The JI
HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
QUICK SEARCH:   [advanced]


     
 

The Journal of Immunology, 2007, 179, 901-909
Copyright © 2007 by The American Association of Immunologists, Inc.
This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
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 Download to citation manager
Right arrow Request Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Segain, J.-P.
Right arrow Articles by Pacaud, P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Segain, J.-P.
Right arrow Articles by Pacaud, P.

Urotensin II is a New Chemotactic Factor for UT Receptor-Expressing Monocytes1

Jean-Pierre Segain2,*,{dagger}, Malvyne Rolli-Derkinderen2,{ddagger},{dagger}, Nadine Gervois§,¶, Diane Raingeard de la Blétière*,{dagger}, Gervaise Loirand{ddagger},{dagger},|| and Pierre Pacaud3,{ddagger},{dagger}

* Institut National de la Recherche Agronomique, Unité Mixte de Recherche 1280-Physiologie des Adaptations Nutritionnelles, Nantes, France; {dagger} Faculté des Sciences, Université de Nantes, l’Institut du Thorax, Nantes, France; {ddagger} Institut National de la Santé et de la Recherche Médicale, Unité 533, Nantes, France; § Institut National de la Santé et de la Recherche Médicale, Unité 601, Nantes, France; Faculté de Médecine, Université de Nantes, Nantes, France; and || Centre Hospitalier de l’Université Nantes, l’Institut du Thorax, Nantes, France


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
Urotensin II (U-II), a vasoactive cyclic neuropeptide which activates the G protein-coupled receptor UT receptor, exerts various cardiovascular effects and may play a role in the pathophysiology of atherosclerosis. In this study, we report that the UT receptor is expressed and functional on human PBMC and rat splenocytes. PBMC surface expression of the UT receptor was mainly found in monocytes and NK cells, also in a minority of B cells, but not in T cells. Stimulation of monocytes with LPS increased UT receptor mRNA and protein expression. Cloning and functional characterization of the human UT receptor gene promoter revealed the presence of NF-{kappa}B-binding sites involved in the stimulation of UT receptor gene expression by LPS. Activation of the UT receptor by U-II induced chemotaxis with maximal activity at 10 and 100 nM. This U-II effect was restricted to monocytes. Analysis of the signaling pathway involved indicated that U-II-mediated chemotaxis was related to RhoA and Rho kinase activation and actin cytoskeleton reorganization. The present results thus identify U-II as a chemoattractant for UT receptor-expressing monocytes and indicate a pivotal role of the RhoA-Rho kinase signaling cascade in the chemotaxis induced by U-II.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 Disclosures
 References
 
It is well-established that neuropeptides play a role in the regulatory mechanisms between neuroendocrine and immune systems both in physiological and pathological conditions such as autoimmune diseases and chronic inflammation (1, 2). Studies have shown that binding of neuropeptides to specific seven transmembrane domain G protein-coupled receptors expressed in various types of immune cells can activate or suppress different immune functions such as cytokine production (3), dendritic cell (DC)4 maturation (4), Th2 cell polarization (5), and chemotaxis (6, 7). These features and their implication in several inflammatory and autoimmune diseases have led to the development of receptor-selective agonist and antagonist for the treatment of several diseases (8).

The human neuropeptide urotensin II (U-II) is a cyclic peptide of 11 aa that was identified as the ligand for the orphan G protein-coupled receptor 14, recently renamed UT receptor (9, 10). Expression of U-II and UT receptor has been reported in kidney, CNS, and in the cardiovascular system (10, 11, 12, 13, 14). Pharmacological studies have shown that U-II exerts potent vasoactive activity in different mammal species, including humans and rats (13, 15, 16, 17, 18, 19). The contractile activity of U-II is mediated by the activation the small GTPase RhoA and its effector, Rho kinase (20). In addition to its vasoactive effect, U-II also displays mitogenic effects on vascular smooth muscle cells through activation of the RhoA/Rho kinase pathway (20). U-II has been shown to act synergistically with mildly oxidized low-density lipoprotein in inducing vascular smooth muscle cell proliferation suggesting a possible role of U-II in the pathophysiology of atherosclerosis (21). In agreement with this hypothesis, recent immunohistochemical studies have reported increased expression of U-II and UT receptor in atherosclerotic human aorta and coronary arteries (19, 22, 23). Interestingly, U-II immunoreactivity localized to regions of macrophage infiltration within the atherosclerotic lesions (19), and U-II and UT receptor mRNA were respectively detected in lymphocytes and in monocytes/macrophages isolated from human PBMC of healthy subjects (23). However, the significance of the presence of U-II and UT receptor in inflammatory regions of atherosclerotic lesions and the possible role of U-II/UT receptor in immune and inflammatory reactions are not yet elucidated.

In the present study, we investigated a potential immune function of U-II. We found that in PBMC, the majority of monocytes and a large portion of NK cells express UT receptor. Moreover, we demonstrated that the UT receptor functions as a chemoattractant receptor for U-II in human PBMC and rat splenocytes. U-II-induced chemotaxis was blocked by inhibition of the RhoA effector Rho kinase. Accordingly, U-II induced RhoA activation, actin cytoskeleton reorganization, and myosin L chain (MLC) phosphorylation. Finally, the finding that UT receptor gene expression is up-regulated by inflammatory stimuli, together with the chemotactic activity of U-II, reinforce the idea of a possible role of this peptide and its receptor in the pathophysiology of atherosclerosis.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 Disclosures
 References
 
Reagents

Tissue-culture products were obtained from Invitrogen Life Technologies. TNF-{alpha}, IL-1beta, and IFN-{gamma} were obtained from PeproTech. The PE-conjugated mAbs, anti-CD3 (clone HIT3a), -CD19 (clone 4G7), -CD56 (clone B159), and the allophycocyanin-conjugated mAbs anti-CD4 (clone RPA-T4), -CD8 (clone RPA-T8), -CD16 (clone 3G8) were obtained from BD Biosciences. The PE-conjugated mAbs anti-CD14 (clone RMO52), anti-TCR{alpha}beta (clone BMA031), and anti-TCR{gamma}{delta} (clone IMMY510) were obtained from Beckman Coulter. The rabbit anti-human UT receptor Ab (H-90), the mouse anti-RhoA mAb (26C4), and the goat anti-MLC2 Ab (A-20) were obtained from Santa Cruz Biotechnology. The rabbit anti-phospho-MLC2 (Ser19) Ab was obtained from Cell Signaling Technology. Human U-II, the inactive U-II analog (Asp-Cys-Phe-Ala-Lys-Tyr-Cys-Val), was provided by Prof. H. Vaudry (Institut National de la Santé et de la Recherche Médicale, Unité 413, Rouen, France) (24). All other reagents were purchased from Sigma-Aldrich.

Cell isolation

The protocols have been reviewed and approved by our institutional review committee. Human PBMC were isolated from heparinized blood of healthy volunteers by Ficoll density gradient centrifugation and suspended in RPMI 1640 medium supplemented with 2 mM L-glutamine, 100 IU/ml penicillin, and 100 µg/ml streptomycin.

Polymorphonuclear cells (PMN) were recovered from the lower phase of Ficoll gradient and separated from RBC by addition of 6% dextran (m.w. 500 000, final concentration: 1% dextran). After sedimentation, the supernatant was centrifuged, then the PMN pellet was suspended in RPMI 1640. The purity of the PMN preparation assessed by Giemsa coloration was 98%.

Monocytes were isolated by elutriation (Aventi J-20; Beckman Coulter). CD3+ T cells and NK cells were purified using the CD3 and NK isolation kits, respectively, according to the manufacturer’s instructions (Miltenyi Biotec). Purity of cell populations exceeded 95% as assessed by flow cytometry.

To generate immature monocyte-derived DCs (iDC), elutriated monocytes were cultured in culture medium containing 500 IU/ml GM-CSF and 200 IU/ml IL-4 (AbCys) for 6 days. Mature DC (mDC) were obtained by an additional 48-h treatment of iDC with 1 µg/ml LPS.

Splenocytes were isolated from Sprague-Dawley rats as previously described with slight modifications (25). Briefly, the spleen was perfused with the above RPMI 1640 medium and the cell suspension was centrifuged at 1500 rpm for 5 min. The pellet was resuspended and incubated in 0.17 M ammonium chloride for 10 min at room temperature to lyse erythrocytes. Splenocytes were then washed and suspended in RPMI 1640 medium.

Cell culture

To analyze the modulation of UT receptor expression by several stimuli, PBMC (5 x 106) were cultured in RPMI 1640 medium supplemented with 10% FCS at 37°C, in the absence or presence of LPS (1 µg/ml), TNF-{alpha} (10 ng/ml), IL-1beta (10 ng/ml), IFN-{gamma} (500 U/ml), or PMA (50 ng/ml) for 6 or 18 h. Cells were then recovered for the analysis of mRNA and protein expressions by real-time RT-PCR and flow cytometry, respectively. Cells were then recovered for RNA isolation. To analyze RhoA activation and MLC phosphorylation, splenocytes (60 x 106) were preincubated at 37°C in 30 ml of RPMI 1640/0.1% BSA containing or not the Rho-kinase inhibitor Y-27632 (10 µM) for 20 min, before addition of 10 nM U-II or 1 µg/ml LPS for 1 more hour. Cells were then centrifuged and washed twice with ice-cold PBS and pellets were rapidly frozen in liquid nitrogen and lysed as described below.

Flow cytometry analysis

Flow cytometry analysis of surface expression of the UT receptor was analyzed on freshly isolated PBMC by a two- or three-color method using mAbs directed to human leukocyte Ags (i.e., CD3, CD4, CD8, CD14, CD16, CD19, CD56, TCR{alpha}beta, and TCR{gamma}{delta}), and an anti-human UT receptor Ab. Cells were also stained with appropriate isotype-matched control Abs. FcRs and nonspecific binding sites were blocked during a saturation step by incubating cells with PBS containing 3% normal donkey serum and 2% human AB serum (Sigma-Aldrich). PBMC were stained for 20 min with the anti-UT receptor Ab. Cells were then washed with PBS-0.1%BSA, and incubated for 20 min with a donkey anti-rabbit FluoProbes 488 Ab (FluoProbes; Interchim) and with the anti-CD3, -CD4, -CD8, -CD14, -CD19, or -CD56-PE mAb. After washing, cells were resuspended in PBS and analyzed on a FACScan flow cytometer using the CellQuest software (BD Biosciences). Compensation was checked before each acquisition and 105 events were collected.

RNA isolation and real-time RT-PCR

Total RNA was isolated using TRIzol reagent (Invitrogen Life Technologies) and treated for 45 min at 37°C with RQ1 DNase (Promega). One microgram of RNA was reverse transcribed using random primers and the Superscript III-Reverse Transcriptase (Invitrogen Life Technologies) at 50°C for 45 min, according to the manufacturer’s instructions. The resulting cDNA was subjected to PCR in a Bio-Rad iCycler iQ system using the QuantiTect SYBR Green PCR kit (Qiagen) and specific primers for human or rat UT receptor and beta-actin mRNA. Human (h) primers are as follow (5' to 3'): hUT receptor sense, CACGGGCACCATTGGGACTC; hUT receptor antisense, CGCCAGGTTGACCACGTAGAC; hbeta-actin sense, TGCTATCCAGGCTGTGCTATCC; hbeta-actin antisense GCCAGGTCCAGACGCAGG. Rat (r) primers (5' to 3'): rUT receptor sense, CCTCGTGCCCACCGTACTTAC; rUT receptor antisense, CCGCCGTGTCTGCTTGAAAG; rbeta-actin sense, CTATCGGCAATGAGCGGTTCC; rbeta-actin antisense, GCACTGTGTTGGCATAGAGGTC. The expression level of beta-actin was used as a reference value to normalize UT receptor gene expression in each sample. Relative quantitative gene expression among different PBMC treatment was calculated by the 2–{Delta}{Delta}Ct method (26), using untreated controls as the calibrator samples (where Ct is the cycle threshold). The validity of this approach was confirmed in preliminary experiments showing equivalent amplification efficiencies of primer pairs.

Cloning and site-directed mutagenesis of the human UT receptor promoter

The sequence of 1500 bp upstream of the ATG codon of the human UT receptor gene (chromosome 17 contig. NT-010663+) was cloned into the pDRIVE vector (Qiagen) by PCR using the 5'-GCACGCTTGTCATTTTCTGA-3' sense primer (up), the 5'-GTGGGGAAAGAAGCAAATCA-3' antisense primer (down), and human genomic DNA as matrix. This sequence was fused to the luciferase coding gene in the pGL2-Basic vector (Promega) between KpnI and HindIII restriction sites. The entire sequence was verified by sequencing the insert with pGL1 and pGL2 primers (Promega).

In vitro site-directed mutagenesis of the four NF-{kappa}B-binding sites

The first (distal to ATG), second, third, and fourth (proximal to ATG) NF-{kappa}B-binding site of the human UT receptor promoter was mutated to inactive ones according to the QuikChange site-directed mutagenesis kit instruction manual (Stratagene) using the following PAGE-purified primers: N1: up 5'-CCCTGACTGGCATTATCCTGCTTGTTGGC-3', down 5'-GCCAACAAGCAGGATAATGCCAGTCAGGG-3'; N2 up 5'-TGTCCTTTTCATGACGTCCACGGTCACGGTTTTGG-3', down 5'-CCAAAACCGTGACCGTGGACGTCATGAAAAGGACA; N3 up 5'-TCTGCCCCAGAGTCTGCACTCCTGTGTTCCGG-3', down 5'-CCGGAACACAGGAGTGCAGACTCTGGGGCAGA-3'; N4 up 5'-TGGCTTCCAGAGAGTCTAGAGAGTTGGAGGGC-3'; N4 down 5'-GCCCTCCAACTCTCTAGACTCTCTGGAAGCCA-3' to obtain the N1 to N4 mutants, respectively.

Transfections and reporter assays

Splenocytes freshly isolated from rat spleen were electroporated with wild-type (WT) UT receptor or N1 to N4 mutants (Amaxa Nucleofector). One day after transfection, cells were pretreated or not with MEK1/2 (U0126, 10 µM) or NF-{kappa}B (caffeic acid phenethyl ester (CAPE), 10 µM) inhibitors and then stimulated with LPS (LPS 2 µg/ml)) for different times (as indicated). The pIRES-EGFP vector was always cotransfected to estimate the level of transfection (fluorescence measure from lysates with Victor2) and to normalize the luciferase activity measured with the luciferase reporter reagent (Promega) in a LB96V luminometer (Berthold Technologies).

Chemotaxis assay

Chemotaxis assays were performed using Transwell cell culture chambers (Costar) with 6.5-mm diameter, 5-µm pore size polycarbonate membranes essentially as described (27). PBMC or splenocytes (0.5 x 106) suspended in 100 µl of RPMI 1640/0.1% BSA were added to the upper chamber and U-II or the inactive analog S28981-1 (at the indicated doses) were added to the lower chamber. In checkerboard control experiments, U-II was also added in the upper chamber. When indicated, cells were pretreated with 0.1 ng/ml IL-1beta at 37°C for 18 h or with 10 µM Y-27632 or 20 µM ML-7 at 37°C for 30 min. Cells were allowed to migrate for 3 h at 37°C. After this period, migrated cells onto the lower well were counted in four high-power fields under light microscopy and also recovered and counted with a Malassez counting chamber in triplicates under light microscopy. Cell migration was expressed as the chemotactic index calculated by the following ratio as previously described (number of cells migrating to U-II/cells migrating to vehicle) (7).

Calcium [Ca2+] measurements

[Ca2+] was measured fluorometrically in monocytes loaded with the intracellular probe fura 2-AM (Molecular Probes/Invitrogen Life Technologies). Briefly, 3 x 105 monocytes were plated on 35-mm diameter glass-bottom culture dishes (MatTek) and incubated with 1 µM fura 2-AM in HBSS at 37°C for 30 min, then washed, and further incubated in fresh HBSS at 37°C for 30 min in the experimental cuvette. Cells were stimulated with 0.1 µM U-II or 10 µM ATP (used as control) and fluorescence was recorded by videoimaging (Leica). Fluorescence intensities (340 and 380 nm) were stored and analyzed using Metafluor software (Roper Scientific).

Actin staining

Polymerized (F) actin staining was performed as previously described (28), with slight modifications. PBMC or splenocytes (0.5 x 106) were plated in culture chambers (Lab-Tek; Nunc) and preincubated at 37°C without or with the Rho-kinase inhibitor Y-27632 (10 µM) in 400 µl of RPMI 1640/0.1% BSA, for 30 min. Different concentrations (as indicated) of U-II were then added to the culture medium and cells were maintained at 37°C for 1 h. After washing with PBS, cells were fixed in PBS containing 2% paraformaldehyde, washed, and permeabilized with PBS/0.1% Triton X-100 for 5 min. Cells were then stained with Alexa Fluor 488 phalloidin (Molecular Probes) (1 U/ml in PBS) for 20 min at room temperature. After four to five washes, cells were mounted in Vectashield medium (Vector Laboratories/AbCys) and examined with a fluorescence microscope (Nikon).

Analysis of RhoA activity and MLC phosphorylation

Cells were homogenized in ice-cold lysis buffer containing 50 mM Tris-HCl (pH 7.5), 500 mM NaCl, 10 mM MgCl2, 1% Triton X-100, 0.5% sodium deoxycholate, 0.1% SDS, 10 µg/ml aprotinin, 10 µg/ml leupeptin, 1 mM PMSF, and 1 mM sodium orthovanadate. RhoA activity was assessed by a pull-down assay, using the Rho-binding domain of the Rho effector protein rhotekin as described previously (29). Precipitated GTP-bound RhoA and total RhoA were then analyzed by Western blot using a mouse anti-RhoA mAb. Cell lysates were also analyzed by Western blot with Abs directed to total or phosphorylated MLC. Immunoreactive bands were visualized using HRP-conjugated secondary Abs and subsequent ECL detection (Amersham Pharmacia Biotech).

Statistical analysis

The significance of differences was determined by the Mann-Whitney U test and Kruskall-Wallis ANOVA using the Statview software (Abacus Concepts). A difference with p < 0.05 was considered significant.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 Disclosures
 References
 
Expression of UT receptor mRNA in human and rat mononuclear cells

UT receptor mRNA expression was analyzed by real-time RT-PCR using total RNA extracted from freshly isolated, noncultured human PBMC and rat splenocytes. Expression was detected in both human and rat mononuclear cells, with a higher interindividual variability observed in human subjects (Fig. 1). UT mRNA expression was also detected in RNA extracted from Jurkat U-937 or THP1 leukemia cell lines (data not shown).


Figure 1
View larger version (14K):
[in this window]
[in a new window]

  		FIGURE 1. Expression of UT receptor mRNA. UT receptor mRNA expression has been analyzed by quantitative real-time RT-PCR in human PBMC and rat splenocytes. Boxes represent the median ± quartiles of the relative UT receptor mRNA expression calculated by the 2–{Delta}{Delta}Ct method in different cell population (PBMC, n = 6; splenocytes, n = 5); vertical bars indicate the 10th and 90th percentiles.

 
Cell surface expression of UT receptor by PBMC

We next wanted to phenotype the subpopulation of PBMC-expressing cell surface UT receptor protein by flow cytometry. The analysis of surface expression of UT receptor was performed on freshly isolated human PBMC using a polyclonal Ab directed against the N-terminal domain of the human UT protein. The proportion of UT receptor-positive cells in total PBMC varied from 15 to 30% among healthy individuals (Fig. 2A). This variability among individuals was similar to that that observed at the level of mRNA expression (Fig. 1). We next combined the analysis for UT receptor with the analysis of Ags associated with monocytes (CD14), NK cells (CD56), B cells (CD19), and T cells (CD3, CD4, CD8, TCR{alpha}beta, and TCR{gamma}{delta}). We found that the UT receptor was mainly expressed by monocytes since near 98% of the cells positive for the CD14/LPS receptor were UT receptor positive (Fig. 2B and Table I). The UT receptor was also consistently expressed by NK cells (68% of CD56+ cells) and to a lesser extent by B lymphocytes (10% of CD19+ cells) (Fig. 2B and Table I). Only <2% of CD3+ T lymphocytes were positive for the UT receptor, with similar proportions of CD4+ and CD8+ T cells (Fig. 2B and Table I). In accordance, T cells expressing either {alpha}beta or {gamma}{delta} T cell receptors barely expressed the UT receptor (Fig. 2B). The population expressing low levels of CD4 (15% of PBMC) and positive for the UT receptor (Fig. 2B) corresponds to CD14+ cells as confirmed by three-color flow cytometry analysis (data not shown). Similarly, the UT receptor was also found in cells expressing low levels of CD8 (9% of PBMC) that were shown to be CD56+ NK cells (data not shown). Thus, all the monocytes and a high fraction of NK cells present in human peripheral blood express the UT receptor on their cell surface. These two subsets account for ~95% of total UT receptor-positive cells in PBMC (calculated from data in Table I). To further characterize monocyte populations expressing the UT receptor, we performed a three-color flow cytometry analysis of PBMC stained for the UT receptor, CD14, and CD16 (Fc{gamma} RIII) receptors. Indeed, two main populations of human blood monocytes can be distinguished on the basis of CD14 and CD16 expression (30). Four main subpopulations could be identified: CD14highCD16– (region R1), CD14lowCD16– (R2), CD14highCD16+ (R3), CD14lowCD16+ (R4) cells, representing, respectively, 67, 11, 14, and 8% of the total monocytes (Fig. 2C and Table II). All these monocyte populations express the UT receptor. However, CD16-positive cells (21% of CD14+ cells) expressed higher levels (X3) of the UT receptor, as assessed by the mean fluorescence intensity (MFI), than CD16-negative cells (Table II).


Figure 2
View larger version (50K):
[in this window]
[in a new window]

  		FIGURE 2. Flow cytometry analysis of UT receptor cell surface expression in human PBMC. Freshly isolated PBMC from three healthy donors were stained with anti-human UT receptor/FluoProbeR488-conjugated secondary Abs and with PE/allophycocyanin-anti-CD3, -CD4, -CD8, -CD14, -CD16, -CD19, or -CD56, -TCR{alpha}beta, -TCR{gamma}{delta} mAbs and analyzed by flow cytometry. A, Representative histogram of UT receptor expression in total PBMC. B, Panels show representative dot plots of expression of UT receptor vs CD14, CD56, CD16, CD3, CD4, CD8, TCR{alpha}beta, TCR{gamma}{delta}, and CD19. Percentages of single-positive and double-positive cells among total PBMC are indicated. C, Representative dot plots of expression of CD14 vs CD16. Analysis of UT receptor expression on gated CD14highCD16– (R1), CD14lowCD16– (R2), CD14highCD16+ (R3), and CD14lowCD16+ (R4) populations is reported in Table II.

 

View this table:
[in this window]
[in a new window]

  		Table I. Proportion of UT receptor-positive cells in PBMC subsets

 

View this table:
[in this window]
[in a new window]

  		Table II. Expression of UT receptor by monocyte subsetsa

 
We also analyzed UT receptor expression in PMN and monocyte-derived DC. All PMN and iDC expressed the UT receptor at very low levels in comparison to monocytes (PMN: MFI = 9.23; iDC: MFI = 4.74; representative of three individuals). Maturation of iDC by treatment with LPS stimulated by 2-fold UT receptor expression (mDC: MFI = 9.06, representative of three individuals).

Inflammatory stimuli up-regulate UT receptor mRNA expression

Modulation of membrane receptor function and expression influences immune cell responses (31). We therefore assessed whether stimulation of PBMC with proinflammatory cytokines modulates UT receptor mRNA expression. As shown in Fig. 3A, stimulation for 6 h with LPS (2 µg/ml), IL-1beta (10 ng/ml), TNF-{alpha} (10 ng/ml), and IFN-{gamma} (500 U/ml) increased UT receptor mRNA expression by 10-, 8-, and 4-fold over control, respectively.


Figure 3
View larger version (17K):
[in this window]
[in a new window]

  		FIGURE 3. Up-regulation of UT receptor by inflammatory stimuli. A, UT receptor mRNA expression has been analyzed by quantitative real-time RT-PCR in human PBMC in the absence (control) and the presence of LPS (2 µg/ml), IL-1beta (10 ng/ml), TNF-{alpha} (10 ng/ml), and IFN-{gamma}, (500 U/ml). Results are expressed relative to beta-actin expression and normalized to the control as taken as 1. B, Flow cytometry analysis of the effect of LPS (2 µg/ml) on UT receptor expression in monocytes (CD14+), NK cells (CD56+), T lymphocytes (CD3+), and B lymphocytes (CD19+). Results are expressed as mean fluorescence intensity (MFI) ± SEM of three individuals (*, p < 0.05 and **, p < 0.01 vs control).

 
LPS up-regulates UT receptor expression in monocytes

The cell subpopulations responsible for LPS-induced UT receptor up-regulation have been further characterized by flow cytometry analysis of PBMC stained for the UT receptor and, CD14 (monocytes), CD56 (NK cells), CD3 (T lymphocytes), or CD19 (B lymphocytes). Stimulation by LPS (2 µg/ml) for 18 h induced a 2.6-fold-increase in UT receptor expression in monocytes cells but did not affect U-II receptor expression on CD3+ T cells, CD16+ NK cells, and CD19+ B cells (Fig. 3B). In agreement with the stimulatory effect of LPS on UT receptor mRNA, this result indicates that LPS up-regulated UT receptor expression in monocytes. Although TNF-{alpha} (10 ng/ml, 6 h) stimulates U-II receptor mRNA expression, we did not observe such an effect at the protein level by flow cytometry analysis after 18 h of stimulation with TNF-{alpha} (10 ng/ml; data not shown).

Four NF-{kappa}B-binding sites are identified in the human UT receptor gene promoter sequence

To gain additional insight into the mechanisms of transcriptional regulation of UT receptor expression, we examined the human UT receptor promoter using Genomatix tools (32). A promoter structure was found in a sequence of 1500 bp upstream of the ATG codon of the human UT receptor gene on chromosome 17. Using MatInspector (33, 34), we located potential transcription factor-binding sites for early growth response gene 1 (Egr), Elk1, GATA, CRE, AP1–2-4, RasREBP1, or NF-{kappa}B in this sequence (Fig. 4).


Figure 4
View larger version (65K):
[in this window]
[in a new window]

  		FIGURE 4. Identification of the human UT receptor promoter sequence. The first 1500 bp of the 5' upstream sequence of the human UT receptor gene was analyzed using MatInspector to find promoter structure. Egr, Elk1, GATA, CRE, Ap1–2-4, RasREBP1 (shaded), or NF{kappa}B (underlined) potential binding sites were found in this sequence. Up and down boxes indicates the primers used to fuse the promoter to the luciferase gene for functional experiments.

 
LPS induces UT receptor promoter activity through MAPK and NF-{kappa}B activation

To identify the transcriptional regulation involved in LPS-induced stimulation of UT receptor mRNA expression, functional analysis of the human UT receptor promoter was performed. Splenocytes were transfected with the 1500-bp fragment upstream of the ATG codon of the human UT receptor gene fused to the reporter gene firefly luciferase. Measurement of luciferase activity showed that LPS (2 µg/ml) induced a transient 8-fold increase in UT receptor promoter activity after 4–6 h of stimulation. After 24 h of LPS treatment, the promoter activity was nonsignificantly different from that measured under control condition. As MAPK and NF-{kappa}B pathways are major signaling pathways involved in the transcriptional effect of LPS, a pharmacological analysis has been performed to assess their involvement in LPS-induced stimulation of UT receptor promoter activity. Treatment of UT receptor promoter expressing splenocytes with the MEK1/2 inhibitor U0126 (10 µM) or the NF-{kappa}B inhibitor CAPE (20 µM) suppressed the stimulatory effect of LPS on UT receptor promoter activity (Fig. 5A). These results thus indicate that the stimulation of UT receptor promoter activity by LPS depends on MEK1/2 and NF-{kappa}B activation. This observation is in agreement with the presence of four predicted NF-{kappa}B-binding sites (N1-N4) in the UT receptor promoter (Fig. 4). To identify the NF-{kappa}B-binding sites involved in the LPS induction of UT receptor promoter activity, four UT receptor promoter constructs containing inactivating mutation of each putative NF-{kappa}B-binding sequence were transfected to splenocytes. Mutational ablation of the N1 and N3 NF-{kappa}B-binding sites did not modify the effect of LPS on UT receptor promoter activity (Fig. 5B). In contrast, in cells expressing UT receptor gene promoter carrying the mutational ablation of the N2 or N4 NF-{kappa}B-binding sites, LPS had no effect on promoter activity. These results provide evidence that NF-{kappa}B-binding sites N2 and N4 of the UT receptor promoter are necessary to induce UT receptor gene transcription in response to LPS stimulation.


Figure 5
View larger version (24K):
[in this window]
[in a new window]

  		FIGURE 5. LPS induces UT receptor promoter activity through ERK and NF-{kappa}B activation. Luciferase activity of splenocytes expressing UT receptor gene promoter constructs fused to the luciferase gene. A, Twenty-four hours posttransfection, cells were stimulated with LPS (2 µg/ml) for indicated times with or without pretreatment with of MEK1/2 (U0126, 10 µM) or NF-{kappa}B inhibitor (CAPE, 10 µM). B, Splenocytes expressing WT UT receptor promoter or NF-{kappa}B-binding site inactive mutants (N1–N4) were stimulated with LPS (2 µg/ml) for 4 h. Luciferase activity was measured as described in Materials and Methods and normalized by measuring the fluorescence of enhanced GFP (EGFP) expressed by the cotransfected pIRES-EGFP vector. The data are the means ± SD of four independent experiments performed in duplicate (A and B). (*, p < 0.01 vs control (t = 0); #, p < 0.005 vs LPS in A and LPS/WT in B).

 
U-II is a chemoattractant for human monocytes

The next question to address was therefore to assess the functional role of UT receptor activation by its ligand U-II in PBMC. In preliminary experiments, the effect of U-II has been tested on several immune functions. Treatment of PBMC with different doses of U-II (0.1 nM to 10 µM) for 18 h did not affect cell viability (trypan blue exclusion test), cell proliferation (colorimetric MTT assay), or cytokine production (TNF-{alpha} and IL-8, biological and ELISA, respectively) (data not shown).

The fact that the UT receptor is structurally related to chemokine receptors, together with studies showing that several neuropeptides have chemoattracting properties (6, 7) prompted us to examine whether U-II could induce chemotaxis of PBMC. We observed that U-II (1 nM-1 µM) induced PBMC chemotaxis with typical bell-shaped dose-response curves, indicating a receptor-mediated effect (Fig. 6A). Notably, the majority of the cells migrated onto the lower well were adherent to plastic (Fig. 6A). The maximum activity obtained with 10 and 100 nM U-II is similar to that obtained in response to 10 nM IL-8 (chemotactic index = 2.18 ± 0.6, n = 3; data not shown). In checkerboard control experiments, when U-II was also added in the upper chamber, PBMC chemotaxis was not observed. The inactive U-II analog had no effect on PBMC chemotaxis (data not shown). Furthermore, pretreatment of PBMC with IL-1beta for 12 h potentiated the chemotactic activity of 100 nM U-II (Fig. 6B). This result is thus in agreement with the observed IL-1beta-induced increase in UT receptor mRNA expression (Fig. 3).


Figure 6
View larger version (27K):
[in this window]
[in a new window]

  		FIGURE 6. U-II is a chemoattractant for human PBMC. A, Under basal conditions, U-II (1 nM-1 µM; 3 h) induced PBMC migration. B, Pretreatment of PBMC with IL-1beta (0.1 ng/ml; 12 h) potentiated U-II-mediated chemotaxis (*, p < 0.01 vs control). C, Effect of U-II (100 nM) on chemotaxis of purified monocytes, NK cells, and T lymphocytes. D, Effect of U-II (100 nM) on chemotaxis of mDC and iDC. E, Effect of U-II (100 nM) on chemotaxis of PMN. The data are the means ± SD of four independent experiments (*, p < 0.01 vs control).

 
To further define the cell population that migrated in response to U-II, we next analyzed the chemoattracting properties of U-II on purified monocytes, NK cells, and T lymphocytes (Fig. 6C). U-II (100 nM) had no effect on NK and T cells, but was efficient in monocytes, stimulating monocytes chemotaxis by up to 3-fold over control (Fig. 6C).

The chemoattracting properties of U-II have also been assessed in monocyte-derived DC. Whereas iDC did not respond, U-II strongly stimulated mDC chemotaxis (Fig. 6D), in agreement with the induction of UT receptor expression by LPS treatment. In PMN, in agreement with the low level of UT receptor expression, U-II failed to stimulate migration (Fig. 6E).

U-II induces Rho kinase-dependent actin organization

Actin polymerization and actomyosin contraction are key determinants of monocyte migration (35). Labeling of polymerized F-actin with Alexa 488-conjugated phalloidin has therefore been used to analyze the effect of U-II on actin cytoskeleton organization (Fig. 8). In rat splenocytes, U-II (10 and 100 nM) induced actin polymerization, particularly located at the cell periphery and in cytoplasmic extensions (Fig. 7A). This effect was associated with an increase in cell area. Higher U-II concentration had almost no effect on actin cytoskeleton (Fig. 7A). The RhoA-signaling pathway plays a major role in the regulation of actin polymerization and actomyosin contraction (36). RhoA regulated-actomyosin contraction involves Rho kinase-mediated MLC phosphatase phosphorylation, which leads to inhibition of its phosphatase activity and increased MLC phosphorylation (37, 38, 39). To address the role of the RhoA/Rho kinase pathway in the effect of U-II on actin cytoskeleton, we used the Rho kinase inhibitor Y-27632. In the presence of 10 µM Y-27632, the effect of U-II (10 nM) on actin cytoskeleton organization was abolished (Fig. 7B), indicating that U-II-mediated actin organization involved Rho kinase activation.


Figure 8
View larger version (21K):
[in this window]
[in a new window]

  		FIGURE 8. U-II-induced chemotaxis is mediated by the RhoA/Rho kinase pathway. A, The Rho-kinase inhibitor Y-27632 (10 µM) abolished U-II (0.1 µM)-induced chemotaxis in PBMC and in splenocytes. B, U-II (0.1 µM) stimulated RhoA activity and induced an increase in MLC phosphorylation which was blocked by Y-27632 (10 µM).

 

Figure 7
View larger version (53K):
[in this window]
[in a new window]

  		FIGURE 7. U-II induces actin cytoskeleton reorganization. A, F-actin labeling with Alexa 488-conjugated phalloidin in rat splenocytes showed that U-II (10 nM-1 µM) induced actin cytoskeleton reorganization in a concentration-dependent manner. B, The effect of U-II (10 nM) on actin cytoskeleton organization was inhibited by Y-27632 (10 µM).

 
The RhoA-Rho kinase signaling pathway is involved in U-II-induced chemotaxis

We therefore next examined whether Rho kinase activation was also involved in U-II-induced chemotaxis in human PBMC and in rat splenocytes. As shown in Fig. 8A, inhibition of Rho kinase by Y-27632 (10 µM) abolished U-II-induced chemotaxis in both types of cells indicating that it depends on Rho kinase activation. Direct analysis of RhoA activity by pull-down assay indicated that stimulation of splenocytes by U-II (10 nM) induced a 3- to 4-fold increase in the amount of active, GTP-bound RhoA (Fig. 8B). U-II stimulation also induced a rise in intracellular calcium concentration measured as a 2.6-fold increase in fura-2 fluorescence ratio (F340/F380) (data not shown). U-II-induced RhoA activation was associated with increased phosphorylation of MLC, which was blocked by Y-27632 (10 µM). Similar results were also observed with human PBMC (data not shown). These data thus indicate that U-II-induced chemotaxis is mediated by activation of the RhoA/Rho kinase pathway.


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
Disclosures
 References
 
Recent immunohistochemical studies have reported the presence of inflammatory cells expressing both U-II and UT receptor in atherosclerotic lesions of the human aorta (23). UT receptor mRNA was mainly detected in monocytes by RT-PCR, suggesting a potential immune function for U-II (23). In the present study, we show that UT receptor is up-regulated by inflammatory stimuli and identify U-II, the UT receptor ligand, as a novel chemoattractant peptide for monocytes.

Analysis of cell surface expression of UT receptor in PBMC isolated from healthy volunteers by flow cytometry indicated an important interindividual variability (15–30% of total PBMC) also observed at the level of the UT receptor mRNA. Three-color flow cytometry analysis revealed that in human peripheral blood, UT receptor is essentially expressed in monocytes and NK cells. Furthermore, we found that among CD14+ (high and low) monocytes, those expressing the CD16/Fc{gamma}RIII receptor had the higher levels of UT receptor expression. The CD14+CD16+ subset represents 10–15% of monocytes in healthy individuals, whereas the major subset CD14highCD16– accounts for the remaining (30). It has been described, in a model of transendothelial trafficking, that the CD16+ subset of human monocytes had a higher potential to become migratory DCs than CD14+CD16– monocytes (40). This population of CD14lowCD16+ monocytes, which exhibits characteristic of tissue macrophages, is found increased in several inflammatory conditions such as sepsis (30). Accordingly, it has been shown that CD14lowCD16+ cells also express TLR2 and TLR4 and secrete TNF-{alpha} and IL-6 upon stimulation with LPS (41). Of particular interest, an increased proportion of CD14lowCD16+ monocytes has been detected in the blood of patients with low levels of HDL cholesterol, a risk indicator for the development of atherosclerosis (42). Since the development atherosclerosis also appears to be associated with the hallmarks of a systemic inflammatory reaction (43), it would be interesting to verify whether the proportion of CD14lowCD16+ monocytes expressing the UT receptor could be increased in PBMC of patients at risk for atherosclerosis. Interestingly, our finding that UT receptor expression in monocytes is up-regulated by LPS could be relevant to the CD14lowCD16+monocyte population with increased levels of UT receptor surface expression. After PCR cloning and sequencing of the UT receptor gene promoter, we have identified four NF-{kappa}B-binding sites involved in LPS-induced UT receptor gene transcription. Indeed, site-directed mutagenesis revealed that two NF-{kappa}B-binding sites mediated LPS-induced stimulation of UT receptor gene promoter activity. NF-{kappa}B-dependent up-regulation of UT receptor expression in response to inflammatory stimuli supports an increased role for UT receptor and UT receptor-expressing cells in inflammatory conditions.

The structural relationship between UT and chemokine receptors prompted us to assess the potential chemotactic activity of U-II. Chemokines are a family of small proteins, which plays a critical role in immune and inflammatory reactions by directing leukocyte migration to inflammatory sites (44). Binding of chemokines to G protein-coupled receptor triggers intracellular signal pathways involved in cell migration, in particular Rho protein signaling which regulates the dynamic regulation of actin cytoskeleton organization (35, 39, 45, 46). We demonstrate for the first time that U-II is a chemoattractant for human PBMC and rat splenocytes, acting at concentrations similar to most chemokines. The inactive U-II analog that did not bind the UT receptor (24) did not display chemoattractant activity, indicating that U-II-induced chemotaxis is specific of UT receptor activation. Furthermore, we showed that U-II specifically induced the chemotaxis of CD14+ monocytes. Although a great proportion of NK cells express the U-II receptor, we did not observe cell migration of this subset in response to U-II. Thus, the function of the U-II receptor in NK cells remains to be elucidated. The stimulatory effect of U-II on proliferation previously described in vascular smooth muscle cells (20) and renal epithelial cells (14, 21) was not observed in PBMC suggesting that the mitogenic effect of U-II might involve cell-specific signal transduction pathways.

As previously shown in rat smooth muscle cells (20), we found that U-II activates the RhoA-Rho kinase pathway in rat splenocytes and that inhibition of Rho kinase by Y-27632 blocks U-II-induced actin reorganization and chemotaxis. Video-imaging analysis showed that U-II did not stimulate chemokinesis (data not shown). This is in agreement with the observation that monocyte migration, in Boyden chamber experiments, was inhibited when U-II was added both in the upper and lower chambers and suggest that U-II-induced monocyte migration was essentially due to chemotaxis (data not shown). Our data thus suggest that the RhoA-Rho kinase signaling cascade plays a major role in the chemotaxis of monocytes induced by U-II. This signaling pathway has already been shown to be responsible for the chemotaxis of PBLs induced by the chemokine stromal cell-derived factor-1 (27). The fact that UT receptor is up-regulated by proinflammatory stimuli suggests that U-II could play a role in directing leukocyte migration to inflammatory sites. In addition, the potentiation of U-II activity by IL-1beta could also be explained by a synergistic activation of RhoA-Rho kinase pathway. Indeed, we have shown that IL-1beta stimulates RhoA-Rho kinase in human PBMC and that this pathway is involved in inflammatory responses (47).

In view of the present results, it is possible that U-II, which could be released from endothelial cells and vascular smooth muscle cells (23), contributes to the recruitment of monocyte expressing the UT receptor to inflamed tissues. This is particularly relevant to the atherosclerotic lesion formation in which monocyte/macrophage emigration from blood plays a central role (43). Several studies have shown that chemokine production, in particular by vascular endothelial and smooth muscle cells, are essential to this process (43, 48, 49, 50). This is well-established for MCP-1 since it has been reported that mice deficient for MCP-1 or its receptor CCR-2 have an overall decrease in atherosclerotic lesion size associated with a reduced number of monocytes and macrophages in aortic walls (48, 49). Thus, blocking monocyte and macrophage recruitment in the vessel wall is considered a promising therapeutic approach against atherosclerosis development. The increased expression of U-II observed in endothelial and smooth muscle cells of aorta, carotid, and coronary arteries of patients with atherosclerosis might contribute to the attraction of monocyte/macrophage-expressing UT receptors to vessel walls (22, 23). In addition to U-II, other vasoactive peptides could also participate in monocyte recruitment, either directly such as endothelin (51) or indirectly, such as angiotensin II, through stimulation of MCP-1 expression in arterial smooth muscle cells (52).

Our hypothesis for a role of U-II-mediated RhoA/Rho kinase-dependent monocyte recruitment in atherosclerotic lesions is in agreement with the previously described role of the RhoA/Rho kinase pathway in the development of atherosclerotic plaques (53). We have shown that inhibition of Rho kinase significantly reduces early atherosclerotic plaque development in the low density lipoprotein receptor-deficient mice. Inhibition of Rho kinase by Y-27632 should block most of the cellular processes induced by U-II including, vasoconstriction, smooth muscle cell proliferation, and also, as we show here, chemotaxis.

In conclusion, our data suggest that U-II could attract blood monocytes expressing the UT receptor to inflammatory sites and reinforce the idea of a possible role of U-II in the pathogenesis of atherosclerosis.


   Acknowledgments
 
We thank Dr. Martin Schwartz for the gift of the plasmids encoding the rhotekin Rho-binding domain, R. Seuret and N. Vaillant for technical assistance, and P. Hullin for intracellular calcium measurements.


   Disclosures
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
References
 
The authors have no financial conflict of interest.


   Footnotes
 
The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1 This work was supported by grants from Institut National de la Recherche Agronomique, Institut National de la Santé et de la Recherche Médicale, and the Région Pays de la Loire. M.R.-D. was supported by Centre National de la Recherche Scientifique. Back

2 J.-P.S. and M.R.-D. contributed equally to this work. Back

3 Address correspondence and reprint requests to Prof. Pierre Pacaud, Institut National de la Santé et de la Recherche Médicale Unité 533 l’Institut du Thorax, Université de Nantes, Faculté des Sciences, 2 rue de la Houssinière, F-44322 Nantes, France. E-mail address: pierre.pacaud{at}univ-nantes.fr Back

4 Abbreviations used in this paper: DC, dendritic cell; U-II, urotensin-II; MLC, myosin L chain; PMN, polymorphonuclear cell; DC, dendritic cell; iDC, immature DC; mDC, mature DC; h, human; r, rat; Ct, cycle threshold; WT, wild type; MFI, mean fluorescence intensity; CAPE, caffeic acid phenethyl ester. Back

Received for publication July 19, 2006. Accepted for publication May 5, 2007.


   References
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
 Disclosures
 References
 

  1. Straub, R. H., J. Westermann, J. Scholmerich, W. Falk. 1998. Dialogue between the CNS and the immune system in lymphoid organs. Immunol. Today 19: 409-413. [Medline]
  2. Frieri, M.. 2003. Neuroimmunology and inflammation: implications for therapy of allergic and autoimmune diseases. Ann. Allergy Asthma Immunol. 90: 34-40. [Medline]
  3. Lotz, M., J. H. Vaughan, D. A. Carson. 1988. Effect of neuropeptides on production of inflammatory cytokines by human monocytes. Science 241: 1218-1221. [Abstract/Free Full Text]
  4. Makarenkova, V. P., G. V. Shurin, I. L. Tourkova, L. Balkir, G. Pirtskhalaishvili, L. Perez, V. Gerein, J. M. Siegfried, M. R. Shurin. 2003. Lung cancer-derived bombesin-like peptides down-regulate the generation and function of human dendritic cells. J. Neuroimmunol. 145: 55-67. [Medline]
  5. Delgado, M., D. Ganea. 2001. Cutting edge: is vasoactive intestinal peptide a type 2 cytokine. J. Immunol. 166: 2907-2912. [Abstract/Free Full Text]
  6. Dunzendorfer, S., A. Kaser, C. Meierhofer, H. Tilg, C. J. Wiedermann. 2001. Cutting edge: peripheral neuropeptides attract immature and arrest mature blood-derived dendritic cells. J. Immunol. 166: 2167-2172. [Abstract/Free Full Text]
  7. Serhan, C. N., I. M. Fierro, N. Chiang, M. Pouliot. 2001. Cutting edge: nociceptin stimulates neutrophil chemotaxis and recruitment: inhibition by aspirin-triggered-15-epi-lipoxin A4. J. Immunol. 166: 3650-3654. [Abstract/Free Full Text]
  8. Hokfelt, T., T. Bartfai, F. Bloom. 2003. Neuropeptides: opportunities for drug discovery. Lancet Neurol. 2: 463-472. [Medline]
  9. Maguire, J. J., A. P. Davenport. 2002. Is urotensin-II the new endothelin. Br. J. Pharmacol. 137: 579-588. [Medline]
  10. Ames, R. S., H. M. Sarau, J. K. Chambers, R. N. Willette, N. V. Aiyar, A. M. Romanic, C. S. Louden, J. J. Foley, C. F. Sauermelch, R. W. Coatney, et al 1999. Human urotensin-II is a potent vasoconstrictor and agonist for the orphan receptor GPR14. Nature 401: 282-286. [Medline]
  11. Coulouarn, Y., I. Lihrmann, S. Jegou, Y. Anouar, H. Tostivint, J. C. Beauvillain, J. M. Conlon, H. A. Bern, H. Vaudry. 1998. Cloning of the cDNA encoding the urotensin II precursor in frog and human reveals intense expression of the urotensin II gene in motoneurons of the spinal cord. Proc. Natl. Acad. Sci. USA 95: 15803-15808. [Abstract/Free Full Text]
  12. Maguire, J. J., R. E. Kuc, A. P. Davenport. 2000. Orphan-receptor ligand human urotensin II: receptor localization in human tissues and comparison of vasoconstrictor responses with endothelin-1. Br. J. Pharmacol. 131: 441-446. [Medline]
  13. Douglas, S. A., A. C. Sulpizio, V. Piercy, H. M. Sarau, R. S. Ames, N. V. Aiyar, E. H. Ohlstein, R. N. Willette. 2000. Differential vasoconstrictor activity of human urotensin-II in vascular tissue isolated from the rat, mouse, dog, pig, marmoset and cynomolgus monkey. Br. J. Pharmacol. 131: 1262-1274. [Medline]
  14. Matsushita, M., M. Shichiri, T. Imai, M. Iwashina, H. Tanaka, N. Takasu, Y. Hirata. 2001. Co-expression of urotensin II and its receptor (GPR14) in human cardiovascular and renal tissues. J. Hypertens. 19: 2185-2190. [Medline]
  15. Behm, D. J., S. M. Harrison, Z. Ao, K. Maniscalco, S. J. Pickering, E. V. Grau, T. N. Woods, R. W. Coatney, C. P. Doe, R. N. Willette, et al 2003. Deletion of the UT receptor gene results in the selective loss of urotensin-II contractile activity in aortae isolated from UT receptor knockout mice. Br. J. Pharmacol. 139: 464-472. [Medline]
  16. Bohm, F., J. Pernow. 2002. Urotensin II evokes potent vasoconstriction in humans in vivo. Br. J. Pharmacol. 135: 25-27. [Medline]
  17. Camarda, V., A. Rizzi, G. Calo, G. Gendron, S. I. Perron, E. Kostenis, P. Zamboni, F. Mascoli, D. Regoli. 2002. Effects of human urotensin II in isolated vessels of various species; comparison with other vasoactive agents. Naunyn. Schmiedebergs. Arch. Pharmacol. 365: 141-149.
  18. MacLean, M. R., D. Alexander, A. Stirrat, M. Gallagher, S. A. Douglas, E. H. Ohlstein, I. Morecroft, K. Polland. 2000. Contractile responses to human urotensin-II in rat and human pulmonary arteries: effect of endothelial factors and chronic hypoxia in the rat. Br. J. Pharmacol. 130: 201-204. [Medline]
  19. Maguire, J. J., R. E. Kuc, K. E. Wiley, M. J. Kleinz, A. P. Davenport. 2004. Cellular distribution of immunoreactive urotensin-II in human tissues with evidence of increased expression in atherosclerosis and a greater constrictor response of small compared to large coronary arteries. Peptides 25: 1767-1774. [Medline]
  20. Sauzeau, V., E. Le Mellionnec, J. Bertoglio, E. Scalbert, P. Pacaud, G. Loirand. 2001. Human urotensin II-induced contraction and arterial smooth muscle cell proliferation are mediated by RhoA and Rho-kinase. Circ. Res. 88: 1102-1104. [Abstract/Free Full Text]
  21. Watanabe, T., R. Pakala, T. Katagiri, C. R. Benedict. 2001. Synergistic effect of urotensin II with mildly oxidized LDL on DNA synthesis in vascular smooth muscle cells. Circulation 104: 16-18. [Abstract/Free Full Text]
  22. Hassan, G. S., S. A. Douglas, E. H. Ohlstein, A. Giaid. 2005. Expression of urotensin-II in human coronary atherosclerosis. Peptides 26: 2464-2472. [Medline]
  23. Bousette, N., L. Patel, S. A. Douglas, E. H. Ohlstein, A. Giaid. 2004. Increased expression of urotensin II and its cognate receptor GPR14 in atherosclerotic lesions of the human aorta. Atherosclerosis 176: 117-123. [Medline]
  24. Chatenet, D., C. Dubessy, J. Leprince, C. Boularan, L. Carlier, I. Segalas-Milazzo, L. Guilhaudis, H. Oulyadi, D. Davoust, E. Scalbert, et al 2004. Structure-activity relationships and structural conformation of a novel urotensin II-related peptide. Peptides 25: 1819-1830. [Medline]
  25. Feve, B., J. P. Segain, B. Charbonnel, P. Sai. 1990. B cell-adherent splenocytes precede the onset of diabetes in low-dose streptozotocin-treated mice. Diabetologia 33: 9-14. [Medline]
  26. Livak, K. J., T. D. Schmittgen. 2001. Analysis of relative gene expression data using real-time quantitative PCR and the 2–{Delta}{Delta}CT method. Methods 25: 402-408. [Medline]
  27. Vicente-Manzanares, M., J. R. Cabrero, M. Rey, M. Perez-Martinez, A. Ursa, K. Itoh, F. Sanchez-Madrid. 2002. A role for the Rho-p160 Rho coiled-coil kinase axis in the chemokine stromal cell-derived factor-1{alpha}-induced lymphocyte actomyosin and microtubular organization and chemotaxis. J. Immunol. 168: 400-410. [Abstract/Free Full Text]
  28. Sauzeau, V., H. Le Jeune, C. Cario-Toumaniantz, A. Smolenski, S. M. Lohmann, J. Bertoglio, P. Chardin, P. Pacaud, G. Loirand. 2000. Cyclic GMP-dependent protein kinase signaling pathway inhibits RhoA-induced Ca2+ sensitization of contraction in vascular smooth muscle. J. Biol. Chem. 275: 21722-21729. [Abstract/Free Full Text]
  29. Ren, X. D., W. B. Kiosses, M. A. Schwartz. 1999. Regulation of the small GTP-binding protein Rho by cell adhesion and the cytoskeleton. EMBO J. 18: 578-585. [Medline]
  30. Ziegler-Heitbrock, H. W.. 1996. Heterogeneity of human blood monocytes: the CD14+ CD16+ subpopulation. Immunol. Today 17: 424-428. [Medline]
  31. Lombardi, M. S., A. Kavelaars, C. J. Heijnen. 2002. Role and modulation of G protein-coupled receptor signaling in inflammatory processes. Crit. Rev. Immunol. 22: 141-163. [Medline]
  32. Scherf, M., A. Klingenhoff, T. Werner. 2000. Highly specific localization of promoter regions in large genomic sequences by PromoterInspector: a novel context analysis approach. J. Mol. Biol. 297: 599-606. [Medline]
  33. Quandt, K., K. Frech, H. Karas, E. Wingender, T. Werner. 1995. MatInd and MatInspector: new fast and versatile tools for detection of consensus matches in nucleotide sequence data. Nucleic Acids Res. 23: 4878-4884. [Abstract/Free Full Text]
  34. Cartharius, K., K. Frech, K. Grote, B. Klocke, M. Haltmeier, A. Klingenhoff, M. Frisch, M. Bayerlein, T. Werner. 2005. MatInspector and beyond: promoter analysis based on transcription factor binding sites. Bioinformatics 21: 2933-2942. [Abstract/Free Full Text]
  35. Vicente-Manzanares, M., F. Sanchez-Madrid. 2004. Role of the cytoskeleton during leukocyte responses. Nat. Rev. Immunol. 4: 110-122. [Medline]
  36. Etienne-Manneville, S., A. Hall. 2002. Rho GTPases in cell biology. Nature 420: 629-635. [Medline]
  37. Fukata, Y., M. Amano, K. Kaibuchi. 2001. Rho-Rho-kinase pathway in smooth muscle contraction and cytoskeletal reorganization of non-muscle cells. Trends Pharmacol. Sci. 22: 32-39. [Medline]
  38. Kimura, K., M. Ito, M. Amano, K. Chihara, Y. Fukata, M. Nakafuku, B. Yamamori, J. Feng, T. Nakano, K. Okawa, et al 1996. Regulation of myosin phosphatase by Rho and Rho-associated kinase (Rho-kinase). Science 273: 245-248. [Abstract]
  39. Ridley, A. J.. 2001. Rho GTPases and cell migration. J. Cell. Sci. 114: 2713-2722. [Abstract/Free Full Text]
  40. Randolph, G. J., G. Sanchez-Schmitz, R. M. Liebman, K. Schakel. 2002. The CD16+ (Fc{gamma}RIII+) subset of human monocytes preferentially becomes migratory dendritic cells in a model tissue setting. J. Exp. Med. 196: 517-527. [Abstract/Free Full Text]
  41. Skinner, N. A., C. M. MacIsaac, J. A. Hamilton, K. Visvanathan. 2005. Regulation of Toll-like receptor (TLR)2 and TLR4 on CD14dimCD16+ monocytes in response to sepsis-related antigens. Clin. Exp. Immunol. 141: 270-278. [Medline]
  42. Rothe, G., H. Gabriel, E. Kovacs, J. Klucken, J. Stohr, W. Kindermann, G. Schmitz. 1996. Peripheral blood mononuclear phagocyte subpopulations as cellular markers in hypercholesterolemia. Arterioscler. Thromb. Vasc. Biol. 16: 1437-1447. [Abstract/Free Full Text]
  43. Ross, R.. 1999. Atherosclerosis–an inflammatory disease. N. Engl. J. Med. 340: 115-126. [Free Full Text]
  44. Murphy, P. M.. 1994. The molecular biology of leukocyte chemoattractant receptors. Annu. Rev. Immunol. 12: 593-633. [Medline]
  45. Allen, W. E., D. Zicha, A. J. Ridley, G. E. Jones. 1998. A role for Cdc42 in macrophage chemotaxis. J. Cell. Biol. 141: 1147-1157. [Abstract/Free Full Text]
  46. Honing, H., T. K. van den Berg, S. M. van der Pol, C. D. Dijkstra, R. A. van der Kammen, J. G. Collard, H. E. de Vries. 2004. RhoA activation promotes transendothelial migration of monocytes via ROCK. J. Leukocyte Biol. 75: 523-528. [Abstract/Free Full Text]
  47. Segain, J. P., D. Raingeard de la Bletiere, V. Sauzeau, A. Bourreille, G. Hilaret, C. Cario-Toumaniantz, P. Pacaud, J. P. Galmiche, G. Loirand. 2003. Rho kinase blockade prevents inflammation via nuclear factor {kappa}B inhibition: evidence in Crohn’s disease and experimental colitis. Gastroenterology 124: 1180-1187. [Medline]
  48. Boring, L., J. Gosling, M. Cleary, I. F. Charo. 1998. Decreased lesion formation in CCR2–/– mice reveals a role for chemokines in the initiation of atherosclerosis. Nature 394: 894-897. [Medline]
  49. Gu, L., Y. Okada, S. K. Clinton, C. Gerard, G. K. Sukhova, P. Libby, B. J. Rollins. 1998. Absence of monocyte chemoattractant protein-1 reduces atherosclerosis in low density lipoprotein receptor-deficient mice. Mol. Cell. 2: 275-281. [Medline]
  50. Bjorkbacka, H., V. V. Kunjathoor, K. J. Moore, S. Koehn, C. M. Ordija, M. A. Lee, T. Means, K. Halmen, A. D. Luster, D. T. Golenbock, M. W. Freeman. 2004. Reduced atherosclerosis in MyD88-null mice links elevated serum cholesterol levels to activation of innate immunity signaling pathways. Nat. Med. 10: 416-421. [Medline]
  51. Grimshaw, M. J., J. L. Wilson, F. R. Balkwill. 2002. Endothelin-2 is a macrophage chemoattractant: implications for macrophage distribution in tumors. Eur. J. Immunol. 32: 2393-2400. [Medline]
  52. Funakoshi, Y., T. Ichiki, H. Shimokawa, K. Egashira, K. Takeda, K. Kaibuchi, M. Takeya, T. Yoshimura, A. Takeshita. 2001. Rho-kinase mediates angiotensin II-induced monocyte chemoattractant protein-1 expression in rat vascular smooth muscle cells. Hypertension 38: 100-104. [Abstract/Free Full Text]
  53. Mallat, Z., A. Gojova, V. Sauzeau, V. Brun, J. S. Silvestre, B. Esposito, R. Merval, H. Groux, G. Loirand, A. Tedgui. 2003. Rho-associated protein kinase contributes to early atherosclerotic lesion formation in mice. Circ. Res. 93: 884-888. [Abstract/Free Full Text]




This Article
Right arrow Abstract Freely available
Right arrow Full Text (PDF)
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Right arrow Citation Map
Services
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 Download to citation manager
Right arrow Request Permissions
Citing Articles
Right arrow Citing Articles via Google Scholar
Google Scholar
Right arrow Articles by Segain, J.-P.
Right arrow Articles by Pacaud, P.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Segain, J.-P.
Right arrow Articles by Pacaud, P.

HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS