Around 8- to 10-week-old C57BL/6 male mice (20–25 g) were obtained from the Center for Experimental Models Development for Medicine and Biology (CEDEME) of the Federal University of São Paulo (UNIFESP). Additionally, B6:129P CX3CR1^+/GFP and B6:129 (Cg) CCR2^+/RFP mice were obtained from Jackson Laboratory (Bar Harbor, ME, USA), and these inbred strains were backcrossed to generate CCR2^+/RFP CX3CR1^+/GFP mice. All mice were housed in a clean room with constant temperature (22°C) and a light/dark cycle of 12/12 h. The mice received a solid diet and filtered water ad libitum. All animal protocols and experiments were approved by the Ethics Committee on Animal Experimentation of UNIFESP with accession number CEP 685598.

Adipose-derived MSCs were isolated from epididymal fat of C57BL/6 mice (n = 10) as previously published (20). Briefly, adipose tissue was washed with PBS-containing penicillin/streptomycin 50 U/mL (Gibco, Thermo, USA) and minced with surgical scissors. Subsequently, the tissue was incubated with collagenase type IA solution 0.075% (Sigma-Aldrich, USA) at 37°C for 30 min. After digestion, the enzyme activity was neutralized by adding DMEM Low Glucose culture medium (Gibco) with 10% fetal bovine serum (FBS, Hyclone, Thermo Fisher Scientific). The digested material was centrifuged at 300 g for 10 min, and the pellet was subjected to red blood cells lyse solution treatment with NH₄Cl 0.83%. Then, the cell pellet was twice washed with PBS and centrifuged at 300 g for 5 min. The cells were incubated at 37°C in humid atmosphere with 5% CO₂ with DMEM Low Glucose with antibiotics (penicillin/streptomycin 100 U/mL) and 10% FBS. At confluence of 80–90% of the cell monolayer, MSCs were trypsinized (0.05%, Trypsin-EDTA, Gibco) and further expanded to 10–12 passages. The immunophenotypic characterization and of multilineage differentiation of MSCs was performed according to previous works from our group (20, 21).

Mesenchymal stromal cells were expanded until 80% of confluence with DMEM low glucose (10% FBS). At high density confluence, culture growth medium was changed to serum-free DMEM low glucose. After 48 h, the conditioned medium was collected and centrifuged at 2,000 g for 30 min at 4°C to remove cells and cellular debris. Then, the conditioned medium was subjected to ultracentrifugation at 100,000 g for 2 h at 4°C, using a SW28 rotor (Optima L-90K, Beckman Coulter, USA). Finally, the pellet containing MVs-MSCs was re-suspended in PBS and stored at −80°C. MVs-MSCs protein content was quantified by Pierce method (Thermo Fisher Scientific) and characterized using flow cytometry (BD LSRFortessa, BD biosciences) and nanoparticle tracking analysis (NTA)-based methods (NanoSight, Malvern, UK). Transmission electron microscopy was performed using protocols established in our center of facilities for electron microscopy at Federal University of São Paulo (CEME).

Microvesicles-MSCs were analyzed by NTA using NanoSight LM10 (NanoSight, UK). The laser was set at 405 nm and the speed of the high-sensitivity camera shutter was set at 30.01 ms (OrcaFlash2.8). The videos were analyzed using NTA software (version 2.3 NanoSight), and recordings were made for 30 s per sample. All samples were diluted in ultrapure PBS (Gibco, Thermo, USA) to achieve a concentration of 2 × 10⁸ and 9 × 10¹⁰ particles/mL. Samples were administered and registered under controlled flow, using NanoSight syringe pump system. Finally, histograms were generated as results of NTA analyzes to evaluate particle concentration and average size of MVs-MSCs.

After ultracentrifugation, MVs-MSCs containing pellet was resuspended in 100 μL of 2% buffered formaldehyde solution. Then, drops of 20 μL were placed on parafilm in a flat surface. Formvar-carbon-coated electron microscopy grids were placed on the drops for 10 min for MVs adsorption. Grids were washed with 0.1M sodium cacodylate buffer, post fixed in 1% buffered glutaraldehyde, and washed with deionized water. Negative-positive staining was performed according to Théry et al. (22). Briefly, grids were contrasted in oxalic uranyl, then embedded in a mixture of saturated uranyl and methylcellulose 2% (1:9) on ice. Fluid excess was gently blotted on filter paper and air dried. Images acquisition and observation was run under a JEOL 1200 EX II (Japan) transmission electron microscope at 80 kV.

Femurs and tibia of C57BL/6 male mice were dissected, and the epiphyses were surgically removed with scissors to perform internal flushing with sterile PBS. The cells thus obtained from bone marrow were filtered using Cell Steiner 70 μm filter (Corning, USA), and the flow-through cells were twice centrifuged and washed at 300 g for 5 min. Subsequently, cells were lysed with 0.83% NH₄Cl (3 min/4°C) and cultured in DMEM high glucose medium (Gibco) containing 10% FBS (Gibco) and 30% of supernatant medium from L929 cells (ATCC, Cell Lines) (23). The culture medium was refreshed on day 4 and subsequently maintained until day 7 to promote BMDM differentiation. On day 7, 2 × 10⁵ Mϕs/cm² were polarized to M1 by 5 ng/mL IFN-γ (R&D Systems, USA) and 50 ng LPS/mL (E. coli-LPS, Sigma-Aldrich, USA) incubation for 24 h. In contrast, M2 polarization was induced by 10 ng IL-4/mL and 10 ng IL-13/mL (R&D Systems) for 24 h.

M1-Mϕs were seeded at a concentration of 2 × 10⁵ Mϕs/cm² and co-cultivated with 1 × 10⁵ MSCs in cell-to-cell contact systems (1:2) for 48 h in DMEM high glucose with 10% FBS and samples collected with 72 h of total incubation (Figure S1E in Supplementary Material). On the other hand, in experiments using MVs-MSCs, we treated M1-Mϕs (2 × 10⁵ Mϕs/cm²) with serial doses of 4.19 × 10⁹ particles of MVs-MSCs every 8 h for a total period of 48 h in DMEM high glucose with 10% FBS and samples collected with 72 h total incubation (Figure S1F in Supplementary Material). As control, M1-Mϕ was treated with vehicle PBS (Gibco, Thermo, USA) using similar conditions as the above mentioned. M1-Mϕs control was cultured for 48 h in DMEM high glucose with 10% SFB but without stimulus or MSCs/MVs-MSCs. Finally, unstimulated Mϕs were maintained in culture for 72 h as an internal control of M1 polarization. All experiments conducted with MSCs or MVs-MSCs in coculture assays with Mϕs were evaluated after 72 h of incubation (Figures S1E,F in Supplementary Material). In addition, to observe MVs-MSCs internalization into M1-Mϕs, a single dose of MVs-MSCs (4.19 × 10⁹ particles) were pre-labeled with PKH26 (Sigma-Aldrich) and administered onto monolayers of 2 × 10⁵ Mϕs/cm². Images and videos were acquired during 4 h in LSM 880 Laser Scanning Microscope Zeiss (Carl Zeiss, Germany).

For Mϕs multi-step analyses by flow cytometry (FACS), the following markers were used: F4/80, CD11b, CD86, CCR7, CD206, Ly6C (BioLegend, USA) Siglec-F (BD biosciences, USA), and iNOS (BD biosciences, USA). Before staining, Mϕs were incubated with mouse Fc Block (BD Biosciences, USA) for 5 min at 4°C to prevent non-specific antibody interactions. Cells were then washed with staining buffer (BD Biosciences, USA) and centrifuged at 300 g for 5 min. For surface markers detection, Mϕs were incubated with staining buffer containing the respective mix of antibodies for 30 min at 4°C, protected from light. Next, cells were washed and acquired by FACSCanto II our LSRFortessa (BD biosciences, USA). For intracellular staining, the cells were previously incubated with GolgiStop (BD biosciences, USA) by 4 h following the manufacturer’s recommendations. Subsequently, for iNOS detection, Mϕs were treated with fixation/permeabilization solution (BD Biosciences, USA) and incubated with the respective antibody. Next, cells were washed and samples were acquired by FACS. Compensation controls were performed by CompBeads (BD Biosciences, USA), and acquisition controls were based on the fluorescence minus one control method. For MVs-MSCs immunophenotyping, the following antibodies were used: CD90, CD105, CD73, CD31, CD45, CD44 (BD biosciences, USA), and primary CD9 antibody (Abcam, USA), which was subsequently conjugated with Alexa fluor 488 (Abcam, USA). To detect surface antigens in MVs-MSCs, the particles were incubated with the respective antibody in single color staining for 30 min at 4°C. MVs-MSCs were then ultracentrifuged and washed twice in PBS, and subsequently analyzed by FACS. MVs-MSCs populations were set up with the assistance of sizes defined beads (3.8 and 7 μm) and additionally with annexin V antibody (BD biosciences, USA). All data were collected by BD FACSDiva and analyzed using FlowJo V10 software (Tree Star, USA).

Thioglycollate medium 3% (Sigma-Aldrich) was prepared in sterile conditions and protected from light at least 6 months before experiments. Mice were injected intraperitoneally (IP) with 2 mL of thioglycollate solution and euthanized on day 4. Mice treated with MVs-MSCs were injected IP with four doses of MVs-MSCs (8.38 × 10⁹ particles) at 0, 24, 48, and 72 h. In contrast, control mice were treated at the same time-points but using vehicle (PBS). Finally, on the 4th day after thioglycollate infusion, an peritoneal lavage was performed with 5 mL of cold sterile PBS. The peritoneal exudate was centrifuged at 300 g for 5 min and cells were processed by FACS and the supernatant was analyzed by MILLIPLEX (Merck Millipore, USA). During the standardization of the model of peritonitis, washes at 24, 48, 72, and 96 h were performed to identify the cell cluster with GR1, CD19, and CD11b phenotype by FACS (Figures S4A,B in Supplementary Material). Furthermore, the predominant presence of Mϕs in the last day of the induction of peritonitis was confirmed by morphological analysis (HE) and immunohistochemistry for CD11b (Figure S4C in Supplementary Material).

The concentrations of IFN-γ, IL-1α, IL-1β, TNF-α, IL-6, and IL-10 cytokines in the mouse peritoneal lavage were measured using MILLIPLEX MCYTOMAG-70K kit (Millipore Corporation, USA). Controls and samples were processed according to manufacturer’s instructions. The sample detection was performed by Luminex SD device (Luminex, USA) using the software MILLIPLEX Analyst (Millipore Corporation).

Total RNA derived from Mϕs was isolated using Trizol Reagent (Thermo Fisher Scientific), and its concentration and purity were determined using Nanodrop (Thermo Fisher Scientific). The cDNA was then synthesized using M-MLV reverse transcriptase (Promega, USA) following the manufacturer’s specifications. Subsequently, qPCR reactions were performed using the 7300 Real-Time PCR System (Applied Biosystems, USA) with TaqMan or SYBR green assays. For the TaqMan method, the following probes were used: IL-10 (Mm00439616_m1), IL-6 (Mm00446190_m1), Nos2 (Mm01309902_m1), IL-1β (Mm00434228_m1), irf4(Mm01165980_m1), Arg1 (Mm00475990_m1), Hprt (Mm00446968_m1). For the SYBR green assays, these previously validated pair primers were used: Fizz1 5′-TCCCAGTGAATACTGATGAGA-3′ and 5′-CCACTCTGGATCTCCCAAGA-3′; Ym1 5′-GGGCATACCTTTATCCTGAG-3′ and 5′-CCACTGAAGTCATCCATGT-3′; SOCS3 5′-GGGTGGCAAAGAAAAGGAG-3′ and 5′-GTTGAGCGTCAAGACCCAGT-3′; SOCS1 5′-ACCTTCTTGGTGCGCGAC-3′ and 5′-AAGCCATCTTCACGCTGAGC-3′; CXCL9 5′-TGCACGATGCTCCTGCA-3′ and 5′-AGGTCTTTGAGGGATTTGTAGTGG-3′. The results were analyzed by SDS Software (Applied Biosystems, USA), and gene expression was normalized by the expression of the internal control gene Hprt. The analysis of each sample was performed in triplicates and data were expressed by relative quantification method URE (10,000/2^ΔCt) (24). Furthermore, for miRNAs analysis, the total RNA was extracted from Mϕs using the miRNeasy Mini Kit (Qiagen, DEU) according to manufacturer’s instructions. Around 2 μg of RNA was used to amplify mature miRNAs present in the total RNA. This technique was performed using the miScript II RT kit (Qiagen) according to manufacturer’s instructions. The miR-155, miR-21, and miR-146a (Primer Assays, Qiagen) primers were used for miRNAs expression. The qPCR reaction for miRNAs was performed with SYBR Green PCR kit (Qiagen) using 7300 Real-Time PCR System.

The enzymatic activity of arginase was measured using an in house method. Total protein lysate from Mϕs was diluted 1:10 in 50 mM Tris-HCl buffer pH 7.4 containing 0.5 M PMSF and protease inhibitors. Then, 120 μL of each sample was transferred to 1.5 mL tubes and 120 μL of MnCl₂ 10 mM were added to each tube, which were further heated to 55°C for 10 min. After incubation, the tubes were placed on ice and 240 μL of L-Arg (0.5 M) were added. Then each sample was divided into additional four 0.6 mL tubes with 100 μL per tube. Two tubes were kept at 37°C for 1 h and the others left in ice for 1 h (control reaction). After this second incubation period (1 h), all tubes were taken to a fumes hood and 500 μL of the mix of acid solution (H₂SO₄:H₃PO₄:H₂O) (1:3:7) were added, as well as 25 μL of ISPF 9%, always protected from light. Subsequently, tubes were sealed with parafilm. Samples were heated for 45 min at 100°C in a water bath, with 200 μL of each reaction product then pipetted on an ELISA plate. Samples were read in a spectrophotometer at 540 nm. The final results were expressed in nmol of protein (urea/min/mg).

Nitric oxide estimation was performed by indirect colorimetric Griess method. First, a standard curve was constructed with NaNO₂ using concentrations of 5, 10, 30, and 60 μM (Sigma-Aldrich, USA). Then, 50 μL of Mϕs cell lysate was incubated with 50 μL of Griess reagent for 1 h. After incubation, the plates were analyzed spectrophotometrically at 550 nm. The results were expressed in micromolar and normalized by the total protein concentration (μM/μg).

For statistical analyses to compare two samples we used Student’s t-test, and in analyses that comprised more than two independent samples one-way ANOVA was performed followed by post hoc Tukey test. The results are presented as mean and SD for parametric variables. Differences were considered significant if P < 0.05. Statistical analyses and graphs were performed with GraphPad Prism version 5.0 (GraphPad Software, USA).

Previous studies have clearly demonstrated the effectiveness of MSCs to educate and reprogram M1-Mϕs and M2-Mϕs (11, 25). Considering this perspective, we tested the ability of adipose-derived MSCs to regulate M1-Mϕs (Figure S1 in Supplementary Material). Using the gate exclusion strategy with Mϕs specific marker (F4/80), it was possible to discriminate the Mϕs and MSCs independent sub-populations (Figure S1A in Supplementary Material). Further coculture analysis confirmed that MSCs decreased the expression of costimulatory molecules (CD86/B7-2) and chemokine receptor (CCR7) in M1-Mϕs previously stimulated with LPS/IFN-γ (Figures S1B,C in Supplementary Material). Additionally, the MSCs also increased the mannose receptor (CD206) frequency, a molecule associated with M2 profile, in M1-Mϕs cocultures (Figure S1D). Interestingly, unstimulated Mϕs presented similar behavior to MSCs-modulated M1-Mϕs (Figures S1B,D). These findings indicate that MSCs in direct contact with M1-Mϕs can promote a prominent immunomodulatory effect. According to these results, we isolated MVs from adipose-derived MSCs supernatants to investigate if MVs-MSCs could assist a similar regulatory function as that observed for its parental cells.

Initially, we characterized the MVs-MSCs population and defined its size, morphology, and immunophenotypic characteristics. Transmission electron microscopy analysis revealed that MVs-MSCs exhibit double membrane structures and present spheroid shape with a diameter inferior to 200 nm (Figure 1A). Additionally, to confirm size and concentration of the MVs-MSCs, we performed a NTA using the NanoSight equipment. This assay showed that purified MVs-MSCs samples had a size around 147.1 ± 13.26 nm and an approximate concentration of 1.0 × 10¹¹ particles/mL (Figures 1B,C). Finally, we investigated if markers associated with MSCs also could be found in MVs-MSCs. We used predetermined particle size (beads with 3.8 and 7 μm) to discriminate the MVs-MSCs population location in FACS workflow (Figure 1D). MVs-MSCs constitutively expressed the classical MSCs antigens such as CD44, CD105, CD90, CD73, but not endothelial (CD31) and hematopoietic (CD45) molecules (Figure 1E). Additionally, as an internal control, we found that MVs derived from macrophages (MVs-Mϕs) do not express classical MSCs antigens (i.e., CD73, CD90, and CD105) on its surface, suggesting the high specificity of our MVs-MSCs immunophenotypic profile (Figure S2 in Supplementary Material). Moreover, MVs-MSCs presented annexin V (microvesicle marker) and tetraspanin (CD9, an EXOs marker) surface expression (Figure 1F). In summary, our structural and immunophenotypic analysis suggest that our EVs-MSCs population, defined here as MVs-MSCs, can be comprised as a heterogeneous subpopulation predominantly enriched with MVs (particles with 100–1,000 nm).

To investigate the capacity of MVs-MSCs to interact and regulate the pro-inflammatory profile of M1-Mϕs, we initially isolated bone marrow-derived Mϕs and extensively differentiated and characterized these cells to M1 and M2 phenotypes (Figure S3 in Supplementary Material). We observed by fluorescent confocal microscopy the ability of MVs-MSCs to associate/incorporate into M1-Mϕs. Live cell imaging showed that MVs-MSCs pre-labeled with PKH26 (red fluorescent dye) had the ability to attach and partially incorporate into the M1-Mϕs cytoplasm right after their administration to M1-Mϕs (i.e., in less than 30 min) (Figure 2A). Visible MVs-MSCs aggregates were detected on M1-Mϕs surface or in intracytoplasmic space during the timecourse (Figure 2A) (Video S1 in Supplementary Material). Moreover, we also confirmed that a single dose of MVs-MSCs suspension can be rapidly incorporated into a confluent macrophage culture (Figure 2A) (Video S3 in Supplementary Material). As a control, the PKH26 red dye alone did not incorporate into M1-Mϕs (Figure 2A) (Video S2 in Supplementary Material).

To confirm the transfer of vesicles into the cytoplasm, we evaluated granularity in M1-Mϕs treated with serial doses of MVs-MSCs. Pre-treated M1-Mϕs had their side scatter (SSC, granularity) parameter increased when compared with untreated and activated M1-Mϕs (Figure 2B). This suggests that M1-Mϕs may incorporate MVs-MSCs and subsequently enhance its internal granularity, probably by the transfer or indirect production of bioactive molecules such as mRNA, miRNAs, proteins, and lipids (26).

We then investigated the ability of MVs-MSCs to modulate pro-inflammatory markers in M1-Mϕs. The treatment with MVs-MSCs decreased M1-related molecules such as CCR7, but not CD86, a classical marker of activation (Figure 2C). On the other hand, MVs-MSCs administration increased the expression of M2-associated markers in M1-Mϕs such as mannose receptor (CD206) (Figure 2C). Furthermore, MVs-MSCs potentially reduced the mRNA expression of inflammatory cytokines such IL-1β and IL-6 in M1-Mϕs, while increasing anti-inflammatory molecules such as IL-10 (Figures 3A–C). Interestingly, the expression of CXCL9, a strong chemoattractant protein, remained unaltered in M1-Mϕs treated or not with MVs-MSCs (Figure 3D). Finally, we attempted to show the MVs-MSCs ability to reduce oxidative stress in M1-Mϕs. We identified that MVs-MSCs treatment increased arginase activity (M2 marker) in M1-Mϕs and reduced endogenous NO production (M1 marker) (Figures 3E,F). In most experiments, unstimulated Mϕs presented a pattern similar to the observed in MVs-MSC-treated M1-Mϕs. Thus, our results suggest that MVs-MSC were effective to reduce activation, oxidative stress, chemokine receptor expression, and pro-inflammatory cytokines on M1-Mϕs, thereby inducing a switch from M1 to a M2-like phenotype.

After we had identified that MVs-MSCs exerted a regulatory effect on M1-Mϕs through modulation of mRNAs and cell signaling proteins, we studied the ability of MVs-MSCs to modulate some important miRNAs related to M1/M2 phenotype (Figure 4). Analysis of these miRNAs showed that miR-155 and miR-21 were significantly more expressed in M1-Mϕs than in unstimulated Mϕs, considering that miR-146 expression, a regulatory miRNA, was not different between both Mϕs groups (Figures 4A–C). Surprisingly, we detected that MVs-MSCs-treated M1-Mϕs reduced miR-21 and miR-155 expression when compared with M1-Mϕs alone (Figures 4B,C). Thus, these data show that MVs-MSCs may modulate M1-Mϕs via downregulation of miR-155 and miR-21 but not by upregulation of miR146a.

In the next step, we set to investigate the predicted target genes of MVs-MSCs-modulated miRNAs in M1-Mϕs. In this sense, we evaluated the expression of SOCS1 and SOCS3 molecules, which possess immunomodulation potential and have been reported as potential targets of these altered miRNAs (Figures 4D,E) (27, 28). Indeed, M1-Mϕs exposed to MVs-MSCs increased the expression of SOCS3 transcript, suggesting that MVs-MSCs upregulate SOCS3 expression by downregulating its associated miR-155, which is also related to M1 activation (Figure 4E). In contrast, the expression of SOCS1 was slightly increased in M1-Mϕs when pulsed with MVs-MSCs, but this was not statistically significant (Figure 4A).

In order to elucidate the immunoregulatory effect of MVs-MSCs on activated Mϕs in vivo, we examined the influence of MVs-MSCs treatment in an experimental model of thioglycollate-induced peritonitis. This classic acute model is characterized by intense Mϕs recruitment to the peritoneal cavity and release of several inflammatory mediators such as IFN-γ, IL-6, MCP-1, IL-17A, and IL-1β (29). We verified 96 h after thioglycollate injection that there was a massive recruitment of CD11b⁺ cells (macrophages) in the peritoneal space (Figure S4A in Supplementary Material). Complementary, we observed a considerable increase in the total number of inflammatory cells in the peritoneal cavity of animals exposed to thioglycollate over the timecourse (Figure S4B in Supplementary Material).

To validate the predominant presence of Mϕs at day 4, we performed H.E. and IHC staining for CD11b, which confirmed the consistent presence of macrophages in the peritoneal cavity (Figure S4C in Supplementary Material). We also observed that around 55% of neutrophils (GR1⁺ cells) were recruited within 24 h of thioglycollate infusion, but these numbers abruptly decreased over time (20% at 48 h and 5% at 72 h) (Figure S4A in Supplementary Material). In contrast, we detected that the number of peritoneal Mϕs increased considerably after thioglycollate infusion, whose highest concentration in the peritoneal cavity (>80% of CD11b⁺ cells) was observed at 96 h. Together, these results show that at day 4 after peritonitis induction there was a predominant population of Mϕs in the peritoneal cavity with little interference of other inflammatory cells. For this reason, this time point was considered to be suitable to investigate the physiological effects of MVs-MSCs on activated peritoneal Mϕs.

Subsequently, we assessed the role of MVs-MSCs on peritoneal Mϕs in two experimental groups: the control group, which was submitted to thioglycollate and treated with vehicle (named: THIO), and the treated group, which was submitted to thioglycollate and exposed to serial doses of MVs-MSCs (named: THIO+MVs-MSCs). Initially, we compared the total number of peritoneal cells present in both groups. Mice administered MVs-MSCs surprisingly showed a reduction in the absolute number of total cells present in the peritoneal exudate (Figures 5A,B). We observed that MVs-MSCs inhibited the recruitment of approximately 20 × 10⁶ inflammatory cells in peritoneal space (Figures 5A,B). Consistently, we found that mice treated with MVs-MSCs also had reduction in the relative number of peritoneal Mϕs (F4/80⁺CD11b⁺) when compared to the control animals (Figures 5C,E). Next, we built a global profile of the inflammatory cells in the peritoneum, considering the absolute and relative number of cells, and we found that mice treated with MVs-MSCs had 7-fold less peritoneal Mϕs than untreated mice (Figure 5F). These data show that MVs-MSCs were effective o inhibit macrophage recruitment after thioglycollate stimulus (Figure 5F). Information on the absolute and relative numbers of peritoneal Mϕs, as well as of eosinophils in mice not treated with thioglycollate (NON-THIO) can be consulted in the Figures S5A–C in Supplementary Material.

To demonstrated the ability of MVs-MSCs to modulate the function of infiltrating Mϕs in the peritoneal cavity, we evaluated the expression of M1 and M2 markers on these specific Mϕs sub-populations (F4/80⁺CD11b⁺). We found that MVs-MSCs decreased the expression of CD86 in peritoneal Mϕs (Figure 5G). Similarly, Mϕs exposed to MVs-MSCs also reduced the expression of iNOS (Figure 5H). On the other hand, the CD206 and CCR7 levels had a slight reduction after MVs-MSCs treatment but did not reach statical significance (Figures 5I,J). Importantly, when we analyzed the arginase activity in the peritoneal exudate of mice submitted to MVs-MSCs treatment, we found a significant increase when compared to the control group (Figure 5K). Furthermore, we investigated the release of pro-inflammatory cytokines in the peritoneal lavage. Multiplex analysis revealed that, in fact, mice treated with MVs-MSCs have reduced concentrations of most inflammatory mediators in the peritoneal lavage, such as IFN-γ, IL-1β, and TNF-α, while that of IL-1α did not change (Figure 5L). Interestingly, IL-6 was not detected in the washout of the peritoneal cavity from mice treated with MVs-MSCs (Figure 5L). These results suggest that MVs-MSCs were highly effective in regulating classically functional M1-Mϕs during peritonitis in vivo, promoting an anti-inflammatory microenvironment.

In addition, we also identified the participation of an eosinophil population that has been reported as interfering cells in this model (30). Here, these cells were detected in the peritoneal exudate as a F4/80^lowCD11b^-SSC-A^high gate population in Figures 5D,E, and as F4/80^lowSSC-A^highSIGLEC F⁺ gate population in Figures 6A,B,D. Although we have observed that MVs-MSCs provided slight increase on relative number of eosinophils in the peritoneum, this difference was not statistically significant (Figure 5D) and, more importantly, these eosinophils were positive for Ly6C but did not exhibit chemokine-associated receptors (CX3CR1 and CCR2) as monocytes, suggesting a non-pivotal participation in inflammatory peritonitis (Figure 6C).

Currently, it is known that two different monocyte subtypes have opposite roles in murine inflammation: Ly6C⁺ CCR2⁺ CX3CR1^low and LY6C⁻ CCR2⁻ CX3CR1^hi monocytes (31). Previous findings suggest that thioglycollate-induced peritonitis promote infiltration of inflammatory Ly6C⁺ monocytes, which subsequently differentiate into peritoneal activated Mϕs (32). In this context, we decided to investigate the influence of MVs-MSCs in the expression of CX3CR1 and CCR2 receptors on activated peritoneal Mϕs. Thus, we performed the thioglycollate-induced peritonitis in heterozygous mice with a florescent gene report for chemokine receptors CX3CR1^+/GFP (green) and CCR2^+/RFP (red) in inbred mice strains CCR2^+/RFP CX3CR1^+/GFP. Flow cytometry analysis showed that MVs-MSCs were again effective in reducing the relative frequency of peritoneal Mϕs in these fluorescent report mice (Figures 6A,B,E). Next, we found that MVs-MSCs decreased the absolute number of infiltrated cells in the peritoneum cavity (Figure S5F in Supplementary Material), whereas the overall profile of inflammatory cells was similar to the previous experiments depicted in Figure 6F. In addition, we similarly documented some baseline information on macrophage and eosinophil frequencies in NON-THIO mice CX3CR1⁺/^GFPCCR2⁺/^RFP (Figures S5D–I in Supplementary Material).

We have also observed that peritoneal Mϕs (F4/80⁺ gate), both treated and untreated with MVs-MSCs, had high expression of the monocyte marker Ly6C⁺ (Figure 6F,G). These data indicate that MVs-MSCs did not alter the subset of monocytes that infiltrated the peritoneum (Ly6C⁺ or Ly6C⁻) during inflammatory stimulus. Furthermore, we observed the expression of chemokine receptors CCR2 and CX3CR1 in this Mϕs population gated to F4/80⁺Ly6C⁺ (Figure 6H). We detected that Ly6C⁺ Mϕs were positive for CCR2 independently of treatment (Figure 6H, Histograms). This information is consistent with the innate ability of Ly6C⁺ monocytes to exhibit CCR2 in order to migrate from bone marrow to inflammatory tissues. In this context, the MVs-MSCs treatment did not alter the expression of CCR2 in Ly6C⁺ Mϕs (Figure 6H). In addition, peritoneal Mϕs from NON-THIO mice showed low expression of CCR2 and CX3CR1 (Figures S5G,H in Supplementary Material).

Surprisingly, when we evaluated the expression of fractalkine receptor (CX3CR1) on Ly6C⁺ Mϕs population, we could identify three major additional sub-populations based on the fluorescence intensity expression of this molecule: (1) population G1 (Ly6C⁺CCR2⁺CX3CR1⁻ Mϕs), population G2 (Ly6C⁺CCR2⁺CX3CR1^low Mϕs), and population G3 (Ly6C⁺CCR2⁺CX3CR1^high Mϕs) (Figures 6H–M). Importantly, the frequency of the G3 population was significantly increased in mice treated with MVs-MSCs (Figure 6M), compared to the G1 population, that was slightly elevated, and the G2 population, that did not change. In this context, we may partially conclude that MVs-MSCs regulate peritoneal Mϕs to both M1 and M2 phenotype, as well as upregulate fractalkine receptor (CX3CR1) in specific Mϕs subsets.