Abstract
Prorenin is activated without proteolysis by binding of prorenin receptor to the pentameric “handle region” (HR) of prorenin prosegment. It was hypothesized that such activation occurs in the kidneys of hypertensive rats and causes tissue renin-angiotensin system (RAS) activation and end-organ damage. Because the HR’s binding to its binding protein made the adjacent tetrameric “gate region” (GR) accessible to its specific antibody, immunohistochemistry of the GR was performed to test the hypothesis. Methods also were devised specifically to inhibit the nonproteolytic activation by the decapeptide corresponding to the HR as a decoy. Immunohistochemistry of the GR demonstrated that the majority of nonproteolytically activated prorenin is present in podocytes of the kidneys from stroke-prone spontaneously hypertensive rats, in which activation of renal tissue RAS, proteinuria, and glomerulosclerosis occurred. Continuous subcutaneous administration of the HR decoy peptide completely inhibited both nonproteolytic activation of tissue prorenin and activation of tissue RAS without affecting circulating RAS or arterial pressure and significantly attenuated the development and progression of proteinuria and glomerulosclerosis. These studies clearly demonstrated that nonproteolytic activation of prorenin in glomeruli is critically involved in renal tissue RAS activation, leading to renal damage in hypertensive animals.
Activated tissue renin-angiotensin system (RAS) significantly contributes to the development and the progression of end-organ damage. We and others found that a prorenin binding protein exists on the surface of plasma membrane in humans (1–5) and in rat tissues (6) and that its binding to prorenin activates proenzyme by a conformational change without proteolytically cleaving the 43–amino acid prosegment off the main body of active (mature) renin. Because the prorenin receptor localizes to the kidney (4), the locally activated prorenin by prorenin receptor can generate angiotensin I (AngI) and II (AngII) locally and exert its local action, leading to renal damage. In the kidneys of hypertensive animals, however, the nonproteolytic activation of prorenin has not been demonstrated in vivo.
In earlier studies, we found that an N-terminal prosegment of human prorenin has a protruding pentameric segment termed “handle region” (HR) and an adjacent tetrameric segment termed “gate region” (GR) that is not accessible by its specific antibodies until it is loosened from the active site cleft (5). Rat prorenin contains analogous pentameric HR (I11PLLKK15P) and tetrameric GR (S7PFGR10P) in positions that are identical to those of human prorenin. If the nonproteolytic activation mechanism is functional, then the prorenin will gain renin activity by binding of prorenin receptor to its HR, and tissues that are bound by the nonproteolytically activated prorenin can be visualized by immunohistochemical staining of GR by its specific antibody, which provides persuasive evidence that tissue prorenin binds to the prorenin receptor and activation is mediated by a nonproteolytic mechanism.
Here we present evidence that the prorenin receptor activates prorenin in the glomeruli of the hypertensive rat kidneys, where end-organ damage occurs, and markedly elevates tissue AngI and AngII concentration. Furthermore, the prorenin activation was almost completely blocked by HR decoy peptide (HRP), which inhibits the binding of prorenin to prorenin receptor (6), resulting in the normalization of AngI and AngII in the kidneys of hypertensive animals to the levels of normotensive animals without lowering the circulating RAS. These results may be taken as evidence for important roles of nonproteolytically activated prorenin in tissues for hypertensive renal damage.
Materials and Methods
Animals
We maintained male stroke-prone spontaneously hypertensive rats (SHRsp) and normotensive control Wistar-Kyoto rats (WKY; Charles River Labs, Yokohama, Japan) in a temperature-controlled room that was set at 23°C and on a 12:12-h light-dark cycle. Rats had free access to 1% NaCl water and a normal-salt-diet rat chow (0.4% NaCl; CE-2, Nihon Clea, Tokyo, Japan). The Keio University Animal Care and Use Committee approved all experimental protocols. At 4 wk of age, we subcutaneously implanted an osmotic minipump (model 2004 for 28-d use; Alzet, Cupertino, CA) that contained saline or a decapeptide, NH2-RILLKKMPSV-COOH, as an HRP of rat prorenin (0.1 mg/kg) under sodium pentobarbital anesthesia (50 mg/kg intraperitoneally) and divided the rats (100 to 150 g body wt) into four groups: SHRsp, SHRsp+HRP, WKY, and WKY+HRP. At 8 wk of age, we replaced the minipump with a pump that was filled with the same solution, thus the daily dose of HRP was 1.8 to 3.6 μg/kg, considering the body weight gain. Continuous subcutaneous infusion of FITC-conjugated HRP (0.1 mg/kg) for 4 wk by the minipump caused a significant uptake of HRP in brain, heart, liver, adrenal gland, aorta, adipose tissue, and kidneys of rats. No uptake of HRP was observed in small, resistant arteries. In the kidneys, FITC-conjugated HRP was detected in the glomeruli and tubular lumen but not in the extraglomerular vascular system (Figure 1). Four weeks later, six to eight rats were decapitated at 12 wk of age to collect their blood and kidneys.
In vivo uptake of the FITC-conjugated “handle region” (HR) decoy peptide (FITC-HRP; 0.1 mg/kg) in the kidneys from rats that received subcutaneous infusion for 28 d with an osmotic minipump; renal localization of rat prorenin receptor mRNA was assessed by in situ hybridization with antisense and sense probes. Bars = 25 μm. The photomicrographs show the presence of FITC-HRP in the glomeruli and tubular lumen and the presence of rat prorenin receptor mRNA in glomerular epithelium and tubular cells.
Arterial Pressure and Urinary Protein Excretion
At 4 wk of age, we implanted a telemetry transmitter probe (model TA11PA-C40; Data Sciences International, St. Paul, MN) into rats under sodium pentobarbital anesthesia (50 mg/kg intraperitoneally), and the flexible tip of the probe was positioned immediately below the renal arteries. The transmitter then was sutured into the abdominal wall, and the incision was closed. The rats then were returned to their home cages and allowed to recover for 6 d before the start of the measurements. We monitored conscious mean arterial pressure (MAP), heart rate (HR), and the activity in unrestricted and untethered animals with the Dataquest IV system (Data Sciences International), which consisted of the implanted radiofrequency transmitter and a receiver placed under each cage. The output was relayed from the receiver through a consolidation matrix to a personal computer. Individual 10-s MAP, HR, and activity waveforms were sampled every 5 min throughout the course of the study, and daily averages and SD then were calculated. The 24-h urine was collected in a metabolic cage, and urinary protein and creatinine excretion were determined with a Micro TP test kit (Wako, Osaka, Japan) and a Creatinine HA test kit (Wako), respectively.
Renal Morphology
Part of the kidney that was removed from each animal was fixed in 10% formalin in phosphate buffer (pH 7.4), and paraffin-embedded sections of the kidney were stained by the periodic acid-Schiff method. In the kidneys, we quantitatively determined the total area of sclerosis within the glomerular tuft by a semiquantitative scoring system (6,7).
RAS
Immediately after decapitation, a 3-ml blood specimen was collected into a tube that contained 30 μl of EDTA (500 mM), 15 μl of enalaprilat (1 mM), and 30 μl of o-phenanthrolene (24.8 mg/ml) and pepstatin (0.2 mM), and plasma samples were obtained by centrifugation. Plasma levels of components of the circulating RAS were determined as described previously (8). Total renin content and the AngI and AngII levels in renal cortex that was harvested from the kidneys were determined as previously reported (9,10).
We also extracted total RNA from part of the renal cortex that was removed from each animal with an RNeasy Mini Kit (Qiagen, Tokyo, Japan) and performed a real-time, quantitative reverse transcription–PCR with the TaqMan One-Step RT-PCR Master Mix Reagents Kit; an ABI Prism 7700 HT Detection System (Applied Biosystems, Foster City, CA); and probes and primers for the rat genes encoding renin, angiotensinogen, angiotensin-converting enzyme (ACE), cathepsin B, and glyceraldehyde-3-phosphate dehydrogenase, as described previously (6,11,12). We designed the probe and primers for the rat prorenin receptor (forward 5′-CATTCGACACATCCCTGGTG-3′; reverse 5′-AAGGTTGTAGGGACTTTGGGTG-3′; probe 5′-FAM-AAGTCAAGGACCATCCTTGAGACGAAACAA-TAMRA-3′), on the basis of its cDNA sequence reported in the GenBank database (accession no. AB188298 in DNA Databank of Japan).
Preparation of Specific Antibodies and Immunoblot Analysis
A rabbit anti-rat GR antibody was raised against peptide SFGRC conjugated with keyhole limpet hemocyanin. The GR peptide also was used to determine the titer of the antiserum with a Vectastain ABC-AP rabbit IgG kit (Vector Laboratories, Burlingame, CA) and as the ligand of the affinity column for purification of the antibody. High-titer antisera were obtained 8 wk after the first immunization. The affinity gel was prepared by conjugating the NH2-terminal amino group of the antigen peptide to Biogel 102 (amine coupling gel; Bio-Rad, Tokyo, Japan). The antibody was purified with the affinity column, and the concentration of the purified antibody (1.70 mg/ml) was calculated using an extinction coefficient of 1.35 per 1 mg/ml IgG and 280 nm. For immunoprecipitation analysis, the recombinant rat prorenin (13) before and after acidification (pH 3.3) and renin were incubated with 2 μl of anti-rat GR antiserum for 1 h at 4°C in 500 μl of the reaction mixture. For separation of active or inactive prorenin/renin that was bound to the antibody from the unbound prorenin/renin fraction, 50 μl of Protein A Sepharose-CL4B beads (Pharmacia, Peapack, NJ) was suspended for 1 h at 4°C, and then the Sepharose beads were sedimented. After washing, the precipitates were resuspended in SDS-gel sample buffer that contained 2-mercaptoethanol, boiled for 5 min, and centrifuged. The supernatant was subjected to 12% SDS-PAGE and electrophoretically transferred to the polyvinylidene difluoride membrane. Renin was prepared by trypsin treatment of the recombinant prorenin. Prorenin was incubated with 200 μg/ml trypsin for 20 min at pH 7.4 and 25°C, and trypsin action was stopped with 1 mmol/L PMSF. Prorenin-containing medium was acidified to pH 3.3 for 2 wk at 4°C in the presence of 5 mM EDTA, 1 mg/ml thermally treated BSA, and 0.02% sodium azide, as described previously (14). The membrane was incubated with 3 nM purified rabbit anti-rat GR antibody for 1 h at room temperature. After washing, the immunocomplex on these sheets was visualized with horseradish peroxidase–conjugated streptavidin and diaminobenzidine.
Immunohistochemical Evaluation
For immunohistochemical staining, deparaffinized sections were pretreated with proteinase K, and after the sections were boiled in citrate buffer with microwaves to unmask antigenic sites, endogenous biotin was blocked with a Biotin Blocking System (X0590; Dako Corp., Carpinteria, CA). Next, the sections were immersed in 0.3% H2O2 in methanol to inhibit endogenous peroxidase and then precoated with 1% nonfat milk in PBS to block nonspecific binding. For immunohistochemical staining of nonproteolytically activated prorenin, the anti-rat GR antibody was applied to the sections as the primary antibody. The sections were incubated with a biotin-conjugated anti-rabbit IgG as the secondary antibody, and the antibody reactions were visualized with a Vectastain ABC Standard Kit (Vector Laboratories) and an AEC Standard Kit (Dako) according to the manufacturers’ instructions. For immunohistochemical staining of collagen IV and TGF-β1, the monoclonal mouse anti-human collagen type IV antibody (Chemicon, Temecula, CA) and the polyclonal rabbit anti–TGF-β1 (Santa Cruz Biotechnology, Santa Cruz, CA) were applied, respectively, to the sections as the primary antibody. The anti-mouse IgG and anti-rabbit IgG were used as the secondary antibody. We quantitatively determined the immunoreactive nonproteolytically activated prorenin-positive area or the immunoreactive phospho–mitogen-activated protein kinase–positive area in each glomerulus at ×200 magnification with a Mac SCOPE (Version 2.5; Mitani Corp., Fukui, Japan) and expressed it as a percentage of the whole cross-sectional area of the glomerulus.
Double Immunofluorescence
Dual immunofluorescence labeling was performed on 3-μm frozen sections from the kidneys of hypertensive SHRsp. The sections were incubated simultaneously with rabbit anti-rat GR antibody (1:100 dilution) and mouse anti-nephrin antibody (1:50 dilution), mouse anti-desmin antibody (1:100 dilution), mouse anti-Thy1.1 antibody (1:100 dilution), or mouse anti–reca-1 antibody (1:40 dilution). After washing in PBS, the sections were incubated with TRITC-conjugated goat anti-rabbit IgG (1:200 dilution; Sigma) and FITC-conjugated goat anti-mouse Ig antibody (1:200 dilution; Sigma). Slides were examined with a Zeiss confocal microscope.
In Situ Hybridization
Glomerular localization of rat prorenin receptor was assessed by in situ hybridization, as described previously (15). Briefly, a digoxigenin-labeled cRNA probe was synthesized using a 123-bp fragment of the rat prorenin receptor mRNA as template (5′-A A A G G G A U U C G A U C U C C U G G U A U A G G C C A A U U U C C A U U U C G G A A A A C A A C A G A C C C U G G C G A U C U U A A U A U G C U A A A U U C A U U C G C U A A A G C A C U C G A C A C C A G A G A A G A G A G G A G A A C G A C A-3′).
Statistical Analyses
Within-group statistical comparisons were made by one-way ANOVA for repeated measures followed by the Newman-Keuls post hoc test. Differences between two groups were evaluated by two-way ANOVA for repeated measures combined with the Newman-Keuls post hoc test. P < 0.05 was considered significant. Data are reported as means ± SEM.
Results
BP and Circulating RAS
MAP was determined by a telemetry method in conscious, unrestrained animals in the SHRsp group (n = 7), the SHRsp+HRP group (n = 7), the WKY group (n = 4), and the WKY+HRP group (n = 4). MAP in the SHRsp group significantly increased from 118 ± 5 at 4 wk of age to 156 ± 4 at 8 wk of age and further increased to 228 ± 6 at 12 wk of age. However, MAP in the WKY rats did not change during the 8-wk observation period (96 ± 3, 98 ± 2, and 98 ± 3 mmHg at 4, 8, and 12 wk of age, respectively). HRP did not affect MAP in either the SHRsp (116 ± 4, 154 ± 6, and 224 ± 6 mmHg at 4, 8, and 12 wk of age, respectively) or the WKY group (97 ± 3, 96 ± 3, and 96 ± 3 mmHg at 4, 8, and 12 wk of age, respectively).
At 12 wk of age, plasma renin activity and plasma prorenin level in the SHRsp group averaged 3.6 ± 1.0 and 6.7 ± 0.4 ng/ml per h, respectively, and were significantly higher than those in the WKY group (1.7 ± 0.6 and 3.9 ± 0.8 ng/ml per h, respectively), and HRP did not affect them in either the SHRsp (3.5 ± 0.8 and 7.0 ± 0.4 ng/ml per h, respectively) or the WKY group (2.0 ± 0.6 and 3.6 ± 0.5 ng/ml per h, respectively). At 12 wk of age, plasma AngI and AngII levels also were higher in the SHRsp group (249 ± 74 and 169 ± 10 fmol/L, respectively) than in the WKY group (132 ± 26 and 38 ± 8 fmol/L, respectively), and HRP did not influence them in either the SHRsp (255 ± 72 and 173 ± 14 fmol/L, respectively) or the WKY group (124 ± 18 and 32 ± 7 fmol/L, respectively).
Kidney Damage
We investigated changes in renal morphology and urinary protein excretion by implanting minipumps that contained an HRP or saline for 8 wk in SHRsp and WKY, in the treatment and control groups, respectively. Figure 2A shows severe glomerulosclerosis in the kidneys of the SHRsp group at 12 wk of age and its mitigation by HRP. We did not observe any histologic changes in the kidneys of the WKY or the WKY+HRP rats at 12 wk of age. The glomerulosclerosis index in the 12-wk-old SHRsp group averaged 2.68 ± 0.13 and was significantly greater than in the WKY group (0.22 ± 0.03) and the WKY+HRP group (0.20 ± 0.03). The glomerulosclerosis index in the 12-wk-old SHRsp+HRP group averaged 1.07 ± 0.12 and was significantly lower than that in the SHRsp group but still greater than that in the WKY group and the WKY+HRP group (Figure 2B). In the SHRsp group, urinary protein excretion began to increase at 10 wk of age and exceeded 250 mg/d (256.5 ± 19.3 mg/d at 12 wk versus 11.3 ± 0.7 at 6 wk of age). HRP markedly attenuated it to 96.5 ± 13.5 mg/d in the 12-wk-old SHRsp+HRP group. No increases in urinary protein excretion were observed in the WKY or WKY+HRP groups during the 8-wk treatment period (Figure 2C).
Development of hypertensive glomerulosclerosis and its inhibition by the HR decoy peptide (HRP). (A) Periodic acid-Schiff–stained kidney sections. Bars = 25 μm. (B) Quantitative analysis of glomerulosclerosis in the SHRsp, SHRsp+HRP, WKY, and WKY+HRP groups (n = 6 in each). The photomicrographs and graph show the significant glomerulosclerosis at the 12-wk-old SHRsp group and its attenuation by HRP. (C) Urinary protein excretion in the SHRsp group (n = 6), SHRsp+HRP group (n = 8), WKY group (n = 5), and WKY+HRP group (n = 6). The graph shows a progressive increase in urinary protein excretion in SHRsp. HRP significantly attenuated the development and progression of proteinuria in SHRsp. *P < 0.05 versus WKY; †P < 0.05 for SHRsp+HRP versus SHRsp.
Renal AngII Synthesis
At 12 wk of age, the renal AngI content in the SHRsp, SHRsp+HRP, WKY, and WKY+HRP groups averaged 299 ± 42, 108 ± 4, 105 ± 8, and 91 ± 15 fmol/g, respectively, and their renal AngII content averaged 274 ± 35, 91 ± 17, 88 ± 14, and 84 ± 20 fmol/g, respectively (Figure 3, A and B). Thus, HRP completely inhibited the increase in renal AngI and AngII contents in the SHRsp group. Renal levels of angiotensinogen mRNA and ACE mRNA were similar in the four groups (Figure 3, C and D).
Changes in components of the angiotensin system in the kidneys of the SHRsp group (n = 8), SHRsp group treated with the HR peptide (SHRsp+HRP; n = 8), WKY group (n = 6), and WKY group treated with the HR peptide (WKY+HRP; n = 6) at 12 wk of age. (A) Renal angiotensin I (AngI) level. (B) Renal AngII level. The graphs show higher renal levels of AngI and AngII in the SHRsp group than in the SHRsp+HRP, WKY, and WKY+HRP groups, which had similar renal levels of AngI and AngII. HRP significantly inhibited the increases in renal levels of AngI and AngII in the SHRsp group. (C) Renal angiotensinogen mRNA level. (D) Renal angiotensin-converting enzyme (ACE) mRNA level. The graphs show similar renal levels of angiotensinogen mRNA and ACE mRNA in all four groups. *P < 0.05 for SHRsp versus WKY or for SHRsp+HRP versus WKY+HRP.
Renal Prorenin and Prorenin Receptor
At 12 wk of age, renal levels of renin mRNA and total renin protein were similar in the SHRsp, SHRsp+HRP, WKY, and WKY+HRP groups (Figure 4, A and B), but the renal cathepsin B mRNA level was lower in the SHRsp group than in the WKY group, and HRP did not alter its levels in either group (Figure 4C). At 8 wk of age, the renal level of prorenin receptor mRNA, which was present in glomerular epithelium and tubular cells (Figure 1), was higher in the SHRsp group than in the WKY group, and HRP did not alter its levels in either group (Figure 4D).
Changes in components of the renin system in the kidneys of the SHRsp group (n = 8), SHRsp group treated with the HR peptide (SHRsp+HRP; n = 8), WKY group (n = 6), and WKY group treated with the HR peptide (WKY+HRP; n = 6). (A) Renal renin mRNA level at 12 wk of age. (B) Renal total renin content at 12 wk of age. The graphs show similar renal levels of renin mRNA and total renin content in all four groups. (C) Renal cathepsin B mRNA level at 12 wk of age. The graph shows a lower renal cathepsin B mRNA level in the SHRsp group than in the WKY group. HRP did not affect the renal cathepsin B mRNA level in either the SHRsp or the WKY group. (D) Renal prorenin receptor mRNA level at 8 wk of age. The graph shows a higher renal prorenin receptor mRNA level in the SHRsp group than in the WKY group. HRP did not affect the renal prorenin receptor mRNA level in either the SHRsp or the WKY group.
Nonproteolytically Activated Prorenin in the Kidneys
An antibody to rat GR bound only to nonproteolytically activated prorenin but not to inactive prorenin or proteolytically activated renin (Figure 5A). Therefore, we used immunoreactivity to the anti-GR antibody to demonstrate nonproteolytically activated prorenin in the renal tissues of the 12-wk-old rats that were treated with HRP (Figure 5B). The nonproteolytically activated prorenin-positive area that was stained with anti-GR antibody increased in the glomeruli of the kidneys from the SHRsp group, but their number was significantly decreased by HRP infusion. The level of nonproteolytically activated prorenin in the glomeruli of the kidneys from the SHRsp+HRP group was similar to the level in the glomeruli of the kidneys from the WKY and WKY+HRP groups (Figure 5, B and C). These results suggested that the SHRsp kidneys contain higher levels of nonproteolytically activated prorenin.
Immunoprecipitation with the antibody against rat “gate region” (GR). (A) Recombinant rat prorenin (lanes 1 and 2) and renin (lane 3) were analyzed by immunoprecipitation with the antibody against rat GR, which usually is buried in the main body of prorenin in the inactive state and exposed in the active state. The image shows that the anti-rat GR antibody binds to nonproteolytically activated prorenin but not to inactive prorenin or proteolytically activated renin. (B) Immunohistochemistry of nonproteolytically activated prorenin in the kidneys of SHRsp with anti-rat GR antibody. The photomicrographs show an increase in nonproteolytically activated prorenin in the glomeruli of the kidneys from the 12-wk-old hypertensive SHRsp group. HRP completely inhibited the enhanced nonproteolytic activation of prorenin. Bars = 25 μm. (C) Quantitative analysis of nonproteolytically activated prorenin-positive area in glomerulus. The graph shows an increase in nonproteolytically activated prorenin in the SHRsp group and its inhibition by HRP. *P < 0.05 versus WKY.
Glomerular Localization of Nonproteolytically Activated Prorenin
Figure 6 shows glomerular localization of nonproteolytically activated prorenin. The red-colored immunofluorescence with polyclonal antibody against rat GR was merged in the green-colored immunofluorescence with mAb against nephrin but not desmin, Thy1.1, or reca-1. These results suggest that the majority of nonproteolytically activated prorenin was present in the podocytes.
Glomerular localization of nonproteolytically activated prorenin. Dual immunofluorescence with polyclonal antibody against rat GR (red) and mAb against nephrin, desmin, Thy1.1, or reca-1 (green) suggests the superimposition of nonproteolytically activated prorenin and signals of nephrin.
Collagen IV and TGF-β1 Expression in the Kidneys
Figure 7 shows the immunostaining to collagen IV and TGF-β1 in the glomeruli of the kidneys from the SHRsp, SHRsp+HRP, WKY, and WKY+HRP groups at 12 wk of age. The glomerular staining of collagen IV and TGF-β1 were enhanced in the SHRsp group compared with the WKY group. HRP treatment completely inhibited the enhanced staining of collagen IV and TGF-β1 in the SHRsp group.
(A) Immunohistochemistry of collagen IV in the kidneys of SHRsp with monoclonal anti–collagen IV antibody and quantitative analysis of collagen IV–positive area in glomerulus. The photomicrographs show an increase in collagen IV in the glomeruli of the kidneys from the 12-wk-old hypertensive SHRsp group. HRP completely inhibited the enhanced staining of collagen IV. Bars = 25 μm. *P < 0.05 versus WKY. (B) Immunohistochemistry of TGF-β1 in the kidneys of SHRsp with anti–TGF-β1 antibody and quantitative analysis of TGF-β1–positive area in glomerulus. The photomicrographs show an increase in TGF-β1 in the glomeruli of the kidneys from the 12-wk-old hypertensive SHRsp group. HRP completely inhibited the enhanced staining of TGF-β1. Bars = 25 μm. *P < 0.05 versus WKY.
Discussion
These studies using SHRsp show that the novel mode of nonproteolytic prorenin activation mechanism has a specific role in hypertensive end-organ damage in tissues in which prorenin is activated by its binding protein, such as its specific antibodies or prorenin receptor (4), instead of traditional activation by proteolytic cleavage of the 43–amino acid prosegment. Because the pentameric HR of rat prorenin protrudes from the main body of the prorenin molecule (5), it is readily bound by a specific binding protein, such as antibodies that are used as a model of the binding protein or the prorenin receptor. Whereas the adjacent tetrapeptide segment S7PFGR10P, termed GR, is buried in the active site cleft of inactive prorenin, its nonproteolytic activation releases GR from the cleft and makes it accessible to its specific antibody (5). Because the binding of tissue prorenin receptor to HR is prerequisite to the prorenin activation presumably by conformational changes, blockade of its binding to HR by the short synthetic HRP used as a decoy inhibits the nonproteolytic activation, a specific interventional inhibitor, and thus an indicator of the novel activation mechanism. Also, as stated above, nonproteolytic activation made GR accessible to anti-GR antibodies and permitted us to detect the activated prorenin in tissues by an immunohistochemical method. This in vivo approach is supported by the present in vitro observation that the anti-GR antibody binds exclusively to nonproteolytically activated prorenin but not to inactive prorenin or proteolytically activated renin.
Immunohistochemical studies of tissues with anti-GR antibody showed that HR binding generated immunoreactivity to anti-GR antibodies in the glomeruli of the kidneys in the hypertensive SHRsp that was not detectable in the normotensive WKY group. This in vivo observation indicates tissue-specific activation of prorenin by the nonproteolytic mechanism and corroborates previous data showing that high pressure inhibits prorenin processing to renin in juxtaglomerular cells (16). We previously demonstrated in vitro evidence that binding of a prorenin-binding protein, such as a prorenin receptor or the anti-HR antibody, to HR in the prorenin prosegment caused an activation of prorenin without proteolytic cleavage of prorenin and obtained in vitro and in vivo evidence that HRP, used as a decoy, competes out the HR binding and inhibits the nonproteolytic activation of prorenin (6). In our study, chronic administration of the decoy peptide HRP significantly decreased the number of nonproteolytically activated prorenin-positive cells to a level similar to that in WKY rats. These results suggest that the greater amount of prorenin was nonproteolytically activated, probably by intrinsic prorenin-binding proteins prorenin receptor, in the damaged kidneys of hypertensive animals.
Within the glomerulus, podocytes play an important role in the maintenance of the glomerular filtration barrier, and podocyte injury leads to proteinuria and initiates glomerulosclerosis, resulting in progressive loss of renal function (17). There were some indirect pieces of evidence that AngII disturbs podocyte biology. Overexpression of the AngII type 1 receptor in rat podocytes caused structural podocyte damage and proteinuria, leading to glomerulosclerosis (18). In experimental diabetic nephropathy, podocyte foot process broadening was attenuated by the RAS inhibitors (19). In addition, a recent study showed that human podocytes express mRNA for angiotensinogen, renin, ACE, and AngII receptors (20). These findings suggest that the podocyte RAS may significantly contribute to proteinuria and renal injury; however, the mechanism for the podocyte RAS activation in hypertensive animals remained unclear. AngII production has been reported to occur when the podocyte was exposed to mechanical strain (21), suggesting that a high perfusion pressure in the glomerulus stimulates the AngII production in podocytes, leading to proteinuria and glomerulosclerosis. Because double-immunofluorescent stainings in our study showed that activated prorenin is present in the podocyte of hypertensive SHRsp, nonproteolytic activation of prorenin may contribute to the RAS activation of podocytes in hypertensive animals. This also was supported by the results of our study that inhibition of nonproteolytic activation of prorenin by HRP decreased renal AngII levels and attenuated the development and progression of proteinuria and glomerulosclerosis in SHRsp.
Huang et al. (22) recently provided novel evidence that the prorenin receptor mRNA is present in cultured rat mesangial cells, whereas our study showed that the majority of activated prorenin is present in the podocytes but not in the mesangial cells. Therefore, the prorenin in the podocytes may be activated by other prorenin-binding proteins, or rat mesangial cells may have greater affinity for renin than prorenin, because recombinant renin has been reported to increase glomerular TGF-β1 expression in cultured rat mesangial cells (22). In addition, a recent preliminary study reported that the prorenin receptor mRNA also is present in glomerular epithelial cells (23). Further in vivo or ex vivo studies will be needed to determine the glomerular localization of prorenin-binding proteins, including the prorenin receptor.
Glomerular expression of collagen IV and TGF-β1 was enhanced in the kidneys of SHRsp, and the enhancement was significantly inhibited by treatment with HRP. The enhanced expression of matrix proteins may be due to the increased levels of angiotensin peptides, because the tissue levels of AngI and AngII also were higher in the kidneys in SHRsp than in WKY. The angiotensin peptide levels were completely normalized by treatment with HRP, presumably by preventing the binding of prorenin HR to the prorenin receptor protein. There were no changes in other RAS components, suggesting that the nonproteolytic activation of prorenin plays a key role in renal tissue RAS activation in hypertensive animals. It is interesting that we observed significant increases in prorenin receptor mRNA levels in the kidneys of 8-wk-old SHRsp. In addition, the kidney prorenin level should be higher in SHRsp than in WKY because the proportion of proteolytic activation of prorenin by cathepsin B, the proteolytic activator of prorenin, should be reduced in SHRsp, whereas total renin plus prorenin is similar between SHRsp and WKY. Therefore, it is likely that both increased tissue levels of prorenin and prorenin receptor are the determining factors in enhancing the nonproteolytic activation of prorenin by a prorenin receptor.
Renal levels of renin mRNA and total renin protein were similar in treated and untreated SHRsp and WKY groups at 12 wk of age. Because the renal cathepsin B mRNA level was lower in the SHRsp group than in the WKY group, the renal prorenin level was expected to be higher in the SHRsp group than in the WKY group, and the renal renin level was expected to be lower in the SHRsp group than in the WKY group. On the basis of the expected results and the concept that most of circulating renin and prorenin come from the kidneys, the SHRsp group was expected to have low plasma renin levels and high plasma prorenin levels but actually had a high plasma renin activity and high AngI and AngII levels. The detailed mechanism is unknown, although the increased plasma prorenin may be activated by binding to its receptor in small arteries (4) and thus have generated AngI and AngII in the plasma and contributed to the elevated BP in the SHRsp group. Further studies will be needed to determine whether the increased plasma renin activity in the SHRsp group may be attributable to the activated prorenin in the plasma by prorenin receptors.
Conclusion
We observed higher levels of nonproteolytically activated prorenin, activated tissue RAS, and histologic damage in the kidneys of SHRsp. Inhibition of prorenin binding to prorenin receptor completely prevented both the nonproteolytic activation of prorenin and activation of the tissue RAS without affecting the circulating RAS or arterial pressure. These findings suggest a crucial role of nonproteolytic activation of prorenin in tissue RAS activation leading to renal damage in individuals with hypertension.
Acknowledgments
This work was supported in part by grants from the Ministry of Education, Science and Culture of Japan (1503340, 16613002, 16790474, and 17390249) and research grant HL58205 from the National Institutes of Health (Bethesda, MD).
We gratefully acknowledge Dr. Irwin Landon, Department of Pharmacology, Vanderbilt University, for helpful discussion and critical reading of this manuscript.
Footnotes
Published online ahead of print. Publication date available online at www.jasn.org.
- © 2006 American Society of Nephrology
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- Renin, (pro)renin and receptor: an update
- (Pro)renin Receptor-Mediated Signal Transduction and Tissue Renin-Angiotensin System Contribute to Diabetes-Induced Retinal Inflammation
- Critical Review of Prorenin and (Pro)renin Receptor Research
- The Putative (Pro)renin Receptor Blocker HRP Fails to Prevent (Pro)renin Signaling
- Prorenin and Renin-Induced Extracellular Signal-Regulated Kinase 1/2 Activation in Monocytes Is Not Blocked by Aliskiren or the Handle-Region Peptide
- (Pro)Renin Receptor Peptide Inhibitor "Handle-Region" Peptide Does Not Affect Hypertensive Nephrosclerosis in Goldblatt Rats
- The Intrarenal Renin-Angiotensin System: From Physiology to the Pathobiology of Hypertension and Kidney Disease
- Renin and Its Putative Receptor Remain Enigmas
- Regression of Nephropathy Developed in Diabetes by (Pro)renin Receptor Blockade
- Slowly Progressive, Angiotensin II-Independent Glomerulosclerosis in Human (Pro)renin Receptor-Transgenic Rats