells that had not been loaded with fura 2 was measured and subtracted from the results obtained in fura 2-loaded cells. This had no effect on the bandwidth of the measurements. A calibration of the fura 2 fluorescence signal was attempted at the end of each experiment by using Ca2+ ionophore ionomycin (1 μM) and low- and high-Ca2+ buffers. Cai2+ concentration ([Ca2+]i) was calculated from the fluorescence ratio 340:380 nm according to the equation described by Grynkiewicz et al. (15).
Measurements of intracellular cAMP. Podocytes were cultured in six-well plates. They were kept at 37°C and rinsed with a physiological Ringer solution. After preincubation with 0.5 M 3-isobutyl-I-methylxanthine (IBMX; RBI, Cologne, Germany) for 5 min, cells were exposed to the added agents. To terminate the assay, the supernatants were rapidly removed and cells were rinsed with ice-cold ethanol. After an ethanol extraction, cAMP concentrations were measured with an ELISA (Amersham Buchler, Braunschweig, Germany).
Measurements of ANG II secretion. Podocytes were cultured in six-well plates. They were kept at 37°C, and the medium was changed from 10 to 0% FCS. The cells were then incubated either with vehicle or with remikiren (a kind gift of Dr. D. Müller, Berlin, Germany), captopril, chymostatin (both Sigma), or a combination of the three. After 24 h, the supernatant was taken and centrifuged at 10,000 rpm for 5 min. Concentrations were then measured with an ELISA (Spibio, Massy Cedex, France).
Mechanical stress experiments. Differentiated human podocytes were seeded in six-well plates with a flexible bottom (Bioflex, Flexcell International, Hillsborough, NC). The flexible silicone membranes were coated with collagen IV to facilitate cell attachment. After 3 days, the six-well plate was mounted on a Plexiglas manifold connected to a custom-built apparatus, which induced cyclic variations in air pressure below and above atmospheric pressure. Cyclic pressure variations caused upward and downward motion of the silicone membranes. Pressure amplitude was chosen to give a maximum up- and downward deflection of the membrane center of 6 mm, being equal to an increase in membrane area by 11%, or 5% mean linear cell strain. Cycle frequency was adjusted to 0.5 Hz.
Measurements of lactate dehydrogenase release from podocytes. To assess cytotoxicity, lactate dehydrogenase (LDH) release was measured with a routine autoanalyzer (Modular I, Hitachi, Tokyo, Japan). Total LDH content of the cells was measured after incubation of cells in 1% Triton X-100.
Quantification of apoptosis and necrosis by flow cytometry. Apoptosis was determined by detecting annexin V binding using flow cytometry as described previously (24). Evaluation of cell necrosis was performed by simultaneous detection of propidium iodide (PI) uptake by the nonpermeabilized cells. Immortalized podocytes, after 24-h incubation with the indicated stimuli, were labeled with annexin V-FITC (Bender Medsystems, Vienna, Austria) and PI (5 μg/ml) in staining buffer (containing 1% BSA in 50 mM HEPES buffer, pH 7.4) for 15 min on ice. After being stained, cells were washed in PBS and fixed in 4% paraformaldehyde to apply flow cytometry. Cells were analyzed on a FACScan flow cytometer (Becton Dickinson, Mountain View, CA).
TUNEL procedure. DNA strand breaks were identified by TUNEL (TdT-mediated dUTP nick-end labeling) reaction using an in situ cell death detection kit (Roche) according to the manufacturer's instructions. First, podocytes were cultured on coverslips incubated for indicated times, then washed with PBS fixed in 4% paraformaldehyde at room temperature for 30 min. Subsequently, the cytoplasmic membrane was permeabilized with 0.1% Triton X-100 for 2 min on ice. After being washed two times, cells were incubated with TUNEL reaction mixture for 60 min at 37°C. Samples were directly analyzed under a fluorescence microscope by detection at 550 nm. Cells labeled in the absence of TdT were always used as negative controls, whereas cells pretreated with DNase after permeabilization were used as positive controls.
Chemicals. Unless otherwise indicated, chemicals were purchased from Sigma.
Statistical analyses. Data are given as means ± SE, where n refers to the number of experiments. Student's t-test was used to compare mean values within one experimental series. A P value 0.05 was considered statistically significant.
RESULTS
mRNA expression of key components of the RAS in differentiated human podocytes. Differentiated human podocytes express mRNA for angiotensinogen, renin, ACE type 1, and both the AT1 and AT2 angiotensin receptor subtypes (Fig. 1). PCR primers and conditions are depicted in Table 1.
AT receptor expression in differentiated human podocytes. Western blot analyses consistently revealed protein expression of both the AT1 and AT2 receptor in human podocytes (AT1 receptor: Fig. 2A, lanes 1 and 2; AT2 receptor: Fig. 2B, lanes 1 and 2), the specificity of which was confirmed using respective blocking peptides (AT1 receptor: Fig. 2A, lanes 3 and 4; AT2 receptor: Fig. 2B, lanes 3 and 4). In additional immunofluorescence experiments, expression of AT1 and AT2 receptors was detected (Fig. 3). Here, expression of the AT1 receptor was more prominent than AT2 receptor expression. Immunostainings most often detected angiotensin receptors as intracellular clusters. Similar expression patterns of angiotensin receptors have also been recently shown in various other cell types (37, 50).
Angiotensin receptor expression in human kidney cortex. Merged pictures of serial sections of human kidney cortex stained with antibodies against AT1 receptors (blue) and the podocyte-specific protein WT1 (red) demonstrate colocalization of AT1 receptors and WT1 in human podocytes ex vivo (Fig. 4, A-C).
In separate experiments, AT2 receptor expression was detected. Double stainings of human kidney cortex revealed colocalization of AT2 receptors (red) and WT1 (brown) in human podocytes ex vivo (Fig. 4, D-E).
ANG II increases [Cai2+] in differentiated human podocytes via AT1 receptors. In microfluorescence experiments, ANG II reversibly increased [Ca2+]i in podocytes (original experiment, Fig. 5A). This [Ca2+]i increase was concentration dependent (Fig. 5B; EC: 60 nM). The ANG II-induced [Ca2+]i increase was reversibly blocked by the AT1 receptor antagonist losartan (Fig. 5C).
ANG II does not increase the cAMP concentration in differentiated human podocytes. Recently, it has been reported that ANG II elevates the cAMP concentration in rat glomerular epithelial cells in culture (46, 47). To address this issue in differentiated human podocytes, cAMP concentration was measured in separate experiments. Within a time course of 2 h, ANG II (1 μM) did not increase cAMP production in human podocytes (Fig. 6A). To exclude rapid cAMP degradation by phosphodiesterases during the experiments, the specific phosphodiesterase inhibitor IBMX (500 μM) was added. Again, no elevation of cAMP levels was observed (Fig. 6B). In the presence of IBMX, the membrane-permeable adenylate cyclase activator forskolin concentration dependently increased cAMP production. In this setting, ANG II had no additive effect (Fig. 6C).
Differentiated human podocytes secrete ANG II. After having detected mRNA expression of key components of a local RAS, we were interested in secretion of ANG II as the final RAS effector by differentiated human podocytes. As detected by ELISA, human podocytes secrete ANG II (Fig. 7A). However, neither the specific renin inhibitor remikiren (10 μM), the ACE inhibitor captopril (10 μM), the chymase inhibitor chymostatin (100 μM), nor a combination of the three inhibitors significantly reduced ANG II secretion of podocytes (Fig. 7A). Furthermore, ANG II secretion of podocytes was not altered by mechanical stress (Fig. 7B).
ANG II does not increase LDH release in differentiated human podocytes. To test for podocyte damage induced by ANG II, LDH release experiments were performed. Even on prolonged stimulation at high ANG II concentration (24 and 48 h, 1 μM), LDH release was not significantly increased (n = 6–12; Fig. 8).
ANG II increases staurosporine-induced apoptosis in differentiated human podocytes. As measured by flow cytometry, staurosporine (1 μM, 24 h) induced apoptosis of differentiated human podocytes. Stimulation of podocytes with ANG II (1 μM) in addition to staurosporine increased the rate of apoptosis induced by staurosporine only (1 μM). However, stimulation with ANG II only did not alter the rate of apoptosis (Fig. 9). These findings were confirmed by TUNEL assay (data not shown).
DISCUSSION
Based on both experimental and clinical studies, there is clear evidence that the RAS plays a pivotal role in the pathophysiology of cardiovascular and renal disease (48, 51). Furthermore, to its prominent hemodynamic actions, the key effector ANG II causes numerous nonhemodynamic renal effects: ANG II is a growth factor of glomerular mesangial and endothelial cells, tubular epithelial cells, and fibroblasts (18, 51). ANG II causes cellular hypertrophy (51) and induces apoptosis (4). ANG II increases the expression of TGF- and stimulates synthesis of extracellular matrix proteins such as collagen type IV (18, 51). ANG II acts as proinflammatory cytokine and promotes oxidative stress (48, 51). Given this multitude of ANG II effects, local angiotensin-generating systems are of special interest (1).
Within the glomerulus, podocytes are critically involved in maintaining the glomerular filtration barrier and in counteracting glomerular capillary wall distension (23, 40). Podocyte damage leads to proteinuria and initiates glomerulosclerosis, finally resulting in progressive loss of kidney function (22, 40).
Indirect evidence suggests that ANG II is capable of disturbing podocyte biology. Spontaneous proteinuria in Munich- Wistar-Froemter rats has been attributed to an altered distribution of the slit diaphragm protein zonula occludens-1, both of which were prevented by the ACEI lisinopril (30). In experimental diabetic nephropathy, podocyte foot process broadening was attenuated by both the ACEI ramipril and the ARB valsartan (32). In animal models of diabetic nephropathy, reduction of nephrin expression was attenuated by the ACEI perindopril (19) and by the ARB valsartan and irbesartan (5, 8). Selective overexpression of the AT1 receptor in podocytes in rats has most recently been reported to lead to protein leakage and structural podocyte damage, progressing to glomerulosclerosis (17). In a recent renal biopsy study, placebo-treated patients with diabetic nephropathy showed a marked reduction of glomerular nephrin expression compared with nondiabetic control subjects, whereas nephrin expression in patients treated with the ACEI perindopril was similar to the control group (25). Another renal biopsy study revealed reduction of glomerular nephrin expression in both type 1 and type 2 diabetic patients compared with controls (10). These findings suggest direct rather than indirect ANG II effects on podocytes, which furthermore are independent of its hemodynamic actions.
We show for the first time that differentiated human podocytes express all RAS components required to generate ANG II. Furthermore, secretion of ANG II by resting human podocytes was observed. ANG II production has most recently been reported to occur in primary mouse podocytes exposed to mechanical strain (11). We exposed podocytes to mechanical strain under conditions known to affect podocytes (12) but did not find a significant rise in ANG II secretion. It is of note that ANG II production in both studies was not inhibited by the ACEI captopril, suggesting predominant involvement of non-ACE pathways (9) to generate ANG II in podocytes. Similar findings have been reported in both renal and extrarenal tissues (1). As neither a renin inhibitor nor a chymase inhibitor significantly changed ANG II secretion, in podocytes ANG II seems to be produced via an as yet unidentified alternative pathway.
Within the proximal tubular fluid, ANG II concentrations of 6–10 pmol/ml have been measured (7, 36). These high concentrations cannot be explained by filtration of circulating ANG II (36, 45). Given the data provided here, ANG II secretion by podocytes may contribute to the high intratubular ANG II concentration. The extent of this contribution remains to be estimated using a suitable experimental approach. However, ANG II in the tubular fluid appears to be predominantly formed from precursors secreted into the tubular lumen by proximal tubular cells (36).
ANG II secreted by podocytes may well exert autocrine effects. 1) ANG II secreted by podocytes may directly activate respective membrane angiotensin receptors. 2) AT1 receptor-mediated endocytosis of intact ANG II may occur, leading to intracellular ANG II accumulation (36). 3) Intact intracellular ANG II may further regulate gene transcription (41). 4) ANG II might stimulate angiotensinogen mRNA expression under certain conditions (36).
Ex vivo and in vitro studies in rodents revealed functional expression of the AT1 angiotensin receptor in podocytes (11, 13, 16, 39, 46, 49). In some studies, AT2 receptor expression similarly was detected in podocytes (11, 46, 49). We demonstrate by RT-PCR, Western blotting, and immunostaining experiments that differentiated human podocytes express both the AT1 and AT2 angiotensin receptor subtype. As the Cai2+ increase elicited by ANG II in human podocytes was completely blocked by the AT1 receptor antagonist losartan, we conclude from these findings that AT2 receptor activation is not significantly involved in the functional responses to ANG II investigated here. We speculate that in vivo effects of ANG II in podocytes under physiological and pathophysiological conditions are predominantly mediated by AT1 receptor activation.
Recently, ANG II has been reported to induce a concentration-dependent increase of cAMP in rat podocytes, which could only be blocked by simultaneous use of both AT1 and AT2 receptor blockers (46). In patch-clamp experiments of podocytes in intact rat glomeruli, we recently were not able to demonstrate cellular responses to forskolin or 8-(4-chlorophenylthio)-cAMP (13). In the current study, differentiated human podocytes were stimulated either with ANG II alone or in the presence of the phosphodiesterase inhibitor IBMX to exclude rapid cAMP degradation. cAMP production was repetitively measured over a time course of 120 min. In these experiments, we did not observe induction of significant cAMP production by ANG II in differentiated human podocytes. Using the adenylate cyclase activator forskolin in combination with IBMX, marked cAMP production was observed. However, this again was not augmented by addition of ANG II. We conclude from these experiments that ANG II does not induce cAMP production in human podocytes.
Within the plethora of its biological effects, ANG II has been claimed to induce cell growth, proliferation, and apoptosis (9, 48, 51). Here, it has been shown most recently that ANG II enhanced DNA synthesis in differentiated mouse podocytes, an effect that was markedly inhibited by the AT1 receptor antagonist losartan, whereas the AT2 receptor antagonist PD-123319 was without effect (49). In contrast, using a similar model of differentiated mouse podocytes another group reported exogenous ANG II to induce apoptosis (11). Podocyte apoptosis induced by mechanical strain was significantly ameliorated by the AT1 receptor antagonist losartan (11). In the current study, LDH release and cell-sorting experiments did not indicate podocyte damage even on prolonged ANG II stimulation. However, ANG II did increase staurosporine-induced apoptosis. One may therefore speculate that once a pathological, e.g., inflammatory, environment has induced podocyte damage, this may be markedly enhanced by local ANG II. In this context, it is of specific note that intrarenal often exert systemic ANG II concentrations under pathophysiological conditions (45). In these circumstances, a functional RAS in human podocytes may transmit damage to this unique cell type.
Podocytes play a key role in the maintenance of the glomerular filtration barrier. Here, we show for the first time that human podocytes possess a functionally active local RAS. They not only respond to ANG II by activation of AT1 receptors, but are further capable of directly producing ANG II. This RAS appears to be involved not only in physiological mechanisms but also in pathological responses (11). Inhibition of ANG II effects in podocytes might lower podocyte tonus, decrease the rate of apoptosis, and increase glomerular permselectivity (21). Based on these grounds, podocytes constitute an important target for ACE inhibitors and AT1 receptor blockers (5, 8, 19, 25, 30, 32). The combination of these two pharmacological drug classes has recently been shown to result in additive renoprotective effects in both nondiabetic and diabetic disease (33, 35).
Compelling evidence has accumulated that ANG II exerts a number of pathophysiological events in podocytes. Here, nephroprotection by ACE inhibitors and AT1 receptor blockers, in part, can be spelled "podocyto-protection."
GRANTS
This work was supported by Deutsche Forschungsgemeinschaft Grant Pa 483/5–2.
ACKNOWLEDGMENTS
We thank P. Daemisch, C. Meyer, T. Kilic, C. Kocksch, P. Kulick, and B. Weinhold for excellent technical assistance.
FOOTNOTES
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.
H. Pavenstdt and K.-G. Fischer contributed equally to this work.
REFERENCES
Bader M, Peters J, Baltatu O, Muller DN, Luft FC, and Ganten D. Tissue renin-angiotensin systems: new insights from experimental animal models in hypertension research. J Mol Med 79: 76–102, 2001.
Berger ED, Bader BD, Ebert C, Risler T, and Erley CM. Reduction of proteinuria; combined effects of receptor blockade and low dose angiotensin-converting enzyme inhibition. J Hypertens 20: 739–743, 2002.
Blantz RC, Konnen KS, and Tucker BJ. Angiotensin II effects upon the glomerular microcirculation and ultrafiltration coefficient of the rat. J Clin Invest 57: 419–434, 1976.
Bonnet F, Cao Z, and Cooper ME. Apoptosis and angiotensin II: yet another renal regulatory system Exp Nephrol 9: 295–300, 2001.
Bonnet F, Cooper ME, Kawachi H, Allen TJ, Boner G, and Cao Z. Irbesartan normalises the deficiency in glomerular nephrin expression in a model of diabetes and hypertension. Diabetologia 44: 874–877, 2001.
Brenner BM, Cooper ME, de Zeeuw D, Keane WF, Mitch WE, Parving HH, Remuzzi G, Snapinn SM, Zhang Z, and Shahinfar S. Effects of losartan on renal and cardiovascular outcomes in patients with type 2 diabetes and nephropathy. N Engl J Med 345: 861–869, 2001.
Cervenka L, Wang CT, Mitchell KD, and Navar LG. Proximal tubular angiotensin II levels and renal functional responses to AT1 receptor blockade in nonclipped kidneys of Goldblatt hypertensive rats. Hypertension 33: 102–107, 1999.
Davis BJ, Cao Z, de Gasparo M, Kawachi H, Cooper ME, and Allen TJ. Disparate effects of angiotensin II antagonists and calcium channel blockers on albuminuria in experimental diabetes and hypertension: potential role of nephrin. J Hypertens 21: 209–216, 2003.
Dinh DT, Frauman AG, Johnston CI, and Fabiani ME. Angiotensin receptors: distribution, signalling and function. Clin Sci 100: 481–492, 2001.
Doublier S, Salvidio G, Lupia E, Ruotsalainen V, Verzola D, Deferrari G, and Camussi G. Nephrin expression is reduced in human diabetic nephropathy: evidence for a distinct role for glycated albumin and angiotensin II. Diabetes 52: 1023–1030, 2003.
Durvasula RV, Petermann AT, Hiromura K, Blonski M, Pippin J, Mundel P, Pichler R, Griffin S, Couser WG, and Shankland SJ. Activation of a local tissue angiotensin system in podocytes by mechanical strain. Kidney Int 65: 30–39, 2004.
Endlich N, Kress KR, Reiser J, Uttenweiler D, Kriz W, Mundel P, and Endlich K. Podocytes respond to mechanical stress in vitro. J Am Soc Nephrol 12: 413–422, 2001.
Gloy J, Henger A, Fischer KG, Nitschke R, Mundel P, Bleich M, Schollmeyer P, Greger R, and Pavenstdt H. Angiotensin II depolarizes podocytes in the intact glomerulus of the rat. J Clin Invest 99: 2772–2781, 1997.
Greiber S, Münzel T, Kstner S, Müller B, Schollmeyer P, and Pavenstdt H. NAD(P)H oxidase activity in cultured human podocytes: effects of adenosine triphosphate. Kidney Int 53: 654–663, 1998.
Grynkiewicz G, Poenie M, and Tsien RY. A new generation of Ca2+ indicators with greatly improved fluorescence properties. J Biol Chem 260: 3340–3350, 1985.
Henger A, Huber T, Fischer KG, Nitschke R, Mundel P, Schollmeyer P, Greger R, and Pavenstdt H. Angiotensin II increases the cytosolic calcium activity in rat podocytes in culture. Kidney Int 52: 687–693, 1997.
Hoffmann S, Podlich D, Hahnel B, Kriz W, and Gretz N. Angiotensin II type 1 receptor overexpression in podocytes induces glomerulosclerosis in transgenic rats. J Am Soc Nephrol 15: 1475–1487, 2004.
Inagami T. Molecular biology and signaling of angiotensin receptors: an overview. J Am Soc Nephrol 10, Suppl 11: S2–S7, 1999.
Kelly DJ, Aaltonen P, Cox AJ, Rumble JR, Langham R, Panagiotopoulos S, Jerums G, Holthofer H, and Gilbert RE. Expression of the slit-diaphragm protein, nephrin, in experimental diabetic nephropathy: differing effects of anti-proteinuric therapies. Nephrol Dial Transplant 17: 1327–1332, 2002.
Kim S and Iwao H. Molecular and cellular mechanisms of angiotensin II-mediated cardiovascular and renal diseases. Pharmacol Rev 52: 11–34, 2000.
Kriz W. Podocytes as a target for treatment with ACE inhibitors and/or angiotensin-receptor blockers. Kidney Int 65: 333–334, 2004.
Kriz W, Elger M, Nagata M, Kretzler M, Uiker S, Koeppen-Hagemann I, Tenschert S, and Lemley KV. The role of podocytes in the development of glomerular sclerosis. Kidney Int 45: S-64–S-72, 1994.
Kriz W, Hackenthal E, Nobiling R, Sakai T, and Elger M. A role for podocytes to counteract capillary wall distension. Kidney Int 45: 369–376, 1994.
Lang D, Dohle F, Terstesse M, Bangen P, August C, Pauels HG, and Heidenreich S. Downregulation of monocyte apoptosis by phagocytosis of platelets: involvement of a caspase-9, caspase-3, and heat shock protein 70-dependent pathway. J Immunol 168: 6152–6158, 2002.
Langham RG, Kelly DJ, Cox AJ, Thomson NM, Holthofer H, Zaoui P, Pinel N, Cordonnier DJ, and Gilbert RE. Proteinuria and the expression of the podocyte slit diaphragm protein, nephrin, in diabetic nephropathy: effects of angiotensin converting enzyme inhibition. Diabetologia 45: 1572–1576, 2002.
Lapinski R, Perico N, Remuzzi A, Sangalli F, Benigni A, and Remuzzi G. Angiotensin II modulates glomerular capillary permselectivity in rat isolated perfused kidney. J Am Soc Nephrol 7: 653–660, 1996.
Laverman GD, Navis G, Henning RH, De Jong PE, and de Zeeuw D. Dual renin-angiotensin system blockade at optimal doses for proteinuria. Kidney Int 62: 1020–1025, 2002.
Lewis EJ, Hunsicker LG, Bain RP, and Rohde RD. The effect of angiotensin-converting-enzyme inhibition on diabetic nephropathy. The Collaborative Study Group. N Engl J Med 329: 1456–1462, 1993.
Lewis EJ, Hunsicker LG, Clarke WR, Berl T, Pohl MA, Lewis JB, Ritz E, Atkins RC, Rohde R, and Raz I. Renoprotective effect of the angiotensin-receptor antagonist irbesartan in patients with nephropathy due to type 2 diabetes. N Engl J Med 345: 851–860, 2001.
Macconi D, Ghilardi M, Bonassi ME, Mohamed EI, Abbate M, Colombi F, Remuzzi G, and Remuzzi A. Effect of angiotensin-converting enzyme inhibition on glomerular basement membrane permeability and distribution of zonula occludens-1 in MWF rats. J Am Soc Nephrol 11: 477–489, 2000.
Maschio G, Alberti D, Janin G, Locatelli F, Mann JF, Motolese M, Ponticelli C, Ritz E, and Zucchelli P. Effect of the angiotensin-converting enzyme inhibitor benazepril on the progression of chronic renal insufficiency. The angiotensin-converting enzyme inhibition in progressive renal insufficiency study group. N Engl J Med 334: 939–945, 1996.
Mifsud SA, Allen TJ, Bertram JF, Hulthen UL, Kelly DJ, Cooper ME, Wilkinson-Berka JL, and Gilbert RE. Podocyte foot process broadening in experimental diabetic nephropathy: amelioration with renin-angiotensin blockade. Diabetologia 44: 878–882, 2001.
Mogensen CE, Neldam S, Tikkanen I, Oren S, Viskoper R, Watts RW, and Cooper ME. Randomised controlled trial of dual blockade of renin-angiotensin system in patients with hypertension, microalbuminuria, and non-insulin dependent diabetes: the candesartan and lisinopril microalbuminuria (CALM) study. BMJ 321: 1440–1444, 2000.
Mundel P, Reiser J, and Kriz W. Induction of differentiation in cultured rat and human podocytes. J Am Soc Nephrol 8: 697–705, 1997.
Nakao N, Yoshimura A, Morita H, Takada M, Kayano T, and Ideura T. Combination treatment of angiotensin-II receptor blocker and angiotensin-converting-enzyme inhibitor in non-diabetic renal disease (COOPERATE): a randomised controlled trial. Lancet 361: 117–124, 2003.
Navar LG, Harrison-Bernard LM, Nishiyama A, and Kobori H. Regulation of intrarenal angiotensin II in hypertension. Hypertension 39: 316–322, 2002.
Nithipatikom K, Holmes BB, McCoy MJ, Hillard CJ, and Campbell WB. Chronic administration of nitric oxide reduces angiotensin II receptor type 1 expression and aldosterone synthesis in zona glomerulosa cells. Am J Physiol Endocrinol Metab 287: E820–E827, 2004.
Nitschke R, Frbe U, and Greger R. Antidiuretic hormone acts via V1 receptors on intracellular calcium in the isolated perfused rabbit cortical thick ascending limb. Pflügers Arch 417: 622–632, 1991.
Nitschke R, Henger A, Ricken S, Gloy J, Müller V, Greger R, and Pavenstdt H. Angiotensin II increases the intracellular calcium activity in podocytes of the intact glomerulus. Kidney Int 57: 41–49, 2000.
Pavenstdt H, Kriz W, and Kretzler M. Cell biology of the glomerular podocyte. Physiol Rev 83: 253–307, 2003.
Re RN. On the biological actions of intracellular angiotensin. Hypertension 35: 1189–1190, 2000.
Remuzzi G and Bertani T. Pathophysiology of progressive nephropathies. N Engl J Med 339: 1448–1456, 1998.
Ruggenenti P, Perna A, Gherardi G, Garini G, Zoccali C, Salvadori M, Scolari F, Schena FP, and Remuzzi G. Renoprotective properties of ACE inhibition in nondiabetic nephropathies with nonnephrotic proteinuria. Lancet 354: 359–364, 1999.
Saleem MA, O'Hare MJ, Reiser J, Coward RJ, Inward CD, Farren T, Xing CY, Ni L, Mathieson PW, and Mundel P. A conditionally immortalized human podocyte cell line demonstrating nephrin and podocin expression. J Am Soc Nephrol 13: 630–638, 2002.
Seikaly MG, Arant BS Jr, and Seney FD Jr. Endogenous angiotensin concentrations in specific intrarenal fluid compartments of the rat. J Clin Invest 86: 1352–1357, 1990.
Sharma M, Sharma R, Greene AS, McCarthy ET, and Savin VJ. Documentation of angiotensin II receptors in glomerular epithelial cells. Am J Physiol Renal Physiol 274: F623–F627, 1998.
Sharma R, Lovell HB, Wiegmann TB, and Savin VJ. Vasoactive substances induce cytoskeletal changes in cultured rat glomerular epithelial cells. J Am Soc Nephrol 3: 1131–1138, 1992.
Volpe M, Savoia C, De Paolis P, Ostrowska B, Tarasi D, and Rubattu S. The renin-angiotensin system as a risk factor and therapeutic target for cardiovascular and renal disease. J Am Soc Nephrol 13, Suppl 3: S173–S178, 2002.
Wang L, Flannery PJ, and Spurney RF. Characterization of angiotensin II receptor subtypes in podocytes. J Lab Clin Med 142: 313–321, 2003.
Wennemuth G and Aumüller G. Angiotensin II-mediated calcium signals and mitogenesis in human prostate stromal cell line hPCPs. Br J Pharmacol 144: 3–10, 2005.
Wolf G, Butzmann U, and Wenzel UO. The renin-angiotensin system and progression of renal disease: from hemodynamics to cell biology. Nephron Physiol 93: 3–13, 2003.(Max C. Liebau, D. Lang, J. Bhm, N. Endli)