Essential role of complex II of the respiratory chain in hypoxia-induced ROS generation in the pulmonary vasculature

Renate Paddenberg1, Barat Ishaq1, Anna Goldenberg1, Petra Faulhammer1, Frank Rose2, Norbert Weissmann2, Ruediger C. Braun-Dullaeus2, and Wolfgang Kummer1

1 Institute of Anatomy and Cell Biology, and 2 Department of Internal Medicine, Justus Liebig University, 35385 Giessen, Germany


    ABSTRACT
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

In the pulmonary vasculature, the mechanisms responsible for oxygen sensing and the initiation of hypoxia-induced vasoconstriction and vascular remodeling are still unclear. Nitric oxide (NO) and reactive oxygen species (ROS) are discussed as early mediators of the hypoxic response. Here, we describe a quantitative analysis of NO- and ROS-producing cells within the vascular walls of murine lung sections cultured at normoxia or hypoxia. Whereas the number of NO-producing cells was not changed by hypoxia, the number of ROS-generating cells was significantly increased. Addition of specific inhibitors revealed that mitochondria were the source of ROS. The participation of the individual mitochondrial complexes differed in normoxic and hypoxic ROS generation. Whereas normoxic ROS production required complexes I and III, hypoxic ROS generation additionally demanded complex II. Histochemically demonstrable succinate dehydrogenase activity of complex II in the arterial wall decreased during hypoxia. Inhibition of the reversed enzymatic reaction, i.e., fumarate reductase, by application of succinate, specifically abolished hypoxic, but not normoxic, ROS generation. Thus complex II plays an essential role in hypoxic ROS production. Presumably, its catalytic activity switches from succinate dehydrogenase to fumarate reductase at reduced oxygen tension, thereby modulating the directionality of the electron flow.

lung slices; nitric oxide; reactive oxygen species; mitochondria; electron transport chain


    INTRODUCTION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

IN THE LUNG, REDUCED OXYGEN TENSION induces vasoconstriction to match perfusion to ventilation. Sustained generalized alveolar hypoxia results in vascular remodeling, leading to pulmonary hypertension and, consecutively, to cor pulmonale. Up to now, the molecular mechanisms of oxygen sensing and subsequent intracellular signaling pathways leading to vasoconstriction and vascular remodeling are still under debate. Two candidates for early mediators of the hypoxic response are nitric oxide (NO) (26) and reactive oxygen species (ROS) (31).

Studies on hypoxia-regulated NO production have provided conflicting data. In chronically hypoxic rats, upregulation of the endothelial isoform of nitric oxide synthase (eNOS) in small pulmonary vessels correlates with the onset of pulmonary vascular remodeling (42). In agreement with these findings, Quinlan and coworkers (26) have shown that eNOS-deficient mice exhibit reduced vascular proliferation and muscularization after exposure to hypoxia. In contrast, Fagan et al. (11) found in hypoxic mice with targeted disruption of the eNOS gene an increase in the number of partially and fully muscularized pulmonary arteries and a marked hypoxic pulmonary hypertension.

In addition to or instead of NO, ROS such as superoxide anion (O<UP><SUB>2</SUB><SUP>−</SUP></UP>·), hydrogen peroxide (H2O2), and hydroxyl radical (·OH) may function as intracellular second messengers to initiate or modulate signaling pathways leading to vasoconstriction, cell growth, and differentiation (30). In isolated perfused rat lungs, hypoxia induced a decrease in ROS production, as measured by the light-emitting superoxide trap lucigenin (1). Exposure of cultured porcine pulmonary artery endothelial cells to hypoxia results in a significant reduction of H2O2 formation (43). In contrast, various other cell types, including pulmonary artery myocytes (38), Hep 3B cells (7), and cardiomyocytes (10) respond to hypoxia with increased ROS production. Beyond that, conflicting results about the identity of the oxygen sensor and the cellular source of ROS generation have been presented. For example, a role as oxygen sensor is discussed for nonphagocytic NAD(P)H oxidases (20, 33, 39) and the mitochondrial electron transport chain (7, 8). Among those favoring the mitochondrial hypothesis, various groups suggest different complexes of the respiratory chain to be responsible for hypoxic ROS generation. Leach and coworkers (21) propose that complex III of the mitochondrial respiratory chain acts as hypoxic sensor responsible for pulmonary vasoconstriction. For hepatocytes (6) and cardiomyoctes (10), it has been suggested that cytochrome c oxidase of complex IV may contribute to oxygen sensing. More recently, the same group presented data favoring an important role of complex III in hypoxic vasoconstriction (8).

The present study sought to determine the effect of hypoxia on NO and ROS production by vessels of the lung. For these experiments, murine precision-cut lung slices were employed in which on one hand, the cellular architecture of the vessel walls was preserved, and on the other hand, shear stress was avoided. The lung sections were exposed to normoxia or hypoxia in the presence of 4,5-diaminofluorescein diacetate (DAF-2DA) or 2',7'-dichlorofluorescein diacetate (DCF-DA) as fluorescent indicators for NO and ROS, respectively, and the positive cells within the vascular walls were quantified. The source of normoxic and hypoxic ROS generation was characterized by application of inhibitors of flavoproteins and of the individual complexes of the mitochondrial respiratory chain. These investigations included complex II, which often was not taken into account by other groups.


    MATERIALS AND METHODS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Animals. FVB mice were obtained from Harlan Winkelmann (Paderborn, Germany).

Culture medium and chemicals. Minimal essential medium (MEM) was supplied by PAA Laboratories (Coelbe, Germany); DMEM, penicillin, streptomycin, L-glutamine, and pyruvate was supplied by PAN Biotech (Aidenbach, Germany); DMEM NUT MIX F-12 was supplied by GIBCO (Karlsruhe, Germany); and fetal calf serum (FCS) was provided by Greiner (Frickenhausen, Germany). Monoclonal antibodies for characterization of cultured cells were provided by Sigma-Aldrich (Deisenhofen, Germany; anti-smooth muscle actin, clone 1A4 and anti-smooth muscle myosin, clone hSM-V) and by Dako (Hamburg, Germany; anti-von Willebrand factor, clone F8/86). DAF-2DA and diphenyleneiodonium (DPI) were obtained from Alexis Biochemicals (Gruenberg, Germany), DCF-DA was from Calbiochem-Novabiochem (Bad Soden, Germany), rotenone, 3-nitroproprionic acid (3-NPA), thenoyltrifluoroacetone (TTFA), and antimycin A were from Sigma-Aldrich, sodium azide (NaN3) was from Merck (Darmstadt, Germany), and low-melt preparative grade agarose was from Bio-Rad (Munich, Germany). All other reagents were of analytical grade and obtained from Sigma-Aldrich.

Tissue processing. Precision-cut lung slices were prepared in a similar fashion as described previously (14, 22). Four- to six-week-old mice were killed by cervical dislocation and exsanguinated by cutting the vena cava inferior. The chest cavity was opened, and the airways of the lungs were filled with low-melting agarose at 37°C via the trachea. Heart and lungs were removed en bloc and transferred into ice-cold HEPES-Ringer solution (10 mM HEPES, 136 mM NaCl, 5.6 mM KCl, 1 mM MgCl2, 2.2 mM CaCl2, 11 mM glucose, pH 7.4). Precision-cut lung slices (200 µm) were prepared from single lung lobes with a vibratome and cultured in serum-free MEM.

NO assay. DAF-2DA was used to detect NO-producing cells within the pulmonary vasculature. After entering the cell, DAF-2DA is hydrolyzed to DAF-2, which is converted in the presence of NO into the highly fluorescent triazolofluorescein (DAF-2T). Freshly prepared lung sections were transferred into MEM containing 10 µM DAF-2DA and exposed for 3 h to either normoxia (21% O2, 5% CO2, and balance N2) or hypoxia (1% O2, 5% CO2, and balance N2). The lung slices were fixed with 4% buffered formaldehyde for 15 min, washed with PBS, and coverslipped in carbonate-buffered glycerol. DAF-2T-positive cells within the vascular walls were visualized by confocal laser scanning microscopy (Zeiss LSM 410; Zeiss, Jena, Germany) using an excitation wavelength of 488 nm. For a quantitative analysis of NO-generating cells, cross sections of blood vessels (inner diameter: 20-150 µm) were identified by phase-contrast optics, and, subsequently, the fluorescence images were recorded at the following settings: contrast 400 (scale 2-999), brightness 8,925 (scale 0-9,999), and pinhole 30 (scale 0-255). At 400-fold magnification, each vessel was followed in its three-dimensional course within the microscopical image section by scanning the lung slices in steps of ~3 µm in depth. A cell was classified as positive when the intensity of the fluorescence was >150 on a scale of 0-255. The number of DAF-2T-positive cells within such a vessel segment was defined as "DAF-2T-positive cells/vessel". Per condition, two lung sections were analyzed, and for both, the mean was calculated. Finally, the obtained data were statistically analyzed as described below.

ROS assay. Generation of ROS was assayed with DCF-DA. After intracellular cleavage of the diacetate by endogenous esterases and oxidation by ROS, the compound turns into a highly fluorescent product. For detection of ROS-producing cells, freshly prepared lung slices were exposed to 20 µM DCF-DA in MEM for 3 h under either normoxic or hypoxic conditions. Further processing of the organ sections was as described for the NO assay.

Quantification of the intensity of the DCF signal in individual cells. To estimate the intensity of the DCF-fluorescence signal, positive cells (gray value >150 on a scale of 0-255) in vascular walls were outlined, and the average brightness of the enclosed area was measured. Per condition, two lung sections and 15 positive cells/lung section were evaluated and the means calculated. Further processing of the obtained data was performed as described below. In addition, smooth muscle cells were isolated from rabbit pulmonary arteries by carefully preparing <1-mm3 pieces of media, devoid of adventitial tissue, as assessed by microscopic control. These pieces were placed into 12-well culture plates with 500 µl of culture medium (16% DMEM, 64% DMEM NUT MIX F-12, 20% FCS with 100 IU/ml penicillin, 100 µg/ml streptomycin, 2 mM L-glutamine, and 1 mM sodium pyruvate) at 37°C in 95% air-5% CO2. Smooth muscle cell identity was verified by characteristic appearance in phase-contrast microscopy, indirect immunofluorescent labeling for smooth muscle-specific isoforms of alpha -actin and myosin, and lack of immunolabeling for von Willebrand factor. Cells were studied for normoxic and hypoxic (1% O2) ROS production at passage 3 according to a protocol described in detail earlier (16).

Inhibition of the respiratory chain. The following inhibitors were applied simultaneously with DCF-DA: 10 µM DPI for inhibition of flavoproteins, 10 µM rotenone for inhibition of complex I (NADH-ubiquinone oxidoreductase), 5 mM 3-NPA or 10 µM TTFA for inhibition of complex II (succinate-ubiquinone oxidoreductase), 3 µg/ml of antimycin A for inhibition of complex III (ubiquinol-cytochrome c oxidoreductase), and 1 mM NaN3 for inhibition of complex IV (cytochrome c oxidase) (Fig. 1). To investigate the effect of an excess of substrate of succinate dehydrogenase (SDH) on ROS generation, incubations of the lung sections were performed in the presence of 15 mM succinate.


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Fig. 1.   Scheme illustrating the mitochondrial electron transport chain and sites of inhibition. Electrons are transferred from complexes I and II via ubiquinol to complex III and further to complex IV. Within complex III, a bifurcation for the electron shuttle exists: the first electron from ubiquinol is transferred to the iron-sulfur (Fe-S) cluster of complex III, resulting in the formation of ubisemiquinone, and the second electron is shuttled from ubisemiquinone to cytochrome bL (cyt bL) and further to cytochrome bH (cyt bH). Reoxidation of cyt bH occurs by ubiquinone and ubisemiquinone. Roman numerals: complexes I-IV. AA, antimycin A; cyt c and cyt c1, cytochromes c and c1, respectively; DPI, diphenyleneiodonium; 3-NPA, 3-nitroproprionic acid; Q, ubiquinone; QH2, ubiquinol; Q*o and Q*i, ubisemiquinones generated at the outer or inner site of the inner mitochondrial membrane; TTFA, thenoyltrifluoroacetone. Sites of inhibition are indicated in green.

Histochemical detection of SDH activity in murine lung sections. Agarose-filled murine lungs were rapidly frozen in isopentane. To assess the specificity of the SDH assay, agarose-filled lungs were cut into 500-µm-thick sections and preincubated for 15 min at 37°C in MEM and 5 mM 3-NPA. After being washed with MEM, the sections were also frozen in isopentane. Subsequently, 10-µm-thick frozen sections were prepared from the lungs and the organ sections, dried for 5 min at 37°C either by air or in a chamber gassed with N2, fixed for 5 min with cold acetone at 4°C, and dried again as described above. For detection of SDH activity, the sections were incubated for 60 min in a freshly prepared solution of 30 mM potassium phosphate buffer, pH 7.6, 30 mM succinic acid disodium salt, 3 mM MgCl2, 0.57 mg/ml nitro blue tetrazolium, 0.33 mM phenazine methosulfate, and 0.01 mM NaN3. Incubations were performed for 60 min either by air or in a chamber flowing with N2. In an additional set of experiments, incubations were performed in chambers flowing either with 21% O2, 5% CO2, and balance N2 for normoxia or with 1% O2, 5% CO2, and balance N2 for hypoxia. In this case, the reagents for the assay were dissolved in bicarbonate buffer at pH 7.6. The reaction was stopped by a 30-s wash in 0.9% NaCl followed by two 5-min washes in H2O. Finally, the sections were embedded in Kaiser's glycerol gelatin. Staining intensities of vascular walls and of Clara cells of the bronchiolar epithelium were evaluated with an Axioplan 2 microscope (Zeiss, Göttingen, Germany) employing AxioVision 3.0 software and expressed as relative values at a scale of 0-255.

Statistical analysis. Statistical analysis was performed using SPSS Base 8.0 (SPSS Software, Munich, Germany). Percentiles 0, 25, 50, 75, and 100 are presented in box plots. Differences among experimental groups were analyzed with the Kruskal-Wallis and the Mann-Whitney's tests, with P < 0.05 being considered significant and P < 0.01 highly significant.


    RESULTS
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

NO and ROS production within the vascular wall in response to hypoxia. We used precision-cut lung slices cultured either at 21 or 1% O2 to study the effect of hypoxia on NO and ROS production by cells of the pulmonary vasculature. Confocal laser scanning micrographs of NO-producing cells detected by DAF-2DA and of ROS-generating cells detected by DCF-DA are given in Fig. 2A. A quantitative analysis of the number of NO-producing cells within the walls of small- to medium-sized blood vessels (inner diameter: 20-150 µm) revealed no significant effect of hypoxia on the number of positive cells. In contrast, hypoxia induced a distinct increase in the number of ROS-generating cells (Fig. 2B). The relative intensity of the DCF-fluorescence of individual cells with suprathreshold fluorescence intensity was not changed by hypoxia (data presented below together with those obtained with specific inhibitors). On the basis of their morphology and their localization within the vascular wall, we identified the majority of the positive cells as either endothelial cells or smooth muscle cells. In addition, we detected large, irregularly formed cells located within the vascular wall or in the immediate neighborhood that could not be classified unequivocally.


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Fig. 2.   Hypoxia induced increase in reactive oxygen species (ROS) generation but not in nitric oxide (NO) production by cells of intrapulmonary vessels. Generation of NO was assayed with 4,5-diaminofluorescein diactetate (DAF-DA) and that of ROS with 2',7'-dichlorofluorescein diacetate (DCF-DA). Positive cells within the vascular walls of murine lung sections were visualized by confocal laser scanning microscopy (A). The box plot (B) gives the data of a quantitative analysis of the number of positive cells per vessel in lung sections cultured at normoxia (N) or hypoxia (H). The middle horizontal lines indicate the median, the top and bottom of each box identifies the upper and lower quartiles of the distribution, and the top and bottom horizontal lines give the total distribution; n = number of independent experiments. **P < 0.01 (Mann-Whitney's test).

To ascertain that ROS production was indeed increased in vascular smooth muscle cells and to obtain more detailed information of ROS production within individual cells that showed subthreshold DCF-fluorescence in lung slice preparations, ROS production was quantified in cultured smooth muscle cells isolated from rabbit intrapulmonary arteries exposed to either 21 or 1% O2 for 1 h. Because the outlines of individual cells could be clearly traced in these cultured cells, quantification was not restricted to counts of cells with suprathreshold fluorescence intensity, but intensities could be recorded for each individual cell (Fig. 3). In five independent series of experiments, an average and median 1.7-fold increase (SD = 0.25) in ROS production under hypoxia became evident (P < 0.01, Mann-Whitney's test).


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Fig. 3.   A and B: increased ROS production by cultured rabbit pulmonary artery smooth muscle cells under hypoxia (1% O2, 1-h exposure) compared with normoxia (21% O2) as assessed by the fluorescent indicator DCF-DA and confocal laser scanning microscopy.

Generation of ROS within the pulmonary vessels occurs within mitochondria. Several enzymes such as plasmamembrane-located NADPH oxidases, the xanthine/xanthine oxidase system, and the mitochondrial respiratory chain are known to generate ROS in a controlled way (37). To identify the source of normoxic and hypoxic ROS production within the pulmonary vasculature, we exposed lung sections to DPI or rotenone and quantified the number of DCF-positive cells within the vascular walls. DPI is a frequently used inhibitor of flavoproteins, as they exist in complexes I and II of the respiratory chain but also outside of mitochondria, as in NADPH oxidases. In contrast, rotenone is a selective inhibitor of the mitochondrial complex I, blocking the electron transfer from complex I to ubiquinone (Fig. 1). Both at normoxia and hypoxia, ROS generation was almost entirely blocked by DPI and rotenone (Fig. 4, A and B). These results indicate that normoxic and hypoxic ROS production occurs within mitochondria and demands electron influx into the respiratory chain via complex I. 


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Fig. 4.   Effects of inhibitors of the respiratory chain on ROS production by pulmonary vascular cells. Data are presented as box plots with percentiles of 0, 25, 50 (= median, indicated by solid line within the box), 75, and 100. The mean of normoxia of each experiment was normalized to 1. Lung sections were cultured at normoxia or hypoxia in absence or presence of the flavoprotein inhibitor DPI (A), the complex I inhibitor rotenone (ROT; B), the complex II inhibitors 3-NPA (C) and TTFA (D), the complex III inhibitor AA (E), or the complex IV inhibitor NaN3 (F). Inhibition of complexes I and III induced decreases in ROS production both at normoxia and hypoxia, whereas inhibition of complex IV had no effect. Inhibition of complex II selectively suppressed the hypoxic ROS generation; n = number of independent experiments. *P < 0.05, **P < 0.01 (Mann-Whitney's test). Rel., relative.

Inhibition of complex II by 3-NPA or TTFA selectively suppresses hypoxia-induced ROS production. 3-NPA and TTFA are frequently employed inhibitors of complex II (Fig. 1). Coles et al. (9) have shown that 3-NPA is slowly oxidized by SDH, and its oxidation product inhibits the SDH instantaneously and irreversibly. TTFA interferes with the ubiquinone interaction of complex II. In vitro analysis of enzyme preparations obtained from bovine heart revealed that TTFA inhibits both the reduction of ubiquinone to ubisemiquinone and the reduction of ubisemiquinone to ubiquinol (41). Both inhibitors were tested for their effect on normoxic and hypoxic ROS generation.

Under normoxic conditions, the number of ROS-producing cells within the vessel walls was not changed by 3-NPA. In contrast, the hypoxia-augmented ROS generation was entirely prevented by 3-NPA (Fig. 4C). In the presence of TTFA at normoxia, the number of ROS-producing cells per vessel (Fig. 4D) as well as the DCF-fluorescence intensity of those cells with suprathreshold fluorescence intensity (median gray value: 231, range 228-240; after TTFA application vs. a median of 245, range 235-248, in normoxic control; n = 6 slice preparations, in each preparation there were 30 positive cells analyzed) was slightly, although significantly (P < 0.05, Mann-Whitney's test), reduced. This may be due to the recently decribed inhibitory effect of TTFA on esterases responsible for intracellular cleavage of DCF-DA. Inhibition of both complex II and esterases operates in the same concentration range of TTFA (45). At hypoxia, TTFA decreased the number of ROS-generating cells to the level observed at normoxia (Fig. 4D), and the reduction of DCF-fluorescence intensity of individual cells was approximately twice as much as under normoxia (median gray value: 219, range 218-228; after TTFA application vs. median: 244, range 231-245, in hypoxic control; n = 6 slice preparations, in each preparation there were 30 positive cells analyzed; P < 0.01, Mann-Whitney's test).

SDH activity of complex II is reduced under hypoxia. Because the data obtained by complex II inhibitors implied that this complex of the respiratory chain plays an essential role in hypoxia-induced, but not in normoxic, ROS generation, we investigated its enzymatic activity under normoxic and hypoxic conditions. Histochemical detection of SDH activity combined with a quantitative analysis revealed distinct differences in the enzymatic activity of various cell types of the lung. Whereas Clara cells of the bronchial epithelium and cardiac myocytes within the wall of pulmonary veins were intensely stained, indicative of high enzymatic activity, cells of the arterial vascular walls showed only weak labeling (Fig. 5A). Obviously, complex II makes only a minor contribution to the mitochondrial electron chain in vascular smooth muscle cells compared with Clara cells. When the SDH reaction was performed in a nitrogen atmosphere, the signal was significantly reduced, indicative of a hypoxic reduction of the enzymatic activity in both arterial vascular cells (median of the relative staining intensity of vessel walls exposed to nitrogen: 73% of the air control) and Clara cells (median of the relative staining intensity of Clara cells exposed to nitrogen: 25% of the air control) (Fig. 5B). A comparable decrease to 62% was obtained when the incubations were performed in bicarbonate-buffered solutions at 21% O2 or 1% O2 (21% O2: median of the relative staining intensity of vessel walls was 69, range 37.5-76.5; 1% O2: median of the relative staining intensity of vessel walls was 43, range 27.5-63.5; n = 5, in each preparation there were 10 vessels analyzed). The specificity of the SDH assay was proven by the application of 3-NPA. Lung sections preincubated with 3-NPA were nearly unstained (Fig. 5A).


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Fig. 5.   Histochemical detection of hypoxia-induced reduction of succinate dehydrogenase (SDH) activity in lung sections. For normoxia, the enzymatic reaction was performed at air, and for hypoxia, it was performed in a chamber gassed with N2. To assess the specificity of the assay, lung sections were preincubated with 3-NPA before exposure to the reaction solution (N+3-NPA). SDH activity at normoxia, hypoxia, or normoxia in the presence of 3-NPA in vascular walls and in Clara cells of bronchial epithelium is evident in original micrographs (A). All pictures were taken at the same camera settings. The quantitative data are presented as box plot (B). n = number of independent experiments. *P < 0.05 (Mann-Whitney test).

Because, on one hand, the data obtained with complex II inhibitors pointed to an increased activity of this complex under hypoxia, whereas on the other hand, its SDH activity was diminished under this condition, we tested whether the inverse enzymatic reaction, i.e., fumarate reductase instead of SDH, contributed to hypoxic ROS production. To this end, we applied an excess of succinate (15 mM, i.e., threefold molar excess compared with glucose) to the medium that would facilitate SDH activity while inhibiting fumarate reductase activity. At normoxia, succinate had no effect on the number of ROS-generating cells while it resulted in a significant diminution of ROS-generating cells at hypoxia (Fig. 6).


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Fig. 6.   Effect of succinate (Succ), the substrate of SDH, and end product of fumarate reductase, on ROS production. The mean of normoxia of each experiment was normalized to 1; data are presented as box plot. The relative number of DCF-positive cells per vessel remained unaltered in the presence of 15 mM succinate at normoxia, whereas it was drastically reduced under hypoxic conditions. n = number of independent experiments. *P < 0.05 (Mann-Whitney's test).

Inhibition of complex III by antimycin A blocks both normoxic and hypoxic ROS generation. Complexes I and II supply the ubiquinone-ubisemiquinone-ubiquinol cycle (Q-cycle) of complex III with electrons via ubiquinol. Its oxidation at the Qo site entails an obligatory bifurcation: the first electron is transferred from ubiquinol to the "high potential" chain consisting of an iron-sulfur protein and cytochrome c1 (cyt c1), leaving an unstable ubisemiquinone species at the Qo site. This ubisemiquinone is oxidized by the "low potential" chain consisting of cytochromes bL (cyt bL) and bH (cyt bH), which sends the electron to the Qi site where, by two turnovers, ubiquinone is reduced successively to ubisemiquinone and ubiquinol. Antimycin A is an inhibitor of the Q-cycle; it blocks the reoxidation of cyt bH (32, 35) and, thereby, the generation of ubisemiquinone and ubiquinol at the Qi site (Fig. 1). Quantification of DCF-positive cells in vessels of lung sections exposed to antimycin A revealed a highly significant decrease of both normoxic and hypoxic ROS generation (Fig. 4E). Simultaneously, the DCF-fluorescence intensity of cells with suprathreshold fluorescence intensity was greatly attenuated both at normoxia (median gray value: 171, range 163-204; after antimycin A application vs. median: 235, range 223-248, in normoxic control; n = 4 slice preparations, in each preparation there were 30 positive cells analyzed) and at hypoxia (median gray value: 173, range 167-193; after antimycin A application vs. median: 238, range 216-245, in hypoxic control; n = 4 slice preparations, in each preparation there were 30 positive cells analyzed). Hence, complex III appears to be critical for ROS production at both conditions.

Inhibition of complex IV by NaN3 has no effect on normoxic and hypoxic ROS production. NaN3 is a noncompetitive inhibitor of cytochrome c oxidase of complex IV (10). At neither normoxia nor hypoxia was the number of ROS-generating cells within the vascular walls influenced by NaN3 (Fig. 4F), indicating that under both conditions ROS production occurred upstream of complex IV.


    DISCUSSION
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ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
REFERENCES

Lung sections have previously been used to study the response of individual airways and vessels to various drugs (14, 22), and they have been applied to determine the metabolism of a tobacco-specific nitrosamine and its proximal metabolite (28). Because these and other reports have shown that lung slices behave, in many aspects, like intact lungs, we decided to use them to study hypoxia-regulated NO and ROS production by cells of intrapulmonary vessels. From one murine lung, ~15 sections could be obtained. This enabled us to compare the effect of different oxygen concentrations on neighboring areas of the same lung, and it drastically reduced the number of mice necessary for the investigations.

In contrast to cell culture systems, the cellular architecture of the vascular wall is preserved in precision-cut lung sections, and the vessels are kept in their natural histological environment. Thereby, it is possible to selectively quantify NO- or ROS-generating cells within those areas of the lung responsible for hypoxic vasoconstriction and vascular remodeling, i.e., the walls of small intrapulmonary arteries. Compared with isolated perfused lungs, the use of organ sections allows investigation of the effect of hypoxia under conditions free of shear stress, which by itself modulates NO and ROS production. In cultured endothelial cells, shear stress results in increased ROS generation (17) and NO release (24). In the vasculature of lung sections exposed for 3 h to normoxia or hypoxia, we detected no difference in the number of NO-generating cells. Therefore, we suppose that NO is not involved in direct short-term reactions to hypoxia in the absence of shear stress.

In contrast, the number of ROS-generating cells within the vascular walls was significantly increased by hypoxia. Employing specific inhibitors, we identified mitochondria as the cellular source of ROS. These findings are in agreement with various other reports describing an enhanced mitochondrial ROS production at reduced oxygen supply (7, 10, 38).

The mitochondrial electron transport chain is composed of four complexes (Fig. 1). Complex I and complex II oxidize NADH and succinate, respectively, to reduce ubiquinone to ubiquinol. One electron of ubiquinol is transferred to iron-sulfur protein and, subsequently, to cyt c1 of complex III. The reduced cyt c1 donates the electron via cytochrome c to complex IV. Ultimately, molecular oxygen reacts as final electron acceptor, and, in the presence of protons, H2O is formed. The second electron from ubiquinol flows into the Q-cycle where it is transferred successively to cyt bL and cyt bH. The free radical intermediate ubisemiquinone may occur at all sites where reduction of ubiquinone or oxidation of ubiquinol takes place, i.e., complexes I, II, and III. This single-electron carrier is considered to donate electrons also to O2 for the generation of ROS (36). It has been estimated that at normoxia, 1-2% of all electrons passing through the respiratory chain end up as O<UP><SUB>2</SUB><SUP>−</SUP></UP> · that are subsequently converted into other ROS (4). In line with the potential occurrence of ubisemiquinone at complexes I, II, and III, but not at complex IV, we observed alterations of ROS production when agents interfering with the first three complexes were applied, i.e., DPI, rotenone, 3-NPA, TTFA, and antimycin A, but not when complex IV was inhibited by NaN3. Our present results, however, implicate differences in the participation of the individual complexes in the electron flow and ROS generation at normoxia and hypoxia.

At normoxia (Fig. 7A), electrons contributing to ROS formation almost exclusively entered the respiratory chain via complex I since ROS generation was 1) nearly completely abolished in presence of rotenone, which blocks electron entry via complex I, and 2) unaffected by inhibition of complex II by 3-NPA and only minimally reduced by the complex II inhibitor TTFA. This slight reduction in the number of ROS-generating cells by TTFA may well be explained by its recently described inhibitory effect on esterases responsible for intracellular cleavage of DCF-DA. Inhibition of both complex II and esterases operates in the same concentration range of TTFA (45). Also in agreement with the assumption of a rather neglectable role of complex II in pulmonary vessels is the faint signal in the vascular wall obtained for SDH activity observed by histochemistry, fully in line with previous reports in which clear SDH activity was described in bronchiolar epithelial cells but not in the pulmonary arterial wall (34).


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Fig. 7.   Proposed model describing an oxygen-dependent change in the directionality of the electron flow at complex II of the mitochondrial respiratory chain. At normoxia (A), electrons are mainly transferred from complex I via ubiquinol to complex III and further to complex IV. At hypoxia (B), the activity of complex II is switched from SDH to fumarate reductase; now fumarate also acts as terminal electron acceptor, receiving its electrons also from the ubiquinol pool fueled by complex I.

The site at which ROS are generated during normoxia cannot be exactly deduced from the present experiments, but it is clear that it requires electrons fed by complex I, because of its sensitivity to rotenone, and an intact Q-cycle at complex III, because of its sensitivity to antimycin A, which blocks the reoxidation of cyt bH by ubiquinone and ubisemiquinone (35). Both inhibitors rotenone and antimycin A may either reduce or enhance normoxic ROS production, depending on cell type and organ investigated. In most cell types, rotenone reduces ROS production, as also observed in the present study, but it causes a marked increase in ROS generation in rat sensory neurons (15), rat pheochromocytoma cells (16), and rat renal arteries (23). Antimycin A, on the other hand, increases ROS production in most cell types, e.g., cardiomyocytes (10, 27), hepatoma cells (7), and rat renal arteries (23), and based on these observations, it has been discussed that the antimycin A-induced contraction of cultured pulmonary vascular smooth muscle cells and the transient increase of pulmonary pressure in perfused lungs may be related to increased ROS production (38). This interpretation, however, is difficult to match with the observation that rotenone, which is generally accepted to decrease mitochondrial ROS production in vascular cells, shares the transient pulmonary vasoconstrictor effect with antimycin A (29), and direct measurements have shown that antimycin A reduces normoxic ROS generation in rat (1, 23) and murine pulmonary arteries (this study), as it also does in HL-60 cells (2). Differences in methodologies employed by various research groups, rather than cell type or organ-specific differences, are unlikely to account for the observed discrepancies in the rotenone- and antimycin A-induced effects because they could be validated by identical methodology in a single study, and further evidence for organ-specific diversity in smooth muscular mitochondrial function has also been presented (23).

At hypoxia (Fig. 7B), our data provide evidence for an essential role of complex II in ROS formation, and this complex possibly acts as oxygen sensor. Complex II is a component of both the Krebs cycle and the mitochondrial electron transport chain. It is the smallest complex of the respiratory chain and consists of four proteins: the catalytic core is composed of two subunits containing a FAD (SDHA) and three iron-sulfur clusters S1-S3 (SDHB), respectively. It is anchored to the matrix side of the mitochondrial inner membrane by a large (SDHC) and a small (SDHD) subunit that together comprise the membrane-spanning heme protein cyt b (12). In the Krebs cycle, this complex oxidizes succinate to fumarate, thereby providing two electrons to the respiratory chain by reducing ubiquinone. A high SDH activity can be demonstrated in several organs and cell types, e.g., cardiomyocytes and Clara cells, whereas it is low or even absent at other sites such as axon terminals in the nervous system or pulmonary vessels (13, 25, 34, this study). Under certain conditions, electrons flow in the reverse direction so that this complex acts, then, as fumarate reductase rather than as SDH (12). For complex II from bovine heart, this has been observed when the purified enzyme is analyzed under anaerobic conditions or when the mitochondrial membrane is in a highly energized state (44). These in vitro data from bovine heart are supported by a reduction of SDH activity in hypoxic rat hearts (25). Wiesner et al. (40) performed measurements of metabolites in rat hearts perfused with buffer equilibrated with 95% N2, 5% CO2. The contents of succinate increased from 0.08 ± 0.03 µmol/g wet wt of control hearts to 0.45 ± 0.11 µmol/g wet wt of hypoxic hearts. Because this is distinctly lower than the succinate concentration (15 mM) applied in our experiments for inhibition of ROS generation, we suppose that complex II can act as fumarate reductase even at persistent hypoxia. Although the properties of fumarate reductase in mammalian mitochondria are not fully investigated, the fumarate reductase of Escherichia coli is an extremely effective generator of O<UP><SUB>2</SUB><SUP>−</SUP></UP> (18).

These data suggest that complex II may switch its function from SDH to fumarate reductase under hypoxia, thereby generating ROS and accumulating succinate, and our present findings provide strong evidence that this occurs indeed in pulmonary vessels. First, inhibition of complex II specifically reduced hypoxia-induced ROS generation, demonstrating that some kind of activation of complex II under hypoxia shall exist. This activation is not represented by SDH activity since 1) decreased instead of increased SDH activity under hypoxia could be directly demonstrated histochemically, and 2) addition of the substrate for SDH, i.e., succinate, led to a decreased instead of increased ROS formation. This inhibitory effect of succinate on hypoxic ROS formation can best be explained if complex II acts as fumarate reductase, which would be inhibited by excess of its metabolite, succinate. For acting as fumarate reductase, complex II has to be supplied with electrons. It has been demonstrated that these electrons can be provided by NADH (44). Fully in line with this model is our present finding that inhibition of NADH dehydrogenase (= complex I) by rotenone also blocked the hypoxia-induced increase in ROS production. Obviously, an intact Q-cycle of complex III is also required under these conditions, since its inhibition by antimycin A also blocked hypoxia-induced ROS generation. This implies that not all electrons fed into the transport chain are directly transferred via complex II to fumarate as terminal electron acceptor, and electron transfer via the iron-sulfur protein and cyt c1 toward complex IV may occur in parallel. The exact role of complex III under these conditions, however, remains to be determined.

Mutations in genes encoding the proteins of complex II and the resulting phenotypes also argue for an essential role of complex II in oxygen sensing and/or signaling. A mutant of the nematode Caenorhabditis elegans, which shows an inverse correlation between lifespan and oxygen concentration, exhibits a missense mutation in the SDHC gene (19). Cawthon et al. (5) employed various inhibitors of the mitochondrial respiration to detect a site-specific defect in the electron transport chain of hepatic mitochondria isolated from broilers with pulmonary hypertension syndrome. Their results indicate the presence of a site-specific defect at complex II and/or ubiquinone. Hereditary tumors of the carotid body, an organ that senses arterial oxygen tension, are rare, usually benign, and slow-growing tumors sharing several features with carotid bodies from individuals dwelling at high altitudes with chronic hypoxemia. These tumors are caused by mutations within the genes of SDHB, SDHC, and most commonly in SDHD (3).

In summary, we were able to demonstrate that hypoxia induces an increase in ROS generation by cells of the pulmonary vasculature and that mitochondrial complex II plays an essential role in this process. Now it will be important to clarify which cellular pathways are activated by this mechanism.


    ACKNOWLEDGEMENTS

We thank Christian Schuldt for isolating rabbit pulmonary artery smooth muscle cells and K. Michael for excellent technical assistance.


    FOOTNOTES

The financial support of the Deutsche Forschungsgemeinschaft (Sonderforschungsbereich 547, projects C1 and B7) is gratefully acknowledged.

Address for reprint requests and other correspondence: R. Paddenberg, Institute of Anatomy and Cell Biology, Justus-Liebig-Univ., Aulweg 123, 35385 Giessen, Germany (E-mail: Renate.Paddenberg{at}anatomie.med.uni-giessen.de).

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.

First published January 10, 2003;10.1152/ajplung.00149.2002

Received 14 May 2002; accepted in final form 20 December 2002.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
MATERIALS AND METHODS
RESULTS
DISCUSSION
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