Melanocortin-1 Receptor Positively Regulates Human Artery Endothelial Cell Migration
Federica Saporitia Luca Piacentinia Valentina Alfieria,b Elisa Bonoa
Fabrizio Ferraria Mattia Chiesaa Gualtiero I. Colomboa
aUnit of Immunology and Functional Genomics, Centro Cardiologico Monzino IRCCS, Milan, Italy, bDepartment of Pharmacological and Biomolecular Sciences, Università degli Studi di Milano, Milan, Italy
Key Words
Melanocortin receptors • α-MSH • Human artery endothelial cells • Cell migration
Abstract
Background/Aims: Melanocortin receptors (MCRs) belong to a hormonal signalling pathway with multiple homeostatic and protective actions. Microvascular and umbilical vein endothelial cells (ECs) express components of the melanocortin system, including the type 1 receptor (MC1R), playing a role in modulating inflammation and vascular tone. Since ECs exhibit a remarkable heterogeneity, we investigated whether human artery ECs express any functional MCR and whether its activation affects cell migration. Methods: We used reverse transcription real-time PCR to examine the expression of melanocortin system components in primary human artery ECs. We assessed MC1R protein expression and activity by western blot, immunohistochemistry, cAMP production, and intracellular Ca2+ mobilization assays. We performed gap closure and scratch tests to examine cell migration after stimulation with alpha-melanocyte-stimulating hormone (α-MSH), the receptor highest-affinity natural ligand. We assessed differential time-dependent transcriptional changes in migrating cells by microarray analysis. Results: We showed that human aortic ECs (HAoECs) express a functionally active MC1R. Unlike microvascular ECs, arterial cells did not express the α-MSH precursor proopiomelanocortin, nor produced the hormone. MC1R engagement with a single pulse of α-MSH accelerated HAoEC migration both in the directional migration assay and in the scratch wound healing test. This was associated with an enhancement in Ca2+ signalling and inhibition of cAMP elevation. Time-course genome-wide expression analysis in HAoECs undergoing directional migration allowed identifying dynamic co-regulation of genes involved in extracellular matrix-receptor interaction, vesicle-mediated trafficking, and metal sensing – which have all well-established influences on EC motility –, without affecting the balance between pro- and anticoagulant genes. Conclusion: Our work broadens the knowledge on peripherally expressed MC1R. These results indicate that the receptor is constitutively expressed by arterial ECs and provide evidence of a novel homeostatic function for MC1R, whose activation may participate in preventing/healing endothelial dysfunction or denudation in macrovascular arteries.
Introduction
The melanocortin receptors (MCRs) are a family of rhodopsin-like G protein-coupled receptors (GPCRs) that are activated by different melanocortin peptide ligands, derived from the tissue-specific cleavage of a common preprohormone precursor, the proopiomelanocortin (POMC) [1]. These molecules, together with a number of endogenous antagonists and accessory proteins, constitutes the so-called melanocortin system [2]. To date, five MCRs have been identified, with different tissue distribution and a diverse affinity for their natural ligands. MCRs mainly signal through intracellular cAMP increase or, alternatively, transient intracellular elevation of cytosolic free Ca2+ [3]. The melanocortin system has been studied for its ability to regulate several physiological processes, including pigmentation, adrenocortical steroidogenesis, energy homeostasis, and exocrine gland secretion. In particular, the prototypical melanocortin peptide, the alpha-melanocyte stimulating hormone (α-MSH), possesses a wide spectrum of anti-inflammatory [4], immunoregulatory [5], and cytoprotective activities, including protection and repair after organ damage (i.e. cerebral and myocardial ischemia/reperfusion injury, nephrotoxicity, and acute lung injury) [6]. As a consequence, targeting melanocortin system is considered a promising strategy for new therapeutic approaches in various inflammatory conditions [7].
The melanocortin system has been involved in the modulation of oxidative stress [8] and vascular endothelial damage [9]. A local melanocortin system has been described in endothelial cells (ECs) of the cutaneous microcirculation [10]. Moreover, the MC1R (and no other MCR) has been detected both on murine brain microvascular ECs [11], and on human dermal microvascular ECs (HDMECs) [10, 12] and umbilical vein ECs (HUVECs) [13], with possible modulatory effects on endothelium homeostasis. In particular, α-MSH has been shown to modulate blood vessel tone by enhancing nitric oxide-cyclic guanosine monophosphate dependent relaxation responses through endothelial MC1R [13]. Nonetheless, a formal demonstration that human artery ECs of the macrovasculature express functional MCR(s) is currently missing. This is substantial because ECs exhibit a remarkable heterogeneity and show specific structure and functions associated with the blood vessel they belong to, i.e. large and medium arteries, veins, or capillaries [14, 15]. At the molecular level, ECs display phenotype markers that are cell type-restricted, and exhaustive genome-wide expression studies have shown unique gene expression patterns in ECs derived from different tissues [16, 17]. This heterogeneity accounts for many human vascular diseases restricted to specific types of vessels. Nevertheless, our knowledge of EC biology has been mostly inferred by studies on HUVECs, which are cells that originate from a vessel type that is rarely affected by vascular disorders [18]. HDMECs and HUVECs do not recapitulate the physiology of all the vascular ECs and, most importantly, their ability to activate specific cell functions in response to MCR ligands may not overlap those of artery ECs.
A recent report showed that treatment with MCR agonists was able to prevent the development of vascular dysfunction and attenuate plaque inflammation in a mouse model of pre-established atherosclerosis [19]. Artery endothelial dysfunction and/or injury are prominently linked to the pathogenesis of atherosclerosis, thrombosis, or surgery procedure complications [20]. An essential biological process involved in endothelial healing upon vascular injury is EC migration. When a blood vessel is damaged, the restoration of endothelium and vessel integrity is achieved through migration of healthy ECs to the site of the lesion and subsequent proliferation. Hence, EC migration has a key role, besides angiogenesis, in vascular repair and tissue regeneration [21]. In this work, we investigated whether human artery ECs express any functional MCR and whether MCRs activation through α-MSH can affect artery EC migration.
Materials and Methods
Primary human artery endothelial cells
We purchased primary human artery ECs from the European Collection of Authenticated Cell Cultures (ECACC, Salisbury, UK), Lonza (Allendale, NJ), and Promocell (Heidelberg, Germany). We obtained three adult human aortic ECs (HAoECs) and recoded them as c1, c2 and c3, namely: (c1) HAoEC (304-05a) from ECACC (catalogue no. 06090729), (c2) HAoEC from Lonza (catalogue no. CC-2535), and (c3) HAoEC from Promocell (catalogue no. C-12271). We also obtained three adult human coronary artery ECs (HCAECs) and recoded them as c4, c5 and c6, namely: (c4) HCAEC (300-05a) from ECACC (catalogue no. 06090727), (c5) HCAEC from Lonza (catalogue no. CC-2585), and (c6) HCAEC from Promocell (catalogue no. C-12221). Primary ECs were tested for cell-type specific markers by the manufactures. Cells were positive for Factor VIII-related antigen or von Willebrand factor and CD31 expression, positive for acetylated low-density lipoprotein uptake, and negative for α-actin expression. Cells were seeded in 75 mL plastic flasks (Corning, Tewksbury, MA) at a density of 2.5 × 103 cells/cm2 and cultured following manufactures’ instructions. We performed all experiments at cell passages 4–8. We tested cell cultures for mycoplasma contamination before any experiments, using the PCR-based Mycoplasma detection kit Venor GeM OneStep (Minerva Biolabs, Berlin, Germany).
Chemicals
The α-MSH peptide was obtained from Phoenix Pharmaceuticals (Burlingame, CA); the peptide 153N-6 (H-[Met5,Pro6,D-Phe7,D-Trp9,Phe10]-MSH(5-13)) from Bachem (Bubendorf, Switzerland); isobutyl methylxanthine (IBMX), forskolin, PD0332991 isethionate, and thapsigargin from Sigma-Aldrich (St. Louis, MO); 1, 2-Bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid tetrakis(acetoxymethyl ester) (BAPTA-AM) and 1-[6-[[(17β)-3-Methoxyestra-1, 3,5(10)-trien-17-yl]amino]hexyl]-1H-pyrrole-2, 5-dione (U-73122) from Tocris Bioscience (Bristol, UK). The peptides α-MSH and 153N-6 were dissolved in water. IBMX, forskolin, PD0332991, thapsigargin, BAPTA-AM, and U-73122 were dissolved in dimethyl sulfoxide (DMSO).
Reverse transcription quantitative PCR (RT-qPCR) for melanocortin system components
Total RNA was extracted from ECs grown to confluence, adding TRIzol Reagent (Invitrogen, Carlsbad, CA) directly to the culture dishes. Given that some MCRs are single-exon intronless genes (i.e., MC3R and MC4R), while others are multi-exon genes with several splice variants (e.g., MC1R [22]), we treated RNA samples with RNase-free DNase-I to eliminate genomic contamination and prevent amplification of genomic DNA. This allowed us to use a single-exon probe qPCR design to detect the canonical primary transcripts of the MCR genes. RNA quantification and purity assessment were performed by micro-volume spectrophotometry on an Infinite M200 PRO multimode microplate reader (Tecan, Männedorf, Switzerland). RNA quality and integrity were checked by microfluidics electrophoresis with the RNA 6000 Nano Assay Kit on a 2100 Bioanalyzer (Agilent Technologies, Santa Clara, CA). Complementary DNA (cDNA) for single target gene expression analysis was synthesized from 2 μg of total RNA for each sample using the High Capacity cDNA Reverse Transcription Kit (Applied Biosystems, Foster City, CA). TaqMan Array Human Endogenous Controls 96-Well Plate PCR assay (Applied Biosystems) was preliminarily employed to identify the most appropriate endogenous control gene. Analysis of gene expression stability and selection of the best reference gene was performed using the NormFinder v0.953 Excel Add-In [23]. We used single tube TaqMan Gene Expression Assays (Applied Biosystems) for evaluating mRNA expression of the melanocortin receptors (MCRs), proopiomelanocortin (POMC), prohormone convertases, and the endogenous constitutive gene (see details Supplemental Methods – for all supplemental material see www.cellphysiolbiochem.com). Assays IDs for the melanocortin system components and the reference gene were Hs00267168_s1 (MC1R), Hs00265039_s1 (MC2R), Hs00252036_s1 (MC3R), Hs00271877_s1 (MC4R), Hs00271882_s1 (MC5R), Hs01596743_m1 (POMC), Hs01026107_m1 (proprotein convertase subtilisin/kexin type 1, PCSK1), Hs01037347_m1 (PCSK2), Hs00159829_m1 (furin, PCSK3), Hs00159844_m1 (PCSK6), Hs00161638_m1 (secretogranin V, SCG5), and Hs99999902_m1 (ribosomal protein large P0, RPLP0). We run three replicates of each assay for each sample (20 ng/well of cDNA) on a ViiA 7 Real-time PCR System (Applied Biosystems). Experimental threshold and baseline were imputed by algorithms implemented in the ViiA 7 software v1.2 (Applied Biosystems), and data were analysed by the Pfaffl’s corrected ΔΔCt method [24].
α-MSH assay
Quantification of α-MSH release by primary HAoECs and HCAECs was performed using an ultrasensitive fluorescent enzyme immunoassay (EIA) kit (Phoenix Pharmaceuticals), following manufacture’s instruction. The EIA sensitivity, i.e. the minimum detectable concentration, was 8.9 pg/mL. Cross-reactivity with the adrenocorticotropic hormone (ACTH) was zero: α-MSH shares the sequence of ACTH (1–13), but α-MSH is acetylated at the N-terminus and amidated at the C-terminus [7]. Cells were seeded to confluence in 96-well culture plates, in complete endothelial growth medium (EGM2; Lonza), and supernatants were collected and stored at -80 °C until measurement.
Genomic DNA sequencing for MC1R
We used a 3500 Genetic Analyzer (Applied Biosystems) to perform DNA sequencing of the MC1R gene open reading frame (ORF) for all HCAECs and HAoECs. Genomic DNA amplicons of the MC1R ORF were produced by PCR with the following primers: MC1R_Forward(1) (-25) 5’-TCCTTCCTGCTTCCTGGACA-3’, MC1R_Reverse(1) (+980) 5’-CACACTTAAAGCCGCGTGCAC-3’. The amplified fragments were purified using the Agencourt AMPure XP kit (Beckman Coulter). Sequencing reactions were carried out using the BigDye Terminator v3.1 Kit (Applied Biosystems) in both strand directions to allow the production of four overlapping fragments. Sequencing primers used were the MC1R_Forward(1), the MC1R_Reverse(1) and: the inner MC1R_Forward(2) (+449) 5’-TGCGCTACCACAGCATCGTG-3’, and the inner MC1R_Reverse(2) (+510) 5’-CACCCAGATGGCCGCAAC-3’. Unincorporated fluorescent dideoxynucleotides and salts were removed with the BigDye XTerminator Purification Kit (Applied Biosystems). The purified sequencing reaction products were electrokinetically injected into a 50 cm Capillary Array filled with the POP-7 Polymer (Applied Biosystems). Electropherograms were analysed by the Variant Reporter software v1.1 (Applied Biosystems).
Antibodies
Primary antibodies used were: anti-MC1R rabbit polyclonal antibody, supplied with the specific control peptide antigen (Alomone Labs, Jerusalem, Israel); anti-ATPase Na+/K+ transporting subunit alpha 1 (ATP1A1) rabbit polyclonal antibody (Cell Signaling Technology, Danvers, MA); anti-β-actin mouse monoclonal IgG1 (Novus Biologicals, Littleton, CO); and anti-Ki67 rabbit polyclonal IgG (Abcam, Cambridge, UK). Secondary antibodies were: donkey anti-rabbit or anti-mouse IgG conjugated, respectively, to IRDye 800CW and IRDye 680RD infrared dyes (LI-COR Biosciences, Lincoln, NE), for immunoblotting; donkey anti-rabbit IgG conjugated to the DyLight 488 fluorochrome (Jackson ImmunoResearch Laboratories, West Grove, PA), for immunocytochemistry.
Western blotting
HAoECs and HCAECs (1.2 × 106) were lysed in Milliplex MAP Lysis buffer (Millipore, Billerica, MA) with a complete protease inhibitor cocktail (Roche, Mannheim, Germany) to obtain whole extracts, or with the FractionPREP Cell Fractionation Kit (BioVision, Milpitas, CA) to obtain plasma membrane extracts. Proteins were quantified using the Pierce BCA Protein Assay Kit (Thermo Fisher Scientific). Thirty μg of each protein extract were mixed with the Novex Tris-Glycine SDS sample buffer 2× and the Novex sample reducing agent 10× (Invitrogen). Samples were loaded onto 4-12% gradient Novex WedgeWell precast Tris-Glycine polyacrylamide gels (Invitrogen) and run in Novex Tris-Glycine SDS running buffer for 40 min at 200 V. Samples were blotted on nitrocellulose membranes using an iBlot system (Invitrogen). Membranes were blocked in the Odyssey blocking buffer (LI-COR Biosciences) for 1 h. Pre-absorption was performed by incubating the anti-MC1R antibody for 30 min at room temperature with the inhibitory MC1R peptide (two-fold excess of the peptide by weight). Primary or the pre-absorbed antibodies were diluted (1:1000) in the Odyssey blocking buffer (LI-COR Biosciences), and membranes were incubated overnight at 4°C. Anti-β-actin and anti-ATP1A1 antibodies (1:5000) were used as reference controls for whole or membrane extracts, respectively. Membranes were incubated with IRDye secondary antibodies (1:10000) for 20 min at room temperature. Immunoreactive bands were detected by an Odyssey Infrared Imaging System (LI-COR Biosciences).
Anti-MC1R antibody specificity testing
The anti-MC1R antibody was raised against an epitope corresponding to amino acid residues 217-232 in the 3rd intracellular loop of human MC1R. To test the specific binding of the anti-MC1R antibody to the MC1R protein, we generated a positive control for subsequent analyses by transiently transfecting human HEK293 cells with the MC1R full-length cDNA by a C-terminal fusion of tGFP tag in a pCMV6 vector (Origene, Rockville, MD). Cells were grown in RPMI 1640 with 10% foetal bovine serum (FBS), penicillin 100 U/mL and streptomycin 10 μg/mL (Sigma-Aldrich) to approximately 50% confluence, and then transfected by incubation with the TransIT-LT1 Transfection Reagent (Mirus Bio, Madison, WI) for 48 h. Specificity of the MC1R antibody was demonstrated by pre-absorption with the specific blocking peptide supplied with the primary antibody, which abolished MC1R signal in Western immunoblot (see Supplementary Fig. S1).
Intracellular cAMP assay
Quantification of intracellular cAMP levels was performed using the cAMP Biotrak enzyme immunoassay (GE Healthcare Life Sciences, Piscataway, NJ). HAoECs were seeded to confluence in 96-well plates or 24-well plates with IBIDI culture inserts (Martinsried, Germany). Cells, prior to 5-min stimulation with α-MSH 10-8 M, were pre-treated for 30 min with IBMX 0.1 mM, to inhibit cAMP degradation by phosphodiesterases (PDEs). Cells treated with IBMX alone were used as negative controls, whereas cells stimulated with the activator of eukaryotic adenylyl cyclase forskolin (10 μM) served as positive controls. As control for receptor-binding specificity, cells were pre-treated with the MC1R-selective competitive α-MSH antagonist 153N-6 (10-5 M) [25, 26] for 15 min in separate experiments. The abovementioned concentration of α-MSH was selected for all functional assays based on previous publications on HDMECs [10, 12] and on pilot experiments with 100-fold scalar concentrations of peptide (10-6 M, 10-8 M, and 10-10 M) that showed its effectiveness (see Results below).
Immunohistochemistry for MC1R
Formaldehyde-fixed paraffin sections of a normal human aorta were incubated with the primary anti-MC1R antibody overnight at 4°C. As control of the staining specificity, the anti-MC1R antibody was pre-incubated 30 min with its specific blocking peptide. Slides were incubated with a biotinylated goat anti-rabbit IgG secondary antibody (1:200; Vector Laboratories, Burlingame, CA) and signals were revealed using the VECTASTAIN Elite ABC-HRP kit combined with the ImmPACT DAB EqV peroxidase (HRP) substrate (Vector Laboratories). Images were recorded using an AxioSkop microscope equipped with an AxioCam camera (Carl Zeiss).
Directional cell migration assay
The live-cell staining, lipophilic, near-infrared fluorescent membrane probe 1, 1'-dioctadecyl-3, 3,3',3'-tetramethylindotricarbocyanine iodide (DiR) was used for imaging of gap closure in a cell migration assay [27]. HAoECs were treated with a solution of 2.5 μM DiR (Biotium, Hayward, CA) in complete EGM2 medium for 20 min at 37°C, washed and seeded onto 24-well plates with culture inserts (IBIDI, Martinsried, Germany). Inserts were removed to create a cell-free gap of approximately 500 µm, and HAoECs were allowed to migrate for 12h at 37°C and 5% CO2 in the presence of 10-8 M α-MSH or in medium alone. As control for receptor-binding specificity, cells were pre-treated with the MC1R-selective antagonist 153N-6 (10-5 M) for 15 min. In addition, to dissect the calcium-dependency of the α-MSH-induced cell migration, experiments were repeated pre-treating cells for 15 min with either the intracellular Ca2+ chelator BAPTA-AM (10-5 M) or the phospholipase C (PLC) inhibitor U-73122 (5 × 10-5 M). Plates were scanned with the Odyssey imaging system (LI-COR Biosciences) at 0, 3, 9, and 12 h, at 84 μm resolution and high quality setting (emission, 800 nm). Scans were converted to 8-bit images and analysed with the NIH ImageJ software v1.38x. For time-course gene-expression analysis, 2 × 104 HAoECs were plated in high 35-mm dishes with culture inserts (IBIDI) and treated with α-MSH 10-8 M.
Scratch wound healing assay
HAoEC migration was also assessed using a scratch migration assay. Briefly, HAoECs were seeded onto 6-well tissue culture plates at a density of 2.5 × 103 cells/cm2 and grown to confluence. A gap of approximately 1 mm was created in the adherent layer of confluent ECs by using a sterile 0.1-mL pipette tip. After treatment with medium alone or α-MSH 10-8 M, with or without 15-min pre-treatment with the MC1R antagonist 153N-6 (10-5 M), the closure extent of the cell-free gap was detected by confocal microscope imaging (Zeiss, Jena, Germany) at 6 and 24h and measured using the NIH ImageJ software v1.38x.
Immunostaining for Ki67
HAoECs were plated in 8-chamber μ-slides (IBIDI) at a density of 1.0 × 103 cells/cm2, treated with medium alone, α-MSH 10-8 M, or α-MSH plus 153N-6, and allowed to migrate for 24 h. Cells were then fixed for 20 min in 4% paraformaldehyde solution in PBS and permeabilised with 0.1% Triton X-100 (Sigma-Aldrich). Non-specific antibody binding was prevented by using a blocking solution of 10% normal donkey serum (Jackson ImmunoResearch Laboratories) for 1h. Cells were incubated with the anti-Ki-67 primary antibody (1:100) overnight at 4°C and, then, with the DyLight-conjugated species-specific secondary antibody (1:500) for 2h at room temperature. Slides were finally incubated with DAPI (Sigma-Aldrich; 1:1000) for 5 min to stain cell nuclei, mounted in a fluorescence mounting medium (Dako, Glostrup, Denmark), and examined with an ApoTome fluorescence microscope (Carl Zeiss, Jena, Germany). Images were acquired using the ZEN software v.5.0 SP1.1 (Carl Zeiss) and analysed with the ImageJ software, counting the percentage of Ki-67 positive cells over the total number of nuclei in 10 different fields for each treatment conditions in 4 independent experiments.
Cell morphology assessment
HAoECs were plated in 8-chamber μ-slides (IBIDI) at a density of 1.0 × 103 cells/cm2, incubated with α-MSH 10-8 M or medium alone for 6h, fixed for 10 min in 4% paraformaldehyde solution, and permeabilised with 0.1% Triton X-100 for 1h. Non-specific binding was prevented using a blocking solution of 5% bovine serum albumin. Cells were stained for 1h at room temperature with phalloidin, a high-affinity probe for polymeric F-actin, conjugated to the red-orange fluorescent dye tetramethylrhodamine B isothiocyanate (TRITC) (Sigma-Aldrich). Slides were then stained with DAPI and images were acquired with an ApoTome fluorescence microscope (Carl Zeiss). Images were then analysed using the ZEN software and cell shape and stress fibres alignment were assessed. Changes in cell morphology were assessed by the ImageJ software measuring the major and minor cellular axis. Cells with axial ratios (long axis/short axis) larger than 3 were counted in randomly selected fields in 3 separate experiments and expressed as percentages of the total cells counted (250 cells on average).
Intracellular Ca2+ mobilization assay
Intracellular Ca2+ levels were measured using the Fluo-4 NW Calcium Assay Kit (Invitrogen). HAoECs, seeded onto a 24-well plate with IBIDI culture inserts in a calcium free medium, were loaded with 400 μL of Fluo-4 NW for 30 min at 37°C and 5% CO2. Fluorescence was measured for 300 sec after treatment with α-MSH 10-8 M using the Infinite M200 PRO plate reader (excitation, 494 nm; emission, 516 nm). Thapsigargin (10-8 M), an inhibitor of sarco/endoplasmic reticulum Ca2+-ATPases that causes a rapid raise of cytosolic Ca2+ by depleting endoplasmic reticulum stores [28], was used as positive control for Ca2+i release, treating HAoECs for 120 sec before stimulation with α-MSH (10-8 M). MC1R specific activation was assessed pre-treating cells with the MC1R-selective antagonist 153N-6 (10-5 M) for 15 min. Ca2+i changes were calculated as the difference between the area under the curve (AUC) before (resting levels) and after addition of stimuli.
Time-course gene expression analysis by microarray
To isolate RNA from cells undergoing directional migration assay, we used the Agencourt RNAdvance cell v2 kit (Beckman Coulter, Beverly, MA), following manufacturer’s instructions. RNA extracted from migrating HAoECs at 0.5, 3, 6, and 12h was used for microarray analysis. Labelled, linearly amplified complementary RNA (cRNA) was generated by Illumina Total Prep RNA Amplification Kit (Life Technologies), according to manufacturer’s manual. Briefly, 200 ng of total RNA was reverse-transcribed to cDNA using an oligo(dT) primer containing a T7 promoter sequence. Second-strand cDNA was subsequently synthesized, and then in vitro transcribed adding biotin-dNTPs. After column-based purification and ammonium acetate/ethanol precipitation, cRNA was quantified by the Infinite M200 PRO plate reader. cRNA profile of all samples was checked by the RNA 6000 Nano Assay kit in an Agilent 2100 Bioanalyzer. cRNA (750 ng per sample) was hybridized at 58°C for 18h on HumanHT-12 v4 Expression BeadChips (Illumina, San Diego, CA), followed by detection signal reaction with the fluorolink streptavidin-Cy3 (GE Healthcare Life Sciences) as recommended by manufacturer’s instructions. Each array on the BeadChips was scanned using an iSCAN System (Illumina). Array data export and quality control analysis were performed with the GenomeStudio Software v2011.1 (Illumina). Pre-processing of raw data was done by importing and analysing them with the lumi package [29], in the R software environment v2.15.2. Data variance stabilization was performed by variance stabilizing transformation (VST). Transformed data were normalized by robust spline normalization (RSN) algorithm, which combines the features of quantile and loess normalization. For subsequent analysis, we retained probes with a detection p-value < 0.01 in at least 10% samples.
Raw and normalized, MIAME compliant microarray data are available in the NCBI’s GEO repository under the accession number GSE49348 (http://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc=GSE49348).
Microarray data were validated investigating mRNA expression of 84 wound healing related genes at different time points by RT-qPCR, using the Human Wound Healing RT² Profiler PCR Arrays (Qiagen Sciences, Frederick, MD) following manufacturers’ recommendations. The concordance of microarray hybridization intensities (log2 transformed) with PCR data (Ct) was measured computing the Pearson correlation coefficient and assessing its statistical significance.
Statistical analysis and bioinformatics
Data from functional assays passed the Shapiro-Wilk test for normality. Differences among groups were assessed by one-way ANOVA followed by Tukey’s multiple comparisons test, or two-way ANOVA followed by Bonferroni post-hoc test, as appropriate. P-values<0.05 were considered statistically significant.
To analyse the time-course microarray gene-expression experiments, we used the Short Time-series Expression Miner (STEM) v1.3.8 algorithm [30] (see online suppl. material, Table S1 for analysis parameters). STEM implements a clustering method that lines up two steps. First, it selects a set of unique representative temporal profiles that, independently of the data, cover every possible expression profiles that can be generated in the experiment for a given set of parameters; second, it assigns to one of these temporal profiles only those gene profiles that pass the filtering criteria, as determined by a correlation coefficient. A permutation test was used to identify which profiles had a statistically significant enriched number of genes (for a false discovery rate, FDR < 0.05), and significant profiles were grouped into larger clusters by their correlation degree (≥ 0.7). As data input for STEM we used the log2 ratio of the α-MSH-treated to non-treated HAoEC gene-expression values at each time point. Data were from three independent experiments, and each input derived from the average expression values of two technical hybridization replicates. The extent of the regulation is calculated as the maximum-to-minimum fold-change showed by the gene during the time-course.
Hierarchical clustering was performed using the GENE-E software v3.0.206 (https://software.broadinstitute.org/GENE-E). Unsupervised analysis of the average gene expression ratios of α-MSH stimulated cells to non-stimulated cells, at the four time points considered, was performed using one minus Pearson correlation distance and the average linkage method.
Analysis of functional relations among regulated genes was made using the DAVID Bioinformatics Resources v6.7 (http://david.abcc.ncifcrf.gov/home.jsp) [31], testing for multiple annotations, i.e. Gene Ontology (GO) terms, KEGG pathways, and the Swiss Prot (SP)-Protein Information Resource (PIR) keywords. Redundant GO terms were removed using the web-based tool REViGO [32]. A network map of the enrichment analysis was produced by the Cytoscape program v2.8.2 [33], using the Enrichment Map app [34], a network-based visualization method for gene-set enrichment results.
Results
Human macrovascular endothelial cells constitutively express a functional MC1R, but not POMC
To determine which elements of the melanocortin system are expressed in human ECs from large vessels, we first performed real-time PCR analysis for detecting specific mRNAs in six human primary cells grown to confluence, i.e. three aortic ECs (HAoECs) and three coronary artery ECs (HCAECs). All the macrovascular ECs clearly expressed MC1R, but no other known MCRs (Table 1). Cell lines expressed the receptor mRNA at comparable levels, with the exception of one HCAEC which showed levels twice as high as the other HCAECs. At variance with human dermal microvascular EC [10], POMC was undetectable in HAoECs and HCAECs. We detected the prohormone convertases (PCSK1, PCSK6, FURIN, and SCG5) [35] that process POMC into most of the derived peptides, but not PCSK2, which is needed to produce α-MSH. Consistently, α-MSH was undetectable in culture supernatants of all six macrovascular ECs (see Supplementary Fig. S2).
To verify whether the three HAoECs express the MC1R protein, we performed immunoblot analysis. Specific immunoreactive bands, corresponding to the molecular weight of the canonical fully active receptor [36], were detected both in total cell lysates (see Supplementary Fig. S3A) and in membrane extracts (Fig. 1A and S3A), showing that the MC1R receptor was expressed on the plasma membrane of the HAoECs. Bands consistent with the MC1R dimeric forms were also detected (see Supplementary Fig. S1 and S3A). The level of expression of the MC1R monomer in total cell lysates was very similar among the three primary cell lines, whereas the HAoEC no. c3 appeared to express half the quantity of the other two lines (see Supplementary Fig. S3B and S3C). Finally, the canonical monomeric form of the receptor was detected in all three HCAECs as well (see Supplementary Fig. S4).
Given that MC1R is a highly polymorphic gene and that many variants are known to affect its signal transduction [37], we sequenced the MC1R ORF of the six primary human artery ECs to identify and exclude from subsequent functional analysis those cells with gene variants that may interfere with the cellular response to MC1R ligands. All the three HCAECs bear a variant allele, whereas two HAoECs did not present any polymorphism (see online suppl. material, Table S2). The variant alleles found have been associated with a decrease in cAMP production in response to α-MSH stimulation [38-40]. For functional testing, we elected to use the HAoEC no. c2, which carried the wild-type receptor, due to its shorter doubling time.
We measured the changes in intracellular cAMP levels after treatment with α-MSH, to test whether confluent HAoECs express a functionally active MC1R. Indeed, 5-min stimulation with α-MSH 10-8 M induced a significant increase of intracellular cAMP in cells grown to confluence (Fig. 1B); cAMP elevation occurred in a concentration-dependent manner, showing a typical inverted U-shaped dose-response curve (see Supplementary Fig. S5A) [41, 42]. Co-incubation with the receptor antagonist 153N-6 (10-5 M) abolished the elevation of cAMP, indicating that MC1R is specifically activated by α-MSH (Fig. 1B).
Finally, to test whether HAoECs express MC1R in vivo, we performed immunohistochemistry staining for the receptor in formaldehyde-fixed paraffin sections of a normal human aorta. We observed a positive staining of endothelial cells, confirming the in vitro observations (Fig. 1C).
α-MSH promotes migration of HAoEC via MC1R activation
To determine whether MC1R activation has any influence on HAoEC migration and/or proliferation, we used a directional cell migration assay. Stimulation with α-MSH 10-8 M enhanced HAoEC migration (Fig. 2A), and this effect too occurred in a concentration-dependent manner (see Supplementary Fig. S5B): in comparison with cells cultured in growth medium only, migration speed appeared to accelerate after 3h of treatment and became significantly higher at 9 and 12h in α-MSH-treated cells. Consistently, concomitant use of 153N-6 10-5 M was able to abolish the pro-migratory effect of α-MSH, whereas treatment with 153N-6 alone did not alter EC migration speed (also see representative images in Supplementary Fig. S6). Gap closure assays were then performed in the presence of the proliferation inhibitor PD0332991: as expected, blocking cell proliferation increased gap closure time, but the higher speed in α-MSH-treated cells confirmed the enhancement in cell migration after MC1R activation, which was still significant at 9 and 12h (Fig. 2B and representative images in Supplementary Fig. S7A). To prove the generalizability of the pro-migratory effect of MC1R stimulation on macrovascular ECs, we showed that treatment with α-MSH 10-8 M significantly enhanced cell migration also in the other HAoEC line (no. c1) bearing the wild-type receptor, although at a lower speed (Fig. 2C and Supplementary Fig. S7B). Finally, to ascertain whether the observed responses were specifically dependent on the MC1R receptor subtype, we performed the same directional cell migration assay with the HAoEC line that was found to carry a loss-of-function allele in the MC1R gene (no. c3): interestingly, these cells showed an attenuated response to α-MSH (10-8 M), with a slight non-significant acceleration in cell migration (Fig. 2D and Supplementary Fig. S7C). Of note, accelerated HAoEC migration upon activation of MC1R was confirmed in in vitro scratch wound healing assays (see Supplementary Fig. S8A and S8B): again, pre-treatment with the receptor antagonist 153N-6 (10-5 M) abolished the effect. Conversely, there was no clear-cut effect on cell proliferation following MC1R activation, as documented by the number of Ki-67 positive cells, which was not significantly different between treated and untreated HAoECs (see Supplementary Fig. S9A and S9B). MC1R expression did not significantly change over time during cell migration (not shown).
As cell migration is preceded by changes in cell morphology and actin filament remodelling, we evaluated whether these modifications occurred in HAoECs after 6h from stimulation with α-MSH 10-8 M. Indeed, we observed that MC1R activation through α-MSH induced an accelerated shift from a "cobblestone", polygonal shape to an elongated shape in these ECs, with rearrangement of actin filaments (Fig. 3A). Phalloidin-TRITC staining showed formation of aligned stress fibres in α-MSH-treated cells compared to untreated cells, whose actin filaments were mostly organized in short, unaligned stress fibres. HAoECs were then quantified for cell elongation, and cells stimulated with α-MSH showed a significantly higher number of elongated cells in comparison with control cells (Fig. 3B).
Since MC1R may signal through either cAMP increase or intracellular elevation of free cytosolic Ca2+ [43], we tested which signal transduction pathway was active in the enhancement of the HAoEC migration. Stimulation with α-MSH 10-8 M did not lead to an increase of intracellular cAMP in migrating HAoECs compared to control cells (see Supplementary Fig. S10). On the contrary, MC1R activation resulted in a significant, rapid, and sustained increase in intracellular Ca2+ levels over the control, early after the removal of the insert in the cell migration assay (Fig. 4A). This rise was completely abolished when HAoECs were pre-treated with the α-MSH antagonist 153N-6 10-5 M, which in turn alone did not affect Ca2+ signalling. Comparisons of the AUCs confirmed that the α-MSH-induced rise in Ca2+ levels was highly significant (Fig. 4B). Incubation with thapsigargin (10-8 M), a non-competitive inhibitor of sarco/endoplasmic reticulum Ca2+-ATPases (SERCAs) that causes a rapid raise of cytosolic Ca2+ by depleting endoplasmic reticulum stores [28], did not prevent a further significant rise of Ca2+ in response to a subsequent stimulus with α-MSH (Fig. 4C and 4D). This was almost completely inhibited by pre-treating HAoECs with the α-MSH antagonist 153N-6 10-5 M. Intriguingly, α-MSH 10-8 M was able to induce Ca2+ mobilization also in confluent HAoECs, to levels comparable to those produced by thapsigargin (see Supplementary Fig. S11). In this case, thapsigargin almost completely hindered a further rise of Ca2+ in response to a subsequent stimulus with α-MSH.
To further explore the Ca2+-dependency of the α-MSH-induced cell migration, we repeated the directional cell migration assay in the presence of the membrane-permeant Ca2+ chelator BAPTA-AM, which is widely used as an intracellular Ca2+ sponge to control intracellular calcium ion concentration ([Ca2+]i) [44]. The pro-migratory effect of α-MSH 10-8 M, once more significant at 9 and 12 h, was indeed completely inhibited by pre-treating HAoECs (no. c2) with BAPTA-AM 10-5 M (Fig. 5A), confirming that Ca2+ mobilization via MC1R activation was involved in modulating cell migration. Finally, since α-MSH-stimulation of MC1R may induce [Ca2+]i elevation via activation of the PLCβ pathway [45, 46], which in turn has a key role in mediating EC functions and angiogenesis [47], we carried out the directional cell migration assay also in the presence of the PLCβ inhibitor U-73122. Again, the pro-migratory effect of α-MSH 10-8 M was inhibited by pre-treating HAoECs with U-73122 5 × 10-5 M (Fig. 5B), suggesting that α-MSH evoked calcium mobilization/cell migration was dependent on the activation of PLCβ.
Time-course expression analysis revealed several gene modules dynamically regulated by MC1R activation
To evaluate the effects of MC1R activation at the transcriptional level, we assessed genome-wide gene expression profiles at 0.5, 3, 6, and 12h after stimulation with α-MSH 10-8 M vs. control in HAoECs cultured in the directional cell migration assay. Applying stringent filtering parameters, we deemed expressed 18936 of the 47231 measured transcripts (40%). Comparative time-course analysis, using the STEM algorithm [30], identified 637 genes whose expression consistently showed a median change ≥ 30% over time as an effect of stimulation with α-MSH. These genes fitted 57 of the 625 possible model profiles computed by the clustering algorithm (see Supplementary Fig. S12). Five hundred and six transcripts were associated with 15 distinct temporal profiles that showed a statistically significant enriched number of genes at a FDR<0.05 (see Supplementary Fig. S12 and Table S3a). The remaining 131 genes were associated with model profiles that had a FDR>0.05 (see online suppl. material, Table S3b) and, thus, were deemed as potentially arising from noise by random chance and excluded from subsequent analysis. Interestingly, we did not observe any significant change in MC1R expression level in migrating ECs after stimulation with α-MSH at any time point. The 15 significant temporal profiles of differential expression were grouped, based on their similarity by a correlation coefficient ≥ 0.7, to form 6 different clusters (Fig. 6A and 6B). Overall, genes belonging to clusters 1 and 4 showed a marked increase in expression at 6h in treated vs. untreated cells, whereas genes in clusters 2 and 3 displayed a marked decrease at the same time point; conversely, genes in clusters 5 and 6 appeared to be upregulated at 3h.
Time-course expression data of control and α-MSH stimulated cells at 3 and 6h were validated using PCR-based arrays profiling key genes involved in wound healing. Forty-eight genes were detected by both microarray and real-time PCR (see Supplementary Fig. S13), with a strong positive correlation between their average signal intensities (r ≥ 0.8, p<0.0001 for all pairwise correlations).
To uncover the biological meaning beneath these transcriptional effects, we performed a functional enrichment analysis of the 506 regulated genes (see online suppl. material, Table S4). Forty-four terms were significantly enriched at p<0.01 and FDR<0.20 and were used to draw a network to visually interpret biological data (Fig. 7). The most significant gene sets included the phosphoprotein class (FDR<0.00002), the endomembrane system (FDR<0.015), and the ECM-receptor interaction (FDR<0.025). Notably, 197 of the 506 regulated genes encode for phosphoproteins, 188 produce variant proteins by alternative splicing, and 65 are transcription factors or regulators. Importantly, 11 genes, i.e. AGRN, COL1A1, COL1A2, COL4A5, COL5A1, DAG1, ITGA2, ITGA10, LAMB1, LAMC1, and SPN, belonged to either the ECM-receptor interaction pathway or the extracellular matrix cellular component.
To further analyse gene expression changes in a structured fashion, functionally enrichment analysis was performed associating annotated gene sets with the 6 different clusters of temporal profiles (see online suppl. material, Table S5). Co-expressed gene subsets were visualized as temporal clustered profiles sharing functional annotations (Fig. 8). Coupling time-course gene expression analysis to enrichment analysis allowed identifying significantly regulated genes that have never been associated with MC1R signalling before, including genes involved in ECM-receptor interaction, vesicle-mediated transport, SNARE protein complex formation, and metal ion binding through metal-thiolate cluster structures (metallothioneins, MTs).
Discussion
In this work, we provide the first formal demonstration that human artery ECs express constitutively a functional MC1R and present evidence that activation of this melanocortin receptor drives faster EC migration and wound closure. Besides, time-course gene expression analysis allowed us identifying downstream molecular pathways associated with the enhanced cell motility following α-MSH engagement of the MC1R. This observation adds to the known functions of the melanocortin system, which is known to regulate homeostasis and possess cellular and tissue protective effects [2]. It is worth mentioning that our data are apparently in contrast with the notion that α-MSH inhibits migration of several cell types, such as immune/inflammatory [6, 7] or melanoma cells [48]: our findings pinpoint that the spectrum of action of MC1R signalling is wider and more diversified than previously thought, possibly depending on the cell type and pathophysiological context. Similarly, α-MSH binding to the MC1R stimulates or inhibits the proliferation respectively of cultured human melanocytes [49] and mesothelioma cell lines [50].
We detected both the MC1R mRNA and protein and demonstrated that it was functionally active, since resting confluent HAoECs were able to produce cAMP in response to exogenous α-MSH. No other component of the melanocortin system was detectable in HAoECs. Consistently, we did not detect α-MSH in cell culture supernatants. Thus, HAoECs are not a source of melanocortin peptides, but may be targeted by endocrine secretion (by the pituitary gland) or paracrine release of endogenous agonists (e.g. by immune cells at injured sites) [6, 7]. These findings represent a peculiar difference between macrovascular and microvascular ECs, as it has been reported that HDMECs express POMC and release melanocortin peptides upon stimulation [10]. In addition, unlike the microvascular ECs [12], MC1R expression did not significantly changed upon exposure of the HAoECs to α-MSH. This finding suggests that in macrovascular ECs MC1R expression levels are not influenced by pathways that depend on its activation and that HAoEC migration is not dependent on MC1R upregulation.
These results underline that artery ECs present cell type-restricted gene expression, which may account for a specific physiological function for MC1R, other than anti-inflammatory actions. This hypothesis led us to investigate whether MC1R activation could affect macrovascular EC migration after injury. EC migration is a fundamental process primed by damage and involved in vascular homeostasis and repair. Our data revealed that MC1R activation by α-MSH increased the rate of HAoEC migration, both in gap-closure and in injury-induced wound-healing assays, without significantly affecting cell proliferation. This effect was specifically induced by MC1R activation, since the 153N-6 peptide antagonist at MC1R [25] abolished the α-MSH-driven migration of HAoECs, reverting it to the same rate of control cells. Consistently, ECs carrying a loss-of-function mutation in the MC1R gene did not show a significant acceleration in cell motility upon challenge with α-MSH. In seeming contrast to what we observed, it has been recently reported that α-MSH inhibits in vitro migration of HUVECs [51]; but this reinforces our idea that the pro-migratory effect of α-MSH via MC1R activation is restricted to arterial ECs of the macrovasculature. Accordingly, we observed a prompt elevation of intracellular free Ca2+ after α-MSH stimulation in migrating cells, but not of cAMP, suggesting that MC1R activation enhances EC migration through the Ca2+ signalling cascade. Indeed, artificial intracellular calcium buffering by pre-treating cells with the cell-permeant Ca2+ chelator BAPTA-AM completely abolished the α-MSH-evoked acceleration in EC migration. Increase of calcium levels in the cytosol is an evolutionary conserved signal involved in the regulation of EC motility. Ca2+ mobilization can both stabilize and weaken cell-ECM interactions responsible for the asymmetry between cell front and rear adhesions, which finally results in cellular directed movement [52-54].
Intriguingly, our experiments showed that HAoEC MC1R might signal by increasing cAMP and/or intracellular Ca2+ depending on the cellular state: resting confluent HAoECs responded to MC1R engagement with α-MSH through cAMP and [Ca2+]i increase, while migrating cells responded through Ca2+ mobilization without any cAMP increase. To date, only a few reports support an involvement of calcium as a second messenger in MC1R signalling, besides cAMP. Ca2+ responses has been reported in HEK 293 cells transfected with mouse Mc1r [55] and in human melanoma cell lines [48], keratinocytes [56], and basophils [57] expressing MC1R. No elevation in cAMP was detected in keratinocytes and basophils in response to α-MSH [56, 57], whereas in melanoma cells and keratinocytes intracellular Ca2+ release was observed only in the presence of a pharmacological adenosine agonist that inhibits the cAMP pathway [48, 56]. Conversely, our findings provide evidence that MC1R couples to both cAMP and Ca2+ signalling systems in HAoECs and suggest that different functional states may direct alternative signalling pathways in macrovascular ECs. This is remarkable: MC1R appears to be one of those GPCRs that may simultaneously couple to distinct unrelated G-proteins and alternatively activate multiple intracellular effectors [58] depending on cell type, physiological condition, and the availability of G-protein (Gαs or Gαq) adaptors [59]. Of note, in the MCR family, alternative G-protein coupling has been reported for MC4R [60].
In HEK 293 cells transfected with Mc1r, complete depletion of intracellular Ca2+ stores following pre-treatment with thapsigargin 10-8 M abolished a further rise in [Ca2+]i in response to α-MSH, suggesting that the thapsigargin-sensitive endoplasmic reticulum Ca2+ stores were the source of [Ca2+]i increase [55]. This appears to be the case also for confluent HAoECs. On the contrary, in migrating HAoECs this concentration of thapsigargin was not able to inhibit a further rise in [Ca2+]i in response to α-MSH, suggesting that either SERCAs were only partially inhibited or α-MSH may increase intracellular Ca2+ from other sources. Indeed, angiogenic factors induce cytosolic calcium rises through either entry from extracellular space, by opening Ca2+ permeable channels in the plasma membrane, or release from intracellular organelles [61]. On the other hand, it has been shown that thapsigargin 1 µM may activate Ca2+ entry both by store-dependent and store-independent pathways in the HUVEC line EA.hy926 [62]. Here, we provide mechanistic insight showing that α-MSH-induced intracellular Ca2+ mobilization and accelerated migration are dependent on the activation of the PLC signalling pathway: in fact, the blockage of PLC by the specific inhibitor U-73122 completely abrogated the increase in HAoEC migration rate via MC1R activation. GPCR-mediated PLCβ activation cleaves membrane-bound phosphatidylinositol 4, 5 bisphosphate (PIP2) into inositol (1, 4,5) trisphosphate (IP3) and diacylglycerol; IP3 binding to IP3 receptor (IP3R) channels promotes Ca2+ release from the endoplasmic reticulum [63]. Consistent with our observation, MC1R has been recently shown to transduce through the PIP2-PLCβ pathway in sebocytes [45] and B16-F10 melanoma cells [46].
Time-series genome-wide gene-expression analysis on migrating HAoECs showed that large gene sets were significantly affected by α-MSH treatment: i.e., genes involved in the regulation of RNA transcription, encoding for proteins for which isoforms exist due to pre-mRNA splicing events (alternative splicing), and genes belonging to the phosphoprotein category. This indicates that MC1R activation has a wide influence on pathways playing a prominent role in regulating cellular activity. MC1R activation also modulated genes associated with the endomembrane system and intracellular organelle lumen, suggesting a role in controlling cellular trafficking and molecule mobilization. Remarkably, MC1R engagement with α-MSH affected the ECM-receptor interaction pathway, which is known to be critical for the directional haptotactic EC migration [64]. Conversely, MC1R activation did not affect the expression of cell cycle-related genes, which was consistent with the apparent lack of effect on HAoEC proliferation in the gap closure assay. Our findings suggest that the regulation of the ECM components, i.e. collagens and laminins, and of their receptors, i.e. integrins and dystroglycans, through MC1R may drive higher HAoEC motility. α-MSH appears to boost HAoEC migration regulating the interaction between the cellular receptor integrins (ITGA2 and ITGA10) and DAG1 to their ECM counterpart collagens (COL1A1, COL1A2, COL4A5, and COL5A1), laminins (LAMB1 and LAMC1) and AGRN. The directed motility of ECs is strictly dependent on cell adhesion to ECM [64, 65]. Integrins and interstitial collagen mediate haptotactic cell migration, which is of primary importance in driving EC migration during large vessel repair [21, 66]. Furthermore, time-course analysis evidenced that 9 genes of the ECM-receptor interaction pathway had a similar temporal expression profile, with a peak induction at 3h followed by a reversion at 6h, suggesting that common factor(s) may control their co-expression. Conversely, ITGA2 and SPN showed a specular temporal profile, with a later peak expression at 6h, which is suggestive of a sequential upregulation of ECM-receptor interaction genes [67]. Remarkably, we showed that α-MSH enhances EC migration along with actin filament remodelling and changes in cell architecture. Binding of integrins to type-I collagen suppresses cAMP production and the activity of cAMP-dependent protein kinase A: consequently, actin polymerization is induced, contributing to the formation of stress fibres and to EC contractility, which finally generates the directional movement [68]. This is consistent with the idea that the fine-tuning of integrins and their binding molecules promoted by MC1R activation plays a key role in conditioning HAoEC migration rate. In addition, MC1R stimulation induced an early upregulation of SNARE proteins (which mediate vesicle-membrane fusion) and cytoplasmic vesicle genes, followed by a later overexpression of metal-binding proteins. Importantly, trafficking and delivery/fusion vesicle proteins are essential for the regulation of front-rear polarity during directional cell migration [69, 70]. Likewise, MTs enhance EC motility [71] and angiogenesis [72] through transcriptional regulation of various vascular growth-factors, and their modulation drove suppression of reactive oxygen species production in ECs exposed to elevated laminar shear stress [73]. Such a pattern of temporal dynamics in gene expression (i.e. ECM-receptor interaction, SNARE, or MT genes) is expected as an "impulse response" to a transient signal, namely a single pulse of α-MSH [74]. This typical oscillating wave of co-expressed genes may reflect a highly ordered temporal organization in gene transcription, which ultimately results in the subsequent, coordinated translation into the corresponding effector proteins that drive the α-MSH-mediated increase in EC migration speed. Consistently, the peaks in gene expression were followed at 6-12 hours by a transition to the steady state.
In summary, MC1R activation via α-MSH appears to accelerate directional HAoEC migration through the following steps: (a) binding of melanocortin hormones to MC1R induces (b) an increase in cytosolic Ca2+ while preventing a rise in cAMP biosynthesis, through a putative alternative G-protein coupling and PLC-pathway activation, and subsequently (c) the coordinate modulation of genes of the ECM-receptor interaction, vesicle- and SNARE-mediated trafficking pathways, and metal sensing proteins, (d) possibly regulating the cell front-rear polarity. These responses reflect an unrecognized protective function of the melanocortin system, which is fostered by previously unreported α-MSH-activated, MC1R-mediated signalling and molecular pathways.
MC1R tonic signalling and pro-migratory action may be relevant for the homeostatic functions of the arterial endothelium. The endothelium monolayer lining in the luminal side of blood vessels plays a pivotal role in the regulation of the haemostatic balance, prevention of vascular inflammation, and protection against vascular injury [75]. Normal ECs express a number of inhibitors of platelet and leukocyte activation, vasodilators, and anticoagulant and procoagulant molecules. Damage to these cells is associated with a shift in the haemostatic balance to the procoagulant side [15], loss of protective molecules and expression of adhesive, inflammatory and mitogenic factors, leading to the development of thrombosis, pathologic remodelling, and atherosclerosis [75]. Endothelial dysfunction is characterized by an imbalance between procoagulant and anticoagulant mediators and regenerated arterial endothelium may be functionally incompetent with reduced expression of antithrombotic molecules [15, 76]. EC migration is a key event in wound healing and tissue regeneration, including reendothelialisation after stent implantation [76]. Remarkably, MC1R activation in migrating HAoEC did not alter the balance between pro- and anticoagulant genes, i.e. expression of procoagulant (such as VWF, F2R, and F3) and anticoagulant genes (THBD, HSPG, EPCR, and TFPI) was not affected by α-MSH. Thus, MC1R activation may have beneficial effects both ameliorating HAoEC motility properties and maintaining the equilibrium between pro- and anticoagulant signals.
Conclusion
Our work broadens the knowledge on MCR regulatory roles and supports the concept of a novel function for peripherally expressed MC1R, whose signalling may participate in preventing/healing of artery endothelial dysfunction, vascular repair, and reendothelialisation. Endothelial artery MC1R could represent a target for original therapeutic strategies aimed at preventing/repairing endothelial injury in a variety of cardiovascular pathological conditions associated with endothelial denudation [20].
Acknowledgements
We thank Dr Chiara Speroni for help and excellent technical assistance. We thank Fondazione Banca di Treviso ONLUS for kindly providing us with the human aorta specimen. This study was supported by Institutional Research Funds (Italian Ministry of Health, Funds 5‰ 2009-11; to G.I.C.).
Disclosure Statement
The authors declare no conflict of interests.
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