Monocytes educated by cancer-associated fibroblasts secrete exosomal miR-181a to activate AKT signaling in breast cancer cells

The peripheral blood samples from 40 invasive breast ductal carcinoma patients who had not received any chemotherapeutic treatment before surgery were collected in the study. A cohort of 35 age-matched healthy control women with no evidence of any personal or family history of BC participated in this study. A record of the clinicopathological parameters of BC patients is summarized in Additional file 1: Table S1. To harvest the plasma samples, around 5 mL of blood samples from each participant were centrifuged at 3000×g for 10 min at 4 °C, and then stored at − 80 °C until use. To isolate the stromal fibroblasts, primary cancer tissues were obtained from three female BC patients with histological grade III invasive ductal carcinoma who had undergone mastectomy. Normal breast tissues were obtained from three healthy women undergoing reduction mammoplasty. This study was approved by the Ethics Committee of Tarbiat Modares University and written informed consent was obtained from the participants.

Fibroblasts were isolated enzymatically from both normal and cancerous breast tissues using collagenase A as previously described [21] and were maintained in Dulbecco’s Modified Eagle’s medium nutrient mixture F12 (DMEM/F12) supplemented with 10% fetal bovine serum (FBS). Immunophenotyping of patient-derived CAFs was performed for positive expression of α-smooth muscle actin (α-SMA), fibroblast-activation protein (FAP), and negative expression of CD31 to exclude endothelial cell contamination using flow cytometry (BD Biosciences). To prepare the conditioned media (CM) of cancerous and normal cultured fibroblasts (CAF- and NF-CM, respectively) for monocyte treatments, stromal fibroblasts derived from tissue specimens were passaged 4–6 times. The expressions of CAF markers (α-SMA and FAP) and CAF-derived cytokines interleukin (IL)-6 and transforming growth factor (TGF)-β as were also measured by western blotting over the course of passaging the cells. When fibroblasts reached a confluency of > 80%, the cells were serum starved. After 48 h, the CM were collected, pooled, and centrifuged at 300×g for 10 min and then further centrifuged at 10,000×g for 30 min to eliminate residual cells and cellular debris, respectively.

Low-density mononuclear cells were first separated from the peripheral blood of healthy volunteers using a Ficoll-Hypaque density gradient. CD14⁺ monocytes were isolated using a magnetic bead-based positive selection system (Miltenyi Biotech, Germany) with a purity of > 90%, as confirmed by flow cytometric analysis. CD14⁺ monocytes were cultured in Roswell Park Memorial Institute (RPMI)-1640 medium supplemented with 10% FBS alone as control monocytes or were educated with CM derived from CAFs or NFs (CM:RPMI, 1:1) for 7 days. Cultured monocytes were stained with antibodies against CD163, CD206, PD1, CD14, as well as HLA-DR and then analyzed by flow cytometry (BD Biosciences).

Peripheral blood mononuclear cells (PBMCs) were isolated from healthy individuals’ peripheral blood by Ficoll-Hypaque density gradient separation. The isolated PBMCs were then cultured in RPMI-1640 medium containing 10% FBS at 37 °C for 2 h. Subsequently, the adherent cells were removed and T-cells were isolated by nylon wool columns. T-cells were stimulated with 5 μg/mL of phytohemagglutinin (PHA) and expanded in RPMI-1640 medium supplemented with 10% FBS for 7–10 days.

The human BC cell lines MDA-MB-231 and MCF-7 were cultivated in DMEM supplemented with 10% exosome-depleted FBS. The human monocyte cell line THP-1 was cultured in RPMI medium containing 10% FBS. All cells were cultured with 100 U/mL penicillin and 100 μg/mL streptomycin at 37 °C in a 5% CO₂ humidified atmosphere. The monocytic THP-1 cells were differentiated into macrophages by 24 h incubation with 150 nM phorbol 12-myristate 13-acetate (PMA, Sigma-Aldrich) in RPMI medium. Macrophage M2 polarization was obtained by incubation of THP-1 M0 macrophages with 20 ng/mL of interleukin (IL)-4 for 48 h.

Exosomes were isolated from the supernatant of monocytes educated with the CM derived from CAFs or NFs by differential centrifugation as we previously described [17]. Briefly, monocytes were maintained in bovine serum albumin (BSA) or serum-free medium 48 h before supernatant collection. The cell culture supernatants were collected, and centrifuged at 300×g for 10 min to eliminate residual cells and at 10,000×g for 30 min to further remove cells and debris. The supernatant was filtered through a 0.22-μm filter to remove any vesicles larger than 200 nm. The filtered supernatant was subjected to ultracentrifugation at 100,000×g for 70 min at 4 °C. To further eliminate contaminating protein, the exosome pellet was re-suspended in PBS and centrifuged again at 100,000×g for 70 min at 4 °C. Finally, the exosome-enriched pellets were re-suspended in PBS and stored at − 80 °C until use.

The quantity of exosomes was expressed as exosome-associated proteins using the BCA method. Exosome-specific surface markers CD9 and CD81 were detected by western blotting as we previously described [17]. The morphology of exosomes was observed using transmission electron microscopy (TEM, LEO 906 Zeiss 100 kV, Germany). The size distribution of the purified exosomes was also determined by dynamic light scattering (DLS) using a Zetasizer Nano ZS (Malvern Instrument, UK).

Fluorescent labeling of purified exosomes was performed using a PKH26 Red Fluorescent Cell Linker Kit (Sigma-Aldrich) according to the manufacturer’s instructions with some modifications. The labeled exosomes were added to a subconfluent layer of BC cells and incubated at 37 °C for 6 h. Then, the cells were washed twice with PBS and fixed with 4% paraformaldehyde. For nuclear staining, DAPI (4′, 6-diamidino-2-phenylindole, Sigma-Aldrich) was used. The uptake of exosomes was imaged using a spectral confocal microscope (Nikon Eclipse TiE).

To assess the effects of CAF- or NF-educated monocytes on T lymphocyte proliferation, T-cells were labeled with carboxyfluorescein succinimidyl ester (CFSE) and were co-cultured with either control monocytes or with monocytes educated with CAFs or NFs. In parallel, T-cells were incubated with exosomes derived from NFs or CAFs. After 72 h, the proliferation of CFSE-labeled T cells was evaluated by flow cytometry and compared to the control T-cells.

BC cells were transiently transfected with 25 nM miR-181a mimic or negative controls (NC); miR-181a inhibitor or scramble (Sc) using Lipofectamine^® 2000 (Invitrogen, USA). To perform a luciferase reporter assay, BC were transfected with 2 µg of the psi-CHECK2 luciferase reporter plasmid (Promega). Then, luciferase-expressing BC cells were co-cultured with monocytes and were maintained in the control medium, NF-CM, or CAF-CM. After 24 h, cell lysate was collected and added into a 96-well plate. The luciferase activity was measured using the Luciferase Reporter Assay System (Promega Corp., Madison, WI, USA).

BC cells were transfected with miR-181a mimic, miR-181a inhibitor, or the corresponding negative controls as described above. Additionally, another group of BC cells was treated with 100 µg/mL exosomes derived from CAFs or NFs. After 36 h, BC cells were harvested, treated with Triton X100 and RNase A, and then stained with propidium iodide (PI, Sigma). Afterward, cell cycle distribution was analyzed via a flow cytometer (BD Biosciences).

A confluent monolayer of serum-starved BC cells plated into a 12-well plate was subjected to a single-scratch wound by using a sterile pipette tip. The cells were incubated with 100 µg/mL exosomes derived from control monocytes, NEMo, or TEMo and compared to PBS-treated control cells. The cell migration distance was measured and imaged. A cell transwell assay was also performed using 24-well transwells (Corning, USA). The corresponding treated or transfected cells were seeded into the upper chambers in 100 µL of FBS-free medium. As a chemoattractant, the medium with 10% FBS was added to the bottom part of the chambers. Cells were fixed and stained using 1% crystal violet dissolved in methanol 24 h after incubation. The cells that migrated through the membrane and stuck to the lower surface of the membrane were imaged and counted.

The intracellular reactive oxygen species (ROS) production levels were measured by adding the 2′, 7′-dichlorofluorescein diacetate (DCFDA) (ab113851, Abcam) to the cell suspension according to the manufacturer’s protocol. The fluorescent intensity was measured by flow cytometry (BD FACS Canto II, BD Bioscience) and analyzed by FlowJo Software 7.6.2.

Quantitative measurements of secreted IL-10 and IL-12 cytokines were performed on the culture supernatants of THP-1 M0 macrophages, M2 macrophages (serve as positive control), and NF- or CAF-educated M2 macrophages for 48 h, using enzyme-linked immunosorbent assay (ELISA). All cell culture supernatants were used undiluted.

In order to predict the potential targets of miR-181a, we used TargetScan 8.0 (http://​www.​targetscan.​org), miRTargetLink Human (https://​ccb-web.​cs.​uni-saarland.​de/​mirtargetlink) and RNAhybrid (https://​bibiserv.​cebitec.​uni-bielefeld.​de/​rnahybrid) online tools. The DNA Intelligent Analysis (DIANA)-miRPath v3.0 (http://​diana.​imis.​athena-innovation.​gr/​DianaTools) algorithm was used to show the miRNA regulatory roles and to find the significant Kyoto Encyclopedia of Genes and Genomes (KEGG) molecular pathways.

Total RNA was isolated using TRIzol reagent (Invitrogen, USA) according to the manufacturer’s recommendation and treated with RNase-free DNase (Fermentase, Lithuania). RNA was then reverse-transcribed into complementary DNA (cDNA) using the PrimeScript 1st strand cDNA synthesis kit (TAKARA, Japan). To quantify miRNA, a poly(A) tail was initially added to the extracted total RNA by using polyA polymerase enzyme (NEB), and cDNA was then synthesized by using an anchored oligo (dT) primer as described previously [22]. Reverse transcription quantitative PCR (RT-qPCR) was conducted on an ABI Step One Detection System (Applied Biosystems, USA). The relative expression was normalized to U48 small nuclear RNA (snRNA) and GAPDH using the 2^−ΔCt and 2^−ΔΔCt methods [23].

Total protein was extracted from the cells or exosomes using radioimmunoprecipitation (RIPA) lysis buffer. Equal amounts of proteins were separated by 12% SDS-polyacrylamide gel electrophoresis (SDS-PAGE) and transferred to polyvinylidene difluoride (PVDF) membranes. For membrane blocking, 5% skim milk was used for 1 h at room temperature. The primary antibody incubation was performed for 12 h at 4 °C and then followed by horseradish peroxidase (HRP)-conjugated secondary antibody incubation for 1 h at room temperature. The membranes were subjected to chemiluminescence using an ECL Kit (Amersham, UK). β-actin was used as a loading control.

All animal experimental procedures were approved by the Committee for Animal Research of the University. A total of 5 × 10⁶ MDA-MB-231 BC cells alone or mixed with 200 µg/mL exosomes (derived from TEMo or NEMo) were suspended in 100 µL serum-free DMEM and matrigel (1:1 ratio) and were subcutaneously injected into the oxter of 6-week old female BALB/C athymic nude mice. Tumor volume was monitored at every 7-day interval. The tumor volume was calculated using the formula: V (mm³) = (L × W²)/2. After 4 weeks, the mice were sacrificed to evaluate tumor growth.

Formalin-fixed tumor tissues were embedded in paraffin and cut into thin sections. The tissue sections were stained with hematoxylin and eosin (H&E) using standard procedures. For immunohistochemical (IHC) staining, tissue sections were dewaxed with xylene and then rehydrated in graded series of ethanol. Antigen retrieval was subsequently performed by microwave heating in sodium citrate buffer at pH 6.0. To block endogenous peroxidase activity, the sections were immersed in 3% H₂O₂ for 30 min. The sections were incubated with primary monoclonal antibodies against α-SMA (ab7817, Abcam) and Ki-67 (sc-23900, Santa Cruz Biotechnology) at 4 °C overnight. After washing, the sections were incubated with HRP-conjugated secondary antibody for 1 h, and reactive products were visualized by staining with 3, 3′-diaminobenzidine (DAB). The images were captured using an inverted fluorescence microscope (Olympus CKX41, Japan) with appropriate magnification.

Data are expressed as the mean ± standard deviation (SD) of at least three experiments. Statistical significance was calculated by Student’s t-test when comparing two groups or by one-way or two-way analysis of variance (ANOVA) when comparing three or more groups. A p-value of < 0.05 was considered statistically significant.

To investigate the relationship between CAFs and the malignance phenotype, we first studied the expression of α-SMA (a myoepithelial cell marker) and Ki67 (a marker of proliferating cells) in tumor and non-tumor tissues obtained from BC patients. IHC assay performed on tissue samples confirmed that α-SMA is localized, particularly in adjacent stromal cells, which are mainly comprised of stromal myofibroblasts. Results also indicated that the expression of α-SMA has a significant association with histopathological grade (Fig. 1A). As CAFs appear to be major tumor microenvironment components facilitating tumor progression, we aimed to clarify the function of CAFs isolated from patients who have been diagnosed with advanced grade tumors. To this end, CAFs were isolated from three primary BC tissues (Grade III) obtained from patients undergoing mastectomy (Fig. 1B). As healthy counterparts, we used normal fibroblasts (NFs) isolated from normal breast tissues obtained from patients undergoing reduction mammoplasty. Primary cultures of both of those isolated fibroblasts were established. The evaluation of cell surface markers by flow cytometry indicated the positive expression of α-SMA and FAP but the lack of expression of vessel marker CD31 in isolated CAFs (Fig. 1C). Additionally, western blot analysis revealed the expression of CAF-specific markers (α-SMA and FAP) and CAF-derived cytokines (IL-6, and TGF-β) over the course of passaging, confirming CAF activation status throughout the experiments (Additional file 1: Fig. S1).

Stromal cells have been demonstrated to affect monocyte recruitment and polarization in addition to altering their functions in the TME [24]. To evaluate whether CAFs recruit monocytes, we isolated monocytes from the peripheral blood of healthy individuals and performed transwell migration assays. The bottom chambers contained conditioned media (CM) collected from CAFs and NFs, thus serving as chemoattractants. We observed that monocytes migrate toward CAF-CM in the bottom chambers, suggesting that CAFs may effectively recruit monocytes (Fig. 1D, E). Additionally, to explore whether CAFs were able to induce a pro-tumoral phenotype in monocytes, CD14⁺ cells isolated from healthy individuals were subjected to differentiation following treatment with CAF-CM. Hereafter, monocytes educated by CM derived from tumoral CAFs are denoted as tumor-educated monocytes or TEMo, and monocytes co-cultured with CM derived from NFs are referred to as normal fibroblast-educated monocytes or NEMo. After 7 days of culture, the expression M2 macrophage markers CD163 and CD206 were higher in TEMo than in NEMo. In addition, the expression of programmed cell death protein 1 (PD-1) was higher in TEMo than in NEMo. In fact, NF-educated cells’ expressions of PD-1, CD163, and CD206 were similar to control monocytes. On the other hand, the expression of CD14 was much higher in NEMo than in TEMo. In addition, the major histocompatibility complex (MHC) class II (HLA-DR) expression of TEMo is much lower than that of NF-educated cells (P < 0.05, Fig. 1F, G). All of these results suggest that CAFs may recruit monocytes and affect their polarization states.

To explore the capability of CAF to induce M2 polarization of macrophages, the human monocyte cell line THP-1 was differentiated into macrophages by incubation in the presence of 150 nM PMA. After 24 h, cells became adherent and displayed an irregular cell shape characteristic of cells undergoing differentiation. As shown in Fig. 2A, qRT-PCR revealed that the expression levels of recognized macrophage markers CD36, CD68, and CD71 were found to be higher as the macrophages differentiated (P < 0.001). The expression of CD14 was also decreased (P < 0.001), further confirming the monocyte-to-macrophage differentiation (Fig. 2A). Afterward, THP-1 M0 macrophages were incubated with either CAF- or NF-CM and compared to positive control cells (i.e., THP-1 M0 macrophages stimulated with 20 ng/mL of IL-4 for 48 h). The M2 polarization of macrophages was assessed by studying the transcript and protein levels of several M2 macrophage markers. After 48 h of incubation, the transcript levels of CD163 (P < 0.01) and CD206 (P < 0.001) were found to be significantly increased in CAF-CM-incubated THP-1 macrophages (Fig. 2B). Consistently, protein secretion level of anti-inflammatory cytokine IL-10 was increased in culture supernatants of CAF-educated macrophages (P < 0.001), whereas protein secretion level of pro-inflammatory cytokine IL-12 was found to be decreased in macrophages incubated with CAF-CM (P < 0.001) (Fig. 2C). These findings shed light on the potential of CAFs in promoting THP-1 polarization into anti-inflammatory M2 macrophages.

Additionally, as it was demonstrated that oxidative stress plays a role in modulating macrophage phenotype, we aimed to determine ROS status during macrophage polarization. Data showed that when monocytes were incubated with PMA, ROS production in differentiated THP-1 macrophages was markedly elevated (P < 0.001; Fig. 2D, E). Importantly, following CAF-CM incubation, ROS production was abolished in M2-like macrophages (P < 0.01; Fig. 2D, E). Even though ROS levels dropped during M1/M2 macrophage polarization, CAF-induced M2-like macrophages still seem to produce more ROS than THP-1 control monocytes (P < 0.05; Fig. 2D, E). Overall, these findings suggest that CAFs contribute to M2 polarization in part by increasing ROS production in monocytes, as M2 macrophages require a level of ROS for proper polarization.

To uncover the paracrine effect of exosomes derived from CAF-educated monocytes on BC cells, we first isolated and purified exosomes from the conditioned media of monocytes educated with CAF or NF. The morphologically round and spherical shapes of purified exosomes with an approximate diameter ranging from ~ 50–150 nm were observed by TEM (Fig. 3A). Also, the particle size distribution of exosomes by DLS revealed a single bell-shaped size distribution with a peak at ~ 90 nm (Fig. 3B). Additionally, western blotting demonstrated that the exosome-specific tetraspanin markers CD9 and CD81 were enriched in the exosome preparation but not in the monocyte lysate. Conversely, the negative maker of Calnexin (the endoplasmic reticulum protein marker) was almost undetectable in the isolated exosomes, but highly expressed in cells (Fig. 3C). To examine whether the monocyte-derived exosomes could be taken up by BC cells, PKH26-labeled exosomes were incubated with sub-confluent MDA-MB-231 BC cells for 18 h. Confocal microscopy imaging showed that monocyte exosomes can be incorporated and internalized into the cytoplasm of MDA-MB-231 BC cells (Fig. 3D).

CAFs are found to be a major cause of immunosuppression, partly through disrupting T-cell function in the tumor microenvironment. First, in order to explore the functional role of CAF-educated monocytes (TEMo) in promoting immunosuppression, human autologous peripheral T lymphocytes were labeled with carboxyfluorescein succinimidyl ester (CFSE) and co-cultured with control monocytes, NEMo, or TEMo. After 72 h, flow cytometry data showed that T-cell proliferation was inhibited by TEMo more markedly than control (P < 0.01) and NEMo (P < 0.001; Fig. 4). Importantly, we found that 100 µg/mL exosomes derived from TEMO (TEMo-Exo) had a similar inhibitory effect on the proliferation of T-cells compared to that of cells co-cultured with control monocytes or treated with exosomes derived from NEMo (NEMo-Exo) (P < 0.01; Fig. 4). These data suggest that CAFs could exert their immunosuppressive effects on T-cells, at least in part and indirectly, through educating monocytes.

M2 macrophages are typically pro-tumoral and lack the tumoricidal properties of M1 macrophages [25]. Because CAF-CM was able to induce the up-regulation of M2-specific markers, suggesting a conversion of naive monocytes into M2 macrophages (Fig. 1F, G), we first sought to investigate whether CAF-induced polarization of M2 macrophages is concomitant with the loss of tumoricidal properties of monocytes. In order to assess this phenotype, we co-cultured luciferase-expressing MDA-MB-231 BC cells (MDA-MB-231-luc) with monocytes maintained in the control medium, NF-CM, or CAF-CM. As shown in Fig. 5A, in co-cultured conditions with the control medium, the luciferase activity was remarkably diminished, suggesting the effective tumoricidal properties of monocytes. On the contrary, when monocytes were incubated with CAF-CM, these cells lost their ability to kill BC cells and instead enhanced the proliferation of BC cells. These data propose that CAFs may be able to influence the phenotype of monocytes by polarizing them to an M2 pro-tumoral phenotype, promoting BC cell proliferation.

To investigate the effects of TEMo-Exo on BC cell proliferation and migration in vitro, MDA-MB-231 BC cells were incubated with exosomes derived from monocytes educated by CAF or NF. As shown in Fig. 5B, treatment of different concentrations of TEMo-Exo markedly resulted in a significant increase in the proliferation rate of BC cells in a dose- and time-dependent manner. On the contrary, the proliferation rate of BC cells was not significantly affected following incubation with 100 µg/mL exosomes derived from monocytes educated by NF, compared to the cells treated with vehicle control PBS. As expected, BC cells incubated with 100 µg/mL control monocyte exosomes had the lowest rate of proliferation. Using a wound healing assay, we also found that the migration of BC cells into the scratched areas of monolayers was reduced in the presence of exosomes derived from control monocytes compared to PBS-treated BC cells. Importantly, we observed that when incubated with 100 µg/mL TEMo-Exo, BC cells migrated significantly faster than BC cells incubated with NEMo-Exo or vehicle control PBS (Fig. 5C). Consistent with this observation, we found through a transwell migration assay that incubation of BC cells with 100 μg/mL TEMo-Exo led to significantly higher rates of migration through transwell membranes than those incubated with exosomes derived from control monocytes or NF-educated monocytes (Fig. 5D, E). Additionally, we showed that exposure to TEMo-Exo induced the protein expression of epithelial-to-mesenchymal transition (EMT) markers in MDA-MB-231 BC cells. As shown in Fig. 5F, the expressions of EMT markers including N-cadherin, Vimentin, and Snail were found to be up-regulated and the expression of epithelial marker E-cadherin was found to be down-regulated in BC cells by the effects of TEMo-Exo. On the contrary, neither EMT markers nor E-cadherin expression were significantly changed by the effects of NEMo-Exo, compared to that of control monocyte exosomes or PBS vehicle control. Overall, it seems that CAFs educate monocytes to secrete exosomes, which up-regulate the EMT markers and impart more migratory behavior in BC cells.

To further assess the role of TEMo-Exo in tumor growth in vivo, we established tumor models in 6-week-old BALB/c nude mice by subcutaneously injecting MDA-MB-231 cells alone or MDA-MB-231 cells mixed with 200 µg/mL exosomes derived from monocytes educated by CAF. Tumor sizes were measured three times a week, and the observations lasted over 28 days after tumor challenges. As shown in Fig. 6A–D, tumor volume and weight in mice implanted with MDA-MB-231 BC cells mixed with 200 µg/mL TEMo-Exo were significantly higher than in those implanted with BC cells alone. To test whether exosomes derived from NEMo also exert tumor-promoting effects, tumor growth in mice implanted with BC cells mixed with 200 µg/mL NEMo-Exo was measured, and the results showed that NEMo-Exo had no effect on promoting tumor growth compared to those of mice implanted with BC cells only. Furthermore, H&E staining of tumor sections revealed that cells from mice implanted with TEMo-Exo arranged more densely with a more irregular cell shape and had increased cell size when compared to the NEMo-Exo co-implantation group or negative control tumor group (Fig. 6E). As Ki-67 indicates the proliferative ability of tumors, we examined Ki-67 expression in xenograft tumor sections. Consistently, IHC results revealed that tumor tissue from mice implanted with TEMo-Exo exhibited stronger positive staining than that in the cell-only group or cells co-implanted with NEMo-Exo (Fig. 6E). Collectively, these results indicate that TEMo-Exo enhance the tumorigenicity of MDA-MB-231 BC cells in vivo, while exosomes derived from control-educated monocytes have no significant effect on the growth of xenograft tumors.

Mounting evidence has confirmed that miRNAs play a substantial role in the initiation and progression of cancer. In this regard, the dynamic expression pattern of miRNAs may be associated with the progression of tumors [26]. To identify circulating miRNAs that may be involved in BC progression, we performed a survey of public BC data sets for miRNA expression and found that miR-181a is among miRNAs to be frequently up-regulated in BC. To explore the clinical significance of circulating miR-181a in BC, we measured the expression level of this miRNA in BC plasma samples. RT-qPCR results revealed that miR-181a expression level was significantly up-regulated in BC patients compared with healthy individuals and, importantly, miR-181a expression level was positively correlated with tumor aggressiveness (Fig. 7A). These findings confirm that plasma-derived miR-181a has the potential to be a diagnostic biomarker for BC patients.

Being enriched in exosomes, miRNAs secreted from activated monocytes in the peritumoral stroma of tumor cells may contribute to inducing the malignant behavior of tumor cells [20]. To describe the molecular mechanisms by which exosomes derived from monocytes educated with CAF promote the EMT response, we conjectured that miRNAs secreted by TEMo-Exo might account for the oncogenic effects of tumoral monocytes on BC cells. We first measured the expression of miR-181 family members in two groups of monocytes educated by CAF or NF. Fold change analysis in miRNA expression levels revealed a higher level of miR-181a in TEMo, while the expression of this miRNA did not show a significant difference compared to control-educated monocytes (Fig. 7B). We also detected a high level of miR-181a in exosomes derived from TEMo as compared to those derived from NEMo (Fig. 7C).

Since exosomes facilitate the communication between adjacent cells by transferring miRNAs [20], we sought to examine the exosomal transfer of miR-181a from educated monocytes into BC cells. To this end, MDA-MB-231 cells were incubated with 100 μg/mL exosomes derived from monocytes educated with either CAF or NF at different time points and compared to control cells. Additionally, to confirm that miR-181a is transferred from monocytes into BC cells and not transcriptionally induced, we incubated BC cells with either monocyte exosomes or PBS in the presence of α-amanitin, which is an inhibitor of transcriptional activation. As shown in Fig. 7D, a gradual increase in the level of miR-181a was observed in MDA-MB-231 cells stimulated with 100 μg/mL TEMo-Exo, while the level of this miRNA did not show a significant difference in control BC cell groups (i.e., non-treated cells or BC cells treated with NEMo-Exo). These data confirm that miR-181a is enriched in exosomes derived from monocytes educated by CAFs and is shuttled by exosomes into BC cells.

To determine whether exosomes derived from TEMo promote the malignant phenotype of BC cells by transferring miR-181a, functional rescue experiments were performed. The miR-181a inhibitor was first transfected into exosome-treated BC cells. Flow cytometric analysis of the cell cycle distribution revealed that the proportion of BC cells in the S and G2/M phases in TEMo-Exo-treated BC cells was significantly higher than that in NEMo-Exo-treated control cells. Upon miR-181a silencing, the cells accumulated in the sub G1 phase, with a concomitant decrease in the proportion of cells in the S and G2/M phases. Consistently, transfection of miR-181a mimic recapitulated the promoting effects of TEMo-Exo on BC cell progression. Importantly, reintroducing the miR-181a inhibitor abolished the TEMo-Exo-caused increases in the proportions of cells in the S and G2/M phases (Fig. 7E, F). Additionally, the stimulating effect of TEMo-Exo on BC cell migration was recapitulated by miR-181a overexpression. However, the promoting effects of TEMo-Exo on the migration of BC cells were effectively weakened by exogenous inhibition of miR-181a expression (Fig. 7G, H). As miR-181a overexpression recapitulated the phenotypes induced by TEMo-Exo in BC cells, it is suggested that miR-181a shuttled by TEMo-Exo plays a vital role in promoting BC cell progression.

We next analyzed the intracellular signaling that might be regulated by miR-181a secreted from TEMo in BC cells. As shown in Fig. 8A, western blot analysis demonstrated a significant increase in phosphorylated AKT levels in MDA-MB-231 BC cells that were incubated with 100 μg/mL TEMo-Exo compared to control cells. To investigate the functional role of exosomal miR-181a in AKT activation, we utilized a miR-181a inhibitor and found that the promoting effect of TEMo-Exo on AKT activation was partially abrogated when MDA-MB-231 BC cells were transfected with miR-181a inhibitor. To confirm that the exosomal transfer of miR-181a functionally correlates with AKT signaling, the phosphorylated levels of mTOR were measured in BC cells. Results showed that inhibition of miR-181a reduced phosphorylated levels of mTOR in BC cells. As expected, the promoting effects of exosomes derived from TEMo on mTOR activation were partially rescued in the presence of the miR-181a inhibitor (Fig. 8A). To further investigate whether Akt signaling is critically involved in miR-181a function in BC, we inhibited the Akt activity with MK-2206, which is clinically used for the treatment of human BC [27]. Results showed that although transfection of miR-181a mimic led to the phosphorylation of Akt–mTOR, MK-2206 treatment blocked miR-181a overexpression-induced activation of Akt–mTOR signaling in BC cells (Fig. 8B). Importantly, MK-2206 blocked the effects of miR-181a on the proliferation and migration of BC cells because miR-181a overexpression did not significantly increase the proliferation and migration rates in the group of cells treated with MK-2206 (Fig. 8C–E).

Next, we explored the potential mechanism underlying miR-181a-mediated activation of Akt–mTOR signaling. Among the predicted targets of miR-181a, PTEN is a key tumor suppressor gene, dysregulation of which is frequent in BC and is associated with poor prognosis [28]. The PTEN encoding mRNA contains a putative miR-181a binding site within its 3′-UTR (Additional file 1: Fig. S2). Since PTEN has been previously identified as a direct target of miR-181a and is a key upstream inhibitor of Akt–mTOR signaling [29], we conjectured that the enhancing effects of TEMo-secreted miR-181a on AKT activation may be attributed to modulating PTEN. To this end, we transfected MDA-MB-231 BC cells with miR-181a mimic and indicated that miR-181a overexpression led to a significant reduction of PTEN expression at both mRNA and protein levels (Fig. 8F), showing that miR-181a may regulate PTEN in BC cells. To highlight the functional paracrine effects of monocytes, we first measured the expression level of PTEN in MDA-MB-231 cells that were incubated with 100 μg/mL exosomes derived from CAF- or NF-educated monocytes. RT-qPCR results showed that incubation of MDA-MB-231 cells with TEMo-Exo led to a significantly lower level of PTEN expression than those incubated with exosomes derived from control monocytes or NEMo (Fig. 8G). Next, to reveal the functional effect of TEMo-secreted miR-181a on PTEN expression, MDA-MB-231 cells were transfected with miR-181a inhibitor (100 nM) and stimulated with 100 μg/mL CAF-educated monocyte exosomes for 24 h. Importantly, following transfection with the miR-181a inhibitor, the inhibitory effect of CAF-educated monocyte exosomes on the PTEN mRNA expression level was partially rescued in MDA-MB-231 cells (Fig. 8G), suggesting that down-regulation of PTEN is specific and largely dependent on the transfer of this exosome-shuttled miRNA. Next to MDA-MB-231 cells, we also examined the effect of exosomal transfer of miR-181a on another BC-derived cell line, MCF-7. The qRT-PCR results revealed that incubating α-amanitin-treated MCF-7 with 100 μg/mL TEMo-Exo resulted in a gradual increase in the level of miR-181a at different time points, whereas there was no significant difference in the level of this miRNA in control cell groups. Moreover, results showed that the exosomal transfer of miR-181a from TEMo reduced PTEN expression in MCF-7 cells (Additional file 1: Fig. S3). Altogether, these findings support the notion that exosomes derived from TEMo play a role as an activator whereby they activate AKT signaling. We also came up with a plausible model in which TEMo-secreted miR-181a activates AKT signaling through regulating PTEN in BC cells.