Abstract

Abdominal aortic aneurysm (AAA) is a degenerative vascular disease with a complex aetiology that remains to be fully elucidated. Clinical management of AAA is limited to surgical repair, while an effective pharmacotherapy is still awaited. Endoplasmic reticulum (ER) stress and mitochondrial dysfunction have been involved in the pathogenesis of cardiovascular diseases (CVDs), although their contribution to AAA development is uncertain. Therefore, we aimed to determine their implication in AAA and investigated the profile of oxysterols in plasma, specifically 7-ketocholesterol (7-KC), as an ER stress inducer.

In the present study, we determined aortic ER stress activation in a large cohort of AAA patients compared with healthy donors. Higher gene expression of activating transcription factor (ATF) 6 (ATF6), IRE-1, X-binding protein 1 (XBP-1), C/EBP-homologous protein (CHOP), CRELD2 and suppressor/enhancer of Lin-12-like (SEL1L) and greater protein levels of active ATF6, active XBP1 and of the pro-apoptotic protein CHOP were detected in human aneurysmatic samples. This was accompanied by an exacerbated apoptosis, higher reactive oxygen species (ROS) production and by a reduction in mitochondrial biogenesis in the vascular wall of AAA. The quantification of oxysterols, performed by liquid chromatography-(atmospheric pressure chemical ionization (APCI))-mass spectrometry, showed that levels of 7-KC were significantly higher while those of 7α-hydroxycholesterol (HC), 24-HC and 27-HC were lower in AAA patients compared with healthy donors. Interestingly, the levels of 7-KC correlate with the expression of ER stress markers.

Our results evidence an induction of ER stress in the vascular wall of AAA patients associated with an increase in circulating 7-KC levels and a reduction in mitochondrial biogenesis suggesting their implication in the pathophysiology of this disease.

Introduction

An abdominal aortic aneurysm (AAA) is defined as a focal dilation of 50 percent greater than the normal average diameter of an aorta. It is a complex disease involving the transmural infiltration of inflammatory cells, the production of reactive oxygen species (ROS), an up-regulation and activation of degradative proteases, stimulation of apoptosis and degradation of elastin [1]. Even though the histopathological features of AAA are well documented, cellular and molecular mechanisms underlying AAA pathogenesis are not fully understood. Well-defined risk factors that are associated with the development of AAA include advanced age, male gender, smoking, hypertension, Caucasian race, family history, presence of other large vessel aneurysms and atherosclerosis [2,3]. In 2013, according to the Centers for Disease Control, AAA was the 15th leading cause of mortality in the United States in adults between the age of 60 and 64 years [4]. Numerous pharmacologic interventions have been proposed to limit AAA growth but none have provided convincing results in clinical trials [5–9]. Therefore, it is imperative to better understand the mechanisms of this important disease to provide new specific drug targets for the treatment of AAA.

We and others have reported the critical role of endoplasmic reticulum (ER) stress in the pathogenesis of hypertension, atherosclerosis, heart failure and other cardiovascular diseases (CVDs) [10–12]. Environmental factors such as ischaemia, hypoxia, ROS, aging and genetic factors disrupt ER function leading to an accumulation of misfolded and unfolded proteins in the ER lumen which promotes ER stress [13]. Unfolded protein response (UPR) is the adaptive response to ER stress, which is regulated by three ER transmembrane proteins triggering three effector branches: inositol requiring enzyme 1 (IRE1), PKR-like ER kinase (PERK) and activating transcription factor (ATF) 6 (ATF6). In turn, the activation of these effectors trigger transcriptional programmes mediated by spliced X-binding protein 1 (XBP-1), ATF4 and cleaved ATF6, respectively. Under prolonged ER stress the UPR can lead to cell pathology and apoptosis orchestrated by the C/EBP-Homologous Protein (CHOP). Dysregulation of the UPR pathway has been involved in several diseases such as diabetes, inflammation, neurodegenerative disorders or CVD [14–16]. The UPR is often closely linked to mitochondrial stress, ROS responses and autophagy [17,18]. Under pathological conditions the cross-talk between mitochondria and the ER promotes mitochondrial ROS generation and mitochondrial dysfunction which is characterized by a decrease in mitochondrial biogenesis and by an increase in oxidative damage [19,20]. Therefore, pathways that ameliorate ER stress and/or regulate mitochondrial biogenesis are potential therapeutic targets for diminishing vascular dysfunction and CVD.

Oxysterols are oxygenated derivatives of cholesterol that are present at low levels in the circulation and have shown to accumulate in the plasma and tissues in some pathologies [21]. They have been demonstrated to induce numerous deleterious effects by perturbing cholesterol homoeostasis in vascular and immune cells [21,22]. Specifically, 7-Ketocholesterol (7-KC) is a well-studied oxysterol formed by the oxidation of cholesterol and cholesterol esters present in lipoprotein deposits. Free 7-KC levels are abundant in the circulation of patients with hypercholesterolaemia and 7-KC also accumulates in atherosclerotic lesions to a greater extent than any other oxysterol [23]. Moreover, 7-KC is an inducer of ER stress and autophagy in vascular cells [15,24,25].

Due to increasing evidence involving oxysterols in the pathophysiological processes mediating CVDs [26,27], we hypothesized that the oxysterols levels can be altered in plasma of AAA patients. The clinical significance of circulating 7-KC in the setting of AAA and its relationship with the induction of ER stress in the vascular wall has not yet been evaluated.

The purpose of the present study is to elucidate the implication of ER stress and its deleterious relationship with mitochondria in AAA disease. Moreover, we analysed circulating levels of several oxysterols in the plasma focussing on 7-KC. Our findings demonstrate the enhanced expression of ER stress markers and alterations in mitochondrial biogenesis, autophagy and in the oxysterols profile of AAA patients, and suggest that therapeutic approaches aiming to reduce ER stress and to increase mitochondrial biogenesis could be useful to manage the progression of AAA disease.

Materials and methods

Human samples

Human aneurysmatic and blood samples were obtained from patients undergoing open repair for AAA at the Hospital de la Santa Creu i Sant Pau (HSCSP; Barcelona, Spain), while healthy aortas were obtained from multi-organ donors. Approval to use the discarded human tissue was given by the Ethics Committee of the HSCSP. Research has been carried out in accordance with the Declaration of Helsinki. Participation in the study of patients and control subjects was based upon informed consent of patients or legal representatives.

Abdominal aorta segments were obtained from patients (n=94) and control subjects (n=17), following strict standard operating procedures and ethical guidelines. Samples of control subjects had no post-mortem evidence of AAA. Samples were rapidly collected and stored at −80°C for subsequent RNA and protein studies or processed for immunohistochemical analysis.

The blood samples from healthy donors included in the present study were provided by the Barcelona Tissue Bank and accomplished the following inclusion criteria: healthy subjects aged 55–65 years, without type 2 diabetes, non-hypertensive (or pharmacologically controlled hypertension individuals) and without any cardiovascular complication.

Cell culture

Human vascular smooth muscle cells (VSMCs) were isolated from abdominal aorta of multi-organ donors by an explant procedure as previously described [28]. Briefly, endothelium denuded medial tissue was cut into 2–4 mm cubes that were transferred to a 25-cm2 culture flask containing 5 ml of pre-warmed culture medium M199 (Gibco, Carlsbad, CA, U.S.A.) supplemented with 10% foetal calf serum (FCS; Biological Industries, Kibbutz Beit-Haemek, Israel) and antibiotics (100 U/ml penicillin and 0.1 mg/ml streptomycin). VSMCs migrate out from the explants within 2–3 weeks. Then, after removing the explants from the flask surface, cells were trypsinized, used as P1 stage cells and routinely subcultured. Human VSMCs were cultured in M199 supplemented with 20% FCS, 2% human serum, 2 mmol/l l-glutamine (Invitrogen) and antibiotics.

For experimental procedures, cells between passages 3 and 6 were seeded in multiwell plates and subconfluent cells were starved in medium supplemented with 1% FCS for 24 h prior to the addition of 15–30 µM of 7-KC (Sigma–Aldrich) or vehicle (ethanol; 0.1% for 24–48 h in the presence or absence of human Angiotensin II (Ang II) (10−7 M) for 48 h (Sigma–Aldrich). In another set of experiments, VSMCs were pre-incubated with two antioxidants, Tempol (10 µM) or N-acetylcysteine (NAC, 10 mM), 1 h prior to exposure to 7-KC for 24 h.

Assessment of apoptosis in VSMCs

Viable, apoptotic and necrotic cells were evaluated by using Annexin V FITC Apoptosis Detection Kit I with 7-AAD (BD, Pharmingen). Briefly, treated cells (1 × 106/ml) were resuspended in binding buffer (1×) and stained with Annexin V FITC and 7AAD according to the manufacturer’s instructions. Cytometric experiments were carried out with a Beckman Coulter’s Epics XL flow cytometer (Beckman Coulter, Brea, CA) equipped with a 488-nm argon laser. For each sample, at least 10000 events were acquired. Samples incubated with the vehicle were used as a negative control. Data were analysed with the Expo32 software (Beckman Coulter, Brea, CA).

Total mRNA and protein isolation from tissue and cells

The RNeasy Fibrous Mini Kit (Qiagen, Venlo, Netherlands) was used to isolate total RNA from human aortic samples following the manufacturer’s recommendation. Total RNA isolation from human VSMCs was performed using the Tripure reagent (Roche Diagnostics; Indianapolis, IN, U.S.A.) following the manufacturer’s instructions. RNA integrity was determined by electrophoresis in agarose gels and was quantified by a NanoDrop 1000 Spectrophotometer (Thermo Scientific). Protein lysates from human aortic samples were prepared in a RIPA buffer (150 mM NaCl, 1% (v/v) Triton X-100, 0.5% (w/v) sodium deoxycholate, 0.1% (w/v) SDS, 2 mM EDTA, 50 mM Tris/HCl pH 8) by using a tissue homogenizer and following a standard protocol.

Quantitative real-time PCR

DNase I-treated total RNA (1 μg) was reverse transcribed into cDNA using the High Capacity cDNA Archive Kit (Applied Biosystems, Foster City, CA, U.S.A.) with random hexamers. Quantification of mRNA levels was performed by real-time PCR using specific primers and probes provided by the Assay-on-Demand system (Applied Biosystems) for human ATF6 (Hs00232586_m1), CHOP or DDIT3 (Hs99999172_m1), XBP-1 (Hs00231936_m1), heat shock protein 5 (HSPA5) or GRP78 (Hs99999174_m1), IRE1 or ERN1 (Hs00176385_m1), ATF4 (Hs00909569_g1), Suppressor/Enhancer of Lin-12-like (SEL1L; Hs01071406_m1), cysteine-rich with EGF-like domains 2 (CRELD2; Hs00360923_g1), mitochondrially encoded cytochrome c (Cyt C) oxidase III (MT-CO3; Hs02596866_g1), CYBB (NOX2, Hs00166163_m1), CYBA (p22phox, Hs00609145_m1), NOX4 (Hs01379108_m1), nuclear factor erythroid 2-related factor 2 (NRF2 or NFE2L2; Hs00975961_g1), metalloproteinase 2 (MMP2) (Hs01548727_m1) and IL1β (Hs01555410_m1). Human peroxisome profilerator-activated receptor coactivator 1 α (PGC1α), nuclear respiratory factor 1 (NRF1), mitochondrial transcription factor A (TFAM) and Cytochrome B (Cyt B) mRNA was determined by using real-time PCR with SYBR Green PCR Master mix (Applied Biosystems, Thermo Fisher Scientific Inc, Waltham, MA, U.S.A.) and the following primers (www.Biomers.net, Donau, Germany): Cyt B forward 5′-AGACAGTCCCACCCTCACAC-3′ and reverse 5′-AAGAGAAGTAAGCCGAGGGC-3′; PGC1 forward 5′-CCTGTGGATGAAGACGGATT-3′ and reverse 5′-TAGCTGAGTGTTGGCTGGTG-3′; TFAM forward 5′-GATGCTTATAGGGCGGAG-3′ and reverse 5′-GCTGAACGAGGTCTTTTTGG-3′; NRF1 forward 5′-GATCGTCTTGTCTGGGGAAA-3′ and reverse 5′-GGTGACTGCGCTGTCTGATA-3′; and COX1 forward 5′- GGCCTGACTGGCATTGTATT-3′ and reverse 5′- TGGCGTAGGTTTGGTCTAGG-3′ as previously described [28]. In order to appropriately apply this method, we checked that the target genes were amplified with comparable efficiencies (>95%, calculated on the basis of the slope of calibration curves). As endogenous controls glyceraldehyde-3-phosphate dehydrogenase (GAPDH; Hs02758991_g1) and β-actin (Hs99999903_m1) were used. Each sample was amplified in duplicate. Similar results were obtained after normalization to either housekeeping gene. Quantitative RT-PCR was carried out in an ABI PRISM 7900HT Sequence Detection System (Applied Biosystems). Relative mRNA levels were determined using the 2−ΔΔCt method.

Immunostaining and histology

Aorta samples were fixed in 4% paraformaldehyde/0.1 M PBS (pH 7.4) for 24 h and embedded in paraffin or transferred to a cryomold containing OCT embedding medium (Tissue Tek, Sakura, Leiden, NL) and snap-frozen in liquid nitrogen. The histological characterization of aortic samples was performed by Masson’s trichrome and by orcein stain. Immunostaining assays were performed in aortic sections from human AAA and donor samples as previously described [29].

Briefly, for immunohistochemistry assays aortic sections (5 μm) were deparaffinized in xylene, rehydrated in graded ethanol, and treated with 0.3% hydrogen peroxide for 30 min to block peroxidase activity. Then, samples were blocked with 10% of normal serum and incubated with antibodies against CHOP, cleaved-caspase 3 (9661, Cell Signaling; Boston, MA, U.S.A.) and CD68 (M0876, Dako) overnight at 4°C. After washing, samples were incubated for 1 h with a biotinylated secondary antibody (Vector Laboratories, Peterborough, U.K.). After rinsing three times in PBS, standard Vectastain (ABC) avidin–biotin peroxidase complex (Vector Laboratories) was applied, and the slides were incubated for 30 min. Colour was developed using 3,3′-diaminobenzidine (DAB) and sections were counterstained with Haematoxylin before dehydration, clearing and mounting. Negative controls, in which the primary antibody was omitted, were included to test for non-specific binding. Results were quantified and expressed as percentage of positive area versus total area or as positive cell number per area in independent sections of AAA for CHOP, cleaved-caspase 3 and CD68.

For antigen colocalization studies, double-fluorescence immunostaining was performed using a sequential method. After deparaffinization, antigen retrieval and permeabilization with PBS-0.1% Triton X-100, sections were blocked with PBS containing 5% albumin for 1 h at room temperature. Incubation with mouse polyclonal antibodies against ATF6, XBP-1 or CHOP (4°C overnight), and a rabbit polyclonal antibody against smooth muscle α-actin (ab5694, Abcam) or CD3 (SK202, Dako), were then sequentially applied. After washing the primary antibodies, donkey anti-mouse IgG conjugated to Alexa Fluor 594 and chicken anti-rabbit IgG conjugated to Alexa Fluor 488 (Molecular Probes, Life Technologies) were applied for 1 h at room temperature. Negative controls, in which the primary antibody was omitted, were also included. Samples were then mounted with ProLong Gold antifade reagent with DAPI (Molecular Probes, Life Technologies Co., Eugene, OR). Images were obtained using an SP5 Leica confocal microscope.

Western blot

Tissue lysates were separated by SDS/PAGE and transferred to 0.45-μm polyvinylidene difluoride membranes (Immobilon, Millipore. Merck KGaA, Darmstadt, Germany). Blots were incubated with antibodies directed against ATF6 (NBP1-40256), XBP-1 (NBP1-75514), CHOP (NB600-1335), dynamin-related protein 1 (DRP-1; NB110-55288) and p62 (NBP1-48320) purchased from Novus Biologicals (Bio-Techne LD-R&D Systems Europe Ltd, Abingdon, U.K.); rabbit polyclonal antibodies against Beclin 1 (PA1-16857) and LC3A/LC3B (PA1-16931) were obtained from Pierce (Thermo Fisher Scientific). Equal loading of protein in each lane was verified by β-actin (A5441, Sigma–Aldrich).

Recombinant terminal deoxynucleotidyl transferase mediated nick-end labelling assay

Recombinant Terminal Deoxynucleotidyl transferase (rTdT) mediated nick-end labelling (TUNEL) was performed using the Dead End Fluorometric Tunel System (Promega) according to the manufacturer’s guidelines. Four micrometre paraffin sections from AAA and donor aortas were deparaffinized, fixed in methanol-free paraformaldehyde before and after proteinase K treatment at 20 μg/ml for 8–10 min at room temperature. The sections were incubated with the nucleotide mixture (which included fluorescein-tagged dUTP) and rTdT enzyme for an hour at 37°C. The slides were mounted using SlowFade Gold antifade reagent with DAPI (Molecular Probes, Invitrogen) and immunofluorescent signals were viewed using a fluorescence microscope (Nikon Eclipse 55i).

Detection of superoxide anion production in situ

The oxidative fluorescent dye Dihydroethidium (DHE; Molecular Probes™ Invitrogen, Carlsbad, CA, U.S.A.) and the fluorogenic dye MitoSOX™ Red were used to evaluate superoxide anion (O2•−) production in frozen aorta sections preserved in OCT. The fluorescent dye MitoTracker™ Green FM (Molecular Probes, Thermo Fisher Scientific) was used to stain mitochondria.

Briefly, frozen tissue embedded in OCT were cut into 10-μm thick sections, placed on a glass slide and equilibrated under identical conditions for 30 min at 37°C in Krebs-HEPES buffer (in mM: 130 NaCl, 5.6 KCl, 2 CaCl2, 0.24 MgCl2, 8.3 HEPES, 11 glucose, pH = 7.4). Fresh buffer containing DHE (5 × 10−6 M) was applied topically on to each tissue section, coverslipped and incubated for 30 min in a light-protected humidified chamber at 37°C. For MitoSox red and Mitotracker green stainings, a solution of 1 × 10−6 M of each dye in phosphate buffer was applied on to the tissue sections and incubated for 15 min in a light-protected humidified chamber at 37°C. Slides were subsequently washed with phosphate buffer to remove the excess of dyes and mounted using SlowFade Gold antifade reagent (Molecular Probes, Invitrogen).

Aorta images were viewed by fluorescent laser scanning microscope (20× objective in a Leica DMI 3000 microscope; Leica AG, Germany) and quantified using the Leica LAS, 4.3 analysis system (Leica AG, Germany). Histological sections of aortas from ten different patients and different fields in each section per sample were quantified, averaged and expressed as relative fluorescent units (RFU).

Determination of oxysterols in plasma

Plasma was separated from EDTA blood collection tubes within 2 h of the blood draw. Plasma was frozen in aliquots at −80°C promptly after separation. Absolute quantification of oxysterols (24-hydroxycholesterol (HC); 25-HC; 27-HC; 7α-HC; 7β-HC and 7-KC was performed by liquid chromatography coupled to tandem mass spectrometry (LC-MS/MS) with atmospheric pressure chemical ionization (APCI) interface (UHPLC-(+)APCI-MS/MS) in the Centre for Omic Sciences (Reus, Tarragona, Spain) as previously described [30,31].

Briefly, two extraction methods were developed and validated in plasma samples: one to quantify free oxysterols (7-KC and 27-HC) based on Helmschrodt et al. [30] containing the internal standards deuterated and the other to quantify total oxysterols introducing a saponification step based on Mukhopadhyay et al. [31]. Simultaneously, a development and validation of LC-(APCI)-MS/MS methodologies for each of the compounds were carried out. An UHPLC 1290 Infinity II Series coupled to a QqQ/MS 6490 Series, both Agilent Technologies (Agilent Technologies, Santa Clara, CA, U.S.A.) was used to analyse the extracts. 25-HC was found <MQL (ranged from 1 to 5 ng/ml) in all the samples.

Analytical standards 7-β-HC, 25-HC and 7-KC were obtained from Sigma–Aldrich. 24-S-HC and 27-HC from Enzo life Sciences and 7-KC-d7 from Toronto Research Chemicals.

Determination of the lipid profile in plasma

The lipid profile was determined enzymatically using commercial kits adapted to a COBAS 6000 autoanalyzer (Roche Diagnostics, Bassel, Switzerland). The lipid profile included total cholesterol (TC), triglycerides (TGs), high-density lipoprotein-cholesterol (HDL-C), LDL cholesterol and VLDL cholesterol. LDL and VLDL cholesterol were calculated with the Friedewald’s equation, except when TG were higher than 3 mmol/l, where VLDL was separated by ultracentrifugation according to the National Cholesterol Education Program (NCEP) recommendations [32].

Statistical analysis

GraphPad Prism 4.0 software (GraphPad, U.S.A.) was used for statistical analysis. Data were expressed as mean ± SEM and values of P≤0.05 were considered significant. When data fitted a normal distribution, differences between two groups were assessed using the Student’s t test (two-tailed) and one-way ANOVA, and the Bonferroni test for more than two groups. When normality failed, we used the Mann–Whitney rank sum test to compare two groups and Kruskal–Wallis one-way analysis of variance on ranks for multiple comparisons (Dunn’s method). Pearson product-moment correlation coefficient was used to determine the association between variables. Multivariate logistic regression analysis using the statistical software package R (www.r-project.org) was performed to assess potential confounding factors. The results were expressed as odds ratio (OR) and 95% confidence intervals (CI).

Results

ER stress is present in AAA

The histological examination of abdominal aorta samples from AAA patients and donors showed the disturbance in overall abdominal aorta wall morphology and the extracellular matrix (ECM) disorganization frequently observed in AAA (Supplementary Figure S1A,B). Table 1 shows demographic and clinical data of patients and donors included in the present study.

Table 1
Demographics and clinical data of individuals included in the study
 mRNA Plasma 
 AAA Normal aorta AAA Blood donors 
n 96 17 94 47 
Age (years) 70.9 ± 6.5 60.9 ± 13.4 71.0 ± 6.7 59.3 ± 4.3 
Women (%, n) 5.2 (5) 5.9 (1) 5.3 (5) 30 (14) 
Aortic diameter (mm) 64.1 ± 1.4 64.1 ± 1.4 
Dyslipidemia (%, n) 63.5 (61) 17.6 (3) 64.9 (61) 
HTN (%, n) 85.4 (82) 41.2 (7) 84.6 (80) 7.8 
Diabetes (%, n) 22.9 (22) 29.4 (5) 23.4 (22) 
Smokers/ex-smokers (%, n) 83 (78) 41.2 (7) 82 (76) 9.8 
Systolic blood pressure (mmHg, n=67) 139.8 ± 19.3 139.8 ± 19.3 
Heart rate (n=65) 73 ± 11.6 73 ± 11.6 
Glucose (mM, n=79) 5.9 ± 1.3 5.5 ± 0.9 
TC (mM) 3.9 ± 1.2 5.6 ± 0.8 
HDL-C (mM) 1 ± 0.3 1.5 ± 0.3 
LDL-C (mM) 2.3 ± 0.9 3.5 ± 0.7 
VLDL-C (mM) 0.6 ± 0.3 0.6 ± 0.3 
TGs (mM) 1.3 ± 0.7 1.4 ± 0.7 
PAD 47.9 0.0 48.4 
BVD 9.4 11.8 15.4 
IHD 24.0 0.0 27.5 
COPD 17.7 0.0 18.7 
Antiplatelet users 59.4 11.8 62.6 
Statin users 69.8 5.9 68.1 1.8 
NSAID users 4.2 0.0 2.2 
Corticoid users 7.3 0.0 4.4 
Immuno-supressors users 4.2 0.0 3.3 
 mRNA Plasma 
 AAA Normal aorta AAA Blood donors 
n 96 17 94 47 
Age (years) 70.9 ± 6.5 60.9 ± 13.4 71.0 ± 6.7 59.3 ± 4.3 
Women (%, n) 5.2 (5) 5.9 (1) 5.3 (5) 30 (14) 
Aortic diameter (mm) 64.1 ± 1.4 64.1 ± 1.4 
Dyslipidemia (%, n) 63.5 (61) 17.6 (3) 64.9 (61) 
HTN (%, n) 85.4 (82) 41.2 (7) 84.6 (80) 7.8 
Diabetes (%, n) 22.9 (22) 29.4 (5) 23.4 (22) 
Smokers/ex-smokers (%, n) 83 (78) 41.2 (7) 82 (76) 9.8 
Systolic blood pressure (mmHg, n=67) 139.8 ± 19.3 139.8 ± 19.3 
Heart rate (n=65) 73 ± 11.6 73 ± 11.6 
Glucose (mM, n=79) 5.9 ± 1.3 5.5 ± 0.9 
TC (mM) 3.9 ± 1.2 5.6 ± 0.8 
HDL-C (mM) 1 ± 0.3 1.5 ± 0.3 
LDL-C (mM) 2.3 ± 0.9 3.5 ± 0.7 
VLDL-C (mM) 0.6 ± 0.3 0.6 ± 0.3 
TGs (mM) 1.3 ± 0.7 1.4 ± 0.7 
PAD 47.9 0.0 48.4 
BVD 9.4 11.8 15.4 
IHD 24.0 0.0 27.5 
COPD 17.7 0.0 18.7 
Antiplatelet users 59.4 11.8 62.6 
Statin users 69.8 5.9 68.1 1.8 
NSAID users 4.2 0.0 2.2 
Corticoid users 7.3 0.0 4.4 
Immuno-supressors users 4.2 0.0 3.3 

Nominal variables are presented as %. Continuous variables are presented as mean ± SD. Due to the nature of normal aorta samples and plasma from donors some of the clinical characteristics are not always recorded or are unknown and infra-evaluation of them is probable. Abbreviations: BVD, brain-vascular disease; COPD, chronic occlusive pulmonary disease; HTN, chronic hypertension; IHD, ischaemic heart disease; LDL-C, low-density lipoprotein-cholesterol; NSAID, non-steroidal anti-inflammatory drugs; PAD, peripheral artery disease.

The expression of ER stress markers that represent the three pathways of the UPR was analysed in abdominal aorta samples from AAA patients (n=96) and healthy donors (HD, n=17). IRE1, XBP-1, ATF6 and CHOP mRNA levels were significantly higher in AAA, however, no differences were observed for ATF4 and HSPA5 in comparison with aortas from donors (Figure 1A–F). After adjusting for age, sex, smoking and comorbidities (hypertension and type 2 diabetes), the differences between AAA patients and donors remained statistically significant except for CHOP expression (Supplementary Table S1).

ER stress markers expression is altered in human AAA

Figure 1
ER stress markers expression is altered in human AAA

(AH) Human abdominal aortic mRNA levels of IRE-1, XBP-1, ATF4, ATF6, HSPA5, CHOP, CRELD2 and SEL1L measured by quantitative real-time PCR analysis and normalized to β-actin in healthy donors (n=17) and patients (AAA) (n=96). Results are expressed as mean ± SEM. **P<0.01 vs. AAA; ***P<0.001 vs. AAA; NS, non-significant.

Figure 1
ER stress markers expression is altered in human AAA

(AH) Human abdominal aortic mRNA levels of IRE-1, XBP-1, ATF4, ATF6, HSPA5, CHOP, CRELD2 and SEL1L measured by quantitative real-time PCR analysis and normalized to β-actin in healthy donors (n=17) and patients (AAA) (n=96). Results are expressed as mean ± SEM. **P<0.01 vs. AAA; ***P<0.001 vs. AAA; NS, non-significant.

Additionally, we studied the expression of two recently discovered ER stress-inducible genes: CRELD2 and SEL1L (Figure 1G,H). Our results showed higher mRNA levels of CRELD2 (Figure 1G) and SEL1L (Figure 1H) in AAA, indicating an activation of the ER-associated degradation (ERAD) process.

The higher expression of XBP-1, ATF6 and CHOP in aneurysmatic samples was confirmed by Western blot (Figure 2A–C) and was further assessed by immunofluorescence. A strong immunostaining for ATF6, XBP-1 and CHOP was observed in the media layer of aneurysmatic samples, specifically in VSMCs (Figure 2D and Figure S3). Double-staining of ER stress markers with CD3 revealed that only ATF6 and XBP-1 co-localized in lymphocytes in areas of inflammatory infiltrates (Figure 2D and Figure S3).

ER stress markers expression in human AAA and co-localization with VSMC and T lymphocytes

Figure 2
ER stress markers expression in human AAA and co-localization with VSMC and T lymphocytes

(A–C) Representative Western blot analysis of cleaved ATF6, CHOP and XBP-1 (unspliced and spliced isoforms) and their corresponding quantitative histogram below (AAA: n=15 and Donors: n=10). Results are expressed as mean ± SEM. ***P<0.001 vs. AAA. (D) Representative images of immunofluorescence assays performed in abdominal aorta sections targeting ATF6, XBP-1 or CHOP (red) and α-actin or CD3-positive cells (green) as indicated in AAA patients (upper panels) and donors (lower panels) analysed by confocal microscopy (n=8–10; scale bars: 25 µm).

Figure 2
ER stress markers expression in human AAA and co-localization with VSMC and T lymphocytes

(A–C) Representative Western blot analysis of cleaved ATF6, CHOP and XBP-1 (unspliced and spliced isoforms) and their corresponding quantitative histogram below (AAA: n=15 and Donors: n=10). Results are expressed as mean ± SEM. ***P<0.001 vs. AAA. (D) Representative images of immunofluorescence assays performed in abdominal aorta sections targeting ATF6, XBP-1 or CHOP (red) and α-actin or CD3-positive cells (green) as indicated in AAA patients (upper panels) and donors (lower panels) analysed by confocal microscopy (n=8–10; scale bars: 25 µm).

As a pro-apoptotic effector of the UPR, CHOP expression in VSMC from AAA was associated with a higher expression of cleaved caspase 3. Additionally, an enhanced presence of apoptotic cells was detected in aneurysmatic wall assessed by TUNEL assay (Supplementary Figure S2A–C).

Oxidative stress is higher in the aortic wall of human AAA than in healthy abdominal aorta

In aneurysmatic abdominal aortas the expression of members of the nicotinamide adenine dinucleotide phosphate-oxidase (NADPH oxidase) family was altered. Specifically, NOX2 and p22phox mRNA levels were higher, while NOX4 expression was lower compared with donor samples. The expression of NRF2 was not altered in AAA patients (Figure 3A–D). Superoxide anion production measured by DHE staining was greater in AAA compared with donors (Figure 3E). Because mitochondria are a target of superoxide anion produced by NADPH oxidase, we stained abdominal aortic samples with Mitosox red probe. As observed in Figure 3F an increase in mitochondrial ROS was also detected in AAA patients. The staining of mitochondrial mass by using the fluorescent probe Mitotracker green revealed that mitochondria abundance (green fluorescence detected between the autofluorescent elastic lamellae; please see negative controls) was lower in aneurysmastic samples that in the arterial wall from donors (Figure 3G). Further, inmunostaining analysis revealed a partial co-localization of superoxide anion (DHE staining) with XBP-1, ATF6 and CHOP in frozen aortic samples of AAA patients (Supplementary Figure S4).

Oxidative stress is exacerbated in AAA

Figure 3
Oxidative stress is exacerbated in AAA

(AD) NOX2, p22phox, NOX4 and NRF2 mRNA levels quantified by real-time PCR analysis and normalized to β-actin in abdominal aortas from donors (n=17) and AAA patients (n=96). (E,F) Representative abdominal aortic sections from frozen tissue preserved in OCT stained with fluorescent probes DHE (E) or mitosox red (F). The histograms show the quantification of the fluorescence intensity (n=10; bar: 50 μm). (G) Representative abdominal aortic sections stained with the fluorescent probe Mitotracker green labelling mitochondrial mass (n=10; bar: 75 μm). Results are expressed as mean ± SEM. ***P<0.001 vs. AAA; ****P<0.0001 vs. AAA; NS, non-significant.

Figure 3
Oxidative stress is exacerbated in AAA

(AD) NOX2, p22phox, NOX4 and NRF2 mRNA levels quantified by real-time PCR analysis and normalized to β-actin in abdominal aortas from donors (n=17) and AAA patients (n=96). (E,F) Representative abdominal aortic sections from frozen tissue preserved in OCT stained with fluorescent probes DHE (E) or mitosox red (F). The histograms show the quantification of the fluorescence intensity (n=10; bar: 50 μm). (G) Representative abdominal aortic sections stained with the fluorescent probe Mitotracker green labelling mitochondrial mass (n=10; bar: 75 μm). Results are expressed as mean ± SEM. ***P<0.001 vs. AAA; ****P<0.0001 vs. AAA; NS, non-significant.

Mitochondrial biogenesis and mitophagy are disturbed in AAA

To analyse mitochondrial biogenesis we determined the ratio of the expression of Cyt B and Cyt C oxidase vs. that of β-actin as indicator of mitochondrial biogenesis. These ratios were significantly reduced in AAA patients compared with donors supporting a decrease in mitochondrial biogenesis (Figure 4A,B). As expected, the mRNA levels of PGC1α, a major regulator of the mitochondrial biogenesis, and its downstream targets: NRF1 and mitochondrial TFAM, a direct regulator of mitochondrial DNA replication and transcription, were dramatically reduced in human AAA samples compared with donors (Figure 4C–E).

Mitochondrial biogenesis and mitophagy/autophagy alterations in AAA

Figure 4
Mitochondrial biogenesis and mitophagy/autophagy alterations in AAA

(A,B) Graphs showing the ratios between Cyt B or Cyt C oxidase (MT-CO3 gene) and β-actin. (CE) PGC1α, TFAM and NRF1 mRNA levels quantified by real-time PCR analysis and normalized to GAPDH in abdominal aortas from donors (n=17) and AAA patients (n=96). (F) Representative Western blot analysis of DRP-1, p62, Beclin-1 and LC3II and their corresponding quantitative histograms (AAA: n=15 and Donors: n=10). Results are expressed as mean ± SEM. *P<0.05 vs. AAA; **P<0.01 vs. AAA; ***P<0.001 vs. AAA.

Figure 4
Mitochondrial biogenesis and mitophagy/autophagy alterations in AAA

(A,B) Graphs showing the ratios between Cyt B or Cyt C oxidase (MT-CO3 gene) and β-actin. (CE) PGC1α, TFAM and NRF1 mRNA levels quantified by real-time PCR analysis and normalized to GAPDH in abdominal aortas from donors (n=17) and AAA patients (n=96). (F) Representative Western blot analysis of DRP-1, p62, Beclin-1 and LC3II and their corresponding quantitative histograms (AAA: n=15 and Donors: n=10). Results are expressed as mean ± SEM. *P<0.05 vs. AAA; **P<0.01 vs. AAA; ***P<0.001 vs. AAA.

Because mitochondrial autophagy (or mitophagy) plays an important role in mitochondrial turnover, we analysed the expression of mitophagy and autophagy markers in tissue lysates of AAA patients and donors. The expression of DRP1 was lower while protein expression of LC3II, Beclin-1 and p62, three autophagy regulatory proteins, was higher in AAA compared with donors (Figure 4F).

The oxysterols profile is altered in the plasma of AAA patients

We measured the levels of free 7-KC levels and other oxysterols (24S-HC, 25-HC, 27-HC, 7-β-HC and 7-α-HC) in the plasma of AAA patients (n=94) and healthy donors (n=47) to determine their profile. We observed that 7-KC levels were significantly higher in aneurysmatic patients (Figure 5A). However, 7-α-HC, 24 (S)-HC and 27-HC (analysed as free oxysterol and as a hydrolysed oxysterol extracted from plasma) were significantly lower in AAA patients compared with donors (Figure 5C–E). 7-β-HC plasma levels were similar between donors and AAA (Figure 5B), while 25-HC levels were undetectable. After adjusting for age and sex, the differences for 7-KC, 24-S-HC and 7α-HC plasma levels between AAA patients and donors remained statistically significant (Supplementary Table S2).

Oxysterols levels are altered in AAA patients

Figure 5
Oxysterols levels are altered in AAA patients

(AE) Histograms representing the values of 7-KC, 7α-HC, 7β-HC, 24-HC and 27-HC in nanogram per millilitre in plasma of healthy donors and AAA patients. Results are expressed as mean ± SEM. The P-values from a Mann–Whitney test are provided on the donors versus AAA comparison: ***P<0.001 vs. AAA. (FH) Graphs showing the correlation analysis between IRE1, ATF6 and CHOP mRNA levels and 7-KC levels in AAA patients (n=75). (I,J) Graphs showing the correlation analysis between NOX2 or p22phox mRNA levels and 7-KC levels in AAA patients (n=78). The r and P-values are obtained by performing the Pearson’s correlation coefficient test.

Figure 5
Oxysterols levels are altered in AAA patients

(AE) Histograms representing the values of 7-KC, 7α-HC, 7β-HC, 24-HC and 27-HC in nanogram per millilitre in plasma of healthy donors and AAA patients. Results are expressed as mean ± SEM. The P-values from a Mann–Whitney test are provided on the donors versus AAA comparison: ***P<0.001 vs. AAA. (FH) Graphs showing the correlation analysis between IRE1, ATF6 and CHOP mRNA levels and 7-KC levels in AAA patients (n=75). (I,J) Graphs showing the correlation analysis between NOX2 or p22phox mRNA levels and 7-KC levels in AAA patients (n=78). The r and P-values are obtained by performing the Pearson’s correlation coefficient test.

We then studied possible correlations between levels of the different oxysterols with TC, HDL-C, low-density lipoprotein-cholesterol (LDL-C), VLDL-cholesterol (VLDL-C) and TGs levels in the plasma of patients and donors and the results are summarized in Table 2. A positive correlation was observed between 24S-HC or 27-HC levels and TC and LDL-C levels. Furthermore, only 24(S)-HC levels correlated with TC and LDL-C in either donors or AAA patients. LDL-C levels positively correlated with 27-HC levels in the AAA group but not in donors. In AAA patients HDL-C levels were significantly associated with 24S-HC and 7α-HC levels but they did not correlate in donors. A positive correlation was also found between 24(S)-HC or 27-HC levels and TG and VLDL-C levels in AAA patients but not in donors. None of the lipid profile variables were significantly associated with 7-KC levels in either AAA patients or donors. No associations were found between values of abdominal aortic diameter and oxysterols levels in AAA patients (Table 2). Interestingly, in AAA patients, 7-KC plasma levels positively correlated with the aortic expression of IRE1, ATF6 and CHOP (Figure 5F–H). Further associations of 7-KC levels were established with the local expression of NAPDH oxidase subunits in AAA (Figure 5I,J).

Table 2
Associations between oxysterols and lipid profile variables in healthy donors and AAA patients
Variable AAA vs HD n 7-KC 24-S-HC 27-HC 7α-HC 
TC (mmol/l) AAA 94 −0.0323 (P<0.7573) 0.5762 (P<0.0001) 0.4627 (P<0.0001) 0.1711 (P<0.1010) 
 HD 47 0.07300 (P<0.6258) 0.5470 (P<0.0001) 0.03066 (P<0.8379) −0.0715 (P<0.6326) 
HDL-C (mmol/l) AAA 94 −0.0023 (P<0.9825) 0.2976 (P<0.0036) 0.1385 (P<0.1832) 0.2294 (P<0.027) 
 HD 47 0.07150 (P<0.6330) 0.1355 (P<0.3637) 0.2013 (P<0.1749) −0.0193 (P<0.8974) 
LDL-C (mmol/l) AAA 94 −0.0145 (P<0.8896) 0.5730 (P<0.0001) 0.4839 (P<0.0001) 0.1003 (P<0.3388) 
 HD 47 0.07491 (P<0.6168) 0.5590 (P<0.0001) −0.04272 (P<0.7756) −0.0663 (P<0.6577) 
TG (mmol/l) AAA 94 −0.0842 (P<0.4197) 0.2828 (P<0.0057) 0.2699 (P<0.0085) 0.1322 (P<0.2066) 
 HD 47 −0.03966 (P<0.7913) 0.1003 (P<0.5024) −0.008408 (P<0.9553) −0.0260 (P<8621) 
VLDL-C (mmol/l) AAA 94 −0.0819 (P<0.4326) 0.2856 (P<0.0053) 0.2698 (P<0.0085) 0.1328 (P<0.2046) 
 HD 47 −0.03609 (P<0.8097) 0.1011 (P<0.4988) −0.01043 (P<0.9445) −0.0266 (P<8591) 
AA diameter AAA 94 −0.191 (P<0.15) −0.119 (P<0.246) −0.196 (P<0.107) −0.129 (P<0.208) 
Variable AAA vs HD n 7-KC 24-S-HC 27-HC 7α-HC 
TC (mmol/l) AAA 94 −0.0323 (P<0.7573) 0.5762 (P<0.0001) 0.4627 (P<0.0001) 0.1711 (P<0.1010) 
 HD 47 0.07300 (P<0.6258) 0.5470 (P<0.0001) 0.03066 (P<0.8379) −0.0715 (P<0.6326) 
HDL-C (mmol/l) AAA 94 −0.0023 (P<0.9825) 0.2976 (P<0.0036) 0.1385 (P<0.1832) 0.2294 (P<0.027) 
 HD 47 0.07150 (P<0.6330) 0.1355 (P<0.3637) 0.2013 (P<0.1749) −0.0193 (P<0.8974) 
LDL-C (mmol/l) AAA 94 −0.0145 (P<0.8896) 0.5730 (P<0.0001) 0.4839 (P<0.0001) 0.1003 (P<0.3388) 
 HD 47 0.07491 (P<0.6168) 0.5590 (P<0.0001) −0.04272 (P<0.7756) −0.0663 (P<0.6577) 
TG (mmol/l) AAA 94 −0.0842 (P<0.4197) 0.2828 (P<0.0057) 0.2699 (P<0.0085) 0.1322 (P<0.2066) 
 HD 47 −0.03966 (P<0.7913) 0.1003 (P<0.5024) −0.008408 (P<0.9553) −0.0260 (P<8621) 
VLDL-C (mmol/l) AAA 94 −0.0819 (P<0.4326) 0.2856 (P<0.0053) 0.2698 (P<0.0085) 0.1328 (P<0.2046) 
 HD 47 −0.03609 (P<0.8097) 0.1011 (P<0.4988) −0.01043 (P<0.9445) −0.0266 (P<8591) 
AA diameter AAA 94 −0.191 (P<0.15) −0.119 (P<0.246) −0.196 (P<0.107) −0.129 (P<0.208) 

The Pearson product-moment correlation coefficient is shown (r). The significant P-values are shown in bold. Abbreviation: HD, healthy donor.

ER stress activation and mitochondrial biogenesis alteration in human VSMC after exposure to 7-KC

To investigate the effects of 7-KC on vascular cells, human VSMCs were incubated with 15 μM of 7-KC, concentration established on the basis of preliminary studies revealing that primary VSMCs are very sensitive to 7-KC-induced apoptosis at a higher dose (data not shown). The percentages of early apoptotic, late apoptotic cells, necrotic cells and live cells in 7-KC-induced cells were quantified by flow cytometry (Supplementary Figure S5).

The expression of the ER stress markers ATF4, ATF6, CHOP, HSPA5, IRE1, SEL1L and CREDL2 was significantly induced by 7-KC in human VSMCs. When cells were co-incubated with Ang II, as a vasoactive peptide and an important mediator of the aneurysmatic process, we discovered that Ang II triggered an additional increment in CHOP mRNA levels which suggests a synergistic effect (Figure 6A–G).

The exposure to 7-KC triggers ER stress response in human VSMCs and the addition of Ang II has a synergistic effect

Figure 6
The exposure to 7-KC triggers ER stress response in human VSMCs and the addition of Ang II has a synergistic effect

(AG) mRNA levels of HSPA5, ATF4, ATF6, IRE1, CHOP, SEL1L and CREDL2; (HK) mRNA levels of PGC1α, TFAM and NRF1 normalized to GAPDH in human VSMCs stimulated with 7-KC (15 × 10−6 M) for 24 h in the presence or absence of Ang II (AII, 10−7 M) for 48 h. Values are shown as mean ± SEM (n=6; *P<0.01 vs. vehicle; **P<0.001 vs. vehicle; #P<0.05 vs. 7-KC).

Figure 6
The exposure to 7-KC triggers ER stress response in human VSMCs and the addition of Ang II has a synergistic effect

(AG) mRNA levels of HSPA5, ATF4, ATF6, IRE1, CHOP, SEL1L and CREDL2; (HK) mRNA levels of PGC1α, TFAM and NRF1 normalized to GAPDH in human VSMCs stimulated with 7-KC (15 × 10−6 M) for 24 h in the presence or absence of Ang II (AII, 10−7 M) for 48 h. Values are shown as mean ± SEM (n=6; *P<0.01 vs. vehicle; **P<0.001 vs. vehicle; #P<0.05 vs. 7-KC).

Additionally, we studied whether mitochondrial biogenesis biomarkers, MMP2 and the pro-inflammatory marker IL-1β were altered in VSMCs treated with 7-KC and co-incubated or not with Ang II. We observed that 7-KC reduced PGC1α and TFAM expression, while mRNA levels of NRF1 were not significantly altered (data not shown) (Figure 6H,I). Furthermore, MMP2 and IL-1β mRNA levels were up-regulated after exposure to 7-KC (Figure 6J,K). The co-incubation with Ang II did not either change the effects of 7-KC on mitobiogenesis markers or in IL-1β expression but triggered a further increment in MMP2 expression (Figure 6H–K).

Because it has been well-established that 7-KC triggers oxidative stress in vascular cells [33,34], we determined whether ROS production contributes to 7KC-mediated responses. The pre-incubation of VSMCs with the antioxidants Tempol and/or NAC attenuated the 7-KC-mediated induction of several ER stress and oxidative stress markers expression (Supplementary Figure S6).

Discussion

Currently, therapeutic strategies in AAA are restricted to invasive surgical repair for those AAA patients with a high risk of rupture. No effective pharmacological strategies are yet established to suppress the development of AAA and to prevent the need for invasive aneurysm repair [5–9,35]. Therefore, the search for new therapeutic targets to limit AAA progression is a challenge that requires further research.

In the last decade, an increasing number of studies have reported that an exacerbated induction of ER stress and mitochondrial stress could be detrimental in multiple pathologies including CVDs. Emerging evidence for protein trafficking dysregulation and ER abnormalities compatible with ER stress have been described in cardiovascular pathologies [36–38]. Nevertheless, ER stress has not been considered as a therapeutic target in AAA and little is known about its role in the vascular wall degeneration that occurs during AAA formation, expansion and rupture. Our results show evidence of the involvement of ER stress and its deleterious cross-talk with mitochondria, oxidative stress generation and apoptosis in human AAA.

In human aneurysmatic samples from our patient cohort, the UPR response was activated as demonstrated by the higher expression of IRE1-XBP1 axis, ATF6 and CHOP compared with abdominal aortas from donors, while no differences between were observed for ATF4 and HSPA5, indicating a differential activation of the three axes of the ER stress response at the end stage of AAA. To reinforce our data, we studied the expression of two recently reported players involved in ER stress known as CRELD2 and SEL1L. Interestingly, their expression was higher in aneurysmatic samples. CRELD2 is implicated in the processing and trafficking of proteins through the ER–Golgi apparatus and it is up-regulated by ATF6 [39] while SEL1L is involved in ERAD [40] and targets misfolded secretory and membrane proteins in the ER for proteasomal degradation [41]. Therefore, their induction is consistent with the enhanced expression of ATF6 detected in aneurysmatic patients.

According to the immunofluorescence studies, the induction of ER stress markers was observed mainly in VSMCs and to a lesser extent in the inflammatory infiltrate of the aneurysmatic wall. These results suggest that ER stress could be related to arterial wall degeneration occurring during AAA progression. Recently, it has been reported that inhibition of ER stress with Tauroursodeoxycholic acid (Tudca), with intermedin 1-53 peptide or by the treatment with statins reduced the incidence of AAA in a murine model, supporting the role of ER stress in AAA formation [42–44]. However, to our knowledge, this is the unique study that has exhaustively analysed the three effector branches of the ER stress response in a big cohort of AAA patients.

We found that the profile of oxysterols was clearly altered in our cohort of aneurysmatic patients. Interestingly, a similar oxysterol profile was recently reported in a study performed with patients suffering from multiple sclerosis (MS) [31]. Other studies have shown that oxysterols levels are altered not only in MS but also in other neurodegenerative diseases such as Alzheimer’s and Parkinson’s diseases, in which ER stress seems to play a critical role [31,45,46]. 7-KC levels were significantly higher in plasma of AAA patients compared with healthy donors and more interestingly, our data show a positive correlation between 7-KC plasma levels and the vascular expression of ER stress markers in AAA. In this context, recent publications emphasize on the importance of circulating oxysterols in the development of CVDs, and in particular the augmentation of 7-KC levels has been associated with CVD events [26,27]. In support to our results, other authors previously reported that cigarette smoking increases plasmatic 7-KC levels in humans [47]. Since smoking is considered one of the main risks factors for the development of AAA disease, an imbalance in the oxysterols levels could be considered as a causative factor. However, the pathophysiological repercussion of the higher 7-KC levels in aneurysmatic patients remains to be defined.

In human VSMCs, 7-KC evoked the induction of ER stress and apoptosis as reported before by Pedruzzi et al. [48]. It has been described that 7-KC amplifies inflammatory processes and regulates the expression of inflammatory cytokines and MMPs [21]. Accordingly, we found an induction of MMP-2 and IL-1β expression in human VSMCs triggered with 7-KC supporting the deleterious effect that this oxysterol could trigger in the vascular wall during AAA disease. In our cohort of patients, the levels of 7β-HC remained unchanged compared with donors. In contrast, 7α-HC, 24(S)-HC and 27-HC levels were significantly lower in AAA. Compared with other oxysterols, 24-HC levels are found to be evidently dependent upon age and they are strikingly decreased in the sixth decade of life. A decrease in 24-HC levels has been reported before in MS and Alzheimer’s patients [49]. However, no correlations of either oxysterols levels and the age of individuals were found, which is anticipated due to the advanced age of our cohort of AAA patients and donors. 24-HC and 27-HC are ligands of liver X receptors (LXRs) which controls the transcription of genes involved in cholesterol homoeostasis and innate immunity [50]. We hypothesize that a decrease in 24-HC, 27-HC and 7α-HC levels could disrupt cholesterol homoeostasis and exacerbate an immunological response in the vascular wall through a reduction in LXRs activation. Altogether, these findings suggest that a disruption in the oxysterols network could contribute to AAA disease progression.

Increased circulating 7-KC could entail the unique intersection of enhanced intracellular free cholesterol and increased oxidative stress. Among the oxysterols, 7-KC and 27-HC, both induce ROS in VSMCs and non-vascular cells [51–53]. Interestingly, we observed that the pre-incubation of VSMC with antioxidants attenuated the increased expression of oxidative and ER stress markers induced by 7-KC. In this context, we noticed a positive correlation between NOX2 or p22phox expression and 7-KC levels in AAA patients, supporting the negative impact of this oxysterol on vascular homoeostasis. Oxidative stress is closely related to ER stress induction and it is also known to be involved in the pathological mechanisms underlying human aortic abdominal aneurysm [54]. ER stress up-regulates NADPH oxidase expression and activity, the major source of vascular ROS [12]. In turn, O2 modulates inflammatory reactions and can enhance leucocyte recruitment, which also contributes to AAA progression [55]. In our cohort of AAA patients, we observed an increased expression in NOX2 and p22phox subunits of NADPH oxidase and an exacerbated production of superoxide anion and mitochondrial ROS in aneurysmatic tissue. In contrast, NOX4 expression was lower in AAA aorta than in donor samples as previously described by Guzik et al. in a small cohort of AAA patients [56].

Moreover, ER stress is associated with mitochondrial dysfunction by promoting mitochondrial ROS generation and apoptosis [19]. Mitochondria-dependent ROS/apoptosis activation plays a critical role in the progression of AAA in animal models of the disease [57]. It has been shown that induction of mitochondrial biogenesis leads to a decrease in apoptosis and ROS production. Conversely, mitochondrial biogenesis is down-regulated by ROS [58,59]. In our cohort of AAA patients we observed that the expression of transcription factors that activate mitochondrial biogenesis and regulate mitochondrial function (PGC1α, NRF1 and TFAM) as well as Cyt B and Cyt C oxidase were all decreased in aneurysmatic tissue, suggesting a reduction in mitochondrial content. Consistent with this, a previous study enrolling a limited number of patients reported a decrease in PGC1α in the medial wall of aneurysmatic aorta [60]. Moreover, Mitotracker staining evidenced a reduction in the mitochondrial mass in aneurismatic wall. Therefore, our results reinforce the concept that the maintenance of mitochondrial homoeostasis could be an interesting therapeutic strategy in AAA and support that pharmacological approaches increasing mitochondrial biogenesis could ameliorate vascular degeneration.

Activation of ER stress can trigger changes not only in mitochondrial function but also in autophagy [61] and 7-KC induces autophagy and apoptosis in advanced atherosclerotic plaques [23]. Autophagy is a major catabolic process that delivers proteins, cytoplasmic components and organelles to lysosomes for degradation and recycling. While autophagy is a critical cytoprotective mechanism, it has been suggested to also lead to cellular death depending on cell circumstances [17,62]. In this study, we found a higher expression of Beclin-1, LC3II and p62 in human aneurysmatic aortas. In turn, mitophagy eliminates the accumulation of damaged mitochondria that otherwise result in excessive ROS production and apoptosis [61,63]. Protein levels of DRP1, a marker for mitophagy, was lower in AAA than in donors. This reduction could be derived from the decrease in mitochondrial content, but could also suggest that this protective mechanism was no longer active in the aneurysmatic wall.

We found the inherent restrictions of the work with human specimens. The availability of aortic specimens is hampered by the requirement of multiorganic donors, from which it is almost impossible to obtain blood samples. Therefore, healthy aorta and blood donors are not from the same group of individuals. Further, on the basis of our data, we could not definitively demonstrate a direct role of ER stress in human AAA and we can only speculate that ER stress is related to AAA development as part of the pathophysiological process. However, it should be noted, that inhibition of ER stress limits AAA development in animal models, supporting the contribution of ER stress to this disease [42–44,64]. Finally, there are important differences between AAA patients and healthy donors regarding clinical data. Full clinical information and pharmacological treatments that might interfere in our study are lacking for blood donors. Nevertheless, after performing the corresponding statistical analysis we could exclude age, sex, smoking, hypertension and type 2 diabetes as confounding factors of our current observations.

In summary, our study highlights the importance of ER stress and mitochondrial biogenesis in AAA and demonstrates, for the first time, the associated disturbance of the oxysterols profile in this disease. Our data support that the increase in 7-KC levels and the decrease in plasma concentrations of several HCs could contribute to ER stress activation and chronic inflammation in the aorta of AAA patients. Most importantly, the present study suggests that ER stress and mitochondrial dysfunction could be potential targets for novel therapeutic strategies to limit aneurysm progression in patients who are diagnosed at an early stage of AAA in screening programmes.

Clinical perspectives

  • The surgical treatment of AAA is only recommended for aneurysms that are greater than 5.5 cm in diameter. Moreover, for those AAAs which are first diagnosed when their diameter is less than 5.5 cm, pharmacologic treatment could be beneficial in slowing or reducing AAA expansion and rupture. Numerous pharmacologic interventions have been proposed to limit AAA growth and rupture but the negative results obtained with these therapies evidence the need of new pharmacological approaches to manage this disease.

  • The results of the present study support an important role of ER stress activation and reduction in mitochondrial biogenesis in AAA and indicate a strong association between an enhanced expression of ER stress markers, oxidative stress and 7-KC in AAA.

  • Our findings suggest that ER stress and mitochondrial biogenesis could be potential therapeutic targets to limit AAA progression.

Acknowledgments

The authors are grateful to Sonia Alcolea for her technical assistance; to Dr. Nuria Canela for her technical support at the COS center (Reus, Tarragona) and to Dr. David de Gonzalo-Calvo for his assistance with the statistical analysis using the software package R.

Competing Interests

The authors declare that there are no competing interests associated with the manuscript.

Funding

This work was supported by the Spanish Ministerio de Economía y Competitividad (MINECO)-Instituto de Salud Carlos III (ISCIII) [grant numbers CP15/00126, PI17/08137 (to M.G.), PI18/0919 (to C.R.) and RTI2018-094727-B-100 (to J.M.G.)]; the CIBERCV [grant number CB16/11/00257]; the ISCIII, Miguel Servet I program (grant number CP15/00126 (to M.N.M. and M.G.)]; and by Agencia de Gestio d’Ajuts Universitaris i de Recerca (AGAUR; program of support to Research Groups. Ref. 2017-SGR-00333). The study was co-founded by Fondo Europeo de Desarrollo Regional (FEDER)-The way to build Europe.

Author Contribution

M.G. conceived, designed and supervised the study. M.-N.M. and M.G. performed experiments, analysed and interpreted data. J.F., J.R.E., M.C. and M.G. were responsible for the clinical aspects of the study; participated in patients material collection and analysis. C.R., M.K., L.C. and J.M.-G. conceived specific experiments, revised and carried out results interpretation. M.G. drafted the manuscript. All the authors revised the manuscript for important intellectual content and gave their final approval of the submitted version.

Abbreviations

     
  • AAA

    abdominal aortic aneurysm

  •  
  • APCI

    atmospheric pressure chemical ionization

  •  
  • Ang II

    angiotensin II

  •  
  • ATF

    activating transcription factor

  •  
  • CHOP

    C/EBP-homologous protein

  •  
  • CVD

    cardiovascular disease

  •  
  • Cyt B

    Cytochrome B

  •  
  • Cyt C

    Cytochrome c

  •  
  • DHE

    Dihydroethidium

  •  
  • ER

    endoplasmic reticulum

  •  
  • ERAD

    ER-associated degaradation

  •  
  • HC

    hydroxycholesterol

  •  
  • HDL-C

    high-density lipoprotein-cholesterol

  •  
  • HSCSP

    Hospital de la Santa Creu i Sant Pau

  •  
  • HSPA5

    heat shock protein 5

  •  
  • IL

    interleukin

  •  
  • IRE1

    inositol requiring enzyme 1

  •  
  • LDL-C

    low-density lipoprotein-cholesterol

  •  
  • LXR

    liver X receptor

  •  
  • MMP2

    metalloproteinase 2

  •  
  • MS

    multiple sclerosis

  •  
  • NAC

    N-acetylcysteine

  •  
  • NADPH oxidase

    nicotinamide adenine dinucleotide phosphate-oxidase

  •  
  • NRF1

    nuclear respiratory factor 1

  •  
  • NRF2

    nuclear factor erythroid 2-related factor 2

  •  
  • PGC1α

    peroxisome profilerator-activated receptor coactivator 1 α

  •  
  • ROS

    reactive oxygen species

  •  
  • rTdT

    recombinant terminal deoxynucleotidyl transferase

  •  
  • SEL1L

    suppressor/enhancer of Lin-12-like

  •  
  • TC

    total cholesterol

  •  
  • TFAM

    mitochondrial transcription factor A

  •  
  • TG

    triglyceride

  •  
  • TUNEL

    rTdT-mediated nick-end labelling

  •  
  • UPR

    unfolded protein response

  •  
  • VLDL-C

    VLDL-cholesterol

  •  
  • VSMC

    vascular smooth muscle cell

  •  
  • XBP-1

    X-binding protein 1

  •  
  • 7-KC

    7-ketocholesterol

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