Atherosclerosis (hardening of the arteries) leads to high blood pressure, blockage, heart attack, stroke, and death in millions. More Mcr1 stimulation will decrease/stop this. Msh/Scenesse therapy here could be relavent for tens of millions patients
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Deficiency in Melanocortin 1 Receptor Signaling Predisposes to Vascular Endothelial Dysfunction and Increased Arterial Stiffness in Mice and Humans.
Abstract
OBJECTIVE:
The melanocortin 1 receptor (MC1-R) is expressed by vascular endothelial cells and shown to enhance nitric oxide (NO) availability and vasodilator function on pharmacological stimulation. However, the physiological role of MC1-R in the endothelium and its contribution to vascular homeostasis remain unresolved. We investigated whether a lack of functional MC1-R signaling carries a phenotype with predisposition to vascular abnormalities.
APPROACH AND RESULTS:
Recessive yellow mice (MC1Re/e), deficient in MC1-R signaling, and their wild-type littermates were studied for morphology and functional characteristics of the aorta. MC1Re/e mice showed increased collagen deposition and arterial stiffness accompanied by an elevation in pulse pressure. Contractile capacity and NO-dependent vasodilatation were impaired in the aorta of MC1Re/e mice supported by findings of decreased NO availability. These mice also displayed elevated levels of systemic and local cytokines. Exposing the mice to high-sodium diet or acute endotoxemia revealed increased susceptibility to inflammation-driven vascular dysfunction. Finally, we investigated whether a similar phenotype can be found in healthy human subjects carrying variant MC1-R alleles known to attenuate receptor function. In a longitudinal analysis of 2001 subjects with genotype and ultrasound data (The Cardiovascular Risk in Young Finns Study), weak MC1-R function was associated with lower flow-mediated dilatation response of the brachial artery and increased carotid artery stiffness.
CONCLUSIONS:
The present study demonstrates that deficiency in MC1-R signaling is associated with increased arterial stiffness and impairment in endothelium-dependent vasodilatation, suggesting a physiological role for MC1-R in the regulation of arterial tone.
© 2015 American Heart Association, Inc.
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Excerpt:
What Are the Clinical Implications?
• The removal of excess cholesterol from macro- phage-derived foam cells in atherosclerotic plaques is the critical first step of reverse cholesterol trans- port, which facilitates the transport of peripheral cholesterol to the liver for excretion.
• Promoting macrophage reverse cholesterol trans- port limits the progression of atherosclerosis and may ideally complement other lipid-lowering ther- apies to control for the residual risk that is often present in medically treated patients with athero- sclerotic disease.
• Therefore, the identification of MC1-R in lesional macrophages and its role in regulating reverse cholesterol transport, combined with the estab- lished anti-inflammatory effects of MC1-R, could serve as an attractive new approach for preventing atherosclerosis.
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Inflammation and its resolution in atherosclerosis: mediators and therapeutic opportunities
Magnus Bäck, Arif Yurdagul Jr, Ira Tabas, Katariina Öörni & Petri T. Kovanen
Nature Reviews Cardiology (2019) | Download Citation
Abstract
Atherosclerosis is a lipid-driven inflammatory disease of the arterial intima in which the balance of pro-inflammatory and inflammation-resolving mechanisms dictates the final clinical outcome. Intimal infiltration and modification of plasma-derived lipoproteins and their uptake mainly by macrophages, with ensuing formation of lipid-filled foam cells, initiate atherosclerotic lesion formation, and deficient efferocytotic removal of apoptotic cells and foam cells sustains lesion progression. Defective efferocytosis, as a sign of inadequate inflammation resolution, leads to accumulation of secondarily necrotic macrophages and foam cells and the formation of an advanced lesion with a necrotic lipid core, indicative of plaque vulnerability. Resolution of inflammation is mediated by specialized pro-resolving lipid mediators derived from omega-3 fatty acids or arachidonic acid and by relevant proteins and signalling gaseous molecules. One of the major effects of inflammation resolution mediators is phenotypic conversion of pro-inflammatory macrophages into macrophages that suppress inflammation and promote healing. In advanced atherosclerotic lesions, the ratio between specialized pro-resolving mediators and pro-inflammatory lipids (in particular leukotrienes) is strikingly low, providing a molecular explanation for the defective inflammation resolution features of these lesions. In this Review, we discuss the mechanisms of the formation of clinically dangerous atherosclerotic lesions and the potential of pro-resolving mediator therapy to inhibit this process.
Key points
Modified lipoproteins and cholesterol crystals accumulate in the arterial intima and induce foam cell formation and inflammation.
Defective efferocytosis of apoptotic foam cells leads to necrotic core formation.
Defective efferocytosis is a sign of failure in the resolution of inflammation.
Inflammation resolution is mediated by specialized pro-resolving lipid mediators, proteins and signalling gases.
Improvement of the balance between pro-inflammatory and pro-resolving processes enables the resolution of inflammation.
Pro-resolving mediator therapy could be a novel approach to suppressing the formation of clinically dangerous atherosclerotic lesions.
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Protection of cultured brain endothelial cells from cytokine-induced damage by α-melanocyte stimulating hormone
Research articleCell BiologyNeurosciencePharmacology
András Harazin1,2, Alexandra Bocsik1, Lilla Barna1,2, András Kincses1, Judit Váradi3, Ferenc Fenyvesi3, Vilmos Tubak4, Maria A. Deli1, Miklós Vecsernyés3
Published May 15, 2018PubMed 29780671
Author and article information
Abstract
The blood–brain barrier (BBB), an interface between the systemic circulation and the nervous system, can be a target of cytokines in inflammatory conditions. Pro-inflammatory cytokines tumor necrosis factor-α (TNF-α) and interleukin-1β (IL-1β) induce damage in brain endothelial cells and BBB dysfunction which contribute to neuronal injury. The neuroprotective effects of α-melanocyte stimulating hormone (α-MSH) were investigated in experimental models, but there are no data related to the BBB. Based on our recent study, in which α-MSH reduced barrier dysfunction in human intestinal epithelial cells induced by TNF-α and IL-1β, we hypothesized a protective effect of α-MSH on brain endothelial cells. We examined the effect of these two pro-inflammatory cytokines, and the neuropeptide α-MSH on a culture model of the BBB, primary rat brain endothelial cells co-cultured with rat brain pericytes and glial cells. We demonstrated the expression of melanocortin-1 receptor in isolated rat brain microvessels and cultured brain endothelial cells by RT-PCR and immunohistochemistry. TNF-α and IL-1β induced cell damage, measured by impedance and MTT assay, which was attenuated by α-MSH (1 and 10 pM). The peptide inhibited the cytokine-induced increase in brain endothelial permeability, and restored the morphological changes in cellular junctions visualized by immunostaining for claudin-5 and β-catenin. Elevated production of reactive oxygen species and the nuclear translocation of NF-κB were also reduced by α-MSH in brain endothelial cells stimulated by cytokines. We demonstrated for the first time the direct beneficial effect of α-MSH on cultured brain endothelial cells, indicating that this neurohormone may be protective at the BBB.
Cite this as
Harazin A, Bocsik A, Barna L, Kincses A, Váradi J, Fenyvesi F, Tubak V, Deli MA, Vecsernyés M. 2018. Protection of cultured brain endothelial cells from cytokine-induced damage by α-melanocyte stimulating hormone. PeerJ 6:e4774 https://doi.org/10.7717/peerj.4774
Main article text
Introduction
The neuropeptide α-melanocyte stimulating hormone (α-MSH) belongs to the family of melanocortins, which are created from pro-opiomelanocortin (Catania et al., 2010). The α-MSH, a 13 amino acid long peptide hormone, is mainly produced in the hypothalamic region (Brzoska et al., 2008). Induction of melanogenesis in pigment cells and immunomodulation are among its main physiological functions (Catania, 2008). There are five melanocortin receptors (MCRs), from which four, MC1R, MC3R, MC4R, and MC5R, bind α-MSH (Brzoska et al., 2008). MCRs are G-protein-coupled and exert their effects via cyclic 3′,5′-adenosine monophosphate (cAMP)-dependent signaling pathways. The predominant receptor of α-MSH is MC1R, which binds the neuropeptide with high affinity, and is expressed in both the brain and periphery. MC1R was demonstrated in several tissues, such as brain, skin, immune system, and gut (Brzoska et al., 2008).
Melanocortins are evolutionary conserved defense molecules against tissue injury and bacterial invasion (Catania, 2008). The immunomodulatory and anti-inflammatory effects of α-MSH are well known and widely investigated. The anti-inflammatory action of α-MSH was proven in animal models of systemic or local inflammation, like sepsis, arthritis, uveitis, dermatitis, pancreatitis, and colitis (Brzoska et al., 2008). The protective effect of α-MSH was also shown in ischemic conditions in the heart (Vecsernyés et al., 2003, 2017), retina (Varga et al., 2013), kidney and intestine (Brzoska et al., 2008). In addition to animal models of gut inflammation (Rajora et al., 1997a), the effects of α-MSH were examined on cell cultures. Our groups have recently described that pro-inflammatory cytokines tumor necrosis factor-α (TNF-α) and interleukin-1β (IL-1β) disrupted the barrier integrity of Caco-2 human intestinal epithelial cells, which was attenuated by α-MSH via the inhibition of the NF-κB pathway (Váradi et al., 2017).
In addition to systemic inflammatory conditions, the beneficial effect of α-MSH was also investigated in models of acute and chronic injuries of the central nervous system (CNS; Catania, 2008). Treatment with α-MSH was neuroprotective in cerebral ischemia, traumatic spinal cord injury, kainic acid induced excitotoxic brain damage, and lipopolysaccharide (LPS)-induced cerebral inflammation in animal studies. One of the main mechanisms of action of α-MSH is the down-regulation of pro-inflammatory cytokines TNF-α and/or IL-1β in blood and brain tissue, as it was described in LPS-induced brain inflammation (Rajora et al., 1997b) and unilateral middle cerebral artery occlusion (Huang & Tatro, 2002) in mice, and in kainic acid-induced brain damage in rats (Forslin Aronsson et al., 2007). Studies on cultured astrocytes, neurons, and microglia revealed that, similarly to the periphery, inhibition of the NF-κB pathway and downstream blockade of pro-inflammatory cytokine release and nitric oxide overproduction contribute to the protective effects of α-MSH (Catania, 2008).
Dysfunction of the blood–brain barrier (BBB) has been described in many systemic and CNS inflammatory diseases (Erickson & Banks, 2018) and plays a central role in the pathomechanism of many neurological diseases (Zhao et al., 2015; Liebner et al., 2018). TNF-α and IL-1β are the two most studied major pro-inflammatory cytokines in neuroinflammation observed in CNS pathologies (Sochocka, Diniz & Leszek, 2017; Liebner et al., 2018), and induced by systemic inflammation (Hoogland et al., 2015). The elevated levels of these two cytokines, confirmed in chronic neurodegenerative diseases (Sochocka, Diniz & Leszek, 2017), open the BBB (Liebner et al., 2018). TNF-α directly increases the permeability of the BBB for marker molecules in animal studies (Megyeri et al., 1992; Ábrahám et al., 1996), as well as in culture models (Deli et al., 1995) and a similar effect was described for IL-1β in BBB in vitro experiments (for review see Deli et al., 2005). Since these two pro-inflammatory cytokines are major players in both systemic and CNS inflammation and induce BBB dysfunction, TNF-α and IL-1β were selected for our experiments.
A binding site was found on murine brain endothelial cells by radiolabeled α-MSH (de Angelis et al., 1995), suggesting the presence of MCR(s) at the BBB, but they were not identified. MCRs expressed on cells of the neurovascular unit could mediate the neuroprotective actions of melanocortins (Catania, 2008), however, the effects of α-MSH on the BBB, especially on brain endothelial cells have not been investigated yet.
Therefore, our aim was to investigate the effects of α-MSH peptide on a culture model of the BBB, its possible protective action against barrier disruption induced by TNF-α and IL-1β cytokines by measuring cellular viability, permeability for marker molecules, structure of interendothelial tight junctions, production of reactive oxygen species (ROS) and the nuclear translocation of NF-κB p65 subunit.
Materials and Methods
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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).
Table 1. Expression of the MCRs, POMC, and prohormone convertases in primary human macrovascular endothelial cells. HAoEC: human aortic endothelial cells; HCAEC: human coronary artery endothelial cells; HA: human astrocytes (positive control). Three different primary lines for each type of EC were analysed by RT-qPCR: from c1,c4ECACC, c2,c5Lonza, and c3,c6Promocell. In the upper panel, detection levels are reported as: -, undetected; +, <35 Ct; and ++, <30 Ct. In the lower panel, MC1R expression levels in each cell line are shown as means ± SEM of triplicate technical replicates
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).
Fig. 1. HAoECs express a functional MC1R. (A) Immunoblot analysis of membrane extracts showed that all three studied primary cells express MC1R on the plasma membrane. An anti-ATPase Na+/K+ transporting subunit alpha 1 (ATP1A1) antibody was used as membrane-specific loading control (upper light grey arrow). Absence of a β-actin immunoreactive band (lower light grey arrow) excluded contamination with cytoplasmic proteins in this preparation. The 37 kDa MC1R-specific immunoreactive band is indicated by a grey arrow. Lanes are: M, molecular weight marker; C+, HEK293 cells transiently transfected with the MC1R full-length cDNA (positive control); c1, c2, c3, primary HAoECs from ECACC, Lonza, and Promocell, respectively. (B) Intracellular cAMP concentrations were measured in confluent HAoECs after treatment with α-MSH for 5 min, with or without the MC1R-selective α-MSH antagonist 153N-6. Results are shown as scatter dot plots with mean ± SD (n = 5-6 per treatment group). Statistical significance of differences was assessed by one-way ANOVA [F(3,18) = 7.900, p=0.0014] followed by Tukey's post-hoc test (*p<0.05, **p<0.01). (C) Immunohistochemical detection of MC1R in a normal human aorta specimen (10×) confirmed that HAoECs express the receptor in vivo. To control for staining specificity, we used secondary antibody alone (left), anti-MC1R antibody pre-adsorbed with the specific blocking peptide and secondary antibody (centre), and anti-MC1R antibody with secondary antibody (right): only the latter showed an intense staining.
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).
Fig. 2. MC1R activation enhances HAoEC migration. (A) After insert removal, HAoEC (no. c2) monolayers were treated with α-MSH 10-8 M, with or without the MC1R-antagonist 153N-6 10-5 M, and allowed to migrate for 3, 9, and 12h: gap closure was quantified using DiR cell staining and near-infrared fluorescence scanning. Results are shown as mean ± SEM (n = 6). Statistical significance of differences was assessed by two-way ANOVA [F(9,80) = 2.957, p=0.0044, interaction time × treatment; F(3,80) = 10.85, p<0.0001, treatment effect] with Bonferroni post-hoc test [**p<0.01, ***p<0.001, α-MSH vs. medium alone (C)]. (B) Directional migration assay was repeated in the presence of the proliferation inhibitor PD0332991. Results are shown as mean ± SEM (n = 10). Statistical significance of differences was assessed by two-way ANOVA [F(3,72) = 3.018, p=0.0353, for interaction; F(1,72) = 9.074, p=0.0036, treatment effect] with Bonferroni post-hoc test (*p<0.05, **p<0.01). (C) The migration assay was repeated with HAoECs no. c1, the other cell line bearing wild-type MC1R alleles. Results are shown as mean ± SEM (n = 4). Statistical significance was assessed by two-way ANOVA [F(1,24) = 6.016, p=0.0218, for treatment effect] with Bonferroni post-hoc test (**p<0.01). (D) The migration assay was finally repeated with HAoECs no. c3, carrying a loss-of-function mutation in the MC1R gene. Results are shown as mean ± SEM (n = 5). Two-way ANOVA with Bonferroni post-hoc test showed no statistically significant differences.
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).
Fig. 3. MC1R activation enhances actin filament remodelling and cell elongation in migrating HAoECs. (A) Non-confluent ECs were stimulated with α-MSH for 6 h, then fixed and stained with TRITC-labelled phalloidin for actin filament visualization, using DAPI for nuclear counterstain (40×). Aligned stress fibres and cellular elongation are pronounced in treated vs. untreated HAoECs. (B) Quantification of cell elongation. Cells with axial ratios > 3 were counted in randomly selected fields and expressed as percentages of the total cells counted. Results are shown as scatter dot plots with mean ± SD (n = 3 per group). Statistical significance of differences was assessed by two-tailed unpaired t test (*p=0.0356).
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.
Fig. 4. MC1R activation increases intracellular calcium levels in migrating HAoECs. (A) Treatment with α-MSH after insert removal in the cell migration assay induced a prompt increase in intracellular Ca2+ levels (as detected by Fluo-4 NW fluorescent calcium indicator), which was completely abolished by pre-treatment with the MC1R-antagonist 153N-6. (C) Rise of intracellular Ca2+ in response to the stimulus with α-MSH was not prevented by prior stimulation with thapsigargin (THAPS). This was inhibited by pre-treating HAoECs with 153N-6. Arrows indicate thapsigargin or α-MSH stimulation. Curves present the mean ± SEM of n = 5-6 independent experiments. RFU, relative fluorescence unit. (B, D) The areas under the curve (AUC) were used to compare α-MSH-induced effects with control treatments. Results are shown as scatter dot plots and mean ± SD Statistical significance of differences was assessed by one-way ANOVA [(B) F(3,17) = 14.56, p<0.0001; (D) F(3,16) = 12.50, p=0.0001] with Tukey's post-hoc test (*p<0.05, **p<0.01, ***p<0.001, ****p<0.0001).
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.
Fig. 5. Calcium chelation or PLC inhibition prevent MC1R-mediated enhancement of HAoEC migration. (A) After insert removal, HAoEC (no. c2) monolayers were treated with α-MSH 10-8 M, with or without pre-treatment with the intracellular Ca2+ chelator BAPTA-AM 10-5 M, and allowed to migrate for 3, 9, and 12h: gap closure was quantified using DiR cell staining and near-infrared fluorescence scanning. Results are shown as mean ± SEM (n = 5-6 per group). Statistical significance was assessed by two-way ANOVA [F(9,68) = 2.337, p=0.0233, interaction time × treatment; F(3,68) = 5.493, p=0.0019, treatment effect] with Bonferroni post-hoc test [*p<0.05, **p<0.01, α-MSH vs. medium alone (C)]. (B) Directional migration assay was carried out also pre-treating cells with the PLC inhibitor U-73122. Results are shown as mean ± SEM (n = 5-6). Statistical significance was assessed by two-way ANOVA [F(9,68) = 2.607, p=0.0120, for interaction; F(3,68) = 7.102, p=0.0003, treatment effect] with Bonferroni post-hoc test (*p<0.05).
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.
Fig. 6. Significant temporal expression profiles of genes modulated by MC1R activation in migrating HAoECs. (A) List of significantly enriched temporal expression patterns identified by STEM analysis. Expression profiles are grouped into six clusters based on their similarity (r ≥ 0.7) and ordered by p-value significance within each cluster profile. The number of genes belonging to a profile is reported. (B) Heatmap depicting temporal expression of genes within each cluster. Genes hierarchically clustered into 6 groups using one minus Pearson correlation distance and the average linkage method. Data are the average log2 gene expression ratio of α-MSH stimulated cells to non-stimulated cells (n = 3 independent experiments, with two technical replicates each). Normalized expression ratios are shown as a gradient colour ranging from lower (blue) to higher (gold) values.
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.
Fig. 7. Modules of co-regulated genes in migrating HAoECs upon MC1R activation. The enrichment map of modulated genes was drawn as a network of the most significant functionally annotated gene sets (p<0.01 and Benjamini FDR<0.20). Nodes represent gene sets. Node colour intensity is relative to enrichment significance, from lower (light) to higher (dark red). Node size is proportional to the gene set size. Gene sets are connected by green edges based on their similarity. Edge thickness measures the degree of the overlap between two gene sets (using a cut-off of the Jaccard plus Overlap combined coefficient = 0.375). Clusters of tightly, functionally related gene sets are circled and assigned an overall label. Heat maps of temporal expression patterns of relevant gene sets and pathways are displayed. Hierarchical clustering of genes was performed using one minus Pearson correlation distance and the average linkage method. Row normalized expression values are shown as a gradient colour ranging from lower (blue) to higher (gold) values.
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).
Fig. 8. Time-course gene cluster profiles. The 6 different expression profiles represent time-dependent dynamic gene modulation as the mean of significant temporal profiles grouped on the basis of their similarity. Each cluster profile is associated with gene sets and pathways (coloured rectangles) significant at the enrichment analysis. For cluster 5, a gene set with a nominal p<0.05 is indicated. On the y-axes is the log2 mean fold change (FC) relative to control cells, i.e. the log2 gene expression ratio of α-MSH stimulated cells to non-stimulated cells; on the x-axis is the experimental time scale (hours).
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.
Uhohinc
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- 1Research Centre for Integrative Physiology & Pharmacology, Institute of Biomedicine, University of Turku, Turku, Finland
- 2Drug Research Doctoral Programme (DRDP), University of Turku, Turku, Finland
- 3MediCity Research Laboratory, University of Turku, Turku, Finland
- 4Turku Center for Disease Modeling, University of Turku, Turku, Finland
Melanocortin receptor 1 (MC1-R) is expressed in leukocytes, where it mediates anti-inflammatory actions. We have previously observed that global deficiency of MC1-R signaling perturbs cholesterol homeostasis, increases arterial leukocyte accumulation and accelerates atherosclerosis in apolipoprotein E knockout (Apoe-/-) mice. Since various cell types besides leukocytes express MC1-R, we aimed at investigating the specific contribution of leukocyte MC1-R to the development of atherosclerosis. For this purpose, male Apoe-/- mice were irradiated, received bone marrow from either female Apoe-/- mice or MC1-R deficient Apoe-/- mice (Apoe-/- Mc1re/e) and were analyzed for tissue leukocyte profiles and atherosclerotic plaque phenotype. Hematopoietic MC1-R deficiency significantly elevated total leukocyte counts in the blood, bone marrow and spleen, an effect that was amplified by feeding mice a cholesterol-rich diet. The increased leukocyte counts were largely attributable to expanded lymphocyte populations, particularly CD4+ T cells. Furthermore, the number of monocytes was elevated in Apoe-/- Mc1re/e chimeric mice and it paralleled an increase in hematopoietic stem cell count in the bone marrow. Despite robust leukocytosis, atherosclerotic plaque size and composition as well as arterial leukocyte counts were unaffected by MC1-R deficiency. To address this discrepancy, we performed an in vivo homing assay and found that MC1-R deficient CD4+ T cells and monocytes were preferentially entering the spleen rather than homing in peri-aortic lymph nodes. This was mechanistically associated with compromised chemokine receptor 5 (CCR5)-dependent migration of CD4+ T cells and a defect in the recycling capacity of CCR5. Finally, our data demonstrate for the first time that CD4+ T cells also express MC1-R. In conclusion, MC1-R regulates hematopoietic stem cell proliferation and tissue leukocyte counts but its deficiency in leukocytes impairs cell migration via a CCR5-dependent mechanism.
Atherosclerosis is characterized by deposition of cholesterol and chronic inflammation within arterial walls (1, 2). Cholesterol-rich lipoprotein particles accumulate in the sub-endothelial layer of medium- and large-sized arteries, which, in turn, promotes endothelial cell activation and consequent entry of leukocytes into growing atherosclerotic lesions. It has been demonstrated that the development of atherosclerotic lesions correlates directly with an increased number of circulating leukocytes, particularly monocytes, and their enhanced recruitment to inflamed arteries (3, 4). Monocytes in the intima differentiate into macrophages, take up modified lipoprotein particles via scavenger receptors and eventually transform into foam cells. These cells have impaired migratory capacity and tend to persist in atherosclerotic plaques, thus promoting disease progression (5–8). Hypercholesterolemia is an important risk factor for atherosclerosis, since it not only drives lipid deposition in lesions but it also induces an expansion of circulating monocytes (6). This is attributable to an increase in the classical, pro-inflammatory Ly6Chigh monocyte subset and is primarily mediated by increased proliferation of hematopoietic stem cells and their release from the bone marrow. In addition to cholesterol, hematopoiesis is induced by sympathetic activation and production of proinflammatory cytokines that follow acute cardiovascular events such as myocardial infarction (9). While hematopoiesis occurs mainly in the bone marrow under healthy steady-state conditions, atherosclerosis-linked inflammation can stimulate myeloid cell production in other lymphoid organs, particularly in the spleen (10). Monocytes from the bone marrow and spleen are released into the circulation, extravasate and accumulate as macrophages in the plaque. Although macrophages predominate in the lesions, T cells are also abundant and contribute significantly to the initiation and progression of atherosclerosis (11). CD4+ T cells are the most frequent T cell subset in the plaque and their disease-promoting activity is especially ascribed to the interferon-γ (IFN-γ)-producing T helper type 1 (Th1) subset (12, 13). In contrast, regulatory T cells (Tregs) are atheroprotective (14), while the role of Th2 cells is more controversial based on observations of both pro- and anti-atherosclerotic effects (11). Evidence thereby strongly supports the involvement of both innate and adaptive immunity in atherogenesis.
We have recently found that melanocortins are implicated in the regulation of macrophage function and cholesterol homeostasis in atherosclerosis (15, 16). Melanocortins are products of post-translational processing of the pro-opiomelanocortin (POMC) precursor protein and include melanocyte-stimulating hormones (α-, β- and γ-MSH) and adrenocorticotropic hormone (ACTH) (17). They interact with five closely related G-protein coupled melanocortin receptor (MC1-R - MC5-R), which regulate important physiological functions including skin pigmentation, steroidogenesis and energy homeostasis (18). MC1-R is the best-characterized regulator of skin and hair coloration but it also plays a significant role in the control of inflammation (19–21). It has a wide expression profile in the immune system and is present virtually in all major leukocyte subpopulations including neutrophils, monocytes, macrophages, dendritic cells, cytotoxic T cells and B lymphocytes (22–25). It is well-established that MC1-R exerts potent anti-inflammatory actions (21), making it an attractive therapeutic target in inflammatory diseases such as atherosclerosis. Indeed, MC1-R activation was found to be protective in experimental models of atherosclerosis (15, 26, 27), but the underlying mechanism is likely to be multifactorial and extends beyond the immunoregulatory function of MC1-R. We recently identified a novel role for MC1-R in macrophage cholesterol transport, which provides protection against atherosclerosis by inhibiting foam cell formation (15). Conversely, phenotyping of recessive yellow Mc1re/e mice that carry a single base deletion mutation in the Mc1r gene revealed that MC1-R deficiency exacerbates atherosclerosis by disturbing cholesterol and bile acid metabolism and by increasing arterial accumulation of pro-inflammatory Ly6Chigh monocytes (16). However, the observed phenotype cannot be solely attributed to loss-of-function of MC1-R in leukocytes, since this receptor is distributed also in other tissues relevant to atherosclerosis such as the liver, endothelium and adipose tissue (28–31). Therefore, the exact role of leukocyte MC1-R in the development of atherosclerosis remains to be determined. In this study, we set out to address this and transplanted apolipoprotein E knockout (Apoe-/-) mice with MC1-R deficient bone marrow and then analyzed tissue leukocyte profiles and plaque phenotype of this mouse model.
Materials and MethodsMiceC57BL/6 Mc1re/e recessive yellow mice (Jackson Laboratory, strain# 000060, Bar Harbor, ME, USA) were intercrossed with C57BL/6 Apoe-/- mice (Jackson Laboratory, strain# 002052) to generate double mutant Apoe-/- Mc1re/e mice. Mice were housed in groups of 3-5 littermates on a 12 h light/dark cycle. The experiments were approved by the local ethics committee (Animal Experiment Board in Finland, License Numbers: ESAVI/6280/04.10.07/2016 and ESAVI/1260/2020) and conducted in accordance with the institutional and national guidelines for the care and use of laboratory animals.
Bone Marrow TransplantationBone marrow (BM) cells were isolated from the femurs and tibiae of female Apoe-/- Mc1re/e mice or age-matched Apoe-/- mice. Eight- to ten-week-old recipient Apoe-/- male mice were lethally irradiated with two doses of 5 Gy 3 hours apart in a Faxitron MultiRad 225 X-ray irradiator (32). Three days later, recipient mice were reconstituted intravenously with 1x107 BM cells from either Apoe-/- or Apoe-/- Mc1re/e mice. Mice received acidified and autoclaved water from 1 week prior to the BM transplantation until 4 weeks after. Mice were allowed to recover for 6 weeks and were thereafter either maintained on a normal chow diet or fed a Western-type diet (D12079B, Research Diets Inc, NJ, USA) for 10 weeks. At the end of the experiment, mice were euthanized via CO2 asphyxiation and whole blood was obtained via cardiac puncture. Aorta, spleen, liver and bone marrow (femur) were collected for further analyses.
To determine the efficiency of BM reconstitution, genomic DNA was extracted (QIAamp DNA Blood Mini Kit, Qiagen, USA) from peripheral blood of recipient mice at the end of the experiment. Samples were quantified by real-time PCR (Applied Biosystems 7300 Real-Time PCR system) for the expression of the Y chromosome-specific gene Zfy1 and a reference gene (Bcl2) (33). The engraftment of female donor cells in the recipient males was calculated using a standard curve generated from samples with known percentages of male and female DNA. At 16 weeks after BM transplantation, donor cell engraftment level in the peripheral blood ranged from 95% to 99%. The level of chimerism was 97.2 ± 0.3% in Apoe-/- and 97.6 ± 0.4% in Apoe-/- Mc1re/e BM transplanted mice (P=0.48).
Flow CytometryTotal leukocytes and leukocyte subsets in the aorta, spleen, bone marrow (femur) and whole blood were quantified by flow cytometry as previously described (16). Aortic samples were digested with an enzymatic cocktail (Collagenase I, 450 U/ml; Collagenase XI, 250 U/ml; Hyaluronidase, 120 U/ml; DNase I, 120 U/ml; Sigma Aldrich) for 60 min at 37°C and then filtered through a 50-µm cell strainer (BD Biosciences). Single cell suspensions were stained for 30 min at 4°C with fluorochrome-conjugated antibodies against CD45.2 (clone 30-F11, BD Biosciences), CD11b (clone M1/70, BioLegend), CD115 (clone AFS98, BioLegend), Ly6C (clone AL-21, BD Biosciences) and Ly6G (clone 1A8, BD Biosciences). To quantify lymphocyte subsets, cells were stained with CD45.2 (clone 30-F11, BD Biosciences), CD11b (clone M1/70, BioLegend), CD19 (clone 6D5, Biolegend) CD4 (GK1.5, Biolegend), CD8a (clone 53-6.7, Biolegend), NK1.1 (clone PK136, Biolegend) and TCR-β (clone H57-597). To stain intracellular antigens, splenocytes were harvested and incubated in RPMI 1640 medium supplemented with cell stimulation and protein transport inhibitor cocktail (eBioscience, catalogue number: 00-4975-03), 10% fetal bovine serum and 100 U/100 μg/ml penicillin-streptomycin (Gibco Life Technologies, NY, USA) for 18 hours. Thereafter, cells were washed with PBS, stained for surface antigens (CD45, CD11b, TCR-β and CD4), fixed and permeabilized (eBioscience, catalogue number: 00-5523-00) and then stained with antibodies against IFN-γ (clone XMG1.2, Biolegend) and FoxP3 (clone JJK-16s, eBioscience). To analyze hematopoietic stem cells, bone marrow suspensions were stained with antibodies against lineage markers (cocktail of Ter119, B220, CD11b, CD3, and GR1), c-Kit (clone 2B8), Sca-1 (clone D7), CD48 (clone HM48-1) and CD150 (clone TC15-12F12.2, all from BioLegend). Data were acquired on an LSR Fortessa (BD Biosciences) and the results were analyzed with FlowJo software (FlowJo, LLC, Ashland, USA).
In Vivo Homing AssaySplenocytes or bone marrow cells from male Apoe-/- and Apoe-/- Mc1re/e mice were harvested and red blood cells were lysed (BD Pharm Lyse™, BD Biosciences) and thereafter, the samples were washed and filtered through a 70-μm cell strainer. Splenocytes and bone marrow cells were stained with eFluor™ 670 (eBioscience, catalogue number: 65-0840-85) or carboxyfluorescein succinimidyl ester (CFSE, Invitrogen™, catalogue number: C34554) at 37°C for 15 min. Cells were washed with RPMI 1640 medium containing 10% FBS, resuspended in PBS and mixed at 1:1 ratio. Ten million splenocytes or bone marrow cells were injected via tail vein into each recipient male Apoe-/- mouse. Twenty-four hours after the injection, mice were sacrificed and blood, spleen and para-aortic lymph nodes were harvested for staining with antibodies against CD45, CD11b, TCR-β, CD4 and CD8 or CD45, CD11b, CD115, Ly6C and Ly6G. Samples were analyzed by flow cytometry (LSR Fortessa, BD Biosciences) and the results are expressed as percentage of injected CD45+ cells.
CD4+ T Cell IsolationSpleens from Apoe-/- and Apoe-/- Mc1re/e mice were harvested and processed into single cell suspensions as previously described (15). Splenic CD4+ T cells were then isolated by positive selection (Invitrogen, catalogue number: 8802-6841-74) and the resulting cell fraction was subjected to total RNA or protein extraction.
Chemotaxis AssaySplenocytes from Apoe-/- and Apoe-/- Mc1re/e mice were harvested and subjected to chemotaxis assay using 6.5 mm Transwell® polycarbonate membrane cell culture inserts (5.0-μm pore size, Corning, NY, USA). In brief, splenocytes were harvested as described above and suspended in RPMI1640 medium supplemented with 0.5% fatty acid-free BSA. Chemokine-containing (murine recombinant CCL3, CCL4 or CCL5, 100 ng/mL, PeproTech, London, UK) migration medium (RPMI-1640 + 0.5% fatty acid-free BSA) was placed in the bottom of the well and thereafter, an aliquot (100 μL, 1x106 cells) of cell suspension was seeded in the top chamber. After 3-hour incubation at 37°C in a 5% CO2 atmosphere, the top chamber was removed and migrated cells were stained (CD45, TCR-β, CD4, CD8, CD115, Ly6C and Ly6G) and counted by flow cytometry. The number of migrated cells is expressed as percentage of CD45+ cells that were seeded in the top chamber.
CCR5 Internalization and Recycling AssaySplenocytes were harvested from Apoe-/- and Apoe-/- Mc1re/e mice and suspended in RPMI1640 medium containing 0.5% fatty acid-free BSA at a density of 1 x 106 cells/mL. After 60 min incubation at 37°C, cells were left untreated or stimulated with recombinant murine CCL5 (400 ng/mL, 60 min, PeproTech) to evoke internalization of CCR5. To evaluate recycling of CCR5, CCL5-stimulated cells were washed three times with RPMI1640 medium, resuspended in RPMI1640 medium containing 0.5% fatty acid-free BSA and incubated for 60 min at 37°C. Cells were harvested, stained (CD45, TCR-β, CD4, CD8 and CCR5, clone HM-CCR5, Biolegend) and analyzed for the expression of cell surface CCR5 by flow cytometry.
RNA Isolation, cDNA Synthesis and Quantitative RT-PCRSpleen, bone marrow and CD4+ T cell samples were isolated and homogenized in QIAzol Lysis Reagent using the Qiagen TissueLyser LT Bead Mill (QIAGEN, Venlo, Netherlands). Total RNA was extracted (Direct-zol RNA Miniprep, Zymo Research, CA, USA) and reverse-transcribed to cDNA with PrimeScript RT reagent kit (Takara Clontech) according to the manufacturer’s instructions. Quantitative real-time polymerase chain reaction (RT-PCR) was performed with SYBR Green protocols (Kapa Biosystems, MA, USA) and a real-time PCR detection system (Applied Biosystems 7300 Real-Time PCR system). Samples were run in duplicate. Target gene expression was normalized to the geometric mean of two housekeeping genes (ribosomal protein S29 and β-actin) using the delta-Ct method and results are presented as relative transcript levels (2-ΔΔCt). Primer sequences are presented in Supplementary Table 1.
Histology and ImmunohistochemistryAortic roots were fixed in 10% formalin overnight followed by embedding in paraffin and cutting into in 4 μm-thick serial sections. Sections were stained with hematoxylin and eosin (H&E) and Masson’s trichrome to measure atherosclerotic plaque area, the size of necrotic core and plaque collagen content at the level of the aortic sinus. For immunohistochemistry, sections were incubated in 10 mM sodium citrate buffer (pH 6) for 20 min in a pressure cooker for antigen retrieval. Thereafter, sections were blocked in 1% H2O2 for 20 min and then in 10% normal horse serum. Samples were incubated overnight with primary antibodies against Mac-2 (1:300, Abcam) or alpha smooth muscle actin (α-SMA, 1:200, Sigma-Aldrich, St. Louis, MO, USA) followed by horseradish peroxidase-conjugated secondary antibody incubation and detection with diaminobenzidine (ABC kit, Vector Labs, Burlingame, USA) to estimate macrophage- and smooth muscle cell-positive plaque areas, respectively. For immunofluorescence, frozen 6 μm-thick spleen sections were stained with antibodies against CD4 (1:50, clone RM4-5, Biotechne) and MC1-R (1:50, Elabscience) followed by detection with fluorochrome-conjugated secondary antibodies (anti-rabbit Alexa Fluor 647 or anti-rat Alexa Fluor 488 Jackson ImmunoResearch, West Grove, USA). For the multicolor immunofluorescence imaging, negative and single-stain controls were included and are presented in Supplementary Figure 1. Sections were counterstained with hematoxylin (CarlRoth) or DAPI (Fluoroshield mounting medium, Abcam), cover-slipped and then scanned with Pannoramic 250 or Pannoramic Midi digital slide scanner (3DHISTECH Kft, Budapest, Hungary). Image analyses was perform using ImageJ software (NIH, Bethesda, MD, USA).
Plasma Cholesterol Assay and Cytokine AnalysisPlasma lipids and lipoproteins were obtained from EDTA-anticoagulated whole blood and measured with commercial enzymatic colorimetric assays (CHOD-PAP and GPO-PAP, mti Diagnostics, Idstein, Germany) according to the manufacturer’s protocols. Plasma total cholesterol concentration was determined. Plasma pro-inflammatory cytokines and chemokines as well as plasma antibodies were quantified with ProcartaPlex™ Multiplex Immunoassays (High Sensitivity 5-Plex Mouse Panel, catalogue number: EPXS050-22199-901, eBioscience & Mouse Antibody Isotyping Panel, catalogue number: EPX070-20815-901, Thermo Fisher Scientific) (34).
Western BlottingCD4+ sorted cell samples were lysed in RIPA containing a protease inhibitor cocktail (Complete Mini, Roche). Aliquots of total protein were separated by SDS-PAGE and transferred to a nitrocellulose membrane. Blots were probed with antibodies against CCR5 (Novus Biologicals, Bio-techne Ltd, UK) and MC1-R (Alomone Labs, Jerusalem, Israel). Horseradish peroxidase-conjugated anti-IgG (Cell Signaling Tech, Frankfurt, DE) secondary anti-body detection was applied and membranes were developed using a chemiluminescence system (ECL detection reagent: Pierce ECL Western (Thermo Scientific, USA).
StatisticsStatistical analyses were performed with GraphPad Prism 8 software (La Jolla, CA, USA). Statistical significance between the experimental groups was determined by unpaired Student’ t-test or two-way ANOVA followed by Bonferroni post hoc tests. The D’Agostino and Pearson omnibus normality test method was employed to test the normality of the data. Possible outliers in the data sets were identified using the regression and outlier removal (ROUT) method at Q-level of 1%. Data are expressed as mean ± standard error of the mean (SEM). Results were considered significant for P<0.05.
ResultsHematopoietic MC1-R Deficiency Induced Leukocytosis in Apoe-/- MiceTo determine the contribution of leukocyte MC1-R to atherosclerosis, we transplanted Apoe-/- or Apoe-/- Mc1re/e BM into lethally-irradiated Apoe-/- recipient mice. After transplantation and recovery period, mice were maintained on chow diet or fed a cholesterol-rich high fat diet (HFD) for 10 weeks to enhance hypercholesterolemia and atherosclerosis. Hematopoietic MC1-R deficiency did not affect body weight or plasma cholesterol concentration in either of the diet groups (Supplementary Table 2). Given the importance of leukocytes and inflammation in the development of atherosclerosis, we did a thorough immunophenotyping for the blood, spleen and BM by flow cytometry. Of note, total leukocyte count was significantly increased in the blood of Apoe-/- Mc1re/e BM transplanted mice (Figures 1A, B). This effect was observed in both diet groups and was largely attributable to increased lymphocyte count (Figure 1C). The numbers of circulating Ly6Chigh monocytes and neutrophils were also elevated (P=0.008 and 0.006, respectively, for genotype effect by 2-way ANOVA) in Apoe-/- Mc1re/e chimeras (Figures 1D, E). Further phenotyping of lymphocyte subsets revealed increased B and CD4+ T cell counts in the blood of Apoe-/- Mc1re/e chimeric mice (Supplementary Figure 2), while the numbers of NK cells, NK T cells and CD8+ T cells were comparable between the experimental groups. In agreement with increased B cell count, we observed an elevation in plasma antibody concentrations particularly in HFD-fed Apoe-/- Mc1re/e chimeric mice (Supplementary Figure 2). This effect was evident across different antibody subclasses including IgG1, IgG2a, IgG2b, IgA and IgM.
Figure 1 Hematopoietic MC1-R deficiency induced leukocytosis in Apoe-/- mice. (A) Representative dot plots for the gating of total leukocytes (CD45+), lymphocytes (CD45+, CD11b-), neutrophils (CD45+, CD11b+, CD115- Ly6G+) and Ly6Chigh monocytes (CD45+, CD11b+, CD115+, Ly6Chigh) in the peripheral blood of Apoe-/- and Apoe-/- Mc1re/e BM transplanted mice. (B–E) Quantification of total leukocyte, lymphocyte, Ly6Chigh monocyte and neutrophil counts in the blood of Apoe-/- and Apoe-/- Mc1re/e BM transplanted mice fed a chow or high-fat diet (HFD) for 10 weeks. Data are mean ± SEM, *P < 0.05, **P < 0.01 versus Apoe-/- mice. Each dot represents individual mouse. SSC-A indicates side scatter area; FSC-A, forward scatter area; Apoe, apolipoprotein E.
To investigate whether induction of hematopoiesis accounts for the observed leukocytosis in Apoe-/- Mc1re/e chimeric mice, total leukocytes and different leukocyte subsets as well as hematopoietic stem and progenitor cells were quantified in the bone marrow. We found that HFD feeding had triggered an expansion of total leukocyte, Ly6Chigh monocyte and neutrophil populations in the bone marrow of Apoe-/- Mc1re/e BM transplanted mice, while the numbers of these cells were unchanged in chow-fed mice (Figures 2A, C, D and Supplementary Figure 3). Chimeric Apoe-/- Mc1re/e mice did not show any change in the frequency of Lin− Sca1+ cKit+ cells (LSK+) that are considered to represent the hematopoietic stem/progenitor cell (HSPC) population (Figures 2E, F). However, the number of true HSPCs that are capable of repopulating the hematopoietic system and called long-term hematopoietic stem cells (HSCs; defined as CD150+ CD48-LSK+) was markedly increased in the bone marrow of Apoe-/-Mc1re/e chimeric mice after feeding HFD (Figure 2G). This was mechanistically associated with down-regulation of ATP-binding cassette transporter ABCA1 (P=0.08), vascular cell adhesion molecule 1 (VCAM-1) and C-X-C motif chemokine ligand 12 (CXCL12) (Supplementary Figure 4), which regulate HSC proliferation and release of progenitor cells into circulation (35–37). On the other hand, chow-fed Apoe-/- Mc1re/e chimeric mice displayed more multipotent progenitors (MPP; defined as CD150- CD48-/low LSK+) (Supplementary Figure 4) that are derived from short-term HSCs and can support hematopoiesis transiently (38). This, in turn, might explain the increase in lymphocyte count that was observed only in chow-fed Apoe-/- Mc1re/e chimeric mice (Figure 2B) and stemmed from higher CD4+ and CD8+ T cell counts (Supplementary Figure 4).
Figure 2 Hematopoietic MC1-R deficiency increased leukocyte and hematopoietic stem cell counts in the bone marrow of Apoe-/- recipient mice. (A–D) Quantification of total leukocytes (CD45+), lymphocytes (CD45+, CD11b-), neutrophils (CD45+, CD11b+, Ly6G+) and Ly6Chigh monocytes (CD45+, CD11b+, CD115+and Ly6Chigh) in the bone marrow of Apoe-/- and Apoe-/- Mc1re/e chimeric mice. (E) Representative dot plots for the gating of LSK+ (Lin-, Sca-1+, c-Kit+), MPP (CD48+, CD150-), HPC (CD48-, CD150-), HSC (CD150+, CD48-) cells in the bone marrow of Apoe-/- recipient mice. (F, G) Quantification of LSK+ and HCS (Lin1-, Sca-1+, Kit+; and CD150+, CD48-) cells in the bone marrow. Data are mean ± SEM, *P ≤ 0.05, **P < 0.01, ***P < 0.001 versus Apoe-/- mice. Each dot represents individual mouse. Lin- indicates lineage-negative; Sca-1, stem-cell antigen-1; c-Kit, proto-oncogene receptor tyrosin kinase; MPP, multi-potent progenitor; HSC, hematopoietic stem cell; HPC, hematopoietic progenitor cell.
Based on the heightened leukocyte count, particularly lymphocyte count, observed in the blood of Apoe-/- Mc1re/e chimeric mice, we next sought to quantify leukocytes and their subsets in the spleen, which constitutes an important source of inflammatory leukocytes infiltrating atherosclerotic plaques. Consistently, flow cytometry analysis revealed an increased leukocyte count in the spleen of Apoe-/- Mc1re/e chimeric mice (Figure 3A and Supplementary Figure 5). This effect was also reflected as higher spleen weight in HFD-fed Apoe-/- Mc1re/e chimeric mice (Supplementary Table 2). The increased leukocyte count was attributable to augmented total lymphocyte count in Apoe-/- Mc1re/e BM engrafted mice (Figure 3B), while Ly6Chigh monocyte and neutrophil counts were comparable between the genotypes (Figures 3C, D). Identification and enumeration of different lymphocyte subsets revealed that MC1-R deficient mice had significantly higher B cell and CD4+ T cell counts in the spleen (Figures 3E, F and Supplementary Figure 6). It is worth noting that CD4+ T cell count was consistently increased in the blood, bone marrow and spleen of Apoe-/-mice receiving MC1-R deficient BM. Since this cell population comprises different effector cells, including Th1, Th2, Th17 and Treg cells, that can have opposing effects on atherosclerosis, we did a qPCR analysis and screened the spleen samples for the signature markers of different CD4+ T cell subtypes. Apoe-/- Mc1re/e chimeric mice displayed upregulation of the Th1 cytokine interferon gamma (IFN-γ) and the transcription factor T-bet that drive the differentiation of CD4+ T cells into Th1 effector cells (Figure 3G). The expression of the Th2 cytokine interleukin 4 (IL-4) was also upregulated but it was not accompanied by transcriptional changes in GATA3 that polarize cells to a Th2 phenotype. None of the Th17 or Treg signature genes were affected in Apoe-/- Mc1re/e chimeric mice. These phenotypic characteristics were further confirmed by flow cytometry analysis that indicated an increase in the proportion of IFN-γ-expressing CD4+ T cells (Figures 3H, I). Additionally, plasma concentrations of the T cell growth factor interleukin 2 (IL-2) and IL-4 were significantly increased in Apoe-/- Mc1re/e chimeric mice on HFD (Figure 3J). IL-6 and IFN-γ concentrations were also moderately elevated (P=0.06) in the plasma (Figure 3J) of these mice. Taken together, bone marrow transplantation of Apoe-/- Mc1re/e BM into Apoe-/- mice expanded CD4+ T cell population and the Th1 effector cells appeared to be the primary CD4+ T cell subtype responsible for this expansion.
Figure 3 Increased leukocyte and lymphocyte counts in the spleen of Apoe-/- mice reconstituted with MC1-R deficient BM. (A–D) Quantification of total leukocytes (CD45+), lymphocytes (CD45+, CD11b-), Ly6Chigh monocytes (CD45+, CD11b+, CD115+and Ly6Chigh) and neutrophils (CD45+, CD11b+, Ly6G+) in the spleen of Apoe-/- recipient mice. (E) Representative dot plots for the gating of NK T cells (CD45+, TCRβ+, NK1.1+), NK cells (CD45+, TCRβ-, NK1.1+), CD4+ T cells (CD45+, TCRβ+, CD4+), CD8+ T cells (CD45+, TCRβ+, CD8+), B cells (CD45+, TCRβ-, CD19+, CD11b-) and CD11b+ B cells (CD45+, TCRβ-, CD19+, CD11b+) in the spleen of Apoe-/- BM transplanted mouse. (F) Quantification of splenic lymphocyte subsets in HFD-fed recipient mice. (G) Quantitative real-time PCR (qPCR) analysis of effector CD4+ T cell markers in the spleen of Apoe-/- and Apoe-/- Mc1re/e BM transplanted mice. n=8-9 mice per genotype. (H, I) Representative flow cytometry results for the gating and quantification of IFN-γ+ and Foxp3+ CD4+ T cells (CD45+, TCRβ+, CD4+) in the spleen of Apoe-/- and Apoe-/- Mc1re/e BM transplanted mice. n=4 mice per genotype. (J) Pro-inflammatory cytokine concentrations in the plasma of chow- and HFD-fed Apoe-/- mice. *P < 0.05, **P < 0.01, ***P < 0.001 versus control Apoe-/- mice. Data are mean ± SEM.
We next aimed to address whether enhanced leukocytosis in Apoe-/- Mc1re/e chimeric mice affects atherosclerosis, and analyzed the development of atherosclerotic lesions at the aortic root. Unexpectedly, Apoe-/- Mc1re/e BM transplantation did not change plaque size in chow- or HFD-fed mice (P=0.10 for genotype effect by 2-way ANOVA) (Figures 4A, C). Further characterization of atherosclerotic lesions in HFD-fed mice did not reveal any significant changes in terms of plaque macrophage (Figures 4B, D) or smooth muscle cell (Figures 4B, E) content as judged by the expression levels of Mac-2 and α-smooth muscle actin (α-SMA). In addition, plaque collagen content and necrotic core area were comparable between the genotypes (Figures 4B, F, G). Lastly, flow cytometric analysis of aortic lysates showed that Apoe-/- Mc1re/e BM transplantation did not result in enhanced accumulation of total leukocytes, lymphocytes, Ly6Chigh monocytes or neutrophils in the aorta (Figure 4H). These findings contradict the observed phenotype in the blood, BM and spleen of Apoe-/- Mc1re/e chimeric mice, characterized by markedly increased leukocyte counts.
Figure 4 Hematopoietic MC1-R deficiency does not affect plaque phenotype but significantly alters homing behavior of T cells. (A) Representative images of hematoxylin and eosin (H&E) staining of the aortic sinus of Apoe-/- and Apoe-/- Mc1re/e BM engrafted mice on chow or HFD for 10 weeks. (B) Representative images of Mac-2 (galectin-3), α-SMA (α-smooth muscle actin), Masson trichrome staining and necrotic core areas in the aortic sinus. Necrotic core areas are indicated with dashed lines in H&E-stained images. (C) Quantification of plaque area in aortic sinuses. (D–G) Quantification of Mac-2- and α-SMA – positive areas, plaque collagen content and acellular necrotic core areas as percentage of total plaque area. (H) Quantification of total leukocytes (CD45+), lymphocytes (CD45+, CD11b-), Ly6Chigh monocytes (CD45+, CD11b+, CD115+, Ly6Chigh) and neutrophils (CD45+, CD11b+, Ly6G+) by flow cytometry in the aorta of Apoe-/- recipient mice. (I) Experimental design for analyzing homing of different lymphocyte subsets to the para-aortic lymph nodes (paLN) and spleen. Cells were isolated from the spleen of Apoe-/- and Apoe-/- Mc1re/e mice and injected into recipient Apoe-/- mice. (J–L) Quantification of CD4+ T cells (CD45+, TCRβ+, CD4+), CD8+ T cells (CD45+, TCRβ+, CD8+) and double positive (DP) T cells (CD45+, TCRβ+, CD4+, CD8+) by flow cytometry in the blood, paLNs and spleen as percentage of injected CD45+ cells. Data are mean ± SEM, **P < 0.01, ***P < 0.001 versus control Apoe-/- mice. Each dot represents individual mouse.
The apparent mismatch between the expected and observed plaque phenotype raised a question whether MC1-R deficient leukocytes have an altered migratory behavior. We therefore performed an in vivo homing assay with CD45+ cells isolated from the spleen of male Apoe-/- and Apoe-/- Mc1re/e mice (Figure 4I). Cells were fluorescently labeled and injected into recipient male Apoe-/- mice and quantified by flow cytometry in the blood, para-aortic lymph nodes (paLNs) and spleen 24 hours after the injection. In the blood and paLNs, we observed a significant reduction in the number of CD4+ and CD8+ T cells that were isolated from Apoe-/- Mc1re/e mice compared to control Apoe-/- cells (Figures 4J, K and Supplementary Figure 7). In contrast, no difference was noted in double positive CD4+ CD8+ T cells (DP T cells) that were used as internal control to confirm that the same effect is not universally appearing in all cell types (Figures 4J–L). Instead of being retained in the blood and homing into paLNs, CD4+ and CD8+ T cells from Apoe-/- Mc1re/e mice were trafficking more into the spleen (Figure 4L). We also performed a similar homing assay using myeloid cell-enriched BM from Apoe-/- and Apoe-/- Mc1re/e mice to track the migration of monocytes and neutrophils (Supplementary Figure 8). Mimicking the behavior of CD4+ and CD8+ T cells, classical Ly6Chigh and patrolling Ly6Clow monocytes from Apoe-/- Mc1re/e mice migrated more readily into the spleen in comparison with the corresponding cell types from Apoe-/- mice (Supplementary Figure 8). Ly6Chigh and Ly6Clow monocytes from Apoe-/- Mc1re/e mice were also retained more in the circulation (Supplementary Figure 8), while in the paLNs, they were detected in equal amounts as control Apoe-/- cells (Supplementary Figure 8). On the other hand, neutrophils did not show any genotype difference in their migratory behavior (Supplementary Figures 8). Collectively, MC1-R deficient CD4+ and CD8+ T cells as well as monocytes preferentially home to the spleen and are thereby less likely to migrate to the aorta.
MC1-R Is Expressed on CD4+ T Cells and Regulates mRNA and Protein Level of the Chemokine Receptor CCR5Having noted significantly increased CD4+ T cell counts and altered homing behavior of these cells in the Apoe-/- Mc1re/e genotype, it raised a question of whether these cells express MC1-R. It has been previously reported that MC1-R is expressed in mouse and human CD8+ T cells but not in CD4+ T cells (25), while another study found high MC1-R mRNA levels particularly in human CD4+ T cells (39). It was therefore crucial to establish the possible presence of MC1-R in CD4+ T cells. Indeed, double immunofluorescence staining revealed localization of MC1-R in CD4+ cells in the murine spleen (Figure 5A). Supporting this finding, Western blot analysis using sorted CD4+ T cells from the spleen demonstrated a clear expression of MC1-R protein in these cells. (Figure 5B). The specificity of the signal was validated by preadsoprtion with a blocking peptide (Figure 5B). Furthermore, MC1-R mRNA level was significantly higher in sorted CD4+ T cell samples compared to whole spleen or liver (Supplementary Figure 9). To find a potential explanation for the different homing behavior, we sorted CD4+ T cells from the spleen of Apoe-/- and Apoe-/- Mc1re/e mice and analyzed the expression of various chemokine receptors and adhesion molecules that are known to regulate lymphocyte migration. Among all screened genes by qPCR, the expression of CCR5 stood out by showing significant upregulation in Apoe-/- Mc1re/e CD4+ T cells (Figure 5C). Likewise, protein expression of CCR5 was increased in Mc1r deficient CD4+ T cells (Figure 5D, E) Given that CCR5 guides T cell recruitment and infiltration into sites of tissue inflammation such as atherosclerotic plaques (40, 41), increased CCR5 mRNA and protein expression in MC1-R deficient CD4+ T cells was unexpected and contradicts the finding of preferential homing of these cells into the spleen. To investigate this issue further, we analyzed CCR5 expression at the cell surface of splenic T cells by flow cytometry. Interestingly, CCR5 surface expression as well as percentage of CCR5+ cells was significantly reduced in Apoe-/- Mc1re/e CD4+ T cells (Figure 5F and Supplementary Figure 9). These changes were not evident in CD8+ T cells from Apoe-/- Mc1re/e mice (Figure 5F and Supplementary Figure 9).
Figure 5 | MC1-R is expressed on splenic CD4+ T cells and its deficiency selectively modulates CCR5 expression. (A) Immunofluorescence staining of MC1-R and CD4 in the mouse spleen. White arrows indicate co-localization of MC1-R and CD4. RP indicates red pulp; WP, white pulp. (B) Western blot analysis of MC1-R protein expression in isolated CD4+ T cell samples from the spleen. The expression of vinculin is shown as loading control. (C) Quantitative real-time PCR (qPCR) analysis of chemokine receptor and adhesion molecule expression in isolated CD4+ T cells from Apoe-/- or Apoe-/- Mc1re/e mice. Lanes on the right were incubated in anti-Mc1r antibody solution that was premixed with a molar excess of a blocking Mc1r peptide (D, E) Representative Western blots and quantification of CCR5 and β-actin (loading control) in isolated CD4+ T cells lysates from Apoe-/- or Apoe-/- Mc1re/e mice. (F) Quantification of CCR5 surface expression by flow cytometry in CD4+ and CD8+ T cells from the spleen of Apoe-/- or Apoe-/- Mc1re/e mice. Data are mean ± SEM, *P < 0.05, **P < 0.01 versus control Apoe-/- mice. Each dot represents individual mouse.
To determine whether the distinct CCR5 expression profile has functional consequences, we employed a chemotaxis assay on isolated splenocytes from Apoe-/- and Apoe-/- Mc1re/e mice and examined the migration of these cells towards the known ligands of CCR5 including CCL3, CCL4 and CCL5. Migration of Apoe-/-Mc1re/e CD4+ T cells towards CCL4 and CCL5 was significantly reduced (Figure 6A). Similar tendencies were also noted for Mc1r deficient CD8+ T cells (Figure 6B). Ly6Chigh monocytes showed most drastic changes and CCL3-induced migration of these cells was particularly blunted in the Apoe-/- Mc1re/e genotype (Figure 6C). Finally, to address whether the intracellular trafficking of CCR5 is compromised due to Mc1r deficiency, isolated and cultured splenocytes were stimulated with CCL5 followed by withdrawal of the ligand to evoke receptor recycling to the cell membrane. The experiment unveiled that CCR5 internalization as well as recycling was impaired in CD4+ T cells from Apoe-/- Mc1re/e (Figure 6D). Likewise, CD8+ T cells from the same mice showed a lack of proper internalization and recycling response (Figure 6E). Overall, these results demonstrate a defect in CCR5 cell surface expression and recycling in the Apoe-/-Mc1re/e genotype, thus proving a possible explanation for the altered migratory behavior of T cells and monocytes.
Figure 6 MC1-R deficient CD4+ T cells show impaired CCR5-dependent migration and recycling of the CCR5 receptor. (A–C) Transwell migration assay using splenocytes from Apoe-/- and Apoe-/- Mc1re/e mice. Cell were allowed to migrate for 3 hours towards the indicated chemokines and migrated CD4+ T cells, CD8+ T cells and Ly6Chigh monocytes were quantified by flow cytometry as percentage of input CD45+ cells. n=4 mice per genotype (D, E) Quantification of CCR5 surface expression on CD4+ cells and CD8+ T cells that were isolated from Apoe-/- or Apoe-/- Mc1re/e mice. CCR5 surface expression was quantified during baseline, after CCL5-induced internalization and after withdrawal of CCL5. n=4 mice per genotype. Data are mean ± SEM., *P < 0.05, **P < 0.01 versus Apoe-/- mice.
The present study shows that MC1-R is intrinsically involved in the regulation of leukocyte production and migration, especially in conditions of excess dietary cholesterol. Hematopoietic MC1-R deficiency drastically increased tissue leukocyte counts in Apoe-/- mice without affecting atherosclerosis. We also found that MC1-R is present in CD4+ T cells and the migration of these cells is disrupted by MC1-R deficiency.
In recent years, MC1-R has been recognized as a promising therapeutic target in various inflammatory diseases including atherosclerosis. Leukocytes, particularly monocytes and their descendant macrophages, contribute to the initiation and progression of atherosclerosis by accumulating in the subendothelial space and by destabilizing atherosclerotic lesions. Most of leukocyte subpopulations have been found to express MC-1R, which activation evokes a multitude of anti-inflammatory and pro-resolving responses (21). The evidence relies mainly on gain-of-function approaches that have utilized either the endogenous ligand α-MSH or more stable, synthetic agonists to address the therapeutic potential of targeting MC1-R for various inflammatory diseases. Owing to the multifaceted role of MC1-R in modulating inflammation, chronic activation of the receptor has been shown to provide protection against atherosclerosis in both pharmacological and genetic models (15, 26, 27). In macrophages, MC1-R has an additional regulatory function by promoting the clearance of excess intracellular cholesterol (15). This happens via induction of the ATP-binding cassette transporters ABCA1 and ABCG1, which initiate macrophage reverse cholesterol transport and prevent atherosclerosis. Although several studies have consolidated the favorable effects of MC1-R activation in leukocytes, little is known about the intrinsic role of MC1-R in these cells. Therefore, it remains largely an open question whether silencing of leukocyte MC1-R carries a reverse phenotype and leads to overproduction and accumulation of leukocyte subsets in tissues. To address this question, we employed bone marrow transplantation to generate a model of hematopoietic MC1-R deficiency and characterized the immunophenotype of this model in the context of atherosclerosis.
Strikingly, hematopoietic MC1-R deficiency led to a generalized increase of total leukocytes in the blood, bone marrow and spleen. This was mainly a result of expanded CD4+ T cell, B cell, monocyte and neutrophil populations and could have derived from induction of hematopoiesis. From a methodological standpoint, LSK+ cell count in the bone marrow was comparable between the genotypes indicating that hematopoietic MC1-R deficiency had not affected reconstitution efficiency following irradiation and transplantation. Likewise, the level of chimerism was comparable between the genotypes. Thus, the observed leukocyte profiles in Apoe-/- Mc1re/e BM transplanted mice were not biased by a difference in BM reconstitution efficiency.
Of note, HFD-fed Apoe-/- Mc1re/e chimeras showed increased frequency of total leukocytes, Ly6Chigh monocytes and neutrophils in the bone marrow. These observations corroborate our earlier study on Apoe-/- mice with global MC1-R deficiency (16). These mice displayed elevated total leukocyte and Ly6Chigh monocyte counts in the bone marrow in response to HFD-induced hypercholesterolemia. Although we did not quantify stem cell count in that study, increased Ly6Chigh monocyte frequency most likely reflects induction of hematopoiesis. It is well-established that hypercholesterolemia causes gradually developing monocytosis as a result of enhanced Ly6Chigh monocyte production and increased survival of these cells in peripheral tissues (6). The development of monocytosis often requires elevation of plasma cholesterol concentration and this could have been the trigger for the monocytosis in Apoe-/- mice with global MC1-R deficiency, which had elevated plasma cholesterol level after HFD feeding. However, in the present study, hematopoietic MC1-R deficiency caused monocytosis and enhanced HSC proliferation in the absence of exaggerated hypercholesterolemia. Some kind of an interaction between the genotype and cholesterol seem to exist since total leukocyte and HSC count were only increased in the bone marrow of HFD-fed Apoe-/- Mc1re/e chimeric mice. Considering that cholesterol efflux pathways are disrupted due to MC1-R deficiency (15, 16), intracellular cholesterol levels might be accumulating in HSCs of Apoe-/- Mc1re/e BM transplanted mice, rendering these cells more sensitive to hematopoiesis-triggering signals. Supporting this notion, we have previously observed reduced expression of ABCA1 and ABCG1 in the bone marrow of Apoe-/- Mc1re/e mice (16), which could explain the hypersensitivity to HFD-induced hematopoiesis. Down-regulation of ABCA1 was also observed in this study along with reduced transcript levels of VCAM-1 and CXCL12, which are known to regulate proliferation and release of HSCs (35, 36). Collectively, these findings suggest that MC1-R is involved in HSC proliferation but it remains to be determined whether HSCs or nonhematopoietic cell types that form the HSC niche such as adipocytes, osteoblastic cells and endothelial cells express MC1-R and could MC1-R on those cells be functionally active and regulate hematopoiesis.
Exaggerated hypercholesteremia can induce myeloid-cell bias and lead to expansion of monocytes and neutrophils in the bone marrow and their release into the circulation (9). This occurs at the expense of suppressed lymphopoiesis, which could explain the present finding of increased bone marrow lymphocyte count in chow-fed Apoe-/- Mc1re/e chimeric mice and the abolishment of this effect after HFD. Nevertheless, hematopoietic MC1-R deficiency was associated with significantly elevated lymphocyte counts in the circulation and spleen regardless of the dietary regimen. This was largely attributable to expansion of CD4+ T cells and B cells, which was accompanied by elevated plasma concentrations of different antibody subclasses as well as IL-2, IL-4 and IFN-γ. We also obtained evidence suggesting that the increase in CD4+ T cells is to some extent confined to pro-atherogenic Th1 effector cells. The principal Th1 cytokine IFN-γ and the transcription factor T-bet were significantly up-regulated in the spleen of Apoe-/- Mc1re/e chimeric mice. Supporting the pro-atherogenic role of these markers, genetic deficiency of IFN-γ or T-bet attenuate lesion formation (42–44) while injection of recombinant IFN-γ accelerates atherosclerosis (45). Given this, combined with the fact that circulating monocyte numbers correlate with atherosclerosis progression, it was unexpected that the increased monocyte and CD4+ T cells did not aggravate atherosclerosis in Apoe-/- Mc1re/e chimeric mice. On the other hand, global deficiency of MC1-R associated with larger and more vulnerable atherosclerotic plaques. However, this phenotype could have been primarily driven by increased plasma cholesterol level that arose as a consequence of disturbed bile acid metabolism (16). Hematopoietic MC1-R deficiency did not affect plasma cholesterol level, which thus allows to dissect the role of leukocyte MC1-R without confounding by a difference in cholesterol level. Even if elevated plasma cholesterol was the decisive factor for the observed phenotype in global MC1-R deficiency, increased numbers of leukocytes and particularly monocytes, neutrophils and CD4+ T cells in the circulation of Apoe-/- Mc1re/e chimeras should have migrated into the atherosclerotic lesion and accelerated disease progression. The lack of effect on arterial leukocyte counts, lesional macrophage coverage and plaque size led us to investigate whether MC1-R deficiency influences the migratory behavior of leukocytes. In vivo homing assay revealed that CD4+ and CD8+ T cells as well as Ly6Chigh monocytes from MC1-R deficient mice were preferentially entering the spleen. It is thought that T cells are trafficking between the spleen and inflamed atherosclerotic arteries through the circulation. Although we were unable to reliably track T cells that had entered the aortic wall, the reduced T cell count in the blood and paLNs suggest that MC1-R deficient cells are less likely to infiltrate aortic adventitia or atherosclerotic plaques.
When exploring the mechanistic explanation for the disturbed migratory behavior of leukocytes, we turned our attention to CD4+ T cells since previous studies have not provided conclusive evidence for the expression of MC1-R on these cells (25, 39). CD8+ T cells, B cells, monocytes and neutrophils, which were also increased in Apoe-/- Mc1re/e BM transplanted mice, have been demonstrated to carry functional MC-1R (24, 25, 46, 47). To our knowledge, this is the first study to prove that MC1-R is present in CD4+ T cells, which opens the possibility that silencing leukocyte MC1-R signaling can directly affect this cell population as well.
We found that MC1-R deficient CD4+ T cells had up-regulated CCR5 mRNA and protein expression, while cell surface CCR5 level was reduced. Furthermore, CCR5-dependent cell migration and recycling of the receptor back to the cell surface after internalization were impaired in MC1-R deficient CD4+ T cells. Likewise, MC1-R deficiency drastically hampered the migration of Ly6Chigh monocytes towards the CCR5 ligand CCL3. These findings imply that MC1-R deficient cells are upregulating CCR5 expression as a compensatory response to a defect in its recycling mechanism. CCR5 is expressed on various leukocytes subsets including monocytes and pro-inflammatory Th1 cells, where it mediates ligand-triggered (CCL3, CCL4 and CCL5) arrest and transendothelial migration (48). Genetic deletion or pharmacological inhibition of CCR5 reduces monocyte and T cell migration into lesions and protects against diet-induced atherosclerosis in Apoe-/- mice (49, 50). Against this background, impairment of CCR5 function is a plausible explanation for the observed plaque phenotype in Apoe-/- Mc1re/e chimeric mice that in the context of enhanced leukocytosis, would have otherwise showed signs of aggravated atherosclerosis. It remains to be still determined what is the exact mechanism behind disrupted CCR5 recycling. It is known that mutations in the MC1-R gene do not often block transcription or translation of the gene product but rather interfere with the intracellular traffic and lead to retention of the misfolded protein (51). Although it is purely speculative, MC1-R and CCR5 might be interacting and forming heteromers, which is a unique feature of GPCRs. Disturbed trafficking of mutated MC1-R might thereby also affect the internalization and recycling of CCR5.
In conclusion, the present study demonstrates that MC1-R is critically involved in the regulation of leukocyte production and migratory behavior. Hematopoietic MC1-R deficiency markedly increased tissue leukocyte counts and induced hematopoiesis in response to excess dietary cholesterol. However, CCR5-dependent migration was impaired in MC1-R deficient leukocytes. Opening a completely new avenue for investigation, we found that MC1-R is also present in CD4+ T cells, which are important for immune responses during host defense and play a central role also as drivers of autoimmune diseases and chronic inflammatory diseases such as atherosclerosis.
Uhohinc
- afamelanotide evaluated as safe in mild to moderate arterial ischaemic stroke (NIHSS 1–15, n=6)
- NIHSS1 scores improved in five patients
- brain scans (MRI-FLAIR2) in all patients show reduction of affected tissue
- strong functional recovery in all five surviving patients
CLINUVEL today released positive final results of the open label pilot study (CUV801) in arterial ischemic stroke (AIS), evaluating multiple doses of CLINUVEL’s drug afamelanotide in six adult patients. Afamelanotide was shown to be well tolerated, with five of the six patients showing considerable clinical and functional recovery up to 42 days after treatment.
“Final analyses from the CUV801 study show that surviving patients who received treatment with afamelanotide all seemed to have recovered well in the six weeks following their brain injury,” CLINUVEL’s Head of Clinical Operations, Dr Pilar Bilbao said. “Our clinical team often publicly emphasise the significance of afamelanotide as a safe drug in patients, and in this study, we obtained further data that patients with longstanding cardiovascular disease seem to tolerate afamelanotide well. The significance of these findings is of benefit to all our current and future programs.”
Study Results CUV801CUV801 is the first clinical study assessing afamelanotide as a treatment for a life-threatening brain injury.
All six patients enrolled in the study carried an increased risk of stroke due to their history of cardiovascular disease, elevated blood pressure or diabetes type II, and all suffered a stroke (blood clot) in the left half of the brain. The study was conducted at the specialist stroke unit of the Alfred Hospital in Melbourne, Australia.
Safety was the primary endpoint of the pilot study with afamelanotide administered up to four times over ten days following the stroke. This frequency of dosing seemed not to affect patient safety, with no drug-related adverse events reported during or after the study completion. One patient with a complex cardiovascular history passed away following a second stroke on day 5, which was assessed as unrelated to afamelanotide treatment.
Treatment efficacy was measured using computer imaging to assess the volume of the area affected by the stroke, and validated clinical assessments of function, neurological impairment, and disability.
Analyses of the brain scans (MRI-FLAIR2) performed at days 3 and 9 showed a reduction in size of the affected area in five of the six patients.
Analyses of the NIHSS scores1 up to day 42 indicated that all five surviving patients showed an improvement in neurological functions and reduction in overall impairment (p=0.0625). Four out of five surviving patients showed an improvement of 4 points or more on the scale, regarded as significant, and all five patients reported a clinically meaningful reduction of 3 points. Two patients were symptom free at day 42.
The modified Rankin Scale, a non-stroke specific tool used to determine global disability, proved not sensitive enough as an instrument for the short study period.
Ischaemic strokes account for around 85% of the estimated 15 million strokes suffered worldwide each year. Stroke is the leading cause of serious, long-term disability in the United States. Considering the staggering prevalence of stroke, the burden of post-stroke recovery and ongoing disability is of primary public health importance.
“Despite the considerable impact of strokes on individuals and society, the treatment options available, even at specialist stroke units, are tragically limited,” Dr Bilbao said. “We are seeking to prove that afamelanotide can provide a safe, effective treatment option which can improve the overall prognosis post-stroke and reduce patient disability long-term.”
“The first steps are to gain comfort that the intervention with afamelanotide poses no harm to patients, while obtaining objective measures of impact of treatment on the course of the patients’ disease. With CUV801 we have achieved both these outcomes and can now pursue further studies and regulatory interactions with a degree of confidence that the drug performs as expected.”
“The gain for stroke patients, but also for society as a whole, lies in the improvement in neurological functions, since the ability to resume independent living saves high costs to our healthcare systems,” Dr Bilbao said.
- The National Institutes of Health Stroke Scale (NIHSS) consists of 15 tests to evaluate neurologic functioning and impairment caused by acute cerebral infarction (stroke). A clinical assessment is made on the basis of consciousness, language, neglect, visual-field loss, extraocular movement, motor strength, muscle control, speech, and sensory loss. A trained clinician assesses the patient’s ability to answer questions and perform specific activities. In general, the evaluation is made in less than 10 minutes.
- The standard diagnosis of stroke patients is made upon hospital admission through computed tomography perfusion (CTP) images to assess the brain damage caused by the clot. The CTP holds some predictive value to assess whether further brain damage will occur if the clot persists. However, days after the stroke, brain scans are made by magnetic resonance (MRI-FLAIR) providing actual information on the extent of brain damage and recovery.
The primary objective of the study CUV801 was to evaluate the safety of patients, who were first time administered afamelanotide within 24 hours of suffering a stroke, while a secondary assessment was made of the recovery of brain tissue calculated from the volume of area affected, neurological function assessments and of the overall disability over 42 days.
Validated evaluations were made using:
- National Institutes of Health Stroke Scale (NIHSS)
Evaluation of the patients’ condition through functional assessment on days 0, 1, 2, 3, 4, 7, 8 and 42. - Brain scans
Brain scans (CTP and MRI-FLAIR) were made at various time intervals (day 0, 3, 9) to assess dead brain tissue and areas at risk of irreversible damage, due to an arterial clot. - Evaluation of disability was made using the modified Rankin Scale (mRS) (pre-stroke assessment on day 0, post-stroke assessment on days 7 and 42).
Scientific progress has demonstrated melanocortins, including afamelanotide, provide a positive effect on the central nervous system (CNS). Afamelanotide is known to offer neuroprotection and act as a potent anti-oxidative hormone. The drug possesses further therapeutic benefits, activating vessels, reducing fluid formation, protecting critical nerve and brain tissue, and restoring the blood brain barrier (BBB: a critical defence mechanism protecting the brain). The drug therapy is thought to improve blood flow and increase the delivery of oxygen and nutrients to deprived brain tissue.
CLINUVEL (ASX: CUV; ADR LEVEL 1: CLVLY; XETRA-DAX: UR9) is a global specialty pharmaceutical group focused on developing and commercialising treatments for patients with genetic, metabolic, systemic, and life-threatening, acute disorders, as well as healthcare solutions for the general population. As pioneers in photomedicine and the family of melanocortin peptides, CLINUVEL’s research and development has led to innovative treatments for patient populations with a clinical need for systemic photoprotection, DNA repair, repigmentation and acute or life-threatening conditions who lack alternatives.
CLINUVEL’s lead therapy, SCENESSE® (afamelanotide 16mg), is approved for commercial distribution in Europe, the USA, Israel and Australia as the world’s first systemic photoprotective drug for the prevention of phototoxicity (anaphylactoid reactions and burns) in adult patients with erythropoietic protoporphyria (EPP). Headquartered in Melbourne, Australia, CLINUVEL has operations in Europe, Singapore and the USA. For more information, please go to https://www.clinuvel.com.
SCENESSE®, PRÉNUMBRA®, and NEURACTHEL® are registered trademarks of CLINUVEL.
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