Microparticles (MPs) are membrane vesicles released during cell activation and apoptosis. We have previously shown that MPs from apoptotic T cells induce endothelial dysfunction, but the mechanisms implicated are not completely elucidated. In this study, we dissect the pathways involved in endothelial cells with respect to both NO and reactive oxygen species (ROS). Incubation of endothelial cells with MPs decreased NO production that was associated with overexpression and phosphorylation of endothelial NO synthase (eNOS). Also, MPs enhanced expression of caveolin-1 and decreased its phosphorylation. Microparticles enhanced ROS by a mechanism sensitive to xanthine oxidase and P-IκBα inhibitors. PI3K inhibition reduced the effects of MPs on eNOS, but not on caveolin-1, whereas it enhanced the effects of MPs on ROS production. Microparticles stimulated ERK1/2 phosphorylation via a PI3K-depedent mechanism. Inhibition of MEK reversed eNOS phosphorylation but had no effect on ROS production induced by MPs. In vivo injection of MPs in mice impaired endothelial function. In summary, MPs activate pathways related to NO and ROS productions through PI3K, xanthine oxidase, and NF-κB pathways. These data underscore the pleiotropic effects of MPs on NO and ROS, leading to an increase oxidative stress that may account for the deleterious effects of MPs on endothelial function.

Microparticles (MPs)4 are vesicles shed from the blebbing plasma membrane of various cell types, such as platelets, T and B cells, monocytes, and endothelial cells during activation by agonists, shear stress, or apoptosis (1, 2). MPs constitute a heterogeneous population, differing in cellular origin, number, size, antigenic composition, and functional properties (3). MPs bear cell surface proteins and the cytoplasmic component of the original cell and exhibit negatively charged phospholipids such as phosphatidylserine, and tissue factor (3, 4), accounting for their procoagulant character and proinflammatory properties, including alteration of vascular function (3, 4). MPs may transfer bioactive molecules to other cells, different than those from which they have been produced, thereby stimulating cells to produce cytokines, cell adhesion molecules, growth factors and tissue factor, and modulate endothelial functions (3). However, the mechanisms involved in the effects produced by MPs are not fully elucidated.

Elevated levels of circulating MPs have been detected in pathological states (5, 6, 7, 8, 9, 10) associated with vascular dysfunction, including decrease of endothelium-dependent vasodilatation and/or alteration of responsiveness of vascular smooth muscle to vasoconstrictor stimuli. Because MPs usually accumulate in areas of disordered blood flow (1), the enhanced level of MPs may have pathological consequences. Besides, the above effects of MPs are related to their components, which depend on cell origin and on stimuli used to generate them (11, 12). Thus, in previous studies, we have shown that MPs generated from activated/apoptotic lymphocytes induced cell differentiation and NO production by a morphogen-dependent mechanism (12, 13). In contrast, when MPs were produced from apoptotic lymphocytes, they did not bear morphogen, resulting in a lack of cell differentiation (12) and they induced vascular dysfunction (8, 14). To date, little is known about the effects of MPs on the function of cells that form the vessel wall, such as endothelial cells, which are known to play a crucial role in the regulation of vasomotricity and in inflammation. The endothelium can release relaxant factors, including NO, prostacyclin, and endothelium-derived hyperpolarizing factor. A change in the ability of endothelium to release these factors can lead to the development of cardiovascular and/or inflammatory diseases. Furthermore, endothelial function is usually defined as NO production and/or bioavailability. It is well-established that endothelium-derived NO undergoes a very rapid reaction with superoxide anion (O2), thereby reducing NO bioavailability (15). Thus, increased oxidative stress impairs endothelial function, promoting inflammatory diseases including hypercholesterolemia, atherosclerosis, and hypertension. MPs can affect endothelial function by altering the balance between NO and reactive oxygen species (ROS) production and release. In this way, it has been reported that MPs decrease endothelial NO-dependent relaxation (eNOS) expression (8) and enhance ROS production (16, 17).

Recently, we have shown that MPs from apoptotic T cells induce endothelial dysfunction through the alteration of NO and prostacyclin pathways (8), but the molecular mechanisms implicated are not completely elucidated. The present study was designed to investigate the signaling pathways of these MPs in endothelial cells with respect to both NO pathway and ROS production.

The human lymphoid CEM T cell line was used for MP production. Cells were seeded at 106 cells/ml and cultured in serum-free X-VIVO 15 medium (Cambrex). MPs were produced as previously described (12). Briefly, cells were treated with actinomycin D (0.5 μg/ml; Sigma-Aldrich) for 24 h. Supernatant was obtained by centrifugation at 750 × g for 15 min, and then at 1,500 × g for 5 min to remove cells and large debris, respectively. MPs from the supernatant were washed after three centrifugation steps (45 min at 14,000 × g) and recovered in 400 μl of NaCl 0.9%. Last washing medium was used as control. The determination of the amount of MPs was conducted by measuring MP-associated proteins using the DC Protein Assay (Bio-Rad). MPs were used at 10 μg/ml medium. Ten micrograms of MPs correspond to 1,330 ± 312 × 103 MPs. This concentration of lymphocyte MPs (0–2,000 × 103 MPs/ml plasma) is found in plasma from patients undergoing carotid endarterectomy (18). MPs isolated from at least four independent preparations were analyzed for each experimental condition.

Human endothelial cell line (Eahy 926) was maintained in culture in medium culture (DMEM, Ham’s F-12 1:1 (Cambrex), 1% l-glutamine, 1% hypoxanthine/aminopterin/thymidine, 1% nonessential amino acids, 1% sodium pyruvate and 1% streptomycin/penicillin) supplemented with 10% of FBS (Invitrogen Life Technologies). Cells were grown for 24 h in the absence or presence of 10 μg/ml MPs after preincubation with or without inhibitors (PI3K inhibitor, LY294002 (20 μM; Calbiochem); MEK inhibitor U0126 (2 μM; Calbiochem); NOS inhibitor, nitro-l-arginine (l-NA, 100 μM; Sigma-Aldrich); inhibitor of IκB-α phosphorylation, Bay 11-7082 (Bay, 7.5 μM; BioMol Research); xanthine oxidase inhibitor, allopurinol (50 μM; Sigma-Aldrich); NADPH oxidase inhibitor, apocynin (100 μM; Sigma-Aldrich); mitochondrial inhibitor, rotenone (10 μM; Sigma-Aldrich); or with the superoxide dismutase mimetic, manganese(III) tetrakis (1-methyl-4-pyridyl) porphyrin pentachloride (MnTMPyP 100 μM; Calbiochem). MPs and all agents were used at concentrations at which no cytotoxicity was observed, as deduced from trypan blue exclusion.

The detection of NO production was performed using the technique with Fe2+ diethyldithiocarbamate (DETC; Sigma-Aldrich) as spin trap. Briefly, after 24 h of MP treatment, cells were stimulated with either vehicle or bradykinin (20 μM; Sigma-Aldrich) for 45 min at 37°C. Then, the medium was replaced with 250 μl of Krebs solution, and then treated with 250 μl of colloid Fe(DETC)2 and incubated for 45 min at 37°C. NO detection was measured in situ by EPR. Values are expressed in units per microgram per microliter of endothelial cell proteins.

NO measurement was performed on a table-top x-band spectrometer Miniscope (MS200; Magnettech). Recordings were made at 77°K, using a Dewar flask. Instrument settings were 10 mW of microwave power, 1 mT of amplitude modulation, 100 kHz of modulation frequency, 150 s of sweep time and five scans. Signals were quantified by measuring the total amplitude, after the correction of baseline as previously described (14). The quantitative measurement of the NO-Fe(DETC)2 signal amplitude was reported to the relative units for protein concentration (amplitude per microgram per microliter).

The production of NO was also quantified by measuring the released NO metabolites (nitrite and nitrates) with Griess reagent (Promega). The culture medium samples were collected from control and MP-treated cells and prepared cell-free by centrifugation. The medium was processed according to the manufacturer’s protocol.

After incubation of endothelial cells with MPs at 10 μg of proteins/ml for 24 h in absence or presence of inhibitors, they were washed three times with phosphate buffer salt solution, and then incubated with the oxidative fluorescent dye DHE (3 μM; Sigma-Aldrich). Fluorescence (at 620 nm) was measured and the number of DHE-positive cells after treatment with MPs was measured by flow cytometry (Beckman Coulter). This method allows the determination of labeled-positive cells (19). Thus, if the number of positive cells increases with the treatment, that means that fluorescence of these cells is enhanced as well. Fluorescent-positive cells were determined by flow cytometry.

In another set of experiments, aortae from male Swiss mice (8–10 wk old) were treated in vitro with MPs and in situ production of O2 in the vessel wall was evaluated with DHE as previously described (9). After washing, vessel sections were mounted on glass slides. A Solamere confocal instrument with a DLS-300 laser mounted on a Nikon Eclipse TE 2000-S inverted microscope was used for the optical sectioning of the tissue. Digital image recording was performed using QED in vivo software. All animal studies were conducted using approved institutional protocols.

After treatment, cells were homogenized and lysated. Proteins (20 μg) were separated on 10% SDS-PAGE. Blots were probed with anti-eNOS, caveolin-1 (BD Biosciences), phospho-caveolin-1 Tyr14, phospho-eNOS Ser1177, phospho-eNOS Thr495, phospho-ERK 1/2 (Cell Signaling), phospho-IκB-α (U.S. Biological), and Akt (Santa Cruz Biotechnology) Abs. A polyclonal rabbit anti-human β-actin Ab (Sigma-Aldrich) was used at 1/5000 dilution for visualization of protein gel loading. Also, 20 μg of MPs were separated on 10% SDS-PAGE, and blots were probed with anti-NADPH oxidase subunits NOX1 and NOX4 Abs. The membranes were then washed at least three times in Tris-buffer solution containing 0.05% Tween 20 and incubated for 1 h at room temperature with the appropriate HRP-conjugated secondary Ab (Amersham Biosciences). The protein-Ab complexes were detected by ECL plus (Amersham) according to the protocol of the manufacturer.

siRNA duplexes specific for human Akt and control nonsilencing siRNA were obtained from Santa Cruz Biotechnology. Transient transfection of Eahy 926 endothelial cells was done according to the manufacturer’s protocol. Briefly, cells were seeded in 6-well plates, grown for 24 h (60% confluence), and then transiently transfected with 100 nM Akt-specific or control siRNA using the transfection reagent provided. Medium was replaced 24 h later by fresh medium and cells were grown for an additional 24 h. Then, cells were treated with MPs and NO measurements were performed.

Swiss mice (8–10 wk old) were treated in vivo by i.v. injection into the tail vein of MPs (10 μg/ml) as previously described (13). After 24 h, aortic rings were isolated and mounted on a wire myograph. Endothelium-dependent vasodilatation was studied in aortas with functional endothelium precontracted with U46619 (Sigma-Aldrich).

Data are represented as mean ± SEM; n represents the number of experiences. Statistical analysis was performed by a one-way ANOVA, Mann-Whitney U, or ANOVA for repeated measures with a subsequent Bonferroni post-hoc test. A value of p < 0.05 was considered to be statistically significant.

Both control and MP-treated cells exhibited an EPR feature of signals derived from NO-Fe(DETC)2. The NO-Fe(DETC)2 EPR signal was significantly reduced in cells treated by MPs. Although no significant difference was observed between the three concentrations of MPs used (5, 10, and 20 μg/ml), the decrease in NO production was greatest at 10 μg/ml, and this concentration was used for all further studies. In addition, the NO-Fe(DETC)2 EPR signal results for NOS activation as demonstrated by the abrogation of the signal in the presence of the NO inhibitor, l-NA (Fig. 1,A). To further demonstrate that the NO-Fe(DETC)2 EPR signal reflects NO generation from endothelial cells, nitrite and nitrate, both main metabolites of NO, were determined by Griess assay. As shown in Fig. 1,B, MP treatment reduced the nitrite/nitrate concentration. Moreover, NO production was significantly reduced in cells treated with MPs as compared with nontreated cells, and this in the absence or in the presence of bradykinin (Fig. 1,C). The MP-induced NO decrease was almost completely reversed when cells were incubated in the presence of the selective PI3K inhibitor LY294002. Also, incubation with the MEK inhibitor U0126 partially reversed the decrease of NO induced by MPs (Fig. 1,C). It should be noted that both inhibitors alone did not modify NO release from endothelial cells (data not shown). Furthermore, silencing Akt signaling using siRNA to Akt reversed MP-induced NO decrease (Fig. 1 D). Taken together, these data demonstrate that the PI3K/Akt and MAPK pathways play a major role in the decrease of NO induced by MPs.

FIGURE 1.

MP treatment decreases NO production in endothelial cells. A, Quantification of the amplitude of the NO-Fe(DETC)2 complex signal in human endothelial Eahy 926 cells. Cells were incubated for 24 h in the presence of either vehicle (CTL), 5, 10, or 20 μg/ml MPs, or in the presence of 10 μg/ml MPs plus the NO synthase inhibitor (l-NA, 100 μM). B, Quantification of nitrite and nitrate productions in human endothelial Eahy cells. Cells were incubated for 24 h in the presence of either vehicle (CTL) or MPs (10 μg/ml). Supernatants were collected and assayed for nitrite and nitrate production. C, Quantification of the amplitude of the NO-Fe(DETC)2 complex signal in human endothelial Eahy 926 control cells (CTL), 10 μg/ml MP alone, or in the presence of PI3K inhibitor LY294002 (LY, 20 μM) or MEK inhibitor U0126 (10 μM) and MPs (MPs + LY or MPs + U0126, respectively). After 24 h, cells were incubated without or with bradykinin (BK, 20 μM for 45 min) and in situ NO production was determined by EPR. D, Quantification of the amplitude of the NO-Fe(DETC)2 complex signal in either control cells (CTL), 10 μg/ml MP- or MPs + siRNA of the Akt-treated human endothelial Eahy 926. Insert, Western blot for Akt expression in the control (lane 1), siRNA scrambled-(lane 2), or Akt siRNA (lane 3)-treated endothelial cells. Values are expressed in units of amplitude per microgram per microliter of proteins of samples. The results are expressed as mean ± SEM of five independent experiments; ∗, p < 0.05, ∗∗, p < 0.01, ∗∗∗, p < 0.001 vs their respective CTL; †, p < 0.05, ††, p < 0.01 vs MPs alone.

FIGURE 1.

MP treatment decreases NO production in endothelial cells. A, Quantification of the amplitude of the NO-Fe(DETC)2 complex signal in human endothelial Eahy 926 cells. Cells were incubated for 24 h in the presence of either vehicle (CTL), 5, 10, or 20 μg/ml MPs, or in the presence of 10 μg/ml MPs plus the NO synthase inhibitor (l-NA, 100 μM). B, Quantification of nitrite and nitrate productions in human endothelial Eahy cells. Cells were incubated for 24 h in the presence of either vehicle (CTL) or MPs (10 μg/ml). Supernatants were collected and assayed for nitrite and nitrate production. C, Quantification of the amplitude of the NO-Fe(DETC)2 complex signal in human endothelial Eahy 926 control cells (CTL), 10 μg/ml MP alone, or in the presence of PI3K inhibitor LY294002 (LY, 20 μM) or MEK inhibitor U0126 (10 μM) and MPs (MPs + LY or MPs + U0126, respectively). After 24 h, cells were incubated without or with bradykinin (BK, 20 μM for 45 min) and in situ NO production was determined by EPR. D, Quantification of the amplitude of the NO-Fe(DETC)2 complex signal in either control cells (CTL), 10 μg/ml MP- or MPs + siRNA of the Akt-treated human endothelial Eahy 926. Insert, Western blot for Akt expression in the control (lane 1), siRNA scrambled-(lane 2), or Akt siRNA (lane 3)-treated endothelial cells. Values are expressed in units of amplitude per microgram per microliter of proteins of samples. The results are expressed as mean ± SEM of five independent experiments; ∗, p < 0.05, ∗∗, p < 0.01, ∗∗∗, p < 0.001 vs their respective CTL; †, p < 0.05, ††, p < 0.01 vs MPs alone.

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We examined its probable implication in MP effects on endothelial cells. Thus, we studied eNOS and caveolin-1 pathways with respect to their possible regulation by PI3K using the inhibitor LY294002. As shown in Fig. 2,A, MP treatment increased eNOS expression. Furthermore, eNOS phosphorylation either on its activator (Ser1177) or inhibitor (Thr495) site was markedly enhanced (Fig. 2, B and C) by MP treatment. However, the phosphorylation on the inhibitor site was significantly greater (Fig. 2,D). Interestingly, the blockade of PI3K almost suppressed the effect of MPs on both expression and phosphorylation of eNOS (Fig. 2, A and C).

FIGURE 2.

Regulation of NO pathway by MPs: involvement of PI3K. Endothelial cells were incubated for 24 h with either vehicle (CTL), 10 μg/ml MP alone, PI3K inhibitor LY294002 (LY, 20 μM) alone or plus MPs (MPs + LY). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against: eNOS (A), phospho-eNOS Ser1177 (B), phospho-eNOS Thr495 (C), caveolin-1 (E), or phospho-caveolin-1 Tyr14 (F). D, Phosphorylation level of eNOS at Ser1177 and Thr495. Quantitation of immunoblots was done by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05; ∗∗, p < 0.01 vs CTL; †, p < 0.05; ††, p < 0.01 vs MP alone.

FIGURE 2.

Regulation of NO pathway by MPs: involvement of PI3K. Endothelial cells were incubated for 24 h with either vehicle (CTL), 10 μg/ml MP alone, PI3K inhibitor LY294002 (LY, 20 μM) alone or plus MPs (MPs + LY). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against: eNOS (A), phospho-eNOS Ser1177 (B), phospho-eNOS Thr495 (C), caveolin-1 (E), or phospho-caveolin-1 Tyr14 (F). D, Phosphorylation level of eNOS at Ser1177 and Thr495. Quantitation of immunoblots was done by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05; ∗∗, p < 0.01 vs CTL; †, p < 0.05; ††, p < 0.01 vs MP alone.

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Caveolin-1 plays a key role in negative regulation of eNOS activity at the level of caveolae (20). As shown in Fig. 2,E, MP treatment increased caveolin-1 expression. However, caveolin-1 phosphorylation on Tyr14 was markedly reduced by MP treatment (Fig. 2,F). Interestingly, the inhibitor LY294002 had no effect on the capacity of MPs to increase caveolin-1 expression (Fig. 2,E) or to decrease its phosphorylation (Fig. 2 F).

Taken together, these data indicate that MPs regulate eNOS activity, mainly via PI3K, by acting on its expression and phosphorylation. In addition, MPs modulate caveolin-1 expression and activity in a PI3K-independent manner.

The ERK 1/2 MAPK pathway may play a role in the effects of MPs on endothelial cells. Indeed, we investigated the implication of this pathway using the MEK inhibitor U0126. As shown in Fig. 3,A, blockade of MEK did not modify the effect of MPs on the expression of eNOS. However, it almost suppressed MP-induced phosphorylation of eNOS on both Ser1177 and Thr495 (Fig. 3, B and C), to the same extent as the PI3K blockade shown above (Fig. 2, B and C). As obtained with LY294002, the MEK silencing by U0126 had neither effect on the enhancement of caveolin-1 expression nor on the reduction of its phosphorylation upon MP treatment (Fig. 3, D and E). Taken together, these data show that both LY294002 and MEK inhibitor U0126 had a similar suppressive effect on MP-induced eNOS phosphorylation.

FIGURE 3.

Regulation of NO pathway by MPs: involvement of ERK 1/2 MAPK. Endothelial cells were incubated for 24 h with either vehicle (CTL), 10 μg/ml MP alone, MEK inhibitor U0126 (U, 10 μM) alone or plus MPs (MPs + U). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against: eNOS (A), phospho-eNOS Ser1177 (B), phospho-eNOS Thr495 (C), caveolin-1 (D), or phospho-caveolin-1 Tyr14 (E). Immunoblots were quantified by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05; ∗∗, p < 0.01 vs CTL; †, p < 0.05; ††, p < 0.01 vs MPs alone.

FIGURE 3.

Regulation of NO pathway by MPs: involvement of ERK 1/2 MAPK. Endothelial cells were incubated for 24 h with either vehicle (CTL), 10 μg/ml MP alone, MEK inhibitor U0126 (U, 10 μM) alone or plus MPs (MPs + U). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against: eNOS (A), phospho-eNOS Ser1177 (B), phospho-eNOS Thr495 (C), caveolin-1 (D), or phospho-caveolin-1 Tyr14 (E). Immunoblots were quantified by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05; ∗∗, p < 0.01 vs CTL; †, p < 0.05; ††, p < 0.01 vs MPs alone.

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As shown in Fig. 4, the incubation of cells with MPs significantly enhanced ERK 1/2 MAPK phosphorylation. As expected, the U0126 treatment completely suppressed MP-induced phosphorylation of ERK (Fig. 4,A). Interestingly, PI3K inhibitor partially suppressed the MP-induced phosphorylation of ERK 1/2 (Fig. 4 B). These findings suggest a cross-talk between the PI3K and MAPK pathways, PI3K being upstream of ERK 1/2 phosphorylation.

FIGURE 4.

MPs induce ERK 1/2 phosphorylation. Endothelial cells were incubated for 24 h with either vehicle (CTL), 10 μg/ml MPs alone, MEK inhibitor U0126 (U, 10 μM) alone or plus MPs (MPs + U) (A), PI3K inhibitor LY294002 (LY, 20 μM) alone or plus MPs (MPs + LY) (B). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting, using Abs raised against phospho-ERK 1/2. Immunoblots were quantified by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05 vs CTL; †, p < 0.05 vs MPs alone.

FIGURE 4.

MPs induce ERK 1/2 phosphorylation. Endothelial cells were incubated for 24 h with either vehicle (CTL), 10 μg/ml MPs alone, MEK inhibitor U0126 (U, 10 μM) alone or plus MPs (MPs + U) (A), PI3K inhibitor LY294002 (LY, 20 μM) alone or plus MPs (MPs + LY) (B). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting, using Abs raised against phospho-ERK 1/2. Immunoblots were quantified by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05 vs CTL; †, p < 0.05 vs MPs alone.

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NF-κB is a transcription factor that is complexed with IκB-α and -β in the cell cytoplasm, which are selectively phosphorylated, ubiquinated, and degraded, leading to NF-κB translocation. Phosphorylation of IκB-α is known to result in the activation of NF-κB and to allow its translocation to the nucleus. Because NF-κB activation might regulate increase of several protein expressions, we have studied this pathway using an inhibitor of the phosphorylation of IκB-α, Bay. This inhibitor completely reversed the increase in eNOS (Fig. 5,A) and caveolin-1 (Fig. 5,B) expressions induced by MP treatment. In addition, although MPs did not affect the level of phosphorylation of IκB-α, as expected, Bay decreases its phosphorylation (Fig. 5 C).

FIGURE 5.

The NF-κB pathway is involved in the MP effects on endothelial cells. Human Eahy endothelial cells were treated for 24 h with either medium (CTL) or 10 μg/ml MPs in absence or in presence of the inhibitor of IκB phosphorylation, Bay 11-7082 (7.5 μM). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against: eNOS (A), caveolin-1 (Cav; B), or P-IκB-α (C). Immunoblots were quantified by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05 vs CTL; †, p < 0.05 vs MPs alone.

FIGURE 5.

The NF-κB pathway is involved in the MP effects on endothelial cells. Human Eahy endothelial cells were treated for 24 h with either medium (CTL) or 10 μg/ml MPs in absence or in presence of the inhibitor of IκB phosphorylation, Bay 11-7082 (7.5 μM). Twenty micrograms of proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against: eNOS (A), caveolin-1 (Cav; B), or P-IκB-α (C). Immunoblots were quantified by densitometric analysis. β-actin control was included. Data are representative of four separate blots, and the densitometry values are expressed in arbitrary units (AU) as mean ± SEM. ∗, p < 0.05 vs CTL; †, p < 0.05 vs MPs alone.

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ROS play a well-documented role in endothelial dysfunction, thus we investigated the possible involvement of oxidative stress in MP effects in endothelial cells and the vascular wall. First, we evaluated the production of ROS in endothelial cells after MP treatment. The percentage of DHE-positive cells determined by flow cytometry was greater by up to 20% in MP-treated cells vs control cells (Fig. 6,A). In contrast with NO, the PI3K inhibition potentiated the MP-induced increase of ROS production (Fig. 6,A), suggesting a negative control of PI3K pathway on MP-induced ROS production. In addition, MEK inhibition had no effect on the MP-induced increase of ROS production (Fig. 6,A), indicating that MP effects on ROS production are ERK 1/2 independent. To determine the sources of ROS production, cells were incubated in the presence of inhibitors of either NADPH oxidase (apocynin), mitochondrial complex I (rotenone), NO synthase (l-NA), or xanthine oxidase (allopurinol) and O2 production was evaluated. ROS generation was independent of NADPH oxidase, mitochondrial complex I activity, or NO synthase (Fig. 6,A). By contrast to other inhibitors, allopurinol blunted the increase of DHE-positive cells induced by MPs (Fig. 6,A). It should be noted that the increase of ROS production after rotenone treatment may be associated with the cytotoxicity of this drug on cells (21). Interestingly, the inhibitor of phosphorylation of IκB-α, Bay, decreased the production of ROS induced by MPs suggesting that the NF-κB pathway is also involved in ROS generation. As expected, the superoxide dismutase mimetic MnTMPyP abolished MP-induced ROS production (Fig. 6 A).

FIGURE 6.

Mechanisms involved in the increase of ROS induced by MPs in endothelial cells. A, Measurement of ROS production in endothelial cells. Cells were treated for 45 min with LY294002 (LY, 20 μM), U0126 (U, 10 μM), apocynin (100 μM), rotenone (10 μM), l-NA (100 μM), allopurinol (50 μM), Bay (7.5 μM) or MnTMPyP (100 μM) and then with 10 μg/ml MPs for 24 h. After this, 30-min incubation with the fluorescent dye DHE (3 μM) was performed and the percentage of fluorescent-positive cells was determined by flow cytometry. The results are expressed as mean ± SEM of five independent experiments. ∗, p < 0.05; ∗∗, p < 0.01; ∗∗∗, p < 0.001 vs CTL; †, p < 0.05 vs MPs alone. B, Twenty micrograms of MPs proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against NOX1 and NOX4. C, Endothelial cells were treated for 24 h with MPs, and then NOX1 and NOX4 expression was detected by Western blotting. β-actin control was included. D, Detection of ROS production in the vascular wall. Mice aortas were treated by 10 μg/ml MPs or vehicle and then vessel sections were incubated 30 min with DHE and visualized by confocal microscopy. Results are representative pictures of three independent experiments.

FIGURE 6.

Mechanisms involved in the increase of ROS induced by MPs in endothelial cells. A, Measurement of ROS production in endothelial cells. Cells were treated for 45 min with LY294002 (LY, 20 μM), U0126 (U, 10 μM), apocynin (100 μM), rotenone (10 μM), l-NA (100 μM), allopurinol (50 μM), Bay (7.5 μM) or MnTMPyP (100 μM) and then with 10 μg/ml MPs for 24 h. After this, 30-min incubation with the fluorescent dye DHE (3 μM) was performed and the percentage of fluorescent-positive cells was determined by flow cytometry. The results are expressed as mean ± SEM of five independent experiments. ∗, p < 0.05; ∗∗, p < 0.01; ∗∗∗, p < 0.001 vs CTL; †, p < 0.05 vs MPs alone. B, Twenty micrograms of MPs proteins were subjected to 10% SDS-PAGE and analyzed by Western blotting using Abs raised against NOX1 and NOX4. C, Endothelial cells were treated for 24 h with MPs, and then NOX1 and NOX4 expression was detected by Western blotting. β-actin control was included. D, Detection of ROS production in the vascular wall. Mice aortas were treated by 10 μg/ml MPs or vehicle and then vessel sections were incubated 30 min with DHE and visualized by confocal microscopy. Results are representative pictures of three independent experiments.

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Although MPs expressed NOX1 and NOX4 isoforms of NADPH oxidase at their surface (Fig. 6,B), they were not able to either transfer these proteins to endothelial cells or to increase their expression (Fig. 6 C). Altogether, these data suggest that MPs enhanced ROS production via xanthine oxidase and NF-κB pathways.

In another set of experiences, we evaluated the in situ production and the topographical distribution of ROS in mice aortas. Tissue sections from vessels treated with MPs displayed a marked increase in ethidium bromide fluorescence in the endothelium and the medial layer (Fig. 6 C), reflecting an increase in vascular wall oxidative stress compared with the control.

To evaluate the in vivo physiological significance of MPs in NO production, MPs were injected into mice for 24 h. Under these conditions, MPs decreased the acetylcholine-evoked endothelial relaxation of precontracted vessels (Fig. 7).

FIGURE 7.

MPs impair endothelial function. Acetylcholine-induced relaxation in control (CTL) and MP-injected mouse aorta. Results are expressed as a percentage of relaxation of U46619-induced precontraction. ∗∗∗, p < 0.001 between the two curves.

FIGURE 7.

MPs impair endothelial function. Acetylcholine-induced relaxation in control (CTL) and MP-injected mouse aorta. Results are expressed as a percentage of relaxation of U46619-induced precontraction. ∗∗∗, p < 0.001 between the two curves.

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Taken together, these findings suggest that endothelial dysfunction induced by MPs from apoptotic T cells is linked to the capacity of MPs to decrease NO and increase ROS in endothelial cells.

It is well-accepted that MPs generated under certain conditions possess deleterious properties. Indeed, circulating MPs from patients with cardiovascular and/or inflammatory diseases are able to impair endothelial function (6, 7, 8, 10). We have also reported that MPs from cultured apoptotic T lymphocytes induce endothelial dysfunction similar to that of circulating MPs (8). Although different mechanisms have been proposed to explain how MPs can evoke endothelial dysfunction such as the modifications of the eNOS expression (8) or the endothelial NO transduction pathway (6), little is known about the mechanisms involved in the effects of MPs on enzymes involved in NO generation.

In the present study, we demonstrate that MPs from apoptotic T lymphocytes decreased the NO production and increased oxidative stress in endothelial cells. These effects were associated with a reduction of eNOS activity, which was dependent on PI3K, ERK 1/2, and NF-κB pathways. The increase of ROS production that was down-regulated by PI3K pathway involved xanthine oxidase and NF-κB pathways. Also, caveolin-1 expression and phosphorylation were increased and reduced, respectively, by MP treatment, these effects being independent of PI3K and ERK 1/2 cascade. These data suggest that NO bioavailability is reduced during MP treatment and can account for their capacity to promote endothelial dysfunction, as shown in isolated aorta from mice treated with MPs.

Endothelial production of NO was initially considered to be dependent on increases in intracellular calcium and the binding of calcium/calmodulin to eNOS. However, eNOS regulation is more complex and is determined by a cascade of events, including changes in eNOS mRNA and protein levels (22), association of eNOS with regulatory proteins in the signaling complex (23), changes in intracellular location of the enzyme (24), and phosphorylation of Ser, Thr, and Tyr residues (25). In line with this concept, we have previously reported that MPs from apoptotic T lymphocytes reduce eNOS expression in HUVECs, indicating that these MPs are able to induce endothelial dysfunction (8). Here, using the same MPs but Eahy cell line to avoid differences due to donor of HUVECs, we found that NO production was reduced, whereas eNOS expression was increased. The differential effect induced by MPs on eNOS expression might be due to the use of different cell types. Nevertheless, NO production was decreased in both cell types, and caveolin-1 expression was increased, suggesting that the cell model used in the present study is comparable to the other endothelial cells previously used at least in terms of reduction of NO. In the present study, we also showed that MPs enhanced eNOS phosphorylation on both activator and inhibitor sites, but the eNOS phosphorylation on the latter was higher than in the former, resulting in a decrease of NO produced. Altogether, these results suggest that MPs evoke inhibition of eNOS activity and contribute to the decrease of NO production and endothelial dysfunction.

PI3K/Akt signal pathway is important for regulation of eNOS expression and/or activity. It has been shown that the effects of estrogen (26), VEGF (27) and shear stress (28) on eNOS pathway involve PI3K/Akt signal pathway. In the present study, the inhibition of PI3K by LY294002 prevented the up-regulation of eNOS expression and phosphorylation in MP-treated endothelial cells, as well as the reduction of NO. Also, silencing Akt expression with siRNA to Akt abolished the reduction of NO induced by MPs. Taken together, these findings indicate that MPs modify the eNOS pathway through PI3K/Akt signal transduction.

Interestingly, MPs increased ERK 1/2 phosphorylation, which was partially reversed in the presence of the PI3K inhibitor. In addition, the present study using the MEK inhibitor U0126 clearly showed the involvement of the MAPK pathway in the MP-induced eNOS phosphorylation at both Ser1177 and Thr495 and in NO production. Consistent with these results, ERK 1/2 has recently been shown to be involved in estrogen-induced eNOS phosphorylation (29). Thus, the findings of the present study suggest the existence of cross-talk between the PI3K and MAPK pathways in the MP-induced eNOS phosphorylation. Cross-talk between the PI3K and MAPK pathways in vascular endothelial cells has been observed with insulin-like growth factor-1 (30), but not with either vascular endothelial growth factor (31) or sphingosine 1-phosphate (32) stimulation. To our knowledge, the present work is the first to describe the MP-mediated cross-talk between the MAPK and PI3K pathways in endothelial cells.

The activity of eNOS is also regulated by subcellular localization and/or protein-protein interactions. In this respect, eNOS activity is inhibited when it interacts with caveolin-1 through the caveolin-1-scaffolding domain (20). Data in the literature (33, 34, 35), including our previous report (8), suggest that caveolin-1 could modify not only eNOS activity, but also its expression. In the present study, we observed an increase of caveolin-1 expression in MP-treated cells, which was associated with a decrease of eNOS activity. Caveolin-1 is phosphorylated on Tyr14 by Src family kinases (36, 37, 38), which leads to caveolae fission (39). Here, we reported that MPs increased caveolin-1 expression and decreased its phosphorylation on Tyr14 by a mechanism insensitive to PI3K and MAPK inhibitors, while the NO production is decreased as reported above. Collectively, these findings suggest that, in addition to Thr495 phosphorylation of eNOS reflecting a decrease of eNOS activity, the phosphorylation of caveolin-1 on Tyr14 may have a determinant role in regulating caveolin-1, namely in caveolin-1/eNOS interaction. However, further studies are necessary to decipher the exact role of caveolin-1 phosphorylation in MP-mediated regulation of NO pathway.

Accumulating evidence suggests that ROS play major roles in the initiation and development of vascular diseases such as hypertension, diabetes, or atherosclerosis, acting directly on vascular cells (40). Indeed, ROS act by reducing the bioavailability of NO, impairing endothelium-dependent vasodilatation and endothelial cell growth, causing apoptosis, stimulating endothelial cell migration, and activating adhesion molecules and inflammatory reaction. Here, we examined ROS production in both mouse aortic rings and endothelial cells incubated with MPs. Interestingly, MPs increased ROS production in both endothelial cells and mouse aorta, which may be involved in the impairment of the endothelial function in response to agonist and to flow in both conductance and resistance arteries (8). ROS production in MP-treated endothelial cells was enhanced in the presence of the PI3K inhibitor, but was not influenced by MEK blockade, giving evidence that the PI3K, but not the ERK 1/2, pathway partially and negatively regulates ROS generation. Among the main sources of ROS, NADPH and xanthine oxidases play a major role in vascular cells (41), although eNOS and complex I in mitochondria can also be involved in ROS production. Here, we showed that among the pharmacological inhibitors used, inhibition of xanthine oxidase reduced the ability of MPs to produce ROS. These results suggest that MPs exert their effects on ROS production mainly via the xanthine oxidase pathway. In accordance with our study, the inhibition of xanthine oxidase by allopurinol attenuates endothelial dysfunction associated with diabetes and hypertension (42). In addition, MnTMPyP treatment prevented the increase in ROS evoked by MPs, supporting a role for endothelial cell O2 production as a mechanism underlying the early reduction in NO bioavailability. Thus, inhibition of ROS production through xanthine oxidase activity may represent a potential therapeutic target in vascular pathologies associated with an increase in the circulating level of deleterious MPs.

Finally, we have observed that mechanisms regulating MP effects on endothelial cells are controlled by the NF-κB pathway. Indeed, pharmacological inhibition of this pathway with Bay blunted the enhancement of eNOS and caveolin-1 expression, as well as, the reduction of ROS increase associated with MP treatment.

In summary, MPs activate multiple pathways related to NO and ROS production, mainly through PI3K. Besides, PI3K controls the activation of ERK1/2 cascade, which counteracts the increase of xanthine oxidase-derived ROS production by the former. Furthermore, the NF-κB pathway regulates both NO and ROS production associated with the effects of MPs. All these effects of MPs lead to in vivo endothelial dysfunction into mice. Altogether, these data underscore the pleiotropic effects of apoptotic MPs on NO and ROS in endothelial cells leading to an increase oxidative stress in the vessel wall. The latter may account for the deleterious effects of MPs, resulting in endothelial dysfunction.

We thank Dr. C. Porro for help with MP preparation.

The authors have no financial conflict of interest.

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1

This work was supported in part by grants from Fonds Européen pour le Développement Régional (R.A. 8891), Fondation pour la Recherche Médicale (R.A. INE20050303433 and MCM INE20060306500), Centre National de la Recherche Scientifique, Institut National de la Santé et de la Recherche Médicale Unité et Université d’Angers. H.A.M. and A.A. are recipients of a doctoral fellowship from Conseil Régional du Pays de la Loire and French Education Ministry, respectively.

4

Abbreviation used in this paper: MP, microparticle; ROS, reactive oxygen species; eNOS, endothelial NO synthase; EPR, electronic paramagnetic resonance; DETC, diethyldithiocarbamate; DHE, dihydroethidine; siRNA, small-interfering RNA; MnTMPyP, manganese(III) tetrakis (1-methyl-4-pyridyl) porphyrin pentachloride; O2, superoxide anion; l-NA, nitro-l-arginine.

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