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Vol. 18, Issue 6, 2002-2012, June 2007
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*Institute for Environmental Medicine, University of Pennsylvania, Philadelphia, PA 19104-6068; and Institute of Pathology, Johannes-Gutenberg University, D-55101 Mainz, Germany
Submitted September 18, 2006;
Revised January 10, 2007;
Accepted March 2, 2007
Monitoring Editor: John Cleveland
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). It is now appreciated that a major source of ROS in the vasculature is through one or more isoforms of the phagocytic enzyme NADPH oxidase, a membrane-localized protein which generates the superoxide (O2.–) anion on the extracellular surface of the plasma membrane (Lambeth, 2004). As a charged and short-lived anion, it is believed that O2.– flux is insufficient to initiate intracellular signaling due to the combination of poor permeability through the phospholipid bilayer (Tanabe et al., 2005) and a rapid dismutation to its uncharged and more stable derivative, hydrogen peroxide (Finkel, 2003). However, recent evidence has indicated discrete signaling roles for both O2.– and H2O2 (Madesh and Hajnoczky, 2001; Devadas et al., 2002). In our studies, we have found that extracellular O2.–, but not H2O2, leads to Ca2+ signaling and apoptosis in pulmonary endothelial cells (Madesh et al., 2005). This indicates that extracellular O2.– produced by NADPH oxidase or other sources either crosses the plasma membrane or modifies cell surface proteins to mediate cell signaling.
Previous studies of erythrocytes and amniotic cells have provided evidence for O2.– transport through anion channels, which could be effectively blocked by 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid (DIDS; Lynch and Fridovich, 1978; Ikebuchi et al., 1991). DIDS also effectively blocked release of O2.– from mitochondria into the cytosol (Han et al., 2003) without affecting ROS production (Korchak et al., 1980). Despite these reports, whether O2.– crosses the cell membrane to elicit a discrete intracellular signal remains controversial (Babior, 1999; Mikkelsen and Wardman, 2003).
The present study evaluated the response of pulmonary microvascular endothelial cells (PMVECs) to extracellular O2.–. Our findings using a fluorophore trap demonstrate that O2.– enters the cell through a chloride channel-3 (ClC-3)-dependent mechanism. Further, extracellular O2.–, through a Ca2+-mediated signaling event, stimulates the production of O2.– by the mitochondria. This observation provides a model by which extracellular O2.– can propagate intracellular ROS signaling.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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-actin, ClC-3, and ClC-4 were obtained from Operon Biotechnologies (Huntsville, AL). Rabbit polyclonal anti-ClC-3 antibody was obtained from Santa Cruz Biotechnology (Santa Cruz, CA). pEYFP-Mito was purchased from Clontech (BD Biosciences, Mountain View, CA). KO2, apocynin, angiotensin II, thapsigargin, and all other chemicals were purchased from Sigma (St. Louis, MO). Mice were obtained from The Jackson Laboratory (Bar Harbor, ME).
Cell Culture
Immortalized human pulmonary microvascular endothelial cells (HPMVEC clone ST1.6R) were generated as described previously (Krump-Konvalinkova et al., 2001) and cultured in Medium-199 supplemented with 15% FBS, glutamax, antibiotics, and endothelial cell growth supplement. Isolation, characterization, and propagation of mouse pulmonary microvascular endothelial cells (MPMVEC) from wild-type (C57BL/6) and gp91phox gene-targeted mice have been previously described (Milovanova et al., 2006). Primary cells were cultured in DMEM supplemented with 10% FBS, nonessential amino acids, endothelial cell growth supplement, and antibiotics and used between passages 6 and 20. For some experiments cells were transfected with various DNA constructs by electroporation (Amaxa Biosystems, Gaithersburg, MD) using programs T-23 or S-05 according to the manufacturer's instructions.
Imaging of O2.– Flux
PMVECs cultured on 0.2% gelatin-coated 25-mm diameter glass coverslips were loaded with the O2.–-sensitive dye HE (10 µM) in DMEM for 10 min at 37°C. Cells were then placed on a temperature-controlled stage, and images were recorded every 5 s for 5 min using LaserSharp software (Bio-Rad Laboratories, Hercules, CA) on a Bio-Rad Radiance 2000 imaging system (Bio-Rad Laboratories) equipped with a Kr/Ar-ion laser source at 568- and 605-nm excitation and emission, respectively, using a 60x oil objective. A bolus of KO2 was added after 1 min of baseline recording. KO2 was prepared in a 1.8 mM concentration as described previously (Reiter et al., 2000). KO2 was not directly applied to the image field to avoid alterations in microscope focus. For inhibitor studies, DIDS (200 µM) was present during HE loading and KO2 addition. Antioxidant enzymes were added immediately before imaging and mitochondrial inhibitors were added similarly as KO2. HE fluorescence was quantified by nuclear masking of all cells in the field. For angiotensin II (Ang II) and thrombin experiments, HPMVECs were cultured on coverslips, and the medium was replaced with M-199 containing 2% FBS 18 h before study. HPMVECs were pretreated with DIDS (300 µM) or apocynin (Apo; 2 µM) for 10 min before addition of 2 µM Ang II or 0.5 U/ml thrombin. For imaging, cells were loaded with HE (10 µM) and five independent fields were recorded by confocal microscopy.
Cell-free HE Oxidation Measurement
HE fluorescence (40 µM) in a 2 ml solution of PBS was monitored in a multiwavelength-excitation dual wavelength-emission fluorimeter (Delta RAM, PTI, Birmingham, NJ) using 510- and 568-nm excitation and emission, respectively. Briefly, KO2 or the xanthine/xanthine oxidase (X/XO; X-100 µM; XO-50 mU/ml) O2.–-generating system was added to the solution after 60 s of baseline recording. Total recording time was 3 min. DMSO, H2O2, and KOH were added in a similar manner. For dismutation studies, KO2 was added to a solution of PBS containing 1000 U superoxide dismutase (SOD), mixed briefly, and then added to the HE solution. Results were normalized to the baseline fluorescence before addition of O2.–. The stable oxidation product was assessed in intact MPMVECs loaded with HE. Briefly, cells were treated for 20 min with either antimycin A or a 10 µM bolus of KO2 and a spectral scan of emission wavelengths was performed using an excitation wavelength of 494 nm.
O2.– -induced HE Fluorescence and Mitochondrial ROS Production
PMVECs cultured on coverslips were loaded with HE and mounted on a confocal microscope stage as described earlier. After measurement of HE baseline fluorescence, KO2 (10 µM) or X/XO (X-100 µM; XO-20 mU/ml) was added to the medium evenly across the coverslip and gently agitated to mix the solution. After 20 min, five fields were chosen for imaging and quantitation. To measure O2.– in mitochondria, MPMVECs were transfected with 2 µg/ml pEYFP-Mito (Clontech, BD Biosciences) and cultured in complete medium. Colonies were selected and passaged to increase the number of green fluorescent protein (GFP)-positive cells and plated on gelatin-coated coverslips. Cells after loading with the mitochondrial-O2.––sensitive fluorophore MitoSOX Red (Molecular Probes; 1.25 µM) were exposed to KO2, Tg, and DIDS as described above. In some experiments, MPMVECs were pretreated with BAPTA-AM (50 µM) for 30 min before KO2 application.
ClC-3 Knockdown
Confluent MPMVECs (5 x 106) were washed and placed in serum-free DMEM before transfection by electroporation with 250 pmol of either ClC-3 or negative control siRNA. To establish transfection efficiency, PMVECs were also transfected with Cy3-labeled GAPDH siRNA. Cells were then transferred to the appropriate culture vehicle and cultured in RPMI medium supplemented with 10% FBS, essential amino acids, endothelial cell growth supplement, and antibiotics. To confirm transfection, cells at 24 h after transfection were counterstained with the nuclear marker DAPI and images were acquired using MetaMorph software (Molecular Devices, Downingtown, PA) via epifluorescence microscopy (TE2000U, 10x objective; Nikon, Melville, NY). After 24 h, medium was replaced with standard growth medium and changed daily for an additional 48 h. Cells at 60 h after transfection were lysed and evaluated for ClC-3 mRNA by RT-PCR or imaged via confocal microscopy, respectively. Cells at 72 h after transfection were lysed and ClC-3 protein level was assessed by Western blotting using a rabbit polyclonal anti-ClC-3.
Total RNA Extraction and RT-PCR
Total RNA was prepared from wild-type and siRNA transfected MPMVECs using an RNeasy Mini Kit (Qiagen, Valencia, CA). The Transcriptor first-strand cDNA synthesis kit (Roche Applied Science, Indianapolis, IN) was used to reverse transcribe cDNA from 2 µg of RNA using both random hexamer and anchored-oligo(dT)18 primers. For ClC-3, the forward and reverse primers were GCGTGAGAACCGCGTTACT and GCTTTCAGGAGAGGTTACGT, respectively. For ClC-4, the forward and reverse primers were GATGGGCATTATTTTGAGAAG and CAGTAGCATGCGAATACCCC, respectively. For -actin, the forward and reverse primers were ATGGATGACGATATCGCTGC and CTTCTGACCCATACCCACCA, respectively. The PCR amplification profile consisted of an initial denaturation at 95°C for 2 min followed by 35 cycles of a 30-s denaturation at 95°C 30 s, annealing at 55°C for 30 s, and 1-min extension at 72°C, followed by a final 10-min extension step at 72°C using GoTaq DNA Polymerase (Promega, Madison, WI). PCR products were separated by electrophoresis on a 2% agarose/TBE gel and visualized by ethidium bromide staining.
Mitochondrial Membrane Potential
Cells cultured on coverslips were incubated with the cationic potentiometric fluorescent dye rhodamine 123 (25 µM) for 20 min at 37°C. After dye loading, the cells were washed and resuspended in DMEM. Images were recorded every 5 s for 5 min using the Bio-Rad Radiance 2000 imaging system with excitation at 488 nm. A decrease in mitochondrial membrane potential (
m) results in loss of rhodamine 123 from the mitochondria into the cytoplasm and the nucleus. Quantitation of the m change was determined by nuclear masking for fluorescence of all cells in the field. Treatment with DIDS and other agents was performed as described above.
Measurement of [Ca2+]i Mobilization
Endothelial cells adherent to 25-mm-diameter glass coverslips were loaded with the cytosolic Ca2+ indicator Fluo-4/AM (5 µM; Invitrogen, Carlsbad, CA) at room temperature for 30 min in extracellular medium (ECM) containing 121 mM NaCl, 5 mM NaHCO3, 10 mM Na-HEPES, 4.7 mM KCl, 1.2 mM KH2PO4, 1.2 mM MgSO4, 2 mM CaCl2, 10 mM glucose, and 2.0% bovine serum albumin (BSA), pH 7.4, in the presence of 100 µM sulfinpyrazone and 0.003% pluronic acid. After dye loading, the cells were washed and resuspended in the experimental imaging solution (ECM containing 0.25% BSA) and images recorded every 3 s at 488-nm excitation using the Bio-Rad Radiance 2000 imaging system.
Annexin V Imaging
To determine phosphatidylserine externalization as an indication of early apoptosis, cells were exposed to KO2 for 3 h and incubated with the conjugate annexin V Alexa-Fluor-488 (Molecular Probes) for 15 min in annexin V binding buffer. PI (0.5 µg/ml) was added 5 min before imaging. After treatment, annexin V– and PI-positive cells were excited at 488 and 568 nm, respectively, and were counted in 10 independent fields. The normally impermeable PI is internalized as the plasma membrane loses integrity. Positive PI staining indicates either late stage apoptosis or necrosis.
Data Analysis
Either nuclear (HE, rhodamine 123) or perinuclear (MitoSOX Red) masking of all cells in a given field was used to quantitate the cellular response using Spectralyzer (custom software provided by Paul Anderson, Thomas Jefferson University) image analysis software. Tracings indicate the mean fluorescence value of all cells in one field and are indicative of n independent experiments. Multiple experiments were normalized to baseline average and expressed as fold change. Data are expressed as mean ± SEM for n independent experiments.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Johnston et al., 1978). The bolus of O2.– caused rapid and transient HE oxidation in MPMVECs (Figure 1A), which was eliminated by pretreatment with DIDS (200 µM; Figure 1B). HPMVECs responded similarly to bolus O2.– addition (Figure 1C). Subsequent experiments with MPMVECs exposed to varied O2.– concentrations revealed a concentration-dependent response that was blocked either by SOD (2500 U/ml) or by DIDS (200 µM); catalase (1,000 U/ml) had relatively little effect (Figure 1D). Application of O2.– also resulted in a dip in fluorescence below baseline values after the transient peak (Figure 1B). Because this dip corresponded to brief gap formation in the endothelial monolayer as observed by simultaneous recording of differential interference contrast (DIC; data not shown), it most likely represents a transient alteration of focus. We postulate that the transient gaps in the endothelial monolayer are associated with intracellular Ca2+ release as described below. H2O2 (500 µM) added as a bolus had no effect on HE fluorescence (Figure 1D).
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). Mitochondrial-derived O2.– resulting from either FCCP or AA treatment resulted in a rapid and progressive increase in HE fluorescence (Figure 1E). The fluorescence spectrum at 494-nm excitation was similar after treatment with either extracellular O2.– or AA, suggesting the same HE oxidation product (Supplementary Figure 1). The present studies do not allow us to differentiate between oxyethidium and ethidium in these experiments although previous studies suggest oxyethidium as the major stable metabolite (Zhao et al., 2005).
O2.– Causes Rapid and Transient HE Oxidation in a Cell-free System
The reaction of HE with O2.– creates a stable product in a multistep process (Fink et al., 2004; Zhao et al., 2005). We therefore hypothesized that the HE fluorescence transient (Figure 1) may be an HE oxidation intermediate. A cell-free system was used to investigate the chemical nature of the transient response of HE to O2.– observed in PMVECs. The HE fluorescence changes were monitored after delivery of either a bolus of KO2 (200 µM) or a bolus of X/XO (50 mU/ml) delivered to HE (40 µM) dissolved in PBS (Figure 2A). Similar to findings with PMVECs, a rapid HE fluorescence transient was observed after KO2 application, whereas the X/XO O2.– generating system resulted in a progressive increase. HE fluorescence was unaltered after addition of DMSO vehicle (200 µl), KOH (200 µM), H2O2 (5 mM), or KO2 (100 µM) that had been predismutated into H2O2 by SOD (1000 U/ml). Increasing concentrations of O2.– correlated with the magnitude of both the initial peak and the stable postpeak HE fluorescence (Figure 2B).
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; Li and Shah, 2004). O2.– production was monitored by the increase in HE fluorescence after 1 h of agonist stimulation. Both Ang II and thrombin increased O2.– production in HPMVECs (Supplementary Figure 2 and Figure 4). Pretreatment with the NADPH oxidase inhibitor Apo (2 µM) or with DIDS (300 µM) decreased HE oxidation in Ang II–stimulated cells (Supplementary Figure 2 and Figure 4), compatible with Ang II–mediated extracellular O2.– generation via NADPH oxidase. Thrombin-stimulated O2.– release, on the other hand, was insensitive to Apo and DIDS, suggesting an intracellular source of O2.– for this agonist.
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5.5-fold higher for O2.– versus untreated (vehicle only) cells (Figure 5B). Treatment with the sarcoplasmic/endoplasmic reticulum Ca2+-ATPase inhibitor thapsigargin (Tg) in the absence of extracellular O2.– resulted in transient elevation of intracellular Ca2+ and also triggered mitochondrial ROS production. Chelation of intracellular Ca2+ by BAPTA abolished the increase in MitoSOX Red fluorescence in response to bolus O2.– (Figure 5B). This result provides evidence that mitochondrial ROS generation occurs as a result of intracellular Ca2+ mobilization (Madesh et al., 2005). These data indicate that inhibition of O2.– flux across the plasma membrane by DIDS or ablation of O2.– -triggered Ca2+ release by chelation prevents mitochondrial ROS production associated with extracellular O2.–.
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) and is inhibitable by DIDS (Hirotsu et al., 1999), we chose to assess whether knockdown of ClC-3 elicits a reduction in HE fluorescence similar to that demonstrated by DIDS. Electroporation efficiently delivered siRNA to MPMVECs as evidenced by microscopic evaluation of Cy3-labeled GAPDH siRNA (Figure 6A). siRNA sequences targeting exon 10 (sequence 1) and exons 2 and 3 (sequence 2) resulted in a marked decrease in ClC-3 mRNA expression by RT-PCR at an equivalent cycle amplification as compared with wild-type and negative control siRNA-transfected MPMVECs (Figure 6B). No differences in -actin or ClC-4 expression were observed, indicating specificity of the siRNA effect. A reduction in ClC-3 protein expression was also noted at 72 h after delivery of ClC-3 siRNA (Figure 6B, bottom panel). Both the rapid peak (Figure 6C) and the subsequent increased nuclear HE fluorescence at 20 min after O2.– exposure (Figure 6, D and E) were markedly inhibited in ClC-3 siRNA treated cells (sequence 1) compared with negative control–transfected cells. These siRNA knockdown experiments provide additional evidence that O2.– membrane flux is mediated by ClC-3 and are consistent with the effect of DIDS on O2.–-mediated HE oxidation.
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mm changes after addition of 5 µM O2.– (Figure 7, D and E). However, HPMVECs appeared to be more sensitive to O2.–, as addition of a 10 µM bolus produced irreversible m loss (data not shown). m changes were consistently delayed in comparison to the HE fluorescence transient. However, the biphasic phenomenon of m alteration is similar to that observed with HE after O2.– exposure (see Figures 2 and 7).
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m, we investigated the causal role of intracellular Ca2+ in m alterations in MPMVECs pretreated with Tg (2 µM). This pretreatment with Tg prevented mitochondrial depolarization after bolus addition of O2.– (Figure 7D) without having a direct effect on m (Supplementary Figure 3), indicating that the effect of increased extracellular O2.– on m requires Ca2+ derived from intracellular stores. The specificity of the Ca2+ effect was evaluated by measuring the responses to an uncoupler of mitochondrial respiration and to depolarization of the plasma membrane. Addition of the mitochondrial uncoupler FCCP (2 µM) facilitated irreversible m loss in contrast to the biphasic response to extracellular O2.–. To investigate a possible interaction between plasma membrane and mitochondrial membrane potentials, 20 mM KCl was added to rhodamine 123–loaded HPMVECs in order to partially depolarize the plasma membrane (Zhang et al., 2005). KCl addition had no effect on m (Figure 7E). This excludes a possible effect of this cation when added with O2.–.
Anion Channel Blockade Prevents O2.–-induced Apoptosis
Addition of O2.– as a single bolus led to a subsequent significant increase in the number of MPMVECs that stained positively for annexin V (Figure 8, A and B). Many of the annexin V–positive cells also stained positively for PI. There was no significant population of PI-positive but annexin V–negative cells. These results are compatible with early to later events of apoptosis of MPMVECs associated with the O2.– bolus.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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). However, specific oxidants may result in discrete effects on endothelial cells (Finkel, 2001; Devadas et al., 2002). Endogenous O2.– produced by NADPH oxidase has been implicated in cell proliferation (Milovanova et al., 2006), whereas phagocyte-derived O2.– is associated with endothelial apoptosis (Madesh et al., 2005). Although it is possible that the disparate functions of extracellular O2.– may be attributed to varying mechanisms including rapid dismutation to H2O2, modification of cell-surface proteins, or release of cytokines secondary to endothelial cell activation, we hypothesize a unique signaling cascade specific to the extracellular presence of this anion. The biological effects of extracellular ROS have been studied primarily by adding exogenous H2O2 to target cells (Wright et al., 1994) or by examining autocrine effects on the ROS-producing cells themselves (Thannickal and Fanburg, 2000). No studies, to our knowledge, have demonstrated a role for paracrine-derived O2.– in triggering intracellular ROS generation in a physiological/pathophysiological relevant context. In the present study, we demonstrate that extracellular O2.– can cross the cell membrane via ClC-3 and initiate an intracellular signaling cascade resulting in Ca2+-mediated O2.– production by the mitochondria. These results suggest an important role for extracellular O2.– in endothelial cell biology.
In this study, we investigated O2.– membrane flux by utilizing a previously unpublished property of the O2.–-sensitive fluorophore HE. The transient intracellular fluorescence peak associated with HE oxidation was O2.– concentration-dependent in both live cell and cell-free models. Specificity of this transient to O2.– is indicated by its inhibition with SOD, whereas catalase had no effect. Addition of KO2-treated HE to the extracellular milieu did not alter HE fluorescence (data not shown), excluding the possibility that HE oxidized outside the cell rapidly traverses the plasma membrane. Abrogation of the effect by DIDS suggests that this intracellular HE fluorescence transient results from membrane flux of O2.– through an anion channel. ClC-3 is the most abundant chloride channel in endothelial cells (Lamb et al., 1999) and knockout of this channel results in compensatory changes in cell membrane protein expression and function (Yamamoto-Mizuma et al., 2004). Selective knockdown of ClC-3 using siRNA resulted in a significant reduction in the HE transient similar to that observed by anion channel inhibition with DIDS. Therefore, we conclude that ClC-3 is the primary channel that supports transmembrane O2.– flux in endothelial cells.
After the HE transient with addition of O2.–, we observed a progressive increase in nuclear HE fluorescence that was blocked by DIDS and ClC-3 knockdown. It seems unlikely that this delayed response is due to the extracellular O2.– because of the expected short lifetime of O2.– in solution. A possibility for this finding is that extracellular O2.– triggered a secondary response in the cells leading to O2.– generation from a cellular source. Intracellular O2.– production by the mitochondrial inhibitor AA and the uncoupler FCCP resulted in progressive increase of nuclear HE fluorescence. This led us to hypothesize that the mitochondria may be a secondary source of O2.– after addition of O2.– to the extracellular medium. Measurement of nuclear HE fluorescence has been suggested as an indicator for O2.– derived from NADPH oxidase (Sun et al., 2005). However, O2.– generated by the mitochondria elicits a similar response (Becker et al., 1999). The failure of NADPH oxidase deficient cells to change the response and the use of the mitochondrial O2.– specific dye MitoSOX Red in the present experiments indicate that mitochondrial production of O2.– is primarily responsible for the progressive increase in nuclear HE fluorescence associated with extracellular O2.–. These results suggest that nuclear HE fluorescence associated with activation of NADPH oxidase and consequent extracellular O2.– generation may actually reflect mitochondrial-derived ROS resulting from intracellular Ca2+-mediated signaling.
Addition of Ang II or thrombin was used to initiate endogenous NADPH oxidase activity in endothelial cells in order to test whether mitochondrial O2.– production was activated by physiological levels of extracellular O2.–. We observed that Ang II triggered a significant increase in HE fluorescence that was blocked by both Apo and DIDS. Ang II–induced endothelial cell O2.– production has been linked to endothelial dysfunction and associated hypertension (Lassegue et al., 2001), and a link has been demonstrated between Ang II stimulated NADPH oxidase-derived O2.– and mitochondrial ROS production (Kimura et al., 2005). It has been proposed that ROS produced by mitochondria in endothelial cells serve an intracellular signaling function (Quintero et al., 2006). Oscillations in mitochondrial ROS production due to a localized production of ROS by a small number of mitochondria have provided evidence for mitochondrial-mediated signaling via ROS (Zorov et al., 2000; Aon et al., 2003). However, the possibility that extracellular ROS also could stimulate intracellular ROS production by the mitochondria has not been previously reported. The present study demonstrates that extracellular O2.– produced by NADPH oxidase can permeate the cell membrane to trigger intracellular (mitochondrial) ROS production.
Transmembrane O2.– flux has previously been shown in membranes highly enriched with anion channels, such as the erythrocyte (Lynch and Fridovich, 1978). However, under normal conditions, the diffusion distance of O2.– before spontaneous dismutation to H2O2 is estimated at 0.5 µm (Mikkelsen and Wardman, 2003). The rate of dismutation would be increased within the cell by cytosolic SOD (Fridovich, 1995). This precludes extracellular O2.– from traveling much beyond the plasma membrane to react with potential intracellular signaling proteins (Finkel, 2001). Nonetheless, we demonstrate that O2.–-mediated signaling can be attenuated by both molecular inhibition of ClC-3 and anion channel blockade by DIDS, indicating a discrete role for O2.– membrane flux in endothelial function. The question therefore arises as to the mechanism through which the short-lived O2.– anion leads to cell signaling. The experimental findings are that extracellular O2.– triggered rapid Ca2+ mobilization and that mitochondrial ROS production was preceded by Ca2+-dependent changes in m and was prevented by passive depletion of ER Ca2+ stores. These results are in agreement with studies using activated macrophages or the X/XO O2.–-generating system (Madesh et al., 2005). Loss of m in isolated mitochondria as a result of Ca2+ overload has been demonstrated previously (Galindo et al., 2003). Thus, we propose that the mechanism by which extracellular O2.– triggers mitochondrial O2.– production is through cell signaling secondary to Ca2+ release from intracellular stores. The present evidence supports this hypothesis, because mitochondrial O2.– generation occurred in association with increased intracellular Ca2+ after Tg treatment and was abolished by chelation of the increased Ca2+ mediated by extracellular O2.–. Based on our previous studies, O2.–-mediated Ca2+ release occurs via an inositol trisphosphate receptor-dependent mechanism (Madesh et al., 2005).
In summary, transmembrane O2.– flux occurs in PMVECs through ClC-3 channels and results in m alterations and mitochondrial O2.– production. This novel finding elucidates a potential mechanism by which extracellular O2.– is propagated to the intracellular milieu to trigger endothelial cell signaling or dysfunction associated with oxidative stress. We postulate that endothelial cell injury via paracrine O2.– signaling may represent a basis for pulmonary vascular remodeling.
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ACKNOWLEDGMENTS |
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Footnotes |
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The online version of this article contains supplemental material at MBC Online (http://www.molbiolcell.org). 
Address correspondence to: Aron B. Fisher (abf{at}mail.med.upenn.edu).
Abbreviations used: ROS, reactive oxygen species; SOD, superoxide dismutase; ClC-3, chloride channel-3; m, mitochondrial membrane potential; DIDS, 4,4'-diisothiocyanostilbene-2,2'-disulfonic acid; HE, hydroethidium; HPMVEC, human pulmonary microvascular endothelial cells; Ang II, angiotensin II; MPMVECs, murine pulmonary microvascular endothelial cells; Apo, apocynin; FCCP, carbonyl cyanide p[trifluoromethoxy]-phenyl-hydrazone; Tg, thapsigargin.
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