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Published ahead of print on September 23, 2004, doi:10.1165/rcmb.2004-0202OC
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American Journal of Respiratory Cell and Molecular Biology. Vol. 32, pp. 44-51, 2005
© 2005 American Thoracic Society
DOI: 10.1165/rcmb.2004-0202OC

Hypoxia Decreases Cellular ATP Demand and Inhibits Mitochondrial Respiration of A549 Cells

Kristin Heerlein, Andreas Schulze, Lorenz Hotz, Peter Bärtsch and Heimo Mairbäurl

Medical Clinic VII, Sports Medicine, University of Heidelberg; and Division of Metabolic and Endocrine Diseases, Department of General Pediatrics, University Children's Hospital, Heidelberg, Germany

Correspondence and requests for reprints should be addressed to Heimo Mairbäurl, Medical Clinic VII, Sports Medicine, University of Heidelberg, INF 410, 69120 Heidelberg, Germany. E-mail: heimo.mairbaeurl{at}med.uni-heidelberg.de


   Abstract
Top
 Abstract
Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
Hypoxia inhibits activity and expression of transporters involved in alveolar Na reabsorption and fluid clearance. We studied whether this represents a mechanism for reducing energy consumption or whether it is the consequence of metabolic dysfunction. Oxygen consumption (JO2) of A549 cells and primary rat alveolar type II cells was measured by microrespirometry during normoxia, hypoxia (1.5% O2), and reoxygenation. In both cell types, acute and 24-h hypoxia decreased total JO2 significantly and reoxygenation restored JO2 after 5 min but not after 24 h of hypoxia in A549 cells, whereas recovery was complete in type II cells. In A549 cells under normoxia Na/K-ATPase accounted for ~ 15% of JO2, whereas Na/K-ATPase–related JO2 was decreased by ~ 25% in hypoxia. Inhibition of other ion transporters did not affect JO2. Protein synthesis–related JO2 was not affected by acute hypoxia, but decreased by 30% after 24-h hypoxia. Acute and 24-h hypoxia decreased JO2 of A549 cell mitochondrial complexes I, II, and III by 30–40%. Reoxygenation restored complex I activity after acute hypoxia but not after 24-h hypoxia. ATP was decreased 30% after 24-h hypoxia, but lactate production rate was not affected. Reduced nicotinamine adenine dinucleotide was slightly elevated in acute hypoxia. Our findings indicate that inhibition of the Na/K-ATPase by hypoxia contributes little to energy preservation in hypoxia. It remains unclear to what extent hypoxic inhibition of mitochondrial metabolism affects ATP-consuming processes.

Key Words: energy metabolism • hypoxia • mitochondrial electron transfer chain • Na-transport • respirometry


   Introduction
Top
 Abstract
 Introduction
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 
A tight coupling exists between oxygen consumption (JO2) and energy demand. A decrease in ATP-utilizing pathways translates, therefore, into a decrease in cellular respiration rates and oxygen demand. This strategy is applied by a variety of hypoxia-tolerant species, such as hibernators, turtles, and various fish, when they experience tissue hypoxia (1). Adjustments might be the result of low inspiratory and tissue oxygen pressures due to pulmonary edema, as in disease states characterized by ventilatory and cardiovascular insufficiency, ischemia, by a decreased oxygen transport capacity, and during exposure to high altitude. A decrease in oxygen demand is achieved by decreasing the activity of major ATP-consuming pathways, such as ion pumping and protein synthesis (2, 3). Also, an upregulation of the energetic efficiency of ATP-producing pathways has been described previously (4).

One such mechanism might be hypoxia-induced inhibition of Na transport in cultured alveolar epithelial cells and an inhibition of ion transport and a reduced rate of water reabsorption in the lung in hypoxia (5–7), which has been suggested to enhance the formation of hypoxic pulmonary edema (8) because a high capacity for fluid reabsorption has been associated with recovery from pulmonary edema (9). In contrast, when the diffusing capacity of nonperfused dog lungs was measured after 30 to 60 min of recovery from several hours of hypoxia, there was no observed difference from control conditions (10).

Long-term adjustments to hypoxia involve changes in gene expression that are aimed at an improvement of the tissue oxygen and energy supply, and which is mainly stimulated by the hypoxia-inducible factor (HIF)-1{alpha} encoding for bcl-2, Bax, erythropoietin, vascular endothelial growth factor, and glycolytic enzymes, such as glyceraldehyde-phosphate dehydrogenase (11). However, the overall rate of protein synthesis is reduced (1). Examples of proteins, the expression of which is decreased, are ß-actin (12) and transport molecules, such as Na/K-ATPase, cystic fibrosis transmembrane regulator, and ENaC (5, 13).

Ischemic or hypoxic cell injury is often due to structural alterations and dysfunction of mitochondria (14). Mitochondrial metabolism of hepatocytes was found to be decreased by hypoxia, a process termed "oxygen conformance" (15). In addition, Taylor and colleagues (16) reported an almost instant decline in mitochondrial respiration upon hypoxia of hepatocytes. It has further been shown that expression of both mitochondrial and nuclear genome–encoded proteins is downregulated by hypoxia (17) and that hypoxia causes damage in mitochondrial DNA (18).

It was the aim of this study to find out whether hypoxic ion transport inhibition of alveolar epithelial cells is aimed at reducing the cellular JO2 by decreasing the activity of ATP-consuming processes, such as protein synthesis and ion transport in this state of limited oxygen supply. Because oxygenation of the alveolar epithelium in vivo is the highest among all tissues, care was taken to study cellular metabolism by microrespirometry within a range of PO2 where the oxygen supply to mitochondria is not limited.


   MATERIALS AND METHODS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 References
 
Chemicals/Reagents
Ham's F-12 medium, penicillin/streptomycin, fetal calf serum (FCS), and N-2-hydroxyethylpiperzine-N'-ethanesulfonic acid (HEPES) were from Gibco (Eggenstein, Germany). Ouabain, malate, pyruvate, succinate, ADP, rotenone, antimycin A, and sucrose were from Sigma-Aldrich (Munich, Germany). Cycloheximide, mannitol, digitonin, and bovine serum albumin were from Serva Chemicals (Heidelberg, Germany). Decylubiquinone was a gift from Dr. Jürgen G. Okun (University Children's Hospital, Heidelberg, Germany).

Cells
A549 cells (American Type Culture Collection), which are similar in many functions to alveolar type II cells (19), passages 81–90, were grown in flasks on Ham's F-12 medium supplemented with 7% FCS, penicillin/streptomycin, 10 mM HEPES, and 15 mM sodium bicarbonate. Ham's F-12 medium is well suited for measurements of cell metabolism, as it contains a wide spectrum of substrates for energy metabolism. In normoxia, A549 cells were kept in an incubator (Heraeus, Hanau, Germany) at typical tissue culture conditions (37°C; room air with 5% CO2). Cells were exposed to hypoxia (37°C; 2% O2 in N2) for 24 h in a modular incubation chamber (Billups Rothenberg, Del Mar, CA).

Confluence was reached after ~ 3–4 d in culture and all experiments were performed on 5- to 8-d-old confluent A549 cells, which were trypsinated and spun at 600 rpm for 10 min. Cells from one 75 cm2 flask were suspended in 1.8 ml Ham's F-12 medium supplemented with 10 mM HEPES.

Rat primary alveolar type II cells were prepared as previously described (7). Briefly, lungs from rats anesthetized by intraperitoneal injection with 100 mg/kg pentobarbital (Trapanal; Byk Gulden, Konstanz, Germany) were perfused with phosphate-buffered saline while being ventilated with air. Alveolar epithelial type II cells were isolated by elastase digestion, mincing of lung tissue, filtration, and differential adhesion on immunoglobulin (Ig) G–coated plates (6). Nonadherent cells were suspended in Dulbecco's modified Eagle's medium supplemented with 10% FCS, glutamine (4 mM), and gentamicin (50 µg/ml), and plated on 6-well plates (Transwell, Costar, Cambridge, MA) at a seeding density of 4 x 105 cells/cm2. Both purity and viability of alveolar epithelial type II cells were > 85%. Cells were maintained in normoxia (room air supplemented with 5% CO2) until they had reached confluence (typically on Day 3 after plating). For exposure to hypoxia, confluent monolayers were placed in an incubator, adjusted to 1.5% O2 and 5% CO2 at 37°C for 24 h. Hypoxia did not affect cell viability, as measured by trypan blue exclusion, compared with that observed for normoxic cells.

High-Resolution Respirometry
Rates of JO2 were measured in a high-resolution respirometer (Oxygraph; OROBOROS Instruments, Innsbruck, Austria) at 37°C. DatLab software (OROBOROS Instruments) was used for data acquisition. Cellular JO2 was calculated from the recorded data as the time derivative of the oxygen content in the chamber, which was corrected for the response time of the oxygen sensor and for instrumental background oxygen flux (back-diffusion of oxygen, JO2 by the oxygen sensor). JO2 was corrected for protein to account for differences in the number of cells among experiments. Protein was measured with a test kit from BioRad (Hercules, CA). Exposure of the cells to shear stress by stirring at 480 rpm in the respirometer for up to 2 h did not affect respiration rates nor vitality as measured by trypan blue exclusion.

After heating at 37°C and equilibration with room air 2 ml medium in the chamber (Ham's-12 buffered with 10 mM HEPES), 0.5 ml of the cell suspension was added. PO2 was lowered to ~ 100 mm Hg by gassing with humidified N2. The system was then closed to start the JO2 measurement in normoxia (PO2 range, 70–85 mm Hg). A typical profile of JO2 and O2 content in the chamber is shown in Figure 1A. Next, the chamber was opened and the cell suspension was equilibrated with a humidified gas composed of 2% O2 and 98% N2 (hypoxia), which took ~ 5 min to reach equilibrium. The system was then closed to determine JO2 in the PO2 range of 12–9 mm Hg (Figure 1A). In this range of PO2, mitochondrial respiration was not oxygen-limited (Figure 1B). For reoxygenation, the system was opened again for 5 min to raise PO2 to ~ 100 mm Hg, and JO2 was measured after this short-term reoxygenation. When JO2 was measured on cells that had been exposed to hypoxia for 24 h, the medium in the respirometer was equilibrated with a gas composed of 2% O2 and 98% N2 before adding the cells. JO2 was then measured under hypoxic conditions and after short-term reoxygenation only (Figure 1A). Control cells for the 24-h hypoxia experiment were cells from the same batch that was measured on the same day as the hypoxia-exposed cells.



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  		Figure 1. (A) Representative recording of medium PO2 and JO2 in normoxia, acute hypoxia, and reoxygenation; (B) change in JO2 with PO2. (A) Typical tracing of changes in PO2 (solid line) during an experiment. The medium was equilibrated to room air, cells were added (a), and the oxygen content of the medium was adjusted by gassing with N2 to obtain the desired PO2 (b) when the system was closed (c) to record JO2,N in normoxia (PO2 75–90 mm Hg). The system was opened again to equilibrate with gas containing 2% O2 and 98% N2 (hypoxia; d), and to record JO2,H in acute hypoxia (e) in the closed chamber, opened again and equilibrated with room air to reoxygenate to a PO2 of ~ 90 mm Hg (f) and record JO2,R (reoxygenation; g), and equilibrated with N2 to obtain the zero-point calibration (h). Transient changes in JO2 during step changes in PO2 have been removed. (B) Medium was equilibrated with room air before adding the cells when the system was closed. JO2 was recorded until it dropped to zero. The dark shaded areas on the curve indicate the range in which JO2 was recorded in normoxia, hypoxia, and upon reoxygenation. The steep decrease in JO2 at low PO2 values indicates oxygen limitation of mitochondria.

 
To study the effects of inhibition ion transport on JO2, ouabain (0.1 mM; activity of the Na/K-ATPase [JO2,{Delta}ouab]), amiloride (10 µM), benzamil (1 µM), bumetanide (50 µM), and KB-R7943 mesylate (mesylate; 2-[2-[4-(4-Nitrobenzyloxy)phenyl]ethyl]isothiourea mesylate; 50 µM) were present in the medium. Cycloheximide (2.5 mM) was used to determine JO2 related to protein synthesis. Inhibitors were added to the medium before the addition of A549 cells. In case of cycloheximide, a commonly used inhibitor of protein synthesis, when JO2 was measured (as in Ref. 20) pretreatment of the cells with the inhibitor for 1 h was required to achieve full inhibition. Experiments with inhibitors were performed in a paired manner with control cells in Chamber A of the respirometer and inhibitor-treated cells in Chamber B.

Permeabilized Cells
The nonionic detergent, digitonin, was used to solubilize membrane-bound proteins, which make cells permeable to metabolites required to measure the activity of the mitochondrial electron transport chain (mETC) (4, 21). The permeabilization medium, which was also used to measure ETC activity, contained 75 mM sucrose, 100 mM KCl, 10 mM KH2PO4, 0.5 mM EDTA, 5 mM MgCl2, 20 mM TRIS-HCl, and 1 mg/ml bovine serum albumin (fatty acid–free). In titration experiments (data not shown), permeabilization was visualized by trypan blue staining, and it was determined that the optimal digitonin concentration for permeabilization was 40 µg/106 cells. The suspension (0.5 ml) containing the digitonin-permeabilized cells was transferred to the respirometer, where 2 ml of medium had been equilibrated to the respective gas, thereby diluting digitonin 5-fold. Substrates and inhibitors of mitochondrial respiration were added as required to measure the activity of the mETC complexes I, II, and III at the following concentrations: 2 mM malate; 10 mM pyruvate; 2 mM ADP; 2.5 µM rotenone; 10 mM succinate; 50 µM decylubiquinone; 1 mM antimycin A.

Measurements of Enzyme Activity and Concentration of Metabolites
Citrate synthase, a mitochondrial matrix enzyme, is commonly used as a marker for content of intact mitochondria (22). Its enzyme activity was measured in A549 cells that had either been exposed to normoxia or to hypoxia for 24 h.

Lactate and ATP were measured from perchloric acid extracts of cells exposed to normoxia and acute hypoxia in the respirometer. Lactate was measured spectrophotometrically using a test kit from Sigma-Aldrich. ATP was measured by chemiluminescence with a luciferin/luciferase assay (Roche, Mannheim, Germany).

Relative changes in reduced nicotinamine adenine dinucleotide (NADH) were determined using a fluorescence microscopy (23, 24). Briefly, A549 cells grown to confluence on untreated 1-cm2 glass cover slips were placed in a perfusion chamber (POC; Saur, Reutlingen, Germany). This chamber is covered with a glass slide and allows oxygen-controlled perfusion. The perfusion medium was composed of 140 mM NaCl, 20 mM HEPES, 10 mM glucose, 5 mM KCl, 1 mM NaH2PO4, 1 mM MgCl2, and 0.2 mM CaCl2, pH 7.4 at room temperature and was equilibrated with the appropriate gas. Bubbling with pure nitrogen resulted in a PO2 of 10 mm Hg in the perfusion chamber (hypoxia). The perfusion rate was ~ 1.5 ml/min. Measurements were performed at room temperature using an inverted microscope (Axiovert 35; Zeiss, Göttingen, Germany) equipped with a high-pressure mercury short-arc lamp (HBO 100 W/2; Osram, Germany). The excitation wavelength was 340 nm, and emission was measured at 510 nm by photon counting (photomultiplier; Thornton EMI Gencom Inc., Ruislip, UK) and recorded on a computer (Mega STE; Atari Corporation, Frankfurt, Germany).

Statistical Analysis
Results are shown as mean values ± SD of the number of experiments indicated in the figure legends. Statistical analysis was done by Student's t tests, one-way analysis of variance and Tukey's post hoc tests using SigmaPlot and SigmaStat (SPSS Science, Erkrath, Germany). Level of significance was P <= 0.05.


   RESULTS
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 References
 
Intact Cells
Average JO2 of A549 cells in normoxia was 290.7 ± 60.3 µmol/s/mg protein and that of primary rat alveolar type II (AII) cells was only about one third of this value. Figure 2 shows that total JO2 had decreased in acute hypoxia by ~ 25% in A549 cells and 33% in AII cells. After reoxygenation, 91% of the normoxic JO2 values were regained in A549 cells, whereas AII cells recovered fully. In cells exposed to hypoxia for 24 h, JO2 decreased by ~ 35% in comparison with normoxic control cells, whereas in AII cells the decrease was 45%. Reoxygenation after 24 h hypoxia caused no recovery of JO2 in A549 cells, whereas in AII cells complete recovery was seen.



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  		Figure 2. Effects of acute and 24-h hypoxia on JO2 of A549 cells (A) and primary rat alveolar type II cells (B). Measurement of total JO2 was performed as described in legend to Figure 1. Mean values ± SD are shown as percent of normoxic control cells (A549 cells: acute hypoxia, n = 12; 24 h of hypoxia, n = 6; AII cells: n = 10). *P < 0.05 as compared with normoxic controls; #P < 0.05 between hypoxia and reoxygenation.

 
In normoxia, JO2,{Delta}ouab amounted to ~ 15% of total JO2. Figure 3A shows that both in acute and 24-h hypoxia, JO2,{Delta}ouab had decreased to ~ 11% of the total JO2 of normoxic control cells, which corresponds to a decrease of ~ 30% in Na/K-ATPase–related JO2. Only after acute hypoxia did short term reoxygenation restore initial JO2,{Delta}ouab. JO2 was also measured in the presence of inhibitors of secondary active Na transporters and channels, such as Na/K/2Cl cotransport, Na/H exchange, Na/Ca exchange, and epithelial Na channels. However, none of the inhibitors used affected JO2 (data not shown).



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  		Figure 3. Effects of acute and 24-h hypoxia on (A) Na/K-ATPase and (B) protein synthesis–dependent JO2. Na/K-ATPase–related JO2 was measured in the absence and presence of 100-µM ouabain (JO2,{Delta}ouab). JO2 related to protein synthesis was measured after equilibration of A549 cells with cycloheximide (2.5 mM; JO2,{Delta}cycl) for 1 h. This long equilibration was required to achieve complete inhibition. Mean values ± SD of inhibitor-sensitive JO2 relative to total JO2 of untreated, normoxia-exposed cells of ouabain (n = 6 ) and cycloheximide (n = 9) experiments. *P < 0.05 as compared with normoxic control cells.

 
Cycloheximide, an inhibitor of protein synthesis, reduced JO2 in a time-dependent manner, reaching maximal inhibition after 1 h of exposure to the inhibitor (not shown). Therefore, measurements were performed after cycloheximide pretreatement of the cells in suspension. Cycloheximide blocked ~ 20% of A549 cell JO2 in normoxia (Figure 3B). JO2,{Delta}cycl decreased significantly, by ~ 27% in acute hypoxia and by ~ 60% after 24 h of exposure to hypoxia. Although upon short term hypoxia, normoxic JO2,{Delta}cycl was fully restored by short-term reoxygenation, it did not recover upon reoxygenation of cells that had been exposed to hypoxia for 24 h.

The ouabain- and cycloheximide-insensitive (residual) JO2 (JO2,res) was decreased significantly by exposure to acute (–28%) and 24-h hypoxia (–61%). JO2,res recovered after acute hypoxia, but did not recover in cells exposed to hypoxia for 24 h (data not shown), indicating the presence of additional hypoxia-sensitive metabolic pathways.

Mitochondrial Respiration
To study whether hypoxia affected the function of the mETC, JO2 was also measured in digitonin-permeabilized cells with substrates chosen to establish ADP-stimulated (state 3) respiration of mETC complexes I, II, and III. Results are summarized in Figure 4.



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  		Figure 4. Hypoxia inhibits mitochondrial JO2. JO2 was measured on digitonin-permeabilized A549 cells (40 µg/106 cells) that were cultured in normoxia and hypoxia (1.5% O2, 24 h). Permeabilized cells were suspended, and JO2 was measured in a medium composed of 75 mM sucrose, 225 mM mannitol, 10 mM KCl, 5 mM KH2PO4, 10 TRIS-HCl and 2 malate, as well as 1 mg/ml bovine serum albumin (fatty acid–free). State 3 respiration was initiated by adding ADP (2 mM). JO2,C1 was measured in the presence of 10 mM pyruvate, JO2,C2 with succinate (10 mM) in the presence of rotenone (2.5 µM); substrate for JO2,C3 was decylubiquinone (50 µM). Mean values ± SD (n = 6) are presented. *P < 0.05 as compared with normoxic controls; #P < 0.05 between hypoxia and reoxygenation.

 
Complex I respiration (JO2,C1) was initiated by adding ADP to permeabilized cells in a suspension that also contained pyruvate and malate as metabolic substrates (state 3 respiration), which was 1.4-fold higher than the ADP-limited state 4 respiration. In normoxic control cells, state 3 JO2,C1 was 108.7 ± 59.7 pmol · s–1 · mg protein but showed quite large variability. Acute hypoxia decreased JO2,C1 by ~ 40% (Figure 4A). During reoxygenation upon acute hypoxia, JO2,C1 increased again and reached ~ 78% of normoxic control values. In cells exposed to hypoxia for 24 h, JO2,C1 was decreased by ~ 40%, but reoxygenation did not cause recovery.

Complex II respiration (JO2,C2) was measured in the presence of rotenone and succinate, and was started by addition of ADP, which stimulated respiration 2.5 ± 0.6–fold. JO2,C2 was 632.1 ± 87.8 pmol · s–1 · mg protein in normoxic control cells. Hypoxia decreased JO2,C2 by ~ 30%, and reoxygenation values were not significantly different from hypoxia (Figure 4B). After 24-h hypoxia, the JO2,C2 was reduced by only ~ 15%. In this case, also, reoxygenation failed to restore respiration.

Decylubiquinone was used as a specific substrate for mETC complex III. Because of its hydrophobic nature, which allows measurement of JO2 for a short time period only, paired measurements were not possible. JO2,C3 in normoxic cells was 773.9 ± 95.1 pmol · s–1 · mg protein. After acute hypoxia, JO2,C3 decreased by 37% of normoxic levels; reoxygenation did not recover control values of JO2,C3 (Figure 4C). Exposure to hypoxia for 24 h decreased JO2,C3 by ~ 51% of that observed in normoxia. Again, there was no recovery upon reoxygenation.

Citrate synthase activity was measured in A549 cells exposed to normoxia and hypoxia for 24 h to explore whether the decreased respiration in hypoxia was due to a decrease in the number of mitochondria (22). Figure 5 shows that, in fact, citrate synthase activity had decreased by ~ 23% after a 24-h exposure to hypoxia.



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  		Figure 5. Effects of hypoxia on citrate synthase activity. Citrate synthase activity was determined as a measure of the number of mitochondria in A549 cells exposed to hypoxia for 24 h. Mean values ± SD (n = 6). *P < 0.05 compared with normoxia.

 
Extramitochondrial JO2
Due to the presence of oxygen-consuming oxidases and oxygenases in the cytosol and in the plasma membrane, JO2 is not restricted to mitochondria. It was therefore important to determine the contribution of extramitochondrial JO2 (JO2,ex) to the decrease of JO2 in hypoxia. Inhibition of mitochondrial respiration with rotenone abolished ~ 75% of JO2 in normoxic cells. At levels of PO2 higher than 3 mm Hg, JO2,ex decreased almost linearly with PO2, whereas, at lower PO2, JO2,ex decreased more rapidly (data not shown). Figure 6 shows that during acute hypoxia, JO2,ex had decreased to ~ 20% of the total JO2 of normoxic cells and was decreased to ~ 15% in cells that had been exposed to hypoxia for 24 h. However, in normoxia as well as in acute and chronic hypoxia, the proportion of JO2,ex was always ~ 75% of the respective total JO2.



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  		Figure 6. Effects of hypoxia on extramitochondrial JO2 of A549 cells. JO2 was measured in normoxia as well as in acute and 24-h hypoxia in A549 cells in the absence and presence of rotenone (2.5 µM). Mean values ± SD (n = 6) of total JO2 and rotenone-insensitive JO2 (% JO2 of normoxic control cells) are presented. *P < 0.05 compared with normoxic controls; #P < 0.05 between acute and 24 h hypoxia. Black bars, total; gray, hatched bars, rotenone.

 
Metabolic Substrates
It was further tested whether the hypoxia-induced decrease in mETC activity shown in Figure 4 affects the concentration of metabolic intermediates. The cellular ATP concentration (Figure 7A) did not change significantly during acute hypoxia and subsequent reoxygenation, whereas it had decreased significantly by ~ 30% after exposure of cells to hypoxia for 24 h. In the latter, 15 min of reoxygenation were insufficient to restore normal ATP values. During incubation in the respirometer, the lactate concentration began to increase at a rate of ~ 1.7 µM/mg protein per 15 min. This rate did not change when cells were exposed to acute and 24-h hypoxia (data not shown). Figure 7B shows a slight but statistically significant increase in NADH during acute hypoxia. However, this increase was less than 10% of the increase induced by rotenone, an inhibitor of complex I of mETC.



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  		Figure 7. Effects of acute and 24-h hypoxia on the concentration of (A) ATP and (B) NADH of A549 cells. (A) ATP was measured in cell suspensions exposed to normoxia, hypoxia, and reoxygenation in the respirometer for 15 min at 37°C. Cells exposed to hypoxia for 24 h were washed and then transferred into the respirometer, in which the buffer had already been equilibrated with gas containing 2% O2 and 98% N2. Mean values ± SD (n = 6) are presented. *P < 0.05 relative to normoxic controls. (B) NADH was measured by fluorescence microscopy on A549 cells grown on glass slides mounted in a gas-tight perfusion chamber in the presence of rotenone (2.5 µM). Mean values (n = 5) of changes in fluorescence relative to the fluorescence in normoxic control cells are presented. *Significant difference between normoxia and hypoxia; #significant effect of rotenone (P < 0.05).

 

   DISCUSSION
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
References
 
It was the goal of this study to determine whether the inhibition of lung ion transport by hypoxia is an adequate mechanism for conserving energy during hypoxia, and which other mechanisms might also decrease ATP demand. This inhibition has been observed in native rat lung (8) in primary and SV-40–transformed alveolar type II cells, as well as in A549 cells, a human lung–derived adenocarcinoma cell line with characteristics of alveolar type II cells (19). The major finding of this study is that alveolar epithelial cells in fact decrease their JO2 rapidly upon exposure to hypoxia, which can be interpreted as an adaptive mechanism for conserving energy in a situation of reduced oxygen availability. In A549 cells, this is in part due to a reduced proportion of JO2, attributable to reduced activity of Na/K-ATPase, protein synthesis, and other metabolic pathways that have not been identified in this study. This decrease in JO2 might also be caused by inhibition of mitochondrial respiration. However, inhibitors of ion transport pathways that mediate secondary active Na entry into the cell do not affect cellular JO2.

A possible limitation of this study needs to be addressed: inhibition of ion transport, protein synthesis, and ATP production by hypoxia has been observed in lung tissues of different origin and primary rat alveolar epithelial cells, but also in the A549 tumor-derived cell line. It needs to be pointed out, however, that tumor cells are particularly resistant to hypoxia (for review, see Ref 25). The pattern of adjustments of A549 tumor cells to hypoxia appears similar to that observed in native tissue. It remains to be clarified whether the restriction by hypoxia of metabolic activity and mitochondrial function observed in A549 cells justifies any generalization on specific adaptive mechanisms of alveolar epithelial cell or on lung function in clinical situations that cause alveolar hypoxia. The results on primary rat alveolar type II cells indicating a hypoxia-induced inhibition of JO2,tot may point toward the validity of such conclusions.

Exposure of A549 cells to a PO2 of ~ 10 mm Hg decreases the PO2 of the surrounding medium to ~ 7% of the typical PO2 of normoxic tissue culture conditions, and by almost 90% of a typical alveolar PO2 in normoxia. Yet this PO2 is well above oxygen saturation of enzymes of the mETC, indicating that mitochondrial ATP production should not be limited. Nevertheless, cells initiate mechanisms to decrease ATP demand—mechanisms that are probably aimed at protection from a likely decrease in ATP upon further decrease in PO2. This process has been termed "oxygen conformance" (15). In A549 cells, cellular respiration is inhibited by ~ 20% by acute hypoxia. Simon and colleagues (26) showed that prolonged anoxic exposure of cloned alveolar epithelial cells decreases JO2 by a similar degree to that found in hypoxia in the present study. We show here in cancerous A549 cells and in rat primary alveolar type II cells that adjustments are found at higher levels of oxygen than in native tissue and that they seem to occur within minutes. In tissues of hypoxia-adapted animals, as well as in a variety of cultured cells of different origin, inhibition of respiration is achieved by inhibition of a variety of metabolic pathways (27); active ion pumping is one of them (1, 21).

In normoxia, ~ 15–20% of the total JO2 of A549 cells can be attributed to the activity of Na/K-ATPase. Similar numbers have been reported in other studies (5, 28). At the typically low cellular Na levels, the activity of Na/K-ATPase is not maximal (29), and any increase in cellular Na will therefore increase Na/K-ATPase activity and JO2, indicating a reserve capacity of ATP production. Preliminary results on nystatin-permeabilized A549 cells indicate a 2- to 3-fold increase in JO2,{Delta}ouab when respiration is measured in high-Na medium (data not shown). Hypoxia decreases the activity of the Na/K-ATPase in plasma membrane preparations of lung alveolar epithelial cells (6, 28), ouabain-sensitive 86Rb-uptake (6), and ouabain-sensitive short-circuit current (7) by ~ 20–45%. The ~ 20–30% inhibition of ouabain-sensitive JO2 of A549 cells in suspension upon exposure to hypoxia (Figure 3) is in accordance with the above-mentioned results. Interestingly, JO2 was not affected by inhibitors of other Na transporters in the plasma membrane of A549 cells, indicating that the activity of Na/K-ATPase is the rate-limiting step in transcellular Na –transport, and that uptake of Na into the cell is limited by the transmembrane Na gradient generated by the Na/K-ATPase. This is in accordance with measurements of short-circuit currents on primary rat alveolar epithelial cells, where it has been found that hypoxia inhibited in a coordinated manner both the basolateral Na/K-ATPase and apical amiloride-sensitive Na entry pathways without affecting amiloride-insensitive currents (7).

The question arises whether inhibition of Na pumping is an adequate mechanism to save significant amounts of ATP (and oxygen) in states of limited oxygen supply. Calculations indicate that it is not. Based on a proportion of 20% of all oxygen consumed to supply the ATP for nonstimulated Na pumping in normoxia, it can be calculated that cellular JO2 can be reduced, at most, by ~ 4% when the activity of the Na/K pump is reduced by 20%, as found in this study. Therefore, additional ATP-consuming pathways have to be inhibited as well to reduce cellular oxygen demand to a degree that matches the decreased oxygen supply. Such pathways include Ca-ATPases and other ion pumps in the plasma membrane and in the membranes of intracellular organelles, phosphorylation processes, and protein synthesis. The lack of dramatic changes in cellular ATP concentrations indicates that this pathway inhibition might, in fact, be the case.

Mitochondria
Whereas the above described mechanisms imply that adjustment of cells to hypoxia occurs through inhibition of ATP-consuming processes, there is also the possibility that cellular functions, such as Na pumping, are restricted by a diminished ATP supply. Vinogradov (30) found that complex I of the mETC is shifted into an inactive form during hypoxia, and that ubiquinol levels increase. It was also found that inactivation was reversible after short-term but not after long-term ischemia (30). Our results (Figure 4A) are consistent with this finding, because they also show a hypoxia-induced inhibition of pyruvate-dependent respiration of digitonin-permeabilized A549 cells and a lack of reversibility after 24 h of hypoxia. Similar results have also been obtained in cardiomyocytes and in renal tubules (31, 32).

Results on hypoxia effects on complexes II and III are divergent. We found activity of succinate-stimulated respiration of permeabilized A549 cells decreased by 15–30%. However, we have no explanation as to why the inhibition of mETC complex II respiration was less pronounced after 24 h than after acute hypoxia. In contrast, no effects of ischemia on mitochondrial respiration of myocardial cells (33) or on renal tubules (32) have been found. With regard to mETC complex III, however, our data agree well with those of Rouslin (33), who also found pronounced inhibition and lack of reversibility of mETC complex III activity in myocytes from ischemic hearts.

The mechanisms that cause dysfunction of mitochondria in hypoxia are not clear. We found that the activity of the enzyme citrate synthase was decreased by ~ 20% in cells that were exposed to hypoxia for 24 h. This enzyme has been used as an indicator of the number of mitochondria (22). It is also possible that hypoxia-mediated alterations in NO affect mitochondrial function (34). Because the activities of mETC shown in Figure 4 are normalized to whole-cell protein, this result indicates that, after 24 h of hypoxia, the mETC activity might actually be decreased by more than 60% due to a parallel decrease in both the number of mitochondria and mETC activity.

The decrease in citrate synthase and the capacity of mETC enzymes might be caused by inhibition of protein synthesis—the predominant mechanism of energy conservation in many hibernators and hypoxia-resistant species (1, 35). Control of protein synthesis in hypoxia is complex. The expression of proteins, which are important for glycolytic ATP production and for oxygen transport to tissues, are typically upregulated by HIF-1{alpha}–dependent processes (11). A549 cells follow this pattern, because they also show elevated levels of nuclear HIF-1{alpha} and increased expression of glyceraldehyde phosphate dehydrogenase (data not shown). The expression of many more proteins, such as Na transporters (5, 36), is suppressed, which contributes to the decreased rate of JO2.

It is unclear whether this decrease in mETC capacity is of significance for cellular ATP supply. The slight decrease in NADH, which amounts to ~ 10% of the NADH increase achieved by complete inhibition of mETC complex I with rotenone, might be taken as evidence for a reduced level of mETC activity. However, the lack of increase in lactate production and the lack of pronounced decrease in ATP in hypoxia-exposed A549 cells indicates that the mETC still supplies sufficient ATP, at least in this state, in which the activity of ATP-consuming processes, such as Na/K-ATPase and protein synthesis, are also decreased. This parallel decrease in ATP consumption and ATP production points to coordinated mechanisms that balance both processes—although the triggering factors are not known—and implies the question of cause and effect: is the inactivation of ATP-consuming pathways the reason for the decrease in mitochondrial activity, or does the latter decrease because of limited oxygen supply, which in turn reduces activity and expression of enzymes that catalyze ATP-consuming processes? Mitochondria are thought to play a role in oxygen sensing (37, 38), in which cytochrome oxidase plays an important role (39). In hypoxia, the maximal activity of this enzyme decreases without a change in substrate affinity, probably by direct action of molecular oxygen, which does not affect cellular JO2, but affects the redox state of the cell (39).

Oxygen radicals have been discussed as possible hypoxia signals (38), although they are also known to damage lipids and proteins (40), such as in reoxygenation injury (41). The reduced metabolic activity and/or decreased activity of ATP-consuming processes could therefore also be due to damage by oxygen radicals. However, the nearly complete reversibility of short-term hypoxia effects argues against this hypothesis and points to a strategy of defense against damage subsequent to restricted oxygen supply.


   Acknowledgments
 
The authors thank Mrs. Sonja Engelhardt, Mrs. Martina Haselmayr, and Mrs. Christiane Herth for excellent technical assistance.


   Footnotes
 
Supported by Deutsche Forschungsgemeinschaft grant Ma 1503/14-1 (to H.M.).

Conflict of Interest Statement: K.H. has no declared conflicts of interest; A.S. has no declared conflicts of interest; L.H. has no declared conflicts of interest; P.B. has no declared conflicts of interest; and H.M. has no declared conflicts of interest.

Received in original form June 23, 2004

Received in final form September 8, 2004


   References
Top
 Abstract
 Introduction
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 References
 

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