THP-1 cell line (TIB-202) was purchased from ATCC and cultured in Roswell Park Memorial Institute (RPMI) 1640, supplemented with 2-mercaptoethanol (0.05 mM), 10% fetal bovine serum (Gibco), and 1% penicillin/streptomycin antibiotic (Gibco), and incubated in 37°C at 5% CO₂ in humidified incubator. To differentiate cells, we stimulate them with 100 nM phorbol-12-myristate 13-acetate (PMA) (Millipore/Sigma) and ionomycin (1 µg/ml; Sigma) for 3 days. The PMA-supplemented media was removed by washing two times with phosphate-buffered saline (PBS) and incubated with fresh PMA-free media, then treated with Ang II (1 µM) for another 24 h. We start our experiments with 100 nM and 1 µM, but consistent results for all markers were not found. We chose 1 µM to boost to maximum the effect of Ang II that may be hidden for some markers. To address for relevance to human cells, additional studies were performed using human-derived macrophages. The monocytes isolated by plastic adherence from human PBMCs after incubation at 37° for 3 h in RPMI 1640 were stimulated for 5 days with rhM-CSF (50 ng/ml) (Thermo Fisher Scientific) to complete the macrophage differentiation. For M1 polarization, the cells were further incubated for 24 h with 100 ng/ml of LPS and 20 ng/ml of human IFN-γ.

To evaluate the cytotoxicity of Ang II, the cells were resuspended in 400 µl of PBS, stained with propidium iodide (PI) (10 µg/ml) for 15 min at room temperature, and immediately analyzed by flow cytometer (BD FACSCanto II). An immunofluorescence imaging using the EVOS® FL Auto Imaging System (Thermo Fisher Scientific) was performed to confirm the flow cytometry experiment. The PI/Hoechst 33342 (10 µg/ml) combination staining was used to assess cell necrosis and apoptosis. After 15 min incubation with Hoechst at 37°C, the cells were centrifuged, and the pellets were resuspended in 300 µl PI (10 µg/ml) staining solution at room temperature for another 15 min. Immediately, the cells were analyzed by flow cytometry using (470/50 nm) emission light for Hoechst and (570/30 nm) for PI detection.

Some of the cells were treated with 1 µM Ang II for 24 h, and other cells without Ang II treatment. After washing with PBS, the cells were resuspended in FACS buffer (100 µl), then stained with corresponding conjugated antibodies at 1 µg/ml for the following markers: PE-conjugated mAb specific for CD15, CD33, and CD44; FITC-conjugated mAb specific for CD49F, CD64, and CD116; and Percp-conjugated mAb specific for CD38. The PE-conjugated mAb for TNF-α and CD11C were used as M1 markers in a separate experiment, while to identify M2 cells, CD206 was used as the specific antibody. After incubation for 30 min–1 h in the dark at 4°C, the cells were washed to remove unspecific staining and transferred to FACS tubes, mixed gently in 400 µl of FACS buffer, and then analyzed by flow cytometry.

To measure ROS production, the cells were incubated with CellROX™ Deep Red reagent (5 µm) for 25 min at 37°C. ROS fluorescence was then detected at 644⁄665 nm of excitation/emission light using a flow cytometer (BD FACSCanto II), and the values were reported as median fluorescence intensity (MFI) and percent of positive cells.

Fluo4/AM fluorescence (Biotium, Fremont) was used to measure intracellular calcium. In the presence of esterases, Fluo4/AM ester is intracellularly hydrolyzed into Fluo4. When Fluo4 bound to Ca²⁺, the fluorescence increases and becomes detectable. Control and treated cells were resuspended in a 5 mM CaCl₂ buffer, before incubation with 5 µM of Fluo4/AM for 30 min at 37°C. The cells were washed with PBS, resuspended in 400 µl of CaCl₂ buffer, and then analyzed by a BD FACSCanto II flow cytometer. The fluorescence was detected at 488/530 nm excitation/emission wavelengths.

Control or Ang II-treated cells (1 × 10⁶/ml) were washed with PBS, fixed in ethanol (70%) for 24 h at −20°C. After that, the cells were centrifuged for 5 mins at 400 g, and then washed two times with PBS to remove ethanol and resuspended in 400 µl PBS supplemented with 50 µg/ml of PI. To avoid false RNA staining, the cells were further incubated for 15 min at room temperature in the presence of RNase A (100 µg/ml). Flow cytometry at low speed was performed to analyze the distribution of cells in G1, G2, and S phase of the cell cycle.

To determine the localization of intracellular calcium, THP1-derived macrophages with and without Ang II treatment were stained by Fluo4/AM. The immunofluorescence imaging was performed using EVOS® system as previously described with modifications (8).

The experiment was performed as previously described (11). Briefly, monocytes with and without Ang II treatment were stained with 1,1′-dihexadecyl-3,3,3′,3′-tetramethylindocarbocyanine perchlorate (Dil) (2 µM)). After 2 min, the excess of Dil was washed four times. The images were analyzed using the confocal laser scanning microscope (Zeiss) with excitation at 543 nm for Dil and 488 nm for photobleaching, and the emission was collected at 600–700 nm. The PMT voltage was adjusted on the detectors to detect the high background signal with zero measurement following bleaching.

Human umbilical vein endothelial cell (HUVEC) cells were cultured in Dulbecco's Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum and 1% pen-strep until they created a monolayer on the bottom of the 6-well plate. The THP-1 cells were cultured in RPMI supplemented with 10% fetal bovine serum and 1% pen-strep; the cells were treated with 100 nM Ang II and/or 100 µM angiotensin receptor blocker (Losartan) and incubated in a humidified chamber at 37°C for 48 h. For labeling, the treated THP1 cells were stained with lipophilic fluorescent stains (DIL), and the cells were then washed and resuspended in complete RPMI and seeded on the attached HUVEC cells in a 6-well plate. The 6-well plate containing attached HUVEC cells and stained THP1 cells was incubated for 30 min at 37°C on shaker at 25 rpm; the plate was then washed three times with 1× phosphate-buffered saline. The images of adherent monocytes were acquired using the EVOS microscope. This experiment was repeated at least three times.

To measure the relative protein phosphorylation, the human phospho-kinase array kit (ARY003C; R&D Systems) was used according to the instruction of the manufacturer (12). Briefly, THP1-derived macrophages treated or not with Ang II (1 μM) were harvested and equal cell lysate (30 μg) from each group was used, than signal intensity was analyzed using the Image Lab software.

The Prism® 5.0 (GraphPad Prism Software Inc., La Jolla, CA, USA) was used for the statistical data analysis. The data are presented as means ± standard deviation (SD). The one-way analysis of variance (ANOVA) was used to compare more than two groups, and the Student t-test was used to determine the significance between two groups.

In our study, Ang II increased the expression of CD116 and Fc-gamma receptor I (CD64) (Figures 1A,B), a specific macrophage marker that differentiates it from dendritic cells. Moreover, THP-1-like macrophage cells were highly activated after the treatment with Ang II by an increase in the CD38 marker (Figure 1C). Furthermore, we observed an upregulation of the CD44 marker following Ang II stimulation, which may play a role in proinflammatory phenotype. It has been reported that monocytes differentiated into macrophages that are CD33-deficient increase phagocytosis function and proinflammatory signatures (13). The treatment with Ang II upregulates CD15, CD33, and integrin CD49f (Figures 5A–C), which are known to be involved in monocyte/macrophage differentiation and cell adhesion. The results were confirmed by an adhesion assay showing an increase in monocyte adhesion to endothelial cells after treatment with Ang II and inhibition of this effect by Los treatment (Figures 5E,F). Most of these results were confirmed in human-derived macrophages (Supplementary Figure S2). We therefore hypothesized that Ang II promotes M1-like-THP-1-macrophage phenotype, while suppressing M2 markers. Consistent with this, we observed an upregulation of CD11c, TNF-α, and HLA-DR (HLA class II) (Figures 1D,E,G) as M1 markers and a decrease in the mannose receptor (CD206) (Figure 1H) as an M2 marker. The treatment with angiotensin II type 1 receptor (AT1R) blocker (Losartan) blunted CD11c upregulation (Figure 1F) without any effect on the other markers. The results were confirmed in human-derived macrophages; however, the effect was significantly inhibited by Losartan treatment for both CD11c and TNF-α expression (Supplementary Figure S1).

To evaluate Ang II effect on cell viability and apoptosis, we treated THP-1 macrophages with 1 µM Ang II for 24 h, 48 h, and 72 h. Apoptosis, measured by Annexin V staining, was significantly detected only after 48 h following treatment with Ang II (Figure 2A). After 24 h of exposure to Ang II, the cell cycle analysis showed G1 arrest in a significant part of macrophages compared with controls (Figure 2C). The apoptosis status was restored after blocking AT1R. The same results of apoptosis were confirmed in human-derived macrophages (Supplementary Figure S5).

Reactive oxygen species (ROS) production is a hallmark of macrophage activation (14). During cellular metabolism, ROS modulates inflammatory macrophage phenotype (15). To evaluate the effect of Ang II on macrophage cellular redox balance in our study, we used the CellROX™ Deep Red for ROS generation analysis. We found that Ang II treatment significantly increased ROS production (Figure 3A). The same results were confirmed in human-derived macrophages (Supplementary Figure S3).

Intracellular calcium modulates the transition of macrophages from a resting state into an activated phase (16). Our results showed that treatment of THP-1-derived macrophages with Ang II increased cytosolic calcium production (Figure 4C). The same experiment was performed and confirmed in human-derived macrophages (Supplementary Figure S4). In addition, the fluorescence recovery after photobleaching (FRAP) technique revealed a delay in calcium expression after UV stimulation (Figures 4A,B). After inducing a Ca²⁺ signal in macrophages, it was noted that the recovery of FRAP signals was faster in the cells treated with Ang II (Figure 4A) compared with control cells (Figure 4B). The full recovery time for treated cells was 20 s, whereas control cells typically completed FRAP recovery no more than 26 s. In a separate experiment, treatment of cells with Ang II induces a decrease in intracellular calcium, partially restored after pretreatment of cells with Los (Figure 4E).

Preincubation of monocytes with Ang II induced an increase in binding to endothelial cells (Figures 5E,F). Adhesion of monocyte to the endothelium is a signal of major change in cell phenotype and a first step in initiating inflammation. Treatment with Losartan blunted this effect compared with the Ang II group (Figures 5E,F).

We used the Human Phospho-Kinase array to screen the activated proteins in response to Ang-II-stimulated THP-1-macrophages and detect the pathways dependent on AT1R activation. This approach revealed a significant activation of the key proteins of different signaling pathways, such as ERK1/2, p38a, endothelial nitric-oxide synthase (eNOS), STAT1, STAT2, MSK1/2, RSK1/2, Lyn, HSP27, HSP60, and PRAS40 (Figures 6A,B). The activation was blunted after Losartan treatment. In addition, a considerable increase occurred in other phosphoproteins (such as AKT1/2/3, GSK-3α/β, Chk-2, RSK 1/2/3, and STAT3) after treatment with Ang II without any effect of AT1R blocker treatment (Figure 6C).