# Extracellular Vesicles From LPS-Treated Macrophages Aggravate Smooth Muscle Cell Calcification by Propagating Inflammation and Oxidative Stress

Linda Yaker, Abdellah Tebani, Céline Lesueur, Chloé Dias, Vincent Jung, Soumeya Bekri, Ida Chiara Guerrera, Saïd Kamel, Jérôme Ausseil, Agnès Boullier

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## To cite this version:

Linda Yaker, Abdellah Tebani, Céline Lesueur, Chloé Dias, Vincent Jung, et al.. Extracellular Vesicles From LPS-Treated Macrophages Aggravate Smooth Muscle Cell Calcification by Propagating Inflammation and Oxidative Stress. Frontiers in Cell and Developmental Biology, 2022, 10, pp.823450. 10.3389/fcell.2022.823450. hal-03654211

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# Extracellular Vesicles From LPS-Treated Macrophages Aggravate Smooth Muscle Cell Calcification by Propagating Inflammation and Oxidative Stress

Linda Yaker¹, Abdellah Tebani²,³, Céline Lesueur², Chloé Dias⁴, Vincent Jung⁵, Soumeya Bekri²,³, Ida Chiara Guerrera⁵, Saïd Kamel¹,⁶, Jérôme Ausseil⁴,⁷† and Agnès Boullier¹,⁶*†

OPEN ACCESS

**Edited by:**

Youhua Tan,
Hong Kong Polytechnic University,
Hong Kong SAR, China

**Reviewed by:**

Yvonne Döring,
Universitätsspital Bern Departement
Herz und Gefässe Universitätsklinik für
Angiologie, Switzerland
Ulrike Resch,
Medical University of Vienna, Austria

*Correspondence:

Agnès Boullier
Boullier.agnes@chu-amiens.fr

†These authors have contributed
equally to this work and share last
authorhip

**Specialty section:**

This article was submitted to
Molecular and Cellular Pathology,
a section of the journal
Frontiers in Cell and Developmental
Biology

**Received:** 27 November 2021

**Accepted:** 09 February 2022

**Published:** 09 March 2022

**Citation:**

Yaker L, Tebani A, Lesueur C, Dias C,
Jung V, Bekri S, Guerrera IC, Kamel S,
Ausseil J and Boullier A (2022)
*Extracellular Vesicles From LPS-Treated Macrophages Aggravate Smooth Muscle Cell Calcification by Propagating Inflammation and Oxidative Stress.*
*Front. Cell Dev. Biol.* 10:823450.
doi: 10.3389/fcell.2022.823450

¹MP3CV-UR7517, CURS-University of Picardie Jules Verne, Amiens, France, ²INSERM U1245, CHU Rouen, Normandie University, UNIROUEN, Rouen, France, ³Department of Metabolic Biochemistry, Rouen University Hospital, Rouen, France, ⁴Infinity, INSERM UMR1291, CNRS UMR5051, University of Toulouse III, Toulouse, France, ⁵INSERM US24/CNRS UAR3633, Proteomic Platform Necker, University of Paris—Federative Research Structure Necker, Paris, France, ⁶Laboratory of Biochemistry, CHU Amiens-Picardie, Amiens, France, ⁷Service de Biochimie, Institut Fédératif de Biologie, CHU Toulouse, Toulouse, France

**Background:** Vascular calcification (VC) is a cardiovascular complication associated with a high mortality rate among patients with diseases such as atherosclerosis and chronic kidney disease. During VC, vascular smooth muscle cells (VSMCs) undergo an osteogenic switch and secrete a heterogeneous population of extracellular vesicles (EVs). Recent studies have shown involvement of EVs in the inflammation and oxidative stress observed in VC. We aimed to decipher the role and mechanism of action of macrophage-derived EVs in the propagation of inflammation and oxidative stress on VSMCs during VC.

**Methods:** The macrophage murine cell line RAW 264.7 treated with lipopolysaccharide (LPS-EK) was used as a cellular model for inflammatory and oxidative stress. EVs secreted by these macrophages were collected by ultracentrifugation and characterized by transmission electron microscopy, cryo-electron microscopy, nanoparticle tracking analysis, and the analysis of acetylcholinesterase activity, as well as that of CD9 and CD81 protein expression by western blotting. These EVs were added to a murine VSMC cell line (MOVAS-1) under calcifying conditions (4 mM Pi—7 or 14 days) and calcification assessed by the o-cresolphthalein calcium assay. EV protein content was analyzed in a proteomic study and EV cytokine content assessed using an MSD multiplex immunoassay.

**Results:** LPS-EK significantly decreased macrophage EV biogenesis. A 24-h treatment of VSMCs with these EVs induced both inflammatory and oxidative responses. LPS-EK-treated macrophage-derived EVs were enriched for pro-inflammatory cytokines and CAD, PAI-1, and Saa3 proteins, three molecules involved in inflammation, oxidative stress, and VC. Under calcifying conditions, these EVs significantly increase the calcification of VSMCs by increasing osteogenic markers and decreasing contractile marker expression.
**Conclusion:** Our results show that EVs derived from LPS-EK-treated-macrophages are able to induce pro-inflammatory and pro-oxidative responses in surrounding cells, such as VSMCs, thus aggravating the VC process.

**Keywords:** extracellular vesicles, vascular calcification, inflammation, oxidative stress, macrophages

# 1 INTRODUCTION

Vascular calcification (VC) is a cardiovascular complication found among patients with diseases such as diabetes, atherosclerosis, and chronic kidney disease (CKD) (Lee et al., 2020). VC is an active process (Demer and Tintut, 2008; Voelkl et al., 2019), characterized by an imbalance of calcium/phosphate homeostasis and hydroxyapatite mineral deposition, both in the intimal and medial layers of the artery (Demer and Tintut, 2008; Drüeke and Massy, 2011; Lee et al., 2020). Vascular smooth muscle cells (VSMCs), which represent the most abundant cell type in vessels, play a pivotal role in the initiation and development of VC (Jaminon et al., 2019). VSMCs undergo a phenotypic switch, with modification of osteogenic, contractile, and synthetic marker expression (Durham et al., 2018). Other cell types may be involved in the induction of the VC process, such as macrophages (Li Y. et al., 2020) and endothelial cells. Indeed, macrophages play an important role in the progression of VC by secreting inflammatory factors and inducing oxidative stress (Hénaut et al., 2019). As a consequence, macrophage-induced inflammation can reduce the production of VC inhibitors, such as fetuin-A (Moe and Chen, 2005), a protein that can bind excess mineral and increase their plasma solubility (Komaba and Fukagawa, 2009). Furthermore, an increase in reactive oxygen species (ROS) production is involved in VSMC osteochondrogenic trans-differentiation during the VC process (Tóth et al., 2020; Hu et al., 2021). Recent studies have highlighted the role of extracellular vesicles (EVs) in VC (Hodroge et al., 2017; Mansour et al., 2020; Yaker et al., 2020; Qin et al., 2021). These membrane-bound vesicles, secreted by prokaryotic and eukaryotic cells (Woith et al., 2019), can be of various origins, depending on their mode of biogenesis. For example, exosomes (50–150 nm) originate from endosomes and microvesicles (50–500 nm) generated by budding of the plasma membrane and apoptotic bodies (van Niel et al., 2018). Several studies have showed macrophage-derived EVs to promote VC (New et al., 2013; Chen et al., 2016; Kawakami et al., 2020). Analysis of their content identified a subset of molecules involved in inflammation and oxidative stress, such as pro-inflammatory cytokines (Fitzgerald et al., 2018; Aiello et al., 2020) and oxidant machinery proteins (Bodega et al., 2019). New et al. showed that macrophages can release calcifying EVs enriched for S100A9, a calcium-binding protein involved in mineralization (New et al., 2013). Furthermore, Kawakami et al. recently showed that calcifying EVs released by macrophages contribute to the formation of microcalcification (Kawakami et al., 2020). In addition, Chen et al. demonstrated that the cytokine HMGB1 can induce the secretion of macrophage-derived EVs involved in ectopic mineralization (Chen et al., 2016). Here, we aimed to investigate the role of macrophage-derived EVs in the

propagation of inflammation and oxidative stress during the VC process.

# 2 MATERIALS AND METHODS

## 2.1 Cell Culture, Molecular, and Biochemical Reagents

Fetal boyine serum (FBS) and glutamine were purchased from Eurobio® (Les Ulis, France). Lipopolysaccharide from *Escherichia coli* K12 (LPS-EK) was obtained from InvivoGen® (San Diego, California, United States) and inorganic phosphate (Pi) from Merck® (Darmstadt, Germany). Exosome-free FBS, TRIzol™ Reagent, RNase/DNase-free water, High-Capacity RNA-to-cDNA™ kits, BCA™ Protein Assay kits, and dihydroethidium (DHE) were purchased from Thermo Fisher Scientific® (Waltham, Massachusetts, United States). Takyon™ was obtained from Eurogentec® (Liège, Belgium). 2', 7'-dichlorofluorescein diacetate (DCFH-DA) was purchased from Molecular Probes® (Eugene, Oregon, United States). All other molecular and biochemical reagents were obtained from Sigma-Aldrich® (Saint-Louis, Missouri, United States).

## 2.2 Culture and Treatment of Cells

### 2.2.1 Murine Macrophage Culture

Murine macrophages (RAW 264.7 ATCC® TIB-71™, Manassas, Virginia, United States) were maintained in DMEM 6546 medium supplemented with 10% FBS, 4 mM glutamine, 100 UI/ml penicillin, and 100 µg/ml streptomycin at 37°C in a 5% CO₂ humidified atmosphere. We first assessed RAW cell viability in the presence of 0.1 and 1 µg/ml LPS-EK, concentrations that are generally used to activated RAW macrophages (Pi et al., 2014). A 24-h treatment with LPS-EK induced 25% cell cytotoxicity (data not shown). RAW cells are known to be particularly sensitive to LPS, which could explain the cell growth inhibition observed in our experiment (Raschke et al., 1978). We then tested the treatment of RAW cells for only 6 h with LPS-EK and detected no cell cytotoxicity (Supplementary Figure S1). We thus used these experimental conditions for all further experiments.

### 2.2.1 Murine Aortic Vascular Smooth Muscle Cell Culture

Murine aortic VSMCs (MOVAS-1 ATCC® CRL-2797™, Manassas, Virginia, United States) were maintained in DMEM 6546 medium supplemented with 10% FBS, 4 mM glutamine, 100 UI/ml penicillin, 100 µg/ml streptomycin, and 200 µg/ml geneticin® (G418) at 37°C in a 5% CO₂ humidified atmosphere. To induce VSMC calcification, cells were treated in DMEM 6546 containing 1% FBS with 4 mM Pi for 14 days. The
media was changed twice a week. For each experiment, VSMCs were treated with macrophage-derived EVs isolated from the same volume of cell-culture medium.

## 2.3 Extracellular Vesicle Preparation and Characterization

### 2.3.1 Isolation of Extracellular Vesicles

Macrophages were seeded in 10-cm petri plates at a density of 30,000 cells/cm² and cultured in DMEM 6546 medium supplemented with 10% FBS. After 48 h, cells were washed with PBS then treated with various concentrations of LPS-EK in DMEM 6546 medium supplemented with 10% exosome-free FBS for 6 h. Cell debris was removed from cell-culture supernatants by centrifugation at 800 × g for 5 min at 4°C.

EVs were isolated from cell-culture supernatants of untreated (EV-CT) or LPS-EK treated (EV-LPS) macrophages by sequential centrifugation, as described previously by Chen et al. (2008). Briefly, supernatants were first ultracentrifuged at 100,000 × g for 50 min at 4°C. The pellet was then resuspended in cold Dulbecco's phosphate buffered saline (D-PBS) and centrifuged at 100,000 × g for another 50 min at 4°C. The final pellet was resuspended in D-PBS or RIPA buffer and the protein concentration determined using the BCA™ Protein Assay kit. EV samples were stored at -80°C for future analysis.

### 2.3.2 Characterization of Extracellular Vesicles

#### 2.3.2.1 Transmission Electron Microscopy

EVs were isolated as described previously and resuspended in 50 µL Tris base buffer (100 mM, pH 7.4). EV samples were prepared for TEM using the conventional negative staining procedure. Briefly, 10 µL EV samples were absorbed for 2 min on formvar-carbon-coated copper grids preliminarily ionized using the PELCO easiGlow™ Glow Discharge Cleaning System (Ted Pella Inc., Redding, California, United States). Preparations were then blotted and negatively stained with 1% uranyl acetate for 1 min. Grids were examined using an 80 kV JEM-1400 electron microscope (JEOL Inc., Peabody, Massachusetts, United States) and images acquired with a digital camera (Gatan Orius, Gatan Inc., Pleasanton, California, United States).

#### 2.3.2.2 Cryo-Electron Microscopy

To analyze the morphology of EVs by cryo-EM, 3 µL of EV sample was first deposited onto a glow-discharged 200-mesh lacey carbon grid. Prior to freezing, the grid was loaded into the thermostatic chamber of a Leica EM-GP automatic plunge Freezer, set to 20°C and 95% humidity. Excess solution was blotted from the grid for 1-2 s with a Whatman n°1 filter paper and the grid immediately flash frozen in liquid ethane cooled to -185°C. Specimens were then transferred into a Gatan 626 cryo-holder. Cryo EM was carried out using a Jeol 2,100 microscope equipped with a LaB6 cathode operating at 200 kV under low-dose conditions. Images were acquired using SerialEM software (Mastronarde, 2005), with the defocus ranging from 600 to 1,000 nm, using a Gatan US4000 CCD camera. This device was placed at the end of a GIF quantum energy filter (Gatan Inc. Berwyn, Pennsylvania, United States)

operating in zero-energy-loss mode, with a slit width of 25 eV. Images were recorded at a magnification corresponding to a calibrated pixel size of 0.87 Å.

#### 2.3.2.3 Nanoparticle Tracking Analysis (NTA)

EVs were resuspended in 50 µL D-PBS. Particle-size distribution and concentration were analyzed using a NanoSight LM10-HS instrument (Malvern Instruments Ltd., Malvern, United Kingdom) according to the manufacturer's instructions. Briefly, EV samples were diluted 100-fold in D-PBS and the diluted preparation injected into the chamber. Samples were analyzed at room temperature for 60 s. Three replicates were performed for each sample. Data were acquired and analyzed using NTA 2.2 Build 127 software (Malvern Instruments Ltd., Malvern, United Kingdom).

#### 2.3.2.4 Specific Extracellular Vesicle Markers

EVs were characterized by analyzing tetraspanin (CD81, CD9) and β-actin protein expression by western blotting as recommended by the International Society for Extracellular Vesicles (ISEV) (Théry et al., 2018).

#### 2.3.2.5 Measurement of Acetylcholinesterase Activity

Acetylcholinesterase activity was measured using a colorimetric assay as previously described by Ellman et al. (1961). Briefly, 200 µL D-PBS containing 1 mM acetylcholine and 0.1 mM 5,5'-Dithiobis 2-nitrobenzoic acid (DNTB) was added to 100 µL EV sample. After a 15 min incubation at room temperature, the absorbance was read at 450 nm using an Envision microplate reader. Data are expressed as the percentage difference in absorbance compared to the control (assay diluent D-PBS).

## 2.4 Biochemical Assays

### 2.4.1 Cell Viability Assay

Cell viability was assessed using the WST-1 assay. Cells were seeded in 96-well plates at a density of 7,500 cells/well. After 48 h, cells were treated with LPS-EK or macrophage-derived EVs for 6 or 24 h, respectively. Ten percent dimethyl sulfoxide (DMSO) was used as a positive control for viability loss. The medium was then changed and the cells incubated in 100 µL DMEM 6546 medium containing 10 µL WST-1 reagent for 1 h at 37°C. Absorbance was measured at 450 nm using an Envision microplate reader (Perkin Elmer®, Waltham, Massachusetts, United States).

### 2.4.2 Calcification Assay

VSMC calcification was assessed by measuring the intracellular calcium concentration using the o-cresolphthalein assay as previously described (Ray Sarkar and Chauhan, 1967). The total cell protein concentration was assessed by the method of Peterson (Peterson, 1977) and used to normalize the intracellular calcium concentration.

### 2.4.3 Measurement of Oxidative Stress (ROS, O₂⁻, and NO Production)

The three fluorescent probes DCFH-DA, DHE, and DAF were used to measure ROS, O₂⁻, and NO production, respectively. Macrophages were seeded in white 96-well plates at a density of
7,500 cells/well. After 48 h, cells were washed twice with D-PBS and incubated at 37°C with 10 µM DCFH-DA for 30 min and then 10 µM DHE or 0.1 µM DAF for 1 h. Next, cells were washed twice with D-PBS and treated with LPS-EK or macrophage-derived EVs. Hydrogen peroxide (H₂O₂) at 50 and 500 µM was used as a positive control of ROS production. For certain experiments, the antioxidants N-acetyl-L-cysteine (NAC, 10 mM) and α-tocopherol (vitamin E, 10 µg/ml) were added to each well 1 h prior to treatment. For NO production, the iNOS substrate L-arginine was added to each well at 50 µM. All solutions were prepared in Krebs-Ringer-phosphate buffer (KRP). Fluorescence was measured using an Envision microplate reader (λ<sub>Ex</sub> 492 nm, λ<sub>Em</sub> 535 nm for DCFH-DA and DAF; λ<sub>Ex</sub> 492 nm, λ<sub>Em</sub> 615 nm for DHE).

2.4.4 Western Blot Analysis

After treatment, cells were washed twice with cold D-PBS and
lysed with RIPA buffer, sonicated, and centrifuged (16,000 × g, 5
min, 4°C). Supernatants were collected in a new tube and the
protein concentration determined using the BCA™ Protein Assay
kit according to the manufacturer's instructions. Proteins were
precipitated with methanol/chloroform (1/0.25; v/v) and
centrifuged for 5 min at 16,000 × g at room temperature. The
pellet was then resuspended in 4X Laemmli buffer and heated to
99°C for 5 min. Fifty µg of each protein sample was separated on a
12% SDS-PAGE gel and transferred onto a nitrocellulose
membrane. After blocking, membranes were incubated
overnight at 4°C with primary antibodies: rabbit anti-CD9
polyclonal antibody (1/1,000, GeneTex® GTX55564), goat anti-
CD81 polyclonal antibody (1/1,000, Santa Cruz Biotechnology,
Inc.® sc-31234), mouse monoclonal anti-β-actin antibody (1/
5,000, Sigma-Aldrich® A1978, clone AC-15), rabbit anti-
SMPD3 (1/10,000, Sigma-Aldrich® SAB2102245), or rabbit
anti-p62/SQSTM1 (1/1,000, Sigma-Aldrich® P0067). After
several washes, membranes were incubated with goat anti-
rabbit IgG-HRP (1/5,000, Santa Cruz Biotechnology, Inc.® sc-
2004), goat anti-mouse IgG-HRP (1/5,000, Santa Cruz
Biotechnology, Inc.®, sc-2005), or rabbit anti-goat IgG (H +
L)-HRP (1/5,000, Southern Biotech®, 6,160-05) for 1 h at room
temperature. Then, proteins were visualized using ECL™ Western
Blot Detection Reagents and a ChemiDoc™ MP Imaging System
(Bio-Rad®, Hercules, California, United States). β-actin protein
levels were quantified to normalize protein levels.

**2.5 RNA Extraction and Quantitative**
**Real-Time PCR**

After treatment, cells were washed twice with D-PBS. RNA
extraction was then performed using a mixture of TRIzol™
and chloroform (1/0.2; v/v). After a 15-min centrifugation
(12,000 × g, 4°C), RNA was collected and precipitated with
isopropanol. The RNA pellet was then washed twice with 70%
ethanol and resuspended in 40 µL RNase/DNase-free water.
The RNA concentration was determined using a NanoVue™
Plus device (Thermo Fisher Scientific®, Waltham,
Massachusetts, United States). cDNA was synthesized
using a High-Capacity RNA-to-cDNA™ kit according to

the manufacturer's instructions. Quantitative real-time
PCR was carried out using Takyon™ and specific primers
(Supplementary Table S1). A CFX96 Touch Real-Time PCR
Detection System (Bio-Rad®, Hercules, California,
United States) was used with the following steps: 95°C for
15 s, followed by 40 cycles of 95°C for 15 s, 60°C for 1 min and
72°C for 30 s. β-actin was used as a housekeeping gene to
normalize gene expression.

**2.6 MSD Multiplex Immunoassay**

Proinflammatory cytokine protein levels were measured in
macrophages and EVs derived from macrophages using a
proinflammatory panel multiplex kit (V-PLEX® K15048D,
Meso Scale Diagnostics®, Rockville, Maryland, United States)
according to the manufacturer's instructions. Briefly, 50 µL/well
of sample or calibrators was added to a 96-well plate pre-coated
with capture antibodies. After a 2-h incubation with shaking at
room temperature and three washes (D-PBS with 0.05% Tween
20), 25 µL 1X detection antibody solution was added to each
well. After another 2-h incubation under the same conditions
and three washes, 150 µL 2X Read Buffer T was added to each
well. The plate was then analyzed using the MSD instrument
(Meso Scale Diagnostics®, Rockville, Maryland, United States).

**2.7 Mass Spectrometry Proteomic Analysis**

**2.7.1 Proteomic Digestion for Mass Spectrometry**

An S-TrapTM microspin column digestion was performed on
10 µg of macrophage-derived EVs according to the
manufacturer's instructions (Protifi, Hutington, United States).
Briefly, samples were reduced with 20 mM tris (2-carboxyethyl)
phosphine and then alkylated with 50 mM chloroacetamide for
15 min at room temperature. Aqueous phosphoric acid was then
added to a final concentration of 1.2%, followed by the addition of
S-Trap binding buffer (90% aqueous methanol, 100 mM
tetraethylammonium bromide, pH 7.1). The mixtures were
then loaded onto S-Trap columns. Two extra washing steps
were performed to eliminate SDS. Samples were then digested
with 1 µg trypsin (Promega, Madison, Wisconsin, United States)
at 47°C for 1 h. After elution, peptides were vacuum dried and
resuspended in 45 µL 2% acetonitrile/0.1% formic acid mixture in
HPLC-grade water prior to MS analysis. A volume of 1 µL of the
peptide suspension was injected into a nanoelute high-performance
liquid chromatography (HPLC) system coupled to a timsTOF Pro
mass spectrometer (Bruker Daltonics, Germany). HPLC separation
was performed using a mixture of 0.1% formic acid in water, 2%
acetonitrile (Solvent A), and 0.1% formic acid in acetonitrile
(Solvent B) at 250 nL/min using a packed emitter column (C18,
25 cm × 75 µm 1.6 µm) (Ion Optics, Australia) with a gradient
elution (2–11% solvent B over r19 min, 11–16% over 7 min, 16–25%
over 4 min, 25–80% over 3 min, and, finally, 80% for 7 min to wash
the column). Mass-spectrometric data were acquired using the
parallel accumulation serial fragmentation (PASEF) acquisition
method. The measurements were carried out over an m/z range
from 100 to 1700 Th, with ion mobility from 0.8 to 1.3 V s/cm² (1/
k0). The total cycle time was set to 1.2 s and the number of PASEF
MS/MS scans was set to 10. A total of 2,762 proteins was identified
in at least 60% of all samples.
The mass spectrometry proteomics data have been deposited
in the ProteomeXchange Consortium via the PRIDE (Perez-
Riverol et al., 2019) partner repository with the dataset
identifier PXD029441 and 10.6019/PXD029441.

## 2.7.2 Data Analysis

The data were analyzed using MaxQuant version 1.6.14.0 (Max-Planck, Munich, Germany) and searched using the Andromeda search engine against the UniProtKB/Swiss-Prot *Mus musculus* database (release 02-04-2020, 17040 entries). Mass deviations of 3 and 20 ppm were used to search parent mass and fragment ions, respectively. The minimum peptide length was set to seven amino acids and strict specificity for trypsin cleavage was required, allowing up to two missed cleavage sites. Carbamidomethylation (Cys) was set as a fixed modification, whereas oxidation (Met) and N-term acylation were set as variable modifications. The false discovery rate (FDR) for both proteins and peptides was set to 1%. Scores were calculated in MaxQuant as previously described (Cox and Mann, 2008). The reverse and common contaminant hits were removed from the MaxQuant output. Proteins were quantified according to the MaxQuant label-free algorithm using LFQ intensities; protein quantification was obtained using at least two peptides per protein. Matches between runs were allowed. Statistical and bioinformatic analysis, including heatmaps, profile plots, and clustering, were performed using Perseus software (version 1.6.14.0) freely available at www.perseus-framework.org (Tyanova et al., 2016). For statistical comparisons, we set up five groups, each containing up to five biological replicates. We then filtered the data to keep only proteins with at least four valid values in at least one group. Next, the data were imputed to fill missing data points by creating a Gaussian distribution of random numbers with a standard deviation of 33% relative to the standard deviation of the measured values and a 1.8 standard deviation downshift of the mean to simulate the distribution of low-signal values. We performed an ANOVA test, FDR <0.05, S0 = 0.5. Hierarchical clustering of proteins that survived the test was performed using Perseus on log-transformed LFQ intensities after z-score normalization of the data using Euclidean distances.

## 2.8 Statistical Analysis

Statistical analyses were performed using GraphPad Prism
(version 7.0, San Diego, California, United States). Wilcoxon-
Mann-Whitney and Kruskal-Wallis tests were performed with a
significance threshold of 0.05. Data are expressed as the mean ±
standard error of mean (SEM) from at least three independent
experiments.

Principal Component Analysis has been performed on log
transformed and scaled values from targeted MSD multiplex
immunoassay using the R package pcaMethods.

# 3 RESULTS

## 3.1 Effects of LPS-EK on Macrophages

### 3.1.1 Lipopolysaccharide From *Escherichia coli* K12
Induces Inflammation in Macrophages

In setting up our cellular model of inflammation, we first verified the effect of LPS-EK on inflammation in RAW macrophages by measuring the mRNA levels of various pro-inflammatory cytokines (IL-6, IL-1β, TNF-α, and MIP-2), as well as that of the NLRP3 inflammasome marker, before and after treatment. The mRNA levels of these markers were all significantly higher after LPS-EK treatment than those of unstimulated macrophages (CT) (*p* < 0.05 vs. CT, **Figure 1**). We also analyzed the levels of 10 proinflammatory cytokines in LPS-EK-treated or untreated macrophages using an MSD multiplex kit. Macrophages treated with LPS-EK showed significantly higher levels of these proinflammatory cytokines than untreated control cells (*p* < 0.05 vs. CT, **Supplementary Figure S2**). These results show that LPS-EK induces an inflammatory response in our cellular model.

### 3.1.2 Lipopolysaccharide From *Escherichia coli* K12
Induces Oxidative Stress in Macrophages

As oxidative stress is also involved in the calcification process, we next determined the effect of LPS on reactive oxygen species (ROS) production using the fluorescent probe DCFH-DA. ROS production by cells treated with LPS-EK was significantly higher than that of untreated cells (*p* < 0.05 vs. CT) (**Figure 2A**). In addition, mRNA levels of two components of the antioxidant system, Keap1 and Nrf2, were lower after LPS-EK treatment, which could partially explain the increase in oxidative stress (*p* < 0.05 vs. CT, **Figures 2E,F**). Among the various ROS, superoxide anions ($O_2^{•-}$) play an important role in oxidative stress. We therefore investigated the effect of LPS-EK on $O_2^{•-}$ production using the fluorescent probe DHE. Surprisingly, $O_2^{•-}$ production was significantly lower in LPS-EK treated cells than in untreated cells (*p* < 0.05 vs. CT, **Figure 2B**). We investigated this seeming discrepancy by measuring the mRNA levels for both Nox-2, the enzyme that synthesizes $O_2^{•-}$, and superoxide dismutase (SOD-2), which is responsible for $O_2^{•-}$ degradation. We observed significantly higher Nox-2 and lower SOD-2 mRNA levels in LPS-EK stimulated macrophages than in untreated macrophages (*p* < 0.05 vs. CT, **Figures 2C,D**). Our results thus show that LPS-EK decreases superoxide anion production by decreasing Nox-2 expression and increasing SOD-2 expression. Superoxide anions can rapidly react with nitric oxide (NO) to form peroxynitrite (ONOO⁻). Thus, we next investigated the effect of LPS-EK on NO production using the fluorescent probe DAF. NO production was significantly lower after LPS-EK treatment than in untreated cells (*p* < 0.05 vs. CT, **Figure 2G**), despite significantly higher levels of iNOS (*p* < 0.05 vs. CT, **Figure 2H**). These results suggest that both decreases in NO and $O_2^{•-}$ production may be partially explained by the formation of peroxynitrite.
<figure><img src="image_4.png" /></figure>

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<figure><img src="image_7.png" /></figure>

<figure><img src="image_8.png" /></figure>

**FIGURE 1** | Lipopolysaccharide-EK (LPS-EK) induces inflammation in macrophages. RAW cells were incubated with LPS-EK for 6 h. Gene expression of (A) IL-6, (B) IL-1β, (C) TNF-α, (D) MIP-2, and (E) NLRP3 was quantified by quantitative real-time PCR and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. CT, Mann-Whitney test. LPS 0.1: 0.1 µg/ml LPS-EK, LPS 1: 1 µg/ml LPS-EK.

### 3.1.3 Lipopolysaccharide From *Escherichia coli* K12 Decreases Extracellular Vesicle Biogenesis in Macrophages

The aim of this study was to determine whether EVs can propagate inflammation and oxidative stress to recipient cells. Thus, we first studied EV biogenesis in RAW cells after LPS-EK treatment by analyzing the mRNA levels of various EV biogenesis markers (phospho-1, TNAP, and SMPD3) in LPS-EK-treated macrophages. The mRNA levels of these markers were all significantly lower in LPS-EK-treated cells than in unstimulated macrophages (*p* < 0.05 vs. CT, **Figures 3A-C**), suggesting a decrease in EV biogenesis. SMPD3 protein levels were also significantly lower after 1 µg/ml LPS-EK treatment than in untreated macrophages (*p* < 0.05 vs. CT, **Figure 3D**).

### 3.1.4 Lipopolysaccharide From *Escherichia coli* K12 Decreases Autophagy in Macrophages

Recent studies have highlighted novel functions of autophagy in the biogenesis and secretion of EVs (Yang et al., 2021).

Indeed, it has been shown that not only exosome biogenesis and autophagy share molecular machinery but also that substantial crosstalk exists between these two processes. We therefore analyzed the effect of LPS-EK on autophagy markers, such as ULK1, Beclin-1, Atg5, LC3a, and LC3b. The mRNA levels of all these markers were significantly lower after LPS-EK treatment than in unstimulated macrophages (*p* < 0.05 vs. CT, **Figures 4A-E**), suggesting a decrease in the autophagy process. p62, also known as sequestome-1 (SQSTM), is an autophagy cargo receptor (Lamark et al., 2009) that can be used as a sensor of autophagic flux. Indeed, p62 accumulates when autophagy is inhibited and decreased levels of p62 can be observed when autophagy is induced. After LPS-EK treatment, p62 mRNA levels were higher than in unstimulated macrophages (*p* < 0.05 vs. CT, **Figure 4F**), confirming inhibition of the autophagic flux. p62 protein levels were not modified after LPS-EK treatment relative to untreated cells (**Figure 4G**). Overall, our results show that LPS-EK not only induces inflammation and oxidative stress
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**FIGURE 2** | Lipopolysaccharide-EK (LPS-EK) induces oxidative stress in macrophages. ROS (A), O2⁻ (B), and NO (G) production in RAW cells was measured using the fluorescent probes DCF, DHE, and DAF, respectively. Gene expression of (C) Nox-2, (D) SOD-2, (E) Keap1, (F) Nrf2, and (H) iNOS was quantified by quantitative real-time PCR and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. CT, Mann-Whitney test. LPS 0.1: 0.1 µg/ml LPS-EK, LPS 1: 1 µg/ml LPS-EK.

in RAW cells but also decreases autophagy and EV biogenesis.

## 3.2 Effects of EV-LPS on Smooth Muscle Cells

### 3.2.1 Characterization of Macrophage-Derived EVs

We characterized macrophage-derived EVs by their morphology, size, and concentration (Figure 5) using transmission electron

microscopy (TEM) (Figure 5A), cryo-electron microscopy (cryo-EM) (Figure 5B), and nanoparticle-tracking analyses (NTA) (Figure 5C). The particle size for exosomes is between 50 and 150 nm and between 50 and 500 nm for microvesicles (van Niel et al., 2018). The average particle size detected by NTA was 139 nm for EVs isolated from untreated macrophages (EV-CT) and 130 nm for EVs isolated from LPS-EK-treated macrophages (EV-LPS) (Figure 5C). Thus, our EV samples were likely enriched for both exosomes and microvesicles. Furthermore,
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**FIGURE 3** | Lipopolysaccharide-EK (LPS-EK) decreases EV biogenesis in macrophages. RAW cells were incubated with LPS-EK for 6 h. Gene expression of EV of the biogenesis markers (A) phospho-1, (B) TNAP, and (C) SMPD3 was then quantified by RT-qPCR and normalized to that of the housekeeping gene β-actin. (D) SMPD3 protein expression was studied by western blotting and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. CT, Mann-Whitney test. LPS 0.1: 0.1 µg/ml LPS-EK, LPS 1: 1 µg/ml LPS-EK.

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**FIGURE 4** | Lipopolysaccharide-EK (LPS-EK) decreases autophagy in macrophages. RAW cells were incubated with LPS-EK for 6 h. Gene expression of the autophagy markers (A) ULK1, (B) Atg5, (C) LC3a, (D) LC3b, (E) Beclin-1, and (F) p62 was then quantified by RT-qPCR and normalized to that of the housekeeping gene β-actin. (G) p62 protein expression was studied by western blotting and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. CT, Mann-Whitney test. LPS 0.1: 0.1 µg/ml LPS-EK, LPS 1: 1 µg/ml LPS-EK.
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**FIGURE 5** | Characterization of macrophage-derived EVs. EVs were isolated from the culture media of RAW cells incubated with (EV-LPS) or without (EV-CT) 1 µg/ml lipopolysaccharide-EK (LPS-EK) for 6 h. Evaluation of EV morphology by (A) transmission electron microscopy (TEM) and (B) cryo-electron microscopy (cryo-EM). (C) Particle-size distribution and (D) the total concentration of macrophage-derived EVs were measured by nanoparticle-tracking analysis (NTA). (E) CD9, CD81, and β-actin protein expression were assessed by western blotting. (F) Quantification of the enzymatic activity of acetylcholinesterase (AChE) in macrophage-derived EVs. Control AChE activity in the assay diluent 1X-D-PBS (CT) was defined as 100%. Data are expressed as the mean ± SEM of seven independent experiments performed in triplicate (n = 7). \*\*\*p < 0.001 vs. CT; \$p < 0.05\ vs. EV-CT, Mann-Whitney test.

we verified EV membrane integrity by cryo-EM (Figure 5B). Indeed, we could observe a discernible lipid bilayer and internal vesicular structures (Figure 5B). The diameter of EVs can vary, as well as the content. The International Society for Extracellular Vesicles (ISEV) has published recommendations for EV characterization (Théry et al., 2018). Indeed, two categories of markers must be analyzed in sample preparations to confirm the presence of EVs: transmembrane proteins, such as tetraspanins, and cytosolic proteins recovered in EVs (Théry et al., 2018). Thus, we performed western blotting to determine the presence of EV-specific tetraspanin (CD9, CD81) and β-actin and detected all of these EV markers in our EV samples (Figure 5E). Acetylcholinesterase (AChE) is expressed in macrophages (Fujii et al., 2017) and can also be found in macrophage-derived EVs. According to the ISEV, AChE activity can also be used to characterize EVs (Figure 5F). Overall, these results confirm the

presence of EVs in both preparations. LPS-EK treatment of macrophages resulted in a lower number of EVs than in control untreated cells (Figure 5D). Indeed, we detected 7.54 × 10¹⁰ particles in the EV-CT sample, whereas the EV-LPS preparation contained 3.20 × 10¹⁰ particles. Furthermore, we observed significantly less AChE in the EV-LPS preparation than in the EV-CT preparation. Both results could be explained by the lower amount of secreted EVs due to decreased EV biogenesis after LPS-EK treatment ($p < 0.05$ vs. EV-CT, Figure 5F). Importantly, the EVs were isolated from the same volume of cell-culture media.

### 3.2.2 Effect of Macrophage-Derived EVs on MOVAS Cell Viability

We next studied the effects of macrophage-derived EVs on the smooth muscle cell line MOVAS-1. Cell viability was first assessed after 24 h of treatment with macrophage-derived EVs. EVs secreted
<figure><img src="image_34.png" /><figcaption>**FIGURE 6** | Effect of macrophage-derived EVs on smooth muscle cell viability. MOVAS-1 cells were incubated for 24 h with EVs secreted by either treated (1 µg/ml LPS-EK; EV-LPS) or untreated macrophages (EV-CT) or without (CT). Cell viability was measured using the WST-1 assay. The viability of cells incubated without EVs (CT) was defined as 100%. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). *p* < 0.05 is considered significant, Wilcoxon- Mann-Whitney test.</figcaption></figure>

by macrophages did not affect smooth muscle cell viability relative to untreated cells (CT), regardless of the treatment (Figure 6).

### 3.2.3 EV-LPS Induce Inflammation in Smooth Muscle Cells

Our objective was to study the propagation of inflammation from macrophages to smooth muscle cells. Thus, we measured the mRNA levels of proinflammatory cytokines (IL-6, IL-1β, and

TNF-α) in MOVAS cells treated with macrophage-derived EVs. We observed significantly higher proinflammatory cytokine mRNA levels after 24 h of treatment of smooth muscle cells with EV-LPS than in untreated cells (CT) (*p* < 0.05 vs. CT, **Figure 7**). There was no effect of EVs derived from control cells (EV-CT) on cytokine levels. A comparison of the effects of the two populations of EVs showed only the increase in IL-6 mRNA levels to be significant ($p < 0.05$ vs. EV-CT, **Figure 7A**). These results show that EV-LPS are able to induce inflammation mainly via an increase in IL-6 expression.

### 3.2.4 EV-LPS Induces Oxidative Stress in Smooth Muscle Cells

We investigated the effect of macrophage-derived EVs on oxidative stress in smooth muscle cells by measuring intracellular ROS production. Both EV preparations (EV-CT and EV-LPS) significantly increased ROS production over that of untreated smooth muscle cells (*p* < 0.05 vs. CT, **Figure 8**). This result suggests that the effect of macrophage-derived EVs on oxidative stress in smooth muscle cells is independent of macrophage treatment. We next measured mRNA levels of antioxidant enzymes (SOD-1, SOD-2) and nuclear receptors (Nrf2, and Keap1) involved in the antioxidant response. Surprisingly, the mRNA levels of all markers were significantly higher in cells incubated with EV-LPS than in untreated cells (CT) (*p* < 0.05 vs. CT, **Figure 9**). Only SOD-2 mRNA levels were higher in MOVAS cells treated with EV-LPS than those treated with EV-CT ($p < 0.05$ vs. EV-CT, **Figure 9B**). Overall, these results suggest that EV-LPS can affect the oxidative stress response of smooth muscle cells.

<figure><img src="image_35.png" /><figcaption>**FIGURE 7** | EV-LPS increases cytokine gene expression in smooth muscle cells. MOVAS-1 cells were incubated with RAW cell-derived EVs for 24 h. Gene expression of (A) IL-6, (B) IL-1β, and (C) TNF-α was then quantified by quantitative real-time PCR and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). *p* < 0.05 vs. CT, $p < 0.05$ vs. EV-CT, Mann-Whitney test. EV-CT: EV-derived from untreated macrophages, EV-LPS: EV-derived from LPS-EK-treated macrophages.</figcaption></figure>
<figure><img src="image_36.png" /><figcaption>**FIGURE 8** | Macrophage-derived EVs induce ROS production in smooth muscle cells. MOVAS-1 cells were incubated with 10 µM DCFH-DA in D-PBS at 37°C for 30 min and then treated with macrophage-derived EVs for 24 h. ROS production was determined by measuring fluorescence (λEx 492 nm, λEm 535 nm). ROS production by untreated control cells (CT) was defined as 100%. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. CT, Mann-Whitney test. EV-CT: EV-derived from untreated macrophages, EV-LPS: EV-derived from LPS-EK-treated macrophages.</figcaption></figure>

### 3.2.5 Analysis of the Content of EV-LPS

Inflammation is known to play an important role in vascular calcification. We thus first analyzed the inflammatory content of EVs by measuring the expression of 10 proinflammatory

cytokines in EV-LPS and EV-CT using an MSD multiplex kit. Proinflammatory cytokine levels were, indeed, significantly higher in EV-LPS than EV-CT6 (\*p < 0.05 vs. CT, **Supplementary Figure S3**). We further analyzed the data using an unsupervised analysis approach to visualize any samples that clustered based on their inflammatory profile. The principal component analysis score plot showed a clear separation between the inflammatory profiles of EV-CT6 and EV-LPS (**Figure 10**). These results show that there is a specific inflammatory profile that distinguishes between these two conditions. We then analyzed the EV protein content by mass spectrometry and detected several EV-specific proteins in our samples, confirming the presence of EVs in our preparations (**Supplementary Table S2**). Furthermore, the level of three proteins in EV LPS were higher than in EV CT6: cis-aconitate decarboxylase (CAD), encoded by the *immunoresponsive gene 1* (*Irg1*); plasminogen activator inhibitor-1 (PAI-1), encoded by the *Serpine1* gene; and serum amyloid A-3 protein (Saa3), encoded by the Saa3 gene (*p* < 0.01 vs. EV CT6, **Figure 11**).

### 3.2.6 EV-LPS Increase Pi-Induced Calcification by Inducing the Osteogenic Switch of Smooth Muscle Cells

Oxidative stress and inflammation are two processes known to play an important role in vascular calcification. Our results show that EV-derived macrophages can induce both oxidative stress and inflammation. Thus, we next determined the effect of these EVs on Pi-induced calcification in smooth muscle cells. MOVAS cells were simultaneously treated with 4 mM Pi and macrophage-derived EVs for 14 days (**Figure 12A**) or macrophage-derived

<figure><img src="image_37.png" /><figcaption>**FIGURE 9** | EV-LPS increase antioxidant marker gene expression in smooth muscle cells. MOVAS-1 cells were incubated with EVs-derived RAW cells for 24 h. Gene expression of (A) SOD-1, (B) SOD-2, (C) Nrf2, and (D) Keap1 was then quantified by quantitative real-time PCR and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. CT, \$p < 0.05\ vs. EV-CT, Mann-Whitney test. EV-CT: EV-derived from untreated macrophages, EV-LPS: EV-derived from LPS-EK-treated macrophages.</figcaption></figure>
<figure><img src="image_38.png" /><figcaption>**FIGURE 10** | Principal component analysis score plot of EV-CT6 and EV-LPS. EVs were isolated from the culture media of RAW cells incubated with (EV-LPS) or without (EV-CT6) LPS-EK. Proinflammatory cytokine protein levels (IL-1β, IL-2, IL-4, IL-5, IL-6, IL-10, IL-12p70, TNF-α, IFN-γ, and KC-GRO) were measured in EVs derived from the macrophages using an MSD multiplex immunoassay. PCA analysis score plot is performed using four independent experiments (n = 4).</figcaption></figure>

EVs for the last 7 days only (Figure 12B) of the induction of calcification. The intracellular calcium concentration in the smooth muscle cells was significantly higher after 14 days of treatment with Pi and EV-LPS than in cells treated with Pi and EV-CT ($p < 0.05$ vs. EV-CT, **Figure 12A**). Moreover, the intracellular calcium concentration was significantly higher after 7 days of treatment of Pi-treated smooth muscle cells with EV-LPS than Pi treatment alone ($*p < 0.05$ vs. 4 mM Pi, **Figure 12B**). These results suggest that EV-LPS can significantly induce calcification in MOVAS-1 cells. We can rule out that this effect was due to the presence of LPS-EK in EVs, as LPS-EK alone, with or without 4 mM Pi, had no effect on calcification (data not shown). We next measured the mRNA level of matrix gla protein (MGP), an inhibitor of vascular calcification. The level of MGP mRNA was significantly lower after 7 days of treatment of smooth muscle cells with 4 mM Pi and EV-LPS than in cells treated with Pi alone ($*p < 0.05$ vs. 4 mM Pi, **Figure 13**). The observed increase in calcification may therefore be partially due to a decrease in the level of calcification inhibitors, such as MGP.

Smooth muscle cells can adopt a contractile, synthetic, or osteochondrogenic phenotype, depending on their environment (Durham et al., 2018). During vascular calcification, smooth muscle cells undergo an osteogenic switch to become osteoblast-like cells (Durham et al., 2018). We investigated whether this phenotypic change occurred in our experiments by measuring the mRNA levels of several osteogenic markers, such as osterix (Osx) and osteocalcin (OCN), as well as alpha-smooth muscle actin

<figure><img src="image_39.png" /><figcaption>**FIGURE 11** | Volcano plot representing the effects of lipopolysaccharide-EK (LPS-EK) on the protein content of macrophage-derived extracellular vesicles (EVs). EVs were isolated from the culture media of RAW cells incubated with (EV-LPS) or without (EV-CT6) lipopolysaccharide-EK (LPS-EK). High-performance liquid chromatography (HPLC) coupled to mass spectrometry analysis was performed. An unpaired Student's t test was conducted and a p-value < 0.01 and a false discovery rate (FDR) < 5% were used. Five independent experiments were performed.</figcaption></figure>
<figure><img src="image_40.png" /><figcaption>**FIGURE 12** | EV-LPS enhance Pi-induced calcification in smooth muscle cells. Calcification in MOVAS-1 cells was induced by incubation with 4 mM Pi with or without RAW cell-derived EVs for (A) 14 days or (B) during the last 7 days of a 14-day calcification induction. Calcification was then measured using the OCP method. Data are expressed as the mean ± SEM of four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. 4 mM Pi, \$p < 0.05$ vs. EV-CT, Mann-Whitney test. EV-CT: EVs derived from untreated macrophages, EV-LPS: EVs derived from LPS-EK-treated macrophages.</figcaption></figure>

<figure><img src="image_41.png" /><figcaption>**FIGURE 13** | EV-LPS decrease MGP gene expression in smooth muscle cells. MOVAS-1 cells were incubated for 7 days with 4 mM Pi with or without EVs secreted by RAW cells. Gene expression of MGP was then quantified by quantitative real-time PCR and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of seven independent experiments performed in triplicate (n = 7). \*p < 0.05 vs. 4 mM Pi, Mann-Whitney test. EV-CT: EVs derived from untreated macrophages, EV-LPS: EVs derived from LPS-EK-treated macrophages.</figcaption></figure>

(α-SMA), a marker of the contractile phenotype. Osx and OCN mRNA levels were significantly higher in Pi-treated smooth muscle cells incubated with EV-LPS than Pi-treated smooth muscle cells, suggesting an osteogenic switch of the MOVAS-1 cells (*p* < 0.05 vs. 4 mM Pi, **p** < 0.01 vs. 4 mM Pi, **Figures 14A,B**). We also observed higher Osx and Ocn mRNA levels after treatment of Pi-treated smooth muscle cells with EV-CT, but without reaching significance. This result can be explained by the higher amount of EVs in the EV-CT preparation. At the same time, α-SMA mRNA levels were significantly lower than in Pi-treated smooth muscle cells (**p* < 0.01 vs. 4 mM Pi, **Figure 14C**). Overall, these results show that EV-LPS induce an osteogenic switch of smooth muscle cells, which could, in turn, promote calcification.

## 4 DISCUSSION

Vascular calcification (VC) is a complex process involving various molecular and cellular mechanisms, such as the VSMC osteogenic switch, loss of VC inhibitors, cell death, and dysregulation of Ca²⁺/Pi homeostasis, as well as matrix degradation and modification (Lee et al., 2020). Macrophages are among the main sources of inflammation and oxidative stress (Castaneda et al., 2017) and associated with arterial calcification (Agharazii et al., 2015). Indeed, Aikawa et al. showed co-localization of calcification and macrophages in atherosclerotic plaques of apolipoprotein E-deficient mice by image analysis, suggesting a prominent role of these cells and inflammation in calcification (Aikawa et al., 2007). In addition, a number of studies have shown macrophage-derived EVs to be mediators of VC (New et al., 2013; Chen et al., 2016; Li Y. et al., 2020; Kawakami et al., 2020). In this context, we hypothesized that macrophage-derived EVs secreted under pro-inflammatory and pro-oxidative conditions may increase VSMC calcification by propagating inflammation and oxidative stress. We tested our hypothesis by treating a murine VSMC cell line with EVs derived from LPS treated-murine macrophages. We then analyzed the effect of macrophage-derived EVs on the calcium content and levels of inflammatory, oxidative stress, and osteogenic markers in VSMCs. Moreover, we analyzed the protein content of EVs secreted by such activated macrophages.

We show that LPS-EK induces an inflammatory response in macrophages, as demonstrated by the significant increase in mRNA levels of pro-inflammatory cytokines (IL-6, IL-1β, and TNF-α). These results are in accordance with those in the literature, as activation of TLR4 by LPS triggers activation of the MyD88-dependent pathway, inducing pro-inflammatory cytokine gene transcription. NLRP3, an activator protein of the inflammasome has also been shown to be significantly upregulated after LPS treatment (He et al., 2016). Our results show that LPS-EK induces pro-inflammatory M1 polarization of RAW cells, in accordance with the results of a study of Li et al. performed in the U937 monocyte cell line (Li et al., 2017). LPS-EK also induced oxidative stress, as shown by the significant increase in ROS production, as previously reported. In parallel,
<figure><img src="image_42.png" /><figcaption>**FIGURE 14** | EV-LPS increase osteogenic marker gene expression and decrease contractile marker gene expression in smooth muscle cells. MOVAS-1 cells were incubated with 4 mM Pi, with or without RAW cell-derived EVs, for 7 days. Gene expression of (A) osterix, (B) osteocalcin, and (C) α-SMA was then quantified by quantitative real-time PCR and normalized to that of the housekeeping gene β-actin. Data are expressed as the mean ± SEM of at least four independent experiments performed in triplicate (n = 4). \*p < 0.05 vs. 4 mM Pi, \*\*p < 0.01 vs. 4 mM Pi, Mann Whitney test. EV-CT: EVs derived from untreated macrophages, EV-LPS: EVs derived from LPS-EK-treated macrophages.</figcaption></figure>

we showed a significant decrease in the expression of genes involved in the antioxidant system, such as Nrf2/Keap. Nrf2 is a transcriptional factor that induces the expression of antioxidant genes. Under basal conditions, Keap1 forms a complex with Nrf2 to induce its degradation by the proteasome. Under conditions of oxidative stress, modifications of the cysteine residues of Keap1 inhibit its interaction with Nrf2, allowing its nuclear translocation and protection of cells against oxidative stress. The observed decrease in Keap1 mRNA levels could be explained by the induction of protective mechanism activated by the cells to counteract ROS production. Surprisingly, despite an increase in ROS production, we showed a significant decrease in O₂⁻- production, which can be explained by both a decrease in Nox-2 levels and an increase in its degradation by SOD-2. This discrepancy can be partially explained by the fact that O₂⁻ can rapidly react with NO to form peroxynitrite (ONOO⁻), which mediates oxidative responses. Indeed, under LPS treatment, we observed a decrease in NO production, despite an increase in iNOS, an enzyme that produces NO.

As already shown (Liu et al., 2017; Bell et al., 2019), LPS-EK inhibited EV biogenesis by decreasing the expression of several markers of EV biogenesis, such as sphingomyelin phosphodiesterase 3 (SMPD3), phospho-1, and tissue nonspecific phosphatase alkaline phosphatase (TNAP). These results were confirmed by NTA analysis, which showed a reduced number of secreted EVs after LPS treatment. A

similar result was reported by Bell *et al.* in AC16 human cardiomyocytes (Bell *et al.*, 2019). In our cellular model, LPS-EK significantly decreased the expression of autophagic markers (Atg5, beclin1, ULK). Autophagy has already been shown to play an important role in EV biogenesis (Yang *et al.*, 2021). This was confirmed by a significant increase in p62 expression after LPS treatment, as it has been shown that p62 protein accumulates upon inhibition of autophagy (Lamark *et al.*, 2009). Interestingly, Liu *et al.* recently demonstrated a link between autophagy and macrophage polarization, showing that inhibition of autophagy induces the polarization of macrophages towards a pro-inflammatory M1 phenotype (Liu *et al.*, 2015), a phenotype that we observed in our cells after treatment with LPS-EK. Moreover, Zhao *et al.* have shown that autophagy contributes to redox homeostasis, not only by clearing oxidized cellular components, but also by promoting antioxidant defenses *via* the p62/Keap1/Nrf2 pathway (Zhao *et al.*, 2019). Therefore, a deficiency in autophagy, as observed in our cellular model, would decrease antioxidant defenses, which could also explain the observed increase in oxidative stress.

We investigated whether EVs secreted by LPS-treated cells play a role in VC by incubating VSMCs with EV-LPS. We show that EV-LPS induces inflammation and oxidative stress in VSMCs, with a significant increase in the expression of pro-inflammatory cytokines (IL6, IL1β, TNFα). Assessment of the inflammatory profile of the secreted EVs showed that EV-LPS are
in fact enriched in these proinflammatory cytokines, consistent
with the M1 polarization of RAW cells observed after treatment
with LPS-EK. Cytokines are key players in cell-cell
communication and play important roles in several biological
processes, such as cell differentiation and inflammation. Various
secretory pathways are responsible for their release, such as the
classical ER/Golgi route and the unconventional pathway, also
known as unconventional protein secretion (UPS). Recently,
cytokines have been shown to be capable of reaching the
extracellular milieu *via* a novel secretory pathway through
EVs. Fitzgerald et al. thus showed that cytokine encapsulation
into EVs is a general biological phenomenon observed *in vitro*
and *in vivo* (Fitzgerald et al., 2018). Interestingly the profile of
such encapsulated cytokines changes in response to various
stimuli, suggesting that cytokine association with EVs is not
specific to any particular cytokine, as all cytokines can be
encapsulated. Tokarz et al. showed, for example, that the
association of specific cytokines with EVs is strongly
influenced by disease duration and treatment in diabetes
(Tokarz et al., 2015). Such an association between cytokines
and EVs appears to result from a specific physiological need,
depending on whether the cytokines act near the Ev-secreting
cells or at a distance. Indeed, Fitzgerald et al. (2018) showed that
tissue explants, in which cells are in proximity with each other,
secrete more soluble cytokine than cells in suspension, in which
cytokines are more highly associated with EVs to allow their
interaction with recipient cells at a distance. It is therefore
possible that pro-inflammatory cytokines from M1
macrophages are carried by EVs secreted by these cells to
induce a pro-inflammatory microenvironment for recipient
cells, such as VSMCs. Moreover, several studies have shown
that inflammatory cytokines have biological effects in VSMCs
(Nilsson, 1993). For example, Barillari *et al.* showed that IL-1β,
TNF-α, and IFN-γ released by activated immune cells enhance
the expression of α5β1 integrin, a fibronectin receptor, leading to
an increase in VSMC proliferation and migration, two
mechanisms required for the formation of atherosclerotic
lesions (Barillari *et al.*, 2001). Our results also show that EV-
LPS induce oxidative stress, as demonstrated by increased ROS
production and decreased expression of antioxidant enzyme
genes. However, EVs from non-treated cells (EV-CT) were
also able to induce oxidative stress. Nevertheless, it is
worthwhile noting that we treated cells with equivalent
volumes of EV preparations. As LPS-treated cells secreted
fewer EVs than control cells, it is likely that using the same
volumes to treat VSMCs resulted in treating them with a lower
number of EV-LPS than EV-CT, thus underestimating the effects
of EV-LPS on VSMCs. These results confirm our hypothesis that
EVs secreted under pro-inflammatory and pro-oxidative
conditions are able to propagate inflammation and oxidative
stress to surrounding cells, such as VSMCs. Finally, EV-LPS
enhanced Pi-induced calcification of VSMCs by inducing the
VSMC osteogenic switch and decreasing expression of the
calcification inhibitor, MGP. These effects were intrinsic to
EVs, as LPS alone had no effect on calcification. Moreover, in
a similar manner, the study of Li *et al.* showed that the
conditioned medium of LPS-treated macrophages induces an

osteogenic switch of valve interstitial cells by increasing OPN,
BMP-2, and ALP expression (Li et al., 2017). Furthermore, the
aforementioned study showed that the increase in IL-6, IL-1β,
and TNF-α in the conditioned medium was associated with an
increase in the production of MMPs, which contribute to
extracellular matrix degradation, remodeling, and valve
calcification (Li et al., 2017).

Proteomic analysis highlighted the upregulation of three proteins
in EV-LPS relative to EV-CT: cis-aconitate decarboxylase (CAD),
plasminogen activator inhibitor-1 (PAI-1), and serum amyloid A-3
protein (Saa3). CAD, a mitochondrial enzyme encoded by
immunoresponsive gene 1 (Irg1) and involved in itaconate
production, is known to be upregulated in macrophages under
pro-inflammatory conditions (Basler et al., 2006; Németh et al.,
2016; Tallam et al., 2016; Song et al., 2020). Itaconate has been
shown to be an immunoregulatory and anti-oxidant molecule
(Lampropoulou et al., 2016; Mills et al., 2018; Li R. et al., 2020;
Song et al., 2020). Itaconate has also been shown to promote IL-1β
production and inflammatory apoptosis when administered at high
doses to bone marrow-derived dendritic cells (Muri et al., 2020). In
addition, the production of ROS can be mediated by Irg1 induction
(Tan et al., 2016). In our study, EV-LPS were enriched in CAD. It is
thus possible that this enzyme may induce the production of itaconate
in VSMCs when transferred to the recipient cells via EVs, leading to
inflammation and oxidative stress. PAI-1 is known to inhibit the
action of plasminogen activators, such as tPA and uPA. First, several
studies have already shown the upregulation of PAI-1 expression in
LPS-treated cells (Wang et al., 2014; Ren et al., 2015) and an
immunoregulatory role for PAI-1 through the TLR4 signaling
pathway (Gupta et al., 2016). PAI-1 levels also increase under
oxidative conditions (Vulin and Stanley, 2004). The higher amount
of PAI-1 found in EV-LPS could be explained by macrophage
activation. Numerous studies have shown PAI-1 to be associated
with atherosclerotic lesions. Indeed, PAI-1 levels are high in
atherosclerotic coronary arteries (Schneiderman et al., 1992; Lupu
et al., 1993; Raghunath et al., 1995; Padró et al., 1997). PAI-1
expression was also found to be higher in CKD patients than
healthy individuals (Ouyang et al., 2013). Furthermore, a number
of studies have shown that PAI-1 is linked to VC. The upregulation of
PAI-1 was, indeed, shown to be proportional to the calcium content in
65 calcified aortic valves (Kochtebane et al., 2014). Another study
showed PAI-1 to positively correlate with vascular media thickness
and calcification (Ouyang et al., 2013). PAI-1 transported by EV-LPS
could thus participate in the aggravation of the VC process, as
observed under our experimental conditions. Finally, Saa3, a
member of apolipoproteins associated with high-density
lipoprotein (HDL) in plasma, was only overexpressed in EV-LPS.
As for CAD and PAI-1, Saa3 was upregulated after LPS treatment
(Meek et al., 1992; Reigstad et al., 2009). Several studies have shown
Saa proteins to have cytokine-like activity and to be able to activate
several receptors, such as TLRs, and transcription factors, such as NF-
κB (Ye and Sun, 2015). In addition, a number of studies have
highlighted the role of Saa proteins in calcification (Tanaka et al.,
2011; Ebert et al., 2015; Zhang et al., 2017). Saa proteins induce the
production of pro-inflammatory cytokines and the osteogenic
differentiation of mesenchymal stem cells *via* the TLR4 receptor
(Ebert et al., 2015). Zhang *et al.* also showed that Saa proteins can
induce the VSMC osteogenic switch through the p38 MAPK signaling pathway (Zhang et al., 2017). Furthermore, Saa proteins can increase calcium entry in human coronary artery smooth muscle cells (Tanaka et al., 2011). Thus, it is possible that Saa3 transported by EVs increases Pi entry into VSMCs and enhances the VSMC osteogenic switch.

In conclusion, we show a direct contribution of macrophages in the microcalcification process *via* EV secretion, an alternative pathway, in addition to the VSMC osteogenic switch. Indeed, EV-LPS, enriched for molecules involved in inflammation, oxidative stress, and VC mechanisms, were able to create an inflammatory microenvironment for surrounding cells, such as VSMCs, which in turn underwent an osteogenic switch, leading to increasing calcification. As atherosclerotic plaques containing microcalcifications are more susceptible to rupture and cardiovascular accidents, this study suggests that EVs could be used as non-invasive biomarkers to better stratify patients with a high risk of CV. Moreover, such EVs could therefore also be a therapeutic target to limit VC in patients.

## DATA AVAILABILITY STATEMENT

The data presented in the study are deposited in the PRIDE repository, accession number PXD029441(http://www.proteomexchange.org).

## AUTHOR CONTRIBUTIONS

AB and JA designed the experiments and performed the relevant literature research and data analysis. LY contributed

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## FUNDING

LY was financially supported by the French Research Ministry. This work was financially supported by the Université de Picardie Jules Verne.

## ACKNOWLEDGMENTS

This work benefited from the assistance of Stephanie Balor and Vanessa Soldan from the Multiscale Electron Imaging platform (METI) of the Centre de Biologie Intégrative (CBI), Toulouse, France.

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