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Am J Physiol Heart Circ Physiol 288: H336-H343, 2005. First published August 26, 2004; doi:10.1152/ajpheart.00025.2004
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Membrane depolarization and NADPH oxidase activation in aortic endothelium during ischemia reflect altered mechanotransduction

Ikuo Matsuzaki, Shampa Chatterjee, Kris DeBolt, Yefim Manevich, Qunwei Zhang, and Aron B. Fisher

Institute for Environmental Medicine, University of Pennsylvania Medical Center, Philadelphia, Pennsylvania

Submitted 12 January 2004 ; accepted in final form 24 August 2004


   ABSTRACT
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 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
We previously showed that "ischemia" (abrupt cessation of flow) leads to rapid membrane depolarization and increased generation of reactive oxygen species (ROS) in lung microvascular endothelial cells. This response is not associated with anoxia but, rather, reflects loss of normal shear stress. This study evaluated whether a similar response occurs in aortic endothelium. Plasma membrane potential and production of ROS were determined by fluorescence microscopy and cytochrome c reduction in flow-adapted rat or mouse aorta or monolayer cultures of rat aortic endothelial cells. Within 30 s after flow cessation, endothelial cells that had been flow adapted showed plasma membrane depolarization that was inhibited by pretreatment with cromakalim, an ATP-sensitive K+ (KATP) channel agonist. Flow cessation also led to ROS generation, which was inhibited by cromakalim and the flavoprotein inhibitor diphenyleneiodonium. Aortic endothelium from mice with "knockout" of the KATP channel (KIR6.2) showed a markedly attenuated change in membrane potential and ROS generation with flow cessation. In aortic endothelium from mice with knockout of NADPH oxidase (gp91phox), membrane depolarization was similar to that in wild-type mice but ROS generation was absent. Thus rat and mouse aortic endothelial cells respond to abrupt flow cessation by KATP channel-mediated membrane depolarization followed by NADPH oxidase-mediated ROS generation, possibly representing a cell-signaling response to altered mechanotransduction.

ATP-sensitive potassium channels; KIR6.2; gp91phox; fluorescence microscopy; flow adaptation

ENDOTHELIAL CELLS EXPOSED to blood flow are known to express mechanosensors that convert shear stress-related mechanical forces on the plasma membrane to specific cellular signals (5, 8). Increased shear stress can modulate endothelial cell function by initiating a wide range of responses (5, 9, 29), including activation of flow-sensitive ion channels (5–7), altered gene expression (14, 26), and cytoskeletal reorganization (10, 24). In contrast, the effects of decreased shear due to various causes, such as embolism, thrombosis, or shock, are relatively poorly understood.

We previously used imaging techniques with fluorescent indicators in an isolated rat lung model to detect pulmonary endothelial responses in situ to loss of shear stress. Flow cessation resulted in depolarization of the endothelial cell plasma membrane and generation of reactive oxygen species (ROS) in isolated perfused lungs (2–4, 28) and in flow-adapted pulmonary vascular endothelial cells in vitro (21). This response was shear dependent and independent of changes in intravascular pressure (4).

Vascular heterogeneity among vessel types from different organs has been well recognized. Thus it is not clear whether the responses observed in the pulmonary vasculature would be seen in a systemic vascular bed. The goal of this study was to evaluate the reaction of aortic endothelial cells to altered shear stress with use of in situ and cell culture systems. The present study shows that these cells respond to flow cessation with cell membrane depolarization followed by generation of ROS similar to that described for pulmonary endothelium. Using gene-targeted mice, we determined that an ATP-sensitive K+ (KATP) channel is responsible for the plasma membrane depolarization and that the plasma membrane NADPH oxidase is the ROS generator.


   MATERIALS AND METHODS
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 ABSTRACT
 MATERIALS AND METHODS
RESULTS
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Materials. Bis-(1,3-dibutylbarbituric acid)trimethine oxonol (bis-oxonol), Amplex red, and 1,1'-dioctadecyl-1,3,3',3'-tetramethylindocarbocyanine perchlorate acetylated LDL were purchased from Molecular Probes (Eugene, OR). Cromakalim, a K+ channel agonist, diphenyleneiodonium chloride (DPI), a flavoprotein inhibitor, bovine erythrocyte superoxide dismutase (SOD), and ferricytochrome c (cyto c) from horse heart were purchased from Sigma (St. Louis, MO). Catalase was obtained from Boehringer Mannheim (Indianapolis, IN). Rat aortic endothelial cells (RAEC) and complete RAEC culture medium were purchased from VEC Technologies (Rensselaer, NY); cells from passages 3–11 were used.

Sprague-Dawley male rats weighing 180–220 g were obtained from Charles River Breeding Laboratories (Kingston, NY). KATP channel (KIR6.2)-knockout mice (22) were obtained originally from Dr. S. Seino (Dept. of Pharmacology, Chiba University) and bred in our institutional Animal Care Facilities. NADPH oxidase (gp91phox)-knockout mice and wild-type C57BL/6 mice were purchased from Jackson Laboratories (Bar Harbor, ME). KIR6.2- and gp91phox-knockout mice had been backcrossed to the C57BL/6 background. All animal study protocols were approved by the University of Pennsylvania Institutional Animal Care and Use Committee.

Vessel preparation and laminar flow chamber. A parallel plate chamber was used to study flow effects in aortic tissue. Animals (rats or mice) were anesthetized with pentobarbital sodium (50 mg/kg ip), tracheotomized, and placed on a ventilator. After the chest was opened, the thoracic aorta was immediately removed and carefully trimmed to remove excess adventitial tissue. The aorta was cut longitudinally into 5-mm-long sections (~20–30 mg each) and then immediately fixed to a glass slide with instant glue on one corner, with the endothelial cell layer facing up (Fig. 1A). The edges of the tissue were sealed with adhesive tape to keep the preparation flat. The slide with affixed tissue was placed in the tissue flow chamber (Fig. 1B). The total ischemic time from death of the animal to the start of reflow adaptation was <5 min.



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  		Fig. 1. Experimental setup. A: thoracic aorta was removed from rats or mice and cut longitudinally into ~5-mm-long sections. Aortic section was fixed to a glass slide at one corner with instant glue, and edges were sealed with tape. B: schematic of perfusion circuit. Chamber containing aortic tissue fixed to the glass slide was placed in the stainless steel chamber. C: flow chamber was placed on the stage of an inverted fluorescence microscope for imaging studies. CCD, charge-coupled device.

 
The flow chamber consisted of a steel plate (7.5 x 3.5 x 0.4 cm) with a central hollow slot. The glass slide containing aortic tissue was placed on the top, and the bottom was sealed with a coverslip (2.2 x 2.2 cm) to create a rectangular flow channel (1.5 x 1.5 x 0.02 cm). The flow chamber was installed between two reservoirs containing culture medium; after passage through the flow chamber, medium collected in the first reservoir and was recirculated to the second reservoir by means of a peristaltic pump (Fig. 1B). This double-reservoir system resulted in laminar flow to the aortic preparation. The volume of recirculating perfusate was ~10 ml, and the volume of the chamber was ~200 µl. The perfusate was Krebs-Ringer bicarbonate solution (in mmol/l: 118 NaCl, 4.7 KCl, 1.2 MgCl2, 1.2 MgSO4·7H2O, 1.3 CaCl2·2H2O, 1.2 KH2PO4, and 24.9 NaHCO3) with 10 mM glucose + 25 mM HEPES, pH 7.4, equilibrated with 5% CO2 in air. For experiments with high K+, NaCl was replaced equivalently with KCl. Aortic tissue in the flow chamber was subjected to shear stress, generally at 10 dyn/cm2, for 1 h (reflow adaptation) before experiments. The shear stress ({tau}) to which the cells were exposed was calculated as follows: {tau} = (6µ/h2b), where µ is the dynamic viscosity, b is the flow chamber width, h is the flow chamber height, and is the flow rate. Ischemia was produced by abrupt cessation of perfusate flow. During the ischemic period, the upper plate was loosened, so that the static perfusate was exposed in part to air to maintain oxygenation. In some experiments, a graded alteration of shear stress was produced by decreasing, rather than stopping, the perfusate flow.

Cell culture. RAEC in complete medium (pH 7.4) were plated on a glass slide (44 x 20 mm) that had been coated with 0.2% gelatin. Cells were allowed to grow for ≥24 h until they became fully confluent. Cells were then cultured in a laminar flow chamber under static conditions or continuous laminar flow as described above for aortic preparations. The chamber used for flow adaptation of RAEC in monolayer culture was a commercially available apparatus (Confocal Imaging chamber RC-30, Warner Instruments, Hamden, CT). For adaptation to flow, a slide with adherent cells was perfused in the flow chamber with growth medium supplemented with 25 mM HEPES (pH 7.4) at 37°C generally for 24 h at an estimated shear stress at the cell surface of 5 dyn/cm2. A similarly prepared slide was maintained under static culture conditions for the same duration (i.e., static cells). Microscopic evaluation showed that cells exposed to laminar flow reoriented with the long axis in the direction of flow (not shown) as described previously for endothelial cell flow adaptation (9, 10). After flow adaptation, the growth medium was substituted with a standard Krebs-Ringer bicarbonate buffer supplemented with 10 mM glucose, 25 mM HEPES, and 3% dextran (Sigma), pH 7.4, and the cells were incubated as described above.

Microscopy. An inverted epifluorescence microscope (Diaphot TMD, Nikon) equipped with an optical filter changer (Lambda 10-2, Sutter Instrument, Novato, CA), a digital camera (model ORCA-100, Hamamatsu), and MetaMorph imaging software (Universal Imaging, West Chester, PA) was used for imaging as previously described (28). The flow chamber with intact aorta or the cultured cell monolayer was mounted on the stage of the microscope. Tissue or cells in the flow chamber were preperfused with the membrane potential-sensitive fluorophore bis-oxonol (200 nM) with or without inhibitors for 30 min or with the H2O2-sensitive fluorophore Amplex red (2.5 µM) + horseradish peroxidase (0.01 U/ml) for 10 min before flow cessation. Excitation for fluorescence imaging was accomplished with a mercury lamp fiber-optic light source and appropriate filter set: for bis-oxonol, HQ-41001 with 480 ± 20 nm excitation, 505LP dichroic, and 535 ± 25 nm emission; for Amplex red or 1,1'-dioctadecyl-1,3,3',3'-tetramethylindocarbocyanine perchlorate acetylated LDL, HQ-41002b with 545 ± 15 nm excitation, 570 LP dichroic, and 610 ± 37.5 nm emission (Chroma Technology, Brattleboro, VT). For quantitation, areas of interest were randomly selected, and the fluorescence intensity of each was measured. Fluorescence intensity was normalized as a percent change in intensity level from baseline.

Biochemical measurements. O2–· production by aortas during graded decrease of flow was measured by reduction of cyto c. Cyto c (100 µM) was added to the perfusate during the 60-min period of reflow adaptation at 10 dyn/cm2 shear stress; catalase (50 µM) was added to prevent reoxidation of cyto c by H2O2. The flow was then abruptly reduced to give a shear stress of 0 ("ischemia"), 0.25, 0.5, or 1 dyn/cm2. A different aorta preparation was used for each reduced shear experiment. The perfusate was collected during the first min of reduced flow or aspirated from the chamber for the zero-flow condition, and its absorbance and that of a sample of recirculated perfusate were measured at 550 nm. O2–· production during the 60-min reflow-adaptation period was calculated from the change in absorbance at the start and end of reflow adaptation. The absorbance reading of the 0 and 0.25 dyn/cm2 samples exceeded 10 times the threshold sensitivity of the spectrophotometer. O2–· production was calculated from the change in absorbance using an extinction coefficient of 21 mM–1·cm–1 for reduced cyto c and normalized to the wet weight of the aorta preparation.

To determine adequacy of oxygenation during ischemia, the PO2 phosphorescent probe PdP1 (50 µM) was added to the perfusate (21). The medium in the chamber was aspirated after 10 min of ischemia, and phosphorescence (excitation at 524 nm, emission at 690 nm) of the aspirate was measured using a luminescence spectrometer (model LS50B, Perkin-Elmer, Buckinghamshire, UK) as described previously (21). Full-scale deflection obtained by addition of glucose oxidase to the sample to deplete O2 was 950 counts/s above background. This method shows increased phosphorescence at PO2 <45 mmHg.

To measure ATP content of the cells, the chamber was flushed at the end of the experiment with 100 µl of ice-cold 2 M perchloric acid, and the cell extracts were incubated and neutralized as described previously (17). ATP content of the endothelial cell lysate was measured with an ATP chemiluminescence assay kit (Molecular Probes) using a luminometer (model TD-20/20, Turner Designs, Sunnyvale, CA). Cellular protein content was measured by Lowry assay (20).

Statistical analysis. Results are expressed as means ± SE or means ± ranges for n = 2. Comparison among conditions was made using ANOVA with Bonferroni's test (SigmaStat, Jandel Scientific, San Rafael, CA). Differences were considered to be significant at P < 0.05.


   RESULTS
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
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The relation between magnitude of shear stress during 1 h of reflow adaptation of the aorta and the subsequent ischemic response was studied. Aortas were flow adapted with shear stress varying from 0 to 15 dyn/cm2, and ROS generation with ischemia was measured with Amplex red as a fluorophore. Aortas flow adapted to 1 dyn/cm2 showed a significant increase in Amplex red fluorescence with ischemia and a greater response when reflow adaptation was 2 dyn/cm2 (Fig. 2). There was no significant difference in ischemic response with shear stress that varied from 2 to 15 dyn/cm2 during reflow adaptation (Fig. 2). Further studies of ischemic response were carried out using 10 dyn/cm2 shear stress during reflow adaptation.



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  		Fig. 2. Relation between magnitude of shear stress during 60 min of "reflow adaptation" and subsequent ischemic response, with Amplex red fluorescence used as indicator. Data represent fluorescence intensity after 10 min of ischemia. In zero-shear stress condition (zero flow), fluorescence intensity decreased by 11% during the subsequent 10-min observation period, and increased fluorescence was noted if tissue had been exposed to flow. Values are means ± SE; n = 3. *P < 0.05 vs. preceding time point.

 
An increase in ROS generation indicated by increased Amplex red fluorescence was observed after ~2–3 min of flow cessation in reflow-adapted aortas and increased linearly over the next 10 min (Fig. 3A). Pretreatment with catalase (1,000 U/ml) to scavenge H2O2 markedly suppressed the change in Amplex red fluorescence (Fig. 3A). ROS generation with flow cessation was not seen in aortic tissue during continuous flow (Fig. 3A). Preincubation with 10 µM DPI or 30 µM cromakalim significantly suppressed the changes in Amplex red fluorescence, indicating inhibition of ROS generation (Fig. 3B). The use of Amplex red to detect cellular generation of ROS primarily reflects H2O2 and is supported by the inhibiting effect of catalase. Although the Amplex red probe and catalase are extracellular, H2O2 could arise from intra- or extracellular sources, because H2O2 readily crosses cell membranes.



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  		Fig. 3. Reactive oxygen species (ROS) generation in endothelium of rat aorta detected by Amplex red fluorescence measured at 610 ± 37.5 nm (545 ± 15 nm excitation). A: increase in Amplex red fluorescence with cessation of flow in reflow-adapted (10 dyn/cm2) aortic tissue indicates ROS generation. Fluorescence in tissues pretreated with catalase (1,000 U/ml) or subjected to continuous flow did not change significantly with flow cessation. Values are means ± SE; n = 3. *P < 0.05 vs. other conditions. B: effect of diphenyleneiodonium chloride (DPI, 10 µM) or cromakalim (30 µM) on Amplex red fluorescence in reflow-adapted aortic endothelium. Amplex red fluorescence is shown 5 and 10 min after flow cessation. Perfusion for 30 min with DPI or cromakalim significantly inhibited H2O2 generation. Values are means ± SE; n = 3. *P < 0.05 vs. control.

 
ROS generation in reflow-adapted aortas also was evaluated by reduction of cyto c added to the perfusate. This method indicates extracellular generation of O2–·, because cyto c remains in the extracellular space and O2–· crosses cell membranes relatively slowly. O2–· generation was minimal under flow conditions, but production increased markedly with ischemia (Table 1). The graded decrease in flow showed a threshold effect with no change in O2–· production until shear stress was reduced by ~95% from the reflow-adaptation value (Table 1). Reduction of cyto c with ischemia was completely abolished by addition of SOD (100 µM) to the perfusate (not shown).


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  		Table 1. O2–· generation by reflow-adapted rat aorta as detected by cytochrome c reduction under various shear stress conditions

 
Increased ROS generation with flow cessation also was seen in RAEC that had been flow adapted at 5 dyn/cm2 shear stress in cell culture (Fig. 4). The increase in Amplex red fluorescence with flow cessation was completely blocked by the presence of catalase (100 U/ml). Control RAEC that had been cultured under static conditions and exposed to flow for only 10 min to load the dye showed no significant change in Amplex red fluorescence with flow cessation (Fig. 4).



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  		Fig. 4. Effect of flow cessation on Amplex red fluorescence in cultured rat aortic endothelial cells (RAEC). Fluorescence was increased significantly with cessation of flow in flow-adapted (5 dyn/cm2 for 24 h) RAEC; fluorescence change was not significant in RAEC grown under static conditions. Pretreatment with catalase prevented increased Amplex red fluorescence with flow cessation. Values are means ± SE; n = 3. *P < 0.05 vs. static or catalase-treated cells.

 
Bis-oxonol was used as a fluorescent indicator for changes in endothelial cell membrane potential. RAEC in situ that had been reflow adapted showed a rapid initial increase in bis-oxonol fluorescence with flow cessation followed by a slower rate of increase to an apparent plateau value at ~2–3 min (Fig. 5). The increase in fluorescence indicates plasma membrane depolarization. The change in bis-oxonol fluorescence with flow cessation was markedly attenuated by pretreatment of aortas with cromakalim, a KATP channel agonist (Fig. 5). To assess tissue oxygenation during ischemia, the perfusate in contact with the aorta was aspirated 10 min after flow cessation. Analysis of the sample by phosphorescence assay showed PO2 >45 mmHg, indicating adequate oxygenation during the ischemic period.



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  		Fig. 5. Change in plasma membrane potential after ischemia in rat aortic endothelium as measured by bis-(1,3-dibutylbarbituric acid)trimethine oxonol (bis-oxonol) fluorescence. Rat aortas were subjected to reflow adaptation by perfusion for 1 h at shear stress of 10 dyn/cm2. Aortas were then perfused for 30 min with 200 nM bis-oxonol in the presence or absence of cromakalim (30 µM). Fluorescence was monitored at 535 ± 20 nm (480 ± 25 nm excitation) during continuous flow or with acute flow cessation (no treatment and cromakalim traces). Increase in fluorescence with cessation of flow indicates plasma membrane depolarization. Values are means ± SE; n = 3–4. *P < 0.05 vs. continuous flow.

 
The membrane depolarization response of endothelium to ischemia was also studied with RAEC in monolayer culture. Cells were cultured under static conditions or were flow adapted at 5 dyn/cm2 for 24 h before study. Cells that were flow adapted showed an increase in bis-oxonol fluorescence after flow cessation (Fig. 6), indicating depolarization similar to that in the intact aorta. There was no significant change in bis-oxonol fluorescence with cessation of flow in static (control) RAEC that were subjected to flow for only 30 min (to load the dye) or in continuously perfused flow-adapted cells (Fig. 6B). ATP content of RAEC was measured to determine whether decreased ATP could be responsible for the membrane depolarization. ATP content was 13.7 ± 2.1 nmol/mg protein in control cells adapted to 5 dyn/cm2 and 15.4 ± 3.2 nmol/mg protein in flow-adapted cells 10 min after cessation of flow (mean ± SE, n = 3); this difference was not statistically significant. The lack of change in ATP content indicates that cellular oxygenation was adequate during the stop-flow period.



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  		Fig. 6. Change in membrane potential in RAEC with flow cessation as detected by bis-oxonol fluorescence. A: fluorescence images of RAEC loaded with bis-oxonol (200 nM) during continuous flow (a) and bis-oxonol fluorescence intensity (shown in pseudocolor) at time 0 and 1 and 5 min after flow cessation with flow-adapted (5 dyn/cm2) cells and cells grown under static conditions (b). B: bis-oxonol fluorescence at 1-min intervals in continuously perfused flow-adapted cells and after flow cessation in flow-adapted cells and cells cultured under static conditions. Values are means ± SE; n = 3–4. *P < 0.05 vs. continuous flow.

 
To confirm that a change in cell membrane potential of aortic endothelium could result in ROS production, aortas were perfused with an isotonic solution containing 24 mM K+, instead of the usual 5.9 mM K+, to depolarize the endothelium during continuous flow. ROS production by rat aortic tissue during continuous flow was detected 5 and 10 min after the solution was switched from normal to high K+ and was markedly inhibited by the presence of DPI (Fig. 7).



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  		Fig. 7. High-K+-induced ROS generation in rat aortic endothelium as a positive control for effect of cellular depolarization. After they were loaded with Amplex red for 10 min with or without DPI (10 µM), aortas were subjected to perfusion with isosmotic buffer containing 24 mM K+ under continuous-flow conditions in the absence (control) or presence of DPI (10 µM) added during the loading period. Values are means ± SE; n = 3. *P < 0.05 vs. control.

 
To further investigate the link between endothelial cell membrane depolarization and ROS generation after flow cessation, we evaluated aortas from mice with knockout of KIR6.2, the pore-forming unit of the KATP channel in endothelial cells (22). We showed previously in pulmonary vascular endothelial cells in culture that induction of this channel occurs during flow adaptation and is required for membrane depolarization with cessation of flow (7). Membrane potential change in aortas from KIR6.2-knockout mice was significantly diminished compared with that from wild-type mice (Fig. 8A). ROS generation with cessation of flow also was significantly smaller in the KIR6.2-knockout than in the wild-type mice (Fig. 8B).



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  		Fig. 8. Membrane potential change and ROS generation in endothelium of isolated aortic sections from wild-type mice and mice with knockout (KO) of KIR6.2 or gp91phox. A: membrane potential change detected with bis-oxonol. Values are means ± SE; n = 3. *P < 0.05 vs. continuous flow. B: ROS generation at 5 and 10 min of ischemia as detected by Amplex red fluorescence. Values are means ± SE; n = 3–5. *P < 0.05 vs. wild type.

 
On the basis of our previous studies with lungs (3), we postulated that a cell membrane NADPH oxidase is responsible for ROS production with flow cessation. We used mice with knockout of gp91phox (the integral membrane flavoprotein component of NADPH oxidase) to evaluate the role of the enzyme complex in the aortic endothelium. Membrane depolarization following flow cessation was similar for wild-type and gp91phox-knockout aortic endothelium (Fig. 8A), but ROS generation was markedly diminished in the NADPH-deficient aortas (Fig. 8B).


   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
We reported previously that cessation of flow (ischemia) results in endothelial cell membrane depolarization and subsequent generation of ROS in isolated lung in situ (3, 4, 28) and flow-adapted pulmonary vascular endothelial cells in vitro (21). We have termed this model "oxygenated" or "normoxic" ischemia, because the studies are carried out with continued oxygenation from ambient sources, in contrast to most ischemia models, where anoxia results. Anoxia would, of course, abolish ROS production because of the absence of O2 and could independently influence membrane potential through change in ATP. This lung model has enabled us to establish an endothelial cell response paradigm related to altered shear stress. Previously published evidence for a similar response in organs other than the lung is the finding of ROS generation during the early phase of ischemia when O2 is available. ROS generation in ischemia has been indicated by electron paramagnetic resonance studies of the heart (13) and by the salicylate hydroxylation method for the gut (30), although the emphasis in those studies was on the more robust ROS production with reperfusion. To directly study the response of a systemic vascular bed, we have developed a method using isolated perfused rat or mouse aorta that allows imaging of the endothelial cell layer. Although it is unlikely that aortic endothelium would be normally subjected to the large decrease in shear necessary to elicit the ROS response, these cells can serve as a convenient model for other systemic endothelia. We have used the isolated aorta preparation and cultured aortic endothelial cells that have been flow adapted in vitro to investigate a shear stress-associated pathway for ROS generation with flow cessation.

Membrane depolarization was observed with cessation of flow in aortic endothelial cells in situ. A similar response was observed in flow-adapted RAEC in vitro but did not occur in cells that had been maintained in culture under static conditions. On the basis of our previous studies with pulmonary microvasculature, we propose that membrane depolarization with ischemia results from inactivation of cell membrane KATP channels (7). Endothelial cells from rat aorta have demonstrated the presence of membrane currents associated with KATP channels (16). In the present experiments, a KATP channel agonist, cromakalim, suppressed the bis-oxonol response to flow cessation, providing evidence that depolarization is associated with KATP channel inactivation. To further evaluate whether a KATP channel is the element responsible for membrane depolarization, we studied aortas isolated from mice with knockout of KIR6.2 (22). Compared with aortas from wild-type mice, endothelial cell membrane potential change after ischemia in the KIR6.2-knockout mice was greatly diminished. Thus results obtained with the KATP channel agonist and the KATP channel-knockout mice indicate that this channel is involved in the cell membrane depolarization response to cessation of flow. Our recent studies showed that rat pulmonary microvascular endothelial cells in culture demonstrate only low levels of KATP channel expression, and channel expression and inwardly rectified membrane current increased significantly with flow adaptation (7). Increased channel expression may be responsible in part for the augmented response to ischemia of aortic endothelial cells after flow adaptation. Endothelial cells in situ would be expected to be in a flow-adapted state.

We used Amplex red as an indicator of ROS generation. This probe reacts with H2O2 in the presence of horseradish peroxidase to form the fluorescent product resorufin. The studies using cyto c reduction indicate that O2–· is generated extracellularly during ischemia. Therefore, extracellular H2O2 detected by Amplex red likely arises from dismutation of extracellular O2–·, either spontaneously or catalyzed by extracellular SOD (25). ROS generation with flow cessation was observed in flow-adapted endothelial cells but not in cells that had been cultured under static conditions. Recently published studies indicate that endothelial cells possess a plasma membrane NADPH oxidase-like enzyme system (3, 12). Studies with the flavoprotein inhibitor DPI are compatible with NADPH oxidase as the ROS generator, but this inhibitor is not specific. The absence of ROS generation with ischemia in gp91phox-knockout mice confirms that NADPH oxidase is normally required for ROS generation. Extracellular generation of superoxide is consistent with the presumed localization of NADPH oxidase to the plasma membrane, as has been demonstrated for neutrophils and macrophages (12) and in our previous studies with bovine pulmonary artery endothelial cells (21).

In the present study, neither addition of DPI nor knockout of gp91phox altered the cell membrane depolarization response to flow cessation, indicating that electron transfer by the oxidase was not the mechanism for altered membrane potential. On the other hand, addition of a KATP channel agonist (cromakalim) or knockout of KIR6.2 resulted in a greatly diminished response, indicating that endothelial cell membrane depolarization is required to activate NADPH oxidase with flow cessation. Oxidant generation by other nonexcitable cells, such as the neutrophil and the macrophage, also has been shown to involve membrane depolarization associated with activation of the membrane-bound oxidase system (18, 19). We previously showed that K+-induced membrane depolarization in bovine pulmonary artery endothelial cells in vitro and microvascular endothelial cells in the intact rat lung results in ROS production (1–3). A subsequent study showed high-K+-induced ROS production in human umbilical vein endothelial cells (27). Although the precise mechanism for coupling of depolarization and activation of NADPH oxidase is not clear, protein tyrosine phosphorylation and Rac translocation appear to be involved (27).

As postulated for fibroblasts, macrophages, and now other cells, generation of ROS appears to represent an important cell-signaling mechanism (11, 15). Our previous studies with pulmonary endothelium showed that ROS generation with endothelial cell ischemia results in cell signaling through the activation of transcription factors, e.g., NF-{kappa}B and activator protein-1, and endothelial cell proliferation (23, 31). The present study with aortic cells shows that a systemic vascular endothelium also responds to acute reduction in shear stress with ROS production. The change in shear required to elicit the response is ~80% of the control flow in the pulmonary vasculature (4) and slightly greater in the present study of the aorta. Thus this response is elicited with severe impairment of flow and not with normal physiological flow variation. Because O2 is required for ROS production, this response would not occur during established anoxia but could be observed during the initial stages of vascular occlusion or during partial vascular obstruction when PO2 remains adequate for O2–· production by the NADPH oxidase.

In summary, we have found in rat and mouse aortic endothelium that simulated ischemia results in ROS production subsequent to membrane depolarization. Depolarization requires the presence of a cell membrane KATP channel, and ROS generation results from activation of a cell membrane NADPH oxidase. ROS production with ischemia may represent an important signaling mechanism associated with altered mechanotransduction.


   GRANTS
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 ABSTRACT
 MATERIALS AND METHODS
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This work was supported by National Heart, Lung, and Blood Institute Grant HL-75587.


   ACKNOWLEDGMENTS
 
We thank J. Rossi for typing the manuscript.

Present address of I. Matsuzaki: Dept. of Surgery, Akita University, Akita, Japan.


   FOOTNOTES
 

Address for reprint requests and other correspondence: A. B. Fisher, Institute for Environmental Medicine, Univ. of Pennsylvania School of Medicine, 1 John Morgan Bldg., Philadelphia, PA 19104-6068 (E-mail: abf{at}mail.med.upenn.edu)

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


   REFERENCES
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
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 REFERENCES
 

  1. Al-Mehdi AB, Ischiropoulos H, and Fisher AB. Endothelial cell oxidant generation during K+-induced membrane depolarization. J Cell Physiol 166: 274–280, 1996.[CrossRef][Web of Science][Medline]
  2. Al-Mehdi AB, Shuman H, and Fisher AB. Oxidant generation with K+-induced depolarization in the isolated perfused lung. Free Radic Biol Med 23: 47–56, 1997.[CrossRef][Web of Science][Medline]
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