Smooth Muscle Specific Rac1 Deficiency Induces Hypertension by Preventing p116^RIP3‐Dependent RhoA Inhibition

Abstract

Background

Increasing evidence implicates overactivation of RhoA as a critical component of the pathogenesis of hypertension. Although a substantial body of work has established that Rac1 functions antagonize RhoA in a broad range of physiological processes, the role of Rac1 in the regulation of vascular tone and blood pressure is not fully elucidated.

Methods and Results

To define the role of Rac1 in vivo in vascular smooth muscle cells (vSMC), we generated smooth muscle (SM)‐specific Rac1 knockout mice (SM‐Rac1‐KO) and performed radiotelemetric blood pressure recordings, contraction measurements in arterial rings, vSMC cultures and biochemical analyses. SM‐Rac1‐KO mice develop high systolic blood pressure sensitive to Rho kinase inhibition by fasudil. Arteries from SM‐Rac1‐KO mice are characterized by a defective NO‐dependent vasodilation and an overactivation of RhoA/Rho kinase signaling. We provide evidence that Rac1 deletion‐induced hypertension is due to an alteration of cGMP signaling resulting from the loss of Rac1‐mediated control of type 5 PDE activity. Consequently, cGMP‐dependent phosphorylation and binding of RhoA with its inhibitory partner, the phosphatase‐RhoA interacting protein (p116^RIP3), are decreased.

Conclusions

Our data reveal that the depletion of Rac1 in SMC decreases cGMP‐dependent p116^RIP3/RhoA interaction and the subsequent inhibition of RhoA signaling. Thus, we unveil an in vivo role of Rac1 in arterial blood pressure regulation and a new pathway involving p116^RIP3 that contributes to the antagonistic relationship between Rac1 and RhoA in vascular smooth muscle cells and their opposite roles in arterial tone and blood pressure.

Introduction

The Ras homologous (Rho) family of small GTPases contains 20 members in humans among which RhoA, Rac1, and Cdc42 are the best characterized. One of the major functions of these proteins is the spatial and temporal regulation of the actin cytoskeleton assembly during essential cell processes such as migration, division, and adhesion.^1–3 To precisely modulate these cellular functions, the activity of Rho GTPases is finely regulated by mechanisms that include reciprocal negative or positive feedbacks. Typically, numerous studies have revealed a link between RhoA and Rac1 activities. It was initially proposed that activated Rac1 positively regulates RhoA and this finding has been validated in cells from Rac1‐deficient mice.^4–5 However, in most situations, Rac1 and RhoA display an antagonistic relationship^6–9 that could result from several different mechanisms affecting their activity, expression and stability, and their downstream signaling pathways.⁷

In the vascular system, Rho GTPases have been increasingly implicated in the regulation of different physiological and mechanoelastic responses. In endothelial cells, RhoA and its main effector Rho kinase negatively regulate NO production by decreasing endothelial nitric oxide synthase (eNOS) expression and activity.^10–11 Rac1 has exactly opposite effects, enhancing eNOS activity and increasing eNOS expression through elevated eNOS mRNA stability and promoting eNOS gene transcription via its effector p21‐activated kinase 1 (Pak1).^12–13 This role of Rac1 in endothelial cells was further confirmed in vivo in endothelial Rac1 haploinsufficient mice (EC‐Rac1^+/−). These mice developed mild hypertension correlated with a decreased expression and activity of eNOS.¹² Endothelial Rac1 was also described to control the production of reactive oxygen species (ROS) and the mobilization of cortical actin network to modulate endothelial permeability.^14–16

In vascular smooth muscle cells (vSMC), RhoA and Rho kinase are recognized as major regulators of the contraction by inhibiting myosin light chain phosphatase (MLCP), thereby increasing MLC phosphorylation.¹⁷ Accordingly, RhoA/Rho kinase signaling is a key regulator of vascular tone and arterial pressure, and overactivation of RhoA/Rho kinase signaling has been identified as a critical mechanism in the pathogenesis of pulmonary or systemic hypertension.¹⁸ Several vasoconstrictors have also been shown to activate Rac1 in vSMC, however its role in the control of vascular tone is unclear.¹⁹ Rac1/Pak1 signaling has been described to phosphorylate and inhibit MLC kinase, thereby decreasing MLC phosphorylation and smooth muscle cell contraction.^20–21 However, opposite effects have also been reported^22–23as Pak3, another Rac1 effector, has been shown to enhance smooth muscle contraction through phosphorylation of the thin filament regulatory protein caldesmon.^22–23 In the absence of relevant animal models, the in vivo role of vSMC Rac1 in the control of vascular tone and blood pressure remains unknown. In fact, mouse models of ubiquitous deletion of Rac signaling components such as exchange factors of the Vav family display complex cardiovascular phenotypes resulting from multiple defects including sympathetic nervous system and neurohormonal dysfunctions.^24–25

We therefore generated a new animal model with inducible inactivation of the Rac1 gene in SMC. We demonstrated that Rac1 deletion in SMC induces high systolic blood pressure in mice through an alteration of cGMP signaling that decreases NO‐induced p116^RIP3/RhoA interaction, leading to a defective NO‐mediated RhoA inhibition and vasorelaxation.

Methods

Animals Use

All experimental procedures and animal care were performed in accordance with the European Community Standards on the Care and Use of Laboratory Animals and approved by the local ethics committee (Comité d'Ethique en Expérimentation Animale des Pays de Loire). We mated a transgenic mouse line carrying floxed alleles of the gene coding for Rac1 (Rac1^lox/lox, Jackson Laboratories) to mice expressing a fusion protein of the Cre recombinase with a modified estrogen receptor–binding domain (CreER^T2) under the control of the smooth muscle myosin heavy chain (SMMHC) promoter (SMMHC‐CreER^T2 mice)^26–27 to produce SMMHC‐CreER^T2;Rac1^lox/lox mice (SM‐Rac1^lox/lox mice). The recombinase Cre was activated in 2‐month‐old mice by intraperitoneal injection of tamoxifen (1 mg/day dissolved in sun‐flower oil, T5648, Sigma) for 5 consecutive days during 2 weeks. Male mice were analyzed 1 month after tamoxifen treatment. Rac1^lox/lox mice treated with tamoxifen and SM‐Rac1^lox/lox mice without tamoxifen treatment were used as control animals. In this study, 85 SM‐Rac1^lox/lox and 95 Rac1^lox/lox mice were used for functional analyses.

Immunoblot Analysis

vSMC or cleaned aortas were incubated on ice with lysis buffer supplemented with proteases and phosphatases inhibitors cocktails (Sigma) and sodium orthovanadate. Lysates were subjected to SDS‐PAGE, transferred to nitrocellulose membranes and incubated with specific antibodies. Immune complexes were detected with appropriate secondary antibodies and enhanced chemiluminescence reagent (ECL plus; GE Healthcare). Protein band intensities were quantified using ImageJ Software (NIH software).

Histological Analysis

Hearts, kidneys, and aortas were fixed in 4% paraformaldehyde in PBS and embedded into paraffin. We stained 6 μm sections with hematoxylin and eosin (Sigma). Aorta media wall thickness and cellular density were quantified in a blind manner using Metamorph‐Metaview software (Universal Imaging).

Arterial Pressure Measurements

Blood pressure was measured in conscious and unrestrained mice using a radiotelemetry system as described previously²⁸ (PA‐C10 and Dataquest software; Data Sciences International). L‐NAME treatment (N5751; Sigma) was administrated in the drinking water (300 mg/kg of body weight/day) and renewed every 3 days. The recording room was maintained with a 12‐hour‐light/12‐hour‐dark cycle. Blood pressure was also measured in restrained mice with a noninvasive tail‐cuff device (BP 2000; Visitech Systems). Fasudil treatment (F4660; LC laboratories) was administrated by intraperitoneal injection (5 or 30 mg/kg of body weight) 20 minutes before measurements.

Echocardiography

Two‐dimensional (2‐D) echocardiography was performed on mice using a Vivid 7 Dimension ultrasonography (GE Healthcare) with a 14‐MHz transducer. Left ventricular free wall thickness, anterior wall, and posterior wall thickness were measured during diastole and systole from long‐ and short‐axis images obtained by M‐mode echocardiography. Transmitral flow measurements of ventricular filling velocity were obtained using pulsed Doppler, with an apical four‐chamber orientation. Thus, early diastolic (E), late diastolic (A), and the ratio, E/A were obtained to assess diastolic dysfunction. To avoid bias in the analysis, blind experiments were done and scored by members of our research team.

Renal Function

Physiocages were used to evaluate urine production during 48h after a 3‐day acclimatizing period. Urines were collected and analyzed at the ONIRIS laboratory facility (Nantes, France) to determine creatinine clearance and Na⁺ concentration.

Arterial Reactivity

To analyze the pressure and the flow‐dependent tone, segments of first‐order mesenteric arteries were removed and cannulated at both ends in a video‐monitored perfusion system (Living System Instrumentation) as previously described.²⁹ Briefly, arteries were bathed and perfused with a physiological salt solution. To assess myogenic tone, diameter changes were measured when intraluminal pressure was set from 10 to 125 mm Hg without intraluminal flow (active diameters). At the end of each experiment, arterial segments were superfused with a Ca²⁺‐free physiological salt solution containing EGTA (2 mmol/L), sodium nitroprusside (10 μmol/L) and papaverin (10 μmol/L), and pressure steps (10 to 125 mm Hg) were repeated (passive diameter). Myogenic tone was expressed as active tone (passive diameter − active diameter) in percentage of passive diameter. For flow‐dependent tone, arteries were submitted to a 75 mm Hg pressure and preconstricted with phenylephrine (Phe; 0.3 to 1 μmol/L) to maintain a stable tone (≈50% of the maximum response). Intraluminal flow was increased by step (0 to 100 μL/min) and diameter measured. Flow‐mediated relaxation was expressed as the percentage of dilation of the active tone (preconstriction).

To analyze aortic and mesenteric arterial vasoreactivity, 2‐mm‐long rings were cut, cleaned, and mounted on a multichannel isometric myograph (Danish Myo Technology) containing a Krebs‐Henseleit solution (in mmol/L: 118.4 NaCl, 4.7 KCl, 2 CaCl₂, 1.2 MgSO₄, 1.2 KH₂PO₄, 25 NaHCO₃, and 11 glucose). We constructed cumulative concentration‐response curves in response to KCl, Phe, and the thromboxane A2 receptor agonist (U46619). Relaxation‐response curve in response to carcachol (CCh), S‐Nitroso‐N‐Acetyl‐DL‐Penicillamine (SNAP), and isoprenaline (IsoP) were constructed after Phe precontraction. When indicated, rings were pretreated with L‐NAME (100 μmol/L) 1 hour before contraction measurements. Chamber wire myograph was connected to a digital data recorder (MacLab/4e; AD Instruments) and recordings were analyzed using LabChart v7 software (AD Instruments).

NO Signaling Pathway Analysis

Plasmatic nitrites concentration was determined using Griess Reagent Kit (Molecular probes; Invitrogen). cGMP and cAMP concentrations were evaluated in aorta and transfected vSMC according to the supplier's specifications (Cyclic GMP and AMP EIA Kit; Cayman Chemical).

Analysis of Rac1 and RhoA Activity

Pull‐down assay using GST‐PBD or GST‐RBD fusion proteins were performed on fresh vSMC or aorta lysates to evaluate Rac1 and RhoA activities, respectively.³⁰

Cell Culture, Transfection and Actin Staining

Primary aortic smooth muscle cells were isolated and cultured as described previously.³¹ cDNA coding for wild‐type Rac1 (Rac^WT), wild‐type RhoA (GFP‐RhoA^WT), constitutive inactive form of Rac1 (Rac^N17), constitutive active Rac1 (Rac^V12), phospho‐mimetic RhoA (GFP‐RhoA^S188E), and phospho‐resistant RhoA (GFP‐RhoA^S188A) were transfected in vSMC with jetPEI (Polyplus transfection) according to the manufacturer's instructions. To selectively knockdown the expression of endogenous p116^RIP3, two siRNA oligonucleotides were designed (Eurogentec): siRNA1 5′‐GGUCCCAGGUAAUUGAGAA+dTdT‐3′ and siRNA2 5′‐GAGCACAUGGAAACCAACA+dTdT‐3′. RNA duplexes or a siRNA negative control oligonucleotide (SR‐CL000‐005; Eurogentec) were transfected by Lipofectamine RNAiMAX (Invitrogen) according to the supplier's specifications. When indicated, cells were treated 1 hour with SNP (10 μmol/L), IsoP (10 μmol/L) or 8‐Br‐cGMP (100 μmol/L). After treatments, vSMC were fixed with 4% paraformaldehyde in PBS, permeabilized in PBS 0.5% Triton X‐100, and incubated with 130 μg/mL of FITC‐conjugated phalloidin (Sigma) or 5 mg/mL of Texas red‐DNaseI (Molecular Probes; Invitrogen) to visualize F‐actin and monomeric G‐actin, respectively. After staining, images were captured by a fluorescence microscope (Nikon) and quantified using Metamorph‐Metaview software (Universal Imaging). The ratio of fluorescence of FITC‐phalloidin and Texas red‐DNaseI (F‐actin/G‐actin ratio) was used to quantify the actin cytoskeleton organization.

Co‐Immunoprecipitation

Immunoprecipitation of RhoA in aorta to reveal p116^RIP3 was carried out with a polyclonal RhoA antibody (1/500) using A‐protein magnetic beads (μMACS 130‐071‐001; Milteny) according to the supplier's specifications. Anti‐GFP‐beads (μMACS 130‐091‐125; Milteny) were used for GFP immunoprecipitation. The coimmuniprecipitations for the others were performed with anti‐Pak1 (1/500) or anti‐PKG (1/1000) antibodies and G‐Sepharose beads (fast‐flow; GE Healthcare) overnight at 4°C followed by washes with the lysis buffer.

Chemicals and Antibodies

Antibodies against Rac1 (610651) and eNOS/NOS (610297) were from BD Biosciences. Anti‐MYPT1 (2634), P‐Pak (2605), Pak1 (2602), PDE5 (2395), PKG (3248), and RhoA (2117) antibodies were from Cell Signaling Technology. Anti‐P‐MYPT (sc17556), p116^RIP3 (sc135494) and P^Ser188‐RhoA (sc32954) antibodies were from Santa Cruz Biotechnology. Anti‐soluble guanylyl cyclase sGCα1 (G4280) and sGCβ1 (G4405) antibodies were from Sigma. Anti‐eGFP antibody was from Clontech (632569). All agonists used for the analysis of blood vessels reactivity and vSMC treatment were from Sigma when not specified.

Statistics

Results are expressed as the mean±SEM of sample size n. Statistical analysis were performed with either the 2‐tailed Student's t test or the 1‐way ANOVA test. For small sample sizes, a nonparametric test (Mann‐Whitney) was applied. A value of P<0.05 or less was considered to be statistically significant.

Results

Generation of Mice With Smooth Muscle‐Specific Rac1 Deficiency

Mice specifically lacking Rac1 in SMC (SM‐Rac1‐KO mice) were obtained by treating SM‐Rac1^lox/lox mice with tamoxifen. Wild‐type and floxed Rac1 alleles were identified by PCR (WT, 115 bp; floxed, 242 bp) and Cre‐mediated recombination generated a 140 bp band allowing the identification of SM‐Rac1‐KO mice (Figure S1A). In situ hybridization and RT‐PCR confirmed the inhibition of Rac1 mRNA expression in vSMC from SM‐Rac1‐KO mice (Figure S1B and S1C). Likewise, Rac1 protein was poorly detected by immunoblot and immunohistochemistry in aortic vSMC from SM‐Rac1‐KO mice (Figure S1D and S1E).

Rac1‐Deficient Mice Develop High Systolic Blood Pressure

The impact of SM Rac1 deletion on blood pressure and heart rate was evaluated by radiotelemetry. Before treatment with tamoxifen, baseline heart rate, systolic, diastolic and mean arterial blood pressures were similar in control and SM‐Rac1^lox/lox mice (Figures 1A, 1B, S2A, and S2B). Tamoxifen treatment was rapidly followed by an increase in systolic blood pressure in SM‐Rac1‐KO mice (Figure 1A). One month after induction of Rac1 deletion in SMC, systolic blood pressure was significantly higher in SM‐Rac1‐KO mice compared to tamoxifen‐treated Rac1^lox/lox mice (Figure 1B; Table) and SM‐Rac1^lox/lox mice without treatment (data no shown), during both 12‐hour‐dark and 12‐hour‐light periods. This difference in systolic blood pressure was maintained over time (Figure S3B) and was not associated with change in diastolic blood pressure and heart rate (Figures S2A and S2B and Table). SM‐Rac1‐KO mice developed thickening of the aortic wall typical of the hypertensive remodeling observed in mice models,^24–25 and patients with high blood pressure^24–25,32(Figure S3A; and Table). At the cardiac level, echo Doppler measurements revealed that left ventricle thickness was slightly enlarged indicating cardiac hypertrophy initiation in response to the elevated systolic blood pressure (Table). However, the heart weight to body weight ratio was similar in SM‐Rac1‐KO and Control mice (Table). Measurements of blood pressure in 14‐month‐old SM‐Rac1‐KO mice showed persistent high blood pressure, without any sign of myocardial fibrosis (Figure S3B and S3C).

Figure 1.

Rac1 deficiency in SMC induces an increase of systolic arterial blood pressure. A, Telemetric records of systolic blood pressure in Rac1^lox/lox (n=8) and SM‐Rac1^lox/lox (n=9) mice. Tamoxifen treatment period to induce Rac1 deletion is indicated. B, Profiles over 24 hours of systolic blood pressure. Measurements were performed under a 12:12‐hour light/dark schedule. Left panels represent analysis before induction of Rac1 deletion and right panels after tamoxifen treatment (n=8 to 9). *P<0.05. SM‐Rac1 indicates smooth muscle Rac1 knockout.

Table 1.

Cardiovascular Parameters in Rac1^lox/lox and SM‐Rac1‐KO Mice

	Rac1lox/lox	SM‐Rac1‐KO	P Value	n
Body weight, g	27.2±0.6	27.1±0.8	0.46	22
Restrained systolic ABP, mm Hg	102±3	120±4	0.01*	10
Telemetric systolic ABP (mm Hg) (24 hours)	127±2	142±3	0.02*	9
Telemetric diastolic ABP (mm Hg) (24 hours)	95±1	97±2	0.19	9
Relative aorta media wall thickness	1.0±0.07	1.33±0.07	0.02*	4
Relative aortic cellular density	1.0±0.06	1.23±0.03	0.01*	4
Aortic diameter, μm	417±16	407±19	0.29	8
Mesenteric artery diameter, μm	188±12	192±8	0.35	8
Aortic elastance (% control)	100±8	104±5	0.28	8
Mesenteric artery elastance (% control)	100±4	120±6	0.01*	8
Heart (mg)/body weight (g)	4.9±0.3	4.8±0.1	0.24	7
LV systolic thickness, mm	1.78±0.06	2.08±0.17	0.12	8
LV diastolic thickness, mm	1.26±0.07	1.69±0.15	0.02*	8
Heart rate, bpm	532±21	564±9	0.20	8
E/A ratio	1.47±0.07	1.18±0.07	0.01*	8
Kidney weight, mg	180±1	183±4	0.30	4
Creatinine clearance, mL min−1 kg−1	4.29±0.53	3.82±0.45	0.34	6
FENa, %	0.44±0.03	0.45±0.05	0.38	6

Arterial characteristics were monitored either by tail‐cuff, telemetry, histological or myograph measurements. Cardiac parameters were obtained by echocardiography. Renal function was assessed by sodium and creatinine related dosages. All parameters were measured 1 month after Rac1 deletion. ABP indicates arterial blood pressure; E/A, early diastolic (E), late diastolic (A), LV, left ventricle; Fe_Na, fractional sodium excretion; n, sample size per group; SM‐Rac1, smooth muscle Rac1 knockout.

^*P<0.05 Rac1^lox/lox vs SM‐Rac1‐KO mice.

Analysis of the renal function that plays a major role in the regulation of blood pressure, showed that glomerular filtration rate (creatinine clearance) and excretion rate of Na⁺ were not altered in SM‐Rac1‐KO mice (Table). Moreover, histological analysis did not reveal any structural remodeling such as renal fibrosis or glomerulosclerosis in 14‐month‐old SM‐Rac1‐KO compared with control mice (Figure S3C). These data thus suggest that the rise in systolic pressure in SM‐Rac1‐KO mice is not related to renal dysfunction.

Rac1 Deficiency in SMC Leads to Impaired NO‐Dependent Vasodilation

To shed light on the origin of high systolic blood pressure, we explored the functional properties of SM‐Rac1‐KO arteries. Arterial diameters were similar in both group of mice, but a slight increase of elastance was observed in mesenteric arteries from SM‐Rac1‐KO mice compared to control mice (Table). Pressure‐induced tone (myogenic tone) was not affected by SMC Rac1 deletion (Figure 2A). The contractile response to KCl was similar in the aorta, mesenteric, and renal arteries from SM‐Rac1‐KO mice compared with control mice (Figures S4, S5A, and S6A). The responses to the thromboxane A2 analog (U46619) and phenylephrine (Phe) were slightly increased in mesenteric arteries from SM‐Rac1‐KO mice (Figures S4 and S5A). In contrast, flow‐induced relaxation was significantly reduced in mesenteric resistance arteries from SM‐Rac1‐KO mice as compared to Rac1^lox/lox mice (Figure 2B). Flow‐mediated dilation mainly depends on the capacity of vSMC to respond to flow‐induced endothelium‐derived vasodilator agents such as nitric oxide (NO) and cyclooxygenase‐mediated prostacyclins. The altered relaxation observed in arteries from SM‐Rac1‐KO mice suggests that Rac1 might be involved in response to these vasodilator factors.

Figure 2.

NO‐dependent relaxation is impaired in SM‐Rac1‐KO mice. A, Pressure‐induced myogenic tone and (B) flow‐induced vasodilation in mesenteric arteries from Rac1^lox/lox and SM‐Rac1‐KO mice (n=8). C, Representative real‐time recordings of the response of Phe‐constricted mesenteric arteries to cumulative administration (arrows) of CCh (10⁻¹⁰ to10⁻⁵ mol/L). D, Percentage of vessel relaxation induced by the indicated doses of CCh, SNAP and IsoP on Phe‐constricted mesenteric arteries from Rac1^lox/lox and SM‐Rac1‐KO mice pretreated or not with L‐NAME (1 hour, 100 μmol/L; n=5 to 12). E, Systolic blood pressure in Rac1^lox/lox (n=8) and SM‐Rac1‐KO (n=9) mice chronically treated with L‐NAME (300 mg/kg of body weight/day in drinking water). F, Plasma nitrite measurements 5 and 15 days after tamoxifen treatment (n=5 to 10). Log M, logarithm of the molar concentration used for each agent. *P<0.05 compared with controls. CCh indicates carbachol; IsoP, isoprenaline; NO, nitric oxide; SM‐Rac1, smooth muscle Rac1 knockout; SNAP, S‐Nitroso‐N‐Acetyl‐DL‐Penicillamine.

To test this hypothesis, the endothelial‐mediated relaxation was assessed by testing the vasodilatory effect of carbachol (CCh) in arterial rings pre‐contracted with Phe. This relaxation response was impaired in both aorta and mesenteric arteries from SM‐Rac1‐KO mice (Figures 2C and 2D, S5B). CCh promotes vasodilation by inducing NO release and endothelium‐derived hyperpolarization factors (EDHF) production. To determine the relaxing pathways affected by Rac1 deletion, we first inhibited NO signaling by the NO synthase (NOS) inhibitor, L‐NAME. L‐NAME reduced the relaxing effect of CCh in mesenteric arteries from control but not in those from SM‐Rac1‐KO mice. In the presence of L‐NAME, the relaxing effect of CCh was the same in WT and SM‐Rac1‐KO mice arteries (Figure 2D). Similarly, in vivo treatment with L‐NAME increased systolic blood pressure in control mice but had no significant effect in SM‐Rac1‐KO mice (Figure 2E). Consequently, after L‐NAME treatment, systolic arterial blood pressure was similar in both groups of mice (Figure 2E), supporting that NO signaling deficiency contributes to the vascular defects and the increased systolic pressure observed in SM‐Rac1‐KO mice. These ex vivo and in vivo data thus suggest that EDHF signaling pathway was not altered in SM‐Rac1‐KO mice and that the reduced CCh‐induced relaxation in arteries from SM‐Rac1‐KO‐mice likely resulted from a defective vSMC response to endothelial NO.

This hypothesis was confirmed by the use of the NO donor SNAP. The relaxing effect of increasing concentrations of SNAP was strongly decreased in arterial rings from SM‐Rac1‐KO mice compared to that obtained in control rings (Figures 2D, S5B, and S6B). In contrast, the relaxation induced by the β‐adrenergic receptor agonist isoprenaline (IsoP), that is mediated by increased production of cAMP in vSMC, was similar in arteries from control and SM‐Rac1‐KO mice (Figures 2D and S5B). All these results indicate that Rac1 deletion causes a specific primary defect in signaling pathways responsible for relaxing response to endothelial NO in vSMC. This conclusion is further supported by the observation that the NO production, estimated by measuring the plasma nitrite concentration, was similar in control and SM‐Rac1‐KO mice at 5 and 15 days after induction of Rac1 deletion in SMC, while systolic blood pressure was already increased (Figure 2F).

Rac1 Deletion Leads to Defective Production of cGMP in vSMC

We next wanted to understand the mechanism linking Rac1 deletion in SMC to a defective response to NO. Western‐blot analysis revealed that the expression of the main molecular components of the NO pathway, endothelial NOS (eNOS), soluble guanylyl cyclases (sGC) α1 and β1, type 5 phosphodiesterase (PDE5), and cGMP‐dependent protein kinase (PKG), was not affected by Rac1 deletion in vSMC (Figure 3A). However, 15 days after the end of tamoxifen treatment, we observed a 50% reduction of the cyclic Guanosine MonoPhosphate (cGMP) content in SM‐Rac1‐KO mouse arteries compared to those of control animals (Figure 3B). The concentration of cAMP, another second messenger leading to vasodilation, was similar in both groups of mice (Figure 3B). These results suggest that the defective response to NO in SM‐Rac1‐KO mice arteries resulted from a reduction of NO‐induced rise in cGMP in vSMC. This hypothesis was confirmed by the use of the cell‐permeable cGMP analog (8‐Br‐cGMP) that induced a similar vasodilation in mesenteric arteries rings from control and SM‐Rac1‐KO mice (Figure 3C).

Figure 3.

Rac1 is essential for NO‐dependent cGMP production in vSMC. A, Immunoblot analysis of eNOS, sGCα1, sGCβ1, PDE5, and PKG protein expression in Rac1^lox/lox and SM‐Rac1‐KO aortas (n=3). B, Level of cGMP and cAMP contents in aorta from the indicated mouse strains 15 days after tamoxifen treatment (n=6 to 7). C, Percentage of vessel relaxation induced by the indicated doses of the cGMP analogue 8‐Br cGMP on Phe‐constricted mesenteric arteries from Rac1^lox/lox and SM‐Rac1‐KO mice (n=5). Log M, logarithm of the molar concentration used. D, Left panel, representative images of the F‐actin cytoskeleton in Rac1^lox/lox and SM‐Rac1‐KO vSMC after treatment with the indicated drugs (scale bar 20 μm). Right panel, corresponding quantification of F‐actin/G‐actin ratio (n=3). E, Pull‐down analysis of Rac1 activation by SNP in vSMC. F, Production of cGMP after SNP treatment in vSMC transfected with the indicated Rac1 mutants (n=3). G, Detection by western blot of PDE5 in anti‐Pak1 immunoprecipitates obtained from Rac1^lox/lox and SM‐Rac1‐KO aortas treated or not with SNP (n=6). ^#P<0.01 compared with Rac1^lox/lox control; *P<0.05 compared with the same condition without drug; ^§P<0.05 compared with Rac^WT with SNP. cGMP indicates cyclic Guanosine MonoPhosphate; eNOS, endothelial nitric oxide synthase; IP, immunoprecipitation; Ig, Immunoglobulin; IsoP, isoprenaline; NO, nitric oxide; PAK1, p21‐activated kinase 1; PDE5, type 5 phosphodiesterase; PKG, protein kinase G; sGC, soluble guanylyl cyclases; SM‐Rac1, smooth muscle Rac1 knockout; TCL, total cell lysate; vSMC, vascular smooth muscle cells.

To directly address the role of Rac1 downstream of NO in vSMC, we assessed the ability of NO to promote cGMP‐dependent disassembly of F‐actin in vSMC.³³ In control condition, F‐actin level was similar in vSMC from Rac1^lox/lox and SM‐Rac1‐KO mice (Figure 3D). As expected, the NO donor sodium nitroprussiate (SNP) induced stress fiber disassembly in vSMC from Rac1^lox/lox mice, but had no effect on vSMC from SM‐Rac1‐KO mice (Figure 3D). In contrast, the cAMP‐elevating agent IsoP or 8‐Br‐cGMP induced a same stress fiber disassembly in both control and Rac1‐KO vSMC (Figure 3D), thus confirming a role of Rac1 in NO signaling upstream of cGMP production in vSMC. In fact, we observed that SNP induced Rac1 activation in vSMC (Figure 3E) and that expression of a dominant negative Rac1 mutant (Rac1^N17) significantly inhibited SNP‐induced increase in cGMP in vSMC (Figure 3F), indicating that Rac1 activation is necessary for NO‐induced rise in cGMP. The effector of Cdc42 and Rac1 proteins Pak1 has been shown to interact in vitro with PDE5 thereby limiting its cGMP‐hydrolyzing activity.³⁴ To assess the potential role of PDE5 in the reduced cGMP response to NO donors in Rac1‐deficient vSMC, we analyzed the Pak1/PDE5 interaction in control and SM‐Rac1‐KO mice arteries. Activation of NO signaling by SNP induced the formation of the Pak1/PDE5 complex in aortas from control mice but not in those from SM‐Rac1‐KO mice (Figure 3G). These results suggest that Rac1 deletion in vSMC prevents Pak1‐mediated PDE5 inhibition, thus leading to a reduced cGMP level and vasodilation in response to NO.

Rac1 Deletion Induced RhoA/Rho Kinase Signaling Overactivation

Inhibition of RhoA/Rho kinase signaling is one of the main mechanisms by which NO mediates vasodilation through cGMP/PKG‐dependent RhoA phosphorylation on serine 188 (P^Ser188‐RhoA).³³ To determine the role of RhoA/Rho kinase activation in the hypertensive SM‐Rac1‐KO phenotype, mice were treated with the Rho kinase inhibitor fasudil (at 5 or 30 mg/kg of body weight). Twenty minutes after fasudil injection, systolic arterial pressure was decreased to the same level in both groups of mice, suggesting that the rise in blood pressure in SM‐Rac1‐KO mice was mediated by an increase in RhoA/Rho kinase activity (Figure 4A). Immunoblot analysis in aorta revealed that the depletion of Rac1 in SM‐Rac1‐KO aorta was associated with a loss of phosphorylated (active) Pak1 (P‐Pak1) but had no effect on RhoA or MYPT expression (Figure 4B). However, the amount of P^Ser188‐RhoA was decreased and the level of P‐MYPT was increased in SM‐Rac1‐KO mice aorta compared with control (Figure 4B). These results indicate an upregulation of RhoA/Rho kinase activity in arteries of SM‐Rac1‐KO mice suggesting that NO‐mediated Rac1 activation tonically antagonizes RhoA/Rho kinase activity.

Figure 4.

Over activation of RhoA/Rho kinase signaling in Rac1 deficient arteries. A, Tail‐cuff measurements of systolic arterial blood pressure 20 minutes after i.p. vehicle or fasudil treatment in vivo (5 or 30 mg/kg of body weight). B, Top panel, immunoblot analysis of Rac1/Pak1 and RhoA/Rho kinase pathways in aortas from Rac1^lox/lox and SM‐Rac1‐KO mice (n=3 to 4). Bottom panel, corresponding quantification of Ser188 RhoA and MYPT phosphorylation. *P<0.05. PAK1 indicates p21‐activated kinase 1; Rho, Ras homologous; SM‐Rac1, smooth muscle Rac1 knockout.

Rac1 Deletion Disrupts p116^RIP3‐RhoA Association

In order to identify Rac1 interacting partners or effectors other than Pak1 involved in NO‐induced RhoA/Rho kinase inactivation, we performed Rac1‐GST pull‐down assay in control and SNP‐treated vSMC. After two‐dimensional gel electrophoresis and Coomassie blue staining, spots modified by SNP treatment were identified by mass spectrometry. We thus caught the Myosin Phosphatase Rho‐Interacting Protein (p116^RIP3), whose interaction with Rac1 was decreased by SNP (Table S1). Western blot analysis of p116^RIP3 in Rac1 immunoprecipitate confirmed this result (Figure 5A). Interaction of p116^RIP3 with Rac1^WT was similar to that observed with the dominant active Rac^V12 and the dominant negative Rac^N17 mutants (Figure 5B), indicating that p116^RIP3‐Rac1 association did not depend on SNP‐induced Rac1 activation.

Figure 5.

NO signaling modulates association between RhoA and its inhibitor p116^RIP3. A, Left panel, p116^RIP3‐Rac1 and p116^RIP3‐RhoA complexes were analyzed by Western blotting of each GTPase immunoprecipitates (IP GFP) from GFP‐Rac1WT and GFP‐RhoAWT transfected vSMC treated or not with SNP (n=3 to 4). Right panel, quantification of p116^RIP3 interaction with Rac1 and RhoA. B, Co‐immunoprecipitation of endogenous p116^RIP3 with the indicated Rac1 mutants transfected in vSMC (n=4). C, Co‐immunoprecipitation of endogenous p116^RIP3 with PKG in murine aortas stimulated with SNP. D, Time‐course of SNP‐induced RhoA phosphorylation on Ser188 and p116^RIP3‐RhoA complex formation assessed in immunoprecipitated RhoA from vSMC. E, Co‐immunoprecipitation of endogenous p116^RIP3 with the phosphomimetic (GFP‐RhoA^S188E), phosphoresistant (GFP‐RhoA^S188A) or wild type (GFP‐RhoA^WT) RhoA in transfected vSMC in the absence or presence of SNP (n=5). *P<0.05. GFP indicates Green Fluorescent Protein; Ig, Immunoglobulin; IP, immunoprecipitation; PKG, protein kinase G; Rho, Ras homologous; TCL, total cell lysate; vSMC, vascular smooth muscle cells; WT, wild type.

Interestingly, p116^RIP3 was identified as a RhoA inhibitor that directly interacts with RhoA in a nucleotide‐independent manner and also with PKG in human vSMC.³⁵ As a mirror image of its dissociation from Rac1, SNP promoted p116^RIP3‐RhoA interaction (Figure 5A). In contrast, although the p116^RIP3‐PKG complex was not modified by SNP stimulation (Figure 5C), the kinetic of SNP‐induced RhoA‐p116^RIP3 complex formation paralleled SNP‐induced RhoA phosphorylation (Figure 5D). Expression of wild‐type (RhoA^WT), phospho‐mimetic (RhoA^S188E) or phospho‐resistant (RhoA^S188A) RhoA mutant in vSMC showed that under resting condition, p116^RIP3 was more strongly bound to RhoA^S188E than to RhoA^WT (Figure 5E). Furthermore, SNP increased the interaction of p116^RIP3 with RhoA^WT but not with the phospho‐resistant RhoA^S188A mutant (Figure 5E). These results indicate that phosphorylation of RhoA on Ser188 favors its interaction with its inhibitor p116^RIP3.

We then wanted to assess the role of p116^RIP3 in NO‐induced RhoA inactivation by siRNA‐mediated p116^RIP3 silencing in vSMC (Figure 6A). The down‐regulation of p116^RIP3 expression did not affect NO‐induced RhoA phosphorylation but strongly reduced NO‐induced actin stress fiber disruption (Figures 6B and 6C) suggesting that the interaction of phosphorylated RhoA with p116^RIP3 is required for the inhibitory action of NO on RhoA/Rho kinase signaling. The next question was therefore to know whether p116^RIP3 was involved in the alteration of NO signaling and the up‐regulation of RhoA/Rho kinase pathway induced by Rac1 deletion in SM‐Rac1‐KO mice arteries. Co‐immunoprecipitation experiments demonstrated that p116^RIP3 was associated with RhoA in aorta of control mice but not in those from SM‐Rac1‐KO mice (Figure 6D). In agreement with the positive effect of NO donors observed in vitro, the inhibition of NO production by L‐NAME decreased p116^RIP3‐RhoA interaction in control mice, but had no effect in SM‐Rac1‐KO mice (Figure 6D). Thus, these results suggest that Rac1 is required for NO‐induced RhoA‐p116^RIP3 interaction and are consistent with the involvement of the disruption of the RhoA‐p116^RIP3 complex in the activation of RhoA/Rho kinase signaling in SM‐Rac1‐KO mice arteries.

Figure 6.

Down‐regulation of p116^RIP3 expression in vSMC prevents NO‐dependent RhoA inactivation. A, Inhibition of p116^RIP3 protein expression by RNA interference (siRNA) assessed by immunoblot in vSMC (n=3). B, Immunoblot analysis of NO‐dependent RhoA phosphorylation (P^ser188) in p116^RIP3 siRNA transfected vSMC (n=3). C, Effect of p116^RIP3 siRNA on F‐actin cytoskeleton organization with or without SNP treatment in vSMC (n=4). D, Endogenous p116^RIP3 in RhoA immunoprecipitates obtained from aortas of Rac1^lox/lox and SM‐Rac1‐KO mice treated or not with L‐NAME (3 g/L in drinking water) (n=8). *P<0.05. IP indicates immunoprecipitation; NO, nitric oxide; Rho, Ras homologous; SM‐Rac1, smooth muscle Rac1 knockout; TCL, total cell lysate; vSMC, vascular smooth muscle cells.

Discussion

Our study identifies a new role of Rac1 as a regulator of blood pressure and a key component of the signaling pathway that mediates the blood pressure‐lowering effect of NO in vSMC. We demonstrate that ablation of the Rac1 gene in vSMC is sufficient to induce high blood pressure in mice, supporting a physiological role of vSMC Rac1 in blood pressure homeostasis. Moreover, we found that Rac1 is a molecular link connecting NO to cGMP and RhoA signaling inhibition, and discovered the key role of p116^RIP3 in this pathway essential for NO‐mediated vasodilation (Figure 7).

Figure 7.

Schematics representations of the new regulatory mechanism reported in this work. MLC indicates myosin light chain; PAK1, p21‐activated kinase 1; PDE5, type 5 phosphodiesterase; Rho, Ras homologous; sGC, soluble guanylyl cyclases; SM‐Rac1, smooth muscle Rac1 knockout; WT, wild type.

SM‐Rac1‐KO mice display a ≈15 mm Hg increase in blood pressure associated with a specific loss of NO‐mediated increase in cGMP and vasodilation. Moreover, mutant mice are resistant to L‐NAME‐induced rise in blood pressure. The role of Rac1 in the regulation of NO/cGMP signaling and the causative effect of Rac1 deletion‐induced defective NO signaling in the pathogenesis of hypertension in SM‐Rac1‐KO mice are supported by previous studies showing that eNOS or PKG gene disruption in mice similarly led to the abrogation of NO/cGMP‐dependent arterial relaxation and an increase in arterial pressure of about 10 to 15 mm Hg.^36–39 In addition to an increase in arterial pressure, this alteration of NO signaling could also account for the rise of arterial stiffness observed in SM‐Rac1‐KO mice as a chronic reduction of NO production induced vascular stiffening.^40–41 This close relationship between high systolic blood pressure, endothelial dysfunction, and vascular elastance was also found in hypertensive patients⁴² and aging population.^43–45 All these data are thus consistent with an essential role of Rac1 to mediate NO effect in vSMC and consequently, NO‐dependent regulation of vascular tone and blood pressure. In agreement with our finding, it was recently found that the Ala370Ser polymorphism in the ARHGAP9 gene coding for a RacGAP inactivating Rac1, is associated with coronary artery spasm in human ascribed to a decreased GAP activity towards Rac1.⁴⁶

In mice, overexpression of a dominant‐negative or a constitutively active mutant of Rac1 gene in SMC has shown that Rac1 is an essential regulator of NADPH oxidase/ROS pathway in blood vessels.^47–48 While blood pressure has not been measured in transgenic mice overexpressing the dominant‐negative form of Rac1, animals that overexpressed the active Rac1 mutant in vSMC developed moderate hypertension due to an excessive amounts of O₂^•−,⁴⁸ Therefore, it is surprising that overexpression of active Rac1 mutant and deletion of endogenous Rac1 in SMC lead to the same blood pressure phenotype. A possible explanation is that the massive artificial ROS production due to Rac1 overexpression completely masks the physiological role of endogenous Rac1 as a component of the NO/cGMP signaling in vSMC.

In SM‐Rac1‐KO mice, the inability of vSMC to normally respond to NO resulted from a defective production of cGMP.^34,49–50 Although Rac1 has been shown to positively control sGC expression in adipocytes,⁵⁰ we did not observe any modification of sGCα1 and sGCβ1 expression in SM‐Rac1‐KO aorta suggesting that the low level of cGMP in SM‐Rac1‐KO mice arteries was not due to a defect in cGMP generation. However, we show that NO induces activation of Rac1 and formation of Pak1/PDE5 complex in vSMC. The loss of Pak1/PDE5 interaction in response to NO donor observed in SM‐Rac1‐KO mice arteries is likely responsible for the decrease in cGMP level by preventing NO‐induced inhibition of cGMP hydrolysis.

Arterial blood pressure depends in part on the regulation of blood vessel lumen diameter, determined by the contractile activity of vSMC. The RhoA/Rho kinase signaling pathway plays a major role in the Ca²⁺‐independent contraction of vSMC and the regulation of vascular tone and blood pressure by vasoconstrictors.^18,51–53 Conversely, inhibition of RhoA/Rho kinase signaling by NO, through PKG‐mediated phosphorylation of RhoA on Ser188, plays a major role in the relaxing effect of NO in vSMC.^54–56 In agreement with the involvement of this pathway in vivo, the depression of NO signaling in SM‐Rac1‐KO mice arteries is associated with an increase in RhoA/Rho kinase activity attested by a rise of MYPT phosphorylation and a decreased phosphorylation of RhoA on Ser188. Deletion of Rac1 in SMC thus mimics the activation of RhoA/Rho kinase signaling observed in hypertensive eNOS‐deficient mice.^57–58 This higher activity of RhoA/Rho kinase in SM‐Rac1‐KO mice arteries is consistent with the potentiation of contractile responses of resistance arteries to the thromboxane A2 analog U47619 and Phe, whereas Ca²⁺‐dependent K⁺‐contractions are unchanged.

In an attempt to decipher the molecular pathway linking Rac1 to RhoA regulation, we identified the protein p116^RIP3. p116^RIP3 was initially described as a component of MLCP complex that directly binds RhoA.³⁵ Overexpression of p116^RIP3 disrupts the actin cytoskeleton and inhibits contractility by activating MLCP and inactivating RhoA.^59–63 The interaction between RhoA and p116^RIP3 occurs independently of the nucleotide binding state of RhoA and the regulation of this complex formation as well as its physiological role are totally unknown.³⁵ Here we demonstrate that stimulation of NO pathway promotes Rac1‐p116^RIP3 dissociation and RhoA‐p116^RIP3 complex formation that is favors by RhoA phosphorylation. In addition, we demonstrated that despite its association to PKG, p116^RIP3 is not involved in NO‐induced RhoA phosphorylation but is essential to mediate the inhibitory action of NO on RhoA/Rho kinase activity. These data thus identified p116^RIP3 as a new and essential intermediate in the signaling pathway responsible for cGMP/PKG‐mediated inhibition of RhoA cellular functions. By preventing NO‐induced p116^RIP3‐RhoA interaction, Rac1 deletion disrupts NO‐mediated inhibition of RhoA/Rho kinase effects in vSMC, leading to RhoA/Rho kinase‐dependent increase in arterial tone and blood pressure. Vascular SMC Rac1 thus constitutively acts as a brake that limits RhoA/Rho kinase activity in response to NO.

Finally, our present data highlight a functional significance of the reciprocal antagonistic relationship between Rac1 and RhoA at a physiological level, ie, blood pressure regulation. The blood pressure‐lowering effect of vSMC Rac1 opposes the hypertensive effect of vSMC RhoA. Rac1 is required for NO/cGMP/PKG/p116 ^RIP3‐mediated inactivation of RhoA that in turn activates Rac1.⁹ This positive feedback creates a virtuous circle that contributes to the balance between vasodilation and vasoconstriction and efficiently regulates blood pressure.

Supplementary Material

Appendix S1 Supplemental Methods.

Acknowledgments

We thank N. Vaillant, V. Aillerie, M. Rio, and S. Heurtebise‐Chretien for excellent technical assistance. We also value the support provided by the animal facility units of the University of Nantes. We thank Cardiex platform for the cardiovascular expertise and the INRA Research Unit 1268 for the mass spectrometry analyses.