American Journal of Respiratory Cell and Molecular Biology

Lung capillary endothelial cells (ECs) are a critical target of oxygen toxicity and play a central role in the pathogenesis of hyperoxic lung injury. To determine mechanisms and time course of EC activation in normobaric hyperoxia, we measured endothelial concentration of reactive oxygen species (ROS) and cytosolic calcium ([Ca2+]i) by in situ imaging of 2′,7′-dichlorofluorescein (DCF) and fura 2 fluorescence, respectively, and translocation of the small GTPase Rac1 by immunofluorescence in isolated perfused rat lungs. Endothelial DCF fluorescence and [Ca2+]i increased continuously yet reversibly during a 90-min interval of hyperoxic ventilation with 70% O2, demonstrating progressive ROS generation and second messenger signaling. ROS formation increased exponentially with higher O2 concentrations. ROS and [Ca2+]i responses were blocked by the mitochondrial complex I inhibitor rotenone, whereas inhibitors of NAD(P)H oxidase and the intracellular Ca2+ chelator BAPTA predominantly attenuated the late phase of the hyperoxia-induced DCF fluorescence increase after > 30 min. Rac1 translocation in lung capillary ECs was barely detectable at normoxia but was prominent after 60 min of hyperoxia and could be blocked by rotenone and BAPTA. We conclude that hyperoxia induces ROS formation in lung capillary ECs, which initially originates from the mitochondrial electron transport chain but subsequently involves activation of NAD(P)H oxidase by endothelial [Ca2+]i signaling and Rac1 activation. Our findings demonstrate rapid activation of ECs by hyperoxia in situ and identify mechanisms that may be relevant in the initiation of hyperoxic lung injury.

Since its discovery in 1770 by Scheele and Priestley, oxygen has become one of the most effective, widely available, and cheap therapeutic agents. In a wide range of hypoxic and/or hypoxemic disorders, normobaric hyperoxia can improve tissue oxygenation and thus frequently provides life-saving benefits. Furthermore, hyperoxic ventilation can diminish the risk of surgical wound infections after colorectal resection (1, 2) and reduce ischemic brain damage in experimental stroke (3). However, exposure to increased oxygen concentrations for hours or days results in deleterious effects on lung function in humans (4) and ultimately in death from hyperoxic lung injury in laboratory animals (5). The morphologic changes in hyperoxic lung injury share many common features with other forms of acute lung injury in that an initial exudative phase, characterized by inflammation, atelectasis, and edema formation, is followed by a fibroproliferative phase with irreversible loss of respiratory function (6). Sequestration and infiltration of circulating neutrophils and platelets are early hallmarks of the inflammatory phase and indicate the activation of pulmonary microvascular endothelial cells (ECs) by hyperoxia (7). Early changes in EC ultrastructure (8) and increased microvascular leakage (9) identify lung ECs as a primary target in hyperoxic stress. The length of the initiation phase preceding changes in global lung function and morphology varies inversely with the concentration of oxygen (7) and has been reported to range between 14 and 30 h at O2 concentrations of 70% or higher in healthy human subjects (4, 10). Shorter episodes of normobaric hyperoxia are therefore generally considered as clinically safe. In contrast, in vitro data suggest that lung ECs respond to hyperoxia within 30–60 min (11, 12), and biochemical alterations may occur within seconds. Hence, EC activation and/or damage may significantly precede overt morphologic and/or functional changes and could initiate long-term effects on lung structure and function.

The pathogenesis of hyperoxic lung injury has been attributed to the generation of reactive oxygen species (ROS), including superoxide (), hydrogen peroxide (H2O2), and hydroxyl radical (HO•), and subsequent formation of potent oxidants such as peroxynitrite (ONOO) (13). These highly reactive intermediates cause cellular and subcellular damage by initiating oxidative and peroxidative chain reactions and serve as intracellular signaling molecules to elicit active cell responses. , in particular, seems to play a predominant pathophysiologic role in hyperoxic lung injury. Overexpression of any of the three mammalian isoenzymes of superoxide dismutase (SOD), which rapidly catalyze the dismutation of to H2O2, provides partial or complete protection from hyperoxic lung injury (1416). In contrast, mice lacking extracellular SOD are more sensitive to hyperoxia, showing an earlier onset of edema and reduced survival (17).

A variety of mechanisms have been proposed to contribute to enhanced production during hyperoxia. In lung homogenates and slices exposed to 100% O2, Crapo and coworkers detected an increased electron leak from the mitochondrial respiratory chain that results in production (18). In contrast, Parinandi and coworkers recently reported hyperoxia-induced ROS generation in cultured human pulmonary artery ECs by NAD(P)H oxidase (11). Although this enzyme was long considered to be a unique feature of phagocytes by which they can rapidly generate vast amounts of to kill invading pathogens, recent studies demonstrate its expression in endothelial and vascular smooth muscle cells (19). In these nonphagocytic cells, NAD(P)H oxidase-derived may act as intracellular signaling molecule and/or scavenger of NO (20).

In the intact pulmonary microcirculation, the onset of endothelial responses to hyperoxia and underlying cellular mechanisms are unclear. A difficulty in this understanding lies in the dynamic quantification of ROS formation in intact lungs during normobaric hyperoxia. Using our recently developed in situ fluorescence imaging techniques (21), we determined endothelial ROS formation during normobaric hyperoxia and underlying cellular signaling pathways in the isolated blood-perfused rat lung.

Animals

Male Sprague-Dawley rats (423 ± 17 g bw) were obtained from the local breeding facility of the academic institution. All animals received care in accordance with the “Guide for the Care and Use of Laboratory Animals” (NIH publication no. 86-23, rev. 1985). The study was approved by the animal care and use committees of the local government authorities.

Fluorescent Probes, Antibodies, and Drugs

The fluorescent dyes fura 2-AM, 2′,7′-dichlorodihydrofluorescein diacetate (H2DCF-DA), hydroethidine (HE; dihydroethidium), and HOECHST 33342 were purchased from Molecular Probes, Inc. (Eugene, OR). Rabbit anti-Rac 1 polyclonal antibody and rabbit anti-CD4 polyclonal antibody were purchased from Santa Cruz Biotechnology, Inc. (Santa Cruz, CA), and r-phycoerythrin-conjugated sheep anti-rabbit IgG were from DPC Bierman (Bad Neuheim, Germany). Other agents used were the flavoenzyme inhibitor diphenyleneiodonium (DPI), the serine protease inhibitor 4-(2-aminoethyl)-benzenesulfonyl fluoride (AEBSF), the mitochondrial complex I electron transport inhibitor rotenone, the NO synthase inhibitor Nω-Nitro-L-arginine methyl ester hydrochloride (l-NAME), hydrogen peroxide (H2O2, 30%) (all Sigma-Aldrich, Taufkirchen, Germany), the intracellular Ca2+ chelator 1,2-bis-(o-aminophenophenoxyethane)-N,N,N',N'-tetra-acetic acid tetra-(acetoxymethyl)-ester (BAPTA-AM), and the Ca2+ ionophore 4-bromo A-23187 (both Molecular Probes, Inc.). Unless stated otherwise, agents were infused in 2% dextran (70 kD) (Sigma-Aldrich), 1% FBS (Gemini BioProducts, Calabasas, CA) HEPES solution containing (in mmol/liter) 150 Na+, 5 K+, 1.5 Ca2+, and 20 HEPES (Serva, Heidelberg, Germany) at pH 7.4 and osmolarity of 295 mosM.

Lung Preparation

Experimental procedures have previously been described (21, 22). In brief, lungs excised from anesthetized Sprague-Dawley rats were continuously pump-perfused with 14 ml/min autologous heparinized (500 U) rat blood at 37°C. At baseline, lungs were constantly inflated with a gas mixture of 21% O2, 5% CO2, balance N2 at a positive airway pressure of 5 cm H2O. Left atrial pressure was adjusted to 3 cm H2O, yielding pulmonary artery pressures of 10 ± 1 cm H2O. Pulmonary artery pressure and left atrial pressure were continuously monitored and recorded (Recomed; Hellige, Freiburg, Germany). Lungs were positioned on a custom-built microscope stage and superfused with normal saline at 37°C to prevent drying. For local delivery of fluorescent probes and drugs to pulmonary microvessels in situ, a microcatheter (Ref. 800/110/100; SIMS Portex Ltd., Kent, UK) was advanced through the left atrium and wedged in a pulmonary vein draining a small capillary area on the lung surface.

In Situ Fluorescence Microscopy

Fluorophores loaded to rat lung capillary ECs were visualized on an Olympus BX50 upright microscope (Olympus, Hamburg, Germany). Light from a 75-watt xenon source was narrowed to a near monochromatic beam by a digitally controlled galvanometric scanner (Polychrome II; TILL Photonics, Martinsried, Germany). Fluorescence emission was collected through an apochromat objective (UAPO 40× W2/340; Olympus) and appropriate dichroic and emission filters (FT 425 and BP 505–530 or DCLP 500 and LP 515) by a CCD camera (Sensicam; PCO, Kelheim, Germany) and subjected to digital image analysis (TILLvisION 4.0; TILL Photonics). Single venular capillaries were viewed at a focal plane corresponding to maximum vessel diameter (17–25 μm).

Capillary Imaging and Analysis

Intracellular ROS concentrations in ECs were measured in situ using two intracellular fluorescent probes, H2DCF-DA and hydroethidine. H2DCF-DA (2 μmol/liter) was continuously infused into lung capillaries via the venous microcatheter as previously described (23). Image acquisition was started 30 min after the onset of fluorophore infusion. Capillaries were excited at 488 nm, and emission > 515 nm was recorded. HE (20 μmol/liter) was loaded to lung ECs in situ for 30 min, and fluorescence of hydroethidine and its oxidation products ethidium (E+) and hydroxyethidium (2-OH-E+), respectively, were measured at excitation/emission wavelengths of 360 nm/425 nm and 470 nm/ > 515 nm according to the respective spectra (24, 25). Our fura 2 ratio imaging technique for the quantification of intracellular calcium concentrations ([Ca2+]i) in ECs of lung venular capillaries has been described previously (22). In brief, membrane-permeant fura 2-AM (5 μmol/liter), which de-esterifies intracellularly to impermeant fura 2, was infused into lung capillaries for 20 min via the venous microcatheter. Fluorescence images at excitation wavelengths of 340, 360, and 380 nm were recorded, and endothelial [Ca2+]i was determined from the 340/380 ratio based on a Kd of 224 nmol/liter and appropriate calibration parameters (22). Fluorescence images were obtained at 5-s intervals and background corrected, with background determined in images captured before fluorophore loading. Capillary fluorescence was quantified in 4 μm2 areas along the capillary wall, representing single lung ECs. [Ca2+]i oscillations were analyzed for amplitude and frequency using fast Fourier transformation.

Rac1 Translocation Assay

Activation of endothelial NAD(P)H oxidase involves activation and translocation of the small GTPase Rac1 from the cytosol to the cell membrane. In isolated-perfused lungs, we monitored Rac1 translocation in ECs by adaptation of a previously established immunofluorescence assay (22). After in situ fixation with 3.7% paraformaldehyde and permeabilization of EC membranes by 0.5% Triton X-100, indirect immunofluorescence labeling by sequential infusion of primary anti– Rac-1 mAb (20 μg/ml) for 10 min and secondary rPE-conjugated mAb (100 μg/ml) for 1 min was performed. Membrane permeabilization permits cell loading with Abs and causes loss of unbound cytoplasmic molecules, such as nonactivated Rac1, when cells are washed by capillary infusion of HEPES solution. Hence, residual fluorescence indicated the presence of activated, membrane-bound Rac1. Counterstaining of EC nuclei in situ was performed by capillary infusion of HOECHST 33324 (5 μg/ml) followed by wash.

Experimental Protocols
Ventilation.

After baseline recordings were made, lungs were ventilated with inspiratory oxygen fractions (FiO2) of 0.21 (normoxia) or 0.7 (hyperoxia), 5% CO2, balance N2 for 90 min, after which normoxic ventilation was re-established for another 45 min. For image acquisition, airway pressure was kept constant at 5 cm H2O for 150 s, allowing for acquisition of 30 images in 5-s intervals. In between these imaging periods, airway pressure was varied cyclically between 2 and 10 cm H2O for 30 s to allow for adequate alveolar gas mixture and exchange.

Dose response.

To determine ROS formation at different FiO2, lungs were ventilated with 21%, 50%, 70%, and 95% O2; 5% CO2; balance N2 for 90 min.

Inhibitor studies.

Inhibitors were added to the perfusate after baseline recordings, 15 min before the start of hyperoxic ventilation. Mitochondrial electron transport was blocked by the complex I inhibitor rotenone (1 μmol/liter) and NO synthase by l-NAME (100 μmol/liter). For inhibition of NAD(P)H oxidase, the flavoenzyme inhibitor DPI (10 μmol/liter) or AEBSF (300 μmol/liter), which interferes with the multicomponental assembly of the NAD(P)H oxidase complex (26), was applied. Endothelial Ca2+ signaling was blocked by infusion of the intracellular Ca2+-chelator BAPTA, AM (40 μmol/liter in Ca2+-free HEPES solution).

Blood cell-free perfusion.

Infusion of Ca2+-rich HEPES solution was started 15 min before hyperoxic ventilation.

[Ca2+]i elevation.

Lungs were infused with HEPES solution containing 200 nmol/liter free Ca2+ in the absence or presence of the Ca2+ ionophore 4-bromo A23187 (5 μmol/liter).

Statistics

All data are mean ± SEM. For statistical analysis, imaging data acquired over 15-min intervals were averaged. Values of different groups were compared by Kruskal Wallis and Mann-Whitney U-Test. Spearman's coefficient of correlation (rs) was calculated to test for correlation of parameters. Linear and nonlinear regression analyses were performed (SigmaPlot; Jandel Scientific, San Rafael, CA). Statistical significance was assumed at P < 0.05.

For in situ determination of endothelial ROS production, lung venular capillaries were continuously infused with membrane-permeant H2DCF-DA, which de-esterifies intracellularly to H2DCF, the substrate that is oxidized to fluorescent DCF by ROS. Images of subpleural venular capillaries revealed DCF fluorescence in the capillary wall but not in the pericapillary space (Figure 1A). The fluorescent cells stained positive for the endothelial marker Alexa Fluor 488 AcLDL, confirming their endothelial phenotype (n = 4). Under physiologic conditions, the oxidation of H2DCF to DCF is irreversible. Due to the rapid leakage of the dye from ECs (27), continuous capillary infusion of H2DCF-DA can establish a steady state of dye delivery and removal that allows for near-online monitoring of endothelial ROS production (28, 29). Accordingly, endothelial fluorescence was stable during continuous dye infusion but decreased reversibly to background levels upon discontinuation of dye delivery (Figure 1B). Capillary infusion of a single bolus of H2O2 rapidly increased endothelial DCF fluorescence, which returned to background levels within minutes after washout of the bolus (Figure 1C). H2O2-induced fluorescence increase occurred in a concentration-dependent manner (Figure 1D) and was repeatable throughout experiments at normoxic and hyperoxic conditions (data not shown). Continuous imaging of DCF in 5-s intervals did not alter endothelial fluorescence over the time course of experiments, ruling out illumination-induced autoxidation or photobleaching of the dye.

Although fluorescence of H2DCF-loaded ECs was relatively low at baseline, capillaries became intensely fluorescent during hyperoxic ventilation (Figure 2A). Fluorescence increased in all ECs of the imaged field, indicating a homogeneous endothelial response to hyperoxia in lung capillaries. The slow but progressive increase of fluorescence over the 90-min hyperoxic interval indicated that ROS production did not result from spontaneous autoxidation of cellular molecules at high oxygen partial pressure but involved enzymatic reactions and active cellular responses (Figure 2B). Enhanced DCF fluorescence was evident within < 30 min, and its intensity increased continuously during the hyperoxic interval, demonstrating a time-dependent amplification of ROS production. After return to normoxic conditions, the hyperoxia-induced response was rapidly reversible, and capillary fluorescence decreased to baseline values within 30 min. Furthermore, DCF fluorescence was not attributable to intermittent cyclic ventilation, which was applied to warrant the admixture of hyperoxic gas, or to effects of illumination because a similar ventilation and imaging regimen performed at normoxia did not increase fluorescence (Figure 2B).

To establish a dose–response relationship between normobaric hyperoxia and endothelial ROS formation, we measured endothelial DCF fluorescence after 90 min of ventilation at different inspiratory oxygen fractions. Blood gas analysis in the perfusate confirmed adequate alveolar admixture and gas exchange (data not shown). DCF fluorescence increased progressively with higher FiO2, indicating an increased ROS formation at higher oxygen fractions that is reflected by an exponential curve fit described by the equation y = 84.7 e0.023x, with x representing FiO2 and y representing DCF fluorescence (Figure 2C).

Hyperoxia-induced ROS formation in lung capillary ECs was further verified by a second fluorophore, HE. HE readily permeates into ECs and yields blue cytosolic fluorescence upon ultraviolet excitation (Figure 3A). In the presence of , HE is oxidized to red fluorescent E+ and orange fluorescent 2-OH-E+, which are compartmentalized to the nucleus (Figures 3B and 3C) due to intercalation into DNA (25, 30).

After endothelial loading, HE fluorescence decreased at a slow rate, whereas the combined fluorescence of its oxidation products E+ and 2-OH-E+ increased, indicating oxidation of HE by basal ROS production (Figure 3G). During 90 min of hyperoxia, the rate of decrease in HE fluorescence and the increase in fluorescence of its oxidation products accelerated markedly (Figures 3D–3F), consistent with the progressive generation of under these conditions, but returned to rates similar to baseline within 20 min after return to normoxia (Figure 3G). In normoxic controls, fluorescence of HE and its oxidation products changed linearly at baseline rates over the observation period.

We investigated mechanisms underlying endothelial ROS formation in hyperoxia by inhibiting potentially involved cellular pathways. The mitochondrial inhibitor rotenone, which blocks electron transport at complex I, significantly reduced baseline DCF fluorescence and completely blocked the hyperoxia- induced fluorescence increase (Figure 4A). These findings suggest that baseline ROS levels and hyperoxia-induced ROS were, at least in part, of mitochondrial origin. Attenuation of the fluorescence increase was also achieved by DPI (Figure 4B), an inhibitor of flavoprotein oxidoreductases that blocks NAD(P)H oxidases, nitric oxide synthases, and xanthine oxidase. Although DPI may also block mitochondrial complex I and thus reduce (31) or even increase (32) mitochondrial production, it does not seem to interfere with mitochondrial electron transport in situ at the given concentrations (28). In accordance with this notion and unlike rotenone, DPI did not reduce baseline DCF fluorescence in lung capillaries. The fluorescence response to hyperoxia was also attenuated by AEBSF (Figure 4C), a serine protease inhibitor that interferes with the assembly of the multicomponental NAD(P)H oxidase complex (26). In contrast, inhibition of endothelial nitric oxide synthase (eNOS), which is also sensitive to DPI, by l-NAME had no effect, demonstrating that eNOS products were not involved in the fluorescence response (n = 5, data not shown). The effects of DPI and AEBSF were virtually identical in that neither inhibitor prevented the initial increase of DCF fluorescence during the first 30 min of the hyperoxic interval but significantly attenuated the subsequent progressive increase in fluorescence. This similarity suggests an involvement of NAD(P)H oxidase that seems to contribute in particular to progressive ROS production after more than 30 min of hyperoxia.

ECs express the NAD(P)H oxidase isoform Nox2 (gp91phox) and several of its regulatory subunits, such as p47phox and p67phox (11). Agonist-induced NAD(P)H oxidase-dependent generation and downstream effects critically depend upon GTP binding and membrane translocation of Rac1 in ECs (33). After 60 min of hyperoxic ventilation, immunofluorescence of membrane-bound Rac1 was detectable in lung capillary walls (Figure 5A). Counterstaining for endothelial nuclei (Figure 5B) and merged images (Figure 5C) revealed Rac1 translocation in all ECs of the imaged capillary. After 60 min of normoxic ventilation, very little Rac1 staining was detected (Figure 5D), although nuclear staining was positive (Figures 5E and 5F), indicating the absence of a Rac1 preassembly with the membrane-bound NAD(P)H oxidase subunits. Taken together, these findings suggest that, in addition to mitochondrial electron transport, Rac1-dependent activation of NAD(P)H oxidase contributed to hyperoxia-induced ROS production in lung capillary ECs. The fact that the portion of ROS production that was inhibitable by DPI and AEBSF was completely blocked by rotenone suggests that NAD(P)H oxidase activation occurred downstream of mitochondrial ROS production. Consistent with this notion, hyperoxia-induced immunofluorescence of translocated Rac1 was reduced to normoxic baseline levels in the presence of rotenone (Figure 5G). Translocation of Rac1 was also blocked by AEBSF, which is consistent with its proposed mode of action as inhibitor of NAD(P)H oxidase complex assembly (26), and by the intracellular calcium chelator BAPTA, indicating requirement for endothelial [Ca2+]i in this context (Figure 5G).

[Ca2+]i signaling may regulate endothelial NAD(P)H oxidase by triggering the translocation of Rac (34), via Ca2+-dependent PKC-isoforms and subsequent phosphorylation of the p47phox subunit (35), or by activation of Ca2+-sensitive gp91phox homologs such as NOX5 or DUOX1/2 (36, 37). Using the fura 2 ratio imaging technique, we determined that hyperoxia induced a rise in mean endothelial [Ca2+]i in lung capillaries (Figure 6A). This response increased with the duration of hyperoxic ventilation and was associated with a concomitant increase in the amplitude of endothelial [Ca2+]i oscillations, whereas oscillation frequency did not change from baseline (Figure 6B). Return to normoxia re-established the baseline [Ca2+]i profile within 30 min. In lungs ventilated at normoxia, no [Ca2+]i signal other than baseline [Ca2+]i oscillations was detected, demonstrating that the endothelial [Ca2+]i signal was in response to hyperoxia (Figure 6C).

Plotting mean endothelial [Ca2+]i, determined in 15-min intervals over the experimental time course, against capillary DCF fluorescence, which was measured at similar time points but in a separate set of experiments, revealed a linear correlation between both responses (Figure 6D). This finding demonstrates a close temporal and, presumably, functional relationship between both endothelial responses. To differentiate cause from effect, we infused the membrane-permeable intracellular Ca2+ chelator BAPTA-AM into lung capillaries for 15 min before hyperoxic ventilation. In a separate set of fura 2 imaging experiments (n = 5), we verified that [Ca2+]i signaling was completely blocked in BAPTA-treated ECs (data not shown). In capillaries infused with H2DCF-DA, BAPTA did not affect baseline endothelial fluorescence and, unlike rotenone, did not prevent the fluorescence increase during the initial 30 min of hyperoxia (Figure 6E). However, BAPTA effectively blocked the subsequent progressive increase in fluorescence, indicating that the late component of ROS production was Ca2+ dependent. Therefore, BAPTA elicited a similar pattern of inhibition as DPI and AEBSF (Figure 6E), indicating that hyperoxia-induced ROS production by NAD(P)H oxidase is regulated by endothelial [Ca2+]i.

To test whether endothelial Ca2+ fluxes of the observed magnitude can activate ROS formation by NAD(P)H oxidase in situ, we infused lungs with HEPES solution containing 200 nmol/liter free Ca2+ and rotenone to block potential Ca2+ effects on mitochondrial ROS production (38). Addition of the Ca2+ ionophore 4-bromo A23187 increased endothelial [Ca2+]i from 94.5 ± 4.5 to 186.3 ± 6.8 nmol/liter, as demonstrated by fura 2 ratio imaging (P < 0.05, n = 4) (data not shown), and increased DCF fluorescence 4-fold within 15 min, verifying that Ca2+ fluxes can induce ROS formation in lung capillary ECs (Figure 6F). The increase in DCF fluorescence was blocked by DPI, indicating that ROS were generated by activated NAD(P)H oxidase. Because mitochondrial ROS production may be a potential trigger of [Ca2+]i responses, we determined the effect of the mitochondrial inhibitor rotenone on the hyperoxia-induced [Ca2+]i increase in ECs. Rotenone did not change the baseline endothelial [Ca2+]i profile, but it completely blocked hyperoxia-induced increases in mean endothelial [Ca2+]i (Figure 7A) and oscillation amplitude (Figure 7B). To characterize the potential of ROS to induce [Ca2+]i signals in lung ECs, we infused capillaries with 100 μmol/liter H2O2 for 15 min. H2O2 caused a rapid increase in mean endothelial [Ca2+]i and amplified the amplitude of [Ca2+]i oscillations (Figure 7C). With subsequent washout of H2O2, [Ca2+]i returned to baseline profile within 15 min. Hence, H2O2 elicited a [Ca2+]i signal that had a profile similar to the response to hyperoxia. The NAD(P)H oxidase inhibitors DPI and AEBSF did not block the early [Ca2+]i response within the initial 30 min of hyperoxia, but both inhibitors significantly attenuated the subsequent progressive [Ca2+]i increase (Figure 7D). The analogy of this inhibition pattern with the effects of DPI and AEBSF on ROS production is consistent with the notion that NAD(P)H oxidase–derived ROS production is predominantly confined to the late phase (> 30 min) of the hyperoxic response and, accordingly, contributes only to the late [Ca2+]i signal. Microvessel perfusion with HEPES buffer instead of blood did not diminish this [Ca2+]i increase, demonstrating that NAD(P)H oxidase activity was in the vessel wall and not in circulating blood cells (n = 4) (data not shown).

In normobaric hyperoxia, we determined the production of ROS in pulmonary ECs and identified underlying signaling mechanisms. By in situ imaging of lung endothelial DCF fluorescence, ROS production was detectable within < 30 min of hyperoxia and increased progressively with time. Hyperoxia-induced formation of superoxide was verified by oxidation of HE to E+ and increased exponentially with higher FiO2. ROS production was triggered by a sequence of events that originated from the mitochondrial respiratory chain, as evident from complete inhibition by rotenone. Using fura 2 ratio imaging, we identified an endothelial [Ca2+]i response to hyperoxia that was blocked by rotenone, indicating its induction by mitochondrial ROS. In conjunction, we used an immunofluorescence assay to demonstrate hyperoxia-induced translocation of Rac1, which was blocked by rotenone and BAPTA and thus occurred downstream of the [Ca2+]i signal. After > 30 min, production of endothelial ROS was attenuated by DPI and AEBSF, indicating their generation by NAD(P)H oxidase, which presumably was activated by Ca2+-dependent Rac1 translocation. To our knowledge, this is the first report linking ROS from the mitochondrial electron transport chain to ROS production by NAD(P)H oxidase. Data demonstrating that DPI and AEBSF attenuated the endothelial [Ca2+]i response to hyperoxia support the notion of a positive feedback mechanism with reciprocal amplification of [Ca2+]i signaling and ROS generation by NAD(P)H oxidase. These findings provide new insights into the pathophysiology of hyperoxic lung injury by identification of a signaling cascade, which involves mitochondrial electron transport, endothelial [Ca2+]i, Rac1, and NAD(P)H oxidase and results in progressive ROS generation.

Methodologic Considerations

To establish normoxic and hyperoxic conditions, isolated perfused lungs were ventilated with 21% O2 or 70% O2, respectively. Between periods of image acquisition, cyclic lung ventilation was required to establish stable hyperoxic conditions. However, cyclic ventilation itself did not contribute to endothelial ROS generation or [Ca2+]i signaling as confirmed in normoxic control experiments. For the same reason, the effects of heparin, which was used as anticoagulant, can be excluded as mechanism underlying the detected endothelial responses. Because Pco2 and pH were constant throughout experiments, differences between hyperoxic and normoxic ventilation can be directly attributed to differences in Po2.

Although endothelial [Ca2+]i signaling was measured using our previously described fura 2 ratio imaging technique (22), ROS production was determined by in situ imaging of two intracellular fluorescent probes, DCF and HE. H2DCF-DA readily enters the cells where esterases cleave the acetate group, trapping the nonfluorescent H2DCF inside. Colocalization with an endothelial marker confirmed the endothelial phenotype of H2DCF-loaded microvascular cells, consistent with the notion that lung microvessels of diameters < 30 μm lack smooth muscle cells and pericytes. Oxidation of H2DCF by ROS yields the fluorescent product DCF. Therefore, DCF fluorescence provides a quantitative measure for ROS, as demonstrated by its linear response to different H2O2 concentrations. However, intracellular retention of DCF is poor, and the dye leaks from ECs within minutes (27). Therefore, we infused H2DCF-DA continuously into lung capillaries to establish a steady state between dye delivery and extrusion. This approach facilitated near-online monitoring of ROS production because endothelial fluorescence was stable at baseline, decreased rapidly to background levels upon discontinuation of dye delivery and returned to baseline intensity upon reinitiation of dye infusion and because bolus injections of H2O2 evoked transient and repeatable fluorescence increases in a concentration-dependent manner. Fluorescence responses to discontinuation of dye infusion or H2O2 challenge were conserved throughout the experimental protocols, demonstrating that hyperoxic conditions or applied inhibitors did not influence the measurement.

HE, the second fluorescent probe applied for detection of intracellular ROS, is uncharged and readily permeates the cell membrane (39). Oxidation of HE by , but not by O2, H2O2, or hydroxyl radicals yields the charged dye E+, which is trapped inside the cell where it binds to DNA (30, 40). Recently it was reported that HE may also react with to form fluorescent 2-OH-E+, which exhibits a left-shifted emission spectrum as compared with E+ upon excitation with blue light (25, 41). Therefore, a long-pass emission filter with a low cut-off was applied to allow for detection of both fluorescent products.

ROS Production by the Mitochondrial Electron Transport Chain

Hyperoxic ventilation for 90 min increased endothelial DCF fluorescence significantly yet reversibly, demonstrating that a rather short hyperoxic interval suffices to induce active cellular responses in lung capillaries. Because the fluorescence increase was conserved during buffer perfusion (i.e., in the absence of circulating blood cells), ROS production occurred in the alveolo-capillary wall. ROS generation in less than 90 min was confirmed by a marked decrease in HE fluorescence and a mirror-image increase in the combined fluorescence of its oxidation products, E+ and 2-OH-E+, suggesting as a major intracellular oxidant formed in hyperoxia. Capillary ECs were identified as the source of hyperoxia-induced ROS production because the response was blocked by inhibitors infused via the capillary route, which under absorptive conditions target the endothelial but not the epithelial cell layer (22). Contribution of NO production to the fluorescence signal was ruled out by pretreatment with l-NAME, which did not attenuate the DCF response.

Inhibition by rotenone demonstrates that the hyperoxia- induced increase in DCF fluorescence depends upon mitochondrial electron transport. The mitochondrial respiratory chain is composed of a series of electron carriers in spatial arrangement according to their redox potentials and is the main source of ROS during normal metabolism because ∼ 2–4% of electron flow through the electron transport chain results in only partial reduction of O2, yielding (42). This notion of a physiologic “electron leak” is consistent with basal ROS production in lung ECs reflected by a slow rate of HE oxidation at baseline (24). This basal ROS production was largely derived from mitochondria because inhibition of electron transport at complex I by rotenone reduced baseline DCF fluorescence by more than 50%.

Hyperoxic ventilation increased formation by the respiratory chain, which was equivalent to the fraction of DCF fluorescence that was insensitive to the NAD(P)H oxidase inhibitors DPI or AEBSF but blocked by rotenone. This DPI-insensitive fluorescence was constant throughout the hyperoxic interval but rapidly dropped to baseline values upon return to normoxia. This pattern is characteristic for a stoichiometric effect of oxygen tension on production by the respiratory chain, which is primarily controlled by mass action (43). Oxygen-dependent production from the electron transport chain has previously been detected in vitro in rat lung slices and isolated mitochondria (18), in cultured sheep pulmonary microvascular ECs (12), and in submitochondrial membranes (44). To our knowledge, this is the first evidence demonstrating hyperoxia-induced mitochondrial ROS production in ECs of intact lung microvessels in situ.

Endothelial Calcium Signaling

Although the mitochondrial respiratory chain was the predominant initial source of ROS and its inhibition by rotenone blocked all subsequent cellular events, additional enzymatic sources sensitive to DPI and AEBSF contributed to hyperoxia-induced ROS formation. Activation of these mechanisms depended critically on mitochondrial ROS and endothelial [Ca2+]i signaling because rotenone and Ca2+ chelation by BAPTA blocked DPI-sensitive ROS production.

Hyperoxia induced a characteristic endothelial [Ca2+]i response that consisted of an increase in mean endothelial [Ca2+]i and in [Ca2+]i oscillation amplitude. These [Ca2+]i transients occurred distal to mitochondrial ROS generation because they were effectively blocked by rotenone and could be mimicked by direct stimulation with H2O2. Direct interference of rotenone with endothelial [Ca2+]i signaling can be ruled out in this model because rotenone does not block endothelial [Ca2+]i transients induced by arachidonate (29). Mitochondrial ROS have been shown to trigger [Ca2+]i increases and evoke Ca2+ sparks in vascular smooth muscle cells, but the underlying molecular mechanisms are poorly understood (45, 46). Potential pathways involve Ca2+ release from the endoplasmic reticulum because exogenous H2O2 or the –generating system xanthine oxidase/hypoxanthine activate phospholipase C (47), enhance formation of inositol-1,4,5-trisphosphate (IP3), increase the sensitivity of the endoplasmic reticulum Ca2+ store to IP3 (48), and activate ryanodine-sensitive Ca2+ release channels (49). Ca2+ signals may also originate from mitochondria due to mitochondrial membrane depolarization and subsequent nonspecific Ca2+ leak (50) or by specific NAD+-linked Ca2+ release (51) or may reflect influx of extracellular Ca2+ via redox-sensitive ion channels (52). Further studies are required to dissect the contribution of these potential mechanisms to endothelial [Ca2+]i signals in hyperoxia. Here, the detected endothelial [Ca2+]i transients link mitochondrial electron transport to nonmitochondrial, DPI-sensitive ROS production. Elevation of [Ca2+]i stimulates Rac activation via protein kinase C and induces the translocation of cytosolic Rac to the plasma membrane (34). By immunofluorescence assay, we show that hyperoxia induced membrane translocation of Rac1, which was blocked by BAPTA and hence was Ca2+ dependent.

Activation of NAD(P)H-Oxidase

In ECs, GTP-binding and translocation of Rac1 can activate NAD(P)H oxidase and thus stimulate ROS production. Overexpression of a dominant negative form of Rac1 abrogates NAD(P)H oxidase–dependent endothelial responses, such as the expression of mitogen-activated protein kinase phosphatase-1, in response to atrial natriuretic peptide (23). In hyperoxia, AEBSF inhibited translocation of Rac1 and attenuated DCF fluorescence to a similar degree as the NAD(P)H oxidase inhibitor DPI, indicating a contribution of Rac1 and NAD(P)H oxidase to endothelial ROS formation. Activation of NAD(P)H oxidase was Ca2+ dependent because Ca2+ chelation blocked DPI-sensitive ROS production. This is consistent with the notion that Ca2+-dependent Rac1 translocation triggered NAD(P)H oxidase activation in hyperoxia.

Nonphagocytic cells express several members of the large family of NAD(P)H oxidases, and expression of gp91phox (NOX2), NOX4, and NOX5 has been demonstrated in ECs (11, 53, 54). In cultured human pulmonary artery ECs, Parinandi and coworkers recently reported hyperoxia-induced production that was inhibited by transfection with a p22phox antisense plasmid, indicating involvement of NOX2 rather than NOX4. In ECs, preassembly of the Phox regulatory proteins p47phox, p67phox, and p40phox with gp91phox and p22phox has been proposed (19), leaving Rac1 translocation as a major mechanism of post-translational NOX2 regulation. On the other hand, Ago and coworkers recently suggested that the major catalytic site of endothelial NAD(P)H oxidases is NOX4, which may also assemble with p22phox (55). Dominant negative Rac1 inhibits NOX4-dependent ROS generation and downstream stimulation of extracellular signal-regulated protein kinase 1/2 in response to angiotensin II (56). NOX2 and NOX4 can be regulated by Rac1 and thus potentially by [Ca2+]i, whereas NOX5 can be directly activated by a Ca2+-induced conformational change (36). Inhibitors of NAD(P)H oxidase also attenuated the endothelial [Ca2+]i response to hyperoxia, demonstrating that not only mitochondrial ROS, but also NAD(P)H oxidase-derived ROS can stimulate endothelial [Ca2+]i transients. In human aortic ECs, NAD(P)H oxidase activation was shown to mediate histamine-induced [Ca2+]i oscillations (20) and to increase the sensitivity of intracellular Ca2+ release to IP3 (48). NAD(P)H oxidase–triggered [Ca2+]i transients may establish a positive feedback mechanism that could account for the close correlation between endothelial [Ca2+]i and DCF fluorescence in hyperoxia and underlie the progressive increase of both responses during the hyperoxic interval. Positive feedback between [Ca2+]i signaling and ROS generation may result in a vicious circle amplifying cell activation and oxidative stress and initiating and/or promoting lung microvascular injury.

Clinical Significance

The clinical relevance of our findings relates to the controversial question of whether hyperoxic intervals of only a few hours may result in or contribute to deleterious lung vascular effects. Although increased generation of ROS is evident in lung ECs in vitro within 30–60 min of hyperoxia (11, 12), clinical use of normobaric hyperoxia for several hours is frequently considered harmless or even recommended (e.g., to reduce the risk of postsurgical wound infections) (1, 2). In humans, the first respiratory symptoms have been reported after 6 h of oxygen exposure (57), and ultrastructural alterations such as EC swelling are evident within 14 h at 70% O2 (10). In rats, animals die within 60–72 h of exposure to 100% O2, whereas an FiO2 of 0.85 is sublethal but may cause platelet accumulation within 3 d and increase lung weight within 5 d of exposure (5, 7). In isolated perfused rat lungs, confocal microscopy revealed microvascular expression of the adhesion molecule ICAM-1 and leukocyte sequestration in lung capillaries after 48 h of exposure to 90% O2 (58). Platelets and leukocytes may accumulate even earlier in hyperoxia because the reported data were measured at the first investigated time points in these studies. The onset of underlying cellular responses can be expected to further precede these proinflammatory effects, but have not been characterized in vivo or in situ.

The severity of hyperoxic lung injury is time- and dose dependent (7). As compared with 95% O2, an FiO2 of 0.7 results in less ROS formation but does not protect from lung injury when applied over longer periods because rats exposed to 60% O2 for 7 d show reduced lung compliance and perivascular edema formation (59), and 14-d exposure to 60% O2 causes low-grade epithelial injury and interstitial fibrosis in baboons (60).

The findings of our study may link early cellular activation in vitro to the delayed onset of lung injury in vivo. In isolated lungs ventilated at 70% O2, we detected endothelial activation within 30 min, which may contribute to the infiltration of inflammatory cells because endothelial [Ca2+]i and ROS signaling can initiate the expression of vascular adhesion molecules (e.g., P-selectin) (28, 61). These early endothelial effects themselves do not constitute an injury because endothelial responses were fully reversible after 90 min of hyperoxia. However, their progression with time may initiate functional and structural changes if hyperoxia persists.

The authors thank Prof. Dr. med. Dr. h.c. mult. K. Messmer for helpful comments and Dr. F. Ringel for technical support.

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Correspondence and requests for reprints should be addressed to Prof. Dr. Wolfgang M. Kuebler, Institute of Physiology Charité – Universitätsmedizin Berlin, Campus Benjamin Franklin, Arnimallee 22, 14195 Berlin, Germany. E-mail:

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