Advanced glycation end products upregulate lysyl oxidase and endothelin-1 in human aortic endothelial cells via parallel activation of ERK1/2–NF-κB and JNK–AP-1 signaling pathways

Abstract

Endothelial dysfunction involves deregulation of the key extracellular matrix (ECM) enzyme lysyl oxidase (LOX) and the vasoconstrictor protein, endothelin-1 (ET-1), whose gene expression can be modulated by the transcriptional activators nuclear factor kappa B (NF-κB) and activator protein-1 (AP-1). Advanced glycation end products (AGEs) present an aggravating factor of endothelial dysfunction which upon engagement to their receptor RAGE induce upregulation of mitogen-activated protein kinases (MAPKs), leading to NF-κB and AP-1 potentiation. We hypothesized that AGEs could induce NF-κΒ- and AP-1-dependent regulation of LOX and ET-1 expression via the AGE/RAGE/MAPK signaling axis. Western blot, real-time qRT-PCR, FACS analysis and electrophoretic mobility-shift assays were employed in human aortic endothelial cells (HAECs) following treatment with AGE-bovine serum albumin (AGE-BSA) to investigate the signaling pathway towards this hypothesis. Furthermore, immunohistochemical analysis of AGEs, RAGE, LOX and ET-1 expression was conducted in aortic endothelium of a rat experimental model exposed to high- or low-AGE content diet. HAECs exposed to AGE-BSA for various time points exhibited upregulation of LOX and ET-1 mRNA levels in a dose- and time-dependent manner. Exposure of HAECs to AGE-BSA also showed specific elevation of phospho(p)-ERK1/2 and p-JNK levels in a dose- and time-dependent fashion. AGE administration significantly increased NF-κΒ- and AP-1-binding activity to both LOX and ET-1 cognate promoter regions. Moreover, LOX and ET-1 overexpression in rat aortic endothelium upon high-AGE content diet confirmed the functional interrelation of these molecules. Our findings demonstrate that AGEs trigger NF-κΒ- and AP-1-mediated upregulation of LOX and ET-1 via the AGE/RAGE/MAPK signaling cascade in human endothelial cells, thus contributing to distorted endothelial homeostasis by impairing endothelial barrier function, altering ECM biomechanical properties and cell proliferation.

Introduction

Vascular complications affect most human organs contributing to accelerated tissue aging as well as to a wide range of pathological entities. Endothelial dysfunction that involves functional alterations of the normal endothelial phenotype of arteries is a major contributor to vascular aging, diabetes and cardiovascular disease [1, 2]. Several molecular mechanisms have been proposed to underlie endothelial dysfunction and contribute to the impairment of vascular homeostasis including loss of vasodilators such as prostacyclin and nitric oxide (NO), generation of vasoconstrictors (e.g., endothelin-1, ET-1), expression of adhesion molecules (e.g., vascular cell adhesion molecule-1, VCAM-1 and intercellular adhesion molecule-1, ICAM-1), deregulation of inflammation, coagulation factors and cell growth [3, 4].

Glycation as another aggravating factor of endothelial dysfunction refers to the non-enzymatic reaction of reducing sugars with the amino groups of proteins, lipids and nucleic acids leading to the formation of irreversible end products known as advanced glycation end products (AGEs). Formation of these protein adducts can also occur from exogenous sources (mainly diet) with a negative impact to human health.

The role of AGEs and their precursors in causing endothelial dysfunction has been previously addressed with excessive AGE levels in circulation and tissue deposition being implicated in the underlying inflammatory and oxidative stress processes, augmentation of cytokines production and glycation of low- and high-density lipoproteins (LDL and HDL, respectively) [5]. However, AGE-induced molecular targets and associated intracellular pathways that impair endothelial function have not been fully elucidated. It has been demonstrated that AGE adducts can affect the structure and function of extracellular matrix (ECM) components as well as induce intracellular signaling through the receptor for AGEs (RAGE), a member of the immunoglobulin superfamily of cell surface molecules [6]. Upon receptor engagement several key signal transduction pathways, mainly the mitogen-activated protein kinase (MAPK) pathways (extracellular signal-regulated kinase, ERK; c-Jun N-terminal kinase, JNK and p38) are known to be activated in endothelial cells, monocytes and vascular smooth muscle cells (VSMCs). This is followed by potentiation of the respective proinflammatory transcription factor nuclear factor kappa B (NF-κB), leading to upregulation of proinflammatory and oxidative stress mediators, and ultimately deregulation of endothelial homeostasis with detrimental vascular effects [5, 7–11]. Moreover, the activator protein-1 (AP-1) transcription factor can also be activated via the AGE/RAGE/MAPK signaling axis and in concert with NF-κB induces expression of proinflammatory genes such as cyclo-oxygenase-2 (COX-2), interleukin-1 beta (IL-1β) and tumor necrosis factor alpha (TNFα) [12–14].

A key enzyme that participates in the maintenance of ECM stability and the elastic properties of connective tissue is lysyl oxidase (LOX), a copper-dependent amine oxidase that catalyzes the covalent cross-linking of ECM components such as collagen and elastin [15]. LOX downregulation has been associated with endothelial dysfunction observed in early stages of atherosclerosis [16–18] and has been linked to hypoxia-induced vascular endothelial growth factor (VEGF) regulation as well as promotion of angiogenesis in tumor cells [19, 20]. On the other hand, LOX upregulation has been associated with advanced atherosclerosis, neointimal hyperplasia and restenosis that follow proliferation and migration of VSMCs [15, 21].

ET-1 is a well-known endothelial dysfunction biomarker that acts as a vasoconstrictor and regulates cell proliferation, apoptosis, vascular remodeling and fibrosis. ET-1 exhibits powerful functional and physical association with NO and has been linked to a variety of pathological mechanisms in the vasculature, chiefly presenting increased expression in atherosclerosis and diabetic vascular complications [22, 23].

Previous studies have shown a connection of increased AGE levels to LOX and ET-1 expression in various cell types. High AGEs have been shown to stimulate LOX activity and subsequent collagen deposition in ovarian tissue of polycystic ovary syndrome (PCOS) [24], a clinical entity that has been strongly associated with endothelial dysfunction. Additionally, high glucose intake seems to increase LOX expression in rat retinal endothelial cells [25]. Furthermore, elevated serum AGEs have a strong positive correlation with ET-1 levels in PCOS [26] while exogenous administration of AGEs has been shown to induce ET-1 transcriptional activation in bovine aortic endothelial cells (BAECs) in an NF-κB-dependent manner [27, 28].

Consistently, computational analysis has revealed that LOX and ET-1 gene promoters harbor putative NF-κB- and AP-1-binding sites [24, 28–30]. Therefore, it is tempting to speculate that AGEs may trigger the ERK1/2–NF-κB and JNK–AP-1 signaling cascades ultimately leading to modulation of LOX and ET-1 target-gene expression.

Taken together all the above evidence, the aim of our study was to explore the potential impact of AGE/RAGE/MAPK signaling axis on the regulation of LOX and ET-1 gene expression in human aortic endothelial cells (HAECs). An experimental in vivo rat model exposed to high and low in AGEs content diet was further evaluated to establish functional relations of these molecules.

Materials and methods

General

All reagents were purchased from Life Technologies (Carlsbad, CA, USA) unless otherwise stated. AGE-bovine serum albumin (AGE-BSA) was obtained from Abcam plc (ab51995; Cambridge, UK). Following the manufacturer’s instructions, glycated BSA was produced by reacting BSA with glycoaldehyde under sterile conditions followed by extensive dialysis and purification steps. Fluorescence of AGEs was confirmed by fluorescence spectrophotometry with Ex./Em. = 370/440 nm. Glycated BSA showed 7000 % increase in fluorescence when compared to control BSA. The purity of the final stock is >95 % as analyzed by SDS-PAGE and filter sterilized using a 0.22-μm filter.

Cell culture media (including fetal bovine serum, FBS) were purchased from Biochrom (Berlin, Germany) while inhibitors PD98059 and SP600125 were obtained from Sigma Aldrich (P215; Steinheim, Germany) and Cayman Chemical (10010466; Ann Arbor, MI, USA), respectively. Antibodies against LOX (goat polyclonal, sc-32410), ET-1 (goat polyclonal, sc-21625), phospho(p)-ERK (mouse monoclonal, sc-7383), p-c-Jun (mouse monoclonal, sc-822), NF-κB p65 (mouse monoclonal, sc-8008) were obtained from Santa Cruz Biotechnologies (Santa Cruz, CA, USA). The anti-p-JNK rabbit monoclonal antibody (ab76572) was provided by Abcam plc (Cambridge, UK). The anti-AGE (clone 6D12) mouse monoclonal and anti-RAGE goat polyclonal (RDI-RAGEabG) were obtained from Fitzgerald Industries Int (Acton, MA, USA). The anti-RAGE blocking (neutralizing) antibody (goat polyclonal, sc-8230) was purchased from Santa Cruz Biotechnologies. The anti-Actin mouse monoclonal (MAB-1501) was from Millipore (Bedford, MA, USA), the anti-mouse IgG-HRP conjugate goat polyclonal (12–349) and anti-rabbit IgG-HRP conjugate goat polyclonal (12–348) were from Sigma Aldrich (Steinheim, Germany). The anti-goat IgG-FITC conjugate rabbit polyclonal (F2016) was from Research Diagnostics (Concord, MA, USA).

Cell culture

Human aortic endothelial cells were obtained as cryopreserved cells from European Collection of Cell Cultures (ECACC) and were cultured in endothelial cell growth medium M200 supplemented with Low Serum Growth Supplement containing FBS (2 % v/v), hydrocortisone (1 g/mL), human epidermal growth factor (10 ng/mL), basic fibroblast growth factor (3 ng/mL), heparin (10 g/mL), gentamicin (50 μg/mL) and amphotericin B (50 ng/mL). Cells were cultured at 37 °C in a humidified 95 % air–5 % CO₂ atmosphere and were split according to standard procedures. HAECs were used between passage four and eight in all assays. HAECs were treated with variable concentrations of AGE-BSA (100, 200 μg/mL) for several time points (1, 6, 12, 24, 72 h). Specific inhibitors SP600125 (dissolved in DMSO; 20 μM), PD98059 (dissolved in DMSO; 50 μM) and anti-RAGE blocking antibody (dilution 1:100) were also used for treatment of the cells (for 90 min) prior to incubation with AGE-BSA. Control HAECs in these experiments were pre-treated with DMSO alone or goat anti-serum, respectively, for 90 min.

Real-time quantitative PCR

Real-time quantitative PCR was performed using an iCycler real-time instrument (Biorad). RT-PCR product was amplified using the iQ SYBR Green Supermix (Biorad) in a total reaction volume of 20 μL. Primer efficiencies were calculated from a standard curve of serially diluted cDNA. Product identity was confirmed by a single pick in the melt curve. Relative expression values were calculated using the $2^{-(\mathrm{\Delta }\mathrm{\Delta }{C}_{\text{t}})}$ formula. The data are presented as fold change in gene expression normalized to GAPDH and relative to the untreated control. The primers used for the amplification of ET-1 were: forward primer: 5′-CCAAGGAGCTCCAGAAACAG-3′ and reverse primer: 5′-GATGTCCAGGTGGCAGAAGT-3′; for the amplification of LOX: forward primer: 5′-CCAGAGGAGAGTGGCTGAAG-3′ and reverse primer: 5′-CCAGGTAGCTGGGGTTTACA-3′; and for the amplification of GAPDH: forward primer: 5′-GGGTGTGAACCATGAGAAGT-3′ and reverse primer: 5′-CATGCCAGTGAGCTTCCCGTTC-3′. All primer pairs were designed using Primer3 software [31].

Western immunoblotting

For immunoblot analysis cells were solubilized with ice-cold RIPA buffer (Thermo Scientific, Rockford, IL, USA) supplemented with protease inhibitor cocktail (Thermo Scientific). The protein concentration in the lysates was determined by using Bradford assay (Biorad). Equal amounts of total protein were resolved by SDS-PAGE and immunoblotted with anti-p-ERK (dilution 1:200), anti-p-JNK (dilution 1:5000) and anti-Actin (dilution 1:5000) antibodies. Relative protein amounts were evaluated by a densitometric analysis using Image J software and normalized to the corresponding actin levels. All experiments were performed at least three times and representative results and the corresponding quantification data of one experiment are shown.

Flow cytometry analysis

For the assessment of RAGE expression, HAECs were seeded in 6-well plates at a density of approximately 2 × 10⁵ cells per well and then treated with AGE-BSA at a concentration of 200 μg/mL for 72 h. Cells were then harvested, rinsed twice with PBS, collected by centrifugation at 2000 rpm for 5 min and stained with anti-RAGE (dilution 1:400) antibody. Fluorescence labeling was performed in a second step with anti-goat IgG-FITC conjugate (dilution 1:1000) antibody. Samples were scanned with a fluorescence-activated cell sorter (FACSCalibur, Becton–Dickinson, USA) and the data were analyzed with CellQuest software (Becton–Dickinson). A total of 10,000 events were measured per sample. Each experiment was performed in triplicate.

Electrophoretic mobility-shift assay (EMSA)

Nuclear protein extracts from HAECs (30–40 µg) untreated or treated with AGE-BSA at a concentration of 200 μg/mL for 72 h were mixed with 50 fmol of biotin-labeled probe in a total reaction volume of 20 µL containing: 10 mM Tris, 170 mM KCl, 0.5 mM MgCl₂ (for the reactions with the AP-1 probe), 2.5 % glycerol, 0.04 % NP40, 50 ng/μL poly(dI·dC), 2 mM EDTA, 2.6 mM DTT, 50 mM NaF, 1 mM PMSF, 1 mM Na₃VO₄ and 0.5 μL protease inhibitor cocktail (Sigma Aldrich). Nuclear protein extracts from HAECs treated with AGE-BSA were also incubated with antiserum (1 µL of anti-NF-κB p65 or anti-p-c-Jun antibody) and the reaction mixture was left for 30 min at room temperature prior to addition of the labeled probe. Reactions were allowed to equilibrate for 25 min at room temperature. DNA–protein complexes were resolved on 5 % native polyacrylamide gels in 1× Tris–borate-EDTA (TBE) buffer (125 V, room temperature) and electrotransferred onto nylon membrane at 380 mA (~100 V) for 60 min. The biotin-labeled DNA complex was detected by chemiluminescence using the LighShift Chemiluminescence EMSA kit (Pierce Biotechology, Rockford, IL, USA).

In vivo experimental model

Thirty (30) female Wistar rats at 12–20 weeks of age were allocated to the study, and were equally subdivided in two groups. Subgroup A (n = 15) was fed commercial chow low in AGEs content (LA) for 3 months, while subgroup B (n = 15) was fed commercial chow high in AGEs content (HA) for the same period.

The animals were housed four to five per cage under controlled conditions (21–22 °C, 55–65 % humidity, 12-h light/12-h dark cycle) and were given pelleted food and water ad libitum at ELPEN S.A, Experimental-Research Center, Pikermi-Athens, Greece. Animal care and experimental procedures conformed to the “Guide for the Care and Use of Laboratory Animals” (Department of Health, Education and Welfare, Athens, Greece) and were approved by the Institutional Animal Care and Use Committee.

The diets used were derived from a single standard rat chow (AIN-93G) purchased from Bioserve (Frenchtown, NJ, USA), consisting of 18 % protein, 58 % carbohydrate, 7.5 % fat and 3.73 kcal/g. Regular AIN-93G chow is normally prepared by heating at 190 °C for 30 min. Analysis of this preparation showed that it contained 76.0 ± 15.3 mg carboxymethyllysine (CML)/100 g sample (or 436.9 ± 88.1 mg CML/100 g protein), 205.32 ± 22.25 mg fructoselysine/100 g sample (or 1.179.98 ± 127.90 mg fructoselysine/100 g protein) and 52.68 ± 5.71 mg furosine/100 g sample (or 302.78 ± 32.82 mg furosine/100 g protein), and it was considered as high-AGE (HA) diet.

The same rodent mix was also prepared without heating. This preparation was of equivalent macro- and micronutrient and energy content but contained 1.3 ± 0.4 mg CML/100 g sample (or 7.7 ± 2.2 mg CML/100 g protein), 104.58 ± 3.08 mg fructoselysine/100 g sample (or 601.01 ± 17.7 mg fructoselysine/100 g protein) and 26.83 ± 0.79 mg furosine/100 g sample (or 154.22 ± 4.54 mg furosine/100 g protein), and it was considered as low-AGE (LA) diet.

Body weight was monitored weekly. The study was concluded after 3 months and rats were killed with administration of 20 mg/mL xylazine hydrochloride and 100 mg/mL ketamine hydrochloride, under anesthesia with ether, allowing blood sample collection and tissue retrieval.

Biochemical and hormonal assays

Insulin was quantified using ELISA immunoassays purchased by Biovendor Laboratory Medicine (Czech Republic). Serum AGE levels (U/mL) were measured by CML-specific competitive ELISA as described previously. Serum glucose, fructosamine, total cholesterol and triglycerides were measured by spectrophotometry using commercially available kits (Bayer Hellas).

Immunohistochemical analysis

Paraffin-embedded sections of formalin-fixed thoracic aortas were deparaffinized by xylene and dehydrated in graded ethanol. Sections were treated in 3 % hydrogen peroxide in phosphate-buffered saline (PBS) for 15 min and then rinsed in PBS. To increase the immunoreactivity of AGEs, the sections were placed in 500 mL of 0.01 M citric acid buffered solution (pH 7.0) and microwaved at 500 W for 5 min. After thorough washing, the sections were incubated with normal rabbit serum for 20 min at room temperature to avoid nonspecific binding of the antibodies. The sections were then incubated overnight at 4 °C with the anti-AGE monoclonal antibody 6D12 (dilution 1:50), anti-RAGE (dilution 1:400), anti-LOX (dilution 1:100), anti-ET-1 (dilution 1:100) in PBS containing 1 % BSA. Immunoreactivity was detected by the streptavidin–biotin–peroxidase method according to the manufacturer’s protocol. The final reaction product was visualized with 3,3′-diaminobenzidine tetrahydrochloride (LSAB detection kit; Dako, Carpinteria, CA, USA). Lung tissue sections from diabetic rats were used as positive controls for AGE antibody. Negative controls (e.g., rat aortic tissue in which the primary antibody was substituted with nonimmune mouse or goat serum) were also stained in each run. The percentage of positive cells was estimated using light microscopy. The evaluation of the immunostained slides was performed blindly and independently by two pathologists. AGE, RAGE, LOX and ET-1 expression was evaluated in four levels according to the percentage of positive cells as follows: minimum (staining 1–10 % of cells), low (staining 11–30 % of cells), moderate (31–60 % of cells) and extensive (more than 60 % of cells) immunostaining. The staining intensity was also assessed in four levels: 1, very weak; 2, weak; 3, moderate; and 4, strong. H-score was estimated by multiplying the intensity score with the percentage of the immunostained cells.

Statistical analysis

The data are expressed as mean ± standard error (SE) of mean and data analysis was performed using the SPSS 18.0 software for Windows. Two-way analysis of variance (ANOVA) was used for comparison between parameters and when P < 0.05 the LSD test was used to analyze differences among groups. The Spearman rank correlation test was performed to examine the associations between parameters tested. Statistically significant difference was defined as a P value <0.05.

Results

LOX and ET-1 expression levels in AGEs-treated HAECs

Earlier studies have demonstrated a positive impact of AGEs on upregulation of LOX activity in human ovarian cells while high glucose levels augment LOX expression in rat retinal endothelial cells [24, 25]. Moreover, AGEs induce ET-1 transcriptional activation in BAECs and a strong positive correlation has been observed between AGEs and ET-1 in PCOS [26–28].

Consistently, we investigated whether LOX and ET-1 expression in HAECs may be affected by AGEs at mRNA level. To this end, confluent HAECs were incubated overnight with starvation medium and then subjected to variable concentrations of AGE-BSA (100, 200 μg/mL) at different time points (1–72 h) (Fig. 1a, b). Total RNA was isolated from HAECs and subjected to quantitative real-time PCR analysis. As depicted in Fig. 1a, b, HAECs exposed to AGE-BSA for various time points displayed upregulation of LOX and ET-1 mRNA levels in a dose- and time-dependent manner (P < 0.05). Total RNA isolated from untreated HAECs cultivated for the entire 72-h time course was used as control. Pre-incubation of cells with anti-RAGE blocking (neutralizing) antibody (dilution 1:100) for 90 min prior to treatment with AGE-BSA (200 μg/mL) for 72 h revealed a significant decrease of LOX and ET-1 mRNA levels (Fig. 1c, d), confirming that RAGE was mediating the observed changes in gene expression. Total RNA isolated from HAECs pre-incubated with goat anti-serum prior to AGE-BSA treatment (72 h) was used as control.

Fig. 1.

LOX and ET-1 mRNA levels following treatment of human aortic endothelial cells (HAECs) with increasing concentrations of AGE-BSA at different time points. LOX (a) and ET-1 (b) elevated mRNA levels following treatment with AGE-BSA in a time- and dose-dependent manner. Control, untreated HAECs cultivated for the entire 72-h time course. LOX (c) and ET-1 (d) mRNA downregulation following pre-incubation with ERK and JNK inhibitors (PD98059—50 μM and SP600125—20 μM, respectively) as well as with anti-RAGE blocking (neutralizing) antibody (dilution 1:100) for 90 min. Control, HAECs pre-treated with DMSO alone or goat anti-serum, respectively, for 90 min prior to incubation with AGE-BSA (200 μg/mL, 72 h). *P < 0.05, **P < 0.01, ***P < 0.001

ERK1/2 and JNK status in AGEs-treated HAECs

Previous work in endothelial cells has implicated the activation of MAPK/JNK signaling pathways by AGEs [32–35]. Based on these data, we pre-treated HAECs with ERK1/2 and JNK inhibitors (PD98059—50 μM, SP600125—20 μM, respectively), for 90 min prior to incubation with AGE-BSA (200 μg/mL) at the time point of maximum response (72 h). Upregulation of LOX and ET-1 mRNA expression was fully abolished in the presence of inhibitors indicating the involvement of these kinases in AGE-induced regulation of LOX and ET-1 in human aortic cells (P < 0.05) (Fig. 1c, d).

To further validate the above result, lysates from HAECs exposed to variable AGE-BSA concentrations for various time points were analyzed by western immunoblotting employing specific antibodies against the activated (i.e. phosphorylated, p) forms of ERK1/2 and JNK (p-ERK1/2 and p-JNK, respectively). Exposure of HAECs to different concentrations of AGE-BSA for variable time points showed augmentation of the p-ERK1/2 species in a dose- and time-dependent manner, starting at 1 h and remaining elevated up to 72 h (Fig. 2a). P-JNK levels were increased at 6 h post exposure and displayed a progressive elevation up to 72 h (Fig. 2a). Densitometric analysis data confirmed the immediate-early elevation of p-ERK1/2 levels compared to p-JNK levels upon AGE-BSA treatment (Fig. 2a). Pre-incubation of HAECs with ERK1/2 and JNK inhibitors (PD98059 and SP600125, respectively), prior to treatment with AGE-BSA (200 μg/mL) for 72 h reduced significantly both p-ERK1/2 and p-JNK species, implicating these two kinases in AGE-induced intracellular signaling in HAECs (Fig. 2b). HAECs pre-incubated with solvent alone (DMSO) prior to AGE-BSA treatment (72 h) was used as control. Pre-incubation of cells with anti-RAGE blocking (neutralizing) antibody (dilution 1:100) for 90 min prior to treatment with AGE-BSA (200 μg/mL) for 72 h showed a significant reduction in the abundance of p-ERK1/2 and p-JNK species (Fig. 2c), verifying that RAGE was mediating the observed changes in ERK1/2- and JNK-funneled signaling. HAECs pre-incubated with goat anti-serum prior to AGE-BSA treatment (72 h) was used as control.

Fig. 2.

Effect of AGE-BSA administration on p-ERK1/2 and p-JNK levels. a Western immunoblotting and corresponding densitometric analysis showing abundance of p-ERK1/2 and p-JNK in human aortic endothelial cells (HAECs) after treatment with increasing concentrations of AGE-BSA (100, 200 μg/mL) at various time points (1–72 h). Control, untreated HAECs cultivated for the entire 72-h time course. b Western immunoblotting and corresponding densitometric analysis showing specificity of AGE-BSA-mediated p-ERK1/2 and p-JNK activation (200 μg/mL, 72 h) after pre-treatment with p-ERK1/2 and p-JNK inhibitors (PD98059—50 μM and SP600125—20 μM, respectively). Control, HAECs pre-incubated with solvent alone (DMSO) prior to AGE-BSA treatment (200 μg/mL, 72 h). c Western immunoblotting and corresponding densitometric analysis showing reduction in p-ERK1/2 and p-JNK abundance after pre-incubation of cells with anti-RAGE blocking (neutralizing) antibody (dilution 1:100) for 90 min prior to treatment with AGE-BSA (200 μg/mL) for 72 h. Control, HAECs pre-incubated with goat anti-serum prior to AGE-BSA treatment (200 μg/mL, 72 h). d Detection of RAGE protein expression in HAECs by FACS analysis before and after AGE-BSA administration (200 μg/mL, 72 h). Representative results and corresponding quantification data of one experiment are shown

RAGE protein expression in HAECs

In-as-much-as RAGE is the only receptor of AGEs that mediates intracellular signaling in several cell types including endothelium, we monitored RAGE expression in untreated HAECs using FACS analysis (Fig. 2d). The relative protein expression levels of RAGE were not altered considerably after exposure of cells to 200 μg/mL AGE-BSA for 72 h, suggesting the absence of an autoregulatory feedback mechanism of AGEs upon RAGE (Fig. 2d).

Regulation of LOX and ET-1genes by AGEs via NF-κB and AP-1 transcription factors

The upregulation of LOX and ET-1 by AGEs in human endothelial cells prompted us to investigate the existence of a possible interaction between the AGE-induced transcription factors NF-κB, AP-1 and LOX, ET-1 promoter regions. Based on computational analysis (Genomatix, TFSEARCH) and literature [28, 29], putative NF-κB- and AP-1-binding sites are present in the 5′-regulatory region of the ET-1 promoter in the vicinity of the transcription start site (Fig. 3a). Similarly, our previous work has revealed two functional NF-κB- and AP-1-binding sites in the 5′-regulatory region of the LOX promoter near the transcription start site [24] (Fig. 3a).

Fig. 3.

Binding of AGE-induced transcription factors NF-κΒ and AP-1 on EΤ-1 and LOX gene promoter regions. a Schematic diagram depicting NF-κB and AP-1 functional binding sites in LOX and ET-1 promoter regions. b Electrophoretic mobility-shift assays (EMSAs) showing formation of NF-κB- (i, iii) and AP-1-containing complexes (ii, iv) using respective ET-1- (i, ii) and LOX-derived probes (iii, iv) after treatment with AGE-BSA (200 μg/mL, 72 h). Lanes 1, 6, 7, 12 free probe; lanes 2, 8 nuclear extracts from untreated HAECs; lanes 3, 9 nuclear extracts from AGE-BSA treated HAECs; lanes 4, 11 nuclear extracts from AGE-BSA treated HAECs pre-incubated with anti-NF-κB antibody; lanes 5, 10 nuclear extracts from AGE-BSA treated HAECs pre-incubated with anti-p-c-Jun antibody

Towards this end, biotin-labeled double-stranded oligonucleotides derived from the human LOX and ET-1 promoters encompassing the NF-κB- and AP-1-binding motifs, respectively, were used as probes in a standard EMSA employing nuclear-cell protein extracts from untreated and AGE-BSA-treated HAECs. A concentration-dependent DNA–protein complex was obtained for all binding reactions (Fig. 3b, i–iv). The specificity of complex formation was verified by competition assays in all cases (data not shown).

To demonstrate the presence of NF-κB p65 and p-c-Jun (phosphorylated hence activated component of AP-1) in the above specific DNA–protein complexes, anti-NF-κB p65 and anti-p-c-Jun antibodies were included in the corresponding reaction mixtures prior to addition of the labeled probe (Fig. 3b). Pre-incubation with the anti-NF-κB p65 antibody hampered complex formation with ET-1- and LOX-derived NF-κB probes, reducing drastically the abundance of complexes in extracts from AGE-BSA-treated HAECs (85.31 and 97.92 %, respectively) vs. the decrease observed following pre-incubation with non-specific anti-p-c-Jun antibody (47.01 and 64.15 %, respectively) (Fig. 3b, i, iii). In corroboration, pre-incubation with the anti-p-c-Jun antibody impaired complex formation with the ET-1- and LOX-derived AP-1 probes (55.02 and 64.65 %, respectively) vs. the reduction noticed following pre-incubation with non-specific anti-NF-κΒ antibody (27 % increase and no change, respectively) (Fig. 3b, ii, iv). These data provide evidence that NF-κB and AP-1 are engaged in the transcriptional control of LOX and ET-1 expression.

Immunohistochemical analysis of LOX and ET-1 protein expression in aortic endothelium of low- and high-AGE-fed animals

In order to examine the functional relevance of the above data, we evaluated the effect of glycative stress over LOX and ET-1 protein expression levels in thoracic aortas of rats exposed to low (LA) or high in AGEs (HA) diet for 3 months. Physiological parameters and blood analyses of the rats are listed in Table 1. HA animals presented a distorted metabolic and lipid profile, which is known to modulate endothelial function compared to LA. More specifically, higher levels of serum AGEs (6.72 ± 0.26 U/mL) were observed in HA animals compared to LA (4.48 ± 0.24 U/mL; P < 0.0001). Furthermore, HA animals exhibited higher insulin levels (1.96 ± 0.14 vs. 1.22 ± 0.04 μU/mL, respectively; P < 0.0001), elevated total cholesterol (45.4 ± 3.0 vs. 36.8 ± 2.0 mg/dL, respectively; P = 0.014) and increased triglycerides (54.6 ± 3.0 vs. 44.6 ± 1.3 mg/dL, respectively; P = 0.004) compared to LA. Notably, elevated C-reactive protein (CRP) levels indicative of inflammation were also present in HA animals compared to LA counterparts (356.3 ± 11.1 vs. 176.3 ± 10.9 μg/mL, respectively; P < 0.0001).

Table 1.

Somatometric, metabolic and biochemical parameters of the two study groups

Parameters	Group LA (mean ± SE)	Group HA (mean ± SE)	P value (LA vs. HA comparison)a
Body weight at baseline (g)	197.0 ± 4.3	199.7 ± 4.2	0.851
Body weight at 3 months (g)	206.0 ± 4.5	193.3 ± 4.4	0.050
Serum glucose (mg/dL)	119.0 ± 7.9	112.2 ± 6.6	0.548
AGEs (U/mL)	4.48 ± 0.24	6.72 ± 0.26	<0.0001*
Fructosamine (μmol/L)	201.2 ± 33.1	159.4 ± 8.8	0.663
Insulin (μU/mL)	1.22 ± 0.04	1.96 ± 0.14	<0.0001*
CRP (μg/mL)	176.3 ± 10.9	356.3 ± 11.1	<0.0001*
Total cholesterol (mg/dL)	36.8 ± 2.0	45.4 ± 3.0	0.014*
Serum triglycerides (mg/dL)	44.6 ± 1.3	54.6 ± 3.0	0.004*

Values are presented as mean ± standard error (SE) of mean

LA low-AGE-fed animals, HA high-AGE-fed animals

^aDerived from Mann–Whitney Wilcoxon test for independent samples (level of statistical significance 0.007, due to the Bonferroni correction)

* Denotes statistically significant differences

Immunohistochemical analyses of the thoracic aortas of the two experimental groups revealed no apparent histological changes in the aortas of LA and HA animals during the experimental period. However, AGEs immunoreactivity was markedly increased in the cytoplasm of endothelial cells of HA animals compared to LA (P = 0.05, Fig. 4a, b) followed by a trend showing elevation of RAGE staining in the same areas (Fig. 4c, d).

Fig. 4.

LOX and ET-1 protein expression in aortic endothelium of low- and high-AGE-fed animals. Immunohistochemical expression of AGEs (a, b), RAGE (c, d), ET-1 (e, f), LOX (g, h) and corresponding quantitation graphs showing significant elevation of AGEs and LOX in aortic endothelial and SMCs of high-AGE-fed animals (b, d, f, h) compared to low-AGE-fed animals (a, c, e, g). *P < 0.05; original magnifications (a–h): ×400

Moreover, HA animals seem to overexpress ET-1 protein in the cytoplasm of endothelial and VSMCs compared to LA (Fig. 4e, f), whilst higher LOX immunoreactivity was observed in endothelium and media of HA rats (P = 0.02, Fig. 4g, h).

Taken together, these findings confirm the functional impact of high AGEs on the regulation of endothelial markers ET-1 and LOX thus affecting endothelial homeostasis.

Discussion

Vascular endothelial dysfunction is envisioned as a key precursor of clinical arterial diseases, serving as a biomarker of the inherent risk to develop cardiovascular disease (CVD) [36, 37]. Given its central role in the development of cerebrovascular, coronary and peripheral artery diseases, vascular endothelial deregulation is considered an important therapeutic target for reducing CVD morbidity and mortality risk [37, 38].

Arterial endothelial dysfunction refers to functional changes in the normal endothelial phenotype of arteries that may contribute to the clinical manifestations and development of atherosclerosis and other vascular disorders [37, 39, 40]. These alterations trigger a shift to a vasoconstrictor, proliferative, proinflammatory and pro-coagulation state [41, 42].

The vascular endothelium is a single layer of cells lining blood vessels that regulates the strength and function of arteries [37, 39]. Vascular endothelial cells synthesize and deliver a wide range of biologically active molecules that act in an autocrine or paracrine manner to modulate arterial structure as well as vasodilatory, thrombolytic and vasoprotective functions. Endothelial barrier integrity and ECM maturation is achieved by LOX, a key enzyme that is strongly expressed in the endothelium of coronary arteries of humans and animal models [43, 44]. ET-1 is the most potent vasoconstrictor with mitogenic properties released from endothelial cells upon stimulation by cytokines, angiotensin, thrombin and reactive oxygen species (ROS) [45–49].

Excessive formation and accumulation of AGEs in arteries is believed to contribute to vascular dysfunction, perhaps via fibrosis and remodeling [50, 51]. Administration of AGE albumin (AGE-BSA) to healthy animals was followed by vascular dysfunction as observed in experimental diabetes models [52]. This cellular response is thought to be mediated through the binding of AGEs to their receptor RAGE [53]. In humans, administration of alagebrium, a breaker of AGE cross-links, to adults with isolated systolic hypertension has been shown to improve brachial artery flow mediated dilation (FMD) [54].

The present study aimed at elucidating the potential effect of AGEs in the regulation of LOX and ET-1 expression in HAECs and in an in vivo experimental model of high-AGE-fed animals. Our data demonstrate that HAECs exposed to AGE-BSA for various time points displayed upregulation of LOX and ET-1 mRNA levels in a dose- and time-dependent manner. Upregulation of LOX activity and expression has been previously detected in rat retinal endothelial cells being associated with compromised ECM-barrier function [25]. In accordance, a previous study from our group has shown that AGEs deposition in ovarian tissue regulates LOX gene expression and activity via an NF-κB-dependent pathway [24]. Since AGEs signaling is mediated via interaction with the receptor RAGE, we investigated RAGE expression in HAECs using flow cytometry. Further treatment with AGE-BSA did not altered RAGE expression in aortic cells, excluding any autoregulatory feedback mechanisms of RAGE upregulation upon AGE exposure.

In this vein, we carried on to investigate the intracellular signaling pathways that are implicated in LOX and ET-1 overexpression. We treated arterial endothelial cells with ERK1/2 and JNK inhibitors prior to glycation treatment at the highest dose (200 μg/mL) and incubation time (72 h). The observed upregulation of LOX and ET-1 mRNA levels was completely abolished upon inhibitors presence, indicating the involvement of the aforementioned kinases and their associated signaling cascades in AGE-induced regulation of LOX and ET-1 in human aortic cells. This finding was further validated by Western immunoblotting analysis of lysates from HAECs exposed to different AGE-BSA concentrations for various time points against the activated (phosphorylated, p) forms of ERK1/2 and JNK, respectively. Early activation of ERK1/2 in a dose- and time-dependent manner was revealed and remained elevated up to 72 h. P-JNK levels were increased at 6 h post exposure showing a progressive elevation up to 72 h. Moreover, employment of an anti-RAGE blocking (neutralizing) antibody prior to treatment of cells with AGE-BSA (200 μg/mL, 72 h) corroborated that RAGE was mediating the observed alterations in ERK1/2 and JNK activation. It is well established that activation of ERK1/2 and JNK pathways leads to NF-κB and AP-1 activation and previous reports have demonstrated that AGEs treatment of different cell types enhances NF-κB and AP-1 activation [55].

Given the above data, the next step was to investigate the possible regulation of the transcriptional activity of LOX and ET-1 promoter by AGEs. Using homology analysis and published literature, we identified at the functional part of LOX and ET-1 promoters highly conserved sequence motifs for NF-κB and AP-1 binding [24, 28, 29]. EMSAs revealed that these sites bear functional role. Specific DNA–protein complexes on these sites were detected, whose composition was verified by employing specific antibodies to NF-κB p65 and AP-1/p-c-Jun. Recognition and binding of both transcription factors to LOX and ET-1 promoter regions indicates that it is highly likely for AGE molecules to regulate LOX and ET-1 gene expression using at least one of the two signaling pathways. In fact, for many types of stress a concurrent activation of NF-κB and AP-1 has been documented, albeit the signal transduction pathways differ for the two transcription factors [56, 57]. It is, therefore, reasonable to postulate that accumulation of AGEs in the aortic cells may lead to potentiation of both NF-κB and AP-1 and ultimately stimulation of LOX and ET-1 expression/activity.

To further investigate the functional significance of our findings, we proceeded with the assessment of LOX and ET-1 protein expression levels in thoracic aortas of rats exposed to low (LA) or high in AGEs (HA) diet for 3 months. At the end of the three-month period, the HA animals exhibited a deregulated metabolic and lipid profile compared to LA that could potentially induce early vascular changes. More specifically, HA animals displayed elevated serum AGE levels, hyperinsulinemia, hypercholesterolemia, hypertriglycemia and early inflammation, known risk factors for impaired endothelial function (Table 1). Circulating AGEs have been shown to induce profound effects on cell function, by enhancing ROS liberation and reactive nitrogen species [58]. ROS are linked to the overexpression of pro-inflammatory mediators that play a critical role in endothelial activation and atherosclerosis [59–61]. Increased levels of ROS reduce bioactive NO through chemical inactivation, forming toxic derivatives such as peroxynitrite. These compounds can further couple endothelial NO synthase to form dysfunctional superoxide-generating enzyme that fuels oxidative stress, diminishing endothelium-dependent vascular relaxation.

Furthermore, CRP has been proposed to play a pivotal role in vascular inflammation and dysfunction [62]. It has been shown to induce RAGE expression in HAECs and is accompanied by ROS overproduction, ERK1/2 phosphorylation/activation and NF-κB potentiation [35].

Histological analysis of thoracic aortas of both animal groups revealed stronger LOX immunoreactivity in endothelial cells and media of HA-fed animals compared to LA. This is in agreement with multiple studies suggesting that disturbances of LOX expression could be related to onset and progression of atherosclerosis as well as to the instability of atherosclerotic plaques. More specifically, in a rabbit animal model exposed to a diet rich in cholesterol and peanut oil (2 % cholesterol, 8 % peanut oil) an increase in vascular LOX activity has been reported in advanced stages of atherosclerosis [63]. The same animals exhibited cholesterol levels over 1000 mg/dL within a month and highly fibrotic lesions, while an increase in vascular LOX was observed exclusively in advanced lesions from the aortic arch. Moreover, LOX has been associated with the migration and proliferation of VSMCs, being essential for the insolubilization of ECM components, since LOX is the main isoform accounting for the 80 % of LOX activity in aortic SMCs [64].

Other studies also correlate the disturbances of LOX expression with ECM remodeling associated with restenosis and aneurysm development [15]. Similarly, overexpression of ET-1 was observed in the cytoplasm of endothelial and SMCs of HA-fed animals compared to LA. This finding is in harmony with the high abundance of ET-1 mRNA in atherosclerotic tissue and the ET-1 positivity of almost all endothelial cells in vivo [65]. Furthermore, inhibition of ET-1 signaling with an ETA receptor antagonist improves endothelium-dependent dilation (EDD) in arteries of mice [66] and this is partly mediated by increased endothelial formation and exocytotic release of ET-1 [67].

Conclusion

In summary, our findings pose that AGEs can potentially regulate LOX and ET-1 genes, leading to increased LOX and ET-1 expression that is associated with distorted endothelial homeostasis. Excessive AGEs in circulation and their deposition in vascular tissues have a detrimental effect in overall endothelial physiology by impairing endothelial barrier function, altering ECM biomechanical properties and cell proliferation. It is, therefore, evident that regulation of glycative stress is a promising approach for the prevention of endothelial dysfunction and the management of vascular diseases. Given the need to identify strategies that can be used to prevent and treat vascular endothelial dysfunction, more clinical research studies are demanded to establish the value of novel interventions. Among these new approaches that are effective in reversing oxidative stress and inflammation in the vascular endothelium, anti-glycation agents would seem particularly compelling.

Abbreviations

AGEs

Advanced glycation end products

AP-1

Activator protein-1

BAECs

Bovine aortic endothelial cells

COX-2

Cyclo-oxygenase-2

CRP

C-reactive protein

CVD

Cardiovascular disease

ET-1

Endothelin-1

HDL

High-density lipoprotein

ICAM-1

Intercellular adhesion molecule-1

IL-1β

Interleukin-1 beta

LDL

Low-density lipoprotein

LOX

Lysyl oxidase

MAPK

Mitogen-activated protein kinase

NF-κB

Nuclear factor kappa B

NO

Nitric oxide

PCOS

Polycystic ovary syndrome

RAGE

Receptor for AGEs

ROS

Reactive oxygen species

VEGF

Vascular endothelial growth factor

VSMCs

Vascular smooth muscle cells

Compliance with ethical standards

Conflict of interest

The authors declare that there are no conflicts of interest with any financial organization regarding the material discussed in the manuscript.