Skip Navigation

Cardiovascular Research 2006 72(2):339-348; doi:10.1016/j.cardiores.2006.07.017
This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Gouëffic, Y.
Right arrow Articles by Loirand, G.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Gouëffic, Y.
Right arrow Articles by Loirand, G.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

Copyright © 2006, European Society of Cardiology

Hyaluronan induces vascular smooth muscle cell migration through RHAMM-mediated PI3K-dependent Rac activation

Yann Gouëffica,b,*, Christophe Guilluya, Patrice Guérina,b, Philippe Patrab, Pierre Pacauda and Gervaise Loiranda,b,*

aInserm, Université de Nantes, U533, l'institut du thorax, Nantes, F-44000, France
bCentre Hospitalier Universitaire de Nantes, Nantes, F-44000, France

* Corresponding authors. Inserm U-533, Université de Nantes, L'institut du Thorax, Faculté des Sciences, 2 rue de la Houssinière, BP 92208, Nantes F-44000, France. Email address: yann.goueffic{at}chu-nantes.fr gervaise.loirand{at}univ-nantes.fr

Received 6 April 2006; revised 18 July 2006; accepted 24 July 2006


   Abstract
Top
 Abstract
1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
 References
 
Objective: Hyaluronan (HA) is an important constituent of the extracellular matrix and is known to regulate cellular events through binding to CD44 and the receptor for HA-mediated motility (RHAMM). Here we investigated the role of these receptors and the signaling pathways involved in HA-mediated effects in arterial smooth muscle cells (ASMC).

Methods: Effects of high-molecular weight HA (1 to 5 mg/ml) were analyzed in cultured ASMC from rat aorta.

Results: HA promoted actin stress fiber and lamellipodia formation and dose-dependently induced ASMC migration without effect on proliferation. Pull-down assay of Rho protein activity indicated that HA activated RhoA and Rac. HA-induced ASMC migration was not affected by the RhoA inhibitor Tat-C3 (10 µg/ml), the Rho kinase inhibitor Y-27632 (10 µM) and blocking anti-CD44 antibody, but was reduced by the non-selective Rho protein inhibitor simvastatin (10 µM), the Rac inhibitor LT-toxin (1 µg/ml), small interfering RNA (siRNA) targeting Rac and the phosphatidyl inositol 3-kinase (PI3K) inhibitor LY294002 (25 µM), which also blocked HA-induced Rac activation. CD44 knockdown by siRNA inhibited HA-mediated RhoA activation without effect on ASMC migration. In contrast, siRNA targeting RHAMM inhibited both HA-induced migration and Rac activation.

Conclusions: High-molecular weight HA independently activates RhoA and Rac through CD44 and RHAMM, respectively. HA-induced migration depends exclusively on RHAMM-mediated PI3K-dependent Rac activation.

KEYWORDS Smooth muscle; Extracellular matrix; Signal transduction; Receptors; G-proteins


   1. Introduction
Top
 Abstract
 1. Introduction
2. Materials and methods
 3. Results
 4. Discussion
 References
 
Vascular proliferative disorders such as atherosclerosis, restenosis or allograft rejection are characterized by arterial smooth muscle cells (ASMC) proliferation and migration associated with marked changes into extracellular matrix (ECM) [1]. Hyaluronan (HA) is a high molecular weight polysaccharide (1000–10,000 kDa) widely distributed in ECM of a variety of tissues including the arterial wall. Studies have clearly shown that HA not only accumulates at different stages in atherosclerosis and restenosis, promotes atherosclerosis, but also regulates ASMC functions including proliferation, migration and secretion [2–4]. HA could also be located inside the cells and some HA effects have been ascribed to intracellular HA [5,6].

Numerous studies that analyzed in vitro the biological effects of HA in various cell types have yielded to mixed results. In addition to the different cell types used, several influences directly related to the complexities of HA binding, HA receptors and HA size could underlie the observed variability [7]. The biological functions of HA are essentially mediated by cell surface HA receptors or hyaladherins. Among hyaladherins, CD44 and the receptor for HA-mediated motility (RHAMM) have been identified in ASMC [8]. CD44 comprises a large family of transmembrane glycoproteins that exhibit extensive molecular heterogeneity. The CD44 ectodomain is responsible of the binding of HA, and the cytoplasmic domain regulates specific signaling and interacts with cytoskeletal proteins such as ankyrin or ERM (ezrin, radixin, and moesin) [8,9]. RHAMM is an 85-kDa protein expressed in most tissues, distributed in multiple compartments including cell surface, cytoskeleton, mitochondria and cell nucleus [8]. RHAMM binds HA and also acts as a microtubule-associated protein interacting with the actin [8,10]. The relative contribution of the two types of HA receptors and the intracellular signaling pathways involved in HA-mediated effects in ASMC remain unknown.

The small G proteins of the Rho family are identified as tightly regulated molecular switches that cycle between an inactive GDP-bound and an active GTP-bound conformation that interacts with downstream targets (effectors) to elicit cellular responses [11]. The best characterized members of the Rho protein family, RhoA, Rac and Cdc42, are recognized as major regulators of the actin cytoskeleton [11]. A large body of evidence has now been obtained regarding the important functions of Rho proteins in the vasculature [12,13]. Rho protein signaling has been demonstrated to be a critical mechanism for controlling vascular smooth muscle differentiation [14,15], proliferation [16,17], and migration [18,19]. In addition, mainly through their effect on actin cytoskeleton organization, Rho proteins regulate essential cell functions such as adhesion, cytokinesis and transcription. In mammary epithelial cells and in breast tumor cells, it has been shown that HA binding to CD44 promotes activation of Rac and that the association of CD44V3 isoform to RhoA is involved in metastatic tumor cell migration [20–22].

In this study we investigated the role of the HA receptors CD44 and RHAMM, and the involvement of Rho protein signaling in the effects mediated by high-molecular weight HA in ASMC. We show that HA stimulates the actin cytoskeleton organization and dose-dependently induces ASMC migration. Our results demonstrate that HA activates two independent signaling pathways in ASMC and that although HA binding to CD44 induces RhoA activation, HA-induced migration exclusively depends on phosphatidyl inositol 3-kinase (PI3K)- and Rac-dependent signaling pathway downstream to RHAMM activation.


   2. Materials and methods
Top
 Abstract
 1. Introduction
 2. Materials and methods
3. Results
 4. Discussion
 References
 
2.1. Cell culture
Rat ASMC were isolated from aorta by enzymatic dissociation and cultured as previously described [23]. Only smooth muscle cells at passage 3 were used in this study. The investigation conforms with the Guide for the Care and Use of Laboratory Animals published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996).

2.2. Vascular smooth muscle cells migration assays
For in vitro wound healing assay, a sterile pipette tip was used to make a 0.5-mm-wide wound by streaking across a monolayer of ASMC serum-starved for 24 h. Multiple photographs of the wound were taken at various time post-wounding in serum-free medium (control) or in the presence of various HA concentrations. The areas of cell recoverage were determined with image analysis software (Metamorph, Universal Imaging Corp, West Chester, PA). The percentage of cellular recoverage area to the whole wound area was measured to evaluate migration. The effect of various agents such as Tat-C3, Y-27632, LY 294002 and blocking CD44 antibody were analyzed by addition to culture medium 24 h before adding HA. For Boyden chamber assay [24], ASMC were trypsinized and washed in serum-free DMEM before plating into a 12-well Transwell plate (12 mm diameter, 8 µm pore size, Corning Incorporated Costar) at 5x104 cells per/well. DMEM containing various concentrations of HA (1–5 mg/ml) was placed in the lower chamber and ASMC were allowed to migrate for 8 h at 37 °C. After incubation, the filter was removed, and ASMC on the upper side of the filter were scraped off. ASMC that had migrated to the lower side of the filter were fixed in methanol at 4 °C for 15 min, stained with Toluidin blue and counted under a microscope for quantification of ASMC migration. Migration was expressed as the number of cells that had migrated per high-power field (x400).

2.3. Actin staining
Serum-starved (24 h) ASMC, stimulated or not with HA, were fixed for 30 min in 4% paraformaldehyde, permeabilized for 10 min in 0.5% Triton X-100, and then rinsed in phosphate-buffered saline (PBS). Polymerized F-actin was stained with FITC-conjugated phalloidin (5 µg/ml, 45 min at room temperature). After washing with PBS, coverslips were mounted on a glass slide and examined with a fluorescence microscope (Eclipse E-600, Nikon, Champigny-sur-Marne, France). Images were collected with a cool-SNAP camera (Princeton Instruments, Evry, France) and stored for analysis (Metamorph, Universal Imaging Corp).

2.4. Western blot analysis
Serum-starved (24 h) ASMC, stimulated or not with HA, were harvested and homogenized in lysis buffer containing 50 mM Tris, pH 7.2, 500 mM NaCl, 10 mM MgCl2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, protease inhibitor cocktail (Sigma), phosphatase inhibitor cocktail (Sigma). Nuclei and unlysed cells were removed by low speed centrifugation at 10,000xg for 15 min at 4 °C. Proteins were resolved on SDS-PAGE and transferred to nitrocellulose membranes, which were incubated with specific antibodies for detection of RhoA, Rac, CD44, RHAMM, β-actin, phosphorylated P21-activated kinase 1 (PAK1) (used to monitor the activation of the Rac/PAKs signalling) and PAK1. Immunoreactive bands were visualized using horseradish peroxidase conjugated secondary antibody and subsequent ECL detection (Amersham Pharmacia Biotech, Orsay, France), then quantified by densitometric analysis using QuantityOne (Biorad, Hercules, CA, USA).

2.5. Measurement of RhoA and Rac activities
Activity of RhoA and Rac was analyzed by a pull-down assay using the Rho binding domain of the Rho protein effectors Rhotekin and PAK, respectively [25]. cDNA encoding for the rhotekin and PAK RBD was kindly provided by Dr. Martin Schwartz (University of Virginia, Charlottesville, Virginia). ASMC were serum-starved for 24 h, stimulated or not with HA, then washed with ice-cold PBS and lysed in a lysis buffer (50 mM Tris, pH 7.2, 500 mM NaCl, 10 mM MgCl2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, protease inhibitor cocktail (Sigma)). After centrifugation 12,000xg at 4 °C for 10 min, the extracts were incubated at 4 °C for 45 min with glutathione-Sepharose 4B beads coupled with glutathione-S-transferase (GST)-rhotekin RBD fusion protein (for Rho activity) or GST-PAK RBD fusion protein (for Rac activity). Precipitated GTP-bound and total RhoA and Rac were then analyzed by Western blotting using a mouse monoclonal anti-RhoA or anti-Rac antibody. Immunoreactive bands were detected using ECL detection and quantified by densitometric analysis.

2.6. siRNA
The siRNA sequence targeting rat CD44 (mRNA sequence accession number NM_012924 [GenBank] ) and RHAMM (mRNA sequence accession number NM_012964 [GenBank] ) were selected using the software developed by Eurogentec (Seraing, Belgium) and verified in a BLAST search. The sense strand sequences of siRNAs (Eurogentec) used were: CD44, 5'-GAAGGGCGAGUAUAGAACAdTdT-3'; and RHAMM, 5'-AGAGAAGGAUCGUGAGAUUdTdT-3'. To inhibit Rac1 synthesis, we used the following sense strand sequences of siRNA: 5'-GUUCUUAAUUUGCUUUUCCdTdT-3' [26]. A "universal" scrambled siRNA has been used for control (sense: 5'-CGACUUCCUUACUCUAUACdTdT-3').

The siRNAs were introduced into ASMC with JetPEI reagent (Polytransfection, Illkirch, France) according to the manufacturer's recommendations. Cells were incubated with or without 1.2 µg of specific or scrambled siRNA for 48 h before functional assays were conducted. The efficiency of siRNA-mediated gene silencing has been assessed by real-time quantitative PCR and Western blot.

2.7. Real-time quantitative PCR
Total RNA was extracted from control (scrambled siRNA), CD44 siRNA- and RHAM siRNA-treated ASMC using TRIzol reagent then treated with RNase-free DNase. Total RNA (1 µg) was used as template for the reverse transcriptase (RT) reaction using MMLV, 2 µg random hexamers, 0.5 mM dNTPs and 5 mM DTT (Life Technologies). Real-time quantitative PCR was performed in the iCycler iQTM Detection System (Bio-Rad Laboratories) using Sybr green detection (Molecular Probes) and TitaniumTM Taq DNA polymerase (Clontech), according to the manufacturer's recommendations. The primer pairs used were: CD44 (sense: 5'-TGGATGCGAGGAGGATATACAC; anti-sense: 5'-CTGCGAGGCTTTCAACACC); RHAMM (sense: 5'-TGCCCTGGATGAGCTGGA; anti-sense: 5'-GCATGTGCAGCGGTTCTTT); GAPDH (sense: 5'-TCCATTTACAACCACAACGATTC; anti-sense: 5'-CTGAGAGGCAAGGATGAATGATT). The following temperature profile was used: 40 cycles of 15 s at 95 °C and 1 min at 60 °C. Cycle numbers obtained at the log-linear phase of the reaction were plotted against a standard curve prepared with serially diluted control cDNA samples. Expression of target genes was normalized to GAPDH levels. The delta Ct (Cycle threshold) method was used to calculate relative expression levels.

2.8. Chemical and drugs
Mouse monoclonal RhoA and Rac antibodies, blocking CD44 antibody (sc-7946) [27] and goat polyclonal RHAMM antibody were purchased from Santa Cruz Biotechnology (Santa Cruz, CA). Rabbit polyclonal anti-phospho-PAK1 and anti-PAK1 antibodies were from Cell Signaling Technology (Ozyme, Saint Quentin, France). Rat monoclonal CD44 antibody was from Chemicon (Hampshire, UK). LY 294002 was purchased from Calbiochem (France Biochem, Meudon, France). The permeant RhoA inhibitor Tat-C3 protein was produced in E. coli and purified as already described [17,28]. HA from rooster comb was used in this study (Sigma, Saint-Quentin Fallavier, France). Its average molecular weight was 2100 kDa [29]. All other reagents were purchased from Sigma (Saint-Quentin Fallavier, France).

2.9. Statistics
All results are expressed as the means±S.E.M. of sample size n. Significance was tested by Student's t-test. Data were considered statically significant when P<0.05.


   3. Results
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
4. Discussion
 References
 
3.1. High-molecular weight HA dose-dependently induces ASMC migration but not proliferation
The in vitro model of ASMC monolayer wounding and restitution was first used to assess the effect of high-molecular weight HA on ASMC migration (Fig. 1). After scrape wounding, ASMC migrated across the wound edge to fill in the area that was denuded of cell by the scrape (Fig. 1A). Exposure to HA (1–5 mg/ml) resulted in a dose- and time-dependent stimulation of ASMC migration, leading to acceleration of the wound closure (Fig. 1A–C). The HA concentration required to induce 50% of maximal effect (EC50) is closed to 1.8 mg/ml whatever was the time post-wounding (Fig. 1C). In similar experiments, the rate of cell proliferation, determined by cell counting or BrDU incorporation, in migrating and stationary ASMC over the initial 24 h period, was not modified in the presence of HA (data not shown), indicating that the accelerated wound closure induced by HA is proliferation-independent. Similar results have been obtained in Boyden chamber assays.


Figure 1
View larger version (82K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 1 HA stimulates ASMC motility. (A) Typical scratch-wound repair at 0, 16, and 24 h post-wounding in control ASMC and in ASMC stimulated by 1 and 5 mg/ml HA. (B) Time course of the wound closure in control ASMC (circle) and in the presence of 1 mg/ml (square), 3 mg/ml (down triangle) and 5 mg/ml (up triangle). (C) Concentration–response curve of HA-induced wound closure at 24 h post-wounding (n=4–6 independent experiments).

 
3.2. HA stimulates the actin cytoskeleton organization
Since actin cytoskeleton reorganization is required for cell shape modification and cell migration, we examined the effect of HA on actin cytoskeleton organization in ASMC. Phalloidin staining revealed that HA induced reorganization of actin cytoskeleton at the wounded edge, promoting lamellipodia and actin stress fiber formation (Fig. 2). This effect was observed from the first hour to up to 24 h post wounding.


Figure 2
View larger version (82K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 2 HA stimulates actin cytoskeleton organization. FITC-phalloidin labelled F-actin of ASMC at the wound edge at 16 h and 24 h post-wounding in the absence (control) and in the presence of HA 3 mg/ml HA. HA stimulates actin cytoskeleton organization leading to actin stress fiber (asterisks) and lamellipodia formation (arrowheads).

 
3.3. HA induces sequential activation of the Rho proteins RhoA and Rac
RhoA and Rac protein activity controls stress fibers and lamellipodia formation, respectively [30]. We therefore next asked whether HA induced RhoA and/or Rac activation in ASMC. The activity of RhoA and Rac was assessed by the detection of the GTP-loaded forms of these proteins by pull down assays. As shown in Fig. 3A, HA transiently activated RhoA in ASMC stimulated with HA for 10 min. RhoA activity returned to basal level after 1 h of HA stimulation. HA also induced a sustained Rac activity, observed at 10 min and 1 h (Fig. 3B). Increased Rac activity was maintained up to 6 h (not shown). Our results thus provide evidence that HA-induced migration and actin cytoskeleton reorganization is associated with activation of RhoA and Rac.


Figure 3
View larger version (37K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 3 HA stimulates the activity of the small G-proteins RhoA and Rac. Western blot and corresponding densitometric analyses for RhoA activity (GTP-RhoA) and RhoA level (A), Rac activity (GTP-Rac) and Rac level (B) in ASMC stimulated for 10 min and 1 h with 1 mg/ml HA. RhoA and Rac activity is normalized to RhoA and Rac level, respectively, and expressed relative to the control in the absence of HA taken as 1. The data presented are representative of three independent experiments (#P<0.01 and *p<0.001 vs control).

 
3.4. Inhibition of Rac prevents HA-induced migration
To determine whether HA-mediated ASMC migration was dependent on Rho protein activity, we analyzed the effects of non-selective and selective Rho inhibitors on ASMC migration. In Boyden chamber assay, the non-selective Rho protein activity inhibitor simvastatin (10 µM) dramatically decreased the migration induced by HA (Fig. 4A). In contrast, selective inhibition of RhoA or its effector Rho kinase by Tat-C3 (10 µg/ml) and Y-27632 (10 µM), respectively, did not alter HA-induced ASMC migration (Fig. 4A). These concentrations of inhibitors efficiently inhibited RhoA/Rho kinase-dependent functions in ASMC [17]. To further investigate the signaling pathways involved in HA-induced ASMC migration, we next assessed the effect of inhibition of Rac by the Clostridium sordellii lethal toxin (LT-toxin) and by siRNA targeting Rac. When ASMC were treated with LT-toxin (1 µg/ml), HA-induced migration was almost totally inhibited (Fig. 4B). Similarly, Rac siRNA, that produced a 60% depletion of Rac protein (48 h, not shown), efficiently inhibited HA-induced migration (Fig. 4B). PI3K inhibition by LY-294002 (25 µM; Fig. 4B) or wortmanin (100 nM, not shown) also significantly decreased HA-mediated ASMC migration. Scratch-wound repair assays gave similar results. These results thus provide evidence that RhoA and Rho kinase activity are not involved in the effect of HA but that Rac and PI3K activation are required for HA-mediated migration.


Figure 4
View larger version (15K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 4 Inhibition of Rac but not RhoA prevents HA-induced migration. (A) In Boyden chamber assays, HA (3 mg/ml, 8 h) induced-ASMC migration was inhibited by the non-selective Rho protein inhibitor simvastatin (10 µM). RhoA and Rho kinase inhibition by Tat-C3 (10 µg/ml) and Y-27632 (10 µM) respectively, did not alter ASMC migration. (B) The Rac inhibitor LT-toxin (1 µg/ml), siRNA targeting Rac (48 h) and the PI3K inhibitor LY-294002 (25 µM) prevents HA-induced ASMC migration. Results are expressed as a percentage of control (*p<0.001 vs control).

 
3.5. RHAMM knockdown inhibits HA induced ASMC migration
Both CD44 and RHAMM are established signal-transducing receptors that influence several cell functions, including cell motility [7]. We therefore analyzed the ability of CD44 function-blocking antibody and siRNA targeting RHAMM and CD44, to prevent HA-induced ASMC migration. The efficiency and the selectivity of siRNA targeting RHAMM and CD44 have been assessed by real time RT-PCR. After 48 h, siRNA targeting CD44 decreased the expression of CD44 mRNA expression by 88.0±2.0% (n=3) without effect on RHAMM mRNA. Similarly, siRNA targeting RHAMM decreased the expression of RHAMM mRNA expression by 52±10% (n=3), without effect on CD44 mRNA level. At protein level, siRNA targeting CD44 and RHAMM induced a 60–70% decrease in CD44 and RHAMM expression, respectively (Fig. 5A).


Figure 5
View larger version (68K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 5 RHAMM knockdown inhibits HA-induced ASMC migration. (A) Western blot analysis of CD44 and RHAMM expression 48 h after transfection with siRNA-scrambled or siRNA targeting CD44 (left) or RHAMM (right). (B) Wound closure induced by 3 mg/ml HA at 0 h and 24 h post-wounding, in the absence (control) or presence of blocking anti-CD44 antibody, siRNA targeting CD44 and siRNA targeting RHAMM. (C) Quantitative effect of anti-CD44 antibody, siRNA-scrambled, siRNA-CD44 and siRNA-RHAMM on the wound closure at 24 h post-wounding. Results are expressed as a percentage of the wounded area at t=0. (D) Effect of blocking anti-CD44 antibody, siRNA-scrambled, siRNA-CD44 and siRNA-RHAMM analyzed in Boyden chamber migration assay in the presence of 3 mg/ml HA (8 h). The number of migrating cells is expressed as a percentage of control (n=4–6 independent experiments; *p<0.001 vs control).

 
In wound-healing assay, HA-induced ASMC motility was not modified by treatment within CD44 blocking antibody (10 µg/ml) or by siRNA targeting CD44 (Fig. 5B, C). Boyden chamber migration assays confirmed the absence of effect of CD44 blocking antibody and CD44 siRNA on HA-induced ASMC migration (Fig. 5D). In contrast, both HA-stimulated wound closure and HA-induced migration in Boyden chamber assay were significantly inhibited in RHAMM siRNA-treated ASMC (Fig. 5B–D). These data thus provide evidence that CD44 is not involved in HA-mediated ASMC migration and that only RHAMM is required for HA-mediated ASMC motility.

3.6. HA-induced Rac activation is mediated by RHAMM
Both Rac and RHAMM are required for HA-induced migration. We therefore hypothesized that stimulation of Rac activity is mediated by HA-induced RHAMM activation, while activation of RhoA would involve CD44. To assess this hypothesis, we therefore performed pull-down assay of RhoA and Rac activity in the absence and presence of siRNA targeting CD44 and RHAMM. As shown in Fig. 6A, siRNA targeting CD44 inhibited HA-induced RhoA activation by 61±7% at 10 min (n=3, P<0.001), without any effect on Rac activity. In contrast, siRNA targeting RHAMM decreased HA-mediated Rac activation by 67±6% (n=3, P<0.001), but did not affect RhoA activation (Fig. 6A). These findings indicate that stimulation of RhoA and Rac activity by HA is independently mediated by CD44 and RHAMM activation, respectively.


Figure 6
View larger version (48K):
[in this window]
[in a new window]
[Download PowerPoint slide]
  		Fig. 6 RHAMM-mediated Rac activation is PI3K-dependent. (A) RhoA and Rac expression and activity, and corresponding densitometric analysis after 10 min and 60 min of stimulation with 3 mg/ml HA for RhoA and Rac, respectively, in the absence and presence of siRNA targeting CD44 and siRNA targeting RHAMM. Control conditions, with and without HA corresponded to ASMC treated with siRNA-scrambled. Rho protein activity is normalized to Rho protein level and expressed as percentage of control. (B) Western blot for Rac expression and activity, expression of PAK and phosphorylated PAK, and corresponding densitometric analysis after 60 min of stimulation with 3 mg/ml HA, in the absence and presence of 25 µM LY-294002. Rac activity and PAK phosphorylation are expressed as percentage of control. The data presented are representative of at least three independent experiments. *p<0.001 vs control.

 
3.7. HA-induced Rac activation is dependent of PI3K
The inhibition of PI3K prevented HA-induced ASMC migration. We therefore assessed whether PI3K is involved in HA-mediated activation of Rac signaling pathway in ASMC. HA-induced activation of Rac signaling was monitored by measuring both the amount of GTP-bound Rac, and the level of activation/phosphorylation of the Rac target PAK in the absence and presence of the PI3K inhibitor LY-294002. Pull-down assay and western blot analysis show that HA-induced Rac and PAK activation was inhibited by LY-294002 (25 µM) (Fig. 6B) or wortmannin (100 nM, not shown). These results indicate that activation of Rac by HA-mediated stimulation of RHAMM is a PI3K-dependent process.


   4. Discussion
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
References
 
Despite several observations have suggested a major role for HA in the development of atherosclerosis and restenosis, little is known about the signaling pathways responsible for HA effects in ASMC. Herein, we demonstrate that high-molecular weight HA leads to independent activation of the actin cytoskeleton-regulating proteins RhoA and Rac through CD44 and RHAMM, respectively. ASMC migration induced by HA stimulation depends exclusively on RHAMM-mediated PI3K-dependent Rac activation.

In normal physiological conditions, HA consists of 2000–25,000 disaccharides, which corresponds to polysaccharides with relative mass of 1–10·106 Da [7]. Biological HA effects have been reported to depend on HA molecular mass [31,32]. Here we addressed the effect of HA of approximately 2·106 Da, typically found in injured arterial walls [33]. Although it is not possible to correlate the concentrations used with the in vivo physiological or pathological HA concentrations, the effective HA concentrations inducing ASMC migration (EC50~1.8 mg/ml) corresponded to active concentrations typically reported for high-molecular weight rooster comb HA, that ranged between 0.1 and 4.0 mg/ml [29,34–36].

It is recognized that migration of ASMC from the media plays a central role in progressive intimal thickening leading to arteriosclerosis or restenosis after arterial injury [37]. ASMC motility could be altered not only by several soluble factors but also by ECM proteins [38]. There is increasing evidence that HA facilitates ASMC locomotion [4]. Several mechanisms could participate to HA-induced ASMC motility. Around ASMC, formation of dynamic HA- and versican-rich pericellular matrix facilitates migration by diminishing cell surface adhesion and affecting cell shape [3]. HA also regulates cell motility through interaction with cell surface receptors such as CD44 and RHAMM, and activation of downstream intracellular signaling. The HA binding to CD44 affects cell adhesion to ECM components and is implicated in the stimulation of aggregation, proliferation, migration, and angiogenesis [8]. HA binding to CD44 regulates interaction of the cytoplasmic domain of CD44 with tyrosine kinases and with the cytoskeleton, and modulates the activity of the Rho proteins RhoA and Rac1 [20,22]. Activation of RhoA and its downstream effector Rho kinase by HA binding to CD44 has been proposed to be necessary for membrane–cytoskeleton interactions and tumor cell migration during the progression of breast cancers [22]. Selective interaction of the CD44 cytoplasmic domain with Rho protein guanine nucleotide exchange factors (p115RhoGEF, Tiam1, and Vav2) has been shown to play an important role in HA-mediated tumor cell migration [8,39]. In addition, regulation of Cdc42–cytoskeleton interaction by HA-CD44 binding is involved in ovarian tumor progression [40]. It is important to mention that the most part of studies analyzing intracellular signaling pathways coupling HA receptors to HA-mediated effects has been performed in transformed/tumor cells, and therefore probably cannot be extended to untransformed/normal cells. In this study, we demonstrate that in ASMC, HA induces activation of RhoA and Rac. However, siRNA-mediated HA receptor knockdown and blocking antibody indicate that only RhoA activation is mediated by CD44 and that, contrary to results obtained in tumor cells, this signaling pathway is not involved in HA-induced ASMC migration under our experimental conditions. Our observation is however in agreement with previous reports showing the major role of RHAMM in HA-stimulated migration of untransformed vascular cells, such as bovine aortic smooth muscle cells [4], and endothelial cells [41]. The varied role of CD44 in HA-induced cell migration may thus be attributable to differences in experimental design but other obvious differences include cell type (untransformed versus transformed), cell origin, cell passage, animal species, HA origin and size.

The role of HA-CD44-mediated RhoA activation has not been investigated here. However, our data suggest that this signaling pathway does not stimulate ASMC proliferation. As RhoA is a key regulator of ASMC contraction, CD44-mediated RhoA activation could induce ASMC contraction, which could participate to the previously described HA-induced enhancement of collagen contraction by ASMC [42]. This hypothesis is supported by (i) the observation of HA-induced stimulation of actin stress fiber formation, which is a RhoA-dependent phenomenon associated to cell contraction [11], and (ii) the inhibition of HA-induced stress fibers in cells treated with siRNA targeting CD44 (not shown).

Our results do not reveal a role for HA in the regulation of vascular smooth muscle cell proliferation. Although high concentrations of HA of unknown size have been reported to stimulate DNA synthesis in vascular smooth muscle cells [43], similarly to our results, high molecular weight HA applied for 24 h had no effect on aortic smooth muscle cells from monkey [42]. Very recently, fragmented, but not high molecular weight HA (>2·106 Da) has been shown to stimulate rat aortic smooth muscle cell proliferation in inverse correlation with fragment size [44].

Although RHAMM is identified as microtubule-associated protein that interacts with actin [8,10], the molecular bases of RHAMM-mediated cell motility are not clearly identified. Here we show that in ASMC, HA–RHAMM interaction activates Rac and that Rac activation is required for HA-mediated ASMC motility. To our knowledge, this is the first demonstration of the involvement of Rho protein in intracellular signaling downstream of RHAMM. An essential role of Rac in cell migration has been established in a large variety of cell types [45,46]. Rac is important for the formation of lamellipodia at the leading edge of the cells and forward movement, and is also involved in cell retraction at the trailing edge via its effector PAK. The observation that HA induced lamellipodia formation in ASMC is therefore in agreement with Rac-mediated cytoskeletal rearrangement and migration in response to HA. Inhibition of Rac by LT-toxin and siRNA targeting Rac almost completely prevented HA-induced ASMC migration indicating that Rac activation is a critical step in the intracellular pathway activated by HA-RHAMM mediating cell motility.

Results obtained with LY-294002 show that activation of PI3K is required for HA to stimulate ASMC motility. Furthermore, the analysis of Rac/PAK activation by HA indicated that PI3K is upstream to Rac activation. PI3Ks lipid products have been widely implicated in controlling cell migration and polarity [47,48]. The production of PI(3,4,5)P3 has been shown to lead to an increase in GTP-bound Rac in many cell types. The mechanism by which this lipid promotes GTP loading on Rac is thought to be through a direct interaction with Rac exchange factors. All members of the Dbl family of Rho protein exchange factor contain a pleckstrin homology domain and at least some of these can bind phospholipids. Tiam, PIX and Vav have been shown to be activated by PI(3,4,5)P3 [47,49]. It is therefore likely that HA-induced PI3K-dependent Rac activation and migration occurred through the activation of a PI(3,4,5)P3-dependent exchange factor.

Previously, drugs such as paclitaxel have been used to inhibit ASMC proliferation in in-stent restenosis by acting on the cytoskeleton [50]. Our results, showing that RHAMM-associated PI3K/Rac signaling is responsible for HA-induced actin cytoskeleton remodeling and cell migration might represent a new signaling pathway that could participate to the development of in-stent restenosis.


   Acknowledgements
 
This work was supported by grants from the Institut National de la Santé et de la Recherche Médicale (INSERM) and from la Fondation de l'Avenir. We thank Dr. Martin Schwartz for the gift of the plasmids encoding for the Rhotekin and PAK RBD (University of Virginia, Charlottesville, Virginia). We thank Cédric Boularan for his help in siRNA design and validation; Isabelle Suard and Nathalie Vaillant for excellent technical assistance.


   Notes
 
Time for primary review 19 days


   References
Top
 Abstract
 1. Introduction
 2. Materials and methods
 3. Results
 4. Discussion
 References
 

  1. Chung I.M., Gold H.K., Schwartz S.M., Ikari Y., Reidy M.A., Wight T.N. Enhanced extracellular matrix accumulation in restenosis of coronary arteries after stent deployment. J Am Coll Cardiol (2002) 40:2072–2081.[Abstract/Free Full Text]
  2. Chai S., Chai Q., Danielsen C.C., Hjorth P., Nyengaard J.R., Ledet T., et al. Overexpression of hyaluronan in the tunica media promotes the development of atherosclerosis. Circ Res (2005) 96:583–591.[Abstract/Free Full Text]
  3. Evanko S.P., Angello J.C., Wight T.N. Formation of hyaluronan- and versican-rich pericellular matrix is required for proliferation and migration of vascular smooth muscle cells. Arterioscler Thromb Vasc Biol (1999) 19:1004–1013.[Abstract/Free Full Text]
  4. Savani R.C., Wang C., Yang B., Zhang S., Kinsella M.G., Wight T.N., et al. Migration of bovine aortic smooth muscle cells after wounding injury. The role of hyaluronan and RHAMM. J Clin Invest (1995) 95:1158–1168.[ISI][Medline]
  5. Hascall V.C., Majors A.K., De La Motte C.A., Evanko S.P., Wang A., Drazba J.A., et al. Intracellular hyaluronan: a new frontier for inflammation? Biochim Biophys Acta (2004) 1673:3–12.[Medline]
  6. Collis L., Hall C., Lange L., Ziebell M., Prestwich R., Turley E.A. Rapid hyaluronan uptake is associated with enhanced motility: implications for an intracellular mode of action. FEBS Lett (1998) 440:444–449.[CrossRef][ISI][Medline]
  7. Toole B.P. Hyaluronan: from extracellular glue to pericellular cue. Nat Rev Cancer (2004) 4:528–539.[CrossRef][ISI][Medline]
  8. Turley E.A., Noble P.W., Bourguignon L.Y. Signaling properties of hyaluronan receptors. J Biol Chem (2002) 277:4589–4592.[Free Full Text]
  9. Kohda D., Morton C.J., Parkar A.A., Hatanaka H., Inagaki F.M., Campbell I.D., et al. Solution structure of the link module: a hyaluronan-binding domain involved in extracellular matrix stability and cell migration. Cell (1996) 86:767–775.[CrossRef][ISI][Medline]
  10. Assmann V., Jenkinson D., Marshall J.F., Hart I.R. The intracellular hyaluronan receptor RHAMM/IHABP interacts with microtubules and actin filaments. J Cell Sci (1999) 112(Pt 22):3943–3954.[Abstract]
  11. Etienne-Manneville S., Hall A. Rho GTPases in cell biology. Nature (2002) 420:629–635.[CrossRef][Medline]
  12. Loirand G., Guerin P., Pacaud P. Rho kinases in cardiovascular physiology and pathophysiology. Circ Res (2006) 98:322–334.[Abstract/Free Full Text]
  13. van Nieuw Amerongen G.P., van Hinsbergh V.W. Cytoskeletal effects of rho-like small guanine nucleotide-binding proteins in the vascular system. Arterioscler Thromb Vasc Biol (2001) 21:300–311.[Abstract/Free Full Text]
  14. Liu H.W., Halayko A.J., Fernandes D.J., Harmon G.S., McCauley J.A., Kocieniewski P., et al. The RhoA/Rho kinase pathway regulates nuclear localization of serum response factor. Am J Respir Cell Mol Biol (2003) 29:39–47.[Abstract/Free Full Text]
  15. Mack C.P., Somlyo A.V., Hautmann M., Somlyo A.P., Owens G.K. Smooth muscle differentiation marker gene expression is regulated by RhoA-mediated actin polymerization. J Biol Chem (2001) 276:341–347.[Abstract/Free Full Text]
  16. Seasholtz T.M., Zhang T., Morissette M.R., Howes A.L., Yang A.H., Brown J.H. Increased expression and activity of RhoA are associated with increased DNA synthesis and reduced p27(Kip1) expression in the vasculature of hypertensive rats. Circ Res (2001) 89:488–495.[Abstract/Free Full Text]
  17. Sauzeau V., Le Mellionnec E., Bertoglio J., Scalbert E., Pacaud P., Loirand G. Human urotensin II-induced contraction and arterial smooth muscle cell proliferation are mediated by RhoA and Rho-kinase. Circ Res (2001) 88:1102–1104.[Abstract/Free Full Text]
  18. Ai S., Kuzuya M., Koike T., Asai T., Kanda S., Maeda K., et al. Rho–Rho kinase is involved in smooth muscle cell migration through myosin light chain phosphorylation-dependent and independent pathways. Atherosclerosis (2001) 155:321–327.[CrossRef][ISI][Medline]
  19. Liu B., Itoh H., Louie O., Kubota K., Kent K.C. The signaling protein Rho is necessary for vascular smooth muscle migration and survival but not for proliferation. Surgery (2002) 132:317–325.[CrossRef][ISI][Medline]
  20. Oliferenko S., Kaverina I., Small J.V., Huber L.A. Hyaluronic acid (HA) binding to CD44 activates Rac1 and induces lamellipodia outgrowth. J Cell Biol (2000) 148:1159–1164.[Abstract/Free Full Text]
  21. Bourguignon L.Y., Zhu H., Shao L., Chen Y.W. CD44 interaction with tiam1 promotes Rac1 signaling and hyaluronic acid-mediated breast tumor cell migration. J Biol Chem (2000) 275:1829–1838.[Abstract/Free Full Text]
  22. Bourguignon L.Y., Zhu H., Shao L., Zhu D., Chen Y.W. Rho-kinase (ROK) promotes CD44v(3,8-10)–ankyrin interaction and tumor cell migration in metastatic breast cancer cells. Cell Motil Cytoskeleton (1999) 43:269–287.[CrossRef][ISI][Medline]
  23. Guibert C., Pacaud P., Loirand G., Marthan R., Savineau J.P. Effect of extracellular ATP on cytosolic Ca2+ concentration in rat pulmonary artery myocytes. Am J Physiol (1996) 271:L450–L458.[ISI][Medline]
  24. Fukui R., Shibata N., Kohbayashi E., Amakawa M., Furutama D., Hoshiga M., et al. Inhibition of smooth muscle cell migration by the p21 cyclin-dependent kinase inhibitor (Cip1). Atherosclerosis (1997) 132:53–59.[CrossRef][ISI][Medline]
  25. Ren X.D., Kiosses W.B., Schwartz M.A. Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J (1999) 18:578–585.[CrossRef][ISI][Medline]
  26. Deroanne C., Vouret-Craviari V., Wang B., Pouyssegur J. EphrinA1 inactivates integrin-mediated vascular smooth muscle cell spreading via the Rac/PAK pathway. J Cell Sci (2003) 116:1367–1376.[Abstract/Free Full Text]
  27. Teramoto H., Castellone M.D., Malek R.L., Letwin N., Frank B., Gutkind J.S., et al. Autocrine activation of an osteopontin-CD44-Rac pathway enhances invasion and transformation by H-RasV12. Oncogene (2005) 24:489–501.[CrossRef][ISI][Medline]
  28. Nagahara H., Vocero-Akbani A.M., Snyder E.L., Ho A., Latham D.G., Lissy N.A., et al. Transduction of full-length TAT fusion proteins into mammalian cells: TAT-p27Kip1 induces cell migration. Nat Med (1998) 4:1449–1452.[CrossRef][ISI][Medline]
  29. Coleman P.J., Scott D., Mason R.M., Levick J.R. Characterization of the effect of high molecular weight hyaluronan on trans-synovial flow in rabbit knees. J Physiol (1999) 514(Pt 1):265–282.[Abstract/Free Full Text]
  30. Mackay D.J., Hall A. Rho GTPases. J Biol Chem (1998) 273:20685–20688.[Free Full Text]
  31. McKee C.M., Lowenstein C.J., Horton M.R., Wu J., Bao C., Chin B.Y., et al. Hyaluronan fragments induce nitric-oxide synthase in murine macrophages through a nuclear factor kappaB-dependent mechanism. J Biol Chem (1997) 272:8013–8018.[Abstract/Free Full Text]
  32. McKee C.M., Penno M.B., Cowman M., Burdick M.D., Strieter R.M., Bao C., et al. Hyaluronan (HA) fragments induce chemokine gene expression in alveolar macrophages. The role of HA size and CD44. J Clin Invest (1996) 98:2403–2413.[ISI][Medline]
  33. Chajara A., Raoudi M., Delpech B., Leroy M., Basuyau J.P., Levesque H. Increased hyaluronan and hyaluronidase production and hyaluronan degradation in injured aorta of insulin-resistant rats. Arterioscler Thromb Vasc Biol (2000) 20:1480–1487.[Abstract/Free Full Text]
  34. Kikuchi T., Yamada H., Fujikawa K. Effects of high molecular weight hyaluronan on the distribution and movement of proteoglycan around chondrocytes cultured in alginate beads. Osteoarthritis Cartilage (2001) 9:351–356.[CrossRef][ISI][Medline]
  35. Pilloni A., Bernard G.W. The effect of hyaluronan on mouse intramembranous osteogenesis in vitro. Cell Tissue Res (1998) 294:323–333.[CrossRef][ISI][Medline]
  36. Sheehan K.M., DeLott L.B., West R.A., Bonnema J.D., DeHeer D.H. Hyaluronic acid of high molecular weight inhibits proliferation and induces cell death in U937 macrophage cells. Life Sci (2004) 75:3087–3102.[CrossRef][ISI][Medline]
  37. Clowes A.W., Reidy M.A., Clowes M.M. Kinetics of cellular proliferation after arterial injury: I. Smooth muscle growth in the absence of endothelium. Lab Invest (1983) 49:327–333.[ISI][Medline]
  38. Wight T.N., Merrilees M.J. Proteoglycans in atherosclerosis and restenosis: key roles for versican. Circ Res (2004) 94:1158–1167.[Abstract/Free Full Text]
  39. Bourguignon L.Y., Singleton P.A., Zhu H., Diedrich F. Hyaluronan-mediated CD44 interaction with RhoGEF and Rho kinase promotes Grb2-associated binder-1 phosphorylation and phosphatidylinositol 3-kinase signaling leading to cytokine (macrophage-colony stimulating factor) production and breast tumor progression. J Biol Chem (2003) 278:29420–29434.[Abstract/Free Full Text]
  40. Bourguignon L.Y., Gilad E., Rothman K., Peyrollier K. Hyaluronan–CD44 interaction with IQGAP1 promotes Cdc42 and ERK signaling, leading to actin binding, Elk-1/estrogen receptor transcriptional activation, and ovarian cancer progression. J Biol Chem (2005) 280:11961–11972.[Abstract/Free Full Text]
  41. Savani R.C., Cao G., Pooler P.M., Zaman A., Zhou Z., DeLisser H.M. Differential involvement of the hyaluronan (HA) receptors CD44 and receptor for HA-mediated motility in endothelial cell function and angiogenesis. J Biol Chem (2001) 276:36770–36778.[Abstract/Free Full Text]
  42. Travis J.A., Hughes M.G., Wong J.M., Wagner W.D., Geary R.L. Hyaluronan enhances contraction of collagen by smooth muscle cells and adventitial fibroblasts: role of CD44 and implications for constrictive remodeling. Circ Res (2001) 88:77–83.[Abstract/Free Full Text]
  43. Jain M., He Q., Lee W.S., Kashiki S., Foster L.C., Tsai J.C., et al. Role of CD44 in the reaction of vascular smooth muscle cells to arterial wall injury. J Clin Invest (1996) 97:596–603.[ISI][Medline]
  44. Joddar B., Ramamurthi A. Fragment size- and dose-specific effects of hyaluronan on matrix synthesis by vascular smooth muscle cells. Biomaterials (2006) 27:2994–3004.[CrossRef][ISI][Medline]
  45. Kjoller L., Hall A. Signaling to Rho GTPases. Exp Cell Res (1999) 253:166–179.[CrossRef][ISI][Medline]
  46. Nobes C.D., Hall A. Rho GTPases control polarity, protrusion, and adhesion during cell movement. J Cell Biol (1999) 144:1235–1244.[Abstract/Free Full Text]
  47. Raftopoulou M., Hall A. Cell migration: Rho GTPases lead the way. Dev Biol (2004) 265:23–32.[CrossRef][ISI][Medline]
  48. Stephens L., Ellson C., Hawkins P. Roles of PI3Ks in leukocyte chemotaxis and phagocytosis. Curr Opin Cell Biol (2002) 14:203–213.[CrossRef][ISI][Medline]
  49. Wymann M.P., Marone R. Phosphoinositide 3-kinase in disease: timing, location, and scaffolding. Curr Opin Cell Biol (2005) 17:141–149.[CrossRef][ISI][Medline]
  50. Grube E., Silber S., Hauptmann K.E., Mueller R., Buellesfeld L., Gerckens U., et al. TAXUS I: six- and twelve-month results from a randomized, double-blind trial on a slow-release paclitaxel-eluting stent for de novo coronary lesions. Circulation (2003) 107:38–42.[Abstract/Free Full Text]

Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?



This Article
Right arrow Abstract Freely available
Right arrow FREE Full Text (PDF) Freely available
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrowRequest Permissions
Google Scholar
Right arrow Articles by Gouëffic, Y.
Right arrow Articles by Loirand, G.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Gouëffic, Y.
Right arrow Articles by Loirand, G.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?