Cell culture.

HCMVEC were purchased from Lonza and cultured in complete EGM-2MV medium containing 5% FBS. Passages between 5 and 8 were used for all experiments. Cells were grown at 37°C in a humidified atmosphere containing 5% CO₂. For all signaling and transendothelial electrical resistance measurement experiments, cells were starved for 1 h in serum-reduced EBM-2 medium containing either 0.02% FBS or 0.1% BSA and then treated in the same medium.

Perfusion system and media composition.

HCMVEC grown on 25-mm glass coverslips were placed inside a Sykes-Moore chamber (1.2 ml; Bellco Glass, Vineland, NJ) as described previously (53). The chamber and inflow tubing were maintained at 37°C. Phenol red-free EBM-2 medium with 0.02% FBS was prebubbled with 5% CO₂, 21% O₂, and 74% N₂ gas for 30 min before each experiment. The medium was continuously channeled into the chamber at a flow rate of 0.25 ml/min. Oxygen tension (PO₂), pH, and the temperature of the perfusate were controlled as previously described (23, 25). Tubing supplying medium to the chamber was made of stainless steel and PharMed polymer (Cole-Parmer Instrument, Chicago, IL).

Video/fluorescent microscopy.

A Nikon TE 2000-U inverted phase/epifluorescent microscope was used for cell imaging. Phase contrast Hoffman modulation optics and a charge-coupled device (CCD) camera were used to monitor morphologic changes over time in the same field of cells (~70 × 90 μm). Fluorescent images were acquired from a cooled Hamamatsu slow-scanning PC-controlled camera (Coolsnap; Photometrics), and changes in fluorescent intensity were quantified with MetaMorph software (Universal Imaging, Downington, PA).

Cell viability.

Cell viability was assessed with the fluorescent exclusion dye PI (5 μM) with excitation of 540 nm and emission of 590 nm. PI fluorescence was used to quantify cell death throughout the entire experiment in a selected field of endothelial cells. At the end of the experiment, all cells were permeabilized with digitonin (100 μM). Cell death was expressed as the PI fluorescence at any given time point relative to the maximal fluorescence observed after cell permeabilization with digitonin.

Measurement of intracellular NO production.

NO generation by HCMVEC was evaluated in the perfusion system by monitoring fluorescence of the NO-selective probe DAF-2 DA (1 μM) following cell treatment with 500 μM H₂O₂ for 10 min. Images were taken at 30 min following H₂O₂ washout with excitation of 480 nm and emission of 520 nm. Fluorescence intensities of cells were measured using MetaMorph software and expressed as arbitrary units.

Measurement of HCMVEC transendothelial electrical resistance.

HCMVEC were grown to confluence in polycarbonate wells containing gold microelectrodes, and transendothelial cell electrical resistance (TEER) measurements were performed using an electrical cell substrate impedance sensing system (Applied Biophysics, Troy, NY) as described previously (27). These measurements provide a highly sensitive biophysical assay that indicates the state of cell shape and focal adhesion (19). TEER values from each microelectrode were pooled at discrete time points and plotted versus time as means ± SE. Each experiment was initiated with HCMVEC monolayer resistance of no less than 1,200 ohm. H₂O₂ (100, 250, or 500 μM, as indicated) was present all course during these experiments.

Caspase activity assay.

HCMVEC treated with or without H₂O₂ were harvested at selected times, washed twice with cold phosphate-buffered saline (PBS), lysed, and stored at −80°C until analysis. Cells stimulated with 10 μM staurosporine for 3 h were used as a positive control. Caspase-3 activity was measured by the ApoAlert caspase profiling assay, as described previously (41).

Western blotting.

After H₂O₂ or control treatments, HCMVEC were washed twice with cold PBS and incubated in lysis buffer (Cell Signaling, Danvers, MA) on ice for 5 min followed by brief sonication. Protein concentrations were determined with a Bradford protein assay. Proteins (50 μg) were separated on 7.5% or 10% gels by SDS-PAGE, transferred to a nitrocellulose membrane, and subjected to immunoblotting with appropriate primary antibody (1:1,000 dilution) overnight at 4°C. After washing and incubation with appropriate secondary antibody, bands were detected with enhanced chemiluminescence kit (Amersham Pharmacia). Densitometric analysis of Western blots was carried out using Quantity One Software (Bio-Rad, Richmond, CA).

Akt activity assay.

The activity of Akt was assessed using a nonradioactive Akt kinase assay kit (Cell Signaling), according to the manufacturer's instructions. Briefly, immunoprecipitation of endogenous Akt was performed overnight on 250 μg of endothelial cell extract with immobilized anti-Akt antibody at 4°C. Following precipitation, the immunoprecipitate was centrifuged, washed two times with lysis and kinase buffers, and then incubated with 1 μg of GSK fusion protein and ATP for 30 min at 30°C in a final volume of 50 μl. The reaction was stopped by adding loading buffer, and the proteins were separated on 10% gels by SDS-PAGE, transferred to nitrocellulose membrane, and probed with anti-phospho-GSK3-α/-β Ser-21/9 antibody. Western blots were developed with the enhanced chemiluminescence kit (Amersham Pharmacia).

High concentrations of H₂O₂ do not induce HCMVEC caspase-3 activation or cell death.

To evaluate whether 500 μM H₂O₂ induced cell death in HCMVEC, we measured caspase-3 activation and cell viability following 30 min H₂O₂ exposure. We found that 500 μM H₂O₂ does not increase but rather decreases the background level of caspase-3 activity in HCMVEC (Fig. 2A). Cell death, as determined by PI exclusion, did not exceed 5% during 30 min of exposure to 500 μM H₂O₂ or during the subsequent 60 min in replacement medium (Fig. 2B). Taken together, these data indicate that the concentrations of H₂O₂ used did not increase cell death in HCMVEC.

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Fig. 2.

H₂O₂ does not induce cell death in HCMVEC. A: HCMVEC were treated with 500 μM H₂O₂ for 30 min and then with basal medium for 2.5 h or with 10 μM staurosporine for 3 h, and caspase-3 activity was assessed in cell lysates using caspase-3-selective fluorigenic substrate (means ± SE; n = 3). B: HCMVEC were treated with 500 μM H₂O₂, and cell death was monitored by propidium iodide (PI) exclusion. Data are presented as means ± SE of 6 independent experiments. *P < 0.05; ***P < 0.001.

H₂O₂-induced activation of NOS3 via Ser-1177 phosphorylation and Thr-495 dephosphorylation.

Maximal activation of eNOS (NOS3) requires phosphorylation at Ser-1177, which is downstream of Akt signaling, and concomitant dephosphorylation of Thr-495 via an Akt-independent process (34, 48, 49). As shown in Fig. 3, A and B, H₂O₂ induces maximal NOS3 phosphorylation in HCMVEC at Ser-1177 and dephosphorylation at Thr-495 sites by 1.5 min after treatment. The phosphorylation-to-dephosphorylation ratio between these two sites peaked at 1.5 min relative to the ratio seen at 10 min after H₂O₂ cell stimulation (Fig. 3C). We also measured H₂O₂-induced NO production using the NO-specific fluorescent probe DAF-2 DA. As shown in Fig. 4, NO generation was substantially upregulated by H₂O₂ and blocked by the NOS inhibitor l-NAME. The selective Akt inhibitor API-2 also attenuated H₂O₂-induced NO production. These results implicated Akt as a regulator of NOS3 in ROS-stressed HCMVEC. To further confirm the involvement of the PI3K-Akt pathway in the regulation of NOS3, we tested the effect of the PI3K inhibitor LY-294002 and the Akt inhibitor API-2 on H₂O₂-induced Akt phosphorylation. Both inhibitors blocked Akt phosphorylation at its Ser-473 and Thr-308 sites (Fig. 5, A and B). In addition, both PI3K and Akt inhibitors blocked H₂O₂-induced NOS3 phosphorylation at Ser-1177 (Fig. 5C), the site regulated by Akt (13, 33). PI3K and Akt inhibitors, however, did not significantly affect the Thr-495 dephosphorylation of NOS3 caused by H₂O₂ (Fig. 5D). This confirms that, as in other endothelial cell types, the PI3K-Akt pathway is not involved in the regulation of the NOS3 Thr-495 site. This site has been found to be phosphorylated by protein kinase C and dephosphorylated by protein phosphatase 1 and 2A and calcineurin (5, 15, 16).

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Fig. 3.

H₂O₂ affects phosphorylation and activation of nitric oxide (NO) synthase (NOS)3 in HCMVEC. HCMVEC were treated with 500 μM H₂O₂ for the indicated time period with or without intermittent medium replacement 10 min after stimulation, and the status of NOS3 Ser-1177 (A) and Thr-495 (B) phosphorylation was determined by Western blotting. C: experimental data from A and B are presented as the ratio between NOS3 p-Ser-1177 and p-Thr-495. Blots represent 1 out of 3 separate experiments. Results are means ± SE from 3 independent experiments. *P < 0.05; **P < 0.01; ***P < 0.001 vs. untreated control cells. eNOS, Endothelial NOS.

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Fig. 4.

Effect of NOS and Akt inhibitors on H₂O₂-induced NO generation. HCMVEC on coverslips were pretreated with the NOS inhibitor N^ω-nitro-l-arginine methyl ester (l-NAME; 200 μM) or the Akt inhibitor Akt/PKB signaling inhibitor-2 (API-2; 10 μM) for 2 h and then treated with 500 μM H₂O₂ for 10 min. Cells were washed with endothelial basal medium-2 (EBM-2), and coverslips were mounted in the perfusion chamber and perfused with EBM-2 containing 1 μM 4,5-diaminofluorescein diacetate (DAF-2 DA). DAF-2 fluorescence was recorded at 30 min with excitation of 480 nm and emission of 520 nm. Fluorescence intensity was quantified using MetaMorph software.

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Fig. 5.

Effect of phosphatidylinositol 3-kinase and Akt inhibitors on Akt and NOS3 phosphorylation. HCMVEC were pretreated with 10 μM API-2 or 10 μM LY-294002 for 2 h before exposure to 500 μM H₂O₂ for 10 min. Lysates were probed for Akt Ser-473 (A) and Thr-308 (B) phosphorylation. Membranes were also probed with anti-Akt antibody to determine total Akt. The same lysates were also analyzed for eNOS phosphorylation at Ser-1177 (C) and Thr-495 (D) and total eNOS. Densitometric analyses represent means ± SE of 3 independent experiments. NS, not significant.

ROS and endothelial cell barrier integrity.

The effect of H₂O₂ on vascular permeability and signaling in endothelial cells of different origin in vitro has been studied, but HCMVEC have only recently become available. In other cell types it is known that H₂O₂ can cause endothelial cell monolayer permeability due to the disruption of cell tight junctions (9, 31, 44) with ERK (14, 24) and p38 (25, 36, 51) MAP kinases mediating these effects of H₂O₂. The current data suggest that HCMVEC adapt after oxidant stress and restore TEER. In our experiments, HCMVEC could restore TEER within 30 min after treatment with 100 μM H₂O₂ and within 3 h after stimulation with 500 μM H₂O₂. Importantly, HCMVEC restored both adherence (VE-cadherin based) and tight (ZO-1 based) junctions, confirming a strong adaptive potential of these cells (Fig. 7). Similar to HCMVEC, rat lung microvascular endothelial cells could restore TEER within 30 min after stimulation with 100 μM H₂O₂ (26). In contrast, bovine pulmonary artery endothelial cells (50) and bovine lung microvascular cells (51) do not restore monolayer integrity for several hours after exposure to as low as 100 μM H₂O₂. Collectively, this indicates relatively high adaptive potential of HCMVEC during exposure to relatively high oxidant stress.

Role of NO in endothelial cell integrity.

H₂O₂ has been shown to activate NOS3 and NO production in the endothelial cell, and NO appears to play some role in both permeability and restoration of barrier integrity in some cell types (34, 48, 49). A direct role for NOS3 in preserving the normal structure of endothelial cell junctions in multiple vascular beds, including cardiac vasculature, was shown using electron microscopy in eNOS^−/− mice (40). Conversely, NO has been demonstrated to be involved in ischemia-reperfusion, platelet-activating factor, histamine, and VEGF-induced microvascular permeability (7, 37). However, despite evidence of NOS3 activation and NO generation, NO in this HCMVEC model appeared to play no major role in either the initial permeability or recovery phases. In HCMVEC, NOS3 seems not to be involved in the regulation of H₂O₂-induced permeability. l-NAME, at concentrations that inhibit NO production (Fig. 4), failed to affect the H₂O₂-elicited changes in TEER (see Fig. 6D). It is likely that the NO release seen in these cells could play other important roles in the heart. The release of NO by endothelial cells results in vasorelaxation and in overall improvement of circulation through the effect of NO on vascular wall smooth muscle cells (3) and NO modulation of cardiomyocyte contractility (22). In our own heart cell models, increased cardioprotective NO generated early during reperfusion can modulate ROS generation by the mitochondria and attenuate cardiomyocyte injury (29, 45). In addition, exogenous NO (as could be provided by neighboring endothelial cells) is highly protective against ischemia-reperfusion injury (21). Thus the NO generation seen in HCMVEC could serve many cardioprotective roles apart from the regulation of microvascular barrier integrity.

Role of Akt in endothelial cell integrity.

The present study examined the role of Akt because it has recently been implicated in endothelial cell barrier function regulation (31, 44). Since it can be simultaneously activated by levels of ROS that activate p38 (39) and induce barrier disruption (51), it is reasonable to think that Akt plays a role in regulating barrier dysfunction in the setting of oxidant stress. In addition, Akt is upstream of NOS3 activation (13) and cardioprotective NO generation (18, 22). Others have shown that H₂O₂ can activate PI3K-Akt signaling pathways and activate NOS3 via downstream NOS3 Ser-1177 phosphorylation (9, 34, 48). Our results suggesting that 250–500 μM H₂O₂ activates PI3K-Akt signaling in HCMVEC (see Fig. 1) are consistent with this previously reported work. The maximal stimulation of NOS3 Ser-1177 phosphorylation in porcine artery endothelial cells was achieved by 600 μM H₂O₂ (49). In other reports, NOS3 phosphorylation in the same cell type was achieved by 100 μM H₂O₂ but not with 500 μM H₂O₂ (48). In both reports, Akt was shown to be critical for eNOS Ser-1177 phosphorylation. Our finding that ROS-mediated Akt Ser-473 and Thr-308 phosphorylation in HCMVEC is mediated by the upstream activation of PI3K (see Fig. 5) is also consistent with work by others using different cell types (9, 49).

In human umbilical vein endothelial cells, Akt has recently been described as a regulator of endothelial chemotaxis that can increase barrier integrity (31). Other work in human pulmonary artery endothelial cells suggests that Akt can improve barrier integrity by phosphorylating the barrier-promoting sphingosine-1-phosphate receptor 1, inducing downstream RhoA/Rac1 signaling to the cytoskeleton (46). Conversely, Akt may mediate ROS stress-induced permeability changes in rat brain endothelial cells (44). In contrast to the current work, this study exposed cells to prolonged superoxide generation using xanthine/xanthine oxidase exposure. Our own results suggest for the first time that Akt in HCMVEC helps restore H₂O₂-induced permeability, given that the Akt inhibitor API-2 completely blocked HCMVEC ability to regain monolayer integrity (see Fig. 6B). Interestingly, API-2 had no effect on the basal TEER, indicating that Akt is not required to maintain the integrity of established cell contacts.

We thank our colleagues Kimberly Wojcik and Michael Retzer for expert medical writing and editorial assistance. We also acknowledge Lynne Harnish for substantial administrative help.