Pivotal role of the renin/prorenin receptor in angiotensin II
production and cellular responses to renin
Genevieve Nguyen,1 Françoise Delarue,1 Céline Burcklé,1 Latifa Bouzhir,1 Thomas Giller,2
and Jean-Daniel Sraer1
1Institut
National de la Santé et de la Recherche Médicale (INSERM) U489, and Association Claude Bernard,
Hopital Tenon, Paris, France
2Hoffman–La Roche Ltd., Basel, Switzerland
Address correspondence to: Genevieve Nguyen, INSERM U489, Hopital Tenon, 4 rue de la Chine, 75020, Paris, France.
Phone: (331) 56 01 83 17; Fax: (331) 43 64 54 48; E-mail: genevieve.nguyen@tnn.ap-hop-paris.fr.
Latifa Bouzhir’s present address is: INSERM U451, Ecole Polytechnique-ENSTA, Palaiseau, France.
Thomas Giller’s present address is: AXOVAN Ltd. Innovation Center, Allschwil, Switzerland.
Received for publication September 24, 2001, and accepted in revised form April 22, 2002.
Renin is an aspartyl protease essential for the control of blood pressure and was long suspected to
have cellular receptors. We report the expression cloning of the human renin receptor complementary DNA encoding a 350–amino acid protein with a single transmembrane domain and no homology with any known membrane protein. Transfected cells stably expressing the receptor showed
renin- and prorenin-specific binding. The binding of renin induced a fourfold increase of the catalytic efficiency of angiotensinogen conversion to angiotensin I and induced an intracellular signal
with phosphorylation of serine and tyrosine residues associated to an activation of MAP kinases
ERK1 and ERK2. High levels of the receptor mRNA are detected in the heart, brain, placenta, and
lower levels in the kidney and liver. By confocal microscopy the receptor is localized in the mesangium
of glomeruli and in the subendothelium of coronary and kidney artery, associated to smooth muscle cells and colocalized with renin. The renin receptor is the first described for an aspartyl protease.
This discovery emphasizes the role of the cell surface in angiotensin II generation and opens new perspectives on the tissue renin-angiotensin system and on renin effects independent of angiotensin II.
J. Clin. Invest. 109:1417–1427 (2002). doi:10.1172/JCI200214276.
Introduction
The renin-angiotensin system (RAS) is critical for the
control of blood pressure and salt balance in mammals.
Renin is an aspartyl protease synthesized as prorenin, a
proenzyme that contains an additional 43–amino acid
N-terminal fragment. The physiological maturation of
prorenin into active renin takes place exclusively in the
juxtaglomerular cells of the kidney (1). Renin has high
substrate specificity, and its only known substrate is
angiotensinogen. Renin cleaves the N terminus of circulating angiotensinogen to angiotensin I (Ang I; a
decapeptide), which is then transformed in angiotensin
II (Ang II; an octapeptide) by soluble or endothelial
cell–associated angiotensin-converting enzyme (ACE).
In the heart, the majority of Ang I is converted by chymase (2). The rate-limiting step in the RAS is Ang I generation, even though the major biologically active peptide is Ang II. Ang II acts on vascular smooth muscle
cells as a potent vasoconstrictor via Ang II receptors.
These receptors are widely distributed and expressed by
many cell types (3). Components of the RAS and Ang II
receptors are found in the brain (4) and in many peripheral tissues such as the heart (5) and kidney (6), but also
placenta (7), testis (8), adipose tissue (9), and eye (10, 11).
Recent studies have shown that the RAS is involved in
diverse physiological and pathological processes such as
growth and remodeling (12), development (13), inflamThe Journal of Clinical Investigation
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mation (14), vascular hypertrophy, and thrombosis (15).
If Ang II and Ang IV receptors are well characterized, the
demonstration of a functional renin receptor is still
missing. Several proteins able to bind renin have been
reported. The widely distributed mannose-6-phosphate
receptor has been shown to bind renin and prorenin on
rat cardiac myocytes (16) and on human endothelial
cells (17). Another renin-binding protein called RnBp
has been identified in rat, porcine, and human tissues
and was shown later to be identical to the N-acyl-D-glucosamine 2-epimerase (18). Several binding sites have
also been described on membranes from different rat
organs but no functional effects of renin binding were
reported (19, 20). In contrast to these studies (19, 20), we
have shown that renin could bind to human mesangial
cells in culture and that the binding induced an hypertrophic effect and an increase of plasminogen activator
inhibitor-1. Renin bound to the receptor was neither
internalized nor degraded (21, 22).
The receptors of proteases may play several roles. They
serve to focus the enzymatic activity on a cell surface or
at a cell-extracellular matrix interface, as for urokinase
and plasminogen/plasmin (23). The urokinase receptor
binds not only the active enzyme but also the inactive
proenzyme. The binding of the proenzyme induces a
conformational change that unmasks the active site in
the absence of proteolytic activation of the proform.
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The receptor-bound enzyme has a much more efficient catalytic activity on the substrate (24). The receptor of a protease may also activate an intracellular signaling pathway leading to a change of cell phenotype
and proliferation, as for the receptors of urokinase
(25) and thrombin (26).
In this report we describe the identification of a renin
receptor by expression cloning. Transfection of renin
receptor cDNA into cells lacking specific binding of
renin results in the expression of a 45-kDa membrane
protein that specifically binds renin and prorenin. The
binding of renin to its receptor induces: (a) an increase
of the catalytic efficiency of angiotensinogen conversion into Ang I by renin bound to the receptor compared with renin in soluble phase, and (b) a rapid phosphorylation of the receptor on serine and tyrosine
residues associated with an activation of MAP kinases
ERK1 and ERK2. Immunofluorescence and confocal
analyses on normal human kidney and cardiac biopsies
show that the receptor is localized within the mesangial area of glomeruli and in the subendothelium of
kidney and coronary artery. Double-staining with antirenin and anti–smooth muscle α-actin showed that, in
the arteries, the receptor is present on smooth muscle
cells. Furthermore, the staining of mature fetal placenta showed that renin and the receptor were colocalized
in vascular structures and in syncytiotrophoblast cells.
Taken together, our results provide evidence for the
existence of a functional renin receptor. This receptor
is responsible for renin cellular effects, independent of
the generation of Ang II. On the other hand, by increasing the efficiency of angiotensinogen cleavage by membrane-bound renin, this receptor also plays a role in the
renin effects dependent on Ang II generation.
Methods
Reagents. Human recombinant renin, prorenin (27), and
the polyclonal Ab to human renin were a gift of Walter
Fischli (Hoffmann-La Roche Ltd., Basel, Switzerland).
The polyclonal Ab to Ang I was a gift from Jean-Baptiste Michel (INSERM U460, Paris, France).
Expression cloning and 5′ rapid amplification of cDNA ends
PCR. A human kidney cDNA-phage library (CLONTECH Laboratories Inc., Palo Alto, California, USA) was
screened with 125I-renin. Twelve positive clones were subcloned in pGEM-T Easy Vector (Promega Corp., Madison, Wisconsin, USA) and sequenced. One clone, N14F
(GenBank accession number: AF291814), was selected,
and 5′ rapid amplification of cDNA ends (RACE) was
performed using the 5′ RACE PCR kit of Life Technologies Inc. (Rockville, Maryland, USA). Specific RT was performed on polyA+ mRNA from primary mesangial cells
with the antisense primer 5′-GAGCGTCAACAAGGATCTTAGAA-3′ complementary to nucleotides 785–819. Nested PCR was done using the antisense primers 5′GGGGGAGTGAACTGAGAACA-3′ and 5′-GCATTCTCCAAAGGGTACGA-3′ complementary to nucleotides
577–597 and 371–391. The nucleotide and the deduced
amino-acid sequences were analyzed using NetPhos 2.0
1418
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prediction server (Center for Biological Sequence Analysis, Technical University of Denmark, Copenhagen,
Denmark) and PSORT II prediction server (prediction
of protein sorting signals and localization sites in amino
acid sequence, Human Genome Center, Institute of
Medical Science, University of Tokyo, Japan).
In vitro translation of N14F and coprecipitation experiments. The coding region of N14F with the FLAG tag
was translated in vitro (TnT-Coupled Reticulocyte
Lysate; Promega Corp.) in the presence of 35S-methionine and was analyzed by SDS-PAGE and fluorography. For coprecipitation experiments, the translated
product was incubated with 25 nM renin for 2 hours
at 37°C and precipitated with anti-renin Ab and protein G-Sepharose (Sigma-Aldrich, St. Louis, Missouri,
USA). The proteins were eluted and analyzed by SDSPAGE and fluorography.
Transfection of mesangial cells and renin-binding assay. A
human fetal mesangial cell line (HMC) (28) that did
not express N14F either by PCR or Northern blot
analysis was selected and transfected by the Fugene
method (Roche Diagnostics, Meylan, France) and with
the coding region of N14F subcloned in pcDNA 3.1
Zeo (Promega Corp.) tagged at the 3′ end with the
FLAG peptide (Asp-Tyr-Lys-Asp-Asp-Asp-Asp-Lys) by
PCR, introducing GACTACAAGGACGACGATGACAAG
before the stop codon. Stable transformants were
selected with Zeocine. Two clones, HMC2 and HMC4,
were selected. The binding experiments were performed as described (21), and the results were analyzed
by the Scatchard method.
Ang I generation by cell-associated renin and prorenin or in
soluble phase. Membranes from HMC and HMC2 were
prepared as described (21). Five or ten micrograms of
membrane was incubated with 5 nM renin for 1 hour at
37°C. After washing, the membranes were resuspended
in 10 µl Krebs buffer containing 2% BSA and incubated
1 hour at 37°C with increasing concentrations (1 nM–1
µM) of angiotensinogen (Sigma-Aldrich) in 110 µl
acetate buffer, pH 5.7. The membranes were centrifuged, and the supernatant was assayed for Ang I (29).
At the end of the incubation time, aliquots of membranes were treated with 100 mM glycin buffer, pH 3.5,
and the renin eluted was measured with the Renin III
Generation kit (Bio-Rad Laboratories Inc., Hercules,
California, USA). Identical amounts of renin in solution
were incubated alone or with 10 µg of HMC membranes
together with angiotensinogen. Similar experiments
were performed with prorenin bound to HMC2 membranes or in solution, except the incubation was performed in Kreb’s buffer, pH 7.4, for 4 hours to prevent
a possible activation of prorenin. These experimental
conditions were determined previously for fully active
renin to obtain identical Ang I generation by renin incubated in buffer at pH 5.7 and pH 7.4. Experiments with
prorenin were performed twice in duplicate.
Analysis of the receptor expressed by transfected cells and
cross-linking with renin. Control cells and HMC2 cells
were metabolically labeled with 35S-methionine for 24
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hours, lysed with 10 mM Tris buffer, pH 8.0, 140 mM
NaCl (TSA) containing Triton X-100 1%, 1 mM PMSF,
500 U/ml aprotinin (TSA-TX). The receptor was
immunoprecipitated, and the proteins eluted were analyzed by SDS-PAGE and fluorography. For cross-linking experiments, the cells were incubated with or without 25 nM renin for 1 hour at 37°C and cross-linked as
described (21). The cells were lysed, and the reninreceptor complex was immunoprecipitated with antireceptor Ab. The proteins eluted were analyzed by SDSPAGE and fluorography.
Intracellular free [Ca2+] and cyclic AMP changes induced by
renin. Briefly, subconfluent cells in a 10-mm Petri dish
were loaded with 5 µM Fura-2 for 45 minutes at 37°C,
washed, and detached by mild trypsinization. Cells were
resuspended in PBS buffer containing 1% BSA and transferred into a quartz cuvette under constant stirring at
37°C. Fluorescence was monitored in a Quanta Master
1 spectrofluorometer, and [Ca2+] was calculated using
Felix 1.1 software program (Photon Technology International, Brunswick, New Jersey, USA).
For cAMP study, cells in 24-well plates were serum
deprived for 18 hours and incubated 30 minutes in culture medium containing 100 µM isobutyl-methyl-xanthine. After addition of 10 nM renin or 1 µM PGE2 or 1
µM of isoproterenol, the reaction was stopped after 5
minutes by addition of 250 µl of ethanol 95%-formic acid
5%. The supernatant was evaporated overnight, and the
cAMP content of each well was measured with cAMP[125I] assay (Amersham Pharmacia Biotech, Piscataway,
New Jersey, USA). Measurements of cAMP activity were
made two times in triplicate, and statistical analysis was
performed using the Wilcoxon nonparametric test.
Analysis of the pattern of the receptor phosphorylation and
MAP kinases ERK1/ERK2 activation induced by renin.
HMC2 cells were incubated with 10 nM renin for 0, 3,
and 10 minutes and in the presence of 100 nM Captopril. The cells were lysed with TSA-TX containing 2 mM
Na3VO4, 50 mM NaF, and 5 mM EDTA. The receptor
was immunoprecipitated with anti-FLAG agarose.
Bound proteins were analyzed by Western blotting with
anti-phosphotyrosine (Chemicon International, Temecula, California, USA), anti-phosphoserine, or anti-phosphothreonine Ab’s (Sigma-Aldrich) and revealed by the
ECL kit (Amersham Pharmacia Biotech). Control cells
were incubated with renin, lysed, and analyzed by Western blotting with anti-phosphotyrosine and anti-phosphoserine Ab’s without prior immunoprecipitation of
the receptor with anti-FLAG agarose.
To study MAP kinases ERK1 (p44) and ERK2 (p42)
activation, cells were serum deprived for 18 hours and
stimulated with 10 nM renin for various times in the
presence of 1 µM Losartan. The cells were lysed with 100
µl lysis buffer as described. Proteins were measured with
the Bio-Rad Laboratories Inc. DC protein assay and analyzed by Western blotting using Ab’s to total and active
ERK1/ERK2 (Promega Corp.) and revealed by the ECL
kit. The blots were scanned and plotted to obtain a bar
graph representation of the ratio of active-to-total
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ERK1/ERK2 activation using the NIH IMAGE program
(Bethseda, Maryland, USA). To confirm MAP kinases
ERK1/ERK2 activation, MAP kinase activity of cell
lysate was also measured using the Biotrak p42/p44
MAP kinase enzyme assay system (Amersham Pharmacia Biotech). Measurements of MAP kinase activity were
made two times in triplicate, and statistical analysis was
performed using the Wilcoxon nonparametric test.
Northern blot analysis. A cDNA fragment containing
the Not1-Xba1 coding region of N14F subcloned into
pcDNA 3.1 was radiolabeled (PrimeIt labeling kit;
Stratagene, La Jolla, California, USA) and hybridized to
a membrane containing the polyA+ mRNA from
human tissues (CLONTECH Laboratories Inc.). The
amount of RNA was adjusted so that the β-actin
hybridization signal was of comparable intensity in
every lane. The blots were hybridized and washed using
ExpressHyb (CLONTECH Laboratories Inc.).
Immunofluorescence studies on transfected cells, on human
normal kidney and heart, and on mature placenta. Rabbits
were immunized by coinjection of synthetic peptides
corresponding to amino acids 221–235 and 337–350 of
the receptor (Eurogentec SA, Herstal, Belgium). Specimens of kidney and heart were obtained from the operating room after total nephrectomy for tumor and during mitral valve replacement, respectively, with informed
consent of the patient. Mature placenta was obtained
from the delivery room, with informed consent of the
patient. Permeabilized cells and the cryopreserved tissues were fixed in 4% paraformaldehyde and incubated
with rabbit antiserum to the receptor (1:1,000 dilution)
with mAb’s to CD31 or to smooth muscle α-actin
(Sigma-Aldrich). FITC or Texas red (TRITC) secondary
Ab’s were used to visualize the staining. The specificity
of the staining for the renin receptor was assessed by
preincubation of the primary Ab with the peptides. To
study the colocalization of renin and renin receptor,
staining was performed on sequential sections of placenta using the polyclonal Ab to the receptor (1:1,000
dilution) revealed by FITC-secondary anti-rabbit Ig or
the polyclonal Ab to human renin (1:50 dilution)
revealed by TRITC-secondary anti-rabbit Ig.
Results
Expression cloning of renin receptor. We have screened
1,000,000 clones of a commercial human kidney expression library in λgt11 with recombinant human renin
labeled with 125I. The positive clones were amplified by
PCR using λgt11 primers and subcloned in pGEM-T
Easy for sequencing. The sequence of one clone (N14F,
2,034 bp) represented a new protein with no homology
with any known membrane-receptor family (Figure 1a).
The initiation codon was surrounded by the optimal
Kozak nucleotide consensus sequence (GCCACC), and
the open reading frame encoded a 350-amino acid protein (molecular weight 39.008 kDa). The integrity of the
5′ end was assessed by rapid amplification of complementary cDNA 5′ end by PCR and by comparison with
an expressed sequence tag data base. The deduced
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Figure 1
Nucleotide and amino acid sequence of N14F. (a) The nucleotide sequence is numbered on the right. The ATG of GCACCATGG is assigned
as codon 1 on the basis of its close match to the C/GCACCATGG Kozak consensus sequence for optimal initiation of translation in eukaryotic cells. An in-frame TGA stop codon is located 858 nucleotides before the AATAAA cleavage and polyadenylation sequence, followed by
the poly(A)+ sequence 15 nucleotides after the AATAAA. (b) The amino acid numbering begins with the first methionine of the longest open
reading frame. The hydropathy profile of the deduced amino acid sequence of the N14F computed according to Kyte and Doolittle hydropathicity analysis is shown in the lower panel. The region with hydropathic index consistent with formation of a transmembrane spanning
segment of 21 amino acids is boxed. Tyr 335, the amino acid most likely to be phosphorylated, is underlined.
amino acid sequence and the analysis of hydrophobicity of the translated protein (30) predicts that the protein has two hydrophobic regions, one located in the
amino terminus from amino acids 1–16, which likely
represents a signal peptide, and another in the carboxy
terminus from amino acids 306–326, which likely represents a transmembrane region (Figure 1b). Of the 24
amino acids of the cytoplasmic tail, Ser 337, Thr 343,
Tyr 335, and Tyr 340 represent possible sites for regulatory phosphorylation, of which Tyr 335 appears to be
the most likely phosphorylated site (NetPhos 2.0 prediction). The predicted structure of this protein was
characterized by the absence of a N-linked glycosylation
site, a pI of 5.76, no PEST (proline, glutamic acid, serine,
and threonine) sequence, and an estimated half-life of
30 hours (PSORT II prediction).
In vitro translation and coprecipitation of the FLAG receptor
with renin. The construct N14F-FLAG was translated in
vitro using a rabbit reticulocyte lysate system and labeled
with 35S-methionine. Analysis by SDS-PAGE and fluo1420
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rography showed one band at the expected molecular
weight of 39 kDa (Figure 2a, lane A). Prior to performing
cell transfection, we tested the interaction of N14F with
renin by coprecipitation experiments. The translated
product labeled with 35S-methionine was incubated with
renin (25 nM) for 2 hours at 37°C, and the complexes
precipitated with anti-renin Ab. SDS-PAGE and fluorography analysis of the precipitate showed that the
receptor was recovered in the precipitate, confirming
that N14F was able to interact with renin (Figure 2b,
lane B). In the absence of renin, anti-renin Ab did not
react with in vitro–translated receptor (Figure 2b, lane C)
or by Western blot analysis (Figure 2c).
Renin binding and membrane expression of N14F protein by
HMC cells transformed with N14F. Next, the identity of the
cloned cDNA was demonstrated by stable transfection
of immortalized fetal HMCs (28) that did not express
N14F either by PCR using primers specific for N14F or
by Northern blot analysis (not shown). Two clones of
stable transformants, HMC2 and HMC4, were analyzed
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Figure 2
In vitro transcription/translation of N14F/FLAG protein and coprecipitation with renin. (a) N14F/FLAG fusion protein (lane A) or
control empty vector (lane B) transcribed in vitro and labeled with
35S-methionine are analyzed by SDS-PAGE and fluorography. (b)
Coprecipitation of the N14F/FLAG fusion protein incubated with
renin and immunoprecipitated with anti-renin Ab. The proteins
were eluted and analyzed by SDS-PAGE and flurography. Lane A:
N14F/FLAG fusion protein immunoprecipitated with anti-FLAG
agarose; lane B: elution of N14F/FLAG protein bound to renin; lane
C: N14F/FLAG protein immunoprecipitated with anti-renin Ab. (c)
Renin alone (lane A) was immunoprecipitated with Ab to the renin
receptor and analyzed by Western blotting with anti-renin Ab. Lane
B: recombinant renin.
by binding assays. The binding of renin on transformed
cells is saturable and characterized by a Kd of 5.0 nM and
a maximal binding site (Bmax) of 2.8 fmol/100,000 cells,
corresponding to approximately 17,000 sites/cell for
HMC2, and a Kd of 7.8 nM and a Bmax of 1.8
fmol/100,000 cells, approximately 13,000 sites/cell for
HMC4. In contrast, HMC control cells do not display
any specific renin binding (Figure 3a). Renin-specific
binding to HMC2 cells was inhibited by an excess of
cold prorenin (100 nM), suggesting that the receptor
also binds prorenin. In contrast, renin binding was not
inhibited by an excess of other aspartyl proteases, such
as pepsin and cathepsin D, or modified by nonrelated
proteins, such as hemoglobin, hepatocyte growth factor, or tissue-plasminogen activator (not shown). To
further assess the membrane expression of the renin
receptor on HMC2 cells we performed immunofluores-
cence staining with Ab to the receptor and analysis by
confocal microscopy. The results showed that the renin
receptor was cell surface associated and that no staining
could be detected in control cells transfected with
empty vector (Figure 3b).
Increased catalytic efficiency of angiotensinogen cleavage by
membrane-bound renin and prorenin. The kinetics of
angiotensinogen cleavage by HMC2 membrane-bound
renin and renin in solution were compared. The results
showed that the amount of Ang I generated by membrane-bound renin was much higher than that by
renin in solution (Figure 4a). The double-reciprocal
(Lineweaver-Burk) transformation of data resulted in
Km values of 1 µM and 0.15 µM for renin in solublephase and membrane-bound renin, respectively (Figure
4b). From the slope of the individual double-reciprocal
plots and the values of renin eluted from the membranes, the catalytic constant (kcat) values were 2.4 s–1
and 1.4 s–1 with 0.5 nM renin for solution phase and
membrane-bound renin, respectively. For 1 nM renin
immobilized on the membranes or in solution, the kcat
values were 2.6 s–1 and 1.8 s–1 for solution-phase and
membrane-bound renin, respectively. Therefore the catalytic efficiency of the reaction of renin with
angiotensinogen measured by the ratio kcat/Km was
lower for renin in solution, 2.2 to 2.4 µM–1 s–1, than for
renin bound to the receptor, 9.3 to 11.2 µM–1 s–1 (Figure
4c). These results indicate that angiotensinogen cleavage by membrane-bound renin is five times more efficient than by renin in solution. When membranes from
HMC control cells were tested for renin binding, an elution of bound renin, the amount of renin eluted from
HMC membranes, was under the limit of detection of
the renin antigen assay, precluding the comparison of
angiotensinogen conversion by HMC membranebound renin to renin in solution. In addition, Ang I generation was identical in buffer and in the presence of 10
µg of HMC membrane (not shown), indicating that
phospholipids alone, in the absence of receptor, do not
Figure 3
Membrane expression of N14F protein and renin binding by HMC cells transformed with N14F. (a) Saturation of binding on the HMC clone 2
(HMC2) expressing the receptor (squares, total; circles, nonspecific) and on HMC control cells (+, total; ×, nonspecific). The nonspecific binding was determined in the presence of 100 nM cold renin. Inset: Scatchard plot of the binding of renin to HMC2. (b) Immunofluorescence and
confocal analysis of the expression of N14F on HMC2 cells stably expressing N14F (upper panel) and by HMC control cells transfected with
empty vector (lower panel). The cells were stained with anti-receptor Ab (dilution 1:1,000) and with TRITC–conjugated secondary Ab.
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Figure 4
Ang I generation by soluble-phase and membrane-bound renin and prorenin. (a) Ang I generated by membrane-bound renin (squares, 1
nM; triangles, 0.5 nM) and by renin in solution phase (circles, 1 nM; diamonds, 0.5 nM). (b) Lineweaver-Burk plots of 1/Ang I generated
vs. 1/angiotensinogen (Agen) by 1 nM renin bound to the membranes (squares) or in solution (circles), and by 0.5 nM renin on membranes
(triangles) or in solution (diamonds). (c) Kinetic parameters of Ang I production by renin. (d) Comparison of Ang I generated by 0.5 nM
renin and prorenin incubated with angiotensinogen (Agen) 1 µM under different conditions. Bars 1, 2, and 3 are renin in solution, pH 5.7,
for 1 hour; pH 7.4 for 1 hour; and pH 7.4 for 4 hours, respectively. Bars 4 and 5 are prorenin pH 7.4 for 4 hours, in solution or membrane
bound, respectively. Experiments were performed twice in duplicate, and the results represent the mean of the two experiments.
alter renin activity. However, we cannot exclude the possibility that, since renin is more closely apposed to
membrane phospholipids, they may be important for
the increase activity of renin. Next, we investigated the
capacity of membrane-bound prorenin to activate
angiotensinogen. The determination of recombinant
prorenin antigen in our preparation of prorenin with
the RIA Renin III generation kit, which uses an antirenin active site Ab as secondary Ab, showed that about
25% of prorenin could be detected, indicating that the
active site of prorenin was accessible in about 25% of the
molecules. This prorenin preparation was used for binding to HMC2 membranes and for measuring the extent
of Ang I generation as described. Prorenin bound to
membranes was eluted by pH 3.5 treatment and measured with the RIA Renin III generation kit. Acidification
resulted in a total activation of prorenin (not shown).
Accordingly, in the control experiments identical
amounts of prorenin were incubated with angiotensinogen, in the absence of membranes. The results showed
that at pH 7.4, little Ang I–generating activity was
observed with 0.5 nM prorenin in solution after 4 hours
of incubation, compared with active renin under similar conditions. In contrast, when prorenin was immobilized on HMC2 membranes, the amount of Ang I generated was measurable and almost comparable to that
generated by fully active renin in solution studied under
similar experimental conditions (Figure 4d).
Cross-linking with renin and phosphorylation of the receptor.
Immunoprecipitation of the receptor from transfected
cells labeled with 35S-methionine and with the poly1422
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clonal anti-receptor Ab showed that the receptor has an
apparent molecular weight of 45 kDa, greater than the
in vitro translation product (39 kDa), indicating that
posttranslational modifications have occurred in the
eukaryotic cell (Figure 5a, lane A). Our previous results
have suggested a molecular weight of 70–80 kDa for
the receptor on primary human mesangial cells (21).
Therefore, we performed cross-linking experiments of
renin bound to transformed cells. The results showed
the appearance of a 110- to 120-kDa band, confirming
our previous study (Figure 5a, lane C). Precipitation of
cell lysate with anti-FLAG Ab during the phosphorylation experiments confirmed a molecular weight of 45
kDa for the receptor as found by immunoprecipitation
of 35S-methionine–labeled cell lysate with the antireceptor Ab. Renin stimulation induced a phosphorylation of serine and tyrosine residues of the receptor,
but not of threonine (not shown). The phosphorylation of serine residues occurred more rapidly than for
tyrosine and was already detectable after 3 minutes in
the presence of renin, whereas the phosphorylation of
tyrosine residues appeared after 10 minutes (Figure 5b,
upper panel). No phosphorylation corresponding to a
45-kDa band could be observed in control cell lysate
(Figure 5b, lower panel).
Intracellular [Ca2+] and cAMP changes, MAP kinases
ERK1(p44)/ERK2(p42) activation induced by renin. To further support the receptor function of the cloned renin
receptor, we studied the effect of renin binding on
[Ca2+] concentration change, on cyclic AMP, and on
MAP kinases ERK1(p44)/ERK2(p42) activation using
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two different techniques: Western blot analysis of total
and active phosphorylated ERK1 and ERK2 and a MAP
kinase activity assay. No changes in [Ca2+] or in cyclic
AMP could be evidenced after renin addition (Figure
6a). On the contrary, renin addition induced a rapid
activation of ERK1/ERK2 resulting in a twofold
increase of active ERK1/ERK2 after 0.5 min (P < 0.05)
and a three- to fourfold increase after 10 minutes and
for at least 60 minutes (Figure 6b, left panel). The blot
of total ERK1/ERK2 confirmed an even loading of the
gels and the ratio of active-to-total ERK1/ERK2 was
made using the NIH IMAGE program (Figure 6b, left).
These data were confirmed by a MAP kinase activity
assay (Figure 6b, left bottom). No activation of
ERK1/ERK2 was observed in cells not expressing the
receptor (Figure 6b, right panel).
Expression of renin receptor mRNA in human adult tissues
by Northern blot analysis. A single transcript of 2.4 kb was
found in the brain, placenta, heart, liver, kidney, and
pancreas. Because the amount of polyA+ mRNA immobilized on the membrane was adjusted for β-actin, the
very strong signal observed in the brain, heart, and placenta indicates very high levels of expression of the
receptor mRNA in those tissues. The mRNA of the
renin receptor was barely detectable in lung and skeletal muscle (Figure 7).
Immunofluorescence staining and confocal microscopy on
normal human kidney, heart, and placenta. Immunofluorescence staining of kidney cortex with the anti-receptor Ab showed labeling of glomeruli and vascular structures and no staining of the tubules and, at higher
magnification, confirmed that the labeling was restricted to the mesangium (Figure 8a, left). The specificity of
the staining was assessed by preincubation of the
immune serum with the peptides (Figure 8a, right).
Double-labeling with anti-CD31 Ab specific for
endothelial cells and with anti-renin receptor Ab and
analysis by confocal microscopy of kidney cortex
showed that the renin receptor did not colocalize with
CD31 in endothelial cells in glomeruli but was localized
in the mesangial area. In kidney arteries, the renin
receptor was not associated with the endothelium
either (Figure 8, middle panel). To compare the vascular localization of the renin receptor in another tissue,
we also studied the distribution of the receptor in the
heart. The results showed that in coronary arteries, as
in kidney arteries, the renin receptor did not colocalize
with CD31 on endothelial cells but was found in the
subendothelium (Figure 8, middle panel). To better
localize the renin receptor in the subendothelium of
kidney and coronary arteries, we performed a doublestaining with anti-receptor and anti–smooth muscle
α-actin Ab. The results showed that the renin receptor
colocalized with smooth muscle α-actin, indicating
that renin receptor was present on smooth muscle cells
(Figure 8, lower panel). The absence of evidence for a
colocalization of both renin receptor and smooth muscle α-actin in normal glomeruli can be attributed to the
absence of smooth muscle α-actin expression by
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human mesangial cells in normal, noninflammatory
glomeruli (31). Finally, we looked for a colocalization
of renin and of the receptor in normal tissue. Placenta
is known to secrete large amounts of renin and
prorenin throughout gestation (7), and Northern blot
analysis also showed abundant expression of the receptor in placenta. Immunofluorescence staining showed
that the receptor was found in mature placenta, associated to vascular structures and to syncytiotrophoblast
cells (Figure 9). In addition, staining of sequential section of placenta with renin Ab showed that renin and
renin receptor had a similar pattern of distribution in
arteries and in syncytiotrophoblast cells (Figure 9).
Figure 5
Cross-linking with renin and phosphorylation of the receptor. (a)
HMC and control cells were labeled with 35S-methionine, and the
receptor was immunoprecipitated with anti-receptor Ab (lane A and
lane B for HMC and control cells, respectively) or after incubation
with renin and cross-linking (lane C and lane D for HMC and control cells, respectively). In lane E, HMC cells cross-linked in the
absence of renin. (b) In the upper panel HMC2 cells were stimulated with renin in the presence of 100 nM Captopril. The receptor was
immunoprecipitated with anti-FLAG agarose, and the eluate was
analyzed by Western blotting using Ab’s to phosphotyrosine or to
phosphoserine. In the lower panel control cells were stimulated with
renin, and the cell lysate was analyzed by Western blotting using
Ab’s to phosphotyrosine or to phosphoserine. The right lanes of the
two lower blots are the receptor immunoprecipitated from HMC
cells stimulated by renin.
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Figure 6
Intracellular calcium and cAMP changes, MAP kinases ERK1(p44)/ERK2(p42) activation induced by renin. (a) Cells expressing the receptor were stimulated by 100 nM renin, and the intracellular calcium changes were analyzed by spectrofluorometry (top). Cell stimulation
by human thrombin (10 nM) was used as control. Analysis of intracellular cAMP changes by cells stimulated by 10 nM renin and by 1 µM
PGE-1 or by 1 µM isoproterenol (Isop), as positive controls (bottom). The results represent the mean ± SD of two experiments performed
in triplicate. *P < 0.05 compared with basal value. (b) HMC2 cells (left panel) or control cells (right panel) were stimulated with renin in
the presence of 1 µM Losartan. At intervals, the cells were lysed and the lysate analyzed by Western blotting using Ab’s to active or to total
ERK1 and ERK2. The blots were scanned and the ratio of active, phosphorylated ERK1/ERK2 to total ERK1/ERK2 was plotted using the
NIH IMAGE program. MAP kinase activity assay was performed on HMC2 renin-stimulated cells (left bottom). The results are expressed
as 32P incorporated and represent the mean ± SD of two experiments performed in triplicate. *P < 0.05 compared with basal value.
Discussion
This study reports, we believe for the first time, the existence of a functional receptor of renin. The evidence
that N14F cDNA is identical to the receptor of renin is
as follows: (a) The N14F protein expressed in vitro is
able to bind renin in coprecipitation experiments; (b)
cells transfected with N14F cDNA construct and
expressing the protein bind renin with high affinity,
and the binding is specific for renin and prorenin; (c)
renin binding induces the activation of MAP kinases
ERK1 and ERK2 associated with a phosphorylation of
tyrosine and serine residues.
The localization of the receptor in the mesangium of
glomeruli suggests that this receptor is the one we
described on human mesangial cells in culture. In pri-
mary mesangial cells (21) we have observed a molecular weight of 70–80 kDa for the receptor, whereas the
cloned renin receptor here is 45 kDa. This discrepancy
may be explained by a dimerization of the receptor
induced by renin binding as described for receptors of
the tyrosine-kinase family, which are also single-transmembrane domain receptors (32), or a complex formation with another, yet-unidentified protein. Part of this
clone was identical to the previously reported “M8-9,”
a truncated protein of 8.9 kDa that copurified with a
proton-ATPase of chromaffin granule membranes (33).
While the sequencing of our N14F clone was completed, two identical cDNA sequences from human fetal
brain and human hypothalamus appeared in Genbank
(DKFZ p56400582, accession number AL049929.1 and
Figure 7
Expression of the receptor mRNA. Northern blot analysis. After
hybridization of the human blot, a strong signal corresponding to a
2.4-kb mRNA band was detected in heart, brain, and placenta, a
weaker signal was seen in liver, pancreas, and kidney. The mRNA of
the receptor was hardly detected in lung and skeletal muscle. The
amount of RNA on the membrane was adjusted so that β-actin
hybridization is comparable in every lane.
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Figure 8
Expression of the renin receptor on
human kidney and heart. (a) At the
left, immuofluorescence staining of
kidney cortex with anti-receptor Ab
(upper panel magnification ×20)
and a glomerulus at higher magnification (lower panel, ×40). Yellow
staining is due to autofluorescence
of degradation products in tubular
cells. The immune serum was preincubated with the peptides (right).
(b–m) Double-staining and analysis
by immunofluorescence and confocal microscopy of kidney cortex
labeled with anti-renin receptor (b,
e, h), anti-CD31 (c, f, i), and antireceptor plus anti-CD 31 (d, g, j).
e–g represent the glomerulus in b–d,
and h–j represent the artery in b–d at
higher magnification. (k–m) A coronary artery labeled with anti-receptor, anti-CD31, anti-receptor plus
anti-CD 31 Ab, respectively. (n–v)
Double-staining and analysis by
immunofluorescence and confocal
microscopy of kidney cortex labeled
with anti-renin receptor (n and q) or
smooth muscle α-actin Ab (SM-αactin; o and r) and anti-receptor
and anti–SM-α-actin (p and s); q–s
represent the kidney cortex artery in
panels n and q at higher magnification. (t–v) A coronary artery stained
with anti-receptor, anti–SM-α-actin,
and anti-receptor plus anti–SM-αactin Ab, respectively.
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Figure 9
Expression and colocalization of renin and of the renin receptor in
placenta. Sequential sections of placenta were stained with the Ab
to the receptor and FITC-coupled secondary anti-rabbit Ab (a–c) or
with anti-renin Ab and Texas-red–coupled secondary Ab (e–g). (d)
FITC-secondary Ab alone. (h) TRITC–coupled secondary Ab alone.
The receptor and renin have similar distribution in vascular structures
and in syncytiotrophoblast cells.
AF248966), confirming the high level of expression of
the receptor in the human CNS.
The physiological plasma concentration of renin is in
the picomolar range, but it is admitted that in tissues,
especially in interstitial fluids, the renin concentration
may be 100-fold greater. Since we found that the dissociation constant of the renin-receptor complex was in
the nanomolar range, we postulated that it could not be
considered as a binding protein responsible for retaining renin in the interstitium because only about 1% of
the interstitial renin (10 pM) would be bound by the
receptor. Alternatively, it could function as an effective
receptor. Therefore, we looked for [Ca2+] change and for
cyclic AMP and MAP kinases ERK1(p44)/ERK2(p42)
activation. Our results showed that renin binding did
not modify either intracellular calcium or cAMP, but
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provoked a rapid activation of the ERK1/ERK2 pathway, associated with a phosphorylation of serine and
tyrosine residues. The presence of either an ACE
inhibitor or an AT1 receptor antagonist during the
incubation of transformed cells with renin confirms
that these events are independent of a possible generation or action of Ang II. The pathway involved in the
tyrosine phosphorylation of the receptor, the respective
roles of serine and tyrosine phosphorylation in the
receptor function, and the early phase of renin signal are
actually under investigation.
We have shown that receptor-bound renin activates
angiotensinogen with kinetics different from those
observed for renin in solution and, in particular, we
showed a reduction in the Km for angiotensinogen from
1 µM in solution to 0.15 µM with receptor-bound renin.
This Km is significantly below the normal plasma concentration of angiotensinogen, approximately 1 µM
(29), suggesting that conversion of angiotensinogen by
receptor-bound renin may be of physiological importance, especially in tissues in which the concentration of
angiotensinogen is much lower than in plasma. We
hypothesize that receptor-bound renin is able to activate
angiotensinogen while it is actually bound to the cell
surface and with higher efficiency than renin in solution. The slight decrease of kcat (1.5-fold reduction)
observed concomitantly might reflect the constraints
imposed by the immobilization of renin on the receptor. The overall catalytic efficiency kcat/Km increased four
to five times, indicating that the cell surface is an important site for angiotensinogen activation. Taken together with the recent report of the existence of a vascular
smooth muscle chymase (34), our results suggest that
the smooth muscle cell surface may play an essential
role in tissue generation of Ang II. The binding of renin
to its receptor, thereby increasing angiotensinogen
cleavage efficiency, would facilitate Ang I cleavage in
Ang II by vascular smooth muscle chymase.
We also hypothesized that bound prorenin would activate angiotensinogen. Although our prorenin preparation was reactive against Ab’s to the active site of renin,
suggesting that some molecules have an accessible active
site, prorenin showed very little Ang I–forming activity
when incubated at pH 7.4 for 4 hours. However, the
extent of Ang I generated by prorenin bound to membranes was comparable to the extent of Ang I generated
by fully active renin under these experimental conditions. Whether the increase of the catalytic activity of
membrane-bound prorenin should be attributed to the
fraction of prorenin with an already exposed active site
or to inactive prorenin that underwent conformational
changes induced by receptor binding is an important
issue to investigate in the near future. Supporting evidence for catalytically active prorenin in vivo, in the
absence of prorenin cleavage, exists in the literature (35).
Methot et al. generated double-transgenic mice expressing human angiotensinogen and a mutated, noncleavable human prorenin in the pituitary gland (35). These
animals have elevated pituitary Ang I content in the
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absence of prorenin cleavage as shown by Western blot
analysis, indicating that prorenin was enzymatically
active even with the prosegment still in place.
Knockout mice for angiotensinogen, Ang converting
enzyme, Ang II receptors, and nullizygotes for the renin
gene, display low blood pressure and severe renal vascular lesions associated with high or low plasma renin
concentrations (36–39). The vascular lesions have been
attributed to the absence of Ang II during development,
but these data do not exclude a role of renin in the
pathogenesis of vascular lesions. Moreover, rats expressing the prorenin transgene exclusively in the liver suffer
severe nephroangiosclerosis, cardiac and aortic hypertrophy, and liver fibrosis in the absence of hypertension
(40). This observation suggests that the effects observed
in vitro on the activation of ERK1/ERK2 pathway
involved in cell hypertrophy and proliferation, as well as
the cellular hypertrophy and the increase of PAI1 (21)
synthesis, may be relevant in vivo.
In conclusion, our results show that renin receptor
exerts dual effects. The renin receptor is able to trigger
intracellular signal by activating the ERK1/ERK2 pathway, and it also acts as a cofactor by increasing the efficiency of angiotensinogen cleavage by receptor-bound
renin, therefore facilitating Ang II generation and action
on a cell surface. This novel concept may provide a new
approach to a better understanding of the pathogenesis of vascular diseases associated with RAS activation.
Acknowledgments
We would like to thank Walter Fischli and Volker Breu
for their constant intellectual and material support. We
thank Oliver Nayler for helpful advice in the construction of the FLAG receptor. We also wish to thank Pierre
Corvol, François Alhenc-Gelas, Eric Rondeau, and Raymond Ardaillou for stimulating discussions, Tam-Tam
Guyenne for her advice on the Ang I assay, and
Madeleine Delauche and Philippe Fontanges for expert
technical assistance for confocal microscopy studies.
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