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In adult respiratory distress syndrome (ARDS) pulmonary perfusion failure increases physiologic dead-space (VD/VT) correlating with mortality. High VD/VT results in alveolar hypocapnia, which has been demonstrated to cause edema formation, atelectasis, and surfactant depletion, evoked, at least in part, by apoptosis of alveolar epithelial cells (AEC). However, the mechanism underlying the hypocapnia-induced AEC apoptosis is unknown. Here, using fluorescent live-cell imaging of cultured AEC type 2 we could show that in terms of CO2 sensing the tricarboxylic acid cycle enzyme isocitrate dehydrogenase (IDH) 3 seems to be an important player because hypocapnia resulted independently from pH in an elevation of IDH3 activity and subsequently in an increase of NADH, the substrate of the respiratory chain. As a consequence, the mitochondrial transmembrane potential (ΔΨ) rose causing a Ca2+ shift from cytosol into mitochondria, whereas the IDH3 knockdown inhibited these responses. Furthermore, the hypocapnia-induced mitochondrial Ca2+ uptake resulted in reactive oxygen species (ROS) production, and both the mitochondrial Ca2+ uptake and ROS production induced apoptosis. Accordingly, we provide evidence that in AEC type 2 hypocapnia induces elevation of IDH3 activity leading to apoptosis. This finding might give new insight into the pathogenesis of ARDS and may help to develop novel strategies to reduce tissue injury in ARDS.
The adult respiratory distress syndrome (ARDS) is a typical complication of ventilated intensive care patients with an incidence of almost 10% and a mortality rate of ~40%. There are many predisposing factors known, including pneumonia, sepsis, or trauma. Inflammation, alveolar-capillary barrier dysfunction, development of lung edema, and formation of atelectasis typically in the dorsal regions of both lungs are the hallmarks of ARDS. Accordingly, opening of atelectatic lung regions, avoiding their re-collapse by applying a positive end-expiratory pressure, reducing the tidal volume during ventilation, and, if necessary, extracorporeal oxygenation and CO2 removal are the treatment approaches in ARDS.
Although the pathogenesis of ARDS has been extensively studied over the last decades, a causal therapy has not been found yet. During the last years it became clear that in both the ARDS and its experimental equivalent, the acute lung injury, intravascular coagulation, and perfusion disorders lead to an increase of the physiologic dead-space fraction (VD/VT), which is associated with increased mortality.1 In contrast to the CO2 accumulation in the arterial blood, the CO2 concentration in alveoli may drop, especially in lung regions with high VD/VT. It has been shown that alveolar hypocapnia may contribute to tissue injury, including depletion of surfactant, which is produced by alveolar epithelial cells (AEC) type 2 and normally opposes the alveolar-collapsing tendency by lowering the air–liquid surface tension. Furthermore, reduced secretion of surfactant is commonly seen in ARDS patients and is associated with worse outcomes especially in critically-ill patients (which has been excellently reviewed by J Laffey et al.2, 3, 4, 5, 6). Thus, the alveolar hypocapnia-induced AEC type 2 damage seems to be likely in ARDS. However, the underlying mechanism has not been elucidated yet.
Apoptosis of AEC potentially has an important role in ARDS. Decreased size and condensation of the chromatin of AEC type 2 have been shown early in human ARDS.7 Moreover, caspase-cleaved cytokeratin-18, a marker for epithelial cell apoptosis, increases in bronchoalveolar lavage fluids during ARDS.8
Several pathways of apoptosis induction are known, also in the context of ARDS. One important trigger for apoptosis is the mitochondrial Ca2+ overload, especially in combination with increased mitochondrial production of reactive oxygen species (ROS).9 Importantly, mitochondrial Ca2+ uptake and ROS production can mutually be dependent. The driving force for mitochondrial Ca2+ uptake is the negative transmembrane potential gradient (ΔΨ) across the mitochondrial inner membrane.10, 11 There are reports that ΔΨ can be increased by stimulation of the respiratory chain with its substrate NADH, which is produced in the tricarboxylic acid (TCA) cycle by the enzymes isocytrate dehydrogenase (IDH) 3 and α-ketoglutarate dehydrogenase12, 13, 14, 15 together with CO2.
According to the law of mass action the accumulation of the product may inhibit, whereas a rapid clearance of the product may accelerate its enzymatic reaction rate. We therefore hypothesize that a decrease of the alveolar partial pressure of CO2 (pCO2), which occurs in lung regions with high VD/VT, increases IDH3 activity, NADH production, and thus ΔΨ in AEC.
Using real-time fluorescence microscopy of AEC type 2 we could identify that mitochondrial IDH3 is a key player in CO2 sensing. Accordingly, under hypocapnic conditions the resulting increase of IDH3 activity elicits an increase of ΔΨ and subsequently a shift of Ca2+ from the cytosol into mitochondria. In consequence, the hypocapnia-induced mitochondrial Ca2+ uptake leads to elevation of mitochondrial ROS production, and thus apoptosis. The understanding of this context may help to develop strategies to reduce AEC type 2 injury, including formation of atelectasis, in ARDS patients with elevated VD/VT. Furthermore, the opening of atelectasis, a central element of ADRS therapies, should be re-evaluated in non-perfused lung regions to prevent hypocapnic conditions and thus apoptosis of AEC.
Using extracts of primary isolated AEC type 2 we were able to show that hypocapnia at constant pH of 7.4 did not induce a detectible increase of IDH3 activity (Figure 1a). We assumed that the IDH3 product NADH, which was measured for IDH3 activity determination, was oxidized by the complex 1 of the respiratory chain before its colorimetric assay detection. Therefore, we inhibited the complex 1 with rotenone. Under these conditions, hypocapnia-induced NADH production was detectable. Furthermore, the respective NADH production rate was higher in comparison with rotenone alone (Figure 1a). Importantly, the used biochemical assay was specific for the mitochondrial isoenzyme IDH3, because the added co-substrate NAD+ does not react with the non-mitochondrial IDH isoforms 1 and 2.
In addition, using biexponential 2-photon fluorescence lifetime imaging microscopy (FLIM) we could show that lifetimes for free (τfree) or bound (τbound) NAD(P)H were 438±47 or 2156±295ps, respectively (Supplementary Figure 1a). In total, 91.8±0.5% of the detected NAD(P)H was bound (Supplementary Figure 1b). According to Blacker et al.16 τbound measurements can be used to discriminate between NADH and NADPH. These authors predicted that τbound values for NADH and NADPH were 1.5 and 4.4ns, respectively. Hence, the measured τbound values of ~2 indicated that under both normo- and hypocapnic conditions primary isolated AEC type 2 produced rather NADH than NADPH (Figures 1b and c). We could detect the NAD(P)H fluorescence predominantly in small granules located around the non-fluorescent nucleus, which are ascribed to mitochondria,17 (Figure 1a Supplementary Figure 1b and d) indicating that under our experimental condition the main NADH production site is the TCA cycle. In line with the enzymatic IDH3 activity assay results, hypocapnia alone did not change NAD(P)H fluorescence in comparison with normocapnic conditions (Figures 1e and f). However, after rotenone pretreatment NAD(P)H fluorescence steadily increased and further increased after additional hypocapnia (Figures 1d–f) in comparison with hypocapnia alone. Because after rotenone alone or in combination with hypocapnia NAD(P)H fluorescence increased, whereas after both interventions τbound stayed stable at 2ns we assume that under both conditions, rather NADH than NADPH was produced (Figures 1b and c). All experiments were conducted at constant pH of 7.4.
In accordance with the FLIM experiments on primary isolated AEC type 2, in rotenone-treated A549 cells the NAD(P)H auto-fluorescence increase was higher under hypo- than normocapnic conditions (Figures 1g–i). Importantly, the NAD(P)H auto-fluorescence in A549 cells was confined to mitochondrial localization, indicating again the mitochondrial origin of the NAD(P)H production (Figure 1g). As auto-fluorescence is not specific to NADH we further measured NADH concentration in A549 cells extracts by using an enzymatic assay. We revealed that NADH concentration 15min upon onset of hypocapnia was only slightly increased in comparison to normocapnia (Supplementary Figure 2). However, after rotenone hypocapnia doubled the NADH concentration within 30min (Supplementary Figure 2), indicating again that without rotenone treatment the NADH produced under hypocapnia is immediately consumed by the complex 1 of the respiratory chain.
To further elucidate the specificity of IDH3 in the hypocapnia-induced NADH production process we downregulated this enzyme in A549 cells. The mitochondrial IDH3 knockdown blocked the hypocapnia-induced increase of the NAD(P)H auto-fluorescence (Figures 1g–i). The IDH3 knockdown was verified on mRNA and protein level (Figures 1j and k, respectively). Taken together, we conclude first that, independent from pH, hypocapnia increases the mitochondrial IDH3 activity and thus the NADH production rate and second that, directly after its synthesis, NADH is totally consumed by the complex 1 of the respiratory chain.
We next determined whether the hypocapnia-induced increase of IDH3 activity, NADH production rate and subsequent respiratory chain NADH consumption elevated ΔΨ. The characterization of the primary isolated AEC type 2 was revealed by the detection of LTG-stained lamellar bodies (Figure 2a). In these cells, hypocapnia at constant pH of 7.4 led to an increase of the ratio between the mitochondrial and cytosolic tetramethylrhodamine methyl ester (TMRM) fluorescence intensity equivalent for an increase of ΔΨ (Figures 2b). Under normocapnic conditions, rotenone decreased, as expected, the TMRM ratio (Figures 2d and e) verifying the correctness of the method. In these rotenone-treated cells, hypocapnia did not increase the TMRM ratio and thus ΔΨ (Figures 2d and e). All these results could be reproduced in A549 cells (Figures 2f–i). Furthermore, in these cells the IDH3 knockdown inhibited the hypocapnia-induced increase of ΔΨ (Figures 2f, g and i). Summarizing, the hypocapnia-induced increase of IDH3 activity, NADH production rate, and subsequent respiratory chain NADH consumption caused an elevation of mitochondrial ΔΨ.
As mentioned before, ΔΨ can be the driving force for mitochondrial Ca2+ uptake but it is also well known that mitochondrial Ca2+ regulates IDH3 and respiratory chain activity. Accordingly, we conclude that mitochondrial Ca2+ is not responsible for the observed ΔΨ changes. We could demonstrate that the silencing of the mitochondrial Ca2+ uniporter (MCU) did not inhibit the hypocapnia-induced ΔΨ increase (Figure 2i). The MCU knockdown was verified on mRNA and protein level (Figures 2j and k, respectively).
We next determined in primary isolated AEC type 2 the hypocapnia-induced mitochondrial Ca2+ uptake. The characterization of these cells was again revealed by the detection of LTG-stained lamellar bodies (Figure 3a). Switching from normo- to hypocapnic conditions induced a rhod-2 fluorescence intensification, equivalent to an increase of the mitochondrial Ca2+ concentration [Ca2+]mito (Figures 3b). This response was completely reversible after switching back to normocapnic buffer. All experiments were performed at a constant extracellular pH of 7.4. The mitochondrial localization of the rhod-2 fluorescence was proofed by its co-localization with the mitochondrial marker MitoTracker green (MTG) (data not shown) and by the observation that in AEC type 2 the rhod-2 fluorescence decreases, as expected, after inhibition of the mitochondrial electron chain by rotenone (Figures 3d and f). Furthermore, in primary isolated AEC type 2 cells the hypocapnia-induced increase of the rhod-2 fluorescence intensity could be blocked by either rotenone or the MCU inhibitor ruthenium red (Figures 3d–f). In line with these findings hypocapnia induced an increase of rhod-2 fluorescence in A549 cells at constant pH of 7.4 (Figures 3g). In addition, the hypocapnia-induced mitochondrial Ca2+ uptake could be verified by FRET (Supplementary Figure 3A–C). The measurement of mitochondrial Ca2+ with FRET was validated by ATP or rotenone as it is well known that ATP increases and rotenone decreases [Ca2+]mito.18 Moreover, in A549 cells the MCU silencing prevented the hypocapnia-induced increase of rhod-2 fluorescence intensity (Figures 3i and j). We therefore assumed that hypocapnia shifts Ca2+ from the cytosol into mitochondria. Importantly, IDH3 silencing inhibited the hypocapnia-induced increase of rhod-2 fluorescence intensity indicating that the elevated IDH3 activity and ΔΨ were responsible for the hypocapnia-induced mitochondrial Ca2+ uptake (Figures 3i and j).
To further test if mitochondria incorporate Ca2+ from the cytosolic compartment we measured cytosolic Ca2+ concentration [Ca2+]cyt in primary isolated AEC type 2. A switch of pCO2 from 40 to 0mmHg decreased [Ca2+]cyt by 25±5% versus baseline conditions within 10min in a reversible manner (Figures 4a–c, e). Exposing primary isolated cells to hypocapnic conditions for 30min revealed that the hypocapnia-induced [Ca2+]cyt decrease is transient (Supplementary figure 4). Interestingly, switching back to normocapnic conditions causes an overshooting increase of [Ca2+]cyt assuming that cells compensate the hypocapnia-induced [Ca2+]cyt decrease by mechanisms, as e.g., decrease of cell membrane Ca2+ pump activity. The pretreatment of AEC type 2 cells with ruthenium red or rotenone inhibited the hypocapnia-induced [Ca2+]cyt decrease (Figures 4d and e). In accordance to these findings, in A549 cells hypocapnia reduced [Ca2+]cyt 20±2%, which was inhibited by rotenone (Figures 4f). In addition, both the MCU and IDH3 silencing inhibited the hypocapnia-induced decrease of [Ca2+]cyt (Figure 4h).
We could demonstrate that in native A549 cells ATP alone caused an increase of [Ca2+]cyt, which was comparable to that after ATP in combination with rotenone. Further, the ATP response in native A549 cells did not differ from the ATP responses in scRNA- or IDH3-siRNA-treated cells. This indicates that both rotenone and IDH3 knockdown did not affect cell viability at least in terms of receptor-mediated Ca2+-signaling cascades (Supplementary figure 5).
Furthermore, we were also interested whether pH may play a role in Ca2+ response induced by hypocapnia. Without additional buffering, the removing of the dissolved CO2 increased the pH of the superfusion buffer from 7.4 to 7.8. Under these conditions hypocapnia induced a similar reduction of [Ca2+]cyt in A549 cells as at constant extracellular pH of 7.4 revealed by buffering (Supplementary figure 6A). In contrast, an alkalosis of 7.8 under normocapnic conditions did not induce an alteration of [Ca2+]cyt (Supplementary figure 6A). Furthermore, the cytosolic pH was not affected in unbuffered cells (Supplementary figure 6B). We therefore conclude that the hypocapnia-induced [Ca2+]cyt responses were pH independent.
Since Ca2+ can be shifted directly from the endoplasmatic reticulum (ER) to mitochondria, we measured Ca2+ in the ER [Ca2+]ER. In contrast to [Ca2+]cyt and [Ca2+]mito, hypocapnia did not change [Ca2+]ER (Supplementary figure 7A–C). However, this ratio was clearly elevated after ATP application indicating the correctness of the method (Supplementary figure 7A–C). Taken these results together, we conclude that hypocapnia and thus acceleration of IDH3 activity induces a Ca2+ shift from the cytosol to the mitochondria, whereas, the ER seems to have a minor role in this scenario.
To investigate whether hypocapnia causes mitochondrial ROS production in primary isolated AEC type 2 cells we used the fluorescent MitoSOX Red mitochondrial superoxide indicator for live-cell imaging. We could show that under hypocapnia the ROS production, expressed as delta increase of the dye, was enhanced in comparison with normocapnia (Figure 5). To evaluate the respiratory chain as potential source of ROS we used rotenone under both normo- and hypercapnia. Rotenone almost diminished the hypocapnia-induced delta increase, whereas rontenone with normocapnia did not change ROS production in comparison with normocapnia alone (Figure 5). In addition, ROS production was also inhibited after ruthenium red incubation, which blocked the mitochondrial calcium uniporter and subsequently the mitochondrial Ca2+-uptake. Mitochondrial Ca2+ and ROS are potential trigger for apoptosis. Therefore, we used a CellEvent Caspase-3/7 fluorogenic substrate to detect apoptotic cells upon hypercapnia in a live-cell image setting. As depicted in Figure 6a, the percent of apoptotic cells increased from 20.1±5.9 under normocapnia to 48.8±6.6 under hypercapnia. Treatment with rotenone or ruthenium red prevented the initiation of apoptosis (23.8±8.1% apoptotic cells and 16.4±4.6% apoptotic cells, respectively) (Figure 6a). In order to validate these results we measured as an additional apoptosis marker the expression of cleaved poly-(ADP-ribose) polymerase (PARP) in primary isolated AECs type 2.Western blot analysis of this protein revealed that only cells cultured under hypocapnic conditions express cleaved PARP, which was inhibited by rotenone or ruthenium red (Figure 6b). Under normocapnic condition alone or in combination with rotenone or ruthenium red cleaved PARP was not detectable (Figure 6b).
We provide evidence that in AEC type 2 hypocapnia increases IDH3 activity and NADH production, ΔΨ, [Ca2+]mito, and mitochondrial ROS concentration, which in turn induces apoptosis (Figure 7).
For the first time, at least to our knowledge, we were able to measure an increase of the IDH3 activity in primary isolated AEC type 2 in response to hypocapnia. In this context, we have to critically emphasize that we could not see any increase of NADH concentration in AEC type 2 subjected solely to hypocapnia. Instead, after rotenone we could measure a steady rise of NADH concentration, which could further be increased in combination with hypocapnia. We concluded that NADH produced under hypocapnic conditions is immediately consumed by complex 1 and can thus not been detected. Inhibition of complex I however blocked the NADH consumption and after this intervention the hypocapnia-induced NADH production was detectable.
The dependency of the respiratory chain activity from NADH has been demonstrated by Kuznetsov et al.19 Hence, as a consequence of hypocapnia-induced elevation of NADH production, ΔΨ and thus the driving force for the mitochondrial Ca2+ uptake increased. This hypothesis was supported by the IDH3 knockdown. However, the IDH3 knockdown did not completely blocked the hypocapnia-induced NADH and ΔΨ responses. Reason for this result could be either that the IDH knockdown was not efficient enough or, more importantly, other mechanisms for the hypocapnia-induced NADH and ΔΨ responses are involved. Hypothetically, low pCO2 could also act on mitochondrial Ca2+ handling, e.g., on increase of mitochondrial Ca2+ influx or decrease of mitochondrial Ca2+ efflux. In this context another aspect has to be discussed: because IDH3 is one of the first enzymes of the TCA cycle and therefore upstream of all other TCA cycle enzymes, e.g., the CO2 producing alpha-ketoglutarate dehydrogenase, the IDH3 knockdown would affect these enzyme activities, which could also contribute to the hypocapnia-induced changes of in NADH and ΔΨ. This might be the reason why the IDH knockdown was highly efficient. In contrast, it could be assumed that the IDH3 knockdown would affect the overall flux through the TCA cycle. Unfortunately, we cannot exclude this hypothesis. However, we measured cytosolic Ca2+ responses to the purinergic receptor agonist ATP in IDH3-siRNA-treated AEC and the revealed Ca2+ responses were comparable to native and even to rotenone-treated AEC indicating that, at least, G-protein coupled receptor Ca2+ signaling is intact.
To determine whether the hypocapnia-induced increase of NADH and ΔΨ elicits a Ca2+ shift into mitochondria from the cytosol we measured both [Ca2+]mito and [Ca2+]cyt. Accordingly, we could demonstrate on both primary isolated AEC type 2 and A549 cells that hypocapnia was accompanied not only by a [Ca2+]mito increase but also by a [Ca2+]cyt decrease. In order to differentiate whether the Ca2+ responses in AEC type 2 were CO2 rather than pH mediated, we additionally measured [Ca2+]cyt primarily in A549 cells under several pH conditions and found that in AEC type 2 [Ca2+]cyt response to CO2 is independent from pH.
Moreover, we could show that the hypocapnia-induced [Ca2+]cyt response was inhibited by MCU knockdown or pharmacological blockade of the MCU with ruthenium red confirming a Ca2+ shift from the cytosol into mitochondria. From these experiments we further conclude that the driving force for the mitochondrial Ca2+ uptake came from the mitochondria and not from the cytosol since in response to hypocapnia [Ca2+]cyt rather de- than increased. According to the literature, the [Ca2+]mito increase can also be evoked by a Ca2+ shift from the ER directly into mitochondria. In our scenario this pathway seems to be unlikely because [Ca2+]ER was unaffected by hypocapnia. However, we cannot exclude the possibility that the ER was synchronously re-filled with cytosolic Ca2+, whereas Ca2+ was shifted from the ER into mitochondria. It has also been described that mitochondria can uptake Ca2+ from the extracellular space. Indeed, this possibility, at least in part, cannot be excluded by our experiments, too.
The inhibition of the CO2-induced Ca2+ shift by rotenone and the unaffected [Ca2+]ER suggests that the driving force for the mitochondrial Ca2+ uptake was an increase of the respiratory chain activity and hence ΔΨ rather than ER. This hypothesis was confirmed by our finding that the fluorescence of TMRM, which accumulates in mitochondria in a potential-dependent manner, was enhanced in both primary isolated AEC type 2 and in A549 cells under hypo- compared with normocapnic conditions. Addition of rotenone revealed, as expected, a quick decrease of the TMRM fluorescence indicating that the respiratory chain activity is the important determinant of ΔΨ. Further, hypocapnia induced an increase of the respiratory chain substrate NADH and the knockdown of the NADH-producing IDH3 inhibited NADH, ΔΨ, [Ca2+]mito, and [Ca2+]cyto response under low CO2 levels.
Taken together, we suggested that CO2, a by-product of IDH3, inhibits IDH3 activity via a negative feedback loop. Under hypocapnic conditions, this inhibitory effect of CO2 on the IDH3 activity is reduced which subsequently increases NADH production and in turn ΔΨ which, at least in part, lead to a Ca2+ shift into mitochondria.
The hypocapnia-induced mitochondrial Ca2+ uptake was associated with AEC type 2 apoptosis. Ca2+ enters mitochondria via the Ca2+ uniporter. As its pharmacological inhibition by ruthenium red prevented both Ca2+ uptake and apoptosis we conclude that the hypocapnia-induced Ca2+ uptake was, at least in part, responsible for the apoptosis of AEC type 2. We can only speculate about the underlying mechanism. Increase in [Ca2+]mito has been shown to induce apoptosis by alterations in mitochondrial membrane permeability, formation of the permeability transition pore, cytochrome c release, and activation of caspase-9 and -3 and disruption of ΔΨ. We could demonstrate that immediately after onset of hypocapnia ΔΨ increases. However, we have no data on ΔΨ 24h after hypocapnia because TMRM measurements are useful only for a short time period. There are reports that especially Ca2+ coming from the ER might be one trigger for apoptosis. As stated above, this way of apoptosis induction might be possible in our scenario but we were not able to proof this hypothesis.
We could show that hypocapnia is associated with mitochondrial ROS production. As this response could be inhibited by rotenone we conclude that the hypocapnia-induced ROS was produced in the respiratory chain. In addition, the mitochondrial ROS production could be inhibited by ruthenium red. We therefore conclude that, at least in part, the hypocapnia-induced mitochondrial Ca2+ uptake was responsible for the ROS response. Although the Ca2+-dependent mitochondrial ROS production is well established we cannot give any proof for the mechanism of the Ca2+-dependent ROS production. Theoretically, it could be the case that the hypocapnia-induced increase of NADH production results in a higher amount of ubisemiquinone on complex III of the respiratory chain. This might statistically increases the probability that the electron will instead be donated to O2, generating superoxide. Furthermore, a reverse electron transport may occur under hypocapnia and be responsible for the hypocapnia-induced ROS production. Moreover, we cannot differentiate whether apoptosis was evoked directly by Ca2+, ROS, or both of them.
In summary, we identify a novel CO2-sensing mechanism of AEC type 2 in which the mitochondrial TCA enzyme IDH3 has a key role. Under hypocapnic conditions, IDH3 activity and thus NADH production increases, which subsequently induces mitochondrial Ca2+ uptake and ROS production leading to apoptosis. We propose that this mechanism plays an important role in the pathogenesis of ARDS because alveolar hypocapnia may occur in lung regions with high VD/VT. According to our results one could consider a novel ARDS therapy, namely CO2 inhalation in order to prevent hypocapnia-induced apoptosis of AECs and tissue injury in ARDS.
Male Sprague-Dawley rats (350g) were purchased from Charles River (Sulzfeld, Germany). After review and approval by the local ethics committee and the government of Hamburg, all animals were treated humanely in accordance with 'principles of laboratory animal care' (NIH Publication No. 86-23, revised 1985) as well as with the German legislation on protection of laboratory animals.
Fura 2-AM, LysoTracker green (LTG), MTG, rhod-2, 2′, 7'-bis-(2-carboxylmethyl)-5-(and 6-) carboxyfluorescin-AM and TMRM were purchased from Molecular probes (Thermo Fisher Scientific, Waltham, MS, USA). Both rotenone and ruthenium red were purchased from Sigma-Aldrich (St. Louis, MO, USA). Vehicle for dyes and other agents was HBSS buffer (150 mmol/l Na+, 5 mmol/l K+, 1.0 mmol/l Ca2+, 1 mmol/l Mg2+, and 20 mmol/l HEPES at pH 7.4). All esterified fluorescent probes were prepared as stock solutions in dimethylsulfoxide.
Isolation and culture of AEC type 2 from adult male Sprague-Dawley rats according to procedures described previously.20, 21 In brief, after injection of 100mg/kg ketamine and 20mg/kg xylazinum (rompun), the lungs were surgically exposed and perfused with 3 × 10mlF-12K medium (ATCC, Manassas, VA, USA) supplemented with 25mM HEPES (Sigma-Aldrich) to remove blood. After multiple lavages (5 × 10ml) via the cannulated trachea with 5mM ethylenediaminetetraacetic acid (EDTA, Sigma-Aldrich) and 5mM ethylene glycol-bis (ß-aminoehtylether)-N,N’,N’-tetraacetic acid (Sigma-Aldrich) in a phosphate-buffered salt solution the lungs were removed from the body. Upon instillation of 20ml warm F-12K medium (ATCC) enriched with 50mM HEPES and 4.5 units/ml elastase Grade II (Worthington, Lakewood NJ, USA) the lungs were incubated at 37°C for 30min. After dissection of the large airways the lungs were added to 10ml F-12K medium containing 25mM HEPES, 20% fetal bovine serum (FBS, Biochrom AG, Berlin, Germany), and 100μg/ml DNAse (Typ IV, Sigma-Aldrich) quickly minced and incubated at 4°C for 10min. Then the solution was mixed by end-over end-rotation for 4min and filtered successively through cotton gauze (1, 2, and 4 ply, respectively) following filtration through nylon mesh (100 and 40μm, BD Falcon, Corning, NY, USA). Upon centrifugation of the cell suspension at 150 × g for 10min and resuspension in 20ml warm DMEM (Biochrom AG) the cells were applied to plates covered with rat immunoglobulin (Ig) G (Sigma-Aldrich) for 1h at 37°C. The nonadherent cells were removed, collected in sterile tube, twice centrifuged at 150 × g for 10min and resuspended in 3ml of culture medium (DMEM with 10% FBS, 1% penicillin/streptomycin and 10ng/ml keratinocyte growth factor (Sigma-Aldrich)). After determination of cell count and viability, cells were plated on collagen-treated coverslips and incubated at 37°C and 5% CO2.
Human lung carcinoma A549 cells were grown in F-12K Medium (ATCC) with 10% FBS, 10000U/ml penicillin and 10000μg/ml streptomycin (Biochrom AG). A549 is an alveolar epithelial cell line with type 2 cell characteristics including production of surfactant proteins. It has been widely used as a model to study the response of AEC in several conditions.22, 23
For fluorescent imaging, a coverslip with fluorophore-loaded primary isolated AEC type 2 or A549 cells was mounted in an imaging/perfusion chamber on the stage of a conventional epifluorescence microscope (Olympus, Hamburg, Germany). The imaging chamber was perfused continuously with warm HEPES buffer (37°C).
Fluorophores were excited by a mercury arc lamp illumination directed through appropriate interference filters and filter sets (Semrock, Rochester, NY, USA). Fluorophore exposure was controlled by a filter wheel (Sutter Lambda 10-C, Sutter Instrument Co). The fluorescence emission was collected using an objective lens (40 × water immersion, numerical aperture 0.8, Zeiss, Oberkochen, Germany) and captured with a charge-coupled device camera (Photometrics Coolsnap HQ2, Tuscan, AZ, USA).
[Ca2+]cyt determination. For [Ca2+]cyt measurements, cells were loaded for 45min with 10μM fura 2-AM in physiological solution containing 0.02% pluronic. The cells were excited at 340 and 380nm. The fluorescence emissions at 510nm were recorded and [Ca2+]cyt was calculated from a computer-generated 340:380 emissions ratio based on a dissociation constant of 224nmol/l and appropriate calibration parameters.24
Fluorescence images were recorded under normo- or hypocapnic conditions in combination with an extracellular pH of 7.4 or 7.8 at 10s intervals.
[Ca2+]mito determinations. For [Ca2+]mito measurements, cells were loaded with rhod-2-AM for 15min followed by a Ringer's flush. To improve the specifity of [Ca2+]mito measurement mitochondria were additionally localized by the fluorophore MTG, which binds specifically to the inner mitochondrial membrane.25 The wavelength for excitation was 560nm, whereas monitoring the emission at 590nm.
For mitochondrial membrane potential assessment, primary isolated AEC type 2 or A549 cells were loaded with TMRM. This indicator dye is a lipophilic cation accumulated by mitochondria in proportion to the electrical potential across their inner membrane (Δψ).26 The wavelength for excitation was 560nm while monitoring the emission at 590nm. Addition of the mitochondrial depolarizer FCCP (50nM) resulted in a rapid loss of the mitochondrial TMRM fluorescence.
NADH concentration was quantified by measuring its auto-fluorescence intensity. Primary isolated AEC type 2 or A549 were excitated with 360nm while monitoring the emission at 510nm. For determining the mitochondrial production of NADH, its auto-fluorescence was co-localized with the mitochondria-specific marker MTG.
Mitochondrial ROS production was measured using MitoSOX Red (Thermo Fisher Scientific). Primary isolated AEC type 2 were stained with 10μM MitoSOX Red and 10μM MTG (staining of mitochondria) for 10min at 37°C. Upon washing with normocapia buffer, cells were then imaged with a conventional epifluorescence microscopy (Olympus) in normo- and hypocapnia puffer, respectively. Changes in the fluorescence intensity of MitoSOX Red were measured at 550nm excitation and 600nm emission wavelengths. In an additional set of experiments we preincubated the cell with 1μM rotenone to inhibit Complex I of the electron chain and/or 10μM ruthenium red to inhibit the MCU. As a positive control we used Antimycin AA (2μg/ml).
We used the Cell Event Caspase-3/7 Green Detection Reagent (Thermo Fisher Scientific) according to the manufacture’s instruction to determine apoptotic cells. In brief, upon incubation with normo- und hypocapnia puffer for 30min at 37°C primary isolated AEC type 2 were stained 24h later with the caspase-3/7 reagent (10μM) and counterstained with the nuclei dye Hoechst 33342 (0.5μg/ml). Cells were then image with a conventional epifluorescence microscopy (Olympus) and caspase-3/7-positive cells were counted in four independent fields per well.
To determine changes of NADH / NAD(P)H during inhibition of complex I and/or under hypocapnia treatment, FLIM was performed on a single-beam 2-photon microscope setup described elsewhere.27 In brief, the excitation wavelength of the pulsed Ti:Sa Laser (MaiTai, Spectra Physiscs) was 750nm and sent through a water-dipping lens (20 ×, NA 0.95, WD 2 mm – Olympus) to the lung surface. Emission photons passed a band-pass filter (475/47nm) and were finally detected at the Ultra-sensitive-port by a 16-channel time-correlated single photon counter, with a maximum rate of 78 MHz (LaVision Biotec, Bielefeld, Germany). Evaluation of the multi-exponential fluorescence signal was performed by the self-developed software RINIFLIM programmed in Matlab (MathWorks). For evaluation, Levenberg-Marquardt algorithms were used.
The NAD+-dependent IDH3 activity was determined using the commercial available IDH activity assay kit according to the manufactures protocol (Sigma-Aldrich). In brief, the IDH3 activity in cell extracts was measured using the substrate isocitrate and the cofactor NAD+ in an enzyme reaction, which results in a colorimetric (450nm) product proportional to NADH concentration. The enzyme activity was calculated according to the standard curve. One unit of IDH3 is the amount that will generate 1.0μmol NADH per minute at pH 8.0 at 37°C.
A549 cells were transfected with 10nM stealth siRNA of IDH3 (Thermo Fisher Scientific) and/or 10nM siRNA of MCU (Quiagen, Hilden Germany) and scrambled siRNA (Thermo Fisher Scientific and Quiagen) by using Lipofectamine RNAIMAX (Thermo Fisher Scientific) according to the manufacturer’s protocol. In brief, siRNA was diluted in 500μl Opti-MEM I medium (Quiagen) without serum in a well of the tissue culture plate. Lipofectamine RNAiMAX was added to each well containing the diluted siRNA. Cells were diluted in complete growth medium without antibodies so that the cell density after 24h would be 30–50% confluent after plating.
Total RNA was isolated using the RNeasy Kit (Quiagen). A total of 2μg of RNA were used for cDNA synthesis using the Maxima First Strand cDNA Synthesis Kit (Thermo Fischer Scientific) according to the manufacturer's instructions. qRT-PCR was performed with gene- and species-specific primers. cDNA template was amplified using the QuantiNova SYBR GreenPCR Kit (Quiagen) on a Rotor-Gene (Quiagen) according to the manufacture’s protocol. Quantitative analysis of data were performed by using the deltadeltaCT method.
For immunoblotting, cells were washed twice with PBS and then scraped into RIPA buffer (Sigma-Aldrich) supplemented with the protease inhibitor cocktail complete Mini EDTA-free (Roche Diagnostics, Mannheim, Germany). Upon incubation on ice for 30min lysed cells were centrifuged at 12000 × g for 20min (4°C). The supernatants with the cytoplasmatic protein were collected, and protein concentrations were determined by using the bicinchoninic acid (BCA protein assay kit, Thermo Fisher Scientific), assay according to the manufacturer’s instructions. A total of 20μg samples were reduced in sample buffer (NuPAGE sample buffer, Thermo Fisher Scientific) plus reducing agent (NuPAGE sample reducing agent, Invitrogen) at 95°C for 5min and run on a 4–12% Bis-Tris polyacrylamid gel (NuPage, Thermo Fisher Scientific) and transferred to nitrocellulose filters (Thermo Fisher Scientific). The filters were blocked for 1h at room temperature in blocking puffer (PBS plus 0.1% Tween 20 containing 5% milk powder and/or 5% bovine serum albumin (BSA)) with agitation. After application to the polyclonal rabbit IDH3A antibody (1:800, GenWay Biotech Inc., San Diego, CA, USA) or to the monoclonal rabbit MCU antibody (1:1000, Cell Signaling Technology, Danvers, MA, USA) overnight at 4°C, the filters were exposed to an ECL horseradish peroxidise-conjugated donkey anti-rabbit IgG (1:10000 and/or 1:20000), GE Healthcare, Little Chalfont, UK) for 1h at room temperature with agitation. After washing with washing puffer (PBS plus 0.1% Tween 20), the filters were developed with the ECL Plus Western Blotting Detection Reagents (GE Healthcare) according the manufacture’s protocol. For molecular weight standards, we used a MagicMark XP protein ladder (Thermo Fisher Scientific).
Upon incubation with normo- und hypocapnia puffer for 3h at 37°C primary isolated AEC type 2 were collected 24h later and prepared for immunoblotting as described in the section 'Western blot IDH3 and MCU knockdown analysis’. After gelelectrophoresis and immunoblotting, filter were blocked for 1h at room temperature in blocking puffer (PBS plus 0.1% Tween 20 containing 5% BSA) with agitation. Upon application of the monoclonal rabbit Anti-cleaved-PARP antibody (1:500, Abcam, Cambridge, UK) overnight at 4°C, the filters were exposed to an ECL horseradish peroxidise-conjugated goat anti-rabbit IgG (1:5000, Jackson ImmunlResearch Labaroties, Inc, West Grove, PA, USA) for 1h at room temperature with agitation. After washing with washing puffer (PBS plus 0.1% Tween 20), the filters were developed with the ECL Plus Western Blotting Detection Reagents (GE Healthcare) according the manufacture’s protocol. For molecular weight standards, we used a MagicMark XP protein ladder (Thermo Fisher Scientific). As a positive control, we used staurosporine (1μM) treated cells.
All data are represented as means±S.E. Statistical data analysis was performed using SigmaStat (Systat Software, Chicago, IL, USA). Comparisons between two groups were tested using RankSumTest. Comparisons between more groups were tested by ANOVA on ranks followed by a pairwise multiple comparison. Repeated measurements were tested using the Mann–Whitney Rank Sum Test. Statistical significance was accepted at P<0.05.
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We wish to thank Kirsten Pfeiffer-Drenkhahn, Andrea Pawelczyk, Claudia Luechau, and Monika Weber for their great technical assistance. Without it, this work would not have been possible. This research work was funded by a grant from Else Kröner-Fresenius Stiftung (grant number 2011_A207) and Johanna und Fritz Buch-Gedächtnis-Stiftung.
Supplementary Information accompanies this paper on Cell Death and Disease website (http://www.nature.com/cddis)
Edited by P Agostinis
The authors declare no conflict of interest.